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1993

Seed Dormancy in Red Rice (Oryza Sativa):Changes in Embryo pH and Metabolism Duringthe Dormancy-Breaking Process.Steven FootittLouisiana State University and Agricultural & Mechanical College

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O rder N u m b er 9401522

Seed dorm ancy in red rice ( Oryza sativa): Changes in embryo pH and m etabolism during th e dorm ancy-breaking process

Footitt, Steven, Ph.D.

The Louisiana State University and Agricultural and Mechanical Col., 1993

U M I300 N. ZeebRd.Ann Arbor, MI 48106

SEED DORMANCY IN RED RICE (ORYZA SATIVA): CHANGES IN EMBRYO pH AND METABOLISM DURING THE DORMANCY-

BREAKING PROCESS.

A Dissertation

Submitted to the Graduate Faculty of the Louisiana State University and

Agricultural and Mechanical College in partial fulfillment o f the

requirements for the degree of Doctor of Philosophy

in

The Department of Plant Pathology and Crop Physiology

bySteven Footitt

B.Sc. (Hons), North East London Polytechnic, London, England, 1985May 1993

Dedicated to the memory o f my beloved friend,

Suzanne Packer (1958-1989).

ACKNOWLEDGMENTS

For his advice, encouragement, and patience, I would like to thank my long

suffering major professor, Dr. Marc Alan Cohn. I also wish to thank the members of

my advisory committee: Drs. JP Jones, J Lynn, K Morden and M Musgrave, for their

advice and encouragement.

Financial support was provided by the Department of Plant Pathology and Crop

Physiology, LSU, (research assistantship) and the American Seed Research Foundation

(travel). I am indebted to Dr. John Baker for his continued support during this

research.

I am grateful to my good friend, Dr. Robin Probert of the Royal Botanic

Gardens, England, who awakened my interest in seed dormancy, then promptly got

appendicitis and left me to work it out for myself. Many thanks to Fujiko Signs and

Esther Quintinella for translating manuscripts from Japanese and Italian, and Dr.

David Vargas, for his invaluable assistance with the NMR experiments.

For their help, good humour, and companionship, I especially wish to thank

my friends and fellow participants in this mad pursuit: Gloria Balagtas, Begona

Gutierrez, Karen Jones, Mark Signs, Drs. John Constable, Christine Daugherty, Mark

Evans, Sathasivan Kanagasabapathi, Bill Odom, and Jim Smith.

I would especially like to acknowledge the love and support of my parents who

have suffered my long absence without complaint.

TABLE OF CONTENTS

PAGE

DEDICATION ii

ACKNOWLEDGEMENTS iii

LIST OF TABLES vi

LIST OF FIGURES x

ABSTRACT xiv

CHAPTER

1 SO YOU WANT TO BREAK DORMANCY?CROSS-KINGDOM SIMILARITIES IN CHEMICAL TREATMENTS

Introduction 1Early development of chemical based dormancy-breaking treatments 2Common features of chemical based dormancy-breaking treatments 3Is there a mechanistic basis for the parallels observed? 10

2 DORMANCY-BREAKING IN SEEDSIntroduction 11Physiological dormancy 13Contemporary dormancy-breaking hypotheses 14Components of metabolism as markers 15Markers arising from dormancy-breaking chemical treatments 17

3 EMBRYO ACIDIFICATION DURING DORMANCY-BREAKING AND SUBSEQUENT GERMINATION

Introduction 20Materials and Methods 22Results 27Discussion 44

4 LEVELS OF FRUCTOSE 2,6-BISPHOSPHATE IN RED RICE EMBRYOS DURING DORMANCY-BREAKING AND GERMINATION

Introduction 50Materials and Methods 52Results 57Discussion 66

5 A 13C NMR STUDY OF THE METABOLIC FATE OFDORMANCY-BREAKING CHEMICALS IN DORMANT RED RICE EMBRYOS

Introduction 72Materials and Methods 73Results 78Discussion 86

6 31P NMR OF DORMANT RED RICE EMBRYOS: THE OBSERVATION OF INTERNAL 31P USING A SPIN ECHO SEQUENCE

Introduction 89Materials and Methods 90Results 95Discussion 103

7 CONCLUSION 107

LITERATURE CITED 111

APPENDIX A: ACCOUNTING FOR DIFFERENCES BETWEEN 24H PULSE AND OH POST-PULSE EMBRYO pH 128

APPENDIX B: EMBRYO pH DURING DRY AFTERRIPENING 131

APPENDIX C: HARVEST COMPARISON 134

APPENDIX D: DATA TABLES FOR CHAPTER 3 136

APPENDIX E: DATA TABLES FOR CHAPTER 4 160

APPENDIX F: DATA TABLES FOR CHAPTER 5 173

APPENDIX G: DATA TABLE FOR CHAPTER 6 178

VITA 180

V

LIST OF TABLES

TABLE PAGE

1.1 Chemicals showing dormancy-breaking activity and the kingdoms inwhich they are found to be active. 4

3.1 Moisture content(%) of dry intact nondormant red rice seeds and ofseed components. The contribution to whole seed MC of each component is also shown. 28

3.2 Changes in [H+] of embryo homogenates detected after a 24 hourchemical pulse. 43

5.1 13C NMR assignments of standards and metabolite signals. 84

5.2 Effect of pH on 3-hydroxypropionate assignments. 85

A .l Embryo homogenate pH as measured in 1989, recalculated forextraction in 916.8 fxL, and determined again in 1990 using the same harvest (SH 87-1). 130

D .l Embryo data for hydrating dehulled seeds of dormant red rice at30°C. 137

D.2 Embryo data for hydrating dehulled seeds of nondormant red rice at30°C. 138

D.3 Endosperm data for hydrating dehulled seeds of dormant red rice at30°C. 139

D.4 Endosperm data for hydrating dehulled seeds of nondormant red riceat 30°C. 140

D.5 Effect of nitrite at pH 3.0 on embryo pH and germination, as in figure3.2. 141

D.6 Effect of nitrite at pH 7.0 on embryo pH as in figure 3.2. 142

D.7 Effect of 10 mM nitrite at pH 3.0 on endosperm pH as in figure 3.2. 143

D.8 Effect of nitrite at pH 7.0 on endosperm pH as in figure 3.2. 144

D.9 Effect of 25 mM citrate/phosphate buffer at pH 4.9 on embryo pH and germination as in figure 3.3.

D. 10 Effect of 20 mM propionate at pH 4.9 on embryo pH and germination as in figure 3.3 & 3.5.

D .l l Effect of 70 mM propanol at pH 4.9 on embryo pH and germination.

D. 12 Effect of 25 mM citrate/phosphate buffer at pH 7.0 on embryo pH and germination as in figure 3.3 & 3.5.

D.13 Effect of 70 mM propanol at pH 7.0 on embryo pH and germination as in figure 3.3 & 3.5.

D.14 Effect of 40 mM propionaldehyde at pH 7.0 on embryo pH and germination as in figure 3.3 & 3.5.

D.15 Effect of 30 mM methyl propionate at pH 7.0 on embryo pH and germination as in figure 3.3 & 3.5.

D. 16 Effect of 25 mM citrate/phosphate buffer at pH 4.9 on endosperm pH as in figure 3.4 & 3.6.

D.17 Effect of 20 mM propionate at pH 4.9 on endosperm pH as in figure3.4 & 3.6.

D. 18 Effect of 70 mM propanol at pH 4.9 on endosperm pH.

D.19 Effect of 25 mM citrate/phosphate at pH 7.0 on endosperm pH as in figure 3.4 & 3.6.

D.20 Effect of 70 mM propanol at pH 7.0 on endosperm pH as in figure 3.4 & 3.6.

D.21 Effect of 40 mM propionaldehyde at pH 7.0 on endosperm pH as in figure 3.4 & 3.6.

D.22 Effect of 30 mM methyl propionate at pH 7.0 on endosperm pH as in figure 3.4 & 3.6.

D.23 Embryo pH in the dormant 1990-1 harvest following 24 h/30°C exposure to 22 mM propionate or 75 mM «-propanol in 25 mM citrate/phosphate buffer at pH 4.9.

E. 1

E.2

E.3

E.4

E.5

E.6

E.7

E.8

E.9

E. 10

E .l l

Radicle and shoot emergence after hydration of nondormant seeds for 8 to 12 h/30°C followed by drying for 7 d/30° then rehydration for 7 d/30°C. 161

Seed respiration and embryo [Fru 2,6-P2] data for hydrating dormant seeds in figure 4.1 and 4.2. 162

Seed respiration, embryo [Fru 2,6-PJ and germination data for hydrating nondormant seeds in figure 4.1 and 4.2. 163

Endosperm [Fru 2,6-PJ data for hydrating dormant and nondormant seeds. 164

Embryo [Fru 2,6-PJ, real time germination, and contact timegermination for the combined controls during and post contact as in figure 4.3. 165

Embryo [Fru 2,6-PJ, real time germination, and contact timegermination resulting from a 24 h/30°C pulse of 32 mM methyl propionate/ 25 mM citrate/phosphate buffer at pH 7.0 as in figures 4.3 and 4.5. 166

Embryo [Fru 2,6-PJ, real time germination, and contact timegermination during a 24 h/30°C pulse of 4 mM sodium nitrite/ 25 mM citrate/phosphate buffer at pH 3.0 as in figures 4.3 and 4.5 167

Embryo [Fru 2,6-PJ, real time germination, and contact time germination when dry dehulled dormant seeds are exposed 75 mM n-propanol/ 25 mM citrate/phosphate buffer pH 7.0 168

Embryo [Fru 2,6-PJ, real time germination, and contact time germination resulting from a 24 h/30°C pulse of 75 mM n-propanol/25 mM citrate/phosphate buffer at pH 7.0 as in figure 4.3 and 4.5. 169

Embryo [Fru 2,6-PJ, real time germination, and contact timegermination as a result of a 24 h/30°C pulse of 40 mM propionaldehyde/ 25 mM citrate/phosphate buffer at pH 7.0 as in figure4.3 and 4.5. 170

Embryo [Fru 2,6-PJ, real time germination, and contact timegermination resulting from a 24 h/30°C pulse of 22 mM propionate/25 mM citrate/phosphate buffer at pH 4.9 as in figure 4.3 and 4.5. 171

viii

E.12 Embryo [Fru 2,6-PJ on plateaux during chemical contact vs. time to30 and 40% germination as in figure 4.4 . 172

