The Use of the Oxygen Electrode and Fluorescence Probes in Simple Measurements of Photosynthesis

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Robert Hill Institute

University of Sheffield

THE USE OF THE OXYGEN ELECTRODE AND FLUORESCENCE PROBES IN SIMPLE MEASUREMENTS OF PHOTOSYNTHESIS

By David Walker with illustrations by Richard Walker

Robert Hill Institute, The University of Sheffield, SHEFFIELD, S10 2TN, U.K.

“Where we might soon have fainted in that Enchanted Ground but now and then a cluster of pleasant grapes we found”

PREFACE

Like most biologists of my generation, I was raised on the measurement of gaseous exchange by Warburg manometry and indeed my first research of any consequence involved an elegant Warburg apparatus which, in those days (1953), dominated Harry Beevers’ laboratory in the “Peirce Conservatory and Small Animal House” of Purdue University. Here everything was a masterpiece of time and motion but even at its best, manometry was scarcely a technique which was likely to win friends. Calibration was frightful, taking measurements tedious, and cleaning Warburg flasks a pain. Happily, after Purdue and Newcastle my research took me away from manometry and it was not until 1962 that I was obliged to consider using it again. By then (with Bill Cockburn, Carl Baldry and Chris Bucke), chloroplasts had been isolated from peas and spinach which could fix 14CO2 at rates approaching those of the parent tissue. CO2-dependent O2 evolution (except for a report by Amen and Whatley of very slow rates under anaerobic conditions) had not, however, been demonstrated. At that time I was at Imperial College, travelling one day a week to the Biochemistry Department at Cambridge to continue a long-standing co-operation with Robin Hill. It was he who introduced me to the Clark-type O2 electrode. Soon after we were able to purchase a Gilson electrode at Imperial College and, immediately it was in operation, to show that our CO2-fixing chloroplasts did indeed evolve O2 at equally fast rates.

The switch to polarography was both a delight and a challenge. At one stroke we were freed of the need to base assays on 14C samples and introduced to the pleasures of continuous measurement. The advantages of using isotopes were not lost because we could also add or withdraw samples, at will, during an. experiment. Suddenly, manometry seemed to be as old fashioned as Pettenkofer tubes. Oxygen electrodes, however, proved to be a mixed blessing. Those that we could afford were less than perfect and not really suited to our needs. It was for this reason that Tom Delieu and I tried to come up with something better. We have been trying ever since. In spare moments I make crude pencil drawings. Tom somehow contrives to convert these into material reality. Very often my ideas lack feasibility. Always progress has been painfully slow. Until recently, electrode design and construction was largely a spare-time occupation. Each prototype has had to earn its living in our day-to-day work Nevertheless, by the late 60’s we had a Clark-type electrode which has served us very well. Since 1974 it has been produced commercially by Hansatech. More recently Tom Delieu joined the A.F.R.C. staff in what is now “The Robert Hill Institute” in the University of Sheffield. Within the Institute one of our aims is to develop new (and, ideally, simpler) means of measuring photosynthesis.

The main purpose of this book is to describe some of the equipment which Tom Delieu and I have developed (often with the help of colleagues in Sheffield and elsewhere). Much of this apparatus is now available commercially so that anyone who has a mind to do so can put together experimental systems very similar to the ones described here. Clearly I am not addressing those who are already familiar with such apparatus, or those from whom we ourselves learn on an almost daily basis. On the contrary, this was originally intended to be a beginner’s guide but what started its existence as an introductory manual, largely restricted to the leaf-disc electrode, has grown in a way which has threatened the simplicity of the original concept. Some of the more complex phenomena (such as oscillations) which can be studied with the help of the leafdisc electrode, are barely understood and difficult to explain. New ways of analysing fluorescence signals have been devised and could not be neglected. There are classic experiments, like the Hill Reaction, which will continue to be an important part of practical teaching into the foreseeable future and for this and other reasons a section on aqueous-phase measurements has been added. Inevitably, there will be errors and incorrect assumptions. In some places I have probably erred on the side of over-simplification and, in others, complexity may have been compounded by brevity. There is no doubt, however, that polarographic measurement of oxygen has much to offer those involved in photosynthesis, whether in teaching or research, and that there is advantage to be derived from simultaneous measurements of chlorophyll a fluorescence. It is hoped that what follows will constitute a useful practical introduction to these topics.

David Walker, January 1987.

Second Edition, Second Impression In the second impression we have endeavored to correct errors which slipped through in the first impression and have taken the opportunity to incorporate information concerning the “Leafdisc” software as an appendix to the main text. We have also added a laminated cover which we trust will be as attractive its predecessor and more hard-wearing.

David Walker, March 1990.

Acknowledgements: To Tom Delieu who by translating inadequate pencil drawings of apparatus into splendid mechanical reality made the whole thing possible, to Rick Walker who illuminated an otherwise dull text, to Richard Leegood, Uli Schreiber and Mirta Sivak for invaluable advice, criticism and contributions to specific sections. To Barry Osmond for endless warm encouragement, to Christa Critchley, Robyin Cleland, Simon Robinson and Steve Powles for their participation in Australian measurements, to Ross Lilley for an excursion, with the apparatus, into a rain forest. To Margaret and John Humby. David Thody and Richard Poole (of Hansatech) for continuing and fruitful cooperation. To John McAuley for his contribution to the first computer software and to George Seaton who has taken it to its present state of sophistication, to Anne Dawson, Jackie Rowed and Diane Wright for their work on the manuscript and to Iris Walkland for the index. To Marney and Shirley Walker for grapes. Much of the earlier work on electrodes was supported by grants from the Royal Society. Most recently, work on the automated leaf-disc electrode system has been supported by the Society’s Paul Instrument Fund.

My colleague and dear friend, Tom Delieu, died after a brief illness, on Nov 12 1987 aged 58. Tom was a legendary figure to everyone who knew him - a master craftsman who could make almost anything out of almost anything; a man of great warmth and humour who will be sorely missed not only by his wife and his family but by his friends and colleagues the world over.

CONTENTS

PART (A) OXYGEN 1 1. INTRODUCTION 1 2. THE PRINCIPLE OF OXYGEN MEASUREMENT 1 3. ASSEMBLY 4

4. CALIBRATION

5

(a) Experiment 1. Does it work? Checks for function and the determination of values required for calibration (b) Calculations relating to volume and calibration (c) Introducing Carbon Dioxide (d) Experiment 2. Does it work with a leaf? (e) Quick Calibration

5. EXPRESSION OF RATES 13

Part (B) FLUORESCENCE 17 6. INTRODUCTION 17 7. PRINCIPLE OF MEASUREMENT 18 8. IS FLUORESCENCE INFLUENCED BY CARBON ASSIMILATION? 19 9. FLUORESCENCE INDUCTION 22 10. INDUCTION 24 11. THE FLUORESCENCE “M-PEAK” 25 12. COMPLEX FLUORESCENCE KINETICS AND OSCILLATIONS 27 (a) Gas Transients (b) Experiment 3 (c) Oscillations (d) Factors which Favour Oscillations

13. QUENCHING ANALYSIS 35 (a) Introduction (b) The “DCMU Method” (c) Light-scattering (d) “Light-doubling” (e) The Pulse-Saturation Method (f) Modulated Light (g) The Procedure (h) The Analysis (i) Examples

Part (C) LIGHT SOURCES 47 14. (a) Slide-Projectors and Heat Filters (b) A High Intensity Light Source (c) Light-Emitting Diodes (d) The Bjorkman Lamp

Part (D) MAXIMUM EFFICIENCY OF PHOTOSYNTHESIS 51 15. INTRODUCTION 51 16. MEASUREMENT OF QUANTUM YIELD 52 17. THEORETICAL BACKGROUND 53

18. MEASUREMENT OF APPARENT QUANTUM YIELD WITH THE LEAF DISC ELECTRODE AND THE BJORKMAN LAMP 54 19. DOING IT IN OTHER WAYS 56 (a) According to Bjorkman and Demmig (i) Introduction (ii) Photon Yield Measurements (iii) Leaf Absorptance Measurements (iv) Further Considerations (b) Doing it with Chloroplasts and Algae (c) Doing it in the Dark

Part (E) COMPUTERISED MEASUREMENT OF RATE AS A FUNCTION OF PHOTON FLUX DENSITY 63 20. UNDERLYING PRINCIPLES 63 (a) Why do it at all? (b) Roots and Ceilings (c) The Roof (d) The Ceiling (e) How Steep is the Roof? (f) Relative and Absolute Quantum yield (g) Why Trouble to Automate Measurement? (h) How Does it Work? (i) What Are The Drawbacks? (j) What Are The Advantages?

21. MEASURING RATE AS A FUNCTION OF PFD 72 (a) Choice of CO2 Concentration etc. (b) The Unsteady State (c) The Closed Refrigerator Door Conundrum

22. LIGHT ENHANCED RESPIRATION 73 23. THE “KOK EFFECT” 74 24. IS THERE A RECOMMENDED PROCEDURE? 76 25. HOW TO GET THE MOST OUT OF YOUR DATA 78 26. RATE V PFD UNGILDED 78 27. THE INITIAL SLOPE 79 28. THE SHAPE OF THE CURVE 80 29. ANALYSIS OF STRESS 81 (a) Stress and Performance (b) Shifting the Arbitrary Ceiling (c) Shifting the Ceiling and the Perpendicular

30. LUX ET VERITAS - A REMINDER ABOUT LIGHT 83 (a) Measurement (b) The Nature of Light (c) The Energy Content of Light (d) Can We Compare Micromoles with Foot Candles? (e) Energy, Plants and Man - an Aside About Energy Utilisation

(d) Experiment 3. CO2 and PGA-dependent O2 evolution (e) Experiment 4. The Requirement for Orthophosphate (f) Experiment 5. Orthophosphate Inhibition and its Reversal (g) Inorganic Pyrophosphate (h) Experiment 6. Carbon Dioxide Dependence

42. PROTOPLASTS 136 (a) Digestion (b) Pretreatment (c) Enzymes and Incubation (d) Isolation (e)Purification

43. CHLOROPLASTS FROM PROTOPLASTS 140 44. THE RECONSTITUTED CHLOROPLAST SYSTEM 141 (a) The Simplest System (b) Systems with Additional Stroma

45. EXPERIMENTS WITH THE RECONSTITUTED SYSTEM 142 (a) Experiment 1. CO2-dependent O2 evolution (b) Experiment 2. PGA-dependent O2 evolution (C) Experiment 3. Uncoupling

46. SOME FURTHER PRACTICAL CONSIDERATIONS 146 (a) Twin Electrodes (b) Illumination

Part (G) DEFECTS AND PRECAUTIONS 152 47. (a) The Sensor (b) Use (c) Membrane Application (d) The Leafdisc Electrode (e) Leaks (f) Membranes and Electrolytes

APPENDIX 1 PREPARATION OF FERREDOXIN 155 APPENDIX 2 SUPPLIERS 157 APPENDIX 3 LEAFDISC(CONTENTS) 160 LEAFDISC (TEXT) 163 APPENDIX 4 LATEST HANSATECH EQUIPMENT 197 INDEX 198

ABREVIATIONS

ADP ATP BSA C4 C3 CAM CF0 CF1 Chi CP CPK DBMIB DCMU DHAP DPGA EDTA E4P F6P FAD FBP FMN F26BP G1P C3P G6P GBP HEPES hv IRGA KCN

adenosine diphosphate adenosine triphosphate bovine serum albumin plants having the C4 dicarboxylic acid pathway plants fixing atmospheric CO2 directly through the RPP-pathway Crassulacean Acid Metabolism proton channel of coupling factor ATP synthesising component of coupling factor chlorophyll creatine phosphate creatine phosphate kinase dibromothymoquinone-2,5-dibromo-3-methyl isopropyl-p-benzoquinone 3-(3’ ,4’-dichlorophenyl)-l,l-dimethylurea dihydroxyacetone phosphate 1,3-diphosphogylcerate ethylenediaminetetrascetic acid erythrose 4-phosphate fructose 6-phosphate flavin adenine dinucleotide fructose 1,6-bisphosphate flavin mononucleotide fructose 2,6-bisphosphate glucose l-phosphate glyceraldehyde 3-phosphate glucose 6-phosphate glycerate 1,3-bisphosphate N-2-hydroxyethylpiperazine-N’-2-ethanesulfonic acid light (Planck’s constant x frequency of radiation) infra-red gas analysis potassium cyanide

LED mV NADP NADPW nm P680 P700 PAR PC PFD PGA, 3PGA Pi PMF PMS PPI PQ PS1, PS2 PSI, PSII PT Q QA QB QE qQ R5P RPPP Ru5P RuBP,RBP S7P SBP Xu5P

light emitting diode millivolts nicotinamide adenine dinucleotide phosphate (oxidised) nicotinamide adenine dinucleotide phosphate (reduced) nanometre, 10-9 metres reaction centre of PS2 - with an absorption maximum at 680 nm reaction centre of PS1 - with an absorption maximum at 700 nm photosynthetically active radiation plastocyanin photon flux density 3-phosphoglycerate orthophosphate, inorganic phosphate proton motive force N-methylphenazonium methosulfate pyrophosphate plastoquinone photosystem I and 2 photosystem I and II phosphate translocator “Quencher” primary electron acceptor of PSII secondary electron acceptor of PSII high energy quenching photochemical quenching ribose 5-phosphate reductive pentose phosphate pathway, Benson-Calvin cycle ribulose 5-phosphate ribulose 1,5-bisphosphate sedoheptulose 7-phosphate sedoheptulose 1,7-bisphosphate xylulose 5-phosphate

PART A

OXYGEN

1. INTRODUCTION The measurement of oxygen evolution in a closed system is one of the easiest and cheapest means of demonstrating, or following, the process of photosynthesis in a leaf. The account which follows is based on a device designed by Delieu and Walker (1981) and now manufactured by Hansatech (Appendix 2). As such it complements “Notes for Users” supplied with the Hansatech LD2. It includes descriptions of simple experiments which can be used in teaching or for gaining experience with the apparatus. Little or no knowledge of photosynthesis, or of oxygen measurement by polarography, is assumed .

2. THE PRINCIPLE OF OXYGEN MEASUREMENT In photosynthesis, light energy is absorbed by chlorophyll and used to drive the reduction of carbon dioxide to carbohydrate. The major end-product of photosynthesis in higher (flowering) plants is usually sucrose. Starch is also often formed as a temporary storage product but both of these carbohydrates (sucrose and starch) are formed from three-carbon sugar derivatives. For simplicity, all of these substances can be represented by a purely nominal carbohydrate, CH20, and the overall process by the equation:-

“an easy way of following photosynthesis”

hv CO2 + H2O

CH2O + O2 ........Eqn. 2.1

in which the light energy needed to drive this process is represented by “hv” (h = Planck’s constant and the Greek letter “v” the symbol used to represent the frequency of light). Accordingly, if a leaf is enclosed in a chamber and provided with carbon dioxide (or bicarbonate as a source of carbon dioxide) and then illuminated, oxygen will be evolved. In the Hansatech LD2 (Fig. 2.1.), a leaf-disc is used and CO2 is provided in the gas-phase or in the form of sodium bicarbonate (which dissociates in solution):-

NaHCO3

NaOH + CO2

........Eqn. 2.2

The oxygen which accumulates in the gas-phase during photosynthesis is then detected, polarographically, by a “Clark-type electrode (Clark, 1956). The “regular” Hansatech version of the Clark-type electrode (Fig 2.1) comprises a relatively large (2mm) platinum cathode and a silver anode immersed in, and linked by, an

“a temporary storage product”

electrolyte. Both electrodes are set in a plastic (epoxy resin) disc; the cathode at the centre of a dome and the silver anode in a circular groove (the well, or electrolyte reservoir). The electrodes are protected by a thin teflon or polythene membrane which is permeable to oxygen and the purpose of the dome is to stretch the membrane smoothly over the surface of the platinum cathode and to allow it to be secured in position by an O-ring. The membrane also traps a thin layer of electrolyte.

Fluorescence probe

LED SOURCE Top window

Water

Water in/out

O-rings Thin glaas window PP

Leaf disc well Air in

Air out Electrode disc

PP

Water

Water in/out

Figure 2.1. Schematic diagram of a gas-phase oxygen electrode and fluorescence probe. The leaf-disc, or leaf pieces are supported on a stainless steel mesh in a chamber which is located in the middle section of the apparatus. The O2 sensor (Clark-type electrode) lies beneath the leaf chamber with its Pt cathode exposed to the atmosphere within it. The leaf tissue is pressed lightly against the temperature-controlled roof of the chamber by a foam disc which also separates it from carbonate/bicarbonate buffer carried on capillary matting. The leaf is illuminated through this window which also allows fluorescence to reach a probe (inserted at an angle of 40 degrees) where it is monitored by a photodiode. Actinic light is delivered to the top of the apparatus, by an array of light-emitting diodes, as shown, or from an appropriate light source such as the Hansatech LS1 or LS2. The fluorescence probe is a photodiode protected from the actinic light by optical filter or filters. The clips which draw the top section on to the middle section (so that the roof of the leaf chamber is sealed against an O-ring) are not shown. The taps (with luers) are for calibration and adjustment of the gas phase (after Delieu and Walker, 1983).

O2 Membrane Cathode (Pt) O2+H2O+2eH2O2+2OHH2O2+2e2OH-

KCl bridge KCl solution in well

-

+

0.6 - 0.7v

4e-

+ + + + + Ag (anode) 4Ag 4Ag+ + 4e4Ag+ + 4Cl4AgCl

Figure 23. Diagrammatic representation of oxygen electrode reactions. When a potentiating voltage is applied across the two electrodes, the platinum (Pt) becomes negative (i.e. becomes the cathode), and the silver (Ag) becomes positive (the anode). Oxygen diffuses through the membrane and is reduced at the cathode surface so that a: current flows through the circuit (which is completed by a thin layer of KCl solution or other electrolyte). The silver is oxidised and silver chloride deposited on the anode. The current which is generated bears a direct, stoichiometric, relationship to the oxygen reduced and is usually recorded, as a voltage, on a pen-recorder. (a solution, which usually contains potassium chloride) over the surface of the electrodes. A paper “spacer” is usually placed beneath the membrane in order to provide a uniform layer of electrolyte between anode and cathode. When a small voltage is applied across these electrodes, so that the platinum is made negative with respect to the silver, the current which flows is at first negligible and the platinum becomes polarized (i.e. it adopts the externally applied potential). As this potential is increased to 600-700 mV, oxygen is reduced at the platinum surface, initially to hydrogen peroxide H2O2 (Fig. 2.2) so that the polarity tends to discharge as electrons are donated to oxygen (which acts as an electron acceptor). The current which then flows is stoichiometrically related to the oxygen consumed at the cathode. As indicated in Fig. 2.2, the electrolyte in the vicinity of the cathode rapidly becomes alkaline during oxygen measurements (because of the generation of hydroxyl ions in the cathode reaction) and carbon dioxide can permeate the membrane and dissolve in this solution causing “drift” in the electrical output. For this reason, electrolytes containing carbonate ions and buffers (Section 47d) are frequently used for gas-phase measurements in atmospheres containing high concentrations of CO2. These salts facilitate equilibration between the electrolyte and the gas-phase and enhance electrical stability. Electrolytes containing 95% ethylene glycol and 5% KCl resist desiccation and may be preferred. The electrical current generated by the reduction of oxygen at the cathode is converted to a voltage output signal by the Hansatech CB1 or CBI-D control box. (The latter also has a digital display of the output voltage). The output signal is sufficiently large to be monitored in the 1 volt range.

3. ASSEMBLY Take a clean electrode (cleaning is best accomplished by polishing with fine-grade aluminium oxide paste) and place a drop of electrolyte on the dome which bears the platinum cathode. Place a “spacer” approximately 2 cm square on to the drop of electrolyte. Nut

Spring Cylinder (1) add electrolyte

Membrane

Cone O-ring

Spacer o-ring membrane spacer

(2) fit membrane

(3) membrane fitted

check for smoothness

KCl soin in well Figure 3.1. The membrane applicator. This is used as follows. The nut is loosened so that the cone emerges from the cylinder, pushed by the spring. The O-ring is slipped over the cone and the nut tightened so that the cone retracts into the cylinder, pushing the O-ring downwards as it does so, until it comes to rest on the lip of the cone (as shown) in a groove made for this purpose. The applicator is then placed in the position shown and the cylinder pressed downwards. As the descending O-ring touches the membrane it tends to stretch it slightly and, as the cylinder continues to move downwards, it pushes the O-ring off the lip of the cone and over the dome so that the membrane is stretched smoothly over the surface of the dome. The membrane is held in this position as the O-ring contracts on to the neck. (The “spacer” is used to provide a uniform layer of electrolyte between the anode and the cathode. Cigarette paper, which is thin and uniform is often used for this purpose - Caution, avoid the gummed part!). A hole, cut in the spacer immediately above the cathode, heightens sensitivity but can lead to unevenness of the electrolyte film and, malfunction. Place a similarly sized piece of membrane on top of the’ spacer and secure the membrane and spacer in position, over the dome, with the O-ring provided for this purpose. The O-ring is best applied with an applicator (Fig. 3.1). In this case, the O-ring is first placed on the end of the applicator and the stem of the applicator is turned. This advances the stretched O-ring to the tip of the applicator. If the applicator is now held by its centre in a vertical position on top of the dome and pressed downwards, the O-ring will be transferred to the dome. When the membrane is in position it should be checked for smoothness. If it is greatly wrinkled it should be rejected and the procedure repeated. At this stage, you may wish to connect the electrode to its potentiating circuit and the recorder. If you breathe directly an to the electrode you should see an excursion in the recorded signal and you will know that the electrode is functioning.

Disconnect the electrode and place it, dome upwards, on the bottom section of the chamber (Fig. 2.1). Place the central section of the assembly over the electrode and thread it on to the base. Avoid over-tightening because this can cause damage to the membrane. Finally, place the top section of the chamber into position and secure it, in place, by tightening the clips. Reconnect the electrode to its potentiating circuit. The chamber is normally used with temperature-controlled water circulating through the top and bottom sections and this should be examined to ensure that there are no air bubbles trapped in these compartments (tilt each section, starting with the base, so that the water exit assumes a vertical position and repeat until any bubbles are dislodged). You should ensure, by using filtration if necessary, that the circulating water is clean. You may wish to include a “water wheel” or similar device in your plumbing (on the exit side) so that you can see, at a glance, that water is flowing freely at a suitable rate.

4. CALIBRATION The electrochemical reactions generate a minute ‘residual current’ even in the absence of oxygen and if accurate calibration is required the discrepancy between zero oxygen and the electrical zero should be identified by passing nitrogen over the electrode. Alternatively a small drop of dithionite solution may be placed on the cathode. This consumes O2 according to the equation Na2S2O4 + O2 + H2O

NaHSO4 + NaHSO3 .....Eqn. 4.1

and, since the reaction goes virtually to completion, this procedure is equivalent to flushing with oxygen-free gas. Unfortunately, dithionite is not pleasant to use (CAUTION, DITHIONITE IS CORROSIVE) and it can also cause damage to, or poisoning of, the membrane. For these reasons it is best avoided. If there is no alternative, it should be removed from the membrane as quickly as possible by using an aspirator with a soft tip (to prevent mechanical damage). In this way, the true zero is established and, if preferred, this can be “backed-off” to coincide with electrical zero. Once the dithionite solution is removed (or N2 flushing is stopped), and the cathode is exposed to air, the signal generated by 21% oxygen (the oxygen content of air) can be determined. In a closed system the amount of O2 is governed by the partial pressure of O2 and the volume of the chamber. Thus 1 ml (1000 µl) of air will contain 210 µl of oxygen. Similarly, if 1 ml of the space, in a 5 ml chamber, is occupied by a leaf-disc the increase in signal which follows the introduction of a given volume of air into the chamber will be correspondingly greater. In fact, the difference between the two signals can be used to determine the volume of a leaf-disc with some accuracy (4b and Fig. 4.2). In general, however, although allowances can be made for differences in atmospheric pressure etc., the instrument was designed for simplicity rather than absolute measurement of oxygen. Although it has proved to be much more versatile than was originally imagined it was originally intended for the measurement of the oxygen evolved when a relatively large piece of leaf was illuminated in a relatively small chamber. For most purposes, therefore, the excursions following the introduction (or

“flushing with O2- free gas”

removal) of small volumes of air into the chamber (using a gas-tight syringe) can be taken as the basis of calibration and all that remains to be done is some simple arithmetic (see Fig. 4b). Experiment 1 incorporates several aspects of the calibration procedure but this can be shortened (4e) or extended according to need and the degree of accuracy sought.

4(a)Experiment l. Does it work?

a large leaf and a small chamber

Using a 1 ml gas-tight syringe, introduce successive 200 µl aliquots of air into the leaf-disc chamber through one of the two gas-vents. (The other should be kept closed during this procedure). As each volume of air is pushed into the chamber the reading on the pen-recorder should rise quickly to a new level and stay there (Fig. 4.1). You will see that the initial rise is fast but that it decreases as it approaches its final level. This is because the rate of diffusion of oxygen to the detector will become limiting as the difference in concentration between the atmosphere in the chamber and that at the electrode surface approaches equilibrium. If equilibrium is not approached within about one minute (i.e. if the response is sluggish) the electrode may require cleaning. If the signal rises rapidly but then falls equally quickly, the chamber is leaking and you should check to see that you have remembered to close the other gas-tap, that the electrode disc is pressed securely against its O-ring, that the O-ring which seals the top of the chamber to the bottom is in place and that the clips which hold the two parts of the chamber together are properly closed. If the response seems too slow for the experiments that you have in mind, it can be speeded by using a thinner membrane, by cutting a hole in the spacer immediately above the cathode or by dispensing with the spacer altogether. If no spacer is used, departure from linearity may sometimes result (see below).

3.70

Changes in signal (mV)

0.6 0.5

3.56

0.4

3.403

3.435

0.3 3.31

3.315

0.2 3.202 3.20

0.1 3.100

3.095

1 2 3 4 5 6 7 8 9 10 11 12 13 14 Time (min) Figure 4.1. Excursions produced by introducing and removing aliquots of air.

A sluggish response

A 1 ml, gas-tight, syringe, with its plunger fully withdrawn (to the 1 ml position) was attached to an open tap on the leaf chamber and the other tap closed. The figure illustrates the responses which were observed as the plunger was then pressed inwards in 200 µl stages and finally withdrawn again in a similar fashion. Note that each successive addition produced a larger signal (for explanation see text).

(N.B.) Care should be taken to avoid over-tightening the threaded base of the chamber which pushes the electrode disc against its sealing O-ring. Once an effective seal is established, excessive tightening can constrict the layer of electrolyte between the anode and the cathode, leading to loss of linearity of response. It should also be noted that if this procedure is followed with a leaf already in position in the chamber, the reading will decline with time because of dark respiration. Even with an empty and properly sealed chamber there will be a small decline with time as oxygen in the chamber is consumed in the cathode reaction. In normal use this will be negligible but the actual size of all signals will, of course, vary with the degree of pre-amplification used and the channel on the pen-recorder which you have selected. If you suspect a slight leak, subject the chamber to negative pressure by attaching an empty syringe to the chamber and withdrawing aliquots of air. Clearly the excursions of the pen-recorder will now be negative and a leak will be indicated by an increase in signal rather than the converse.

Subject to negative pressure

You may note, and possibly be alarmed by, the fact that there is an increase in the signal caused by each successive aliquot of added air. On reflection, however, you will correctly conclude that this is to be expected because if you attach a 1 ml syringe to the chamber you effectively increase its volume by 1 ml and every time you push the plunger further into the barrel of the syringe this effective volume becomes smaller. Accordingly, 200 µl added to, say, 4 ml will cause a larger excursion than if the same volume were added to 5. In normal use, the electrode response is linear because oxygen is being consumed or produced within a fixed volume rather than one which changes in relation to the position of the plunger in the syringe. You may use the data that you have derived from Experiment 1 to plot relationships similar to those illustrated in Fig. 4.2. This, or individual values, can also be used to calculate the effective volume of the chamber and therefore to relate the electrical output on the chart to the oxygen evolved by a leaf (4b). 4(b) Calculations relating to volume and calibration At s.t.p. (standard temperature and pressure) the amount of oxygen in 1 ml of air (containing 21% by volume) is 210 µl and, since 1 µmole of gas occupies 22.414 µl, this is equivalent to 210/22.414 or 9.37 µmoles. At any other temperature (T), the amount can be derived by multiplying by 273/(273 + T) so that, for example, at 20oC the corresponding value would be 8.73 µmole per ml. Because the oxygen electrode measures concentration (or, more strictly, activity) it is also necessary, for some purposes, to know the effective volume of the chamber itself. This, of course, is variable because the leaf-disc and any other material enclosed within the fixed volume of the chamber will occupy a significant space. However, the increase in signal (i.e. the increase in the electrical output) from the electrode circuit can be used to calculate both the effective volume of the chamber and the linearity of the electrode response. Suppose, for example, that the initial electrode output (R1) in Experiment 1. was 1.5 mV and that, after the plunger was fully depressed, this reading

“each successive increase produced a larger signal”

had increased to 1.8 mV(R2). The effective volume (v) of the chamber can be calculated from the relationship:v=

R1 R2 - R1

=

1.5 1.8 - 1.5

= 5 ml

...Eqn. 4.2

but this only applies when 1 ml of air is introduced. The general form of this equation:-

(a) 5.0 Chamber volume (ml)

if sufficiently large excursions result

v= R1 x [R2 x (1 - volume injected)] ...Eqn. 4.3 R2 - R1 can also be used provided that sufficiently large excursions result from the introduction of volumes of air smaller than 1 mi. This has been done in Fig. 4.2 (using the data from Fig. 4.1). Similar data, derived from the excursions which were observed when this procedure was repeated after the insertion of plastic discs of 1 ml volume into the chamber, has also been plotted in Fig. 4.2. This provides a measure of the degree of accuracy which can be expected and also illustrates the way in which the chamber can be used to measure the volume of any small body enclosed within it. Normally, of course this “small body” will be a leaf-disc. Moreover, it will be respiring and therefore consuming oxygen. The rate of oxygen consumption by a leaf-disc of 10 sq. cm area (the maximum that can be accommodated) will not, however, be so large that the calibration procedure will be invalidated but, clearly, an appropriate correction for O2 uptake should be applied. For most purposes, the respiratory loss during the excursion(s) can be ignored but, if greater accuracy is required, the decrease in signal due to respiratory uptake of oxygen (during the calibration procedure) can be measured and a correction applied.

(b) 4.0

(c) 3.0 200

400 600 800 1000 Air added (µl)

Figure 4.2. Data from Figure 4.1 re-plotted as a function of volume of air added. Additional readings were obtained following thee insertion of a body of 1 ml volume into the chamber (b) and after the insertion of a second body of equal volume (c). This indicates the degree of accuracy which can be expected and illustrates the use of the chamber in measuring volume (i.e. the experimental points permit the horizontals a, b and c to be drawn at 1 ml intervals, as would be expected).

Now, let us suppose that we had established that the volume of the leaf chamber was 5.17 ml, that the volume of the 10 sq. cm leaf-disc that we wished to use was 500 µl, that the sum total of all other inclusions (supporting grids, capillary matting etc.) was 1.17 ml and that the insertion of 1 ml of air at 20oC caused an excursion of 0.775 mV (as it might have done in the experiment illustrated in Fig. 4.1, in which the initial reading was 3.1 mV). We could then have calculated, from Equation 4.1, that the effective volume of the chamber was 4.0 ml. If we had inserted a leaf-disc with a volume of 500 µl and repeated this procedure we would have observed an excursion of 0.885 mV, from which, with the help of Equation 4.3 (above) we would have correctly calculated the volume of the leaf. We can also see, however, that we do not need to know the volume of the leaf itself for purposes of calibration since the size of the excursion must inevitably adjust to the effective volume of the chamber. Accordingly we may simply equate 8.73 µmoles of O2 in 1 ml of air at 20oC with a signal of 0.885 mV and say that 1.0 mV = 9.86 µmoles = approx 10 µmoles. The establishment of the relevant volumes does, however, allow us to calculate a more precise initial setting because each ml of otherwise unoccupied space in the leaf-chamber contains 8.73 µmoles of O2. We can, therefore, use Equation 4.3 to calculate a more desirable setting of the initial reading (“the air line”). For example, if the final effective volume of the chamber was 3.5 ml and we make the denominator (R2 - R1) = 0.873, then:R1 0.873 and R1 = 3.05 3.5 =

....Eqn. 4.4

Accordingly if the air-line were re-set, electrically, from 3.1 mV to 3.05 mV the relationship would be 1.0 mV = 10 µmoles of O2, precisely.

4(c) Introducing Carbon Dioxide Having assembled and calibrated the apparatus you are now presumably anxious to see that it will record oxygen evolution from an illuminated leaf. At this stage you must remember that photosynthesis requires CO2 and that air contains only about 350 parts per million. This means that the carbon dioxide content of the closed chamber will not sustain rapid photosynthesis for long. For example 4 ml of air will contain approximately 4 x 0.35 µl = 14 µl of CO2. A healthy spinach leaf would, under favourable conditions, have little difficulty in maintaining a rate of CO2 fixation in excess of 240 µmoles/mg chl/hr. This is equivalent to about 2 µmoles/min for a disc of 10 sq. cm area (containing, 500 µg chlorophyll). At this rate, the CO2 consumption would be 2 x 22.4 = 44.8 µl per minute - enough to deplete the chamber of CO2 in less that 20 seconds. Of course, this rate would not be sustained as the CO2 concentration fell but it points to the need for a CO2 source, if linear rates are to be maintained. There are also problems associated with too much CO2. Although these remain ill-defined it is clear that high concentrations of carbon dioxide may adversely affect many cellular processes and it is unlikely that photosynthesis will prove to be exceptional in this respect. Moreover, high concentrations of CO2 are known to induce stomatal closure. Nevertheless external concentrations in the region of 5%, or less, will

there is a certain comfort in being safely confused

apparently force CO2 through closed stomata because of the greatly steepened diffusion gradient. With photorespiration then largely, if not entirely, suppressed, the rate of consumption of CO2 within the chloroplast at these high external CO2 concentrations will, at least in high light, be so rapid that it will prevent CO2 concentrations reaching inhibitory levels and, if tire leaf is not left for long periods in the dark, will allow reasonable estimates of maximal rates of photosynthesis to be derived.

a “time-honoured method”

Gas mixtures can be generated from carbonate/bicarbonate buffers (which have been allowed, if so required, to equilibrate with N2) or by the time-honoured method of adding a non-volatile acid, such as sulphuric acid (H2SO4), to a known quantity of carbonate in a closed vessel of known volume. The CO2 concentration can then be checked by IRGA (infra-red gas analysis) or by titration and gas mixtures transferred to the chamber using a gas-tight syringe. If gas mixtures are available in cylinders (tanks) the chamber can be flushed, from this source, prior to use. Similarly, gases can be mixed using one of a number of commercial devices now available for this purpose (e.g. Wosthoff pumps, Signal blenders, mass-flow controllers). As will be seen below (Experiment 2), carbon dioxide can also be introduced into the leaf chamber by the simple expedient of breathing into it. Alternatively it can be generated within the chamber by adding a small volume (say 0.2 ml) of approximately molar sodium or potassium bicarbonate solution to capillary matting or coarse filter paper on the floor of the chamber. The leaf-disc is then prevented from contacting this solution by making a “sandwich” using the stainless-steel grids and foam (sponge) discs provided (see also 4e). Air-saturated water is about 10 µM with respect to CO2 and although the solubility of carbon dioxide is not affected by pH as such, an increasing amount of bicarbonate is built up as the pH is increased. Thus, at OoC and pH zero, virtually all of the CO2 will be present as dissolved gas. From pH 4.0 to 9.0, HCO3- increases from about 0.2% to about 96.5%, and CO3-- starts to appear at about pH 7.0 and increases to 97% at pH 12. These relationships are the basis of the carbonate/bicarbonate buffers which have been used for many years in Warburg manometry. They are less useful for the present purpose because, although at a given concentration, temperature and pH, each buffer is in equilibrium with a known partial pressure of carbon dioxide in the gas-phase, they can only be used with reasonable precision if the gas-phase is small or if the mixtures are pre-equilibrated with the appropriate gas mixture, thereby largely defeating the object of the exercise. Where less accuracy is required, pure bicarbonate solutions can be used instead (see below). One mole of carbon dioxide at standard temperature and pressure (s.t.p.)occupies 22.414 litres. At 25oC, it expands to All the larger space of 24.45 litres (V2 = V1.298/273). Accordingly, 1 millimole of CO2 (24.45 ml at 25oC) in 1 litre of gas will equal 2.445% and 1% CO2 in such a gas mixture would be 1/ 2.445 = 0.409 mM, or approximately 400 µM· At 25oC, in the concentration range of 1 to 100 mM, bicarbonate solutions equilibrate to a more or less constant pH of 8.37 and a similarly constant ratio of [HCO3-]/[CO2] = 90. Thus if n/400 = 90, n = 3,600 µM or 36 mM. This means that if 36.4 mM bicarbonate is left to equilibrate with an equal volume of air at 25oC the final

concentration will approximate to 1%. Equilibration can be speeded by the addition of a little phosphate ion (at the same pH) or carbonic anhydrase. If you are in a hurry and all you need is a shot of CO2, throw an excess of sodium bicarbonate into distilled water, shake in a half-filled closed flask and the atmosphere within the flask will soon contain about 5% CO2. 4(d). Experiment 2. Does it work with a leaf? An experiment of this nature is illustrated in Fig. 4.3. Cut a 10 sq. cm area disc from a broad leaf, such as spinach, with the cutter provided. To avoid dessication place it, on damp (not wet) capillary matting supported by the stainless-steel grid provided (use the grid with the unperforated centre which prevents light shining directly on to the cathode). This is best supported, in turn, by a sponge disc and a second stainless-steel grid. Close the chamber with the clips but leave both taps open. Attach a tube to the tap and breathe heavily into the chamber. (In so doing you are taking advantage of the fact that you are possibly a portable, and almost certainly a peripatetic, CO2-generator and O2-consumer). Close both taps. You should have succeeded in replacing air (21% O2, 0.035% CO2) with expired air (say 14% O2, 5% CO2).

Take a leaf

16 14

Fluorescence

12

Breath

10 8 6 4 Oxygen

2 5

10

15

20

Figure 4.3. Simultaneous recording of oxygen evolution and chlorophyll a fluorescence. Chart showing CO2- dependent O2 evolution by a spinach leaf and the changes in chlorophyll a fluorescence associated with the onset of illumination, the depletion of CO2 and the addition of CO2 following depletion. Note that the fluorescence signal should be shifted to the right by approx 0.6 of the distance between two vertical divisions in order to correct the displacement caused by the pens on the chart-recorder. Offset the signal on the pen-recorder using “back-off” until the trace occupies the lower 10% of the chart. Record the dark respiration for 2 or 3 minutes (until this reaches a steady O2 consumption as the temperature within the chamber adjusts to that of the circulated water).

“breathe heavily”

Switch on the light and allow the leaf to photosynthesise at high irradiance. If the leaf-disc has been taken from the dark or from low light you will observe an initial lag or induction period (Section 10). Thereafter, the rate should approach linearity and it illumination is interrupted by about 30 sec to 1 min darkness you should observe enhanced oxygen uptake and little or no lag following re-illumination. After some time, the rate of O2 evolution will decline (abruptly in C3 species, less abruptly in C4 species). If you then open both taps and displace the atmosphere in the chamber with more expired air you will observe an abrupt fall in the recorded reading as the oxygen content is returned to about 14% and (possibly after some temperature re-equilibration) a return to a higher rate of photosynthesis. If measured simultaneously, as in Fig. 4.3 the fluorescence signal (see Part B) will fall from an initial high, often in a fairly complex fashion, to a quasi steady-state value. In this experiment, as the CO2 within the chamber ran out, there was a change in the fluorescence signal reminiscent of the change seen in air to CO2-free air gas transients (Section 12a). Following the re-introduction of CO2, dampening oscillations in fluorescence were observed (see also Fig. 11.1 and Section 12b). 4(e). Quick Calibration Much of the above is intended to illustrate principles and to familiarise the user with the leaf-disc electrode (the LD2). If you are already one of the cognoscenti you may wish to proceed, with less ado, to the quickest possible calibration procedure. If so continue as follows:(i) Establish an electrically convenient “air-line” by closing the chamber and adjusting the recorder reading to some arbitrary value in the top half of the chart. Now establish the “N2 line” (Fig. 4.4) by flushing with nitrogen, or, if none is available, by using dithionite (Equation 4.1). If you must use dithionite, take care and remove it as quickly and as gently as possible. (ii) Place a leaf-disc on the stainless steel grid and sponge “sandwich” which presses the leaf against the temperature controlled roof of the chamber. Close the chamber, thus establishing a second arbitrary “air-line”. This will fall, with time, because of respiratory uptake of oxygen but the required values can be obtained, as indicated in Fig. 4.4, by extrapolation if the rest of the procedure is not delayed. (iii) Introduce air from a gas-tight syringe. This gives the value R2-R1 needed for the calculation (from Equations 4.2 and 4.3) of the effective volume of the chamber should you need it for other purposes, and the increment itself (Fig. 4.3) gives the equivalence between electrical output and lunges of oxygen generated. This, if you are impatient, is all that you need. “leaf-disc sandwhich” N.B. Grid with eintire centre should be beneath the first layer of capillary matting

If you wish to add a little elegance and the undoubted advantages that derive from running consecutive experiments on the same absolute scale, calculate the effective volume, multiply it by the number of micromoles of oxygen in this volume at the temperature being used, and adjust R1 so that the scale on the recorder gives a convenient direct “read-out”.

- leaf R -R 1

- matting - grid

2

air line

adjusted air-line - sponge - grid - sponge

R1

- matting N2 line

“leaf-disc sandwhich”

Figure 4.4 “Quick” calibration - for explanation see text.

5. EXPRESSION OF RATES Photosynthesis is often expressed as a rate of CO2 uptake per unit leaf area. This is convenient, although it can be very misleading if it is used as a basis for comparison. Thus two plants which display identical rates of photosynthesis per unit area, may grow (accumulate dry matter) at markedly different rates. This is readily understood if one is capable of diverting a fraction more of its photosynthetic product into new photosynthetic machinery than the other. On a “compound interest” basis this would soon account for the observed differences in weight. Even so, this sort of observation has led to statements such as “there is no clear correlation between photosynthesis and growth”. Since green plants grow (i.e. accumulate dry weight) almost entirely as a consequence of photosynthesis, such statements are self-evident nonsense. In the above example there is correlation, by definition, between photosynthesis per plans and growth. Provided, however, that we do not equate photosynthesis per unit area with “photosynthesis” per se we can proceed to express it on this basis. Our arithmetic is then made easier by the thoughtful designer who provided us with a tool which cuts a disc of precisely 10 sq. cm from broad leaved species. (An equivalent approximation can be made for some narrow-leaved species by attaching individual leaves, in rows to 2 strips of adhesive tape and then cutting one disc, comprising segments from several leaves. There are also more sophisticated commercially available devices for measuring the area of irregularly shaped leaves, or pieces of leaves). We are, however, immediately confounded by lack of agreement about units. The plant biochemist may prefer µmoles per mg chl per hr (µmoles. mg-1chl. hr-1) or (Appendix 3) µmoles. m-2. s-1 whereas

“there is correlation between photosynthesis per plant and growth”

the crop physiologist may go for mg CO2 per sq. decimeter per hour (mg.dm-2.hr-1). Suppressing a sigh, we can (if actual values are not available) arrive at some approximate equivalence by ignoring the differences between pale and dark green leaves and settling for an “average” chlorophyll content of 3 mg chlorophyll per sq. decimeter (i.e. 300 µg/leaf-disc). We must also remember that one mole of CO2 weighs 44 gm (i.e. that 1 µmole weighs 44 µg). Accordingly, the rate of photosynthesis by a 10 cm2 leafdisc which, contained 0.3 mg of chlorophyll and evolved O2 at 150 µmole/hr could be expressed as:150 = 500 µmoles.mg-1 chl. hr-1 ...Eqn. 5.1 0.3 or as 150 x 44 x 10 = 66mg. dm-2. hr 1000

...Eqn. 5.2

from which we can derive a conversion factor such that 1 µmole. mg-1chl. hr-1 = approx 0.13 mg. dm-2. hr-1 or 1 mg. dm-2. hr-l = approx 7.6 µmoles. mg-1. hr. It should be noted that we have used 44 (the molecular mass of CO2) rather than 32 (the corresponding value for O2) in the above conversion because the older values in the Literature were based on CO2 rather than O2. Since there is a 1 to 1 relationship between CO2 fixed and O2 evolved, at least in photosynthesis as described by the classic overall equation (Equation 2.1), we can take this liberty. We multiplied by 10 because 1dm = 1/10 metre = 10 cm and 1 dm2 =100 sq. cm, whereas our actual measurement was based on a disc of 10 sq. cm area. (We divided by 1000 in order to convert µg to mg). According to Zelitch, we can expect C4 crop species to give rates in the region of 3565 mg. dm-2, hr-1 (265-475 µmoles mg-1.chl. hr-1), C3 crop species to give 12-45 (91-335) and woody plants to be in the region of 10(75). These values were at high illuminance, in air, whereas our athletic spinach leaf which gave the convenient rate of 66(500) must have been provided with sufficiently high CO2 to suppress photorespiration and to overcome stomatal and boundary layer resistances. It should be noted that if the leaves contain more chlorophyll (say 0.05 mg/sq. cm rather than 0.03 mg/sq. cm) the values and the conversion factors based on chlorophyll would be decreased ‘ correspondingly (in this case by 3/5). To convert µmoles per disc per hr into µmoles. m-2.s-1 we are on somewhat safer ground with our oxygen measurements because an are recording the O2 evolved, in µmoles, from 10 sq. cm and there are 10,000 sq. cm in lm2. Accordingly, our (10 cm2) spinach leaf-disc which evolved O2 at 150 µmoles/hr, would give us a value of:150 x 10,000 = 41.66 µmoles.m-2.s-1 3,600 x 10

...Eqn. 5.3

and, in the general case, we would simply record the change (in µmoles) per min (a second is too short for accuracy) and multiply by 10 3/60 = 16.67 e.g. 150 x 16.67/60 = 41.66 µmoles m-2 .s-1

References

Photosynthesis in General Edwards, G.E. and Walker, D.A. (1983) C3, C4, Mechanisms, and Cellular and Environmental Regulation in Photosynthesis. Blackwell Scientific Publications Ltd, Oxford. 1-542. Hall, D.O. and Rao, K.K. (1987) Photosynthesis (4th ed.). Edward Arnold, London. pp 1-122. Rabinowitch, E.I. (1945 and 1951) Photosynthesis and Related Processes. Volumes 1 and 2. Inter Science New York, 1-2088. Walker, D.A. (1979) Energy, Plants & Man. Packard Publishing Ltd, Chichester. 1-31. Zelitch, I. (1971) Photosynthesis, Photorespiration and Plant Productivity. Academic Press, New York, 1-347

Polarography in General Aiba, S. and Huang, S.Y. (1969) Oxygen permeability and diffusivity in polymer membranes immersed in liquids. Chem. Eng. Sci. 24, 1149-1159. Benedek, A.A. and Heideger, W.J. (1970) Polarographic oxygen analyzer response: The effect of instrument lag in the non-steady state reaction test. Water Res. 4, 627-640. Berkenbosch, A. and Riedstra, J.W. (1963) Temperature effects in amperometric oxygen determinations with the Clark electrode. Acta Physiol. Pharmacol. Neerl. 12, 131-143,144-156. Clark, L.C. Jr (1956) Monitor and control of blood and tissue oxygen tension. Trans. Am. Soc. artif. internal Organs, 2, 41. Fatt, I. (1976) Polarographic oxygen sensors. CRC Press, Cleveland. 1-278 Gnaiger, E. and Forstner, H., eds. (1983) Polarographic Oxygen Sensors. Springer, Berlin, Heidelberg. 1-370. Hahn, C.E.W., Davis, A.H. and Albery, W.J. (1975) Electromechanical improvement of the performance of pO2 electrodes. Respir. Physiol. 25, 109-133. Hitchman, M.L. (1978) Measurement of dissolved oxygen. John Wiley & Sons, New York, pp. 255. Jensen, O.J., Jacobsen, T. and Thomsen, K. (1978) Membrane covered oxygen electrodes. I. Electrode dimensions and electrode sensitivity. J. Electroanal. Chem. 87, 203-211. Keidel, F.A. (1960) Coulometric analyser for trace quantities of oxygen. Ind. Eng. Chem 52, 490-493. Koltoff, I.M. and Lingane, J.J. (1952) Polarography, Vols. I and II. Interscience, New York and London. Lucerno, D.N. (1969) Design of membrane-covered polarographic gas detectors. Anal. Chem. 41, 613. Mergenhagen, D. Schweiger, H.G. (1973) Recording the oxygen production of a single Acetabularia Cell for a prolonged period. Exp. Cell Res. 81, 360-364. Oeseburg, B., Kwant, G., Schut, J.K. and Zijlstra, W.G. (1979) Measuring oxygen tension in biological systems by means of Clark-type polarographic electrodes. Proc. K. Ned. Akad. Wet Ser. C. 82, 83-90. Weast, R.C. (ed) (1969) Handbook of Chemistry and Physics 50th ed. Chemical Rubber Co., Cleveland, p151.

Delieu, T. and Walker, D.A. (1972) An improved cathode for the measurement of photosynthetic oxygen evolution by isolated chloroplasts. New Phytol 71, 201-225. Delieu, T. and Walker, D.A. (1981) Polarographic measurement of photosynthetic O2 evolution by leaf discs. New Phytol 89, 165-175. Delieu, T. and Walker, D.A. (1983) Simultaneous measurement of oxygen evolution and chlorophyll fluorescence from leaf pieces. Plant Physiol. 73, 534-541.

Never admit to working with spinach!

PART B

FLUORESCENCE

6. INTRODUCTION Chlorophyll looks green in white light because it absorbs light in the blue (round about 420 nm) and in the red parts (round about 660 nm) of the visible spectrum and transmits and reflects in the green. Light can be regarded as a stream of particles, or parcels, of energy. Each particle (quantum or photon) can bring about a single photochemical event provided that it carries sufficient energy to drive that specific event. Each quantum of red light which is absorbed by a chlorophyll molecule raises an electron from a ground state to an excited state and all of its energy is transferred in this process. This excitation is, essentially, an oxidation. Electron transport is initiated, as the electron is lifted into a higher energy orbital and a positively charged “hole” is left behind. Absorption of blue light causes even greater excitation (because of the higher energy content of the blue quantum) but the elevated electron then falls back into the “red orbital too quickly to permit useful chemical work. Thus, whatever the quality of the light absorbed, the electron reaches the same energy level more or less immediately after excitation and all subsequent events derive from this common starting point (“excited state one”). Chlorophylls a and b } Excited state (two)

H2C = CH H3C

Radiationless de-excitation

N

blue hv

red hv

Chemistry hv fluorescence }Ground state

Figure 6.1. The excitation of chlorophyll by light. The parallel lines represent energy sub-states or electronic orbitals. Thus the energy delivered by the absorption of a blue photon (left) is sufficient to raise on electron to “excited state two” from where it rapidly returns by a process of radiationless de-excitation, “cascading” through sub-states, to excited state “one”. A photon of red light (centre) only has enough energy to raise an electron to excited state “one” but this excited state is sufficiently stable to permit useful chemical work and is, in effect, the starting point of all other events in photosynthesis. “Excited state one” con also dissipate energy by re-emitting light as (deep red) fluorescence.

N

CH3(in a) OR - C (in b) O CH2CH3

Mg

} Excited state (one)

N N H3C H CH2 H C=O O CH2 OCH3 C=O O Phytyl

H

CH3

As indicated in Fig. 6.1 some of the excitation energy of chlorophyll is lost by radiationless de-excitation and some is used to drive the “chemical” reactions of photosynthesis (ATP synthesis, NADP reduction etc). Some energy is also dissipated as fluorescence. This, it should be emphasised, is not reflected or transmitted light. It is light which is created in the leaf, just as electron transport (electric current) through the filament of an incandescent lamp leads to the emission of light. Fluorescence (which takes about 10-9 sec to discharge) derives from the lower more persistent “excited state one” rather than “excited state two” which decays in about 10-13 sec. For this reason chlorophyll fluorescence is red, regardless of the quality of the exciting light and it is a deeper (longer wavelength) red than the red absorption peak because of the “Stokes shift” (the rapid cascade of heat dissipation which occurs within “excited state one”) so that electrons fall to the ground state from the lowest levels of excitation and must therefore give rise to photons of lower energy content (i.e. longer wavelength light).

7. PRINCIPLE OF MEASUREMENT

Pale

Dark

690 740 Fluorescence spectra of pale and dark leavesafter French and Young, 1952

The fraction of excitation energy which is dissipated as fluorescence in vivo is very small (3-5%). In solution this fraction is much larger (up to 30%) and if a solution of chlorophylls in ethanol, or acetone, is illuminated in a rectangular vessel it will look green from the front and deep red from the side. (Chlorophyll reflects in the red and the blue and transmits in the green but, viewed from the side, the retina is no longer flooded with green photons and the deep red fluorescence emanating from the excited chlorophyll is readily perceived). Photomultipliers are often used in light measurements but, in most circumstances, fluorescence from a leaf can be adequately detected with a photodiode. As in photosynthesis, each photon which falls on the detector initiates one photochemical event in which an electron is raised to a higher energy level and a positively charged “hole” is created. In photosynthesis, the “hole” accepts an electron from water (Figs. 8.1 and 8.2) and the electron is passed (via photosystem II and I) to NADP. In the photodiode, the corresponding electron transport (electric current) is amplified and can be applied, as a voltage, to a pen-recorder. The main problem is to prevent the detector “seeing” light which is not fluorescence and, inevitably, a relatively large fraction of the “actinic” light (the light used to drive photosynthesis) will be reflected from the leaf surface into the detector. Accordingly, the detector is protected by optical filters which, ideally, exclude all of the reflected actinic light and transmit all of the fluorescence. In practice a compromise is needed because in terms of “energy-effectiveness” (Section 6) red light is best for photosynthesis and the peak of chlorophyll a absorption in the red (about 680 nm) is not far removed from the peak of chlorophyll a fluorescence (about 685 nm). For this reason blue exciting (actinic) light is sometimes used because it is readily separated from the fluorescence peak. For many purposes, however, fluorescence signals at longer wavelengths (i.e. wavelengths about 740 nm, where there is a smaller fluorescence maximum) are sufficiently strong (and sufficiently similar in quality to those at shorter wavelengths) to allow the employment of filters which will exclude most of the actinic light and transmit most of the fluorescence. In many experiments, fluorescence

signals resulting from excitation with blue light and detected at about 695 nm are similar to fluorescence resulting from excitation with red light and detected at about 740 nm.

