No title

Advances in BOTANICAL RESEARCH VOLUME 10 This Page Intentionally Left Blank Advances in BOTANICAL RESEARCH Edited...

Author: Author Unknown

7 downloads 301 Views 16MB Size Report

This content was uploaded by our users and we assume good faith they have the permission to share this book. If you own the copyright to this book and it is wrongfully on our website, we offer a simple DMCA procedure to remove your content from our site. Start by pressing the button below!
Report copyright / DMCA form

DOWNLOAD PDF

Advances in

BOTANICAL RESEARCH VOLUME 10

This Page Intentionally Left Blank

Advances in

BOTANICAL RESEARCH Edited bj1

H. W. WOOLHOUSE John Innes Institute Norwicli, Englund

VOLUME 10

1983

ACADEMIC PRESS, INC. (Harcourt Brace lovanovich. Publishers)

London Orlando San Diego New York Toronto Montreal Sydney Tokyo

ACADEMIC PRESS INC. (LONDON) LTD. 24/28 Oval Road, London NWl 7DX

U S . Edition published by ACADEMIC PRESS, INC.

Orlando, Florida 32887

Copyright

8 1983 by Academic Press Inc. (London) Ltd

AN Rights Reserved No part of this book may be reproduced in any form by photostat, microfilm, or any other means, without written permission from the publishers

ISBN 0-12-005910-X

PRINTED IN THE UNITED STATES OF AMERICA

84 85 86 87

9876 5 432

CONTRIBUTORS TO VOLUME 10 J. BARRETT, Department of Plant Industry, CSIRO, Canberra, ACT 2601, Australia R. D. GRAHAM, Department of Agronomy, Waite Agricultural Research Institute, The University of Adelaide, Glen Osmond, South Australia 5064 A. W. D. LARKUM, School of Biological Sciences, University of Sydney, NSW 2006, Australia


PREFACE The papers in this volume both come from Australia. In the first Larkum and Barratt consider light-harvesting mechanisms in algae. This is a subject of astonishing complexity for, as the authors point out, the algae have evolved a much greater variety of mechanisms for the capture and conservation of light energy than have other groups of plants. The reasons for this diversity are to be found in the light climate, which is considered in an early section of the review. The authors then explore the structure and function of the photosynthetic membranes. From the membrane biochemistry their analysis leads on to the photosynthetic pigments, the reaction centre complexes, pigment-protein complexes involved in light harvesting and the principles of the light-harvesting process. The authors conclude with an analysis of the adaptive and evolutionary aspects of the light-harvesting systems of algae. The article illustrates an important principle followed by successiveeditions of these reviews, that of allowing the authors freedom to explore their subject in full and consider the wider implications. The article by R. D. Graham sits interestingly between plant pathology and plant nutrition, two subjects which are rarely considered together. Graham indicates that this separation is probably a mistake and suggests a number of potentially interesting links between nutrient status, biochemical functions and susceptibility to diseases which should provide fertile ground for further research. The authors are to be congratulated on making the Editor’s task a simple one. I also wish to thank my indexers for their patient endeavours and Mrs Jeni Fox for invaluable secretarial assistance. John Innes Institute, 1983

H. W. Woolhouse


CONTENTS CONTRIBUTORS T O VOLUME 10 PREFACE . . . . . .

. .

. .

. .

. .

. .

. .

.

.

.

.

.

v . vii

Light-harvesting Processes in Algae A. W. D . L A R K U M A N D JACK BARRETT . . . . . . . . . . I . Introduction . . . . . . . . . . 11. Taxonomic Basis . . . . . . . Ill . The Light Climate for Algae . A. Introduction . . . . . . . . . . . . . . . . . . B. Sunlight . C. The Air-Water Interface . . . . . . D. Light Attenuation in Water . . . . . . . . . . . . E. Ultraviolet-B Irradiance . F. The Effect of Algae on Attenuation . . . . . . . . . G. Underwater Light Climates . IV . Structure and Function of the Photosynthetic Membrane . V. Strategies of Light Harvesting . . . . . . . . . . . . . . . A. Introduction . . . . . . B. General Ecological Aspects . C. Taxonomic Aspects . . . . . . . . . . . . . . D. Morphological Aspects . . . . . . . . E. Cytological Aspects F. Biochemical Strategies of Light Harvesting . . G. Physical Strategies of Light Harvesting . . . . . . . . . . VI . Photosynthetic Pigments . Chlorophylls . . . . . . . . . A. . . . . . . . . B. Carotenoids . . . . . . . . Phycobiliproteins . C. D. Action Spectra and Quantum Yields . . . . . . . . . . . VII . Reaction Centre Complexes PSI Reaction Centre Complex . . . . . A. B. PSI1 Reaction Centre Complexes . . . . . C. Size of Antenna Reaction Centres . . . . D. Optical Spectral Analysis of Chlorophyll Proteins . . . . VIII . Pigment Protein (Light-harvesting) Complexes A. Chlorophyll Protein Complexes . . . . . B. Phycobilisomes and Biliprotein Aggregates . . C. Carotenoid-Protein Complexes . . . . . IX . Principles of Light Harvesting . . . . . . . A. Quantum Chemistry and Transfer of Excitation Energy

. . . . . . . . . .

. . . . . . . . . .

. . . . . . . .

. . . . . . . .

.

.

.

.

. . . . . . .

. .

. . . .

. . .

. . .

. . .

. . .

.

.

. .

3 4 5 5

6 8 9 12 12 14 17 20 20 21 23 24 28 36 42 49 49 54 63 68 76 76 88 94 95 102 102 109 1 18 129 129

CONTENTS

X

Structure and Function . . . . . . . . . Distribution of Excitation Energy Between the Photosystems . Interaction of the Light-harvesting Apparatus with other Photo. . . . . . . . . synthetic Processes X . Chromatic Adaptation . . . . . . . . . . A. Historical Aspects . . . . . . . . . . B. Ontogenetic Complementary Chromatic Adaptation . . C. Phylogenetic Complementary Chromatic Adaptation . . D. Other Types of Chromatic Adaptation . . . . . XI . Photo-control of Biosynthesis of Light-harvesting Proteins . . . XI1. Evolutionary Aspects . . . . . . . . . . . A. Evolution of Photosynthesis . . . . . . . . B. Evolution of Photosynthetic Pigments . . . . . . C. Evolution of Early Photosynthetic Prokaryotes . . . . D. Evolution of Eukaryotic Algae . . . . . . . E. Evolution of Thylakoid Stacking . . . . . . . Acknowledgements . . . . . . . . . . . References . . . . . . . . . . . . . B. C. D.

138 I43 149 165 165 166 167 171 173 174 174 171 180 183 187 188 189

Effects of Nutrient Stress on Susceptibility of Plants to Disease with Particular Reference to the Trace Elements ROBIN D . GRAHAM I . Introduction . . . . . . . I1. Adaptation to Nutrient Stress . . . 111. Macronutrients . . . . . . A. The General Pattern . . . B. Phosphorus . . . . . C. Potassium . . . . . . D. Nitrogen . . . . . . E. Sulphur, Magnesium and Calcium F. Summary of Macronutrient Effects IV . Micronutrients . . . . . . A. Copper . . . . . . B. Boron . . . . . . C. Manganese . . . . . D. Iron . . . . . . . E. Zinc . . . . . . . F. Nickel . . . . . . G. Silicon . . . . . . H. Other Elements . . . . I. Summary of Micronutrient Effects J. Guidelines for Experimentation . Epilogue . . . . . . . Acknowledgements . . . . . References . . . . . . . AUTHOR INDEX SUBJECT INDEX

. .

. .

. .

. .

. .

. . .

. . .

. . .

.

.

.

.

.

.

.

.

.

.

.

.

.

.

.

. . .

. . .

.

.

. . . .

. . . .

. . . .

. . . .

. .

. .

. .

. .

. .

. .

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

.

.

.

.

.

.

. . . .

. . . .

. . . .

. . . .

. . . .

. . . .

222 222 223 223 224 225 226 228 228 229 229 238 243 254 257 260 261 263 265 268 269 269 270

. .

. .

. .

. .

. .

. .

277 297

. .

. .

. . .

. .

. . .

. .

. . .

. .

. . .

. .

Light-harvesting Processes in Algae

A . W . D. L A R K U M

JACK BARRETT

School of Biologicul Sciences Universiry o f Sydney N S W 2006 Australia

Division of' Plant Industrj . CSIRO Cunbvrru ACT 2601 Austruliu

1. Introduction . . . . . . . . . . . . 11 . Taxonomic Basis . . . . . . . . . . . . 111. The Light Climate for Algae . . . . . . . . . A. Introduction . . . . . . . . . . . . B. Sunlight . . . . . . . . . . . . . C. The Air-Water Interface . . . . . . . . D. Light Attenuation in Water . . . . . . . E. Ultraviolet-B Irradiance . . . . . . . . F. The Effect of Algae on Attenuation . . . . . G. Underwater Light Climates . . . . . . . IV . Structure and Function of the Photosynthetic Membrane . V . Stategies of Light Harvesting . . . . . . . . . A. Introduction . . . . . . . . . . . . B. General Ecological Aspects . . . . . . . C. Taxonomic Aspects . . . . . . . . . . D. Morphological Aspects . . . . . . . . . E. Cytological Aspects . . . . . . . . . . F. Biochemical Strategies of Light Harvesting . . . G. Physical Strategies of Light Harvesting . . . . VI . Photosynthetic Pigments . . . . . . . . . . A. Chlorophylls . . . . . . . . . . . . B. Carotenoids . . . . . . . . . . . . C. Phycobiliproteins . . . . . . . . . . D. Action Spectra and Quantum Yields . . . . .

.

.

.

. . . . .

. . . . .

. . . . .

. . . . . .

. . .

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

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

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

. . . .

.

.

3 4 5 5 6 8 9 12

12 14 17

20 20 21 23 24 28 36 42 49 49 54

63 68

2

.

.

A W . D LARKUM AND JACK BARRETT

VII . Reaction Centre Complexes . . . . . . . . . . . . PSI Reaction Centre Complex . . . . . . . . . A. B. PSI1 Reaction Centre Complexes . . . . . . . . . Size of Antenna of Reaction Centres . . . . . . . . C. Optical Spectral Analysis of Chlorophyll Proteins . . . . D. VIII . Pigment Protein (Light-harvesting) Complexes . . . . . . . A. Chlorophyll Protein Complexes . . . . . . . . . B. Phycobilisomes and Biliprotein Aggregates . . . . . . C. Carotenoid-Protein Complexes . . . . . . . . . IX. Principles of Light Harvesting 1 . . . . . . . . . . . A. Quantum Chemistry and Transfer of Excitation Energy . . B. Structure and Function . . . . . . . . . . . C. Distribution of Excitation Energy Between the Photosystems . D. Interaction of the Light-harvesting Apparatus with other Photosynthetic Processes . . . . . . . . . . . . X . Chromatic Adaptation . . . . . . . . . . . . . . A. Historical Aspects . . . . . . . . . . . . . B. Ontogenetic Complementary Chromatic Adaptation . . . C. Phylogenetic Complementary Chromatic Adaptation . . . D. Other Types of Chromatic Adaptation . . . . . . . XI . Photo-control of Biosynthesis of Light-harvesting Proteins . . . XI1. Evolutionary Aspects . . . . . . . . . . . . . . A. Evolution of Photosynthesis . . . . . . . . . . B. Evolution of Photosynthetic Pigments . . . . . . . C. Evolution of Early Photosynthetic Prokaryotes . . . . . D. Evolution of Eukaryotic Algae . . . . . . . . . E. Evolution of Thylakoid Stacking . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . References . . . . . . . . . . . . . . . . .

ABBREVIATIONS ATP APC BChl BPh CD Chl ETC IK

Km LiDS LHCP MLWS Mr NADP

Adenosine triphosphate Allophycocyanin Bacteriochlorophyll Bacteriopheophytin Circular dichroism Chlorophyll Electron transport chain Light saturation onset parameter Michaelis-Menten constant Lithium dodecyl sulphate Light-harvesting chlorophyll a/b protein Mean low water spring (tide) Molecular mass Nicotinamide adenine dinucleotide phosphate

76 76 88 94 95 102 102 109 118 129 129 138 143 149 165 165 166 167 171 173

174 174 177 180 183 187 188 189

LIGHT HARVESTING PROCESSES IN ALGAE

ORD Pmax P-680 P-700 PAR PBS PC PCP PCRC PE Ph PQ PSI PSI1 PSU RC I RC I1 RuBP RuBPc’ase SDS UV-B

3

Optical rotatory dichroism Light saturated photosynthetic rate Photoreactive Chl a of RC I1 Photoreactive Chl of RC I Photosynthetically active radiation Ph ycobilisome Phycocyanin Peridinin-Chl a protein Photosynthetic carbon reduction cycle Phycoerythrin Pheophytin Plastoquinone Photosystem I Photosystem I1 Photosynthetic unit Reaction centre I Reaction centre I1 Ribulose Bisphosphate Ribulose bisphosphate-carboxylase Sodium dodecyl sulphate Ultra-violet B radiation (320-280 nm) I. INTRODUCTION

Land plants have evolved a remarkably uniform strategy for harvesting and converting light energy into chemical energy. This is based on processes using the Mg-tetrapyrroles Chla and Chlb, together with a limited number of carotenoids. In sharp contrast, the diverse assemblage of organisms grouped together under the general classification of algae have evolved a greater diversity of pigments and many elaborate strategies for light-harvesting and energy conversion. Yet algae have not attracted as much photosynthesis research despite their inherent interest, as have land-based plants, perhaps because of the agricultural importance of the latter. As a result our current understanding of photosynthesis is largely influenced by a limited “land-plant’’ viewpoint, notwithstanding the deeper and more balanced perspective that can come from an examination of the algal systems. The importance of algae, both as a contribution to our understanding of living things and in practical terms, hardly needs stressing today. Two-thirds of the earth’s surface is covered by the oceans, fresh water lakes and rivers, and in these waters approximately half of the world’s photosynthesis is carried out. Diversification of the primary apparatus of the primitive photosynthetic organisms probably occurred in the more sheltered seas, and later oxygenic photosynthetic systems were evolved, leading to the present day aquatic and

4

A. W. D. LARKUM AND JACK BARRETT

marine algae. It is only recently that the conceptual and practical importance of algal photosynthesis has been reflected in a commensurate increase in research effort, stimulating over the last decade an upsurge in publications in this area of algal science. Thus, despite the previous emphasis on photosynthesis research in land plants there is now a large corpus of work on algae. We have been aware for some years of the absence of any previous review on algae which covered the many aspects of light-harvesting and energy conversion, from the ecological scale down to the molecular level. Consequently it has been our intention to bring much of the dispersed literature together, so as to achieve an integrated framework from which conclusions can be drawn to further stimulate research. It therefore seemed important to us to give space to the many aspects of the topic. Some of these, depending on one’s own discipline, may seem trivial or of no significance, or perhaps experimentally intractable in the light of current scientific knowledge. We have not attempted to review the extensive literature exhaustively. Rather, we have deliberately chosen to emphasize significant data and important hypotheses, and the arguments that relate to them. 11. TAXONOMIC BASIS

This review deals with organisms from the borderline of groups loosely called prokaryotes, plants and animals. It would be logical to use one internationally recognized system to describe these taxonomic groups and subgroups. However no such system exists and each of the three groups (including some overlap, e.g. Cyanobacteria or Cyanophyta) is codified according to different principles and using different hierarchical levels. An attempt has been made by Whittaker and Margulis (1978) to set up a unified classificatory system of the living world. For simplicity this system is adopted here and the relevant parts are set out as follows: (i) Kingdom Monera. Prokaryotic cells Superphylum Photomonera, photosynthetic prokaryotes Phylum Photobacteria, non-oxygen-elimjnating photosynthetic bacteria Phylum Prochlorophyta, green-oxygen-eliminating prokaryotes Phylum Cyanophyta or Cyanobacteria, blue-green algae Superkingdom Eukaryota. Nucleate organization. (ii) Kingdom Protista or Protoctista. Eukaryotic cells with solitary and colonial unicellular organization (Protista) or also including simpler multicellular forms (Protoctista) Branch Protophyta, plant-like protists (or Protoctists) Superphylum Chromophyta or Chromobionta, yellow and brown flagellate algae and allies


5

Phylum Chrysophyta, s.s., Golden algae (including Prymnesiophyta and Chloromonadophyta) Phylum Bacillariophyta, diatoms Phylum Xanthophyta, yellow-green algae Phylum Haptophyta, haptophyte or coccolithophores Phylum Eustigmatophyta, eustigmatophytes Phylum Dinoflagellata or Pyrrophyta, S.S. dinoflagellates Phylum Cryptophyta, cryptomonads (Phylum Phaeophyta) Form-Superphylum Chlorophyta, s.p. or Chlorobionta, green algae Phylum Chlorophyta, S.S. Grass-green algae Phylum Siphonophyta, siphonaceous, syncytical green algae Phylum Prasinophyta, prasinophytes Phylum Zygnematophyta or Gamophyta, conjugating green algae Phylum Charophyta, stoneworts Phylum Euglenophyta, euglenoid flagellates (Form -Superphylum Rhodophyta) Phylum Rhodophyta There will, no doubt, continue to be much argument concerning this and other attempts at classification. However it is sufficiently close to other schemes (e.g. Bold and Wynne, 1978) to cause little confusion in the case of the algae and avoids the strictly correct botanical, but cumbersome, usage of divisions and the suffix “-phyceae”. It recognizes the existence of Prochlorophyta (Lewin, 1976) which has been challenged (Antia, 1977). It recognizes Zygnematophyta which probably belongs with the Charophyta and does not recognize the Ulvaphyta (Stewart and Mattox, 1978). It recognizes Siphonophyta which many workers place in the Chlorophyta. These and other minor problems can be expected to remain for some time to come. 111. THE LIGHT CLIMATE FOR ALGAE A. INTRODUCTION

The majority of algae live in water and are therefore influenced by the light transmitting properties of water and any dissolved or suspended matter. Intertidal and terrestrial algae however live partly or wholly in a terrestrial light climate which is brighter and less complex than that underwater. However, in such places as the understorey of forests and in caves, both of which are good habitats for terrestrial algae, the light climate may be very dim and complex (Section 1X.D: cave algae). Terrestrial light climate studies have been reviewed by Anderson (1 966) and Gates ( 1980).

6

A. W. D. LARKUM A N D JACK BARRETT B. SUNLIGHT

Almost all light of biochemical importance comes from the sun. Although the sun has a complex structure and radiates energy in a complex way, the spectrum of light reaching the earth approximates that of a black body at 5794°K and has a peak at 500nm. This spectrum probably has remained reasonably constant throughout earth history. Theoretical considerations indicate that the luminosity of the sun may have increased by as much as 30 per cent since the formation of the solar system (Owen et al., 1979). Since the radiation of a black body is proportional to the fourth root of the temperature (Stefan-Boltzmann Law) the spectrum of sunlight is unlikely to have shifted by more than 30 nm, towards the blue, during the last 4.5 billion years. The flux of solar energy (irradiance) impinging on a surface perpendicular to the sun's rays just outside the atmosphere is known as the solar constant (1.36 KW m -2) but this varies by 7 per cent during the year due to the elliptical orbit of the earth. Approximately 65 per cent of the energy reaches the ground on a clear day at noon with the sun nearly overhead (air mass = l), and about 45 per cent of this is in the visible range of the spectrum of electromagnetic radiation. Visible radiation approximates closely to photosynthetically active radiation (PAR) which has been defined by the SCOR Working Group No 15 (1965) as energy between 350-700 nm, although the region 350-400 nm is relatively weak photosynthetically, and is often excluded in current PAR measurements. The spectra for sunlight passing through various air masses are shown in Fig. 1 . Sunlight varies sinusoidally during the day according to the sun's elevation. This can be calculated according to the equation:

s,= S,& 1 + cos 2n t/N) where t = time (h) before or after solar noon, s, = irradiance (Wm-2 h-') at time t, s,,=maximum noon irradiance (Wm-2 h-'), N=day length (h) and the cosine function is in radians. In practice this means that approximately 80 per cent of daily irradiance occurs during the middle 50 per cent of daylight hours. For biological purposes light is currently measured as irradiance which is the amount of light energy falling on unit area of a flat surface collecting from a solid angle of 180°, for measuring sunlight the surface should be perpendicular to the sun's rays, unless cosine corrections are made, but for underwater light or under plant canopies other configurations or other units may be preferable (cf. Gates, 1980). Other units for measuring sunlight are: radiance which is the radiation flux (of energy) from a particular, specified direction and solid angle and scalar irradiance which is the radiation flux (of energy) from all directions (solid angle of 360" or 471 steradians) and which is a


7

Wavelength ( F m )

Fig. I . Spectra of light incident at the earth’s surface after passing through various air masses. An airmass of I occurs when the sun is vertically overhead. Spectra plotted from data of the US. Bureau of Meteorology. (For further spectra see Gates, 1980.)

very important parameter at depth underwater. The proper SI units for I Wm-2 or even mW cm-2). However for irradiance are joules m-’ S K(or photosynthetic considerations the flux of photons or quanta is often important (quanta of blue light contain over twice as much energy as quanta of red light but are no more effective in photosynthesis). As a result many values are reported in Einstein’s mK2s-’ or pE mP2s K 1(an Einstein being Avogadro’s number-6.02 x 1 0 2 3 - ~ fquanta). However the Einstein is not an SI unit and some authors therefore prefer moles of quanta of PAR as a more appropriate terminology. Recently there have been attempts to adopt generally a system very similar to scalar irradiance but based on the flux rate of quanta. The new unit would be “photon flux fluence rate” and it seems likely that this will be adopted by the international regulatory body, the Commission Internationale de I’Eclairage. Photosynthetic photon flux fluence rate (PPFFR) is defined as the number of quanta of PAR incident at a point from all directions (i.e. a solid angle of 360”)in unit time. The ideal sensor for this unit would have a spherical collecting surface having the properties of a cosine collector at every point on its surface and responding equally to all photons in the 400-700 nm spectral region. While the new system will be most useful for studies of phytoplankton photosynthesis in the oceans it can also be used for benthic algae even though

8


light here is more vectorial, i.e. is from a solid angle of 180".The new unit has the value that it is defined in terms of photons and not energy. C. THE AIR-WATER INTERFACE

Even when a light beam is normal to a flat water surface some light is reflected and scattered back into the air (Fig. 2(a)). As the angle of incidence increases so does the reflected light. The transmitted light is refracted as it enters the water and if there are waves present a lens effect is produced (Fig. 2(b)) causing concentration of light in various patterns which is known as glitter (Weinberg, 1976; Drew, 1983). Glitter can be important down to depths of 20 m in clear water. It is an important consideration for underwater photosynthetic studies and a major difficulty for the measurement of irradiance in shallow water.

'--

0

10

20

3040

5 0 6 0

70

8090

a(o)

Fig. 2. (a) Loss of light at the surface ofthe sea, a. apparent albedo on a sunny day, b. reflection on a cloudy day, c. reflection on a sunny day (redrawn from Weinberg, 1976); (b) The lens effect of surface water waves which produce glitter.

Above a sea state of 4 on the Beaufort scale, white caps and small air bubbles in the surface water increase backscattering and reflection of incident light. Under these conditions the amount of light entering the water column may be reduced by as much as 50 per cent. However when the sun is low in the sky (angle of incidence > 53") almost all light is reflected from a calm sea and any kind of surface roughening will increase the penetration of light into the water column (Cox, 1974), Fig. 2(a).


9

D. LIGHT ATTENUATION IN WATER

The general properties of light transmission in water have been recognized for many years (e.g. Rabinowitch, 1951). However, the quantitative studies of Jerlov (1951, 1974, 1976) and others (cf. Tyler and Smith, 1970; Jerlov and Nielsen, 1974) have led to a widely accepted classification of water types. Although this work has dealt largely with seawater it is also applicable to freshwater (Talling, 1971; Kirk, 1976b). As shown in Fig. 3 Jerlov (1951) classified seawaters according to the curves of irradiance versus depth and according to the "colour" of the water (Fig. 4). Oceanic water may approach the attenuation properties of pure water (Morel, 1974); the term "attenuation" is used in preference to absorption to give recognition to the large part played by light scattering in all types of seawater (Morel, 1974). The Sargasso Sea is an example of a region with very clear seawater of a type classified by Jerlov as oceanic water type I. It has a

lrrodionce

("/o

of surfoce, 350-700nrn)

Fig. 3. The attenuation of light in seawaters, as classified by Jerlov (see text for details)

10

A. W. D. LARKUM A N D JACK BARRETT Downward irradiance

" I

400

500

600

400

500

600

Wavelength (nm)

Fig. 4. Spectra of downwelling light at various depths for (a) oceanic water and (b) coastal water. (Data taken from Jerlov, 1976.)

maximum transmission at 475 nm in shallow water and 465 mm in deeper water (Jerlov, 1976); red light is screened out at shallow depth (Fig. 4) and the high relative transmission of blue light has led to the description of this water as blue in colour (the surface colour of the sea, although related to the type of seawater, is influenced by a number of considerations other than the wavelength of maximum downward irradiance: Jerlov, 1976; Morel and Prieur, 1977). Two further types of oceanic water are also blue in colour according to Jerlov's classification-types I1 and I11 (and these may be further subdivided-see Jerlov 1976); attenuation of light is greater than that of type I (Fig. 1) and the maximum wavelength of downward irradiance is between 475 and 500nm, due to the presence of some dissolved and particulate matter which screens out or scatters some blue light. The other types of seawater shown in Fig. 3 are found in inshore waters which are influenced by the presence of dissolved pigments as well as suspended matter. The pigments involved have been only poorly defined and have been found to be abundant in the breakdown products of terrestrial vegetation; carbohydrates and humic matter form a large component (Kalle, 1966; Seiburth and Jensen, 1969). Thus it has often been proposed that these substances are brought to the sea through run-off from the land and hence are common in inshore waters. However Seiburth and Jensen (1 969) suggested that a proportion of these substances are produced in situ in the inshore zone


11

by benthic algae. These substances screen out blue light giving a yellowishbrown appearance to waters in which they are abundant and for this reason such pigments have often been referred to collectively by the German term “gelbstoff’ (Kalle, 1966) or “yellow substance”, although Kirk (1977) argues for “gilven” (from the Latin). Yellow substance contributes to the spectral characteristics of inshore seawater (see Figs 3 and 4), according to concentration, from type 1 to type 9. It is also found in inland freshwater (e.g. Kirk, 1976b). In all such waters the attenuation of light is much greater than in oceanic water and the maximum wavelength of downward irradiance is found between 500-600 nm (in that order going from type 1 to type 9 seawater).

400

450

500

1 550A,n6,0

Fig. 5 . Spectral attenuation of various types of seawater as measured by Pelevin and Rutkovskaya (1977) (continuous curves) compared with spectra given by Jerlov (broken curves), see text.

While the classification of Jerlov (195 1) has been useful, the recent development of submersible monochromators with great sensitivity and bandwidth separation has revealed significant differences from the spectral attenuation curves first presented by Jerlov. In particular, much greater attenuation in the blue region is found for all types of water, particularly for coastal waters (Fig. 5). It has been proposed that a new type of classification be adopted based on the vertical extinction coefficient of light (a,) at 500 nm (Pelevin and Rutkovskaya, 1977).

12

A. W. D. LARKUM A N D JACK BARRETT

E. ULTRAVIOLET-B IRRADIANCE

Ultraviolet-B radiation (UV-B, 280-320 nm region) or middle ultraviolet radiation (MUV 280-340 nm region) penetrates to the surface of the earth in significant amounts in sunlight, but is rapidly attenuated in water (Zanefeld, 1975; Smith and Calkins, 1976). Such radiation is harmful to a variety ofplant processes (cf. Trocine et al., 1981 and the references contained therein). Photosynthetic reactions in a number of algae are inhibited by UV-B irradiation (Bell and Merinova, 1961; Halldal, 1964; Van Baalen, 1968; Smith et al., 1980) under dosage levels equivalent to those at the air-water interface and just below. Glass containers, which have been the preferred type of vessel for measuring photosynthesis in the field, screen out UV-B irradiation so there is little satisfactory evidence on the significance of UV-B irradiation in causing photo-inhibition of photosynthesis, which certainly occurs at high irradiance levels (Harris, 1978). Recent work suggests that UV-B does play an important role in the inhibition of photosynthesis of phytoplankton (Smith et al., 1980). This raises the question as to its importance for photoinhibition in intertidal algae, and other benthic algae in shallow waters such as mud flats and coral reefs. Photoinhibition, however, is not just an effect of high dosages of UV-B irradiation and is discussed further in Section 1X.C. F. THE EFFECT OF ALGAE ON ATTENUATION

The inherent properties of water, the presence of yellow substance and nonliving particulate matter largely determine the attenuation of light in a given body of water. However, the contribution of algae (particularly microalgae) cannot be ignored. In many marine and freshwater situations the scarcity of phytoplankton causes a negligible effect on light attenuation. At the other extreme, very dense populations can reduce the irradiance to zero within a depth of 0.6 m (Talling et al., 1973). Lorenzen (1972, 1976) has attempted to quantitate attenuation in terms of the above three components, the water itself, the dissolved and particulate matter and the phytoplankton. If an “average” extinction coefficient (400-700 nm) is assigned to each of these three components (kl,k, and k,, respectively) the contribution of each component to the attenuation of light at any depth in a particular water column can be determined. These factors can be related to the depth of the photic zone (that is the depth at which light is attenuated to 1 per cent of surface light). A narrow photic zone will be caused by either large amounts of dissolved or suspended matter or by high densities of phytoplankton. Figure 6 shows a typical relationship between absorption by each of the three components and the extent of the photic zone. It is assumed in Fig. 6 that in a very narrow photic zone the major contribution to attenuation is by phytoplankton (as was so in the extreme case of Talling et ul., 1973) although


13

this is not necessarily so. In photic zones of intermediate extent (1 5-50 m) dissolved and particulate matter account for the greater part of the attenuation; and in very clear waters (Oceanic Type I, 11 and 111) the water component predominates. A very important conclusion from the Lorentzen model is that for all but the most shallow algal communities the potential for utilization of sunlight will fall far short of terrestrial systems.

Euphotic zone (m)

Fig. 6. The contribution (KJK,) to the absorption of light in the sea made by the water itself (LI-IJ) and by phytoplankton (A- -A). (From Lorenzen, 1972, 1976.)

(0--0) by dissolved substances and inorganic particulate matter -

-

However, the model of Lorenzen does not take into account the effect of light scattering. Light scattering which is enhanced by the presence of dissolved or particulate matter (Kullenberg, 1974; Timofeeva, 1974; Jerlov, 1976) increases the effective path length of light in the water column, thus increasing light attenuation. Monte Carlo treatments of light attenuation offer a better approach to the real situation (Zanefeld, 1974; Kirk, 1981) and are likely in the future to yield more precise information on the attenuation of light in natural water columns and the proportion of light harvested by algae. Phytoplankton in a water column will affect the spectral quality as well as the attenuation of downward irradiance. The exact spectral change will depend on the species composition: the presence of green algae would result in light in deeper water with a maximum in the green region (500-560nm) whereas Dinoflagellata or Bacillariophyta would result in light with a maximum in the yellow region (550-620 nm). However it is difficult to obtain quantitative results on such spectral changes in the field, so that evidence has

14


been taken from laboratory studies of the passage of light through algal suspensions. Rabinowitch (1951) reviewed some of the earlier results (with Chlorophyta, Phaeophyta, Rhodophyta, Cyanobacteria, Bacillariophyta) including the important work of Emerson and Lewis (1942, 1943) on Chlorellu (Chlorophyta) and Chroococcus (Cyanobacteria). Yentsch (1962) investigated spectra from Phueductylum tricornutum (Bacillariophyta) and Nunnochloris utomus (Chlorophyta) using an opal glass method and obtained other spectra for microalgae deposited on a filter disc (see also Yentsch, 1980). All the results on light transmitted through microalgal suspensions show that the spectral modification is less than would be calculated from a summation of the spectra of the individual pigments (this effect was particularly marked in the study of Chlorellu by Emerson and Lewis, 1943). The reason for the apparent discrepancy is the “package” effect (sometimes called the “sieve” effect) (Duysens, 1956; Rabinowitch, 1956; Kirk, 1977). This is the phenomenon by which the presence of pigments in discrete packets (as in algal cells) rather than as a homogeneous dispersion throughout the water column leads to a less efficient capture of light at the absorption maxima of the pigments. The mechanism is discussed in greater detail in Section V.F.4. The result is a flattened spectrum. Further flattening may result from light scattering within algal cells (Section V.F.4). Kirk (1975a,b, 1976a) has presented a theoretical treatment of the effect of cell size and shape on the “package” effect for microalgal populations. Large, spherical cells produce the greatest effect i.e. the least efficient absorption, whereas small rods produce the least effect, i.e. the most efficient absorption, although even with the latter package effects are still appreciable (Fig. 11) (see Section V.D.). G. UNDERWATER LIGHT CLIMATES

An important consideration for underwater irradiance is that the variability of underwater light regimes is an order of magnitude greater than that normally encountered in the terrestrial environment (cf. Luning and Dring, 1979; Weinberg, 1976). Variability occurs on both a day-to-day and seasonal basis. Luning and Dring (1979) found that virtual darkness occurs for long periods in the winter at moderate depths in waters near Helgoland (North Sea). The seasonal decline, in winter, is brought about by (i) storms which firstly decrease the amount of light entering the water column due to surface turbulence and secondly increase turbidity due to stirring up of bottom sediments and (ii) the decrease in the sun’s declination which markedly diminishes the amount of light entering the water column (Subsection C, above). The quality of underwater irradiance is also seasonally variable, being influenced by seasonal phytoplankton blooms and by rains which bring large amounts of run-off from the land. This water has a high content of yellow substance and particulate matter. On a day-to-day basis, storms, heavy rains


15

and phytoplankton blooms also produce sudden short-lived changes in the light climate. These variable conditions of irradiance impose the need for a high degree of flexibility on the photosynthetic strategies of freshwater and marine algae. These strategies must be more adaptable than those of terrestrial plants. An even greater flexibility is required of micro-algae which may be taken in currents from the top of the photic zone to the bottom within a short period. Despite the importance of characterizing the underwater light climate few comprehensive studies have been done. The most complete study at present is that of Luning and Dring (1979), in which instantaneous readings of downward irradiance were taken every 20 min for one year at two depths (2.5 and 3.5m below MLWS) and in three spectral regions (400-500nm, 500-600 nm and 600-700 nm). The site was in the sublittoral of the rocky island of Helgoland (North Sea). Even this study has shortcomings because as the authors note, there can be wide variation in the irradiance within seconds and therefore a single reading every 20 min is inadequate. Nevertheless much useful information was provided by the study. Figure 7 shows the daily photon flux density in the visible range (400-700 nm) for a complete year at various depths. Approximately 90 per cent of the annual total light reaching the sublittoral region was received in the six months from April to September.

Fig. 7. Annual variation in the photon flux density of PAR (400-700 nm) at various depths in near inshore water in the North Sea. (Data from Luning and Dring, 1979.)

16


During this period the water quality corresponded to Jerlov water type 7; approximately 50 per cent of the irradiance was in the green region (500-600 nm), 30 per cent in the red region (600-700 nm) and 20 per cent in the blue region (400-500 nm), see Fig. 7. More studies of this kind need to be done in order to characterize the various water types and underwater light climates. In regions where smaller variations in downward irradiances occur, such as in the tropics, a reasonable estimate could be obtained from continuous above-water measurements and occasional measurements of spectral transmittance, using calculations based on the water types of Jerlov (1976). Weinberg and Cortel-Breeman (1978) used this method, but changes in climatic conditions may cause large errors in such estimates. Another feature of underwater irradiance that needs consideration is the large short-term changes that can occur in underwater irradiance. Figure 8 illustrates variation in the underwater irradiance of a shallow sublittoral site at Long Reef (Sydney). Such observations stress the fact that submerged algae must not only adapt to an average underwater light regime, but must be able to withstand periods when daily light levels are very much lower or sometimes higher, than normal. Work in freshwater lakes (Talling and Spence, 1975) and rivers (Westlake, 1965) suggests that similar variations occur in their light climates. The light-harvesting properties of algae are presumably adapted in general to this wide variability in photon flux (but see Section XC).

0600

0800

1000 1200 TIME

1400

1600

1800

Fig. 8. Diurnal change in underwater irradiance at Long Reef Sydney in spring on a day with a high tide at noon; (a) change in water depth (b) underwater irradiance; top and bottom lines indicate maximum range of irradiance, centre line is the mean and hatched area is the 70% range; broken line shows maximum irradiance on a day with low tide at noon.The decrease in irradiance near midday occurred when the rising tide caused detritus and other materials to be washed off the nearby rock platform and to drift across the study site.


17

IV. STRUCTURE AND FUNCTION O F THE PHOTOSYNTHETIC MEMBRANE The photosynthetic carbon reduction cycle (PCRC) or Benson-Calvin cycle is powered by ATP and NADPH which are produced by the photochemical and electron transport reactions of photosynthesis. There is abundant evidence that all the components for these photochemical reactions are localized in the photosynthetic membrane while the PCRC is located in the soluble stroma (chloroplasts) or cytoplasm (prokaryotes). The PCRC is universal to all photosynthetic organisms including photosynthetic bacteria (Fuller, 1978). The reduction of NADP + requires a source of hydrogen. In all oxygenic photosynthetic organisms this source is normally water, oxygen being released as a by-product. However there is some evidence that in certain cyanobacteria other reductants may at times be used (Krogmann, 1981). The splitting of water and the set of reactions which lead to the reduction of NADP are part of what is called the photosynthetic electron transport chain (ETC). This chain involves the transfer of reducing equivalents (in the form of electrons) against the prevailing thermodynamic gradient (which is towards oxidation) and requires an input of energy. To reduce one equivalent of NADP to NADPH ( + H +) a free energy change must take place at least equivalent to that necessary to drive an electron across the 1.63 eV span from water to NADPH. It is light energy converted to chemical energy that provides the energy. Theoretically a photon of red light with an energy of 1.83 eV has the ability to power the ETC with a single photochemical event. However as explained by Radmer and Kok (1977) only about 1.13 eV of the energy of each photon can be supplied as free energy. Furthermore energy is needed not only for the formation of NADPH but also for ATP. Thus in fact, there is a requirement for two sites of photochemical conversion on the ETC between water and NADP+. A scheme involving two such sites was formulated by Hill and Bendall (1960) and is commonly known as the “Z” scheme. The “Z” scheme has received wide experimental support and a current version (somewhat simplified) is shown in Fig. 9 (for further details see Golbeck et al., 1977; Junge, 1977; Williams, 1977). The two photochemical reactions are carried out by photochemical units composed of pigment-proteins and non-coloured proteins which are embedded in a lipid bilayer. The major pigment is Chl u. However, only a small fraction of the Chla present in each unit is involved at the reaction centre where the photochemical reaction occurs. The remainder is antenna Chl which harvests light and passes excitation energy on to the reaction centre. Each reaction centre of PSI or PSII contains Chl a (or possibly phaeophytin in PSII reaction centres) probably in the form of a dimer, denoted as P-700 for PSI and P-680 for PSII, since each centre undergoes oxidation after excitation with accompanying bleaching at about 700 nm or 680 nm. +

+

18

A. W. D . LARKUM A N D JACK BARRETT

Eb (volts) -0.4/ /

Pc

‘0.8

3

Photosystem

I

-.---4

Photosystwn JI

Fig. 9. The “Z” scheme for photosyntheticelectron transport based on the original suggestion of Hill and Bendall (1960) but modified according to later information (see text for details).

The function of the reaction centre Chl P-680 is to cause charge separation so as to form a strong oxidant, Z (E’o -0.8 V) and a weak reductant, Q -(E’o 0.0V). The nature of Z is not known but it is sufficiently oxidized to extract electrons from water. However the splitting of water requires the extraction of four electrons since molecular oxygen is formed. This probably occurs sequentially (Bouges-Bocquet, 1980) although it is possible that two reaction centres could be involved in each photolytic act. Thus there must be a direct 1 : 1 or possible 1 : 2 proportioning between the water-splitting apparatus and P-680. The primary reductant Q -(Section V1I.B) interacts with a pool of plastoquinone (PQ) which in turn interacts with other intermediate electron carriers (the cytochrome b-f complex, and plastocyanin) which connect to oxidized P-700. The primary electron acceptor of PSI, X,may be a special ferredoxin which is an intermediate electron carrier between X and NADP +(SectionVI1.A). The ratio of intermediate electron carriers for P-700 is not fixed and suggests that the whole photosynthetic ETC is not a discrete unit in the membrane (Section IX). The “Z” scheme does not account for the formation of ATP, nor does it fully explain why the reactions occur in a membrane. The membrane involved is the inner membrane of chloroplasts which is vesiculate, that is, it has an inside space delimited from an outside space (the stroma). Each unit is called a thylakoid (Menke, 1962). In many chloroplasts the thylakoids appear to be interconnected and may be folded and arranged in complex ways (see Section IV.E.3). The structuring of the thylakoids is even more complex in Phaeophyta (Greenwood, personal communication).

-

+

+


19

Mitchell (1966) first suggested that the reaction centres of PSI and PSII were arranged across the thylakoid membrane in such a way as to cause charge separation and the generation of a proton gradient. There is now much evidence for such a membrane scheme of photosynthetic ETC (Trebst, 1974; Junge, 1977) which is shown in Fig. 10. The proton gradient is generated by the release of protons by water-splitting on the inner side of the thylakoid membrane and by the translocation of protons to the intrathylakoid space by reduced plastoquinone. The energy for the generation of the proton gradient

Fig. 10. A model of the arrangement of components of the thylakoid membrane (nonappressed). (Redrawn from Hinkle and McCarty, 1978.)

(strictly an electrochemical gradient for protons involving both a pH difference and an electrical potential difference) comes from the charge separation generated by the photochemical conversions of P-700 and P-680. ATP is generated by coupling the proton gradient to an ATP synthase located partly in the thylakoid membrane (Junge, 1977; McCarty, 1979). Thus part of the energy from each of the photo-acts of PSI and PSII is used to form ATP as well as NADPH. According to the membrane model there is coupling of PSII and PSI through plastoquinone and between charge separation and ATP formation via a proton gradient. There is thus no requirement for a close physical proximity of the PSI, PSII and ATP synthase apparatus. In photosynthetic bacteria there appears to be only one type of reaction centre associated with a BChl-Bpheophytin dimer with a light-induced absorption decrease, between 870 and 960 nm depending on the particular organism. Charge separation drives a proton gradient by which ATP is generated. This process involves a cyclic electron transport system in which ubiquinone is involved in the return of electrons to the oxidized side. However

20

A. W . D . LARKUM AND JACK BARRETT

NADPH may be produced by a secondary ATP-driven process involving reversed electron flow. The bacterial photosystem therefore has some similarities to, but also some differences from PSI. V. STRATEGIES OF LIGHT HARVESTING A. INTRODUCTION

In this section the various kinds and levels of light harvesting available to algae are reviewed briefly. A more detailed analysis of some biochemical and biophysical aspects of light harvesting is to be found in Section IX. Light is essential to all photosynthetic autotrophs. But it is only to the extent that light is limiting to growth that light harvesting strategies become important. It is therefore necessary to consider under what conditions light does become limiting for algal growth. A simple calculation shows that a single green plant cell has the capacity to absorb only a small fraction of incident sunlight on a clear summer day in tropical and temperate regions of the earth. Thus sunlight at midday near the equator has an energy of approx. 1 KW m at the earth’s surface [Solar Constant (above the atmosphere), 1.36 KW m -2]. The photosynthetic thylakoid membranes have the capacity to convert light to chemical energy at a rate of about 0.05 W m -2 (Raven, 1978). In an algal cell, such as Chlorellu, the chloroplast membranes are folded many times and this gives an amplification factor of some ten fold, that is, a capacity to fix energy at 0.5 W m-’ (Atkinson et ul., 1974). In higher plant cells, folding of the membranes and a higher concentration of the photosynthetic apparatus leads to a further two-fold amplification (Raven, 1978) or a capacity to fix light energy at about 1 W m -2. Therefore in the most efficient green cell there is a thousand fold excess of incident light energy under full sunlight. Such a simple calculation may ignore many important factors such as (i) that 50 per cent of incident light is in regions of the spectrum unavailable to photosynthetic pigments; (ii) that large losses due to reflection, scattering and heat transfer occur; (iii) that the photosynthetic apparatus is at best 35 per cent efficient (Nobel, 1974; Radmer and Kok, 1977); (iv) that incident light levels are generally much lower than 1 KW m -’, especially for algae (Section 111). Nevertheless, it is apparent that under the best conditions there is an oversufficiency of incident PAR and that, ignoring self-shading problems, there is enough available energy for a layer of cells between 10 and 100 cells deep (from rather different assumptions Raven (1978) concluded that such a layer could be at least 10 cells deep). The basic reason for this poor matching of incident and captured energy lies in the inability of photosynthetic organisms to provide a greater density of photosynthetic apparatus per unit of thylakoid membrane, or to increase the

-’


21

amount of thylakoid membrane per unit volume beyond a critical limit. Raven (1977) has discussed the strategy of increasing the number of layers of chloroplasts within a cell, and came to the conclusion that diffusion of carbon dioxide limited the number of layers that could photosynthesize effectively to no more than two. Therefore the only means that a photosynthetic organism has to increase its light harvesting capacity beyond these basic limits is to provide a number of cell layers. In evolutionary terms, multicellular organisms are a relatively recent development (Section XII). Basic light harvesting mechanisms in unicellular organisms must have evolved long before this. It is probable that the mechanisms were evolved under conditions in which light was not the major limitation for growth, and these have been conserved ever since. B. GENERAL ECOLOGICAL ASPECTS

Consideration of the adaptations for light-harvesting at the ecological level has not been treated well for any group of plants. For terrestrial plants many factors complicate the issue, especially water usage (Mooney and Gulman, 1979). Perhaps the best terrestrial example of an ecological adaptation for light-harvesting is the development by trees of a large photosynthetic canopy which overtops competitors. This in turn leads to understorey plants that have developed shade strategies, and are able to exist in a low light environment (Boardman, 1977). Grime (1974, 1977) includes these strategies within a general hypothesis which describes the major ecological principles involved in terrestrial vegetation, and Raven et al. (1979) have extended this hypothesis to include algae. Grime identified three basic groups of land plants, (a) canopy dominants, (b) ruderal plants (opportunistic, pioneer types), and (c) stresstolerant plants.

Canopy dominant plants have overtopping characters, involving large aerial structures and also generally reach reproductive maturity very slowly. Ruderalplants have a fast growth rate but develop less massive structures and rapidly reach reproductive maturity. Stress-tolerant plants are of variable form and structure but have special adaptations which enable them to survive stress, such as heat, cold, low light intensity, aridity and low nutrient supply. In terms of ecological succession the canopy dominants will tend to dominate in stable, favourable environments, ruderal plants will dominate in unstable, disturbed environments and stress-tolerant plants will be found in situations where any particular stress exists. In the underwater environment the number of possible factors is reduced considerably. Wave and storm damage will give rise to the greatest instability and the major stresses will be shading, low nutrient status and variable salinity, of which shading will be by far the most common.

22


Thus it is possible to predict three major ecological types of benthic algae; (a) canopy dominants with large photosynthetic laminae and overtopping features, (b) ruderal algae with small thalli which form a “turf’ in waveexposed situations and (c) shade algae which form an understorey beneath canopy dominants or at great depths. These ideas are not entirely new and some have been incorporated in other ecological models. (For example Odum (1969), Pianka (1970) and Dayton (1975) proposed almost identical ecological categories of algae: canopy dominants, opportunists and shade algae). There are a number of points which arise from Grime’s approach. Amongst benthic marine algae, canopy dominants are formed almost exclusively of kelp (Laminariales) or fucoid (Fucales) algae. Ruderal algae are found in the four major phyla (Chlorophyta, Phaeophyta, Rhodophyta and Cyanobacteria). Shade-tolerant algae are also found in all four groups, although the Rhodophyta have more species of shade algae (Sears and Wilce, 1968), but they may be less abundant in terms of biomass (Larkum et al., 1967). Canopy dominant algae are restricted to the upper, well lit benthic zone. Below this zone, sciaphilous (shade-loving) algae dominate. Great care must be taken in defining this sciaphilous zone. In a number of reports it has been shown that apparent changes in vertical zonation, from canopy dominants to turf and crustose (encrusting) algae, are not due to changes in irradiance (Kain, 1962; Kain et al., 1975). Many significant factors may change with increasing depth apart from irradiance e.g. turbulence, wave-action, nutrient status, slope, substratum and faunal interactions. Only in a few studies have such complicating factors been eliminated, and a true sciaphilous community been demonstrated (Molinier, 1960a,b; Larkum et al., 1967; Drew, 1969; Lang, 1974; Sears and Cooper, 1978). In particular it has often been observed that the transition from rocky to sandy substratum, which normally occurs on any rocky coastline between a depth of 10 and 100 metres has a profound effect on the communities living on the rock substratum near the transition (Kain, 1962; Kain et al., 1975). This “bottom effect” may generate what appears to be a sciaphilous community, that is a community of red algal turfs and crustose coralline algae, which may occur anywhere between 10 and 50 metres depth. Analysis of the change of vegetation on isolated boulders occurring at different depths is subject to similar difficulties. Other ecological groupings of algae have also involved light-harvesting characteristics. Many attempts have been made to relate pigmentation to ecological strategies in benthic algae, particularly in terms of vertical zonation. This aspect is discussed in the Subsection C and in Section X. The mobility of phytoplankton in the water column would appear to run counter to any specific ecological light-harvesting adaptations (Raymont, 1980). Nevertheless, some unicellular algae are adapted to high irradiance conditions in shallow lagoons or salt pans e.g. Dunaliella sp. (Chlorophyta) (cf. Falkowski and Owens, 1980; Section 1X.D).


23

C. TAXONOMIC ASPECTS

The traditional separation of algae into two major functional groups-the benthic algae and the planktonic microalgae-has focused attention in terms of light-harvesting on the benthic macroalgae. Since the latter are for the most part marine, attention has been directed specifically to zonation on rocky coastlines. The phytoplankton, subject as they are to passive movement in currents, which may drastically affect their vertical distribution, are less easy to study, although modern culture techniques have enabled simulation of some of these conditions in the laboratory. Engelmann proposed in 1883-1 884 that algae of the groups, Chlorophyta, Phaeophyta, Rhodophyta and Cyanobacteria, had distinctly different light harvesting characteristics because of their different photosynthetic pigments (Section VI). Engelmann, and later Gaidukov (1903), suggested that this pigmentation might be related to the light climate of the algae. Red algae, in particular, were well adapted to living in deep water where green light predominates. The hypothesis of complementary chromatic adaptation is discussed in Section X. The simplified assumptions on which the hypothesis was based have undergone great changes in the last twenty years; the great complexity of pigmentation in algae has been established (Section VI); the emphasis on macrobenthic algae has receded following recognition of the many other phylogenetic groups; and above all, there has come about a better understanding of the wide variation in underwater irradiance, qualitatively and quantitatively (Section 111). In oceanic waters which are predominantly “blue”, it is now widely accepted that no benthic or planktonic algal group have an advantage in terms of light-harvesting at the lower limit of the photic zone (Crossett ef al., 1965; Larkum et al., 1967; Drew, 1969; Levring, 1966; Luning and Dring, 1979; Dring, 1981). The predominance of red algae near the lower limit (Molinier, 1960a,b; Larkum el al., 1967) is probably an adaptation to low light conditions per se rather than to the quality of the light. Members of the Rhodophyta and Cyanobacteria are also found to predominate in caves at the limit of algal growth although the light changes very little spectrally (Larkum ef al., 1967; Norton et al., 1971). In the mid-photic zone the dominance by species of the Phaeophyta, which occurs in temperate and polar regions (Mann, 1973), is related to the successful growth form of the kelps and fucoids which itself may be regarded as a light harvesting strategy. In tropical waters where these algae dominate only rarely, no such zone of brown algae in the mid-photic zone occurs (Gilmartin, 1958). In coastal waters which are yellow or yellow-green and have a narrow photic zone there is an advantage for deeper-living algae in having the ability to harvest green and yellow light. However, in these waters there are few

24


convincing reports of dominance by Rhodophyta and Cyanobacteria at the deeper limit of algal growth (Section X).Other ecological considerations seem to override any constraint imposed by pigmentation. Furthermore, the very variable underwater light climate (Section 111) and the ability of most microalgae and benthic algae, to adapt their pigmentation and chloroplast structure to increase their light-harvesting in yellow-green light (Subsection F) tend to lessen the apparent advantage of Rhodophyta and Cyanobacteria. The pigmentation of algal phyla is shown in Table I. D . MORPHOLOGICAL ASPECTS

I. Unicellular Algae The shape of plants may have profound effects on their light harvesting properties (Gates, 1980). It might be assumed that the microscopic unicellular algae of the phytoplankton would be less affected but this is probably quite incorrect. Kirk (1975a,b, 1976a, 1977a) made a theoretical treatment of the effect of size and shape of unicellular algae on their light harvesting properties. His conclusions may be summarized broadly as follows. In terms of populations of cells, small non-spherical cells are more efficient than spherical cells (see Fig. 11). However due to the package effect (Subsection G) populations of even the smallest cells can never harvest light as efficiently as a homogeneous dispersion of an equivalent amount of pigment. As stressed above Kirk’s calculations treat populations of cells. Presumably selection pressure operates at this level to favour small, non-spherical unicells (see Parsons et al., 1977). However the range of shape, size and form within phytoplankton indicates that other factors may be involved (cf. Malone, 1980). Kirk (ibid) assumed that cells were not vacuolated and had a uniform distribution of pigment. In practice this assumption is not correct since many unicellular algae have large vacuoles and the pigments are located in chloroplasts which are distributed about the periphery of the cell. A large, vacuolated, cylindrical cell may be reasonably efficient in its light harvesting properties. This morphology may have other advantages: for buoyancy control (Smayda, 1970;Subsection F), for reducing the diffusion resistance for COz between the plasmalemma and the chloroplast (Raven, 1977; Subsection F) and for expanding the light harvesting surface area under shade conditions. 2. Macroalgae Light harvesting is important in any consideration of the morphology of macroalgae even though morphology is related to a number of ecological strategies. For instance, many macroalgae are canopy dominants and these algae, with a large photosynthetic canopy raised above the substrate, are susceptible to damage by wave-action. Canopy dominance is thus constrained by wave-action, and in very exposed situations other types of algae may


25

i 350

I

1

1

1

I

I

I

400

450

500

550

600

650

700

Wavelength (nm)

Fig. 1 I . The effect of cell size and shape on the absorption cross-section of algae. The curves relate to the calculated absorption cross section of randomly oriented blue-green algal colonies. The data apply to 100 000 pm3 of algal volume which corresponds to one particle in the case of 57.6 pm diameter spheres (a), to one particle in the case of 230.4 by 28.8 pm prolate spheroids (A). to one particle in the case of 3537 by 6 pm cylinders (A)and to 884 particles in the case of 6 p m diameter spheres (0). (Data from Kirk 1978a.)

dominate. The latter may be turf algae (tough, erect thalli of few centimeters length), microscopic filamentous algae or encrusting algae (which may be Cyanobacteria, Chlorophyta, Phaeophyta or Rhodophyta). In intermediate sites of wave-exposure, many fucoid algae (e.g. Surgassum spp.) may dominate. Canopy dominants are mainly brown algae. In temperate and polar regions Laminariales are the most prevalent group (Mann, 1973), with fucoid algae restricted to sites of greater wave-action. In tropical subtidal waters fucoid algae are the more abundant. In the intertidal region where because of high wave-action there are few Laminariales, fucoid algae are also abundant. However, members of the Chlorophyta and Rhodophyta compete effectively for co-dominance. Here light-harvesting strategies are of less importance because of the higher levels of PAR and morphological and physiological adaptions for resisting waveaction and desiccation are of critical importance. The relationship between photosynthesis and morphology in macroalgae has rarely been treated experimentally. Kanwisher (1966) speculated about the optimum shape of an alga and concluded that a high surface area to volume ratio would be ideal. Algae with a greater percentage of structural (non-photosynthetic) tissue should show lower photosynthetic capacities. Odum el al. (1969) indeed found that gross photosynthetic capacity was

TABLE I Pigment Type, Thylakoid Arrangement and Nuclear Organization in Different Groups of Algae Chlorophylls Cyanoph yta

a

Prochlorophyta

a, b

No. of appressed thylakoids

Major light-harvesting pigment-protein

Carotenoid"

Organization of nuclear material

PC, PE

non-appressed

P

LHCP?

2-10

P

~~

Rhodophyta

a

PC, PE

non-appressed

E

Cryptophyta

a, c2

PC,PC, Chl a/c

2

E

Dinoflagellata

a, c2

Peridinin

PCP

3

M

Prymnesiophyta

a, c l , c2

Fucoxanthin

Chl a/c- Fucoxanthin

3

E

Chrysophyta

a, c,, c2

Fucoxanthin

? not known

3, girdle lamella

E

Xanthophyta

a, c,*, c2*

? not known

3 fgirdle lamella

E

? not known

3 fgirdle lamella

E

? not known

3 fgirdle lamella

E

+c,

3. girdle lamella

E

Fucoxanthin +Chl a, c l , + c2

3, girdle lamella

E

a, b

LHCP

3, grana?

E

Chloroph yta

a, b

LHCP

2 4 , grana?

E

Prasinophyta

a, b

LHCP

2 4 , grana?

E

Charoph yta

a, b

LHCP

2 4 , grana?

E

Chloromonadophyta

a, c,, c2

Fucoxanthin

Eustigmatoph yta

a, (c?)

Bacillariophyta

a, c l , c2

Fucoxanthin

Fucoxanthin+Chl

Phaeoph yta

a, cl, c2

Fucoxanthin

Euglenophyta

a, clr

a See Table 11, &carotene present in all groups. Lutein may be a light-harvesting pigment in Chlorophyta, cf. higher plants (Siefermann-Harms, 1980b). See text for further details. * Signifies present in trace amounts. P=Prokaryotic M = Mesokaryotic E = Eukaryotic (For further division of eukaryotic types, see Stewart and Mattox, 1975).

28


proportional to the surface area to volume ratio. Further confirmation of this point has been provided (Littler and Murray, 1974; Ramus, 1978; Littler, 1980; Arnold and Murray, 1980; Littler and Littler, 1980). Thin, sheet-like construction as found in Ulva spp. and Enteromorpha spp. (Chlorophyta), yields the highest photosynthetic capacity. However, such algae are, like the Laminariales, susceptible to damage or loss through wave-action; they do not often dominate benthic algal vegetation although they are omnipresent species in most coastal habitats. Littler and Littler (1980) and Hay (198 1) have discussed photosynthetic performance in a variety of marine benthic algae in relation to form and function. Macroalgae with thicker and more complex thalli than those with thin sheets may under certain conditions exhibit high photosynthetic performance. Ramus (1978) showed that the green alga Codium fragile which has a thick, branched, tubular thallus can, under low light conditions, give a greater photosynthetic performance than Ulva lactuca. This is because the light harvesting capacity (approaching 100 per cent) of the thallus of Codium is much greater than that of Ulva for an equal concentration of photosynthetic pigments. Ramus ascribed the greater light absorption efficiency to the special structure of the Codium thallus, which enhances internal absorption of light. However, apart from this single report no other investigations have been made in this important area. Clearly, a disadvantage of canopy dominance or the presence of a tough, thick thallus is the increased respiratory activity incurred by cells which are non-photosynthetic or are highly shaded. Thus although light harvesting may be efficient, productivity (the net accumulation of organic carbon) may be less so. Nevertheless as discussed above such types of algae dominate in many littoral and sublittoral situations due to various ecological factors (cf. Littler and Littler, 1980). As light is attenuated with increasing depth such canopy dominants are adversely affected (Luning, 1969; Kain et al., 1975) and other communities take their place (Subsection B). Luning (1969) claimed that the leaf area index (the ratio of the surface area of photosynthetic laminae to projected surface area of the site) is closely correlated with depth and presumably with underwater irradiance. E. CYTOLOGICAL ASPECTS

1. Cell Morphology The importance of cell shape and size in phytoplankton has already been discussed (Subsection D. 1). In multicellular algae cell shape and size also have an important bearing on light harvesting although this has received little attention. As mentioned above Ramus (1978) has shown that light absorption in Codium fragile is very efficient and proposed that this is due to a special anatomical feature. In this coenocytic alga terminal branches (ramuli)


29

fuse to form an unbroken surface. Chloroplasts are located in these surface ramuli. Ramus (1978) suggested that this arrangement acts as a light guide and also leads to multiple scattering from the internal non-photosynthetic tissue especially when these become air-filled, which overall results in an almost black-body absorption of PAR (Fig. 17). The enhancement of scattering and light absorption by air-filled tissue has been discussed earlier (Seybold, 1932, 1933, 1934). A similar increase in the efficiency of light absorption perhaps occurs in other algae such as crustose coralline algae. 2. Chloroplast Movement A powerful means of varying light-harvesting in plants is by displacement and reorientation of chloroplasts. This tactic response of plants has been known for over a hundred years, and the early work is discussed by Rabinowitch (1951) who stated that “The largest variety of chloroplast movement has been observed by Senn in green and brown algae and in diatoms.” Senn (1908, 1919) carried out detailed studies on Dictyota dichotoma (Phaeophyta) and other algae. Two basic responses occur. In bright light chloroplasts may reorientate along the cell membranes parallel to the incident light (parastrophe) causing much self-shading. In dim light chloroplasts reorientate along membranes perpendicular to the incident light (diastrophe). Seybold (1933) estimated that a single higher plant chloroplast absorbed between 30-60 per cent of incident light, transmitted between 30-60 per cent and scattered about 10 per cent. Kirk and Goodchild (1972) have largely confirmed these estimates, and also further provide evidence of the absorption and scattering of light of various wavelengths. Theoretically reorientation of chloroplasts should be able to modify greatly light absorption in plants. Evidence of this in practice is not substantial. The most conspicuous example was observed by Britz and Briggs (1976) in Ulva lactuca (Chlorophyta) where the absorbance at 681 nm increased from 0.3 in darkness to 0.9 near midday. However this change is the reverse of those observed above. The absorbance changes in Ulva were indeed accompanied by chloroplast movement from side walls (darkness) to face-on walls (light) and bqth followed a circadian rhythm. However the following evidence indicated a minimal relationship with light-harvesting or protection against high light intensities, (i) chloroplast orientation was the same in low and high light intensities (up to 80 klux of white light), (ii) photosynthetic capacity was not well correlated with chloroplast orientation, (iii) no such movements occurred in the closely related chlorophytes, Enteromorpha intestinalis and Monostroma grevilfei.Similar circadian rhythms of chloroplast movement or conformational change have been observed in Caulerpa spp. (Chlorophyta) (Larkum, unpublished), Acetabularia mediterranea (Chlorophyta) (Van den Driessche, 1966), Pyrocystis sp. (Dinophyta) (Swift and Taylor, 1967). Gonyaulax polyedra (Dinophyta) (Herman and Sweeney, 1975).

30


Light-induced chloroplast movements of a photo-protective kind (i.e. less absorbance in high light) have been observed in Fucus vesiculosus (Phaeophyta) (Rufferet al., 1978) in Dictyota dichotoma (Pfau et al., 1979; Ruffer et al., 1981) and in Mougeotia and Mesotaenium (Chlorophyta) (see Haupt and Thiele, 1961; Haupt and Bock, 1962; Schonbohm, 1965, 1978). In Dictyota and Fucus absorbance increased between bright and dim light by 11-20 per cent and increased further in darkness to 40 per cent in Fucus. These changes were accompanied by chloroplast movements: in Dictyota from sidewalls to face-on walls, and in Fucus, in cortical but not epidermal cells, from a position near the centre of the cell around the nucleus to the face-on walls. In Mougeotia and Mesotaenium the rectangular chloroplast moves to present its maximum profile area in weak light and minimum in strong light. These movements are triggered by phytochrome located in the cytoplasm (Haupt and Thiele, 1961; Haupt and Bock, 1962; Schonbohm, 1965, 1978). The presence of phytochrome and light-triggered responses in deep-water algae is discussed in Section XI. Diurnal movements of another kind occur in various species of Siphonales (Chlorophyta). These algae are formed of coenocytic branched hyphae which end at the surface in a fused cortical layer. Chloroplasts move towards the surface in light, and away from it in darkness. This occurs in Caulerpa spp. (Dawes and Barilotti, 1969) and in Halimeda tuna (Larkum, unpublished). Light-induced changes in chloroplast shape and size (conformational changes) are found in Ulva (Chlorophyta) (Murakami and Packer, 1970), Acetabularia (Chlorophyta) (Van den Driessche and Hars, 1972) and Fucus vesiculosus (Phaeophyta) (Ruffer et al., 1978). The only study of the effect of such changes on transmission properties (Packer et al., 1967),carried out only at one wavelength (546 nm), showed a transmittance decrease of 0.3-18 per cent in the light, due mainly to decreased absorption but also to decreased light scattering (Murakami et al., 1975). In summary, present evidence on chloroplast movement and conformational change do not support the idea that such changes greatly affect the light harvesting properties of algae cf. Ruffer et al. (1982). At least two poorly understood processes are involved, as well as that of light harvesting: photoprotection against damagingly high levels of irradiance and diurnal changes. The relationship of these two processes to light-harvesting mechanisms requires more investigation. Conformational changes in chloroplasts are undoubtedly related to osmotic changes and changes in membrane thickness (Murakami and Packer, 1970; Murakami et al., 1975), but it is doubtful whether the underlying mechanism (ion transport and energy transduction in the thylakoid membrane) plays a significant role in controlling the light harvesting properties of algae. 3. Chromatophore and Chloroplast Structure and Arrangement ( a ) Introduction. Chloroplast structure shows greatest variation throughout the mrimir nhvla nf aloae. A t nresmt there i s no simnle formulation of the


31

principles governing this structural variation; evolutionary diversification appears to have arisen at an early stage (Section XII). The structure and rearrangement of chloroplasts within cells is undoubtedly related to lightharvesting but aside from phylogenetic affinities other constraints such as diffusion of inorganic carbon species, the concentration of carbon-fixing enzymes and the supply of ATP and NADPH influence structure (Raven, 1977,1978; Krause and Heber, 1976; Farquhar and Von Caemmerar, 1981; see Subsection F). ( h ) Chromatopkores. Chromatophores occur in four groups of photosynthetic prokaryotes. These are the purple bacteria, the green bacteria, the Cyanobacteria and the Prochlorophyta. The first two groups have BChl and perform anoxygenic photosynthesis whereas the two latter groups contain Chl a as the major photosynthetic pigment and carry out oxygenic photosynthesis. A full account of chromatophore structure is to be found in Remsen (1978). In purple bacteria the chromatophore membranes are extensions of the plasma membrane. They contain a part or all of the photosynthetic pigments (Gibbs et ul., 1975). Chromatophores may form small vesicles attached to the plasma membrane, or long sacs extending from the plasma membrane, or they may form stacks of lamellae with no obvious connection with the plasma membrane. However in all cases the structure is vesicular, that is a membrane divides off an inside compartment (which, theoretically, should be connected to the outside of the organism) and an outside compartment, the cytoplasm. Thus chromatophores are thylakoids, as defined by Menke ( 1 962). The thylakoid membranes of photosynthetic bacteria are approximately 7-8 nm in thickness. Folding of the membrane may be quite complex. In many types of purple bacteria the lamellae lie parallel to one another separated by a cytoplasm-filled space of approximately 10 nm (Remsen, 1978). However appression of lamellae can occur (Hickman and Frenkel, 1965; Holt et al., 1966;Gibbs et a / . ,1975; Remsen, 1978) such that the lamellaecome into much closer contact (approx. 1.5 nm). This is an important point since appression of lamellae in chloroplasts of many algal divisions has been thought to represent an advanced characteristic (Coombs and Greenwood, 1976); it is also related to light-harvesting characteristics (Section IX). Gibbs et al. (1975) showed that the degree of appression increased with shading in cultures of the purple bacterium, Rhodospirillum molischianum. Thus the same relationship exists here between appression and shading as in algae and higher plants (Section IX). The carbon-fixation enzymes, now thought to be those of reductive pentose pathway (cf. Fuller, 1978) are in the soluble cytoplasm. RuBP carboxylase exists either in soluble form or as polyhedral inclusions (Fuller, 1978), which presumably act as storage sites. However the suggestion has recently been made that these “carboxysomes” are primitive organelles which carry out the complete cycle of photosynthetic CO, fixation (Beudeker and Kuenen, 198 I). In Cyanobacteria, which carry out oxygenic photosynthesis, chromat-

32


ophores occur as folded sheets of thylakoids which have little obvious connection with the plasma membrane. The thylakoid or lamella membranes are 7-8 nm in thickness. Thylakoids may be arranged in a variety of ways and often aggregate in parts of the cell to form regions where the membranes align closely parallel to one another (Figs 12b and 13a). However the membranes are never appressed: the phycobilisomes (PBS) which contain the phycobiliproteins (Section VI) are located on the outer face of the thylakoid membranes and restrict the approach of neighbouring membranes to about 40 nm. In Prochlorophyta (Lewin, 1976, 1977), which are also oxygenic prokaryotes but which contain Chl b in addition to Chl a (see Section XII.C), the thylakoids lie in close proximity to one another to form a chromatophore which fills a large proportion of the cell (Fig. 12) (Whatley, 1977; Giddings et al., 1980; Cox and Dwarte, 1981). A large proportion of the thylakoid membranes are appressed, and as many as 20 thylakoids may be stacked together. This is particularly marked in the Prochloron sp. from Didemnum molle (Fig. 12) (Cox and Dwarte, 1981). The thylakoid membrane in unstacked regions is about 8-10 nm in thickness. The thylakoids may be separated by much cytoplasm in unstacked regions. In stacked regions thylakoids are separated by less than 1 nm at most, to form a fused double layer of about 15 nm thickness. (c) Green algal phyla. In those phyla of algae in which Chl b occurs (e.g. Chlorophyta, Euglenophyta, Prasinophyta, Charophyta), the thylakoids of chloroplasts have a membrane structure and arrangement (Fig. 13) not unlike that of the chromatophore of Prochloron. Thylakoids lie parallel to one another and are appressed at intervals to form stacks of from two to many thylakoids. There are many arrangements of stacking from very regular to very irregular, from long fused thylakoids to short stacks (similar to the grana of higher plants) and from stacks of two to stacks of five or more thylakoids (Coombs and Greenwood, 1976; Kirk and Tilney-Bassett, 1978). The thylakoid membranes are approximately 10 nm thick in non-appressed regions and 12nm in appressed regions. However the thickness of the membranes is not static; shrinkage occurs during the transition from dark to light and the reverse on return to darkness (Murakami and Packer, 1970).The centre-to-centre distance of appressed thylakoid membranes is approximately 18 nm. (4 Chl c-containing algae (except Cryptophyta). Algae of the superphylum Chromophyta, with the exception of the Cryptophyta, are characterized by a thylakoid system in which thylakoids are arranged basically in groups of three (Fig. 13) and appression of adjacent thylakoid membranes occurs along much of the length of each group (cf. Coombs and Greenwood, 1976). There may be some variation with groups of two, four, five and six occurring less commonly. Humphrey (1983) found in two phytoplankton species that the number of thylakoids in stacks was variable and could be

Fig. 12. (a) An electron microscope section of a cell pf Prochloron (Prochlorophyta) from Dzdemnum molk ( x 15 000). (b) An electron microscope section of Gloeocupsa NS4 (Cyanobacteria), an extreme shade alga found in caves (Cox et d.,1982). Both cells were fixed in

glutaraldehyde and osmium tetroxide and post-stained in uranyl acetate and lead citrate. (Micrographs by G . Cox, Electron Microscope Unit, University of Sydney.)

Fig. 13. Electron microscope sections to show characteristic thylakoid structure of (a) Lyngbya sp (Cyanobacteria), (b) Haliptilon cuvieri (Rhodophyta), (c) Chroomonas sp (Cryptophyta), (d) Prorocentrum micans (Dinoflagellata), ( e ) Ecklonia radiata (Phaeophyta), (f) Dunaliella tertiolecta (Chlorophyta) ( x 60 000). Materials fixed in glutareldehyde and stained in osmium tetroxide. (Micrographs by M. Vesk, Electron Microscope Unit, University of Sydney.)


35

correlated with the growth irradiance and its spectral composition. In general the thickness of the thylakoid membranes is approximately 1 1 nm for nonappressed and 15 nm for appressed membranes. The centre-to-centre distance of appressed membranes is very variable but is approximately 20 nm: which is greater than for the green algal types and may reflect a greater thickness of the inter-thylakoid space. It has been claimed that in the Phaeophyta the thylakoids are not truly appressed but are separated by a space approximately 4 nm in width (cf. Kirk and Tilney-Bassett, 1978). Nevertheless disrupted chloroplasts from Phaeophyta still retain the characteristic arrangement of thylakoids in threes indicating that the mechanism of appression involves tight binding of neighbouring membranes. A further feature of the Phaeophyta, the Bacillariophyta and the Chloromonadophyta, but not the other phyla, is a peripheral or girdle lamella or thylakoid which encircles the plastid just inside the chloroplast envelope. Girdle lamellae are found also in many Rhodophyta and some Chlorophyta. ( e ) Rhodophj-fa.The arrangement of thylakoids in the Rhodophyta (Fig. 13) is somewhat similar to that in Cyanobacteria except that they lie within a chloroplast and there is often a girdle lamella present. The thylakoids lie singly in the stroma, often parallel to one another but separated by a space 40-50 nm thick. The spacing is occasioned by the presence, as in Cyanobacteria, of PBS, with a diameter of 3 0 4 0 nm. The PBS are arranged in regular arrays and are attached by a stalk to the adjacent thylakoid membrane. In close-packed configuration (in shade algae) the PBS, attached to one membrane, alternates with the phycobilisome attached to the neighbouring membrane (Fig. 28; Section 1X.B). However much looser arrangements are seen in sun plants. PBS of two types occur: the cyanobacterial semi-discoid type occurs in some Protoflorideae and the globular type in Euflorideae (Section VI1.B). The close-packing of these two types is therefore quite different (Section 1X.B). The thickness of the thylakoid membrane is approximately 5 nm (Wanner and Kost, 1980). ( f ) Crjptophjlta. In the Cryptophyta, thylakoids are characteristically found in pairs (Fig. 13) although triplets also occur. The thylakoids are rather loosely held together in many species but more tightly in others. The thylakoid membrane thickness is quite variable and may be as thick as 36nm (Wehrmeyer, 1970). Although phycobiliproteins are present in Cryptophyta, PBS are absent and the phycobiliproteins are probably present in the intra-thylakoid compartment which is filled with a finely granular, electron-dense material (Gantt et al., 1971; Faust and Gantt, 1973). (g) Chloroplast arrangement. Light harvesting on a cell basis can theoretically be increased by increasing the number or size of chloroplasts per cell. However, beyond the limit of a single layer of chloroplasts around the periphery of the cell this is a process of diminishing returns. Self-shading is

36

A. W . D. LARKUM AND JACK BARRETT

clearly a factor but diffusional-limitation of inorganic carbon supply is probably of greater importance (Nobel, 1974; Raven, 1977). Under high light conditions the carboxylating machinery of the outermost layer of chloroplasts can effectivelyutilize all the carbon dioxide diffusing into a cell (Subsection F). Thus a second layer (or larger chloroplasts) would be of little use. Under low light conditions much more carbon dioxide is available inside the first layer and it may be “profitable” in terms of light harvesting for plants to arrange a second layer of chloroplasts. However, in terms of overall economy of cell protein and other constituents it may not be a profitable adaptation. Most photosynthetic cells have the equivalent of at most a single layer of chloroplasts (arranged immediately inside the plasmalemma), although this may occasionally increase to the equivalent of two layers in low light conditions. When chloroplasts are found several layers deep in only a part of a cell and not in a layered arrangement e.g. in the lower part of epidermal cells of Fucus vesiculosus (Ruffer et al., 1978), it is possible that other factors such as photoprotection are involved. F. BIOCHEMICAL STRATEGIES OF LIGHT HARVESTING

I . Pigments Although photosynthetic pigments are dealt with in detail in Section VI, it is necessary to make some general points here. All algae have Chl a as an obligate component of PSI and PSII, and in addition show a great diversity of photosynthetic pigments. All these pigments mainly absorb light only in the 300-700 mm region of the spectrum. This narrow spectral range contrasts with the spectral limits of photosynthetic bacteria where certain BChls absorb light up to at least 900 nm (Thornber et al., 1978). No clear reason can be given for the narrower range in algae, although the upper limits may have been determined by the fact that above 700 nm the quantum energy may be too low for the “Z” scheme (Section IV) to operate. The presence of a water-splitting oxygen-evolving system in algae imposes constraints that do not apply to the anaerobic photosynthetic bacteria. Evolutionary events have almost certainly been of importance (Section XII). Since water is the source of hydrogen equivalents in all such organisms it is reasonable to assume that the earliest forms existed where water was plentiful, and since water strongly absorbs infrared and ultraviolet light (Section 111) the evolutionary pressure for pigments absorbing in the visible region of the spectrum would have been considerable. Chl a-proteins in vivo have absorption maxima at 435 nm and 675 nm but absorb poorly in the green and yellow regions of the spectrum. In all algae there is evidence for the evolution of accessory pigments to cover the green and yellow “windows” in the Chl a spectrum. Chl b, c,, and c2 fulfill this role but not very efficiently. Some carotenoids are more effective (Section V1.B)


37

but leave the yellow and orange regions of the spectrum largely untapped. Phycobilins fill this remaining gap by absorbing green, yellow and orange light (Section VII1.B). Thus there are photosynthetic pigments which absorb all regions of visible light. Yet in only three groups of algae, Cyanobacteria, Rhodophyta and Cryptophyta, is this ability exploited to the full, by the presence of phycobiliproteins. At the other extreme there are the Chlorophyta (and related groups) and all higher plants which, with rare exceptions (Kageyama et al., 1977; Anderson et al., 1980), possess no photosynthetic pigments which can absorb efficiently between 500 and 620 nm, a region which contains 55 per cent of visible irradiance in terrestrial habitats and between 55 per cent and 95 per cent in shallow underwater habitats. There are a number of possible explanations for this apparent deficiency:

(i) other mechanisms such as pigment concentration or chloroplast structure lead to strong absorption of green and yellow light (see Section V.G); (ii) the light harvesting system based on Chl a and b and carotenoids alone is more efficient than those involving either Chl c or phycobilins as well; (iii) early evolution of photosynthetic pigment arrays became “fossilized” after the endosymbiotic developments involved in the evolution of eukaryotes (see Section XU). A further explanation might be that such plants live in environments whose light is never limiting, but this is certainly not a convincing argument for algae since Chlorophyta as well as other algae are found in a number of shaded habitats (Larkum et al., 1967). All three explanations may have validity. 2. Pigment Arrays A basic condition of photosynthetic pigments in all photosynthetic organisms, including photosynthetic bacteria, is the arrangement of pigments in cooperative arrays (Junge, 1977). These arrays, composed of pigment-proteins (Section VII), greatly increase the probability of lightcapture of each unit by increasing the optical cross-sectional area. This allows a greater turnover of each reaction centre, which carries out photochemical conversion (Section IV), and results in a more efficient use of the photosynthetic machinery. A further consideration for PSI1 is the need for four photochemical events in quick succession for the efficient release of oxygen from water (Kok et al., 1970; Diner and Joliot, 1977)and this is clearly closely related to the pigment arrays available to PSII (Anderson, 1981). Pigment arrays may be classified into two broad groups (i) antenna systems which are core units of PSI and PSII, and (ii) light-harvesting pigment systems that do not possess a reaction centre and transfer their absorbed energy to PSI or PSII or both (Section VIII; also Hiller and Goodchild, 1982).

38


3. Photosynthetic Unit The concept of the photosynthetic unit (PSU) is based on the work of Emerson and Arnold1 (1932a) hhich indicated that approximately 2500 Chl molecules were involved in the evolution of each molecule of oxygen. In its original definition the PSU was simply a statistical unit of cooperative chlorophyll molecules. However there have been various attempts to show that the PSU is a structural entity in the thylakoid membrane (e.g. Park, 1965), since the general adoption of the series formulation of two photosystems (Section IV). The PSU concept would assign some 300 chlorophyll molecules to each reaction centre (assuming 8 photoacts are involved in the evolution of each oxygen molecule). A great amount of work, especially on algae, has attempted to relate the size of the photosynthetic unit to the light-climate experienced by a plant (Section 1X.C). Much work was based on the ratio of Chl a molecules to P-700 and assumed that the number of PSI and PSII reaction centres were equal. However it has become clear that the ratio of PSI and PSII reaction centres is not unity (Melis, 1978; Kawamura et al., 1979; Melis and Brown, 1980; Anderson, 1981; Falkowski et al., 1981). As a result the photosynthetic unit concept is of limited usefulness and it is more appropriate to consider PSI and PSII units separately, defined in terms of the number of Chl a molecules and other light harvesting pigment molecules per unit (cf. Falkowski et al., 1981). Attempts to relate PSI and PSII units to structural entities are discussed in Section 1X.B. 4 . Membrane Structure Chlorophylls and other photosynthetic pigments are located in proteincomplexes (Sections VII and VIII). These complexes are integral or peripheral proteins of the thylakoid membrane and their arrangement in appressed and non-appressed regions is probably of prime importance to the distribution of absorbed energy, and this matter is discussed in detail in Section 1X.B and XI1.D and in the following Subsection. 5. Electron Transport

The absorbed light energy of photosynthesis is stored temporarily in the form of NADPH and ATP, both of which are generated by an electron transport chain; NADPH is produced directly and ATP indirectly through a gradient of protons across the thylakoid membrane (Section IV; Fig. 9). The proton gradient appears to be generated largely by the flow of electrons between PSII and PSI involving the electron carriers, Q, plastoquinone and cytochrome b6-f complex (Trebst, 1974; Junge, 1977; Hurt and Hauska, 1981). The simplest hypothesis to account for these various processes is an interconnected linear sequence of PSII, PSI and electron transport components operating as a structural entity in the thylakoid membrane (Fig. 14). Such an hypothesis fits well with the widely-held view that PSI and PSII units have approximately


hVTl

qhV'

-I

H,O

Q

39

-

-,-

I

-0 - -r +Q -

I

0 2ms

-1-

+PQ - -1-

I

I

540 545 > 563 (575) 545 > 563 > 498 567 > 538 >498 541-565

Fluorescence emission max (nm) 680 660 680 637 636

619d 577 570 575 578

Chromophore and number achain B-chain 1PCB 1PCB 1PCB 1PCB lPCB 1PCB 1PXB' ZPEB ZPEB ZPEB ZPEB PEB

IPCB" 1PCB

1PCB 1PCB (1PCBIPEB) IPCB, lPEB 2PCB 4PEBb 4PEB 4PEB" 4PEB" PEB

Monomer

Protein structure aggregation

a3B3

.B aB aB a@ aa'h aB

aB aB %B6Ya

abP6Y"

aB

1 (1)3,6 (1)3,6 1,3,6 3,6 1

3 1,3,6 3 1 1 1

In B-Phycoerythrin and R-Phycoerythrin, the additional y-subunit carries 2 PEB and 2 PUB, which results in the distinctive 498 nm peak.

* Muckle and Rudiger (1977) reported an arrangement ZPEB, 3PEB for one cyanobacterium.

Chromophore undetermined (see text). Nies and Wehrmeyer (1980); MacColl er a/. (1981). ' An A, at 575 nm sometimes found (Zilinskas et a/., 1978). b-PE may be a glycoprotein (Chapman, 1973). For presence of the PUB chromophore in certain cyanobacterial phycoerythrins see MacColl, 1982. Note: The phycobiliproteins of Cryptophyta (K-PC and K-PE) have been less studied and cannot easily be compared with those of Cyanobacteria and Rhodophyta (Gantt, 1979; Glazer, 1981). The phycoerythrins contain only PEB chromophores but the phycocyanins contain PCB and possibly a phycobiliviolin similar to that found in Phycoerythrocyanin (Glazer, 1981). References to the original literature to be found in Glazer (1977, 1981), Rudiger (1980) and Scheer (1981).


65

Anabaena variubilis and Mu.ytigocludus laminosus (Bryant et al., 1976, 1979; Nies and Wehrmeyer, 1980; MacColl et a/., 1981). Minor chromophores of unknown structure are the third chromophore (Psq0)of the PC from a species of Hemiselmis (Cryptophyta) (Glazer and Cohen-Bazire, 1975) and the blue chromophore of Jung et a/. (1980). A phycobilin with an interrupted conjugation, the yellow phycourobilin (PUB), occurs in the /$chain of B- and R-phycoerythrin and in certain Cyanobacteria (McColl, 1982). Other chromophores have been tentatively detected in red algae (O'Carra and O'Heocha, 1976; Glazer and Cohen-Bazire, 1975). It should be noted that the structures of the free phycobilins are not identical with those of the covalently bound chromophores. In Callitliamnion roseum (Rhodophyta) the ratio of phycoerythrobilin to phycourobilin is modulated by variations in light intensity (Yu et al., 1981). Table 111 summarizes the types, occurrence and properties of phycobiliproteins. There are basically three types: phycerythrin (PE), absorbing in the green region (495-570 nm), phycocyanin (PC) absorbing in the green to yellow region (550,630 nm) and allophycocyanan (APC) absorbing in the orangeered region (650-670 nm). An extensive array of PC, PE and APC are known to have their phycobilins covalently bound by thioether linkage, through cysteine, to the pyrrole ring B of the tetrapyrrole (cf. Scheer, 198 1) Fig. 2 1. The chromophore PUB may have a second thioether link to pyrrole ring D.

H

H

H

Fig. 21. Schema of attachment of (a) phycocyanobilin to polypeptide back-bone in phycocyanin, and (b) phycoerythrobilin to protein in phycoerythrin.

66


The involvement of a second attachment site between the chromophore and the protein is less certain. The propionic carboxyl of pyrrole ring C in the PC of Spirulina platensis (Cyanobacteria) was demonstrated to be attached to the polypeptide by chromic acid degradation to the imide (Scheer, 1981). In a different approach the free carboxyls of chromopeptides from PE of Anabaena variabilis (Cyanobacteria) were conjugated with glycine, and the PCB obtained on hydrolysis of the chromopeptide had only one free propionic carboxyl (Barrett, unpublished). Evidence has been presented that serine is the amino-acid involved in the binding of phycobilins (cf. O’Carra and O’Heocha, 1976; Muckle et al., 1978).A PCB-dipeptide containing cysteine and glutamic acid was obtained from PC of A . variabilis (Barrett, 1968).Bindings of PEB to a chromopeptide through glutamate was found in a cryptophyte (Brooks and Chapman, 1972). A PCB-tyrosine ester bond was the second attachment to the a-chain of C-phycocyanin in Oscillatoria agardhii (Cyanobacteria) (Wallin et al., 1978). Complete amino-acid sequences are known for the PC from Mastigocladus laminosus (Cyanobacteria) (Frank et al., 1978), the PC ci and P subunits of Cyanidium caldarium (Rhodophyta) (Troxler et al., 1981; Offner et al., 1981) and the APC of M . laminosus (Zuber, 1978; Sidler et al., 1981). The amino acid sequence of the P-chain of PC of Synechococcus spec. 6301 (Cyanobacteria) has been determined (Freidenreich et al., 1978). The polypeptide sequences of the respective a- or P-chains of M . laminosum and of Synechococcus (Cyanobacteria) both thermophilic algae, show homologies of 80 and 78 per cent, respectively. Comparison of the primary structure of the APC with the PC from M . laminosum shows that the P-chain has an extra peptide sequence of ten amino acids to accommodate the second PCB. There are differences in the homology of the amino acid sequences of the four subunits of either APC and PC, and between the subunits of both APC and PC. Sequence homologies between the a- and P-chains of PC from these two thermophilic algae and that of a marine Cyanobacterium Agmenellum quadruplicatum (80 per cent of sequence determined: Gardner et al., 1980) is only about 30 per cent. A high degree of homology has been found in the chromophore binding sequence of PC chromopeptides from several sources (Byfield and Zuber, 1972; Bryant et al., 1978; Freidenreich et al., 1978; Williams and Glazer, 1978; Lagarias el al., 1979) and in a PE chromopeptide (Muckle et al., 1978). A similar level of homology of the N-terminal regions has been noted (Glazer, 1977). The geometry of phycobilins attached to proteins differ from that of free bile pigments, such as urobilin which forms a cyclohelical porphyrin-like structure in solution (Moscowitz et al., 1964), or the extended and twisted linear form of biliverdin revealed by X-ray crystallography (Bonnett et af., 1978).ORD studies showed that the bound chromophore of undenatured PC of Anacystis nidulans (Cyanobacteria) was in a chiral configuration and this


67

chirality was dependent on the helicity of the protein (Barrett, 1968). The high absorbance of the red peak of PC is due to the chromophore being rigidly fixed in an extended conformation (Scheer and Kufer, 1977), Fig. 22. C D studies of the geometry of PE suggests a chiral conformation for the bound chromophore (Langer et al., 1980).

Protein

7

COO-

f

CO-Protein

(

coo-

Fig. 22. Conformation of the chromophore of phycocyanin; (a) cyclohelical (b) extended, sterically unhindered (Scheer, 1981).

