SECTION ONE—
PLANT CELL STRUCTURE AND FUNCTION

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Figure 1.1
A diagram of an undifferentiated cell showing the principal components. Some
of the constituents are illustrated by only a few examples (e.g. ribosomes). The
components may be identified by letters which refer to those given in the text.
Figure taken from Ultrastructure and the Biology of Plant Cells by B.E.S.
Gunning and M.W. Steer, published by Edward Arnold, London. The
drawing was generously provided by Dr. Steer and is reproduced
with the kind permission of the authors and copyright holders.

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Introduction

The term cell, as first used by Robert Hooke in 1665 signified an apparently empty space or lumen, surrounded by walls. We now know, of course, that the space is far from empty, and that rigid cell walls as seen by Hooke in thin slices of cork, are not ubiquitous in multicellular organisms. Indeed, the wall became to be regarded as the definitive structure of the cell, and when in the 1830s, the zoologist Schwann was able to recognise structures in cartilage resembling plant parenchymatous cell walls, the concept of the cell as the basic biological unit common to all organisms was born. Definitions have changed considerably in the subsequent century and a half, and, in particular, the cell wall is now seen in its proper perspective as being a structure, albeit of great importance, but restricted to plants and existing only outside the true cell. Nevertheless, the general concept of the cell as the basic minimlum unit of life remains.

Since all organisms need to perform a number of essential functions merely in order to survive, both as individuals and as species, it should not be surprising to find a basic unity between the cells of all organisms. Each cell, at least in the early stages of its development, possesses the capacity to synthesize complex substances from simple ones, to liberate and transform the potential chemical energy of highly reduced compounds, to react to internal and external stimuli, to control the influx and efflux of materials across the limiting cell membranes and to regulate its activities in relation to the information contained in its individual store, or stores, of hereditary genetic material. Evolution has solved the problems posed by these requirements in more or less identical ways in all organisms, and thus the basic processes, activities, and structures of each individual plant cell are similar, not only to other plant cells, but also to all other eucaryotic cells. This book concentrates on the unifying features of plant cells and relates them to present knowledge and general theories of molecular biology. It should not be forgotten, however, that cells are characterised as much by their diversity as their unity. A wide range of different cell types with varying specialized functions are necessary for the life of the higher green plant; however, the origin of cell heterogeneity is a topic outside the scope of this present book.

The basic structures of an undifferentiated plant cell can be seen in Fig. 1.1. The cell proper is delimited by the plasma membrane (or plasmalemma ) which is of unit membrane construction (chapters 2 and 8). Outside the plasma membrane, and thus actually extra-cellular, is the cell wall (chapter 1). The cell wall is normally closely appressed to the plasma membrane and in meristematic cells is thin and relatively weak. During differentiation various specialized

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wall structures develop; depending on the function of the mature cell, the walls may become relatively massive and extremely strong through the deposition of rigid, highly cross-linked polymeric substances. Adjacent protoplasts (i.e. the cells proper) are connected across the cell walls by narrow cytoplasmic channels, bounded by the plasma membrane, known as plasmodesmata (PD).

Within the cell a number of separate compartments, and interconnecting compartments, delimited by membranes, may be recognised (chapter 8). Vacuoles (V) are prominent, apparently empty spaces, spherical and numerous in the meristematic cell but irregular, very large, and coalescent in the mature expanded cell. Vacuoles serve as intracellular dust-bins—repositories for unwanted and often toxic byproducts of metabolism—and may also have functions similar to the lysosomes of animal cells. They are bounded by a single membrane known as the tonoplast (chapters 2 and 8).

The nucleus (N) (chapter 9), a major compartment in most cells, comprises a nuclear envelope possessing many large nuclear pores (NP) and nucleoplasm, the ground substance in which the hereditary material, chromatin, and the nucleolus (NU) lie. The nucleus is the principal site of the hereditary material of the cell, although both plastids and mitochondria also contain DNA. The material outside the nuclear envelope is commonly known as cytoplasm.

Ramifying throughout the cytoplasm, and occasionally connected to the outer membrane of the nuclear envelope, the cisternae of the endoplasmic reticulum act to integrate the biosynthetic functions of the cell (chapter 8). The endoplasmic reticulum is generally classified into two types: rough endoplasmic reticulum (RER), which has ribosomes attached to its outer face (chapter 10); and smooth endoplasmic reticulum (SER) which is not involved in protein synthesis. The endoplasmic reticulum may also, on occasion, be seen to be associated with stacks of vesicles (VE) known collectively as dictyosomes (D) or Golgi bodies. The endoplasmic reticulum and the dictyosomes are responsible for the formation and secretion of cellular membranes.

Three other membrane-bound compartments remain, each concerned with an aspect of energy or intermediary metabolism. Plastids (P), undifferentiated in meristematic cells and present only as proplastids, represent a general class of organelle in which the chloroplast is the characteristic member (chapters 3 and 4). Mitochondria (M) are smaller, but also bounded by a double membrane, and similarly involved in energy metabolism (chapter 5). As mentioned above, both mitochondria and plastids contain their own stores of hereditary material (chapter 11. The final compartments, in contrast, are bound by only a single membrane and do not contain hereditary material; these are known as microbodies (MB) and often contain dense, granular, or even crystalline contents (chapter 6). Within the cytoplasm just inside the plasma membrane lie long narrow cylinders known as microtubules (MT); microtubules function in a number of processes in which orientation of cellular components is important (chapter 7). Finally, plant cells contain many fine fibrils, known as

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microfilaments, which appear to be contractile in function and to be composed of a material similar to actin, one of the contractile components of muscle.

The 'typical' plant cell does not exist, of course, and the meristematic cell shown in Fig. 1.1 has only been chosen since it possesses all the essential characteristics of plant cells. Many of the cellular components are only present in very simple forms in meristematic cells, however, and the subsequent chapters in Section I necessarily involve a consideration of a variety of more specialized cell types.

Chapter 1—
Plant Cell Walls

1.1—
Introduction

Plant cell walls establish a home, and indeed a city, for plant protoplasts. They serve many specialized functions in plant tissues, and form the skin, the skeleton, and the circulatory system of plants.

There are many variations in the form and substance of plant cell walls. The walls may be plastic or they may be rigid, permeable or impermeable, impregnated with plastics or coated with slime, cemented in layers to form fibres or dissolved in spots to form pores. These variations are of vital importance to the proper biological functioning of plant cells and organs, and thus the structure of the cell wall is often our best indication of the nature of the protoplast which dwells inside.

1.2—
The Molecular Structure of Plant Cell Walls

Polysaccharides are the principal components of all plant cell walls. The polysaccharides of the cell wall are made up of sugars which are linked to each other by glycosidic bonds to form the polymer chains. Each polysaccharide contains particular kinds of sugars which are joined to each other in characteristic patterns of linkage position and sequence. It is now known that the secondary, tertiary and quaternary structures of cell wall polysaccharides are determined by the structures of the component sugars and the linkages between them, just as the three dimensional structure of a protein is determined by the sequence and structures of its component amino acids (Rees, 1972).

The various polysaccharide chains of the plant cell wall are connected to each other in specific ways, and they form an integrated network. The properties of this network depend not only on the amounts, characteristic properties and orientations of the individual polysaccharides, but also on the nature and frequency of the interconnecting linkages between them.

The conformational structures of the nine sugars commonly found in plant cell walls are shown in Fig. 1.1. The three types of polysaccharide normally found in plant cell walls (cellulose, hemicelluloses, and pectic polysaccharides), and the structural protein of primary walls, are described briefly below.

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Figure 1.1
Sugars of plant cell walls.
Conformational line drawings indicate approximate bond angles. b -L arabinose is shown in its
preferred planar furanose ring form. The other sugars are shown in their most stable pyranose
chair form. Carbon atoms are numbered as indicated for b -D -glucose. Ring hydrogens are
indicated by bonds only. Note that groups attached to a ring may be either axial (projecting
above or below the ring) or equatorial (projecting to the side of the ring). Substituents at C₁
project equatorially in the b configuration, but are axial in the a configuration. All 'bulky'
groups (–OH, –CH₂ OH, & –COOH) are in equatorial positions in b -D -glucose, b -D -
glucuronic acid, and b -D-xylose. Note that these sugars differ only in the group
attached to C₅ . Galactose, galacturonic acid and fucose are similarly related
(axial –OH group at C₄ ), as are mannose and rhamnose (axial –OH group at C₂ ).

1.2.1—
Cellulose

Cellulose occurs as a crystalline, fibrillar aggregate of b -1,4-linked glucan chains (Frey-Wyssling, 1969). Cellulose fibrils give plant cell walls most of their enormous strength, much as glass fibres embedded in an epoxy resin give strength to a fibreglass composite (Northcote, 1972).

The basic structure of the b -1,4-linked glucan chains of cellulose is illustrated in Fig. 1.2 by conformational line drawings and in Fig. 1.3 by molecular models. Residues of b -D -glucose (Fig. 1.1) are glycosidically linked to each other, from carbon 1 of one residue to carbon 4 of the adjacent residue. The upside-down inversion of every second residue in the chain minimizes contact between atoms of adjacent residues. Close inspection of the models in Fig. 1.3 shows that the –OH groups at carbon 3 are in very close proximity to the ring oxygens (O₅ ) of adjacent residues. Hydrogen bonds between O₃ and O'₅ help to stabilize the flat, straight, ribbon-like structure of b -1,4-linked glucan chains.

The flat, ribbon-like structure allows the chains to fit closely together, one on top of the other, over their entire lengths. These interchain associations are stabilized by hydrogen bonds between O₆ of a glucose residue in one chain and

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Figure 1.2
b -1,4–linked glucan chains of cellulose.
Portions of two associated chains are illustrated by
conformational line drawings. Distances between atoms
are not accurately indicated in this illustration, but see Fig. 1.3.

Figure 1.3
Hemicellulosic xyloglucan associated with cellulose.
The repeating subunit of a hemicellulosic xyloglucan is shown in
association with a portion of a b –1,4 –linked glucan chain of cellulose
(Bauer et al., 1973). Molecular models have been used to accurately indicate
interatomic distances and bond angles. Hydrogen bonds from the cellulosic
glucan chain to the glucan backbone of the hemicellulose are indicated by arrows.

the oxygen of the glycosidic bond (O₁ ) between glucose residues in an adjacent chain. Since the glucan chains of cellulose are very long (8,000 to 15,000 residues), the number of hydrogen bonds between adjacent chains is very large. The resultant crystal is extremely stable and so tightly packed that there is no room for water molecules in the crystal structure.

Although there is some controversy as to whether native cellulose fibrils are 3.5 nm or 10 nm in diameter, it is clear that the glucan chains of cellulose

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aggregate to form stiff crystalline rods of very considerable length and mechanical strength. The molecular structure of b -1 ,4-linked glucose thus neatly determines the secondary and tertiary structures of cellulosic glucan chains, and establishes the quaternary, interchain associations—although not the dimensions—of the microfibrils. The stiff, crystalline rods of cellulose are clearly well suited to their biological function in the plant cell wall.

1.2.2—
Hemicelluloses

Xylans, arabinoxylans, galactomannans, glucomannans and xyloglucans are common types of hemicelluloses (Timell, 1965). The basic repeating sequence of a hemicellulosic xyloglucan molecule is shown in Fig. 1.3, adjacent to a cellulosic glucan chain.

Although different hemicelluloses have different component sugars, all have two structural features in common which bear importantly on their biological function. (1) All hemicelluloses have straight, flat b -1,4-linked backbones. Any side chains attached to the backbone are short—usually just one sugar long—and stick out to the sides of the backbone (cf. Fig. 1.3). (2) All hemicelluloses have some structural feature which prevents the chains from extended self-aggregation of the type which exists between the b -1,4-linked glucan chains of cellulose. Xyloglucans, for example, have a b -1,4-linked glucan backbone (just as cellulose), but most of the glucose –CH₂ OH groups (C₆ ) are substituted with xylose side chains (see Fig. 1.3) and are thus not available for the formation of interchain hydrogen bonds. Similarly, since xylose is a pentose (i.e. a five carbon sugar), the b -1,4-linked xylose backbones of xylans and arabinoxylans have no–CH₂ OH groups available for interchain hydrogen bonding. The glucomannans have b -1,4-linked backbones containing both glucose and mannose. The –CH₂ OH groups of these sugars are unsubstituted, but the axial conformation of the –OH groups at carbon 2 of the mannose residues (cf. Fig. 1.1) prevent close interchain associations wherever mannose residues occur in the backbone.

Although hemicelluloses cannot self-aggregate to form long, close-packed crystalline fibrils in the manner of cellulose, the chains of hemicellulosic polysaccharides can form important hydrogen bonded associations with each other in the cell wall, particularly between regions of the chains which have few side branches or axial hydroxyl groups (Blake & Richards, 1971; McNiel et al., 1974). Even more importantly, hemicelluloses can co-crystallize with cellulosic glucan chains at the surface of the cellulose microfibrils (Bauer et al., 1973, Northcote, 1972). The cocrystallization probably involves the formation of hydrogen bonds from the –CH₂ OH groups present in cellulose chains to the glycosidic oxygens in the adjacent hemicellulose chains (see Fig. 1.3). This association would form a tightly bound monolayer of the hemicellulose on the surface of the cellulose microfibril, and would function as part of the 'glue' which holds the microfibrils together in the cell wall.

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1.2.3—
The Pectic Polysaccharides and Structural Protein

1.2.3.1—
Rhamnogalacturonans

The rhamnoglacturonans are long polymers of a -1,4-linked galacturonic acid interspersed with a few residues of 1,2-linked rhamnose (Aspinall, 1973). There is some evidence that the rhamnosyl residues may occur at definite positions in the galacturonan chain, giving a subunit structure to the rhamnogalacturonan polymer (Talmadge et al., 1973; see Fig. 1.4).

Figure 1.4
Pectic rhamnogalacturonan.
CPK models illustrate the repeating subunit of a rhamnogalacturonan,
with a short sequence of b -1,4-linked galactan attached to C₄ of one
of the rhamnosyl residues (Talmadge et al., 1973). The sequence
of the subunit is GalUA₈ Rha GalUA Rha GalUA₄ .

The diaxial conformation of the a -1,4 glycosidic linkages between galacturonic acid residues (see Fig. 1.1) causes the orientation of adjacent rings to be twisted. Thus, the galacturonan polymer forms a tight, stiff, rod-like helix with three residues per turn (Rees & Wight, 1971; Fig. 1.4). The insertion of 1,2-linked rhamnosyl residues in the galacturonan chain creates 'kinks' or right angle bends.

Divalent cations, particularly calcium, form complexes with the carboxyl and hydroxyl groups of galacturonic acid residues in the polymer. Complex formation of this type occurs primarily between adjacent residues in a galacturonan polymer, but could serve to create ionic ligand bridges between adjacent galacturonan chains.

1.2.3.2—
Arabinogalactans

Two distinct types of arabinogalactans are known to occur in plant cell walls (Aspinall, 1973). The first type has a b -1,4-linked galactan backbone with

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highly branched arabinose side chains. The second type has a b -1,3-linked galactan backbone with many short side chains containing galactose and arabinose.

1.2.3.3—
Structural Protein

Primary cells walls (i.e. the type of walls characteristic of actively growing cells) contain a structural protein component. While the structural protein of these walls has not yet been isolated as an intact molecule, the analysis of peptide fragments from the primary walls of dicotyledonous plants has revealed several interesting characteristics (Lamport, 1970; Lamport et al., 1973). This structural protein contains over 25% hydroxyproline, which is an unusual amino acid known to break the continuity of a -helical structures. In animals, hydroxyproline occurs almost exclusively in the proteins of connective tissue (collagen and gelatin). Several tryptic peptides of the structural protein have been isolated,

Figure 1.5
Interconnections between cell wall components.
Schematic representation of the polymeric components of
sycamore primary cell walls and their interconnections (Keegstra et al.,
1973). Hemicellulosic xyloglucan polymers are cocrystalized with cellulosic
glucan chains on the surface of the (two) microfibrils. The reducing ends of some
—but not all—of the xyloglucan chains are glycosidically attached to some—but not
all—of the b -1,4-linked (arabino) galactan side chains of the rhamnogalacturonan polymers.
The rhamnogalacturonan polymers and the structural protein are interconnected by 1,3-linked
(arabino) galactan bridges. Arrowheads indicate the reducing ends of polysaccharide chains.

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and all appear to contain the sequence Ser-Hyp₄ . Arabinose tetrasaccharides are glycosidically linked to virtually all of the hydroxyproline residues in the protein. In addition, many of the serine residues are glycosylated with galactose. As a result of this extensive glycosylation, the structural protein is very resistant to degradation by proteases, and is likely to have an extended, rod-like shape.

1.3—
Interconnections between Cell Wall Components

The polysaccharide components of the primary cell walls of dicotyledonous plants are specifically attached to one another to form an interconnected network (Keegstra et al., 1973; see Fig. 1.5). The b -1,4-linked glucan chains of cellulose are aggregated to form crystalline microfibrils. The hemicellulosic xyloglucan chains cocrystallize with the cellulose chains on the surface of the microfibrils as described above (1.2.2). The reducing ends of the xyloglucan chains are covalently (glycosidically) attached to b -1,4-linked arabinogalactan. The arabinogalactan chains, in turn, are glycosidically linked to the backbone of the rhamnogalacturonan, probably to the 4 positions of the rhamnosyl residues. The 1,4 arabinogalactan is thus a side chain of the rhamnogalacturonan, and forms an interconnecting bridge between the xyloglucan and the rhamnogalacturonan. The other type of arabinogalactan (b -1,3-linked) also seems to serve as an interconnecting bridge, glycosidically linking the reducing ends of rhamnogalacturonan chains to amino acid residues in the structural protein of the cell wall.

1.4—
The Universality of Plant Cell Wall Structures

Studies of cell walls isolated from suspension-cultured plant cells have demonstrated quite clearly that dicotyledonous plants from taxonomically diverse species have very similar primary walls (Albersheim, 1974). Similar studies on the primary walls of aspen cambial tissue and potato tuber tissue have shown that these walls are quite similar to the walls of suspension-cultured cells (Timell, personal communication; Bauer, unpublished results). Thus, all dicotyledons probably have the same basic primary cell wall.

The primary cell walls of monocotyledons have a structure which is somewhat different from that of dicotyledons, although the walls from various monocotyledon species appear to be quite similar to each other. The hemicellulose of the primary monocotyledon walls is an arabinoxylan instead of a xyloglucan, and the structural protein appears to contain very little hydroxyproline (Burke et al., 1974).

Most cells, after they stop active growth, synthesize 'secondary' cell wall material which is quite different from the primary wall. Secondary cell walls

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consist almost exclusively of large amounts of cellulose and one (or more) of the hemicellulosic polysaccharides. Lignification is very common for cells with secondary walls.

Within broad taxonomic groups, related plants appear to have very similar cell walls. However, it should be recognized that relatively few types of cell walls from relatively few plant species have been carefully analysed. The diversity of specialized cell wall functions is likely to be reflected in a diversity of cell wall structures.

1.5—
Cell Wall Plastics

1.5.1—
Lignin

Lignin is a biological plastic formed in plant cell walls by the enzymic dehydrogenation of coumaryl, coniferyl and synapyl alcohols (Fig. 1.6) followed by a free radical polymerization (Freundenberg, 1968; Northcote, 1972). Since the polymerization is not enzymically controlled, and the monomeric free radicals can react with each other in a variety of ways, lignin does not have a unique structure. However, the lignin from a particular plant species or tissue does usually have a characteristic monomer composition. Some covalent linkages are formed between lignin and the polysaccharides of the cell wall during lignin biosynthesis (Morrison, 1974).

Figure 1.6
Lignin precursors.
R₁ = H, R₂ = H, coumaryl alcohol.
R₁ = H, R₂ = OCH₃ , coniferyl alcohol.
R₁ = OCH₃ , R₂ = OCH₃ , synapyl alcohol.

Lignin formation is initiated very soon after secondary wall synthesis begins, and proceeds from the region of the middle lamella (the pectin-rich layer between adjacent cell walls) inward towards the plasmalemma. Thus, both the primary and secondary walls become fully impregnated with a rigid, hydrophobic plastic which is covalently linked to the polysaccharide matrix. The resultant structure is extremely strong and resistant to degradation.

1.5.2—
Cutin

Cutin is a biological plastic which coats the cell walls on the outer surface of plant epidermal cells. Although little is known about the detailed structure of

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cutin, it is clear that the principal monomeric components of this material are mono-, di-, and trihydroxyfatty acids (C₁₆ –C₁₈ ). These hydroxyfatty acids are linked to each other mainly through ester bonds, although the presence of both ether and peroxide bonds have been reported (Kolattukudy & Walton, 1972). The formation of ester linkages between monomeric hydroxyfatty acids and preformed cutin appears to involve the enzymic transacylation of the hydroxyfatty acids from a Coenzyme A intermediate to free hydroxyl groups in cutin (Croteau & Kolattukudy, 1974).

The film of cutin polymer is impregnated—and frequently coated—with a complex mixture of waxes. The cutin-wax structure (cuticle) merges gradually with the normal polysaccharide components of the epidermal cell wall. There is evidence for the existence of hydrophylic channels (ectodesmata) in the hydrophobic cuticle (Franke, 1967). These channels may occur in morphologically distinct patterns on the epidermal surface.

The cuticular waxes and the cutin polymer form a tough hydrophobic skin over the surface of the plant which is important in minimizing water loss and preventing mechanical injury or invasion by pathogens.

1.6—
Biosynthesis of Plant Cell Walls

It is generally believed that cell wall polysaccharides are synthesized from the appropriate sugar nucleotides (e.g. UDP-glucose) by specific enzymes or enzyme complexes. Many—though not necessarily all—cell wall polysaccharides are synthesized in the golgi bodies (dictyosomes) and transported to the wall in golgi-derived vesicles which are able to fuse with the plasmalemma (O'Brien, 1972). The cell wall polysaccharides are then presumably interconnected while in the wall.

There are many reports of isolated enzymes or particulate complexes which incorporate sugars from the sugar nucleotides into polysaccharide material. However, the amount of polysaccharide material formed by these in vitro enzyme systems is often a miniscule fraction of the amount formed in vivo. Thus the relationship between these isolated enzymes and cell wall biosynthesis is uncertain.

Isolated enzyme preparations have been reported to synthesize alkaliinsoluble b -1,4 glucan ('cellulose') at rates comparable to the rate of in vivo cellulose synthesis (Rollit & Maclachlan, 1974). However, these enzymes have not as yet been shown to form the microfibrils characteristic of cellulose. There is some evidence that cellulose microfibrils are synthesized at the plasmalemma rather than in the golgi (Bowles & Northcote, 1972), perhaps by plasmalemmafused vesicles which contain rows of membrane-bound, microfibril-synthesizing particles (Kiermayer & Dobberstein, 1973).

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1.7—
Cell Wall Ultrastructure

1.7.1—
Cellulose Microfibrils

One of the most important and intriguing aspects of plant cell wall ultrastructure is the orientation of the cellulose microfibrils within the wall (Albersheim, 1965). Cellulose microfibrils are deposited (or possibly synthesized) at the inner surface of the wall, adjacent to the plasmalemma. They have a particular orientation when deposited. On the other side of the plasmalemma there is a thin layer of microtubules. The orientation of the microtubules in this layer exactly parallels the orientation of the cellulose microfibrils being deposited on the opposite side of the plasmalemma (Newcomb, 1969). Disruption of the microtubules (e.g. by colchicine) affects the orientation of the microfibrils.

In the primary walls of elongating cells, the most recently deposited cellulose microfibrils (those closest to the plasmalemma) lie parallel to the plasmalemma and are oriented perperndicular to the long (growth) axis of the cell, like the hoops which hold a barrel together. Cellulose microfibrils deposited at earlier times (and thus further from the plasmalemma) are still parallel to the plasmalemma, but are oriented at decreasing angles (i.e. more nearly parallel) to the growth axis of the cell. It is as though the process of cell wall elongation pulls the ends of the microfibrils towards the ends of the cell. In the end walls of a cell, or in the newly forming cell wall created by cell division (i.e. the cell plate), the cellulose microfibrils again lie parallel to the plasmalemma, but are randomly crossed and not parallel to each other.

The cellulose microfibrils in secondary walls are usually deposited in discrete, concentric layers. The microfibrils in a given layer are all parallel to each other, but are oriented at a considerable angle to both the microfibrils in adjacent layers and to the long axis of the cell. This cross-hatching of microfibrils in adjacent layers of secondary walls adds considerably to the overall strength of the wall, and is the same principle used in the manufacture of fibreglass-epoxy fishing rods, where great strength for weight is essential.

Secondary wall material may also be deposited in very localized regions to form highly specialized structures such as the rings or spirals of secondary wall found in xylem cells. In such cases the microtubules on the inside of the plasmalemma have the same localization and orientation as the cellulose microfibrils being deposited on the opposite side of the plasmalemma.

1.7.2—
Specialized Structures

Many of the modifications of cell wall structure which occur are related to communication or transport between cells. In cells with primary walls, intercellular communication is facilitated by plasmadesmata. Plasmadesmata are small pores of approximately 40 nm diameter which extend through the wall between two adjacent cells (Ledbetter & Porter, 1970; Fig. 1.7). The plasmalemma

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Figure 1.7
Plasmadesmata of plant cell walls.
Plasmadesmata (Pd) in the end walls of collenchymal cells of a wheat
stamen filament. The 'core' material in the plasmadesmata can be seen
as an electron-dense dot in the centre of the plasmadesmatal pores.
(See Ledbetter & Porter, 1970. Reproduced courtesy of the authors.)

of the cell(s) also extends through these pores, so that the cytoplasm of one cell is continuous with that of adjacent cells. There appears to be a core of electron dense material in the center of the plasmadesmatal tubes (see Fig. 1.7). This core may be a specific (though as yet speculative) structure capable

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Figure 1.8
Pits in tracheids of Taxus canadensus.
A full bordered pit (BP) can be seen in cross-section in the upper left portion of the figure.
The faint line in the centre of the pit is the disc. The pits lie between tracheids (Tr), which are
long, cylindrical, cytoplasmless tubes that carry water from the roots to the leaves. The primary
cell walls and middle lamella can be seen as electron dense material running in a thin line between
cells and in the corners where 3 or 4 cells meet. Layering of the thick secondary wall material is
evident. Several half-bordered pits (BP/2) between tracheids and ray cells (RC) can be seen.
(See Ledbetter & Porter, 1970. Reporduced courtesy of the authors.)

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Figure 1.9
Collenchymal cells.
The walls of these cells are unusually thick primary walls, heavily
hydrated and unlignified. Cellulose microfibrils can be seen throughout
the walls. These cells elongate very rapidly, and the walls are quite plastic.
(See Ledbetter & Porter, 1970. Reproduced courtesy of the authors.)

of regulating what can and cannot be passed between cells. Regardless of the nature of the core, it is clear that the plasmadesmata are an aspect of plant biology which has no obvious counterpart in animal tissues. Each cell may have thousands of plasmadesmata, either randomly scattered as individual

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pores or grouped in distinct fields where the primary wall is often thinner than usual.

In cells with secondary walls, and particularly in tracheids, intercellular communication and transport is facilitated by pits (Ledbetter & Porter, 1970; Fig. 1.8). In the full, bordered type of pit, the primary wall and middle lamella are largely dissolved. The hole thus formed is surrounded and partly overgrown by depositions of secondary wall material which create a raised, overhanging border. The primary wall and middle lamella in the pit is replaced by a relatively impermeable disc or torus of thickened wall material. The disc is suspended by an easily permeable, radial network of cellulose microfibrils. The diameter of the disc is slightly larger than the aperture of the overhanging pit border. Thus the disc can act as a valve to close the pit when pressed against the border by a large pressure differential (Albersheim, 1965). Gas bubbles, which would break the flow of water through the tracheids, can be sealed off by the action of such valves.

Where the tracheids are adjacent to ray cells, the pits are differentiated only on the tracheid side, giving half-bordered pits (Fig. 1.8). The walls of the ray cells in the pit region appear to remain intact or only slightly modified.

A further important example of a cell wall modification which facilitates intercellular communication or transport is the sieve plate of primary phloem. The phloem cells have primary cell walls with many plasmadesmata. Some of the plasmadesmata of the crosswalls between adjacent phloem cells enlarge considerably and become lined with callose (a b -1,3 glucan) on the wall side of the plasmalemma. The pores of the sieve plate thus formed become filled with a fibrous protein (which may be analogous to the core material of plasmadesmata).

Many other specialized cell wall structures or modifications could be described (see also Fig. 1.9). Several examples may be mentioned just to indicate the range of possibilities: the walls of pollen cells, which are so indestructible that they are used by archeologists to characterize the flora of dwelling sites many thousands of years old; the walls of bark cells, made largely of suberin, a wall material similar to cutin that can form a protective coating over wounds so as to prevent water loss or pathogen invasion; and the walls between adjacent endodermal cells, which have very dense Casparian strips which prevent the movement of ions through the walls and into the xylem stream.

The wall structures and modifications described in this section may all be seen with the microscope—there are undoubtedly many other important differentiations of the cell wall which we cannot see.

1.8—
Hormonal Control of Cell Wall Biosynthesis and Differentiation

We know that cell wall biosynthesis and the modification of cell walls are important aspects of cellular differentiation, and that cellular differentiation can

― 20 ―

be controlled—or at least affected—by plant hormones. However, the mechanisms by which particular plant hormones affect the form, substance and synthesis of particular cell walls are almost wholly unknown.

Significant progress has been made recently in elucidating the role of auxins in cell wall elongation. Indoleacetic acid and several similar compounds cause a marked increase in the rate of coleoptile cell elongation (Cleland, 1971). Coleoptiles normally receive auxin from the apical region of the shoot. However, excised coleoptiles can respond to auxin exogenously supplied in solution, elongating at a rate of 10–30% per hour. The mechanism of auxin-stimulated elongation of coleoptile cells has been proposed to involve the activation of a hydrogen ion 'pump' in the plasmalemma (Cleland, 1973; Rayle, 1973). The hormonal activation of this pump results in a lower pH in the cell walls which, in turn, appears to activate enzymes in the cell wall which are capable of selectively 'loosening' the polysaccharide network so that the walls can elongate more rapidly. Auxin must also stimulate cell wall biosynthesis (by some unknown mechanism) since the walls retain a constant thickness while more than doubling their length during prolonged auxin treatment.

A further important aspect of hormonal effects on plant cell wall biosynthesis and differentiation has been revealed by studies on the changes in microtubule orientation caused by exogenously supplied hormones (Shibaoka, 1974). In bean epicotyl segments, kinetin inhibits elongation and promotes a thickening or lateral expansion of the cells. Gibberellins, on the other hand, promote elongation and inhibit lateral expansion. The microtubules adjacent to the plasmalemma in cells of epicotyl sections supplied with kinetin and auxin are found to be oriented parallel to the cell axis. However, in sections supplied with gibberellin and auxin the microtubules are oriented transverse to the cell axis. The microtubules are randomly oriented in sections supplied with auxin alone. Thus, kinetin and gibberellins (but not auxins) appear to be able to control the direction of cell growth by somehow determining the orientation of the microtubules. The orientation of the microtubules, in turn, determines the orientation of the cellulose microfibrils being deposited in the cell wall—which determines the direction in which the walls can most easily expand. (See Chapter 13 for further discussion of hormone action.)

1.9—
The Role of the Plant Cell Wall in Interactions with Other Organisms

Plants are beset by many pests and pathogens and helped by a variety of symbionts. The walls of epidermal cells may be specialized in a number of ways to form a protective skin over the entire plant (e.g. cutin, wax, gums and mucilages, bark and thorns, etc.). Many cell walls throughout the plant can become resistant to degradation by pathogens by means of lignification.

Quite apart from such modifications, however, the structure of the cell wall

― 21 ―

Figure 1.10
Penetration of epidermal cell walls of broad bean by Botrytis cinerea.
The fungal germ tube (G) is attached to the surface of the cuticle (CU) by
a layer of mucilage (M) secreted by the fungus. The fungus appears to
have dissolved a hole through the cuticle and to have begun dissolving
the plant cell wall (CW) beneath this hole. A membrane-bound infection
peg (P) penetrates through the pore in the fungal cell wall and the cuticle.
(Courtesy Prof. W. E. McKeen. See Phytopathology (1974), 64, 461–67.)

― 22 ―

can itself present a formidable barrier to pathogens. In order to penetrate the cell wall and utilize its component sugars, pathogenic fungi and bacteria have evolved sophisticated, inducible batteries of enzymes which can hydrolyze components of the plant cell wall (Bateman & Basham, 1975). Figure 1.10 illustrates the initial stages of cell wall penetration by a fungal pathogen. The pathogen appears to have dissolved a hole through the cuticle and to have started degrading the cell wall.

Plants, in turn, have evolved countermeasures to the hydrolytic enzymes of the pathogens. In primary walls, the molecular architecture is such that only the pectic polysaccharides are accessible to hydrolytic enzymes (Bauer et al., 1973). It is probably for this reason that the first enzymes to be secreted by an invading pathogen are the pectin-hydrolyzing enzymes (Bateman & Basham, 1975). This is also likely to be the reason why the walls of many plants contain proteins which specifically inhibit the pectin-degrading enzymes (and only the pectin-degrading enzymes) of pathogenic microorganisms (Anderson & Albersheim, 1971). In other plants the pectic polysaccharides are heavily acetylated, and thus resistant to enzymic attack.

Plants have evolved mechanisms for counterattack as well as defence. Inducible enzymes are present in the cell walls of several plants which hydrolyze the wall polysaccharides of invading fungal pathogens (Ables et al., 1970; Pegg & Vessey, 1973). The pathogens react by secreting proteins which specifically inhibit the attacking plant enzymes (Albersheim & Valent, 1974). The cell walls of a variety of other plants contain glycoside-hydrolyzing enzymes which can release hydrogen cyanide from cyanogenic glycosides. The release of hydrogen cyanide by the plant occurs in response to attack by a pathogen. The pathogen (sometimes) avoids cyanide poisoning by an inducible enzyme which converts the cyanide to harmless formamide (Fry & Munch, 1975).

From these and other examples it is clear that the cell wall is a most important battleground in the contest between plants and their pathogens. The plant cell wall is not just a strong but passive barrier to invasion. It is impregnated with a host of molecules which can recognize a pathogen, modify the defences, or mount a counterattack.

Chapter 2—
Membrane Structure and Transport

2.1—
Introduction

The control of metabolism and the development of cells frequently depends on the right substance being present in the right amount at a specific location in the cell at the right time. This may be achieved by regulation of the passage of materials from the external environment into the cell or from one compartment of the cell to another. All compartments of the cell, and its external surface, are bounded by membranes. It is clear, therefore, that any complete understanding of control mechanisms in metabolism or development must include a precise knowledge of the structure and composition of membranes and of the mechanisms whereby materials move through them. While it would not be true to suggest that all of this knowledge is available at present, the pace at which new information and insight has been gathered in the last decade is most impressive. In this short chapter it will not be possible to trace the history of the way in which ideas about membrane structure have developed, but it is worth mentioning that a (substantially correct) view of the basic structure of biological membranes was advanced in the 1930's, long before it was possible to visualize membranes in the electron microscope or to examine their detailed structure by X-ray deffraction techniques (Danielli & Davson, 1935). The simple trilaminar appearance of biological membranes in the transmission electron microscope (Fig. 2.1) is now familar to elementary students of biology and is known as the unit membrane; its occurrence is ubiquitous and this very fact has impressed on biochemists and others that this apparently uniform structure cannot explain the diverse properties of different membranes. This chapter covers the chemical composition of membranes and how these components are arranged. From this it will become apparent that the membrane is composed of a matrix, whose design is broadly similar in all cases, and a sub-structure on which many of the specific properties of the membrane probably depend. With this picture in mind it will then be possible to explore the basic types of transport which can occur across membranes and to relate them to the structures described.

2.2—
Chemical Composition of Membranes

Biological membranes are composed primarily of two main classes of compounds, lipids and proteins, which interact in several ways with water to bring

― 25 ―

Figure 2.1
Electronmicrograph showing the plasmalemma of two endodermal cells
separated by a cell wall (c.w.) in the root of barley. The plasmalemma (pl.) is
prominently stained in the central region of the picture and is clearly trilaminar.
Notice that the tonoplast (t) bounding the vacuole (vac.) is much less distinct
but also appears trilaminar at the point indicated by the arrow. Other symbols
er = endoplasmic reticulum; c = cytoplasm. Total magnification about × 200,000.
(Micrograph by courtesy of Dr. A. W. Robards).

― 26 ―

about a characteristic trilaminar structure. As a very broad generalization it might be said that lipids make up the supporting matrix of the membrane while the proteins thus supported determine its specific properties. This is, of course, merely a convenient simplification as it is becoming plain that some of the characteristic properties of a membrane, particularly those which determine its effectiveness as a barrier to the diffusion of water and electrolytes, depend on the nature of the membrane lipids. Conversely some proteins have a structural role. For the present purposes, however, the classes of compounds will be considered separately and a synthesis attempted later on.

2.2.1—
Lipids

Of the lipids present in plant cells the various phospholipids, glycolipids and sterols are of the greatest significance in membrane construction. The relative abundance of the components can be quite variable (Table 2.1) depending on the part of the plant analysed, the species and prevailing environmental conditions (see p. 29).

Table 2.1. Relative abundance of lipids of various classes in plant organs and organelles.
	Phospho- lipids	Glyco- lipids (% of total lipid)	Neutral lipid*	Ref.
Bean leaves	22	38	40	Wilson & Crawford (1974)
Spinach
chloroplasts	15	47	38	Allen et al., (1964)
Grape root	34	10	56	Kuiper (1968)
Oat root
mitochondria	38	10	52	Keenan et al., (1973)
* Sterols and sterol esters usually make up more than 50% of the neutral lipid fraction.

2.2.1.1—
Phospholipids

The phospholipid molecule can be separated into a charged or polar 'head' region and an uncharged or non-polar 'tail'. Such a molecule is described as amphipathic, and as we shall see later on, this property is of crucial importance in determining membrane structure (p. 36). Phospholipids are generally thought to be restricted to membranes but the extremely rapid rate at which membranes can be taken apart and re-assembled, as in cell plasmolysis and de-plasmolysis, makes it probable that there are stores of phospholipid within the cell.

Phospholipids are readily extracted from macerated plant tissues by a mixture of chloroform and methanol (2:1) and can be separated by thin layer chromatography using a variety of solvent systems (see Hitchcock & Nichols, 1971, for a review of techniques).

― 27 ―

The commonest phospholipids in plant membranes are derivatives of phosphatidic acid (Fig. 2.2); thus, lecithin is the choline ester of phosphatidic acid. Other common derivatives are also shown in Fig. 2.2. Phosphatidic acid (PA) itself is generally said to occur only in minute quantities in membranes or not at all, indeed, its presence in an extract is often taken as an indication of the activity of phospholipase D (Mazliak, 1973). There is a report, however, in which phosphatidic acid is said to be one of the major constituents of the plasmalemma of oat (Avena sativa ) root, (Keenan et al., 1974). Unfortunately,

Figure 2.2
Structural formulae of phospholipids commonly found in plants.

detailed analyses of the plasmalemma from other plants are not available for comparison. In passing it might be noted that a great deal remains to be done, firstly in preparing pure sub-cellular fractions of the plasmalemma and of other membranes from plants and subsequently in determining their lipid composition. Table 2.2 presents some of the available information on the distribution of the different types of phospholipid. The information on the composition of the

― 28 ―

Table 2.2. Relative abundance of phospholipids in tissues and organelles.
	PA*	PI	PC	PE	PG	DPG	Other	Ref.
		(% of total phospholipid)
Whole leaves
Sugar Beet	ND	11	47	23	19	ND	—	(1)
Maize	ND	8	30	16	31	15	—	(2)
Bean	5	7	42	20	26	ND	—	(3)
Chloroplasts
Sugar Beet	10	9	35	13	33	ND	—	(1)
Tobacco	ND	4	23	16	58	ND	—	(4)
Spinach (lamellae)	trace	10	20	13	57	ND	—	(5)
Mitochondria
Cauliflower (floret)	ND	7	44	34	3	12	—	(6)
Potato (tuber)	ND	13	44	26	ND	17	—	(6)
Microsomes
Cauliflower (floret)	ND	6	50	35	8	1	—	(6)
Potato (tuber)	ND	19	44	18	ND	19	—	(6)
Plasmalemma
Oat (root)	14	4	18	11	4	ND	11**	(7)
(1) Wintermans (1960) (2) Roughan & Batt (1969) (3) Wilson & Crawford (1974) (4) Ongun & Mudd (1968) (5) Allen et al., (1964) (6) Moreau et al., (1974) (7) Keenan et al., (1973) Notes * possibly indicates the activity of phospholipase D during preparation (see Mazliak, 1973). ** detected lysolecithin and lysophosphatidyl-ethanolamine—indicates possible activity of phospholipase A during preparation (see Nachbaur & Vignais, 1968)

membranes of mitochondria and chloroplasts is the most detailed and reliable since these organelles can be separated with relative ease and high purity during cell fractionation. The predominant phospholipid in a given membrane may be characteristic, e.g. phosphatidylglycerol (PG) is a major component of chloroplast membranes while it is only a minor component of the inner mitochondrial membrane where diphosphatidylglycerol (DPG) is predominant. In general extracts of shoots and roots neither of these phospholipids is as abundant as lecithin (PC) or phosphatidyl ethanolamine (PE). A small quantity of phosphatidyl inositol (PI), usually less than 10% of the total phospholipid, is found in all membranes.

In Fig. 2.2 the exact chain length of the acyl groups R₁ and R₂ which make up the hydrophobic tail, is not defined precisely. In nature it can vary considerably even in one type of phospholipid from a given tissue. The chain may be made from 12 to 22 carbon atoms and may contain up to three or, rarely, six double bonds. The chain is straight in all eukaryotic organisms and has been found to be branched only in certain bacteria (Asselineau, 1966). Variation in both the length and unsaturation (i.e. the number of double bonds) of the hydrocarbon chain influences its melting point; shorter and unsaturated chains melt at much

― 29 ―

lower temperatures than longer and saturated ones. As an example of this consider the effect of double bonds on the melting of free fatty acids containing 18 carbon atoms; the saturated stearic acid (C_18:0 ) melts at 69ºC, the monounsaturated oleic acid (C_18:1 ) at 5°C and the double unsaturated linoleic acid (C_18:2 ) at –12ºC. Organisms which live in warm conditions and warm blooded animals are generally found to have phospholipids with an abundance of fatty acids which tend to be fully saturated (e.g. the thermophilic alga Cyanidium caldarium, see Kleinschmidt & McMahon, 1970). By contrast, organisms which are exposed to lower temperatures have either more unsaturated acids or ones with shorter average chain lengths or a combination of both of these (e.g. in Acholeplasma laidlawii, see Huang et al., 1974) to give phospholipids whose tails remain fluid. The significance of the maintenance of membrane fluidity will become apparent later (p. 41). The process of hardening plants against injury from frost or chilling is accompanied by changes in the degree of unsaturation of the membrane lipids (Wilson & Crawford, 1974).

Table 2.3 shows fatty acid analyses of individual phospholipids extracted from various sources. Bearing in mind that there is a great deal of room for manoeuvre in selecting the fatty acids to suit the prevailing environmental temperature the values for the relative abundance of fatty acids should be considered only as very general guides to the types of acid found in nature. Thus, the predominant fatty acids have even numbers of carbon atoms, the saturated acids found most frequently are palmitic (16:0) and stearic (18:0), and the principal unsaturated acids are linoleic (18:2) and the triply unsaturated linolenic (18:3). The fatty acid composition of lecithin can depend very strongly on its origin. For example, the lecithin in the outer mitochondrial membrane is much richer in palmitic acid (16:0), and perhaps is a less fluid component than in the inner mitochondrial membrane where triply unsaturated linolenic (18:3) is the most abundant fatty acid.

2.2.1.2—
Glycolipids

In several respects the glycolipids resemble phospholipids. The molecule is amphipathic, the polar group being a galactosyl derivative of a diglyceride, the non-polar part of the molecule being a pair of long, straight-chain fatty acids. Glycolipids are unusually rich in the triply unsaturated linolenic acid (C_18:3 ) which may make up more than 90% of the fatty acid (Table 2.3).

The two most abundant glycolipids are mono- and di-galactosyl diglyceride, the structural formulae of which are illustrated in Fig. 2.3. They are characteristic of photosynthetic tissues since they are the major lipid component of chloroplast lamellae, largely replacing the phospholipids. Ongun et al., (1968) showed that more than 80% of all of the glycolipid in leaf cells was present in the chloroplasts. The probable orientation of these molecules in the chloroplast lamellae is much like that described for phospholipids (see 2.2.1.1.) with the fatty acid tails inserted into the central region of the membrane with the polar

― 30 ―

Table 2.3. Fatty acid composition of phospholipids and glycolipids extracted from various plant membranes.
				Fatty Acid* (% of total)
	14:0	15:0	16:0	16:1	16:3	18:0	18:1	18:2	18:3	Ref.
Lecithin
Bean leaf	—	—	27	trace	—	6	4	38	26	1
Spinach leaf	trace	—	20	trace	trace	—	11	30	40	2
Barley root	1	< 1	19	1	(3)*2	3	7	40	25	3
Cauliflower mitochondria
—outer membrane	trace	trace	61	trace	trace	7	17	7	8	4
—inner membrane	trace	trace	20	trace	—	1	9	13	57	4
Phosphatidyl ethanolamine
Spinach leaf	trace	—	46	—	2	1	2	7	43	2
Barley root	9	< 1	20	1	(6)*	2	6	35	21	3
Grape root	5	—	49	—	—	12	12	11	3	5
Monogalactosyl diglyceride
Bean—whole leaf	—	—	2	trace	—	trace	trace	2	96	1
—chloroplast envelope	—	—	10	trace	—	6	11	9	62	6
—chloroplast lamellae	—	—	1	2	trace	1	1	3	94	6
Digalactosyl diglyceride
Bean—whole leaf	—	—	5	trace	—	1	trace	1	93	1
—chloroplast envelope	—	—	9	1	—	2	6	17	65	6
—chloroplast lamellae	—	—	4	trace	—	3	2	2	89	6
(1) Sastry & Kates (1964) (2) Allen et al., (1964) (3) Clarkson (unpublished) (4) Moreau et al., (1974) (5) Kuiper (1968) (6) Mackender & Leech (1974). * Fatty acids are given as carbon chain length and number of double bonds. ** There is uncertainty about the exact length of this component—it may possibly be 17:2.

― 31 ―

galactosyl groups at the membrane surface protruding into the stroma (Weier & Benson, 1967). Because the fatty acid is so highly unsaturated the membranes of lamellae probably remain fluid even at sub-zero temperatures—thus any photosynthetic reaction, or molecular reorientation, which depends on membrane fluidity may have a wide temperature range in which it can occur.

A sulphur-containing glycolipid is found as a minor component of most membranes. It is known as sulpholipid (Fig. 2.3) and its structure and occurrence in chloroplasts was reported by Benson (1963); the acyl groups are mainly palmitic with a preponderance of linolenic acid, thus resembling the other

Figure 2.3
Structural formulae of glycolipids commonly found in plants.

― 32 ―

galactosyl lipids. Sulpholipid represents only 1% of the total lipid in most tissues and organelles but in chloroplasts it may be as much as 10–15% of the lipid (Ongun & Mudd, 1968).

2.2.1.3—
Sterols

A number of sterols can be extracted from plant tissues and fungi as well as from isolated membrane fractions. The conventional example of a sterol of common biological origin is cholesterol (Fig. 2.4); in practice plant cells contain relatively little of this sterol in comparison with animal cells. Such meagre quantitative data as is available show that sterols having 29 carbon atoms, e.g. b -sitosterol (Fig. 2.4), are the most abundant in higher plants, while in fungi the C-28 sterol, ergosterol (Fig. 2.4) is often dominant. All of these molecules have an extended concertina-like configuration (known as the 'chair' or 'boat'),

Figure 2.4
Structural formulae of sterols commonly found in biological membranes.

seem metabolically inert and are synthesized and turned over very slowly, especially in comparison with the other lipid components of membranes (Nes, 1974). Their function in membranes is not well understood but it is likely that they have an architectural role concerned with the maintenance of structure or order in the lipid domain. In this respect all of them probably function in the same was as cholesterol (see p. 38) because Butler et al., (1970) found that the structural order of bilayer membranes synthesized from lipids of ox brain tissue was stabilized equally well by cholesterol, b -sitosterol of plant origin and ergosterol. In the plasmalemma of the animal and plant cell there is a much higher proportion of sterols and sterol esters relative to phospholipid than in other

― 33 ―

membranes (Table 2.4). It should be noted, however, that the membranes of intracellular organelles contain much more protein than do plasmamembranes (see p. 35). To some extent this protein, much of ₂ which is bonded hydrophobically to the lipid, may function in a way similar to sterol in maintaining the structural order of the membrane interior.

Table 2.4. Relative proportions of sterols and phospholipids from membrane fractions ofAvena root.
	Proportion of total lipid*(%)
	Plasmalemma	Mitochondria
Sterol	24	18
Phospholipid	29	38
Sterol/Phospholipid	0.82	0.47
The remaining lipid is composed largely of triglycerides and some free fatty acid and glycolipid. (Based on data from Keenan et al.,* (1973)).

2.2.2—
Proteins

In many membranes, particularly in those of chloroplasts and mitochondria proteins make up most of the weight. The proteins found are many but as a first step in classifying them integral and peripheral proteins may be distinguished. This classification anticipates the subsequent discussion of membrane structure on page 38 but the terms clearly suggest that proteins in the two classes are associated with other components in the membrane in different ways. The recognition that certain proteins are embedded deeply in the lipid membrane represents a departure from the view, often advanced in earlier texts, that all of the membrane protein is located in the two peripheral bands which stain darkly with osmium and are visualized in the electron microscope (see Fig. 2.1). Whereas some of the protein is certainly located in this way and is probably bonded to the polar regions of the phospholipids electrostatically, it has become apparent that much protein is associated with the non-polar regions of the lipid by hydrophobic bonding. Peripherally located protein can be easily separated by washing with salt solutions or chelating agents but integral proteins are attached very strongly to the membrane and can be removed only after drastic treatment with organic solvents or detergents; even then, the isolated protein usually has some lipid attached to it.