F .l Relative signal intensities (RSI) and carbon number of metabolitesignals in perchloric acid extracts and germination in real time

of propionate-2-13C labelled embryos in figure 5.2 and 5.3. 174

F.2 Total relative signal intensities (TRSI) of the combined relative signalintensities of the signals from C2 of both 3-hydroxypropionate-2-13C and propionate-2-,3C in figure 5.3B. Data from Table F .l. 175

F.3 Relative signal intensities (RSI) and carbon number of metabolitesignals in perchloric acid extracts and germination in real time of n- propanol-l-13C labelled embryos (100.6 MHz in 85% H20/15% D20) in figure 5.5 and 5.6. 176

F.4 Relative signal intensities (RSI) and carbon number of metabolitesignals in perchloric acid extracts propionate-2-13C labelled endosperms. 177

G. 1 31P NMR assignments (ppm) of the presumed P; signal in perfusedred rice embryos in a single pulse spectra (Fig. 6.4) and in spin echo spectra during exposure to nitrite (Fig. 6.5), alkaline pH (Fig. 6.6). 179

ix

LIST OF FIGURES

FIGURE PAGE

2.1 The hydration phases of seeds broken down into those phases seen in A) nondormant (ND) seeds, and B) those proposed to occur in dormant(D) seeds as a result of a dormancy-breaking treatment. 12

3.1 A) Moisture content (fresh weight basis) of embryo and endospermtissue from dehulled red rice seeds during hydration at 30°C. B) Embryo pH of dehulled red rice seeds during hydration, and germination (%) of nondormant seeds. (C) Endosperm pH of the same seeds. 29

3.2 Response of embryo and endosperm pH and germination of dormant dehulled red rice seeds to 10 mM nitrite 4h/30°C, followed by transferto H20 12h/30°C. 31

3.3 Embryo pH of dormant dehulled red rice during contact with membersof the propionic acis series. Germination percentages shown are after 7d/30°C on H20 following increasing periods of contact with each dormancy-breaking compound under the same conditions. 33

3.4 Endosperm pH of dormant dehulled red rice during a 24h/30°C pulse with a dormancy-breaking compound at A) pH 4.9, and B) pH 7.0 (asin Fig 3.3.). 35

3.5 Embryo pH of dormant dehulled red rice and percent germinationduring 24h/30°C on H20 following a 24h/30°C pulse with each dormancy-breaking compound as in Fig 3.3. 37

3.6 Embryo pH vs. germination (%) of dormant dehulled red rice during24h/30°C on H20 following a 24h/30°C pulse with a dormancy- breaking compound (data from Fig 3.5.). 39

3.7 Endosperm pH of dormant dehulled red rice during 24h/30°C on H20 following a 24h/30°C pulse with a dormancy-breaking compound at A)pH 4.9, and B) pH 7.0 (as in Fig 3.5.). 40

3.8 A) pH titration curves of filtered embryo and endosperm homogenates of dormant red rice. B) Buffer capacity of embryo ‘sap’ calculated from (A). 42

4.1 A) Respiration of dehulled dormant and nondormant seeds over 24 h/30°C. B) Fru 2,6-P2 content of embryos in dehulled dormant and nondormant seeds during hydration at 30°C. C) Utilization of pericarp splitting, radicle and shoot emergence as the criteria for % visible germination of nondormant seeds in Fig. IB.

4.2 Correlation between embryo [Fru 2,6-PJ and seed respiration.Data from Fig. 4.1.

4.3 Dormant seeds hydrated for 24h/30°C, then exposed to each dormancy- breaking chemical. A) Dormancy-breaking after increasing periods of contact. Buffer controls represent 0 hours. B) Fru 2,6-P2 content of embryos in dehulled dormant seeds during and post-chemical contact. C) Germination (pericarp splitting) in real time of seeds in Fig. 4.2B.

4.4 Correlation between mean plateau [Fru 2,6-PJ and the time to 30% germination. Data are from Fig. 4.3B and 4.3C.

4.5 Correlation between % germination and [Fru 2,6-PJ. Data are from Fig. 4.3B and 4.3C.

5.1 Pulse sequences used for acquisition of 13C NMR spectra. (A) Broad band proton decoupling, and (B) gated proton decoupling. Abbreviations: AQ, acquisition; PW, pulse width; RD, recycle delay.

5.2 13C NMR spectra of perchloric acid extracts. Time course of propionate-2-13C metabolism in red rice embryo during (0 to 24 h) and post-chemical contact (28 to 36 h).

5.3 (A) Visible germination percentages after 7 d/30°C following increasing periods of contact with propionate and visible germination percentages in real time. (B) Time course of signal intensities (arbitrary units) of 13C-resonances from citrate-2,4-13C, 3-hydroxypropionate-2- 13C, propionate-2-13C, and an unidentified resonance at 32.1 ppm.

5.4 Total relative intensity of the signals from C2 of both 3- hydroxypropionate-2-13C and propionate-2-13C during and post chemical pulse. Data from Fig. 5.3B.

5.5 13C NMR spectra of perchloric acid extracts. Time course of n- propanol-l-13C metabolism in red rice embryos during chemical contact (0 to 24 h).

59

61

62

64

65

77

79

80

81

82

xi

(A) Visible germination percentages after 7 d/30°C following increasing periods of contact with n-propanol and visible germination percentages in real time. (B) Time course of relative signal intensities (arbitrary units) of 13C-resonances from 3-hydroxypropionate-l-13C and n-propanol- 1-13C.

Airlift perfusion system used to acquire 31P NMR spectra of red rice embryos.

(A) Single pulse spectra (64 scans) and (B) spin echo spectra (64 scans) of phosphate/phytate solution at pH 7.0. The inorganic phosphate peak is (a), all other peaks are phytate signals.

(A) Single pulse spectra (127 scans) and (B) spin echo spectra (352 scans) of CDTA amended embryo sap. The inorganic phosphate peak is (a).

(A) Single pulse spectra and (B) spin echo spectra of perfused red rice embryos.

Spin echo spectra of perfused red rice embryos following increasing contact with 20 mM NaN02 at pH 2.2. Spectra represent, (A) control,(B) 5 min contact, (C) 30 min contact, (D) 60 min contact. Control peak (a); new emerging peak (b).

Spin echo spectra of perfused red rice embryos following increasing periods at pH 8.3. Spectra represent, (A) 5 min contact, (B) 15 min contact, (C) 30 min contact. Peak produced by acidification in figure6.5 is (a); peak produced by treatment at pH 8.3 is (b).

Spin echo spectra of perfused red rice embryos following the addition of 20 mM inorganic phosphate (a) at pH 7.9.

Hydrated dormant seeds are exposed to a dormancy-breaking chemical pulse (A). Following dormancy-breaking visible germination is seen (B). Embryos are acidified during chemical contact (C) and during visible germination (D). Embryo Fru 2,6-P2 levels increase in response to some chemicals (E) and does increase with visible germination (F). The higher the embryo [Fru 2,6-PJ the shorter the germination phase and the greater the subsequent rate of germination.

Embryo and endosperm pH of dehulled seeds hydrated for 6h/30°C, and germination (7d/30°C) after increasing periods of dry afterripening at room temperature of dormant seeds.

Comparison of embryo and endosperm pH of dehulled seeds from four separate harvests of dormant red rice.

ABSTRACT

Hydrated, dehulled, dormant seeds of red rice (Oryza sativa L.) were exposed

to chemical treatments (nitrite, propionic acid, methyl propionate, propionaldehyde,

and n-propanol) that saturate the dormancy-breaking response. Dormancy-breaking

chemicals were (a) metabolized (n-propanol and propionate) by embryos in dormant

seeds to weak acids; (b) decreased embryo pH; and (c) increased embryo [Fru 2,6-P2]

prior to dormancy-breaking. Embryo acidification, but not increased embryo [Fru 2,6-

P2], was associated with the chemical contact interval required for the onset of

dormancy-breaking. During chemical contact, embryo [Fru 2,6-P2] increased

independent of dormancy-breaking and was inversely correlated with the elapsed time

to 30% germination. On subsequent transfer to H20 , further embryo acidification and

increased embryo [Fru 2,6-PJ were significantly correlated with visible germination

irrespective of the dormancy-breaking chemical employed. These data suggest that

dormancy-breaking chemicals lacking a dissociable proton are metabolized to weak

acids, leading to embryo acidification that results in dormancy-breaking; increased

embryo [Fru 2,6-PJ is related to the subsequent germination rate. Embryo

acidification may be analogous to that associated with the termination of

developmental arrest in other multicellular systems (Brine shrimp and nematodes).

In hydrating dormant and nondormant seeds, seed respiration, embryo pH and

[Fru 2,6-PJ were similar. In phase 2, these parameters diverge as nondormant seeds

enter the germination phase. In dormant seeds, embryo [Fru 2,6-PJ was transient in

phase 2 suggesting the imposition of a block on the germination phase.

xiv

CHAPTER 1

SO YOU WANT TO BREAK DORMANCY?

CROSS-KINGDOM SIMILARITIES IN CHEMICAL

TREATMENTS

Introduction

To survive adverse conditions many species enter an arrested state; this may

be metabolic arrest as seen in diving or hibernating animals (Hochachka and Guppy,

1987) or developmental arrest. Organisms entering developmental arrest generally do

so before attaining their adult form. Developmental arrest is seen in the egg (Arbacia

sp.) (Epel, 1989), blastocyst (Capreolus capreolus L.) (Renfree, 1978), cyst (Artemia

sp.) (Drinkwater and Crowe, 1987), larval (Haemonchus contortus Cobb) (Petronijevic

et al., 1986), pupal (Sarcophaga crassipalpis Macquart) (Denlinger et a l., 1980), seed

(Oryza sativa L.) (Cohn, 1989), and spore (Phycomyces blakesleeanus Burgeff)

(Thevelein et al., 1979) stages. However, developmental arrest is not obligatory

(Sussex, 1978).

What then is developmental arrest? Developmental arrest occurs when the

organism enters a form resistant to adverse conditions. Once development is arrested,

metabolism is reduced to minimal levels in order to retain viability. Generally, an

ametabolic state does not occur; slow metabolism can be detected in both dry and

1

hydrated systems (Bewley and Black, 1982; Hand and Gnaiger, 1988). Development

is not resumed when optimum environmental conditions return. An activating stimulus

is required to terminate developmental arrest (i.e., break dormancy). This activating

stimulus is not required for further growth after dormancy is broken.

