International Review of Cytology: A Survey of Cell Biology, Volume 113

INTERNATIONAL

REVIEW OF CYTOLOGY

VOLUME113

ADVISORY EDITORS H. W. BEAMS HOWARD A. BERN DEAN BOK GARY G. BORISY PIET BORST BHARAT B. CHATTOO

KEITH E. MOSTOV AUDREY MUGGLETON-HARRIS DONALD G. MURPHY ANDREAS OKSCHE MURIEL J. ORD VLADIMIR R. PANTIC STANLEY COHEN W. J. PEACOCK RENE COUTEAUX DARRYL C. REANNEY MARIE A. DIBERARDINO LIONEL I. REBHUN BERNDT EHRNGER JEAN-PAUL REVEL CHARLES J. FLICKINGER L. EVANS ROTH NICHOLAS GILLHAM JOAN SMITH-SONNEBORN M. NELLY GOLARZ DE BOURNE WILFRED STEIN YUKIO HIRAMOTO RALPH M. STEINMAN YUKINORI HIROTA HEWSON SWIFT MARK HOGARTH K. TANAKA K. KUROSUMI DENNIS L. TAYLOR ARNOLD MITTELMAN TADASHI UTAKOJI ALEXANDER YUDIN

INTERNATIONAL

Review of Cytology A SURVEY OF CELLBIOLOGY

Editor-in-Chief

G. H. BOURNE (Deceased)

Editors

K. W. JEON

M. FRIEDLANDER

Department of Zoology University of Tennessee Knoxville, Tennessee

UCLA School of Medicine

Jules Stein Eye Institute Los Angeles, Galflornia

VOLUME113

ACADEMIC PRESS, INC. Harcourt Brace Jownovich, Publishers

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0 1988 BY ACADEMICPRESS, INC.

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9 8 7 6 5 4 3 2 1

Contents Micromorphology and Structure Research: Application of Principles Valid a Priori RAINERH . LANCE(REVISED BY KEVINLEONARD) I . Introduction

..........................................................

I1. The Dissymmetric Biomacromolecule .....................................

I11. Symmetric Arrays ..................................................... IV Application of Structure Principles ...................................... V. Concluding Remarks ................................................... References ............................................................

.

1 2 4 14 29 32

Functional Inclusions in Prokaryotic Cells

. .

. .

J . M . SHIVELY. D. A . BRYANT.R C FULLER.A E KONOPKA. S . E. STEVENS. JR., AND W . R . STROHL

I. I1. I11. IV. V.

Introduction .......................................................... Inclusions as Metabolic Machinery ....................................... Inclusions as Adjusters of the Environment ............................... Inclusions as Metabolic Products (Reserves) ............................... Concluding Remarks ................................................... References ............................................................

35 36 59 67 87 88

Microtubules in Cardiac Myocytes L. RAPPAPORT AND J . L . SAMUEL

I . Introduction .......................................................... I1. Proteins Constitutive of Cardiac Microtubules .............................

111. Distribution of Microtubules in Cardiac Muscle ........................... IV. Roles of Microtubules in the Cardiac Myocyte ............................ V. Conclusion ........................................................... References ............................................................ V

101 102 108 123 136 139

vi

CONTENTS

Functional Morphology of the Thyroid HISAOFUJITA

I. I1. 111. IV. V VI . VII. VIII . IX X.

.

.

Introduction .......................................................... Synthesis and Release of Thyroglobulin .................................. Iodination of Thyroglobulin ............................................ Reabsorption of Colloid ................................................ Hydrolysis of Thyroglobulin and Release of T. and T. ..................... Connective Tissue Space and Vascularization .............................. Nerve Supplies ........................................................ Follicular Cell Polarity and Inverted Follicles .............................. Why Does the Thyroid Need Follicle Structures? .......................... Concluding Remarks ................................................... References ............................................................

145 147 152 155 163 166 171 172 179 180 181

Bacterial Surface Polysaccharides: Structure and Function IANW. SUTHERLAND

I. What Are They? Introduction and Definition ............................. I1 Appearance-Light Microscopy and Transmission and Scanning Electron Microscopy ....................................... I11 Physiological Influences ................................................ IV Chemical Structures .................................................... V Physical Properties .................................................... VI . Function ............................................................. VII . Conclusions ........................................................... References ............................................................

. . . .

187 189 194 197 209 219 226 226

Reorganization of the Egg Surface at Fertilization FRANK J . LONGO I. I1. 111 IV. V. VI . VII . VIII .

.

Introduction .......................................................... Egg Cortical Structure ................................................. Interaction and Fusion of Sperm and Eig ................................ Cortical Granule Reaction .............................................. Plasma Membrane Changes Attending the Cortical Granule Reaction ........ Microvillar Elongation ................................................. Endocytosis ........................................................... Concluding Remarks ................................................... References ............................................................

233 233 239 250 252 257 260 261 263

CONTENTS

vii

Ultrastructural Modifications and Biochemical Changes during Senescence of Chloroplasts

u . c. BISWALAND BASANTIBISWAL I . Introduction .......................................................... I1. Senescence-Induced Structural Modifications .............................. 111. Loss of Primary Photochemical Reactions ................................ IV. Loss of RuDPCase Activity ............................................. V. Senescence of Cell-Free Chloroplasts ..................................... VI . Regulation of Chloroplast Senescence .................................... VII . Conclusion ........................................................... References ............................................................

271 273 285 291 297 304 311 316

INDEX ......................................................................

323

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INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 113

Micromorphology and Structure Research: Application of Principles Valid a Priori RAINERH.

LANGE*”

(REVISEDB Y KEVINLEONARD?)

*Institute for Anatomy and Cell Biology, University of Giessen, Giessen, Federal Republic of Germany, ?European Molecular Biology Laboratory, 6900 Heidelberg, Federal Republic of Germany

I. Introduction The term micromorphology as used here signifies the reproducible and often aesthetically pleasing aspect (Fawcett, 1964) of biostructure at a level at which macromolecules have completely lost their individuality, with very few exceptions (e.g., glycogen; Revel, 1964). Micromorphology is, therefore, superficial to the structural level, as is always the case with levels of low resolution with respect to levels of better resolution. With the establishment of discrete macromolecular models for micromorphoIogical elements-one of the first being the lipid bilayer membrane (Danielli, 1936)-an evolution (and more recently a revolution) began that caused us to look at biostructures in a manner quite different than before. The present article aims at illustrating relationships between micromorphology and the structural level or, more precisely, the implications for micromorphology of principles governing symmetric aggregates. These principles possess a priori validity, at least from the standpoint of the microscopist. We shall particularly be concerned with studies on biological specimens in their natural context, i.e., with electron microscopy. Here, then, the very complex and delicate structure of such specimens presents extreme difficulties (Beer el nl., 1975), resulting in a large gap between the resolution accessible by instrumental power and that related to specimen organization. This gap is difficult to bridge, but in the case of symmetric aggregates it can provisionally be filled by the application of a priori knowledge: the consequences of a symmetry concept are very specific and serve to promote the investigation to prove or disprove the concept, thereby providing a control mechanism of its own. The severe limitation of the approach is that strict structural symmetry can be spoken of only when the analysis has gone to the atomic level I

Deceased August 4, 1984. I Copyright 8 1988 by Academic Press, Im. All rights of reproduction in any form regerved.

2

RAINER H. LANGE

(Fig. 1). This dependence on resolution has the result that, using morphological methods, we can prove only apparent symmetry. Any additional data (biochemical, X-ray diffraction, spectroscopic) are, therefore, important for as correct as possible an interpretation of the system under study. As long as this limitation is kept in mind, the approach remains meaningful and powerful. Facing the limited regard given in micromorphological studies to this well-known and truly morphological discipline, it appears worthwhile to illustrate the application of such principles to a number of biological specimens. Although this era of powerful computer application (Frank et ul., 1981) might seem to diminish their relevance, emphasis will lie on understanding principles-and thus on explanation-rather than on technical procedures. 11. The Dissymmetric Biomacromolecule

In structure research we are concerned with assemblies of biomacromolecules, especially protein and nucleic acid molecules. The most important feature of such molecules is their dissymmetry (Barry and Barry, 1969), and this property is a fundamental principle of living systems (Bernal, 1966). In importance, this principle comes up to those formerly defined as principles of living systems as a whole, e.g., metab-

FIG. 1. Mirror pseudosymmetry of the Bilateralia at the macrolevel. The molecules are the same (right) and not enantiomers in the two body halves (left). Symmetry labels depend on the level of resolution.

MICROMORPHOLOGY AND STRUCTURE RESEARCH

3

o h m , susceptibility to external stimulation, and change of form (Hartmann, 1953). A dissymmetric (or chiral) molecule is not congruent with its mirror image. Thus, for a dissymmetric molecule, a mirror image can be constructed by reversing the chirality of the molecule. However, for biomacromolecules only one of these two so-called enantiomers exists in living systems. (Structures built of such dissymmetric monomers are called enantiomorphic.) This fact implies three important principles of a priori validity: 1. Structural polarity is a genuine property of most supramolecular structures. 2. Biostructures cannot have mirror symmetry. 3. A structure having n-fold rotational symmetry must consist of n or n-m macromolecules, rn and y1 being integers.

The first principle is best illustrated with elongate structures (e.g., filaments) and means that such a filament when reversed in its course is not the same as before (Fig. 2), although micromorphology may not allow a distinction to be made between the two orientations. The only exception to this is the presence of a 2-fold axis of rotation perpendicular to the structure axis, which occurs very rarely. From the second principle the well-known fact follows that the macroscopic mirror symmetry of the Bilateralia is, indeed, only an apparent symmetry (Fig. 1). We dwell upon this trivial notion because it demonstrates so clearly the above mentioned fact that our symmetry labels depend on the resolution obtained. The third principle enables the estimation of (minimum) numbers of macromolecules building up a given structure (see Section IV, E). The asymmetry of macromolecules renders shape determinations of isolated molecules by electron microscopy very difficult because image

Fie. 2. Structural polarity of a biomolecular structure illustrated in an example of helical symmetry s. Although not apparent at the common level of resolution, there is no congruency of the two structures unless one is rotated by 180" about an axis perpendicular to the paper plane.

4

M I N E R H. LANGE

noise and destructive effects such as electron beam damage combine to hide every detail, except for coarse features (Beer et al., 197.5). This is often taken into account by depicting such unknown shapes using arbitrary fanciful shapes (Fig. 3 ) that are later refined step by step with progress in structural analysis. The situation is quite different when identical particles form an array with degrees of symmetry. In such a case with structural redundancy, methods of image detail enhancement are easy to apply to refining the image of a single particle either by averaging the image directly, i.e., in real space (Horne and Markham, 1973), or in reciprocal space by optical filtering of the diffraction pattern (Klug and DeRosier, 1966); the latter method is now most often performed with digital computers. Images of isolated particles have likewise been refined by mounting them in a symmetric array and subsequently subjecting this array to optical filtering (Ottensmeyer et al., 1977) and computer digital solutions of this correlation problem have found application (Frank et a / . , 1981).

111. Symmetric Arrays

A.

SYMMETRY

ELEMENTS

A symmetric array made of one particle species is described by giving the geometrical operations (or symmetry elements) used in its construction. Such operations transform a given structure into itself. For example, a single asymmetric particle has 1-fold rotational symmetry (symbol l), thus it is transformed into itself by rotation for 360"/1 or an array of OUTSIDE

INSIDE FIG 3 Representation of individual macromolecules using deliberate fanciful shapes to express their largely unknown structural features on the one hand, and their polarity on the other hand. The drawing shows the erythrocyte membrane. From Steck (1974). Courtesy of the author and The Rockefeller University Press. New York.

MICROMORPHOLOGY AND STRUCTURE RESEARCH

5

asymmetric particles with n-fold rotational symmetry (symbol n) is transformed into itself by rotation about the rotation axis for 360"ln. Symmetry elements are defined in textbooks of crystallography. The following list comprises only those elements occurring in arrays of (dissymmetric) biomacromolecules:

1. Translation (Fig. 4a-c). This is the (vectorial) displacement along a straight line by a given length, the period. Translation is not present in all symmetric arrays but only in those having, in principle, infinite length in one, two, or three (generally not orthogonal) directions of space. Those arrays possessing translation(s) are characterized by a point lattice, which is one-, two-, or three-dimensional according to the number of primary translation vectors (Fig. 4a-c). Each lattice point stands for the discrete

t t

a

C

-ah , -bk , -2cl

=

[112]

d

FIG. 4. One- (a), two- (b), and three-dimensional (c) point lattices and definition of crystallographic screw axis operation (d). In a-c the primary translation vectors are set off by heavy lines; dashed lines in b indicate a different but equivalent choice of the unit cell, and lattice centering is illustrated in the top right-hand unit cell; in c, angles and faces of the unit cell in the standard setting and derivation of the indices hkl of a direction in the crystal are illustrated. A crystallographic screw axis operation (d) is an inseparable combination of rotation and translation, defined in this drawing for a screw axis 4, along unit-cell side a.

6

RAINER H . LANGE

matter complex upon which translation acts and may be conceived of as the center of gravity of this mass. The symmetry of a point is infinite and so may be thought of as an atom; however, biomacromolecules, and complexes thereof, always possess very limited symmetry. 2. Rotational symmetry (Fig. 5 ) . This is characterized by a rotation axis (with unique position in the case n # 1) and rotation for 360"ln (n-fold rotational symmetry; symbol n). If n is not a prime number (Lea, n = nl'n2*n3 . . .), n implies also the lower symmetries n1,n2,n3. . . . 3 . Screw symmetry (Fig. 4d). This is an inseparable combination of rotation and translation. The screw axis is identical with the rotation axis and the direction of translation.

0'

90'

180'

270'

36C

hQAAA%

FIG. 5. Poin group symmetries. The symmetry symbol is given in the black boxes. (a, b) A pair of enantiomers, only one of which is available in biostructures. A cone serves as a model and is shown as seen from different directions. (c-f) Point groups as occurring with dissymmetric particles. (g, h) Point groups that comprise a mirror plane and, therefore, cannot be formed by one species of dissymmetric particles. Whereas the cone from (a) is exclusively used in c-f, both cones (a, b) are needed for the construction of the point groups in g and h.

MICROMORPHOLOGY AND STRUCTURE RESEARCH

7

Symmetric aggregates have one or several of the above symmetry elements. Symmetry elements can be additively combined without contradiction only according to certain rules, and these possible combinations form the second body of principles with a priori validity. By acting upon the aggregated particles, symmetry elements also act, of course, on the other symmetry elements present in the array. A rotation axis not doing such but having only limited influence is not a proper symmetry element of the structure and is called a local axis. The reference made to crystallography may, however, lead us to think along the wrong lines since crystallographic rotation and screw axes are limited to special cases, whereas in noncrystallographic cases they can have all degrees of freedom not forbidden by the symmetry definition. Thus, for crystallographic rotation axes, II can only be 1, 2, 3, 4, or 6, whereas noncrystallographic rotation axes can be any number (e.g., 5 , 7 , or 13). Curiously, the crystallographic numbers seem to have influenced authors of structural schemes (e.g., the 12-protofilament model of the microtubule; Krstic, 1979). B. RIGIDSYMMETRY VERSUS FREEDOM OF AGGREGATION In the above sections we have stressed symmetry and shall have to do so in the following sections. However, upon increasing resolution it may come to light that supramolecular structures seen as being symmetric aggregates of one monomer species have lower symmetry by either containing identical particles at not strictly equivalent sites or by being composed of monomeric species not truly identical. The former case has been discussed in detail by Caspar and Klug (1962) and the latter is illustrated by terms such as heterodimer and the a-p-tubulin complex (for a review, see Dustin, 1978) and heterotetramer for isozymes of lactate dehydrogenase (Markert, 1963). Such deviations from strict symmetry illustrate the immense freedom realized in nature, which goes beyond limits of traditional geometrical concepts (Bernal, 1966), and render the description of structures very complicated. It is, therefore, useful to speak of quasi-symmetry and quasi-equivalence of position, meaning that the symmetry label applies only in a qualified sense; Matthews and Bernhard (1973) use the term pseudoisologous association in this context. Pseudosymmetry, a condition also related to our approach, has a very different meaning; it denotes the lack of correspondence between point lattice symmetry and space group symmetry (see Section II1,C ,3).

8

RAINER H . LANGE

C. SYMMETRY GROUPS Natural symmetric aggregates are classified as having point symmetry, line symmetry, or space symmetry (Caspar, 1966; Klug, 1969). 1 . Point Groups A point symmetry group is characterized by one or more rotational symmetries, all rotation axes intersecting in one point. There are severe restrictions in the combination of several rotation axes; screw axes and translations cannot occur in point symmetry goups. Point symmetry groups describe discrete arrangements of matter such as protein oligomers and virus capsids. As a consequence of the restricted possibilities in the combination of rotation axes alluded to above, only three kinds of point groups occur: cyclic, dihedral, and cubic (see Crick and Watson, 1956; Klug, 1969; Klotz et al., 1970; and textbooks of crystallography). Cyclic symmetry (Figs. 5c and 8b. f, and g) is defined by one n-fold rotation axis (symbol n). Dihedral symmetry (Figs. 5e and f and 8d) has one ( n odd) or two ( n even) sets of 2-fold rotation axes (or rotation diads) (symbols: n2, n22; reads “n-two-two”) Perpendicular to the n-axis (as in cyclic groups). The second set of rotation diads follows necessarily from n even and the first diad. If n is odd (Fig. 5f), the diads perpendicular to the n axis are polar because they are not also present in the reversed sense. The dihedral point group 222 signifies a tetramer and has wrongly been likened to a tetrameric tetrahedron (Klotz et al., 1970), which would represent a cubic point group (see below) possessing, by definition, 4 3-fold rotation axes (rotation triads). Therefore, although a tetramer of dissymmetric molecules cannot be tetrahedral, it can display tetrahedral pseudosymmetry (see Section 111,B). Cubic point groups are combinations of rotation axes including more than one axis different from a diad; they always contain four rotation triads in the directions of the cube diagonals and, in addition, 5,4,and/or 2 axes. In their symbols (tetrahedral: 23; octahedral: 432; icosahedral: 235 or 532; Fig. 6) a fixed sequence of these axes is observed (as in space group symbols): the first position gives the symmetry along the edges of a cube, the second position that along the cube diagonals, and the third (except for the icosahedral case) that along the face diagonals of a cube; the symbols accordingly read, e.g., “four-three-two” (octahedral). Cubic point group symmetries if realized by dissymmetric molecules have the following number of particles occupying equivalent positons: 12 in the tetrahedral (see above), 24 in the octahedral, and 60 in the icosahedral case, an interesting illustration of the numerical considerations made above (see Section 11).

MICROMORPHOLOGY AND STRUCTURE RESEARCH

FIG. 6. The icosahedral point group 235. (a) Stereographic projection (see geographic atlases). The numbers 2 , 3 , and 5 mark the intersection with the sphere surface of 2-, 3-, and 5-fold rotation axes, respectively. The orthogonal 2 axes of the cubic point lattice are marked by brackets, the triads falling on cube diagonals, by triangles. (b) lcosahedral surface symmetry in three examples set correspondingly; left, icosahedron; middle, ball; right, pentagonal dodecahedron. In the latter, the orthogonal cube axes are represented as rods. From Lange and Blodorn (1981). Courtesy of Thieme Verlag, Stuttgart.

2. Line Groups A line symmetry group is characterized by a general screw axis (Fig. 4d), which may in special cases be combined with an n-fold rotation axis along it and a 2-fold rotation axis perpendicular to it (Fig. 8c, e, and h), and leads to a structure of principally infinite length: helical symmetry is a line symmetry and microtubules, filamentous structures, and certain virus capsids fall into this group. Among the four line groups shown by Klug et al. (1958) to apply to dissymmetric molecules (s, s2, sr, sr2; see Fig. S), the primitive helix (symbol s) is the most important one in micromorphology .

3 . Space Groups Space groups are characterized by translation vector(s) (Fig. 4a-c) (in combination with other elements of symmetry) and describe structures having principally infinite extension in one, two, or three dimensions. Accordingly, they are classified as one-, two-, and three-dimensional space groups in the very important reference book, “International Tables for X-Ray Crystallography,” Vol. 1 (Henry and Lonsdale, 1969). The symmetry elements of space groups other than the translation vectors describe the symmetry of the matter complex (unit cell) repeating in the point lattice formed by the translation vectors and, at the same time, that

10

RAINER H. LANGE

of the whole arrangement. Referring to the above definition of a crystal in the strict sense (Section III,A), we can now make it precise by stating that thcrc is a corrcspondcncc between space group symmetry and symmetry of the point lattice. In order to provide a fuller understanding of (two- and) three-dimensional space groups at this point, two concepts must be introduced here: (1) the relationship between point groups and space groups, and ( 2 ) lattice centering (Fig. 7). First, space groups result from those point groups having the following symmetry elements: 1-, 2 - , 3-, 4-, and 6-fold rotation axes; furthermore, the rotation axes mentioned can be modified in space symmetry to screw axes (Fig. 4d), e.g., 2 to 2 1 , 3 to 3 , 3 ? , etc. Second, the point lattices defining vectorial translation in two- and three-dimensional space groups may be either primitive (symbol p and P for two- and three-dimensional lattices, respectively) or may possess various degrees of centering. Thus, two-dimensional point lattices may have centering of their basic rectangular lattice elements (symbol c), and three-dimensional point lattices may have the following kinds of centering: body centering (one additional lattice point in the center of the elementary parallelepiped of the lattice; symbol: I ) , centering in one face of the parallelepiped (one additional lattice point in the center of this face; symbol C or other face), or centering in all faces (additional lattice point in the center of all three faces; symbol: F). Let us consider consequences and applications of the above generalities. The cubic point groups 23 and 432 (see Section III,C,l) occur as three-dimensional space groups, e.g., 23 as P 2 3 , F 2 3 , 123, P 2 1 3 , I 2 , 3 , whereas the icosahedral point group 235 cannot (because it possesses a 5 axis). Whereas in the primitive lattice ( P ,p ) there is one lattice point per unit cell, I - , C-, and c-lattice have two and the F-lattice has four lattice points in one unit cell. One-dimensional space groups (according to “International Tables for X-Ray Crystallography”) are very simple aggregates, possibly too simple for the complex functions and regulations of biostructures. If the string of mRNA was neglected, ribosomes forming polysomes could be thought of as being arranged in a one-dimensional space group. One-dimensional space groups have a relationship with line groups. Two-dimensional space groups describe the symmetry of planar aggregates, and they also have an important role in the description of projected views of three-dimensional periodic aggregates. As described in “International Tables for X-Ray Crystallography,” two-dimensional space groups have symmetry elements perpendicular only to the reference plane (rotation axes, mirror and glide planes; the latter two can in biological specimens apply only to projected views of three-dimensional periodic aggregates). However, description of “planar” aggregates of biomacro-

MICROMORPHOLOGY AND STRUCTURE RESEARCH

11

molecules requires the use of rotation and screw axes lying in the reference plane and an extension to “two-sided” planes. For this purpose, a listing like that given by Holser (1958; plane groups) is important. The plane group symbol has four positions: the first position refers to centering ( p or c ) , the second position gives the symmetry of rotation (axis perpendicular to the reference plane), the third the symmetry element coinciding with the first elementary vector, and the fourth that coinciding with the second elementary vector of the two-dimensional point lattice (Fig. 4b). Symbols are abridged if unequivocal; the full notation frequently contains symmetry 1 for formal reasons. Examples of such two-dimensional space groups are macromolecular aggregates associated with membranes or produced in uitro, forming so-called twodimensional crystals (2D-crystals) (see Baumeister and Vogell, 1980). Three-dimensional space groups characterize three-dimensional crystals of proteins (Blundell and Johnson, 1976), as well as those of lipids (Hauser et al., 1980) and proteoglycans (Winter el al., 1978). These examples represent single crystals grown in uitro for tertiary structure determination by X-ray crystallography. Do three-dimensional crystals that occur in uiuo, e.g., in secretion granules of polypeptide-secreting cells (Lange, 1981a) as well as other sites, play a biologically active role? We think we have observed such crystals in oocytes where they show a high evolutionary conservation and thus must have physiological relevance (anamniote yolk-platelet crystals; see Section IV,E). From the generalities above, the 14 Bravais lattices demonstrated in Fig. 7 follow. The key problem for somebody inexperienced in crystallography is to understand that although a crystal is a three-dimensional periodic aggregate, it is also a symmetric aggregate. The translational periodicity is defined by one of the 14 point lattices, and the symmetry by one of the 65 three-dimensional space groups (there are 230 threedimensional space groups in total, but only the 65 enantiomorphic space groups apply to biomacromolecules because of the dissymmetry of the latter; due to this restriction, crystallography of biomacromolecules is greatly simplified). Point lattice symmetry and space group symmetry correspond, the former being duplicated only by space groups of the highest symmetry, which can never. be obtained in biomacromolecular crystals. This very important property of crystals will be illustrated now for a number of cases (Fig. 7). The general triclinic point lattice has symmetry 1 (which means symmetry 1 and center of inversion = reflection at a point). It is made by nonorthogonal elementary vectors all of different lengths. It is the only lattice that can be realized by one dissymetric molecule per lattice point

RAINER H . LANGE

12

P

C

I

F

monoclinic

tetragonal

trigonal

R

hexagonal

cubic F I G .7. The 14 crystallographic point lattices having three dimensions. One unit cell is shown. only angles differing from 90"are indicated, and equal unit-cell sides have the same label (this does not fully apply to the trigonal-hexagonal group including the rhombohedra1 cell, R). See text. From Lange and Blodom (1981). Courtesy of Thieme Verlag, Stuttgart.

(or per unit cell); all other lattices have higher symmetry. Let us consider the case that arises when the interaxial angles become 90" and the unit-cell sides become equal; the lattice would then be a cubic P-lattice, however, if it is made up of one dissymmetric molecule per lattice point it is triclinic by symmetry. In this case the triclinic crystal has cubic pseudosymmetry (see Section II1,B).

MICROMORPHOLOGY AND STRUCTURE RESEARCH

13

The monoclinic point lattice has twofold rotational symmetry and a mirror plane perpendicular to it: 2lm. The standard setting of the cell is with the 2 axis along b , hence p # 90" = a = y , all unit-cell sides principally differing from one another. In the monoclinic P-lattice there are only two enantiomorphic space groups, P2 and P 2 1 ,with a minimum of two dissymmetric molecules per cell to realize monoclinic symmetry. The C-lattice brings in one more space group, C2, with a minimum number of four monomers per unit cell. The orthorhombic point lattices ( P , C , I, F ) are orthogonal with a # b # c # u and have 2-fold rotational axes coinciding with the unit-cell sides and mirror planes perpendicular to them: 21m 21m 21m or simply mmm. This high symmetry (mirror planes!) cannot be attained by dissymmetric molecules, but the dihedral point group 222 is the basis for all enantiomorphic orthorhombic space groups. If we consider only primitive orthorhombic lattices, there are four space groups for dissymmetric molecules, P222, P2221,P21212,and P 2 , 2 1 2 with ~ a minimum of four monomers per unit cell. The first position following P indicates the symmetry along a , the second along b, and the third along c. Of course, P2,22, P22,2, and P222, are only different notations (and settings of the unit cell) with principal identity because there is no unique axis. With the tetragonal point lattice a unique 4-fold rotational axis is introduced by two of the three orthogonal axes being equal (point lattice symmetry 4lmmm). With the trigonal, rhombohedra], and hexagonal point lattices true 3-fold or 6-fold rotation axes, which cannot occur in the aforementioned lattices, appear for the first time. If we confine ourselves to the hexagonal point lattice, there is the unique 6-fold axis perpendicular to two equal unit-cell sides forming an angle of 60" (point lattice symmetry 6lmmm). The unique axis has the first position in the symmetry symbol of space groups. The cubic point lattice (orthogonal with identical unit-cell sides) is that of the highest symmetry: The positions in the symmetry notation have been explained (Section III,C,l). The cubic point lattice has the symmetry 4lm 3 2lm or, abbreviated, m3m. Again, this high symmetry cannot be realized by dissymmetric molecules, and the monomer numbers required to realize cubic symmetries are considerable. The enantiomorphic cubic space groups arise from the cubic point groups 23 and 432 (see Section III,C,l). Space group P23 requires a minimum number of 12, space group P432 of 24,1432 of 48, and F432 of 96(!)dissymmetric molecules per unit cell. The characteristic symmetry elements of the cubic system are the four triads along the cube diagonals. In considering the symmetry of three-dimensional space groups, we

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RAINER H. LANGE

have seen that a large body of geometrical rules of a priori validity exists from which practically important conclusions follow: that crystallographic labels are primarily symmetry labels and-as it were-only secondarily lattice data (see pseudosymmetry above), that the dissymmetry of biomacromolecules greatly limits the symmetry realizable in aggregates of such molecules, and that, in contrast, symmetric aggregates of biomacromolecules force considerable minimum numbers of such molecules being present. We shall have to apply these principles in Section IV.

D. (SELF-)ASSEMBLY OF SYMMETRIC AGGREGATES Symmetry in biological structures is closely linked to assembly mechanisms, which are beyond the scope of the present article. If symmetric structures arose by self-assembly without the intervention of other molecules, the final product would be fully determined by the tertiary structure of the assembling monomer and assembly conditions. Selfassembly is a very interesting concept, although it is hardly a proper description of reality. Despite this shortcoming, it helps considerably in understanding the construction of symmetric aggregates. Green (1972) worked out a system of closed structures on the basis of asymmetric monomers and two types of bonding, symmetric and asymmetric (isologous and heterologous according to Monod’s terminology), between then (Fig. 8). This system comprises the same point and line symmetry groups as are arrived at when starting from the dissymmetric macromolecule and the symmetry elements (rotational, screw symmetry) possible in aggregates of such molecules (Section 111,A). The kinds of intermolecular contacts considered in Fig. 8 have been felt to be incomplete and an “axial” connection has been added (Matthews and Bernhard, 1973). Self-assembly would severely restrict regulatory events so abundant in nature. In fact, the assembly of symmetric structures has turned out to involve intervention of other molecules at several levels of the biogenetic process (Anderson, 1980).

IV. Application of Structure Principles OF T H E PROJECTED VIEW A. SYMMETRY

The primary result of thin-section electron microscopy is a projected view of the three-dimensional section contents; in the case of freezefracture replicas (Severs and Warren, 1978) it is the (distorted) projection

MICROMORPHOLOGY AND STRUCTURE RESEARCH

15

Two-dimemmal

Q

n2, n22

FIG. 8. A system of structures made of dissymmetric monomers according to Greene (1972). The packing unit has a rectangular triangle as a basis and one such face of black color. Symmetry symbols are given in bold type. S and A denote symmetric and asymmetric bonding according to Green (1972). a, monomer; b, symmetric dimer; c , antiparallel double helix; d, dihedral ring; e, helix; f, ring; g, trimer; h, parallel helices.

of a more or less two-dimensional face. In projected views of threedimensional structures, new symmetry elements occur that are not present in the structures themselves: mirror plane and glide plane. An image contains a mirror plane when the line of view is perpendicular to a 2-fold rotation axis in the specimen, and a glide plane when it is perpendicular to a 2-fold screw axis in the specimen (Fig. 17). These planes are oriented at right angles to the image plane and intersect it at the projection of the specimen 2-fold axis. A glide plane is an inseparable combination of mirror reflection and translation for one period along the

16

RAlNER H. LANGE

reflecting plane. See Fig. 5 for an illustration of rotation and mirror symmetry.

B. MICROSCOPY o k SYMMETRIC AGGREGATES A symmetric aggregate reveals its symmetry by the scattering interaction with electromagnetic radiation. The specimen in its natural state (alive. hydrated, unstained) can be studied by light, X-ray, and spectro5copic methods. In micromorphology, however, structure and topographic connections of tiny subcellular formations are dealt with and there is no or only limited access to this organizational level using the above probes. Hence, electron microscopy is the primary tool. For this purpose, the biological specimen in situ normally requires fixation, dehydration, embedding, thin sectioning, and contrast enhancement by heavy metal staining with few alternative techniques (e.g., cryoelectron microscopy, freeze fracturing, negative staining of disrupted specimens) being available. And yet, there has been considerable progress in techniques related to thin sectioning and this approach has not lost importance (Sjdstrand and Barajas, 1968; Pease and Peterson, 1972; Carlemalm e f al., 1980). What eventually limits electron microscopy is, first, the gap between specimen resolution obtained and the atomic level and. second, the fact that the specimen has been altered to a variable extent during preparation. With respect to the latter restriction, we hold that symmetry at the macromolecular level is not necessarily unrepresented in electron microscopic images. Support for this assumption comes from a number of studies in which additional X-ray data have been used for comparison (e.g., Labaw and Rossrnann, 1969; Lange er al., 1979; Langer et al., 1975; Longley, 1967; Ohlendorf et al., 1975). In our model study on insulin crystals we observed that a number of reflections in the electron diffraction patterns changed their intensity in a reproducible manner depending on the "electron stain" applied (Lange et al., 1979). In studies on symmetric aggregates the first goal is the determination of the intrinsic coordinate system, i.e., of the point lattice present (Fig. 4a-c). For this purpose electron and optical diffraction are used. Diffraction is an interference phenomenon that leads to a specimen-dependent redistribution of radiation such that specific amplitude and phase relations with respect to the primary beam can be attributed to segments of space behind the specimen. In diffraction photographs this is expressed by the distribution of the squared amplitudes. i.e., intensities. Confining ourselves to these intensities and their geometrical distribution in the case of a three-dimensional periodic aggregate, we can formulate that a point

MICROMORPHOLOGY AND STRUCTURE RESEARCH

17

lattice in a real specimen (real space) has a corresponding point lattice in diffraction space (reciprocal space, Fourier space). In the case of primitive lattices, both real and reciprocal lattices are of the same geometrical type, e.g., a primitive monoclinic lattice in real space has a primitive monoclinic lattice in reciprocal space. Reciprocal lattice points are represented in diffraction patterns as reflections, the sites of high intensity. The reciprocal lattice participates in a rotation of the real lattice, however, both lattices are invariant with respect to translations. A given diffraction pattern is a central section through the reciprocal lattice (containing its origin, 000, where the undiffracted or direct beam intersects the plane of the diffraction pattern) and every set of equivalent planes (parallel to the beam) in the specimen produces a pair of reflections in the' pattern; these two reflections lie symmetric to the direct beam, perpendicular to the scattering plane, and have a distance from the direct beam of d*hki =

Cfdm

where C is a constant of the diffraction system (hence the importance of calibration) and dhkl the interplanar spacing of the Scattering planes (hkl). The indices h , k, 1of lattice planes are the reciprocal fractions of unit-cell sides between intersections of the planes (hkl) with the latter (as shown in Figs. 4c and 9), h is so for unit-cell side a , k for b, and 1 for c . Thus, unit-cell face A is (lOO),.B is (OlO), and C is (001). Crystal planes (hkl) are represented in diffraction patterns by a pair of symmetric reflections having the symbols hkl and hkl (Fig. 16a and b). In order to obtain the three-dimensional reciprocal lattice of a threedimensional periodic aggregate, diffraction patterns are collected from different orientations of the specimen. This can be achieved by electron

FIG. 9. Lattice planes (hkl) in a cubic lattice. See text. From Lange and Blodorn (1981). Courtesy of Thieme Verlag, Stuttgart.

18

RAlNER H. LANGE

diffraction in combination with specimen tilting in the electron microscope (the most direct method) or by optical diffraction of electron micrographs (projected views). Both methods supply similar, though not identical information (“projection theorem”; see Lake, 1972),however, a number of advantages make electron diffraction the preferred approach wherever possible (calibration, zonal tilting, three-dimensional instead of two-dimensional scattering object etc. ; Lange, 1982b). From the reconstruction of the reciprocal lattice we obtain the geometrical type of the real point lattice, point lattice symmetry, and, using known formulas (see Henry and Lonsdale, 1969), the real unit-cell sides. Thus, the first goal important for an appropriate description of a three-dimensional periodic aggregate is reached. It should be added that the analysis of diffraction patterns yields further information about lattice centering, the presence of screw axes (by the application of extinction rules), and actual symmetry (interesting in cases of pseudosymmetry). The second step of structural analysis aims at revealing the symmetry of molecular packing (by studying defined projections) and at determining coordinates of components. In simple cases it may not be too difficult to arrive at reasonable conclusions. However, we may have to accept that three-dimensional periodic aggregates do not have the high symmetry of crystals, e.g., microtubular arrays or liquid crystals (for review, see Brown and Woiken, 1979). In such cases, our approach does not work or does so only in a limited sense. Cases of apparent or possible crystaI nature should, at any rate, be seriously investigated using the above principles in order to not lose specimens providing easy access to the molecular level. In general. a three-dimensional computer-aided reconstruction of the specimen will finally establish symmetry at the molecular level with more precision, probably so in several steps along with progress made with respect to resolution.

C. POINTGROUPS Oligomeric enzymes (see for review Matthews and Bernhard, 1973; Klotz et a l . , 1970; Green, 1972) provide ample illustrations of point groups; however, they are not central to micromorphology. It is quite interesting that unusual values of cyclic symmetry may occur, e.g., 5 , 7, 13, and 17. There are three well-known examples of point symmetry groups in micromorphology , the connexon and gap junction complex (Staehelin, 1974; Caspar er a l . , 1977; Makowski er of., 1977), the shell of spherical viruses (Caspar and Klug, 1962), and-possibly so-the clath-

MICROMORPHOLOGY AND STRUCTURE RESEARCH

19

rin coat of coated vesicles (Kanaseki and Kadota, 1969; Crowther et al., 1976; Heuser, 1980; Crowther and Pearse, 1981). Based on electron microscopy and X-ray diffraction of isolated gap junctions (Caspar et al., 1977, Makowski et al., 1977), a gap junction consists of regularly arranged channels spanning adjacent plasma membranes, each channel consists of two units, connexons, one per cell membrane. The connexon has 6-fold rotational symmetry as does the channel, which arises by a rotation diad perpendicular to the 6 axis of the aligned connexons (Fig. 10). What then results is the dihedral symmetry 622 for each channel. Interestingly, the association of the channels in the gap junction area is governed by the same symmetry elements so that a plane group (symmetry p622) is present and long-range interaction of the constituent monomers of the connexon is to be taken into account (Caspar et al., 1977). These symmetry labels should not obscure the existence of short- and long-range disorder and the observation of systematic differences in connexon structure between samples (Caspar et al., 1977; Zampighi and Unwin, 1979). The shell of spherical viruses possesses cubic point symmetry 235 (icosahedral; Caspar and Klug, 1962). Hence the number of equivalent

FIG. 10. Section of a gap junction. The drawing takes account of the dissymmetry of the six monomers forming one connexon, of point.group symmetry 622 of one channel, and of plane group symmetry p622 of the whole arrangement. From Makowski et a / . (1977). Courtesy of Dr. D.L.D. Caspar and The Rockefeller University Press, New York.

20

RAINER H. LANGE

particles is 60. The classical article by Caspar and Klug is a key article in structure research and details of it will not be discussed here. We shall, however, be interested in the problem of how particles of a noncrystallographic point symmetry can assemble into highly symmetric threedimensional aggregates, especially in cubic crystals with only one or two particles per unit cell (see Section III,C,3). This is possible in the cubic space groups related to point group 23 because 3 of the 15 icosahedral 2-fold, and 4 of the icosahedral 3-fold axes coincide with cube edges and body diagonals, respectively (Fig. 6 ) , and has been considered since the early days of virus structure research (Klug and Caspar, 1960). The clathrin skeleton of coated vesicles with its hexagonal and pentagonal pattern (Fig. 1 la), like the icosahedral shell (Fig. 6), reminds us of the fact that a sphere cannot be covered by a closed hexagonal array. Heuser (1980) has illustrated the simultaneous presence of planar hexagonal clathrin nets (ideally p 6 , but containing pentagons and heptagons as well) underlying the cell membrane and of mixed mainly hexagonal-pentagonal nets covering single vesicles in the stage of pinching-off, thus demonstrating in this case the transformation from a planar to a vaulted structure accompanied by the necessary change in the coat pattern. We can also discuss the conditions under which coated vesicle skeletons take icosahedral symmetry. For this consideration the observation of Crowther and Pearse (1981) is important in that the molecular packing model for the edge between two vertices of the clathrin cage has 2-fold rotational symmetry when it best fits the aspect of negatively stained preparations. Furthermore, the quasi-3-fold vertices of the structure cannot have strict 3-fold symmetry because they border on hexagons and pentagons. However, 60 clathrin triskelions could form a cage with exact icosahedral symmetry. This cage would have 90 edges, 60 vertices, 20 hexagons, and 12 pentagons and a diameter of approximately 95 nm. In this icosahedral array, the triads would pierce the hexagon centers and the 5-fold axcs the pentagon centers; the icosahedral 2-fold axes would halve those edges connecting two pentagons (separating simultaneously two hexagons), and all other diads (those lying between hexagon and pentagon) would be local (Fig. 6). D. LINEGROUPS Among the well-known examples of line symmetry, microtubules (for review, see Dustin, 1978), actin filaments (F-actin; Huxley, 1972), and virus capsids (Caspar, 1963), the first two are of particular interest to micromorphology and, being of symmetry s, their structural polarity (Fig. 2) has played an important role in function-related considerations. In

MICROMORPHOLOGY AND STRUCTURE RESEARCH

21

FIG. 11. (a) Coated vesicle cages, isolated from brain homogenates and demonstrated by the quick-freeze, deep-etch, rotary-replication method with contrast reversed during printing. From Heuser (1980). Courtesy of Dr. J. Heuser and The Rockefeller University Press, New York. (b-k) Rhombic dodecahedra1 morphology of a crystal in a glucagon producing cell from cod, Gadus morrhua. as revealed by tilting a thin-sectioned crystal (h) in a systematic manner. Since the section contains only a slice of the crystal, the tilted aspects are incomplete. d+, g+, and f+ are models of a rhombic dodecahedron oriented in the same way as the corresponding crystal sections. Tilt angles and projcctions: b, [ l l l ] , -5Y"; c, [loll, -54";d, [1I11, -54";e, [Oll], +47";f, [OOl],O";g, [Olll], -46";h, [ i l l ] , +54";j, [IOI], +40°; k, [Ill], +5P. Theoreticdl angles for [ l l l ] , 54.7"; for [loll, 45". From Lange (1977). Bars, 100 nm.

73

L I

RAINER H . LANGE

low-resolution studies, structural polarity of these helical structures can be visualized by decoration. The polarity of actin filaments is well known in a number of situations (Tihey, 1977) including the antiparallel arrangement in the sarcomere. The idea that an antiparallel neurotubule arrangement in axons might be the basis for a centrifugal and a centripetal neuronal transport using the same mechanism (an interesting concept relating structure and function) apparently has to be abandoned in view of the results of decoration experiments that show equal structural polarization to be present in cases with a high proportion of decorated neurotubules (Heidemann et al., 1981). According to Borisy (1978), using the different growth rates for the two microtubule ends as indicators of polarity, microtubules grow unidirectionally (with respect to polarity) from organizing centers, more specifically at their (+)-end defined as the free end of ciliary microtubules. It will be interesting to know the molecular structure of other components of the cytoskeleton; they will necessarily have structural polarity unless they possess a 2-fold axis as in line group s2 (Fig. 8c). Looking at the above helical structures just as a monomeric particle array with helical symmetry is an oversimplification; this is demonstrated by a major contribution from micromorphology dealing with the attachment of associated structures not following the intrinsic repeat of microtubules (see the thin-section analysis of ciliary structure by Warner and Satir, 1974; or the review by Amos at ul., 1976).

E. SPACEGROUPS Naturally occurring protein crystals and numerical implications of their structure will be considered in this section. With respect to twodimensional space groups, a topical collection of reports is available in the volume edited by Baumeister and Vogell (1980), to which the reader is referred. Although intracellular crystalline aggregates cannot normally be considered as representing biologically active inclusions, their occurrence is not rare and their structural analysis and adequate description is possible, as well as desirable, in view of the interesting implications. There are, for example, the frequent intramitochondrial inclusions as analyzed by Berger (1969) and Sternlieb and Berger (1969), or the trigonal intracellular or even intranuclear crystals reminiscent of the trigonal form of catalase described by Langley (1967); in situ, such crystals have been studied in testicular interstitial cells (Nagano and Ohtsuki, 1971; crystals of Reinke) and in the frog parathyroid (Lange el ul., 1974). Similarly, the protein-

MICROMORPHOLOGY AND STRUCTURE RESEARCH

23

aceous contents of secretion granules frequently crystallize in insulin- and glucagon-secreting cells of the endocrine pancreas. Whereas apparent insulin crystals occur in amphibians, reptiles, birds, and mammals (for review see Lange, 1973), glucagon crystals seem to be confined to teleost islet cells (Lange and Klein, 1974; Lange, 1977; Lange and Kobayashi, 1980). Although the question is still open as to whether these crystals are pure hormone crystals, it is noteworthy that, by structural comparison, identity of intracellular crystals with crystal forms known from in uitro studies was not suggested in the case of insulin (Lange et al., 1972; Lange, 1976, 1980a). On the other hand, intracellular glucagon crystals from teleosts resemble the cubic form of porcine glucagon (King, 1959; Sasaki et af.,1975) so closely that they have tentatively been interpreted as being pure crystals of teleost glucagon (Lange, 1980a). Similarity includes external morphology (rhombic dodecahedron; Fig. 1lb-k) and cubic unit-cell length (electron microscopic values of a = 4.1-4.7 nm in three teleost species as compared to X-ray values of a =4.7-4.8 nm in porcine glucagon), as well as certain diffraction features fitting space group P2,3 of the porcine crystals (Lange, 1979, 1980a). The small dimensions of intracellular glucagon crystals (diameter approximately 100 nm) points to the importance of electron microscopy in this case and makes application of selected area diffraction impossible, so that microdiffraction (Lange and Kobayashi, 1980) and optical diffraction were the only useful techniques. On the basis of the cubic interpretation of intracellular glucagon crystals, a frequency distribution of exocytotic glucagon quanta (Fig. 12) could be derived by crystal measurements (Lange, 1979). Numerical restrictions for molecule numbers in crystals (see Section III,C,3) are such that the only alternative to the numbers given in Fig. 12 would be doubling them. When related values for transmitter quanta or quanta of other hormones-derived using biochemical methods-are compared with our data (Table I), the advantage of the morphological approach is obvious: it needs little material and yields a discrete distribution indicating more than one size class. Estimation of size classes in histograms requires the use of bias-free histogram techniques (moving histogram; see Victor, 1978). Whereas in the above cases physiological interpretation of crystal occurrence is difficult, such an interpretation is forced by high evolutionary conservation of crystal architecture in anamniote yolk platelets. Structural analysis of these oocyte inclusions also provides a good illustration of the general principles dealt with in the foregoing sections. Previous interpretations of the structure of the crystals in cyclostomes

24

RAINER H . LANGE

1 I

I

I I

2

1

I

I I,

I

I

I

6

I

I

I

6

I

1 I

I 1

10

I I

1;

[los Molecules1

C r y s t a l size FIG. 12. Frequency distribution of exocytotic glucagon quanta from a teleost (Xiphophorus hvllrri Heckel) as derived from crystal measurements and unit-cell size determination in thin-sectioned material. Moving histogram. The upper abcissa scale (unit cells) depends only on the presence of a cubic lattice. the lower scale (molecules) also depends on the correct space group (P2,3 was suggested in this case). Values are from nine fish divided into three groups (represented by white. black, and white areas). Two maxima of the frequency distribution are apparent, at approximately 400,000 and 800,000molecules. From Lange (1979). Ordinate scale is l i n , where n = 203 crystals.

and in an amphibian led to the claims that a cubic F-lattice (Fig. 13, left) should be present in cyclostomes (Karasaki, 1967) and a “simple hexagonal lattice” (Fig. 13, right) in an amphibian (for review, see Honjin, 1976). Such models are difficult to arrive at and to check unless these highly ordered aggregates are treated as crystals. Then, the cubic model cannot apply because it has diagonal particles (in this case probably phosvitin dimers) on the 3-fold cube that do not possess 3-fold rotational symmetry. If one reconstructs the reciprocal lattice by diffraction in a number of cyclostomes (both myxinoidea and petromyzontida, separated since some 250 million years) one obtains a reciprocal lattice-identical in all cases-with symmetry 2 / m (Fig. 24); therefore, a monoclinic point lattice is present in the crystals (Lange and Richter, 1981; Lange 1982a). It can be shown to be a C-lattice (Lange and Richter, 1981) made up principally of obvious dimers of lipovitellin pierced by the crystallographic 2-fold axes of rotation (Fig. 14). The model proposed by Honjin (1976; see Fig. 13) is not truly crystallographic. Low-angle X-ray patterns provided by this author have been discussed by Ohlendorf et al., (1975) and the latter authors have interpreted the structure of amphibian yolk platelets on the

1 NUMBER OF MOLECULES I N SECRETION GMNULES".~ Number of molecules/secretion granule Cell type, molecule, species Neurohypophysis, vasopressin, and oxytocin, rat B cell, insulin, rat

B cell, insulin, Natrix natrix A cell, glucagon, Myoxocephalus scorpius A cell, glucagon, Gndus rnorrhua A cell, glucagon, Fugu rubripes A cell, glucagon, Xiphophnrus helleri

Basis of estimation

Mean

Median

Sample size

Range

Biochemistry + morphometry

6 x lo4

-

Biochemistry + morphometry Crystal volume (~2~3) Crystal volume

1.5-3 x 105

-

2 x 106

1.5 x 106

5

2.1 x 105

2 x 105

8.5

X

104-6.2 x lo5

45

Crystal volume

3.3 x 105

3 x 105

1.8

X

1OS-6.1 x lo5

38

Crystal volume

2.7 x 105

2.4 x 105

3.8 x lo4-7 x los

I16

Crystal volume

7.1 x 105

4.7 x los

8.5

203

X

105-6.7 x lo6

X

104-4.5

X

lo6

32

" From Lange (1979). Estimated either by biochemical or by crystal packing considerations. In the latter case, more accurate values of the variation in sample size can be made.

26

RAINER H. LANGE

13

FIG. 13. Yolk-platelet structure models according to Karasaki (1967) (left) (see also Wallace. 1963) and Honjin (1976) (right). For a discussion of these models see text.

basis of X-ray powder patterns,negatively stained crystal fragments, and a three-dimensional molecular reconstruction as orthorhombic crystals, space group P 2 , 2 2 , , with four dimeric lipovitellin-phosvitin complexes per unit cell (Ohlendorf et al., 1978). When one studies such higher anamniotes like ancient bony fishes (Lange et al., 1981j, teleosts (Lange, 1980bj, and amphibians (Lange, 198la)-separated since some 400 million years-by thin-section electron microscopy, one easily reconstructs a reciprocal lattice-identical in all cases-with symmetry m m m (Fig. IS), that is, the crystals have an orthorhombic point lattice. In addition, a true 3-fold symmetry necessarily required in hexagonal arrays (see Section III,C,3) could not be found. The study of crystal projections in electron micrographs reveals three interesting results: 1 . Positively stained thin sections have much the same absolute density distribution as negatively stained thin crystal fragments (Fig. 16d and e). Thus the contrast is not reversed in the two preparations as should have been expected; this points to an irregular staining behavior of the constituent protein molecules. It is clear that such comparisons can only be made when the intrinsic coordinate system of the crystal is known. 2 . The density distribution-like the reciprocal lattice-in a number of projections studied is very much the same in ancient bony fishes, teleosts,

MICROMORPHOLOGY AND STRUCTURE RESEARCH

27

..

..a.

FIG. 14. The cyclostome yolk crystal. Reconstruction of the reciprocal lattice by arranging electron diffraction patterns according to their angular relationship in a stereographic projection. The front surface of a sphere is shown, projection labels are valid for the view from sphere surface to center. Symmetry 2/m of the reciprocal lattice; the mirror plane is set off by a heavy line, the 2 axis pierces the sphere at [OlO]. Bottom left: Unit cell and packing model [compare with Fig. 13 left, suggested by Karasaki (1967) for cyclostomes]. Bottom right: Interpretation of the symmetry of the unique projection along unit-cell side b with indication of rotation ( 4 ) and screw ( 4 ) diads. From Lange and Richter (1981). Compare this drawing with the micrograph in Fig. 16c.

RAINER H . LANCE

28

...... ....

. . . . .

...... ......

.10011.-

-

.,:. 0

...... . ......

. .ioioi.

-m-

. . . -.

...... .. ......

FIG. 15. The yolk crystal of fishes and amphibians. Reconstruction of the reciprocal lattice (symmetry tntnm) as in Fig. 14.

and amphibians, demonstrating the high evolutionary conservation of crystal architecture. 3 . According to the symmetry of the axial projections of the crystal, the space group should be P 2 , 2 , 2 , (see Figs. 16f and g and 17).

The yolk platelet shows the extreme usefulness of applying a priori knowledge to the analysis of an apparently crystalline aggregate; in this case it enabled an adequate description of the crystals, their interspecies comparison, model building (Lange, 1982a), and even the production of evidence for an extremely long evolutionary conservation of the struc-

MICROMORPHOLOGY AND STRUCTURE RESEARCH

29

ture. Since biochemically (DeVlaming et al., 1980) and genetically (Wahli et al., 1980), yolk proteins have changed in the course of evolution much like other proteins, the architecture of the crystalline aggregate must have a physiological significance, which has been interpreted as being related to the storage of cations necessary for embryonic development (Lange, 1981c, d). Since this property would be shared by cyclostome and higher anamniote platelets, since yolk proteins should be homologous in both groups of vertebrates, and since the volume available for a lipovitellinphosvitin complex is almost the same in the monoclinic and orthorhombic crystals (Lange and Richter, 19Sl>, both crystal forms should share structural features. This indeed seems to be the case, and the difference in molecular packing appears to be related to the presence of a symmetric lipovitellin dimer in cyclostomes and of a heterodimer in the higher anamniotes. The suggestions so far are based on a comparative study at the 2- to 3-nm level of resolution. However, their corroboration by high resolution studies (the course of which is outlined by the available data) will again need great efforts.

V. Concluding Remarks Biostructures are efficient and cannot escape certain restrictions given by their history. The occurrence of symmetric aggregates appears reasonable in connection with assembly processes, but oversymmetry would not be expected to occur in functional aggregates since there is no use in high symmetry as such. If symmetry is required at a macroscopic level, it is achieved, although it may not be realizable in a strict sense owing to molecular properties. Also at the near molecular level, considerable freedom of nature in constructing symmetric aggregates is always visible. In spite of these observations, which emphasize the dependence of our symmetry labels on the resolution obtained, it is felt and has been illustrated that the application of principles valid a priori for symmetric aggregates is meaningful in mircomorphology as long as the limitation by resolution is observed; this approach can be used provisionally to fill the gap between specimen resolution and the near atomic level in those not so rare instances in which access to the macromolecular basis of micromorphology is facilitated by the presence of structural redundancy. Furthermore, low-resolution data gain considerably in precision when a comparative study is performed in which use can be made of the clarifying effect of small morphological variations.

FIG. 16. (a,b) Electron diffraction patterns from yolk crystals of Myxine gQtinosa when the beam ha9 the direction of unit-cell side c (a) and of b (b). Fron Lange and Richter (1981). (c) Projected view of the yolk crystal from the cyclostome Lumgetru plnnrri along

MICROMORPHOLOGY AND STRUCTURE RESEARCH

OO

31

OO

0 OO

0

o b o ,o

o 0

0

0

0

o

Q

0

0

o 0

o 0

o 0 0 0

0

0

0

0

‘P2,.2,20 0

0

0 0

0

0 0

0 ‘0 0

FIG. 17. Orthorhombic yolk-platelet crystal. Interpretation of the projection along unit-cell side II (top; see also Fig. 16f) and along unit-cell cide c (bottom; see also Fig. 16g). Densely stained particles are indicated and numbered in the same way as in Fig. 16. The matter distribution in the upper projection can be explained by screw diads (half arrow) along unit-cell sides h and c. In the direction of the third side, a, there could be either a screw diad (space group P212121,left) or a rotation diad (space group P22,2,, right). The decision is made in favor of P212121 by the interpretation of the lower projection, which does not allow a 2-fold axis along unit-cell side Q (dashed arrows). ( +), rotation diad; ( $), screw diad perpendicular to paper plane. From Lange (1981a). Reprinted with permission from Macmillan Journals Ltd.

ACKNOWLEDGMENTS This manuscript was revised by K. R. Leonard, EMBL, Heidelberg, after Dr. Lang’s untimely death. We would like to thank Mr. G. Magdowski for technical assistance and Mrs. B. Wildner (Giessen) and N. van der Jagt (EMBL) for preparing the manuscript.

the unique unit-cell side b (parallel to all diads; compare with Fig. 14, bottom right). (d-g) Orthorhombic yolk crystal from a teleost (f, Peluicuchromis pulcher) and an amphibian (d,e,g, Xenopus lueuis) as seen in different projections (d, [212];e, (1011; f, [loo]; g, [Ool]). d and e are composed of one half (bottom left) from a positively stained thin section and one half (top right) from a negatively stained platelet fragment. The irregular staining behavior of the crystal components finds its expression in very little difference between the two preparations. f and g are explained with respect to symmetry and packing in Fig. 17. All electron images are averaged according to Lange (1981b). Bars, 10 nm.

32

RAINER H . LANGE

REFERENCES Amos. L.A.. Linck, R.W., and Klug. A.( 1976). Cell Mofil. 3, 847. Anderson, R.G. W.( 1980). Cell Bio. 4, 393. Barry. J.M., and Barry, E.M.(1969). “An Introduction to the Structure of Biological Molecules.“ Prentice-Hall, New York. Baumeister, W., and Vogell, W., eds.( 1980). “Electron Microscopy at Molecular Dirnensions.” Springer-Verlag Berlin. Beer. M., Frank. J.. Hanssen, K.-J., Kellenberger, E., and Williams. R.C. (1975). Rev. Biophys. 7, 211. Berger, J.E. (1969). J . Cell Biol. 43, 442. Bernal. J.D. (1966). In ”Principles of Biomolecular Organization” (G.E.W. Wolstenholine, and M.O’Connor. eds.), p.1. Churchill, London. Blundell. T.L.. and Johnson. L.N. (1976). “Protein Crystallography.”Acadernic Press, New York. Bor‘isy. Ci.G.(1978). J . M o l . B i d . 124, 565. Brown. G.H.. and Wolken, J.J. (1979). ”Liquid Crystals and Biological Structures.” Academic Press, New York. Carlemalm, E.. Garavito, M.. and Villiger, W.(1980). Eur. Congr. Electron Microsc., 7th Lriden Vol. 2. p. 656. Caspar. D.L.D. (1963). Adu. Protein Chem. 18, 37. Caspar. D.L.D.. and Klug. Ad1962). Cold Spring Harbor S y m p . Quant. Biol. 27, 1. Caspar, D.L.D., Goodenough, D.A.. Makowski. L., and Philips, W.C.( 1977). J . Cell Biol. 74, 605. Crick, F . H . C . . and Watson, J.D.(1956). Narirre (London) 177,473. Cross. B.A., Dyball, R.E.J., Dyer, R.G., Jones, C.W., Lincoln, D.W., Morris, J.F., and Pickering. R.T.( 1975). Recent. Prog. Horm. Res. 31, 243. Crowther. R.A., and Pearse, B.M.F.11981 ), J . Cell Biol. 91, 790. Crowther, R.A., Finch, J.T., and Pearse. B.M.F.( 1976). J . Mol. Bid 103, 785. Danielli. J.F. ( 1936). J . Cell. Comp.Physio1. 7, 393. DeVlaming, V.L... Wiley, H.S., Delahunty, G . , and Wallace, R.A.(1980). Comp. Biochem. Pliysiol. 67B, 6 I 3. Dustin. P.( 1978). “Microtubules.” Springer-Verlag. Berlin. Fawcett, D.W. (1964). In ”Modern Developments in Electron Microscopy” (B.M. Siegel, ed.), p. 257. Academic Press, New York. Frank, J . , Verschoor. A , , and Boublik. M.(1981). Science 214, 1353. Green, N.M.( 1972). In “Protein-Protein Interactions’’ (R. Jaenicke and E. Helmreich, eds.), p.183. Springer-Verlag, Berlin. Hartmann, M.( 19531. “Allgemeine Biologie.” Fischer, Stuttgart. Hauser, H.. Pascher. I.. and Sundell, S.(1980). J. Mol. Biol. 137,249. Heidemann, S.R.. Landers. J.M.. and Hambourg, M.A.( I981). J . Cell Biol. 91, 661. Henry. N.F.M., and Lonsdale. K., eds. (1969). “International Tables for X-Ray Crystallography.” Vol. I . Kynoch Press, Birmingham. Heuser, J.(1980). J . Cell Biol. 84, 560. Holser, W.T.(1958). 2. Krisfallogr. 110, 266. Honjin, RK1976). In “Recent Progress in Electron Microscopy of Cells and Tissues” (E. Yamada, V. Mizuhira. K . Kurosurni. and T. Nagano, eds.), p. 95. Thieme, Stuttgart. Home. R.W., and Markham,R.(1973). In “Practical Methods in Electron Microscopy”(A.M.Glauert. ed.),Vol. I . p.325. North-Holland Publ., Amsterdam. Howell. S.L.(1974). Adu.Cytopharmaco1. 2, 3 19.

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Huxley, H.E.(1972). In “The Structure and Function of Muscle” (G.H.Bourne, ed.), 2nd Ed., Vol. 1, Part 1 , p. 302. Academic Press, New York. Kanaseki, T., and Kadota, K.(1969). J. Cell Biol. 42, 202. Karasaki, S. (1967). J. Ultrastruct. Res. 18, 377. King, M.V. (1959). J. Mol. Biol. 1, 375. Klotz, I.M., Langerman, N.R., and Darnall, D.W.(1970). Annv. Rev. Biochem. 39, 25. Klug, A.(1969). In “Symmetry and Function of Biological Systems at the Macromolecular Level” (A.Engstrom and B. Strandberg, eds.), p.425. Alrnqvist & Wiksell, Stockholm. Klug, A., and Caspar, D.L.D.(1960). Adu. Virus Res. 7, 225. Klug, A., and De Rosier, D.J. (1966). Nature (London) 212, 29. Klug, A., Crick, F.H.C., and Wyckhoff, H.W.(1958). Actu Crystallog. 11, 199. Krstic, R.V.(1979). “Ultrastructure of the Mammalian Cell.” Springer-Verlag, Berlin. KuHer, S.W., and Yoshikami, D.(1975). J. Physiol. (London) 251, 465. Labaw, L.W., and Rossrnann, M.G.(1969). J. Ultrastruct. Res. 27, 105. Lake, J.A.(1972). In “Optical Transforms” (H. Lipson, ed.), p. 153. Academic Press, London. Lange, R.H. (1973). Hand. Histochem. 8, 141. Lange, R.H. (1976). In “Endocrine Gut and Pancreas” (T. Fujita, ed.), p. 167. Elsevier, Amsterdam. Lange, R.H. (1977). Gen. Comp. Endocrinol. 32, 208. Lange, R.H. (1979). Eur. J. Cell Biol. 20, 71. Lange, R.H.(1980a). In “Insulin-Chemistry, Structure and Function of Insulin and Related Hormones” (D. Brandenburg and A. Wollmer, eds.), p. 665. De Gruyter, Berlin. Lange, R.H.(198Ob). Cell Tissue Res. 209, 511. Lange, R.H.(1981a). Nature (Londonj 289, 329. Lange, R.H.(1981b). Mikroskopie (Vienna) 38, 142. Lange, R.H.(1981c). Z . Nafurforsch. 36c, 686. Lange, R.H.(1982a). J. Ultrastruct. Res. 79, 1 . Lange, R.H.(1982b). Mikroskopie (Viennaj 39, 207. Lange, R.H., and Blodorn, J.(1981) “Das Elektronenmikroskop: TEM + REM.” Thieme, Stuttgart. Lange, R.H., and Klein, C.(1974). Cell Tissue Res. 148,561. Lange, R.H., and Kobayashi, K.(1980). J. Ultrastruct. Res. 72, 20. Lange, R.H., and Richter, H.-P.(1981). J . Mol. Biol. 148, 487. Lange, R.H., Boseck, S . , and Syed Ali, S.(1972). Z . Zellforsch. 131, 559. Lange, R.H., Soames, A.R., and Coleman, R.(1974). Cell Tissue Res. 153, 167. Lange, R.H., Blodorn, J., Magdowski, G., and Trampisch, H.J.(1979). J . Ultrastrucf.Res. 68, 81. Lange, R.H., Grodzinski, Z., and Kilarski, W.(1981). Cell Tissue Res. 222, 159. Langer, R., Poppe, C., Schramm, H.J., and Hoppe, W.(1975). J. Mol. Biol. 93, 159. Longley, W.(1967). J. Mol. Biol. 30, 323. Makowski, L., Caspar, D.L.D., Phillips, W.C., and Goodenough, D.A.(1977). J . Cell Biol. 74, 629. Markert, C.L. (1963). Science 140, 1329. Matthews, B.W., and Bernhard, S.A.(1973). Annu. Reu. Biophys. Bioeng. 2, 257. Nagano, T., and Ohtsuki, I. (1971). J. Cell Biol. 51, 148. Ohlendorf, D.H., Collins, M.L., Puronen, E.O., Banaszak, L.J., and Harrison, S.C.( 1975). J . Mol. Biol. 99, 153. Ohlendorf, D.H., Wrenn, R.F., and Banaszak, L.J.(1978). Nature (London) 272, 28.

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Ottensrneyer, F.P., Andrew, J.W., Bazett-Jones, D.P., Chan, A.S.K., and Hewitt, J.(1977). J . Microsc. 109, 259. Pease, D.C., and Peterson, R.G.(1972). J . Ultrusfruct.Res. 41, 133. Revel, J .P.( 19641. J.Histochem.Cytochem. 12, 104. Sasaki, K., Dockerill, S., Adamiak, D.A., Tickle, I.J., and Blundell, T. (1975). Nature (London) 257, 75 1. Severs. N.J., and Warren, R.C.(1978). J. Ulrrastrucr. Res. 64, 124. Sjostrand, F.S., and Barajas. L.(1968). J. Ulrrustruci. Res. 25, 121. Staehelin, L.A. (1974). f n t . Reu. Cyrd. 39, 191. Steck. T.L.(1974). J . CellBiol. 62, 1. Sternlieb, I., and Berger. J.E.(1%9). J. Cell B i d . 43, 448. Tilney, L.G.(1977). In “International Cell Biology” (B.R.Brinkley and K.R. Porter, eds.), p. 388. Rockefeller Univ. Press, New York. Victor, N . (1978). Merhods fnf. Med. 17, 120. Wahli, W., David, I.B., Ryffel, C.U., and Weber, R.(1981). Science 212, 298. Wallace. R . A . (19631. Biochirn. Biophys, Acro 74, 505. Warner, F.D.. and Satir, P.(1974). J . Cell B i d . 63, 35. Winter. W.T.. Arnott, S., Isaac, D.H., and Atkins, E.D.T.(1978). J. Mol. B i d . 125, 1 . Zarnpighi, C., and Unwin, P.N.T.(1979). J. Mol. B i d . 135, 451.

INTERNATIONAL REVIEW OF CYTOLOGY, VOL. I13

Functional Inclusions in Prokaryotic Cells J. M. SHIVELY,* D. A. BRYANT,?R. C. FULLER, $ A. E. KONOPKA,§ S. E. STEVENS, J R . , ~AND W. R. STROHL** * Department of Biological Sciences, Clemson University, Clemson, South Carolina 29634,

t Department of Cell and Molecular Biology, Pennsylvania State University, University Park, Pennsylvania 16802, $ Department of Biochemistry, University of Massachusetts, Amherst, Massachusetts 01003, § Department of Biological Sciences, Purdue University, West Lafayette, Indiana 47907, and ** Depurtment of Microbiology, Ohio State University, Columbus, Ohio 43210

I. Introduction Inclusions are visible expressions of cell metabolism. The inclusion might be an integral part of the cell’s metabolic machinery, it might be important in adjusting the environment of the cell, thereby regulating certain metabolic events, or it might represent a product of the cell’s metabolism. In some instances, the inclusion might encompass more than one of these characteristics, e.g., poly-p-hydroxybutyrate granules. This review is organized along these functional lines. Consequently, a review on cell inclusions should contain a discussion of relevant metabolism, occurrence, ultrastructure, composition, formation, disappearance or loss, function, etc., and potentially the genetics of all of these processes. The point is that a review on inclusions will, out of necessity, be quite complex. Whether an inclusion is covered or omitted from this review as well as the depth and detail of the information given on those covered depends on the attention they have been given in other reviews, the availability of new data, the biases of the authors, and space limitations. The less common inclusions and those covered in detail elsewhere have been omitted; this omission should in no way minimize their interest and/or importance. The organization of information within the review is the sole responsibility of J. M. Shively. The inclusions of prokaryotes were reviewed by Shively in 1974, and those of cyanobacteria by Allen in 1984. A number of other reviews (see individual sections) have been published that cover one or more of the topics included in this review. 35

Copyright 0 1988 hv Academic Presr, Inc All nghts of reproduction in any form rescrved

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J. M. SHIVELY ET AL.

11. Inclusions as Metabolic Machinery

A. CARBOXYSOMES Polyhedral bodies have been observed in the cyanobacteria, thiobacilli, ammonia- and nitrite-oxidizing bacteria, Pseudornonas thermophilia K2, Rhodotnicrobhrn uannielii, and Prochloron (Codd and Marsden, 1984). As seen in thin section, the bodies are 40-900 nm in diameter and possess a surrounding barrier (shell) 3-4 nm thick. These inclusions were first isolated from Thiobacillrts neapolitanus, shown to contain ribulosebisphosphate carboxylase/oxygenase (Rubisco), and consequently called carboxysomes (Shively et al., 1973a). Cells also contain a soluble “nonparticulate” form of Rubisco; the enzymes appear to be identical (Lanaras and Codd, 1981a; Cannon and Shively, 1983). The polyhedral bodies of Anabaena cylindrica (Codd and Stewart, 1976), Nirrobacter agifis (Shively e f al., 1977), Nitrosornonas sp. (Harms et al., 1981), and Chlorogfoeopsis fritschii (Lanaras and Codd, 1981b) have since been shown to contain Rubisco. Presumably, all of the polyhedral bodies with similar size and structure will be carboxysomes. The carboxysomes were reviei:red extensively by Codd and Marsden (1984). Therefore, this report will only briefly summarize older data, and concentrate on recent contributions. Holthuijzen el al. (1986a) proposed that the carboxysomes of T. neapolitanus are pentagonal dodecahedrons possessing 12 pentameric planes. They indicated that this differed from the structure proposed by Peters (1974) for the polyhedral bodies of Nitrobacrer winogradskyi. According to Holthuijzen et al. (l986a), Peters’ model, “an icosahedron composed of 72 identical subunits,” would account for the regular hexagonal appearance of carboxysomes but would not explain the inability of the bodies (at least those of T . neapolitanus) to form two-dimensional crystals. Holthuijzen et al. (1986a) additionally proposed that the Rubisco is organized as a monolayer on the internal surface of the shell, the small subunit of Rubisco being the connector. This theory was primarily based on the analysis of carboxysome composition (see below) and is in sharp contrast to the general concept that the carboxysomes are “filled” with Rubisco. The authors do not explain how their proposed structure could account for the uniform granular substructure of the bodies in thin section (Shively et al., 1973b). Cannon and Shively (1983) reported that the carboxysomes of T . neapoliranus were composed of 12- 15 polypeptides as demonstrated by sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDSPAGE). Earlier, using SDS-PAGE. Biedermann and Westphal(1979) and

FUNCTIONAL 1NCLUSIONS IN PROKARYOTIC CELLS

37

Lanaras and Codd (1981b) showed seven and eight polypeptides for the carboxysomes of N . agilis and C. fritschii, respectively. In all of these reports, more than 50% of the protein was attributed to Rubisco. Cannon and Shively (1983) also identified two of the polypeptides as shell components. Holthuijzen et al. (1986b) recently confirmed and extended the composition studies of Cannon and Shively (1983);they stated that the bodies are composed of “8 proteins and at the most 13 polypeptides.” In addition to the large and small subunits of Rubisco, these authors reported the presence of four glycoproteins as shell components. One of these glycoproteins with an M,.of 54,000 comigrated with the large subunit of Rubisco in SDS-PAGE. This glycoprotein plus Rubisco, not Rubisco alone, accounted for 60% of the carboxysomes [see above). The relative abundance of these components along with electron microscopy data led the authors to propose the internal structure described above. Beudeker and Kuenen (1981) reported the presence of all Calvin cycle enzymes as well as several other enzymes in the carboxysomes of T. neapolitanus. This communication was obviously erroneous; Holthuijzen et al. (1986b) declared the absence of all of these enzymes with the exception of Rubisco. The enzymes shown to be absent in carboxysome preparations from a number of organisms (including those in the above report) are phosphoribulokinase, phosphoriboisomerase, Dglyceraldehyde-3-phosphate:NAD+ oxidoreductase, ATP:3-phospho-~glycerate 1-phosphotransferase, D-fructose-1,6-bisphosphate D-gjyceraldehyde-3-phosphate-lyase,sedoheptulose-l,7-bisphosphate1-phosphohydrolase, L-ma1ate:NAD’ oxidoreductase, ~-aspartate:2-oxoglutarate aminotransferase, ATRAMP phosphotransferase, and carbonic anhydrase (Cannon and Shively, 1983; Codd and Marsden, 1984; Hawthornthwaite et al., 1985; Lanaras and Codd, 1981b; Lanaras et al., 1985; Marsden et al., 1984). Westphal et al. (1979) reported the presence of extrachromosomal DNA in the carboxysomes of N . agilis and N . winogradskyi. It now seems likely that this observation was the result of plasmid DNA sticking to the inclusions during purification. Vakeria et al. (1984) and Holthuijzen et al. (1986c), studying different organisms, could not find any evidence for plasmid DNA in their preparations. Chromosomal DNA appears to stick to the surface of the bodies. The significance of this observed “sticking” DNA, if any, awaits further experimentation. New information on the formation, disappearance, and function of the carboxysome has not been forthcoming. Preliminary evidence [Shively , unpublished) has confirmed the earlier observations of Purohit et al. (1976). The carboxysomes of Thiobacillus intermedius disappear when Rubisco is repressed; this loss appears to be by dilution during cell

38

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M .SHIVELY ET AL.

growth. When the enzyme is derepressed, the carboxysomes return. Furthermore, the inclusions appear to form rather quickly after derepression, i.e., the formation is not delayed until the enzyme reaches an appropriate “high” level. It is the opinion of this author (J.M.S.) that the carboxysome will, in the future, be shown to be active in carbon dioxide fixation, i.e., a functional prokaryotic organelle.