Most of the membrane-bound enzymes, transport proteins (e.g. monovalent cation stimulated ATPase), drug and hormone receptors (in animal cells) and antigenic proteins are integral and are revealed when membranes are split open in freeze-fracturing (see p. 38). In many instances the enzymes arenonfunctional in vitro in the absence of lipid. It is thought that the non-polar parts of the polypeptide chains are associated with the hydrophobic tails of the fatty

― 34 ―

acids and, this being so, several types of conformation are possible (see Singer, 1974 for a review). Proteins, like phospholipids, are amphipathic and their polar regions will arrange themselves so that their contacts with the hydrophobic regions of the membrane will be minimized; to ensure this, a protein could be arranged so that its polar, hydrophilic region lies among the phospholipid 'heads', or projects through them into the protein at the membrane periphery (Fig. 2.5).

Figure 2.5
Possible orientations of proteins in a membrane. (a) peripherally bound protein
with polar groups all over its surface. (b) and (c) non-polar regions of the protein
bonded hydrophobically to lipid but with different numbers of polar groups. (d) polar
groups at either end of a long molecule with a non-polar central region. (e) a pair of
proteins as in (d) making up a polar pore or channel in the membrane. (f) a hydrophobic
globular protein wholly in the lipid domain—polar groups, if any, directed inwardly.

Long polypeptide chains with charged groups at either end may actually lie across the width of the membrane, or groups of them might lie with polar groups directed inwardly to form a hydrophilic pore across the membrane. An alternative conformation would be provided by the formation of a globular structure in which all of the polar groups would be directed towards the centre of the globule so that a hydrophobic surface would be presented to the lipid. This latter kind of conformation is probably least common.

The peripheral proteins can be attached to the polar groups of either phospholipids or integral proteins; examples which might be taken include cytochrome c which is located on the outer surface of the inner mitochondrial membranes (Schneider et al., 1972; see also Chapter 5), the chromoprotein, phytochrome, which is thought to be attached to the plasmalemma (Marmé et al., 1974; see also Chapter 12) and the sulphate and other ion-binding proteins on the outer surface of bacterial membranes (see Oxender, 1972, for a review).

From a quantitative point of view, certain generalizations about the relative abundance of peripheral and integral proteins can be made. The greater the metabolic activity which centres upon a given membrane system the greater amount of protein integrated into it. Thus, it might be anticipated that chloroplast lamellae and inner mitochondrial membranes would be relatively rich in these proteins, whereas membranes whose role is more concerned with providing

― 35 ―

a diffusion barrier, e.g. the plasmalemma and the tonoplast, would be less so; evidence from electron microscopy shows that this is so (see Table 2.5).

2.2.3—
Water

Water is an important, if neglected, constituent of membranes for several reasons. In a general way it determines their basic design since, in its presence, amphipathic lipids assume a bilayered configuration (see p. 36). There are, however, other specific associations of water molecules and membrane components which are not fully understood.

It has been estimated that water of hydration accounts for about 30% of the weight of membranes. Much of this water will almost certainly not be in a liquid state but will exist in ordered layers around the hydrophilic parts of lipids and proteins. Immobilized by hydrogen bonding these water molecules are in a liquid-crystalline condition and cannot be frozen to form ice. Water layers bound at the surface of the membrane have been estimated to have viscosity of 39 times that of pure water and to have a thickness of at least 2.2 nm (Schultz & Asunmaa, 1970). They must contribute to the mechanical stability of membranes and add significantly to their barrier properties to diffusing solutes. Expressing an extreme view, Ling (1973) has proposed that it is not lipid but these polarized multilayers of water which provide the cell with its selective surface barrier.

Hydrophobic bonding between the non-polar regions of lipids and integral proteins (see p. 34) is favoured thermodynamically by the interactions of their polar regions with water (Tait & Franks, 1971).

Much experimental evidence points to the fact that water molecules are not restricted to membrane surfaces but cross the hydrophobic regions in numerous water-filled pores. These pores are thought to conduct water and small solutes (diameter <0.4 nm) to which membranes are highly permeable (see p. 60). Some water lining these pores is fully 'organized' and should, therefore, be regarded as a structural feature but there is indirect evidence to suggest that some of it must be free water in transit.

2.3—
Membrane Structure

It is convenient to discuss membrane structure under two headings; the organization of the membrane matrix which is largely a matter of the relationships of the lipid components, and the substructure of the protein in the membrane.

2.3.1—
The Membrane Matrix

2.3.1.1—
Phospholipids

The structure of phospholipid molecules considered earlier provides the key to understanding why it is that the membranes as seen in transverse sections have

― 36 ―

a characteristic trilaminar appearance. The hydrophilic head regions make hydrogen bonds with water and may become cross-linked to other heads and to proteins through ionic bridges, e.g. by calcium ions; thus they are organized into a lattice-like structure. By contrast the long acyl chains of the two fatty acids attached to each phospholipid are strongly hydrophobic, loosely organized and, above their melting point, are relatively fluid. If phospholipid is dispersed in water the 'tails' will take on a conformation which will minimize their contacts with water. The 'heads' will, of course, react favourably with water. If the available water surface is large relative to the amount of lipid, the molecules will arrange themselves as a film-like monolayer with the heads at the water surface and the 'tails' protruding from it at right angles (Fig. 2.6a). If more phospholipid molecules are added to this system so that there are more than can be fitted into a tightly packed monolayer over the water surface, a second type of arrangement occurs quite spontaneously. The phospholipids form two ranks with the heads facing outwards in both and the tails directed inwards to form a non-polar hydrophobic layer sandwiched between them (Fig. 2.6b). This bilayer arrangement, which is common to all biological membranes, can also be formed from mixtures of phospholipids under laboratory conditions. The synthetic membranes thus produced have helped in arriving at an understanding of many of the structure/function relationships of natural membranes (see Goldup et al., 1970, for a readable review).

Figure 2.6
An illustration of how a monolayer of dispersed phospholipid (a) in water, forms
into a bilayer, (b) on contraction of the water surface area. The phospholipid
heads have water bound to them in polarized multilayers (see p. 35).

The selected analyses in Table 2.2 show that a given membrane may contain several types of phospholipid as well as appreciable quantities of sterol. It is probable that there is a great deal more organization of phospholipids in natural membranes than can be demonstrated positively at present. Lipids of one kind may be associated into clumps so that the membrane surface may be very heterogeneous with lipids of differing physical properties arranged in a mosaic.

― 37 ―

A mosaic of charged and uncharged areas might occur because some phospholipids carry a net electrostatic charge at normal pH values, e.g. phosphatidyl glycerol, while others, like phosphatidyl choline (lecithin) are neutral. This is of significance because it has been shown that, in synthetic bilayers, the surface charge on the phospholipid heads can partly determine both the ion-selectivity and the cation permeability of the membrane (Papahadjopoulos, 1971), and it may also be relevant in determining regions of the surface of the plasmalemma where endo-cytosis may occur (see p. 61).

Local variations in the packing of sterols may render some parts of the membrane less fluid than others and thus determine areas where diffusion may be severely restricted (Papahadjopoulos et al., 1973).

More recently researchers have begun to investigate the possibility that the inner and outer halves of the bilayer may differ in their phospholipid composition. Should this prove to be the case, then it is possible that the barrier properties of the membrane to solute diffusion may be different when the membrane is approached from different sides.

In some special circumstances phospholipid molecules may become arranged into globular micelles in which the polar groups are directed towards the periphery of the sphere, the surface being hydrophilic. This state of affairs can be induced by dehydration in synthetic membrane systems, and in nature by viruses which create membrane instability, e.g. sendai virus, and by certain phospholipids (e.g. lysolecithin) with wedge-shaped head regions which tend to induce curvature of layers of closely packed phospholipids when they are introduced into a bilayer (Lucy, 1970). It has been suggested that rapid local transitions from the predominant bilayer to the micellar state are important in membrane fusion and in pinocytosis (see Lucy, 1970). If these transitions do occur then it is possible that they may cause transient gaps or pores to be created in the membrane; much physiological evidence points to the conclusion that membranes do have very fine pores in them (see p. 59).

2.3.1.2—
Sterols

The insertion of sterol molecules into the membrane increases the structural order of the hydrophobic region. These molecules lie with their long axes parallel to the hydrocarbon chains of the fatty acids with their more rigid ring structures directed towards the outside and their open chain ends towards the centre. The mobility of the hydrocarbon chains nearest to the outside of the membrane is, therefore, restricted by these stiffening structures but they remain pliant at their ends so that the central region is fluid (Caspar & Kirschner, 1971). The rigidity conferred on the membrane by the inclusion of sterols slows down the diffusion of materials through the outer part of the lipid domain in synthetic bilayers (Papahadjopculos et al., 1973).

― 38 ―

2.3.1.3—
A Model of the Membrane Matrix

Figure 2.7 provides a basic interpretation of the ideas on the membrane matrix discussed so far. A fact, which it is important to understand but which is difficult to illustrate, is that the centre of the membrane is fluid while the periphery is semi-crystalline. Although it is a stable structure, it is known that phospholipid molecules can be inserted into, and withdrawn from the matrix rapidly and that the structure illustrated in Fig 2.7 represents a dynamic steady state when it is part of a biological membrane.

2.3.2—
Membrane Sub-Structure

When a cell or a piece of tissue is frozen and then fractured with a suitable blade, the fracture plane will follow lines of weakness in the structure. Since the central region of the membrane matrix contains no ice it is a potential line of weakness, along which the membrane tends to fracture (Fig. 2.8). Where the fracture line passes tangentially across a cell or organelle, sheets of membrane material become apparent; since membranes tend to be cleaved down the middle it is obvious that the surface exposed is not the true membrane surface but is the membrane interior. Figure 2.9 shows a relatively smooth sheet of plasmalemma from an onion root tip on which numerous round particles and depressions can be seen. Some of the particles are arranged in files while others are randomly distributed. These particles, which are usually 6–9 nm in diameter, are embedded in the membrane and are not resting on its true surface. This was demonstrated by Pinto da Silva & Branton (1970) who etched away the ice from the fracture plane by leaving the specimens under a high vacuum for some minutes after fracturing them. Using this method, the true surface of membranes which lay obliquely to the fracture plane was eventually revealed as the ice from the surrounding cytoplasm sublimed. The true surface had a much smoother appearance, the undulations of which gave the impression of a blanket lying over the embedded particles. The particles in the membrane can be removed by treatment with proteolytic enzymes and can be re-created in synthetic membranes which have been made in the presence of a hydrophobic protein. Vail et al., (1974) reported that a synthetic bilayer membrane into which a hydrophobic protein had been incorporated had intercalated particles of 8.5 to 9 nm diameter occupying 12% of the internal membrane surface whereas in bilayers lacking the protein there were none.

The frequency of particles varies according to membrane type (Table 2.5); chloroplast and mitochondrial membranes have the highest frequency as one might anticipate from the many metabolic events which are centred upon them. One of the two fracture faces is usually more densely populated than the other. Bearing in mind the amphipathic nature of membrane proteins (see p. 34) it is possible that the more densely populated face may reflect the principal orientation of the polar groups of the integral proteins, i.e. if most of the polar groups were directed towards the outside of the cell or compartment, the outer fracture

― 39 ―

Figure 2.7
Orientation of polar lipids, cholesterol and peripheral protein in a model of the membrane matrix.
Based on X-ray diffraction analysis of nerve myelin. (From Caspar & Kirschner, 1971). In many
ways myelin has been an unfortunate choice for detailed structure of cell membranes since it is
not typical in having virtually no integral protein and has no intercalated membrane particles
(see Table 2.5). It is, however, very appropriate as a general model of the membrane matrix.

Figure 2.8
Line of fracture when a frozen membrane is split open in freeze-fracturing. Particles
embedded in the membrane become apparent in the cleavage plane either as
projections or as hollows if they are removed on the upper half of the bilayer.

― 40 ―

face would be the most densely populated. It should be noted that in synthetic lecithin membrane and in nerve myelin, which both have a very low permeability to ions, there are no particles.

Figure 2.9
Particles seen on the inner fracture face of the plasmalemma from a root tip
cell in onion. The fracture face exposed as described in Fig. 2.8 and metal
shadowed to produce a replica; the arrow represents the direction of
shadowing. Note that there are distinct files of particles as well as randomly
distributed individuals. The membrane matrix appears to be relatively smooth.
(Micrograph by courtesy of Professor D. Branton).

― 41 ―

Table 2.5. Density of intercalated particles in various membranes.
	Number of particles m m^–2
Type of membrane	Densely populated face	Thinly populated face	Membrane area covered with particles %	Ref.
Lecithin: synthetic	0	0	0	(1)
Nerve myelin sheath	0	0	0	(1)
Plasmalemma—cambial cells of willow	1,608	282	16	(2)
Plasmalemma—root tip of onion	2,030	550	15	(1)
Tonoplast—cambial cells of willow	695	140	7	(2)
Tonoplast—root tip of onion	3,300	2,480	32	(1)
Nuclear envelope—root tip of onion	1,790	420	12	(1)
Endoplasmic reticulum—root tip of onion	1,700	380	12	(1)
Mitochondria
—outer membrane	2,806	770	23	(3)
—inner membrane	4,208	2,120	40	(3)
Chloroplast—lamellae*	3,860	1,800	80	(1)
(1) Data cited in Branton & Deamer (1972) (2) Parish (1974) (3) Wrigglesworth et al., (1970) * The particles in the chloroplast lamellae were 16–20 nm in diameter while in most other membranes examined they were 7–9 nm.

2.3.2.1—
A Model of Membrane Structure

The representation of membrane structure shown in Fig. 2.10 is one which commands wide acceptance at the present time. In it we see that large masses of integral protein are inserted to varying extents in the membrane matrix while some are restricted to the surface and others protrude right across the width having a dumbell configuration. The model, which has become known as the fluid mosaic (Singer & Nicholson, 1972), has a disorderly appearance in contrast to the neat pictures used to illustrate the unit membrane; the membrane sub-structure has been likened to numerous protein icebergs in a sea of lipid.

2.3.2.2—
Membrane Fluidity

An essential feature of the model in Fig. 2.10 is that the various components of the membrane can move relative to one another. Rather than being fixed points, many of the proteins are best thought of as drifting around in the lipid, the viscosity of which will determine the rate at which they move. Evidence that membrane particles can move in the plane of the membrane comes from freeze

― 42 ―

Figure 2.10
A representation of the fluid mosaic model of membrane structure
showing peripheral and integral proteins in a 'sea' of lipid molecules.
Not shown are the polarized water layers which are bound to the polar
regions of the lipids and proteins. The round-headed objects with twin
tails represent phospholipids, the obovoid structure with the single tail
represents sterol and the large irregularly shaped objects are proteins.
(Based on ideas in Singer & Nicholson, 1972 and Capaldi, 1974).

fracture studies of the plasmalemma of Mycoplasma mycoides, in which it was seen that particles became aggregated as the membrane was cooled down but dispersed again when it was warmed up (Rottem et al., 1973). A second line of the evidence comes from studies where two animal cells were induced to fuse with one another (Edidin & Farnbrough, 1973). One of the cells has a specific antigen in its plasma membrane which the other cell lacked. The distribution of the antigen could be visualized by the binding of a fluorescent antibody. The fusion of the cells to form a heterocaryon and the subsequent redistribution of the fluorescent antibody were observed using a fluorescence microscope. At first the fluorescent marker was restricted to one half of the heterocaryon, but quite quickly, and in an orderly progression the fluorescence spread right around the coat indicating that the antigenic protein, known to be integrated into the membrane matrix, had diffused in the plane of the membrane. For it to have done this it must have drifted through the lipid. If the heterocaryon was cooled below the transition temperature of its lipids, no mixing of the antigen occurred.

From what we have said above and from earlier comments (p. 29) it is clear that temperature will have a very important influence on the diffusion of materials through, and in the plane of, the membrane because of its effect on the viscosity of the lipid (Edidin, 1974). If the membrane is cooled to the extent that the lipids freeze then such diffusion will become very restricted; the biological

― 43 ―

significance of this is reflected in the fact that the rates of most physiological processes examined over a range of temperatures are found to have a sharp transition at a temperature which is close to the transition of membrane lipids from a liquid crystalline condition to the gelled condition (Simon, 1974).

Most workers agree that there must be some proteins whose position in the membrane relative to others must be maintained, e.g. components of electrontransport chains. Such proteins may require anchoring points either in parts of the membrane which are less fluid than others, or by association with some extra-membrane protein. It is possible that aggregations of sterol molecules might serve to create more viscous patches, and there is evidence, again from synthetic lipid bilayers, that cholesterol can be concentrated in association with certain phospholipids (de Kruyff et al., 1974).

2.3.2.3—
Membrane Synthesis and Flow

The intermediary metabolism concerned with the synthesis of the components of the membrane is beyond the scope of this chapter but it is believed that the components themselves may be centrally assembled and then distributed to the various membranes of the cell. This flow of membrane material can be detected in experiments where cells are provided with a radioactive precursor to a common membrane protein for a short while, and then returned to non-radio-active medium. This type of analysis has not been performed on plant cells but in animal cells, harvested at intervals after pulse-labelling, the radioactive label appears first in the endoplasmic reticulum and then in the Golgi cisternae. Subsequently, the radioactivity of these compartments declines followed by an increase in label associated with the plasmamembrane some hours later (see Table 2.6). Since mitochondria and chloroplasts probably do not have all of the metabolic apparatus to assemble their own membranes it is thought that they too may obtain partly finished membranes from the endoplasmic reticulum (see also chapter 8).

Table 2.6. Estimated time constants for the appearance of L -(guanido-¹⁴ C) arginine, administered as a pulse label to living rats, in various fractions of the liver (data of Franke et al., 1971).
Fraction	Labelling time (min.)
	Half max.	Max.
Endoplasmic reticulum	5.5–6.5	10
Golgi apparatus	8.5	30
Plasmalemma	>60	>180

There is evidence that the cytoplasm contains numerous membrane-bounded vesicles which frequently appear to be in conjunction with the major membranes

― 44 ―

of the cell (e.g. Mahlberg et al., 1974). They are particularly prominent in cells synthesizing walls and, since it is known that precursors of wall synthesis are formed inside the cell and elaborated outside, it is reasonable to conclude that the vesicles contain precursors which are discharged after fusion with the plasmalemma (Heyn, 1971). It has also been found that particulate material from the external medium can be detected in vesicles within the cytoplasm (Mayo & Cocking, 1969; Robards & Robb, 1974). These results suggest that vesicles can both fuse with, and be formed from the plasmalemma and other membranes, thus allowing materials to pass out of or into the cell without their having to cross a membrane (see p. 61); such movements are known as exoand endo-cytosis, respectively. Membrane fusion also presents opportunities for the transfer of blocks of membrane from place to place as is implicit in the observations in Table 2.6.

Since adjacent membranes can frequently be found to be in contact over comparatively large areas and yet show no tendency to fuse with one another, it is believed that special proteins or phospholipids (e.g. lysolecithin) in the vesicle membrane may trigger fusion where they make contact with the larger membrane sheet (see Lucy, 1970).

2.4—
Transport of Substances Across Membranes

In a healthy cell there is a continuous interchange of water, ions, uncharged solutes, metabolites and dissolved gases across the plasmalemma. As one might expect, not all of these substances move through the membrane in the same manner. Firstly, some substances diffuse into a cell down a gradient of potentia energy; such movement is spontaneous and is the thermodynamic equivalent 0¹ heat passing from a warmer to a cooler body. There are, however, many substances which are accumulated by cells against a gradient of potential hence their movement into the cell is 'uphill' and is equivalent to the flow of heat from a cooler to a warmer body. 'Uphill' transport requires work to be performed and thus consumes energy. 'Downhill' transport is frequently described as passive while 'uphill' transport is described as active and directly involves the participation of cellular metabolism.

In Fig. 2.11 some further sub-divisions of transport processes have been made. Thus, passive movements may occur by at least three types of pathway, whereas active movements must be linked to some energy-consuming mechanism, referred to as a 'pump', in the membrane. The third type of movement occurs because of the undulation and vesicularization of the membrane in endocytosis (see p. 61). Clearly this is a process which depends at some point on metabolism but, as is discussed below, the substances which move into the cell do not necessarily cross the membrane at all. In such circumstances the observed transport is not strictly active in a thermodynamic sense.

― 45 ―

Figure 2.11
Types of active ('uphill') and passive ('downhill')
transport across a membrane. For discussion see text.

2.4.1—
Passive ('Downhill') Transport

Solute molecules in more concentrated solutions possess more free energy than those in lower concentrations; in other words they are at a higher potential. If two solutions of different concentration are mixed, solute molecules will diffuse from areas of high to areas of low potential. In any given situation the chemical potential of an uncharged solute is dependent on its activity as shown in equation 2.1.

where m_s	= the chemical potential of the solute in joules.mole^–1 .
m_s^*	= the chemical potential of the solute in a standard state—it is essentially a reference point and is a constant which can usually be cancelled out in calculations.

― 46 ―

R	= the ideal gas constant, usually taken as 8.314 joules °K^–1 mole^–1 .
T	= the absolute temperature, °K.
ln a_s	= the natural logarithm (log_e ) of the activity of the solute in moles 1^–1 .

For many practical purposes it is assumed that the activity of a solute moving freely in dilute solution is the same as its concentration so that the chemical potential is more frequently written

where C_s = the concentration in moles 1^–1 .

If the solute is charged, its movement from place to place can also be influenced by differences in electrical potential. An ion, therefore, has an electrochemical potential which is related to its concentration (strictly, its activity) and the electric potential of the medium in which it is moving. Thus,

where

and

are the electrochemical potentials of the ion j in a given set of conditions and in a standard state respectively, C_j is the concentration of the ion j, Z_j is its algebraic valency (i.e. K⁺ would be + 1, and Cl^– , –1), F is the Faraday constant which defines the amount of electric charge (approx. 10⁵ coulombs) carried by 1 gram equivalent of the ion and y is the electric potential of the medium. It is important to appreciate that y is a property of the medium and not of the ion j in particular.

Because C_j and y can influence

independently it is easy to see that a difference in electrical potential across a membrane can promote the diffusion of an ion in the absence of any difference in concentration or even against a gradient of concentration. This situation is illustrated in Fig. 2.12 where we can see that the concentration of an ion within a membrane-bounded compartment may be much greater than in the surroundings and yet be at electrochemical equilibrium providing that a sufficiently large electric potential is maintained. It is wrong, therefore, to conclude, because an ion is more concentrated within a cell than in the outside solution, that it has been actively transported.

2.4.2—
Criteria For Active ('Uphill') Transport

To decide whether or not an ion or solute is actively transported across a membrane we need to know its activity or concentration in the two solutions separated by the membrane and for ions, in addition, we must know the electrical potential difference across the membrane. It is frequently difficult to measure the concentrations, and more difficult to measure the activity, of substances within cells with any accuracy, especially in those of higher plants.

In the giant cells of several sorts of algae, which may be 5,000 to 10,000 times the volume of a parenchyma cell in a root, such measurements are made

― 47 ―

Figure 2.12
An explanation of the way in which a decrease in electrical potential, y , across a membrane
can result in the diffusion of an ion against a gradient of concentration. Note that, in the initial
situation, in spite of concentration of j being greater in B, the electrochemical. potential gradient is
still directed 'downhill' towards B. As B fills with ion j, µ_j flattens out and, at equilibrium becomes zero—
at this point the 'uphill' concentration gradient and the 'downhill' electrical gradient are equal and opposite.

routinely. The electrical potential difference across membranes can be measured if a small glass micro-electrode, with a tip diameter of 1 to 3m m, can be inserted into the membrane-bounded compartment (see Clarkson, 1974).

Having made the necessary measurements a simple test can be applied to see if a given ion or solute within the compartment is at a higher or lower potential than in the surrounding solution. The principal snag in this analysis is that the cell or compartment should be in a steady state and that no net movement of solute should be occurring. In nature this condition is infrequently met.

Let us suppose that the ion j is at electrochemical equilibrium between the two compartments i.e.:

re-writing equation 2.3 and cancelling out

we have

― 48 ―

gathering the electrical terms to the left-hand side we get

yⁱⁿ —y^out is the electrical potential difference across the membrane where the ion j is at equilibrium and is given a special name, the Nernst Potential, and is usually symbolized Ej^N . We now compare this calculated equilibrium potential with the potential difference which is actually measured by the electrodes on either side of the membrane. If the calculated and observed values coincide we would conclude that, in spite of any differences in C_j across the membrane, the system was at equilibrium. If, however, the observed potential was lower than the equilibrium potential we would conclude that the electrical driving force was not sufficiently large to support the observed asymmetry of C_j and we would suspect that active transport was occurring. The example worked out in Table 2.8 may make this clearer. For each ion the appropriate Nernst Potential has

Table 2.8. Comparison of Nernst Potentials and equilibrium concentrations of ions with observed values from a hypothetical cell
Observed potential E = – 116 mV Temperature 293°K (20°C)
	Concentration		E_jⁿ (mV)	E_j (mV)	Equilibrium concentrat (mM )
Ion	Outside (mM )	Inside	E_jⁿ (mV)	E_j (mV)	Equilibrium concentrat (mM )
K⁺	1	100	–116	0	100
Na⁺	10	50	–41	–75	1,000
Cl^–	11	150	+65	–181	0.11
The Nernst Potential for each ion was calculated from equation 2.5. The column E_j is the difference between the Nernst Potential and the observed potential. Where the observed potential E is the same as the Nernst Potential the ion is at electrochemical equilibrium. If the sign of this difference is negative it means that, for cations, the electrochemical potential is 'downhill' into the cell; for anions, however, it signifies an 'uphill' gradient. In the last column, the equilibrium concentration inside the cell, has been computed on the assumption that the observed potential was equal to the Nernst Potential for each ion. Departures between this computed value and the observed concentrations indicate how far the cell is from being in a passive equilibrium with respect to the ion.

been calculated from the observed concentrations on the outside and inside of the membrane using equation 2.5. The exact correspondence of the Nernst Potential for potassium,

, with the measured electric potential difference shows that potassium ions inside the cell are at electrochemical equilibrium with those outside. For sodium, however,

is much less negative than E, hence Na⁺ is at a lower electrochemical potential inside; there is, therefore, a strong tendency for sodium to diffuse into the cell, and the fact that the observed

― 49 ―

concentration is only 1/20 th of the equilibrium concentration strongly suggests that metabolic energy must be coupled to a Na⁺ -efflux pump. Chloride ions in the cell are a very long way indeed from being in electrochemical equilibrium with their surroundings, being more than 1,000 times greater than the equilibrium concentration, thus their movement into the cell is steeply uphill.

In theory this type of analysis can be applied to any ion, although it is difficult to apply to minor ionic constituents, e.g. trace elements, because they may be complexed with organic ligands within the cell so that their ionic activity may be very much lower than their concentration as measured by chemical analysis.

If an analysis of the kind described in Table 2.8 shows that the transport of an ion in a given direction is 'uphill', one should not conclude necessarily that the membrane is equipped with a special pumping mechanism for that ion. In some cases it may be, but in others the 'uphill' transport of the ion may be coupled with the 'downhill' transport of another via a common carrier; this latter possibility is described under the heading Co-transport on p.56.

2.4.3—
The Nature and Origin of the Membrane Potential

It is clear that electrical potential differences across membranes are of great importance in generating driving forces on ions. It is important, therefore, to try to understand how these potentials arise and how they are maintained.

An electric potential difference arises because positive and negative charges become separated. Since the cytoplasm of most cells is electrically negative relative to the surroundings it is very slightly enriched in anions relative to cations. This can be attributed to the differential permeability of the cell membrane and to the activity of ion pumps. First let us examine how differential permeability can create an electrical potential difference.

2.4.3.1—
Diffusion Potential

Imagine a simple system of two compartments separated by a membrane which has a much higher permeability to K⁺ than to Cl^– (Fig. 2.13). If the compartments are filled with potassium chloride solutions of different concentration, initially K⁺ will move through the membrane out of the more concentrated compartment, and for a very brief period, the more concentrated cell will lose K⁺ faster than Cl^– leaving it enriched in negative charge. The negative diffusion potential thus created slows down the further escape of K⁺ by attracting it back into the more concentrated compartment. When the potential has developed, a large concentration difference can be maintained between the compartments. Since the membrane has a finite permeability to Cl^– , albeit a low one, over a long period of time both the electrical potential difference (the diffusion potential) and the concentration difference would run down as Cl^– leaked through the membrane. If the membrane were completely impermeable to Cl^– , the

― 50 ―

potential, once established, would be maintained indefinitely. In nature, membrane permeability to anions is a tenth to one hundredth of that for the monovalent cations; thus, the maintenance of a potential of the kind just considered depends on topping up the cell with anions at a rate comparable with their leakage into the surroundings. This is an 'uphill' transport and therefore requires the mediation of some ion pumping mechanism. Active transport is, therefore, necessary to maintain a diffusion potential.

Figure 2.13
Development of charge separation and a diffusion potential
in a model system containing a membrane selectively
permeable to cations. For further explanation see text.
(From Clarkson. 1974.)

In nature it is frequently possible to find cells where the electrical potential difference across the plasmalemma is, indeed, a diffusion potential of the kind just described which depends very closely on the concentration of either K⁺ or H⁺ in the medium and in the cytoplasm. In such circumstances its value can be predicted from the Goldman equation (2.6) which relates the concentration ratios of ions across the membrane and their permeabilities (P_K , P_Na , P_Cl etc.).

― 51 ―

This relationship will apply strictly only to situations where the cell and the surroundings are in a steady state and hence limits its application to mature and non-growing cells. The last pair of terms in equation 2.6 has been put in to emphasize that other diffusing ions can be added to the equation; clearly H⁺ has an important effect on the membrane potential in some instances (Kitasato, 1968). Since the concentration ratio is multiplied by the permeability coefficient, the value of E will be most strongly influenced by the ionic asymmetry of the most rapidly diffusing ion. In the system considered above the value of E would have been given by

and be governed almost entirely by K⁺ since P_Cl was very small compared with P_K . Notice that the ratio of the chloride terms is inverted relative to the cationic terms. This is explained because the chloride concentration differential will tend to reduce any negative electrical potential set up by the asymmetric distribution of the cations.

2.4.4—
Membrane Pumps

Pumping mechanisms which allow cells to accumulate solutes up gradients of potential can contribute to the membrane potential in both an indirect and a direct way. The former type are referred to as neutral exchange pumps in which the cell dumps an unwanted ion of equal and like charge into the external environment in a one-to-one exchange for an ion which is more useful, e.g. there are well known exchanges of cellular Na⁺ or H⁺ for K⁺ from the surroundings. The second type transports an ion in one direction only without coupled exchange and is known as electrogenic since charge is separated. They can, therefore add to, or subtract from the diffusion potential, described above, depending on which ion is carried. These two types of mechanisms are outlined in Fig. 2.11.

2.4.4.1—
Neutral Ion Pumps

Neutral ion pumps are present in the plasmalemma of all cells and by their activity they create the ionic asymmetry necessary to set up the diffusion potential described above. As a much simplified illustration of this, consider a cell which, because of its synthetic and respiratory activity, is generating H⁺ and HCO₃^– internally. A pair of exchange pumps could swap H⁺ and HCO₃^– for K⁺ and Cl^– from the surroundings quickly enriching the interior in these ions and thus setting up the conditions in which the differential rates of diffusion

― 52 ―

of K⁺ and Cl^– out of the cell give rise to a membrane potential. But how does such a pump actually work? There are fewer detailed examples than one would like but the best known is of the membrane-bound ATPase which exchanges intracellular Na⁺ for extracellular K⁺ in many animal and plant cells (see Hall, 1971; Hodges et al., 1972). In the red blood cell it is known that Na⁺ is one of the cofactors which is essential for the binding of ATP to the ATP-ase enzyme. In vivo the active centre of the enzyme is accessible only from the cytoplasm, so that the ATP and the Na⁺ must be inside the cell (Fig. 2.14). Once bound, the ATP

Figure 2.14
A highly simplified illustration of the working of a sodium-potassium exchange pump based on a
membrane-bound ATPase. The cross hatched area on the ATPase is its active centre. The large
re-orientation of the molecule is for illustrative purposes only—quite subtle molecular re-arrangement
may be all that is necessary to expose the Na⁺ - binding site to the outside and for the step called relaxation.

is hydrolysed, ADP is released into the cytoplasm leaving the cleaved terminal phosphorus atom attached to the active centre to form a phosphoenzyme. These reactions result in some molecular re-orientation of the phosphoenzyme and its attached Na⁺ which exposes the ion-binding site to the different chemical environment of the external medium. It is proposed that this change of environment alters the ion-specificity of the binding site so that K⁺ is favoured; K⁺ thus replaces Na⁺ . This done, there is a second re-orientation (referred to as 'relaxation' in Fig. 2.14) which carries the bound K⁺ to the inside. The phosphorus is released from the active centre and Na⁺ , which is preferentially bound on the cytoplasmic side exchanges for K⁺ and the pump is ready for a second cycle. The pump has used the free energy released on hydrolysis of ATP as fuel to exchange K⁺ and Na⁺ against their respective electrochemical potential gradients. The two ions in the appropriate orientations are essential cofactors in the enzyme reaction; in vitro, ATPase of this kind will not hydrolyse ATP unless Na⁺ and K⁺ are both present.

In theory many pumps based on ATPase are possible with only subtle modifications of the ATPase molecule to provide binding sites of varying field

― 53 ―

strength which will select various ions, e.g. a Ca²⁺ transporting ATPase is found in mitochondria and in sarcoplasmic reticulum (Racker, 1972).

2.4.4.2—
Electrogenic Pumps

The unidirectional transport of an ion across a membrane separates charges and in so doing provides a driving force for the passive diffusion of a similarly charged ion in the opposite direction or an oppositely charged ion in the same direction. The molecular details of exactly how an electrogenic pump is put together remain uncertain although in one instance it is highly likely that an electrogenic H⁺ -efflux pump is based on an ATPase (Slayman et al., 1973). It is possible, nevertheless, to deduce certain general consequences of their operation. If, for instance, there was an outwardly directed pump at the plasmalemma which actively pumped hydrogen ions (protons) out of the cell thus making the interior electrically negative, this could contribute to the electrical driving force on the diffusion of K⁺ from the external medium. Indeed the rate at which charge is extruded and the rate at which it leaks back into the cell must be very nearly in balance unless a dangerously large potential is to accumulate. Examples of both proton extrusion pumps and anion influx pumps of the electrogenic kind are well documented from research on plant tissues (Higinbotham & Anderson, 1974; Spanswick, 1972). In the giant alga, Acetabularia, an electrogenic chloride influx pump contributes more than half of the potential of –170mV found across the plasmalemma when the cell is kept in the light. Almost immediately the cell is put in the dark the pump stops working (since it is closely linked with photosynthesis) and the membrane potential abruptly depolarizes to –80mV (Saddler, 1970). A similar light-dependent electrogenic pump is found in Nitella translucens (Spanswick, 1972, 1974). Electrogenic pumps are not, however, restricted to green tissues but have been reported in plant roots (Higinbotham et al., 1970) and fungal hyphae (Slayman, 1970). In every instance, however, inhibition of the pump caused an immediate depolarization of the membrane potential, indeed this is often used to detect the activity of such a pump. The inhibition of a neutral ion pump gives rise to gradual depolarization as the ionic asymmetry runs down (see equation 2.6).

2.4.5—
Membrane Carriers

Many ions and uncharged solutes cross membranes more rapidly than could be expected if they passed through the membrane lipids without some assistance. Since pores in membranes are too narrow to accommodate many substances which are transported, it is widely believed that membranes contain carrier molecules which, in combination with the solute, facilitate diffusion. The ion pumps considered above are carriers of a special kind since they are linked to

― 54 ―

the metabolic activity of the cell; the carriers we shall now consider promote net movements of solutes only down the prevailing potential gradient and are not capable of 'uphill' transport even if in some instances (see p. 57) they appear to be doing so.

2.4.5.1—
Evidence from Kinetics

One widely used approach to gather information about carriers has been the application of kinetics first derived from enzyme reactions. The evidence for these carriers is obtained by placing a cell or tissue in a range of solute concentrations and measuring the initial rate of uptake. As illustrated in Fig. 2. 15 the uptake rate shows a tendency to saturate at higher concentrations and can thus be used to calculate the maximum velocity, V_max , possible under the conditions used in the experiment. By making a double reciprocal plot of the data (Fig. 2.15) the concentration of solute at which half maximal velocity is achieved can

Figure 2.15
Saturation kinetics of solute uptake versus concentration. Such results are used
as evidence for the association of the solute and a carrier. The double reciprocal plot of
the data gives a more accurate estimate of V_max and K_m when the number of points is limited.

be estimated. This is known as the Michaelis Constant, K_m . The K_m measures the affinity of the carrier for the solute it carries; if the affinity is high then the concentration, K_m will be low and vice versa. For most ions in plant tissue K_m is quite low, concentrations ranging from 5–100 mM , but for sugars and other metabolites K_m values are usually greater than 300 mM . Much work of this kind is summarized by Epstein (1972) who shows that at concentrations less than 0.1 mM , the uptake of a given ion is not subject to serious interference from other

― 55 ―

common ions in solution. There is, however, competitive inhibition between related ions of similar molecular dimensions, e.g. K⁺ uptake is inhibited competitively by Rb⁺ but not by Na⁺ ; Ca²⁺ is inhibited by Sr²⁺ but not by Mg²⁺ . Thus the carriers which bind the major nutrient ions at low concentrations appear to be highly ion-selective. At higher concentrations (more than 1.0–10.0 mM ) this selectivity begins to decline. The interpretation of this observation is contentious and beyond the scope of this chapter but can be pursued in Epstein (1972), Laties (1969) and Clarkson (1974).

The limitation of the kinetic approach is that it can tell us nothing about the nature of the carrier. One can observe similar uptake kinetics for ions whose transport into the cell must be mediated by ion pumps e.g. H₂ PO₄^– and Cl^– (see p. 48) as for ions which probably diffuse into the cell passively e.g. Na⁺ and Ca²⁺ and for those which are completely exotic and toxic, e.g. Tl⁴⁺ (Barber, 1974). Indeed, it has been pointed out that saturation kinetics of this kind would also be found if salt movement was observed across a synthetic membrane containing nothing but pores (Stein & Danielli, 1956), where the system would saturate when all of the pores were filled with solute at any moment in time; V_max is, after all, merely a measurement of capacity to react or transport and K_m is derived from it (Fig. 2.15).

2.4.5.2—
Ionophores as Lipophilic Carriers

A more illuminating approach to the nature of carriers has come from studies on the ionic conductance of synthetic membranes which have been modified in various ways. A bilayer of pure phospholipids has a very low conductance to ions, usually only 10^–7 to 10^–8 ohm^–1 cm^–2 . The addition of very small amounts of ionophores (i.e. ion-carrying antibiotics) like monactin or valinomycin to the solutions bathing the synthetic bilayer causes a huge increase in the conductance. Figure 2.16 shows that 10^–6M monactin changes the membrane conductance to K⁺ nearly a million-fold and that even at 10^–10 M its effect is quite strong. The conductance change for a given monactin concentration is greatest for K⁺ and for Rb⁺ and is much less for Na⁺ , Cs⁺ and Li⁺ . Monactin is, therefore, acting as a selective carrier of K⁺ and Rb⁺ and valinomycin, another bacterial product, behaves similarly. Since these two compounds differ chemically it is instructive to see what they have in common. Both of them are amphipathic ring-structured molecules which have their non-polar groups on the outside of the ring and their polar groups directed towards the space at the centre of the molecule. The outside of the ring interacts favourably with lipid while the hydrophilic core, 0.7 nm in diameter, provides room for several hydrated potassium ions to be bound. Evidence from a variety of sources shows that this complex diffuses across the membrane so that the ions never leave a polar environment (Eisenman et al., 1968).

Other substances are known which select for divalent cations, e.g. the unnamed compound A23187 which is a carboxylic acid antibiotic found in

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Figure 2.16
Influence of the ionophore, monactin, on the electrolytic conductance of a
phospholipid bilayer in the presence of single salt solutions of alkali cations.
(Redrawn from Eiseman et al., 1968.)

cultures of Streptomyces chartreusensis (Reed & Lardy, 1972). This compound carries Ca²⁺ and Mg²⁺ across bilayers and natural membranes but has no effect on monovalent cations.

Apart from a few synthetic analogues all of the ion-carrying antibiotics are natural products of bacteria and fungi. There are many who believe that compounds of a similar kind may act as ion carriers in all membranes but the technical difficulty of isolating what are probably minute quantities of such compounds from tissues appears to be formidable and so the belief may rest on faith for some time yet.

2.4.5.3—
Co-transport

As suggested earlier, carrier-assisted diffusion can sometimes appear to go in an 'uphill' direction, thus giving the impression of active transport. In many animal tissues and micro-organisms, sugars, amino acids, organic acids and vitamins move into the cell up a concentration gradient. This transport is, however, almost completely dependent on having Na⁺ or H⁺ in the external medium; other ions such as K⁺ , Rb⁺ or Li⁺ cannot be substituted. It has been

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found that the metabolite is carried into the cell along with an ion-carrier complex which is diffusing 'downhill' (Fig. 2.17). In both Chlorella and Neurospora, glucose is transported in this way along with protons, H⁺ (Komor & Tanner, 1974; Slayman & Slayman, 1974).

Figure 2.17
Scheme to illustrate co-transport of protons and sugar. The proton extrusion pump is electrogenic
and thus makes the inside electrically negative. Protons diffuse back into the cell passively via the
carrier which also binds a sugar molecule. The protonated carrier plus sugar diffuses towards the
inner face of the membrane where it dissociates and releases the sugar molecule.

Co-transport depends on active transport in an indirect way (as indeed does all diffusion, see p. 50) because energy-dependent extrusion pumps ensure that the cytoplasm is kept well below its equilibrium concentration in H⁺ and Na⁺ . These ions tend to diffuse back into the cell and, in doing so, decrease their free energy. The energy they give up is coupled, via the carrier, to the co-transport of the solute whose free energy is increased as it moves into the cell.

Co-transport may also assist the 'uphill' movement of inorganic ions into the cell; it may be a more common process than is generally realized. Recent

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evidence by Lowendorf et al., (1974) suggests that the active transport of phosphate into the hyphae of Neurospora depends on (a) the activity of a proton extrusion pump at the plasmalemma which is sensitive to the pH of the external medium, and (b) the formation of a ternary complex between a proton and a phosphate ion from the external medium with a membrane carrier. The protonated phosphate carrier diffuses to the cytoplasmic side of the membrane down the electrochemical gradient of the proton, releasing the proton and phosphate ion into the cytoplasm. This, and other examples of inorganic ion co-transport (see Raven & Smith, 1974) suggest a reason why so little progress has been made in elucidating the molecular details of certain 'pumps' particularly of those which transport anions. Put most simply, it may be that these influx pumps do not exist and that the uphill transport is driven by a combination of active extrusion and re-entry by diffusion of protons or perhaps sodium ions.

Co-transport illustrates the ingenious way in which nature can turn necessity to its own advantage. The active excretion of H⁺ and Na⁺ is essential to maintain pH control and osmo-regulation in the cell, but the energy expended is partly recovered in the transport of essential metabolites and ions into the cell.

2.5—
Correlation of Structure and Function

Having a picture in mind of the way membranes are assembled and of the driving forces which operate across them it is now possible to consider how their design, particularly that of the plasmalemma, manages to reconcile two conflicting sets of priorities. This conflict may be summarized as follows: the cell usually contains solutes at a far higher concentration than in the external medium; without any barrier these solutes would disperse by diffusion. Many solutes are accumulated by mechanisms which consume metabolic energy and it is sensible for nature to minimize their subsequent leakage and thereby economize in the use of 'fuel'. Insulation of this type might be most effective if the cell were surrounded by a barrier impenetrable to virtually everything except water and dissolved gases. Such a barrier would, however, isolate the cell from the outside world and render it insensitive to stimulae and incapable of growth, it might eliminate its interactions with other cells in adjacent or remote tissues and prevent the excretion of harmful excesses of substances generated during metabolism. The cell membrane must provide a balance, therefore, between minimizing losses and permitting a selective interchange of substances between the cell interior and the surroundings. The balance is achieved, in many cases, by providing channels or other more elaborate carrier mechanisms whereby materials crossing the membrane may essentially by-pass the hydrophobic region.

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2.5.1—
Membrane Pores and Channels

Lipid or oil is a very effective electrical insulator (it is used in high voltage underground electric cables for this purpose) and it is not surprising to find that the electrical resistance of synthetic bilayers made from pure phospholipid is very high, being 10⁷ to 10⁹ ohms cm² . In nature, the current conducted across membranes is carried by ions and we can see that an unmodified lipid bilayer with such a high resistance is a poor material across which to conduct this essential current of electrolyte. It is not surprising to find, therefore that the electrical resistance of natural membranes is very much less than that of bilayers, usually lying in a wide range of 10² to 10⁵ ohms cm² .

As described on p. 55, lipophilic carriers can greatly reduce the resistance (and hence increase the c onductance) of synthetic membranes and may resemble the carrier molecules in membranes, but it is also widely believed that membranes contain water-filled pores which must contribute to their relatively high conductance by allowing water and selected solutes to by-pass the lipid domain of the membrane. Certain antibiotic molecules, such as nystatin, appear to condense cholesterol molecules in both synthetic (Holz & Finkelstein, 1970) and natural membranes (de Kruijff & Demel, 1974) to form pores with a radius of ca. 0.4 nm. The presence of such pores greatly increases the electrical conductance and hydraulic conductivity of the membrane. In passing we might note that many antibiotics have their effect by enormously increasing the passive permeability of cell membranes causing non-resistant cells to lose their contents or to lyse. Interest in these substances stems from the experimental evidence that cell membranes also possess pores of similar size and that the antibiotic merely induces an extreme expresssion of the normal condition.

Although the word 'pore' is often used to describe channels through which solutes and water can move, we should resist the temptation to conclude that all 'pores' are definable structural entities like the ones induced by nystatin (see above). In many instances a 'pore' may be more like a transient imperfection in membrane structure. This latter type may have a certain statistical probability but have no fixed position.

Proteins which traverse the lipid layer may give rise to hydrophilic channels or pores (see Fig. 2.5e). Indirect evidence supporting this idea comes from a study in which red blood cell membranes were exposed by deeply etching frozen cells under vacuum (Pinto da Silva, 1973). Shrinkage of the membrane surface was observed in areas overlying groups of membrane particles possibly due to sublimation of ice from within the embedded particles; for this to have occurred this water would be a free liquid in the thawed condition (cf. bound water p. 35).

The first circumstantial evidence for the existence of pores come from studies by Collander and Barlund (1933) on the permeation of the giant internodal cells of the alga, Chara ceratophylla, by a number of uncharged solutes and water. They found that in almost every case the rate of movement of a substance into a cell depended on its molecular weight and dimensions and on its solubility

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in oil relative to water; substances with high oil solubility permeated most rapidly. This strongly suggested that movement across cell membranes involved the movement of the solute out of the water, its solution in lipid and subsequent diffusion through it, and its re-entry into the aqueous phase at the inside face of the membrane. The authors found, however, that several small molecules permeated the membrane far more rapidly (more than 100 times faster in the case of water) than their relative solubility in oil suggested. The upper size limit for molecules which behaved anomalously was a radius of 0.4 nm and it was suggested that the membrane was constructed as a very fine sieve containing pores of 0.4 nm radius through which water (0.25 nm radius) and certain solutes could move. Since this early work a great deal has been done and the equivalent pore radius in many plasma membranes has been confirmed as 0.4 nm (see Solomon & Gary-Bobo, 1972). It must be said, however, that some authorities are reluctant to accept that pores can provide channels for the bulk movement of water and solutes; the arguments for and against pores have been clearly discussed in Oschman et al., (1974).

There has been much discussion about whether pores admit ions, and if so, which ones. Membranes are known to control very precisely the relative rates at which various ions will diffuse across them; potassium ions will diffuse ten to one-hundred times faster than sodium. Ions in solution are hydrated by binding one or more water molecules; it requires a great deal of energy to dehydrate an ion and for this reason we should think of ions in all natural circumstances as being hydrated. Sodium binds 5 water molecules, whereas potassium binds 3, the former is, therefore, the more bulky ion whose diffusion into the narrow water filled pores would be slower than for the smaller hydrated potassium ion. On the other hand the anion chloride, which has only one water molecule in its hydration shell, diffuses much more slowly than either K⁺ or Na⁺ . This is probably due to the fact that 'pores' carry a predominantly negative charge so that cations would be attracted to them, while anions would be repelled—a small number of positively charged pores would handle the flow of anions. Polyvalent cations and anions have much more water in their hydration shells, e.g. Ca²⁺ has 10 and SO₄^2– has 8 and it seems likely that these would be totally excluded from the pores.

2.5.2—
Ion Pumps and Membrane Substructure

It is most unlikely that the intercalated membrane particles described earlier (p. 38) are simple ion-carriers—they are much too large—but they may be ion pumps, or groups of pumps. The best evidence for this assertion comes from work on the ATPase which pumps Ca²⁺ across the membranes of sarcoplasmic reticulum. Racker (1972) was able to isolate this enzyme and put it back into a synthetic bilayer membrane made from soybean phospholipids, and thus reconstitute a membrane which could actively transport Ca²⁺ when ATP and Mg²⁺ were present in the medium. Working with a slightly different system

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Packer et al., (1974) showed that freeze etched fractures of both the original tissue membrane and reconstituted membrane containing the purified ATPase were densely studded by numerous particles 8.5 nm in diameter. There can be little doubt that the particles were the calcium-pumping ATPase. Similar, less complete, evidence is also available for the Na⁺ /K⁺ -dependent ATPase of the red blood cell (see Branton & Deamer, 1972). There is no insuperable difficulty in repeating such observations with plant tissues but, at the time of writing there is no report that this has been done.

One should not conclude that all of the membrane substructure is concerned with solute transport, especially in mitochondria and chloroplasts where other biochemical activities are centred on membranes.

2.5.3—
Endocytosis and Vesicular Transport

The presence of abundant vesicles in the vicinity of the plasmalemma of plant cells has been pointed out earlier (p. 44); however, it is not yet clear what contribution endocytosis makes to the total solute transport across this, or any other membrane. There is kinetic evidence from several sources which is consistent with the notion that a measurable fraction of various ions in the cytoplasm of plant cells is sequestered into a compartment separate from the bulk of the cytoplasm. MacRobbie (1969) found that labelled Cl^– transported into the vacuole of Nitella translucens could be easily resolved into a fast and a slow component. The fast component was envisaged as being delivered to the vacuole formed in vesicles which had been found at the plasmalemma. The chloride ions in these vesicles would not have mixed with the unlabelled chloride in the bulk cytoplasm, whereas labelled Cl^– delivered directly to the cytoplasm via a pumping mechanism would have to mix with a much larger pool of unlabelled chloride ions. At the beginning of the experiment, therefore, the labelled Cl^– in the vacuole increased more rapidly than expected. There are other examples of this kind discussed in MacRobbie (1971) and Baker and Hall (1973).