The nature of dormancy has been reviewed in a number of model systems such

as eggs (Loeb, 1913; Epel, 1989), insects (Jungreis, 1978), seeds (Bewley and Black,

1982; Simpson, 1990), and spores (Sussman and Halvorson, 1966). The events during

and after the termination of developmental arrest are best understood in sea urchin

(Epel, 1990) and South African toad eggs (Charbonneau and Grandin, 1989). From

the large number of studies, it is possible to discern commonalities in activating

treatments, especially for chemical treatments.

Early development o f chemical based dormancy-breaking

treatments

Towards the end of the 19th century, the properties of chemicals capable of

perturbing biological systems attracted increasing attention. Acid toxicity in fungal

spores was related to the presence of the undissociated acid (Clark, 1898). Cell

permeability to weak bases was equated to their degree of dissociation and

lipophilicity (Harvey, 1911). Alcohol partition coefficients (lipophilicity) and

anaesthetic potency were found to increase with carbon number (Overton, 1901). At

the same time, developmental biologists recognized that the above properties affected

the activity of chemicals (Loeb, 1913) that terminated developmental arrest.

Common features of chemical based dormancy-breaking treatments

By surveying dormancy-breaking studies on a cross-kingdom basis, it has been

possible to identify a large number of dormancy-breaking chemicals and a surprisingly

diverse number of model systems, some of which are presented in Table 1.1. From

this survey, the common structural features of dormancy-breaking chemicals can be

identified. In red rice, consideration of these features has been found to increase both

the successful use and number of dormancy-breaking chemicals. Lipophilicity plots

have been especially useful for predicting the concentration range for dormancy-

breaking activity of untested chemicals (Cohn et al., 1989; Cohn et al., 1991).

The pH effect: A number of reports have shown that inorganic acids at pH 2

break dormancy (Loeb, 1913; Lillie, 1926; Sibilia, 1930; Bingham and Meyer, 1979;

Adkins et al., 1985; Petronijevic et al., 1986). Whether this is a result of acid

scarification/mechanical injury or acid loading is not clear.

Contact time: The application of dormancy-breaking chemicals as a pulse is

more effective than continuous contact (Loeb, 1913; Lillie, 1926; Zagorski and

Lewak, 1984; Cohn and Hughes, 1986). This is almost certainly due to the

detrimental effect on normal development of prolonged exposure to dormancy-

breaking chemicals (Loeb, 1913; Lillie, 1926; Mayer and Evenari, 1953)

Table 1.1. Chemicals showing dormancy-breaking activity and the kingdoms in which they are found to be active. In each kingdom, the number designates a species where the appropriate chemical shows activity. The chemicals and species identified are representative only; other examples can be found in the cited literature. Species are identified in the species key below. Within each kingdom, species are arranged by phyla.

KingdomChemical Monera Protista Fungi Plantae Animalia

AlkanesButane 2Pentane 2 10Hexane 2 9,10Heptane 10Isooctane 10Cyclohexane 10AlkenesEthylene _ 10,11,13,14Propylene 2 13,14Propadiene 141-Butene 21-Hexene 10Monocarboxylic acidsMethanoic acid 2 3 7 4,12,15Ethanoic acid 2 2,3 3 1,3,7 6,12,13,15Propanoic acid 2 3 7 12,152-Propenoic acid 3Butanoic acid 2 3 1,2,7 12,13,15Isobutanoic acid 3 7 12Isopentanoic acid 3 7 12Pentanoic acid 2 3 7 12,15Hexanoic acid 2 3 7 12,15Isohexanoic acid 12Heptanoic acid 15Octanoic acid 15Nonanoic acid 15Decanoic acid 15Palmitic acid 2Oleic acid 1,2Linoleic acid 1,2 10Linolenic acid 1Dicarboxylic acidsFumaric acid 7Malonic 7Oxalic acid 7 6,15Succinic acid 7 7Tricarboxylic acidsCitric acid 7 15

ChemicalTable 1.1. cont.

Monera Protista Fungi Plantae Animalia

HydroxyacidsGlycollic acid 7Ascorbic acid 3,4Lactic acid 7 12/3-Hydroxybutyric acid 7 15AldehydesFormaldehyde 6Acetaldehyde 3,7Propionaldehyde 7Pentanal 2Hexanal 1,2 152,4-Hexadienal 1Heptanal 2,7Octanal 1,2,7 15Nonanal 2,7Decanal 2Hexadecanal 1EstersMethyl formate 7Methyl propionate 7Ethyl acetate 7 10Ethyl butyrate 4KetonesPropanone 4 7*,9 9,10Butanone 102-Pentanone 103-Pentanone 10l-Penten-3-one 1,22-Hexanone 6,72-Heptanone 6,7 152-Octanone 5,6,7 15,162-Nonanone 5,6,7 15,16AlcoholsMethanol 3 3,4 7,9,10,11Ethanol 3,4 3,4,5,7,8,11 8,12Propanol 3,4 3,4,5,7,8 12Isopropanol 4 7,11,162-Propene-l-ol 4Butanol 2 3,4 3,4,5,7 10,12Isobutanol 7“Pentanol 2 3,4 5,7,8Hexanol 2 3,4 THeptanol 3,4 TOctanol 3,4 T3-Octanol 1l-Octene-3-ol 1Nonanol 72-Nonanol 7 15,16

6

Chemical MoneraTable 1.1. cont.

Protista Fungi Plantae

3-Nonanol 16l-Nonen-3-ol 16AminesHydroxylamine 2 3,7,10Methylamine 1Ethylamine 2Propylamine TButylamineHeptylamine 2Octylamine 2Decylamine 2Dodecylamine 2EthersDiethyl ether 9,11AromaticsBenzeneBenzoic acid 7Benzyl acetate 5,7Benzaldehyde 5,7 16Benzyl alcohol 5,8Benzyl aminePhenol 3Salicylic acid 3 7Salicylhydroxamic acid 15Methyl salicylic acid 5,7 10Salicylaldehyde 7TolueneXyleneXylolInorganicsAmmonia 1 3 3,7% 12Azide 3,7,10Carbon dioxide 1,2,3,4,5,6 5,7,9Cyanide 7,10,13Hydrogen sulphide 5 TNitrate 2 2,3 3,10Nitrite 3,7,10Nitric oxide 5Nitrogen dioxide 7

Animalia

55

5,13

8 , 10,11

1012,15

5,13

12

10108,11

2,3,4,5,13

2,3,4,7,12,13,14,1512,14412

* unpublished data

7

Key to species:MONERASchizophyta1. Bacillus cereus Frankland & Frankland Endospore2. Bacillus megaterium de Barry Endospore

3. Thermoactinomyces vulgaris Tsiklinsky Endospore

PROTISTASarcomastigophora1. Giardia muris Grassi Cyst2. Hartmannella rhysodes Singh Cyst3. Schizopyrenus russelli Singh Cyst

Apicomplexa4. Eimeria bovis Zublin Cyst5. Eimeria stiedai Lindemann Cyst6. Eimeria tenella Railliet & Lucet Cyst

Ciliophora7. Pleurotricha lanceolata Ehrenberg Cyst

FUNGIAmastigomycota1. Alternaria alternata Fries Conidiospore2. Fusarium solani Mart Conidiospore3. Phycornyces blakesleeanus Burgeff Conidiospore

4. Neurospora tetrasperma Shear 8c Dodge Ascospore5. Puccinia helianthi Schw. Uredospore6. Uromyces vignae Barcl. Uredospore7. Uromyces rumicis Schum. Uredospore

PLANTAEPhaeophycophyta1. Fucus vesiculosus L. Egg2. Sargassum piluliferum C. Agardh Egg

Anthophyta3. Avena fatua L. Seed

4. Avena sativa L. Seed5. Echinochloa crus-galli (L.) Beauv. Seed6. Hordeum vulgare L. Seed7. Oryza sativa L. Seed

8. Panicum capillare L. Seed9. Panicum dichotomiflorum Michx. Seed

10. Amaranthus albus L. Seed

(Preston and Douthit, 1988)(Levinson and Sevag, 1953; Rode and Foster, 1961 & 1965)(Kirillova et al., 1974)

(Schaefer et al., 1984)(Datta, 1979)(Datta, 1979)

(Jensen et al., 1976)(Jensen et al., 1976)(Nyberg et al., 1968; Jensen et al., 1976)

(Jeffries, 1956 & 1962)

(Harman et al., 1979)(Harman et al., 1979)(Thevelein et a l., 1979; Thevelein et a l., 1983; Van Mulders et al., 1986) (Belmans et al., 1983)(French, 1984)(French, 1984)(French et al., 1986)

(Overton, 1913)(Hiror and Inoii, 1954)

(Adkins et al., 1984a& b, 1985; Cairns and De Villiers, 1986)(Corbineau et al., 1991)(Taylorson, 1988; Leather et al., 1992) (Hareland and Madson, 1989)(Tseng, 1964; Major and Roberts, 1968; Cohn et al., 1983; Cohn and Castle, 1984; Cohn and Hughes, 1986; Cohn et al., 1987 & 1989)(Taylorson, 1989)(Taylorson and Hendricks, 1979;Taylorson, 1980)(Hendricks and Taylorson, 1974;Taylorson, 1979)

8

11. Amaranthus retroflexus L. Seed

12. Berbera verna (Mill.) Aschers Seed13. Lactuca saliva L. Seed14. Portulaca oleracea L. Seed15. Rumex acetosella L. Seed16. Rumex crispus L. Seed

ANIMALIANematoda1. Bursaphelenchus xylophilus Nickle Juvenile2. Ascaris suum Goeze Juvenile3. Haemonchus contort us Cobb Juvenile

4. Nematospiroides dubius Baylis Juvenile Annelida5. Polynoe sp. Egg6. Thalassema mellita Ranzani Egg

Arthropoda7. Artemia franciscana Kellog Cyst8. Melanoplus differentialis Uhler. Egg9. Manduca sexta Johansson Pupae

10. Sarcophaga crassipalpis Macquart Pupae11. Loxostege sticticalis L. pupae Echinodermata12. Asterias forbesii Desor Egg13. Arbacia sp Egg