B. CHLOROSOMES Chlorosomes, originally designated “chlorobium vesicles” (Cruden and Stanier, 1970; Holt er al., 1966; Cohen-Bazier et al., 1964), are found exclusively in the two families of green photosynthetic bacteria, the Chlorobiaceae, obligate anaerobic sulfur bacteria, and the Chloroflexiaceae facultative anaerobic nonsulfur bacteria. The term “chlorobium vesicle,” which was used in the only previous review on this topic (Shively , 1974), has been deemed inappropriate because these structures have now been found in all green phototrophic bacteria and the term vesicle implies a bilayer boundary that is not characteristic of either the morphological or biochemical topology of this structure. The discovery of chlorosomes and particularly their morphological description was originally observed in only two of the Chlorobiaceae, namely, Chlorobiurn and Prosthecochloris. The discovery of the facultative green photosynthetic bacterium Chlorojexus aurantiacus by Pierson and Castenholz (1974) stimulated a vast research effort in a number of laboratories on the structure, function, and development of the chlorosome in that organism (Staehelin et al., 1978; Schmidt, 1980; Sprague e l al., 1981a; Bruce e f al., 1982). In parallel, research on the topology and function of the chlorosome of the Chlorobiaceae has been pursued by several laboratories and recently reviewed by Gerola and Olson (1986) and Van Dorssen el a f . (1987). The initial structural observations on chlorosomes have been reviewed by Shively (1974). At that time it was thought that the chlorosome of Chlorobiurn was the photosynthetic apparatus of the green bacteria containing both a bacteriochlorophyll c (Bchl c ) and a bacteriochlorophyll a (Bchl a ) antenna and reaction center. The antenna Bchl a was identified as a water-soluble protein having an absorbance at 800 nm in viuo (Olson, 19801, which was subsequently crystallized and structurally analyzed by X-ray diffraction by Matthews et al. (1979). This antenna chlorophyll contains seven Bchl a’s per 40,000 M , polypeptide. The reaction center Bchl a , with an in uiuo absorbance at 840 nm, was thought to be associated with the chlorosome but has been shown since to be an integral part of the cytoplasmic membrane. The status of the topological arrange-

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

39

ment of the photochemical apparatus in Chloroflexus remained unclear until the publication by Staehelin et al. (1978) of a complete freezefracture-electron microscopic study. Early cell purification technology did not allow the definitive separation of cytoplasmic membrane material and chlorosomes, although enzymatic analysis of the chlorosomes and cytoplasmic membrane by Cruden and Stanier (1970) showed a clear distribution between respiratory enzymes associated with the cytoplasmic membrane and the compounds associated with the chlorosome. With the discovery of the facultative green bacterium Chlorojlexus, detailed structural analysis of the chlorosome has been undertaken using the organism’s potential for developing a photosynthetic apparatus subsequent to dark aerobic growth (Sprague et al., 1981b). k o r e recently, Staehelin et al. (1980), Olson (1980), and Van Dorssen et al. (1986) have analyzed electron microscopic, biochemical, and physicochemical data from experiments on Chlorobium and Prosthecochloris and have shown that these anaerobic green sulfur bacteria have similar but distinct chlorosome assemblies from Chlorojkxus. Structural parameters, chemical properties, and models of both systems have been published by Gerola and Olson (1986) and Feick and Fuller (1984). Table I shows a comparison of the properties of the chlorosome assemblies in the two families of green bacteria (Sprague and Varga, 1986). Thus, both families of green bacteria have their photochemical apparatus arranged in a manner that is distinctly different from other photosynthetic prokaryotes. The major antenna pigment, Bchl c , is associated within the nonunit membrane structure, the chlorosome, which, in turn, is located in the cytoplasmic compartment and attached to, but not derived from, the cytoplasmic membrane. The chlorosome with Bchl c acts as a light-harvesting antenna, whereas charge separation, energy transduction, and secondary electron transport occur in the cytoplasmic membrane. Although the discovery of chlorosomes and their isolation and chemical characterization occurred in the mid-1960s, it was not until the 1980s that their molecular topology and their relationship to the photosynthetic energy-transducing cytoplasmic membrane became clear. It now appears that the total photosynthetic systems in the Chlorobiaceae and the Chloroflexiaceae are quite distinct; however, the chlorosomes of both families have similar structure and functions (light harvesting and energy transfer). The initial freeze-fracture studies of chlorosomes on both Chlorobium and Chloroflexus were published by Staehelin et al. (1978, 1980). These studies showed clearly that the chlorosome was an independent cytoplas-

40

J. M. SHIVELY E T AL.

TABLE I STRUVI U R A L PARAMETERS OF Chlorohirttn A N D ChloroJexrrs CHI.OROSOMES" Chlorosome parameter or structure Length Width Thickness Envelope (Lipid-like layer with no substructure) Core Baseplate

Membrane attachment site

Chlorohiumb

ChlaroJe..uits

70-260 nm 40-100 nrn 20-60 nm 3 nm thick

90-150 nm'; 106 t 24 nmd 25-70 nmc ; 32 2 10 nmd 12 2 2 nmd 2 nm thick"

10-30 rod elements with I@-nrn diameter 5-6 nm thick 6-nm repeat that is oriented 40-60" from long axis of c hlorosorne 20-30 very large ( > I 2 nmJ particles in field of 10- to 12-nm particles

Rod elements -5.2-nm diameter with 6 n m globular subunits" Crystalline with -6-nm periodicity that is oriented 90" from long axis of chlorosome'

-

-5-nm particles in crystalline lattice' with 7-nm particles along the perimetef'

From Sprague and Varga (1%). Measurements were from C. limirolo strain 6230 by freeze-fracture electron microscopy (Staehelin el nl., 1980). c Measurements were made on negatively strained cells of five atrains of C . crrrrcrntiwus: J-10-fl, OK-70-fl, Y11oofl, 39&1, and 254-2 (Madigan and Brock. 1977). Measurements were made on freeze-fractured cells of C . ouronriacus J-104(Staehelin ef d.,1978). ' Measurements were made on freeze-fractured cells of C. iritruntircrus J - 1 M during development of the photosynthetic apparatus (Sprague et a / . , 1981a). "

mic structure not bound by a classic unit membrane. In both cases, the chlorosomes were oblong structures attached to the cytoplasmic surface of the cytoplasmic membrane of the cell. In Chloroflexus, the structure varies in thickness depending on the stage of development. In both families, the internal structure is made up of linear arrays of rod-like elements probably containing the Bchl c (Feick and Fuller, 1984; Gerola and Olson, 1986). The Bchl c is entirely contained within the chlorosome along with several carotenoids and other nonpigmented lipids. In addition, there is a small amount of Bchl a absorbing at around 790 nm associated with the chlorosomes of both families (Schmidt, 1980; Betti et af.. 1982). The association of the cytoplasmic membrane and the chlorosome can be visualized in freeze-fracture micrographs as particulate arrays and have been designated as attachment sites o r base plates (Staehelin et a(., 1978, 1980). In general, the attachment of chlorosome to

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

41

cytoplasmic membrane is fairly rigorous, making their separation rather difficult during purification procedures. Taking advantage of the ability of ChZorofexus to grow in a respiratory mode in the dark, where neither chlorosomes nor cytoplasmic membrane reaction center and Bchl a antenna are made, a thorough study of the topological structure and the development of the chlorosome has been made (Sprague et ul., 1981a; Feick and Fuller, 1984; Van Dorssen et al., 198613). On the basis of this work the assembly, function, and molecular arrangement of both structural and functional components of the chlorosome in ChforoJexus have been analyzed. Similar studies on the reaction center components in the cytoplasmic membrane have also been accomplished, thus allowing a thorough study of the association and development of the two systems. Physicochemical studies on isolated chlorosomes and chlorosomemembrane complexes have been carried out by Betti et af. (1982). Linear dichroism, circular dichroism, and electron spin resonance (ESR) studies all have indicated that Bchl c is arranged as at least a dimer at an organizational level. Fluorescence studies in this work also indicated an energy transfer system involving primary capture by Bchl c with excitation transfer among the Bchl c molecules and subsequent transfer to the B790 pigment, which then transfers its exited states to the membrane antenna Bchl u and to the reaction center. The function of this B790 Bchl a pigment is of interest since it may serve as a bridging or energy transfer structural component between the chlorosome and the cytoplasmic membrane (Betti et ul., 1982). SDS-PAGE analysis of purified chlorosomes has revealed only three major and one minor polypeptide species in the chlorosome. Biochemical analyses using proteolytic enzyme treatment, photolabeling with 3-azido2,7-naphthalenedisulfonic acid, and chemical cross-linking with N ethylmaleimide followed by two-dimensional PAGE have yielded both a molecular-topographical and a protein-oligomeric definition of the chlorosome in ChforoJexus (Feick and Fuller, 1984). The results of these experiments are summarized in Fig. 1. Of the three major proteins from purified chlorosomes, two are located on the surface of the chlorosome. The first has an M, of 11,000 and appears to be present as a trimer; the second has an M , of 18,000 and appears as a cross-linked compound with the M, 11,000 trimer. The two polypeptides are clearly closely associated on the surface of the chlorosome. The third major polypeptide has an M , of 5600 (formerly described as having an M , of 3700) and appears to be associated with the major 740-nm-absorbing Bchl c. This polypeptide is present as a dimer con-

42

.I. M. SHIVELY ET AL.

envelope

I

.

membrone

reocfron cenier polypephdes ‘ontenno 5808-866 polypepfides (M, 26000)

( M, 5300)

FIG. I . Topological diagram of the chlorosome-membrane complex of Chlorojexus aumntiacrrs. This model is derived from the freeze-fracture micrographs of Staehelin ef d. ( 1980), physicochemical parameters observed by Betti ef nl. (1982), biochemical analysis by Feick and Fuller (1984), and Bchl c primary and secondary structure by Wechsler e f d. (1985a). All of these diverse analyses complement and confirm each other. The chlorosome contains the Bchl c polypeptide arranged as a dimer in a tubular array of globular subunits ( 5 . 8 x 5.2 nm) with the seven interacting porphyrin rings linearly arranged along the helix of the monomer. The baseplate 8790 energy transfer polypeptide bridges the area betwen the chlorosome and the cytoplasmic membrane. The transmembrane P865 reaction center and a hexameric Bchl a antenna polypeptide complete the primary light-harvesting and energytransducing photosynthetic complex.

firming previous ESR studies (Betti et al., 1982) and is associated with 10-16 Bchl c molecules per dimer (Feick and Fuller, 1984). The fourth minor chlorosome protein constituent (B790) has an M , of 5800 and is sequestered from protease digestion and photolabeling only when the chlorosomes are still attached to the cytoplasmic membrane, suggesting its bridging location between chlorosome and membrane. This peptide is of extreme interest because if may be associated with the new spectral species of Bchl a described above that absorbs at 790 nrn in uiuo

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

43

(Schmidt, 1980; Sprague et al., 1981a) and serves as a conduit for energy transfer from Bchl c in the chlorosome to the antenna Bchl a B808-865 in the cytoplasmic membrane (Betti et al., 1982). The role of this chlorosome-cytoplasmic membrane-bridging protein was first suggested by the appearance of an organized paracrystalline array at the chlorosome attachment sites on the cytoplasmic membrane (Staehelin et al., 1978). These authors suggested that this “attachment” site could play a role in energy transfer from the chlorosome antenna to the cytoplasmic membrane. This supposition again was strengthened by the discovery that the new Bchl a species, B790, is associated with the chlorosome and by the observation of fluorescent emission at 802 nm ascribed to this pigment species. The 802-nm emission is observed both in isolated chlorosomes and whole cells, suggesting that B790 is an in uiuo structural reality (Betti et al., 1982). The chlorosome attachment site in the Chlorobiaceae appears in an entirely different arrangement consisting of a crystalline array of the water-soluble Bchl a protein, whose crystal structure has been determined (Matthews et al., 1974). Chlorujexus appears to lack this protein entirely and utilizes instead the B790 pigment in the base plate and a B808-865 antenna protein similar to that found in the purple bacteria as the acceptor of the excitation in the membrane (Wechsler et al., 1985a). The previously cited reviews by Van Dorssen et al. (1987) and Gerola and Olson (1986) on the organization of the chlorosome and its relation to the cytoplasmic membrane of Chlorobiaceae emphasize the rather different arrangement of those systems in that family. Another important aspect of the structure of the chlorosome is the small protein content relative to the large amount of Bchl c . Olson (1980) suggested on the basis of geometrical considerations that 12-14 Bchl c molecules are associated with an M, 15,000 D-peptide to form the linear rods observed in electron micrographs of chlorosomes in Chlorobium limicola f. thiosulfatuphilum. Feick and Fuller (1984) have shown that 10-16 Bchl c molecules are associated with a dimer of M , 5600 peptides, giving a weight ratio of pigment to protein of greater than one. This suggests that the organization of pigment in the chlorosome may be quite different from that found in all other chlorophyll proteins that have been isolated from a wide variety of photosynthetic organisms. In all these other systems the mass of peptide is much larger than that of pigment. Recent spectroscopic evidence strongly suggests, however, that the Bchl c molecules in chlorosomes may form oligomeric arrays in organic solvents in which the hydroxyethyl group at position 2, ring 1 forms a proton-sharing H bond with the keto group of ring 5 of another Bchl c ‘Smith et al., 1983b). This unusual arrangement is entirely in agreement

44

J . M. SHIVELY ET AL.

with the structural and spectroscopic data now available on the in uiuo chlorosome structure (Feick and Fuller, 1984). Recent work of Wechsler et id. (1985) has elucidated the complete amino acid sequences of the chlorosome Bchl c polypeptide in Chloroflexus. The structure with a molecular mass of 5600 (51 amino acids) is similar to light-harvesting Bchl a polypeptides in purple bacteria. However, the stoichiometric relationship between Bchl and the polypeptide is strikingly different. On the basis of previous measurements (Feick and Fuller, 1984) it has been shown that around 6-8 Bchl c’s (actually 7) are bound to each 51 amino acid monomer. This is a most curious structure with a higher amount of Bchl c than protein per molecular monomer. A similar ratio has been described in the pigment protein of the chlorosomes of the Chlorobiaceae by Smith et a f . (1983b) and Matthews rt al. (1979). In a helical wheel model of the pigment-protein complex (Wechsler et ul., 1985) it was shown that 5 glutamines and 2 asparagines are distributed vertically along the helix between tryptophan 5 and isoleucine 42 and are located on the outer edge of one side of the helix. It is suggested that the amide residues are possible interaction sites with 7 Bchl c molecules, via the central magnesium atoms. This model presents a surface array of porphyrin rings along the helix that can also interact at the hydroxy and keto groups in rings 1 and 5 as suggested by Smith et al. (1983b). On the basis of a peptide dimer subunit and the arrangement of the location of the amide-containing amino acids, one can suggest a chlorosome subunit consisting of 12 polypeptide chains (6 dimers) associated with 84 Bchl c molecules located on the surface of such a unit. This topography is diagramed in Fig. 1. The molecular dimensions of this arrangement correspond to the electron microscopically demonstrated globular subunits first described by Staehelin et al. (1978). This model of the Bchl c polypeptides in the chlorosomes presents a unique pigment-protein complex that suggests a highly efficient, primitive, and indeed unique photochemical antenna arrangement present in the chlorosomes in green bacteria.

C. PHYCOBILISOMES Phycobilisomes (PBS) are supramolecular protein structures found in cyanobacteria, the chloroplasts of red algae, and the chloroplast-like “cyanelles” of certain dinoflagellates such as Cyanophora purudo.xa; they function as the light-harvesting antennae for Photosystem I1 (Manodori and Melis, 1984, 1985; Manodori et uf., 1984). PBS are primarily composed of the brilliantly colored phycobiliproteins, a class of water-soluble proteins that bears covalently attached open chain tetrapyr-

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

45

role chromophores (phycobilins). In addition, PBS also contain smaller amounts of proteins, most of which do not bear chromophores and which are referred to as “linker polypeptides” (8000-120,000 Mr). These polypeptides are required for proper assembly and functional organization of the structure. The PBS is the photosynthetic antenna system that has been studied in greatest detail. Many recent reviews provide detailed information and comprehensive lists of original references for this field (Bryant, 1986; Gantt, 1980, 1981; Glazer, 1982, 1983, 1984; Glazer et al., 1985a; Cohen-Bazire and Bryant, 1982; Tandeau de Marsac, 1983; Wehrmeyer, 1983b; Zuber, 1985; Scheer, 1981, 1982). The photosynthetic apparatus in the cyanobacteria is housed in and upon a complex membrane system, referred to as the thylakoid membranes, which likely results from infoldings and differentiation of the cytoplasmic membrane. Although this is difficult to discern in many cyanobacteria, there are certain cyanobacteria for which this interpretation is clearly correct (e.g., Arthrospira jenneri; see micrographs in Wildman and Bowen, 1974). PBS are typically located on the stromal (external) surfaces of the thylakoid membrane pair and are characteristically absent from the cytoplasmic membrane surface. In cyanobacteria, PBS are arranged in short or long rows that are displaced in parallel on the stromal surfaces. Remarkably regular arrays of PBS are observed in some species (cf. Lichtle and Thomas, 1976; Gantt, 1980). This regularity suggests that the chlorophyll-protein complexes of Photosystem 11, to which the PBS deliver their excitation energy and to which the PBS are presumably attached (see Manodori and Melis, 1985; Khanna er al., 1983), must likewise be regularly displaced in the plane of the membrane. Evidence supporting this contention can be found from freeze-fracture studies of the thylakoids of PBS-containing organisms (Giddings et al., 1983; Golecki and Drews, 1982). Figure 2 presents a model for the thylakoid membrane of the cyanelle of C. paradoxa (Giddings et al., 1983); this model applies equally well to the cyanobacterial thylakoid. The morphology of phycobilisomes can vary significantly and is dependent upon the source organism (Gantt, 1980). Three types of PBS have been described among the cyanobacteria: (1) bundle-shaped, (2) hemiellipsoidal, and (3) hemidiscoidal. A fourth class of PBS structure, “block-shaped,” has been reported for the PBS of the red alga GrifJithsia pacifica (Gantt and Lipschultz. 1980) but has not yet been reported for a cyanobacterium. Gloeobacter violaceus, an unusual cyanobacterium that lacks thylakoids (Rippka et al., 1974), is the only organism known to have bundle-shaped PBS (Guglielmi et al., 1981). The PBS of G. violaceus consist of bundles of six rods, 50-70 nm long, 10-12

J. M. SHIVELY ET AL.

46

c3

= P S I I +phycobiIisome ottatchment sites (-1Onm EF particles)

0

= P S I , cytochromes, CFo (-7nm

PF particles)

FIG. ?. Model for the cyanohacterial thylakoid membrane. The hemidiscoidal PBS, fihich usually occur a s regularly spaced rows. are attached t o the stoma1 (protoplasmic) surface of the thylakoid membrane. Each PBS is in contact with two Photosystem I1 (P680) reaction centers. Also shown, in regions not covered by the PBS. arc the other three major photosynthetic complexes: the Photosystem I (P700) reaction center complex; the plastoquinol-plastocyanin oxidoreductase (cytochrome .f,hh complex), and the ATP synthase (CF,-CF, coupling factor) complex. EF, exoplasmic fracture face: PF. protoplasmic fracture face. Taken from Giddings ct NI. (1983). ~

nm in diameter, and composed of 8-12 disks 6 nm in thickness. These rods are attached to a poorly defined basal structure and presumably serve to attach the structure to the photosynthetically active cytoplasmic membrane. In C. uiulaceus cells these PBS are attached to the inner surface of the cytoplasmic membrane and stand with their long axes perpendicular to the plane of that membrane (Rippka et uf., 1974; Guglielmi ef a / . , 1981). In thin-section electron micrographs, the PBS appear as an electron-dense cortical layer at the inner surface of the cytoplasmic membrane (Rippka et a / ., 1974). The most common PBS structure is the hemidiscoidal form that has been observed for numerous cyanobacteria (Bryant et ul., 1979; Glazer et

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

47

al., 1979; Nies and Wehrmeyer, 1980; Rosinski et al., 1981; Ohki and Gantt, 1983; Guglielmi and Cohen-Bazire, 1984; Ohki et a f . ,1985;Raps et al., 1985) and for the red alga, Porphyridium aerugineum (Gantt et ul., 1968) and for the cyanelles of the dinoflagellate C . paradoxu (Giddings et al., 1983; see Figs. 2, 3, and 4). The appearance of these PBS in thin-section electron micrographs is dependent upon the plane of sectioning. When the rows of PBS are viewed in cross section, the PBS have a semicircular outline. When the rows of PBS are viewed in longitudinal or tangential (relative to the plane of the membrane surface) section, these PBS appear as regularly spaced, electron-opaque rods. Hemidiscoidal PBS are 45-75 nm wide at their base, about 14 nm thick, and about 30-40 nm high. Each hemidiscoidal PBS consists of two distinct substructures that are constructed from eight or nine cylindrical building blocks. The “core” is composed of two (Glazer et al., 1979) or, more commonly, three (Bryant el a f . ,1979) cylinders that have a diameter of about 11 nm and a length of 14-17 nm. Each of the core cylinders is composed of a stack of four disks about 3.5 nm in thickness. In PBS with tricylindrical cores, these cylinders are stacked along their long axes to produce a structure that approximates a regular triangular prism (see Fig. 4). Radiating from each of two surfaces of this core substructure are six “peripheral rod” substructures. Each peripheral rod is a cylindrical stack of disks with a diameter of 11-12 nm and a thickness of about 6 nm. The number of disks per rod can range from two to six and is dependent upon both the source organism and the growth conditions employed (Bryant et al., 1979;Glazer et al., 1979). It should be noted that each component disk of the peripheral rod is, in turn, composed of two disks approximately 11-12 nm in diameter and 3 nm thick. Until recently, hemiellipsoidal PBS had been reported only in red algae such as Porphyridium cruentum (Gantt and Lipschultz, 1972) and Gustrocfonium coufteri (Glazer et a f . , 1983). However, Guglielmi and CohenBazire (1984) have shown that the PBS of LPP Group sp. PCC 7376 are hemiellipsoidal (63 nm, 40 nm high, and 25 nm long). An accurate model for the hemiellipsoidal PBS, the first PBS type to be isolated and examined (Gantt and Lipschultz, 1972), has not yet been developed because their large size produces a complex superpositioning of stain layers that precludes a straightforward analysis of the electron microscopic images. Nonetheless, some views of these PBS closely resemble those of hemidiscoidal PBS as observed in face view (see above and Fig. 2) and it is possible that these structures simply represent a double thickness of that structure. PBS are believed to be entirely composed of two types of proteins:

48

J . M. SHIVELY ET AL.

phycobiliproteins and linker polypeptides. The phycobiliproteins generally comprise 85-90% of the total PBS protein, with the linker polypeptides accounting for the difference. As routinely isolated, PBS do not contain chlorophyll. The absorbance properties of the phycobiliproteins allow their classifi= 490-570 nm), cation into three major groups: phycoerythrins (A,, = 610-630 nm), and allophycocyanins (A, = 650phycocyanins (A, 670 nm). The basic structure of all phycobiliproteins is a protomer composed of two dissimilar subunits, (Y and @, with molecular masses in the 17,000-21,000 M , range. The complete primary structures of a variety of phycobiliproteins representing all spectroscopic classes and derived from phylogenetically distant sources have been determined (cf. Wehrmeyer, 1983a). These studies lead to several conclusions. First, all primary sequences can be assigned to one of two classes, a-type or @-type. Second, within any given spectroscopic class, a very high degree of primary sequence conservation exists no matter how distantly related the source organism may be. Third, a significant degree of homology exists when comparing a or @ subunits from different spectroscopic families. Finally, the (Y and @ subunit families are sufficiently homologous to establish that the phycobiliproteins constitute a family or proteins descended from a single ancestral gene. Each phycobiliprotein subunit carries at least one linear tetrapyrrole chromophore, which is covalently attached to the polypeptide chain via one or two thioether linkages and that gives the proteins their characteristic and intense visible absorption properties (Glazer, 1985). Four chemically distinct chromophores have been identified among cyanobacterial biliproteins: phycocyanobilin, phycoerythrobilin, phycourobilin, and a biliviolin-type chromophore of as yet undetermined structure (CohenBazire and Bryant, 1982; Glazer, 1985). In general, the absorbance bands of native proteins that occur in the range 590-670 nm are assigned to phycocyanobilin, those that occur in the range 540-570 nm to phycoerythrobilin, those that occur in the range 490-5 10 to phycourobilin, and those that occur at around 565-575 in the subunit of phycoerythrocyanin to the biliviolin-type chromophore (Bryant el af., 1976; Bryant, 1982). Clearly, much of the spectroscopic diversity exhibited by the phycobiliproteins is the direct result of these chemically distinct chromophores that have different numbers of conjugated double bonds in their orbital systems (see Glazer 1985, for a summary of the structures of the various chromophores). However, the chemically distinct tetrapyrrole chromophores are not the only elements contributing to the generation of spectroscopic diversity. Although all covalent linkages are thioether linkages to vinyl

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

49

substituents carried by the pyrrole rings, there are at least three distinct modes of linkage for phycocyanobilin, phycoerythrobilin, and phycourobilin chromophores. Linkages through vinyl substituents on the A pyrrole ring, the D pyrrole ring, or both the A and D pyrrole rings are possible (Glazer, 1985). Finally, the protein environment of the chromophore is extremely important in determining the absorption properties of a given chromophore (compare the absorbance maxima of allophycocyanin and phycocyanin, both due to phycocyanobilin chromophores, in Table 11). It should be noted that protein-protein interactions, which occur during the assembly of the phycobiliproteins, play prominent roles in specifically altering the chromophore environments during the assembly processes that lead to the formation of the PBS. Table I1 summarizes some of the spectroscopic, chemical, and physical properties of the phycobiliproteins that have been described from various cyanobacteria. The basic unit upon which all higher assembly forms of phycobiliproteins are constructed is a disk-shaped (a& trimer, about 3 x 12 nm (Bryant et al., 1976). A notable advance in understanding PBS structure was the solution of the three-dimensional structure of the Mastigocladus laminosus phycocyanin at 0.3 nm resolution by X-ray analysis (Schirmer et al., 1985). The structure solved was that of phycocyanin trimers (ap)3. The molecule is torus shaped and has a diameter of 11.O nm, a thickness of 3.0 nm, and a central hole about 3.5 nm in diameter. This structure agrees nicely with many electron micrographic images of phycocyanin and allophycocyanin trimers (Morschel et al., 1980). Although the structure can eventually be refined to at least 0.21 nm, the present degree of resolution clearly defines the polypeptide backbones of the a and p subunits, establishes the extent and location of eight a-helices for each subunit, and defines the positions and conformations of the nine phycocyanobilin chromophores that occur in the trimer. An interesting point concerning the structure of phycocyanin is that six a-helices (A, B, and E-H) of each biliprotein subunit are similar in three-dimensional arrangement to the equivalent helices of myoglobin and hemoglobin (Schirmer et al., 1985). This observation suggests that the biliproteins and these oxygen-binding heme proteins are members of a superfamily of proteins descended from a common ancestral gene. The advance promises to provide significant new insights for other biliproteins as well. A solution for the phycocyanin of Synechococcus sp. PCC 7002, which crystallizes as stacks of hexarners (arpk that resemble the arrangement of this phycocyanin in PBS, has recently been achieved (Schirmer et al., 1986). Crystals of phycoerythrocyanin from M . laminosus have also been obtained that are isomorphous with those of the phycocyanin trimer (Rumbeli et al., 1985). One imagines that phycoery-

Aggregation state"

Protein Allophycocyanin B

PAP)?

(aAI'13

Bilin content per subunit' a: I PCB 1 PCB (Y: I PCB p: I PCB a: I PCB p : 2 PCB

Molecular mass ( X 10 Da)

Visible absorption maximum (nm)

Fluorescence emission maximum (nm)

89

670 1 618

675

I (X)

650

660

36.5-220

620

630-6445

I20

568, SW'

625

40-240

565

575-580

p:

Allophycocyanin

(d" PA"),

Ph ycoc yanin

(aW

Ph ycoer ythroc yanin

p,,, (n =

1-6)

a: I PXB

pPii(jI

p: 2 PCB Phycoerythrin"

p PL:'La ( n

= 1-6)

a: 2 PEB

B : 3 PEB " Data taken from Cohen-Bazire and Bryant (1982) and Glazer (198.5).

* The molecular masses and spectroscopic

properties are thobe of the aggregates specified in this column. ' PCB, phycocyanobilin: PEB, phycoerythrobilin: PXB, phycobiliviolin-type chromophore. Shoulder Some cyanobacterial phycoerythrins are spectroscopically similar to those found in red algae (R-phycocrythrins). Their subunit structure is carry phycourobilin chromophores (Ong rt a/.,1984: Glazer, 198%.

''

(u,!&y

and they

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

51

thrins have structures that resemble those for phycocyanins, since phycoerythrins share extensive sequence homology with phycocyanins (Sidler et a f . , 1986; Maze1 et af., 1986; Dubbs and Bryant, 1987). The data from the M. faminosus phycocyanin structure might allow refinement of the crystal structure for P. cruentum B-phycoerythrin (Fisher et al., 1980). At this point, diffraction quality crystals of allophycocyanin are required to complete the structure of the PBS at a first, crude level. Even when mixed at very high protein concentrations, highly purified biliproteins do not interact with one another to form higher order aggregates (Bryant et al., 1976). Instead, the assembly of biliproteins into PBS requires a small family of polypeptides, which have been named “linker polypeptides” to reflect their essential role in the assembly process. Linker polypeptides were first shown to be components of cyanobacterial PBS by Tandeau de Marsac and Cohen-Bazire (1977). These workers found that in addition to the phycobiliproteins the linker polypeptides varied when cells capable of chromatic adaptations (Tandeau de Marsac, 1983) were grown under appropriate light wavelength conditions. They postulated that these polypeptides might play roles in PBS assembly and membrane attachment. The number of linker polypeptides required for the assembly of a given PBS depends upon the cyanobacterial species and the number of spectroscopically different biliproteins that comprise the PBS. Typically, 6 to 10 distinct polypeptides are required. The linker polypeptides can be arbitrarily divided into two groups; those that participate in the assembly of the core (Lc) and those that participate in the assembly of the peripheral rod (LR). Glazer (1985) has proposed a systematic nomenclature with abbreviated symbols to represent the linker polypeptides as well as the subunits of the phycobiliproteins; this system will be employed in the remainder of this discussion. Since most of the linker polypeptides do not carry prosthetic groups, they have been referred to as “colorless polypeptides” by some authors (e.g., Cohen-Bazire and Bryant, 1982). However, more recent studies indicate that certain linker polypeptides carry bilin chromophores and hence can participate in the light energy capture and transfer processes. Examples include the y subunits of certain phycoerythrins (Glazer and Hixson, 1977; Ong et al., 1984) and the large core-linker phycobiliproteins, which play roles in membrane attachment, core assembly, and energy transfer to chlorophyll a (Lundell et a f . , 1981b; Redlinger and Gantt, 1982). Lundell et al. (1981a) first purified four linker polypeptides from the PBS of Synechococcus sp. PCC 6301 and determined their amino acid compositions and peptide maps. From the results of these studies and in uitro assembly experiments, these workers concluded that each of the

52

J. M . SHIVELY ET AL.

linker polypeptides was a distinct protein. The linker polypeptides are basic polypeptides suggesting that electrostatic interactions play an important role in their interactions with the acidic biliproteins. In this regard it is interesting to note that the acidic residues that line the inner surface of the phycocyanin trimer are conserved residues in all phycobiliprotein classes (T. Schirmer, personal communication). Glazer (1985) has suggested that the linker polypeptides (1) determine the aggregation state and geometry of the protein with which they interact, (2) modulate the spectroscopic properties of the biliprotein, and (3) determine the location of the biliproteins within the PBS and form bridges between biliprotein complexes within the structure. Structural information has recently become available for several linker polypeptides. Two recent studies (Gantt et al., 1985; Zilinskas and Howell, 1986) have shown that the large core-linker phycobiliproteins (LFK 2o 1 of diverse organisms exhibit a high degree of structural Gmilarity by immunological techniques. Moreover, the amino terminal 5equence9 of the L& and L?,, polypeptides of Synechococcns sp. PCC 6301 and PCC 7002, respectively, are 75% homologous in spite of the apparently large difference in the molecular masses of the polypeptides (G. Guglielmi and D. Bryant, unpublished results). Complete primary structure information has become available for several linker polypeptides and additional partial amino acid sequences have been determined for the amino termini of several other linker polypeptides. Fuglistaller et af. (1984, 1985) have determined the com lete amino acid sequences of 8f and Li9 as well as the amino two M . laminosirs linker polypeptides, LC terminal sequences of Lk4' PEC and Li45 pc and the carboxyl terminal sequence of Li4 ' PEC. The complete amino acid sequences of two linker polypeptides, L i 9 pc and Li3 pc, have been determined from a translation of the nucleotide sequence of the genes from Synechococcus sp. PCC 7002 (Bryant et a!., 1987). Finally, the complete amino acid sequences of the core linker L:8 of Synechococcus sp. PCC 6301 (Houmard et al., 1986) and Synechococcus sp. PCC 7002 (Bryant er al., 1987) have also been determined from a translation of the nucleotide sequences of the genes. Comparisons of these data reveal several new and intriguing aspects about the linker polypeptides. First, these data indicate that the linker polypeptides from different cyanobacteria share considerable sequence homology. For example, the two Li9 proteins are 50% homologous in sequence. Second, the sequence data indicate that the linker polypeptides Lc8, Li9, L? ' ", LF PEC, and Li3 pc share considerable amounts of sequence homology and thus form a family of related proteins. These proteins share a region of very high homology at their carboxyl termini. The conserved domain might be required for interactions with the

'

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

53

equally highly conserved biliproteins. This conserved domain in the linker polypeptides is rich in basic amino acids, and these amino acids could interact with acidic amino acids lining the inner surface of the central hole in the biliprotein trimer. Finally, and perhaps most striking of all, sequence comparisons between the 30,000 M , linker polypeptides and biliproteins with which they interact suggest that the linker polypeptides share some distant homology with the a and P subunits of the biliproteins (Fuglistaller et al., 1985). Thus, all components of the PBS can reasonably be argued to be the descendants of a single ancestral gene (Fuglistaller et al., 1985). It should be noted that Schirmer et al. (1985) have also found evidence that biliproteins and globins are descendants of a common ancestral gene. If this is correct, the globin and biliprotein-linker gene families form one of the largest “supergene” families known. Figures 3 and 4 show diagrammatic representations of the PBS of Synechococcus sp. PCC 6301 and Synechocystis sp. PCC 6701 (Glazer, 1984, 1985). The peripheral rods of the PBS of Synechococcus sp. PCC 6301 are composed of phycocyanin-linker polypeptide complexes as shown. Each of these complexes possesses unique spectroscopic properties that contribute to the unidirectional flow of excitation energy toward the PBS core and ultimately to the chlorophyll a of the Photosystem 11 reaction centers (Lundell et al., 1981a). The organization of the peripheral rods of Synechocystis sp. PCC 6701 is similar to that observed for Synechococcus sp. PCC 6301. The phycoerythrin-linker polypeptide complexes are localized at the distal ends of the peripheral rods as shown (Bryant et al., 1979; Gingrich et al., 1982a, b). The core structure of the PBS is complex and must necessarily be asymmetric to accommodate its interaction with both the thylakoid membrane surface and the peripheral rods. Glazer and co-workers have shown cores to be composed of several distinct multiprotein subcomplexes (Yamanaka et al., 1982; Lundell and Glazer, 1983a-c). Each of the two cylinders of the Synechococcus sp. PCC 6301 core is composed (aAPPAP>* of four subcomplexes as shown in Fig. 3: (aAPfA%, L&, aAPBa:’ p;” L&0.5,and (aAP L&.’. The two central and L& polypeptides, contain the complexes, which contain the aAPB terminal energy acceptors for the PBS and probably are responsible for the delivery of excitation energy to chlorophyll a molecules associated with the Photosystem 11 reaction centers. The tricylindrical core structure of Synechocystis sp. PCC 6701 has not been as extensively characterized as that of Synechococcus sp. PCC 6301 and, at this time, the precise organization cannot be defined. Gingrich et af. (1983) succeeded in isolating a series of subcomplexes from this core that closely resembled the subcomplexes from the Synechococcus sp.

54

J. M. SHIVELY E T AL.

A

(aPCBPC)6

LZ

(aPcppc)6

(apcPpc)6L ’;

nm

622.5

622.5

620

Emox mM-’cm-’

2370

2370

2364

nm

652

648

643

Amox

hiox

B

ayp,”P

a;PpAPflle3L75

2

CM

APB AP AP L10.5 a2 p 3 C

a:p~~L~05

652.5

nm

650

655

652.5

Emox mM-’ cm-‘ G a x nm

7 70

1100

820

1020

660

680

680

662

‘max

FIG. 3. Diagrammatic representation of the hemidiscoidal PBS of Synechococcus sp. PCC 6301. The composition and some properties of the subcomplexes comprising the peripheral rods and the core are shown in parts A and B. respectively. The superscripts AP, APB, and APB denote the biliproteins allophycocyanin, allophycocyanin B, and phycocyanin, respectively. The symbols LR,LRC.Lc, and LCMdenote linker polypeptides associated with the peripheral rods, the rod-core junction, the core, and the core-thylakoid membrane junction, respectively; the superscript numbers indicate the molecular masses for these polypeptides in kilodaltons. Taken from Glazer (1983, which provides additional details of the model.

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

55

"FACE V I E W " OF

PHYCOBILISOME

@ (apEpPE)6 'L.:

576

@ (apcppc),L~

642

fig

"SIDE V I E W " OF CORE

FIG.4. Diagrammatic representation of the hemidiscoidal PBS of Synechocystis sp. PCC 6701. The composition and fluorescence emission properties of the subcomplexes of this PBS are shown. The superscripts AP,APB, PC,and PE denote the biliproteins allophycocyanin, allophycocyanin B, phycocyanin, and phycoerythrin, respectively. The symbols LR, LRC, Lc, and LCMdenote linker polypeptides associated with the periphelal rods, the rod-core junction, the core, and the core-thylakoid membrane junction, respectively; the superscript numbers indicate the molecular masses for these polypeptides in kilodaltons. The number of phycocyanobilin (PCB) and phycoerythrobilin (PEB) chromophoresof the subcomplexesare also shown. Taken from Glazer and Clark (1986).

56

J . M. SHIVELY ET AL.

PCC 6301 core by chromatography of dissociated core fractions on U-(diethylaminoethyl) cellulose (DEAE-cellulose) columns. On the basis of the relative stoichiometries of the subcomplexes obtained, these workers suggested that the third cylinder was composed of two (aAP PAP)3and two (aAP Lko complexes and that the overall structure was probably the same as for Synechococcus sp. PCC 6301 with the addition of the third cylinder. Anderson and Eiserling (1986) have recently isolated core substructures from the Synechocystis sp. PCC 6701 core, which are not consistent with the interpretations of Gingrich et al. (1983). These workers suggest that the subcomplexes of the bottom cylinders are not arranged in the antiparallel fashion shown in Fig. 4 but are asymmetrically distributed. Early work on the photosynthetic apparatus of the cyanobacteria demonstrated that light absorbed by chlorophyll a was less efficient in driving oxygen evolution than light absorbed by the phycobiliproteins (Clayton, 1980). Other studies showed that chlorophyll a fluorescence was more efficiently sensitized by light absorbed by biliproteins than by light absorbed by chlorophyll a itself (Duysens, 1951; French and Young, 1952). These observations played important roles in the formulation of the "Z scheme" for oxygenic photosynthesis, which postulated two interacting photosystems. More recent studies suggest that the Photosystem I reaction center (P700 center) of cyanobacteria contains a chlorophyll a antenna composed of about 120-130 chlorophyll a molecules (Myers et al., 1980; Williams et a / ., 1983; Lundell et al., 1985). The Photosystem 11 reaction center (P680 center) carries a much smaller chlorophyll a antenna composed of about 50 chlorophyll a molecules (Myers et al., 1980; Yamagishi and Katoh, 1985). A variety of studies indicate that most (>95%) of the light energy absorbed by the PBS is initially transferred to Photosystem I1 (Manodori and Melis, 1984, 1985; Manodori et al., 1984). Typical hemidiscoidal PBS carry 300-700 phycobilin chromophores; therefore, PBS greatly augment the absorption cross section for Photosystem 11. Since light energy absorbed by the phycobiliproteins can be delivered to the Photosystem I1 reaction centers with an overall efficiency of 90%, the energy transfer process must proceed rapidly to avoid energy losses by competing radiative or nonradiative decay processes. Hence, it is important to consider briefly those aspects of PBS that contribute to the rate and efficiency of the energy transfer process. Excitation energy transfer has been extensively studied by picosecond fluorescence spectroscopy in the PBS of Synechocystis sp. PCC 6701 (see Fig. 4). and in mutants of this organism that produce PBS deficient in either phycoerythrin alone or both phycocyanin and phycoerthrin (Glazer er al., 1985a. b; Glazer and Clark, 1986). When isolated, PBS are excited

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

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with light absorbed by any component biliprotein; the fluorescence emission occurs via the terminal acceptors L:’M or aAPBat about 676-680 nm. When these PBS are excited with light at wavelengths primarily absorbed by phycoerythrin, fluorescence emission at 680 nm occurs in 56 8 psec; excitation with light primarily absorbed by phycocyanin results in fluorescence at 680 nm in 28 ? 4 psec; and excitation of isolated core substructures results in emission at 680 nm in 6.6 2 3.6 psec. The transfer of energy among the chromophores within an isolated biliprotein disk occurs in < 8 psec. Thus, these results indicate that it is the disk-to-disk transfer time, about 24 psec which is the rate-limiting step in the energy transfer process. Model-building experiments (Bryant et a / ., 1979) suggested that the chromophores of the peripheral rods would be essentially noninteracting. This reduces by nearly a factor of six the random walk possibilities available to excitation energy after it enters the chromophore bed of the PBS (Glazer, 1984). In considering energy flow dynamics, each PBS substructure (peripheral rod disks and cores) can approximately be regarded as a single chromophore since intradisk transfer events are rapid relatie to disk-to-disk transfers. Mimuro et af. (1986) have carefully studied the phycocyanin trimer of M . laminosus whose crystal structure has been solved. From their analyses they conclude that within the trimer disk, light energy is concentrated from chromphores on the periphery of the trimer (the a subunit phycocyanobilin and p subunit phycocyanobilin attached to cysteine-155) to the chromophores that extend into the central hole of the trimer (the phycocyanogilin attached to cysteine-84). Light energy thus concentrated could quickly migrate through the peripheral rods through a “chromophore pipeline” to the allophycocyanin of the core substructure (Schirmer et al., 1986; Mimuro et a / . , 1986). The energy transfer process in the Synechocystis sp. PCC 6701 PBS (see Fig. 4) can be approximated by five intersubstructure transfer events (Glazer and Clark, 1986). Energy flow through the PBS is unidirectional because the component complexes are arranged in order of decreasing energy from the periphery of the structure to the terminal acceptors of the core. This is true even for those PBS that have only phycocyanin in their peripheral rods since the linker polypeptides produce subtle modulations of the chromophore energy levels that cause otherwise identical chromophores to reside at different energy levels within the rods (Lundell et al., 1981a). The relatively large energy differences between phycoerythrin and phycocyanin and between phycocyanin and allophycocyanin, as well as the photochemical reactions associated with the Photosystem 11 reaction center, all serve to drive the energy through the system in an essentially unidirectional fashion.

*

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The amount of phycobiliprotein per cell as well as the specific phycobiliprotein composition of cyanobacteria are affected by a variety of physical and chemical parameters. These include temperature (Halldal, 1958; Anderson et al., 1983), light intensity (Myers and Kratz, 1955; Allen, 1968: Raps et al., 1983; Lonneborg et al., 1985), light wavelength (Jones and Myers, 1965; Myers et al., 1980; Bogorad, 1975; Tandeau de Marsac, 1983), carbon dioxide concentration (Eley, 1971), and the availability of nitrogen (Allen and Smith, 1969), sulfur (Schmidt et al., 1982), phosphate (Ihlenfelt and Gibson, 1975; Stevens et al., 1981b), and iron (Hardie et a / . , 1983b; Guikema and Sherman, 1983). Alterations in the specific phycobiliprotein composition of an organism can be affected by changing the composition and generally the length of the peripheral rod substructures. The best studied example of this is complementary chromatic adaptation, that has been recently reviewed by Tandeau de Marsac (1983). Conditions that favor a decreased total biliprotein content of cells (high light intensity or nutrient deprivation) cause a reduction in the number of PBS but may also cause a decrease in the length of the peripheral rod substructures of some species (Yamanaka and Glazer, 1980, 1981). It should be noted that the phycobiliproteins appear to represent a storage form of reduced nitrogen and perhaps carbon that can be mobilized to provide a pool of amino acids for new protein biosynthesis during conditions of nutrient limitation. This aspect of PBS has recently been reviewed by Allen (1984). The structural genes encoding several PBS components have recently been isolated and characterized. They include the cpcA and cpcB genes encoding the a and p subunits of phycocyanin from Synchococcus sp. PCC 7002 (de Lorimier el al., 1984; Pilot and Fox, 1984), Synechococcus sp. PCC 6301 (Lind et al., 1985; Kalla et al., 1985), C . paradoxa (Lemaux and Grossman, 1984, 1985), Pseudanabaena sp. PCC 7409 (Bryant et ul., 1986), and Calothrjr sp. PCC 7601 (Conley et al., 1985); the apcA and apcB genes encoding the a and p subunits of allophycocyanin from Synechococcus sp. PCC 7002 (Bryant et al., 1986), Synechococcus sp. PCC 6301 (Houmard et al., 1986), C . paradoxa (Bryant et al., 1985a, b; Lemaux and Grossman, 1985), and Cafothrix sp. PCC 7601 (A. Grossman, personal communication); the cpeA and cpeB genes encoding the (Y and p subunits of phycoerythrin from Calothrix sp. PCC 7601 (Maze1 et al., 1986) and Pseudoanabaena sp. PCC 7409 (Bryant et a / . , 1986; Dubbs and Bryant, 1987); the acpC gene encoding the allophycocyanin-associated core-linker Lt9 polypeptide in Synechococcus sp. PCC 6301 (Houmard et al., 1986) and Synechococcus sp. PCC 7002 (Bryant et a/., 1987); and the cpcC and cpcD genes of Synechococcus sp. PCC 7002 33 Pc encoding the LR and L i 9 pc phycocyanin-associated linker polypeptides (Bryant ct al., 1986, 1987).

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In Synechococcus sp. PCC 7002 the genes encoding the PBS components are organized into two apparent operons: a “rod” operon and a “core” operon. The rod operon consists of the cpcB, cpcA, cpcC, and cpcD genes, which are transcribed in that order (Bryant et al., 1985b, 1986, 1987. Two unidentified open reading frames, one of 38 codons upstream from the cpcD gene, have also been found in the cluster. Mutational analysis suggests that the latter may be a trans-acting regulatory element; the translation products of both open reading frames do not correspond to those of known structural components of the PBS (R. de Lorimier, D. A. Bryant, G. Guglielme, and S. E. Stevens, Jr., unpublished results). The core operon of Synechococcus sp. PCC 7002 consists of the upcA, upcB, and apcC genes, which are transcribed in that order (Bryant et a / . , 1987). This arrangement has also been found in Synechococcus sp. PCC 6301 (Houmard et al., 1986) and Calothrix sp.PCC 7601 (A. Grossman, personal communication). The availability of the cloned genes will allow a complete analysis of PBS structure and assembly through the construction of defined mutations. To date, four structural mutations have been characterized in Synechococcus sp. PCC 7002: (1) a deletion of cpcA and cpcB genes, (2) a deletion of the cpcD genes, ( 3 ) an insertion mutation of the cpcC gene (Bryant et al., 1986, 1987), and (4) a deletion of the apcA and apcB genes (V. Stirewalt and D. Bryant, unpublished results). In general, these studies have confirmed the existing notions concerning PBS assembly that were developed by Glazer and co-workers through in uitro experiments (Glazer, 1982, 1984, 1985). The cloned genes are also being employed to investigate the control of biliprotein gene expression by light intensity (e.g., see Conley et al., 1985), and by light wavelength and nitrogen availability (Bryant et al., 1986). These latter two effects are being evaluated in Synechococcus sp. PCC 7002 through the use of a lacZ-cpcB translational fusion that contains all upstream regulatory sequences for the rod operon. Preliminary experiments indicate that the cpc promoter does indeed respond to both light intensity and nitrogen availbility (Bryant et al., 1986; Gasparich et al., 1987).

HI. Inclusions as Adjusters of the Environment

GAS VESICLES Gas vesicles are rigid shells of protein that are found in a wide variety of prokaryotic organisms. The protein subunits of the gas vesicle have unique properties that enable their association to result in a structure that is impermeable to water. The interior contains gases that diffuse through

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the proteinaceous layer. The gas vesicle is permeable to all gases that have been tested (Walsby, 1969, 1971, 1982), therefore, it cannot function as a storage reservoir for metabolic gases. The function of gas vesicles is to provide buoyancy. The hollow structure has a density much less than that of water. Gas vesicles can be identified (Walsby, 1974) and their abundance quantified (Walsby, 1973) by their refractility. The structures are susceptible to rupture by rapid application of pressure Walsby, 1971). The term “gas vacuole” refers to irregularly shaped refractile areas observed in prokaryotes containing gas vesicles when examined in the light microscope. These areas are most prominent when phase contrast optics are used and are due to light scattered by gas vesicles. Gas vesicles occur in phylogenetically diverse species of bacteria. Lists of cyanobacteria (Walsby, 1981a) and other bacteria (Walsby, 1981b) reported to have gas vesicles have been published. The common characteristic of gas-vacuolate strains is that they are aquatic microbes. A large number of gas-vacuolate organisms have never been isolated but have been observed during microscopic examination of natural samples (Skuja, 1956, 1964; Caldwell and Tiedje, 1975; Walsby, 1974; Clark and Walsby, 1978). In a few species, gas vesicles are not produced in vegetative cells but only in differentiated cells involved in dispersal. This has been shown in several cyanobacteria that form hormogonia (Armstrong e f al., 1983; Singh and Tiwari, 1970). In Clostridium, the gas vesicles are produced in the exosporium that surrounds the endospore (Oren, 1983; Duda and Makar’eva, 1978). Although gas vesicles occur in unrelated prokaryotic species, their structures are quite similar. They have the form of cylinders capped by conical endpieces (Bowen and Jensen, 1965; Cohen-Bazire et al., 1969; Smith and Peat, 1967). There are regularly spaced ribs that run perpendicular to the long axis of the gas vesicle (Jost and Jones, 1970; Konopka et a / . , 1977; Walsby and Eichelberger, 1968). Although the morphology is similar, the size varies among organisms. In general, cyanobacteria have longer gas vesicles (with maximum lengths of SOO-1000 nm) than eubacteria (maximum lengths less than SO0 nm). The eubacterial vesicles tend to be wider (100-125 nm) than those of cyanobacteria (40-70 nm). The gas vesicles of several archaebacteria have been examined. The gas vesicles of two methanogenic species are morphologically similar to those of cyanobacteria (Archer and King, 1984; Zinder et af., 1987). In Hulobacferium strains (Simon, 1981) and a halophilic square bacterium (Parkes and Walsby, 1981), two types of gas vesicles have been reported; the majority are spindle shaped with widths up to 300 nm and a small proportion are similar in size and morphology to those of eubacteria. The strength of gas vesicles is inversely related to their width (Walsby, 1972).

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61

Given the similarities in molecular structure of different gas vesicles (see below), the basis for the size differences remains an intriguing, unresolved question. Gas vesicles from Anabaena Jlos-aquae, two Halobacterium species (Krantz and Ballou, 1973; Falkenberg, 1974), Microcystis aeruginosa (Weathers et al., 1977), and Ancylobacter aquaticus (Konopka et a l . , 1977) [formerly known as Microcyclus aquaticus (Raj, I983)I have been isolated and chemically characterized. All of these studies concluded that the gas vesicles were composed of protein. Gas vesicles contain only one type of protein subunit. The initial evidence for this conclusion came from polyacrylamide gel electrophoresis of purified gas vesicles run under denaturing conditions (Jones and Jost, 1971; Konopka et al., 1977; Walker and Walsby, 1983). However, treatments such as suspension in sodium dodecyl sulfate, which solubilize most other proteins, do not always yield satisfactory results with gas vesicles (Hayes et al., 1986). Thus the evidence for a single type of protein was not unequivocal, and the molecular mass of the protein was uncertain. Walker et al. (1984) reported continuous Nterminal amino acid sequences of six gas vesicle proteins (GVP) from six organisms. The sequences ranged in length from 26 to 64 residues. A single, unambiguous sequence was obtained in each case suggesting that the gas vesicles were composed of a single protein. This conclusion also held for Halobacterium salinarium strain 5 , whose gas vesicles had been reported by Falkenberg (1974) to contain two proteins. The complete amino acid sequence of GVP isolated from Anabaena JEos-aquaehas been determined (Hayes et al., 1986). It contains 70 amino acids and has a molecular mass of 7388. An oligonucleotide corresponding to a portion of the amino acid sequence of the Anabaena GVP was synthesized and used to isolate the gene from Calothrix PCC 7601 that codes for this protein (Tandeau de Marsac et al., 1985). Nucleotide sequence analysis indicated that the gupA gene produced a protein of 70 amino acids and had a molecular mass of 7375. Furthermore, the sequence was almost identical to that of the Anabaena GVP. The sequence data together with earlier reports of the amino acid composition of purified gas vesicles illustrated that about 50% of the amino acid residues in GVP were nonpolar, the content of aromatic amino acids was low, and that sulfur-containing amino acids were absent in most preparations (but see Krantz and Ballou, 1973). The N-terminal portion (residues 1-10) of the Calothrix and Anabaena proteins contains a high proportion of polar amino acids, whereas the middle third of the molecules is enriched in nonpolar residues. Thus, gas vesicles are composed of one type of protein subunit that is arranged to form the ribs seen in electron micrographs. X-Ray diffraction

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studies of gas vesicles from Halobacterium halobium (Blaurock and Wober, 1976) and Anabaenajos-aquae (Blaurock and Walsby, 1976) led to a proposed structure for the gas vesicle of Anabaena (Walsby, 1978). The gas vesicle wall is 1.8 nm thick and the ribs have a periodicity of 4.57 nrn. The diffraction patterns suggested that part of the protein was folded into a p sheet and that there was a repeating interval of 1.15 nrn along the ribs. The volume of the unit cell defined by these results would accommodate a protein with a molecular mass of 7500 (Hayes ef al., 1986), very close to the value derived from sequence analysis of Anabaena protein and the Calothrix gvp gene. Although gas vesicles are found in phylogenetically diverse prokaryotes, GVP is conserved. Antisera prepared against the gas vesicles of Aizubaenaflos-aquae (Walker et al., 1984) and A . aquaticus (Konopka et al., 1977) were able to agglutinate gas vesicles from any organism tested including examples of archaebacteria, chemoheterotrophic bacteria, and cyanobacteria. The N-terminal sequences of the first 46 amino acids of GVP isolated from three planktonic cyanobacteria were identical in more than 90% of the positions and the sequence from a halophilic cyanobacterium was 85% identical (Walker et al., 1984). Halobacterial GVPs were sequenced in the same study and these were broadly similar to the cyanobacterial protein. Many of the differences in sequence were caused by the substitution of one nonpolar amino acid for another. The conservation of GVP sequence in divergent species may be a consequence of the constraints involved in fulfilling its unique requirements-to maintain a structure that excludes water. The proteinaceous shell of a gas vesicle is not inflated by gas; the structure is assembled in a way that excludes water from the interior so that a space into which gas can diffuse is created. Gas vesicle assembly has been examined by electron microscopy (Waaland and Branton, 1969; Lehmann and Jost, 1971; Konopka et al., 1975). Small biconical structures were initially observed. These increased in length and width until the width equaled that of the “mature” gas vesicle. Then, the cylindrical midsection was assembled. Waaland and Branton (1969) suggested that new protein subunits were added at the central rib of the gas vesicle, and intuitively this seems most likely, but there are no conclusive data supporting the idea. A basic, unresolved question regarding gas vesicle formation is whether the protein subunits can self-assemble. In uitro experiments with purified G V P cannot be done as long as there is no satisfactory way to solubilize isolated gas vesicles. Konopka et al. (1975) inferred that GVP did not self-assemble because of two observations. First, they isolated a mutant of A . aquaticus that produced gas vesicles lacking the cylindrical midsection. Walsby (1978) suggested that the amino acid sequence of GVP was

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63

altered so that the protein-protein interactions necessary for cylinder (but not cone) assembly could not occur. It seems more Iikely that there was a mutation in another gene whose product either modified GVP or was necessary for assembly. The second relevant observation of Konopka et al. (1975) concerned the kinetics of gas vesicle assembly. When the existing gas vesicles in wild-type cells were collapsed, the assembly of about 20 biconical vesicles was initiated. Subsequently, very few new initiations were found until the first vesicles had completed cone assembly and begun cylinder assembly. At this time, a new set of small biconical structures was initiated; this suggested that there was a site (such as an assembly enzyme) necessary for cone assembly but not for cylinder assembly. Many of the experiments studying gas vesicle assembly were done by collapsing the intact gas vesicles in cells and monitoring the appearance of new gas vesicles. Although the collapsed gas vesicles could not be reinflated, Hayes and Walsby (1984) have shown that the GVP in newly assembled vesicles are from this source. In contrast, when protein synthesis was inhibited in A. aquaticus that contained collapsed gas vesicles, the number of gas vesicles assembled was only 2% of the control value (Konopka et al., 1975). Another fundamental problem regarding assembly is how one type of protein subunit can associate to form the structure. Although proteinprotein interactions can be identical in any portion of the cylinder, they seem likely to vary between subunits in different ribs of the cone (Walsby, 1978). This problem has been extensively studied in the case of spherical virus particles and although the domains (Rossmann and Argos, 1981) of viral protein subunits appear to be very rigid, flexibility in protein structure can be attained by hinge movements between protein domains or by conformational changes due to ligand binding (Rossmann, 1984). Furthermore, assembly pathways may be controlled by the temporary presence of scaffolding proteins, as in the case of bacteriophage P22 (King, 1980). Clones in which gas vesicle production is reduced or abolished have been reported to spontaneously arise at a high frequency in several organisms (Larsen et al., 1967; Das and Singh, 1976a,b; Thomas and Walsby, 1985). These high frequencies and the absence of GVP in a mutant of Anabaena flus-aquae prompted Walsby (1977) to suggest that the GVP gene was located on a plasmid. There is no conclusive evidence to support this hypothesis. In Calurhrix PCC 7601, it is clear that GVP is not encoded on plasmids because a DNA probe specific for the gupA gene did not hybridize to any endogenous Calothrix plasmids but did hybridize to chromosomal DNA (Tandeau de Marsac et a f . , 1985). Extensive studies of genetic variability have been done with two

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species of Halobacterium. Spontaneous mutants of H . salinarium strain 5, in which gas vesicle production is delayed and reduced, occur at a frequency of about 1% (Simon, 1981). These mutants do not revert back to wild type even when selection pressure for reversion is applied. The mutants also lack a plasmid that is found in the wild type (Simon, 1978). Obviously, this plasmid does not contain the gene for gas vesicle protein; Simon suggested that it encoded a system that processed gas vesicle protein prior to assembly. Halobacterium halobium NRC 81 7 produced nonvacuolate clones at a frequency of lop2.However, revertants also arose at a high frequency (Heifer et al., 1981). This genetic variability was due to the insertion and removal of transposable elements (Heifer e f al., 1983) into a large plasmid. It is not known if the plasmid contained all the genetic information for gas vesicle production. Thus, genetic instability is not due to extrachromosomal location per se but rather to the action of transposable elements. Two types of variability have been found in A. aquaticus. If batch cultures are allowed to reach late stationary phase, nonvacuolate clones are found at a frequency of lo-’. Motile variants can also be selected (Lara and Konopka, 1987) and these motile cultures do not usually produce gas vesicles. The phenomenon seems analogous to flagellar phase variation in Salmonella typhimuriurn (lino, 1977). Motile variants revert to nonmotile, vacuolate forms at frequencies of lo-* to transitions to the motile form occur at least lo00 times less frequently. Walsby (1976) suggested that Prosthecomicrobium pneumaticum might contain two copies of the gene for gas vesicle protein. He thought the phenotype of a mutant containing approximately 50% of the wild-type level of gas vesicles was due to gene dosage. Although this conjecture has not been confirmed in P. pneumaticum, use of a DNA probe specific for the GVP gene has demonstrated that there are three copies of this gene, two of which are located 100 base pairs apart on the Calothrix PCC 7601 chromosome (Tandeau de Marsac et al., 1985). The nucleotide sequences of the two linked genes differ in only 18 base pairs, all of which represent silent mutations that do not alter the polypeptide product (Damerval et al., 1987). The function of gas vesicles appears to be buoyancy; their density i s 0.12 g/cm3 (Walsby and Armstrong, 1979). If a cell contains sufficient gas vesicles to counterbalance the weight of molecules denser than water, it will be buoyant. Other functions have been suggested, but Walsby (1978) maintains that providing buoyancy is their primary purpose. Relatively little is known about the regulation of gas vesicle formation, Until recently there were few absolute measurements of cellular gas

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vesicle content. Gas vesicles can occupy as much as 10-20% of a cyanobacterium’s volume and GVP may comprise 6-15% of total cell protein (Oliver and Walsby, 1984; Utkilen et a f . , 1985; Konopka et al., 1987; Thomas and Walsby, 1985). Thus, a significant proportion of the biosynthetic resources can be invested in GVP and 20-50% of the total number of gas vesicles can be in excess of what is needed to provide buoyancy in cyanobacteria (Thomas and Walsby, 1985; Kromkamp et af., 1986). Presumably, there would be a strong selective advantage to regulating the synthesis of such a major protein. There are several instances where gas vesicle production is known to be regulated in cyanobacteria. Nosfoc muscurum filaments do not contain gas vesicles until hormogonium formation is induced (Armstrong et al., 1983). The gas vesicle content was almost 3-fold higher in Aplzanizomenon Jlos-aquae at low Jight-limitedgrowth rates than at high growth rates (Kromkamp et a f . , 1986). Now that a gup gene has been isolated (Tandeau de Marsac et af., 1985) and could serve as a probe of transcriptional expression from gup genes, sensitive measurements of changes in GVP production could be made. Regulation of gas vesicle formation has been easier to demonstrate in chemoheterotrophic bacteria than in cyanobacteria because gas vesicles are completely absent under repressing conditions and the cells transport exogenous organic compounds that can affect gas vesicle production. In A . aquaticus, gas vesicles were not formed in glucose salts media supplemented with L-lysine (Konopka, 1977). In several aerobic and facultatively anaerobic chemoheterotrophic bacteria, gas vesicles seem to be formed only if oxygen is required for the catabolism of the energy source and only when the oxygen concentration is suboptimal (R. L. Irgens and J. T. Staley, personal communication). However, there are species in which gas vesicles are produced under all growth conditions (Staley, 1968; Walsby, 1976). Most studies of buoyancy regulation in cyanobacteria have concentrated on short-term changes (that is, those that can occur within 1 day). The availability of light, inorganic macronutrients, and CO2 is the important environmental factor to which the regulatory mechanisms respond. It has been hypothesized that cells become more buoyant when the rate of photosynthetic energy generation exceeds the capacity to use that energy’ productively for growth (Konopka, 1984). There are two mechanisms by which organisms have been shown to lose buoyancy after exposure to high light intensities. The first is the collapse of weaker gas vesicles due to the 1-1.5 bar increase in turgor pressure observed when many cyanobacteria are shifted from light-limiting to light-saturating irradiances (Dinsdale and Walsby, 1972; Konopka et al., 1978; Konopka,

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1982; Oliver and Walsby, 1984). The rise in turgor in Anabaenajflos-aquae was due to the accumulation of low-molecular-weight organic compounds (Grant and Walsby, 1977) and K+ ions transported by a light-dependent pump (Allison and Walsby, 1981). This mechanism cannot operate in species where the pressure required to collapse the weakest gas vesicles exceeds the turgor pressure the cells can generate (3-5 bars). In these organisms a second mechanism was shown; the accumulation of polysaccharides could serve as “ballast” to compensate for the excess buoyancy provided by gas vesicles (Utkilen et al., 198s; Kromkamp and Mur, 1984; Thomas and Walsby, 1985). Kromkamp et al. (1986) have suggested that, even in species with weak gas vesicles, ballast accumulation will be the primary mechanism for buoyancy loss in nature. In cultures of Aphanizomenon jos-aquae grown under continuous light, there was a rapid rise in turgor and a relatively small rate of polysaccharide accumulation when irradiance was increased. In cultures grown under ecologically relevant light-dark cycles, the turgor rise was insufficienl to collapse gas vesicles and polysaccharide ballast accumulated rapidly. Polysaccharide synthesis and the accumulation of osmotically active molecules can be viewed as alternative mechanisms for dissipating excess energy. The physiological state of the organism will determine which mechanism predominates. Because gas vesicles collapse under pressure, they can be used as probes of intracellular turgor pressure in prokaryotes (Walsby, 1971). This was initially demonstrated in cyanobacteria (Dinsdale and Walsby , 1972). To test predictions of two theories of cell wall growth, the stress on gram-negative cell walls during the cell cycle was monitored in gas vacuolate cells and was found to be constant (Pinnette and Koch, 1988). By varying intracellular turgor, the elasticity of the cyanobacterial cell wall has been determined (Walsby, 1980). The strength of gram-negative cell walls has been estimated using gas vesicles (Hemmingsen and Hemmingsen, 1980). If the external pressure was raised slowly, the pressure could equilibrate across the gas vesicles and large pressures could be introduced into cells. When suspensions were rapidly decompressed, cells ruptured if the pressure was 50-100 atm. By determining the rate at which gas pressure had to be increased to collapse gas vesicles in heterocysts, Walsby (1985) was able to calculate the permeability coefficient of heterocysts to O2 and N 2 . Changes in turgor pressure, monitored using gas vesicles, can also be used to measure solute uptake in prokaryotes. Walsby (1980) found that sugar alcohols entered cyanobacteria by passive diffusion but that other substances, such as K’ and glycine betaine, were actively transported. Furthermore, K’ uptake was dependent upon light energy (Reed and Walsby, 1985).

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67

IV. Inclusions as Metabolic Products (Reserves) A. POLYGLUCOSE (GLYCOGEN) GRANULES The glycogen of prokaryotes is a polymer of D-glucose monomers linked by a-1,4-glucosidic bonds and branched through a-l,6-glucosidic bonds. Its synthesis and accumulation is generally most dramatic at the onset of, or during, unbalanced growth of a wide variety of bacteria and cyanobacteria when the energy and carbon supplies are in excess but some other critical nutrient is deplenished. Depletion of iron (Hardie et al., 1983a,b; Sherman and Sherman, 1983), nitrogen (Lehmann and Wober, 1976; Sigal et al., 1964; Stevens et al., 1981a; Zevenhuizen, 1966), phosphate (Zevenhuizen, 1966), or sulfur (Zevenhuizen, 1966) from a culture medium is usually sufficient to result in the accumulation of glycogen in either bacteria or cyanobacteria. However, exceptions have been reported (Eidels and Preiss, 1970; Konig et al., 1982). When sufficient glycogen accumulates intracellularly , it usually forms granules of variable diameters (typically 20-100 nm) with uneven to rough appearances that are readily visualized by thin-section electron microscopy. Numerous reviews (Dawes and Senior, 1973; Krebs and Preiss, 1975; Merrick, 1978; Preiss, 1969, 1972, 1973, 1978, 1984; Preiss and Walsh 1981; Preiss et al., 1983) on bacterial glycogen and a recent review (Allen, 1984) on cyanobacterial glycogen have appeared. According to recent compilations (Preiss and Walsh, 1981; Preiss, 1984) 47 species of bacteria and cyanobacteria have been reported to form glycogen-like reserves. Frunkia sp. HFPArl3 (Lopez et al., 1985) may be added to this list. In most bacteria, glycogen is freely dispersed within the cytoplasm. In cyanobacteria, glycogen granules are localized within the region of the photosynthetic thylakoid membranes (Stevens et ul., 1981a). Use of the PAT0 poststaining procedure greatly increases the contrast of glycogen granules in relation to the rest of the cytoplasmic contents of the cell (Hardie et al., 1983b; Stevens et al., 1981a). A new technique relying on the specific staining of glycogen on sections prepared from aqueousembedded material has promise as a direct analytical tool for this polymer (Westphal et al., 1985). The generally accepted scheme for the synthesis of glycogen begins with the formation of a sugar nucleotide in a reaction catalyzed by the enzyme ADP-glucose pyrophosphorylase (ATP:a-D-glucose- l-phosphate adenylyltransferase, EC 2.7.7.27) as follows (Preiss, 1984): ATP

+ a-D-glucose-1-P ADPglucose + PP,

The glucosyl unit is then transferred from ADPglucose to a preexisting a- 1,4-glucan or maltodextrin primer forming a new a-l,4-glucosyl bond.

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The reaction is catalyzed (Fox et al.. 1976) by an ADPglucose-specific glycogen synthase (ADPglucose: 1,4-a-~-glucan4-a-D-glucos yltransferase, EC 2.4.1.21) as follows: ADPglucose

+ I ,4-cu-o-glucan

1,4-~-o-glucosyI-glucan+ ADP

Branching of the 1,4-a-~-glucosyl-glucan(Greenberg el al. , 1983) is accomplished in a reaction catalyzed by branching enzyme (1,4-a-Dglucan: I ,Ca-D-glucan 6-a-D-(1.4-cw-D-glucano)-transferase, EC 2.4.1.18) forming 1,6-a-D-glucosyl bonds that account for about 10% of the total linkages found in glycogen. At least 46 strains of bacteria (Preiss, 1984) have been shown to contain significant levels of both ADPglucose pyrophosphorylase and glycogen synthase. Much less attention has been given to glycogen degradation than to its synthesis. The 1,4-a-D-glucosyl-glucan core of glycogen can be hydrolyzed (Chen and Segel, 1968a,b) by the enzyme glycogen phosphorylase ~1,4-a-D-glucan:orthophosphate a-D-glucosyltransferase, EC 2.4.1. I ) as follows: ( I .4-a-D-glucosyl)~+ P, e (1,4-a-D-glucosyl),I + a-D-glucose I-P

A soluble debranching enzyme (isoarnylase) has been found in a few bonds with bacteria (Palmer er al., 1973) that hydrolyzes 1,6-a-~-glucosyl the formation of maltodextrins as the major product. The combined action of maltodextrin phosphorylase and amylomaltase will convert maltodextrins into monomers of glucose, thereby completing the cycle of synthesis and degradation. The activity of ADPglucose pyrophosphorylase, which is the first unique enzyme in he pathway of glycogen synthesis, is subject to allosteric regulation (Preiss er al., 1983; Preiss, 1984). Seven different groups of bacteria can be formed based on the activator specificity of ADPglucose pyrophosphorylase. The four most common activators are fructose 6-phosphate, fructose- 1 ,Qbisphosphate, 3-phosphoglycerate and pyruvate. The activators tend, in general, to increase the apparent affinity of the substrates ATP and glucose I-phosphate for the enzyme. Depending on the source of the enzyme, the presence of AMP, ADP, or P, may inhibit the activity of ADPglucose pyrophosphorylase (Preiss 1984). Increasing the concentration of the appropriate activator may either reverse. or prevent inhibition by AMP, ADP, or P,. Thus, high rates of glycogen synthesis may occur only in the presence of excess carbon supply and at high energy charge, leading to questions about what prevents high rates of glycogen synthesis during the exponential growth of many bacterial cells.

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

69

Several compounds have been proposed as physiological inhibitors of glycogen synthesis during exponential growth. These include 5-amino-4imidazolecarboxamide ribonucleotide (Leckie et al., 198 l), ppGpp and pppGpp (Dietzler et al., 1979), and more recently, PP, (Preiss and Greenberg, 1983) and GTP (Dietzler et al., 1984). Resolution of this problem has been complicated by the stringent response of Escherichia coli. When a K-12 strain of E . coli is grown in a medium with a high concentration of valine, a severe limitation for isoleucine results (Leavitt and Umbarger, 1962) that triggers the stringent response (Cashel, 1975). The refA gene mediates the stringent response and is apparently required for glycogen to accumulate during amino acid starvation if glucose (Leckie et al., 1980; Taguchi et al., 1980) is the carbon source, but not if glycerol is the carbon source (Leckie et a f . , 1980). Levels of cAMP are high during growth on glycerol but low during growth on glucose (Epstein et al., 1975). This suggested to Leckie et al. (1980) that high cellular levels of cAMP might replace the refA gene requirement for glycogen accumulation during amino acid starvation. Leckie et af. (1985) have now shown that triggering the stringent response increased glycogen synthesis in mutants unable to synthesize cAMP and in mutants lacking cAMP receptor protein. Moreover, cAMP addition stimulated glycogen synthesis in relA mutant strains. Thus, cAMP and refA exert dual but independent regulation of glycogen synthesis in E. coli. They also showed that physiological concentrations of GTP inhibited ADPglucose pyrophosphorylase. Because the stringent response causes an abrupt decrease in the cellular level of GTP, Leckie et al. (1985) reasoned that modulation of the activity of ADPglucose pyrophosphorylase by GTP could account for most of the step-up in the rate of glycogen synthesis observed when the stringent response is triggered. Modulation of the activity of ADPglucose pyrophosphorylase by PPi may account for the remainder of the rate increase (Preiss, 1984). Transduction tests in E . coli K-12 have suggested that the genes for the glycogen biosynthetic enzymes are arranged in the order: mafA-gfgA (glycogen synthase)-gfgC (ADPglucose p yrophosphory1ase)-glgB (branching enzyme)-asd, at 75 minutes on the linkage map (Latil-Damotte and Lares, 1977). A 10.5-kb DNA fragment containing the glgA, B , C , and asd genes of E . coli K-12 has been cloned and substantially sequenced (Preiss, 1984). The available evidence suggests that at least the structural genes for the glycogen biosynthetic enzymes exist in an operon. A slightly different gene order has been reported for Salmonella fyphimurium (Steiner and Preiss, 1977). It is generally believed that glycogen serves as a readily mobilized reserve of intracellular carbon that can be rapidly transduced into

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biological energy under the appropriate conditons. Glycogen-containing cells might respond to a condition of more balanced nutrient supply more rapidly than their glycogen-free competitors. Glycogen accumulation has been adduced to support spore maturation in Streptomyces viridochromogenes (Brana et al., 1980) and Bacillus cereus (Slock and Stahly, 1974). Glycogen synthesis and degradation are thought to be important in the development of dental caries (Tanzer et al., 1976). As discussed elsewhere in this review, an important function of glycogen accumulation may be in the negative regulation of buoyancy where it may serve to offset the “excess” buoyancy of too many gas vesicles (Krompkamp and Mur, 1984; Thomas and Walsby, 1985; Utkilen et a l . , 1985; Van Rijn and Shilo, 1985). Another intriguing possible function for glycogen accumulation comes out of studies on the nitrogen starvation of the cyanobacterium Agmenellum quadrupficatum strain PR-6. Kollman et al. (1979) used nitrogen starvation of PR-6 in conjunction with 13C nuclear magnetic resonance (NMR) spectroscopy to follow the time course of polysaccharide accumulation. As cellular nitrogen decreased there was a concomitant decrease in the protein content of the cell and an increase in cellular D-glucose up to 38% of the dry cell weight. About midway in the time course of nitrogen starvation three lowmolecular-weight carbohydrates, glucosylglycerol, glucosylglycerate, and sucrose, became detectable by NMR. Kollman et al. (1979) noted that these compounds were probably involved in osmoregulation. Osmoregulation in cyanobacteria by low-molecular-weight carbohydrates is now firmly established (Borowitzka et al., 1980; Mackay et al., 1983a,b; Reed et al., 1984). Glycogen is the probable source of these low-molecularweight carbohydrates.

B. POLY-P-HYDROXYBUTYRIC ACIDGRANULES Pol y-P-hydroxybutyric acid (PHB) is a linear polyester of D-( -)-phydroxybutyric acid that is formed in a variety of bacteria, most notably by members of the genera Bacillus, Alcaligenes, Azotobacter, Beggiatoa, Spirillum, Sphaerotilus, Caulobacter, and Rhodobacter. PHB is not produced by plants, animals, or eukaryotic single-celled organisms; thus it is limited in its disposition to the prokaryotes. Phylogenetically, it may be significant that PHB is found in both eubacteria and archaebacteria (Fernandez-Castillo et a / ., 1986). The ability of bacteria to deposit PHB is considered a taxonomically important trait especially among the species of the genus Pseirdomonas (Dawes and Senior, 1973); however, the quantity of PHB deposited under certain growth conditions is physiologically and environmentally controlled.