Exciton-interaction (Section 1X.A) between chromophores on the same polypeptides, or adjacent polypeptide, may be responsible for the significant differences in the spectra between phycobiliproteins which have the same molecular structure and similar chromophore geometry. Fluorescence properties are dependent on these interactions. In picosecond pulse fluorescence studies with Nostoc (Cyanobacteria) phycobiliproteins the results indicated the occurrence of singlet-singlet annihilation being shared among the several phycobilin chromophores within the individual phycobiliprotein molecules (Wong et al., 1981). The phycobiliproteins are sensitive to perturbation of their absorption and fluorescence properties by modification of their chemical and physical environment to a greater degree than is the case with the Chl-proteins (Grabowski and Gantt, 1978a,b; Schreiber, 1979, 1980; Fisher et al., 1980; Glazer, 1981; Kost et al., 1981; Zickendraht-Wendelstadt et al., 1980). This susceptibility to perturbation of the fine structure of the phycobiliproteins is extremely pertinent to their energy transfer properties (Section 1X.A). Secondary and tertiary structure of phycobiliprotein monomers has not been investigated extensively (cf. Scheer, 1981). Fisher et al. (1980) have reported on the structure of C-PC and R-PE from X-ray diffraction studies at

68

A. W. D . LARKUM A N D JACK BARRETT

5.0 A resolution. However the C-PC was in oligomeric form. Both had a 3-fold symmetry. B-PE appeared to be a 32-D, symmetric stack of two (a& trimers with a y-chain fitted in the centre. C-PC appeared to have a 3-C, symmetry with three (a/?)2dimers arranged around a central solvent channel. In the CPC there were long columnar regions of density which were presumed to be uhelix arrangements. Low-resolution X-ray and solubilization studies of the CPC from the cyanobacterium Agmenellum quadriplicatum suggests that the monomer (34 KD) forms dodecamers in which the hydrophobic /?-chains of the subunits of the monomer are oriented to the inside of the dodecamer (Hackert et al., 1977; Gardner et al., 1980). Circular dichroism measurements on several phycobiliproteins indicate an cc-helix content of about 60 per cent in the a- and 40 per cent in the /?-chains (Brown et al., 1975; Langer et al., 1980). Canaani and Gantt (1980) studied the APC species from Nostoc sp. (Cyanobacteria) for circular dichroism; and concluded that these molecules contained 2-13 per cent pleated sheet and 18-43 per cent cr-helix arrangements. The quaternary structure of phycobiliproteins has been extensively studied (cf. Gantt, 1980; Glazer, 1981; Scheer, 1981). Although each phycobiliprotein is basically composed of two polypeptides, an cc-chain of 16-19 K D and a 8chain of 19-21 KD, oligomers appear to predominate in vivo (Table 111) with the trimer ( a m , as a common arrangement. As shown by electron microscopy (e.g. Morschel et al., 1977; Koller et al., 1978; Morschel er al., 1980) and by Xray diffraction (Fisher el al., 1980) the trimeric arrangement gives rise to a planar ring-shaped or disc-shaped molecule. However other arrangements also occur (Table 111) and in the Cryptophyta dimers are common (MacColl et al., 1976). Immunochemistry has shown that all the APCs from Cyanobacteria and Rhodophyta are closely related and the same applies to PC and PE (cf. MacColl and Berns, 1979; Scheer, 1981). In contrast APC, PC and PE do not undergo cross-reactions. In the Cryptophyta both PC and PE cross-react with R-PE but not with C-PE indicating a relationship with the Rhodophyta (MacColl and Berns, 1979)and suggesting the possibility of the late evolution of Cryptophyta from the Rhodophyta (Section X1I.E). D. ACTION SPECTRA A N D QUANTUM YIELDS

1 . Action Spectra

(a) Oxygen evolution. Action spectra for photosynthesis in algae were first obtained a century ago (Engelmann, 1883, 1884). Action spectra have been used to determine which pigments are active in harvesting light for photosynthesis. However, the recent identification of a number of partial reactions of photosynthesis has allowed a more refined probing of the role of lightharvesting pigments in PSI and PSII. A photosynthetic action spectrum is defined as “the photosynthetic rate per


69

unit of irradiance as a function of wavelength". If the irradiance is in the region where photosynthetic rate is linear with intensity, then the effectiveness of various wavelengths can be assessed and by comparison with the absorption spectra of the pigments present, those pigments contributing to the photosynthetic response can be identified. By this means Engelmann (ibid) was able to demonstrate that Chls, carotenoids and phycobiliproteins are major light-harvesting pigments. The modern era was introduced with the use of the bare platinum electrode to measure action spectra in Chlorophyta, Phaeophyta, Rhodophyta and Cyanobacteria (Haxo and Blinks, 1950). This technique is only semiquantitative since not all the oxygen evolved is measured and the algae are held under unstirred conditions; furthermore it is restricted to chloroplasts, unicellular algae and some thin membranous algae. Nevertheless the technique yielded valuable results and has been used by a number of workers (Fork, 1963; Jones and Myers, 1965; Joliot and Joliot, 1968; Prezelin et al., 1976; Wang et al., 1977). The spectra obtained before 1960 suffer from a lack of appreciation of the need to balance the light absorption of PSI and PSII. This is specially true with Rhodophyta where the wavelength maxima of the various light harvesting pigments are widely separated and the distribution of Chl between the two photosystems may be very unequal. As a result Haxo and Blinks (1950) found that Chl was largely "inactive" in harvesting red light in both Rhodophyta and Cyanobacteria-whereas dual wavelength studies (Fork, 1963; Jones and Myers, 1965; Larkum and Weyrauch, 1977; Wang et al., 1977; Ley and Butler, 1977b), (Fig. 24), have shown that Chl a is a light harvesting pigment mainly of PSI in the majority of species (but see Ley and Butler, 1980a and Myers et al., 1980 for examples of a more equal distribution of Chl). The Clark-type oxygen electrode (Delieu and Walker, 1972) allows quantitative measurement of photosynthesis (oxygen evolution) and has been used in studies of action spectra of algae (e.g. Larkum and Weyrauch, 1977). In summary the large body of evidence in this area has established, along with other lines of evidence, a role in light-harvesting for the following (see Fig. 23): Chl a Chl b Chl c

in all algae in Chlorophyta, Euglenophyta and Prasinophyta (Haxo and Blinks, 1950; Haxo, 1960). (probably) in Phaeophyta, Bacilliarophyta and Dinoflagellata (Haxo and Blinks, 1950; Haxo, 1960; Mann and Myers, 1968; Iverson and Curl, 1973; Prezelin et al., 1976). The area of doubt arises from the small peaks of absorption in the yellow and red spectral region-at 579 and 628 nm and the larger peak at 453 mm in the blue region which is masked by the activity of fucoxanthin or peridinin.

70


in many algae (Haxo and Blinks, 1950) with a high efficiency for fucoxanthin in Phaeophyta and Bacillariophyta (Haxo and Blinks, 1950; Tanada, 1951; Mann and Myers, 1968) and peridinin in Dinoflagellata (Prezelin ef al., 1976). in Rhodophyta, Cryptophyta and Cyanobacteria (Haxo Phycoerythrin and Blinks, 1950; Haxo and Fork, 1959; Haxo, 1960; Jones and Myers, 1965; Larkum and Weyrauch, 1977; Ley and Butler, 1977a,b; Lichtle et al., 1980). in Rhodophyta, Cryptophyta and Cyanobacteria (rePhycocyanin ferences as for -phycoerythrin) Allophycocyanin in Cyanobacteria and -Rhodophyta (Lemasson et al., 1973; Larkum and Weyrauch, 1977). Carotenoids

In general, it may be said that the action spectrum of an alga is similar to the absorption spectrum in the visible range provided the activities of the two photosystems are balanced. Thus the presence of a large absorption band generally indicates a light-harvesting pigment (or pigments). However, Ulvo toeniata t*thalius absorption

Coiloderme thollus absorptioi action specrrum

60 40

40

20

20

400440 480 520560 600 640 680720760 400 440480 520 560 600640680 720 Porphym nereocystis

p thallus absorption * oc1mn spectrum

..,,.,... absorption action

40

g 400

500

600

700

400440480 520 570600640 680 720

*

-*actonspectrum

60

20 400

500

600

Wovakqth,nm

700

400

500

600

Wavelength, nm

700

0.2

0.I

B 669 nm I

J

0.2

1%

C 545nm

0.I

700 60

-

546 m p

-

0-

'

400

L,

I

L

500

I

600

.A',

700

Wavelength, nm

Fig. 24. Action spectra for red algae supplied with background monochromatic light. Top four curves Grifithsia monilis from Larkum and Weyrauch ( 1977). Bottom graph Porphyru perforalu from Fork (1963). Fig. 23 (opposite page). Action and absorption spectra for six algae: Ulva raeniara (Chlorophyta) (Haxo and Blinks, 1950); Coilodesme (Phaeophyta) (Haxo and Blinks, 1950); Laminaria sacchnrina (Phaeophyta) (Halldal, 1974); Rhodomonas lens (Cryptophyta) (Haxo and Fork, 1959); GIenodinium (Dinoflagellata) (Prezelin el a / . , 1976).

72


discrepancies can occur especially in the blue region (e.g. Prezelin et a!., 1976; Thielen and von Gorkom, 1981).The occurrence of plastoglobuli containing carotenoids, and faulty technique (French, 1977) can lead to large discrepancies in the blue region. (6) Fluorescence. Fluorescence excitation spectra are a form of action spectra. This rests on the implicit assumption that Chl a is the photoactive pigment (Section IV) and that other light-harvesting pigments pass on absorbed energy efficiently to a small number of Chl a molecules. As early as 1943, Dutton et al. showed that light absorbed by fucoxanthin in Phaedoactylum sp. (Bacillariophyta) caused Chl a to fluoresce and concluded that fucoxanthin was a light-harvesting pigment. Haxo and Blinks (1950) also showed from fluorescence studies that in Rhodophyta and Cyanobacteria light absorbed by phycoerythrin was transferred to Chl a with high efficiency. In 1952, Duysens put forward a generalized scheme of “sensitized fluorescence” for a number of algae and pigments. This early work confirmed that various pigments, already indicated by oxygen-evolution techniques, had a role in light harvesting (e.g. Goedheer, 1972). Fluorescence action spectra have been used recently to show that siphonoxanthin harvests green light very efficiently (Kageyama et al., 1977; Anderson et al., 1980) in Ulva japonica (Ulvales) and Caulerpa cactoides (Chlorophyta). Fluorescence studies have recently proved to be a very elegant tool for investigating the fine structure of pigment assemblies (see reviews by Papagiorgiou, 1975; Butler, 1978; Lavorel and Etienne, 1977; Breton and Geancintov, 1980) and some of the major points will be dealt with in the following subsections and in Sections V1I.C and IX. (c) Partial Reactions in Photosynthesis. The ability to isolate, chemically or physically, PSI and PSII has allowed the measurement of action spectra for each of these photosystems, and has thus provided evidence of the association of light-harvesting pigments with one or other photosystem (e.g. Duysens and Amesz, 1962; Amesz and Duysens, 1962; Gott and Kok, 1962; Jones and Myers, 1965; Joliot and Joliot, 1968; Ludlow and Park, 1969; Wang et al., 1977; Mimuro and Fujita, 1977; Diner, 1979); the literature in this field is too extensive to cover individually and is dealt with in a number of reviews (Haxo and Fork, 1959;Govindjee and Zilinskas, 1974; Breton and Geacintov, 1980). Numerous reactions have been measured, such as reduction of dyes (DCPIP, ferricyanide, methyl viologen), the production of reducing or oxidizing equivalents (NADPH, hydrogen peroxide, oxygen), redox changes of cytochromes, bleaching of P700, fluorescence emission, fluorescence yields, fluorescence lifetimes and luminescence. In summary such studies have led to:

(i) the generalization that the Chla associated with PSI has a peak of absorption in the red region slightly but significantly red shifted in comparison with PSII (e.g. Junge, 1977).


73

(ii) The association with PSII of Chlb (Cho and Govindjee, 1970a, Govindjee and Zilinskas, 1974; Ludlow and Park, 1969), Chl c (Goedheer, 1970), phycobiliproteins (Duysens and Amesz, 1962; Amesz and Duysens, 1962; Ludlow and Park, 1969; Cho and Govindjee, 1970b; Wang et a/., 1977; Lichtle et al., 1980) and fucoxanthin (Goedheer, 1969); while ,Ocarotene is associated with RCI and RCII (Sections VI1.A and B). However, the finding (Murata 1969a,b) that the amount of fluorescence induced in either PSI or PSII by light harvesting pigments is variable and dependent on environmental effects led to the hypothesis of transfer of excitation energy between the two photosystems (e.g. Butler, 1978) often called spillover; this phenomenon is dealt with in greater detail in Section 1X.B. Cho and Govindjee (1969a) showed that some PSI fluorescence was sensitized by Chl h in Chlorellu sp. (Chlorophyta). I t is also clear that in Rhodophyta and Cyanobacteria, green light, 90 per cent of which is absorbed by phycobiliproteins (Emerson and Lewis, 1943; Jones and Myers, 1965; Ley and Butler, 1980a,b), supports efficient photosynthesis (Wang et al., 1977; Larkum and Weyrauch, 1977; Ley and Butler, 1976); this implies equal sharing of the green light energy between PSI and PSII and demonstrates the difficulties inherent in assigning phycobiliproteins to one photosystem or the other (for further discussion see Sections 1X.B and X). The identification of short-wave red fluorescence (685, 695 nm maxima) with PSI1 and long-wave red fluorescence (710-740 nm) with PSI (e.g. Cho and Govindjee 1970a,b; Rijgersberg and Amesz, 1980) at 77°K has provided a useful tool for identifying the connections of light harvesting pigments to the photosystems. However interpretations on the basis of such evidence have to be made with care. Satoh and Butler (1 978b) found that the 695 nm (PSII) and 71C-740 nm (PSI) fluorescence are products of the very low temperatures at which the measurements are made. They suggested that energy migration was affected at these temperatures by changes in the interaction of the various species of light-harvesting (antenna) Chl a (Section VII). Thus one can expect that other delicate interactions such as the transfer of energy in either PSI or PSI1 may be affected by low temperature. Therefore, assignment of a pigment to one photosystem or the other on the basis of fluorescence measurements alone should be treated with caution (Wangetul.. 1980). Useful information is now being provided from pigment-protein complexes and detailed mapping of thylakoid membranes (Sections VII and 1X.B). 2. Quuntum Eficiencjl Quantum efficiency spectra differ from action spectra only in that the quanta absorbed, not the total number of incident quanta, are measured and their effect determined. The first work was carried out on Chlorella (Emenon and Arnold, 1932b) and action spectra were obtained for Chroococcus sp.

74


(Cyanobacteria) by Emerson and Lewis (1943). The claim by Warburg (cf. Kok, 1960) of quantum efficiences of 0.25 (that is 0.25 carbon dioxide molecules fixed per quantum of absorbed light) in Chlorella sp. stimulated a large body of research with various unicellular Chlorophytes and algae of a few other phyla, which indicated quantum efficiences in red light of between 0.08 and 0.125 (see Kok, 1960; Gaffron, 1960; Govindjee et ul., 1966; Radmer and Kok, 1977). Most of this work has been based on the saturating-flash techniques of Emerson and Arnold (1932a), see Fig. 25. Apart from the many studies with Chlorophyta, quantum efficiency spectra have been measured in Porphyridium cruentum (Rhodophyta) (Brody and Brody, 1962),in Chroococcus sp. (Cyanobacteria) (Emerson and Lewis, 1943) and in Navicula minima (Bacillariophyta) (Tanada, 1951), and in a number of marine benthic macroalgae (Yocum and Blinks, 1954, 1958).Some spectra are shown in Fig. 25. Unlike action spectra, spectra of quantum efficiency are rather flat in the visible range up to about 680 nm (after which efficiency declines rapidly-the “red drop”). This finding would seem to indicate that light absorbed in the visible region is used effectively no matter what the wavelength (below 680 nm) or the pigment. This last statement has to be qualified, since it has been shown that light absorbed by phycobiliproteins is used with 80-90 per cent efficiency, that by p-carotene with as little as 20 per cent efficiency and that by many xanthophylls (but not fucoxanthin, peridinin or siphonaxanthin) with zero efficiency (Govindjee and Govindjee, 1975; Goedheer, 1979). Thus the spectrum of quantum efficiency may have peaks and troughs depending on the type of alga and its pigments (see e.g. Fig. 25) and these may be more significant than formerly admitted. In Rhodophyta and Cyanophyta a similar problem exists as for the action spectra: that of unequal activation of PSI and PSI1 at certain wavelengths when a single actinic wavelength is used. Wang et al.( 1977) and Myers et al. (1980) have overcome this problem by the use of modulated actinic light with a background light of variable wavelength, adapting a technique first used for Chlorellu sp. by Joliot and Joliot (1968). It should be remembered that quantum yields are measured under very low light fluxes-either at low irradiance or with saturating flashes of light. At higher light fluxesthe same efficiency is not maintained (Section 1X.C) but also changes in the arrangement and interaction of light-harvesting pigments may arise (Section 1X.B). Thus the quantum yield evidence merely suggests that most algae are highly efficient at utilizing irradiant energy at most wavelengths of visible light under low light conditions. Under high irradiances or under variable irradiance, other factors come in to play (Section 1X.B and C); see also Nultsch and Rueffer (1981).

0 02.-

(Cyanobacteria)

400

500

0 02'- (Bacillariophyta)

600

\"

\

700

Wovelength, nm

Fig. 25. Spectra of quantum yield for five algae (a) Chlorellu (Emerson and Arnold, 1942). top left, and Govindjee et a / . (1968) top right; Chroococcus (Emerson and Arnold. 1943); Nitschia (Tanada, 1951). Ihl cruentum cultured under white light (broken line) and green light (solid line). \-I -Pnvnhvridium -.r (Redrawn from Brody and Emerson 1959a ) _I

76


VII. REACTION CENTRE COMPLEXES A. PSI REACTION CENTRE COMPLEXES

I. Introduction Algae of all classes have a Chla-protein supracomplex which is similar in spectral, other physical properties and photoactivity with P-700-Chl a-protein complex (RCI) of higher plants (cf. Brown et al., 1974; Barrett and Anderson, 1980). Identification of RCI complexes in algae have largely rested on absorption and fluorescence spectra of the complexes and their electrophoretic properties in polyacrylamide gels in the presence of SDS or LiDS. Frequently the P-700 activity of the isolated RCI complex has been affected by this technique. Few preparations have been rigorously identified as a RCI complex, by light or chemical-induced oxidation of P-700. Nevertheless, the assumption is that all wild algae and photosynthetically competent mutants contain a PSI reaction centre complex which has the principal characteristics and molecular structural features of the more thoroughly characterized P-700 Chl a-protein complex of higher plants. Algal RCI complexes of particular note, because of the isolation procedure, extent of characterization or other feature are listed in Table IVa. The similarity of the thylakoid organization of green algae, the relative ease of rupture of the cells and isolation of the membranes, has led t o early use of the Chlorophyta (Grimme and Boardman, 1972; Table IVa). The ease of removal of the light-harvesting phycobilisomes of blue-green algae (Section V1II.B) and, latterly, the recognition of the increased stability of membrane-proteins of thermophilic Cyanobacteria has promoted research on the RC complexes of these organisms (cf. Nakayama et al., 1979; Stewart and Bendall, 1979). A much exploited property of both Chlorophyta and Cyanobacteria is their capacity to yield mutants lacking some mechanistic or molecular feature of the photosynthetic systems (Bishop and Senger, 1971). Such mutants have provided much confirmatory evidence about the molecular structure of both RCI and RCII (Table IV). 2. Properties o j ~P-700-Chl a-Protein Complex Overall, measurements of the Chl a/P-700 ratios of the PSI reaction centre

complexes of various algal classes agree with those obtained for higher plants (Thornber et al., 1979). The ratio of Chl a/P-700 is about 40:l provided an innocuous detergent is used (cf. Markwell et al., 1980). A small amount of /?carotene is present in all the algal preparations. Usually, its presence has been inferred by the appearance of a small shoulder in the absorption spectrum at about 500-510 nm; only in a few instances has the /?-carotene been extracted and measured. Thornber (1969) quotes a molar ratio of Chl a//?-carotene of 30:l for a P-700-Chl a-protein complex from a blue-green alga, close to that


77

for several CI preparations from higher plants (Shiozawa et al., 1974). Ogawa et al. (1966) obtained a Chl alp-carotene molar ratio of 6: 1 for Chlorella ellipsoida (Chlorophyta) and certain green plants. These RCI preparations were however not as free of other Chl-proteins as that of Thornber. Barrett and Anderson (1980) extracted and measured the ,!I-carotene of a P-700-Chl a complex from Arrocarpia paniculuta (Phaeophyta). The molar ratio of Chl a to p-carotene was l O : l , and that of [$carotene to P-700 was 4:l. This latter ratio may be significant in relation to the Chl a 684 of the P-700-Chl acomplex, which comprises 20 per cent of the Chl a of this complex. The spectra of Chl a 684 and the j3-carotene show a concomitant blue shift when the P-700Chl a-complex I s treated with long chain neutral or zwitterionic detergents (Barrett, unpublished). This may indicate a close structural relationship between the two pigments in situ. One molecule of ,&carotene to two of Chl a 684 are involved in this blue-shift. The P-700lP-carotene molar ratio may depend on the detergent regime used in the purification of the P-700 Chl a complex. P-700 Chl a-protein coniplexes from Anubaena (Cyanobacteria), Scenedesmus (Chlorophyta) and some green plants which had Chl a/P-700 molar ratios as low as 20-25 had no detectable /?-carotene (Alberte and Thornber, 1978). The mutant Scendesmus C-6E had high PSI activity but lacked any p-carotene (Senger and Strassberger, 1978; Wellburn ef al., 1980). Minor lipids, other than carotenoids, have been detected in P-700-Chl uprotein complexes. Traces of phosphatidyl-ethanolamine, phosphatidic acid and a quinone were detected in the P-700-Chla complex of Phormidium luridium (Cyanobacteria) (Dietrich and Thornber, 1971). Ogawa el al. (1966) also detected quinones in the P-700-Chl a complexes from other algae. As SDS has been used in many modern preparations it is probable that lipid depletion will have occurred with these. However, from digitonin fragmented thylakoids of Nicotiaria tabacum a P-700-Chl a-complex was obtained having phylloquinone, in molar ratio to Chl N of 1: 100 and molar ratio ofp-carotene to lutein of 6 (Interschick-Niebler and Lichtenthaler, 1981). Removal of lipid from the P-700-Chl a complex facilitates removal of a part of the antenna complex by subsequent use of detergents. Alberte and Thornber (1978) obtained a Chl a/P-700 ratio of 20: 1 with preparations from heterocysts of Anaebenu low in carotenoids, and from Scenedesmus mutant 6E, which lacks carotenoids. Carotenoidless mutants of photosynthetic, bacteria. incidentally, have been used to provide purified bacterial RC complexes (Gingras, 1978). Organic solvents have been used to remove antenna Chl a. Sane and Park (1970) using acetone obtained a Chl u/P-700 ratio of 16:I . Extraction of a P-700-Chl a preparation with an initial ratio of 13O:l with wet ether lowered this ratio to 1O:l (Ikegami and Katoh, 1975). Surprisingly the red peak was no further to the red than 673nm. The P700/protein ratios were not given for these P-700 complexes.

TABLE IV(a) PSI Reaction Centre C o r n p k e s Source

Method

Measurements

Authors

Polypeptides ~~~~

Chlamydomonas reinhardrii Wild and mutant strains Chlamydomonas reinhardtii y-1

Chloroplast membranes solubilized in SDS+ dithiothreitol. PAGE SDS

Weight ratios : Chl a :Chl h = 5.3 Protein :Chl=8.8

(1) Membranes disrupted by French press : fractionated by sucrose gradient centrifugation. (2) PAGE Chlamydornonas From chloroplast fragments reinhardrii (Hoober, 1970). wild SDS-urea/PAGE Scenedesmus obliquus Mixed Triton X-100-LDAO- Chl a/P-700 ratio = 16-20 mutant 6E SDS system for solubiliza- A,, 677,438 nm. A 438 tion : hydroxylapatite -= 1.26 chromatography A 677 Acrocarpia Triton X-100 solubilized Chl qF-700; paniculata chloroplasts fractionated by 3811 and sucrose gradient 6011 Ecklonia radiata centrifugation 674.436

Molecular weight 66 KD. Associated with 8-9 Chl a molecules. Absent from mutants lacking P-700 Two polypeptides M, of 63 KD M, of 65 KD. Each having 15 Chl a molecules attached Min. mol. weight of PS I complex = 154 KD. Two polypeptides of M, of 60 KD each derived from PSI polypeptide of 110 KD Polypeptides of M, 50 KD one major and one minor. Also two very small polypeptides

Chua et

a / . ( I 975)

Bar-Nun et a / . (1977)

Anderson and Levine (1974) Alberte and Thornber (1978)

Barrett and Anderson (1980)

Ibid. (1977) (a) Glenodinium sp. 5M29 (b) Gonvuulax polyedra Oscillatoria limosa

Membranes fragmented by French press. Triton extract Chromatographed on hydroxylapatite. SDS-PAGE Cells broken with French

A,, 675.438 nm ratio of Chl a/P-700 (a) 5011 (b) 55/1 A,, 677 nm (77°K)

Prezelin and Alberte (1978)

M, of P-700 complexes:

Thomas and Mousseau (1981)

FI. emission 77°K 726 nm (sh. 680 nm) Difference spectra at 77°K shows presence of 686,695 and 707 nm forms of Chl a Cells extracted with Bound allophycocyanin Digitonin and Triton X-100 Chl a/P-700=35 Mossbauer, EPR and Fluorescence A,, 673 FI. emission SDS-PAGE of phosphate 735 nm (77'K) major, washed membranes 685 minor. Chl a/P-700 ratio 40 : 1 Digitonin fractionation

Oscillatoria splendida Dress : extracted with LiDS p MEL

+

Chlorogloea frirschi

Nostoc sp. Synechococcus cedrorum

Synechococcus sp.

Fractionation of membranes by digitonin. Then chromatography on DEAE-cellulose

Anabaena Jos-aquae

Solubilization with Triton X-100. Sucrose gradient centrifugation and chromatography on DEAE-cellulose

Phormidium luridium Extracted from sonicated var. olivaceae cells by TRIS-HCL pH 8.0 Then chromatography on hydroxylapatite, and precipitation of P700 complex with (NH,),SO,

Chl a/P-700 ratio of 200: 1 A,,,, 25 C. 677 nm A,,,, 77 K 679 nm sh 672 nm Sh. at 686. 710 nm Weak FI emission at 730 nm Chl n'P-700 ratio 36 Chl alprotein w w 0 056

667 nm. 437 nm. 4 9 W 9 5 nm @-carotene) 77'K 676 nm, sh. 710 nm. FI. emission at 690, 77'K Chl a/P-700 ratio, 6 1 3 0 . Ems at pH 8.0= +405 mV

258 K D 181 K D 139 K D 170 subunits recorded Evans er u l . (1979)

Major polypeptide 58 K D lesser of M, 75 K D ; 58 K D

Rusckowski and Zilinskas ( 1980)

Polypeptides of M, 62 K D 18 K D 17 K D 15 K D 14 K D

Newman and Sherman (1978)

Nakayama et nl. (1979)

Klein and Vernon (1977) M, of polypeptides : 1120KD I1 S2 K D 111 46 K D IV S K D Also 2C30 K D derived from 1 Major polypeptide. Dietrich and Thornber (1971) M, 48 K D ; minor, 46 K D .

TABLE IV(b) PSII Reaction Centre Complexes Source Acetabularia mediterranea Chlarnydomonas reinhardtii

Chlarnydomonas reinhardtii

Method Triton X-100 and sucrose gradient centrifugation SDS-PAGE Membranes fragmented by French press. Solubilized in Digitonin and Triton-X100, then sucrose gradient centrifugation. Then chromatography on DEAESephadex A50 Fractionated thylakoid membranes solubilized with LiDS dithiothreitol PAGE Li Dod SOa

Measurements

Polypeptides

Authors

FI. emission at 77"K, peaks Major unit 67 KD consisting Ape1 et al. (1975) at 678 and 694 nm. Enriched of subunits 23 KD and in Chl b, DCIP activity 21.5 KD in a ratio of 2 : 1 Diner and Wollman (1980) Specific activity (AA of M,-45 KD, -50 KD C550/unit Chl a ) 4-7 times Other -polypeptides in __ varying amounts. that of whole algae. Antennae size < 40 Chls. Chl a/Chl b molar ratio, 4.

M, 45-50 KD A,, 674,438 nm ; pronounced sh. 495 nm. Spectrum at 77°K exhibits shoulder at 682,641 nm on red peak. Polypeptides have 4 5 mols Chl a per mole. 1 mole of /3-carotene. Complex accounts for 1% of total Chl. Chlorella fusca SDS-PAGE solubilization CPa migrated between Wild and 2 under mild conditions LHCP and dimer . A, mutants strains 671 nm. CPa accounts for 15% of Chl a Acrocarpia Digitonin extraction of 15.1 KD predominant; A,, 674 and 438 nm well paniculata chloroplasts followed by some 34 KD polypeptide defined peak at 420 nm. Sargassum sp. sucrose gradient Pronounced shoulder at and Scytosiphon sp. centrifugation. Then 496 nm due to /3-carotene. treatment of PSI1 complex FI. emission at 694 nm only. with glycholate and sucrose Extremely low fluorescence. gradient centrifugation.

Delepelaire and Chua (1979)

Wild and Urschel (1980)

Barrett and Thorne (1980) Barrett and Thorne (1981)

Spirulina platensis

Fragmented thylakoids fractionated by sucrose density gradient centrifugation then SDS-PAGE Pellet from French press. LIDS-PAGE give CPA, CPA

A,, 674 nm &carotene present

M, 42 K D

A,,, 6 7 M 7 2 sh. at 5 M 5 Polypeptides of 61 and 45 due to carotenoids. FI. KD. Both absent from N, emission at 685 nm sh. at starved cells and 696 nm. ZHD-grown cells. Membrane passed through A,, 669 nm. Nostoc sp. M, 48 KD. 44 K D French press. Pellet collected FI,,, 685 nm. at 200 OOO g. PAGE-SDS Lysozyme treated cells A,,,, 77 K, 672 nm (sh. 685) Synechococous sp. Highly active in DCIP subjected to French press. Digitonin ertract photoreduction. Intense FI. emission peaks at 685. fractionated on DEAEcellulose 695 nm One cytochrome b,,,/100 Chl a Synechococcus Spectrum at 77-K. A,,, 675, Particles from digitonin cedrorurn 1U 1191 treatment of spheroplasts 437 nm, smaller peak 499 nm centrifugated on sucrose(sh. 465 and 428 nm). gradient followed by DEAE- Nearly symmetrical FI. cellulose chromatography. emission peak at 685 nm. DPIP activity inhibited by DCMU. Synechococcus Miranol S2M-SF solubilized Chl u,cytochrome b,,, Major polypeptides thylakoids fractionated by molar ratio 50:l. Dichloro- molecular weight of 50 K D sp. strain 6301 sucrose gradient phenyldimethyl urea and 48 KB; minor. centrifugation. inhibited light induced a 38 K D and 31 KD. flow from DPC-DCIP Oscillatoria limoso 0 .splendida

Remy and Hoarau (1978)

Thomas and Mousseau (1981)

Rusckowski and Zilinskas ( 1980) Nakayama er ul. (1979)

Newman and Sherman (1978)

Koenig and Vernon (1980, 1981)

TABLE IV(b) --continued

Source Phormidium laminosum

Method Lysozyme-induced. Spheroplasts membranes Solubilized in LDAO. Particles chromatographed on 6B Sepharose.

Measurements A 676.438 nm (sh 465). Carotenoid shoulder 495 nm. F1. emission max at 685-695 nm. Highly active in 0, evolution. Enriched in Mn. 1 Mn/13-17 Chl a molecules. High potential cytochrome b,,,/Chl a 1 :60.C-550 present. P680/antennae Chl a 1 :W 7 0 . In absence of MgCI, sensitive to hypoosmotic media.

Polypeptides

Authors Stewart and Bendall (1979)

Major polypeptides : Ibid. (1981) M, 46.4 KD, 40.1 K D Also a total of 14 other bands ranging from M, of 15.6 to 87.1 KD. Reinman and Thornber (1979) Phormidium luridium Membranes sonicated or put A,, 671,438 nm Phormidium: var. olivaceae and through French press. Then, Carotenoid shoulder 500 nm. Major polypeptides Anabaena cylindrica SDS-PAGE M, 49 KD, 51 KD. Pemmerman Utex 377. Anabaena : 8 polypeptides ranging from M, 15-72 KD. Cyanobacteria : French press, then LDAO Highly active in O2 evolution. England and Evans (1981) various mesophilic High cytochrome b-559/Chl a strains ratio. SDS-solubilized membranes or particles

Fl.=fluorescence. sh.=shoulder.


83

3. Chlorophyll of P-100 Until recently the prevailing view was that P-700 is a dimer of Chl a, coupled to protein in a specific way to generate the spectral and redox properties of P700 and correctly orienting the electron-donating Chl a pair to the primary electron acceptor. The dimeric structures proposed (Shipman et al., 1976; Boxer and Closs, 1976; Junge and Schaffernicht, 1979; cf. Katz et a[., 1979) however would result in a quenching of energy in Chl a dimers, and a lowered efficiency ofenergy transfer (Beddard and Porter, 1976)both inconsistent with the properties of P-700. Furthermore, a reassessment of EPR and ENDOR spectra of P-700 did not support the dimer hypothesis (Davis et al., 1979; Hoff, 1979; Rutherford and Mullet, 1981). The large difference between the Em of Chl a (a+ 860 mV) (Felton, 1978) and P-700 ( +420 mV) was not plausibly explained by these models. The proposal of Wasielewski et al. (1980), that the ring V ester of Chl a is enolized and that the enol form is stabilized by the protein of P-700-protein, could explain the drop in redox potential, but does not account for the 40 nm difference between the absorption maxima of Chl a and P-700 in the red region. Co-ordination of histidine to the fifth and sixth position of the Mg of Chl a could markedly lower the Emof Chl a, depending on the degree of polarity of the protein environment (Kassner, 1973; Xavier et al., 1978; cf. Stellwagen, 1978). However the complex of Chl a with pyridine-a stronger n electron donor than histidine-has its A,, at 440 and 670 nm only (McCartin 1963). Excited states of Chla enol are discussed by Petke et al. (1981). The recent isolation of Chl RCI from Scenrdesmus, wild type and mutants, and from spinach chloroplasts and P-700-Chl a-complex preparations (Dornemann and Senger, 1981, 1982), raises the possibility that the chemical and physical properties of P-700 are attributable to a Chl different from Chl a. Chl RCI has a lower redox potential than Chl a, estimated by I, titration, and the fluorescence yield of Chl RCI is one third that of Chl a. Chl RCI has its red absorption maxima at about 10 nm to the red compared to Chl a. Structural differences between Chla and Chl RCI are not yet determined. The red spectral shift could arise from the presence of a second vinyl group in Chl RCI, or by the replacement of the vinyl at /?-position 2 of Chla (Fig. 19) by an oxygen-containing electron-withdrawing group. Chl RCI has an increase in mass of 36 over Chl a, and this in part may be due to an extension of the alkyl sidechains, as occurs in the Chlorobium chlorophylls (Holt, 1966). The hydrophobic regions of the protein of the P-700 complex may have a part in determining the spectral and redox properties of P-700. Hoarau and Remy (1978) observed that Triton X-100 caused the long wavelength forms of Chla (684682nm) in P-700 complexes and LHCP from Nicotiana, Chlorella and Porphyridium (Rhodophyta) to assume configurations with A,,, at 662 nm. Disturbance of the hydrophobic environment by the milder long chain alkyl-detergents, LDAO, Zwittergen and Tweens, caused similar

84

A. W. D . LARKUM AND JACK BARRETT

spectral effects in P-700 complexes and PSII core reaction centre complexes of brown algae (Barrett, in preparation). It is significant that a Chl a-protein with an absorption maximum of661 nm (Murata, 1981; Barrett and Thorne, 1980) has similar spectra to those of Chl a in polar solvents, and that a hydrophobic environment can induce larger spectral changes in Chl a than is obtained by co-ordinating pyridine to Chl a. Chunaev et al. (1980) observed that in mutants of Chlamydomonas reinhardtii which are low in carotenoid the dominant Chl a spectral form was Chl a 681, but in those mutants which accumulate carotenoids the longer wavelength forms of Chl a predominate. 4. Antenna Chls of’ The P-700-Chl a Complexes The P-700-Chl a protein complex accounts for 15-30 per cent of the Chl a of green algae and higher plants (Shiozawa et al., 1974; Brown et al., 1974; Anderson et al., 1980); and about 20 per cent in brown algae (Barrett and Anderson, 1977, 1980) and 10 per cent in the dinoflagellates, Glenodinium sp. and Gonyaulax polyedra (Prezelin and Alberte, 1978). In algae which have phycobiliproteins the relative amount of Chl a associated with PSI is higher. Hiller and Goodchild (1981) estimate that in Grifiithisia monilis (Rhodophyta) 50 per cent of the Chl a is in PSI. Mimuro and Fujita (1977) estimated the distribution of Chl a between PSI and PSII by delayed light emission for intact cells, and by DCIP reduction and cytochrome c photooxidation. The first method indicated that 19 per cent of the total Chl a was associated with PSII and the second method, 12 per cent. In an extensive study with Anacystis nidulans (Cyanobacteria) and several mutants, 70 per cent of the Chl a was found to be associated with the PSI complex when the algae were grown in red light, and 80 per cent when they were grown in green light (Wang et al., 1977; Myers et al., 1980; Section IX and X). There is uncertainty as to the number of Chl a molecules which contribute to each of the spectral species Chl683, Chl67 1 and Chl661, resolved by curve deconvolution of the P-700-Chl a complex (cf. Brown, 1977a). It is not known whether these species arise from the interaction of Chl a molecules with different polypeptides or from coupling interaction between close Chl a molecules (Shipman, 1980). The only Chl-protein which permits some insight into this problem is the water soluble BChl a-protein from Prosthecochloris aestuarii (Section 1X.B). X-ray crystallographic analysis at 2.8 A of this BChl a-protein shows that seven BChl a molecules are arranged in a protein chain of 35 K D molecular weight (Fig. 42). Six of the BChl a molecules have an amino-acid liganding to the fifth co-ordination position of the central Mg atom: two or three histidines are ligands (Fenna and Matthews, 1979). This BChl a-protein has a simple spectrum (Olson, 1980), whereas the Chl a-P-700complex has a compound spectrum. Comparison with the cytochromes and haemoglobins (Lemberg and Barrett, 1973; Antonini and Brunori, 1971) and the phycobiliproteins (Gantt,


85

1980; Glazer, 1981) suggests that there is unlikely to be more than one molecule of Chl a for each polypeptide unit mass of 5 KD. This is more so as the protein moiety has to accommodate the phytyl chain of the Chls. Refined SDS-PAGE procedures have provided evidence of P-700 complexes which have Chl a/P-700 ratio of 120 in higher plants (Anderson er al., 1978; Mullet et al., 1980a,b). Similar complexes from the marine alga Caulerpa cactoides had a Chl a/P-700 ratio of 1 10 (Anderson et al., 1980).The red alga Grjfithisia monilis (Hiller and Larkum, 1981) has also yielded a similar P-700-Chl a-complex. Chromatography has been used to purify the native complex isolated from higher plants (Mullet et al., 1980a,b). The fluorescence emission of the latter preparations is at 735 nm rather than at 685 nm as for the core P-700-Chl a-complexes. I t is not known whether the extra 70 to 80 Chls a comprise one spectral species or give rise to each of the major spectral species, Chl 66 1, Chl 67 1, Chl 682. The Chl a/b ratios of these larger P-700-Chl a complexes in higher plants are 10. Haworth et al. (1981) find that most of the Chl band half of the Chl a is associated with polypeptides of molecular weight (MW) 19-24 KD. These P700-Chl a complexes (Chi u/P-700= 110) show in their absorption and fluorescence spectra evidence for the presence of Chl b (Haworth et al., 1981; Interschick-Niebler and Lichtenthaler, 1981 ; Ryrie, personal communication), and their emission peak is at 748 nm.

-

-

5. Molecular Weights of RCI Complexes and of' the

Component Polypeptides Estimates of the molecular weight of core P-700 Chl a-complex range from 140-260 K D for higher plants and blue-green algae (cf. Table IV). Some of the differences in MW may be attributable to methodological errors due to anomalous binding of SDS (cf. Grefarth and Reynold, 1974; Chua et al., 1975). In any case the isolated complex may be a sub-aggregate of the in vivo form and different oligomers can be obtained. Three P-700-Chl a-complexes isolated by LiDS-PAGE from two cyanobacteria, Oscillatoria limosa and Oscillatoria splendida (Thomas and Mousseau, 1981) had apparent MW of 258, 181 and 139 KD. The P-700 complexes from a wide range of macrobenthic brown algae have a large MW, but may exhibit disaggregation in the presence of steroid detergents (Barrett, unpublished). Most preparations of the P-700-Chl a-complex have three polypeptides detectable by Coomassie blue (Table IV(a)). Preparations of the P-700-Chl acomplex which have Chl a/P-700 ratios of 120 or greater (native complexes) have extra polypeptides, arising from the peripheral antennae Chl a and colourless peptides. In an intensive investigation using a Chl b-less barley mutant, developing cucumber cotyledons, and mature normal strains of barley, Mullet ef al. (1980b) have shown that the peripheral Chl a is associated with polypeptides of MW 21.5 and 24.5 KD, not present in the core P-700-

86


Chla-complex (ChlalP-700 of 40). The core complex had two major polypeptides of 66 and 68 KD; the number of minor polypeptide bands on SDS-PAGE increased with the increase of the Chl a/P-700 ratio (Mullet et al., 1980a). There is no decisive evidence on how the molecules of Chl a are distributed between the polypeptides of the P-700-Chl a-complex (P-700/Chl a = 40), or whether all the polypeptides carry Chl a. More refined methods of separation of the polypeptides are likely to increase the number detectable. For example, the use of chloral hydrate as a dissociating agent has led to the detection of 15 polypeptides in cytochrome oxidase instead of the customary 7-9 (Griffin and Landon, 1981). Lagoutte et al. (1980) have used a limited proteolysis technique, coupled with SDS-PAGE, for characterizing the polypeptides in the 60 KD region: these authors claim that CFl is co-purified with the P-700Chl a-complex by most methods used to prepare the latter from spinach. Estimation of the molar ratio of Chl a to protein is vitiated by dissociation of Chla from the polypeptides by SDS or Triton X-100, especially during PAGE. Nevertheless, the current use of milder electrophoretic conditions has improved the credibility of recent estimates of the number of Chls associated with polypeptides. Chua er al. (1975) found that the 66 KD polypeptide of the P-700-Chl acomplex of a wild strain of C. reinhardrii carried 8-9 Chls a; this polypeptide is absent from P-700-less mutants. Bar-Nun et al. (1977) found that 15 Chlsa were associated with each of two polypeptides of 63 KD and 65 KD respectively obtained from the P-700-Chl a-complex of C. reinhardtii y-1. The MW of these two polypeptides are close to those for two polypeptides (-60 K) from a 1 10 KD subunit of this complex from a wild strain of C. reinhardtii (Anderson and Levine, 1974). In particularly pertinent studies, Setif et al. (1 980) and Lagoutte et al. (I98 1) isolated P-700 complex from spinach in high yield. Only polypeptides of MW 65 and 62 KD were detected, apart from an iron-sulphurpolypeptide with MW of I 1 KD. One mole of P-700 and 40 moles of Chl a were coupled to about 140 x lo3 g of protein. This would suggest that either the 65 or 62 KD polypeptide carried the P-700 Chl molecule and that the antenna Chl was shared between both of the polypeptides. The P-700-complex polypeptides of Cyanobacteria and Phaeophyta have lower molecular weights than are found for Chlorophyta and higher plants (Table IV(a)), so that it can be expected that more molecules of these polypeptides will be present, each carrying fewer Chls. 6. The Primary Electron Acceptor and Donor of RCI The P-700-Chl a complexes undergo reversible photo-oxidation, indicating that the primary electron acceptor is present. The complexes have a very low fluorescence yield because of the quenching of the excitation energy by the


87

electron acceptor. Evidence of the nature of the acceptor comes from EPR studies on Cyanobacteria, green algae and higher plants (Malkin and Bearden, 1971; Cammack and Evans, 1975; Evans et a/., 1976; Malkin and Bearden, 1975, 1978) and from Mossbauer spectroscopy (Evans et al., 1977, 1979). PSI has two iron-sulphur centres, A and B. Centre A has a slightly less positive redox midpoint potential and is reduced first. For barley and spinach, the Emof centre A is - 550 mV, and of centre B it is - 594 mV for barley and -585 mV for spinach. The variation in the Em of centre B with species is considerable (Evans et a/., 1974; Cammack and Evans, 1975). The PSI iron-sulphur centres in the halophile, Dunuliella parva (Chlorophyta) exhibit unusual temperature dependence (Hootkins et al., 1981). EPR studies of membranes from Phormidium laminosum (Cyanobacteria) reveal that there is spin-spin interaction between the Fe of the two centres, complicating the redox situation (Cammack et al., 1979). Nugent et al. (1980a,b, 1981), from EPR studies with Scenedesmus obliquus (Chlorophyta), Phormidium laminosum and higher plants, propose that the electron flow from P-700 passes through an acceptor prior to Centres A and B, which can act in parallel, and then to ferredoxin (Em- -420mV). The presence of substantial amounts of iron in the P-700-complex of the chlorophyte Chlorogloea jritschii (Chl alp-700 = 35: 1) has been shown by Mossbauer spectroscopy (Evans et al., 1979, 1981); some of the iron is in an environment similar to that of the 4Fe-4S centres of ferredoxin. Recent EPRflash photolysis studies have placed the Em of membrane-bound acceptor higher, at - 540 mV and - 590 mV, while the Emof the intermediate acceptors was more negative than - 700 mV (Mclntosh et d., 1981).The Emof P-700 in Chforella thylakoids is +450mV, measured by EPR, while for P-700 in Anacystis lamellae it is + 430 mV: in purified P-700 complexes the Em was 2&30 mV lower than those for lamellar preparations (Hoarau et a / . , 1981). These values are higher than that for spinach (Em+ 375 mV), determined also by EPR (Nugent et al., 1980a,b, 1981), but redox potentials are sensitive to structural changes induced in the complexes during purification procedures. A prime factor in the regulation of the reduction of P-700 is the electrostatic interaction between electron-transfer components, dependent on net charges of the individual components and the surface of the membrane (Wood and Bendall, 1975; Davis et al., 1980; Tamura et al., 1980, 1981). The study on Chl a sensitized reduction in non-ionic micelles is pertinent (Kalyanasundarum and Porter, 1978), as are those of Ilani and Mauzerall (1981) and Krakover et al., (1981). For many algae plastocyanin, with similar properties to that in higher plants, is the primary stable electron donor to P-700. For example, the crystallized plastocyanin of Enteromorphaprol[fera(Chlorophyta) has a M W of 12 KD, pl of 4.1 and an Em of +0.37 mV (Fuminori et al., 1981). The amino acid composition is close to that of the plastocyanin of higher plants.

88


Many of these alga possessing plastocyanin have an alternative primary electron donor cytochrome c-552 or 553. The molecular size, solubility and redox properties of this cytochrome c are similar to those of plastocyanin, and there is little difficulty in this cytochrome substituting for plastocyanin, particularly in cultures of Scenedesmus (Bohner and Boger, 1978) and other copper-deficient algae (Wood, 1978; Bohner et al., 1980). Cytochrome c 552-3 may replace plastocyanin entirely in certain algae, e.g. Euglena gracilis (Euglenophyta) (Wildner and Hauska, 1974) Bumilleriopsis Jiliformis (Xanthophyta) (Kunert and Boger, 1975; Kunert et al., 1976) and Chlamydomonas mundana (Chlorophyta) (Wood, 1978; Sandman and Boger, 1980). The Chromophyta, which contains many algae possessing cytochrome c 552-3, merit a wider survey for the occurrence of plastocyanin. B. PSII REACTION CENTRE COMPLEXES

1 . General Properties PSII reaction centre (RCII) complexes can broadly be divided into two classes. The first contains the macromolecular systems which have retained the capacity for photolysis of water (oxygenic complexes). The second class is that comprising the core RCII complex which has P-680, with its associated antenna pigments Chla and &carotene, and can only mediate transfer of electrons from a donor to a suitable acceptor in the presence of light. Several workers in the past decade have obtained oxygenic preparations from algae (Table IV) and the manganese involved in water-splitting has received special attention (cf. Mauzerell and Piccioni, 1981). It is only in the last few years that it has been possible to isolate and identify the RCII core complex. The rather harsh conditions for solubilization of the thylakoids, using either Triton X-100 or SDS, and hydroxylapatite chromatography or PAGE employed by the earlier workers were not conducive to the retention of the Chl a by the somewhat labile RCII core complex. With the general use of milder conditions or PAGE, where SDS is frequently replaced by LiDS, octylglucoside (Camm and Green, 1980; Green and Camm, 1981), digitonin (Wessels et a[., 1973; Satoh and Butler, 1978a) or other steroids (Barrett and Thorne, 1981), a Chl a-protein complex has consistently been obtained from many types of algae, as well as green plants. The Chl a-protein ascribable to the RCII complex has been designated CPa by Hayden and Hopkins (1977) and since this designation has been generally used by workers with higher plants (Anderson et al., 1978), this term will be used for the algal RCII core complex here as its properties are clearly similar to the CPa of higher plants. CPa in SDS-PAGE is more mobile than CPI a, CPI, LHCP, and LHCP2, but less so than LHCP3 (Anderson et al., 1978; Henriques and Park, 1978; Wessels and Borchert, 1978) (LHCPs are Chl a/bproteins: see Section VII1.A). CPa has been isolated from the green


89

siphonaceous alga Caulerpa cactoides and Codiumfragile where it comprised 6 per cent of the total Chl distribution (Anderson et al., 1981). Both CPa's had low fluorescence yields, as expected of a reaction centre complex. A CPa has been isolated from Acetabularia, octylglucoside being the preferred detergent (Green et a[., 1982). A potentially valuable source of CPa is Chlorella fusca, where in its mutant S-36, the CPa accounts for 15 per cent of the total Chl a in the thylakoids (Wild and Urschel, 1980). Delepaire and Chua (1979) used LiDS both to solubilize the thylakoid membranes of a wild strain of Chlamydomonas reinhardtii, and in the subsequent PAGE separation of the Chl-complexes where two RCII complexes were obtained. Unlike some other CPa preparations obtained by PAGE, these CPa had a shoulder at 682 nm and a minor peak at 641 nm. A pronounced shoulder at 495 nm was confirmed to be due to p-carotene. C . reinhardtii mutants with impaired PSI1 function have been studied by PAGE (Marco and Gamier, 1981). Several workers have demonstrated the presence of a CPa in SDS or LiDS solubilized thylakoids of various Cyanobacteria and it has been isolated from a rhodophyte Griffithsiu monilis (Hiller and Larkum, 1981). Spectral characteristics and in certain cases the principal polypeptides of these preparations are given in Table IVb. Such core RCII complexes have not been assayed for DCIP photoreduction activity . Preparations which used sucrose density gradient centrifugation rather than PAGE for isolation ot KCII complexes include, amongst the green algae, that from Acetabularia mediterranea (Ape1 et a / . , 1975) and a Chlamydomonas reinhardtii mutant (Diner and Wollman, 1980), Table IV. The specific activity (AA of C550/unit Chl a ) of the latter preparation was 4-7 times that of the whole alga. Considerable and growing attention has been given to the Cyanobacteria as a source of RCII complexes. This is mainly because of their high oxygenic capacity, but also because thermophilic strains are available which provide more stable preparations, and this feature has been exploited by Katoh (Nakayama et al., 1979) and Stewart and Bendall (1979, 1980, 1981). Highly oxygenic preparations have been obtained also from mesophilic cyanobacteria (England and Evans, 1981) comparable in activity to preparations from spinach (Berthold et al., 1981). The preparation of Stewart and Bendall (1981) was enriched in Mn (1 Mn/13 Chla). The removal of this Mn by mild heating, Tris or EDTA treatment caused complete loss of oxygen evolution (Stewart and Bendall, 1980, 1981). This oxygenic preparation was also enriched 5-6 fold in cytochrome h-559Hp and cytochrome c-549, but to a lesser extent in cytochromes b-559Lp, b-563 and f . An essential factor in preserving the oxygenic-activity uf the particles from Phormidium laminosum, was correct osmolarity and the inclusion of glycerol or sucrose in the preparation media. Maintenance of hydrophobic interactions of the complex in the presence of

90


detergent could have contributed to the stability. Measurement of the P-680 to antenna Chla by Stewart and Bendall (1981) gave a ratio of 40-70, comparable to the value obtained for spinach RCII complex by Satoh (1979a,b). Other particulate preparations which had the capacity to photoreduce DCIP but were not oxygenic have been obtained from various Cyanobacteria. The preparations from Synechococcus of Newman and Sherman (1978) used digitonin as the solubilizing agent, as did of Nakayama et al. (1979). Koenig and Vernon (1981) used Miranol to isolate RCII from Anacystis nidulans, and Stewart and Bendall (1980, 1981) used lauryl-dimethylamine oxide to obtain the oxygenic particles from P. larninosurn. All these preparations contained cytochrome b 559. A thylakoid membrane preparation from a thermophilic cyanobacterium had oxygen evolving activity (Yamaoka et al., 1978), but this activity was destroyed by digitonin. Protoplasts of Anacystis nidulans (Indiana str. 625) exhibited electron transfer from water to DPIP, but this activity was lost on disrupting the protoplasts; however, electron transfer activity from hydroxylamine to DPIP was retained (Sigalat and de Kouchkovsky, 1974). Core RCII complexes have also been obtained from the brown seaweeds Acrocarpia paniculata, Phyllospora cornosa and Padina commersonii (Table IV), Fig. 35. These RCII complexes are distinctive in that their long-wave fluorescence emission at 77°K has a single peak, at 694 nm, with no emission peak at 684 nm (Barrett and Thorne, 1980). A distinctive component of the RCII complexes is p-carotene, and absorption spectra of the RCII algal complexes show that it is present in at least twice the amount as in the RCI complexes. Moreover the absorption maximum in the RCII complexes is at 495 nm, rather than the 510 nm of the RCI complexes. Measurement of the P-carotene by chromatography followed by spectroscopy indicate that there is 1 mole of P-carotene to 4 5 Chla molecules (Delepaire and Chua, 1979; Barrett, unpublished). This carotenoid has a role in establishing the longwave forms of the Chl a of the RCII complex. In Scenedesmus obliquus (Chlorophyta) xanthophylls, mainly lutein, have been implicated in the structural organization of RCII (Senger and Strassberger, 1978). The influence of cations on the structure and photochemistry of the RCII complex has been studied (Lazlo and Gross, 1980). The integration of the RCII complex into the photosynthetic apparatus of S . obliquus has been explored by Brinkman and Senger (1981) on the basis of the isolation and assignment of function to six thylakoid Chl-protein complexes. 2. Polypeptides OJ RCII Complexes In common with the RCII complexes of higher plants (Satoh, 1979a,b; Wessels and Borchert, 1978; Koenig et al., 1977) many preparations have two polypeptides of MW of between 44-51 K D (cf. Table IV). With algae lacking


91

PSII, e.g. N,-starved Oscillatoria sp. (Cyanobacteria) (Thomas and Mousseau, 1981), Chlamydomonas reinhardfii mutants (Chua and Bennoun, 1975), or a PSII dificient mutant of barley (Machold and Hoyer-Hansen, 1976), the polypeptides of -40 K D are obscure on PAGE of the thylakoid preparations. On the other hand, the CPa obtained by PAGE from lettuce thylakoids had a major polypeptide of about 40 K D (Henriques and Park, 1978). It is probable that the polypeptides of MW-50 K D may consist of smaller subunits. Indeed the oxygenic complex from P . laminosum contains 16 polypeptides in the MW range 15.6-87.1 K D (Stewart and Bendall, 1981), while the cyanobacterial PSII preparation of Reinman and Thornber (1979) gave 8 polypeptides in the MW range 15-72 KD. The RCII-core complex from Acrocarpia paniculafa (Phaeophyta)contains two peptides of 27 and 15 KD, respectively. As peptides from thylakoids of brown algae stain very weakly with the conventional stains, it is not excluded that other polypeptides are present. Improvements in the resolving power of the analytical PAGE system and dissociation of the polypeptide complexes e.g. by the use of hydrophobic-bond breaking reagents, has resulted in the detection of additional polypeptides, some of low M, in the RCIl complex of spinach (Satoh, 1981) and that of C . reinhardtii (Bennoun ef al., 1981). 3. Primary Electron Donor and Electron Acceptors (a) Electron donor. The Chl a species P-680 can be regarded as the primary electron donor (D) in PSII since photooxidation of this photoreactive Chl generates the Chl a cation, P-680+, with ejection of an electron (cf. Amesz and Duysens, 1977). The redox potential of P-680 is well above + 810 mV. On the water side of the electron flow the electron donor, D,, to P-680 has not been identified chemically, but it is assumed to have a high Emand the t/, of reduction is 30ns (van Best and Mathis, 1978). The rate of transfer of electrons from D , to P-680 is very sensitive to pH and temperature in P . laminosum and in spinach chloroplasts (Reinman et al., 1981; Conjeoud and Mathis, 1980), consequently this stage in the electron flow might be a regulator of photosynthesis via the proton flux in the membrane. P-680 can be reduced in a back reaction from Q1 (Bouges-Bocquet, 1980), see below. Q1 ijself is subject to reverse electron flow, which is associated with a fluorescent transient dependent on a midpoint potential of + 345 mV, at pH 6.9 (Hardt, 1981). Properties of the electron donors associated with Q, and Qs have been described (Melis and Homann, 1976; Thielen and Van Gorkom, 1981a,b). (b) Primary stable acceptor. As P-680, but not P-680 +,is a quencher of fluorescence the reduction of the primary stable acceptor, Q1, can be monitored by measuring the fluorescence yield of the antenna Chl a of PSII. The reduction of Q, (and Q,, a secondary acceptor), may be inhibited by DCMU which combines with a specific polypeptide. In Spirodelu oligorrhiza (Mattoo et al., 1981) this polypeptide has a M, of 33 KD; in the cyanobac+

+

+

+

92


terium Aphanocapsa str 6714 it has an M, of 33 K D (Astier and JosetEspardellier, 1981), while the DCMU-reactive polypeptide in pea chloroplasts has a M, of 32-34 KD (Mullet and Arntzen, 1981). Renger et af.(1981) have identified a similar protein which they suggest acts as a regulator of the electron flow between P-680 and Q l . Q l is a special bound form of plastoquinone (van Gorkom, 1974) which Witt (1973) proposed to be complexed with a transition metal. Klimov et a f . (1980) have observed a photoinducible ESR doublet signal, plastoquinone A (Q ,) and Fe dependent, centred at g=2.00 with a splitting of 52 g at 77°K in addition to a narrow signal at g = 2.0033. The absence of a signal for the radical anion PQ - in the ESR spectrum had previously been interpreted as indicating that PQ interacts with a paramagnetic species (cf. Mathis and Paillotin, 1981). Another electron acceptor, Q2, has been detected by Joliot and Joliot (1981). An absorbance change occurs at 550 nm (the C-550 shift, Knaff and Arnon, 1969) during reduction of the plastoquinone. It is thought that this (2-550 shift is due to p-carotene molecules sensitive to the redox state of the plastoquinone, since restoration of plastoquinone a alone to chloroplasts which had been fully depleted of lipid restored the primary photochemistry at 77°K but not the C-550 shift. The reduction of P-680+ is a heterogeneous process, with a fast phase followed by a slow phase (Joliot and Joliot, 1979) and consisting of several redox levels, each separated by a few hundred millivolts (Melis, 1978; Horton, 1981; Bouges-Bocquet, 1980, for review). The fluorescence-yield redox titration of the redox properties of Q1 (Cramer and Butler, 1969) and the absorbance changes at 5 18 nm, the electrochromic band associated with the primary charge separation across the thylakoid membrane, both indicate the existence of two primary acceptors in PSII. The redox potentials of these two centres differ by about 250-300 mV (Melis, 1978). Thielen and van Gorkom (198 1) and Horton and Croze (1979) extended the study of these centres, using fluorescence-redox titration. In the presence of DCMU, Q, is reduced at - 300 mV, but in its absence Q, is reduced at + 30 mV. The second centre Q, is reduced at +115mV. Two types of antenna systems have now been demonstrated in tobacco chloroplasts: antenna-Q, consisted mainly of Chl 683, while antenna-Qa had absorption maxima at 650 and 672 nm (Thielen et af.,1981). Q, and Qa cannot be correlated biochemically with Q l and Q2, the RCII electron acceptors described by Joliot and Joliot (1981). Properties of the electron donors associated with Q, and Qa have been described (Melis and Homann, 1976; Thielen and von Gorkom, 1981). Q, and Qp are assigned respectively to PSII, and PSII, by Thielen and Gorkom (1981b), which function with high quantum efficiency, as does PSI. PSII, and PSII, are thought to be different and independent structures, the former located in stroma membranes and the latter in appressed thylakoid +


93

'membranes. Biophysical evidence supports these assignments (Anderson and Melis, personal communication). (c) Intermediate acceptor I . Although Q I , the special plastoquinone (or its Fe-complex) with an E, of 0 to -200mV has been designated the RCII primary acceptor, substantial evidence points to there being an intermediate acceptor (I) between P-680 and Q I (van Best and Duysens, 1977). EPR of RCII preparations and redox titrations of these indicate an E, of 610 mV for I (Klimov et al., 1979; Rutherford et al., 1981a). Interpretation of the physicochemical evidence for the identify of I has relied to an extent on analogies with the photochemistry of bacterial RC, which however correspond more closely to RCI centres than to RCII centres. These bacterial RC contain 4 molecules of BChla and 2 of Bpheophytin. Resonance Raman spectroscopy shows that the pheophytins are each bound differently to protein (Lutz, 1980). Fluorescence studies in the pico-second range provide evidence that Bpheophytin is an intermediate electron acceptor (cf. Codgell, 1979), but Swarthoff et al. (1981b) deduce from similar fluorescence studies that in green photosynthetic bacteria the intermediate acceptor is BChl. Evidence that pheophytin maybe the intermediate acceptor, I, in PSII comes largely from Klimov and his associates and is based on optical studies of the reversible photo-reduction of I (Klevanik et al., 1977; Klimov et al., 1977). Negative difference A,,, were observed at 422, 518, 545 and 685 nm, with concomitant increases in absorption, principally at 450 nm and 660 nm, these two A,,, were related to the radical anion of pheophytin (Ph'-). The differences of about 15nm between the position of the four A,,, of the difference spectra and the A,,, of pheophytin in ether were ascribed to association with other molecules or aggregation. A comparison of the spectra of Pha and Pha'- and Chl a and Chl a - in dimethylformamide with that of reduced PSII reaction centre complex led Fujita et al. (1978) to concur with this interpretation. However, the spectra are not completely convincing. Other evidence for the nature of I comes from the detection of a triple state in photoexcited PSII particles (Rutherford er al., 1981a) and from the magnetic field induced increase of fluorescence emission at 685 or 695 nm in chloroplasts of Chlorella vulgaris and spinach (Rademaker et al., 1979). Evstigneev and Gavrilova (1979) ,have demonstrated a photochemical intereaction of Chl a and phaeophytin a mediated by quinones. The identification of I as pheophytin a by isolation is difficult. RCII complexes used in these studies have molar ratios of Chl a/P-680 of from 40 (Shuvalov et al., 1979) to 120 (Rutherford et al., 1981a) unlike bacterial centres which have only 4 molecules of BChl and 2 of the putative pheophytin. Loss of Mg from Chl a may be difficult to avoid during preparation of RCII core complexes. If I in the RCII complex is pheophytin a, two questions are posed: what is the mechanism whereby there is selective demetallation of a few +

94


Chl a molecules, and how are the ubiquitous Zn or Cu excluded from chelation with the pheophytin in vivo? Another view of the identification of pheophytin with 1 is given by Vermaas and Govindjee (1981). The redox bridge between P-680 and Q , may have an earlier acceptor than I. Rutherford (1981) has detected by EPR a light-induced radical signal when preparations are held at an Em of -645 mV, at which potential I is fully reduced. Swarthoff et a f . (1981a,b) have likewise detected a spin polarized triplet under relative redox conditions in reaction-centre preparations from green photosynthetic bacteria. +

C. SIZE OF ANTENNA OF REACTION CENTRES

Although Cryptophyta and Rhodophyta have phycobiliproteins as their major light-harvesting pigment complexes, they also have antenna Chlproteins which capture and funnel light to the RC. The absence of intrinsic light-harvesting proteins in these algae confers an advantage for studying RCantenna relationships. In the Cryptophyta and Rhodophyta most of this antenna Chl a contributes to RCI, as in Porphyridium cruentum Rhodophyta (Ley and Butler, 1976). For Anabaena variabifis(Cyanobacteria) Mimuro and Fujita (1977) estimated a ratio of 140 Chl a for each P-700, but only 20 Chl a for each P-680. The smallness of the core antenna of PSI1 units in Cyanobacteria and Rhodophyta has been demonstrated by a number of workers (Diner and Mauzerall, 1973; Diner and Wollman, 1977a; Ke and Dolan, 1980; Myers et al., 1980; Vierling and Alberte, 1980) and this topic is discussed in Section 1X.D. In other algae also the PSII unit size may be small (Falkowski et a/., 1981) presumably because light-harvesting complexes pass on excitation energy to PSII (Section V1.D). Diner and Wollman (1980) isolated highly active PSII particles from a mutant of the green alga, Chfamydomonas reinhardtii lacking a P-700 Chl a complex: the antenna size of 4&50 Chl’s contained little Chl @-protein. Antenna size in algae can vary with light-shade adaptation (Section 1X.D). Grown under high light Skeletonema costatum (Bacillariophyta) had a Chl a/P-700 ratio of 650, but low-light increased the ratio of this algae to 1100 (Falkowski and Owens, 1980). Similar results were obtained with other diatoms. In contrast, the Chl a-b/P-700 ratio (-470) varied little in Dunafiella tertiofecta (Chlorophyta) in response to light-shade adaptation, instead the number of P-700 units was increased (Falkowski and Owens, 1980). Falkowski et a f .(1981) found with D . tertiolecta that the increase in Chl a content in cells grown under low levels of light parallels the increase in the number of both RCl and RCII, so that effectively there was no increase of antenna size with S. costatum: the Chl a/RCIl ratios stayed constant over the range of irradiation used, but in contrast, the levels of P-700 decreaecd as the


95

Chl a increased in response to lower levels of light. The RC1:RCII ratio of S. costaturn decreased from 1 in high light to 0.4 in low light growth. Intermittent light during growth also can affect the RC/Chl ratio. The antenna optical cross-section of Euglenu grarilis was reduced by a factor of about 4 in intermittent light cultures. (Dubertret and Lefort-Tran, 1981). The RC antenna size does not vary always with changes in the light regime. Fleischhacker and Senger (1978) found that in synchronous culture of Scenedesmus obliquus the Chl/P-700 ratio of 500 was constant under high and low light growth, and that the decrease of photosynthetic capacity found under a low light regime was due to a lowering of the molar ratio of cytochrome f/P-700 (see Section 1X.D). D. OPTICAL SPECTRAL ANALYSIS OF CHLOROPHYLL PROTEINS

I . Introduction Chlorophylls and carotenoids in their complexes with proteins exhibit multiple spectral forms which differ from those observed for the individual pigments in organic solvents. Moreover each chromophore, in a particular Chl a protein, may generate more than one spectral species. The composite absorption spectra are encountered both in the intact chloroplast, isolated reaction centre complexes and light-harvesting complexes. Further, there may be extensive interactions between both homologous and heterogenous chromophores. Knowledge of the number and types of pigment-protein species is of crucial importance in defining the sequence of electronic events that comprise the primary photochemistry of photosynthesis (Section IX). Discussions of the theoretical molecular origin of the spectra of Chls and porphyrins have been presented by Gouterman et al. ( I 963), Weiss ( 1972, 1979). Reviews on spectra of Chls are those of Goedheer (1966) and Gurinovich et al. (1 968); Seely (1977) provides a concise summary of the topic. Important studies on solvent effects in the spectroscopy of Chls have been made by McCartin (1963) and Seely and Jensen (1965). The analysis of the composite light absorbance spectrum of the pigments is not easy, and conclusions are sometimes debatable. Four methods are principally used, and though these may provide approximate rather than precise definitions of the component spectral species, they provide insight into the complexity of the molecular basis of the photochemical processes within the chloroplast. There are three physical methods; difference absorption spectroscopy, fourth derivative spectroscopy and fluorescence excitation spectroscopy. The fourth method is the mathematical analysis of the absorption or flourescence excitation curves. Analysis of composite absorbance spectra of Chl-protein complexes has concentrated on the absorption at the red end of the spectrum since the absorption maxima of Chls a, b and c are well separated, and because of the

96


absence of the obscuring effects of carotenoids. The carotenoids contribute in all Chl-protein complexes to the absorption spectra in the near-UV region. None the less, valuable information about the electronic state of the components Chls (Weiss, 1972),and thus their molecular environment, which could be obtained by resolution of these near UV spectra warrants determined efforts to apply to this problem the resources of computer analysis. 2. Curve Deconvolution Analysis The defining of composite absorption spectra or fluorescence excitation and emission curves as being composed of the sum of hypothetical gaussian, or lorentz-gaussian components, is termed curve deconvolution (French et al., 1968; Katz et al., 1977) (Fig. 26). The mathematical components discerned do not necessarily correspond to the actual spectrum of any individual spectral species generated by a Chl-protein. There is no reason apriori to assume that such spectral species, arising from a specific environment, must possess a symmetrical absorption peak; also harmonics from another spectral species may cause distortion. Consequently the number of gaussian symmetrical curves adduced to define the absorption peak may exceed the number of real spectral species. Nevertheless, curve deconvolution provides a means of identifying spectral components common to different Chl-protein complexes. The far-red absorption peak (A,,, 674-6 nm) of P-700-Chl a-protein complexes of tobacco and Euglena has been deconvoluted into four major gaussian components with centre wavelengths (CW) at 662, 669, 677 and

-

1

630

I

I

I

1

1

I

I

I

I

I

I

I

I

I

I

640

650

660

670

680

690

700

710

720

Wavelength (nrnl

Fig. 26. Curve deconvolutionanalysis of RCI complex from Pisum sarivum (Chi a/P-700 ratio of 127:Chlalb ratio of 6.6). (From Brown and Schoch, 1981.)


97

686 nm and 663,670,677 and 685 nm, respectively, and minor components at 652 and 690 nm (Brown et a/., 1974).Curve analysis ofthe P-700 Chl a-protein complex of a cyanobacterium (Thornber et a/., 1977) gave four gaussian components with CW at 663, 670, 677, 685 nm and (692) nm; a minor component, CW 708 nm was not due to P-700. These gaussian components accord well with the findings of fourth derivative spectroscopy (Butler and Hopkins, 1970) which reveals A,,, at 673, 679,686 and 695 nm (Thornber et al., 1977). The flash-induced difference-absorption spectrum of P-700-Chl a-complex has been deconvoluted by Schaffernicht and Junge (1981) into two components with CW at 695.5 and 689.9 nm, plus one bandshift. These authors noted this deconvolution also fits the spectra of P-700 from other sources, and that the major variable in the fitting of the curves was the electrochromic component. Mendelian mutants of Chlamydomonas reinhardtii (Chlorophyta) deficient in P-700-Chl a-complex were essentially devoid of gaussian components CW 691 and CW 704 (77 per cent) present with the four major components in the wild strain of this alga (Bennoun and Jupin, 1976). PSII particles prepared from thylakoids using digitonin have less perturbed spectra than those of the P-700 complex or LHCP, since the latter complexes are generally prepared using SDS or Triton X-100, which can cause diminution of their red spectral components. Sugiyama and Murata (1978) analysed the red peak (A,,, 679 nm) of such PSII particles from spinach: four major components at 25’C were obtained, CW 652.4 (Chl b), 662.9,672.1 and 681.6 nm, with minor bands (643.4 and 693.0) a minor 691.0 band was not seen in spectra taken at 77 K. A PSII reaction centre complex from Acrocarpia paniculara (Phaeophyta) (A682, FL692) gave on deconvolution of the red peak (672 nm at 77 K ) major components CW 668 and 679 nm, and two lesser ones CW 660 and 693 nm (Barrett and Duniec, unpublished). The Chl alb-protein LHCP (see Section VIII) isolated from various algae by SDS extraction, and from higher plants has, been deconvoluted into components CW 649,650,670,677 and (640,684) nm (Thornber et al., 1977; Brown et al., 1974; Brown, 1977a): fourth derivative peaks were at similar wavelengths. The far-red peak of a highly purified extrinsic Chl a/b-protein from Lepidium has been deconvoluted into gaussian components with CW at 651.7 nm (Chl b), 661.4 and 669.3 nm (Chl a) with (630.8 and 744), at 77°K (Sugiyama and Murata, 1978). Lepidium Chl alb-protein-663 gave the same component bands, but the relative amounts were different. The CW at 25°C were at -1 nm to the red compared to CW at 77°K. For comparison the spectrum of Chl a in diethyl ether at 25°C had a major gaussian component at 660.5 and a minor one with CW at 647.5 nm. An extrinsic ChI a/b-protein (Chl a/b = 8.0) from Brassica was resolved into a 653.7 nm component (Chl b) and four major components 662,670,677 and 684 nm (Chl a) and 629.9 and 643.1 nm).

98


For a correct evaluation of the contribution of the various spectral forms of Chls in situ to the photosynthesis absorption spectrum, spectra should be determined at temperatures at which photosynthesis proceeds. At 77°K some spectral forms may be attenuated or absent. Further as in most studies at 77°K. glycerol (66 per cent) is used to suspend the Chl-complexes and disturbance of the fine structure of the proteins and lipids, and that of the lipid bilayers and Chl-proteins, cannot be excluded. 3. Dflerence Absorption Spectra This method of optical analysis of composite spectra is useful in determining the A,,, of the main absorption bands of the spectrum of the component pigment-proteins of multiple pigment complexes. Because of excitation coupling of pigment molecules (Shipman, 1980) the absorption maxima revealed may differ from those of the isolated component pigment-proteins. Hiller et al. (1977) demonstrated the presence of Chl a 683 in the P-700Chl a complex of greening barley and pea leaves. The Soret A,,, of Chl a 683 was at 445 nm, at 77°K and 25°C. The contribution of Chl a 683 to the A,,, 675 nm absorption band of the PSI and PSI1 core reaction centre complexes was estimated to be from 18-22 per cent for various Phaeophyta (Barrett and Anderson, 1977, 1980; Barrett and Thorne, 1980). This method established the A,,, of the protein complexes of Chls a, c I and c2and of fucoxanthin in the light harvesting supramolecular complexes of PSI and PSI1 of these algae (Section VIILC), Fig. 37. Difference spectra enable the monitoring of changes in chromophores by chemical agents, light’ or other factors affecting the physical state of the pigment proteins, e.g. pH, specific ion concentration, hydrophobic agents. Kok (1961) established the existence of P-700 by light-induced oxidation of photo-active particles; the absorption difference spectrum having peaks at 698 and 432nm. A heterogeneity in the photoinduced spectra of membrane preparations of the cyanobacteria Spirulina and Fremyella, Porphyridium (Rhodophyta), and in Nicotiana tabacum, was observed in difference spectra at 77°K (Hoarau et al., 1977) The maximum of complexity was seen with Spirulina membranes, where three bands (A,,, 718, 705 and 695 nm) were revealed. Fourth derivative difference spectra of phycobilisomes and thylakoids of various cyanobacteria and of Chlorella (Chlorophyta) at 29°C and 77°K showed significant splitting of the Chl a 680 band (Leclerc et al., 1979). Absorption spectra of cyanobacteria at 77°K have been analyzed by second derivative spectroscopy (Shubin et al., 1979). The conversion of the long wave form (683 nm) of Chl a to a 663 nm form by Triton X-100 has been studied in Chlorella and Porphyridium and in the P700-Chl a complex (Hoarau and Remy, 1978). Similar studies on Chl a 683 with a wide range of detergents has been carried out on the Chla/c,fucoxanthin-protein complex of Acrocarpia paniculata (Barrett, in prepara-


99

tion). Changes in the absorption properties of the p-carotene component were observed in RCI and RCII complexes in the blue-green region of the spectrum. Light-induced changes in phycobilisomes in the presence of dithionite have been studied with difference spectroscopy (Bekasova et al., 1981). Duniec and Thorne (1981) have calculated the effect on absorbancy of light-scattering due to conformational changes. These studies have included slow absorbancy changes around 5 15 nm and the fluorescence changes at about 683nm, both of which arise in chloroplasts from several different chemical and physical causes (Thorne et d., 1983). Circular dichroism (CD) provides another means of identifying forms of Chl. C D spectra of both chloroplasts and detergent isolated Chl-complexes have been described (Scott and Gregory, 1975; Setif et al., 1981). P-700-Chl a complexes gave a negative peak at 684 nm and a positive at 694 nm. A minor C D band at 682nm seen in isolated Chl-protein complexes at 77°K was assigned to PSII; another minor band at 675 nm remained unassigned (Gregory e t a / . , 1980, 1981). A C D band at 639 nm, observed with Chl b rich fractions by Canaani and Sauer ( 1 978) was not seen by Gregory et al. (1980) in their preparations, but a large C D signal observed in aggregated Chl a/bprotein-complex was attributed by them to macromolecular association. Linear dichroism studies, in conjunction with polarized light absorbance and polarized fluorescence, on cyanobacteria and pea chloroplasts have given insight into the specific orientation of the Chl in the thylakoid (Gulyaev and Teten’kin, 1981). Polarized absorption and reflection spectra of Chl in bimolecular lipid membranes have been studied by Krawczyk ( 1981) to clarify concepts of the structure of the photosynthetic membranes. 4 . Fluorescence Spectra Fundamental aspects of the fluorescence of aromatic molecules and their complexes are comprehensively discussed by Birks (1 970) and collectively in Birks ( 1 976). Harnischfeger ( 1 977) has discussed the fluorescence analysis at 77°K of the photosynthesis apparatus. Kinetic aspects of fluorescence in photosynthesis are presented in Section 1V.G.Fluorescence spectra in relation to the forms of Chls are discussed hereunder. Chls have extremely low fluorescence in the absence of HzO or a n electron donor, e.g. lysine or histidine, but when Chls are solvated, as in acetone, their fluorescence is sensitive to the presence of water due to polymer formation (Gurinovich et a f . , 1968). Quenching of fluorescence by H 2 0 probably has little effect in Chl-proteins, where the Chl is held in a hydrophobic cage. Quenching of fluorescence by 02,however, occurs in Chl-proteins (Thorne and Boardman, 1971) and porphyrin-globins (Alpert and Lundquist, 1976). Fluorescence is also sensitive to subtle differences in the molecular interactions of identical chromophores induced by pH changes, cations and buffer components (cf. Brown, 1977b). Concentration effects (Mohanty et al.,

100


1972; Yuen el al., 1980) and the temperature of measurement (Brown, 1977b) affect resolution of fluorescence components and their F,,,. The replacement of H 2 0 by D 2 0 in Chlorella vulgaris does not appreciably alter either the emission or the excitation spectra of the cells (Ghosh et al., 1966). The use of appropriate wavelengths of exciting light can cause fluorescence emission of a particular pigment to be preferentially increased, leading to identification of the species. This has been exploited by Brown (1977b) with preparations of P-700 complexes from Euglena gracilis. Emission at 670 nm was generated preferentially by 430 nm light, while 680 and 690 F,,, were generated by 440nm light. A pronounced shoulder, F,,, 696 was preferentially generated by 450 nm light in these Euglena preparations. With E. gracilis P-700 complex which had been purified by hydroxylapatite in the presence of 1 per cent Triton X-100, distinct emission peaks, at 68 1 and 696 nm were seen in 450nm light (Brown, 1980). These observations accord with excitation energy pathway found in C. reinhardtii (Delepaire, 1980). In E. gracilis an emission band, F,,, 652 Chl b was generated maximally at 77°K by 465 nm light, together with minor band at 705 nm due to the upper harmonic. The relative strength of the fluorescence emission correlated with the concentrations of P-700 in the P-700 complex from a variety of plants (Brown, 1977b). A P-700-Chl a complex with a Chl a/P-700 ratio of 90 when excited at 436 nm had an emission band, F,,, 676 which resolved into 670 and 680 nm components, 450 nm light generated only the F,,, 680 band. The yield of the P-700-Chl a complex (90) was 20 times as great as that of the P-700 complex (30) which had very low fluorescence. The F,,, of the P-700-Chl a complex of Chlamydomonas reinhardtii, was a t 715 nm (Bennoun and Jupin, 1976). In contrast the F,,, of the P-700-Chl a complex of Acrocarpiapaniculara was at 728 nm (Barrett and Anderson, 1981) while Fucus serratus and Cystoseira mediterranea (Phaeophyta) had a substantial band, F,,, 727 nm (Berkaloff etal., 198 1). Rather broader bands were found for Bumilleriopsis (Xanthophyta) and Anabaena (Cyanobacteria) (Brown, 1977b). At 77"K, in addition to the 685 and 695 nm emission bands a broader emission band at longer wavelengths is seen on excitation of intact tissues of higher plants or in intact Cyanobacteria, Chlorophyta, Rhodophyta and Euglenophyta (Murata et al., 1966a,b). In the algae this band is a t 705-720 nm, while in higher plants it is at 720-750 nm. The 685 and 695 nm bands were generally assigned to PSII, and the far red emission to PSI only (cf. Goedheer, 1970; Boardman et al., 1978). This assumption has been questioned by Goedheer (1968) and Lavorel and Etienne (1977). It is now recognized that both Chla and b contribute to the far red emission. This fluorescence is in part due to the excitation to a different vibrational level of the ground state but mainly to the degree of exciton coupling imposed by the geometry of the Chl-protein complexes.


101

Thalli of Phaeophyta and cells of micro-chromophytes exhibit at 77' K two emission bands, F,,, 690-695 and F,,, 705-720 (Brown, 1967; Sugahara et al., 1971; Shimura and Fujita, 1973; Barrett and Thorne, 1981), but there is a great variability amongst species and dependence on the physiological state of the algae. Phorphyra species (Rhodophyta) such as P . perforata are exceptional amongst most plants in having measurable far-red fluorescence (730 nm) from PSI at physiological temperatures (Fork et al., 1982). PSII reaction centre complexes of Acrocarpia paniculata and other Phaeophyta also exhibit 700-750 nm fluorescence (Barrett and Thorne, unpublished), at 77"K, as also does the PSII reaction centre of spinach (Anderson et al., 1978; Satoh, 1980; Larkum and Anderson, 1982).When PSII reaction centre complexes were incorporated into reconstituted liposomes the far red emission was intensified (Larkum and Anderson, 1982); the intensity was dependent on the lipid to protein ratio of the liposomes. The PSI reaction centre complex incorporated into the liposomes was the major contributer to the 735 nm emission. Fractionation of sub-chloroplast particles and isolation of specific Chl-proteins shows that Chl a-protein complexes of both photosystems contribute in different degrees to the far red emission. Minor amounts of Chl with A,,, 690-700nm also contribute to the far red emission (Goedheer, 1981): this may be of two forms, one of which may be involved in energy coupling between PSI reaction centres, the other a participant of PSII. The fluorescence of chloroplasts in situ may be different from that of isolated complexes. The emission spectra (77°K) of brown algae of seven different genera had emission bands at 705-7 15 nm greater than the 694 nm band (Barrett and Thorne, in preparation). The 705-715 nm fluorescence is dramatically decreased in chloroplasts isolated from these thalli in a wide range of osmotic media. Furthermore, a digitonin-prepared supramolecular complex containing the light harvesting Chl a + c,-fucoxanthin protein complex plus PSII reaction centre gave an emission band at 688-720 nm; a distinct emission band, F,,, 71 6 nm, more intense than the 694 nm band was obtained with 436 nm light, but not 450 nm light; a shoulder at 720 nm was also present. This far red emission band may be an expression of the higher organization of the supramolecular complexes. Exposure to detergents such as Triton X-100, known to decrease far red fluorescence (Ogawa and Vernon, 1970), and octylglucoside caused a marked diminution of the 710nm emission. In contrast the anionic steroids cholate and glycholate, tended to sharpen the spectrum of the double emission band. Under some circumstances PSII preparations can give a large emission at 736 nm (Fuad et al., 1983), indicating that great care is necessary in interpreting 735 nm emission as being associated with PSI only. It is clear that several factors affect the shape and intensity of the far red emission bands. One is the number of associated antenna Chls. A Euglena P700-Ch, a complex (30) had a substantial emission band, (F,,, 718 nm),

102


while the P-700-Chl a complex (90) had a very weak emission band at 730 nm (Brown, lY77b). Glycerol (50 per cent) causes a shitt to shorter wavelengths, and the structure of the far red band is dependent also on the temperature of measurement. Lipids can be a determinant in the longwave fluorescence of particles. Kochubei et al. (1981) have studied the effect of galactolipase on such fluorescence from PSI particles from peas. VIII. PIGMENT PROTEIN (LIGHT-HARVESTING) COMPLEXES A. CHLOROPHYLL-PROTEIN COMPLEXES

1 . Association of Chlorophylls with Protein Functional chlorophyll in vivo is now believed to be wholly complexed to

specific proteins through non-covalent linkages (cf. Thornber et al., 1979; Thornber and Barner, 1979; Hiller and Goodchild, 1982; ArgyroudiAkoyunoglou and Castorinis, 1980) and thus analogous to the haemoglobins and cytochromes b. This view contrasts with that generally held until a decade ago that most of the Chl is located in uncomplexed form in the lipid bilayers. The lipid-Chl interactions were considered to be prime determinants in establishing the various in vivo forms of Chls. This view was substantially based on UV-visible, infra-red and NMR spectroscopy studies of Chls in monomeric and different states of aggregation (Katz et al., 1966; Goedheer, 1966; Ballschmitter and Katz, 1969; Katz et al., 1977). Supportive evidence was drawn from studies of the behaviour of Chls at lipid-solvent interfaces (Ke, 1966) and X-ray diffraction studies (see Section V1.A). The Chl-lipid hypothesis, though no longer tenable, led to much experimentation which yielded valuable information about the interactions of the various chemical groups of Chl a with one another, both intra- and inter- molecularly, and with solvent molecules. Such interactions indeed may have a role in the finer organization of Chls in their complexes with protein. An example is the demonstration that the NMR spectra of hydrated Chl a are similar to that of photosynthetically active Chl a, while those of anhydrous Chl a are not (Katz et al., 1968). Convincing proof that essentially all Chls are bound to specific proteins is based largely upon the results of electrophoresis in polyacrylamide gels (PAGE) in the presence of a detergent, commonly SDS. The “free Chl”, amounting to about 40 per cent of the total Chl, found by most workers until the mid 1970susing SDS-PAGE is not present, except for a few per cent, in the current milder PAGE procedures. (Henriques and Park, 1978; Anderson et al., 1978, 1981; Markwell et al., 1979; Bennett et al., 1981; Wild and Urschel, 1980). An alternative procedure, differential extraction of the Chl-complexes with a series of detergents coupled with density gradient centrifugation, shows


103

that in the brown macro-algae the Chl is entirely associated with protein (Barrett and Thorne, 1981). It should be noted that electrophoresis is not initially a mild procedure, since under the influence of an electric field any Mgjamino-acid bond is likely to be disrupted in buffers of neutral or acidic pH, or those containing an anion which can complex with the Mg atom of Chl. I3C-NMR spectroscopy reveals a heterogeneity in the Chl environment of the thylakoid. About 30 per cent contributes to the high resolution I3C spectrum (Eigenberg et al., 1981) and the NMR data suggest that part of the Chl is loosely associated with protein, or at least that the phytyl chains are associated with lipids. This 30 per cent of the Chl may correspond to that portion of Chl a which is easily dissociated during SDS-PAGE. Separate evidence of the bonding of Chl to protein is derived from resonance Raman spectroscopy (Lutz, 1977; Lutz et al., 1978; Lutz et ul., 1982). The X-ray crystallographic structure determination of a BChl aprotein (Fenna and Mathews, 1979; Mathews et d . , 1979) provides a precise view of one arrangement of BChl ascomplexed to protein (Fig. 42). This BChl a-protein, however, is not a typical membrane bound Chl-protein. 2. Types of Chl-protein Complexes The Chl-proteins comprise three major groups: (i) the reaction centre complexes of PSI and PSI1 (Section VII); (ii) the core antenna Chl a which is immediately associated with the reaction centres and transmits photoenergy from the light-harvesting complexes to P-700 or to P-680; to these may be added peripheral antennae Chl a-proteins (cf. Mullet et al., 1980a,b) (iii) the supramolecular complexes which harvest light energy, principally for PSI1 (Section VIII). In certain algae xanthophylls may be as important as Chls for light-harvesting. These Chl-complexes comprise the Chl alb-siphonaxanthinprotein amongst the Chlorophyta and Chl u + c,-fucoxanthin-protein complex and the Chl a + c I + c,-violaxanthin-protein complex in the Chromoph yta. A water-soluble peridinin-protein has been obtained from some dinoflagellates and this has only Chlu apart from the xanthophyll (see Section VII1.C. I), but here the peridinin is the major light harvesting pigment. Rhodophyta and Cyanophyta have water-soluble phycobiliproteins for lightharvesting, but some of the Chl a gathers light for PSI. In certain Cryptophyta Chl a2 is an adjunct to the predominant phycoerythrin or phycocyanin (Section VI.C, VII1.B). Except for the water-soluble Chl 661 and Chl 663 of Chenopodium album and Lepidium virginicum (Yakushiji et al., 1963; Murata and Ishikawa, 1981) most Chl-proteins so far isolated are hydrophobic proteins which in vivo are embedded in the lipid matrix of the thylakoid membranes through protein-lipid and protein-protein interactions. Detergents are required for their release from the thylakoids and with the more vigorous detergents this

104


results in the concomitant fragmentation of the membranes. Depending on the type of alga and the detergent used, selective release of some Chl-proteins can be effected (Sections VII and V1II.C) leaving the gross structure of the membrane intact; this provides some insight into the relationship of the Chlcomplexes in situ (Barrett and Thorne, 1981; Green and Camm, 1981). Lipids, especially carotenoids and phospholipids, may be an integral part of the complexes (Siefermann-Harms, 1980a,b;Anderson et al., 1981; Tremolieres et al., 1981). 3. Solubilization of Chl-protein complexes by detergents and their fractionation The two detergents most frequently used over the last decade have been the anionic sodium dodecyl sulphate (SDS) or the non-ionic Triton X-100 (polyoxyethylene p t-octylphenyl) (cf. Table IV). LiDS was introduced by Delepelaire and Chua (1979) because of its higher solubility than SDS at 4°C. The zwitterionic detergent, lauryldimethylamine oxide (LDAO), much used for cytochrome oxidase (Saunders and Jones, 1975) and for bacterial reaction centres (cf. Olson, 1981) has surfactant properties similar to Triton X-100 but is less harsh and does not cause the shifts of the A,,, of the red peak of Chl a/bprotein commonly observed with Triton X-100. Deriphat 160 (disodium Nlauryl-P-imindopropionate) and Zwittergen (TM 12-N-dodecyl-N, Ndimethyl-3-ammonia-1 -propanesulfonate) have been added to the range of detergents used (Markwell et al., 1980; Barrett and Thorne, 1981). The Tweens (polyoxyethylene sorbitans) are useful as mild detergents for the differential solubilization of thylakoids. The glycosteroid digitonin was exploited to split the two photosystems rather than to obtain individual protein-complexes (Anderson and Boardman, 1966). Digitonin is much favoured now for the separation of Chlalb-proteins from RCI and RCII complexes (Yamaoka et al., 1978; Satoh, 1980; Larkum and Anderson, 1982; cf. Jennings et al., 1980). Recently octyl-l)-D-glucopyranoside has been used for differential extraction of thylakoids in Acetabularia (Chlorophyta) and a variety of green plants (Camm and Green, 1980). Deoxycholate has been used for fractionating the chloroplast membranes from Chfamydomonas reinhardtii (Chlorophyta) (Bar-Nun and Ohad, 1977). Octa-ethyleneglycol mono-ndodecyl ether (Nikko Chemicals Co. Ltd., Tokyo) has been recommended as a mild replacement for Triton X- 100. Valuable background knowledge of selection and conditions of use of detergents for the solubilization of membranes, as well as for the preparation of vesicles for model studies, is provided by the reviews of Helenius and Simons ( I 975), Reynolds (1980) and Helenius et al. (1980). Detergent-solubilized Chl-protein-complexes are commonly fractionated by PAGE in the presence of SDS, LiDS, or other detergents, and a reductant of S-S bonds (cf. Machold et al., 1979). PAGE in urea is sometimes useful to


105

dissociate Chl-protein oligomers. Density gradient centrifugation (Argyroudi-Akoyunoglou and Thomou, 1981) or chromatography on DEAE-cellulose or Sepharose (Nakayama et al., 1979; Ilina and Borisov, 1980) are useful. Isoelectric focusing has come to the fore recently in the separation of Chl a/b-proteins and RCII and RCI complexes (Satoh, 1979; Heinz and Siefermann-Harms, 1981; Larkum and Anderson, 1982). 4. Chl alb-protein Complexes The name light-harvesting Chl a/b-protein (LHCP) was proposed by Thornber and Highkin (1974) for the Chl alb-protein which previously had been variously termed Complex 11, CPII or PSII Chl-protein; these terms recognized that the Chl a/b-protein was associated with PSII. Recently Haworth et al. (1981) have claimed that a small amount of a Chl alb-protein is associated with the outer antenna of RCI. The light-harvesting Chl a/b-protein complexes of higher plants have been reviewed generally (Thornber, 1975; Thornber et al., 1977) and in respect of their structural organization (Hiller and Goodchild, 1981) and in relation to model Chl-protein complexes (Thornber and Barber, 1979). The ensuing discussion will refer to higher plants only where lacunae exist in our knowledge of the algal Chl a/b complexes. Kan and Thornber (1976) chromatographed SDS extracts of Chlamydomonas reinhardtii thylakoids on hydroxylapatite and obtained a Chl albprotein of M, 28 K D f2 KD. On the basis of amino acid analysis and Chl assay the M, of this Chl-protein was about 35 K D (the protein being 29 KD), in accord with the value of 4530 g protein per mole of Chl calculated on the basis of the content of several different amino acids. This Chl alb protein had a sedimentation coefficient of 23.8. On average, a mole of protein of 29.4 K D was associated with 3 moles of Chl a and 3 moles of Chl b and only one mole of carotenoid. As the Chl a/b complex contained neoxanthin, violaxanthin, lutein and p-carotene, in close proportion, it was concluded that there must be a heterologous distribution of the carotenoids amongst the polypeptides of the Chl a/b-protein. It was neither established whether any of the carotenoids were on the same polypeptides as the Chl, nor whether the Chl a and the Chl b were together on the same peptide. The Chl a/b-protein had absorption peaks at 670, 652,470 and 437 nm. The millimolar extinction at 670 nm was 73 f4 (based on Chl a), and at 652 nm the extinction was 59 f6 (based on Chl b). It is of significance for understanding the bonding of the Chl to the protein that this Chl-protein has a sloping shoulder on the blue-side of the 437 nm peak of the Chla, similar to Chla in monomeric state in organic solvents. This contrasts with the shoulder-peak seen in the spectra of reaction centre complexes of both PSI and PSII (Alberte and Thornber, 1978; Stewart, 1980; Barrett and Anderson, 1980; Barrett and Thorne, 1981). The amino acid analyses of the Chlalb-protein of the chlorophytes

106


Chlamydomonas reinhardtii and Acetabularia and of higher plants when compared to those of the corresponding P-700-Chl a-protein complexes reveals a crucial difference. The histidine content is low in the Chl a/b-proteins (Kan and Thornber, 1976; Apel, 1977), and is three or four times higher in the P-700-Chl a-complexes (Kan and Thornber, 1976; Apel, 1977; Lagoutte et al., 1980). Low histidine content has been found also in the water-soluble lightharvesting Chl a/b-proteins CP661 and CP663 (Murata and Ishikawa, 1981). Ohad and associates in a study of the Chl a/b-proteins of Chlamydomonas reinhardtii y- 1 during chloroplast membrane biosynthesis obtained two polypeptides of M, 24 KD and 22 K D by SDS-PAGE (Bar-Nun et al., 1977). Both polypeptides had equal molar ratios of Chla and b. These findings accord with those of Anderson and Levine (1974), that the peptides of 24 and 22 KD were absent from Chlarnydomonas lacking the Chl a/b-proteins. BarNun er al. (1977) were able to show that upon denaturation the 28 K D polypeptide could be separated into two pigment-proteins of 24 and 22 K D and a colourless protein of 28 KD which, though co-migrating with the Chl a/b-protein, was not an active component of the Chl a/b-protein complex. Both the 24 and 22 KD-polypeptides carried 6 Chl a and b molecules, an average of 1 mole of Chl for each 4 KD of polypeptide. This contrasts with the 4 moles (Chl a/b = 1) bound to proteins of M, 78-80 KD in CP661 and CP663 (Murata and Ishikawa, 1981).The Chl a/b-protein complex was absent from a 37°C grown C . reinhardtii 7 mutant and from a Euglena sp. lacking Chl b. However the RCII complex was still active. Further mutant studies showed that different polypeptides are required for the stabilization of LHCP and RCII complex. The molecular weight of the native Chl a/b-protein complex was calculated to be 57 KD. A Chl a/b-protein complex has been isolated from Acetabularia mediterranea chloroplast by SDS-PAGE after solubilizing the membranes with Triton X-100 combined with EDTA (Apel, 1977a,b). The M, of the complex was 67 KD, but this was resolved by electrophoresis into 23 K D and 21.5 K D subunits. The molar ratio of these two polypeptides was 2 : 1. This discrepancy between the apparent M, and that calculated is due to anomolous migration in SDS-PAGE (cf. Chua et al., 1975). Each of the subunits contains about 1 per cent of sugar, largely glucose moieties. This is of note in respect to the presence of sugar components in the light-harvesting complexes of brown algae (Section VIII.C.2) and the finding that galactolipids are associated with Chl-protein complexes of spinach (Heinz and Siefermann-Harms, 1981). Recently several workers using milder PAGE procedures have detected the existence of minor Chl a/b-protein complexes having Chl a/b ratios which differ from the classical LHCP (Tremolieres et al., 1981). Wild and Urschel (1980) isolated a normal LHCP from Chlorella fusca with a Chl a/b ratio of 1.3. This value accords with Chi a/b ratios of LHCP obtained from higher plants by PAGE or by chromatographic procedures (Anderson et al., 1978;


107

Boardman and Anderson, 1978) and for Sinapsis alba (Wild et al., 1981). The LHCP accounted for 40-60 per cent of the total Chl. In a Chl-deficient mutant of Chl.fusca, the Chl a/b ratio of LHCP was 1.5-1.8, but this Chl comprised only 33 per cent of the chloroplast Chl. By limited extraction of thylakoids of a variety of plants and of Acetabularia, using octylglucoside followed by SDS-PAGE of the extract, two polypeptides of M, 26 and 27 K D were obtained (Green and Camm, 1981). The Chl a/b ratios were the same for both polypeptides, in a range of 1 .0-1.3. The LHCP was isolated mainly in its oligomeric form. A minor Chlalbprotein, designated C29, consisting of a single polypeptide of M, 29-30 KD, was also isolated from Acetabularia and some higher plants (Green and Camm, 1981). 5 . Forms of LHCP and Adjuvant Lipids

Multiple forms of Chl a/b-proteins with different electrophoretic mobilities have been isolated by various systems of SDS-PAGE. It is difficult to relate these forms because of variations in technique amongst the different workers. There is not agreement on the origin of the three forms of LHCP, designated by indices 1-3 (Anderson et al., 1978; Machold and Meister, 1980) or ABl, AB2 and AB3 by Markwell et al. (1979). Most workers have considered these to be forms of a single Chl a/b-protein. Bennett et al. (198 1) now propose that the predominant LHCP, and the most stable on electrophoresis, is generated from the additional forms of Chl a/b-proteins, AB-1, AB-2 and AB3, during electrophoresis of SDS extracts of thylakoid membranes. The problem is further complicated by the possibility of glycosylation (Apel, 1977a,b), the copresence of colourless peptides, the role of cations in oligomeric transformations (Argyroudi-Akoyunoglou and Thomou, 1981) and reversible phosphorylation of the main apoprotein (Bennett, 1979), which latter process may determine distribution of excitation energy between PSI and PSI1 (Bennett et al., 1980; Horton and Black, 1981; Allen et a f . . 1981; Section 1X.C). Complex lipids appear to be involved in the organization of the LHCP oligomers (Sieferman-Harms, 1980). Tremolieres et a f . (1981) obtained a higher content of phosphatidyl-diacylglycerol in the oligomeric forms of LHCP than in the monomer. Trans-hexadecenoic acid was present in the diacyl lipid: the trans-fatty acid is only present in biological membranes that have LHCP and is not found in membranes of Cyanobacteria. Spinach LHCP contains five diacyl lipids; diacylgalactosylglyceride accounts for half of the total amount (Ryrie et al., 1980). Douce and Joyard (1979) suggest that carotenoids may stabilize the conformation of the complexes within the thylakoid membrane. 6. LHCP of Other Marine Algae In four different marine green algae the Chlalb ratios of the LHCP were

108


0.8-1.1 (Nakamura et a f . , 1976). The oligomeric forms of the LHCP were contaminated with the P-700-Chl a-complex judging from the Chl alb ratio of 4.8. The ratio of Chl P-700 was 27 per cent less on average in these four marine algae compared to the Chl P-700 ratio of four higher plants also studied. Thus algae which are found to have different Chl alb ratios for their LHCP may have varied the proportion of the total Chl associated with LHCP, or have changed the Chl b composition of the LHCP. A Chl alb-protein complex has been obtained from a unicellular prokaryotic marine alga (Prochforon;Section XII) which is associated with dideminid ascidians inhabiting coral reefs and other locations in the Pacific ocean (Kott, 1980).The Chl alb ratios ranged from 26-12.0 for samples from the Barrier Reef (Thorne et al., 1977) and from 4.46.9 for Prochlorons from a wider variety of locations (Withers et al., 1978). The Chl a/b-proteins account for 26 per cent of the total Chl of the Prochloron and show similarities to the LHCP of higher plants. There are 260 Chl’s per P-700. 7. Chl alb-siphonaxanthin-proteinComplex A light harvesting complex that is unusual in having a xanthophyll, siphonaxanthin, as an integral part of the Chl alb-protein has been isolated from two chlorophytes Caulerpa cactoides (Anderson et al., 1980) and from Codiumfragile (Anderson et af., 1981). This Chl a/b-siphonaxanthin complex accounts for 60 per cent of the total Chl in Caulerpa. The Chl a/b ratio was 0.62-0.74, similar to the LHCP of Acetabularia mediterranea. The bands

Emission ot 685 nm

I

400

450

I

I

500 550 Wovelength (nm)

600

Fig. 27. Fluorescence excitation spectra of Wlva (Chlorophyta) showing contribution of siphonaxanthin. (From Kagayama et al., 1977.)