14. Paracentrotus lividus Lamark Egg15. Strongylocentrotus purpuratus Stimpson Egg

(Taylorson, 1979 & 1989; Schonbeck and Egley, 1980) (Hendricks and Taylorson, 1974) (Brooks et al., 1985; Abeles,1986) (Taylorson, 1979)(French and Leather, 1979)(French and Leather, 1979;Taylorson, 1984; French et al., 1986)

(Matsumori et a l., 1989) (Petronijevic and Rogers, 1987a) (Petronijevic et al., 1986; Petronijevic and Rogers, 1987a & b)(Petronijevic et al., 1986)

(Loeb, 1913)(Loeb, 1913)

(Drinkwater and Crowe, 1987) (Slifer, 1946)(Denlinger et a l., 1980)(Denlinger et al., 1980)(Pepper, 1937)

(Lillie, 1910, 1913, 1926 & 1927) (Lyon, 1903; Loeb, 1913; Harding, 1951)(Lyon, 1903; Loeb, 1913)(Loeb, 1913)

Dissociation constant: The pH-dependent response to acidic or basic

dormancy-breaking chemicals is related to their pKs. Dormancy-breaking activity

requires the presence of the uncharged species. This has been recognized in several

model systems (Loeb, 1913; Lillie, 1926; Toole and Cathey, 1961; Palevitch and

Thomas, 1976; Van Mulders et al., 1986; Cohn and Hughes, 1986; Cohn et al.,

1987; Petronijevic et al., 1986; Petronijevic and Rogers, 1987a). This dependence

upon pH for activity is reflected in weak acid uptake (Van Mulders et al., 1986).

Lipophilicity: In several model systems the activity of dormancy-breaking

activity of organic acids and their derivatives is correlated with their lipophilicity

(Loeb, 1913; Thevelein et al., 1979: Bel mans et al., 1983; Taylorson, 1988; Cohn

et al., 1989). This correlation is modified by a functional group effect, with acids

being more active (Cohn et al., 1989).

Molecular size: The activity of some compounds (eg. azide and formic acid)

appears to be related to molecular size rather than lipophilicity (Cohn et al., 1989).

It is somewhat disappointing that the features identified by Loeb (1913), i.e.,

contact time, dissociation constant, and lipophilicity, have taken so long to be

recognized in other model systems.

10

Is there a mechanistic basis for the parallels observed?

Of the dormant forms shown, the majority are activated by representatives

from many of the chemical classes shown in Table 1.1. A notable exception seems to

be the alkanes which appear to have no activity in seeds (Taylorson, 1979; Abeles,

1986; Cohn et al., 1989). It would at first appear strange that such a wide array of

chemicals should be active in species as diverse as Bacillus megaterium de Barry,

Oryza sativa, and Sarcophaga crassipalpis Macquart (Table 1.1). However, this can

be accommodated if the same, or similar dormancy-breaking mechanisms operate in

all five kingdoms. Evidence is presented herein that dormancy-breaking chemicals act

via a common mechanism in red rice.

CHAPTER 2

DORMANCY-BREAKING IN SEEDS

Introduction

When dry, viable, nondormant seeds are hydrated, they recommence

development and ultimately germinate. During this process, water uptake kinetics can

be used to denote three distinct phases (Fig. 2.1 A). Phase I is the physical process of

imbibition. Phase II is a lag phase in water uptake. In nondormant seeds, phase II also

represents the germination phase, during which the metabolic events required for

visible germination occur. Phase III marks an increase in water uptake, coincident

with visible germination (classically denoted as radicle emergence). This phase marks

the end of the germination process and the onset of seedling growth. Dormant seeds

enter phase II in terms of water uptake, and metabolism is active. But, they do not

progress to the visible germination phase (Bewley and Black, 1978).

In the case of red rice (Oryza sativa), the main difference between dormant

and nondormant seeds is that the germination phase is arrested in the former. In

hydrated, dormant seeds, phase II can be theoretically split into three overlapping

phases during a dormancy-breaking chemical treatment (Fig. 2. IB). The length of a

dormancy-breaking treatment is the initial phase, during which a finite period is

required for seeds to perceive the dormancy-breaking treatment. Following perception,

11

12

HYDRATION PHASES

II III

ANONDORMANT

IMBIBITION GERMINATION PHASE GROWTH

_________ ^ _____DORMANT

..... ...B

IMBIBITION DORMANCY-BREAKINGTREATMENT

D/ND TRANSITION

GERMINATION PHASE GROWTH

TIME

Figure 2.1. The hydration phases of seeds broken down into those phases seen in A) nondormant (ND) seeds, and B) those proposed to occur in dormant (D) seeds as a result of a dormancy-breaking treatment.

13

the dormancy-breaking process is initiated, and the seed enters the

dormant/nondormant transition. This transition may occur during or following the

dormancy-breaking treatment. Once this transition has occurred, seeds are committed

to the germination phase. The germination phase is terminated by visible germination

and is followed by seedling growth.

Physiological dormancy

Dormant seeds are in a state of developmental arrest, providing them with a

means to survive adverse conditions. The red rice used in this study exhibits

physiological dormancy as opposed to that resulting from a) an impermeable seed

coat; b) mechanical restraint of the embryo; or c) embryo immaturity as seen in many

other species (Bewley and Black, 1982). The term, physiological dormancy, may be

interpreted as that resulting not from the effects of seed structure, but from the

consequences of a developmental program.

Physiological dormancy can be broken in response to environmental (Bewley

and Black, 1982), physical (Sung et al. 1987) and chemical agents (Bewley and Black

1982; Cohn, 1989; Cohn et al. 1989). Several hypotheses attempt to explain the

dormancy-breaking ability of these agents (Cohn, 1987). In the case of chemical

agents, hypotheses tend to center on specific components of metabolism and/or related

groups of compounds (Roberts and Smith, 1977; Taylorson and Hendricks, 1980;

Esashi et al., 1981a & b).

14

Contemporary dormancy-breaking hypotheses

Based on the dormancy-breaking properties of glycolytic, tricarboxylic acid

cycle, and electron transport inhibitors, Roberts (1969) proposed that dormancy was

maintained by the consequences of respiratory imbalance: the pentose shunt

hypothesis. Operation of the pentose shunt is prevented thus maintaining dormancy.

This is achieved by competition between a high affinity oxidase (cytochrome oxidase)

and low affinity oxidases (unidentified). Operation of the low affinity oxidase is

needed to oxidize the nicotinamide adenine dinucleotide phosphate (NADPH) required

by the pentose shunt. Inhibition of the glycolytic route favors the low affinity oxidase

and operation of the pentose shunt. Carbon isotope discrimination ratios (C6/C,) were

used to evaluate carbon flow in glycolysis vs. the pentose shunt, with low ratios

indicating operation of the pentose shunt and loss of dormancy (Roberts and Smith,

1977). However, the technique is confounded by interchange of the label between

intermediates and dilution by intermediate pools of varying size (Ap Rees, 1980). The

level and activity of pentose shunt enzymes could not be related to dormancy-breaking

(Adkins and Ross, 1981; Upadhyaya et al., 1981; Cairns and De Villiers, 1986).

This was followed by the alternative respiration hypothesis of Esashi (Esashi et al. ,

1981a & b), which was based on the dormancy-breaking action of cyanide and the

requirement for a cyanide-resistant oxidase, as recognized in the pentose shunt

hypothesis. This hypothesis requires a balance between cyanide-sensitive and -resistant

respiration, with dormancy being lost as cyanide-resistant respiration increases with

15

respect to the sensitive pathway. However inhibitors of cyanide-resistant respiration

also break dormancy (Brooks et al., 1985).

The anesthetic release hypothesis was proposed based on the reversal by

increased pressure of the dormancy-breaking activity of alcohols. Such pressure

reversal is a characteristic of anesthetics and is usually taken to indicate the membrane

as a site of action (Taylorson and Hendricks, 1980; Taylorson, 1988). This hypothesis

is flawed in that some secondary alcohols do not break dormancy (Cohn et al., 1991)

despite their anesthetic properties (Alifimoff et al., 1987). These hypotheses attempt

to explain the dormancy-breaking action of specific groups of compounds. They are

not, however, able to address the fundamental question of why a wide range of

apparently unrelated chemicals are able to break dormancy.

Components of metabolism as markers

As metabolic activation occurs in the germination phase (Bewley and Black,

1978), indicators of increased metabolic activity are potential markers for the onset

of the germination phase. In the context of dormancy-breaking chemical treatments,

such markers have potential for identifying the dormant/nondormant transition.

Adenosine triphosphate: In dry wild oat (Avenafatua L.) seeds the adenosine

triphosphate (ATP) concentration is lower in dormant than in nondormant seeds

(Adkins and Ross, 1983). On hydration, differences in developmental state were not

reflected in the [ATP] or energy charge (Adkins and Ross, 1983; Van Larondelle et

al., 1987; Come et al., 1988). Gibberellic acid (Adkins and Ross, 1983) and ethanol

(Van Larondelle et al., 1987) broke dormancy but had no effect on [ATP] over that

in dormant seeds.

Fructose 2,6-bisphosphate: In the germination phase, glycolytic activity is

expected to increase. Glycolysis is stimulated by the signal molecule, fructose 2,6-

bisphosphate (Fru 2,6-Vj). Fra 2,6-P2 acts by stimulating pyrophosphate: fructose 6-

phosphate 1-phosphotransferase (PPr PFK) and inhibiting cytosolic fructose 1,6-

bisphosphatase (Fra 1,6-P2ase) (Van Schaftingen, 1987; Stitt, 1990). PPr PFK specific

activity increases when oxygen is limiting (Mertens et al., 1990; Mertens, 1991),

which may be the case in the dense tissues of a germinating seed. In germinating

seeds of Phaseolus vulgaris L. and Citrullus lanatus Thunb., PPr PFK specific activity

was higher than the phosphofractokinase (PFK) activity, but did not itself increase

prior to visible germination (Botha and Small, 1987; Botha et al., 1989; Botha and

Botha, 1990).

In the germination phase, increased Fra 2,6-P2 levels may reflect increased

glycolytic activity. In dormant tubers and roots, restoration of metabolic activity led

to an increase in [Fra 2,6-PJ (Van Schaftingen and Hers, 1983; Kowalczyk, 1989).