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PHB is the ideal carbon and energy storage compound due to its insolubility, its lack of exertion of osmotic pressure on the cells, and its degree of reduction (Nickerson, 1982). PHB also has been considered as a potentially important biologically degradable bioplastic (Lafferty and Heinzle, 1977; Hardman, 1981). Several organisms can accumulate up to 5040% of their dry weight as PHB (Dawes and Senior, 1973); however, under certain growth conditions, Alcaligenes eutrophus can accumulate up to 96% of its dry weight as PHB (Pedros-Alio et al., 1985). Heinzle and Lafferty (1980) developed a mathematical model to describe the fermentation of A . eutrophus using C02, €32, and 0 2 substrates for production of PHB. Recently, Suzuki et al. (1986) developed a computer-controlled fermentation method for growing a methylotrophic Pseudomonas sp. to 233 g dry weight per liter, at which time the concentration of PHB in the fermentor was 149 g/liter. They obtained a maximal yield of 0.2 g PHB per gram of methanol utilized. Considering that Wakisaka et al. (1982) thought that their yield of 0.9 g/liter of PHB was significant, the results of Suzuki et al. (1986) are phenomenal. Their results further increase the potential for use of PHB as an industrially important bioplastic. PHB and similar p-hydroxy polymers are found free in soils, estuarine sediments, and ground waters (Findlay and White, 1983; White et al., 1983; Findlay et ul., 1985). High concentrations of PHB in oligotrophic environments have yielded the suggestion that PHB is important to bacterial survival in starvation conditions (White et al., 1983). PHB is the polymerized form of D-(-)-P-hydroxybutyric acid. The polymer has an empirical formula of (C4H602),,and a theoretical composition of 55.81% carbon, 7.03% hydrogen, and 37.16% oxygen (Dawes and Senior, 1973). PHI3 typically is soluble in hot chloroform or dichloromethane but not in alkaline sodium hypochlorite, benzene, or ether. The molecular masses of PHB vary from 1000 to 256,000 (Dawes and Senior, 1973) and the degree of polymerization can be calculated from the melting points, of which the highest reported is 188°C (Dawes and Senior, 1973). PHB is considered to be the simplest known biologically important polymer (Dawes and Senior, 1973; Nickerson, 1982), however, considering the recent studies of Findlay and White (1983), it is probable that “PHB” is not as simple as previously thought. PHB apparently is deposited in cells in crystalline fibrils. These polymer crystals were suggested to have “folded chain” lamellar morphology (Alper et al., 1963). Nicolay et al. (1982), however, recently observed with I3C NMR that the PHB of Rhodopseudomonas sphaeroides had a greater mobility in uiuo than in closely packed crystalline polymers. Moreover, Pedros-Alio et al. (1985) and Mas et al. (1985) showed that intact PHB granules are hydrated by about 40% of their weight (see below).

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Cornibert and Marchissault ( 1972) used X-ray and conformational analysis to derive a molecular model for PHB in which the molecule contained a compact right-hand helix with a 2-fold screw axis along the chain. Ellar et al. (1968) used light-scattering techniques to demonstrate a weight-average particle weight of 3.57 x lo9 glmol of particles, indicating that each PHB granule contains several thousand individual PHB molecules. Purified PHB has an apparent density of 1.19-1.25 g/cm3 (Nickerson, 1982; Mas et ul., 1985). Calculations of PHB density in uiuo yielded a figure of 1.1549 pg/pm’, considerably less than the value for purified polymer (Mas et al., 1985). From this difference, Mas et al. (1985) demonstrated that 40% of the PHB weight in uiuo is due to water. The accumulation of PHB by A . eutrophus increased the density of the organism from 1.100 pg/pm3 (non-PHB accumulating mutant) to 1.120 pg/pm3(wild type) (Pedros-Alio et al., 1985). The specific relationship between PHB content and cell density was hyperbolic. Accumulation of PHB also caused an increase in cell volume that was linear with respect to PHB content (Pedros-Alio el al., 1985). Mas et al. (1985) developed a mathamatical model to describe the relationships between PHB accumulation and cell mass. Wallen and Rohwedder (1974) demonstrated the presence of poly-phydroxyalkanoates other than PHB in samples of activated sludge. This was the first demonstration of any major potential heterogeneity in the lipid form of PHB. Findlay and White (1983) tested sediment samples as well as “PHB” from Bacilliis megaterium for the presence of polymers of P-alkanoates. They observed that B . megaterium “PHB” actually was composed of 95% PHB, 3% poly-/3-hydroxyheptanoate (PHH), trace amounts of poly-P-valerate (PHV), and three other uncharacterized poly-p-alkanoates (Findlay and White, 1983). Estuarine sediment samples included 30% of both PHB and PHV, 10% of PHH, and from less than 1% to 14% of eight other uncharacterized poly-p-alkanoates. This led the authors to propose that PHB be discarded in favor of poly-p-alkanoates (PHA) (Findlay and White, 1983). Furthermore, Findlay et af. (1985) have proposed the use of the PHAlphospholipidifatty acid (PLFA) ratio to measure the nutritional/metabolic status of microorganisms in estuarine sediments. Tunlid et a/. (1985) also used the PHB/PLFA ratio to show that root-colonizing bacteria were under balanced growth conditions whereas the same bacteria in nearby sands experienced unbalanced growth. PHB typically exists in cells as discrete inclusions of the hydrated polymeric ester and are bound by a proteinaceous single-layered envelope (Lundgren e l al., 1964; Dawes and Senior, 1973; Shively, 1974; Merrick,

-

FUNCTIONAL INCLUSIONS I N PROKARYOTIC CELLS

73

1978; Pedros-Alio et al., 1985). Intact PHB inclusions were purified and found to contain 97-98% PHB, -2% protein, and 0.5% lipid (Griebel et af., 1968). The protein envelope has a width of 2-8 nm, depending on the organism in which it is observed (Lundgren et al., 1964; Merrick, 1978). PHB synthetase is apparently a major component of the proteinaceous envelope (Griebel et al., 1968). Similarly, PHB depolymerases and a proteinaceous depolymerase inhibitory factor may be associated with the protein envelope layer (Shively, 1974; Merrick, 1978). Recently, PHB also was found to be a lipidic component of the membranes of certain bacteria (Reusch and Sadoff, 1983). Both the presence (Reusch and Sadoff, 1983) and the de nouo biosynthesis (Reusch et al., 1986) of PHB in bacterial cell membranes has been linked to the transformability of the bacteria by DNA. In those cases, the PHB was not present as an “inclusion” but rather as a biochemical polymeric component of the cell membrane (Reusch et al., 1986). The freeze etch structure of PHB, with its characteristic “pull-out” morphology, has been described in detail by Dunlop and Robards (1973). They developed a model of PHB granules showing a central core that occupies less than 50% of the granule volume and stretches during freeze etching. The central core is surrounded by a coat (Dunlop and Robards, 1973) that is enclosed in the envelope. Their analysis suggests that the inner core and the coat may represent different physical forms of the PHB polymer. PHB inclusions can be identified in whole cells by their distinctive appearance under phase and dark-field microscopy, their affinity with Sudan black B, and, more recently, by their orange fluorescence when stained with Nile blue A (Ostle and Holt, 1982). These, however, should only be considered as presumptive tests; further chemical analyses should be used to confirm the presence of PHB. PHB typically has been extracted by traditional chemical methods, i.e., the alkaline sodium hypochlorite method (Williamson and Wilkinson, 1958) or by differential extraction with chloroform (Dawes and Senior, 1973). Griebel et al. (1968) combined differential centrifugation, fractionation in a polymer two-phase system, and density gradient centrifugation to purify intact PHB granules. Nickerson (1982) recently developed a simple density gradient procedure using NaBr gradients for the purification of PHB, which yielded two purified fractions of PHB with densities of 1.19 and 1.23 g/cm3. The less dense fraction apparently still was associated with the protein envelope (Nickerson, 1982). PHB has been quantitated gravimetrically (cf. Dawes and Senior, 1973), turbidimetrically (Williamson and Wilkinson, 1958), and spectrophotometrically (Law and Slepecky, 1961). The spectrophotometric

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method (Law and Slepecky, 1961), which has been used most widely, recently has been updated so that PHB and polyphosphates can be measured simultaneously (Poindexter and Eley, 1983). Gas chromatographic methods have been devised recently that are more specific and sensitive (Braunegg et al., 1978; Findlay and White, 1983). The added advantage of these chromatographic methods is that they also separate the various poly-p-alkanoic acid derivatives (Findlay and White, 1983). Karr et al. (1984) also recently developed an ion exclusion highperformance liquid chromatography (HPLC) procedure to quantitate and analyze PHB from Rhizobium japonicirm. Other important methods used to measure the metabolism of PHB include "C isotope incorporation (Strohl et al . , 1981a) and I3C NMR, the latter being noninvasive and more informative (Nicolay et af., 1982; Fernandez-Castillo et al., 1986). Jacob et al. (1986) also recently used cross-polarization magic angle spinning (CPMAS) NMR to measure PHB directly in a pseudomonad. PHB from a variety of sources also had characteristic IR spectra with major absorption peaks at 1730-1735 cm-' (Dawes and Senior, 1973). X-Ray diffraction patterns on PHB from several different sources also snowed remarkable consistencies (Lundgren et al., 1965; Dawes and Senior, 1973).The ability to measure PHB by flow cytometry was recently demonstrated by Srienc et al. (1984), who showed that the light scattering due to PHB was significantly and measurably different from cytoplasmic material. This technique may be usable for studying the metabolism of PHB during fermentations in situ using flow-through technology. The synthesis and degradation of PHB have been the subjects of a great number of studies that are summarized by Dawes and Senior (1973), Shively (19741, and Merrick (1978). Since few data have been obtained on the metabolism of PHB since 1978, it will not be covered here. PHB typically is accumulated by bacteria in response to certain noncarbon-energy nutrient limitations and other conditions that may bring about unbalanced growth. Nitrogen limitation (Dawes and Senior, 1973; Suzuki et al., 1986), oxygen limitation (Senior et al., 1972; Ward et c i l . , 19771, or limitation by ions such as SO:-. Mg", Fe2+, or Mn2+ (Suzuki et nl., 1986) all have stimulated the accumulation of PHB by certain organisms. Limitation of Kc to Bacillus thuringiensis caused the accumulation of PHB whereas high potassium concentrations caused &endotoxin to be synthesized preferentially over PHB (Wakisaka et a l . , 1982), demonstrating the importance of ion balance in the control of PHB formation (Suzuki et al., 1986). Photosynthetic bacteria usually accumulate PHB in light conditions in the presence of hydrogen gas but sometimes under dark conditions after starvation (Sirevag and Casten-

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

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holz, 1979). The accumulation of PHB is thought to be due to high internal NAD(P)H concentrations, conditions that are found under most of the limitations (e.g., NH;, K', or 0,) shown above (Suzuki et al., 1986). The accumulation of PHB then would serve as an electron sink for the excess reducing power (Senior et al., 1972). White et al. (1983) recently showed that microbes in nutrient-poor groundwater contained the highest concentrations of PHB of any organisms studies in situ, suggesting an important role for PHB in the survival of those organisms under starvation conditions. Physiological studies also indicate that PHB is synthesized by several organisms such as Spirillum sp. (Matin et al., 1979), Sphaerotilus sp. (Stokes and Parson, 1968), Azospirillum brasilense (Tal and Okon, 1985), and Beggiatoa sp. (Strohl et al., 1981a) that thrive under oligotrophic conditions (Dawes, 1976; Merrick, 1978). In these cases, PHB recalcitrance apparently is part of a mechanism to stave off starvation (Merrick, 1978; Matin et al., 1979). Beggiatoa alba can accumulate up to 50% of its dry weight as PHB but when placed under starvation conditons the PHB was not metabolized significantly even after 4 days (W. R . Strohl, unpublished data). B . alba (Gude et al., 1981) and Spirillum sp. (Matin et al., 1979) also accumulated PHB in continuous culture under apparent energy and carbon-energy limitations, respectively. PHB accumulated in all of these organisms under conditions of excess oxygen and energy limitation suggesting that mechanisms controlling PHB accumulation in these organisms were different than for organisms using the PHB as an electron sink. In another example of PHB influence on survival, the presence of PHB in A . brasilense increased the resistance of that organism during starvation to several environmental stress factors such as UV irradiation, desiccation, and osmotic pressure (Tal and Okon, 1985).

C. CYANOPHYCIN GRANULES The cyanobacteria as a group are rich in subcellular structures and inclusions (Allen, 1984; Fuhs, 1968; Geitler, 1960; Jensen, 1984, Lang, 1968; Wolk, 1973). Among the latter, many investigators using light microscopy have noted conspicuous and highly light-refractive inclusions (Fritsch, 1945; Geitler, 1960) that have come to be called cyanophycin granules. Fogg (1951) used the Sakaguchi test on vegetative cells and heterocysts of A . cylindrica and noted that prominent granules in vegetative cells and polar plug material of heterocysts stained deeply with this reagent. He concluded that the stained material was cyanophycin and that cyanophycin contained arginine. Simon (1971) first isolated and chemically characterized cyanophycin granules showing them to consist

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of arginine and aspartic acid in equimolar concentration. Structural studies (Simon and Weathers, 1976) indicated that the cyanophycin granule polypeptide (CGP) consisted of a backbone of polyaspartate with an arginine residue linked by its a-amino group to the p-carboxyl group of aspartate leading to its designation as multi-L-arginyl-poly(aspartic acid). In its native state, it is suggested that CGP exists primarily as a p sheet structure (Simon et ul., 1980). Cyanophycin granules are quite variable in size with irregular margins, whether viewed in cross section by electron microscopy or within whole cells by light microscopy. Under some conditions of growth, cyanophycin granules may attain sizes of 1 .O pm in diameter in vegetative cells to nearly 2 pm in akinetes. A roughly spherical inclusion in this size range would occupy about 20-40% of the volume of a normal E . cofi cell. With some fixation protocols for electron microscopy, these granules appear as electron-transparent regions in the cell that have the rough appearance of a vacuole (Lang, 1968). With other fixatives and poststaining procedures, they have a distinctively structured appearance and look, by analogy, rather like dried prunes. CGP is widely distributed among cyanobacteria although probably absent from a few species (Codd e f al., 1979; Lawry and Simon, 1982). It also appears to be a polymer unique to the cyanobacteria. Members of unicellular, filamentous, and true-branching groups of cyanobacteria have been shown to produce cyanophycin granules. Although they misidentified the structures, Van Baalen and Brown (1969) observed cyanophycin granules in the planktonic cyanobacterium Trichodesmiurn erythraeum. Especially prominent cyanophycin granules are formed in akinetes (Braune, 1980; Clark and Jensen, 1969; Miller and Lang, 1968; Sutherland er al., 1979; Wildman et al., 1975) and in motile hormogonia (Castenholz, 1982; Hernandez-Muniz and S. E. Stevens, Jr., unpublished). Cyanophycin granules have been observed in the cyanobacterial partner of the symbiotic associations between cyanobacteria and both lower and higher plants (Grilli-Caiola and de Vecchi, 1980; Honegger, 1980; Neumuller and Bergman, 1981; Obukowicz et al.. 1981; Sharma e? af., 1982; Spector and Jensen, 1977). Rodgers and Stewart (1977) noted the absence of cyanophycin granules in the Anfhoceras punctatus-Nostoc and Blasia pusillu-Nostoc associations but Honegger (1980) observed many of these granules in the Nosfoc symbiont of both liverworts in material collected in Iceland in September. Honegger explained the difference by suggesting that a slowed metabolism of the plant partner, due to shorter day length, lowered the nitrogen demand of the plant and allowed the cyanobacterial partner to accumulate nitrogen in the form of CGP. Obukowicz et al. ( 19811 independently confirmed Honegger’s explanation in the Cycas

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revoluta-Anabaena symbiosis. Even the cyanobacteria found in the hollow hairs of green polar bears have cyanophycin granules (Lewin et a f . , 1981). Unbalanced growth of cyanobacteria seems to be the requisite condition for accumulation of CGP and formation of cyanophycin granules. CGP is at a low to undetectable level during the balanced growth of cyanobacteria (Allen, 1984). However, it accumulates in response to starvation for light (Allen et al., 1980; Van Eykelenburg, 19791, phosphorus (Allen et al., 1980; Lawry and Simon, 1982; Stevens e t a f . , 1981b), or sulfur (Allen et a f . , 1980; Lawry and Simon, 1982). Inhibitors of RNA and protein synthesis such as rifamycin (Rodriques-Lopez et al., 1971), rifampin (Lawry and Simon, 1982), and chloramphenicol (Ingram et al., 1971; Obukowicz and Kennedy, 1980; Simon, 1973b) cause CGP accumulation. Likewise, 5-methyltryptophan, casein hydrolysate, urea, ammonium chloride, and cysteine induce CGP accumulation in some cyanobacteria (Lawry and Simon, 1982). A temperature that is suboptimal for growth induces CGP accumulation in Spirulina platensis (Van Eykelenburg, 1979) but not in Aphanocapsa 6308 (Allen et a f . , 1980). Stressed growth in the presence of sulfur dioxide also leads to CGP accumulation (Sharma et al., 1982). An enzyme, denoted trivially as Arg-poly(Asp) synthetase, which catalyzes the addition of arginine and aspartic acid to a preformed primer of CGP, was purified by Simon (1976) and confirmed by Gupta and Carr (1981a). The enzyme activity was insensitive to inhibitors of protein synthesis and nucleases. These results were supportive of a previous interpretation of an in viuo study (Simon, 1973b) concluding that CGP was nonribosomally synthesized. Stevens et a f .(1981b) and Allen and Hawley (1983) have suggested that CGP is synthesized from the turnover of cellular protein. An exopeptidase given the trivial name cyanophycinase was observed in extracts of Anabaena sp. (Gupta and Carr, 1981a). The substrate of cyanophycinase was CGP and the product of the reaction was the dipeptide aspartic acid-arginine. Allen et al. (1984) also observed a CGP peptidase in extracts of Aphanocapsa 6308 but the products of hydrolysis were primarily free arginine and possibly some aspartic acid-artinine dipeptide. Early workers suggested that cyanophycin granules were reserve or storage materials (summarized by Fritsch, 1945; and Geitler, 1960). The appearance of the granules in A . cylindrica during the stationary phase of growth and their subsequent disappearance upon dilution into fresh medium (Simon, 1973a) supported this notion. Stanier and Cohen-Bazire (1977) suggested that cyanophycin granules also might serve as energy

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M. SHIVELY ET AL.

reserves if the arginine dihydrolase pathway was present in cyanobacteria. Weathers er ul. (1978) and Gupta and Carr (1981b) have confirmed the presence of the arginine dihydrolase and arginase pathways for the catabolism of arginine in Aphanocapsa 6308 and two Anahaena sp. The presence of these two catabolic pathways and the reportedly higher activity of cyanophycinase relative to Arg-poly(Asp) synthetase led Carr (1983) to postulate a dynamic role for CGP accumulation, in which CGP was continuously synthesized and degraded to ornithine and citrulline. The extenstive I4C radiolabeling of citrulline first observed by Norris et al. (19%) and identified by Linko ef a f . (1957) was adduced as further support for the hypothesis of Carr (1983). Removal of free arginine by synthesis of CGP might be necessary for continued arginine biosynthesis because of the strong feedback inhibition exerted by this amino acid on its own synthesis (Hoare and Hoare, 1966).

D. POLYPHOSPHATE GRANULES Polyphosphates are linear polymers of orthophosphate residues linked by energy-rich phospho-anhydride bonds. They vary widely in size from two residues in pyrophosphate to perhaps thousands of residues in polyphosphate granules. Upon purification from the bacterium Desulfuuibrio gigas, these granules proved to be composed of magnesium tripolyphosphate (Jones and Chambers, 1975). In may bacteria and cyanobacteria, polyphosphates accumulate rapidly in cells grown in phosphate-replete media, a situation accentuated by a prior limitation of phosphate (Healey , 1982). The rate of polyphosphate accumulation is generally several times that of the cellular growth rate. At the onset of phosphate starvation, polyphosphate granules disappear with time indicating their ready mobilization for the metabolic needs of the cell. Rapid accumulation during times of plenty and rapid disappearance during times of need support the notion that polyphosphate granules serve a reserve or storage function. However, there are reports of bacteria that do not synthesize polyphosphates during exponential growth but do so only under conditions of unbalanced growth (Merrick, 1978; Kulaev and Vagabov. 1983). Polyphosphate granules are easily visualized by thin-section electron microscopy but their appearance varies considerably depending on fixation and poststaining procedures (Jensen et a l . , 1977). Recently, polyphosphate granules were visualized in air-dried, unfixed cyanobacteria (Jensen and Baxter, 1985). They have been analyzed by X-ray microprobe (Kessel, 1977) and through use of 3'P NMR spectroscopy (Florentz et ul., 1984). They are also readily characterized by chemical or

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enzymatic means (Kulaev and Vagabov, 1983; Poindexter and Eley, 1983). Polyphosphate accumulation into granules is very widespread among both bacteria and cyanobacteria (Kulaev and Vagabov, 1983; Allen, 1984). Recently, members of the archaebacteria also were shown to contain polyphosphate granules (Scherer and Bochem, 1983). Most of the bacterial isolates from subsurface aquifers were shown to accumulate polyphosphates (Balkwill and Ghiorse, 1985). In general, polyphosphates are associated with the nuclear region of prokaryotic cells. Application of extensive serial thin-sectioning and computer graphics techniques allowed Nierzwicki-Bauer et al. (1983) to unequivocally place polyphosphate granules within the central nuclear core of a cyanobacterial cell. In E . coli, phosphate is transported by two independent systems termed Pit and Pst (Rosenberg et al., 1984). The Pit system is energized by the proton motive force whereas the Pst system and its periplasmic-binding protein are energized by chemical bond energy. Moreover, Pit is a low-affinity, high-velocity transport system in contrast to Pst, which is a high-affinity, low-velocity system (Willsky and Malamy, 1980). The Pst system is thought to consist of the closely linked genes pst, phoS, phoT, and probably a fourth gene termed phoU (Zuckier and Torriani, 1981). The phoU genotype renders E . coli constitutive for alkaline phosphatase without the requirement for an organic phosphorous source such as glycerol 3-phosphate. Rao et al. (1985) took advantage of this to show that E. coli grown anaerobically could accumulate polyphosphates of at least 200 residues in length. They also used NaF, a competitive inhibitor of the acid phosphatase found in E. coli (Dassa and Boquet, 1981) during extraction, fractionation, and analysis, to prevent polyphosphate degradation. Consideration of the synthesis and degradation of polyphosphates is complicated by the number of enzymes that may be involved and by the fact that five of these enzymes are reversible under physiological conditons. The first enzyme described was polyphosphate kinase, which catalyzed the following reaction: ATP

+ (phosphate), e ADP + (phosphate), +

Other enzymes involved in the transfer of an orthophosphate residue are po1yphosphate:AMP phosphotransferase, polyphosphate-dependent NAD' kinase, polyphosphate glucokinase, and 1,3-bisphosphoglycerate:polyphosphate phosphotransferase (Kulaev and Vagabov, 1983). In addition, polyphosphate depolymerase and a group of enzymes called polyphosphatases hydrolyze polyphosphates. These last two enzymes are

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degradative in activity whereas the previous five may be synthetic or degradative depending on the physiological state of a cell. Many bacteria and cyanobacteria react to the addition of phosphate following phosphate starvation by its very rapid intracellular accumulation. This has been termed the “phosphate overplus” phenomenon. The rapid accumulation of intracellular phosphate has been associated with nuisance blooms of cyanobacteria in many inland lakes and also may be important in the metabolism of microorganisms in wastewaters. Species of Acinetobacrer, Aerornonas, and Pseudomonus have been shown to accumulate polyphosphates to as much as 10-2096 of their dry weight during wastewater treatment (Lotter and Murphy, 1985). It is also of interest that several metals are sequestered in polyphosphate granules (Jensen et al.. 1982; Rachlin et al., 1985).

E. SULFUR GLOBULES Although a wide variety of prokaryotic microorganisms can metabolize elemental sulfur (Lackey rt al., 1965; Laishley et ul., 1986), including heterotrophic organisms (Okada et al., 1982; Wainwright et al., 1984), only a limited number of organisms deposit sulfur internally. The term “internally,“ as used here, denotes deposition of sulfur globules within the confines of the cell wall. This distinguishes organisms like Chromatium sp. or Beggiatoa sp., which deposit sulfur within internal characteristic globules, from organisms such as Thiohacillus, Thiomicrospira, or Chlorobiirm. which can synthesize sulfur but do not deposit it within their cell walls (La Riviere and Schmidt, 1981; Van Gemerden, 1984). Sulfur globules are the only prokaryotic “inclusions” that appear in markedly different morphological forms, depending on the organism in which they are observed (Table 111; Fig. 5). Sulfur-depositing bacteria fall into three categories. The first group is represented by the purple photosynthetic sulfur bacteria, including organisms within the genera Chrornatium, Thiocupsa, and Thiocystis but excluding members of the genus Ectothiorhodospira (Truper, 1978). The colorless, filamentous gliding sulfur bacteria, including members of the genera Beggicitoa (Maier and Murray, 1965; Strohl et al., 1981b, 1982), Thiorhrir (Bland and Staley, 1978; Larkin and Shinabarger, 1984; Nielsen, 1984), Thioploca (Maier and Murray, 1965; Maier and Gollardo, 1984), and Thiospirillopsis (Lackey et al., 1965), make up the second group. Included also in this second group are the Thiothrix-like activated sludge bacteria designated as “Type 021”’ (Williams and Unz, 1985). The final group of sulfur bacteria consists of “morphologically conspicuous” coiorless sulfur bacteria, including organisms of the genera Achromatium,

TABLE I11 INTERNAL SULFUR DEPOSITS IN BACTERrA

Type of

Groups and genera of bacteria Colorless filamentous (&ding) bacteria Beggiafoa Thiothrir Thioploca (freshwater)

Thiuploca (marine) Thiospirillopsis 021N-Type Purple photosynthetic bacteria Chrornatiurn

sulfur deposita

CMEP-ENV

Method of observationb

TS and FE

CMEP TS CMEP TS CMEP or UMC TS ND LMO ND LMO

References

Strohl et al. (1981b); Strohl el al. (1982) Bland and Staley (1978) Maier and Murray (1965) Maier and Gallardo (1984) Lackey et al. (1%5) Williams and Unz (1985)

SMC

TS, FE, and P

CMAP-ENV

TS

Schmidt and Kamen (1970); Hageage ei al. (1970); Nicolson and Schmidt (1971); Remsen and Truper (1973); Remsen (1978) Eimhjellen et al. (1967)

SMC

TS

Remsen (1978)

Morphologically conspicuous nonphotosynthetic sulfur bacteria Thiovulurn SMC

TS

Faure-Fremiet and Rouiller (1958); deBoer et nl. (1961); Wirsen and Jannasch (1978) La Riviere and Schmidt (1981) La Riviere and Schmidt (1981) La Riviere and Schmidt (1981) La Riviere and Schmidt (1981)

Thiocapsa (Thiococcus) Thiocystis

A chromafiurn

ND

LMO

Macrornonas

ND

LMO

Thiobacteriurn

ND

LMO

Thiospira

ND

LMO

a CMEP, sulfur deposit located outside of cytoplasm in a pocket formed by invaginated cytoplasmic membrane (no other envelope observed); CMEP-ENV, sulfur deposit bound by a distinct (usually single-layered) envelope located outside of cytoplasm in a pocket formed by invaginated cytoplasmic membrane; UMC, sulfur deposit located in the cytoplasm and enclosed within a unit-type membrane; SMC, sulfur deposit located in cytoplasm and bound only by a single-layered envelope; ND, fine structure of inclusion not done. * TS,thin-section electron miscoscopy; FE, freeze etch electron microscopy; P, sulfur deposits purified; LMO, sulfur inclusions viewed only by light or phase microscopy.

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0

CYTOPLASM^

SMC

a

CMEP-ENV

FIG. 5. Model showing the four different types of internal sulfur deposits (S) found in bacteria. CW, cell wall; CM, cell membrane. Refer to Table 111 for the abbreviations for each sutfur inclusion type and the organisms in which they are found.

Mucromonus, Thiobacteriurn, Thiospira, and Thiovulum (La Riviere and Schmidt, 1981). Sulfur deposition and sulfur metabolism biochemistry are best characterized with the photosynthetic sulfur bacteria, primarily because pure cultures of these bacteria have been available longest. Several strains of Begqiatoa and Thiothrix have been available for study in the past 10 years, whereas members of the genus Thioploca and all of the “morphologically conspicuous” bacteria (La Riviere and Schmidt, 1981) remain unpurified. Determining whether or not the photosynthetic sulfur bacteria contain representative sulfur globules will require further purification of strains and further analysis of sulfur inclusions from less known strains.

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

83

About 30 solid allotropes of elemental sulfur exist, although the most common and stable form under normal temperature and pressure is orthorhombic sulfur, which exists in the Sg ring form (Roy and Trudinger, 1970). Intracellular elemental sulfur from Chromatiurn was determined to be arranged in “spherically symmetrical aggregates of radially arranged arrays of S: molecules” that were in a “liquid” modification (Hageage et al., 1970). When these liquid sulfur globules were dried, they slowly passed through a previously uncharacterized unstable crystallized state until they were completely converted to crystalline orthorhombic sulfur (Hageage et al., 1970). Dried preparations of sulfur from the bacteria Thiovulum majus (La Riviere, 1963), as well as Thiocystis violacea and Chromatium 81 1 1 (Triiper and Hathaway, 1967), also were observed to be in the orthorhombic crystalline state. Elemental sulfur has a density of 1.957 g/cm3for monoclinic crystals to 2.07 g/cm3 for rhombic crystals (20°C) as determined by specific gravity (Weast, 1972). The density of non-sulfur-containing bacteria ranges from 1.05 to 1.30 g/cm3 depending on the species (Guerrero et al., 1984). The density of sulfur in globules extracted from two Chromatium species, however, was calculated to be 1.219 g/cm3 (Guerrero et al., 1984), far below the density of orthorhombic sulfur. This led Guerrero et a f . (1984) to postulate that the deposited sulfur was complexed with another, less dense component. They further suggested that if the globules contained “hydrated sulfur,” then the degree of hydration would be 65% of the wet weight of the sulfur globule. Although orthorhombic sulfur typically is not characterized as water soluble (Roy and Trudinger, 1970), hydrophilic forms of elemental sulfur exist that apparently are metabolically active (Roy and Trudinger, 1970). Moreover, Laishley et al. (1986) recently showed that the rate of sulfur metabolism was dependent on the molecular crystalline structure of the sulfur. Taken as a whole, these observations are consistent with the concept of microbial deposition of intracellular “wetted” (Hageage et al., 1970), “hydrated” (Guerrero et al., 1984), and hydrophilic elemental sulfur species. Moreoever, morphological evidence using polarized microscopy (Hageage et af., 1970) and freeze etch microscopy (Remsen, 1978; Strohl et al., 1981b) supports the premise that intracellularly deposited sulfur is not a form that is typical of orthorhombic sulfur. Sulfur can comprise -20% (Nelson and Castenholz, 1981) to 25% (Guerrero et al. 1984) of the cell dry weight of Beggiatoa sp. and Chromatium sp., respectively. Deposition of sulfur by cells of Chromatium vinosum increases their density from 1 . 1 150 to 1.2281 g/cm3(Mas et af., 1985). For C. warrningii, the depositions of sulfur increased the density of the cells from 1.0890 to 1.1321 g/cm3 (Mas et al., 1985) and for

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B. alba BI8LD, sulfur deposition increased the density from 1.095 to I . 115 g/cm’ (Schmidt, 1985). These increased densities could have ecological as well as physiological consequences on the cells (Guerrero et al., 1984). The sulfur globules appear highly refractile in light and denser than cellular material when unfixed organisms are placed under an electron beam (Nicolson and Schmidt, 1971; Lawry et al., 1981). Both of these properties may be attributed, in part, to the density of the sulfur globules as well as to the state of the sulfur contained within. Sulfur globules are extractable by carbon disulfide, acetone, ethanol, benzene, pyridine, and a variety of other organic solvents (Windholz, 1983). Methods for extraction and measurement of deposited sulfur are described by Van Gemerden (1968) and Nelson and Castenholz (1981). Vargas and Strohl (1985), Schmidt et al. (1986), and Strohl er a f . (1986) describe methods for determining sulfur deposition by Beggiatoa using Na?5S isotope. This latter method allows for the determination of sulfur deposition rates as well as the calculation of total sulfur content. Sulfur globules from Chromatium were purified by repeated low-speed centrifugation after breakage of the cells (Schmidt ef al., 1971; Guerrero er al., 1984). Partial purification of B. alba sulfur globules also was achieved after breakage by sonication in the presence of lysozyme, DNase, RNase, and phospholipase, followed by low-speed centrifugation washes (Schmidt et al., 1986). Further purification of the sulfur globules may be achieved by density gradient centrifugation using PercoU or a similar matrix (Guerrero et al., 1984). In thin-section electron micrographs, sulfur deposits are identified by a conspicuous, empty, and electron-translucent space surrounded by an envelope or membrane (or both) (Strohl et al., 1986). The sulfur is dissolved from the space during dehydration (Schmidt and Kamen, 1970; Remsen and Triiper, 1973; Strohl ef af., 1981b). Freeze etch preparations of sulfur inclusions show a characteristic smooth appearance when the cleavage plane passes through the sulfur (Remsen and Triiper, 1973; Remsen, 1978; Strohl P t al., 1981b). Occasionally, the cleavage plane passes along the outer edge of the sulfur globule, yielding some information about the limiting membrane (Remsen and Truper, 1973; Strohl et al., 1981b). The sulfur inclusions of Chromatiurn are bound by a single electrondense envelope of 2.5-3 nm (Schmidt et a f . , 1971). This envelope is constructed from a single peptide with a molecular mass of 13,500. After solubilization and subsequent reaggregation of this protein, it formed sheets (Schmidt et al., 1971). It was postulated that the envelope

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

85

functioned as a barrier to separate the sulfur from the cellular constituents, as well as to provide binding sites for potential enzymes involved in sulfur metabolism (Schmidt et al., 1971). The sulfur globules and envelopes of Chromatium were found closely associated with the chromatophores (Nicolson and Schmidt, 1971; Remsen and Triiper, 1973; Remsen, 1978). It is apparent from several studies that the sulfur globules in Chromatium are located in the cytoplasm, separate from and internal to the cytoplasmic membrane (Schmidt and Kamen, 1970; Nicolson and Schmidt, 1971 ; Remsen and Truper, 1973; Remsen, 1978; Van Gemerden, 1984). In order for Chromatium to deposit sulfur from sulfide, the sulfide must traverse the cytoplasmic membrane and enter the cell (Van Gemerden, 1984). Van Gemerden (1984) developed a model to describe this oxidation system and showed that photosynthetic bacteria that deposited sulfur inside the cells had lower affinities for sulfide than those that deposited sulfur outside the cells. The sulfur globules of Beggiatoa (Morita and Stave, 1963; Maier and Murray, 1965; Strohl et al., 1981b, 1982; Lawry et al., 1981), Thiothrix (Bland and Staley, 1978; Larkin and Shinabarger, 1983; Nielsen, 1984), Thioploca (Maier and Murray, 1965; Maier and Gallardo, 1984), Thiovulum (Faure-Fremiet and Rouiller, 1958; Wirsen and Jannasch, 1978), Thiocapsa (Eimhjellen et al., 1963, Thiocystis (Remsen, 1978), and Achromatium (deBoer et al., 1971) all have been observed by thinsection electron microscopy (Table 111). In some of these cases, it is difficult to discern the actual structure of the surrounding envelope and (or) membrane. However, where enough information is available, the data suggest that four types of sulfur deposits may be formed by the different organisms (Table 111; Fig. 5). The sulfur globules may be enclosed within a single electron-dense envelope (Strohl et al., 1981b; Lawry et al., 1981; Nicolson and Schmidt, 1971); the envelope may be located directly in the cytoplasm (Schmidt and Kamen, 1970;Remsen and Triiper, 1973) or in the periplasm within an invagination of the cytoplasmic membrane (Fig. 5 ; Lawry et al., 1981; Strohl et al., 1981b). Similarly, Maier and Murray (1965) described sulfur globules that were delineated by the invaginated cytoplasmic membrane alone. These too were located external to the cytoplams but internal to the cell wall. In some cases, the sulfur deposition may appear as a unit membrane-bound inclusion in the cytoplasm with no apparent connection to the cytoplasmic membrane (Strohl et al., 1981b; Maier and Gallardo, 1985; Fig. 5). This type of structure may be observed as a result of the sectioning plane, as shown by Strohl et al. (1986), or may be the result of sulfur globule formation in the cytoplasm, analogous to the inclusions in Chromatium (Remsen and Triiper , 1973).

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In the characterization of the sulfur globules of Beggiatoa strain B 15LD, Strohl et al. (1982) observed two interesting features of the sulfur inclusions in this strain. First, the sulfur globules were surrounded by an unusual 12 to 14-nm-thick pentalaminar envelope consisting of three electron-dense layers, 3.5, 2.1, and 3.5 nm thick (Strohl et a f . , 1982). Second, rudimentary structures of these unusual envelopes also were observed in cells grown in the complete absence of reduced sulfur (Strohl et al., 1982). It was hypothesized that these envelopes were present in collapsed form until a reduced sulfur source became available. Upon exposure to sulfide, the envelopes would quickly expand to deposit the sulfur from the oxidation of sulfide (Strohl er al., 1982). Sulfide oxidation to deposited sulfur by B. alba Bl8LD recently was shown to be constitutive, further indicating the need for premade sulfur globule envelopes (Schmidt et a f . , 1986). The sulfur inclusions of B. ulbu B18LD are enclosed by a single 3 to 4-nm-thick electron-dense envelope that is located outside of the cytoplasmic membrane within invaginations (Strohl er al., 1981b). Partial purification of the B. albu sulfur inclusions led to the enrichment of a few peptide bands visualized by SDS-PAGE (Schmidt et al., 1986). A band corresponding to an approximate M , of 15,000 was enriched with the sulfur inclusions and was the major protein synthesized in response to addition of sulfide (Schmidt et al., 1986). Because sulfide oxidation is constitutive in B. alba. this suggests that this peptide may be important to the structure of the B. alba sulfur globule. As more sulfur is deposited, there may be a need for increased sulfur globule envelope synthesis to enclose the deposited sulfur. The biochemical mechanisms for the oxidation of sulfide to sulfur are described in detail elsewhere (Triiper, 1978; Truper and Fischer, 1982; Fischer, 1984). In brief, it appears that for C. uinosum a flavocytrochrome c552 accepts electrons from sulfide with sulfur as the product (Fukumori and Yamanaka. 1979; Gray and Knaff, 1981). The oxidation of sulfide by Chromatiurn is apparently constitutive (Van Gemerden, 1984) and Chromaiium also is capable of reducing sulfur to sulfide under anaerobic conditions in the dark (Van Gemerden, 1968b). Beggiaroa couples the oxidation of sulfide with oxygen (Vargas and Strohl, 1985; Schmidt et al., 1988). The oxygen-dependent oxidation of sulfide to sulfur by B. alba B18LD can be inhibited by several electrontransport inhibitors although it has not been demonstrated whether or not a coupling site is involved (Strohl and Schmidt, 1984; Schmidt el al., 1988). Recently, Hooper and Dispirit0 (1985) described a mechanism for external oxidation of inorganic molecules. This model would fit the oxidation of sulfide by organisms by Beggiatoa (Strohl and Schmidt, 1984; Hooper and DiSpirito, 1985). B . alba B18LD apparently cannot

FUNCTIONAL INCLUSIONS IN PROKARYOTIC CELLS

87

oxidize the deposited sulfur to sulfate, a biochemical mechanism that can be carried out by Thiothrix and Chromatium (Schmidt et al., 1988). Instead, Beggiatoa, like Chromatiurn, can reduce sulfur to sulfide in the absence of oxygen (Nelson and Castenholz, 1981; Schmidt, 1985; Schmidt et a / ., 1988). This may have ecological significance because Beggiatoa sp. live at the interface of the oxic-anoxic zone in which they are exposed to sulfide from below and oxygen from above (Jorgensen and Revsbech, 1983). Under conditions where this zone moves, the beggiatoas can experience time periods of anoxia in which they would require an internal electron-acceptor source, particularly because they do not reduce nitrate in a dissimilatory manner (Vargas and Strohl, 1985). V. Concluding Remarks

If the information in this review is compared to that of earlier reviews (see individual sectons for references), various degrees of progress in the understanding of prokaryotic inclusions will be found. Obviously, the amount of progress depends on many factors including the interest of researchers and the availability of support. Furthermore, the most significant and rapid advances have been made by researchers using all available avenues and tools, e.g., cytological, nutritional, physiological, biochemical, and genetic. Although good, “hard-nosed” biochemistry has been a major contributor in some instances, the use of genetics and recombinant DNA technology have been especially fruitful in adding a new dimension to this understanding. The genes encoding the structural components of the phycobilisome and gas vesicle as well as the genes for the enzymes involved in glycogen biosynthesis have been cloned and sequenced. Attention being given to these inclusions is now being directed toward the regulation of gene expression. This is likely to involve complex processes in all of these structures, but especially so in the phycobilisome. Many questions will need to be considered. How do light intensity and wavelength as well as nutrient availability regulate phycobiliprotein synthesis? How is the correct phycobiliprotein-linker polypeptide stoichiometry achieved? How is the synthesis of chromophores and apoproteins coordinated? How and when do posttranslational modifications occur, e.g., chromophore attachment? Several laboratores are pursuing these and other regulatory questions; significant progress can be expected in the near future. For the gas vesicle, in addition to experiments dealing with regulation, an intriguing question sure to be addressed is how this supramolecular

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structure is constructed such that water and other materials are excluded from the interior. Since the enzymes of glycogen biosynthesis have been cloned and sequenced, many new avenues of research can be undertaken. The enzymes can be produced in much larger quantity allowing for much greater detail in the study of their protein chemistry. Site-directed mutagenesis will provide valuable information relating to the molecular mechanisms of catalysis and allosteric regulation. Obviously, the molecular approach needs to be applied to the study of the other prokaryotic inclusions. Experimentation has already been initiated in many instances. Proteins will be isolated and sequenced; polynucleotide probes will be made and genes isolated, sequenced, and expressed in a suitable organism. The organization of the genes for a given inclusion will be determined and may be found to be clustered on the chromosome. Ultimately, it would be of interest to show the expression of a total inclusion, e.g., the carboxysome in an organism like E. coli. Basic research on the inclusions is likely to pay considerable dividends in the industrial sector where useful products and/or processes are the major goal. For example, PHB and other recently discovered poly-palkanoates are degradable bioplastics. PHB can now be produced at over 150 g/liter. which makes it economically feasible. The properties of PHB as well as the other poly-/3-alkanoates need to be carefully determined. The genes for their synthesis can then be engineered to improve these properties. A whole new level of research will be fostered using the molecular approach. Many unanswered questions will be answered and many new questions posed for tomorrow’s researchers.

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INTERNATIONAL REVIEW OF CYTOLOGY, VOL. I I3

Microtubules in Cardiac Myocytes L. RAPPAPORTAND J. L. SAMUEL INSERM U 127, H6pital Larihoisisre, 75010 Paris, France

1. Introduction Components of the cytoskeleton-mainly referred to as actin filaments, intermediate filaments, and microtubules-reflect the distinct spatial geometry of the eukaryotic cells, which is recognized by the position of the organelles. Microtubules (MTs) have mainly been studied in brain and in cultured cells, whereas very few studies have been carried out on adult cardiac myocytes in which MTs are present in small numbers. MTs, like actin filaments, are made up of globular protein subunits (a- and ptubulin) that can assemble and disassemble rapidly in the cell. Tubulin, the more prominent protein in MTs, is ubiquitous, whereas other proteins associated with the MTs-microtubule-associated proteins (MAPS)appear to be more tissue specific. The MAPS regulate MT assembly from pools of unpolymerized tubulin subunits and influence the MT interactions with other cellular structures. Mechanisms for controlling MT assembly are crucial since MTs are generally considered to have a unique part in determining the overall organization of a cell. A major constituent of the mitotic spindle, MTs influence the distribution of actin filaments, especially during the process of cell division, and of intermediate hlarnents during cell interphase. A wide variety of studies using pharmacological and histochemical techniques attest that MTs play an essential role in the maintenance of cellular shape, in cell movement, and in cellular polarity. Evidence for a role of MTs as a temporary scaffold to organize the other cytoplasmic components emerged from studies carried out on plant cells, developing sperm tail, and insect flight muscle in the early developmental stages. However, most of these processes are not directly relevant to adult cardiac myocytes that do not divide and whose shape, polarity, and movements appear to depend mainly on the highly organized arrangement of the myofibrils. Participation of MTs in the intracellular processes of mature cardiac myocytes was analyzed using the model of cardiac hypertrophy in the rat. Cardiac hypertrophy is one of the adaptive responses of the heart to the increase in hemodynamic load that develops during postnatal life or when the overload is repeated or long lasting. Striking changes in intracellular 101

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architecture and arrangements of organelles occur during the early period following cardiac overload. Some characteristics of MTs in adult heart and the distribution of MTs within myocytes have been described and the involvement of MTs in several cellular processes has been analyzed. It emerges that MTs, although few in number, are, as in embryonic or neonatal cells, involved in the organization of adult myocytes.

11. Proteins Constitutive of Heart Microtubules

The study of MTs in adult cardiac muscle has been hampered by their relatively low abundance (Limas, 1979; Samuel et al., 1983b; Rappaport ct a l . , 1986). Recently, improvement in purification techniques has made possible the analysis of MT components in this tissue. Analysis of MT proteins is of interest since as a result of recent studies carried out in other tissues, it emerged that MAPs are tissue specific and are involved in defined functions of MTs (Olmsted et al., 1986) and that tubulin exists in many isoforms, the presence of which varies with development (Ginzburg and Littauer, 1984; Denoulet et a / . , 1982).

OF MICROTUBULES FROM CARDIAC TISSUEAND A. PURIFICATION FROM ISOLATED MYOCYTES

Various methods for isolating and purifying tubulin have been reported (for a review see Dustin, 1984) and include purification by DEAE chromatography (Weisenberg el al., 1968), cycles of assembly and disassembly with or without glycerol (Shelanski et al., 1973), affinity chromatography on a colchicine- or peroxidase-loaded Sepharose column (Rousset and Wolff, 1980), and vinblastine precipitation (Bryan and Wilson, 1971). All these techniques yield purified tubulin when applied to brain tissue, which is particularly rich in MTs; however, they require that a critical concentration of tubulin be present that readily occurs in the brain but not in the adult heart, which has relatively few MTs (Samuel et al., 1983b). Using the antimitotic drug taxol-a substance isolated in 1971 from Tuxus breuifolia-Vallee improved the MT purification procedure so that the critical tubulin concentration could be reduced to only 0.01 mg/ml (Vallee, 1982). Taxol binds to tubulin and MTs, favors the formation of MTs, and increases their stability, hence preventing the disassembly of tubulin and MAPs. The binding of taxol is reversible and assembly can take place in the absence of MAPs (for a review, see De Brabander et af., 1986).

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The procedure of Vallee (1982) adapted by Parysek et al. (1984a) has made it possible to purify MTs from small amounts of cardiac tissue (such as 1 g of whole adult heart or 6-8 x lo6 isolated adult myocytes). Hearts are rapidly removed and immediately homogenized in 0.1 M PIPES, pH 6.9, 0.1 mM MgClz, 1.0 mM EGTA, 1.0 mM phenylmethylsulfonyl fluoride (PMEG buffer) (ratio of 1 g wet wtfl.5 buffer). Leupeptin (2 pg/ml) and trypsin inhibitor (0.1 mg/ml) are included in the buffer since cardiac tissue is rich in proteases. Extracts are prepared by an initial centrifugation at 4°C for 40 minutes at 60,000 g and then by centrifugation of the resultant supernatants for 90 minutes at 150,000 g . MTs are assembled by a 30-minute incubation at 37°C in the presence of 40 p M taxol and 2.5 mMGTP. The MTs are then sedimented by centrifugation at 250,000 g for 30 minutes at 30°C on a 0.5% sucrose cushion in PMEG buffer containing 20 p M taxol and 2.5 mM GTP. The pellet is washed with the same buffer and recentrifuged using the same conditions onto the sucrose cushion. The final pellet is frozen in liquid nitrogen and stored at -80°C.

B. IDENTIFICATION OF MICROTUBULE PROTEINS I N CARDIAC TISSUE 1. Tubulin Identification MTs are obviously very conservative structures and their main constitutive protein is tubulin. Tubulin exists as heterodimer composed of a! and p subunits (Luduena, 1979) that differ in their amino acid compositions (Krauhs el al., 1981; Ponstingl et al., 1981) and electrophoretic and immunological properties. The a! and p subunits of the dimer have molecular weights of 55,000 and 53,000, respectively (for a review, see Dustin, 1984). MTs from cardiac tissue were analyzed by Western blot analysis using polyclonal or monoclonal immunoglobulins directed against brain tubulin. After reduction and alkylation, cardiac tubulin resolves into two subunits of about 50,000 Da differing slightly in their electrophoretic mobilities as do those of brain tubulin. Both polyclonal and monoclonal tubulin antibodies recognized tubulin purified from total heart or from isolated cardiac myocytes. No differences in cardiac tissue and isolated myocytes that could arise from the presence of nonmuscle cells in heart were observed (Fig. 1). Isoelectric focusing enables the resolution of as many as 14 different isoforms of tubulin in the mouse brain (Wolff et al., 1982). Although tubulin has been highly conserved during evolution, intra- and interspecies differences exist. At the present time, no data are available concern-

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FIG. 1 . Imrnunoblot probed with antibodies affinity purified from brain tubulin. Brain ( I ) , heart tissue (2). and cardiac myocyte extracts (3) were electrophoresed on an 8% polyacrylamide SDS gel, blotted, and then (A) stained with india ink or (B) incubated with antibodies directed against tubulin. In collaboration with C. Delcayre (INSERM. Paris, France).

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ing the different tubulin isoforms present in cardiac cells nor their similarities and/or differences with those of other tissues. It appears that the tubulin isoforms are encoded by a multigene family. Fifteen to 20 tubulin genes have been described in the mammalian genome, however, in human and rat genomes, half of the a- and p-isotubulin genes appear to be pseudogenes (Cleveland and Sullivan, 1985). In chicken, seven ptubulin genes have been identified by Sullivan et al. (1984, 1985) and were shown to be independently regulated and expressed in a tissue-dependent fashion. However, although a preferential expression of a particular gene may be observed in some tissues, it appears that an individual P-tubulin isoform is not enough to constitute a complete cellular /I-tubulin set and must be present together with other p-isotypes in the cell (for a review, see Cleveland, 1987). The major differences in amino acid sequences of various p-tubulin isoforms reside within the last 15 amina acids of the carboxyl-terminus. This implies that this region of the protein must be involved in some important function within MTs (Farmer et al., 1984). a-Tubulins are encoded by approximately six functional genes and are expressed in specific but complicated programs during differentiation (for a review, see Cleveland, 1987; Miller et al.., 1987). 2 . IdentiJication of Microtubule-Associated Proteins (MAPs) in Cardiac Tissue

Following microtubule depolymerization in uitro, the major components isolated from brain MTs besides tubulin are polypeptides with an approximate M , of 280,000-300,000 known as microtubule-associated proteins (MAPs) that are described as MAP, to MAP4, and a group of proteins with an approximate M , of 60,000 called Tau factors (for a review, see Dustin, 1984). MAPz, like Tau factors, stimulate MT assembly in uitro. They form long protrusions on the polymer surface and are strong candidates as factors for polymerization regulation and crosslinking (Vallee et al., 1984; Drubin and Kirschner, 1986). In the heart, MAPs were identified in MTs purified from myocytes or cardiac tissue using the taxol-driven MT assembly (Fig. 2). However, brain anti-MAP1 antibodies do not react with mouse cardiac tissue (Fig. 2C), thus suggesting that MAPI is specific to the brain (Parysek et al., 1984a). In contrast, Wiche et al. (1984) have described the presence of both MAP, and MAP2 in rat cardiac tissue. The discrepancy concerning the presence of MAP, in the heart may result from subtle differences in the polypeptides of the MAP complexes or from species specificity of the protein. A small amount of MAP2has also been observed in isolated adult cardiac myocytes by radioimmunolabeling (A. Fellous, personal communication). Interestingly, Parysek et al. (1984a), using mouse heart,

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Fro. 2. Immunoblots probed with antibodies to the MAP4 region and MAP, region. Neuroblastoma (Nb), brain (Br). and heart (He) extracts were electrophoresed on a 7% gel, blotted, and then incubated with whole MAPl antiserum (A), antibodies affinity purified from the MAP, band ( B ) ,and antibodies affinity purified from the MAP, region ( C ) .Note the absence of MAP, in the heart extract. Two dots. MAP,; arrowhead, MAP,; dash, band 4. Reproduced from Parysek et a / . , J. C e / / B i d . , 99. 1309, 1984a, by copyright permission of The Rockefeller University Press and authors.

identified consistent amounts of MAP, designated as 215,000 Da (Fig. 2B). It is worth noting that MTs purified from certain classes of skeletal muscle fiber types show no reaction with the antiserum against MAP4 (Parysek et al. , 1984a,b). Among the subgroups of MAP4, cardiac MT pellets contain (as in brain, liver, and lung but in contrast to kidney and

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spleen) the MAP4A subunit and varying amounts of MAP4B and C. Interestingly, MAP4 has been shown to be similar in mobility to a-spectrin (240,000 MW) (Olmsted and Lyon, 1981). Even if data in cardiac tissue are limited and still controversial, the expression of MAP complexes appears to be tissue specific. Their study will be important in ultimately understanding how MAPS function in situ in the heart.

c. QUANTITATION O F TUBULIN I N CARDIAC MYOCYTES 1 . Characteristics of the [3H]Colchicine-Binding Reaction in Heart

Although many chemicals combine specifically with MT subunits and prevent their assembly, colchicine is one of the most powerful and specific (for a review, see Dustin, 1984). The characteristics of binding have permitted the identification of tubulin in a large number of organs and species. The characteristics of [3H]Colchicine binding to myocardial cell extracts, first described by Limas (1979), are similar to those described by others for various tissues. [3H]Colchicine binding is both temperature and time dependent. The colchicine-binding activity is unstable, the half-life for decay is approximately 4 hours. The [3H]colchicine-binding reaction permitted quantitation of tubulin in rat heart myocytes. A few days after birth, when both colchicine binding and extent of polymerization are maximal, free tubulin represents approximately 0.04% of the total myocyte proteins and half of the total tubulin pool (Klein, 1983). From day 5 to adulthood, colchicine-binding capacity declines from 49 to 21 pmol/mg protein and is accompanied by a decrease in the extent of tubulin polymerization from 39 to 19% (Limas, 1979).

2. Quantitation of Tubulin in Mature Cardiac Myocytes As tubulin is a ubiquitous and conserved protein, polyclonal antibodies raised against brain tubulin usually have very little tissue and species specificity. Therefore, tubulin quantitation can be carried out using a sensitive immunoenzymatic technique according to the sandwich ELISA (enzyme-linked immuno-sorbent assay) method (Engvall, 1980). This technique relies on obtaining antibodies against tubulin in two different species. The ELISA method allows the quantitation of purified brain tubulin from 0.20 to 10 pg tubulin/ml, which means that 10-500 ng of tubulin can be quantitated. In adult myocytes, tubulin is particularly insoluble since only 35% of the tubulin is extracted in the 26,000 ,g supernatant instead of the 75% extracted from brain; the remaining tubulin can be solubilized only with NP-40. Using the ELISA method, the

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tubulin concentration in adult cardiac myocytes was found to be 0.01% of the total myocyte protein (Samuel et al., 1983b) whereas in other tissues and cells it represents about 0.2-10% of the total protein (Hiller and Weber. 1978; Fulton and Simpson, 1979). The lower concentration found in the adult (0.01% of the total protein) as compared to the newborn (0.08% of the total protein) (Klein, 1983) confirms, using a different approach, the decrease in tubulin concentration described during postnatal development by Limas (1979). 111. Distribution of Microtubules in Cardiac Muscle

Mts were first visualized in striated muscle using transmission electron microscopy (Fardeau, 1965; Page, 1967; Hatt et al., 1970). However, the immunocytochemical techniques that have been developed over the last few years appear to be the best methods to reveal the MT network in individual whole cells (Weber and Osborn, 1981) and to allow the analysis of the distributions of their constitutive proteins such as a- or P-tubulin and MAPS subunits. The ultrastructural analysis, whether or not it is associated with immunolabeling, permits the visualization of the relationship between MTs and cell organelles (Goldstein and Entman, 1979; Watkins et al., 1987).

A. METHODS

FOR THE

ANALYSIS OF MICROTUBULE DISTRIBUTION CARDIACMYOCYTES

WII HIN

1. Techniques of Immunolabeling Using the Light Microscope a . Labeling of Isolated Myocytes. Immunolabeling of MTs within mature rod-shaped cardiac cells cannot be obtained using the classical techniques described for cultured cells (Weber and Osborn, 1981). One of the technical problems inherent in adult myocytes is the poor penetration of the primary specific antibodies even following permeabilization (Samuel et a l . , 1983a,b). Another problem is that MTs are difficult to distinguish in mature myocytes (30 pm of thickness) because they are either superimposed upon each other or are in a direction perpendicular to the plane of focus. An efficient labeling of MTs within the rod-shaped myocytes was obtained by treating the cells with 1% Triton X-100 in an MT-protective buffer (100 mM PIPES, pH 6.9, 10 mM EGTA) (Schliwa et a / . , 1982) at room temperature for 1 minute, followed by 3.7% formaldehyde fixation (Samuel er al., 1983a,b). Slight modifications proposed by Zernig and Wiche (1985) gave similar results.

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Techniques for labeling immature, cultured cardiocytes are identical to those described for other types of cultured cells (for a review, see Weber and Osborn, 1981). Fixation with cold methanol containing 10 mM EGTA was found to be the best method for the immunocytochemical study of microtubules together with other cytoskeletal components present in myocytes (Samuel et al., 1985). b. Labeling of Cryostat Sections. No specific labeling was obtained with antitubulin antibodies in cardiac tissue when rat heart was frozen in isopentane at the temperature of liquid nitrogen and then cut in a cryostat. However, following the method of Wiche et al. (1984), M M , and MAP2 can be localized in this tissue using a posttreatment with acetone. c. Labeling of Ultrathin Sections of in Situ Fixed Heart. Visualization of cytoskeletal structures in cardiac muscle was recently improved by using a technique for immunocytochemical labeling of ultrathin frozen sections (80 nm) that allows precise localization of specific intracellular proteins (Tokuyasu at al., 1981, 1983a,b; Tokuyasu, 1983). For tubulin labeling, mild fixation of the tissue with glutaraldehyde (0.2%) and formaldehyde (2%) at room temperature prior to ultrathin cryosectioning was the only method that permitted us to obtain high resolution combined with good preservation of both the polymerized state of the MTs and the antigenicity to antibodies (Watkins et al., 1987). With light microscopy, fluorescent labeling of ultrathin sections allowed an overview of tubulin distribution throughout the tissue section. Since the thickness of the sections was less than the depth of the focus of the objective, high-magnification and high-resolution images were readily obtained. A similar technique (Wolosenvick and De Mey, 1982) allowed Parysek et al. to study the localization of MAP4and tubulin in the mouse heart at the light fluorescence level (Parysek et a f . , 1984b). 2. Methods for Improving the Ultrastructural Analysis of Microtubules in Heart Muscle MTs were first visualized in the heart using classical fixation procedures for electron microscopy (Page, 1967; Hatt et al., 1970; Ferrans and Roberts, 1973; Page and Callister, 1973; Simpson et af., 1974). Goldstein and Entman (1979) further demonstrated that fixation in a polymerizing buffer (5% glutaraldehyde, 100 mM PIPES, 2 mM EGTA, 1 mM MgS04, 1 mM GTP) at 25” C preserves MTs as well as the overall cell morphology. Furthermore, MTs in heart were found to be best preserved and the most obvious in tissues that have undergone “conventional” fixation with aldehyde solution and postfixation in cacodylate-

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buffered osmium tetroxide as opposed to OsFeCN treatment, which renders relatively low electron opacity to cellular components other than the elements of membrane systems (Forbes and Sperelakis, 1983). Recently, the visualization of cytoskeletal elements in striated muscle at the electron microscopic level was improved by using the technique for immunocytochemical labeling of ultrathin frozen sections. This method enables the precise localization of specific intracellular proteins without compromising the integrity of the tissue (Tokuyasu et al., 1981, 1983a,b; Tokuyasu, 1983). The fixation procedure is identical to that described above for immunofluorescent labeling of ultrathin sections. These fixation conditons ensured minimal ultrastructural changes when used for immunoelectron microscopy. The tubulin distribution can be analyzed together with that of other proteins such as desmin by double indirect immunolabeling using two different sizes of gold particles (5 and 15 nm) coupled to the secondary antibodies in thin sections of muscle (Watkins et af., 1987). This method is a particularly useful tool for studying the distribution of components in cardiac myocytes that are either present at a very low concentration (as MTs) or sometimes difficult to identify by their size only (i.e., IO-nm desmin filaments).

B.

MICROTUBULE DISTRIBUTION IN CARDIAC

TISSUEAT

THE

LIGHT

MICROSCOPIC LEVEL 1 . Microtubule Distribution in Isoluted Myocytes In isolated myocytes. the absence of artifacts resulting from the permeabilization and/or fixation procedures that may distort cell organization is ensured by the double labeling of myocytes with antitubulin and either antimyosin or antidesmin (Fig. 3). In addition, a good correlation is found to exist between the cross-striations observed by phase-contrast microscopy and the staining with antimyosin in mature myocytes that confirms ( 1 ) the preservation of both the whole morphology of cells and the antigenicity of proteins and (2) the accessibility of antibodies throughout the cell (Samuel et al., 1983b). In rod-shaped myocytes, an abundant tubulin label is seen in the perinuclear area with a more intense staining at the poles (Fig. 3b and d). Few bright fluorescent spots extend from the nuclear pole in the conal region, an area free of myofibrils. A thin and tortuous network forming a looping structure, never superimposable on the striated pattern of myofibrillar components such as desmin or myosin, extends throughout the sarcoplasm. Interestingly, the microtubule network varied from one myocyte t o the other suggesting the dynamic state of this structure. MTs

FIG. 3. Double indirect immunofluorescence of myocytes isolated from 8-week-old rat hearts. Cells were incubated with antitubulin (b,d) and with either antimyosin (c) or antidesmin antibodies (e). Note that tubulin network (b,d) is unrelated to the striated pattern revealed by either phase-contrast optics (a) or by myosin (c) and desmin (e) labeling (bar = 10 pm).

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are also characterized by either the disorganization of their network by colchicine, nocodazole, and cold treatment o r by their stabilization by taxol, which enhances the polymerization of the cytoplasmic microtubules and induces resistance to cold (for a review, see Dustin, 1984). These agents were used as tools to ensure that the fluorescent label was truly associated with MT structures. When permeabilized mature myocytes are incubated for 3 hours at 4°C o r are treated with 10 p M colchicine, the MT network completely disappears although the pattern of myosin staining is not modified (Samuel et af., 1983a). However, the effect of nocodazole ( 10-6M) on MT depolymerization, which occurs quite rapidly in cultured cells (Tassin et al., 1985a), is often incomplete in adult myocytes and observed only after 4 hours of treatment in a minor population (Fig. 4). Conversely, when permeabilized myocytes are incubated with 10 pM taxol, the number of MTs does not increase significantly, but the depolymerization of the MT network, normally induced by a cold treatment (20 hours PC), is prevented. Thus, as shown in other cells with definitive morphology (Kirschner and Mitchison, 1986), MTs in mature myocytes are less sensitive to depolymerizing drugs than those in immature cells. In immature embryonic and neonatal cultured cardiac myocytes, the MT network, concentrated around the nucleus, disperses as a meshwork toward the cell margins (Fuseler et a / . , 1981; Nath et al., 1978). The MT network in freshly isolated neonatal myocytes (Fig. 5a) more closely resembles that of mature myocytes (Fig. 3) than that of cultured cells (Fig. 5c and e). After only I day of culture there is a dramatic change in MT distribution. The MTs, although still craddling the nucleus, form abundant longitudinal arrays in the sarcoplasm (Fig. 5c). With increased time of culture, the MT network develops along with the spreading of cells. MTs extend throughout the whole sarcoplasm (Fig. 5e), thus giving a pattern similar to that observed in fibroblasts (Fig. 5c), except that they never appear to emerge from a microtubule-organizing center (MTOC). 2 . Microtubule Distribution within the Myocardium Tubulin-labeled ultrathin transverse sections of adult rat ventricular myocardium reveal that the labeling is much more intense in nonmuscle cells and blood vessels than in myocytes (Fig. 6), thus confirming de uisu data previously obtained with the aid of immunoquantitation (see Section (11). In longitudinal sections, tubulin labeling was visualized as a ring around the nuclei and as punctuate elements or short strands with no particular orientation in the sarcoplasm. However, some MTs that have an axial orientation reflect their loose, helicoidal arrangement in whole cells (compare Fig. 3 and 6). In mouse, helicoidally arranged MTs are

FIG. 4. Effects of the MT-depolymerizing drug nocodazole on the MT pattern in cultured mature myocytes. After treatment for 4 hours with nocodazole (5 x M ) , MTs are either completely (a) or partially (c) depolyrnerized. MTs depolymerization has no affect on the rod shape or the striated pattern of the myocyte, which are easily visualized with phase-contrast optics (b) (bar = 10 jm).

FIG. 5 . Double immunolabeling of neonatal heart myocytes with antibodies directed against tubulin (a.c.e) and desmin (b,d.f ). Myocytes were freshly isolated (a,b) and cultured for 1 day (c.d) and for 4 days (e.f). Within elongated myocytes, MTs are distributed around the nuclei and longitudinally throughout the sarcoplasm whereas desmin has a striated pattern. After 4 days of culture (e), the MT pattern resembles that of a fibroblast (arrow in c) (bar = 10 pm). 114

FIG. 6 . Fluorescence micrograph of a longitudinal ultrathin frozen section of rat ventricular muscle immunolabeled with tubulin antibodies (a) and observed with phasecontrast optics (b). Tubulin labeling is abundant in the connective and perivascular tissues (arrow) and appears as little spots and dotted lines within myocytes (bar = 10 pm).

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seen more frequently in cardiac muscle than in skeletal muscle (Fig. 7a and cl.

3. Distribution oj’Microtubule-Associated Proteins in Myocardium The presence of an MT-associated protein, MAP4, in cardiac tissue has recently been demonstrated by Parysek et al. (1984b). Using specific antibodies, these authors have demonstrated that in mouse heart the patterns of distribution of MAP4 and tubulin are similar (Fig. 7), however MAPj labeling is much weaker than that of tubulin. As was previously shown for tubulin (Fig. 6), connective tissue cells and blood vessels of muscle are also more strongly labeled than muscle cells by MAP4 antibodies (Fig. 76). In brain, MAP4 is a minor component of the MAP proteins compared to the dominant MAP, and MAP: (Olmsted and Lyon, 1981 ; Parysek et al., 1984a). MAP4 appears to be highly tissue specific and mainly present in connective and support tissues (Parysek et al., 1984a,b). According to Parysek et al. (1984b) and also to our recent results (Fig. 8), no specific MAP, and MAPz labeling is visualized either in mouse or rat cardiac muscle. However, in both rat skeletal o r cardiac muscle sections and in isolated myocytes, Wiche and co-workers (Wiche et 01.. 1984; Zernig and Wiche, 1985) described cross-striations stained with anti-MAP, and anti-MAP, antibodies that are unrelated to tubulin pattern. The huge dissimilarity between tubulin and MAPl and MAP2 distribution observed by Wiche and co-workers and the discrepancy of their results with those of Parysek et af. (1984b) and ours can be explained by epitopes shared between unrelated structures. Antibodies raised against MAPS are found to stain organelles other than MTs (Hill et af., 1981; Sherline and Mascardo, 1982). In these cases, it was suggested that the antibodies recognize an antigen common to MTs and to multiple MT-associated structures. The absence of the MT-binding site for a given MAP is likely t o result in an MAP distribution characteristic of some other structure such as actin o r intermediate filaments (Vallee et al., 1984). More experiments are required to affirm the presence or absence of MAP, and MAPl in mammalian cardiac myocytes. The subtle differences in the distribution and specificity of the different MAP subunits is important in understanding the cell and/or tissue-specific functions of MTs. C. MICROTUBULE DISTRIBUTION I N CARDIAC MYOCYTES AT ELECTRON MICROSCOPIC LEVEL

THE

MTs in cardiac muscle range from 24 to 30 nm in diameter and may reach lengths of several micrometers (Forbes and Sperelakis, 1980;

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FIG.7. Sections of mouse skeletal anterior thigh (a, a’), eye (b, b’), and cardiac (c, c’, and d) muscle labeled with MAP4 (b and d) or tubulin (a and c ) antibodies. Although weaker in intensity, the MAP4 antibody labeling patterns are similar to those seen with tubulin antibody. In (d), a perivascular cell (arrowhead) and presumptive connective tissue (arrow) are also labeled with MAPI antibody. Corresponding contrast images (a’, b’, c‘) are shown for a, b, and c. Bar = 10 pm. Reproduced from Parysek et al., 1.Cell B i d . , 99,2287, 1984b, by copyright permission of The Rockefeller University Press and authors.

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FIG. 8. Isolated adult rat cardiac myocytes incubated with MAP2 antibodies (a) and observed with phase-contrast optics (b). An insignificant pattern is observed even at the level of nuclei (bar = 10 p m ) (MAP2 antibodies were kindly provided by A. Fellous, INSERM. Paris. France).

Goldstein and Entman, 1979). Transverse sections of these tubules show them to have typical rounded profiles further enhanced by Markham rotation of n = 13 (Forbes and Sperelakis, 1983; Goldstein and Entman, 1979). They correspond to the typical longitudinal arrangements of 13 protofilaments with a regular assembly of alternating a and p subunits in a helicoidal structure. In adult dog and guinea pig (Goldstein and Entman, 1979), mouse (Forbes and Sperelakis, 1980), human (Ferrans and Roberts, 1973), or rat (Watkins et ul., 1987) ventricle, myocardial cells yield images of MTs dispersed longitudinally or transversely in the rnyoplasm adjacent to the I

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band and at the periphery of nuclear profiles. Longitudinal MTs surround the nuclei without displaying any connections with the myofibrils in this area (Fig. 9) (Goldstein and Entman, 1979; Watkins et al., 1987). MTs are also present in the conal area where the Golgi apparatus and the rough endoplasmic reticulum are concentrated. In longitudinal sections, MT profiles seem to wrap around the myofibrils and go across their surface at various angles (Fig. 10) (Goldstein and Entman, 1979; Forbes and Sperelakis, 1983; Watkins et al., 1987). MTs are longitudinally oriented in the intermyofibrillar spaces and sometimes pass through the desmin filaments located between the Z disks when extending along several sarcomeres (Fig. 11) (Watkins et al., 1987). Occasionally, MTs appear to surround the Z disks and the sarcomere, but MTs never display a striated pattern. Undulating profiles are also found to follow the contour of sarcoplasmic reticulum tubules (Fig. 10). Some MTs run closely apposed to the outer membranes of mitochondria (Fig. 11) and along the sarcolemma1 borders, but they are absent from the intercalated disk area (Ferrans and Roberts, 1973; Watkins et al., 1987). In other types of cardiac muscle cells such as Purkinje cells (Rybicka, 1978) or atrial cells (V. J. Ferrans, personal communication), MTs were

FIG. 9. Immunoelectron micrograph of a straight longitudinal profile of MTs (arrows) next to the smooth-shaped nucleus (bar = 0.3 pn). In collaboration with S . Watkins, D.F.C. Institute, Boston.

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Fro. 10. Electron micrograph of a mouse ventricular myocyte. MTs run both longitudinally and obliquely in this field with the individual MTs tending to bend over the level of the I band (bar = 0.2 p m ) . Reproduced from Forbes and Sperelakis (1984). “Physiology and Pathophysiology of the Heart.” p. 3 by copyright permission of Martinus Nijhoff Publishing and the authors.

shown to have a distribution similar to that observed in ventricular cells and are said to be more numerous. However, no morphometric data are available to confirm this latter observation that may be secondary to the relative lower density of myofibrils present in these types of cardiac cells (Forbes and Sperelakis, 1984).

D. RELATIONSHIP BETWEEN MICROTUBULES AND INTERMEDIATE FILAMENTS MTs and intermediate filaments have been described as functionally associated in different types of cultured cells (Geiger and Singer, 1980; Summerhayes er al., 1983). This concept was supported by the observation of a codistribution of these two cytoskeletal elements that could be distinguished only after disruption of the MT network by an antimitotic drug that induces intermediate filaments to collapse around the nuclei (De Brabander et al., 1982; Weber and Osborn, 1981; Price and Sanger, 1983). In ventricular myocardium, ultrastructural studies have shown the absence of connections between MTs and intermediate filaments (Ferrans and Roberts, 1973; Behrent, 1977; Thornell and Eriksson, 1981).

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FIG. 11. Electron micrograph of a longitudinal ultrathin frozen section of rat heart muscle double immunolabeled with desmin (large gold particles) and tubulin (small gold particles). Note that the tubulin label is intermingled with the desmin label (I.F.) at the level of the Z disk ( Z ) . M, mitochondria (bar = 0.2 pm).In collaboration with S. Watkins, D.F.C. Institute, Boston.

However, the high density of myofibrils and of other cellular components within striated muscle cells and the sparsity of MTs make it very difficult to use transmission electron microscopy to study the interaction of MTs with other cytoskeletal elements. Immunoelectron microscopic techniques allow the identification of MT constitutive proteins. We have already pointed out the clear differences in the respective distributions of tubulin and desmin at the light microscopic level. From immunoelectron microscopic studies, it emerges that desmin filaments are particularly concentrated in the intermyofibrillar space of the Z disk area where MTs

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are few and at the intercalated disk areas devoid of MTs. Desmin label never appears to reach the nuclear membrane. MTs and intermediate filaments, quite distinct in their distribution, appear to be poorly connected throughout the mature myocyte even if the two labels are occasionally seen to be intermingled (Fig. 11). Connections between MTs and desmin filaments in adult rat ventricle are thus probably more coincidental than functional. Possible interrelations between MTs and intermediate filaments were further studied by analyzing the effect of microtubule-depolymerizing drugs on immature or mature cultured myocytes. In mature myocytes. desmin associated with the 2 disks displays the typical cross-striated pattern (see Fig. 3). In immature myocytes, with time of culture, desmin loses its striated pattern to form a meshwork of cables and foci more or less similar to that of vimentin in nonmuscle cells (see Fig. 5 ) . In both types of myocytes when desmin is associated with the Z disks, depolymerization of the MT network does not affect the desmin distribution. In contrast, when desmin is no longer associated with the Z disks, i.e., within immature myocytes after 3 days of culture, the MT disruption by colchicine induces, as in nonmuscle cells, the collapse of desmin into a perinuclear mass (Schwartz and Rappaport, 1987). Consequently, we suggest that in immature cardiac myocytes, desmin filaments and MTs may be interconnected in some way and that, as in nonmuscle cells, the intermediate filaments are supported in the cell upon a MT scaffold (Bloom et al., 1985). However, the interconnections between MTs and desmin filaments, when they occur, are much weaker than those of desmin within Z disks. According to Vallee et al. (1984), the specificity of the cross-linking of MTs and intermediate filaments would mainly depend on MAP?. Thus, a parallel could be made between the poor association of MTs and desmin filaments that we observed in adult heart and the small amount of MAP? detectable in this tissue. Similarly, it could be hypothesized that in immature cardiac myocytes the linking of MTs with desmin filaments might depend on a relatively higher amount of MAPz. In conclusion, the distribution of MTs in cardiac tissue appears to be rather similar among the different mammalian species studied-mouse, rat, human-and between the different types of cells-conductive fiber and ventricular myocyte. The fact that few studies have been carried out on MTs in cardiac tissue seems to be secondary to technical difficulties in the preservation of tissues and/or microtubular structures mainly by fixation procedures (Goldstein and Entman, 1979; Rybicka, 1978; Samuel et al., 1983b). To sum up, MTs in cardiac myocytes are rather sparsely distributed around the myofibrils and constitute the main cytoskeletal structure in the areas close to the mitochondria, plasma, and nuclear membranes.