109

Qbtained on SDS-PAGE corresponded to LHCP', LHCP' and LHCP3 of higher plants (Section 4). The fluorescence excitation spectra of the complexes resembled the action spectra for photosynthesis of the intact alga (Kageyama et al., 1977) (Fig. 27). B. PHYCOBILISOMES AND BILIPROTEIN AGGREGATES

1 . Discovery and General Features The first suggestion that phycobiliproteins of Cyanobacteria and Rhodophyta might be aggregated into discrete bodies was made by Myers et al. (1956) when 22 nm granules were observed between the thylakoids in chloroplasts of Grifithsia josculosa (Rhodophyta). Gantt and Conti ( 1 965, 1966a) observed similar bodies in the unicellular red alga Porphyridium cruentum (Protojorideae, Rhodophyta) and then showed that they contained PE and PC, and were attached to the stroma surface of thylakoids (Gantt and Conti, 1966b). Since then phycobilisomes (PBS) have been observed in and isolated from many Cyanobacteria and Rhodophyta (cf. Gantt, 1980, 1981; Glazer, 1981) and have been shown to contain APC (Gantt and Lipschultz, 1972). In the cyanobacteria Mastigocladus laminosus and strains of Anabaena variabilis phycoerythrocyanin is also present (see Section V1.C for references) PBS are not found in Cryptophyta. In species of the Cryptophyta the phycobiliproteins are probably contained in the intra-thylakoid space rather than attached to the outer surface (Gantt, 1979); yet the efficient transfer of light energy absorbed by PE or PC to the reaction centres of PSI and PSI1 (Gantt, 1979; Lichtle et al., 1980) indicates a specific arrangement of the biliproteins in the intra-thylakoid space and a molecular connection to the thylakoid membrane. 2. The Structure of Phycobilisomes Two basic types of PBS have been observed: disc-shaped and globular (or spherical). Disc-shaped PBS occur mainly in Cyanobacteria but are also found in some Rhodophyta; globular PBS occur predominantly in Rhodophyta (Gantt, 1980, 1981), see Fig. 28. PBS have been isolated from many Cyanobacteria and Rhodophyta (Gantt and Conti, 1965, 1966a; Wildman and Bowen, 1974; Koller e t a l . , 1978; Bryant et al., 1979; Gantt et al., 1979; see also references in Gantt 1980, 1981 and Glazer, 1981). High K-phosphate (-0.75M) media have proved efficacious, in conjunction with Triton X-100, for isolating PBS (Gantt et al., 1979). A sucrose (0.5 M), phosphate (0.5 M) and citrate (0.3 M) medium has been found to preserve the attachment of PBS to the thylakoid membrane (Katoh and Gantt 1979). In all types of PBS the constituent phycobiliproteins and colourless polypeptides (see below) appear to be held together by noncovalent intermolecular forces, and solutions of low ionic strength cause the

110


PBS to fall apart (Gantt and Lipschultz, 1972; Gantt et al., 1979; Zilinskas and Glick, 1981). Zilinskas and Glick (1981) and Siegelman (1982) concluded that hydrophobic interactions are most important. Dispersion forces were also a significant factor, but the requirement of high salt concentrations for PBS preservation was not simply explained as the countershielding of charges on proteins (Zilinskas and Glick, 198 1). Although high phosphate concentrations preserve intactness in vifro this does not imply that such conditions pertain in vivo. Lilley and Larkum (1981) have shown that carbon dioxide fixation in intact chloroplasts of red algae is inhibited above 5 mM phosphate, but the ionic composition of the stroma in these algae is not known. Siegelman and Kycia (1982) has shown that if PBS are kept in dense suspension, high phosphate concentrations are not necessary for their integrity. There is strong evidence (Section V1.C; Table 111) that disc-shaped PBS are constructed of ( a m 3 units of phycobiliprotein. These in turn aggregate in pairs to form ( a R 6 discs. According to the model of disc-shaped PBS in red algae (Koller et al., 1978), APC discs form a triangular base for radiating rods in which PC is proximal and PE is distal; the PC and PE forming tripartite units of 1 PC + 2 PE of ( a m 6 discs. A similar model has been proposed for cyanobacterium LPP-7409 (Bryant et al., 1979) (Fig. 29) and for Fremyelfa diplosiphon (Rosinski et al., 198 1). In these cyanobacteria the ratio of PE to PC is not fixed and can change with complementary adaptation (Section X.B); changes in length of the radiating rods have been related to changes in the amount of PC and PE. Further investigation has shown that both PE and PC are differentiated into two different types in certain chromatically-adapting cyanobacteria, forming distinct (c$)6 discs (Bryant and Cohen-Bazire, 198 1; Gingrich et af., 1982). Each rod of a PBS of Synechocystis 6701 appears to consist of four such discs in the order -27 KD PC, - 33.5 KD PC,

Fig. 28. Models for the possible arrangement of (a) hemi-discoid phycobilisomes and (b) globular phycobilisomes on the thylakoid membrane. See text for further details.


111

(a/$6-31.5 K D PE and (a& -33.5 K D PE starting from the core. Thus the arrangement of the biliproteins parallels the pathway of energy transfer (see below) in the intact PBS: PE+PC+APC-+Chl. Koller et al. (1978) estimated that a disc-shaped PBS of the Rhodophyta units and six tripartite was composed approximately of three APC PE-PC units with a total M, of approximately 6000 KD. Globular PBS have about the same diameter (30-40 nm) as disc-shaped PBS (Gantt, 1980). but being prolate spheroids they occupy a much larger volume. In Porphyridiuni cruentum there is a total of 60-80 (aP)3units in the ratio of 50-70 PE, 8R-PC and 4 APC, giving a total M, of approximately 10 000 K D (Gantt, 1980; Dilworth and Gantt, 1981). The largest PBS is found in the rhodophyte Grzjithithsia pac$ca (Gantt and Lipschultz, 1980) with dimensions of 63 nm x 38 nm x 38 nm. Studies of disaggregation (Gantt et al., 1976a,b) and immunochemical evidence (Gantt et al., 1976a,b) and immuno-electron microscopy (Gantt and Lipschultz, 1977) again suggest that PE is arranged towards the outside of globular PBS and APC nearest the thylakoid membrane. Two recent studies (Wanner and Kost, 1980; Dilworth and Gantt, 1981) have documented the topography and distribution of PBS on the thylakoid membrane in Porphyridium cruentum. In an interdigitated configuration there is much unused space on any single thylakoid membrane (Fig. 28) (Wanner and Kost, 1980; Section 1X.B). Negative-staining measurements indicate that in a single cell of Porphyridium cruentum there are 5 to 7 x 10’ PBS on a total thylakoid area of 1.1 to 1.6 x lo3 pm’ (Dilworth and Gantt, 1981). There are approximately 450 PBS pm i.e. 50-60 per cent of the available surface area. This figure is probably reduced in an interdigitated configuration (Wanner and Kost, 1980). Until recently it was assumed that the PBS was composed only of phycobiliproteins which had interacted by non-covalent bonds during assembly to form PBS. There are a number of colourless polypeptides in PBS (Tandeau de Marsac and Cohen-Bazire, 1977; Koller et al., 1978; Yamanaka et al., 1978; Lundell et al., 1981; Bryant and Cohen-Bazire, 1981) and at least some of these are involved in PBS assembly. In Synechococcus 6301

-’

Fig. 29. Model for the molecular arrangement of phycobiliproteins in a hemi-discoid phycobilisome. (Redrawn from Bryant er al., 1979.)

112


polypeptides of M, 27 KD, 30 K D and 33 K D have been shown to mediate the assembly of phycocyanin into discs and rods (Lundell et al., 1981). Another colourless “linker” polypeptide of 75 K D may be involved in linking the PC rods to APC or the APC to the thylakoid membrane. Bryant and CohenBazire (1981) have shown that in Pseudanabaena 7409 (Cyanobacteria) the number of colourless polypeptides is 8 when growth is in green light and 6 in red light and that this change is related to other changes involved in chromatic adaptation (Section X.B). Partial or complete reassembly of PBS is accompanied by fluorescence emission changes suggesting that polar energy transfer occurs along rods towards the core (Canaani et al., 1980; Lundell et al., 1981). In vitro partial reassembly of phycobilisomes has recently been demonstrated in Porphyridium cruentum (Canaani et al., 1980) with APC and a PC-PE complex, and in Porphyridium sordidum (Lipschultz and Gantt, 1981)with PE and PC. However under certain undefined conditions APC was unrecombinable and it is possible that this was due to the loss of a mobile linking polypeptide. Gantt et al. (1981) have further studied the role of such polypeptides in P . cruentum, and Gantt et al. (198 1) and Redlinger and Gantt (1981) have suggested that a 95 KD polypeptide which is common to both PBS and thylakoids is involved in anchoring the PBS to the thylakoid membrane. 3. Fluorescence Evidence of Eaergy Transfer Isolated phycobiliproteins exhibit strong fluorescence emission (Fig. 30). However fluorescence from these components is very low in vivo. Haxo and Blinks (1950) showed that light absorbed almost exclusively by the phycobiliproteins led to high rates of photosynthesis in Rhodophyta (Section V1.D) and yet gave rise to fluorescence from Chl a only. They concluded that light energy is passed on from the biliproteins to Chl with almost 100 per cent efficiency. More recent estimates are slightly lower (Govindjee and Govindjee, 1975; Grabowski and Gantt, 1978a,b). Isolated phycobilisomes also exhibit low fluorescence from most of the biliproteins, but have a characteristic fluorescence at about 670 nm, at room temperature, in the region of the fluorescence maximum of Chl and APC (see below). Yet the partially disaggregated PE-PC complex has strong fluorescence at the PC emission peak (644nm) when the PE is excited at 546nm (Koller et al., 1978) indicating intermolecular transfer of excitation energy from PE to PC. The fully disaggregated PE and PC show only fluorescence at their individual emission peaks of 572 and 644nm respectively (Gantt and Lipschultz, 1973). This is evidence for the migration of energy from PE+PC+(APC, Chl). Direct evidence for such a route has come from a study of the small residual fluorescence from all these components when PE is excited (Porter et al., 1978; Searle et al., 1978). The rise-times of fluorescence from all these components (Fig. 31) is entirely consistent with the route


113

565

Einission 683 nm

Absorbance

I

Ii I \

Emtation

/

500nm

1

\/

I I

I /

'P /

420

493

I

500

Wavelength

I

I'

600

nm

Wa ve l e n g th nm

Fig. 30. Absorption spectrum and fluorescence excitation (a) and emission spectra (b) for isolated phycobilisomes of Grifithsiu monilis (Euflorideae, Rhodophyta). The fluorescence spectra are for phycobilisomes at 77-K. A Huorescence excitation and emission spectrum is shown (broken curve) for disrupted phycobilisomes (in 1 mM phosphate buffer). (Data of Hiller and Larkum, unpublished).

PE+PC-+APC-+Chl. Furthermore, the kinetics of the energy migration suggest an ordered system in which random walk processes are restricted, that is, consistent with the rod structure of the phycobilisome (Section 1X.B). Theoretical support for this scheme of energy migration comes from the Forster theory of inductive resonance energy migration (Section 1X.A). Such energy transfer depends on close association between the donor and acceptor molecules, and falls off with the sixth power of the distance between the chromophores. For photosynthetic pigments efficient migration also depends on an adequate lifetime of the excited state of the donor molecule, and a good overlap between the fluorescence emission spectrum of the donor molecule and the absorption spectrum of the acceptor molecule. For these reasons Duysens as long ago as 1952 proposed that energy migration was from PE+PC+Chl a. Much evidence is consistent with the migration of energy in phycobiliprotein aggregates by inductive resonance (Teale and Dale, 1970; Dale and Teale, 1970; Grabowski and Gantt, 1978a,b; MacColl and Berns, 1978). However, exciton migration may also be involved (Section 1X.A). For phycobilisomes the most recent information suggests that the predicted

114

A . W. D. LARKUM A N D JACK BARRETT

route is PE+PC+APC+APC-B (or APC-l)+Chl a (Gantt 1980), and for Cryptophyta the predicted route is PE or PC-Chl a or PE+Chl c2 (MacColl and Berns, 1978). The late advent of APC into the proposals is due to the more recent characterization of these biliproteins (Gysi and Zuber, 1976; Glazer and Bryant, 1975; Zilinskas et al., 1978; Canaani and Gantt, 1980) despite earlier identification (Haxo el al., 1955). Four forms of APC have been isolated: APC I, I1 and 111 and APC B (Glazer and Bryant 1975; Canaani and Gantt 1980; Bryant and Cohen-Bazire, 1981), but they do not always occur together. A

0

Picoseconds

Fig. 31. The rise-times of fluorescence from phycobiliproteins and Chl a in dark-adapted Porphyridium cruentum. The upper set of curves represent experimental results. The lower set represent estimates based on a kinetic analysis for direct excitation of B-phycoerythrin (Porter et al., 1978). (Redrawn from Searle et al., 1978.)

Both APC 1 and APC B in their hexameric form, (a& have fluorescence emissions close to those of intact, isolated PBS (Clement-Metral and LefortTrans, 1971; Gantt et al., 1979); that is at 670-675 nm at room temperature and at 68CL685 nm at 77°K. In Rhodophyta evidence suggests the presence of APC B but not APC 1 (Ley et al., 1977; Gantt, 1980). This indicates that APC B which forms only a small proportion of the total APC in these algae is the bridging pigment between the PBS and the thylakoid membrane (Gantt, 1980; Glazer, 1981)and that the remainder of APC has a light-harvesting role as first


115

shown by Lemasson et al. (1973). Thus the route of energy migration in these algae appears to be PE+PC+(APC)LH+APC B-,Chl a. In Nostoc however both APC 1 and APC B can occur together with the other forms of APC (Canaani and Gantt, 1980). Here both APC 1 and APC B could act in parallel in channelling energy to the thylakoid membrane or there could be two types of PBS with one or other form of these APCs as the bridging pigment. Mimuro and Fujita (1980) have found that the contents of APC 1 and APC B vary widely in the Cyanobacteria, Anabaena cylindrica, Anacystis nidulans and Anacystis variabilis. They suggest three types of interaction: (i) that in which APC B is abundant enough to be the bridging pigment, (ii) in which APC 1 replaces APC B and (iii) in which APC 1 and APC B are rare and the bulk light-harvesting APC is the bridging pigment. These proposals remain speculative for lack of critical evidence. A further question which has not been fully resolved is the presence of Chl a in isolated PBS. The fluorescence of intact, isolated PBS at 670-675mm (680-685 nm at 77°K) is close to the emission bands of some forms of Chl a. Disaggregated Chl a has an emission peak at 680-682 nm, at 77^K,especially in the presence of Triton X-100 (Satoh and Butler, 1978a) and the PSII RC complex, has an emission at 685 nm at 77°K (Govindjee and Zilinskas, 1974). The lowering of the Chl concentration to one Chl a to ten PBS by exhaustive extraction in cold acetone, causing no diminution of the 685 nm emission, suggests that Chl a is not a natural component of PBS (Gantt et al., 1976a,b; Katoh and Gantt, 1979). Mimuro and Fujita (1980) disagreed with this view. From experiments with three species of Cyanobacteria they concluded that the PBS fluorescence arises from Chl a, because the fluorescence from APC 1 and APC B was at a lower wavelength and the fluorescence from PBS was diminished when Chl a was removed by cold methanol. 4 . Phycobiliprotein-ThylakoidInteractions One of the least understood aspects of phycobiliproteins is the interaction of PBS (in Cryptophyta, the PE or PC) with the thylakoid membrane and PSI and PSII (Fig. 32). The previous discussion of PBS structure and energy migration has raised some of the problems. The discussion here concerns the transfer of energy absorbed by phycobilins to PSI and PSII units in the thylakoids. There is much evidence from fluorescence and action spectra studies (Haxo and Blinks, 1950; Brody and Brody, 1962; Haxo and Fork 1959; Fork, 1963; Jones and Myers, 1964; Cho and Govindjee, 1970b; Larkum and Weyrauch, 1977; Mimuro and Fujita, 1977; Wang et al., 1977; Ley and Butler, 1977a,b, 1980a,b; Diner, 1979) to suggest that light energy absorbed by phycobiliproteins is passed on with high efficiency to RCI and RCII, but the evidence on the bridging of the molecules is scanty. Indirect evidence comes from studies of the numbers and arrangement of PBS, freeze-fracture particle distribution

116


2

Fig. 32. Models for the distribution ofexcitation energy from phycobilisomes to PSI and PSII.

and fluorescence studies. The density of disc-shaped PBS on the thylakoid stroma face ranges from 1200-1400 per nm2 (Lichtle and Thomas, 1976) whereas, because of greater size the packing of globular PBS ranges from 165-400 per nmz (Neushul, 1971; Waalund et al., 1974; Lichtle and Thomas, 1976; Staehelin et al., 1978). Such a difference by itself indicates that there are very different degrees of interaction between the two types of PBS and the PSI and PSII units. Other evidence comes from freeze-fracture studies. Bourdu and Lefort (1967) observed from certain profiles that PBS were arranged on the thylakoid lamellae in rows approximately 50 nm apart, and this was confirmed for a number of algae (Guerin-Dumartrait et al., 1970; Neushul, 1971; Lefort-Tran et al., 1973; Wollman, 1979). Lefort-Tran et al. (1973) found that large (10 nm) freeze-fracture particles on the exoplasmic faces (EF particles-Section 1X.B) were also arranged in rows 50 nm apart and these were next to the rows of PBS. Lefort-Tran et al. (1973) suggested an arrangement (Fig. 33) which has received support from other investigations (cf. Gantt, 1980). There is good evidence that the large EF particle is the site of PSII (Section 1X.B). The freeze-fracture evidence further suggests that PBS are attached to the thylakoid membrane adjacent to the PSII units and allows a comparison of the density of PBS and putative PSII particles. Gantt (1980) has tabulated the evidence on this aspect. In Cyanobacteria the ratio of PBS to E F particles is between 1 and 2, but in Rhodophyta this ratio is between 0.2-0.5. This means either that in Rhodophyta only some PBS are connected


117

to PSII units or that PBS can be associated with more than one PSII unit. Thus Cyanobacteria and Rhodophyta may exhibit two different strategies for light-harvesting: in Cyanobacteria (and some Prorojlorideae of the Rhodophyta) the PBS is small and is approximately equal in numbers to the large E F particles, but in the majority of Rhodophyta the PBS are larger and spatial constraints prevent a 1 : 1 connection with the large EF particles (Section IX.B, for further discussion).

50-6Cnm

Fig. 33. A model for the arrangement of globular phycobilisomes on the thylakoid membrane in relation to the large EF freeze-fracture particles (putative PSII units). (Redrawn from LefortTran er al., 1973.)

Green light absorbed by PBS contributes with high quantum efficiency to photosynthesis (Brody and Brody, 1962). From fluorescence data Ley and Butler (1976, 1977a,b, 1980a,b) calculated that under normal conditions all the light energy absorbed by PBS (excitation energy) migrates to PSII and 50 per cent migrates from PSII to PSI. Their data on Porphyridium cruentum grown under white light indicate that only 5 per cent of the Chl a is in PSII with the other 95 per cent in PSI (in agreement with Amesz and Duysens, 1962). However according to Ley and Butler (1980a,b) the light quality and intensity may alter both the distribution of Chl and the energy transfer between the photosystems (Section X.D). Some of the assumptions in this work have been challenged (Wang et af., 1980; Section IX.A,B). The evidence from fluorescence therefore supports a model similar to that suggested by the freeze-fracture evidence; i.e. a close connection between the PBS and PSII and a secondary link between PSII and PSI. Larkum and Weyrauch (1977) pointed out, however, that it is difficult to conceive of an efficient distribution of energy through PSII which had only 5 per cent of the total Chl a. A different model was suggested in which non-fluorescent Chl was the distributor of PBS energy. Harnischfeger and Codd (1978) and Schreiber

118


(1979, 1980) found that preillumination and low temperature affected the distribution of energy from the PBS to the thylakoid membrane, presumably by disrupting some of the molecular (APC?) connections to the photosystem units. Evidence other than fluorescence is needed to establish the size and structure of PSI and PSII in Cyanobacteria and Rhodophyta. The study of Chl-protein complexes provides an alternative approach (Section V1I.B). Hiller and Larkum (1981) have isolated the RCII complex (which may account for 25 per cent of the total Chla from GrifJithsia monilis (Rhodophyta). In this alga, where the fluorescence evidence suggests that a small proportion of the Chla is in PSII (Larkum and Weyrauch, 1977), a much higher amount of Chl a has been shown to be in the RCII complex. Possibly some antenna Chla is not fluorescent, so remaining unaccounted for in fluorescence studies. This Chl a is perhaps a bridge, as suggested by Larkum and Weyrauch (1977). The present evidence on Chl a-phycobiliprotein interactions is summarized as follows: (i) PBS pass on absorbed light energy, by inductive resonance or exciton transfer (Section IX.A), via APC, to the Chl a associated with PSII, from there it is distributed to RCI and RCII; (ii) evidence suggests that some PBS are associated with PSI (Pullin et al., 1979; Peterson et al., 1981) but most of the present evidence indicates association of PBS with PSII only; (iii) the amount of energy redistributed to PSI from PSII is variable and is dependent on the type of irradiance regime under which the plants are grown and the type of material (see Section 1X.B); (iv) with Crytophyta, where phycobiliproteins occur within the intra-thylakoid space and are not aggregated into PBS, less is known of the molecular interactions. In these algae light absorbed by the biliproteins may pass directly to the Chl a of PSII, or perhaps via Chl c2 to Chl a (MacColl and Berm, 1978; Lichtle et al., 1980). C. CAROTENOIItPROTEIN COMPLEXES

1 . Peridinin-Chl a Protein Peridinin, an allenicxanthophyll (Strain et al., 1971; Section V1.B) is the major carotenoid of Dinoflagellata. Schutt as early as 1890 suggested that peridinin is conjugated to a protein, but only recently has the existence of a watersoluble peridinin-Chl a protein (PCP) been demonstrated (Bode and Hastings, 1963; Haidak et al., 1966; Haxo et al., 1976; Prezelin and Haxo, 1976; Siegelman et al., 1977) and this PCP is singular amongst the carotenoChl proteins in being water-soluble. Peridinin in ethanolic solution has a broad absorption maximum at 470 nm but in the pigment-protein this maximum is shifted to 478 nm (Fig. 34) with a slight shoulder around 525 nm. At 77°K the shoulder is sharpened. This shoulder was also seen in the action spectrum of photosynthesis by Prezelin et al. (1976) which led them to suggest


1I9

that the interaction of PCP with the thylakoid membrane in vivo enhances the absorption of the 525 nm shoulder. Thus PCP is efficient at harvesting blue and green light, 470-560 nm, but less so in harvesting violet and near blue light, 400-470nm (Prezelin el al., 1976). It is notable that the major absorption maximum of PCP (478 nm) corresponds closely to the wavelength of light penetrating deepest in clear ocean waters (465480 nm; Section HI).

L

1

0-6

0.4 n SI

a

0.2

0

300

400

500

600

700

Wavelength (nm )

Fig. 34. Absorption spectrum of the water-soluble peridinin-Chl a protein (PCP) from Glenodinium sp. (Dinoflagellata). (Redrawn from Prezelin and Haxo, 1976.)

Freeze-thaw treatment or sonication releases up to 65 per cent of the total peridinin and 5-25 per cent of the total Chl a as pigment-protein from cells of Glenodinium sp., Gonyaulax polyedra and Amphidinium carterae (Haxo et al., 1976; Prezelin and Haxo, 1976; Prezelin and Sweeney, 1978). With Amphidinium carterae a single polypeptide was found of 3 1.8 KD (Haxo et al., 1976). Six types were distinguished using isoelectric focusing, although 90 per cent of the material was attributable to a type with a PI of 7.5. The peridininChl a protein and the major isoelectric species were enriched in alanine. Each pigment-protein molecule had a M, of 39 KD and contained 9 peridinin and 2 Chl a chromophores non-covalently bound to the protein. Somewhat similar results were obtained for Glenodiniumsp. and Conyaulax polyedra (Prezelin and Haxo, 1976). Whereas G . polyedra PCP gave a single polypeptide with M, of 32 KD, Glenodinium sp. gave subunits of 15.5 KD after treatment with SDS. The ratio of peridinin to Chla was about 4 : 1, although because of lack of a precise protein analysis it could not be decided whether there were 8 peridinins and 2 Chl a molecules in each molecule of protein, or possibly 9 : 2 as in Amphidinium carterae. In the above studies fluorescence analysis showed an effective transfer of

120


energy from peridinin to Chl a (fluorescence emission major peaks at 672 nm, 709 nm and 733 nm, at 77°K). Song et al. (1976) and Koka and Song (1977) have investigated the fluorescence and circular dichroism of PCP, and have proposed a model in which two peridinin dimers are aligned to allow efficient transfer of energy from the peridinin to Chl a despite the fact that peridinin has an unfavourably short excited state lifetime, to 10-'3s (Fig. 45; Section IX). Two important points remain unresolved concerning peridinin and PCP: firstly, the nature of its placement in the thylakoid membrane and secondly, the role of peridinin in other pigment-protein complexes. It is assumed that PCP is associated with the thylakoid membrane even though it is a watersoluble pigment protein complex in vitro. The lamellae of Dinoflagellata are grouped in threes with little space between; their thickness is comparatively large (about 24nm; Dodge, 1968). Koka and Song (1977) judged from the binding of hydrophobic probes such as anilinonaphthalene sulphate, that the surface of the protein was highly polar. Consequently this pigment could be loosely bound to the thylakoid membrane, but specifically oriented so as to provide efficient transfer of energy to PSI and PSII. Prezelin and Sweeney (1978) came to the conclusion that all the peridinin in Gonyaulax polyedra was in the form of PCP on the basis that this complex accounted for two-thirds of the total peridinin in an extraction in which the remaining third of the total was membrane-bound. In other Dinoflagellata much smaller amounts of the peridinin can be extracted in the form of this complex (Prezelin and Haxo, 1976) and in Amphidinium carterae (PY-I) whose total peridinin content was high none of the complex could be extracted. This finding may mean that the amino acid composition or arrangement of some complexes results in the exposed protein having a less hydrophilic surface causing reduced extractability. Alternatively some of the peridinin may be in other pigment-protein complexes. Pertinent to this conjecture is the finding of two bands containing peridinin and Chl a on SDS-PAGE gels from Glenodinium sp. (Boczar et al., 1980; Prezelin and Boczar, 1981). These presumably represent strongly membrane-bound peridinin, but possibly still in the form of the peridinin-Chl a protein. 2. Chlorophyll c and Fucoxanthin-Containing Complexes (a) Introduction. The Chromophyta, which encompasses the macrobenthic and micro-benthic algae, the many forms of marine phytoplankton and the fresh water alga Vaucheria and other xanthophytes all containing Chl c, were largely ignored by photosynthesis research workers before the mid-l970s, except by the photophysiologists Haxo and Blinks (1950) and Goedheer (1970), and the biochemists Allen (1966) and Jeffrey (1968). Yet the Chromophyta are responsible for a substantial amount of the world's photosynthesis and are the predominant photosynthesizers in the marine


121

area. In themselves they provide a remarkable solution to the problem of achieving maximum collection of light in the inshore waters, where rapid light attenuation and spectral restriction occurs with depth (see Section 111). On an evolutionary level, the point of divergence of the Chromophyta from the other major line of algae, the Chlorophyta, which also have their light harvesting pigments organized into intrinsic protein complexes, provides an important challenge to the systematist (Anderson and Barrett, 1979; Section XII). (b) Distribution of chls c1 and c2. Chl c, and Chl c2 occur in Phaeophyta, Bacillariophyta, Chrysophyta, Haptophyta, Xanthophyta, and in Dinoflagellata that have fucoxanthin as an photoaccessory carotenoid, but Chl c1 is absent from peridinin-containing dinoflagellates (Jeffrey, 1976, 1980). In Gyrodinium (Dinoflagellata) Chl c occurs with a derivative of fucoxanthin (Section V1.B). Outside of the Chromophyta Chlc is found only in Cryptophyta, sometimes included with Chromophyta (Whittaker and Margulis, 1978), where Chl c2 is a minor accessory pigment bound to an intrinsic protein (Jeffrey, 1972; Lichtle et al., 1980; Ingram and Hiller, 1983),and represents an evolutionary minor mystery (Section XII). Jeffrey has established the distribution of the Chls c in 86 species of phytoplankton (Jeffrey, 1976; Jeffrey et al., 1975). Quantitative relationships of Chls c, and c2 for several phyla of Chromophyta algae are given by Jeffrey (1969). Barrett and Anderson (1 980) determined the Chla/c ratios for five different orders of Phaeophyta from coastal waters of south-eastern Australia: the range of Chl a/c was 3.44.0, except for the surface-exposed Phyllosporum which had a Chl alc ratio of 6-9. The molar ratio of Chl cJc, was around 3.2 in these seaweeds, determined with a sensitive fluorescence method at 77°K. In Phaeophyta chloroplasts about 75-80 per cent of the Chl c2 is associated with the major Chl a-c2fucoxanthin-protein complex of PSII, while the entire Chl c , is in a minor complex together with the remaining 20 per cent of the Chl c2 in a 1 : 1 ratio. This complex appears to be associated with PSI (Barrett and Anderson, 1980; Barrett and Thorne, 1981; Barrett, in preparation). The Chlc content of Chromophyta has been reported to increase under low light conditions such as in caves or in deeper waters. Titlyanov and Lee (1 978) found the increase to be less than that for the Chl a of the benthic algae investigated. However for Ascophyllum nodosum and Fucus vesiculosus the Chl alc ratios were constant at 5 between 0 and 4 m depth (Ramus et al., 1977). The fucoxanthin content of these algae also increases with depth. With A . nodosum and F. vesiculosus the fucoxanthinlchl a ratios decreased with depth (Ramus et al., 1977) but for Laminaria cichorioides and Chordafilum the converse was the case (Titlyanov and Lee, 1978). The increase in the fucoxanthin/Chl a ratio with these two algae may be attributable to the deeper water conditions used by Titlyanov and Lee. With the dinoflagellate Glenodinium, Prezelin (1976) found that the Chlc content of the cells stayed constant over a wide range of growth irradiance, but in contrast the content of Chl a and peridinin increased so that

122


the Chl a/c ratios rose from 0.83 under low light conditions to 1.33under high light. In experiments with the neritic diatom Skeletonema costatum grown under different intensities of light, the synthesis of Chl c increased considerably when the light was at its lowest intensity; the Chl a/c ratio dropped from 5.6 to 1.9 at the same time as there was a twofold increase in the Chl a/P700 ratio (Falkowski and Owens, 1980). In macroalgae variations of pigment content can occur along the thallus. In Macrocystis pyrifera the content of Chl a, Chl c and fucoxanthin increased from the apical meristem, reaching a maximum 2-3m below the apex; the pigment ratios remained relatively constant (Wheeler, 1980). ( c ) Chl c and carotenoid protein-complexes of micro-chromophytes. Early attempts to separate the light harvesting complexes from other Chl-proteins in extracts of brown algae and pigment related phytoplankton, although yielding the P-700 complex, did not give a complex analogous to the Chl a/bprotein complex of green plants (Brown et al., 1974; Prezelin, 1976). This is a measure of the difficulties imposed with brown macro-algae and the related Chl c2-phytoplankton. Milder conditions of electrophoresis must be used but these inhibit the breaking of protein-protein interactions within the supracomplexes. Boczar et al. (1980) with Glenodinium replaced SDS with Deriphat 160-C in PAGE, and limited electrophoresis to 30 mins. However, it was necessary to use SDS as a solubilizing agent. A series of four pigmented bands were obtained, containing Chls a and c2 in different amounts. One of the bands, which had a major polypeptide of M, 20 KD, had a Chl c2/aratio of about 4.8. The absorption maxima were at 636,585 and 453 nm, close to the A,,, of the Chl c2-polypeptide isolated free of Chl a and carotenoid from Acrocarpia paniculata (Barrett and Thorne, 1981) (Fig. 35). From the spectra of the Chl c2/a-protein small amounts of peridinin appear to be associated with this Glenodinium Chl c,/a-protein fraction. As other Chl a-containing proteins were obtained having lesser amounts of Chl c2, but increasing amounts of peridinin, it is possible that the Chl c,/a-protein of ratio 4-8 is a fragment of a supra-polypigment protein complex. Other fractions contained the bulk of the peridinin in association with Chl a mainly (Section XI1I.C). No supracomplex containing Chl a-Chl c2 and peridinin-protein analogous to the Chl a-Chl c2 fucoxanthin complex of brown macro-algae was found. Three Chl-protein complexes have been obtained from the Prymnesiophyta, Pavlova lutheri(Droop), using digitonin for solubilization and PAGE or density gradient centrifugation (Romeo, 1981). An apple-green fraction had spectral and biochemical characteristics of the P-700-Chl a-protein complex. An orange-brown complex which contained most of the fucoxanthin had fluorescence characteristics at 77°K which compared closely to those for the Chl a/c2-fucoxanthin-proteincomplex isolated from Acrocarpia paniculata (Barrett and Anderson, 1980). A green complex, intermediate to the

123


other two on gels and density gradients, had strong fluorescence emission at 685 nm and may contain the PSII reaction centre.

90

I

8ol\

_-

70

'0 Wavelength ( n m)

Fig. 35. Absorption spectrum at 20°C of RClI core complex from the brown alga Acrocarpia paniculuta.

A Chl a + c,-fucoxanthin-protein complex has been isolated from cells of the diatom, Phaeodactylum tricornutum using a French press and chromatography on Ultragel, then DEAE-Sephadex (Holdsworth and Arshad, 1977). This pigment-complex had a M, of 850KD and probably consisted of 40 subunits. The macro-complex contained 40 mol of Chl a, 20 mol of Chl c,, 20 mol of fucoxanthin, 8 g-atoms of Cu and between 0.6 and 2.0 g atoms of Mn. Detergents split the macro-complex into subunits of M,-25 KD. Photoenzymic and EPR properties of the complex indicate that it is a component of Phaeodactylum PSII. (d) Phaeophyta. Brown macro-algae (Phaeophyta) are probably the dominant algal class in terms of annual photosynthetic biomass per mz, being comparable to rain forests (Mann, 1973). These algae have evolved lightharvesting and structural features (Section V.B and C ) enabling them to flourish in a wide range of environments, yet few investigations into the molecular structure and organization of the photosynthetic pigment assemblies of brown algae have been made. This is largely because of the refractory nature of their thalli. The high cellular osmolarity, 1.8 Osmol Kg -' for Fucus serratus (Nordhorn et al., 1976), requires preparative buffers of matching osmolarity, and the massive muco-polysaccharide secretion necessitates a high ionic concentration during the isolation of chloroplasts, which are difficult to disrupt, probably because of polysaccharides intimately involved in the thylakoid structure (see below).

124


Triton X-100 has been used to disrupt the thylakoids and various fractions have been obtained containing the light harvesting complexes, and RCI and RCII complexes (see Section VILA and B). Sucrose density gradient centrifugation of Triton X-100 extracts of Ecklonia radiata chloroplasts yielded three major fractions and a minor one (Barrett and Anderson, 1977). The major fraction of the highest density contained the P-700-Chl a-protein complex, the next densest a Chl a-Chl c2-fucoxanthin protein complex and the lightest zone, a Chl a-Chl c2-violaxanthin protein-complex. The minor upper zone was enriched in 8-carotene protein. Kirk (1977b) fractionated Triton X100 extracts of Hormosira into two or three pigment-protein complexes, but the separation was neither as clear cut or as reproducible as by the centrifugation method. SDS extracts of Ecklonia chloroplasts when chromatographed on hydroxylapatite gave the green, modified form of fucoxanthinprotein (A,,, 485 nm instead of 520-540 nm) in the Chl a-Chl c2-fucoxanthin supramolecular complex, but also the complex enriched in P-carotene-protein (Barrett, unpublished). Importantly, fluorescence excitation spectra of the brown Chl a-Chl c2-fucoxanthin protein complex confirmed energy transfer from Chl c2 and fucoxanthin to Chl a (Barrett and Anderson, 1977; Anderson and Barrett, 1979). Improvements in technique coupled with the use of a seaweed with fibrillar thalli, e.g. Acrocarpia paniculata, or thin lamellae e.g. Padina commersonii or Lobophora ( = Pocockiella) variegata, improved yields of chloroplasts and gave superior fractionation. With Acrocarpia and several other species of brown algae the Chl a/c2-fucoxanthin complex was obtained as a heavy sucrose gradient zone, evidently an oligomer of the lighter orange-brown zone obtained from all the brown algae investigated. The properties of the major light-harvesting complexes are given in Table IV. Similar properties were found for the complexes from Ecklonia radiata, Phyllospora comosa, Sargassum sp., Scytothamnus australis, Hormosira banksii, Padina commersonii, Cladostephus spongiosus, Colpomenia sinuosa, Lobophora variegata and Scytosiphon sp.. The absorption and difference absorption spectra of the Chl a-Chl c2fucoxanthin complex reveals that the A,, of Chl c2-protein is at 465 nm, well to the red of the A,, of Chl c2 in solvents; while in the Chl a-Chl c I + c2violaxanthin complex, the combined Chl c I c 2 A,,, is at 457 nm; which may imply that the A,,, of the Chl c,, is at 450 nm or so, since the two Chls c are present in a 1:1 ratio. Fluorescence excitation spectroscopy at 77°C supports these assignments (Fig. 36). The fucoxanthin-protein may exist in two confirmations, or there may be two separate fucoxanthin-proteins in the supracomplex. The absorption spectra of the supracomplex at 77"K, and the difference absorption spectra have broad maxima centred at about 520 nm and 540 nm (Fig. 37). These two components are more distinctly revealed in the fluorescence excitation spectra (Barrett and Anderson, 1980).

+

LIGHT HARVESTING PROCESSES IN ALGAE 0.8

I I 4 6Chlg

3

I -Fwxanthin-Chl g/Chl --ChlQ/Chl El+&/

I

1

I

0.8

~

I

125 1

I

so

~2 protein

-violamnthin-pMii

Wovelength (nm)

Fig. 36. Fluorescence excitation and emission spectra, 77"K, of the brown alga Acrocarpia paniculafa light-harvesting complexes.

-02

661

~

:4 . 0400

450

--

500

__

I

-~

550

600

650

700

750

Wovelength (nm)

Fig. 37. Difference absorption spectra, at 2 0 T , of light-harvesting assemblies of PSII versus PSI of the brown alga Acrocarpia paninclura, demonstrating A,,,,, of the different pigment components.

The existence of two forms of fucoxanthin-protein has been detected also in Fucus serratus chloroplasts in studies of their variable fluorescence (Duval, personal communication). Fluorescence emission spectra at 77°K of A. paniculata chloroplasts excited at 520 and 540 nm (Fig. 40) suggests that part of the 520 nm component in the fucoxanthin complex is associated with PSI, and the 540 nm component mainly with PSII.This conclusion is supported by the finding of a fucoxanthin protein with excitation peak at 540 nm in association with PSII reaction centre

126


isolated using cholate (Barrett and Thorne, 1980). Furthermore, differential extraction of the thylakoids with steroid detergents and LDAO, leaves a residue of fucoxanthin-protein, together with the entire PSI pigment assembly in the lamellae. The european brown seaweeds Fucus serratus and Cystoseira mediterranea have yielded two major light-harvesting complexes by the Triton X-100 gradient-centrifugation method (Berkaloff et al., 1981). The absorption and fluorescence properties of the complexes were generally in accord with those from Ecklonia and Acrocarpia. Curve deconvolution analysis of the absorption band with A,,, 670 nm of Fucus chloroplasts showed that Chl a 668 and Chl a 678 accounted for 36 per cent and 32 per cent respectively of the Chl a present, but curve analysis of the two light-harvesting complexes indicated that only Chla 668 was present. This analysis does not accord with the demonstration of two Chl a components given by the fluorescence spectra. Although it is certain that Chl c2 and fucoxanthin in the PSII supramolecular complex transfer light energy to Chl a, it is not fully established whether the energy transfer is to a common pool of Chla, or whether each pigment-protein complex has its own cluster of antenna Chla. Current evidence however supports the latter case. Firstly, the Chla, Chlc, and fucoxanthin of the supracomplex can be isolated as polypeptide complexes each having a single pigment (Barrett 1978; Barrett and Thorne, 1980) (Figs 38, 39). Secondly, Alberte et al. (1981) have isolated from SDS-extracts of French press-fragmented chloroplasts by SDS-PAGE a Chl a-fucoxanthinprotein complex, with molar ratio of Chl a to fucoxanthin of 1 : 5 , and a Chl a Chl c2-protein complex with a molar ratio of Chl a/Chl c2 of 1 : 2 ; neither

Wavelength ( nm)

Fig. 38. Absorption spectrum, at 20"C, of fucoxanthin-polypeptideisolated from PSII lightharvesting supracomplexes of the brown alga Acrocarpia paniculara.


127

binary complex was free of the third pigment. Energy transfer to Chl a from the accompanying pigment was observed at 29°C but the proportion of the pigments effectively coupled to Chla cannot be assessed because of the absence of absorption spectra. Thirdly, the Chl a/c,-fucoxanthin complex (molar ratio 2 : 1 : 2), obtained by digitonin extraction could be fractionated by SDS-PAGE, into a fucoxanthin-Chl a complex (molar ratio, 6 : 1) and Chl aChl c2 fractions differing in Chl c2-content (Barrett, unpublished). A minor Chl a-Chl c,-protein-complex retaining some fucoxanthin, was also obtained.

\.:zxI

L;

--Ap 1 -

350

550

450

650

Wavelength (nm)

Fig 39 Absorptlon spectrum, at 20 C, o f Chlc,-polypeptlde Isolated from PSI1 Iightharvesting supracomplex of the brown alga Acrocarpra panrculafa

C

.Q

e

60-

=d

20-

L

r

'400

440

400

520

560

620 660 700 740 700

Wavelength (nm 1

Fig. 40. Fluorescence excitation spectra, at 77"K,of chloroplasts of the brown alga Acrocurpiu puniculuru, in glycerol 66%. Emission measured, at 735 nm-; at 694 nm- - - - -

128


This has A,,, attributable to Chl c2 at 472 nm and 650 nm instead of the usual A,,, of 465 nm and 642 nm. The A472nm-A,,, nm-complex appears to be a device for extending light harvesting into the green spectral region. The evidence discussed for the direct transfer of energy from Chl c2 and fucoxanthin to separate pools of Chl a does not exclude the possibility that, because of the absorption overlap in the near-red region, some transfer of energy from fucoxanthin to Chl a occurs via Chl c2 in the supracomplex. (e) Organization of the photosystems within the chloroplast. The organization of the thylakoids in Phaeophyta differs markedly from that in Chlorophyta (see Section V). The thylakoids largely occupy the brown algal chloroplasts with interconnected sets of three appressed lamellae (Section V.E.3). A basic difference in the molecular structure of the thylakoids is shown by their resistance to dissolution by detergents. Electron microscopy reveals that multiple extraction of chloroplasts with 1 per cent Triton X-100 leaves a residue which consists of chains of vesiculate residues which conform in geometry to the contours of the triple appressed lamellae (Fig. 13) from which they arise (Barrett and Goodchild, in preparation). A less extensive loss of the triple-lamellar structures is observed when chloroplasts are extensively extracted with digitonin. Colorimetric analysis and chromatography of the products of methanolysis of pigment protein complexes, chloroplasts and thylakoid residues after detergent extraction point to polysaccharides having an intimate role in the thylakoids. The thylakoids are probably stabilized by a skein of polysaccharides against the weakening effect of high ionic strength. Little is known of the lipid composition of the Phaeophyta, but differences have been found in the proportions of mono-, di- and tri- enoic fatty acids present in thalli compared to green plant tissues (Jamieson and Reid, 1972) and similarly for chloroplasts (Barrett and Bishop, unpublished), The studies of Berkaloff and Duval, (1977,1981) on the effect of cations on chloroplasts of Fucus serratus are of considerable significance with regard to structural and chemical differences noted between brown and green chloroplasts. Firstly, the fluorescence induction, FvDt-MU, of Fucus is much weaker than in spinach requiring concentrations of Na >>200mM, whereas Mg does have a noticeable effect on F v ~ M uthough , less than for spinach. Secondly, the relative effect of Mg++ on the rate of reduction of DCIP is greater in Fucus than in spinach chloroplasts, especially in high levels of light. These properties of the brown algal chloroplasts raise questions concerning the location of the two photosynthetic assemblies. Several pieces of evidence point to more of PSII being located at, or near, the surface of the thylakoids, while PSI is more embedded in the thylakoid. Firstly, Triton X-100 at a concentration of 0.02 per cent at pH 8.0 selectively removes Chl cz- and fucoxanthin-polypeptides from the thylakoids. Secondly, 1 per cent cholate, pH 8.0, removes PSII reaction centre complex, followed by Chl cZ-, then +

++


129

fucoxanthin polypeptides from the thylakoids. Further treatment with deoxycholate removes the remainder of the PSII pigment assembly, leaving P-700-Chl a complex and its associated Chl u-Chl c, + c2-violaxanthin complex in residual lamellae from which it can be released by Triton X-100 (Barrett and Thorne, 1980). Thirdly, digitonin removes the entire PSII reaction centre and light-harvesting complex intact (Barrett, in preparation), while passage of Phaeophyta chloroplasts through the Yeda Press yields fragments of thylakoids which are composed of RCII and associated lightharvesting complexes. So far it has not been possible to establish to what extent lateral heterogeneity (Section 1X.B)applies to the organization of PSI and PSII in the Phaeophyta. Application of the aqueous two phase polymer partition fractionation (Andersson and Anderson, 1980) is fraught with problems peculiar to the Phaeophyta, but preliminary investigations have provided some partitioning of the pigment complexes (Barrett and Andersson, unpublished). A freeze-fracture study of algae from six Divisions shows that PSII particles are aggregated in stacked regions of the thylakoid membrane (Dwarte and Vesk, 1982), suggesting the occurrence of lateral heterogeneity (Section 1X.B) in these algae. A schema of the light-harvesting pigment-protein assemblies of PSI and PSII together with their relevant reaction centres, in A . paniculutu, is shown in Fig. 41. ~

IX. PRINCIPLES O F LIGHT HARVESTING A. QUANTUM CHEMISTRY A N D TRANSFER OF EXCITATION ENERGY

1. Introduction Photosynthetic pigments are arranged in thylakoid membranes in a number of pigment-protein arrays (Section VII and VIII) which provide a means of transferring absorbed light energy to RCI and RCII with high efficiency. Apart from the photochemically reactive P-700 and P-680 molecules, each reaction centre is closely associated with a number of antenna pigments comprising Chlu and a limited number of carotenoids (Section VII and V1II.A). Further, there are the light harvesting complexes which contain a variety of other photoaccessory pigments, and these also communicate efficiently with the reaction centres. Two important questions arise (i) what is the mechanism for energy transfer? and (ii) how are the pigment-proteins arranged? More specifically, do the light harvesting complexes connect to the reaction centres via the antenna pigments? A quantum of light energy absorbed by a pigment molecule, either an antenna or light-harvesting pigment, moves as an excited state (exciton), from pigment molecule to pigment molecule until it is trapped by a reaction centre

-.?Ig. 41. Schema of spalial arrangemen1 of pigment-protein complexes in RCI and RCIl supracomplexes of thc brown alga Acrocarpiu puniculara


131

(or until lost by fluorescence or thermal decay). Two mechanisms of energy transfer are possible in photosynthetic pigment systems (cf. Knox, 1975); Forster resonant energy transfer (Forster, 1946, 1965) and exciton migration (Davidov, 1962).

2. Forster Resonance Transfer Resonance energy transfer as proposed by Forster (1946, 1965) involves a weak interaction between donor and acceptor molecules in which the transfer time is long in comparison with the time for intramolecular vibrational relaxation. The excited donor molecule relaxes to the vibrational ground state before transferring its energy to the acceptor. After transfer the acceptor molecule also relaxes and is no longer in resonance with the donor. The process is therefore essentially unidirectional. For resonance to take place the donor and acceptor must be closer than 100 A, must h a w transition dipoles which are aligned and must show overlap between the fluorescence band of the donor and absorption band of the acceptor. In Forster resonance transfer the molecules concerned do not show vibrational resonance: the process is one of incoherent energy transfer. Some loss of energy is incurred and the transfer is therefore from pigments with shorter to those with longer A,,,. The rate of transfer is inversely related to the sixth power of the distance between donor and acceptor molecules (which limits transfer to about 100 A distance). 3. Exciton Migration For exciton migration the excited state is considered to be "delocalized" within a group of pigment molecules (Davidov, 1962, Knox, 1975). The relaxation time is of the same order as the transfer time (10 -I2s). This is called coherent energy transfer but because no single pair of donor-acceptor molecules can ever be identified the term exciton migration is preferred (Knox, 1975). It is characterized by bidirectional movement and little loss of energy. However it does require a close proximity and strong interaction between molecules of the pigment array (Shipman, 1980). 4 . Evidence on the Type o j Energy Transfer in Photosynthetic Systems Forster resonance energy transfer and exciton migration are extremes of a range of interactions which may show characteristics of both types under intermediate conditions. They are also concepts derived from different mathematical descriptions: Forster resonance deals only with the interaction of two molecules whereas the exciton concept deals with the probability of locating the excited state in any one of a number ofclosely interacting pigment molecules (Knox, 1975). It is only very recently that evidence for the critical evaluation of the two approaches has been available as a result of the development of picosecond

132


spectroscopy (Breton and Geacintov, 1980) and circular dichroism studies (Sauer, 1975). However, the situation is still largely unresolved although there is growing support for limited exciton migration in some systems (Knox, 1975, 1977). Other evidence has come from X-ray crystallography or electron diffraction. These approaches have established the three dimensional structure of three important light-harvesting proteins, BChla - protein at 2.8 A resolution (Fenna and Matthews, 1976, 1979) as well as C-Phycocyanin and BPhycoerythrin at 5 A resolution (Fisher et al., 1980),all of which are peripheral membrane proteins. In addition the electron diffraction study of the purple membrane of Halobacrerium rubrum has yielded a 7 A resolution map of the light-activated proton-pumping protein, bacteriorhodopsin (Henderson and Unwin, 1975). The four proteins have 3-C3 symmetry which gives an optimal molecular orientation for absorption of light and energy transfer (Fisher et al., 1980). 5 . BChl-Protein

The X-ray crystallographic structure determination of BChl a-protein from Prosthecochloris aestuari (Fenna and Matthews, 1975,1979) (Fig. 42) shows

Fig. 42. The structure of BChl-protein from Prosthecochloris aesruari determined by X-ray crystallography at 2.8 A resolution. (Redrawn from Fenna and Matthews, 1977.)


133

seven BChl a molecules linked non-covalently to protein, some via the magnesium of the BChl a to the polypeptide backbone. Further stabilization of the orientation of the BChl a molecules within the protein is by interaction of the ring carbonyls to the magnesium of an adjacent BChl a and through hydrophobic bonds between the phytyl chains and protein. The BChl a molecules are 9-14 A from their nearest neighbour BChl a . 6. Phycobilisomes The mechanism of excitation transfer in phycobilisomes is also unresolved. Here C D studies confirm the existence of exciton interactions (Pecci and Fujimori, 1969; Glazer et al., 1973; Canaani and Gantt, 1980). However, the arrangement of phycobiliproteins in the order PE+PC+APC-+Chl a (Section VIII) could just as well fit a Forster resonance energy transfer mechanism because of the presence of overlapping fluorescence and absorption bands (see e.g. Teale and Dale, 1970; Gannt, 1975). Grabowski and Gantt (1978a,b) analysed the evidence for a Forster-type mechanism and found it to be generally consistent with the facts. Many of the assumptions used were based on the properties of isolated phycobiliproteins and not of the intact phycobilisomes. The structure and interaction of subunits in the phycobilisome may be different from that of isolated subunits (Morschel et al., 1980a,b; Fisher et af., 1980; Glazer, 1981). The study of picosecond fluorescence kinetics in phycobilisomes of Porphyridium cruentum (Rhodophyta) (Porter et al., 1978; Searle et al., 1978) discussed in Section VII1.B supported the above sequence of phycobiliproteins, but found that the transfer times were an order of magnitude faster than those predicted by Grabowski and Gantt (1978b). Thus a strong interaction of the exciton type is indicated. Evidence for such strong exciton interactions has been obtained for cryptophyte PC (Kobayashi er al., 1979; Jung et al., 1980) and cyanobacterial APCI and B (Canaani and Gantt, 1980). However, as pointed out by Canaani and Gantt (1980), even within one molecule of APC there may be both exciton interaction and Forster resonance depending on the distance and coupling between the various chromophores. The model of the phycobilisome which is emerging (Section IX, Fig. 28) also confirms that strong interaction exists between chromophores: each rod of a or hexamers phycobilisome is composed of discs which are trimers (a& (a6p6)of PE, PC or APC. The exact alignment of subunits is not known but a convincing view is that tl and jl subunits alternate within the disc and lie in a close-packed overlapping configuration along the rod (Fig. 43). Teale and Dale (1970) and Dale and Teale (1970) first made the suggestion that the chromophores on the tl and p subunits might serve different roles. They obtained evidence from fluorescence polarization to suggest the presence of two different types of chromophores in a single protein. designated “sensitizing” (“s”) and “fluorescing” (‘T’)

134


From the work of Glazer et al. (1973) it is tempting to assign the ‘‘s” chromophores to the p subunit and the “f’ chromophore to the a subunit. However that evidence was not conclusive (Glazer, 1981) nor is it supported by the work of Doukas et al. (1981) and Wong et al. (1981). Zuber (1978) on the basis of amino acid sequencing and studies of tertiary structure of C-PC and APC has put forward a model of subunit and chromophore arrangement (Fig. 43). Although the evidence is far from conclusive the model does provide a working hypothesis. In C-PC the chromophores are between 15 and 40 A apart (cf. average of 25.5. A,Dale and Teale. 1970). In APC the chromophores are even closer together. Thus the structure suggests the strong feasibility of exciton migration. In B- or R-PE there IS a greater density ofchromophores both on the a and p subunits (Table 3) and by the addition of the y subunit, with 4 chromophores, located in the centre of the disc (Fisher et al., 1980). Here the average chromophorechromophore distance may be less and the exciton interaction consequently greater. It is therefore possible to envisage the existence of exciton migration between discs of the same kind. Between the discs of one type and another (viz. PE-PC or PC-APC) it is possible that a Forster type of transfer occurs. Such an arrangement would give a rapid and overall unidirectional movement of excitation energy down the phycobiliprotein rod to Chl a. It will also be noted that the arrangement of chromophores suggested by Zuber (1978) (Fig. 43) fits well with the models of PBS structure (Fig. 28) put forward by Koller et al. (1 978) and Bryant et al. (1 979), where the APC discs are end-on to the rods of PC and PE. 7. Intrinsic Membrane Chl a-Antenna Proteins The structure of intrinsic Chl a-protein complexes of Cyanobacteria or C-Phycocyanin

Hexamer

Fig. 43. A possible arrangement of phycobiliprotein tl and /Isubunits and chromophore orientation in phycobilisomes. (Redrawn from Zuber, 1978.)


135

eukaryotes have not been established. Some evidence of the mechanism of energy transfer can be adduced from fluorescence lifetimes (Junge, 1977) and from circular dichroism (Sauer, 1975). Fluorescence lifetimes are 1.9 ns for PSI1 and 30 ps for PSI (Borisov and Il’ina, 1973). Assuming a random walk process amongst 300 Chl molecules in an antenna unit of PSI, Borisov and Il’ina (1973) estimated that it would take an average of 125 steps before the excitation reached the reaction centre. Thus each step could be no longer than 0.25 ps to complete effectively with the 30 ps fluorescence lifetime of this photosystem. Such a fast rate indicates an exciton mechanism (Junge, 1977), although the conditions could also be satisfied by exciton interaction between a number of closely-spaced Chl molecules in the same sub-complex and a Forster type of transfer between subcomplexes. Sauer (1975) from circular dishroism evidence supports the latter model which he called the “pebble-mosaic model”. 8. Light-Harvesting Pigment-Proteins Located in Thylakoid Membranes Our knowledge of the molecular topology and the finer interaction between various light-harvesting pigment-protein species located in membranes cannot compare with that of the BChl-protein or the phycobiliproteins. Consequently less firm conclusions can be made about the mechanism of excitation energy transfer within, for example, the light-harvesting Chl a/b protein (LHCP), the Chl a + c2-fucoxanthin-protein complex and the watersoluble peridinin-Chl a-protein complex which, in vivo, must be membrane bound (Section VII1.C). Nevertheless, some insights can be obtained from fluorescence and C D studies. From such studies Van Metter (1977) and Knox and Van Metter (1979) put forward a model of the arrangement of Chl in LHCP in which there are three closely-associated Chl b molecules which exhibit exciton interaction and three Chl a molecules to which the excitation energy is transferred. The Chl a weakly interact with one another and with Chl a molecules of Chl a-proteins (Fig. 44). Using similar techniques, Song et al. (1976) deduced a model for the

Fig. 44. Proposed model for the arrangement of Chla and Chlh in the light-harvesting Chl a/b-protein (LHCP). (From Van Metter, 1977.)

136


pigment arrangement in the peridinin-Chl a-protein (PCP) in which there are probably 4 peridinin molecules and 2 Chla molecules (Section VIII). According to this model there are two pairs of peridinin molecules, each pair being separated by a distance of 12 A and further separated from the other pair by a Chl a molecule (Fig. 45). Exciton interaction between the pairs of peridinin molecules lengthens the fluorescence lifetime of peridinin over that observed in organic solvents. This prolonged fluorescence and the topology of the pigment molecules change the probability of energy transfer to the Chl a

P

Protein

Fig. 45. Proposed arrangement of Chl a and peridinin molecules in the water soluble Chl aperidinin proteincomplex of dinoflagellates. (From Song et d., 1976.)

molecule from zero in isolation to 100 per cent in the complex. The fluorescence lifetime of the Glenodinium PCP (Section VII1.C. I ) is Se53f0.14ns, a value which is similar to that of Chla of PSII (Section IX.A.7).The PCP complex is thought to have a hydrophobic crevice in which the pigment molecules are located and liganding of pigments to tryptophan and tyrosine residues may be involved (Koka and Song, 1977). 9. Interconnection of Core Antenna Chl a Units There has been speculation but little firm evidence on the arrangement of reaction centres and their antenna Chl a-protein subcomplexes in the thylakoid membrane. Two extreme models have been proposed-the puddle and lake models (Section V.G.l, Fig. 15). Studies of PSI1 have provided much of the evidence. It has often been assumed that cooperativity is needed between several reaction centres of PSII because the photolysis of water requires the withdrawal of four reducing


137

equivalents in close succession (Williams, 1977). Joliot and Joliot (1964) and Kok et al. (1970), from oxygen yield experiments using modulated or flashing light, have suggested a four-step reaction. The model which has been generally accepted is that of Kok et af. (1970) and is summarized as follows:

so hv --so*-s \

I

hv -s

1

*-s

hv hv 2-S**-S3-S3*-(S4) /

There is a requirement in this scheme for four photochemical events in close succession, to prevent back reactions which reduce the efficiency of the process (Diner and Joliot, 1977; Radmer and Kok, 1977). The photochemical steps can be carried out by a single reaction centre or by the cooperation of more than one centre. Present evidence does not support cooperation (BougesBocquet, 1980). Joliot and Joliot (1964, 1968) working with Chlorella vulgaris (Chlorophyta) proposed a connection between antenna Chls belonging to different reaction centres. From fluorescence induction kinetics and oxygen yields in modulated light they concluded that excitation energy must be able to migrate from PSII units with closed traps to neighbouring units with open traps. This proposal has received wide support (cf. Williams, 1977). In Chlorella, at least four units appear to be interconnected (Dubertret and Joliot, 1974; Diner and Wollman, 1979a,b), while in a mutant of Chlamydonionas (Chlorophyta) at least three units were shown to be interconnected (Joliot et al., 1973).Further support comes from theoretical considerations (Palliotin, 1976a,b) and picosecond fluorescence spectroscopy (Breton and Geacintov, 1980). The identity of the intercommunicating units has still to be resolved. Results from higher plants (Akoyunoglou, 1977) and algae (Diner and Wollman, 1979a,b) suggest that a core unit containing only 30 to 40 antenna Chla molecules is sufficient to act as a unit and this may well have its physical expression in the small EF freeze-fracture particle of 80-100 A diameter (Subsection B). In Cyanidium caldarium (Rhodophyta) there is evidence (Diner and Wollman, 1979a; Diner, 1979; Wollman, 1979) for greater aggregation of EF particles and greater interconnection of PSII in the phycocyanin-less mutant compared to the wild type. This suggests that in Cyanobacteria and Rhodophyta the PBS impose spatial constraints on PSII units which lower the cooperativity of units (see Section 1X.B for further discussion). In those plants containing Chl b it appears that increase in size of the PSII unit is at least partly due to the addition of LHCP to the PSII complex, or close association with it (Armond et al., 1977). Two possibilities for intercommunication of PSII units exist in this situation. Either four or

138


more complexes are grouped together along with LHCP complexes, or there is a central PSII complex surrounded by LHCP complexes which act as the channel of intercommunication with neighbouring PSII complexes. The work of Wollman et al. (1980) on wild type and mutants of Chlamydomonas reinhardtii (Chlorophyta) supports the latter proposal. According to this work communication is most likely by means of protein complexes that correspond with PF freeze-fracture particles (probably LHCP complexes-see Section 1X.B). Intercommunication between complexes of PSI is also a possibility although it has less experimental support (Williams, 1977). B. STRUCTURE AND FUNCTION

1 . Introduction Physical studies have shown (Subsection A) that in photosynthetic pigment arrays the chromophores are specificallyarranged to be in close proximity and to interact with one another to give directional and rapid transfer of excitation energy to the reaction centres. Two important points remain: firstly, the arrangement of the photosystems in the thylakoid membranes; secondly, the distribution of excitation energy between PSI and PSII.

2. Freeze-fracture Particle Evidence Much of the evidence on the arrangement of pigment-protein complexes in thylakoid membranes is based on intra-membrane particles revealed by freeze-fracturing techniques of electron microscopy (Boardman et af., 1978; Staehelin et al., 1978). Early freeze fracture studies indicated that small particles (70-80 A diameter) could be assigned to PSI complexes, since they occurred on stroma lamellae which exhibited only PSI activity, and that large particles (140-180 A diameter) could be assigned to PSII complexes, as they occurred in appressed thylakoids enriched in PSII activity (Boardman et af., 1978). More recent studies (Armond et af.,1977; Staehelin et a f . ,1978; Miller, 1976; Simpson, 1979; Wollman, 1979) have shown that PSII activity can be found in thylakoid membranes where no large particles are present. Furthermore, other protein complexes such as LHCP, the cytochrome f-b, complex and the F, complex of the ATP synthase may give rise to small particles (e.g. Simpson, 1979). Simpson (1979) has proposed a model (Fig. 46)in which large particles on the exoplasmic fracture face (EF) are formed of PSII complex and LHCP complexes. These are found in appressed regions of thylakoids. The smaller particles, also found in appressed regions but on the protoplasmic fracture face (PF), may be formed to some extent also of LHCP complexes. In non-appressed thylakoid membranes the small particles which are found in both E F and P F may represent PSI complex or other complexes. Armond et af. (1977) obtained strong evidence that the LHCP complex is a part of the large E F particle by following changes in size of the E F particle from 80 A to


139

A

164 and correlating this change with the formation of LHCP in greening pea leaves. Similar evidence has since been obtained for Euglena gracilis (Euglenophyta) (Dubertret and Lefort-Tran, 1981). In Cyanobacteria and Rhodophyta, where the major light-harvesting complex, the phycobilisome, lies in the stroma space, there are no large EF particles and the PSII complex, with a small antenna unit, can be assigned to a 100 A EF particle (Fig. 33). (Lefort-Tran et al., 1973; Staehelin et al., 1978; Wollman, 1979; Gannt, 1980). Dwarte and Vesk (1982) have found that EF particles (100-160 A) are largely located in appressed regions of thylakoid membranes of several chromophyte algae. In summary the freeze-fracture evidence, taken together with the evidence presented in Section IX.A, supports the hypothesis that, in green algae (including Euglenophyta), in higher plants and possibly in chromophyte algae, PSII units intercommunicate in the lateral plane of the membrane, either by close proximity or via complexes such as LHCP present as small P F particles. An alternative hypothesis, with less experimental support, is that PSII units in appressed systems intercommunicate in the vertical plane from one appressed membrane to the other (Miller, 1976; Arntzeq.1978; Dubertret and Lefort-Tran, 1981). 24 nm

+ k

17nm

3

nn

f

PSI

ESs

24 nm

3

Fig. 46. Proposed arrangement of PSI, PSII and LHCP in the thylakoid membrane as revealed by freeze-fracture electron microscopy. (Simpson, 1979.)

3. Lateral Heterogeneity of the Photosystems and Thylakoid Appression Physical separations have played a major role in elucidating the structure and function of chloroplast membranes (Sane et ul., 1970; Andersson et al., 1978a,b). Using the phase-separation technique of Andersson et al. (1978), Andersson and Anderson (1980) separated appressed thylakoid membranes (appressed membranes from grana stacks) from non-appressed membranes (external membranes of grana stacks or stroma lamellae) and investigated the distribution of PSI and PSII complexes and LHCP complex in spinach chloroplasts. In their model (Fig. 47) there is extreme lateral heterogeneity of

140


0 Photoryrtem 1 complex

$ Coupling I factor

Photosystem 2 complex and light - horverting complex

Fig. 47. Lateral heterogeneity model for PSI, PSI1 and LHCP in appressed and non-appressed thylakoids of green plants. (Redrawn from Andersson and Anderson, 1981.)

the thylakoid membranes with PSII complexes and LHCP restricted largely to the appressed membranes and PSI complex in non-appressed membranes. Taken together with the evidence that the ATP synthase and NADP reductase are also on the non-appressed membranes (Section V.F), this model has important consequences which are discussed by Anderson (1981, 1982), but no clear role has as yet been assigned to the phenomenon of lateral heterogeneity and membrane appression. Anderson (198 1) suggested that restriction of PSII and LHCP complexes to appressed regions may increase the efficiency of PSII; presumably by increasing light-harvesting efficiency, possibly by promoting intercommunication between reaction centres, or by protecting the water-splitting apparatus against backreactions (see Section 1X.A). The fact that the proportion of appressed membranes increases in shade plants (Boardman et al., 1978) is then easily explained, since under these light-limited conditions PSII activity is rate-limiting. In algae, thylakoid appression occurs in all phyla except Cyanobacteria and Rhodophyta. However it is not known whether thylakoid appression is accompanied by lateral heterogeneity in any alga, although this seems certain for Chlorophyta and is also indicated for chromophyte algae from freezefracture particle evidence (Dwarte and Vesk, 1982 and see Fig. 48). The theoretical advantages of lateral heterogeneity -cooperativity of PSII units and control of spillover, put forward by Anderson (1981) are not the only possible advantages of thylakoid appression. Other advantages may be, +

(i) more efficient packing of light-harvesting complexes (Section V.F). (ii) more efficient maintenance of the light-driven proton pump at low irradiance (Subsection D). (iii) increased light-scattering with enhanced absorption of green light (Bialek et al., 1977; Section V.G).


141

(iv) more efficient harvesting of light enriched in green-yellow light deep inside chloroplast stacks by light-harvesting complexes which absorb such light more efficiently and are preferentially located in appressed regions. The major disadvantage of the appressed thylakoid system as it exists in algae today appears to be the lack of a suitable light-harvesting complex that is both located in the membrane and harvests light efficiently throughout the 5W630 nm region.

Fig. 48. Freeze-fracture thylakoids from the chromophyte alga Isochrysis galbana. (Mirrograph by D. Dwarte and M. Vesk, Electron microscope Unit University of Sydney.)

I42


4 . Non-appressed versus Appressed Thylakoid Systems Any discussion of the role of thylakoid appression must be set against the fact that thylakoid appression does not occur in Cyanobacteria and Rhodophyta, whose members survive under a wide range of light environments and are especially successful in extreme shade conditions (Section X.C). Thus appressed thylakoids cannot confer any unique property. Rather the two approaches, appressed thylakoids with membrane-located light harvesting complexes and non-appressed thylakoids with large phycobilisomes located in the stroma space, seem to serve a similar end although employing different mechanisms. The phycobilisome system offers the advantage that most PAR can be harvested efficiently. The disadvantage of such a system is the presence of relatively large phycobilisomes (Section V1II.B) in the stroma space adjacent to the outer surface of thylakoids which imposes a number of constraints upon both the membrane and the stroma (see Section V.F). Even under conditions of light-limitation, where spatial constraints in the stroma (Section V.F) would be less important, the phycobilisome system may still impose important constraints on light-harvesting, as is shown below. Current evidence does not support a 1 : 1 ratio of phycobilisomes to PSII units in Rhodophyta, at least. Diner (1979) estimated from repetitive flash experiments that only half the phycobilisomes in Cyanidium caldarium (Rhodophyta) were connected to RCII. Evidence also comes from the density of large EF freeze-fracture particles, the putative PSII units (see above). In Cyanobacteria, there appear to be 0.5-1 .O E F particles per phycobilisome but in Rhodophyta there are between 2 and 5 particles per phycobilisome (Section VII1.B). There is also evidence for linear rows of phycobilisomes overlying rows of EF particles in Cyanobacteria and Rhodophyta (Lefort-Tran et al., 1973; Neushul, 1974; Lichtle and Thomas, 1976; Wollman, 1979); in Rhodophyta the rows are not frequent. Based on the above evidence a possible interpretation of structure and function is as follows. In Cyanobacteria the discoid phycobilisomes are arranged in rows, with their long axes at right angles to each row, overlying rows of PSII units which are sufficiently closely spaced to allow for close cooperativity between any PSII unit and its 2 neighbours. In Rhodophyta the arrangement of rows of spherical phycobilisomes, with each phycobilisome overlying a single PSII unit, would lead to poor usage of membrane space and a more plausible arrangement is the connection of each phycobilisome to 2-5 PSII units, thus allowing for close cooperativity between these 2-5 PSII units. Clusters of EF particles, in groups of 2 4 , have been observed in GriBthsia pacijica (Rhodophyta) (Staehelin et al., 1978) and Cyanidium caldarium (Wollman, 1979). Optimal packing, in terms of the greatest concentration of phycobiliproteins per thylakoid would be the arrangement of phycobilisomes, either discoid or spherical, in B closepacked configuration on a single thylakoid (cf. Guerin-Dumartrait et d.,


143

1973). It should be noted that under extreme shade this arrangement is not possible since the phycobilisomes of one thylakoid interdigitate with those of the adjacent thylakoid (Fig. 12; Subsection D.2). Even with optimal packing, it seems unlikely that there could be efficient communication between the phycobilisomes and the photosystem units. This particularly applies to the Rhodophyta where each phycobilisome has a diameter of 3 W O nm (Section VII1.B). Each phycobilisome covers a projected area which is 5 times (for discoid phycobilisomes) to 16 times (for spherical phycobilisomes) the maximum projected area of a PSI or PSII unit, based on freeze-fracture particle evidence (see above). If each phycobilisome were to connect only to a single PSI or PSII unit or even if phycobilisomes were to connect only to PSII units, as evidence suggests (Section VIII.B), but in a 1 : 1 ratio of phycobilisomes to PSII units, much of the thylakoid membrane would be unoccupied by photosystem units. Furthermore the regular, low-density spacing of PSII units would preclude cooperativity between PSII units. Thus even with such a packing system as that outlined above there would be much excess, unoccupied space in the thylakoid membrane; as a result vacant space may be taken up by unattached PSII units (Diner, 1979) or by extra PSI units (Subsection D.2). In summary, an ideal solution to the problem imposed by the spatial constraints of a phycobilisome system may not be possible. A compromise solution for efficient light-harvesting under shade conditions appears to be either to align each (discoid) phycobilisome, with a relatively small optical cross section, in a row overlying an equal number of PSII units, as in Cyanobacteria, or to connect more than one PSII unit to a phycobilisome with a much larger optical cross-section, as in Rhodophyta. However under extreme shade, with interdigitation of phycobilisomes from different thylakoids, even less efficient packing may occur. This would result in poor use of membrane space and loss of energy through the decay of the S3 state of oxygen evolution (see above) but also possibly through decay of the trans-thylakoid proton gradient (Subsection D). Since phycobilisomes are also expensive in terms of protein (Section V.E) it is apparent that the disadvantages of a phycobilisome system may well offset the advantages of wide spectral absorption (for further discussion see Sections V.F and XI1.D). C. DISTRIBUTION OF EXCITATION ENERGY BETWEEN THE PHOTOSYSTEMS

1 . Activity of PSI and PSII Units. Implicit in the hypothesis that the two photosystems act in series (Fig. 9) is the assumption that efficient photosynthesis depends on nearly equal activity of the two photosystems, although cyclic electron flow around either photosystem may result in some differences in rate. Quantum efficiency measure-

144


ments suggest that in low light conditions all plants are very efficient (Section V.D). Equal activity of both photosystems does not necessarily imply equal turnover rate. The minimum reduction time by the PSI reaction centre (RCI) as measured by P-700 bleaching (Junge, 1977; Section V.F) is S(L100 times faster than the minimum turnover time for 0, production as measured in benthic macroalgae (Mishkind and Mauzerall, 1980); also evidence from at least three species of unicellular algae suggests a faster turnover of RCI compared with RCII (Falkowski et al., 1981). It has become apparent, too, that the number of RC of both photosystems may not be equal: there may be as many as twice the number of RCII as RCI in some plants (Kawamura el al., 1979; Melis and Brown, 1980; Myers et al., 1980; Falkowski et af., 1981; Malkin et al., 1981). Under some conditions RCI may thus turn over at twice the rate of RCII. Two control mechanisms, termed here “fast” and “slow”, control the activity of the two photosystems under light-limiting conditions. The slow mechanism involves changes of the light-harvesting apparatus: changes that affect the number of light-harvesting complexes, the number of antenna Chl, the density of reaction centres and structural variation such as the number of appressed thylakoids or density of thylakoids per unit chloroplast volume. Such adaptations which are known to accompany variation in ambient lightclimate (Section 1X.D) are brought about by nuclear and ribosomal control systems involving protein synthesis, which may take hours or days to respond to changed conditions (Boardman et al., 1978; Schiff, 1978). The fast mechanism controls the distribution of excitation energy from the lightharvesting complexes to the core units of the two photosystems and possibly also intercommunication between core units. The mechanism of fast control is often called spillover a term used by Myers and Graham (1963). However, the evidence that variations in growth irradiance cause large changes via the slow mechanism (Section 1X.D) indicates that spillover is at best only a poor substitute for the longer-term changes. Nevertheless, for plants that live in conditions of variable PAR, particularly algae, such control mechanisms may be of great significance. The slow control mechanism is discussed further in Sections IX.D, X.Band XI. 2. Spillover The first significant evidence for spillover came from the work of Murata (1969a,b, 1970) who showed in Rhodophyta and in spinach chloroplasts that the amount of variable fluorescence (assigned to PSII) was affected by the previous conditions of illumination. These conditions were first defined by Bonaventura and Myers (1969) and may be restated, as follows: state I in which there is an excess of light absorbed by PSI and a decrease in the amount of excitation energy distributed from the light-harvesting pigments to PSI; and


145

state 2 in which there is an excess of light absorbed by PSII and an increase in

the amount of excitation energy distributed from the light-harvesting pigments to PSI. Current evidence indicates strongly that spillover involves the control of excitation energy from light-harvesting complexes, and PSII core units, to PSI and not from PSI to PSII (Williams, 1977; Butler, 1978). A number of models have been proposed to describe spillover and state 1state 2 transitions (e.g. Thornber and Barber, 1979). However, as a result of recent evidence on pigment-protein complexes and their distribution in thylakoid membranes (Sections VIII and 1X.B) only two such models, for green algae and higher plants, seem plausible (Fig. 49). Firstly, Butler and coworkers (Butler, 1978) developed a quantitative model in which excitation energy is distributed from light-harvesting complexes to the nearest RCII. If

Fig. 49. Two schemes for transfer of excitation energy between PSI, PSII and LHCP. (a) The model based on the work of Butler and co-workers (see Butler, 1978): (b) The model proposed by Boardman er al. (1978).

146


the trap is closed the excitation energy is then distributed to PSI (Fig. 49a). For this mechanism to work the PSI and PSII core units need to be in close proximity and the extreme lateral heterogeneity of the Andersson and Anderson model (Section 1X.B) would reduce the possibility of spillover to a low level. However evidence suggests that the amount of spillover in green algae and higher plants is not great (see Butler, 1978). In Cyanobacteria and Rhodophyta spillover is much greater (Murata, 1969a,b, 1970; Wang and Myers, 1976a,b; Ley and Butler, 1977a,b; Ried and Reinhardt, 1977, 1980) and this fact is consistent with the greater membrane homogeneity of the (nonappressed) thylakoids of these algae (Section 1X.B). Ley and Butler (1976, 1977a,b, 1980a,b) have shown that the Butler model can be applied to Porphyridium cruentum (Rhodophyta) effectively. Ried and Reinhardt (1977, 1980) have also demonstrated that many red algae show strong state 1-state 2 transitions. The Butler model (Fig. 49 (a)) may be criticized on the dependence it places on close proximity of the core units of the two photosystems and the necessity for excitation energy to “visit” the reaction centre of PSII before it can be transferred to PSI. Also the evidence is based on low-temperature fluorescence characteristics about which there is currently much argument (Bishop and Oquist, 1980; Wang et al., 1980). Wang et al. (1980) have presented evidence which calls into question the validity of the fluorescence indicators upon which the Butler model is based. The second model (Fig. 49(b)) of Boardman et al. (1978) proposes that any spillover in higher plants, and presumably green algae, is via LHCP. This model fits well with conclusions derived from structural considerations (Section 1X.B) (Wollman et al., 1980; Dubertret and Lefort-Tran, 1981). The model could be extended to accommodate other integral membrane lightharvesting pigment-proteins and, with somewhat more difficulty, to the phycobilisome system in Cyanobacteria and Rhodophyta (Fig. 5 l(b)). While evidence supports a close link between the phycobilisome and PSII through a “stalk” of allophycocyanin B (Section V1II.B) it is possible that there are two “stalks” one to each photosystem. Alternatively mediation may occur between the core units of PSI and PSII by antenna Chl-protein(s) common to both photosystems, as in the schemes of Thornber et al. (1977) and Larkum and Weyrauch (1977). A counterview to the concept of spillover is argued by Chow et al. (1982) and Thorne et al. (1982) on the theoretical and experimental grounds that saltinduced fluorescence changes, A F683, relate mainly to light-scattering changes which arise from alterations in chloroplast structure, giving rise to direct, differential absorbancy changes between PSI and PSII. 3. Control Mechanisms for the Distribution of Excitation Energy The basis for control of spillover in the short-term state 1-state 2 transitions

147


has been largely unexplored. Allen et al. (1981) and Horton et af. (1981) have put forward a mechanism for higher plant systems involving LHCP. They proposed that phosphorylation of LHCP (Bennett, 1977,1979; Bennett et al., 1981) is dependent upon the redox state of plastoquinone with phosphorylation taking place under reduced conditions, which in turn promotes the association of PSII-LHCP complex with PSI (Fig. 50, see also Barber (1983)). The model of Allen et al. (1981) incorporates a Butler-type of spillover model from PSII to PSI and does not take into account the evidence for extreme lateral heterogeneity of the two photosystems (Subsection B). However these criticisms can be met by a modified scheme, as shown in Fig. 5 1a incorporating membrane appression. Although the mechanism of appression of thylakoids is not fully understood, it is known that in higher plants it is controlled by salts, especially divalent cations (Sculley et al., 1980; Chow et al., 1980; Chow et al., 1982) and by pH (Arntzen, 1978; Gerola et al., 1979, 1981; Barber, 1980; Jennings et al., 1981; Grouzis et al., 1982). These effects may be mediated through an exposed polypeptide loop (2 KD) of LHCP which carries negative charges and magnesium binding sites (Steinbeck et af.,1979; Mullet and Arntzen, 1980; Ryrie et al., 1980): the interaction of the exposed LHCP loops with those of an adjacent membrane may be the operational factor in causing membrane appression and lateral heterogeneity. Barber (1980) suggested that PSI units are charged and are electrostatically repelled from appressed regions of thylakoids. The fluorescence characteristics of chloroplasts, upon which much of the evidence for spillover is based, have been correlated by Barber (1980) with the degree of thylakoid appression. Barber (1980) proposed, on theoretical grounds, a model of photosystem distribution in appressed and non-

I1

I 1- 1 1

hght

6 . / \ phorphotam.Mg2+ b

ps,,

LHC

protein

4

m

excitotian energy transfer

kinarc, Alp, Mg2'

m

c x o t a t ~ o n energy transfer

-z(.pr)L,/

excitation energy transfer

octwolion

m

NADP

\

P700

m m

p680

'-

c -W

20

electron transport

Fig. 50. Scheme for the control of the distribution of excitation energy between PSI and PSII. (Redrawn from Allen e r a / . , 1981.)

148


appressed regions of thylakoids broadly similarly to the model of extreme lateral heterogeneity of Anderson and Anderson (1980, Fig. 47). The primary cause of this lateral heterogeneity has been shown by Chow et al. (1982) to arise from PSI carrying a higher negative charge than PSII in vivo, at neutral pH in the presence of Mg2 +.This results from Mg2 ions adsorbing selectively to the membrane surfaces of PSII, with no apparent binding to PSI. +

Phae 7

ATP

(b)

-[@ = ~% Fig. 51. A model for the control of distribution of energy in a chloroplast system with lateral heterogeneity, (a) and a phycobilisome-thylakoid membrane system, (b).