8. IS FLUORESCENCE ASSIMILATION?

INFLUENCED

BY

CARBON

The short answer to this question is “yes”. This has been known, or believed, for many years. The original concept was one of “gainful employment”. In short, if excitation energy could be “gainfully employed” by a plant to drive photosynthesis, fluorescence would represent some fraction of the wasted energy. If this were so, a broadly reciprocal relationship might be expected - e.g. if photosynthesis were constrained by some external factor (such as CO2 limitation) more energy would be “wasted” and more fluorescence might be seen. B

Catalase 1/2 O2 + H2O

H2O2 O2

A Antymicin A F d DCMU Cyt b6 QA PQ

ATP

Fp Fd

NADP

C NADPH

PMS or pyocyanine

DBMIB

ATP

X

P700 PSI

P680

H2O 1/2 O2 PSII Figure 8.1. The arrangement of carriers in the photosynthetic electron transport pathway (the Z-scheme). This is an energy diagram showing the “up-hill” transport (in two stages) from H2O to NADP. (A) is linear or non-cyclic electron transport (B) is pseudo-cyclic (the Mehler reaction - see Section 41c) and (C) is cyclic electron transport. The site of action of 3 inhibitors (Antimycin A, DBMIB and DCMU,) (see Trebst, 1980) is indicated and also the action of PMS (or pyocyanine), which creates its own cycle. The spatial organisation of electron carriers is outlined in fig. 8.2. In recent years this concept has been modified many times. Duysens and Sweers (1963) formalised and extended the original notion by applying the name “Q” (for “quencher”) to an electron-acceptor in photosystem II (Figs. 8.1 and 8.2).

The first stable electron acceptor in PSII (“the primary quinone acceptor”) is now called “QA”. When QA is oxidised, it can accept electrons from P680 (the reaction centre in PSII). When fully reduced it can no longer accept electrons which would be passed, in turn (via QB, the plastoquinone pool and PSI) to NADP and beyond. The probability of a fraction of chlorophyll a excitation energy being dissipated as fluorescence is therefore increased. We can, therefore, talk about “qQ quenching” (quenching directly related to the oxidation status of QA) and its relaxation. We can see that if QA is kept oxidised by transferring electrons to NADP and finally to CO2 it will continue to “quench fluorescence” whereas, if its reoxidation is limited, fluorescence will increase as qQ quenching relaxes. For these reasons there are obvious links between the oxidation status of QA, fluorescence, and carbon assimilation. DCMU 2H+

Stroma Thylakoid membrane 2 photons

fp QA

QB

PSII

oxid

PQ shuttle

cyt f PC P700

P680

Mn Inside H2O

NADP

X PSI

reduc tase

2 photons

red

1/2 O2 +

2H+ 2H+

DBMIB KCN Figure 82. Organisation of electron carriers in the membrane. The same carriers as in 8.1, suggesting the manner in which they might be organised within the thylakoid membrane. This figure also indicates the inward movement of protons from the stroma. The proton gradient, discharging through the ATPase (Fig. 8.3) drives ATP formation from ADP and Pi. Note that 2 photons (quanta) are required in each photosystem to transfer 2 electrons from H20 to NADP. Accordingly, a total of 8 are required to transfer the 4 electrons associated with the release of one molecule of O2 (see “quantum requirement” in Sections D and E). The decrease in fluorescence yield brought about by electron transfer to QA (i.e. qQ quenching) is now often referred to as “photochemical quenching” to distinguish it from other quenching mechanisms (see e.g. Krause and Weis, 1984, Oxborough and Horton, 1986) of a “non-photochemical” nature. These include quenching by oxidised plastoquinone (qp), quenching resulting from “state-transitions” (qT) and quenching associated with photoinhibition (qI). The major non-photochemical quenching mechanism, however, is high energy state quenching, or “qE”·

Following an original observation by Murata and Sugahara (1969), work by Krause (for a review see Krause and Weis, 1984 and Barber et al, 1974) has shown that qE is closely related to the proton gradient which develops across the thylakoid membrane as a result of illumination. No one yet knows precisely how this works but it seems that cation exchange processes (involving protons and magnesium ions) bring about changes in the ultra-structure of thylakoid membranes which switch excitation energy dissipation from fluorescence into thermal channels so that energy is lost as heat rather than light. Light-scattering (see Heber, 1969 and Section 13c) is also related to energisation and may be used to monitor changes in the proton gradient and the contribution of qE to fluorescence in vivo (see e.g. Sivak et al, 1985). [It should be noted that, in experiments with isolated chloroplasts, qE may be inhibited by antimycin A although ∆pH is unaffected (see e.g. Oxborough and Horton, 1986)]. When chloroplasts are illuminated, electron transport brings about movement of protons (hydrogen ions, H+) into-the thylakoid compartment so that a gradient of protons is established between the thylakoid compartment (which becomes more acid) and the stroma (which becomes more alkaline). According to Mitchell’s Chemiosmotic Theory this proton gradient and its associated membrane potential (the “proton motive force” or “p.m.f.”) drives ATP synthesis as the proton gradient discharges through the ATPase. The proton gradient also exerts a “back-pressure” on electron transport (it is more difficult to push a proton into a space already full of protons) and this is why electron transport runs faster if it is “uncoupled” (e.g. by detergents which make artificial holes in the thylakoid membrane, or by compounds which act as artificial proton sinks - see e.g. Good et al, 1966; McCarty, l980). In this respect, electron transport is like a railway engine which could go faster if it were uncoupled from its trucks or carriages. Electron transport will also run faster if it is “coupled”, however. This is not as contradictory as it may at first seem because “coupled”, in this sense, means provided with an ample supply of ADP and inorganic phosphate. This allows the proton gradient to discharge through the ATPase, bringing about ATP synthesis. Thus both “coupling” and *uncoupling” allow the proton gradient to discharge. It is only in the absence of ADP, inorganic phosphate, (Pi) or uncouplers that the proton gradient exerts its maximal back-pressure. This is why qE quenching is affected by CO2 assimilation. If CO2 is being rapidly fixed, ATP consumption will be high and there will be plenty of ADP to discharge the proton gradient. Conversely, if CO2 assimilation is limited, ATP consumption will also be limited, ADP will be in short supply, and the proton gradient will increase. As it increases, there will be a shift from energy dissipation as light, to energy dissipation as heat. Thus, when the proton gradient is high, qE will be high and fluorescence will be quenched. The effect, on fluorescence, of changes in ATP and NADPH consumption can be extremely complex but the more simple interactions are probably best understood in regard to air --> CO2 --> free air transients (Section 12a).

H+ X Chloroplast stroma PSI PSII H+

Inside Thylakoid

H+ Fo CF1

H+ H+ (uncoupler)

ADP ATP H+ + Pi Figure 8.3. Electron transport linked to proton transfer into the thylakoid compartment. The proton gradient across the thylakoid membrane can be dissipated by transport through the coupling-factor, which facilitates ATP synthesis, or artificially by uncouplers. CF1 is the ATPase component of the coupling factor and CFo (Fo in this figure) forms a channel in the membrane through which protons are transported.

9. FLUORESCENCE INDUCTION If a leaf is kept for a few minutes in darkness (or low light) and then brightly illuminated, fluorescence rises (in fractions of a second) to an initial peak and then declines (in seconds or minutes) in a more or less simple fashion to a steady-state value. Fluorescence induction kinetics (the Kautsky effect 1931) are, however, often much more complicated than this. The induction curve may be regarded as variable fluorescence superimposed on a “constant” background fluorescence (Fo). The curve has often been labelled OIDPSMT (Fig. 9.1). On illumination, fluorescence rises immediately (in picoseconds but limited by the opening time of the shutter) to O (the level of the “constant fluorescence”) and, thereafter, there is an initial rise to a level “I” in the variable fluorescence (Fv), followed by a dip (D), a peak (P) and a fall, via a quasi-steady-state (S) to a terminal value (T). Sometimes, particularly when 3 or 4 minutes darkness is followed by re-illumination at very low light intensities, a secondary maximum (M) is seen between S and T (Section 11). In other circumstances, particularly in high light and high CO2, oscillations may follow. The initial changes (O to P) are very rapid (about 1 sec) compared with the onset of carbon assimilation and, at least in leaves, there are often additional complexities within the first few seconds which make this nomenclature inadequate.

Figure 9.1. Fluorescence induction kinetics. Schematic representation of fast (left) and slow (tight) fluorescence kinetics during a dark to light transition showing the use of the OIDPSMT terminology (after Lavorel and Etienne, 1977). Note that “0” is designated “Fo” in this figure. The initial fluorescence (Fo) is believed to come from excited chlorophyll a molecules in the antennae of PSII before the excitons have migrated to the reaction centres. Its level is determined by illumination after a period of darkness in order to ensure that QA is fully oxidised (otherwise the rise to O cannot be easily separated from the onset of variable fluorescence and the subsequent rise to I). QA is believed to exist in a state of quasi-equilibrium with plastoquinone to which it passes electrons via QB and the pause at I, and the dip at D, are thought to reflect imbalances between the rates of reduction and reoxidation of QA. Thus, from O to I, reduction is thought to exceed oxidation. Thereafter, increased reoxidation will cause the fluorescence rise to falter at I or even to reverse momentarily, causing the dip (D). In strong light, P may approach Fm (the maximal fluorescence) and the ratio of Fv/Fo may be 4 to 6. The changes O to P are complete within one or two seconds and are best followed on a transient recorder. P to S, on the other hand, takes about 10 seconds and the “M peak” (Section 11), if it develops, may be a minute or so after the initial rise. For this reason some of what occurs could be called induction fluorescence (the fluorescence associated with induction - see Section 10) rather than fluorescence induction (the fluorescence induced by a period of darkness). If the period of darkness is extended (i.e. if the leaf is “dark adapted”) the initial rise approaches a maximum value and the subsequent decline is similar to that observed when illumination in low light is interrupted by short periods of darkness (a minute or so) as in Figure 9.2. In these circumstances, however, the initial rise is small and fluorescence returns very quickly to its steady-state or terminal value. The initial rise can be mostly attributed to the rapid reduction of QA which occurs upon re-illumination.

Oxygen (µmoles)

10.00 9.00 P 8.00 7.00 6.00 5.00 4.00 light 3.00light 2.00

O2

Fluorescence

T

1.00 0.50 1.00 1.50 2.00 2.50 3.00 3.50 4.00 4.50 5.00 Time (minutes) Figure 9.2. Simple Fluorescence Kinetics Following Re-illumination After Darkness. The sequence of events is as follows. Upon illumination, QA is rapidly reduced, qQ quenching is therefore suppressed and fluorescence rises to an “immediate” peak. Thereafter, qQ quenching slowly increases as QA re-oxidises to its steady-state value. At the same time a proton-gradient builds up accross the thylakoid membrane with a consequent increase in qE quenching. During this period. both quenching mechanisms combine to push fluorescence down, from the initial peak (P) to its steady-state or terminal value (T). The subsequent fall results from QA becoming more oxidised (as electrons move on to NADP and beyond) and to the generation of a proton-gradient (the initiation of qE quenching). Between these two extremes (i.e. very long darkness and relatively short darkness) all manner of complexities may appear in the fluorescence kinetics. One is the appearance of the “M-peak” (see above and Section 11). It now seems likely that this particular rise in fluorescence is associated with the termination of the induction period in carbon assimilation (Section 10).

10. INDUCTION As noted above (and in Section 4d), there is often a lag between first illumination and the attainment of maximal photosynthesis, i.e. carbon assimilation does not reach its full rate immediately on illumination but only after an interval. This lag is often evident after a few minutes darkness. After long periods (hours) of darkness, the induction period may last for many minutes. Haas studied induction

The reasons for induction are complex and still a matter for argument (see e.g. Edwards and Walker, 1983). More than 50 years ago, when Osterhout and Haas first studied induction in Ulva they suggested two possibilities. One was, light-activation (and, by

implication, dark-deactivation) of catalysts and, the second, depletion (during darkness) of metabolite pools. These explanations are still valid and there is evidence for both. Light-activation of enzymes is often so rapid, however, that it is unlikely to make much contribution to long periods of induction and these are probably best explained in terms of the known depletion of metabolites such as ribulose bisphosphate (RuBP) (which often falls to very low concentrations in darkened leaves). Some enzymes, are activated by their substrates, as well as by light, so that there can be complex interactions between the two principal underlying factors which lead to induction. In leaves, stomatal opening may also contribute to lags in gaseous exchange following re-illumination after darkness.

11. THE FLUORESCENCE “M-PEAK” In Fig. 9.1, the slower changes which follow re-illumination after a period of darkness, include a decline from a peak (P) to a terminal (T) value via a secondary maximum (M). Perhaps inevitably, this “M-peak” has meant different things to different observers. One approach (Horton, 1985) is to describe any secondary maximum, or maxima (because there may be several) as “M-peaks”. This has the virtue of simplicity but it must be immediately emphasised that it can create a pitfall for the unwary if it is assumed that all “M peaks” are causally related in precisely the same way. A comparison of A and B in Fig. 13.2 shows that fluorescence signals which are superficially very similar may mask underlying components which are quite different and the same may be true of “M-peaks” if they are not additionally defined. For example, re-illumination after a dark interval may give rise to a “CO2-gulp” and an associated “O2 burst” prior to the termination of induction (Section 10). In some circumstances, the termination of induction may be followed by oscillations. All three events (the gulp and burst, the termination of induction and oscillations) may give rise to secondary maxima or “M-peaks” but the underlying causes, the relative contributions of qQ and qE and their timing may differ substantially. Fig. 11.1 illustrated an “M-peak” of the sort which may sometimes be observed when a leaf is abruptly re-illuminated with low intensity light following a period of darkness long enough to permit appreciable depletion of metabolites and some dark-deactivation of enzymes but not so long that the leaf is fully “dark-adapted”. This particular “M-peak” was attributed (Walker, 1981) to the termination of induction (i.e. the onset of more rapid carbon assimilation after an initial lag). According to this view, QA will be rapidly reduced as soon as the leaf is illuminated because, although it must accept electrons from excited chlorophyll, it is denied the possibility of reoxidation by the remainder of the electron transport chain. This is because carbon assimilation has been “switched-off” in the dark and requires time (spent on light-activation of enzymes and building-up metabolites) to get going again. Some CO2 assimilation will soon occur, however, and electrons will also be passed to O2 as an alternative acceptor so that QA Oxidation will become detectable within seconds and there will be a decline in fluorescence from the original peak value as qQ quenching increases in parallel with O2 evolution. This decline will be reinforced

10

30

150 Wm-2

Oxygen (µmoles)

M Fluorescence

1.40 1.20 1.00 0.80 0.60 0.40 0.20

S

d[O2]/dt O2

20.00 17.00 14.00 11.00 8.00 5.00 2.00 -1.00 -4.00

Rate (µmoles.O2.m-2.s-1)

Chl a fluorescence Effect of increasing light Sivak et al 1985

P

2.00 1.80 1.60

Light

light

Dark 4

by qE Quenching as cyclic and linear electron transport lead to the establishment of a proton-gradient across the thylakoid membrane (Figs. 8.2 and 8.3) which is not yet being discharged at its maximal rate. Such a discharge of protons through the ATPase will increase as the flux of metabolites through the Benson-Calvin cycle (Fig. 12.6) increases and ATP consumption by its PGA and Ru5P kinases (Eqns. 12.2, 12.4) makes more ADP available for photophosphorylation. In relatively low light, this new drain on the transthylakoid proton gradient will bring about a significant relaxation of qE quenching (or a significant slowing of the rate of increase of qE) and cause fluorescence to rise from S (Fig. 9.1) towards M. Almost immediately further changes will combine to reverse this trend. Any decrease in the proton gradient will tend to accelerate electron transport as will the increasing availability of NADP reoxidised in PGA reduction (Fig. 12.3). Associated increases in qQ will once again push fluorescence downwards from M and qQ will once again be reinforced by an increasing qE caused by an increasing proton gradient. In higher light (Sivak et al, 1985), the drain on the proton gradient associated with the termination of induction will have a smaller impact on the fluorescence kinetics and the associated M-peak will be smaller.

-7.00 0.50 1.00 1.50 2.00 2.50 3.00 3.50 4.00 4.50 5.00 Time (minutes)

Figure 11.1. “M-Peak” fluorescence kinetics following re-illumination, in very low light after darkness. The initial events are similar to those in Fig. 9.1 but the subsequent differences relate to the fact that induction has been established (i.e. that the period of darkness has been long enough to allow metabolite pools to be partially depleted and enzymes to be inactivated). When the leaf is re-illuminated, therefore, it is not fully prepared for photosynthetic carbon assimilation at maximum rates, and there is a initial lag. As this lag terminates, some of the newly developed proton-gradient is discharged in ATP synthesis, qE relaxes and fluorescence starts to rise again As this happens, electron transport to CO2 causes a simultaneous increase in the proton gradient so that it pushes fluorescence down from its “M-Peak”. At higher light intensities (Sivak et al, 1985 the M-Peak is lost or modified - after Walker (1981).

12. COMPLEX OSCILLATIONS

FLUORESCENCE

KINETICS

AND

12(a) Gas Transients When a leaf is allowed to photosynthesise for some minutes in air, photosynthesis will often rise to a constant rate and chlorophyll a fluorescence will fall to a steady-state value. If the gas-phase is then changed in a way which will affect photosynthesis the fluorescence signal will also respond. The air to CO2-free air transient provides a good example and one which can be explained with some degree of confidence. It will be seen (Fig, 12.1) that, in the steady-state, if the gas-phase surrounding a leaf is changed, abruptly, from air to CO2-free air, the fluorescence signal will rise quickly and then fall again, almost as quickly, to a value which (at least in moderate light intensities) is often lower than it was at the outset.

Figure 12.1. “Gas-transients” in spinach. This is a facsimile of a chart showing fluorescence transients in spinach induced by changes in the gas-phase. Air to CO2-free air transients alternate with similar transients in which the O2 was dropped from 21% to 2% at the sane moment as the CO2 was removed. The theoretical interpretation of some of the gas-transients is straightforward but possibly over-simplified. For air to CO2-free air,it goes as follows. When the leaf is deprived of CO2, carboxylation of RuBP ceases and so does the consumption of ATP and the oxidation of NADPH in the following sequence.

RuBP + H2O + CO2 --> 2 x 3PGA ......Eqn. 12.1 3PGA + ATP --> 1,3DPGA + ADP

......Eqn. 12.2

1,3DPGA + NADPH --> G3P + NADP ......Eqn. 12.3

A “gas-transient”

Initially, therefore, fluorescence will rise because QA (the primary electron acceptor in PSII) will become reduced as soon as it can no longer pass electrons to NADP (or when electron transfer to NADP is diminished). This relaxation of qQ quenching may be accompanied by an initial relaxation of qE quenching (cf. Fig. 13.2B) because ATP utilisation will continue in the absence of CO2 so long as there is 3PGA and Ru5P available for phosphorylation (equations 12.2 above and 12.4 below). Ru5P + ATP --> ADP + RuBP

Any relaxation of qE quenching will reinforce the rise in fluorescence brought about by the relaxation of qQ. Very quickly, however, QA will arrive at a more reduced state and fluorescence would level out at this higher value if qE had not already started to increase and push it downwards again. This increase in qE quenching results from lack of ADP regeneration (caused by ATP consumption in reactions 12.2 and 12.4) and a switch to increased cyclic or pseudocyclic electron transport once linear electron transport to CO2 has stopped. If O2 is decreased to 2% at the same time as the CO2 is removed (Fig. 12.1), the initial rise in fluorescence is greater because of decreased transfer of electrons to O2 (which can substitute, albeit less effectively, for CO2). Accordingly, qQ will relax more quickly and to a greater extent than it does when CO2 alone is removed. In these circumstances, qE will also increase less rapidly because the rate of electron transfer to O2 will be less rapid in low O2 but the interactions are extremely complex. For example, consumption of ATP in photorespiration will also be diminished in decreased O2.

[CO2] ATP [O2] PGA

RBP

[CO2] C3

......Eqn. 12.4

product

photorespiration

Figure 12.2. Effect of Pi and a phosphate-sequestering sugar on gas-transients displayed by a young barley leaf. From left to right the gas transients were 1000 ppm CO2 (5 min) --> air (O.5 min) --> air (3 min). This sequence was then repeated (4 times in this illustration). The change from air to 1000 ppm CO2 (in air) induced oscillations but, when Pi (50mM) was fed, as indicated at the end of the first cycle, the oscillations were rapidly suppressed. They were equally rapidly re-initiated and increased when 2-deoxyglucose (DG) was fed, as indicated, in the last cycle (from Walker and Sivak, 1985)

The switch back to air from CO2-free air is followed by the reverse of these changes (Fig. 12.1) but the entire transient is asymmetric. This is because the resumption of linear electron transport (when CO2 is restored) occurs in the presence of the high proton-gradient which has developed in CO2-free air and although the light-generated “pressure” which drives electrons from chlorophyll to CO2 is unchanged, the “back-pressure” exerted by the proton gradient is larger (Section 8) and electron transport is consequently slower. Gas transients have been used recently (see e.g. Walker and Sivak, 1985) to help to define the relationship between oscillations and cytosolic Pi status. A chamber similar to the Hansatech LD2 was used but the design allowed young barley or spinach leaves to be fed, through the cut-end of the leaves, with water or solutions containing Pi or Pi-sequestering sugars (such as mannose or 2-deoxyglucose - see Chen-she et al, 1975; Herold et al, 1976). These enter the cytosol and are phosphorylated, (in a reaction catalysed by hexokinase) but not further metabolised. This leads to a decrease in cytosolic Pi. If gas-transients from air to high CO2 (say 1000 ppm) are used, oscillations (see also Section 12c) will follow (Fig. 12.2) if the light intensity is sufficiently high or the temperature sufficiently low. Thresholds in CO2 concentration, light intensity, O2 concentration and temperature can be defined but these can all be raised by Pi-feeding and lowered by Pi-sequestering sugars (Walker and Osmond, 1986; Laisk and Walker, 1986, Sivak and Walker, 1986). This suggests that Pi supply is the common feature in most oscillatory behaviour because Pi-recycling in sucrose synthesis will tend to be limiting in high light, low temperature, high CO2 and low O2 (and vice verse). These relationships are explored more fully in Section 12c. 12(b) Experiment 3. Attach a small soda-lime column to one vent of one of the 3-way taps of the Hansatech LD2 and a small pump to the other tap so that when the pump is switched on it is possible to draw air directly through the chamber or through the chamber via the soda-lime column. Place a leaf in the chamber and allow it to photosynthesise, with air being drawn through the chamber by the pump, until fluorescence has fallen to a steady-state value. Change from air to CO2-free air by changing the position of the 3-way tap. After 30 seconds change back to air. Transients similar to some of those illustrated in Fig. 12.1 should be observed. 12(c) Oscillations In certain circumstances, photosynthesis can be induced to oscillate (see for example, Figs 12.2, 12.3). These oscillations can sometimes be seen in the O2 evolution trace but become very much more evident if an electronic “differentiator” is employed. This converts “change in oxygen” into “rate of change”. The fluorescence signal which you record is already a “rate” measurement and, if photosynthesis does oscillate, associated oscillations in fluorescence become immediately apparent. Oscillations (Fig. 12.3) are most readily observed when steady-state photosynthesis is perturbed under conditions which normally favour rapid rates (e.g. high light, high CO2). Much remains to be elucidated

Envelopes

A

CO2 Carbon Chloroplast cycle stroma

Starch<

HP< TP

Pi Cytoplasm Envelopes CO2

B

Carbon Chloroplast cycle stroma Starch< HP< TP

Pi

Mannose > Mannose

Cytoplasm Mode of action of mannose A: normal export from the chloroplast B: diminished export resulting from phosphorylation of mannose

Ru5P

RuBP

but there seems to be little doubt that dampening oscillations are a manifestation of regulatory mechanisms struggling to regain control (and over-reacting) after being thrown off-balance (see e.g. Sivak and Walker, 1984; 1985). One control mechanism which is thought to be implicated involves the [ATP]/[ADP] ratio. As we have seen (Section 8), a low [ATP]/[ADP] ratio favours electron transport because there is then ample ADP to discharge the proton gradient by ATP synthesis. However, a hey reaction in the Calvin cycle is inhibited by low [ATP]/[ADP] ratios (Robinson and Walker, 1979) This is the reaction in which 3-phosphoglycerate (3PGA) is phosphorylated to give 1 ,3-diphosphosphoglycerate (1 ,3DPGA).

CO2

ATP ADP PGA ATP

ATP ADP

ADP DPGA

Pi

NADPH

1/2 O2

NADP

H2O

G3P

control by [ATP]/[ADP] ratio

3PGA + ATP --> 1,3DPGA + ADP

......Eqn. 12.5

This is a freely reversible reaction but the equilibrium position favours PGA formation from DPGA and, in photosynthesis, it has to be pushed to the right by high [PGA] and a favourably high [ATP]/[ADP] ratio. Carbon assimilation, therefore, must inevitably strike a compromise between the low [ATP]/[ADP] ratio which favours electron transport (Section 8) and the high [ATP]/[ADP] ratio which favours PGA phosphorylation . CO2

18 16 CO2

14 12 10 8

Oxygen

6 4 Fluorescence 2 5

10

15

20

25

Figure 12.3. Dampening oscillations in fluorescence. This is essentially the same experiment as that illustrated in Fig. 4.3, except that a confrey (Symphytum officinale L.) leaf was used instead of spinach Dampening oscillations follow the re-introduction of C02 once the CO2 has been exhausted by photosynthesis in a closed system.

Let us now suppose that steady-state photosynthesis has been attained and we suddenly raise the CO2 concentration. This will cause a transient increase in PGA as the high CO2 discharges the ribulose 1,5-bisphosphate pool. RuBP + CO2 + H2O --> 2 x 3PGA

........Eqn. 12.6

This, in turn, will be associated with a transient increase in electron transport and O2 evolution as more PGA is reduced. Fluorescence will start to fall as the proton gradient and qE increase but this fall will be rapidly overtaken by an increase in fluorescence as increased ATP consumption (in the phosphorylation of PGA and Ru5P - Eqns. 12.2. and 12.4) makes more ADP available to discharge the proton gradient. The consequent decrease in the [ATP]/[ADP] ratio will become unfavourable to PGA phosphorylation and reduction (and therefore to O2 evolution and CO2 fixation) and the rise in fluorescence will be reinforced by relaxation of qQ as NADP and QA become more reduced. Each time the [ATP]/[ADP] ratio becomes unfavourably low, the interruption of PGA phosphorylation will tend to diminish the “waves” of metabolites transversing the cycle. As the metabolite “waves” subside (aided by discharge to the cytosol), via the translocator, in exchange for incoming Pi see Figs. 12.4 and 12.7) the oscillations in fluorescence will also dampen (Fig. 12.3). Because of the intimate relationship between the cytosol and the chloroplast, Pi-supply has a major impact on oscillatory behaviour. High cytosolic Pi will facilitate ATP formation and the discharge of metabolites to the cytosol, thereby suppressing oscillatory behaviour. Conversely, any conditions or events which limit Pi-supply (and hence retain metabolite transients within the stroma) will tend to favour oscillations (see e.g. Section 12a and eg. 12.2). Pi

ATP + NADPH

O2

Carbon cycle

THYLAKOID STROMA Starch

HP

CO2

TP

Triose phosphate

Figure 12.4. “High cytosolic Pi will facilitate ATP formation and the discharge of metobolites to the cytosol”.

This explanation is consistent with the fact that the oscillations in fluorescence bear a broadly reciprocal, but phase-shifted relationship, to the oscillation in CO2 and O2 (Fig. 12.5). The phase-shift (the fact that fluorescence changes anticipate O2 and CO2 changes) can be accounted for by the fact that O2 evolution would be tightly linked to the oxidation status of QA but that changes in O2 evolution would be preceded by change in qE. Other measurements, such as light-scattering (Section 13c), suggest that the proton gradient does, in fact, oscillate in advance of O2 and CO2 (Sivak et al, 1985).

E

Fv

1 min

CO2

Fo Time Figure 12.5. Oscillatory behaviour in spinach showing relationship between Carbon dioxide fixation, fluorescence and energisation. Oscillations were induced by re-illumination in 0.55% CO2, 2% O2. The fall in fluorescence (Fv) anticipates the rise in CO2 fixation and vice versa. Similarly, the increase in energisation (E), which is largely responsible for the fall in fluorescence, anticipates Fv. “E” was defined by q-analysis (Section 13a) and is the difference between maximal and variable fluorescence (E = qE(Fv)m = (Fv)m-(Fv)s - see Eqn 13.7). (After Sivak and Walker 1986). It should be noted that while oscillatory behaviour may be initiated by sudden illumination following a period of darkness (a circumstance in which induction will be more or less re-established depending on the duration of the dark interval) it can also be initiated equally readily by almost any perturbation of steady-state photosynthesis. During induction, interpretation is complicated by “switching-off” of light-activated enzymes and depletion of metabolites in the dark whereas, in continuous light, such factors might be expected to make a much smaller contribution.

12(d) Factors which favour oscillations (i) Dark to light (ii) Low light to high light (iii)Low CO2 to high CO2 (e.g. air --> 1000 ppm CO2) (iv) Low temperature --> high temperature (v) High O2 to low O2 (e.g. 21% --> 2% O2) (vi) Low O2 to high O2 Oscillations occur more readily in shade-grown leaves or following illumination of the under-surface of leaves (Walker and Osmond, 1986). Young cereal leaves, which do not form much starch, oscillate readily, whereas Soya, which is a massive starch former, is only induced to oscillate with great difficulty, if at all. Oscillations occur most readily at low temperatures, in high light and in high CO2 but the thresholds for light, CO2, etc., below which oscillations will not normally occur may be lowered or raised by experimental lowering or raising of cytosolic orthophosphate concentration (Section 12a and Fig. 12.2). In all of these regards, oscillatory behaviour may be regarded as an over-reaction, in conditions of limiting phosphate supply, of the regulatory mechanisms which control photosynthetic carbon assimilation. An analogy which has been used is that of a car being driven up a road. If the driver is obliged to swerve violently to avoid an accident he will tend to over-react and his vehicle, following such a perturbation, will proceed down the road in a series of dampening over-reactions until a straight course is recovered. Leaves photosynthesise in order to produce the wherewithal for maintaining respiration and growth. The chloroplast (Section 41e) is an orthophosphate-requiring, triose phosphate-exporting organelle (Walker and Herold, 1977; Walker and Robinson, 1978) and a balance has to be struck between the export of triose phosphate to the cytosol (where it is mostly converted to sucrose) and internal utilisation in the regeneration of ribulose bisphosphate, RuBP, the CO2 acceptor (Fig. 12.6). Five out of six molecules of triose phosphate formed in the Benson-Calvin cycle must be consumed in the regeneration of RuBP simply to maintain the status quo as five C3 compounds (triose phosphates) are re-arranged to give three C5 compounds (ultimately RuBP) . Even more triose phosphate must be consumed internally to allow for the autocatalytic increase in RuBP, at the expense of triose phosphate, without which photosynthesis could not increase and growth could not occur (Fig. 12.6). Only the “spare” triose phosphate can be exported or consumed within the chloroplast during starch synthesis (if it occurs). Export, via the phosphate translocator (Fig. 12.7) is by strict, one-to-one stoichiometric exchange with orthophosphate in the cytosol. Cytosolic Pi seems to be controlled within fine limits and the utilisation of triose phosphate in sucrose synthesis 4 triose phosphate --> 1 sucrose + 4 Pi ...Eqn.12.7 in the cytosol is the principal mechanism of Pi recycling. This allows the synthesis of triose phosphate in the chloroplast to continue (see Section 41e) 3CO2 + 1 Pi --> 1 triose phosphate ...Eqn.12.8

“a car being driven up a road”

Figure 12.6. The Benson-Calvin Cycle as an autocatalytic sequence. Five out of six C3 molecules (G3P) are rearranged to give three C5 molecules (RuBP). The sixth is available for feedback or export to the cytosol via the Pi-translocator as DHAP. Sucrose synthesis in the cytosol (like starch synthesis in the stroma) is regulated by a complex mechanism involving a modulator (fructose 2,6-bisphosphate) and is favoured by high concentrations of triose phosphate and low concentrations of Pi. (see e.g. Cseke et al, 1984; Stitt, 1986) Orthophosphate enters photosynthetic carbon assimilation through photosynthetic phosphorylation in which ADP + Pi --> ATP

......Eqn. 12.9

and it is ADP and Pi which bring about the discharge of the proton-gradient through the ATPase (Section 8). In this way, Pi-supply embraces both electron transport and photosynthetic carbon assimilation. Much remains to be determined about the manner in which all of these events inter-act and are regulated but it is evident that a number of compromises must be established. The maintenance of these checks and balances is evidently central to regulation and It is these which clearly cannot cope, without over-reaction, when steady-state photosynthesis is violently perturbed. Hence the

Outer envelope

Inner envelope

CO2

ADP + NADP

DHAP Sucrose permeable space

DHAP Pi

RPPP

ATP + NADPH

Light reaction

Pi Stromal compartment

Tylakoid membrane

Figure 12.7. The Phosphate Translocator. Located in the inner envelope of the chloroplast, the Pi-translocator permits a strict, one-to-one exchange between internal DHAP and external Pi. oscillations in O2 and CO2 and metabolites, which are most easily followed by measurement of chlorophyll fluorescence and which, undoubtedly, involve rapid interaction between stroma and cytosol, with Pi acting as a chemical messenger.

13. QUENCHING ANALYSIS 13(a) Introduction No contemporary consideration of chlorophyll a fluorescence in its relation to photosynthetic carbon assimilation would be complete without a mention of q-analysis (the analysis of fluorescence quenching - see e.g. Schreiber et al, 1986). Once again the underlying concept is that fluorescence emission (in this context) is largely determined by two quenching mechanisms. As already noted (Section 8) the first of these, Q-quenching (qQ) derives from photochemical energy conversion at PSII reaction centres such that if the primary acceptor, QA, is oxidised it accepts electrons (thereby quenching fluorescence) whereas, if it is reduced, some of the excitation energy is dissipated as fluorescence. Energy is also dissipated by radiationless de-excitation (e.g. as heat) and an increase in the rate of dissipation via this channel is believed to be associated with the build-up of the proton gradient across the thylakoid membrane (Figs. 6.1, 8.2 and 8.3), hence qE or “energyquenching” . 13(b) The “DCMU Method” The action of the two quenching components qQ and qE can be resolved in experiments with isolated chloroplasts (Krause et al, 1982) by the addition of the inhibitor DCMU which blocks the reoxidation of QA by PSI and the rest of the electron transport chain (Fig. 8.2).

Accordingly, if 10 to 15 µM DCMU is added to a chloroplast suspension which has been illuminated for some minutes, an abrupt rise in fluorescence is observed (Fig. 13.1). This relates to the difference in the degree of reduction of QA brought about by photosynthesis (the balance between reduction by electrons from water and reoxidation by PSI) and the complete reduction which follows once re-oxidation of QA is blocked. The fast rise in fluorescence is followed by a slower increase which is attributed to the decrease in the proton gradient.

<-1 min->

Fluorescence rel. units

<-3 min-> P

R slow R fast Fv Fo

Light on

DCMU

Figure 13.1. Fluorescence changes consequent upon the addition of DCMU to isolated chloroplasts. The significance of the bi-phasic rise in fluorescence following the addition of DCMU to intact isolated chloroplasts in the presence of KHCO3 (2 x 10-5M) is explained in the text (from Krause et al. 1982). 13(c) Light-scattering Ulrich Heber made a major contribution (Heber, 1969) to photosynthetic measurements when he devised a method of following changes in (pH based on “light-scattering”. A weak beam of 535 nm green light is passed through a leaf and increases in the signal, detected by a photomultiplier, have been related to changes in the thylakoid membrane during energisation. Fig. 13.2 shows changes in fluorescence and light-scattering during gas-transients (Section 12a). If it is accepted that light-scattering is an indicator of the proton gradient (some caution is necessary because light-scattering is also affected by water status) the respective contributions of qQ and qE are immediately apparent. Upon withdrawal of CO2, QA re-oxidation is constrained and fluorescence rises steeply. This rise is soon over-taken by a fall because, in the absence of CO2, ATP consumption (and associated ADP generation) no longer discharges the proton gradient whereas cyclic and pseudocyclic electron transport continue. It is this increase (Fig. 13.2) in the proton gradient (and the associated imposition of qQ quenching) which pushes fluorescence back down following its initial rise. When CO2 is restored everything is reversed.

1 min

(B)

(A)

(C)

Figure 13.2. Changes in Fluorescence During Gas-transient (A) air --> CO2-free air --> air transient (cf. Section 12a) showing associated change in qE quenching as indicated by light-scattering. (B) and (C) as for (A) but involving higher CO2 concentrations and therefore more complex kinetics, including the onset of oscillations in (C). At higher CO2 concentrations, the behaviour is more complex (e.g. ATP may continue to be consumed at appreciable rates for some seconds after CO2 is withdrawn so that the proton gradient may fall before it rises). Comparison of Figs. 13.2A and 13.2B show this situation and emphasises how important “q-analysis” is (see below) because fluorescence kinetics which are superficially very similar can disguise quite different underlying changes in quenching. 13(d)”Light-doubling” An alternative approach (“light-doubling”) was introduced by Bradbury and Baker (1981) and, in experiments with protoplasts, Quick and Horton (1984), used modulated light (Section 13f) in demonstrating an acceptable correlation between qQ determination by the “light-doubling method” and by the “DCMU method”. Their procedures (like those of Dietz et al.,1985) involved periodic illumination by saturating light, superimposed for brief intervals on the normal “actinic” (or “driving” light) during photosynthesis measurements. An apparatus, designed by Neil Baker, which combines

a modulated measuring beam and lock-in amplifier detection is available from Hansatech Instruments Limited. The rationale behind this approach is that the pulses of saturating light will drive QA fully reduced (qQ will be zero) and that the contribution of qE will then be given by subtraction. It also presupposes that brief, intermittent, pulses will not have a significant impact on other photosynthetic processes. 13(e) The Pulse-Saturation Method “Q-analysis”, as it is now known, has been based largely on pulse-saturation apparatus designed by Schreiber and manufactured by Walz (Appendix 2). The Schreiber apparatus (Schreiber et al 1986) also utilises a modulated measuring beam, but one of such low intensity (O.12 µmole quanta m-l s-1) that, to all effects and purposes, the leaf is still in darkness when this is applied. The Schreiber apparatus also employs a novel and sophisticated detection system which “is free of the switching-on and switching-off artefacts found with conventional lock-in amplifier systems” 13(f) Modulated Light In simple measurements of fluorescence, actinic light is used at one wavelength (e.g. blue) and (red) fluorescence measured at another (say 685 nm or above). This, in itself, limits the range of experiments which can be undertaken (measurements in natural light would be precluded) and there are also changes in “constant” fluorescence emission (changes in Fo) which result from changes in light intensity per se rather than changes in the rate of photosynthesis. These complicate interpretation and comparison. For all of these reasons, there are advantages in using a modulated measuring beam at an intensity which is so low that the leaf scarcely notices that it is not in the dark. A “lock-in” amplifier tuned (or “locked-in”) to the same frequency of modulation is then used as an integral part of the fluorescence detector so that it effectively disregards all changes in fluorescence which are not also modulated. (Such a detector may also be protected by optical filters to prevent it becoming saturated with reflected actinic light). Modulation can be achieved mechanically by “chopping” (a “chopper” is essentially a revolving disc fitted with slits) or electrically. Light emitting diodes are easily modulated by an electrical pulse generator and are used for this purpose in the commercially available apparatus. 13(g) The Procedure A dark adapted leaf or a suspension of chloroplasts etc is first illuminated by the extremely weak measuring beam. This gives Fo, the so-called “dark” or initial fluorescence (Fig. 13.3). A single saturating pulse then establishes (Fv)m (Fig. 13.3) the “maximal” variable fluorescence (i.e. the fluorescence seen when all of the quenching mechanisms are at zero). Thereafter, illumination is started using actinic light of the desired intensity (photon flux density) and pulses of, say, 300 msec duration of saturating light are super-imposed at appropriate intervals (say every 10 to 20 seconds).

At the time of writing, there are incompatibilities between various types of existing commercial apparatus but both the Schreiber (Walt) and Hansatech Instruments equipment have, for example, been successfully used with modified Hansatech LD2 and DW2 electrode systems (e.g. by bringing the fibre optic which carries the actinic, saturating and measuring Light and collects the fluorescence nearer to the sample). Hansatech Instruments have plans to market new apparatus, in the near future, which will be compatible with the Schreiber system. 13(h) “Q” Analysis

The variable fluorescence (Fv) varies between zero at Fo and a maximum, (Fv)m. Each saturating pulse drives QA fully reduced so that a pulse applied soon after the actinic light is switched on will give a fluorescence peak height close to that of (Fv)m. Thereafter, such pulses will give smaller fluorescence peaks, largely because of the development of qE (quenching associated with the proton gradient). If it is assumed that neither (Fv)m nor Fo change, the rest is a matter of simple but tedious arithmetic best left to a computer. It is becoming increasingly clear that this assumption is not valid in all circumstances but this does not detract from the general usefulness of the procedure and the analysis - it simply means that you should, as always, proceed carefully and avoid pitfalls.

These relationships, as defined by Schreiber et al (1986), are as follows (Fig. 13.3). During induction (Section 9 and 10) the variable fluorescence (Fv) falls short of the maximum (Fv)m because of quenching so that Fv = (Fv)m -q(Fv)m ......Eqn. 13.1 In Eqn. 13.1 “q” is the quenching coefficient [q = l-(l-qE)(1-qQ)]. This can vary between q = O (qQ = O because QA is 100% reduced and qE = O because there is no proton gradient) and q=l (when variable fluorescence (Fv) is fully suppressed). Following each saturating pulse, fluorescence is pushed to (Fv)s because qQ is eliminated (Fv)s = Fv + qQ(Fv)s

......Eqn. 13.2

(Fv)s fall short of the maximum, (Fv)m, because of the generation of a proton gradient and its associated quenching (qE) so that:(Fv)s = (Fv)m - qE(Fv)m ......Eqn. 13.3 Thus, combining Equations 13.2 and 13.3:proceed carefully and avoid pitfalls

Fv = (Fv)m - qE(Fv)m - qQ(Fv)s

.....Eqn. 13.4

By replacing (Fv)s in 13.4 with 13.3 and rearranging

saturating pulse

measuring beam

actinic beam

Figure 13.3. Q-analysis. Definition of quenching coefficients etc by Schreiber et al (1986). To is the fluorescence displayed by a dark-adapted leaf in very weak modulated light. (Fv)m is the maximal variable fluorescence first seen when a pulse of saturating actinic light is applied. Fv is the variable fluorescence seen, once continuous actinic light is applied and (Fv)s are the peaks to which this fluorescence is then driven by pulses of saturating light. The derivation of qQ and qE quenching is explained in the text.

Fv = (1 - qE)(1 - qQ)(Fv)m ......Eqn. 13.5 From Eqn 13.5 four further expressions can be derived qQ = (Fv)s - Fv ....Eqn. 13.6 (Fv)s qE = (Fv)m - (Fv)s ....Eqn. 13.7 (Fv)m 1 - qQ = Fv ....Eqn. 13.8 (Fv)s 1 - qE = (Fv)s ....Eqn. 13.9 (Fv)m The two quenching mechanisms (qQ and qE) have already been explained. These bear an inverse relationship to fluorescence so that emission decreases as quenching increases. Conversely, the two expressions 13.8 and 13.9 bear a direct relationship to fluorescence so that, as first approximations, (1-qE) follows the lowering of fluorescence yield by the development of the proton gradient and (1 -qQ) reflects the degree of reduction of Q. Schreiber and Bilger (1986) have recently re-emphasised the problems involved in q-analysis in certain circumstances in which changes in the initial or “dark” fluorescence (Fo) are regarded as constant, become appreciable. Variable fluorescence may also fall below the Fo level and some allowance must be made if an analysis is to be attempted along the lines described in Section 13h. Schreiber and Bilger have defined a quenching coefficient go and shown that there is a relationship between qo and qE which, when fully resolved, is likely to provide the basis of an appropriate correction. 13(i) Examples. Like light-scattering (Heber, 1969), q-analysis has allowed a fuller understanding of the manner in which fluorescence is related to photosynthetic carbon assimilation. A “causal connection” between the M-peak (Section 11) and the onset of carbon assimilation (the termination of induction) was first proposed by Walker (1981) and subsequently by Ireland et al (1984) who used “light-doubling” techniques. Both light-scattering (Sivak et al, 1985) and recent q-analysis have corroborated this, showing that the M-peak is associated with relaxation of “non-photochemical quenching” (qE) and an increase in photochemical quenching. During oscillations (Section 12a) fluorescence anticipates the changes in O2 and CO2. Light-scattering measurements were consistent with the proposal that the phase-shift was attributable to changes in the proton gradient (Sivak et al, 1985) and q-analysis (Fig. 12.4) allows a similar conclusion to be drawn. Quick and Horton (1984) have determined the changes in qQ and qE in experiments with isolated protoplasts and, like Schreiber et al (1986), have found that the fluorescence induction curve is dominated by qQ at low light intensities whereas at higher light intensities there is

a greater influence of qE. The reduction status of QA in the light is largely dependent on the balance which is established between electron transfer from H2O and electron transfer to CO2. Not surprisingly, therefore, qQ declines at a given light intensity as carbon assimilation declines but this relationship is affected by other factors and is best seen in high [CO2] and low [O2], i.e., when the situation is not complicated by photorespiration and electron transport to oxygen. Quenching-analysis has been shown to have important applications in studies on photoinhibition, heat stress, water-stress etc (for reviews, see Schreiber and Bilger, 1986; Schreiber et al, 1986). Fig. 13.4 for example shows the relationship between Q-quenching and assimilation rate as the water content of Arbutus unedo was decreased from 100% to 40%.

100%

A, % of control

80% 70%

60%

50% 40% Q - quenching Figure 13.4. Correlation between Q-quenching and assimilation rate at varying relative water contents in leaves of Arbutus unedo. Fluorescence and rate were measured simultaneously in a leaf disc electrode at high CO2 concentration. (after Shreiber and Bilger, 1986). As already noted, cuvettes and leaf-chambers which will allow q-analysis to be combined with O2 measurement are not yet commercially available although much can be achieved (see e.g. Stitt, 1986) with minor modification of existing apparatus. New prototypes have been designed by Delieu and Walker in the University of Sheffield Research Institute for Photosynthesis and it is hoped that these will soon be made commercially available to a wider public. When incompatibility does exist it lies in the fact that the Schreiber apparatus is partly based on fibre optics. One optic carries the modulated measuring beam, a second the actinic light, a third the saturating pulses

and a fourth the fluorescence signal from the leaf (or plant material) to the detector. These four optics are combined to form a single, larger, optic with an effective diameter of approximately 12 mm. Accordingly if this optic is required to illuminate an area much larger than 12 mm in diameter or to collect fluorescence at a distance from a source it will be more difficult to achieve light saturation on the one hand or to optimise the fluorescence signal on the other. The Sheffield prototypes overcome these problems. Fig. 13.5 illustrates signals obtained with the Schreiber apparatus used in conjunction with a leaf-disc electrode specifically designed for this purpose.

Figure 13.5. Pulse-saturation combined with oxygen measurements in a specifically designed leaf-disc electrode. Both parts of the figure are facsimiles of chart recordings of oxygen and fluorescence traces. Left: Oscillatory behaviour induced by abrupt illumination and re-illumination after a brief dark interval. Note oscillations in the fluorescence maxima elicited by saturating pulses and the inverse relationship between the rate of oxygen evolution and fluorescence emission. Right: Decline in photosynthetic oxygen evolution and corresponding decline in fluorescence maxima associated with depletion of carbon dioxide. Note that re-illumination after a dark interval immediately before this (when carbon dioxide was nearing depletion) did not induce the oscillatory behaviour seen in the upper facsimile. An alternative is to use a conventional leaf-disc electrode fitted with an adaptor now manufactured by Hansatech for this purpose. Walz have recently marketed a device designed by Schreiber for the evaluation of the oxidation status of P700 (Fig. 8.2) so that it is now possible to measure P700 plus O2 in one system and undertake Q-analysis plus O2 simultaneously in a parallel experiment.

References

General References Baker, N.R. and Horton, P. (1986) Chlorophyll fluorescence quenching during photoinhibition. In: Photoinhibition (Kyle, D.J., Arntzen, C.J. and Osmond, C.B., eds). Topics in Photosynthesis, Vol. 9, Chapter 7. Elsevier Science Pubs. pp. 145-168 Clayton, R.K. (1970) Light and living matter. Vol. 1. McGraw-Hill, New York. Hipkins, M.F. and Baker, N.R. (eds) (1986) Photosynthesis energy transduction a practical approach. IRL Press, Oxford, Washington DC. pp 1-199. Horton, P. (1985) Interactions between electron transfer and carbon assimilation. In: Photosynthetic Mechanisms and the Environment. (Barber, J. and Baker, N.R., eds). Topics in Photosynthesis Vol. 6. Elsevier Science Pubs., pp. 135-187. Krause, G.H., and Weis, E. (1984) Review: Chlorophyll fluorescence as a tool in plant physiology. II. Interpretation of fluorescence signals. Photosynthesis Research, Vol 5. Martinus Nijhoff/Dr. W. Junk, The Hague, pp 139-157. Lavorell, J. and Etienne, A.L. (1977) In vivo chlorophyll fluorescence. In: Primary Processes in photosynthesis (Barber, J., ed). Elsevier/North Holland Biomedical Press, Amsterdam pp 203-268. Mitchell, P. (1966) Chemiosmotic coupling in oxidative and photosynthetic phosphorylation. Biol. Rev. Cam. Philos. Soc. 42, 445-502 Renger, G and Schreiber, U (1986) Practical applications of fluorometric methods to algae and higher plant research. In: Light Emission by Plants and Bacteria. (Govindjee, Amesz, J and Fork, D.C., eds) Academic Press, in press San Pietro, A. (ed) (1980) Photosynthesis and Nitrogen Fixation. Methods in Enzymol. Academic Press, London and New York, Vol 69. pp 1-894. Schreiber, U. (1983) Technical review. Chlorophyll fluorescence yield changes as a tool in plant physiology. 1. The measuring system. In: Photosynthesis Research 4, 361-373. Schreiber, U. (1986) Detection of rapid induction kinetics with a new type of high frequency modulated chlorophyll fluorometer. Photosynthesis Research 9 (1-2), 261-272. Schreiber, U and Bilger, W (1986) Rapid assessment of stress effect on plant leaves by chlorophyll fluorescence measurements. NATO workshop Sesimbra Portugal Oct 1985. Schreiber, U., Schliwa, U. and Bilger, W. (1986) Continuous recording of photochemical and non-photochemical chlorophyll fluorescence quenching with a new type of modulation fluorometer. Photosynthesis Research 10, 51-62. Walker, D.A. (1976) Plastids and intracellular transport. In: Encyclopedia of plant physiology, transport in plants II. (Stocking, C.R. and Heber, U., eds) New Series Vol III, Springer-Verlag, Berlin, Heidelberg, New York. pp 85-136. Walker, D.A. and Sivak M.N. (1985) Can phosphate limit photosynthetic carbon assimilation in vivo? Physiologie Vegetale 23, 829-841.

Specific Barber, J., Telfer, A., Mills, J.D. and Nicolson, J. (1974) In: Proc. III Int. Cong. Photosynthesis (Avron, M., ed) Elsevier, Amsterdam. pp 281-288. Bradbury, M. and Baker, N.R. (1981) Analysis of the slow phases of the in vivo chlorophyll fluorescence induction curve. Biochim. Biophys. Acta 63, 542-551.