In Phycomyces blakesleeanus spores and seeds of Avena sativa L ., dormancy-breaking

chemical treatments induced a transient rise in [Fra 2,6-PJ (Van Laere et al., 1983;

Van Larondelle et al., 1987). However, in nondormant Avena sativa, a transient

increase was also seen in the germination phase but not coincident with radicle

17

protrusion (Van Larondelle et al., 1987). The [Fru 2,6-PJ increased during Phaseolus

vulgaris germination (Botha and Small, 1987). Clearly, Fru 2,6-P2 has potential value

as a marker for increased glycolysis.

Enzyme activity: Enzyme levels and activities would be expected to increase

in both the germination phase and the dormant/nondormant transition. Enzyme

activities differ between tissue from dormant tubers vs. tissue slices aged in vitro to

induce respiration. In aged tissue, the activity of glycolytic enzymes increased, and

their distribution changed from the soluble to the particulate fraction (Moorhead and

Plaxton, 1988). In permeabilized sea urchin eggs, the activity of glucose-6-phosphate

dehydrogenase increased rapidly following fertilization (Epel, 1989). Similar studies

in seeds may provide important information regarding markers of developmental state.

Markers arising from dormancy-breaking chemical treatments

A number of dormancy-breaking chemicals have been identified for seeds

(Roberts, 1973; Roberts and Smith, 1977; Bewley and Black, 1982) and other model

systems (Chapter 1). These include inorganic and organic weak acids and bases,

alcohols, aldehydes, alkanes, esters, and ketones (Table 1.1). This provokes the

question: how does a dormant seed perceive the presence of a dormancy-breaking

chemical? Alternatively, based on the properties of dormancy-breaking chemicals,

what signals might they generate? Such signals may be used as markers to identify the

point at which the developmental pattern is altered, i.e. the transition from the

dormant to the nondormant state. Identification of a series of markers would allow the

construction of a testable dormancy-breaking hypothesis.

Intracellular pH: Many developmentally arrested systems exhibit a change

in internal pH upon activation. In unicellular systems such as sea urchin and Xenopus

eggs, one of the early events of fertilization and artificial activation is an increase in

intracellular pH (Grainger et al., 1979; Whitaker and Steinhardt, 1982; Busa and

Nuccitelli, 1984; Charbonneau and Grandin, 1989; Epel, 1989). By contrast,

intracellular pH decreases upon activation of dormant multicellular systems, such as

embryos of the brine shrimp, Artemia franciscana Kellog (Drinkwater and Crowe,

1987), and larvae and juveniles of the nematodes, Caenorhabditis elegans Daugherty

and Haemonchus contortus (Petronijevic and Rogers, 1987b; Wadsworth and Riddle,

1988). In plants, the intracellular pH of dormant buds of Jerusalem artichoke tubers

is higher than in nondormant buds (Gendraud and Lafleuriel, 1983). These

observations identify changes in intracellular pH as a potential marker for the change

in developmental pattern.

In red rice, the most active dormancy-breaking chemicals are inorganic and

organic weak acids. Their common feature is the dissociable proton (Cohn, 1989;

Cohn et al., 1989). Movement of the undissociated acid into the model system of

interest, where it presumably dissociates, will decrease the internal pH. This

implicates changes in tissue pH with dormancy-breaking. If so, dormancy-breaking

19

chemicals lacking a dissociable proton may only act on conversion to the parent acid

(Cohn et al., 1991).

By identifying a series of markers during and following a dormancy-breaking

chemical treatment, a time-line of events can be constructed. The position of events

on the time-line will suggest other associated events as markers. Hence, the time-line

will also fulfill a predictive function. Ultimately, this approach will make it possible

to unravel the events that make up the dormancy-breaking process and identify the

point of transition between the developmental states. Such a strategy is deemed more

productive, flexible and less prone to bias as it is an approach driven by observation,

rather than by hope.

In the following chapters, the dormancy-breaking activity of chemicals with

different functional groups is reported. By use of conditions that saturated the

dormancy-breaking response, it was possible to identify several events occurring

during a dormancy-breaking chemical treatment. This in turn may help explain why

a large number of chemicals show cross-kingdom dormancy-breaking activity.

CHAPTER 3

EMBRYO ACIDIFICATION DURING DORMANCY-BREAKING

AND SUBSEQUENT GERMINATION

Introduction

Dormancy-breaking chemicals of seeds are predominantly weak acids or their

derivatives (Bewley and Black, 1982; Cohn et al., 1987; Cohn, 1989; Cohn et al.,

1989). Weak acids also terminate developmental arrest in other model systems, such

as eggs of the alga (Fucus vesiculosus L.) (Overton, 1913), sea urchin

(Strongylocentrotuspurpuratus Stimpson) (Loeb, 1913), starfish eggs (Asteriasforbesii

Desor) (Lillie, 1926), Phycornyces blakesleeanus spores (Van Mulders et al., 1986),

and juvenile nematodes (Petronijevic et al., 1986). The mechanism by which these

compounds act is unclear. However, weak acids have been demonstrated to induce cell

acidification (Felle, 1989; Petronijevic and Rogers, 1987; Roos and Boron, 1981).

In the animal kingdom, many developmentally arrested systems exhibit a

change in internal pH upon activation. In unicellular systems such as the sea urchin

egg, internal pH increases during fertilization and artificial activation (Epel, 1990;

Shen, 1982). In multicellular systems, internal pH decreases as a result of activation

in embryos of the brine shrimp, Artemia salina L. (Crowe et al., 1987), as well as

20

21

in larvae and juveniles of the nematodes, Caenorhabditis elegans (Wadsworth and

Riddle, 1988) and Hacmonchus contortus (Petronijevic and Rogers, 1987).

Changes in internal pH upon activation of developmentally arrested systems

in the plant kingdom have been neglected, in comparison to those in animal systems.

In the older seed literature there are several reports of changes in seed tissue pH

during the dormancy-breaking process and germination. During cold stratification,

embryo pH decreased in dormant seeds of Crataegus gloriosa Sargent (Eckerson,

1913), Acer saccharinum L. (Jones, 1920), Juniperus virginiana L. (Pack, 1921), and

in whole seeds of Tilia americana L. (Rose, 1919) and Heracleum sphondylium L.

(Stokes, 1953). As a result of dry afterripening, embryo pH also decreased in dormant

Avena fatua (Atwood, 1914). Embryo pH decreased during germination (Eckerson

1913; Pack, 1921; Rose, 1919); similar observations were also made in Cornus florida

L. (Davis, 1927) and Spinacia oleracea L. (Sifton, 1927). This decrease in embryo

tissue pH during germination appeared to be initially located in the outer layer of

embryo cells (Davis, 1927; Pack, 1921) and in the sieve tubes (Davis, 1927). The role

of internal pH, as regards the transition from the dormant to the nondormant state, has

not been rigorously addressed in seeds since these early studies. More recently, in

Jerusalem artichoke tubers, the internal pH was found to be higher in dormant vs.

nondormant buds (Gendraud and Lafleuriel, 1983). A common theme between

developmentally arrested, multicellular systems of the plant and animal kingdoms

seems to be a decrease in internal pH upon activation. Herein, I report the effect of

dormancy-breaking compounds on embryo pH. It was demonstrated that embryo pH

22

is higher in dormant than in nondormant seeds of red rice. Evidence is also presented

that embryo acidification is a prerequisite for the termination of dormancy. Part of this

chapter has published elsewhere (Footitt and Cohn, 1992).

Materials and Methods

Mature, dormant red rice (Oryza sativa) seeds (strawhulled, awnless) were

obtained from the South Farm, Rice Research Station, Crowley, LA in 1987. Seeds

were harvested by hand shattering of individual plants. Moisture content at harvest

was 17.8%. After drying for two days in open trays at 22°C, the final moisture

content was 12.6%. Seeds were stored in Mason jars at -15°C. Nondormant red rice

was obtained by afterripening dormant seeds at 22 °C for 60 days after which they

were stored at -15°C. Seed moisture content did not change as a result of dry

afterripening. Seeds were manually dehulled immediately prior to sowing or not more

than 14 hours before hand, in which case they were stored in paper packets over

desiccant at -16°C. No difference between these two procedures was seen.

Germination and viability tests were performed with each experiment (buffer capacity

determinations excepted). Freshly prepared solutions were used for each experiment.

All experiments were repeated four times, and errors represent the standard error of

the mean.

23

Homogenate pH measurement

One hundred dehulled seeds were sown on 9 cm Petri plates containing three

sheets of germination paper and 10 mL distilled H20 . Seeds were covered with a

double layer of tissue paper (Kimwipe) to ensure even hydration during incubation at

30°C in darkness. Plates were incubated on an incline. Nondormant and dormant

seeds were sampled during hydration. Germination was also recorded.

In studies with dormancy-breaking chemicals, dehulled seeds were incubated

as above for 24 hours, rinsed from the germination paper, washed copiously with

H20 , and briefly blotted with tissue paper. Germinated seeds (approx. 5%) were

replaced with similarly hydrated dormant ones. Seeds were then transferred to 250 mL

wire clasp storage jars (Heritage Industries, Millville, NJ) containing two sheets of

Whatman N°1 filter paper and 10 mL of test solution. Seeds were covered with tissue

paper, and jars were sealed. Dormancy-breaking compounds were applied as either

a 4 h/30°C (nitrite) or a 24 h/30°C (all others) pulse in 25 mM citrate/phosphate

buffer. Dormancy-breaking chemicals were used at concentrations giving a saturating

response i.e. dormancy was broken in 90% of the population, as determined by

germination after a subsequent 7 d H20 incubation at 30°C. Sodium nitrite (10 mM)

was applied at pH 3.0 (pK 3.3). Propionic acid (20 mM) was applied at its pK of 4.9.

n-Propanol (70 mM) was applied at pH 4.9 and pH 7.0; propionaldehyde (40 mM)

and methyl propionate (30 mM) were applied at pH 7.0. At the end of the chemical

pulse, seeds were transferred to Petri plates containing H20 (as described above) for

24

up to 12 h/30°C (nitrite) or 24 h/30°C (all others). Samples were taken throughout

the chemical pulse and post-pulse phases.