Barber pointed out that spillover would be greatly decreased by lateral separation of the photosystems, and suggested that the spillover changes accompanying state 1 and state 2 changes were brought about by changes in the ratio of appressed to non-appressed membranes. Extending this proposal, it is possible that phosphorylation of the exposed loop region of the LHCP complex reduces the interaction with a neighbouring LHCP complex in the opposite membrane. Phosphorylation would thus lead to an increase in the proportion of non-appressed membranes and an increase in the amount of PSII and LHCP complexes in non-appressed membranes; that is, greater spillover would be favoured. This model is set out in Fig. 51(a). How far such a process based on phosphorylation of LHCP could alter the structure of the chloroplast depends on what other factors affect thylakoid appression in vivo. Both the model of Allen et al. (1981) and the modified model (Fig. 51(a)) are self-regulatory; as the LHCP complex becomes phosphorylated more excitation energy is diverted to PSI and less to PSII, thereby oxidizing the pool of plastoquinone and inhibiting the phosphorylation reaction. The modified model takes into account structural as well as functional control of energy distribution in the thylakoid membrane. 4 . Control Mechanisms in Algae Although Chlorophyta and related algae do not possess typical grana, the


149

ratio of appressed to non-appressed thylakoid membranes is very variable (Section V.D.3), and the spillover model set out above may be extended to these algae. With algae of other phyla where thylakoid appression does occur, the degree of appression has been hitherto regarded as invariant (Section V.E.3). This fixity of appression implies a rather inflexible short-term control of spillover. However, Humphrey (1982) has found that thylakoid appression in two chromophytes, Amphidinium and Biddulphia, is more variable than was previously thought, and is related to the spectral climate. For Cyanobacteria and Rhodophyta, spillover of excitation energy from the phycobilisomes, situated on thylakoids which are never appressed, may require a different mechanism. Ried and Reinhardt (1980) concluded that state 1-state 2 transitions in Rhodophyta were controlled by the oxidation-reduction state of a component of the electron transport chain. It has become important to test for phosphorylation of the phycobilisome, and to investigate whether the redox state of plastoquinone controls, via phosphorylation, the distribution of excitation energy between the two photosystems (see Fig. 51(b)). D. INTERACTION OF THE LIGHT-HARVESTING APPARATUS WITH OTHER PHOTOSYNTHETIC PROCESSES

I . Photosynthetic Rate Versus Light Intensity ( a ) The P versus I curve. (i) Introduction. The general relationship of photosynthesis to incident PAR is shown in Fig. 52a. The general form of such a P versus I curve has been known for many years (for early work, see Rabinowitch, 1951), but no widely accepted mathematical description of the curve exists even today (see below). The curve can be divided into four regions (see Fig. 52b): (i) the initial slope (often assumed to be linear) where incident light is the major limiting factor and the slope is determined mainly by the lightharvesting capacity of the alga; (ii) a region near light saturation where the supply of ATP and NADPH or the activation of RuBP carboxylase (Perchorowicz et al., 1981) may possibly be the major limitation (Section V.F.5); (iii) a light-saturated region where enzyme turnover (most probably of RuBP c’ase-Section V.F.6) is the major limitation; and (iv) a region where photoinhibition occurs (see Subsection 3). A fifth region may exist at extremely low light intensities where the quantum efficiency of photosynthesis is reduced (Radmer and Kok, 1977; Raven and Beardall, 1982; see Subsection (b) below). (ii) The initial slope. In theory the initial slope is determined by the quantum

150

A. W . D. LARKUM A N D JACK BARRETT

efficiency (Section V1.D) and the per cent absorption of light by the photosynthetic pigments, which approximates to the per cent absorption or absorptance of the alga or algal suspension. Thus the light-harvesting apparatus, at the molecular level, should largely determine the initial slope (cf. Herron and Mauzerall, 1972). However two complications make a quantitative assessment difficult. Firstly the photosynthetic rate must take into account the rate of respiratory (mitochondrial-linked) processes. Such processes are difficult to estimate in the light (Raven, 1972) and may show light-inhibition at low irradiances, the so-called Kok effect, (Kok, 1949; Hoch el al., 1963; Healy and Myers, 1971).Secondly, photorespiration occurs in the light (Hatch, 1976) and this process increases with increased irradiance (Hoch et al., 1963; Jassby, 1978). (a)

lrradiance

lrradiance

Fig. 52. Photosynthesisversus irradiance curves. (a) Theoretical curve; I,, compensation light level Ik, light level where the linear slope curve intersects with the P,,, line. (b) The major factors influencing the P versus I curve at different light levels. (c) Typical P versus I curves for sun,shade and intermediate types.

These factors probably account for a non-linear initial slope in many algae. In addition, such factors as internal multiple light scattering (Section V.G.4), surface reflectance (Drew, 1982), chloroplast distribution (Section V.E.3) and cell density (Rabinowitch, 1951; Kok, 1960) may affect the initial slope. Thus it is not surprising that in comparing the P versus I curves of a wide variety of algae (Table V) a wide range of characteristics is found. Nevertheless, it is

TABLE V The I, and PmaX of P versus I Curvesfor Various Unicellular Algae (see Fig. 52 for explanation 0.f these parameters)

Species Chlorophyta Chlorella vulgaris Scenedesmus obliquus Dunaliella tertiolecta Dunaliella euchlora

Bacillariophyta Skeletonema costaturn Chaetoceros gracilis

Dinoflagellata Glenodinium sp. Gonyaulax polyedra Zooxanthellae

Chrysophyta Isochrysis galbana Cyanobacteria Anacystis nidulans

Growth irradiance or Photon flux density

pmax

Reference

prnole CO,/rng Chl/hr

28 (Wrne2) 5 (Wrn -2) 400 2 300 4

115 (Wm-') 30 (Wm -2) 230 30 150 25

458 667" 221" 390" 155" 380 130

130 0.I 300 300 4

70 45 100 160 45

190" 155" 430 590 210

loo0 100 250 300

133" 63" 255" 149" 193" 53"

35 (Wrn -') 22(Wm-') 45 (Wrn -') 30 (Wm -') 2100 150

25 (Wm -2) 2.5 ( W m-2) 25 (Wm - 2 ) 4 (Wm - 2 ) - 1700 - 250

28 (Wrn -2) 14 (Wm-') 4O (Wm -') 30 (Wm -*)

400 (Wm -2) 200 ( w m - 2 ) 700 150 250 120

150

9 8 8 (see also 2) 4 4 5 5 3 3 5 5 5 6 6 7 7 1 1

4

180 70

670 170

700 120

5 (see also 2-3) 5

300 4

300 130

830" 420"

700 400

10 10

300

0

Recalculated from values of photosynthetic oxygen evolution assuming a photosynthetic quotient of 1.2. (Ryther, 1956). (AOd References: (1) Falkowski and Dubinsky (1981); (2) Dunstan (1973); (3) Falkowski and Owens (1978); (4) Falkowski and Owens (1980); (5) Perry et a / . (1981); (6) Prezelin (1976); (7) Prezelin and Sweeney (1978); (8) Senger and Fleischhacker (1978); (9) Steeman-Nielsen (1961); (10) Vierling and Alberte (1980). @

152


possible to categorize algae broadly into sun and shade types, along with intermediates as shown in Fig. 52(c). This can be understood in terms of lower densities of PSI and PSI1 units, but with larger light-harvesting arrays, in shade algae (see Subsection 2). It has often been assumed that the initial slope of the P versus I curve is a constant characteristic for a specific alga grown under a given set of conditions (Parsons et al., 1977). However short-term changes in the initial slope have recently been recorded (Prezelin and Matlick, 1980; Ramus and Rosenburg, 1980) which seem to depend on the stage of life cycle, on diurnal changes and on the conditions of previous illumination. Diurnal patterns of photosynthetic activity, due to endogenous factors, have previously been discussed by Harris (1980). In synchronously-grown Scenedesrnus (Chlorophyta) quantum efficiency has been shown to change diurnally (Senger and Bishop, 1969). In synchronously-grown Chlorella vulgaris (Chlorophyta) marked changes in Chl a, Chl b, RCI, RCII and cyt.f occur during the light period (Venediktov et al., 1979) and these changes would be likely to affect the quantum efficiency. (iii) Mathematical descriptions. A number of mathematical descriptions of the P vs I curve exist (for more extensive discussions see Thornley, 1976, Parsons et al., 1977, Chalker, 1980). These range from very simple to very complex functions involving four or more variables. In some cases attempts have been made to include terms for the effects of carbon dioxide concentrations, respiration and photorespiration (Charles-Edwards and Ludwig, 1974). However all these treatments predict curves which lack an initial linear region. Also few models can easily account for the region of photoinhibition, with the exception of Steele (1962) and Fee (1969). As a result, for modelling purposes, many workers prefer the simpler formulation of Steele (cf. Parson et al., 1977). Jassby and Platt, 1976, found that the best fit to many curves was a tan -' function and Chalker (1980) has extended this idea with a theoretical treatment, although neither approach allows for photoinhibition. Talling (1957) was the first to propose that the intercept of the initial slope and the P,,, (Fig. 52), called I,, is a useful characteristic. Since that time it has been widely used in algal studies (see Steeman-Nielsen, 1977, Parsons et al., 1977), together with P,,, and the initial slope (k) to define algal photosynthesis. The basic premise of such an empirical approach is that the dark reactions of photosynthesis can be considered as a single enzymatic reaction over the range of irradiance and that light can be considered to be a substrate. This leads to a Michaelis-Menten type of approach where the curve is treated as hyperbolic. However, IK is not related to the K, of the Michaelis-Menten equation. Probably a more fruitful but difficult approach would be to model all the limiting reactions in photosynthesis in the manner attempted by Farquhar and von Caemmerer (1981). These workers have set out mathematical

153


descriptions of CO, supply, respiration. photorespiration, the enzymatic reactions of the photosynthetic pentose reduction pathway, the levels of ADP/ATP and NADP/NADPH and the rate of electron transport. (6) Quantum yield, turnover and pigment concentration. Quantum yield is defined as the number of CO, molecules fixed in photosynthesis per quantum of light absorbed (Radmer and Kok, 1977). It is now generally agreed that the maximum quantum yield is between 0.1 0 and 0,125 (Kok, 1960; Radmer and Kok, 1977). It is clear from the P versus I curve that the highest quantum yields are obtained from the initial region of the curve : in fact the quantum yield can be calculated from this slope if the proportion of absorbed light is calculated (Senger and Fleischhacker, 1978) although quantum yields are generally calculated by flash photolysis (Emerson and Arnold, 1932a,b). However, at the extreme lower limit of irradiance there may be an initial region of non-linearity and low quantum yield, where the irradiance is too low to promote efficient photosynthesis. Radmer and Kok (1977) put forward an hypothesis in which the lower limit was determined byt the S-states involved in oxygen evolution (Section IX.A.9). They proposed that the back reaction of the couple S2p S 3 becomes rate limiting at very low levels of turnover of RCII (Fig. 53) i.e. under very low levels of irradiance. Figure 53 shows how the dynamics of the S-states can be seen to determine the efficiency, and therefore the quantum yield of photosynthesis over a range of irradiance. Taking an “average” cell with “average levels” of pigmentation Seconds between hots per chbrophyll or per centre: 0.2 0.02 20 2 Chlorophyll 200 4 A I centre 1 1b-3 i04 01 b 2

1

10

102

103

T

I

lo4

sunlight Photon Fluence Rate

/

~~Erri~s-‘

Fig. 53. Working range of photosynthesis, and some of the limiting parameters as suggested by Radmer and Kok (1977). The S- states are discussed in the text. (Redrawn from Radmer and Kok. 1977.)

154


the half-life of the least stable precursor (S,) is about 3 s. Thus the efficiency of photosynthesis is half maximal at an irradiance at which each RCII receives a quantum every 3 s. For a PSII unit with 200 Chl molecules per RC this corresponds to the absorption of one quantum per Chl moleculeevery 10 min, or about 1/1000th the intensity of full sunlight (it will be noted from Section 111 that this is about the absolute lower limit for benthic algae). Above this lower limit the efficiency of photosynthesis approaches a maximum at about one quantum per molecule of Chl every 30s: this corresponds to about 1/100th the intensity of sunlight and a turnover, on average, of each RC every 150 ms. At a turnover rate of 20 ms the oxidation of reduced plastoquinone becomes rate limiting (Section V.F). Thus, at a light intensity of about 1/15th sunlight, where photosynthesis is half-saturated, the light-harvesting apparatus is no longer a limiting factor. Above this level of irradiance excess excitation energy absorbed by the light-harvesting apparatus is deactivated via triplet-triplet interaction of Chl, carotenoids and oxygen and lost as heat (Section VI.B.8). The hypothesis of Radmer and Kok based on the above example using average concentrations of Chl and plastoquinone leads to the conclusion that the photosynthetic apparatus of a typical plant is adapted to operate efficiently under low light conditions (between 1/100th and 1/1000th of full sunlight). This range presumably best matches the natural fluctuations of irradiance during a diurnal cycle. Shade plants adapt to a lower dynamic range by increasing the size of PSI and PSII units and light-harvesting complexes (Boardman, 1977; Wild, 1979; Malkin and Fork, 1981; Falkowski et al., 1981). Raven and Beardall (1982) have recently put forward an alternative hypothesis to explain the lower limits to photosynthesis. They point out that photosynthetic bacteria without the oxygen-evolving machinery have similar lower limits to photosynthesis and suggest that it is the inability to maintain a sufficient gradient of protons across the thylakoid membrane, to power the formation of ATP (Section IV), which ultimately limits photosynthesis under extreme shade. 2. Effects of Shading in Plants ( a ) General characteristics. The characteristic effects of shading in higher plants are set out in Table VI. Much of the evidence used here comes from higher plants but algae show similar trends which are discussed in detail below. In general shading of plants leads to larger chloroplasts, greater concentrations of Chl a and of light-harvesting complexes per chloroplast. The number of chloroplasts per cell may also increase. However, in multicellular plants there is a marked decline in the number of layers of photosynthetic tissue and the amount of the primary carboxylase, RuBPc’ase, decreases.

155


TABLE VI General Characteristics of Shading in Plants Based on Work on Vascular Plants and Algae Functional level

Photosynthetic reaction

Effect of shading

Leaf or frond anatomy

Stomata per unit area Leaf/frond thickness Number of palisade cells Size of cells Dry wt per fr.wt Chl per g fr.wt Chl per unit surface area

Decreased greatly Decreased greatly Decreased greatly Decreased greatly Decreased greatly Increased Decreased

Chloroplasts

Number per cell Number per cross-sectional area Size Chl per Chloroplast Grana Stroma volume

Decreased Decreased Increased Increased Increased greatly Decreased

R,

Stomata1 resistance

Increased greatly

CO, fixation

RUBP Other soluble stroma enzymes Glycollate oxidaseidehydrogenase Soluble protein per Chl

Decreased greatly Decreased Decreased greatly Decreased greatly

Electron transport

Cytochrome f per Chl Ferredoxin per Chl Plastoquinine per Chl Rate of electron transport NADP reductase per ATP’ase

Decreased greatly Decreased greatly Decreased greatly Decreased greatly Decreased ?

Photochemical units

P-700 per Chl P-700 per cyt f P-700 per ferredoxin P-700 per plastoquinone

Little change

Chl a/b ratio LHCP Accessory pigment-protein complexes in algae

Decreased Increased greatly Increased greatly

Light harvesting complexes

Increased greatly

Clearly under low light conditions the supply of CO, becomes less critical and the “payload” of the tissue is reduced by reducing the number of cells. (b) Shading eflects in algae. Much work has been carried out on the effect of shading in various divisions of algae. Table VII summarizes some of the of shade algae is major evidence. In general the photosynthetic capacity (PmaX)

TABLE VII Effects of Shading on some Unicellular Algae ~~

Species Chlorella pyrenoidosa (Chloroph yta) Dunaliella tertiolecta (Chlorophyta) Scenedesmus obliquus (Chlorophyta)

Anacystis nidulans (Cyanobacteria) Anabaena variabilis (Cyanobacteria)

Porphyridium cruentum (RhodOPhYta) Grifithsia pacifica (Rhodophyta)

Shade adaptation Increase in Chl a and Chl b content and Hill activity, per cell. Decrease in Chl a/b ratio and cell volume. Increase in Chl a and b per cell and P-700 per cell. Decrease in Chl a/b ratio,,,P I, cell volume, (N ratio, respiratory rate) Increase in Chl a and b per cell. Decrease in Chl a/b ratio, P, Ik cell volume, Hill activity per mg. Chl., Ps 1 activity, cytochrome f+ b, plastoquinone activity, respiratory rate.

~~

Shade irradiance or photon flux density

40 f.c. 2 pEm -'s

Reference Brown and Richardson (1968)

-'

Falkowski and Owens (1980)

(a) Fleischhacker and Senger (1978) (b) Senger and Fleischhacker (1978)

S Wm-'

-'

(a) Vierling and Alberte (1980) (b) Brown and Richardson (1968) (c) Myers and Kratz, (1955)

Increase in Chl a per cell, phycocyanin per cell P-700 per cell volume. Decrease in P, and I,. Little change in Chl alphycocyanin ratio. Increase in P-700 per cell. Little change in RCII per cell.

(a) 10 pEm -'s (b) 50 f.c. 500 Ix

Kawamura et al. (1979)

Increase in Chl a and phycobilins per cell, thylakoids per chloroplast, chloroplast volume. Decrease in cell volume. Little change in Chl a/phycobilin ratio. Increase in Chl a and phycobilin content, phycobilisomes per unit area of thylakoids. Decrease in Chl alphycobilin ratio. Little change in chloroplast size or number of thylakoids per chloroplast.

(b) 55 f.c.

(a) Brody and Emerson (1959a,b) (b) Brown and Richardson (1968) (c) Ley and Butler (1980a) Guerin-Dumartrait et al. (1973) (a) Waalund et a / . (1974) (b) Staehelin et al. (1978)

(a) 4 f.c. (b) 50 f.c.

Cyanidium caldarium (Rhodophyta)

Increase in Chl a per cell and Chl a/phycocyanin ratio. Decrease in cell volume.

(a) 55 f.c.

(a) Brown and Richardson (1980) (b) Halldal and French (1958)

Sphacelaria sp. (Phaeophyta)

Increase in Chl a and fucoxanthin per cell, Chl alfucoxanthin ratio, Hill activity per cell.

42 f.c.

Brown and Richardson (1968)

Skeletonema costaturn (Bacillariophyta)

Increase in Chl c per cell. 0.7 pEm -’s Decrease in P-700 per cell, P,,,, cell volume, C/N ratio, respiratory activity. Little change in Chl a per cell, Ik. Increase in Chl a and fucoxanthin per cell, Chl 50 f.c. alfucoxanthin ratio, cell size, chloroplast volume, Hill activity per cell.

Phaeodacrylum (Nitschia) Closterium (Bacillariophyta) Glenodinium sp. (Dinoflagellata)

-’

Falkowski and Owens (1978) 1980

Brown and Richardson (1968)

2.5 Wm-’ Increase in Chl a and peridinin per cell, peridinin-Chl a-protein, activity of green light in action spectrum. Decrease in Chl a/c per cell. Chl a/peridinin per cell. P,,,. I,. Little change in P,, per cell, Chl c per cell. 4 Wm Complex changes in Chl a, Chl c and peridinin per cell-these begin to increase with shading but decline at very low irradiance. Decrease in P,,,. I,, photosynthetic performance, cell volume, respiratory rate.

(a) Prezelin (1976) (b) Prezelin et a/. (1976)

Cryptomonas ovata Cryptomonas rufescens (CrYPtoPhYta)

Increase in Chl a and phycobilins per cell. Hill activity per cell, chloroplast size. Decrease in Chl a/c ratio.

(a) 36 f.c. (b)4 Wm-’

(a) Brown and Richardson (1968) (b) Lichtle (1979)

Chroomonas sp. (Cry ptophyta)

Increase in Ch/a and phycocyanin per cell. intrathylakoid width. No change in Chl a/c ratio.

1 Wm -2

Faust and Gantt (1973)

Gonyaulax polyedra (Dinoflagellata)

-’

Prezelin and Sweeney (1978)

158


low (see, for example, Steeman-Nielsen, 1975; Prezelin, 1976;Jorgensen, 1969; Vierling and Alberte, 1980; Perry et al., 1981).Jorgensen (1969) suggested that there were a number of algae in which the reverse was true, i.e. photosynthetic capacity increases with shading and one of these algae was Scenedesmus obliquus (Chlorophyta). However in a detailed study of S. obliquus, Senger and Fleischhacker (1978) have shown clearly that in weak light 5 W m the photosynthetic capacity on a Chl basis, was only 32 per cent of that for cells grown in strong light, 28 W m -2. Lower capacity therefore appears general on a Chl basis, but because cells of shade algae are larger, photosynthetic capacity on a per cell basis may show little change (e.g. Vierling and Alberte, 1980). Prezelin and Sweeney (1978) demonstrated that another characteristic, photosynthetic performance, is better correlated with photo-adaptation than photosynthetic capacity in Gonyaulax polyedra (Dinoflagellata). Photosynthetic performance is defined as the photosynthetic rate occurring at the level of irradiance under which the cells have been grown. Amongst all the photosynthetic characteristics examined by Prezelin and Sweeney (1978) only photosynthetic performance showed a consistent, but not exact correspondence, with growth rates. Respiration in algae is usually depressed under shade conditions (Prezelin and Sweeney, 1978; Senger and Fleischhacker, 1978; Falkowski and Owens, 1980; Raven and Beardall, 1982). Thus the compensation point (the level of irradiance at which gross photosynthetic rate equals the respiratory rate) is low in shade algae. This adaptation allows algae to survive at greater depths in the water column, but is not in the strict sense a light-harvesting adaptation. (c) Adaptation of the light-harvesting apparatus. From the discussion of the rate-limiting steps of the photochemical apparatus (Subsection 1.b, above) it is apparent that under intermediate conditions of light limitation (in the “linear” region of the P versus I curve) the oxidation of reduced plastoquinone is most likely the critical step in photosynthesis (cf. Raven and Beardall, 1982). Here, optimal rates of photosynthesis can be maintained by matching the levels of plastoquinone, and other intermediates between the two photosystems, such as cytochrome f, with the levels of irradiance. Evidence that this does occur has been obtained in algae by Fleishhacker and Senger (1978) (for higher plants, see Boardman, 1977; Wild, 1979), who found that the most significant changes to occur when Scenedesmus obliquus was grown at 5 W m as compared with 28 W m -2, were a 50 per cent decline in cytochrome f and cytochrome b, and a 70-75 per cent decline in the pool size (or turnover) of plastoquinone (as judged from inhibition studies with dibromothymoquinol). These decreases were accompanied by increases in the levels of Chla and Chlb per cell and in PSI1 activity. Thus the light-harvesting apparatus also changes in the intermediate region of light limitation, presumably in order to match the turnover rates of the reaction centres with electron transport through plastoquinone. Thus in algae as in higher plants

-’,

-’


159

(Fig. 54) the proportion of reaction centres to electron transport components may change according to the irradiances at which the plants are grown.

Fig. 54. A proposed arrangement of RCI and RCII units and other components of the photosynthetic electron transport chain in shade plants. (Redrawn from Boardman et al., 1978.)

At extremely low light intensities the major rate-limiting step of the photochemical apparatus may be the S , d S , step of oxygen production or passive dissipation of the trans-thylakoid proton gradient (Subsection 1 .b). Under these conditions only increased activity of the PSII reaction centres (RCII) can improve the efficiency of photosynthesis. This can be brought about in the following four ways: (i) increase in the size of PSII core units and an increase in the number of light-harvesting complexes servicing each unit, i.e. increase in the optical cross-section; (ii) Increase in the concentration of PSII core units either by increase in the density of units on the thylakoid membrane or by increase in the thylakoid content of the chloroplast; (iii) increase in the intercommunication of PSII units (by either core or light-harvesting complexes); (iv) increase in the spectral width of PAR harvested by each PSII unit. At present there is only meagre evidence concerning these four possible adaptations in algae. However there is sufficient to suggest that the algae may be divided into at least two groups. The first group consists of the Chlorophyta and allied phyla and possibly the Chromophyta. This group is similar to the higher plants in that all four adaptations probably occur, although the evidence at present is fragmentary. The following points are important considerations. (i) under shade conditions the relative proportions of pigments of lightharvesting complexes increases relative to Chl a (Table VII); (ii) the number of thylakoids per unit volume of chloroplast increases with shading (Table VII) and in Chl b-containing algae the number of appressed thylakoids increases (Table VII);

160


(iii) the photosynthetic unit (the number of pigment molecules or chromophores per C 0 2 fixed in flash yield experiments) remains relatively constant in sun and shade algae (Myers and Graham, 1971; Mishkind and Mauzerall, 1980; Falkowski et al., 1981) as well as in higher plants (Boardman, 1977), at values between 1500-2500; but there is sometimes a trend for lower values in sun plants and higher in shade plants (Myers and Graham, 1971; Mishkind and Mauzerall, 1980; Malkin and Fork, 1980; Falkowski et ul., 1981); (iv) the ratio RCII/RCI (number of reaction centres in PSII to those in PSI) is near to 1 ( 1 .I-1.2) in Dunaliefla tertiolecta (Chlorophyta) under high and low light conditions and much greater than 1 (2.3) in Skefetonemacostaturn (Bacillariophyta) under low light conditions (Falkowski et al., 1981); in higher plants the ratio is known to be greater than 1 in spinach and much greater than 1 in appressed thylakoids (Melis and Brown, 1980), and is influenced by light quality (Melis and Harvey, 1981); (v) higher plants under shade conditions may have up to 80 per cent of pigments in PSII or LHCP (Anderson et al., 1973; Boardman et al., 1975). These facts lead to the conclusion that shading results both in an increase in the RCII/RCI ratio and an increase in the number of pigment molecules servicing each RCII. Evidence for increased intercommunication between PSII units comes from fluorescence induction kinetics in Scenedesmus obliquus (Chlorophyta) (Fleischhacker and Senger, 1978). Finally increased spectral width of harvested light would accompany the increased concentration of pigments, especially of light-harvesting complexes, and greater numbers of appressed lamellae (Section V.G). The second group of algae are the Cyanobacteria and the Rhodophyta. In these algae, too, low light conditions result in a great increase in the number of thylakoids and phycobilisomes per unit volume of chloroplast (Table VII). However in contrast to the first group there is good evidence that the ratio RCII/RCI is < 1 (Fujita, 1976; Mimuro and Fujita, 1977; Kawamura et al., 1979; Melis and Brown, 1980; Myers et a f . , 1980). Unfortunately only the study by Kawamura et al. (1979) of Anabaena variabilis (Cyanobacteria) was concerned with shading effects; it was found that the ratio RCII/RCI was 0.67 in low light (500 lux) and 1.43 in high light (4000 lux). The change in ratio was brought about mainly by an increase, under shading, of the number of P700 molecules per cell and there was little change in the RCII number. The phycobilisome system and its response to shade conditions can be understood in terms of the packing constraints on a system where the major light-harvesting pigment protein is located in the stroma space (Section 1X.C) (Fig. 28). Even under optimal packing, with phycobilisomes close-spaced on one membrane, it seems unlikely that there could be a one-to-one relationship between the photosystem units and phycobilisomes, since each phycobilisome


161

occupies a projected area which is 5 times (for discoid phycobilisomes) or 16 times (for spherical phycobilisomes) the maximum projected area of a PSI or PSII unit, based on freeze-fracture evidence (Section 1X.C). In fact from the freeze-fracture evidence there are between 1 and 5 PSII units per phycobilisome (Section V1II.B). Since, under shade conditions, the phycobilisome connects preferentially to PSII units (Amesz and Duysens, 1962; Wang et al., 1977; Mimuro and Fujita, 1980; Ley and Butler, 1980a) there would be much unoccupied space in the thylakoid membrane if the ratio of RCII/RCI is unity. The evidence from the RCII/RCI ratio suggests that at least a part of the space is filled with excess PSI units. Under conditions of extreme closepacking where the phycobilisomes of one thylakoid interdigitate with those of the neighbouring thylakoid (Fig. 28) as occurs under extreme shade the packing efficiency of phycobilisomes in relation to PSII units must be even poorer. I t should be noted that such a model still allows for some cooperative interaction of PSII units especially in Rhodophyta with spherical phycobilisomes (Section 1X.C). The finding of Yu e f al. (198 1) that the ratio of PEB and PUB chromophores of the PE of the marine rhodophyte Callithamnion roseum is modulated by shading, demonstrates another form of light intensity adaptation in red algae. This type of adaptation allows for some control over the spectral range of light-harvesting. (4 Dynamic range of light-harvesting. From the discussion of the lightclimate of algae (Section 111) it is apparent that underwater irradiance may vary greatly from hour to hour from day to day and from season to season. Therefore in algae, more so than in other plants, it is essential to control the levels of C02-fixing enzymes, electron transport components and lightharvesting apparatus to give an optimum level of photosynthesis over a fairly long time-period (days or weeks). Presumably there are detector and control systems to effect such a response but little is known of these (see Section 1X.C). The stimulus could be the average daily irradiance or the maximum irradiance over a period of days or some other function of irradiance. In response to such a stimulus the control system presumably sets the levels of the various parts of the photosynthetic apparatus. Such considerations and their elucidation may lead to an understanding of why “normal plants” under high growth irradiance in the field, approach saturation of photosynthesis at about 1/15th sunlight irradiance and algae growing under shaded conditions approach saturation at much lower light levels (Table V). Many attempts have been made to grow algae in the laboratory under defined, constant conditions of irradiance and thereby investigate the effect of shading. Apart from the matter of light quality (Section XI), such studies are difficult to interpret without some theoretical framework for the stimulus and control mechanisms involved. For example, on what basis should one compare cultures grown under constant, high, daytime irradiance with algae

162

A. W.

D. LARKUM AND JACK BARRETT

growing under high midday irradiance in the field? And is an alga grown at very low constant irradiance adapted specifically to that irradiance or is it adapted as if it will receive a range of irradiance? As pointed out by Myers (1946) cells grown at a given, constant, irradiance have a P,,, at much higher irradiance than the growth irradiance. Since Myer’s work these questions have not been entirely ignored (Marra, 1978; Harris, 1980; Savidge, 1980) but they still remain largely unanswered. (e) Sun andshade algae. There is little doubt that in ecological terms there are sun and shade communities of algae composed of different species (Section V.B). The question arises then as to whether there are special light-harvesting mechanisms which characterize the algae of these communities. Little is known on this matter since there is little work on deep-water benthic algae. The occurrance of siphonaxanthin in deep-water chlorophytes (Kageyama et al., 1977; Anderson et al., 1980) is one clear example of a light-harvesting adaptation. At present no evidence exists on the size or number of PSI or PSII units in such deep-water algae. Falkowski and Owens (1980) have suggested that there may be two strategies of shade adaptation in phytoplankton, involving light-harvesting (but see Falkowski et al., 1981). In the first or sun type the number of PSI (and PSII) units increases with shading whereas in the second or shade type the size of the PSI or PSII units increases. There is however no clear explanation as to why one strategy should be better than the other for sun or shade types. As pointed out above, both these and other adaptations probably occur together as a shade response of some algae. The results of Falkowski and Owens (1980) based on work with only one “sun type”, DunaIiella tertiolecta (Chlorophyta), and one “shade type”, Skeletonema costatum (Bacillariophyta), should be treated with caution, especially since changes in C0,-fixing enzymes and electron transport components were not considered. In most phytoplankton, subject to large passive vertical movements in the water column the development of definite sun and shade species would be puzzling. In the case of Dunaliella tertiolecta, which grows in shallow, saline lagoons the development of sun-type characteristics seems more reasonable. However it is probable that the basis, if any, for any such adaptation is based on the characteristics of the C0,-fixing apparatus and photoprotection (Section V.F) rather than light-harvesting properties. (f) Cave and other deep shade algae. A number of studies have been carried out on algae found growing in caves under the extreme low light conditions of below 1 W m-’(Norton et al., 1971; Cox and Marchant, 1977; Cox etal., 1982; Leclerc etal., 1981) or in other equivalent situations (Halldal, 1968; Vincent, 1980; Parker et al., 1981; Leclerc et al., 1981). Care must be exercised in making conclusions from such studies unless the algae are regrown under defined conditions, since heterotrophy (Droop, 1974) may occur under natural conditions.


163

Studies on such extreme shade algae have revealed few novel features, of the light-harvesting apparatus, so far. The proliferation of thylakoid membranes is extreme (Fig. 12; Cox et al., 1982) as is the concentration of antenna and light-harvesting pigments (Leclerc et al., 1981). Leclerc et al. (1981) found that many cave algae were characterized by forms of Chl a with long-wavelength absorption (685-720 nm) and a good quantum yield in this region; Halldal (1958) also demonstrated that the boring alga Ostreobium sp. (Siphonophyta), which lives deep within the skeleton of living Favia corals, can photosynthesize efficiently in near infrared light. Leclerc et al. (1981) found that some algae, e.g. Spirulina platensis (Cyanobacteria), could repress respiration very effectively (the Kok effect) at low irradiance ( < 10 p E m -2 s -I), by an unknown photocontrol mechanism that was wavelength-dependent, while other algae e.g. Ch1orobotr.w sp. (Xanthophyta) showed no such effect but were equally efficient up to 700 nm. They tentatively suggested that, in the latter group, reversed electron transfer towards PSI1 might occur (cf. Van Ginkel and Kleinem-Hammas, 1980). Recently, Raven and Beardall (1982) have suggested that increased resistance to the passive movement of protons in the thylakoid membrane may be an important property of algae that survive in extremely shaded conditions. ( g ) Summary. (i) Light-harvesting processes are not the major limitation on photosynthetic activity except at very low irradiance levels. Other parts of the photosynthetic apparatus such as the electron transport chain and the pools of ATP + ADP and NADPH + NADP are the sites of limitation in the middle and upper linear region cf the P versus I curve. (ii) The efficiency of light-harvesting per cell at very low irradiance levels can be enhanced by four means which increase the activity of the two photosystems. (iii) Chlorophyta, and possibly algae in the Chromophyta group, may show all four photoadaptations. Cyanobacteria and Rhodophyta appear to be distinct in that the ratio RCII to RCI decreases instead of increasing under shading. This can be related to the presence of the phycobilisome which transfers energy mainly to PSI1 and places constraints on the packing of the thylakoid membrane. (iv) All algae that have been studied adapt to shade conditions. No clear distinction in terms of light-harvesting can be made at present between “sun” and “shade” algal species. +

3. Photoinhibition of Photosynthesis The provision of an efficient light-harvesting apparatus is only beneficial when the other processes of the cell have the capacity to use the absorbed energy. At high irradiance such a situation may not exist for a number of reasons and

164

A. W . D . LARKUM AND JACK BARRETT

dissipation of the excess energy via non-harmful processes (see, e.g. Section VI.C.8) may not be possible. The results will then be photoinhibition, defined by Jones and Kok (1966) as “The debilitating effect of high intensities of visible light upon photosynthetic capabilities of green organisms”. Photoinhibition has been found in all oxygenic photosynthetic organisms so far studied (Harris, 1980; Osmond, 1981). However its molecular basis is not fully understood and a number of effects seem to be involved. In algae the occurrence of photoinhibition has long been known (see Harris, 1980). As shown by Harris and Piccinin (1977) it is most marked in experiments using in situ incubation bottles and is less apparent where a stirred experimental technique, such as the Clark-type oxygen electrode, is used. Ultraviolet (UV) light has often been implicated as a major factor in photoinhibition (see Harris, 1980; Smith et al., 1980; Section 111). However photoinhibition has been observed in glass bottles and in deeper layers of the sea where UV light is screened out (Steeman-Nielsen, 1975; Harris, 1980). In isolated chloroplasts and lyophilised Anacystis niduluns (Cyanobacteria), Jones and Kok (1966) found evidence for the involvement in photoinhibition by both UV light and light absorbed by the major photosynthetic pigments. In higher plants photoinhibition occurs under high irradiance when C 0 2 supply is restricted and turnover of the reaction centres is reduced (Powles et al., 1979) or when absorption of light exceeds the capacity of the electron transport chain to accept reducing equivalents from the reaction centres (cf. Critchley, 1981). Under such conditions it has been suggested that damage occurs to the reaction centres (e.g. Jones and Kok, 1966). Evidence for damage to both RCI and RCII has been obtained (Kok ef al., 1965: Satoh, 1970b; Powles el al., 1979; Critchley, 1981), but in algae there is convincing evidence only for an effect on RCI (Harvey and Bishop, 1978; Gerber and Burris, 1981). It should be pointed out that bulk bleaching of photosynthetic pigments occurs mainly at higher irradiances (IO-fold higher) and after longer times than those necessary for photoinhibition and impairment to the reaction centres (Satoh, 1970a; Abeliovich and Shilo, 1972). It seems probable that photoinhibition is an important factor in determining the concentration of the light-harvesting apparatus and the optical crosssection of PSI and PSI1 units. For example, shade-adapted phytoplankton (that is, algae taken from depth) suffer rapid photoinhibition at surface irradiance (cf. Ryther and Menzel, 1959). Such shade adaptation can only be of substantial benefit in thermally stratified waters since where mixing occurs the algae will suffer from photoinhibition when they are brought into surface waters. It is probable therefore that a compromise situation exists even in shade-adapted phytoplankton. Deep-living benthic algae, on the other hand, offer the possibility for extensive shade adaptation (Section IX.C.3). It has been suggested that in higher plants there may be specific processes for reducing photoinhibition (Powles and Osmond, 1978; Osmond, 1981).


165

One suggestion is that photorespiration is a mechanism for maintaining electron transport when CO, supply is limited (Powles et al., 1979; Osmond, 1981). Another is that the Mehler reaction (the flow of electrons through the photosynthetic electron transport chain to oxygen, forming hydrogen peroxide, instead of NADPH) also maintains electron transport under similar circumstances (Osmond, 1981). The extent of photorespiration in algae is an open question but most workers now agree that photorespiration does occur (cf. Harris, 1980). Harris and Piccinin (1977) obtained results which were consistent with significant photorespiration in phytoplankton at high irradiance and suggested that this was a stress response of algae to high irradiance. The presence of such protective processes in algae would permit a much higher concentration of the light-harvesting apparatus and electron transport components. X. CHROMATIC ADAPTATION A. HISTORICAL ASPECTS

There is no doubt as discussed in Section IX that algae show chromatic adaptation; that is, both the levels of photosynthetic pigments and their ratios to one another change under different light regimes. However for historical reasons chromatic adaptation has been used to describe the much more narrowly defined hypothesis of complementary chromatic adaptation. Since this hypothesis was first put forward in the infancy of photosynthetic studies it is necessary to understand what was meant by it in its original context and then to assess it in the context of present knowledge. Engelmann ( 1883, 1884)in a classic series of experiments obtained evidence for the role of pigments other than Chl in the photosynthesis of algae of various groups (see Section V1.D). The question then arose as to why different groups of algae should produce different sets of photosynthetic pigments. It is a question which even today begs a definitive answer. The answer by Engelmann was that the algal groups were pigmented to suit the quality of light which predominated in the environment in which each group lived. As a result the hypothesis of complementary chromatic adaptation arose, though not formally set out before Gaidukov (1902). Work at that time, showed that in some algae the ratios of the photosynthetic pigments responded quickly to changes in growth irradiance (ontogenetic effects) e.g. Cyanobacteria, whereas in other algae, the pigment ratios were invariable and therefore phylogenetically determined. Two types of adaptation were therefore possible, (i) ontogenetic adaptation, where algae adapt to changes in the spectral quality of light in a particular environment by specific and complementary pigment changes and (ii) phylogenetic adaptation where the optimal photo-

166


synthesis and therefore the habitat of an alga is determined by the spectral characteristics of particular depths in the ocean. B. ONTOGENETIC COMPLEMENTARY CHROMATIC ADAPTATION

The hypothesis of ontogenetic complementary chromatic adaptation was tested by Gaidukov (1903, 1906) and Boresch (1919) who showed clearly that several Cyanobacteria responded to light of various colours by producing an overall pigmentation approximately complementary to the colour of the incident light. Similar experiments by many workers (see Rabinowitch, 1945) including Halldal (1958) have confirmed these findings. However, an alternative hypothesis of “intensity adaptation” was put forward by Berthold (1 882) and Oltmanns (1892), who ascribed changes in pigmentation as solely due to light intensity. A number of workers have shown that pigment changes of a similar kind to those induced by colour changes can be induced by changes in growth irradiance or temperature or nutrient levels (see Rabinowitch, 1945). Halldal(l958) who investigated the effects ofcrossed gradients of light quality and quantity was unable to decide whether pigment changes could be ascribed solely to complementary chromatic adaptation in Anacystis nidulans (Cyanobacteria). A clear case of complementary chromatic adaptation occurs in some, but not all, Cyanobacteria. Fujita and Hattori (1959) showed that reversible changes in the amounts of phycoerythrin and phycocyanin could be induced by green or yellow light in Tolyporhrix tenuis. Bennett and Bogorad (1973) established this fact in quantitative terms, and recently such complementary changes in the ratio PE to PC have been correlated with changes in phycobilisome rods in Synechocystis 6701 (Bryant et al., 1979; Williams et al., 1980). In Pseudanabaena 7409 red light suppresses the formation of PE and specifically induces an extra pair of PC subunits which are chemically distinct and are the products of different genes from those for the pair of PC subunits produced in green light (Bryant and Cohen-Bazire, 1981). Tandeau de Marsac (1977) investigated complementary chromatic adaptation in 44 strains of Cyanobacteria and found that 12 strains did not adapt chromatically (i.e. there were fixed proportions of PE, PC and APC) while 7 strains showed adaptation by variation of PE alone and 25 strains showed variation of both PE and PC. Bryant (1981) confirmed and extended these observations to 69 more strains of cyanobacteria. Thus ontogenetic complementary chromatic adaptation has been shown to occur unequivocally in a number of Cyanobacteria. In Rhodophyta such adaptation does not occur (Bogorad, 1975). Apparently in the evolutionary development of eukaryotes from prokaryotes the ability to vary the ratio of PE to PC was lost; in the Cryptophyta, only PC or PE occurs in any one species, with one possible exception (Gantt, 1979).


I67

The absence of specific complementary ontogenetic adaptation in the ratio of PE to PC in Rhodophyta has not prevented the formulation of general proposals for complementary adaptation in Rhodophyta and other phyla, on the basis of physiological or pigment adaptation (Ramus et al., 1976; Ley and Butler, 1980a,b). Such proposals are very difficult to test since it is necessary to show that specific complementary changes take place and the experimental attempts are confounded by the presence of light intensity, as well as light quality, effects. The observation that in some benthic algae lightharvesting pigments increase with depth is not proof of complementary chromatic adaptation since light quantity as well as light quality changes with depth. It is well established (Section 1X.D)that shading causes an increase in the amount of light-harvesting pigment proteins e.g. LHCP, peridinin-Chl-protein, fucoxanthin-Chl-protein and phycobiliproteins. Unfortunately these are also the pigments which would increase under complementary chromatic adaptation. Lee and Titlyanov (1978) have shown that similar effects to those found at depth also occur when benthic algae (Chlorophyta, Phaeophyta, and Rhodophyta) are placed in the shade of caves, where light quantity but not light quality is affected. It must be concluded that ontogenetic complementary chromatic adaptation has been demonstrated unequivocally only in certain Cyanobacteria. C. PHYLOGENETIC COMPLEMENTARY CHROMATIC ADAPTATION

1 . The Hypothesis In reviewing this hypothesis in 1945, Rabinowitch stated “The concept of phylogenetic adaptation of plants to the prevailing intensity and colour of light has its origins in observations of the vertical distribution of marine algae which is characterized by the predominance of the green Chlorophyta in shallow waters and of the red Florideae in deep waters, with the brown Phaeophyta in an intermediate position”. The concept originates in Engelmann (1883, 1884) and was supported by a number of workers between 1893 and 1960 including Gaidukov (1 903, 1904, 1906)-for further references see Rabinowitch (1945) and Levring (1966). However as early as 1892, Oltmanns, citing the earlier work of Berthold ( 1 882), challenged the hypothesis, by suggesting that light intensity rather than light quality determined the vertical distribution of algae, a view which has been supported by a number of subsequent workers (Rabinowitch, 1945, who cites earlier references; Larkum et al., 1967; Lee and Titlyanov, 1978; Dring, 1981). Care must be exercised in treating the hypothesis of phylogenetic complementary chromatic adaptation, because different types of seawater have different spectral characteristics (Section 111). The original hypothesis was based on the proposal that downwelling light becomes progressively more green during its passage through the sea; and this supposition was contrasted

168


with the poor absorption properties of Chl and carotenoids for green light (see Rabinowitch, 1945).These suppositions were unjustified simplifications of the natural situation. As discussed in Section 111, the work of Jerlov (1951, 1976) and subsequent workers has shown that the colour of downwelling irradiance in natural waters varies considerably depending on the type of water-from the blue (475 nm maximum) of Type I oceanic water to the yellow-green (575 nm maximum) of Type 9 coastal water. As shown by Larkum et al. (1967) and confirmed for a greater number of algae by computer modelling (Dring, 1981) the set of photosynthetic pigments in a chlorophyte alga are at least equally as efficient as a rhodophyte alga in absorbing incident light at all depths, in Type I oceanic water. In fact at extreme depth, calculations show that Chlorophyta photosynthesise at a greater rate than Phaeophyta which in turn have a much greater rate than Rhodophyta (Larkum et al., 1967; Dring, 1981)(see Table VIII). The same is not true of the yellow-green coastal waters, where the Rhodophyta should have an advantage over the other benthic algae. It should be pointed out that the modelling of Dring (1981) (Table VIII), was based on the earlier values of Jerlov which require some modification (see Section 111) and that the simplifying assumption of equal irradiance levels was made for all wavelengths at the sea surface (an assumption not made by Larkum et al., 1967). Nevertheless it seems clear that the phylogenetic hypothesis cannot be supported on theoretical grounds, for oceanic waters. 2. Evidence f r o m Zonation

On practical grounds the evidence for the hypothesis is even less compelling TABLE VIII Predicted Photosynthetic Rate ( P r ) Based on Action Spectra for 8 Sublittoral Algae near the Photic Limit in Various Types of Seawater. Data Taken from Dring (1981). Water Type ClassiJed According to Jerlor (1976)--see Section III Water type

I

11

111

1

3

5

7

9

Chlorophyta Ulva taeniata U . lactuca

1.483 1.255 0,915 1.558 0.524 0.553 0.583 0.616 1.297 1.193 0.983 0.749 0.731 0.756 0.801 0.846

Phaeophyta Coilodesme sp. Laminaria saccharina

1.466 1.322 1.104 0.867 0.813 0.800 0.691 0.611 1.364 1.327 1.213 1.080 1.017 0.982 0.838 0.733

Rhodoph yta Porphyra umbilicus Chondrus crispus Delesseria sanguinea

0.511 0.754 1.112 1.464 1.499 1.486 1.529 1.561 0.717 0.907 1.128 1.322 1.346 1.345 1.387 1.422 0.894 1.149 1.412 1.634 1.635 1.577 1.430 1.345

Cyanoph yta Phormidium sp.

0.639 0.819 1.169 1.575 1.627 1.575 1.493 1443


169

than the theoretical evidence. As shown above it is in yellow-green coastal waters and freshwater bodies (Kirk, 1976b, 1979) that evidence for the hypothesis should be sought. For phytoplankton, mixing and seasonal succession make it difficult to draw conclusions concerning their depth preference and experimental evidence gives little support to the hypothesis (Humphrey, 1982). Freshwater bodies provide little evidence since the number of freshwater members of the Rhodophyta is very small and benthic algae are not a common feature of freshwater lakes. Therefore, the pertinent evidence is restricted to marine benthic algae in yellow-green coastal waters. Unfortunately the evidence here is very scanty. Such waters are usually very turbid and the photic zone is narrow but seasonally variable (Luning and Dring, 1979). In these waters, on rocky coasts, the kelp forest of shallower depths gives way near the photic limit to a turf community of shade algae (Kitching, 1941; Kain, 1962; Larkum, 1972; Shepherd and Womersely, 1976). The composition of this community, although abundant in Rhodophyta. also includes Chlorophyta and sometimes even Phaeophyta. Sears and Cooper (1978) have observed that Rhodophyta are the main algae at the lower limit (or extinction depth) in the temperate Western N. Atlantic Ocean. In their study, off the coast of Massachusetts, the lower limit was at 44-45 m and, although the water type was not described, it may reasonably be assumed (on the basis of 0.1-1 per cent of surface light at this level) to be oceanic water type 111. Only one study, carried out in Australia, in Westernport Bay, Victoria (Millar and Kraft, 1983), has shown a clear zone of only Rhodophyta near the lower limit of algal growth; in this case on concrete pilings of a pier. Clearly careful study of algal communities near the lower limit in coastal waters is an important area of future research.

3. Difjiculties with the Hypothesis Several considerations make the hypothesis of phylogenetic complementary chromatic adaptation difficult to formulate precisely and therefore difficult to test. The underlying premise of the hypothesis is that photosynthesis should be greater in members of the Rhodophyta compared with Chlorophyta or Phaeophyta, at least in deeper coastal waters. As discussed in Sections V and IX, all algae, no matter what phylum, can be perfect absorbers of PAR by (i) increasing the concentration of photosynthetic pigments (ii) by increasing tissue thickness or (iii) by increasing internal reflection and scattering (cf. Ramus et al., 1976a,b). All such adaptations, however, require more expenditure of organic carbon and nitrogen per quantum of absorbed light, especially in algae that do not possess phycobiliproteins. On a daily basis, it is necessary to consider primary productivity rather than photosynthetic rate alone since a critical factor in determining the survival of an alga is the daily increment in stored carbon. Gross gains in carbon fixed by photosynthesis are offset by carbon losses due to respiration, photore-

170


spiration and loss of dissolved organic carbon. Under normal conditions it is probable that respiration is the most important factor in carbon loss. Thus it is important to know whether respiratory rates differ between the various groups of algae, with particular reference to Rhodophyta. In fact many Rhodophyta do have low respiratory rates in the dark and the rates are lower for deep-living forms (Drew, 1969; Vooren, 1981) but there is evidence to suggest that other algae can have as low rates of respiration as the Rhodophyta (King and Schramm, 1976; Burris, 1977; Vooren, 1981). Nevertheless, Rhodophyta, would still possess an advantage in photosynthesis in deeper coastal waters because of their more efficient absorption of PAR per unit of pigment-protein and this should be reflected in higher rates of primary productivity and greater compensation depths. There is a certain amount of evidence to support such a conclusion (Rabinowitch, 1951; Levring, 1966; Millar and Kraft, 1982). Greater primary productivity should give a member of the Rhodophyta greater ability to grow and establish once settlement has taken place. However ecological considerations indicate that many physical and biotic factors bear upon the chances of establishment and survival of an attached plant (Harper, 1977). This is especially true in the comparatively shallow sublittoral benthic environment of coastal waters (cf. Dayton, 1975; Littler and Littler, 1980; Hay, 1981). Temperature, wave action, siltation, herbivorous activity as well as light quality and quantity all vary so greatly that the conditions under which the single factor of higher primary productivity will outweigh all other factors will be rare. Other factors such as reproductive activity, dispersal, ability to withstand wave-action or herbivory must often determine what benthic algae are present near the lower limit of the photic zone. Rhodophyta are known to predominate in littoral and sublittoral caves. In three careful studies of algal zonation in caves the only algae found at the limit of algal growth were macroalgal members of the Rhodophyta (Dellow and Cassie, 1955; Larkum et al., 1967; Norton etal., 1971).Under these conditions light intensity is the major variable; light quality in surface or shallow sublittoral caves presumably changes little although the spectral characteristics of light in caves need further study. Therefore the advantage which Rhodophyta enjoy here is consistent with their broad light-harvesting properties and low compensation points. Although Cyanobacteria are present in marine caves (see e.g. Dellow and Cassie, 1955) they are not found at the limit of algal growth. This contrasts with the situation in aerial caves where freshwater Cyanobacteria occupy the most shaded habitats (Section XI.D.2). Apparently marine Cyanobacteria are not as well adapted as Rhodophyta for surviving under extreme shade. 4. Conclusion In summary, Rhodophyta are not predetermined by their phylogenetic


171

characteristics to live in the lower depths. They compete successfully with many other algae in both the intertidal region and in all zones of the subtidal region. However, their efficient light-harvesting properties throughout the visible spectrum give the Rhodophyta an extra advantage in shallow, shaded habitats, especially in yellow-green coastal waters. Cyanobacteria, which exist under extreme light-limitations in caves on land (Section IX.D.2) also enjoy a similar advantage, although much less is known of their distribution amongst marine benthic algae. As discussed in Sections IX and XII, the apparent spectral advantages of Rhodophyta and Cyanobacteria are probably reduced by the spatial constraints of a phycobilisome arrangement. D. OTHER TYPES OF CHROMATIC ADAPTATION

1. Distribution of Excitation Energy Ontogenetic complementary chromatic adaptation as usually defined involves only the production of photosynthetic pigments which complement the colour of incident irradiation. Myers and coworkers (Jones and Myers, 1965; Wang et al., 1977; Myers et al., 1978; Myers et al., 1980) and Ghosh and Govindjee (1966) working on Anacystis nidulans (Cyanobacteria) and Ley and Butler (1976, 1977a,b, 1980a,b) working on the unicellular rhodophyte, Porphyridium cruentum have investigated adaptation involving redistribution of energy to the two photosystems once light has been absorbed. In both algal species growth under light which is predominantly absorbed by phycobilins (green or yellow light), and under light predominantly absorbed by Chl a, causes marked changes in the amount of Chl in each photosystem and in the distribution of energy from the phycobilisomes. In such experiments it should be remembered that the experimental conditions where light is predominantly absorbed by Chl a is not a situation likely to be encountered in the natural environment. Even in the deep ocean where light is mainly blue the peak is at 475 nm where absorption would be shared between carotenoids, phycoerythrin B (in Floridexe, and much less by other phycoerythrins in other Rhodophyta and Cyanobacteria) and to a smaller extent by Chl a (see e.g. Jones and Myers, 1965).In shorter wavelength violet light (436 nm) or in red light (670 nm), absorbed maximally by Chl a, a peculiar phenomenon occurs which has been called counter complementary chromatic adaptation, in which large amounts of phycobilins are produced (Jones and Myers, 1965; Ley and Butler, 1980a). It is as if the algae receive a signal similar to that for low light intensity. However the response is different from that induced by low levels of green or white light (Ley and Butler, 1980a): in all treatments light absorbed by phycobilins was transferred initially to PSII, however in cells grown in green or white light a large fraction (55 per cent) of this excitation was redistributed to PSI whereas in cells grown in red or blue light only 20-38 per cent was transferred to PSI. Furthermore Ley and

172


Butler (1980a) estimated that the amount of Chl a in PSII was much greater following red or blue growth irradiance (approx. 40 per cent of total Chl a ) as compared with green or white irradiance (approx. 5 per cent of total Chl a). Myers et al. (1978, 1980) found much smaller changes in the amount of Chl a in PSII in Anacystis nidulans. Wang et al. (1980) have also recently challenged the basis on which Ley and Butler (1980a) calculated the distribution of excitation energy from PSII and PSI (see further discussion in Section 1X.C). No clear conclusions can be drawn at present concerning light quality and the control cf the distribution of energy between the two photosystems. It is likely that the light-harvesting mechanism is influenced by growth irradiance in ways other than simple changes in the ratio and concentrations of photosynthetic pigments. Future research in this area is required and should be directed to answering questions relating to natural light fields-white, green (500-570 nm) and blue (475 nm) light-of varying irradiance levels. 2. Blue-Light Effects Blue light induces a number of changes in algae and higher plants (Sundquist et al., 1980;Senger, 1980,1982) including increase in chloroplast size or rate of division, increase in respiratory rate and increase in pigment concentration. A blue-light system, whose photoreceptor has not been identified, has been implicated in the development of these changes (Voskresenskaya, 1979; Brinkmann and Senger, 1980; Senger, 1980). Chloroplast movement and phototropic curvature in Vaucheria (Xanthophyta) is regulated by a blue light receptor (Briggs and Blatt, 1980). Recently it has been shown (Jeffrey and Vesk, 1977,1981; Vesk and Jeffrey, 1977) that a large number of phytoplankton species show large structural and pigment changes when grown under low intensity blue light (A max 480 nm) compared with the same levels of white light. Pigment content, protein content, chloroplast size and ultrastructure and photosynthetic capacity all showed large changes. The light used here was meant to approximate to that found in the clearest oceanic water (Section 111) but in fact had a much broader spectral range. Humphrey (1 982) has grown Amphidinium carterae (Dinoflagellata) and Biddulphia aurita (Bacillariophyta) under carefully defined spectral conditions, including violet light (A max 435 nm), blue-green light (A max 510 nm) and green light (A max 535 nm). Of all the spectral bands (excluding white light) these algae grew and photosynthesized best under violet light and next best under blue-green light. At these wavelengths their pigment contents or structural features were not greatly changed, at the growth irradiances used (80 p E m s - I ) , although marked changes were observed in green and red light. Clearly more work is needed on a broad range of algae under defined light conditions before any firm conclusions can be drawn.

-'


173

XI PHOTO-CONTROL O F BIOSYNTHESIS O F LIGHT-HARVESTING PROTEINS It is clear from Sections IX and X that algae are able to adjust their photosynthetic apparatus according to the quality and intensity of incident irradiation. Little is known of the detector systems involved or of the mechanisms for the control of Chl or protein synthesis, despite some detailed studies with algae (Senger, 1980). Phytochrome is present in a number of Chlorophyta and may also be present in other groups (see review; Bjorn, 1979), although the latter has not been established beyond a doubt (Mohr, 1980). In higher plants phytochrome has been implicated in both Chl synthesis and its destruction (cf. Mohr, 1980). Ape1 has shown that phytochrome in barley is involved in the formation of LHCP, through a specific effect on the mRNA of LHCP (Apel, 1979; Ape1 and Kloppstech, 1980) and the translatable mRNA for the NADH: protochlorophyllide oxidoreductase (Apel, 1981). The lack of convincing evidence for the presence of phytochrome in many algal groups, together with the clear demonstration of photomorphogenetic effects of blue light (cf. Senger, 1980) has stimulated research into the blue light detector systems (see Brinkmann and Senger, 1980; Hase, 1980; Schiff, 1980; Song, 1980). A difficulty with the hypothesis of phytochrome-mediated control systems for many algae living in deep water concerns the apparent lack of sufficient red light to activate the phytochrome system. This assumption has recently been challenged (Duncan and Foreman, 1980) and needs more investigation. A further difficulty concerns the mechanism of ontogenetic complementary chromatic adaptation in Cyanobacteria. Here the detector system must distinguish between green and orange to red light which a phytochrome system cannot do (PRmax, 660 nm, P, max 730 nm). For these reasons a search for other detector substances had been made by a number of workers. Moreover since the phytochrome chromophore is closely related to phycobilin chromophores, other phycobilin-type substances have seemed likely candidates. Action spectra have been used to try to identify a photoreversible pigment in a number of algae. For the detector system in Cyanobacteria the experimental procedure of Fujita and Hattori (1959, 1962)has been followed. The procedure involves pretreatment with strong white light in a medium lacking nitrogen sources, which causes a drastic decrease in the levels of PE and PC. The algae are then given the experimental irradiation followed by incubation in darkness in a complete medium with carbon and nitrogen sources, when resynthesis of phycobilins is assessed. Fujita and Hattori (1959, 1962) found that 541 nm irradiation was most effective for promoting PE production and 641 nm irradiation was most effective for promoting PC

174


synthesis in Tolypothrix tenuis. In this alga and other cyanobacteria photocontrol continues in the dark long after illumination (Ohki et al., 1980). Diakoff and Scheibe (1973) found maxima at 550 nm and 660 nm in the same organism with smaller responses at 350 and 360 nm. In Fremyella diplosiphon (Vogelmann and Scheibe, 1978) a similar action spectrum has been established although there is some disagreement as to whether PC production may be stimulated by blue light (Haury and Bogorad, 1977). These results indicate that a photoreversible pigment called adaptochrome (Scheibe, 1972) with photoaction maxima at about 550nm and 650nm is involved in photocontrol of phycobilins. Scheibe (1972) found the A fraction of an extract of T. tenuis to increase at 650 nm and decrease at 520 nm when irradiated with green light and these effects were reversible in red light. Bjorn and Bjorn (1976) and Bjorn (1978) identified a number of “phycochrome” substances in Cyanobacteria (Tolypothrix luridum, Nostoc muscorum, Anacystis nidulans) which had photoreversible effects in green or red light. These results have been reviewed by Bjorn (1979). Phycochrome b(A,,, 570 nm) and phycochrome d(A,,, 650 nm) have been found in phycoerythrocyanincontaining cyanobacteria (Bjorn, 1980), and it was suggested by Bjorn that phycochrome b-type absorbance changes are due to changes in the a-subunit of PEC. The technique of Bjorn and Bjorn has been extended by Ohad et al. (1979, 1980) who suggest that phycochrome is an allophycocyanin which shows photoreversible changes at 620-640 nm and 547 nm. The relationship of phycochrome to adaptochrome has not been resolved. Ohki and Fujita (1979) and Ohki et al. (1980) have recently concluded that although phycochromes exist and are probably allophycocyanins they are not present under all the conditions when complementary chromatic adaptation is known to take place. Photoreversible reactions of C-phycocyanin from Synechococcus sp. (Cyanobacteria) in model systems have been reported (Bekasova et al., 1981; De Kok et al., 1981). A membrane-bound pigment-protein supracomplex containing PEB and PUB, with absorption maxima at 568 and 490nm has been isolated from chloroplasts obtained from a wide range of brown seaweeds (Barrett, in preparation). This is a putative candidate for a regulatory photoreceptor in these algae. XII. EVOLUTIONARY ASPECTS A. EVOLUTION OF PHOTOSYNTHESIS

I. Introduction It is likely that anoxygenic photosynthesis arose in prokaryotes very early in the evolution of life on the earth in an atmosphere which initially was anaerobic (Walker et al., 1982). There is good evidence that oxygenic


175

photosynthesis emerged about 3.8 billion years (Schidlowski et al., 1979; Schidlowski, 1980), that is only 700 million years after the formation of the earth. Few geological clues exist at present concerning the course of early photosynthetic evolution, but the disciplines of molecular genetics and biochemistry are currently providing much evidence on the possible course of events (Schwartz and Dayhoff, 1978; Simionescu et al., 1978; Mauzerall, 1978; Fox et al., 1980; Luehrsen et al., 1981; Langridge, 1982).Given the widespread occurrence of photosynthetic phenotypes among Eubacteria, Fox et al. (1980) suggested that the ancestral phenotype for Eubacteria was photosynthetic. This would place the evolution of anoxygenic photosynthesis as probably very closely following the evolution of the common ancestor for all bacteria. As mentioned above oxygenic photosynthesis probably emerged about 3.8 billion years ago. Stromatolites with fairly advanced forms of structural complexity have been found in the 3.5 billion year old Warrawoona groups of the Pilbara Block, Western Australia (Dunlop et al., 1978; Walter etal., 1980). Although these stromatolites do not conclusively prove the existence of photosynthesis at that time, they are consistent with the evidence from geological and atmospheric changes (Schidlowski, 1980) and other evolutionary evidence. Despite the evidence for the early evolution of oxygenic photosynthesis it is probable that the earth’s atmosphere remained largely anaerobic until about 2 billion years ago (Cloud, 1976; Schidlowski et al., 1979) at which point the rate of supply of reducing substances, e.g. sedimentary carbon and sulphur (Garrels and Lerman, 1981) was overtaken by the rate of production of molecular oxygen by water-splitting photosynthetic organisms. The oxygen level in the primitive atmosphere is estimated at between l o p 2 and l o p 3 present atmosphere levels (Walker, 1978; Carver, 1981).Carbon dioxide levels are estimated to have been higher than at present (Owen et al., 1979; Carver, 1981) and an early hydrosphere is probable (Henderson-Sellers and Cogley, 1982). The pH of the oceans has, however, remained more or less constant over geologic time (Walker et al., 1982). Eukaryotic organisms, including algae, arose less than 2 billion years ago (Cloud, 1976) coinciding with the transition to an aerobic atmosphere. The evolution of eukaryotic organisms would have been protected by a Precambrian ozone shield (Carver, 1981). Thus the evidence suggests that for at least 1.8 billion years, one third of the total duration of life on earth, oxygenic photosynthetic prokaryotes existed in the absence of eukaryotic competitors. Much diversification of phenotypes could have taken place during this period. 2. Evolution of the Reaction Centre The harnessing of light energy to drive chemical reactions, which by evolutionary selection became biologically significant, occurred at a very early

176


stage in the surface chemistry of the juvenile earth (Cammack et al., 1981; Cloud, 1982). Preceding the most primitive form of photosynthesis was possibly the generation of ATP by light-driven proton pumping similar to that utilizing bacteriorhodopsin in the archebacterium Halobacterium cutirubrum (El-Sayed et af., 1981). This system is relatively simple compared to that generating ATP in the most primitive anaerobic photosynthesizer, where not only is light translated to chemical energy but molecular C 0 2 must be assimilated. The appearance of water-splitting photosynthesis requiring PSII as well as PSI with consequent elaboration of membrane structure, added another order of complexity. The capability of certain cyanobacteria and heterocysts to photosynthesize in the absence of PSII (Section X1I.C; Kerfin and Boger, 1982) suggest that PSI evolved earlier than PSII. Granick (1949,1957) suggested that the earliest form of photosynthesis was built around the octa-carboxylic uroporphyrin, as the Zn or Mg complex. Iron ( 5 per cent in the earth’s crust) does not have photoactive complexes. Ca, K and Na though giving fluorescent species are readily hydrolysed in water; Mg complexes are photochemically but not catalytically active. Uroporphyrin is the first porphyrin in the natural series, being formed from the condensation of four molecules of porphobilinogen. Strong evidence points to it being amongst the earliest organic molecules on the Earth. Light facilitates condensation of the four porphobilinogens, and so do metals such as Cu and Zn. Magnesium, probably because of its hydration shell, is not known to do so. Consequently, it is likely that a simple Zn-porphyrin, having the right fluorescenceproperties preceded Mg-porphyrin as the photochemical reactive species in the primitive photoreaction centre. The photo-driven separation of charge in the presence of suitable reductants could generate ATP (cf. Mauzerall, 1978). Evolution of the primitive photosynthesis reaction centre would have required availability of the aromatic amino acids, essential for formation of hydrophobic zones around the reaction-centres, and of histidine as a coordination species for establishing efficient geometry in the metalloporphyrin-protein complex. These amino acids are thought not to have been abundant in the early stages of biotic evolution. A critical step in the evolution of photosynthesis, especially oxygenic, was the development of devices to cope with the toxicity of oxygen, either as the molecular species or as the triplet state. The principal enzyme for disposing of molecular oxygen is superoxide dismutase [SOD] a metallo-protein, having Fe3 +,Mn3 or Cu/Zn, according to species (Okada et al., 1979; Cammack et al., 1981). Fe3 SOD is found in photosynthetic bacteria and some cyanobacteria and eukaryotic algae. Mn3 SOD is also found in cyanobacteria, algae and mitochondria. Cu/Zn SOD is present in higher plants and certain chlorophycean algae, but rarely in prokaryotes. Selection of Fe3 or Mn3 by the progenitors of today’s blue green algae may represent differences +

+

+

+

+


177

in availability of these metals in the habitats of the ancient and primitive oxygenic prokaryotic algae. Polyenes are thought to have arisen later than the porphyrins, though derivatives of the polyenes are found in present day Archaebacteria (Ragan and Chapman, 1978; Chapman and Ragan, 1980). Cyclization of phytoene derivatives is ancient, occurring in the green sulphur bacteria. The introduction of double bonds into the phytoene chain conferring photoprotective capacity was probably a prerequisite for the evolution of oxygenic photosynthesis, leading to the ubiquitous presence of &carotene for this purpose. The question of the evolutionary precedence of Chla and BChl is unresolved. Mauzerall(l973,1978) has pointed out that the oxidation level of the two tetrapyrroles is the same despite one being a dihydro- and the other a tetrahydro-porphyrin. BChl is seen to be especially adapted to efficient photosynthesis where the redox properties of BChl and its primary electron acceptor, whether BChl or bacteriopheophytin, are best fitted (Olson, 1981a,b). In contrast the redox span inherent in Chl a is sufficient to provide the electromotive potential to split water, or conversely the combining of the nascent oxygen atoms (Mauzerall, private communication: cf. also Olson 1980, 1981a,b). The absence of BChl in the oxygenic line of photosynthetic organisms can also be attributed to the greater likelihood of this tetrahydroporphyrin, compared to Chl a, being photoxidized if free oxygen was present in proximity to the photoreaction centres. Compared to the biosynthesis of Chl a extra steps are required for BChl: the stereospecific addition of two hydrogens to ring B of the macrocycle and oxidation of the 2-vinyl group. The biosynthesis of the acetyl group would be more efficient where molecular oxygen is available. Significantly although BChl is present in the reaction-centres, the dominant Chl in the strict anaerobes (green photosynthetic bacteria), is Chlorohium Chl650 or Chl660, essentially similar in the structure of the macrocycle to Chl a, whereas in the facultative photoaerobes, only BChl is present. In the surface waters of their natural habitat, which permit the gathering of far-red light, total exclusion of molecular oxygen from the facultative photoaerobes could not occur. The presence of cytochrome oxidase in the facultative aerobes also points to these photosynthetic bacteria emerging at a later stage of evolution. B. EVOLUTION OF PHOTOSYNTHETIC PIGMENTS

At the time of ancestral photosynthesis the sun was colder (Walker et al., 1982) with a luminosity about 25 per cent less than now (Newman and Rood, 1977)and a somewhat higher red component in the incident light (Section 111). The UV light content however would have been greater, either because of the absence of an ozone layer or because of high UV radiation from the juvenile

178


sun (Canuto et al., 1982). Since these conditions were probably unfavourable for the further development of photosynthesis, the early photosynthetic organisms may well have moved away from the zone of photodestruction into more shaded habitats, where there would have been evolutionary pressure for the development of light-harvesting accessory pigment systems. Some insight into evolution of these accessory pigment systems is gained by consideration of the chemistry of the pigments. The Chromophyta are the only group of algae in which polyoxygenated polyenic pigments (xanthophylls) are used extensively. An aerobic atmosphere, at least within the illuminated cells, would have been required for their emergence as a major light-harvesting component of chloroplasts (Ragan and Chapman, 1978; Chapman and Ragan, 1980;Nes and Nes, 1980).Their widespread occurrence in marine algae has occurred in close association with Chlc, since in Phaeophyta a supracomplex is found composed of units of Chl a and Chl c2 and units of Chl a and fucoxanthin (Section VII1.C). Chl c2 is not always associated with fucoxanthin. In Cryptophyta the Chl c2 is in a membranebound complex comprising Chls a + c2 (Section VII1.Q while the phycobiliproteins, which replace fucoxanthin, are located in the intrathylakoid space. This may indicate that Chls c preceded the evolution of polyoxygenated xanthophylls. Chlsc, and c2, Mg-porphyrins, are relatively easily derived from Mgprotoporphyrin the stem precursor for all chlorophylls, and which Granick (1957) has suggested to have preceded Chl a or BChl as the photoreactive species in the reaction centres. The more abundant Chl c2 in its biosynthetic sequence does not lose the 4-vinyl, as happens with Chls a, b and cI, and the BChls. But the major simplificationis the absence of a phytyl or farnesyl ester of the 7-acrylic side-chain in Chls c, which has the important consequence of conserving twenty C I units for carotenoid formation. The evolution of Chl b, since it has a formyl group, may not have been favoured in the early development of photosynthetic organisms. The haem a of cytochrome oxidase, which is present in all eukaryotic mitochondria, requires molecular oxygen for its formation (Lemberg and Barrett, 1973) and this is probably so for the formyl of Chl b. Consequently, we advance the thesis that Chls cI and c2, which are similar in their absorption to Chl b in the blue region of the spectrum but lack a formyl group and thus a biosynthetic oxygen requirement, evolved earlier than Chl 6. Phycobilins, linear tetrapyrroles, may have arisen from the photooxidation of Mg-porphyrins (Barrett, 1968; Hudson and Smith, 1975), but as photosynthesizing organisms adapted to deeper waters, not only was the requirement for phycobilins increased to harvest the dim light, but the need for a dark pathway of biosynthesis would have exerted evolutionary pressure to develop the catalytic oxidation of haem rather than a Mg-porphyrin. This dark biosynthesis of phycocyanobilin is found in present day Cyanobacteria and


179

eukaryotic algae (Troxler, 1972). Molecular oxygen is required for the opening of the haem ring (King and Brown, 1978) and for formation of the phycobilins (Troxler et al., 1979) and this also argues against the early appearance of the phycobilins. Unlike pigments based in the porphyrin ring which are inherently rigid and thus can fluoresce, linear tetrapyrroles must be attached firmly to a polypeptide in order to exhibit significant fluorescence and therefore could hardly have any photo-function in a free state as would have been possible with porphyrins. The covalent attachment of the phycobilins to apoprotein, and the adoption of the correct conformation of the linear tetrapyrrole to give the required spectral species (A,,, ranging from 560-660 nm) represents a sophisticated chemical evolutionary sequence. Because of the need in excitation transfer (Section 1X.A) for overlap between the fluorescence and absorption spectra of Chl a and a photoaccessory pigment, phycobilin evolution logically would have proceeded from phycocyanin (yellow-light absorber) to phycoerythLin (green-light absorber). The excitation transfer may also have developed along the line exemplified in Cryptophyta, where Chl c2 is possibly an intermediary excitation transfer species (Section VII1.B). In either case phycoerythrin appears to be a late evolutionary development. The phycourobilin present in B- and R-phycoerythrins which allow extension of light-harvesting into the blue-green region, probably came even later in the evolution of the phycobiliproteins, as an increase in complexity of the quaternary structure of the phycobilisome is present where phycourobilin is found (Section 1X.B). This is consistent with its presence only in the more “advanced” Rhodophyta, the Euflorideae. The presence of certain phyco-erythrins containing phycourobilin in some Cyanobacteria (MacColl, 1982) shows that PUB evolved in cyanobacteria. Thus the absence of PUB from most cyanobacteria and the Protoflorideae indicates a polyphyletic origin of the red algae. A decisive step in the evolution of pigments based on the porphyrin nucleus was the formation of the asymmetric uroporphyrin 111 instead of the symmetric uroporphyrin I. The synthesis of the former is enzyme-directed. The juxtaposition of the 6 and 7 propionic acid side chain conferred specificity of orientation not only on the derivative side chain e.g. vinyl or formyl, but also the topography of the porphyrin-protein complexes. It is likely that iron porphyrins preceded magnesium-porphyrins since iron will chelate to porphyrin in an aqueous phase, while the insertion of magnesium into porphyrins requires a more complex chemical process. The haem formed in this early phase of photosynthesis evolution would have provided the basis for an electron transport chain, and provided the immediate precursor for the dark synthesis of open chain phycobilins (Brown et al., 1980). Clostridium tetanomorphum and other obligate anaerobic Archaebacteria have no haem but contain small amounts of vitamin B,,, a nucleotide adduct

180

A.

W. D. LARKUM AND JACK BARRETT

of a cobalt-corrinoid complex, biochemically derived from uroporphyrin 111 (Bykhovsky, 1979). Presumably the early photosynthetic pigments may have been attached to such nucleotides, but the amphipathic flexibility of proteins conferred an evolutionary advantage on pigments complexed to proteins. It is the attachment of the various chromophores to specific proteins that endows the pigment-protein complexes with their special properties of light absorption and excitation transfer. It is now clear that the light-harvesting Chlproteins carrying Chl b or Chl c are very different from the inner antenna Chlprotein complexes (Section, VII1.A). However, until the polypeptides have been better resolved and then sequenced any homologies will remain unknown, but when this is done it should be a fruitful means of tracing evolutionary lines. C. EVOLUTION OF EARLY PHOTOSYNTHETIC

PROKARYOTES

It is probable that in the early stages of photosynthetic evolution light was not a limiting factor and this may explain the poor efficiency of the basic photosynthetic apparatus of all photosynthetic organisms (Section V.A). However during the 1.8 billion years between the appearance of the first oxygenic prokaryote and the evolution of eukaryotic algae, it is likely that photosynthetic prokaryotic organisms diversified and colonized a variety of habitats including shaded environments. It is probable therefore that a variety of light harvesting systems arose during this period. Evidence suggests the salinity of the oceans was much lower during this period (possibly a salinity of 10 per thousand) compared to the present salinity of 33 per thousand of the oceans (Schopf, 1980).This implies that there was a greater similarity then between freshwater and marine environments and between the corresponding organisms than is the case today. A greater availability of inorganic nitrogen sources (NO;, NO,, NH;) and phosphate in the hydrosphere is suggested by present evidence (Schopf, 1980). These conditions lead to the conclusion that following the evolution of water-splitting apparatus of photosynthesis there arose conditions for massive photosynthesis in the photic zone of Precambrian seas and freshwater bodies. Algal blooms would have arisen such as have been witnessed only rarely under eutrophic conditions in modern times. Talling et al. (1973) describes a situation in the phosphate lakes of Ethiopia which might be equivalent; here very high productivity was sustained within a short column of water: the phytoplankton suspensions were so dense that light was reduced to zero within a vertical distance of 0.6 m. Under such conditions there would be great evolutionary pressure for diversification of light-harvesting systems. Assuming that at the earliest stages only chlorophyll a and some carotenoids existed then predominantly yellow and green light would penetrate the upper algal layer. Furthermore, without


181

the later evolutionary adaptations to harvest light effectively by (i) modification of cell shape (Section V.D. 1) or (ii) increased light scattering (Section V.G.4), the amount of green and yellow light would be greatly enhanced. Thus the conditions for evolutionary development of systems to harvest green and yellow light would be optimal. Other ecological systems which may be similar to those found in Precambrian times are salt flats (Reimer et al., 1979; Bauld et al., 1980)and stromatolites (Avramik et a/., 1976; Parker et ul., 1981);in both these systems prokaryotic organisms predominate and light is filtered rapidly. No convincing evidence exists as to which prokaryotic oxygenic organisms evolved first, although chemical considerations (discussed in Subsection B) serve to limit the possibilities. It is possible that filling of the "green window" began on either side (blue-green and orange regions) with the evolution of Chlsh and c, and carotenoids such as fucoxanthin and peridinin. The evolution of Cyanobacteria with phycobilisomes may have been a relatively late development as the evolutionary pressure for light-harvesting reached its climax (Subsection B). Evidence from amino-terminal sequences of phycobiliproteins and immunological studies point to these pigment-proteins being descended from a common ancestral gene (Glazer and Apell, 1977; MacColl and Berns, 1979; Glazer, 1980). Troxler et al. (1981) have argued from a matrix comparison of the complete LY and B amino acid sequences of phycocyanins and allophycocyanins from the rhodophyte Cyanidium caldarium and cyanobacteria Mastigocladus laminosus and Svnechoccus sp., that an allophycocyanin was close to the ancestral phycobiliprotein. A /?-type allophycocyanin precursor is more evident when the homology between the NH, terminal, middle and carboxyl terminal thirds of the sequences of the LX and p subunits are compared. No phycocyanin, however, is closely related antigenically to phycoerythrin, even from the same organism; but within each pigment class all members are antigenically related, whatever the taxonomic classification of the algae (MacColl and Berns, 1979). Although the evolution of pigments to harvest green and red light in Cyanobacteria and Rhodophyta has stabilized with phycoerythrin and phycocyanin, pigments whose absorption maxima are between those of the predominant phycobiliproteins have been found, e.g. phycoerythrocyanin (Section V1.C) in various cyanobacteria, and a novel phycoerythrin and phycocyanin (A,,, 640nm), in a marine cyanobacterium, Synechococcus sp. (Kursar et al., 1981). These may represent residual evolutionary intermediates. The possible early evolution of oxygenic photosynthetic prokaryotes is depicted in Fig. 55. Intracellular oxygen is extremely toxic to anaerobic organisms, so that the earliest water-splitting oxygenic algae must also have evolved mechanisms for disposing of toxic levels of oxygen. There must have been transitional oxygen tolerant amphiaerobes which selectively survived the build up of atmospheric oxygen to the level at which water-splitting prokaryotes evolved into obligate aerobes. A number of present day Cyanobacteria can photosynthesize under

I82


RHODOPHYTA

T 1

CHROMOPHYTA

eukaryote

CHLOROPHYTA

---

prokaryote

1

CYANOBACTERIA Chl a + Phycobilinr PROCHROMOPHYTE

Chl a +c

OXYGENIC PHOTOSYNTHETIC PROKARYOTE Chlorophyll 9

PHOTOSYNTHETIC PROKARYOTE Archaschlarophyll

Fig. 55. A simplified scheme for the early evolution of oxygenic photosynthetic prokaryotes and the origin of eukaryotic algae.

anaerobiosis, forming no oxygen and having PSI (for CO, fixation) coupled to H,S or other primary electron donors (Padan, 1979). Filamentous cyanobacteria belonging to Nostocales produce heterocysts which provide an anoxygenic environment; these photosynthesize using PSI only, coupled to molecular hydrogen, the biological production of which is light induced (Kerfin and BBger, 1982). Such algal systems may be in a direct evolutionary line with the ancient microaerophilic photosynthesizers. Further, certain green algae under anaerobiosis also exhibit hydrogen metabolism and this adaptation may be an evolutionary residue (Kessler, 1974). As the oxygen levels in the atmosphere climbed to near present levels, a point reached about 1 billion years ago (Cloud, 1976; Schidlowski et al., 1979), the activity of RuBP carboxylase-oxygenase (Hatch, 1976) would have increased making C3 photosynthesis less efficient. Thus after this period of favourable growth for prokaryotic oxygenic organisms, depletion of nutrients, rise in oxygen and lowering of CO, levels would lead to conditions similar to those at present where the oceans and oligotrophic lakes sustain a low algal biomass. Under these later conditions selection pressure on lightharvesting mechanisms would have virtually disappeared and the possibility for the evolution of eukaryotic sublittoral benthic algae would have been


183

enhanced and this in turn may have provided the stimulus for the development of the characean algal line leading to higher plants. D. EVOLUTION OF EUKARYOTIC ALGAE

A major question concerning the evolution of eukaryotic algae is the origin of the chloroplasts and mitochondria. One view is that they have an episomal origin (Raff and Mahler, 1972).A more widely accepted hypothesis is that of a symbiotic origin from prokaryotic progenitors (Schimper, 1885; Margulis, 1970; Raven, 1970) and convincing arguments and evidence have been put forward for such an origin (Broda, 1975; Cavalier-Smith, 1980; Wolk, 1980; Whatley and Whatley, 1981; Schiff, 1981a,b; Kiintzel and Kochel, 1981). However, it has been postulated that some eukaryotic algae have chloroplasts that are the result of at least two serial endosymbioses one of which involved a eukaryotic alga (Gibbs, 1978; 1981a,b; Whatley et al., 1979; Whatley, 1981; Whatley and Whatley, 1981). The existence of the glycine-succinate (mitochondrial) and the glutamate (plant) pathway to the biosynthesis of Saminolaevulinic acid (cf. Porra and Grimme, 1978; Beale, 1978) also supports the symbiosis hypothesis. The glutamate pathway is dominant in Cyanobacteria (Laycock and Wright, 1981; McKie et al., 1981) in Cyanidium caldarium (Troxler el a[., 1978;Troxler and Offner. 1979) and in Scenedesmus (Oh-hama et al., 1982). In the eukaryote Euglena gracilis the chloroplasts have the glutamate pathway while the mitochondria possess the glycine-succinate pathway (Beale et al., 1981). Evidence that chloroplasts and cyanobacteria have a common ancestor is the occurence of phytochrome, a phycobiliprotein, in Chlorophyta, and higher plants; a similar pigment is associated with chloroplasts of various Phaeophyta (Barrett, unpublished). The close similarity of the chemical structure of phytochrome chromophore, the binding to protein and the amino acid sequence about the chromophore (Lagarias and Rapoport, 1980) with those of phycocyanin argue for a common genetic ancestry of phytochrome and phycocyanin, rather than for a divergent evolution of these pigments. Most of the earlier work assumed a monophyletic origin of chloroplasts via the initial capture of a free-living cyanobacterium. According to this view all algae evolved from ancestral forms of Rhodophyta. There is little doubt that the chloroplasts of Rhodophyta arose from Cyanobacteria (Bonen and Doolittle, 1975; Fox et al., 1980) but no good evidence that other algae evolved from early rhodophytes (Fox et al., 1980). However, a multiple prokaryotic origin of chloroplasts as suggested by Raven (1970) has recently received support by the discovery of another photosynthetic prokaryote, Prochloron (see below) which contains Ch1 b. A comparison of 16s ribosomal RNA sequences shows no specific relationship between Prochloron and the chloroplasts of the green algae Euglena gracilis and Chlamydomonas reinhardii

184


(Seewaldt and Stackebrandt, 1982). Rather Prochloron shows highest sequence homology with cyanobacteria of several different genera. Similarly Chl c containing algae (Chromophyta) may have arisen from a prokaryote alga, Fig. 55. We therefore propose the scheme shown in Fig. 55 for the origin of eukaryotic algae (as first suggested by Raven, 1970)which is broadly similar to that of Olson (1981a,b). According to this scheme the three major lightharvesting pigment complexes phycobiliproteins, Chl a c2-fucoxanthin complex and LHCP-arose at different times from prokaryotic organisms which had Chla and an oxygenic photosynthesis (as shown in Fig. 55). However such a scheme by no means recognizes all the structural and biochemical evidence now available. Pertinent evidence at present available includes pigmentation, thylakoid arrangement, number of membranes surrounding the chloroplast, storage products, site of storage products, flagellar structure, spindle structure, DNA and RNA and their location, and polypeptide sequences of important proteins (Schwartz and Dayhoff, 1978; Dodge, 1979; Dayhoff and Schwartz, 1980; Whatley and Whatley, 1981). At present the powerful evidence based on homologies amongst DNA, 16srRNA or 5s-rRNA and proteins is as yet at an early stage but can be expected to support much stronger conclusions in the near future (cf. Fox et al., 1980; Seewaldt and Stackebrandt, 1982). Based on current evidence various schemes of algal evolution have been proposed (e.g. Taylor, 1978; Dodge, 1979; Whatley and Whatley, 1981). Interestingly, all the schemes envisage the evolution of all lines from an ancestral cyanobacterium. Those of Dodge (Fig. 56) and Whatley and Whatley (1981) incorporate proposals for multiple, serial symbioses for some groups, which can explain the existence of four membranes around the chloroplasts of chromophyte algae (and cryptophytes) and the existence of two types of chloroplasts in some Dinoflagellata. A serious limitation of all these schemes are the difficulties with the evolution of chromophytes from the Rhodophyta, i.e. the loss of phycobilins and the acquisition of Chlc. Whatever the scheme the Cryptophyta, which have both phycobilins and Chl c2, are particularly difficult to place. The phycobiliproteins of Cryptophyta are very different from those of Cyanobacteria or Rhodophyta (Gantt, 1979) implying the possibility of greater evolutionary divergence (Glazer, 1980). In particular, the presence in almost all Cryptophyta of phycocyanin or phycoerythrin only and the absence of allophycocyanin and phycobilisomes are striking features. On the other hand there is a strong similarity between 15 of the first 19 amino acid residues on the B-subunit of cryptophyte and rhodophyte phycoerythrin (Glazer and Apell, 1977) and there are also other affinities (see Gantt, 1979). The latter evidence would suggest that gene transfer has occurred during evolution but whether at an early or late stage is difficult to decide (Glazer, 1980).Dodge (1979; see Fig. 56) concluded from other characteristics, that the Cryptophyta arose rather early

+


(

Ancestral

185

)

Fig. 56. Evolutionary scheme for the diversification of eukaryotic algae based on an origin from Cyanobacteria. (Redrawn from Dodge, 1979.)

from the Rhodophyta and later gave rise to the many Chl c-containing flagellate algae (Chromophyta). Gillott and Gibbs (1980), Whatley and Whatley (1 98 1) and Greenwood (personal communication) argue for a similar evolutionary development. However, it is difficult in such a pathway to accommodate the sudden appearance of Chl c2 and then Chl c , and the equally sudden disappearance of phycobilins. It would also be difficult to explain why such an apparently successful formula as that of C h l a + c together with a phycobiliprotein and appressed thylakoids has not been more fully exploited in algal evolution. Another possibility is that the Cryptophyta appeared rather late in evolution by transfer of phycobilin genes to a chromophyte alga which did not possess Chl c, , or in which the gene for the latter Chl had been lost.

186


Prochloron (Prochlorophyta; Lewin, 1976) is the best evidence yet for a multiple, prokaryotic origin of the algae. Organisms of this genus have been found as unicellular extracellular symbionts in a number of didemnid ascidians in tropical waters (Kott, 1980). The cells of Prochloron are found in the external radial grooves or in internal cavities where they are firmly attached by mucilaginous threads (Whatley et al., 1979). For a prokaryotic organism, Prochioron species are very large, ranging in size from 8-25 pm in diameter, but there is little doubt of their prokaryotic nature (Newcomb and Pugh, 1975; Lewin, 1976; Whatley, 1977; Giddings et al., 1979; Cox and Dwarte, 1981; Seewaldt and Stackebrandt, 1982). Prochloron species contain Chl b with ratios of Chl a/b from 2.6 to 12.0 (Thorne et al., 1977). Chl b was reported to be present in LHCP (Withers et al., 1978) but Hiller and Larkum (in preparation) could find no Chl-protein complex strictly comparable with LHCP. The Chl/P-700 ratio is similar to that of higher plants (Withers et al., 1978). The carotenoids present are fl-carotene, zeaxanthin, echinenone, cryptoxanthin and mutatochrome which are neither typical of cyanobacteria nor of Chlorophyta (Withers et al., 1977; Johns er al., 1981). A possible objection to the evolution of a prokaryotic organism with Chl c is the lack of evidence for the existence of such an organism. However, the same was said of a Chl 6-containing prokaryote (Lee, 1972) until recently. We believe that evolution of the three lines of algal development at the prokaryote stage (Fig. 56) is the simplest, and therefore the best, scheme. The eukaryotic algae can then be seen to have arisen as a result of from one to three endosymbiotic stages. The Cryptophyta are seen as a more recent development from a Chromophyte algae and this accounts for the retention of the nucleomorph organelle which can be regarded as a degenerate second nucleus (Greenwood et al., 1977; Gillott and Gibbs, 1980; Gibbs, 1981b). Present evidence suggests the present day Cyanobacteria are a young group (Fox et al., 1980),coming from a very ancient stock, which may have given rise also to the chloroplasts of eukaryotic algae. Evolutionary relationships between algae may be assessed also by comparison of components of the redox chains of the photosystems. Algal ferredoxins diverged at an early stage of evolution (Matsubara et al., 1980). An unusual ferrodoxin amino acid sequence is shared by Porphyra umbilicus (Rhodophyta) and the unicellular Cyanidium caldarium, and this feature definitively places C. caldarium amongst the red algae. Plastocyanin is present in many Chlorophyta, Rhodophyta and some cyanobacteria (Crofts and Wood, 1978), but is undetectable in Euglena gracilis (Euglenophyta), the and several cyanobacteria. Cytochrome Xanthophyte Bumifleriopsisjif~oforrnis c-552 replaces plastocyanin in those algae lacking the gene for this Cu-protein or in Cu-deficient growth (Wood, 1978; Bohner et al., 1980a,b). Cu-proteins appeared early in evolution. The Cu-protein of Archaebacteria contains amino-acid sequences similar to those in plastocyanin (Cammack et al., 1981).