Briantais, J.M., Vernotte, C, Krause, G.H. and Weis, E. (1985) Chlorophyll fluorescence of higher plants: Chloroplasts and leaves. In: Light Emission by Plants and Bacteria. (Govindjee, Amesz, J and Fork, D.C., eds). Academic Press, New York. Chen-She, S-H., Lewis, D.H. and Walker, D.A. (1975) Stimulation of photosynthetic starch formation by sequestration of cytoplasmic orthophosphate. New Phytol 74, 383-392. Cseke, C., Balogh, A., Wong, J.H., Buchanan, B.B., Stitt, M., Herzog, B., and Heldt., H.W. (1984) Fructose 2,6-bisphosphate: a regulator of carbon processing in leaves. Trends in Biochemical Sciences Vol 9, 533-535. Deitz KJ, U Schreiber, U Heber (1985) The relationship between the redox state of QA and photosynthesis in leaves at various carbon dioxide, oxygen and light regimes. Planta 166: 219-226 Duysens, W.L.N.M. and Sweers, H.E. (1963) Mechanism of two photochemical, reactions in algae as studied by means of fluorescence. In: Studies on Microalgae and Photosynthetic Bacteria (Jap. Soc. of Plant Physiol., eds) Tokyo, University of Tokyo Press. pp. 353-372. French, S.C. and Young, V.K. “Pigment Spectra” In: Biological Effects of Radiation (Duggar,ed. 2nd ed). Vol II. McGraw-Hill, N.Y. Good, N., Izawa, S. and Hind, G. (1966) Uncoupling and energy transfer inhibition in photophos-phorylation. In: “Current Topics in Bioenergetics”. Vol I. 75-112. Heber, U. (1969) Conformational changes of chloroplasts induced by illumination of leaves in vivo. Biochim. Biophys. Acta 180, 302-319. Herold, A., Lewis, D.H. and Walker, D.A. (1976) Sequestration of cytoplasmic orthophosphate by mannose and its differential effect on photosynthetic starch synthesis in C3 and C4 species. New Phytol 76, 397-407. Hill, R. and Bendall, F. (1960) Function of the two cytochrome components in chloroplasts: a working hypothesis. Nature, London 186, 136-137. Hipkins, M.F. and Baker, N.R. (1986) Spectroscopy. 1 Photosynthesis energy transduction a practical approach. IRL Press, Oxford, Washington DC. pp 51-101. Ireland, C.R., Long, S.P. and Baker, N.R. (1984) The relationship betwen carbon dioxide fixation and chlorophyll a fluorescence during induction of photosynthesis in maize leaves at different temperatures and carbon dioxide concentration. Planta 160, 550-558. Kautsky, H. and Hirsch, A. (1931) Neue versuche zur kohlenstoffassimilation. Naturwissenschaften 19, 964. Kitajima, M. and Butler, W.L. (1975) Quenching of chlorophyll fluorescence a primary photochemistry in chloroplasts by dibromothynoquinone (DBMIB). Biochim. Biophys. Acta 376, 105-115. Krause, G.H., (1974) Changes in chlorophyll fluorescence in relation to light dependent cation transfer across thylakoid membranes. Biochem. Biophys. Acta 333, 301-313. Krause, G.H., Briantis, J-M. and Vernotte, C. (1982) Photoinduced quenching chlorophyll fluorescence in intact chloroplasts and algae. Resolution in two components. Biochem. Biophys. Acta 679, 116-124. Laisk, A. and Walker, D.A. (1986) Control of phosphate turnover as a rate-limiting factor and possible cause of oscillations in photosynthesis: a mathematical model. Proc. R. Soc. Lond. B 227, 281-302. Leegood, R.C. and Malkin, R. (1986) Isolation of sub-cellular photosynthetic systems. In: Photosynthesis energy transduction a practical approach. IRL Press, Oxford, Washington DC. pp 9-26. McCarty, R.E. (1979) Roles of a coupling factor for photophosphorylation in chloroplasts. Ann. Rev. Plant Physiol. 30, 79-104. McCarty, R.E. (1980) Delineation of the mechanism of ATP synthesis in chloroplasts. Use of uncouplers, energy transfer inhibitors and modifiers of CF1. In: Methods in Enzymol. (San Pietro, A., ed) Academic Press, New York and London. Vol 69 p719-728. Murata, N. and Sughara, K (1969) Control of excitation transfer in photosynthesis. III. Light-induced decrease of chlorophyll a fluorescence related to photophosphorylation system in spinach chloroplasts. Biochim. Biophys.Acta 189, 182-192. Olson, J.H. and Hind, G. (eds) (1961) Chlorophyll-proteins, reaction centers and photosynthetic membranes. Brookhaven Symp. in Biol no 28. Brookhaven Nat. Lab. Assoc. Universities Inc. Upton, New York.

Oxborough, K. and Horton, P. (1987) An investigation of high energy state quenching in spinach and pea chloroplasts. In: “Progress in Photosynthesis Research” Proc. VII Int. Congress on Photosynthesis (Barber, J., ed) Vol II. August 1986, Providence, Rhode Island, U.S.A. Martinus Nijhoff/Dr. W. Junk, The Netherlands. pp 489-492. Quick, W.P. and Horton, P. (1984) Studies on the induction of chlorophyll fluorescence in barley protoplasts I. Factors affecting the observation of oscillations in the yield of chlorophyll fluorescence and the rate of oxygen evolution. Proc. R. Soc. Lond. B. 220, 361-370. Robinson, S.P. and Walker, D.A. (1979) The control of 3-phosphoglycerate reduction in isolated chloroplasts by the concentrations of ATP, ADP and 3-phosphoglycerate. Biochim. Biophys. Acta 545, 528-536. Sharkey, T.D., Stitt, M., Heineke, D., Gerhardt, R., Raaschke, K. and Heldt, H.W. (1986) Limitation of Photosynthesis by Carbon Metabolism1 II. O2-insensitive CO2 uptake results from limitation of triose phosphate utilization. Plant Physiol. 81, 1123-1129. Sivak, M.N. and Walker, D.A. (1984) New perspectives in the understanding of the regulation of photosynthesis and its relation to chlorophyll fluorescence kinetics through the study of oscillations. In: Oscillations in physiological systems: Dynamics and control (Proc. Symp., Oxford, September 1984) Inst. of Measurement and control, London. pp 91-96. Sivak, M.N. and Walker, D.A. (1985) Chlorophyll a fluorescence; Can it shed light on fundamental questions in photosynthetic carbon dioxide fixation? Plant Cell and Environment 8, 439-448. Sivak, M.N. and Walker, D.A. (1986) Photosynthesis in vivo can be limited by phosphate supply. New Phytol. 102, 499-512. Sivak, M.N., Heber, U. and Walker, D.A. (1985) Chlorophyll a fluorescence and light-scattering kinetics displayed by leaves during induction of photosynthesis. Planta 163, 419-423. Sivak, M.N., Lea, P.J. and Walker, D.A. (1986) New developments in the measurement of photosynthesis in vivo. Implications for plant genetic engineering. Biochemical. Soc. Trans. 14, 63-64. Stitt, M., (1986) Limitation of Photosynthesis by Carbon Metabolism1 I. Evidence for excess electron transport capacity in leaves carrying out photosynthesis in saturating light and CO2. Plant Physiol. 81, 1115-1122 Trebst, A. (1980) Inhibitors in electron flow: Tools for the functional and structural localisation of carriers and energy conservation sites. In: Methods in Enzymol. (San Pietro, A., ed) Academic Press, New York and London. Vol 69 p 675-715. Walker, D.A. (1981) Secondary fluorescence kinetics of spinach leaves in relation to the onset of photosynthetic carbon assimilation. Planta 153, 273-278. Walker, D.A. (1985) Measurement of oxygen and chlorophyll fluorescence. In: Techniques in Bioproductivity & Photosynthesis, 2nd Edition. (Coombs, J., Hall, D.O., Long, S.P. and Skurlock, J.M.O, eds). Pergamon Press, Oxford. pp 95-106. Walker, D.A. and Herold, A. (1977) Can the chloroplast support photosynthesis unaided? In: Photosynthetic Organelles: structure and function (Fujita, Y., Fatoh, S., Shibata, K. and Miyachi. S. eds). Special issue of plant and cell Physiol., Japanese Society of Plant Physiologists and Centre for Academic Publications, Japan pp 295-310. Walker, D.A. and Osmond, C.B. (1986) Measurement of photosynthesis in vivo using a leaf disc electrode: Correlations between light dependence of steady-state photosynthetic O2 evolution and chlorophyll a fluorescence transients. Proc. Roy. Soc. B 227, 267-280. Walker, D.A. and Robinson, S.P. (1978) Regulation of photosynthetic carbon assimilation. In: Photosynthetic carbon assimilation. Basic life sciences. (Hiegelman, H.W. and Hind, G. eds) Vol ll. Proc. Brookhaven Symposium in Biology 1978, Plenum press, New York, U.S.A. pp 43-59. Walker, D.A. and Sivak M.N. (1986) Photosynthesis and phosphate: a cellular affair? Trends in Biochemical Sciences 11, 176-179. Walker, D.A., Sivak, M.N., Prinsley, R.T. and Cheesbrough, J.K. (1983) Simultaneous measurement of oscillations in oxygen evolution and chlorophyll a fluorescence in leaf pieces. Plant Physiol 73, 542-549. Walker, D.A. and Slabas, A.R. (1976) Stepwise generation of the natural oxidant in a reconstituted chloroplast system. Plant Physiol. 57, 203-208.

PART C

LIGHT SOURCES

NEW LAMPS FOR OLD 14(a). Slide-projectors and Heat Filters One of the easiest ways to illuminate a small area of leaf in a laboratory is to use a slide-projector. Sadly, there is also a grave danger of inadvertently cooking the leaf unless adequate heat filtration is also used. These days it is possible to buy excellent “hot” and “cold” mirrors (for example from O.C.L.I. Ltd, High Wycombe, Bucks, U.K.). The former transmit visible light and reflect radiant heat (near far-red, far-red and infra-red radiation). The latter, if held at 450 to the incident light will transmit incident long-wave radiation and reflect the visible. Even using various combinations of such filters as in Fig. 14.1 it is, however, still not easy to stop an illuminated leaf getting hot. This is partly because conventional light sources often emit much more long-wave radiation than visible radiation and a lot still sneaks through the filters. For this reason water filters are often inserted into the light path. Water is a good infra-red filter but it must be remembered that, when it is used in this capacity, its heat absorbing prowess is directly related to its depth. A shallow, flowing, layer of water will conduct heat away from a surface but it will not absorb as much radiant heat as a deeper one. In leaf chambers such as the LD2 there is a built-in water path which will absorb some of the heat and also help to keep the leaf-disc, pressed against it, close to the required temperature. Here again, however, it is important to remember that high-intensity visible light (i.e. light from which all infra-red radiation has been eliminated)

will still cause some heating. It is therefore very difficult to avoid any heating within a leaf as a result of very bright illumination even in a system through which air is being rapidly circulated. Where heating is thought, or known, to be a problem it is best to make measurements with a suitable thermocouple in order to ascertain the extent of the problem. In addition to slide-projectors, and sources such as xenon are lamps, a number of commercially available light sources are now available, some specifically designed for work on photosynthesis. Several firms, for example (e.g. Schott Glass, Stafford, U.K.) produce light-sources which permit fibre or liquid optics to be attached but few of these allow easy insertion of additional optical filters for adjusting light quality. It must also be borne in mind that despite claims to the contrary, fibre optics do transmit radiant heat as well as visible light and that if the heat coming out of the optic is really significantly less than that going in, the same will apply to the light. Hansatech Instruments Ltd now manufacture several light sources, specifically designed for work on photosynthesis. 14(b) A High Intensity Light Source. This is a powerful lamp, of a type similar to that employed in the Research Institute for Photosynthesis in Sheffield but not commercially available. It comprises a quartz-halogen source with a built-in dichroic reflector which preferentially reflects visible light and therefore allows some heat to escape through the reflector. 4 combination of hot and cold Grid Heat-sink

Heat-sink FAN Infra-red Cold mirror Aspheric lens Cooling jacket

Water Path

Reflective cone

Dichroic reflector Quartz-halogen element hot (heat reflecting) mirrors Aspheric lens Mirrored surface Electronic shutter and iris diaphragm

Visible light Fig. 14.1 A High Intensity Light Source.

mirrors pass the visible light through a water path and finally through an iris diaphragm and an electronic shutter. The shutter box also incorporates a system of mirrors which allows additional lights to be added, or fluorescence signals to be deflected. 14(c) Light-Emitting Diodes Light-emitting diodes (LED’s) are becoming increasingly useful as light sources in photosynthetic applications (Rurainski and Mader, 1980). They are small and do not require high amperage currents. They do not produce a great deal of heat and the extensive heat dissipation built into high intensity sources such as that described in Section 14a is no longer necessary. The red versions, which have peak output at 635 nm or above, produce light which is not far removed from the peak absorption of chlorophyll in the red (Part B) and such LED’s can often be used (e.g, in fluorescence measurements) without the need to define the light quality by using expensive optical filters. Similarly, their very rapid “switch-on” time (about 0.25 µsec) eliminates the requirement for rapidly opening shutters which are mandatory when conventional light sources are used as actinic light in the measurement of early events in fluorescence induction etc. Their output can be varied electrically with little effect on light quality and their output can be readily modulated electronically for applications involving lock-in amplifiers. They are relatively cheap, relatively stable, and relatively long-lived.

Aluminium Case Array of LED’s

Figure 14.2 Light Emitting Diodes (The Hansatech Instruments LH36/2R) An array of LED’S (Fig. 14.2) which give enough light to support about one third of maximal photosynthesis by most C3 leaves is currently marketed by Hansatech (the Hansatech LH36/2R). This light source was specifically designed for use with the leaf-disc electrode and will work off mains-electricity or car batteries. It has all of the intrinsic properties and advantages of LED sources. The field of illumination is not as uniform, nor the output as high, as that of the Bjorkman lamp (Section 14d, below) but uniformity can be improved (at some cost in output) by judicious use of diffusing screens and it is a versatile and extremely useful light source for many applications involving the leaf-disc electrode. Hansatech offer a high intensity LED head (LH36/2R). The LH36/2R operates in conjunction with a computer controlled electrode control box (The Hansatech Oxylab system. Appendix 3) which is also available for incorporation into the measurement of quantum yield (Part D and Appendix 3). The light output is varied automatically and may be configured from the control software of the Hansatech Oxylab control box. This can be calibrated to give pre-selected photon flux

densities which are digitally displayed. When activated, a series of light intensities may be switched automatically by the Hansatech Oxylab system. 14(d)The Bjorkman Lamp This was designed by Olle Bjorkman of the Carnegie Institute for use with the Hansatech LD2 leaf-disc electrode and is now also marketed by Hansatech (the LS2). It is an elegantly simple lamp (Fig. 14.3) which gives a very uniform field of illumination. It accommodates neutral density filters which slip into slots, thereby providing a range of light intensities.

Fan Tungsten-halogen bulb

socket for fibre optic cable Heat filter adjustable bulb mounting

Heat filter lenses Slots for neutral density filters

neutral density filters

Figure 14.3. The Bjorkman Lamp. This fits directly on to the top of the Hansatech leaf-disc electrodes but may also be used as an actinic source for a variety of purposes. This makes it particularly suitable for measuring the rate of photosynthesis as a function of light intensity (see Part D). It is portable, robust and will work off a 12-volt car battery. It does not have a rapid switch-on time like the LED’s (the light source itself is a tungsten-halogen bulb) and (unlike the Hansatech LSl)could not be used for following the earliest changes in induction fluorescence unless it was used in conjunction with a fast shutter and appropriate optical filters to define the light quality.

Reference Rurainski, H.J. and Mader, G. (1980) Light-emitting diodes as a light source in photochemical and photobiological work. In: Methods in Enzymology (San Pietro, A., ed) Vol 69, 667-675. Academic Press. Inc., New York, London.

PART D

MAXIMUM EFFICIENCY OF PHOTOSYNTHESIS

15. INTRODUCTION Few scientific controversies have raged as long, or as fiercely, as that surrounding the determination of the maximum efficiency of photosynthesis i.e. the relationship between CO2 fixed (or O2 evolved) and light energy utilised. For many years the minimum quantum requirement has been taken to be 8, but, during his lifetime, this view was steadfastly rejected by Otto Warburg who put the value at 4 or even less. Warburg, of course, was a Nobel laureate, the man who first used spectrophotometry and manometry in biochemistry and the discoverer of NAD and NADP. In short, his expressed opinion, based on meticulous experiments, was not to be taken lightly. One who did have the temerity to question the great man’s conclusions, however, was a young American called Robert Emerson who studied in Warburg’s laboratory in Berlin and then returned to Urbana, Illinois, where he was to make major contributions to photosynthesis (e.g. the Emerson enhancement effect). Emerson found the requirement to be 8, but Warburg insisted that it was 4 and nothing, seemingly, would bridge the gap. Eventually Warburg traveled to the United States with a view to undertaking definitive experiments but, in the end, nothing was resolved. Emerson himself believed that he wasted years of his life in attempting to arrive at the definitive answer. Warburg’s measurements of quantum yield requirement were made on Chlorella, by manometry. Over the years, the procedures that he first described in 1923 were greatly simplified and even, he said, “performed as simple classroom experiments” (Warburg and Burke, 1950). Nevertheless, it was never an easy matter to obtain low values. The algae had to be carefully grown in well water or spring water and in experiments undertaken in 1949 “it took us about two and a half months to find out the conditions under which high efficiencies could be obtained in every experiment”. “Only gradually did we come to clarify and identify the reasons for the irregularities: the sedimentation of the cells in the cultures, the not always adequate shaking of the manometer vessels, the too few cells in the manometer vessels, the too great time differences between manometric readings of the two vessels, the damage done to the cells when shaken too long in the dark, the too high intensity of the measured red-light beam, etc. The experiments themselves took several hours”. After all this effort Warburg believed that he had answered his critics, that high efficiencies could be routinely measured and “that in a perfect nature, photosynthesis is perfect too”. He was wrong. Why he was wrong remains uncertain to this day. Perhaps James Franck (1953) came nearest to the mark in his erudite discussion of this problem in which he implicated respiratory intermediates in an attempt to rationalise the

results of Warburg and Burke. Here the suggestion was that the lowest values were not a measure of photosynthetic efficiency at all but of some mixture of respiration and incomplete photosynthesis (reduction of 3-phosphoglycerate not linked by one-to-one stoichiometry with CO2 uptake) induced by an unusual combination of conditions (shaking of the algae in the manometer vessels resulted in only 5% of the algae being illuminated at any one time, spending 0.1 or 0.2 seconds in bright light and a further second or so in darkness) and compounded by the sluggishness of manometry (“it has been repeatedly shown that manometry is the most inadequate technique for following closely the quick rate changes which occur during the induction period”). Whatever the real explanation of Warburg’s low quantum values, Franck was, however, undoubtedly correct when he said “We do not feel that it is justified to disregard all other evidence only to make possible an unusual -explanation of certain brief, transient anomalies of the gas exchange, which occur under special conditions”. Since he made this statement in 1953 the “other evidence” has come to be regarded as compelling by almost everyone in the field. Photosynthesis is an imperfect phenomenon in an imperfect world. In higher plants the quantum requirement might approach 8 but a value of 4, or less, is inconceivable.

4 or 8? 30

photosynthetic O2 evolution / (µmol O2 m-2 s-1)

control

Using the leaf disc electrode, it is now really possible to “perform simple classroom experiments” to measure apparent quantum yield by the procedures described below. Using a computerised version of this procedure a value can be obtained in 20 minutes, using the simplest of apparatus. Absolute values can be derived from the apparent value by applying a correction obtained by determining light absorption by the leaf disc in an integrating sphere. For some purposes, however, the apparent values will allow useful comparisons to be made without this correction.

25

16. MEASUREMENT OF QUANTUM YIELD 20

15

10

5

Quantum yield is a measure of the costs of converting light energy into chemical energy in photosynthesis. It is measured as the number of moles of O2 evolved (or +20mM CO2 fixed) per mole quanta of absorbed photosynthetically active radiation (PAR; λ phosphate 400 - 700mn). It can be · calculated from the initial slope of the light response curve of photosynthesis and correcting this slope for the absorbance of the particular plant tissue (usually 80-85% of the incident light between, 400 - 700 nm, is absorbed). In healthy plants, quantum yield varies according to the pathway of photosynthetic +20mM carbon metabolism, temperature and prevailing O2 and CO2 concentrations. In plants D-mannose exposed to stress, quantum yield varies with the extent and duration of a particular stress, and whether the stress is experienced in light or darkness. Unchanging quantum yield may be equally informative. The slope of the light response curve (left) from which quantum yield is derived, remained unchanged despite massive changes in photosynthetic rate brought about by experimental manipulation of cytosolic Pi (Section 12a). Thus measurements of quantum yield are useful indicators of a range of different photosynthetic parameters in healthy and stressed plants. 200 400 600 800 Incident PFD /(µmol quanta m-2 s-1)

17. THEORETICAL BACKGROUND The light dependent production of chemical energy in photosynthesis requires co-operation of two photochemical acts (Figs. 8.1 and 8.2). Production of one molecule of NADPH requires removal of 2 electrons from water and the movement of these through the electron transport chain driven by the two photochemical acts (one in PSII and the second in PSI), each of which requires one quantum of light energy (one photon) to function (Fig. 8.1). Thus production of one mole of NADPH requires the absorption of 4 quantum moles of light energy. As each electron moves through the electron transport chain, two protons are pumped to the interior of the thylakoid. Production of one molecule of ATP by noncyclic photophosphorylation requires movement across the thylakoid of 3 protons. Thus the theoretical stoichiometry of NADPH, ATP production, electron transport and quantum absorption is 1 NADPH : 1.33 ATP : 2 electrons : 4 quanta absorbed Removal of two electrons from water produces half a molecule of O2 so that the molar ratio for O2 evolution in photosynthesis is 1 O2 evolved : 2 NADPH : 2.66 ATP : 4 electrons : 8 quanta absorbed A theoretical minimal quantum requirement can also be estimated from the known stoichiometry of the photosynthetic carbon reduction cycle in C3 plants. In the absence of photorespiration, reduction of one molecule of CO2 requires 2 NADPH and 3 ATP. Note that there is a shortfall of 0.33 ATP between this estimate and that estimated on the basis of O2 evolution above. Because the stoichiometry of photosynthetic CO2 and O2 exchange in vive is 1, we must account for this additional ATP requirement, and it is thought to be produced in processes of cyclic photophosphorylation (Fig. 8.1). Thus the minimum quantum requirement for photosynthetic CO2 fixation is slightly greater than 9 quanta absorbed per CO2 fixed. Quantum yield is the reciprocal of quantum requirement. Accordingly, the theoretically maximum quantum yield is about 1/9 or 0.111 molecules of O2 evolved (or CO2 absorbed) per photon of PAR absorbed. This calculation refers to red light which is used with greater efficiency than light of other wavelengths. Because most plants are exposed to the full solar spectrum and because it is sometimes more practical to use white light for photosynthetic measurements, a correction can be applied for the lower efficiency of white light overall. The data of McCree (1972) suggested that quantum yields in white light could be 25% lower than those in red light, because blue and green light are less effective than red light (but see Evans, 1986 and below). This would have put the maximum theoretical quantum yield for C3 plants in white light at about 0.83 (a quantum requirement of about 12). Many literature values approach this (0.081 in Encelia and Agropyron: Ehlringer and Bjorkman, 1977; Ehlringer and Pearcy, 1983) but, more recently, values equivalent to an average quantum requirement of 9.47 have been obtained for more than 30 C3 species (Section 19) in measurements undertaken with the leaf-disc electrode. “the full solar spectrum”

In C3 plants, in air, a certain amount of photorespiration takes place, depending on the temperature and the stomatal control of intercellular partial pressure of CO2. Photorespiration involves the release of CO2 from recently fixed carbon, so that net fixation per quantum absorbed is reduced. The reduction of quantum yield in C3 plants due to photorespiration which was controlled by varying O2 concentrations was demonstrated by Ehlringer and Bjorkman (1977). For a variety of reasons, photorespiration in air tends to increase with leaf temperature and these authors also sheared that quantum yield of C3 plants in air decreases with temperature. All of these complications are removed when quantum yield is measured at CO2 saturation in the O2 electrode. In C4 plants, carbon reduction takes place under saturating CO2 concentrations in bundle sheath cells and is insensitive to O2 concentrations up to air levels. However, the CO2 concentrating process involves preliminary carboxylation of phosphoenolpyruvate and requires additional energy. The known stoichiometrics of C4 photosynthesis suggest that 2 NADPH and 5 ATP are required, per CO2 fixed. This corresponds to a quantum requirement of 15 quanta per molecule of O2 evolved, or a quantum yield of 0.067 mol O2 evolved, per mol quantum absorbed. Correcting for red light vs. white light, values of 0.050 would be expected in air. In practice, the quantum yields of C4 plants are higher than this (0.053-0.065, Ehlringer and Pearcy 1983 and 0.069, Bjorkman and Demmig, 1987).

Quantum Yield

“several intriguing explanations can be advanced”

In CAM plants, depending on the pathway of decarboxylation, the quantum yield of photosynthesis should be marginally less than in C4 plants. However, recent experiments show that quantum yields of CAM plants are the same as those of C3 plants, when measured in the O2 electrode. Although several intriguing explanations can be advanced to explain these discrepancies, no direct evidence in support of them is currently available. John Evens (1986) has recently re-examined the dependence of quantum yield on wavelengths. His results support earlier findings that the quantum yield is lower in the blue than in the red and this is attributed to the difference in the absorption spectra of the antennae of PSI and PSII (PSII, being richer in chlorophyll b, will absorb more quanta than PSI at wavelength shorter than 680 nm and vice versa at longer wavelength). This will affect the correction for absorbance and will also lead to an imbalance between the two photosystems such that at wavelengths below 680 mm, PSI receiving fewer quanta than PSII, will limit whole-chain electron flow. Evan’s results showed a smaller difference between red and white light than those of McCree (1972). His values of quantum yield for spinach were 0.098 (a requirement of 10.2) and 0.112 (a requirement of 8.9) for white and red light respectively - a difference of about 12%.

0.08

0.05

18. MEASUREMENT OF APPARENT QUANTUM YIELD WITH THE LEAF DISC ELECTRODE AND THE BJORKMAN LAMP 440

520 600 Wavelength (nm)

Quantum yield as a function of light quality.

680

Quantum yield measurements using the LD2 require high sensitivity of photosynthetic rate measurement at low light intensity, uniformity of illumination over the leaf and precise control of light intensity. All of these can be obtained with the Bjorkman lamp. A 50W bulb is normally most suitable for quantum yield measurements.

First calibrate the light-output with a suitable set of neutral density filters and a sensitive quantum meter. (A “Skye” quantum meter which can also be supplied by Hansatech (QSPAR) (Appendix 2) and which is specifically designed for use with the LD2 is ideal for this purpose). This is done by inverting the lamp, fitting the top chamber of the leaf disc electrode (filled with clean water) and placing a disc of white paper on the plexiglass plug which presses against the leaf disc when the chamber is in use. Take care that the fan is not obstructed and switch the lamp on. Fix the quantum-sensor in one place on the disc of white paper (or move it about carefully if an integrating meter is available) and insert combinations of filters to give light of appropriate intensities. If the “Skye” sensor is used it fits directly over the plexiglass plug and integrates the photon flux density over this surface (without need for a paper disc). It is advisable to have four or more photon flux densities between 0 and 100, 2 or more between 100 and 200 and 3 or more between 200 and 800 µmol quanta m- 2 s-l , for most purposes REPEAT THIS PROCEDURE whenever bulbs are exchanged. Check also the alignment of the bulb (Hansatech LS2 Notes for Users page 3). Check the light output of the lamp at frequent intervals. If changes in the mains voltage cause significant fluctuations in light output it may be necessary to use a regulated power supply or car batteries. TAKE PARTICULAR CARE to ensure that cooling water in the chamber is clear at all times. An in-line filter helps. Quantum yield measurement is commenced by placing a leaf disc in the chamber just as if it were to be used for photosynthetic or fluorescence transient measurements. Calibrate the chamber volume, and O2 content as described previously, for each leaf disc used (Section 4). For most purposes it is best to measure quantum yields in saturating CO2. This can be done by using a pad soaked in bicarbonate/carbonate buffer, by breathing into the chamber until a standard, say 5%, CO2 concentration is attained, or by purging the chamber from a bottle of 5% CO2, passing the gas through an aspirator to saturate it with water vapour. Having charged the chamber with CO2 the leaf disc should be exposed to several cycles of dark, followed by full light output from the Bjorkman lamp until a steady rate of photosynthesis is attained and oscillations in O2 evolution are minimised (Section 12; Fig. 12.1). If no bicarbonate buffer has been used be sure to recharge the chamber with CO2 , after this procedure. In practice, the chamber filled with 5% CO2 will sustain only about 5 min high rate photosynthesis. Illumination of leaves which are inadequately supplied with CO2 may lead to photoinhibition. Similarly, if shade leaves are used, it is prudent to use lower light intensities (say 200 µmol quanta m-2 s-1) during this conditioning phase. For some purposes, it is convenient to use this conditioning period to measure room temperature fluorescence transients. In this case a blue filter may be used for the conditioning light, and removed before commencing the

quantum yield measurements. Alternatively, the Hansatech LED light source can be used to elicit the fluorescence responses. The electrode back-off should be applied to obtain the maximum sensitivity available with recording system. The conditioned leaf disc should then be kept in the dark for 5-10 min until a steady rate of respiratory O2 uptake is obtained. Starting with the neutral density filter combination which gives the lowest light intensity selected, a light response curve is then constructed. Filters should be changed at frequent intervals, but only after a steady rate of O2 exchange has been established. Experience suggests that a single charge of 596 CO2 is adequate to complete the linear portion of the light response curve (to 200 µmol quanta m-2 s-1 approximately). Depending on the photosynthetic capacity of the species, and on whether or not information is to be sought on saturation characteristics, it may be necessary to recharge the chamber CO2 supply at higher light intensities. Note that when the chamber is recharged by breathing, some time for temperature adjustment will be required before steady rates can be obtained. Rate of O2 evolution is then calculated for each light intensity and the “apparent” quantum yield read from the slope of rate vs. incident quanta. Actual quantum yield requires measurement of transmission and reflectance of the leaf surface presented to the light in the electrode chamber. This may be done with an integrating sphere and reflectance standards. For most leaves without reflective hairs or wax, a value of 80-85% absorbance can be assumed. Changes in absorbance following stress are small so that comparisons of apparent quantum yield, before and after stress treatment, are sufficient to indicate damage.

19. DOING IT IN OTHER WAYS 19(a) According to Bjorkman and Demmig (i) Introduction. Sections D(18) and E described procedures for measuring quantum yield. Since they were written a paper has been published by Bjorkman and Demmig (1987) which has such important implications that the methods used by these authors are reproduced here in their entirety (Section 19b below). The importance of this work, which was concerned with the measurement of absolute quantum yield, lies in the remarkable constancy of the values (derived with the leaf-disc electrode) for plants possessing the same pathway of CO2 assimilation (Table 19.1). Together, the constancy and magnitude of the C3 values suggests that they may be the best values yet achieved. Even the most careful previous measurements, such as those made by Ehleringer and Bjorkman (1977) and Sharp et al (1984) would appear to have under-estimated the real quantum yields (the latter authors obtained a value of 0.087 (11.49) for Helianthus leaves using IRGA at 0.205% CO2 and 21% O2).

Table 19.1. Quantum Yield (from Bjorkman and Demmig, 1986). Pathway

No of Species

Mean QY

Mean QR

C3* 37 0.106(+/- 0.001) 9.43 C4 5 0.0692(+/- 0.004) 14.45 * The values for 2 CAM species were similar to those of the C3 species

If this is the case, the difference may lie in the difficulty in achieving CO2 saturation without adversely affecting measurement by IRGA (because of stomatal closure and loss of sensitivity at high CO2 concentrations). As noted previously, the leaf-disc electrode is not affected in this way because, even if high CO2 induces stomatal closure the diffusion gradients are such that measurement is not impaired. Similarly, the new values prompt questions about the magnitude of the data of McCree (1972) which suggested that quantum yields in white light might be 25% lower than those in red light. Bjorkman and Demmig used white light and a 25% higher quantum yield in red light (i.e., a quantum yield of 0.1325 or a quantum requirement of 7.54) would be highly questionable. Recent work by John Evans (Section 17) suggests, however, that the difference between red and white might be smaller than previously supposed. Al of the leaves that were used were of healthy appearance and had not been subjected to stress. The procedure was then as follows:(ii) Photon Yield Measurements “Rates of photosynthetic O2 exchange were measured at 25oC and CO2 saturation with a system incorporating a leaf disc electrode (Model LD2, Hansatcch Ltd, Kings Lynn, Norfolk, UK) designed by Delieu and Walker (1981. 1983), and a fan-cooled quartz-lamp housing, designed and built in this laboratory. A similar commercial light source in now manufactured (Model LS2; Hansatech). The light from a quartz-halogen lamp is collimated by 2 condenser lenses and filtered through a hot mirror (OCL1, Santa Rosa, California, U.S.A). This source provide a collimated light bean at the position of the leaf disc immediately below the lower window in the leaf disc chamber. The lamp (Type Bellaphot 50W; Osram) was operated at 1l.60 +/- 0.02 VDC, provided by a regulated power supply (Vista Model XRD; Clifford Industries, Amarillo, California, U.S.A) that was modified to permit voltage adjustment and to provide accurate display of the output voltage. The PFD at the leaf position was approximately 600 µmol m-2 s-1. Desired attenuation of the light was obtained by inserting in the light beam neutral density screens (made by sandwiching fine-mesh nylon screens between two 50 x 50 mm slide mounting glasses) and/or neutral density filters (Melles Griot, Irvine, California, U.S.A). Optimal uniformity and reproducibility of the light field was obtained when the nylon-mesh screens were placed at a position normal to the light beam, 1 cm below the lenses, and the neutral density filters were placed close to the top of the leaf chamber window at a 20 to the horizontal plane. The PFD incident on the leaf disc with each of the filter combinations used in the photon yield measurements was measured at 17 different positions in the light field with a quantum sensor (Model LI 190SB; Li-Cor, Lincoln Nebraska, U.S.A), recently calibrated at the factory. The variation in PFD over the light field was +/-3 to +/- 6%, depending on the filter combination wed. The mean PFD values obtained at the 17 positions were used for calculations of photon yield.

Comparisons made with three other quantum sensors from the same manufacturer gave values which agreed within +/- 2%. The spectral photon distribution of the light incident on the leaf disc was measured with a spectral radiometer (Model 1800; Li-Cor). Polarization voltages and signal readouts from the leaf disc electrodes were provided by a read-out control box (Model CB1-D; Hansatech) a by a laboratory-built unit of similar design, and the signals were displayed on a chart recorder (Recordall 5000; Fischer Scientific Co., Pittsburgh, Pa, USA). Temperature control was provided by a controlled-temperature water circulator, capable of keeping the temperature constant within +/- 0.02oC. The influence of variations in ambient air temperature was minimised by insulation of the lower part of the electrode chamber with a cylinder of polyurethane foam. The electrode was covered with an electrolyte gel (part no. 77948D; Beckman Instruments Inc., Fullerton, California, USA) followed by a 25 x 25 an piece of cigarette paper (Rizla) and a teflon membrane of the same size (part no. 76438, Bechman). This procedure proved superior to others that were tried in terms of ease of application, electrode signal noise and stability, and useful lifetime of each application. The electrode signal usually reached adequate stability within 1 h after the polarization voltage (0=100 V) had ban applied. The electrode output was checked in an air stream of known O2 pressure (usually 20 kPa) and then in a stream of pure N2; the residual signal in N2 was usually < 1% of the signal in air. After adjustment of the signal and calibration of the O2 electrode, the signal amplification was increased by a factor of 10, and an appropriate bias signal was applied to give desired sensitivity for the photon yield measurements. In most uses the signal was adjusted so that the difference between zero and full scale deflection on the chat recorder was equal to a change in O2 pressure from 20 to 22 kPa O2. Rates of O2 exchange at 25oC and standard barometric pressure were calculated from the expression P = 68.2.V.∆S/SA where P is the rate of O2 exchange in µmol O2 m-2 s-1; V the net volume of the electrode chamber in ml; ∆S the change in the electrode signal per min caused by the leaf disc; S the signal obtained for each kPa change in O2 pressure using the same amplification; and A the exposed leaf area in cm2. The volumes of the two electrode chambers used in this study (with the electrode in place, and the 3 metal discs, the foam disc and the fibre matting supplied with the chamber present) was 4.44 and 4.54 ml. To obtain the net chamber volume, any addition of water to the chamber and the volume of the leaf disc were subtracted from the chamber volume. The leaf area exposed to the Light beam (A) was 8.86 cm2 for both of the electrode chambers. The total leaf area was 10.0 cm2 In the measurements at Stanford, CO2-containing air was provided from a gas cylinder containing 20 kPa O2, 5 kPa CO2 (balanced N2). A slow stream of this gas whose flow rate was controlled with a mass flow-controller (Model FC260; Tylan, Carson, California, U.S.A.) was humidified by passing it through a porous ceramic cylinder immersed in water at 25oC and then was passed through the electrode chamber. After placing the leaf disc in the chamber, the flow rate was adjusted in relation to the expected rate of O2 evolution so that the rate of O2 exchange could be monitored in an open flow mode. When a constant signal had been reached, the inlet and outlet valves of the chamber were closed, and the rate of rise in O2 concentration was then followed in closed mode until it could be ascertained that the slope bad reached a constant value. The leaf disc was then darkened and the rate of O2 uptake recorded, usually for 4 min. The rate of O2 evolution measured just before darkening the leaf was then added to the value of O2 uptake measured in the interval 1 to 4 min after darkening the leaf. The light intensity from the light source was then changed, an open-flow mode was resumed and when a new constant signal had been reached, the valves were again closed and the cycle repeated. The main advantage with this alternation between open and closed modes in photon yield measurements is that this procedure avoids undesirable build-up of O2 and depletion of CO2 while waiting

for the establishment of a constant rate. At high rates of O2 evolution open-mode measurements are superior to measurements in the closed mode; however, quantative determinations in the open mode require that the O2 concentration be measured in the exit gas stream from the leaf disc chamber, not in the centre of this chamber. Very close agreement was obtained when rates measured in open and closed modes on the same leaf discs were compared. In general, photon yield measurements were started at a PFD of 125 µmol m-2 s-1 and the PFD was then decreased in steps of approximately 125 µmol m-2 s-1, until the sum of the rate of O2 evolution in the light and the rate of O2 uptake measured in the subsequent dark period (P + R) was linearly dependent on PFD. The photon yield on the basis of incident PFD (ϕi) was then calculated as ϕi = (P + R)/PFD. Except for extreme shade leaves which had very low light-saturated rates, (P + R) was generally a linear function of PFD up to PFD values as high as 75 to 125 µmol m-2 s-1. In the photon yield measurements made in Australia, CO2 was supplied from a 1M sodium bicarbonate buffer or by the operator blowing a slow stream of air from his lungs through the leaf disc chamber. This procedure was not as satisfactory as that later employed at Stanford, and many more replicate determinations of the rate of O2 exchange had to be made. Also, in these photon yield determinations the rate O2 uptake in the dark was not measured after each light step; it was usually only measured after the O2 exchange measurement at lowest light level had been completed (iii) Leaf Absorptance Measurements After completion of the photon yield measurement a 4 cm2 sample was punched from the leaf disc and placed in the centre of an Ulbricht integrating sphere with the upper surface facing the light source. The spectral absorptance (αλ) was determined at 25 nm intervals from 400 to 750 nm as described by Ehleringer (1981). The absorptance (α) of the photons incident on the leaf in the photon yield determinations and in natural daylight was then calculated for the waveband 400 to 700 nm according to the expression 700

α=

Σ

400

αλ . eλ

700

Σ

400

eλ

where αλ is the spectral absorptance of the leaf, eλ the spectral photon emittance of the light source and λ the wavelength in nm”. (iv) Further Considerations It will be noted that Bjorkman and Demmig’s calibration procedure was different from that described in Sections 4 and see Appendix 3 which is based on the difference in signal between air and a second concentration of O2 rather than the difference between air and nitrogen. The relationship P = 68.2 x V x ∆S/S.A contains a value of 68.2. This is because the rest of the equation expresses the rate as a change in the partial pressure of O2 per minute per sq cm of leaf. In order to express it in µmol O2 m-2 s-1 it is necessary to multiply by 10,000 (to convert cm2 to m2), by 1/60 (to convert minutes to seconds) and by 0.4087 (to convert 1 ml of 1% O2 into µmoles/ml). This latter value (0.4087) comes from the fact that air (21% O2) contains 9.37 µmoles of O2 at STP and at 25oC this becomes 9.37 x 273/298 = 8.58 µmoles/ml (Section 4) or 8.58/21 (= 0.4087 for 1% O2).

In Bjorkman and Demmig’s experiments only 8.86cm2 of the 10 cm2 leaf disc was exposed to light because of shading by O-rings round the circumference of the perspex (plexiglass) roof of the chamber. Ifs roof without O-rings (or O-ring grooves) is used, the full 10 cm2 of the disc is illuminated. Open mode measurements were made by using a second leaf disc chamber in series with the first. The second chamber contained an electrode but no leaf and was simply used (in connection with a sophisticated gas-flow device) to monitor the O2 in the gas-stream from the first chamber.

“sophisticated gas-flow”

It will be noted that corrections were made for respiratory O2 uptake whereas this is not done in the procedures described in Appendix 3. The validity of this subtraction is a matter of judgement. If “dark” respiratory O2 uptake occurs at an unchanging rate in the light, subtraction of a constant “dark rate” will not affect the slope of the rate v light curve from which the quantum yield is derived. If measured respiratory uptake changes as a function of light intensity, it will affect the slope (and hence the quantum yield) but it has to be assumed that the “dark” rate is an accurate measure of the “light” rate and it is by no means certain that this is so. Either way, the correction is unlikely to have a substantial effect on the quantum yield values, especially if these are estimated from the slope of the line above the light compensation point. Quantum yield determination based on points above the compensation point (i.e. on net O2 evolution) have sometimes been preferred because, in many tissues, a pronounced “Kok effect” (a change in slope at very low light intensities) may be observed. This is believed to reflect an interaction between respiration and photosynthesis (Section 23). 19(b) Doing it with Chloroplasts and Algae Although the leaf-disc electrode was obviously designed with leaf discs in mind it has been used for other purposes. Zoran Cerovic (Institute of Botany, University of Belgrade, Yugoslavia) for example, has supported intact spinach chloroplasts on filter discs and used these as you would a disc. Similarly, Avigard Vonshak (of the Ben-Gurion University of the Negev, Jacob Blaustein Institute for Desert Research, Israel) has undertaken quantum yield measurements on Spirulina platensis (on filter discs) before and after photoinhibitory treatment. 19(c) Doing it in the Dark If we look at Fig. 22.1 it is clear that O2 evolution in the light is replaced by O2 uptake in the dark as photosynthesis is replaced (or ceases to be masked) by “dark” respiration. Whether or not “dark” respiration occurs in the light as well as in the dark is a bit like trying to decide if the light inside the refrigerator is on permanently or only when the door is open. As usual, much depends on what is meant by dark respiration. There is no doubt that if 14C labelled intermediates of the tricarboxylic (Krebs) cycle are fed to green leaves, the label spreads to all of the other intermediates in the light as well as the dark (Graham and Walker, 1962). Conversely, it is difficult to conceive that such a major metabolic event as photosynthesis is without effect on respiration (cf Section 19a(iv)). This, however, is not the place to re-kindle old controversies and the intention of this section is simply to remind the

reader that the leaf-disc can be used for following respiration as well as photosynthesis. Oxygen uptake in the dark immediately following illumination is faster than it is after prolonged darkness. This may be due, in part, to increased availability of respiratory substrate but it may also reflect the operation of control mechanisms. Temperature artefacts must also be considered. These fall into two categories. One is to do with inadequate temperature control because it is difficult to avoid any heating of the chamber during bright illumination (Section 14) and a small artefact of this nature (caused by temperature changes in the sensor) can often be observed if a leaf-disc is replaced by damp capillary matting. The second is to do with heating of the leaf which is also difficult to avoid entirely and this might be expected to affect the rate of respiration. With due regard to these points, however, respiration may be measured and, provided it will fit in the chamber, anything which respires (e.g. a seed or an insect or a politician) can be monitored. Alternatively, a slow gas-stream can be passed straight through a larger vessel, containing respiring tissues or organisms and then through a leaf-disc chamber, for measurement of oxygen. Given the fact that research funding in some parts of the world is fast approaching a nadir the fact- that polarography of this sort is generally less expensive than many alternatives may not be an unimportant consideration in this context.

References

References Bjorkman, O. and Demmig, B. (1987) Photon yield of O2 evolution and chlorophyll fluorescence characteristics at 77K among vascuolar plants of diverse origins. Planta, 170, 489-504. Ehleringer, J. (1981) Leaf absorptance of Mojave and Sonoran Desert plants. Oecologia 49, 366-370. Ehleringer, J. and Bjorkman, O (1977) Quantum yields for CO2 uptake in C3 and C4 plants. Plant Physiol. 59, 86-90. Ehleringer, J. and Pearcy, R.W. (1983) Variation in quantum yield for CO2 uptake among C3 and C4 plants. Plant Physiol. 73, 555-559. Evans, J.R. (1986) The dependence of quantum yield on wavelength and growth irradiance. Aust J. Plant Physiol, Vol 14 (1). In press. Franck, J (1953) Participation of respiratory intermediates in the process of photosynthesis as an explanation of abnormally high quantum yields. Arch. Biochem. Biophys. 45, 190-229. Gaastra, P. (1959) Photosynthesis of crop plants as influenced by light, carbon dioxide, temperature, and stomatal diffusion resistance. Mededelingen van de Landbouhogeschool te Wageningen 59, 1-68. Graham, D. and Walker, D.A. (1962) Some effects of light on the interconversion of metaoblites in green leaves. Biochem. J. 82, 554-560. McCree, K.J. (1972) The action spectrum, absorbance and quantum yield of photosynthesis in crop plants. Agric. Meterol 10, 443-453. Sharp, R.E., Matthews, M.A. and Boyer, J.S. (1984) Kok effect and the quantum yield of photosynthesis. Plant Physiol. 75, 95-101. Walker, D.A. and Osmond, C.B. (1986) Photon yield of O2 evolution photosynthesis in vivo with a leaf disc electrode: correlations between light dependence of steady-state photosynthetic O2 evolution and chlorophyll a fluorescence transients. Proc. R. Sec. Lond. B 227, 267-280. Warburg, O. and Burke, D. (1950) The maximum efficiency of photosynthesis. Arch. Biochem. Biophys. 25, 410-442.

PART E

COMPUTERISED MEASUREMENT OF RATE AS A FUNCTION OF PHOTON FLUX DENSITY

The previous chapter describes a procedure for the determination of quantum yield in which the leaf-disc electrode is used to measure oxygen, and light intensity is changed by employing a quartz halogen source and neutral density filters. What follows is a general description of an automated procedure for the determination of the rate of photosynthesis as a function of PFD and the importance of this relationship. Details of a computer program used for this purpose are given in Appendix 3.

20. UNDERLYING PRINCIPLES 20(a) Why do it at all? Photosynthesis is, by definition and in fact, a light driven process. It is self-evident that there is no photosynthesis in the dark and a good bet that the rate of photosynthesis will increase as the light intensity increases. Thereafter, supposition and inference must give way to measurement if the relationship between photosynthesis and light is to be properly understood. Surprisingly, although it has had a great deal of attention, there is still much to be learned about it. What is even more important is the fact that this relationship is not invariable and that the manner in which it changes can tell us a great deal. If we plot rate as a function of “light intensity” (more properly “photon flux density”) we can learn something about “dark” respiration and the way that it is affected by light. We can put a value on the light compensation point (the point at which photosynthetic O2 evolution balances respiratory O2 uptake). The initial slope of the rate v PFD plot gives us a measure of quantum yield (maximum photosynthetic efficiency). We can, within the scope of our measuring system, arrive at an approximation of how fast a plant might photosynthesise under given conditions. Hidden within the broad relationship there is additional information, which can be revealed by appropriate analytical methods (Walker, 1988). 20(b) Roofs and Ceilings Many houses have a roof and a ceiling. The roof keeps out the weather and is usually pitched at an angle so that rain runs off it. The ceiling is usually horizontal, serving to isolate the roof space from the room below for purposes of insulation and aesthetics. Blackman (1905) in particular, sought to define the factors which limit photosynthesis in similar terms and there is still much to be learned from a comparable exercise.

20(c) The Roof In the present context light intensity is now usually defined as photon flux density (PFD) and expressed as µmole quanta.m-2.s-1. In other words it is a rate measurement - expressing the rate of arrival of photons (quanta or “parcels” of electromagnetic radiation in the visible wavelength range - Section 26b) as the number which would be intercepted, by one square meter of surface, each second. Photosynthetic rate can be expressed in similar terms i.e. as µmoles O2 (or CO2) evolved (or fixed) per square meter of leaf surface per second (µmoles.O2.m-2.s-2). If photosynthesis is measured at different light intensities the rate of photosynthesis can therefore be plotted as a function of light intensity (or PFD) in these terms so that one rate can be directly related to the other. The initial slope of this relationship is a measure of maximal efficiency or quantum (photon) yield. This is the roof of our photosynthetic house. The maximum rate is the ceiling (Fig. 20.1).

Figure 20.1 Roofs and Ceilings. A typical rate v PFD plot. The intercept on the vertical axis is a measure of dark respiration, that on the horizontal axis is the light compensation point. The initial slope is a measure of quantum yield and its reciprocal, quantum requirement. The curve lies within two constraints, a sloping “roof” and a “horizontal ceiling” (the “true ceiling” or maximum rate). The roof is a thermodynamic constant pitched at an angle dictated by the maximal photosynthetic efficiency (in this instance a nominal quantum yield value of 0.111). The ceiling represents the absolute maximum rate of carbon assimilation. For purposes of comparison, a lower or “nominal” ceiling, imposed by the rate of carbon assimilation at 800 µmole.quanta.m-2.s-1 is employed. To some, it will be immediately apparent why the initial slope of the rate v PFD plot is a measure of efficiency. Others may be helped by an everyday analogy. The efficiency of a motor vehicle can be expressed in terms of its fuel consumption. One that does 30 miles to a gallon of fuel would be more efficient than one which did 10. Such efficiency would be most easily be measured by putting precisely one gallon (or one litre) of fuel into the vehicle’s tank and seeing how far it would go before the tank

Figure 20.2. Illustrates the distribution of incident light between four layers of chloroplasts in a leaf which is illuminated from above and the corresponding rate v PFD plots, assuming a strict Blackman (roof and ceiling) relationship. Curve A, bottom left, is a simulation of the relationship which would be observed if the leaf was illuminated from the lower (abaxial) surface and the departure from the Blackman relationship consequent upon the differences in photosynthetic capacity between the chloroplasts in the 4 layers. Curve B, bottom right, shows the extent to which this effect would be diminished if the chloroplasts from the 4 layers were homogenously distributed in one layer (Terashima, personal communication). was empty. If accurate instruments were available which would measure both the speed (i.e. the rate of progress of the vehicle) and the rate of fuel consumption, it would also be possible to plot one rate against the other. The initial slope would then be a measure of efficiency. For example, if the vehicle used 1 gallon of fuel in one hour and traveled 30 miles during the same period the fuel consumption efficiency could be expressed as 30 miles per gallon. The corresponding value for photosynthetic efficiency might be 0.1 molecules of O2 evolved (or CO2 fixed) per quantum (or photon) of light absorbed. Leaves and motor vehicles have a common feature in that these relationships are non-linear in both - i.e. there is a departure in linearity (a decrease in efficiency) at higher rates, because more and more energy is being dissipated as heat rather than accomplishing useful work.

20(d) The Ceiling As we press the accelerator in a motor vehicle we increase the rate at which fuel is supplied to the engine and therefore the rate at which the vehicle can travel. Eventually, however, the engine will reach its maximum capacity for utilising fuel, no matter how rapidly the fuel is supplied. Green leaves behave in a broadly similar fashion. They will photosynthesise faster and faster as they are given more and more light but eventually the increment in rate per increment in light will become imperceptible. A ceiling will have been reached. This ceiling is not set at a finite height like that of a house but will move upwards or downwards according to environmental factors such as temperature and CO2 availability (C4 plants, which incorporate a CO2-concentrating facility, behave differently from C3 plants for this reason). Moreover, the ceiling is not usually horizontal but tilted upwards at higher PFD’s. It does not necessarily follow, however, that the ceiling is really as tilted as it might seem. A given photosynthetic machine must have a finite capacity under given conditions. Once reached, under unchanging conditions, there is no reason why this value should be exceeded. This implies a horizontal ceiling. One reason why it is often not horizontal, is that a leaf usually contains several layers of chloroplasts so that when the photosynthetic machinery in the upper layer (i.e. the layer facing the incident light) has already reached its maximum capacity, the chloroplasts in the lowermost layer may still be less than saturated. Moreover, the chloroplasts which are located in the loafer layers of cells will function like those in a “shade” leaf because they have developed in a shaded environment (Laisk, 1977; Terashima and Saeki, 1985; Terashima et al, 1986; Terashima and Takenaka, 1986). Complex regulatory factors , such as those which bring about “long induction” (Rabinowitch, 1956; Walker, 1976), may also contribute to the tilting of the ceiling. Nevertheless, while recognising this inevitable gulf between theory and practice it is still useful to regard these limiting factors in these terms. Accordingly we shall visualise the rate v PFD plot as being constrained by a roof and a ceiling. As we shall see, the pitch of the roof is a thermodynamic constraint and is determined by the efficiency of the photosynthetic machinery. The ceiling is a metabolic constraint and is determined by the capacity of the photosynthetic machinery to utilise light energy. 20(e) How Steep is the Roof? As already noted (Part D) little in biology has occasioned so much argument as the answer to this question. Here we shall start with the simple premise that photosynthetic electron transport from water to NADP is in general accord with the Z-scheme of Hill and Bendall (1960) and that Einstein’s Law of photochemical equivalence demands that one photon will initiate one photochemical event. Given that the Z-scheme incorporates two photosystems and that it is necessary to transport 4 electrons through each photosystem in order to evolve one molecule of O2 2H2O + 2NADP

2NADPH2 + O2

it follows that the minimal quantum requirement is 8 and that the maximal quantum efficiency (the reciprocal of the requirement) is 1/8 (or 0.125). This defines the pitch of the roof. If the Z-scheme is an adequate description of photosynthetic electron transport and if what is measured accurately evaluates its function, a quantum efficiency of 0.125 cannot he exceeded. In practical terms it is also difficult to believe that maximal

efficiency can ever be achieved. There is more to photosynthesis than NADP reduction. Except in the very short term, what is measured embraces cytosolic events such as sucrose synthesis. For the present, at least, there is much merit in accepting a nominal quantum requirement of 9 (a quantum efficiency of 0.111). Bjorkman and Demmig (1987) reported an average value of 9.37 (a quantum efficiency of 0.106) for 37 different

Figure 20.3. The Z-scheme by analogy. Two hammer blows (quanta) are needed to project each ball (electron from the starting point (the ground state) to the net (the electron acceptor). Lifting four balls from ground to net would require 8 hammer blows. species. This important finding not only adds credibility to a value of about 9 (which is consistent with the underlying biochemistry as it is presently understood) but also has important implications. Much has been said about the need to improve plant productivity and the extent to which this might be achieved by changing metabolic processes to advantage. Unless Bjorkman and Demmig’s results are substantially refined by future experience (which seems most unlikely) any such improvements will not result from greatly improved efficiency. Evolution, it would seem, has made photosynthetic energy transduction about as efficient as possible, despite the energy losses involved. The molecular biologist seeking improvement would have little or no genetic variation to start from since each species seems to be about as efficient as the next. Redesigning plants to advantage by modifying existing mechanisms is feasible. Redesigning de novo, in entirety, is still in the realms of science fiction. However, if it is not a practical proposition to steepen the pitch of the roof it does not mean that all hope of improvement is gone. Raising the height of the ceiling is not unthinkable nor is improving the extent to which photosynthesis approaches the constraints imposed by the roof on the one hand and the ceiling on the other. 20(f) Relative and Absolute Quantum yield If we plot rate v PFD for incident light the initial slope gives us a measure of relative quantum yield. If we plot rate v PFD for absorbed light we get absolute quantum yield. Bjorkman and Demmig’s (1987) values were corrected for transmission through the leaf and for reflection from the surface. If measurements are made in white light without this correction the values for (apparent) quantum yield may be as much as 15% lower and, if absolute values are needed, corrections for transmission and reflection

based on the use of an integrating sphere must be applied. For most purposes, such corrections may not be necessary. Relative (uncorrected) quantum yield is a valid measurement in its own right. For example, if the consequences of stress, or experimental intervention, are to be examined it will be the change in quantum yield which will attract attention and, in most circumstances this change will be the same irrespective of whether or not corrections for transmission and reflectance are made. When 660 nm light, from LEDs (14c) is used in measurement, the difference between relative and absolute quantum yield is unlikely to be much more than 10% and changes within this correction (such as increased reflection or decreased transmission) will be accordingly less, if they occur at all. 20(g) Why Trouble to Automate Measurement? Another immediate and obvious question is why any one should feel that there is a need for automation. A driver, born and bred to manual gear changes, may disdain a vehicle with an automatic shift because all automatic shifts incur some loss of power and, what is much worse, they take some of the decision making out of the driver’s hands. Their only obvious advantage is that they require less skill to operate. Happily, automated measurement of quantum yield has many more advantages to offer. The non-computerised procedures described in Sections 18 and 19 are not difficult. Anyone can learn how to do these in an hour or so but they are tedious and call for a great deal of concentration if they are to be done in the most effective way. Sitting and waiting to change neutral density filters is not the most exciting way of brightening an otherwise dull afternoon and if your mind wanders to some consideration of the meaning of life or if you try to catch up on your correspondence you are inviting what could be a modestly disastrous lapse in concentration.