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IV. Roles of Microtubules in the Cardiac Myocyte

Considerable evidence exists that indicates that in most cells MTs are implicated in cellular morphogenesis and provide a framework to which membranous organelles are attached. Depending on the tissue and on its developmental stage, MTs were shown to be associated with intermediate filaments (Geiger and Singer, 1980) and actin microfilaments (Pollard et al., 1982, 1984), and to be involved in many tissue-specific roles such as myofibrillogenesis in muscle (Auber, 1969; Toyama et al., 1982; Warren, 1968). In cardiac tissue, the role of MTs has, as yet, been poorly investigated; however, recent studies tend to demonstrate that they are involved in several membranous processes and could have a more dynamic rather than mechanical role within myocytes. A. MICROTUBULES A N D MYOCYTE SHAPE PRESERVATION The shape of nonmuscle cells depends greatly on the MT network (Weber and Osborn, 1981; De Brabander et al., 1982; Dustin, 1984). In certain embryonic muscle cells, disruption of the MT network induces spherical cells in which the myofibrils are arranged in a concentric pattern around centrally located nuclei (Warren, 1968; Ishikawa et al., 1968). In differentiated cardiac muscle, several authors (Ferrans and Roberts, 1973; Goldstein and Entman, 1979) have suggested that the microtubule network might influence the shape of the myocyte and, more particularly, provide a structural support to maintain the shape of nuclei. This latter concept was essentially based on the observations of longitudinal MTs paralleling the smooth contour of the nuclei and of changes in their orientation when nuclear profiles are irregular. Additional data suggest that MTs are poorly involved in the maintenance of myocyte shape. Beating immature cardiac myocytes maintained in culture develop, with time, changes in their shape that are associated with a random distribution of new myofibrils, and a progressive alteration of the MT patterns (see Section III,B, 1). However, according to Kulikowski (1981), the changes in shape of cultured immature myocytes are mainly secondary to the absence of unique direction of strain during the cycle of contraction and relaxation. Alterations in the MT network could be secondary to the later event. On the other hand, a rounding of mature rod-shaped myocytes has been described as being related to a hypercontraction of myofibrils secondary to the Ca2+ paradox (for reviews, see Dow et al., 1981a,b; Jacobson and Piper, 1986). A few of the mature rod-shaped myocytes, when maintained in culture, sometimes exhibit a rounding of their shape (Schwartz et af., 1985; Eckel et af., 1985). Hypercontraction in these

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myocytes is evidenced by anti-a-actinin staining (Fig. 12d). A clear MT pattern is displayed throughout these cells (Fig. 12b). A minor role of MTs in the maintenance of myocyte shape is also suggested by the absence of an effect of MT depolymerization on the shape of the whole myocyte and nuclei in both mature (Fig. 4) and immature (Schwartz and Rappaport, 1987 myocytes. These results confirm ultrastructural observations suggesting that the round nucleus of neonatal cardiac myocytes takes on an oval almost rectangular shape in response to the molding influence of the developing myofibrils (Legato, 1979). Therefore, in both immature and mature cultured myocytes, changes in shape seem to be more dependent on myofibrillar organization than on microtubule network. The compacted, registered collection of myofilaments might be maintained by the arrangement of various combinations of several cytoskeletal components among the myofibrils distributed either transversely as intermediate filaments (Lazarides, 1980; Tokuyasu et al., 1983a.b, 1984, 1985), or longitudinally as titin and nebulin (Wang, 1982; Wang et al., 1984; Horowits et al., 1986). Finally in situ, the shape of the cardiac muscle cell is also dictated to some degree by its surroundings, the ventricular cell's profile conforming to the contours of the numerous vessels of the heart (Forbes and Sperelakis, 1984).

B. INVOLVEMENT OF MICROTUBULES I N MYOCYTE FUNCTION 1. Microtubules und Contraction Rates of Myocytes The relationship between cellular contraction and cytoskeletal elements has been studied using cultures of immature rat myocytes (Nath et al., 1978: Klein, 1983; Lampidis et al., 1986). Nath et al. (1978) reported that the contraction of immature myocytes is inhibited by 80% after incubating cell cultures for 2.5 hours in the presence of 0.2 mM dBcAMP. This inhibition is reversed by washing the treated cells or by the addition of 1 p M colchicine or 1 p M colcemid. By light microscopy, dBcAMP treated myocytes show the MTs to be organized into thick parallel arrays rarely observed in untreated myocytes (Bollon et al., 1982). The alignment of MTs into parallel arrays might confer a physical constraint on the myocytes that prevents beating (Nath et al., 1978). These authors suggest that the effect of dBcAMP on the rate of contraction is due to some FIG. 12. lrnrnunofluorescent labeling of cultured adult cardiac rnyocytes with antibodies directed against either tubulin (a,b) or a-actinin (c,d). In both rod- (a) and round- (b) shaped rnyocytes the MT network is easily detected. Hypercontraction of myofibrils in a roundshaped myocyte is demonstrated by the a-actinin pattern (d) (bar = 10 prn). The anti-aactinin antibodies were kindly provided by B. Jockush, Bielefeld, RFA.

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indirect action of the cyclic nucleotide such as changes in endogenous calcium availability for contraction. However, the positive chronotropic effect observed when the MT network is destroyed would argue in favor of an MT role in the control of myocyte function. The spontaneous rate of myocyte beating increases by 84% when cells are treated with colchicine M> or with nocodazole (1 X M ) (Klein, 1983; Lampidis et (1 x al., 1986). The chronotropic effect of the drug is dose dependent, obtained after 10 minutes (Lampidis et al., 1986), and reaches a maximum after 100 minutes (Klein, 1983). According to Klein (1983), the chronotropic effect of colchicine is unrelated to alterations of the basal level of adenyl cyclase activity or of adrenergic receptor sensitivity since the effects of isoprenaline (P-adrenergic agonist) on increased heart rate occur after 3 minutes and are transitory. Furthermore, propanolol, a P-adrenergic receptor blocker, does not inhibit the colchicine effect. Since the spontaneous rate of myocyte contraction is dependent upon the properties of membrane conductance, it is possible that changes in the MT network affect sarcolemmal components that control ionic fluxes. However, according to the results of Lampidis et al. (19861, the MT equilibrium may interfere with the P-adrenergic system of myocytes since colchicine, in addition to its positive chronotropic effect, causes a reduction of the amplitude of contraction that is reversed (within 5 minutes) upon the addition of the p-adrenergic agonist, isoproterenol (Lampidis et al., 1986). In conclusion, the state of polymerization of MTs clearly influences, either directly or indirectly, myocyte function. 2 . Microtubule Involvement in the Process of Receptor Internalization In nonmuscle cells, different types of membranes, such as Golgi apparatus, mitochondria, and endoplasmic reticulum, were shown to possess high-affinity, large capacity tubulin-binding sites (Berlin et nl., 1982; Bernier-Valentin et a l . , 1983) that might initiate micropolymerization of tubulin (Bloom et al., 1984). It is clear from the morphological data shown above that in the heart, MTs are mainly found in areas rich in these structures. The hypothesis may be raised that in heart, as in nonmuscle cells, MTs are involved in the intracellular movements and protein synthetic processes related to these structures (Heggeness et nl., 1978; Ball and Singer, 1982). Biochemical data have reinforced the concept of a role of MTs in the regulation of sarcolemmal receptor number. Using a system of isolated adult rat heart myocytes, Eckel and co-workers have shown that various MT-depolymerizing agents inhibit the binding of insulin to cardiac myocytes without any significant effect on cell viability. The decrease in insulin binding is most probably related to the MT disruption and would

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be secondary to an MT involvement in the intracellular transport of insulin receptors to and from the sarcolemma (Eckel and Reinauer, 1980, 1983). According to Limas and Limas (1983), the MTs are involved in the process of down-regulation of p-adrenergic receptors by isoproterenol since MTs in a polymerized state inhibit the receptors’ translocation to the cytoplasm at the sarcolemmal level. It cannot yet be excluded that these results are secondary to a direct effect of the MT-depolymerizing drug on the plasma membrane. An alternative explanation is that these drugs disrupt the connections between the MT network and the plasma membrane. However, the biochemical investigations together with the ultrastructural observations argue that MTs play a role in the physiological regulation of cell receptor number at the level of the sarcolemma and/or sarcoplasm in the heart. AND MYOCYTE HYPERPLASIA C. MICROTUBULES

The process of mitosis is highly dependent upon MTs (McIntosh and Landis, 1971; Mitchison et al., 1986). However, in cardiac myocytes the mitotic index is very low during the first days after birth, and images of the mitotic apparatus in the heart are exceptional (for a review, see Zak, 1984). In the adult, some mitosis has been observed in atrial tissue in response to myocardial infarction (Rumyantsev, 1977). According to Limas, myocardial MTs are involved in the early steps of the initiation of DNA synthesis (Limas, 1979). The author describes in cultured neonate myocytes, a dose- and time-dependent inhibitory effect of colchicine on thyroxine (T&induced enhancement of [3H]thymidine incorporation that is no longer observed when colchicine addition is delayed for more than 10 hours after T4 administration. Colchicine would act during the period of commitment to mitosis. Since both [3H]colchicine binding and tubulin polymerization capacity decline in myocytes after birth, Limas proposed that the MT system may contribute to the age-dependent decline in the proliferative capacity of myocytes.

D. INVOLVEMENT OF MICROTUBULES IN CARDIAC HYPERTROPHY AND SARCOMEROGENESIS Hypertrophy of the heart develops as a result of hemodynamic overload. In the adult, the cardiac hypertrophy is mainly the result of the hyperplasia of nonmuscle cells and of the hypertrophy of the myocytes. Enlargement in myocyte size is accompanied by a structural remodeling including alterations in the relative proportions and ultrastructure of

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cellular components, such as myofibrils, mitochondria, and sarcoplasmic reticulum (Ferrans, 1984; Legato et al., 1983, 1984), that have previously been shown in Section I11 to be more or less surrounded by MTs. The exact nature of the process of cellular remodeling is dependent upon the stimulus to myocardial hypertrophy (hormonal, pressure overload, volume overload) (Oparil et ul., 1984). Thus, questions arise as to whether MTs are involved in the process of myocyte hypertrophy, and if they are involved, does their involvement differ depending on the nature of the stimulus? 1. Microtubules and Myogenesis

Hypertrophy of adult myocytes in response to an increase in hemodynamic load shares mechanisms common to all striated muscles (Legato et al., 1984). Therefore, a review of the work performed on skeletal myogenesis is important for our understanding of the roles of MTs in hypertrophy of cardiac myocytes. A remarkable action of MTs has been reported in myofibrils of embryonic striated muscle cells that opened new paths in the study of muscle organization (Antin et al., 1981; Toyama et al., 1982): the myosin thick filaments assemble regularly, as myonemes, along the MTs without any participation of actin microfilaments. Twenty years ago, Fischman (1967) and Auber (1969) observed in embryonic chick skeletal muscle and in insect striated muscle, respectively, that MTs are always oriented parallel to the long axis of the cell during myofibrillogenesis; thus, Auber proposed that MTs provide a scaffold for new myofibrils. During in uiuo and in uitro myofibrillogenesis, new myofibrils are thought to be formed under the sarcolemma. The translation products of the myosin and actin genes are rapidly assembled into thick and thin filaments to form sarcomeres that are laterally apposed to the preexisting myofibrils (Legato, 1979; Holtzer et af., 1982a,b). In addition, Holtzer and co-workers proposed that MTs may be involved in the addition of myosin filaments to form the A band (Antin et al., 1981; Toyama ct al., 1982). They demonstrated that myotubes previously depleted of their myofibrils, by a pretreatment with TPA (the cocarcinogen 12-0tetradecanoylphorbol-13-acetate), which were allowed to regenerate in the presence of the MT-polymerizing drug taxol, form “A bands” of aligned thick filaments interdigitating with long MTs and “I bands” consisting only of MTs and devoid of actin filaments (Fig. 13). The interaction between myosin and MTs may be an intracellular artifact. However, in uitro studies have shown that the affinity of tubulin dimers for myosin filaments equals that of actin, but is unaffected by Ca2+-or Mg2+-ATPin contrast to actin-myosin binding (Shimo-Oka et al., 1980). On the other hand, when regeneration of myotubes occurs in the presence

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FIG. 13. Electron micrographs of myofibrils assembled in TPA-treated myotubes that recovered in taxol for 4 days. (a) The individual thick filaments are -1.6 pm in length and the M and pseudo-H bands have the same dimensions as the corresponding bands in normal A bands. The same microtubules that constitute the only visible structures of the “I bands” (arrowheads) interdigitate with the flanking thick filaments, which form a remarkable facsimile of normal A bands. Single microtubules extend through several consecutive sarcomeres. The arrow points to a cluster of parallel microtubules of indefinite length. Elsewhere in this myotube, the lateral alignment of the thick filaments may be less precise. (b) A cross section illustrating the absence of all thin filaments in interdigitating arrays consisting only of thick filaments and microtubules. (c) A cross section through a sarcomere displaying interdigitation of thick and thin filaments and microtubules. The number of thin filaments in such arrays varies directly with the size of the Z band material. (d) A longitudinal section through a solitary A band. The precise lateral alignment of thick filaments reveals the M and H bands. Reproduced from Toyama et a/. (1982), by copyright permission of the authors.

of colcemid, i.e., in the absence of MTs, all the myofibrillar components are formed but do not achieve their rigorous lateral alignment. Consequently, MTs in a constant assembled and dissassembled equilibrium might influence the polymerization and/or spatial organization of myofibrillar structure.

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2 . Microtubules and Cardiac Myocyte Hypertrophy a. Microtubules during Postnatal Development. During the hyperplastic phase of myocardial growth (from birth to 4 days of life), the heart enlarges by increasing the number of mononucleated cells as in fetal life. The myocardial cells then lose the capacity of dividing, and further growth of the heart is due to myocardial cell hypertrophy and nonmuscle cell hyperplasia. The transition from hyperplastic to hypertrophic growth phases is related to the formation of binucleated myocytes (Clubb and Bishop, 1984). Cartwright and Goldstein (1982) reported that during postnatal development of the soleus skeletal muscle, MTs are added together with myofilaments, but an upper limit to the MT density is reached before the arrest of myofibril growth. To further assess if such an upper limit to the MT numeric density exists in the cardiac myocyte, and if the frame for development is similar when hyperplasia occurs together with hypertrophy, these authors analyzed the number of MTs in papillary muscle during the first weeks of postnatal life (Cartwright and Goldstein, 1985; Park et a / . , 1984). In cardiac myocytes, the number of MTs/pm2 increases after birth to reach a maximum value at 5-9 days of age, and then abruptly decreases to a constant level characteristic of the adult. According to these authors, MTs in cardiac myocytes can be divided into two subpopulations: ( I ) those adjacent to the nuclear membrane in the pennuclear space, and (2) those distributed randomly between the myofilament bundles. During the first 2 weeks of postnatal life, the numeric density of the perinuclear MT subpopulation is essentially stable and does not fluctuate significantly with age. Therefore, the fluctuation in the total MT numeric density should reflect changes in the number of MTs mostly scattered randomly among the myofilament bundles (Park et al., 1984; Cartwright and Goldstein, 1985). An increase in the MT density has been shown by immunofluorescence to occur within a part of the myocyte population during postnatal development (Samuel et al., 1986). These myocytes are binucleated, i.e., are in the hypertrophic phase of growth (Anversa et al., 1980). Taken together, these results suggest an organizational role for MTs during postnatal myocyte hypertrophy.

b. Microtubules during Rapidly Induced Hypertrophy. Involvement of MTs in the process of cardiac myocyte hypertrophy was further investigated in two models of in uivo cardiac hypertrophy induced either by pressure overload secondary to aortic stenosis or by thyroxine administration to severely hypothyroid rats. Cardiac hypertrophy may be defined, among other parameters, by myocyte enlargement (Korecky and Rakusan, 1978) and increased heart weight (for a review, see

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Swynghedauw and Delcayre, 1982). Hypertrophy of the rat heart may also be characterized biochemically by the shift in the expression of isomyosin heavy chain genes (MHC). During the adaptation to a pressure overload, a down-regulation of a-MHC occurred in the ventricle in such a way that the P-MHC became the predominant form 4 weeks after imposition of the overload (Lompre et al., 1981; Mercadier et al., 1981; Rappaport et al., 1984). On the other hand, thyroid hormone has a much more rapid effect and induces in few hours the expression of a-MHC in the ventricles (Lomprk et al., 1984; Chizzonite and Zak, 1984; Everett et al., 1984). Thus, these two isoforms of MHC provide sensitive markers for evaluating the response of the myocytes to different stimuli of cardiac hypertrophy and allow the investigation of the relationship between the alteration in the pattern of distribution of MTs and the process of cardiac hypertrophy. In both models of cardiac hypertrophy-mechanical and hormonal-a rearrangement of MTs into dense parallel arrays within the myocytes (Fig. 14) coincides with the early state of cardiac growth and myocyte hypertrophy (Rappaport et al., 1984, 1985, 1986; Samuel et al., 1984, 1986). This alteration does not occur simultaneously in all myocytes, which at a given time do not exhibit an identical MT network. The population of myocytes with a dense MT pattern increases for a period of 5 days reaching a maximum of 35% of the myocyte population, and then decreases toward zero 2 weeks after the induction of hypertrophy (Fig. 15). The MT reorganization lasts about 4-5 days in each myocyte and gradually affects the whole population (Samuel et al., 1984). This process is also observed during rat postnatal development, but its rate is slower since the rate of hypertrophy is less rapid than that obtained after hormonal or mechanical stimulation. The maximum percentage of the myocyte population with a modified MT pattern is mainly related to the rate of myocyte hypertrophy (Samuel et al., 1986). The process of increasing MT density is independent of the growth stimulus and of the specific response of the heart since it precedes the appearance of the newly synthesized P-MHC when hypertrophy is mechanically stimulated, but follows that of a-MHC when hypertrophy is hormonally stimulated (Rappaport ef al., 1984; Samuel et al., 1984, 1986). MT involvement in the remodeling of myocytes associated with the process of hypertrophy was investigated using a high-resolution immunolabeling technique (Watkins et al., 1987). Alterations in tubulin labeling obviously occur in myocytes randomly distributed throughout a tissue section of hypertrophied cardiac muscle when analyzed with the light microscope. They reveal the heterogeneity of the myocyte population, which can also be depicted using other cellular markers such as desmin

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FIG. 14. Different patterns of tubulin labeling observed within myocytes isolated at various times after stimulation of hypertrophy secondary to an aortic stenosis to 3-week-old rats (a-c) or a daily thyroxine treatment (4 pg/lOOg body wt) to 10-day-old hypothyroid rats (d-f). Myocytes were isolated 1 day (a,d), 7 days (b,e), and 15 days (c,f) after the onset of mechanical or hormonal stimulation. Note the more dense MT pattern in hypertrophying myocytes (b,e) (bar = 10 pm).

(Thornell et ul., 1984; Watkins et a l. , 1987) or isomyosins (Gorza et ul., 1981). This heterogeneity is also clearly evidenced by studies carried out using immunoelectron microscopy in which certain cells have a normal appearance (described in Section 111) whereas others exhibit the morphological characteristics of hypertrophic myocytes, i.e., alterations in intercalated disks, mitochondria, and myofibrils. In the pressure-

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n P

a

c 0

Days a f t e r stimulation FIG. 15. Percentage of myocytes exhibiting a dense microtubule pattern as a function of time after the onset of a mechanical (@) or hormonal (M)stimulation of hypertrophy; (0), controls. Experimental conditions were those of Fig. 14.

overloaded rat heart, the relative number of MTs running along the mitochondria and parts of the sarcoplasmic reticulum appears unchanged when compared to that of the control heart (Watkins et al., 1987). Thus, the MTs found along the mitochondria and plasmalemma are probably involved in cell organelle distribution and in the transport of substances through the sarcoplasm, but are not associated with the increased activity of the cell. The intercalated disks are altered to various degrees and sometimes display increasingly deep convolutions (Bishop, 1983; Ferrans, 1984; Legato et al., 1983, 1984). It is striking how MTs are absent from this area although numerous alterations in the intermediate filament distribution are observed (Ferrans, 1984; Thornell et al., 1984; Watkins et a)., 1987). The intense tubulin staining observed by light microscopy throughout the sarcoplasm is correlated, in electron microscopy, with an increased MT number in some myofibrillar areas (Watkins et al., 1987). Numerous MTs are seen either tightly apposed in a longitudinal direction to some sarcomeres (Fig. 16a) or in areas showing disorganized sarcomeres. The

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increase in the number of MTs apposed to the myofibrils probably reflects their possible organizational role in the myofibrillogenesis that occurs soon after aortic stenosis (Hatt, 1977; Legato et a/., 1983, 1984). Their role would thus be similar to that described in growing myoblasts (Warren, 1968; Auber, 1969; Toyama et al., 1982) or in postpartum rats (Cartwright and Goldstein, 1985). It also emerges that in hypertrophied myocytes, MTs are still absent from the Z disks and intercalated disks, both of which are supposed to provide a template for the organization of actin filaments (Sanger et al., 1984, 1986). Connections between actin filaments and MTs (Sattilaro, 1986) have been reported to form, in uitro, a lattice permitting the aggregation of myosin filaments (Pollard et al., 1982, 1984; Toyama et a / . , 1982). Such connections might also occur during in uiuo myofibrillogenesis. In myocytes that display a loss of myofibrillar register, desmin labeling is found to be distributed longitudinally between two laterally displaced Z disks, but in this area no MT profiles are observed (Fig. 16b). According to studies on skeletal myogenesis, desmin filaments are mainly involved in the process of myofibril registration (Price and Sanger, 1983) and their association with the Z disks occurs only after myofibrillogenesis (Tokuyasu et al., 1984, 1985). Thus, in hypertrophying myocytes, the transient increase in the MT number along the myofilaments might provide a scaffolding for the assembly of myosin filaments and precede the appearance of the longitudinal desmin filaments linking the Z disks of sarcomeres out of register. An increase in the number of MTs streaming away from the nuclei towards the sarcoplasm is observed in active hypertrophying myocytes in both rabbit (Hatt et al., 1970; Schwartz and Rappaport, 1987) and rat (Watkins et al., 1987) hearts. The higher density of MTs in this area, which is rich in membrane bound ribosomes and Golgi apparatus (Hatt et al., 1970; Ferrans and Roberts, 1973; Hatt, 1977), has to be related to their recently reported role in the processes of protein synthesis (Walker and Whitfield, 1985; Rogalski and Singer, 1984), and particularly in the translation of cytoplasmic mRNA (Bonneau et al., 1985). Therefore, the increase in density of MTs around nuclei may be associated with the transient rise in protein synthesis that occurs during the onset of cardiac hypertrophy (Schreiber et al., 1978; Moalic et al., 1984; Zak, 1984). ~~

~

FIG. 16. Immunoelectron micrograph of a frozen ultrathin section of hypertrophied rat ventricle double labeled with antitubulin (small particles, thin arrows) and antidesmin (large gold particles, head of arrow). In (a), a large number of MTs run along myofibrils. Note that MTs are absent from the intercalated disk (ID). In (b), longitudinal desmin labeling forms a longitudinal array connecting one Z disk with two laterally displaced Z disks (bar = 0.5 pm). In collaboration with S. Watkins, D.F.C. Institute, Boston.

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The new orientation of the MTs and the increase in their density in the nuclear area and along myofibrils during the onset of cardiac hypertrophy are in favor of a dynamic role of MTs in sarcomerogenesis. Therefore, in striated muscle, the state of assembly of MTs is correlated with the activity of the cell and more especially with the process of myofibrillogenesis, whether the muscle cells are in the myoblast (Auber, 1969; Toyama et al.. 1982; Tassin et al., 1985a,b), neonatal (Cartwright and Goldstein, 1985) or growing adult stage (Rappaport et al., 1984; Samuel et al., 1984; Watkins et al., 1987). V. Conclusions Great strides have recently been made in our understanding of the distribution of MTs in various physiopathological states and of their involvement in a broad spectrum of cellular processes. Furthermore, the dynamic nature of MTs appears to be highly controlled (Ginzburg and Littauer, 1984; Williams er al., 1986) and their function directly coupled to the pattern of expression of the different isotypes of tubulins and MAPs. Among the possible different mechanisms controlling MT dynamics, one may invoke the expression of multiple genes coding for the various tubulin isoforms (Ginzburg et al., 1985; Sullivan et al., 1984; Farmer el a/., 1984; Miller r t ul., 1987) and the different MAP subunits (Olmsted et al., 1986). Additional mechanisms may involve differential-splicing events as well as posttranslational modifications of these proteins. Functionally different MTs may arise from variabilities within the 15 carboxyl-terminal amino acids that could determine the specificity of interactions between /3-tubulin isoforms and the various MAPs (Cleveland and Sullivan, 1985). In rat brain, the number and pattern of isotubulins (Denoulet er al., 1982, 1986; Ginzburg and Littauer, 1985; Francon et al., 1982; Drubin et al., 1984; Olmsted et al., 1986), as well as the ability of different MAPS to promote in uitro tubulin polymerization (Olmsted, 1986), vary during development. However, in contrast to the large amount of research performed on brain tissue or on invertebrate animals, the constitutive elements of MTs in the heart have been poorly investigated. The pattern of tubulin isoforms has not yet been described in the heart and the nature of MAP subunits is still unclear. Moreover, nothing is known about the genomic expression of these proteins, nor the nature of the posttranslational events that most likely play a major role in MT growth within myocytes. The analysis of the behavior of cardiac MTs, in light of studies performed

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on other cell types, will help to provide a framework into which various hypotheses and/or various future lines of research may be incorporated. Overall synthesis of tubulin decreases rapidly during postnatal development in the two organs-heart and brain-that become almost mitotically quiescent after birth. An increase in the number of tubulin isoforms and a change in their relative proportions have been observed during brain maturation (Denoulet et al., 1982, 1986; Ginzburg and Littauer, 1984; Ginzburg et al., 1985). Changes in the genomic expression of many proteins such as actin (Minty et al., 1982; Mayer at af.,1984; Schwartz et al., 1986), myosin (LomprC et al., 1984), and creatine kinase (Ingwall et al., 1985) occur in the heart during postnatal development. It is likely that changes in the isotubulin pattern also occur in the heart during this period even if the pattern of tubulin isotypes in the heart is probably, as in other tissues, less heterogeneous than in brain (Gozes and Littauer, 1978). Change in brain tubulin microheterogeneity are regulated, at least in part, at the mRNA level (Cleveland and Sullivan, 1983). As yet, only the RNA sequences containing the translated region have been described in cardiac tissue, but the DNA sequence analysis has revealed that in many tissues the specificity of each p-tubulin mRNA sequence resides within its 3' untranslated region (Farmer et a l . , 1984). According to Cleveland and his co-workers (1983, tubulin heterogeneity associated with the specific expression of MAPS must be essential to specify the types of interactions in which MTs may participate in different cellular events. Vallee and co-workers (1984) have proposed that MAP2 are involved in the interactions between MTs and intermediate filaments. The very small amount of MAP2 observed in myocardium by Parysek et al. (1984a) agrees with our observations that in the mature myocyte there are only a few connections between MTs and desmin filaments (see Section 111). Among the microtubule-associated proteins, MAP4 are predominantly expressed in cardiac tissue and their relatively high abundance is probably related to the function of MTs common to supportive and connective tissue types (Parysek et al., 1984b). In most of the cells, new MTs arise from a centrosome or nucleation site. Nucleation sites are like permanently stabilized GTP ends (Kirschner and Mitchison, 1986) and allow MTs to grow toward the cell periphery with a uniform polarity (Euteneuer and McIntosh, 1981). It has been proposed that nucleation sites allow appreciable polymerization to occur below the critical concentration of tubulin and thus can act not only as a catalyst, but also to determine the MT number and hence the polymer mass (Kirschner and Mitchison, 1986). Centrosomes are rarely seen in adult cardiac myocytes, but are observed during early postnatal

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development (Forbes and Sperelakis, 1984). The prominent number of MTs arising from the whole nuclear area in hypertrophying adult cardiac myocytes suggests that nucleation sites are distributed on the whole surface of the nuclear membrane as in skeletal muscle cells (Tassin P I al., 1985a). Heterogeneity among MTs was detected using both their pharmacological properties (Behnke and Forer, 1967) and their immunological properties (Thompson er a l. , 1984). More recently, Schulze and Kirschner (1986, 1987) were able to distinguish two MT types that differed in the turnover of their subunits. One representing 10% of the whole population appears to be highly stable. The authors suggest that kinetic differentiations are likely to reflect functional distinctions. It was recently shown that alterations in tyrosination (or detyrosination) may effect rapid changes in the behavior of tubulin and MTs (Gundersen et af., 1987). In this article we have shown that in cardiac tissue, the state of assembly of some MTs correlates with the activity of the hypertrophying myocytes. One population of MTs distributed along sarcolemma and mitochondria does not vary in number with the activity of growth of the cell. whereas the second that consists of MTs localized around the nuclei and along myofibrils may change in density depending on the growth activity of the myocytes. Therefore, MTs distributed around nuclei (Watkins et al., 1987) and/or along developing myofibrils (Cartwright and Goldstein, 1985; Watkins et al., 1987) might correspond to an MT class more dynamic than the one composed of those MTs distributed along mitochondria and sarcolemma. Whatever the relative stability of each population, the observation of different MT families within myocytes suggests that in heart, as in other tissues, (Schulze and Kirschner, 1986, 1987) tubulin polymerizes into a diverse number of MT arrays whose assembly and functional properties are defined by specific programs of cellular differentiation (Cleveland and Sullivan, 1985). Neither genetic diversity nor a single mechanism can account for cellular control over the enormous versatility of MT functions. Although most of the extrapolations on the various roles of MTs in heart may turn out to be incorrect, they may nevertheless provide a good framework for further investigation into the role of MTs and more generally of the cytoskeleton within cardiac cells.

ACKNOWLEDGMENTS l h e authors are grateful to Mrs. F. Marotte for technical assistance and Miss P. Cagnac for secretarial assistance. Research was supported by INSERM and CNRS.

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INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 113

Functional Morphology of the Thyroid HISAOFUJITA Department of Anatomy, Osaka University Medical School, Osaka 530, Japan

I. Introduction Thyroxine (tetraiodothyronine, T4) and triiodothyronine (Ti), secreted from the thyroid follicular epithelial cell, are characteristic hormones because they are not peptides, amines, or steroids, but special amino acid derivatives (diphenyl ethers) derived from only two molecules of iodotyrosine linked by an ether bond (-0-) (Fujita, 1975, 1980). T4 and T3 are not synthesized directly in the cell from their constituents, tyrosine and iodine; formation of the hormones requires more complicated steps. Parts of their synthetic process take place in the special extracellular region named the follicular lumen. In addition, calcitonin and serotonin are also secreted from the thyroid gland, not from the follicular epithelial cell but from the parafollicular cell; this is not dealt with in this review. The thyroid gland is also a characteristic endocrine organ morphologically. The organ consists of an accumulation of characteristic ball-like structures called follicles and interfollicular connective tissues containing blood capillaries. Each follicle is composed of numerous follicular epithelial cells arranged as a simple epithelium, a follicular lumen surrounded by the epithelium, and a few parafollicular cells located singly or in groups outside of the follicular epithelium. The secretory cells in most other endocrine organs are not arranged in an epithelial manner and do not show any distinct cell polarity, but the thyroid follicular epithelial cells are arranged to form a typical simple epithelium and show a typical cell polarity quite similar to that of the usual exocrine-type cell. Follicular epithelial cells have two kinds of important activities for secretion; the secretion of thyroglobulin (high-molecular-weight glycoprotein) into the follicular lumen from the apical part of the cell by exocrine secretion, and the secretion of hormones (T4 and T3) into the connective tissue space from the basal part of the cell by endocrine secretion. The outline of the secretory process of thyroid hormones (T4 and T3) is summarized as follows (Fig. 1): (1) synthesis of thyroglobulin (glycoprotein) in the follicular epithelial cell, (2) release of thyroglobulin into the follicular lumen, (3) iodination of tyrosyl residues in the thyroglobulin molecule, (4) coupling of iodotyrosines in the thyroglobulin 145

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FIG. 1 . A schematic representation of the synthesis and secretory process of thyroxine (T,) and triiodothyronine (Ti). Synthesis and release of thyroglobulin into the follicular lumen (left). and reabosorption and hydrolysis of thyroglobulin with release of T4and T3 from the basal part of the cell are depicted (right). (R)Rough endoplasmic reticulum; (G) Golgi apparatus: (S) secretory granule: (F) follicular lumen: (M) micropinocytosis; (PI phagocytosis: ( V ) coated vesicle; (D) colloid droplet: (L) lysosome; (T) thyroxine: (B) basal lamina: ( C ) connective tissue space; (El endothelial cell.

molecule, ( 5 ) reabsorption of thyroglobulin from the follicular lumen into the follicular epithelial cell, (6) fusion of reabsorbed thyroglobulin (colloid droplet) with lysosomes. (7) hydrolysis of reabsorbed thyroglobulin and liberation of T4 and Tq, and (8) release of the hormones from the basal part of the cell into the connective tissue space and then into the blood capillary or lymph vessel. The present review deals with the characteristic ultrastructural and cytochemical functions of the follicular epithelial cell. The relationship between the uniqueness of the secreted hormones and that of the functional morphology of the thyroid gland and the follicular epithelial cell is also discussed.

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11. Synthesis and Release of Thyroglobulin

There are many molecular sizes of thyroglobulin: 3-8 S, 12 S, 19 S, and 27 S. Among them, 19 S (660 kDa), consisting of 5500 amino acids and 300 molecules of monosaccharide, is the thyroglobulin most commonly distributed in the thyroid gland throughout higher vertebrates. It is well known that 19 S thyroglobulin is composed of two subunits (Vassart et al., 1975; Edelhoch and Robbins, 1978). By electron microscopic (EM) examination of negatively stained monoclonal antibody-linked chains of hog thyroglobulin molecules, Kondo et al. (1985) determined that the two subunits, translated from a common mRNA (Vassart and Brocas, 1980), are equal in size and have a similar structure. Except for Cyclostomes, the follicular epithelial cells of vertebrates are characterized by well-developed rough endoplasmic reticula (RER) distributed in a manner similar to that of the exocrine pancreatic cell and stomach chief cell (Fig. 2). The elements of this organelle are widely distributed in the cytoplasm especially in the basolateral region of the cell, and their cisternae are usually enlarged in size and irregular in shape. This means that the synthesis of the exportable protein in this cell is as active as that in exocrine cells such as pancreatic acinar cells and stomach chief cells. Incorporation of amino acids and sugar components into this protein was demonstrated morphologically as follows. Five minutes after injection of [3H]leucine into the peritoneal cavity of the normal mouse, silver grains appear in the cisternae of RER by autoradiography (Fujita, 1975). The cisternae of this organelle are weakly positive to silver methenamine staining for carbohydrates (Tamura and Fujita, 1978). Thyroglobulin is biochemically known to be an N-linked glycoprotein. To make an N-linked glycoprotein, N-acetylglucosamine is first bound to asparagine in the peptide synthesized at the ribosome attached in the cisternae of RER, and then mannose is linked to the N-acetylglucosaminyl residue. By autoradiography, it was shown that [3H]mannose appears in the cisternae of the rough endoplasmic reticulum soon after injection (Whur et al., 1969). It thus appears that protein slightly glycosylated by mannose and N-acetylglucosamine occurs in the cisternae of rough endoplasmic reticula that are weakly positive for silver methenamine staining. Mannosyltransferase and N-acetylglucosaminyltransferase, which are necessary for mannosylation and N-acetylglucosaminylation of protein, have been demonstrated biochemically in the microsome fraction of sheep thyroids (Bouchilloux et al., 1970). The glycoprotein in the cisternae of RER is transported to the Golgi apparatus. The membrane of the rough endoplasmic reticulum facing the Golgi apparatus lacks ribosomes; small coated pits called buddings arise

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FIG.2. A part of thyroid showing follicular epithelium, follicular lumen, and interfollicular connective tissue with blood capillary ( B ) in a mouse 30 minutes after injection of thyroid-stimulating hormone (TSH). Note reabsorbed colloid droplets (D), lysosomes accumulating around the droplets, and well-developed rough endoplasmic reticulum in the cell. x4750.

from this membrane and transporting vesicles produced by pinching off the buddings are usually seen in the area between the RER and Golgi apparatus (Fujita, 1975) (Fig. 1). The glycoprotein in the cisternae of RER is believed to be transported to the Golgi apparatus through this pathway. By autoradiography, it has been shown that [3H]galactose and [3H]fuc o ~ are e bound to exportable glycoprotein in the Golgi apparatus of the thyroid cell (Whur ef al., 1969; Haddad et al., 1971). However, it is difficult to detect the specific site of the Golgi apparatus for glycosylation of the exportable glycoprotein by this method because of inadequate resolution. It was demonstrated that the 1-3 saccules of the trans (maturing) side of the Golgi apparatus were strongly positive to silver methenamine staining for carbohydrates (Tamura and Fujita, 1978) (Fig. 4) and were also positive for thiamine pyrophosphatase (TPPase) activity (Sawano and Fujita, 1981). Roth and Berger (19821, using HeLa cells, reported that

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uridine diphosphate (UDP) linked with galactose enters the cisternae of the Golgi saccules from the cytoplasmic matrix passing through the Golgi membrane and transfers galactose to the exportable glycoprotein by the enzymatic activity of galactosyltransferase. Then, the liberated UDP is disintegrated to uridine monophosphate (UMP) and phosphoric acid by uridine diphosphatase (UDPase) that is regarded as equivalent to TPPase. Finally, UMP returns to the cytoplasmic matrix passing through the Golgi membrane. If this view is generalized, the sites for galactosylation of protein in the thyroid cell must be the cisternae of trans-side Golgi saccules where TPPase is localized. Although galactosyltransferase has been demonstrated both biochemically and cytochemically in the Golgi saccules of the trans compartment in HeLa cells (Roth and Berger, 1982; Strous et al., 1983; Rothman, 1985) and sialyltransferase has been found in the trans-side Golgi tubular networks in kidney cells, fibroblasts, and hepatocytes (Berger and Hesford, 1985; Roth et al., 1985), morphological studies are needed to determine the site of these glycosylations for thyroid cells. As described above, thyroglobulin synthesized in the RER and Golgi apparatus system involves carbohydrate chains consisting of mannose, N-acetylglucosamine, galactose, fucose, and sialic acid. There are two kinds of carbohydrate chains in thyroglobulin, unit A and unit B. Both are linked with asparagine in the protein and 1 molecule of thyroglobulin contains 9 molecules of unit A and 14 of unit B. Using autoradiography of 35S04,Herzog (1985) noticed that thyroglobulin is sulfated in the Golgi apparatus and postulated that the sulfate residues contribute to the anionic state of thyroglobulin, operate in the regulation of thyroglobulin transport in the cell, and function in the tight packaging of thyroglobulin in the follicular lumen. From the trans-side Golgi saccules arise thyroglobulin-secretory granules that are round in shape and 150-200 nm in diameter. By autoradiography of [3H]leucine, it has been shown that the silver grains are localized on the Golgi apparatus and secretory granules 15-30 minutes after injection of the normal mouse and appear in the follicular lumen 45 minutes after injection (Fujita, 1975). Microtubules are considered to be necessary for intracellular transport of thyroglobulin-secretory granules based on [3H]fucose autoradiography data showing that treatments of colchicine or vinblastine inhibit the migration of the secretory granules to the cell apex (Bennett et af., 1984; Wild and Bennett, 1984). But, the detailed mechanism of the transport and release of the secretory granule and the role of microtubules and other cytoskeletons are problems for the future. In our laboratory, the occurrence of numerous fine filamentous cross-linkages between the limiting membrane of secretory granule and microtubule has been recognized in the anterior pituitary cell by a rapid

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freeze-deep etching method (Senda and Fujita, 1987); this finding could be generalized and applied to the thyroid cell. Using autoradiography of ['H]leucine, Ofverholm and Ericson (1984~) reported that the transport and release of thyroglobulin are regulated by thyroid-stimulating hormone (TSH). They calculated that in normal rats the half-life of exportable protein in the cisternae of the rough endoplasmic reticulum and Golgi apparatus is 28 minutes and in the secretory granules it is 18 minutes: in animals with hypofunctional thyroids, which are treated with thyroxine for 2 days, the half-life of exportable protein in the RER-Golgi apparatus cisternae is 63 minutes and in the secretory granules it is 62 minutes; when TSH is given to the thyroxine-treated animals 20 or 90 minutes before sacrifice, the half-life of exportable protein in the RER-Golgi apparatus is 25-33 minutes and in the secretory granules it is 9 minutes. The effect of cooling and monensin on the intracellular transport and secretion of thyroglobulin was studied by Ring et al. (1987a) using autoradiography of [3H]leucine and [3H]galactose. They stated that the transport of thyroglobulin from the RER to the Golgi apparatus, from the Golgi apparatus to the secretory granules, and from the secretory granules to the follicular lumen is markedly inhibited at 20°C as compared to 37"C, and speculated that this phenomenon is due to the inhibition of membrane fission and fusion. Monensin, an ionophore of Na'/K', has been known to perturb the structure of the Golgi apparatus (Tartakoff, 1983). In the thyroid cell treated with this agent, the Golgi saccules become vacuolated and granule formation is stopped. Ring et a/. (1987b) reported that this agent arregts transport of protein in the Golgi apparatus but does not inhibit transport from the Golgi apparatus to the follicular lumen of the thyroglobulin granules already produced. It is well known that peroxidase, which is localized at the periluminal region, is necessary for iodination of tyrosyl residues in the molecule of thyroglobulin. The localization of peroxidase has been detected by many investigators using histochemical methods. as discussed in Section 111. Although many different opinions have been reported, this enzyme is now believed to be localized in the periluminal colloid or apical plasma membrane. Both cytochemical and histochemical data suggested that this enzyme is derived from the rough endoplasmic reticulum and Golgi apparatus system (Nakai and Fujita, 1970; Strum and Karnovsky, 1970; Sawano and Fujita, 1981). If so, this enzyme should be transported like thyroglobulin, from the Golgi apparatus to the apical plasma membrane. It ha? not been determined whether or not this enzyme coexists with thyroglobulin in the secretory granule.

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Using the thyroid from rats pretreated with thyroxine for 2 days to inhibit endocytosis, Bjorkman et al. (1976) analyzed electrophoretically the exocytotic vesicles isolated by centrifugation. They determined that the uniodinated thyroglobulin and thyroalbumin contained in these vesicles are positive for peroxidase activity. We obtained the same results by EM histochemistry (Sawano and Fujita, 1981). In our study, all the secretory granules that accumulated in the apical cytoplasm after thyroxine treatment were strongly positive for peroxidase. From these facts, it is understood that the secretory granules contain thyroglobulin as well as peroxidase, both of which are derived from the rough endoplasmic reticulum and Golgi apparatus system. In aged mouse thyroids, cold follicles appear (Studer et al., 1978) that consist of attenuated epithelial cells and large follicular lumen. These follicles do not respond lo intensive stimulation from injections of exogenous TSH and lack the ability to accumulate iodine into the follicular lumen (Tamura and Fujita, 1981). Our observations show that in 10-month-old mice, the cold follicles occupy about 4.5% of the total number of follicles and in 19 to 20-monthold animals, about 20% (Tamura and Fujita, 1981) (Fig. 3).

FIG. 3. Cold follicles, which do not response to exogeneous TSH, in 10-month-old mouse. Note attenuated follicular epithelial cells. ~ 3 3 3 .

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111. Iodination of Thyroglobulin

The author wishes to emphasize that tyrosyl residues in the molecule of thyroglobulin are iodinated in the follicular lumen, especially in the peripheral region of the lumen (Fig. 4). For a long time, this problem has been discussed by numerous investigators using autoradiographic, cytochemical, and biochemical data (Leblond and Gross, 1948; Doniach and Pelc, 1949; Nadler and Leblond 1954; Stein and Gross, 1964; Ibrahim and Budd, 1965; Lupulescu and Petrovici, 1965; Simon and Droz, 1965; Nadler, 1965, 1971; Ekholm, 1966. 1981; Ekholm and Wollman, 1975; Fujita, 1969, 1972, 1975; Nunez, 1980: Ericson, 1983, and so on). By autoradiography of it was shown by most of these authors that silver grains appear almost entirely in the follicular lumen, especially in its peripheral region, within a few minutes after injection. Reviews on this problem have been published by Fujita (1972, 1975, 1976), Nunez (1980) and Ekholm (1981).

FIG. 4. An autoradiograph of cultured mouse thyroid 1 minute after immersion in a medium containing 600 pCi/ml of Na"'1 (A). and the thyroid of a hagfish, Eptatreius burger;. 1 hour after injection of 1 mmCi of Na"'1 (B). Silver grains are localized in the follicular lumen. ( A ) x 11.160; (B) x 10,450.

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Inorganic iodide is washed away by autoradiographic procedures such as washing, fixation, and dehydration of the tissue and only iodide that is bound to protein is detected by this method. In mouse thyroid cultured in a medium containing lZ5I, silver grains appear in the follicular lumen within 30 seconds and increase rapidly with time, but no grains were detected in the cell (Fujita, 1981) (Fig. 4). To detect the site for accumulation of inorganic iodide, the author used ''I autoradiography of freeze-dried sections of mice treated with methylmercaptoimidazole, which blocks the iodination of thyroglobulin, and showed that inorganic iodide trapped in the thyroid gland is localized mainly in the follicular lumen, although a few grains are found in follicular epithelial cells (Fujita, 1976). This indicates that most of the trapped inorganic iodide is transported to the follicular lumen where it is primarily stored. It is reasonable to assume that iodination of thyroglubulin does not occur in the cell but in the follicular lumen. In lower vertebrates, the main site of iodination of thyroglobulin is also the follicular lumen as reported by Nakai et al. (1970) for amphibia, by Nakai and Gorbman (1969) for chimeric fish, and by Fujita and Shinkawa (1975) for hagfish (Fig. 4). In larval lampreys and some amphioxi and ascidians, there is no typical thyroid gland showing follicle structure, but some thyroid-homologous cells having an activity for T4 and T3 synthesis are localized in certain regions of the endostyle, of a hollow or grooved structure of the pharynx. The main site of iodination of thyroglobulin or thyroglobulin-like protein is extracellular at the apical plasma membrane region of these thyroid-homologous cells in the endostyle, as reported by Fujita and Honma (1969) for larval lampreys, by Ericson et al. (1985) and Fredriksson et al. (1984, 1985b) for amphioxi, and by Fujita and Nanba (1971), Thorpe et at. (19721, Dunn (1974), and Fredriksson et al. (1985a) for ascidians. It is now believed that the iodinated protein is reabsorbed into the multivesicular bodies and lysosomes of these cells and that the hormone is released from the basal part of the cell (Fujita and Honma, 1969; Fredriksson et al., 1984). The enzyme necessary for iodination of tyrosyl residues in thyroglobulin is peroxidase. Although morphological aspects emphasize the occurrence of iodination in the follicular colloid, the microsome is considered the main site for thyroglobulin iodination by some biochemists, based on data showing that it is also the main site of peroxidase localization (Hosoya et al., 1972). Morphological studies for localization of peroxidase have been done by many investigators chiefly using the 3,3'diaminobenzidine tetrahydrochloride (DAB) method (Nakai and Fujita, 1970; Strum and Karnovsky, 1970; Tice and Wollman, 1972,1974; Nanba, 1972, 1973; Ofverholm and Ericson, 1984b; Yamashita et af., 1984). Their

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data are summarized as follows: peroxidase exists in the luminal colloid, apical plasma membrane, RER, Golgi apparatus, and subapical secretory granules. From these studies, we noticed that there are inconsistencies with the thyroglobulin iodination site and peroxidase localization. The problem arises as to why protein in the RER, Golgi apparatus, and secretory granule is not iodinated, although peroxidase activity is demonstrated in them. Strum and Karnovsky (1970) speculated that intracellular peroxidase might be in an immature form and becomes mature only after being released into the follicular lumen. There is another problem with the localization of peroxidase: determining whether the enzyme is strictly bound to the apical plasma membrane or is soluble in the colloid. Some biochemists (De Groot and Niepomniszcze, 1977; Matsukawa et af.,1981) and morphologists (Tice and Wollman, 1972, 1974; Ofverholm and Ericson, 1984a) speculated that peroxidase in the thyroid cell is membrane bound, and that luminal and cisternal peroxidase reactions do not show a soluble enzyme but a diffusion artifact produced by histochemical procedures. If so, there is the problem of whether or not the positive reactions for peroxidase in the contents of the secretory granule and cisternae of the RER and Golgi apparatus are also diffusion artifacts. If the enzyme is contained in the granules and cisternae of the cytoorganelles, it is reasonable to consider that the peroxidase being released by exocytosis is localized in the luminal colloid. The histochemical study €or peroxidase in the thyroid-homologous cell (endostylic cell) of the protochordate is interesting. Peroxidase activity has been demonstrated in the iodine-binding cells of the endostyle along the luminal plasma membrane and in the RER, Golgi apparatus, and subapical vesicles in amphioxi (Fredriksson et a / ., 1985b) and ascidians (Fujita and Sawano, 1979; Fredriksson et al., 1985a) (Fig. 5). The findings for the iodination site described above and for peroxidase localization are principally the same as those for the thyroid of the higher vertebrates. Hydrogen peroxide (H20~)is also necessary for iodination of thyroglobulin. An iodide ion taken up into the follicular epithelial cell is oxidized I?. the active form of iodine. In this reaction, H2O2acts as the electron acceptor. The active form of iodine binds with the tyrosyl residue in thyroglobulin. Bjorkman el a / . (1981) and Labato and Briggs (1985) using NADPH oxidase showed histochemically that H202 is generated on the apical surface of the follicular epithelial cell in the rat thyroid. Comparison of data from '"I autoradiographic and histochemical tests for peroxidase and HzOz emphasized that iodination of tyrosyl residues thyroglobulin takes place mainly at the peripheral region of the follicular lumen. Although about 120 tyrosyl residues are contained in a thyroglobulin

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FIG. 5. Peroxidase activity in the iodine-binding cell (type 7 cell) of an endostyle in the ascidian Ciona intesrinalis. Luminal plasma membrane, apical vesicles, and rough endoplasmic reticulum are positive. X 18,240.

molecule, only a few of them are iodinated and coupled and only a few molecules of tetraiodothyronine (T4) are made. A paper chromatogram of mouse thyroid using I3'I autoradiography showed that I3'I first appears in monoiodotyrosine (MIT) and diiodotyrosine (DIT) and then much later in T3 and Tq. The coupling of two molecules of tyrosyl residue in thyroglobulin occurs in the follicular lumen after the iodination of tyrosyl residues.

IV. Reabsorption of Colloid The luminal colloid thyroglobulin is reabsorbed into the follicular epithelial cell by micropinocytosis as well as phagocytosis (Fig. 1). After injection of TSH, pseudopods suggesting phagocytosis, in addition to micropinocytotic features, often appear at the apical surface of the follicular epithelial cell. In normal conditions, pseudopods and phagocytotic features are few although coated pits for micropinocytosis are usually seen.

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The micropinocytotic features could be principally distinguished from the exocytotic figures of the thyroglobulin secretory granules by the presence of a coat around the micropinocytotic pits or vesicles and the absence of a coat in the secretory granules. Herzog and Miller ( 1979a) and Denef and Ekholm (1980) suggested that micropinocytosis may be involved in membrane retrieval in addition to the reabsorption of colloid. Using the freeze-fracture method, we demonstrated aggregates of membrane particles on the P-face of small depressions corresponding to the initial site for coated pits (Ishimura et al., 1976a) (Figs. 6 and 7). Orci et al. (1978), who found accumulations of membrane particles at the initial site for pinocytosis of very-low-density lipoprotein (VLDL) in the fibroblast. believed these particles were implicated with the receptor protein for VLDL. However, it is not known whether or not the thyroid cell has a site to recognize and reabsorb only the mature thyroglobulin molecules in which tyrosyl residues and their couplings have been iodinated. The relationship between the cholesterol contents in the membrane and the distribution of intramembranous protein particles is of interest. Freeze-fracture images of filipin (antibiotic)-treated tissue have been used to detect the distribution of cholesterol in the membrane because filipin is bound with membrane cholesterol to make filipin-sterol complexes that show 25- to 30-nm protuberances on the fractured plasma membrane (Tillack and Kinsky, 1973; Verkleij et al., 1973). By this method, it has become clear that the membrane of the pinocytotic pit where the membrane particles aggregate, as mentioned above, lacks the filipinsterol complexes although most other parts of the plasma membrane are very rich in cholesterol (Fujita et al., 1981). Similar results have been reported for cultured fibroblasts and hepatocytes by Montesano et ul. (1979). This suggests that in the thyroid cell the initial site for micropinocytosis of the luminal colloid is an active region of the plasma membrane. By immunohistochemistry, actin was demonstrated in the apical cytoplasm of the follicular epithelial cell of the rat thyroid (Fujita et a l . , 1984) (Figs. 8 and 9). It is well known that cytoplasmic actin filaments are decorated by treatment with heavy meromyosin (HMM) showing arrowhead structures with a constant polarity (Ishikawa et al., 1969). Using this method, we demonstrated numerous actin filaments at the apical cytoplasm of the follicular epithelial cells of mice (Miyagawa et al., 1982b). The actin filaments attached to the apical plasma membrane run perpendicularly toward the network of actin filaments in the subapical region. Coated pits and coated vesicles are often connected with these filaments running toward the deeper regions of the cytoplasm (Fig. 8). The coated vesicles are therefore considered to be pinched off and internal-

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FIG. 6. Thin section image (A) and freeze-fracture image (B) of micropinocytosis of the luminal colloid in a TSH-treated mouse. Note accumulation of membranous particles at the initial site of rnicropinocytosis in the freeze-fracture image. (A) x 110,000; (B) x 120,000. From Miyagawa et al. (1982b).

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FIG. 7. A schematic representation of micropinocytosis. Membranous particles are accumulated at the initial site of micropinocytosis.

ized to coated pits. The actin filament system might be involved in this process. Gabrion et a / . (1980) have also reported a close association between actin filaments and apical vesicles. We clearly demonstrated by immunocytochemistry that caldesmon is located in the apical cytoplasm of almost all follicular epithelial cells in the rat (Fujita et af., 1984)(Fig. 9). Immunoreactivities for both actin and caldesmon showed almost the same pattern of localization. Caldesmon, discovered by Sobue et a / . (1981,

FIG. 8. Decorated actin filaments in contact with a subapical coated pit (A) and coated vesicles ( B ) in TSH-treated mouse thyroid immersed in heavy meromyosin. (A) x 11,520; (B) <9,600.

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FIG. 9. Immunofluorescent staining of caldesmon (A) and actin (B) in mouse thyroid. Both are positive at the apical cytoplasm of follicular epithelial cells. (A) X 108; (B) x 135.

1982) in the smooth muscle cell, is a calmodulin-binding protein composed of M4 150K and 147K subunit polypeptides that are able to bind to F-actin filaments. It is well known that the binding of caldesmon to F-actin is inhibited by Ca2+-dependent binding of calmodulin to caldesmon and that the binding of caldesmon to calmodulin or F-actin can alternate depending upon the concentration of Ca2+, resulting in the flip-flop binding hypothesis (Kakiuchi and Sobue, 1983). Biochemically, both myosin and troponin C-like phosphodiesterase activators (calmodulin) were purified in the thyroid gland by Kobayashi et al. (1977, 1979). From these results, it is easily understood that caldesmon, actin, and calmodulin play a major role in the regulation of cell motility such as micropinocytosis, phagocytosis, and exocytosis of the follicular epithelial cell (Fujita et a/., 1984). Recently, calspectin (fodrin, a nonerythroid spectrin-like protein) was immunocytochemically demonstrated in the follicular epithelial cell, especially along the basolateral plasma membrane in normal rats (Ishimura et al., 1987). Spectrin underlying the plasma membrane of the erythrocyte is known to participate in the maintenance of that cell’s unique shape (Branton at al., 1981); calspectin in the follicular epithelial cell might function to maintain its follicular structure. In TSH-treated animals, the apical plasma membrane region of follicular epithelial cells

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becomes positive for calspectin (Ishimura et al., 1987). Calspectin may also play a role with calmodulin and actin in the conjugation activities of the subapical region of this cell, such as secretion and reabsorption of thyroglobulin. Phagocytosis of the luminal colloid by follicular epithelial cells is also involved in the reabsorption of colloid. After injection of thyroidstimulating hormone (TSH), cytoplasmic protrusions, like pseudopods, appear in the apical cytoplasm (Wetzel et al., 1965; Seljelid, 1967; Fujita, 1975). Pseudopods projecting into the follicular lumen are very rare in normal animals but occur often in animals receiving large doses of TSH. This suggests that the luminal colloid is reabsorbed mainly by micropinocytosis under normal conditions and that phagocytotic activity appears in this cell only during the strongly stimulated state. In addition to TSH, dbcAMP and forskolin have the same effect on the follicular epithelial cell. It is believed that the TSH receptor is in the basolateral plasma membrane and that the hyperactive reaction of the follicular epithelial cell to TSH is mediated by cyclic AMP. The role of actin filaments in phagocytosis of luminal thyroglobulin (colloid) has been reported, Neve et al. (1972) showed that in TSH- or dbcAMP-stimulated slices of dog thyroid the release of thyroid hormone and the phagocytosis of colloid are inhibited by cytochalasin B, which is an inhibitor of the polymerization of actin. We also demonstrated the appearance of actin and caldesmon in the pseudopod of the rat follicular epithelial cell after injection of TSH (Fujita et al., 1984). Microtubules may also be implicated in colloid phagocytoid activity in this cell. The phagocytosis-inducing effect of TSH in the luminal colloid is inhibited by treatment with colchicine, vinblastine, vincristine, ethanol, and deuterium oxide, which are microtubule inhibitors (Neve et a / . , 1972). Although these data suggest a role for actin filaments and microtubules in phagocytosis, further studies are needed to clarify the mechanism. Inverted follicles are useful for studying the mechanism of phagocytotic activity. It is well known that isolated follicles of the thyroid are made by digestion of interfollicular connective tissue elements with collagenase and trypsin, and that the epithelial cells of isolated follicles cultured in suspension reverse their cell polarity under some conditions (Maucharnp et al., 1979; Inoue et al., 1980; Nitch and Wollman, 1980a; Herzog and Miller, 1981; Garbi and Wollman, 1982; Miyagawa et al., 1982a). In our scanning and transmission EM studies, it was demonstrated that the apical surface facing the culture medium, which showed numerous microvilli and a central cilium, is phagocytotic for erythrocytes and latex beads (Miyagawa et af., 1983). When the inverted follicles were incubated in a medium containing TSH (50 mU/ml) and human red blood cells or

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TSH and polystyle latex beads (2.02 pm in diameter) for -1-3 hours, numerous red cells or latex beads were taken up by the follicular epithelial cells (Fig. 10). Zeligs and Wollman (1977a) reported erythrophagocytosis by thyroid cells in rats fed a diet containing thiouracil, which underwent a microhemorrhagic process resulting in deposition of red cells in the follicular lumen. In addition, Rodesch et al. (1970) and Kowalski et al. (1972) observed the phagocytosis of latex beads by isolated thyroid cells. These facts demonstrate that the phagocytotic activity of the thyroid epithelial cell is not specific for lurninal thyroglobulin and suggest that thyroglobulin, either with or without iodinated and coupled tyrosyl residues, might undergo random phagocytosis by the follicular epithelial cell, although mature thyroglobulin might be selectively reabsorbed by micropinocytosis as coated pits. Ericson el al. (1979), Engstrom and Ericson (1981), and Johanson et al. (1984) reported that the apical plasma membrane, after treatment with TSH, is enlarged by exocytosis of secretory granules then reabsorbed by endocytosis, by either micropinocytosis or phagocytosis. Using rats treated with T4, whose thyroid cells show retention of numerous secretory granules in the apical cytoplasm due to suppresion of secretory

FIG.10. Endocytotic erythrocyte (A) and latex beads (B) in the follicular epithelial cells of inverted follicles isolated from a porcine thyroid. Lysosomes (dense granules) are in contact with the latex bead (right). (A) X4800; (B) X 17,000. From Miyagawa et a / . (1983).

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activity by feedback mechanism, Ericson et al. (1979) and Engstrom and Ericson (1981) demonstrated stereologically that TSH treatment induces an increase in exocytosis of these granules and an increase in the quantity of apical plasma membrane that is soon reused for endocytosis of the luminal colloid. Engstrom and Ericson (1981) showed that exocytosis precedes endocytosis, that the rate of exocytosis is linearly correlated with the logarithm of the TSH dose, and that the total membrane surface area of endocytotis structures (pseudopods, colloid droplets, and micropinocytotic vesicles) is related to the surface area added to the apical plasma membrane by exocytosis. Aminopeptidase N has been demonstrated at the apical plasma membrane of the porcine thyroid cell (Feracci et al., 1981; Hovsepian et al., 1982). This enzyme might be used in the catabolism of thyroglobulin after undergoing endocytosis (Hovsepian e? al., 1982). It was speculated that the luminal thyroglobulin molecule is resistant to aminopeptidase N degradation because the N terminal group is masked (Marriq et al., 1977), and that this enzyme, which is localized at the limiting membrane of the reabsorbed colloid droplet, might be partly responsible for the hydrolysis of reabsorbed thyroglobulin. Another interesting phenomenon is transcytosis, found by Herzog (1983) using the inside out follicle and either a ferritin cation or thyroglobulin-gold complex tracer. In the inside out follicle, in which epithelial cell polarity is reversed, the tracer undergoes endocytosis by the apical cell membrane of the follicular epithelial cell facing the culture medium. Herzog and Miller (1979a, b, 1981) demonstrated that a part of this membrane is fused with endosome and lysosome membranes and becomes inserted into the membrane of the Golgi apparatus within 30 minutes. In addition to this main pathway, Herzog (1983, 1984) found that ferritin cations and colloid particles in the culture medium undergo endocytosis into the coated pits of the basal and lateral plasma membrane within 16 and 1 1 minutes, respectively, which demonstrates transcytosis, the direct pathway from apical to basolateral. This suggests that thyroglobulin partly enters the tissue fluid in the connective tissue space directly passing through the cell, and then enters the blood capillary and lymph vessel. As described in the review by Herzog (1984), thyroglobulin was demonstrated by radioimmunoassay in the blood circulation (Roitt and Torrigiani, 1967) and lymphatic system (Daniel et al., 1967; Kotani et af., 1968)of the normal rat and human. The thyroglobulin level in the blood is raised by stimulation with TSH and hyperthyroidism (Uller er a / . , 1973; van Herle et al., 1975) and suppressed by the addition of thyroid hormone (van Herle et al., 1973). Because the follicular epithelium is tightly sealed

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by well-developed zonula occludens (Fujita et al., 1975; Tice et al., 1975; Thiele and Reale, 1976; Zeligs and Wollman, 1977a,b; Ishimura and Fujita, 1979), the main pathway for release of thyroglobulin from the follicular lumen into the blood circulation and lymph vessel is by transcytosis, the vesicular transport across the follicular epithelial cell proposed by Herzog (1984).

V. Hydrolysis of Thyroglobulin and Release of T4 and T3 Reabsorbed colloid (thyroglobulin) droplets, round, oval, or irregular in shape and variable in size (0.5-5.0 pm in diameter), are sometimes seen in the apical cytoplasm of follicular epithelial cells of normal and TSHtreated animals. They increase in number and size after injection of TSH, often fusing with each other to become larger. The micropinocytotic vesicles derived from the coated pits become uncoated and fused with one another and also with the large colloid droplets (Fig. 11). By the rapid freeze substitution method, we observed the fusion of colloid droplets to proceed as follows: at the site of fusion, the outer layers of limiting membranes of two adjacent colloid droplets form a pentalaminar structure that becomes trilaminar, then disappears at the contact area (Miyagawa er al., 1982b). The colloid droplets eventually link with lysosomes to be hydrolyzed and liberate T4 and T3 (Fig. 12). The colloid droplets are fused directly with lysosomes, sometimes becoming enwrapped by them before fusion,

FIG. 11. A schematic representation of the fusion of reabsorbed colloid droplets.

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Fic;. I?. Acid phosphatase activity in the follicular epithelial cell of the TSH-treated mouse. Note primary lysosomes localized around colloid droplets and a phagolysosome (colloid droplet positive for acid phosphatase). ~ 3 3 , 6 0 0 .

as reported in histiocytes and hepatocytes during the formation of autophagolysosomes described by Mayahara (1972) and Abe and Ogawa (1980). The fresh colloid droplets reabsorbed from the follicular lumen are relatively low in electron density, whereas the primary lysosomes are homogeneously electron dense; the colloid droplets fused with lysosomes, which could be named phagolysosomes, are usually heterogeneously electron dense. The acid phosphatase activity is positive in both primary lysosomes and phagolysosomes, but the fresh colloid droplet is negative. I n addition to lysosomes, 1-3 trans saccules of the Golgi apparatus and vesicles in the trans region are also positive for acid phosphatase (Sawano and Fujita, 1981). The localization of this enzyme activity almost overlaps that of TPPase. This suggests that lysosomes are derived from trans saccules of the Golgi apparatus. Reabsorbed thyroglobulin is hydrolyzed

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by proteolytic enzymes originated from lysosomes, and tetraiodothyronine (T4), triiodothyronine (T3), and various kinds of amino acids are liberated. This specialized hydrolytic reaction for reabsorbed thyroglobulin produces hormones (T4 and T3) from prohormones (thyroglobulin). The secretory vesicles containing T4 and T3 have not yet been detected around the phagolysosome or in the basal cytoplasm of the follicular epithelial cell. The hormones may be transported from the phagolysosome to the basal plasma membrane passing through the cytoplasmic matrix and released into the connective tissue fluid. T4 and T7,which are amino acid derivatives of 777 and 651 molecular weight, respectively, seem not to have granular structure because of their small molecular size compared to other proteinaceous secretory materials having granular form. Even if the hormones are bound with carrier protein, they are not packed by a limiting membrane as a vesicle, and might be transported through the cytoplasmic matrix. Secretory vesicles and exocytotic figures are difficult to find at the basal part of the cell. T4 and T3, which are not easily solubilized by water but are soluble in organic solvent, could pass through the plasma membrane. It is believed that T4 and T3, liberated from the carrier protein at the basal plasma membrane region, pass directly through the membrane into the connective tissue space. If this view is correct, the release mechanism of T4 and T3 corresponds to a diacrinous (transfusion) type proposed by Kurosumi (1961). However, further studies are needed on the liberation of T4 and T3 from the phagolysosome and on their release mechanism. The fate of other amino acids liberated by hydrolysis of reabsorbed thyroglobulin is also obscure; some of them might be reused for synthesis of new thyroglobulin, and others might be released from the basal part of the cell into the tissue fluid, When we inject [3H]leucineinto the animal, a fairly large portion is incorporated into the follicular epithelial cell. This suggests that the liberated amino acids are not sufficiently reused in this cell. In young mice, reabsorbed colloid droplets seem to be quickly and completely hydrolyzed, and a few primary lysosomes that are small in size and homogeneously electron dense are seen in the apical cytoplasm. Lysosomes increase in number and size with age; many large secondary lysosomes, containing heterogeneously dense materials such as lipid-like droplets and fibrous or lamellar structurs, are usually seen in old mice (Fujita et al., 1980). In addition, the specific activities of acid phosphatase, p-glucuronidase, and p-galactosidase show a relative decresae with age after 6 months. Some of the lysosomes observed in old animals, especially in those over 20 months of age, are residual bodies that could

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not be digested further (Fujita er af., 1980). Cell fragments derived from degenerated cells are sometimes seen in the follicular lumen of old animals. In addition, shedding of apical plasma membrane and dense cell fragments was found in the follicular lumen, especially in animals with hyperplastic goiter (Tachiwaki and Wollman, 1982; Nilson er al., 1984). These fragment formations are mechanisms for disposal of excess plasma membrane deposited by exocytotic vesicles during rapid secretion of thyroglobulin into the follicular lumen (Tachiwaki and Wollman, 1982). These cell fragments also undergo endocytosis and become phagolysosomes or residual bodies.

VI. Connective Tissue Space and Vascularization There are two basal laminae (one belonging to the thyroid cell and the other to the endothelial cell) and a connective tissue space between the basal plasma membrane of the follicular epithelial cell and that of the endothelial cell. Thyroxine (T4)and triiodothyronine (T3), released from the basal part of the follicular epithelial cell, pass through the basal lamina and enter the loose connective tissue fluid. They then enter the blood or lymph stream by passing through the endothelial fenestrations of capillaries in the region. Light microscopy clearly shows that blood capillaries are localized in the interfollicular connective tissue. However, the mode of distribution, namely, the three-dimensional relationship between blood capillaries and follicles, is difficult to understand by light or transmission electron microscopy (TEM). Our scanning electron microscopic (SEM) studies of methacrylate resin corrosion casts have shown that each follicle is encapsulated by a well-defined, basket-like capillary network that is generally independent of adjacent networks, although a few anastomoses or common capillaries are sometimes seen between them in monkeys (Fujita and Murakami, 1974; Shimada, 1981) and rats (Imada et al., 1986a,b) (Fig. 13). This suggests that each follicle functions somewhat independently of other follicles, and whole cells in each follicle function synchronously. In TSH-treated, low iodine-treated, or propylthiouraciltreated rats, whose thyroids are overstimulated by TSH overproduced by feedback mechanism, we demonstrated that the network of capillaries becomes markedly dilated and fused (Imada et al., 1986a) (Fig. 14), whereas in levothyroxine sodium-treated animals, whose thyroids are hypofunctional by feedback mechanism, the capillaries become markedly poor in distribution and anastomosis and conspicuously small in diameter (Imada et al., 1986b). In all of these experimental animals, the capillary of the parathyroid does not change under these experimental conditions.

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FIG. 13. A scanning electron micrograph of a corrosion cast of blood vessels in a rat thyroid. Note the basket-like network of blood capillaries for each follicle and for a lobule (arrow). X67.

The capillary endothelial cells of the thyroid are fenestrated as those of other endocrine organs in higher vertebrates. It is believed that hormones secreted from endocrine cells enter the capillary lumen through the endothelial fenestrations. Each fenestration, about 50 nm in diameter, is closed by a diaphragm having a small central hole of about 7 nm in diameter. The number and density of fenestrations are changed by the

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functional state and secretory activity of the organ. Our freeze-etching studies showed that both number and density are markedly increased in TSH-treated mice and decreased in Thyradin (powdered thyroid)-treated animals (Ishimura et al., 1976b) (Fig. 15). In the thyroid of the hagfish and its homologous organ, the endostyle, of the larval lamprey, whose secretory activities are very weak, blood capillaries are poorly distributed and endothelial cells are nonfenestrated (Fujita and Honma, 1966, 1968, 1969; Fujita, 1980). Thus, fenestrations in the endothelial wall are closely implicated in the secretory activity of the thyroid cell. In the endocrine organ, the blood capillary is morphologically flexible and changeable and reflects the functional state of the gland’s secretory activity. Wollman and his co-workers (1978) found fusion of capillary walls, in the form of partial septa, in the capillary lumen of the thyroid in rats fed a low-iodine diet containing 0.25% thiouracil. The mitotic process of endothelial cells, the increase of RER in these capillary cells and pericytes, and the fusion and enlargement of blood capillaries during the development of thyroid hyperplasia in these experimental animals have been described in detail (Zeligs and Wollman, 1981; Ericson and Wollman, 1980a,b). There are fewer reports on the lymph vessels of this organ (Rienkoff, 1931; Bargmann, 1939; Nakachi, 1972; Shimada, 1981) than on its blood vessels. By SEM microscopy of HCI-macerated and collagenase-digested thyroid, Shimada (I98 1) observed the three-dimensional architecture of lymph vessels of the monkey and compared it with thin section images. He described the lymph capillaries, which are lined by very thin endothelial cells, as quite variable (20-100 pm) in diameter and forming a loose network among the follicles. He calculated that each mesh of the lymph capillary network in monkey thyroid contains 1-6 follicles. This finding suggests that the fluid in the connective tissue could enter lymph vessels as well as blood vessels. The papers by Daniel et al. (1967) and Kotani et al. (1968), demonstrating thyroglobulin in the lymph vessel, support the finding tht thyroid hormones could also partly enter the lymph stream. TEM of thin sections shows a very thin (usually less than 2 pm in width) connective tissue space between the basal laminae of the thyroid epithelial cell and the capillary endothelial cell and very few cellular elements in the connective tissue, although nonmyelinated nerve fibers, thin processes of the fibroblasts, and collagen fibrils are rarely recognized. The other side of the blood capillary faces a large connective tissue

-

FIG. 14. Scanning electron micrographs of a corrosion cast of blood capillaries in the thyroid of a normal rat (A) and in a rat fed a low-iodine diet for 20 days (B). (A) x428; (B) x608. From Imada et al. (1986a).

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FIG. 15. Freeze-fracture images of endothelial fenestrations in a normal mouse (A) and a TSH-treated (1 .S Uiday for 6 days) mouse (B). Note an increase in number of fenestrations in the TSH-treated animal. (A) ~30,030;(B) x I1,550.

space that is rich in formed elements such as fibroblasts, mast cells, and collagen fibrils, which corresponds to the interfollicular connective tissue space in strict meaning. The fibroblasts, having long, thin, and branching cytoplasmic processes, are often in contact with the outside of the capillary endothelium. There are both a basal lamina and narrow space between this cell and the endothelial cell, and the fine structure is similar to a typical fibroblast having both a well-developed rough endoplasmic reticulum and Golgi apparatus. In the pericapillary region, these fibroblasts have been called pericytes based on SEM observation of thyroid treated with HCl and collagenase for removal of connective tissue fibers

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(Shimada, 1981). Their cytoplasmic processes are in contact to form a network. Recently, the occurrence of a network of cytoplasmic processes of fibroblasts has been noted in the lamina propria of the intestine (Desaki et al., 1984)and also in the corneal stroma (Fujita et af., 1987;Ueda et aE., 1987). Cytoplasmic processes of fibroblasts in the cornea are connected by gap junctions (Fujita et al., 1987;Ueda et al., 1987). The modes of distribution and connection of fibroblasts and their cytoplasmic processes in the thyroid gland remain to be studied.

VII. Nerve Supplies The thyroid is innervated by three kinds of nerve fibers, sympathetic, cholinergic, and peptidergic, although few fibers actually enter this organ. Sympathetic fibers that terminate at both the blood vessel wall and the follicular epithelial cell have been reported by many investigators (Melander et al., 1974, 1975;Tice and Creveling, 1975;Uchiyama et al., 1985). Using EM autoradiography of ~-[~H]norepinephrine, Melander et al. (1974) found the sympathetic nerve terminal labeled by this radioisotope to be in close contact to the follicular epithelial cell, via the basal lamina in human thyroid, and the smooth muscle of the blood vessel. They also demonstrated immunohistochemically that the sympathetic nerve fibers terminating around the blood vessel exist as a network, whereas those at the thyroid cell exist as a single fiber in rat, mouse, hamster, dog, sheep, and pig (Melander et al., 1975). Among these species, thyroid innervation by sympathetic nerves has been reported to be relatively poor in rat and mouse but poorest in pig. Using false transmitter (6-hydroxydopamine and 5,7-dihydroxytryptamine)-treated mice, Tice and Creveling (1975) ultrastructurally demonstrated direct contact of the sympathetic nerve terminal to the follicular epithelial cell. However, they had difficulty observing this contact because of its inconspicuous appearance, and concluded that only one to three nerve endings were present per innervated follicle. The author agrees with this opinion, having found it very hard to locate a nerve terminal in contact with a secretory cell during 30 years of thyroid research. Melander et al. (1972, 1974) reported that the sympathetic nerve may stimulate the thyroid follicular epithelial cell to secrete thyroxine. Cholinergic nerve endings containing noncored small vesicles were rarely observed in the interfollicular connective tissue of the mouse thyroid (Fujita, 1975). Although the nerve terminal is closely associated with the follicular and capillary endothelial cells via basal laminae, direct contacts between them are difficult to recognize. Acetylcholine might be

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released into the pericapillary tissue fluid and indirectly stimulate the follicular epithelial cell and the capillary endothelium (Fujita, 1975). Peptidergic nerve fibers containing vasoactive intestinal peptide (VIP), substance P, neuropeptide Y, and peptide HI were demonstrated in the interfollicular connective tissue by light microscopic immunohistochemistry, and VIP neurons were reported to be characteristically numeroub if Brattleboro rats that lacked vasopressin synthetic activity and suffered from diabetes insipidus (Hedge ef al., 1984). Since it is known that VIP enhances thyroid secretion (Ahren et al., 1982) and that thyroids of diabetes insipidus rats lack responsiveness to TSH. Hedge et al. (1984) reported that a striking increase of VIP fibers in these thyroids would indicate compensation. In normal rat thyroid, Uchiyama et al. (1985) found only very few nerve terminals containing flattened and large-cored vesicles of 95.1 nm in mean diameter, which are directly in contact with the follicular epithelial cell, and regarded them as peptidergic nerve fibers. Using serial sections, they estimated that only one to two nerve endings are in contact with the follicle. The occurrence of the subsurface cisterna-like endoplasmic reticulum immediately under the plasma membrane that faces the membrane of the nerve ending was reported (Uchiyama rt al., 1985), although its functional significance remains to be determined. Few thyroid follicular epithelial cells and blood vessels are innervated by sympathetic, cholinergic, and peptidergic nerve fibers compared with those of the adrenal medulla and pancreatic islet, in which direct contacts between nerve terminals and secretory cells are frequently seen. It should be emphasized that the main route for stimulating the follicular epithelial cell to secrete T4 and T3 is humoral, through the thyrotropin-releasing hormone (TRH)-TSH system. VIII. Follicular Cell Polarity and Inverted Follicles

The follicular epithelial cell of the thyroid gland is different from the secretory cells of almost all other endocrine organs in cell polarity. No distinct cell polarity is found in most endocrine cells, such as the anterior pituitary, pancreatic islet, parathyroid, and adrenal, whereas a clear polarity exists in the follicular epithelial cell. The cell surface facing the follicular lumen is apical, and that facing the basal lamina and connective tissue is basal. In the thyroid cell, many microvilli are protruded from the apical surface. well-developed zonula occludens (tight junctions) are present at the apical end of the lateral plasma membrane, RER is well

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developed in the infranuclear and lateronuclear cytoplasm, Golgi apparatus is usually situated at the supranuclear cytoplasm, and secretory granules are localized in the subapical cytoplasm. These structures and their organization are those of a typical exocrine cell, and thyroglobulin is secreted from the apical part of the cell by the mode of exocrine secretion. The thyroid cell is also similar to the intestinal absorptive cell in absorptive function. Luminal thyroglobulin is reabsorbed from the apical surface into the cell, and the hormones Tq and T3 are first released from the basal part of the cell into the connective tissue and then into circulation, whereas amino acids, carbohydrates, and triglycerides are released into circulation from the basal part of the intestinal absorptive cell. The occurrence of zonula occludens between thyroid follicular cells was described by Fujita et al. (1975) in rabbits, by Tice el al. (1975) in rats, and by van Uijen et al. (1985) in goats (Fig. 16). In freeze replica

FIG. 16. Freeze-fracture image of well-developed zonula occludens (0)and gap junctions (G) between follicular epithelial cells in the thyroid of a 19-day-old chick embryo. ~ 4 1 , 2 8 0From . lshimura and Fujita (1979).