For each pH determination, 100 seeds were washed thoroughly in H20 and

placed on moist tissue paper. Embryos were excised with a scalpel, weighed and

immediately ground to a powder in liquid nitrogen. This powder was homogenized in

a cold glass tissue homogenizer with 1 mL of ice cold H20 that had been purged with

N2 to remove C 02. The homogenate was transferred to a microfuge tube, centrifuged

(16,000 xg for 5 min, Eppendorf 5414) and the supernatant filtered (0.22 /xm filter,

Millipore GSWP 01300) at 4°C. The filtrate was collected in a microfuge tube and

placed in an ice bath. The pH was measured using a cold (1 to 3°C) Ross 8103 pH

electrode (Orion) in conjunction with an Orion EA 920 lonalyzer and chart recorder

(Cole Palmer 8373-20). The pH was recorded when the electrode registered a stable

pH (<0.01 units min'1); this was generally achieved in two to three minutes. The

procedure was repeated using the same weight of endosperm tissue. The pH data were

analyzed according to Stevens (1955). Tissue pH measurements were highly

reproducible. For all time points (n = 168) embryo mean standard errors were

+0.020 ± 0.001, and -0.019 ± 0.001, and endosperm mean standard errors were

+0.029 ± 0.001, and -0.027 + 0.001. Unless otherwise stated the effect of

dormancy-breaking compounds on embryo [H+] was expressed as embryo [H+]

dormancy-breaking treatment minus embryo [H+] control at the same time point.

25

Buffer capacity

One hundred dehulled dormant seeds were hydrated on HaO in Petri plates for

24 h/30°C as above. After 24 h germinated seeds were replaced as described

previously. Filtered embryo and endosperm homogenates were prepared as above, of

which 400 fiL was taken for pH determinations. When a stable pH was obtained, 2.5

HL of ice cold NaOH or HC1 (0.05N) was added and the sample mixed before the pH

was remeasured. This was repeated up to pH 9.0 for alkali titrations and pH 5.0 for

acid titrations, using fresh filtrates in each case. The electrode was not rinsed between

additions. Embryo buffer capacity was determined from the slope between adjacent

points on the titration curve (aH +/apH ) (Takeshiga and Tazawa, 1989).

Electrode calibration and maintenance

The electrode was calibrated at pH 7.09/3°C, using a chilled, pH 7.0 standard

buffer purged of C02 as above. The electrode was sealed in the buffer using parafilm

to reduce access to C 02. This one point calibration gives an error of 0.0037 pH units

°C'1 at 0°C with a reading one unit from calibration (Westcott, 1978). When checked

against a two point calibration, no significant differences were found between the pH’s

of standards or the slopes. Between measurements the electrode was kept in ice cold

storage buffer (Orion). At the end of each day, the electrode was cleansed of lipid and

protein contaminants by rinsing with 1% detergent (Micro 8790-00, Cole-Parmer),

and soaking in 1% pepsin/100 mM HC1 for 15 min. The reference electrode-solution

was then replaced.

26

Germination tests

Each treatment used five replicates of 20 dehulled seeds in 50 mL Erlenmeyer

flasks with two layers of Whatman N°1 filter paper and 2 mL of test solution. Flasks

were sealed with rubber septum caps. Nondormant seeds were incubated for 7

d/30°C. For studies using dormancy-breaking chemicals, seeds were incubated in

Erlenmeyer flasks as for tissue pH determinations and transferred to H20 for 7

d/30°C, when germination was measured. Germination was measured as rupture of

the pericarp and aleurone over the embryo (criterion used in time course experiments)

or radicle protrusion. Viability of ungerminated seeds was tested by incubation of

excised embryos on H20 at 30°C. In dormancy-breaking experiments seed viability

was 98% (n = 180); for imbibition experiments viability was 97% for dormant and

100% for nondormant seeds. Germination data are presented in the appropriate figure

legends.

Moisture content

The moisture content of four replicates of 100 intact dry seeds and seed

components (hull, embryo and endosperm) was determined. The moisture content of

seed components (embryo and endosperm) during the hydration of dehulled dormant

and nondormant seeds was determined in four replicates of 50 seeds at each time

point. Seeds were incubated in 9 cm Petri plates containing three sheets of

germination paper and 10 mL of H20 . Two samples of 50 seeds were sown in each

plate. Samples were physically separated and each was covered by a double layer of

27

tissue paper. Seeds and components were dried for 7 d at 100°C. Dry material was

placed in a desiccator for 30 minutes prior to the final weighing. Percent moisture

content was determined based on the fresh weight at the time of sampling.

Results

Tissue hydration kinetics

There was no significant change in moisture content as a result of dry

afterripening (Table 3.1). The moisture content of individual seed components was

comparable. In whole seeds, moisture content is contributed mainly by the endosperm

and least by the embryo. During hydration embryo moisture content was the same in

both dormant and nondormant seeds until the onset of germination. Endosperm

moisture contents were the same in both dormant and nondormant seeds (Fig. 3.1 A).

Comparison of dormant and nondormant embryos and endosperms showed no

differences in dry or fresh weight during hydration (Appendix D, Table D .l, D.2,

D.3, D.4). Imbibition was essentially complete after four hours.

Tissue pH during hydration

Before investigating the role of pH during dormancy-breaking, it was essential

to determine if there were differences in tissue pH during the hydration of dormant

and nondormant seeds under conditions leading to germination in the latter.

28

Table 3.1. Moisture content(%) on a fresh weight basis of dry intact nondormant red rice seeds3 and of seed components following drying for 7 days at 100°C. The contribution to whole seed MC of each component is also shown.

Sources of MC(%)b

MC(%) in whole seeds

Intact seeds 12.66 ± 0.15

Seed components

Hull 8.45 ± 0.49 2.03 ± 0.12

Embryo 10.87 ± 0.97 0.21 ± 0.2

Endosperm 13.74 ± 0.08 10.16 ± 0.05

a) Moisture content of dormant seeds was 12.6%.

b) Determined as

1 - (fresh wt comp a + b + dry wt comp c l x 100 Ifresh wt comp a + b + c J

= contribution of component c to whole seed MC(%)

29

60.0

c 40 .0 --

<u 30

20.0- / —I

e* 10.0<>

*11= 1= 1 -□

Embryo o — O Nondormant • — ® Dormant

Endosperm □ — □ Nondormant Dormant

1 1 ! 1-----!—

w 7 .0 -

6 .8 / i A A A

$ : ‘ CL / a s03Q - 7 4(0Oc 7 .3 -I

Ld7.2-1

7.1

1 □—tjl □-

i

H h4 6 8 10 12 14

Hours

Figure 3.1. A) Moisture content (fresh weight basis) of embryo and endosperm tissue from dehulled red rice seeds during hydration at 30°C. B) Embryo pH of dehulled red rice seeds during hydration, and germination (%) of nondormant seeds. Germination after 7 d/30°C on H20 was 8% for dormant and 98 + 1% for nondormant. C) Endosperm pH of the same seeds. Error bars represent one standard error of the mean; no error bar indicates the symbol is larger than the error.

30

The pH of dry dormant and nondormant embryos was 7.37 and 7.25 (Fig.

3. IB). During imbibition embryo pH became stable after two hours in dormant (pH

7.28) and after four hours in nondormant (pH 7.16) seeds. When imbibition was

complete (4 h), the increase in embryo [H+] over that measured at Oh in the respective

embryos was 13.4 (dormant) and 14.4 nM (nondormant). After six hours the embryo

pH of nondormant seeds started to decrease. Germination commenced after 12 h.

Endosperm pH declined during imbibition in both cases, after which nondormant

endosperm exhibited a slow decline (Fig. 3.1C).

The effect of nitrite on tissue pH

Initial experiments on the effects of dormancy-breaking compounds on tissue

pH utilized nitrite, which is highly effective at low concentration over a four-hour

period. As nitrite is a weak acid, it was expected that a decrease in tissue pH would

be easily detected. At an incubation medium of pH 3.0, the pH of dormant embryos

decreased more rapidly and to a greater extent in nitrite solution than in the buffer

control (Fig. 3.2A). The overall drop in embryo pH for nitrite and the control was

0.47 and 0.12 pH units, respectively, during chemical contact between -4 to 0 h. On

transfer to water at 0 h, embryo pH in the control (pH 3.0) recovered to that found

at -4 h.

During the post-pulse phase after nitrite treatment, a slight decline in embryo

pH occurred as germination commenced. Endosperm pH was higher than embryo pH,

31

Control pH 7 .0 Nitrite pH 7 .0

□ Control pH 3 .0 Nitrite pH 3 .0

□ □

Hours

2 0 .0 roXCL

I - - 10 . 0

cooc

0.0

H o u r s

Figure 3.2. Response of embryo and endosperm pH and germination of dormant dehulled red rice seeds to 10 mM nitrite 4 h/30°C, followed by transfer to H20 12 h/30°C. Germination in each treatment after 7 d/30°C on H20 was: pH 3.0 control, 5 + 1%; pH 7.0 control, 4+1% ; nitrite at pH 3.0, 98%; nitrite at pH 7.0, 5 + 1%.

32

and followed a similar pattern (Fig. 3.2B). Nitrite applied at pH 7.0 had little effect

vs. its buffer control on embryo pH and did not break dormancy.

The effect of propionate and its derivatives on tissue pH:

During the chemical pulse

Use of the propionic acid series allowed the comparison of compounds with

and without a dissociable proton that have similar octanol/water partition coefficients

(Cohn, 1989; Cohn et al., 1989). In the pH 4.9 control, embryo pH decreased by

0.08 units after two hours then remained stable (Fig. 3.3A). The pH 7.0 control

increased embryo pH by 0.08 units over 24 h (Fig. 3.3B). Propionate decreased

embryo pH by 0.27 units by 12 h vs. the dormant control, after which the pH

remained stable (Fig. 3.3A). Propanol decreased embryo pH significantly over 24 h,

by 0.05 at pH 4.9 (Table D.9 & D. l l ) and 0.15 at pH 7.0 vs. the dormant controls

(Fig. 3.3B). These data at pH 4.9 were confirmed with seeds from a later harvest

(Appendix D.23). Propionaldehyde at pH 7.0 decreased embryo pH by 0.14 over the

first 8 h vs. the dormant control. Propionaldehyde elicited initial germination 16 hours

into the pulse. Germination was 9% by 24 h. No other compound used in this study

elicited germination during chemical contact. Methyl propionate decreased embryo pH

by 0.18 over the first 8 h v.v. the dormant control. In the case of endosperm tissue,

pH values were higher. Endosperm pH decreased in response to all dormancy-

breaking treatments, but only methyl propionate produced a change in endosperm pH

of similar magnitude to that in the embryo (Fig. 3.4B).