“brightening an otherwise dull afternoon”

So automation ends all that. You offer a piece of leaf to a machine and it does the rest. You can write that letter, long overdue, to your mother in Falmouth, Lucknow, Alice Springs, Wurzburg or West Lafayette without fear of error. Moreover, while computers are not yet renowned for their intelligence, they can do some simple tasks much better than people.

One such task is split-second timing and this is very important in the present context because it allows the timing of repeated measurements to be very precise. Reproducibility is of the essence in following the change in quantum yield which results from experimental intervention. This, indeed, goes to the heart of the automated approach. If you wish to make an occasional measurement of quantum yield the manual procedure is fine and has infinite flexibility. In general, however, quantum yield measurement for its own sake is an esoteric pursuit best left to the expert whose work currently indicates that, in the very best circumstances, you might expect your C3 or CAM leaf (Adams et al, 1986) to give a value close to nine. Like Warburg and Burke before them (1950) Bjorkman and Demmig (1987) may be proved wrong but unless we are prepared to reject the entire basis of contemporary theory of photosynthetic electron transport (see e.g. Figs. 8.1,8.2 and 20.3) it seems unlikely that they will be far wrong. For many workers, much more interest lies in the way in which quantum yield might be affected by environmental stress or by experimental intervention of one sort or another. The entire shape of the rate v PFD plot is also full of information but this information can only be properly elicited by comparing one species, or variety, or ecotype, with another and by examining the manner in which it changes with circumstance. Such comparisons demand a great many measurements and minimal variation in procedure. Here the computer comes into its own, not only because it revels in these simple repetitive tasks, which would soon make you feel that you were working on an assembly line, but also because of the many analytical procedures which it can be programmed to undertake. 20(h) How Does it Work? How, you might ask, does the automated system work? As before (Sections 18 and 19), the leaf-disc electrode is used to measure photosynthetic oxygen evolution but now a high-intensity LED source is substituted for the Bjorkman lamp (p. 49). As we have already seen (Section 14c) LED arrays have much to recommend them in photosynthesis work. They are a relatively cool source of light and this simplifies temperature control. They switch on very quickly, facilitating fluorescence measurement. In the present context they have additional advantages. In order to follow the way in which the rate of photosynthesis varies in its relationship to PFD it is necessary to measure PFD at the leaf surface as accurately as possible. Quantum sensors are now technically advanced but it helps if all of the light which is being offered to the leaf is “photosynthetically active”. Red LEDs have peak emission at about 660 nm, close to the red absorption maximum of chlorophyll at 680 nm. Quantum sensors can be calibrated to measure light of this quality without uncertainties about what fraction of the incident light is non-photosynthetic. Unlike a tungsten source which, although admirable for many purposes, delivers more heat than light, LEDs emit very little red light at wavelengths which will not be absorbed by the photochemical apparatus of the leaf. Another major advantage is that there is a near-linear relationship between light emitted and current supplied to the LEDs. This means that the computer has a very easy job if it is asked to change PFD by controlling the electrical supply. A tungsten source is not suitable for this purpose even though the changing quality of the light which it gives out at different currents can be defined by optical filters. (The quality of LED light does not change significantly as the electrical supply is varied). The main problem with a tungsten source is not merely the fact that the relationship between light and supplied current is non-linear but also because it changes, in a very complex way as the filament changes

“crafty electronic circuitry”

temperature. The LED also suffers from instability as a result of heating but to a much lesser extent because the ratio of emitted light to heat is much more favorable than it is for a tunsten or quartz halogen source. Moreover, it is relatively easy to put together crafty electrical circuitry which incorporates feed-back control so that changes of this nature can be largely corrected. This then is a large part of the automated measurement but, as we shall see, much less than the full story. What we do, in effect, at this stage is to offer a leaf-disc to the apparatus and sit back and do whatever we please while it (with the help of the computer) measures photosynthetic O2 evolution over a range of pre-selected PFDs (see 20k). 20(i) What Are The Drawbacks?

“offer a leaf to the apparatus”

LEDs have many virtues but they were not, in common with almost all other light sources, developed for photosynthetic work. Existing LEDs (or combinations of LEDs) are not yet capable of supplying as much light as ace would wish. It is true that they will draw much more current and give much more light at very low temperatures (e.g. in liquid N2) but this is not yet a practical proposition. Even so the Hansatech high-intensity LED source will give about 950 µmole.quanta.m-2.s-1 of red light at the leaf surface (i.e. after transmission through the water filter which is an integral feature of the leaf-disc chamber) and this is more than adequate for many purposes. High intensity LEDs are still relatively expensive, however, and the cost of a suitable array is about two to three times greater than a more conventional light source. Secondly the LED array is intrinsically less uniform than (for example) the Bjorkman lamp, comprising, as it does, separate sources rather than one source and a combination of lenses designed to yield maximal uniformity. This drawback is largely offset if it is used in conjunction with a specially designed quantum sensor (Skye Instruments - see Appendix 2) which effectively integrates the PFD over the surface of the window against which the leaf-disc is pressed during measurement. The O2-electrode, of course, measures total O2 evolution from the leaf-disc and it is largely immaterial if some cells receive a little more or a little less light than their neighbors provided that both incident light and oxygen production measurements are integrated over the full 10cm2 area of the leaf-disc. Problems of reproducibility may arise if leaf pieces are used but these can be largely overcome if care is taken to position such pieces in the same place within the chamber each time a measurement is made. Within the 0-125 µmole.quanta.m-2.s-1 range (i.e. within the largely linear part of the rate v PFD plot) much greater uniformity can be achieved by inserting a diffuser between the LEDs and the top window of the chamber. 20(j) What Are The Advantages? The trivial advantages have already been touched upon. Every research scientist learns, as an apprentice, that tedium will be part of his life and that routine and repetitive work will not only be inescapable but will tar exceed those rare moments of insight or accomplishment which makes it all worth while. That is not to say that tedium should be acorn like a hair-shirt, in order to mortify the flesh. There is everything to be said for making research a pleasure because there is little likelihood that your salary will encourage you to do it for money. Automated measurement of quantum yield is not only more precise than manual measurement but a great deal less boring. One particular selection of PFDs and time intervals, which has proved useful and productive, takes only 16 minutes to run.

“every research scientist learns”

This is time enough to think about the purpose of the experiment, to take a coffee or write the first paragraph of that letter. Nevertheless, these are the trivial advantages. The teal advantages are in data analysis as well as in data acquisition and storage. First of all, you will certainly wish to plot rate of photosynthesis as a function of light intensity (PFD). In the manual procedure, rates have to be derived from a pen-recorder chart and plotted by hand. This will take you 60 to 90 minutes (according to your aptitude for mental arithmetic and your dexterity with pen or pencil). The computer does this virtually immediately. Quantum yield is derived from the initial slope of this relationship (Section 27 and Appendix 3). The PFD for 50% and 90% of the maximal rate can be derived, at the press of a button, as can the “light utilisation capacity” etc (Appendix 3). Plots can be immediately re-drawn on different scales (the computer always selects what it believes to be the most appropriate scale but, lacking intuition or telepathy it is not to know what is in your mind). An extremely useful facility is the ability to call up previous data so that this can be “over-plotted” for purposes of comparison (Fig. 20.4). These and other devices, which will be described below, extend the versatility and usefulness of measurements enormously.

time to write that letter

Figure 20.4. Rate v PFD for Sun and Shade leaves of Avocado. (a) To illustrate both the differences which can often be observed in the behaviour of leaves of the some species if they have been grown in a different light environment (in this instance, leaves from outside and inside of the canopy of a single tree) and the ability of the computer to facilitate such comparisons (sec Appendix 3) by allowing previously stored data to be “over-plotted”, on the same axes, using any chosen scales Automation puts rate v PFD determinations into the category of a routine measurement which can be undertaken by the non-specialist. It becomes possible, really for the first time, to examine how this relationship changes with time, to make comparisons which could not otherwise be contemplated and to extract the last jot of information from the results. PFD for 50% of “maximal” rate

21. MEASURING RATE AS A FUNCTION OF PFD 21 (a) Choice Of CO2 Concentration etc. “Entia non sunt multiplicanda”

If appropriate carbonate/bicarbonate buffers are employed the leaf-disc electrode can be used to measure photosynthesis in concentrations of carbon dioxide close to those in air but the instrument was originally designed for use with saturating CO2 and there is much to be said for using it in this way for this particular purpose. The most compelling reason is Occam’s razor which, loosely interpreted, ask “why make life more difficult than it is already?”. If we wish to examine the manner in which the rate of photosynthesis responds to increasing light intensity interpretation will be simplified if co-limitation by other facts is avoided. How much CO2 is needed to achieve optimal saturation in these circumstances will, in the end, have to be determined by experiment for each and every leaf but 5% is often appropriate. As previously noted (4c) this concentration is readily achieved by enclosing capillary matting moistened with 1.0 M NaHCO3 in the chamber or by using exhaled air or both. If a mixture containing 5% CO2, 21% O2 and 74% N2 is available on tap it is probably to be preferred because it can be pre-humidified at a selected temperature and this simplifies everything. The use of bicarbonate buffers as a CO2-source has implications for calibration which should be borne in mind. High CO2 can effect the electrode response and therefore the instrument should be calibrated in the same CO2 concentration as that employed in the assay. If CO2 is generated within the chamber, O2 can be displaced if the taps to the external atmosphere are not immediately closed and injection of 1 ml of gas containing (e.g.) 5% CO2 and (20% O2 rather than 21% O2) during calibration could lead to corresponding over-evaluation of the O2 subsequently evolved in photosynthesis. Carbon dioxide at concentrations as high as 5% may induce stomatal closure but the concentration gradient is so steep that the closed stomata may offer no sensible barrier to gaseous exchange. Photorespiration will also be almost entirely suppressed because of the competitive nature of the oxygenase reaction (Edwards and Walker, 1983; Walker et al, 1986). [The enzyme Rubisco (ribulose l,5-bisphosphate carboxylase/ oxygenase) which catalyses the carboxylation of ribulose 1,5-bisphosphate (RuBP) in photosynthetic carbon assimilation (Walker et al, 1986) also catalyses the oxygenation of the same substrate. Carboxylation leads to the formation of two molecules of 3-phosphoglycerate This is used both to regenerate the CO2 -acceptor (RuBP) and to form the immediate end products of photosynthesis such as starch and sucrose. Oxygenation yields one molecule of PGA and one molecule of phosphoglycolate which enters a complicated sequence of reactions (Tolbert, I971) which finally ensure that some newly fixed carbon which would otherwise be lost is returned to the chloroplasts. Because oxygen and CO2 compete at the active site of Rubisco, these photorespiratory reactions are largely repressed by high concentrations of CO2 or by low concentrations (2%) of O2]. 21(b) The Unsteady State

“The Unsteady State”

The rate of photosynthesis changes with time. If a leaf is abruptly and brightly illuminated after a prolonged period of darkness there may be little discernible O2 evolution at all for the first few minutes and, after this initial lag or short incubation period, a gradual increase in photosynthesis

(“long” induction) may continue for many minutes or even hours. Induction may also follow abrupt upward or downward changes in light intensity. The problems which are experienced in going from prolonged darkness into full light can be largely avoided by pre-illumination or “wake up”. However, prolonged photosynthesis may result in feed-bad inhibition or other changes in the physiological status of the leaf. It is obviously desirable to avoid measurement during such changes but the true steady-state may be largely illusory. For the present purpose it is pointless to measure during the period of negligible photosynthesis which characterised “short” induction but a leaf which has been too rudely awakened may be as misleading in some regards as one which is still half asleep. This has to do with the impact of light on “dark” respiration. 21(c) The Closed Refrigerator Door Conundrum. Does the light in the refrigerator stay on when the refrigerator is closed? Answering this question would not call for inspired experimental science but the impact of light on “dark” respiration is still more of a matter for argument and conjecture than definitive statement (Graham, 1980; Graham and Chapman, 1979). If respiratory O2 uptake remained unchanged on going from darkness through progressively increasing light intensity it would have no impact on the slope of the rate v PFD plot from which quantum yield is derived. At present no one knows for sure what does happen in these circumstances but there is no doubt that photosynthetic carbon assimilation has an impact on cytosolic events and vice versa (Sivak and Walker, 1986). For example “long” induction can be shortened by feeding Pi and may be largely attributable to the changes in Pi supply associated with the regulation of sucrose synthesis (Cseke et al, 1984; Stitt et al, 1987) the principal mechanism for recycling of Pi. Oxygen evolution below the light compensation point can also be influenced by the “Kok effect” (Section 23) and “light enhanced” respiration (Section 22).

22. LIGHT ENHANCED RESPIRATION Oxygen uptake is low if measured in a leaf which has been kept in darkness for several hours. After induction (10) and a period (say 5 minutes) of strong illumination, respiration is much faster (than before) if the leaf is then abruptly darkened. Thereafter the rate of O2 uptake declines rapidly during the first few minutes of darkness (Fig. 22.1) and then more slowly until it eventually falls to its original value. As measured with the leaf-disc electrode, part of this “O2 uptake” is artefactual and related to the changes in temperature which are an inevitable consequence of strong illumination. Thus black cloth or dead leaf tissue exhibits a small apparent O2 evolution in the light and a corresponding decrease in the O2 signal on darkening. Nevertheless, when due allowance has been made for this artefact, light enhancement of dark respiration can still be discerned. The nature of this enhancement remains unexplained. In bright light, triose phosphate will pour through the Pi-translocator (Fig. 12.7, Sections 41g and h) on its way from stroma to cytosol. Once there, it will undoubtedly influence cytoplasmic events in complex ways because of its impact on metabolite status, ATP/ADP ratios etc. Whether or not these changes lead to enhanced dark respiration (as the O2 uptake data suggests) or to a light inhibition of “dark” respiration, as some believe, is still uncertain. It is clear, however that a “Kok effect” (Section 23) can sometimes be observed.

Figure 22.1. Light-enhanced “Dark” Respiration. This figure shows the course of O2 evolution following illumination at zero time. Following an initial lag or induction period lasting about 3 mins (seen most clearly in the differential, dO2/dt) O2 evolution reached a quasi steady-state of 30 µmoles.O2.m-2.s-1. Following a brief dark interval of 1 min, photosynthesis was resumed after a scarcely perceptible lag (differentiation was over +/- 12.5 sec). Finally, after 6.5 min, the leaf was darkened and during the next 3 minutes the apparent rate of dark O2 uptake declined from -6.84 (at the point that the dotted horizontal coincides with the differential) to –2.37. It is this light-enhancement of the apparent rate of dark O2 uptake which contributes to the departure from linearity, near the light compensation point, in Fig. 23.1. The “Kok Effect”

(after Sharp et al, 1984)

23. THE “KOK EFFECT” This is named after Bessel Kok who was the first to observe what he interpreted as an inhibitory effect of light (Section 19a(iv)) on dark respiration in Chlorella (Kok, 1948; 1949; 1951). Fig. 23.1 shows a departure from linearity in O2 evolution below the light compensation point which was most pronounced when measurements were made, firstly in darkness and then in increasing PFDs, immediately following a period of bright illumination. It could be argued that this is not a true Kok effect but there is no doubt that it could be easily mistaken for one even though the relationship is a smooth curve rather than a discontinuity (see left). In turn, this immediately prompts other questions. Is the “Kok effect” real or an artefact of measurement? Is it a universal phenomenon and, if not, why not? Is there only one effect or several? None of these questions can be properly answered yet but some things are clear. Firstly the Kok effect is by no means a universal phenomenon. Emerson (1958) was even sceptical about its existence, stating “The small amount of positive evidence for the “Kok effect”, together with negative results in several instances of search for confirmatory evidence, leads to the conclusion that a substantial difference in quantum yield above and below the compensation point is not a general phenomenon. If it is something associated with

special conditions, these conditions have yet to be specified. It seems equally possible that the apparent positive evidence arises from changes in rate of respiration between the dark and light intervals chosen for calculation of rate of photosynthesis”. Decker (1957), like Van der Veen before him (1949), observed a discontinuity but preferred the interpretation that dark respiration was constant below the light compensation point and thereafter increased with increasing light intensity. Sharp et at (1984) found the discontinuity to be present even when respiration was constant in the dark (i.e. when effects such as those illustrated in Fig. 23.1 could not be advanced in support of an alternative explanation). Of course, it would be most unwise to seek to reinterpret changes in CO2 exchange, made in very different circumstances, in terms of the changes in O2 illustrated in Fig. 23.1.

Figure 23.1. Rate v PPD in quantum yield range (0-125 µmole.quanta.m-2.s-1 ) for cape weed Arctotheca candula (L.). Single leaf pre-illuminated for 5 minutes at high (900 µmole.quanta.m-2.s-1) moderate (125 µmole.quanta.m-2.s-1) and low light (12.5 µmole.quanta.m-2.s-1). Measurements were made consecutively in the order “moderate”, “high”, “low”. Note the departure from linearity at low PFDs (which is most marked when measurements are commenced immediately after pre-illlumination at high PFDs) but otherwise the reproducibility of the data. At the same time it must be noted that experimental approach is central to this issue. Sharp et al (1987) avoided the rapid decline in the rate of CO2 efflux from sunflower leaves which occured in the first 2 hours of darkness (after a 14 hour photoperiod) and measured respiration in a “relatively stable” subsequent period, reporting values of net photosynthesis (after as much as 45 minutes in PFDs up to 30 µmole.quanta.m-2.s-1 only when “the rate of respiration in the dark was similar at the beginning and the end of the experiment”. This was an admirably rigorous attempt to preclude the “temporal changes in dark respiration” which caused Emerson (1958) and Heath (1969) to question the reality of the Kok effect. Whether or not anything like normal photosynthesis would occur in such circumstances is another matter. Induction (Section 10) is still not fully understood (Walker, 1981) but it is clear that it involves light activation of enzymes, build-up of dark depleted substrates, and the supply of Pi from the cytosol to the chloroplast (Walker, 1976; Sivak, 1987;

Sivak and Walker, 1985, 1986; Walker and Sivak, 1987). If a sunflower leaf grown in relatively high light (900 µmole.quanta.m-2.s-1 in the experiments of Sharp et al, 1984) is illuminated at 30 µmole.quanta.m-2.s-1 (or below) following 2 hours or more of preceding darkness it is most unlikely that cytosolic pools will become filled during the period of measurement even if apparent photosynthesis has become constant. The changes in slope reported by Sharp et al (1984) are too large to be explained in terms of a shift, at high PFDs, from Benson-Calvin cycle activity alone (Fig. 12.6) to Benson-Calvin cycle plus sucrose synthesis. Nevertheless the values of Sharp et al (1984) for quantum yield of about 0.086 (measured in high CO2 above the light compensation point) imply that photosynthesis in these circumstances was, for whatever reason, operating at less than maximal efficiency. Notwithstanding the continuing uncertainty about the nature of the “Kok effect”, results such as those in Fig. 23.1 will have to be borne in mind in making quantum yield measurements. As in the past, the investigator must face an inevitable dilemma. If “dark” respiration is to be measured in the steadystate, several minutes of darkness will be needed in order to avoid light-enhanced O2 uptake and several hours, in many species, before the decline in dark respiration, following illumination, becomes imperceptible. However, even 1-3 minutes darkness will normally be enough to re-establish “short induction” in photosynthetic O2 evolution (Walker 1981). Moreover, re-illumination at high PFDs after a period of darkness, often initiates a “burst” (Fig. 22.1) of O2 (and an associated “gulp” of CO2) within the induction period (Sivak and Walker, 1985). Opinions about the origin of the O2 gulp still differ but there is no doubt that 3-phosphoglycerate often persists in darkness in circumstances in which RuBP falls to near zero, or that pre-formed PGA will serve as an oxidant in suspensions of isolated chloroplasts. If the Benson-Calvin cycle is “switched off” and if the reduction of PGA exists, even as a possibility, there is an obvious danger in attributing gas exchange, during induction, to “real” photosynthesis. Conversely, if induction is avoided by pre-illumination, the possibility of light enhancement of respiration cannot be disregarded. No doubt, these problems will continue to pre-occupy those who wish to continue to refine the measurement of maximal photosynthetic efficiency. If, on the other hand, the initial slope of the rate v PFD plot is primarily of interest as an indicator of environmental stress or experimental intervention there is much to be said for a simple standard procedure (e.g. measuring from 125 to darkness after pre-illumination at 125 µmole.quanta.m-2.s-1). In unstressed leaves, this approach (Appendix 3) can yield reproducible quantum yield values close to those demanded by our present understanding of photosynthetic electron transport. The methods described here permit such determinations to be made, routinely, in less than half an hour.

24. IS THERE A RECOMMENDED PROCEDURE? As usual, there are “horses for courses” and what may be ideal for one leaf may be less than ideal for another. If you take a previously illuminated leaf and measure its rate of photosynthesis as a function of PFD by making 15-20 measurements each lasting one minute you will immediately learn a great deal about its physiology. Some would insist that such short time

Intervals will not permit the achievement of steady-state photosynthesis and they would be right. On the other hand, enzymologists (similarly concerned with the determination of rate as a function of substrate concentration) have traditionally ignored the changes in substrate concentration which occur during measurement. As always, measurement is rarely non-intrusive (p. 95) and compromise is inevitable. In the present context the use of a relatively large number of relatively small PFD increments avoids the creation of disturbances, such as oscillations (Section 12c and Fig. 12.5) which are often associated with major perturbations. The computer is also programmed to ignore the first 10 seconds of the O2 signal following a change in PFD during which time minor perturbations, consequent upon this change, will tend to subside. How to ensure “wake-up” on the one hand and induction on the other is more difficult. After prolonged darkness some leaves display such prolonged induction that the term “short” induction becomes a misnomer and the observer searches, in vain, for a prince to kiss this sleeping beauty into wakefulness. Other leaves may wake up quite readily in the sense that the initial slope is unchanged or largely unchanged if the light intensity is increased without pre-illumination but longer illumination, or repeated assay (Fig. 24.1), may bring about pronounced increases in the rate of photosynthesis at high PFDs.

Figure 24.1. To illustrate the usefulness of the automated approach in following a rapidly changing situation. Six measurements (3 not shown, for clarity) were made successively on a disc from a shade avocado leaf harvested the previous day and kept in darkness overnight. Prior to the first measurement, the disc was illuminated for 15 minutes in 90 µmole.quanta.m-2.s-1 light. The second, third and fourth measurements were consecutive. Between the fourth and fifth there were 6 minutes darkness followed by 1 minute in 900 µmole.quanta.m-2.s-1 light and between the fifth and sixth a further 10 minutes darkness and 2 minutes in 900 µmole.quanta.m-2.s-1 light. Such behaviour could be indicative of reversal, by light, of dark inactivation of Rubisco. Whatever its cause it adds to the difficulties in arriving at an acceptable procedure. The Kok effect and photoinhibition also constitute pitfalls. Prolonged illumination in high light is the obvious way to ensure that the leaf is fully awake but it can induce light enhanced “dark” respiration and/or photoinhibition.

How to ensure “wake up”

As usual, there is no real alternative to being familiar with the idiosyncracies of your leaf. In the absence of such knowledge the best way of achieving wake-up is probably to pre-illuminate the leaf at the highest PFD that you can provided that this does not exceed that which the leaf normally experiences in its immediate natural environment. Once fully awake, the leaf will not usually relapse into deep sleep in 5-10 minutes darkness but this will be sufficient to allow light enhanced dark respiration to subside. Thereafter the leaf should be pre-illuminated at the highest PFD to be employed in determining the initial slope (100-125 µmole.quanta.m-2.s-1) and measurements made by following the rate of O2 evolution as the PFD is decreased from this maximal value. Wake-up will be ensured, light enhanced “dark” respiration will be minimal and will decrease throughout the period of measurement so that, as a percentage error, it will have least impact where it might otherwise lead to greatest distortion (c.f. Fig. 20.6).

25 HOW TO GET THE MOST OUT OF YOUR DATA It is one thing to ask a computer to make life easier for you but quite another matter to conjure real science out of what you might be offered. Measurement is descriptive. Experimental science starts with measurement. Thereafter it is a matter of asking the right questions. Plants are unable to communicate but that is not a major disadvantage. Ask your friend what causes your migraine (or his or hers) and see if you get a helpful answer. If, on the other hand, you were able to ask your friend the right questions, physiologically and biochemically, by perturbation and measurement, you could get the right answers and, at one fell swoop, rid the world of fearful torment and rejoice in new-found wealth, prestige and appreciation. Ask a plant the right questions and you are unlikely to be rewarded so well (although if you came up with a really nice new herbicide you might find yourself in a same league as Croesus, Getty, Gulbenkian and QE2). In any event, asking the questions is down to you. The only way to get answers from a plant is to make measurements. Measuring rate v PFD is only one of a large spectrum of measurements currently available (see e.g. Sections 13c and e) but at least you should ensure that you derive as much benefit from your measurements as subsequent analysis will allow.

26. RATE v PFD UNGILDED As we have already noted, the rate v PFD plot offers a wealth of information in its simplest form. The intercept on the vertical (Y) axis is a measure of O2 uptake mostly attributable to “dark” respiration. This value is by no means invariable and the effect of pre-illumination on “dark” respiration and the relationship between photosynthesis and respiration and how this varies from species to species, from variety to variety, according to environmental influences is still one of the major unquantified and uncharted areas of plant physiology (Graham, 1980; Graham and Chapman, 1979; Graham and Walker, 1962). The intercept on the horizontal (Y) axis is no less important. This is the light compensation point. Shade species or shade adapted leaves of sun species may be expected to have low light compensation points but there is more to it than

“a wealth of information”

that. If the pitch of the roof of the rate v PFD plot is as steep as thermodynamics permit (i.e. if the quantum requirement is in the region of 9) a low light compensation point demands a low rate of dark respiration or a pronounced Kok effect or both. If the relationship is linear and the light compensation point remains unchanged despite a change in quantum requirement (as in photoinhibition - Section 32) then the change in photosynthesis must also have affected dark respiration. In short, there is more to the light compensation point than immediately meets the eye - it merits evaluation.

27. THE INITIAL SLOPE As we have seen, the initial slope is of fundamental importance because it is a measure of maximal photosynthetic efficiency (i.e. of quantum requirement and its reciprocal, quantum yield). Some ways of carrying out the actual experiment (see “Is there a recommended procedure?” - Section 24) are better than others but how do we recognise the initial slope when we see it? The human hand guided by the human eye is still as good as anything and the computer programme offers you a “line” or “rule” to help you draw the line of best fit. If you prefer to put your faith in statistical analysis use a least squares regression (Appendix 3) but bear in mind that whatever procedure you use will involve subjective judgement and that there is never a substitute for precise data which will confine errors in judgement. A third possibility is to use a more sophisticated line-fitting procedure which has been instructed to recognise and ignore departures from linearity above and below the PFDs at which such departures might be expected to occur.

28. THE SHAPE OF THE CURVE

Quantum requirement by “Line”

Put aside all impure thoughts and contemplate the nature of the rate v PFD curve. In the first place it is not really a curve at all, certainly not the rectangular hyperbola of enzyme kinetics fame. Often it looks more like two straight lines joined by a curvilinear region and indeed this would be expected if its photosynthetic performance is constrained by a thermodynamic roof (Section 20c) and a metabolic ceiling (Section 20d). Let us suppose that this is the case. It would follow that an efficient or successful leaf would take all that it was offered, i.e. that its rate v PFD curve would be close to the roof and ceiling. Attempts have been made to quantify the realisation of this potential which is partly constrained by thermodynamics and partly by metabolic capacity. Terashima and Saeki (1985), Terashima et. al. (1986) and Terashima and Takenaka (1986) have suggested the convexitivity of the curve as an index. (When the convexitivity index is 1 there would be an abrupt transition between roof and ceiling, i.e. a Blackman relationship. When the index is zero the curve would be a rectangular hyperbola).

One alternative is “nominal light utilisation capacity”. This defines an area bounded by the roof, the ceiling, the horizontal (X) axis and a perpendicular erected at a convenient, high PFD (800 µmole.quanta.m-2.s-1)for measurements made with the high intensity Hansatech LED array). “Nominal light utilisation capacity” is then the area under the rate v PFD curve expressed as a percentage of the area defined by these constraints (see also Appendix 3). In this analysis the roof is pitched at an angle equivalent to a quantum requirement of 9 and rises directly from the origin (Fig. 20.1). Because of dark respiration, no leaf will ever accomplish zero oxygen exchange at zero PFD so that even if the convexitivity of the curve were maximal the nominal light utilisation capacity would be less than 100%. A roof that rises directly from the origin is therefore arbitrary and it will be immediately obvious that a leaf with a low light compensation point (LCP) will approach this particular constraint more closely than one which requires a higher PFD to balance dark respiration. In this regard “nominal light utilisation capacity” is descriptive rather than carrying implications of efficiency. In terms of survival strategy a leaf which gets little light and therefore photosynthesises slowly may be best served by a commensurately low rate of dark respiration. Conversely, a fast growing species in full light might easily do best if it can grow rapidly in order to maximise its light interception and compete most successfully with rival species. In this situation a very high rate of “dark” respiration and a correspondingly high LCP might simply be an indicator of rapid metabolism rather than inefficient utilisation of photosynthetic product. Similar considerations apply to the convexitivity of the curve. A fraction (about 5%) of incident photosynthetically active light is transmitted through leaves. Shade leaves characteristically contain high chlorophyll and this has, correctly or otherwise, been regarded as an attempt by the leaf to maximise light interception. Be this as it may, there must be an optimal response in a given situation. Utilising as much light as possible in a deeply shaded environment would have obvious advantages but a point Photo Flux Density must be reached when excessive investment of valuable resources in chlorophyll Convexitivity: Rectangular hyperbola (θ = 0) could “cost” more than would be gained by increased light interception. Whether or not quantification of the rate v PFD relationship on the basis of nominal light and Blackman (θ = 1) for identical initial utilisation capacity, convexitivity (Terashima and Saeki, 1985; Terashima et. al, 1986; slope and asymptote. Terashima and Takenaka, 1986) or whatever (After Terashima and Saeki, 1985)

will come to assume importance in ecophysiological work remains to be seen but, on the face of it, it would appear to offer more information than some alternatives.

29 ANALYSIS OF STRESS 29(a) Stress and Performance Björkman and Demmig’s (1987) work confirmed the tacit assumption that a leaf is unlikely to photosynthesise well if it is stressed. Much the same can be said of poorly trained athletes, tired executives or ageing professors. Different types of stress must obviously affect leaves in different ways but little has been done to qualify and quantify stress in terms of the rate v PFD relationship. Again nominal light utilisation capacity affords some promise of quantification, particularly if combined with other measurements such as Q-analysis of chlorophyll a fluoresence (Sections 13e and h). Wilting affects the rate at high PFDs more than at low PFDs (Ben et. al, 1987) thereby leaving the initial slope largely intact. Stresses which lead directly or indirectly to photoinhibition (Section 32) will affect the initial slope and, eventually, the maximum rate. Such responses, the manner in which they are imposed, and the form which recovery or acclimation takes if the inflicted damage is not irreversible, can be quantified in terms of quantum yield, nominal light utilisation capacity. 29(b) Shifting the Arbitrary Ceiling As we have seen in Section 28 above, “nominal light utilisation capacity” offers a means of quantifiying the extent to which the photosynthetic and respiratory performance of a leaf allows it to

Figure 29.1. Shade leaf before and after photoinhibition which diminished the “nominal light utilisation capacity” (i.e. the area under the rate v PFD plots expressed as a percentage of the area defined by the roof, ceiling and perpendicular) from 83.5% to 54%.

Figure 29.2. As for Fig. 29.1 showing further progression of photoinhibition to 42% despite removal from bright light and an intermediary stage in recovery at which the area under the curve (the nominal light utilisation capacity) has increased from 42% to 62%. approach a potential defined by thermodynamics on the one hand and metabolic capacity on the other. This analysis can also be used in a comparative way. Figs. 29.1 and 29.2, for example, show ceilings which have been shifted in order to calculate the nominal light utilisation capacity of a leaf before and after photoinhibition. If the arbitrary ceiling is drawn for the lower (photoinhibited) curve the nominal light utilisation capacity is 80% but if it is now raised to the ceiling that would be appropriate for the control (prior to photoinhibition) the nominal light utilisation capacity is only 54%. The control nominal light utilisation capacity was 83.5% so that photoinhibition has, in this case, decreased the nominal light utilisation capacity from 83.5% to 54%. Following removal from high light, photoinhibitory damage continued to progress at first, falling a further 12% to 42% but thereafter maintenance of the leaf disc in low light eventually brought about complete recovery. Fig. 29.2 shows an intermediary stage in recovery at which time nominal light utilisation capacity had increased from 42% to 62%. 29(c). Shifting the Ceiling and the Perpendicular. In some circumstances it may be useful to ask questions such as “What is the nominal light utilisation capacity at low PFDs?” Figs.29.1 and 29.2 illustrate this point. Fig. 29.3(a) is a rate v PFD plot for a sun leaf and if the perpendicular (see also Appendix 3) is moved from 800 to 100µmole.quanta.m-2.s-1) the area under the curve (the nominal light utilisation capacity within the constraints defined by the roof, the shifted perpendicular and the lower ceiling) is only 25.75%. When the same exercise is repeated for a shade leaf off the same tree (Fig. 29.3(b)) the corresponding value is 67%. Comparisons made in this way, which take into account the rate of photosynthesis, the rate of dark respiration (and therefore the light compensation point) provide an extremely convenient basis for evaluating the extent to which a leaf is accommodated to a particular light environment.

Figure 29.3(a). Sun leaf of avocado showing analysis in terms of nominal light utilisation capacity in the 100 µmole.quanta.m-2.s-1 range. The roof is the same as before but the perpendicular in both has been moved from 800 to 100 µmole.quanta.m-2.s-1 ). The ceiling has been lowered to coincide with the rate at 100 µmole.quanta.m-2.s-1 ) perpendicular. The areas under the curves expressed as a percentage of the area confined by roof, ceiling and perpendicular was 25.75%.

Figure 29.3(b). Shade leaf of avocado from same tree as that used in the experiment illustrated in Fig. 29.3(a). Corresponding analysis, in terms of nominal light utilisation capacity, gave a value of 67%.

30. LUX ET VERITAS, A REMINDER ABOUT LIGHT 30(a) Measurement If it is our aim to measure the absolute efficiency of light utilisation in photosynthesis according to the above procedure, it is necessary to know how much light is absorbed by the leaf. This is equal to the incident light,

less the reflected and transmitted light. Corrections for reflection and transmission can be made by using an Ulbricht or integrating sphere. This is a sphere coated with a totally reflecting surface (such as a deposit of magnesium oxide) into which light is introduced. A detector then records the difference in brightness within the sphere when a leaf is substituted for a standard reflecting surface within the sphere. For all of these measurements (and less accurate ones in which a loss of, say, 15% by reflection and transmission is assumed) it is necessary to measure incident light. This is not an easy matter, though it has been made easier by recent advances in instrumentation. Most of the older instruments for measuring radiation were thermocouples, consisting of a number (a ‘pile’ of thermocouples - i.e. a “thermopile”) of alternate junctions between two dissimilar metals such as copper and constantan. An increase in temperature, at such a junction, when it is exposed to light, generates a voltage which is proportional to the difference in temperature between that junction and another which is protected from the light. Such a device is, in fact, an electric thermometer. Thermistors, which are also electric thermometers (but which are based on electronic devices which vary in resistance according to temperature) may also be used. Both are stable, reproducible and independent of wavelength but, because of their intrinsic heat capacity they respond slowly and, at very low light intensities they are less accurate because of heat gain or loss from the surrounding environment which is not engendered by incident light. Radiation meters based on thermocouples or thermistors may be calibrated in absolute units such as watts.m-2 (the unit of “radiant power”). The modern alternative to the thermopile is the photodiode which is a quantum (see Sections 31b and c) sensor. In this instrument, each photon of light which strikes the surface of the detector releases an electron (thus initiating an electron flow or electric current). Because such detectors are less sensitive at short-wave visible wavelengths this compensates for the fact that these (high-energy) wavelengths would otherwise generate more current and this, together with the use of appropriate filters, allows the construction of what is, in effect, a photon counting meter. Such a sensor, screened by filters which exclude light at wavelengths below 400 nm and above 700 nm, will record “photosynthetically active photon flux density” (PPFD). Photon flux density (PFD) is now usually expressed as the number of photons (mole quanta) striking an area of 1 square metre in 1 second i.e. µmole quanta. m-2.s-1 (see Section 30c). 30(b) The Nature of Light “God said, let Newton be, and all was light”. Newton himself suggested that light was comprised of corpuscles travelling through space at a constant speed and that there were different corpuscles for different colours. However, Thomas Young’s experiments, in which two beams of light were used to partially cancel each other (producing interference patterns) showed that light had the characteristics of a waveform. Then, in 1900, Max Planck concluded that electromagnetic radiation is absorbed or emitted only in discrete bundles or particles of energy or quanta. (Quanta in the visible-light range are called photons). The particulate (corpuscular) nature of light was confirmed by Lenard in 1902. He studied the photoelectric effect in which radiation of a given frequency will cause electrons to be ejected from a metal surface. He found that there was a threshold, or critical frequency, below which electrons were never ejected. Above this frequency, increasing intensity only increased the number of electrons ejected, whereas increasing frequency increased the energy of

each electron but not the number. Einstein’s explanation, in 1905, of this effect contributed to his Nobel laureate in 1921. This is that light consists of quanta (or photons) and that, in the photoelectric effect, a single electron can only acquire the energy of a single photon. Increasing the energy content of the incident photons (i.e., decreasing the wavelength of the incident light) will then increase the energy per electron. Increasing the light intensity will increase the number of photochemical events and therefore the number of electrons emitted. This, of course, is the basis of the quantum sensor (Section 30a). 30(c) The Energy Content of Light Quantum theory implies that light must behave simultaneously as a waveform and a stream of particles. The relationship between wavelength (λ), frequency (v) and the speed of light (c) can be expressed as

joules are a girl’s best friend

λ (nm) x v (per second) = c (nm per second) where a nanometer (nm) is 1 x 10-9 metres in length and the speed of light, c, is 3 x 1010cm/sec or 3 x 1017 nm/sec. The energy content (E) of one photon of light can be calculated from the relationship E = hv or E = hc

λ

where h is Planck’s constant (= 6.625 x 10-34 joule. sec) and v is the frequency (= c/λ). Thus for red light at 680 nm, which approximates to the red absorption peak of chlorophyll in vivo, the energy content of one photon is 1 E = 6.625 x 10-34 x 3 x 1017 nm x sec 680 nm = 2.92 x 10-19 joules Clearly the energy content of one photon is a very small value indeed and for many purposes the “einstein” i.e. the energy content of Avogadro’s number (6.023 x 1023) of quanta has become the basis of expression. Similarly, the term “quantum mole” is used to describe Avogadro’s number of quanta, just as a “gram molecule” or “mole” refers to Avogadro’s number of molecules. Thus the energy content of 1 quantum mole of red photons (at 680nm) = 2.92 x 10-190 x 6.023 x 1023 = 176 Kjoules and that of l µmole quantum (i.e. 1 microeinstein) = 1.76 Kjoules x 10-6 = 0.176 joules

Half an Avogadro Pear

Accordingly (since 1 joule. sec = 1 watt), it would take 500 = 2,840 0.176 µmole quanta of 680 nm light to deliver as much radiant power as the photosynthetically active component (wavelengths between 400 and 700 nm) of full sunlight (approximately 500w. metre-2 s-1) - see Section 30d and Table 30.1.

micromoles and microeinsteins

The “einstein”, however, is, in turn, too large a unit for many calculations because full sunlight only delivers a fraction of an einstein per square metre of surface per sec. Accordingly, “micro-einsteins” (µE = 10-6 einsteins) and “micromole quanta” are the preferred units and “photon flux density” (PFD) and “photosynthetic photon flux density” (PPFD) are expressed in µmole.quanta.m-2.s-1 (Section 30a). It follows from the relationship E = hc/λ (above) that the energy content of light is inversely related to its wavelength. Blue light (at, say, 450 nm) will have a much higher energy content than red but, as we have seen in Section 6, this additional energy is not used in photosynthesis because of the extremely rapid radiationless decay of the blue excited state of chlorophyll. This is why, when we are concerned with energy relations, we think in terms of red rather than blue light. It is the red excited state which is the “driving force” of photosynthesis. Even green photons, if absorbed and utilised, give rise to the red excited state of chlorophyll and are therefore just as effective as red or blue photons. Incident green light is less effective than blue or red because it is more likely to be reflected from the surface of the leaf or transmitted through it (see also Evans in Section 17). 30(d) Can We Compare Micromoles with Foot Candles? “Shall I compare thee to a summer’s day?” While it is difficult to compare chalk with cheese we are often called upon to do so. The intensity of light was a matter of interest long before quantum mechanics were

thought of and it is not surprising that nineteenth century man thought in terms of “candle power” just as he thought in terms of “horse power”. There is a great deal in the older literature, which is still of considerable interest today, in which light is not expressed in the units which are now favoured. It is essential, therefore, that the early terminology is not forgotten. However, it follows from some of the foregoing (Section 30c) that absolute equivalence between some units is impossible to calculate because the spectral composition of the light employed cannot be accurately defined. Table 30.1 attempts to compare chalk and cheese by assuming that visible sunlight has a mean wavelength of 575 nm (whereas, in actuality, its wavelengths range from 400 to 700 nm and, although it contains a large blue, component, its spectral composition will change from hour to hour). It also ignores the fact that the spectral composition of “electric light”, though possibly not too dissimilar from “natural” light, will vary with the source, the age of the source, the current etc.

Table 30.1 Full Sunlight Expressed in Several Ways Illuminance

lux 100,000

foot candles 10,000

Photometric

Photosynthetically active component per square metre per second Watts per square metre ergs 500 5x109

joules 500

radiometric (400-700nm)

cals 120

photons 1.3x1021

µmole quanta 2200

quantum (400-700nm)

These comparisons are only valid for sunlight (see text). All of the values in Table 30.1 are measures of flux density at an illuminated surface. Light “intensity”, as such, is a property of the radiating source. For example, a “standard candle” emits a flux of one “candle power” or 4π (= 12.566) lumens. The corresponding value of light received (or flux density) at the surface of a sphere of 1 foot radius with a standard candle at its centre is “1 foot candle” (1 lumen/foot2). If the radius is increased to 1 metre the light is spread more thinly, the flux decreasing according to the square of the distance. Since 1 metre = 3.2808feet and 3.2808 squared = 10.8, it follows that 1 foot candle = 10.764 lux (or metre candles or lumens/sq metre). Similarly 1 phot = 1 lumen/sq cm or 10-4 lux and 1 foot candle = 1,0746 milliphots. In S.I. units the standard candle becomes the “candela” which, similarly acting as a point source, emits 12.566 lumens. The standard source is now taken as a black-body radiator at the melting point of platinum and the candela emits one sixtieth of the intensity of 1 sq cm of such a black-body. Because light is measured (Section 30a) in different ways, and because flux density is expressed in units of amount, time and area, other variations on this theme may be encountered but translation from one to another should be simply a matter of arithmetic.

31. ENERGY, PLANTS AND MAN AN ASIDE ABOUT ENERGY UTILISATION In terms of light power one square metre of the earth’s surface (at sea level on the equator) is rated at approximately 1 kw, of which approximately half (500 w) can be used in photosynthesis. The corresponding mean annual value (24 hours a day, 365 days a year) varies with latitude, altitude, aspect and cloud cover. In the Red Sea area it is about 300 w, in Australia and the United States about 200 w and in the United Kingdom about 100 w. In terms of energy consumption, man is also rated at 100w so that, even in the U.K., he could meet his metabolic energy requirement from 1 square metre, given 100% efficiency of conversion of incident light energy. Photosynthetic conversion of light energy to chemical energy may approach 33%. This is based on the calorific content of sugar (one mole of glucose releases 672 Kcal when burnt in a calorimeter), a minimal quantum requirement of eight (Section 15), and a value of 176joules (= 42 Kcals) per quantum mole of red light at 680 nm. Accordingly, if it is assumed that 672/6 (= 112) Kcals would be required to convert CO2 to “CH2O”, the efficiency of conversion would be 112 x 100 = 33% 8 x 42 If the quantum requirement were 10 (see below) instead of 8, the value would be a more realistic 26%. Since the photosynthetically active component of sunlight accounts for only half of the total radiation and has an average energy content of about 50 Kcal, this decreases the efficiency to 112 x 100 = 11.2% 10 x 50 x 2 If appropriate allowances are made for transmission, reflection, respiration etc., this theoretical value falls again to about 5% and this percentage conversion has actually been approached in the field. (For example, Pennisetum typhoides grown in a closed canopy at Katherine, West Australia, with a daily energy input of 5,100 Kcal/sq metre, showed a daily dry weight increase of 54g/sq metre. This is an efficiency of some 4.5%). The best recorded yields in western agriculture are equivalent to a conversion efficiency of about 1.25% and average yields about 0.4%. It follows from the above that if the quantum requirement were to increase from 10 to 20 (plants need to be at peak performance to approach the theoretical maximum of about 9) there would be correspondingly large decreases in yield. Several types of environmental stress can lead to decreases in quantum yield (i.e. increases in quantum requirement). Such stresses often involve photoinhibitory damage.

32. PHOTOINHIBITION 32(a) What Is It? When a chlorophyll molecule absorbs a photon it becomes excited. As we have already seen this excitation energy is dissipated in a variety of ways. It is dissipated as light (as fluorescence), as heat, or in the accomplishment of chemical work (photosynthesis and related processes). One such related process is breakdown of the photochemical apparatus itself. Electric light-sources can be regarded as the converse of the photochemical apparatus of photosynthesis in the sense that they convert electrical energy into light. They are not able to do this without damage to themselves and, despite being protected by inert gas, electric filaments have a finite life. The photochemical apparatus is also thought to be protected in various ways. The fact that the rate v PFD plot usually departs from linearity at PFDs a little above 100µmole.quanta.m-2.s-1 itself implies the existence of a safe mechanism for diverting excitation energy into channels of thermal dissipation. This seems to be related (Section 13a) to changes in ∆pH (Krause and Weis, 1984) and associated changes in the organisation of the thylakoid membrane (Demmig and Björkman, 1987 and Section 13c). In addition, the thylakoid membranes may need to be kept in a state of more or less continuous repair in order to cope with the rigours of energy transduction as it converts light-energy into electrical-energy. The part of the photochemical apparatus which is most susceptible to photoinhibitory damage is PSII (Fig. 32.1).

“when a chlorophyll molecule absorbs a photon”

Figure 32.1. A diagrammatic representation of PSII and its association with components of the light harvesting complex in the thylakoid membrane. The oxidising (water-splitting) side gives rise to powerful oxidising species which are harboured in the D1 polypeptide which binds QB. Photoinhibition involves damage to this QB binding protein (Courtesy of Jim Barber). Precisely which part of PSII is damaged (and why) is still a matter for argument and conjecture but there is no doubt that one component of this reaction system undergoes more or less continuous replacement in the light or that inhibition of its synthesis (by agents such as chloramphenicol) can exaggerate photoinhibitory damage. The component which turns over rapidly is a 32-kDa protein (i.e. one with a molecular weight of 32 kg)

“it becomes excited”

which is located in the reaction centre of photosystem II and binds QB, the plastoquinone which constitutes the secondary electron acceptor of PSII (Fig. 8.2). Damaged reaction centres are still capable of trapping excitation energy and quenching fluorescence but the excitation energy is preferentially dissipated as heat, rather than initiating photosynthetic electron transport so that both the rate of electron transport, and the rate of fluorescence emission are diminished. (The QB-binding protein is normally involved in the transfer of electrons from the primary electron acceptor, QA, in PSII to the plastoquinone pool. It is also often referred to as the “atrazine-binding protein” because this is the site of action of herbicides such as atrazine, which block the transfer of electrons from PSII to plastoquinone and therefore to the rest of the electron transport system). New protein synthesis is involved in maintaining the integrity of P680 (the PSII reaction centre) implying that damage to the QB-binding protein cannot be brought about in situ but rather that the damaged protein has to be replaced, in the thylakoid membrane, by newly synthesised protein. Such replacement can be regarded as an inevitable part of photosynthetic electron transport just as tyre replacement is an inevitable part of road transport. On a perfect road the rate of tyre replacement is normally so low that it would not be apparent to a casual observer but if a vehicle is pushed to its limits, as in Grand-Prix racing, or if a tyre is stressed by rough terrain, replacement will become more frequent thereby diminishing transport efficiency.

PSII (courtesy of J. Barber)

“pushed to it’s limits” In some circumstances, failure to effect adequate repair may even lead to complete failure and breakdown. Much the same is true of this repair of P680 (or, to be specific, the repair of the QB-binding protein within that reaction centre of PSII). It is in such circumstances that “photoinhibition” becomes apparent. A decrease in variable fluorescence can be detected together with a decline in the rate of photosynthesis which is often most marked at low light intensities - i.e. there is a decrease in the initial slope of the rate of photosynthesis v PFD plot (Fig. 32.2, p. 92) from which quantum yield is calculated. Accordingly, photosynthetic efficiency and quantum yield decrease whereas the quantum requirement (the reciprocal of quantum yield) increases. As photoinhibition progresses, even the rate of photosynthesis at higher PFDs may also become markedly depressed.

32(b) What Causes Photoinhibition? Damage to P680 is almost certainly an inescapable aspect of photosynthetic energy transduction but it may only manifest itself as photoinhibition when repair (of the QB-binding protein) fails to keep pace with wear and tear. Photoinhibition is often initiated, in contemporary laboratory practice, by strongly illuminating a leaf at temperatures between 5 and 10oC, for about 1 to 3 hours. Not all leaves will show the marked decline in quantum yield and maximum rate which characterises photoinhibition as a result of such treatment. Much depends upon the species and the environment in which it has been grown. “Sun species” (i.e. those which normally grow in non-shaded environments) are often very resistent to photoinhibition. On the other hand, if they have been obliged to grow in the shade, they may be more susceptible to photoinhibitory damage. Similarly, “shade species” (i.e. those which do well in shaded environments or normally grow in the shade) are more likely to show photoinhibitory damage if they are brought from deep shade in to bright light. The role of temperature in this relationship is complex. Shade species are not noted for high rates of photosynthetic carbon assimilation and some have been demonstrated to lack the enzymic capacity to maintain high rates of carbon cycle activity.

“Particularly at low temperatures” Particularly at low temperatures it would seem evident that such plants would not be able to dissipate high levels of excitation energy, via carbon assimilation, at rates which would avoid photoinhibition. This notion is also strengthened by the fact that photoinhibition may be induced by bright illumination in CO2-free atmospheres (Powles et. al, 1984). Although photosynthetic carbon assimilation may help to protect against photoinhibition this is only one part of a larger story, however. Tolbert, as long ago as 1971, suggested that photorespiration might dissipate excess photosynthetic energy (photorespiration consumes substantially more ATP and NADP than photosynthesis as such) and Osmond & Björkman (1972), Björkman (1981) and Osmond (1981) have developed this concept in detail. It has also been demonstrated that 1% O2 in N2 (which suppresses photorespiration as well as photosynthetic carbon assimilation) is very effective in inducing photoinhibition in bright light.

While there is little doubt, therefore, that a combination of photosynthesis and photorespiration can partially protect the thylakoid membranes from photodamage it does not follow that these are the sole channels of energy dissipation normally available to the leaf. Sequestration of cytosolic Pi, by feeding mannose or 2-deoxyglucose (Section 16), can bring about marked depression in the rate of photosynthesis without affecting quantum yield (Walker and Osmond, 1986) and phosphate feeding is finally unable to protect against photoinhibition even though it initially increases the rate of carbon assimilation and may delay the onset of a decline in quantum yield (Walker, Osmond and Cleland, 1988).