If the vesicles contained nothing more than a small volume of the outside solution, their net contribution to the solute content of the cell would be very small and they could not exercise the ion-selectivity which characterizes cell membranes. Baker and Hall (1973) suggest that endocytosis becomes a much more plausible transport process if it is assumed that ions become selectively bound to the membrane surface and thus become concentrated from the dilute external medium. This assumption is entirely reasonable because both proteins and phospholipid heads can provide ion-selective binding sites. These authors also relate membrane-bound ATPase activity with the subsequent invagination of the plasmalemma to form a vesicle but this implies that vesicle formation is energy-dependent. This is against the substantial evidence from animal cells which shows that micro-vesicle formation results from Brownian movement of the membrane and does not need to be energized by ATP (Casely-Smith, 1969); ATP is required for the very large vesicles formed by re-orientation of micro-

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fibrils in phagocytosis, but the type of vesicle under discussion here is much smaller, probably 70–100 nm in diameter and is not dependent on microfibrils. There is visual evidence that these micro-vesicles can discharge their contents directly into lysosomal vacuoles (Casely-Smith & Chin, 1971).

The important advance of thinking required to deal with endocytosis will provide a severe strain on biophysical theorists accustomed to thinking of cells as homogeneous phases separated by diffusion barriers, partly because materials can reach the interior of the cell without crossing a membrane at all. Although endocytosis is probably a common phenomenon it is likely to remain an 'unknown quantity' until new experimental techniques are devised.

2.5.4—
Concluding Remarks

In the light of recent discoveries we must discard the notion that membranes are neat semi-crystalline rigid lattices in favour of a more dynamic view. The diffusion of large protein molecules in the plane of the membrane emphasizes its fluid character and we see that there may be great heterogeneity both in the lipid matrix and in the proteins embedded in it. In the past it is possible that we have over-emphasized the importance of electrostatic interactions between phospholipid heads and underestimated the role of the surrounding polarized water layers in maintaining the familiar bilayered arrangement of membranes.

This more 'fluid' view of membranes has influenced the way in which we think about transport processes and diffusion, particularly those workers with interests in endo- or pino-cytosis. It seems probable that some of the processes which have hitherto been accepted as active transport may be, in reality, examples of co-transport which ultimately depend on the active transport of some other ion. This would allow a simplification of the model of the cell membrane which at present is uncomfortably cluttered by numerous hypothetical ion-pumps. Perhaps in time we shall be able to reduce this picture to one or two pumps and many smaller carrier molecules in a membrane which constantly cuts off vesicles from its undulating surface.

Chapter 3—
Chloroplasts—Structure and Development

3.1—
Introduction

Plastids are organelles which are bounded by double membranes and which occur, as far as is known, in all cells of eukaryotic green plants at some stage, usually becoming modified according to their function. In their undifferentiated form they may remain as proplastids, which are characteristic of epidermal and meristematic cells, for example. In the green parts of plants the proplastids normally develop into chloroplasts, which are the site of photosynthesis, while in starchstoring organs they form amyloplasts which produce the starch grains. However these two functions are not mutually exclusive as most chloroplasts will form starch under appropriate physiological conditions and the exposure of starch-storing organs to illumination results in the amyloplasts forming some thylakoids and chlorophyll. In certain plant parts, such as flowers, fruits and some leaves, the thylakoids of the chloroplasts become degraded, forming chromoplasts, which contain large amounts of carotenoids, the pigments responsible for 'autumn colouration' and the characteristic colours of certain flowers and fruits.

As the whole range of plastid structure, composition, genetics and development has been extensively covered in the monograph by Kirk and Tilney Bassett (1967) this chapter will confine itself to the consideration of chloroplasts, on which recent work has centred. The work of the author's laboratory has concentrated on a study of the structure and development of chloroplasts in Phaseolus vulgaris (bean) and Zea mays (maize) and thus many of the examples are drawn from work with these plants in order to make a consistent account. This approach must induce some lack of balance, which is acknowledged, but a properly balanced account of this topic would virtually require a volume of its own. References have been selected primarily for their clarity and not necessarily for their priority.

It has been postulated that the eukaryotic cell gained its autotrophic capacity by the capture of a prokaryotic organism at an early stage in the evolutionary process (Stanier, 1970), that the trapped prokaryote became the chloroplast, and that the nucleic acid of the resultant chloroplast still retains an important genetic function (see also chapter 9). Although the genetic evidence indicates that the plastid DNA is transmitted from generation to generation independently of the nuclear genes, Bell (1970) has published electron microscopic evidence that proplastids may arise anew each generation from the nucleus. However, it is hard to reconcile this latter proposal with the evidence that both nuclear and plastid DNA contribute to the plastid genotype (see chapter 9).

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3.2—
Chloroplast Structure

3.2.1—
Chloroplast Dimensions and Number

Beans possess chloroplasts typical of the sun leaves of higher plants in that they are discoid or lens-shaped with a diameter of about 5µm and maximum thickness of about 1 µm as may be seen in the electron micrograph in Fig. 3.1B, which is a vertical section through the disc. On the other hand observations of chloroplasts in living cells by phase contrast microscopy suggest that the maximum thickness of the chloroplast may be rather less (S. G. Wildman, personal communication). Under optimum conditions for growth, Phaseolus vulgaris averages 45 chloroplasts per palisade mesophyll cell and 32 per spongy mesophyll cell, giving about 8.3 × 10⁸ chloroplasts per leaf and about 2.3 × 10⁷ chloroplasts per cm² of leaf. This latter number is similar to that of mature spinach leaves (Possingham & Saurer, 1969) although the number of chloroplasts per spinach mesophyll cell is more than ten times greater than for bean.

3.2.2—
Chloroplast Fine-Structure

The following interpretation of chloroplast fine structure is based on transmission electron microscopy of thin sections of leaf material prefixed in 3% glutaraldehyde for 24 hours at 5°C, followed by fixation and staining with osmium tetroxide and post-staining with uranyl acetate and lead citrate. This technique is the most widely used, and, as glutaraldehyde is a relatively mild reagent, is considered to produce images most closely resembling the living plastid. Heslop-Harrison (1966) has written a critical account of the interpretation of chloroplast electron micrographs while the results of freeze-etch studies are considered in chapter 4.

The chloroplast envelope consists of two membranes (Figs. 3.1 and 3.2) the outer of which is unspecifically permeable to crystalloidal solutes. In contrast the inner membrane shows very specific permeability and has so far been shown to be the site of three specific anion translocation systems; (a) the phosphate translocator, facilitating a counter exchange of inorganic phosphate, 3-phosphoglycerate and dihydroxyacetone phosphate; (b) the dicarboxylate translocator, facilitating a counter exchange of dicarboxylic acids; and (c) the ATP translocator which is less active than the previous two and may be responsible for the entry of ATP in the dark (Heldt et al., 1972).

Within the inner membrane is a complex system of flattened sacs of membrane which were first termed thylakoids by Menke (1962). The thylakoids are closely associated in stacks (grana) which are shown in section for chloroplasts of bean and maize mesophyll in Fig. 3.1 (B and C). When seen from above the plane of the membrane the grana are essentially circular in outline with an average diameter of about 0.5 µm. Under optimum growing conditions bean grana

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Figure 3.1
Plastids of bean and maize. A, proplastid from the primary leaf of a six day-old dark-grown
bean (× 42,000); B, chloroplast from the primary leaf of a bean grown for 14 days in the dark
and then transferred to continuous illumination (3 mW. cm^–2 ) for 48 hours (× 29,000); C, bundle
sheath and mesophyll chloroplasts of maize grown under normal diurnal illumination (×12,500).
Scale lines represent 1 µm. Key to lettering: BS, bundle sheath; E, plastid envelope; EOL, electron
opaque layer; G, granum; I, invagination of envelope inner membrane; M, mesophyll; OG, osmiophilic
globule; PLS, porous lamellar sheet; PR, peripheral reticulum; S, starch grain; T, thylakoid.

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contain up to 8 thylakoids while those of maize have up to 40. The thylakoids in each granum are continuous with those in other grana through intergranal thylakoids. In bean the granal thylakoids average 78%, and the intergranal thylakoids 22%, of the total thylakoid membrane (Bradbeer et al., 1974a). During chloroplast development the inner membrane of the envelope appears to give rise to the thylakoids by invagination (see Fig. 3.1A and p. 75) and it is possible that the space between the inner and outer membranes of the envelope is continuous with the whole of the thylakoid space of the chloroplast. Although the magnification of Figs 3.1B and C is such as to allow an overall impression of chloroplast structure, enough detail is visible to show that the intra-thylakoid space is electron-transparent, the thylakoid membrane is electron-opaque and the inter-thylakoid space seen in the grana between adjacent thylakoids is densely electron-opaque.

In the chloroplast the thylakoids are embedded, or suspended, in a matrix, the stroma, which has a somewhat granular appearance (Fig. 3.1). Within the stroma may be seen DNA fibrils and ribosomes (chapter 9), starch grains, osmiophilic globules and occasional extensive crystal-like structures. Such crystals can often be induced to appear in chloroplasts suspended in hypertonic media or in plants subjected to stress, although they have also been observed in plants grown under normal environmental conditions. It is thought likely that such crystals are composed of ribulosebisphosphate carboxylase (Larsson et al., 1973).

3.2.3—
The Mobile and Stationary Phases of Chloroplasts

From observations of chloroplasts in living cells by phase-contrast microscopy Wildman (1967) distinguished two components of the chloroplasts, a stationary phase which he equated to the thylakoid system and a mobile phase which he equated to the stroma. The mobile phase surrounds the grana and also penetrates the intergranal regions of the chloroplast. The mobile phase is always in some kind of motion though the intensity of the activity shows variability, even in the same chloroplast. In the reported investigations, observations were always made in cells which could be seen to be living by the presence of visible protoplasmic streaming. In such cells the mobile phase formed protuberances which sometimes broke away from the chloroplast and became indistinguishable from

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mitochondria, and also mitochondria-like bodies appeared to fuse with the mobile phase. Subsequently Wildman and coworkers (1974) have reported that mitochondria-like bodies became stationary below tobacco chloroplasts in living cells and that starch grains subsequently appeared in similar positions in the chloroplasts. In the opinion of the present writer the interpretation that mitochondria give rise to starch grains cannot be substantiated. We have found that in plants grown under low light intensities each chloroplast has one or more mitochondria embedded in deep pockets close to the chloroplast margin but that no break in the chloroplast envelope occurs and there is always at least a thin layer of cytoplasm between chloroplast and mitochondrion (Montes & Bradbeer, 1976). It is possible that some of the observations of Wildman and colleagues may be explained by the development of the latter phenomenon.

3.2.4—
The Range of Chloroplast Structure

Quite early in the application of electron microscopy to plant structure the algae were found to show an interesting range of structural diversity of their chloroplasts. Subsequent studies on vascular plants established that a number of structural types were to be found in both cultivated and non-cultivated plants and furthermore that structural modification might be induced by mutation or by variation of the environmental conditions.

3.2.4.1—
The Algae

A monograph such as that of Dodge (1973) should be consulted for details of the range of chloroplast structure of this group. The characteristics of the chloroplasts have provided an important part of the basis for the taxonomic classification of the algae. In the green algae (Chlorophyceae and Prasinophyceae) the arrangement of the thylakoids tends to be basically similar to that in vascular plants and many of the differences may be associated with the non-discoid shape of the chloroplasts in most green algae. The simplest arrangement is found in the Rhodophyceae where the thylakoids occur as single sheets. In the Cryptophyceae the thylakoids tend to occur as pairs but the individual thylakoids in the pair do not appear to be fused to each other. The other algal classes have their thylakoids arranged in threes; in some cases (Dinophyceae, Euglenophyceae and Haptophyceae) the component thylakoids appear to be fused to each other while in other cases (Chrysophyceae and Phaeophyceae) the thylakoids do not appear to be fused. In two algal classes (Dinophyceae and Euglenophyceae) the chloroplast envelope consists of three membranes instead of two. In a number of classes (Bacillariophyceae, Chloromonadophyceae, Chrysophyceae, Cryptophyceae, Haptophyceae, Phaeophyceae and Xanthophyceae) the chloroplast is also surrounded by a sheath of endoplasmic reticulum which is usually continuous with the nuclear membrane. To complete this brief list of

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the major peculiarities of algal chloroplasts it should be noted that the chloroplasts of many algae possess pyrenoids and eye spots.

3.2.4.2—
The Dimorphic Chloroplasts of C₄ Plants

C₄ plants form oxalacetate, malate and aspartate as the primary products of their photosynthetic CO₂ -fixation in contrast to the more 'usual' C₃ plants whose primary fixation product is 3-phosphoglycerate. Laetsch (1974) lists the following families of flowering plants as containing C₄ species: Amaranthaceae, Aizoaceae, Chenopodiaceae, Compositae, Cyperaceae, Euphorbiaceae, Gramineae, Nyctaginaceae, Portulaceae and Zygophyllaceae. All of these families also contain C₃ plants and there is the case of the genus Atriplex where the C₄A. rosea will hybridize with the C₃A. patula (Björkman et al., 1970). Apart from the difference in primary photosynthetic CO₂ -fixation products C₄ and C₃ plants show other substantial differences in their biochemistry, physiology, anatomy and fine structure. Maize and bean will be discussed here as typical examples of C₄ and C₃ plants respectively

In the maize leaf the chloroplasts are concentrated in two concentric sheaths of cells around each vascular bundle. The inner sheath of cells is described as the bundle sheath and consists of equal numbers of large barrel-shaped cells and smaller cells which can be divided into two sorts on the basis of their dimensions (Montes & Bradbeer, 1975). Laetsch (1974) points out that the thick walls of bundle sheath cells adjoining mesophyll cells in C₄ grasses possess an electronopaque layer (Fig. 3.1C). The bundle sheath chloroplasts possess abundant thylakoids which do not associate into grana, (Fig. 3.1C) and they normally contain starch grains. The plant from which the material was taken for Figure 3.1C had been stored in the dark for 24 hours prior to fixation to remove the starch so as to obtain a clear electron micrograph. In most cases, the bundle sheath chloroplasts of maize have been found to be completely agranal, although by modification of the environmental conditions the formation of grana can be induced (Bradbeer & Montes, 1976). In contrast Laetsch (1974) considers it normal for bundle sheath chloroplasts to show a small amount of thylakoid appression.

The outer sheath of cells is known as the mesophyll sheath and it contains chloroplasts similar to those of C₃ plants in that they have grana. They are, however, different in that they do not normally contain starch grains. Not all C₄ plants show such structural dimorphism of their chloroplasts. However all chloroplasts of C₄ plants possess a system of tubules, the peripheral reticulum, (PR in Fig. 3.1C), which is associated with the inner membrane of the chloroplast envelope. There are reports of peripheral-reticulum-like membrane systems in chloroplasts of some cells of C₃ plants (Laetsch, 1974). Circumstantial evidence assembled by the latter author suggests that the function of the peripheral membrane in C₄ plants may be to facilitate transfer of metabolites between chloroplast and cytoplasm.

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3.2.4.3—
The Effect of Environmental Conditions on Chloroplast Structure

The fact that plants grown in the complete absence of illumination form etioplasts (Fig. 3.2A and section 3.4.2) while those grown under diurnal illumination form chloroplasts establishes that light is an essential requirement for chloroplast development. When Björkman et al., (1972) compared plants of Atriplex patula which had been grown under three different irradiances: 20, 6.3 and 2 mW. cm^–2 (in the waveband 400–700 nm), which are referred to as high, intermediate and low, they found that the high irradiance treatment yielded thicker leaves than the low, with more cells per leaf section and more chloroplasts per cell. The chloroplasts from plants grown under low irradiances were larger than those from high irradiance conditions and they contained more thylakoids and larger grana. The intermediate illumination gave intermediate results. The low irradiance-grown Atriplex plants were compared with plants adapted for growth on the floor of the Queensland rainforest where the daily quantum flux was about one-twentieth of that provided by the low irradiance treatment. The chloroplasts of these plants, Alocasia macrorrhiza, Cordyline rubra and Lomandra longifolia were found to be dark green, unusually large and irregular in outline and to contain very well developed grana of enormous size (Anderson et al., 1973). The grana were also irregularly arranged and apparently adapted for a maximum efficiency in light-trapping.

When maize plants were grown under very low irradiances (0.3 mW. cm^–2 ), which nevertheless provided about six times the daily quantum flux received by the rainforest plants, some chlorophyll and photosynthetic CO₂ -fixation developed even though the light compensation point was not exceeded and the leaves showed a net loss of CO₂ followed by senescence and premature death (Bradbeer & Montes, 1976). Grana did not develop and both bundle sheath and mesophyll chloroplasts developed closely parallel arrangements of thylakoids which did not become appressed. Transfer of plants greened under very low irradiances to higher irradiances resulted in a partial recovery towards normal structure.

Research work in Belgium has shown that exposure of etiolated seedlings to electronic flashes of 1 millisecond in duration and given at 15 minute intervals brought about greening in which the resultant plastids were agranal with parallel unfused thylakoids. These flashed leaves showed the interesting property that, on transfer to continuous illumination, they initially did not show any photosynthetic oxygen evolution, but within 2 minutes of the commencement of illumination oxygen evolution was detected and a maximum rate was found after 6 minutes (Strasser & Sironval, 1972). When etiolated leaves are transferred to continuous illumination without any pretreatment with light the onset of oxygen evolution tends to be considerably delayed. An alternative treatment which has produced almost completely agranal plastids is exposure of dark-grown seedlings to continuous far-red irradiation at wavelengths longer than 700 nm (De Greef et al., 1971). For further discussion see page 80.

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3.2.4.4—
Chloroplast Mutants

Many chloroplast mutants are known with defects in pigments or other components of structure or function. Some of the mutations are nuclear and show normal Mendelian inheritance whilst others show non-Mendelian inheritance and are considered to be mutations of the chloroplast DNA. Chloroplast mutations in algae can be maintained in heterotrophic culture and have been used for important research on chloroplast genetics, development and function (see e.g. Levine, 1969). In contrast, although there have been numerous investigations of individual chloroplast mutations in higher plants, the only substantial research collection of higher plant material with mutant chloroplasts is the barley mutant collection of D. von Wettstein's group in Copenhagen (von Wettstein et al., 1971). This collection has so far not been made generally available for studies on chloroplast development.

3.3—
Isolation of Chloroplasts

Although Hill in 1937, obtained chloroplast suspensions from Stellarsia media which were capable of O₂ evolution under illumination, it was not until 1954 that Arnon et al., reported the occurrence of photosynthetic phosphorylation in isolated chloroplasts. These latter chloroplast preparations were capable of rates of photophosphorylation and electron transport similar to those assumed to occur in intact leaves, but they showed very low rates of CO₂ fixation (<1 µmole CO₂ fixed/hour/mg chlorophyll) compared with the rates found in intact leaves (200 µmole CO₂ fixed/hour/mg chlorophyll). Little improvement in the rates of CO₂ fixation by chloroplast preparations was obtained until Walker (1964) devised procedares which initially gave spinach chloroplasts able to fix 24.3 µmole CO₂ /hour/mg chlorophyll. Subsequent modifications gave chloroplasts with improved rates of CO₂ fixation and in the author's laboratory the following adaptation of Walker's method has been used routinely (Reeves & Hall, 1973). Washed spinach leaves are pre-illuminated for 30 minutes before use. 50 g of deribbed blades are rapidly cut up with a sharp knife to give pieces of about 2 cm^–2 which are placed in a cooled perspex grinding vessel 6 cm × 5 cm × 25 cm. The leaves are then covered with 200 ml of fresh grinding medium which has been partially frozen to a slushy consistency. The grinding medium consists of 400 mM sorbitol, 10 mM NaCl, 5 mM MgCl₂ , 1 mM McCl₂ , 2 mM EDTA, 2 mM isoascorbic acid, 0.4% (w/v) bovine serum albumin and 50 mM 2-(N -morpholino) ethanesulphonic acid (MES) adjusted to pH 6.5 (at room temperature). The leaves are ground for 3 seconds with a Polytron PT₂₀ with a PT₃₅ head (Willems Kinematica GbmH, Lucerne, Switzerland) at a speed setting of 3.5. The resultant slurry is squeezed through two layers of butter muslin (cheese cloth) and the filtrate poured through eight more layers of muslin. The final filtrate is centrifuged in an MSE Super Minor bench centrifuge with a precooled

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head. Rapid acceleration up to 4,000 × g, followed by braking by hand, (only to be attempted after training and with adequate safety precautions) enables the total centrifugation time to be less than 1 minute. The supernatant is discarded and the pellet gently resuspended with the aid of cotton wool and a glass rod in about 1 ml of solution identical to the grinding solution except that the isoascorbic acid is omitted and the MES is replaced by 50 mMN -2-hydroxyethylpiperzaine-N' -2-ethanesulphonic acid (HEPES) as buffer adjusted to pH 7.5 (at room temperature). The procedures are carried out at 0–4°C and the resultant chloroplast preparation is kept on ice. The total preparation time from cutting the leaves should be about 4 minutes. Most of the chloroplasts (60–80%) in such a preparation should be complete chloroplasts with a morphologically and functionally intact envelope and a high rate of light-dependent CO₂ -fixation and O₂ -evolution, similar to that found in vivo. They are considered to possess a full complement of unimpaired photosynthetic reactions although most of the reactions cannot be measured directly as the inner membrane of the envelope is impermeable to most of the intermediates which might be added to test these reactions. S. G. Wildman has demonstrated to me, by phase contrast microscopy, that most of the chloroplasts in this sort of preparation have lost their starch grains and therefore presumably possess resealed membranes.

The literature contains a multiplicity of published methods for obtaining a range of chloroplast preparations. To bring some order to the situation, Hall (1972) devised a scheme of nomenclature for these preparations, the main features of which are presented in Table 3.1. In this scheme the complete chloroplasts of Walker are classified as type A and the remaining types represent a succession of increased degradation. To the present writer the criteria for type B chloroplasts seem to be somewhat obscure and unsatisfactory and Walker (personal communication) is also sceptical about the validity of type B. Type B seems to have been proposed by Hall on the basis of older publications which had reported the preparation of apparently unbroken chloroplasts with impaired CO₂ fixation. The other types of chloroplast preparation listed in Table 3.1 are more clearly defined and they correspond with the present state of knowledge.

For most chloroplast preparations investigators would seem to be best advised to prepare type A chloroplasts as the first step. Disadvantages of this procedure are that type A chloroplast preparations (a) contain a proportion of damaged and fragmented chloroplasts (normally 20–40% of the chloroplasts are not type A), (b) are contaminated with other organelles, (c) are contaminated with a substantial amount of cytosol, and (d) are obtained with a fairly low yield. In any further purification stages such as gradient centrifugation or washing there may be further damage to the chloroplasts and reduction of the yield. Consequently it is not yet possible to obtain a pure preparation of type A chloroplasts which is completely free from all other cellular components.

At the present time the application of similar techniques to those described above have yielded active preparations of type A chloroplasts from a very limited number of species (see e.g. Walker, 1971). For successful chloroplast

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Table 3.1. Types of chloroplast preparations after the classification of Hall (1972) together with an outline of their main properties.
Chloroplast type	Description	Preparation method	Envelope	Rate of CO₂ fixation µmole CO₂ /mg chlorophyll/hour	Electron transport and photophosphorylation capacity
A	Complete chloroplasts	Rapid in isotonic or hypertonic sugar, 1 centrifugation	Intact	50–250	Presumed to be unimpaired.
B	Unbroken chloroplasts	In isotonic or hypertonic sugar or salt with 2 or 3 centrifugations.	Morphologically but not functionally intact	<5	Good
C	Broken chloroplasts	Vigorous preparation in isotonic sugar or salt.	Removed during preparation	Little or none	Good, but addition of ferredoxin is necessary for NADP reduction.
D	Free-lamellar chloroplasts	Osmotic shock of type A chloroplasts immediately followed by return to isotonic medium.	Lost from chloroplasts but retained in medium.	Appreciable if carbon-pathway intermediates added.	Good
E	Chloroplast fragments	Resuspend chloroplasts in hypotonic medium.	Lost	None	Ferredoxin required for NADP reduction. Better rates of electron transport and lower rates of photophosphorylation than by types B and C.
F	Sub-chloroplast particles	By sonication or treatment with detergent or French press. Fractionation by differential centrifugation	Lost	None	Photophosphorylation (cylic) low or absent. Limited electron transport when electron donors added.

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preparation it is usually necessary to modify the standard procedures, but despite modification some species have failed to yield appreciably active chloroplasts.

There are also two very different ways of obtaining plastid preparations which are worth noting. Wellburn and Wellburn (1971) devised a method of purifying structurally intact etioplasts by re-suspending the centrifugation pellet and passing this material through a loosely packed column of coarse Sephadex G-50. In vitro developmental studies have been carried out with such preparations (Wellburn & Wellburn, 1973), which show structually intact envelopes, although the biochemical properties of such etioplasts and the functional intactness of the envelopes do not appear to have been established so far. Alternatively, fractions rich in chloroplast material have been obtained by the non-aqueous homogenization and fractionation of freeze-dried leaf material (Stocking, 1971). Although important data have been obtained from non-aqueous preparations, the method is both technically difficult and hazardous, the preparations are impure and many of the biochemical reactions of the chloroplast are destroyed during the preparation.

3.4—
Chloroplast Development

Although the study of chloroplast development and the onset of photosynthesis in seedlings has attracted attention during essentially the whole of the twentieth century (Irving, 1910) it is in recent years that most interest has been shown. There have been two main methods of conducting this study, of which the first is essentially concerned with chloroplast development in plants grown under natural environmental conditions. Since natural conditions are normally neither constant nor consistent such studies have often been conducted in controlled environmental chambers set to a standard day length and standard conditions for day and night. When it became evident that illumination was the critical environmental factor controlling chloroplast development it became fashionable to study chloroplast development by allowing the seedlings to grow for an initial period of several days in complete darkness prior to their transfer to continuous illumination. Both approaches have their merits and it is clear to the present author that both are necessary to obtain an understanding of chloroplast development. Obviously seedling growth under natural environmental conditions shows the normal state of chloroplast development but it provides two main experimental difficulties. Firstly, under diurnal conditions there is a gradient of chloroplast development both within the plant and within the leaf, such that any experimental analysis is either very difficult or impossible. The second difficulty is that diurnal conditions make it difficult to distinguish the effects of illumination on chloroplast development. If seedlings are grown in continuous darkness for a sufficiently long period, their development reaches a stationary phase in which all of the developing plastids in a leaf or a section of a leaf show

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the same stage of growth. On transfer to illumination the subsequent development then tends to occur in a synchronous manner, thus facilitating microscopic analysis and making biochemical analysis feasible. Such treatments may enable leaf chloroplast development to show synchrony like that obtainable in certain microbial cultures. It should also be pointed out that much progress in the study of photomorphogenesis has depended on illumination treatments of such darkgrown seedlings (Mohr, 1972). The behaviour of dark-grown seedlings in response to illumination does differ in a number of respects from seedlings grown under diurnal conditions of illumination, as discussed by Schiff (1975) for example.

3.4.1—
The Proplastid

Figure 3.1A shows an electron micrograph of a proplastid in a section of a primary leaf of a 6-day-old dark-grown bean seedling. The invaginations of the inner membrane of the envelope are interpreted as representing the formation of porous sheets of membrane which are shown in section. The proplastid also contains a starch grain and scattered ribosomes while the cytoplasmic ribosomes are more prominent and are apparently mostly present as polysomes.

3.4.2—
Etioplast Formation

Between 4 and 14 days of dark growth of Phaseolus vulgaris seedlings, the primary leaf primordia, which are already present in the dry seed, show a considerable amount of growth in increasing from a fresh weight of less than 1 mg to 30–40 mg while the cell number increases by more than 10 times and plastid number increases by 18 times (Bradbeer et al., 1974c). During this time the amount of membrane within the plastid increases considerably so that etioplasts like that in Fig. 3.2A are formed. The term etioplast was used first by Kirk and Tilney-Bassett (1967) and defines a structure which is typical of dark-grown seedlings, not normally found in plants grown under diurnal conditions of light and dark. In the bean etioplast about half of the membrane is organised in a regularly-arranged network of tubules called the prolamellar body with the remainder in concentrically arranged porous lamellar sheets (thylakoids). The prolamellar body shows a para-crystalline form and a knowledge of crystallography has contributed to the interpretation of its basic structure. In one plane the tubules form a mesh of hexagons, each individual hexagon being connected to the one immediately above by tubules arising from three alternate nodes, and to the one immediately below by tubules arising from the other three nodes (Weier & Brown, 1970). In planes cutting the hexagonal plane at 90º the arrangement of the tubules is approximately rectangular. Prolamellar bodies are frequently large and complex structures with evident discontinuities, but there are no published reports which account exactly for the structure of these large pro-

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Figure 3.2
Stages of chloroplast development during the greening of the primary leaves
of 14-day-old dark-grown beans under continuous illumination of 3 mW. cm^–2 A, no illumination; B, 105 minutes illumination; C, 4 hours illumination; D, 5
hours illumination; E, 15 hours illumination. Magnification × 25,000. Key
to lettering: PB, prolamellar body; other details as in Fig. 3.1.

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lamellar bodies. Calculations based on measurements in the author's laboratory of electron micrographs of the prolamellar body structure described above show that 1 µm³ of the prolamellar body of 14-day-old dark-grown bean leaves should contain the equivalent of 44 µm² of membrane.

From measurements of electron micrographs and the dimensions of the plastids determined by light microscopy it has been possible to follow the changes in the area of the lamellar sheets and of the volume of the prolamellar bodies during etioplast development. Determination of the number of plastids in the leaf then enables quantities per plastid (Bradbeer et al., 1974c) to be expressed on a per leaf basis as shown in Fig. 3.3. In Fig. 3.3 the area of membrane in the prolamellar bodies has been calculated on the basis stated above that 1 µm³ of prolamellar body contains 44 µm² of membrane. As this factor is likely to vary during development according to the degree of contraction of the prolamellar body some inaccuracy is inevitable from this source for samples other than the 14-day-old one. In particular the slight apparent fall in the total membrane, shown in Fig. 3.3, after 14 days of dark growth may represent a contraction of the prolamellar body without any change in its membrane content. In the experiment shown in Fig. 3.3 membrane formation occurred between 6 and 14 days of growth. The porous lamellar sheets were formed first and the data are consistent with the presumed formation of the prolamellar bodies by some sort of condensation of these sheets (Weier & Brown, 1970). The area of the lamellar sheets reached a peak at 12 days after which the continued formation of the prolamellar bodies seems to have resulted in some consumption of the lamellar sheets.

During dark development of the etioplast most of the chemical components of the chloroplast are formed; for example all of the photosynthetic carbon cycle enzymes are present in the etioplast (Bradbeer et al., 1974c). Substances not yet detected in etioplasts which have received no illumination are chlorophyll, the chloroplast pigment-protein complexes and cytochrome b –559_HP . The etioplasts of beans appear to reach the peak of their development by 14 days of dark growth while retaining an ability to form chloroplasts on illumination; however 17-day-old leaves with 75% of the etioplast membrane in the prolamellar body green only feebly and 21-day-old dark-green bean leaves fail to survive when transferred to illumination. It should also be pointed out that more rapid greening occurs in leaves younger than 14 days.

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Figure 3.3
The formation of internal membrane during etioplast development in the
mesophyll of primary leaves of Phaseolus vulgaris during growth in
continuous darkness at 23°C

, total membrane;

, membrane in
porous lamellar sheets;

, membrane in prolamellar body.

3.4.3—
The Conversion of Etioplasts into Chloroplasts

The formation of chloroplasts in dark-grown leaves requires the provision of an appropriate amount and quality of illumination. Continuous illumination of 3 mW. cm^–2 by fluorescent tubes in a growth cabinet was used for the electron microscopic study of 14-day-old dark grown beans shown in Fig. 3.2. For this material fixed with glutaraldehyde-osmium tetroxide, the paracrystalline appearance of the prolamellar body is retained for about 30 minutes after the beginning of illumination, and then between 30 and 60 minutes there is a change from a regular to an irregular appearance like that shown in Fig. 3.2B (Bradbeer et al., 1974a). Transformation of the prolamellar body appears to occur much more rapidly, usually within less than 1 minute of illumination, if the leaves are fixed with permanganate but this rapid change is commonly regarded as an artefact in structural terms. On the other hand, it may result from an early photochemical reaction in the etioplast which renders the paracrystaline nature of the prolamellar body less stable (Berry & Smith, 1971). Subsequently the volume of the reacted prolamellar body falls and the area of the thylakoid sheets increases as shown in Fig. 3.2C & D for 4 and 5 hours of illumination

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Figure 3.4
The effects of the transfer of 14-day-old dark-grown beans to continuous
illumination of 3 mW. cm^–2 on the prolamellar body volume and the thylakoid area
of the plastids of the primary leaves.

, prolamellar body volume;

, thylakoid area.

respectively and in Fig. 3.4. Figure 3.4 shows that the fall in the prolamellar body volume and the increase in the area of the thylakoid sheets approximately correspond with each other with the actual increase of 21.5 µm² thylakoid/plastid (43 µm² membrane) accounting for the loss of 0.99 µm³ of prolamellar body per plastid (43.6 µm³ membrane). After 10 hours of illumination, appression of the thylakoids becomes more obvious, the formation of grana occurs, and the further increase in thylakoid membrane may be presumed to have resulted from de novo membrane synthesis. Figure 3.2E shows the stage of development after 20 hours of illumination and Fig. 3.1B that after 48 hours of illumination. During the course of bean chloroplast development under these conditions, de novo membrane synthesis results in an 8-fold increase in thylakoid area.

The illumination of 14-day-old dark-grown beans results in a synchronous development of the etioplasts in the primary leaf mesophyll cells. There is no division of these cells, less than 10% of the plastids divide, and the rather amoeboid etioplasts double in diameter to give typical discoid chloroplasts. The small amount of chloroplast division does not seem to be typical of developing leaves in which replication of chloroplasts is usual. Greening leaves show a lag before the onset of photosynthesis. The length of this lag depends upon the nature of the plant material, the conditions and duration of dark growth, the

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conditions for greening, the intensity of the illumination used to assay photosynthesis and the sensitivity and nature of the method of assay. Consequently, published reports on the duration of this lag show a range of values; however, all agree that at least one hour of illumination, and usually more, is required before photosynthetic CO₂ -fixation can be detected. For the bean leaves studied in the author's laboratory, the lag has been at least 5 hours, after which the photosynthesis of the developing chloroplasts becomes increasingly important as the source of the energy and the carbon requirements, of their own development (Bradbeer, 1976).

The effects of different light treatments on the form of the resultant chloroplast have been considered in an earlier section (3.2.4.3). There have been a number of studies in which light of different wavelengths has been used for the illumination of dark-grown seedlings in an attempt to determine an action spectrum of the light dependent reactions involved in chloroplast development. Henningsen (1967) obtained a sharp peak at 450 nm in the action spectrum for a stage termed vesicle dispersal in the development of bean plastids. Unfortunately this developmental stage may be another artefact of permanganate fixation as thylakoid extrusion does not involve such a stage when seen in glutaraldehydeosmic acid fixed material and studies with this latter fixative have not so far observed any corresponding action spectrum. However Henningsen's experiments have not been exactly replicated with the latter fixative and his data may well indicate a so far undefined photoresponse. Most studies of the effects of light quality on chloroplast development have used rather wide bands of wave-lengths and have concentrated on the role of phytochrome in chloroplast development. It has been shown in a number of laboratories that phytochrome has an important role in chloroplast development, and a number of aspects of chloroplast development are promoted by short treatments with red light and reversed by short exposure to far-red. In bean, the presumably-active form of phytochrome, Pfr (see chapter 12) promotes plastid expansion, plastid division, the formation of plastid membrane and the synthesis of chloroplast proteins (Bradbeer, 1971; Bradbeer et al., 1974b). These reactions may be regarded as 'slow' reactions in that they require several hours to become evident. In addition to phytochrome, 'slow' reactions may possibly also be effected by a red-light-absorbing photoreceptor other than phytochrome, and by a blue-light-absorbing photoreceptor. Most of the 'rapid' changes in etioplasts, which occur within three hours of illumination, do not seem to involve phytochrome; such changes as prolamellar body transformation and loss are sensitive to a wide range of wavelengths whilst thylakoid extrusion seems to depend on a red-absorbing photoreceptor other than phytochrome. Only two 'rapid' effects of phytochrome on etioplast fine structure have so far been defined; they are the Pfr-promoted crystallization of the prolamellar body in mustard cotyledons (Kasemir et al., 1975) and the Pfr-inhibited recondensation of the prolamellar body in bean primary leaves (Bradbeer & Montes, 1976). Although Wellburn and Wellburn (1973) have implicated Pfr in 'rapid' changes in isolated and

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in situ etioplasts their method of analysis does not permit them to define these changes with any precision and their conclusions seem to require reconsideration.

3.4.4—
The Formation of Chloroplast Components in Greening Leaves

The transfer of dark grown plants to continuous illumination results in substantial increases in the amounts of most of the etioplast constituents which are also found in the chloroplasts (see for example Bradbeer et al., 1969; Gregory & Bradbeer, 1973). A few components, namely chlorophyll, the chloroplast pigment-protein complexes and cytochrome b– 559^HP , which cannot be detected in dark-grown etioplasts, appear as a result of illumination.

The photosynthetic enzymes have almost exclusively been studied by determination of enzyme activity and thus any change in activity may result from either a change in the amount of enzyme protein or a change in the activity of the enzyme protein. Since, during the greening process, there is considerable protein synthesis it was considered that the simultaneous increases in enzyme activity probably resulted from protein synthesis. Subsequent investigations have established that enzyme activation is also responsible for a substantial part of the increase in the activities of certain enzymes in greening leaves.

Ribulosebisphosphate carboxylase is the most abundant protein in the chloroplast and probably consists of 8 large subunits (molecular weight about 5.2 × 10⁴ ) and 8 small subunits (molecular weight about 1.3 × 10⁴ ) which give a molecule of about 5.2 × 10⁵ in molecular weight. By the use of two-dimensional polyacrylamide gel electrophoresis, Arron and Bradbeer (1975) found that the commencement of the synthesis of both subunits in bean leaves coincided with the beginning of illumination but that there was a lag before enzyme activity increased. Smith et al., (1974) reached a similar conclusion from experiments with barley in which they measured newly-synthesized ribulosebisphosphate carboxylase by labelling it during its biosynthesis and trapping the labelled substance with a specific antibody. In both bean and barley, synthesis of the enzyme occurs early in greening with a corresponding increase in enzyme activity, while late in greening after the enzyme synthesis has ceased considerable activation occurs. The mechanism of the activation is not known although the effect of illumination may be mediated by a small (molecular weight 5 × 10³ ) constituent (Wildner et al., 1972).

In contrast, rather more is known about the activation of phosphoribulokinase and triosephosphate dehydrogenase, where a pretreatment of the extracted enzyme with 6 mM ATP prior to assay brings about activation, (Wara Aswapati et al., 1977). Activation may also be brought about by NADPH and sulphydryl reagents. On illumination of etiolated leaves, the activity of each of these enzymes shows a lag of several hours before it increases; however, preincubation of these extracts with ATP shows that the increases in activity seem to commence from the beginning of illumination (Fig. 3.5). It is concluded that synthesis of

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Figure 3.5
The development of phosphoribulokinase activity in primary
leaves of 14-day-old dark-grown beans on transfer to
continuous illumination of 3 mW.cm^–2 .

, assayed
after pretreatment of the extract with 6 mM ATP;

,
no pretreatment;

, leaves illuminated;

,
leaves left in dark. After Wara-Aswapati (1973).

these two enzymes commences with the beginning of illumination but that activity does not develop until photophosphorylation commences, thus accounting for the lag in Fig. 3.5 for the extracts which were not activated prior to assay. Thus, for the phosphoribulokinase in greening bean leaves Fig. 3.5 shows that there was a 10-fold increase in enzyme protein and a 9-fold increase in the enzyme activity of the protein, thus accounting for a 90-fold increase in total enzyme activity.

There is the further example of ATPase, which is a polypeptide complex in which illumination brings about an increase in enzyme activity with comparatively little new synthesis of the protein being apparent (Gregory & Bradbeer, 1975). In this case the effect of illumination seems to be more direct and is probably necessary for the onset of photophosphorylation rather than a result of it.

3.5—
Summary

The chloroplasts of higher plants are normally lens-shaped organelles with a diameter of about 5 µm and a maximum thickness of about 1 µm. They are

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bounded by an envelope consisting of two membranes the outer of which is freely permeable to solutes while the inner shows very specific permeability. Within the inner membrane is a complex system of thylakoids (flattened sacs of membrane) which are closely associated in a number of stacks (grana), the thylakoid system itself being embedded in a matrix (stroma). Considerable variation in chloroplast structure is found in the algae. In plants which possess the C-4 pathway of photosynthetic CO₂ fixation, dimorphism of chloroplasts is usual with agranal starch-containing chloroplasts occurring in the bundle sheath, which is a single layer of cells immediately surrounding the vascular bundles, while in the next outermost layer of cells (mesophyll sheath) are found most of the remaining chloroplasts, which possess grana and do not normally accumulate starch. Chloroplast structure may also be modified by environmental conditions or mutation.

Techniques for the isolation of chloroplasts for physiological and biochemical studies have been developed and the properties of the main types of chloroplast preparation are summarized in tabular form.

Proplastids occur in meristematic cells and are small with little membrane within the envelope. The direct conversion of proplastids to chloroplasts has been studied in plants grown under diurnal conditions of illumination although in most developmental studies seedlings have been grown in complete darkness at first so that the proplastids have developed into etioplasts. Etioplasts possess a substantial amount of internal membrane, some of which occurs as porous thylakoid sheets with the remainder condensed to give one or more regular lattices of tubules, the prolamellar bodies. On transfer of dark-grown seedlings to illumination the prolamellar body loses its regular structure and becomes converted into thylakoids. Subsequently de novo thylakoid synthesis occurs together with the formation of grana. Most of the chemical constituents of chloroplasts are already present in etioplasts. On illumination the missing components are synthesized, some enzymes are subjected to activation, most components undergo further de novo synthesis and photosynthetic activity develops after a lag. Some preliminary investigations have also been made of the effects of quality and intensity of irradiation on chloroplast development.

Chapter 4—
Chloroplasts—Structure and Photosynthesis

4.1—
Introduction

Chapter 3 dealt with the ultrastructure of the higher plant chloroplast and the differentiation of the chloroplast's internal membrane system into grana and stroma thylakoids. It outlined the structure of the etioplast and examined the sequence of structural and biochemical changes during the maturation of an etioplast into a chloroplast. This chapter examines the thylakoid membrane at the molecular level in relation to its prime function of converting light energy into chemical energy in the form of NADPH and ATP. The formation of a functional thylakoid from the prolamellar body membranes of the etioplast during the greening process will also be considered at the molecular level. Finally, we briefly examine the relationship of the chloroplast to cytoplasm in terms of energy metabolism.

4.2—
Thylakoid Structure

4.2.1—
Thin Sectioning and Heavy Metal Shadowing

When thin sections of fixed and stained leaf material are examined in the electron microscope, the chloroplast thylakoid membrane appears as a single electron dense line or as a tripartite structure (two dense lines with an intermediate light line) characteristic of the unit membrane (chapter 2). In high resolution electron micrographs of thin sections, fixed in permanganate, the thylakoid membranes give the appearance of containing globular subunits, 9 nm in diameter. It is possible, however, that these are an artefact since it is difficult to understand how several layers of overlapping 9 nm particles can be clearly resolved in a section thicker than 40 nm (Branton, 1968).

Some of the earliest electron micrographs of thylakoid membranes were obtained simply by drying isolated chloroplast fragments of Spirogyra and Mougeotia on an electron microscope grid and shadowing the preparations with chromium (Steinmann, 1952). The membranes showed a granular or particulate structure. Similar observations were made subsequently with spinach thylakoid membranes (Park & Pon, 1961). The particles usually existed in a random array but sometimes they were seen in highly ordered arrays. Their dimensions were 15 × 18 × 10 nm and they appeared to contain subunits. Park (1962) suggested

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that the particle, which he termed quantasome, might be the morphological expression of the so-called photosynthetic unit, defined as the minimum number of chlorophyll molecules needed to fix one molecule of carbon dioxide per intense flash of light (Emerson & Arnold, 1932). The quantasomes were seen on the inner surface of the thylakoid membrane, suggesting that there was some distortion of the thylakoid membrane during drying.

4.2.2—
Freeze-Etching

The freeze-etch technique is a different approach to the study of the substructure of the thylakoid membrane, because chemical fixatives are not used. Leaf material or isolated chloroplasts are frozen and fractured (Moor et al., 1961). The fracture plane in the frozen sample occurs along hydrophobic regions within the thylakoid membrane exposing complementary faces. Platinum-carbon replicas of the fracture planes are made and examined in the electron microscope.

Two major sizes of particles are seen when mature plant chloroplasts are freeze-fractured; large particles of average diameter, 17.5 nm and small particles of average diameter, 11 nm (Fig. 4.1a). The small and large particles occur on different fracture faces (Fig. 4.1b). Fracture face C, which is towards the stroma

Figure 4.1a
Freeze-fracture electron micrograph of a spinach grana thylakoid, showing
the large and small particle fracture faces, B and C, and the surfaces, A' and D.
(By courtesy of Dr. D.J. Goodchild.)

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of the chloroplast contains tightly packed small particles, and the complementary fracture face B contains the large particles at about half the density per µm² . The large particles are only observed within grana stacks, where the adjacent thylakoid membranes are in close contact, while the small particles are observed in grana and stroma thylakoids. In the unstacked stroma thylakoids the complementary fracture face (designated B_u ) is relatively smooth and shows only a few small particles (Park & Sane, 1971).

Figure 4.1b
Model of the thylakoid membrane, showing grana and stroma regions.
The particles on the A' surface represent the coupling factor (CF₁ ).
(By courtesy of Dr. D. J. Goodchild.)

The exterior thylakoid surface, A', and the interior surface, D, are observable by the deep etch technique, in which the ice covering the surfaces is sublimed off before the replica is made. In well-washed thylakoids the A' surface shows little surface relief, but is covered with proteins in unwashed lamellae. The interior surface is covered with particles about 18.5 × 15.5 nm in size which show slight surface relief. These particles on the D surface contain 3 or 4 subunits and they may correspond to the quantasomes as seen by metal shadowing.

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The freeze-etch data suggest that the particles within the thylakoid membrane are arranged somewhat asymmetrically. The larger particles are located towards the inner surface and partly protrude into the intrathylakoid space, and the smaller particles are near the outer surface. The model of the thylakoid membrane shown in Fig. 4.2 appears to be consistent with the freeze-etching results and also with X-ray diffraction studies of thylakoid membranes (Sadler et al., 1973). The membrane is viewed as a lipid bilayer in which the particles are embedded, in accordance with the fluid mosaic model for biological membranes (Singer & Nicolson, 1972; see chapter 2).

Figure 4.2
Structural model of the thylakoid membrane showing an asymmetric
distribution of large and small particles. The two membranes belong to
one thylakoid. The lipid molecules are indicated as a bilayer, the circles
representing the polar head groups. Lipid asymmetry is indicated by the
shading of the head groups. The model appears to be consistent with freeze
-etch electron microscopy and X-ray diffraction of thylakoid membranes.
(From Sadler et al., 1973.)

4.2.3—
Negative Staining

Two sizes of particles are observed on the surface of unwashed thylakoids by negative staining with phosphotungstic acid. The larger particles (11 nm in diameter) are removed by water washing the thylakoids and correspond to the CO₂ -fixing enzyme, ribulose bisphosphate carboxylase. The smaller particles (9 nm) are removed by washing with the chelating agent, ethylenediamine tetraacetate, and are identical with the coupling factor (CF₁ ) which catalyses the terminal steps in the energy coupling process of ATP formation. These surface particles are not related to the membrane particles seen by freeze-etching (Park & Sane, 1971).

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4.3—
Thylakoid Composition

4.3.1—
Lipids

The thylakoid membrane consists of 50% protein and 50% lipid. The feature of its lipid composition is the high percentage of the neutral galactolipids, digalactosyl diglyceride (27%) and monogalactosyl diglyceride (13%), compared with the phospholipids (9%) (Table 4.1). Chlorophyll a plus chlorophyll b account for 21% of the lipids. The thylakoid membrane also contains an anionic sulpholipid. The other unique feature of the thylakoid lipids is their high content of the polyunsaturated fatty acid, linolenic acid (18:3) (Table 4.2).

Table 4.1. Lipid composition of spinach thylakoid membranes (adapted from Lichtenthaler & Park, 1963).
	% Weight of total lipid	Number of lipid moles per 400 chlorophyll moles
Monogalactosyl diglyceride	26.8	602
Digalactosyl diglyceride	13.4	198
Sulpholipid	4.1	42
Phospholipid*	9.1	201*
Chlorophyll a	15.0	290
Chlorophyll b	5.8	110
Carotenoids**	2.8	82**
Quinone compounds	3.3	80
Sterols	2.2
Unidentified lipids	17.0
* Phosphatidyl glycerol (91); phosphatidylcholine (73); phosphatidyl inositol (24); phosphatidyl ethanolamine (10); phosphatidic acid (3). ** b -carotene (24); lutein (38); violaxanthin (10); neoxanthin (10).

Table 4.2. Fatty acids of spinach thylakoid lipids (percent composition) (from Allen & Good, 1971).
Fatty acid	14:0	16:0	D₃ –trans 16:1	16:2	16:3	18:0	18:1	18:2	18:3
Monogalactosyl diglyceride	–	–	–	–	25	–	–	2	72
Digalactosyl diglyceride	–	3	–	–	5	–	2	2	87
Sulpholipid	–	39	–	–	–	–	––	6	52
Phosphatidyl glycerol	1	11	32	–	2	–	2	4	47
Phosphatidyl choline	–	12	–	–	4	–	9	16	58
Phosphatidyl inositol	4	34	6	–	3	2	7	15	27

― 90 ―

In spinach thylakoids, trans-D³ -hexadecenoic acid is a major constituent of phosphatidylglycerol. Lipophilic quinones, of which PQA₉ (2, 3 dimethyl-p -benzoquinine with a 45-carbon side chain at the 5-position) is the major constituent, account for 3 % of the thylakoid lipids.