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images from these animals, this structure of the thyroid cell consists of 5-15 strands and belongs to the “very tight form” in the classification by

Claude and Goodenough (1973); the luminal colloid is strongly sealed outside of the connective tissue fluid. In addition, well-developed gap junctions are usually seen between follicular epithelial cells. This suggests that all cells in each follicle may work synchronously by exchanging information through these structures. In ontogenetic studies, it has been shown that the zonula occludens is present between the cells before the primitive follicular lumen appears (Hilfer. 1964; Fujita and Tanizawa, 1966; Calvert and Pusterla, 1973; Tice el al., 1977: Ishimura and Fujita, 1979). Our studies showed that the primitive follicular lumen first appears within the region of a macula occludens formed between two adjacent cell cord cells in 8-day-old chick embryos, and that the macula occludens becomes the zonula occludens when the primitive follicular lumen appears in it (Ishimura and Fujita, 1979). The cell surface facing this lumen protrudes microvilli and the Golgi apparatus is usually situated between the nucleus and cell surface. Thus. it is reasonable to conclude that the exocrine cell-like polarity of the cell begins to form when the primitive follicular lumen appears in the cell cord (Fujita, 1984). Each follicle is continously surrounded by a basal lamina. It is well known that this structure, consisting of laminin, type 4 collagen, fibronectin, and so on, is synthesized in the rough endoplasmic reticulum and released via small vesicles of the follicular epithelial cell. The follicular epithelial cell is believed to interact with mesenchymal elements, such as type 1 collagen and fibroblasts, to produce the basal lamina (Alquier et al., 1979). An explanation of the inversion of cell polarity follows (Fig. 17 and 18). The experimental method to obtain the special type of follicle with reversed polarity epithelial cells has been established (Mauchamp et ul., 1979; Nitch and Wollman, 1980a,b; Inoue et af., 1980; Herzog and Miller, 1981; Miyagawa rt al., 1982a; Garbi and Wollman, 1982). These follicles are classifed into two types: one is a true inverted follicle and the other is an inside out follicle. Inversion of thyroid follicles takes place constantly when they are isolated by digestion of connective tissue elements and cultured in cuspension in Eagle’s medium containing fetal calf serum. inverted follicles are made by the following method. The thyroids of large animals, such as pigs, are cut into small pieces with razor blades and put into Hanks’ balanced salt solution (BSS) oxygenated at 4°C. The tissues are incubated in Eagle’s minimum essential medium (MEM) containing 0.1% collagenase (Warthington, type 11) for 30 minutes at 37°C in an atomosphere of 95% 0 2 - 5 % CO?, then reincubated in Ca2+-and Mg’+-

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FIG. 17. A schematic representation of a normal (left) and inverted (right) follicle. Note changes in the situation of microvilli (M), nucleus (N), Golgi apparatus (G), and zonula occludens.

free Hanks’ BSS containing 0.2% trypsin in (DIFCO) and 0.004% EDTA for 30 minutes at 37°C. After this enzymatic digestion procedure is repeated three times, the tissues are sifted by Cellector tissue sieve (Bellco) (mesh size 230 pm) and isolated follicles are collected by centrifugation (15 g for 5 minutes). Inside out follicles are obtained in the following way (Herzog and Miller, 1979a): thyroid tissue is dissociated into segments of follicle wall by digestion with collagenase and application of shearing force, these segments are cultured in suspension for 24 hours, and all of the segments are transformed into follicles with reversed polarity. In the inverted follicle, the luminal surface loses its microvilli, whereas the outer surface facing toward the culture medium protrudes numerous microvilli, the zonula occludens moves from the luminal end to the culture medium side of the cell’s lateral margins, and the Golgi apparatus becomes located in the cytoplasm between the nucleus and outer surface of the cell (Miyagawa et al., 1982a) (Fig. 18). The migration of strands of zonula occludens is initiated by the reversal of cell polarity (Kitajima el al., 1985). It is clear that the elements of zonula occludens, located at the luminal end of the lateral plasma membrane, spread in a basal direction and disperse in the lateral plasma membrane during 3 hours of incubation. During the next 24 hours, they gather toward the culture medium side of the lateral membrane and protrude microvilli; the movement of the Golgi apparatus is complete by that time (Kitajima et al., 1985). It is believed that either basal lamina or collagen is the most important factor for maintaining normal cell polarity, and that this polarity becomes unstable after digestion of this structure and the connective tissue. As reported by Nitch and Wollman (1980a, b) and Miyagawa et af. (1982a), polarity of isolated follicular cells is retained in medium containing 0.5%

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FIG. 18. A scanning electron micrograph of an inverted follicle of porcine thyroid. Note numerous microvilli protruding from the outer suiface of the follicle. ~ 5 0 0 0 .From Miyagawa et nl. (1983).

fetal calf serum and reversed in medium containing 5 or 10% fetal calf serum; therefore, the concentration of the serum in the culture medium plays an important role in reversing cell polarity, although the mechanism is not clear. When inverted follicles are embedded in a collagen gel suspension culture, extensive reorganization occurs (Chambard et al., 1984; Garbi et al., 1986). Cell polarity returns to normal, which shows the important role of collagen in determining the basal pole of the cell. The inverted follicle was used in the detection of the endocytotic mechanism (Miyagawa et al., 1983), and the inside out follicle contributed to in the discovery of transcytosis (Herzog, 1983, 1984). Both have been described in Section IV. When isolated thyroid cells or preformed monolayers of thyroid cells are cultured in a collagen gel, the follicular structure is reconstituted (Chambard et al., 1981; Kitajima et al., 1987). On the basis of data from Kitajima et ul. (1987), the process of follicle formation has been determined (Figs. 19 and 20). When isolated porcine thyroid cells are incubated for 1 week on collagen gel or culture dish, monolayered epithelium is formed. All of the cells maintain normal polarity; the surface facing the type 1 collagen gel or culture dish is basal and the opposite side (free surface) is apical. When monolayered epithelium on the type 1 collagen

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FIG. 19. Light micrographs of cultured thyroids. Thyroid cell monolayer formed on a collagen gel (A), thyroid cells grown in a collagen sandwich for 1 day showing proliferation and migration of cells to form two layers (B), and follicle-like cavities after 3 weeks of culture in a collagen sandwich containing TSH, insulin, and dexarnethasone (C). (A) ~ 5 6 0 ; (B) x430; (C) ~ 4 8 0 From . Kitajimd et a / . (1987).

gel is overlaid with a second layer of type 1 collagen, changes occur in the polarity and arrangement of the cells. Within 1 day of culture in the collagen sandwich, some cells in the monolayer proliferate, migrate, and form two cell layers. Cell surfaces facing the upper and lower collagen gels always show basal characteristics. The primitive follicular lumen sealed by zonula occludens appears between two cells (Fig. 20). Microvilli protrude into the lumen. Within 2 days of culture in the collagen sandwich, thyroid cells form two cell layers with large follicle-like cavities between them, even in the absence of TSH. These cavities are developed by proliferation of cells surrounding the lumen and luminal fusion. Numerous actin filaments are located along the apical and basal plasma membrane whereas microtubules are well developed in the cytoplasm. Colloid-like substances stored in the cavity are positive for both periodic acid-Schiff (PAS) and immunofluorescent staining of thyroglobulin, and laminin is imrnunohistochemically demonstrated at the outer surface facing the collagen gel. The formation of the follicle-like cavity is inhibited by colchicine. Pic et al. (1984) demonstrated that anticytoskeletal agents, such as cytochalasin B and colchicine, alsoi nhibit folliculogenesis in organ-cultured fetal rat thyroid. These facts suggest the following: surface facing type 1 collagen usually obtains basal characteristics, microtubules are important for organization of cell polarity and formation of a

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FIG. 20. Electron micrographs of monolayered thyroid cells cultured for I day in a collagen sandwich (A and B). Some areas of cultured cells consist of two cell layers, whereas others remain in a monolayer showing a mitotic cell (A). Appearance of a primitive follicular lumen sealed by a zonula occludens between upper and lower cell layers (B). (A) ~ 2 8 0 0 (B) ; x6600. From Kitajima ef al. (1987).

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follicle-like cavity, and cultured cells secrete thyroglobulin into the cavity and laminin into the basal part of the cell. IX. Why Does the Thyroid Need Follicle Structures? In vertebrates, the thyroid is a morphologically unique organ consisting of many follicles. The follicular epithelial cell shows typical exocrine cell polarity and undergoes three functional steps in the process of hormone synthesis and secretion: (1) synthesis and secretion of protein in exocrine cell manner, (2) reabsorption and hydrolysis of protein, and (3) release of hormone from the basal part of the cell. This complicated process for synthesis and secretion of thyroid hormones T4 and T3 is also due to their unique chemical structure. They are iodinated diphenyl ethers, not peptides, which are not synthesized directly from their constituents, tyrosine and iodine, in the cell. lodination of tyrosine and the coupling of two molecules of iodotyrosine take place in the extracellular lumen, not in the cell cytoplasm. The follicular epithelial cell synthesizes high-molecular-weight glycoprotein (thyroglobulin) in the cytoplasm and releases it into the follicular lumen in typical exocrine cell secretory manner. In thyroglobulin molecules, tyrosyl residues are iodinated and coupled in the follicular lumen of most vertebrates and in the endostylic lumen of larval lampreys and protochordated. Thyroglobulin stored in the follicular lumen in reabsorbed into the cell by stimulation of TSH. Hence, the follicular lumen is believed to be the site for iodination and coupling of tyrosyl residues in thyroglobulin and for storage of thyroglobdin. Thyroglobulin reabsorbed from this lumen is hydrolyzed to liberate T4and T3. This hydrolytic activity is regarded as a specialized process for making thyroid hormones (T4 and T3), and thyroglobulin is regarded as a specialized prohormone in this process. T4 and T3 amino acid derivatives are specialized ethers and, because they are not directly synthesized by the RER and Golgi apparatus system, do not seem to be stored in granular form in the cytoplasm. When TSH is injected, the follicular epithelial cell takes up a large quantity of thyroglobulin by phagocytosis and pinocytosis, and many large colloid droplets, some of which are fused with lysosomes, appear in the cytoplasm. The liberated hormones are believed to be released quickly from the basal part of the cell. This means that luminal colloid (prohormone) stored in the common lake (follicular lumen) is mobilized by TSH stimulation and that hormones are not stored in the cell. In this sense, the luminal colloid is regarded as an extremely large,

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specialized prohormone granule commonly surrounding follicular epithelial cells.

X. Concluding Remarks

The secretory activity of the thyroid follicular epithelial cell, its relationships to vascularizations and nerve supplies, and some problems concerning its cell polarity and inverted structure were reviewed in structurally, histochemically, and cytochemically. The follicular epithelial cell morphologically shows typical exocrine gland cell polarity, synthesizes high-molecular-weight glycoprotein (thyroglobulin) by the RER and Golgi apparatus system, and releases secretory granules containing thyroglobulin into the follicular lumen in exocrine cell manner. Iodination and subsequent coupling of tyrosine residues in thyroglobulin primarily take place in the follicular lumen, especially at the periluminal region. Luminal thyroglobulin is usually reabsorbed into the cell by pinocytosis and in stimulated conditions, by phagocytosis as well as pinocytosis. Actin and caldesmon localized at the apical cytoplasm are implicated in reabsorption of the colloid (thyroglobulin). Intramembranous protein particles accumulate at the initial site of pinocytosis in freeze replica images. Reabsorbed colloid droplets and vesicles are fused to become larger droplets that, in turn, are fused with primary lysosornes to be hydrolized and liberate thyroxine (T4) and triiodothyronine (T4. T4 and T3 are released from the basal part of the cell into the connective tissue space probably by a diacrinous mode. Each follicle structure is encapsulated by a well-defined, basket-like capillary network that is generally independent of adjacent networks, although a few anastomoses or common capillaries are sometimes seen between them in monkeys and rats. In each follicle, gap junctions are well developed between the follicular epithelial cells. This suggests that each follicle functions almost independently of other follicles, and whole cells in each follicle function synchronously. Blood capillaries become markedly dilated and fused in the overstimulated state of the gland, whereas they become markedly small in diameter and poor in distribution and anastomosis in the hypofunctional state. The number of capillary endothelial fenestrations is increased in the hyperfunctional state of the gland and reduced in the hypofunctional state. These facts suggest that the blood capillary is flexible and changeable in both morphology and distribution, reflecting the functional state of the gland's secretory activity . The thyroid is innervated by three kinds of nerve fibers, sympathetic,

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cholinergic, and peptidergic, although few fibers actually enter the organ. The main route for stimulating the follicular epithelial cell to secrete T4 and T3 is humoral, through the TRH-TSH system, rather than neuronal. The polarities of follicular epithelial cells in the isolated follicle are reversed by some experimental conditions, which were reviewed. Using the inverted follicle, it is easy to show that the thyroid cell takes up foreign materials, such as latex beads and erythrocytes, in addition to thyroglobulin. Finally, the reasons why the thyroid needs follicle structures were considered and discussed. REFERENCES Abe, S., and Ogawa, K. (1980). Biomed. Res. 1, 47-58. Ahren, B., Hakanson, R., and Rerup, C. (1982). Acta Physiol. Scand. 114, 471-473. Alquier, C., Fayet, G., Hovsepian, S., and Michel-Bechet, M. (1979). Cell Tissue Res. 200, 69-81. Bargmann, W. (1939). In “Mollendorffs Handbuch der Mikroskopischen Anatomie des Menschen” (W. von Mollendorff, ed.), Vol. V1, Part 2. Springer-Verlag, Berlin. Bayliss, W. M., and Starling, E. H. (1902). J. Physiol. (London) 28, 325-358. Bennett, G., Parsons, S., and Carlet, E. (1984). A m . J. Anat. 170, 521-530. Berger, E . G., and Hesford, F. J. (1985). Proc. Nail. Acad. Sci. U.S.A. 82, 4736-4739. Bjorkman, U., Ekholm, R., Ericson, L. E., and Ofverholm, T. (1976). Mol. Cell. Endocrinol. 5, 3-17. Bjorkman, U., Ekholm, R., and Denef, J.-F.(1981). J. Ulrrastruct. Res. 74, 105-115. Bouchilloux, S., Chabaud, O., Michel-Bechet, M., Ferrand, M., and Athouel-Haon, A. M. (1970). Eiochern. Biophys. Res. Commun. 40, 314-320. Branton, D., Cohen, C. M., and Tyler, J. (1981). Cell 24, 24-32. Calvert, R., and Pusterla, A. (1973). Gen. Comp. Endocrinol. 20, 584-597. Chambard, M., Gabrion, J., and Mauchamp, J. (1981). J. Cell B i d . 91, 157-166. Chambard, M., Varrier, B., Gabrion, J., and Mauchamp, J. (1984). B i d . Cell. 51, 315-326. Claude, P., and Goodenough, D. A. (1973). J. Cell Eiol. 58, 390-400. Daniel, P. M., Pratt, 0. E., Roitt, I. M., and Torrigianai, G. (1967). Immunolozy 12, 489-504. De Groot, L. J., and Niepomniszcze, H. (1977). Metabolism 26, 665-718. Denef, J. F., and Ekholm, R. (1980). J. Ultrasfruct. Res. 71, 203-221. Desalu, J., Fujiwara, T., and Komuro, T. (1984). Arch. Histo!. J p n . 47, 179-186. Doniach, I., and Pelc, S. R. (1949). Proc. R . Soc. Med. 42, 957-959. Dunn, A. D. (1974). J. Exp. Zool. 188, 103-124. Dunphy, W. G., and Rothman, J. E. (1985). Cell 42, 13-21. Edelhoch, H . , and Robbins, J. (1978). In “The Thyroid” (S. C. Werner and S . H. Ingbar, eds.), pp. 62-76. Harper & Row, New York. Ekholm, R. (1966). J. Ultrastruct. Res. 14, 419-420. Ekholm, R. (1981). Mol. Cell. Endorinol. 24, 141-163. Ekholm, R., and Wollman, S. H. (1975). Endocrinology 97, 1432-1444. Engstrom, G., and Ericson, L. E. (1981). Endocrinology 108, 399-405. Ericson, L. E. (1983). J . Endocrinol. Inuest. 6, 311-324.

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INTERNATIONAL KEVIEW OF CYTOLOGY, VOL. 113

Bacterial Surface Polysaccharides: Structure and Function IAN W. SUTHERLAND Department of Microbiology, Edinburgh University, Edinburgh EH9 3JG, Scotland

I. What Are They? Introduction and Definition At the surface of the bacterial cell, as with other cell types, polysaccharide molecules are found. These polymers can take various physical forms; they may be associated with other surface components or they may be totally dissociated from the microbial cell. It is now clear that in some examples, the polysaccharides may bear a strong chemical similarity to components of the cell wall, but in others, they are distinct chemical entities in no way related to cellular components. Surface polysaccharides of one type or another are widely observed in prokaryotic isolates, including those that are pathogenic for humans, animals, and plants and also those that are free-living saprophytes. The definition of polysaccharides and related macromolecules found at the prokaryotic cell surface provides a wide variety of nomenclature. There is no confusion about polymeric components of the cell wall such as peptidoglycan, lipopolysaccharide, and teichoic or teichuronic acid. The term exopolysaccharide (EPS) has been widely used to cover polysaccharides found external to the surface of the microbial cell and has the advantage of including polymers of different physical types (and of very diverse chemical composition). This term will be used in the present review to embrace all polysaccharides considered. The term glycocalyx, introduced by Costerton and co-workers (1981), suffers from the disadvantage that it has been used to represent an even more complex array of molecules including both extracellular components and wall polysaccharides along with other chemical species observed in and closely associated with the cell wall. The term glycocalyx also fails to provide an adequate chemical definition of these polymers, as it may include a number of non-carbohydrate-containing materials. Surface polysaccharides may form structures recognizable per se by light or electron microscopy, or they may form components of more complex structures. These structures may, in turn, be involved in complex morphogenetic cycles, such as those observed in Azotobacteraceae or Myxobacteraceae, where EPS in the form of microcysts are 187

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associated with both normal vegetative bacteria and resting cells. The microcysts are found free in the former group, but in the latter are contained within exotic and complex fruiting bodies. The production of EPS may indeed be a feature of a specific phase of growth, or it may be a constant feature during bacterial growth and division. A number of bacteria are known to synthesize several different EPS, but it is not yet clear whether more than one polymer is secreted throughout the growth cycle. As yet, few attempts have been made under controlled physiological conditions to study the growth of bacteria yielding such mixtures. The presence of EPS associated with bacterial cells grown on solid surfaces is frequently recognizable from the mucoid colonial morphology. At times, this may be associated with unusual appearance (Fig. 1). It does not. however, indicate whether the EPS forms part of a capsule firmly attached to the bacterial cell surface or is in the form of loose slime secreted by the bacterium, but not directly attached to it. On solid surfaces within the human or animal body exposed to aqueous environments, either freshwater or ocean,bacterial growth is seen as biofilms in which bacterial cells are associated with large amounts of EPS. Unfortunately, many of the descriptive reports of EPS in the laboratory and in natural environments have failed to recognize the relationship between particular forms of these macromolecules and the physiological conditions present. Changes in physiological conditions can drastically alter the composition. physical properties, and organization of polysaccharides at the bacterial surface. Some techniques such as scanning electron microscopy (SEM) frequently provide strong evidence for the presence of EPS, but are not capable of providing confirmatory chemical evidence. Equally, modern techniques of transmission electron microscopy (TEM) can reveal considerable information on the surface structures involved, including polysaccharides, but are unable to distinguish between different chemotypes such as homopolysaccharides and heteropolysucchurides. They do not have sufficient definition to provide much information on the microstructure of the bacterial capsule. However, improved methods can now be used to study the structure of pure polysaccharides when prepared in aqueous solution, and this may provide supporting evidence for their conformation in solution. The aim of this review is to attempt to correlate the chemical and physical information we now possess on bacterial surface polysaccharides and determine to what extent it matches and amplifies the cytological observations of these polymeric structures. It makes no attemp to be comprehensive, but rather reflects specific groups of extracellular polysaccharides that have provided much of the current

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FIG. I . Colonial morphology of a bacterial isolate from the interior of a fungal sporocarp (see Chard et a / . , 1987).

information. There are a number of reviews providing extensive bibliographies of the chemical structures (e.g., Lindberg et af., 1975), therefore, structures will be illustrated only when they are strictly relevant to the material under discussion.

II. Appearance-Light Microscopy and Transmission and Scanning Electron Microscopy

A. POLYSACCHARIDE FIBERS The direct examination of polysaccharides and the observation of fiber-like structures can be accomplished in either of two ways. The structures may be seen during electron microscopic examination of

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bacterial cultures. Alternatively, bacterial polysaccharides may be isolated from cultures, purified either partially or extensively, and then examined by electron microscopy. Because of their relatively amorphous structure, EPS strands are not normally visualized as :hey are formed. An exception to this is seen in cellulose production by Acetobacter xyfinum. A ribbon of cellulose formed from a number of microfibrils has been observed (Zaar, 1979). Beneath the polysaccharide strands, a linear array of pores having apparent diameters of 12-15 nm could be seen on the outer membrane of the bacteria. It was suggested that such pores might represent export sites for bacterial cellulose. The results were in good agreement with an earlier study, indicating production of a ribbon of microfibrils elongating at a rate of 2 pmlminute (Brown et al., 1976). These observations also indicated that approximately 50 synthetic sites were organized in a row along the longitudinal axis of the bacterial cell. This row of sites was apparently duplicated prior to division, each daughter cell receiving a comparable number of sites. Each ribbon of cellulose was composed of about 46 microfibrils, perhaps resulting from the products of the pores coming together. This synthetic mechanism is under the control of a complex regulatory mechanism in which the synthase is affected by an unusual nucleotide activator (Ross et af., 1987). The acthator has recently been characterized as bis(3’ + 5’) cyclic diguanylic acid. The synthesis of cellulose or cellulose-like EPS has been attributed to various other bacteria as well as Acetobacter xylinum. These include strains from activated sludge and other floc-forming types, the presence of cellulose apparently conferring on them their flocculent growth characteristics (Deinema and Zevenhuizen, 1971). This has mainly been determined on the basis of dissolution of flocs or fibrils with cellulase; as enzyme preparations are liable to contain various contaminating endo-pglucanase activities, such conclusions must be regarded with caution. By utilizing the ability of 1,4-@-glucans to interact with Congo red, Zevenhuizen et al. (1986) have provided stronger evidence for the synthesis of cellulose in the form of microfibrils by Rhizohiurn trifolii mutants (see, e.g., Wood, 1980). The mutant bacteria strongly absorbed the dye and, in addition, grew as flocs during the exponential phase, whereas electron microscopy revealed extracellular fibrils (Fig. 2). Dye uptake of this type was confined to a small number of mutant strains, but was also found in Rhizobium legurninosarurn and R . phaseoli, again being limited to only a few strains among those tested. Electron microscopic observation of the ultrastructure of curdlan, either after purification or directly from Alcaligenes faecafis var. rnyxogenes (Koreeda ef al., 1974), revealed long microfibrils 10-20 nm wide.

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FIG. 2. Cellulosic fibrils associated with Rhizobiurn trifolii cells. (Reproduced from Zevenhuizen et a / . , 1986, with permission.)

The microfibrils appeared to be formed from clusters of single fibrils. Partial degradation, reducing the degree of polymerization (DP) from 400 to 260, resulted in shorter fibrils. It was suggested that maintenance of an ordered helical conformation (see Section V> was necessary for microfibril formation and that disordered portions inhibited the formation of longer microfibrils. A recent electron microscopic study of xanthan from Xanthornonas campestris showed the effect of polysaccharide contour length on conformation (Stokke et al., 1986, 1987). The appearance was rod-like at short lengths, but was in the form of a random coil at longer lengths (>1 pm). In the presence of salt, a double-stranded conformation was apparent with chain splitting or separation seen at lower salt concentrations. Strain differences were also observed; the polysaccharide from one strain with higher acetate and low pyruvate was predominantly single stranded under conditions where EPS from a normal preparation revealed a mix of double- and single-stranded conformation.

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IAN W. SUTHERLAND

B.

CAPSULES A N D

RELATEDS T R U C r U R E S

Recognition of a discrete layer of EPS surrounding many bacterial cells as a capsule has been achieved initially through light microscopy and then through steadily improving electron microscopic techniques. Because the capsule is a highly hydrated structure, many early electron microscopists encountered the problem of its structural collapse with resultant loss of definition and integrity. A milestone in electron microscopic studies of EPS capsules was the paper by Bayer and Thurow (1977) on Escherichia coli K29 capsule. After pretreatment with a relatively high concentration of anticapsular IgG and freeze etching, the EPS was seen in an uncollapsed state. Under the physiological conditions used, this strain produced a capsule that appeared as a 250- to 300-nm-thick homogeneous layer. Ultrathin sections suggested a fibrous composition, and alkali treatment of the capsule yielded fine filaments 250-350 nm long and 3-6.6 nm wide. A knob-like structure was seen at one end of many fibers, perhaps indicative of the mode by which they had been attached to the cell surface. Examination of thin sections of gelatin-enrobed bacteria showed that the fibrous strands extended radially through the capsule and were embedded in a fine granular matrix (Bayer et al., 1985), as can be seen in Fig. 3. There was apparent association between the fibers, perhaps representing the ordered form of EPS solutions discussed below, but capsules of adjacent cells remained distinct and did not coalesce. Other bacteria, with different EPS composition and physical properties, may well behave differently from E. coli K29. Indeed, Bayer et af. (1986) found that even in strain K29, lipopolysaccharide (LPS) extended 40-60 nm into the capsulsar matrix and is clearly nonuniform, perhaps trapping macromolecules such as LPS if they slough off the bacterial cell surface. Excretion of EPS is not necessarily uniform. In a species that forms a capsule surrounding the bacterial cell, such as E . cufi, Bayer and Thurow (1977) noted that the sites of new polysaccharide emergence were distributed randomly over the bacterial surface. However, in Asticacaulis biprosthecum, acid polysaccharide, as recognized by ruthenium red staining, was found in the center of rosettes, but not in other large aggregates of these prosthecate bacteria (Umbreit and Pate, 1978). Between these extremes are the slow-growing Rhizobium species, such as Rhizobium japonicum, in which polarity of capsulation was recognized by Tsien and Schmidt (1977). Vasse et a / . (1984) observed that capsulation FIG. 3. Esclierichia coli K29 capsulate cell. The bacteria were fixed with glutaraldehyde, enrobed with gelatin. and embedded. The capsular material can be seen forming a layer 600 nrn thick around the bacterium. The gelatin is recognizable by its coarse grain. (Reproduced from Bayer et 01.. 1985, with permission.)

BACTERIAL SURFACE POLYSACCHARIDES

193

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IAN W. SUTHERLAND

developed only at one pole of the cell in early exponential phase growth (Fig. 4). Later, there was extension and some loss of this material from the surface into the culture medium, where polysaccharide was also found as loose fibrils. Care had to be taken that the capsular material was not lost during centrifugation for electron microscopy. It is thus possible that nonuniform capsulation, as seen in Rhizobium juponicum, might be of wider occurrence than has hitherto been recognized.

111. Physiological Influences

A. CARBONSUBSTRATE Most studies on the effect of varying physiological conditions have been made with gram-negative bacteria. It is clear, however, that

FIG. 4. Asymmetric capsule synthesis in Rhiiobzum juponicurn visualized with ferritin. In the early exponential phase (A), the material is associated with one pole of the cell, but by

BACTERIAL SURFACE POLYSACCHARIDES

195

physiological conditions can have a profound effect on polysaccharide production in all types of bacteria. In a study of Lactobacillus casei subsp. rhamnosus, Wickenbet al. (1983) observed that whereas the cell wall poly saccharide remained constant under different culture conditions, capsular polysaccharide production in batch culture was dependent on the carbon substrate. In continuous culture, capsule synthesis was affected by the dilution rate and by the nature of the limiting carbon substrate. There was no indication of changes in the composition of the capsular material. Although alginates are now known to be synthesized by a number of plant-pathogenic Pseudomonas spp., production is dependent on the availability of glucose or gluconate as carbon substrate (Fett et ul., 1986). When sucrose was used as carbon substrate, some strains produced only alginate, but others yielded a mixture of levans and alginate. A similar finding was made for Erwinia amylouora, the causative bacterium of plant

the mid-exponential phase (B), it shows wider distribution. (Reproduced from Vasse et a / . , 1984, with permission.)

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IAN W . SUTHERLAND

fireblight diseases. Two different EPS were produced in uitro, a heteropolysaccharide containing glucose, galactose, mannose, and uronic acid, and a homopolymer, a fructan (Bennet and Billing, 1980). There was a close similarity in the chemical composition of the acid heteropolysaccharide produced on glucose or sorbitol medium and that isolated from infected plants, this being true of a single E . umylouora strain in different plant hosts or of different bacterial strains in the same plant host. B. NUTRIENT LIMITATION

The effect of nutrient limitation on the capsular polysaccharide morphology of Azotobacter chroococcum has recently been studied by Ferrala et ul. (1986). The composition and structure of this polymer have not been determined, but studies from other strains of this bacterium indicate that it is probably an acid heteropolysaccharide. Growth of these nitrogen-fixing bacteria under iron or molybdenum limitation caused changes in the appearance of ruthenium red-stained capsular material examined by electron microscopy. Care was taken to avoid changes that might otherwise be due to the encystment process. The bacteria grown under nonlimiting conditions revealed electron-dense capsular material closely associated with the outermost layers of the walllmembrane complex. Under iron limitation, the capsule was much more extensive and diffuse. The effect of molybdenum limitation was much less marked, whereas simultaneous limitation of both ions revealed little difference from samples obtained under iron limitation. By contrast, cells allowed to encyst differed little when grown under metal ion sufficiency or deficiency. It can, perhaps, be considered that in a nitrogen-fixing species such as A . chroococcum, limitation of either iron or molybdenum is solely an extreme form of nitrogen limitation. If, as is likely, the EPS comprises an acidic macromolecule, it would probably bind metal ions that could even assist in conferring structures seen in normal cells. However, addition of iron to iron-limited cultures did not restore the original structure, indicating that the polymer synthesized under iron limitation was probably altered in one or more of the three variable parameters described on pages 197-202. It is now clear that the chemical composition and physical properties of EPS synthesized in batch cultures depend on the physiological conditions present at the time of synthesis. Unfortunately, many studies have used batch culture material that represents the total synthesis of the bacterium, i.e., a pool of macromolecular types that probably varies considerably in both carbohydrate composition and acylation. There are wide variations in the extent of polysaccharide synthesis in different bacteria, some commencing EPS production during late exponential phase and mostly

BACTERIAL SURFACE POLYSACCHARIDES

I97

yielding actual polymer in the stationary phase, Others, such as alginate in A . uinelandii, are synthesized throughout both exponential and stationary phases (Horan et al., 1981). Although early studies on the effects of nutrient limitation, on which many subsequent studies were based, were performed using batch culture, a considerable amount of information has been obtained in recent years through chemostat experiments. Thus, Davidson (1978) found relatively slight differences in glucose :mannose ratios for xanthan produced under various nutrient limitations, but found reduced mannose, glucuronic acid, and pyruvate under magnesium limitation. This would indicate apparent deficiency of many side chains of molecules synthesized under such conditions. Phosphate limitation had similar effects. Suprisingly, limitation of the carbon substrate did not cause any marked difference in the composition of the polysaccharide. Tait etal. (1986) found that under sulfate limitation, the carbohydrate composition of xanthan remained relatively constant over a range of dilution rates, but the acyl content tended to fall with increased dilution rate, as did the consistency index. In parallel studies performed in batch culture, the acyl content was seen to reach a maximum immediately subsequent to termination of logarithmic growth. Large differences were observed in the maximum consistency indices obtained during and after logarithmic growth, probably due to lower relative molecular mass of the polysaccharides as the cells grew rapidly. Evans et al. (1979) also found that the viscosity and chemical composition of the xanthan from X . campestris pathovar juglandis depended on the physiological conditions used. In contrast to the results for xanthan, EPS production by Rhizobium trifolii in carbon-limited cultures was strongly dependent on the dilution rate (De Hollander et al., 1979). Almost certainly, each species has to be regarded as a unique system, and considerable experimentation is needed before the different parameters affecting EPS synthesis can be determined. In general, good polysaccharide synthesis is permitted in nutrient limitations where nitrogen, sulfur, or phosphate provide the limiting nutrient, and a utilizable carbon source is in excess. However, it is now clear that in a number of different bacteria, even under carbon limitation, appreciable amounts of polysaccharide can be formed. IV. Chemical Structures

A. COMPOSITION The components of EPS comprise a wide range of sugars together with a limited number of acyl groups and, occasionally, inorganic residues such as phosphate. In general terms, the polymers can be divided into

198

IAN W. SUTHERLAND

homopolysaccharides and heteropolysaccharides, the former being mainly glucans, fructans, or polymers of N-acetylamino sugars (e.g., sialic acid and Vi antigen of Salmonella spp.). The major components of heteropolysaccharides are usually hexoses of the D series: the 6deoxyhexoses, L-rhamnose and L-fucose; N-acetylamino sugars ; and D-hexuronic acids. most commonly D-glucuronic acid. However, with improved analytical procedures and examination of a much wider range of prokaryotic species, the range of monosaccharides identified continues to increase. Some sugars previously thought to exist solely in LPS have now been found in EPS. The E. coli K14 antigen contains repeating units 1 + 5)-0-(3-deoxy-pof [ O-(2-acetamido-2-deoxy-~-~-galactopyranosyl)-( D-mannooctulopyranosyluronic acid)(2+6) 1, an example of a number of E . cofi EPS containing 3-deoxy-~-mannooctu~osonic acid (KDO) ( Jann et al., 1983). Indeed, these authors showed that this serotype was one of a group of E . coli EPS that owed their negative charge to components such as KDO, phosphate, or N-acetylneuraminic acid instead of containing hexuronic acids. A further four E. coli serotypes, K6, K13,K20,and K23, all contained equimolar amounts of ribose and KDO (Vann et al., 1981, 1983; Jennings er al., 1982). Although ribose has frequently been found associated with EPS, almost certainly because of contamination with RNA, it is now seen as a genuine component of certain EPS structures. Phosphate has long been known to be part of the EPS structure from various Streptococcus pneumoniae strains that excrete polymers bearing structural similarities to teichoic acids. It is also present in E . cofi K51 EPS a5 an 0-acetylated poly(a- 1,3-N-acetylglucosamine phosphate) (Jann and Jann, 1985) and in the teichoic acid-like K2 EPS, which contains D-galactose and D-glycerol phosphate (Jann et al., 1980; Fischer et al., 1982). A further phosphate-containing polymer is the E . coli K52 capsular polysaccharide, which has a D-galactosyl phosphate backbone to which are attached D-fructosyl residues (Fig. 5). Also of interest in this structure are 0-acetyl substituents on the C2 position of the galactose residues, whereas 10% of the fructose molecules carry both 0-acetyl and 0-propionyl groups on the C1 position (Hofmann et al., 1985). Although 0-acetyl groups and pyruvate ketals have been recognized acyl substituents of EPS for many years, it is now clear that several other such substituents may also be present. 0-Propionyl esters are rare and, as yet, have been identified only in E. coli K52 material and in E. cofi K14 (Jann et al., 1983), where they are located on D-fructose residues and KDO. respectively. The EPS of Rhizobium trifolii as well as similar polymers from Agrobacteriurn spp. have been known for some time to be unusual in that they carry succinyl groups in addition to pyruvate and acetate (Hisamatsu et al., 1978; Amemura and Harada, 1983). The

199

BACTERIAL SURFACE POLYSACCHARIDES

OH

FIG. 5. The structure of Escherichia coli K52 EPS. R1, Acetyl; R2, acetyl and propionyl.

succinyl groups were ester linked, the content varying from 0.4-7.4%. A further acyl substituent , 3-hydroxybutanoic acid, was identified in the polysaccharide from R . trifolii strain 0403 (Hollingsworth et al., 1984). This was later identified in polysaccharides of some strains of both R . trifolii and R . leguminosarum. In both of these EPS, the hydroxybutanoate was esterified to the C3 position of a terminal D-galactose residue, which also carried a 4,6-pyruvate ketal (Kuo and Mort, 1986). R . leguminosarum E P S also carried two 0-acetyl groups on a D-glucosyl branch point (Fig. 6). There are probably several EPS in which multiple acylation of sugars can be found, including at least one from an X. campestris strain ( I . W . Sutherland, unpublished results) and some bacterial alginates. Acetyl groups in bacterial alginate are found exclusively associated with Dmannuronosyl residues (Davidson et al., 1977; Skj2k-Braek et al., 1986). In a study of alginates from Pseudomonas aeruginosa, Sherbrock-Cox et

-4)-/3-o-GlcpA-(l-4)-~-~-GlcpA-(l-4)-/3-o-Glcp-(l-4)-~-D-(

CD

3

-

. 2

t 0t

0

I 1. .

R

R

FIG. 6. The multiply acylated EPS structure of Rliizobium leguminosarum.

200

IAN W . SUTHERLAND

al. ( 1984) obtained O-acetate : mannuronic acid ratios ranging from 0.76I .46, indicating that some strains contained multiply acetylated Dmannuronic acid. This was confirmed in a subsequent study by SkjgkBraek et al. (1986), who reported that the acetyl content of P . ueruginosa alginates ranged from 37-57%, whereas that of a P. mendocinu EPS was 30%. In contrast, preparations from P . putidu and from P . , f u o r e s c ~ n s contained only 3 - 4 s acetate. In alginates from both Azotobacrer uinelandii and P . aerrrginosu, the O-acetyl groups were carried on the C2 and C3 positions of the D-mannuronic acid, 3-11% of these residues carrying acetylation on both carbon atoms (Table I). Glycerol is present in a number of teichoic acid-like EPS, mainly from gram-positive bacteria such as Streptococcus pneumoniae. In these polymers. it is linked through phosphate groups to carbohydrate residues. It has now also been detected as an unusual acyl substituent of "gellan," the polysaccharide produced by an Alculigenes sp. (Kuo et al., 1986). Another recent addition to the list of noncarbohydrate components of EPS is serine, which is located in E. coli K40 capsular polysaccharide (Dengler et ul., 1986). O-Methyl sugars are present in a number of EPS, but the largest source of such sugars so far identified has been the genus Rhizobium. The EPS from Rhizobium japonicum contains 5- 13% 4-O-methylgalactose (Dudman, 1976), whereas .t-O-methyI-~-glucoseand 3-O-rnethyl-~-ribosehave also been detected in EPS from different Rhizobium species (Kennedy, 1976; Kennedy and Bailey, 1976). In one polymer, two O-methyl sugars, 3-O-methyl-~-glucoseand 6-0-methyI-~-galactose, were present together with 6-deoxy-~-taloseand other sugars (Kennedy, 1978). 4-O-Methyl-~glucuronic acid has also been found as a side-chain terminus in EPS from R . juponicuin strains (Dudman. 1978), whereas a further rare sugar, D-riburonic acid, occupied a similar role in the more complex repeating unit of a R . meliroti EPS (Amemura et al., 1981). It has been reported that the degree of O-methylation of 3-O-methyl-~-rhamnosein the EPS from one Rhizobium strain varied with the composition of the growth medium (Jackson et al., 1982), but it is not clear whether this is a general feature of O-methylation. The effect of the acyl groups, as with O-methyl groups, is to increase the hydrophobicity of parts of the molecule, and this presumably affects the structure of polysaccharides in solution. The acylation also has marked effects on both inter- and intrachain associations and, thus, greatly alters the physical conformation of polymers in solution together with their resultant properties. In some EPS, the acylation varies, but in others, it appears to be totally unaffected by culture conditions. The ester-linked groups, such as O-acetyl, are highly labile to alkaline

TABLE 1 BACTERIAL ALGINATE COMPOSITION^ Percentage of total alginate composition Strain

Guluronic acid :

rnannuronic acid ratio

Acetyl

C2 Acetyl

C3 Acetyl

2,3 Diacetyl

0.67 0.05 0.45 0.94 0 0.26 0.4 0.37

0.33 0.95 0.55 0.06 1 .o 0.74 0.6 0.63

21 52 22 4 37 30 3 4

13 30

8 22

3 11

23

14

5

Azotobacter vinelandii I A . vinelandii I1 A . vinelandii I11 A . vinelandii IV Pseudomonas aeruginosa I P . mendocina 10541 P . fluorescens 10255 P . putida 1007 Results from Skjak-Braek et al. (1986).

202

IAN W. SUTHERLAND

conditions, and it is possible that some preparations may be partially degraded during isolation if appropriate precautions are not taken.

B. STRUCTURE

I , General Findings The structures of EPS so far studied vary considerably. Heteropolysaccharides, with the exception of bacterial alginates, are composed of repeat units varying upward in size from two monosaccharides. A few bacterial genera or species have been exhaustively studied with respect to their EPS structures, but for many isolates, little is known beyond the monomeric composition and, occasionally, analysis of component sugars and acyl groups. Provided that pure material is studied, it usually reveals stoichiometric ratios for the carbohydrate and noncarbohydrate components. In the case of the latter, a few examples are now known in which substituents such as acetate or pyruvate are only found on half of the repeat units, usually on alternate repeat groups. However, it is possible, especially with some of the more complex polysaccharide structures, that replication of the repeat units may not be totally accurate. Consequently, some polysaccharides may be nonuniform in the addition of side chains to a main chain of constant composition. An example of this is xanthan from Xanthomonas carnpestris in which degradation of unordered polymer solutions using endo-P-glucanases yielded an excess of glucose thought to be derived from portions of the macromolecule lacking side chains (Sutherland, 1984a). One of the most thorough series of investigations has been that on Klebsiellu (Enterobncter) aerogenes. Systematic examination of the EPS from about 80 serotypes of these bacteria revealed a limited range of structural types composed of simple sugars and a few acyl substituents. In general terms, the repeat units have, for the most part, been characterized a5 charged structures of three to six sugar residues, one of which is an uronic acid-most commonly mglucuronic acid-and two to three neutral monosaccharides together with ester-linked acetate and/or pyruvate ketals. This is the most uniform series of polysaccharide structures so far detected and is in marked contrast to the EPS from Escherichia coli or Streptococcus pneurnoniae, which vary widely in both composition and structure. All the Klebsiella EPS are anionic polysaccharides. Some are linear polymers without side chains, exemplified by K1 and K5, each of which are formed from trisaccharide repeat units (Dutton and Yang, 1973; Erbing et al., 1976). Both of these polysaccharides carry pyruvate ketals attached to D-glucuronic acid and D-mannose, respectively, although K5

BACTERIAL SURFACE POLYSACCHARIDES

203

is also acetylated. Other serotypes from Kfebsielfaare represented by repeat units with side chains of various sizes, frequently a single sugar, but sometimes a trisaccharide as in K18 (Dutton et al., 1978) and K41 (Joseleau et al., 1978). In a few cases, two side chains are present. These may be attached to the same main-chain sugar residue, as is seen in K30, K33, and K64 (Lindberg et al., 1979a,b; Ravenscroft et al., 1985). Another unusual structure is found in K60 in which there is a heptasaccharide repeat unit of which three sugars form single glucosyl side chains (Dutton and DiFabio, 1980). Whereas the great majority of Kfebsiella polysaccharides contain an uronic acid, usually D-glucuronic acid, in a few, the only acid component is a pyruvate ketal, as in K32 where the acetal group is exceptionally attached to an L-rhamnosyl residue (Bebault et al., 1978). 2 . Polysaccharides with Common or Closely Related Structures Inevitably, considering the limited number of potential component sugars, close similarities may be found between some EPS from different genera. Another characteristic of EPS is that considerable portions of the carbohydrate repeat unit structure are shared. Thus, extensive serological cross-reaction between different EPS is a relatively common occurrence. This conservation of structure may be found in closely related species such as Rhizobium, many of which yield EPS with an octasaccharide structure, or in more disparate groups. For example, many of the recently developed “commercial” polysaccharides, from a range of gram-negative bacteria including Alcaligenes and Pseudomonas strains, have structural features in common. They reveal a remarkably conserved main-chain structure. To this are attached various side chains and substituents (Fig. 6). As already indicated, this group of polymers has been investigated because of their potentially exploitable rheological properties. Gellan from Pseudomonas elodea has a straight-chain tetrasaccharide repeat unit but, in addition to the recently discovered glyceryl esters, is also O-acetylated on approximately 25% of the tetrasaccharides (Fig. 7) (Jansson et a / . , 1983). The Alcaligenes S130 and ,588 polymers, although sharing the same backbone structure as gellan, are branched with either cw-L-mannosyl or a-L-rhamnosyl residues, the former not having been previously described in other bacterial EPS (Jansson et al., 1985, 1986a,b; O’Neill et al., 1986b). S88 polysaccharide has a disaccharide (gentibiosyl) side chain attached to the corresponding main-chain glucose residue, which in gellan is acylated. An unusual feature of the polysaccharide is that one of the main-chain sugar residues can apparently be either L-rhamnose or L-mannose. It is not yet clear whether this repre-

204

IAN W . SUTHERLAND L

- _ _ -_3

p

- Glyc

4 D-Glc(1-4)

p

D-GlcA(1-4)

p

D-Glc(1-4)

q - L - R h o ( 1 .....-..

P 0 - Ac ........ - 3

p

p

G-GIc(1-4)

GELLAN

p

D-GlcA(1-4)

f3 D - G l c ( 1 - 4 )

q - L - R h a ( l ........

Itb G - G t c (1-6)

........ - 3

S I94

q-D-Glc

p

G - G l c (1-4)

p

D-GkA(1-4)

p

D - G k (1-4)

,I' q L - Rho

q -L-Rha(l.....__. or Man

588

F ~ G 7. . The structure of "gellan" from Pseudomonos elodea, and related polymers.

sents a random sequence of the sugars, a regulated and regular sequence, or a mixture of two polymeric types, one containing L-rhamnose and the other L-mannose. This last possibility is probably unlikely. Another polysaccharide belonging to the gellan group, S198 from an Afcaligenes sp., resembles S88 in the variability of its main-chain monosaccharides. Again, the main-chain sequence of the repeat unit terminates in either L-rhamnose or L-mannose (Chowdhury et al., 1987b). A further unusual feature of this polymer is that it carries an a-L-rhamnosyl side chain on about 50% of the main-chain D-glucosyl residues distal to the uronic acid. The polymer is also acylated. A Xanthomonas polysaccharide (S657) has a main chain identical to the gellan structure, but carries an a-linked rhamnosyl disaccharide on the glucose situated between the uronic acid and main-chain L-rhamnose (Chowdhury et a f . , 1987a). It is of particular interest that this family of polysaccharides, with potentially interesting physical characteristics and highly conserved chemical structure, should extend over a range of gram-negative bacterial species. All pathovars of X . campestris that have been thoroughly studied appear to produce a polysaccharide with essentially the same carbohydrate structure, a repeat pentasaccharide unit (Jansson et a / . , 1975; Melton ef al., 1976). Variations are found in the acyl groups, depending on both the strain used and the physiological conditions employed for growth (Sutherland, 1981). The other members of the genus Xanthornonas also produce EPS, although generally in lower yields than X . campestris, but their EPS contains additional sugars, such as D-galactose, and the polymers are not degraded by xanthanases. A number of other reports are mainly from strains of diverse origin of which the species' authenticity has not necessarily been confirmed. These preparations

BACTERIAL SURFACE POLYSACCHARIDES

205

possessed both components and partial structures common to xanthan, but whereas one also contained galactose (Fareed and Percival, 1976), the other contained KDO (Yadomae et al., 1978). In neither case was a complete structure reported. An Acetobacter species (strain AM2) has been studied by Tayama et al. (19851, who reported that it produced an EPS with remarkable similarity to xanthan. It contained the same p-1,4-glucan main chain, whereas alternate glucose residues carried a pentasaccharide side chain. This portion had the same two sugars adjacent to the glucose and in the same linkages as xanthan. However, instead of a terminal mannose residue, a trisaccharide of L-rhamnosyl- 1,6-p-~-glucosyl1,6-~-glucosewas attached to the uronic acid. 0-acetyl groups were present but no pyruvate. A second EPS from an Acetobacter species contained many of the structural features of AM2 polymer but was rather more complex, carrying two different side chains, one as in AM2 and the other a pentasaccharide composed of glucose and galactose (Tayama et al., 1986). Another report on an EPS from an Acetobacter species indicated that the same sugars were present and that many of the linkages were common to Japanese studies, but a rather different structure was deduced (Weng et al., 1984). The polymer structure may thus be a reflection of strain differences. On the basis of the structure reported by the Japanese group, it would be expected to be sufficiently similar to xanthan to be degradable by xanthanases, although the specificity of the enzymes has not been completely elucidated. However, all of these A . xylinum polysaccharides are resistant to degradation by xanthanases (Sutherland, unpublished results). There have also been reports of the simultaneous synthesis of cellulose and an EPS containint glucose, mannose, rhamnose, and glucuronic acid by A . xylinum (Savidge and Colvin, 1985). Thus, in reality, there may be a series of soluble polysaccharides with many common structural features produced by these bacteria. In the same way that Xanthomonas species and strains yield very similar EPS, the polysaccharides from Rhizobium spp. show close similarities. Indeed, McNeil et al. (1986) showed that in all discernible features of the polymers from several strains of R . trifolii, R . leguminosarum, and R . phaseoli, no differences could be found. In this study, the generation of fragments through a phage-induced lyase cleaving the main chain produced octasaccharides from the EPS from different sources. These then could be readily characterized by modern analytical techniques. Using other strains of I?. trifolii and R . leguminosarum, Kuo et al. (1986) found that although the carbohydrate structures were identical, differences did exist in the acyl substituents. An aberrant polysaccharide from R . trifolii strain 4 s yielded a heptasaccharide rather than an

206

IAN W. SUTHERLAND

octasaccharide repeat unit (Amemura ef af., 1983). It shared the structural features of other EPS. but lacked the nonreducing galactosyl terminus on the side chain. This work also used a phage-induced enzyme, in this case an endoglucosidase. The heptasaccharide unit found in an R. trifofii strain studied by Jansson et d.(1979) possessed two adjacent ketalated sugars on the side-chain terminus, but some of the repeating units were incomplete and lacked the terminal pyruvylated galactose. The Rhizobium polymers also include one polymer from another strain of R . trifafii that possesses an octasaccharide repeat unit but, instead of the normal terminal sugar, contains D-riburonic acid in a furanosyl linkage (Amemura el u l . , 1981). The EPS from many of the Rhizobiurn isolates can, therefore, be considered as a series of very similar polymers, identical in many strains but revealing occasions1 “incomplete” or altered versions where either the side-chain terminus is missing or the galactose is replaced by another sugar. The presence of more than one uronic acid residue and/or pyruvate ketal provides a relatively high charge : mass ratio, whereas various ester-linked components can add to the hydrophobicit y . A second polymer is common to a number of bacteria including Rhizobiiirrz species, Alcaligvnrs jirecalis var. rnyxogenes, and Agrohucteriirm species. Because it contains succinyl groups, it has been termed “succinoglycan” and was first obtained from an A. faecalis strain capable of growth on a wide range of carbon sources. Various aspects of the polymer have been reviewed by Harada and Amemura (1981). Elucidation of the polysaccharide structure from various sources, through the use of specific degradative enzymes, revealed a polymer with D-ghcose, D-galactose, and pyruvate in the molar ratio 7 : 1 : 1, i.e., formed as is the Rhizobium polysaccharide from an octasaccharide (Ghai et d., 1981). The two types of octasaccharide show considerable similarities, particularly in their main-chain structures. The succinoglycan from Arthrohacter srahilis also had the same composition (Knutson et al., 1979). The succinogiycan is synthesized simultaneously with other EPS including curdlan; changes in relative yields were obtained through variation of culture conditions or through mutation (Hisamatsu el al., 1977). Alginates differ from other bacterial heteropolysaccharides in lacking a regular repeat unit. Instead, they again represent a conserved structure composed of irregular sequences of two uronic acids. Bacterial alginates are of much wider occurrence than was originally realized. They are found in AzotobarrcJr uinelundii as both capsule and slime produced by the vegetative cell, and as an essential component of the “exine” and “intine” layers of the microcyst resting stage (Eklund et al., 1966). This

207

BACTERIAL SURFACE POLYSACCHARIDES

material is similar to the product of marine algae in containing a range of different sequences of D-mannuronic acid and L-guluronic acid residue (Fig. 8), except that 0-acetyl groups are also present, associated with some of the mannuronic acid. The molar ratio of the two uronic acids varies considerably, reflecting the presence of the three types of regions within the molecule (Fig. 8). A number of Pseudomonas species are also known to produce alginates. A relatively small number of Pseudomonas aeruginosa strains, primarily isolates from cystic fibrosis patients, were found to synthesize an EPS similar in composition to alginate (Evans and Linker, 1973). Production of this type of EPS was extended to other Pseudomonas species-P. putida, P. Juorescens, and P. mendocina together with further P. aeruginosa strains-by selection with aminoglycoside antibiotics or with carbenicillin, following ethane methane sulfonate mutagenesis (Govan and Fyfe, 1978; Govan at al., 1981). A further source of bacterial alginates was discovered when O m a n et al. (1986) characterized the EPS from Pseudomonas syringae var. glycinaegrown in glucose-containing media as an acetylated polyuronide composed of D-mannuronic acid and L-guluronic acid. Other plant-pathogenic Pseudomonas species also yield similar polymers (Fett et al., 1986). Products from the plant pathogens differed from Pseudomonas aeruginosa alginate, being of lower molecular weight and higher polydispersity. This could, perhaps, be due to endogenous alginase production under the growth conditions used, as material produced in infected plants was of higher molecular weight. The Pseudomonas alginates show a further marked difference from both Alginate Structures Algal

- M - M - M - G - G - G - M - G - M - G -

A.vinelandii

-M-M

P. aeruginosa

- M\ - M - M\ - M\ - M\ - M - M\ - M - M

\ OAc

OAc

P. putida, P. fluorescens

-M -G -G -G -M-G

\ OAc

\ OAc

OAc

OAc

OAc

-M-G \ OAc

OAc

- M - M - M - M - G - M - G - M - G \

OAc

P. phaseolicola

- M - M - G - M\ - M - M - M - M - M OAc

FIG. 8. The sequence of uronic acid found in various types of “alginate.” M, Mannuronic acid; G , guluronic acid; OAc, acetyl ester. Esters on C2, C3, or both.

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IAN W. SUTHERLAND

Azotobucter uinelandii alginate and the product of marine algae in that they lack polyguluronic acid sequences (Skjlk-Braek et al., 1986) and possess only single residues of L-guluronic acid. This may be due to a postpolymerization reaction involving an epimerase with a different specificity from that found in A . uinelandii, which generates the eventual block copolymer by multiple attacks on the poly-D-mannuronic acid excreted from the bacterium (Larsen et al., 1986). It also affects interaction of the alginate with divalent cations (see Section V), as contiguous sequences of L-guluronic acid are necessary for this interaction. An analogous system to bacterial alginates, production of polymers of the same chemotype but different structure, is found in sialic acid. This is secreted as an EPS by various bacteria, and most work has been on Escherichia coli and Neisseria rneningitidis. Production of sialic acid is found in E. coli serotypes K1 (where it has been named colominic acid) and K92. The K1 material is structurally identical to sialic acid from N . meningitidis group B, being a (2 + 8)-a-linked homopolysaccharide (McGuire and Binkley, 1964). Group C meningococcal sialic acid is also a homopolysaccharide but is linked (2 -+ 9)-a (Bhattachajee et ul., 1975). An unusual feature of the meningococcal and E. coli K92 EPS has been the identification of hydrophobic termini, 1,2-diacylglycerol groups, on each polysaccharide (Gotschlich et al., 1981). The presence of these terminal groups cause the polysaccharide to aggregate in the form of micelles and might provide a mechanism for attaching capsular polymer to the bacterial surface, perhaps through linkage to a component of the outer membrane. It would seem unusual if such an attachment mechanism is confined only to these two bacterial species but, as yet, reports of similar terminal groups from other capsular-derived polysaccharides obtained from different bacterial genera or species are lacking. Another possible example of a conserved backbone structure with different side chains can be seen in the structures of E. coli K28 (Altman and Dutton, 1985) and Klebsielfa aerogenes type K54 (Dutton and Merrifield, 1982). In each of these polysaccharides, a trisaccharide sequence of 1.3-a-n-glucose- 1,4-p-~-glucuronicacid- 1,4-a-~-fucose forms the backbone. In both the E. coli and K . aerogenes polymers, there is an 0-acetyl group attached to the fucose, but in neither strain is it in stoichiometric amounts. The E . coli tetrasaccharides were 70% acetylated, whereas in the K . aerogenes EPS, digestion with phage-induced enzymes has shown it to be on the fucose of alternate repeat units (Sutherland, 1967; Dell et a/., 1983). The E. coli polysaccharide carries a /3-D-galactose residue attached as a side chain to the main-chain glucose, whereas the K54 polymer has a p-D-glucosyl residue in the corresponding position. Another E. coli serotype, K27, also resembles these two

BACTERIAL SURFACE POLYSACCHARIDES

209

polysaccharides in its major structural features (Jann et al., 1968). The similarity of these structures to K . aerogenes K54 is sufficient to permit a phage-induced enzyme, now known to be an endoglucosidase, to hydrolyze both polymers (Sutherland et al., 1970).

V. Physical Properties A. ORDER/DISORDER Many of the microbial EPS that have found industrial usage are now known to undergo transition from ordered to disordered states as the temperature of aqueous solutions is raised. The transition process can be studied by various physical techniques such as circular dichroism, measurement of gel strength, and viscosity. Such transitions also undoubtedly occur in most other EPS. As the transition temperatures of those polysaccharides studied so far fall into the range of 30°C and above, the normal state seen in aqueous solution, either in cultures or for isolated polymers, is that of order for the majority of bacterial EPS. In only those species capable of growth at higher temperatures would disordered polysaccharide solutions normally be found. For some EPS, the start of transition has been observed at temperatures as low as lO"C, but completion was generally at 50-90°C (Wolf et al., 1978). Normally a single transition is seen, but Klebsiella type 8 polysacharide is unusual in that it displayed two distinct transition steps at approximately 30" and 42"C, respectively. This was interpreted as a change from an initial helical conformation, characterized by interaction between uronic acids and main-chain hexoses, to a cooperative reorientation of the uronic acid. Subsequently, the backbone reoriented in another helical conformation, which then underwent transition to a less-ordered state as the temperature was raised. The structures of EPS seen associated with bacterial cells, thus, inevitably reflect the ordered conformation adopted by these macromolecules in solution. The chemical structures of the polysaccharides confer specific properties that result in various degrees of inter- and intrastrand association. Such conformation in solution also depends on the physical environment in which it is studied. The majority of EPS possess anionic groups deriving their charge from uronic acids, pyruvate ketals, or phosphate groups, which are capable of interacting with various cations. This too can show marked effects on the solution properties of macromolecules. Gel formation can be a phenomenon occurring either with or without the involvement of cations. In some polymers, specific

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I 4 N W . SUTHERLAND

cations may be necessary, as is seen with alginates of marine algal origin; alternatively, a range of different cations may function in gelation. Even if gels are not formed, the presence of cations can have profound effects on the physical properties of polysaccharide solutions. Polymers that may yield highly viscous solutions in distilled water may show greatly reduced viscosity in the presence of salts, which is due to the altered conformation of the molecules. The effect of anions on transition of EPS has received less attention. A recent study by Clarke-Sturman et a / . (1986) indicated that anions, through their interaction with the solvent water, strongly influence the order-disorder transition. High concentrations of Br- or SCN- decrease polymer stability by lowering the transition temperature, whereas SO:and PO:- have the opposite effect provided they do not cause precipitation of the polymer. The conformation of several EPS has been studied using X-ray fiber diffraction methods. Considerable similarity was seen in several Klebsiella polysaccharides, which adopted helical structures. Isaac et al. (1978) deduced that Klebsiella K57, an EPS with a tetrasaccharide repeat unit, formed a 3-fold helix with an axially projected repeat of 1.143 nrn-a value that correlated directly with the chemical repeat unit. The polysaccharide chain was highly extended, as was that of K9-a polymer with a pentasaccharide repeat unit (Isaac et al., 1981b). Klebsiella K25, another polysaccharide composed of tetrasaccharide repeat units, also adopted a 3-fold helical conformation, the projected chemical repeat unit being 0.97 nm (isaac er al., 1981a). An earlier study on Escherichia coli K29 capsular polysaccharide and on the corresponding material from two mutant strains, all of which contained the same hexasaccharide repeat unit, utilized a model with antiparallel 2-fold helices in which the cations and solvent molecules present were bound only weakly (Moorhouse et a f . , 1977). The molecular configuration was stabilized by three intramolecular and two intermolecular hydrogen bonds for each hexasaccharide unit. It was proposed that the extensively hydrated conformation might persist in uiuo and thus play an important role in the function of EPS as a cell envelope component. Curdlan, a p- 1,3-glucan synthesized by a number of Agrobacterium and related bacterial strains, including some Rhizobium species (Saito et af., 1968), yielded an X-ray diffraction diagram for wet fibers indicative of a simple helical structure (Takeda et a/., 1978). However, some of the molecules were possibly in a triple-stranded-helical form; after heat treatment or other chemical manipulations, single-helical molecules were converted to triple-stranded helices. It is not clear whether the mixture of (predominantly) single and triple helices found in the native material

BACTERIAL SURFACE POLYSACCHARIDES

211

corresponds to the material observed as microfibrils by Harada (1977). From studies of 13C nuclear magnetic resonance (NMR) spectra, the resonance peaks were also ascribed to a single helix (Takeda et al., 1977). Kasai and Harada (1981) used 13C NMR and other techniques to thoroughly study the ultrastructure of curdlan and demonstrate the conversion of the polysaccharide to triple-stranded helices following heating. In the triple-stranded form, curdlan yielded irreversible gels. Although it is insoluble in water, curdlan could be dissolved in sodium hydroxide solutions (Ogawa et al., 1972; Saito et al., 1977). In such solutions, the polymer conformation depended on the NaOH concentration, a reversible change occurring from the helical structure to a random coil conformation. From these results, a model for an orientational gel consisting of micelles interspersed with water molecules was proposed. Despite the large number of studies on xanthan, there is still disagreement over its conformation. Milas and Rinaudo (1979) noted that in dilute aqueous solutions, the melting temperature was directly correlated to the total ionic strength of the solution and was independent of polymer concentration. Since the specific rotation and circular dichroism spectra of chromophores represented lateral side chains, the conformational transition was considered to be helix-coil with simultaneous release of the side chains. This was followed by decreased rigidity of the p-1,4glucan chain as the temperature was raised above the melting point. It was thought possible that spreading of the lateral chains could also increase the swelling and hydration of the main polysaccharide chain. Morris et al. (1983), using a combination of various techniques, obtained results that led to the proposal that xanthan dispersions were weak, gel-like networks in which the polysaccharide molecules were highly associated. Such structure was disrupted by centrifugation or heat treatment, yielding a solution of individual molecules together with “microgel” aggregates-side-by-side associations of up to 47 individual molecules. In true solutions, the molecules were thought to be rod-like with a length of 0.2-2 pm and a molecular weight of 1.1 X lo6. Xanthan molecules of different size have also been produced following degradation by ultrasonic treatment and size exclusion chromatography (Paradossi and Brant, 1982), and then used for study by NMR, optical rotary dispersion (ORD), and UV absorption chromatography. The higher molecular weight material yielded a linear mass density of -2000 Da/nm, whereas lower molecular weight fragments behaved consistently and, in the view of the authors, supported the concept of xanthan solutions existing as double-stranded-helical chains. For single-stranded material, a molar mass per unit contour length of -1000hm would be expected. Further support for the concept of xanthan as a five to one double-

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IAN W. SUTHERLAND

stranded-helical structure in salt solution came from light-scattering studies by Sato et al. (1984), uusing native and ultrasonically treated polysaccharide. When dissolved in cadoxen [tris(ethylenediamine)cadmium dihydroxide], values were consistent with single flexible chains. Maret et al. (1981) compared the long-range order of highly concentrated (7.5% viv) xanthan solutions to cholesteric liquid crystals and found a close similarity using birefringence, optical rotation, and laser light scattering. The left-handed cholesteric screw was interpreted as a single molecule with rod-like conformation and right-handed helical structure. This structure was considerably affected by the counterion present; in the absence of excess salt, the sodium and calcium salts of xanthan corresponded to two different chain conformations (Lambert et al., 1985). The divalent calcium salt represented the ordered conformation whereas sodium xanthan was disordered, their conformation being a stretched coil and a Gngle 5-fold helix, respectively. The divalent counterions were strongly bound to the polysaccharide. A diagrammatic representation of the xanthan molecular transition in aqueous solution is shown in Fig. 9. The strong interaction of xanthan with calcium ions was confirmed by rheological studies. The apparent viscosity of the calcium salt was very large at low temperatures but decreased with increasing temperature. This was interpreted as suggesting very strong intramolecular association of the xanthan molecules, probably mediated by the divalent cations and adjacent charged trisaccharide side chains of different molecules (Tako and Nakamura, 1987). Molecular modeling of xanthan chain conformation has also been attempted. Based on an assumed single polymer chain with a well-defined repeat unit, several stable conforming molecular families were obtained. These generated either hollow or extended helices, one of which appeared to correspond to the ordered conformation of xanthan in the solid state (Perez and Vergelati, 1987). One of the few polysaccharides from gram-positive bacteria to be extensively studied is the polymer from Streptococcus pneurnoniue type 3. The conformation of this material with its relatively simple disac] was charide repeat unit [poly( 1,3-j3-~-glucuronosyl-1,4-P-~-glucose) found to be a 3-fold helix with a repeat of 0.923 nm (Marchessault et al., 1980). T w o possible models for left-handed helices were evaluated, but lack of conclusive X-ray intensity data precluded a choice between them; however, the possibility of a right-handed helix was excluded.

B. MOLECULAR WEIGHT Southwick et uf. (1980) employed quasi-electric light scattering to determine the nature of xanthan in aqueous solution. They concluded that

BACTERIAL SURFACE POLYSACCHARIDES

+

500-1500 nm

213

w

Heat

E

0 0

Salts

1------.)

4 nm

FIG. 9. A diagrammatic representation of the reversible disaggregation of xanthan molecules in aqueous solution. (Reproduced from Tait, 1984 with permission.)

xanthan undergoes self-association in aqueous solution but is inhibited by 4 M urea, indicating a possible role for hydrogen bonding in the process. They reported that the molecular weight of the unassociated form of the polysaccharide was 2.16 x lo6, and that a value very similar to this has been obtained in earlier studies, in addition to a similar value for xanthan dissociated in urea. (Dintzis et al., 1970). In contrast to these estimates of the molecular weight, a study using the sedimentation coefficient of xanthan labeled with a fluorescent group yielded a much higher value for native material-15 X lo6 (Holzwarth, 1978). Apart from xanthan, molecular weight estimations have been performed on a number of other EPS, including many from the genus Kfebsiella. The majority of the polymers were in the molecular weight range 1.13-2.94 X lo6,but the value for one preparation was as low as 1.7 x 104 when determined by light scattering (Wolf et al., 1978). These values should probably be regarded with caution, as it is not clear whether they represent monomeric or oligomeric conformations. Further, since

214

IAN W . SUTHERLAND

they were products of single batch cultures on solid media, values from other culture conditions might differ especially in shaken or stirred fluid cultures where the polymer solutions would be exposed to shear forces. The molecular weight of EPS is one of the properties that is thought to be affected by the physiological conditions under which the bacteria are grown (e.g., Sutherland, 1985). This has been confirmed recently for xanthan from Xanthomonas carnpestris at various stages of batch culture (Lecacheux et al., 1986). The product from 10-hour cultures had a molecular weight of 1.22 x lo6, as opposed to 2.2 X lo6 over the time range of 20-60 hours. Although it was suggested by these authors that molecular weight was not very sensitive to fermentation conditions, the results presented did not investigate different nutrient limitations or growth rates. Variation in the latter showed that the intrinsic viscosity of the same polysaccharide depended on the nutrient limitation employed and in batch culture, on the growth phase sampled (Tait et al., 1986), with very marked differences being found. Similar results had been reported earlier from other laboratories. The conformation of an EPS in solution may affect its susceptibility to enzymatic degradation as well as other physical properties. Xanthan from Xanthomonas carnpestris is now known to be degraded by a number of enzymes obtained from either pure or mixed heterologous bacterial cultures (e.g., Cadmus et al., 1982: Sutherland, 1982). These enzymes function in the presence of moderate salt concentrations or in one case (Hou e? al., 1986), at relatively high salt concentrations, thus cleaving the ordered form of the polysaccharide. They are all 1,4-endo-/3-glucanases that also degrade carboxymethylcellulose. Xanthan is not hydrolyzed by other cellulases despite its cellulosic backbone except in salt-free aqueous solutions (Rinaudo and Milas, 1980). The rate of hydrolysis depended on the solution quality, being reduced by the presence of aggregates or microgels. Hydrolysis thus occurred only on the unordered chain, and the side chains could be considered to protect the /3-1,4-glucan backbone from the enzyme. Alternatively, the observation that “excess” glucose was released by fungal cellulases during hydrolysis of unordered xanthan led us to suggest that, at sectors deficient in side chains, hydrolysis was favored under such conditions (Sutherland, 1984a). A further effect of the conformation change in xanthan solutions was seen in studies on their shear stability. The ordered form found in the presence of salts exhibits relatively high shear stability, but in the disordered form, conformation suffered degradation after 30-minute shear at 5000isecond (Chen and Sheppard, 1980). The changes in physical properties occumng in the ordered and disordered states make it very likely that any observation of the appearance of capsules, fibers, or

BACTERIAL SURFACE POLYSACCHARIDES

215

related material is highly dependent not only on the physiological conditions employed for bacterial culture, but also on the way in which the sample has been handled during preparation. This includes the effects of both chemical treatment and any physical degradation, such as shear force, to which it has been exposed.

C. GELFORMATION A number of EPS synthesized by prokaryotes are water insoluble. This is a consequence of their molecular size and chemical structure. Such polymers, of course, include bacterial cellulose from Acetobacter xylinum and also the flocs formed by Zoogloea spp. There are also numerous examples of EPS that are found as insoluble gels, including “mutan” from Streptococcus mutans, curdlan from Agrobacterium turnefaciens, and other species in addition to the neutral polymer recently discovered in Rhizobium (Zevenhuizen, 1984). This EPS is of particular interest in that it is common to certain strains of Rhizobium leguminosarum and R . trifolii, but is absent from R. phaseoli and R . melitoti. Production of capsular material occurs in addition to synthesis of acid slime polysaccharides, their monomeric repeat units, and cyclic @-I,Zglucans. Synthesis of capsular gels occurred in the stationary phase of growth. The structure was shown to represent a neutral hexasaccharide repeat unit (Fig. 10) in which the main chain was predominantly 1,3 and 1,4 linked. This polysaccharide formed gels at relatively low concentrations, and no ions were necessary to promote gelation. The gelation of alginates has been studied using marine algal alginates, but many of the findings can be extrapolated to A . uinelandii alginate with similar poly-D-mannuronic acid and poly-L-guluronic acid sequences. Gelation of the polyuronic acid alginate, promoted by the presence of D-GAL p

’t

t“

1

0-GAL p 14 11

D-GALp

FIG. 10. The neutral hexasacchariderepeat unit of gel-forming EPS produced by various Rhizobium species.

216

IAN W. SUTHERLAND

Ca”, involves cooperative binding of the cations by polyguluronic acid sequences in microcrystalline junction zones (Rees, 1969; Bryce er al., 1974). An “eggbox” model was proposed by Grant et a / . (1973) in which there were regular packing and coordination of the cations in interstices of associated extended sections of the polysaccharide chains. Polyguluronic acid sequences played the major role, but circular dichroism (CD) studies also indicated a possible function for polymannuronic acid portions of the macromolecule. The extent of the change in CD spectra reflected the polyguluronate content of the alginate, decreasing as the polyguluronate in the polymer sample decreased (Thom er al., 1982), and supported the view that such sequences are primarily responsible for interchain association and consequent gel formation. The polyguluronic acid sequences of alginate possess a higher selectivity for Sr?’ than Ca”, which could also be demonstrated by comparison of CD spectra. In the presence of Mg2+, which does not induce gelation, relatively limited formation of junction zones was postulated. Interaction with monovalent cations also occurs to an extent sufficient reflect the polyguluronate content of the alginate (Seale ef a / . , 1982), but insufficient to cause gel formation. The alginates synthesized by A . uinelundii, insofar as they have been investigated, show marked similarity to algal polymers, including gel formation with 1980). However, the effect of acetyl groups on the Cu’- (Paoletti et d., gel-forming capacity of mannuronosyl residues of these polysacchrides is uncertain. Pseudornonas alginates, because of their lack of contiguous L-guluronic acid residues (SkjHk-Braek et al., 19861, do not readily form gels in the presence of Ca2+ or S8’. Unlike A . uinelundii and algal alginates, the proton form also fails to yield gels. However, at higher polymer concentrations, some Pseudornonas EPS of the alginate type can gel in the presence of Ca” (1. W. Sutherland, unpublished results) . Such gel-forming bacterial polysaccharides are probably more common than is realized, as most attention has until now been paid to watersoluble polymers because of their ease of isolation and purification. A series of recent isolates from our laboratory (Chard et a/., 1987) includes examples of bacteria yielding water-insoluble gels that could be recovered from culture supernatants by centrifugation. These bacteria included some with colonial morphology such as that shown in Fig. 1 , although it is not clear whether such appearance is necessarily due to the water insolubility of the polymers. The chemical structure of these polysaccharides is as yet unknown, but may resemble mutan and curdlan in being predominantly 1,3 linked or, perhaps, 1,4 linked as in cellulose, chitin, and other structural molecules such as peptidoglycan.