Figure 32.2. Photoinhibition in Helleborus niger without loss in maximum rate.A leaf-disc Helleborus niger growing in deep shade(2 µmole.quanta.m-2.s-1) was assayed at 20oC before and after 2.5 hours in 10,000 µmole.quanta..m-2.s-1 at 8oC. The initial slope is affected (i.e. the quantum yield has fallen significantly from 10.5 in the control to 24 following photoinhibition) but there has been no decline in rate at the highest light intensities. Krause and Cornic (1987) have drawn attention to the fact that in experiments by Powles et. al (1984) photoinhibition only occurred when CO2 was depleted below a critical limit of about 50 bar. At this concentration of CO2 Phaseolus vulgaris was only able to maintain 40% of its rate of photosynthesis in air, implying that at least 60% of the excitation energy was safely dissipated in ways other than carbon assimilation. A mechanism for safe dissipation has been suggested recently by Krause and Behrend (1986), Krause and Laesch (1987a,b) and Weis et. al (1987). It is related to qE quenching (Section 8) and is believed to be associated with acidification of the intrathylakoid space as a result of light-driven proton transport. Related changes in the ultrastructure of the membranes are are thought to lead to an increase in the rate of thermal dissipation. Björkman and Demmig (1987) have drawn attention to the possibility that the violaxanthin-zeaxanthin cycle may play a specific role in bringing about changes in the rate of thermal dissipation. For much the same reasons it would be a mistake to leap to the conclusion that photoinhibition of shade leaves in strong light is attributable to their inability to fix CO2 at high rates. Some shade leaves (see below) can support rates of photosynthesis

which are equal to their “sun grown” counterparts. Even if they photosynthesise more slowly, the limiting factor is as likely to be the rate of electron transport to NADP as the removal of electrons from NADPH2. To an extent, reoxidation of PSI by O2 will also provide an alternative safe means of energy dissipation if the mechanisms for detoxification of super-oxide radicals do not become overloaded. Finally it might be supposed that the switch from assimilatory dissipation to thermal dissipation might occur as readily (or more readily) in shade leaves. All of this, together with the role of low temperature in photoinhibition, suggests that, although inadequate CO2 assimilation may sometimes contribute to damage, the crucial factor is more likely to be the strain imposed on (already inadequate) repair of the QB-binding protein or on replacement of the damaged protein within the thylakoid membrane. If, on the other hand, the upper limit of “safe” dissipation of excitation energy is normally about 60% of the energy used in CO2 fixation, factors affecting CO2 reduction (such as the availability of cytosolic Pi) will increase in importance as external constraints (such as low temperatures) bring CO2 fixation below the 40% level in bright light.

Figure 32.3. Photoinhibition in Helleborus niger. Sun leaf taken from full Australian summer light, shade leaf from 2 µmole.quanta..m-2.s-1. Both leaf discs were assayed before and after the same “photoinhibitory” treatment (4.5 hours at 1000 µmole.quanta..m-2.s-1 and 8oC). The shade leaf now showed loss of rate at high PFDs (not seen after shorter periods of photoinhibition - c.f. Fig. 32.1) and even greater decline in quantum yield. Conversely the sun leaf photosynthesised faster. Figure 32.3 bears on these relationships. Leaves from Christmas Rose (Helleborus niger) growing on the campus of the Australian National University in Canberra, performed in a very similar way (in regard to quantum yield and maximal rates) whether they were taken from full sun or from an immediately adjacent site in deep shade (0.5%, or less, of full sun). As expected, the shade leaf was strongly photoinhibited by strong

light 1000 µmole.quanta.m-2.s-1 (at 8oC) whereas the sun leaf showed a significant increase in rate after such treatment and a decrease only after very prolonged exposure (36 hours or more) to high light and low temperature. In the early stages of photoinhibition, however, there was often no decrease in maximal photosynthetic rate of the shade leaf despite a decline in quantum yield and clearly, in these circumstances, the leaf’s capacity for energy dissipation cannot have been much inferior to that of its “sun” counterpart. Similarly, following photoinhibition and subsequent overnight exposure to very low light, the shade leaf not only showed complete recovery but an increase in rate which brought it very close to the performance of the “sun” leaf. Such results are best explained by a rate of repair which is so rapid in the sun leaf, despite chilling, that it can increase its rate of photosynthesis (see Fig. 32.3) during a treatment which is markedly deleterious to the shade leaf. The rate of repair in the shade leaf on the other hand is not sufficiently rapid to allow it to cope with the continuing damage caused by high light at low temperature but, once returned to low light and normal temperature, its capacity for CO2 fixation not only returns but increases (Fig. 32.4).

Figure 32.4. Photoinhibition (1000 µmole.quanta..m-2.s-1 at 8oC) and recovery from photoinhibition (2 µmole.quanta..m-2.s-1 at 25C) in Helleborus niger. Improvement in the rate at which shade leaves of sun species photosynthesise following long term exposure to higher light is referred to as “acclimation” (see, for example, Anderson and Osmond, 1987) and, in some species, will undoubtedly involve new synthesis of enzymes concerned in carboxylation as well as changes to the photosynthetic apparatus. In Christmas Rose as we have already noted, the difference in photosynthesis between sun and shade leaves was small and the increase in rate during the high-light illumination of the sun leaves could be reasonably placed into

the category of “long induction” (Rabinowitch 1956). This gradual increase in rate can be shortened by orthophosphate feeding (Walker and Sivak 1986) and must, in part at least, relate to the availability of cytosolic Pi. It seems likely that similar changes in rate would also be set in train in the shade leaf but that these would be masked, by photoinhibitory damage so long as strong illumination at low temperatures was continued. Such short-term responses to sun and shade are important because many plants exist in unstable and changing environments to which they must adjust. In addition, the short-term changes themselves may well be part of the complex signalling process by which a plant perceives that it is in a “permanently” changed environment and should, if it can, proceed to long-term adjustment. Such adjustment would eventually manifest itself as “acclimation”.

“measurement is rarely non-intrusive”

Although it is still possible to go to parts of S.E. Asia and find new genera, let alone a new species, taxonomy is so comprehensive that it can offer authoritative identification and descriptions of most of the plants that we normally encounter. By comparison, our knowledge of the physiology of plants is virtually non-existent. In many respects the physiology of the green leaf is dominated by photosynthesis but there has not yet been any serious attempt to evaluate the photosynthetic behaviour of most species. This is largely because it has been such a difficult thing to do until now. As a result of recent advances in molecular biology, our need to improve our knowledge of plant physiology has assumed new practical importance. Genetic engineering allows the possibility of transferring a desirable ability from species to species. The identification of what is desirable resides in improved evaluation of physiological performance. Procedures such as those described above, when used together with techniques such as Q-analysis (Sections 13e and h) can make a significant first contribution to such sorely needed evaluation.

REFERENCES

General References Anderson, J.M. and Osmond, C.B. (1987) Sun-shade responses: compromises between acclimation and photoinhibition. In: Topics in Photosynthesis. Vol 9, Photoinhibition (Barber, J. ed.). Elsevier, Amsterdam, New York, Oxford. pp 1-37. Cseke, C., Balogh, A., Wong, J.H., Buchanan, B.B., Stitt, M., Herzog, B. and Heldt, H.W. (1984) Fructose 2,6-bisphosphate: a regulator of carbon processing in leaves. Trends in Biochem. Sci. 9, 533-535. Evans, J.R. (1986) The dependence of quantum yield on wavelength and growth irradiance. Aust. J. Plant Physiol. 14, 69-79. Graham, D. and Chapman, (1979) Interactions between photosynthesis and respiration in higher plants. In: Encyclopedia of Plant Physiology: Photosynthesis II. Vol 6 (Gibbs, M., Latzko, E. eds) Springer Verlag, Berlin. pp. 150-162. Laisk, A. (1977) Kinetics of photosynthesis and photorespiration in C3 plants. Nauka, Moscow (In Russian). pp 1-196. Rabinowitch, E.I. (1956) Photosynthesis and related processes, vol. II, part 2. Interscience, New York. pp. 537-1432. Walker, D.A., Leegood, R.C. and Sivak, M.N. (1986) Ribulose bisphosphate carboxylase oxygenase. Phil. Trans. Roy. Soc. B. 313, 305-324. Walker, D.A. and Sivak, M.N. (1986) Photosynthesis and phosphate: a cellular affair? Trends in Biochem. Sci. 11, 176-179. Specific References Adams, W.W., Nishida, K. and Osmond, C.B. (1986) Quantum yields of CAM plants measured by photosynthetic O2 exchange. Plant Physiol. 81, 297-300. Ben, G-Y., Osmond, C.B. and Sharkey, T.D. (1987) Comparisons of photosynthetic responses of Xanthium strumarium and Helianthus annuus to chronic and acute water stress in sun and shade. Plant Physiol. 84, 476-482. Björkman, O. (1981) Responses to different quantum flux densities. In: Physiological plant ecology Vol 12a: Interactions with the physical environment (Lange, O.L., Nobel, P.S., Osmond, C.B., Ziegler, H. eds.). Springer, Heidelberg, Berlin, New York. pp 57-107. Björkman, O. and Demmig, B. (1987) Photon yield of O2 evolution and chlorophyll fluorescence characteristics at 77K among vascular plants of diverse origins. Planta 170, 489-504. Blackman, F.F. (1895a) Experimental researches on vegetable assimilation and respiration. I. On a new method for investigating the carbon acid exchanges of plants. Phil. Trans. Roy. Soc. B. 186, 485-502. Blackman, F.F. (1895b) Experimental researches on vegetable assimilation and respiration. II. On the paths of gaseous exchange between aerial leaves and the atmosphere. Phil. Trans. Roy. Soc. B. 186, 503-562. Blackman, F.F. (1905) Optima and limiting factors. Ann. Bot. 19, 281-295.

Decker, J.P. (1957) Further evidence of increased carbon dioxide production accompanying photosynthesis. J. Solar Energy Sci and Engng. 1, 30-33. Demmig, B. andBjörkman, O. (1987) Comparison of the effect of excessive light on chlorophyll fluorescence (77K) and photon yield of O2 evolution in leaves of higher plants. Planta 171, 171-184. Edwards, G.E. and Walker, D.A. (1983) C3, C4, Mechanisms, and Cellular and Environmental Regulation of Photosynthesis. Blackwell Scientific Publications Ltd., Oxford. pp. 1-542. Emerson, R. (1958) The quantum yield of photosynthesis. Ann. Rev. Plant. Physiol. 9, 1-24. Graham, D. (1980) Effects of light on dark respiration. In: Biochemistry of Plants. Vol 2. (Davies, D.D. ed). Academic Press, New York. pp. 526-580. Graham, D. and Walker, D.A. (1962) Some effects of light on the interconversion of metabolites in green leaves. Biochem. J. 82, 554-560. Heath, O.V.S. (1969) The Physiological Aspects of Photosynthesis. Heinemann Ed. Books Ltd., London. pp. 1-310. Hill, R. and Bendall, F. (1960) Function of the two cytochrome components in chloroplasts: a working hypothesis. Nature 186, 136-137. Kok, B. (1948) A critical consideration of the quantum yield of Chlorella - photosynthesis. Enzymologia 13, 1-56 Kok, B. (1949) On the interrelation of photosynthesis and respiration in green plants. Biochim. Biophys. Acta 3, 625-631. Kok, B. (1951) Photo-induced interactions of metabolism of green plant cells. Symp. Soc. Exp. Biol. 5, 209-221. Krause, G.H. and Behrend, U. (1986) ∆pH-dependent chlorophyll fluorescence quenching indicating a mechanism of protection against photoinhibition of chloroplasts. FEBS Letters 200, 298-302. Krause, G.H. and Cornic, G. (1987) CO2 and O2 interactions in photoinhibition. In: Topics in Photosynthesis. Vol 9, Photoinhibition (Barber, J. ed.). Elsevier Science Pubs BV (Biomedical Division) pp.1-37. Krause, G.H. and Laasch, H. (1987a) Photoinhibition of photosynthesis. Studies on mechanisms of damage and protection in chloroplasts. In: Progress in Photosynthesis Research (Biggins, J. ed) Vol 4. Martinus Nijhoff, Dordrecht. pp 19-26. Krause, G.H. and Laasch, H. (1987b) Energy-dependent chlorophyll fluorescence quenching in chloroplasts. Correlation with quantum yield of photosynthesis. Z. Naturforsch.C 42(5), 581-584. Osmond, C.B. (1981) Photorespiration and photoinhibition: some implications for the energetics of photosynthesis. Biochim. Biophys. Acta 639, 77-98. Osmond, C.B. (1987) Photosynthesis and carbon economy of plants. New Phytol (Suppl) 102, 161-175. Osmond, C.B. and Björkman, O. (1972) Effects of CO2 on photosynthesis. Simultaneous measurements of oxygen effects on net photosynthesis and glycolate metabolism in C3 and C4 species of Atriplex. Carnegie Inst. Washington Yearb. 71, 141-148. Powles, S.B. (1984) Photoinhibition of photosynthesis induced by visible light. Ann. Rev. Plant Physiol. 35, 15-44. Powles, S.B., Cornic, G. and Louason, G. (1984) Photoinhibition in vivo photosynthesis induced by strong light in the absence of CO2: an appraisal of the hypothesis that photorespiration protects against photoinhibition. Physiol. Veg. 22, 437-446. Sharp, R.E., Matthews, M.A. and Boyer, J.S. (1984) Kok effect and the quantum yield of photosynthesis. Plant Physiol. 75, 95-101. Sivak, M.N. (1987) The effect of oxygen on photosynthetic carbon assimilation and quenching of chlorophyll fluorescence emission in vivo: phenomenology and hypotheses concerning the mechanism involved. Photobiochem. Photobiophys. Suppl, 141-156. Sivak, M.N. and Walker, D.A. (1985) Chlorophyll a fluorescence: can it shed light on fundamental questions in photosynthetic carbon dioxide fixation? Plant Cell and Environment 8, 439-448. Stitt, M., Huber, S.C. and Kerr, M. (1987) Control of photosynthetic sucrose formation. In: The Biochemistry of Plants. Vol 13. (Hatch, M.D., Boardman, N.K., eds). Academic Press, New York. pp 327-409.

Terashima, I. and Saeki, T. (1985) A new model for leaf photosynthesis incorporating the gradients of light environment and of photosynthetic properties of chloroplasts within a leaf. Ann. Bot. 56, 489-499. Terashima, I., Sakaguchi, S. and Hara, N. (1986) Intra-leaf and intracellular gradients in chloroplast ultrastructure of dorsiventral leaves illuminated from the adaxial or abaxial side during their development. Plant Cell Physiol. 27(6), 1023-1031. Terashima, I. and Takenaka, A (1986) Organisation of photosynthetic system of dorsiventral leaves as adapted to the irradiation from the adaxial side. In: Biological control of Photosynthesis. (Marcelle, R., Clijsters, H., Van Poucke, F., eds). Martinus Nijhoff Pubs., Dordrecht. pp 219-299 Tolbert, N.E. (1971) Microbodies, peroxisomes and Glyoxysomes. Ann. Rev. Plant Physiol. 22, 45-76. Van der Veen, R. (1949) Induction phenomena in photosynthesis. I. Physiologia Plant. 2, 217-234. Walker, D.A. (1976) CO2 fixation by intact chloroplasts: photosynthetic induction and its relation to transport phenomena and control mechanisms. In: The Intact Chloroplast, Chapter 7 (Barber, J. ed). Elsevier, Amsterdam. pp 235-278. Walker, D.A. (1988) Automated measurement of rate v PFD. In: “New Vistas in Photosynthesis”. Proc. Phil. Trans. Roy. Soc. B. Meeting, London May 1988. **now avail Walker, D.A. and Osmond, C.B. (1986) Measurement of photosynthesis in vivo with a leaf disc electrode: correlations between light dependence of steady-state photosynthetic O2 evolution and chlorophyll a fluorescence transients. Proc. R. Soc. Lond. B 227, 267-280. Walker, D.A., Osmond, C.B. and Cleland, R.E. (1988) Concerning photoinhibition of shade leaves of Helleborus niger. AFRC meeting Imperial College, London. paper no. 23. Warburg, O. and Burke, D. (1950) The maximum efficiency of photosynthesis. Arch. Biochem. Biophys. 25, 410-442. Weis, E., Ball, J.T. and Berry, J. (1987) Photosynthetic control of electron transport in leaves of Phaseolus vulgaris evidence for regulation of photosystem 2 by the proton gradient. In: Progress in Photosynthesis Research, Vol 2 (Biggins, J. ed) Martinus Nijhoff, Dordrecht. pp 553-556.

PART F

AQUEOUS PHASE MEASUREMENTS

33. INTRODUCTION Although Sections A-E inclusive have been concerned with measurements of O2 in the gas-phase, polarographic measurement of O2 is more usually undertaken in the aqueous phase and is currently such an every-day technique that familiarity will largely obviate the need for explanation. For completeness, however, some details and descriptions will be given.

34. PRINCIPLE The principle is the same as before (Section 2) except that the rate of diffusion of O2 through water is slower (by 3 orders of magnitude) than it is through air and this normally necessitates stirring of the solution, or suspension, if rapidity of response is required (as it will be in any study of kinetics). Stirring is usually achieved by the use of a revolving magnet (below) and a magnetic follower or “flea” within the vessel containing the reaction mixture. (If a single microcathode detector is used the rate of consumption of O2 at the electrode surface may be so small that stirring may not be necessary).

35. THE “FLEA” Most reaction mixtures are small in volume (1 to 3 ml) and the magnetic followers which are employed are accordingly small in size and easily lost. Like real fleas, they can be immobilised by pressing them into a bar of wet soap but are more normally given safe keeping on a bar magnet. They can be readily made out of glass capillary tubing and thin iron wire, although an element of skill is required and a good flea is therefore a thing to be cherished. A “good” flea is one which is completely sealed but without conspicuously heavy glass beads at either end. It is precisely the right length (i.e., a shade shorter than the diameter of the reaction vessel) and encloses (for buoyancy) about three to four times as much air as iron. It must be pleasing to the eye on the basis of these criteria but it must also live up to promise by spinning freely and evenly in the reaction mixture. Uneven stirring can generate electrical “noise” on pen-recorder traces. Some electrode circuits incorporate electronic filters to limit this effect and a small capacitor (say 10 F) is often put across the recorder terminals for the same purpose. However, unless extra sensitivity is required, (greater amplification can obviously contribute to greater “noise”) time

“a good flea is a thing to be cherished”

invested in procuring a good flea and a smoothly revolving magnet (avoid sparking electric motors) is often rewarded by essentially “noise-free” traces.

36. CALIBRATION. 36(a) Setting an Arbitrary “Air-line”

effect of stopping stirrer

If a pen-recorder with a 10 inch (25 cm) chart is used, calibration normally starts by establishing electrical zero and then adjusting the electrical output of the O2 sensor until, in distilled water, it draws a line across the chart about three quarters way from zero (Fig. 36.1). Clearly, the precise electrical value will depend upon the electrode and its circuitry. With a standard Hansatech electrode and CB1 control box with the “output” switch set to x1, a 1v full-scale setting would be appropriate. If the stirrer is stopped, the trace should immediately fall towards zero but return, equally readily, to its starting point once stirring is resumed. (This is a consequence of limiting diffusion of oxygen in unstirred solution - Section 35, above). 36(b) Establishing the N2 Line If a few crystals of dithionite are dropped from a spatula into the stirred water the trace should fall, rapidly at first and then more slowly, to near zero (Fig. 36.1). This is because O2 is consumed according to Eqn. 36.1: NA2S2 O4 + O2 + H2O

NaHSO4 + NaHSO3 ...Eqn. 36.1

Alternatively, (and preferably) the empty chamber can be flushed with N2 gas (at the operating temperature). Both procedures establish the so-called “N2 line” (i.e. the line that the pen traces in a N2 atmosphere - Delieu and Walker 1972). 36(c) Residual Current If the electrode were perfect the N2 line would coincide with the electrical zero of the recorder but there is always a small “residual current” (Section 36c, and Fig. 36.1), which should not exceed 10%, at most, of the “air-line” (see 36d below). Some residual current can be diminished by careful cleaning and by periodically storing electrodes in a dessicator. (Many electrodes are embedded in epoxy resins and therefore tend to take up a certain amount of water. This can usually be eliminated by dry storage. For this reason it is good practice to store a spare electrode in silica gel and to alternate this, on a weekly basis, with one that has been in use).

Response of electrode to N2, O2 and air

A large residual current is an indication of malfunction and, if it cannot be corrected by cleaning or desiccation, the electrode will have to be discarded or returned to the manufacturer for repair. (Some manufacturers will replace or refurbish damaged electrodes for less than the price of a new electrode). Malfunction is also indicated by a ring of crystals forming around the junction between an electrode and the epoxy resin in which it is sealed, reflecting the difficulties involved in sealing one material (platinum) to another (epoxy resin) over a range

of temperatures. Electrolyte (which usually contains KC1) penetrates this junction and subsequently crystallises. The crystals fracture the seal and eventually give rise to an unacceptably high residual current. Microcathode electrodes sealed in glass are less vulnerable to damage of this sort than those with cathodes sealed in epoxy resins but are likely to be more expensive (sometimes very much more expensive) than those based on resins. 36(d) Establishing a New “Air-line” Once the “residual current line” or “nitrogen line” has been established either by adding dithionite or, ideally, by flushing the empty reaction chamber with nitrogen, calibration can proceed (Fig.36.1). If dithionite has been used, the vessel and the electrode should be washed repeatedly (about 10 times) with distilled water. This is most easily done with a wash-bottle and an aspirator which is used to remove the added water as soon as the vessel is full. The aspirator tube should be fitted with a narrow flexible tip so that the membrane of electrodes inserted into the base of the chamber is not damaged and the “flea” is not swallowed. Finally, distilled water which has been shaken with air (at the operating temperature) is added. This establishes the “air-line”. The air-line can be set electrically to any arbitrary value (Delieu and Walker, 1981) but there is merit in using a convenient and constant value. For example, if 2 ml is the preferred volume and 20oC the preferred temperature, it makes life easier if the difference between the N2 line and the air-line is set at 5.6 divisions of the chart recorder paper. This is because air-saturated water at 20oC contains approximately 0.28 µmoles of O2 per ml (Table 36.1) and each large division of the chart will then be equivalent to 0.1 µmole of O2. Of course, if recording is started without displacing the air-line, the O2 trace will soon reach the top of the chart. So it also helps (if O2 evolution is to be recorded) to displace the air-line (following calibration) to the foot of the chart using the “back-off” control on the O2 electrode “box”. At one time, a 1.0 mV recorder was almost mandatory but, over the years, advances in signal amplification have made it possible to record at larger voltages and also to use a computer rather than a pen-recorder.

try to ensure that “the flea is not swallowed”

Table 36.1. The Oxygen Content of Air Saturated Water Temp.(oC)

O2(ppm)

O2(µmole/ml)

0 5 10 15 20 25 30 35

14.16 12.37 10.92 9.76 8.84 8.11 7.52 7.02

0.442 0.386 0.341 0.305 0.276 0.253 0.230 0.219

The values in column 2 may also be derived from the empirical formula of Truesdale and Downing (1954) which states that Cs = 14.16-0.3943T + 0.0077142T2 - 0.0000646T3 where Cs = saturation concentration (ppm) and T = temperature (C)

“it makes life easier”

It should be noted that while air-saturated water is a very convenient standard for calibration it only serves as a first approximation and, although it is ideal for comparative work, absolute calibration necessitates a different approach. This is because other solutes affect the solubility and hence the “activity” (or concentration of O2) in media (Section 36g) which are commonly employed in work with isolated organelles such as chloroplasts. One method of absolute calibration is to generate O2 in the medium by the action of catalase on pre-calibrated hydrogen peroxide (see Delieu and Walker, 1972 and Figure 36.1). 36(e) Normal Calibration Procedure Summarised The above procedure (Section 36a-d) is summarised in Fig. 36.1.

Figure 36.1. Calibration Summarised. (i) With water in the vessel set an arbitrary “air-line” by adjusting the electrical output of the electrode until the recorder pen is near the top of the chart. (ii) Add dithionite crystals (or flush empty vessel with N2 gas) to establish the “N2 line”. (iii) If dithionite has been used, wash the chamber several times with distilled water. (iv) Add water, saturated with air at the operating temperature, to obtain a true “air-line”. (v) Adjust this true air-line electrically so that it is a convenient multiple of the number of moles of O2 per ml of water (e.g. at 20C there are 2.8 µmoles/ml therefore if a reading of 5.6 is selected as the calibrated “air-line” and a reaction mixture of 2 ml is used, each division of the chart will be equivalent to 0.1 µmoles of O2).

36(f) Absolute Calibration Additional checks on calibration and linearity of response may be carried out as follows: (i) A small quantity of catalase is added to a reaction mixture which is allowed to equilibrate in the closed vessel. (ii) Hydrogen peroxide is standardized by adding it to an acidic solution of potassium iodide and titrating the iodine released with sodium thiosulphate (freshly prepared, e.g., from a newly opened BDH analytical ampoule or standardized, in turn, against an iodine solution). Sodium thiosulphate (Na2S2O3) is a reducing agent by virtue of the following reaction (Eqn. 36.2) 2S2O3 = → S4O6 = + 2e-

......Eqn.36.2

It will therefore reduce iodine to iodide: I2 + 2e- → 2I-

......Eqn.36.3

so that 2Na2S2O3 + I2 → Na2S4O6 + 2NaI

......Eqn.36.4

and, since hydrogen peroxide reacts with iodide in acid solution as follows: 2I- + H2O2 + 2H+ → I2 + 2H2O

......Eqn.36.5

and because H2O2 also yields O2, by catalatic dismutation, according to equation 36.6: 2H2O2 → 2H2O + O2

.......Eqn.36.6

the relationships between thiosulphate, hydrogen peroxide and oxygen evolved are 1 ml of 0.01 N Na2S2O3 ≡ 5µmoles H2O2 ≡ 2.5µmoles O2. The titration procedure (Vogel, 1961) is to add 0.1-2 ml of hydrogen peroxide solution to a mixture containing 0.5 ml of 10% KI and 2 ml of H2SO4 (in a total of 4.5 ml) to which one drop of 1% ammonium molybdate solution is added (as a catalyst) and then to titrate after allowing 3 minutes for the release of iodine to be completed. Thiosulphate is added until the solution is a pale straw colour, at which point two or three drops of starch glycollate are also added. As the titration is continued a green colour develops which is followed by the appearance of the characteristic blue, of the iodine-amylose complex immediately prior to the colourless end point. Titration of the H2O2 used in the experiment illustrated in Fig. 36.2 gave a value of 0.1345 µmoles for 0.2 ml of H2O2 calculated from a mean titration value of 5.38 ml of thiosulphate. When 2 µl aliquots were subsequently added to 2 ml of deionized water at 20C in the electrode vessel, a mean recorded value of 0.1380 µmoles was obtained. Fresh, “10-volume” commercial H2O2 is approximately molar, so that 1 µl of such a solution would yield approximately 0.5 µmoles of oxygen by catalatic dismutation (Equation 36.6).

(iii) Small aliquots of standardized H2O2 are added (in sequence) to the vessel, using a microsyringe. The recorder registers the O2 evolution following each addition and a series of uniform steps are obtained (Fig. 36.2) indicating both the extent and the linearity of the response. If this method is used with catalase in distilled water at 20oC, the oxygen recorded corresponds very closely to that obtained by using the value of 0.28 µmoles in setting the air-line. With other aqueous solutions the recorded values will depend on the effect of the solutes on the activity of oxygen within the solution at a given concentration. For example, Fig. 36.2 illustrates the traces obtained using H2O2 and catalase in twin vessels containing either 2 ml of deionized water or 2 ml of resuspending medium (Section 37g). It will be seen that each electrode gave virtually the same response in each case but that the response in water was about 11% smaller than that in the assay medium.

Figure 36.2. Serial additions of H2O2 (0.25 µmoles in 2µl) to duplicate 2 ml reaction mixtures containing water and catalase (left) and resuspending medium and catalase (right). The “real” values obtained with water were, on average, 11% lower than the “apparent” values obtained with assay medium (after Delieu and Walker, 1972). For some purposes, absolute calibration is inescapable but it should be noted that the catalase method is tedious and exacting. Not only is it necessary to determine, with great precision, the concentration of H2O2 in the solution added, but it is also important to prevent bubble formation during dismutation and to add the peroxide solution (inevitably containing excess dissolved oxygen) in very small but very accurate aliquots. For most purposes, therefore, relative calibration (Section 36e) is to be preferred in conjunction with a correction factor which can be derived (as in Fig.36.2) for any given solution and temperature. If isolated chloroplasts are at hand, an additional check can be applied by using a ferricyanide solution, freshly prepared from clean crystals and protected from strong white light. With such a solution,

and washed chloroplasts (free of endogenous electron donors such as ascorbate) which do not exhibit appreciable dark or post-illumination O2 uptake, 1 µmole of O2 is evolved for every 4 µmoles of ferricyanide added (Section 41a). Except under well-defined conditions, however, the possible errors are substantial (see e.g. Whitehouse et. al., 1971) and the effort involved is no less considerable than the peroxide-catalase calibration. 36(g) Using Constant Chlorophyll If a prolonged series of experiments is envisaged, there is also merit in working at a constant and pre-defined chlorophyll concentration. Chlorophyll is readily determined by adding 50 µl of chloroplast suspension to 20 ml of 80% acetone (16 ml of acetone plus 4 ml of H2O) and quickly filtering or centrifuging (avoid evaporation or exposure to bright light). The optical absorbance (1cm light-path) of the chlorophyll solution at precisely 652 nm (Bruinsma, 1961) is then multiplied by 100/9 to give µg chlorophyll per ml of original suspension (check the calibration of the spectrophotometer as necessary). Alternatively, the reciprocal of the absorbance x 9 approximates to the number of µl of chloroplast suspension required to give 100 µg of chlorophyll (Walker, 1980). The amount of chlorophyll to be used will, of course, vary with the nature of the experiment but 100 µg/ml is about the upper limit for light saturation in a conventional O2 electrode vessel. 36(h) Calculation of Rate An advantage of using constant chlorophyll in a series of experiments is that it eliminates a variable and that it also facilitates both comparisons (within and between experiments) and the determination of rates. Table 36.2. Rate for a mixture containing 0.1 mg chlorophyll. Angle 68 66 64 54 51.5 50

1cm = 1min 150 135 123 82 75 71

1cm=0.05 µmoles 300 270 246 164 150 142

Thus the angle of the trace (of change in O2 against time) can be measured (using a protractor) and multiplied by an appropriate factor. For example in Fig. 36.3 the angle θ is 680 and tan θ = 2.5. Thus if time and ∆ O2 have been recorded on the same scales as in Fig. 36.3 (in which the vertical scale is 0.1 of the horizontal scale), the value 2.5 will be a measure of the number of µmoles of O2 evolved in 10 minutes. This, in terms of O2 per hour could become 15 (2.5 x 6) and on a 1 mg of chlorophyll basis (since there was only 0.1 mg of chlorophyll in the reaction mixture) would become 150 (15/0.1). Had the same trace been recorded at 2 cm/min, the angle θ would have

been 51.5, the tangent 1.25 and this value would be a measure of the O2 evolved in 5 rather than 10 minutes so that the final rate (1.25 x 12. x 0.1) would remain the same (Table 36.2.)

Figure 36.3. Calculation of rate. In this figure the vertical scale has been electrically set to display the change () in oxygen in one tenths of moles according to the procedures described in Section 36. The horizontal scale has been selected on the chart recorder to display time in minutes and the reaction mixture (a suspension of intact chloroplasts engaged in CO2-dependent O2 evolution and displaying a characteristic lag) contain 100 µg of chlorophyll. Traditionally, rates have been expressed as moles O2 per mg chlorophyll per hour and the rate here is readily seen to be 150 µmoles oxygen mg-1 Chl.hr-1 from the fact that 0.5µmole of O2 was evolved in 2 min by chloroplasts containing 100 µg chlorophyll (i.e. 0.5 x 60). Alternatively, tables can be compiled 0.1 x 2 (see Table 36.2) based on the tangent of the angle. These save on arithmetic and, once correctly compiled, eliminate errors in subsequent calculations. Table 36.2 is part of a larger table (for all angles) based on the relationship tan θ x n = rate of O2 evolution expressed in µmoles mg-1 Chl. hr-1 where n = 60 x Chl (mg) time x oxygen Thus, if chart speed (in minutes per cm) is 1, oxygen (in µmoles per cm) is 0.1 and the reaction mixture contains 0.1 mg chlorophyll the rate for an angle of 68 (tan θ = 1.25) would be 2.5 x 60 x 0.1 = 150 1x 0.1

Pocket calculators make tables of this sort largely superfluous but, once prepared and, particularly for work at constant chlorophyll, they can remove the tedium and minimise the errors of computation.

37. MECHANICAL ISOLATION OF CHLOROPLASTS 37(a) Conventional Wisdoms For the last two decades, mechanical separation of fully competent chloroplasts (i.e. those capable of achieving light-dependent CO2 fixation of the same order of magnitude as the parent tissue) has, in general, only been achieved with spinach and with peas. Even then, this has meant using “good” spinach (Section 38) or “good” peas. It is not possible “to make a silk purse out of a sow’s ear”. Equally it is not possible to isolate chloroplasts, which will display high rates of photosynthesis, from spinach containing large quantities of starch, or calcium oxalate, or from leaves, which are anything other than healthy and actively growing. For many purposes, therefore, there has really been no alternative to “good” spinach. More recently, very respectable chloroplasts have been isolated mechanically from a limited number of additional species and chloroplasts have been enzymically separated (Section 42) from poor spinach and from notoriously difficult species (like sunflower) which have not yet proved susceptible to the mechanical approach.

“a respectable chloroplast”

Mechanical isolation involves disrupting the leaf but not the chloroplast (Section 37b, below and Fig. 37.1) in a grinding medium. An appropriate medium (Robinson, Cerovic and Walker, 1986) always contains an osmoticum (usually 1/3 M sugar or sugar alcohol) to prevent swelling or shrinking. It may also contain a buffer to maintain pH, an anti-oxidant (such as ascorbate) and a variety of other additives (some included for traditional or ritualistic reasons or on the principle of “why change a winning team?”). Following maceration of the leaf, the chloroplasts are separated as quickly and gently as possible by filtration and centrifugation and then resuspended in another medium containing the same, alternative, or additional goodies (Fig. 37.1). 37(b) Grinding Chloroplasts can be prepared by pounding leaves in a pestle and mortar and this has much the same therapeutic value as kneading dough but it is hard work and difficult to do in a reproducible manner. In an increasingly sophisticated world it also smacks of alchemy and is usually despised. If you like old things, however, don’t forget to chill the pestle and mortar, use a small quantity of grinding medium and pound very vigorously for a minute or two, preferably on the floor. All manner of motor-driven homogenisers offer a greater degree of ease and control. Details of procedure vary with the blender and, at the outset, there is no alternative to trial and error. Clearly, there should be sufficient medium to cover the leaves. The medium is best frozen to a slush (the consistency of melting snow) because the latent heat of melting then increases its cooling capacity by an order of magnitude. Similarly, the mechanical resistance of the slush tends to hold the leaves against the blades during the first moments of blending.

“it smacks of alchemy”

If you have a choice, use a blender which will mash your leaves into a coarse “brei” within 10 sec. A really good homogeniser like a Polytron (Kinematica GmbH, CH-6000 Kriens, Luzern, Switzerland) will do this in under 5 seconds. Continued homogenisation will increase the yield but also increase the damage to the chloroplasts. A bad blender or homogeniser is one which causes a lot of frothing. Cavitation effects seem to damage fragile chloroplast envelopes, possibly by smearing nasties (from the vacuole and cell wall) on active surfaces (Leegood et. al., 1981; Robinson et. al., 1979; Walker 1971, 1980). It often helps to stack the leaves and slice them with a sharp knife prior to blending but don’t be tempted, by admiration of skilled chefs, to use a chopping action. This causes detrimental bruising of the leaf tissues. Spinach responds well to 30 minutes pre-illumination in cold water. For some reason, never yet established, this actually increases yield as well as activity. Spinach will also store beforehand and good chloroplasts have been prepared from spinach stored in the dark at 5oC, in a crate, for as long as a month. Conversely, pea leaves often deteriorate very rapidly as soon as they are harvested and it therefore makes sense, if working with peas, to emulate the North American practice of cutting the corn cob only when the water in the pot is already boiling. Once grinding is at an end, the chloroplasts should be separated from the brei in the shortest possible time. 37(c) Squeezing and Filtering Have a beaker (400 ml is a good size) and a couple of thicknesses of cheese-cloth or muslin chilled and waiting. “Hairy” cloth is better than nylon, mira cloth etc. Spread it across the top of the beaker and decant the contents of the blender into it. Gather up the corners of the cloth and twist and squeeze with the fingers so that the juice is expressed from the brei. Filtration through additional layers of muslin or a sandwich of muslin and cotton wool is useful and the juice may be squeezed directly on to such a “sandwich” stretched across a second beaker. Ideally this filter sandwich should be rinsed, immediately prior to use, with a little cold grinding medium and this time there should be no squeezing so that the chloroplast suspension will simply drain through under gravity. As quickly as possible the filtered suspension should be decanted into chilled centrifuge tubes. With practice, these can be adequately “balanced” by eye but use a balance to equalise them prior to centrifugation if you are in any doubt about your ability to do this successfully. 37(d) Centrifugation Like grinding, this is often a matter of trial and error until you know, by experience what your centrifuge will accomplish. Ideally, at this stage of the preparation (Fig. 37.1), you should have about 160 ml of suspension equally distributed between four 50 ml centrifuge tubes. These should be put into a swing-out centrifuge with sufficient acceleration and deceleration to go from rest to about 5000 rpm (and back to rest) in about 90 seconds. Alternatively, you may prefer to accelerate to about 1500 rpm and maintain this speed for 1 min before deceleration. Much longer centrifugation may be imposed upon you by the characteristics of your centrifuge but, if a short spin will give you a sufficiently large pellet for your needs, do not even contemplate prolonging spinning for longer than absolutely necessary.

Figure 37.1. Diagramatic representation of mechanical preparation of isolated chloroplasts. Starting top left: 1. The leaves are pre-illuminated whils floating on cold water (for half an hour or so). 2. They are then shaken dry and sliced, not chopped, on a board using a sharp knife. 3. They are then blended in a domestic blender (or preferably a polytron) and the brei squeezed through muslin into a chilled beaker. 4. The suspension is filtered through a snadwich of cotton wool in muslin. Following filtration the suspension is decanted into centrifuge tubes and centrifuged (preferably in the cold in a swing-out centrifuge). 5. Finally, the supernatant is decanted and the chloroplast pellet is ready for resuspension.

This will increase the yield only at the expense of quality and, unless you can contrive to use a great many chloroplasts in a short time, you will probably discard most of those that you have prepared anyway. If you have access to a refrigerated centrifuge you should use it at (or just below) zero degrees but since the entire centrifugation in a one-stage conventional chloroplast preparation can take as little as 90 seconds it possible to avoid undue warming, even in hot climates, by pre-chilling tubes and holders to be used in an unrefrigerated bench centrifuge. A swing-out centrifuge with new, or carefully treated centrifuge tubes, is better than an angle-head with old tubes. This is because the sedimenting chloroplasts migrate to the outer wall of the tube in an angle-head and, particularly if the wall has become roughened by harsh cleaning, the effect can be rather like pulling a balloon across a concrete floor with a large weight on it. You should avoid washing centrifuge tubes with detergent and indeed it is best, if you can afford it, to set aside a set of tubes and glassware which is never exposed to detergent but simply rinsed in distilled water after use. Depending on the centrifuge, some advantage may also derive from shortening the sedimentation path - i.e. by filling the tubes to one half, or less, of their capacity. If you are likely to be using chloroplasts routinely and wish to secure the best preparation possible, time invested in optimising centrifugation would be well spent because it is one of the most critical parts of the whole procedure. resuspending using cotton wool and a glass rod

37(e) Resuspension Chloroplast pellets are sticky, so that, at the end of centrifugation, the supernatant can be decanted by pouring confidently into a beaker without fear of undue loss. The inside of the tube can then be wiped with a tissue and the chloroplasts are ready for resuspension. For some purposes, the chloroplasts may need to be washed but most procedures are more or less the same until this stage. If it is a one-stage (no second centrifugation) preparation you may wish to rinse the surface of the pellet with a little chilled resuspending medium poured carefully down the side of the tube and then decanted. After that, resuspension is a matter of choice. Ulrich Heber, still probably the unofficial world record holder for “best” chloroplasts, prefers to add a millilitre or two of medium to each tube and, standing in a cold room, agitating the contents with a trembling motion of his hands. Others prefer to dislodge the pellet with a small soft paint-brush or by sucking the suspension in and out of a pipette. Personally, I prefer the “Bob Whatley” method, which is to use a small piece of cotton-wool soaked in chilled medium and pushed to and fro with a glass rod. This requires a modicum of skill but it is gentle and thorough. Ruptured chloroplasts also tend to adhere selectively to the cotton wool. Resuspension for any purpose is best achieved by using small quantities (2 or 3 ml per tube) of resuspending medium (Section 37g) until the pellet is homogenous. Otherwise you are likely to find yourself pursuing pieces of dislodged pellet through a larger volume of medium with little hope of uniform resuspension. Following this step, the volume can be adjusted to whatever is desirable. Relatively large quantities (say 40 ml/tube) of medium will be preferred if you wish to wash the chloroplasts and recover them in a second spin, whereas

relatively small quantities are best if you wish to use your chloroplasts as only one constituent in a conventional reaction mixture containing 50-100 µg chlorophyll (Section 41). You may also find some advantage in storing your chloroplasts as a pellet until immediately prior to use (Jensen and Bassham, 1965) - a procedure which may diminish loss of essential ions, such as K+, to the medium. 37(f) Grinding Medium Chloroplasts contain three main compartments, each surrounded by a membrane. The thylakoid membrane houses the chlorophyll, other pigments and electron carriers. Protons driven, by electron transport, into the compartment bounded by the thylakoid membrane, discharge through the ATPase (Section 41 and Fig. 41.3) into the stroma. The stroma contains the enzymes involved in carbon assimilation and the stroma is contained, in turn, by a double envelope. Movement of inorganic ions and metabolites into the space between the two envelopes appears to occur very freely but the inner envelope incorporates the phosphate translocator which controls the principal movements of metabolites into and out of the chloroplast. This basic structure, which has only become clear in the last 2-3 decades, has many implications for chloroplast isolation and the nature of the grinding medium (see e.g. Edwards and Walker, 1983). Firstly, the chloroplast is a fairly fragile organelle. If it contains starch, centrifugation can be a bit like swinging a paper bag full of potatoes round your head - there is a real risk of the bag bursting. For this reason, it is easier to isolate intact chloroplasts from peas or spinach early in the day before diurnal starch accumulation is well advanced. Otherwise the envelopes are reasonably resistant to mechanical shock but very susceptible to osmotic damage. This is because, like any semi-permeable membrane, the chloroplast envelopes will permit entry of water from a hypotonic solution (i.e. one with a lower osmotic pressure as a consequence of containing fewer solute molecules than the stroma). As already noted (Section 37a), it is necessary to employ an osmoticum in sufficient concentration to make the grinding medium and the chloroplast isotonic (i.e. of equal osmotic pressure) so that there is neither swelling nor shrinking of the chloroplasts during separation. For many years, chloroplasts were isolated by Arnon and Whatley in “tris-NaCl” (i.e. in a medium containing trishydroxymethylaminomethane, as the buffer, and NaCl as the osmoticum). There were sound reasons for doing this. Arnon and Whatley preferred an inorganic osmoticum to a sugar in order to eliminate the possibility of substrate-driven ATP formation in their pioneering studies on photophosphorylation. “Tris” was simply one of a very limited range of relatively inert buffers readily available to the plant biochemist in the 1950’s. For a variety of reasons, these chloroplasts (which played a major role in furthering our understanding of photosynthesis), were very imperfect when it came to CO2 fixation. Increased rates followed a return to Hill’s practice of using one third molar sugar as the osmoticum and to other procedures adopted at that time (Walker, 1964). Sucrose will serve very well as an osmoticum and considering the fact that domestic sugar is both cheap and pure it is surprising that it is not more widely used to make up large quantities of medium which frequently go down the sink within a very short time of being prepared. Originally (Walker, 1964), sorbitol was preferred to sucrose, glucose and fructose for much the same reason as Arnon and Whatley used

salt - i.e., because it was less likely to be metabolised. By now, the use of sorbitol has become time-honoured (and the author wishes that he had negotiated a percentage with the manufacturers) but other sugar alcohols (e.g. manitol) or sugars (e.g. glucose) are equally effective and sometimes to be preferred (sorbitol, for example, will interfere with some molybdate assays for inorganic phosphate and is clearly inappropriate in such circumstances). Ficoll (a sucrose polymer) which gives a relatively viscous solution at an appropriate osmolarity, has also been used as an osmoticum and seems to affect the shape of the isolated chloroplast. It may be used to advantage in spectroscopic studies in which stirring is precluded and sedimentation under gravity needs to be minimised. Choice of pH and buffer has also varied over the years. It is possible to prepare active chloroplasts from spinach in sorbitol alone, and for many purposes almost any pH between 5.5 and 8.5 will suffice. Inorganic phosphate was used in the first preparations of active chloroplasts from peas (Walker, 1964) but was superseded by “tricine” (N-Tris(hydroxymethyl)methylglycine) following Norman Good’s (1966) outstanding gift to the scientific community of a new range of “zwitterion” buffers such as “Hepes” (N-2-Hydroxyethyl-piperazine- N’-2-ethanesulphonic acid). In turn, these have been mostly restricted to use in resuspension and assay because of their cost, whereas inorganic pyrophosphate, which also serves as a chelating agent, has been used extensively. An anti-oxidant is often included and ascorbate is the usual choice. Isoascorbate once also threatened to become mandatory but has no special virtue and was originally preferred to the isomer only because it was freely available in the U.K. in the 1960’s as the sodium salt. Originally both Mg++ and Mn++ were believed to be advantageous and were added in the presence of EDTA to prevent precipitation by phosphate. Surprisingly, their omission can be detrimental although it is difficult to judge why. Protective agents such as Bovine Serum Albumin may also be added. BSA affords some protection against phenolics but may also affect the charge distribution on the envelope in a beneficial way. Relatively inactive chloroplasts from “difficult” species like sunflower can function very well in a “reconstituted system” (Section 44) suggesting that the “difficulty” lies in adsorption of deleterious materials on the envelope whereas the internal stroma and thylakoids remain undamaged (Delaney and Walker, 1976). A traditional grinding medium (Cockburn et. al., 1968) would therefore look something like this: Sorbitol (330 M) Na ascorbate (2 mM) Na pyrophosphate (10 mM) at pH 6.5 (adjusted with HCl) MgCl2 (5 mM) but it should again be emphasised that only the osmoticum is really vital and none of the quantities are really critical. In short, there is no need to waste time ensuring that the sorbitol is 0.33 rather than 0.32 or adjusting the pH to within 0.1 of a unit.

Whether or not this medium is still the most useful for general purposes is quite another matter. One which can be recommended is that of Cerovic and Plesnicar (1984) which contains Sorbitol (340 mM) KCl (0.4 mM) EDTA (0.04 mM) Hepes-KOH (2.0 mM) at pH 7.8 but it should be noted that in order to secure best advantage from this “low ion” and other media (Nakatani and Barber, 1977) it should be employed throughout (i.e., as both grinding and washing medium). Another extremely interesting innovation is that of Robinson (1986) who has used 200 mM KCl as the osmoticum rather than a sugar or sugar alcohol. This is arguably nearer to the in vivo osmoticum and helps to maintain the high K content which appears to be necessary for optimal activity (Kaiser et.al., 1980). 37(g) Resuspending and Assay Medium One suitable medium (RS1) contains Sorbitol (330 mM) EDTA (2 mM) MgCl2 (1 mM) MnCl2 (1 mM) Hepes-KOH (50 mM) at pH 7.6 BSA (0.2%) Again, this is a traditional mixture and it is difficult to say what benefit derives from the inclusion of the MgCl2, the MnCl2 and the chelating agent (EDTA) which they, in part, make necessary (but see Edwards et. al., 1978). An osmoticum is essential and it is convenient to store the chloroplasts at or about the pH that will be used in the reaction mixtures (see Section 41). Tradition, however, is hard to escape and, if you are preparing chloroplasts for the first time, you would be advised to use a well established procedure initially. Once you know that you can prepare chloroplasts from “good” spinach (Sections 37a and 38) which will support CO2-dependent O2 evolution (at 20oC) at rates of about 100 µmoles-1 O2 mg Chl-1 hr-1 (Section 36f and Fig. 36.3) you will know that you have mastered the simple art of preparation and are licensed to take as many liberties as you wish with the system. The rate of CO2-dependent O2 evolution is really the only criterion for full photosynthetic competence (the category “A” chloroplasts of Hall, 1972) and for any serious work a rate of about 100 is almost mandatory and a rate of 200, or more, most desirable. Apart from anything else, isolated chloroplasts do deteriorate both during use and during storage and it is possible to do so much more, so quickly, with “good” chloroplasts rather than “bad”. Similarly, if you come up with some totally new observation you will feel much more confident that it will be “real” rather than artefactual, if it is obtained with active chloroplasts. Many different resuspending media are used for specific purposes. As already noted (in 37f above) chloroplasts prepared in “low ion” grinding medium are resuspended in low ion grinding medium. In Robinson’s (1986) procedure 200 mM KCl may also be substituted, for sorbitol in the resuspending medium (RS1).

37(h) Purification (i) Washing. Chloroplasts prepared by the simplest one-spin procedures are very active and, once the procedures have been mastered and solution etc made ready, it is possible to go from intact leaf to chloroplast suspension in about 15 minutes. For general purposes such chloroplasts are also very pure. The extent of contamination by other cellular constituents, such as mitochondria, etc., is minimal but it does exist. For example, such chloroplasts will readily evolve O2 in the dark if given hydrogen peroxide. The contaminating catalase which is largely responsible for this reaction (Eqn. 41.7, Section 41c) comes from peroxisomes which are ruptured during grinding. Apparently this enzyme then becomes loosely adsorbed to the chloroplast envelope. Washing, i.e. resuspension of the original pellets in relatively large quantities (roughly equivalent to the original volume of grinding medium) of a washing medium, followed by a second centrifugation, will remove a large fraction of such contamination. Washing media can contain almost anything in addition to the osmoticum which (with the possible exception of K+ - see 4f) is the only really essential component. Grinding medium will suffice or resuspending medium if it is essential or affordable. Two washes can be better than one for some purposes but it should be borne in mind that all of the handling procedures carry the risk of mechanical damage to the chloroplast envelope and that prolonged washing may also lead to undesirable losses (e.g. leaching of K+ etc). Similarly, ageing (itself still a very ill-defined process) will continue to occur at an accelerated rate from the moment that the chloroplast is separated from its cellular environment. (ii) Percoll. A number of procedures involving silica-sol gradients have been reported (see e.g. Morganthaler et. al., 1974; Price et. al., 1979) but the following, based on a method introduced by Mills and Joy (1980) is quick and cheap (because it does not call for a great deal of Percoll). All it involves, is resuspending the initial pellets (37e) in about 6ml of medium, transferring this to two 50ml glass centrifuge tubes, and introducing a 4ml “cushion” (below the chloroplast suspension) of 4 parts by volume of “Percoll” (a modified silica gel from Pharmacia) in 6 parts by volume of resuspending medium. This cushion of 40% (v/v) Percoll may be put into position by using a syringe or Pasteur pipette. After centrifugation at 1700g for 1 minute, followed by deceleration without braking, the intact chloroplasts are pelleted whereas the fragmented chloroplasts and other cell debris and organelles are retained at the interface between the Percoll and the added chloroplast suspension. The supernatant and the Percoll layer are aspirated and the pellet resuspended. Purification can be achieved within 15 minutes in this way and gives cleaner and more active preparation. Inevitably, it is not done without cost because the yield is only 30-40% of that in the original pellet, indicating that some intact chloroplasts must be retained at the Percoll interface and thereby causing concern that this procedure might select in favour of particularly dense, as well as intact, chloroplasts. There is also one report that Percoll can be inhibitory (Stitt and Heldt, 1981) although this inhibition (which is not in line with general experience) could be overcome by dialysis. More sophisticated gradients have also been successfully employed e.g. by Mourioux and Douce (1981).

38. SPINACH 38(a) Why Spinach? Spinach (Spinacia oleracia L.) is undoubtedly a trying and, some might even say, a perverse plant but it could be argued that its use has done at least as much to further our understanding of photosynthesis as that of its distant cousin Chlorella or its C4 counterparts, sugar cane and maize. Why, we might ask, was spinach chosen to play such a major role? Part of the answer lies in the fact that spinach is a horticultural crop of importance in Europe, particularly in the Netherlands. European traditions die hard in the United States and there too it has been readily available to the plant biochemist who wished to buy it from his or her corner store. No doubt much of plant science is still totally inadequate because most European and North American scientists remain lamentably uninformed about tropical species but the fact remains that spinach is largely a traditional choice, reflecting the origin of its users. This, of course, is only part of the story. These same scientists would have had even less difficulty in laying their hands on cabbage or lettuce. However, the former is never used and the latter does not easily yield fully competent CO2-fixing chloroplasts and has only rarely been first choice. The fact is that spinach also has some intrinsic virtues. It is soft (and therefore readily disrupted) but, most of all, it is largely free of phenolics and other inhibitory compounds which are prevalent in the majority of cultivated species. Having said that, no one in their right mind would choose to work with it because it is, or can be, so very difficult to grow.

God banishes spinach from heaven 38(b) The Origin of Spinach The genus Spinacia belongs to the family Chenopodiaceae. Compared with a really ancient crop like Amaranthus edulis, (which was grown in Mexico from about 4800 BC and from which the Aztec emperor Montezuma received an annual tribute of 200,000 bushels of grain), spinach is a relative newcomer but it was mentioned in Arab and

Chinese writings as being consumed in and before the 10th century and it was grown in Spain as early as the 12th century. It is not a tropical species but, contrary to fairly widely held views, it can do well in warm climates and is believed to have originated in Iran, or that immediate area (Arab, isfinâj; Pers, isfânâj). 38(c) Growing Spinach

Spinach flowers in long days

There is a famous book called “Growing Madder” which describes the cultivation of Madder (Rubia tinctorum) which was the source of the once important red dye alizarin. Its title would also probably be an apt description of the feelings of anyone who has set out to grow spinach for scientific purposes. One reason is that spinach is a “long-day” plant i.e. one which flowers or “bolts” when the daylength to which it is exposed exceeds a critical length. The critical daylength varies with the cultivar and, especially in the Netherlands, considerable effort has been put, by plant breeders, into the production of cultivars which are best suited to sowing at different times of the year. In general, however, the critical daylength is about 10 hours and in the north of the United Kingdom, where summer days can be as long as 21 hours, there is little hope of growing spinach in the summer without it “bolting”. At worst, this will result in virtually no leaf. Such spinach will give rise to a leaf about as large as a thumb nail and then, presumably feeling fulfilled, will flower and die. Conversely, optimal growth is only achieved at or about the critical daylength, and in fairly high light. Again in the U.K., this means that the daylength must be artificially shortened in the summer and that both light intensity and daylength must be artificially augmented in the winter (Fig. 39.1). Nearer the equator, spinach can be grown in the field for most, or even all, of the year but it remains a crop which requires adequate irrigation and plenty of nitrogen. Ironically, in many countries in which it would undoubtedly grow well it is virtually unknown. “Spinach” may be available in local markets but it will almost certainly turn out to be a totally different species (see Section 38d) which will not easily yield actively competent CO2-fixing chloroplasts. 38(d) Alternatives and What to Avoid It has to be said, immediately, that there is no real alternative to spinach. This is not only because spinach chloroplasts are usually so much better than those prepared mechanically from other species but because, by now, there is such a huge scientific literature on spinach chloroplasts that there would be no point in looking elsewhere unless there was a very good reason for so doing. Very good chloroplasts can be prepared from pea (Pisum sativum) leaves but they are different, in some important ways from spinach chloroplasts (see e.g. Stankovic and Walker, 1977; Robinson and Wiskich, 1976) and rarely match those from Spinacia in terms of activity. Recently, good chloroplasts have been prepared from sugar beet leaves (Robinson, 1983). Many other species have been called “spinach” but they are best avoided and their chloroplasts will certainly not compare with those from spinach if prepared by the procedures described here. “Perpetual spinach”, for example is a cultivar of Beta vulgaris and “New Zealand spinach” is Tetragonia expansa. Neither is a sensible choice.