4.3.2—
Chlorophyll-Proteins

The proteins of the thylakoid membrane include the carriers of the photosynthetic electron transport chain, but these account for only a relatively small percentage. When a thylakoid preparation, from which the coupling factor, lipids and pigments have been removed, is solubilized by the anionic detergent, sodium dodecyl sulphate (SDS), and the extract examined by SDS-polyacrylamide gel electrophoresis, at least 20 polypeptides are obtained (Anderson & Levine, 1974). These range in size from 10,000 to 100,000 daltons. A major fraction of the proteins appear in the region of 25,000 daltons. If the thylakoids are extracted with SDS without prior removal of pigments, two chlorophyllprotein complexes (termed complex I and complex II) are separated by SDS polyacrylamide gel electrophoresis (Thornber, 1975). Complex I which is derived from photosystem 1 (see section 4.4) has a chlorophyll a /chlorophyll b ratio of 12:1 and accounts for about 20% of the thylakoid membrane protein. It has an apparent molecular weight of 110,000 daltons and contains 14 moles chlorophyll a per mole of protein. Complex 11, which accounts for 40–50% of the thylakoid membrane proteins has an apparent molecular weight of 32,000 daltons and contains one mole chlorophyll a and one mole of chlorophyll b per mole of protein.

4.3.3—
Electron Carrier Proteins

The thylakoid membrane contains four spectroscopically distinguishable cytochromes: cytochrome f (a c -type) with an a -band at 554 nm and a midpoint potential of + 0.36 at pH 7, cytochrome b₆ (alternate name b– 563) with an a -band at 563 nm and a mid-point potential below - 0.1V, cytochrome b– 559_HP (high potential form) with an a -band at 559 nm and a mid-point potential (+ 0.37V) close to that of cytochrome f and cytochrome b – 559^LP (low potential form) with an a -band at 559 nm and a mid-point potential of 0.06V (Bendall et al., 1971). The thylakoid cytochromes are not extracted by aqueous solvents. Cytochrome ¦ is extracted into ammoniacal ethanol (Bendall et al., 1971), but the b cytochromes are tightly associated with the thylakoid membranes and are released on extraction of the membranes with detergent (Boardman, 1975).

Plastocyanin is a copper-containing protein with a mid-point potential of + 0.37V at pH 7. It is extracted by sonication of the thylakoid membranes or by detergent treatment. Its molecular weight is 10,500 with 1 gm atom of Cu per mole. The oxidized form of plastocyanin is intensely blue with a main absorption band at 597 nm.

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Figure 4.3
Structure of iron-sulphur group in ferredoxin
(from Hall et al., 1972).

The ferredoxins are stable iron-sulphur proteins in which Fe is coordinated to the sulphur atoms of cysteine and inorganic sulphur (Fig. 4.3). Chloroplasts contain a soluble ferredoxin (E₀ ' = –0.43V at pH 7.5), which is readily released in aqueous media, and one or more membrane-bound ferredoxins. Soluble ferredoxin has a molecular weight of 12,000 and contains 2 gm atoms of Fe and 2 gm atoms of inorganic sulphur per mole. Bound ferredoxin has also been extracted and purified (Malkin et al., 1974). It has a molecular weight of 8,000 and 4 gm atoms of Fe and 4 gm atoms of inorganic sulphur per mole. Its mid-point potential is 0.1V more negative than that of soluble ferredoxin.

The thylakoid membrane contains a flavoprotein, ferred oxin-NADP⁺ reductase, which is extractable with detergents. It has a molecular weight of 40,000.

4.4—
Photosynthesis

In the process of photosynthesis in green plants, light energy is absorbed by the pigments of the chloroplast and converted into chemical free energy in the form of ATP and NADPH, which are then used for the reduction of carbon dioxide and the synthesis of plant materials. The major organic products of photosynthesis are carbohydrate and the overall equation of photosynthesis is:

The oxygen which is evolved is derived from water. Carbon dioxide is converted to carbohydrate via the Calvin cycle of reactions (Calvin & Bassham, 1962). In the carbon reduction cycle, one molecule of ribulose bisphosphate (RBP or RuDP) reacts with CO₂ to form two molecules of 3-phosphoglyceric acid (PGA), which are converted into phosphoglyceraldehyde (triose phosphate) in a reaction which needs two molecules of NADPH and two molecules of ATP. The regeneration of the CO₂ acceptor (RBP) involves a complex series of reactions and requires one molecule of ATP. The overall requirement of the Calvin cycle is 3 molecules of ATP and 2 molecules of NADPH for each molecule of CO₂ reduced to carbohydrate. In the C₄ -dicarboxylic acid pathway of photosynthesis, in which CO₂ reacts initially with phosphopyruvate, 5 molecules of ATP and 2 molecules of NADPH are needed for each molecule of CO₂ reduced (Hatch & Slack, 1970).

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The light reactions of photosynthesis and the formation of NADPH and ATP are performed by the thylakoids, and the carbon reduction cycle occurs in the stroma or soluble phase of the chloroplast.

4.4.1—
The Photosynthetic Unit and Energy Transfer

The light absorbing molecules, i.e. the chlorophylls and carotenoids, are organized into units in the thylakoid membrane. Quanta absorbed by a large number of pigment molecules are transferred by an efficient resonance mechanism to a special molecule of chlorophyll a , called the trap or reaction centre chlorophyll, where the primary conversion of light energy into chemical free energy takes place. This is the concept of the photosynthetic unit which was first proposed to account for the observation that the maximum yield of CO₂ fixed, or O₂ evolved, per single flash of intense light was one mole per 2,500 moles of chlorophyll. The absorption band of the reaction centre chlorophyll a is at a longer wavelength than the absorption bands of the light-harvesting pigments to ensure efficient trapping of the energy at the reaction centre.

The reaction centre chlorophyll is in close association with an electron acceptor molecule (A) and an electron donor molecule (D) in the thylakoid membrane. On excitation of the reaction centre chlorophyll (Chl*) by transfer of energy from the 'antenna' pigments, an electron is donated to A giving A^– and leaving the chlorophyll molecule deficient in an electron (eq. 4.1). The positively charged chlorophyll then receives an electron from the donor D, and is restored to its ground state energy level (eq. 4.2). The net result is that the energy of a photon is used to transfer an electron from D to A, and thus the primary photochemical event of photosynthesis is an oxidation-reduction process.

4.4.2—
Two Photosystems and the Z-Scheme of Electron Transport

Investigations in the 1950's and early 1960's established that the thylakoid membrane contains two types of photosynthetic units, designated photosystem 1 (PS–1)and photosystem 2 (PS–2), which cooperate in a sequential manner to transfer electrons from water to NADP⁺ (Boardman, 1968). Figure 4.4 depicts the electron transport pathway known as the Z-scheme first put forward by Hill and Bendall (1960). Both photosystems contain chlorophyll a , chlorophyll b and the four carotenoids, but in different proportions. Each unit of PS–1 or PS–2 contains 200 light-harvesting chlorophylls and one reaction centre.

Quanta of light absorbed by PS–2 are transferred to a form of chlorophyll absorbing at 682 nm and termed chl a– 682. Excitation of chl a– 682 catalyses

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the transfer of an electron from Y to Q, giving a strong oxidant, Y⁺ , and a weak reductant, Q^– . Oxidation of water and the release of a molecule of O₂ requires the sequential absorption of four quanta and the accumulation of four oxidizing equivalents (Cheniae, 1970). The mechanism of water oxidation is unknown, although it is established that manganese, probably in the form of a manganeseprotein complex, and Cl^– are required (Boardman, 1975). The primary acceptor of PS–2, (Q), has a redox potential around zero volts, and is incapable of reducing NADP⁺ , without an imput of energy into PS–1. Quanta absorbed by PS–1 are transferred to P–700, a form of chlorophyll a absorbing at 700 nm. On excitation, P–700 donates an electron to an acceptor Z, resulting in the formation of P–700⁺ , a weak oxidant with a mid-point potential of + 0.43V, and Z^– , a strong reductant with a redox potential in the vicinity of –0.6V. P–700⁺ interacts with the weak reductant, Q^– , generated by PS–2 via an electron transport chain, which includes plastoquinone A, cytochrome ¦ and plastocyanin. The reduction of NADP⁺ by Z^– is mediated by soluble ferredoxin and ferredoxin-NADP reductase. It seems possible that the primary acceptor of PS–1, Z, is identical to bound ferredoxin, since the latter is photoreduced at very low temperatures (25°K) (Bearden & Malkin, 1974).

Cytochrome b₆ appears to function on a cyclic electron transport pathway around PS–1 and it may play a role in cyclic phosphorylation. There is conflicting evidence concerning the role of cytochrome b– 559_HP (Boardman, 1975). This cytochrome is oxidized by PS–2 at liquid nitrogen temperature and under certain conditions at room temperature, but at high light intensity it can also be reduced by PS–2. The function of cytochrome b– 559_LP is unknown at present.

Much evidence for the scheme of photosynthetic electron transport depicted in Fig. 4.4 has come from difference spectroscopy of chloroplasts. In this method, the change in the absorbance of various chloroplast components is measured on illumination of the chloroplasts with monochromatic light of various wavelengths. By following the change in the spectrum of individual components e.g. cytochrome ¦ or P–700, the role of these components can be deduced. For example, if chloroplasts are illuminated with far-red light of wavelength 720 nm, which is only absorbed by PS–1, there is a decrease in the absorbance of the chloroplasts in the region of 554 nm, due to the oxidation of cytochrome ¦ . If the chloroplasts are then illuminated with 650 nm light, absorbed by PS–2 (as well as by PS–1)there is an increase in absorbance due to the reduction of cytochrome ¦ . The oxidation of P–700 is followed by a decrease in absorption at 700 nm.

Electron transport from water to NADP⁺ in isolated chloroplasts may be intercepted by the addition of artificial electron acceptors (oxidants), which accept electrons from Z^– in PS–1 or from PQ in PS–2. For example, methyl viologen accepts electrons at PS–1 while p -phenylenediamine intercepts the chain at PQ. Ferricyanide or dichlorophenolindophenol can interact at either PS–1 or PS–2, depending on the experimental conditions (Trebst, 1974).

― 94 ―

Figure 4.4
Z-scheme for photosynthetic electron transport and photophosphorylation. The
number beside a component indicates the number of molecules of that component per
photosynthetic unit of 400 chlorophyll molecules. Two sites of ATP formation are located
on the pathway between water and PS–1 (see text); one between water and plastoquinone
(PQ) and the other between PQ and PS–1. A scale of redox potentials is shown on the left.

Photosynthetic electron transport is readily monitored by illuminating isolated chloroplasts in the presence of an electron acceptor, and measuring either the oxygen evolved or the amount of acceptor reduced (Hall & Rao, 1972). This reaction is known as the Hill reaction (Hill, 1939).

The herbicides 3(3,4-dichlorophenyl) 1,1-dimethylurea (DCMU) and 3(p -chlorophenyl)-1,-1-dimethylurea (CMU) inhibit electron transport between

― 95 ―

Q and PQ. Photoreduction of NADP⁺ can be restored in the inhibited chloroplasts by the addition of an artificial electron donor such as reduced 2,6-dichlorophenolindophenol. Electrons from the artificial donor enter the electron transport chain between the light reactions, and photoreduction of NADP⁺ is then driven by PS–1.

PS–1 is known as the far-red system because its absorption spectrum extends to longer wavelengths than that of PS– 2. At wavelengths beyond 700 nm, PS– 1 receives a high fraction of the quanta absorbed by chloroplasts. Chloroplasts contain one molecule of cytochrome ¦ and one molecule of P–700 per 430 chlorophyll molecules, from which it is concluded that the photosynthetic unit contains about 400 chlorophyll molecules. As shown in Fig. 4.4 the chlorophyll molecules appear to be distributed about equally between the two photosystems.

4.4.3—
Photosynthetic Phosphorlation

Electron flow from water to NADP⁺ is coupled to the formation of ATP. Until recently it was uncertain whether there is one or two energy conserving sites, where ATP is formed. This arose because of the conflicting experimental determinations of the amount of ATP formed during the transfer of 2 electrons from water to NADP (the P/e₂ ratio). Earlier measurements gave a P/e₂ ratio of one (Arnon et al., 1958), suggesting one energy conserving site, but more recent determinations indicate P/e₂ ratios between 1.2 and 1.8 (Trebst, 1974). One energy conserving site is associated with the reoxidation of plastoquinone by cytochrome f, and the second (not shown on Fig. 4.4) is associated with the electron transfer from water to plastoquinone. However, the two sites do not necessarily yield two ATP per 2 electrons transferred from water to NADP⁺ , as indicated by the experimental P/e₂ ratios. A P/e₂ ratio of at least 1.5 is needed to provide enough ATP to drive the Calvin cycle, and more ATP is required for the C₄ -pathway.

Another type of phosphorylation has been observed with isolated chloroplasts. Known as cyclic phosphorylation, it requires an exogenous cofactor such as phenazine methosulphate, pyocyanin or ferredoxin. Unlike non-cyclic phosphorylation, cyclic phosphorylation is not accompanied by any net change in oxidation or reduction and it is not inhibited by DCMU. The process is driven by light absorbed by PS–1. Chloroplasts exhibit very low rates of cyclic phosphorylation in the absence of an exogenous electron carrier. It is uncertain, therefore, whether cyclic phosphorylation in vivo contributes a significant amount of ATP. Recent work suggests that oxygen may act as an alternative electron acceptor at PS–1 when NADP is fully reduced, and provide extra ATP during electron flow from water to O₂ (Heber, 1975). The relative amounts of ATP formed by noncyclic electron flow to NADP⁺ or O₂ could be regulated by the demands of the cell for NADPH and ATP.

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4.4.4—
Phosphorylation and Thylakoid Structure

Three main hypotheses have been proposed for the mechanism of ATP formation in mitochondria and chloroplasts (Slater, 1971), but here we will briefly consider only one of these, the Mitchell chemiosmotic hypothesis (Mitchell, 1966; see also chapter 5). The fundamental concept of the chemiosmotic hypothesis is that of vectorial flow of protons across the mitochondrial inner membrane or the thylakoid membrane. Mitchell proposed that the electron transport chains of mitochondria and chloroplasts consist of alternate electron and hydrogen carriers, organized in a vectorial fashion across the membrane. Transport of electrons through the chain of carriers results in the net transfer of protons across the membrane. For the thylakoid membrane, protons are transported from the outside (stroma or matrix) to the inside intrathylakoid space. This is depicted in Fig. 4.5 (Trebst, 1974). In this scheme, reduction of plastoquinone by Q

Figure 4.5
Photosynthetic electron flow, showing vectorial arrangement of carriers and the
movement of protons from outside to inside of the thylakoid. (From Trebst, 1974.)

occurs on the outside of the thylakoid and oxidation by cytochrome ¦ on the inside, resulting in the transfer of 2 protons for each 2 electrons transported from water to NADP⁺ . Water oxidation is considered to take place on the inside to produce 2 protons per 1/2O₂ . According to the Mitchell hypothesis, the protons move back through the membrane through special channels which contain the coupling factor or ATPase, and result in formation of ATP from ADP and inorganic phosphate. As already discussed, part of the ATPase (CF₁ ) is on the outside of the thylakoid membrane, and part (CF₀ ) is embedded in the membrane. It is not yet established whether the proton gradient across the thylakoid membrane is an essential intermediate in the process of ATP formation during photosynthetic electron flow. However, with thylakoid membranes it has been shown

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that ATP formation can be driven by a pH gradient created artificially (Jagendorf & Uribe, 1966). Chloroplasts are suspended in a buffer of low pH (pH < 5) and then transferred to a buffer of pH 8, containing ADP and inorganic phosphate.

According to the Mitchell hypothesis, the energy-conserving sites are identified with the proton-releasing sites on the inside of the thylakoid. Substances such as ammonia which uncouple ATP formation from electron transport in the thylakoid membrane (termed uncouplers) are considered to destroy the proton gradient by transporting the protons across the membrane.

4.5—
Subchloroplast Fragments and the Fractionation of the Photosystems

4.5.1—
Digitonin Method

Methods of isolation of chloroplasts were outlined in chapter 3. In this chapter, we will consider the fragmentation of the thylakoid membrane and the isolation of subchloroplast fragments enriched in PS–1 and PS–2 respectively. The thylakoid membrane is fragmented by a number of treatments including sonication, incubation with detergents, or mechanical shearing on passage through the valve of a French pressure cell (Boardman, 1970). Here, we will outline the digitonin method for obtaining subchloroplast fragments (Boardman, 1971).

Chloroplasts are isolated in a sucrose-phosphate buffer medium and resuspended in 50 mM phosphate buffer + 10 mM KC1. Digitonin., a nonionic detergent is added to a final concentration of 0.5% and the mixture incubated for 30 minutes at 0°C. The resulting subchloroplast fragments are sedimented by differential centrifugation. The first centrifugation is at 1,000 g for 10 minutes, followed by successive centrifugations at 10,000 g for 30 minutes, 50,000 g for 30 minutes and 144,000 g for 60 minutes. The larger fragments which sediment at 1,000 g (D–1) and 10,000 g (D–10) have a lower chl a /chl b ratio than chloroplasts, whereas the smaller fragments (D–50 and D–144) have much higher ratios (Table 4.3). The D–10 fractions accounts for about one-half of the chlorophyll of the chloroplast, while approximately one-third of the chlorophyll is divided about equally between D–50, D–144 and the 144,000 g supernatant.

The smaller fragments (D–144) are highly enriched in PS–1; for example, they have a chl/P–700 ratio of 205, compared with 440 for chloroplast, a low level of manganese and they lack cytochrome b– 559_HP . They contain cytochromes ¦ , b₆ and b– 559_LP , and show the photochemical reactions of PS–1, including cyclic phosphorylation, but little or no PS–2 activity. The larger fragments (D–10) are enriched in cytochrome b –559_HP and manganese and they have less P–700 and cytochrome f on a chlorophyll basis than do chloroplasts. Thus the small fragments have the photochemical properties of PS–1,

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Table 4.3. Distribution of chlorophyll in subchloroplast fragments (from Boardman, 1971).
Fraction	Chla + Chlb (%)	Chla /Chlb (Ratio)
Chloroplasts	100	2.83
D-1	19.0	2.36
D-10	46.2	2.27
D-50	12.3	4.40
D-144	111.7	5.34
Supernatant	10.8	3.76

and the large fragments are enriched in PS–2. The non-ionic detergent Triton X–100, produces a more complete fractionation of the chloroplasts than does digitonin, but it has the disadvantage that the resulting PS–2 subchloroplast fragments are unable to evolve oxygen.

4.5.2—
Mechanical Disruption of Chloroplasts

Mechanical treatment of chloroplasts by passage through the French Press or by brief sonication disintegrates the chloroplasts into grana thylakoids and small vesicular structures derived from the stroma thylakoids (Park & Sane, 1971). The grana thylakoids are separated from the smaller vesicles by differential centrifugation. The composition and photochemical properties of the vesicles are similar to those of the D–144 subchloroplast fragments, i.e. they are highly enriched in PS–1. This led to the conclusion that stroma thylakoids contain only PS–1, whereas grana thylakoids contain both PS–1 and PS–2 (Park & Sane, 1971), gut an alternate explanation is that the PS–1 containing vesicles are derived from stroma thylakoids.

When chloroplasts are incubated with digitonin, PS–1 containing vesicles are first released from the stroma, but digitonin (and Triton X–100) also releases PS–1 fragments from grana thylakoids.

4.6—
Thylakoid Structure in Relation to the Photosystems

Attempts have been made to isolate individual quantasome particles, active in electron transport from water to NADP⁺ or ferricyanide. However, disruption of the thylakoid membrane into fragments comparable in size to the quantasome results in the complete loss of the ability to photo-oxidize water, although the small particles retain PS–1 activity, i.e. the small particles are capable of photoreducing NADP⁺ or methyl viologen with reduced indophenol dye as the electron donor. Biochemical evidence is lacking for the existence in the thylakoid mem-

― 99 ―

brane of a quantasome particle (containing one reaction centre of PS–1 and one of PS–2 and the associated light harvesting pigments and electron transfer components) which functions as a discrete unit independent of neighbouring reaction centres and electron transport chains. On the contrary, there is evidence for interaction between reaction centres and between electron transfer chains (Siggel et al., 1972; Boardman et al., 1974).

Studies with chlorophyll-deficient mutants of higher plants and with plants grown under different light intensities do not support the view that PS–2 is confined to grana thylakoids in the higher plant chloroplast (Boardman et al., 1974). Growth at low light results in greater development of grana, but the quantum efficiency for CO₂ fixation by leaves or for the reduction of 2:6 dichlorophenolindophernol by isolated chloroplasts is the same, irrespective of the light intensity during growth. If PS–2 were confined to grana, then the plants grown at high light intensity should be less efficient than those grown at low intensity. Some chlorophyll-deficient mutants show poor development of grana, but they exhibit good PS–2 activity. Studies on chloroplast development also support the conclusion that grana are not essential for PS–2 activity (section 4.7).

We have already noted that the large particles seen in freeze-etching of mature chloroplasts are confined to the grana thylakoids. The large particles are not seen in developing chloroplasts prior to grana formation. The large particles seem to be related in some way to membrane stacking and to the presence of chlorophyll b, and do not correlate with functional photosystems, either PS–1 or PS–2.

4.7—
Assembly of the Thylakoid Membrane

4.7.1—
Protochlorophyllide

The structural events of chloroplast development were covered in chapter 3. In this chapter we will consider the net synthesis of chlorophyll and the formation of photosystems 1 and 2 on illumination of dark-grown seedlings. The small amount of protochlorophyllide (Mg-2-vinyl pheoporphyrin a₅ ) which is accumulated by the etioplasts of dark-grown seedlings is localized in the prolammelar bodies. Three spectoscopic forms of protochlorophyllide with slightly different absorption maxima (at 628 nm, 637 nm and 650 nm) can be distinguished. Two of these (Pchl-637 and Pchl-650) are converted rapidly to chlorophyllide a on illumination, while Pchl-628 is photo-inactive. The molecular basis for the difference in spectroscopic properties of the protochlorophyllides is not established. Different states of aggregation of the pigment or different modes of binding to protein have been suggested. In the photo-reduction of protochlorophyllide to chlorophyllide a , two hydrogen atoms are added to the 7,8 positions of the porphyrin ring and this requires that the protochlorophyllide

― 100 ―

is bound to a protein, termed holochrome (Boardman, 1966). The photoactive protochlorophyllide holochrome can be isolated from bean and barley seedlings. The origin of the hydrogen atoms for the photo-reduction is not known. In the membranes of the prolamellar body, the protochlorophyllide molecules appear to be organized into small units consisting of at least 4 protochlorophyllides, and there is some evidence that the units may contain as many as 20 chromophores.

4.7.2—
Chlorophyll Accumulation

Chlorophyll accumulation on the illumination of 6-day-old dark-grown barley seedlings with white light is shown in Fig. 4.6. On turning on the light a rapid conversion of protochlorophyllide to chlorophyllide a occurs, followed by a lag of about 30 minutes before additional chlorophyll is formed. During the lag phase, the esterification of chlorophyllide a with phytyl alcohol to chlorophyll a takes place. Rapid chlorophyll synthesis occurs after about 2.5 hours. The ratio of chlorophyll a to chlorophyll b is shown by the broken line. Chlorophyll b cannot be detected immediately after the photo-conversion of protochlorophyllide, but it is formed during the lag phase and the ratio of chlorophyll a /chlorophyll b falls rapidly. It takes four hours of illumination, however, before the chlorophyll a /chlorophyll b ratio approaches that of the mature chloroplast. The enzymic steps in the formation of chlorophyll b are not known although radiotracer studies suggest that chlorophyll b is formed from chlorophyll a (Shlyk, 1971).

4.7.3—
Development of the Photosystems

In the greening barley seedling, photosynthetic oxygen evolution is detected after 30 minutes of illumination. After 2 hours, the rate of oxygen evolution per gm fresh weight of leaf is as high as that in the greened leaf at 45 hours. However, when the photosynthetic rate of O₂ evolution is related to chlorophyll content of the leaf, it is 80-fold greater after 90 minutes than after 45 hours. The photosynthetic units of PS–1 and PS–2 are functional at an early stage of greening, but the size of the units are small compared with the photosynthetic units in the mature thylakoid membrane. During the greening process, chlorophyll a and chlorophyll b are synthesized and incorporated into the light harvesting units of the photosystems.

PS–1 is active ahead of PS–2. In isolated plastids from greening barley seedlings, appreciable PS–1 activity, including cyclic phosphorylation is observed as early as 15 minutes after turning on the light (Henningsen & Boardman, 1973; Plesnicar & Bendall, 1972).

The cytochromes which are localized in photosystem 1 in the mature chloroplast (cytochrome ¦ , cytochrome b₆ and cytochrome b–559_LP ) are already present in the etioplasts of dark-grown seedlings but cytochrome b–559_HP is

― 101 ―

Figure 4.6
Chlorophyll accumulation during the greening of dark-grown
barley seedlings. The solid line indicates total chlorophyll ( a+b )
and the broken line the ratio of chlorophyll a /chlorophyll b.
(From Henningsen & Boardman, 1973.)

formed during greening (Table 4.4). However the synthesis of cytochrome b– 559_HP does not correlate with the onset of oxygen evolution and this cytochrome does not appear to be essential for photosystem 2 activity (Henningsen & Boardman, 1973). In greening pea seedlings, the photo-oxidation of cytochrome ¦ , and the photo-reduction of cytochrome b₆ which require an active photosystem 1, could be detected after 20 minutes of greening.

There are no significant changes in the amounts of the phospholipids or galactolipids of greening pea leaves during the first 5 hours of greening, but the

― 102 ―

Table 4.4. Cytochrome contents of greening barley leaves (from Henningsen & Boardman, 1973).
Time of greening (hr)	Cyt ¦	Cyt b –559_LP + Cyt b₆ (male fresh weight)	Cyt b –559_HP
0	0.73	2.0	0
2	0.71	2.4	0.05
4	0.73	2.0	0.33
6	1.15	2.9	0.38
8	0.81	2.0	0.76
24	1.16	2.7	1.8

galactolipids increase after that time during grana formation. The fatty acid composition is also constant for 5 hours after illumination.

This constancy of the lipid and fatty acid compositions supports the electron microscope evidence that the membranes of the etioplast are the precursors of the first thylakoids. It is apparent that the etioplasts already contain many of the components which are required for functional photosystems (see also chapter 3).

4.7.4—
Thylakoid Structure During Greening

Earlier studies with isolated plastids from bean and pea seedlings suggested that there was a correlation between the onset of grana formation and an active PS–2, but more recent work shows that grana are not essential for photosynthetic oxygen evolution. Grana formation occurs during the phase of rapid chlorophyll synthesis, which suggets that the degree of grana development may be related to the chlorophyll content of the plastid. This view is supported by work on plants grown at different intensities (section 4.6). If dark-grown seedlings are greened in intermittent illumination (2 minute light periods separated by 15 minute dark periods) grana are not formed, but the plastids exhibit good photochemical oxygen evolution.

4.8—
Relationship of Chloroplast to Cytoplasm

Chloroplasts are surrounded by an envelope which consists of two membranes, the outer and inner membranes. The inner membrane is a permeability barrier to the diffusion of many metabolites from the chloroplast to the cytoplasm and vice versa. For example, the inner membrane is practically impermeable to sucrose, which is a major product of photosynthesis. It is very slowly permeable to ATP and impermeable to pyridine nucleotides and hexosephosphates (Heber, 1974). How then are reduced carbon compounds transported into the cyto-

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plasm and how is photosynthetic ATP made available for many other cell processes, such as protein synthesis?

Heldt and his colleagues (1975) have postulated that specific carriers or translocators are involved in the transfer of certain metabolites across the inner membrane of the chloroplast envelope (Heber, 1974). For example, the so-called phosphate trarnslocator facilitates the transfer of 3-phosphoglycerate (PGA), dihydroxyacetone phosphate (DHAP), glyceraldehyde-3-phosphate (GAP) and inorganic phosphate across the inner membrane in a competitive fashion. Figure 4.7 depicts the transfer of carbon and phosphate from the

Figure 4.7
Movement of carbon and phosphate energy from the chloroplasts to
the cytoplasm. ATP is formed in the cytoplasm as the result of
the transfer of dihydroxyacetone phosphate (DHAP) and
oxaloacetate (OAA). Sucrose is formed from DHAP.
(From Heber, 1974.)

chloroplast. It seems likely that sucrose is formed outside the chloroplast although this is not established. ATP and NADPH produced inside the chloroplast by photosynthetic electron flow are used in the conversion of PGA to DHAP. DHAP is exported from the chloroplast to the cytoplasm, where it is converted into sucrose or used for the generation of reducing power and ATP. The PGA can then re-enter the chloroplast.

Reducing power is transported across the inner membrane by a counter exchange of malate and oxaloacetate. NAD-dependent malate dehydrogenase occurs both in the cytoplasm and the chloroplast, while NADP-dependent dehydrogenase occurs in the chloroplast and is activated by light. Heldt et al., (1975) have postulated that the counter exchange of malate and oxaloacetate is facilitated by a dicarboxylate translocator.

Rapid transfer of metabolites between chloroplast and cytoplasm is crucial for the operation of the C₄ -dicarboxylic pathway of photosynthesis. In plants

― 104 ―

with the C₄ -pathway, CO₂ is fixed initially by reaction with phosphoenolpyruvate in the mesophyll cells. The resulting oxaloacetate is reduced to malate or transaminated to aspartate. Depending on the species of C₄ plant, one or other of these acids is transported to the bundle sheath cells, where the C₄ -acid is decarboxylated to give CO₂ and a 3-carbon compound. The CO₂ is then refixed by the Calvin carboxylation cycle (Hatch & Slack, 1970).

Chapter 5—
Plant Mitochondria

5.1—
Introduction

Mitochondria are the sites of non-photosynthetic energy transduction in eukaryotic cells which, carry out aerobic metabolism. Energy transduction includes those processes by which the chemical potential energy of organic substrates is transformed into a readily mobilized form, adenosine-5'-triphosphate (ATP). Organic substrates are oxidized and the free energy of oxidation is conserved by processes which are common to all mitochondria, regardless of source. Thus, with regard to the oxidative and phosphorylative processes, information obtained from studies in animal mitochondria is applicable to plant mitochondria.

Notable differences between plant mitochondria and animal mitochondria do occur, although these differences do not contradict the basic similarities in the mechanism of energy transduction. For example, plant mitochondria possess external reduced nicotinamide adenine dinucleotide (NADH) dehydrogenases which oxidize exogenous NADH; mitochondria from animal sources lack this capability. Mitochondria from many plant sources are relatively insensitive to cyanide inhibition, a feature not found in animal mitochondria. On the other hand, the b -oxidation pathway of fatty acids is located in animal mitochondria, whereas in plants, the enzymes of fatty acid oxidation occur in the glyoxysomes.

In this chapter, the morphology and function of plant mitochondria are discussed. In almost all cases, information is drawn from studies with mitochondria from higher plants. Emphasis is placed on the components of the plant mitochondria respiratory chain and their interactions with each other. Current ideas on oxidative phosphorylation are discussed with reference to knowledge gained from studies with animal and yeast mitochondria. Reversed electron flow and ion transport activities are considered with reference to studies in plant mitochondria. Structure and function relationships are sought, but in many instances, sufficient evidence is not available or available only from studies with mammalian or avian systems; it seems unwarranted, however, to draw exact parallels between animal and plant systems.

5.2—
Morphology

5.2.1—
Morphology in Situ

Mitochondria in living cells are highly pleomorphic, as shown by phase contrast microcinematography by Hongladarom et al., (1965). Pleomorphism is reflected

― 106 ―

also in thin section electron microscopy, in which mitochondria appear as roughly circular profiles as well as highly elongated or irregular cross sections (Fig. 5.1a). The circular sections may represent transverse or oblique sections through an otherwise elongated organelle. The diameter of the elongated mitochondrion appears to be about 0.4 to 0.5 µm, while the length may be several micrometers long. Although rods or apparent spheres are the most common profiles seen, sections derived from branched or cup shaped organelles have also been discerned (Bagshaw et al., 1964). The recent analysis of serial sections of yeast cells by Hoffman and Avers (1973), which showed that yeast contains a single, giant, branched mitochondrion, suggests that the irregular cross sections of mitochondria of other cells might also be sections of a single branched organelle.

The mitochondrion consists of a double membrane system with an inner convoluted membrane enclosing the matrix, and surrounded by a smooth outer membrane (Fig. 5.1a, 5.1b). High resolution electron micrographs of material

Figure 5.1a
Mitochondria in phloem parenchyma cells of a maize leaf. Magnification bar = 1 m m.
(Micrograph courtesy of O. E. Bradfute and Diane C. Robertson.)

fixed with glutaraldehyde and post-fixed with osmium tetroxide show the tripartite nature of both the inner and outer membranes. Each membrane has a thickness of approximately 9 nm (Baker et al., 1968).

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Figure 5.1b
Isolated mitochondria from mung bean hypocotyls. Mitochdria have been
suspended in 0.3 M mannitol prior to fixation. Magnification × 26,000.
(Micrograph courtesy of W. D. Bonner, Jr.)

5.2.2—
Morphology of Isolated Mitochondria

Electron micrographs of isolated mitochomdria show circular cross sections, presumably reflecting a spherical shape when released from their cellular environment. The electron micrographs of the intact isolated mitochondrion show clearly the two membrane systems, as well as the tripartite organization of each membrane. The fine structure of isolated mitochondria is highly dependent upon the osmolarity of the suspending medium (Baker et al., 1968). When mitochondria are suspended in 0.3 to 0.4 M sucrose or mannitol, the matrix appears contracted and electron dense (Fig. 5.1b), but when suspended in 0.2 M sucrose, the matrix appears more expnaded and less electron dense, and resembles that of mitochondria seen in sity . The dense matrix of mitochondria suspended in 0.3 to 0.4 M sucrose or mannitol is due to the hypertonicity of the suspending medium. Since the inner membrane is generally regarded as the osmotic barrier, the dense nature of the matrix reflects a water loss, which is reversible when the organelles are suspended in 0.2 M sucrose.

Negatively-stained water-lysed mitochondria show that the inner membranes have the characteristic stalked particles similar to those reported for mammalian mitochondrial membranes (Fernandez-Moran, 1962; Parsons et al., 1965). The particles have a headpiece with a diameter of 10 nm, attached to a stalk 3.5 to 4.5 nm wide and 4.5 nm long (Fig. 5.2). These resemble the particles identified with ATPase function in heart mitochondria (Racker et al., 1969).

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Figure 5.2
Portion of a surface spread and negatively stained summer squash
mitochondrion. The large areas of membrane (IM) are presumed to be
part of the inner membrane forming the shell of the mitochondrion.
The cristae (C) appear as smaller pieces of membrane of rounded shape
connected together by narrower (possibly tubular) pieces. The membranes
are coated with projecting knob-like subunits which are best seen lying in
the plane of the object at the edge of the pieces of membrane (arrow). The
dimension of the head of the subunit is 10 nm and the stem is 3.5–4.0 nm wide
and 4.5 nm long. (Parsons et al., 1965). (Reproduced by permission of the
National Research Council of Canada from the Canadian
Journal of Botany, Volume 43, 1965. pp. 647–55.)

― 109 ―

5.3—
Isolation and Purification

5.3.1—
Techniques of Isolation and Purification

Mitochondria have been obtained from a large number of plant sources including roots, storage tissue, stems and photosynthetic tissues. The usual problems of isolation, regardless of the source, are (a) the rupture of a rather rigid cell wall and (b) the prevention of damage to organelles through the release of intracellular, particularly vacuolar, contents. Ikuma (1970) listed a number of conditions for successful isolation of tightly-coupled mitochondria. These include (a) gentle tissue disruption, (b) rigorous exclusion of contaminating particles and (c) the use of a buffered grinding medium isotonic with mitochondria and containing a variety of protective reagents. Most investigators employ some device to reduce quickly the tissue to a coarse slurry, which is passed through a cloth filter to remove large debris. The fraction which sediments between 1,000 g and 10,000 g is collected as the mitochondrial fraction. This fraction will oxidize all the intermediates of the Krebs tricarboxylic acid cycle, exhibit respiratory control and yield ADP to O ratios approaching the theoretical value for the substrate used. The mitochondrial fraction can be further purified by density gradient centrifugation. This may be done in discontinuous sucrose gradients (Baker et al., 1968; Douce et al., 1972a) or Dextran-40 gradients (Solomos et al., 1973). Mitochondria form a band at the interface between 1.2 and 1.5 M sucrose (Douce et al., 1972a). This is recovered and diluted slowly to 0.3 M sucrose. This procedure yields mitochondria with intact outer and inner membranes as shown by electron microscopy. The integrity of the outer membrane is also shown enzymatically by the inability to reduce exogenous cytochrome c with NADH or succinate as substrates, unless the mitochondria have been subjected to mild osmotic shock which renders the outer membrane permeable to high molecular weight solutes.

During the disruption of cells and throughout the isolation procedures, a number of protective reagents must be present. Inclusion of sodium ethylenediamine-tetracetate (EDTA) in the isolation medium has been shown to give mitochondria with high respiration rates (Lieberman & Biale, 1955). EDTA probably removes cationic inhibitors although the specific cation complexed is unknown.

Many plant cells release phenolic compounds when ruptured. These phenolic compounds are oxidized in the presence of air and form polymers which are inhibitory to mitochondrial respiration and coupled phosphorylation. To prevent the damaging effects of phenolic compounds, a variety of reagents have been used successfully. Polyvinyl pyrrolidone is a competitive inhibitor of purified phenolase (Walker & Hulme, 1965), and has been used extensively as a protecting agent for mitochondrial isolation (Jones et al., 1965; Hulme et al., 1964; Wiskich, 1966). Other reagents include morpholinopropane sulphonate, cysteine, and sodium metabisulphite (Stokes et al., 1968). Morpholinopropane

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sulphonate is thought to form complexes with phenolic compounds, while sulphydryl reagents inhibit phenoloxidases.

Free fatty acids are known to uncouple oxidative phosphorylation from electron transport (Borst et al., 1962; Baddeley & Hanson, 1967). The uncoupling activity of fatty acids is reversed by the addition of bovine serum albumin. Bovine serum albumin also reverses the uncoupling activity of many other uncoupling agents, such as nitro- and halo-substituted phenols, dicumarol, and carbonyl-cyanide m -chloro-phenylhydrazone in rat liver mitochondria (Weinbach & Garbus, 1966). Dalgarno and Birt (1963) showed that free fatty acids were present in mitochondrial preparations from carrot root tissue. These included oleic, stearic, palmitic and some short chain fatty acids, as well as polyunsaturated C₁₈ acids. Mitochondria isolated from such tissues were uncoupled as shown by a P/O ratio less than 0.1. When bovine serum albumin was included in the isolation medium, mitochondria became well coupled, with a P/O ratio greater than 1.6 with succinate as substrate. As a matter of routine, most isolation procedures include 0.1% (w/v) of bovine serum albumin (Cohn Fraction V, low in free fatty acids). Bovine serum albumin binds fatty acids, and other lipophilic uncoupling agents, but the nature of the binding is not clear.

5.3.2—
Isolation from Green Tissues

Mitochondria isolated from photosynthetic tissues are rarely free from chloroplasts or chloroplast fragments. Rocha and Ting (1970) subjected spinach leaf material to linear sucrose gradients (40 to 80% w/v) and obtained fractions after equilibrium. They found, nonetheless, that the mitochondrial fraction was contaminated with 13% intact chloroplasts and 6% broken chloroplasts. The degree of contamination was estimated from the activities of characteristic marker enzymes. Malate dehydrogenase and cytochrome c oxidase served as mitochondrial markers, while chlorophyll content and triose-phosphate dehydrogenase were chloroplast markers.

5.4—
Mitochondrial Membranes

5.4.1—
Structure of Membranes

Electron micrographs of plant mitochondria show clearly the tripartite nature of both the outer and inner membranes. This may be interpreted as a lipid bilayer with the hydrophobic fatty acid chains oriented toward the interior of the bilayer (see chapter 2). The lipids of mitochondrial membranes are largely phospholipids. It is the current view that phospholipid bilayers are highly dynamic, with a high degree of fluidity in the fatty acid region, as well as high lateral mobility of the phospholipids in the plane of the membrane.

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The membrane proteins may form loose interactions with the lipid bilayer, or may be very tightly associated with the membrane. The loosely associated proteins most likely have exposed hydrophilic side chains and are easily extracted from membranes. The proteins tightly associated with membranes presumably have exposed hydrophobic side chains and are pictured as partially or wholly embedded in the lipid bilayer. These proteins are extracted from membranes with difficulty. Indeed, they may require a lipid environment for optimal activity.

5.4.2—
Membrane Lipids

A survey of the lipid composition of mitochondrial membranes reveals great differences depending upon the source of mitochondria. McCarty et al., (1973) investigated the phospholipid composition of the inner and outer membranes of mung bean (Phaseolus aureus ) and potato tuber (Solanum tuberosum ) mitochondria which were prepared in such a way as to exclude contaminating particles. Phospholipids comprised 90% or more of the mitochondrial membrane lipids. The main phospholipids were phosphatidyl choline, phosphatidyl ethanolamine and phosphatidyl glycerol, except in the outer membrane which did not contain significant amounts of phosphatidyl glycerol. The phospholipid composition is shown in Table 5.1 together with that of beef heart mitochondria

Table 5.1. Phospholipid composition of mitochondrial membranes (McCarty et al., 1973.)
	% total lipids
Phospholipid	mung bean inner membrane	potato inner membrane	potato outer membrane	beef heart mitochondria
Phosphatidyl choline	33	33	36	38
Phosphatidyl ethanolamine	32	33	64	37
Phosphatidyl glycerol	23	19	none detected	16

for comparison. Minor amounts of lysophosphatidyl ethanolamine, phosphatidyl inositol, and phosphatidyl glycerol were also found. The fatty acids of the three principal phospholipids of mung bean mitochondria were palmitic, linoleic and linolenic acids. Stearic acid occurred in conjunction with phosphatidyl glycerol. Schwertner and Biale (1973), by contrast, found that phospholipids comprised only about 50% of the total lipids of mitochondria from avocado (Persea sp.), cauliflower (Brassica oleracea ) and potato tubers, and found more phosphatidyl inositol than phosphatidyl glycerol. The principal fatty acids of the phospholipids of avocado, cauliflower and potato mitochondrial membranes were palmitic, linoleic and linolenic. The fatty acids of the neutral lipids of cauliflower

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mitochondria were C₁₆ and shorter chain acids, while those of potato mitochondria included palmitic, oleic, linoleic and linolenic. The fatty acid composition of the membrane lipids can be highly variable and may reflect the growth conditions of the tissue. Mitochondria from cold grown mung beans (15°C) have a higher amount of unsaturated fatty acids than mitochondria isolated from mung beans grown at 25°C (see also chapter 2).

Sterols comprise about 2.6% of the total lipids of maize shoot mitochondria (Kemp & Mercer, 1968). These may be esterified, of which cholesterol and b -sitosterol are the principal compounds. Of the unesterified sterols, stigmasterol and b -sitosterol are the principal ones. The fatty acids esterified to the sterols include lauric, myristic, palmitic, and linolenic as the most abundant.

5.5—
Enzymes

5.5.1—
Enzymes of the Tricarboxylic Acid Cycle

Mitochondria contain all the enzymes of the tricarboxylic acid cycle. Isolated mitochondria oxidize all of the acids of the cycle, and chromatographic analysis shows that the products are those expected for the reactions of the tricarboxylic acid cycle (Lieberman & Biale, 1956; Avron & Biale, 1957; Bogin & Erickson, 1965).

5.5.1.1—
Citrate Synthetase

Citrate synthetase (Citrate oxalacetate-lyase (CoA-acetylating)) activity is associated with a particulate fraction of leaf and root tissue from tobacco, bean, and soybean, which sediments at 10,000 g. This fraction consisted largely of mitochondria (Hiatt, 1962). Citrate was formed in the presence of acetyl CoA and oxalacetate. Citrate synthetase has been isolated from wheat scutellum mitochondria (Barbareschi et al., 1974) with minimal contamination by glyoxysomes. The mitochondrial citrate synthetase was released by sonic disruption of the mitochondria and recovered in the supernatent fluid. The purified enzyme had a molecular weight of 96,000 daltons as determined by its elution volume in Sephadex G-100 gel filtration. The K_m for acetyl CoA and for oxalacetate were 4 µM and 34 µM respectively. The enzyme was inhibited competitively with respect to acetyl CoA by ATP. In these respects, the mitochondrial citrate synthetase from wheat scutellum mitochondria is similar to that from mammalian sources.

5.5.1.2—
Pyruvate Oxidase

Pyruvate is oxidized by mitochondria with a requirement for catalytic amounts of one of the TCA cycle acids (Millerd, 1953; Walker & Beevers, 1956). TCA

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cycle acids were added to castor bean (Ricinus communis ) mitochondria at a concentration of 0.001 M . Oxygen consumption ceased after 60 minutes. When pyruvate at a concentration of 0.01 M was then added, oxygen consumption continued at rapid rates and linearly up to three hours. Other cofactors required were NAD, coenzyme A, ATP, and thiamin pyrophosphate.

5.5.1.3—
Isocitrate Dehydrogenase

A NAD-specific isocitrate dehydrogenase [L-iso-citrate:NAD oxidoreductase (decarboxylating)] has been purified from pea shoot (Pisum sativum var Alaska) mitochondria (Cox & Davies, 1967). The enzyme was released from pea shoot mitochondria ruptured by extrusion through a French pressure cell at 3,000 lbs in^–2 . The enzyme, whose K_m for NAD was 0.22 µM , was activated by Mn²⁺ and Mg²⁺ and to a lesser extent by Zn²⁺ , and inhibited by NADH, with K_i = 0.19 mM . The mitochondrial isocitrate dehydrogenase was specific for NAD which differentiates it from the cytosolic isocitrate dehydrogenase, which requires NADP as the electron acceptor.

5.5.1.4—
Malate Dehydrogenase

Malate dehydrogenase (L-malate:NAD oxidoreductase) exists in both mitochondrial and cytosolic forms. Moreover, the mitochondrial malate dehydrogenase may occur as several isozymes. Ting et al., (1966) separated two isozymes from young maize (Zea mays ) mitochondria after sonic disruption. Starch gel electrophoresis revealed a faint fast moving (toward anode) band and a major slow moving band. Grimwood and McDaniel (1970) also found a major slow moving band in polyacrylamide gel electrophoresis with several lighter fast moving bands. Boulter and Laycock (1966) attributed the minor bands to complexes of the mitochondrial malate dehydrogenase with other proteins, since re-electrophoresis of the eluted bands always gave a band in the main mitochondrial fraction as well as the original minor band. They determined the molecular weight of the main malate dehydrogenase to be 74,000 daltons.

Plant mitochondria oxidize malate readily, but glutamate must be included in the reaction mixture to remove the accumulated oxalacetate, due to the unfavourable equilibrium of the reaction. The oxidation of malate with endogenous NAD⁺ is inhibited by rotenone and antimycin (Day & Wiskich, 1974), but is insensitive to these inhibitors when exogenous NAD⁺ is supplied.

5.5.1.5—
Malic Enzyme

Malate may be oxidized by a NAD⁺ -dependent malic enzyme [L-malate:NAD oxidoreductase (decarboxylating)] with the formation of pyruvate. Macrae and Moorhouse (1970) showed that pyruvate accumulated during malate oxidation

― 114 ―

by cauliflower bud mitochondria, unless thiamin pyrophosphate was included in the reaction medium. Under the latter conditions, malate was probably oxidized by both malic enzyme and malate dehydrogenase so that in the presence of cofactors for pyruvate oxidation and citrate formation, the latter accumulates. Malic enzyme is the main pathway for malate oxidation by wheat shoot mitochondria, since oxalacetate did not inhibit malate oxidation, although it did inhibit transiently the oxidation of citrate and pyruvate (Brunton & Palmer, 1973). The activities of malic enzyme and malate dehydrogenase differ in mitochondria from various sources (Macrae, 1971b). While the relative activity of malate dehydrogenase has in all cases been greater than the activity of malic enzyme, the accumulation of pyruvate vs oxalacetate may vary considerably. Pyruvate was accumulated in preference to oxalacetate by a ratio of 27.0 in cauliflower bud mitochondria, while the pyruvate/oxalacetate ratio for wheat shoot mitochondria was 0.13. When malic enzyme activity is high, mitochondrial NADH levels are raised, and thereby reduce oxalacetate accumulation by product inhibition of malate dehydrogenase. A strong pH dependence of the activities of the two enzymes was also observed (Macrae, 1971a). At pH 6.0 to 7.0, pyruvate accumulates, while at pH values between 7.0 and 8.0 pyruvate accumulation drops and oxalacetate accumulation rises. The pathway may reflect the pH profiles of malic enzyme and malate dehydrogenase. Below pH 7.0, the activity of malic enzyme would maintain a high internal concentration of NADH which would favour the conversion of oxalacetate to malate; above pH 7.0, the decreased activity of malic enzyme and the consequent drop in the NADH levels would favour the oxidation of malate by malate dehydrogenase.

5.5.2—
Enzymes of Fatty Acid Oxidation

Fatty acids were oxidized by a particulate preparation from peanut cotyledons (Stumpf & Barber, 1956). This fraction was identified as the mitochondrial fraction. Cooper and Beevers (1969a,b) have separated the particulate fraction from castor bean and have shown that the enzymes of the b -oxidation pathway as well as the enzymes of the glyoxylate pathway are associated instead with a heavy particle, the glyoxysome. Mitochondria from castor beans contained less than 5% of the glyoxylate cycle enzymes and virtually none of the b -oxidation enzymes.

5.5.3—
Enzymes of Fatty Acid Biosynthesis

Isolated mitochondria from avocado mesocarp, flowerlets of cauliflower, and from white potato tubers were capable of incorporating radioactive acetate, acetyl-CoA, malonate, or malonyl-CoA into long-chain fatty acids (Yang & Stumpf, 1965; Mazliak et al., 1972). Cofactor requirements included coenzyme A, NADPH, ATP, and Mg²⁺ or Mn²⁺ . The principal acids formed were palmitic

― 115 ―

and stearic acids by avocado mesocarp mitochondria, while mitochondria from cauliflower and white potato tubers synthesized some mono-unsaturated fatty acids as well, i.e., palmitoleic (9-hexadecanoic acid) and, with longer incubation times, oleic (9-octadecanoic) and cis-vaccenic (11-octadecanoic) acids. With the appearance of the C₁₈ mono-unsaturated acids, stearic acid is found only in trace amounts, indicating that the unsaturated C₁₈ acids were formed from stearic acid.