BACTERIAL SURFACE POLYSACCHARIDES

217

GELLING D. SYNERGISTIC A further property of polysaccharides, which has been recognized for some time and is used in a number of industrial applications, is the ability of two dissimilar species to interact. Two different types of interaction between dissimilar polysaccharides have been identified (Dea et a f . , 1977). Incompatible molecules are mutually excluded, resulting in an increase in the effective concentration of each type. Alternatively, there may be an energetically favorable association of the structurally and sterically regular regions of the unsubstituted polysaccharide backbones. Such interchain associations have been reported for several mixtures of polysaccharides and it is possible that polymer/polymer interactions involving EPS may also have various biological roles. There is, as yet, no indication that synergistic interactions occur when a bacterial strain synthesizes more than one EPS. Numerous studies have been made on mixtures involving xanthan and plant gluco- or galactomannans. When such mixtures are heated and allowed to cool, synergistic gels are formed. Whether such gross reactions occur naturally or whether more subtle interactions exist, is by no means clear. Various rheological techniques have been used to study the interchain associations. Galactomannans, especially those in which substitution with a-D-galactosyl side chains is relatively low, interact with xanthan to form firm gels at low total polysaccharide concentrations (Dea et a]., 1986). The EPS : plant galactomannan interaction was highly dependent on the fine structure of galactomannan. On the basis of optical rotation studies, Dea and Morris (1977) concluded that the native, ordered xanthan structure existed in such synergistic gels. The gels were, therefore, thought to represent the result of cooperative association of galactose-free portions of the mannans in regular, ribbon-like conformation with the ordered xanthan structure. When xanthan was mixed with an acetylated glucomannan obtained from the tubers of Amorphophallus konjac, a very strong interaction was obtained resulting in gelation at total polysaccharide concentrations on the order of 0.05% (i.e., within the concentration range expected in capsular polysaccharides). In marked contrast to xanthadgalactomannan gelation, neither EPS from Arthrobacter species, which do possess ordered conformation in solution, nor random coil polysaccharides, such as dextran, show similar interactions (Dea et al., 1977). The physical properties of a polysaccharide can be greatly affected by relatively slight changes in chemical structure. The presence or absence of acyl groups may have considerable effects on the conformational

218

IAN W . SUTHERLAND

association of EPS molecules in solution and, thus, on their physical properties. As pointed out by Morris and Myles (1986), little is known about features of the chemical structure of EPS that are of particular significance in determining the interactions of these polymer molecules and their helical conformation. The existence of a number of natural polymer "families," in which there are slight differences in chemical structure, has enabled preliminary determination of the role of certain carbohydrate and noncarbohydrate substituents. Thus, Atkins e f al. (1987) were able to compare the effect of acetylation on molecular interactions and gelling properties of one bacterial exopolysaccharide. Fiber X-ray diffraction revealed that both had similar helical structures, but in the O-acetylated polymer, association between molecules was inhibited. This could account for the failure of this particular polysaccharide to form gels. Using the gellan series of polysaccharides, Crescenzi e t al. (1987) were able to study the influence of side chains on the properties of these polysaccharides in dilute solution. Increase in ionic strength over a narrow range of salt concentration promoted cooperative conformational transition of the unbranched polysaccharide. It also underwent a thermally reversible conformational transition under the ionic conditions tested. In contrast to these findings, the side chains on related polymers appeared to mediate against conformational ordering. This also explained the lack of gel formation by the polysaccharides possessing side chains. There have been studies on a number of different systems including gellan, xanthan, and Klebsiella (Enterobacter) aerogenes K54, all of which reveal the importance of the presence or absence of acyl groups. The polymer from Pseudomonas elodea and a similar polysaccharide from a closely related bacterium were highly viscous in aqueous solution in the natural state (Kang et al., 1982; Anson e f a/., 1987). Yet, when chemically deacetylated, each formed firm gels at relatively low concentration (approximately 0.8%).This alteration of chemical structure probably does not relate to any observed material physically associated with the respective bacteria, but a family of EPS with and without O-acetyl substituents provides a naturally occurring example amenable to study. The bacteria concerned do not appear to show any observable differences in their slime polysaccharides. They share the same tetrasaccharide carbohydrate repeat unit (Table 11) but in Enterobacfer aerogenes XM6, carry no O-acetyl groups (O'Neill e f al., 1986a). The polysaccharide from this strain forms highly viscous solutions and on the addition of various salts, yields thermoreversible gels (Nisbet et af., 1984). Thus, at temperatures below 30"C, even in the presence of monovalent cations at relatively low concentrations, ordered gel structures are produced at polymer

219

BACTERIAL SURFACE POLYSACCHARIDES TABLE TI GEL FORMATION BY Klebsiella TYPEK54 POLYSACCHARIDES Strain K . aerogenes K54 KS4 deacetylated" K . aerogenes A3SI A3S1 deacetylated" Enferobacferaerogenes XM6

Composition

Molar ratio

Glc :Fuc : GlcA : acetate Glc : Fuc : GlcA Glc : Fuc : GlcA : acetate Glc : Fuc : GlcA Glc : Fuc : GlcA

2 :1: 1: 1 1:l:l 2 : 1 : 1 : 0.5 2:l:l

Gel formation

2:l:l

-

+ + +

' O-Acetyl groups removed by mild alkaline treatment.

concentrations of 0.3%. The corresponding polysaccharide from strain A3(S1) does not gel, but on chemical deacetylation, resembles that of XM6. What is particularly significant is that in the A3 polymer, the O-acetyl groups are present as regular substituents on every alternate repeat unit. Thus, gelation clearly requires a sequence of nonacetylated oligosaccharides, but is inhibited by the presence of a single acyl group on each octasaccharide. In xanthan solutions, macromolecular associations commence at lower concentrations when both ionic strength and degree of pyruvylation are high (Smith et al., 1981). It was proposed that at high-ionic strength, the electrostatic repulsions were diminished and association was promoted by apolar internal interactions of the pyruvate methyl groups. Once the fraction of side chains carrying pyruvate exceeded -0.3 1, the viscosity increased fractionally on addition of KCl to salt-free xanthan solutions and further pyruvylation.

VI. Function

SYMBIOSIS, AND INFECTION A. RECOGNITION, In many phytopathogenic bacteria that are capable of producing EPS, there is a proven correlation between virulence and EPS synthesis. Loss of the ability to produce EPS, frequently found in laboratory subcultures, leads to loss of or decrease in virulence. Thus, avirulent strains of Erwinia stewartii lacked capsules and secreted low levels of EPS into the culture medium when compared with wild-type strains (Bradshaw-Rouse et al., 1981). It is not clear whether the requirement for EPS synthesis is always an attribute of pathogenesis. Rather, it may reflect a requirement for recognition and initial interaction between the prokaryote and its eukaryotic host, as is seen in the Rhizohiumilegume interaction involved in symbiotic nitrogen fixation.

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Some bacterial polysaccharides appear to have one or more nonspecific roles in the infective process. One effect of EPS is to cause wilting, disturbing the supply of water and soluble nutrients through the plant. Even this effect is probably complex, the macromolecules impeding the water flow and rendering the fluid highly viscous. There are also secondary effects due to release of substances from the plant in response to bacterial products. Polysaccharide from X . campestris var. phaseoli induced wilt in seedlings of tomato, sunflower, and bean, the effect being reversible in early stages and correlating with low water uptake by the seedlings (Leach et a ) . , 1957). The effect was apparently nonspecific, all types of seedlings being similarly affected apparently through occlusion of the water-uptake mechanism. Slime from the plant pathogen X . campestris var. oryzae has also been shown to cause wilting of rice cuttings (Choi et af., 1982), the effect being noted with EPS from both virulent and avirulent bacterial strains. The effect was probably caused by decreased vascular flow. The exact mechanism by which bacterial EPS induce wilt symptoms is still not known. As opposed to nonspecific mechanisms, Goodman and White (1981) suggested that there is also a recognition element involving the EPS, which then affects cells in the plant tissue. The possible role of EPS conformation in wilting was indicated by Sijam et al. (1985) using Erwinia amylovora polymer. Addition of salt to the polysaccharide, which greatly reduced the solution viscosity, decreased or eliminated the ability to cause wilting of plant shoots. Less explicable was the observation that oligosaccharides produced by the action of phage-induced enzymes retained the ability to cause wilting, but this effect was also absent when salt was added to the system. When radioactive EPS or fragments interacted with plant material to cause wilting, they were retained at the site of contact with the plant, but salt inhibition caused polysaccharide or oligosaccharides to be distributed throughout the plant shoot. A study of Rhizobium mutants altered in EPS synthesis indicated a lack of correlation between polysaccharide production and the ability to nodulate host bacteria (Sanders et a / . , 1981). An extension to this work using R . melitoti confirmed that most, though not all, of a group of genetically diverse but phenotypically homogeneous EPS- mutants formed non-nitrogen-fixing nodules on alfalfa (Leigh et al., 1985). These results were interpreted to imply that this bacterial EPS was essential for the formation of an effective nodule, the defect probably residing in nodule invasion. There are several possible roles for the EPS, some being indirect-promotion of root hair deformation or infection thread formation-whereas others are direct, e.g., a structural component of the

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22 1

effective nodule. It has also been suggested that R . trifolii EPS deficient in acyl groups (0-acetyl esters or pyruvate ketals) might be ineffective in interactions involving nodulation (Skorupska et al., 1985). These conclusions were based on competition studies using mutants and chemically deacylated polymers, but it was not clear whether more extensive degradation of the polysaccharide or loss of the side chains might have occurred, particularly in the latter material. In another similar study, a strain of R . japonicum and a mutant lacking the symbiotic plasmid both yielded EPS that appeared to be similar in all respects tested (Carlson and Yadav, 1985). Although the glycosidic structure of the EPS was not determined by the plasmid, the possibility remained that it might, in some way, regulate the presence or the location of substituent acyl groups. Recognition of EPS may also play a role in infection of humans and animals through interactions other than those of antigen and antibody. Although alginate-producing strains of P . aeruginosa are widely recognized in cystic fibrosis patients (e.g., Dogget and Harrison, 1972), the role played by this specific EPS in the infection process is not known. A lectin recognizing the bacterial alginate has been found in soluble extracts from rat lung tissues (McArthur and Ceri, 1983). It was postulated that an interaction of this type could represent an adhesion mechanism forming part of the pathogenisis of P . aeruginosa. The lectin recognized P . aeruginosa alginates but not algal preparations, thus exhibiting a possible requirement for the 0-acetylated D-mannuronic acid and/or the single guluronic acid residues of the bacterial polymer. Similarly, binding of both capsular and soluble EPS from Rhizobium japonicum by lectins from soybean plants has been reported (Tsien and Schmidt, 1981). The polysaccharide was thought to differ only in molecular weight, the soluble material being 2-5 x lo6 whereas the capsule represented very highmolecular-weight gel-like material found in close association with the bacteria. Interaction of either polymer preparation with lectin was inhibited in the presence of galactose (or N-acetyl-D-galactosamine), indicating the probable “immune” dominance of galactosyl residues in the EPS (see Fig. 6). Lectin interaction with cell-bound EPS was asymmetric, the polymer being associated with one pole of the bacterium and not with the other, which tended to contain a high poly-phydroxybutyrate level revealed by electron microscopy. Binding of Pseudomonas solanacearum by a lectin from potato tubers was demonstrated by Duvick and Sequiera (1984), but the specificity of the reaction was not entirely clear, as both EPS and LPS were bound. A problem encountered in the study was the possibility that there was nonspecific binding of the anionic polymer to the highly basic lectin; binding was inhibited at ionic strengths >0.03 M . This suggests that either the charge

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on the polymer has some effect or a change in conformation occurs and prevents binding to the lectin. The possible biological activity of Rhizobium polysaccharides including EPS has been discussed in a recent review by Halvorsen and Stacey (1986). Although such molecules undoubtedly play a role in the recognition and establishment of symbiosis, few clear-cut answers have emerged. Polysaccharide-degrading enzymes from the plant root are present and modify the polysaccharides (e.g., Dazzo et d.,1982); this may not always be a requirement for polymeric material, but rather for oligosaccharide fragments derived from it that act as signal molecules. Various results are seen from the interaction between plant cells and polysaccharides or oligosaccharides, including root hair curling, nodule initiation, and infection. In almost all studies reported so far, it has not been possible to determine unequivocally whether EPS or other carbohydrates, such a5 LPS and p-1.?-glucans, are the essential molecules, or whether a combination of these substances is required to achieve the complete symbiotic process. It is now also clear that EPS cannot be determinants of host specificity since the glycosyl sequences and locations of nonglycosyl substituents are identical in octasaccharide repeat units from a number of Rhizobirtrn acid EPS, although the bacterial strains were obtained from different plant hosts.

B. EPS

IN

MORPHOGENESIS A N D DEVELOPMENT

In at least two prokaryotic systems, EPS appears to play a key role in morphogenesis. These systems involve gram-negative bacteria and the formation of microcyst- or myxospore-resting cells. However only single cells in A . uine/andii and related species, the system involves whereas in myxobacteria. changes in single cells can be accompanied by social behavior leading to the production of exotic fruiting body structures in which the resting cells are enclosed. The vegetative cells of A . uinelnrrdii are generally surrounded by relatively large amounts of EPS in the form of bacterial alginate. When the cells convert to the resting microcyst form, TEM of thin sections shows two major polysaccharide-containingstructures, the intine and the exine (Lin and Sadoff. 1969). Both of these forms were shown to contain carbohydrate. D-rnannuronic acid and L-guluronic acid being the only sugars identified, i.e., the polysaccharide alginate was present in both layers. accounting for 32 and 13%, respectively (Page and Sadoff. 1975). Differences were found in the ratio of mannuronic to guluronic acid in intine, exine, and EPS. The ratio was 0.45 in exine, 1.81 in intine, and 1.24 in EPS. but the uronic acid composition depended on the level of Ca2+in

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the medium. This is a reflection of the calcium requirement of the epimerase involved in postpolymerization conversion of mannuronic acid residues to guluronic acid (Larsen el al., 1986). Omission of the Ca2+(or Mg”) leading to abortive encystment yielded uronic acid ratios of 5.94 (or 4.6), respectively. The actual role of the alginate in the encystment process is not clear, but it is essential since alginate-deficient mutants failed to form mature cysts (Sadoff, 1975). Azospirillum brasilense is another nitrogen-fixing bacterium apparently capable of encystment, but lacking the clearly defined intine and exine of Azotobacter species (Sadasiva and Neyra, 1985). As cells aged, they converted from vibroid forms to ovoid cells surrounded by a thick polysaccharide layer. Simultaneously, flocs were found in liquid medium, the major polysaccharide component of these being tentatively identified as cellulose on the basis of cellulase hydrolysis and calcofluor staining. Whether there are other bacterial groups behaving similarly is still not known. Myxobacteria possess EPS associated with vegetative cells, but they also excrete such polymers in the process of fruiting body formation. The EPS slime associated with vegetative bacteria is unusual in apparently being neutral material lacking uronic acids and other charged compounds (Sutherland and Thomson, 1975). It has been suggested that this EPS is essential for gliding motility, being a common feature of myxobacteria and Cytophaga species (Burchard, 1981a, b). EPS in the form of slime or sheath material is a characteristic of most gliding types of prokaryotic cells and can frequently be observed as a “slime trail” on solid surfaces. As well as in any role possibly associated with vegetative bacteria, EPS is present with apparently the same composition both in the fruiting bodies of myxobacteria and as part of the myxospore coat (Rosenberg, 1984). A problem encountered in electron microscopic studies of fruiting body formation has been the presence of very large amounts of EPS. This tends to obscure the actual bacteria. However, Vasquez et al. (1 985) have used the results of SEM to postulate that fruiting body formation by Stigmatella aurantiaca is a complex process occurring in several stages with different motility patterns. As the cells aggregate and form fruiting bodies, they are closely associated with EPS, some of which, in the form of slime trails, may orientate the bacteria. A further role for the EPS was suggested, that of binding to the bacteria and aligning them within the population. The complex mixture of cells and EPS observed in S. aurantiaca fruiting body formation can be seen in Fig. 11A, in which the stalk is fully developed. As demonstrated in Fig. 11B, the bacteria are oriented upward, frequently at an angle, in large amounts of the slime matrix. Unlike A . vinelandii, in which the polysaccharides involved are

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FIG. 1 I . Development of the stalked fruiting body of Srigmatella auruntiucu. (A) shows the fully developed stalk. whereas in ( B ) , the upward orientation of the bacterial cells can be seen. (Reproduced from Vasquez er al., 1985, with permission.)

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well characterized, little is known about the structure or properties of myxobacterial EPS, nor are there readily available EPS- mutants. The exact role of EPS in fructification thus remains to be elucidated. It is, however, of particular interest that this group of bacteria, some of which do not readily hydrolyze polymeric carbohydrate or utilize carbohydrates, should expend so much nutrition and energy on EPS production at a time of apparent starvation.

C. ADHESION Bacteria adherent to surfaces in both freshwater and seawater are frequently associated with large amounts of EPS. Corpe (1970) showed that Pseudomonas atlantica cells were embedded in large masses of EPS, and similar findings have been reported from both natural habitat and in vitro studies. Detailed discussion of different aspects of adhesion can be found in the recent series of reviews edited by Savage and Fletcher (1986). Specific aspects of the role of EPS in adhesion have also been discussed (Sutherland, 1984b). Of the three major stages thought to occur in adsorption-reversible adhesion, irreversible adhesion, and microcolony formation-EPS probably has a major role in only microcolony formation (Allison, 1985) where it provides a nonspecific adhesive, the composition of which obviously depends on the bacterial species present. In a comparison of EPS- and non-EPS-producing bacteria. It was clear that all types tested adhered, but only those capable of secreting EPS developed microcolonies on the submerged surfaces (Allison and Sutherland, 1984). This was true of both capsulate and slime-forming bacterial species. Development of the microcolonies was relatively rapid and was associated with the synthesis of considerable amounts of polysaccharide, a finding also made by Wardell et al. (1980). The polysaccharides involved in adhesion of this type are poorly characterized. The component sugars of a few of them have been identified, but structural details that might permit some correlation with physical properties are lacking. The secondary and tertiary structures of the polymers may have a very important role in maintaining adhesion, especially once microcolonies and multiple layers of bacterial cells have developed. Allison (1985) found that under reduced divalent cation concentrations, EPS-producing bacteria were still capable of adhesion to glass surfaces but failed to produce extensive microcolonies. The lack of divalent cations may well have had a significant effect on the conformation of the polysaccharides and their resultant ability to bind cells to the glass surface. As the surface itself plays an important role in the adhesion of microorganisms, any appraisal of the role of polysaccharides

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has to consider the interrelationship of surface charge, polymer charge, and associated ions. If the surface is subjected to high shear forces, the shear thinning of (mainly pseudoplastic) bacterial EPS may also affect adhesion, reducing viscosity and probably enabling release of bacteria from the surface film or matrix.

VII. Conclusions Bacterial EPS are a very diverse series of macromolecules, some of which show a relatively high degree of conservation of structural features. They produce a highly hydrated and in many cases, an anionic environment immediately surrounding the bacterial cell. This, in turn, provides the bacteria with a means of complexing cations close to the outer surface. In some polysaccharides, there is a marked specificity for the binding of certain cations. The EPS possess various physical characteristics, foremost of which is the ability to undergo transition from an ordered to a disordered conformation in solution. The transition is dependent on factors such as the ionic environment, temperature, and, above all, the chemical composition of the polymer. Thus, the appearance of material observed by elecron microscopy is highly dependent on its prior history and on the conformational state in which it is prepared. The function or functions of EPS pose many unanswered questions. The polysaccharides undoubtedly function in adhesion, recognition, and pathogenesis, as well as in prokaryotic development. The molecular subtleties of many of these processes are far from clear. Equally, it is still not possible to relate the details of chemical structure to the physical properties of EPS in solution. However, the upsurge of interest in these molecules, together with improved methods of chemical analysis and determination of physical characteristics, will hopefully lead to rapid advances in the future. ACKNOWLEDGMENTS The author is grateful to Professor M . E. Bayer, Dr. G . L. Truchet, Professor D. White, and Dr. L. P. T. M . Zevenhuiren for the gift of photographs and for the permission to reproduce them in this review.

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Marchessault. R. H . , Imada, K., Bluhm. T.. and Sundarajan, P. R. (1980). Curbohydr. Res. 83. 287-302. Maret. G.. Milas. M., and Rinaudo. M. (1981). Polxrner Bull. 4, 291-297. Melton. L. D.. Mindt. L . , Rees. D. A.. and Sanderson. G . R. (1976). Carhohydr. Kes. 46, 245-157. Milas. M.. and Rinaudo. M . (1979). Cnrbohydr. Re.s. 76, 189-196. Moorhouse, R.. Winter. W . T . . Arnott. S . . and Bayer. M. E. (1977). J . Mol. B i d . 109, 373-91. Morris. V. J.. and Miles. M . f 1986). / f i / . J. B i d . Marromol. 8, 342-348. Morris. V. J.. Franklin. D.. and I'anson. K . (1983). Cnrbohydr. Res. 121, 13-30. Nishet. B. A , . Sutherland. I. W., Bradshaw. I. J . . Kerr. M. H., Morris. E. R.. and Shepperson. W . A . ( 1984). Carbohydr. Polymers 4, 377-394. Ogawa, K.. Tsurugi. J.. and Watanabe. T. (1972). Cheni. L e u . 95-98. O'Neill. M . A , . Morris. V. J . . Selvendran. R . R., Sutherland. I . W.. and Taylor, 1. T. ( 1986a). C'urhohydr. Re3. 148, 63-69. O'Neill. M. A , . Sclvendran. R. R..Morris. V. J . . and Eagles. J. (l986b). Carbohydr. Res. 147. 195-313. O m a n . S. F . , Fett. W. F.. and Fishman, M. L. (1986). J . Bactcriol. 166, 66-71. Page. W . J.. and Sadoff. H . L . (1975). J . Bncferiol. U2 , 145-151. Paoletti. S . . Cesaro. A,. Ciana, A.. Delben, F.. Manzini, G.. and Crescenzi, V. (1980).A m . (-hem. Soc. Sytnp. 150, 379-386. Paradossi. G.. and Brant. D. (1982). Macromology 15, 874-9. Perez. S., and Vergelati. C. (19871. Ini. J. B i d . Macmtno/. 9, 211-218. Ravenicroft, N..Merrifield. E. H.. and Stephen. A. M. (1985). S. Afr. J . Sci. 81, 380. Rees. D. A . (1969). Adu. Carhohydr. Chem. Biochem. 24,267-332. Rinaudo, M . . and Milas. M. (1980). 1111.J . Bioi. Mrrcromol. 2, 45-48. Rosenberg, E. (1984). "Myxobacteria." Springer. New York. Ross. P., Weinhouse, H.. Aloni. Y.. Michaeli. D., Weinberger-Ohana, P., Mayer, R., Braun. S.. de Vroom. E . . van der Marel. G . A., Van Boom. J. H., and Eenziman, M. ( 1987). ~\,.utrtre (I.ondon) 325, 279-181. Sadasiva. L . . and Neyra. C. A. (1985). J. Bacferiol. 163, 716-723. Sadoff, H . 1.. (1979. Bucieriol. Rev. 39. 516-539. Saito. H.. Misaki. A . . and Harada, T. (1968). Agric. Biol. Clzem. 32, 1261-1269. Saito. H., Ohki. T.. and Sasaki, T. (1977). Biocliemisir? 16, 908-913. Sanders. R.. Raleigh. E . , and Signer, E. (1981). Nafiire (London) 292, 148-149. Sato. T.. Norisuye. T.. and Fujita. H. (1984). Polymer J. 16, 341-350. Savage. D. C . . and Fletcher. M. (198% "Bacterial Adhesion Mechanisms and Physiological Significance." Plenum. New York. Savidge. R. A.. and Colvin, J . R. f 1989. Can. J . Microbid. 31, 1019-1025. Seale. K.. Morris. E. R.. and Rees. D. A. (1982). Ccobohydr. Rcs. 110, 101-112. and Gacesa. P. (1984). Carbohydr. Res. 135, Sherbrock-Cox. V.. Russell. N. .l.. 147- i 54. Sijarn. K.. Goodman. R. N.. and Karr. G. L. (1985). Phjsiol. Plant. Pntlrol. 26,23 12-2332. Skjak-Braek. G.. Grasdalen. H., and Larsen, B. (1986). Carbohydr. Res. 154, 239-250. Skorupska. A.. Deryko. -M., and Lorkiewicz, Z. (1985). Arch. Microbiol. 143, 307-310. Smith. I. H.. Symes. K. C . . Lawson. C . J.. and Morris. E . R. (1981). Int. J . Biol. Macrornol. 3, 129-134. Southwick. J . G . . Lee. H.. Jarnieson, A . M.. and Balckwell. J. (1980). Curbohydr. Res. 84, 287-1,95. Stokkc. B . . Elgsaeter. A . . and Srnidsrod. 0. (1986). I n t . J . B i d . Mocromol. 8. 217-225.

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Stokke, B. T., Elgsaeter, A,, Skjak-Braek, G., and Smidsr@d,0. (1987). Carbohydr. Res. 160, 13-28. Sutherland, I. W. (1967). Biochem. J. 104, 278-285. Sutherland, I. W. (1981). Carbohydr. Polymers 1, 107-115. Sutherland, 1. W. (1982). J . Appl. Bacteriol. 53, 385-393. Sutherland, I. W. (1984a). Carbohydr. Res. l31,93-104. Sutherland, I. W. (1984b). Crit. Rev. Microbiol. 10, 173-201. Sutherland, I. W. (1985). Annu. Reu. Microbiol. 39, 243-270. Sutherland, I. W., and Thomson, S. (1975). J. Gen. Microbiol. 89, 124-132. Sutherland, I. W., Jann, K., and J a m , B. (1970). Eur. J. Biochem. 12, 285-288. Tait, M. I. (1984). Ph.D. thesis, Edinburgh University. Tait, M. I., Sutherland, I. W., and Clarke-Sturman, A. J. (1986). J . Gen. Microbiol. 132, 1483-1492. Takeda, H., Yasuoka, N., and Kasai, N. (1977). Carbohydr. Res. 53, 137-152. Takeda, H., Yasuoko, N., Kasai, N., and Harada, T. (1978). Polymer J. 10, 365-368. Tako, M., and Nakamura, S. (1987). Agric. Biol. Chem. 51, 2919-2923. Tayama, K., Minakami, H., Entani, E., Fujiyama, S . , and Masai, H. (1985). Agric. Biol. Chem. 49,959-966. Tayama, K., Minakami, H., Fujiyama, S., Masai, H., and Misaki, A. (1986). Agric. BioL Chem. 50, 1271-1278. Thom, D., Grant, G . T., Morris, E. R., and Rees, D. A. (1982). Carbohydr. Res. 100,29-42. Tsien, H. C., and Schmidt, E. L. (1977). Can. J. Microbial. 23, 1274-1284. Tsien, H. C., and Schmidt, E. L. (1981). J. Bacteriol. 145, 1063-1074. Umbreit, T. H., and Pate, J. L. (1978). Arch. Microbiol. 118, 157-168. Vann, W. F., Schmidt, M. A., J a m , B., and J a m , K. (1981). Eur. J . Biochem. 116,359-364. Vann, W. F., Soderstrom, T., Egan, W., Tsui, F-P., Schneerson, R., Idrskov, I., and Qrskov, F. (1983). Infect. Zmmun. 39, 623-629. Vasquez, G . M., Qualls, F., and White, D. (1985). J . Bacteriol. 163, 515-521. Vasse, J. M., Dazzo, F. B . , and Truchet, G. L. (1984). J. Gen. Microbiol. 130, 3037-3047. Wardell, J. N., Brown, C. M., and Ellwood, D. C. (1980). Zn “Microbial Adhesion to Surfaces” (R. C. W. Berkeleley, J. M. Lynch, J. Melling, P. R. Rutter, and B. Vincent, eds.), pp. 21 1-30. Horwood, Chichester. Weng, E., Valla, S . , Anthonsen, T., Paulsen, B. S . , and Srnidsr@d,0. (1984). Absrr. Int. Carbohydr. Symp., 12th D 9.5. Wicken, A. J., Ayres, A., Campbell, L. K., and Knox, K. W. (1983). J. BacterioL 153, 84-92. Wolf, C . , Elsaesser-Beile, U., Stirm, S., Dutton, G. G . S., and Burchard, W. (1978). Biopolymers 17, 731-748. Wood, P. J. (1980). Carbohydr. Res. 85, 271-287. Yadomae, T., Yamada, H., Miyazaki, T., Omori, T., and Hirota, T. (1978). Carbohydr. Res. 60,129-139. Zaar, K. (1979). J . Cell Biol. 80, 773-777. Zevenhuizen, L. P. T. M. (1984). Appl. Microbiol. Biotechnol. 20, 393-399. Zevenhuizen, L. P. T. M., Bertocchi, C., and van Neervan, A. R. W. (1986). Antonie uan Leeunenhoek 52. 381-386.

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INTERNATIONAL REVIEW OF CYTOLOGY, VOL. 113

Reorganization of the Egg Surface at Fertilization FRANK J. LONGO Department of Anatomy, University of Iowa, Iowa City,Iowa 52242

I. Introduction As a result of the interaction and fusion of the gametes, processes are initiated that move the previously quiescent egg on to pathways leading to its cleavage, differentiation, and eventual development into an adult organism. It is noteworthy that many of the early processes of fertilization, such as the transient, positive shift in potential (fertilization potential), cortical granule reaction, and intracellular pH increase, are related to or are a direct result of alterations in the function of the egg plasma membrane. Analyses of these early aspects of development, particularly those related to the role of ions in egg activation, have been published (Shen, 1983; Jaffe, 1985; Jaffe and Gould, 1985; Whitaker and Steinhardt, 1985). In this article, structural, chemical, and functional changes of the egg plasma membrane as a result of gamete fusion and cortical granule dehiscence are discussed. Attention is directed to investigations examining the fate of the sperm plasma and cortical granule membranes following their fusion with the egg plasmalemma, and to studies that may lead to an understanding of fundamental mechanisms as they are related to metabolic changes of the activated ovum. Processes related to aspects presented here, including dynamic changes of the egg cortex and secretory functions of cortical granules during fertilization, have been reviewed (Gulyas, 1980; Shapiro and Eddy, 1980; Vacquier, 1981; Schuel, 1985; Kay and Shapiro, 1985; Longo, 1987a).

11. Egg Cortical Structure The cortices of sea urchin eggs, which provide the basis of much that is known regarding egg cortical structure, are lined with a layer of cortical granules about 1 pm in diameter (Fig. 1). In Strungylucentrotus there are about 18,000 of these organelles per egg (Vacquier, 1981). They are manufactured by the Golgi complex and become closely associated with the plasma membrane during oocyte development (Anderson, 1968; 233

Copyright Q 1988 by Academic Press, Inc. All rights of reproduction in any form reserved.

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FM

FIG. 1. Diagrammatic representation of cortical changes in a fertilized sea urchin egg. ( A ) Cortex of an unfertilized egg depicting cortical granules, plasma membrane (PM), vitelline layer (VL). and short microvilli (MV). The cortical endoplasmic reticulum, normally present in association with the plasma membrane. is not depicted. CGM, cortical granule membrane. (B) Cortical granule discharge and vitelline layer elevation. (C and D) A portion of the cortical granule contents has joined with the vitelline layer to form the fertili~ationmembrane (FMj. The remaining cortical granule material remains in the perivitelline space to become the hyaline layer (HL). Immediatly following cortical granule discharge, portions of the plasma membrane become involved in endocytosis as evidenced by endocytotic pits and vesicles (EP and EV). The surface of the fertilized egg is projected into elongate microvilli (EMV) containing a core of actin filaments (AF). Taken from Long (1987a).

Szollosi, 1967). The cortices of some ophiuroid, anthozoa, and polychaete eggs contain a layer of cortical granules five- to six-deep (Pasteels, 1966; Fallon and Austin, 1967; Dewel and Clarke, 1974; Holland, 1979). Eggs of the ascidian Ciona and the salamander Triturus do not have cortical granules (Hope et al., 1963; Rosati el al., 1977; see Elinson, 1986). In Ciona, a population of granules located in the subcortex reportedly functions as cortical granules (Rosati ei al., 1977). Ultrastructurally, cortical granules of sea urchin eggs display variations

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in organization depending on the species. The observed patterns reflect the organization of its components that localize to specific portions of the extracellular coats that form following cortical granule dehiscence (Anderson, 1968; Schuel, 1985). The contents of Arbaciu cortical granules are distinguished by a crenated central mass surrounded by some lenticular material. In Strongfyocentrotus there is a spiral of electrondense material that is associated with an amorphous mass. the organization of cortical granule internal components of the mollusc Mytilus (Humphreys, 1967), and other organisms, has also been described (see Schuel, 1978). The cortical granules in amphibian and mammalian eggs do got show unusually complex structural patterns and are filled with electron-dense granular material (Kemp and Istock, 1967; Gulyas, 1980). Cortical granule contents from sea urchins, amphibians, and mammals have been examined directly by biochemical and cytochemical techniques and indirectly by analysis of the medium following their discharge (Schuel, 1978, 1985). Calcium, serine protease, and sulfated mucopolysaccharides appear to be universal components of these structures. Peroxidase, pl,3-glucanase, hyaline protein, p-glucuronidase, and other proteins are also present in the cortical granules of some organisms. Although the structure of the sea urchin egg cortex has been analyzed by a number of techniques, the manner in which the cortical granules are associated with the plasma membrane remains an enigma (Millonig, 1969; Detering e? al., 1977; Longo, 1981, 1985). Electron microscopy of conventionally prepared eggs shows the cytoplasmic region associated with the cortical granules and plasma membrane as relatively unspecialized, i.e., lacking any apparent modification that might serve to attach the two structures (Anderson, 1968; Millonig, 1969). However, in quickfrozen and freeze-substituted preparations, filaments join cortical granules to the plasma membrane and extend from the granules into the cytoplasm (Chandler, 1984a). These filaments may contribute to the attachment of the cortical granules to the plasma membrane. The connection of cortical granules to the oolemma is sufficiently strong to survive forces encountered during the isolation of plasma membrane-cortical granule complexes (Detering et al., 1977) and preparation of “cortical lawns” in which the cytoplasm of the cortices is sheared away from the plasma membrane and cortical granules (Vacquier, 1975). The normal attachment of cortical granules to the overlying plasma membrane can be disrupted by chaotropic agents, urethane, and tertiary amines, reinforcing the idea that a special attachment exists between the two structures (Longo and Anderson, 1970; Vacquier, 1975; Hylander and Summers, 1981; Decker and Kinsey, 1983). Modifications of the sea urchin egg plasma membrane have been

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observed in areas occupied by cortical granules with freeze-fracture replicas, by scanning electron microscopy, and with filipin staining for the demonstration of 3P-hydroxysterol components (Longo, 1981; Carron and Longo, 1983; Zimmerberg et af., 1985). The plasma membrane modifications seen with freeze-fracture replication are dome-shaped areas lacking intramembranous particles. These modifications appear to form as a result of the association of the cortical granule with the plasmalemma, i.e., they are lacking in the plasma membrane of fertilized and immature eggs in which the cortical granules are absent or not localized to the cortex (Longo, 1981). Furthermore, the dome-shaped areas disappear when the mature egg is treated with amines (Longo, 1981). These specializations are believed to represent specific contacts between the plasma membrane and the cortical granule that produce or stimulate conditions required for bilayer fusion. Unlike the situation that exists in most other cells, in Xenopus oocytes the E face of the plasma membrane is endowed with a higher concentration of intramembranous particles than the P face (Bluemink and Tertoolen, 1978). Although the overall density of intramembranous particles is the same in the animal hemisphere and vegetal hemisphere, there is a preponderance of small intramembranous particles in the animal hemisphere that Bluemink and Tertoolen (1978) suggested might be involved with cortical morphogenesis following insemination. Freezefracture replicas of the plasma membrane of mammalian eggs have also been examined and a lower intramembranous particle density has been noted at those sites of presumptive cortical granule exocytosis (Koehler et al., 1982, 1985; Suzuki and Yanagimachi, 1983). Unique patterning of intramembranous particles possibly induced by structures subjacent to the plasma membrane has been described in numerous cells having secretory activities (Satir et al., 1973; Friend and Fawcett, 1974; Beisson et al., 1976; Satir, 1976; Weiss et al., 1977a,b; Kinsey and Koehler, 1978). In addition, clearings of intramembranous particles have been observed in portions of the plasma membrane associated with secretory vesicles and are generally considered to represent areas depleted of membrane proteins at the fusion zone (Chi et al., 1975; Lawson et af., 1977; Orci et af., 1977; Amherdt et af., 1978; Swift and Murkhejee, 1978; Theodosis et a)., 1978). The plasma membrane of sea urchin eggs is reflected into short microvilli that lack a core of actin microfilaments (Carron and Longo, 1982; Fig. 1). The underlying cortical granules tend to be situated in areas that lack microvilli (Schroeder, 1979; Longo, 1981). In amphibian and mammalian eggs the microvilli are relatively longer and contain a microfilamentous core (Phillips et al., 1985; Longo and Chen, 1985). Attached to the sea urchin oolemma is a glycocalyx, or vitelline layer

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(Anderson, 1968; Kidd, 1978). It is this structure to which sperm adhere via the protein bindin (Vacquier and Moy, 1977) and that, at the time of cortical granule exocytosis, becomes detached from the egg surface to form the fertilization membrane (see Kay and Shapiro, 1985). In many organisms, particularly sea urchins, the structure of the cortex is virtually identical along all regions of the egg surface. However, in eggs that are fertilized at an earlier stage of meiotic maturation (e.g., molluscs, annelids, and vertebrates), the meiotic spindle is positioned within the cortex and the cytoplasm associated with this structure usually differs in composition as compared to the cytoplasm in other regions of the egg cortex. For example, in mammalian eggs, the region that overlies the meiotic spindle is distinguished by the absence of microvilli and cortical granules, a diminished affinity to the plant lectin concanavalin A, and the presence of a dense layer of actin filaments (Johnson et al., 1975; Eager et al., 1976; Nicosia er af., 1977, 1978; Phillips and Shalgi, 1980; Albertini, 1984; Ebensperger and Barros, 1984; Mar0 et al., 1984, 1986a,b; Longo and Chen, 1984, 1985; Karasiewiez and Soltynska, 1985; Van Blerkom and Bell, 1986; Capco and McGaughey, 1986). This region has been referred to as the microvillus-free area (Longo and Chen, 1985). When the meiotic spindle develops in mammalian oocytes it moves to the animal pole. With the arrival of the metaphase spindle in the cortex, a cortical granule-free area appears as a result of the peripheral migration and exocytosis of cortical granules (Okada et al., 1986). In addition, the formation of a thickened layer of actin filaments and the disappearance of microvilli (Longo, 1985; Van Blerkom and Bell, 1986) in the cortex associated with the spindle, suggests that meiotic spindle localization and cortical modifications are linked. Experimental support for this contention has been presented (Longo and Chen, 1985; Van Blerkom and Bell, 1986; Mar0 et al., 1986b; Longo, 1987b). The concurrent events of actin accumulation at the site occupied by the meiotic spindle and actin loss from other regions of the cortex are consistent with a translocation of molecules from one part of the cell to another (Longo, 1987b). Similar transformations in cortical actin distribution have been described for other cell types. For example, during cap formation in Dictyostelium,actin, which was formerly distributed along the cell periphery, becomes concentrated at one locus on the cell surface (Condeelis, 1979). This rearrangement is believed to be a result of a sliding of actin filaments that are bound to membrane receptors, thereby reducing the percentage of protoplasmic surface of the plasma membrane that is associated with actin. A sliding of actin filaments may also occur during oocyte maturation to generate its polar localization, although other possibilities have not been ruled out. Differences in the anchorage of actin filaments via an electron-dense

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material to the plasma membrane of mature mouse eggs and immature oocytes have been identified that are similar to those observed in numerous cell types (Longo, 1987a). In some instances, the electrondense materials have been characterized as specific aetin-binding proteins (Weeds, 1982: Burridge and Feramisco, 1982; Jockusch and Isenberg, 1982: see Geiger, 1983). Changes in the affinity of the actin-binding protein vinculin to the plasma membrane have been proposed to initiate the formation of microfilament bundles in lamellipodia of cultured cells (Geiger, 1983). Comparable alterations in the association of actin-binding proteins with the plasma membrane, leading to the assembly or disassembly of actin filaments at specific loci within the cortex, as well as modifications in actin interactions (Lin et nl., 1982), might also be involved in cortical actin changes characteristic of maturing mouse oocy tes . Although a meiotic spindle is not formed in mouse oocytes treated with colchicine, the chromosomes move to the periphery of the oocyte and a microvillus-free area forms in the region of the cortex associated with the chromo5omes (Longo and Chen, 1985). Moreover, when the meiotic chromosomes are prevented from moving to the oocyte cortex, a microvillus-free area does not develop (Longo and Chen, 1984, 1985: Van Blerkom and Bell, 1986; see also Mar0 er al., 1986a,b). Chromosomes transferred to germinal vesicle-intact or maturing oocytes are also capable of inducing a localized thickening of actin, the loss of microvilli, and reduction of surface glycoproteins (Van Blerkom and Bell, 1986). These observations show that interaction of the meiotic chromosomes with the oocyte cortex brings about the formation of the microvillus-free area and that the capacity for this transformation occurs prior to germinal vesicle breakdown. Moreover, the cortical and plasma membrane polarities that are established at metaphase I and 11 in mouse oocytes are induced rather than preexisting; the entire surface of the oocyte is capable of differentiation in response to the presence of chromosomes (Van Blerkom and Bell, 1986). Subsequent to its migration to the oocyte cortex, the meiotic spingle becomes anchored to the plasma membrane (Chambers, 1917; Conklin, 1917: ShimiLu, 198la). In TirbifLx the meiotic apparatus appears to be tethered to the oocyte surface by structural connections between a filamentous cortical layer, possibly actin, and microtubules of the peripheral aster (Shimizu, 1981a). A similar morphology is also seen in I l y a n u ~ s noocytes (Burgess, 1977). Interestingly, meiotic spindle attachment in Chuetopterus oocytes is colchicine sensitive and unaffected by cytochalasin B (Hamaguchi et al., 1983). In contrast to observations with the eggs of come invertebrates (Longo, 1972; Peaucellier et al., 1974),

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cytochalasin B prevents the cortical localization of the meiotic spindle in maturing mouse oocytes, suggesting that a cytochalasin B-sensitive component of the cytoskeletal system is involved in this movement (Wassarman e? a/., 1976; Longo and Chen, 1985). The roles that microtubules, microfilaments, and possibly other cytoskeletal structures may serve in cortical localization and attachment of the meiotic spindle to the oocyte surface warrants further investigation. With respect to studies in mammals, actin has been demonstrated in nuclei and in mitotic spindles, and has been implicated in force production of chromosome movements during mitosis (Zimmerman and Forer, 1981). In light of these investigations and studies demonstrating the disruptive effects of cytochalasin B on actin (Yahara e? a / . , 1982; Schliwa, 1981), an actin-based system may be responsible for the cortical localization of the meiotic spindle in mouse oocytes (Longo and Chen, 1984, 1985). The association of actin with the meiotic chromosomes in mature mammalian eggs may serve a number of functions, such as to insure the presence of a contractile mechanism necessary for polar body formation (Nicosia et al., 1977; Mar0 et al., 1984, 1986b), to maintain chromosome condensation (Rungger et al., 1979), as well as to insure the peripheral location of the meiotic spindle (Webb et al., 1986; Longo, 1987b). Changes in the distribution of cortical actin during meiotic maturation may also be associated with alterations in surface or membrane-bound components that are linked to the underlying actin network and that become redistributed during meiotic maturation.

111. Interaction and Fusion of Sperm and Egg A. THECHRONOLOGY OF EARLY EVENTS The attachment of the sperm and egg ultimately results in their fusion. In sea urchins, sperm binding to the egg occurs via a hydrophobic protein that coats the acrosomal process, bindin (Vacquier and Moy, 1977). Bindin from sea urchin sperm has been shown to promote fusion of artificial lipid vesicles (Glabe, 1985), and a similar protein from Urechis sperm agglutinates eggs and initiates development (Gould et al., 1986). These results are in agreement with recent studies examining the time at which gamete fusion actually occurs (Hinkley et aE., 1986; Longo et af., 1986). Correlative ultrastructural and electrophysiological observations have been carried out examining sperm-egg interactions in Lytechinus in which eggs voltage clamped at -20 mV were fixed 1-20 seconds after the onset of the conductance increase caused by a single sperm. Continuity

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between the sperm and egg plasma membranes is first detected 5 seconds after the onset of the conductance increase (the earliest known physiological response of the egg following sperm attachment). The earliest stages in the formation of fertilization cones coincide with the establishment of the continuity of the gamete plasma membranes. One possible explanation for the increase in conductance of the egg at fertilization would be the inclusion of channels from the plasma membrane of the acrosome-reacted sperm as the gamete plasma membranes fuse. Measurements of Schackman et ul. (1984) indicate that the membrane conductance of sperm after the acrosome reaction has occurred may be high, since the sperm plasma membrane depolarizes and becomes highly permeable to calcium ions with the acrosome reaction. Consequently, when the gamete plasma membranes fuse, a patch of sperm membrane of high conductance would be inserted in parallel with the egg membrane of low conductance (Chambers and de Armendi, 1979). However, the conductance increase occurs 5 seconds before electron microscopic evidence of gamete membrane continuity can be detected. Consequently, an alternative explanation is that the conductance increase results from the opening of channels in the egg plasma membrane in response to a component released from or on the sperm surface, rather than as a result of sperm-egg fusion. This component could be in the sperm surface membrane or from the acrosomal vesicle (e.g., bindin). Cortical granule discharge is initiated following sperm-egg plasma membrane fusion in the immediate vicinity at which continuity of the gamete plasma membranes occurs 6-8 seconds after the initial membrane conductance increase (Longo et al., 1986). This ultrastructural observation is consistent with capacitance measurements on whole Lytechinus eggs in which an increase in membrane capacitance, inferred to have been due to an increase in surface area from the integration of cortical granule membrane with the plasma membrane, could first be detected (McCulloh, 1985; McCulloh and Chambers, 1985).

B. INTEGRATION OF THE SPERMAND EGGPLASMA MEMBRANES It is assumed in many instances that all of the sperm plasma membrane is incorporated into the egg plasma membrane at fertilization, although experimental evidence has not verified this unequivocally. Electron microscopic studies of some invertebrates and mammals have demonstrated internalized membranous elements at the site of gamete fusion that appear to be derived from the fused sperm and/or the egg plasrnalemma (Franklin, 1965; Colwin and Colwin, 1967; Piko, 1969; Zamboni, 1971; Bedford and Cooper, 1978). Insertion of sperm plasma membrane components into the egg plasma membrane has been demonstrated by

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24 1

O’Rand (1977) who used isoantiserum against whole rabbit sperm. Fertilized rabbit eggs lyse when incubated with the antibody and compliment. Unfertilized eggs are unaffected suggesting that sperm antigen is exposed on the surface of the fertilized egg. Electron microscopic investigations examining the integration of sperm and egg plasma membranes at fertilization, in which one of the gametes has been labeled, have been carried out in both invertebrates and mammals (Yanagimachi et al., 1973; Gabel ef al., 1979; Longo, 1982). Prior to sperm-egg fusion in hamsters, the sperm plasma membrane of the postacrosomal region does not bind colloidal iron hydroxide. Once gamete fusion has been initiated, however, the former sperm plasma membrane is able to bind this marker (Yanagimachi et al., 1973). The rapid increase in colloidal iron hydroxide binding on the incorporating sperm head is believed to be a result of intermixing of sperm-egg membrane components comparable to the intermingling of antigenic determinants after fusion of somatic cells (Frye and Edidin, 1970). These observations, however, do not exclude the possibility that colloidal iron hydroxide-binding receptors are enzymatically added to sperm plasma membrane oligosaccharides after fusion or that colloidal iron hydroxidebinding membrane components are inserted into the sperm plasma membrane following fertilization. Similar experiments have been carried out with the surf clam Spisula (Longo, 1982) in which concanavalin A binding to the egg but not to the sperm plasma membrane has been demonstrated by the horseradish peroxidase-diaminobenzidine reaction (HRP-DAB). Because of this dichotomy in lectin binding, possible changes in the affinity of the sperm plasmalemma following its fusion and integration with components of the egg plasma membrane can be followed. By 1 minute postinsemination, the plasma membranes of fertilized Spisula eggs incubated with concanavalin A-HRP-DAB are uniformly associated with enzymatic precipitate except at sites of sperm incorporation. These portions of unstained plasma membrane are derived from the sperm and are localized to the apex of the fertilization cone. From 2 to 4 minutes postinsemination, the HRP-DAB reaction product gradually becomes associated with all of the membrane delimiting the fertilization cone. By 4 minutes postinsemination, no difference in staining of plasma membranes derived from the egg or the sperm is detected. These observations are consistent with the movement of concanavalin A-binding sites from the egg plasmalemma into the sperm plasma membrane. Similar results have also been obtained using cationized ferritin-labeled gametes (Longo, 1986a) or monoclonal antibodies to sperm-specific surface components (Nishioka et al., 1987; Figs. 2 and 3). These studies provide evidence for the free lateral mobility of sperm membrane

FIGS.1 and 3. Sea urchin (Arbacia pitnctulora) eggs and sperm fixed at different intervals following insemination (Fig. 2. 30 seconds postinsemination; Fig. 3, 5 minutes postinsemination). Following fixation. the samples were washed and incubated with a monoclonal antibody that recognizes sperm surface antigens, followed by goat anti-mouse antibody conjugated to colloidal gold. The reacted specimens were then dehydrated and embedded for electron microscopy. The monoclonal antibody shows exclusive reaction to 242

sperm surface antigens (Fig. 2) that migrate as M,33,000 and 35,000 polypeptide bands in Western blots. Following gamete fusion, colloidal gold (sperm surface antigen) is observed in portions of the egg plasma membrane distant from the site of sperm entry (arrow, Fig. 3) indicating a mixing of egg and sperm plasma membrane components. The fertilization cone (FC) and portions of the adjacent fertilized egg surface show incorporation and diffusion of the sperm surface antigen. MV, microvilli; N, sperm nucleus; M, sperm mitochondria; T, sperm tail. 243

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components in the fertilized egg and demonstrate that significant rearrangements occur in the egg and sperm plasma membranes following gamete fusion giving rise to transient asymmetries in membrane topography. Components of both the sperm and egg plasma membranes become redistributed within the bilayer adjacent to and delimiting the fertilization cone. The relationship between the movement of sperm and egg surface components and chemical and physical changes of the egg cortex is not entirely clear. The sea urchin egg cortex becomes rigid at fertilization (Mitchison and Swann, 1955; Hiramoto, 1974) as a result of dramatic cytoskeletal changes; the possible relation of this alteration to changes in plasma membrane properties has not been demonstrated. Apparently not all components of the sperm and egg plasma membranes intermix rapidly following gamete fusion. Sea urchin and mouse eggs fertilized with fluorescent or '251-labeledsperm reportedly retain a topographically mosaic surface during cleavage, as if the lateral mobility of sperm plasma membrane components was restricted and the membrane was retained as a discrete patch (Gabel et af., 1979; Shapiro et af., 1982). In this regard, it is remarkable that some incorporated sperm surface proteins persist within a localized area, despite rearrangements of the egg plasma membrane involving exocytosis of cortical granules and endocytosis. Similar experiments have also indicated that labeled sperm components (surface and mitochondrial) are internalized after fertilization (Gundersen ef al., 1982); some proteins persist intact whereas others undergo a specific, limited degradation (Gundersen and Shapiro, 1984; Gundersen et al., 1986). These results are consistent with electron microscopic studies indicating the incorporation of portions of the sperm plasma membrane into the egg cytoplasm (Colwin and Colwin, 1967; Bedford, 1972; Bedford and Cooper, 1978; Yanagimachi and Noda, 1970).

C. FERTILIZATION CONEFORMATION At the site of gamete fusion in many organisms a protuberance forms, the fertilization or incorporation cone, through which the contents of the sperm pass (Longo and Anderson, 1968; Longo, 1973; Schatten and Schatten, 1980; Tihey and Jaffe, 1980). Formation of this structure involves a movement of egg cytoplasm into the region surrounding the sperm nucleus, mitochondria, and axonemal complex, resulting in a protrusion at the site of sperm entry (Longo, 1973). The fertilization cone increases in size as more of the egg cytoplasm surrounds incorporated sperm components, which in turn move deeper into the ovum cortex. Based on scanning electron microscopic observations, it has been claimed that microvilli in the region of gamete fusion cluster and engulf the

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spermatozoon (Cline et af., 1983; Schatten and Schatten, 1980). Although elongation of the microvilli is first recognized in the vicinity of gamete fusion, their involvement in sperm incorporation was not apparent by transmission electron microscopy (Longo, 1980). During early stages of sperm incorporation, the fertilization cone is relatively small; it increases greatly in size after the sperm nucleus passes through it and comes to rest within the egg cortex (Longo, 1980; Schatten and Mazia, 1976). The maximum size of fertilization cones varies depending upon the organism in question; in mature Arbacia eggs they measure approximately 6 pm in length by 4 pm in diameter 5-7 minutes postinsemination. They then regress and are reabsorbed by 20 minutes postinsemination. Interestingly, in Arbaciu, the fertilization cones that form on immature eggs are much larger than those that develop on mature ova, e.g., sizes of 25 pm in length by 10 pm in diameter are not unusual (Seifriz, 1926; Tilney and Jaffe, 1980; Dale and Santella, 1985). Surface area measurements of sea urchin oocyte fertilization cones indicate that considerably more membrane delimits this cortical projection of cytoplasm than can be accomodated by the sperm plasma membrane (Longo, 1986a). That is, if all of the sperm plasma membrane becomes a part of the egg plasma membrane, it would comprise less than 10% of the surface area of the fertilization cone. This and the absence of evidence demonstrating a contribution from the spermatozoon other than its plasmalemma indicate that most of the membrane (90%+) delimiting the fertilization cone is derived from the oocyte. Qualitative assessment of oocytes at different times following insemination with different degrees of polyspermy indicate that the presence of microvilli is inversely related to the number and the size of fertilization cones. This suggests that microvilli are retracted into the oocyte surface; the membrane thus produced accommodates fertilization cone expansion (Dale and Santella, 1985; Longo, 1986a). These conclusions are consistent with observations in somatic cells indicating that surface material stored in microextensions of suspended cultured cells is used during spreading (see Trinkaus, 1980; Erickson and Trinkaus, 1976; Rovensky and Vasiliev, 1984). It is uncertain how this might be accomplished, but it would presumably involve a modification of the microvillous cytoskeletal core and the migration of membrane components to the base of the microvilli. The membrane area associated with the microvilli is spread smooth as the supporting cytoskeleton is disrupted; it may then move laterally to the site of the forming fertilization cone (Fig. 4). Recent studies (Longo, 1986a,b) have also demonstrated that the fertilization cones of inseminated sea urchin oocytes have a distinctive crenulated appearance that differs from that of the remainder of the

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Egg'+ Sperm

Egg + Sperm'

Egg + Sperm

+ Gationized Ferritin (GF) P

FIG. 4. Diagrammatic representations, based on results of electron microscopic observations of reciprocally cationized ferritin-labeled sea urchin sperm and oocytes, indicating migrations of sperm and oocyte plasma membrane components at the site of gamete I'usion. (A) Expansion of the oocyte plasma membrane (large arrows) selects for components lacking anionic residues. which contribute to greater than 9Wc of the surface area of the fertilization cone. A source of membrane that is involved in the development of the fertilization cone is derived from the regression of microvilli (small arrows). (B) Cationized ferritin receptors move laterally from the sperm plasma membrane (arrows) into the fertilization cone and microvilli and intermix with oocyte plasma membrane components. (C) The labeling pattern that develops when unlabeled gametes are mixed simultaneously with cationized ferritin is a mix of what is shown in (A) and (B). The membrane delimiting the fertilization cone is lightly labeled whereas the remainder of the oocyte surface is associated with a denser layer of cationized ferritin. Taken from Longo (1986a).

oocyte surface. Membranes delimiting fertilization cones also have a much lower affinity for agents that stain negatively charged and carbohydrate moieties. This difference in surface properties of membranes delimiting the site of sperm-egg fusion is not due solely to the incorporated sperm plasma membrane and does not occur when inseminated oocytes are incubated in cytochalasin B . These observations indicate that following insemination significant rearrangements of surface molecules take place within the egg plasmalemma that give rise to asymmetries in membrane topography. The development of membrane asymmetry of inseminated sea urchin oocytes is comparable to that observed in other cell types. The plasma membrane of epithelial cells shows a striking

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polarization into two morphologically and functionally distinct domains (Rodriquez-Boulan, 1983). It has been well established that many of the surface proteins of spermatoza are restricted to specific domains (Friend, 1982; Primakoff and Myles, 1983). Experiments by Myles and Primakoff (1984) have demonstrated that some of these proteins migrate to new domains either before or during sperm-zona binding. Redistribution of cell surface molecules into or out of specific domains is also evident during capping (Oliver and Berlin, 1982) and endocytosis (Pastan and Willingham, 1981). The possible involvement of oocyte microvilli in fertilization cone expansion implies that the asymmetry in membrane topography may occur as a result of (1) the migration of specific components from the egg plasma membrane into the domain of the fertilization cone or (2) the modification of a more general pool of plasma membrane components as they become a part of the fertilization cone. The latter would involve significant alterations in the charge and carbohydrate environment of egg plasma membrane constituents. Results of fertilization experiments employing reciprocally cationized ferritin-labeled sea urchin gametes are summarized in Fig. 4. It is possible that fertilization cone formation may be related to localized changes in surface properties of the oocyte that are brought about by sperm fusion. McCloskey and Po0 (1984) postulated that components of the cell coat could be excluded or lost from experimentally induced blebs on various cell types. Sugrue and Hay (1981) showed that corneal epithelial cells generate membrane blebs when stripped of extracellular matrix. Although extramembranous constraints are important to the lateral mobility of membrane components, it is possible that lateral motion of integral membrane components is also modulated by structural parameters of the bilayer or by direct interactions with other integral components (see Oliver and Berlin, 1982). It has not been determined how movements of sperm and egg surface components may be modulated during fertilization cone formation. The mobility of the plasma membrane in other cells has been shown to be regulated to some extent by underlying cytoskeletal elements (Elgsaeter and Branton, 1974; Edelman, 1976; Sheetz et al., 1980; Tank et al., 1982; Wu et al., 1982; Jacobson et al., 1984). The findings that the fertilization cone is filled with bundles of actin filaments and fails to form when eggs are incubated in cytochalasin B are consistent with the idea that the cortical cytoskeleton may be involved in the lateral migration of spermand egg-derived surface components (Longo, 1981, 1986a,b; Tilney and Jaffe, 1980). The migration of sperm- and egg-derived components within the plasma membrane of the zygote is also coincident with dramatic

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changes in the physiological state of the egg, which in turn could be a result of functional alterations of the membrane. The movement of surface molecules may be involved in new membrane functions similar, for example, to the activation of adenylate cyclase resulting from the lateral mobility of membrane molecules (Martin, 1983). In sea urchin eggs, fertilization cones are filled with numerous bundles of actin filaments that show the same polarity when reacted with heavy meromyosin or S1 (Tilney and Jaffe, 1980), i.e.. the arrowhead complexes that form are directed to the center of the egg. The microfilaments found in the fertilization cone are polymerized in situ from cortical monomeric actin as few actin filaments are present in the egg cortex (Spudich and Spudich, 1979; Cline et al., 1983; Cline and Schatten, 1986). In addition to actin, fodrin, a spectrin-like protein, has also been identified in sea urchin fertilization cones (Schatten et ul., 1986a). Spectrin has been shown to interact with actin oligomers to form gels that are associated with the plasma membrane (see Branton et al., 1981). The role that fodrin may play during fertilization is not yet clear, however, Schatten et uf. (1986a) suggest that the formation of an actin-spectrin gel at the site of spermegg membrane fusion could serve to anchor the sperm to the egg surface and may account for the required polymerization of actin for successful sperm incorporation. Whether and how actin in the fertilization cone might function to effect sperm incorporation are not entirely clear. Actin filaments of the sperm acrosomal process are also polarized with the heavy meromyosin-actin arrowheads pointing to the sperm nucleus. Consequently, egg myosin could not bridge sliding actin filaments of both the fertilization cone and the acrosomal process to bring about sperm nucleus incorporation (Tilney and Kallenbach. 1979); both sets of actin filaments are polarized in the wrong direction when compared to the orientation of myosin and actin of a sarcomere. It is possible that actin filaments present in the fertilization cone might be primarily involved in the elevation and enlargement of this cortical projection of cytoplasm. Agents that disrupt actin microfilaments inhibit surface activity of fertilized sea urchins eggs, including microvillar elongation and fertilization cone formation (Longo, 1980; Cline and Schatten, 1986; Schatten ef a / . , 1986b). Cytochalasin B-treated eggs undergo a cortical granule reaction, elevate a fertilization membrane, and are metabolically activated (Gould-Somero et al., 1977; Longo, 1978). These observations are consistent with the suggestion that at the site of gamete fusion there is a localized polymerization of actin that participates in the formation of the fertilization cone. In addition, cytochalasin B-treated sea urchin eggs can be activated by sperm, but sperm fail to enter the egg (Gould-Somero et

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al., 1977; Longo, 1978). How the sperm is capable of activating the egg in this instance without entering it has not been determined. It is possible that the acrosomal process fuses with the egg plasma membrane, but, since actin polymerization is impaired, the bridge linking the fused gametes is weak and the sperm is removed from the egg surface by exocytosing cortical granules. Another possibility, consistent with recent observations that ionic activation of Lytechinus eggs precedes sperm-egg plasma membrane fusion (Longo et af., 1986), is that cytochalasin B inhibits fusion of the egg and sperm and that gamete contact/binding in this instance is sufficient for egg activation. In teleosts, the fertilizing spermatozoon enters a region of the egg cytoplasm that is highly specialized. At the base of the micropyle, the plasma membrane of the unfertilized egg is differentiated into a structure apparently designed for sperm binding. It is a short projection in Fundulus (Brummett and Demont, 1979) and a cluster of microvilli in Cyrinus (Kudo, 1980), Rhodeus (Ohta and Iwamatsu, 19831, and Brachydanio (Hart and Donovan, 1983). In the eggs of some teleosts the site of sperm incorporation may lack cortical granules (Brummett and Dumont, 1979; Kobayashi and Yamamoto, 1981; Hart and Donovan, 1983) or granules may be present but smaller than those in other regions of the cortex (Gilkey, 1983). Changes in the teleost egg cortex associated with fertilization cone formation, as determined by scanning electron microscopy, have been described (Iwamatsu and Ohta, 1978; Ohta and Iwamatsu, 1983; Kudo and Sato, 1985; Kudo, 1980; Brummett et al., 1985). At the site of sperm entry in anurans a microvillus-free bleb of cytoplasm forms, presumably functionally equivalent to a fertilization cone (Picheral, 1977; Elinson and Manes, 1978; Picheral and Charbonneau, 1982). Eventually it disappears and is replaced by a small clump of elongate microvilli. The microvillus-free bleb is believed to be pinched off in Pleurodeles leaving the microvilli on the surface of the fertilized egg (Picheral, 1977; Picheral and Charbonneau, 1982). If this is the case, it is possible that plasma membrane components, as well as other spermderived structures, are eliminated from the egg. The site of sperm entry reportedly remains detectable as a clump of microvilli for at least 2 hours. Following the fusion of the egg and sperm plasma membranes in mammals, tongues of cytoplasm surround the anterior portion of the sperm head (Piko, 1969) forming a vesicle that is present for a time within the zygote (Yanagimachi and Noda, 1970). At the site of gamete fusion, a protrusion of cytoplasm forms that is homologous to the fertilization cones seen in invertebrate eggs and is often referred to as an incorporation cone (Shalgi et a f . , 1978; Zamboni, 1971; for reviews, see GaddumRosse, 1985; Mar0 et al., 1986a). As in sea urchins, the incorporation

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cone of fertilized mammalian eggs reaches its maximum only after the sperm head has entered the cortex (Gaddum-Rosse, 1985). In mouse eggs this protrusion is filled with cytoplasmic organelles found in other regions of the zygote, and along its plasmalemma is a prominent layer of actin (Maro et al., 1984, 1986a). Surface movements also occur in fertilized rat and mouse oocytes (Gaddum-Rosse ef al., 1984; Battaglia and GaddumRosse, 1984; Waksmundzka et al., 1984). This transient activity involves localized elevations of the cortical cytoplasm that disappear following incorporation of the sperm tail. Formation of cortical elevations is sensitive to cytochalasin B and occurs when eggs are activated with the calcium ionophore A23187. These observations suggest that the elevations are a manifestation of cytoskeletal changes of the oocyte involving actin and are characteristic of the activation process and not dependent upon the presence of sperm components.

1V. Cortical Granule Reaction Exocytosis of cortical granules has been studied in eggs of invertebrates and vertebrates and appears to involve similar processes (Anderson, 1968; Elinson, 1980; Gulyas, 1980). In sea urchins, sperm-egg binding is followed by the dehiscence of cortical granules that underlie the plasmalemma and exocytosis spreads from the point of gamete contact in a wave to the opposite pole of the egg (Afzelius, 1956; Endo, 1961; Wolpert and Mercer, 1961; Anderson, 1968; Millonig, 1969; see also Holland, 1980; Longo er al., 1982, 1986; Fig. 1). In some pelecypods and annelids, cortical granules are present but do not undergo exocytosis o r a change at fertilization (Pasteels and de Harven, 1962; Rebhun, 1962; Humphreys, 1967). The fate of cortical granule contents and the development of extracellular layers surrounding activated eggs have been reviewed (Kay and Shapiro, 1985). The mechanisms by which the egg plasma and cortical granule membranes fuse, and the nature of the intermediates in this process are unclear. Using freeze-fracture replicas, Chandler and Heuser (1979) were unable to find intermediate stages of cortical granule membrane-plasma membrane fusion, suggesting that the fusion process is completed very rapidly. They indicated that a single pore is formed that increases in size to allow dehiscence of the cortical granule contents. Using transmission electron microscopy of sectioned specimens, Anderson (1968) and Millonig (1969) indicated that the opening of cortical granules may occur via multiple fusions between the cortical granule membrane and oolemma, and thereby a series of vesicles, composed of membrane derived

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from both the cortical granules and oolemma, are released to the perivitelline space. Chandler (1984b) has also observed vesicle formation along the lips of expanding pores of fused cortical granules in thin sections of quick-frozen eggs. He suggested that the vesicles may be a result of an increased lipid mobility and fusibility of membranes. If the vesicles at the site of cortical granule fusion are, in fact, real and not an artifact of tissue preparation, then all of the “original” egg plasmalemma and membrane delimiting the cortical granules may not be incorporated into the plasma membrane of the fertilizediactivated egg during cortical granule dehiscence. The current paradigm regarding the mechanism of cortical granule discharge is that calcium functions as an essential intracellular messenger (for reviews, see Shen, 1983; Jaffe, 1985; Whitaker and Steinhardt, 1985). The release of calcium from different cellular compartments has been demonstrated for sea urchin eggs (Zucker and Steinhardt, 1978), and the store involved with cortical granule discharge appears to be associated with the endoplasmic reticulum located within the egg cortex (Jaffe, 1985; Luttmer and Longo, 1985). Almost all calcium-binding ability of the unfertilized sea urchin egg is found in a large particle fraction (microsomes) isolated by differential centrifugation (Steinhardt and Epel, 1974). Consistent with this observation are demonstrations that preparations of vesicles derived from Xenupus eggs sequester calcium in an ATPdependent manner (Cartaud et al., 1984) and that calcium is associated with the plasma membrane and cortical granules as well as cortical endoplasmic reticulum of sea urchin eggs (Cardasis et al., 1978; Sardet, 1984). In addition, electron microscopic studies have demonstrated specialized regions of the egg endoplasmic reticulum that are associated with the cortical granules in Xenopus, sea urchin, and mouse ova (Campanella and Andreuccetti, 1977; Gardiner and Grey, 1983; Sardet, 1984; Luttmer and Longo, 1985). These observations, as well as (1) the striking morphological similarity of the plasma membrane-endoplasmic reticulum association observed in Xenopus, mouse, and sea urchin eggs to the transverse tubule-sarcoplasmic reticulum association of muscle cells (Endo, 1977; Gardiner and Grey,, 1983; Sardet, 1984; Luttmer and Longo, 1985) and (2) the temporal correlation of cortical endoplasmic reticulum development and the capacity of Xenopus eggs to propagate a wave of cortical granule exocytosis (Charbonneau and Grey, 1984; Campanella et al., 1984), suggest that the close association of the plasma membrane and cortical endoplasmic reticulum transduces the interaction of gametes into an intracellular calcium release that then triggers the cortical granule reaction and the activation of development. At the completion of the cortical granule reaction in the eggs of sea

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urchins, amphibians, and fish, virtually all of the cortical granules have been discharged. In mice, a substantial number of cortical granules (about 25% of the population) are exocytosed before sperm-egg fusion; the remainder are dehisced at fertilization (Nicosia et al., 1977). In Sabellaria, the cortical granule reaction is initiated when the eggs are spawned into sea water (Pasteels, 1965). In Wrechis, a subset of cortical granules is released at insemination; the remainder are discharged later with the elevation of the vitelline layer (Paul, 1975). For organisms whose eggs do not possess cortical granules nor undergo a cortical granule reaction, it is clear that cortical granule exocytosis is not a required feature of fertilization nor of egg activation and development. Furthermore, inhibition of the cortical granule reaction in sea urchins does not impair events of fertilization and cleavage (Longo and Anderson, 1970; Vacquier, 1975; Schmidt and Epel, 1983).

V. Plasma Membrane Changes Attending the Cortical Granule Reaction As a consequence of the cortical granule reaction, cortical granule contents are externalized that have profound structural and physiological effects on the egg (Shapiro and Eddy, 1980; Shapiro et ul., 1981; Kay and Shapiro, 1985). Insertion of cortical granule membrane into the egg plasma membrane is accompanied by dramatic structural changes of the cortex and oolemma. The resultant membrane of the fertilized egg has been referred to as a mosaic, indicating that it is derived from several sources, i.e., the egg plasma membrane, the cortical granule membrane, and the sperm plasmalemma (Colwin and Colwin, 1967; Anderson, 1968; Schroeder, 1979). Although not proven, it is believed that there is essentially a doubling of the surface area of the activated echinoid egg as a result of the cortical granule reaction, i.e., the sum of membranes delimiting all of the cortical granules within the egg is equivalent to the surface area of the egg plasma membrane and both sources of membranes are believed to be completely incorporated with the cortical granule reaction (Schroeder, 1979; Vacquier, 1981). It has not been established that, in fact, all of the membranes delimiting the cortical granules and the plasma membrane of the unfertilized egg become a part of the plasmalemma of the activated ovum. Electron microscopic investigations (Anderson, 1968; Millonig, 1969; Chandler, 1984b) indicate a vesiculation of membrane at sites of cortical granule-plasma membrane fusion. If real, this vesiculation, apparently involving both the plasma and cortical

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granule membranes, may have a pronounced effect on the composition and the surface area of the fertilized egg. Since many aspects of fertilization are membrane-mediated events leading to egg activation it is likely that a change in the plasma membrane is an obligatory step in cellular activation (Pardee e? al., 1974; Campisi and Scandella, 1978). Potential changes in the organization of membrane lipids following insemination have been studied and the results, as yet, are incomplete. Using electron spin resonance spectroscopy, Campisi and Scandella (1978, 1980a) demonstrated an increase in bulk membrane fluidity of sea urchin eggs after fertilization. However, because the spin label (fatty acid) was equilibrated among all subcellular membrane fractions, it could not be determined whether: (1) ovum activation is accompanied by a change in all of the cellular membranes to a more fluid state, or (2) more specialized membranes (such as the plasmalemma) entered a more fluid state and the probe was showing the average change of altered and unaltered membranes. Changes of membrane lipids accompanying activation are probably not a result of the cortical granule reaction as eggs partially activated by ammonia showed a similar effect. In experiments with cortical fractions it has been shown that the fluidity of the fertilized egg cortex is less than that of the unfertilized egg cortex (Campisi and Scandella, 1980b). Adding calcium to cortical fractions from unfertilized eggs resulted in a fluidity decrease in uitro. It has been suggested that this change may represent an alteration in membrane structure rather than a direct interaction of calcium with phospholipid groups (Campisi and Scandella, 1980b). Another approach to the question of possible organizational changes in egg plasma membrane lipids has been explored with the fluorescent probe, merocyanine 540 (Freidus ef al., 1984). These studies indicate that cortical granule fusion results in changes in plasma membrane lipid organization, i.e., membrane lipids become more loosely organized. Translational diffusion in the plasma membranes of sea urchin and mouse eggs using fluorescence lipid probes have been examined (Peters and Richter, 1981; Wolf ef al., 1981a,b; Wolf and Ziomek, 1983). The investigations of Peters and Richter (1981) indicated that protein diffusion is restricted more severely after fertilization than lipid diffusion. Since a decrease in membrane fluidity should act on lipid and protein diffusion in a comparable manner, the observations of Peters and Richter (1981) suggest that the fluidity decrease can not be the only parameter whereby diffusion in the egg plasma membrane is restricted after fertilization. They suggested that changes in the cortical cytoskeleton may regulate diffusion processes as in somatic cells (Peters ef al., 1974a,b; Fowler and Branton, 1977; Golan and Veatch, 1980; Sheetz ef al., 1980).