33

Figure 3.3. Embryo pH of dormant dehulled red rice during contact with: (A) 20 mM propionic acid (pH 4.9), B) 70 mM n-propanol (pH 7.0), C) 40 mM propionaldehyde (pH 7.0), and (D) 30 mM methyl propionate (pH 7.0) for 24 h/30°C. Treatment and control media were buffered at pH 4.9 (A), at pH 7.0 (B, C, D) with 25 mM citrate/phosphate. Germination percentages shown are after 7 d/30°C on H20 following increasing periods of contact with each dormancy-breaking compound under the same conditions. Germination of control treatments after 7 d/30°C on H20 was 2 to 3%.

Embr

yo

pH• — • Control O— O Propionate

-100

7 .4 -

■ — ■ Control □ — □ Propanol

7 .2 -

V1

♦i —yo

7 1

□ □i

7 . 1 * = * = *

- 8 0

- 6 0

- 4 0

-20

0

■ — ■ Control O— O Propionaldehyde

T

7 .4

■ — B Control qA A Methyl propionate

T

8 12 16 2 0 2 4 8 12 16 2 0 2 4

Contact t ime in hours

Ger

min

atio

n on

H

2O 7

d/3

0°C

35

O t

X 7 Q_ • * %

E 7 .4L_CL)CL 7 .3 if)O

~D 7 .2 c

Ld7.1 ■

T . .

'O

T1

T-m.

‘o-

T

T'O .

A

‘O

8 12 16 20

Hours

Figure 3.4. Endosperm pH of dormant dehulled red rice during a 24 h/30°C pulse with a dormancy-breaking compound at A) pH 4.9, and B) pH 7.0 (symbols as in Fig 3.3.).

36

To evaluate whether or not embryo acidification was temporally related to the

dormancy-breaking process, initially dormant red rice seeds were exposed to each

chemical for increasing contact periods (up to 24 h) and transferred to H20 for seven

days to assess the extent of germination. Acidification always occurred prior to or

coincident with chemical contact times necessary to elicit a subsequent germination

response (Fig. 3.3).

Post-chemical pulse

During the post-pulse phase, the embryo pH of propionate treated seeds

increased over 12 h (Fig. 3.5A). Embryo acidification occurred coincident with the

onset of visible germination for all members of the propionic acid series except for

propionaldehyde (Fig. 3.5C). Propionaldehyde elicited visible germination during the

pulse phase, which increased rapidly in the post-pulse phase; this was reflected in a

rapid decrease in embryo pH. In the post-pulse phase embryo pH and percent

germination were highly correlated, irrespective of the dormancy-breaking compound

employed (for all data, y = -0.0053x + 7.04; r = -0.63; P < 0.001; for germination

>3%, y = -0.0062x + 7.066; r = -0.92; P < 0.001) (Fig 3.6). Endosperm pH

showed little or no response to germination in the post-pulse phase (Fig. 3.7).

In the post-pulse phase, control germination commenced after 20 h, being 4%

(pH 4.9) and 3% (pH 7.0) after 24 h. As a result embryo pH decreased to 7.04 and

7.12, respectively. In Figure 3.5 the 20 h control points have been omitted due to the

effect of germination on embryo pH. When germinated seeds were replaced by

37

Figure 3.5. Embryo pH of dormant dehulled red rice and percent germination during 24h/30°C on H20 following a 24h/30°C pulse with each dormancy-breaking compound as in Fig 3.3.

Embr

yo

pH7 .2

• — • Control O— O Propionate

7.1 V7 .0 -

6 .9 -

6.8<fc

T /

t. 9pr/9

O T

1H— -fe” Control □ — □ Propanol7 .3 :

g7 -2 ;

7.1

7 .0

6 .9

' a□i

6.8<>++ — ♦ —""T

' ^ 6.♦ 9

T♦fX

5 0

40

3 0

20

10

060

50

40

30

20

10

0

7 .3Control

O — O Propionaldehyde

V

7 .3 :

■ -----■ ControlA -----A Methyl propionate D ..

•60

■50

•40

-30

■20

•10

060

8 12 16 2 0 2 46.84**=#

♦

12 16 2 0 2 4

Hours postcontact u>oo

♦ %

Ger

min

atio

n

39

Cl

OX

_QE

LU

O Propionate □ Propanol O Propionaldehyde A Methyl propionate

20 30 40% Germina t i on

Figure 3.6. Embryo pH vs. germination (%) of dormant dehulled red rice during 24 h/30°C on H20 following a 24 h/30°C pulse with a dormancy-breaking compound (data from Fig 3.5.).

40

8 12 16

H o u r sFigure 3.7. Endosperm pH of dormant dehulled red rice during 24 h/30°C on H20 following a 24 h/30°C pulse with a dormancy-breaking compound at A) pH 4.9, and B) pH 7.0 (as in Fig 3.5.).

41

identically treated dormant ones at 24 h, embryo pH was 7.16 and 7.18, respectively

(Fig. 3.5). The effect of 3 to 4% germination on embryo pH is as great as some of

the changes seen during the pulse period. Thus, it is important to use populations

where all germinating seeds/seedlings have been removed from the controls.

Buffer capacity of embryo and endosperm

The slope of the endosperm titration curve (Fig. 3.8A) indicated a low buffer

capacity. The embryo titration revealed a shallow sigmoidal curve. It was assumed

that the buffer capacity of the original 1 mL of homogenate was the same as that of

embryos prior to extraction. On this basis, embryo buffer capacity was determined

based on the aqueous volume of 100 hydrated embryos (36 /xL), including both

apoplastic and symplastic contributions. Embryo buffer capacity is lowest in the pH

range found during embryo pH measurements (Fig. 3.8B).

By using the equation of the embryo titration curve (Table 3.2), an estimate

of the gross changes in embryo [H+] required to give the observed net changes could

be determined. In the case of permeant weak acids, differing gross changes in embryo

[H+] produced almost identical net changes in embryo [H+]. At pH 7.0, propionate

derivatives produced similar gross and net changes in embryo [H+].

42

0.75

0.50TDCD

TDTDO

+

0.25

0.00X(0 - ° - 25 0)o -0 .50

E -0 .75

H Embryo a Endosperm

- 1.00

-1 .255.04.0 6.0 7.0 8.0 9.0

pH

Q_OCO

oJ Q

ECD

C3

T.Cl

+I

CO_CD

oEE

Figure 3.8. A) pH titration curves of filtered embryo and endosperm homogenates of dormant red rice. B) Buffer capacity of embryo ‘sap’ calculated from (A).

43

Table 3.2. Changes in [H+] of embryo homogenates detected after a 24 hour chemical pulse.

Treatment Gross change in [H+]a Net change in [H+]b

Control pH 3 137.0 /xM 21.0 nM

Nitrite 289.8 /xM 79.4 nM

Control pH 4.9 120.5 /xM 13.4 nM

Propionate 349.7 /txM 73.7 nM

Propanol 53.8 juM 7.4 nM

Control pH 7.0 -100.0 ixM (-58.3 jtxM 8h) -8.2 nM (-5.1 fxM 8h)

Propanol 182.8 nM 16.6 nM

Propionaldehyde 247.6 /xM (168.0 /xM 8h) 24.5 nM (16.8 nM 8h)

Methyl propionate 273.4 ixM (209.1 /xM 8h) 28.0 nM (22.1 nM 8h)

“Gross changes in [H+]: Gross changes were determined by applying the equation of the embryo titration curve (Fig. 3.8A). y = 3.8307 + 4.4287(x) - 3.081 l(x2) + 0.684(x3) - 0.065195(x4) + 0.0022656(x5). Where x = pH; y = /nmoles H +; r = 0.99985. As titration used a 0.4 mL sample, multiply by 2500 to convert from /xmoles to /xM, assuming volume changes during titration have no significant effect.This equation was applied to the embryo homogenate pH data between 0 and 24 h for the controls or between the 24 h points for controls vs. dormancy-breaking chemical treatments.bNet changes in [H+]: Homogenate data were converted from pH to [H+], and the difference over 24 hours (controls) and between the controls and dormancy-breaking chemical treatments at the same time point (24 h) used to determine net changes.

44

Discussion

In this study, embryo acidification was observed during the activation process

stimulated by dormancy-breaking chemicals applied to dormant seeds. The

homogenate method gave highly reproducible pH values comparable with cytoplasmic

pH determinations obtained with other techniques (Kurkdjian and Guem, 1989) that

are not readily adaptable to dense, seed tissues.

The buffer capacity of embryo homogenates was consistent with values

reported for the cytoplasm of Chora corallim Klein and Neurospora (Takeshige and

Tazawa, 1989; Sanders and Slay man, 1982). In dormant seeds the buffer capacity is

highest at extremes of pH, indicating that the pKs of the predominant ionizable groups

are outside the embryo pH range measured. The low embryo homogenate buffer

capacity near neutrality indicated the likelihood of sensitivity to small endogenous

changes in [H+]. This indicated that in dormant embryos, tissue pH may require active

control if perturbed by dormancy-breaking chemicals. Gross changes in embryo [H+]

clearly reflect the differences in buffering capacity either side of pH 7.0 (ie. pH 3.0

control vs. pH 7.0 control).

Embryo acidification during chemical contact.

Embryo acidification was elicited by direct loading of weak acids. The tissue

pH change induced by nitrite contact was more rapid than that of propionate, and this

may be a function of molecular size related to speed of penetration (Cohn, 1989;

Cohn et al., 1989). However, both compounds increased embryo [H+] by

45

approximately the same amount. Following transfer back to water, embryos did not

fully recover from the acid load imposed; this may partly be due to dissociated acid

trapped in the symplast.

Methyl propionate, /z-propanol, and propionaldehyde treatments also caused

embryo acidification. Incubation at pH 7.0 was used to prevent uptake of any acid

produced by chemical conversion of these derivatives in the incubation media. As such

the observed changes in embryo pH were considered to be due to events internal to

the embryo. These chemicals require relatively broad concentrations to induce small

and consistent changes in embryo [H+]. This indicated that the ability to partition into,

or through a membrane was not the only prerequisite for activity and points to the

importance of the functional group. These derivatives could be internally converted

enzymatically to propionic acid with subsequent weak acid dissociation sufficient to

reduce embryo pH, as has already been demonstrated in yeast, under conditions that

did not increase membrane permeability to H+ (Loureiro-Dias and Santos, 1990).