5.5.4—
Enzymes of Phospholipid Biosynthesis

The synthesis of phospholipids proceeds via the following reaction:

Mitochondria isolated from flowerlets of cauliflower contain all of the enzymes necessary for the formation of CDP-diglyceride from glycerol-3-phosphate, when coenzyme A, ATP, CTP and fatty acids are provided (Douce, 1971). Radioactivity from ³² P-ATP was found in phosphatidic acid in peanut cotyledon mitochondria (Bradbeer & Stumpf, 1960). The incorporation of ³²p into phosphatidic acid was stimulated by the presence of small amounts of a ,b -diglyceride, indicating the presence of a mitochondrial diglyceride phosphokinase. ¹⁴ C-CTP was incorporated into CDP-diglyceride by cauliflower mitochondria (Sumida & Mudd, 1968). The radioactivity of CDP-diglyceride declines in the presence of a -glycerol phosphate or inositol, with the expected formation of phosphatidyl glycerol phosphate or phosphatidyl inositol. Using preparations carefully purified in sucrose density gradients from mung bean hypocotyl mitochondria, Douce et al. (1972b) showed that the CTP:phosphatidic acid cytidyl transferase activity was associated with the inner membrane fraction. Since the activity was not released upon sonication, it was assumed that the transferase was a membrane bound enzyme.

5.6—
Mitochondrial Electron Transport

5.6.1—
Components of the Respiratory Chain

Reducing equivalents derived from the oxidation of the TCA cycle acids are oxidized in a stepwise manner in the mitochondrial electron transport chain. The electron transport chain, or respiratory chain, is a series of functionally

― 116 ―

linked election carriers which undergo alternate reduction and oxidation, with molecular oxygen as the terminal election acceptor. Elections are donated by carriers of low redox potential to carriers of high redox potential. It is via these oxidation-reduction reactions that the main oxidative cellular energy transduction occurs, either through the formation of intermediate states which can be coupled to cellular work, (e.g., ion transport) or through the phosphorylation of ADP to form ATP, which can then mediate cellular endergonic reactions. The main components of the respiratory chain have been identified, both by characteristic reaction toward inhibitors and by spectral analysis. In its basic form, the respiratory chain in plant mitochondria is very similar to that of mitochondria from fungal or animal sources. The chain, (shown in Fig. 5.3) can be functionally separated into (a) NADH: coenzyme Q oxidoreductase; (b) succinate: coenzyme Q oxidoreductase; (c) reduced coenzyme Q: cytochrome c oxidoreductase; and (d) cytochrome oxidase. With refinements in detection systems, additional components have been identified. The concentration and molar ratios of the principal components have been determinded by Lance and

Figure 5.3
The generlized mitochondrial electron transport chain. The components are NAD, nicotinamide
adenine dinucleotide; Fp and Fp_s , the flavoproteins associated with NADH dehydrogenase
and with succinate dehydrogenase respectively; Q, Coenzyme Q or the quinone containing
carrier; cyt b , the complex of b -cytochromes; cyt c₁ , the tightly bound c-cytochrome (cyt
c-549 in plant mitochondria); cyt c, the salt extractable c-cytochrome; and cyt a+a₃ , the
a -cytochromes of cytochrome oxidase. Not shown are non-heme iron proteins which
have been tentatively identified in plant mitochondria, and bound copper of cytochrome
oxidase, which has been identified in plant mitochondria, and bound copper of cytochrome
oxidase, which has been identified in cytochrome oxidase from animal sources only.

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Bonner (1968) for mitochondria isolated from a number of sources and are given in Table 5.2. The concentrations of the cytochromes are quite similar to those of animal mitochondria.

Table 5.2. Concentration and stoichiometry of respiratory chain components in plant mitochondria (Lance & Bonner, 1968.)
	Conc. of carriers
	cyt aa₃	cyt b	cyt c	Fp	PN
Source		nM per mg protein
Helianthus tuberosus	0.10	0.10	0.15	0.38	0.92
Phaseolus aureus	0.11	0.12	0.17	0.58	4.10
Solanum tuberosus	0.17	0.20	0.27	0.89	3.50
Brassica oleracea	0.12	0.15	0.21	0.69	2.04
Symplocarpus foetidus	0.11	0.23	0.36	2.02	—

	Stoichiometry based on cyt c as unity
Helianthus tuberosum	0.69	0.67	1.00	2.90	7.5
Phaseolus aureus	0.65	0.67	1.00	3.64	17.8
Solanum tuberosum	0.65	0.76	1.00	3.30	14.I
Brassica oleracea	0.60	0.75	1.00	3.40	11.5
Symplocarpus foetidus	0.32	0.65	1.00	5.80	—

5.6.1.1—
Nicotinamide Adenine Dinucleotide

Malate dehydrogenase and isocitrate dehydrogenase are NAD⁺ linked enzymes. The oxidation of substrates is accompanied by the reduction of endogenous NAD. This is shown by the strong fluorescence of reduced NAD. Carefully isolated mitochondria contain sufficient endogenous NAD to oxidize malate or isocitrate. The oxidation of malate or isocitrate by endogenous NAD is inhibited by rotenone or amytal. Malate oxidation is stimulated by the addition of NAD, but the oxidation then becomes insensitive to rotenone inhibition (Wiskich et al., 1960; Day & Wiskich, 1974), just as the oxidation of exogenous NADH is insensitive to rotenone or amytal (Wilson & Hanson, 1969; Day & Wiskich, 1974) suggesting more than one pathway of NADH oxidation. Douce et al., (1973) and Day and Wiskich (1974) delineated three mitochondrial NADH dehydrogenases, one located on the outer membrane, a second located on the outer surface of the inner membrane, and a third on the inner surface of the inner membrane. Each dehydrogenase has a characteristic response to inhibitors. The outer membrane dehydrogenase is characterized by an antimycin insensitive NADH: cytochrome c reductase with added cytochrome c. In intact mitochondria, the NADH dehydrogenase of the outer surface of the inner membrane is coupled to cytochrome oxidase and goes through the antimycin sensitive site,

― 118 ―

but by-passes the rotenone sensitive site. This dehydrogenase shows NADH: cytochrome c reductase activity (antimycin sensitive) at low osmolarity only, due to the impermeability of the intact outer membrane to added cytochrome c. These first two dehydrogenases oxidize exogenous NADH. The NADH dehydrogenase of the inner surface of the inner membrane oxidizes the NADH linked to malate and isocitrate dehydrogenases. Electrons must go through the rotenone and the antimycin sensitive sites to cytochrome oxidase. In the intact mitochondrion, these various pathways interact, as demonstrated by the relief of antimycin or rotenone inhibition of malate: cytochrome c reductase activity by added NAD⁺ . Further evidence for the delineation of the NADH dehydrogenases is obtained through the P/O or ADP/O ratios of tightly-coupled mitochondria. Oxidation of malate or isocitrate gives ratios approaching 3.0, while the oxidation of NADH gives ratios approaching 2.0. The by-pass of the rotenone sensitive portion of the respiratory chain results in by-passing one of the phosphorylation sites as well (Wilson & Hanson, 1969).

5.6.1.2—
Flavoproteins of NADH Dehydrogenase

Storey (1970c, 1971a) distinguished a flavoprotein, F_PM , which was rapidly reduced upon addition of malate. The reduction of this flavoprotein was inhibited by amytal. He tentatively assigned a midpoint potential, E_m7.2 = –70mV for this flavoprotein. F_PM is the flavoprotein involved in the first energy conservation site and hence is most likely the flavoprotein involved in the NADH dehydrogenase located in the inner surface of the inner membrane.

F_Pha , a high potential non-fluorescent flavoprotein is rapidly reduced upon the addition of exogenous NADH in mung bean mitochondria (Storey, 1970d). This reduction is insensitive to amytal (Storey, 1970c). Its midpoint potential is approximately +110mV (Storey, 1971a). Flavoprotein F_Pha could be the flavoprotein associated with the dehydrogenases which oxidize exogenous NADH, but it is not possible to determine if it is the flavoprotein of the outer membrane dehydrogenase or the inner membrane dehydrogenase from the information available.

5.6.1.3—
Flavoprotein of Succinate Dehydrogenase

Isolated mitochondria from a number of plant sources oxidize succinate readily, with oxygen consumption rates of about 450 nM O₂ min^–l mg^–l protein (Douce et al., 1972a). Activation by ATP is required to obtain maximal rates of respiration with succinate as substrate, as well as for rapid response to addition of ADP (Drury et al., 1968). The activation by ATP is often attributed to the removal of inhibitory amounts of bound oxalacetate (Wiskich & Bonner, 1963) but the mechanism of the activation is not clear. The ATP effect is not due to phosphorylation mechanisms since neither oligomycin nor dinitrophenol affect the ATP activation (Singer et al., 1973).

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Hiatt (1961) reported the partial purification of succinate dehydrogenase from mitochondria from bean roots and tobacco (Nicotiana tabacum ) leaves. The K_m of the enzyme for succinate was 1 mM . Malonate inhibits competitively, with K_i = 0.24 mM . The apparent Michaelis constant of isolated mitochondria for succinate, in the presence of ADP and phosphate was 0.4 mM (Ikuma & Bonner, 1967). Singer et al., (1973) studied the succinate dehydrogenase of submitochondrial particles prepared by sonication of isolated mung bean and cauliflower mitochondria. The enzyme in submitochondrial particles was activated by a number of agents including substrate or Br^– . With Br^– activation, oxalacetate was removed, although it cannot be assumed that the oxalacetate was uniquely associated with the succinate dehydrogenase of the particles. Other activators included CoQ₁₀ , NADH, NAD-linked substrate (i.e., malate plus pyruvate), and ADP (Oestreicher et al., 1973). Succinate reducible flavoprotein was not detected spectrally (Storey, 1970a), but a flavoprotein associated with succinate dehydrogenase was determined chemically. Singer et al., (1973) found that succinate dehydrogenase contained covalently bound flavin as a histidyl-a -FAD. The flavin content was approximately 0.2 nM per mg protein. The molar ratio of flavin to enzyme was not determined.

5.4.1.4—
Ubiquinone

Studies on the role of ubiquinone in the plant mitochondrial electron transport chain have not been extensive. Beyer et al., (1968) extracted ubiquinone from mung bean submitochondrial particles. A single ubiquinone was found which co-chromatographed with ubiquinone-10. The spectrum of the extracted ubiquinone-10 has an absorption peak at 275 nm in the oxidized form and at 290 in the reduced form. The ubiquinone was reduced by succinate and NADH; at an aerobic steady state, 38% of the ubiquinone was reduced by succinate, while 56% was reduced by NADH. At anaerobiosis induced by succinate or NADH, 88% and 84% respectively were reduced. Sodium hydrosulphite (dithionite) gave additional reduction (i.e., about 93% reduction). The quinones were virtually 100% oxidized in aerobic suspension in the absence of substrates. Ubiquinone is generally acknowledged as part of the mitochondrial respiratory chain and is placed at the juncture of succinate dehydrogenase and NADH dehydrogenase. This placement is based on the considerable work with animal mitochondria. Storey and Bahr (1972), basing their conclusions upon measurements of the half times of reduced to oxidized transitions, and the times for 50% reduced to the fully reduced state, suggested that ubiquinone is in the main respiratory chain of mung bean mitochondria. Ubiquinone is the link between the dehydrogenases and the cytochromes, as usually regarded in animal mitochondria, but Storey and Bahr (1972) in addition placed F_Pha , the high potential flavoprotein, between ubiquinone and the b -cytochromes. F_Pha has no counter-part in the animal mitochondrial respiratory chain.

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5.6.1.5—
Cytochrome b

The cytochromes are hemo-proteins. Three classes of cytochromes, distinguished by their spectral properties, as well as by the nature of their prosthetic groups are found in mitochondria (see Fig. 5.4). The a -cytochrome contains as its prosthetic group, heme a , while the b- and c -cytochromes contain a heme closely related to protoporphyrin IX. In the c -cytochromes, the heme is covalently linked to the protein via sulphur atoms in a thio-ether linkage. In the reduced state, the cytochromes exhibit strong absorption bands in the visible region of the spectrum which have been useful in their identification and in the analysis of their function. In addition, both the oxidized and reduced forms absorb strongly in the region around 400 nm, which is a characteristic of all heme compounds.

The b -cytochromes are best resolved when their spectra are determined at low temperatures, e.g., 77°K. Three b -cytochromes have been identified and two others are suggested. Their spectral properties are summarized in Table 5.3.

Table 5.3. The b- cytochromes of plant mitochondria.
Cytochrome	a -peak, 25°C nm	a -peak, 77°K nm	E_m mV	Reference
b– 556	556	553– 554	+75 to + 100	(a)
		553	+ 75	(b)
b– 560	560	557	+ 40 to + 80	(a)
		557	+42	(b)
b– 558	557– 558	553– 555	– 70 to– 105	(a)
b– 566	566	561– 563	– 75	(a)
		562	– 77	(b)
S₂ O₄^2– reducible	557– 561	554, 560	– 100	(a)
References: (a) Lambowitz & Bonner, 1974; (b) Dutton & Storey, 1971.

Considerable variability exists in the nomenclature of the b -cytochromes. In conformity to the International Union of Biochemistry, the b -cytochromes are designated according to the a -peak of their reduced spectrum at room temperature (25°C). It should be noted that there is a blue shift of about 3 nm in the spectrum at 77°K relative to the spectrum at room temperature. The use of the a -absorption peak is further complicated by the fact that some authors use the absorption maximum at 77°K to designate the various b -cytochromes. In older nomenclature, mammalian cytochrome b –562, as orginally described by Keilin, was designated cytochrome b. As other b -cytochromes were discovered with an a -absorption peak significantly different from 562 nm, these were designated with subscripts. More recently, cytochrome b– 566 was thought to be directly involved in energy transduction and was designated cytochrome b_T , a transducing b -cytochrome, to differentiate it from cytochrome b– 562, or b_K .

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Figure 5.4
Prosthetic groups of the cytochromes.

― 122 ―

The multiplicity of b -cytochromes is due to different b -cytochromes in mitochondria rather than a splitting of the absorption bands at low temperature, since the peak heights do not change in synchrony in the presence of reducing agents, inhibitors or uncouplers (Lance & Bonner, 1968). The b -cytochromes are placed in the respiratory chain according to the following sequence (Storey, 1973):

Cytochrome b– 560 (557) was placed on the oxygen side of cytochrome b –556 (553) as a result of the determination of the rates of oxidation of the 556 and 560 components by an oxygen pulse of an anaerobic suspension of mitochondria. The 560 component was oxidized with a half-time of oxidation of 6 to 8 msec while the 556 component was oxidized with a half-time of 150 to 200 msec. The reduction by succinate of these two components in anaerobiosis showed, however, that b– 560 was reduced more slowly than b– 556 which was contrary to the expected rates in view of the rates of oxidation (Storey & Bahr, 1972; Storey, 1973). The slow reduction was ascribed to the more negative redox potential of b –560. The midpoint potentials of b –560 and b –556 would predict that b– 560 would be on the substrate side of b –556. Further resolution of the sequence of the b -cytochromes is necessary.

Cytochrome b– 566 was thought to be analogous to the b– 566 (b_T ) of mammalian mitochondria. Cytochrome b– 566 from animal mitochondria was found to undergo a midpoint potential shift as well as an enhanced reduction in anaerobic suspension when the respiratory chain was energized (Chance et al., 1970). This was interpreted as the formation of a high energy intermediate of phosphorylation directly involving cytochrome b– 566. In plant mitochondria, the midpoint potential shift of b– 566 was not observed (Dutton & Storey, 1971; Lambowitz et al., 1974). Although enhanced reduction of b– 566 by ATP or by energization of the respiratory chain could be demonstrated in plant mitochondria, it could be explicable by reverse electron flow through the b -cytochromes (Lambowitz et al., 1974; Lambowitz & Bonner, 1974). Thus the status of a transducing b -cytochrome in plant mitochondria is in question. In fact, cytochrome b –566 was excluded from the main sequence of the respiratory chain, since it remains oxidized in anaerobic suspensions (succinate reduced) while other b -components, pyridine nucleotides and fluorescent flavoproteins are reduced (Storey, 1969, 1974). There was a lack of equilibration between the low redox potential carriers with cytochrome b– 566. The function of cytochrome b –566 is left uncertain.

5.6.1.6—
Cytochrome c

Two c -type cytochromes have been detected in plant mitochondria. They have the same relationship as cytochrome c and c₁ in animal mitochondria (Lance & Bonner, 1968). The room temperature spectrum shows a large peak at 550 mn which shifts to 547 nm at liquid nitrogen temperature (77°K). As with cytochrome

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c in animal mitochondria, the cytochrome c (cyt c –547^[*] ) is easily extracted by salt solutions. A second component with a low temperature absorption peak at 549 nm remains after extensive washing with phosphate buffer. The 547 nm absorbing component is recovered in the phosphate buffer extract while the 549 nm absorbing component remains in the pellet. Both are reducible by ascorbate, which differentiates them from the b -cytochromes. The spectral properties of the 549 absorbing c component and its strong binding to mitochondrial membranes relate this c component to cytochrome c₁ of animal and yeast mitochondria. The midpoint potentials of the two c-cytochromes of mung bean mitochondria have been determined to be +235 mV in both cases (Dutton & Storey, 1971). The half-time of oxidation of cytochrome c– 547 and c –549 are 3.0 and 3.1 msec respectively when KCN treated anaerobic mitochondria were pulsed with 14 µM O₂ . The electron transfer sequence of the c -cytochromes was given as cyt c– 549 to cyt c– 547 (Storey & Bahr, 1972).

Cytochrome c (cyt c– 547) is essentially identical to cytochrorne c from all eukaryotic sources. Cytochrome c from one source will react with the reductase and oxidase from quite distantly related sources. The amino acid sequence of cytochrome c from a large number of sources is shown to have a high degree of homology (Nolan & Margoliash, 1968; Dickerson et al., 1971). This homology is all the more striking when the tertiary structure is considered. For example, the amino acids about the heme show a high degree of conservatism among the cytochrome c proteins examined. Those amino acid residues important to the structure and function of the protein have suffered few substitutions in the course of evolutionary history.

5.6.1.7—
Cytochrome Oxidase

The cytochrome oxidase of plant mitochondria contains cytochromes a and a₃ , as does that from animal mitochondria. These are two spectroscopically differentiated components, although two separate chemically different entities have not been isolated. The optical properties are well differentiated in the presence of cyanide or azide, which binds to cytochrome a₃ . The a -band of cytochrome oxidase at room temperature is located at 602 nm; at 77°K, there is a blue shift to 598 nm. The reduced spectrum of cytochrome a is revealed in the difference spectrum of an azide treated aerobic suspension minus an aerobic suspension. All components are oxidized except for cytochrome a, which shows a symmetrical reduced a -band at 598 nm. The a₃ spectrum is shown in a difference spectrum of an anaerobic (succinate reduced) plus azide suspension, minus an aerobic plus azide suspension. The reduced a -peak of cytochrome a cancels and the reduced a -peak of cytochrome a₃ is shown with its maximulm at 603 nm (Lance & Bonner, 1968). The midpoint potentials of cytochromes a and a₃ are

[*] The 77°K absorption peak.

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+190 and +380 mV respectively (Dutton & Storey, 1971). Half-times of oxidation in oxygen pulsed anaerobic suspension are 2.0 msec and 0.8 msec respectively for cytochromes a and a₃ (Storey, 1970b).

5.6.1.8—
Non-Heme Iron Proteins

Few investigations have been carried out on the occurrence and nature of non-heme iron proteins in higher plant mitochondria. Electron paramagnetic resonance (epr) signals characteristic of iron sulphur centres (non-heme iron proteins) were observed by Cammack and Palmer (1973) in mitochondria from Helianthus tuberosus and Arum maculatum with components at g = 2.02 and 1.93, 2.05 and 1.92, and at 2.10 and 1.87. Schonbaum et al., (1971) obtained an epr signal in NADH-reduced skunk cabbage (Symplocarpus foetidus ) mitochondria for a component at g = 1.94 and some complex components near g = 2.00. These iron sulphur centres no doubt participate in electron transport, since they undergo oxidation and reduction. At present, their precise functions are not known. They may be analogous to the iron sulphur centres identified in the NADH dehydrogenase segment of yeast and pigeon heart submitochondrial particles, which are believed to be closely involved in energy coupling (Ohnishi, 1973).

5.6.2—
Cyanide Resistant Respiration

An unusual characteristic of respiration in plant mitochondria is a partial insensitivity to cyanide inhibition. Partial insensitivity is exhibited to inhibition by azide, antimycin and 2-heptyl hydroxy-quinolin-N-oxide, all of which are potent inhibitors of oxygen uptake in animal mitochondria. This cyanide insensitivity may be almost 100% as in the spadix mitochondria from some aroid species, notably of Arum maculatum and Symplocarpus foetidus, partial as in mung bean mitochondria, or completely lacking as in the mitochondria from fresh, dormant white potato tubers (Bahr & Bonner, 1973a). In the latter, a cyanide insensitivity, and indeed a cyanide stimulation of respiration, may be induced upon vigorously aerating slices of potato tuber tissue in water for 24 hours. Mitochondria isolated from such aged potato tuber slices are much less inhibited by antimycin or cyanide (Hackett et al., 1960).

Because of the insensitivity to cyanide at concentrations which completely inhibit the respiration of animal mitochondria, it is unlikely that the cyanide insensitive respiration is due to an incomplete inhibition of cytochrome oxidase. This 'excess' oxidase hypothesis has been considered by various workers and received strong support from the finding that cytochromes a and c were incompletely reduced in the presence of cyanide (Chance & Hackett, 1959). The insensitivity to antimycin, which inhibits electron transport between the cytochrome b to cytochrome c region favours the argument that the cytochrome system is by-passed entirely and that there exists an alternate oxidase in mito-

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chondria showing cyanide insensitivity, although mitochondria of A. maculatum and S. foetidus have the conventional cytochrome complement (Bendall & Hill, 1956; Chance & Hackett, 1959). By and large, cytochrome c and cytochrome oxidase are reduced in the presence of cyanide (Bendall & Bonner, 1971). The incomplete reduction of cytochromes found by Chance and Hackett could be attributed to (a) the fact that the cytochromes are not always reduced by substrates relative to the reduction by dithionite; (b) possible spectral interference in the measurement of cytochrome c due to an oxidized b -cytochrome; or (c) in coupled mitochondria, significant reverse electron transport may cause the carriers on the oxygen side of the respiratory chain coupling site to become partially oxidized (Bonner & Bendall, 1968; Bendall & Bonner, 1971). Chance and Hackett (1959) and Bendall and Hill (1956) reported, however, that a b –type cytochrome becomes oxidized in the presence of cyanide and oxygen. Bendall and Hill called their b -component from A. maculatum cytochrome b₇ (a_max 560 nm), while Chance and Hackett identified an oxidizable b -component with an a_max at 558.5 nm. It was hypothesized that a b -cytochrome functioned as the shunt to the alternate oxidase. Such a role for a b -cytochrome is favoured, since Bahr and Bonner (1973b) reported that the known flavoproteins and ubiquinone had equal access to oxygen by either pathway, based on observations of their oxidation in the presence or absence of cyanide, and Storey and Bahr (1969a) found no identifiable carriers among flavoprotein, ubiquinone, the known b -cytochromes or c -cytochromes which could mediate electron transfer to the alternate oxidase. Cytochrome b₇ was identified with cytochrome b –557 (77°K) (Chance et al., 1968), but since the oxidation rate of cytochrome b– 557 was unaffected by m-chlorobenzhydroxamic acid, an inhibitor of the alternate oxidase (Schonbaum et al., 1971) it was felt that cytochrome b– 557 does not play a part in the alternate pathway, and should not be equated with Bendall and Hill's cytochrome b₇ (Erecinska & Storey, 1970). Bendall and Bonner (1971) showed that thiocyanate and other metal binding agents also inhibit the alternate pathway, suggesting a role for non-heme iron proteins or other metalloproteins. Efforts to identify non-heme iron proteins which may be the alternate oxidase have not been successful (Cammack & Palmer, 1973). Strong epr signals characteristic of iron-sulphur proteins in mitochondria of aged Jerusalem artichoke (H. tuberosus ) and in A. maculatum mitochondria were found. However, these are most likely the iron sulphur proteins of NADH: ubiquinone reductase since they were not reducible by succinate, and were, moreover, unaffected by hydroxamic acids. Schonbaum et al., (1971) found that the epr signals in NADH reduced skunk cabbage submitochondrial particles were enhanced by treatment with m-iodobenzhydroxamic acid. In addition to a signal at g = 1.94, which is characteristic of NADH dehydrogenase, a set of complex signals near g = 2.0 was detected at 77ºK. It was thought that the g = 2.0 signal may originate from the alternate pathway.

The possibility that the alternate oxidase may involve a flavoprotein has been explored. Flavoprotein oxidases which reduce oxygen with the formation of

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hydrogen peroxide are known, but such oxidases have not been identified for cyanide resistant mitochondria (Bendall & Bonner, 1971).

The function of cyanide resistant respiration is unclear. In the spadix tissue of maturing flowers of Arum and Symplocarpus, it may serve the function of thermogenesis. A rise in temperature of the spadix tissue ten degrees above the ambient has been recorded. This thermogenesis may aid these plants in pollination, since flowering occurs in early spring.

5.7—
Energy Linked Reactions of Mitochondria

5.7.1—
Oxidative Phosphorylation

5.7.1.1—
Coupling Sites

Oxidative phosphorylation, the process whereby the reaction

is coupled to the oxidation of reduced electron carriers of the respiratory chain occurs in plant mitochondria in the same manner as does the reaction in animal mitochondria. The sites of phosphorylation are identical (see Baltscheffsky & Baltscheffsky, 1974). These are the sites I in the NADH: ubiquinone reductase segment, II in the cytochrome b: cytochrome c reductase segment and III in the cytochrome oxidase segment of the respiratory chain. It has not been possible to identify precisely those components of the respiratory chain which are the energy transducers in each of the three sites, although these have been approximated from the change in redox potentials (D E₀ ') between adjacent carriers (Lehninger, 1965), by application of the crossover theorem (Chance & Williams, 1955), and from shifts in the midpoint potential of certain electron carriers upon the addition of ATP to uncoupled anaerobic suspensions of mitochondria (Wilson & Dutton, 1970a,b; Lindsay & Wilson, 1972; Chance et al., 1970; Ohnishi, 1973; Devault, 1971). The D E₀ ' indicates the thermodynamic feasibility of coupling between two adjacent components. Thus, coupling sites were thought to be located between endogenous NAD and the flavoprotein of NADH: ubiquinone reductase; between cytochrome b and cytochrome c of the cytochrome b :cytochrome c reductase; and between cytochrome a and oxygen of cytochrome oxidase. The application of the crossover theorem by and large confirms the general location based on thermodynamic grounds. According to the crossover theorem, electron transport through the coupling site is the rate limiting step in tightly coupled mitochondria. The carrier on the substrate side of the coupling site should become reduced, while the carrier on the oxygen side would become oxidized. Upon the addition of the phosphate acceptor, ADP,

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there would be observed a rapid transient oxidation of the carrier on the sub-strate side, and a reduction of the carrier on the oxygen side of the coupling site, concomitant with the release of controlled respiration in the state 4-state 3 transition (see Table 5.4).

Table 5.4. Definition and values of steady states in mitochondria (Chance & Williams, 1955.)
State	(O₂ )	ADP levels	Substrate	Respiratory rates	Rate-limiting substance
1	> 0	low	low	slow	ADP
2	> 0	high	~0	slow	substrate
3	> 0	high	high	fast	respiratory chain
4	> 0	low	high	slow	ADP
5	0	high	high	0	oxygen

Wilson and Dutton (1970a) reported that the midpoint potential (E_m ) of cytochrome a₃ becomes more negative upon energization of an anaerobic suspension of rat liver mitochondria by ATP. Similarly they report that the E_m of one of the b cytochromes (a_max 564 nm) increases upon energization by ATP (Wilson & Dutton, 1970b; Chance et al., 1970). These shifts of the midpoint potentials were attributed to changes in ligand interaction energy of the heme iron upon energization, and that the iron atoms of these cytochromes were directly involved in energy coupling.

Such a change in the midpoint potential was postulated by Wang (1970). In his model, the phosphoimidazole group becomes a much weaker coordinating ligand for the Fe(II) than imidazole, and hence should lower the midpoint reduction potential of the corresponding electron carrier. Similar experiments in the iron-sulphur centres of the site I region of the respiratory chain showed that of the five iron sulphur centres identified, ATP affected the reduction potential of only one of these centres, designated as centre I (Ohnishi, 1973). Addition of ATP caused a partial oxidation of centre I when the reduction potential was poised at a value where the iron was almost completely reduced. These observations suggested that the reduction potential of centre I was dependent upon the phosphate potential, and that the addition of ATP caused a lowering of the midpoint potential of the centre I iron sulphur protein. The significance of the changes in the midpoint potential was questioned, at least for the site II region, since none of the b -cytochromes showed an ATP-induced potential shift when investigated in plant mitochondria (Dutton & Storey, 1971; Lambowitz et al., 1974). One must conclude that there is a fundamental difference in the function of the b -cytochromes in plant and animal mitochondria or that the changes in the midpoint potential do not reflect the formation of an energy transducing species. Lambowitz et al., took the latter view and attributed

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the midpoint potential changes to a reverse electron flow with cytochrome b –566 and cytochrome a₃ equilibrating with the redox mediators by way of cytochrome c, while the iron sulphur centre of site I equilibrated through the nicotinamide adenine nucleotide pool.

5.7.1.2—
ADP:O Ratios

The general location of the three coupling sites means that for tightly coupled mitochondria, predictable ratios of the moles of ADP phosphorylated to the gram-atoms of oxygen consumed (ADP/O or P/O) may be obtained for each substrate. Thus succinate should give a ratio of 2.0, NAD-linked substrates a ratio of 3.0 and a -ketoglutarate a ratio of 4.0 (one phosphorylation via succinyl-CoA, three phosphorylations via the respiratory chain NADH). These ratios are largely confirmed in plant mitochondria. Early attempts gave results much below the predicted ratios, but with refinements in the procedure for the isolation of plant mitochondria, the inclusion of required cofactors, and the exclusion of competing reactions, predicted ratios have been approached, as shown in Table 5.5. Exogenous NADH does not interact with respiratory chain NAD,

Table 5.5. Oxidative phosphorylation by isolated mitochondria (Douce et at., 1972a).
Source	Substrate	P:O	Respiratory control index
Phaseolus aureus	succinate	1.68	3.2
	malate	2.56	5.4
	NADH	1.55	4.5
Solanum tuberosum	succinate	1.64	3.4
	malate	2.59	4.2
	NADH	1.48	2.4

but with flavoproteins of the external dehydrogenase. These ultimately enter the respiratory chain at the level of cytochrome b and hence electrons go through two coupling sites only (Douce et al., 1973).

5.7.1.3—
Energy Coupling in Cyanide Resistant Respiration

Mitochondria of Symplocarpus are fully capable of energy conservation in the uninhibited state. Hackett and Haas (1958) found P/O ratios greater than 3 for a -ketoglutarate oxidation by skunk cabbage mitochondria. In the presence of cyanide, the energy coupling sites associated with the cytochromes are by-passed, and energy coupling involves only the NADH:ubiquinone reductase portion of the respiratory chain. Storey and Bahr (1969b) obtained ADP/O ratios of 1.3 with succinate as substrate in skunk cabbage mitochondria in the absence

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of cynide; the ratio was zero in the presence of cyanide. With malate as the substrate, the ADP/O ratios were 1.9 and 0.7 in the absence and presence of 0.3 mM KCN respectively, showing that coupling sites II and III were effectively by-passed while coupling site I, which could be activated by malate but not by succinate, was still functioning in the presence of cyanide.

Wilson (1970a), however, has proposed that energy coupling occurs in the alternate pathway as well. He found that cyanide treated mitochondria of A. maculatum and Acer pseudoplatanus produced ATP, with an ATP/O ratio of 0.5 at high O₂ concentration (greater than 100 µM ) with succinate as the substrate. Submitochondrial particles of A. maculatum which lacked malate dehydrogenase activity oxidized succinate with P/O ratios of 0.2 to 0.6 (Wilson 1970b). Since malate dehydrogenase-depleted particles were still capable of phosphorylation, the possibility that the phosphorylation was due to the oxidation of a product of succinate oxidation, i.e., malate, could be eliminated, therefore suggesting that a phosphorylation site exists in the alternate pathway which functions only at high C₂ concentration. Cyanide inhibited mitochondria from spadices of A. maculatum and from mung bean hypocotyls still retain an energy related function through the cytochromes. Bonner and Bebdall (1968) showed that a substrate linked reverse electron flow is active in spadix mitochondria. Cytochrome c and cytochrome oxidase of cyanide treated mitochondria were reduced by ascorbate plus, N,N,N',N' -tetramethyl-p -phenylenediamine (TMPD), an electron donor to cytochrome c. Subsequent addition of succinate caused a reoxidation of cytochrome c and cytochrome a with a partial reduction of cytochrome b. The reoxidation of cytochromes c and a was prevented by uncoupling concentrations of carbonylcyanide-phenyl-hydrazone or 2,4-dinitrophenol. Wilson and Moore (1973) also showed that cyanide inhibited mung bean mitochondria could carry out oxidation of ascorbate plus TMPD. The oxidation was thought to involve a reverse electron flow through coupling site II and energized by ATP or the high energy intermediate.

5.7.1.4—
Mechanism of Coupling

The mechanism of energy coupling remains an intractable problem in spite of the impressive array of workers in this area. Three outstanding hypotheses are under active consideration: (a) the chemical intermediate hypothesis; (b) the chemiosmotic hypothesis; and (c) the conformation coupling hypothesis.

The operation of the chemical intermediate hypothesis may be summarized as follows:

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where A and B are adjacent electron carries at the coupling site, X and I are unknown couplers common to all coupling sites (see Greville, 1969). The scheme postulates the existence of both non-phosphorylated and phosphorylated intermediates. A non-phosphorylated intermediate accounts for the action of uncouplers, which is independent of the presence of phosphate. The nonphosphorylated intermediate can also be coupled to work functions such as ion transport, which is sensitive to uncouplers, but not to the phosphorylation inhibitor, oligomycin. The phosphorylated intermediate transfers a phosphoryl group to ADP. Hill and Boyer (1967) showed that the bridge oxygen between the b and g phosphorus of ATP is furnished by ADP. Hence, the mechanism of phosphorylation involves an activation of phosphate.

The chemiosmotic hypothesis in its simplest form as first proposed by Mitchell (1961) involves a vectorial metabolism in which the elements of water are transported to opposite sides of the mitochondrial membrane (Fig. 5.5).

Figure 5.5
Representation of the Mitchell chemiosmotic
scheme for oxidative phosphorylation.
(Redrawn from Mitchell, 1966.)

Thus oxido-reduction reactions create a pH gradient as well as a potential difference (outside positive) which tends to drive H⁺ back across the membrane into the inner compartment. This force is the proton-motive force, and it is this flow of protons through the coupling site (a reversible ATPase) which drives ATP synthesis. The phosphorylation reaction is represented as follows:

It can then be shown that in the absence of a transmembrane electrical potential, a pH differential of 3.5 units is required to poise the ratio of ATP/ADP = 1 while a potential difference of 210 mV would be required in the absence of pH gradients (Mitchell, 1966). The translocation of protons and equivalent OH– is effected by ionizable groups which are designated XH and IOH, corresponding to components of an ATPase. The proposed intermediate X-I of the ATPase must have a sufficiently low hydrolysis constant at the high electrochemical

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potential of H⁺ in the outer phase to come to reverse equilibrium with water according to the reaction

On the other hand, the hydrolysis constant of the intermediate X ~ I must be in the order of 10⁵M , and the intermediate X ~ I must be in equilibrium with the ATP/(ADP + P_i ) couple in the inner phase, so that

The system vibrates between states in which the intermediate X – I is alternately accessible to the outer and inner phases, being X – I when in contact with the outer phase and X – I when in contact with the inner phase. The transition of X – I to X ~ I is due not to the pumping of energy into X–I, but to a lowering of the ground state energy for X – I hydrolysis by some 10,000 calories per mole on translocation through an anisotropic membrane.

While experimental verification of the chemiosmotic mechanism has been realized only in chloroplasts by acid-base trasitions (Jagendorf & Uribe, 1966; see also chapter 4), similar experiments have not been successful in mitochondria. Cockrell et al., (1967) have shown ATP synthesis after establishing a K+ gradient by valinomycin-treated mitochondria. However, Glynn (1967) argued that ATP synthesis via a K⁺ gradient could be explained equally well on the basis of a membrane potential rather than cation transport down a chemical gradient through an ATPase.

In their general aspects, the chemical intermediate hypothesis and the chemiosmotic hypothesis differ primarily in the nature of the initial driving force for coupled phosphorylation. The two hypotheses converge at the level of the ATPase in that both call for an unknown intermediate, X ~ I (Greville, 1969).

The conformational coupling hypothesis of Boyer (1967) differs in that the ATPase has a high affinity for P_i and ADP. Tightly bound ATP is formed via a nucleophilic attack by ADP on orthophosphate in a S_N² reaction with the displacement of water (Korman & McLick, 1970). According to the model, there exists an energy requirement for the release of bound ATP from the complex, by changing the complex from one having a high affinity for ATP to one having low affinity (Boyer et al., 1973). The advantages of this hypothesis are that it explains most of the exchange reactions observed in coupled phosphorylation, and the coupling of ATP to energy yielding reactions of mitochondria.

5.7.2—
Reverse Electron Flow

Electron flow and energy transduction through the coupling sites of the respiratory chain are reversible processes. These are shown by the reduction of endogenous NAD and cytochrome b when substrates of higher reduction

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potentials are oxidized as well as the ATP induced oxidation of reduced cytochrome a + a₃ when the terminal oxidase is inhibited by sulphide. The properties of energy linked NAD reduction by plant mitochondria are similar to those reported in animal mitochondria (Chance, 1961; Chance & Hollunger, 1963). These include the requirement for succinate oxidation and for ATP (Storey, 1971b). Although ATP is required, the reduction of NAD is not sensitive to oligomycin. Hence NAD is reduced by reverse electron flow through the coupling site but without the participation of the ATPase as such. Uncouplers either inhibit the reduction when added before succinate, or reverse the reduction when added after a steady state reduction is attained. This can be interpreted as an effect upon the coupling site in preventing reverse electron flow, as well as a general release of controlled respiration so that the endogenous NADH is now oxidized rapidly through the coupling site. It is not possible to distinguish between the two alternatives.

Reverse electron flow through coupling sites II and III has been demonstrated by Storey (1972) and Lambowitz et al., (1974). When ATP is added to sulphideinhibited or to anaerobic suspensions of mung bean mitochondria, the b -cytochromes become reduced while cytochrome c and cytochrome a + a₃ become oxidized. The effect is reversed by uncouplers or by phenazine methosulphate (PMS) which mediates electron flow between cytochromes b and c. The reverse electron flow through coupling site II involves an ATPase (Lambowitz et al., 1974). Hydrolysis of ATP is observed in an anaerobic suspension of mung bean mitochondria supplemented with ascorbate plus TMPD and ATP. Addition of PMS caused an increase in the rate of ATP hydrolysis which is to be expected if PMS formed a shunt of electrons from reduced cytochrome b to cytochrome c.

5.7.3—
Ion Transport

Mitochondria contain two compartments, one of which is readily accessible to low molecular weight solutes such as sucrose or mannitol, and a second in-accessible to sucrose or mannitol. The former is identified with the inter-membrane space, and the latter, the mitochondrial matrix. The volume enclosed by the inner membrane behaves as an osmometer (Lorimer & Miller, 1969; Overman et al., 1970). Selective permeability to solutes and active transport are properties of the inner mitochondrial membrane. The movement of solutes across the inner membrane may be determined by assay for net increases in intramitochondrial contents. If the movement results in the net increase or decrease in the osmolarity of the sucrose-inaccessible volume, such solute movements will be reflected in volume changes of the inner compartment which can be monitored by changes in the light scattering properties of a mitochondrial suspension. Thus mitochondrial swelling is accompanied by a decrease in light scattering, while a contraction is reflected by an increase in light scattering.

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5.7.3.1—
Monovalent Cations

Permeability of the inner membrane of plant mitochondria to potassium and chloride ions is restricted. Slow, passive permeability to K⁺ and C1^– may be observed on transfer of mitochondria to isosmotic KCl, but when mitochondria are energized by the addition of substrate (NADH), rapid loss of both K⁺ and Cl^– occurs, accompanied by mitochondrial contraction (Wilson et al., 1969; Kirk & Hanson, 1973). In contrast to the energized extrusion of K⁺ and C1^– , mitochondria undergo an NADH induced swelling' in potassium acetate solution, and K⁺ and acetate ions are taken up. This has been interpreted as an active transport of acetate, while K⁺ penetrates along an electrochemical potential and chloride is not transported. The NADH-induced swelling in potassium acetate is inhibited by uncouplers of oxidative phosphorylation, by ADP plus P_i , or by respiratory chain inhibitors (Wilson et al., 1969; Lee & Wilson, 1972), but is restored when oligomycin is present with ADP plus P_i . An intermediate of oxidative phosphorylation possibly mediates active transport of acetate, but not of chloride. This is strengthened by the observation that mitochondria respire with expected respiratory control and ADP/O ratio when suspended in isotonic KCl, but lose respiratory control in potassium acetate.

Ionophorous antibiotics, valinomycin and gramicidin D, increase the permeability of the mitochondrial membrane to potassium ion as shown by the increased rate of passive swelling in potassium chloride solutions (Kirk & Hanson, 1973; Miller et al., 1970a). Valinomycin also increases the rate of NADH-induced swelling in potassium acetate, as well as in chloride, phosphate, sulphate and nitrate salts of potassium (Kirk & Hanson, 1973; Wilson et al., 1972) although the swelling due to acetate and phosphate reverses on exhaustion of NADH. This has been interpreted to mean that the swelling is due to the enhanced permeability of the inner membrane to potassium, as well as an active transport of acetate or phosphate. In the case of the latter anions, swelling is thought to be due to an enhanced permeability toward K⁺ while the anions diffuse along an electric potential. On cessation of NADH-supported respiration, acetate and phosphate leak out according to their chemical potential, followed by potassium ions, while no leakage of chloride, nitrate or sulphate occurrs as they are distributed along their chemical potential (Wilson et al., 1972). The picture which emerges is that the movement of potassium ion is controlled by the movement of anions, but K⁺ flux can be modified by ionophores such as valinomycin or gramicidin D. Valinomycin may stimulate K⁺ uptake via a H⁺ exchange as found by Rossi et al., (1967) in liver mitochondria, but such K⁺ uptake due to H⁺ exchange is not accompanied by swelling of the mitochondria.

5.7.3.2—
Divalent Cations

Ca²⁺ does not cause marked stimulation of respiration in plant mitochondria as it does in animal mitochondria, except with exogenous NADH as substrate

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(Miller et al., 1970b; Miller & Koeppe, 1971). A slight release from controlled respiration is detected with malate plus pyruvate or with succinate as substrates. This is associated with the accumulation of calcium and inorganic phosphate by mitochondria (Miller et al., 1970b). Extensive Ca²⁺ uptake by plant mitochondria occurs only in the presence of inorganic phosphate (Hodges & Hanson, 1965; Elzam & Hodges, 1968; Earnshaw et al., 1973; Chen & Lehninger, 1973). The phosphate dependent Ca²⁺ transport is energy dependent and may be supported by substrate oxidation or by ATP (Elzam & Hodges, 1968). The energy dependence seems to be at the level of an intermediate of oxidative phosphorylation. Substrate-supported Ca²⁺ transport is sensitive to uncouplers and respiratory inhibitors (Chen & Lehninger, 1973) as well as to ADP (Elzam & Hodges, 1968). The ATP supported Ca²⁺ transport is inhibited by oligomycin. Similar observations have been reported for Sr²⁺ transport by mitochondria (Johnson & Wilson, 1972). Other anions, e.g. acetate, arsenate, sulphate, chloride or nitrate promote neither Ca²⁺ nor Sr²⁺ uptake (Hodges & Hanson, 1965; Johnson & Wilson, 1972) although all will produce a metabolically independent swelling, while arsenate and acetate produce an active swelling as well, indicating that mitochondrial membranes are permeable to these anions, and will actively transport arsenate or acetate, as well as phosphate (Hanson & Miller, 1967; Johnson & Wilson, 1972; Lee & Wilson, 1972). Uptake of Ca²⁺ and inorganic phosphate results in the deposition of electron dense material in mitochondria which is dependent upon the concentrations of both Ca²⁺ and phosphate, and the time of incubation (Peverly et al., 1974). The composition of the deposits has not been ascertained, but is believed to be a form of calcium phosphate. The deposition of the phosphate salt of divalent alkaline earth metal ions may account for the contraction of mitochondria induced by Ca²⁺ . Since the volume changes of mitochondria are measured by light scattering changes, the formation of crystals within the mitochondria may increase the ight scattering properties of the suspension, and be interpreted as a contraction.

5.7.3.3—
Anion Transport

The importance of anion transporters in the movement of solutes across the mitochondrial inner membrane has been studied extensively in mammalian mitochondria (Chappell & Haarhoff, 1967; Harris, 1969). In these studies, the role of the phosphate transporter and the malate transporter is emphasized. Similar investigations were carried out by Phillips and Williams (1973b) and by Wiskich (1974). The presence of anion transporters was demonstrated by the spontaneous swelling of mitochondria in solutions of the ammonium salts of phosphate or malate. Ammonium salts were used because the mitochondrial membrane is readily permeable to ammonium ion. Mitochondrial swelling under these conditions is indicative of an osmotic adjustment due to the net influx of solutes into the mitochondrial matrix (Overman et al., 1970; Wilson et al., 1973). The swelling in ammonium phosphate was inhibited by

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N -ethylmaleimide which inhibits the phosphate-hydroxyl antiporter while the swelling in ammonium malate as well as the malate supported respiration was inhibited by 2-butylmalonate, 2-phenylsuccinate, benzylmalonate or p -iodo-benzylmalonate, all inhibitors of the dicarboxylate carrier (Phillips & Williams, 1973a,b). The anion transport system is interpreted as follows: (a) a phosphatehydroxyl antiporter which transports phosphate in exchange for hydroxyl ion; (b) a malate-phosphate antiporter which transports malate in exchange for phosphate; (c) a tricarboxylate-malate antiporter which transports tricarboxylate anions in exchange for malate; (d) other dicarboxylate anions enter by exchange with malate. It is the prevailing view that anions are actively transported, and that cations follow the anion transport along an electric gradient (Hanson & Miller, 1967). The essential role of anion transport in determining cation movement, however, is modified by the presence of cation ionophores.

Chapter 6—
Microbodies

6.1—
Introduction

Most of the organelles of the plant cell were detected by the classical techniques of light microscopy and were described by the turn of this century. Microbodies however were discovered relatively recently with the advent of the electron microscope. They were first recognized in the early 1950's as small spherical bodies in electron micrographs of mammalian kidney and liver tissue and similar organelles were reported in plant tissues in the early 1960's. Later in that decade they were isolated from plant tissues and from studies of their biochemical functions were recognized as being of major importance in plant cell metabolism. It is perhaps interesting to point out that the physiological significance to plant cell metabolism of major organelles, such as mitochondria and chloroplasts, was recognized long before their component biochemical reactions had been studied in detail, while the discovery and isolation of microbodies allowed some already well known metabolic pathways to be ascribed to a specific organelle.

Two types of microbody with identical structures but with distinct physiological functions have been isolated from plant tissues. They have been called peroxisomes and glyoxysomes to distinguish their separate functions in cell metabolism. Peroxisomes occur in the leaves of higher plants and are closely associated with chloroplasts. They are the sites for the oxidation of glycollic acid, a product of carbon dioxide fixation. The oxidation of this compound, results in a release of carbon dioxide and oxygen uptake which is called photorespiration. Glyoxysomes on the other hand, occur abundantly during the germination of those seeds which store fats as a reserve material, and contain the enzymes necessary for the breakdown of fatty acids to acetyl-CoA and the synthesis of succinate from acetyl-CoA.

All microbodies appear to contain flavin-dependent oxidases and catalase. The oxidation of a substrate by a flavin-linked oxidase is accompanied by the uptake of oxygen and the production of hydrogen peroxide, which is broken down by catalase to oxygen and water.

Microbodies, therefore, contribute to the uptake of molecular oxygen by the cell, but unlike mitochondria they do not contain any electron transport system which would be needed for the recovery of energy as ATP.

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6.2—
Structure and Occurrence

Microbodies are usually spherical, but can be ellipsoidal or dumbell shaped, and range from 0.2 to 1.5 µm in cross sectional diameter. Because of the low contrast between the matrix of the microbody and the cytosol they are not detectable by light microscopy but in electron micrographs they can be seen to have a single limiting membrane enclosing a granular matrix of moderate electron density (Fig. 6.1). The core of the microbody commonly contains

Figure 6.1a
A portion of a tobacco leaf cell showing a microbody with a crystalline
inclusion appressed to two chloroplasts. A mitochondrion lies
to the right of the microbody. (Magnification × 33,000).

b. A portion of a tobacco leaf cell, incubated in DAB medium, showing a heavy
deposition of osmium throughout the crystalline inclusion of the microbody.
(Magnification × 30,500). (Reproduced with permission from Frederick and
Newcomb (1969). Original prints supplied by Professor E. H. Newcomb.)

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fibrillar inclusions or a single large crystalline inclusion which may be formed by a reorganization of the fibrillar material. These crystalline bodies have been shown to have catalase activity (Frederick & Newcomb, 1969). No ribosomes or any form of nucleic acids have been detected in the microbody and hence they are not considered to be capable of self-replication. Since they are found in many tissues in association with the endoplasmic reticulum they are thought to be formed by this structure. Microbodies are therefore structurally very simple and cannot be confused in electron micrographs with any organelle except perhaps lysosomes from which they can be distinguished by cytochemical techniques.

The presence of catalase is a distinguishing feature of microbodies and this can be detected cytochemically using the dye 3,3'-diaminobenzidine (DAB). In fixed sections, catalase remains active and in the presence of hydrogen peroxide the dye is oxidized to give an osmiophyllic electron-dense product in the core of the microbody.