38(e) Varieties Some varieties of spinach have become fashionable from time to time and have become widely used in consequence. Perhaps the most famous is Yates Hybrid 102 alias U.S. National Hybrid 74 from the Ferry Morse Seed Company, P.O. Box 100, Mountain View, California 94042. “Virtuosa” from Rijk Zwaan, P.O. Box 40-2678 ZG Burg, Crezeelaan 40-2678 KX De Lier, Holland is particularly well suited to water culture. Growing conditions are often more important than varietal differences, however, and the prospective user should not turn to alternatives unless locally available spinach consistently fails to match the rates reported in the literature. Both Robinson (1983) who used Pisum sativum, var. Massey Gem, in Australia and Cerovic and Plesnicar (1984) who used var., Mali provensalac, in Jugoslavia reported exceptionally good rates but, again, locally available varieties might well have much to offer.

39. HYDROPONIC CULTIVATION OF SPINACH 39(a) General Procedure At what is now the Robert Hill Institute in Sheffield, we have had more than 10 years experience of growing spinach in water culture. While we feel that we could comfortably use another century of experience without even nearing perfection, we have devised a system (Fig. 39.1) which works well. Seeds are sown in polystyrene drinking cups with a perforated base. Regrettably, while such cups are cheap and readily obtainable, there is no great call for cups with holes and the perforations have to be done by hand. The cups are disposable but can be used more than once for economy. They drop into holes in polystyrene supports which are available commercially (e.g. from Accelerated Propagation Ltd, Vines Cross, Heathfield, Sussex, U.K.) for supporting more conventional plastic plant pots (which may be preferred in some systems). If the polystyrene support is now supported in turn by the edges of a tank fashioned out of wood, plastic or whatever, and lined with polythene sheet, the bottoms of the cups will dip by a centimetre or two into a water-culture solution. This is aerated using a compressor and glass tubing (lying on the floor of the tank) fitted with capillary air outlets. A pump raises the level of solution in the tank to that of a large “over-flow” through which the solution returns to a reservoir, below the tank. The pump (activated by a time-clock) raises the level in the tank to that of the main over-flow for an hour or so twice a day. When the pump is switched off the solution falls, through small holes made near the top of the over-flow pipe, to a level just below the cups. This ensures that the contents of the cups are moistened twice daily and also that the cups can drain freely back into the tank. The roots can also grow through the perforations in the floor of the cups into the solution. This device ensures that the hypocotyl region of the seedling is kept relatively dry (and therefore healthy) but that, at the same time, the developing plant is kept well irrigated and well supplied with inorganic nutrients.

a cultivated spinach

Figure 39.1. Detail of hydroponic system for sinach. For full explanation see text. Plants are grown in polystyrene drinking cups and these provide an indication of the size of the nutrient tanks, which can be of any convenient length. The sidth is dictated by the size of the commercially available supports in which the cups are suspended over the tanks. Natural light is augmented by 400W Wotan lamps (metal halide with disprosium from: Wotan Lamps Ltd, Wotan House, 267 Merton Road, London, SW18 5JS, UK.) In the UK., summer daykength necessitates day-shortening to prevent flowering and this is achieved by using an electrically operated, spring tensioned blind which runs the length of the tank and is controlled by a time-clock to give an 11 hour day.

39(b) Sowing Two or three seeds are sown on to a mixture, in each cup, of vermiculite (heat-expanded mica), perlite (alumino-silicate growing medium) and peat, in roughly equal proportions. The number of seedlings in each cup is then decreased to one by hand “weeding”. If small leaves are preferred, seeds can be sown broad-cast, in a similar mixture, in large well-drained trays about 12 cms deep. 39(c). Water Culture Solution. A solution such as the following (Table 39.1) is appropriate Table 39.1. Water Culture Solution for Spinach etc.,

Stock concentration

Volume (ml) per 20 litre nutrient solution made up with distilled water

KNO3 Ca(NO3)2 MgSO4 KH2PO4 MgCl2 Trace elements NaFe-EDTA

1M 1M 1M 1M 1M (B,Mn,Zn,Cu,Mo) (3.86 gm/250 ml)

120 80 40 20 80 20 20

Trace elements

Quantity (mg) in 250 ml H2O

Solution

H3BO3 MnCl2 ZnSO4 CuSO4 NaMoO4

4H2O 7H2O 7H2O 2H2O

715 452 55 20 7.25

Commonly available mixtures (such as “Solufeed” from I.C.I. Plant Protection Division, U.K. Department, Bear Lane, Farnham, Surrey, GU9 7UB, U.K.) have much to recommend them because they are time-saving, relatively inexpensive and minimise errors. 39(d) Cleaning Tanks A good standard of hygiene will obviously minimise disease. Regular sterilisation of tanks with a 5-10% hyperchlorite solution and periodic replacement of the plastic sheeting is advisable. 39(e) Pests and Sprays The worst pests are usually your colleagues who, unless properly trained from a tender age, are likely to decimate a growing crop without prior consultation or use plants which you had reserved for an epoch-making experiment. Other natural disasters will inevitably and invariably occur at the worst possible times so, if you are not the sole user and sole cultivator, be sure to keep your glasshouse in order.

Happily, spinach is not greatly troubled by other pests (at least in temperate climates) except greenfly (aphids). The use of pesticides should always be avoided because even those which are supposedly specific will almost certainly affect some aspect of photosynthesis and many a researcher has had occasion to curse a well-meaning but un-informed sprayer. Biological control is fine and ladybirds (lady bugs, Coccinelliidae sp.) in both the adult and larval form are rapacious devourers of aphids. Sadly, they are so good that they have a tendency to eat themselves out of house and home and the temptation to start breeding aphids to feed ladybirds is one which is probably best resisted. As always, much can be done by good husbandry. The faster the turnover the less likely you are to be troubled by problems of all kinds. Minimise cross infections by growing young plants some distance away from older plants if this is a practical possibility. Try to achieve a pest-free house by limiting access. If all else fails, fumigate with nicotine but bear in mind that nicotine is lethal and that while the spinach will recover in a matter of days you might easily die if you enter a glasshouse during fumigation. Biological control 39(f) Harvesting If someone has always used young spinach leaves to good effect it is of no avail to attempt to persuade them that older leaves may, almost certainly, be equally useful. There is also merit in using young plants as a means of maintaining healthy stock (Section 39b). Otherwise it is a matter for compromise. Depending on light and temperature a spinach plant can reach maturity in about 6 weeks from sowing. For the first couple of weeks it will be really too small to use. In weeks 3-5 it will be in the “grand period of growth”, sometimes increasing in size in an almost alarming manner. Accordingly, if you need a lot of spinach you will either have to settle for older leaves or devote an inordinate amount of time to growing it.

40. THYLAKOIDS If intact chloroplasts are resuspended in media which has been diluted 10-fold, they swell and burst within about 1 min. The stroma is almost entirely released to the medium and the thylakoids or “free-lamellar” fraction may be recovered by centrifugation as before. These so called “broken chloroplasts” are ideal for experiments on photophosphorylation, electron transport etc in which the intact envelope might prevent access to an electron acceptor or co-factor. There is everything to be gained, in fact, by preparing “broken chloroplasts” from intact chloroplasts rather than homogenising leaves in low osmolarity medium. This latter procedure ensures that the thylakoids will be exposed to other cellular components during isolation. Conversely the intact envelope will protect the thylakoids until they need to be “unwrapped” by osmotic shock.

41. EXPERIMENTS WITH ISOLATED CHLOROPLASTS The number of experiments which have been carried out with isolated chloroplasts is legion and it might be reasonable to assume that such experiments will continue to be carried out into the forseeable future.

Here we are concerned specifically with experiments based on O2 measurements and much of what follows simply offers the reader an opportunity to become acquainted with chloroplasts and, if he or she so wishes, to repeat one or two (now ancient) experiments which have contributed to our understanding of photosynthesis. The real classic, of course, is the Hill Reaction (Hill, 1937). When Robert Hill first isolated chloroplasts in the 1930’s the events of the second world war which were to lead to the eventual availability of radioactive isotopes were still just around the corner and nearly twenty years were to elapse before the invention (by Clark in 1956) of the membrane-covered O2 electrode. Many electron acceptors or co-factors which we now take for granted were either not yet described or not commercially available. Against this background, Hill offered his chloroplasts iron, in the form of ferric potassium oxalate, as an electron acceptor and followed O2 evolution in a “spectro-colorimeter” by measuring the change in absorption spectrum as the evolved oxygen combined with muscle haemoglobin to give oxyhaemoglobin. Today, ferricyanide (rather than ferric potassium oxalate) is often used as a “Hill oxidant” and, in Experiment 1, we shall combine nostalgia with present day practicability and use the Hill reaction with ferricyanide as a measure of chloroplast “intactness”. 41(a) Experiment 1. The Hill Reaction as an Intactness Assay. Although ferricyanide is a widely used Hill oxidant it does not cross the intact chloroplast envelope at an appreciable rate (Lilley et. al., 1975) and will therefore only react freely with thylakoids. Happily, this makes it a very convenient material for determining the percentage intactness of the chloroplast envelope. Clearly, if you had succeeded in preparing 100% intact chloroplasts and you illuminated them in the presence of ferricyanide you would not, in theory, expect to see any O2 evolution. Conversely, if you were then to subject these same chloroplasts to osmotic shock, so that their envelopes would rupture, the ferricyanide would have ready access to the thylakoid membranes and O2 evolution should be observed. Preparations approaching 100% intactness are rare, so some O2 evolution is almost always seen but, since intact chloroplasts are readily made 100% envelope-free, comparison of the rates of O2 evolution observed before and after osmotic shock can be used as an intactness assay. This experiment also demonstrates the increase in the rate of electron transport associated with uncoupling (see also Sections 41c and 45c). As electrons from water are passed through the photosynthetic electron transport system, protons are also “picked up” from the outside of the thylakoid compartment and released into the internal space or lumen. This is because some of the electron transport chain can accept or donate electrons whereas others can only accept or donate hydrogen and therefore require both protons (H+) and electrons (e-) - see Edwards and Walker (1983) 2H+ + 2e- → H2

......Eqn.41.1

Chlorophyll, for example, when excited by light of an appropriate wavelength, will emit an electron thereby creating a positively charged hole which can accept an electron from water. Further along the electron transport chain, this electron is offered to plastoquinone (see

Fig. 41.2 and also Part B for further detail). While gracefully accepting this proffered electron, plastoquinone simultaneously grabs a proton from the outside of the thylakoid membrane in order to make up the hydrogen atom that it needs in order to become a fully fledged plastoquinol.

Figure 41.1. The Hill Reaction. Ferricyanide was used as the oxidant or electron acceptor. (Details of the reaction mixture are given in Table 41.1). Note the acceleration caused by the addition of NH4Cl as an uncoupler and the difference in rate between “whole” and “broken” chloroplasts which can be used to evaluate “intactness” (see text). Subsequently, plastoquinol in its turn offers a hydrogen atom to a cytochrome (in the cytochrome b, cytochrome complex). Cytochromes, however, can only handle electrons and, such is the structure of the thylakoid membrane, that the rejected proton is discarded at the inner surface. This is an essential and important feature of photosynthetic electron transport because, according to Mitchell’s Chemiosmotic theory, the protons which accumulate within the thylakoid space then discharge through the ATPase into the stroma (Part B) and it is the movement of protons through this sluice gate which drive the formation of ATP in the process of photophosphorylation (Eqn.41.2). ADP + Pi → ATP + H2O .........Eqn.41.2 Needless to say, this account is a gross over-simplification of what really occurs. Proton gradients are established and can, e.g., be easily observed with a pH electrode if thylakoids are illuminated in an unbuffered mixture containing an appropriate “cofactor” of cyclic electron transport, such as pyocyanine. (If ferricyanide is used as an electron acceptor there is a change in pH associated with its reduction [Eqn. 41.4]. This in itself has nothing to do with the establishment

Figure 41.2. Consequences of Illumination. The pigments and electron transport chain are housed in the thylakoid membrane. On illumination, electrons are donated to excited chlorophyll in photosystem II (PSII). The transfer of electrons to photosystem I (PSI) via plastoquinone (PQ) requires protons that are taken up from the stroma and released into the thylakoid compartment. The stromal pH rises and Mg2+ moves as a counter ion. [ATP] is formed from [ADP] and [Pi] as H+ is discharged through the ATPase in photophosphorylation. of a proton gradient). Pyocyanine returns the electron that it receives to an earlier point in the electron transport chain and this rapid cycling of electrons very quickly establishes a measurable proton gradient. However, the movement of H+ implies a separation of electron charge and, although charge separation undoubtedly occurs and may be regarded as an integral part of photophosphorylation, biological membranes are not thought to be capable of accommodating large electrical imbalances. Movement of Cl- ions into the lumen and Mg++ ions out of the lumen will tend to lessen these imbalances. Similarly, the mechanism by which ATP formation is driven by the discharge of the proton gradient is still unknown. Nevertheless, the concept in its most simple form is still very useful and one which allows us to understand why electron transport goes faster if it is either coupled to photophosphorylation OR uncoupled from it. The picture which then unfolds is as follows. As protons accumulate within the thylakoid space they exert a back-pressure. It becomes more and more difficult to discharge protons into this space as the proton concentration increases, just as it becomes more and more difficult to blow air into a balloon as it expands. Accordingly, electron transport will slow down. If, on the other hand, the proton gradient is free to discharge through the ATPase (Part B, Fig. 8.3) i.e. if there is an adequate supply of ADP and Pi, electron transport (now coupled to photophosphorylation) will accelerate. Similarly, if it is uncoupled, i.e., if an “uncoupler” provides another means of discharging the proton gradient, electron transport will again increase. The most easily understood uncoupling action is that caused by detergents. These simply make holes in the lipid/protein membranes of the thylakoid through which protons can escape (Part B. Fig. 8.3). Ammonium chloride, which is used in this experiment, works differently. Free ammonia (NH3) crosses the thylakoid

NH3 + H+ → NH4+ ...........Eqn.41.3 membrane and is reprotonated (Equation 41.3) in the thylakoid compartment, thereby acting as a proton sink. The experiment itself is based on two reaction mixtures which are identical, except in the order of addition of the reactants (Table 41.1). Simply varying the order of addition in this way, gives two reaction mixtures which contain either chloroplasts which are all ruptured or chloroplasts which maintain the same degree of intactness as before Table 41.1. Reaction Mixture for Intactness Assay. INTACT l.0ml RS1 x 2 0.8ml H2O 0.1ml chloroplasts 10µl D,L-glyceraldehyde(2M) 10µl K Ferricyanide (0.5M)

RUPTURED 0.8ml H2O 0.1ml chloroplasts 1.0 ml RS1 x 2 10µl D,L-glyceraldehyde (2M)

The reaction mixtures contain the resuspending medium listed at the beginning of Section 37(g) (the BSA is optional). It is added at double strength (RS1 x 2). That on the left is diluted to single strength prior to the addition of chloroplasts which therefore experience no change in osmotic pressure when they are added to it. On the right, chloroplasts are added to water so that they experience a 9-fold dilution which causes them to swell and rupture almost immediately. In practice, they are usually left for l min at this osmolarity in order to allow them all to burst. Double strength resuspending medium is then added in equal volume so that the two reaction mixtures are finally identical except for the intactness of the chloroplasts. D,L-glyceraldehyde is added to inhibit CO2 fixation. To obtain fast and more linear rates (i.e. to avoid any retardation caused by the build-up of a proton gradient) NH4Cl (say 10 l of a 0.5 M solution) is added to each after a minute, or two in order to uncouple electron transport. The degree of uncoupling (i.e. the difference between the coupled and uncoupled rates) is also an indicator of competence. Ferricyanide-dependent O2 evolution by well-coupled chloroplasts may increase in rate by a factor of 10 or more, following the addition of an uncoupler. Percentage intactness is calculated from the relationship % intactness = (A - B) x 100 A in which A is the rate after osmotic shock and B is the rate prior to osmotic shock. An ancillary to this experiment is to add a smaller amount of carefully made up ferricyanide solution. (Take a large crystal of potassium ferricyanide and wipe off surface ferrocyanide with a tissue. Dissolve in an appropriate volume of water). The Hill reaction with this oxidant (electron acceptor) will go virtually to completion (i.e. until nearly all of the ferricyanide has been reduced). This adequately demonstrates the need for an acceptor and, depending on how you wish to look at it, will

verify the accuracy of calibration (Section 36) or establish the stoichiometry of ferricyanide reduction by water (Equation 41.4). 2H2O + Fe+++ → 4Fe++ + 4H+ + O2 .....Eqn.41.4 41(b) Some practical points The above experiment has become a routine first step in work with intact chloroplasts and it is always worth doing in a teaching context. It demonstrates Hill’s classic experiment while also introducing the concept of uncoupling (an aspect of photosynthesis which would not have even come to that great man’s attention for another twenty years). It also illustrates one or two useful practical details. The first concerns the use of double strength media. This is a very convenient device because it can be used to avoid large unintentional decreases in osmotic pressure. For example, if you wish to work in a final volume of 2 ml, you would simply ensure that the total volume of all of the other additives was less than 1 ml and then add H2O to bring up the total to 1 ml followed RS 1 x 2 to a final volume of 2 ml (adding the chloroplasts last!). Each additive would, of course, make its own slight contribution to the osmotic pressure but these contributions are small and can usually be safely disregarded. Subsequent addition (e.g., the addition of NH4Cl in Experiment 1) should be made in small volume to avoid a significant effect on the calibration. Ten microlitres added to 2 ml is less than a 1% increase in volume and is therefore of little consequence. When all of the constituents have been added to a reaction mixture, the vessel is “sealed” by a plunger. The depth to which the plunger enters the reaction vessel is dictated by a threaded collar which should be adjusted by hand until the mixture just enters the capilliary at the top of the cone. Further additives, in small volumes, can be made with a micro-syringe via this neck and the collar should always be subsequently readjusted. Particularly if samples are being withdrawn, care should be taken to ensure that the mixture always just enters the capillary. If a bubble of air is left at the top of the cone its oxygen will slowly equilibrate with the rest of the mixture and since it will contain more O2 than there is in the solution, this could give rise to errors in any work which demands a very high degree of precision. The chlorophyll content should not exceed 100 g/ml if light saturation is necessary and smaller amounts (consistent with convenient rates) are to be preferred. Large quantities of active chloroplasts can produce O2 at such high rates that bubbles will form, again giving rise to inaccuracies. A polished, semi-circular cross-section, reflector (which rests against the back of the water-jacketed vessel) will help to shield the contents from laboratory light when the vessel is not illuminated and help to maximise illumination when it is. Ideally, the water circulating through the jacket should be filtered and contain an agent (such as “Myacide AS”, Boots Biocides Group, Chemical Marketing, The Boots Company PLC, Nottingham, NG2 3AA, U.K.) to inhibit algal and bacterial growth.

vessel “sealed” by plunger

In Experiment 1, D,L-glyceraldehyde is used (Stokes and Walker, 1972) to inhibit CO2 fixation because, unless great care is taken to exclude it, there will be enough dissolved CO2 present to allow CO2-dependent O2 evolution to start in the mixture which still contains the intact chloroplasts. Glyceraldehyde is a “nice” inhibitor in the sense that it appears to be reasonably specific for the reductive pentose phosphate pathway (i.e. it inhibits CO2 fixation but does not affect electron transport or photophosphorylation). Glycoaldehyde can be used for the same purpose (Sicher, 1984) other inhibitors (cyanide is an obvious example) and electron acceptors (methyl viologen is more poisonous than cyanide) kill people as well as chloroplasts and should always be used with extreme caution. Yet others (e.g., DCMU) are only rinsed with difficulty from the electrode vessel (a detergent which is not, in itself, desirable may have to be used) and if washing is ineffective can carry over into subsequent reaction mixtures. Some of Good’s buffers (e.g Hepes) are photoxidised in white light in the presence of flavin containing mixtures and red light may be preferred if there is any possibility of this occurring (see Section 41c). Washing, between experiments, is essential to good work. The vessel should be aspirated and flushed with distilled water at least ten times (don’t swallow the flea and don’t forget to wash the plunger).

Inhibition of PGA-dependent O2 by D.L-glyceraldehyde. Continuous line O2, points CO2

If microsyringes are used, always bear in mind that the floor of the electrode vessel is covered by a delicate membrane which should not be damaged. 41(c) Experiment 2. The Mehler Reaction. Before we leave the Hill reaction entirely we ought to remind ourselves of one special version of it. This is named after Mehler (1951) and involves O2 uptake rather than O2 evolution (Figs. 41.3 and 41.4). Mehler reagents (MR in Eqns. 41.5 and 41.6) are Hill oxidants which accept electrons from water but are then autoxidised by molecular oxygen in reactions which lead to hydrogen peroxide formation (Equation 41.5) H2O + MR → MRH2 + 1/2 O2 ......Eqn.41.5 MRH2 + O2 → MR + H2O2

......Eqn.41.6

In the presence of catalase, hydrogen peroxide undergoes rapid dismutation to water and O2 (Equation 41.7) H2O2 → H2O + 1/2 O2

....Eqn.41.7

O2 evolution and O2 consumption are then equal so that no change is observed but, if catalase is removed by washing, or inhibited by azide, the Mehler reaction gives rise to a net O2 uptake. Flavin mononucleotide (FMN) was used as the Mehler reagent in Figs. 41.3 and 41.4 but it should be noted that flavin compounds (including FMN, FAD and flavoproteins) will, bring about the photoxidation of commonly used buffers such as Hepes (Yamazaki and Tolbert, 1970; Andreae, 1955) and, if this is to be avoided it is necessary to use red, rather than blue or white light.

Figures 41.3 and 41.4. The Mehler Reaction. Figure 41.3 (left) compares the Hill Reaction (with ferricyanide as oxidant), with a Mehler Reaction in which FMN (0.04 µmoles) was used as the Mehler reagent. It will be seen that the rates were similar and that both were accelerated by the addition of NH4Cl (20 µmoles) as an uncoupler. In Figure 41.4 (right) several Mehler reagents (all 0.1 µmoles except ferredoxin which was 0.006 µmoles) are compared. Note that the rates did not change appreciably until nearly all of the detectable oxygen in solution had been consumed. The Mehler Reaction is a bit more time consuming to set up than the Hill Reaction because of the probable presence of catalase in the chloroplast preparation but it can be a lot of fun. Catalase is not a chloroplast enzyme but the normal mechanical preparative procedures rupture the peroxisomes in which it is normally located and smears their catalase on the chloroplast envelopes. Washing (Section 37h(i)) will remove about 10% of this contamination and, if these washed intact chloroplasts are osmotically shocked and spun again, there should be another five-fold decrease in catalase. [If you wish to see how effective your washing is, add a little hydrogen peroxide (about 10 µmoles/ml of reaction mixture) to your chloroplasts and record the rate of O2 evolution in the dark]. The last traces of catalase are difficult, if not impossible, to remove but what can’t be cured need not, in this case, be endured because a little (0.5 µmoles/ml) cyanide or azide (“it just takes a smidgeon”) will inhibit what is left. Without inhibitor, nice steady-state relationships can be established in which light-generated H2O2 formation and catalatic dismutation of H2O2 (Eqn. 41.7) come into balance (Fig. 41.5). The manganous ion (Mn++, added as MnCl2) accelerates the Mehler reaction and an oxidation product, possibly manganic ion (Mn+++) will reoxidise H2O2. This can result in post- illumination O2 evolution (Walker et.al., 1970).

“it just takes a smidgeon”

Figure 41.5. Establishment of steady-state between O2 uptake and catalatic dismutation of water (Eqn. 41.7). Commercial catalase (5 units) was added at the outset to both reaction mixtures which contained FMN as the Mehler reagent. In this sort of experiment, the level at which the first steady-state is established can be raised or lowered by lowering or increasing the light intensity (after Whitehouse et al, 1971). The reaction mixture for the Mehler Reaction is much the same as before (Table 41.1) but the order of addition will be unimportant since you will be using chloroplasts which have already been osmotically shocked (see above). Obviously you will not need a “Hill oxidant” (like ferricyanide) but you will require a “Mehler reagent” which is also an electron acceptor but one which, alone or in combination with another, will undergo autoxidation by molecular oxygen (Eqn. 41.5). Methyl viologen (which is the active component of the weed-killer “Paraquat”) is a very good Mehler reagent but it is also horrendously toxic and should NOT IN ANY CIRCUMSTANCES BE PIPETTED BY MOUTH. (Many have survived ingestion of small quantities of cyanide but paraquat poisoning has been almost invariably fatal. Moreover, it can induce a proliferation of cells in the lung which is associated with, or brings about, death after 2 or 3 days; so it can be an extremely unpleasant way to go). For these reasons its use is not recommended. Flavin coenzymes (such as FMN or FAD) are O.K. although they should not, as already noted above, be used in light containing blue wavelengths. The Mehler Reaction, like the Hill Reaction will, of course, respond to uncouplers and to the presence of ADP + Pi. This can be used to demonstrate so-called “photosynthetic control” (West and Wiskich, 1968) in which ADP (added to a mixture already containing Pi) stimulates the rate of O2 evolution by discharging part of the proton gradient (c.f., the action of uncouplers, Section 41c above). The O2 consumed during the period of accelerated O2 uptake (Fig. 41.6) can be used to calculate a P/O ratio, which in this instance (Whitehouse et. al., 1971) was about 1 (i.e., about 1 mole oxygen/0.2 µmoles of ADP added).

Figure 41.6. “Photosynthetic control” in the Mehler Reaction. Both reaction mixtures contained KCN (1 µmole), to inhibit residual catalase, and orthophosphate (2 µmole) “A” contained FMN (0.1 µmole) and “B” contained methyl viologen (0.1 µmole). ADP was added as indicated. 41(d) Experiment 3. CO2 and PGA-dependent O2 evolution. This is another golden oldie. Neither quite so venerable nor epoch-making as the Hill Reaction but certainly important in its own right. The mixture is very much the same as before except that the chloroplasts are kept intact and either CO2 or PGA are used as electron acceptors. Reaction Mixtures CO2-dependent

PGA-dependent

RS1 x 2 (1.0ml) H2O (0.7ml) 0.2M NaHCO3 (0.1ml) chloroplasts (0.1ml) 1.0 mM Pi (0.1ml)

RS1 x 2 (1.0ml) H2O (0.7ml) 200mM PGA (10µl) chloroplasts (0.1ml) 1.0 mM Pi (0.1ml)

It will be seen that upon illumination (Fig. 41.7), CO2-dependent O2 evolution does not immediately reach maximal rate (c.f., ferricyanide reduction, Fig. 41.1) but only after a lag or induction period. The underlying causes are partly light-activation of enzymes but mostly autocatalytic build up of carbon cycle intermediates which have become depleted in the dark (Edwards and Walker, 1983). For this reason the addition of PGA shortens or eliminates the lag - a predictable result but one which nevertheless caused the author to rush naked into the street crying “Eureka” when he did the experiment for the first time.

Figure 41.7. Carbon dioxide-dependent and PGA-dependent oxygen evolution. This figure shows the lag or induction period which is characteristic of CO2-dependent O2 evolution by chloroplasts and leaves (c.f., Fig. 37.3). The shortening of the lag by PGA is typical but the rate with PGA is usually slower than the rate with CO2. In any case, although the reaction mixture containing PGA contained no added bicarbonate, it had not been made CO2-free and hence the O2 evolution observed would be partly CO2-dependent stimulated by PGA. In the complete absence of CO2 (or in the presence of D,L-glyceraldehyde), PGA-dependent O2 evolution soon ceases. The discerning reader will also note that the second part of this experiment is not, strictly speaking, PGA-dependent O2 evolution but a mixture of PGA-dependent and CO2-dependent because although bicarbonate is not added there will still be some CO2 in the various solutions. In fact, if CO2 is rigorously excluded, PGA-dependent O2 evolution soon falls in rate for reasons that have yet to be certainly established. Concomitant oxidation of RuBP accumulating in the absence of CO2 may be a contributory factor but the decline in rate also occurs in the presence of D,L-glyceraldehyde which is believed to block the cycle at the transketolase steps (Stokes and Walker, 1972).

PGA-dependent oxygen evolution

The first demonstration of rapid CO2-dependent O2 evolution under aerobic condition (Walker and Hill, 1967) added the second dimension to the photosynthetic capacity of the isolated chloroplasts. (Originally, Hill could only detect O2 if he provided his chloroplasts with an artificial oxidant and, when Arnon and Whatley first demonstrated CO2-dependent O2 evolution under anaerobic conditions the rates were extremely low). Observations such as these following on the pioneering work of Arnon, Whatley et. al., (see Arnon, 1967 for a review) appeared to confirm the supposition, in doubt since Hills 1930’s experiments, that the chloroplast was a self-sufficient photosynthesising organelle. Subsequent experiments, which you are also invited to repeat (Experiments 4 and 5 below), made it clear, however, that the chloroplast is also a “Pi-consuming, triose phosphate-exporting”, organelle

and that in this and other regards photosynthesis is, after all, really a cellular affair. During the course of CO2-dependent O2 evolution it is easy, for purposes of demonstration, to make some additional points. One is simply to introduce a brief dark interval (Fig. 41.8). It is always

Photosynthesis and phosphate: a cellualr affair

Figure 41.8. Carbon dioxide-dependent oxygen evolution. To show the effect of light intensity and temperature on the induction period. Note the near absence of a lag following re-illumination after a dark interval. reassuring to see that the chloroplasts know when they are in the dark and instructive to observe that there is no lag when illumination is resumed (built-up intermediates do not decline sufficiently rapidly to reinitiate the induction period if the dark interval is limited to a duration of about 1 minute). Changes in light intensity will also be seen to affect the rate. 41(e) Experiment 4. The Requirement for Orthophosphate. When CO2 fixation by isolated chloroplasts was first demonstrated in the 1950’s (Allen et. al., 1955; Arnon, 1967) and when rates were improved in the 60’s (Walker, 1964) chloroplasts normally failed to incorporate radioactivity into sucrose. This was commonly regarded as a malfunction caused by isolation, particularly since key enzymes of sucrose synthesis were found to be associated with chloroplasts prepared in non-aqueous media. Eventually, however, it become clear that sucrose synthesis was a cytosolic event. Two of the experiments (Experiments 4 and 5) which made it clear that chloroplasts imported orthophosphate and exported triose phosphate are described here. The reaction mixture is the same as that used for CO2-dependent O2 evolution in Experiment 2 except that no orthophosphate (Pi) is included. When the mixture is illuminated, photosynthesis normally

Figure 41.9. The Requirement for Orthophosphate. In Pi-deficient chloroplasts, photosynthetic O2 evolution falls to a value commensurate with starch formation. The addition of Pi then gives approximately 3 molecules of O2 per molecule Pi. In a reaction mixture containing no Mg++ there is no response to added PPi unless Mg++ is added. External hydrolysis then leads to Pi formation and a stoichiometry of 6 molecules of O2 per molecule PPi. starts, as usual because there is enough endogenous Pi to support some CO2 fixation. Nevertheless, the rate soon declines until it is commensurate with the internal Pi recycling associated with starch synthesis (Edwards and Walker, 1983). At this stage, rates can be readily restored by the addition of Pi. If small quantities of Pi are used, it will be seen that there is a stoichiometric relationship between added Pi and extent of the restoration in rate (such that 1 Pi yields 3 O2). This is consistent with the formation of triose phosphate according to Equation 41.8.

(Sum) 3CO2 + H2O + Pi → triose phosphate + 3O2 .....Eqn. 41.8 41(f) Experiment 5. Orthophosphate Inhibition and its Reversal The orthophosphate requirement (Experiment 4) for photosynthesis by intact chloroplasts has a very sharp optimum. This can be readily demonstrated (Fig. 41.10) by using the same reaction mixture as that in Experiment 4 but including different amounts of Pi to give final concentrations between 0.5 and 10 mM. It will be seen that the initial

lag is extended as the Pi concentration is increased until the onset of O2 evolution is delayed more or less indefinitely as it is in Fig. 41.11. At any time, this Pi inhibition may be immediately and fully reversed by the addition of PGA or triose phosphate. Slower reversal follows the addition of pentose monophosphates. When first performed (Cockburn et. al., 1967), experiments of this sort led to the suggestion that there must be obligate exchange between external Pi and internal triose phosphate (Walker and Crofts, 1970). This was confirmed by direct measurement (Heldt and Rapley, 1970) and the concept of the phosphate translocator became a reality. Seen in this way, triose phosphate is the principal product of the photosynthesising chloroplast and photosynthesis is therefore severely constrained if it is supplied with inadequate Pi.

Figure 41.10. Inhibition by orthophosphate of photosynthetic carbon assimilation in isolated spinach chloroplasts. Kinetics of CO2-dependent O2 evolution in the presence of increasing quantities of exogenous Pi. The characteristic induction period is shortest in the absence of added Pi but photosynthesis then soon declines. Small increments in [Pi] lengthen the lag and either increase the rate or leave it unchanged. Larger quantities of Pi cause lag extension and rate depression. Pi values (mM) are recorded against each trace and the figures in parentheses are the rates in µmol mg-1 Chl. hr-1 (at 20oC). The broken line shows the extent of alleviation of Pi inhibition (at [Pi] of 5 mM) by the inclusion of 10 mM PPi. Conversely, if there is too much Pi in the external medium, triose phosphate which would have otherwise been used to regenerate the CO2 acceptor (RuBP), is “pulled out” of the chloroplast and the induction period (which, in less extreme circumstances, largely reflects autocatalytic build-up of cycle intermediates) is prolonged indefinitely.

Pi optimum

Externally added metabolites such as triose phosphates then reverse the Pi inhibition either by inhibiting Pi entry, or by entry instead of Pi, or both. In vivo, sucrose synthesis from triose phosphates recycles orthophosphate and there is now good evidence that there is an intimate relationship between events in the cytosol and photosynthetic carbon assimilation in the stroma of the chloroplast. In vitro, photosynthesis by isolated chloroplasts is only possible to the extent that the reaction mixture can imitate, in a limited way, the contribution that is made by the cytosol.

fructose 1, 6-bisphoshate (FBP or FDP)

Figure 41.11. Orthophosphate inhibition and its reversal. On the left, complete inhibition by 10 mM Pi. On the right, the kinetics of reversal of inhibition following the addition of 2 µmoles of DHAP, R5P, PGA and FDP (fructose 1,6-bisphosphate). 41(g) Inorganic Pyrophosphate In the history of chloroplast isolation, inorganic pyrophosphate was once an enigma and therefore deserves a mention in this context. The improvements in rate brought about by Walker (1964) still fell short of the magic 100 µmoles per milligram chlorophyll per hour which, at that time, was regarded as a convincing rate for reasonably competent chloroplasts. Rates in excess of about 25 were only rarely obtained at first and then only when catalytic amounts of Benson-Calvin cycle intermediates were added to the reaction mixtures (Appendix 3). As we have seen (in Fig. 41.7) PGA dependent oxygen evolution occurs without an appreciable lag but much smaller (i.e. catalytic quantities) of PGA or triose phosphate will also abolish induction. Rates of 100 were finally obtained by Bucke et. al., (1966) with such additions but, at about the same time, Jensen and Bassham were able to report that they could routinely obtain as high, or even higher rates, without recourse to additives. The procedures which Jensen and Bassham (1965) used were, in all essentials, exactly the same as those previously introduced by Walker (1964) except in one important detail.

That is they included inorganic pyrophosphate in the reaction mixtures. It took much longer to understand why the presence of pyrophosphate was so beneficial. Similarly, it was very difficult to understand why inorganic PPi would, in some circumstances, act like 2 molecules of orthophosphate as it does in Figure 41.9 whereas excess pyrophosphate does not inhibit in the same way as orthophosphate (Figs. 41.10 and 41.11) but will even afford a degree of protection against orthophosphate inhibition (Fig. 41.10). After a great deal of work, and a great deal of head scratching, everything became clear. Orthophosphate inhibition and its reversal by Benson-Calvin cycle intermediates on the one hand, and the shortening of induction by Benson-Calvin cycle intermediates on the other, led to the concept that there was “a direct obligatory exchange between orthophosphate (outside) and sugar phosphate (inside) phosphates (Walker and Crofts, 1970). In the hands of Heldt and Rapley (1970) this concept became the experimental reality of the “Phosphate Translocator”. The nature of orthophosphate inhibition was now clear because, if isolated chloroplasts were suspended in a medium containing greater than optimal orthophosphate, the lag or induction period would be extended - because of enforced export of internal triose phosphate. This is why isolated chloroplasts show such a sharp orthophosphate optimum. If they are offered too little Pi they become phosphate deficient, if they are offered too much Pi the autocatalytic build-up of intermediates which would otherwise terminate the induction period cannot occur, or is significantly delayed. In the presence of inorganic pyrophosphate, however, the orthophosphate optimum is greatly broadened. It is broadened for two reasons. The first is that PPi cannot, itself, cross the chloroplast envelope but can compete with orthophosphate at the phosphate translocator. For this reason it alleviates orthophosphate inhibition. The second advantage which comes with the use of PPi was remarkably fortuitous. Chloroplasts were invariably prepared at this time in media containing magnesium ions. The intact chloroplast contains an inorganic pyrophosphatase and since some chloroplasts are invariably ruptured, small amounts of pyrophosphatase are released to the external medium. The substrate for this extremely active pyrophosphatase is magnesium pyrophosphate whereas the anionic form of PPi is inhibitory. Put all this together (for a review see Edwards and Walker, 1983) and you have a situation in which inorganic pyrophosphate in the external medium acts, to an extent, like an artificial cytosol. Firstly, it ameliorates the inhibitory action of orthophosphate by competing with the phosphate translocator in much the same way as would triose phosphates and PPi in the cytosol. Secondly, the limited hydrolysis of PPi in the reaction mixtures (catalysed by inorganic pyrophosphatase released from ruptured chloroplasts) brings about cycling of orthophosphate at about the same rates as sucrose synthesis would bring about recycling of orthophosphate in the in vivo situation. It is for these reasons that, if you simply wish to obtain the best possible rates of CO2-dependent oxygen evolution with isolated chloroplasts, it is a good idea to include Mg++ and PPi in your reaction mixtures. By this simple device you will, with good chloroplasts, routinely obtain rates in excess of 100µmoles.mg-1chl.hr-1. If, for any reason, you do not wish to use PPi then you will only obtain the best rates, as a matter of course, if you happen to get the orthophosphate concentration in your reaction mixtures almost exactly right. The optimum is about 0.2 to 0.5 µmole but it does vary from preparation to preparation.

In retrospect, it is clear that the improvements in the rates of photosynthesis displayed by isolated chloroplasts which took place in the 1960’s were only due in part to the improvements in the methods of isolation. The treatment of the chloroplasts, once isolated, was equally important. Learning how to treat the chloroplasts properly proved enormously important because it not only led to the recognition that the chloroplast was an orthophosphate-consuming triose phosphate- exporting organelle but also to the elucidation of the phosphate translocator and the realisation that the cytosolic regulation via Pi has a major impact on photosynthesis. 41(h) Experiment 6. Carbon Dioxide Dependence. This, of course, is what photosynthesis is all about and it is certainly worth demonstrating and it also carries some hidden messages (Fig. 41.12). Again the mixture is as before (Experiment 2) but without added bicarbonate. There should be sufficient endogenous CO2 to allow O2 evolution to commence but, after a few minutes, the rate will approach zero. If CO2 is then added, there will be an immediate resumption of O2 evolution. If addition of CO2 is deferred, however, the rate of O2 evolution following addition of bicarbonate will be diminished. (Much the same sort of response is seen if the addition of Pi is deferred when chloroplasts are running out of phosphate as in Experiment 4, Section 41d). Anything, in fact, which limits CO2 fixation during intense illumination will bring about photoinhibitory damage.

Figure 41.12. Carbon dioxide dependence. The original experiment by Walker and Hill (1967). On the left, CO2-dependent O2 evolution. On the right the same experiment without added bicarbonate until its addition as indicated.

42. PROTOPLASTS Although one major aspect of research in photosynthesis has rested heavily on chloroplasts, there is additional information to be derived from protoplasts (Edwards and Walker, 1983). Moreover, you may wish to look at chloroplasts from species other than spinach or peas because you are interested in that particular species rather than features

of the photosynthetic process which are common to most species. If this is the case you may find that the only realistic option currently available to you is to prepare chloroplasts from protoplasts (Section 43). 42(a) Digestion. The separation of protoplasts from leaves is based on enzyme digestion of the cell walls and cellular membranes (Edwards et. al., 1976; 1978; 1979), (Huber and Edwards, 1975), (Kanai and Edwards, 1973a; 1973b; 1973c). The procedure is a compromise in much the same way as mechanical disruption is a compromise. Thus ineffective and slow digestion will give too few protoplasts to be useful in most experiments. Conversely, if very effective digestion is allowed to go on for too long, everything may be digested or, alternatively, digestion of vital components may proceed to an extent that function is damaged. Much, of course, depends on the enzymes that are used in the digestive process, the nature of the leaf, the conditions of incubation etc, etc, but the guiding principle should be to use the gentlest and shortest treatment which will give you a reasonable and active yield. 42(b) Pretreatment. Here the trick is to make the tissue as accessible to the enzymes as possible without causing undue damage in the meantime. Some leaves will readily part with their lower epidermis if this is pulled away with a pair of forceps but if you wish to become a good stripper you will need patience and nimble fingers. Alternatively you might wish to try rubbing the cuticle with carborundum or a toothbrush (an electric toothbrush has distinct potential). Cereals and spinach respond well to cutting into fine strips (and this may be best done under 0.5M sugar or sugar-alcohol solution). If you have to recourse to cutting, do not economise on razor blades because each preparation will consume several, if the best results are to be obtained. Vacuum infiltration or shaking during incubation can sometimes help but not, apparently, with slices of cereal leaves.

42(c) Enzymes and Incubation There is no one mixture which can be recommended in all circumstances. Some of the better enzymes are costly or difficult to obtain. (If price is a consideration you may also wish to recover your enzyme after use). Each new species will probably respond most readily to a particular cocktail of enzymes (e.g. Rohament P pectinase is particularly good with sunflower leaves). The following is worth trying at the outset: 2% cellulase (source-Trichoderma viride) 0.3% pectinase (Macerozyme R-10, source-Rhizopus sp.) Sorbitol (0.5 M) CaCl2 (1 mM) Buffer (20 mM) at pH 5.5 Bovine serum albumin (defatted) 0.05% - may help (For sunflower try 0.5% Rohament P instead of Macerozyme R-10 and for pea try 0.5% Macrozyme R-10 plus 0.25% Rohament P)

“a good stripper”

Figure 42.1. Isolation and Purification of Protoplasts. Starting top left: 1. Leaves are very finely sliced using an extremely sharp knife or new razor blades (42b). 2. Leaf slices are incubated with digestive enzymes. 3. Dish is rocked to dislodge protoplasts. 4. Protoplasts are decanted through a tea—strainer. 5. Leaf slices (plus those retained in the strainer) are washed with medium from a Pasteur pipette to dislodged more protoplasts. 6. Dislodged protoplasts are combined, recovered by centrifugation and resuspended in new medium by gentle shaking. 7. The resuspended protoplasts are over—layered with 0.5 M sucrose and 0.4 M sucrose + 0.1 M sorbitol resuspending media from a Pasteur pipette. 8. After centrifugation, intact protoplasts are recovered, with a Pasteur pipette, from the interface between the two over—layered media.

Incubation is best carried out in a shallow suspension under low light at 25-30oC and should not be prolonged for more than 2 to 3 hours at the most (unless a lower temperature is used). 42(d) Isolation. On returning after 2 - 3 hours or so to your digesting leaves you will find, if you are lucky, that protoplasts will start to fall out of the leaf material if the incubating dish is gently rocked from side to side Fig. 42.1. If this is not the case you must either wait a little longer or try a different mixture. If things have gone well, decant the protoplast suspension and wash additional protoplasts out of the leaf debris with a solution of sorbitol (0.5 M) and CaCl2 (1 mM) and MES (5 mM at pH 6.0). Pour through nylon mesh (a plastic tea-strainer is ideal and may be used to support finer mesh if necessary). C3 and CAM debris is best retained by mesh with 1 mm and 200 µm openings respectively. C4 bundle sheath strands are usually resistant to digestion and can be collected on sieves with 80 µm holes. The filtrate is then spun in 15 x 90 mm glass tubes at 100 g for 5 minutes and the supernatant discarded (or stored frozen in saturated NH4Cl prior to recovery of the cellulase). Do not spin longer or harder because you are now faced with the need to resuspend the protoplast pellet in a solution of 0.5M sucrose, 1.0mM CaCl2 and 5 mM MES (at pH 6.0) and this is best accomplished, without damage, by gentle agitation of a loosely packed pellet to which a few drops of this mixture has been added. 42(e) Purification Add a further 5 ml of the 0.5 M sucrose resuspending solution (42d, above) to the protoplasts in the centrifuge tubes and then, on top of this suspension, layer a further 2 ml of a second mixture containing 0.4 M sucrose, 0.1 M sorbitol and CaCl2 and MES as before. Finally the whole sandwich can be topped off with a further 1 ml of a third solution containing 0.5 M sorbitol and CaCl2 and MES as before. The tubes are now spun at 250 g for 5 minutes. The intact protoplasts rise and collect at the interface between the two upper solutions from where they can be recovered by gentle aspiration with a Pasteur pipette. If necessary, (i.e. if the protoplasts do not float or if inspection of the “chloroplast” pellet under the microscope shows it to contain lots of protoplasts), the density of the medium can be increased by increasing the concentration of the osmoticum the addition of dextran (5-10% w/v dextran T20 or T40). Once recovered from the gradient, the protoplasts should be diluted 10-fold in: Sorbitol (0.4 M) CaCl2 (1 mM) MES (20 mM) at pH 6.0 and recovered from this medium by centrifugation at 100 g for 5 minutes. A similar medium containing sucrose rather than sorbitol is good for prolonged storage.

C3 mesophyll protoplasts are often assayed in a medium somewhat similar to that used for chloroplasts (Section 41, Experiments 1-4) but containing Sorbitol (0.4 M) Tricine-KOH (5 mM) at pH 7.5, NaHCO3 (10 mM) CaCl2 (5 mM) This Ca++ is added to prevent clumping and to inhibit CO2 fixation by chloroplasts released from ruptured protoplasts. The protoplasts may be resuspended in this medium if they are to be used immediately. For C4 tissues, the procedures are similar but the crude extract is filtered through 500 µm and 80 µm sieves. The bundle sheath protoplasts are retained on the 80 µm mesh and may be resuspended in: Sorbitol (0.5 M) CaCl2 (1 mM) MES (5 mM) at pH 6.0. The mesophyll protoplasts can be purified in the same way as C3 protoplasts (above), with 10% (w/v) T20 dextran added to the flotation medium to compensate for the fact that C4 protoplasts tend to be more dense than C3 (or, ideally, by increasing the molarity of the entire gradient, e.g., to 0.6 M with respect to sucrose and sorbitol Day et. al., 1981).

43.CHLOROPLASTS FROM PROTOPLASTS One reason for preparing protoplasts is that this may be the only way, yet known, of isolating chloroplasts from certain species. For example, considerable effort was once put into the mechanical isolation of chloroplast from sunflower (Delaney and Walker, 1976) and all manner of protective agents were used to prevent damage by phenolics. Although the thylakoids and the stroma from these chloroplasts functioned well, the intact chloroplasts themselves supported CO2-dependent O2 evolution at rates which were only 10% of those displayed by those from spinach, suggesting that irreversible damage had been inflicted on the chloroplast envelope (Section 37b). Certainly, chloroplasts from sunflower protoplasts can match the performance of those from spinach. No one really knows what makes the difference but the cavitation effects produced by mechanical blenders could be a good guess. Certainly the rapid movement of a blender blade through a solution will leave a momentary partial vacuum in its wake and this together with consequent frothing seems to cause the damage by depositing a variety of deleterious compounds on to the chloroplast envelope. Conversely, the separation of chloroplasts from protoplasts is a relatively gentle process. The principle is simple. C3 protoplasts are about 30-40µm in diameter and they will therefore not pass unbroken through 20 µm holes. Accordingly a piece of nylon mesh with pores of this diameter is held over the tip of a 1ml plastic disposable syringe with a plastic collar fashioned out of a 1 ml automatic pipette tip. The syringe tip itself is usually cut short in order to widen its aperture. A protoplast suspension is then drawn into the syringe and expelled

through the mesh two or three times. If this suspension is centrifuged at 250 g for 1 min a chloroplast pellet is obtained and this can be washed and re-centrifuged if so desired. It should be noted that Edwards et. al., (1978) found a requirement for chelation in their work with isolated chloroplasts from wheat and sunflower. At pH 7.8 with 0.5 mM Pi in the assay medium sunflower chloroplasts performed best in the presence of 10 mM EDTA and 5 mM PPi. Wheat chloroplasts, like those from pea leaves (Robinson and Wiskitch, 1977; Stankovic et. al., 1977) are inhibited by PPi and stimulated by a mixture of PPi and ADP (at about 0.2 mM).

44.THE RECONSTITUTED CHLOROPLAST SYSTEM This was an invention of Whatley et. al. (1956) which was largely neglected until it was re-invented, with greatly improved activity, by Stokes and Walker (1971). Since then it has been again largely neglected but is, arguably, still a system of great potential. One of its best features is that it is often possible to obtain an active reconstituted system from less-than-perfect intact chloroplasts and its main recommendation is that it is accessible to intervention in a way which is otherwise precluded by the chloroplast envelope. In its simplest form, the reconstituted chloroplast system (Lilley and Walker, 1979) is simply intact chloroplasts which have been osmotically shocked in a reaction mixture - i.e. a mixture of thylakoids and stroma in the same proportions (though not the same relative “concentrations”) as in the parent chloroplasts. More sophisticated versions have also been used in which the thylakoid and stromal fractions have been separated and the stroma has been subjected to some degree of concentration or fractionation before reconstitution. In all cases, however, some degree of augmentation is necessary. This is because there are changes in concentration of key components such as NADP, ADP and ferredoxin as a consequence of envelope rupture. In the intact chloroplast these are held in intimate contact with the thylakoid membranes, as constituents of the stroma, by the chloroplast envelopes. In the reconstituted system the relative dilution (i.e. the concentration of each of the soluble components in the vicinity of the insoluble thylakoid membranes) is partly made good by their addition as purified, or partly purified, components. 44(a) The simplest system. This is prepared from intact, fully competent, CO2-fixing chloroplasts according to the procedures listed in Section 37 or 43. Reaction mixtures (2 ml in total) could contain, for example, H2O (0.8 ml) Chloroplasts (0.1 ml) RS1 x 2 (1.0 ml) (Section 4g) Ferredoxin (about 40-50 µg) ADP (0.4 µmoles) NADP (0.1 µmoles) Substrate (e.g. PGA, 3 µmoles) KCl (10 mM) Dithiothreitol (5 mM)

ADP

As in Experiment 1 (Section 41a) the water should be added before the chloroplasts and, to be certain, one minute should be allowed for the chloroplast to swell and burst as a result of the ensuing osmotic shock. 44(b) Systems with additional stroma. Several possibilities exist here. One is to osmotically shock a larger batch of chloroplasts in a centrifuge tube and then recover the thylakoids by centrifugation. This allows the re-addition of a much larger amount of stroma. Inevitably, however, an upper limit will eventually be reached. This limit can then be raised by using concentration-dialysis of the chloroplast extract (Lilley and Walker, 1979). Reconstitution from partially purified fractions has also been achieved (Slabas and Walker, 1976).

45. EXPERIMENTS WITH THE RECONSTITUTED SYSTEM Because of the dilution involved in preparation, the concentration of endogenous substrate is low and substrates need to be added in order to initiate a reaction. This, of course, is what gives the system its usefulness because it is possible to add substrates which would not readily penetrate the chloroplast envelope. 45(a) Experiment 1. CO2-dependent O2 evolution.

Figure 45.1. Carbon dioxide dependent oxygen evolution in the reconstituted system. O2 evolution commences on illumination at zero time and ceases when the added NADP (0.1 µmole) is reduced. It recommences on the addition of RuBP or, after a delay (see text) if a pentose monophosphate such as R5P is added.