187

These Cu-proteins probably appeared first in organisms growing in mineralrich pools, rather than in organisms in the open sea, where Cu would have been diluted out but Fe still be abundant. Many Phaeophyta, Rhodophyta and unicellular marine algae possess a cytochrome c-552 (cf. Sugimura et al., 1981). E. EVOLUTION OF THYLAKOID STACKING

The appression of thylakoids is a feature of most photosynthetic organisms from photosynthetic bacteria to higher plants. In photosynthetic bacteria its occurrence is not common. In Cyanobacteria and Rhodophyta it is rare; this may readily be explained by the presence of phycobilisomes on the outer surface of thylakoids but where phycobilisomes are absent appression occurs in Cyanobacteria. In Cryptophyta phycobilins are present but not in the form of phycobilisomes, and appression of thylakoids occurs giving rise to the characteristic pairs of thylakoids (Fig. 13), or sometimes triplets. In all other algae and higher plants appression of thylakoids is invariably found in mature chloroplasts, although under certain conditions such as in the bundle sheath chloroplasts of some C, plants the degree of appression may be much reduced (Laetsch, 1974). There may be some variation in the inter-lamellar distance (Section V.E) although there is no evidence to suggest that such variations have any significance. The general occurrence of appressed thylakoids suggests that this arrangement serves an essential function. However no identification of such a function that is fully convincing has yet been put forward. Possible explanations can be set out as follows: (i) cooperativity between PSII units (either by lateral or trans-membrane communication) (Section IXA; Miller, 1976; Arntzen, 1978); (ii) protection of PSII and water-splitting; (iii) regulation of PSI and PSII activity (Anderson, 1981); (iv) promotion of light-scattering (Section V.G.4); (v) conservation of a proton gradient at low light intensity (Section 1X.D); (vi) spatial economy of the membrane-bound photosynthetic reactions and the soluble stroma systems (Sane, 1977); (vii) stabilization of the thylakoids where there is high cellular osmolarity, as in marine algae. Too little is known at present of the photosynthetic characteristics of many algal groups to make adequate comparisons of the various types of thylakoid arrangement. Nevertheless the Cyanobacteria, the Chlorophyta and higher plants have been studied sufficiently to show that there are no clear distinctions to be made between the photosynthetic response in organisms

188


with and without thylakoid appression (Section X1.B). Such a conclusion seems to rule out explanations (i) and (ii) for the overwhelming advantages of appressed systems and suggests that the possible advantages of (iii), (iv), (v) and (vi) could be achieved in other ways in Cyanobacteria and the Rhodophyta. Consequently, any advantage of appression must be exerted in small ways. Perhaps, some or all of the above points confer small advantages which together provide a more efficient system of thylakoid organization than that based on phycobilisomes (refer to Section 1X.C). According to the evolutionary scheme of Fig. 56 the prokaryotic oxygenic lines evolved their distinctive features between 3.8 and 2 billion years ago and presumably this also involved development of the characteristic thylakoid appression. The evidence from Prochloron supports this (Whatley, 1977; Thinh, 1978; Giddings et al., 1979; Cox and Dwarte, 1981). Is it coincidental that only Cyanobacteria have survived in free-living forms? Perhaps the rise of atmospheric oxygen levels created problems which imposed critical constraints on the other free-living ancestral algae, which were offset by these ancestors entering into symbiotic associations. Some aspect of RuBP oxygenase, e.g. the production of glycollate, may have been a major factor. Whatever the reason, Prochloron is the only prokaryotic alga known to have survived, other than Cyanobacteria. Following the evolution of the eukaryotic algae it is probable that little change in thylakoid structure occurred as shown in the scheme of Fig. 56. The only major development appears to have been in the Characean line with the development of true grana (Stewart and Mattox, 1975). This has taken place in conjunction with the development of the peroxisome and the enzyme glycollate oxidase (Raven, 1977; Floyd and Salisbury, 1977a,b; Raven and Glidewell, 1978). No adequate explanation for these developments in this single group of algae can be given as yet. Another unresolved question is why there has been no wide exploitation of the light-harvesting strategy of appressed thylakoids in association with the presence of phycobiliproteins, as evolved in the Cryptophyta. ACKNOWLEDGEMENTS We should like to thank the following who supplied manuscripts prior to publication: R. S. Alberte, Jan M. Anderson, J. Duniec, A. N. Glazer, R. G. Hiller, S. W. Jeffrey, B. R. Green. Electron micrographs were kindly provided by M. Vesk, D. Dwarte and G. Cox (Electron Microscope Unit, University of Sydney). We gratefully acknowledge the many critical discussions with our colleagues, particularly Jan M. Anderson, J. Duniec, R. G. Hiller, J. T. 0. Kirk, D. Mauzerall, H. Senger, A. Stewart and S. Thorne, some of whom read part of the manuscript. In particular we should like to thank Jan M. Anderson who


189

made many constructive comments on the manuscript. We acknowledge grants in support from the Australian Marine Science and Technology Advisory Committee (to J.B.) and from the Australian Research Grants Committee for J.B. and A.W.D.L. We thank Dr W. J. Peacock, Chief of the CSIRO Division of Plant Industry for making the facilities of the Division available to J.B. while in receipt of an Australian Research Grants Committee Senior Fellowship. We are particularly indebted to Mrs Denese McCann for her skill and patience in the preparation of our manuscript. REFERENCES Abeliovich, A. and Shilo, M. (1972). J. Bacteriol. 111, 682-689. Aizawa, M., Hirano, M. and Suzuki, S. (1978). Electrochimica Acta 23, 1185-1190. Akoyunoglou, G. (1977). Arch. Biochem. Biophys. 183, 571-580. Alberte, R. S. and Thornber, J. P. (1978). FEBS Lett. 91, 126130. Alberte, R . S., Friedman, A. L., Gustafson, D. L., Rudnik, M. S. and Lyman H. (1 98 1). Biochim. Biophys. Acra 635, 304-3 16. Allen, J. F., Bennett, J., Steinbeck, K. E. and Arntzen, C. J. (1981). Nature 291,25-29. Allen, M. B. (1966). In “The Chlorophylls” (L. P. Vernon and G. R. Seeley, eds) pp. 5 1 1-5 19. Academic Press, New York. Allen, M. B., Goodwin, T. W. and Phagpolngarm, S. (1960). J. Gen. Microbiol. 23, 93-103. Allen, M. M. and Smith, A. J. (1969). Arch. Microbiol. 69, 114-120. Alpert, B. and Lundquist, L. (1976). In “Excited States of Biological Molecules” (J. B. Birks, ed.) pp. 425433. Wiley, London. Amesz, J. and Duysens, L. N. M. (1962). Biochim. Biophys. Acta 64, 261-278. Amesz, J . and Duysens, L. N. M. (1977). In “Topics in Photosynthesis” Vol. 2, pp. 149- 186. Elsevier, Amsterdam. Anderson, J. M. (1974). Biochim. Biophys. Acta 333, 378-387. Anderson, J. M. (1975). Nature 253, 536537. Anderson, J. M. (1981). FEBS Lett. 124, 1-10, Anderson, J. M. (1982). FEBS Lett. 138, 62-66. Anderson, J . M . and Andersson, B. (1982). Trends Biochem. Sci. 7,288-292. Anderson, J. M. and Barrett, J. (1979). Ciba Foundation Symposium 61, 81-104. Excerpta Medica, Amsterdam. Anderson, J. M. and Boardman, N. K. (1966). Biochim. Biophys. Acta 112,403421. Anderson, J . M . and Levine, R. P. (1974). Biochim. Biophys. Acta 357, 118-126 Anderson, J. M., Barrett, J. and Thorne, S. W. (1981). In “Photosynthesis 111” (G. Akoyunoglou, ed.), pp. 301-3 16. Balaban International Science Services, Philadelphia, Pennsylvania, USA. Anderson, J. M., Goodchild, D. J. and Boardman, N. K. (1973). Biochim. Biophys. Acts 325, 573-585. Anderson, J. M., Waldron, J. C. and Thorne, S. W. (1978). FEBS Letr. 92, 227-233. Anderson, J. M., Waldron, J. C. and Thorne, S. W. (1980). Plant Sci. Lett. 17, 149-157. Anderson, M. C. (1966). J. Appl. Ecol. 3, 41-54. Andersson, B. and Anderson, J. M. (1981). Biochim. Biophys. Acta 593,426439. Andersson, B., Akerlund, M.-E. and Albertsson, P.-A. (1978). Biochim. Biophys. Acta 423. 122-132.

190


Antia, N. (1977). Br. Phycol. J . 12, 271-276. Antonini, E. and Brunori, M. (1971). “Hemoglobin and Myoglobin in Their Reactions with Ligands.” North Holland, Amsterdam. Apel, K. (1977a). Brookhaven Sypm. Biol. 28, 149-161. Apel, K. (1977b). Biochim. Biophys. Acta 462, 390-402. Apel, K. (1979). Eur. J . Biochem. 97, 183-188. Apel, K. (1981). Eur. J. Biochem. 120, 89-93. Apel, K. and Kloppstech (1980). Planta 150, 426430. Apel, K., Bogorad,-L. and Woodcock, C. L. F. (1975). Biochim. Biophys. Acta 387, 568-579. Argyroudi-Akoyunoglou, J. H. and Castorinis, A. (1 980). Arch. Biochem. Biophys. 200, 326-335. Argyroudi-Akoyunoglou, J. and Thomou, H. (1981). FEBS Leu. 135, 177-181. Armond, P. A., Staehelin, L. A. and Arntzen, C. J. (1977). J . Cell Biol. 73, 4 0 M 1 8 . Arnold, K. E. and Murray, S. N. (1980). J . Exp. Mar. Biol. Ecol. 43, 183-192. Arnon, D. I. (1949). Plant Physiol. 124, 1-15. Arnon, D. I., McSwain, B. D., Tsujimoto, H. Y. and Wada, K. (1974). Biochim. Biophys. Acta 357, 231-245. Arntzen, C. J. (1978). In “Current Topics in Bioenergetics” (D. R. Sanadi and L. P. Vernon, eds) 8, 1 1 1-160. Academic Press, New York. Arpin, N., Svec, W. A. and Liaanen-Jensen, S. (1976). Phytochem. lS, 529-532. Astier, C. and Joset-Espardellier, F. (1981). FEBS Letf. 129, 47-51. Atkinson, A. W. Jr, John, P. C. L. and Gunning, B. E. S. (1974). Protoplasma 81, 77-1 10. Avramik, S. M., Hofman, H. J. and Raaben, M. E. (1976). In ”Stromatolites” (M. R. Walter, ed.), pp. 149-162. Elsevier, Amsterdam. Badger, M. R., Kaplan, A. and Berry, J. A. (1980). Plant Physiol. 66, 407413. Ballschmitter, K. and Katz, J. J. (1969). J . Am. Chem. SOC.91, 2661-2667. Bar-Nun, S., Schantz, K. and Ohad, I. (1977). Biochim. Biophys. Acfa 459,451467. Barber, J. (1980). FEBS Lett. 118, 1-10, Barber, J. (1983). Phofochem. Photobiophys. 5, 181-190. Barrett, J. (1967). Nature 215, 733-734. Barrett, J. (1968). In “Structure and Function of Cytochromes” (K. Okunuki, M . D. Kamen and I. Sekuzu, Eds) pp. 701-712. University of Tokyo Press, Tokyo. Barrett, J. and Anderson, J. M. (1977). Plant Sci. Lett. 9, 275-283. Barrett, J. and Anderson, J. M. (1980). Biochim. Biophys. Acta 590, 309-323. Barrett, J. and Thorne, S. W. (1978). Intern. Union Biochem. (NZBS and ABS) Meeting on “Plant Proteins”, Christchurch, New Zealand, Abstract 25. Barrett, J. and Thorne, S. W. (1980). FEBS Lett. 120, 24-28. Barrett, J. and Thorne, S. W. (1981). In “Photosynthesis” Vol. I11 “Structure and Molecular Organisation of the Photosynthisis Apparatus” (G. Akoyunoglou, Ed.), pp. 347-356. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Bart, J. C. J. and MacGillavry, C. H. (1968). Acta Crystallogr. Sect. B24, 1569-1587. Baszynski, T., Panczyk, B., Krol, M. and Krupa, Z. (1975). Z . Pjlanzenphysiol. 74, 20C207. Bauld, J., Burne, R. N., Chambers, L. A., Ferguson, J. and Skyring, G. W. (1980). In “Biogeochemistry of Ancient and Modern Environments” (P.A. Trudinger, M. R. Walter and B. J. Ralph, Eds), pp. 157-166. Australian Acad. Sci., Canberra. Baumann, Th., Weber, G. and Grime, L. H. (1982). Phofochem.Photobiophys. 4, 1-8. Beale, S. I. (1978). Annu. Rev. Plant Physiol. 29, 95-120. Beale, S. I., Foley, T. and Dzelzkolus, V. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 16661669.


191

Beardall, J. and Morris, 1. (1976). Mar. Biol. 37, 377-387 Beddard. G . S. and Porter, G. (1976). Nature 260. 366367. Bekasova, 0. D., Bukhov, N. G. ’and Karapetyan. N. V . (1981). Biokhirniya 46, 287-295. Bell, L. N. and Merinova, G. L. (1961). Biojizika 6 , 21-26. Bendall, D. S. (1982). Biochim. Biuphy.7. Acra 683, 1 19-1 5 1 . Bennett, A. and Bogorad, L. (1973). J . Cell Biol. 58, 419. Bennett, J. (1977). Nature 269, 344346. Bennett, J. (1979). FEBS Lett. 103, 342-344. Bennett, J. (1980). Eur. J . Biochern. 104, 85-89. Bennett, J., Markwell, J. P., Skrdla, M. P. and Thornber, J. P. (1981). FEBS Left. 131, 325-330. Bennoun, P. and Jupin, H. (1 976). Biochim. Biophys. Acta 440, 122-1 30. Bennoun, P., Diner, B. A., Wollman, F.-A., Schmidt, G. and Chua, N. H. (1981). In “Photosynthesis 111” (G. Akoyunoglou, Ed.), pp. 839-850. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Bensasson, R. V., Land, E. J., Moore, A. L., Crouch, R. L., Dirks, G., Moore, T. A. and Gust, D. (1981). Nature, (London) 290, 329-332. Benson, E. E. and Cobb, A. H. (1981). ”Photosynthesis VI” (G. Akoyunoglou, Ed.), pp. 429434. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Beudeker, R. F. and Kuenen, J. G. (1981). FEBS Lett. 131, 269-274. Benz, J. and Rudiger, W. (1981). Z . P’anzenphysiol. 102, 95-100. BerkalofT, C. and Duval, J. C. (1977). In “Proceedings 4th Int. Congress on Photosynthesis” pp. 29-30. Biochem. SOC.,London. Berkaloff, C. and Duval, J. C. (1981). Photosynth. Res. 1, 127-135. Berkaloff, C., Duval, J. C., Jupin, H., Chrissovergis, F. and Caron, L. (1981). In “Photosynthesis 111” (G. Akoyunoglou, Ed.), pp. 429434. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Berthold, D. A,, Babcock, G. T. and Yocum, C . F. (1981). FEES Lett. 134,231-234. Berthold, G. (1882). Mitt. Zool. Sta. Neapel. 3, 393-536. Bialek, G . E., Horvath, G., Garab, G. I . , Mustardy, L. A. and Faludi-Daniel, A. (1977). Proc. Natl. Acad. Sci. U . S . A .74, 1455-1457. Birks, J. B. (1976). “Excited States of Biological Molecules”. Wiley, New York. Birks, J. B. (1970). “Photophysical Aromatic Molecules’’. Wiley-Interscience, New York. Bishop, N . I. (1961). Photochem. Photobiol. 6, 621-628. Bishop, N. I. and Oquist, G. (1980). Physiol. Plantar 49, 477486. Bishop, N. I . and Senger, H. (1971). Methods of Enzymology Vol. XXIII Part A, 338-351. Academic Press, New York. Bjorn, G. S. (1978). Physiol. Plantar. 42, 321-323. Bjorn, G. S. (1980). Physiol. Plantar. 48, 483485. Bjorn, G. S. and Bjorn, G. 0. (1976). Physiol. Plantar. 36, 297-304. Bjorn, L. 0. (1979). Q . Rev. Biophys. 12, 1-23. Bjornland, T. and Tangen, K. (1979). J . Phycol. 15, 457463. Boardman, N . K. (1977). Annu. Rev. Plant Physiol. 28, 355-377. Boardman, N. K. and Thorne, S. W. (1971). Biochim. Biophys. Acta 253, 222-231. Boardman, N. K., Thorne, S. W. and Anderson, J. M. (1966). Proc. Natl. Acad. Sci. U.S.A. 56, 586593. Boardman, N. K., Bjorkman, O., Anderson, J. M., Goodchild, D. J. and Thorne, S. W. (1975). In “Proc. Third. Intern. Congr. Photosynth” (M. Avron, Ed.), pp. 1809-1 827. Elsevier, Amsterdam. Boardman, N. K., Anderson, J. M. and Goodchild, D. (1978). In “Current Topics in

192


Bioenergetics” (D. R. Sanadi and L. P. Vernon, Eds) 8,36109. Academic Press, New York. Boczar, B. A., Prezelin, B. B., Markwell, J. P. and Thornber, J. P. (1980). FEBS Lett. 120, 243-247. Bode, V. C. and Hastings, J. W. (1963). Arch. Biochem. Biophys. 103, 488-499. Bogorad, L. (1975). Annu. Rev. Plant Physiof. 26, 369-401. Bohner, H. and Boger, P. (1978). FEBS Lett. 85, 337-339. Bohner, H., Bohme, H. and Boger, P. (1980). Biochim. Biophys. Acta 592, 103-1 12. Bohner, H., Merkle, H., Kroneck, P. and Boger, P. (1980). Eur. J . Biochem. 105, 603-610. Bold, H. C. and Wynne, M. J. (1968). “Introduction to the Algae” Prentice Hall, Englewood Cliffs, New Jersey, USA. Bonaventura, C. J. and Myers, J. (1969). Biochim. Biophys. Acta 301, 227-248. Bonen, L. and Doolittle, W. F. (1975). Proc. Natl. Acad. Sci.U.S.A. 72,2310-2314. Bonnett, R., Mallams, A. K., Spark, A. A., Tec, J. L., Weedon, B. C. L. and McCormick, A. (1969). J . Chem. Soc. C , 429-454. Bonnett, R., Davies, J. E., Hursthouse, M. B. and Sheldrick, G. M. (1978). Proc. R. SOC.Ser. B. 202, 249-268. Boresch, K. (1919). Ber Deut. Botan. Ges. 37, 25-39. Borisov, A. Yu and Ilina, M. D. (1973). Biochim. Biophys. Acta 305, 364-371. Boucher, F., van der Rest, and Gingras, G. (1977). Biochem. Biophys. Acta 461, 339-347. Bouges-Bocquet, B. (1980). Biochim. Biophys. Actu 594, 85-103. Bourdu, R. and Lefort, M. (1967). C . R . Acad. Sci. (Paris) 265, 37. Boxer, S. G. and Closs, G. L. (1976). J. Am. Chem. SOC.98, 5406-5408. Breton, J. (1976). Biochim. Biophys. Acta 459, 66-76. Breton, J. and Geacintov, N. E. (1979). Ciba Found. Symp. ( N S ) 61, 217-236. Breton, J. and Geacintov, N. E. (1980). Biochim. Biophys. Acta 594, 1-32. Briggs, W. R. and Blatt, M. R. (1980). In “The Blue Light Syndrome” (H. Senger, Ed.), pp. 261-268. Springer Verlag, Berlin. Brinkman, G. and Senger, H. (1980). In “The Blue Light Syndrome” (H. Senger, Ed.), pp. 526-540. Springer Verlag, Berlin. Brinkman, G. and Senger, H. (1981). In “Photosynthesis” Vol. 111 (G. Akoyonoglou, Ed.), pp. 337-346. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Britz, S. J. and Briggs, W. R. (1976). Plant Physiol. 58, 22-27. Britz, S. J., Pfau, J., Nultsch, W. and Briggs, W. R. (1976). Plant Physiol. 58, 17-21. Brockman, H. (1968). Angew. Chem. 80, 233-234. Broda, E. (1975). “The Evolution of the Bioenergetic Process”. Pergamon Press, Oxford. Brody, M. and Brody, S . S. (1962). Arch. Biochem. Biophys. 96, 354359. Brody, M. and Emerson, R. (1959a). J. Gen. Physiol. 43, 251-264. Brody, M. and Emerson, R. (1959b). Am. J. Bor. 46, 433-440. Brooks, C. and Chapman, D. J. (1972). Phytochemistry 11, 2663. Brown, A. P.(1981). Photochem. Photobiol. 34, 207-214. Brown, A. S., Foster, J. A., Voynow, P. V., Franzblau, C. and Troxler, R. F. (1975). Biochemistry 14, 3581-3588. Brown, J. S . (1967). Biochim. Biophys. Acta 143, 391-398. Brown, J. S. (1977a). Photochem. Photobiol. 26, 319-326. Brown, J. S . (1977b). Photochem. Photobio[. 26, 519-525. Brown, J. S . (1980). Biochim. Biophys. Acta 591, 9-21. Brown, J. S. and French, C. S . (1961). Biophys. J . 1, 539-551.


193

Brown, J. S. and Schoch, I. (1981). Biochim. Biophys. Acta 636, 201-209. Brown, J. S., Alberte, R. S. and Thornber, J. P. (1974). Proc. Third Intern. Phofosynfhesis Cong. 3, 195 1 - 1 961. Elsevier, Amsterdam. Brown, S. B., Holroyd, A. J. and Troxler, R. F. (1980). Biochem. J . 190, 445449. Brown, S. B., Holroyd. J. A., Troxler, R. F. and Offner, G . D. (1981). Biochem. J . 194, 1 37- 147. Brown, T. E. and Richardson, F. L. (1968). J. Phycol. 4, 38-54. Bryant, D. A. (1981). Eur. J . Biochem. 119, 425-429. Bryant, D. A. and Cohen-Bazire, G . (1981). Eur. J. Biochem. 119, 41-424. Bryant, D. A., Glazer, A. N. and Eiserling, F. A. (1976). Arch. Microbiol. 110, 61-75. Bryant, D. A., Hixon, C . S. and Glazer, A. N. (1978). J. B i d . Chem. 253, 220-225. Bryant, D. A., Gugliemi, G., Tandeau De Marsac, N., Castets, A. M. and CohenBazire, G. (1974). Arch. Microbiol. 123, 113-127. Budzikiewicz, A. and Taraz, H. (1971). Tetrahedron 27, 1447-1460. Burrell, J. W. K., Jackman, L. M. and Weedon, B. C. L. (1959). Proc. Chem. SOC. (London) 263-264. Burris, J. (1977). Mar. B i d . 39, 371-379. Butler, W. L. (1977). Proc. Nail. Acad. Scr. U.S.A. 77, 4697-4701. Butler, W. L. (1978). Annu. Rev. Plant Physiol. 29, 345-378. Butler, W. L. and Hopkins, D. W. (1970). Photochem. Phofobiol. 12, 439450. Byfield, P. G. H. and Zuber, H. (1972). FEBS Lett. 28, 3 H 1 . Bykhovsky, V. Y. (1979). In “Vitamin B,,” (B. Zagalak and W. Friedrick, Eds), pp. 293-314. Walter de Gruyter, New York. Camm, E. L. and Green, B. R. (1980). Plant Physiol. 66, 428432. Cammack, R. and Evans, M . C. W. (1975). Biochem. Biophys. Res. Commun. 67, 544-549. Cammack, R., Ryan, M. D. and Stewart, A. C. (1979). FEBS Lett. 107, 422426. Cammack, R., Rao, K. K. and Hall, D. 0. (1981). Biosystems 14, 57-80. Canaani, 0. and Gantt, E. (1980). Biochemistry 19, 2950-2956. Canaani, D. D. and Sauer, K . (1978). Biochim. Biophys Acta 501, 545-551. Canaani, O., Lipschultz, C. A. and Gantt, E. (1980). FEBS Lett. 115, 225-229. Canuto, V. M., Levine, J. S., Augustsson, T. R. and Imhoff, C. L. (1982). Nature 296, 8 16820. Carver, J. H. (1981). Nature (London) 292, 136-138. Cavalier-Smith, T. ( 1 980). In “Endocytobiology; endosymbiosis and cell biology; a synthesis of recent research” (W. Schwemmler and H. E. A. Schenk, Eds), 893-916. Walter de Gruyter, Berlin. Chalker, B. (1980). J . Theor. Biol. 84, 205-215. Chapman, A. R. O., Markham, J. W. and Luning, K. (1978). J. Phycol. 14, 195-198. Chapman, D. J. (1966). Phytochernistry 5, 1331-1 333. Chapman, D. J. and Haxo, F. T. (1963). Plnnr Cell Physiol. 4, 57-63. Chapman, D. J. and Haxo, F. T. (1966). J. Phycol. 2, 89-91, Chapman, D. J. and Ragan, M. A. (1978). “A Biochemical Phylogeny of the Protists”. Academic Press, New York. Chapman, D. J. and Ragan, M. A. (1980). Annu. Rev. Plant Physiol. 31, 639-678. Chapman, D. J., Cole, W. J. and Siegelman, H. W. (1967). J . Am. Chem. Soc. 89, 59765977. Charles-Edwards, D. A., and Ludwig, L. J. (1974). Ann. Bot. 38, 921-930. Chen, G. C., Krieger, M., Kane, J. P., Wu, G . S. C., Brown, M. S. and Goldstein, J. L. (1980). Biochemistry 19, 4 7 0 U 7 1 2 . Cho, F. and Govindjee (1970a). Biochim. Biophys. Acta 205, 37-378.

194


Cho, F. and Govindjee (1970b). Biochim. Biophys. Acta 216, 151-161. Chow, H. C. (1981). Diss. Abstr. B 38, 47994800. 97, 7230-7237. Chow, H. C., Serlin, R. and Strouse, C. E. (1975). J. Am. Chem. SOC. Chow, W. S., Thorne, S. W., Duniec, J. T., Sculley, M. J. and Boardman, N. K. (1980). Arch. Biochem. Biophys. 201, 347-355. Chow, W. S., Thorne, S. W., Duniec, J. T., Sculley, M. J. and Boardman, N. K. (1982). Arch. Biochem. Biophys. 216, 242-256. Chua, N. H. and Bennoun, P. (1975). Proc. Natl. Acad. Sci. U.S.A. 72, 2175-2179. Chua, N. H., Matlin, K. and Bennoun, P. (1975). J . Cell Biol. 67, 361-377. Chunaev, A. S., Lipkind, B. I., Kvitko, K. V. and Giller, Y. E. (1980). Biol. Nuuki (MOSC)0, 45-51. Clement-Metral, J. D. and Lefort-Tran, M. (1971). FEBS Lett. 12, 225-228. Clezy, P. S. and Fookes, C. J. R. (1975). J. C. S. Chem. Comm. 707-708. Cloud, P. E. (1976). Paleobiol. 2, 351-387. Cloud, P. E. (1982). Nature 296, 198-199. Lond. 284, 569-579. Cogdell, R. J. (1978). Phil. Trans. R. SOC. Cogdell, R. J. (1979). Trans. Biochem. SOC. U.K. 7, 1228-1231. Cogdell, R. J., Hipkins, F. M., MacDonald, W. and Truscott, T. G. (1981). Biochim. Biophys. Acta 634, 191-202. Conjeoud, H. and Mathis, P. (1980). Biochim. Biophys. Acta 590, 353-359. Coombs, J. and Greenwood, A. D. (1976). In “The Intact Chloroplast” (J. Barber, Ed.), pp. 1-51. Elsevier, Amsterdam. Cox, C. S. (1974). In “Optical Aspects of Oceanography” (N. G. Jerlov and E. Steeman Nielsen, Eds), 51-75. Academic Press, London. Cox, G. C. and Dwarte, D. M. (1981). New Phytol. 88, 427438. Cox, G. C. and Marchant, H. J. (1977). In “Proc. 7th Int. Speleological Congr.” pp. 131-133. Sheffield, England. Cox, G. C., Benson, D. and Dwarte, D. M. (1982). Arch. Microbiol. 130, 165-174. Crabbe, P., Djerassi, C., Eisenbraun, E. J. and Lia, S. (1959). Proc. Chem. SOC. 264-265. Cramer, W. A. and Butler, W. L. (1969). Biochim. Biophys. Actu 172, 503-510. Crespi, H. L., Boucher, L. J., Norman, G. D., Katz, J. J. and Dougherty, R. C. (1967). J . Am. Chem. SOC.89, 3642-3643. Critchley, C. (1981). Plant Physiol. 67, 1161-1 165. Crofts, A. R. and Wood, P. M. (1978). In “Current Topics in Bioenergetics” (D. R. M. Sanadi and Leo P. Vernon, Eds), Vol. 7, pp. 175-244. Academic Press, New York. Crossett, R. N., Drew, E. A. and Larkum, A. W. D. (1965). N a m e 207, 547-548. Dale, R. E. and Teale, F. W. J. (1970). Photochern. Photobiol. 12, 99-1 17. Dallinger, R. F., Woodruff, W. H. and Rodgers, M. A. J. (1981). Photochern. Photobiol. 33, 275-277. Davidov, A. S . (1962). “Theory of Molecular Excitons”. McGraw Hill,New York. David, M. S., Forman, A. and Fajer, J. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 41 70-4 174. Dawes, C. J. and Barilotti, D. C. (1969). Am. J . Bot. 56, 8-15. Dayhoff, M. 0.and Schwartz, R. M. (1980). In “Endocytobiology; endosymbiosis and cell biology; a synthesis of recent research” (W. Schwemmler and H. E. A. Schenk, eds), 63-83. Walter de Gruyter, Berlin. Dayton, P. K. (1975). Ecol. Monogr. 45, 137-159. De Kok, J., Braslavsky, S. E. and Spruit, C. J. (1981). Photochem. Photobiol. 34, 705-710. Delepelaire, P. (1980). Photobiochem. Photobiophys. 1, 139-1 46.


195

Delepelaire, P. and Chua, N . H. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 1 1 1-1 15. Delieu, T. and Walker, N. A. (1972). New Phytol. 71, 201-225. Dellow, V. and Cassie, R. M. (1955). Trans. R. SOC.N . Z . 83, 321-331. Diakoff, S. and Scheibe, J. (1973). Plant Physiol. 51, 382-385. Dietrich, W. E. and Thornber, J. P. (1971). Biochim. Biophys. Acta 245, 482493. Dilworth, M. F. and Gantt, E. (1981). Plant Physiol. 67, 608-612. Diner, B. A. (1979). Plant Physiol. 63, 30-34. Diner, B. A. and Joliot, P. (1977). In“Ency1. Plant Physiol. Vol. 5 . Photosynthesis Part 1 ” (A. Trebst and M. Avron, eds), pp. 187-205. Springer, Berlin. Diner, B. A. and Mauzerall, D. (1973). Biochim. Biophys. Acta 305, 353-363. Diner, B. A. and Wollman, F.-A. (1979a). Plant Physiol. 63, 20-25. Diner, B. A. and Wollman, F.-A. (1979b). Plant Physiol. 63, 26-29. Diner, B. A. and Wollman, F.-A. (1980). Eur. J . Biochem. 110, 521-526. Dirks, G . A. L., Moore, T. A. and Gust, D. (1980). Photochem. Photobiol. 32,277-280. Dodge, J. D. (1968). J . Cell Sci. 3, 41-48. Dodge, J. D. (1979). Biochemistry and Physiology of Protozoa (M. Levandowsky and S. H. Hutner, Eds) 1, 7-57. Academic Press, New York. Dornemann, D. and Senger, H. (1981). FEBS Leu. 126, 323-327. Dornemann, D. and Senger, H. (1982). Photochem. Photohiol. 35, 821-826. Douce, R. and Joyard, J. (1979). Adv. Bot. Res. 7, 2-1 16. Dougherty, R. C., Strain, H. H., Svec, W. A., Uphaus, R. A. and Katz, J. J. (1966). J . Am. Chem. SOC.83, 5037-5038. Dougherty, R. C., Strain, H. H., Svec, W. A,, Uphaus, R. A. and Katz, J. J. (1970). J . Am. Chem. SOC.92, 2826-2833. Doukas, A. G., Stefanci, V., Buchert, J., Alfano, R. R. and Ziliskas, B. A. (1981). Photochem. Photobiol. 34, 505-5 10. Drew, E. A. (1969). Proc. Infer. Seaweed Symp. 6, 151-159. Drew, E. A. (1983). In “The Sublittoral Environment of the British Isles-In Perspective” (R. Earl1 and D. G. Erwin, Eds). Oxford University Press, Oxford. Drikas, G., Ruppel, G., Dietrich, H. and Sperling, W. (1981). FEBS Lett. 131,23-27. Dring, M. J. (1981). Limnol. Oceanogr. 262, 271-284. Droop, M. R. (1974). In “Algal Physiology and Biochemistry” (W. D. P. Stewart, Ed.), pp. 530-559. Blackwell Scientific, Oxford. Dubertret, G. and Joliot, P. (1974). Biochim. Biophys. Acta 357, 399-411. Dubertret, G. and Lefort-Tran, M. (1981). Biochim. Biophys. Acta 634, 52-69. Dubinsky, Z. and Berman, T. (1979). Limnol. Oceanogr. 24, 652-663. Duncan, M. J. and Foreman, R. E. (1980). J . Phycol. 16, 138-142. Duniec, J . T. and Thorne, S. W. (1981). Photobiochern. Photobiophysics 2, 85-91. Dunlop, J . S. R., Muir, M. D., Milne, V . A. and Groves, D. I. (1978). Nature 274, 676678. Dunstan, W. M. (1973). J . Exp. Mar. Bid. Ecol. 13, 181-187. Dutton, H. J., Manning, W. M. and Duggar, B. M. (1943). J . Phys. Chem. 47,308-313. Duysens, L. N. M. (1952). Ph.D. Thesis, Utrecht. Duysens, L. N. M. (1956). Biochim. Biophys. Acta 19, 1-12. Duysens, L. N. M. and Amesz, J . (1962). Biochim. Biophys. Acta 64, 243-260. Dwarte, D. M. and Vesk, M. (1982). Micron 13, 325-326. Eigenberg, K. E., Croasmun, W. R. and Chant, S. I . (1981). Biochim. Biophys. Acta 642, 438-442. El-Sayed, M. A., Karvaly, B. and Fukumoto, J. M. (1981). Proc. Natl. Acad. Sci. U S A . 78, 7512-7516. Emerson, R. and Arnold, W. (1932a). J . Gen. Physiol. 15, 191420.

196


Emerson, R. and Arnold, W. (1932b). J. Gen. Physiol. 16, 391-420. Emerson, R. and Lewis, C. M. (1942). J. Gen. Physiol. 25, 579-595. Emerson, R. and Lewis, C. M. (1943). Am. J. Bot. 30, 165-178. Engelmann, Th. W. (1883). Botan. Z. 41, 18. Engelmann, Th. W. (1884). Botan. Z. 41, 81-97. England, R. R. and Evans, E. H. (1981). FEBS Lett. 134, 175-177. Evans, M. C. W., Reeves, S. G. and Cammack, R. (1974). FEBS Lett. 49, 11 1-1 14. Evans, E. H., Cammack, R. and Evans, M. C. W. (1976). Biochem. Biophys. Res. Commun. 68, 1212-1218. Evans, E. H., Carr, N. G., Rush, J. D. and Johnson, C. E. (1977). Biochem. J . 166, 547-55 I , Evans, E. H., Rush, J. D., Johnson, C. E. and Evans, M. C. W. (1979). Biochem. J. 182, 861-865. Evans, E. H., Rush, J. D., Johnson, C. E., Rush, J. D. and Evans, M. C. W. (1981). Eur. J . Biochem. 118, 81-84. Evstigneev, V. B. and Gavrilova, V. A. (1979). Bioj?zika 24, 797-800. Falkowski, P. G. and Dubinsky, Z. (1981). Nature 289, 172-174. Falkowski, P. G. and Owens, T. G. (1978). Mar. Biol.45, 289-295. Falkowski, P. G. and Owens, T. G. (1980). Plant Physiol. 66, 592-595. Falkowski, P. G., Owens, T. G., Ley,A. C. and Mauzerall, D. C. (1981). Plant Physiol. 68, 969-973. Farquhar, G. D. and Von Caemmerer, S. (1981). In “Physiological Plant Ecology 11”. Encyclopedia of Plant Physiology New Series, Vol. 12B (0.L. Lange, P. S. Nobel, C. B. Osmond and H. Ziegler, Eds), pp. 549-587. Springer Verlag, Heidelberg. Faust, M. A. and Gantt, E. (1973). J . Phycol. 9, 489-495. Fee, E. J. (1969). Limnol. Oceanogr. 14, 906-91 1 . Felton, R. H. (1978). In “The Porphyrins” (D. Dolphin, Ed.) 5, 53-135. Academic Press, New York. Fenna, R. E. and Matthews, B. W. (1975). Nature 258, 573-577. Fenna, R. E. and Matthews, B. W. (1977). Brookhaven Symp. Biol. 28, 170-182. Fenna, R. E. and Matthews, B. W. (1979). In “The Porphyrins” (D. Dolphin, Ed.) 7 , 473-494. Academic Press, New York. Fischer, H. and Orth, H. (1937). “Die Chemie des Pyrrols” Vol. 1 Akad. Verlagsgesellschaft, Leipzig. Fischer, H. and Stern, A. (1940). “Die Chemie des Pyrrols” Vol. 2 (2). Akad. Verlagsgesellschaft, Leipzig. Fischer, M. S . , Templeton, D. H., Zalkin, A. and Calvin, M. (1972). J . Am. Chem. SOC. 94, 3613-3619. Fisher, R. G., Woods, N. E., Fuchs, H. E. and Sweet, R. M. (1980). J . Biol. Chem. 255, 5082-5089. Fleischer, W. E. (1935). J . Gen. Physiol. 18, 573-597. Fleischhacker, Ph. and Senger, H. (1978). Physiol. Plantar 43, 43-51. Fleming, I. (1968). J . Chem. SOC.C. 2765-2770. Floyd, G. L. and Salisbury, J. L. (1977a). Am. J . Bot. 64, 12941296. Floyd, G. L. and Salisbury, J. L. (1977b). J. Phycol. 13, 21a. Fork, D. C. (1963). In “Photosynthetic Mechanisms of Green Plants”, pp. 352-361. Pub. 1145, Nat. Acad. Sci., Washington, D.C. Fork, D. C., Oquist, G . and Hock, G. E. (1982). Plant Sci. Lett. 24, 249-254. Forster, Th. W. (1946). Naturwiss. 33, 166175. Forster, Th. W. (1965). In “Modern Quantum Chemistry, Part 111: Action of Light and Sinanoglu, Ed.), pp. 93-151. Academic Press, New York. Organic Molecules” (0.


197

Forward, R. B. Jr. (1976). In “Photochemical and Photobiological Review” (K. C. Smith, Ed.) 1, 157-209. Plenum Press, New York. Fox, G. E., Stackbrandt, E., Hespell, R. B., Gibson, J., Maniloff, J., Dyer, T. A., Wolfe, R. S., Balch, W. E., Tanner, R. S., Magrum, L. J., Zablen, L. B., Blakemore, R., Gupta, R., Bonen, L., Lewis, B. J., Stahl, D. A., Luehrsen, K . R., Chen, K. N. and Woese, C. R. (1980). Science 209,457463. Frank, F., Sidler, W., Widmer, H. and Zuber, H. (1978). Hoppe-Seylers Z. Physiol. Chem. 359, 1491-1499. Freidenreich, P., Apell, G. S. and Glazer, A. N. (1978). J. Biol. Chem. 253, 212-219. French, C. S. (1977). Photochem. Photobiol. 25, 159-160. French, C. S., Michel-Wolwertz, M. R., Michel, J. M., Brown, J. S. and Prager, L. K. (1968). In ”Porphyrins and Related Compounds” (T. W. Goodwin, Ed.), 147-1 62. Academic Press, London. Fuad, N., Day, D., Ryrie, I. and Thorne, S. W. (1983). Phorohiochem. Photobiophys. (in press). Fujita, I., Davis, M. S. and Fajer, .I.(1978). J. Am. Chem. SOC.100, 6280-6281. Fujita, Y. (1976). Plant Cell. Physiol. 17, 187-191. Fujita, Y.and Hattori, A. (1960). Plant Cell. Physiol. 1, 293-303. Fujita, Y. and Hattori, A. (1962). Plant Cell. Physiol. 3, 209-220. Fuller, R. C. (1978). In “The Photosynthetic Bacteria” (R. K. Clayton and W. R. Sistrom, Eds), pp. 691-705. Plenum Press, New York. Gaffron, H. (1960). In “Plant Physiology” (F. C. Stewart, Ed.), Vol. IB, 3-277. Academic Press, New York. Gaidukov, N. I. (1903). Ber. Deuf. Botan. Ges. 21, 484. Gaidukov, N. I. (1904). Ber. Reul. Botan. Ges. 22, 23. Gaidukov, N. 1. (1906). Ber. Deur. Botan. Ges. 24, I . Gantt, E. (1975). Bioscience 25, 781-787. Gantt, E. (1979). In “Biochemistry and physiology of protozoa” (M. Levandowsky and S. H. Hutner, Eds) 1, 121-137. Academic Press, New York. Gantt, E. (1980). Intern. Rev. C y f . 66, 45-80. Gantt, E. (1981). Annu. Rev. Plant Physiol. 32, 327-347. Gantt, E. and Conti, S. F. (1965). J . Cell. Biol. 26. 365-381. Gantt, E. and Conti, S. F. (1966a). J. Cell Biol. 29, 423434. Gantt, E. and Conti, S. F. (1966b). Brookhaven Symp. Biol. 19, 393405. Gantt, E. and Lipschultz, C. A. (1972). J. Cell. Biol. 54, 313-324. Gantt, E. and Lipschultz, C. A. (1973). Biochim. Biophys. Acta 292, 858-861. Gantt, E. and Lipschultz, C. A. (1974). Biochemistry 13, 2960-2966. Gantt, E. and Lipschultz, C. A. (1977). J . Phycol. 13, 185-192. Gantt, E. and Lipschultz, C. A. (1980). J . Phycol. 16, 304398. Gantt, E., Edwards, M. R. and Provasoli, L. (1971). J. Cell. Biol. 48, 280-290. Gantt, E., Lipschultz, C. A. and Zilinskas, B. (l976a). Biochim. Biophys. Acta 430, 375-388. Gantt, E., Lipschultz, C. A. and Zilinskas, B. (1976b). Brookhaven S p p . B i d . 28, 347-357. Gantt, E., Lipschultz, C. A., Grabowski, J. and Zimmerman, B. K. (1979). Plant Physiol. 63, 615-620. Gantt, E., Canaani, O., Lipschultz, C. A. and Redlinger, T. (1981). In “Photosynthesis Ill” ( G . Akoyunoglou, Ed.), pp. 143-153. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Gardner, E. E., Stevens, S. E. and Fox, J. L. (1980). Biochim. Biophys. Acta 624, 187- 195.

198


Garrels, R. M. and Lerman, A. (1981). Proc. Natl. Acad. Sci. U.S.A. 78,46524656. Gates, D. M. (1980). “Biophysical Ecology”. Springer Verlag, New York. Gerola, P. D., Jennings, R. C., Forti, G. and Garlaschi, F. M. (1979). Plant Sci. Lett. 16, 249-254. Gerola, P. D., Garlaschi, F. M., Forti, G. and Jennings, R. C. (1981). Biochim. Biophys. Acta 679, 10 1-1 09. Gerber, D. W. and Burns, J. E. (1981). Plant Physiol. 68, 699-702. Ghosh, A. K. and Govindjee. (1966). Biophys. J . 6, 61 1-619. Ghosh, A. K., Govindjee, Crespi, H. L. and Katz, J. J. (1966). Biochim. Biophys. Acta 120, 19-22. Gibbs, S. P. (1978). Can. J . Bot. 56, 2883-2889. Gibbs, S . P. (1981a). Ann. N . Y . Acad. Sci. 361, 193-208. Gibbs, S. P. (1981b). Inter. Rev. Cytol. 72, 49-99. Gibbs, S. P., Sistrom, W. R. and Worden, P. B. (1975). J . Cell. Biol. 26, 395412. Giddings, T. H., Withers, N. W. and Staehelin, L. A. (1980). Proc. Natl. Acad. Sci. U.S.A. 77, 352-356. Gillott, M. A. and Gibbs, S. P. (1980). J . Phycol. 16, 558-568. Gilmartin, M. (1960). Ecology 41, 21C221. Gingras, G. (1978). In “The Photosynthetic Bacteria” (R. K. Clayton and W. R. Sistrom, Eds), 133-160. Plenum Press, New York. Gingrich, J. C., Blaha, L. K. and Glazer, A. N. (1982). J . Cell Biol. 92, 261-268. Glazer, A. N. (1977). Mol. Cell. Biochem. 18, 125-140. Glazer, A. N. (1980). In “Evolution of protein structure and function” (D. Sigman and M. A. B. Brazier, Eds), pp. 221-244. Academic Press, New York. Glazer, A. N. (1981). In “The Biochemistry of Plants” (M. D. Hatch and N. K. Boardman, eds) 8, 51-96. Academic Press, New York. Glazer, A. N. and Apell, G. S. (1977). FEMS Microbiol. Lett. 1, 113-116. Glazer, A. N. and Bryant, D. A. (1975). Arch. Microbiol. 104, 15-22. Glazer, A. N. and Cohen-Bazire, G. (1975). Arch. Microbiol. 104, 29-32. Glazer, A. N., Fang, S. and Brown, D. M. (1973). J . Biol. Chem. 248, 5679-5685. Goedheer, J. C. (1966). In “The Chlorophylls” (L. P. Vernon and G. R. Seely, Eds), 147-185. Academic Press, New York. Goedheer, J. C. (1968). Biochim. Biophys. Acta 153, 903-906. Goedheer, J. C. (1969). Biochim. Biophys. Acta 172, 252-265. Goedheer, J. C. (1 970). Photosynthetica 4, 9 6 1 06. Goedheer, J. C. (1972). Annu. Rev. Plant Physiol. 23, 87-112. Goedheer, J. C. (1973). Biochim. Biophys. Acta 314, 191-201. Goedheer, J. C. (1979). Ber. Deutsch. Bot. Ges. 92, 427436. Goedheer, J. C. (1981). Photosynthesis Res. 2, 49-60. Golbeck, J. H., Lien, S. and Pietro, A. S. (1977). In “Encyclopedia of Plant Physiology N.S.” (A. Trebst and M. Avron, Eds) 5, 94-124. Springer-Verlag, Berlin. Goodwin, T. W. (1971). In “Aspects of Terpenoid Chemistry and Biochemistry” (T. W. Goodwin, Ed.), pp. 315-356. Academic Press, London. Goodwin, T. W. (1974). “Comparative Biochemistry of Carotenoids” 2nd ed. Chapman and Hall, London. Goodwin, T. W. (1 976). In “Chemistry and Biochemistry of Plant Pigments” (T. W. Goodwin, Ed.) 1, 225-261. Academic Press, New York. Gossauer, A. and Weller, J.-P. (1978). J. Am. Chem. SOC.100, 5928-5933. Gossauer, A., Heinz, R. P. and Katschan, R. (1981). Chem. Ber. 114, 132-146. Gouterman, M., Wagniere, and Snyder, L. C. (1963). J . Mol. Spectrosc. 11, 108-127.


199

Govindjee and Govindjee, R. (1975). In “Bioenergetics of Photosynthesis” (Govindjee, Ed.), pp. 1-50, Academic Press, New York. Govindjee, and Zilinskas, B. (1974). In “Physiology and Biochemistry of the Algae” (W. D. P. Stewart, Ed.), pp. 346-390. Blackwell Scientific, Oxford. Govindjee, R., Rabinowitch, E. and Govindjee, (1968). Biochim. Bioph.hjs. Aria 162, 539-544. Grabowski, J. and Gantt, E. (1978a). Photochem. Photobinl. 28, 39-45. Grabowski, J. and Gantt, E. (1978b). Phorochern. Photobiol. 28, 47-54. Granick, S. (1949). In “The Harvey Lectures” pp. 220-245. C. C. Thomas, Springfield, Illinois. Granick, S. (1957). Ann. N . Y. Acad. Sci. 69, 292-308. Greef, J. A. and Couberg, S. R. (1970). Nuturwiss. 57, 673-674. Green, B. R. (1980). Biochim. Biophys. Acta 609, 107-120. Green, E. L. and Camm, B. R. (1981). In “Photosynthesis Ill” (G. Akoyunoglou, Ed.), pp. 675-681. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Green, E. L., Camm, B. R . and Van Houten, A. (1982). Biochim. Biophys. Arm 681, 248-255. Greenwood, A. D., Griffiths, H. B. and Santore, V. J. (1977). Br. Phycol. J . 12, 119. Grefarth, S. P. and Reynolds, J . A. (1 974). Proc. Nail. Acad. Sci. U.S.A.71,39 13-39 16. Gregory, R. P. F. (1975). Biochem. J . 148, 487497. Gregory, R. P. F., Demeter, S. and Faludi-Daniel, A. (1980). Biochim. Biophys. Acta 591, 356-360. Gregory, R. P. F., Borbely, Demeter, S. and Faludi-Daniel, A. (1981). In “Photosynthesis 111” (G. Akoyunoglou, Ed.), pp. 533-538. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Griffin, D . C. and Landon, M. (1981). Biochem. J . 197, 333-344. Grime, J. P. (1974). Nature 250, 26-31. Grime, J. P. (1977). Am. Nut. 111, 1169-1194. Grimme, L. H. (1974). Ber. Dtsch. Bot. Ges. 87, 509-514. Grimme, L. H. and Porra, R. J. (1974). Arch. Mikrobiol. 99, 173-179. Guerin-Dumartrait, E., Sarda, C. and Lacourly, A. (1970). C. R. Acad. Sci. (Paris) Ser. D 270, 1977-1980. Guerin-Dumartrait, E., Moaru, J., Leclerc, J.-C. and Sarda, A. (1973). Phycologirr 12, 119-130. Gulyaev, B. A. and Teten’kin, V. L. (1981). Biophysics 26, 288-294. Gurinovich, G. P., Sevchenko, A. N. and Soloviev, K. N. (1968). In “Spectroscopy of Chlorophyll and related compounds”. U.S. Atomic Energy Commission Translation series AEC-tr-7199. Gysi, J . and Zuber, H. (1976). FEBS Lett. 68, 49-54. Hackert, M. L., Abad-Zaptiero, C., Stevens, S. E. Jr. and Fox, J. L. (1977). J . Mol. Biol. 111, 365-369. Hager, A. (1975). Ber. Deutsch. Bot. Ges. 88, 45-60. Hager, A. (1980). In “Pigments in plants” (Franz-Christian Czygan, Ed.), 2nd ed. G . Fistor, Stuttgart. Hager, A. and Stransky, H. (1970). Arch. Mikrobiol. 71, 132-163. Haidak, D. J. C., Mathews, C. K. and Sweeney, B. M. (1966). Science 152, 212-213. Hall, J. D., Barr, R., Al-Abbas, H. and Crane, F. L. ( 1 972). Plant Physiol. 50,404409. Halldal, P. (1958). Physiol. Plantar. 11, 401420. Halldal, P. (1964). Physiol. Plantar. 17, 414421. Halldal, P. (1968). Biol. Bull. 134, 41 1 4 2 4 .

200


Halldal, P. and French, C. S. (1958). Plant Physiol. 33, 249-252. Hardt, H. (1981). Biochim. Biophys. Acta 635, 631-644. Harnischfeger, G. (1977). Adv. Bot. Res. 5, 2-52. Harnischfeger, G. and Codd, G. A. (1978). Biochim. Biophys. Acta 502, 507-513. Harper, J. L. (1977). “Population Biology of Plants”. Academic Press, London. Harris, G. P. (1978). Arch. Hydrobiol. Beih. Ergebn. Limnol. 10, 1-171. Harris, G. P. (1980). In “Physiological Ecology of Phytoplankton” (I. Morris, Ed.),pp. 129-1 87. Blackwell Scientific, Oxford. Harris, G. P. and Piccinin, B. B. (1977). Arch. fur Hydrogiologie 80, 405457. Harvey, G. W. and Bishop, N. I. (1978). Plant Physiol. 62, 330-336. Hase, E. (1980). In “The Blue Light Syndrome” (H. Senger, Ed.), pp. 512-525. Springer-Verlag, Berlin. Hatch, M. D. (1976). In “Plant Biochemistry” (J. Bonner and J. Varner, Eds), 3rd ed., pp. 797-844. Academic Press, New York. Haupt, W. and Bock, G. (1962). Planta 59, 38-48. Haupt, W. and Thiele, R. (1961). Planta 56, 388401. Haury, J. F. and Bogorad, L. (1977). Plant Physiol. 60,835-839. Haworth, P., Breton, J. and Arntzen, C. J. (1981). Plant Physiol. 67, Suppl. 164. Haxo, F. T. (1960). In “Comparative Biochemistry of Photoreactive Systems” (M. B. Allen, Ed.), pp. 339-360. Academic Press, New York. Haxo, F. T. and Blinks, L. R. (1950). J. Gen. Physiol. 33, 389422. Haxo, F. T. and Fork, D. C. (1959). Nature 184, 1051-1052. Haxo, F. T., O’Heocha, C. and Norris, A. (1955). Arch. Biochem. 54, 162-173. Hay, M. E. (1981). Am. Nut. 118, 52G540. Hayden, D. B. and Hopkins, W. G. (1977). Can. J. Bot. 55, 2525-2529. Healy, F. P. and Myers, J. (1971). Plant Physiol. 47, 373-379. Heinz, E. and Siefermann-Harms, D. (1981). FEBS Lett. 124, 105-1 11. Henderson, R. and Unwin, P. N. T. (1975). Nature 257, 28-32. Henderson-Sellers, A. and Cogley, J. G. (1982). Nature 298, 832-835. Helenius, A. and Simons, K. (1975). Biochim. Biophys. Acta 415, 29-79. Helenius, A., McCaslin, D., Fries, E. and Tanford, C. (1980). “Methods in Enzymology” 63, 734. Academic Press, New York. Henriques, F. and Park, R. B. (1978). Arch. Biochem. Biophys. 189, 44-50, Herman, E. M. and Sweeney, B. M. (1975). J. Ultrastruct. Res. 50, 347-354. Herron, H. A. and Mauzerall, D. (1972). Plant Physiol. 50, 141-148. Hertzberg, S., Mortensen, T., Borch, G., Siegelman, H. W. and Liaaen-Jensen, S. (1977). Phytochemistry 16, 587-590. Hickman, D. D. and Frenkel, A. W. (1965). J. Cell. Biol. 25, 261-278. Hill, R. and Bendall, F. (1960). Nature 186, 136-137. Hiller, R. G. and Goodchild, D. (1982). In “The Biochemistry of Plants” (M. D. Hatch and N. K. Boardman, Eds) 8, 2-50. Academic Press, Sydney. Hiller, R. G. and Larkum, A. W. D. (1981). In “Photosynthesis 111” (G. Akoyonuglou, Ed.), pp. 387-396. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Hiller, R. G., Pilger, T. B. G. and Genge, S. (1977). Biochim. Biophys. Acta 460, 431444. Hinkle, P. C. and McCarty, R. E. (1978). Sci. Am. 238, 104-123. Hjortas, J. (1972). Acta Cryst. B. 28, 2252-2259. Hoarau, J. and Remy, R. (1978). In “Chloroplast Development” (G. Akoyunoglou and J. H. Argyroudi-Akoyunoglou, Eds), 455459. Elsevier, Amsterdam. Hoarau, R. and Remy, J. (1978). In “Chloroplast Development” (G. Akoyunoglou and J. H. Argyroudi-Akoyunoglou, Eds), pp. 235-240. Elsevier, Amsterdam.


20 1

Hoarau, J., Remy, R. and Leclerc, J. C. (1977). Biochim. Biophys. Actu 462, 659-670. Hoarau, J., Phung Nhu Hung, S. and Houlier, B. (1981). It? “Photosynthesis 111” (G. Akoyunoglou, Ed.), pp. 785-794. Balaban Intern. Sci. Serv. Philadelphia, Pennsylvania, USA. Hoard, J. L. (1979). In “Porphyrins and Metalloporphyrins” (Kevin M. Smith, Ed.), pp. 3 17-380. Elsevier, Amsterdam. Hoch, G., Owens, 0. H. and Kok, B. (1963). Arch. Biochem. Biophys. 101, 171-180. Hoff, A. J. (1979). Phys. Rep. 54, 76200. Holden, M. (1976). In “Chemistry and Biochemistry of Plant pigments” (T. W. Goodwin, Ed.) 2, 2-37. Academic Press, New York. Holdsworth, E. S. and Arshad, J. H. (1977). Arch. Biochem. Biophys. 183, 361-373. Holt, A. S. (1961). Can. J . Botutiy 39, 327-336. Holt, A. S. (1966). In “The Chlorophylls” (L. 0. Vernon and G. R. Seely, Eds), pp. 11 1-1 18. Academic Press, New York. Holt, A. S. and Morley, H. V. (1959). Can. J . Chem. 37, 507-514. Holt, S. C., Conti, S. F. and Fuller, R. C. (1966). J . Bucteriol. 91, 31 1-323. Holt, T. K. and Krogmann, D. W . (1981). Biochim. Biophys. Actu 637,408-414. Hootkins, R., Malkin, R. and Bearden, A. (1981). FEBS Lett. 123, 229-234. Horton, C., Allen, J. F., Black, M. T. and Bennett, J. (1981). FEBSLett. 125, 193-196. Horton, P. (1981). Biochim. Biophys. Actu 635, 105-1 10. Horton, P. and Black, M. T. (1981). Biochim. Biophys. Actu 635, 53-62. Horton, P. and Croze, E. (1979). Biochim. Biophys. Actu 545, 188-201. Hudson, M. F. and Smith, K. M, (1975). Chem. Soc. Rev. 4, 363-399. Humphrey, G. F. (1983). J . Exp. Mar. B i d . E d . 66. 49-67. Hurt, E. and Hauska, G. (1981). Eur. J . Biochem. 117, 591-599. Hurt, E., Hauska, G. and Malkin, R. (1981). FEBS Lett. ISM, 1-5. Ikegami, I . and Katoh, S . (1975). Biochim. Biophys. Acru 376, 588-592. Ilani, A. and Mauzerall, D. (198 1 ). Biophys. J . 35, 79-92. Il’ina, M. D. and Borisov, A. Y. (1980). Biochim. Biophys. Actu 590, 345-352. Ingram, K . and Hiller, K. G. (1983). Biochim. Biophys. Actu 722, 310-319. Interschick-Niebler, E. and Lichtenthaler, H. K. (1981).Z . Nuturfbrsch C36.276 283. Isler, 0.(ed.) (1971). “Carotenoids” Birkhauser Verlag, Basel. Iverson, R. L. and Curl, H. (1973). Phyxiol. Pluntor 28, 498 502. Jamieson, G. R. and Reid, E. H. (1972). Phytochrmistry 11, 1423-1432. Janzen, A. F., Bolton, R. J. and Connolly, J. S. (1980). Abstracts of 3rd International Conference on Photochemical Conversion and Storage of Solar Energy, pp. 103-105. Jassby, A. D. (1978). In “Handbook of Phycological Methods, Physiological and Biochemical Methods” (J. A. Hellebust and J. S. Craigie, Eds), pp. 297-303. Cambridge University Press, Cambridge. Jassby, A. D. and Platt, T. (1976). Limnol. Oceunogr. 21, 540-547. Jeffrey, S. W. (1961). Biochem. J . 80, 336342. Jeffrey, S . W. (1968). Biochim. Biophys. Actu 162, 271-285. Jeffrey, S. W. (1969). Biochim. Biophys. Actu 177, 456467. Jeffrey, S. W. (1972). Biochim. Biophys. Actu 279, 15-33. Jeffrey, S. W. (1976). J . Phycol. 12, 349-354. Jeffrey, S. W. (1980). In “Primary Productivity in the Sea” (P. G. Falkowski, Ed.), pp. 33-58. Plenum Press, New York. Jeffrey, S. W. and Humphrey, G. F. (1975). Biochrm. Physiol. fflanzen 167, 191 -194. Jeffrey, S. W. and Vesk, M. (1977). J . Phycol. 13, 271-279. Jeffrey, S. W. and Vesk, M. (I98 I ) . In “Photosynthesis VI” (G. Akoyunoglou, Ed.), pp. 435442. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA.

202


Jeffrey, S. W., Sielicki, M. and Haxo, F. T. (1975). J. Phycol. n, 374384. Jennings, R. C., Garlaschi, F. M., Forti, G. and Gerola, P. D. (1979a). Biochim. Biophys. Acta 581, 87-95. Jennings, R. C., Garlaschi, F. M., Gerola, P. D. and Forti, G. (1979b). Biochim. Biophys. Acta 546, 207-219. Jennings, R. C . ,Garlaschi, F. M., Gerola, P. D. and Forti, G. (1980). FEBSLett. 117, 3 32-3 34. Jennings, R. C., Garlaschi, F. M. Gerola, P. D., Etzion-Katz and Forti, G. (1981). Biochim. Biophys. Acta 638, 100-107. Jensen, A. (1966). Rep. Norw. Inst. Seaweed Res. No. 31. Jensen, N.-H., Wilbrandt, R. and Pagsberg, P. B. (1980). Photochem. Photobiol. 32, 719-725. Jensen, R. G. and Bahr, J. T. (1977). Annu. Rev. Plant Physiol. 28, 379400. Jerlov, N. G. (1951). Rep. Swedish Deep-sea Expedition 3, 1-59. Jerlov, N. G. (1974). In “Optical Aspects of Oceanography” (N. G. Jerlov and E. Steeman Nielsen, Eds), pp. 77-94. Academic Press, London. Jerlov, N. G. (1976). “Marine Optics”, Elsevier, Amsterdam. Jerlov, N. G. and Nielsen, E. S. (1974). “Optical Aspects of Oceanography”. Academic Press, London. Johansen, J. E., Svec,W. A., Liaaen-Jensen, S. and Haxo, F. T. (1974). Phytochemisfry 13, 2261-2271. Johansson, L. B.-A., Lindblom, G., Wieslander, A. and Arvidson, G. (1981). FEBS Lett. 128, 97-99. Johns, R. B., Nichols, P. E., Gillan, F. T., Perry, G. J. and Volkman, J. K. (1981). Comp. Biochem. Physiol. 69B, 843-849. Joliot, A. and Joliot, P. (1964). C.R. Acad. Sci. (Paris) 258 D, 46224625. Joliot, P. and Joliot, A. (1968). Biochim. Biophys. Acta 153, 625-634. Joliot, P. and Joliot, A. (1979). Biochim. Biophys. Acta 546, 93-105. Joliot, P. and Joliot, A. (1981). FEBS Lett. 134, 155-158. Joliot, P., Bennoun, P. and Joliot, A. (1973). Biochim. Biophys. Acta 305, 317-328. Jones, L. W. and Kok, B. (1966). Plant Physiol. 41, 1037-1043. Jones, L. W. and Myers, J. (1964). Plant Physiol. 39, 938-946. Jones, L. W. and Myers, J. (1965). J. Phycol. 1, 7-14. Jorgensen, E. G. (1 969). Physiol. Plantar. 22, 1307-1 3 15. Jung, J., Seng, P.-S., Paxton, R. J., Edelstein, M. S., Swanson, R. and Hazen, E. E., Jr (1980). Biochemistry 19, 24-32. Junge, W. (1977). In “Encyclopedia of Plant Physiology” Vol. 5 (A. Trebst and M. Avron, Eds), pp. 59-93. Springer Verlag, Berlin. Junge, W. and Schaffernicht, H. (1979). In “Chlorophyll Organisation and Energy Transfer in Photosynthesis”. Ciba Foundation Symposium 61, 127-146. Excerpta Medica, Amsterdam. Junge, W., Schaffernicht, H. and Nelson, N. (1977). Biochim. Biophys. Acta462,73-85. Kageyama, A., Yokohama, Y., Samura, S . and Ikawa, T. (1977). Plant Cell Physiol. 18, 477-480. Kain, J. A. (1962). J. Mar. Biol. Ass. U.K. 42, 377-385. Kain, J. A., Drew, E. A. and Jupp, B. P. (1975). In “Light as an Ecological Factor; 11” (G. C. Evans, R. Bainbridge and 0. Rackham, Eds), pp. 63-93. Blackwell, Oxford. Kalle, K. (1966). Oceanogr. Mar. Biol. Annu. Rev. 4, 91-104. Kalyanasundaram, K. and Porter, G. (1978). Proc. R . SOC.London, Ser. A . 36,2944. Kan, K.-S. and Thornber, J. P. (1976). Plant Physiol. 57, 47-52.


203

Kanwisher, J. W. (1966). In “Some Contemporary Studies in Marine Science” (H. Barnes, Ed.), pp. 407420. George Allen and Unwin, London. Kao, H., Berns, D. S., Town, W. R. (1973). Biochem. J. 131, 39-50. Kaplan, A., Badger, M. R. and Berry, J. A. (1980). Planta 149, 219-226. Kassner, R. J. (1973). J. Am. Chem. SOC.95, 2674-2677. Katoh, T. and Gantt, E. (1979). Biochim. Biophys. Acta 546, 383-393. Katz, J. J. (1979). In “Chlorophyll organization and energy transfer in Photosynthesis”, Ciba Foundafion Symposium 61 (new series). Excerpta Medical Amsterdam, 334-348. Katz, J. J., Dougherty, R. C. and Boucher, L. J. (1966). In “The Chlorophylls” (L. P. Vernon and G. R. Seely, Eds), pp. 195-251. Academic Press, New York. Katz, J. J., Strain, H. H., Leussing, D. L. and Dougherty, R. C. (1968). J. Am. Chem. SOC.90,784-791. Katz, J. J., Norris, J. R. and Shipman, L. L. (1977). Brookhaven Symp. Biol. 28,1655. Katz, J. J., Shipman, L. L. and Norris, J. R. (1979). In “Chlorophyll organization and energy transfer in Photosynthesis”, Ciba Foundation Symposium 61, N.S., pp. 1 4 0 . Excerpta Medical Amsterdam. Kawamura, M., Mimuro, M. and Fujita, Y. (1979). Plant Cell. Physiol. 20, 697-705. Ke, B. (1966). In “The Chlorophylls” (L. P. Vernon and G. R. Seely, Eds), pp. 253-282. Academic Press, New York. Ke, B. and Dolan, E. (1980). Biochim. Biophys. Acta 590, 401406. Keast, J. F. and Grant, B. R. (1976). J . Phycology 12, 328-331. Kerfin, W. and Boger, P. (1982). Physiol. Plantar. 54, 93-98. Kessler, E. (1974). In “Algal Physiology and Biochemistry” (W. D. P. Stewart, Ed.), pp. 45W73. Blackwell Scientific, Oxford. Kiefer, D. A., Olson, R. J. and Wilson, W. H. (1 970). Limnol. Oceanogr. 24,664-672. King, R. F. G. and Brown, S. B. (1978). Biochem. J. 174, 103-109. King, R. J. and Schramm, W. (1976). Mar. Biol. 37, 215-222. Kirk, J. T. 0. (1975a). New Phytol. 75, 11-20. Kirk, J. T. 0. (1975b). New Phytol. 75, 21-36. Kirk, J. T. 0. (1976a). New Phytol. 77, 341-358. Kirk, J. T. 0. (1976b). Aust. J . Mar. Freshwater Res. 27, 61-71. Kirk, J. T. 0. (1977a), Aust. J. Mar. Freshw. Res. 28, 497-508. Kirk, J. T. 0. (1977b). Plant Sci. Letts. 9, 373-380. Kirk, J. T. 0. (1979). Ausf. J. Mar. Freshwafer Res. 30, 1-11. Kirk, J. T. 0. (1981). Aust. J . Mar. Freshwater Res. 32, 517-532. Kirk, J. T. 0. and Goodchild, D. J. (1972). Aust. J . Biol. Sci. 25, 215-241. Kirk, J. T. 0. and Reade, J. A. (1970). Aust. J. Biol. Sci. 23, 33-41. Kirk, J. T. 0. and Tilney-Bassett, R. A. E. (1978). “The Plastids”. Elsevier, Amsterdam. Kirst, G. 0. (1981). Planta 151, 281-288. Kitching, J. (1941). Biol. Bull. 80, 324-337. Klein, S. M. and Vernon, L. P. (1977). Biochim. Biophys. Acta 459, 364-375. Klein, S., Vernon, L. and Kent, S. S. (1981). In “Photosynthesis” 111 (G. Akonoglou, Ed.), pp. 175-184. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania USA. Klevanik, A. V., Klimov, V. V., Shuvalov, V. A. and Krasnovskii, A. A. (1977). Dokl. Akad. Nank. SSSR 236, 241-244. Klimov, V. V., Klevanik, A. V., Shuvalov, V. A. and Krasnovsky (1977). FEBS Lett. 82, 183-186. Klimov, V. V., Allakhverdiev, S. I. and Pashchenko, V. Z . (1978). Dokl. Akad. Nank. SSSR 242, 1204-1201.

204


Klimov, V. V., Dolan, E., Shaw, E. R. and Ke, B. ( 1980). Proc. Natl Acad. Sci. U.S.A. 11,7227-7231. Knaff, D. B. and Arnon, D. I. (1969). Proc. Natl. Acad. Sci. U.S.A. 63, 963-969. Knaff, D. B., Malkin, R., Clark-Myron, J. and Stoller, M. (1977). Biochim. Biophys. Acta 459, 4 0 2 4 1 1. Knox, R. S. (1975). In “Bioenergetics of Photosynthesis” (Govindjee, Ed.), pp. 183-221. Academic Press, New York. Knox, R. S. (1977). In “Primary Processes of Photosynthesis” (J. Barber, Ed.), pp. 55-97. Elsevier, Amsterdam. Knox, R. S. and Van Metter, R. L. (1979). ClBA Foundation Symposium 91 (N.S.). “Chlorophyll organization and energy transfer in Photosynthesis”, pp. 177-190. Excerpta Medica, Amsterdam. Koenig. F. and Vernon, L. P. (1981). Z. Naturforsch C. 36, 295-304. Koenig, F., Menke, W., Radunz, A. and Schmid, G. H. (1977). Z. Naturforsch. 32, 8 17-827. Kok, B. (1 949). Biochim. Biophys. Acta 3, 625-63 1. Kok, B. (1960). In “Encyclopedia of Plant Physiology” (W. Ruhland, Ed.) SpringerVerlag, Berlin. Kok, B. (1961). Biochim. Biophys. Acta 48, 527-533. Kok, B. and Gott, W. (1960). Plant. Physiol. 35, 802-808. Kok, B., Gamer, E. and Rurainski, H. J. (1965). Photochem. Photobiol. 4, 215-227. Kok, B., Forbush, B. and McGloin, M. (1970). Photochem. Photobiol. 11, 457475. Koka, P. and Song, P . 4 . (1977). Biochim. Biophys. Acta 495, 22&231. Koka, P. and Song, P.-S. (1978). Photochem. Photobiol. 28, 509-515. Koller. K.-P., Wehrmeyer, W. and Morschel, E. (1978). Eur. J. Biochem. 91, 57-63. K a t , H.-P., Rath, K., Wanner, G. and Scheer, H. (1981). Photochem. Photobiol. 34, 139-143. Kott, P. (1980). Menz. Queenslund Mus. 20, 1-38. Krakover, T., Ilani, A. and Mauzerall, D. (1981). Biophys. J . 35, 93-98. Krause, G. H. and Heber, U. W. (1976). In “The Intact Chloroplast” (J. Barber, Ed.), pp. 171-214. Elsevier, Amsterdam. Krawczyk, S. (1981). Biochim. Biophys. Acta 640, 628-639. Kretzer, F., Ohad, I. and Bennoun, P. (1976). In “Genetics and Biogenesis of Chloroplasts and Mitochondria” Th. Bucher, W. Neupert, W. Sebald and S. Werner, Eds), pp. 25-32. Elsevier/North Holland, Amsterdam. Krogmann. D. W. (1981). Bioscience 31, 121-124. Kulandaivelu, G . and Senger, H. (1976). Biochim. Biophys. Acta 430, 94-101. Kullenberg, G. (1974). In “Optical Aspects of Oceanography” (N. G. Jerlov and E. Steeman Nielsen, Eds), pp. 25-49. Academic Press, London. Kunert, K.-J. and Boger, P. (1975). Z. Naturforsch, C 30, 19C200. Kunert, K.-J., Bohme, H. and Boger, P. (1976). Biochim. Eiophys. Acta 449, 541-553. Kursar, T. A., Swift, H. and Alberte, R. S. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 6888-6892. Laetsch, W. M. (1974). Annu. Rev. Plant Phvsiol. 25. 27-52. Lagarias, J. C.. Glazer, A. N. and Rapoport, H. (1979). J . Am. Chem. SOC.101, 5030-5037. Lagarias, J. C. and Rapoport, H. (1980). J . Am. Chem. SOC.102, 4821-4828. Largeau, C., Casadevall, E. and Berkaloff, C. (1980). Photochemistry 19, 1081-1086. Lagoutte, B., Setif, P. and Duranton, J. (1980). Phorosynrhesis Res. I, 3-16. Lagoutte, B., Setif, P. and Duranton, J. (1981). In “Photosynthesis Vol. 111” (G. Akoyunoglou, Ed.), pp. 237-243. Balaban Intern. Sci. Service, Philadelphia, Pa. Lang, J. C. (1974). Am. Sci. 62, 271-281.


205

Langer, E., Lehner, M., Rudiger, W. and Zickendraht-Wendelstadt, B. (1980). Z . Naturforsch C 35, 367-375. Langridge, J. (1982). I n “Mineral Deposits and the Evolution of the Biosphere” (H. D. Holland and M. Schidlowski, Eds), 83-102. Springer Verlag, Berlin. Larkum, A.,W. D. (1972). J . Mar. Biol. Ass. ( U . K . ) 52, 405-418. Larkum, A. W. D. and Anderson, J. M. (1982). Biochim. Biophys. Acta 679,410-421. Larkum, A. W. D. and Weyrauch. S . K . (1977). Photochem Photobiol. 25, 65-72. Larkum, A. W. D., Drew, E. A. and Crossett, R. N. (1967). J . Ecol. 55, 361-371. Laszlo, J. A. and Gross, E. L. (1980). Arch. Biuchem. Biophvs. 203, 496-505. Latimer, P. and Rabinowitch, E. I. (1957). In “Research in Photosynthesis” (H. Gaffron, Ed.), pp. 100-107. Interscience, New York. Latimer, P., Moore, D. M. and Bryant, F. D. (1968). J . Theor. Biol. 21, 348-367. Lau, R. H., Mackenzie, M. M. and Dolittle, W. F. (1977). J . Bacteriol. 132, 771-778. Lavorel, J. (1980). Biochim. Biophjm Acta 590, 385-400. Lavorel, J. and Etienne, A.-L. (1977). I n “Primary Processes in Photosynthesis” (J. Barber, Ed.), pp. 203-268. Elsevier, North Holland Biomedical Press, Amsterdam. Laycock, M. V. and Wright, J. L. C. (1981). Phytochemistry 20, 1265-1268. Leclerc, J. C., Hoarau, J. and Remy, R. (1979). Biochim. Biophys. Acta 547, 398409. Leclerc, J. C., Coute, A. and Hoarau, J. (1981). I n “Photosynthesis, VI” (G. Akoyunoglou, Ed.), 443454. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Lee, B. D. and Titlyanov, E. A. (1978). Biol. Moryu 2, 47-55. Lee, R . E. (1972). Nature (London) 237, 44-46. Lefort-Tran, M., Cohen-Bazire, G. and Pouphile, M. (1973). J . Ultrastruct. Res. 44, 199-209. Lemasson, C., Tandeau de Marsac, N. and Cohen-Bazire, G. (1973). Proc. Natl. Acad. Sci. U.S.A. 73, 3957-3960. Lemberg, R. (1928). Justus Liebigs Ann. Chem. 461, 4&80. Lemberg, M. R. (1930). Justus Liebigs Ann. Chem. 477, 195-245. Lemberg, M. R. (1954). Rep. Aust. N . Z . Ass. Advmt. Sci. 30,243-64. Lemberg, R. and Barrett, J. (1973). “Cytochromes”. Academic Press, London. Levring, T. (1966). In “Light as an Ecological Factor” (R. Bainbridge, G. C. Evans and 0. Rackham, Eds). Blackwell Scientific, Oxford. Lewin, R. A. (1976). Nature (London) 261, 697-698. Lewin, J., Norris, R. E., Jeffrey, S. W. and Pearson, B. E. (1977). J . Phycol. 13, 259-266. Ley, A. C. and Butler, W. L. (1976). Proc. Natl. Acad. Sci. U.S.A. 73, 3957-3960. Ley, A. C. and Butler, W. L. (1977a). Biochim. Biophys. Acta 462, 290-294. Ley, A. C. and Butler, W. L. (1977b). In “Photosynthetic Organelles, Structure and Function. Special Issue of Plant and Cell Physiol. No. 3” (S. Miyachi, S. Katoh, Y. Fujita and K . Shibata, Eds), pp. 3 3 4 6 . Tokyo Jap. SOC.Plant Physiol; Center Acad. Publ. Japan. Ley, A. C. and Butler, W. L. (1980a). Plant Physiol. 65, 714722. Ley, A. C. and Butler, W. L. (1980b). Biochim. Biophys. Acra 592, 349-363. Ley, A. C., Butler, W. L., Bryant, D. A. and Glazer, A. N. ( 1 977). Plant Physiul. 59, 974-980. Lichtenthaler, A. K. (1968). Planta 81, 104152. Lichtle, C. (1979). Protoplusma 101, 283-299. Lichtle, C. and Thomas, J. C . (1976). Phycologia 15, 393-404. Lichtle, C., Jupin, H. and Duval, J . C. (1980). Biochim. Biophys. Acta 591, 1041 12. Lilley, R. Mc. and Larkum, A. W. D. (1981). Plant Physiol. 67, 5-8.

206


Lipschultz, C. A. and Gantt, E. (1981). Biochemistry 20, 3371-3376. Littler, M. M. (1980). Bot. Marina 23, 161-165. Littler, M. M. and Littler, D. S. (1980). Amer. Nut. 116, 25-44. Littler, M. M. and Murray, S. N. (1974). Mar. Biol. 30,277-291. Lorenzen, C. J. (1972). J . Cens. Int. Explor. Mer. 34, 262-267. Lorenzen, C. J. (1976). In “The Ecology of the Seas” (D. H. Cushing and J. J. Walsh, Eds), pp. 173-185. Blackwell Scientific, Oxford. Ludlow, C. J. and Park, R. B. (1969). Plant Physiol. 44, 540-543. Luehrsen, K. R., Nicholson, D. E., Eubanks, D. C. and Fox, G. E. (1981). Nature 293, 755-756. Lundell, D. J., Williams, R. C. and Glazer, A. N. (1981). J . Biol. Chem. 256, 3580-3592. Luning, K. and Dring, M. J. (1979). Helgolander wiss Meeresunters 32, 403424. Lutz, M. (1977). Biochim. Biophys. Acta 460, 404430. Lutz, M. (1980). CR-Conf. Int: Spectrose Raman 7th (W. F. Murphy, Ed.), pp. 520-523. Lutz, M., Agalidis, I., Hervo, G., Codgell, R. J. and Reiss-Husson, F. (1978). Biochim. Biophys. Acta 503, 287-303. Lutz, M., Hoff, A. J. and Brehamet, L. (1982). Biochim. Biophys. Acta679, 331-341. McCartin, P. J. (1963). J . Phys. Chem. 67, 513-515. McCarty, R. (1979). Annu. Rev. Plant Physiol. 30, 79-104. McCaslin, D. R. and Tamford, C. (1981). Biochemistry 20,5212-5221. MacColl, R. (1982). Photochem. Photobiol. 35,899-904. MacColl, R. and Berns, D. S. (1978). Photochem. Photobiol. 27, 343-349. MacColl, R. and Berns, D. S. (1979). Trends in Biochem. Sci. 44, 44-47. MacColl, R., Berns, D. S. and Gibbons, 0. (1976). Arch. Biochem. Biophys. 177, 265-275. MacColl, R., Csatorday, K., Berns, D. S. and Traeger, E. (1980). Biochemistry 19, 28 17-2820. MacColl, R., O’Connor, G., Crafton, G. and Csatorday, K. (1981). Photochem. Photobiol. 34, 719-723. Machold, 0. and Hoyer-Hansen, G. (1976). Carlsberg Res. Commun. 41, 359-366. Machold, O., Simpson, D. J. and Lindberg-Moller, B. (1979). Carlsberg Res. Commun. 44, 234-235. McIntosh, A. R., Manikowski, H. and Bolton, J. R. (1981). In “Photosynthesis” ( G . Akoyunoglou, Ed.), pp. 687-695. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania. McKie, J., Lucas, C. and Smith, A. (1981). Photochemistry 20, 1547-1549. MacKinney, G. (1941). J . Biol. Chem. 140, 315-322. Malkin, R. and Bearden, A. (1971). Proc. Natl. Acad. Sci. U S A . 68, 16-19. Malkin, R. and Bearden, A. J. (1975). Biochim. Biophys. Acta 396, 250-259. Malkin, R. and Bearden, A. J. (1978). Biochim. Biophys. Acta 505, 147-181. Malkin, R. and Fork, D. C. (1980). Plant Physiol. 67, 580-583. Malkin, S . , Armond, P. A., Mooney, H. A. and Fork, D. C. (1981). Plant Physiol. 67, 570-519. Mallams, A. K., Waight, E. S., Weedon, B. C. L., Chapman, D. J., Haxo, F. T., Goodwin, T. W. and Thomas, D. M. (1967). Proc. Chem. SOC.London, pp. 301-302. Malone, T. C. (1980). In “The Physiological Ecology of Phytoplankton” (1. Morris, Ed.), 433-463. Blackwell Sci. Pub., Oxford. Mandelli, E. F. (1968). J . Phycol. 4, 347-348.


207

Mangel, M., Berns, D. S. and Ilani, A. (1975). J . Membrane Biol. 20, 171-180. Mann, K . H. (1973). Science 182, 975-978. Mann, J . E. and Myers, J. (1968). J . Phycol. 4, 349-355. Margulis, L. (1970). Origin of Eukaryotic Cells. New Haven. Markwell, J . P., Thornber, J. P. and Boggs, R. T. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 1233-1235. Markwell, J. P., Thornber, J. P. and Skrdla, M. P. (1980). Biochim. Biophys. Acta 591, 39 1-399. Markwell, J . P., Nakatani, H. Y., Barber, J . and Thornber, J. P. (1980). FEBS Lett. 122, 149-153. Marco, J . and Gamier, J . (1981). Biochim. Biophys. Actu 637,473480. Marra, J. (1978). Mar. B i d . 46, 191-202. Mathews, B. W., Fenna, R. E., Bolognesi, M. C., Schmid, M. F. and Olson, J. M . (1979). J . Mol. Biol. 131, 259-285. Mathis, P. and Paillotin, G . (1981). In “Biochemistry of Plants” (P. K. Stumpf and E. E. Conn, Eds), 8, 98-161. Academic Press, New York. Matsubara, H., Mase, T., Wakabayaski, S. and Wada, K . (1980). In “The Evolution of Protein: Structure and Function” (D. S. Sigman and M. A. B. Brazier, Eds.) pp. 245-262. Academic Press, New York. Mattoo, A. K., Pick, U . , Hoffman-Folk, H . and Edelman, M. (1981). Proc. Natl. Acad. Sci. U.S.A. 78, 1572-1576. Mauzerall, D. (1973). Ann. N . Y . Acad. Sci. 206, 483--494. Mauzerall, D. (1978). In “The Photosynthetic Bacteria” (R. K. Clayton and W. R. Sistom, Eds), pp. 223-231. Plenum Press, New York. Mauzerall, D. and Hong, F. H. (1975). In “The Porphyrins” (K. V. Smith, Ed.), pp, 701-725. Elsevier, Amsterdam. Mauzerall, D. C. and Piccioni, R. G. (1981). In “Oxygen and Living Processes” (D. L. Gilhert, Ed.). Springer-Verlag, New York, pp. 102-123. Melis, A. (1978). FEBS Lett. 95, 202-206. Melis, A. and Brown, J . S . (1980). Proc. Natl. Acad. Sci. U . S . A .77, 47124716. Melis, A. and Harvey, G. W. (1981). Biochim. Biophys. Acta 637, 138-145. Melis, A. and Homann, P. H. (1976). Photochem. Photobtol. 23, 343-350. Mel’nikov, S. S. and Yevstigneyev, V. B. (1964). Biophysics 9. 447456. Menke, W. (1962). Annu. Rev. Plant Physiol. 13, 27-44. Menke, W. and Schmid, G. M. (1980). Z . Naturforsch. C 35, 461466. Metz, J. and Bishop, N. I. (1980). Biochem. Biophys. Res. Commun. 94, 560-566. Millar, A. and Kraft, G. (1983). Proc. R. SOC. Victoria (in press). Miller, K. R. (1976). J . Ultrastruct. Res. 54, 159-167. Miller, K. R. and Staehelin, L. A. (1976). J . Cell. Biol. 68, 30-47. Mimuro, M. and Fujita, Y. (1977). Biochim. Biophys. Acta 459, 376389. Mimuro, M. and Fujita, Y . (1980). Plant Cell. Physiol. 21, 3 7 4 5 . Mishkind, M. and Mauzerall, D. (1980). Mar. Biol. 56, 261-265. Mitchell, P. (1966). B i d . Rev. (Cambridge) 41, 445-502. Mohanty, P., Braun, B. Z., Govindjee and Thornber, J. P. (1972). Plant Cell Physiol. 13, 81-91. Mohr, H. (1980). In “The Blue Light Syndrome” (H. Senger, Ed.), pp. 97-109. Springer-Verlag, Berlin. Molinier, R. (1960a). Vegetatio 9, 121-192. Molinier, R. (1960b). Vegefatio 9, 217-312. Mooney, H. A. and Gulman, S. L. (1979). In “Topics in Plant Population Biology” (0.

208


T. Solbrig, S. Jain, G. B. Johnson and P. H. Raven, Eds), pp. 316337. Coluinbia Univ. Press, New York. Moore, A. L., Dirks, G., Gust, D. and Moore, T. A. (1980). Photochem. Photobid. 32, 69 1-696. Morel, A. (1974). In “Optical Aspects of Oceanography” (N. G. Jerlov and E. Stcernan Nielsen, Eds), pp. 1-24. Academic Press, London. Morel, A. and Prieur, L. (1977). Limnol. Oceanogr. 22, 702-709. Morschel, E. and Wehrmeyer, W. (1979). Ber. Deutsch. Bot. Ges. 92, 40341 1. Morschel, E., Koller, K.-P., Wehrmeyer, W. and Schneider, H. (1977). Cytobiology 16, 118- 129. Morschel, E., Koller, K.-P. and Wehrmeyer, W. (1980). Arch. Microbiol. 125, 43-51. Morschel, E., Wehrmeyer, W. and Koller, K.-P. (1980). Eur. J . Cell. Biol. 21,319-327. Moscowitz, A., Krueger, W. C., Kay, I. T., Skews, G. and Bruckensteins, S. (1964). Proc. Natl. Acad. Sci. U S A . 52, 1190-1194. Moss, G. P. and Weedon, B. C. L. (1976). In ”Chemistry and Biochemistry of Plant Pigments” 2nd ed. (T. W. Goodwin, Ed.), pp. 149-224. London, Academic Press. Muckle, G. and Rudiger, W. (1977). Z. Naturforsch. 32, 957-962. Muckle, G., Otto, J. and Rudiger, W. (1978). Z. Physiol. Chem. 359, 345-355. Mullet, J. F. and Arntzen, C. J. (1980). Biochim. Biophys. Acta 589, 100-1 17. Mullet, J. E. and Arntzen, C. J. (1981). Biochim. Biophys. Acta 635, 236-248. Mullet, J. E., Burke, J. J. and Arntzen, C. J. (1980a). Plant Physiol. 65, 814822. Mullet, J. E., Burke, J. J. and Arntzen, C. J. (1980b). Plant Physiol. 65, 823-827. Murakami, S. and Packer, L. (1970). Plant Physiol. 45, 289-299. Murakami, S., Torres-Periera, J. and Packer, L. (1977). In “Bioenergetics of Photosynthesis” (Govindjee, Ed.), pp. 556-618. Academic Press, New York. Murata, N. (1969a). Biochim. Biophys. Acta 172, 242-251. Murata, N. (1969b). Biochim. Biophys. Acta 189, 171-181. Murata, N. (1970). Biochim. Biophys. Acta 205, 379-389. Murata, N., Nishimura, M. and Takamiya, A. (1966a). Biochim. Biophys. Acta 120, 23-33. Murata, N., Nishimura, M. and Takamiya, A. (1966b). Biochim. Biophys. Acta 126, 23C243. Murata, T. (1981). In “Photosynthesis 111” (G. Akoyunoglou, Ed.), pp. 397404. Balaban Intern. Sci. Serv. Philadelphia, Pennsylvania, USA. Murata, T. and Ishikawa, C. (1981). Biochim. Biophys. Acta 635, 341-347. Myers, A., Preston, R. D. and Ripley, G. W. (1956). Proc. Roy. Soc. B. 144,450-459. Myers, J. (1946). J. Gen. Physiol. 29, 429440. Myers, J. and Graham, J.-R. (1963). Plant Physiol. 38, 105-1 16. Myers, J. and Graham, J.-R. (1971). Piant Physiol. 48, 282-286. Myers, J. and Kratz, W. A. (1955). J. Gen. Physiol. 39, 11-22. Myers, J., Graham, J. and Wang, R. T. (1978). J. Phycof. 14, 513-518. Myers, J., Graham, J.-R. and Wang, R. T. (1980). Plant Physiol. 66, 1144-1149. Nakamura, K., Ogawa, T. and Shibata, K. (1976). Biochim. Biophys. Acta 423, 227-236. Nakayama, K. and Yamaoka, T. and Katoh, S. (1979). Plant Cell Physiol. 20, 1565-1 576. Nelson, N. (1977). In “Encyclopedia of Plant Physiology N.S.” Vol. 5. “Photosynthesis I” (A. Trebst and M. Avron, E d ) , pp. 393-409. Springer, Berlin. Nelson, N. (1981). Current Topics in Bioenergetics 11, 1-33. Nelson, N. and Neumann, J. J. (1972). J. Biol. Chem. 247, 1817-1824. Nes, W. R. and Nes, W. D. (1980). “Lipids in Evolution”. Plenum Press, New York. pp. 244.


209

Neushul, M. (1967). Ecology 48. 83-94. Neushul, M. (1971). J . Ultrastruct. Res. 37, 532. Newcomb, E. H . and Pugh, T. D. (1975). Nature, (London) 253, 533-534. Newman, M. J. and Rood, R. T. (1977). Science 198, 1035-1037. Newman, P. J. and Sherman, L. A. (1978). Biochim. Biophys. Actu 503, 343-361. Nies, M. and Wehrmeyer, W. (1980). Plarrrlc f Berl.) 150, 330-337. Nobel, P. S. (1974). “Introduction to Biophysical Plant Physiology”. W. H. Freeman and Co., San Francisco. Nordhorn, G., Weidner, M. and Willenbrink, J. (1976). Z. F‘panzen Physiol. 805, 153- 165. Norton, T. A., Ebling, F. J. and Kitching. J . A. (1971). In “Fourth European Marine Biology Symposium (D. J. Crisp, Ed.), pp. 409-432. Cambridge University Press, Cambridge. Nugent, J. H. A.. Moller, B. L. and Evans, M. C . W. (l980a). FEBS Lett. 121,355- 357. Nugent, J . H. A., Moller, B. L. and Evans. M. C . W. (1980b). Biochim. Biophys. Acta 634, 249-255. Nugent, J. H. A., Stewart, A. C. and Evans. M. C. W. (1981). Biochim. Biophys. Actu 635, 488497. Nultsch, W. and Rueffer, U. (1981). Mar. Biol. 62. 111-117. O’Carra, P. and O’heocha, C. (1976). In “Chemstry and Biochemistry of Plant Pigments” (T. W. Goodwin, Ed.) 2nd ed. 1,328-376. Academic Press, New York. Odum, E. P. (1969). Science 164, 262-270. Offner, G. D., Brown-Mason, A. S., Erhardt, M. M. and Troxler, R. F. (1981). J . Biol. Chem. 256, 12167-12175. Ogawa, T. and Vernon, L. P. (1970). Biochim. Biophys. Acra 197, 332-334. Ogawa, T., Obata, F. and Shibata, K. (1966). Biochim. Biophys. Acra 112, 223-234. Ogawa, T., Nakamura, K. and Shibata, K. (1966). Arch. Hydrobiol. Suppl. 49, 37 4 8 . Ohad, I . , Clayton, R. K. and Bogorad, L. (1979). Proc. Natl. Acad. Sci. U.S.A. 76, 5655-5659. Ohad, I., Schneider, H.-J. A. W., Gendel, S. and Bogorad, L. (1980). Plant Physiol. 65, 6-12. O’hEocha, C. (1971). Oceanogruph. Mur. Biol. Annu. Rev. 9,61-82. Oh-hama, T. and Fujita, Y. (1979). Plant Cell Physiol. 20, I341 - 1347. Oh-hama, T. and Hase, E. (1981). Plant CeN Physiol. 22, 747-757. Oh-hama, T., Matsuka, M. and Hase, E. (1968). I n “Comparative Biochemistry and Biophysics of Photosynthesis” (K. Shibata, R. Fuller, A. Takamiya, A. T. Jagendorf, Eds), pp. 279-290. University Park Press, State College, Pennsylvania, USA. Oh-hama, T., Seto, H., Otake, N. and Miyachi, S. (1982). Biochem. Biophys. Res. Commun. 105,647-652. Ohki, K., Isoni, T. and Fujita, Y. (1980). I n “The Blue Light Syndrome” (H. Senger, Ed.), pp. 597-604. Springer Verlag, Berlin. Okada, S., Kanematsa, S. and Asada, K . (1979). FEBS Lett. 103, 106-1 10. Okayama, S. and Butler, W. L. (1972). Plant Physiol. 49, 769-774. Okada, S., Kanematsu, S. and Asada, K. (1979). FEBS Lett. 103, l o b 1 10. O’Kelley, C . J. (1982). Botanica Marina 25, 133-137. Olson, J. M. (1980). Biochim. Biophys. Acta 594, 33-51. Olson, J. M. (1981a). Ann. New York Acad. Sci. 361, 8-19. Olson, J. M. (1981b). Biosystems 14, 89-94. Oltmanns, F. (1892). Jb. Wiss. Bot. 23, 349440. Oquist, G., Samuelsson, G. and Bishop, N. I. (1980). Physiol. Plant 50,63-70. Osmond, C. B. (1981). Biochim. Biophys. Acta 639,77-98.