Microbodies have been seen in electron micrographs of many plant tissues. They have been found in the leaves of angiosperms, gymnosperms and bryophytes where they are numerous and are invariably located near, or appressed to, chloroplasts. Microbodies have also been found in yeast and hyphal fungi and in species representing a wide range of algal phyla. In the fat-storing seeds of higher plants they are numerous at certain stages of germination and are found in the cells in close association with spherosomes which are large lipid storage bodies. In the roots of higher plants a catalase-containing microbody has been found in association with the endoplasmic reticulum but since it does not contain either glycollate oxidase or enzymes of the glyoxyllate cycle, its function in the cell is not clear.

6.3—
Isolation

Microbodies appear to be very fragile, since homogenization of plant tissue damages them to such an extent that their constituent enzymes appear in soluble fractions after removal of chloroplasts and mitochondria by centrifugation. More gentle techniques of tissue breakage however allow microbodies to be isolated at a yield of usually about 10% of the organelles present in the whole tissue as judged by the solubilization of enzymes presumed to be present in the microbody. The method of breakage depends upon the tissue. Leaves are ground for a brief time with a mortar and pestle at 0–4°C in a buffer containing an osmoticum such as sucrose at a concentration of 0.4 to 0.8 M . Other tissues may be chopped or finely sliced with razor blades in a similar medium. The resulting homogenate is squeezed through cheesecloth and the brei centrifuged at a low speed to remove cell debris. The supernatant is then centrifuged at 6,000 to 10,000 g to obtain a pellet containing broken chloroplasts, microbodies and some mitochondria.

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Microbodies are separated from the other organelles by layering this resuspended pellet fraction on a discontinuous or continuous density gradient of sucrose ranging in concentration from 1.3 to 2.5 M , and centrifuging in an ultracentrifuge for 3–4 hr at 40,000 g. Microbodies increase in density during this process by a loss of water and ultimately form a band in the gradient at a density of 1.24 to 1.26 g cm^–3 . This density is higher than that of other organelles and they are clearly separated from mitochondria which sediment at a desntiy of 1.16 to 1.19 g cm^–3 (Fig. 6.2). On the separation of the gradient into fractions

Figure 6.2
The distribution of protein in continuous (A) and
discontinuous (B) sucrose density gradients after
centrifugation of crude particles from castor bean endosperm.
(Reproduced with permission from Cooper & Beevers, 1969a.)

the microbodies are detected by assaying for specific 'marker' enzymes. Typical marker enzymes for peroxisomes are catalase, glycollate oxidase and hydroxypyruvate reductase, while those for glyoxysomes are catalse, ma late synthetase and isocitrate lyase. Mtitochondria are characterized by the presence of cytochrome c or succinic dehydrogenase, and chloroplasts by their chlorophyll pigments.

Much of the activity of the microbody marker enzymes is found in the soluble non-particulate fraction of gradients, indicating that the yield of micro-bodies is low. Despite these low yields, sufficient amounts of intact microbodies have been isolated from several plant tissues to be able to determine their metabolic functions. This has resulted in the distinguishing of at least two types of microbody, peroxisomes and glyoxysomes, with distinctly different enzyme complements and physiological functions.

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6.4—
Glyoxysomes

There are many plant species in which lipid is the main storage material in the cotyledons or endosperm of the seed. During the first few days of seed germination there is a dramatic decrease in the lipid content of the seed and sugars, principally sucrose are formed. These sugars are subsequently translocated to the growing embryo or embryonic axis. This lipid to carbohydrate conversion has been correlated with an increase in the activity of the glyoxysomal enzymes, malate synthetase and isocitrate lyase, and as germination proceeds and the lipid reserves are depleted, the number of lipid storage bodies or spherosomes decrease and there is a drop in the activities of malate synthetase and isocitrate lyase. The elucidation of the metabolic processes involved in this process of gluconeogenesis, ultimately led to the isolation of particles in which were localized the crucial enzymes of this pathway. These particles were found to be morphologically similar to animal peroxisomes and were called glyoxysomes.

The breakdown of lipids is initiated by their hydrolysis to fatty acids. Triglycerides are hydrolized to glycerol and fatty acids by the enzyme lipase while phospholipids are hydrolized by phospholipase. The resultant long-chain fatty acids are subsequently degraded by the successive removal of 2-carbon fragments in the process of b -oxidation.

6.4.1—
b -Oxidation

In the process of b -oxidation the removal of each 2-carbon fragment from a long chain fatty acid involves a succession of five reactions (Fig. 6.3). The sequence is initiated by the activation of the substrate by coenzyme-A, catalyzed by the enzyme fatty acid thiokinase in the presence of ATP. The fatty acyl-CoA is then oxidized by the removal of hydrogen from carbons 2 and 3 of the chain and a double bond between 2 and 3 is formed. In this reaction the hydrogen is transferred to FAD. This reacts with molecular oxygen to produce peroxide which is broken down by catalase, and results in the uptake of one mole of oxygen for every two moles of fatty acyl-CoA oxidized. The unsaturated acyl-CoA produced is hydrated to form 3-hydroxy acyl-CoA, the reaction being catalysed by enoyl hydratase or crotonase, and this product then oxidized, with a concomitant reduction of NAD⁺ , to form a 3-keto acyl-CoA by the action of hydroacyl-CoA dehydrogenase. In the final reaction the 3-keto acyl-CoA is cleaved by the enzyme thiolase into acetyl-CoA and a new fatty acyl-CoA. The fatty acyl-CoA re-enters the reaction sequence and successive acetyl-CoA units are generated.

Fatty acids with an even number of carbons yield only acetyl-CoA units but those with an odd number of carbons result in the formation of acetyl-CoA and propionyl-CoA. In plant tissues propionyl-CoA is degraded by a modified b -oxidation sequence to yield acetyl-CoA and carbon dioxide.

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Figure 6.3
The b -oxidation pathway.

The b -oxidation pathway was unequivocally demonstrated to occur in plant tissues by Stumpf and Barber (1956) who showed that long chain aliphatic acids were oxidized to carbon dioxide by mitochondrial preparations from germinating peanut cotyledons, when these preparations were supplemented with a number of cofactors including ATP, CoA, and NAD⁺ . The rate of release of ¹⁴ CO₂ from specifiically labelled butyric and palmitic acids was consistent with their degradation by b -oxidation and subsequent oxidation by the TCA cycle.

6.4.2—
The Glyoxyllate Cycle

The acetyl-CoA derived from fatty acid breakdown in germinating seeds could be consumed by the TCA cycle, as indicated by the experiments of Stumpf and Barber (1956). However this would not result in the net formation of a glucose precursor since for each molecule of acetyl-CoA consumed two molecules of carbon dioxide would be produced. Furthermore, it was known at the time that little of the fatty acid was oxidized to CO₂ but instead contributed to a net synthesis of sugars.

The problem of how acetyl-CoA was converted to a glucose precursor was solved by the discovery of the glyoxyllate cycle, by Kornberg and Krebs in 1957. This cycle represents a modification of the tricarboxylic acid cycle in which two molecules of acetyl-CoA are consumed and a molecule of succinic acid is formed (Fig. 6.4). Five enzymes are involved in this process three of which,

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Figure 6.4
The glyoxyllate cycle.

citrate synthetase, aconitase, and malic dehydrogenase are components of the TCA cycle. The remaining two enzymes, the key enzymes of the glyoxyllate cycle, are isocitric lyase (isocitratase) and malate synthetase. The first reaction of the glyoxyllate cycle, catalyzed by citrate synthetase, is the condensation of oxaloacetate and acetyl-CoA to form citrate, which is then converted to isocitrate by the action of aconitase. The next reaction, unique to this cycle, is the cleavage of isocitrate into succinate and glyoxyllate catalyzed by isocitrate lyase. One of the products of this reaction, glyoxyllate, is then condensed with a second molecule of acetyl-CoA under the catalytic action of malate synthetase, to produce one molecule of malate. Malate is then oxidized by malate dehydrogenase to oxaloacetate with the concomitant reduction of NAD⁺ . The overall equation for the cycle is therefore:

Succinate produced by the glyoxyllate cycle can then be converted to hexose by conversion to oxaloacetate, by the action of succinic dehydrogenase and fumarase. The oxaloacetate is converted to phosphoenolpyruvate, a reaction catalised by phosphoenolpyruvate carboxykinase,

and the phosphoenolpyruvate converted to glucose by a reversal of the reactions of the Embden-Meyerhof-Parnas pathway. Thus four molecules of acetyl-CoA will give rise to two molecules of succinate and this in turn will result in the

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formation of one molecule of glucose and the loss of two molecules of carbon dioxide (Fig. 6.5).

Figure 6.5
The pathway of incorporation of [¹⁴ C] from [1-¹⁴ C]-acetate (O) or
[2-¹⁴ C]-acetate (

) into glucose during gluconeogenesis in plant tissue.

The glyoxyllate cycle was first demonstrated in the bacterium Pseudomonas (Kornberg & Krebs, 1957) grown on two-carbon compounds, and has since been shown to operate in many micro-organisms and plant tissues. The evidence for the cycle is based on the presence of the two key enzymes, malate synthetase and isocitrate lyase, and on the distribution of ¹⁴ C in organic acids, sugars and carbon dioxide when the tissue is supplied with specifically labelled [¹⁴ C] acetate. Malate synthetase and isocitrate lyase are found in a wide variety of plant tissues (Carpenter & Beevers, 1958), particularly in fatty seedlings where they increase in activity during germination. Similarly the activities of aconitase and citrate synthetase increase in these tissues during germination. Incubation of castor bean endosperm tissue with [I-¹⁴ C] acetate or [2–¹⁴ C] acetate results initially in the formation of [¹⁴ C] malate, and subsequently the radioactivity from [I-¹⁴ C] acetate results in about an equal labelling of CO₂ and sucrose. In the cotyledons of germinating peanut and sunflower seedlings (Bradbeer & Stumpf, 1959) and castor bean endosperm (Canvin & Beevers, 1961), [1-¹⁴ C] acetate was converted to carboxyl-labelled malate and to sucrose in which the glucose moiety was labeled in the 3 and 4 carbons, while [2-¹⁴ ]C acetate gave rise to malate labelled in the methylene carbons and to sucrose where the glucose moiety was labelled in the 1, 2, 5 and 6 carbons. These patterns or labelling of the products of acetate metabolism are consistent with the operation of a glyoxyllate cycle in these tissues (Fig. 6.5).

6.4.3—
Metabolic Functions of the Glyoxysome

The reactions of the glyoxyllate cycle and the pathway of b -oxidation were generally thought to be associated with the mitochondria since the enzymes of these pathways were usually present in the mitochondrial fraction isolated from cell homogenates. Elegant studies by Beevers' group at Purdue University however showed that sucrose density centrifugation of crude particulate fractions of castor bean endosperm resulted in the separation of three distinct bands of

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particles which were identified as proplastids, mitochondria and a new particle sedimenting at a high density. Enzymes of the glyoxyllate cycle, isocitrate lyase and malate synthetase were found exclusively in this dense particle together with catalase and a large proportion of the glycollate oxidase of the gradient. On the other hand citrate synthetase and malate dehydrogenase were associated with both the mitochondrial band and the band containing the new particle, while succinic dehydrogenase, fumarase and NADH oxidase were located exclusively in the mitochondrial band with cytochrome oxidase (Fig. 6.6).

Figure 6.6
The distribution of protein, fumarase and isocitric lyase
after sucrose density gradient separation of the components of
a crude particulate fraction of the endosperm of germinating castor bean.
(Reproduced with permission from Breidenbach & Beevers, 1967.)

The enzyme distribution indicated that the TCA cycle enzymes were located in the mitochondria while the enzymes of the glyoxyllate cycle were compartmentalized in the denser particle. These particles were therefore called glyoxysomes (Breidenbach & Beevers, 1967; Breidenbach et al., 1968). The isolated glyoxysomes were found to be organelles with a single unit membrane bounding a finely granular matrix. Similar structures were recognized in electron micrographs of intact castor bean endosperm tissue indicating that the isolated organelles were not artefacts of the isolation and centrifugation processes. Since the finding of glyoxysomes in endosperm tissue, they have been reported to be the site of the glyoxyllate cycle in the cotyledons of a number of fatstoring seeds including those of watermelon, sunflower, peanut and cucumber. Microbodies containing catalase and enzymes of the glyoxyllate cycle have also been found in yeast (Szabo & Avers, 1969) and although these have been called peroxisomes they clearly have the enzyme complements of glyoxysomes.

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In addition to glyoxyllate cycle activity, the glyoxysomes were shown to be the site of b -oxidation in castor bean endosperm (Cooper & Beevers, 1969b; Hutton & Stumpf, 1969). Glyoxysomes isolated from this tissue oxidized palmitoyl-CoA to acetyl-CoA with a concomitant reduction of NAD⁺ and uptake of oxygen. Addition of [¹⁴ C] oxaloacetate during this reaction resulted in the formation of [¹⁴ C] citrate and [¹⁴ C] malate from palmityl-CoA indicating that the acetyl-CoA produced by the b -oxidation process was consumed in the glyoxyllate cycle also located in the organelle (Cooper & Beevers, 1969b). Similarly, ricinoleate, linoleate and palmitate were oxidized by glyoxysomes of castor bean with the formation of acetyl-CoA, and three enzymes of the b -oxidation complex, crotonase, b -ketothiolase, and b -hydroxyacyl dehydrogenase were found to be located specifically in the organelle (Hutton & Stumpf, 1969). The activity of the b -oxidation complex in this tissue during the germination of the seed was found to parallel the increase in activity of the glyoxyllate cycle enzymes indicating that an integrated system for lipid utilization develops together with the formation of the glyoxysome.

The principal pathways of gluconeogenesis are therefore compartmentalized in a single organelle, the glyoxysome. The complete pathway of gluconeogenesis however involves at least three organelles, the spherosome, the glyoxysome and the mitochondrion. The reactions are initiated by a hydrolysis of lipids in the spherosome by the action of lipase, and the glycerol and fatty acids produced diffuse out of the organelle. Glycerol is utilized directly by the EMP pathway in the cytosol and contributes to sucrose synthesis (Beevers, 1956) while the fatty acids diffuse into glyoxysomes which are located near the spherosome. In the glyoxysome the fatty acid is oxidized in the b -oxidation pathway and the acetyl-CoA released is converted to succinate by the action of the glyoxyllate cycle enzymes located in the organelle. Although the b -oxidation pathway is reversible, the equilibrium is presumably maintained in a catabolic direction by the removal of peroxide, produced in the oxidation step of the pathway, by the catalase present in the glyoxysome. Reduced NAD⁺ produced in these reactions is probably oxidized in the mitochondria. Succinate produced in the glyoxysome is further metabolized to oxalacetate and finally to phosphoenolpyruvate in the mitochondria since succinic dehydrogenase and fumarate are not component enzymes of the glyoxyllate cycle and are absent from glyoxysomes.

This compartmentalization of the b -oxidation complex and the glyoxyllate cycle together in an organelle discrete from the mitochondrion is probably the reason why acetyl-CoA is utilized in gluconeogenesis in plant tissues rather than being oxidized to carbon dioxide and water as in animal tissues. Free acetyl-CoA-is presumably not released from the glyoxysome and made available for oxidation in the mitochondrion, while consumption of succinate, the final product of the glyoxyllate cycle, by the mitochondrion, would not stimulate the rate of oxidation in the TCA cycle since this can only occur by increasing the supply of acetyl-CoA. Some measure of control of competing metabolic pathways is therefore achieved by separation of these reactions within different organelles.

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6.5—
Peroxisomes

Glycollic acid is produced in large amounts in the chloroplast, as a by-product of the reactions of carbon dioxide fixation. The formation of glycollate has been proposed to occur by the oxidation of ribulose-1,5-bisphosphate (RBP) by molecular oxygen, a reaction which would produce a two carbon fragment of phosphoglycollic acid and a three carbon fragment of phospholgyceric acid (PGA) instead of two molecules of PGA resulting from the normal carboxylation of RBP by CO₂ . It has been found that RBP carboxylase acts as an oxygenase in the presence of molecular oxygen and that phosphoglycollate and PGA are produced in this reaction in vitro (Andrews et al., 1973). Thus the enzyme RBP carboxylase can act as an oxygenase or as a carboxylase and the formation of phosphoglycollic acid is favoured by high partial pressures of oxygen. Glycollate is produced from phosphoglycollate by the action of phosphoglycollate phosphatase, a chloroplast enzyme. Glycollate is also produced, in vitro, from fructose-6-phosphate by the action of the chloroplast enzyme, transketolase, which may be due to the oxidation, by hydrogen peroxide, of the glycolaldehyde-thiamine pyrophosphate, an intermediate complex in this enzyme reaction (Bradbeer & Racker, 1961). Glycollate is released from the chloroplast into the cytosol, where it is further metabolized in a specific metabolic pathway, the initial reactions of this pathway being located in the peroxisome.

6.5.1—
The Glycollate Pathway

The pathway of glycollate metabolism in leaves has been elucidated by infiltrating excised leaves or leaf discs with ¹⁴ C-labelled glycollate or other intermediates of the pathway. Glycollate is converted rapidly to glyoxyllate which may be oxidized non-enzymatically to carbon dioxide and formic acid, in the presence of hydrogen peroxide. In this reaction the carbon dioxide is derived from the carboxyl carbon of glycollate and the formate from the methyl carbon (Tolbert & Burris, 1950). However the presence of catalase, which breaks down peroxide, is thought to preclude such a total degradation of glyoxyllate in the peroxisome and the supply of ¹⁴ C-labelled glycollate or glycoxyllate to leaf tissue in the light has been found to give rise initially to labelled glycine and serine and subsequently to labelled glyceric acid hexoses and sucrose (Tolbert, 1963; see Fig. 6.7).

Infiltration of [2-¹⁴ C] glycollate into leaves in the light was found to give [2-¹⁴ C] glycine but serine was found to be labelled in the 2 and 3 carbons (Tolbert & Cohan, 1953; see Fig. 6.7). From this evidence it was concluded that two molecules of glycine give rise to one molecule of serine with a loss of one molecule of carbon dioxide; the 1 and 2 carbons of serine are derived from carbons 1 and 2 of glycine respectively, while the 3 carbon of serine is derived from the 2 carbon of glycine and carbon dioxide arises from carbon 1 of glycine. In wheat leaves in light [3-¹⁴ C] serine was converted to [3-¹⁴ C] glycerate

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Figure 6.7
The glycollate pathway. The labelling pattern of intermediates of the
pathway ane hexose is indicated for when they are derived from
[1-'⁴ C]-glycollate (

) or [2-'⁴ C]-glycollate (

and this was incorporated into hexose presumably by the reactions of the Embden-Meyerhof-Parras pathway (Rabson et al., 1962). This pathway of hexose formation from glycollate has been confirmed by the finding that [2-¹⁴ C] glycollate gives rise to glycose labelled in the 1, 2, 5 and 6 carbons and [3-¹⁴ C] serine to glucose labelled in the 1 and 6 carbons (Jiminez et al., 1.962).

The glucollate pathway is therefore gluconeogenic in light. The conversion of glyceric acid derived from glycollate, to glucose and sucrose is inhibited by DCMU an inhibitor of Photosystem II (Miflin et al., 1966). In the dark, supplied [¹⁴ C]-glycollate is converted into TCA cycle acids rather than sugars which may be due to an oxidation of pyruvate derived from glyceric acid, or may result from a direct conversion of glyoxyllate to malate.

Glycollate produced during photosynthesis in the presence of ¹⁴ CO₂ is usually found to be uniformly labelled i.e. both carbons have the same specific activity. This distribution of radioactivity would be expected if the glycollate was derived from carbons 1 and 2 of ribulose-1,5-bisphosphate. Consequently all the compounds arising from glycollate are uniformly labelled. Serine, in particular, has been found to be uniformly labelled while phosphoglyceric acid was predominately carboxyl labelled, indicating that serine was produced from glycollate rather than directly from phosphoglyceric acid formed in CO₂ -fixation (Rabson et al., 1962). The uniformly labelled glyceric acid formed in this pathway in turn produces uniformly labelled hexoses instead of the 3,4-¹⁴ C-hexoses resulting from incorporation of ¹⁴ C from the photosynthetic carbon cycle.

The operation of the glycollate pathway in leaves has been shown by tracer experiments but it has been confirmed by the detection of enzymes in leaves which are necessary for some reactions of the pathway. The initial reaction, the oxidation of glycollate to glyoxyllate is catalysed by glycollate oxidase, an enzyme first isolated by Zelitch and Ochoa (1953). It has FMN as the prosthetic group and utilizes molecular oxygen as the electron acceptor. The enzyme is competitively inhibited by bisulphite addition compounds of aldehydes, a -hydroxy-sulphonates, which have the general structure R-CHOH-SO₃ H and

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are therefore structural analogues of glycollate. The most commonly used a -hydroxysulphonate is a -hydroxypyridylmethane sulphonate (HPMS) and treatment of leaf discs or infiltration of excised leaves with this compound results in the accumulation of glycollic acid while having no effect upon the rate of CO₂ -fixation. Experiments of this type have shown that 40 to 70: of the carbon fixed in photosynthesis will accumulate as glycollate in tissues treated with HPMS and these results have been interpreted as indicating that a large fraction of the carbon fixed in photosynthesis is metabolized by this pathway.

Transaminases are present in leaves which catalyse two steps of the pathway: a glutamate-glyoxyllate transaminase catalysing the formation of glycine is widespread in plant tissues, and a serine-pyruvate aminotransferase catalysing the formation of serine to hydroxypyruvate is found in leaves. The conversion of glycine to serine is catalysed by serine hydroxymethyltransferase which has been found in pea and wheat leaves (Cossins & Sinha, 1966). This reaction is inhibited by isonicotinyl hydrazide, and treatment of leaf tissue with this compound during photosynthesis in ¹⁴ CO₂ causes an accumulation of [¹⁴ C] glycine and [¹⁴ C] glycollate and a decrease in the incorporation of ¹⁴ C into glucose (Miflin et al., 1966). Use of this inhibitor thus provides additional evidence for the operation of the pathway.

Two types of glyoxyllate reductase have been found to occur in leaves. One, an NADP-linked enzyme, is thought to be located in the chloroplast, while a second NAD-linked enzyme is present in the cytoplasm and is referred to as hydroxypyruvate reductase since the enzyme isolated from some sources has a higher activity to hydroxypyruvate than to glyoxyllate. The enzyme appears to catalyse the reduction of hydroxypyruvate to glycerate in the glycollate pathway.

6.5.2—
Metabolic Reactions of the Peroxisome

It was generally accepted that the enzymes of the glycollate pathway were soluble proteins of the cytosol and the failure of several attempts to localize these enzymes, particularly glycollate oxidase, in discrete organelles confirmed this idea. The first successful localization of enzymes of glycollate metabolism in a discrete organelle was achieved by Tolbert and coworkers at Michigan State University (Tolbert et al., 1968). They demonstrated that sucrose density gradient centrifugation of spinach homogenates separated three bands of particles: broken chloroplasts, mitochondria, and small bodies distinctly separated from, and denser than the other organelles. Electron microscopic examination of these bodies showed them to be organelles bounded by a single unit membrane and since they closely resembled peroxisomes from animal cells in size and morphology, they were referred to as leaf peroxisomes. The most important finding was that these peroxisomes contained the bulk of the activity of the glycollate oxidase, catalase and hydroxypyruvate reductase (NAD-glyoxyllate reductase) of the gradient, while cytochrome c oxidase was specifically located in the mitochondrial fraction. Peroxisomes were later

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isolated from the leaves of nine other plant species including tobacco, maize, and sugarcane, and all contained the same enzyme complement (Tolbert et al., 1969). These studies therefore demonstrated that processes of glycollate oxidation and peroxide breakdown were localized together in an organelle discrete from the chloroplast. Electron microscope studies, particularly by Newcomb, have since revealed the presence of peroxisomes in leaves of many plant species (e.g. Frederick & Newcomb, 1969).

Further studies in Tolbert's laboratory have shown that other enzymes of the glycollate pathway are also localized in the peroxisome. Two aminotransferases, glutamate-glyoxyllate aminotransferase catalysing the conversion of glyoxyllate to glycine, and serine-pyruvate aminotransferase catalysing the conversion of serine to hydroxypyruvate, were found in peroxisomes isolated from leaves of various species. Serine hydroxymethyltransferase is the only enzyme of the glycollate pathway not found in the peroxisome, and is probably located in the mitochondrion. Supply of [¹⁴ C] glycollate and [¹⁴ C] glyoxyllate to isolated peroxisomes gave rise only to [¹⁴ C] glycine and while oxygen uptake occurred, no ¹⁴ CO₂ release was detected (Kisaki & Tolbert, 1969).

The glycollate pathway appears to require enzymic steps located in three subcellular organelles (Fig. 6.8). Glycollate, formed in the chloroplast is

Figure 6.8
The distribution of the reactions of the glycollate
pathway among organelles of the leaf cell.

oxidized to glyoxyllate in the peroxisome. The glyoxyllate may then be exported to the chloroplast where it could be reduced to glycollate by the action of NADP-dependent glyoxyllate reductase which is located specifically in the chloroplast (Tolbert et al., 1970). Such a coupling of alternate oxidation and reduction reactions. a 'glycollate-glyoxyllate shuttle', has been proposed as a mechanism for controlling the levels of reduced NADP in the chloroplast, but no unequivocal evidence for such a reaction in vivo has been found. Glycine is formed from glyoxyllate in the peroxisome and then transferred to the

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mitochondrion where it is converted to serine, with a concomitant loss of carbon dioxide. The conversion of serine to glycerate can then be accomplished by enzymes localized in the peroxisome. Further metabolism of glycerate to hexose appears to be confined to the chloroplast since these reactions would be initiated by the formation of phosphoglyceric acid, a reaction catalysed by phosphoglycerate phosphatase which is located in the chloroplast.

The metabolism of glycollate by leaf tissue results in the release of carbon dioxide, but the exact site of this CO₂ release in the cell is still a controversial question. It has been proposed that CO₂ is evolved by the mitochondria during the conversion of glycine to serine, since [¹⁴ C] glycine is as good a precursor as [¹⁴ C] glycollate for ¹⁴ CO₂ evolution in leaves and the ¹⁴ CO₂ evolved is derived from the carboxyl groups of these compounds. The presence of catalase in the peroxisomes is thought to minimize the non-enzymatic oxidation of glyoxyllate by H₂ O₂ to formate and carbon dioxide and no evolution of CO₂ by isolated peroxisomes from glycollate or glyoxyllate could be detected by some workers (Kisaki & Tolbert, 1970). This loss of CO₂ from glycine, which amounts to only 25% of the total carbon passing through the glycollate pathway, would not account for the large losses of CO₂ which occur as a result of the photorespiration of glycollate in leaves. However the amount of catalase present in the peroxisome may not preclude the non-enzymatic oxidation of glyoxyllate and it has been shown that both [¹⁴ C] glycollate and [1-¹⁴ C] glyoxyllate can be decarboxylated by peroxisomal fractions at pH 8.0 (Halliwell & Butt, 1974). An enzyme is also present in chloroplasts which catalyses the decarboxylation of glyoxyllate to formic acid and CO₂ (Zelitch, 1972). Present evidence indicates therefore that three subcellular organelles have the capacity to decarboxylate components of the glycollate pathway and each may contribute to the production of the carbon dioxide in photorespiration (Fig. 6.8).

6.5.3—
Photorespiration

The oxidation of glycollate in the peroxisome is accompanied by a consumption of oxygen and ultimately results in the release of carbon dioxide. The net result is a respiratory gas exchange where the substrate of respiration is glycollate rather than glucose. Since glycollate is only formed in light this respiration is light-dependent and is called photorespiration.

The direct measurement of oxygen uptake during a net photosynthetic release of oxygen, or CO₂ release during net photosynthetic CO₂ fixation is impossible. However, indirect methods have demonstrated that photorespiration occurs in plants and that the rate of CO₂ loss in light is higher than that in the dark. Measurements of photorespiration rates vary considerably with the assay method used and can only be considered as approximations of the magnitude of the process. Nevertheless, it is becoming clear that the process of photorespiration has great importance in decreasing the rate of net photosynthesis in many plants.

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Photorespiration can be detected by measuring the flux, that is the simultaneous uptake and release, of oxygen or carbon dioxide of photosynthesizing tissue by using isotopic tracer methods. The uptake of ¹⁸ O₂ by leaves during photosynthesis has been detected but experiments using ¹⁸ O₂ have generally given equivocal results which have been difficult to interpret.

The measurement of carbon dioxide flux during photosynthesis is much more convenient and depends upon the accurate measurement of carbon dioxide concentration in the atmosphere using an infra-red gas analyser and a simultaneous measurement of the total activity of supplied ¹⁴ CO₂ with an ion counter or Geiger-Müller counter. When a plant is placed in a closed system in light, there is a rapid uptake of carbon dioxide and the CO₂ concentration of the atmosphere around the plant decreases to a concentration at which the uptake of CO₂ exactly balances the output of CO₂ by the plant (Fig. 6.9). This concentration of carbon dioxide is called the CO₂compensation point of the plant and is usually measured in parts per million (ppm) of CO₂ in air. Plants such as tobacco, sunflower and wheat have compensation points, ranging from 35 to 100 ppm indicating a marked loss of CO₂ (i.e. photorespiration) during photosynthesis, but others such as maize and sugarcane have compensation points of 3 to 10 ppm indicating a low loss of CO₂ or a low photorespiration rate. If photosynthetic carbon fixation is similarly measured in a closed-system but in an atmosphere containing ¹⁴ CO₂ , the CO₂ arising from, the plant by photorespiration will, over short time periods, be ¹² CO₂ . Thus, while a decrease in the CO₂concentration of the atmosphere around the plant will occur, the radioactivity of ¹⁴ CO₂ in the atmosphere will appear to decrease at a faster rate because of the efflux of unlabelled CO₂ from the plant, and continues to decrease even after the compensation point is reached (Fig. 6.9a). In a plant which

Figure 6.9
The concentration of CO₂ and ¹⁴ CO₂ and the specific activity of ¹⁴ CO₂ around
a detached sunflower leaf (a) and a detached maize leaf (b) during illumination
and in subsequent darkness in an atmosphere of 21% oxygen and at 21°.
(Reproduced with permission from Hew et al., 1969.)

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is photorespiring, therefore, a decrease in the specific radioactivity of ¹⁴ CO₂ will occur, while in a plant with no photorespiration the uptake of ¹⁴ CO₂ will not be accompanied by an efflux of ¹² CO₂ and the specific radioactivity of the ¹⁴ CO₂ in the atmosphere will remain constant (Fig. 6.9b). In the dark the specific radioactivity of the ¹⁴ CO₂ decreases with the efflux of ¹² CO₂ and the rate of decrease of this activity is a measure of dark respiration. The rate of photorespiration as measured by these methods has been shown to be 1.5 to 2.5 times that of dark respiration, while in plants with low photorespiration, such as maize, loss of CO₂ in light is only a fraction of the rate of CO₂ loss in the dark. This method clearly indicates that photorespiration occurs in some species while it is absent in others.

Another method which has been used to detect differences in the rate of photorespiration and dark respiration was devised by Zelitch (1968). In this method leaf discs are allowed to fix ¹⁴ CO₂ for a period of 45 to 60 minutes. The remaining ¹⁴ CO₂ is then quickly flushed out of the closed system, and the release of ¹⁴ CO₂ from the tissue into CO₂ -free air is measured over short periods in the light and the dark. The method is based on the assumptions that the rate of ¹⁴ CO₂ loss is a measure of total CO₂ loss i.e. that the specific radioactivity of the CO₂ evolved remains constant over short time periods and that low CO₂ tensions have no effect on the loss of CO₂ . These assumptions may not be valid for all photosynthetic tissues but within these limitations the method is a very rapid and sensitive means for detecting photorespiration. Photorespiratory loss of CO₂ in tobacco has been shown by this method to be 2 to 5 times higher than in the dark while CO₂ loss from maize is not detectable (Fig. 6.10a). This method has been similarly used to detect photorespiration in other species.

Figure 6.10a
A comparison between the release of ¹⁴ CO₂ by tobacco and maize discs
in the light and the dark after previously being supplied ¹⁴ CO₂ in light.
(b). The effect of a -hydroxysulponate on the release of
¹⁴ CO₂ from tobacco leaf discs in the light and dark.
(Reproduced with permission from Zelitch, 1968.)

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Photorespiration is not simply a stimulation of dark respiration since the two process of photorespiration and dark respiration respond differently to changes in oxygen concentration. Dark respiration of leaves has an optimal rate at about 2% oxygen in air and any increase in the O₂ concentration up to 100% does not increase the rate (Fig. 6.11). Net photosynthesis however is inhibited

Figure 6.11
The effect of oxygen concentration on the rate of photorespiration
(PR) and dark respiration (R_D ) of detached soybean leaves.
(Reproduced with permission from Forrester et al., 1966.)

by oxygen, a phenomenon called the Warburg effect, and this is attributable to an increase in the rate of photorespiration rather than to an inhibition of photosynthesis per se . A reduction in the oxygen concentration in the atmosphere around a leaf from the ambient 21% O₂ of air has been shown to lower the compensation point and increase the rate of net photosynthesis, while an increase in concentration raises the compensation point and lowers net photosynthesis (Forrester et al., 1966). Plants with photorespiration have been found to evolve CO₂ at a high rate when they are transferred to darkness after a period of photosynthesis. This post-illumination CO₂ burst lasts for a few minutes before a steady dark respiration is established and the magnitude of the burst has been found to depend on the light intensity and oxygen concentration during the previous period of photosynthesis: the CO₂ -burst increases with an increase in light intensity and decreases with a decrease in O₂ concentration (Tregunna et al., 1961). This burst has been explained as being due to the continued slow oxidation, in the dark, of a product produced in photosynthesis after the uptake of CO₂ by photosynthesis has stopped. Photorespiration therefore appears to have different characteristics than dark respiration and to have a different substrate.

The substrate for photorespiration is thought to be glycollic acid. Glycollic

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acid production in leaves and in chloroplasts is stimulated by an increase in oxygen concentration as is photorespiration, while the inhibition of glycollate oxidation by HPMS has been found to lower the rate of photorespiration in leaf discs to that of dark respiration (Fig. 6.10b). Photorespiratory loss of CO₂ is also stimulated by the infiltration of leaves with glycollic acid while acetate has no effect on this rate. Thus the oxidation of glycollate, mediated by the leaf peroxisomes results in a photorespiratory loss of carbon dioxide by the leaf.

Plants with low compensation points are said to lack photorespiration. These plants are C₄ plants, that is, the primary carboxylation step is catalysed by phosphoenolpyruvate carboxylase rather than by RBP carboxylase. This efficient fixation of CO₂ has been postulated to preclude the formation of glycollate, but maize for example, has been shown to possess a glycollate pathway (Osmond, 1969) and peroxisomes occur in the mesophyll cells of the leaf. Presumably in such plants glycollate may be formed and oxidized, but the efficiency of refixation of the CO₂ resulting from glycollate oxidation is such that no CO₂ is released from the plant and hence no photorespiration is detected.

Estimates from various plant species suggest that from 15 to 40% of the carbon fixed in photosynthesis is lost by photorespiration. There is no unequivocal evidence to show that energy in the form of ATP is recovered during the oxidation of glycollate and the process appears to be a wasteful one in terms of energy conservation. An important question therefore arises: does photorespiration serve a useful function or is it simply an inevitable consequence of photosynthesis being carried on in an atmosphere of 21% oxygen? It has been suggested that photorespiration might act as a 'safety valve' for the plant, in that, under conditions of high light intensity and low carbon dioxide concentration, the oxidation of glycollate would consume both excess reduced NADP⁺ and excess oxygen, which would serve to protect the chloroplast from photo-oxidative damage. If the process of photorespiration imposes limitations on the growth of plants, it would be expected that there would have been some selective pressure to eliminate it by natural selection during the evolution of the higher plants. The occurrence of C₄ plants, which have low photorespiration, may represent such an evolutionary step to correct for the presence of photorespiration by the development of an efficient CO₂ -fixation mechanism. Since variations in the rate of photorespiration occur within a single species (Zelitch, 1971) it may be possible to select artificially for low photorespiration in crop plants and consequently increase net photosynthetic productivity and crop yield.

6.6—
Ontogeny and Turnover of Microbodies

The development of microbodies in plant cells is a difficult process to follow since they usually occur in small numbers and are difficult to characterize

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enzymatically. Small, membrane-bound organelles occur in close proximity to the endoplasmic reticulum but these do not always give a positive reaction to DAB, the usual criterion for the presence of catalase. Nevertheless catalase is invariably the first enzyme to be detected in microbodies and it appears to be the most active enzyme in the developing organelle. Studies of microbody development have been carried out principally on tissues having large populations of microbodies such as the endosperm and cotyledons of fatty seedlings.

In the endosperm of fat-storing seeds, particularly of the castor bean, the process of germination results in the development of large numbers of microbodies which reach numbers per cell twice that of mitochondria (Vigil, 1970). This increase in number is correlated with increases in the activities of isocitrate lyase and malate synthetase in the whole tissue. It has been suggested by several investigators that microbodies develop from the endoplasmic reticulum by a budding process. In castor bean the formation of microbodies has been linked directly to the endoplasmic reticulum and connections between the cisternae of the rough endoplasmic reticulum and developing microbodies have been clearly demonstrated (Vigil, 1970). This connection does not appear to persist during the whole course of organelle development although they increase seven-fold in size and increase in density suggesting a continuous addition of newly synthesized protein. The depletion of lipid in the endosperm or cotyledons of fatty seedlings is accompanied by a decrease in glyoxysomal enzymes which is attributable to a decrease in the number of glyoxysomes as measured by enzyme activity and total protein (Gerhardt & Beevers, 1970). It is not clear how these microbodies are destroyed but some at least become enclosed in autophagic vacuoles and are digested.

Increases in peroxisomal enzymes have been detected in germinating seedlings and etiolated seedlings on exposure to light and some of these changes have been correlated with the size and structure of microbodies. The development of microbodies in, leaves of wheat seedlings in light involves distinct changes in the biochemical and physical characteristics of the organelle. During the first few days of germination an increase occurred in the density of catalasecontaining particles from 1.17 to 1.24 g cm^–3 on separation on sucrose gradients, which was paralleled by an increase in catalase, hydroxypyruvate reductase and glycollate oxidase (Fierabend & Beevers, 1972).

In etiolated leaves small microbodies occur which are rich in catalase but low in other peroxisomal enzymes. These bodies have not been unequivocally characterized as peroxisomes because of the difficulty of isolation. Upon exposure to light there is an activation of glycollate oxidase and hydroxypyruvate reductase, but these changes in enzyme activity have not been correlated with an increase in size and number of peroxisomes. The activity of glycollate oxidase is stimulated by exposure of etiolated tissue to red light and this activation is reversed by -far-red light. These effects probably reflect a stimulation of leaf development rather than a specific phytochrome involvement in peroxisome development. However an increase in glycollate oxidase occurs under

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continuous far-red light in mustard seedlings where chloroplast development is inhibited, suggesting that substrate activation is not involved in this increase of enzyme activity. Similar increases in the activities of glycollate oxidase, and hydroxypyruvate reductase have been found in cotyledons of germinating seedlings on exposure to light (Trelease et al., 1971; Kagawa et al., 1973).

In germinating seeds and in greening leaves the microbodies have easily identifiable functions, those of glyoxysomes or peroxisomes, and only one type of microbody occurs in each tissue. However an interesting situation occurs in some seeds of the Cucurbitacea. In cucumber and watermelon for example, lipid is the main storage material in the seed and during germination there is an increase in activity of glyoxysomal enzymes in the cotyledon. During normal development, the cotyledons become exposed to light and turn green. The glyoxysomal activity of the microbodies, as measured by the activity of malate synthetase and isocitrate lyase, decreases while the peroxisomal activity, as measured by glycollate oxidase activity, increases. In one organ therefore in response to environmental stimulus there is a transition from lipid breakdown and gluconeogenesis, to glycollate metabolism, which is reflected in a change in the microbody population from a glyoxysomal to a peroxisomal function. Since the two microbodies are morphologically indistinguishable the question arises as to whether the structure of the glyoxysome is retained and there is a selective replacement of enzymes to give it a peroxisomal function or whether there is an autolysis of the glyoxysome and a replacement of it by a newly synthesized peroxisome.

The changes in microbody enzyme complements which occur during seedling development have been correlated with changes in the fine structure of the microbodies. It has been reported that in cucumber seedlings the decrease in glyoxysomal enzyme activity is not accompanied by a decrease in the number of microbodies as determined by electron microscopy. The subsequent exposure of the cotyledons to light caused a great increase in the activity of peroxisomal enzymes, particularly that of glycollate oxidase, while there was not a concomitant increase in the number of microbodies (Trelease et al., 1971). No evidence was found for the degradation of microbodies during the loss of glyoxysomal enzyme activity as has been reported in castor bean (Vigil, 1970). These studies suggest that there is a continuity of microbody structure in the transition from a glyoxysomal to a peroxisomal function. On the other hand, in developing watermelon seedlings, the decrease in the total activity of glyoxysomal enzymes in isolated microbodies which occurred after the depletion of lipid, was accompanied by a decrease in the total protein of these microbodies. This was interpreted as indicating a destruction of glyoxysomes in the tissue (Gerhardt & Beevers, 1970). Normally the peroxisomal activities of the microbody increase as glyoxysomal activity decreases, but in this tissue the decrease in glyoxysomal activity of the microbodies could be separated from the increase in the peroxisomal activity. Brief exposure of dark-grown cotyledons to light at an early stage of development stimulated the activities of peroxisomal enzymes, while

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this did not affect the rate of decline of the activity of glyoxysomal enzymes (Kagawa et al., 1973). These results suggest that the formation of peroxisomes can occur while active glyoxysomes are present and that two populations of microbody can occur in the tissue at the same time. The apparent contradiction in the results from these two tissues has still to be resolved, and it is still not clear whether the transition from a glyoxysomal function to a peroxisomal function in the microbodies of these tissues is due to enzyme replacement in a single microbody or to synthesis of a new microbody.

6.7—
Algal Microbodies

The microbodies of photosynthetic higher plants have distinct physiological roles either as peroxisomes or glyoxysomes which are active at different stages of the life cycle of the plant. In the algae however their role is not quite as clear since both functions of the microbody, i.e. glycollate oxidation and gluconeogenesis, may be carried on in a single cell. Microbody-like organelles have been found in a wide range of algae, and they are most numerous in algal cells grown on two-carbon compounds. Catalase has been detected cytochemically by DAB staining in situ but has not been shown to be universally present in algal microbodies.

Algae grown autotrophically, that is, in light on carbon dioxide, produce glycollic acid which is metabolized by a pathway similar to that of higher plant leaves (Bruin et al., 1970; Lord & Merrett, 1970). Glycollate oxidase however is not present in algae, with the possible exception of some members of the Zygnematales, Ulotrichales and Charophyceae (Fredrick et al., 1973) but the oxidation of glycollate is catalysed by glycollate dehydrogenase. This enzyme is not FMN dependent and does not couple to molecular oxygen so that hydrogen peroxide is not formed during glycollate oxidation (Nelson & Tolbert, 1970). This may to some extent account for the low activities of catalase reported in many algae. The natural electron acceptor for glycollate dehydrogenase is as yet unknown.

While the presence of a glycollate pathway has been established in green algae, the extent to which glycollate is metabolized by this pathway during photosynthesis is not known. In contrast to higher plant cells, serine formed during photosynthesis in ¹⁴ CO₂ in several algae is carboxyl labelled indicating its formation directly from PGA rather than by the glycollate pathway (Bruin et al., 1970). There is also a lower rate of CO₂ loss by photorespiration in algae than in higher plants (Cheng & Colman, 1974) suggesting perhaps that little glycollate is formed in algal photosynthesis. Glycollate is excreted by algal cells and this has been interpreted to indicate that the algae have a low rate of glycollate metabolism. This can now be discounted however since little glycollate is excreted during steady-state photosynthesis (Watt & Fogg, 1966; Colman et al., 1974) and all the glycollate formed appears to be metabolized in the algal cell.

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Many unicellular algae can be grown in the dark on two-carbon compounds such as acetate or ethanol as sources of both energy and carbon for growth. Chlorella grown in the dark on acetate, incorporates [¹⁴ C]-acetate into protein and carbohydrate without prior degradation while simultaneously about half of the acetate taken up is oxidized to CO₂ in a manner consistent with the operation of the TCA cycle (Syrrett et al., 1964). These cells have high levels of malate synthetase and isocitrate lyase, thereby allowing the formation, from acetate, of four-carbon compounds which can form precursors of amino acids and hexoses (Syrrett et al., 1963). Autotrophically-grown cells however have little isocitrate lyase activity and [¹⁴ C]-acetate supplied to these cells in the dark is oxidized to CO₂ . Furthermore the activity of isocitrate lyase increases when autotrophically-grown cells are supplied with acetate in the dark and no cell division takes place during this time. Similar increases in malate synthetase and isocitrate lyase in response to growth on two-carbon compounds have been reported in Euglena gracilis.

The enzymes of both the glycollate pathway and the glyoxyllate cycle have therefore been found in several algae and the question arises as to whether these enzymes are compartmentalized in a microbody. The limited number of studies which have been done suggest, on the basis of their enzyme activities, that microbodies isolated from algae have either a glyoxysomal function or are of the non-specialized type often present in non-green plant tissues (Huang & Beevers, 1971). In Euglena gracilis grown on ethanol the clyoxylate cycle enzymes malate synthetase and isocitrate lyase together with glycollate dehydrogenase and glyoxyllate reductase were detected in a particulate fraction distinct from mitochondria and, while no catalase activity was detected, the enzyme complement indicates a microbody with a glyoxysomal function (Graves et al., 1972). However in another flagellate, Chlorogonium elongatum, grown either on acetate in the dark or autotrophically in the light, a catalase-containing microbody was found which had neither glyoxysomal nor peroxisomal enzymes associated with it. In the acetate-grown cells isocitrate lyase and malate synthetase were found to be located in the cytosol (Stabenau & Beevers, 1974), while in autotrophically-grown cells, glycollate dehydrogenase, hydroxypyruvate reductase and glyoxyllate-glutamate aminotransferase were₂ found to be localized in a mitochondrial fraction together with cytochrome oxidase, and malate and isocitrate dehydrogenases (Stabenau, 1974).

It is apparent that algal microbodies do not have a peroxisomal function and that this is correlated with the oxidation of glycollate in these cells by glycollate dehydrogenase, thus eliminating the requirement for the compartmentalization of glycollate oxidation with catalase. However, microbodies have been isolated from only a few algae primarily because vigorous methods are required to break algal cell walls and microbody isolation is therefore difficult to achieve. It remains to be seen whether peroxisomes occur in algae such as Nitella and Spirogyra which have relatively high levels of catalase and also have glycollate oxidase rather than glycollate dehydrogenase.

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Chapter 7—
Microtubules

7.1—
Introduction

Microtubules were first described in spermatozoids of the moss, Sphagnum, by Irene Manton in 1957. Since the introduction, six years later, of glutaraldehyde as a fixative for electron microscopy, their presence has been revealed in a very wide variety of plant and animal cells. They have not been observed in prokaryotes.

These organelles are associated with several processes in the repertoire of movements exhibited by the eukaryotic cell, including; movements of individual cells by flagellar motion or axopod extension; movement of components within cells, as in mitosis, axoplasmic flow or transport of certain pigment granules; and morphogenetic movements involving the generation and maintenance of cell shape. In the case of plant cells, in addition to chromosome movements during cell division, the presence of microtubules has been correlated with definition of the plane of cell cleavage, formation of the cell plate and determination of cell wall architecture.

In certain cell types, microtubules are seen only during mitosis and meiosis, suggesting that they were involved originally in these processes alone, and only later did they come to be used for extranuclear functions in advanced eukaryotic organisms. Indeed, it has been suggested (Margulis, 1970) that acquisition of the microtubule was a major factor in permitting the evolution of the eukaryote cell.

7.1.1—
Description

Under the electron microscope, microtubules appear as long, unbranched cylindrical structures with a diameter of 22–25 nm (Fig. 7.1 and Fig. 7.4). In transection, they show an electron-lucent core, ca. 12 nm in diameter, and a wall of, usually, 13 electron-dense subunits. The subunits are 4–5 nm wide and are organized along the long axis of the microtubule into protofilaments. There is an axial displacement of subunits between adjacent protofilaments, resulting in a visible pitch relative to the long axis of the tubule (Fig 7.2).

Microtubules are usually separated from each other or from adjacent cellular components by a clear space of 10–40 nm. Also, in longitudinal view, an electron-lucent zone is often observed along the sides of the microtubule. These observations suggest that each tubule may be surrounded by a specialized region or layer of material. In certain systems, microtubules show 'arms' projecting from the walls of the tubules at regular intervals along their length.

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Figure 7.1
Negatively stained microtubules prepared from brain, showing axially aligned
protofilaments and their substructural periodicity. Scale marker 0.1 m m.

Figure 7.2
Model microtubule, showing in transection the 13 globular
subunits comprising the wall, and in longitudinal view, the
axial displacement of subunits in adjacent protofilaments. From
Bryan (1974) Fedn. Proc. Fedn. Am. Socs. exp. Biol. 33, 152–57,
reproduced by permission of the author and Fedn. Am. Socs. exp. Biol.

Such lateral projections act, in some instances, as cross-bridges between adjacent microtubules. Alternatively, projections can extend from microtubules to adjacent membranes such as the plasmalemma, the nuclear envelope, the endoplasmic reticulum or associated vesicles.

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7.1.2—
Background

Eukaryote mitotic spindles are a heterogeneous group of intracellular structures characterized by the presence of anisotropically arranged microtubules. The number of microtubules within the spindle of a given species appears to reflect the mass of the chromosomes, and can vary from a few to several thousand. The highly oriented microtubules are responsible for the weak form-birefringence of the spindle under polarized light and there are natural fluctuations in spindle birefringence during the different phases of mitosis.

Mitotic arrest can be achieved with a variety of physical and chemical agents. Spindle birefringence is also affected by such agents. For example, birefringence in dividing Lilium pollen mother cells is abolished within seconds of exposure to low temperature. Upon return to normal temperatures, spindle birefringence reappears in a few minutes, after which mitosis proceeds normally. Exposure to high hydrostatic pressure causes a similar effect. Chemical agents with antimitotic activity, such as the alkaloids colchicine, podophyllotoxin or vinblastine, also abolish spindle birefringence. The important feature of all these effects is that they are reversible upon removal of the inhibitory agent.