The reaction mixture is that listed in Section 44a above, together with NaHCO3 (10 mM) and an additional substrate. The latter can be RuBP, or one of the pentose monophosphates of the Benson-Calvin cycle (Ru5P, Xu5P or R5P) as in Fig. 45.1, or FBP. If any of these is not added at the outset, O2 evolution will still commence because the system will carry out a Hill Reaction in which the added NADP will serve as the oxidant. This O2 evolution soon ceases as the NADP is reduced but can be reinstated by the addition of RuBP etc. If a pentose monophosphate is added there is a lag before O2 evolution starts and, with FBP, a larger lag. These lags can be largely abolished by the addition of creatine phosphate and its kinase which, together, constitute an artificial ATP generating system (Eqn. 45.1) suggesting that most of the lag is attributable to an unfavourable ATP/ADP ratio. CPK ADP + CP → C + ATP

...........Eqn.45.1

This concept is explored in a second experiment below 45(b) Experiment 2. PGA-dependent O2 evolution Everything is done in precisely the same way as in Experiment 1 but no bicarbonate is added. Once the initial O2 evolution has ceased, PGA (2 mM) is added and O2 evolution immediately recommences. If a small quantity (say 0.2 to 2 µmoles) of any of the following is added: ADP, Ru5P, Xu5P or R5P, glucose (plus hexokinase)

Figure 45.2. PGA dependent oxygen evolution. As for Experiment 1 (above) but with PGA as substrate. O2 evolution can be interrupted by any additive (such as R5P, see text) which will serve as an ATP sink. The interruption is lengthened (broken line, right) if the quantity of additive is increased. The interruption is eliminated (broken line, left) if CP + CPK is added as an artificial ATP generator.

O2 evolution quickly ceases (Fig. 45.2) but restarts spontaneously after an interval which is proportional to the amount of the additive. Conversely, the additives have no effect if creatine phosphate (CP) and its kinase (CPK) are also present in the mixture. The explanation of all of this lies in the reaction catalysed by PGA kinase (Eqn. 45.2). This is freely reversible but needs to be pushed in the “photosynthetic” direction (i.e. towards the formation of glycerate 1,3 bisphosphate) by a favourable ratio of reactants to product, O O C C OH OPO(OH)2 Mg2+ HCOH + ATP → HCOH + ADP CH2OPO(OH)2

CH2OPO(OH)2 PGA + ATP → GBP + ADP

....Eqn.45.2

Accordingly, the addition of ADP, or any system which consumes ATP (thereby producing ADP) such as the reaction catalysed by Ru5P kinase (Eqn. 45.3). CH2O

CH2OPO(OH)2

C

C

O + ATP →

HCOH HCOH

O

HCOH

+ ADP

HCOH

CH2OPO(OH)2

CH2OPO(OH)2

Ru5P + ATP → RuBP + ADP

.....Eqn.45.3

or hexokinase (Eqn. 45.4) glucose + ATP → glucose 6-phosphate + ADP

......Eqn.45.4

Conversely, CP + CPK, if present in sufficient quantity, will restore the ATP/ADP ratio and suppress the transitory inhibition of O2 evolution (Robinson and Walker, 1979; Carver, Hope and Walker, 1983). This is an important experiment because it illustrates the need for balance in metabolic turnover and the way in which “feed-back” (or in this case, “feed-forward”) can work. 45(c) Experiment 3. Uncoupling. We have already seen how uncoupling increases the rate of electron transport in the Hill Reaction (Section 41a, Experiment 1). With the reconstituted system we can see another facet of this situation (Fig. 45.3). The mixture is as before (Section 45a, Experiment 1, above) and when O2 evolution has ceased, following the initial reduction of NADP,

PGA is added and O2 evolution is resumed. In a sense, this is also a Hill reaction but it is the “natural” Hill reaction for which ATP is required (Eqns. 45.5 and 45.6) PGA + ATP → GBP + ADP

.......Eqn.45.5

GBP + NADPH → DHAP + NADP O

.......Eqn.45.6 O

C

C

O HCOH

+ NADP H2 → HCOH

CH2OPO(OH)2

H

+ H OPO(OH)2 + NADP CH2OPO(OH)2

2NADP + 2H2O → 2NADPH + O2

.....Eqn.45.7

For this reason, when an uncoupler is added, O2 evolution ceases. If ATP is then added, O2 evolution is at first resumed at a faster pace because the reaction then no longer depends on ATP which is generated within the system (from ADP and Pi) by photophosphorylation (Section B8) and the uncoupled rate of electron transport to NADP is faster than before (c.f., 41a Experiment 1).

Figure 45.3. Uncoupling. PGA-dependent O2 evolution, unlike the Hill Reaction (41a) or the Mehler Reaction (41c) is ATP-dependent. Adding an uncoupler (UC) therefore inhibits rather than causing an acceleration (c.f., Figs. 41.1 and 41.3). Evolution is restored by ATP but the restoration is short-lived unless CP + CPK are present or are added. This is because ADP accumulates (in the uncoupled situation) as ATP is consumed and the unfavourable ATP/ADP ratio becomes inhibitory (c.f., Fig. 45.1).

This new rate is not maintained, however, because, as the added ATP is consumed, ADP increases until the now adverse ATP/ADP ratio starts to inhibit the PGA kinase reaction (Eqn. 45.5) by mass action. If CP + CPK is added, a fast rate is again restored as this ATP regenerating system (Eqn. 45.1) restores a favourable ATP/ADP ratio.

46. SOME FURTHER PRACTICAL CONSIDERATIONS In all of the foregoing (Part F), oxygen evolution has been followed polarographically and all that is required is a single electrode vessel which lends itself to illumination (such as the Hansatech Instruments DW1 and DW2/2). Distinct advantages derive, however, from employing twin electrodes. 46(a) Twin electrodes Chloroplasts have a relatively short “bench-life” and, however active on separation, will suffer changes in activity during use. There is much to be said, therefore, for using two electrode vessels so that experiment and control can be carried out simultaneously. This means that really subtle differences can be regarded with more confidence. Secondly, oxygen electrodes are still not as reliable or stable as pH electrodes and, like any sort of apparatus, they are not immune to malfunction. “Trouble-shooting” is a great deal easier with two electrodes because components can be switched between each system until that responsible for the malfunction is identified. Inevitably, malfunction has also been known to occur during an experiment and there is a natural law (Sod’s Law, or more properly “The Conservation of Chagrin”) which will ensure that, if this does occur, it will coincide with a spinach famine. If, therefore, you have just used the last spinach, or you are leaving the next day on vacation, and only need to do one more vital experiment in order to win fame and fortune, it is best to use two electrodes for the same reason that you would probably prefer to see two engines on each wing. “you are leaving the next day on vacation”

46(b) Illumination Figure 46.1 was first published in 1972 and it illustrates a twin electrode system (Section 45a, above) in which slide projectors were used as light-sources. At the time, there was no cheap and reliable alternative and indeed even today, because of mass production, there is still a lot to be said for slide projectors if there is no particular need for anything more sophisticated. Some alternatives have already been mentioned in Part C and if an electrode system such as the Hansatech DW2/2 is used, a purpose built light-source and a fibre or liquid optic is almost mandatory. If you use fibre optics bear in mind that, even with the best, there is a significant loss of light. Similarly, do not be fooled by claims that fibre optics give “cold” light - they act as heat filters only to the extent that they do not transmit 100% of all that they are offered and in many circumstances it is wise to insert an appropriate heat filter before the optic, such as a wide band hot mirror as those supplied by O.C.L.I. (Part No 6022001) in order to protect it and to diminish what heat is transmitted. Liquid optics

transmit light more effectively than fibres but they are expensive and, at the time of writing, cannot be branched.

Figure 46.1. A twin electrode system. Two electrode vessels, each mounted on its stirrer are sited within similar compartments of a matt-black E-shaped screen (this is open above, but together with the shading offered by the mirrors and relatively subdued laboratory lighting, provides a sufficient approximation to darkness within the reaction vessel for most purposes). Intense illumination is provided by two quartz-iodine slide projectors shining through spherical, water-filled flasks which serve as additional lenses and heat filters. Further filters (see text) are sited in apertures in the screen immediately in front of the electrode vessels. These apertures are closed by a shutter with two openings which align with the holes in the screen as the shutter is pulled to the right, allowing simultaneous onset of illumination. As the shutter opens or closes it actuates a micro-switch so that this event is automatically recorded. New light sources are now coming onto the market which are based on light emitting diodes (Part C) and the brightness of LED’s continues to be improved in manufacture. The red ones are a particularly good source of photosynthetically active light, they are relatively stable, produce little heat and switch on very quickly.

REFERENCES

General References Arnon, D.I. (1967) Photosynthetic activity of isolated chloroplasts. Physiol. Rev., 47,317. Edwards, G.E., Huber, S.C. and Gutierrez, M. (1976) Photosynthetic properties of plant protoplasts. In: Microbial and Plant Protoplasts (J.F. Peberdy, A.H. Rose, H.J. Rogers and E.C. Cocking, eds). Academic Press, New York, pp 299-322. Edwards, G.E. and Walker, D.A. (1983) C3, C4, Mechanisms, and Cellular and Environmental Regulation of Photosynthesis. Blackwell Scientific Publications Ltd, Oxford. pp 1-542. Kalberer, P.P., Buchanan, B.B. and Arnon, D.I. (1967) Rates of photosynthesis by isolated chloroplasts. Proc. Nat. Acad. Sci (Wash.) 57,1542-1549. Leegood, R.C., Edwards, G.E. and Walker, D.A. (1981) Chloroplasts and Protoplasts. In: Techniques in Bioproductivity and Photosynthesis (Coombs, J and Hall, D.O. eds) Pergamon Press Oxford, pp 92-109 Lilley, R.McC., Fitzgerald, M.P., Reinits, K.G. and Walker, D.A. (1975) Criteria of intactness and the photosynthetic activity of spinach chloroplast preparations. New Phytol. 75, 1-10. Price, C.A. Bartolf, M, Ortiz, W and Reardon, E.M. (1979) Isolation of chloroplasts in silica-sol gradients. In: Methodological Surveys in Biochemistry Plant Organelles (Reid, E., ed) Vol. 9. Ellis Horwood, Chichester, pp 25-37 Robinson, S.P., Cerovic, Z.G. and Walker, D.A. (1986) Isolation of intact chloroplasts -General principles criteria of integrity. Methods in Enzymology. In press. *** must be in print now Truesdale, G.A. and Downing, A.L. (1954) Solubility of Oxygen in Water. Nature, Lond. 173, 1236. Vogel, A.I. (1961) Textbook of Quantitative Inorganic Analyses. Longman, London. pp 1-926. Walker, D.A. (1964) Improved rates of carbon dioxide fixation by illuminated chloroplasts. Biochem. J. 92, 22c-23c. Walker, D.A. (1971) Chloroplasts (and Grana) - Aqueous (including high carbon fixation ability). In: Methods in Enzymology (San Pietro, A. ed.) Vol. 23. Academic Press, New York, p 211-220. Walker, D.A. (1980) Preparation of higher plant chloroplasts. In: Methods in Enzymology (San Pietro, A. ed) Vol 69. Academic Press, New York, pp 94-104. Walker, D.A. and Crofts, A.R. (1970) Photosynthesis. Ann. Rev.Biochem. 39, 389-428. Walker, D. A. and Sivak, M.N. (1986) Photosynthesis and Phosphate: A cellular affair? Trends in Biochemical Sciences, 11, 176-179. Specific References Allen, M.B., Arnon, D.I., Capindale, J.B., Whatley, F.R. and Durham, L.J. (1955) Photosynthesis by isolated chloroplasts. III. Evidence for complete photosynthesis. J. Amer. Chem. Soc. 77, 4149-4155. Andreae, W.A. (1955) The photoinduced oxidation of manganous ions. Arch. Biochem. Biophys.55, 584-586. Baldry, C.W, Walker, D.A, Bucke C. (1966) Calvin-cycle intermediates in relation to induction phenomena in photosynthetic carbon dioxide fixation by isolated chloroplasts. Biochem J.101, 642-646 Bjorkman, O. and Demmig, B. (1987) Photon yield of O2 evolution and chlorophyll fluorescence characteristics at 77K among vascular plants of diverse origins. Planta 170,489-504.

Bruinsma, J. (1961) A comment on the spectrophotometric determination of chlorophyll. Biochim. Biophys. Acta. 52, 576-578. Bucke, C, Walker, D.A. Baldry C.W. (1966) Some effects of sugars and sugar phosphates on carbon dioxide fixation by isolated chloroplasts. Biochem J. 101,636-641 Bucke, C, Baldry, C.W., W‘alker, D.A. (1967) Photosynthetic carbon dioxide fixation by isolated chloroplasts in Good’s buffers. Phytochemistry 6, 495-497 Carver, K.A., Hope, A.B. and Walker, D.A. (1983) Adenine nucleotide status, phosphogylcerate reduction and photosynthetic phosphorylation in a reconstituted chloroplast system. Biochem J. 210, 273-276. Cerovic, Z.G. and Plesnicar, M. (1984) An improved procedure for the isolation of intact chloroplasts of high photosynthetic capacity. Biochem J. 223, 543-545. Cockburn, W., Baldry, C.W. and Walker, D.A. (1967) Oxygen evolution by isolated chloroplasts with carbon dioxide as the hydrogen acceptor. A requirement for orthophosphate or pyrophosphate. Biochim. Biophys. Acta 131, 594-596. Cockburn, W., Baldry, C.W. and Walker, D.A. (1967) Photosynthetic induction phenomena in spinach chloroplasts in relation to the nature of the isolating medium. Biochim. Biophys. Acta 143, 606-613. Cockburn, W., Baldry, C.W. and Walker, D.A. (1967) Some effects of inorganic phosphate on O2 evolution by isolated chloroplasts. Biochim. Biophys. Acta 143, 614-624. Cockburn, W., Walker, D.A. and Baldry, C.W. (1968) Photosynthesis by isolated chloroplasts. Reversal of orthophosphate inhibition by Calvin-cycle intermediates. Biochem. J. 107, 89-95. Day, D.A., Jenkins, C.L.D. and Hatch, M.D. (1981) Isolation and properties of functional mesophyll protoplasts and chloroplasts from Zea mays. Aust. J. Plant Physiol. 8, 21-29. Delaney, M.E. and Walker, D.A. (1976) A reconstituted chloroplast system from Helianthus annuus. Plant Science Letters 7, 285-294. Delieu, T. and Walker D.A. (1972) an improved cathode for the measurement of photosynthetic oxygen evolution by isolated chloroplasts. New Phytol. 71, 201-225. Delieu, T, and Walker D.A. (1981) Polarographic measurement of photosynthetic O2 evolution by leaf discs. New Phytol. 89, 165-175 Demmig, B. and Gimmler, H. (1979) Effect of divalent cations on cation fluxes across the chloroplast envelope and on photosynthesis of intact chloroplasts. Z. Naturforsch 34, 233-241. Plant Physiol. 73, 164-174. Edwards, G.E., Robinson, S.P., Tyler, N.J.C. and Walker, D.A. (1978) Photosynthesis by isolated protoplasts, protoplast extracts and chloroplasts of wheat. Influence of orthophosphate, pyrophosphate and adenylates. Plant Physiol. 62, 313-7. Edwards, G.E., Robinson, S.P., Tyler, N.J.C. and Walker, D.A. (1978) A requirement for chelation ion obtaining functional chloroplasts of sunflower and wheat. Arch. Biochem. Biophys., 190, 421-433. Edwards, G.E., Lilley, R.McC. and Hatch, M.D. (1979) Isolation of intact and functional chloroplasts from mesophyll and bundle sheath protoplasts of the C4 plant Panicum miliaceum Plant Physiol. 63,821-827. Gibbs, M. and Calo, N. (1959) Factors affecting light induced fixation of carbon dioxide by isolated spinach chloroplasts. Plant Physiol. 34, 318-323. Good, N.E., Winget, G.D., Winter, W., Conolly, T.N., Izawa, S. and Singh, R.M.M. (1966) Hydrogen ion buffers for biological research. Biochemistry 5, 467-477. Hall, D.O. (1972) Nomenclature for isolated chloroplasts. Nature New Biol. 235,125-126. Heldt, H.W. and Rapley, L. (1970) Specific transport of inorganic phosphate, 3-phosphoglycerate and dihydroxyacetonephosphate, and of dicarboxylates across the inner membrane of spinach chloroplasts. FEBS Letters 10, 143-148. Hill, R. (1937) Oxygen evolved by isolated chloroplasts. Nature Lond., 139, 881-882. Hill, R. (1939) Oxygen production by isolated chloroplasts. Proc. Roy Soc. B: 127, 192-210 Huber, S.C. and Edwards, G.E.(1975) An evaluation of some parameters required for the enzymatic isolation of cells and protoplasts with CO2 fixation capacity from C3 and C4 grasses. Physiol. Plant. 35, 203-209.

Jensen, R.G. and Bassham, J.A. (1965) Photosynthesis by isolated chloroplasts. Proc. Nat. Acad.Sci (Wash) 56, 1095-1101. Kaiser, W.M., Urbach, W. and Gimmler, H. (1980) The role of monovalent cations for photosynthesis of isolated intact chloroplasts. Planta 149, 170-175. Kanai, R. and Edwards, G. (1973) Enzymatic separation of mesophyll protoplasts and bundle sheath cells from leaves of C4 plants. Die Naturwissenschaften 60, 157-158. Kanai, R. and Edwards, G. (1973) Separation of mesophyll protoplasts and bundle sheath cells from maize leaves for photosynthetic studies. Plant Physiol. 51,1133-1137. Kanai, R. and Edwards, G. (1973) Purification of enzymatically isolated mesophyll protoplasts from C3, C4 and Crassulacean Acid Metabolism plants using an aqueous dextran polyethylene glycol two-phase system. Plant Physiol. 52, 484-490. Larkum, A.W.D. and Wyn Jones, R.G. (1979) Carbon dioxide fixation by chloroplasts isolated in glycinebetaine. A putative cytoplasmic osmoticum. Planta 145, 393-394. Leegood, R.C, and Walker D.A. (1980) Autocatalysis and light activation of enzymes in relation to photosynthetic induction in wheat chloroplasts. Arch Biochem Biophys 200, 575-582 Lilley, R.McC. and Walker, D.A. (1979) Studies with the reconstituted chloroplast system. In: Encyclopedia of Plant Physiology - Photosynthesis. (Gibbs, M. and Latzko,E. eds) Vol. II. New Series, Springer-Verlag Berlin, Heidelberg, New York, pp 41-52. Ludwig, L.J. & Whitehouse, D.G. 1970. Oxygen evolution in the dark following illumination of chloroplasts in the presence of added manganese. FEBS Letters 6, 281-284. Maury, W.J., Huber, S.C. and Moreland, D.E. (1981) Effects of Mg++ on intact chloroplasts. II Cation specificity and involvement of the envelope ATPase in (sodium) potassium/proton exchange across the envelope. Plant Physiol. 68, 1257-1263. Mehler, A.H. (1951) Studies on reactions of illuminated chloroplasts. I Mechanism of the reduction of oxygen and other Hill reagents. Arch. Biochem. Biophys. 33, 65-77 Mehler, A.H. (1951) Studies on reactions of illuminated chloroplasts. II Stimulation and inhibition of the reaction with molecular oxygen Arch. Biochem. Biophys. 34, 339-51. Mills, W.R. and Joy, K.W. (1980) A rapid method for isolation of purified, physiologically active chloroplasts, used to study the intracellular distribution of amino acids in pea leaves.Planta 148, 75-83. Morganthaler,J-J., Price,C.A., Robinson,J.M.& Gibbs,M.(1974) Photosynthetic activity of spinach chloroplasts after isopycnic centrifugation in gradients of silica. Plant Physiol. 54, 532-534 Nakatani, H.Y. and Barber, J. (1977) An improved method for isolating chloroplasts retaining their outer membranes. Biochim. Biophys. Acta 461, 510-512. Mourioux, G. and Douce, R. (1981) Slow passive diffusion of orthophosphate betwen intact isolated chloroplasts and suspending medium. Plant Physiol. 67, 470-473. Robinson, S.P. (1977) Pyrophosphate inhibition of carbon dioxide fixation in isolated pea chloroplasts by uptake in exchange endogenous adenine nucleotides.Plant Physiol.59,422-427. Robinson, S.P. (1983) Isolation of intact chloroplasts with high CO2 fixation capacity from sugarbeet leaves containing calcium oxalate. Photosynthesis Research 4, 281-7. Robinson, S.P. (1986) Improved rates of CO2-fixation by intact chloroplasts isolated in media with KCl as the osmoticum. Photosynthesis Research 10, 93-100. Robinson, S.P. and Downton, W.J.S. (1984) Potassium sodium and chloride content of isolated intact chloroplasts in relation to ionic compartmentation in leaves. Arch. Biochem. Biophys. 228, 197-206. Robinson, S.P., Edwards, G.E. and Walker, D.A. (1979) Established methods for the isolation of intact chloroplasts. In: Methodological Surveys in Biochemistry Plant Organelles (Reid,E.,ed) Vol. 9. Ellis Horwood, Chichester, pp 13-24 Robinson, S.P. and Walker, D.A. (1977) Pyrophosphate inhibition of carbon dioxide fixation inisolated pea chloroplasts by uptake in exchange for endogenous adenine nucleotides. Plant Physiol.I59, 422-427 Robinson, S.P. and Walker, D.A. (1979) The control of 3-phosphoglycerate reduction in isolated chloroplasts by the concentrations of ATP, ADP and 3-phosphoglycerate. Biochim. Biophys.Acta. 545,421-433.

Robinson, S.P. and Wiskich, J.T. (1976) Stimulation of carbon dioxide fixation in isolated pea chloroplasts by catalytic amounts of adenine nucleotides. Plant Physiol. 58, 156-162. Robinson, S.P. and Wiskich, J.T. (1977) Pyrophosphate inhibition of carbon dioxide fixation in isolated pea chloroplasts by uptake in exchange for endogenous adenine nucleotides. Plant Physiol. 65, 291-297. Sicher, R.C. (1984) Glycolaldehyde inhibition of photosynthetic carbon assimilation by isolated chloroplasts and protoplasts. In: Advances in Photosynthesis Research (Sybesma,C. ed.) Martinus Nijhoff/ Dr. W .Junk Pubs. Vol.III. p 413-416. Slabas, A.R. and Walker, D.A. (1976) Enzymic reconstitution of photosynthetic carbon assimilation. Pentose phosphate-dependent O2 evolution by illuminated envelope free chloroplasts from Spinacia oleracea Arch. Biochem. Biophys. 175 590-597. Stankovic, Z.S. and Walker, D.A. (1977) Photosynthesis by isolated pea chloroplasts. Some effects of adenylates and inorganic pyrophosphate. Plant Physiol. 59, 428-432. Stitt, M. and Heldt, H.W. (1981) Physiological rates of starch breakdown in isolated intact spinach chloroplast. Plant Physiol. 68, 755-761. Stokes, D.M. and Walker, D.A. (1971) Phosphoglycerate as a Hill oxidant in a reconstituted chloroplast system. Plant Physiol. 48, 163-165. Stokes, D.M. and Walker D.A. (1972) Photosynthesis by isolated chloroplasts. Inhibition by DL-glyceraldehyde of carbon dioxide assimilation. Biochem. J. 128,1147-1157. Stokes, D.M., Walker, D.A. and McCormick A.V. (1972) Photosynthetic oxygen evolution in a reconstituted chloroplast system. In: Progress in Photosynthesis.(Forti, G., Avron, M. and Melandri, A., eds) Proc. II Int. Cong. on Photosynthesis, Stresa 1971, W. Junk, NV Pub, The Hague p 1779-1785. Walker, D.A. (1976) CO2 fixation by intact chloroplasts: photosynthetic induction and its relation to transport phenomena and control mechanisms. In: The Intact Chloroplasts (Barber,J. ed) Chapter 7, Elsevier, Amsterdam pp 235-278. Walker, D.A. and Hill, R. (1967) The relation of oxygen evolution to carbon assimilation with isolated chloroplasts. Biochim. Biophys. Acta 131, 594-596. Walker, D.A., Baldry, C.W. and Cockburn, W. (1968) Photosynthesis by isolated chloroplasts, simultaneous measurement of carbon assimilation and oxygen evolution Plant Physiol. 43,1419-1422. Walker, D.A., Ludwig, L.J. and Whitehouse, D.G. (1970) Oxygen evolution in the dark following illumination of chloroplasts in the presence of added manganese. FEBS Letters 6, 281-284. West, K.R., and Wiskich J.T. (1968) Photosynthetic control by isolated pea chloroplasts Biochem J. 109, 527-32 Whatley, F.R., Allen, M.B., Rosenberg, L.L., Capindale, J.B. and Arnon, D.I. (1956) Photosynthesis by isolated chloroplasts. V. Phosphorylation and carbon dioxide fixation by broken chloroplasts. Biochim. Biophys. Acta 20, 462-468. Whitehouse, D.G., Ludwig, L.J. and Walker, D.A. (1971) Participation of the Mehler reaction and catalase in the oxygen exchange of chloroplast preparations.J. Exp. Botany 22, 772-791. Yamazaki, R.K. and Tolbert, N.E. (1970) Photoreactions of flavin mononucleotide and a flavoprotein with zwitterionic buffers. Biochim. Biophys. Acta. 197,90-92.

PART G

DEFECTS AND PRECAUTIONS

47(a) The Sensor. Most O2 sensors have a platinum or gold cathode which is electrically joined to an anode (usually silver) by an electrolyte (a solution containing potassium chloride). The cathode is then polarised (Part A). All of this constitutes a basically unstable situation. The best electrodes have cathodes which have been very carefully sealed into lead glass which has a coefficient of expansion very similar to that of platinum. Other electrodes are often sealed in “Araldite” or a similar epoxy resin. These are very good for what they cost but they do have a very limited life, partly because they find themselves in an unstable situation (both electrically and physically) and partly because they are mostly maltreated by their users who are usually too preoccupied with paying the rent and solving the mysteries of the universe to pay them the attention that they deserve. Epoxy resins gradually take up water. Seals between cathodes and resin gradually deteriorate. Electrolyte penetrates into crevices, crystallises and widens fissures. The underlying electrochemistry virtually ensures that cathodes will become changed by deposition of other metals, oxidised or poisoned. Contacts or leads become broken. So, if you have an electrode at all, you must resign yourself to the fact that, if it is relatively cheap it won’t last and that if it is relatively expensive you probably won’t be able to afford it. Thereafter, there are things that you can do which will help. 47(b) Use. Do not be tempted to leave a “good” electrode unchanged. You will almost certainly be too idle to change it daily but you should certainly change it weekly. Have two electrodes. At the end of a week, clean the one that has been in use and put it in a desiccator (a plastic box containing silica gel will do). Weekly cleaning should not involve more than polishing the platinum surface with aluminium oxide (polishing grade) on dampened felt or cotton wool. Set up the replacement electrode and let it polarise overnight or over the weekend. The silver may need cleaning periodically if the layer of chloride has become contaminated. With the Hansatech electrode disc, for example, this is best done by subjecting it to a reverse current. This can be done with a simple apparatus comprising an electrode (a silver rod), a 3V D.C. supply (2 x 1 1/2V batteries), 50% saturated sodium sulphate solution, and the electrode disc. The electrode disc is partially submerged in the solution (enough to cover the silver ring but not the connector). A negative (3V) potential is applied to the anode and the rod is used to complete the circuit. Hydrogen is thereby generated at the silver ring, dislodging particles and reducing the oxidised surface layer. A 2 hour treatment is generally sufficient

Even a modestly priced electrode may cost as much as a reasonable camera. It is unlikely that you dip your camera in sea-water every week and leave it to dry on the bench. You are obviously obliged to use an electrolyte but don’t leave your electrode drying out on the bench and then expect it to go on working as well as ever, month after month. 47(c) Membrane Application Use an effective applicator to apply the membrane and ensure that it is correctly in place (neither slack, crinkled nor over-stretched) and that, if it depends on an O-ring, that the O-ring is the correct size and has not perished. Use a lens or a microscope to check that you have put the membrane on correctly. If a membrane is too loose or stretched too tightly, some movement is more or less inevitable and consequent changes in the electrolyte path will lead, or contribute to, “drift” (i.e. the air-line will change, at an unacceptable rate, in an open chamber or vessel). If a “spacer” (Part A) is not used with the Hansatech electrode and if an over-tight O-ring therefore narrows the electrolyte layer, lack of linearity may be experienced. 47(d) The Leaf-disc Electrode. Because this is mostly used in saturating CO2 and measures in the gas-phase, it is important to use a modified electrolyte (e.g., one part saturated KCl solution, one part 0.4M borate buffer at pH 9.0 and two parts 1.0M sodium bicarbonate solution previously adjusted to pH 9.0 by the addition of equimolar sodium carbonate - Delieu and Walker, 1981) or to allow adequate equilibration (hydroxyl ions are generated in use and CO2 will dissolve in the electrolyte - this does not matter but it should be allowed for). 47(e) Leaks. The leaf-disc electrode was designed to measure changes in oxygen in a closed space. If the chamber leaks, it will not stop the electrode working but there will be a decrease in accuracy at best, and total malfunction at worst. The ports and taps can leak and should be checked-out individually. Start with a proven electrode disc in place and put a disposable gas-tightsyringe directly into each port (rather than the taps). Check that each port has its correct O-ring and that it is tightly threaded. Similarly, the O-ringagainst which the sensor seals should be clean, in good condition and properly positioned before the electrode is held firmly in position by tightening the base. Moving either of the syringe plungers should then cause a positive or negative excursion to a new, more or less unchanging value (Section 4). If the chamber seals, repeat the procedure with first one tap and then both taps in position. Each tap should be pushed firmly into the luer on the port before its locking-nut is tightened. Check that the large O-ring which makes the seal between the chamber itself and its upper water-jacket is of the correct size and that it is adequately compressed when the clips which pull the two parts together are tightened. 47(f) Membranes and Electrolytes There are lots of both (see e.g., Sections 2 and 47d) and they are almost as difficult to recommend, in general terms, as are plays to theatre-goers. One which has been used successfully with the

leaf-disc electrode contains: 0.2M KCl, 66mM Na2 HPO4, 0.6M KNO3 + 70mM NaOH at pH 11.2 (with HCl) (N.B. THIS IS CAUSTIC). It has the advantage of speeding stabilisation on applying a polarising voltage. Most manufacturers provide their own membranes and these are always first choices but you may find (as did Bjorkman and Demmig, 1987) that a Hansatech electrode worked best, in their hands, with a Beckman membrane (part no. 77948) and electrolyte (part no. 76438). (Our own experience has been different but the difference could be in the manner in which the membrane is applied - accordingly you should use what suits you best.) In general, thinner membranes are to be preferred (see Appendix 2) if rapidity of response is required and more robust membranes if resistance to damage is more important.

PICTURE OF AN O2 ELECTRODE There was a little girl and she had a little curl right in the middle of her forehead.

And when she was good, she was very, very good.

But when she was bad she was horrid.

References Bjorkman, O. and Demmig, B. (1987) Photon yield of O2 evolution and chlorophyll fluorescence characteristics at 77K among vascular plants of diverse origins. Planta 170 489-504. Delieu, T. and Walker, D.A. (1981) Polarographic measurement of photosynthetic oxygen evolution by leaf discs.New Phytol. 89 165-178. Osmond, C.B. (1986) Research in South-East Asia. Nature 320: 307.

Appendix 1

Preparation of Ferredoxin Preparation of Ferredoxin If you tire of electrodes and wish to remind yourself of what bucket biochemistry is like, use the spinach that you have accumulated in your glass-house deepfreeze (every good glasshouse should have one) to prepare ferredoxin. The procedure involves the preparation of a crude homogenate from spinach leaf and the use of diethylaminoethyl cellulose to absorb the ferredoxin in it. The crude ferredoxin is eluted from the cellulose, partially fractionated by ammonium sulphate precipitation and finally passed through a second DE cellulose column. At this stage of purity it is good for experiments with the reconstituted chloroplast system (Section F12). Procedure Day 1 Prepare the buffer solutions and equilibrate the DE52 cellulose. A. Prepare 2 litres of 1 M KH2PO4 at pH 7.5. Use this in the preparation of the following solutions: i) 15 litres of grinding medium (20 mM KH2PO4 at pH 7.5), ii) 1 litre of 20 mM KH2PO4/0.2 M NaCl at pH 7.5, iii) 1 litre of 20 mM KH+PO4/0.8 M NaCl at pH 7.5. iv) 10 litres of 20 mM KH2PO4 at pH 7.5 (for cellulose equilibration) B. Wash 125 g of Whatman’s DE52 cellulose with 5 litres of distilled water. This is best done by suspending the cellulose in about 2 litres of water and pouring it on to a number 1 filter paper in a large buchner funnel, using the rest of the water to wash the cellulose on the filter paper. Finally wash with 1 litre of 1 M KHPO4 at pH 7.5 followed by 10 litres of 20 mM KH2PO4 at pH 7.5. C. Resuspend the washed cellulose in about 200 ml of grinding medium, and leave this, together with all other solutions, in the refrigerator overnight. D. Leave the equipment required on Day 2 in the refrigerator overnight (homogenizer, centrifuge bottles, chromatography columns, etc). Day 2 A. Use frozen spinach from which the mid-ribs have been removed. Slice the spinach with a sharp knife and homogenize in grinding medium using about 200 ml medium for every 200 g of spinach. B. Squeeze the homogenate through a double layer of muslin into an ice-cooled conical flask.

C. Centrifuge the filtrate at 16,000 g for 10 mins. D. To the supernatant, add solid NaCl 6 g/litre and adjust the pH to 7.5 via NaOH. E. Add DE52 slurry (about 40 ml/litre) and stir briefly, then leave to stand in the cold, for at least ½ hour. F. Decant off the liquid and pour the slurry into a 3 cm (wide) column, and pack the DE52 slowly, wash the column with buffer containing 0.2 M NaCl. Do not run the column too quickly as the fine cellulose is easily dried out. Wash until the eluate is clear. G. When the eluate is clear, elute the ferredoxin with the high salt buffer. A red-brown band begins to descend, and is collected as it emerges from the column. H. Add ammonium sulphate slowly to the collected solution to give a 50% w/v solution (0.5 g ammonium sulphate to 1 ml of solution). The pH is adjusted to 7.5 with NaOH and the solution left to stand for 30 min. I. Centrifuge the suspension at 10,000 rpm for 30 min and retain the supernatant. J. The supernatant is dialysed against 4 litres of 20 mM phosphate buffer at pH 7.5 for 12 hours (overnight). Day 3 A. Pack a column (small Pharmacia K9/15) with DE52, KEEP A LOW FLOW RATE! Load the dialysate on the column, then wash with 0.2 M NaCl in buffer, then slowly elute with 0.8 M NaCl in buffer. B. Estimate the concentration by measuring the O.D. in a 2 mm light path at 420 nm. 420 O.D.

x 1.22 = mg FD ml-1

0.2 C. Store under liquid nitrogen and dialyse before use (store in high salt for long periods). N.B. All operations must be carried out in the cold. Speed is essential from homogenization until the first addition of DE52. Delay at this stage results in a poor yield of ferredoxin.

Appendix 2

Suppliers

Oxygen Electrodes. The measurements described in this manual are largely based on Hansatech Instruments equipment (Appendix 4). This is because much of the apparatus now commercially available from this firm was originally designed and constructed by myself and Tom Delieu, specifically for photosynthetic work. The “leaf-disc” electrode and related apparatus is not available elsewhere. Hansatech equipment includes electrodes, control boxes, cuvettes, light sources, fluorescence accessories, etc., computing software and this manual and is available from Hansatech Instruments Limited, Narborough Road, Pentney, Kings Lynn, Norfolk, PE32 1JL, U.K. Other aqueous-phase O2 electrodes, not specifically designed for photosynthesis, (and accessories) may be obtained, for example from:Yellow Springs Instrument Co., Box 279, Yellow Springs, OH 45387., U.S.A. (U.K. Agent: Clandon Scientific, Lysons Avenue, Ash Vale, ALDERSHOT, Hampshire, GU12 5QR). Beckman Instruments, Inc., Altex Scientific Div. 2350 Camino Ramon, San Ramon, CA 94583., USA (U.K. Agent: Analytical Instruments, Progress Road, Sands Industrial Estate, HIGH WYCOMBE, Bucks, HP12 4JL). A gas-phase electrode which can be recommended, is that supplied by Draeger Medical Ltd, Hertfordshire House, Wood Lane, Hemel Hempstead, Hertfordshire. Miscellaneous (a) Algal and Bacterial Inhibitors Boots Ltd, Nottingham, NG2 3AA, U.K. (b) Chart Recorders Fischer Scientific Co., Pittsburgh, Pa., U.S.A. Rikedenki-Mitsui, Oakcroft Road, Chessington, Surrey, KT9 1SA, U.K. (c) Dextran (T20-Avg. MW 20,000) (T20-Avg. MW 40,000) U.S. Biochemicals Corp., 21000 Miles Parkway, Cleveland, OH 44128, U.S.A. (Dextran T20 and T40) Pharmacia Fine Chemicals, Uppsala, Sweden. (Dextran T40) (d) Enzymes Cellulase from Trichoderma viride (contains cellulose and hemicellulose degrading enzyme) As Onozuka R10 and RS Yakult Biochemical Co. Ltd., Enzyme Products, 8-21 Shingikancho, Nishinomiya, Japan.

As Cellulysin Calbiochem-Behring Corp., Hoechst UK Ltd., Hoechst House, Salisbury Road, Hounslow, Middx. TW4 6JH, U.K. Calbiochem-Behring Corp., P.O. Box 12087, San Diego, CA 92112, U.S.A. As Onozuka R10 Unwin and Co. Ltd., Prospect Place, Welwyn, Hertfordshire, U.K. As Meicelase Meiji Seika Kaisha Ltd., 8-2 Chome Kyobashi, Chuo-Ku, Tokyo, Japan. Pectinase from Rhizopus sp. (contains polygalacturonase) As Macerozyme R10 Yakult Biochemical Co. Ltd. (see above) Unwin & Co. Ltd. (see above) As Macerase Calbiochem-Behring Corp. (see above) Pectinase from Aspergillus sp. As Extractase PC Fermco Biochemics, Inc., 2638 Delta Lane, Elk Grove Village, IL 60007, U.S.A. As Rohament P Rohm GmbH, D-6100 Darmstadt, Kirschenallee, Postfach 4242, West Germany. As pectolyase Y23-Pectinase from Aspergillus japonicus (active components polygalacturonase, pectin lyase and an unidentified protein factor). Seishin Pharmaceutical Co. Ltd., Noda, Chiba, Japan. (e) Glasshouse Lamps Wotan Lamps Ltd, Wotan House, 267, Merton Road, London, SW18 5JS, U.K. (f) Heat Filters OCLI (Europe), Central Way, Hill End Industrial Estate, Dunfermline, Fife, KYL 5JE, U.K. OCLI, Santa Rosa, California, U.S.A. (g) Light Meters/Quantum Sensors Skye Instruments Ltd, Unit 6, Ddole Industrial Estate, Llandrindod Wells, POWYS, Wales. LD1 6DF Li-Cor, Lincoln, Nebraska, U.S.A. (h) Light Sources Phillips Lighting Division, (U.K. Agent: Photographic Lighting U.K., Magnetic Motors Complex, Water Lane, Leeds, LS11 5PR, U.K.) Schott - U.K., Drummond Road, Astonfields Industrial Estate, Stafford, ST16 3EL, U.K. West German Agents: Schott Glaswerke, Works Wiesbaden, PO Box 130367, D-6200, Wiesbaden 13, West Germany. (i) Mass Flow Controllers Chell Instruments Ltd, Tudor House, Grammar School Road, North Walsham, Norfolk, NR28 9JH, U.K. Tylan, Carson, California, U.S.A (U.K. Agents: Epak Electronics Ltd, Pool House, Bancroft Road, Reigate, Surrey, RH27AP, U.K.) (j) Membranes Beckman Ltd. (see above) Draeger Medical Limited, Hertford House, Wood Lane Hempstead, Hertfordshire, U.K. General Electric Company, Membrane Products Operation, Medical Systems Division, 1, River Road, Schenectady, New York, N.Y. 12345, U.S.A. Hansatech Instruments Ltd. (see previous page) Yellow Springs Instrument Co. (see above)

(k) Nylon mesh Henry Simon, Ltd., P.O. Box 31, Stockport, Cheshire, SK3 ORT, U.K. Tetko, Inc., Precision Woven Screening Media, 420 Saw Mill River Road, Elmsford, NY 10523, U.S.A. (l) Optical Filters OCLI (Europe), Central Way, Hill End Industrial Estate, Dunfermline, Fife, KYL 5JE, U.K. West German Agents: Oriel GmbH, Darmstadt, West Germany Balzers, Northbridge Road, Berkhamsted, Herts. Schott - U.K., Drummond Road, Astonfields Industrial Estate, Stafford, ST16 3EL, U.K. Corning, Optical Products Department, Corning Glass Works, Corning NY 14830, U.K. (m) Optics Melles Griot, 15, South Street, Farnham, Surrey, GU9 7QU, U.K. Melles Griot, Irvine, California, U.S.A. Ealing Beck Limited, Greycaine Road, Watford, WD2 4PW, U.K. (n) Percoll Pharmacia G.B., Prince Regent Road, Hounslow, Middlesex, TW31NE U.K. (o) Polystyrene Supports Polystyrene Supports, Accelerated Prop. Ltd, Vines Cross, Heathfield, Sussex, U.K. (p) Polytron Kinematica GmbH, Ch-6000 Kriens, Luzern, Switzerland. (q) Spinach Seeds Ferry Morse Seed Company, P.O. Box 100, Mountain View, California 94042, U.S.A. Rijk Zwaan, P.O. Box 40-2678 ZG Burg, Crezeelaan 40-2678 KX De Lier, Holland. (r) Stabilised Power Supplies Clifford Industries, Amarillo, California, U.S.A. (s) Water Culture Solution I.C.I. Plant Protection Division, UK Department, Bear Lane, Farnham, Surrey, GU9 7UB, U.K.

DIFFERENTIATOR Differentiation of the O2 signal can be achieved by concentrating a 2-10 F capacitor (in series) followed by a 30 resistor (in parallel) between the output of the O2 electrode box and the input of a recorder. If such a device is used to differentiate a change of lv/sec, it will allow “change” in O2 to be recorded as “rate of change” (dO2/dt) in a mv range. Pre-amplification of the O2 signal prior to differentiation will improve the signal to noise ratio.

10µf

31kΩ

LEAF DISC

CONTENTS

INTRODUCTION

163

A. THE SOFTWARE

163

B. THE MENU

164

C. THE OPTIONS

164

1. PEN RECORDER MODE (OPTION 1) 163 (i) (ii)

Changing the Scale of the Vertical, or Y, Axis Changing the Scale of the Horizontal, or X-Axis

2. CALIBRATION (OPTION 2) (i) (ii) (iii) (iv)

168

Calibration in Outline Calibration in Detail Expanding the Vertical Axis Leaks

3. MEASURE RATE V PHOTON FLUX DENSITY (OPTION 3) 170 (i) (ii) (iii) (iv)

Manual Calibration Automated Calibration Data acquisition Bjorkman Lamp

4. OXYGEN AND FLUORESCENCE V TIME (OPTION 4) (i) (ii)

(iii)

Redraw Vertical and Horizontal Lines (a) Grid (b) Horizontal and Vertical lines The Differential (F8)

175

5. PLOT RATE V PHOTON FLUX DENSITY (OPTION 5) (i) (ii) (iii) (iv)

(v) (vi) (vii) (viii)

The relationship between rate and light The Initial Slope, By Eye The Initial Slope by Least Squares Regression Light Utilisation Capacity (a)Light Utilisation Capacity (b)Nominal Light Utilisation Capacity Determination of Rate at Different Percentage Overprint Comparing data in different directories Plotting data not previously recorded

6. EXPERIMENTAL PARAMETERS (OPTION 6) 183 A B C D E F G H I J K L M N

Programme and PFD directory DATA directory Time between AUTO steps (seconds) Rate Window (seconds) Rate Step up Size Area Cut off PFD Traces Wrap Interface Board Present Time between each stored data set Axis settings (Top, Bottom) Weighting to first part of curve in Zin fit Leaf Area in cm2 Verify data saved Real Time Differential

185

7. RETRIEVE (OPTION 7) (i) (ii) (iii)

On entry Change Differential

187

8. STORE DATA ON DISC (OPTION 8) D.ASPECTS OF ILLUMINATION

188

(a) Calibration and Uniformity (b) Transmittance and Reflection

E.DEMONSTRATION DATA

189

F. DEFAULT VALUES

189

G.PRINT OUT

189

H.REMINDERS

189

(i) Leaf Material (ii) Carbon Dioxide (iii) Absolute Calibration of O2 (iv) Capillary Matting

178

(v) Leaf pieces (vi) Pre-illumination

I. COMPUTING NOTES (i) (ii) (iii) (iv) (v) (vi) (vii) (viii) (ix) (x)

191

Software Creating Files Creating Directories Looking for Directories Delete Backup Looking for lost files Searching for a file in a directory Transferring files How best to file data

J. QUITTING

196

LEAFDISC INTRODUCTION “LeafDisc” is a computer programme written by John McAuley and George Seaton in collaboration with David Walker of the Research Institute for Photosynthesis, University of Sheffield. Now available from Hansatech Instruments Ltd (Narborough Road, Pentney, King’s Lynn, PE32 1JL) it allows the leaf-disc electrode (2,7) and associated apparatus to be used in conjunction with IBM-compatible microcomputers. Calibration is simplified and oxygen and fluorescence signals can be displayed on the computer screen rather than on a pen-recorder. Data can be stored, retrieved and displayed in many different ways. In addition the computer can be instructed to measure the rate of photosynthetic O2 evolution as a function of photon flux density (PFD). It then automatically changes the PFD according to a pre-defined programme and plots and analyses the recorded data. What follows is an aid to the computer software.

A. THE SOFTWARE The software (i.e. the computer programme) is on a disc and contains files which are required to run the programme (see Section Ii) and, for purposes of demonstration (Section E), some of the data illustrated in the following pages. Once the disc is correctly installed into the drive of the computer, (usually

drive A:\) then, on typing LD (or an appropriate batch file as in the Hansatech installation procedure) and pressing the “enter” key, the “LeafDisc” logo (see top of previous page) will be displayed. Hit any key to continue and the computer will display a “box” showing time, date, etc., and prompt you to press space bar to continue. A “menu” will now be displayed.

B. THE MENU The Menu invites you to select any one of several options. The meaning of each option will become clear as we proceed. This is what appears on the screen:SELECT OPTION 1. Pen recorder mode 2. Calibrate oxygen 3. Measure rate v photon flux density 4. Oxygen and fluorescence v time 5. Rate v photon flux density 6. Alter parameters 7. Retrieve data from disc 8. Store data on disc Select Option and Press Enter+ [x - Exit from programme] You may select any of these options by pressing numeric keys 1 to 8 followed by the key marked “enter” (or ←) on the computer or by moving the arrow (now pointing to “1”) by pressing “↓” or “↑” until it points to the desired option (followed by “enter”).

C. THE OPTIONS 1. Pen Recorder Mode (Option 1) When this mode is selected (key 1 and enter) it will display O2 (in µmoles - see calibration procedure described in option 2) on the vertical axis and time (in minutes) on the horizontal axis. If a functioning O2 electrode has been linked to the computer via a control box and interface (as in the Hansatech system), the O2 trace will now be seen advancing across the screen exactly as it would on a pen-recorder. Similarly, the position of this trace may be adjusted, at will, using the “back-off” control on the electrode control box. If the chamber contains a leaf-disc and the taps are closed, the O2 trace will fall because of dark respiration. On illumination (after a lag if the leaf has been in darkness for some time), the O2 uptake will give way to O2 evolution as photosynthesis commences (Fig. 1.1a). (The fluorescence trace will be displayed at, or near, the top of the screen at the moment that illumination is commenced and

will then fall, sometimes displaying secondary kinetics). Like the O2 signal, the fluorescence signal will “wrap round” i.e., instead of going off-scale at the bottom of the screen it will reappear at the top of the screen or vice versa (Fig. 1.1c). Either at this time, or subsequently (Option 4) the scale of the horizontal and vertical axes can be changed by using the function keys described below and as listed in Table 1.1 below. Such changes are retained in your exit from the programme via X (see Section J). (i) Changing the Scale of the Vertical, or Y, Axis Change by pressing Function Key F2 followed by F2 for 20 µmoles (of oxygen) full scale, F3 for 10 µmoles full scale, etc (see Table 1 for possible options which appear at the foot of the screen when F2 is pressed) (ii) Changing the Scale of the Horizontal, or X-Axis Change by pressing F3 followed by F2 for time intervals of 0.5min, F3 for time intervals of 1 min, etc., (see Table 1 for options displayed at foot of screen when F3 is pressed). (iii) Fluorescence Expand or contract the fluorescence trace by pressing F7 followed by one of the function keys corresponding to options (x20, x10, etc) which are then displayed at the foot of the screen (see Table 1). The fluorescence signal may also be altered by adjusting the gain on the fluorescence control box and its position on the screen may be altered by adjusting the adjacent “back-off” control.

TABLE 1.1 SCREEN DISPLAYS ON PRESSING FUNCTION KEYS Time

1 menu

2 0.50

3 1.00

4 1.50

5 2.00

6 2.50

7 3.00

8 3.50

9 4.00

10 4.50

F3

O2

1 menu

2 20µM

3 10µM

4 5µM

5 2µM

6 1µM

7

8

9

10

F2

Fl

1 menu

2 x20

3 x10

4 x5

5 x2

6 x1

7

8

9

10

F7

PFD

1 menu

2 max

3 75%

4 50%

5 25%

6 10%

7 5%

8 OFF

9

10

F6

Table 1.1 The above values (centre panel) are displayed at the foot of the screen in Option 1 when F3 (for time), F2 (for O2), F7 (for fluorescence) or F8 (for PFD) are pressed. The values underlined are those currently selected (the default values) and the underlining will move according to changes made. For example, if you wish to change the time interval from the default value of 2 minutes (underlined) you first press F3 to bring up the alternatives shown and then, if you require a 0.5 min interval,press F2. Similarly, to expand the fluorescence signal 10-fold, press F7 (so that the fluorescence option shows x1 underlined)and change to x10,by pressing F3. To switch from darkness to 50% PFD press F8 followed by F4. To return to menu, press F1.

Figure 1.1

Fig 1.1 Data, in file JA1, following manipulation in various ways a) After acquiring data in Option 1, traces such as this will be displayed. They may also be re-drawn (as here) using Option 4, to allow further examination and manipulation of the data. This trace shows a time-course of photosynthetic oxygen evolution and the corresponding changes in chlorophyll fluorescence. The vertical axis is calibrated in µmoles of O2 (1-10), the horizontal axis in minutes (2 to 20). Oxygen evolution from a leaf taken after a short dark interval (as here) commences initially after a very brief lag or induction period. Following a longer dark interval, during which evolution gives way to respiratory uptake, it recommences slightly less rapidly, because of the re-establishment of induction. Note that the rise in fluorescence upon illumination is too rapid for the computer to follow and that the initial fluorescence was greater than the ceiling imposed by the setting of the fluorescence meter (i.e., insufficient back-off). Thereafter oscillations in fluorescence corresponded (in a broadly antiparallel way) to oscillations in O2 evolution. The words and numbers at the bottom of the screen (see also Fig. 3.1) refer to the function keys and what they do. Thus F1 is to return to the menu, F3 to change the horizontal axis (see (b) below), F2 to change the vertical axis (see (c) below) F5 to skip to what is “next” if the trace has gone off screen to the right, F4 to “return” from what is “next” (if anything) to what can be seen now, F7 to expand or contract the fluorescence signal, F8 to differentiate the O2 signal (see 4(iii)), F9 to introduce a grid (as in b). (b) As for (a) but with the horizontal axis expanded by using function key F3 as described in the text. The fluorescence signal has also been amplified by the computer, using F7, so that the expanded trace “wraps around” i.e., it goes off-scale at the bottom of the screen at one point only to re-appear at the top. Such scale expansions can be made before or during recording (Option 1), subsequently (Option 4), or following storage (Option 8). In the latter case use “recall” (Option 7) then Option 4. A grid has also been introduced by pressing F9 and the commands obliterated by pressing |←. (c) As for (a) but with both axes expanded by using F3 and F2 for the horizontal and vertical axes respectively (see Table1). The fluorescence trace has also been amplified as in (b) by using F7 (see Table 1). (d) As for (b) but without the grid. Function key 8 has also been pressed and the differentiator parameters (see 4(iii)) appear one after another each time “enter” is pressed. Before acceptance and entry each parameter can be changed (by “overtyping”). (e) As for (d) after “enter” has been pressed for the third time. The box has gone and the differential of the O2 signal has appeared. Three (broken) horizontal lines have been introduced by pressing “↓”. The horizontal lines have been moved by pressing “↓” or “↑” to obtain readings of the dark uptake rates immediately prior to illumination and the maximum and minimum rates of O2 evolution during the oscillations. Note the initial burst of O2 which is apparent in the differentiated trace but is not so readily detectable in the undifferentiated trace. (f) As for (e) but following time scale expansion. A (broken) vertical line has been introduced by pressing “↓” and moved from the centre by pressing “←” or “→”. When judged to be coincident with peaks or troughs in O2 or fluorescence, they were anchored by pressing “Ins”. The times now displayed show the phase shifts (3,15,8 and 13s respectively) between O2 and fluorescence at different times during the experiment.

“skip to what is next”

(iv) Taking the Rate After a few seconds have elapsed, and at any subsequent time, the rate of O2 change can be displayed by pressing F4. The rate is expressed in moles.m-2.s-1. The rate displayed is the change in O2 recorded over the preceding 50s but, if a different “time window” is preferred, the “default” value in option 6D can be changed. For example it can be changed to 30s by pressing d, typing 30 and pressing enter. Note that this part of the programme is also used for automatic measurements in (Option 3) and that, for this purpose, the autostop value (see Section C6c) must be at least as long as the “time window”.