210


Ovchinnikov, Yu. A., Abdulaev, N. G., Feigina, M. Yu., Kiselev, A. V. and Lobanov, A. (1979). FEBS Lett. 100, 219-224. Owen, T., Cess, R. D. and Ramanathan, V. (1979). Nature (London) 277, 640-642. Packer, L., Barnard, A. and Deamer, D. W. (1967). Plant Physiol. 42, 283-293. Padan, E. (1979). Annu. Rev. Plant Physiol. 30, 27-40. Paillotin, G. (1976a). J . Theor. Biol. 58, 219-235. Paillotin, G. (1976b). J . Theor. Biol. 58, 237-252. Papagiorgiou, G. (1975). In “Bioenergetics of Photosynthesis” (Govindjee, Ed.), pp. 320-371. Academic Press, New York. Park, R. B. (1965). J . Cell Biol. 27, 151-161. Parker, B. C., Simmons, G. M. Jr, Love, F. G., Wharton, R. A. Jr and Seaburg, K. G. (1981). Bioscience 131, 656661. Parsons, T. R., Takahashi, M. and Hargrave, B. (1977). “Biological Oceanographic Processes” 2nd ed. Pergamon Press, Oxford. Pecci, J. and Fujimori, E. (1969). Biochim. Biophys. Acta 188, 230-236. Pelevin, V. N. and Rutkovskaya, V. A. (1977). Oceanology 17, 28-32. Perchorowicz. J. T., Raynes, D. A. and Jensen, R. G. (1981). Proc. Narl Acad. Sci. U.S.A. 78, 2985-2989. Perry, M. J., Talbot, M. C. and Alberte, R. S. (1981). Mar. Biol. 62, 91-101. Peterson, R. B., Dolan, E., Calvert, H. E. and Bacon, K. E. (1981). Biochim. Biophys. Acta 634, 237-248. Petke, J. D., Shipman, L. L., Maggiova, G. M. and Christoffersen, R. E. (1981). J . Am. Chem. Soc. 103, 46224623. Pfau, J., Ruffer, U. and Nultsch, W. (1979). Ber. Deutsch. Bot. Ges. 92, 695-715. Pianka, E. R. (1970). Am. Nut. 104, 592-597. Porra, R. J. and Grimme, L. H. (1974). Ann. Biochem. 57, 255-267. Porra, R. J. and Grimme, L. H. (1978). Int. J . Biochem. 9, 883-886. Porter, G . , Tredwell, C. J., Searle, G. F. W. and Barber, J. (1978). Biochim. Biophys. Acta 501, 232-245. Powles, S. B. and Osmond, C. B. (1978). Aust. J . Plant Physiol. 5 , 619-629. Powles, S. B., Osmond, C . B. and Thorne, S. W. (1979). Plant Physiol. 64, 982-988. Powls, S. R. and Britton, G. (1976). Biochim. Biophys. Acta 453, 270-276. Prezelin, B. P. (1976). Planta 130, 225-233. Prezelin, B. B. and Alberte, R. S. (1978). Proc. Natl Acad. Sci. U.S.A. 75, 1801-1804. Prezelin, B. B. and Boczar, B. A. (1981). In “Photosynthesis 111” (G. Akoyunoglou, Ed.), pp. 417-426. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Prezelin, B. B. and Haxo, F. T. (1976). Planta 128, 133-141. Prezelin, B. B. and Matlick, H. A. (1980). Mar. Biol. 58, 85-96. Prezelin, B. B. and Sweeney, B. M. (1978). Mar. Biol. 48, 27-35. Prezelin, B. B., Ley, A. C . and Haxo, F. T. (1976). Planta 130, 251-256. Pullin, C. A., Brown, R. G. and Evans, E. H. (1979). FEBS Lett. 101, 110-112. Rabinowitch, E. (1945). “Photosynthesis and Related Processes”. Vol. I. Interscience, New York. Rabinowitch, E. I. (1951). “Photosynthesis and Related Processes”. Vol. 11. Pt. 1 . Interscience, New York. Rabinowitch, E. (1956). “Photosynthesis and Related Processes”. Vol. 11. Pt. 2. Interscience, New York. Rademaker, H., Hoff, H. J. and Duysens, L. N. M. (1979). Biochim. Biophys. Acta 546, 248-255. Radmer, R. J. and Kok, B. (1977). In “Encyclopedia of Plant Physiology 5 . Photosynthesis I” (A. Trebst and M. Avron, Eds), pp. 1 2 4 1 35. Springer-Verlag, Berlin.


21 1

Raff, R. A. and Mahler, H. R. (1972). Science 177, 575-582. Ragan, M. A. and Chapman, D. J. (1978). “A Biochemical Phylogeny of the Protists”. Academic Press, New York. Raghavan, N. V., Das, P. K. and Bobrowski, K. (1981). J . Am. Chem. SOC.103, 4569-4573. Ramus, J. (1978). J. Phycol. 14, 352-362. Ramus, J. and Rosenberg, G. (1980). Mar. Biol. 56, 21-28. Ramus, J., Beale, S. I., Mauzerall, D. and Howard, K. L. (1976). Mar. Biol. 37, 223-229. Ramus, J., Beale, S. I. and Mauzerall, D. (1976). Mar. Biol. 37, 231-238. Ramus, J., Lemons, F. and Zimmerman, C. (1977). Mar. Biol. 42, 293-303. Raven, J. A. (1972). New Phytol. 71, 227-247. Raven, J. A. (1977). Adv. Bot. Res. 5, 154-219. Raven, J. A. (1978). Proc. Fourth Int. Congr. Photosynth. (D. 0. Hall, J. Coombs and T. W. Goodwin, Eds), pp. 147-155. Biochemical SOC.,London. Raven, J. A. and Beardall, J. (1982). Plant Cell Environ. 5, 117-124. Raven, J. A. and Glidewell, S. M. (1978). Plant Cell Environ. 1, 185-197. Raven, J. A., Smith, F. A. and Glidewell, S. M. (1979). New Phytol. 83, 299-309. Raven, P. H. (1970). Science 169, 641-646. Raymont, J. E. (1980). “Plankton and productivity in the oceans” Vol. 1 ., “Phytoplankton”. 2nd ed. Pergamon Press, Oxford. Redlinger, T. and Gantt, E. (1981). In “Photosynthesis 111” (G. Akoyunoglou, Ed.), pp. 257-262. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Reger, B. J. and Krauss, R. W. (1970). Plant Physiol. 46, 568-575. Reimer, T. O., Barghoorn, E. S. and Margulis, L. (1979). Precambr. Res. 9, 93-104. Reinman, S. and Mathis, P. (1981). Biochim. Biophys. Acta 635, 249-258. Reinman, S. and Thornber, J. P. (1979). Biochim. Biophys. Acta 547, 188-197. Reinman, S., Mathis, P., Conjeaud, H. and Stewart, A. (1981). Biochirn. Biophys. Acta 635, 429433. Remsen, C. (1978). In “The Photosynthetic Bacteria” (R. K. Clayton and W. R. Sistrom, Eds), pp. 31-61. Plenum Press, New York. Remy, R. and Hoarau, J. (1978). In “Chloroplast Development” (G. Akoyunoglou, Ed.), pp. 235-240. Elsevier, Amsterdam. Renger, G., Hagemann, R. and Dohnt, G. (1981). Biochim. Biophys. Acta 636, 17-26. Reynolds, J. A. (1979). In “Methods in Enzymology” (C. H. W. Hirs and S. N. Timasheff, Eds). Academic Press, New York. Ried, A. B. and Reinhardt, B. (1977). Biochim. Biophys. Acta 460,25-35. Ried, A. and Reinhardt, B. (1980). Biochim. Biophys. Acta 592, 7 6 8 6 . Rijgersberg, C. P. and Amesz, J. (1980). Biochim. Biophys. Acta 593, 261-271. Riley, J. P. and Wilson, T. R. S. (1967). J . Mar. Biol. Assoc. (U.K.) 47, 351-362. Romeo, A. (1981). XI11 Intern. Bot. Congr. Abstracts p. 224. Australian Acad. Sci., Canberra. Rosinski, J., Hainfield, J. F., Rigby, M. and Siegelman, H. W. (1981). Ann. Bot. 47, 1-12. Rudiger, W. (1975). Ber. Deutsh. Bot. Ges. 88, 125-139. Riidiger, W. (1980). In “Pigments in Plants” (F. C. Cyzan, Ed.), 2nd Ed., pp. 314-351. Gustav Fisher, Stuttgart. Ruffer, U., Nultsch, W. and Pfau, J. (1978). Helgolander wiss Meeresuntersuchg 31, 333-346. Ruffer, U., Pfau, J. and Nultsch, W. (1981). Z. Pjanzenphysiol. 101, 283-293. Rusckowski, M. and Zilinskas, B. A. (1980). Plant Physiol. 65, 392-396. Rutherford, A. W. (1981). Biochim. Biophys. Res. Commun. 102, 1065-1070.

212

A. W.D. LARKUM AND JACK BARRETT

Rutherford, A. W. and Mullet, J. E. (1981). Biochirn. Biophys. Acta 635, 225-235. Rutherford, A. W., Mullet, J. E. and Crofts, A. R. (1981a). FEBS Letr. 123, 235-237. Rutherford, A. W., Paterson, D. R. and Mullet, J. E. (1981b). Biochirn. Biophys. Acta 635, 205-214. Ryrie, I. J., Anderson, J. M. and Goodchild, D. J. (1980). Eur. J. Biochem. 107, 345-354. Ryther, J. (1956). Limnol. Oceanogr. 1, 72-84. Ryther, J. H. and Menzel, D. W. (1959). Limnol. Oceanogr. 4, 492-497. Salares, V. R., Young, N. M., Carey, P. R. and Bernstein, H. J. (1977). J . Ramun. Spectrosc. 6, 282-288. Sandmann, G. and Boger, P. (1980). Plant Science Lett. 17, 417-424. Sane, P. V. (1977). In “Encyclopedia of Plant Physiol. N.S. Vol. 5 . Photosynthesis I” (A. Trebst and M. Avron, Eds), pp. 522-542. Springer-Verlag, Berlin. Sane, P. V., Park, R. B. (1970). Biochem. Biophys. Res. Commun. 41, 206-210. Sane, P. V., Goodchild, D. J. and Park, R. B. (1970). Biochim. Biophys. Acta 218, 162-1 78, Satoh, K. (1970a). Plant Cell Physiol. 11, 15-27. Satoh, K . (1970b). Plant Cell Physiol. 11, 187-197. Satoh, K. (1980). FEBS Letr. 110, 53-56. Satoh, K. (1981). In “Photosynthesis 111” (G. Akoyunoglou, Ed.), pp. 607-616. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Satoh, K. and Butler, W. L. (1978a). Plant Physiol. 62, 373-379. Satoh, K. and Butler, W. L. (1978b). Biochim. Biophys. Acra 502, 103-110. Saunders, V. A. and Jones, 0. T. G. (1975). Biochim. Biophys. Acta 3%, 220-228. Sauer, K. (1975). In “Bioenergetics of Photosynthesis” (Govindjee, Ed.), pp. 115-1 81. Academic Press, New York. Savidge, G. (1980). Mar. Biol. Lett. 1, 295-300. Schaffernicht, H. and Wolfgang, J. (1Y8l). Photochem. Photobiol. 34, 223-232. Scheer, H. (1981). Angewandte Chemie 20, 241-261. Scheer, H. and Katz, J. J. (1975). In “Porphyrins and Metalloporphyrins” (Kevin Smith, Ed.), pp. 399-524. Elsevier, Amsterdam. Scheer, H. and Kufer, W. (1977). Z. Naturforsch. C 32, 513-519. Scheibe, J. (1972). Science 176, 1037-1039. Schidlowski, M. (1980). In “Biogeochemistry of Ancient and Modern Environments” (P. A. Trudinger and M. R. Walter, Eds), pp. 47-54. Aust. Acad. Sci., Canberra. Schidlowski, M., Appel, P. W. U., Eichmann, R. and Junge, C. E. (1979). Geochim. Cosmochim. Acta 43, 189-199. Schiff, J. A. (1978). In “Chloroplast Development” (G. Akoyunoglou and J. H. Argyroudi-Akoyunoglou, Eds), pp. 747-767. Elsevier/North Holland, Amsterdam. Schiff, J. A. (1980). In “The Blue Light Syndrome” (H. Senger, Ed.), pp. 495-511. Springer-Verlag, Berlin. Schiff, J. A. (1981a). Ann. N.Y. Acad. Sci. 361, 166-189. Schiff, J. A. (1981b). Biosystems 14, 123-147. Schimper, A. F. W. (1885). Jb. wiss. Botan. 16, 1-9. Schonbohrn, E. (1965). Z. Pjanzenphysiol. 53,344355. Schonbohm, E. (1978). Progress in Botany 40,185-196. Schopf, T. J. M. (1980). “Paleoceanography” Harvard Univ. Press, Cambridge, Massachusetts, USA. Schreiber, U. (1979). FEBS Lett. 107, 4-9.


213

Schreiber, U. (1980). Biochim. Biophys. Acta 591, 361-371. Schutt, F. (1890). Ber. Deut. Bot. Ges. 8. 9-32. Schwartz. R. M. and Dayhoff, M. 0. (1978). Science 199, 395403. Scott, B. and Gregory, R. P. (1975a). Biochern. J . 148, 487497. Scott, B. and Gregory, R. P. (1975b). Biochem. J . 149, 341-347. Sculley, M. J., Duniec, J. T., Thorne, S. W., Chow, W. S. and Boardman, N. K. (1980). Arch. Biochem. Biophys. 201, 339-346. Searle, G. F. W. and Wessels, J. S. C. (1978). Biochirn. Biophys. Acta 459, 40241 1. Searle, G . F. W., Barber, J., Porter, G. and Treadwell, C. J. (1978). Biochim. Biophys. Acta 501, 245-256. Sears, J. R. and Cooper, R. A. (1978). Mar. Biol. 44, 309-314. Sears, J. R. and Wilce, R. T. (1975). Ecol. Monogr. 45, 337-365. Sebald, W. and Hoppe, J. (1981). Current Topics in Bioenergetics 12, 1-64. Seckbach, J. (1972). Microbiol. 5, 133-142. Seely, G. R. (1977). In “Primary Processes of Photosynthesis” (J. Barber, Ed.), pp. 3-53. Elsevier, Amsterdam. Seely, G . R. and Jensen, R. G. (1965). Spectrochimica Acta 21, 1835-1845. Seewaldt, E. and Stackebrandt, E. (1982). Nature 295, 618-620. Seiburth, J. M. and Jensen, A. (1968). J . Exp. Mar. Biol. Ecol. 2, 174189. Seiburrh, J. M. and Jensen, A. (1969). J . Exp. Mar. Biol. Ecol. 3, 275-289. Senger, H. (1980). ”The Blue Light Syndrome” Springer-Verlag, Berlin. Sengcr, H. (1982). Photochern. Photobiol. 35, 91 1-920. Senger, H. and Bishop, N. I. (1967). Nature 214, 140-142. Senger, H. and Fleischhacker, Ph. (1978). Physiol. Plant 43, 3542. Senger, H. and Strassberger, G. (1978). In “Chloroplast Development” (G. Akoyunoglou and J. H. Argyroudi-Akoyunoglou, Eds), pp. 367-377. Elsevier, Amsterdam. Senn, G. (1908). “Die Gestalts-und Lageveranderungen der pflanzenChromatophoren”. Engelmann, Leipzig. Senn, G. (1919). 2.Bot. 11, 81-144. Setif, P., Acker, S., Lagoulto, B. and Duranton, J. (1980). Photosynthetic Res. 1, 17-27. Setif, P., Acker, S., Lagoulte, B. and Duranton, J. (1981). In “Photosynthesis 111”, (G. Akoyunglou, Ed.), pp. 503-5 1 1. Balaban Intern. Sci. Services. Seybold, A. (1932). Planta 18, 479485. Seybold, A. (1933). Planta 20, 577-584. Seybold, A. (1934). Jahrb. wiss. Botan. 79, 593-601. Sewe, K. U. and Reich, R. (1977). Z. Naturforsch. C 32, 161-171. Shepherd, S. A. and Womersley, H. B. S. (1976). Trans. R . SOC.South Aust. 100, 177-191. Shimura, S. and Fujita, Y. (1973). Plant Cell Physiol. 4, 341-352. Shimura, S. and Fujita, Y. (1975). Mar. Biol. 33, 185-194. Shiozawa, J. A., Alberte, R. S. and Thornber, J. P. (1974). Arch. Biochem. Biophys. 165, 388-397. Shipman, L. L. (1980). Photochem. Photobiol. 31, 157-167. Shipman, L. L., Cotton, T. M., Norris, J. R. and Katz, J. J. (1976). Proc. Natl Acad. Sci. U S A . 93, 1791-1794. Shubin, L. M., Bekasova, 0.D. and Evstigneev, V. D. (1979). Biophysics 24,472475. Shuvalov, V. A., Dolan, E. and Ke, B. (1979). Proc. Natl Acad. Sci. U.S.A. 76, 17&173. Sidler, W . , Gysi, J., Isker, E. and Zuber, H. (1981). In “Photosynthesis 111” (G.

214


Akoyunoglou, Ed.), pp. 583-594. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Siefermann-Harms, D. (1980a). Dev. Plant Biol. 6 , 331-340. Siefermann-Harms, D. (1980b). In “Biogenesis and Function of Plant Lipids” (P. Mazliak, P. Benveniste, C. Costes and R. Douce, Eds), pp. 331-340. Elsevier, Amsterdam. Siefermann-Harms, D. and Ninneman, H. (1979). FEBS Lett. 104, 71-77. Siefermann-Harms, D. and Ninnemann, H. (1 982). Photochem. Photobiol. 35, 7 19-732. Siefermann-Harms, D. and Yamamoto, H. Y. (1974). Proc. Natl Acad. Sci. U . S . A .71, 807-810. Siegelman, H. W. (1982). Plant Physiol. 70, 887-897. Sigalat, C. and de Kouchkovksy, Y. (1974). “Proceedings of the 3rd Int. Congress on Photosynthesis” (M. Avron, Ed.), pp. 621-627. Elsevier, Amsterdam. Simionescu, C. I., Mora, R. and Simionescu, B. C. (1978). Bioelectrochemistry Bioenergetics 5 , 1-17. Simpson, D. J. (1979). Carlsberg Res. Commun. 44,305-336. Smayda, T. J. (1970). Oceanogr. Marine Biol. Ann. Rev. 8, 353-414. Smith, F. A. and Walker, N. A. (1980). New Phytol. 86, 245-259. Smith, R. C. and Calkins J. (1976). Limnol. Oceanogr. 21, 746-749. Smith, R. C., Baker, K. S., Holm-Hansen, 0. and Olson, R. (1980). Photochem. Photobiol. 31, 585-592. Song, P . 4 . (1980). In “The Blue Light Syndrome” (H. Senger, Ed.), pp. 157-171. Springer-Verlag, Berlin. Song, P.-S., Koka, P., Prezelin, B. B. and Haxo, F. T. (1976). Biochemistry 15, 4422-4427. Sonneveld, A., Duysens, L. N. M. and Moerdijk, A. (1980). Proc. Natl Acad. Sci. U.S.A. 10, 5889-5893. Spence, D. H. N. (1975). In “Light as an Ecological Factor: 11” (G. C. Evans, R. Bainbridge and 0. Rackham, Eds), pp. 93-133. Blackwell, Oxford. Staehelin, L. A., Giddings, T. H., Badami, P. and Kryzymowski, W. W. (1978). In “Light Transducing Membranes” (D. W. Deamer, Ed.), pp. 335-355. Academic Press, New York. Stellwagen, E. (1978). Nature 275, 73-74. Steele, J. H. (1962). Limnol. Oceanogr. 7, 137-150. Steemann Nielsen, E. (1961). Physiol. Plantar. 14, 868-876. Steemann Nielsen, E. (1975). “Marine Photosynthesis” Elsevier Oceanography Ser. 13, Elsevier, Amsterdam. Steinbeck, K. E., Burke, J. J. and Arntzen, C. J. (1979). Arch. Biochem. Biophys. 195, 546-557. Sterling, C. (1964). Acta Crystallogr. 17, 1224-1228. Stewart, A. C. (1980). FEBS Lett. 114, 67-72. Stewart, A. C. and Bendall, D. S. (1979). FEBS Lett. 107, 308-312. Stewart, A. C. and Bendall, D. S. (1980). Biochem. J . 188, 351-361. Stewart, A. C. and Bendall, D. S. (1981). Biochem. J . 194, 877-887. Stewart, K. D. and Mattox, K. R. (1975). Bot. Rev. 41, 104-125. Stewart, K. D. and Mattox, K. R. (1978). Biosystems 10, 145-152. Strain, H. H. (1958). 32nd Annual Priestley Lectures, 180-191. (Penn. State University, University Park, Pa, USA). Strain, H. H., Svec, W. A., Aitzetmuller, K., Grandolfo, M. C., Katz, J. J., Kjosen, H., Norgard, S., Liaaen-Jensen, S . , Haxo, F. T., Wegfahrt, P. and Rapoport, H. (1971). J. Am. Chem. Soc. 93, 1823-1825. Stransky, H. and Hager, A. (1970a). Arch. Mikrobiol. 71, 164-168.


215

Stransky, H. and Hager, A. (1970b). Arch. Mikrohiol. 72, 8 4 9 2 . Sugahara, K., Murata, N. and Takayima, A. (1971). Plant Cell Physiol. 12, 377-385. Sugimura, Y., Hase, T., Matsubara, H. and Shimokoriyama, M. (1981). Biochemistry 90,1213-1219. Sugiyama, K.-I. and Murata, N. (1978). Biochim. Biophvs. Acta 503, 107-1 19. Sundqvist, C., Bjorn, L. 0. and Virgin, H. I. (1980). In “Results and Problems in Cell Differentiation Vol. 10. Chloroplasts” (J. Reinart, Ed.), pp. 20 1-224. SpringerVerlag, Berlin. Svedberg, T. and Lewis, N. B. (1928). J. Am. Chem. Soc. 50, 525-536. Swarthoff, T., Gast, P., Hoff, A. J. and Amesz, J. (1981a). FEBS Lett. 130, 93-98, Swarthoff, T., Gast. P., van der Veek-Horsley, K . M., Hoff, A. J. and Amesz, J. (1981b). FEBS Lett. 131, 331-334. Swift, E. and Taylor, W. R. (1967). J. Phycol. 3, 77-81. Szalontai, B. and Csatorday, K. (1980). J. Mol. Struct. 60, 269-272. Szalontai, B. and Van de Ven, M. (1981). FEBS Lett. 131, 155-157. Szalontai, B., Bagyinka, Cs. and Horvath, L. I. (1977). Biochem. Biophys. Res. Commun. 76, 660-6155 Talling, J. F. (1957). New Phyrol. 56, 29-50. Talling, J. F. (1971). Mitt. Internat. Verein. Limnol. 19, 2142. Talling, J. F., Wood, R. B., Prosser, M. V. and Baxter, R. M. (1973). Freshwater Biol. 3, 53-76. Tamura, N., Yamamoto, Y. and Nishimura, M. (1980). Biochim. Biophys. Acfa 592, 536-545. Tamura. N., Itoh, S. and Nishimura, M. (1981). Plant Cell Physiol. 22, 603-612. Tanada, T. (1951). Am. J. Bot. 38, 276-283. Tandeau de Marsac, N. (1977). J. Bacteriol. 130, 82-91. Tandeau de Marsac, N. and Cohen-Bazire, G. (1 977). Proc. Natl Acad. Sci. US.A . 74, 1635- 1639. Taylor, F. J. R. (1978). Biosystems 10, 67-89. Teale, F. W. J. and Dale, R. E. (1970). Biochem. J . 116, 161-169. Theodor, R., Zinsmeister, H., Mues, R. and Markham, K. R. (1980). Phytochemistry 19, 1695-1700. Thielen, A. P. G. M. and van Gorkom, H. J. (1981a). FEBS Lett. 129, 205-209. Thielen, A. P. G. M. and van Gorkom, H. J. (1981b). Biochim. Biophys. Acta 635, 1 1 1-120. Thielen, A. P. G. M., van Gorkom, H. J . and Rijgersberg, C. P. (1981). Biochim. Biophys. Acta 635, 121-131. Thinh, L.-V. (1978). Aust. J . Bot. 26, 617-620. Thomas, J . C. and Mousseau, A. (1981). I n “Photosynthesis 11” (G. Akoyunoglou, Ed.), pp. 435444. Balaban Intern. Sci. Serv., Philadelphia, Pennsylvania, USA. Thornber, J. P. (1969). Biochim. Biophys. Acta 172, 236241. Thornber, J. P. (1975). Annu. Rev. Plant Physiol. 26, 423458. Thornber, J. P. and Barber, J. A. (1979). In “Photosynthesis in relation to model systems” (J. Barber, Ed.), pp. 27--70. Elsevier/North Holland, Amsterdam. Thornber, J. P. and Highkin, H. R. (1974). Eur. J . Biochem. 41, 109-116. Thornber, J. P., Alberte, R. S., Hunter, F. A,, Shiozawa, J. A. and Kan, K.-S. (1977). In “Brookhaven Symp. Biol.” 28, 132-148. Thornber, J. P., Trosper, T. L. and Strouse, C. E. (1978). I n “The Photosynthetic Bacteria” (R. K. Clayton and W. R. Sistroni, Eds), pp. 133-160. Plenum Press, New York. Thornber, J. P., Markwell, J. P. and Reinman, S. (1979). Photochem. Photobiol. 29, 1205-121 6. Thorne, S. W. (1981). Biochim. Biophys. Acta 590, 309-323.

216


Thorne, S. W. and Boardman, N. K. (1971). Biochim. Biophys. Acta 234, 113-125. Thorne, S. W., Horvath, G., Kahn, A. and Boardman, N. K. (1975). Proc. Natf.Acad. Sci. U.S.A. 72, 3858-3862. Thorne, S . W., Newcomb, E. H. and Osmond, C. B. (1977). Proc. Natl. Acad. Sci. U.S.A. 74, 575-578. Thorne, S . W., Duniec, J. T. and Lee, J. A. (1983). Photobiochem. Pholobiophys. 5 , 71-78. Thornley, J. H. M. (1976). “Mathematical models in plant physiology”, Academic Press, London. Thrash, R. J., Fang, H. L.-B. and Leroi, G. E. (1979). Photochem. Photobiol. 29, 1049-1050. Timofeeva, V. A. (1974). In “Optical Aspects of Oceanography” (N. G. Jerlov and E. Steeman Nielsen, Eds), pp. 177-219. Tiselius, A. (1930). Inaugural Dissertation. R. Soc. Sci. Upsala, Ser. IV 7 , 1-107. Titlyanov, E. A. and Lee, B. D. (1978). Biol. Morya 4, 36-41. Trebst, A. (1974). Annu. Rev. Plant Physiol. 25, 423-458. Tremolieres, A., Dubacq, J.-P., Ambard-Bretteville, F. and Remy, R. (1981). FEBS Lett. 130, 27-3 I . Troche, R. P., Rice, J. D. and Wells, G. N. (1981). Pfant Physiol. 68, 74-81. Troxler, R. F. (1972). Biochemistry 11, 4235-4242. Troxler, R. F. and Offner, G. D. (1979). Arch. Biochem. Biophys. 195, 53-65. Troxler, R. F., Kelly, P. and Brown, S . B. (1978). Biochem. J. 172, 569-576. Troxler, R. F., Brown, A. S. and Brown, S. B. (1979). J. Biol. Chem. 254,341 1-3418. Troxler, R. F., Ehrhardt, M. M., Brown-Mason, A. S. and Offner, G. D. (1981). J. Biol. Chem. 256, 12176-12184. Tyler, J. E. (1961). Proc. Natl. Acad. Sci. U.S.A. 41, 1726-1733. Tyler, J. E. (1964). Proc. N a d Acad. Sci. U.S.A. 51, 671-678. Tyler, J. E. and Smith, R. C. (1970). “Measurements of spectral irradiance underwater”. Gordon and Breach, New York. Van Baalen, C. (1968). Plant Physiol. 43, 1689-1695. Van Best, J. A. and Duysens, L. N. M.(1977). Biochim. Biophys. Acta 459, 187-206. Van Best, J. A. and Mathis, P. (1978). Biochim. Biophys. Acta 503, 178-188. Van Den Driessche, T. (1966). Exp. Cell. Res. 42, 18-36. Van Den Driessche, T. and Hars, R. (1972). J. Microsc. 15, 85-90. Van de Hulst, H. C. (1957). “Light scattering by small particles”. Wiley, New York. Van Gorkom, H. J. (1974). Biochim. Biophys. Acta 347, 4 3 9 4 2 . Van Ginkel, G. and Kleinen-Hammans, J. W. (1980). Photochem. Photobiof. 31, 385-396. Van Metter, R. L. (1977). Biochim. Biophys. Acta 462, 642-658. Velichko, I. M.(1980). Gidrobiol. Z H 16, 46-51, In Biol. Abstr. 72, 83799. Venediktov, P. S. A., Rubin, A. B., Freidlin, M. I. and Shinkarev, V. P. (1979). Biophysika 24, 1030-1034. Vermaas, W. F. J. and Govindjee (1981). Photochem. Photobiol. 34, 775-793. Vesk, M. and Jeffrey, S. W. (1977). J. Phycol. 13, 280-288. Vierling, E. and Alberte, R. S . (1980). Plant Physiol. 50, 93-98. Vincent, W. F. (1980). Br. Phycol. J. 15, 27-34. Vogelmann, T. C. and Scheibe, J. (1978). Planta 143, 233-239. Vooren, C. M. (1981). Aquatic Botany 10, 143-154. Voskresenskaya, N. P. (1979). In “Encyclopedia of Plant Physiology”, N.S. Vol. 6, “Photosynthesis 11, Photosynthetic carbon metabolism and related processes” (M. Gibbs and E . Latzko, Eds), pp. 174-179. Springer, Berlin. Waalund, J. R.,Waalund, S. D. and Bates, G. (1974). J. Phycol. 10, 193-199.


217

Walker, J . C. G. (1978). Pure Appl. Geophys. 116, 222-231. Walker, J . C. G., Klein, C., Schidlowski, M., Schopf, J. W., Stevendon, D. J. and Walter, M. R. (1982). I n “The Earth’s earliest biosphere: its origin and evolution” (J. W. Schopf, Ed.). Princeton University Press, Princeton, N . J . Wallen, D. G. and Geen, G. H. (1971). Mar. Biol. 10, 3 4 4 3 . Wallen, D. G. and Geen, G. H. (1971). Mar. Biol. 10, 4 4 5 1 . Wallin, R., Selset, R. and Sletten, K. (1978). Biochem. Biophys. Res. Commun. 81, 13 19-1 328. Walsby, A. E. and Booker, M. J. (1980). Br. Phycol. J . 15, 311-319. Walsby, A. E. and Reynolds, C. S. (1980). In “The Physiological Ecology of Phytoplankton” (I. Morris, Ed.), pp. 371412. Blackwell Scientific, Oxford. Walter, M. R., Buick, R. and Dunlop, J. S. R. (1980). Nature 284,4 4 3 4 5 . Wang, R. T. and Myers, J. (1976a). Photochem. Photobiol. 23, 405410. Wang, R. T. and Myers, J . (1976b). Photochem. Photobiol. 23, 411414. Wang, R. T., Stevens, C. L. R. and Myers, J. (1977). Photochem. Photobiol. 25, 103- 108. Wang, R. T., Graham, J.-R. and Myers, J. (1980). Biochim. Biophys. Acta 592, 277-284. Wanner, G. and Kost, H.-P. (1980). Protoplasma 102, 97-109. Wasielewski, M. R., Smith, R. and Kostka, A. G. (1980). J . Am. Chem. SOC.102, 6924-6928. Wasley, J. W. F., Scott, W. T. and Holt, A. S. (1970). Can. J. Biochem. 48, 377-383. Weedon, B. C. L. (1971). In “Carotenoids” (0. Isler, Ed.), pp. 267-324. Birkhauser Verlag, Basel. Wehrmeyer, S. (1970). Arch. Mikrobiol. 71, 367-383. Weinberg, S. (1976). Mar. Biol. 37, 291-304. Weinberg, S. and Cortel-Breeman, A. (1978). Bijdr. Dierk. 48, 35-44. Weiss, A. (1981). Angew. Chem. Int. Ed. Engl. 20, 856860. Weiss, Jr, C. (1972). J. Mol. Spectroscopy 44, 37-80. Weiss, C. (1979). In “The Porphyrins” (D. Dolphin, Ed.), Vol 111, pp. 211-223. Academic Press, New York. Wellburn, A. R. (1976). Biochem. Physiol. Pfanzen. 169, 265-271. Wellburn, F. A. M., Wellburn, A. R. and Senger, H. (1980). Protoplasma 103, 35-54. Wessels, J . S. C. and Borchert, M. T. (1978). Biochim. Biophys. Acta 503, 78-93. Wessels, J . S. C. and Spijkerboer, F. W. J. M. (1981). Biochim. Biophys. Acta 638, 94-99. Wessels, J. S. C., Waveren, V. A,-V. and Voorn, G. (1973). Biochim. Biophys. Acra 292, 741-752. Westlake, D. F. (1965). I n “Light as an Ecological Factor” (R. Bainbridge, G. C. Evand and 0. Rackham, E d ) , pp. 99-1 19. Blackwell, Oxford. Whatley, J . M. (1971). New Phytol. 70. 725-742. Whatley, J. M. (1977). New Phytol. 79, 309-313. Whatley, J. M. (1981). Ann. N . Y . Acad. Sci. 361, 154-165. Whatley, J . M. and Whatley, F. R. (1981). New Phytol. 87, 233-247. Whatley, J. M., John, P. and Whatley, F. R. (1979). Proc. R . SOC.Lond. B. 204, 165-187. Wheeler, W. N. (1980). Mar. Biol. 56, 97-102. Whittaker, R. H. and Margulis, L. (1978). Biosystems 10, 3-18. Wild, A. (1979). Ber. Deutsch. BOI.Ges. 92, 341-364. Wild, A. and Urschel, B. (1980). Z. Naturforsch 35, 627-637. Wild, A., Stuehn, N. and Ruehle, W. (1981). Phofosynth. Res. 2, 105-114. Wildman, R. B. and Bowen, C. C. (1974). J. Bacteriol. 117, 866-881.

218


Wildner, G. F. and Hauska, G. (1974). Arch. Biochem. Eiophys. 164, 127-135. Williams, W. P. (1977). In “Primary Processes of Photosynthesis” (J. Barber, Ed.), pp. 101-147. Elsevier, Amsterdam. Williams, W. P. and Glazer, A. N. (1978). J. Eiol. Chem. 253, 202-21 1. Williams, W. P., Furtado, D., Nutbeam, A. R. (1980). Photobiochem. Photobiophys. 1, 9 1-1 02. Willstatter, R. and Page, H. P. (1914). Ann. 404,237-271. Withers, N. W., Alberte, R. S., Lewis, R. A., Thornber, J. P., Britton, G. and Goodwin, T. W. (1978). Proc. Natl. Acad. Sci. U.S.A. 75, 2301-2305. Witt, K. (1973). FEES Lett. 118, 279-282. Wolk, C. P. (1980). I n “The Biochemistry of Plants”, Vol. 1,654-686. Academic Press, New York. Wollman, F.-A. (1979). Plant Physiol. 63, 375-381. Wollman, F.-A,, Olive, J., Bennoun, P. and Recouvreur, M. (1980). J . Cell. Eiol. 87, 728-735. Wong, H., Pellegrino, P., Alfano, R. R. and Zilinskas, B. A. (1981). Photochem. Photobiol. 33, 651-662. Wood, A. M. (1979). J. Phycol. 15, 330-332. Wood, P. M. (1978). Eur. J. Eiochem. 87, 9-19. Wood, P. M. and Bendall, D. S. (1975). Eiochim. Eiophys. Acta 387, 115-128. Wood, N. B. and Haselkorn, R. (1980). J. Eacteriol. 141, 1375-1385. Woodward, R. B. (1961). Pure Appl. Chem. 2, 3 8 3 4 4 . Xavier, A. V., Czerwinski, E. W., Bethge, P. H. and Mathews Scott, F. (1978). Nature 275, 245-246. Yamanaka, G. and Glazer, A. N. (1980). Arch. Microbiol. 124, 39-47. Yamanaka, G., Glazer, A. N. and Williams, R. C. (1978). J. Eiol. Chem. 253, 8303-83 10. Yamaoka, T., Satoh, K. and Katoh, S. (1978). In “Photosynthetic Oxygen Evolution” (H. Metzner, Ed.), pp. 104-115. Academic Press, New York. Yentsch, C. S. (1962). Limnol. Oceanogr. 7, 202-217. Yentsch, C. S. (1980). In “The Physiological Ecology of Phytoplankton” (I. Morris, Ed.). Blackwells Scientific, Oxford. Yocum, C. S. and Blinks, L. R. (1954). J. Gen. Physiol. 38, 1-16. Yocum, C. S. and Blinks, L. R. (1958). J. Gen. Physiol. 41, 1113-1118. Yokohoma, Y. (1981). Eoranica Marina 23, 637-640. Yokohama, Y., Kageyama, A., Ikana, T. and Shimura, S. (1977). Eotanica Marina 20, 433436. Yoshizaki, F., Sugimura, Y. and Shimokoriyama, M. (1981). J. Eiochem. 89, 1533-1540. Yu, M.-H., Glazer, A. N., Spencer, K. G. and West, J. A. (1981). Plant Physiol. 68, 482-488. Yuen, M. J., Shipman, L. L., Katz, J. J. and Hindman, J. C. (1981). Photochem. Photobiol. 32, 28 1-296. Zanefeld, J. R. (1974). In “Optical Aspects of Oceanography” (N. G. Jerlov and E. Steeman Nielsen, E h ) , pp. 121-134. Academic Press, London. Zanefeld, J. R. V. (1975). Section 2.4 of Climatic Impact Assessment Program, Monograph V (Part 1) United States Department of Transportation, Report No. DOT-TST-75-55, pp. 2-108-2-1 57. Zickendraht-Wendelstadt, B., Friedrich, J. and Rudiger, W. (1980). Photochem. Photobiol. 31, 367-376. Zilinskas, B. A. and Glick, R. E. (1981). Plant Physiol. 68, 447-452.


219

Zilinskas, B., Zimmennan, B. K. and Gantt, E. (1978). Photochem. Photobiol. 27, 587-595. Zuber, H . (1978). Ber. Deutsch Bot. Ges. 91, 459-475.


Effects of Nutrient Stress on Susceptibility of Plants to Disease with Particular Reference to the Trace Elements

ROBIN D . GRAHAM Department of Agronomy. Waite Agricultural Research Institute. The University of Adelaide. Glen Osmond. South Australia 5064

I . Introduction .

. . . . . I1. Adaptation to Nutrient Stress . I11. Macronutrients . . . . . A. The General Pattern . B. C. D. E. F.

. . . . . . . . . . . . . . . . Phosphorus . . . . . . . . Potassium . . . . . . . . Nitrogen . . . . . . . . .

. . . . . . . 222 . . . . . . . 222

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

. . Sulphur, Magnesium and Calcium . . Summary of Macronutrient Effects . .

IV . Micronutrients . . . . . . . . A. Copper . . . . . . . . B. Boron . . . . . . . . C. Manganese . . . . . . . D. Iron . . . . . . . . . E. Zinc . . . . . . . . . F. Nickel . . . . . . . . G. Silicon . . . . . . . . H. Other Elements . . . . . . I. Summary of Micronutrient Effects J. Guidelines for Experimentation .

. . . .

223 223 224 . . . . . 225 . . . . . 226 . . . . . 228 . . . . . 228

. . . . . . . . 229 .

.

.

.

.

.

.

. 229

. . . . . . . . 238

. . . . . . . . 243 . . . . . . . . 254 . . . . . . . . 257

. . . . . . . . 260

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

261 263 . 265 . 268

. . . . . . . Epilogue . . . . . . . . . . . . . . . . . . Acknowledgements . . . . . . . . . . . . . . . References

269 269

. . . . . . . . . . . . . . . . . 270

222

R. D. GRAHAM

I. INTRODUCTION The effect of mineral nutrition on plant diseases has received considerable attention over the years,* but little of this attention has been directed towards the trace elements.? The purpose of this review is to consider the biochemistry, physiology and agronomy of these elements in plants in so far as they may influence host-pathogen relationships. Each of the micronutrients can be identified with specific biochemical pathways, so their effects on disease offer avenues for elucidating mechanisms of resistance in higher plants. Certain general principles may be demonstrated, but the complex nature of many interacting factors influencing plant health limits the extent to which simple patterns emerge. The quest for general principles is stimulated by the increasing prevalence of micronutrient deficiencies in modern agricultural systems, in which the use of the appropriate micronutrients may be seen as a form of biological control. 11. ADAPTATION TO NUTRIENT STRESS

Plants under nitrogen deficiency direct relatively more of their resources into long, thin exploratory roots (Hackett, 1972; Lungley, 19741, which will have survival value if these roots intercept more available nitrogen. This is an example of a homeostatic response which tends to lessen the impact of the nutrient stress. Both wild and domesticated plants have evolved such “active” response mechanisms to overcome or minimize the major edaphic limitations for growth and reproduction. Among these limitations are the deficiencies of nitrogen, phosphorus and potassium. In particular, most plants appear to have evolved mechanisms for the re-mobilization and translocation of these major nutrients from old leaves to the meristems under deficiency, the root meristems representing the source of more of the limiting element and the above-ground meristems eventually becoming the reproductive axis in most cases. Thus, meristems are the means of survival and are preferentially supplied with the limiting nutrient. In contrast to this, many plants, particularly our crop plants, do not seem to have evolved mechanisms for retranslocating the micronutrients (and calcium) from old leaves to the growing points under deficiency conditions. Deficiency symptoms of these elements, for example, iron, boron, manganese, zinc, copper, molybdenum (as well as calcium) appear generally in the young tissues. From this we can conclude that micronutrient deficiencies were not

* For this, the reader is referred to recent general reviews by Black (1968), Borys (1968), Goss (1968), Huber (1980, 1981), Huber and Watson (1974), Jenkyn and Bainbridge (1978) Krauss (1 969), Sadasivan ( 19654, Trolldenier ( I 969), von Uexkuell (1 966). t For the present purpose, the term “trace elements” refers to the (essential) micronutrients plus some others such as Li, Cr, Ni, Pb, F, Si, Cd, Al.

NUTRIENT STRESS AND PLANT DISEASE

223

frequently a significant limitation to plant growth and survival before the agrarian revolution, and indeed probably not in agricultural systems until the advent of mineral NPK fertilizers, plant breeding and quite modern levels of crop productivity. Nitrogen and phosphorus were probably the common limiting nutrients. Since higher plants and their pathogens have co-existed over evolutionary periods of time, most relationships can be said to have achieved some form of delicate balance, neither the host nor the parasite having eliminated the other. The exact point of balance between host and pathogen in a given situation depends on the environment of which nutrition is part. The following discussion assumes that perturbations in host physiology caused by changes in plane of nutrition can shift the balance considerably without affecting all resistance/susceptibility mechanisms known to exist in each case. Small changes in the host may have considerable effects on the severity of disease. The resistance conferred on a host plant by “proper” nutrition is most likely a form of general or “horizontal” resistance quite distinct from race-specific (gene-for-gene) type mechanisms. It is well documented (see next section) that high nitrogen supply may predispose plants both to disease and also, in certain marginal soils, to micronutrient deficiencies not apparent at low nitrogen supply. Micronutrient deficiencies themselves also predispose plants to disease as we shall see later, so that the increased use of nitrogen fertilizers over the last century or so may have shifted the host-pathogen balance significantly from that of either the historical or the evolutionary past. This is the context in which physiology and pathology are viewed in this paper. Nutritionally deficient plants generally have less chance of survival from an attack by a pathogen. This is a general proposition. There is, however, a case for going further. Individual elements differ intrinsically in their effects on the host-pathogen relationships. The differences are rooted in the biochemical roles of the essential (and non-essential) elements, and the differential mineral requirements of host and parasite. 111. MACRONUTRIENTS A. THE GENERAL PATTERN

Before discussing in detail the trace elements, I should like first to describe the general pattern of nutritional effects on disease. This is illustrated by the data of Last (1962) from a NPK fertilizer study of barley infected naturally in the field with powdery mildew (Erysiphe gruminis DC var. hordii). Table I shows the main effects on yield and disease and Fig. 1 the individual elemental effects. This figure depicts four variables, one dependent variable (the disease index, being a percentage of leaf area affected) and three independent nutrient

224

R. D. GRAHAM

variables in two dimensions; yet it is quite clear that the addition of nitrogen, shown as double lines, to any other combination markedly increased the disease index. Likewise phosphorus, in every combination decreased, and potassium had little effect on, severity of disease (see also Table I).

"r

N

I

I

0

I

I

I

2 Number of nutrients applied

3

Fig. 1. Effects of the factorial combinations of nitrogen, phosphorus and potassium on the mean level of powdery mildew infection of barley from May 25 to July 14,1960 (after Last, 1962).

TABLE I Main Effects of N , P and K on Yield and Incidence of Barley Powdery Mildew (after Last, 1962)

P

N

+ Yield of grain (15% moisture) (t ha-') 3.4 Mean level of infection (%) 19.5

K

-

+

-

+

2.3 10.2

3.3 11.7

2.4 18.1

3.0 13.3

-

S.E.

2.8 k0.18 16.4 k0.87

B. PHOSPHORUS

The phosphorus response here seems to be typical of many reports and may be thought of as a form of growth dilution of the number of infections as the plants respond to phosphorus over the range from deficiency to adequacy. The increased vigour of P-adequate plants enables them to outgrow the disease; phosphorus does not confer resistance in supraoptimal amounts. Beneficial effects of phosphorus also occur with soil-borne diseases, e.g. Pythium root rot of wheat (Vanterpool, 1935) and take-all, Gaeumannomyces

NUTRIENT STRESS A N D PLANT DISEASE

225

graminis, of wheat (Huber, 1981). There are numerous reports of a similar nature with many species of both host and pathogen. Graham et al. (1981) proposed that P-deficient membranes leak metabolites to invading fungi. C. POTASSIUM

Although I have described Fig. 1 as a general pattern, we can generalize only so far. Potassium was barely deficient, giving a mean yield increment of only 1/5 that of nitrogen or phosphorus (Table I). However, in situations of moderate to severe deficiency, potassium additions have dramatic effects in decreasing disease, including powdery mildew of barley (Glynne, 1959). Because of the repeated removal of clippings, turf grasses place great demands on soil potassium reserves and are frequently responsive to applications of this element; applications of potassium have been reported to achieve outstanding decreases in disease ratings, for example, Table 11. Effects of potassium on disease are most consistently reported in relation to fungal diseases especially the wilts and root diseases such as Fusarium, Verticillium, Pythium, Phytophthora and Rhizoctonia. In these cases (Table III), the yield response to potassium appears to be due to compounding effects TABLE I1 Efects o f N , P , K and Lime" on Leaf-spot Disease oj' Bermuda Grass (Cynadon sp.) (.from Evans et al., 1964) Fertilizer

Spotsbper leaf (av.)

13.5 19.9 23.1 16.0 19.6 147.5

NPKL (complete) N, PKL NP,,2 KL NKL NPK NPL (minus K)

" N . 178 kg h a - ' ; P, 39 kg ha-'; K. 74 kg ha-'; lime to pH 6.5. Caused by two undisclosed fungi.

TABLE 111 Yield of Cotton and Percentage of Wilted Plants in Response to Applications of Potassium Fertilizer (Miles, 1936) Fertilizer (kg ha -' K)

Yield of bolls (t ha -')

0 22 45

0.56 0.98 1.01

Proportion of wilted plants

(%I 32 10 8

226

R. D. GRAHAM

of concurrent increases in growth of the host and decreases in the level of infection. Potassium fertilization also has decreased numerous foliar diseases, both fungal and bacterial, on many species, rice being a well documented example (von Uexkuell, 1966). The physiology of potassium deficient plants is that of a plant completely disordered and it is difficult to point to specific mechanisms of resistance broken down by the deficiency. The loss of cell turgor (Graham and Ulrich, 1972) may be a physical factor facilitating penetration, both by fungi and insects. Deficient plants accumulate soluble organic acids, amino acids and amines. Glutamine for example is particularly high in potassium deficient plants (Thompson et al., 1960) and has been shown to stimulate germination of spores of at least one fungal pathogen, Piricularia oryzae on rice leaves (Suryanarayanan, 1958). D. NITROGEN

Nitrogen markedly increased the disease index in Last’s study but yield was also increased by about one tonne per hectare (Table I). There is no question of growth dilution here; the susceptibilityof the plants was much enhanced by the nitrogen fertilizer, but the true yield loss from the disease could only be measured against a hypothetical “N + effective fungicide” treatment. It is important to distinguish between the effects of nitrogen on host vigour and on susceptibility to the pathogen, the increase in yield being the net effect. Tolerance is the ability of the plant to grow and produce a yield in spite of the disease burden. In contrast, resistance describes the relative ability of the host to prevent the growth of the pathogen and involves the physiology of both. Both resistance and tolerance may be affected by the plane of nutrition. Few workers have factorially combined nutritional and fungicide treatments in order to measure separately the contributions to yield of the triple effects of nutrition on host plant and pathogen vigour and on host plant resistance. For example, McNew (1953) used virus-free wheat seed to make this distinction and Jenkyn and Moffatt (1975) and Graham (1980) used ethirimol and benomyl respectively to control powdery mildew for this purpose. Below ground, where it is more difficult to make and interpret disease indices, it is likely that there are similar responses. Nitrogen increases the vigour of the plant and the number of roots produced; thus even if there be numerically more infection points, there are probably also more healthy roots to support the crop and ultimately a higher yield. This was the situation observed by Garrett (1948) where nitrogen supplied to a take-all infested field of wheat resulted in greater tolerance of the disease. Another effect of nitrogen appears to be its stimulation of the soil microflora to the extent that competition with the pathogen for nutrients decreases the number of propagules surviving in the soil, resulting in actual


227

decreases in the disease index. This appears to operate on Rhizoctonia solani of beans and buckwheat (Davey and Papavizas, 1960) and has been considered also to be the basis of beneficial effects of nitrogen at sowing on take-all infected wheat (Rovira and Wildermuth, 1981). The effect of nitrogen has been generalized in Fig. 2. Inorganic nitrogen is seen as placing a high demand on carbon from photosynthesis via the Kreb’s cycle, leaving little for secondary metabolism via the pentose shunt and shikimate pathway. Under nitrogen limiting conditions, the latter is enhanced providing for a large pool of phenolics and alkaloids. These are the basis for many of the defence mechanisms of plants: physical (preformed barriers), chemical (oxidized and translocated to the infection) and phytoalexin (induced/synthesized in response to the presence of an invading organism) (see Horsfall and Cowling, 1980; Bell, 1981). Whereas supraoptimal applications of phosphorus or potassium are commonly without effect on disease, excess nitrogen may encourage fungal pathogens (Trolldenier, 1969),especially if phosphorus or potassium is low, as in Fig. 1 . Mengel and Kirkby (1978) quote several examples of this commonly observed response. On the other hand, Pseudomonas syringae (bacterial) and

1Q I Inorganic

Inorganic nitrogen

Sugar

Fig. 2. Carbohydrate and energy economy at (a) high nitrogen supply and (b) low nitrogen supply (after Borys, 1968).

228

R. D. GRAHAM

Helminthosporiun turcicum (fungal) on maize were suppressed by high rates of nitrogen fertilizer. It seems highly likely that interactions of nitrogen with other nutrient elements, clearly established in the plant nutrition and agronomic literature generally, are important to the interpretation of nitrogen effects. An example of the nitrogen and copper interaction effect is discussed later. E. SULPHUR, MAGNESIUM AND CALCIUM

Sulphur and magnesium do not figure strongly in the literature on nutrition and disease (see general reviews) except where sulphur is used directly as a foliar dust or to lower the pH of the soil, an effect we shall return to. We can assume that these elements confer the general tolerance associated with vigour where the soils are deficient in either element. There are, however, quite a number of reports of effects on disease due to calcium, some inhibitory to disease (for example, Davidson and GOSS,1972) and some favourable (Williams, 1961). Calcium has important roles in the integrity of all membranes and cell walls (Epstein, 1972) and these have been invoked as mechanisms for the greater resistance against Pythium, Sclerotium, Botrytis and Fusarium conferred by calcium in a number of cases. The dual effects of lime in providing the nutrient calcium and in increasing pH have not always been separated, still less the multiple effects associated with changing the soil pH. Gill (1972) using a soil of pH 4.3 found that gypsum (CaSO,) effectively decreased Pythium whereas dolomite (CaCO, + MgCO,) or gypsum+dolomite did not, although soil pH was raised by one or two units. Instinctively, we would predict that raising the pH from 4.3 would favour the host since fungi are generally more acid tolerant than are higher plants, but this was not so. There are many similar cases most noteably, take-all (Gaeumannomyces graminis) of cereals and grasses and common scab (Streptomyces scabies) of potato. Particularly with scab, one of the best methods of control in commercial production is to keep the pH low. Since the optimal pH for the host is higher in the absence than in the presence of disease, our attention is directed to the known effects of raising the pH in decreasing the availability of most of the trace elements. F. SUMMARY OF MACRONUTRIENT EFFECTS

We may summarize the principles which emerge from a study of macronutrient effects on incidence of disease in higher plants:

(i) A well nourished host is generally more tolerant of disease. (ii) Some elements appear to affect disease severity simply through greater tolerance, for example, phosphorus and sulphur; while others including


229

nitrogen and potassium clearly alter specific host-plant resistance mechanisms. Nutrition is only one of many factors influencing resistance to disease. Macronutrients increase resistance to disease, if at all, only in the deficiency range; supraoptimal amounts of nutrient do not provide further protection, and may be detrimental in the case of nitrogen. Yield responses to fertilizers often contain two components, due to overcoming both the deficiency per se, and changing the disease burden. Nitrogen may either increase or decrease disease, depending on other interacting factors, but in either case will increase yield where the soil is deficient in the nutrient. Nitrogen effects are associated with effects on the balance between the primary and secondary metabolic pathways in the host.

IV. MICRONUTRIENTS Nitrogen, phosphorus and potassium are involved in so many fundamental plant processes that in deficiency the whole metabolism of the plant is greatly disturbed. With micronutrient deficiencies, it is often possible to point to more specific areas of metabolism to explain the observed effects on host-parasite relationships. Copper, boron and manganese are discussed seriatum as they each interact with nitrogen, are all intimately involved in the synthesis of phenols in plants, and all have major effects on host susceptibility to disease. A. COPPER

Control of fungal diseases with copper has been documented over more than a century of use of copper oxychloride (for example, Lodeman, 1902) and various Bordeaux mixtures. During the first 50 years of use of Bordeaux mixtures, it was gradually realized that copper had beneficial effects on the crop, in some cases in the absence of any obvious pathogen (Felix, 1927).This led to the discovery ofcopper as an essential nutrient for higher plants in 1931 (Sommer, 1931; Lipman and Mackinney, 1931) and for microorganisms as well (Bortels, 1927). Thus in soils of low copper status, Bordeaux mixtures had multiple effects on host plant vigour and yield: (i) on yield from physiological effects of absorbed copper in the leaves, (ii) on disease resistance, and (iii) on the leaf surface where inoculum potential was lowered by the copper precipitates. In the 50 years since then it has become clear that copper deficient plants are

230

R. D. GRAHAM

frequently more susceptible to air-borne fungal diseases than plants with adequate copper. This is the case even where the disease is caused by a foliar pathogen and copper is supplied to the high-copper plants through their roots (Olsen, 1939; Schutte, 1967; Graham, 1980; Bussler, 1981, personal communication). Concentrations of copper in the leaves may be as low as 1-10 ppm Cu on a dry weight basis and cannot be considered to be directly toxic to the pathogen. The response is not a simple one of diluting out the inoculum with increased growth of the copper-supplied plants (Fig. 3); there is a clear suppression both of the number of infection sites and of lesion expansion. A feature of the data in Fig. 3 is the decline in the whole-plant disease index from 7 weeks onwards in the copper sufficient plants. This is known among plant pathologists, particularly in relation to powdery mildew of cereals, as adult plant resistance (APR) (Jones and Hayes, 1971; Jenkyn and Bainbridge, 1978). Each succeeding leaf is more resistant than the one before. The mechanism of APR remains far from clear, but one interpretation of Fig. 3 is that copper deficiency has inhibited the development of APR which may therefore have a copper-requiring step. 50

-

40.

4 .-

30-

;2 0 -

VI .-

c l

10-

0-

P

....... 0 mg Cu per pot

30.

E

.s 20.

r

4

lo-

5 o. c

3

5

7

3

5

7

9

II

13

15

17

19

21

9

II

13

15

17

19

21

10.

c

t

5 .

Time (weeks after emergence)

Fig. 3. The dependence on age of wheat plants and level of copper supply of (a) mean disease index per leaf of powdery mildew (Erysiphe graminis); (b) number of green leaves per pot (three plants); (c) number of tillers per pot. Means of four replicates (Graham, 1980).

23 1


A number of other diseases have been reported to be more severe on copper deficient plants, including such different types of organisms and hosts as Alternuria on sunflowers (W. Bussler, personal communication), Gaeumannomyces graminis on wheat (Wood, 1981), ergot of rye and barley (Tainio, 1961; Simojoki, 1969), Heterodera of sugarbeet (Nesterov and Korol'chuk, 1980), Pucciniu triticina of wheat (Schutte, 1964), Piricularia oryzae of rice (Primavesi and Primavesi, 1970), and Septoria of wheat (Toms, 1958). In addition there are numerous reports of beneficial effects of fertilizer copper in reducing disease under conditions which may not have been copper deficient, for example Gaeumannomyces on wheat (Reis et al., 1982) and Sclerotinia on peanuts (Hallock and Porter, 1981). In these cases the copper at high rates was applied to roots and shoots respectively being the tissues under attack, and was not effective or much less so when applied to the unattacked tissues. A fungistatic effect seems possible in these cases. The apparently simple effect of copper on powdery mildew described by Fig. 3 is shown to be quite complex by a copper and nitrogen interaction study (Fig. 4, Fig. 5(a)). Nitrogen deficient plants were most resistant to powdery mildew regardless of the copper supply. However, at moderate levels of nitrogen supply corresponding to many field situations, copper deficient plants were more susceptible with up to four times as many pustules at midtillering (leaf numbers were not significantly affected by copper supply). At high rates of nitrogen supply (320 and 640 mg/pot) the effect of copper Supply was markedly to decrease resistance. Finally, the highest rate of nitrogen suppressed mildew somewhat in each of two different seasons, regardless of 6 -

5 4 -

3 2 -

1 0 -

40

80 Nitrogen

160 supply

320

640

1280

( mg Nlpot )

Fig. 4. The interaction of nitrogen and copper supply on the severity of powdery mildew on wheat plants in the glasshouse (from unpublished work of R. D. Graham, J . S. Ascher and R. M. Mills).

Fig. 5 . (a) Powdery mildew development on the second youngest leaf with ligule of wheat plants grown at six levels of nitrogen supply (increasing to the right) and at low (left of each pair) and high copper supply (right) in the glasshouse. The senescence of one leaf at high nitrogen supply was consistent across replicates. This photograph corresponds to the data of Fig. 4. (b) Seminal root systems from 60 day old wheat plants grown in calcareous manganese deficient sand inoculated with the take-all fungus. Left: no manganese; right: l00mg manganese per pot. Arrows show the infection site of the main take-all lesions. Only one spot-lesion was found on the entire high manganese root system whereas each root of the unsupplied plant had a debilitating lesion, the root to the right having broken at the lesion during washing out. Some lateral root development can be seen above the lesions (which block phloem and xylem transport) but the combined effects of deficiency and disease were synerigistic in their effects on the host.


233

copper status, an effect associated with very high nitrate concentrations in the leaves, which may be inhibitory to the fungus. This could be an important effect, if confirmed, since nitrate is frequently very high in seedlings (for example, > 10 000 ppm NO,-N, Papastylianou and Puckridge, 1981). Nitrogen supply greatly affects the metabolism of copper in plants owing to the binding of copper by proteins, porphyrins and other organic nitrogen compounds (Loneragan, 1981; Graham and Nambiar. 1981). In the experiment of Fig. 4, the severity of the wither-tip symptom of copper deficiency increased with the rate of nitrogen supply and was worst in the high nitrogen treatments where mildew was less than in the corresponding high-copper plants. It is possible that copper in low-copper high-nitrogen plants is so tightly bound to organic molecules as to be inadequate for the nutrition of the fungus. However, in Table IV, an alternative explanation is suggested. The data in Table IV came from a preliminary experiment to that of Fig. 4, with fewer nitrogen treatments. The results show that the pattern of mildew severity is not correlated with the tissue copper, nitrate or total nitrogen concentrations or to the C u : N ratio. There is, however, an inverse relationship between number of pustules and the amount of lignin in the leaf. A number of other phenolic fractions, and catalase and peroxidase activities, were also determined but did not correlate with the disease index. Lignin is recognized as a physical barrier against a wide range of invading organisms (Beckman, 1980) and a partial defence against powdery mildew (Friend, 1977); it is possible that differences in the lignin content of the plants could be part of the explanation of the differences shown in Fig. 4. The reason for this is developed below. Figure 6 shows the various fates of carbon with emphasis on the shikimate pathway which leads to the synthesis of a suite of phenolic compounds, including ultimately the polyphenolic secondary wall material, lignin. Copper enzymes play an essential role in this synthesis as shown in Fig. 7, PPO (polyphenol oxidase or phenolase) being a copper-requiring enzyme. Lignin and other phenols may constitute a very large part of the leaf biomass as illustrated in Table V. Poor lignification in high nitrogen plants may be explained, as in Fig. 2, by the utilization of more photosynthate by excess nitrogen for the synthesis of amino acids and proteins and less for the synthesis of phenols and lignin via the shikimate pathway. It is also possible, especially under low to moderate levels of copper supply, that the excess nitrogen was binding copper to decrease its mobility and availability (Loneragan, 1981) in the plant to such an extent that its catalytic role in the synthesis of phenols was restricted. In copper deficient plants, lignification is poor (Rahimi and Bussler, 1973; Bussler, 1981), such copper as is available apparently being directed to more critical roles such as electron transport (cytochrome oxidase and plastocyanin both being copper proteins). Lignin and other phenols have been widely

TABLE IV Effect of copper andnitrogen supply on the severity of mildew and the composition of the leaves of wheat in the glasshouse (from unpublkhed work of R. D. Graham, A . C . Jennings and J. S . Ascher) Nutrient supply copper nitrogen

0 0 0

+ + +

L M VH L M VH

Average No. pustules/leaf

Lignin conc. % of DW

Copper conc. in leaves PPm (DW)

Nitrogen conc. in leaves % of DW

4 22 6

26 14 21 33 15 27

1.6 2 -5 2.5 3.2 3.0 8.2

2:o 4.0

2 30 9

Means of three replicates. LM;M:250,VH:IZSOmg N supplied per pot. ND. not detectable. DW. dry weight.

6.5

1.7 42 5.9

Cu/N ratio ( x 104)

Nitrate-N conc. in leaves PPm (DW)

0.79 0 94 0.38 1-92 0.71 1.38

ND 1800 4060 ND 1130 2830

Product

Function

CQZ

t

plysaccharides

Photosynthesis

J-

Hexase - Phosphate Regulation Phosphaenol -Shikimic pyruvic ocid acid

Attraction Coumorins Flovonoids Tannins

Allelopathy Predator protection

+

Environmental resietance

Acetyl coenzyme

I

1

P44dojic

Ter penes

4

Steroid;

. -. ty acids

-$!;

Tie

Pigments

Amino ~

Pro,eins

synthesis

i

Alkaloids

Fig. 6. Selected metabolic pathways of carbon. The shikimate pathway has been expanded to illustrate end-products with diverse functions including defence (after Mooney, 1972). Reproduced with permission, from the Annual Review of Ecology and Systematics, Volume 3. 01972 by Annual Reviews Inc.

TABLE V Ranges in Composition of Loblolly Pine Needles and Axes over the Growing Season (derived from data of Chung and Barnes, 1977) Chemical group Carbohydrates Phenolics Lignin 23-25 Other phenols 15-20 Protein Lipids Soluble acids and minerals

Total biomass (%) 3658 3845 4-5 4-7 4-1

236

R. D. GRAHAM

implicated in resistance of plants to disease (Horsfall and Cowling, 1980) and a low rate of synthesis of phenols, particularly lignin, is proposed as the basis of the greater susceptibility of copper deficient wheat to Erysiphe graminis at moderate nitrogen levels. Bussler (1981) has described how lignification can be turned on and off quite dramatically in tomatoes in solution culture by supplying or withdrawing supply of copper. Such a technique could be used to study the relative importance of lignin, compared to more labile quinones, phytoalexins and non-phenolic inhibitors, to the resistance of plants to invasion. We have embodied in Fig. 2 and Fig. 7 an hypothesis for the interaction of nitrogen and copper on disease resistance: both low nitrogen and high copper favour the synthesis of phenols, which are implicated in a number of ways in resistance to pathogens. Soluble phenols are stored in vacuoles in a reduced state which is non-toxic, but are released in the presence of fungal hyphae and oxidized by the copper enzyme to the more toxic quinones which may kill both the fungus and the nearby host cells, creating a local lesion. As a result of decreased synthesis of the end-product lignin, reduced phenols may build up in copper deficient plants also (Adams er al., 1975; Robson er al., 1981) but they apparently remain ineffective owing to a deficiency of copper-activated polyphenol oxidase (Walker and Webb, 1981). McLaughlin and Shriner (1980) point out “that under the normal (nondisease) circumstances this allocation of carbor. to the production of phenols occurs during periods of the growing cycle when there is not great demand on carbon for structure, storage or reproduction”. In a cereal crop this is perhaps most likely in the later phases of tillering, since the relative growth rate is highest during the seedling stage, placing a large demand on the plant for carbon, and later, reproduction then demands most available carbon. Thus the likely time of phenol synthesis, tillering to stem extension, corresponds to the time of development of APR (Fig. 3), as well as the phase when the most dramatic foliar symptoms of copper deficiency usually occur. APR appears to be an efficient adaptation to defence against powdery mildew as the inoculum is low during the seedling stage (temperatures are also low) and builds up during the season as more infection sites produce spores. Thus, whatever the details of the chemistry of defence are, the demand for carbon and energy is deferred until the inoculum potential is greater and the resources of the plant to supply carbon and energy are also higher. The technique described by Bussler (198 l), mentioned above, may also be suitable for studying the involvement of copper-phenolic systems in the development of APR. An excellent example of beneficial effects from copper fertilizers controlling plant disease in the field is shown in Table VI. In this case, as with the powdery mildew data above, it is certain that the effects are observed over the deficiency range of the element for host plant growth.

L1-, nrci iisinN PER I DE RM FORMATION

nu I NONE INFUSION

.YMERI2AlION IQU NAND r iIiNONI 'InN

cu

L 1 6 NINSUMTING I F I E D RARRIER

Mn'Fe

GROWTH

SHI KINATE PATHWAY

ELEVATED RESPIRATION

TCA ACIDS -CnnPLf

XING

F IXAT ION

B

CELL UALLS PLASTICIZED

PATHWAY

PUMP

T RY PTO-

I

PHANE

IAA

I

t

-

CELL WALL ACID IF IED

QUI NONE

cu

nX IDA1 ION (PPO)

J

RELEASE STORE!, PHENOL I S

POLYmR INFUSION I Z AAND T I ON M Fen ,

CALLOSE

PAPILLAE L IGNITUBERS PROTECTIVE

RECOGNITION ADHESION SURFACE INTERACTION

Fig. 7. A diagrammatic and speculative representation of the sequence of events by which the various structural barriers are formed in response to injury or to the presence of infectiouc agents (after Beckman, 1980, where the reader is referred for details). The approximate locations where copper, boron and manganese participate have been added to this diagram.

R. D. GRAHAM

238

TABLE VI Copper Fertilization Trials with Different Barley Varieties on a Ligno-Carex Peat (after Tainio, 1961) Basal dressing (NPK) Variety Trial 1-1951 Pirkka Tammi Balder Trial 2-1 953 Pirkka Tammi Balder Trial 3-1955 Tammi

Grain yield kg ha - I

Ergot

%

Basal dressing (NPK) + 50 kg ha CuSO, Grain yield Ergot kg ha %

-'

400

8.4

440 760

14.3 7.1

3680 3320 4240

0.2 0.45 0.2

1280 2240 1920

3.8 4.6 3.5

2840 2440 2320

1.0 1.6 3.0

1800

3.0

2670

0

Normally, it is not possible to get fungistatic effects in the leaves from supplying supraoptimal levels of copper to plants via the roots (such as can be achieved by relatively insoluble copper in foliar-applied Bordeaux mixtures); copper absorbed by roots in excess is not normally transported to leaves (unless the concentrations of copper reach the toxic range and the roots are damaged). Because soil-borne diseases are less readily observed, it remains unclear whether there are suppressive effects of high rates of copper fertilizer on root pathogens. Certainly, the tendency of root cortical cell walls to accumulate copper when the latter is in adequate supply (Graham, 1981; Graham and Nambiar, 1981),may cause some inhibition of microbial attack. If so, this mechanism does not operate in copper deficient soils where root copper levels are as low as in the leaves (Chaudhry and Loneragan, 1970). It seems unlikely that copper applied at normal fertilizer rates can be directly toxic to the inoculum in the soil, since most soils react quickly with added copper to maintain its concentration in the solution phase at quite low levels (McBride, 1981). Whether adsorbed copper in cortical cell walls (which may be 50 per cent of the total copper in roots) does inhibit root pathogens may be tested since it is possible to desorb the copper from such sites (Graham, 1981). B. BORON

An hypothesis that plane of nutrition acts as a predisposing factor in disease by depressing phenol synthesis cannot be extended easily to include the effects of boron; boron deficient plants are predisposed to infection yet accumulate large amounts of phenols, and furthermore, appear to have high polyphenol

239


oxidase activity (Dear and Aronoff, 1965; Lee and Aronoff, 1967; Shkolnik, 1974). Fungi, the major pathogens, apparently do not require boron at all (Bowen and Gauch, 1978), nor do they synthesize lignin as a wall material; fungi therefore have a simple advantage over their hosts in boron deficient environments. In spite of these basic differences in metabolism of boron from that of copper, we do find numerous reports, some quite spectacular, of susceptibility of boron deficient plants to disease, and in such deficient conditions we also find the dual benefits of disease tolerance due to enhanced growth and of disease resistance in the host when the limiting element is supplied. Table VII is illustrative of effects of boron on a leached, boron deficient soil, as well as of a positive interaction between copper and boron. Marked increases in yield and decreases in ergot (Claviceps purpurea) accompanied the application of both NPK fertilizers and boron. Addition of the wrong micronutrient, copper instead of boron, reversed completely the beneficial effects of NPK. The explanation for this is not clear but this type of interaction is common among the micronutrients. Table VIII shows marked beneficial effects of B in disease control, and therefore of grain quality, when the yield response was quite small. Other results from Finland show similar effects. This appears to be in contrast to potassium and copper where beneficial effects are not so apparent in cases of marginal deficiency. Boron has been used as a fertilizer for more than 400 TABLE VII Yield of Barley and Incidence of Ergot in Finland (Tainio, 1961) Fertilizer

Yield (kg ha - I )

Ergot (%)

~

nil

NPK NPK + Cu NPK + B NPK +Cu +B

515 900

625 1450 1500

23.2 3 .O 24.1 0.35 0.33

TABLE VIII Yield and Ergot of Barley in Finland (Simojoki, 1969) Fertilizer borate (kg ha -') 0 10

Grain yield (kg ha -') 2800 3100

Ergot (% of grains) 8 0

Boron in plants straw grain (PP~) (PPm) 1.8

5.5

1.o 3.8

240

R. D. GRAHAM

years but it was not until this century that it was shown to be an essential element (Mengel and Kirkby, 1978). It seems possible that its earlier use was associated with disease control. Boron has also been found to confer resistance to powdery mildew in barley (Yarwood, 1938; Eaton, 1930) and wheat (Schutte, 1967) but not oats in one study (Yarwood, 1938). Powdery mildew (Erysiphe cochoracearum DC) of sunflowers in the USA is used as an indicator of boron deficiency (Butler and Jones, 1955) and Fusarium oxysporum var. lini was more severe on boron deficient flax than on boron adequate plants (Keane and Sackston, 1970). Boron deficient flax was also more susceptible to Melampsora (Heggeness, 1942). Boron nutrition has improved the resistance of wheat to rusts (Dennis and O'Brien, 1937; Ismailov, 1954) potatoes to Synchytrium endobioticum (Hampson, 1980), sugarbeet to Sclerotium rolfsii and tomatoes to Fusarium wilt (Edgington and Walker, 1958).Yarwood (1959) pointed out that his 1938 work and that of Heggeness (1942) could not be repeated at will owing to the delicate balances in the chemistry of this element and between host and pathogen. The genotype of both is also important in how boron will affect the disease. Greater susceptibility to disease in boron deficient plants has been reported at least as frequently in monocotyledonous species as in dicots; yet Shkolnik (1974) finds a higher requirement for boron in dicotyledons than in monocots, a difference apparently related to the greater elaboration of secondary metabolites in the former. Boron is present in soils locked up in minerals, bound to organic matter, and free as HsBO3 in the soil solution. Being very weakly dissociated, the uncharged H3B03 is subject to leaching and must be replaced from the soil organic matter. Soils low in organic matter and readily leached, such as acid sands in high rainfall areas, are particularly prone to boron deficiency. Deficiency of this element ought to be considered as a contributing factor in the etiology of disease on such soils, especially as fungal pathogens have the advantage over their hosts of little or no requirement for this element. The wide range of organisms affected whose pathogenicity is enhanced on boron impoverished hosts is impressive, suggesting that general mechanisms of resistance may be involved; however, beneficial effects from applications of boron are not universally recorded even where the element is deficient (Yarwood, 1938; Cherewick, 1944; Williams, 1961; Wood, 1967; Keane and Sackston, 1970; Pobegailo e f al., 1980). It is worth recalling that plant nutritionists generally agree that the concentration range between deficiency and toxicity is particularly narrow for boron (see Mengel and Kirkby, 1978) which is consistent with a regulatory role for boron in plant metabolism. Some of the reports of boron stimulating disease may be due to toxic effects of this element on the host, but this is unlikely to explain all cases. Additionally, there is need to explain the effects summarized in Table VIII where there were


24 1

marked effects on disease when the host, judging from the yield response, was barely deficient in boron, if at all. These variable responses are not surprising when one looks further at the biochemistry of boron deficiency. Boron deficient plants accumulate soluble carbohydrates in actively photosynthesizing leaves (Mengel and Kirkby, 1978) but concentrations are low in growing points. The accumulation of soluble sugars should be attractive to pathogens, especially if there is leakage to the environment as a result of effects of boron deficiency on membrane integrity, as suggested by Pollard et a f . (1977). The defences of boron deficient plants are further weakened at cell wall middle lamellae which are easily cleaved, boron apparently playing a role in stabilization of pectins by C a 2 + (Schmucker, 1934; Spurr, 1957; Mengel and Kirkby, 1978). There seems to be little doubt that the middle lamella is an important target since many pathogens, like Fusarium (Sadasivan, 1965), are equipped with pectinases. The synergistic effects of boron and calcium at the middle lamella must be contrasted with the antagonistic effects of calcium on boron availability in the soil. The antagonism is expressed when calcium is added as lime to neutralize acid soils, the uptake of boron decreasing with increasing pH. For example, lime usually suppresses club root (Pfasmodiophorabrassicae) of cabbage (Williams, 1961; McFarlane, 1958) but not always (Williams, 1961) and in these cases lime may be inducing a deficiency of boron which also appears to promote club root (Utkina et af., 1980; Antonova et af., 1974; Rohde, 1952). Boron seems especially important in hyperplastic diseases. Lee and Aronoff (1967) have shown that boron combines with 6phosphogluconate, the first intermediate in the pentose shunt pathway, to form a 6-phosphogluconate-boratecomplex which cannot be further metabolized, thus inhibiting this pathway of sugar metabolism, and directing carbohydrates more in the “normal” glycolytic pathway. In boron deficiency the pentose shunt pathway operates unregulated resulting in excessive production of phenolic acids characteristic of boron deficient plants (Dear and Aronoff, 1965; Watanabe et a f . , 1964). According to Shkolnik (1974), intense accumulation of low molecular weight phenolics by boron deficient plants occurs from an increased level of a-glucosidase. Polyphenoloxidase is also high in boron deficient plants unlike those deficient in copper, and phenols are assumed to be in their most toxic oxidized form (Shkolnik, 1974; Lee and Aronoff, 1967). Lewis (1980a) has described an involvement of boron in lignification and has also accounted for the greater requirement of this element in dicotyledons through a role in hydroxylation and methylation of ferulic acid to sinapyl subunits which are more frequent in dicotyledonous lignin (Gross, 1980). Lewis has thus outlined one mechanism by which boron may suppress disease on deficient soils. In a subsequent paper Lewis (1980b) outlined a role for boron both in callose synthesis (which may also have a role in defence-see Fig. 7)

242

R. D. GRAHAM

and in suppressing synthesis of phytoalexins, which he suggests may explain the opposite effects of boron on disease (for example, Wood, 1967). In contrast to Lewis’ view, Dutta and McIlrath (1964) reported enhanced synthesis of lignin in boron deficient plants (see Fig. 8). McClure (1979) summarized reports of effects of boron on phenol metabolism and proposed the generalized scheme shown in Fig. 8. Tissue death

B deficiency

-

p

Enhances hexose-Increasedmonophosphateshunt

o

~

~

~

Increased oxidases

phenols

peroxidase

- *1 Increased lignin

B s u f f i c i e n c y k H,BO, complexes-Blocks \ with 6-phosphogluconate H, BOTsequesters soluble phenols

Fig. 8. Suggested

roles for

hexose monophosphateshunt Enhances carbohydrate transport and starch synthesis

boron in controlling

levels of plant phenolics (modified from

McClure, 1979).

Whatever the pathway, Shkolnik (1974) argues that it is the build-up of phenolics, together with excessive production of auxin (also characteristic of boron deficiency) that is toxic to the meristematic tissues of these plants and also explains the blackening of these tissues. Especially high levels of phenolics are observed around necrotic spots in older tissues. It is therefore likely that accumulations of oxidized phenols are toxic to pathogens in some cases so that the pathogenicity of boron deficient plants will depend on the particular host-parasite pair and the normal defence mechanisms which are invoked by the host and are destroyed by the invading organism. Where defence is related to physical barriers in cell walls, the picture is complicated by the variable effect on synthesis of lignin in boron deficiency. Invasion by some pathogens would be promoted in boron deficient plants where weakened middle lamellae and high concentrations of carbohydrates would assist the invader; but enhanced lignification may suppress others. Where defence normally depends upon toxic phenolic compounds, normal host resistance may be maintained or even enhanced. Boron deficiency, like copper deficiency, tends to enhance susceptibility to

~

~

~

~


243

disease but has quite different effects on phenol metabolism from those of copper. It would appear that the roles of boron in defence mechanisms not involving phenolics (for example, in the middle lamella) are of overriding importance when boron is limiting, or that boron plays a vital role in the phenol-based defences themselves which is not at present understood. Rationalization of these variable effects of boron on disease resistance is likely to be found in its interactions with other elements such as nitrogen, copper and manganese. C. MANGANESE

Plants low in manganese have been shown to be more susceptible to a number of pathogens, notably powdery mildews. Powdery mildews were more severe on manganese deficient barley (Vlamis and Yarwood, 1962), wheat (Colquhoun, 1940; Zubko, 1961; Graham, unpublished results), pumpkin seedlings (Abia et al., 1977) and cucumber(Robinson, 1978). Other diseases affected in the same way include Fomes annosus on Norway spruce (Wenzel and Kreutzer, 1971), phylloxera of grapes (Perov et al., 1971), clubroot of cabbage (Antonova et al., 1974), Fusarium udum of pigeon-pea (Sarojini, 1950), Fusarium wilt on cotton (Lakshminarayanan, 1955), and Helminthosporium oryzae (brown spot) of rice (Kaur et al., 1979).Two notable soil-borne diseases, take-all of wheat (Reis et al., 1982; Fig. 11) and common scab of potato (Mortvedt et al., 1961, 1963; McGregor and Wilson, 1964, 1966) have been decreased by applications of manganese and these case histories will be studied in more detail below. It is the experience of our laboratory that manganese deficiency is difficult to induce in soil in the glasshouse even when using soils which are quite deficient in the field, and other laboratories also have this experience though few publish their results. Temperature, moisture content and drying-andwetting effects during soil preparation appear to enhance manganese availability in glasshouse conditions (Leeper, 1970). Moreover, together with many others (for example, Batey, 1971; Samuel and Piper, 1928) we observed that manganese deficiency in the field is often ephemeral, in accordance with seasonal conditions. It is for these reasons, I suspect, that there are not more reports clearly linking manganese nutrition with susceptibility to disease. It has been suggested in Section 1V.A that plants may accumulate high copper concentrations in roots as a means of defence against root diseases, yet maintain relatively low concentrations in shoots. In contrast, manganese is rapidly distributed throughout the plant, and does not accumulate in the roots in this way (Graham, 1979). However, organisms appear to be quite sensitive to Mn2 ions and this sensitivity may be of advantage to the host in that high levels of Mn2 appear to be inhibitory to certain fungal exoenzymes especially pectin methylesterase (Sadasivan, 1965). Iron is antagonistic to manganese in +

+

244

R. D. GRAHAM

this effect, promoting enzyme production so that host resistance is invested, in part, in a tolerance of a higher Mn/Fe ratio than the fungal pectolytic enzymes can tolerate. Manganese also inhibits host enzymes activated by the pathogen, for example, aminopeptidase (Huber and Keeler, 1977). The role of manganese in resistance to two particular diseases, common scab and take-all are discussed in detail in the following sections.

( I ) Common Scab (Streptomyces scabies) of Potato Judging from the voluminous and continuing literature on this disease, it has been and still is a disease of considerable economic importance in many parts of the world. The disease attacks the tubers as they grow, forming dark, unsightly necrotic areas which decrease the amount of usable product. Control of common scab by agronomic means developed over a long period of time. Basically, control is enhanced by low soil pH, irrigation during tuber growth and incorporation of green manure crops (Wenzl, 1975). Soil pH may be lowered by the use of sulphur or ammonium fertilizers in preference to nitrate (Guntz and Coppenet, 1957; Wenzel, 1975). Although these practices are well established, there are exceptions to them all (for example, Wenzl and Reichard, 1974; Weinhold et al., 1964) and no unifying basis for management has been accepted. However, more recently, manganous sulphate has been shown to control scab in a number of situations (Spatz, 1955; Guntz and Coppenet, 1957; Mortvedt et al., 1961, 1963; McGregor and Wilson, 1964, 1966; Barnes, 1972; and Davis et al., 1976). These, and many other similar reports, are possibly derived from a report of Bjorling (1946) of the higher likelihood of scab on manganese deficient soils in Sweden. It is evident from soil chemistry that these factors, low soil pH, ammonium fertilizers, sulphur, high soil moisture and green manures are all factors which tend to increase available manganese in the soil by promoting reduction of insoluble Mn"' and Mn'" oxides to soluble and exchangeable MnZ+ions (Black, 1968; Leeper, 1970); and several workers have proposed that manganese transformations are the key to the mechanism by which the various agronomic practices control scab (for example, Mortvedt et al., 1961, 1963; McGregor and Wilson, 1964, 1966). Streptomyces was inhibited by as little as 10ppm Mn2 (Bromfield, 1978) and l00ppm was severely toxic in another study (Mortvedt et al., 1963). However, since then in spite of further favourable reports, a number of papers have reported contrary or at least unconvincing results (Rodger et al., 1967; Gilmour et al., 1968; Rogers, 1969; Barnes, 1972) and the hypothesis has lost favour. A brief digression into the chemistry/biochemistry of manganese will be made here before looking again at the hypothesis that manganese is a factor in the control of common scab. Figure 9 shows the general trend in soluble Mn2+in a soil as the pH is lowered. Since the reduction of manganese also requires reducing power, and +

245


4

55

50

45

60

65

pH I I

Fig. 9. Concentration of manganese in displaced soil solutions as a function of soil pH (after Bromfield. 1979b; and McLean er a/.. 1972).

may or may not be mediated by microorganisms, the exact shape and critical pH varies from soil to soil. The general shape is, however, always similar. The reaction may be written M n 0 2 +4H

+

+ 2 e - e M n 2 ++ 2 H 2 0

showing the importance of hydrogen ions and reducing power (supplied by the organic matter). A low redox potential (reducing conditions) is favoured by easily oxidizable organic matter such as green manure crops and by restricted supply of oxygen which may be achieved through high soil moisture regimes. The reverse reaction, manganese oxidation, is carried out in soil by numerous microorganisms including bacteria, actinomycetes, fungi and algae and is favoured by the opposite conditions, aerated, non-acid soils (Bromfield, 1979a). However, if soils are very dry, these organisms are inhibited, and the only reaction which can proceed is the chemical reduction of manganese by organic matter (in spite of free oxygen). Thus during a drying cycle, Mn2+ ions may increase although absorption of manganese from dry soils by plants may be low because roots are relatively inactive in dry soils. The whole picture is extremely complicated and difficult to predict for a given soil and a given sequence of events (see Leeper, 1970, for some of the factors involved). The broad generalizations of pH, moisture and organic “green” manures remain, however, in spite of a rapidly changing microflora. As pointed out earlier, there are a few reports which have not supported the hypothesis that manganese is a factor in control of common scab. Some soils tested appear to have too little easily reducible manganese for standard practices to have any effect on soluble Mn2 (McKay, 1949; Rodger et af., +

246

R. D. GRAHAM

1967; Gilmour et al., 1969; and to a lesser extent, Rogers, 1969). McKay (1949) showed that wet conditions on a gravelly soil actually increased scab. Rogers (1969) dried the soil samples before analysis and others have stored dry soils for some time, which, as Leeper (1970) points out, may eliminate any differences (for example, see Barnes, 1972; Davis et al., 1976). Even reports of high concentrations of manganese in the periderm of scabbed potatoes is not sufficient to reject the hypothesis since the concentrations at harvest may not reflect those of the expanding periderm at the time of infection (Davis et al., 1976). Mortvedt et al. (1963) showed that MnSO, mixed through the tuber zone was more effective than in narrow bands, supporting the idea that manganese is toxic to the inoculum. However, Mn2 fertilizers are more effective for host nutrition in bands than when mixed than when broadcast; in bands, the fertilizer is protected from manganese oxidizing microbes in the soil for longer. Thus for control of soil-borne diseases such as common scab by direct toxic effects on saprophytic survival of the inoculum in the soil, we require the fertilizer to be so thoroughly mixed through the soil that it contacts all the inoculum, yet this may promote rapid oxidation of the added manganese. It is not surprising that in many cases, all treatments have proved relatively ineffective (for example, Barnes, 1972). Much higher rates mixed through the soil would seem to be required and perhaps the best way to achieve this may be the application of large amounts (perhaps up to 1 tonne per hectare) of manganese in the form of insoluble but reducible MnO,. This material may also be relatively less expensive than soluble forms, but would need to be a very fine powder. The use of large amounts of MnO, may provide a further test of the hypothesis if the treatment were able to raise the concentration of soluble Mn2+ sufficiently to control scab at a pH higher than is normally required. Measurements of soluble and exchangeable Mn2 would be needed. We can justifiably conclude, bearing in mind the complexities of transformations in soil manganese, thit results to date have not successfully disposed of the hypothesis that manganese plays a key role in control of common scab. It is however, important to look further into this matter. Mortvedt et al. (1963) concluded that it was not manganese in the plant but manganese in the soil that was important, in other words, that water-soluble soil Mn2 was directly toxic to the scab organism. If this is so, the recognized control procedures may be preventing disease by direct effects of Mn2 on the inoculum in the soil. The disease has not been eliminated by this means, however, and it appears that Mn2+must be sub-lethal but inhibitory to the growth or virulence of the organism. Bromfield (1978) also showed that a Streptomyces sp. was sensitive to high Mn2+ which inhibited growth but induced sporulation. Presumably, sporulation thereby ensured survival should the Mn2 concentration be decreased again. A number of actinomycetes (as well as fungi, algae and bacteria) have the +

+

+

+

+


247

capacity to oxidize Mn2 to insoluble, and therefore non-toxic, Mn"' and Mn'" oxides such as MnO,. Bromfield (1978, 1979a) has proposed that this capability is of survival value in acid soils particularly; Streptomyces scabies is capable of oxidizing Mn2 (Mortvedt et a/., 1963) and it is certainly possible that virulent strains of Streptomyces scabies have this capability as a means of survival in acid soils and of attacking the host. It is especially interesting that these manganese-oxidizing microorganisms perform the oxidation at some distance from their colony or mycelium and in no case has an organism been isolated from the MnO, deposits. Bromfield (1 978, 1979a) has shown that in the case of the Streptomyces, the oxidizing principle can be isolated from the host organism by centrifugation at 10 000 g for 10 minutes, but it does not pass a dialysing membrane. Dialysis was important, however, to remove inhibitors of Mn2 oxidation (which may have been ligands with affinity for Mn2+).The macro molecules were capable of oxidizing Mn2 in sodium acetate buffer at pH 5.0 with no other substances present than what was retained against dialysis. Oxidation commenced after 90 minutes. These cell-free extracts did not appear to be simple polypeptides (Bromfield, personal communication); it is conceivable they are plasmids or viruses. Whatever their nature, it is clear that they can act at a distance from the parent microorganism and I propose that they may have an important role in the infection court of any soil-borne pathogen which carries them. This could be important in the common scab-potato relationship. Firstly, Mn2 oxidizing capability could confer virulence on the scab organism simply by ensuring that it can tolerate the high concentrations of MnZt induced by cultural practices designed to suppress it, in the same way as described by Bromfield (1979a). Secondly, oxidation of Mn2+in the infection court may decrease the defensive response of the host tissues. Salomakhina (1978) has reported that Mn2 increased the thickness and speed of formation of wound periderm in potatoes, a process involving suberization and akin to the normal defensive periderm. Copper was also beneficial. Suberization is acknowledged as a partial defence in potato against common scab (Jones, 1931). A second line of defence against the pathogen involving manganese acting through the host plant physiology is supported by the data of Davis et a f . (1976). They showed that foliar manganese significantly reduced scab in a soil which could have been manganese deficient, judging from the pH and lime-at-depth. +

+

+

+

( 2 ) Lignin Biosynthesis An obvious role for manganese in host defence is the synthesis of lignin where Mn2 appears to be involved in two or more steps in the biosynthetic pathway (Gross, 1980). Figure 10 shows the major steps. Mn2+ stimulates the deamination of phenylalanine to cinnamate and the activation of the substituted cinnamic acids. Finally, in the cell walls, a Mn2+-activated peroxidase system is involved in both the production of H 2 0 2and its function +

R. D. GRAHAM

248

in forming free radicals by oxidation of the various cinnamyl alcohol analogues which are then capable of self-condensation eventually to form the polyphenolic structures known as lignin. Consequent on this non-enzymic polymerization, lignin has no fixed sub-unit sequencing (Gross, 1980; Harkin and Obst, 1973). The advantage of random sub-unit sequencing in lignin is in rendering it more immune to enzymic attack; presumably invaders would need a suite of hydrolytic enzymes to degrade such a polymer. A parallel situation has recently been reported involving random polymerization of phenolic subunits in soil to form the biologically inert complex of polyphenols known as humic acids (Shindo and Huang, 1982). This process is also analogous to lignification in higher plant cell walls in that polymerization is catalysed by manganese and proceeds in the absence of biota. Any of these Mn2 steps, especially the final polymerization step mediated by a cell wall-bound peroxidase would be subject to disruption by an extra cellular Mn2+-oxidizing particle from an invading pathogen. The role of a MnZ+-oxidizingfungal exudate in the infection process is hypothetical, based on circumstantial evidence, but appears to be entirely feasible. Lower organisms commonly do not produce Mn2+-activatedperoxidases but oxidize phenols with a Cu2+-containing laccase (Saunders et al., 1964; Harkin and Obst, 1973; Mayer and Harel, 1979), so are less inhibited by their own capability to oxidize Mn2+. Since host and pathogen may differ in their metabolism with respect to Mn2+-activatedperoxidases, availability of Mn2 +

+

Rzz&cH2" no ~ l y m e r ~ z o l ~ a nM",F*

Cinnomyl oICoholi

1

LVW"

Fig. 10. The principal steps in the biosynthesis of lignin (after Gross, 1980). Symbols of the micronutrients have been added near the steps where they are considered to participate.


