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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