C 2. Calibration (Option 2) (i) Calibration in Outline “rate displayed”

The full details of the calibration display are given below, for completeness. They look forbidding. In actuality, however, the calibration procedure is very simple. In brief (see also Section 4), you are offered two horizontals. You adjust the O2 trace to coincide with the lower horizontal using the “BACK OFF” control on the electrode “box”. Then you inject a volume of air (usually 1 ml), which produces an excursion, and adjust the O2 trace to coincide with the upper horizontal using the “GAIN” control (Fig. 2.1). This is repeated until the excursion moves exactly between the two horizontals. Note that, for accurate measurements, the chamber should be re-calibrated each time a new leaf-disc is used and that this should be done with the leaf in the chamber. (ii) Calibration in Detail Select “Option 2” by pressing key 2 followed by enter. The screen will now display a vertical axis (marked 1 to 10 which, after calibration, will register moles of oxygen) and a horizontal axis (marked 2 to 20, which is a time scale, reading in minutes). To the right of the screen, the following statements will appear if the parameters have not been previously changed. Air temperature (ºC) [20.00]: If this is the temperature at which you are working, press enter. (If not, type in the correct temperature and press enter). The screen will now display: Air Temperature (ºC) [20.00]: ADConv Range(µmoles O2) [10.00]: If you do not wish to change the vertical screen range, press enter again. The screen will now display: Air Temperature (ºC) [20.00]: ADConvRange(µmoles O2)[10.00]: Air to be injected (ml)[1.00] Press enter again. If you have changed neither the temperature nor the vertical scale, the screen will display:

Air Temperature (ºC) [20.00]: ADConv Range (µmoles O2) [10.00]: Air to be injected (ml) [1.00]... back off until trace is at 0.5 inject 1.00ml.of air and... adjust gain until trace is at 9.2 repeat as necessary. The screen will also display two horizontals (Fig. 2.1), the upper at 9.2 and the lower at 0.5. Proceed as the screen invites you to by adjusting the O2 reading (which by now will be advancing across the screen from left to right) until it coincides with the lower horizontal using the “BACK OFF” control. Inject 1 ml of air from a gas-tight syringe connected to one of the taps provided for this purpose. An excursion will occur. When this is complete, adjust the “GAIN” until the O2 trace coincides with the upper horizontal. Now withdraw the 1 ml of air (using the syringe which is left in place during this procedure) and the trace should return to the lower horizontal. If it does not, continue to inject and remove air until the excursion moves between the two horizontals. (Remember to open the tap to which you attach the syringe and close the other!)

Figure 2.1. Calibration. The screen displays two horizontals. In this example the O2 value was “backed -off” (left) until it coincided with the lower (0.5) horizontal. One ml of air was then injected into the chamber causing an excursion which, in this case, went above the upper (9.2) horizontal. The trace was now lowered (using “Gain” on the electrode control unit) until it coincided with the upper horizontal and the 1 ml of air withdrawn and reinserted to confirm that the excursions were now of the correct magnitude. Note that the scale of the horizontal axis has been changed by pressing F3 followed by F3 (Section C). (iii) Expanding the Vertical Axis If you wish to expand the vertical axis (e.g. if you wish to display a screen range of 5 µmol), type in “5” when “10” is first displayed and then proceed as invited. If you now return to the menu and another option, the vertical scale will read, correctly, from 1 to 5 instead of from 1 to 10.

(iv) Leaks initial rise will be followed by a rapid fall. A slow leak can be distinguished from other declining signals (such as the initial fall which is always seen when the electrode is first switched on) by withdrawing air from the chamber. If you adjust the backoff to the 9.2 line and withdraw 1 ml of air then air leaking into the chamber in these circumstances will manifest itself as an increasing signal.

C3. Measure Rate V Photon Flux Density (Option 3) This part of the programme has been designed to allow O2 evolution to be measured over a range of different light intensities (photon flux densities) [see Section K]. a slow leak and a fast potato Select Option 3 by keying 3 followed by enter. The screen then displays the sub menu: OPTION 3 - SUB MENU 1. Load/edit PFD information 2. Do RATE v PFD data acquisition 3. Calibrate light meter Quit (back to menu)

(i) Manual Calibration Prior to measurement of quantum yield, the computer needs to know which PFDs to use (up to 40). The computer directly controls the LED output but before it can do this it must be told the LED value (within the range 0-4095) which gives the required PFD. To do this, press Option 1 (in the sub menu) followed by enter. You will be presented with a table (Table 3.1) to complete. Values may be typed in to the table by moving from line to line and column to column using the “→ ← ↓ ↑“ keys (followed by enter) and typing the LED number (followed by enter) and the corresponding PFD value (followed by enter) on each line. For each LED value used, a PFD value must be measured. This is done by applying a suitable quantum sensor, such as the Skye instrument provided by Hansatech Instruments for this purpose, to the window against which the leaf disc is pressed when the chamber is in use, and reading the PFD displayed each time a new LED number is selected or altered. At the foot of each table, additional options are offered (Table 3.1). Once a range of PFDs has been compiled it can be saved to disc using F2 (Save). To recall these values (or any other values which have been previously stored), press F1 (Dir). The screen will display a directory of previously stored PFD tables, (sets of LED values, and corresponding PFDs as in Table 3.1). To load one of the tables, press any key followed by F3. Now type the name of the PFD table, followed by enter. F3 can also be used to create a new table. This can be done by completing the empty table which is offered when the “Leafdisc” programme is first activated, or by changing values in an existing table by over-typing new values. The number of points can be increased or decreased by “Ins” and “Del” respectively.

TABLE 3.1 A typical PFD Table. LED[1] # 1 2 3 4 5 6 7 8 9 10 11 12 13 14 15 16 17 18 19 20

LED (O-4095) 0 39 74 106 138 170 200 260 317 375 460 545 630 750 980 1350 1800 2470 3250 4090

[PFD File Name: PVL20] PFD (M/M 2/s) 0.00 5.53 10.88 15.96 21.30 26.70 31.90 42.70 53.40 64.30 80.10 96.30 112.70 135.00 181.00 257.00 353.00 499.00 680.00 886.00

F1-Dir F2-Save F3-Load F9-Auto Ins-Insert Del-Delete Esc-Done F10-Quit Before entries have been made, both the second and third columns display zero values. Following entry, the second column shows the selected LED number (0-4095) and the third column the corresponding PFD value derived by photon sensor measurement. The additional options at the foot of the table are explained on the previous page and below.

Once compiled (or changed) a table can be saved by pressing F2. Its name will then appear in the directory which is summoned by F1. F9 is used in automated calibration (below), Ins for inserting additional values and Del for deleting. To continue with the experiment press ESC. F10 returns you to the sub-menu of Option 3. (ii) Automated Calibration Although manual calibration is satisfactory it is very time consuming. A Skye photon-counter (available from Hansatech Instruments) may be directly linked to the computer. This permits the sensor to be substituted for the leaf-disc during what would otherwise be the chosen period of data acquisition. Accordingly the sensor not only “sees” the same range of PFD’s as the leaf disc would experience but continues to record them for precisely the same time intervals. This facilitates calibration and increases its accuracy, accommodating as it does, any slight changes in PFD due to changes in temperature of the LED’s, ageing of the LED’s etc. Automated calibration is achieved by selecting Option 3, sub-option 3 (i.e. “calibrate light meter”). Table 3.2 is displayed on pressing enter. In Table 3.2, the default value for “calibration period” is shown as 50s but this will otherwise agree with the value for “RATE WINDOW” (D, in Option 6) and may be altered there as desired.

Table 3.2a Automated calibration of light meter Calibration of Light Meter 1. The light meter must be connected to channel 2 and the apparatus set up to measure the PFD. 2. The LED box should be set to COMPUTER. Calibration period = 50.00 seconds Light meter reading:

Normal practice would now be to enter 0 after “Light meter reading” (assuming that the light source is set to 0 and the quantum sensor is reading 0). On pressing enter the screen will show:

Light Meter Reading: 0

Time: 50.00

Ave ADC: 0.00

and the time displayed will immediately start to decrease towards zero. When it reaches 0, the screen will display: Light Meter Reading: 0 Light Meter Reading:

Time: 0.00

Ave ADC: 0.00

inviting you to enter the new quantum photon reading, e.g. 900. (The LEDs will come on automatically at this moment if the light source control box is switched to “computer”). Light Meter Reading: 0 Time: 0.00 Light Meter Reading: 900

Ave ADC: 0.00

On pressing enter again, the screen will sample ADC numbers (the output from the quantum sensor) as before. Table 3.2b shows the outcome when the computer is not connected to the apparatus and therefore fails to access the output from the quantum sensor. When it is connected and it has derived ADC numbers for zero PFD and for the highest PFD that the LED source will provide, all that is now necessary is to press “Esc” and you will be offered a blank PFD table. At this stage you must type in the LED values you wish to use. Now press F9 (Auto) and enter. The computer will now proceed to accept the quantum sensor readings and

Table 3.2b Automated calibration of light meter Calibration of Light Meter 1. The light meter must be connected to channel 2 and the apparatus set up to measure the PFD. 2. The LED box should be set to COMPUTER. Calibration period = 50.00 seconds Light Meter Reading: 0 Time: 0.00 Light Meter Reading: 900 Time: 0.00 Error - light meter signal too small

Ave ADC: 0.00 Ave ADC: 0.00

Press R to redo or Esc to end

move through the series of LED values as required, recording the PFD values as it goes. At present, the software requires the LED numbers to be pre-selected rather than having the facility to arrive at pre-selected PFD values by changing the LED numbers to give the required light output. Once an appropriate range of LED numbers has been determined manually, however, the automated procedure will repeatedly measure the PFD’s over the range without further intervention by the operator, provided that the numbers are stored and recalled according to Option 3 sub-option 1. Manual determination of LED numbers is done by entering numbers from 0 to 4095 into a blank LED table and changing these until the desired PFDs are recorded by the quantum sensor. (iii) Data acquisition You should now proceed as before (7) - i.e., place a leaf-disc in the chamber, calibrate, charge with CO2, close the taps, and adjust the oxygen trace to a convenient position near the foot of the screen (say 60 - 100 on the digital display of the oxygen electrode box by adjusting the back-off on this box). To initiate the automatic sequence of PFD values press F8. The computer will “bleep”, thereby acknowledging this instruction. It will then continue to record oxygen exchange for a pre-determined number of time intervals. The default value for time intervals is 60 seconds but this may be changed (e.g. to 30s) by using Option 6. At the end of each time interval, the computer will automatically increase the photon flux density and record the rate of O2 exchange over the previous 50s. (This value of 50s is the default value on the computer and can be changed using Option 6D). Note that 6C and 6D should be compatible, like the default values of 60s and 50s. You can change this to (say) 30s and 20s, in which case the duration of illumination at each PFD will be 30s and the period of measurement will be the last 20s of this 30s of illumination. Do not forget, of course, that you cannot require the computer to measure over 20s if you also ask it to change PFD’s every 10s. Figure 3.1 shows the appearance of the screen during measurements. At the end of the experiment, illumination ceases automatically and you can return to

“charge with CO2”

Figure 3.1. Screen display of “raw” data (acquired by using Option 3) as it would appear during an experiment or following retrieval from memory (see also table 5.1). The computer lifts the trace vertically upwards on the screen, each time the PFD is increased, as an aid to clarifying presentation. Function keys F3 and F2 are used to change the horizontal and vertical scales as before (see e.g. Table 1.1 and Fig. 1.1), F4 to take a rate manually, F5 to repeat a measurement just made, F6 to skip forward (i.e. to go directly to the next highest PFD without measurement), F7 to skip backwards and F8 to initiate automatic changes in PFD. the menu by pressing F1. You may then store this data immediately (Option 8), re-draw it (Option 4), or plot it as as rate against photon flux density (Option 5). Storage is preferred at this stage (i.e., prior to any other manipulations) in order to avoid loss. Option 6M offers a safeguard (select as T) against inadvertent loss by the user. (iv) Björkman Lamp (LS2 Light Source)

“skip backwards”

Changes in PFD can also be accomplished by using an alternative light source such as the Björkman Lamp (see ref.2 & section 5) together with neutral density filters. To do this with the present programme, change parameter O in Option 6 (see section 6) from “false” to “true”. Then, on entering Option 3 (2) the screen will display the following. Filters[1] Filters 1 ------2 ------3 ------4 ------5 ------6 ------7 ------8 ------9 ------and etc up to 16 or 20 filters

PFD 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00 0.00

(µM/M2/s)

The normal value for the neutral density filters should be entered in the first vacant column and the actual (measured) PFD values in the second. This PFD table can be saved (F2) in the normal fashion. You should now proceed as you would with the LED source using either manual or automated modes. If you use the automated mode (F8) you may, of course, wish to change the timing (Option 6) so that you have time to change a filter before the computer starts to record the rate. If you retain the existing default values (in Option 6), you will only have 10s to change the filter before recording commences (since the first 10s out of the 60s of “time between autostops” is neglected if the “rate window” is unchanged at 50s).

C 4. Oxygen and Fluorescence v Time (Option 4) (i) Redraw Press key 4 and enter. This allows data recorded in Options 1 and 3 to be “re-drawn” so that the scale of the horizontal and vertical axes can be retrospectively altered to what is most appropriate. Examples of the use of this facility are illustrated in Fig. 1.1. Change vertical (Y axis) by pressing F2 followed by F2 for 20 µmoles (of oxygen) full scale, F3 for 10 µmoles full scale etc., (see possible options at foot of screen listed in Table 1.1). To change horizontal (X axis), press F3 followed by F2 for time intervals of 0.5 min, F3 for time intervals of 1.0 minute etc., (see options displayed at foot of screen and listed Table 1.1). (ii) Vertical and Horizontal Lines These are offered in different ways:(a) Grid The simplest (Fig 1.1b) is a grid (F9) which introduces horizontal and vertical lines in a fixed relationship to one another so that inter-relationships displayed on the screen can be more readily related to one another by eye. (b) Horizontal and Vertical Lines These may be used to measure the line interval between events such as peaks and troughs in fluorescence or oxygen in Option 4. To start, simply press “←” or “→” and broken vertical and horizontal lines will appear in the centre of the screen. The vertical line may be displaced to the left or right by pressing “←” or “→” respectively. As the line is moved, it indicates its position in elapsed time (e.g. if moved to the left until it coincides with the Y-axis, it will register zero seconds). Pressing the “Ins” key will anchor the horizontal line and display the elapsed time again on the axis. If you press “Esc” you will delete the horizontal line and the vertical line at the point of the elapsed time. The whole process can then be re-initiated by pressing “←” or “→”. When this is done the new lines may be moved with the arrow keys without removing previously summoned lines. The use of the vertical and horizontal lines illustrated in Fig. 1.1 (e,f). The horizontal line is moved by pressing “↑” or “↓”. It registers O2 in µmoles and the differential of the O2 trace in µmoles s-1. This is equivalent to µmoles.m-2.s-1 for a disc of 10cm2 area.

(iii) The Differential (F8) A “box” will appear, mid-screen when you first press F8, displaying:Differentiator parameters Time Window (secs) [50]: The “window” refers to the time span, in seconds, over which the rate

Vertical Lines

Horizontal Lines

Figure 5.1 Data recorded using Option 3, illustrating a few of the range of analytical procedures which may be subsequently applied. (File QJA1)

Figure 5.1 Data acquired in Option 3 (File QJA1) (a) Rate v PFD plot for data acquired using Option 3. Such data can be plotted immediately after the experiment has been completed or following storage (Option 8) or after recovery from previous storage (Option 7). In this and what follows changes have been made to the axes (see Table 6) using the sub-option available in Option 5. (b) As for (a) but with points joined (see Table 6). The PFDs required to give 50% and 90% of the max rate (at 800 µmole.quanta.m-2.s-1) have been displayed by pressing F5 and typing in 50 (or 90) followed by “enter” when prompted by an invitation, which appears at the top of the screen, to “Enter % of max rate”. The initial slope, as judged by eye, is arrived at by pressing F7 (to obtain a line) and the direction arrow keys are pressed until the gradient of the line appears to coincide with the initial slope. Values of the intercepts and the quantum yield and requirement are automatically displayed at the top of the screen. Pressing F7 again has fixed the line. Pressing ← has eliminated the commands. (c) As for (b) but without the 50% and 90% values and with a grid introduced by pressing F9. This time the initial slope was determined by least squares regression (F8). F4 twice then erects a vertical at 800 µmole.quanta.m-2.s-1 and draws a horizontal from the intercept of this vertical and the curve to the extrapolated initial slope. The area under the curve is then expressed as a percentage of the area defined by these constraints (see Light Utilisation Capacity - 5(iv)). (d) As for (c) but without the grid and with a nominal initial slope equivalent to a quantum requirement of 9 introduced by (and fixed by) pressing F7 twice. As in (c), now pressing F4 twice produces a vertical, etc. and gives the area under the curve as a percentage of the area defined by these constraints (see Nominal Light Utilisation Capacity - 5(iv)). (e) Is an expansion of (c) (derived by using F2 and F3). The points are not joined because “N” was preferred in sub-option 5 (see Table 6) “Join up the points? (Y/N)” which is asked when the data is first displayed in numerical form (Table 4) in Option 5 prior to plotting. A least square regression on points 2 to 6 was then applied by using F8. (f) Two sets of data are displayed simultaneously by the overplot facility. This was done by plotting the lower curve WASHQ3 using Option 5 and then (after adjusting the vertical scale) using F6 and typing the file name WASHQ6 (followed by enter) in answer to the question. “File title for overplot”. Now F7 is pressed to introduce a line on screen which is fixed in position by pressing F7 again. The area is now found by pressing the F4 key twice which introduces the vertical and the lower of the two horizontals. The second horizontal is introduced by pressing and using to lift the new horizontal so introduced to coincide with the intercept of WASHQ6 with the vertical. It is fixed in position by keying F4 again. This gives a value of 48% for the area under the curve and allows WASHQ3 to be compared in these terms with WASHQ6 (at 79%) - i.e. the nominal light utilisation capacity in this experiment had increased from 48% to 79% (both measurements were made on the same disc, before and after treatment).

is measured i.e., “50s” means that each point on the differential trace will be based on the change in O2 during the preceding 50s. The smaller the window, the more responsive the differential but small windows carry the penalty of increased “noise”. (The longer the period of measurement, the “smoother” the averaged value because minor perturbations caused by electrical interference etc. will be averaged out). If you wish to retain the 50s window, press enter. Alternatively, type in a new time, say 25 or 100s. In either case, the window will now display the following in its original or amended form: Differentiator parameters Time Window (secs) [50]: Screen Range (nanoM/s) [20.00]: The scale may be accepted, or changed, by keying in a new value. Pressing enter then gives the third line as follows: Differentiator parameters Time Window (secs) [50]: Screen Range (nanoM/s) [20.00]: Zero Value (nanoM/s) [-10.00]: Again this may be either accepted (press enter) or modified (key in new values, e.g., zero, followed by enter). Pressing enter for the third time will cause the box to disappear and the differential to appear on the screen in its place. The differential scale will also appear on the right vertical axis. It is calibrated, in nmoles.s-1. Accordingly, for a 10 cm2, disc a value of 10 nmoles.s-1 will be equivalent to a rate of 10 moles.m-2.s-1 i.e. 10 nmoles per 10cm2 per sec (or 10 nmoles 10cm-2.s-1) = 1 nmole.cm-2.s-1 = 10,000 nmoles.m-2.s-1 = 10 µmoles.m-2.s-1 For the general case, divide the differential reading by the area of the leaf (in square centimetres), multiply by 10,000 (to convert cm-2 to m-2) and divide by 1,000 (to convert nmoles to µmoles).

C 5. Plot Rate v Photon Flux Density (Option 5) (i) The Relationship Between Rate and Light This is used to plot data recorded in Option 3 (see Fig. 3.1) or such data retrieved from storage (Option 7). Suppose, for example, you retrieve data from storage by using F7 and typing in (e.g.) QST1, followed by enter. The screen display will be as in Table 5.1a4. If you do not wish to see the trace data, press enter again and you will find yourself in Option 5. Alternatively, answer Y and, on pressing enter, you will be referred to Option 4 and, if you continue in Option 4 by pressing enter, the screen will display your “raw” data in the form illustrated in Fig. 3.1. This data can also be plotted graphically (Fig. 5.1 a-f) in Option 5 but, whether or not you go to 5 directly (or via Option 4), what you will next see is illustrated in Table 5.2.

If you now press F3 followed by F2, the computer will choose the most appropriate scale to accommodate the maximum rate and plot rate (on the vertical axis) against photon flux density (on the horizontal axis). If you wish to alter the vertical axis, press F3 and enter a new oxygen rate. If you wish to change the horizontal axis, press F2 and enter a new PFD value. Expansion of both scales is useful if you wish to obtain an accurate value for the light compensation point (the point at which “dark” respiration and photosynthesis are in balance) or if you wish to examine the Kok effect (changes in linearity below the light compensation point, Ref 4). If you wish to add to the description of the experiment in the second box, use F4 (Comments). F5 (Graph) gives you access to what is illustrated in Table 5.3 and allows you, by overtyping, to change what is listed. Fig (5.2a,b) shows a print-out of data in file QST1 before and after changes of this sort and after over-printing with QST2 data. (ii) The Initial Slope, By Eye To determine the initial slope, press function key 7. A line will be displayed on the screen (see Fig 5.1b). This can be moved in any direction by using the direction arrows (“← → ↓ ↑“) provided that “shift” is pressed simultaneously. If the shift key is not depressed, the line remains anchored to the dark value and can be pivoted about this point and extended by using the direction arrows. Quantum requirement and quantum yield will be automatically displayed at the top of the screen throughout this process. Once the best fit has been obtained, press any function key prior to printing (see Figs. 5.1b and 5.2a). (iii) The Initial Slope By Least Squares Regression The initial slope (and therefore the quantum requirement) can also be determined by least squares regression. Press F8. Enter the points selected separated by a comma e.g. 2,5. Press enter. The X and Y intercepts, the quantum yield and the quantum requirement will now be displayed (see Figs. 5.1c, 5.1e and 5.2a). Figure 5.1 (b,c and e) shows rate v PFD data from darkness up to PFDs which are approaching light saturation for C3 species. If, however, only the initial slope of this relationship is required, greater accuracy can be achieved by taking more readings in the 0-125 µmole.quanta.m2.s-1 range. Figure 7.1 shows such data derived by using appropriate PFD values and a diffusing plate inserted below the LED source. “once the best fit has been obtained” (iv) Light Utilisation Capacity This is an assessment of light utilisation capacity based on the following concept. Let us suppose that a leaf is exposed to a range of PFD’s at constant temperature and constant CO2 concentration, let us also suppose that carbon dioxide is saturating but not deleterious. At first, photosynthesis will be restrained by the quantum yield. It is this which defines the initial slope (the roof) of the rate against light intensity relationship. The constraint is a thermodynamic one. For example, if we accept the Z-scheme for photosynthesis there is a requirement for 8 quanta per 4 electrons transported from water to NADP. In this context, 8 is an irreducible minimum. Similarly, there must be a maximum (ceiling) above which electrons cannot be transported to NADP and beyond at a faster rate under given conditions. This function defines the approach to these maxima in two ways. (a) Actual Light Utilisation Capacity. The constraints are defined by the initial slope of the rate v PFD plot and its extrapolation to intersect with a horizontal (ceiling) drawn through the rate of O2 evolution at 800 µmole.quanta.m-2.s-1 . These are via least squares regression (F8) and then F4 and return. The area under the Rate v PFD curve is then expressed as a percentage of the area beneath the initial slope and the horizontal ceiling (see Fig. 5.1c).

(b) Nominal Light Utilisation Capacity. The maximum is defined by constraints in the same way as in (iv) but the initial slope is a nominal value (i.e a quantum yield of 0.111). Press F4 and return. The area under the curve as a percentage of the area defined by the constraints (roof and ceiling) is then displayed as before. The horizontal ceiling (Fig.5.1d) can also be moved by using keys “ “. [N.B. Nominal Light Utilisation capacity will only work in this way if F7 has been used previously. If not, press F7 twice prior to F4 and return.] Table 5.1 File information : 13:01:04 proving run on spinach. As for st1 but Ratev PFD on PV120 Number of data sets :0 (0.0 minutes long.) Point

Rate

60 116 172 228 284 340 396 452 508 564 620 676 732 788 844 900 956 1012 1068 1124

-2.00 -1.43 -0.94 -0.29 0.08 0.54 1.07 2.16 3.07 4.47 5.64 7.18 8.91 10.42 12.06 14.23 16.03 17.13 17.40 l8.42

Light (rate in µM/Msq/s Light in µE.Msq/ 0.16 5.68 10.63 16.21 21.27 26.36 32.01 41.60 52.12 66.98 82.16 102.11 126.57 152.95 196.56 268.41 357.70 495.37 663.34 854.30

Load trace data as well ? (N) :

Table 5.1 Numerical data derived in Option 3 as it first appears on screen following retrieval from memory. If enter is pressed at this stage the screen display changes to that shown in Table 5.2 (the default answer to the question at the foot of the table is N). If Y is pressed, the screen displays the menu and if Option 4 is then selected the screen will display the “raw” data as it was originally recorded (see Fig 3.1). (v) Determination of Rate at Different Percentage PFDs To obtain PFDs which give any percentage of the maximum rate, press F5 and enter desired percentage. The corresponding PFD and rate will be displayed. This allows, for example, comparisons to be made with data giving a lower ceiling. Such data is recalled from memory (Option 7) and over-printed (Option 5 vi) with data giving a higher ceiling. The lower ceiling is then raised to coincide with the higher.

Table 5 2 Screen displays of rate v PFD data (2)

Table 5.2 Above: Second screen display (c.f. Table 5.1) of data recorded in Option 3 prior to graphic plot (achieved by pressing F2). F1 returns you to menu. F3 autoscales, F4 allows addition to comments (middle box, left). F5 allows access to lower box left (Table 5.3). Table 5.3 Screen display of rate v PFD data (3)

Table 5.3 as for Table 5.2 pressing F5 has given access to lower panel in which parameters may be altered by overtyping. Note retrospective adjustment of leaf area at foot.

Figure 5.2a Rate v PFD plot (Option 5) of raw data (Fig. 3.1) obtained using Option 3 and displayed in numerical form in tables 5.1-5.3 Quantum yield has also been determined by least squares regression (Section 5iii).

Figure 5.2b As for 5.2a but several changes have also been made. This was done, in part, by pressing F5 when the data was first displayed (in Option 5) in the form illustrated in Table 5.2. Pressing F5 at that stage converts the screen display to that illustrated in Table 5.3 and the values in the lower box can then be varied by overtyping. The horizontal and vertical scales can also be changed by using F2 and F3. A second set of data has also been superimposed (see Section C5 vi) as in Fig. 5.1f but in this instance the additional data came from the same leaf-disc recorded under very similar conditions and were therefore so similar that the second set of points can only easily be distinguished from the first at the extreme right.

(vi) Overprint Press F6. You will be asked (at the top of the screen) to enter the file name for previously recorded rate v PFD plots. If you do this and press enter, this data will also be displayed on the screen on the same scale. This allows direct comparisons to be made on the same axes (see Fig. 5.1f and 5.2b). (vii) Comparing data in different directories Suppose that you have recorded rate v PFD in one directory and wish to compare it with similar data previously stored in another directory. Proceed as before but, having pressed F6, type (e.g.) C:\HA\QSUN2. This will overprint with data from a file QSUN2 in a directory HA. (viii) Plotting data not previously recorded If you have data from some other source and you wish to take advantage of the computer’s handling facilities, press F10(New). An empty box will now be offered. If PFD and corresponding rate values are entered in this space this data can be stored and handled in the usual way.

C 6. Experimental Parameters (Option 6) If Option 6 is selected (by keying 6, followed by enter), the screen displays: Parameters A. Programme and PFD directory: B. DATA directory: C. Time Betweeen AUTO steps (seconds): D. Rate Window (seconds) E. Rate Step up size (Moles [O2]): F. Area Cut off PFD (E/M2/s): G. Traces Wrap: H. Interface Board Present [Hansatech ADC/DAC]: I. Time between each stored data set: J. Axis Settings (Top,Bottom) K. Weighting to first part of curve in ZINfit: L. Leaf area in cm2 M. Verify Data Saved: N. Real Time Differential O. Bjorkman Lamp: Esc. Return to Menu Enter a letter:

C:\LD\ C:\LD\ 60.00 :50.00 0.50 800.00 TRUE TRUE 200 40,40 100.00 10.00 FALSE TRUE FALSE

To change any parameter, type in the appropriate letter (i.e., A to N) followed by a new value. To retain such changes, exit via X (see Section J). A: Programme and PFD directory: The main use of this option is to be able to tell the programme where the PFD files (i.e. those illustrated in Table 3.1) reside. B: DATA directory: Use this option to set the current data directory. e.g., If you store your data in a “QYDATA” directory, change B to read

C:\QYDATA\. A new directory with the name QYDATA must first be established using the “md” command. (See Section H). C: Time between AUTO steps (seconds): Sixty seconds is the default value (i.e. the value which will be automatically used each time the computer is switched on). If, in a rate v PFD determination, you wish to illuminate a leaf for a longer (or shorter) period of time at each light intensity, enter the new time interval D: Rate Window (seconds): The number of seconds over which rate is calculated (default being 50). To change, press D, type in new time (e.g., 25) and press enter. Note that the rate window must be smaller (e.g., 50) than the time (see C above) between autostops (e.g., 60). E: Rate Step up Size: This should need to be changed only rarely. If, for example, the rate of photosynthesis is low, during a rate v PFD determination you may wish to have a larger “step up” between one PFD and the next to avoid overwriting of data. In many instances the same effect can be achieved by expanding the vertical scale. F: Area Cut off PFD (µE/m2/s): The default value of 800 is used by the computer to calculate the area under the curve as a percentage of estimated maximum. If, however the maximum PFD is more (or less) than 800 (or if you wish to have calculations based on a lower PFD, e.g., 100), this value can be altered by pressing F and typing in the actual value and pressing enter. Alternatively, once the lines have been displayed (in Option 5, F4 [area]) the cut-off PFD can be located manually using the “ ←↑ ↓ →“ keys. G: Traces Wrap: When either fluorescence or O2 reach the top of the screen the trace “wraps round” and reappears at the bottom of the screen. If, for any reason, this feature is not required, press G and answer FALSE. H: Interface Board Present: Presence (T) or absence (F) is set accordingly by computer. F produces sine waves I: Time between each stored data set: Use this to set the rate at which data sets are stored. Each data set will be an average of A D readings taken during the interval since the last data set. J: Axis settings (Top, Bottom): The location of the axes on the screen can be altered using this option. K: Weighting to first part of curve in Zin fit: This relates to an analytical facility not yet available. L: Leaf Area in cm2: When it is necessary to use less than 10 sq cm of leaf, enter the area here. The computer will then calculate the rate of O2 evolution, using this new area (default is 10 cm2). Retrospective change in leaf area is available in Option 5. M: Verify data saved - default is FALSE: This is an option to prevent careless loss of data. If you change to “true” (by keying m, followed by t and enter) a warning will appear in the menu if you seek to

change an option which would otherwise result in the loss of data which had not been stored. If you wish to ignore this warning, key y. N. Real Time Differential - This allows the possibility of recording oxygen as a rate whilst actually running an experiment in Option l. [It is still possible to use the retrospective differential (F8 in Option 4) if the real-time differential is used]. If the TRUE command is initiated by pressing “t” and enter, then, whilst still in Option 6, the screen will display, in succession, each of the differential parameters in the table shown below: Differentiator Parameters Time Windows (Secs) [50]: Screen Range (nanoM/S) [20.00]: Zero Value (nanoM/S) [-10.00]: Each line comes up with its default value each time enter is pressed. If different values are required then key in the value before pressing enter in each case. After the zero value option, pressing enter will return you to Option 6 and Esc will return you to the menu.

C 7. Retrieve (Option 7) (i) On entry (press key 7 and enter) this option will display all of the previously stored files in the currently selected directory (see e.g. Table 7.1). Table 7.1 File titles displayed in Disc Retrieve Option. Disc Retrieve Option. JA1.INF QJA1.INF KQ1.INF KQ2.INF WASHQ3.INF WASHQ6.INF

QST1.INF KQ3.INF

QST2.INF WASHQ1.INF

In the “disc retrieve option” (Table 7.1), “.INF” indicates “information” (as opposed to “trc” which means “trace”). As we shall see later, this arrangement allows you to discard raw (“trace”) data (which occupies large amounts of memory while retaining processed (“INF”) data which you may wish to compare with other data (e.g., OPTION 5, F6). Key in the appropriate file name (see Option 8) and enter. If you select a file name for a rate v PFD measurement (e.g., QJA1) the data will be first expressed in tabular form (Table 5.1) and you will be asked if you require the “raw” as well as processed data (y/n). If you say “yes” the screen will display the traces as they were recorded (Fig 3.1) when you press key 4 and enter. If you press key 5 and enter the rate of O2 evolution will be plotted as a function of incident PFD (Fig 5.1). (If only the rate data has been retained, i.e., if the raw data has been discarded see section Hv, the screen will give the same display on pressing 5 and enter, but will remain blank after 4 and enter). If the data that you have recalled (e.g JA1) is from “pen recorder mode” (i.e. change in O2 and rate of fluorescence emission as a function of time) the screen will, at first, display any information (Table 7.2) about the experiment which has been entered previously (Option 8 on MENU), the time and date of the experiment and its duration. Press enter once to return to the MENU and twice to have this data plotted on screen.

Table 7.2 Information about an experiment, recorded in Option 1 and displayed in Option 4, following retrieval from disc (Option 7) C:\bg1.INF Reading C:\bg1.INF File information:08:20:3711-20-1987blue gum showing initial burst Number of data sets: 575 (4.8 minutes long).

Figure 7.1. The initial slope of the rate v PFD curve derived by a procedure in which more points were taken in the 0-125 µmole.quanta.m-2.s-1 range. (a) initial slope derived by least squares regression (b) as for (a) but initial slope derived by eye (F7).

If you wish to change the presentation of recalled data, proceed as follows:(ii) Change vertical (or Y) axis by pressing F2 followed by F2 for 20 mmoles (of oxygen) full scale, F3 for 10 µmoles full scale etc (see possible options at foot of screen and Table 1). To change horizontal (or X) axis press F3 followed by F2 for time intervals of 0.5 min, F3 for time intervals of 1.0 min, etc (see options displayed at foot of screen and Table1). (iii) Differential Press F8 to differentiate the oxygen trace so that this becomes expressed as a rate (dO2/dt) rather than change in oxygen against time (see Section 4).

C 8. Store Data on Disc (Option 8) When this option has been entered, the screen will display (Table 8.1) all of the filenames (see Option 7). You may use a total of up to 8 letters and digits to define new data which you wish to record, e.g., ELMQ3 which might mean, to you, that this is the third quantum yield determination which you have made on an elm leaf. Entering this filename will store the data, which you have just recorded, in the currently selected directory on disc. It can be retrieved using Option 7. File names must not contain any full stops, commas, hyphens or slashes. The designation .INF (see OPTION 7) will be added by the computer to indicate that this is an “information” as opposed to a “trace” file. Table 8.1 Screen display in Option 8 Disc Save Option. JA1.INF QJA1.INF KQ1.INF KQ2.INF WASHQ3.INF WASHQ6.INF

QST1.INF KQ3.INF

QST2.INF WASHQ1.INF

When you type in a file name (e.g QJA1) the computer will tell you if this name has been used before and invite you to overprint (y). If you answer N you will be returned to the menu so that you can proceed as you wish. If the name that you have chosen has not been used before and you press enter, you are now offered an opportunity to enter information about the experiment which you have just completed (Table 8.2). Table 8.2 Describing experiments on storing. QST1 Current information : 09:06:33 Tue 23-05-1989 Enter any further info (to max 255 Chars):

This information is subsequently displayed in OPTION 5 (Table 5.1, top, Tables 5.2,5.3, centre box, left) and may also be retrieved by using the procedure described in Section Iviii, which allows you to simultaneously scan information relating to a number of experiments.

D. ASPECTS OF ILLUMINATION (a) Calibration and Uniformity. If we ask the computer nicely, and if the appropriate hardware is available and in place, it will calibrate PFD for us. (The procedures for both manual and automated calibration are described in detail in OPTION 3). In any case, a photon-counter is needed and one supplied by Hansatech Instruments specifically for this purpose (see also Appendix 2 of ref. 7) is recommended because it fits snugly over the window against which the leaf disc is pressed during measurement and integrates the PFD over the whole of this surface. It is also calibrated to detect photons in the same range as those emitted by the LEDs. It should be noted that the LED source does not give a fully uniform field of illumination and that, if the most uniform possible field is required, other steps should be taken. Very much greater uniformity is very easily accomplished by inserting light diffusers between the LED’s and the top window of the chamber. Such diffusers not only make the field of illumination much more uniform but also permit the most effective use of LED numbers (covering a large range) in accurate measurements (see e.g., Fig. 6.1) of initial slope in the 0 - 100 mole quanta m-2 s-1 region of the rate v PFD relationship. At higher PFD’s, diffusers are less desirable because of the associated decrease in light which is welcome in low light measurements. One alternative is to use an insert which drops into the chamber, thereby diminishing its volume and, at the same time, leaving a central field of illumination which is very much more uniform. (b) Transmittance and Reflection. If rate v PFD measurements are based on incident light, i.e on the rate at which photons arrive at the leaf surface, the initial slope of the rate v PFD relationship is a measure of relative quantum yield (and its reciprocal, quantum requirement). Relative quantum yield is a useful and simple measurement which permits, for example, comparisons between varieties or species, or changes in the initial slope consequent upon stress or recovery from stress. For such purposes, absolute values may be less important than comparison of relative values. If, on the other hand, absolute values are needed, corrections must be applied because light which is reflected from the surface of the leaf, and transmitted through it, is obviously not used in photochemical events within the leaf. Traditionally, corrections are made on the basis of values derived with an Ulbricht sphere. This contains a light source and a quantum sensor within its fully reflective inner surface. The difference between measured PFD when the sphere is empty and when a piece of leaf is inside it gives the amount of light absorbed by the leaf. Transmittance of light through a leaf (at the wavelengths emitted by the LED’s) can also be measured, with accuracy, simply by comparing the light measured by the photon-counter with and without a leaf disc between the counter and the lower window of the chamber. Reflectance is more difficult to measure with accuracy but, for many purposes differences in reflectance will have little impact on results because they will account for such a small percentage of the incident light. Approximate allowance for such differences can be made by utilising a simple device which may soon be available from Hansatech Instruments. This incorporates a single LED and a quantum sensor. When placed on a leaf (which in turn has been placed on a piece of black felt or similarly non-reflecting surface) the only light which can enter the sensor is that (from the LED) which is reflected back from the leaf surface. Reference to a calibration curve gives a measure of reflectance from this area of the leaf surface. If reflectance and transmittance measured in these ways are subtracted from incident light,

relative quantum yield becomes absolute quantum yield. If you wish to apply such a correction to a graphic representation of data in OPTION 5, the retrospective leaf-area facility (accessed via F5) can be used for this purpose. Thus light which is not absorbed is equivalent, in this regard, to leaf which does not exist. For example, if transmittance was 10% and reflectance 5%, a 10 cm2 leaf disc is, in absolute terms, equivalent to one of 8.5 cm2. Accordingly, if you apply such a retrospective leaf area correction as though 100% of the incident light had been absorbed, it would be as though the leaf area had been (10 + 1.5 =) 11.5 cm2. Increasing the effective leaf area in this way will give rates of O2 evolution pro rata lower than those calculated on the basis of the actual leaf area and diminish the calculated value of quantum yield from the (higher) relative value to the (lower) absolute number. Bear in mind that, despite crafty electrical circuitry, the output of the LED may change with temperature. If, for example, you wish to measure from high to low PFDs (rather than from low to high) calibrate in that direction. With the automated procedure it is possible to ask the computer to make a calibration run, during which the photon-counter experiences the same PFDs as a leaf assayed in precisely the same way. Even without this feature, however, good accuracy can be achieved. A leaf is the best judge of this. Irregularities in measurements can obviously arise in several ways but, if everything else is as it should be, the majority of rate v PFD points should fall on a smooth line (Fig. 7.1) or curve (Fig. 5.1). If they do not, check your PFD values again.

E. DEMONSTRATION DATA The disc which you purchase from Hansatech will include the data on which most of the figures reproduced here are based. The data may be retrieved from memory according to Option 7 and are amongst those listed in Table 7.1. Once retrieved, these data may be manipulated to give screen displays similar to those illustrated in Figs. 1.1, 5.1, 5.2 etc., by using the procedures described in Options 1 and 5.

F. DEFAULT VALUES Where possible, default values, once changed, remained changed i.e., if you prefer a time-window of 30s to one of 50s and you have changed to 30s from the original default value of 50s, the computer will not revert to 50s on switching off.

G. PRINT OUT Provided a suitable printer is connected and the appropriate DOS printer files have been loaded (see Section I) it should be possible to print by pressing “print screen” or “shift” and “print screen” or “print screen” and enter.

H. REMINDERS Experimental Procedures are described more fully in the text but pay particular attention to the following:

“a leaf is the best judge”

(i) Leaf Material Leaf discs should be cut with a sharp cutter (or leaf pieces with a sharp blade). Avoid bruising and loss of turgor. Cutting under water can be to advantage but, with some leaves, it can lead to unwanted infiltration. Many partially flaccid leaf-discs will regain full turgor if floated on water for a few minutes and most will survive floating on water for 2 or 3 hours but, if it is wished to maintain discs beyond this, it is best done by placing them on a damp paper, tissue, or cloth in an enclosed space in order to avoid gradual infiltration. (ii) Carbon Dioxide For many reasons, 1 M bicarbonate on capillary matting is an adequate source of CO2 but carbonic anhydrase or phosphate ion (which also catalyses the equilibrium between CO2 and bicarbonate) may also be necessary if very rapid rates of photosynthesis are anticipated. The matting which carries the bicarbonate solution should be moist rather than wet. Ideally, the chamber should also be flushed, prior to measurement with a moist gas-stream containing the same equilibrium concentration of CO2. Absolute calibration must be carried out with air containing the same CO2 concentration as that used in the experiment. In our own Laboratory we have used a procedure in which we draw air (using a 10 ml plastic syringe) through a bicarbonate solution in a glass vessel maintained at the same temperature as the chamber by water circulating through an external jacket. The syringe is attached to one of the chamber taps for this purpose and, once filled, is used to push this gas mixture, now humid and containing the same equilibrium concentration of CO2 as the chamber, into the chamber. This procedure obviates the necessity for re-charging the bicarbonate solution on capillary matting within the chamber between consecutive runs with the same leaf-disc. Obviously, gas from a tank, if available, can be used in the same way to flush the chamber. Although procedures such as the above can be used to meet the CO2 requirements of many C3 species it may sometimes be necessary to use higher CO2 concentrations (see e.g., Reference 6). Conversely, inhibition by CO2 may occur (particularly at higher concentrations) because of acidification of the stroma (Reference 5) and these aspects must obviously be borne in mind using the apparatus with unfamiliar leaves or if anomalous results are observed. (iii) Absolute Calibration of O2 The arithmetic in the present calibration procedure is based on the oxygen content of air at 100 KPa. If you are concerned about the error which is inevitably introduced if you are working, for example, at high altitudes or on abnormal barometric pressure an appropriate correction should be introduced (see below). It is convenient to calibrate a chamber already containing CO2. At present, calibration neglects the O2 displaced by high concentration of carbon dioxide in the atmosphere within the chamber on the calibrating syringe. For example, if 5% CO2 is present in both, the excursion consequent upon the insertion of 1 ml of gas would be 8.74 µmoles rather than 9.2 µmoles. This means that rates will be correspondingly over-estimated by a factor of 9.2/8.74. For many purposes this intrinsic error is sufficiently small to be neglected. A correction in “real” time can

be applied by inserting a falsely high temperature, sufficiently large to diminish the excursion (in this particular case) from 9.2 to 8.74. A retrospective correction can be made in Option 5 by inserting a falsely high leaf area (i.e, in this particular case, an area of 10 x 9.2/8.74). (iv) Capillary Matting If the leaf-disc is supported on capillary matting (to facilitate maintenance of turgor) it should be moist rather than wet (to ensure that there is not a continuous aqueous phase which would greatly diminish the rate of diffusion between the leaf-disc and the detector). (v) Leaf pieces If it is not possible to use 10cm2 discs, it is desirable (because of possible lack of uniformity of PFD within the chamber, maintenance of high humidity, etc. to fill as much of the 10cm2 area as possible with leaf tissue. (vi) Pre-illumination Leaves taken from darkness may display short, and/or long induction. Some may display photosynthesis which is partially suppressed by a nocturnal inhibitor of Rubisco. Similarly, darkening or even transfer from one light intensity to another may re-initiate induction or other constraints. Prolonged illumination may bring about photoinhibition or initiate feed-back mechanisms which adversely affect photosynthesis. For all of these reasons it is virtually impossible to measure photosynthesis in a “steady” state. If you wish to measure quantum yield there is much to be said for pre-illumination at about 100 to 125 µmole.quanta.m-2.s-1 until the rate of photosynthesis approaches constancy and then to decrease the PFD in a fairly large number (15-20) of small steps to complete darkness. This avoids inadvertent photoinhibition, provided that pre-illumination is not prolonged more than 5 minutes or so. It also avoids distortion of readings taken at low PFDs by light-enhanced dark respiration which can lead to O2 uptake (on abrupt darkening from bright light) which, at PFDs below the light compensation point, can be greater in magnitude than photosynthetic O2 evolution. This can lead to effects similar to those first described by Kok (see Reference 7, Section 23) in which there is an apparent inhibition, by low light, of dark respiration.

I. COMPUTING NOTES (i) Software The files which are required to run this programme are as follows. LD. EXE this is the leaf-disc programme itself plus DOS files or

GRAFTABL.COM GRAPHICS.COM HGS .COM

If the programme is to be put on a hard disc in a computer then an LD directory must be created (see below). Such a directory is automatically created if the procedure described in Section A is followed. A separate facility called SEARCH, may be used to search for recorded data etc (see viii below).

(ii) Creating Files

Each set of experimental data constitutes a “file”. For example, if you ask the computer, in option 1, to record O2 evolution and fluorescence emission and, at the end of your experiment you seek to record the data (under option 8) you will be asked to create a file by entering a file title which can have up to 8 letters (or digits or a mixture of letters and digits but no spaces) followed by whatever information you care to add about the nature of your experiment (255 characters, which is just up to the beginning of the third line).

(iii) Creating Directories When you wish to store data you will need to create “files” but you will also need a place to put them. Files are kept in a “filing-cabinet” called a directory. In order to create a directory to put your files in, proceed as follows (Note CD means “change directory” and MD means “make directory”) (a) Press X The screen will now display C:\LD> (or C:\DATA>, or whatever, if you are already in an existing directory) (b) Type CD .. and enter (don’t forget the full stops) The screen will now display C:\> (c) Type MD followed by the name of the new directory e.g., MD NEWDATA and enter The screen will now display C:\> (d) Type CD NEWDATA and enter

The screen will now display C:\NEWDATA> Now call up the Leafdisc programme by typing CD.. and enter, which will give C:\> follow this by typing LD (or GO or HGO as appropriate) and enter. This will bring up the LeafDisc Logo and the programme is ready for use. In Leafdisc option 6, enter B followed by >\NEWDATA and enter. Your filing-cabinet will be in place waiting for you to store files in it. Once you have put files in it you can not send it to the scrap-heap before you empty it but, at this stage, if you do not like it, you should exit from the programme (via ESCAPE and X) and go through the above procedure again but this time enter RD (remove directory) instead of MD, followed by enter which will give C:\>. If you now type LD and enter, you will be returned to LeafDisc but you will find that you are now unable to access NEWDATA in option 6B. (iv) Looking for Directories Keying DIR allows you to read all files and/or directories in the directory you are currently in. (v) Delete By keying DEL, followed by the file name, files may be removed from the disc. The use of a * will allow block deletions. e.g

*.* *.trc WASHQ3.*

will delete everything will delete all trace data will delete both trace and information data for the file WASHQ3 A*.* will delete all files beginning with A Note that there is virtue in discarding trace data, where possible, because this will account for 90% of the memory devoted to data handling. (vi) Backup It is useful to copy all data with a backup system in case of accidental deletion or disc fault. (vii) Looking for lost files (a) Exit from the programme by pressing X (b) Enter C:\>tree /f more Tree /f more - this command allows you to search your disc in order to find lost files. The whole disc contents can be printed out by keying tree /f more >PRN (viii) Searching for a file in a directory (a) Exit from the programme by pressing X. If you happen, at this time, to be in directory AD1 for example, in drive D (if you have one), the screen will display D:\AD1>. More usually, it will display something like C:\DATA>

If you wish to find a series of files in this directory (DATA) called WASHQ1, WASHQ2 etc., you should now make this read C:\DATA>Search WASHQ?.inf FOR “:” being careful about the spacing and being sure to put FOR in capital letters. On pressing enter the screen should now display the information about all of these files that you have put into them at the time that you created them. Similarly, if you ask for WASHQ1 you will get this alone. If you ask for ??ASHQ? you will get 2TASHQ1 (for example) but not any of the WASHQ series, whereas ?ASHQ? will yield all files labelled WASHQ1, WASHQ2 etc., plus, (for example) TASHQ1, TASHQ2 etc. If you want to see the description of all of the files that you have put in the directory use SEARCH *.inf FOR “:” but be prepared to wait many minutes before the search is complete if you have put lots of files in the directory. At the end of the search routine a file with all this information is created called f.$$$. This can be printed out by the command type f.$$$ >PRN (ix) Transferring files To copy a file from one disc to another, or between directories use the copy command as shown below: COPY C:\QYDATA\SPINACH\QW1.* A:\*.* This copies file QW1 from the directory QYDATA\SPINACH in drive C:\ onto a floppy disc in drive A:\ without changing its name. OR, if you happen to be to be in D:\AD1, exit from the programme via X and enter so that the screen displays D:\AD1> Make it read D:\AD1>copy d:\AD1\WASHQ3.* d:\can1\*.* and enter. This will transfer file WASHQ3 in directory AD1 (in drive D). If you put WASHQ*.* WASHQ1, WASHQ2 etc., will also be transferred. (x) How best to file data Inevitably, this is to a large extent, a matter of personal choice. A secretary who puts almost everything into a single large file called “Miscellaneous” would have great difficulty in finding things. At the very least, it would make sense to have folders marked AZ and there might be advantage in using many other categories, as sub-categories necessitating the use of several drawers or even several filing-cabinets. A librarian, bent on cataloguing a whole library would have to proceed on some such basis. For you it may be different. You may wish to emulate the secretary, or even the librarian, and create a tree. Starting in drive C, such a tree could look like this.

c:\

QYDATA

LEAFDISC

BEAN BARLEY SPINACH

the programme PFDfiles

PHOTOINHIBITION

ACTIVATION

etc You have already seen how to make a directory called “NEWDATA”. You can make a new sub-directory in the same way e.g., Starting from C:\NEWDATA>

type MD SPINACH (and enter)

followed by CD SPINACH (and enter)

the screen should now display

C:\NEWDATA\SPINACH> (You can use the command CD.. to move you back through this tree, one directory at a time or CD\ to go back to the root directory, C). If you use this approach you have to enter each file name, in full, under option 6b, before embarking on an experiment. An alternative, which you may prefer, is to use the 8 letters and digits which go into a file name to best advantage. Thus a prefix C could indicate a change v time plot (option 1) or Q, a quantum yield plot (option 3 + filter), S, (or SO) could indicate spinach, P or (PI) photoinhibition etc as in QSP1 which still leaves 4 letters or numbers for further description. This approach has the advantage that it is simpler, and allows immediate over-printing (f6 in option S). It may encourage you in the conviction that files are made to be destroyed and that a residuum of worth is better shared with the world in a publication rather than being left to gather dust in some innermost recess of computer chip.

J. QUITTING There are, of course, many ways to quit and we hope that no-one will be prompted by his or her experience with this programme to take the most extreme options which are available. The nicest way to go is via X (see MENU). If you take this route you will find, to your pleasure, that the changes in parameters which you made when you last used the programme will be retained. If, of course, you wish to make an entirely fresh start simply switch off the computer.

REFERENCES. 1.

Björkman, O. and Demmig, B. (1987) Photon yield of O2 evolution and chlorophyll fluorescence characteristics at 77K among vascular plants of diverse origin. Planta 170: 489-504

2.

Delieu, T. and Walker, D.A. (1981) Polarographic measurement of photosynthetic oxygen evolution by leaf discs. New Phytol. 89, 165-178.

3.

Harper, W.M. (1971) Statistics. 2nd Edition. Macdonald and Evans, Plymouth.

4.

Kok, B. (1949) On the interrelation of photosynthesis and respiration in green plants. Biochim. Biophys. Acta 3, 625-631.

5.

Quick, P., Siegl, G., Neuhaus, E., Feil, R., Stitt, M,. Short-term water stress leads to a stimulation of sucrose synthesis by activating sucrose-phosphate synthase. Planta 177. 535-546.

6.

Robinson, S P. Grant, W.J.R. Loveys, B.R. (1988) Stomatal limitation of photosynthesis in abscisic acid-treated and in water-stressed leaves measured at elevated CO2. Aust. J. Plant Physiol. 15, 495-503.

7.

Walker, D.A. (1987) The use of the oxygen electrode and fluorescence probes in simple measurements of photosynthesis. Oxygraphics Ltd, pp 1-145.

Appendix 4

Latest Hansatech Equipment

K. HANSATECH INSTRUMENTS Recent additions to Hansatech Instruments Limited product range 2000/01 (see also Appendix 2). DW2/2

Oxygen electrode unit for simultaneous oxygen and fluorescence measurement in the liquid phase, with improved optical performance

DW3

Large volume oxygen electrode unit for simultaneous oxygen and fluorescence measurement in larger liquid phase samples

OXYG1

PC operated oxygen electrode control unit with integral magnetic stirrer. Multi-channel capability (up to 8 channels) with additional Oxygraph units.

OXYT1

PC operated oxygen electrode control unit with integral magnetic stirrer and solid state temperature control. Multi-channel capability (up to 8 channels) with additional Oxytherm units.

OXYL1

PC operated oxygen electrode control unit with integral magnetic stirrer. Automatic control of light intensity changes using LH36/2R or LH11 LED light sources.

LD1/2

Gas phase oxygen electrode unit with improved calibration and temperature control

LD2/3

Leaf disc electrode unit for simultaneous oxygen and fluorescence measurement in the gas phase, with improved temperature control DW2 and LD series cuvettes can be supplied with adaptors to allow simultaneous use of the Walz PAM fluorimeter during O2 measurement

LH36/2R

Light housing with 36 Ultra-bright red LED array with automatic control from the Oxylab electrode control unit.

LH11/R

Light housing with 11 Ultra-bright red LED array with automatic control from the Oxylab electrode control unit.

QSPAR

Large area PAR quantum sensor

QSRED

Large area PAR quantum sensor (filtered for red waveband 550-750nm)

QRT1

Hand held digital PAR/Temperature sensor

FMS1

Lab-based pulse modulated chlorophyll fluorescence monitoring system

FMS2

Field-portable pulse modulated chlorophyll fluorescence monitoring system

Handy-PEA

Field-portable continuous excitation fluorescence monitoring system

the end