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Using fluorescence recovery after photobleaching, the lateral diffusion coefficient of lipid bilayer probes has been shown to either increase, decrease, o r remain relatively unchanged at fertilization, depending on the structure of the particular probe employed (Wolf et af.,1981a,b; Wolf and Ziomek, 1981). Based on studies of the partitioning of the probes in model mixed-phase membranes (Klausner and Wolf, 1980), these authors proposed the existence of gel and fluid lipid domains within the egg plasma membrane, the proportion and composition of which change upon fertilization. Changes in probe mobility, according to this view, are a reflection of alterations in the proportion and composition of these domains at fertilization. At fertilization there may be a reordering of lipid domains that releases inactive proteins from gel regions of the plasma membrane into fluid regions, in which they would become active. Changes in lipid composition and gel-fluid transformations at fertilization could activate protein functions not requiring the synthesis o r insertion of new materials into the membrane. Studies have been performed in sea urchin (Arbaciu) eggs treated with filipin to detect alterations in membrane sterols at activation (Carron and Longo, 1983). The plasma membranes of treated unfertilized eggs possess numerous filipin-sterol complexes whereas fewer complexes are associated with membranes delimiting cortical granules, indicating that the plasma membrane is relatively rich in /3-hydroxysterols (de Kruijff and Demel, 1974; Jain and Wagner, 1980; Carron and Longo, 1983). This dichotomy does not appear to be related to a filipin impermeability, and differences in filipin staining of the plasma and cortical granule membranes may represent differences in sterol content. Biochemical analysis (Decker and Kinsey, 1983), however, indicated that the cholesterol content of cortical granules is significantly higher than that of the egg plasma membrane, suggesting that following the cortical granule reaction there would be a substantial increase in plasmalemma cholesterol. Analyses of fertilized egg plasma membranes failed to confirm this expectation (Decker and Kinsey, 1983). Following its fusion with the plasmalemma, membrane formerly delimiting cortical granules undergoes a rapid increase in the number of filipin-sterol complexes (Carron and Longo, 1983). Other than regions involved in endocytosis, the plasma membrane of the zygote possesses a homogeneous distribution of filipin-sterol complexes and appears structurally similar to that of the unfertilized ovum. It has not been determined how the cortical granule membrane might acquire an increase in filipinsterol complexes. Lateral displacement of sterols from membranous regions derived from the original egg plasma membrane may be involved (Friend, 1982); however, there is no evidence documenting such a process

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in activated eggs. Sterols have been shown to diffuse rapidly in bilayers (Trauble and Sackermann, 1972), which is consistent with an extremely rapid lateral displacement of sterols into membrane patches derived from cortical granules (Carron and Longo, 1983). Furthermore, Peters and Richter (1980) have estimated that a mean displacement of 1 p m can be accomplished in about 1 second by membrane lipids and in about 35 seconds by membrane proteins of fertilized sea urchin (Paracentrotus) eggs. Hence, morphological observations and theoretical considerations indicate that the mosaic membrane is an intermediate that, by diffusion, may be rapidly transformed into a more homogeneous membrane. Fluorescence photobleaching recovery experiments have been performed with mouse eggs using probes for membrane proteins that suggest that interactions with cytoskeletal components may regulate membrane protein diffusion (Wolf and Ziomek, 1983). As with membrane lipids, the proteins probed demonstrated a heterogeneous distribution. Moreover, although “new” membranes (i.e., cortical granule and sperm plasma membranes) are added to the egg plasmalemma at fertilization, there is no generalized effect on the diffusion of membrane proteins in the mouse egg. Binding studies using plant lectins have also been utilized in an effort to demonstrate possible membrane changes eggs undergo at fertilization. Investigations with mouse and hamster eggs have shown that concanavalin A binding-sites change quantitatively following fertilization (Yanagimachi and Nicolson, 1976; DeFelici and Siracusa, 1981). In ascidian eggs both the agglutinability and number of concanavalin A receptors increase following activation (O’Dell et al., 1973). These changes in lectin binding following fertilization may reflect modifications in the nature and/or structure of the binding sites themselves. Alterations in lectin binding may also be influenced by membrane fluidity and functional states of the cytoskeleton (Karsenti et al., 1977; Marshall and Heiniger, 1979). In the sea urchin Strungylucentrutus, two classes of concanavalin A-binding sites have been identified: a high-affinity site associated with the vitelline layer and a low-affinity site associated with the plasma membrane. The number of low-affinity sites doubles at fertilization, possibly as a result of the insertion of cortical granule membrane (Veron and Shapiro, 1977). Although the increase in lowaffinity binding sites may be due to the appearance of cryptic sites, there is no doubling when eggs are activated with ammonia, supporting the notion that the increase in the number of sites is caused by the addition of cortical granule membrane to the egg plasmalemma. Examination of freeze-fracture replicas of unfertilized sea urchin eggs demonstrates a significant difference in the number of intramembranous

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particles within the plasmalemma and the cortical granule membrane. In Arbncia, the number of intramembranous particles within the P face of the cortical granule membrane is about 30% of that in the P face of the egg plasma membrane (Longo, 1981). Studies h a w been carried out to determine what happens to this dichotomy following cortical granule exocytosis, i.e., whether localized areas, corresponding to patches of cortical granule membrane, are present within the plasma membrane of the fertilized egg, or whether particles within the plasma membrane of the activated egg are homogeneously distributed (Fig. 5). A homogeneous distribution of particles would suggest an intermixing of components within the mosaic membrane. The mosaic pattern of the fertilized egg plasmalemma, in terms of intramembranous particles, is temporary; recognizable differences between the original egg plasma membrane and cortical granule membrane are lost soon after cortical granule exocytosis (Pollack, 1978; Chandler and Heuser, 1979; Longo, 1981). Patches, containing a reduced number of intramembranous particles and corresponding to the former cortical granule membrane, are not found in the

FIG. 5. Diagram depicting the possible fate of membrane components derived from cortical granules (large spheres). Membrane components may persist, in a time-dependent manner, as patches or disperse to varying extents among elements derived from the original egg plasma membrane (small spheres). Observations published thus far are consistent with a mixing of components derived from the egg plasmalemma within membrane domains originating from the cortical granule.

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plasma membrane of the activated egg. This indicates a rapid alteration in the composition of the cortical granule membrane following its fusion with the plasma membrane. By 4 minutes postinsemination, the density of intramembranous particles in the P face of the plasma membrane of the fertilized egg is slightly reduced from that of the membrane of the unfertilized egg, suggesting a possible “flow” of intramembranous particles from the oolemma into membrane derived from the cortical granules. This suggestion is in keeping with the fluid character of membranes and is consistent with schemes reported for other cells (Frye and Edidin, 1970; Singer and Nicolson, 1972). Changes in the distribution of intramembranous particles occur in the plasma membrane of Spisula eggs that do not have a cortical granule reaction. Following activation, there is an approximate 2-fold increase in the density of intramembranous particles within the plasma membrane of Spisula eggs (Longo, 1976a,b). The functional significance of this membrane change and whether or not it is related to the development of a block to polyspermy have not been determined. As a result of cortical granule fusion with the egg plasma membrane, components located on the inner surface of the cortical granule membrane become localized to the outer surface of the plasma membrane. This transformation has been implicated in the activation of transport systems for specific metabolites (Whitely and Chambers, 1960; Epel and Johnson, 1976). Cortical granule proteases have been shown to be involved in detaching the vitelline layer from the plasma membrane and the proteolytic removal of sperm receptors from the vitelline layer (Carroll and Epel, 1975; Carrol, 1976). A limited proteolysis of surface proteins by cortical granule proteases has also been reported in fertilized sea urchin eggs that is believed to play an, as yet undefined, role in activation (Shapiro, 1975).

VI. Microvillar Elongation It has been shown that the total surface area of cortical granule membranes in a Srrungylocentrutus egg is greater than that of the plasmalemma (Schroeder, 1979). Hence, if all the cortical granule membranes are incorporated into the egg plasmalemma, there would be at least a 2-fold increase in surface area of the egg at fertilization. However, by 16 minutes postinsemination, the surface area of the activated egg is only slightly larger than that of the unactivated ovum, indicating a rapid accommodation in surface membrane. The microvillar elongation that

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occurs following insemination may be one means of accommodating a surface increase in the activated sea urchin egg (Eddy and Shapiro, 1976; Schroeder, 1979; Fig. 1). However, surface area measurements indicate that elongated mirovilli cannot compensate for all the cortical granule membrane that might be incorporated, and membrane internalization has been proposed as a mechanism to quantitatively modify the surface area of activated eggs (Schroeder, 1979). Rapid elongation of microvilli is believed to occur primarily in areas occupied by the original plasma membrane (Chandler and Heuser, 1981), and may take place only at sites on the egg surface in which cortical granules have exocytosed (Fisher et al., 1982). However, more recent investigations (Fisher et al., 1985) indicate that it may occur in sea urchin eggs in which cortical granule exocytosis has been inhibited, but after some delay. By 2 minutes postinsemination in sea urchin eggs, cytoplasmic upheavals develop at the bases of elongating microvilli and form mounds that possess two to four microvilli projecting from their apices. By 5 minutes postinsemination, the mounds shrink and are interconnected by ruffles of cytoplasm (Chandler and Heuser, 1981). Similar morphological changes along the bases of microvilli have also been reported for Spisula eggs (Longo, 1976a). In sea urchins, these changes are a result of the reorganization of the cortical cytoskeletal system that gives rise to a cortex that projects into elongate microvilli and contains cytoskeletal elements, endoplasmic reticulum, and ground substance. Although present in high concentration in fertilized ova, few or no actin filaments are found associated with the cortices of unfertilized echinoderm eggs and relatively little (9-20%) of the total egg actin pool is present in the polymerized form (Spudich and Spudich, 1979; Otto e t a / . , 1980; Coffe at al., 1982; Yonemura and Kinoshita, 1986). Biochemical studies (Spudich and Spudich, 1979; Otto et a/., 1980) and fluorescence microscopic observations with NBD-phallacidin (Cline and Schatten, 1986; Yonemura and Kinoshita, 1986) demonstrate that monomeric, cortical actin is induced to polymerize into filaments at fertilization. Actin has also been demonstrated in the cortices of amphibian and mammalian eggs (Clark and Merriam, 1978; Maro er al., 1984; Longo and Chen, 1985; Reima and Lehtonen, 1985). Investigations, with both intact eggs and isolated cortices exposed to different ionic conditions, demonstrate that microvillar elongation is stimulated by the calcium flux characteristic of egg activation (Carron and Longo, 1980, 1982; Begg et a / . , 1982). As a consequence of this process, microvilli increase three to four times their original length and obtain polarized bundles of actin filaments (Burgess and Schroeder, 1977).

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Microvillar elongation does not occur when eggs are incubated in media that induce an increase in intracellular pH (e.g., sea water containing ammonia). However, actin filament bundle formation is triggered by an increase in intracellular pH. Formation of actin filament bundles is not necessary for microvillar elongation but is required for rigid support of microvilli. It has been suggested that events of activation prior to the intracellular pH increase induce the formation of cortical microfilament networks and microvillar elongation (Carron and Longo, 1982). The microfilaments provide the structural and/or contractile framework for support of the egg surface, which is undergoing extensive rearrangement. Microfilament organization within the microvilli, i.e. , bundle formation, may then be a consequence of cytoplasmic alkalinization. Hence, actin filament bundle formation in the cortex of the fertilized sea urchin egg appears to be a two-step process (Tilney and Jaffe, 1980): (1) the polymerization of actin to form filaments randomly oriented but in most cases with one end in contact with the plasma membrane, followed by (2) the association of filaments by macromolecular bridges to form bundles. Microvillar elongation also occurs in fertilized medaka eggs (Iwamatsu and Ohta, 1976). Microvilli elongate starting at the opening of dehisced cortical granules; this is followed by a propagation of microvilli along the surface of the dehisced cortical granule. This change is accompanied by the formation of an electron-dense layer, possibly actin, that underlies the former cortical granule membrane. The mechanisms of cortical reorganization are not known but are likely to involve actin-binding proteins as described in other systems (Craig and Pollard, 1982). The distributin of fodrin and its possible relation to actin localization in fertilized sea urchin and mouse eggs have been examined (Schatten et al., 1986a). Based on their findings that all of the zygote regions containing high levels of antifodrin reactivity also appeared to contain high concentrations of polymerized actin, Schatten et al. (1986a) suggested that fodrin, by its interaction with the plasma membrane and cortical actin, stabilizes cytoskeletal-membrane interactions during fertilization. The distribution of a-actinin during fertilization has been investigated by microinjection of rhodamine-labeled a-actinin into living sea urchin eggs (Mabuchi et al., 1985). This probe is uniformly distributed in the cytoplasm of unfertilized eggs. Upon fertilization, however, it concentrates in the zygote cortex including the fertilization cone. Migration of fluorescently labeled a-actinin into microvilli apparently does not occur. Aggregation of actin filaments and their association with bundling protein, e.g., fascin, may give rise to microfilament bundles in egg microvilli (Spudich and Amos, 1979; DeRosier and Edds, 1980; Otto et al., 1980; Tilney and Jaffe, 1980; Mabuchi and Nonomura, 1981). A

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profilin-like protein may prevent actin from polymerizing in the unfertilized egg (Mabuchi, 1981; Hosoya er al., 1982). Although fascin is found in the unfertilized sea urchin egg and has been localized in microvilli of fertilized ova, its interaction with actin has not been shown to be calcium or pH sensitive (Bryan and Kane, 1982). Hence, other actin-binding proteins may be instrumental in microvillar elongation; cytoplasmic alkalinization may give rise to microfilament bundle formation by promoting actin and actin-binding protein interactions. In this context, Otto and Schroeder (1984) have shown that 1-methyladenine stimulates starfish oocytes to undergo major organizational changes involving actin, fascin, and a 220-kDa protein. In addition to changes in microvillar conformation, the eggs of a number of different animals undergo changes in cortical rigidity and contraction that appear to involve a n actomyosin system (see Vacquier, 1981). Cyclical changes in surface tension and contraction have been correlated with cytoskeletal alterations (Coffe et al., 1982) and also occur in anucleate egg fragments with the same cycle as in normal embryos (Yoneda et al., 1978; Shimizu,, 1981b; Yarnamoto and Yoneda, 1983). These observations indicate that egg activation initiates processes autonomous of the nucleus that regulate, in a cyclical manner, cortical cytoskeletal components and cytoplasmic contraction. In addition to contractile processes, the cortical cytoskeleton of fertilized and unfertilized eggs may also be important for other functions. For example, comparison of sea urchin eggs and zygotes indicates a correlation between the activation of protein synthesis and the association of polysomes with the cortical cytoskeleton (Moon et al., 1983).

VII. Endocytosis Immediately following the cortical granule reaction and concomitant with the elongation of microvilli is the development of endocytotic pits and vesicles (Anderson, 1948;Chandler and Heuser, 1979,1981; Donovan and Hart, 1982; Fisher and Rebhun, 1983; Carron and Longo, 1984; Sardet, 1984). Endocytosis in sea urchin eggs commences as a burst 3-5 minutes postinsemination in which portions of the plasma membrane are taken into the cytoplasm (Fig. 1). Whether or not portions of the original plasmalemma or the cortical granule membrane are preferentially endocytosed has not been determined. In light of observations demonstrating significant changes in the composition of the egg plasma membrane at fertilization, it seems unlikely that discrete patches of membrane persist intact to be selectively endocytosed. When the cortical granule reaction is inhibited by high pressure, the

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endocytotic burst that immediately follows is inhibited (Fisher et al., 1985). Interestingly, in such cases endocytosis occurs much later than observed in untreated zygotes. These results suggest that cortical granule exocytosis is not the only cause of surface transformations involving endocytosis. Since endocytosis follows the cortical granule reaction it can be suggested that a mechanism exists for both surface area reduction and cell surface remodeling that may be relevant to physiological changes characteristic of fertilized eggs. The finding that endocytosis follows the cortical granule reaction is consistent with observations in secretory cells in which after exocytosis, excess membrane may be removed from the cell surface in the form of endocytotic vesicles (Orci et al., 1973; Pelletier, 1973; Kalina and Robinvitch, 1975; Oliver and Hand, 1978). The amount of membrane internalized by endocytosis at fertilization appears to be extensive and persists up to the time of cleavage. Whether endocytosis remains constant over this period has not been established, however, it has been estimated that about 26,300 pm2 of surface membrane per Strongylucentrotus egg is resorbed by this process during the first 4 minutes of fertilization (Fisher and Rebhun, 1983). This represents approximately 46% of the membrane presumably added to the egg surface by cortical granule exocytosis. The relationship between cortical granule exocytosis and endocytosis, in terms of the quantity of membrane in flux, is unclear since: (1) the rate of membrane interiorization is unknown, (2) the amount of cortical granule membrane added to the zygote surface has not been definitely established, and (3) mechanisms other than endocytosis that may contribute to the reduction of surface area have not been eliminated. Following the appearance of tracer in endocytotic vesicles of fertilized sea urchin and zebra danio eggs, label has been observed in lysosomes (Carron and Longo, 1984; Donovan and Hart, 1986). This transition indicates that the tracer travels from one cellular compartment to another. The localization of label in lysosomes of zygotes also suggests that surface membrane may be degraded or modified. Membrane components may then reenter cytoplasmic precursor pools by traversing the lysosomal membrane to be utilized at later stages of embryogenesis (De Duve and Wattiaux, 1966; Holtzman, 1976).

VIII. Concluding Remarks As a result of the interactions and fusion of the sperm and egg, changes are initiated that affect virtually a11 components of the egg cortex. The egg plasmalemma becomes a mosaic composed of membranes derived from

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the egg and sperm, as well as from dehisced cortical granules. Although investigations employing various markers indicate the mixing of membranes following sperm-egg fusion and the cortical granule reaction, the amount of membrane from each source that becomes and remains a part of this mosaic has not been determined. Membrane changes reflecting an integration of sperm and egg plasma membranes along the fertilization cone have been demonstrated using tracers at the ultrastructural level of observation. It is anticipated that further efforts using probes to specific membrane components will demonstrate finer changes in molecular topography along this specialized portion of egg cytoplasm. Such approaches may help to establish how alterations in the egg plasmalemma are related to functional changes of the ovum regarding its turning-on of new developmental processes and to events initiated at later stages of embryogenesis. In addition to the integration of membrane from various sources, other major structural alterations of the fertilized egg cortex include the elongation of microvilli and the initiation of endocytosis. Both of these processes accomodate the increase in membrane surface area of the fertilized egg as a result of the cortical granule reaction. Studies examining microvillar elongation, as well as fertilization cone formation, indicate significant changes in the polymerization and the bundling of cortical actin. Regulation of actin and actin-binding proteins in modifications of the fertilized egg cortex, and possible membrane topographical changes that occur along specialized portions of the egg cortex remain to be determined. The shunting of membrane from the egg surface. in the form of endoc ytotic vesicles, to the Golgi complex and lysosomes has been demonstrated by tracer studies. Whether specific membrane components are internalized and their fate will, no doubt, be topics of future studies concerning the reorganization of the egg surface at fertilization. The “preferential” endocytosis of specific membrane domains (e.g., egg, sperm, and/or cortical granule membranes) that become a part of the plasmalemma of the fertilized egg has not been determined. In light of observations demonstrating significant changes in the composition of the egg plasma membrane at fertilization, it seems unlikely that discrete patches of membrane persist intact to be selectively endocytosed. Further study is needed, however, t o verify such a speculation. Whereas endocytosis may contribute to zygote surface area reduction, the interiorization of the fertilized egg plasma membrane may participate in modifications of surface properties and may represent a mechanism for altering the number and pattern of developmentally significant cell surface components.

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INTERNATIONAL REVIEW OF CYTOLOGY. VOL. 113

Ultrastructural Modifications and Biochemical Changes during Senescence of Chloroplasts u. c. BIsWAL AND BASANTIBISWAL School of’Life Sciences, Sambalpur University, Jyotivihar, Orissa,India

I. Introduction Senescence is the last phase of development of a whole organism, organ, cell, or organelle. It is basically a degenerative process that leads to the death of a living system. Although its precise mechanism is not known, recent experiments suggest that the process is genetically programmed (Woolhouse, 1984a). Senescence, although deteriorative in nature, is a requirement to facilitate turnover processes and is an essential process for growing parts of living organisms. Senescence of green leaves in plants has been extensively investigated in the last two decades (Thomas and Stoddart, 1980; Woolhouse and Jenkins, 1983; U. C. Biswal and Biswal, 1984). Different tissues and cells of the leaves have their own pattern and timing of senescence. The initiation of senescence in mesophyll cells may not necessarily be synchronized with the process in vascular or epidermal tissues. Even different organelles of a single leaf cell, namely chloroplasts, mitochondria, endoplasmic reticulum, ribosomes, and the nucleus do not show synchrony in the induction and progress of senescence (Biswal et af., 1983b). Different organelles of a leaf cell degrade sequentially; the sequence may change depending on many factors (Woolhouse, 1984a). A general pattern of temporal changes in the fine structure of different cellular organelles, including chloroplasts, is briefly outlined in Fig. 1. Chloroplasts are the first organelles to show symptoms of disorganization when all other organelles are normal (Dodge, 1970). This is followed by a change in the structure of the endoplasmic reticulum and loss of ribosomes. The next organelle that is affected is the mitochondrion, although at this time the nucleus, plasma membrane, and plastid envelope are shown to be intact. Mitochondria show symptoms of degradation that are reflected in the alteration and loss of the cristae and the reduction in the size of the organelle. Nuclear contents show shrinkage and the membrane becomes structurally irregular (Butler and Simon, 1971). Finally, senescence results in the rupture of the plas27 1

Copyright 0 1988 by Academic Presi, Inc All rights of reproduction in any form reserved

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Completion of c ~ l l ~ l abreakdown r

i.plasma membrane

I

Vsriculation and breakdown

Time scale iProqrerr ot senescence)

FIG. 1. The temporal disorganization of different organelles of a plant cell. The initiation of senescence of a particular organelle is marked by a downward arrow.

tid envelope followed by the breakdown of the plasma membrane (Woolhouse, 1984a). Of all the cellular organelles, chloroplasts are the most studied during plant cell senescence. During senescence chloroplasts show qualitative and quantitative changes in the pigment and macromolecular compositions. molecular structure and organization of thylakoid membranes, primary photochemical reactions, and in the activity of Calvin enzymes. Reviews on senescence-induced ultrastructural modification of chloroplasts as probed by electron microscopy (Butler and Simon, 1971), damage of the electron transport system and loss in the activity of Calvin enzymes (Biswal et af., 1983b; Woolhouse, 1984b), and the regulation of plastid senescence (Thomas and Stoddart, 1980; U. C. Biswal and Biswal, 1984) are available. However, these reviews project leaf senescence in general with a compilation of data on some isolated aspects of chloroplast degradation. In addition, current ideas of chloroplast senescence are rapidly changing with new and exciting results. This article, therefore, compiles recent findings of chloroplast degradation and provides an integrated story of organelle senescence.

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11. Senescence-Induced Structural Modifications A. STRUCTURE OF CHLOROPLASTS The structure and organization of chloroplasts are described in detail by Kaplan and Arntzen (1982). The lamellar structure of chloroplasts varies between photosynthetic organisms. The structure is relatively simple in less evolved plants such as blue-green algae and photosynthetic bacteria and more complex in higher angiosperms. Electron microscopic studies reveal the following basic features of a typical higher plant chloroplast (Fig. 2). The organelle is lens shaped and the lamellar system is encircled by a double membrane envelope. The envelope acts as a selective barrier to the transport of various metabolites into or out of chloroplasts. The lamellar system of the organelle is basically composed of lipoprotein disks that are stacked at regular intervals to form structures known as grana. The unstacked loose lamellae, which connect stacked grana zones, are intergranal or stroma lamellae (Fig. 2). The entire lamellar system is suspended in a granular stroma that is mainly composed of plastid genetic materials and ribosomes responsible for chloroplast replication and self-regulation. In addition, starch grains also constitute a part of the granular structure. The proteins present in the stroma are mostly the enzymes of the Calvin cycle. The other components of the stroma are plastoglobuli, which in electron micrographs appear as darkly stained structures consisting of lipids (Tevini and Steinmuller, 1985). The grana and stroma lamellae are the structural units for primary photochemical reactions and contain the photosynthetic pigments in addition to lipids and proteins. The lipids of the thylakoid membrane consist of a complex mixture. The major lipids of the thylakoid membrane are galactolipids, phospholipids, and sulpholipids (Siegenthaler, 1982; Chapman et al., 1985). Chlorophylls, carotenoids, and phycobilins are the major photosynthetic pigments and are involved in the process of light absorption by the organelle. Chlorophylls a and b, which are chemically different from each other, are abundantly present in the higher plant chloroplasts. According to light absorption and fluorescence characteristics, chlorophyll a in uiuo can have a number of optical species as well as different chemical species (Rebeiz and Lascelles, 1982). The formation of different optical species of chlorophyll a is attributed to the complex interaction of the pigment with the membrane environment. A change in membrane environment could lead to an alteration in the optical behavior of the pigments, Therefore, light absorption and fluorescence spectroscopy are used as sensitive tools to probe the microenvironment of pigments in the

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velope

A

Margins

Stroma

V

mPS-I

V

v

D PS-11

U

0 ATPase

B

FIG.2 . ( A ) Outline diagram of a typical higher plant chloroplast with double layer envelope, thylakoid system, and plastoglobuli. (B) Schematic representation of enlarged grana and slroma thylakoids. The diagram shows appressed and exposed membranes. the sites for distribution of photosystem I (PS I ) . PS 11. and coupling factor.

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membrane (Govindjee and Whitmarsh, 1982). Carotenoids and phycobilins found in lower plants are accessory pigments that absorb light energy and ultimately transfer it to chlorophyll for photochemical reactions. In addition, carotenoids protect chlorophyll molecules from photodestruction (Gillham and Dodge, 1985). Chlorophyll and carotenoids are bound to thylakoid proteins to form different types of pigment-protein complexes, which have recently been well reported. (Kaplan and Arntzen, 1982; Ortiz et al., 1985). Proteins associated with the thylakoid membrane are coupling factor proteins, light-harvesting chlorophyll-protein (LHCP) complex, proteins of the electron transport carrier, proteins associated with the reaction center and the polypeptides of the oxygen-evolving compolex. The proteins of the coupling factor and some of the polypeptides of the water-splitting system are extrinsic, but most of the thylakoid proteins are intrinsic in nature.

B. Loss OF PIGMENTS, LIPIDS, AND MACROMOLECULES The visible symptom of chloroplast degradation in uiuo is marked by leaf yellowing. This is best demonstrated by nature in temperate regions in which the leaves of all the deciduous plants turn yellow in autumn. Leaf senescence causes a loss of all types of photosynthetic pigments. However, depending on the plant system and environmental conditions, differential rates of loss of chlorophyll a, chlorophyll b , and carotenoids have been demonstrated (Sestak, 1977; Woolhouse and Jenkins, 1983; Thomas, 1983; Grover et al., 1986). Carotenoids were shown to be relatively stable in comparison to chlorophylls. In contrast, equal rates of degradation have been demonstrated for chlorophyll a, chlorophyll b, and carotenoids of the chloroplasts of fern leaves (Biswal el al., 1983). 1 . Loss of Chlorophyll The precise mechanism of chlorophyll degradation during chloroplast senescence is not known, although several possibilities have been proposed. The difficulty in proposing a mechanism of chlorophyll degradation is the lack of knowledge of the intermediates of pigment breakdown. The loss of pigment is attributed to the action of chlorophyllase (Terpstra and Lambers, 1983a,b). It has been reported that the activity of the enzyme enhances during senescence (Sabater and Rodriguez, 1978; Lambers et al., 1984). For several reasons, the participation of chlorophyllase in chlorophyll degradation is questionable (Biswal et al., 1983b). The enzyme supposedly cleaves phytol side chains leading to the formation of

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chloroph yllide and phytol. However, breakdown of chlorophyll and formation of chlorophyllide do not decrease the green color since the absorption spectrum of chlorophyllide is similar to that of chlorophyll. This, therefore, does not explain the disappearance of chlorophyll during senescence. In addition, the removal of Mg2' from porphyrin and consequent formation of pheophytin have been demonstrated to be the first steps in chlorophyll degradation (Schoch and Vielwerth, 1983). The nature, location, and activation of chlorophyllase have been extensively investigated by Terpstra and Lambers (1983a,b) and Lambers el al. (1984) who have suggested that the enzyme is an intrinsic thylakoid protein: therefore its activity is modulated by the membrane environment including the lipids of thylakoid membranes. Chlorophyllase, under normal conditions, seems to remain in an inactive form in the membrane. Senescence-induced changes in thylakoid organization may lead to activation of the enzyme, which subsequently breaks down chlorophyll molecules. The proposal is supported by the observation that a thylakoid breakdown is a prerequisite for breakdown of chlorophyll molecules (Sakaki et al., 1983). This, however, contradicts the electron microscopic findings for the sequential degradation of chloroplasts during senescence in which chlorophyll degradation occus much earlier than the disintegration of the thylakoid membrane system (Biswal ef al., 1983b; N . K. Choudhury and Imaseki, personal communication). These results seem to be confusing; thus the question of whether chlorophyll degradation involves chlorophyllase merits further investigation. The other enzymes investigated as possible candidates for a role in chlorophyll degradation are peroxidase and lipoxygenase. Senescenceinduced alteration in the activity of peroxidase is well reported (Sharma and Biswal, 1976; Kato and Shimizu, 1985). However, the location of the peroxidase enzyme, which is mostly formed in vacuoles, does not support peroxidative chlorophyll degradation. During senescence, the chloroplast envelope remains intact until the complete disruption of the thylakoid membrane and loss of significant amounts of chlorophyll. Until then, vacuolar enzymes, including peroxidase, remain separated from the organelle and therefore may not have any access to chlorophyll. Specific peroxidases have been demonstrated within chloroplasts by Martinoia et al., (1982). However, it is not clear whether the total loss of chlorophyll could be due to these plastid-specific peroxidases. Lipoxygenase mediates oxidation of polyunsaturated fatty acids leading to the formation of free radicals that then oxidize chlorophyll molecules; however, the association of lipoxygenase with senescence has recently been questioned (Kar and Feierabend, 1984; Peterman and Siedow, 1985). These authors have demonstrated a decline in the activity of the enzyme during

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senescence. Dupont and Siegenthaler (1986) have, on the otherhand, suggested nonparticipation of any enzymatic system for chlorophyll breakdown, which may, however, be mediated by free radicals. 2 . Loss of Carotenoids

The existing literature does not propose any precise mechanism of carotenoid loss during senescence. Since carotenoids are the integral components of chlorophyll-protein complexes, it is tempting to suggest that their loss may be due to senescence-induced breakdown of these complexes (Grill and Schraudolf, 1981; Biswal et al., 1983b). The relative stability of carotenoids as compared to chlorophyll might be explained in terms of their association with plastoglobuli formed during senescence. Since carotenoids are lipid soluble, on their release they may be immediately captured and protected by the globuli. It is suggested that during the breakdown of thylakoids, the fatty acids released interact with the liberated carotenoids to form carotenoid esters in plastoglobuli thus rendering the pigments stable (Tevini and Steinmuller, 1985). 3. Loss of Lipids In addition to the loss of pigments, some specific thylakoid lipids are lost during senescence. Senescence results in differential rates of degradation of different types of membrane lipids, namely monogalactosyl diglycerides, digalactosyl diglycerides, sulfoquinovosyl diglycerides, phosphatidyl glycerol, phosphatidyl choline and phosphatidyl inositol (Harnischfeger, 1973; Koiwai et a!., 1981; Tevini and Steinmuller, 1985). The degradation of lipids is attributed to senescence-induced stimulation in the activity of lipases (Leshem et al., 1984; Woolhouse, 1984a; Thomas, 1986).

4. Loss of Macromolecules A differential loss of plastid-specific macromolecules, namely DNA, RNA, and proteins, is demonstrated during senescence (Makrides and Goldthwaite, 1981). The loss of plastid DNA is relatively slow compared to the loss of protein and RNA (Biswal and Mohanty, 1976a). The relative stability of DNA is evident from the recovery of already damaged chloroplasts during regreening. The stability of DNA is also significant because chloroplasts need specific messages, even at the last phase of their own degradation. The proteins that are lost during senescence include electron transport carriers (Woolhouse, 1984b), polypeptides associated with PS I1 activity, chlorophyll-protein complexes (CP) of PS I (Dos Santos and Hall, 1982), coupling factor proteins associated with photophosphorylation (Camp et

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al., 1982; Choudhury and Biswal, 1984a), and enzyme proteins of the Calvin cycle including ribulose 1,Sdiphosphate carboxylase (RuDPCase) (Lauriere, 1983). Senescence is reported to result in a significant decline in the level of plastid-specific ribonucleic acids (Biswal and Mohanty, 1976a: Panigrahi and Biswal, 1979a). The decreased levels of macromolecules could be attributed to senescence-induced stimulation of nuclease and protease activity or a reduction in the capacity to synthesize these macromolecules (Biswal et a / . , 1983b; Wettern and Galling, 1985).

C. ULTRASTRUCTURAL CHANGES Senescence-induced ultrastructural modifications, including changes in the molecular composition of thylakoids, have been probed by electron microscopy, X-ray diffraction, immunotechniques, absorption and fluorescence spectroscopy, and phase properties of membranes, Of all these probes, electron microscopy has been used extensively by many authors to examine the changes in size, shape, and ultrastructural organization of chloroplasts during senescence (Butler and Simon, 1971; Thomas, 1977; Naito et al., 1981; Wittenbach et al., 1982; Biswal ef al., 1983a). Chloroplasts of completely yellow leaves appear oval and exhibit minimal lengths and widths indicating senescence-induced shrinkage of the organelle (Naito et al., 1981). During senescence. the organelle shows disorganization at the level of both the envelope and lamellar structures. Lamellar breakdown, as revealed by electron microscopy, exhibits unstacking of grana thylakoids (Mlodzianowski and Mlodzianowski, 1973). This is followed by the formation of loose, elongated lamellae scattered in the organelle (Fig. 3). In the next phase, these lamellae undergo further degradation with the concomitant appearance of plastoglobuli (Hudak. 1981; Biswal er a / . , 1983a). The size of the globuli increases as senescence advances (Kulandaivelu and Senger, 1976b). Although these ultrastructural modifications have been qualitatively studied, attempts have been made by Naito et al. (1981) to quantify some of the structural parameters. The authors have demonstrated a decrease in the number of grana per chloroplast section and in granum length with an increase in the number, length, and width of plastoglobuli during senescence. They have also noted senescence-induced qualitative Fic. 3. Ultrastructural changes of chloroplasts of mustard cotyledons during senescence induced by darkness. (a) Seven days after sowing in continuous white light. (b) Fourteen days after sowing in continuous white light. ( c and d) Plastids of cotyledons that were irradiated for 7 days with continuous white light and transferred to darkness; c, 4 days of darkness: d, seven days of darkness. From Biswal ~t al. (1983).

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changes such as the shrinkage of chloroplasts, disappearance of intergranal lamellae, and random orientation of grana followed by their disappearance. Chloroplast degeneration at the final stage of senescence is marked by the disruption of the chloroplast envelope, which leads to rapid lysis of the rest of the chloroplast components (Woolhouse, 1984a). The sequence of ultrastructural changes during chloroplast senescence varies depending on the environmental conditions and plant species. For example, the pattern of ultrastructural changes of chloroplasts during senescence of naturally occurring attached primary leaves of wheat is qualitatively different from the sequence of changes observed in the chloroplasts on detachment of leaves (Hurkman, 1979). The initial event in the degradation of chloroplasts of attached leaves undergoing natural senescence is the appearance of plastoglobuli and the simultaneous disorganization of the thylakoid system. This is followed by formation of vesicles and an increase in the number and size of plastoglobuli. In the final stage, the chloroplast envelope ruptures and the plastid contents, namely plastoglobuli and the remains of the thylakoids, fall into the cell lumen. On the otherhand, plastid degradation during senescence of wheat leaves detached from the parent plant begins with the rupture of the envelope followed by the collapse of the thylakoid system. In this case, swelling of the grana and intergranal lamellae is not significant and the lamellar system breaks down without the formation of vesicles and numerous plastoglobuli. Similarly, mustard cotyledons undergoing senescence in different environmental conditions show qualitatively different ultrastructural modifications. Senescent chloroplasts of cotyledons of naturally growing mustard plants exposed to daily light-dark cycles show the formation of giant grana structures (Hudak, 1981) that are not observed in cotyledon chloroplasts during senescence induced by continuous dark treatment (Biswal ef a / . , 1983a). The ultrastructural changes of mustard cotyledons during dark-induced senescence, and the effect of light through phytochromes in retarding these changes are shown in Figs. 3 and 4. The pattern of illtrastructural changes of chloroplasts differs with plant species, even though the experimental conditions remain same. For example, rupture of the plastid envelope is followed by the degradation of the thylakoid system during senescence of wheat leaves (Hurkman, 1979), however in the chicory leaf, the thylakoid system degrades first and is followed by the rupture of the envelope, although in both cases, detached leaves were used for the experiments (Mlodzianowski and Mlodzianowski, 1973). Ultrastructural degradation consists of three major events: thylakoid breakdown, formation of plastoglobuli, and rupture of the envelope.

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28 1

FIG. 4. Phytochrome-induced retardation of ultrastructural changes during senescence of mustard cotyledons. The seedlings were grown in continuous white light and were transferred to darkness followed by different light treatments. (a) Red light pulses (phytochrome action). (b) Continuous red light (phytochrome action). (c) Red plus far-red light pulses (no phyochrome action). (d) Far-red light pulses (no phytochrome action). From Biswal ef al. (1983).

The precise mechanism of senescence-induced thylakoid breakdown is not known. Unstacking of grana, the first step of lamellar disorganization, could be due to the loss of chlorophyll b and LHCP (Biswal et al., 1983a). Chlorophyll b and LHCP are known to be responsible for grana stacking and both of them are reported to degrade during induced senescence (Lichtenthaler et al., 1981; Biswal et al., 1983a). It is also possible that a senescence-induced loss of ions could result in destacking (Raval et al., 1984). Lamellar degradation, the next event of thylakoid breakdown, could be carried out by the specific enzyme systems. The enzymes for the breakdown of thylakoids are reported to be associated with the organelle itself (Dalling et al., 1983; Thomas, 1984; Wettern and Galling, 1985; Martin et al., 1986). The breakdown of thylakoid membranes and the concomitant formation of plastoglobuli have led to the proposal that these globuli are composed of lipids resulting from senescence-induced lamellar

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breakdown. This is further confirmed by the fact that as senescence advances, the increased thylakoid breakdown is accompanied by a corresponding increase in the number and size of the plastoglobuli (Biswal et al.. 1983a). The structural closeness of thylakoids and globuli is distinctly shown in senescent chloroplasts of Phaseofus leaves (Barton, 1966) in which characteristic swelling of each thylakoid terminus is seen adjacent to dense globuli. Cytokinin treatment prevents an increase in the number of plastoglobuli and simultaneously prevents thylakoid disintegration (Naito et a / . , 1981). This also suggests a correlation between the level of plastoglobuli and the structural status of thylakoid membranes. On the other hand, some reports suggest that plastoglobuli may only in part be the product of lamellar breakdown and that the rest of their structure may represent a nonspecific storage of insoluble lipids (Bailey and Whyborn, 1963; Lewington ~t al., 1967). This view is supported by the observation that the globuli from mature chloroplasts of Vicia faba and spinach contain a mixture of lipids and other substances that are not specific to the lamellar membrane system. The metabolism of plastoglobuli with reference to thylakoid breakdown during chloroplast senescence was recently investigated by Tevini and Steinmuller (1985). They have analyzed the dynamic changes in the levels of galactolipids, carotenoids, plastoquinones, and other lipids simultaneously in the thylakoids and plastoglobuli and have proposed a correlation between these two structural units. The fatty acids, hydrolysis products of thylakoid lipids, are suggested to form esters of carotenoids in plastoglobuli. In addition to carotenoids and fatty acids, prenylquinones, components of thylakoids, also deposit in plastoglobuli. Glycolipids, the major constituents of thylakoid lipids, do not deposit in plastoglobuli, and therefore the degradation of these lipids, as that of proteins, is insignificant in the metabolism of plastoglobuli. Although these data provide some basic information on the chemical composition of plastoglobuli, the biochemical significance of plastoglobuli during senescence merits further investigation. The retention of the integrity of the plastid envelope and plasma membrane may suggest the involvement of these membrane systems in finally transporting the degradative products to other growing cells in the plants (Woolhouse, 1984a). Rupture of the envelope, therefore, represents the last sequence of the structural degeneration of chloroplasts. Two tentative mechanisms can be proposed to explain envelope rupture, one of which is mechanical. In this case the rupture is correlated with the senescence-induced extensive vacuolation of chloroplasts (Hudak, 1981). The origin of vacuoles is attributed to the local lysis of chloroplast stroma. The vacuolation could exert pressure on the chloroplast envelope resulting in it5 rupture. The second mechanism proposes that the rupture of the

CHANGES DURING SENESCENCE OF CHLOROPLASTS

283

envelope may occur by the hydrolytic enzymes released by breakage of tonoplasts. The breakdown of tonoplasts is a crucial phase of senescence because it provides for the direct contact of cellular vacuoles and different cell organelles, including chloroplasts (Peoples et al., 1980). The cytoplasmic vacuoles containing relatively high concentrations of phenols, organic acids, and hydrolytic enzymes could act on the boundary envelope of chloroplasts resulting in its breakdown.

D. OPTICALCHANGESAND MEMBRANE PHASESHIFT 1. Optical Changes

Electron microscopy has certain limitations in probing the ultrastructural changes of chloroplasts. Experimental artifacts may arise particularly during fixation and embedding of the senscent samples. It is quite likely that senescent chloroplasts that are already damaged may react differently to these reagents. In addition, electron micrographs can reveal only gross changes of the thylakoid system without providing any clue to the alteration of the molecular composition of membranes. Therefore, optical probes such as light scattering, absorption, and fluorescence of chloroplasts are used to reveal the specific changes in the membrane organization during senescence (Biswal et al., 1983b). These probes are used to examine the level of stacking of thylakoid membranes, distribution of pigments in pigment assemblies, specific orientation of pigments on the membrane surface, conformational changes of pigment protein complexes, and organization of the reaction center. The sensitivity of these optical probes has been previously examined in the investigation of structural changes of chloroplasts treated with heat and surfactants (Biswal and Mohanty, 1976b). It is known that the bands characteristic of chlorophyll absorbance in solution are different from those observed in pigment attached to the membrane surface suggesting an interaction of chlorophyll with membrane ultrastructure (Biswal et al., 1983b). A change in the ultrastructure could therefore be manifested as an alteration(s) in the absorption spectra of pigments. Senescence is known to result in a shift in the peak of the red band of chloroplast absorbance indicating membrane disorganization (Biswal and Mohanty, 1976b; Sayeed et al., 1985). In addition to the change in peak position, the peak height and band width are also altered (Brody, 1983). A significant change in the scattering of chloroplasts has also been demonstrated (Raval et al., 1982). Although the precise nature of the structural changes of the thylakoid membrane that could lead to these optical changes is not known, a differential degradation of photosystem I (PS I) and photosystem I1 (PS 11) pigments may cause a shift in

284

U. C . BiSWAL A N D BASANTI BISWAL

the red band. Senescence may induce changes in the orientation and/or number of thylakoids per granum in chloroplasts as reported by many authors (Dodge, 1970; Mlodzianowski and Mlodzianowski, 1973; Naito et al., 1981). These changes may find their expression in the red absorption band. Unstacking of grana, as revealed by electron microscopy, may also lead to a change in the optical behavior of senescent chloroplasts (Raval e t ul., 1984). Similarly, analysis of changes in chlorophyll a fluorescence of senescent chloroplasts provides information about the structural disorganization of thylakoid systems (Biswal et al., 1979: Woolhouse and Jenkins, 1983; Biswal, 1987). Enhancement or quenching of the fluorescence of endogenous chromophores like chlorophylls or exogenous fluorophores bound t o thylakoid membranes is indicative of alterations in membrane organization (Biswal et d . , 1983b). Fluorescence quenching induced by senescence is attributed to the loss of fluorescing pigments and structural alterations of thylakoid membranes that favor the redistribution of excitation to PS I and thermal relaxation of PS 11 pigments (Jenkins et al., 1981). Fluorescence polarization has been used to probe the structural status of the photosynthetic unit and physical state of membranes (Mar and Govindjee, 1971). Senescence causes a significant enhancement in fluorescence polarization of celi-free chloroplasts or chloroplasts in leaves (Biswal, 1987). The increase in polarization may suggest a disorganization of photosynthetic assemblies that consequently produces a change in the efficiency of quantum migration or uncoupling of the PS I1 reaction center and antenna chlorophyll molecules. The increase in polarization is also indicative of senescence-induced reduction in the size of the photosynthetic units in thylakoid membranes (Biswal, 1987). Alternately, an increase in chlorophylla fluorescence polarization may suggest conversion of a liquid crystalline state of the membrane to a gel phase that restricts the mobility of the chlorophyll dipole (Venkatarayappa et al., 1984). Senescence-induced modifications in the structure and surface organization of thylakoid membranes are also investigated by exogenous fluorophores. A considerable increase in the quenching of Atebrin fluorescence has been recorded during dark-induced senescence of chloroplasts of maize (Choudhury and Biswal, 1984b) and wheat leaves (Raval et af., 1982) suggesting modifications of the surface charge distribution of thylakoid membranes. L,ight scattering by chloroplasts has long been used as a probe to investigate structural changes of thylakoid membranes induced by photoelectron transport and by different treatments. Recently, scattering has been monitored to examine the stacking status of thylakoid membranes during senescence (Raval et al., 1984; N . K . Choudhury and H. Imaseki, personal communication).

CHANGES DURING SENESCENCE OF CHLOROPLASTS

285

2. Membrane Phase Change Senescence of chloroplasts is known to result in alterations of the lipid composition of thylakoid membranes as described earlier. This consequently brings about a change in the physical state of thylakoid membranes. The analysis. of the data from wide-angle X-ray diffraction of chloroplasts suggests that formation of the gel phase occurs with advancing senescence (McKersie and Thompson, 1978). It is reported that the senescence-induced phase transition of thylakoid lipids makes the membrane leaky. A shift from the liquid crystalline to the gel phase is further confirmed by a shift in the phase transition temperature (Ginkel and Fork, 1981) and an increase in chlorophyll a fluorescence polarization (Biswal, 1987). A reduction in membrane fluidity induced by senescence is also confirmed by Ford and Barber (1980)using 1,6-diphenyl-l,3,5-hexatriene as a fluorescent probe. Venkatarayappa et al. (1984) have established a correlation between membrane lipid peroxidation and alterations in the thylakoid phase properties during chloroplast senescence. The changes in the lipid phase are shown to coincide with the loss of chloroplast pigments or proteins, which may suggest that a shift in the lipid phase of the membrane may stimulate chloroplast degradation.

111. Loss of Primary Photochemical Reactions

A. PHOTOELECTRON TRANSPORT SYSTEM The pigments associated with thylakoid membranes are characteristically assembled into two photosystems, namely photosystem I (PS I) and photosystem I1 (PS 11), that are linked by a series of electrontransport carriers (Cramer et ul., 1985). The two photosystems have specific orientations across the thylakoid membrane (Haehnel, 1984). It is proposed that PS I is located in stroma lamellae and is primarily responsible for cyclic electron flow. PS I in grana, on the otherhand, is linked with PS I1 and is largely involved in the reduction of NADP'. Normally, higher wavelength-absorbing pigments are present in PS I that do not provide any significant contribution to the fluorescence emitted by chloroplasts. The reaction center of PS I is known as P700, which is not directly associated with the oxygen-evolving complex. The major electron-transport carriers associated with PS I are plastocyanin on the oxidizing side, ferredoxin, P430, and NADP reductase on its reducing side. The pigments associated with PS I1 are the short wavelength form of chlorophyll a and other accessory pigments that contribute significantly to the total fluorescence of chloroplasts. PS I1 is responsible for oxygen evolution, although the oxygen-evolving complex has been identified as a

286

U. C. BISWAL AND BASANTI BISWAL

separate structural unit (Raval and Biswal, 1985). P680 is the reaction center of PS 11. The major electron-transport carriers of this photosystem are cytochrome b559, pheophytin, and quinones. Various steps of photochemical reactions, namely light absorption by antenna pigments, migration of light quanta to the reaction center, separation of charge, and movement of electrons, are best explained in the Z scheme as shown in Fig. 5 . The scheme explains the participation of PS I and PS I1 in the synthesis of ATP and formation of NADPH. Water is the ultimate natural electron donor that donates electrons to Q. The process is photosensitized by the reaction center of PS 11 that on excitation, loses electrons to Q and is oxidized to a strong oxidant capable of splitting water to liberate oxygen through the oxygen-evolving enzyme system. Similarly, P700, the reaction center of PS I on excitation loses electrons to P430 and is oxidized. The reduced Q and the oxidized P700

-

0.4

0

L

-

EQ"

+0.4

I

+ox

I

I

FIG. 5. Scheme summarizing the electron-transport chain of the chloroplast membranes. The thicker the dotted arrow, the more sensitive the site is to senescence. DCIP, ?,6-dichlorophenolindophenol;DCMU. 3-(3,4-dichlorophenyl)- I , I-dimethylurea; DPC, diphenylcarbazide: Fd. ferredoxin; FRS. ferredoxin-reducing substance; MV, methyl viologen; NADP. nicotinamide adenine dinucleotide phosphate; P680, reaction center of PS 11: P700. reaction center of PS I; PS I. photosystem I: PS 11, photosystem 11: PC. plastocyanin: PQ. plastoyuinone; P430. primary electron accepor of PS I; QA, primary quinone pool: Qs. .secondary quinone pool.

CHANGES DURING SENESCENCE OF CHLOROPLASTS

287

are reoxidized and rereduced, respectively, due to the continuous flow of electrons from water to NADP'. A flow of electrons from P430 through ferredoxin systems results in the reduction of NADP' in the presence of NADP reductase. In addition to water, a number of exogenous electron donors have been identified and are known to donate electrons at specific sites of the electron-transport chain (Fig. 5 ) . Exogenous electron donors, acceptors, and inhibitors are used extensively to characterize the specific sites of the electron-transport chain of the thylakoid membrane. Photoelectron transport in thylakoid membranes is coupled to ATP synthesis through coupling factor (CF,j, a well-characterized extrinsic protein complex of the thylakoid membrane (Strotmann and BickelSandkotter, 1984). Thus, light-driven electron transport in thylakoid membranes generates NADPH and ATP, the two essential molecules required for fixation of carbon dioxide to sugar through a series of enzymes in the Calvin cycle. Although most of the enzymes of the Calvin cycle are well studied, ribulose 1,5-diphosphate carboxylase (RuDPCase) has drawn the attention of many workers because of its complex structure, regulation of its synthesis, and susceptibility to different environmental stresses. The enzyme is also a major protein of green leaves. B. DISASSEMBLY OF P s 1 AND Ps 11 AND LOSS OF ELECTRON-TRANSPORT EFFICIENCY Senescence causes, a reviewed earlier (Sections I1,B and Cj, an extensive disruption of the thylakoid system including the unstacking of grana and disappearance of photosynthetic pigments. This kind of structural degradation could logically lead to a loss in the capacity of chloroplasts for photoinduced electron transport. A decline in the rates of photochemical reactions in chloroplasts undergoing senescence has been demonstrated by many authors using different plant systems (Biswal et al., 1983b; Woolhouse, 1984b). There are, however, different views on the nature of the loss of photochemical reactions. For example, Camp et al. (1982) have shown relative stability in whole chain electron-transport efficiency when it is measured per chloroplast as compared to a leaf area basis at the initial stage of senescence. These findings may suggest that a loss in photochemical potential during senescence could be due to loss of whole chloroplasts rather than the loss of thylakoid organization. On the otherhand, the loss in the capacity for photochemical reactions has also been attributed to senescence-induced temporal disorganization of thylakoid ultrastructure, loss of chlorophyll, and physical changes in thylakoid membranes (Choudhury and Biswal, 1979a, 1984a; Hilditch et al., 1986).

288

L C BISWAL AlvD BASANTI BISWAL

A gradual decay of membrane structure and consequently a decline in the photochemical potential of chloroplasts are, however. compatible with sequential degradation of thylakoid structure as revealed by electron microscopic studies. Senescence induced-loss of photochemical reactions associated with PS I and PS I1 has been demonstrated to be sequential (Biswal and Mohanty. 1976a: Jenkins et 01.. 1981: Choudhury and Biswal, 1984a; Choc and Whang. 1986; Grover Pt a / . , 1986). Senescence of detached barley leaves was shown to cause a relative instability of PS 11 of thylakoid membranes (Biswal and Mohanty, 1976a). A preferential loss of chlorophyll b , a major pigment of PS 11, is demonstrated during senescence of mustard cotyledons treated with continuous darkness (Biswal et “ l . , 1983a) suggesting a loss of LHCP and disassembly of PS I1 of thylakoid membranes (Bennett. 1981; Lichtenthaler et nl., 1981). This could be correlated with the loss of PS I1 photochemical reactions during senescence of leaves (Biswal and Mohanty. 1976a; Choudhury and His\\al, 1984a1, algae (Kulandaivelu and Senger, 1976a), and isolated chloroplasts (Choe and Whang. 1986). Severe damage to the oxygenevolving complex might also be a factor in the loss of the photochemical potential of PS I1 ofthylakoid membranes (Choudhury and Biswal. 1979b; Hilditch rf d . , 1986). On the othcrhand. Sayeed P r r i l . ( 1985) have demonstrated that during senexence of chloroplasts in detached wheat leaves, a relatively rapid loss of chlorophyll LI occurs indicating the disintegration of the PS I chlorophyll-protein complex. This is further confirmed by their observation of the relative instability of PS 1 photochemical reactions. The preferential loss of PS I is correlated with the senescence-induced disappearance of stroma lamellae before damage to the lamellar system associated with grana thylakoids (Woolhouse and Jenkins, 1983). The instability of PS I of thylakoid membranes is attributed to the inactivation or loss of ferredoxin-NADP reductase, FRS. and plastocyanin (Postius, 1971 : Eytan t’t d., 1974) and a loss of the PS I reaction center ( Jenkins et (11.. 198 1). A simultaneous decline in the primary photochemical reactions associated with both PS I and PS I1 of chloroplasts has recently been observed during senescence of a nonyellowing mutant of Fesirrcu pratensi,c (Hilditch ef a / . , 1986). In addition to the differential loss of the photochemical reactions of PS 1 and PS 11, Biswal and Mohanty (1976a) have proposed specific alterations of the sites in the electron-transport chain between water and the PS I1 reaction center of senescent chloroplasts. Their observations suggest that diphenylcarbazide (DPC)and Mn’+ feed electrons at different sites in the electron-transport chain, DPC donating at a site closer to the PS I1

CHANGES DURING SENESCENCE OF CHLOROPLASTS

289

reaction center. The site for entry of electrons from Mnzt remains further away from the reaction center. During senescence, the site for Mn” disappears faster than the site for entry of electrons from DPC. Experiments involving the application of different electron donors and acceptors specific to different sites of the electron-transport chain and quantitative measurement of electron-transport components suggest that a major change limiting electron transfer in senescent chloroplasts lies between the two photosystems rather than within them. An inactivation of plastoquinone (PQ), as suggested by Woolhouse and Jenkins (1983), and loss of the cyt b-cytfcomplex, as indicated by David-Ben et al. (1983), may be the factors involved in the loss of electron-transport efficiency.

C. DAMAGE TO

THE

OXYGEN-EVOLVING COMPLEX

A decline in oxygen evolution and a restoration of the senescenceinduced loss of PS II-mediated DCIP photoreduction with exogenous electron donors are indicative of the limitations imposed by a damaged oxygen-evolving system on the efficiency of the electron-transport system of thylakoid membranes during senescence. Although the precise geometry and the mode of function of the oxygen-evolving complex are not yet clearly known, recent findings propose the participation of a number of polypeptides that constitute the 02-evolvingcomplex (Murata and Miyao, 1985; Renger and Govindjee, 1985). Some of the polypeptides are known to be intrinsic and are embedded in the lipophilic environment of the membrane, and some are extrinsic, obviously vulnerable to externally applied reagents. The model proposed by Raval and Biswal (1985) provides the details of the arrangement of the polypeptides and their association with Mn2+ to form an active oxygen-evolving enzyme complex. Some of the polypeptides in this model are shown to be extrinsic (Fig. 6). The orientation of extrinsic proteins proposed in the model is supported by the experimental data of Murata and Miyao (1985) who have shown three extrinsic proteins, namely 33-, 24-, and 18-kDa polypeptides associated with the oxygen-evolving complex. Mild treatments with Tris buffer or high pH are shown to release these polypeptides into the aqueous phase from the membranes (Kuwabara and Murata, 1982; Murata and Miyao, 1985). These experiments suggest that extrinsic proteins that are exposed to the aqueous phase and not well protected in the membrane milieu are labile and highly susceptible to stress. It is, therefore, logical to suppose that these polypeptides could behave in a manner similar to the soluble stroma proteins in terms of their accessibility to proteolytic enzymes during senescence. The degradation of these

290

U . C. BISWAL AND BASANTI BISWAL

FIG.6. Structural components of the oxygen-evolving complex. A, active site: MET, manganese electron trap: LRP. lysine-rich polypeptie; P680. photon trap of photosystem 11; Z, primary physiological donor to P680+; phe. pheophytin; FeQA,Iron-quinone pool; Qe. a second quinone pool; HBP. herbicide-binding polypeptide; PQ. plastoquinone; LHPC, light-harvesting protein complex; Antenna. antenna chlorophyll-protein complex. The figure shows three extrinsic proteins, namely 32- to 34-, 23- to 2 5 , and 14- to 18-kDa polypeptides. From Raval and Biswal. 1985.

extrinsic proteins could lead to a decline in the efficiency of the oxygen-evolving complex. In fact, N.K. Choudhury and H. Imaseki have recently observed a senescence-induced quantitative loss of 33-, 24-, and 18-kD polypeptides, and consequently a loss in the oxygen evolution capacity of barley chloroplasts (personal communication). Alternatively, senescence-induced loss of Mn'+ ions could be a factor in the damage to the oxygen-evolving complex. Although the precise role of Mn2+ in the oxygen-evolving complex remains unknown, two different pools of the metal ion in the complex are suggested. One pool of the ion is tightly bound to the intrinsic polypeptides and acts as the charge accumulator at the active site of the water-splitting system. The second pool of Mn2+ which is bound to the extrinsic protein, is relatively unstable and seems to be susceptible to different treatments (Raval and Biswal, 1984; Raval et al., 1985). The extrinsic 33-kDa polypeptide has recently been shown to be responsible for stabilizing Mn" (Murata and Miyao, 1985). Heat treatment causes the release of Mn" along with the extrinsic protein. A SO% quantitative loss of Mn2- results in the complete inactivation of oxygen evolution (Nash et al., 1985). It is possible that during senescence a loss in Mn" may result in damage to the oxygen-evolving complex. Supporting this view is the report that senescence causes a quantitative loss of chloroplast manganese (Margulies, 1971).

CHANGES DURING SENESCENCE OF CHLOROPLASTS

29 1

D. Loss OF PHOTOPHOSPHORYLATION A few reports are available on the senescence-induced loss of the ability of plastid membranes to synthesize ATP (Hernandez-Gil and Schaedle, 1973; Biswal and Mohanty, 1978; Camp et al., 1982; Choudhury and Biswal, 1984a). There are several possible explanations for the mechanism of senescence-induced loss of photophosphorylation. A decline in the efficiency of the electron-transport system during senescence as reviewed earlier may limit the ATP-synthesizing ability of the membrane. This is also evident from the observation of the senescenceinduced loss of activity of the thylakoid-bound Mg*'-ATPase (Choudhury and Biswal, 1984a). The activity of this enzyme reflects the energization status of thylakoid membranes induced by photoelectron transport. An inactivation or quantitative loss of coupling factor proteins could also lead to a decline in photophosphorylation (Thomas, 1983; Choudhury and Biswal, 1984a). A change in the aggregation pattern and conformation of the coupling factor may result in a decline in its functional efficiency. In fact, Choudhury and Biswal (1984a) have demonstrated a structural modification on the surface of the coupling factor as probed by Atebrin fluorescence during senescence of maize chloroplasts. They have also demonstrated a quantitative loss of coupling factor protein that contributes to the loss of the ability of chloroplasts to synthesize ATP. A quantitative loss of coupling factor protein has also been demonstrated by Camp et af. (1982) during senescence of wheat chloroplasts. They have shown that chloroplasts isolated from mature leaves contain more a and /3 protein subunits of the coupling factor than chloroplasts from senescent leaves, suggesting a senescence-induced selective loss of coupling factor subunits.

IV. Loss of RuDPCase Activity The modification of enzymes of the stroma has been examined during chloroplast senescence (Batt and Woolhouse, 1975; Gordon et al., 1978; Makino et af., 1985; Grover et al., 1986). The activities of NADPdependent glyceraldehyde 3-phosphate dehydrogenase, phosphoglycerate kinase, ribose 5-phosphate isomerase, phosphoribulokinase, and ribulose 1,5-diphosphate carboxylase reportedly decrease during senescence. Ribulose 1,5-diphosphate carboxylase has been shown to be most susceptible to senescence (Lauriere, 1983). There is, however, controversy concerning the relative susceptibility of RuDPCase compared to that of chlorophyll and photochemical reactions. For example, it has been suggested that the loss of RuDPCase activity occurs earlier and faster

292

U . C . BISWAL AND BASANTI BISWAL

than the loss of chlorophyll or photoelectron transport (Biswall et al., 1983b). On the otherhand, the enzyme shows more stability than chlorophyll during senescence of the chloroplasts of wheat flag leaves (Hall er a / . , 1978). Similarly. Wittenbach er a / . (1982) have demonstrated that photosynthesis declines faster than RuDPCase activity during senescence of chloroplasts isolated from the protoplasts of wheat leaves (Table I). Data on the parallel loss of photoelectron transport and carbon dioxide fixation are also available (Hernandez-Gil and Schaedle, 1973; Gordon et d., 1978). Recently, Grover er al. (1986) have suggested a simultaneous triggering of the loss of chlorophyll and RuDPCase activity during dark-induced senescence of wheat leaves. The review in this section concentrates on the data available for the mechanism of loss of this enzyme during senescence of chloroplasts in different plant systems.

A . MECHANISM OF Loss It is well known that RuDPCase consists of large and small subunits differing in molecular weight. The large subunit is encoded by plastid DNA and is synthesieed on 70 S ribosomes of the organelle. On the otherhand, the small subunit of the enzyme is encoded by the nuclear gene and the protein is synthesized on 80 S ribosomes of the cytoplasm. The synthesis of the small subunit positively controls the synthesis of the large subunit, suggesting a nuclear control over synthesis of chloroplast proteins (Frosch et d . , 1976). The enzyme has been studied extensively during senescence of chloroplasts. The reports on the loss of its activity are rather controversial. Several factors could contribute to the loss of enzyme activity. Loss of enzyme protein could be a factor contributing to the decline in enzyme activity expressed on either a total soluble protein or leaf area basis. Wittenbach (1978) has quantitated the protein irnmunochemically by using a rocket electrophoresis technique and has reported an enzyme loss constituting about 80% of the total soluble protein even at the initial stage of senescence. Interestingly, at this stage of senescence no significant change in enzyme activity per mg of antibody-recognized carboxylase could be obtained. During senescence of rice chloroplasts, Makino e t a / . (1983, 1984) have shown that no change in the specific activity of RuDPCase occurs when expressed on an enzyme protein basis. However, a decline in the activity is obtained when it is calculated on a leaf area basis. Similar results for the senescence-induced loss of enzyme proteins are reported for barley leaves (Peterson and Huffaker, 1975) and wheat leaves (Mae et al., 1984; Grover el al., 1986). The quantitative loss of RuDPCase during senescence has also been examined by immunological

TABLE I COMPARISON OF MATURE AND SENESCENT WHEATLEAFPROTOPLASTS“,~ Component Chloroplasts

Chl

Protein

RUDPCase

Photosynthesis ~~

Days in the dark

0 3 5

(no./protoplast)

(wg/106 protoplasts)

(pg/106protoplasts)

(nmol COz/ min/lo6 protoplasts)

146( 100) 89(6 1) 47(32)

93(100) 64(69) 35(38)

863( 100) 534(62) 290(34)

3 14(100) 85(27) 14(4)

Protease

~

(nmol O,!min/ 1O6 protoplast s )

(nmol NH2/hour/ lo6 protoplasts)

16 1( 100) 3 1( 19) O(0)

159(100) 213(134) 273(172)

RuDPCase activity was determined by “CO? incorporation at 25°C in the presence of NaHI4CO3and ribulose diphosphate. Photosynthesis was measured using an 0 2 electrode. Protease activity was measured using RuDPCase as the substrate. From Wittenbach et al. (1982). Numbers in parentheses represent percentage of day 0.

294

U. C. BISWAL AND BASANTI BISWAL

techniques during chloroplast senescence in wheat flag leaves (Peoples et al., 1980). At initial stages of senescence, the protein degrades at a rate almost equal to that of the degradation of other soluble proteins. Subsequently. the enzyme protien is rapidly lost, however, without any change in the specific activity of the enzyme as expressed on an enzyme protein basis. No change in the number of active sites as probed by the ['4C]cyanide-binding assay could be demonstrated. Although, these reports suggest a quantitative loss of enzyme protein as a major factor for the loss of enzyme activity, a senescence-induced decrease in the specific activity of the enzyme has been demonstrated by many authors in different plant systems (Callow, 1974; Hall and Brady, 1977; Hall et al., 1978; Wittenbach et a/., 1982). The decrease in the specific activity of the enzyme as calculated on the basis of enzyme protein is obviously the consequence of the senescenceinduced qualitative modification of the enzyme. This could be an alteration in the enzyme such as a change in the number of active sites and/or modification of the sites. For example, the senescence-induced rapid decline in the absolute specific activity of RuDPCase has been attributed to a loss of active sites (Wittenbach. 1978; Wittenbach et al., 1982). A change in the kinetic form of the enzyme could also lead to a lowering of enzyme activity during senescence (Hall er al., 1978). The results suggesting a senescence-induced loss of enzyme protein or a qualitative modification of the enzyme structure leading to a decline in its specific activity are rather conflicting, which prevents a generalization about the behavior of the enzyme during senescence. The data should be carefully analyzed with regard to the experimental conditions, which seem to be very divergent. Various experimental models are used by different authors using different plant systems. In addition, the methods used to assay the proteins such as quantitative analysis of protein by electrophoresis and semiquantitative or quantitative analysis using immunochemical techniques are also different. It is also difficult to find suitable probes to characterize the fine structural changes at the active site or other qualitative changes of the enzyme surface induced by senescence. Finally, the system of reference used to express enzyme activity should be considered.

B. ENZYME TURNOVER Senescence-induced changes in RuDPCase protein may reflect a balance of protein degradation and its synthesis. There is controversy over the turnover of RuDPCase during senescence of chloroplasts. A number of reports suggest that the protein does not have any turnover during

CHANGES DURING SENESCENCE OF CHLOROPLASTS

295

senescence (Woolhouse, 1967, 1982; Peterson et al., 1973), and only degrades without any synthesis. However, many authors offer contradictory views suggesting that the protein is synthesized during senescence (Peterson and Huffaker, 1975; Kasemir et al., 1987). Peterson and Huffaker (1975) have shown a significant incorporation of labeled amino acids into RuDPCase during senescence. A recent report of Kasemir et a/. (1987) is particularly remarkable because they have demonstrated full competence of cotyledons undergoing dark-induced senescence to synthesize enzyme protein after reirradiation. In their elegant experiments, they have shown a loss of RuDPCase activity and a decrease in the level of mRNA responsible for the synthesis of the small subunit of RuDPCase during dark-induced senescence of mustard cotyledons. The disappearance of enzyme activity is attributed to the simultaneous cessation of mRNA formation and degradation of enzyme protein. The de nouo appearance of RuDPCase could be the result of a recovery of gene expression for the synthesis of new enzyme protein. The major problem of this work, however, is the treatment of cotyledons with continuous darkness to induce senescence, a system obviously not comparable to natural senescence. It looks logical, however, to suggest a senescenceinduced modification in the turnover rate of the enzyme. Once the chloroplasts are completely mature and ready for senescence, the turnover of the enzyme should preferentially shift toward degradation (Lauriere, 1983; Lamattina et al., 1985). Reports are available on the proteolytic breakdown of RuDPCase (Lauriere, 1983; Thomas, 1984). A temporal correlation between enzyme protein breakdown and senescence-induced activity of proteases may suggest proteolytic degradation of the enzyme. A senescence-induced protease capable of hydrolyzing soluble proteins, including RuDPCase, has been demonstrated in Festuca chloroplasts (Thomas, 1982). The protease is characterized by its ability to hydrolyze different substrates, including RuDPCase protein, at different pHs during senescence (Peoples et al., 1980). Direct evidence for proteolytic degradation has been demonstrated by Thomas and Huffaker (1981) who have shown the release of a series of low-molecular-weight polypeptides from labeled RuDPCase cleaved by an endopeptidase. The substrate specificity of the protease for the degradation of RuDPCase and the kinetics of the protease activity suggest a relatively high susceptibility of RuDPCase to this degradative enzyme as compared to other Calvin enzymes. A 6-fold difference in the K, of casein and carboxylase used as substrates suggests a high affinity of the enzyme to carboxylase. If casein represents a noncarboxylase plant protein, this differential affinity may explain selective degradation of RuDPCase protein during senescence (Wittenbach, 1978).

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Although the proteolytic degradation of RuDPCase is now known to occur. controversy exists over the precise cellular location of the protease. Wittenbach et al. (1982) suggest that vacuoles may be the sites associated with the proteolytic activity that degrades RuDPCase in chloroplasts. Although a vacuolar location of proteases for chloroplast degradation has been recently questioned (Thomas, 1984), a senescenceinduced structural interaction between chloroplasts and vacuoles as revealed by electron microscopy has been shown by Wittenbach ct al. (1981). Vacuolar localization of proteases is further supported by Huffaker and Miller (1978) and Miller and Huffaker (1985) who have suggested the cytoplasm as a source of endoproteolytic activity in barley leaves. It has been suggested that the senescence-induced breakdown of RuDPCase protein may be due to the release of vacuolar hydrolases that comc into contact with chloroplast inclusions. There are, however, some difficulties in rationalizing the vacuolar concept of chloroplast degradation, which is not compatible with the senescence-induced ultrastructural changes. Most of the reports on senescence-induced structural changes suggest that the envelope is relatively stable and is ruptured at a stage when other chloroplast components (both lamellar and stromal) are degraded. Since the envelope is the barrier to the entry of proteolytic enzymes from vacuoles, it is therefore logical to conclude plastid localization of proteases. In addition, the vacuolar model does not explain the sequential degradation of soluble proteins, extrinsic proteins. and intrinsic proteins of thylakoid membranes. It is, however. possible that the structural interaction of chloroplasts with vacuoles during senescence as demonstrated by Peoples et al. (1980) and Wittenbach el al. (1982) may result in lowering the pH of the stroma or that it may provide small molecules of enzyme activator that in turn could stimulate the activity of plastid-specific proteases. Contrary to the vacuolar localization of proteascs, the degradation of protein has been demonstrated during senescence of cell-free chloroplasts (Panigrahi and Riswal, 1979a) obviously without a vacuolar system supporting chloroplast localization of proteases for the degradation of the proteins. Recent reports support the findings that chloroplasts are the likely sites of the 1984; Thomas, 1984;Wettern and breakdown of their proteins (Mae er d., Galling, 1985; Martin er al., 1986).

C . ENZYME Loss, A LIMITING FACTOR Since extensive reports are available on the senescence-induced rapid loss of the protein and activity of RuDPCase, it is quite tempting to question whether a decline in enzyme activity and/or protein could be a

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major factor limiting the photosynthetic rate of chloroplasts during leaf senescence. It is difficult, however, to establish a correlation between photosynthetic rate and enzyme activity assayed in uitro, and it is not yet possible to accurately measure the activity of the enzyme in uiuo. In uitro assays of the enzyme may not necessarily simulate the true picture of enzyme behavior in chloroplasts during natural senescence since the activity of the enzyme in uiuo is significantly modulated by different factors, namely light, pH, carbon dioxide concentration, concentration of cations such as M 2 + , and availability of the substrate (Jensen and Bahr, 1977). It is well established that light-induced transport of protons into the intrathylakoid space during photoelectron transfer is counterbalanced by the entry of Mg2+ions into the stroma. This also results in a change in pH from approximately 7 to 8. The changes in pH and Mg2+ion concentration are known to modify the activity of many Calvin enzymes including RuDPCase, and in fact these changes provide optimal conditions for the activity of RuDPCase in the stroma. Senescence-induced damage to the oxygen-evolving system and a loss of the electron-transport efficiency of thylakoid membranes, as reviewed earlier (Section IIl), could bring about a decline in proton-translocation efficiency. This in turn may retard the increase in pH and accumulation of Mg2+ ions in the stroma thereby resulting in an inhibition of the enzyme activity. This may suggest that the major factor limiting photosynthesis is the senescence-induced decline in electron-transport efficiency of thylakoid membranes of chloroplasts. Other enzymes of the Calvin cycle also control RuDPCase in uiuo by their capacity to generate RuDP; these enzymes are controlled by many factors including the electron-transport efficiency of thylakoid membrane (Woolhouse and Jenkens, 1983). Therefore, the question of whether senescence-induced changes associated with RuDPCase limit photosynthesis during senescence should be considered with caution. V. Senescence of Cell-Free Chloroplasts

The basic symptoms of senescencelike loss of pigments, macromolecules, and photochemical reactions, and a decline in the efficiency of the Calvin cycle largely remain the same whether chloroplasts undergo senescence in the leaves or in uitro. However, the organelle loses functional potential much faster in uitro than in uiuo (Misra and Biswal, 1981; Raval et al., 1982; Panigrahi and Biswal, 1983). Photosynthetic pigments, plastid-specific proteins, and nucleic acids degrade rather slowly in cell-free systems in the dark (Choe and Thimann, 1975; Panigrahi and Biswal, 1979a; Misra and Biswal, 1981, 1982a,b; Panda et

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a / . , 1986). In addition, the regulatory mechanism of senescence in vitro seems different from its counterpart in viuo as demonstrated by the differential responses of the organelle to light and other environmental factors (Biswal et a / ., 1983b). The differential regulatory mechanisms could logically be explained in terms of the absence of the nucleus and the cellular environment during senescence of isolated chloroplasts (Walker and Sivak, 1986). A. NATURE OF DEGRADATION Chloroplasts in cell-free systems quickly lose the ability to synthesize nucleic acids and proteins. This probably imposes limitations on the prolonged survival of the isolated organelle. The organelle shows an immediate loss of its carbon dioxide-fixing ability followed by the loss of photochemical reactions associated with PS I1 and PS I of thylakoid membranes (Leech, 1980; Raval et al., 1984, 1985; Inoue et al., 1986). Senescence causes the loss of membrane polypetides (Dos Santos and Hall. 1982) and denaturation of pigment-protein complexes (Brouers and Wolwertz, 1980). The denaturation of complexes of thylakoid membranes aging in uitro is correlated with the spectral changes of photosynthetic pigments. Structural disorganization of isolated senescent chloroplasts as reflected in the blue shift of the red absorption band of chloroplasts (Panigrahi and Biswal, 1983). and the decrease in the intensity and increase in polarization of chlorophyll a fluorescence (Biswal, 1987) may result in the rapid loss of the functional efficiency of the organelle. Changes in the absorption characteristics and the loss of the DCIP Hill reaction with water and exogenous electron donors during senescence of cell-free chloroplasts of sunflower leaves (Panigrahi and Biswal, 1979b) are shown in Fig. 7A,B. Light has been shown to stimulate these changes. The predominant feature of the in uitro senescence of chloroplasts is the release of free unsaturated fatty acids, the breakdown products of thylakoid lipids (Biswal er a / . , 1983b). The release of fatty acids affects the efficiency of photochemical reactions in two ways. First, the release may bring both qualitative and quantitative changes in membrane composition and second, the unsaturated fatty acid released may bind to certain sensitive sites such as the donor side of PS I, the oxygen-evolving complex, and the reaction center of PS I1 with an inhibitory effect on electron-transport efficiency (Foyer and Hall, 1980). A correlation between the activity of some enzymes, namely monogalactolipase, transacetylase, and lipases, the release of free fatty acids, and a decline in photochemical reactions including the Hill reaction, and photophosphorylation of chloroplasts has been established (Wintermans et al., 1969; Heise and Jacobi, 1973; Siegenthaler et al., 1981; Dupont and

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Siegenthaler, 1986). Further evidence that the senescence-induced release of free fatty acids results in the inactivation of th eelectron-transport system is obtained by adding exogenous free fatty acids to freshly isolated chloroplasts, whereby the fatty acids immediately produce a decline in electron-transport efficiency of thylakoid membranes (Siegenthaler and Rawler, 1977). This finding also proves that in addition to the compositional changes of membrane lipids that occur when fatty acids are released, the released free fatty acids themselves can be toxic and damage chloroplasts during senescence. Shaw et al. (1976) have demonstrated that if the binding of fatty acids is prevented, a removal of some of the lipids from thylakoid membranes does not exhibit any marked effect on the electron-transport capacity of chloroplasts. It is possible, however, that the removal of lipids up to a certain level may not affect chloroplast breakdown because the quantitative loss of plastid-specific lipids such as monogalactosyl diglyceride (MGDG) , digalactos yl dig1yceride (DGDG), and sulfoquinovosyl diglyceride correlates with the loss of chlorophyll during senescence (Koiwai et al., 1981). The major unsaturated fatty acid released during chloroplast senescence in uitro is linolenic acid, Therefore, stimulation of senescence by the exogenous addition of linolenic acid to freshly isolated chloroplasts has been used as a model system to examine the precise mechanism of action of free fatty acids on organelle senescence; the results propose the following possible mechanisms of the toxic action of fatty acids on membranes. The unsaturated fatty acids may bind to the intrinsic proteins of thylakoid membranes consequently effecting the ejection of the proteins from the interior hydrophobic core of the membrane. These proteins, being located in hydrophobic domains, are not normally subject to proteolytic degradation. The ejection of these proteins by the binding of free fatty acids makes them available for protease attack resulting in membrane damage. Plastocyanin, an intrinsic protein, is known to be released by this mechanism (Golbeck et al., 1980). In addition to this, unsaturated free fatty acids such as linolenic acid are also known to stimulate phenol oxidase. The role of free radicals in the senescence of chloroplasts is well established (Inoue et al., 1986; Panda et af., 1987). One of the mechanisms of participation of unsaturated fatty acids in the senescence of chloroplasts is their conversion to free radicals involved in membrane lipid peroxidation (Thomas, 1986).