249

is an obvious target for expression of virulence on the part of the attacking microorganism and for expression of defence reactions on the part of the host. (3) Gaeumannomyces graminis var. tritici (Ggt) of wheat Take-all (Gaeumannomyres gruminis, previously known as Ophiobolus gramink) is also known as hay-die because it induces white-head production before harvest. There are marked similarities in the etiology of take-all and common scab, although the organism, target species and host tissues are all quite different. The effect of pH, green manure crops, ammonium, nitrate, sulphur, phosphate, lime and manganese fertilizers and moisture all have comparable effects in suppressing both take-all and common scab (Wenzl, 1975; Asher and Shipton, 1981; Trolldenier, 1981) and in releasing M n 2 + ions in the soil (Leeper, 1970). What is different is the environment in which infection is established. Soil temperatures during the critical early growth of the potato tuber are usually quite high whereas the temperatures during infection of seminal roots ofwheat by Ggt* are often quite low, perhaps 5-10°C (although the damage to the above ground crop may not manifest itself until maturity approaches in the summer). In many of the world’s wheat crops, infection by Ggr occurs in cool, wet soils not long after emergence (Cook, 1981); these are also the conditions in which cereal crops often show manganese deficiency symptoms in climates as diverse as in the wheat belts of England (Batey, 1971), the USA (Conner, 1932) and Australia (Samuel and Piper, 1928).The severity of both take-all and of common scab is generally worse in dry years, as is manganese deficiency, with the particular exception mentioned above that manganese deficiency (and the initial infection by Ggt) both commonly occur in the seedling stage in cold wet conditions, presumably when the low temperatures (and perhaps the low oxygen tensions) prevent the host plant from absorbing adequate manganese from soils low in this micronutrient. It is commonly observed that as temperatures rise and the root system expands, crops may grow out of manganese deficiency in marginal soils. However, if manganese deficiency is indeed a predisposing factor in the development of this disease, the initial damage may have already been done. That manganese deficiency is one predisposing factor we have established recently. Figure 1 1 shows that high levels of infection by Ggt occurred only where the manganese status was low (8-18 ppm Mn) whether measured as the concentration of manganese in the shoot or the root. With one marginal exception, there were no cases where a high disease index occurred in plants with adequate manganese (the critical level is approximately 18 ppm Mn in shoots of seedlings-Graham and Loneragan, 1981). The data were derived from an experiment in which wheat plants were grown in 300 g of calcareous, manganese deficient sand fertilized with NH,N03 and 0, 1 or 10 mg per pot

* In this paper, Ggr refers to the organism; take-all, the disease.

250

R. D. GRAHAM

manganese as MnSO,. Inoculum of Ggt was supplied as infested, ground oat seed at 0, 0.05 per cent or 0.1 per cent, designed to give levels of infection comparable to field conditions. Plants were grown at 15°C for 25 days before harvest. Manganese deficiency is often difficult to induce in the glasshouse, even using soils which are characteristically deficient in the field. We have found it necessary to maintain the soil below 15"C,conditions which are also necessary for satisfactory glasshouse infection by Ggt in unsterilized soils (A. D. Rovira, personal communication).

0

5

10

15

20

25

30

Concentration o i manganese in plant

35

40

45

( ug.g-'

Fig. 1 1 . The relationship between total length of take-all lesions on seminal roots of wheat growing in a calcareoussand and the concentration of manganese in the roots or the shoots after 25 days growth at 15°C (R. D. Graham and A. D. Rovira, unpublished data). High levels of disease were associated with low levels of manganese and vice versa.

A feature of the plants in this experiment was that at harvest, there were no visible symptoms of either manganese deficiency or of take-all in the shoots. Extrapolated to the field, this observation suggests that the infection may be established before a cereal grower can see anything wrong with the crop and if the weather were then to warm up, symptoms of deficiency may never occur. Another aspect of this study was that in addition to decreasing the total amount of infection, manganese increased the number of seminal roots which were entirely free of lesions (Table IX). When symptoms appeared at a later stage, the differences were greater than those of Table IX (Fig. 5(b)). In another study, take-all was suppressed entirely by concentrations of manganese high enough to cause cylinders of MnO, to be precipitated around the roots. Underneath the MnO, shells, the roots were white and healthy. These

25 1


concentrations were not toxic to the wheat, but were well above those needed for optimal growth rate in the absence of Gaeumannomyces graminis. Several mechanisms may be proposed for the role of manganese in increasing the resistance of wheat to G g t , and are outlined below. These will be described in more detail elsewhere (R. D. Graham and A. D. Rovira, in preparation). (1) Mn2+ may be directly toxic to the inoculum as has been suggested earlier for Streptomyces. This proposal is consistent with the recent report of Reis et al. (1982) who found that take-all was decreased when manganese was increased in sand cultures from lx to 4x that of Hoagland's solution. (ii) As with Streptomyces (Section IV.C(l)) manganese may also act through the physiology of the plant. Manganese is involved in photosynthesis which in turn controls the rate of exudation of materials from the roots. Soluble exudates affect the rhizosphere microflora and through it the ectotrophic growth of Ggt (Cook and Rovira, 1976). In particular, there may be a rise in the number of fluorescent pseudomonads which have been identified with the suppression of the take-all fungus (Rovira and Wildermuth, 1981). It seems not unreasonable to inquire whether the fluorescent pseudomonads have any manganesereducing capability, even though it is clear that they were able to suppress Ggt in relatively simple media (Rovira and Wildermuth, 1981). (iii) A third role for manganese is its role in the synthesis of lignin (Section IV.C(2)), which is recognized as a partial barrier to the take-all fungus, especially in the form of lignitubers (Skou, 1981). As pointed out earlier, lignin is synthesized in situ in cell walls from monophenols oxidized by the manganese-catalysed exoenzyme, peroxidase. Whether lignin is the end result or not, many oxidized phenols are toxic and have been implicated in host defences (Bell, 1981). Since fungi commonly do not utilize a manganese and iron containing peroxidase, but a copperTABLE I X Effect ofhfanganese and Level ojGgt on the Number of Seminal Roots per Plan! with Take-all Lesions. Plants Averaged Seven Seminal Roots in Total (Data of R . D. Graham and A . D. Rovira) Manganese supplied bg/POt)

0

Amount of inoculum 0

0.05%

0.1%

0 2.2 3.1 0 2.1 2.8 0 1.8 2.0 LSD (P< 0.01) Mn, Ggt: 0.46; no interaction 1 10

R. D. GRAHAM

252

containing laccase, virulence could conceivably take the form of a manganese oxidizing particle such as has been demonstrated in Streptomyces. Numerous particles have been described in Gaeumannomyces (Rawlinson and Buck, 1981) but such a role has never been ascribed to them. Loss of particles with such a role during repeated subculturing could explain the attenuation of virulence observed by Chambers (1971), an hypothesis which may readily be tested. The oxidation of Mn2 to insoluble MnO, in the vicinity of the root could induce a local deficiency of manganese in the hose, inhibiting the defence mechanisms mediated by manganese. +

Chambers (1971) points out another interesting detail: only a few workers have ever produced the white-head symptom (associated with take-all in wheat) by inoculation with the fungus. Most of the successes have been with Australian soils suggesting that trace element deficiencies may be a factor in the expression of this particular symptom of the disease. It is important in the context of the possible role of manganese in the epidemiology of this disease (and also of common scab) that although manganese is readily transported in the xylem, it is poorly translocated in the phloem of the plant. Manganese absorbed by a particular root is transported to the xylem and thence to the shoots in the transpiration stream. It is not translocated from one root to another (Nable, 1983).Thus a fertilizer band of manganese in a deficient soil may supply the roots intersecting it and through them the shoot, but the rest of the root system may still be deficient. In this situation, Ggt may successfully invade those roots not intersecting the fertilizer, while the shoot is adequately supplied with manganese. However, during hot weather in spring or summer, this diseased root system may be incapable of supplying the evaporative demands of the crop, and the classical symptoms of the disease would follow in spite of the manganese fertilizer. This may be the situation on a number of manganese deficient soils in Australia as well as elsewhere around the world, both naturally deficient and overlimed. The hypothesis just described lends itself readily to testing in a split-root experiment where manganese is supplied to the shoot via one part of the root system, while the effects of manganese on disease severity is studied in the other part of the root system. The implications of a role for manganese in the control of take-all on soils of high pH and overlimed soils of any pH, regardless of the mechanisms involved, would be considerable. The obvious need to mix the manganese through the whole soil to the depth of infestation is opposed by the fact that mixing soluble manganese through such soils increases the rate of immobilization (Reuter et al., 1973). Extremely high rates of manganese may be needed which, as with common scab, may best be supplied as finely powdered MnO,, together with ammonium fertilizers.


253

It is consistent with the manganese hypothesis that acid soils with serious take-all problems are rare but not unknown (Yarham, 1981). The hypothesis predicts that such soils are devoid of forms of manganese capable of reduction to active Mn2+by the low pH. The disease in these soils should respond to modest amounts of soluble manganese mixed through the soil, and the prediction can be further tested by determining the content of manganese oxides in these anomalous acid soils. The alternative strategy is to breed genotypes more efficient at extracting manganese from less soluble oxide pools. The potential for doing so successfully is considerable (Graham rt d.,1983) and furthermore, preliminary results suggest there is some correlation between resistance to Ggt defined in laboratory tests and tolerance to manganese deficiency in pots and in the field (manganese efficiency). Manganese may either participate in the mechanism of the resistance, the heritable factor being in fact manganese efficiency, or there may be linkage between manganese efficiency and resistance to take-all. Robinson (1978) reported the interesting observation that in cucumber, susceptibility to powdery mildew was genetically linked to interveinal chlorosis apparently due to manganese deficiency. This report raises the question of the genetic nature of resistance. Generally speaking, major genes for nutritional characters among the micronutrients have proved to be dominant for efficiency (see Epstein, 1972) but in Robinson’s report, resistance was due to three recessives. It may be, of course, that the three genes controlled race-specific resistance which was linked to and interacting with a form of general resistance associated with the ability to accumulate manganese in the tissues. In contrast, it seems unlikely that fungi with a wide host range like Rhizoctoniu are capable of breaking down race-specific resistances in all their hosts, but rather, are efficient at breaking down the general defences (such as lignin) common to them all. Since Rhizoctoniu is a common root disease of infertile soils, resistance to it may be associated with differences in ability to maintain balanced nutrition on such soils. The hypothesis is put forward that manganese is a common factor among the many environmental effects on the severity of take-all, all of which have been described earlier. For example, the manganese hypothesis is consistent with the results of Smiley and Cook (1973) on the effects of ammonium and nitrate fertilizers on the rhizosphere pH (pH,) and in fact offers a mechanism of how the pH shift may work. In particular, the difference in “critical” pH, of sterilized and unsterilized soil would be explained differently by this hypothesis: the microbes in unsterilized soil, rather than just providing saprophytic competition, actually lower the redox potential and thereby raise the critical pH at which a given effective concentration of MnZ+ions can be maintained in the soil solution. Iron, zinc and copper appear to be the elements most likely to play a similar

254

R. D. GRAHAM

role to manganese, having, to begin with, the same trends in availability with changing pH. For the manganese hypothesis to stand, it is necessary to show that these trace elements are not involved in this way, or that they do not fit the available data as well. Soil tests show that the availability of most divalent micronutrient cations decreases as the pH rises (Black, 1968; Leeper, 1970). This means that any of several elements may also play a role in the suppression of take-all when soil acidity is increased by ammonium or sulphur fertilizers. However, manganese appears to be the element whose transformations best fit the requirements of the hypothesis. Because of the importance of H in the reduction of MnO, to Mn2+,the availability rises rapidly as the pH is decreased (Fig. 9). Further, manganese toxicity is recognized in soils below pH 5. In contrast, copper and zinc deficient soils have a bi-modal distribution over pH, being more common in both the acid and the alkaline range (Graham and Nambiar, 1981; Kubota and Allaway, 1972). In particular, Piper and Beckwith (1949) reported no effect of pH on uptake of copper by several species over the pH range 4.5-7.5; moreover, the effects of liming are often favourable or nil (Jarvis, 1981). Phosphate fertilizer aggravates copper deficiency and especially zinc deficiency (Chaudhry and Loneragan, 1970) but may decrease manganese deficiency (Conner, 1932). Nitrogen effects on these elements differ qualitatively. Because of the affinity of copper and zinc for nitrogen-containing ligands and through growth dilution (Tills and Alloway, 1981; Chaudhry and Loneragan, 1970; DeKock et al., 1971; Ozanne, 1955) nitrogen in any form commonly aggravates these deficiencies, whereas only nitrate increases the severity of manganese deficiency. Ammonium fertilizers have the opposite effect, often dramatically (Connor, 1932; Reuter et al., 1973). Cobalt deficiency is rarely, if ever recognized in the wheat belts of the world. Iron availability has a pH pattern closer to that of manganese than any of the other micronutrients but iron deficiency is rare in wheat, certainly in Australia where take-all is common. Further, Reis et al. (1982) showed no effect of iron on take-all. The responses of manganese alone parallel those of take-all in a way that is consistent with the hypothesis. In summary, while we may readily accept the conclusion of Reis et al. (1982) that additions of copper or zinc (or other elements) may decrease take-all infection, the environmental effects on manganese deficiency alone parallel closely those predisposing wheat to take-all. +

D. IRON

Iron is so fundamental to photosynthesis and energy metabolism in plants that it is difficult to separate these effects in iron deficient plants from any it may have in secondary metabolism and in specific defence mechanisms. Iron, like zinc, does activate enzymes involved in lignin synthesis (Flaig, 1973)but in


255

most cases, other ions may substitute (Gross, 1980). Iron is, however, important in peroxidases, but major effects of iron deficiency on lignin synthesis have not been noted. Again, like zinc, iron is as essential to the pathogens as it is for the host (Loring and Waritz, 1957; Saraswathi-Devi, 1958; Sadasivan, 1965) and in particular, both elements are essential to the production of pectin methylesterase by Fusarium cultures (Sadasivan, 1965). This enzyme attacks middle lamella pectins and presumably is vital to successful invasion by the fungus. Manganese inhibits the effect of iron, presumably by direct competitive effect on absorption of iron by the fungus. Interactions between metals such as the iron-manganese antagonism just mentioned are common in host pathogen relationships. For example, iron enhanced the toxicity of lycomarismin, a vivotoxin produced by Fusarium oxysporum var. lycopersici in tomatoes, and copper reversed the effect of the iron (Waggoner and Dimond, 1953). The iron and copper treatments did not inhibit the development of the fungus, so only affected the tolerance of the host to the disease; the toxin therefore is not important to the infection process. A similar situation exists with another fusarial toxin in peas (Kern and Naef-Roth, 1966). Application of iron enhanced, and copper inhibited, germination of smut spores in soil (Strakhov and Yaroshenko, 1959) and Phytophfhora spores in vitro (Halsall and Forrester, 1977). In deficient soils, a foliar application of iron increased the resistance of apples and pears to Spaeropsis malorum (Butler and Jones, 1955), wheat to smut (Yaroshenko, 1959) and turf grass to Fusarium patch disease. In contrast, Forsyth (1957) reported that iron at supraoptimal concentrations (for growth of the host alone) in the culture medium killed rust infections in the leaves of wheat. Zinc, manganese and cobalt had the opposite effect, that is, these ions broke down the resistance of a resistant variety; and iron stimulated a resistant reaction in a susceptible wheat. The inhibitory effect was specific to iron but the opposing effect was non-specific, which suggests the iron plays a role in resistance of the host to Puccinia graminis. Iron did not suppress take-all in the study of Reis el a/. (1982). Increased iron supply resulted in a tolerance of cabbage to Olpidium brassicae (Clarkson and Ferguson, 1973). The organism promoted iron deficiency in the host by impeding translocation to the shoot and this could be compensated for by additional supply of iron. Several reports on effects of iron deficiency on virus multiplication (see Nandi and Raychaudhuri, 1966) indicate that virus and host are both decreased, though at severe deficiency virus production may be decreased relatively more. Knowledge of the involvement of iron in resistance to disease is beginning to reflect a new level of sophistication with recent papers on siderophore production by bacteria in the soil (for example, Kloepper et al., 1980; Leong and Nielands, 1981; Hemming et al., 1982). Competition between host and parasite for iron is considered to be an important defence mechanism in

256

R. D. GRAHAM

animals (Weinberg, 1978) and it is clear that plant pathologists are looking for similar systems in the plant kingdom. The fluorescent pseudomonads are a group of agriculturally important gram-negative rod shaped bacteria which Cook and Rovira (1976) noticed were present in greater numbers in soils suppressive of take-all of wheat. Kloepper et al. (1980) have proposed that the fluorescent pseudomonads suppress disease by producing siderophores in the soil to compete with rhizosphere pathogens for iron. While it is clear that some bacteria within this broad group may suppress pathogens in the rhizosphere, others are themselves pathogens. It is not possible to generalize simply from the presence of this group in the rhizosphere (Schroth and Hancock, 1981). Hemming et al. (1982) and Hemming and Strobe1 (1982) have shown that production of antibiotics against fungi by soil bacteria of several classes was greatly stimulated by iron. Indeed, some of the antibiotics are themselves siderophores. They speculate that the ability of plants and pathogens to acquire iron in the presence of these bacteria may determine the course of disease, but since addition of more iron actually stimulated antimycosis in their study and suppressed it in the work of Kloepper et al. (1980), it is not yet clear what role siderophores play in the infection court. It is not clear either whether metals other than iron can in any way substitute for or inhibit iron in its role in antibiosis. Moreover, in the mechanism proposed by Kloepper et al. (1980), addition of iron actually suppressed antibiosis, which supported the hypothesis that siderophores from the fluorescent pseudomonads actually deprived the pathogens of the iron needed for their growth. It was not explained how the plants themselves avoided iron deficiency as well, but there is good evidence that higher plants have evolved their own siderophores and other adaptations (Schreiber, 1982; Kramer et af.,1980)to accommodate this situation and as well, perhaps, to aid in the suppression of the pathogens. One further question relates to a possible connection between the ironrelated stimulation of fungicides from fluorescent pseudomonads and the well established effect of pH on suppression of Gaeumannomyces graminis. As we have seen, lowering the pH suppresses take-all, and also increases the availability of iron (Leeper, 1970). It is difficult to reconcile this presumed involvement of iron with the manganese hypothesis of take-all control presented earlier. It seems unlikely that the iron-pseudomonad hypothesis could explain the manganese effects of Fig. 11 and Table IX, or that manganese has a role in the efficacy of iron-stimulated fungicides from Pseudomonas. The effects of iron and manganese seem to stand as alternatives, but they may work in the same direction in some interconnected way. For example, increasing the concentration of manganese can induce a relative deficiency of iron (Olsen, 1972) and such an antagonism may link the observations on these two elements. It should be possible to study some of these questions in “take-all’’ soils by varying the ratio of available


257

iron : manganese. Chelating agents, including siderophores, many of which bind iron more strongly than manganese, may also be useful but it is important to recognize that while siderophores may lower the activity of iron in the rhizosphere, they may also increase its mobility through the soil matrix. Studies of this kind, as for others mentioned earlier, may be facilitated by separating the nutrition of the shoot from that of the roots subject to infection, by use of the split root technique. E. ZINC

The role of zinc in the defence mechanisms of higher plants is far from clear. While on the one hand there are some reports of beneficial effects of added zinc increasing host resistance, for example, to mildew and leaf spot (Singh and Aggarwal, 1979; Akai, 1962; Bolle-Jones and Hilton 1956; Tomlinson and Webb, 1958), there is also a number showing no effect or the opposite effect, i.e. that zinc deficient plants are more resistant (Basu-Chaudhary, 1967; Meyer, 1951; Hewitt and Jones, 1951; Cherewick, 1944; Spinks, 1913). In multi-element studies with fungal diseases, zinc is commonly the least effective treatment. It is perhaps, unwise to draw conclusions from such generalities. To start with, reports of no effect may simply mean that, in the soil under investigation, no deficiency of zinc existed. Further, reports of zinc stimulating disease may not mean that it is a direct effect but possibly an indirect effect of zinc aggravating another micronutrient deficiency such as of copper or manganese. Interactions caused by competitive inhibition of absorption of one element by another are common among the micronutrients (Epstein, 1972). Zinc deficient plants may, depending on the soil, be very high in manganese, which may exert a dominant effect on susceptibility; addition of zinc to such plants will markedly decrease the manganese content (Rose11 and Ulrich, 1964). Conclusions can be safely drawn only from well designed experiments with proper controls and adequate chemical analyses (Section 1V.J). One of the consistent reports with zinc is its stimulatory effects on germination of fungal spores (McIlveen and Cole, 1979; Halsall, 1977; Strakhov and Yaroshenko, 1959; Agarwal, 1959; Sharp and Smith, 1952; Yogeswari, 1950), but whether this effect of zinc is significant in pathogenesis generally, as it was in the first mentioned case, is not known. Table X shows how critical the conditions are for vesicle formation (a phase in the development of infection which occurs after germination and penetration of the stoma). The optimal pH range is 6.2-6.6 and the optimal zinc concentration (12 ppm) is approximately the critical minimum concentration in a rapidly expanding wheat leaf. Iron, manganese, copper, molybdenum, calcium and magnesium did not substitute for zinc. The results of Halsall (1977) and Halsall and Forrester (1977) are particularly valuable since the

258

R. D. GRAHAM

TABLE X Effect ofZinc Concentration and pH on Induction of Vesicle Formation (an Early Step in Infection) by Puccinia coronata on a Gelatin Medium (Data Drawn from Figure 2 of Sharp and Smith, 1952)

Zinc concentration (ppm)

PH ~~

0

6

12

18

0 0 0

0 6 0

0 21

0 0

3

2

~~

5 *9 6.5 6.9

effects of several elements on formation and infectivity of Phytophthora cinnamoni and P . drechsieri zoospores were compared over meaningful concentration ranges. Both iron and zinc were stimulatory to germination when added in low concentrations such as we may expect in soil solutions, whereas of the seven micronutrients tested, only copper was inhibitory at low or moderate concentrations (10 -'M Cu); this inhibition was suppressed by addition of a chelate as one would predict if absorption of the copper across the plasma membranes were necessary for expression of its toxicity (Graham, 198 1). Zinc was also far less toxic than copper to nematodes (Turlygina, 1978). Of the biochemical roles known for zinc, the most fundamental are its roles in RNA synthesis and protein metabolism. With zinc deficiency, protein synthesis is significantly inhibited and high concentrations of non-protein nitrogen, including amino acids, build up. Such an accumulation would appear to be favourable to invading heterotrophs but this is not universally the case. There are a number of reports of suppressive effects of zinc on soil-borne diseases (Sulochana, 1952; Bolle-Jones and Hilton, 1956; Primavesi and Primavesi, 1964; Mehrotra and Claudius, 1973; Kuan and Erwin, 1980; Reis et af.,1982; Jones, 1981). Nikolaeva (1977) and Daniel1 and Chandler (1976) reported a beneficial effect of zinc on resistance of sorghum and peach respectively to bacterial diseases in zinc deficient soil. Prasad (1 979) reported that zinc at 3 mg 1 in culture solution checked the symptoms of Fusarium oxysporiurn var. fini on flax but did not stop the spread of the fungus. In this case, zinc is conferring a form of tolerance rather than resistance. Remission of symptoms was attributed to inhibition of production of the toxin, fusaric acid. With respect to infection of soil-borne pathogens in particular, the recent findings of Welch et af. (1982) may be highly relevant. They found that zinc deficient roots lost membrane integrity; so leakage of soluble organic substrates into the environment may be attractive to motile spores and also aid the invasion process. It is important that the zinc is required external to the membrane for its integrity and cannot be supplied from elsewhere in the plant.

-'


259

The same seems to be true for manganese, mentioned earlier, and for boron and calcium (Haynes and Robbins, 1948). It is likely, therefore, that in zinc deficient soils, as for manganese, boron and calcium, supply of the element in a fertilizer band may adequately supply the shoot and some roots but the rest of the roots may remain low in zinc and be vulnerable to pathogens. In the context of zinc (and copper) deficiency, mycorrhizal infection is important. Mycorrhizae have been reported frequently to supply roots with a limiting element (Smith, 1980),including zinc; and Mcllveen and Cole (1979) observed that application of zinc increased the germination of spores of Glomus mosseae and promoted its infection of soybean roots. Thus, application of zinc may stimulate mycorrhizal infection which in turn stimulates uptake of zinc by the host. Given that zinc deficiency interferes with RNA and protein synthesis of the host (Mengel and Kirkby, 1978), it would not be surprising if virus multiplication was inhibited in such plants since it depends on exploiting host plant metabolism. In fact, no such simple picture emerges. Helms and Pound (1955) found that zinc supplied to TMV-infected, zinc deficient tobacco plants affected the yield of virus and host proportionally even when added in excess. In contrast, Yarwood (1954) found that zinc in excess greatly increased the susceptibility of bean leaves to the same virus. Others have found inhibitory effects of zinc, including Korant et al. (1974) with rhinoviruses (of animals) in which zinc appears to inhibit the proteolytic cleavages which produce smaller viral polypeptides. In contrast to the above are the striking reports of zinc mitigating the symptoms of virus infections without eliminating the viral particles within. These effects involve regreening, usually, and prolonged leaf duration (hormonal-type effects). In one study (Dabek and Hunt, 1976) of a disease attributed to mycoplasma, regreening was induced in infected leaves by zinc (and copper) but not in healthy leaves; effects of exogenous applications of hormones, kinetin, IAA and GA, were weaker and not specific to diseased leaves. The mechanism of this therapy is unresolved but tolerance of this kind is undoubtedly an important effect, probably operative in leaves otherwise adequately supplied with zinc; that is, supraoptimal applications can be beneficial and zinc therapy has been advocated in a number of situations: court noue of vines (Dufrenoy, 1934), phyllody virus of white clover (Carr and Stoddart, 1963), beet yellows virus (Sutic and Spasic, 1963), and curl virus of hops (Schmidt et al., 1969; Prusa et al., 1965; Prusa, 1963; Zattler et al., 1962). In the study by Carr and Stoddart (1963), zinc status ranged from marginal deficiency to severe toxicity (90 per cent yield deppression by zinc), yet over this whole range, increasing rate of zinc application decreased the expression of virus symptoms and appeared to immobilize the virus such that virus-free seed could be obtained. When virus-suppressed vegetative plants were transferred to soil of normal zinc content, symptoms of the viral disease

R. D. GRAHAM

260

reappeared in three weeks. It seems possible that in some situations, zinc is overcoming a virus-induced zinc deficiency, but not at the high rates described above. Carr and Stoddart's report of zinc immobilizing the virus is consistent with the evidence reported by Welch et al. (1982) of leaky membranes in zinc deficient plants, but again this explanation could not be extrapolated to zinc toxicity. Manganese enhanced the expression of virus symptoms (Lee and Singh, 1972) indicating a competitive effect on zinc action. Regardless of the economics of zinc therapy for virus diseases these reports undoubtedly suggest an important physiological reaction, the significance of which still remains to be discovered. It is interesting that Verderevskii (1964) believed that zineb instead of Bordeaux mixture would have more positive effects by reason of its zinc content. These effects of zinc on symptoms recalls the established view that the best way of preventing a viral disease does not lie in the prevention of the infection, but in making the disease take a form of latent development. F. NICKEL

Rusts are sensitive to nickel salts, a consistent effect that has been confirmed many times since the early work of Sempio (1936) (for example, Forsyth and Peturson, 1959; Isaeva, 1969; Romanova and Gibadullin, 1980). Table XI shows how wheat was protected from two rusts by nickel salts in various soluble forms applied to the leaves. Nickel sulphate has been used in mixtures with conventional organic fungicides (Bachchhav et al., 1978), is effective supplied to the roots or the leaves, and nickel-amine complexes are less readily leached from the leaves by rainfall than is the sulphate (Forsyth and Peturson, 1959; Keil ef al., 1958). Nickel has both eradicative and protective action. Only a few minutes contact is needed for the effect which acts at sub-lethal concentrations suggesting physiological mechanisms. Smirnov (1978) and Smirnov and Krupnikova (I 978) have reported on some biochemical TABLE XI

The Protective Effect of Foliar Applications of Nickel Salts and Complexes Against Leaf and Stem Rust of Wheat (afrer Forsyth and Peturson, 1959)

Average number of infections per leaf Chemical Ni (NO,), Ni C1, Ni acetate Ni CI,-amine" chelate control a

Ni conc. (PPm) 70 70 70 280 0

leaf rust

stem rust

0.1 6 0.4 0-2

0 0.2

I1

amine chelate : bis [N-(2-hydroxyethyl)dodecylbenzylamine].

0.1

0 19


26 1

responses of plants to nickel supplied at 2.5 ppm. In particular, I A A metabolism was disturbed; IAA oxidase decreased and polyphenol oxidase increased. Nickel has been shown to be part of the plant enzyme urease (Dixon et al., 1975) and it seems likely to be established as an essential plant micronutrient as it has already for animals. From the many reports of effects of this element on host-pathogen relationships, we can safely predict that responses to nickel in plants low in this element will be greater when the plants are carrying disease, especially rusts. Nickel (and chromium) increased polyphenol oxidase activity (Smirnov, 1978) suggesting that it may affect phenol synthesis. The fact that nickel is effective in eradicating well established infections (Forsyth and Peturson, 1959), however, suggests a fungitoxic action; but since supply via the roots is effective (Sempio, 1936) and the concentrations in the leaves cannot be very high, it seems unlikely that nickel is directly toxic to the fungus. Rather, the effects on phenol metabolism may be inducing the synthesis of particular phytoalexins. Pisatin, a phenolic phytoalexin of the isoflavanoid class is a case in point. Perrin and Cruickshank (1965) suggested that nickel may bind at an enzyme sulphydryl group inhibiting the alternative isoflavanoid pathway and stimulating pisatin production. Other compounds including copper, iron and mercury as well as recognized sulphydryl inhibitors were even more effective than nickel in this case, but in another report, nickel was one of the most effective metals in stimulating production of fungitoxic diffusates in rice leaves and in protecting the leaves against Helminthosporium (Sinha and Giri, 1979). G . SILICON

Silicon appears to play some role in plant cell wall structure, providing greater rigidity, bht an essential role is not generally recognized. Silica is well known amongst rice agronomists for the piotection it provides against rice blast (Piriculuria oryzae) and brown spot (Helminthosporium oryzue). A typical response is shown in Table XII. Wet rice in particular normally absorbs silicates readily, and absorption is promoted by magnesium. Matsubayashi et al. ( 1 963) described the silicification of the cell walls hindering penetration by the fungus, and pointed to the use of calcium silicate fertilizers and rice straw compost as important sources of silica for rice. Volk et a/. (1958) described a correlation between rate of infection by the fungus which normally attacks the host through the epidermis and changes in the silicon content of the leaves during maturation. Table XI1 shows clearly that susceptibility was better correlated with the concentration of silicon in the leaves than with that of nitrogen. Similar effects of nitrogen and silicon have been reported by Matsuyama (1975), Mitsui and Takatoh (1963), Huang et al. (1980) and others. Matsubayashi et al. (1963) also stressed a high Si : N ratio for healthy rice plants. Matsuyama (1975)

262

R. D. GRAHAM

TABLE XI1 Effect of Nitrogen and Silicon Supply on the Susceptibility of the Top Leafof 41-day-old Rice Plants to Piricularia oryzae (after Volk, Kahn and Weintraub, 1958) Nutrient supply -N -N +N +N

-Si +Si -Si +Si

No. of lesions (cm -') 5.0 2.3 1.4

4.2

Leaf concentration at inoculation %N %Si 5.1 5.3 5.5 5.5

0.48 2.12 0.35

1.32

pointed to the decreased production of lignin, other phenols and hemicellulose to explain the effects of nitrogen on disease but nitrogen may also act through growth dilution of the silica (Table XII). Baba et al. (1956) pointed out that known effects of humidity on the disease may also be explained through effects of transpiration on leaf silica content. Resistance of barley to powdery mildew was increased by application of silicates (Lowig, 1933, 1937; Germar, 1934; Wagner, 1940; GrosseBrauckmann, 1957). Grosse-Brauckmann described an interaction with nitrogen such that increases in N were most damaging in terms of greater infection by Erysiphe graminis when silicon was low. Since manganese also appears to be involved in resistance of cereals to powdery mildew, the curious effect of silicon on the distribution of manganese within leaves (Williams and Vlamis, 1957; Horst and Marschner, 1978) may be relevant in considering the mechanism. Without silicon, manganese is unevenly distributed within the leaf. Kunoh et al. (1975) described bands first of calcium then silicon and manganese around the infection peg of powdery mildew on barley. Certainly, it seems significant that both Piricularia and Erysiphe enter the host by means of an infection peg which punctures the epidermis and therefore physical barriers are likely to be important. Engel (1953) reported that silica in oat leaves was present in a number of forms including stable esterified Si-galactose units loosely associated with other organic constituents and phosphorus. The molar ratio of Si : galactose increased from 1 : 1 to 2 : 1 during maturation and this type of change has been suggested as a basis for the increased resistance in older rice and barley plants. From the above, one may conclude that silicon contributes to a simple preformed physical barrier in the leaves to the penetration of the fungal haustorium and hyphae. A series of careful studies by Heath (1972, 1974, 1979, 1980, 1981) has shown that while the silicon barrier may be largely physical in function, it is a complex picture. In these studies of bean rust (Uromycesphaseoli)it appears that virulent strains are capable of dissolving the silicon deposits and that extracts from infected leaves applied to healthy leaves makes them susceptible to non-virulent strains normally incapable of


263

dissolving the siliceous deposits. In fact, the presence of the non-virulent strains can actually stimulate the deposition of silicon. Thus, far from being a fixed inert barrier, these silicon deposits can be induced by the presence of the fungus unless it has the capability to dissolve them. Metabolism is involved in SiOz deposition since it is inhibited by actinomycin D, cycloheximide and blasticidin S (Heath, 1979). The whole picture emerging is one much more dynamic than previously thought. The involvement of silica in such complex biochemical pathways for defence raises again the whole question of the essentiality of silicon for higher plants at a new level. Silicon is also found in roots where, in the cereals, it is confined to the endodermis (Bennett, 1982). In reporting these interesting findings Bennett speculated that the silicon may play a role in defence against disease, since the endodermis was shown to provide resistance to the penetration by Gaeumannornyces into the stele of wheat (Clarkson et a f . ,1975). While silica is widespread in the earth’s crust, there are of course some soils which are quite low in silica and especially if acid as well, deficiencies of silica which inhibit the plants’ response to stress caused by (fungal) diseases may indeed exist elsewhere than in the rice fields. Deficiency of silicon may also occur in plants grown under controlled conditions. H. OTHER ELEMENTS

Many trace elements-in the broadest sense ofelements which may be present in plants in small amounts-have at one time or another been reported as influencing a host-parasite relationship. These include Cd, Co, Cr, Ge, Pb, Br, I, Cl, Na, Li, Be, Al, Mo, F, Hg. For example, some of the most clear-cut effects of metals on disease are the suppressive effects of lithium and cadmium on powdery mildews. Considering the generally delicate balance between host and pathogen, it is not surprising that spraying or otherwise dosing the host with a reactive element will upset this balance in favour of either organism under some environmental conditions or other. It would be more surprising if it didn’t. Whether any of these isolated records has biological significance in terms of pathogenicity, biochemistry or control, it is usually difficult to say. The effects of these elements, if they have any generality or usefulness would be achieved at concentrations above any possible requirement for the host, and this contrasts with most of the earlier discussion which related to effects observed over the deficiency range of the element for the host plant. In some cases of metals with known toxicities, it is possible that they are more toxic to the pathogen than to the host. This appears to be the case for cadmium which is inhibitory to the germination and further development to haustorium stage of powdery mildew (Sempio, 1939; Meyer, 1951), at concentrations which can be accumulated in the host. Just 5 x 10-’M Cd

264

R. D. GRAHAM

added to sand or solution culture halved the infection (Meyer, 1951). However, Sempio ef al. (1971) following a series of papers on the cadmium effect concluded it was not toxicological, either direct or through phenols, but rather a cadmium-infection synergism on host metabolism which lasts for about 24 hours after infection. Cadmium seems unlikely to be useful as a control measure itself, but if it interacts with another micronutrient, then these reports may yet shed some light on defence mechanisms. Similar arguments apply to other toxic metals, including lead. The study of lithium dates back to Spinks (19 13). The suppressive effect of lithium is much stronger on Erysiphe graminis than on Puccinia triticina (Wortley, 1939), a distinction which occurs frequently, for example, also with copper and manganese, and in reverse with iron and nickel. This distinction may be related to the different modes of entry of these two obligate parasites of the cereals: mildew pushing down through the tough physical barrier of the cuticle and epidermal cell wall and rust generally entering “the easy way” via the stomata1 cavity. These two groups of obligate pathogens fall into different classes taxonomically. Although they form similar types of lesions on leaves and stems of, quite often, the same groups of hosts, it seems clear that the host defences are fundamentally different in some way and this difference is linked to the trace elements which participate. It is interesting to note that the stomata of copper deficient wheat plants do not open normally (Graham, 1976) and this could partly explain their lack of sensitivity to rust compared to mildew. Lithium increases the thickening of the cell wall below the infection peg (Wortley, 1939), an effect which would appear to involve lithium in the biochemical pathways of defence. Crown gall and club root, two hyperplastic diseases, also may be suppressed by lithium under certain conditions (Palm, 1963; Kent, 1941). The role of lithium in disease resistance is perplexing. Not known to be essential for higher plants and at usual concentrations not known to be exceptionally toxic, it may be representative of all those elements present and widespread in the lithosphere in small amounts, and so also in plants, for which no essential role is known. But it is entirely possible that they catalyse or otherwise contribute to a biochemical pathway which provides a defence mechanism against parasites, a role which would not be discovered in the relatively disease-free environment of the plant nutrition laboratory where such essential roles are actively sought. From the literature, the most likely elements in this class appear to be silicon, nickel, aluminium, lithium, chromium, iodine and fluorine. Aluminium is of interest as its concentration in the soil solution rises to toxic concentrations as the pH falls below 5.0 (Black, 1968) in a manner similar to manganese. It is therefore a potential factor in the control of common scab of potatoes and take-all of wheat by lowering the pH. (Other control measures such as irrigation are less likely to raise the aluminium


265

concentration). However, there are few reports of effects of aluminium on host-pathogen relationships: Greis (1951) controlled scab at the very high concentration of 20 ppm Al. Aluminium was much more effective than iron in detoxifying the metabolites of Fusarium martii var. pisi on pea (Kern and Naef-Roth, 1966). Although molybdenum and cobalt have recently been shown to activate a fungal aminopeptidase (Plumbley and Pitt, 1979), they do not occur in the literature frequently in relation to disease, probably reflecting the infrequent occurrence of deficiency of these elements. Low concentrations of molybdenum had some suppressive effect on zoosporangial formation in Phytophrhoru in contrast to zinc, boron, manganese and iron (Halsall, 1977). Cobalt sometimes acts like nickel, though less effectively. A literature exists on the effect of heavy metals and other pseudophysiological elements like fluorine, iodine and aluminium on viruses. Most frequent references are to molybdenum (Verma and Verma, 1967). The effect, as with cadmium and other protein denaturation agents, is directly on the virus and not through host plant susceptibility. A comment must be made here about chlorine which as sodium, potassium or ammonium salts has been reported to control corn stalk rot and take-all (Christiansen et a f . , 1982). Chloride is not acting here as a micronutrient as large amounts are required. Huber and Watson (1974) argued that the mechanism of chloride action is by competitive inhibition of nitrate absorption. Huber (1981) has also suggested this may be the mechanism by which chloride suppressed take-all (Powelson and Jackson, 1978) since nitrate is known to promote infection by Gaeumannomyces graminis. However, this seems unlikely since it is also argued that the mechanism of take-all stimulation by nitrate is the increase in pH, of the rhizosphere caused by excess anion absorption (Smiley and Cook, 1974; Mengel and Kirkby, 1978), and the absorption of another anion, chloride instead of nitrate, should not change materially the pH, effect. 1. SUMMARY OF MICRONUTRIENT EFFECTS

The summary of principles pertaining to macronutrient effects on susceptibility of plants to disease (Section III.F(i) to (v)) generally applies also to micronutrients with the major exception of (iv) relating to effects only in the deficiency range of the element. In contrast to the largely structural, conformational and osmotic roles of the macronutrients, the micronutrients act as catalysts, cofactors and inhibitors. In these roles supraoptimal concentrations are physiologically important. Thus while the supply of micronutrients normally increases the resistance to disease of deficient plants, there are important effects of both essential and non-essential trace elements in luxury, but not necessarily phytotoxic, concentrations. This type of effect is

266

R. D. GRAHAM

presumably due to competition between the element added in excess and another functioning element at the site of absorption of the latter or at its functional site in the processes of defence. Although iron, zinc, nickel and cobalt may catalyse certain biosynthetic pathways leading to phenols and lignin (Gross, 1980; Flaig, 1973), their participation seems less critical than that of copper, boron and manganese. In fact, these two groups of elements appear to act differently in other ways as well, perhaps the most clear-cut being the connection of the former with control of rusts and the latter with the control of powdery mildews. Zinc and iron stimulate spore germination; copper, particularly, and manganese tend to be inhibitory. Iron and zinc stimulate pectin methylesterase from Fusarium; copper and manganese are inhibitory. It seems possible to identify copper, boron and manganese (and silicon and lithium) with the formation of preformed and inducible physical barriers and the other group of elements with phytoalexin production. Such a generalization may be useful but even if the basis of it were more soundly based than it necessarily is, there would be danger in any extrapolation because competitive inhibition of active sites exists between virtually every pair of divalent cations, both between and within these groups. Exceptions to virtually every generalization are possible by changing the balance of these elements, and examples are already to be found in the literature. Special host-plant adaptations for iron absorption become important for tolerance of a high Mn : Fe ratio in the root environment which seems to be unfavourable to root pathogens. It was argued in Section I1 that there appeared to be little evolution of mechanisms in plants to cope with trace element deficiencies; retranslocation in the phloem, for example, is very poor for all the micronutrients. However, iron is the exception: plants have evolved special mechanisms for absorption and transport of iron including secretion of special ligands, development of transfer cells and synthesis of siderophores (Kramer el a/., 1980; Schreiber, 1982). An important question is the role of iron in production and activation of antibiotics and siderophores by various antagonistic bacteria in the soil. Stimulation of antibiosis by excess free iron and by induced iron deficiency through siderophore release have both been mooted. We need to know too whether copper and manganese are antagonistic to these responses to iron and under what conditions. These are important research questions for the future and are certain to influence thinking in micronutrient nutrition of plants as well as in biological control. These issues lend themselves readily to experimentation. Iron deficiency is undoubtedly an important stress on higher plants in many parts of the world, especially in its transient form associated with temporary waterlogging, low root zone temperatures and as a consequence of high pH, manganese, copper or zinc concentrations. Iron appears to be equally limiting


267

for root-zone microflora, judging from their capability for siderophore production. Competition between host and pathogen for this nutrient resource seems to be crucial for host-pathogen relationships, with the other elements as interacting, and sometimes decisive, factors in the point of balance achieved. We have seen that a number of trace elements which are not recognized as essential to plants do strongly influence the host-pathogen balance. Criteria of essentiality (Arnon and Stout, 1939) are usually established in artificial conditions of laboratory and glasshouse. Especially for micronutrients these conditions must be dust-free and so almost sterile. It seems highly probable that if criteria of essentiality are extended to include the normal environment in which plant species must survive, further elements will be established as essential, and more still as beneficial. The normal environment includes the usual microflora of both pathogenic and non-pathogenic microorganisms and other pests. From this review, the elements most in questi6n are silicon, nickel and lithium. We may summarize the principles which seem to emerge from a study of the micronutrient effects on susceptibility to disease: Correction of a micronutrient deficiency generally increases the tolerance and/or resistance of plants to pathogenic diseases. Further protection in some conditions is conferred by a number of trace elements in concentrations above those needed for host plant growth in disease-free conditions. In other situations, addition of a trace element may exacerbate the severity of disease. In these cases it would appear that interactions between trace elements or with nitrogen are involved, and element imbalances in the host are likely to be a predisposing factor. The contribution of nutrition to disease resistance is usually only partial, there being many other factors. Yield responses to an element often may contain two or even three components, due to overcoming the deficiency, changing the host plant’s defence against disease, and having direct toxic effects on the pathogen. Copper, boron and manganese all influence the synthesis of lignin and simple phenols; zinc, iron and nickel have generally different effects, possibly related to phytoalexin synthesis; silicon and lithium appear to affect physical barriers to invasion. Silicon, nickel and lithium may be essential elements in the biochemical pathways of defence. As in animals, iron may be a key element for which host and pathogen compete; an important factor in host-pathogen relationships is the iron :manganese ratio.

268

R. D. GRAHAM

J. GUIDELINES FOR EXPERIMENTATION

Many studies connecting nutrition and disease are difficult to interpret. It is important, for example, to know whether the soil is deficient in the nutrient in question. As a case in point, we noted in Section 1II.C that in Last’s study the soil was only slightly deficient in potassium, if at all, and the effects of potassium on the disease were correspondingly small, whereas large responses to potassium may be associated with marked increase in disease resistance. Likewise, where micronutrients are reported to be ineffective, it may be simply that the test soil was not deficient. For a study to be conclusive on these issues, it is necessary to have adequate controls in the form of “nil”, “complete” and “complete minus the test element” treatments, each with and without the test organism present. Several rates of application of the test element, rather than plus-and-minus only, are essential to a more rigorous interpretation of the relationships involved. Likewise, it is equally desirable to combine factorially the nutrient treatments with a range of levels of the pathogen. Further, it is necessary to present not only a disease index but also plant yields, and highly desirable indeed to have plant analysis data for as many elements as may impinge on the results and its interpretation. It is also necessary to indicate the nature of the disease index and how it may have been affected by the growth response where there is one. There are several ways to apply disease treatments. Inoculum of a soilborne disease may be applied at increasing rates to an uninfested soil, in which case killed inoculum added to soil would be the proper control. Sterilized soil may be used but this is undesirable since there are many side effects, biological, chemical and physical, besides eliminating the target organism. In particular, sterilization releases essential mineral nutrients such that the soil may no longer be deficient, for example, in manganese or copper (Piper, 1942). Where the disease is seed-borne, surface sterilization or disease-free seed may be used as the control, but it is important to note that two sources of seed may differ in trace element content (especially important for manganese and molybdenum) to an extent which may eliminate the deficiency. Mention was in Section II1.D of how a nil-disease treatment may be obtained using an effective fungicide or antibiotic. Because most reports in the literature are the result of inadvertant natural infestation of simple fertilizer experiments, the necessary controls on the disease factor have rarely been included. Moreover, in reporting the disease scores, yields and particularly plant and soil analyses are commonly omitted. There are few studies which have so far met all these criteria. These guidelines are based on the premise, established with the macronutrients, that effects on susceptibility to disease occur over the deficiency range of the element concerned. The guidelines also apply to the micronutrients. Additionally, however, among the trace elements, supraoptimal applications of some elements may increase the resistance of the host plant. In such cases,


269

“complete nutrients” is the appropriate control for test treatments of “complete +extra test element”, preferably again at several rates, and with rates of inoculum or ffungicide etc., in factorial combination. The “complete” treatment should, of course, be just adequate in the test element, which condition may generally be demonstrated by plant analysis. Distinguishing effects in the deficiency and supraoptimal ranges has both physiological and agronomic implications. In considering factorial experiments, yet other factors such as temperature and moisture also interact and choices limiting the scope of study must be made in accordance with the stated aims. Micronutrient deficiency in the shoot usually decreases the export of photosynthates to the roots, compounding the direct effects of deficiency on the roots. For studying diseases of roots in soil, it is possible to separate the two effects using the split root technique. One part of the root system can be used to supply adequately the shoot with the element which is limiting to the other part of the root system. These roots may then be used to study the effects of deficiency on the host-pathogen relationship, since retranslocation of micronutrients in the phloem is generally poor. Deficient and non-deficient leaves can also be produced on the same plant by withdrawing the supply of the test element, a technique which may prove useful for studying the effects of deficiency on the infection processes of foliar pathogens. EPILOGUE Most of what is known of the role of trace elements in disease resistance has come from simple observational studies. Detailed studies, particularly of the effects of interactions between elements, are few and in soil systems, rare. There is a need to understand the roles and interactions of trace elements in the rhizosphere and how these are affected by rhizosphere organisms such as the fluorescent pseudomonads. The hypothesis, based on analogy with animal disease systems, that competition between host and pathogen for iron determines the course of disease will stimulate much research on how this may occur and how other trace elements interact in the process. Manganese, copper and boron appear to have important roles in disease resistance. Additionally, along with the other micronutrients, nickel, silicon and lithium may be essential to particular biochemical pathways. ACKNOWLEDGEMENTS

I am indebted to Professor H. R. Wallace and Dr P. E. Kriedemann for reading the manuscript, to Drs A. D . Rovira and S. M. Bromfield for helpful discussions, to colleagues who have allowed me to use their work or for personal communications, to staff of the Commonwealth Bureau of Soils,

270

R. D. GRAHAM

Harpenden, to colleagues in the Department of Plant Pathology, Rothamsted Experimental Station, especially Dr M. E. Finney, while I was there on study leave, to Miss J. S . Ascher for assistance and to Mrs S. Suter for the typing. REFERENCES Abia, J. A., Hess, W. M. and Smith, B. N. (1977). Naturwissenschaften 64, 437438. Adams, P., Graves, C. J . and Winsor, G. W. (1975). J . Sci. Fd Agric. 26, 899-901. Agarwal, G. P. (1959). Phyton 12, 2, 87-91. Akai, S. (1962). Potash Rev. 23/27, 1-2. Antonova, G. G., Shestiperova, Z . I. and Shuvalova, G. V. (1974). Zap. Leningr. Sel‘-Khoz. Znst. 239, 81-86. Arnon, D. I. and Stout, P. R. (1939). Plant Physiol., Lancaster 14, 371-375. Asher, M. J. C. and Shipton, P. J. Eds (1981). “Biology and Control of Take-All”. Academic Press, London. Baba, I., Iwata, I., Takahashi, Y. and Kittaka, A. (1956). Proc. Crop Sci. Soc., Japan 24, 169-172. Bachchhav, M. B., Hapase, D. G., Patil, A. 0. and Ghure, T. K. (1978). Sugarcane Path. Newslett. 20, 33-35. Barnes, E. D. (1972). Record of Agric. Res. 1971, 20, 35-44. Basu-Chaudhary, K. C. (1967). Neth. J . Plant Path. 73, 49-51. Batey, T. (1971). Tech. Bull. 21, Min. Agric. Fish. Fd., London. Beckman, C. H. (1980). In “Plant Disease” Vol. V, “How Plants Defend Themselves” (J. G. Horsfall and E. B. Cowling, Eds), pp. 225-245. Academic Press, New York and London. Bell, A. A. (1981). Ann. Rev. Plant Physiol. 32, 21-81. Bennett, D. M. (1982). Ann. Eot. 50, 239-245. Bjrarling, K. (1946). Meddn. St. Vaxtsk Anst. Stockholm Nr. 47, 16. Black, C. A. (1968). “Soil-Plant Relationships” 2nd Ed. John Wiley and Sons, New York. Bolle-Jones, E. W. and Hilton, R. N. (1956). Nature, London 177, 619-620. Bortels, H. (1927). Biochern. Z . 182, 301-358. Borys, M. W. (1968). In “Progress in Soil Biodynamics and Soil Productivity” (A. Primavesi, Ed.), pp. 385404. Pallotti, Santa Maria, Brazil. Bowen, J. E. and Gauch, H. B. (1978). Plant Physiol., Lancaster 41, 319-324. Bromfield, S. M. (1978). Aust. J . Soil Res. 16, 91-100. Bromfield, S. M. (1979a). Soil Eiol. Biochern. 11, 115-118. Bromfield, S. M. (1979b). In “‘Workshop on Acid Soils”, pp. 43-54. Dep. Agric. N.S.W ., Rydalmere. Bussler, W. (1981). In “Copper in Soils and Plants” (J. F. Loneragan, A. D. Robson and R. D. Graham, Eds), pp. 213-234. Academic Press, Sydney. Butler, E. J. and Jones, S. G. (1955). “Plant Pathology.” Macmillan, London. Carr, A. J. H. and Stoddart, J. L. (1963). Ann. Appl. Eiol. 51, 259-268. Chambers, S. G. (1971). Phytopath. Z . 71, 169-182. Chaudhry, F. M. and Loneragan, J. F. (1970). Aust. J . Agric. Res. 21, 865-879. Cherewick, W. J. (1944). Can. J . Res. 22C, 52-86. Christiansen, N. W., Jackson, T. L. and Powelson, R. L. (1982). Proc. 9th h!.Planr. Nutr. CON. 1, 111-116. Comm. Agric. Bur., Slough, U.K. Chung, H. and Barnes, R. L. (1977). Can. J . For. Res. 7, 106-Il1. Clarkson, D. T. and Ferguson, I. B. (1973). Rep. Letcomb Laboratory, Wantage, pp. 28-30.


27 1

Clarkson, D. T., Drew, M. C., Ferguson, I . B. and Sanderson, J. (1975). Physiol. Plant Path. 6, 75-84. Colquhoun, T. T. (1940). J. Aust. Inst. Agric. Sci. 6, 54. Conner, S. D. (1932). J. Am. SOC.Agron. 24, 726-733. Cook, R. J. (1981). In “Biology and Control of Take-All’’ (M. J. C. Asher and P. J. Shipton, Eds), pp. 343-352. Academic Press, London and New York. Cook, R. J. and Rovira, A. D. (1976). Soil Biol. Biochem. 8, 269-273. Dabek, A. J. and Hunt, P. (1976). Trop. Agric., Trinidad 53, 115-123. Daniel, J. W. and Chandler, W. A. (1976). Hort. Sci. 1 1 , 402-403. Davey, C. B. and Papavizas, G. C. (1960). Phytopathology 50, 522-525. Davidson, R. M. and Goss, R. L. (1972). Plant Dis. Rep. 56, 565-567. Davis, J. R., McDole, R. E. andcallihan, R. H. (1976). Phytopathology66, 12361241. Dear, J. M. and Aronof, S. (1965). Plant Physiol., Lancaster 40, 458-459. DeKock, P. C., Cheshire, M. V., and Hall, A. (1971). J. Sci. Fd. Agric. 22, 437-440. Dennis, R. W. G. and O’Brien, D. G. (1937). W . Scot. Agric. CON.Res. Bull. 5, 98pp. Dixon, N. E., Gazzola, C., Blakeley, R. L. and Zerner, B. (1 975). J. Am. Chem. SOC.97, 41 3 1-41 32. Dufrenoy, J. (1934). La Potasse 8, 137-139. Dutta, T. R. and McIlrath, W. J. (1964). Bot. Gaz. 125, 89-96. Eaton, F. M. (1930). Phytopathology 20, 967-972. Edgington, L. V. and Walker, J. C. (1958). Phytopathology 48, 324326. Engel, W. (1953). Planta 41, 358-390. Epstein, E. (1 972). “Mineral Nutrition of Plants: Principles and Perspectives”. John Wiley and Sons Inc., New York. Evans, E. M., Rouse, R. D. and Gudauskas, R. T. (1964). Highlights of’Agric. Res. 1 1 , (2). Felix, E. L. (1927). Phytopathology 17, 49-50. Flaig, W. (1973). Agrochemica 17, 1245. Forsyth, F. R. (1957). Nature, London 179, 217-218. Forsyth, F. R. and Peturson, B. (1959). Phytopathology 49, 1-3. Friend, J. (1977). In “International Review of Biochemistry, Plant Biochemistry 11” (D. H. Northcote, Ed.), Vol. 13, pp. 141-182. University Park Press, Baltimore. Garrett, S. D. (1948). Ann. Appl. Eiol. 35, 14-17. Germar, 9 . (1934). Z. PjErnuhr. Dung. Bodenk. 35A, 102-115. Gill, D. L. (1972). J. Am. SOC.Hort. Sci. 97, 467-471. Gilmour, J., Crooks, P., Rodger, J. B. A., Wynd, A. and McKay, A. J. M. (1968). Edinburgh Sch. Agric. Exp. Work, 1967, pp. 36-37. Glynne, M. D. (1959). Plan1 Path. 8, 15-16. Goss, R. L. (1968). In “Role of Potassium in Agriculture” (V. J . Kilmer, S. E. Younts and N. C. Brady, Eds), pp. 221-241. Am. SOC.Agron., Madison. Graham, J. H., Leonard, R. T. and Menge, J . A. (1981). Plant Physiol., Lancaster68, 548-552. Graham, R. D. (1976). Aust. J . Plant Physiol. 3, 229-236. Graham, R. D. (1979). Plant Cell and Environ. 2, 139-143. Graham, R. D. (1980). Plant Soil 56, 181-185. Graham, R. D. (1981). In “Copper in Soils and Plants” (J. F. Loneragan, A. D. Robson and R. D. Graham, Eds), pp. 141-163. Academic Press, Sydney. Graham, R. D. and Loneragan, J. F. (1981). Proc. Nat. Workshop Plant Anal., Goolwa, pp. 95-96. Dept. Agric. S. Aust., Northfield. Graham, R. D. and Nambiar, E. K. S. (1981). Aust. J. Agric. Res. 32, 1009-1037. Graham, R. D. and Ulrich, A. (1972). Plant Physiol., Lancaster 49, 105-109. Graham, R. D., Davies, W. J., Sparrow, D. H. B. and Ascher, J. S . (1983). Plant Soil (in press).

272

R. D. GRAHAM

Greis, G. A. (1951). Phytopathology 41, 15. Gross, G. G. (1980). Adv. Bot. Res. 8, 25-63. Grosse-Brauckmann, E. (1957). Phytopath. 2. 30, 112-1 16. Guntz, M. and Coppenet, M. (1957). Phytiat.-Phytopharm. 6, 187. Hackett, C. (1972). Aust. J. B i d . Sci. 25, 1169-1180. Hallock, D. L. and Porter, D. M. (1981). Peanut Sci. 8, 48-52. Halsall, D. M. (1977). Can. J. Microbiol. 23, 1002-1010. Halsall, D. M. and Forrester, R. I. (1977). Can. J. Microbiol. 23, 994-1001. Hampson, M. C. (1980). Can. J. Plant Path. 2, 148-151. Harkin, J. M. and Obst, J. R. (1973). Science, New York 180, 296298. Haynes, J. L. and Robbins, W. R. (1948). J . Am. Soc. Agron. 40,795-803. Heath, M. C. (1972). Phytopathology 62, 27-38. Heath, M. C. (1974). Physiol. Plant Path. 4, 403414. Heath, M. C. (1979). Physiol. Plant Path. 15, 141-148. Heath, M. C. (1980). Phytopathology 70, 356360. Heath, M. C. (1981). Physiol. Plunt Path. 18, 149-156. Heggeness, H. G. (1942). Plant Physiol., Lancaster 17, 143-144. Helms, K. and Pound, G. S. (1955). Virology 1, 408-423. Hemming, B. C. and Strobel, G. A. (1982). In “Proc. First Int. Symp. Genetic Specificity Min. Nutr. Plants” Serbian Acad. Sci., Belgrade. Hemming, B. C., Orser, C., Jacobs, D. L., Sands, D. C. and Strobel, G. A. (1982). J . Plant Nutr. 5, 683-702. Hewitt, E. J. and Jones, E. W. (1951). “Rep. Agric. Hort. Res. Sta., Bristol” (1949) pp. 58-63. Horsfall, J. G. and Cowling, E. B., Eds (1980). “Plant Disease” Vol. V, “How Plants Defend Themselves”. Academic Press, New York and London. Horst, W. J. and Marschner, H. (1978). Plant Soil 50, 287-303. Huang, Y. T., Yu, C. H. and Tai, L. H. (1980). Taiwan Agric. Bimonthly 16, 57-64. Huber, D. M. (1980). In “Plant Disease” Vol. V. “How Plants Defend Themselves” (J. G. Horsfall and E. B. Cowling, Eds), pp. 38 1406. Academic Press, New York and London. Huber, D. M. (1981). In “Biology and Control of Take-All” (M. J. C. Asher and P. J. Shipton, Eds), pp. 317-341. Academic Press, London and New York. Huber, D. M. and Keeler, R. R. (1977). Proc. Am. Phytopathol. SOC. 4, 163. Huber, D. M. and Watson, R. D. (1974). Ann. Rev. Phyropath. 12, 139-165. Isaeva, G. Y. (1969). Rev. Plant Path. Abstr. 49, 507. Ismailov, Kh. A. (1954). Soils Fertil. Abstr. 19, 202. Jarvis, S. C. (1981). In “Copper in Soils and Plants” (J. F. Loneragan, A. D. Robson and R. D. Graham, E d ) , pp. 265-285. Academic Press, Sydney. Jenkyn, J. F. and Bainbridge, A. (1978). In “The Powdery Mildews” (D. M. Spencer, Ed.), pp. 283-321. Academic Press, London. Jenkyn, J. F. and Moffatt, J. R. (1975). Plant Path. 24, 16-21. Jones, A. P. (1931). Ann. Appl. Biol. 18, 313-333. Jones, G. P. D. (1981). Hons thesis, Faculty of Agriculture, University of Western Australia. Jones, I. T. and Hayes, J. D. (1971). Ann. Appl. Biol. 68, 31-39. Kaur, P., Kaur, S. and Padmanabhan, S. Y. (1979). Indian Phytoparh. 32, 287-288. Keane, E. M. and Sackston, W. E. (1970). Can. J. Plant Sci. 50, 415-422. Keil, H. L.,Frohlich, H. P. and Glassick, C. E. (1958). Phyfopathology 48, 690-695. Kent, N. L. (1941). Ann. Appl. Biol. 28, 189-209. Kern, H. and Naef-Roth, S. (1966). Phyropurhology 57, 289-297.


273

Kloepper, J. W., Leong, J., Teintze, M. and Schroth, M. N. (1980). Nature, London 286, 885-886. Korant, B. D., Kauer, J. C. and Butterworth, B. F. (1974). Nature, London 248, 588-590. Krainer, D., Romheld, V., Landsberg, E. and Marschner, H . (1980). Plntzta 147, 335-339. Krauss, A. (1969). Z . PfiErniihr. Bodenk. 124, 129-147. Kuan, T. L. and Erwin, D. C. (1980). Phytopathology 70, 981-986. Kubota, J. and Allaway, W. H . (1972). In “Micronutrients in Agriculture” ( J . J . Mortvedt, P. M. Giordano and W. L. Lindsay, Eds), pp. 525-554. Soil Sci. SOC., Am., Madison. Kunoh, H., Ishizaki, H. and Kondo, F. (1975). Ann. Plzytopath. Soc. Japan 41, 33-39. Lakshminarayanan, K. (1955). Ph.D. thesis, Univ. Madras. Last, F. T . (1962). Plant Path. 11, 133-135. Lee, S. G. and Aronoff, S. (1967). Science, Neu York 158, 798-799. Lee, C. R. and Singh, R. P. (1972). Phytoparhologj. 62, 516520. Leeper, G. W. (1970). ”Six Trace Elements in Soils”. Melbourne University Press. Leong, S. A. and Nielands, J. B. (1981). J . Bacteriol. 147, 482491. Lewis, D. H. (1980a). New Phytol. 84, 209-229. Lewis, D. H. (1980b). New Phyfol. 84, 261-270. Lipman, C. B. and Mackinney, G . (1931). Plant Pl.vsiol., Lancaster 6, 593-599. Lodeman, E. G. (1902). “The Spraying of Plants”. Macmillan, New York. Loneragan, J. F. (1981). In “Copper in Soils and Plants” (J. F. Loneragan, A. D. Robson and R. D. Grahm, Eds), pp. 165-188. Academic Press, Sydney. Loring, H. S. and Waritz, R. S. (1957). Science, New York 125, 646648. Lowig, E. (1933). Emiihr. P’. 29, 161-167. Lowig, E. (1 937). Pfanzenbau 13, 362-367. Lungley, D. R. (1974). Ph.D. thesis, Univ. Adelaide. McBride, M. B. (1981). I n “Copper in Soils and Plants” (J. F. Loneragan, A. D. Robson and R. D. Graham, Eds), pp. 2 5 4 5 . Academic Press, Sydney. McClure, J. M. (1979). In “Biochemistry of Plant Phenolics”. Recent Advances in Phytochemistry (T. Swain, J. B. Harborne, and C. F. van Sumere, Eds), Vol. XII, pp. 525-556. Plenum Press, New York. McFarlane, I . (1958). J . Gen. Microbiol. 18, 720-732. McGregor, A. J . and Wilson, G. C. S. (1964). PIanf Soil 20, 59-64. McGregor, A. J. and Wilson, G. C. S. (1966). Plant Soil 25, 3-16. Mcilveen, W. D. and Cole, H. (1979). Agric. Environ. 4, 245-256. McKay, R. (1949). Scient. Proc. R . Dubl. Soc. N.S. 25, 65-84. McLaughlin, S. B. and Shriner, D. S. (1980). I n “Plant Disease” Vol. V, “How Plants Defend Themselves” (J. G. Horsfall and E. B. Cowling, Eds), pp. 407431. Academic Press, New York and London. McLean, A. J., Halstead, R. L. and Finn, B. J. (1972). Can. J . Soil Sci. 52, 427438. McNew, G. L. (1953). In ”Plant Diseases, The Yearbook ofAgriculture”. pp. 100-1 14. USDA, Washington. Matsubayashi, M., Ito, R., Nomoto, T., Takase, T . and Yamada, N. (1963). “Theory and Practice of Growing Rice.” Fuji Publishing CO., Tokyo. Matsuyama, N. (1975). Ann. Phytopath. SOC.,Japan, 41, 5 6 6 1 . Mayer, A. M. and Harel, E. (1979). Phytochemistry 18, 193-215. Mehrotra, R. S . and Claudius, G. R. (1973). Plant Soil 39, 695-698. Mengel, K. and Kirkby, E. A. (1978). “Principles of Plant Nutrition.” International Potash Institute, Berne, Switzerland.

R. D. GRAHAM

274

Meyer, H. (1951). Phytopath. 2.17, 63-80. Miles, L. E. (1936). Mississippi Agric. Exp. Sta. Tech. Bull. 308. Mitsui, S. and Takatoh, H. (1963). Soil Sci and Plant Nutr. 9, 7-16. Mooney, H. A. (1972). Annu. Rev. Ecol. System. 3, 315-346. Mortvedt, J. J., Fleischfresser, M. H., Berger, K. C. and Darling, H. M. (1961). Am. Potato J . 38, 95-100. Mortvedt, J. J., Berger, K. C. and Darling, H. M. (1963). Am. Potato J . 40,96-102. Nable, R. 0. (1983). Ph.D. Thesis, Murdoch Univ., W. Aust. Nandi, P. and Raychaudhuri, S. P. (1966). Am. Potato J . 43, 6-9. Nesterov, P. I. and Korol’chuk, V. V. (1980). Soils Fertil., Harpenden 43, (5), Abstr. 4202. Nikolaeva, N. F. (1977). S-Kh. Biol. 12, 46&461. Olsen, C. (1939). Comptes rend. Lab. Carlsberg, (Ser. chim.) 23, 37-44. Olsen, S. R. (1972). In “Micronutrients in Agriculture” (J. J. Mortvedt, P. M. Giordano and W. L. Lindsay, Eds), pp. 243-264. Soil Sci. SOC.Am., Madison. Ozanne, P. G. (1955). Aust. J. Biol. Sci.8, 47-55. Palm, E. T. (1963). Contr. Boyce Thompson Inst. Plant Res. 22, 91-112. Papastylianou, I. and Puckridge, D. W. (1981). Aust. J . Agric. Res. 32, 713-723. Perov, N. N., Mirzaev, M. N., Chepelenko, A. P. and Perova, L. I. (1971). Fiziol. Rust. 18, 1040-1043. Perrin, D. R. and Cruickshank, I. A. M. (1965). Aust. J. Biol. Sci. 18, 803-816. Piper, C. S. (1942). J . Agric. Sci. 32, 143-178. Piper, C. S. and Beckwith, R. S.(1949). In Brit. Comm. Sci. Off.Conf. Spec. Conf. in Agric. Aust. Proc., pp. 144-155. HMSO, London. Plumbley, R. A. and Pitt, D. (1979). Physiol. Plant Path. 14, 313-328. Pobegailo, A. I., Ladeishchikova, E. I. and Ladnykh, L. F. (1980). Biol. Nauki 11, 81-86. Pollard, A. S., Parr, A. J. and Loughman, B. C. (1977). J. Exp. Bot. 28, 831-841. Powelson, R. L. and Jackson, T. L. (1978). Proc. 29th Ann. Fert. Conf. Pacific N.W., Beaverton, Oregon, pp. 175-182. Prasad, Y. (1979). Indian Phytopath. 32, 61-63. Primavesi, A. and Primavesi, A. M. (1964). Z. PflErnuhr. Dung. Bodenk. 105, 22-27. Primavesi, A. M. and Primavesi, A. (1970). Agrochimica 14, 490-495. Prusa, V. (1963). Sb. csl. Akad. zemed. Ved. C., RostIinna Vyroba 9, 645-658. Prusa, V., Pirkl, J. and Bohmova, J. (1965). Biologia PI. 7, 425-436. Rahimi, A. and Bussler, W. (1973). Z . PJErniihr. Bodenk. 135, 183-195. Rawlinson, C. J. and Buck, K. W. (1981). In “Biology and Control of Take-All” (M. J. C. Asher and P. J. Shipton, E h ) , pp. 151-172. Academic Press, London and New York. Reis, E. M., Cook, R. J. and McNeal, B. L. (1982). Phytopathology 72, 224-229. Reuter, D. J., Heard, T. G. and Alston, A. M. (1973). Aust. J. Exp. Agric. Anim. Hush 13, 440445.

Robinson, R. W. (1978). Cucurbit Genetics Cooperative No. 1, 1 1 . Robson, A. D., Hartley, R. D. and Jarvis, S. C. (1981). New Phytol. 89, 361-371. Rodger, J. B. A., Wynd, A. and Gilmour, J. (1967). Edinburgh Sch. Agric. Exp. Work, 1966, pp. 24-25. Rogers, P. F. (1969). Ann. Appl. Biol. 63, 371-378. Rohde, G. (1952). Deut. Landwirt. 3, 642-646. Romanova, A. M. and Gibadullin, M. G. (1980). Zashchita Rust. 9, 23. Rosell, R. and Ulrich, A. (1964). Soil Sci. 97, 152-167. Rovira, A. D. and Wildermuth, G. B. (1981). In “Biology and Control of Take-All”


275

(M. J. C. Asher and P. J. Shipton, Eds), pp. 385415. Academic Press, London and New York. Sadasivan, T. S. (1965). In “Symp. Ecol. Soil-Borne Plant Pathogens” (K. F. Baker and W. C. Snyder, Eds), pp. 460-470. Univ. California Press, Berkeley. Salomakhina, A. L. (1978). Nauch. Tr. Kursh. Ped. In-t. 185,45-50. Samuel, G. and Piper, C. S. (1928). J. Agric. S . Aust. 31,696-705. Saraswathi-Devi, L. (1958). J. Indian Bot. SOC.37, 509-516. Sarojini, T. S. (1950). J. Madras Univ. 19B, 1-32. Saunders, B. C., Holmes-Siedle, A. G. and Stark, B. P. (1964). “Peroxidase.” Butterworths, London. Schmidt, H. E., Dolzmann, H., Borde, K. and Holtz, S. (1969). NachrBI. dt. PflSchutzdienst. 11, 224234. Schmucker, T. (1934). Planla 23, 264-283. Schreiber, K. (1982). 2nd Int. Symp. Biol. System. Solanaceae, pp. 42-43. Missouri Bot. Gard., St. Louis. Schroth, M. N. and Hancock, J. G. (1981). Ann. Rev. Microbiol. 35,453-476. Schutte, K. H. (1964). “The Biology of the Trace Elements.” Crosby Lockwood, London. Schiitte, K. H. (1967). Plant Soil. 27, 450-452. Sempio, C. (1936). Riv. Patol. Veg. Padova 21,201-278. Sempio, C.(1939). Riv. Patol. Veg. Padova 29, 1-69. Sempio, C., Raggi, V., Barberini, B. and Draoli, R. (1971). Phytopath. 2.70,281-294. Sharp, E. L. and Smith, F. G. (1952). Phytopathology 42, 581-582. Shindo, H. and Huang, P. M. (1982). Nature, London 298, 363-364. Shkolnik, M. Y. (1974). Fiziologiya Rust. 21, 140-150. Simojoki, P. (1 969). Maasseudun Tulevaisuus, Supplement “Koetoiminta ja Kaytanto” No. 1. Singh, P. and Aggarwal, R. K. (1979). Indian J. Agric. Sci. 49,459462. Sinha, A. K. and Giri, D. N. (1979). Curr. Sci. 48, 782-784. Skou, J. P. (1981). In “Biology and Control of Take-All’’ (M. J. C. Asher and P. J. Shipton, Edr), pp. 175-197. Academic Press, London and New York. Smiley, R. J. and Cook, R. J. (1973). Phytopathology 63,882-890. Smirnov, Yu. S. (1978). Bot. Zh. 63, 16361639. Smirnov, Yu. S. and Krupnikova, T. A. (1978). Bot. Zh. 63,67&681. Smith, S. E. (1980). Biol. Rev. 55,475-510. Sommer, A. L. (1931). Plant Physiol., Lancaster 6, 339-345. Spatz, L.(1955). Mitt. dt. LandwGes. 69, 17,406. Spinks, G. T. (1913). J. Agric. Sci., Cambridge, 5, 231-247. Spurr, A. R. (1957). Am. J . Bot. 44, 637-650. Strakhov, T. and Yaroshenko, T. V. (1959). Primen. Mikroelem. Se. Khoz. Medits. Baku. (1958). pp. 373-380. Sulochana, C. B. (1952). Proc. Indian Acad. Sci. 36B,229-233. Suryanarayanan, S. (1958). Curr. Sci. 27, 447-448. Sutic, D. and Spasic, P. (1963). Z. PJKrankh. PPSchutz. 70, 339-342. Tainio, A. (1 961). Maasseudun Tulevaisuus. Supplement. “Koetoiminta ja Kaytanto” No. 12. Thompson, J. F., Morris, C. J. and Gering, R. K. (1960). Qualitas PIunta Materiae Veg. 6, 3-1, 261-275. Tills, A. R. and Alloway, B. J. (1981). Plant Soil 62, 279-280. Tomlinson, J. A. and Webb, M. J. W. (1958). Nature, London, 181, 1352-1353. Toms, J. (1958). J. Dep. Agric. W . Aust. 7, 197-203.

276

R. D. GRAHAM

Trolldenier, G. (1969). Potash Rev. 23/34, 1-16. Trolldenier, G. (1981). Phytoparh. Z. 102, 163-177. Turlygina, E. S. (1978). Akad. Nauk. Estonskoi SSR pp, 133-136. Uexkuell, H. R. von (1966). Better Crops wifh Plant Fd. 50, 28-35. Uexkuell, H. R. von (1975). Potash Rev. 9/30 (1 I), 1-8. Utkina, N. I., Nazarova, I. P., Danilova, T. A. and Bazherova, L. N. (1980). Khim. Sefsk. Khoz. 18, 13-15. Vanterpool, T. C. (1935). Can. J. Res. 13C, 220-250. Verderevskii, D. D. (1964). In “Virusiye Bopezii Rastenii” (K. S . Sukhov, Ed), pp. 293-301. Verma, H. N. and Verma, G. S. (1967). Indian Phytopath. 20, 176-178. Vlamis, J. and Yarwood, C. E. (1962). Plant Dis. Rep. 46, 886-887. Volk, R. J., Kahn, R. P. and Weintraub, R. L. (1958). Phytopathology 48, 179-184. Waggoner, P. E. and Dimond, A. E. (1953). Phytopathology 43, 281-284. Wagner, F. (1940). Phytopath. Z. 12, 427479. Walker, C. D. and Webb, J. (1981). In “Copper in Soils and Plants” (J. F. Loneragan, A. D. Robson and R. D. Graham, Eds), pp. 189-212. Academic Press, Sydney. Watanabe, R., Chorney, W., Skok, J. and Wender, S. H. (1964). Phytochem. 3, 391-393. Weinburg, E. D. (1978). Microbiol. Rev. 42, 45-66. Weinhold, A. R., Oswald, J. W., Bowman, T., Bishop, J. and Wright, D. (1964). Am. Potato J. 41, 265-273. Welch, R. M., Webb, M. J. and Loneragan, J. F. (1982). Proc. 9th Int. Plant Nutr. CON. 2, 710-715. Comm. Agric. Bur., Slough, U.K. Wenzel, G. and Kreutzer, K. (1971). Z. PJErniihr. Bodenk. 128, 123-129. Wenzl, H. von (1975). Z. P’anzenkh. 82, 410440. Wenzl, H. von and Reichard, Th. (1974). Bodenkultur 25, 130-137. Williams, C. N. (1961). In “Some Aspects of Trace Elements in Nature” (K. H. Schiitte, Ed.), pp. 3240. Univ. Cape Town. Williams, D. E. and Vlamis, J. (1957). Plant Physiol., Lancaster 32, 404-409. Wood, J. M. (1981). Hons thesis, Faculty of Agriculture, University of Western Australia. Wood, R. K. S. (1967). “Physiological Plant Pathology.” Blackwell, Oxford. Wortley, W. R. S. (1939). Trans. Brit. Mycof. SOC.23, 122. Yarham, D. J. (1981). In “Biology and Control of Take-All” (M. J. C. Asher and P. J. Shipton, Eds), pp. 353-384. Academic Press, London and New York. Yaroshenko, T. V. (1959). Primen. Mikroelem. Se-Khoz. Medits. Baku. (1958) pp. 441-446. Yarwood, C. E. (1938). Phytopafhofogy 28, 22. Yarwood, C. E. (1954). Phytopathology 44,230-233. Yarwood, C. E. (1959). In “Plant Pathology” (J. G. Horsfall and A. E. Dimond, Eds), Vol. 1, pp. 521-562. Academic Press, New York. Yogeswari, L. (1950). Ph.D. thesis, Univ. Madras. Zattler, F., Pfeifer, H. and Chrometzka, P. (1962). Brauwelt 102, 13741378. Zubko, I. Y. (1961). In “The Role of Microelements in Agriculture”, pp. 221-228. Moscow University, Moscow.