Based on: (a) natural fluctuations in spindle birefringence, (b) reversible effects of inhibitors on both birefringence and mitosis and (c) the findings of many earlier studies that the major components of the spindle are synthesized prior to prophase, Inoué and Sato in 1967 proposed a model in which the spindle is envisaged as a labile structure in dynamic equilibrium with a pool of subunits. Mitosis is thus a process which reflects the sequential assembly and disassembly of different structures to perform different functions. Inhibitory agents may act by disrupting the dynamic equilibrium. This model had a profound effect in engendering the concept of certain microtubules being labile structures capable of being polymerized or depolymerized under cellular control (Tilney, 1971).

However, several lines of evidence suggest that there are different classes of microtubules. Morphological variations in arrangement and in associated components have already been briefly described. Perhaps more importantly, microtubules of cilia or flagella are not depolymerized by treatments which reversibly destroy the birefringence of the spindle. Similarly, certain cytoplasmic microtubules of higher plant cells appear to be resistant to anti-mitotic chemicals. The generalization has been made that microtubules be classified as stable, i.e. those in, for example, cilia and flagella, or labile, i.e. cytoplasmic microtubules in many types of animal cells (Margulis, 1973).

7.2—
Biochemical Studies

7.2.1—
Drug Interaction

The responses of certain types of microtubule to anti-mitotic agents, together with their apparent repeating substructure, thus gave rise to the concept of the

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microtubule as an assembled polymer of soluble subunits. Early investigations into the nature of the subunit were hampered by the absence of a method or an assay for the recognition of a soluble microtubule component, even when such enriched sources as flagella or the mitotic apparatus were examined. This problem was overcome in the mid-1960's when it was postulated that the microtubule system was the direct target of the anti-mitotic drug, colchicine, and that the subunit might therefore be recognizable as a drug receptor.

Preliminary experiments with human carcinoma cells in culture established that, at the low levels of colchicine which depolymerized microtubules, there were no effects on other areas of metabolism such as respiration or protein synthesis (Taylor, 1965). Consideration of the kinetics of drug uptake suggested the presence of a single type of binding site. Work in two laboratories confirmed, using radiolabelled drug, that colchicine itself was indeed bound non-covalently to a single, soluble, protein species (Borisy & Taylor, 1967; Wilson & Friedkin, 1967). Since colchicine-binding could be assayed quantitatively, using gel filtration or ion exchange procedures, the reaction assumed great importance in the subsequent isolation and characterization of the subunit (Adelman et al., 1968).

It was quickly established that colchicine is bound in a stoichiometric 1:1 relationship to a dimeric protein with a molecular weight of 120,000 (see Wilson & Bryan, 1974). Denaturation to the protomers results in loss of binding. The binding affinity is high, of the order of 2 × 10⁶ litres mole^–1 , and it is important to note that there is no binding at all of the colchicine isomer, lumicolchicine. Although the protein can be stored in the frozen state, an outstanding feature of the binding site is its instability in solution, where it shows first order decay kinetics and a half-life of the order of hours even under optimal conditions of pH, temperature and ionic environment. Inclusion in the medium of the nucleotide, GTP, or the alkaloid, vinblastine, stabilizes and binding activity to some extent.

Evidence that the colchicine binding component is the microtubule subunit was initially circumstantial. High levels of colchicine binding protein were obtained from material rich in microtubules, e.g. dividing cells, cilia and brain cells. The physical characteristics of the binding protein with regard to its molecular weight and dimeric nature were similar to those of the major protein species in flagella and cilia. Evidence of a more direct nature was obtained through ultrastructural observations of the solubilization of the central pair nucleotide, GTP, or the alkaloid, vinblastine, stabilizes the binding activity to binding activity.

The binding site for colchicine seems to be masked when the subunits are assembled into microtubules. This, together with the instability of the site, was responsible for initial lack of success in obtaining colchicine-binding activity from stable outer doublet microtubules of flagella. However, the presence of the colchicine site in such subunits has been demonstrated using appropriate solubilization procedures (Wilson & Meza, 1973). Thus, the colchicine binding

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protein has been obtained from a variety of types and sources of microtubule. It is now generally accepted that labile microtubules are depolymerized by the binding of colchicine to subunits in the soluble pool, with consequent disruption of the equilibrium.

The colchicine binding moieties isolated from diverse sources of microtubules display similar physical and chemical properties (see Bryan, 1974). The name 'tubulin' has been applied to this dimeric, functional subunit of microtubules. Tubulin is an acidic, globular protein with a sedimentation coefficient of 6s and an agreed molecular weight of 110–120,000, whose behaviour approximates that of a prolate ellipsoid with an axial ratio of 5:7. The amino acid composition prompted initial comparison with actin, but subsequent comparable peptide maps of both proteins argued against any homology between tubulin and actin (Stephens, 1970). Treatment of tubulin with chaotropic agents such as urea or guanidine. HCl results in a reduction of the apparent molecular weight to 50–60,000. The two protomeric polypeptides can be resolved by appropriate electrophoretic analysis and have been named a and b tubulin, the more mobile electrophoretic component being b tubulin. A number of chemically reactive sites have been distinguished on the tubulin heterodimer. Two sites are known to be involved in drug interaction, a site for colchine and podophyllotoxin, and a separate site for vinblastine. There are also sites which bind endogenous

Table 7.1. Colchicine binding activity from various animal and plant sources*.
		Bound ³ H-colchicine
Author	Source	cpm mg^–1 protein in extract supernatant	cpm g^–1 fresh weight
Borisy & Taylor, 1967	squid axoplasm	180,000	—
	pig brain	50,000	—
	rabbit muscle	4,800	—
	HeLa cells	54,000	—
	sperm tail	60–90,000	—
	mitotic apparatus	13,000	—
	slime mould	100	—
Wilson & Friedkin, 1967	grasshopper embryo	40,000	—
Haber et al., 1972	yeast	7,000 (colcemid)	—
Hart & Sabnis, 1973	pea internode	(80)**	220
(and unpubl.)	hogweed petiole	(135)	480
	hogweed vascular tissue	(195)	970
	barley coleoptile	(90)	310
	mustard root tips	(540)	1,390
* Original data, uncorrected for colchicine concentration or radiocounting efficiency. ** Calculated values for comparison with animal sources; colchicine binding activity not detectable in 100,000 × g supernatants of higher plant extracts, without pre-concentration of protein by ammonium sulphate precipitation.

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ligands, including a site where GTP (guanosine triphosphate) is tightly and non-exchangeably bound and another site to which GTP is bound in such a way as to allow exchange with exogenously added GTP. Dephosphorylation of the terminal phosphate of GTP and transphosphorylation between the two sites have been suggested to play a role in microtubule assembly. In addition, other sites may be involved in interactions with Ca²⁺ and Mg²⁺ ions. The relationship of all these sites to the protein-protein contact sites necessarily involved in the vertical and lateral bonding of the subunits in the assembled tubule is unclear.

Recent examinations of plant preparations for a tubulin-like moiety with characteristic colchicine-binding activity have met with limited success. Such activity has either been undetectable or present only to a low degree (Haber et al., 1972; Hart & Sabnis, 1973; Burns, 1973; Heath, 1975a). The stability and specificity of the binding render it unlikely that all such activity is attributable to a plant tubulin. Table 7.1 shows the levels of colchicine binding activity from a variety of tissues, including plants.

7.2.2—
Polymerization

Since the behaviour of microtubules in vivo obviously involves their strictly regulated assembly-disassembly, a primary aim throughout the biochemical investigations was to develop a system in which polymerization could be studied in vitro (see Borisy et al., 1974). Early studies in this area did indeed demonstrate the presence of ordered structures in tubulin preparations. However, the relevance of such systems to the situation in vivo was considered doubtful, either because the process did not show characteristic kinetic responses to colchicine and temperature, or because the structures did not look like microtubules. It was therefore a notable advanace when authentic microtubules from rat brain, sensitive to colchicine and temperature, were first assembled in vitro, the crucial factors being inclusion in the medium of GTP and the calcium-chelating agent, EGTA (ethylene-glycol-amino ethylether tetracetic acid) (Weisenberg, 1972). It was subsequently reported that polymerization, using pig brain tubulin, also requires nucleation sites which are in the form of discs (29 nm diameter, 17 nm lumen). These successful polymerization studies finally confirmed that the colchicine binding protein is the microtubule subunit. In addition, they have provided a means of demonstrating the probable universality of the tubulin moiety, via co-polymerization of tubulin from such disparate sources as Chlamydomonas flagella and pig brain.

The ability to polymerize microtubules in vitro has initiated several exciting new avenues of research. In the first place, it has provided a method for isolating microtubule protein, rather than tubulin, the drug receptor. Cycles of centrifugation at 37°C and 0°C (i.e. with subunits alternately in the polymerized and depolymerized states) allow microtubules to be isolated with a high degree of purity. Both the ultrastructure and the chemical composition of such preparations have been examined. Electrophoretic analyses of purified microtubules by several

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laboratories has confirmed the presence of more than ten other proteins, including some with a molecular weight some with a molecular weight greater than 200,000. The roles of these proteins are the subjects of intense investigation. Some seem to be involved in the regulation of assembly. Others, with a molecular weight close to that of dynein (the ATPase 'arms' of flagellar microtubules) may be involved in the functioning of microtubules.

Secondly, in vitro polymerization has made the assembly process itself amenable to experimental study. The initial test of biological relevance seems settled, with endothermic, reversible, polymerization system responding to many factors known to regulate in vivo assembly of microtubules. The questions of nucleation centres and morphopoeitic proteins have already been mentioned. In addition microtubules have been successfully polymerized in vitro on cellular components such as asters, basal bodies and kinetochores. The pathway of polymerization is being deduced from combined biochemical and ultra-structural examination of experimentally induced intermediates. A commonly observed intermediate is a ring structure of tubulin, varying in dimensions, and complexity with different investigators. Whether the ring structure serves as a nucleation centre for stack-wise growth of microtubules, or whether it uncurls to form protofilaments which then associate laterally, is unclear.

7.3—
Biological Studies

Microtubules appear to be involved in several functional roles within the plant cell (see Newcomb, 1969). For convenience, these can be grouped under five headings: (1) Cytoskeletal role; generation and maintenance of cell shape (2) Cell wall architecture; cell differentiation (3) Intracellular transport (4) Chromosome movements; cell plate formation (5) Cell motility; cilia and flagella. In this section, we shall consider the involvement of microtubules in these phenomena, with emphasis on possible operative mechanisms, and drawing, and drawing, where necessary, on information obtained from studies of animal cells.

7.3.1–
Cytoskeletal Role

Despite the forces of surface tension, animal and some lower plant cells can maintain a non-spherical shape in the absence of a rigid cell wall. Numerous electron microscope studies have revealed that microtubules are present in large numbers and with distinctive orientations in such asymmetric cells or cell extentions. Correlations between cell asymmetry and microtubules have been extensively studied in animal cells, for example in erythrocytes, explanted nerve ganglia or neuroblastoma cells, lens epithelial cells and mesenchyme cells of echinoderm embryos. When these cells are treated with microtubule depolymerizing agents (high hydrostatic pressure, low temperature or colchicine), they assume spherical shapes.

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The unicellular alga, Ochromonas, lacks a cell wall or pellicle, yet has a characteristic pear-shape, with a narrow tail or rhizoplast, and a bulbous anterior bearing a small projection, the beak. Two sets of microtubules are involved in maintaining this shape. The plasma membrane of the main cell body is underlain by a series of curved microtubules which extend into the rhizoplast. The other set of microtubules is associated with the beak complex. Disassembly of the microtubules with colchicine or hydrostatic pressure leads to loss of the characteristic cell shape; removal of the depolymerizing agent allows the normal shape to regenerate even in the absence of protein synthesis (Brown & Bouck, 1973).

The two sets of microtubules in Ochromonas are differentially sensitive to colchicine and pressure. At relatively high concentrations of colchicine (10 mg ml^–1 ) or at high pressure (8,000 psi), both sets of microtubules are affected but at different rates: first, disassembly of rhizoplast tubules is correlated with disappearance of the posterior tail; this is followed by loss of beak tubules and beak asymmetry. Lower concentrations of drug (5 mg ml^–1 ) or lower pressure (6,000 psi) selectively disassemble only the rhizoplast microtubules. These results possibly indicate separate roles for each set of microtubules in determining the shape of this cell. Assuming that different tubulins are not involved, the beak and rhizoplast microtubules may represent two equilibrium systems competing for a common pool of subunits. Alternatively, the differential sensitivity of the two systems, and their sequential reassembly, may reflect characteristics of the sites which initiate microtubule polymerization.

The protozoan, Actinosphaerium, has been extensively used in similar studies on the morphopoietic function of microtubules. The organism is a spherical cell bearing numerous, long, relatively stiff, protoplasmic extensions or axopods, containing a central core, the axoneme (Fig. 7.3a). Each axoneme is composed

Figure 7.3a
Photomicrograph of Actinosphaerium
showing the slender cell extensions
or axopods radiating from the cell
body. Scale marker 100 µ m.

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of several hundred axially aligned microtubules, arranged into two rows that coil about a central axis (Fig. 7.3b). This strict pattern does not seem to be determined by a template of initiating sites, since, during axopod growth, the microtubule array is initially disorganized and becomes progressively more geometrically perfect.

Figure 7.3b
Cross section through an axoneme of Actinosphaerium
near the axopod base. Two spiral rows of microtubules are
coiled about a central axis in a precise geometrical arrangement.
Scale marker 1 µm. From Tilney & Porter (1965) Protoplasma 60,
317–43, reproduced by permission of the authors and Springer-Verlag.

When low temperature, pressure or drugs are employed to depolymerize microtubules of Actinosphaerium, the axopods undergo immediate retraction. In contrast, D₂ O, which stabilizes microtubules, prevents the breakdown of axopods by cold or pressure. Retractions and extensions are a normal feature of individual axopods and serve to move the cell across its substratum. Thus, the cell must exert a fine control over the behaviour of microtubules in each axopod separately (see Gunning & Steer, 1975).

In flowering plants, the male sperm is formed from the generative cell which usually has no cell wall and lies within the pollen grain. Growth of the pollen tube is accomplished by the vegetative cell, while the generative cell adopts a spindle-shape before entering and travelling down the pollen tube. Development of the spindle shape is accompanied by the appearance of microtubules aligned parallel to the long axis of the generative cell. Treatment of the generative cell with drugs leads to a loss of microtubules and the cell adopts a spherical form (Sanger & Jackson, 1971).

Thus, microtubules are apparently involved in both generating and maintaining changes in cell shape. The mechanisms regulating orientation and assembly of microtubules in this role are still unknown. Microtubules in the

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axonemal cores of Actinosphaerium axopods are connected with their neighbours by cross-bridges. Whether these lateral links function in stabilizing the extended structure or in orienting the microtubules has yet to be determined.

7.3.2—
Cell Wall Architecture

During interphase in higher plant cells, cytoplasmic microtubules lie close to the cell membrane and are arranged circumferentially along the lateral walls of the cells but are randomly disposed underlying the transverse walls (Fig. 7.5a). In the earliest paper reporting the presence of microtubules in higher plant cells, Ledbetter and Porter (1963) noted the parallel alignment of cellulosic microfibrils of the wall and the subjacent cytoplasmic microtubules (Fig. 7.4). Numerous subsequent studies have confirmed this observation, in algal cells and in higher plant cells undergoing both primary and secondary wall formation (see Hepler & Palevitz, 1974).

In the long xylem fibres of Salix, wall microfibrils are deposited in two different orientations simultaneously in the middle and at the extremities of the cell. Even in this situation, microtubule alignment mirrors that of the overlying microfibrils, indicating the ability of the cell to maintain two sets of microtubule-microfibril associations. In the lorica stalk of the alga, Poteriochromonas, the primary wall microfibrils are arranged helically and tend to fasciate into ribbonlike fibrils, 20 nm in width. Every such wall fibril coincides precisely with a microtubule, the plasmalemma separating the two sets of linear structures (Schnepf et al., 1975).

Other examples of the association between microtubules and microfibrils are afforded by cells undergoing irregular or sculptured deposition of secondary wall material. During xylogenesis, microtubules are specifically grouped under the developing wall thickenings in vessels, and, once again, are oriented parallel to the microfibrils. Similar situations prevail during the development of wall thickenings in differentiating guard cells and during the deposition of the nacreous walls of sieve elements.

Attempts have been made to confirm the association between wall microfibrils and microtubules by treating cells with colchicine. However, plant tissues seem relatively resistant to a wide variety of drugs (Heath, 1975b) and, even at high concentrations of colchicine, not all cytoplasmic microtubules in plant cells are depolymerized (Pickett-Heaps, 1967; Wooding, 1969). Also, such levels of colchicine are known to affect other cellular processes, in particular, membrane-related phenomena (see Wilson & Bryan, 1974). Therefore, although wall microfibril orientation is often distorted in the presence of colchicine, such results should be interpreted with caution.

7.3.3—
Intracellular Transport

There is much evidence to indicate that wall precursors are transported to the wall in dictyosome-derived vesicles. On the other hand, there is no evidence to

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Figure 7.4
Microtubules running parallel to one another immediately beneath
a primary cell wall in root tips of bean ( Phaseolus vulgaris ). The
microfibrils of the wall are oriented parallel to the microtubules. The
axis of cell elongation (indicated by the arrow) is at right angles to the
orientation of the microtubules and microfibrils. Vesicles can be seen
among the microtubules. Scale marker 0.1 µ m. From Newcomb (1969)
A. Rev. Pl. Physiol. 20, 253–88, reproduced by permission
of the author and Annual Reviews, Inc.

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suggest that microtubules provide the motive force for such transport. However, it has been proposed that the parallel microtubules girdling the cells of higher plants peripherally provide a framework for the guidance and alignment of these vesicles. In some cases the microtubules lie so close to one another that it has been argued that they could function only by excluding vesicles or elements of the endoplasmic reticulum, preventing their fusion with the plasmalemma at certain sites. In either case, a vectorial role for microtubules is indicated.

Ledbetter and Porter suggested that microtubules may be involved in orienting and even driving cyclosis of cytoplasm, a process which might indirectly be responsible for the alignment of wall microfibrils. However, several lines of evidence have recently implicated cytoplasmic microfilaments in this role (Hepler & Palevitz, 1974). These filaments, 5–8 nm in width, are more consistently located and aligned within the zones of streaming cytoplasm. The microfilaments are morphologically identical to F-actin, one of the major contractile proteins of muscle. Recently, microfilaments in cells of both Nitella and higher plants have been decorated with heavy meromyosin, a characteristic test for actin-like filaments. In primitive organisms, such as the slime mould Physarum, there is overwhelming evidence to indicate that similar micro-filaments provide the motive force for rapid cytoplasmic streaming.

However, around the axonemes of Heliozoan axopods described earlier, cytoplasmic particles stream to and from the cell body. Microfilaments have not been seen in this structure. By contrast, in the giant coenocytic alga, Caulerpa, microtubules are located within the zones of, and parallel to, the numerous sluggish cytoplasmic streams. Mitochondria, plastids and other cytoplasmic components are constantly circulated to and from the growing tips of this large, asymmetric cell. In the chromatophores of animal cells, microtubules have been shown to be essential to the movements of pigment granules. Anti-mitotic agents depolymerize microtubules and simultaneously disrupt the alignment and arrest the movement of the pigment granules (Murphy, 1975). Thus, it remains possible that certain types of cytoplasmic transport require a microtubule framework in both a vectorial and an active role.

7.3.4—
Cell Division

At least two types of microtubule can be distinguished in most spindles, those that traverse the spindle from pole to pole (continuous fibres), and those that connect the kinetochores of the chromosomes to the poles (chromosomal or kinetochore fibres). During separation of chromatids in anaphase, the continuous fibres lengthen, increasing the distance between the poles, while the chromosomal fibres usually—but not always—shorten, further separating the chromatids. After telophase, the continuous spindle may persist and be added to in the interzonal region by numerous additional microtubules, leading to formation of the phragmoplast and eventually, the cell plate.

Considerable changes in microtubule organization (Fig. 7.5, A–F) take

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Figure 7.5
Diagram of changes in microtubule orientation and distribution during the phases
of mitosis in a higher plant cell. A—interphase; B—preprophase; C—prophase;
D—metaphase; E—anaphase; F—telophase. From Ledbetter & Porter (1970)
Introduction to the Fine Structure of Plant Cells, p. 44, Springer-Verlag, Berlin,
reproduced by permission of the authors and Springer-Verlag.

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place during the events of a typical mitotic cycle (Fuge, 1974). With the onset of division, the peripheral cytoplasmic microtubules of the interphase cell disappear. Concomitantly, a band of microtubules is formed, several layers deep, lying close to the cell wall and girdling the nucleus perpendicularly to the prospective spindle axis. This preprophase band of microtubules appears to predict the plane of the future cell plate (Pickett-Heaps & Northcote, 1966). This has been demonstrated in divisions destined to give daughter cells of different size, for example in the asymmetric divisions of guard mother cells of wheat.

At the beginning of prophase, the number of microtubules in the preprophase band decreases, and new microtubules appear close to the nuclear membrane, this time running parallel to the prospective spindle axis. The microtubules in this so-called 'clear zone' in turn appear to furnish protein material for the spindle. It is possible that a total transformation of these extranuclear microtubules into spindle microtubules takes place.

Prior to spindle formation, the nuclear envelope either disintegrates or becomes perforated at the prospective poles, and cytoplasmic (clear zone) microtubules enter the karyoplasm at these sites. Some of these microtubules appear to establish connections with the chromosomal kinetochores. Since the number of kinetochore microtubules (kMts) increases as prometaphase proceeds, and since clear zone microtubules can be utilized only immediately after breakdown of the nuclear envelope, it is likely that de novo microtubule assembly also occurs at the kinetochores. Thus, it appears that the kinetochore, which varies in appearance in different organisms from amorphous to highly structured, acts as an initiating site or microtubule morganizing centre (MTOC).

In prometaphase, continuous fibres also begin to form and to increase in number. There is some controversy as to whether in higher organisms the continuous microtubules pass from pole to pole. A large proportion appear to overlap in the interzone between the half-spindles (Fig. 7.6), and have been termed non-kinetochore microtubules (nkMts). In lower plants, and all animal cells, with centrioles, the continuous or nkMts originate near the centrioles. These loci are represented by amorphous, electron-dense aggregates of material, and behave as another set of MTOCs. The essential feature of the centriole in mitosis seems to be in determining the spindle axis. In higher plants that lack centrioles, neither the location nor form of polar MTOCs is clear.

During metaphase, engagement of sister chromatids to opposite poles takes place via microtubules, followed by movement of the chromatids to the equatorial plate. At this stage, the maximum number of spindle microtubules is attained, the numbers often trebling between prometaphase and metaphase. Estimates from microtubule counts in serial sections of the metaphase spindle of the sea urchin, Arbacia, indicate a total microtubule length of 50,000 µm. The very large

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Figure 7.6
Electron and light micrographs of a dividing cell in prometaphase showing
kinetochore and nonkinetochore microtubules. The line indicates the equatorial
plate. Scale marker 1 µm. From Bajer (1968) Symp. Soc. exp. Biol. 22, 285–310,
reproduced by permission of the author and University Press, Cambridge.

spindles of Haemanthus endosperm cells contain considerably more microtubules than those of Arbacia.

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Anaphase separation of the chromatids involves direct translation of the kinetochores at almost uniform velocity (0.2–4 µm min^–1 ) towards their respective poles. Two experimentally separable processes are involved: first, a movement of the chromatids towards their poles, during which the chromosomal fibres shorten to as little as 20% of their metaphase length; secondly, an elongation of the spindle itself, during which the distance between the poles increases, thereby further separating the two groups of chromatids.

Time-lapse films of dividing Haemanthus endosperm cells show that many other objects, such as vesicles and granules, are commonly transported polewards during prometaphase-anaphase. Conversely, in onion root tip cells, the poleward movement of the chromosomes is accompanied by a reciprocal movement of other materials from the poles towards the mid-plate. Elements of the endoplasmic reticulum accumulate at the poles during metaphase, enter the spindle at anaphase, pass between the chromosomes moving in the opposite direction, and eventually aggregate and coalesce at the mid-plate to give rise to the transverse cell wall at telophase.

Dissolution of kMts and nkMts during telophase is accompanied by the formation of new microtubules in the interzone, leading to the formation of the phragmoplast in higher plant cells (Fig. 7.7). Phragmoplast microtubules appear to be involved in the movement, alignment and fusion of ER- and

Figure 7.7
Phragmoplast microtubules at the mid-plate during telophase.
Vesicles contributing to the forming cell plate can be seen aligned among the
microtubules. Scale marker 1 µm. From Hepler & Jackson (1974) J. Cell. Biol. 38, 437–46,
reproduced by permission of the authors and The Rockefeller University Press.

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dictyosome-derived vesicles to form the cell plate—yet another example of microtubule-associated intracellular transport.

Thus, distinct changes in the polarity and distribution of spindle microtubules take place during the various phases of the mitotic cycle. It should be borne strongly in mind that such changes occur under conditions of very low protein synthesis. Consequently, both assembly and functioning of the mitotic apparatus must involve regulation of microtubule polymerization at a physico-chemical level. Interaction of microtubules with other components of the mitotic apparatus must also be regulated at this level. Several hypotheses have been proposed to explain the mechanism of mitotic movements, although little direct experimental evidence to support the models has yet been obtained.

One prevailing hypothesis to account for spindle elongation (Brinkley & Cartwright, 1971) is that the poles are pushed apart by tip growth of continuous microtubules in the course of which subunits from disassembled kMTs may be utilized. Shortening of the chromosomal fibres is thought to occur by depolymerization at the poles.

Another hypothesis based on the sliding of adjacent spindle microtubules was proposed by Mcintosh et al., (1969). Earlier, several workers had reported the existence of arms or cross-bridges between spindle microtubules. The basis of the sliding tubule hypothesis is that the intertubular bridges in the spindle serve as mechano-chemical elements. Sliding of two microtubules against each other is thought to be generated by the successive breaking and reforming of cross-bridge linkages, a process that is energy-consuming and mediated via ATPase activity of the bridges. It has been estimated that the energy released through the hydrolysis of approximately 20 ATP molecules would suffice to move an average chromosome from the metaphase plate to the pole.

A prerequisite for the McIntosh model is an ordered system of long microtubules, emanating from the poles and reaching far into the opposite half-spindles in metaphase. Microtubules from opposite poles are suggested to have antiparallel polarity with regard to the direction in which their lateral arms could produce force. The kMts in each half-spindle would also show polarity; their lateral arms would only produce force directed towards the spindle equator. If one of the long interpolar microtubules and a kMt of antiparallel polarity came into contact at anaphase, the two tubules, due to the directed activity of their arms, would push themselves in opposite directions by sliding. The model is analogous to the generally accepted model for the interaction of actin and myosin in skeletal muscle. The mechanism could cause chromosome separation, as well as spindle elongation when anti-parallel nkMts interact where they overlap.

A third hypothesis involving actin-microtubule interactions (Forer & Behnke, 1972) has derived from observations of actin-like thin filaments in the mitotic or meiotic spindles of both animal and plant cells. Fluorescein-labelled heavy meromyosin binds to isolated sea urchin spindles. At the ultrastructural level, microfilaments within the spindle have been decorated with heavy

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meromyosin, a test for actin-like proteins (Gawadi, 1971; Forer & Behnke, 1972). Several other recent publications have confirmed the presence of actinlike filaments in dividing cells. In kangaroo rat cells, actin was only demonstrable in association with chromosomal fibres (Sanger, 1975), suggesting an actinmyosin type interaction as the force-producing mechanism for chromosome movements, that is, for the autonomous movements of chromosomes in anaphase, as opposed to spindle elongation which may be controlled solely by microtubule assembly-disassembly.

Myosin-like proteins have not yet been located in spindles. Several workers have suggested that microtubules might serve as rigid structures against which the actin filaments could exert a contractile force by the formation and breakage of cross-links. A Ca²⁺ -dependent ATPase (similar in this respect to both myosin and dynein) is present in the isolated spindle in a concentration three times that in the cytoplasm (Mazia et al., 1972).

The theories on the mechanisms involved in chromosome separation have arisen from observations of living or fixed, intact cells. Such studies are hampered by the inability to experiment directly with the mitotic apparatus, except as it exists in the complexity of its cellular environment. Exciting possibilities have been raised by recent successful attempts to isolate and experiment with the mitotic apparatus in vitro. Intact spindles from the eggs of the surf clam show normal sensitivity to temperature and colchicine. These isolated spindles can incorporate brain tubulin during their reassembly and such hybrid spindles are also cold labile, Ca²⁺ -sensitive and capable of considerable increase in overall length (Rebhun et al., 1974). Moreover, early anaphase spindles isolated from kangaroo rat cells will continue chromosome motion, in the absence of exogenous spindle subunits, when ATP is added (Cande et al., 1974). These results already suggest that while spindle growth requires microtubule polymerization, anaphase movements do not.

7.3.5—
Cell Motility; Cilia and Flagella

Microtubules, arranged in a characteristic 9+2 configuration (Fig. 7.8a), are the major structural components of flagella, cilia and sperm tails (Warner, 1974). The flagellar apparatus consists of a membrane-bounded, slender, cylindrical cell extension subtended by its basal body, the intracellular organelle which appears to be its origin and kinetic centre.

Basal bodies are identical in structure and apparently homologous with the centrioles of animal and lower plants cells. During mitosis, centrioles appear to determine the poles of the spindle axis and to organize the assembly and alignment of spindle microtubules. Following mitosis, in flagellated cells, the centrioles migrate to the cell periphery and organise the assembly of the flagellar apparatus. The mechanism whereby centrioles act as microtubule initiating centres, the reasons for their stability during the life cycle of the cell, and the factors controlling basal body replication in multiflagellated cells, are all unknown.

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Figure 7.8a
Diagram of flagella cross section, showing
arrangement of microtubules, arms and cross-links.

Basal bodies are composed of nine sets of triplet microtubules (A, B and C) with lateral connections between the A and C tubules of adjacent triplets. At the proximal end of each basal body, a thin filament runs from the A tubule of each triplet to a central hub, forming a cartwheel pattern. At the distal end is the complex transitional region between the microtubules of the basal body and those of the cilium.

The shaft of the cilium is characterized by a central pair of axially oriented microtubules, surrounded by a ring of nine doublets (Fig. 7.8a). The A sub-fibre of a doublet possesses a complete wall of 13 protofilaments. However, the B subfibre has only 10 protofilaments in its wall, and shares the three protofilaments of the A tubule forming the common partition between them.

From the A tubule of a doublet, two short arms project towards the B tubule of the adjacent doublet. These arms have been isolated and shown to possess Mg²⁺ -dependent ATPase activity (Gibbons, 1965). The arms are approximately 30 nm long, 9 nm wide, and spaced at intervals of 16–22 nm along the length of the A tubule. The enzyme has been called dynein after its postulated role in converting chemical energy into mechanical force. In addition to the dynein arms, cross-bridges have been seen extending between the outer doublets and the flagellar membrane. Radial links, connecting the central pair of tubules with the outer doublets, have also been described. Further, the two microtubules of the central pair differ from one another, one member exhibits two rows of short projections, 18 nm long, and spaced at intervals of 16 nm. Thus, a complex of radially or longitudinally arranged arms, bridges or filaments is present in the flagellar matrix in association with microtubules (Fig. 7.8b).

The most widely accepted hypothesis to account for flagellar motion is the sliding microtubule model (Satir, 1974). Applied to flagella, the hypothesis states that the force responsible for motion is produced when the axonemal

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Figure 7.8b
Longitudinal section of a flagellum showing connections between the central and
outer doublet microtubules, and bridges between the peripheral tubules and the cell
membrane. Scale marker 0.1 µm. Fig. 8.8b. from Ringo (1967 J. Cell Biol. 33, 543–71,
reproduced by permission of The Rockefeller University Press.

microtubules, which do not change in length, tend to slide with respect to one another. Accordingly, the model predicts that in different stroke positions, the morphological relationships of the microtubules will change in a systematic fashion, so that the geometry of the bent flagellum will be reflected in the displacement of the microtubules. This has, indeed, been demonstrated using serial sections of the lateral gill cilia of the freshwater mussel.

Other evidence has also accumulated to support this hypothesis. Glycerinated or demembranated flagella beat normally upon the addition of ATP and Mg²⁺ or Ca²⁺ ions. When such naked axonemes are briefly treated with trypsin, the circumferential and radial links holding the axoneme together are interrupted, whereas the dynein arms are trypsin-resistant. Addition of ATP now causes a sliding of the microtubules, so that the axoneme grows very much longer and thinner as groups of doublets crawl over one another. This sliding must normally be converted to bending by the series of intermicrotubule connections, within the axoneme.

Thus, microtubules are implicated, in both plant and animal cells, in a variety of processes involving the active movement of cells or cellular components. However, the possibility that a single basic mechanism of action underlies their seemingly diverse roles is still in doubt. The dynamic equilibrium model involving assembly and disassembly of microtubules cannot be applied to the stable microtubules of flagella or the cytoplasmic microtubules of higher plant cells. Neither the assembly-disassembly hypothesis, nor the sliding filament model alone appears adequate to explain the complex functioning of

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microtubules in the mitotic apparatus. Even within a process such as spindle elongation, microtubules may function in different ways. For example, the spindles of HeLa cells show an initial rapid phase of elongation, followed by a subsequent slow phase: colchicine only inhibits the slow phase (presumably by interfering with microtubule assembly) but does not affect the rapid phase which may be based on a sliding mechanism. Furthermore, dividing plant and animal cells are remarkably different in their sensitivity to drugs: the spindles of many plant cells require treatment with approximately 1,000-fold higher concentrations of a wide variety of anti-mitotic drugs to achieve mitotic arrest.

7.4—
Concluding Remarks

It is noteworthy that close collaboration between the electron microscopist and the biochemist has been demanded at almost every stage in the development of knowledge concerning microtubules. After the discovery of the microtubule as an ubiquitous organelle of the eukaryote cell, the physiological manipulation of cells in ablation experiments, together with continuing ultrastructural work, confirmed the interaction of microtubules in a variety of cellular activities. Such studies also stimulated development of the concept of the microtubule as a macrostructure in dynamic equilibrium with its constituent subunits. Biochemical investigation into the nature of the subunit initially depended heavily on the electron microscope for evidence that the moiety was relevant to the microtubule. The phase of study culminating in the development of an in vitro polymerization system also relied on the electron microscope to provide evidence of the biological relevance of both the process and final structure.

Knowledge concerning microtubules has been derived mainly from studies involving animal tissues or cells of lower plants. Elucidation of the roles of microtubules in higher plant cells has not proceeded so rapidly. Ultrastructural investigations have confirmed the ubiquity of the organelle and have shown its oriented presence to be correlated with a variety of processes. While it is clear that microtubules are involved in various stages of cell division, their precise role in the earlier events which predict the plane of cleavage is unclear. Similarly, their functioning in other aspects of plant development, including cell wall growth and differentiation, remains conjectural. There are, of course, practical difficulties in studying microtubules in the cells of higher plants. Such problems vary from the general, i.e., lack of tissue specialization and presence of a vacuole and cell wall, to the more specific paucity of microtubules in the meagre protoplasm of the differentiated higher plant cell. In addition, there would seem to be a problem in the resistance of certain plant cytoplasmic microtubules to depolymerizing agents. Since this includes, in some cases, stability to such physical agents as temperature and pressure, it would seem that it is a real property of the microtubule rather than an indirect effect of, say, cellular impermeability towards a drug. This property may be related to the stability of flagellar micro-

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tubules. It remains to be seen whether the stability is due merely to a lack of an equilibrating subunit pool or to some other feature of this type of microtubule.

Much remains to be learned about plant microtubules, their subunits and associated components, not only to gain further understanding of their roles in specific processes of plant growth and development, but also for comparison of their properties with those of equivalent components from animal sources. The latter aspect, in addition to answering questions concerning the evolutionary origins of this important group of proteins, would also seem to be a prerequisite to determining any common modes of functioning. How microtubules function, what factors regulate their activity in various cellular processes, and what relationships exist among these diverse activities are some of the questions for research.

Chapter 8—
The Endomembrane System and the Integration of Cellular Activities

8.1—
Introduction

The invention of ultra-thin sectioning gave electron microscopists the first sight of a new level of organization within the cell: a system of membranes forming microstructures and sub-compartments throughout the protoplasm. Some components of this system are small and discrete: the mitochondria, plastids and microbodies. Because they therefore survive cell breakage intact, biochemists have been able to characterize the specialized functions of each type. Others, the nuclear membrane, the plasma membrane (plasmalemma) and the tonoplast, bound discrete compartments so large that, though they do not survive cell breakage intact, many of their specialized functions have been worked out by cytologists and physiologists using whole cells.

Subtract these organelles from the membrane system of the cell and the remainder, a considerable bulk of material, is arranged as parallel, paired sheets fused at the edges to enclose a narrow and empty-looking lumen. The membranes are also fused to each other around holes in the twin sheet, and these fenestrations are occasionally so frequent as to reduce the paired membrane to a layer of tubules, or even isolated vesicles. A small proportion of this doublemembrane system, the Golgi apparatus, is organized into units of recognizable morphology, the Golgi bodies. The rest, the endoplasmic reticulum (ER), is relatively amorphous and, more often than not, distributed apparently haphazardly about the cytoplasm. Offering few clues for the microscopist and a host of problems for the biochemist, this material and its function in the cell is only just beginning to be understood.

8.2—
Techniques

8.2.1—
Electron Microscopy

Thin (ca 50 nm) sections provide a very limited image of the membrane system. Seen in profile, not much can be told about the way its parts are, or are not, connected. What is described from an isolated thin section as a collection of vesicles may in fact be a network of tubules, anastomosing in three dimensions, or a stack of paired lamellae, extensively fenestrated, or even a single tubule, coiled back on itself many times (Fig. 8.1). There are methods which provide information on structure in three dimensions. Thanks to the tremendous depth

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Figure 8.1
Identical thin sections can be produced from quite different structures:
(a) a collection of vesicles
(b) a three-dimensional network of tubules
(c) a stack of fenestrated lamellae
(d) a single, twisted tubule

of focus of the electron-microscope, using high-voltage electrons it is possible to view all the organelles within a thick (2 µm) section. The specimen can then be tilted slightly to provide stereo-pairs of micrographs.

The freeze-etch technique often provides information on membrane topography. In frozen tissues the hydrophobic bonds holding the two halves of each membrane together are virtually non-existent. The activity of cell water is so low, the lipids of the two halves of the bimolecular leaflet are no longer forced together. The freeze-fracture, which follows planes of weakness, may run through large areas of interconnected membrane. Freeze-etch studies often allow a direct comparison to be made between views of the internal surface and cross-fracture of the same membrane system, as in Fig. 8.2 where the ER is seen as continuous sheets arranged in concentric layers. Fenestrated ER can only be shown by freeze fracture (Fineran, 1973b).

To obtain a truly comprehensive picture of membrane interconnections requires serial sectioning. So lengthy and difficult it is rarely attempted, this technique has nevertheless proved highly rewarding. Not until 1973 was it discovered by serial sectioning that there is only one, highly branched mitochondrion in a yeast cell, and the same is true for the unicellular green alga, Chlorella (Atkinson et al., 1974; Gunning & Steer, 1975). Though freeze-etch pictures of plant cells have already provided much information on the shape of the endomembrane complex, thick section stereomicrography and serial sectioning are as yet techniques of the future.

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Figure 8.2
Freeze-etch preparation of pea root tip. ER is seen in cross-fracture and surface view,
and forms almost continuous sheets arranged in concentric layers. The arrow indicates
the direction from which heavy metal was evaporated and the scale line represents 1 µm.
(From Northcote, 1968.)

Figure 8.3
Autoradiograph of a thin section of a wheat root cap cell exposed to [6-³ H] glucose
for 30 minutes. Silver grains are located over slime- polysaccharide deposited between
the wall and the plasmalemma, and over Golgi bodies in the cytoplasm. The slime
deposit is sited under a gap in the peripheral ER. The scale line represents 1 µm.
(From Northcote & Pickett-Heaps, 1966.)

High resolution autoradiography is a powerful tool in the investigation of the synthetic activities of endomembranes. First, the living tissue is given a pulse of tritium-labelled precursor. At fixed intervals of time afterwards, portions of tissue are prepared for electron-microscopy, sectioned and the stained sections are coated with a very thin layer of radiosensitive, silver halide emulsion. To prevent the development of this emulsion by chemicals in the section, an inert layer of carbon is deposited on the sections before they are coated. From

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the changing distribution of silver grains over the organelles of the cell, the pulse of radioactive precursor can be followed from the site of synthesis of the first insoluble product to the final resting place of the completed polymer (Fig. 8.3). Because the b -particle may have moved in the plane of the section before hitting the emulsion, the practical limit of resolution of the technique is 100 nm and since the organelles in the cell are fairly crowded it may not be possible to decide unequivocally which of them contained the labelled material.

Proper controls for autoradiographic experiments would have to include some method of identifying the radioactive product in the section. For example, pre-treating the section with solutions of degradative enzymes specific for the polymer in question should eliminate the radioactivity associated with the organelles involved in its synthesis. Also, a precursor specific for the product under investigation should be used, e.g. [³ H]-orotic acid for RNA. Nonspecific precursor, such as [³ H]-glucose, is only useful with cells producing a single polymer-type, e.g. slime polysaccharide in root-cap cells or cellulose in differentiating xylem vessels. Always important, but especially necessary in this case, is a back-up biochemical analysis showing the distribution of label from the precursor in all polymeric fractions at the times that the tissue portions were fixed for electron-microscopy. It must be admitted that proper controls of this nature are hardly ever reported.

8.2.2—
Biochemistry

To isolate the various parts of the endomembrane complex, the biochemist breaks open the cells of a lump of tissue, in a pH-buffered solution to neutralize vacuolar acids, and separates membrane fractions from the rest of the cytoplasm by centrifuging. To homogenize is to use high shear forces to fragment all membranes. The little pieces spontaneously re-anneal forming small vesicles or 'microsomes'. Small quantities are homogenized in some sort of pestle and mortar arrangement, large quantities in a blender. Any compartmentalization of soluble substances in organelles is lost by this method, and the in/out orientation of the membranes of the vesicles does not necessarily represent that in vivo.

To prepare intact organelles, shear forces are minimized by crushing or chopping the tissue in a solution which is also buffered osmotically to prevent bursting. A few organelles will inevitably be broken and many of them will be left behind in pieces of more or less intact tissue. So, unlike homogenization, quantitative recovery of membrane from the tissue does not occur and thus it is impossible to relate directly, say, enzyme activities in isolated organelles, to the total activity in the tissue.

Differential centrifugation involves centrifuging at successively higher speeds for longer times to effect a separation on the basis of size. Thus debris and cell wall are spun out at 4,000 g for about 15–20 minutes, organelles at 10,000 g for about 30 minutes, and total membrane by 100,000 g for 1 hour. The membrane

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which pellets between 10,000 g and 100,000 g is referred to as a microsomal fraction and this material is often assumed to be ER, which probably does make up the bulk of it. Differential centrifugation is now only used to prepare organelle or microsomal fractions for subsequent analysis by density gradient centrifugation or, since resuspension of pelleted organelles causes some breakage, to remove debris before the total membrane material plus soluble cytoplasm is layered onto a gradient (Lord et al., 1973).

Density gradients are prepared in the centrifuge tube by pouring in a progressively less concentrated solution of sucrose in the buffered medium. The membrane preparation is added on top and centrifuged into the gradient. In an isopycnic separation, centrifugation is continued until the components of the sample reach their equilibrium positions in the gradient and so separation is effected on the basis of density alone. In a rate zonal separation, centrifugation is stopped before equilibrium is reached. Smaller particles sediment faster and so the ordering of components in the gradient is determined by size as well as density. Continuous gradients in which the density changes linearly with distance over a range selected from 1.06 g cm^–3 (16% w/w sucrose) to 1.23 g cm^–3 (50% w/w sucrose) are conventionally used to separate the components of membrane fractions. When the density of a particular component is known, discontinuous or 'step' gradients are sometimes used for bulk preparation. However this method does not produce fractions as pure as those from continuous gradients. A heavy particle arriving late at an already crowded interface between two densities in a step gradient may be trapped there or, as appears to happen more often, plummet through carrying light membranes down with it. (Note that results from sucrose gradients cannot be compared with those from experiments involving gradients of 'ficoll', a synthetic sucrose polymer sometimes used, since sucrose is osmotically active and organelles and vesicles lose water progressively as they fall through a gradient of it.) Centrifuging organelles and microsomes through linear density gradients produces distinctive patterns in the distribution of protein in the gradient, so it seems the endomembrane complex can be resolved into different types of membrane. The next step is to identify them.

If the gradient fractions are fixed and embedded for electronmicroscopy, stained thin sections can reveal the presence of organelles with a characteristic morphology, such as attached ribosomes. It would involve a lot of work to establish that a particular organelle was not present. Negative-staining is easier and faster, but less reliable since phosphotungstic acid solution is a powerful protein extractant and so, unless the membranes are glutaraldehyde-fixed, alters the appearance of organelles (Mollenhauer et al., 1973). Many membrane fractions, after conventional post-fixation with osmium and uranyl acetate staining of sections, appear as simple vesicles with no recognizable morphology. Efforts have been made to develop electron-dense stains which are specific for individual membranes of the cell, but so far only the plasmalemma has been stained selectively with any success using periodic acid and various heavy metal

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anions, e.g. chromate and phosphotungstate (Leonard & Van Der Woude, 1976). However, the chemical basis of the staining reaction is by no means certain (it is likely that the carbohydrate moiety of plasmalemma glycoproteins is involved) and its specificity for this membrane cannot always be repeated.

Gradient fractions can be characterized by various enzyme activities associated with them. The enzyme proteins may be freely soluble and enclosed by the membrane, or loosely associated with the membrane surface (extrinsic), or firmly bound in the membrane (intrinsic). Obviously only intrinsic enzymes can be trusted as markers. For organelles with unique and well-characterized functions, enzymes provide reliable identification, e.g. succinate dehydrogenase, an enzyme unique to the tricarboxylic acid cycle and very firmly bound, is used to indicate the presence of inner mitochondrial membrane. Otherwise, to determine the distribution in the cell of an enzyme activity localized in a gradient is a major problem.

Sometimes enzymes can be located in thin sections of cells by cytochemical techniques for electron-microscopy, e.g. specific phosphatases can be tracked down if inorganic phosphate released from supplied substrate is precipitated in situ on thin sections by heavy metal cations. Thiamin pyrophosphatase (TPPase) was shown to be a marker for the Golgi body in vertebrate cells in this way. The specificity of the reaction must be checked by comparison of results obtained using a range of alternative substrates. An enzyme localized by cytochemistry on thin sections cannot with certainty be used to establish the identity of membrane fractions which have not been treated with glutaraldehyde or heavy metal cations, both of which are powerful inhibitors of many enzymes.

An association of enzymes with particular membranes of the cell has sometimes been built up as a result of some knowledge of cell physiology, e.g. Na⁺ /K⁺ stimulated ATPase and callose synthetase are thought to be markers for plasmalemma since salt uptake and callose synthesis take place at the cell surface. However, the limitation of callose synthesis to the outer surface of the plasmalemma in vivo may be due to substrate availability and it would seem equally likely that the tonoplast, for which at present there is no marker enzyme, is also involved in salt uptake.

Alternatively, enzyme markers become accepted by long association. Thus, because the activity of NADH-specific cytochrome c reductase correlates well with the proportion of ribosome-bearing fragments in membrane fractions, this enzyme is widely accepted as a marker for ER. (Note that the inner mitochondrial membrane also shows NADH-specific cytochrome c reductase, which differs from that of the ER in being sensitive to antimycin A. mitochondria must be shown to be absent from the fraction and the activity demonstrably antimycin-insensitive when this marker is used for ER.) This sort of association between an enzyme and a specific membrane is only of any use for tissues and organisms in which it has been established as there is no reason to believe that markers are universal. Thus, glucose-6-phosphatase is an ER-marker in certain cells only, e.g. vertebrate liver and kidney, and together with 5'-adenosine

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monophosphatase (AMPase, 5'nucleotidase), a marker for plasmalemma in animals, does not occur in the equivalent membranes of plant cells.

Note that while the detection of marker enzyme activity can indicate which types of membrane are present, the absence of activity may be due to failure of the assay for a number of reasons other than absence of the enzyme. Also, no membrane fraction can be established as 'pure' on the basis of marker enzymes while there are still types, like the tonoplast, for which a marker does not exist.

8.3—
Composition and Characteristics of the Membrane Types

There is considerable evidence for broad homology between the nuclear membrane and the membranes of the ER, and some indications that the outer mitochondrial membrane and the plastid envelope, while essentially similar to the ER, are intermediate in composition between it and the specialized inner membranes of these organelles. From the little that is known of their composition, the membranes of the Golgi apparatus are closely allied to those of the ER, but the plasmalemma is markedly different from this membrane.

8.3.1—
Nuclear Membrane

Nuclear membrane is prepared from isolated nuclei by sonication to fragment the membrane, high salt treatment to remove associated nucleic acid, and density gradient centrifugation to collect the fragments. In a careful study, taking precautions to minimize phospholipid loss by enzyme degradation during isolation, Philipp et al. (1976) found no difference between the lipid and fatty acid composition and the nature and level of enzyme activities in nuclear membrane and ER from plants. Both membranes, for example, had NADH and NADPH-specific cytochrome c reductases at similar levels, and both possessed a cytochrome of the b₅ type. They could be distinguished by the higher mean buoyant density of nuclear membrane (1.20 g cm^–3 ), attributed to a higher content of nucleic acid and protein. (Values between 1.08 and 1.12 g cm^–3 are recorded for the density of ER without attached ribosomes in sucrose gradients, and between 1.16 and 1.18 for the mean buoyant density of 'rough' ER.) In animal cells ER and nuclear membrane appear to be identical except for the higher density of the latter (Franke, 1974). They also resembled their animal counterparts in having a higher proportion (60%) of their phospholipid as phosphatidyl choline ('lecithin') than the other membranes of the cell. Phosphatidyl ethanolamine was the second most abundant phospholipid at 20–24%. They differed from animal nuclear membrane and ER, and resembled other plant membranes in their high sterol content (ca 30%), and high content of unsaturated fatty acids as esters in the lipid. Oleate (18:1) and linoleate (18:2) accounted for 80% of these and palmitate (16:0) made up most of the rest.

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In thin sections, the outer nuclear membrane is often seen to bear ribosomes on its cytoplasmic face, just like both membranes of the ER, and when nuclei were isolated from pea shoots in high concentrations of Mg²⁺ , the outer membrane was densely covered with ribosomes (Stavy et al., 1973).