B. LIPIDPEROXIDATION Peroxidation of thylakoid lipids is a major factor in the degradation of chloroplasts in uitro. The process, however, is relatively slow during senescence of chloroplasts in leaves. It produces a change in pigment

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400 440

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520

600 640 680 length ( n m )

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Wave

720 760

FIG.7. (A) Senescence-induced changes in absorption spectra of isolated chloroplasts incubated in isolation buffer (pH 7.8) in the light or dark. Top solid line, absorption spectrum of chloroplasts freshly isolated from sunflower leaves; dashed line, absorption spectrum of isolated chloropkdstq after 20 hours of light incubation; broken dotted line, absorption spectrum of isolated chloroplasts after 168 hours (7 days) of light incubation. Bottom solid line. absorption spectrum of chloroplasts freshly isolated from sunflower leaves; dashed

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1601

Dark

Incubation time (hr 1

line, absorption spectrum of isolated chloroplasts after 24 hours of dark incubation; broken dotted line, absorption spectrum of isolated chloroplasts after 168 hours (7 days) of dark incubation. (B) Senescence-induced changes in the rates of DCIP photoreduction with or without exogenous electron donors by chloroplasts incubated in isolation buffer (pH 7.8) in the light or dark. The initial zero time rate representing 100% equals 17.6 pmol DCIP reducedim1 of chloroplast suspension/hour with water as the electron donor. Each point represents the average of values from three experiments. (O), DPC; (O),Mn2+;(El), control.

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composition and a modification of the microenvironment of thylakoid systems (Panda et al., 1986; Wolff et al., 1986). An increase in peroxidation is correlated with senescence-induced changes in the optical behavior of pigments and a decline in electron-transport efficiency (Panda et a/., 1987). The primary agents that induce lipid peroxidation of membranes are the oxygen free radicals generated during senescence (Thomas, 1986). Superoxide (OS-) production by chloroplasts during senescence coincides with a rise in lipid peroxidation and a phase change of the membrane. This leads to disruption of membrane integrity (Barber and Thompson, 1980; McRae and Thompson, 1983; Matto ef al., 1984; Wolff et al., 1986). Lipid peroxidation is known to cause protein cross-linking and inactivation of membrane-bound enzymes. Isolated chloroplasts during aging lose their carbon dioxide-fixing capacity very fast. before any significant inactivation of the electron-transport system (Leech. 1980). This could lead to a situation in which oxygen is the most probable candidate to receive the electrons from light reactions of PS I to form superoxide radicals. Later. as senescence progresses, the oxygen-evolving complex is damaged resulting in the accumulation of oxidized species at the donor side of the PS 11 reaction center that again oxidizes superoxide to singlet oxygen, the most toxic form in damaging chloroplast membranes (Panda et ul., 1986). The formation of free radicals and the process of lipid peroxidation during senescence of isolated chloroplasts is stimulated by light. This is the reason why senescence as measured by clorophyll loss is low in cell-free chloroplasts in the dark (Panigrahi and Biswal, 1979a; Raval et al., 1984; Panda et al., 1987). Recently, photoinactivation of sites associated with the PS I electron-transport system during aging of cell-free chloroplasts was examined by Inoue et d.,(1986) who have demonstrated that PS I photoinactivation during aging, as measured by NADP' reduction with 2,6-dichlorophenolindophenoland ascorbate as electron donors, declines significantly along with a loss in the content of P700 and Fe-S centers of thylakoid membranes. These changes, however, require both light and oxygen suggesting participation of oxygen free radicals. Similar results on the damage to cell-free chloroplasts during aging in light were obtained by Dupont and Siegenthaler (1 986) who demonstrated the involvement of free radicals in senescence-induced changes in chlorophyll a , chlorophyll b, carotenoids, and in both cytochrome b ~ s sand cytochrome f. In their experiments with cell-free chloroplasts they have shown a noninvolvement of peroxidase and chlorophyll oxidase in the bleaching of pigments during senescence that is in contrast to the previous findings of Martinoia et nl. (1982)and Luthy et d.(1984) in which the participation of these enzymes in the process is considered. The nature of the protective mechanism of chloroplasts against free

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radicals during senescence is known. Since plants grow in an oxygen environment, they have developed a defensive strategy to avoid the production of oxygen free radicals as much as possible. Senescence could be the result of the failure of a defense mechanism against the toxic action of free radicals, Chloroplasts are the most sensitive organelles to free radicals since chlorophyll acts as a photosensitizer for the formation of free radicals, and oxygen is liberated in the organelle. However, under normal conditions (nonsenescent conditions), catalase, peroxidase, and superoxide dismutase in the cell keep oxygen free radicals at a very low level that does not show any significant harmful effect on normal chloroplast metabolism. Among the enzymes that scavenge active oxygen species, superoxide dismutase and peroxidase are important in regard to chloroplast degradation, since both enzymes are localized in this organelle. Catalase is not an endogenous enzyme of chloroplasts and resides mostly in peroxisomes (Foyer and Hall, 1980). Superoxide dismutase in chloroplasts is largely present in the stroma. The Cu-Zn superoxide dismutase has been shown to be located in both the lumen and stroma and the Mn superoxide dismutase has been shown to be bound to thylakoid membranes (Hayakawa et al., 1984, 1985). Both types of superoxide dismutases are effective in removing reactive oxygen species from chloroplasts. Although chloroplasts exhibit a significant level of nonspecific peroxidase activity (Foyer and Hall, 1980), ascorbic acid-specific peroxidase has been shown to be present in chloroplasts (Hossain and Asada, 1984). What happens to these enzymes during senescence? The activities of these enzymes are known to decrease during senescence (Dhindsa et al., 198 1, 1982). Dhindsa et al. (1982) have shown a correlation between a decrease in the activities of catalase and superoxide dismutase and an increase in lipid peroxidation during chloroplast degradation in uiuo. Treatment with phytohormones such as kinetin and GA inhibits the senescence-induced decline in the enzyme activity with the consequent effect of decreased lipid peroxidation. Although free radical scavengers such as ethanol, vitamin E, or diphenylisobenzolutan strongly inhibit lipid peroxidation, they do not have any effect on enzymes such as catalase or superoxide dismutase, which suggests that peroxidation is induced by free radicals generated during senescence. However, hormones prevent peroxidation by their modulation at the level of superoxide dismutase and catalase. Contrary to these findings, McRae and Thompson (1983) have reported no change in the activity of superoxide dismutase, catalase, and peroxidase during chloroplast senescence of bean leaves. They have suggested that senescence-induced damage to the thylakoid membrane by superoxide radicals in the absence of a change in superoxide dismutase activity is the result of an alteration in membrane

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structure that makes the radical generated in chloroplasts inaccessible to the scavenging enzyme, This would obviously lead to an accumulation of free radicals that produces a disruption of chloroplast structure. Most of the data on the activities of scavenging enzymes are available from in uiuo studies of senescent chloroplasts. It is possible that a decline in the activities of scavenging enzymes such as superoxide dismutase and peroxidase or their inaccessibility to oxygen radicals during senescence could result in the accumulation of reactive oxygen species with the consequent effect of chloroplast degradation in uitro. VI. Regulation of Chloroplast Senescence

The limited scope of this article does not permit the extensive discussion of all the factors that regulate chloroplast degradation during senescence. Factors such as light, inorganic ions, interorgan correlation. phytohormones, and polyamines have drawn much attention in recent days; therefore. this article briefly projects the compilation of the recent data on these factors only. A . PHOTOREGULATION

Light has been shown to either retard or stimulate senescence of chloroplasts depending on the light intensity and the plant species. In some plant systems. light at relatively high intensity results in a significant loss of photosynthetic efficiency. Photostimulation of the loss of pigments is normally attributed to their photooxidation. Lichtenthaler et a / . (1982) and Meier and Lichtenthaler (1983) have demonstrated that strong light. in contrast to low light, causes the formation of sun-type chloroplasts, which show earlier chloroplast degradation. High light intensities not only stimulate chlorophyll breakdown, but also cause ultrastructural changes and a loss of soluble carbohydrates. These findings, therefore, do not allow for the generalization that photooxidation of pigments is the primary factor in the stimulation of plastid senescence. The retarding effect of light on chlorolast degradation during leaf senescence is well recorded. It is quite logical to imagine a senescencedelaying action of light because once the energy has been invested by the plant for the formation of chloroplasts, they should remain in a functional state for as long as possible. The precise role of light in retarding the degeneration of chloroplasts is not yet known. The reports available are rather controversial and mainly present two different views: either a photosynthetic or photomorphogenic action of light. The authors who

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argue in favor of a photosynthetic control, propose the photosynthetic production of organic nutrients that retard chloroplast degradation (Thimann et al., 1977). Involvement of photosynthesis in controlling chloroplast degradation during leaf senescence has been further confirmed by the sensitivity of the light response to 3-(3,4-dichlorophenyl)1,l-dimethylurea (DCMU) (Goldthwaite and Laetsch, 1967), which inhibits electron transport, and consequently produces a loss of the formation of organic nutrients through the Calvin cycle. On the other hand, Malik and Thimann (1980) have demonstrated the participation of cyclic photophosphorylation in retarding plastid degeneration during leaf senescence. Recently, a series of papers has appeared on the photomorphogenic action of light in senescence regulation (Biswal and Biswal, 1984a; Biswal and Choudhury, 1986, 1987). These reports clearly duggest that light retards the senescence of chloroplasts as characterized by the retardation of the loss of leaf chlorophyll by phytochrome and the blue light photoreceptor. Pfeiffer and Kleudgen (1980) have shown that phytochrome not only retards chlorophyll loss, but also inhibits the loss of carotenoids and plastoquinones and inhibits that senescence-induced changes in the variable fluorescence of chloroplasts. The authors, namely Biswal and Sharma (1976), Tucker (1981), Steinitz et al. (1980), Biswal et af. (1982, 1983), and Sen et al. (1984), have shown in different plant systems that pulses of red light retard senescence to aImost the same extent as continuous white light, and the effect of red pulses is nullified with brief exposure to far-red light suggesting the participation of phytochrome. Biswal et al. (1983a) have shown that phytochrome not only retards chlorophyll loss but also maintains the ultrastructure of the lamellar system (Fig. 4). Recently Biswal and Choudhury (1986, 1987) have observed the participation of the blue light photoreceptor in retarding chlorophyll loss during senescence of the papaya leaf disk. The action of the blue light photoreceptor has been shown to be partially mediated by phytochrome. Although it is difficult to conclude whether the mode of action of light is photosynthetic or photomorphogenic, recent reports are convincingly in favor of the latter mode of action. Since senescence is an integral part of chloroplast development, the regulatory factors have to be considered within the framework of photomorphogenesis. The precise mechanism of phytochrome action in retarding chloroplast senescence is not yet known, but the control of the synthesis and degradation of RuDPCase at both the level of transcription and translation by the chromophore during the senescence of mustard cotyledons has recently been proposed by Kasemir et ul. (1987).

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B . R ~ G U L A T I OBNY PHYTOHORMONES, POLYAMINES, A N D

IONS

1 . Phytohormones Plant hormones and growth regulators reportedly modify chloroplast breakdown during leaf senescence (Misra and Biswal, 1974, 1980; Choudhury and Biswal, 1980; Naito et af., 1981; Elkinawi, 1984; Mei and Thimann, 1984; Reddy e l al., 1985; Sayeed et af., 1985; Thimann, 1985a.b; Nooden, 1986; Veierskov and Kirk, 1986). Chloroplast senescence in leaves as characterized by loss of chlorophyll is retarded by hormones such as auxins, gibberellins, and cytokinins. Abscisic acid and ethylene are well-known senescence promotors. Although gibberellins and auxins have been shown to either retard or promote senescence depending on the plant system, cytokinins have been shown to be universal senescence retardants. The involvement of cytokinins in regulating chlorophyll loss was first predicted by the work of Chibnall (1939) who demonstrated a delay in chlorophyll loss by the formation of roots on petioles of detached leaves. He suggested that a root-producing hormone might have some role in maintaining the pigment level in chloroplasts. Subsequent work of Richmond and Lang (1957) on kinetin-induced retardation of chlorophyll loss confirmed the regulatory role of endogenous cytokinins in senescence. Cytokinins not only retard chlorophyll loss, but also prevent structural disorganization of thylakoid membranes (Naito et a l . , 1981). Cytokinin treatment retards the senescence-induced decrease in the length and thickness of chloroplasts, loss in the number of grana per chloroplast, and increase in the number of plastoglobuli. These hormones are also known to inhibit senescence-induced changes in absorption (Biswal and Mohanty , 1976b) and fluorescence characteristics of chloroplasts (Biswal et al., 1979) and loss of primary photochemical reactions associated with PS I1 (Biswal and Mohanty, 1976a; N. K. Choudhury and H. Imaseki, personal communication) and PS 1 of thylakoid membranes and photophosphorylations (Biswal and Mohanty , 1978). The precise mechanism of cytokinin action in regulating chloroplast degradation is not known. However, the action of the hormones is attributed to their capacity to inhibit the activity of senescence-related proteases (Martin and Thimann, 1972; Miller and Huffaker, 1985). Cytokinins have also been reported to stimulate synthetic capacity (Naito et a / . , 1981; Woolhouse and Jenkins, 1983; Martin et al., 1986). A direct action of hormones on chloroplasts during senescence is also suggested (Misra and Biswal, 1980). Benzyladenine, a synthetic cytokinin, has been shown to act at the level of mRNA to control senescence (Watanabe and Imaseki, 1982). The hormone delays the appearance of senescencerelated proteins and disappearance of essential proteins required for

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cellular maintenance. Recent data on the molecular mechanism of cytokinin action confirm the action of hormones at the level of gene expression (Miller and Huffaker, 1985; Martin et al., 1986). The other possibility for cytokinin-induced retardation of senescence is the modulating action of the hormone at the level of membrane lipid structure. Peroxidation of thylakoid lipids is one of the factors that controls chloroplast senescence. Cytokinins have been shown to inhibit the decline in the activity of scavenging enzymes such as superoxide dismutase and peroxidase thus lowering the level of peroxidation, which leads to retardation of senescence (Dhindsa et al., 1982). Hormones also retard the senescenceinduced phase shift of thylakoid membranes consequently delaying the process (Venketarayppa et al., 1984). Gibberellic acid (GA) and indole-3-acetic acid (IAA) might be controlling senescence by increasing or decreasing the level of endogenous cytokinins or senescence-stimulating hormones such as ethylene and abscisic acid (ABA) (Evans, 1984). Recently, gibberellins were shown to retard chlorophyll loss, but their effectiveness is pronounced only in combination with cytokinins (Nooden, 1986). Abscisic acid and ethylene are known for their action in bringing about physiological modifications of the leaves, such as stomata1 closing, that results in senescence of chloroplasts as measured by chlorophyll loss (Thimann, 1985b). These hormones also change the endogenous levels of other hormones and polyamines that behave as secondary messengers and are known to modify the senescence of chloroplasts (Galston, 1983; Biswal and Biswal, 1984a). In certain systems, the senescence-stimulating action of IAA and ABA is attributed to their specific action in changing membrane permeability (Wyse et al., 1980). Ethylene is now known to control senescencerelated mRNA levels. The hormone accelerates the appearance of mRNA for the proteins responsible for senescence induction (Watanabe and Imaseki, 1982). 2. Polyamines The regulatory role of polyamines in chlorophyll loss during leaf senescence is well reported (Woolhouse and Jenkins, 1983; Biswal and Biswal, 1984a). Polyamines, namely spermine, not only retard chlorophyll loss but also stabilize thylakoid membranes (Galston and Kaur-Sawhney, 1980). Although the precise role of polyamines is not known, their action in retarding the senescence-induced activity of proteases and nucleases is suggested (Kaur-Sawhney et al., 1982; Shih et ul., 1982). Their action is also explained through their interference in the metabolism of phytohonnones such as the cytokinins and ethylene, two important categories of hormones regulating senescence (Galston, 1983;

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Preger and Gepstein, 1985). For example, Fuhrer et al. (1982) have suggested that the binding of polyamines to membranes changes the membrane conformation and leads to the induction of ethylene biosynthesis. This obviously results in stimulation of senescence. 3. Iom

Some of the inorganic ions reportedly regulate chloroplast senescence (Poovaiah and Leopold, 1973; Rao and Swamy, 1977; Thompson et al., 1982: Raval et al., 1982, 1984; Thimann, 1985a: Panda et al., 1986; Biswal and Choudhury, 1987). The senescence-delaying action of MgZt is explained in terms of its stabilizing effect on the stacking of thylakoids (Raval and Bicwal, 1984), which are known to unstack during senescence. Cations such as MgZ- and Mn2+have been shown to regulate the level of thylakoid lipid peroxidation and modify the process of chloroplast senescence (Panda et al., 1987). Lieberman and Wang (1982) have suggested that Ca” and Mg” interact with the surface of plasma membranes, rigidify them, and consequently delay the process of cellular senescence. The action of Ca” however, is not comparable with that of Mg‘ because of the specific affinity of the former to calmodulin. Calcium ion is also known to stimulate the process of senescence when it enters inside the cell (Leshem el al., 1984). The data on the senescenceinhibiting and senescence-stimulating actions of Ca2+ are rationalized with the recent findings of an ion effect on detached fern leaves in which the cation retards senescence initially followed by a stimulatory action at a later stage (B. Biswal, unpublisheddata). It is possible that at an initial stage, the membrane is not permeable to the entry of Ca”, which otherwise stabilizes the lipid bilayer surface by charge-charge interactions and delays senescence. But with the progress of senescence, the membrane disorganizes and permits the entry of cations that form complexes with calmodulin. The Ca’+-calmodulin complex inside the cell may regulate senescence in different ways. The complex may activate phospholipases that destroy the lipid structure of the membrane and convert the membrane from the liquid crystalline to the gel phase, leading to rapid cellular degradation (Leshem et al., 1984; Caffrey and Lew, 1986). On the otherhand, the Ca”-calmodulin complex has been shown to regulate senescence by its action at the level of membrane protein phosphorylation (Paliyath and Poovaiah, 1985). A decrease in Ca2+calmodulin-promoted phosphorylation may lead to the biochemical alteration of the membrane, resulting in an acceleration of senescence. Ca’ ’ -calmoddin-regulated plastid-specific kinase, namely NAD kinase, i s known to control the level of NADP in the organelle (Crane and Barr, 1986). A deficiency of Ca2+ induced by senescence may limit the

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electron-transport efficiency by producing low levels of NADP, the terminal electron acceptor in thylakoids. The precise role of Ca”, whether it is as a stimulator or a retardant of chloroplast senescence, needs more elaborative experimentation.

C. INTERORGANCORRELATION The degradation of chloroplasts during senescence of attached leaves may be influenced by the position of the leaves, which compete with other organs of the plant for space, light, inorganic ions, organic nutrients, and hormones. It is well known that chloroplast degradation as characterized by the loss of chlorophyll is delayed with the removal of developing organs such as young leaves, seeds, and fruits (Thomas and Stoddart, 1980; Grover and Sinha, 1985), suggesting interorgan regulation of chloroplast senescence. The influence of different plant organs on chloroplast formation (Biswal and Biswal, 1981, 1984b, 1985; Biswal, 1985) and degradation (Misra and Biswal, 1973; Biswal et al., 1983b) has been extensively studied in our laboratory. The intensity and qualitative pattern of the senescence of chloroplasts are altered if the organelle undergoes senescence in the leaves excised from the parental axis. Excision of leaves not only causes a differential loss of macromolecules (Misra and Biswal, 1982a) and pigments, (Misra and Biswal, 1981) but also results in an alteration in the sequence of the ultrastructural degradation of the thylakoid system (Hurkman, 1979). Since cytokinins are effective senescence retardants and excised leaves lack a root system, a major source of the cytokinins and gibberellins (Nooden, 1986), a different type of chloroplast senescence might take place in excised leaves. The interorgan regulation of chloroplast senescence in leaves could also be explained in terms of temporal changes in the distribution of nutrients during plant development. D. GENETICREGULATION Chloroplast senescence is now known to be an integral part of the developmental program of plant systems. The senescence-induced synthesis of new proteins, alteration in the pattern of isoenzymes, and control of chloroplast degradation by the nucleus definitely suggest that senescence is genetically programmed (Thomas and Stoddart, 1980; Biswal et al., 1983; Lauriere, 1983; Thomas, 1984). With the help of protein synthesis inhibitors, it was also established that senescence involves translational events (Thomas, 1975; Biswal and Mohanty, 1976b; Pjon, 1981; Biswal et af., 1983b). Although application of transcription

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inhibitors such as actinomycin D fails to arrest senescence in some plant systems (Biswal et a[., 1983b), the turning off of certain vital genes and induction of senescence-related genes are reported (Watanabe and Imaseki, 1982). In this case, different categories of mRNA are identified; one category of RNA is present in nonsenescent leaves and disappears during senescence with the simultaneous appearance of new mRNA, suggesting the involvement of genes in controlling senescence. The response of senescence to inhibitors such as cordycepin also suggests the participation of mRNA processing and transport during senescence (Takegami and Yoshida, 1975). As expected, the relative stability of DNA during senescence explains the preservation of genes that are supposed to transcribe late in senescence for the synthesis of hydrolases for the degradation of cells and organelles including chloroplasts. Reports are available that suggest the control of chloroplast senescence by nuclear genes. The relative stability of photosynthetic pigments of isolated chloroplasts as compared to the pigments of chloroplasts in leaves undergoing senescence in the dark supports this view (Panigrahi and Biswal, 1979a). Senescence of chloroplasts is associated with the loss of polysomes and inhibition of chloroplast RNA synthesis (Callow and Woolhouse, 1973). An arrest in chloroplast ribosomal RNA synthesis after the completion of leaf expansion is attributed to the loss of chloroplast RNA polymerase, an enzyme encoded by the nuclear gene that is synthesized on 80 S ribosomes of the cytoplasm. This may be the enzyme that initiates the shut down of chloroplast-specific protein synthesis (Ness and Woolhouse, 1980). A diminished RNA polymerase activity during senescence of chloroplasts has been demonstrated by Wollgiehn et al. (1976). Senescence-induced loss of rRNA and lowering of polysome levels are supported by the fact that chloroplasts isolated from fresh and nonsenescent leaves have been shown to actively incorporate radioactive amino acids and contain high levels of polysomes, but chloroplasts from senescent leaves exhibit very low polysome levels and lack any significant ability to synthesize proteins (Miller and Huffaker, 1985). Recently, however, Gracia er al. (1983), Cuello el al. (1984), and Martin et al. (1986) have demonstrated the de nuuo synthesis in chloroplasts of new proteins required for their own degradation. They have shown that organelles isolated from senescent leaves synthesize proteins that are different from the proteins synthesized by chloroplasts isolated from young nonsenescent leaves. It is possible that chloroplasts containing low levels of polysomes synthesize only a few proteins such as proteases required for their degradation during senescence (Brady and Scott, 1977). Although controversy exists over the site of synthesis of proteases, i.e., a cytoplasmic site (Peterson and Huffaker, 1975) or a

CHANGES DURING SENESCENCE OF CHLOROPLASTS

31 1

plastidial site (Woolhouse and Jenkins, 1983; Thomas, 1984), it is logical to propose the synthesis of a series of proteases, of both cytoplasmic and plastidial origin, for the complete breakdown of the organelle. Proteases of cytoplasmic origin may be required for the breakdown of the plastid envelope, the last phase of organelle breakdown during senescence. However, the possibility of the transport of proteases synthesized in the cytoplasm but acting in the organelle can not be ruled out. These possibilities lead to the following questions: How are the proteases controlled? How are the signals for protease synthesis and/or activation received? If we accept a nuclear control mechanism, then how does the chloroplast communicate with the nucleus for its own degradation? Recent studies on the interaction between the nucleus, cytoplasmic ribosomes, and chloroplasts suggest the requirement of a plastid-specific signal for nuclear gene expression for the synthesis of LHCP and the small subunit (SSU) of RuDPCase (Duysen et al., 1985; Oelmuller and Mohr, 1986), two important proteins of the organelle. The results are quite encouraging and raise the fundamental question of whether the nuclear message for chloroplast senescence is controlled by a signal from the organelle itself; the answer is not available at this time.

VII. Conclusion

A. MODELSFOR THE MECHANISM OF SENESCENCE It is difficult to propose any single mechanism of senescence in chloroplasts for two main reasons. First, it is difficult to define a senescence mechanism because of the contradictions in the accumulated data resulting from system specificity and a lack of uniformity in experimental conditions. Second, our knowledge of chloroplast senescence is still inadequate and the available data are not sufficient to develop concepts, generalize them, and propose a definite explanation of organelle degradation. However, on the basis of major findings as compiled in this review, three possible mechanisms are discussed (Fig. 8). Figure 8A proposes a loss of the efficiency of the Calvin cycle as the first major step during chloroplast senescence. The signal for the loss of RuDPCase, the crucial enzyme of the Calvin cycle, comes from the cytoplasm. A nonfunctional Calvin cycle may divert the path of electrons to oxygen to generate oxygen free radicals (Section V,B) such as superoxide that cause peroxidation of thylakoid lipids including lipids on the oxygen-evolving complex. Damage to the oxygen-evolving system

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FIG. 8. The skeletons of three possible models (A-C) of senescence mechanisms in chloroplasts. The encircled numbers indicate the step in sequence of chloroplast degradation. CF, , coupling factor; OEC, oxygen-evolving complex; O;-, superoxide; ' 0 2 , singlet oxygen, *, senenscene-induced loss of an electron-transport component. (see Figs. 5 and 6 for other abbreviations).

results in the generation of singlet oxygen and other free radicals. Free radical-induced lipid peroxidation may activate membrane-specific proteases and lipases that degrade thylakoid proteins and further the loss of membrane lipids. Alternatively, a loss of lipids may expose intrinsic membrane proteins and make them accessible to already existing active proteases. Lipid peroxidation may inactivate membrane-bound enzymes and electron-transport carriers by oxidation and cross-linking of proteins. This may ultimately lead to complete breakdown of the chloroplast. The difficulty in accepting the Calvin cycle as the first major step of chloroplast degradation as proposed in Fig. 8A is the lack of data on the activity of RuDPCase in uiuo during organelle senescence. Most of the results regarding the rapid loss of the enzyme activity is measured in uitro, which may not simulate the conditions occurring in leaves. In addition, there are data available from some plant systems that suggest that the loss of chlorophyll and the electron-transport system occurs before any significant loss of RuDPCase activity (Section IV), which would mean that a loss in the efficiency of the Calvin cycle may not be a limiting factor for chloroplast senescence. It is possible that a senescencespecific loss of thylakoid lipids triggered by a lipase is the first step of chloroplast degradation. The oxygen-evolving system has a very delicate structural organization and is highly susceptible to even mild stress

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(Section 111,C). The role of thylakoid lipids in oxygen evolution has been established (Akabori et al.. 1984).The loss of lipids may cause the release of extrinsic proteins and/or Mn" from the oxygen-evolving complex. The release of extrinsic proteins under stress and of Mn" during senescence have been reported (Section 111,C).This obviously leads to a decline in electron-transport efficiency and a change in the light-induced concentration of Mg" and pH of the stroma, which in turn affect the efficiency of carbon dioxide fixation in the Calvin cycle (Section IV,C). Figure 8B therefore proposes that th eloss in Calvin cycle efficiency is the consequence of senescence-induced damage to the thylakoid lipid structure and oxygen-evolving complex. The trigger, here is obviously acting at the level of the membrane, not at the stromal level as proposed in Fig. 8A. Figure 8C proposes the loss of some of the components of the electron-transport chain linking the oxygen-evolving complex to NADP reduction. The loss of the efficiency of the electron-transport system is attributed to the quantitative loss of quinones, reaction centers, plastocyanin, Fd-NADP reductase, or the coupling factor for ATP synthesis (Section 1II.B). This may lead to a loss of the efficiency of the watersplitting system or Calvin cycle resulting in the breakdown of the organelle. Of the three proposed schemes, the first one (Fig. 8A) seems to be more logical than the other two. The rapid and early loss of RuDPCase has physiological significance. RuDPCase protein constitutes about 50% of the total protein of green leaves, the major source of nitrogen for other growing parts of the plant system. During chloroplast senescence, the degradation products of this protein move to the growing parts. However, transport of these products needs a continuous supply of ATP. It is therefore reasonable to assume relative stability of the electron-transport system over the Calvin cycle. When the economy of the system is considered, the second target of the degenerative process obviously should be the oxygen-evolving complex, which should degrade earlier than the intrinsic proteins of the electron-transport chain. As long as the oxygen-evolving system remains intact, it logically requires a totally efficient electron-transport chain to link NADP reduction and ATP synthesis. Therefore, in the sequence, the loss of intrinsic electrontransport components should follow the senescence-induced damage to the water-splitting system.

B. LIMITATIONS A N D PERSPECTIVES The data published thus far do not provide any definite answers to the many questions relating to chloroplast degradation during senescence.

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The questions posed include the following:

I . There is no definite answer to the question of the nature of the trigger that induces chloroplast senescence, although a series of papers have appeared suggesting that the nuclear genome controls the induction. In principle this is acceptable, but the precise nature of the nuclear control over organelle breakdown needs to be elucidated. 2. The data for the sequence of degradation of the different components of chloroplasts are rather confusing. In most cases, chlorophyll loss has been used to determine senescence induction, but recent data suggest that the loss of chlorophyll is the second event. Prior to the loss of pigment, thylakoid lipids are lost resulting in the breakdown of chlorophyll. On the otherhand, the loss of the activity of Calvin enzymes, namely RuDPCase, is demonstrated before any loss of membrane organization. Similarly, it is also difficult to suggest a general pattern for the sequential loss of PS I and PS I1 photochemical reactions, which varies from species to species. We do not yet have an explanation for the species-specific factor that determines the pattern of sequence. 3 . Controversy exists over the nature and location of degradative enzymes responsible for chloroplast degradation during senescence. Some of the findings suggest that vacuolar enzymes degrade chloroplasts. This, however, makes it difficult to explain the data of a number of papers suggesting sequential degradation of chloroplasts (Section 11,C). The sequential degradation of different components is physiologically significant for the system, but it necessitates the temporal appearance of specific proteases. The problem encountered in the investigation of this proposal is the lack of precise information on the different types of chloroplast proteases. At the moment, our knowledge of the plastidspecific proteases, their locations, specificity, time-dependent activation, and de nouo synthesis is limited. Chloroplast senescence and the appearance of certain specific proteases have been reported, although these proteases are not well characterized. However, data for the proteases of RuDPCase (Section IV,B) and proteases of some of the thylakoid membrane proteins associated with PS I1 (Section 11,C) are quite encouraging and provide a basis for further study in this direction. 4. Our knowledge of the precise nature of the biochemical changes in thylakoid lipids is inadequate even though extensive reports arc available on the senescence-induced ultrastructural change of thylakoid membranes and formation of lipid globuli. 5. Although data on chloroplast protein turnover (Woolhouse, 1983) and pigments (Biswal er al., 1983a; Woolhouse. 1984b) are available, controversy exists over the turnover of certain specific proteins. For

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example, it has been suggested by many authors in the past that RuDPCase does not have any significant turnover during senescence (Section IV,B), but just degrades without synthesis. On the otherhand, recent data suggest a significant level of de nouo synthesis of this protein during senescence; particularly notable is the work of Kasemir et al. (1987). Logically, membrane polypeptides should be relatively stable compared to stromal proteins. However, a rapid turnover of the 32-kDa membrane polypeptide (Woolhouse, 1984b) needs more elaborative experimentation. 6 . Most of the literature on chloroplast degradation describes only isolated aspects of senescence such as structural modification, changes in the activities of Calvin enzymes, or loss of primary photochemical reactions of thylakoid membranes. Attempts should now be made to examine the structure-function relationships of the organelle during senescence. This obviously can provide an integrated picture of chloroplast senescence. Knowledge of the kinetics of chloroplast degradation at both structural and functional levels may help to derive some useful conclusions that will contribute to the elucidation of the basic mechanism of photosynthesis. 7. It is known that different factors, both intrinsic and environmental, control chloroplast senescence, but the nature of the integrated regulation of the process remains unknown. We do not know how the environmental signals are translated into biochemical events that ultimately regulate senescence. The question still remains as to whether light controls chlorophyll loss by modifying the level of intrinsic senescence retardants such as phytohormones and polyamines. 8. The major obstacles in attempting to gain an understanding of senescence are the limitations imposed on the study of senescence by artificial laboratory conditions. Most of the data compiled in this article on plastid degradation were generated during artificially induced senescence. For example, the process was examined in dark-induced senescence, obviously without a natural light-dark cycle. Senescence has also been studied in the leaves of seedlings grown in distilled water without any nutrients. This does not simulate natural conditions. In most cases, senescence is investigated in excised leaves or leaf disks, systems that lack regulation by the axis and other organs of the plant. These experimental designs, however, have their own logic, but with limited objectives. Further studies should focus on applying the information available on artificially induced senescence to the examination of the process in a natural system; an approach such as this will provide an accurate account

3 16

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of chloroplast senescence and eliminate the controversies arising from the use of different laboratory models. L4CKNOWLEDGMENTS

We thank all of our research colleagues in the Biochemistry group for their help during the preparation of this manuscript.

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Index

A

Actin-binding proteins, 238, 260 a-Actinin, distribution during fertilization, 259 Actinomycin D, 310 Adhesion, EPS and, 225-226 ADP, 68 ADPglucose, 67-68 ADPglucose pyrophosphorylase, 68, 69 Aemmonas, 80 Agmenelium quadrupiicatum, 70

Abscisic acid, 306, 307

Acetobacter, 205 Acetobaciw xyhum cellulose production, 190 EPS, 205, 215 gel formation, 215 N-Acetylglucosamine, 147, 149 N-Acetylglucosaminyltransferase,147 N-Acetylneuraminic acid, 198 Achromatium, 80, 85 Acid phosphatase activity, in follicular epithelial cells, 164

Agrobacterium curdlan synthesis, 210-211 succinoglycan, 206

Agrobacterium tumefaciens, EPS gel

Acinetobacter, 80 Actin fascin and, 260 in fertilization cone, 248 filaments anchorage to plasma membrane, 237-238 bundle formation, 259 distribution, 101 in follicular epithelial cells, 156, 158 line symmetry, 20 in mammalian eggs, 237-238 microtubules and, 101, 123 microvillar elongation and, 258-259 phagocytosis of luminal thyroglobulin and, 160 polarity, 22 in sea urchin egg cortex, 248 genomic expression, 137 thyroglobulin reabsorption and, 180

formation, 215

Aicaligenes PHB formation, 70 polysaccharides with common or closely related structures, 203-204 Alcaligenes eutrophus, PHB and, 71-72 Alcaligenesfaecalk var. myxogenes, 190-191, 206 Alginates Azotobacter vinelandii, 197, 206-207, 208 composition, 201 gel formation, 215-216 bacterial sources, 207-208 uronic acid sequence, 207 vs. other bacterial heteropolysaccharides, 206 Allophycocyanin, 50 Allophycocyanin E3, 50 5-Amino-4-imidazolecarboxamide ribonucleotide, 69 323

324

INDEX

Aminopeptidase N, 162 Ammonium chloride, 77 Amorphophallus konjac, 217 AMP, 68 Amphibians, cortical granule contents from,

235 Anabaena, 77,18 Anabaena cylindrica cyanophycin granules, 75-78 polyhedral bodies in, 36 Anabaena Jlos-aquae gas vesicle protein, 61-62 gas vesicles, 61 mutant, absence of GVP in, 63 polysaccharide accumulation, 66 turgor, 66 Ancylobacter aquaticus gas vesicles chemical characterization, 61,62 genetic variability, 64 GVP inhibition, 63 regulation of formation, 65 Anthoceras punctatus-Nostoc, 76 Anurans, sperm entry in, 249 AphanizomenonJos-aquae, 65 Aphanocapsa 6308 arginine catabolism, 78 CGP peptidase, 77 A priori validity, 3 Arbacia eggs alterations in membrane sterols at activation, 254 cortical granules, 235,256 fertilization cones, 245 fertilized, mobility of sperm membrane components in, 241-244 Arginase, 78 Arginine, catabolism, 78 Arginine dihydrolase, 78 Arg-poly(Asp) synthetase, 77,78 Arthmbacter srabilis, succinoglycan, 206 L-Aspartate:2-oxoglutarateaminotransferase,

37 Asiiccacaulis biprosthecum, 192 ATP:AMP phosphotransferase, 31 ATP:3-phospho-~-glycerate 1-phosphotransferase, 37 ATP synthesis, coupling factor for, 313 Atrial cells, microtubule distribution,

119-120 Autophagolysosomes, 164 Auxins, 306

Azospirillum brasilense, 75,223 A zotobacter, 70 Azotobacter chmcoccum, 196 Azotobacter vinelandii alginate, 197,206-207 EPS, 222-223,225 composition, 200 morphogenesis, 222-223

B Bacillus, 70 Bacillus cereus, glycogen synthesis and degradation, 70 &rcihs megaterium, 72 Bacillus thuringiensb, PHB accumulation and, 74 Bacterial adhesion, EPS and, 226 Bacterial surface polysaccharides capsules and related structures, microscopic appearance, 192-194 chemical composition, 197-202 definition, 187-189 exopolysaccharide, 187-188.See also Exopolysaccharide (EPS) fibers, microscopic appearance, 189-191 function adhesion, 225-226 EPS in morphogenesis and development, 222-225 recognition, symbiosis, and infection, 219-222 glycocalyx, 187 physical properties gel formation, 215-216 molecular weight, 212-215 order/disorder, 209-212 synergistic gelling, 217-219 physiological influences carbon substrate, 194-196 nutrient limitation, 196-197 structure common or closely related, 203-209 general findings, 202-203 ultrastructure, 187-188 Bacteriochlorophyll a (Bchl a), 38,40-43 Bacteriochlorophyll c (Bchl c) amino acid sequence in Chlomflexus, 44 in chlorosomes, 38-43 vs. protein content, 43-44

325

INDEX Bchl a, 38, 40-43 Beggotoo oxidation of sulfide, 86-87 PHB synthesis, 70, 75 sulfur deposition, internal, 80,82, 83 sulfur globules, 86 sulfur metabolism in, 82, 83 Be@toa alba, 75, 84,86 Bemyladenine, 306-307 Bindin, 239 Binding, 240 Biofilms, 188 Biomacromolecules, dissymmetric arrays,

5-7 1,3-Bisphosphoglycerate:polyphosphate phosphotransferase, 79 Blasia pusilk-Nostoc, 76 Bmchydanio, 249 Buddings, 147-148 Buoyancy, glycogen accumulation and, 70

microtubules and, 127-136 models of, 131 rapidly induced, 130-136 Carotenoids during chloroplast senescence, 275, 277,

282, 302 light absorption and, 273 phytochrome and, 305 Casein hydrolysate, 77 Catalase effect on oxygen free radicals, 303 trigonal form, 22

Caulobacter, 70 Centering, 11 CGP (cyanophycin granule polypeptide),

76-78 Chaetopterus oocytes, 238 Chloramphenicol, 77 Chlombium chlorosomes discovery, 38 freeze-fracture studies, 39-40 photosynthetic apparatus and,

C

Ca2+,senescence-stimulating actions of, 308 Ca2+-calmodulin complex, regulation of senescence, 308-309 Cadoxen, 212 Calcium, in cortical granules, 235 Caldesmon in follicular epithelial cells, 158-159 thyroglobulin reabsorption and, 180 Calmodulin, 159, 308 Calothrix PCC 7601, 61-63 Calspectin, 159-160 Calvin cycle control of RuDPCase, 297 efficiency of carbon dioxide fixation, 313 enzymes RuDPCase. See RuDPCase in stroma lamellae of chloroplast, 273 loss of efficiency in chloroplast senescence, 311-312 CAMP, 69 Carbonic anhydrase, 37 Carbon substrate, effect on bacterial surface polysaccharides, 194-196 Carboxymethylcellulose, 214 Carboxysomes, 36-38 Cardiac hypertrophy development, 127-128

38-39 structural parameters, 38-39,40 Chlombium limicola chlorosomes, 43 Chlorobium vesicles, 38.See also Chlorosomes

Chlomflexus Bchl c polypeptides, 44 chlorosome attachment site, 43 chlorosomes freeze-fracture studies, 39-40 structural parameters, 40 Chlomflexusaumntiacus chlorosomes physicochemical studies, 41 SDS-PAGE analysis, 41, 42 structure, function and development,

38-39 Chlomgloeopsis frischii carboxysomes, 37 polyhedral bodies, 36 Chlorophyll light absorption and fluorescence, 273,

275 loss during chloroplast senescence, 275-277 Chlorophyll a fluorescence, 284, 298 light absorbed by, 56 loss, 288 senescence-induced changes, 302

326

INDEX

Chlorophyllase, 275, 276 Chlorophyll b loss, 281, 288 senescence-induced changes, 302 Chlorophyllide, 276 Chloroplasts grana, 278 lamellae, 278 lamellar system, 273, 274 membranes, electron-transport chain of, 286 senescence. See Chloroplast senescence structure, 273-275 Chloroplast senescence carbon dioxide-fixing capacity and, 302 carotenoid loss, 277 in cell-free chloroplasts, 297-304 nature of degradation, 298-299 chlorophyll loss, 275-277, 287 degradation during, 315 lamellar, 281 nature and location of enzymes, 314 sequence, 314 disorganization symptoms, 271 fluorescence and, 305 limitations and perspectives from current data, 313-316 lipid loss during, 277 loss of electron-transport efficiency during, 287-289 macromolecule loss, 277-278 manganese loss, 290 mechanism, models for, 311-313 membrane phase change, 285 in mesophyll cells, 271 optical changes, 283-284 primary photochemical reactions, loss of damage to oxygen-evolving complex, 289-290 photoelectron transport system, 285-287 photophosphorylation loss, 291 protective mechanism against free radicals, 302-303 protein turnover, 314-315 regulation, 304-311, 315 genetic, 309-311 interorgan correlation, 309 by ions, 308-309 by light, 304-305 by photohormones, 306-307

by polyamines, 307-308 RuDPCase activity, loss of, 296-297 enzyme turnover and, 294-296 in mature vs. senescent wheat leaf protoplasts, 292, 293 mechanism of loss, 292-294 structural modifications, 273-285 superoxide production during, 302 trigger, nature of, 314 ultrastructural modifications, 272, 278-283 and biochemical changes, 271-316 sequence, 280 Chlorosomes description, 38-44 proteins from, 41-43 Cholesterol, intramembranous protein particles and, 156

Chromatiurn arrangement of intracellular sulfur, 83 inclusions, 85 internal sulfur deposition, 80 oxidation of sulfide to sulfur, 86 oxidation of sulfur, 87 sulfur deposits, internal, 83 sulfur globules, 84-85 Chromatiurn 8111, internal sulfur deposits, 83 Chromatiurn vinosurn, 83, 86 Chromatiurn warmingii, 83 Chromophores, in phycobiliprotein, 48-49 Ciona eggs, 234 Clathrin coat of coated vesicles, 18-19, 20 Clostridiurn, gas vesicles, 60 Coated vesicle cages, 21 Colchicine folliculogenesis inhibition, 177 inhibition of TSH phagocytosis-inducing effect, 160 myocyte beating and, 126 thyroxine inhibition, 127 ['H]Colchicine-binding, in myocardial cell extracts, 107 Colloidal iron hydroxide binding, 241 Colloid thyroglobulin, reabsorption of, 155-163 Concanavalin A, 237, 241 Connexons, 19 Cordycepin, 310 Cortical granule membrane-plasma membrane fusion, 250

INDEX Cortical granule protcases, 257 Cortical granule reaction, endocytosis after,

260-261 Cotyledon chloroplasts, during senescence,

279-281

Coupling factor (CF,), 287 Crcatine kinax, 137 Cross-polarization magic angle spining, of PHB, 74 crystals 2 4 11 3D, 11 definition, 10 Cubic point lattice, 13 Curdlan, synthesis, 210-211 Cyanobacteria buoyancy regulation, 65-66 cyanophycin granule polypeptide, 76-78 cyanophycin granules, 75-78 gas vesicles, 60. 65 glycogen granules, 67 PBS, linker polypeptides, 48, 51-55 “phosphate overplus” phenomenon, 80 photosynthetic apparatus in, 45 photosystem I, 56 photosystem 11, 56, 57 phycobiliproteins, 48-51 phycobilisomes, morphological types, 45-47 polyhedral bodies in, 36 polyphosphate accumulation, 79 thylakoid membrane, 45, 46 unbalanced growth, CGP accumulation and, 77

Cyanophom pamiom, 44

Cyanophycinase, 77 Cyanophycin granule polypeptide (CGP),

76-78 Cyanophycin granules. in prokaryotic cells,

75-78 CYW rWoluto-Annboennsymbiosis, 76-77 Cyclostome yolk crystal, 27 Cyrinus, 249 Cysteine, 77 Cytochalasin B actin filament formation and, 247 colloid phagocytosis inhibition, 160 cortical elevation formation and, 250 cortical localization of meiotic spindles,

239

folliculogenesis inhibition, 177 formation of cortical elevations and, 249

327

sperm-egg fusion and, 246 treatment of sea urchin eggs, 248 Cytochrome b,,,, 302 Cytochrome f,302 Cytokinins inhibition of scavenging enzymes, 307 plastoglobuli and, 282 regulation of chlorophyll loss, 306 source of, 309 Cytophaga, 223

D dBcAMP, 124 dbcAMP, 160 DCMU (3-(3,Cdichloropheny1)-1,1dimethylurea), 305 Density distribution, 26. 28 3 - k O X y - P ~ M O O C t ~ Oacid ~ C(KDO),198 Desmin filaments in intermyofibrillar space of Z disk

area, 121-122 microtubule distribution, 119 in microtubules, 121 myocyte heterogeneity and, 131-132 Z disks and, 122

aesulfovibriogigas, 78 Deuterium oxide, inhibition of TSH phagocytosis-inducing effect, 160 3J ‘-Diaminobenzidine tetrahydrochloride, 153

2,6-Dichlorophenolindophenol,302 3-(3,4-Dichlorophenyl)-l,ldimethylurea (DCMU),305 Dictyostelium, 237 Diffraction, 16 Digalactosyl diglyceride (DGDG), 299 Dihedral symmetry, 8 Diiodotyrosine, I55 Diphenylcarbazide (DPC), 288-289 Diphenylisobenzolutan, 303 Dissymmetric biomacromolecules, micromorphology, 2-4 Dissymmetric monomers, I5 DNA loss in chloroplast during senescence,

277-278

during senescence, 310 “sticking,” in carboxysomes, 37 DPC (diphenylcarbazide). 288-289

328

INDEX

E Ectothiorhodospira,80 Egg cortical structure, 233-239 Egg plasma membrane alterations, fertilization and, 233 changes attending the cortical granule reaction, 252-257 integration with sperm, 240-244 lipids, 253 Eggs conductance, increase at fertilization, 239-240 fertilization endocytosis, 260-261 microvillar elongation, 257-260 interactions and fusion with sperm, 239-250, 261-262 chronology of early events, 239-240 fertilization cone formation, 244-250 surface at fertilization, cortical granule reaction, 250-252 Electron diffraction, 18 Electron micrography, crystal projections, 26, 28 Electron microscopy, of microtubule distribution in cardiac myocytes, 116, 118-120 ELISA, quantitation of tubulin in cardiac myocytes, 107-108 Enantiomers, 3 Enantiomorphic, 3 Endocytosis, during fertilization, 260-261 1,4-Endo-P-glucanases, 214 Endostylic cell, peroxidase in, 154 Enterobacter aerogenes XM6, 218 Erwinia, virulence, EPS synthesis and, 219 Erwinia amyIovora EPS, production, 195-196 wilting and, 220 Escherichia coli EPS, 202 sialic acid production, 208 genes for biosynthetic enzymes, 69 K52, 198, 199 K14 antigen, 198 K40 capsular polysaccharide, 200 K29 strain, 192, 193, 210 K28 strain, conserved backbone structure, 208 K12 strain, transduction tests, 69

phosphate transport, 79 Pit system, 79 Pst system, 79 serotypes, 198 stringent response, 69 Ethanol inhibition of TSH phagocytosis-inducing effect, 160 lipid peroxidation and, 303 Ethylene, promotion of chloroplast senescence, 306-308 Excitation energy transfer, 56-57 Exocytosis of cortical granules, 250 Exopolysaccharide (EPS) 0-acetyl groups and pyruvate ketals, 198-199 adhesion and, 225-226 alginates, 207-208 definition, 187 effect of anions on transition, 210 electron microscopy, 190 functions, 226 recognition, symbiosis and infection, 219-222 molecular weight, 212-215 morphogenesis and development role, 222-225 multiple acylation of sugars, 199-200 physical characteristics, 226 physical properties, order/disorder, 209-212 physiological conditions and, 196-197 physiologic conditions and, 188, 189 production, 188 by Erwinia amylovora, 195-196 scanning electron microscopy, 188 structure, general findings, 202-203 synergistic gelling, 217-219 synthesis, virulence and, 219 transmission electron microscopy, 188 wilting and, 220 wilting and, 220

F

Fascin, 260 Fd-NADP reductase, 313 Ferredoxin-NADP reductase, 288 Fertilization or-actinin distribution during, 259 cortical granule reaction, plasma

329

INDEX membrane changes after, 252-257 early processes, alterations in egg plasma membrane and, 233 endocytosis, 260-261 microvillar elongation, 257-260 Fertilization cone, formation, 244-250 Festuca chloroplasts, senescence-induced protease, 295 Festuca pratensis, 288 Filipin, 156 Fodrin, 248, 259 Folliculogenesis, inhibition of, 177 Forskolin, 160 Fourier space, 17 Freedom of aggregation, vs. rigid symmetry, 7 FRS, 288 Fructose-1,6-biphosphate, 68

D-Fructose-1,6-biphosphate D-glyceraldehyde-3-phosphate-lyase, 37 Fructose 6-phosphate, 68 Fucose, 149 Fundulus, 249

G GA (gibberellic acid), 303, 307 Galactomannans, 217 Galactose, 149 @-Galactosidase,165 Galactosyltransferase, 149 Gap junction, 19 Gastmlonium coulteri, 41 Gas vesicle protein (GVP), 65 Gas vesicles formation, self-assembly, 62-63 functions, 59-60 buoyancy, 64 genes, 87 genetic variability, 63-64 morphology, 60 occurance, 60-61 in prokaryotic cells, 59-66 properties, 59-60 proteinaceous shell of, 62 protein subunit, 61-62 Gel formation, 209-210, 215-216 Gellan, 203-204, 218 Gelling, synergistic, of bacterial surface polysaccharides, 217-219

Gibberellic acid (GA), 303, 307 Gibberellins, 306, 309 Glactolipids, 282 Glide plane, 15-16 Gloeobacter violaceus, 45-46 ’ Glucagon, intracellular crystals cubic interpretation of, 23 frequency distribution of exocytotic glucagon quanta, 23, 24 rhombic dodecahedral morphology, 21, 23 @1,3-Glucanase, 235 (1,4-a-D-Glucan:1,4-a-D-glucan 6a-Dg1ucano)-transferase, 68 Glucose-1-phosphate adenyltransferase, 67-68 1,4-a-D-Glucosyl-glucan,68 P-Glucuronidase, 165, 235 Glyceraldehyde 3-phosphate dehydrogenase, NADP-dependent, 291

D-Glyceraldehyde-3-phosphate:NAD+ oxidoreductase, 37 Glycerol, in EPS, 200 Glycocalyx, definition, 187 Glycogen genes, 87 granules, in prokaryotic cells, 67-70 inhibitors, 69 Glycogen synthase, 68 Glycolipids, 282 Glycoproteins, in carboxysomes, 36-37 Gram-negative cells, strength of cell walls, 66 Grana, 273 Griffhsia pacifica, 45 GTP, 69 PP gene, 65

H Halobacterium halobium gas vesicle genetic variability, 64 gas vesicle protein, 62 Halobacterium salinarium, gas vesicle genetic variability, 64 Halobacterium, gas vesicles, 60-61 Heavy meromyosin, 156, 158, 248 Heteropolysaccharides alginates, 206 major components of, 198 transmission electron microscopy and, 188

330

lNDEX

Hexagonal point lattices, 13 Homopolysaccharides, 188, 198 Hormones, lipid peroxidation and, 303 Horseradish peroxidase-diaminobenzidine reaction, 241 Hyaline protein, 235 Hydrogen peroxide, thyroglobulin iodination and, 154 3-Hydroxybutanoic acid, 199 0-Hydroxysterols, 254 Hypertrophy, cardiac. See Cardiac hypertrophy

polysaccharides, 210 type I polysaccharide, 209 Klebsiella aerogenes EPS, 202-203 K54, conserved backbone structure, 208-209 K54 polysaccharides, gel formation by, 218, 219

L

Lnctobacillus casei subsp. rhamnosus, 195 I

Im fndvle-3-acetic acid), 307 "'1 autoradiography, of thyroglobulin

iodination, 152-153 Ilyanassa oocytes, 238 Immunolabeling technique, for analysis of microtubule distribution in cardiac myocytes, 108-1 10 Inclusions definition, 35 as metabolic machinery carboxysomes, 36-38 chlorosomes, 38-44 phycobilisomes, 44-59 prokaryotic cells, future research, 87-88 Indole-3-acetic acid (IAA), 307 Intercalated disks, absence in hypertrophied myocytes, 135 Intermediate filaments, microtubules and, 120-122 Intermyofibrillar spaces, microtubule distribution, 119 Interorgan regulation, of chloroplast senescence, 309 Iron hydroxide, colloid, 241 Isomyosin heavy chain genes (MHC), 131 Isomyosins, 132 Isoprenaline, 126

K Kinetin, 303 Kinetin-induced retardation, of chlorophyll loss, 306 Klebsiella EPS, molecular weight, 212-213

Lamellae, stroma, senescence-induced disappearance, 288

Lamellipodia, 238

iuri,IKI

Light-harvesting chlorophyll-protein complex, 275 Line symmetry, 9 Linker polypeptides, in cyanobacterial PBS, 48, 51-55 Lipid bilayer membrane, 1 Lipid peroxidation, 312 Lipids loss in chloroplast senescence, 277 of thylakoid membrane, 273 Lipoxygenase, chlorophyll degradation and, 276 Lytechinus eggs interaction with sperm, 239-240 ionic activation, 249 membrane capacitance of, 240

M

Macromolecules, loss during chloroplast senescence, 277-278 Macromonas, 82 r-Malate:NAD+ oxidoreductase, 37 Mammalian eggs association of actin with meiotic chromosomes, 239 cortical granule contents from, 235 fertilization and development, 237-238 fusion of, sperm plasma membranes in, 249-250 meiotic spindle development, 237 plasma membrane of, 236

INDEX Manganese, loss during chloroplast senescence, 290 Mannosyltransferase, 147 MAP,, 137 MAP,, 137 MAPS. See Microtubule-associated proteins Mastigocladus laminosus phycocyanin, 49-51, 57 MCDG (monogalactosyl diglyceride), 299 Medaka eggs, microvillar elongation, 259 1-Methyladenine, 260 Methylmercaptoimidazole, 153 0-Methyl sugars, in EPS, 200 5-Methyltryptophan, 77 Mg” change in light-induced concentration, 313 senescence-delaying action of, 308 MHC, isoforms, 131 (Y-MHC, 131 0-MHC, 131 Micromorphology definition, 1 discrete macromolecular models for, 1 dissymmetric biomacromolecule, 2-4 symmetric aggregates, 1-2 microscopy, 16-18 symmetric arrays, symmetric elements, 4-7 Micropinocytosis of colloid thyroglobulin, 157 reabsorption of colloid and, 155-156 schematic representation, 158 Microtubule-associated proteins distribution in myocardium, 116-118 function, 101 identification in cardiac tissue, 105-107 MAPS, 105-107 MAP,, 122 subunits, 136 tubulin identification in cardiac tissue, 103-105 Microtubules assembly mechanisms, control of, 101 in cardiac hypertrophy, 127-136 from cardiac tissue and isolated myocytes, purification of, 102-103 colloid phagocytoid activity of follicular epithelial cells and, 160 composition, 101 constitutive elements, 136-137 contraction rates of myocytes and, 124, 126

331

distribution in cardiac myocytes at electron microscopic level, 116, 118-120 immunolabeling using light microscope, 108-110 at light microscopic level, 110-120 methods to improve ultrastructural analysis, 109-110 in different mammalian species, 122 in isolated myocytes, 110-112, 113, 114 in myocardium, 112, 115, 116 functions, 101, 123-126 heterogeneity, 137-138 inhibitors, 160 interconnections with desmin filaments, 122 intermediate filaments and, 120-122 line symmetry, 20 myocyte hyperplasia and, 127 myogenesis and, 128-129 during postnatal cardiac development, 130 during rapidly induced cardiac hypertrophy, 130-136 rat cardiac hypertrophy model, 101 in receptor internalization, 126-127 roles in cardiac myocyte, 123-136 in sarcomerogenesis, 127-136 shape preservation of cardiac myocytes and, 123-124, 125 Mirror plane, 15 Mirror symmetry, 16 Mitochondria, symptoms of degradation and senescence, 271 Mn’+, thylakoid lipid peroxidation and, 308, 313 Monensin, effect on thyroglobulin transport and secretion, 150 Monoclinic point lattice, twofold rotational symmetry, 13 Monogalactosyl diglyceride (MCDG), 299 Monoiodotyrosine, 155 mRNA brain tubulin microheterogeneity and, 137 processing and transport during senescence, 310 Mucopolyaccharides, sulfated, in cortical granules, 235 Myocardium distribution, of microtuble-associated proteins, 116, 117, 118

332

INDEX

microtubule distribution in, 112, 115, 116 Myocytes, cardiac contraction rates, microtubules and, 124, 126 exhibiting dense microtubule pattern, 133 hyperplasia, microtubules and, 127 hypertrophy. See Cardiac hypertrophy isolated, microtubule distribution in, 110-112, 113, 114 microtubules in, 101-138 subpopulations, 130 quantitation of tubulin in, 107-108 receptor internalization, microtubules and, 126-127 shape preservation, microtubules and, 123-124, 125 Myofibrillogenesis, 123, 128, 135 Myonemes, 128 Myosin in egg, 248 postnatal genomic expression. 137 Mytilus cortical granules. 235 Myxobacteria, role of EPS in morphogenesis and development, 223

N Neisseria meningitidis, 208 Nibrobacter agilis carboxysomes, 31 polyhedra1 bodies, 36 Nitmbacter winogmdrkyi, polyhedral bodies, 36 Nitmomonas, polyhedral bodies, 36 Nocodazole MT depolymerization and, 112, 113 spontaneous myocyte beating and, 126 Nodulation, EPS and, 220-221 Noncrystallographic point symmetry, 20 Nostoc muscoturn filaments, 65 Nutrient limitation, effect on bacterial surface polysaccharides, 196-202

0

Optical changes, during chloroplast senescence, 283-284 Orthorhombic point lattices, 13

Oxygen-evolving complex during chloroplast senescence, 289-290 damage during senescence, 311-313 structural components, 289, 290

P

t! cruentum 8-phycoerythrin, 51 PBS. See Phycobilisomes Periodic aggregate, three-dimensional, 17-18 Perivitelline space, 251 Peroxidase chlorophyll degradation and, 276 in cortical granules, 235 effect on oxygen free radicals during nonsenescent conditions, 303 localization of, 150 thyroglobulin iodination and, 153-154 in thyroid-homologous cell, 154 pH, intracellular, actin filament bundle formation and, 259 PHA/phospholipid/fatty acid (PLFA), 72 Phareolus leaves, senescent chloroplasts of, 282 PHB. See Poly-8-hydroxybutyric acid PHB depolymerases, 73 PHB synthetase, 73 pH changes, RuDPCase activity and, 297 phosgene, 79 Phosphate, in EPS structure, 198 “Phosphate overplus” phenomenon, 80 3-Phosphoglycerate, 68 Phosphoglycerate kinase, 291 Phosphoriboisomerase, 37 Phosphoribulokinase, 37, 291 phoT gene, 79 Photophosphorylation, loss during chloroplast senescence, 291 Photoregulation, of chloroplast senescence, 304-305 Photosynthesis, oxygenic, Z scheme for, 56 Photosynthetic bacteria, accumulation of PHB, 74-75 phoW gene, 79 Phycobilins, 273, 275 Phycobiliproteins cyanobacterial, physical and chemical factors affecting, 58 properties, 48-51 Phycobilisomes genes, 87

INDEX hemidiscoidal, 54, 55, 56 morphology, 44-47 properties, 50-58 proteins, 47-49 structural genes, 58-59 Phycocyanin, 49, 50, 51, 57 Phycocyanobilin, 48-49, 57 Phycoerythrine, 50 Phycoerythrobilin, 48-49 Phycoerythrocyanin, 50 Phycourobilin, 48-49 Phytochrome, 305 Phytohormones. See also specific phytohormones effect on senescence-induced lipid peroxidation, 303 polyamines and, 307 regulation of chloroplast senescence, 306-307 Phytol, 276 Pi, 68 Pinocytosis, 180 Pinocytotic pit, 155, 156 Pit system, 79 Plane groups, 11 Plant cell, temporal disorganization of different organelles, 271-272 Plasmalemma, cortical granule and, 236 Plastocyanin, 288, 299, 313 Plastoglobuli, structural status of thylakoid membranes and, 282 Plastoquinones inactivation, 289 phytochrome and, 305 in thylakoids and plastoglobuli, 282 Pleurodeles, 249 PLFA, 72 Point groups description, 18-20 icosahedral, 9 Point lattices defining vectorial translation in two- and three-D space groups, 10 description, 16-17 symmetry, 11, 13 with three dimensions, 12 Point symmetry, 8 Polarity, structural, 3 Poly-0-alkanoates, 72 Polyamines, regulation of chloroplast senescence, 307-308

333

Polyglucose granules, in prokaryotic cells, 67-70 Polyguluronk acid, 216 Poly-8-hydroxyalkanoates,72 Poly-8-hydroxybutyrate granules, 35 Poly-8-hydroxybutyric acid depolymerases, 73 Poly-b-hydroxybutyric acid (PHB) apparent density, 72 granules, in prokaryotic cells, 70-75 quantitation, 73-74 recalcitrance, 75 synthesis and degradation, 74-75 Poly-0-hydroxybutyric acid synthetase, 73 Poly-6-hydroxyheptanoate,72 Polyphosphatases hydrolyze phosphosphates, 79-80 Po1yphosphate:AMP phosphotransferase, 79 Polyphosphate-dependent NAD’ kinase, 79 Polyphosphate glucokinase, 79 Polyphosphates granules in prokaryotic cells, 78-80 synthesis and degradation, 79-80 Polysomes, loss of, during senescence, 310 Poly-&valerate, 72 hrphyridium cruentum, 47 PPi, 69 Prochloron, polyhedral bodies, 36 Projection theorem, 18 Prokaryotic cells bacterial surface polysaccharides. See Bacterial surface polysaccharides inclusions, 35 carboxysomes, 36-38 chlorosomes, 38-44 environmental adjustment and, 59-66 future research, 87-88 gas vesicles, 59-66 as metabolic products, 67-87 cyanophycin granules, 75-78 polyglucose granules, 67-70 poly-8-hydroxybutyric acid granules, 70-75 polyphosphate granules, 78-80 sulfur granules, 80-87 phycobilisomes, 44-59 intracellular turgor pressure, 66 hsthecochloris, chlorosomes, 38, 39 hosthecomicmbium pneutnaticum, 64 Proteases, 311 220-kDa Protein, 260

3 34

INDEX

Protein macromolecules, loss in chloroplast senescence, 277-278 Proteins in chloroplast stroma, 273 chlorosome, 41-43 microtubule-associated. See Microtubuleassociated proteins phycobilisome, 47-49 in phycobilisomes, 45 thylakoid membrane, 275

Aeudomonas alginates, gel formation, 216 carbon substrate availability and, 195 PHB and, 70, 71 polyphosphate accumulation, 80 polysaccharides with common or closely related structures, 203-204 Aeudomonas aeruginosll alginate, 207-208 composition, 201 EPS composition, 199-200 Pseudomonas atlantica, 225 Pseudomonas elodea, 218 Aeudomonas jluorescens alginate, 207 EPS composition, 200 Pseudomonas mendocina alginate, 207 EPS composition, 200 Pseudomonas putida alginate, 207 EPS composition, 200 Aeudomonas solanacearum, binding by lectin, 221-222 Pseudomonas syringae var. glycinae, alginate, 207 Aeudomonas thermophilia K2, polyhedral bodies in, 36 Pseudosymmetry, 2, 7 pst gene, 79 Pst system, 79 Purkinje cells, microtubule distribution, 119-120 Pyruvate, 68

Q Quinones, loss of, 313

R Rat cardiac hypertrophy model, 101 Reaction centers, 313 Real space, 17 Reciprocal lattice points, 17 Reciprocal space, 17 relA gene, 69 RER, 179 Rhizobium capsulation, 192, 194 curdlan synthesis, 210-211 EPS composition, 200 gel-forming, neutral hexasaccharide repeat unit of, 215 synthesis, nodulation and, 220-221 polysaccharides, 205, 222 Rhizobiumjapmicum capsulation, 192, 194-195 EPS nodulation and, 221 Omethyl sugar content, 200 PHB and, 74 Rhizobium Ieguminosarum Congo red dye uptake, 190 EPS, gel formation, 215 polysaccharides, 199 with common or closely related structure, 205 Rhizobium melitoti EPS composition, 200 gel formation, 215 Rhizobium phaseoii Congo red dye uptake, 190 EPS, gel formation, 215 polysaccharides with common or closely related structure, 205 Rhizobium trfoiii cellulosic fibrils, 190, 191 EPS, 198-199 gel formation, 215 nodulation and, 221 production in carbon-limited cultures, 197 polysaccharides with common or closely related structure, 205-206 Rhodobacter, 70 Rhodomicrobium vannielii, polyhedral bodies in, 36

INDEX Rhcdopseudomonas sphaemides, 71 Rhombohedral point lattices, 13 Ribose 5-phosphate isomerase, 291 Ribululose 1,Sdiphosphate carboxylax. See RuDPCase Rifamycin. 77 RNA, loss in chloroplast during senescence, 277-278 Root-producing hormone, 306 Rotational symmetry characteristics, 6 n-fold, 3, 5 in point symmetry group, 8 Rough endoplasmic reticula, of follicular epithelial cells, 147-148 rRNA, senescence-induced loss, 310 Rubisco, 36, 37 RuDPCase cellular location, 2% loss during chloroplast senescence, 278, 291-297, 311-312, 314 rapid and early loss of, 313 susceptibility to degradation, 295 synthesis and degradation, 305 in thylakoid membrane, 287

S Sabellaria, cortical granule reaction, 252 Salmonella typhimurium, 64,69 Sarcomere, microtubule distribution, 119 Sarcomerogenesis, microtubules and, 127-136 scmv axis, 9 Screw symmetry, 6 Sea urchin eggs alterations in membrane sterols at activation, 254-255 concanavalin A-binding sites, 255 cortical granules, 235 membrane, intramembranous particles on P face of, 256 cortical structurg 233-239 ultrastructure, 234-235 disruption of actin microfilaments, inhibition of surface activity, 248 fertilization cones, 245. 248 surface area measurements, 245 fertilized, mobility of sperm membrane components in. 241-244

335

inseminated, fertilization cone of, 245-246 mobility of sperm membrane components in fertilized egg, 241-244 plasma membrane, 236-237 modifications, 235-236 postinsemination,miftwillarelongation, 258 unfertilized, difference in number of intramembranous particles, 255-256 Secrrtion granules,number of molecules in, 25

Sedoheptulose-1.7-bisphosphate 1-phosphodydrolax, 37 Self-assembly, of gas vesicles, 62-63 Senescence, 271. See a h Chloroplast senescence Serine protease, in cortical granules, 235 Sialic acid, 149, 208 Sialyltransferase, 149 Space groups description, 22-29 one-dimensional, 10 symmetry, 11 three-dimensional, symmetry, 13-14 two-dimensional, 10-11 Space symmetry, 9-14 Spcctrin, 159. 248 Sperm integration with egg plasma membrane, 240-244 interaction and fusion with egg, 239-240 fertilization cone formation, 244-250 interactions and fusion with egg, 261-262 Sphaemtilus, 70, 75 Spirilum, 70. 75 Spirulina platens&, CGP accumulation, 77 spinrla eggs changes in distribution of intramembranous particles, 257 concanavalin A binding, 241 microvillar elongation, 258 Stigmntella aumntiaca, fruiting body formation, 223, 224 Stn?pwcocm mutans, 215 Stn?pwcoccuspneumoniae EPS. 200. 202 polysaccharide, 212 Streptomym viridochmmogenes, 70 Stmngylocentmrus concanavalin A-binding sites, 255 cortical structure, 233-239 spiral of electron-dense material, 235

lNDEX egg endocytosis, 261 total surface area of cortical granule membranes, 257-258 Structural analysis, symmetry of moleculalr packing, 18 Structure principles of line groups, 20, 22 of point groups, 18-20 of projected view symmetry, 14-16 of space groups, 22-29 of symmetric aggregate microscopy, 16-18 Structure research dissymmetric biomacromolecule, 2-4 symmetric arrays, rigid symmetry vs. freedom of aggregation, 7 Succinoglycan, 206 Sulfide, oxidation to sulfur, 86 Sulfoquinovosyl diglyceride, 299 Sulfur density, 83 hydrophilic forms, 83 internal deposits in bacteria, 80, 81, 82 orthorhombiG 83 Sulfur dioxide, CGP accumulation, 77 Sulfur globules characterization, 86 extraction, 84 in prokaryotic cells, 80-87 Superoxide dismutase in chloroplasts during senescence, 303 cytokinins and, 307 Superoxide production, by chloroplasts during senescence, 302 Superoxide radicals, senescence-induced damage to thylakoid membrane, 303-304 Symmetric aggregates microscopy of, 16-18 self-assembly of, 14 Symmetry dihedral, 8 line groups, 9 at macromolecular level, 16 point groups, 8 of projected view, 14-16 rigid vs. freedom of aggregation, 7 space groups, 9-14 Synechmysris, linker polypeptides ftom, 53. 55

Stmn&xenrmtus

Synechococcus linker polypeptides from, 51-54 PBS, 56 PBS structural encoding genes, 59 phycocyanin, 49, 51 Synechocystis energy transfer process in, 57 PBS, 56 excitation energy transfer, 56-57

T TI. See Thyroxine T, . See 'Ifiiodothyronine Tau factors, 105 k o l , 102 Tetragonal point lattice, 13 Tetraiodothyroninc See Thyroxine (TI) Thiamine pyrophosphatase, 148 Thiobacillus polyhedral bodies, 36 sulfur synthesis, 80 Thiobacillus intermedius carboxysomes, 37-38 Thiobacillus neapolitanus carboxysomes of, 36-37 polyhedral bodies in, 36 Thiobacterium, 82 Thiocopu, internal sulfur deposition, 80, 85 Thiocystis, internal sulfur deposition, 80, 85 Thiocystis violacea, sulfur deposits, internal, 83 Thiomicmspim, sulfur synthesis, 80 Thioploca, internal sulfur deposition, 80, 82, 85 Thiospim, 82 Thiospirillopsis, 80 Thiorhrix, 80, 82, 85, 87 Thiovulum, 82, 85 Thiovulum w'us,internal sulfur deposits. 83 Thylakoid membrane of cyanobacteria. 45, 46 decline in electron-transport efficiency, 299 light-driven electron transport, 287 lipids, 282 breakdown products of, 298 membrane p h a x change and, 285

INDEX peroxidation of, 299-304,307. 312 role in oxygen evolution, 313 Mg"-ATPax, senescence-induced loss, 291 photoelectron transport in, 287 photosystem I, 285 disassembly of, 287-289 phytohormones and, 306 pigments, 283-284 supcroxide radical formation and, 302 photosystem 11, 285-286 accumulation of oxidized species, 302 disassembly of, 287-289 phytohormones and, 306 pigments, 283-284 pigments, photosystems of, 285 proteins, 275 senescence-induced changes, 276 breakdown, 281-282 stacking status. 284 temporal disorganization, 287 ultrastructural. 314 Thyroglobulin biochemical characterization, 147 in blood circulation, 162-163 colloid fusion of reabsorbed droplets, 163-164 luminal, phagocytosis. 160 reabsorption, 155-163 reabsorption of, 155-163 hydrolysis, and release of T, and T,, 163-166 iodination, 152-155, 179 molecular size, 147 release, 147-151 storage, 179 sulfation, 149 synthesis, 147-151 transport and secretion, effects of cooling and monensin, 150 tyrosyl residues, 154-155 Thyroglobulin-secretory granules micropinocytotic features, 156 peroxidase content, 151 structure, 149 Thyroid gland anatomy and physiology, 145-146 capillary endothelial fenestrations, I80 connective tissue space and vascularization, 166-171 endostylic cell, pcroxidase in, 154 endothelial cell fenestrations, 167, 169-170

337

exocytotic vesicles, 151 follicle structures, need for, 179-180 follicular epithelial cells, 145-146 acid phosphatase activity, 164 actin filaments in, 156, I58 galactosylation of protein, 149 hormone secretion and, 179 inversion of cell polarity, 174-175 morphology, 180 phagocytosis of luminal colloid, 160 polarity and inverted follicles. 172-179 pseudopods, 160 regulation of micropinocytosis. phagocytosis and exocytosis, I59 secretory activity, 180 transcytosis, 162 ultrastructure and cytochemical functions, 146 zonula occludens between, 173-174 inverted follicles, 160-161 cell t w , 174-175 collagen gel culturing, 176-177, 178 zonula occludens. 175 nerve supplies, 171-172, 180-181 reabsorption of colloid, 155-163 secretory process of thyroid hormones, 145-146 synthesis and release of thyroglobulin, 147-151 Thyroid hormone, 131 Thyroid-stimulating hormone (TSH) colloid reabsorption and, 160 regulation of thyroglobulin transport and release. 150 thyroglobulin reabsorption, 179 Thyroxine (T,) entrance into circulation, 166 and hydrolysis of reabsorbed thyroglobulin, 165 liberation from basal part of cell, 180 secretion of, 145 synthesis, 145 synthesis and secretion, 179 Tonoplast breakdown, 283 'Transcytosis. 162 'Translation, 5-6 'Translational periodicity, I I 'Translation vectors, 9 ZVchodesmium erythmm, 76 Piclinic crystal, cubic pseudosymmetry, 12 'Ificlinic point lattice+ symmetry, 11-12

338

INDEX

Trigonal point lattices, 13 Triiodothyronine (T,) release from basal part of follicular epithelial cell, 166, 180 secretion, 145, 179 synthesis, 145, 165, 179 Trturus eggs, 234 TSH. See Thyroid-stimulating hormone lhbrbifx, 237 lbbulin identification in cardiac tissue, 103-105 isolation and purification, 102-103 in microtubules, 101 in postnatal development of heart and brain, 137 quantitation in cardiac myocytes, 107-108 P-lbbulin genes, 105 P-lbbulin isoforms, 136 Tubulin labeling, patterns in myocytes, 132 5 p e 021N Thiothrix-like activated sludge bacteria, 80

U Urea, 77 Urechis cortical granule reaction, 252 sperm-egg interaction, 239 Uridine diphosphatase, 149 Uridine diphosphate, 149 Uridine monophosphate, 149

W Wilting, EPS and, 220

X Xanthan acyl groups and, 218 conformation, 211-212 molecular weight, 212-213 reversible disaggregation of, 212-213 from Xanthomonas carnpestris, 191, 197, 202 molecular weight, 214 Ahthornonas, 204, 205 Xanthornonas campestris polysaccharide, 204-205 xanthan, 191, 197, 202 molecular weight, 214 Xenopus eggs cortical granule reaction, 251 plasma membrane, 236

Y Yolk platelet crystals electron diffraction patterns, 30 orthorhombic, 31 a priori analysis, 28-29 structure models, 23-24, 26

V Z

Vasoactive intestinal peptide (VIP), 172 Vicia faba chloroplasts, 282 Vinblastine, inhibition of TSH phagocytosis-inducing effect, 160 Vincristine, inhibition of TSH phagocytosis-inducing effect, 160 Vinculin, 238 Vitamin E, 303 Vitelline layer, 257 Virus capsids, 20

Z disks absence in hypertrophied myocytes, 135 desmin and, 122 microtubule distribution, 119 Zonula occludens of inverted thyroid follicle, 175 between thyroid follicular cells, 173-174 Zoogloea, gel formation, 215 Z scheme for oxygenic photosynthesis, 56