AUTHOR INDEX The numbers in italics indicate the pages on which names are mentioned in the reference lists A

Abia, J. A,, 243, 270 Adams, P., 236,270 Akoyunsglou, G., 102, 137, 189 Abad-Zaptiero, C., 68, 199 Abdulaev, N . G. 60, 210 Abeliovich, A,, 164, 189 Acker, S., 86, 99, 213 Agalidis, I., 61, 103, 206 Aggarawal, R. K., 257, 270, 275 Aitzetmuller, K., 58, 118, 214 Aizawa. M . , 62, 189 Akai, S., 257, 270 Akerlund, M.-E., 139, 189 Al-Abbas, H., 41, 199 Alberte, R. S.,52, 77, 78, 84, 94, 96, 105, 108, 122, 126, 146, 151, 156, 158, 186, 189, 193, 210, 213, 215, 216, 218 Albertsson, P.-A., 139, 189 Alfano, R. R., 67, 134, 195, 218 Allakhverdiev, S. I., 93, 203 Allaway, W . H., 254, 273 Allen, J . F . , 107, 147, 148, 189, 201 Allen, 54 Allen. M. B., 120, 189 Allen, M . M . , 41, 189 Alloway, B. J., 254, 275 Alpert, 99, 189 Almgren, 63 Alston, A. M. 252, 254, 274 Ambard-Bretteville, F., 104, 106, 107, 216 Amesz, J., 72, 73, 94, 117, 161, 189, 195, 211, 215 Anderson, J. M. 37,38,39,40,44, 51, 52, 72,76,77,78,84,85,86,88,89,98,100, 101, 102, 104, 105, 106, 107, 108, 121, 122, 124, 129, 138, 139, 140, 144, 145,

146, 147, 148, 159, 160, 162, 187, 139, 190, 191, 205, 212 Anderson, M . C . , 5, 189 Andersson, B., 39,44, 129, 139, 140, 148, 189 Antia, N., 5, 190 Antonini, E., 84, 190 Antonova, G. G . , 241, 243, 270 Apel, G. S . 66, 181 Apel, K., 80, 89, 106, 107, 173, 190 Apell, G. S . 184, 197, 198 Appel, P. W . V., 175, 182, 212 Argyroudi-Akoyunoglou, J . , 105, 107, 190 Argyroudi-Akoyunoglou, J. H. 102, 105, 107. 190 Armond, P. A.,47,137,138,144,190,206 Arnold, K. E., 28, 153, 190 Arnold, W . , 38, 73, 74, 195, 196 Arnon, D . I., 92, 190, 204, 267, 270 Arntzen, C . J., 51, 85, 86, 92, 103, 105, 107, 137, 138, 139, 147, 148, 187, 189, 190, 200, 208, 214 Aronoff, S., 239, 241, 271, 273 Arpin, N . , 58, 190 Arshad, J . H. 123, 201 Arvidson, G., 61, 202 Asada, K.,176, 209 Ascher. J . S. 253, 270 Asher, M . J . C., 249, 270 Astier, G., 92, 190 Atkinson, A. W., Jr, 20, 190 Augustsson, T. R. 178, 193 Avramik, S. M., 181, 190

B Baba, I., 262, 270 Babcock, G. T., 89, 191

278

AUTHOR INDEX

Bachchav, M. B. 260, 270 Bacon, K. E., 118, 210 Badger, M. R., 41, 190, 203 Badami, P. 116, 138, 139, 142, 156, 214 Bagyinka, Cs., 61, 215 Bahr, J. T., 40, 202 Bainbridge, A., 222,230,272 Baker, K. S., 12, 164, 214 Balch, W. E., 175, 183, 184, 186, 197 Ballschmitter, K., 102, 190 Barber, J., 112, 114, 133, 190, 207, 213 Barber, J. A., 48, 102, 105, 145, 147,210, 215 Barberini, B., 264, 275 Barghoorn, E. S., 181, 211 Barilotti, D. C., 30, 194 Barnard, A. 30, 210 Barnes, E. D., 244, 270 Barnes, R. L., 235, 246, 270 Bar-Nun, S., 78, 86, 104, 106, 109 Barr, R., 41, 199 Barrett, J., 59, 62, 66, 67, 76, 77, 78, 80, 84, 88, 89, 90, 98, 100, 101, 102, 103, 104, 105, 108, 121, 122, 124, 126, 129, 178, 189, 190, 205 Bart, J. C. J., 58, 190 Basu-Chaudhary, K. C., 257, 270 Baszynski, T., 41, 190 Bates, G., 116, 156, 216 Batey, T., 243, 249, 270 Bauld, J. 181, 190 Baxter, R. M., 12, 180, 215 Bazherova, L. N., 241,276 Beale, S. I., 167, 169, 183, 190, 211 Beardall, J., 40, 149, 154, 158, 162, 190, 211 Bearden, A., 87, 206 Bearden, A. J., 87, 200,206 Beckman, C. H., 233, 270 Beckwith, R. S., 254, 274 Beddard, G. S., 83, 191 Bekasova, 0. D., 98, 99, 174, 191, 213 Bell, A. A., 227, 251, 270 Bell, L. N., 12, 191 Bendall, D. S., 76,82,87,89,90,91,191, 214, 218 Bendall, F., 17, 18, 200 Bennett, A. 63, 166, 191 Bennett, D. M., 263, 270 Bennett, J., 102, 107, 147, 148, 189, 191 Bennoun, P.,78, 85,86,91,97, 100, 106, 137, 138, 146, 191, 194, 202, 204, 218

Bensasson, R. V., 61, 191 Benson, D., 33, 162, 163, 194 Benson, E. E., 54, 191 Benz, J., 51, 91 Berger, K. C., 243, 244, 246, 247, 274 Berkaloff, C., 100, 126, 128, 191, 204 Berman, T. 46, 195 Berns, D. S., 62, 68, 113, 114, 118, 181, 203, 206, 207 Bernstein, H. J., 61, 212 Berry, J. A., 41, 190, 203 Berthold, D. A., 89, 191 Berthold, G., 166, 191 Bethge, P. H.. 83, 218 Beudeker, R. F., 31, 191 Bialek, G. E., 47, 140, 191 Birks, J. B., 99, 191 Bishop, J., 244, 276 Bishop, N. I., 59, 62, 76, 146, 152, 164, 191, 200, 207, 209, 213 Bjorkman, O., 160, 191 Bjotn, G. O., 174, 191 Bjorn, G. S., 174, 191 Bjorn, L. O., 172, 173, 174, 191, 215 Bjerrnland, T., 58, 191 Bjerrling, K., 244, 270 Black, C. A., 222, 244, 254, 264, 270 Black, M. T., 107, 147, 200 Blaha, L. K., 110, 198 Biakeley, R. L., 261, 271 Blakemore, R., 175, 183, 184, 186, 197 Blatt, M. R., 172, 192 Blinks, L. R., 69,70,72,74, 112, 115,120, 200, 218 Boardman, 76 Boardman, N. K., 21,39.48,52,99,100, 104, 107, 138, 140, 144, 145, 146, 147, 148, 154, 158, 160, 189, 191, 194, 213, 216 Bobrowski, K., 63, 211 Bock, G., 30, 200 Boczar, B. A. 120, 122, 192, 210 Bode, V. C., 118, 192 Boger, P.,88,118,176, 182,186,192,203, 204, 212 Boggs, R. T., 102, 107, 207 Bogorad, L., 63, 80, 89, 166, 174, 190, 191, 192, 200, 209 Bohme, H., 88, 186, 192, 204 Bohmova, J., 259, 274 Bohner, H., 88, 186, 192 Bold, H. C., 5, 192

279

AUTHOR INDEX

Bolle-Jones, E. W., 257, 258, 270 Bolognesi, M . C., 103. 207 Bolton, J. R., 87, 206 Bolton, R . J., 50, 201 Bonaventura, C. J., 144, 192 Bonen, L., 175, 183, 184, 186, 192, 197 Bonnett, R., 58, 66, 192 Booker, M. J. 42, 217 Borbely, 99, 199 Borch, G., 58, 200 Borchert, M . T., 88, 90, 217 Borde, K. 259, 275 Boresch, K., 166, 192 Borisov, A. Y . N., 105, 135,192,201 Bortels, H., 229, 270 Borys, M . W., 222, 227, 270 Boucher, F., 60, 62, 192 Boucher, L. J., 63, 102, 194, 203 Bouges-Bocquet, B., 18, 91, 92, 137, 192 Bourdu, R., 116, 192 Bowen, C. C., 109, 217 Bowen, J. E., 239, 270 Bowman, T., 244, 276 Boxer, S. G., 83, 192 Braslavsky, S. E., 174, 194 Braun, B. Z., 99, 207 Brehamet, L., 103, 206 Breton, J., 60, 61, 72, 85, 105, 132, 137, 192,200 Briggs, W. R., 29, 172, 192 Brinkman, G., 90, 172, 173, 192 Britton, G., 59, 108, 186, 210, 218 Britz, S. J., 29, 192 Brockman, H., 50, I92 Broda, E., 183, 192 Brody, M . , 74, 115, 117, 156, 192 Brody, S. S., 74, 115, 117, 192 Bromfield, S. M., 245, 246, 247, 270 Brooks, C., 66, 192 Brown, A. P., 47, 192 Brown, A. S., 68, 179, 192, 216 Brown, D. M., 133, 134, 198 Brown, J. S., 38,52,76,84,96,97,99, 100, 101, 102, 122, 144. 160, 192, 193, 197, 207 Brown, M . S., 60, 193 Brown, R. G.. 118, 210 Brown, S. B.. 63, 179, 183, 193, 203, 216 Brown, T. E., 156, 157, 193 Brown-Mason, A. S., 66, 181, 209, 216 Bruckensteins, S., 66, 208 Brunori, M., 84, 190

Bryant, D. A., 65, 66, 109, 110, 1 1 1, 112, 1 14, 134, 166, 193, 198, 205 Bryant, F. D., 47, 205 Buchert, J., 134, 195 Buck, K. W., 252, 274 Budzikiewicz, A., 53, 193 Buick, R., 117, 175, 217 Bukhov, N. G., 99, 174, 191 Burke, J. J., 51, 85,86, 103, 147, 208,214 Burne, R. N., 181, 190 Burrell, J. W. K., 51, 193 Burris, J., 170, 193 Burris, J. E., 164, 193, 198 Bussler, W., 230, 233, 236, 270, 274 Butler, E. J., 240, 255, 270 Butler, W. L., 39,43,62,69,70,72,73,88, 92,94,97, 114, 115, 117, 145, 146, 156, 161, 167, 171, 172, 193, 194, 205, 209, 212 Butterworth, B. F., 259, 273 Byfield, P. G. H . , 66, 193 Bykhovsky, V. Y . . 180, 193 C Catkins, J. 12, 214 Callihan, R. H., 244, 246, 247, 271 Calvert, H. E., 118, 210 Calvin, M . , 50, 51, 196 Camm, B. R., 89, 104, 107, 199 Camm, E. L., 88, 193 Cammack, R., 87, 176, 186, 193, I96 Canaani, D. D., 99, 193 Canaani, O., 68, 112, 114, 115, 133, 193, 197 Canuto, V. M., 178, 193 Carey, P. R., 61, 212 Caron, L., 100, 126, 191 Carr, A. J. H . 270 Carr, N. G., 87, 196 Carver, J. H., 175, 193 Casadevall, E., 204 Cassie, R. M . , 170, 195 Castets. A . M., 193 Castorinis, A., 105, 190 Castorinis, 102 Cavalier-Smith, T., 183, 193 Cess, R. D., 6, 175, 210 Chalker, B., 152, 193 Chambers, L. A,, 181, 190 Chambers, S. G., 252, 270

280

AUTHOR INDEX

Chandler, W. A., 258, 271 Chant, S. I., 103, 195 Chapman, 64 Chapman, A. R. O., 42, 193 Chapman, D. J., 54, 55, 63, 66, 177, 178, 192, 193, 206, 211 Charles-Edwards, D. A., 152, 193 Chaudhry, F. M., 238, 254, 270 Chen, G. C., 60, 193 Chen, K. N., 175, 183, 184, 186, 197 Chepelenko, A. P., 243, 274 Cherewick, W. J., 240, 257, 270 Cheshire, M. V., 254, 271 Chorney, W., 241, 276 Cho, F., 50, 51, 73, 115, 193, 194 Chow, H . C., 50, 51, 194 Chow, W . S., 146, 147, 148, 194, 213 Chrissovergis, F., 100, 126, 191 Christiansen, N. W . , 265, 270 Christoffersen, R. E., 83, 210 Chrometzka, P. 259, 276 Chua. N. H.. 78. 80. 85. 86. 89. 90. 91. 104, i 0 6 , b r , '194,195 Chunaev, A. S., 84,104 Chung, H., 235, 270 Clark-Myron, J., 62, 204 Clarkson, D. T., 255, 263, 270 Claudius, G. R., 258, 273 Clayton, R. K., 174, 209 Clement-Metral, J. D., 114, 194 Clezy, P. S., 53, 194 Closs, G. L., 83, 192 Cloud, P.E., 175, 176, 182, 194 Cobb, A. H., 54, 191 Codd, G. A., 117,200 Codgell, R. J., 103, 194, 206 Cogdell, R. J., 60, 61, 93, 194 Cogley, J. G., 175, 200 Cohen-Bazire, G., 41, 65, 70, 1 D, 1 1, 112, 114, 115, 116, 117, 138, 139, 142, 166, 193, 198, 205, 215 Cole, H., 257, 259, 273 Cole, W. J., 63, 193 Colquhoun, T. T., 243, 271 Conjeoud, H., 91, 194, 211 Conner, S. D., 249, 254, 270 Connolly, J. S., 50, 201 Conti, S. F., 31, 109, 197, 201 Coppenet, M., 244,272 Cook, R. J., 231, 243,249,251,253, 254, 255, 256, 258, 270, 274, 275 Coombs,J., 31, 32, 194

Cooper, R. A., 169, 213 Cortel-Breeman, A., 8 , 16, 217 Cotton, T. M., 83, 213 Couberg, S. R., 52, 199 Coute, A., 162, 163, 205 Cowling, E. B., 227, 236, 272 Cox, C. S., 8, 194 Cox,G. C., 32,33,162,163,186,188,194 Crabbe, P., 51, 194 Crafton, G., 64, 65, 206 Cramer, W. A., 92, 194 Crane, F. L., 41, 199 Crespi, H. L., 63, 100, 194, 198 Critchley, C., 164, 194 Croasmun, W. R., 103, 195 Crofts, A. R., 93, 186, 194, 212 Crooks, P. 244, 246, 271 Crossett, R. N . , 22, 23, 37, 167, 168, 170, 194, 205 Crouch, R. L., 61, 191 Croze, E., 92, 201 Cruickshank, I..A. M., 261, 274 Csatorday, K., 62, 64, 65, 206, 215 Curl, H . 69, 201 Czerwinski, E. W., 83, 218

D Dabek, A. J., 259, 271 Dale, R. E., 113, 133, 134, 194, 215 Dallinger, R. F., 60, 194 Daniel, J . W., 258, 271 Danilova, T. A., 241, 276 Darling, H. M., 243, 244, 246, 247, 274 Das, P. K., 63, 211 Davey, C. B., 227, 271 Davidov, A. S., 131, 194 Davidson, R. M., 228, 271 Davies, J. E., 66, 192 Davies, W. J., 253, 271 Davis, J. R.,244, 246, 247, 271 Davis, M. S., 83, 87, 93, 194, 197 Dawes, C. J., 30, 194 Day, D., 101, 197 Dayhoff, M. O., 175, 184, 194 Dayton, P. K., 22, 170, 194 Deamer, D. W . , 30, 210 Dear, J. M.,239, 241, 271 DeKock, P. C., 254, 271 De Kok, J., 174, 194 Delepelaire, P., 80, 89, 90, 100, 104, 194, 195

28 1

AUTHOR INDEX

Delieu, T.. 69, I95 Dellow, V., 170, 195 Demeter, S., 99, 199 Diakoff, S.. 174, I95 Dietrich, H., 60, 195 Dietrich, W. E., 77, 79, 195 Dilworth, M. F., 1 1 1, 195 Dimond, A. E., 255, 276 Diner, B. A.. 37,39,72,80,89,91,94, 115, 137, 142, 143, 191, 195 Dirks, G., 61, 191, 195, 208 Dirks, G. A. L., 61, 195 Dixon, N. E., 261, 271 Djerassi. C., 51, 194 Dodge, J. D., 120, 184, 185, 195 Dohnt, G., 92, 211 Dolan, E., 92, 93. 94, 118, 203, 204, 210, 213 Dolzmann, H., 259, 275 Doolittle, W. F., 41, 183, 192. 205 Dornemann, D., 49, 83, 195 Douce, R., 107, 195 Dougherty, R. C., 52, 53, 63, 102, 194, 195, 203 Draoli, R., 264,275 Doukas, A. G., 134, 195 Drew, E. A,, 8, 22, 23, 28, 37, 150, 167, 168, 170, 194, 195, 202, 205 Drew, M. C., 263, 271 Drikas, G., 60, 195 Dring, M. J., 14, 15, 23, 167, 168, 169, 195, 206 Droop, M . R., 162, 195 Dubacq, J.-P., 104, 106, 107,216 Dubertret, G., 95, 137, 139, 146, 195 Dubinsky, Z., 46, 151, 195, 196 Dufrenoy, J., 259, 271 Duggar, B. M . , 72, 195 Duncan, M. J., 173, 195 Duniec, J. T., 48, 99, 146, 147, 148, 194, 195, 213, 216 Dunlop, J. S. R., 117, 175, 195, 217 Dunstan, W . M . , 151, 195 Duranton, J . , 86, 99, 106, 204, 213 Dutta, T . R., 242, 271 Dutton, H. J . , 72, 195 Duval, J. C., 70, 73, 100, 109, 118, 121, 126, 128, 191, 205 Duysens, L. N. M., 14,45,46, 72,93, 1 13, 117, 161, 189, 195, 210, 214, 216 Dwarte, D. M., 32, 33, 39, 40, 129, 139, 140, 162, 163, 186, 188, 194, 195

Dyer, T. A,, 175, 183, 184, 186, 197 Dzelzkolus, V., 183, 190

E Eaton, F. M., 240, 271 Ebling, F. J., 23, 162, 170, 209 Edelman, M., 91, 207 Edelstein, M. S., 65, 133, 202 Edwards, M. R., 35, 197 Eichmann, R., 175, 182, 212 Eigenberg, K. E., 103, 195 Eisenbraun, E. J., 51, 194 Eiserling, F. A., 65, 193 El-Sayed, M. A,, 176, 195 Emerson, R., 14,38,73, 74, 153, 156, 192, 195, 196 Engel, W., 262, 271 Engelmann, Th. W . , 23,68, 165, 167, 196 England, R. R., 82, 89, 196 Epstein, E., 228, 253, 257, 271 Erhardt, M. M . , 66, 181, 216 Erwin, D. C., 258, 273 Etienne, A-L., 72, 100, 205 Etzion-Katz, 147, 202 Eubanks, D. C., 175, 206 Evans, E. H . , 79,82,87,89, 118,196,210 Evans, E. M., 225, 271 Evans, M. C . W., 87, 193, 196, 209 Evstigneev, V . B., 93, 98, 196, 213

F Fajer, J., 83, 87, 93, 194, 197 Falkowski, P. G., 22, 38, 94, 122, 144, 151, 154, 156, 157, 158, 160, 162. 196 Faludi-Daniel, A,, 47, 99, 140, 191, 199 Fang, H. L-B., 61, 198, 216 Fang, S.,'133, 134, 198 Farquhar, G. D., 31, 40, 152, 196 Faust, M. A., 35, 157, 196 Fee, E. J . , 152, 196 Feigina, M. Yu.,60, 210 Felix, E. L., 229, 271 Felton, R. H., 83, 196 Fenna, R. E., 51, 84, 103, 132, 196, 207 Ferguson, I . B., 255, 263, 270 Ferguson, J., 181, 190 Finn, B. J., 245, 273 Fischer, H., 49, 63, 196

282

AUTHOR INDEX

Fischer, M. S., 50, 51, 196 Fisher, R. G., 67, 68, 132, 133, 134, 196 Flaig, W . , 254, 266, 271 Fleischer, W . E., 41, 196 Fleischfresser, M. H., 243, 244, 274 Fleischhacker, P. H., 52, 95, 151, 153, 156, 158, 160, 196, 213 Fleming, I., 50 Floyd, G. L., 188, 196 Foley, T., 183, 190 Fookes, C. J . R., 53, 194 Forbush, B., 37, 136, 137, 204 Foreman, R. E., 173, 195 Forman, A., 83, 87, 194 Fork, D. C., 47, 69, 70, 71, 72, 101, 115, 144, 154, 160, 196, 200, 206 Forrester, R. I., 255, 257, 272 Forsyth, F. E., 255, 271 Forsyth, F . R., 260, 261, 271 Forti, G., 40, 104, 147, 198, 202 Forward, R. B., Jr, 42, 197 Forward, R. B., 42, 197 Foster, J . A,, 68, 192 Foster, Th. W . , 131, 196 Fox, G. E. 175, 183, 184, 186, 197, 206 Fox, J . L., 66, 68, 198, 199 Franzblau, C., 68, 192 Frank, F., 66, 197 Freidenreich, P., 66, 197 Freidlin, M. I., 152, 216 French, C. S., 52, 72, 96, 157, 192, 197, I99 Frenkel, A. W., 31, 200 Friedman, A. L., 126, 189 Friend, J., 233, 271 Fries, E., 104, 200 Friedrich, J., 67, 218 Frohlich, H. P., 260, 272 Fuad, N., 101, 197 Fuchs, H. E., 67, 68, 132, 133, 134, 196 Fujimori, E., 133, 210 Fujita, I., 93, 197 Fujita, Y.,38,42,72,84,93,94, 115, 144, 156, 160, 161, 173, 174, 197, 203, 207, 209, 213 Fukumoto, J. M., 176, 195 Fuller, R. C., 17, 31, 197, 201 Fuminori, 87 Furtado, D., 166, 218

G

Gaffron, H., 74, 197 Gaidukov, N . I., 23, 165, 166, 167, 197 Gantt, E., 35, 63, 64,67, 68, 84, 85, 109, 110, 111, 112, 113, 114, 115, 116, 133, 139, 157, 166, 184, 193, 195, 196, 197, 199, 203, 206, 211, 219 Garab, G. I., 47, 140, 191 Gardner, E. E., 66, 68, 198 Garlaschi, F. M., 40, 104, 147, 198, 202 Gamier, J., 89, 207 Garrels, R. M., 175, 198 Garrett, S. D., 226, 271 Gasner, E., 164, 204 Gast, P., 93, 94, 215 Gates, D. M., 5, 6, 7, 24, 198 Gauch, H. B., 239, 270 Gavrilova, V . A., 93, 196 Gazzola, C., 261, 271 Geacintov, N . E., 61, 72, 132, 137, 192 Geen, G. H., 21 7 Gendel, S., 174, 209 Genge, S., 98, 200 Gerber, D. W . , 164, 198 Gering, R. K., 226, 275 Germar, B., 262, 271 Gerola, P. D., 40, 104, 147, 198, 202 Ghosh, A. K., 100, 171, 198 Ghure, T . K., 260, 270 Gibadullin, M. G., 260, 274 Gibbons, O., 68,206 Gibbs, S. P., 31, 183, 185, 186, 198 Gibson, J., 175, 183, 184, 186, 197 Giddings, T . H., 32, 116, 138, 139, 142, 156, 186, 188, 198, 214 Gill, D. L., 228, 271 Gillan. F. T., 186, 202 Giller, Y.E., 84, 194 Gillott, M. A., 185, 198 Gilmartin, M.,23, 198 Gilmour, J . 244, 245, 246, 271, 274 Gingras, G., 60, 62, 77, 192, 198 Gingrich, J . C., 110, 198 Giri, D. N., 261, 275 Glassick, C. E., 260, 272 Glazer, A. N., 41, 63, 64, 65, 66, 67, 68, 85, 109, 110, 111, 112, 114, 133, 134, 161, 181, 184, 193, 197, 198, 204, 205, 206, 218 Glick, R. E., 110, 218 Glidewell, S. M., 21, 188, 211

283

AUTHOR INDEX

Glynne, M. D., 225, 271 Goedheer, J . C., 59,60,61, 72,73,74,95, 100, 101, 102, 120, 198 Golbeck, J. H., 17, 198 Goldstein, J. L., 60, 193 Goodchild, D., 37, 39, 84, 100, 102, 144, 145, 146, 191 Goodchild, D. J., 29, 44, 45, 46, 47, 52, 105, 107, 138, 139, 140, 147, 158, 160, 189, 191, 200, 203, 212 Goodwin, T. W., 54,55,57, 108, 186,189, 198, 206, 218 Goss, R. L., 222, 228, 271 Gossaver, A., 63, 198 Gott, 72, 204 Gouterman, M., 95, 199 Govindjee, 73, 74, 94, 100, 112, 115, 171, 193, 194, 198, 199, 207, 216 Govindjee, R., 72, 74, 99, 112, 199 Grabowsi, J., 67, 109, 110, 112, 113, 114, 133, 197, 199 Graham, J. 171, 208 Graham, J-R., 39,69,73,74,84,94, 144, 146, 160, 171, 172, 208, 217 Graham, R. D., 226,230,233,238,243, 249,253,254,258,264,241 Grandolfo, M. C., 58, 118, 214 Granick, S., 53, 176, 199 Grant, B. R., 52, 203 Graves, C. J., 236, 270 Greef, J. A., 52, 199 Green, B. R.,88, 193, 199 Green, E. L., 89, 104, 107, 199 Greenwood, A. D., 31, 32, 186, 194, 199 Grefarth, S. P., 85, 199 Gregory, R. P. 99, 213 Gregory, R . P. F., 99, 199 Greis, G. A,, 265, 271 Griffin, D. C., 86, 199 Griffiths, H. B., 186, 199 Grime, J. P., 21, 199 Grimme, L. H., 42, 52, 59, 76, 183, 199, 210 Gross, E. L., 90, 205 Gross, G. G., 241,247,248,255,266,272 Grosse-Brauckmann, E., 262, 272 Grouzis, 199 Groves, D. I., 175, 195 Gudauskas, R. T., 225, 271 Guerin-Dumartrait, E., 116, 142, 156, 199 Gugliemi, G., 193

Gulman, S. L., 21, 207 Gulyaev, B. A,, 99, 199 Gunning, B. E. S., 20, 190 Guntz, M., 244, 272 Gupta, R., 175, 184, 186, 197 Gurinovich, G. P., 95, 99, 199 Gust, D., 61, 191, 195, 208 Gustafson, D. L., 126, 189 Gysi, J., 66, 114, 199, 213

H Hackert, M . L., 68, 199 Hackett, C., 222, 272 Hager, A., 55, 62, 199, 214 Hagemann, R., 92, 211 Hainfield, J. F., 110, 211 Haidak, D. J. C., 118, 199 Hall, A., 254, 271 Hall, D. O., 176, 186, 193 Hall, J. D., 41, 199 Halldal, P., 12, 157, 162, 163, 166, 199 Halldol, 70 Hallock, D. L., 231, 272 Halsall, D. M., 255, 257, 265, 272 Halstead, R. L., 245, 273 Hancock, J. G., 256, 275 Hapase, D. G., 260, 270 Hardt, H., 91, 200 Harel, E., 248, 273 Hargrave, B., 24, 40,152, 210 Harkin, J. M., 248, 272 Harnischfeger, G., 99, 117, 200 Harper, J. L., 170, 200 Harris, G. P., 12, 152, 162, 164, 165,200 Hars, R., 30, 216 Hartley, R. D., 236, 274 Harvey, G. W., 160, 164, 200, 207 Hase, E., 41, 173, 200, 209 Hase, T., 187, 215 Haselkorn, R., 41, 218 Hastings, J. W., 118, 192 Hatch, M. D., 150, 182, 200 Hattori, A,, 173, 197 Haupt, W., 30, 200 Havry, J. F., 174, 200 Hauska, G., 38, 39, 88, 201, 218 Haworth, P., 85, 105, 200 Haxo, F. T., 54, 55, 58, 69, 70, 72, 112, 114, 115, 118, 119, 120, 121, 135, 136, 157, 193, 200, 202, 206, 210, 214

AUTHOR INDEX

Hay, M. E., 28, 170,200 Hayden, D. B., 88,200 Hayes, J . D., 230,272 Haynes, J . L., 259, 272 Hazen, E. E., Jr, 133, 202 Healy, F. B., 150, 200 Heard, T. G., 252, 254, 274 Heath, M. C., 262, 263, 272 Heber, U. W., 31, 204 Heggeness, H. G., 240, 272 Heinz, E., 105, 106, 200 Heinz, R. P., 63, 198 Helenius, A., 104, 200 Helms, K., 259, 272 Hemming, B. C., 255, 256, 272 Henderson, R., 132, 200 Henderson-Sellers, A., 175, 200 Henriques, F., 88, 91, 102, 200 Herman, E. M., 29, 200 Herron, H. A., 150, 200 Hertzberg, S., 58, 200 Hervo, G., 61, 103, 206 Hespell, R. B., 175, 183, 184, 186, 197 Hess, W. M.,243,270 Hewitt, E. J., 257, 272 Hickman, D. D., 31, 200 Highkin, H. R., 105, 215 Hill, R., 17, 18, 200 Hiller, R. G., 37, 84, 85, 89,98, 102, 105, 118, 121, 200, 201 Hilton, R. N., 257, 258, 270 Hinckle, P. C., 19, 200 Hindman, J. C., 100, 218 Hipkins, F. M . , 60 Hirano, M., 63, 189 Hixon, C. S., 66, 193 Hjortas, J., 58, 200 Hoarau, J., 81,83,87,98, 162, 163,200, 201,205,211 Hoarau, R., 87,98,200 Hoard, J. L., 50, 201 Hoch, G., 150, 201 Hock, G. E., 101, 196 Hoff, A. J., 83, 93, 94, 103, 201 Hoff, H. J., 93, 206, 210, 215 Hofman, H. J., 181, 190 Hoffman-folk, H., 91, 207 Holden, M.,52,201 Holdsworth, E. S., 123, 201 Holmes-Siedle, A. G., 248, 275 Holm-Hansen, O., 12, 164, 214 Holroyd, A. J., 179, 193

Holroyd, J. A., 63, 193 Holt, A. S., 53, 83, 201, 217 Holt, S. C., 31, 53, 201 Holt, T. K., 60, 201 Holtz, S., 259, 275 Homann, P. H., 91, 92, 207 Hong, F. H., 50,207 Hootkins, R., 87,201 Hopf, 50 Hopkins, D. W., 97, 193 Hopkins, W. G., 88, 200 Hoppe, J., 39,40, 213 Horsfall, J. G., 227, 236, 272 Horst, W. J., 262, 272 Horton, C., 147, 201 Horton, P., 92, 107, 201 Horvath, G., 47, 48, 140, 191, 216 Horvath, L. I., 61, 215 Hovlier, B., 87, 201 Howard, K. L., 167, 169, 211 Hoyer-Hansen, G., 90, 206 Huang, P. M., 248, 272, 275 Huang, Y.T., 261, 272 Huber, D. M., 222, 225, 265, 272 Hudson, M.F., 178, 201 Humphrey, G. F., 32, 53, 149, 169, 172, 201 Hunt, P., 259, 271 Hunter, F. A., 97, 105, 146, 21s Hursthouse, M. B., 66, 192 Hurt, E., 38, 39, 201

I Ikana, T., 54, 59, 218 Ikawa, T., 37, 52, 59, 72, 108, 109, 162, 202 Ikegami, I., 77, 201 Ilani, A., 62, 87, 201, 204, 207 Ilina, M.D., 105, 135, 192, 201 Imhoff, C. L., 178, 193 Ingram, K., 121, 201 Interschick-Niebler, E., 77, 85, 201 Isaeva, G. Y.,260, 272 Ishikawa, C., 103, 106, 208 Ishizaki, H., 262, 273 Isker, E., 66, 213 Isler, O., 57, 201 Isoni, T., 174, 209 Ito, R., 261, 273

AUTHOR INDEX

Itoh, S., 87, 215 Iverson, R. L., 69, 201 Iwata, I., 262, 270

J Jackman, L. M., 51, 193 Jackson, T. L., 265, 270, 274 Jacobs, D. L., 255, 256, 272 Jamieson, G. R., 128, 201 Janzen, A. F., 50, 201 Jarvis, S. C., 236, 254, 272, 274 Jassby, A. D., 150, 152, 201 Jeffrey, S. W., 51, 52, 53, 57, 58, 59, 120, 121, 172, 201, 202, 205, 216 Jenkyn, J. F., 222, 226, 230, 272 Jennings, R. C., 40, 104, 147, 198, 202 Jensen, A., 10, 55, 202, 213 Jensen, N-H., 61, 202 Jensen, R. G., 40, 95, 149, 202, 210, 213 Jerlov, N. G., 9, 10, 11, 13, 16, 168, 202 Johansen, J. E., 202 Johansson, L., B-A., 61, 202 John, P. 183, 186, 217 John, P. C. L., 20, 190 Johns, R. B., 186, 202 Johnson, C. E., 79, 87, 196 Joliot, A., 69, 72, 74, 92. 136, 137. 202 Joliot, P., 37-39, 69, 72, 74, 92, 136, 137, 195, 202 Jones, A. P., 247, 272 Jones, E. W., 257, 272 Jones, G. P. D., 258, 272 Jones, I. T., 230, 272 Jones, L. W., 69,70,72,73, 115, 164, 170, 202 Jones, 0. T. G., 104, 212 Jones, S. G., 240, 255, 270 Jorgensen, E. G . , 158, 202 Joset-Espardellier, F., 92, 190 Joyard, J., 107, 195 Jung, J., 13, 202 Junge, C. E., 175, 182, 212 Junge, J., 65 Junge, W., 17, 19, 37, 38, 39, 60, 61, 83, 135, 144, 202 Junge, 97, 202 Jupin, H., 70, 73, 97, 100, 109, 118, 121, 126, 191, 205 Jupp, B. P., 22, 28, 202

K Kageyama, A., 37, 52, 54, 59, 72, 108, 109, 162, 202, 218 Kain, J. A., 22, 28, 169, 202 Kahn, A., 48, 216 Kahn, R. P., 261, 262, 276 Kalyanasundaram, K., 87, 202 Kalle, K., 10, 11, 202 Kan, K-S., 97, 105, 106, 146, 202, 215 Kane, J. P., 60, 193 Kanematsu, S., 176, 209 Kanwisher, J. W., 25, 203 Kao, H., 203 Kaplan, A., 41, 190, 203 Karapetyn, N. V., 99, 174, 191 Karvaly, B., 176, 195 Kassner, R. J., 83, 203 Katoh, S., 76, 77, 79,81, 89,90, 104, 105, 115, 201, 203, 208, 218 Katoh, T., 109, 203 Katschan, R.. 63, 198 Katz, J. J., 50, 51, 52, 53, 58, 63, 83, 96, 100, 102, 118, 190, 194, 195, 198, 203, 212, 213, 214, 218 Kauer, J. C., 259,273 Kaur, P., 243, 272 Kayr, S., 243,272 Kawamura, M., 38, 144, 156, 160, 203 Kay, I. T., 66, 208 Ke, B., 92, 93, 94, 102, 203, 204, 213 Keane, E, M., 240, 272 Keast, J. F., 52, 203 Keeler, R. R., 244, 272 Keil, H. L., 260, 272 Kelly, P., 183, 216 Kent, N. L., 264, 272 Kent, S. S., 203 Kerfin, W., 176, 182, 203 Kern, H., 255, 265, 272 Kessler, E., 182, 203 Kiefer, D. A., 47, 48, 203 King, R. F. G., 179, 203 King, R. J., 170, 203 Kirby, E. A., 227, 240, 241, 259, 273 Kirk, J. T. O., 9, 1 I , 13, 14,24,25,29, 32, 35, 44, 45, 46, 47, 124, 169, 203 Kirst, G. O., 203 Kiselev, A. V., 60, 210 Kitching, J., 169, 203 Kitching, J. A,, 23, 162, 170, 209 Kittaka, A., 262, 270

286

AUTHOR INDEX

Kjosen, H., 58, 118, 214 Klein, C., 174, 175, 177, 217 Klein, S., 203 Klein, S. M., 79, 203 Kleinen-Hammans, J. W., 163, 216 Klevanik, A. V., 93, 203 Klimov, V. V., 92, 93, 203, 204 Kloepper, J. W., 255, 256, 272 Kloppstech, 173, 190 Knaff, D. B., 62, 92, 204 Knox, R. S., 43, 131, 132, 204 Kobayashi, 133 Kochel, H. G., 183, 204 Koenig, F., 81, 90, 204 Kok, B., 17, 20, 37, 72, 74, 98, 136, 137, 149, 150, 152, 153, 164, 201, 202, 203 Koka, P., 61, 120, 135, 136, 204, 214 Koller, K-P., 68, 109, 110, 111, 112, 133, 134, 204,208 Kond, F., 262, 273 Korant, B. D., 259, 273 Korol’chuk, V. V., 231, 274 Kost, H-P., 35, 67, 11 1, 204, 21 7 Kostka, A. G., 83, 217 Kott, P., 108, 186, 204 Kouchkovksy, de-Y., 90,214 Kraft, G., 169, 170, 207 Krakover, T., 87, 204 Kramer, D., 256, 266, 273 Krasnovskii, A. A., 93, 203 Kratz, W. A., 63, 156, 208 Krause, G. H., 31, 204 Krauss, A., 222, 273 Krauss, R. W . , 52, 211 Krawczyk, S., 99, 204 Kretzer, F., 204 Kreutzer, K., 243, 276 Krieger, M., 60, 193 Krol, M., 41, 190 Krogmann, D. W., 17, 60, 201, 204 Kroneck, P., 186, 192 Krueger, W. C., 66, 208 Krupa, Z., 41, 190 Krupnikova, T. A., 260, 275 Kryzymowski, W. W., 116,138,139, 142, 156, 214 Kuan, T. L., 258, 273 Kubota, J., 254, 273 Kuenen, J. G., 31, 191 Kufer, W., 67, 212 Kulandaivelu, G., 62, 204 Kullenberg, G., 13, 204

Kunert, K-J., 88, 204 Kunoh, H., 262, 273 Kuntel, H., 183, 204 Kursar, 181 Kvitko, K. V., 84, 194 1

Lacourly, A., 116, 199 Ladeishchikova, E. I., 240, 274 Ladnykh, L. F., 240, 274 Laetsch, W. M., 187, 204 Lagarias, J. C., 66, 183, 204 Lagoutte, B., 86, 99, 106, 204, 213 Lakshminarayanan, K., 243, 273 Land, E. J., 61, 191 Landon, M., 86, 199 Landsberg, E., 256, 266, 273 Lang, J. C., 204 Langer, E., 67, 68, 205 Langridge, J., 175, 205 Largeav, C., 204 Larkum, A. W . D., 22,23, 37,69, 70, 71, 73,8589, 101, 104, 105, 110, 115, 117, 118, 146, 167, 168, 169, 170, 194, 200, 205, 206 Last, F. T., 223, 224, 273 Laszlo, J. A., 90, 205 Latimer, P., 47, 48, 205 Lav, R. H., 41, 205 Lavorel, J., 72, 100, 205 Laycock, M. V., 183, 205 Lee, C. R., 260, 273 Lee, B. D., 121, 167, 205, 216 Lee, J . A., 48, 99, 146, 216 Lee, S. G., 239, 241, 273 Lee, R. E., 186, 205 Leeper, G. W . , 243, 244, 245, 246, 249, 254, 256, 273 Leclerc, J. C., 98, 142, 156, 162, 163, 199, 200, 205 Lefort, M., 116, 192 Lefort-Tran, M., 95, 114, 116, 117, 139, 142, 146,192,194,195,205 Lehner, M., 67, 68, 205 Lemasson, C., 41, 70, 205 Lemmasson, C., 115, 205 Lemberg, M. R., 50, 63, 178, 205 I,emberg, R., 63, 84, 205 Lemons, F., 121, 211 Leong, J., 255, 256, 272, 273

AUTHOR INDEX

Lerman, A , , 175, 198 Leroi, G. E., 61, 216 Leussing, D. L., 102, 203 Levine, R. P., 78, 86, 106, 189 Levine. J. S., 178, I93 Levring, T., 23, 167, 170, 205 Lewin, J., 58, 205 Lewin, R. A , , 5, 32, 186,205 Lewis, B. J., 175, 183, 184, 186, I97 Lewis, C. M.. 14, 73, 74, I96 Lewis, D. H., 241, 273 Lewis, N. B., 63, 215 Lewis, R. A,, 108, 186, 218 Ley, A . C., 38, 39, 69, 70, 72, 73, 94. 114, 115, 117, 118, 119, 144, 146, 154, 156, 157, 160, 161, 162, 167, 171, 196. 205, 210 Lai, S.,51, 194 Liaaen-Jensen, S., 58, 118, 190, 200, 202, 214 Lichenthaler, H. K., 77, 85, 201 Lichtenthaler, A. K., 52, 205 Lichtle, C.,70,73, 109, 116, 118, 121, 142, 157, 205 Lien, S., 17, 198 Lilley, R. M. C., 110, 206 Lindberg-Moller, B., 104, 206 Lindblom, G., 61, 202 Lipkind, F. M., 84, 194 Lipman, C . B., 229, 273 Lipschultz, C. A., 63, 109, 110, 1 1 I , 112, 114, 115, 193, 197, 205 Littler, D. S., 28, 170, 206 Littler, M. M., 28, 170, 206 Lobanov, A,, 60, 210 Lodeman, E. G., 229, 273 Lorenzen, C. J., 12, 13, 206 Loring, H. S., 255, 273 Loneragan, J. F., 233,238,249,254,260, 270, 271, 273, 276 Loughman, B. C., 241, 274 Lowig, E., 262, 273 Love, F. G., 162, 181, 210 Lucas, C.. 183, 206 Ludlow, C. J., 72, 206 Ludwig, L. J., 152, 193 Luehrsen, K. R.. 175, 183, 184. 197, 205 Lundell, D. J., I 1 I , 112, 205 Lundqvist, 99, I89 Lungley, D. R., 222, 273 Luning, K., 14, 15, 23, 28, 42, 169, 193, 205

287

Lutz, M., 61, 93, 103, 206 Lyman, H., 126, 189

M MacColl. R., 64, 65, 68. 113, 114, 118, 179, 181, 206 MacDonald, W., 60, 61, 194 MacGillavry, C. H., 58, 190 Machold, O., 90, 104, 107, 108, 206 Mackenzie, M . M., 41, 205 MacKinney, G., 206, 229, 273 Maggiova, G. M., 83, 210 Magrum, L. J.. 175, 183, 184, 197 Mahler, H. R., 183, 211 Malkin, R., 39, 62, 87, 160, 201, 204, 206 Malkin, S., 47, 144, 154, 206 Mallams, A. K., 55, 58, 66, 192, 206 Malone, T. C., 24, 206 Mandelli, E. F., 58, 206 Mangel, M., 62, 207 Manikowski, H., 87, 206 Maniloff, J . , 175, 183, 184, 186, 197 Mann, J. E., 69, 70, 207 Mann, K. H., 23, 25, 123, 207 Manning, W. M., 72, 195 Marchant, H. J., 162, 194 Marco, J., 89, 207 Margulis, L., 4, 121, 181, 183, 207, 211, 217 Markham, J. W., 42, I93 Markum, K. R., 215 Markwell, J. P., 62,76, 102, 104, 107, 120, 122, 147, 191, 192, 207, 215 Marra, J., 162, 207 Marschner, H., 256, 262, 266, 272, 273 Mase, T., 186, 207 Mathews, B. W., 51, 84, 103, 132, 196, 20 7 Mathews, G. K., 118, 199 Mathews Scott, F., 83, 218 Mathis, P., 49, 91, 92, 194, 207, 211, 216 Matlick, H. A., 152, 210 M a t h , K., 78, 85, 86, 106, 194 Matsubara, H., 186, 187, 207, 215 Matsubayashi, M.. 261, 273 Matsuka, M., 41, 209 Matsuyama, N., 251, 273 Mattoo, A. K., 91, 207 Mattox, K. R., 5, 26, 188, 214

288

AUTHOR INDEX

Mauzerall, D., 50, 87, 94, 144, 149, 160, 167, 169, 175, 176, 195, 200, 201, 204, 207, 211 Mauzerall, D. C., 38, 88, 144, 154, 160, 162, 177, 196, 207 Mayer, A. M., 248, 273 Mehrotra, R. S., 258, 273 Meister, 107, 206 Melis, A., 38, 91, 92, 144, 160, 207 Mel’nikov, S. S., 52, 207 Mengel, K., 227, 240, 241, 259, 265, 273 Menke, W., 18, 31, 59, 90, 204, 207 Menzel, D. W . , 164, 212 Merinova, G. L., 12, 191 Merkle, H., 186, 192 Metz, J., 207 Meyer, H., 257, 263, 264, 273 Michel, J. M., 96, 197 Michel-Wolwertz, M. R., 96, 197 Miles, L. E., 225, 273 Millar, A., 169, 170, 207 Miller, K. R., 40, 138, 139, 187, 207 Milne, V. A., 175, 195 Mimuro, M., 38,72,84,94,115,144, 156, 160, 203, 207 Mirzaev, M. N., 243, 274 Mishkind, M., 144, 160, 207 Mitchell, P., 19, 207 Mitsu, S., 261, 273 Miyachi, S., 183, 209 Moaru, J., 116, 142, 156, 199 Moerdijk, A., 214 Moffatt, J. R., 226, 272 Mohanty, 99, 207 Mohr, H., 173, 207 Molinier, R., 22, 23, 207 Moller, B. L., 87, ,209 Mooney, H. A., 21,47,144,206,207,235, 2 74 Moore, 60 Moore, D. M., 47, 205 Moore, A. L., 61, 191, 207 Moore, T . A., 61, 191, 195, 208 Mora, R., 175, 214 Morel, A., 9, 10, 208 Morley, H. V., 53, 201 Morris, C. J., 226, 275 Morris, I., 40,190 Morschel, E.,68, 109, 110, 111, 112, 133, 134, 204, 208 Mortensen, T., 58, 200 Mortvedt, J . J., 243, 244, 246, 247, 274

Moscowitz, A., 66, 208 Moss, G. P., 58, 208 Mousseau, A., 78, 81, 85, 91, 215 Muckle, G., 63, 64, 66, 208 Mues, R., 215 Muir, M. D., 175, 195 Mullet, J. E., 51, 83, 85, 86, 92, 93, 103, 147, 208, 212 Murakami, S., 30, 32, 47, 208 Murata, N., 73, 97, 100, 101, 144, 146, 208, 215 Murata, T., 84, 103, 106, 208 Mustardy, L. A., 47, 140, 191 Murray, S. N., 28, 190, 206 Myers, A., 109, 208 Myers, J., 39,63,69,70,72,73,74,84,94, 109, 115, 144, 146, 150, 156, 160, 161, 162, 170, 171, 172, 192, 200, 202, 207, 208. 217 M C

McBride, M. B., 238, 273 McCaslin, D., 104, 200 McCaslin, D. R., 60, 206 McCartin, P. J., 83, 95, 206 McCarty, R., 19, 39, 40,206 McCarty, R. E., 19, 200 McClure, J. M., 242, 273 McCormick, A., 58, 66, 192 McDole, R. E., 244, 246, 247, 271 McFarlane, I., 241, 273 McGloin, M., 37, 136, 137, 204 McGregor, A. J., 243, 244, 273 McIlrath, W. J., 242, 271 McIlveen, W. D., 257, 259, 273 Mclntosh, A. R., 87, 206 McKay, A. J . M., 244, 246, 271 McKay, R., 245, 246, 273 McKie, J., 183, 206 McLaughlin, S. B., 236, 273 McLean, A. J., 245, 273 McNeal, B. L., 231, 243, 251, 254, 258, 2 74 McNew, G. L., 226, 273 McSwain, B. D., 190

N Nable, R. O., 252, 274

AUTHOR INDEX

Naef-Roth, S., 255, 265, 272 Nakamura, K., 52, 108, 208, 209 Nakatini, H. Y., 207 Nakayama, K., 76,79,81,89,90,105,208 Nambiar, E. K . S., 233, 238, 254, 271 Nandi, P., 255, 274 Nazarova, 1. P., 241, 276 Nelson, N., 39, 40, 60, 61, 202, 208 Nes, W. D., 178, 208 Nes, W. R., 178, 208 Nesterov, P. I., 231, 274 Neumann, J. J., 39, 208 Neushul, M., 116, 142, 209 Newcomb, E. H., 52, 108, 186, 209, 216 Newman, M. J., 177, 209 Newman, P. J., 79, 81, 90, 209 Nichols, P. E., 186, 202 Nicholson, D. E., 175, 206 Nielands, J. B., 255, 273 Nielsen, E. S., 9, 202 Nies, M., 64, 65, 209 Nikolaeva, N. F., 258, 274 Ninnemann, H., 54, 62, 214 Nishimura, M., 87, 100, 208, 215 Nobel, P. S., 20, 36. 209 Nomoto, T., 261, 273 Nordhorn, G . , 123, 209 Norgard, S., 58, 118, 214 Norman, G . D., 63, 194 Norris. A., 114, 200 Norris, J. R., 83, 96, 203, 213 Norris, R. E., 58, 102, 205 Norton, T. A., 23, 162, 170, 209 Nugent, J. H. A., 87, 209 Nutbeam, A. R., 166, 218 Nultsch, W., 30,36,74,192,209,210,21 I 0

Obata, F., 77, 209 Obst, J. R., 248, 272 O'Carra, P., 63, 65, 66, 209 O'Connor, G . , 64, 65, 206 Odum, E. P., 22, 25, 209 Offner, G. D., 63, 66, 181, 183, 193,209, 216

Ogawa, T., 52, 77, 101, 208, 209 Ohad, I., 78, 86, 104, 106, 174, 190, 204. 209

Oh-hama, T., 41, 183, 209 O'Heocha, C., 53,63,65,66,114,200,209

289

Ohki, K., 174, 209 O'Kelley, C. J., 59, 209 Okada, S., 176. 209 Okayama, S., 62, 209 Olive, J., 138, 146, 218 Olsen, C., 230, 274 Olsen, S. R., 256, 274 Olson, J . M . , 103, 177, 184, 207, 209 Olson, R., 12, 164, 214 Olson, R. J., 47, 48, 84, 104, 203 Oltmanns, F., 166, 167, 209 Oquist, G., 59.62, 101, 146, 191, J96,209 Orser, C., 255, 256, 272 Orth, H., 63, 196 Osmond, C. B., 52, 108, 164, 165. 186, 209, 210, 216

Oswald, J. W., 244, 276 Otake, N., 183, 209 Otto, J., 66, 208 Ovchinnikov, Yu. A , , 60, 210 Owen, T., 6, 175, 210 Owens, 0. H., 150, 158, 201 Owens, T. G . , 22, 38, 94, 122, 144, 151, 154, 156, 157, 160, 162, 196

Ozanne, P. G . , 254, 274

P Packer, L., 30, 32, 47, 208, 210 Padan, E., 182, 210 Padmanabhan, S. Y., 243, 272 Page, H. P., 52, 218 Pagsberg, P. B., 61, 202 Paillotin, G., 49, 92, 137, 207, 210 Palm, E. T., 264, 274 Panczyk, B., 41, 190 Papagiorgiou, G . , 72, 210 Papastylianou, I., 233,274 Papavizas, G. C., 227, 271 Park, R. B., 38, 72, 77, 88, 91, 102, 139, 200, 206, 210, 212

Parker, B. C., 162, 181, 210 Parker, L., 30 Pam, A. J., 241, 274 Parsons, T. R., 24, 40, 152, 210 Pashchenko, V. Z., 93, 203 Paterson, D. R., 93, 212 Patil, A. O., 260, 270 Paxton, R. J., 65, 133, 202 Pearson, B. E., 58, 205 Pecci, J., 133, 2fO

290

AUTHOR INDEX

Pelevin, V. N., 11, 210 Pellegrino, P., 67, 134, 218 Perchorowicz, J. T., 149, 210 Perov, N. N . , 243, 274 Perova, L. I., 243, 274 Perrin, D. R., 261, 274 Perry, G. J., 186, 202 Perry, M. J., 151, 158, 210 Peterson, B., 260, 261, 271 Peterson, R. B., 118, 210 Petke, J . D., 83, 210 Pfau, J., 30, 36, 192, 210, 211 Pfeifer, H., 259, 276 Phagpolngarm, S., 189 Phung Nhu Hung, S., 87, 201 Pianka, E. R., 22, 210 Piccinin, B. B., 164, 165, 200 Piccioni, R. G., 88, 207 Pick, U., 91, 207 Pietto, A. S., 17, 198 Pilger, T. B. G., 98, 200 Piper, G. S., 243, 249, 254, 268, 275 Pirkl, J., 259, 274 Pitt, D., 265, 274 Platt, T., 152, 201 Plurnbley, R. A., 265, 274 Pobegailo, A. I., 240, 274 Pollard, A. S., 241, 274 Porra, R. J., 42, 52, 183, 199, 210 Porter, D. M., 231, 272 Porter, G., 83,87, 112, 114, 133,191,202, 210, 213 Pound, G. S., 259, 272 Pouphile, M., 116, 117, 138, 139, 142,205 Powelson, R. L., 265, 270, 274 Powles, S . B., 164, 165, 210 Powls, S . R., 59, 210 Prager, L. K., 96, 197 Prasad, Y., 258, 274 Preston, R. D., 109, 208 Prezelin, B. B., 69,70,72,78,84,118,119, 120, 121, 122, 135, 136, 151, 152, 157, 158, 192, 210, 214 Prezelin, B. P., 59, 210 Primavesi, A., 231, 258, 274 Primavesi, A. M., 231, 258, 274 Prosser, M. V., 12, 180, 215 Provasoli, L., 35, 197 Prusa, V., 259, 274 Puckridge, D. W., 233, 274 Pugh, T. D., 186, 209 Pullin, C. A., 118, 210

Prievr, L., 10, 208

R Raaben, M. E., 181, 190 Rabinowitch, E., 44, 45, 46, 74, 166, 167, 168, 199, 210 Rabinowitch, E. I., 9, 14,29,49, 149, 150, 205, 210 Rademaker, H., 93, 210 Radmer, R. J., 17, 20, 74, 137, 149, 153, 210 Radunz, A., 90, 204 Raff, R. A., 183, 211 Ragan, M. A., 54, 177, 178, 193, 211 Raggi, V . , 264, 275 Raghaven, N. V., 63, 211 Rahimi, A., 233, 274 Ramanathan, V., 6, 175, 210 Ramus, J., 28, 29, 44,45, 121, 152, 167, 169, 211 Rao, K. K., 176, 186, 193 Rapoport, H., 58,66, 118, 183,204,214 Rath, K., 67, 204 Raven, J. A., 20, 21, 24, 31, 36, 149, 150, 154, 158, 163, 183, 188, 211 Raven, P. H., 184, 211 Rawlinson, C. J., 252, 274 Raychaudhuri, S. P., 255, 274 Raymont, J. E., 22, 211 Raynes, D. A., 149, 210 Reade, J. A., 45, 203 Recouvreur, M., 138, 146, 211, 218 Redlinger, T., 112, 197,211 Reeves, S . G., 87, 196 Reger, B. J., 52, 211 Reich, R., 60, 213 Reichard, Th., 244 Reid, E. H., 128, 201 Reimer, T. O., 181, 211 Reinhardt, B., 146, 149, 211 Reinman, S . , 62, 76, 82, 91, 102,211,215 Reis, E. M . , 231, 243, 251, 254, 255, 258, 2 74 Reiss-Husson, F., 103, 206 Remsen, C., 31, 211 Rtmy, J., 98, 200, 211, 216 Rtmy, R., 81, 83, 98, 104, 106, 107, 200, 205, 21 I Renger, G., 92, 211 Rest, van der, 60, 62, 192

AUTHOR INDEX

Reuter, D. J., 252, 254, 274 Reynolds, J. A,, 85, 104, 199. 211 Reynolds, C. S., 42, 217 Rice, J. D., 12, 216 Richardson, F. L., 156, 157, 193 Ried, A., 146, 149, 211 Ried, A. B., 146, 211 Rigby, M., 110, 211 Rijgersberg, C. P., 46, 73, 92, 211, 215 Riley, J. P., 58, 211 Ripley, G. W., 109, 208 Robbins, W. R., 259, 272 Robinson, R. W., 243, 253, 274 Robson, A. D., 236, 274 Rodger, J. B. A., 244, 245, 246, 271, 274 Rodgers, M. A. J., 60, 194 Rogers, P. F., 244, 246, 274 Rohde, G., 241, 274 Romanova, A. M., 260, 274 Romeo, A,, 122, 211 Romheld, V., 256, 266, 273 Rood, R. T., 177, 209 Rosell, R., 257, 274 Rosinki, J . , 110, 211 Rosenberg, G., 152, 211 Rouse, R. D., 225, 271 Rovira, A. D., 227, 251, 256, 266, 271, 2 74

Rubin, A. B., 152, 216 Rudiger, W., 51, 63, 64, 66, 67, 68, 191, 205, 208, 211, 218

Rudnik, M. S., 126, 189 Ruehle, W., 107, 217 Ruffer, U., 30, 36, 74, 209, 210, 211 Ruppel, G., 60, I95 Rurainski, H. J., 164, 204 Rusckowski, M., 79, 81, 211 Rush, J . D., 79, 87, 196 Rutherford, A. W., 83, 93, 94, 211, 212 Rutkovskaya, V. A., 1 I , 210 Ryan, M. D., 87, 193 Ryrie, T., 101, I97 Ryrie, I. J., 107, 147, 212 Ryther, J., 151, 2 / 2 Ryther, J . H., 164, 212

S Sackton, W. E., 240, 272 Sadasivan, T. S., 222, 241, 243, 255, 274 Salares, V. R.,61, 212

29 1

Salisbury, J . L., 188, 196 Samuel, G., 243, 249, 275 Samuelsson, G., 59, 209 Samura, S., 37, 52, 59, 72, 108, 109, 162, 202

Sandman, G., 88, 212 Sands, D. C., 255, 256, 272 Sane, P. V., 40, 77, 139, 187, 212 Santore, V. J . , 186, 199 Saraswathi-Devi, L., 255, 275 Sarda, A., 116, 142, 156, 199 Sarojini, T. S., 243, 275 Satoh, K., 73, 88, 90, 91, 101, 104, 105, 115, 164, 212, 218 Sauer, K., 99, 135, 193, 212 Saunders, B. C., 248, 275 Saunders, V. A,, 104, 212 Saunderson, J., 263, 271 Savidge, G., 162, 212 Schaffernicht, H., 60, 61, 83, 97, 202, 212 Schantz, K., 78, 86, 106, I90 Scheer, H., 50, 51, 52, 63, 64, 65, 66, 67, 68, 204, 212 Scheibe, J., 174, 195, 212, 216 Schidlowski, M., 174, 175, 177, 182, 212, 21 7 Schiff, J. A., 144, 173, 183, 212 Schimper, A. F. W., 183, 212 Schmid, G. H., 90, 204 Schmid, G. M.. 59, 207 Schmid, M. F., 103, 207 Schmidt, G., 91, 191 Schmidt, H . E. 259, 275 Schmucker, T., 241, 275 Schneider, H . , 68, 208 Schneider, H-J., A. W., 174, 209 Schoch, I., 96, 193 Schonbohm, E., 30, 212 Schopf, J. W., 174, 175, 177, 180, 212 Schramm, W., 170, 203 Schreiber, K., 256, 266, 275 Schreiber, U., 67, 117, 212, 213 Schroth, M. N., 255, 256, 272, 275 Schutt, F., 213 Schutte, K. H . , 230, 231, 240, 275 Schwartz, R. M., 175, 184, 194,213 Scott, B., 213 Scott, W. T., 53, 217 Sculley, M. J., 146, 147, 148, 194, 213 Seaburg, K. C., 162, 181, 210 Searle, G. F. W., 112, 114, 133,210,213 Sears, J. R., 22, 169, 213

292

AUTHOR INDEX

Simojoki, P., 231, 239, 275 Sebald, W., 39, 40, 213 Simpson, D. J., 104, 138, 139, 206, 214 Seckbach, J., 213 Sinha, A. K., 261, 275 Seely, G. R., 95, 213 Singh, P.,257, 275 Seewaldt, E., 184, 186, 213 Singh, R.P., 260, 273 Seiburth, J. M . , 10, 213 Sistrom, W. R., 31, 198 Selset, R., 66, 21 7 Skrdla, M. P., 76, 102, 104, 147, 191,207 Sempio, C., 260, 261, 263, 264, 275 Skews, G., 66, 208 Seng, P-S., 65, 133, 202 Senger, H., 49, 52, 58, 62, 76, 77, 83, 90, Skok, J., 241, 276 95, 151, 152, 153, 156, 158, 160, 172, Skov, J. P., 251, 275 173, 191, 192, 195, 196, 204, 213, 217 Skyring, G. W. , 181, 190 Sletten, K., 66, 217 Senn, G., 29, 213 Smayda, T. J., 24, 42, 214 Serlin, R., 50, 194 Smiley, R. J., 253, 265, 275 Seto, H., 183, 209 Smirnov, Yu.S., 260, 261, 275 Setif, P., 86, 99, 106, 204, 213 Smith, A., 183, 206 Sevchenko, A. N., 95, 99, 199 Smith, A. J., 41, 189 Seybold, A., 29, 213 Smith, B. N., 243, 270 Sewe, K. U . , 60, 213 Sharp, E. L., 257, 258, 275 Smith, F. A., 21, 41, 211, 214 Smith, F. G., 257, 258, 275 Shaw, E. R., 92,204 Sheldrick, G. M., 66, 192 Smith, K. M.,178, 201 Shepherd, S. A., 169, 213 Smith, R. C., 9, 12, 164, 214, 216 Sherman, L. A., 79, 81, 90, 209 Smith, R., 83, 21 7 Shestiperova, Z. I., 241, 243, 270 Smith, S. E., 259, 275 Shibata, K., 52, 77, 108, 208, 209 Snyder, L. C., 95, 199 Shilo, M., 164, 189 Soloviev, K. N., 95, 99, 199 Shimokoriyama, M., 218 Sommer, S. E., 229, 275 Shimokoriyama, M., 187, 215, 218 Song, 60, 214 Shimura, S., 42,54, 59, 101,212,213,218 Song, P. S., 61, 120, 135, 136, 173, 204, Shindo, H., 248, 275 214 Shinkarev, V. P., 152, 216 Sonneveld, A., 214 Shiozawa, J. A,, 77, 84,97, 105, 146,213, Spark, A. A., 58, 66, 192 215 Sparrow, D. H . B., 253, 271 Shipman, L. L., 83, 84, 96, 98, 100, 102, Spasic, P., 259, 275 131, 203, 210, 213, 218 Spatz, L., 244, 275 Shipton, P. J., 249, 270 Spence, D. H. N., 16, 214 Shkolnik, M. Y., 239, 240, 241, 242, 275 Spencer, K. G., 65, 161, 218 Shriner, D. S., 236, 273 Sperling, W., 60, 195 Shubin, L. M., 98, 213 Spijkerboer, F. W . J. M., 217 Shuvalova, G. V., 241, 243, 270 Spinks, G. T., 257, 264,275 Shuvalov, V. A., 93, 203, 213 Spruit, C. J., 174, 194 Sidler, W . , 66, 197, 213 Spurr, A. R., 241, 275 Siefermann-Harms, D., 26, 54,59,60,61, Stackbrandt, E., 175, 183, 184, 186, 197, 62, 104, 105, 106, 107, 200, 214 213 Siegelman, H. W., 58, 63, 110, 118, 193, Staebelin, L. A., 32, 40, 116, 137, 138, 200, 211, 214 139, 142, 156, 190, 198, 207, 214 Sielicki, M., 121, 202 Stahl, D. A., 175, 183, 184, 186, 197 Sigalat, C., 90,214 Stark, B. P., 248, 275 Simons, K., 104,200 Steele, J. H., 152, 214 Simionescu, C. I., 175, 214 Steeman-Nielsen, E., 151, 152, 158, 164, Simionescu, B. C., 175, 214 214 Simmons, G. M. Jr, 162, 181, 210 Stefanci, V., 134, 195

AUTHOR INDEX

Stellwagen, E., 83. 214 Steinbeck, K. E., 107. 147, 148. 189, 214 Sterling, C., 58. 214 Stern, A,. 49, 196 Stevendon, D. J . , 174. 175, 177 Stevens, C. L. R., 69, 72, 73, 74, 84, 115, 161, 171, 217 Stevens, S. E. Jr, 68, 199 Stevens. S. E., 66, 68, 198 Stewart, A., 89, 90, 91, 211, 214 Stewart. A. C., 76, 82, 87, 89, 90. 91, 105, 193, 209, 214

Stewart, K. D., 5, 26. 188, 214 Stoddart, J. L., 259, 270 Stoller, M., 62. 204 Stout, P. R., 267, 270 Strain, H. H., 52,53,54,58, 102. 118, 195. 203, 214

Strakhov, T., 255. 257. 275 Stransky, H., 55, 62, 199. 214 Strassberger, G., 58, 77, 90, 213 Strobel, G . 4., 255, 256, 272 Strouse, C. E.. 36, 50, 194, 215 Stuehn. N., 107, 217 Sugahara, K., 101, 215 Sugimura. Y., 187, 215, 218 Sugiyama, K-I., 97, 215 Sulochana, C. B., 258, 275 Sundqvist, C . . 172, 215 Suryanarayanan, S., 226, 275 Sutic, D., 259, 27.5 Suzuki, S., 63, 189 Svedberg, T., 63, 215 Svec, W. A.. 52,53.58, 118, 190,195,202, 214 Swanson. R., 65. 133, 202 Swarthoff, T., 93, 94, 215 Sweeney, B. M., 29, 118, 119. 120, 151, 157, 158, 199, 200, 210 Sweet, R. M., 67, 68, 132, 133, 134, 196 Swift, E.. 29, 215 Szalontai, B., 61, 62, 63, 215 T Tai, L. H., 261, 272 Tainio, A., 231, 238, 239, 275 Takahashi, M., 24,40, 152,210 Takahashi, Y., 262, 270 Takamiya, A., 100, 208, 215 Takase, T., 261, 273

293

Takatoh, H . , 261, 273 Takayima, A , 101 Talbot. M. C.. 151, 158, 210 Talling. J . F., 9, 12, 16. 152, 180, 215 Tamford, C., 60, 206 Tamura. N., 87, 215 Tandeau, De, Marsac, N., 166, 193, 205 Tanada, T., 60, 70, 74. 215 Tandeau de Marsac, N., 41,70. I I 1 , 115, 215

Tanford, C., 104, 200 Tangen, K., 58. I91 Tanner, R. S., 175, 183, 184, 186. 197 Taraz. H., 53, I93 Taylor, F. J. R., 184, 215 Taylor, W. R., 29, 215 Teale, F. W. J., 113, 133, 134, 194, 215 Tec, J. L., 58, 66, 192 Teintze, M., 255, 256, 272 Templeton, D. H., 50, 51, I96 Teten’Kin, V. L . , 99, 199 Theodor, R., 215 Thiele, R., 30, 200 Thielen, A. P. G. M., 46, 72, 91, 92, 215 Thinh, L. V., 188, 215 Thompson, J. F., 226, 275 Thomas, 63, 215 Thomas, D. M., 55, 206 Thomas, J . C., 78, 81, 85, 91, 116, 142, 205, 215

Thomou, H., 105, 107, 190 Thornber, J . P., 36.49, 52, 62, 76, 77, 78, 79,82,84,91,97,99, 102, 104, 105, 106, 107, 108, 120, 122, 145, 146, 147, 186, 189, 191, 192, 193, 195, 202, 207, 211, 213, 215, 218 Thorne, S. W., 37, 48, 52, 53, 59, 62, 72, 80, 84, 85, 88, 89, 90, 98, 99, 101, 102, 103, 104, 105, 106, 107, 108, 121, 122, 126, 129, 146, 147, 148, 160, 162, 164, 165, 186, 189, 190, 191, 194, 195, 197, 210, 213, 215, 216 Thornley, J . H. M . , 152, 216 Thrash, R. J., 61, 216 Tills, A. R., 254, 275 Tilney-Bassett, R. A. E., 32, 35, 203 Timofeeva, V. A,, 13, 216 Tiselius, A., 63, 216 Titlyanov, E. A., 121, 167, 205, 216 Tomlinson, J. A,, 257, 275 Torres-Periera, J., 30, 47, 208 Toms, J., 231, 275

294

AUTHOR INDEX

Town, W. R., 203 Traeger, E., 206 Treadwell, C. J., 112, 114, 133, 210, 213 Trebst, A., 19, 38, 216 Tremolieres, A,, 104, 106, 107, 216 Troche, R. P., 12, 216 Trolldenier, G., 222, 227, 249, 275 Trosper, T. L., 36, 215 Troxler, R. F., 63, 66, 68, 179, 181, 183, 192, 193, 209, 216 Truscott, T. G., 60, 61, 194 Tsujimoto, H. Y . , 190 Turlygina. E. S., 258, 275 Tyler, J. E., 9, 46, 216 U Uexkvell, H. R. Von, 222, 226, 275 Ulrich, A., 226, 257, 271, 274 Utkina, N. I., 241, 276

V Van Baalen, C., 12, 216 Van Best, J. A., 91, 93, 216 Van Den Driessche, T., 29, 30, 216 Van Ginkel, G., 163, 216 Van Gorkom, H. J., 46, 72, 91, 92, 215, 216 Van Houten, A,, 89, 104, 199 (in press) Van de Hulst, H. C., 47, 216 Van Metter, R. L., 135, 204 Van de Ven, M., 62, 63, 215 Vanterpool, T. C., 225, 276 Veek-Horsley, K. M., van der, 93,94,215 Velichko, I. M., 54, 216 Verrnaas, W. F. J., 94, 216 Venediktov, P. S. A., 152 Verderevskii, D. D., 260, 276 Verrna, G. S., 265, 276 Verma, H. N., 265, 276 Vernon, L., 203 Vernon, L. P., 79, 81, 90, 101, 203, 204, 209 Vesk, M., 39, 40, 129, 139, 140, 172, 195, 201, 216 Vierling, E., 94, 151, 156, 158, 216 Vincent, W. F., 162, 216 Virgin, H. I., 172, 215 Vlamis, J., 243, 262, 276

Vogelmann, T. C., 174, 216 Volk, R. J., 261, 262, 276 Volkman, J. K., 186, 202 Von Caemmerar, S., 31, 40, 152, 196 Vooren, C. M., 170, 216 Voorn, G., 88, 217 Voskresenskaya, N. P., 172, 216 Voynow, P. V., 68, 192 W Waalund, J. R., 116, 156, 216 Waalund, S. D., 116, 156, 216 Wada, K., 186, 190, 207 Waggoner, P. E., 255, 276 Wagner, F., 262, 276 Wagniere, 95, 199 Waight, E. S., 55, 206 Wakabayaski, S., 186, 207 Waldron, J. C., 37, 52,72,84,85,88, 101, 102, 106, 107, 108, 162, 189 Walker, C. D., 236, 276 Walker, J. C. G., 174, 175, 176, 217 Walker, N. A., 41, 69, 195, 214 Wallen, D. G., 217 Wallin, R., 66, 217 Walsby, A, E., 42, 217 Walter, M. R., 174, 175, 177, 217 Wang, R. T., 39, 69, 72, 73, 74, 84, 94, 115, 117, 144, 146, 160, 161, 171, 172, 208, 2 I 7 Wanner, G., 35, 67, 1 1 1, 204, 217 Waritz, R. S., 255, 273 Wasielewski, M. R., 83, 217 Wasley, J. W. F., 53, 217 Watanabe, R., 241, 276 Watson, R. D., 222, 265, 272 Waveren, V. A-V., 88, 217 Webb, J., 236, 276 Webb, M. J., 260, 276 Webb, M. J. W., 257, 275 Weedon, B. C. L., 51,55,58,66,192, 193, 206, 208, 217 Wegfahrt, P., 58, 118, 214 Wehrmeyer, S., 35, 217 Wehrmeyer, W., 64,65,68, 109, 110, 1 1 1, 112, 133, 134, 204, 208, 209 Weidner, M., 123, 209 Weinburg, E. D., 256, 276 Weinberg, S., 8, 14, 16, 217 Weinhold, A. R., 244, 276

AUTHOR INDEX

Weintraub, R. L., 261, 262, 276 Weiss, A,, 217 Weiss, C., Jr, 95, 96, 217 Welch, R. M., 258, 260, 276 Wellburn, A. R., 51, 77, 217 Wellburn, F. A. M . , 77, 217 Weller, J-P., 63, 198 Wells, G. N., 12, 216 Wender, S. H., 241, 276 Wenzel, G., 243, 276 Wenzel, H., von, 244, 249, 276 Wessels, J. S. C., 88, 90, 213, 217 West, J. A,, 65, 161, 218 Westlake, D. F., 16, 217 Weyrauch, S. K., 69.70, 71, 73, 1 15, 1 17, 118, 146, 205 Wharton, R. A., Jr, 162, 181, 210 Whatley, F. R., 183, 184, 185, 186, 217 Whartley, J. M., 32, 41, 183, 184, 185, 186, 188, 217 Wheeler, W. H., 122, 217 Whittaker, R. H., 4, 121, 217 Whitten. 50 Widmer, H., 66, 197 Wieslander, A,, 61, 202 Wilbrant, R., 61, 202 Wilce, R. T., 22, 213 Wild, A,, 80, 89, 102, 106, 107, 154, 158, 21 7 Wildermuth, G. B., 227, 251, 274 Wildman, R. B., 109, 217 Wildner, G. F., 88, 218 Williams, C . N., 228, 240, 241, 276 Williams, D. E., 262, 276 Williams, R. C., 1 1 1, 112, 206, 218 Williams, W. P., 17, 66, 136, 137, 138, 145, 166, 218 Willenbrink, J., 123, 209 Willstatter, R., 52, 218 Wilson, G. C . S., 243, 244, 273 Wilson, T. R. S., 58, 211 Wilson, W. H., 47, 48, 203 Winsor, G. W., 236, 270 Withers, N. W., 32, 108, 186, 198, 218 Witt, K., 92, 218 Woese, C. R., 175, 183, 184, 186, 197 Wolfe, R. S., 175, 183, 184, 186, 197 Wolfgang, J., 212 Wolk, C. P., 183, 218 Wollman, F. A,, 80, 89, 91, 94, 116, 137, 138, 139, 142, 146, 191, 195, 218 Womersley, H. B. S . , 169, 213

295

Wong, H., 67. 134, 218 Wood, A. M., 52, 218 Wood, J. M., 231, 276 Wood, N. B., 41, 218 Wood, P. M., 87, 88, 186, 194, 218 Wood, R. B., 12, 180, 215, 218 Wood, R. K. S., 240, 242, 276 Woodruff, W. H., 60, 194 Woodcock, C. L. F., 80, 89, 190 Woods, N. E., 67, 68, 132, 133, 134, 196 Woodward, R. B., 48, 50, 218 Worden, P. B., 31, 198 Wortley, W. R. S., 264, 276 Wright, D., 244. 276 Wright, J. L. C., 183, 205 Wu. G. S . C., 60, 193 Wynd, A,, 244, 245, 246, 271, 274 Wynne. M . J., 5, 192

X

Xavier. A. V., 83, 218

Y

Yakushiji, 103 Yamada, N., 261, 273 Yamamoto, Y.. 87, 215 Yamamoto, H. Y., 62, 214 Yamanaka, G., 41, I 1 I , 218 Yamaoka, T., 76, 79, 81, 89,90, 104, 105, 208, 218 Yarham, D. J . , 253, 276 Yaroshenko, T. V., 255, 257, 275, 276 Yarwood, C. E., 240, 243, 259, 276 Yentsch, C. S., 14, 47, 218 Yevstigneyev, V. B., 52, 207 Yocum, C. F., 89, 191 Yocum. C. S., 74, 191, 218 Yogeswari, L., 257. 276 Yokohama, Y., 37, 52, 54, 59, 72. 108, 109, 162, 202, 218 Yoshizaka, F., 218 Young, N. M., 61, 212 Yu, A,, 60, 209 Yu, C. H., 261, 272 Yu, M-H., 65, 161, 192, 218 Yuen, M. J., 100, 218

296

AUTHOR INDEX

Z Zablen, L. B., 175, 183, 184, 197 Zalkin, A., 50, 51, 196 Zanefeld, J. R., 13, 218 Zanefeld, J. R. V., 12, 218 Zattler, F., 259, 276 Zerner, B., 261, 271 Zickendraht-Wendelstadt, B., 67, 68, 205, 218 Zilinskas, B., 64, 66, 72, 111, 115, 197, 199, 219

Zilinskas, B.A.,67, 79,81, 110, 114, 115, 134, 195, 211, 218 Zimmerman, C., 121, 211 Zimmerman, B. K., 64, 109, 110, 1 14, 197, 219 Zinsmeister, H., 215 Zuber, H., 66, 114, 134, 193, 197, 199, 213, 219 Zubko, I. Y.,243, 276

Subject Index A Acetabularia chlorophyll-protein analysis, 105-106 chloroplast shape and size changes, 30 photosystem I1 reaction centre complex, 89 thylakoid extraction, 104 A . mediterranea chlorophyll-protein analysis, 106, 108 chloroplast movement, 29 photosystem reaction centre complexes, 80, 89 Acrocarpia paniculata chlorophyll-fucoxanthin complex, 122, I24 photosystem reaction centre complexes, 80, 90, 91, 130 spectral analysis, 97, 98, 100, 101, 123, 125, 126, 127 Action spectra for photosynthesis in algae, 68-73 Adult Plant Resistance (APR) and the effect of copper, 230, 236 Agmenellum quadruplicatum, phycobiliproteins, 66, 68 Alteneria, effect of copper level on host infection, 23 1 Aluminium, effect on plant disease, 264265 Aminopeptidase, effect of manganese on enzyme activity, 244 Amphidinium carterae blue light effects, 172 peridinum-chlorophyll-protein complex, 119, 120 Anabaena chlorophyll-protein complex, 77 fluorescence spectra, 100 A . cylindrica photosystem reaction centre complexes, 82

phycobiliprotein, 1 15 A . Jos-nquae buoyancy control, 42 photosystem reaction centre complexes, 79 A . variabilis antenna chlorophyll, 94 phycobiliproteins, 63, 66, 109, 115 shading effects, 156, 160 Anacystis nidulans chromatic adaptation, 166, 171, 172 photosynthetic rate, 151 photosystem reaction centre complexes, 84, 90 phycobiliproteins, 66, 1 15 shading effects, 156 Aphanocapsa photosystem primary electron acceptor, 92 phycobiliproteins and nitrogen starvation, 41 Ascophyllum nodosum chlorophyll a/c ratio, 121 ATP synthesis in evolution, 176 in photosynthetic bacteria, 19-20, 132, 176 in photosynthetic electron transport, 19. 38

B Bacteriorhodopsin, ATP synthesis, 132, 176 Barley, nutrient effects on powdery mildew infection, 223, 224 yield and ergot, 237-238, 239 Bermuda grass, nutrient effects on leaf spot disease, 225 Biddulphia aurita, blue-light effects, 172 Blue-light effects in algae, 172

298

SUBJECT INDEX

chloroplast membrane fractionation, 104 photosystem reaction centre complexes, 78, 80, 84, 86, 89, 9194 Chlorella chlorophyll, 52, 83 difference absorption spectra, 98 energy fixation, 20 quantum efficiency spectra, 73, 74, 75 spectral modification, 14 C. elfipsoida, chlorophyll alb-carotene C Cadmium, effect on plant disease, ratio, 77 C . fusca 263-264 Calcium, effect on plant disease, 228,241 chlorophyll-pfotein complex, 106, 107 Callithamnion roseum photosystem reaction centre comphycobiliproteins, 65 plexes, 80, 89 C . pyrenoidosa, shading effects, 156 shading effects, 161 Canopy dominant algae and light har- C . vulgaris vesting, 21-22, 28 fluorescence spectra, 100, 137 Carotenoids in algae photosynthetic rate, 151 apoprotein binding, 59-60 quantum efficiency, 152 chemistry, 58 reaction centre electron acceptor, 93 configuration and conformation, Chlorobium, chlorophyll, 83, 177 60-6 1 Chlorobotrys, shading effects, 163 distribution in algae, 2627, 5 6 5 7 Chlorogloea fritschii, photosystem reacenergy transfer, 61-62 tion centre complexes, 79, 87 photoprotective role, 62 Chlorophylls spectra, 96 assay and distribution, 2627, 51-52 structure, 55, 58 chlorophyll a, 49-5 1 Carotenoid-protein complexes chlorophyll b, 51 chlorophyll c and fucoxanthinchlorophylls c and c2, 52-53 containing complexes, I2&129 chlorophyll d, 53-54 peridinin-chlorophylla-protein,1 18evolution, 178 120 Chlorophyll proteins Caulerpa, chloroplast movements, 30 curve deconvolution analysis, 9 6 9 8 C. cactoides difference absorption spectra, 98-99 light harvesting by siphonoxanthin, 72, fluorescence spectra, 99-1 02 108 Chloroplast photosystem complexes, 85, 89 evolution, 183-184 Chaetoceros gracilis, photosynthetic rate, light scattering, 48 151 movement and light harvesting, 29-30 Chenopodium album, chlorophyll-protein Chondrus crispus, photosynthetic rate, complex, 103 168 Chlamydomonas Chorda .filmurn,fucoxanthin/chlorophyll carotenoid type, 54 a ratio, 121 photosystem structure, 137 Chromatic adaptation in algae C . mundana, electron donor to P700, 88 historical aspects, 165-166 C . reinhardtii ontogenic complementary adaptation, absorption spectra, 97, 100 166167 analysis of chlorophyll-protein comphylogenic complementary adapplex, 105-106 tation, 167-171 Bordeaux mixture and control of plant disease, 229, 238 Boron biochemistry of deficiency, 241 effect on plant infection, 238-243 Botrytis, effect of calcium on infection, 228 Bumilleriopsisjiliformis fluorescence spectra, 100 photosynthetic electron donor, 88

SUBJECT INDEX

Chroococcus quantum efficiency of spectra, 73,7475 spectral modification, 14 Chroomonas shading effects, 157 thylakoid structure, 34 Cladophora, chlorophyll a/b ratio, 52 Cladostephus spongiosus, light-harvesting complex, 124 Clark-type oxygen electrode, algal action spectra, 69 Claviceps purpurea copper and boron effects on host infection, 239 copper fertilizer effects on host infection, 237-238 Closterium, shading effects, 157 Clostridium tetranomorphum, evolution of photosynthetic pigments, 179 Cnacystis nidulans, photoinhibition of photosynthesis, 164 Codium cactoides, chlorophyll a/b ratio, 52 C . fragile carotene type, 54 morphology and light harvesting, 28 photosystem reaction centre complex, 89 siphonaxanthin, 108 Coilodesme action spectrum of photosynthesis, 70 photosynthetic rate, 168 Colpomenia sinuosa, light-harvesting complex, 124 Common scab of potato, see Streptomyces scabies Copper effect on plant disease, 229-238 interaction with other nutrients, 231-234, 239, 255 involvement in phenol synthesis, 236, 237 Copper oxychloride and control of plant disease, 229 Cotton, effect of potassium on yield and wilt infection, 225 Cryptomonas ovata, shading effects, 157 C . rufescens, shading effects, 157 Cryptophyta chlorophyll, 52, 53 thylakoid structure, 34, 35 xanthrophylls, 55

299

Cyanidium caldarium photosystem structure, 137 phycobiliprotein evolution, 181 phycobilisomes, 66, 142 shading effects, 157 Cyanobacterium antenna chlorophyll, 94 buoyancy control, 42 distribution and light quality, 23, 24 photosystem reaction centre complexes, 82 phycobiliproteins, 48, 64, 109 thylakoid arrangements, 33, 35 Cystoseira mediterranea, fluorescence spectrum, 100, 126 Cytological aspects of light harvesting in algae cell morphology, 28-29 chloroplast movement, 29-30 chromatophores, 3&36

D Delesseria sanguinea, photosynthetic rate, 168 Deriphat 160, solubilization of chlorophyll-protein complexes, 104 Detergents for fractionation of chlorophyll-protein complexes, 104-105 Dictyota dichotoma, chloroplast movement, 29, 30 Digitonin, fractionation of reaction centres, 104 Dunaliella adaptation to high irradiance, 22 chlorophyll a/b ratio, 52 D. euchlora, photosynthetic rate, 151 D. parva, photosystem reaction centre, 87 D. tertiolecta antenna chlorophyll, 94 photosynthetic rate, 151 shading effects, 156, 160, 162 thylakoid structure, 34 E Ecklonia radiata chlorophyll-fucoxanthin-proteincomplex, 124 photosystem reaction centre complexes, 78 thylakoid structure, 34

300

SUBJECT INDEX

Ecological aspects of light harvesting in algae, 21-23 Enteromorpha, morphology and light harvesting, 28 E. prolifera, properties of plastocyanin, 87 Ergot, see Claviceps purpurea Erysiphe cochoracearum, boron deficiency and host infection, 240 E. graminis lithium deficiency and disease, 264 silicon/manganese in disease, 262 E. graminis DC var. hordii macronutrient effects on host infection, 223, 224 micronutrient effects on host infection, 230, 231, 232, 240 phenol synthesis and infection, 236 Euglena gracilis absorption spectra analysis, 96 antenna chlorophyll, 95 chloroplast evolution, 183 electron donor to P700, 88 fluorescence spectra, 100, 101-102 Evolution of early photosynthetic prokaryotes, 18&183 eukaryotic algae, 183-187 photosynthesis, 174-177 photosynthetic pigments, 177-180 thylakoid stacking, 187-188

F Fluorescence spectra of chlorophylls, 99-102 of phycobiliproteins, 112-1 15 Fomes annosus, manganese deficiency and infection, 243 Forster resonance in algal photosynthetic systems, 131, 133, 134, 135 Fremyella, difference absorption spectra, 98 F. diplosiphon phycobilisome structure, 1 10 synthesis of light-harvesting proteins, 174 Fucoxanthin absorption of light, 59, 72 distribution in algal groups, 5 6 5 7 energy transfer, 60 evolution in prokaryotes, 181 structure, 55, 59

Fucus serratus chloroplast isolation, 123 fluorescence spectrum, 100, 125, 126, 128 F. vesiculosus chlorophyll a/c ratio, 121 chloroplast arrangement, 36 morphology, 30 movement, 30 Fusarium micronutrient effects on infection, 225, 227, 240, 255 pectinases, 241, 255 F. martii var. pisi, effect of micronutrients, 265 F. oxysporum var. lini effect of boron deficiency, 240 effect of zinc on host infection, 258 F. oxysporum var. lycopersici, nutrient effects on lycomarismin toxicity, 255 F. udum, manganese deficiency and infection, 243 G Gaeumannomyces graminis chloride action mechanism, 265 host infection and concentration of copper, 231 lime, 227 manganese, 249-254 manganeseliron, 256257 phosphorus, 225 Gigartina papillata, chlorophyll d , 53 Glenodinium action spectrum of photosynthesis, 71 chlorophyll c and fucoxanthin, 121, 122 peridinin-chlorophylla-protein, 7 1, 119, 136 photosynthetic rate, 151 shading effects, 157 Gloeocapsa NS4, thylakoid structure, 33 Glutamine and infection of potassium deficient plants, 226 Gonyaulax polyedra chloroplast movement, 29 peridinin-chlorophyll a-protein, 119, 120 photosynthetic rate, 151

30 1

SUBJECT INDEX

photosystem reaction centre complexes, 78 shading effects, 157, 158 Grifithsiaflosculosa, phycobilisomes, 109 G . monilis action spectra for photosynthesis, 71 phycobilisomes, 113, I18 photosystem reaction centre complexes, 84, 85, 89 G . pacifica phycobilisome structure, 1 I I , 142 shading effects, 156 Gyrodinium, chlorophyll c-fucoxanthin, 121

H Halimeda tuna, chloroplast movements, 30

Haliption cuvieri, thylakoid structure, 34 Halobacterium rubrum, bacteriorhodopsin, 132, 176 Helminthosporium, nickel stimulation of host fungitoxins, 261 H . oryzae host infection and manganese level, 243 silica level, 261 H . turcicum, nitrogen supply and host infection, 228 Hemiselmis, phycobiliproteins, 65 Heterodera, copper level and host infection, 231 Hormosira banksii, light-harvesting complex, 124

I IAA metabolism and nickel concentration, 261 Iron level and plant disease, 254257,267 Isochrjsis galbana EM of thylakoids, 141 photosynthetic rate, I5 1

L Laminariu cichorioides, fucoxanthin/ chlorophyll u ratio, 121 L . sarcharina action spectrum for photosynthesis, 70 photosynthetic rate, 168 pigment levels and nitrogen, 42 Lauryldimethylamine oxide, fraction-

ation of cytochrome oxidase, 104 Lepidium, absorption spectrum analysis, 97

L . virginicum, chlorophyll-protein complex, 103 Light climate for algae air-water interface, 8 effect of algae on light attenuation, 12-14

light attenuation in water, 9-1 1 sunlight composition, &8 ultraviolet-B irradiance, 12 underwater light climates, 14-16 Light harvesting strategies in algae biochemical strategies buoyancy control, 42 electron transport, 3 8 4 0 membrane structure, 38 nitrogen level, 4 1 4 2 photosynthetic unit, 38 pigments, 3 6 3 7 spatial constraints in chloroplasts, 40-41

cytological aspects, 28-36 ecological aspects, 2 1-22 morphological aspects, 24-28 physical strategies energy migration, 4 2 4 light scattering, 47-49 package effect, 4 U 7 Lignification, involvement of boron, 241, 267 iron, 254255, 267 manganese, 247-249, 267 Lithium and plant disease, 264 Lycomarismin toxicity, effect of iron and copper, 255 Lyngbya, thylakoid structure, 34 M Macrocystis pyrifera, photosynthetic pigments, 122 Manganese level and plant disease, 243-254 in lignin biosynthesis, 247-249 Musfigocladus laminosus, phycobiliproteins, 65, 66, 109, 181 Melampsora, boron deficiency, 240 Mesotaenium, chloroplast movements, 30 Micronutrients and plant diseasr boron, 238-243 copper, 229-238

302

SUBJECT INDEX

Micronutrients and plant disease-cont. iron, 254257 manganese, 243-254 nickel, 260-261 silicon, 261-263 zinc, 257-260 Mongeotia chlorophyll a/b ratio, 52 chloroplast movements, 30 Morphological aspects of light harvesting in algae macroalgae, 24-25, 28 unicellular algae, 24 Mycorrhiza, effect of zinc on development, 259

manganese, 243-254 nickel, 260-261 silicon, 261-263 zinc, 257-260 0

Octyl-P-D-glucopyranoside, thylakoid extraction, 104 OIpidium brassicae, iron and host resistance, 255 Ophiobolus graminis, see Gaeumannomyces graminis Oscillatoria agardhii, phycobiliprotein structure, 66 0. limosa, photosystem reaction centre complexes, 78, 8 1, 85 0.splendida, photosystem reaction centre complexes, 79, 8 1, 85 Osterobium, effects of shading, 163

N Nannochloris atomus, spectral modification, 14 Navicula minima, quantum efficiency spectra, 74 P Neottia nidus-avis, green light harvesting, Padina commersonnii, photosystem re59 action centre complex, 90 Nickel, effect on plant disease, 260-261 Pavlova lutheri, chlorophyll/fucoxanthin Nicotiana, chlorophyll content, 83 complex, 122 N. tabacum Pectin methyl esterase, effect of nutrient difference absorption spectra, 98 stress on activity, 243-244,255,266 photosystem reaction centre com- Peridinin plexes, 77 absorption, 59 Nitrogen distribution in algal groups, 5 6 5 7 concentration and host susceptibility, evolution in prokaryotes, 181 224, 226, 228, 231, 233, 262 Peridinin-chlorophyll-protein complex, deficiency and plant adaptation, 118, 120, 135-136 222-223 Peroxidase in lignin synthesis, 248, 251, supply and RuBPc’ase, 41 255 Nostoc Phaeodactylum photosystem reaction centre comlight absorption by fucoxanthin, 72 plexes, 79, 81 shading effects, 157 phycobiliproteins, 67, 68, 115 P. tricornutum Nutrient stress and plant disease chlorophyll-fucoxanthin-protein,123 guidelines for experimentation, nitrogen starvation and fucoxanthin, 268-269 42 macronutrients spectral modification, 14 nitrogen, 226228 Phaeophyta phosphorus, 224-225 light harvesting, 23 potassium, 225-226 photoprotective system, 62 sulphur, magnesium and calcium, Phenolics synthesis and resistance to in228 fection, 233, 235, 236, 237, 242, micronutrients 247-249, 261 boron, 238-243 Phormidium, photosynthetic rate, 168 copper, 229-238 P. laminosum, photosystem reaction iron, 254255 centre complexes, 82,87,89,90,91

SUBJEC'T INDEX

P. luridium, photosystem reaction centre complexes, 77, 79, 82 Phosphorus deficiency and plant adaptation, 222-223 involvement in plant disease, 224225 Photoinhibition of photosynthesis, 163-165 Photosynthetic carbon reduction cycle, 17 Photosynthetic electron transport chain, 17 Photosynthetic membrane structure, 17-20 Photosynthetic pigments action spectra and quantum yields, 68-73 carotenoids apoprotein binding, 59-60 chemistry, 58 configuration and conformation, 6 6 61 distribution, 5 4 5 8 energy transfer, 6 1-62 photoprotection, 62 principle function, 58-59 chlorophylls chlorophyll a, 49-5 1 chlorophyll b, 51 chlorophyll c , and c2, 52-53 chlorophyll d, 53-54 distribution in algal groups, 2 6 2 7 phycobiliproteins, 63-68 quantum efficiency, 73-75 Photosynthetic pigment-protein complexes carotenoid-protein complexes fucoxanthin-containing complexes, 120-129 peridinin-chlorophylla-protein,1 18120 chlorophyll-protein complexes chlorophyll a/b-protein, 105-107 chlorophyll a/b-siphonaxanthin, 108- 109 LHCP and adjuvant lipids, 107 solubilization and fractionation, 104-105 types of complex, 103- 104 phycobilisomes and biliprotein aggregates discovery, 109

303

fluorescence, 1 12-1 15 interactions with thylakoids, 115-1 18 structure, 109-1 12 Photosystem reaction centre complexes analysis of chlorophyll proteins curve deconvolution, 9 6 9 8 difference absorption spectra, 98-99 fluorescence spectra, 99-102 distribution of excitation energy, 143-149 PSI antenna cells, 8 4 8 5 chlorophyll of P700, 83-84 molecular weight, 85-86 primary electron acceptor, 8 6 8 8 properties, 7 6 8 2 PSI1 general properties, 88-90 polypeptides, 9&91 primary electron donor and acceptors, 91-94 size of antenna and reaction centres, 9495 Phycobilins evolution, 178 structure, 63 Ph y cobilisomes and biliprotein aggregates, 109-1 18 mechanism of excitation transfer, 133- 1 34 occurrence in Cryptophyta, 35 Cyanobacteria, 32 Rhodophyta, 35 shading effects, 16&161 structure, 134, 139 Phycocyanibilin, distribution, 63, 64 Phycoerythrobilin, distribution, 63, 64 Phyllosporum, chlorophyll a/c ratio, 121 P. comosa light-harvesting complex, 124 photosystem reaction centre complex, 90 Phytoalexins and nutrient status, 242, 26 I , 266, 267 Phyrophthora copper and iron effects on germination, 255 molybdenum and zoosporangia formation, 265 potassium and host infection, 225

304

SUBJECT INDEX

P . cinnamoni, effect of micronutrients on infectivity, 258 P. drechsleria, effect of micronutrients on infectivity, 258 Phytoplankton blooms and irradiance, 14, 15 buoyancy control, 42 light attenuation, 12-14 package effect, 46 thylakoid structure, 33, 35 Piricuiaria oryzue copper effect on infection, 231 glutamine and spore germination, 226 silica and infection, 261 Pisatin production and nickel, 261 Pisum sativum, curve deconvolution analysis, 96 Plasmodiophora brassicae, effect of boron and calcium on host infection, 241 Plastocyanin, electron donor to P700, 87-88 Polyoxyethylene sorbitan, differential solubilization of thylakoids, 104 Polyphenol oxidase and nutrient status, 233, 236, 238-239, 241, 261 Porphyra perforata action spectra for photosynthesis, 71 fluorescence spectra, 101 Porphyridium chlorophyll of P700, 83 difference absorption spectra, 98 P. cruentum antenna chlorophyll, 94 chromatic adaptation, 171 fluorescence from phycobiliproteins, 114, 117, 133 phycobilisomes, 109, 1 1 1, 1 I2 quantum efficiency spectra, 74, 75 shading effects, 156 spillover, 146 P. sordidum, phycobilisomes, 1 I2 Potassium deficiency and plant adaptation, 222-223 involvement in plant disease, 224225 Procentrum micans, thylakoid structure, 34 Prochioron algal evolution, 183-184, 185, 188 thylakoid membrane, 32, 33 Prosthecochloris aestuarii chlorophyll-protein structure, 132-1 33

P700 complexes, 84 Pseudoanabaena chromatic adaptation, 166 phycobilisomes, 112 Pseudomonas syringae, host infection and nitrogen supply, 227-228 Puccinia coronata, zinc concentration and vesicle formation, 258 P. graminis, iron concentration and host resistance, 255 P . triticinu copper concentration and host infection, 231 lithium concentration and host infection, 264 Pyrocystis, chloroplast movement, 29

Q

Quantum efficiency spectra in algae, 73-75

R Rhizoctonia potassium concentration and infection, 225 resistance mechanism, 253 Rhodomonas lens, action spectrum of photosynthesis, 70 Rhodophyta action spectra, 69 distribution and light quality, 23, 24 phycobiliproteins, 64 thylakoid arrangement, 34, 35 Rhodopseudomonas spheroides, configuration of carotenoid component, 61 Rhodospirilium molischianum, thylakoid membrane, 31 R. robrum, configuration of carotenoid component, 60-61 Ribulase bisphosphate carboxylase (RuBPc’ase) nitrogen level, 41 photosynthetic bacteria, 31 radiation level, U 1 shading effect, 154-1 55 RNA synthesis and zinc deficiency, 259 Ruderal algae and light harvesting, 21-22 S Sargassum light-harvesting complex, 124

SUBJECT INDEX

photosystem centre complexes, 80 Scenedesmus B-carotene, 58-59 chlorophyll, 52, 83 electron donor to P700, 88 light attenuation and the package effect, 4&47 photosystem reaction centre complex, 77 quantum efficiency, 152 RuBPc’ase, 41 S. obliquus carotene-polypeptide, 59 photosynthetic rate, 151 photosystem reaction centre complexes, 78, 87, 90, 95 shading effects, 156, 158, 160 Sclerotinia, copper level and infection of peanuts, 231 Sclerotium, calcium level and host infection, 227 S. rolfii, boron nutrition and disease resistance, 240 Scytosiphon light-harvesting complex, 124 photosystem reaction centre complexes, 80 Shade algae and light harvesting, 21-22, 155, 165 Siderophore production and iron concentration, 256, 257, 266267 Silicon concentration and disease, 261-263 Sinapsis alba, chlorophyll-protein complex, 107 Siphonaxanthin chlorophyll-protein complex, 108--109 distribution in algal groups, 5 6 5 7 light absorption, 59, 72 structure, 55 Skeletonema costatum antenna chlorophyll, 94 chlorophyll c, 122 photosynthetic rate, 151 shading effect, 157, 160, 162 Sodium dodecyl sulphate, solubilization of chlorophyll-protein complex, 104 Spaeropsis malorum, iron concentration and host infection, 255 Spartina alternijlora, chlorophyll alb ratio, 52

305

Sphaceluria, shading effects, 157 Spirodela oligorrhiza, photosystem electron acceptor, 91 Spirulina, difference absorption spectra, 98 S. maxima, isolation of caroteno-protein, 60 Spirulina platensis photosystem reaction centre complexes, 81 phycobiliprotein structure, 66 shading effects, 163 Streptomyces scabies lime and host infection, 227 manganese and disease development, 243, 244247, 251, 252 Sulphur level and disease, 228, 244 Superoxide dismutase in the evolution of photosynthesis, 176 Syctotharnnus australis, light harvesting complex, 124 Synchytrium endobioticum, boron nutrition and disease resistance, 240 Synechococcus glutamate pathway, 183 photosystem reaction centre complex, 79, 8 1, 90 phycobiliprotein, 41, 66 phycobilisome structure, 11 1-1 12, 181 Synechocystis chromatic adaptation, 166 phycobilisome structure, I10

T Take-all, see Gaeumannomyces graminis Thylakoid arrangement of pigment-proteins, 135-1 36, 138-143 evolution of stacking, 187-188 in algal groups, 2 6 2 7 , 32, 34, 35 in photosynthetic bacteria, 31-32, 33, 34 membrane electron transport, 19, 3 8 4 0 fractionation, 104 light harvesting, 20-2 1 light scattering, 48 phycobilisomes, 109-1 12, 115-118 PSU, 38 Tolypothrix tenuis, chromatic adaptation, 166, 174

306

SUBJECT INDEX

Triton X-100 isolation of phycobilisomes, 109 solubilization of chlorophyll-protein, 104, 106, 124, 126 Tween, see polyoxyethylene sorbitan U

Ultraviolet irradiance and algal photosynthesis, 12, 36 Ulva japonica, light harvesting by siphonaxanthin, 72 U.lactuca chloroplasts and light harvesting change in morphology, 30 movement, 29 photosynthetic rate, 168 U.taeniata action spectrum for photosynthesis, 70 photosynthetic rate, 168

V

Vaucheria, chlorophyll c, 52, 120 Viloxanthin distribution in algal groups, 56-57, 59 structure. 55

X

Xanthophylls in algae, 55, 59 Z "Z" scheme of photosynthetic electron transport, 17, 18, 36

Zinc, effect on plant disease, 257-260 Zooxanthellae, photosynthetic rate, 151 Zwittergen, solubilization of chlorophyllprotein complexes, 104

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

No title

Recommend Documents