On the basis of lipid composition, enzymic activities and ability to bind ribosomes, nuclear membrane cannot be distinguished from ER. It differs morphologically in possessing nuclear pore complexes which probably contribute to its higher buoyant density.

8.3.2—
Outer Mitochondrial Membrane

The outer membrane can be released from isolated mitochondria either by a slightly hypotonic medium (osmotic lysis) or digitonin to which the inner membrane is relatively resistant. The heavy, intact inner membrane vesicles and their enzymic contents are then easily separated from the light, fragments of outer membrane.

Turnip outer mitochondrial membrane was found to have NADH-specific cytochrome c reductase and a b₅ -type cytochrome, just like the microsomal fraction, from which it differed in lacking firmly bound, NADPH-specific cytochrome c reductase (Day & Wiskich, 1975). Compared with the bulk microsomes, the inner membrane of cauliflower mitochondria had a low (40 %) proportion of its phospholipid as phosphatidyl choline due to replacement by the specialized mitochondrial phospholipid, diphosphatidyl glycerol ('cardiolipin'). The outer membrane resembled the inner membrane in that only 40% of its phospholipid is phosphatidyl choline, but was intermediate between this membrane and the microsomes in its content of diphosphatidyl glycerol (Moreau et al., 1974). Isolated plastids and mitochondria have been shown to contain less of the plant sterols sitosterol and stigmasterol than the other membranes, but about the same concentration of cholesterol in µg mg^–1 protein. As a result, the sterol content of these organelles is only one-tenth that of the bulk membranes of the cell (Hartmann et al., 1973). Mannella & Bonner (1975) estimate the ratio of sterol:phospholipid in the outer mitochondrial membrane as 0.17 for mung bean hypocotyl and 0.06 for potato tuber. These authors report that the degree of unsaturation of the fatty acids in outer membrane lipids was intermediate between that of light microsomes and the value for the highly unsaturated lipids of the inner membrane. For mung bean hypocotyl, the average number of double bonds per fatty acid was 1.3 for light microsomes, 1.7 for the outer and 2.0 for the inner membrane. The same pattern was seen in experiments on potato tuber, although in cauliflower Moreau et al. (1974) reported that the outer membrane was more saturated than the microsomal membranes.

In summary, the plant outer mitochondrial membrane has enzyme activity which shows it to be ER-like, but in its lipid composition it has characteristics intermediate between the ER and the very specialized inner membrane.

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8.3.3—
Chloroplast Envelope

This membrane is prepared from isolated chloroplasts by osmotic rupture. Chloroplast lamellae are chiefly composed of galactolipid (92% of the lipid), especially monogalactosyl diglyceride and the low content of phospholipid is largely made up of phosphatidyl glycerol. The fatty acids are highly unsaturated, with linolenate (18:3) as the major type at 83 mole %. Trans-D³ -hexadecenoate is a minor fatty acid component unique to the lamellae. The envelope was found to differ from the microsomes and resembled the lamellae in its high galactolipid content, in the complete absence of phosphatidyl ethanolamine, and relatively high proportion of linolenate (18:3). It also shared a number of features with the microsomal fraction in respect of which the envelope differed from the lamellae. These were: phosphatidyl choline as major phospholipid, digalactosyl diglyceride as the major galactolipid, and a relatively high proportion of palmitate (16:0) amongst the esterified fatty acids (Mackender & Leech, 1974).

Like the outer mitochondrial membrane, the chloroplast envelope appears to be a curious chimaera, intermediate between the ER and the specialized membrane system it encloses.

8.3.4—
Golgi Membranes

Membrane fractions enriched in Golgi bodies are prepared by density-gradient centrifugation of organelles obtained after chopping plant tissues. Glutaraldehyde is routinely added to the extraction medium to 'stabilize' the structure of the Golgi bodies though it now seems that the organelle survives preparation intact and only requires this fixative for electron microscopy (Bowles & Kauss, 1976). Golgi bodies from plant sources sediment with the fraction of buoyant density 1.12 to 1.15 g cm^–3 in sucrose gradients and are identified in gradient fractions by their distinctive morphology. Thin section studies of whole cells in higher plants show the Golgi body as a stack of 5 to 8 closed membrane sacks. Each sack, or 'cisterna', is flattened to a disc, 0.5 to 1 m m in diameter. Around the perimeter of the disc there is progressive fenestration through, and localized swelling of, the cisternal membranes, grading into isolated vesicles beyond the edge of the disc (Fig. 8.4). In some cases a single layer of parallel fibrils has been seen in the 10–15 nm wide, electronlucent region between the cisternae. Freeze-etch pictures (Fineran, 1973a) confirm this structure for the organelle. Electron-microscope sections and negative-stain images of glutaraldehyde-fixed material show that this structure is maintained in isolated organelles. The cisternae remain joined in a stack as they are in the cell, but separate in media of high ionic strength (Mollenhauer et al., 1973).

Inosine diphosphatase (IDPase) has been used as a marker for Golgi bodies and, for some plant tissues, a single peak of activity is observed on sucrose gradients, coinciding with the Golgi body-rich fraction. However, IDPase activity

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Figure 8.4
The structure of the Golgi body. Based on the
freeze-etch pictures of Fineran (1973a) which
are probably close to the real shape of this
organelle in the cell. Conventional electron
microscope methods involving fixation and
dehydration are more likely to cause distortion.

was found in purified nuclear membrane (Philipp et al., 1976), and using mung bean hypocotyl a second peak of activity was observed in light membrane characterized as ER (Bowles & Kauss, 1976).

Thin sections of Golgi body fractions from sucrose gradients show that a variety of less organized vesicles is also present, and the possibility that these preparations are contaminated with other membrane types cannot be excluded. The purest Golgi body fractions to date have been obtained from rat liver. Giving a rat ethanol induces the liver Golgi bodies to fill up with light lipoprotein particles which cause a distinctive change in the buoyant density of the organelle and serve as internal structural markers for electron-microscopy. The purified Golgi body membranes contained the cytochrome b₅ electron-transport chain (NADH-specific cytochrome c reductase and cytochrome b₅ ) characteristic of ER, but lacked NADPH-specific cytochrome c reductase and some other ER markers. UDP galactose:N-acetyl-glucosamine galactosyltransferase was restricted to the Golgi body fractions (Bergeron et al., 1973). Other evidence from animal cells indicates that fucosyl and sialyl transferases are primarily associated with the Golgi apparatus. 'Pure' Golgi fractions have not yet been prepared from plant material, but it has been shown that maximum activities of a UDP galactose:N-acetyl-glucosamine galactosyltransferase involved in glycoprotein biosynthesis (Powell & Brew, 1974), and UDP glucose:sterol glucosyltransferase (Bowles et al., 1976) were found in the Golgi body-enriched fraction of sucrose density gradients. The evidence then, such as it is, suggests that Golgi membranes have some affinity with the ER, but are specially enriched in glycosyltransferases involved in the synthesis of glycoprotein and glycolipid.

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8.3.5—
Plasma Membrane

Plasmalemma is obtained from tissue homogenates by sucrose density gradient centrifugation after removing mitochondria whose density range overlaps that of this membrane. Reported mean buoyant densities range from 1.16 to 1.18 g cm^–3 . This fraction has been identified as plasmalemma on the basis of specific staining, and characterized by the presence of Na⁺ /K⁺ stimulated ATPase activity, and glucan synthetase activity at high UDP glucose concentrations (callose synthetase), and the absence of ER and Golgi body markers (NADH-specific cytochrome c reductase and IDPase) (Leonard & Van Der Woude, 1976). Since none of the markers can be considered exclusive to the plasmalemma and non-staining membrane is always seen in the fraction, firm characterization of this membrane in plants will depend on better means of separation or identification. Investigators have tried to label the external surface of this membrane in intact cells, e.g. using radioactive iodine, hoping to trace the membrane in a gradient by its label. The most successful attempt has involved the carbohydrate-binding protein, concanavalin A (conA), which has been used to prepare plasmalemma from wall-less mutants of Neurospora and yeast protoplasts. Treatments with conA just before lysis stabilizes the membrane as large sheets and in this form it is easily separated from small vesicles. Removing the conA causes the sheets to vesiculate and now the plasmalemma can be separated from large contaminants which had co-sedimented with it (Scarborough, 1975).

Plasmalemma fractions from animal cells have a much higher content of glycoprotein and glycolipid than the other membranes of the cell and the carbohydrate moieties of these compounds form a glycocalyx over the external surface. Scarborough's work with conA suggests a similar layer is present on the surface of the fungal plasmalemma, and the specific staining of the plant plasmalemma after treatment with periodic acid, which creates reactive aldehyde groups on carbohydrate residues, implies that this is also true for the plant plasmalemma. When thin sections of protoplasts in plasmolysed onion tissue were stained for carbohydrate, only the outer surface of the plasmalemma reacted. Since this response was unaffected by treatment with concentrated cellulase and pectinase, it is unlikely that it was due to wall polysaccharide (Roland & Vian, 1971).

Bailey and Northcote (1976) have obtained a pure preparation of plasmalemma from the green alga, Hydrodictyon, by syringing out the contents of these huge cells, leaving the plasmalemma firmly attached to the wall. Since there was insufficient material for direct chemical analysis, the composition of the membrane was estimated from the radioactivity in each phospholipid after feeding [³² P_i ] to algae for 20 days, sufficient to saturate the membranes with label. Phosphatidyl choline was very much the major phospholipid, accounting for 58% of the radioactivity in identified phospholipids. Phosphatidyl serine, though only traces were found in whole cells, was a substantial component of the plasmalemma at 8%. Sterol:phospholipid molar ratios between 1.0 and 1.2

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have been recorded for plant membranes isolated on a density gradient and identified as plasmalemma-rich by staining. This represents a considerable enrichment in sterol over the total (0.4) and light membrane (0.2) fractions (Hodges et al., 1972). High sterol content will tend to reduce the fluidity of the membrane and so stabilize the cell boundary. In animal cells also phosphatidyl serine is predominantly associated with the plasmalemma and, to a lesser extent, the Golgi apparatus. Again, in animal cells, there is an increase in the proportion of sterols from ER to Golgi apparatus to plasmalemma, but they differ from Hydrodictyon in that this sequence is associated with a marked decline in the contribution of phosphatidyl choline to the phospholipids.

In summary, plasmalemma-enriched membrane fractions have a set of enzyme activities very different from that of the ER. There is some evidence that this membrane, like its animal counterpart, is more glycosylated than other membranes, from which it also differs in lipid composition.

8.4—
Functional Relationships between Membranes

8.4.1—
Membrane Synthesis

Since mitochondria and chloroplasts appear to make only a small fraction of their protein and lipid, and the plasmalemma none at all, there seems little doubt that most of this material is ultimately derived from the ER which has been shown to be the only site in the cell competent to synthesize the major membrane constituents.

Microsomal fractions from onion stem and spinach leaves synthesized phosphatidyl choline from CDPcholine and phosphatidic acid and phosphatidyl ethanolamine from CDPethanolamine and phosphatidic acid (Marshall & Kates, 1973). Beevers' group have shown that the terminal enzymes for the synthesis of phosphatidyl choline, phosphatidyl serine and phosphatidyl inositol are found only in a light membrane fraction (buoyant density 1.12 g cm^–3 ) prepared from chopped endosperm of germinating castor beans (Beevers, 1975). The whole cytoplasm, soluble enzymes as well as membranes, was centrifuged into a linear gradient containing 1mM EDTA. Electron-microscope sections showed that the membranes carried no ribosomes and they had sedimented separately. In gradients containing 3 mM Mg²⁺ , activity at 1.12 g cm^– 3 largely disappeared and the membrane, NADPH-specific cytochrome c reductase, and phosphorylcholine glyceride transferase (the terminal enzyme in phosphatidyl choline synthesis) activities were all spread around 1.16 g cm^–3 . This membrane was now coated with ribosomes. It was concluded that the final steps in the synthesis of these phospholipids are located exclusively in the ER (Lord et al., 1973). A key enzyme of sterol biosynthesis, transfarnesyl pyrophosphate-squalene synthetase, is almost all associated with fractions rich in ER (Hartmann et al., 1973).

Polyribosomes, the groups of ribosomes linked by mRNA and active in protein synthesis, occur free in the cytoplasm and attached to the surface of the

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ER and outer nuclear membrane. They can be seen in glancing sections of these membranes as folded or spiral chains of ribosomes. The potential of the ER to synthesize membrane proteins is therefore not in doubt. There is good evidence from mammalian cells that secretory proteins are synthesized by ribosomes attached to ER and pass rapidly through this membrane into the lumen (Jamieson, 1975). Liver cells synthesize protein mainly for secretion and 70 % of the polysomes are bound. Puromycin released most of the ribosomes from rough ER isolated from liver, indicating that it is the attachment of the growing peptide to the membrane which holds the ribosomes on. Now, this does not mean that proteins synthesized by rough ER are inevitably incorporated into or through the membrane. In non-secretory animal tissue-culture cells the ribosomes were bound to ER only by their attachment to mRNA. Interestingly, for rough ER from bean hypocotyls, 80% of the bound ribosomes required puromycin for release and only 10% were released by RNase alone (the others could be removed by high salt) (Dobberstein et al., 1974). Since only a few plant cell types secrete large amounts of protein, e.g. cells of the aleurone layer of cereal grains, it appears that most of the protein made by the rough ER is being incorporated into the membrane rather than through it. However, only 20 to 25% of the cytoplasmic ribosomes were recovered in the rough ER fraction isolated from bean hypocotyls (cf liver cells) and it is quite possible that membrane proteins are made on free polyribosomes and subsequently incorporated into the appropriate membrane.

To find out more about the synthesis and movement of membrane in the cell, a number of investigators have fed radioactive precursors of lipid and protein and traced the pattern of distribution of labelled product in various cell fractions.

In rat liver cells, nuclear membrane and ER incorporate a variety of [¹⁴ C]-amino acids into membrane protein with identical kinetics, and the labelling of the phosphatidyl choline of these membranes by [¹⁴ C]-choline rules out either synthesis in the nuclear membrane and transfer to the ER, or synthesis in the ER and transfer to the nuclear membrane (Franke, 1974). These results confirm the close similarity of these two membranes. No comparable findings have been reported for plants.

When [¹⁴ C]-choline was supplied to castor bean endosperm, phosphatidyl choline in the ER was labelled before that of any other membrane and part of this labelled phosphatidyl choline appears in mitochondrial membrane within a short time (Beevers, 1975). As the enzyme studies suggested, mitochondria are dependent on the ER to synthesize the bulk phospholipids of their membranes. In similar experiments with animal tissues, the specific activity of phosphatidyl choline and phosphatidyl ethanolamine in the outer membrane was intermediate between that of the ER and that of the inner membrane. This is consistent with the movement of these phospholipids from the ER to the outer membrane to the inner membrane (not a very startling idea since access to the inner membrane from outside the mitochondrion must be via the outer). The

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dependency of this organelle on the ER for phospholipids is not total. The terminal enzyme for the synthesis of phosphatidyl glycerol was found in both ER and mitochondria of castor bean endosperm. This phospholipid is a precursor for diphosphatidyl clycerol (DPG), in which the inner membrane is especially rich, and isolated animal mitochondria have been shown to synthesize phosphatidyl glycerol and DPG.

Whether the lipids of the chloroplast lamellae are made 'on site', or imported from somewhere else in the cell, is still a matter of controversy (Morré, 1975).

Another mystery is the site of synthesis of phospholipid precursors. Although fatty acid synthetase is soluble, recent work has indicated that all the activity is membrane-enclosed in the cell (Weaire & Kekwick, 1975; Donaldson, 1976), and at least part of it was found in 'heavy' organelles, proplastids and mitochondria. Certainly, when [¹⁴ C]-acetate was supplied to castor bean endosperm, lipid in organelles was labelled at the earliest measurement (5 minutes) making it unlikely that fatty acid synthesis was restricted to the ER. If mitochondria and plastids are able to synthesize some of their own fatty acids, this could explain how they maintain a high level of unsaturation against the influx of lipid from the ER, since phospholipid molecules in membranes are dynamic structures in which acyl moieties can be replaced or exchanged. In addition it is possible that the fatty acids of ER-derived phospholipid could be desaturated in the plastid or mitochondrion. Desaturases are always intimately associated with membrane systems since the participation of a special electron transport chain is required, and phospholipid appears to be the preferred substrate for the production of linoleate (18:2) from oleate (18:1) in Chlorella (Sedgwick, 1972).

Independent, internal synthesis of special components, e.g. phosphatidyl glycerol and unsaturated fatty acids, is one way the distinctive composition of organelle membranes may be maintained. Another way would be to use the intermembrane space, e.g. between inner and outer mitochondrial membranes, to control the movement of individual lipid types. The differences in lipid composition between these two membranes support the idea and there are reports of 'exchange' proteins in the soluble cytoplasm which can ferry specific phospholipids in solution between membranes (McMurray & Dawson, 1969; Morré, 1975). The fact that the ER and the outer membrane differ in composition suggests that lipid must also be 'filtered' as it passes between these two membranes. For this reason, if no other, it is unlikely that the ER is structurally continuous with these outer membranes in the cell. Although movement between the two halves of the bimolecular leaflet is restricted (see chapter 2), lipids and many membrane proteins are freely mobile in the plane of the membrane and any differences in composition would soon disappear. Conventionally prepared, electron microscope sections have provided occasional observations of interconnections between all the membrane types of the cell, but phenomena seen only infrequently by electron microscopy should be treated with suspicion. Membrane conformations are stabilized by weak bonds, and the covalent cross-linking of proteins by glutaraldehyde could force lipidic structures to

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re-organize to create continuities, for example where two membranes were closely appressed. For certain species, e.g. the fern Pteris, an unusually large number of conjoinings between membranous organelles has been reported (Crotty & Ledbetter, 1973) which makes it unlikely that these observations have universal significance. The close similarity, biochemically, between nuclear membrane and ER would support observations that these two membranes are in clear luminal continuity, but can also be explained as the result of the total re-structuring of the nuclear membrane from ER at every mitosis. It has been suggested that membrane could be transferred from the ER to the outer organelle membranes by intermittent direct connections, and lipid components might also move in solution on 'carrier' proteins (Morré, 1975).

From thin section studies, electron microscopists suggested that the ER and the plasmalemma are linked functionally via the Golgi apparatus, the cell's complement of Golgi bodies. Golgi bodies can be seen adjacent to a sheet of ER or the outer nuclear membrane, aligned with the cisternae parallel to the plane of these membranes. The portion of membrane apposed to the Golgi body bears no ribosomes but projects as small tubules into the cytoplasm between. It is hypothesized that the tubules vesiculate and the 'primary' vesicles formed then re-fuse to form a new Golgi body cisterna (Morré & Mollenhauer, 1974; Gunning & Steer, 1975). Also, circular membrane profiles, similar to those in the vicinity of the vesiculated rim of the cisternae, are found close by and apparently fused with the plasmalemma. The membrane of these 'secretory' vesicles is stained heavily after treatment of sections with barium permanganate and the plasmalemma stains in the same way. ER does not stain heavily. The membranes of the Golgi cisternae show a progressive increase in staining intensity from ER-like at one pole to plasmalemma/'secretory' vesicle-like at the other (Grove et al., 1968). There is, then, ultrastructural evidence to suggest that the Golgi apparatus is linked to both ER and plasmalemma by alternating membrane vesiculation and fusion, and some aspect(s) of the transition between ER and plasmalemma membranes takes place within the Golgi apparatus. Of course, equivalent fiffinity for heavy metal is no real indication of biochemical similarity, no more than the intermediate density of the Golgi body can be considered as evidence for its function as an intermediate between ER and plasmalemma.

After feeding labelled precursors, the specific radioactivity of phospholipid in membrane fractions was reported to be highest in ER, intermediate in Golgi bodies and lowest in plasmalemma (Morré, 1975), a pattern consistent with the proposed function of the Golgi body.

Studies of the incorporation of a pulse of radioactive amino acid, [guanido-¹⁴ C]-arginine, into membrane protein in rat liver have shown that membrane protein in a 'rough microsomal' fraction was labelled very early on. The kinetics of labelling suggested that some membrane protein passed from ER to Golgi apparatus to plasmalemma. Thus, the rough ER fraction reached maximum specific activity within 10 minutes of feeding. By 20 minutes the

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specific activity had fallen by one-fifth, as if part of the newly labelled protein had left the fraction. At the same time the specific activity of a Golgi bodyenriched fraction, already higher than that of the ER, continued to rise reaching a maximum at 30 minutes and declining until 60 minutes to five-sixths the maximum value. Now, between 10 and 60 minutes, the membrane protein of a plasmalemma fraction accumulated label in two phases, firstly as label was lost from the rough ER and then as label disappeared from the Golgi fraction (Fig. 8.5, Franke et al., 1971). These results indicate that:

Figure 8.5
Changes in the specific activity of membrane proteins in organelle
fractions from rat liver after feeding a pulse of [guanido-¹⁴ C] arginine.

———

Golgi body-enriched fraction

. . .

microsomes
× - - - - × plasmalemma-enriched fraction (from Franke et al., 1971.)

(a) proteins of the ER may contribute to the plasmalemma without the mediation of the Golgi apparatus,

(b) only a part of the newly labelled ER and Golgi apparatus membrane protein turns over sufficiently rapidly to be able to give rise to the observed increase in plasmalemma labelling,

(c) if the newly labelled material of the Golgi apparatus is derived from the ER (and this is by no means obvious from the data since there is no lag between incorporation into ER and into Golgi apparatus fractions) then it must be transferred from a region of synthesis and rapid turnover within the bulk ER since the specific activity of the Golgi apparatus was higher than that of the ER fraction.

Similar experiments have been performed with onion stem tissue, feeding [U-¹⁴ C]-leucine (Morré & Van Der Woude, 1974). In this case a lag of 30 to 60 minutes was observed after incorporation into microsomal and nuclear membranes had begun and before Golgi apparatus and plasmalemma fractions

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began to be labelled. The authors discern a second phase of incorporation into plasmalemma, lagging 30 minutes behind the initial simultaneous labelling of Golgi apparatus and plasmalemma. Their results are consistent with transfer of newly synthesized protein from microsomal to Golgi membranes in plant cells and, as in rat liver, the ER may contribute material directly as well as via the Golgi apparatus. Morré and Van Woude assumed the label in membrane fractions represented incorporation into membrane protein only. Though the cells of onion stem do not secrete protein on the same scale as rat liver, this assumption is not entirely justified. A hydroxyproline-rich glycoprotein is a substantial component of cell walls and its synthesis probably takes place in the ER and the Golgi apparatus (Gardiner & Chrispeels, 1975).

If, as this work suggests, the bulk of the membrane protein in the ER and the Golgi bodies is not transferred, this would explain how these membranes and the plasmalemma maintain distinctive enzyme activities.

However, there is considerable evidence that the membranes in the Golgi body stack are moving rapidly through it. Certain unicellular algae, e.g. Prymnesium and Chrysochromulina, have only one Golgi body per cell. By way of compensation, there may be as many as 30 or more cisternae per stack. The organelle elaborates complex scales which coat the outer surface of the alga and the polarity of the stack is obvious from the gradation in complexity of scale precursors seen inside the cisternae in thin sections (Manton, 1966). This is the most direct evidence that whole cisternae are shuttled along the stack in one direction, from the ER, which apposes the immature ('proximal') end of the stack, to the plasmalemma with which the cisternae full of mature scales fuses. The Golgi body in higher plants and animals differs from that of these algae in that the cisternae vesiculate before fusing with the plasmalemma, but evidence from higher organisms also suggests the whole stack turns over. The number of cisternae per stack decreases in starvation or when 'secretory' vesicle production is stimulated and increases when vesicle production is inhibited (Morré et al., 1971). The alteration in staining properties across the Golgi stack affects the whole cisternal membrane and not just the peripheral vesiculations (Grove et al., 1968), and in isolated Golgi bodies this property spreads within a short time to the ER-like cisternae of the stack so that all the membrane resembles plasmalemma (Frantz et al., 1973).

Golgi-specific enzyme proteins could be retained in the organelle while the lipidic sack and plasmalemma-specific proteins pass on if there is movement of membrane protein between cisternae. The centre of the cisterna where it is closely appressed to its neighbours is the most likely site for this activity and the intercisternal space here is filled by a lipoprotein plaque (Mollenhauer et al., 1973), a hydrophobic environment in which lipophilic membrane proteins would be mobile. It is apparent from the staining evidence and the intermediary status of the Golgi body between ER and plasmalemma that there is a gradient in the concentration of substrate, protein and lipid, for the membrane-modifying glycosyltransferases of the Golgi, and the cisternal membrane at the proximal

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pole of the stack has the highest concentration. Substrate binding alone could ensure that the net movement of Golgi enzymes is in the direction of the proximal pole. The glycosylation of the products, Golgi body-modified plasmalemma-specific proteins and lipids, would make them too hydrophilic to move rapidly between cisternae in this hypothetical scheme.

8.4.2—
Synthesis and Secretion of Extracellular Material

While investigations into the function of the Golgi apparatus in animals have concentrated on protein glycosylation and other membrane modifications, plant biochemists have been more concerned with a role for this organelle in polysaccharide biosynthesis.

Gardiner and Chrispeels (1975) have presented evidence that the glycosylation of the hydroxyproline-rich glycoprotein takes place in the Golgi apparatus. The completed polymer is a cell wall rather than a plasmalemma component and carries arabinosyl side-chains on its hydroxyproline residues. Organelle fractions were prepared on density gradients from carrot root tissue pulse-labelled with [¹⁴ C]-proline. Most of the label in hydroxyproline was associated with a Golgi body-enriched fraction. This fraction coincided with peak activity in the gradient of an enzyme which transferred arabinosyl residues from UDP-arabinose onto endogenous protein acceptors. (It should be noted, though, that the gradients used to prepare organelles contained glutaraldehyde and the distribution of enzyme activities on glutaraldehyde-free gradients is not strictly comparable).

From studies of thin sections, electron-microscopists proposed that the Golgi apparatus was involved in the synthesis and secretion of cell wall polysaccharides. The increase in luminal volume at the cisternal periphery was thought to represent the polymerization or decantation of a batch of wall material which travelled to the plasmalemma in a secretory vesicle and was discharged into the wall as the vesicle membrane fused with the plamalemma. This evidence has been critically reviewed by O'Brien (1972).

Much of the biochemical evidence has come from work on the root cap. It seems that to follow polysaccharide biosynthesis by autoradiography requires the extremely high rates of uptake and incorporation found only in certain differentiating or differentiated cells. The root cap is a very rapidly growing and metabolizing tissue—the entire cap of ca 10, 000 cells in maize is replaced every 24 hours. The cap cells secrete large quantities of slime to ease the passage of the cap through the soil. This material is a hydrated polysaccharide with a chemical composition similar to the matrix components of primary cell walls, pectin, but the structure of the polymer has been modified to reduce gelation, e.g. maize root slime-.polysaccharide has a substantial content of residues of fucose, a sugar absent from maize root pectin. Slime producing root cap cells each have several hundred Golgi bodies (root parenchyma cells have only ca 30) with a characteristic 'hypertrophied' morphology: the vesiculations of the cisternal periphery are very swollen.

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Northcote and Pickett-Heaps (1966) fed [6-³ H]-glucose to wheat roots and analysed the pattern of incorporation of this label into polymeric material of root cap organelles by high resolution autoradiography. After 5 minutes of feeding, silver grains were confined mainly to the immediate vicinity of the Golgi bodies. By 10 minutes, both Golgi bodies and secretory vesicles were labelled, but very little radioactivity was associated with the wall. Subsequent incubation of labelled roots in non-radioactive glucose solution for 10, 30 and 60 minutes provided a time-series of sections showing progressive loss of radioactivity from the Golgi bodies and secretory vesicles and its accumulation outside the plasmalemma (Fig. 8.3). Chemical analysis of the high molecular weight polysaccharide in the root cap after 15 minutes exposure to radioactive glucose showed that more than 70% of the label in this material was in galactosyl residues. Galactose occurs only in pectin-type polymers in angiosperm cell walls and is the only major unit of these polysaccharides to retain a hydrogen atom on carbon 6. Since less than 3% of the polysaccharide label was in glucose units of a -cellulose at this time, the labelled material represents slime-polysaccharide or matrix components of the wall.

This is strong evidence that the Golgi apparatus is responsible for the secretion of pectin-type polymers. Though high molecular weight material appeared first of all in the Golgi apparatus, this does not establish that it was synthesized here. The early stages in polymerization must involve oligosaccharides which may be lost from the sections.

The differing functions of ER and Golgi apparatus in the synthesis and secretion of slime-polysaccharide in the root cap have been investigated by Bowles and Northcote (1972), and compared with the synthesis and secretion of the ordinary matrix components of the wall in the rest of the root. They fed [U-¹⁴ C]-glucose to maize roots and, taking different parts of the root, prepared organelle fractions by differential and 'step' density gradient centrifugation. These were characterized from thin sections. Radioactive polysaccharide was found in a rough ER fraction and a fraction rich in Golgi bodies. Chemically, the labelled material in membranes from the root tip resembled slime polysaccharide and that in membranes from mature root tissue had the composition of the cell wall matrix. It could have been slime or pectin from the wall which was mixed with and bound to, or enclosed in, membranes when the tissue was chopped. To test this, membrane fractions were prepared from unlabelled tissue chopped in an extraction medium with radioactively labelled, soluble slime and wall components. The membrane fractions were essentially unlabelled.

When [U-¹⁴ C]-glucose was fed to maize roots over times ranging from 5 to 90 minutes, radioactivity was incorporated steadily into slime, wall and membrane polysaccharide up to 30 minutes. After this time the slime and wall continued to accumulate label at the same rate but there was no further increase in the radioactivity of each of the sugar residues of the polysaccharide in membranes, confirming that these polymers are precursors of the slime and wall matrix (Bowles & Northcote, 1974).

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Bowles and Northcote (1976) investigated this precursor material after labelling it to saturation from supplied [U-¹⁴ C]-glucose. Some of the polysaccharide was freely soluble with a high molecular weight (>40,000), most of the rest was so firmly membrane-bound it required protease digestion for release. These types were found in both membrane fractions, but the Golgi body-rich fraction had more of the former, whereas the ER had much more of the latter. Most of the membrane bound polysaccharide of the ER was as short chains (MW < 4,000), but all the polysaccharide segments bound to Golgi body membranes had molecular weight greater than 4,000. Interestingly, though all the other sugars of slime-polysaccharide were found in labelled residues of the short chains, fucose, which always occupies a terminal position on pectin side-chains, was absent.

The earliest polymeric precursors of polysaccharides will have the lowest molecular weights and it seems from this work that they are membrane-bound. The glycosyltransferases involved in chain-extension during pectin synthesis are also all bound to membranes (Villemez et al., 1965; McNab et al., 1968; Odzuck & Kauss, 1972). This early precursor is associated chiefly with the ER. Chain extension, addition of terminal fucosyl groups to slime polysaccharide and release from the membrane occur progressively as this material leaves the ER and passes through the Golgi apparatus and there is no evidence that synthesis is restricted to a particular section of the endomembrane complex.

ER fractions for both cap and mature root contained much more radioactive polysaccharide than the equivalent Golgi body fraction. However, since a whole-root ER fraction had 40 times as much lipid, the radioactivity in polysaccharide per unit quantity of membrane (i.e. weight of lipid) for the Golgi body-rich fraction was twice the value for ER. This could be because a smaller fraction of the ER is devoted to polysaccharide synthesis, or because the polymer chains are longer in the Golgi body, or both.

With the biosynthetic machinery saturated from [¹⁴ C]-glucose, Bowles and Northcote (1974) compared the rate of production of radioactive wall and slime with the steady-state levels in membrane polysaccharide. This enabled them to calculate the rates of turnover of polysaccharide in the membrane compartments expected for different models of secretion. For example, labelled wall polysaccharide is produced at 2,000 cpm per minute and the steady state level of radioactivity in wall polysaccharides in the Golgi bodies in 5,000 cpm. Therefore the entire polysaccharide content of the Golgi body will be replaced every 2.5 minutes if all the wall material has to pass through the Golgi apparatus.

Supposing polysaccharide and membrane sack move through the stack as a single entity, then for the average stack of 5 to 6 cisternae, one cisterna is released roughly every 0.5 minutes. This seems very fast, but it is remarkably close to values determined microscopically in other plants. Working with the Chrysophycean alga Pleurochrysis, Brown (1969) showed by time-lapse cinephotomicrography that the single Golgi body and all the other organelles rotate inside the cell wall, 360° in 15 to 20 minutes. Now the lateral displacement

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observed in thin sections between successive cisternae released from the Golgi body was 15°, indicating that the time between the release of successive cisternae is about 0.75 minutes. Schnepf (1961) measured the volume of slime produced by glands on the leaves of the carnivorous plant Drosophyllum lusitanicum over set time periods. From the size and number of Golgi vesicles in the gland, he estimated that the rate of production of vesicles by each Golgi body necessary to maintain the observed rate of secretion was 3 per minute at 28°C.

Now there is good reason to suspect that 2 cisternae per min is an over-estimate of the speed required to shift all the matrix to the wall via the Golgi apparatus in maize roots. Firstly, while the recovery of slime and wall components is probably close to 100%, the recovery of Golgi bodies is certainly much lower and so the steady-state level of radioactivity in this organelle is underestimated. Secondly, the wall fraction from mature root tissue was shown to contain a large amount of labelled glucose polymer which was only a minor component in the membrane fractions. It could be cellulose or contaminant starch, but it means that the measured rate of increase in polysaccharide label of the wall fraction is probably an overestimate of the rate of production of wall matrix by the endomembranes. Nevertheless, observed rates of turnover of cisternal stacks can account even for this overestimate of the rate at which polysaccharide would have to pass through the Golgi body and there is no need, therefore, to invoke secretion via the vesiculating cisternal periphery independent of the turnover of the stack, or transfer direct from ER to the wall.

Similar calculations indicate that if slime-polysaccharide is secreted only via Golgi bodies, the entire polysaccharide content of these organelles is displaced every 20 seconds, a figure corrected for cellulose. The changing pattern of label in polysaccharide monomers after feeding [¹⁴ C]-glucose for different times confirmed that slime-polysaccharide was synthesized faster than wall matrix polymers. Some of it had been secreted from the cells within 2 minutes of supplying the labelled precursor. These results pay tribute to the furious synthetic activity of root cap tissue. Even using conservative estimates of the rate of turnover of cisternae, Morré et al. (1971) calculate that secretory vesicles contribute enough new membrane to the plasmalemma of a maize root cap cell to replace it entirely every 4 to 8 hours. How the excess is recycled is not known. Careful studies of Golgi bodies and numbers of secretory vesicles in thin sections of maize root caps fixed at different times have shown the existence of rhythmic fluctuations in secretory activity. The organelles reach a peak of activity every 3 hours, synchronized over the whole cap. Whole batches of roots can be synchronized by transfer to fresh solution, which induces a peak of activity at around 1 hour later (Morré et al., 1967). Since the results obtained by Bowles and Northcote refer to the first three quarters of an hour after transfer of roots to [¹⁴ C]-glucose solution, they probably represent peak activity for root cap Golgi bodies.

Though still in debate, it now seems unlikely that the cellulose microfibrils of the wall are synthesized in the ER or Golgi apparatus in higher plant cells

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(see also chapters 1 and 7). Low incorporation into glucose relative to other wall monomers in the polysaccharides of ER and Golgi body-rich fractions of maize root confirms the earlier autoradiographic evidence from developing xylem vessels. In this tissue only the plasmalemma was labelled from [³ H]-glucose at the time it was being incorporated exclusively into cellulose (Wooding, 1968). However, Golgi body-enriched membrane fractions synthesized radioactive b -1,4 glucan when supplied with UDP-[¹⁴ C]-glucose (Van Der Woude et al., 1974; Shore & MacLachan 1975). The b -1,4 glucans synthesized from UDP-glucose have been shown to be cellulose and not b -1,4 glucan sections in glucomannan, a matrix component synthesized from GDP sugars (Villemez, 1974). Golgi body cellulose synthetase showed distinctive kinetics when compared with the activity in a plasmalemma-enriched fraction, and was much less active than the plasmalemma enzyme at high (1 mM ) substrate concentration. It seems, then, that the activity in Golgi bodies was not due to contamination by plasmalemma and that the Golgi apparatus may be ferrying the enzyme to the plasmalemma in a less active form. A similar mechanism has been shown for chitin synthetase in yeast and Mucor. Chitin is structurally similar to cellulose and is produced as a microfibril on the outer surface of the plasmalemma by an enzyme particle which spans this membrane. Chitin synthetase is made in an inactive zymogen form, found in microsomal membranes, which can be converted to an active enzyme by protease digestion. An inhibitor of protease found in the soluble cytoplasm is thought to control this transition, preventing the activation of the enzyme: before it reaches its operational site in the plasmalemma (McMurrough & Bartnicki-Garcia, 1973; Durán et al., 1975). The cellulose component of the scales of Chrysophycean algae appears to be made in Golgi cisternae, but this difference between them and higher plants may well amount to nothing more fundamental than the timing of cellulose synthetase activation.

8.4.3—
Differentiation

Inevitably the elements of the endomembrane complex are heavily involved in the differentiation of plant cells. Normal cell growth and development require the biosynthesis of intracellular membrane and extracellular wall material, both of which involve the endomembranes. When a cell differentiates the specialized nature of these products must be the result of altered biosynthetic activity of the endomembrane complex in response to information received, ultimately, from the nucleus. For example, by comparison with a parenchyma cell of the same root, slime-producing root cap cells synthesize more Golgi body membrane, more polysaccharide, and the biosynthetic machinery includes different enzymes, e.g. fucosyltransferases.

As the source of bulk membrane components the ER plays a unique role in the development of specialized organelles in differentiating cells. Undoubtedly the best characterized example is the development of glyoxysomes in castor bean. These organelles are found only in endosperm tissue and develop between

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2 and 4 days after germination, by de novo synthesis of protein and lipid. Glyoxysomes do not contain the enzymes for phospholipid synthesis. When [¹⁴ C]-choline is supplied to endosperm tissue, the first membrane fraction to be labelled is the ER and this label subsequently appears as phosphatidyl choline in the glyoxysome membrane. In the early stages of germination, glyoxysomal enzymes can be detected in the ER fraction (Beevers, 1975). Because there is no evidence that glyoxysomes can synthesize protein, the kinetics of incorporation of radioactive amino acid into these organelles was interpreted as the result of synthesis in the ER followed by transfer to the glyoxysome. Thus the incorporation of [³⁵ S]-methionine into glyoxysomes lagged 30 minutes behind the labelling of the ER. During a 'chase' with unlabelled methionine, the radioactivity in ER membranes decreased at the same time as the glyoxysome membrane continued to accumulate label (Bowden & Lord, 1976). In thin sections, the coincidence of a sheet of ER abutting end-on to a developing microbody is reported as frequent, both for glyoxysomes and for leaf peroxysomes. Although their membranes are seen to touch, the lumina of the two organelles appear quite separate. When segments of juvenile leaves are incubated in a solution of 3,3'-diaminobenzidine and H₂ O₂ (to localize catalase activity) and prepared for electron microscopy, the developing microbodies show a strong electron-opaque deposit but no such product can be detected in the contiguous ER (Gruber et al., 1973).

From electron-microscope sections, the vacuole of plant cells is thought to begin with the distension of portions of ER to form numerous 'pro-vacuolar' vesicles which then coalesce and further expand. A combined thin section and freeze-etch study of later stages in vacuole formation showed that each expanding pro-vacuole had a sheet of fenestrated ER lying close to its bounding membrane over the whole external surface (Fineran, 1973b).

These observations suggest that the microbodies and the vacuole develop in two phases. A primary event is the formation of a separate compartment from part of the ER. This pro-organelle must already differ from the bulk ER because its subsequent development is so different, but it is not known to what extent any of the components for its specialized function are present at this stage. Further development of this structure appears to be by transfer of lipid and protein from apparently undifferentiated ER close to or touching its bounding membrane.

Some correlations between the distribution of the ER and spatial patterns of wall biosynthesis in differentiating cells have suggested a morphogenetic role for this membrane. For example, in the differentiation of sieve-tubes, the siting of the sieve-pores in the end walls is pre-patterned by plates of ER, and in the differentiation of xylem vessels sheets of ER cover the internal surface of the plasmalemma between wall thickenings (Northcote, 1968). In both cases, the result of this pattern is a reduced deposition of secondary wall against the section of plasmalemma persistently overlain by ER. In many thin section electron micrographs of plant cells, large areas of plasmalemma appear to be covered closely by a continuous sheet of ER and it is difficult to see how

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secretory vacuoles could get to the plasmalemma. Figure 8.3 shows secretory vacuoles apparently pouring through a gap in such a barrier. Juniper and Pask (1973) have shown that the deposition of slime polysaccharide is restricted to the outer tangential wall of maize root cap cells and their micrographs show that fenestration of the peripheral ER is most apparent on this side of the cell. The ER, therefore, may not only produce the polysaccharide of the wall but also control the site of its deposition. How the ER is organized in the correct pattern is a central problem of cell morphogenesis. Certainly the membrane is well placed to receive and is competent to interpret information from the nucleus and the possibility that it also 'responds' to chemical information from other organelles or outside the cell cannot be discounted.

The role of endomembrane complex in differentiation is undoubtedly not restricted to the 'motor' function of assembling new structures in response to a changing supply of nucleus-derived information. The effects of plant growth regulators in the tissue environment of the cell on nuclear activity must be mediated by the cytoplasm and, because of its position in the front line, the plasmalemma is strongly implicated. We know virtually nothing about the mechanisms involved in this 'sensory' function of the endomembrane complex. The first indication of the nature of this interaction may be the discovery that RNA polymerase in plants is activated by a factor released from plasmalemma by auxin (Cherry, 1974, see also chapter 13).

8.5—
Conclusions

The working definition of the endoplasmic reticulum is essentially negative. All the membrane of the cell not obviously specialized is relegated to this category. Biochemical investigation of its activities, stimulated and supervised by electron microscopy, has gone some way toward explaining the low level of its organization and replacing the negative definition with something more positive. The ER is the meristem of the cell. It is membrane in the process of synthesizing membranes, creating a pool of components and distributing them about the cell for the maintenance and enlargement of the specialized organelles. For this purpose the outer mem branes of plastids, mitochondria and the nucleus can be considered as rather more specialized 'limbs' of the ER, the two former membranes acting as placentas for the very specialized membranes they enclose. The new protein and lipid components must somehow cross the placental gap in a solubilized form. Material from the ER apparently contributes to the expansion of the tonoplast, microbodies and the plasmalemma in the same way (Fig. 8.6).

However, the glycoproteins, glycolipids and polysaccharides intended for the plasmalemma and wall are not transferred in this way. Probably they are too hydrophilic to be shuttled in and out of different sides of lipid bilayers. Once synthesized, they never leave the membrane (and its lumen) which is incorporated as a unit into the plasmalemma (and the wall). This vesicular transfer cannot

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Figure 8.6
Diagram of a plant cell early in development showing the relationships
between the ER and the other membranes of the cell proposed
from electron microscope and biochemical evidence.

provide the selectivity of carrier proteins. Therefore all the membrane which is subsequently incorporated into the plasmalemma by fusion passes through a phase of close appression (the Golgi body) to allow the selective redistribution of membrane components.

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In that the endomembrane complex is the cell's machinery for organelle and wall synthesis, the specialized nature of these ER products in differentiated cells can only come about as a result of reprogramming the synthetic machinery. Essentially it is the endomembrane complex which builds the plant cell and the nucleus which stores, selects and supplies designs for the appropriate tools.

SECTION ONE— PLANT CELL STRUCTURE AND FUNCTION

Introduction

Further Reading

Chapter 1— Plant Cell Walls

1.1— Introduction

1.2— The Molecular Structure of Plant Cell Walls

1.2.1— Cellulose

1.2.2— Hemicelluloses

1.2.3— The Pectic Polysaccharides and Structural Protein

1.2.3.1— Rhamnogalacturonans

1.2.3.2— Arabinogalactans

1.2.3.3— Structural Protein

1.3— Interconnections between Cell Wall Components

1.4— The Universality of Plant Cell Wall Structures

1.5— Cell Wall Plastics

1.5.1— Lignin

1.5.2— Cutin

1.6— Biosynthesis of Plant Cell Walls

1.7— Cell Wall Ultrastructure

1.7.1— Cellulose Microfibrils

1.7.2— Specialized Structures

1.8— Hormonal Control of Cell Wall Biosynthesis and Differentiation

1.9— The Role of the Plant Cell Wall in Interactions with Other Organisms

Further Reading

Chapter 2— Membrane Structure and Transport

2.1— Introduction

2.2— Chemical Composition of Membranes

2.2.1— Lipids

2.2.1.1— Phospholipids

2.2.1.2— Glycolipids

2.2.1.3— Sterols

2.2.2— Proteins

2.2.3— Water

2.3— Membrane Structure

2.3.1— The Membrane Matrix

2.3.1.1— Phospholipids

2.3.1.2— Sterols

2.3.1.3— A Model of the Membrane Matrix

2.3.2— Membrane Sub-Structure

2.3.2.1— A Model of Membrane Structure

2.3.2.2— Membrane Fluidity

2.3.2.3— Membrane Synthesis and Flow

2.4— Transport of Substances Across Membranes

2.4.1— Passive ('Downhill') Transport

2.4.2— Criteria For Active ('Uphill') Transport

2.4.3— The Nature and Origin of the Membrane Potential

2.4.3.1— Diffusion Potential

2.4.4— Membrane Pumps

2.4.4.1— Neutral Ion Pumps

2.4.4.2— Electrogenic Pumps

2.4.5— Membrane Carriers

2.4.5.1— Evidence from Kinetics

2.4.5.2— Ionophores as Lipophilic Carriers

2.4.5.3— Co-transport

2.5— Correlation of Structure and Function

2.5.1— Membrane Pores and Channels

2.5.2— Ion Pumps and Membrane Substructure

2.5.3— Endocytosis and Vesicular Transport

2.5.4— Concluding Remarks

Further Reading

Chapter 3— Chloroplasts—Structure and Development

3.1— Introduction

3.2— Chloroplast Structure

3.2.1— Chloroplast Dimensions and Number

3.2.2— Chloroplast Fine-Structure

3.2.3— The Mobile and Stationary Phases of Chloroplasts

3.2.4— The Range of Chloroplast Structure

3.2.4.1— The Algae

3.2.4.2— The Dimorphic Chloroplasts of C4 Plants

3.2.4.3— The Effect of Environmental Conditions on Chloroplast Structure

3.2.4.4— Chloroplast Mutants

3.3— Isolation of Chloroplasts

3.4— Chloroplast Development

3.4.1— The Proplastid

3.4.2— Etioplast Formation

3.4.3— The Conversion of Etioplasts into Chloroplasts

3.4.4— The Formation of Chloroplast Components in Greening Leaves

3.5— Summary

Further Reading

Chapter 4— Chloroplasts—Structure and Photosynthesis

SECTION ONE—
PLANT CELL STRUCTURE AND FUNCTION

Chapter 1—
Plant Cell Walls

1.1—
Introduction

1.2—
The Molecular Structure of Plant Cell Walls

1.2.1—
Cellulose

1.2.2—
Hemicelluloses

1.2.3—
The Pectic Polysaccharides and Structural Protein

1.2.3.1—
Rhamnogalacturonans

1.2.3.2—
Arabinogalactans

1.2.3.3—
Structural Protein

1.3—
Interconnections between Cell Wall Components

1.4—
The Universality of Plant Cell Wall Structures

1.5—
Cell Wall Plastics

1.5.1—
Lignin

1.5.2—
Cutin

1.6—
Biosynthesis of Plant Cell Walls

1.7—
Cell Wall Ultrastructure

1.7.1—
Cellulose Microfibrils

1.7.2—
Specialized Structures

1.8—
Hormonal Control of Cell Wall Biosynthesis and Differentiation

1.9—
The Role of the Plant Cell Wall in Interactions with Other Organisms

Chapter 2—
Membrane Structure and Transport

2.1—
Introduction

2.2—
Chemical Composition of Membranes

2.2.1—
Lipids

2.2.1.1—
Phospholipids

2.2.1.2—
Glycolipids

2.2.1.3—
Sterols

2.2.2—
Proteins

2.2.3—
Water

2.3—
Membrane Structure

2.3.1—
The Membrane Matrix

2.3.1.1—
Phospholipids

2.3.1.2—
Sterols

2.3.1.3—
A Model of the Membrane Matrix

2.3.2—
Membrane Sub-Structure

2.3.2.1—
A Model of Membrane Structure

2.3.2.2—
Membrane Fluidity

2.3.2.3—
Membrane Synthesis and Flow

2.4—
Transport of Substances Across Membranes

2.4.1—
Passive ('Downhill') Transport

2.4.2—
Criteria For Active ('Uphill') Transport

2.4.3—
The Nature and Origin of the Membrane Potential

2.4.3.1—
Diffusion Potential

2.4.4—
Membrane Pumps

2.4.4.1—
Neutral Ion Pumps

2.4.4.2—
Electrogenic Pumps

2.4.5—
Membrane Carriers

2.4.5.1—
Evidence from Kinetics

2.4.5.2—
Ionophores as Lipophilic Carriers

2.4.5.3—
Co-transport

2.5—
Correlation of Structure and Function

2.5.1—
Membrane Pores and Channels

2.5.2—
Ion Pumps and Membrane Substructure

2.5.3—
Endocytosis and Vesicular Transport

2.5.4—
Concluding Remarks

Chapter 3—
Chloroplasts—Structure and Development

3.1—
Introduction

3.2—
Chloroplast Structure

3.2.1—
Chloroplast Dimensions and Number

3.2.2—
Chloroplast Fine-Structure

3.2.3—
The Mobile and Stationary Phases of Chloroplasts

3.2.4—
The Range of Chloroplast Structure

3.2.4.1—
The Algae

3.2.4.2—
The Dimorphic Chloroplasts of C₄ Plants

3.2.4.3—
The Effect of Environmental Conditions on Chloroplast Structure

3.2.4.4—
Chloroplast Mutants

3.3—
Isolation of Chloroplasts

3.4—
Chloroplast Development

3.4.1—
The Proplastid

3.4.2—
Etioplast Formation

3.4.3—
The Conversion of Etioplasts into Chloroplasts

3.4.4—
The Formation of Chloroplast Components in Greening Leaves

3.5—
Summary

Chapter 4—
Chloroplasts—Structure and Photosynthesis

4.1—
Introduction

4.2—
Thylakoid Structure

4.2.1—
Thin Sectioning and Heavy Metal Shadowing

4.2.2—
Freeze-Etching

4.2.3—
Negative Staining