Circumstantial evidence from structure-activity studies indicates a requirement for

organic acid analogues that can be converted to the parent acid in order to exert their

dormancy-breaking effect (Cohn et a l , 1991). Such a proposal may explain: (a) the

inhibition of the dormancy-breaking action of ethanol by 4-methylpyrazole (an alcohol

dehydrogenase inhibitor) in Avena sativa (Come and Corbineau, 1989); and (b) the

ineffectiveness of some secondary alcohols as dormancy-breaking chemicals of red

rice (Cohn et al., 1991). Neither of these observations, nor the conflicting effects of

increased air pressure (Taylorson, 1991) are consistent with the anesthetic-like

46

(pressure sensitive) model proposed for the dormancy-breaking action of alcohols

(Taylorson and Hendricks, 1980; Taylorson, 1988).

During chemical contact, the onset of embryo acidification occurred prior to

or coincident with the contact interval necessary for subsequent visible germination.

This change in embryo pH may represent an early stage of a signal transduction chain

leading to an alteration of the developmental pattern (germination). Changes in both

internal pH and free intracellular Ca2+ are associated with cell activation in sea urchin

(Epel, 1990) and Xenopus laevis Daudin eggs (Charbonneau and Grandin, 1989).

Decreasing intracellular pH led to increased cytosolic Ca2+ in Xenopus embryos (Rink

et al. , 1980), Riccia fluitans L. rhizoids, and Zea mays L. root hairs (Felle 1988). In

Onoclea sensibilis L. spores, increasing intracellular free Ca2+ by use of the

ionophore A23187 or red light increased dark germination (Wayne and Hepler, 1984;

1985). While no change in pH; was detected via nuclear magnetic resonance in this

phytochrome response (Wayne et al., 1986), the standard errors were greater than the

magnitude of changes seen in red rice. In dormant red rice seeds, pH may be acting

as part of a secondary messenger system involving calcium (Felle, 1989).

While controls at pH 3.0 and 4.87 did not break dormancy, embryo pH

declined to a similar or greater extent than did the propionic acid derivatives and the

pH 7.0 control. Following transfer from buffer back to H20 embryo pH partially

recovers. This suggests than at least part of the effect seen in the controls results from

buffer in the free space (apoplast). Therefore, homogenate pH values obtained from

chemically-treated seeds should not be regarded as absolute and can only be

47

meaningfully viewed as indicators of change in relation to respective buffer controls

at the same pH. Analogous, still incompletely explained observations are seen during

egg activation (Epel, 1990).

Embryo acidification and moisture content during germination.

Water uptake by embryos in dehulled nondormant seeds showed the typical

triphasic increase in moisture content associated with seed germination (Bewley and

Black, 1982). Embryos in both dehulled dormant and nondormant seeds showed no

significant differences in moisture content before germination (splitting of the pericarp

and aleurone)(Fig. 3.1 A; Appendix D, Table D .l, D.2). If radicle emergence is taken

as the marker for germination (as in Raju et al., 1988), significant increases in the

moisture content of nondormant over dormant embryos would occur prior to

germination.

During hydration, differing changes in the embryo pH of dormant and

nondormant seeds reflected their different developmental patterns (Fig. 3.1). During

imbibition identical hydration-dependent increases in embryo [H+] occurred.

Following imbibition (> 4 h) the embryo pH of dormant and nondormant seeds was

significantly different. In nondormant seeds, metabolic activation occurs in phase two

(Bewley and Black, 1982), and embryo pH also decreased. This was interpreted as a

physiological event: a marker for the onset of the germination process before it was

physically expressed. In phase three, cell expansion of the radicle leads to germination

(Bewley and Black, 1982). Nondormant embryo moisture content increased

48

significantly (12 h) with splitting of the pericarp and aleurone but before radicle

emergence. Embryo pH continued to decrease as germination commenced. In dormant

seeds, embryo moisture content remained in phase two, and embryo pH remained

stable.

Dormant seeds in contact with propionaldehyde begin to visibly germinate

before transfer to H20 . In the case of the remaining compounds, the onset of visible

germination is delayed during chemical contact. Such a delay was also observed in

nondormant Phacelia tanactifolia Benth. seeds after contact with butyric acid (Cocucci

et al., 1989). On transfer to H20 , the onset of germination is coincident with embryo

acidification (Fig. 3.5). When figure 3.5 was replotted as embryo pH vs. %

germination for the combined data, embryo pH and germination were significantly

correlated (Fig. 3.6). This indicates that changes in embryo pH are associated with

the germination process.

Comparison of pH changes between the embryo and endosperm were difficult

due to differences in bulk. Although endosperm tissue of equivalent weight to that of

the embryo was used, the rate of penetration and leaching out of the test solutions was

likely to be different. Changes in endosperm pH could not be correlated with

dormancy-breaking or germination. Oxidation of dormancy-breaking compounds in

the endosperm that were not subsequently removed on rinsing may have influenced

the changes seen.

49

In summary, it was found that: (a) embryo pH is higher in dormant than in

nondormant red rice seeds, consistent with that found in dicot seeds and the

parenchyma cells of dormant and nondormant buds of Jerusalem artichoke tubers, (b)

In hydrating nondormant seeds embryo pH decreased before visible germination. In

dormant seeds after the chemical pulse, embryo pH decreased coincident with

germination, (c) Dormancy-breaking compounds induced a decrease in embryo pH.

(d) Embryo acidification during dormancy-breaking was consistent with that seen in

the activation of developmentally arrested, multicellular systems in the animal

kingdom. It is proposed that embryo acidification is an indicator of the termination of

developmental arrest in dormant seeds and may play a fundamental role in this

process.

C H A PT E R 4

LEVELS OF FRUCTOSE 2,6-BISPHOSPHATE IN RED RICE

EMBRYOS D U R IN G D O R M A N C Y -B R E A K IN G A N D

G E R M IN A T IO N

Introduction

Embryo acidification occurs during dormancy-breaking chemical treatments

(Chapter 3). This acidification phase can be used as a marker for the dormancy-

breaking process. However, dormancy-breaking events must be separated from those

associated with the germination phase. The germination phase represents the activation

of processes in the embryo leading to but not including morphological changes

(germination sensu stricto). Visible germination marks the termination of the

germination phase. Identification of a biochemical marker for the germination phase

would aid in identification of the dormant/nondormant transition.

When hydrated seeds enter the germination phase, they become metabolically

active (Bewley and Black, 1978), and rates of glycolysis and respiration increase.

Glycolysis is stimulated by the signal molecule, fructose 2,6-bisphosphate (Fru 2,6-

P2). Fru 2,6-P2 is synthesized by 6-phosphofructo-2-kinase (PFK2) and degraded by

fructose-2,'6-bisphosphatase (FBPase2). Fru 2,6-P2 increases glycolytic activity by

stimulating pyrophosphate: fructose 6-phosphate 1-phosphotransferase (PPr PFK) and

50

51

inhibiting cytosolic fructose 1,6-bisphosphatase (Fru 1,6-P2ase) (Van Schaftingen,

1987; Stitt, 1990). PPr PFK has been identified in the radicle, scutellum and aleurone

but not in the starchy endosperm of dormant and nondormant A vena sativa (Corbineau

et al., 1989). In rice seedlings, the activity of PPr PFK increases when oxygen is

limiting (Mertens et al., 1990), with a consequent increase in its glycolytic importance

(Mertens, 1991). Such conditions may prevail in the dense tissues of a hydrated seed.

So, an increase in the [Fru 2,6-PJ may act as a marker for increased glycolytic

activity in the germination phase.

In nondormant Avena sativa, Fru 2,6-P2 levels increased in the germination

phase but were not correlated with radicle emergence (Van Larondelle et al., 1987).

Fru 2,6-P2 levels did increase with radicle emergence in nondormant Phaseolus

vulgaris (Botha and Small, 1987). In both dormant spores of Phycomyces

blakesleeanus and Avena sativa seeds, dormancy-breaking chemical treatments induced

transient increases in [Fru 2,6-P2] (Van Laere et al., 1983; Van Larondelle et al.,

1987) prior to visible germination. Slicing of dormant tubers and roots led to

increased levels of glycolytic intermediates indicating the restoration of metabolic

activity. This also led to an increase in [Fru 2,6-PJ (Van Schaftingen and Hers, 1983;

Kowalczyk, 1989). Hence, Fru 2,6-P2 has value as an indicator of increased

glycolysis.

Embryo [Fru 2,6-PJ was investigated in red rice to see if Fru 2,6-P2 could be

used as a marker for the onset of the germination phase, which is taken to indicate the

transition from the dormant to nondormant state. I report the effect of dormancy-

52

breaking chemical treatments on embryo [Fru 2,6-PJ. Evidence is presented that (A)

dormancy-breaking chemicals can induce an increase in embryo [Fru 2,6-PJ after only

2 h contact, and (B) embryo [Fru 2,6-P2] is primarily related to the subsequent

germination rate rather than dormancy-breaking.

Materials and Methods

Mature, dormant red rice (Oryza sativa) seeds (straw-hulled, awnless) were

obtained from the South Farm, Rice Research Station, Crowley, LA in 1990. Seeds

were harvested by hand shattering of individual plants. Moisture content at harvest

was 24.2%. After drying for four days in open trays at 22°C, the moisture content

was 13.8%. Seeds were stored in Mason jars at -15°C. Nondormant red rice was

obtained by afterripening dormant seeds at 22°C for 76 days after which they were

stored at -15°C. Freshly prepared solutions of dormancy-breaking chemicals were

used for each experiment. Experiments were repeated at least three times. All data are

presented as the mean + the standard error of the mean.

Seed incubation for determination of [Fru 2,6-PJ

One hundred manually dehulled seeds were sown on 9 cm Petri plates

containing tliree sheets of germination paper, 10 mL distilled H20 and covered with

a double layer of tissue paper (Kimwipe) and incubated at 30°C in darkness.

Nondormant and dormant seeds were sampled over 24 h. Germination was also

53

recorded. In the case of dormant seeds, escapes were replaced with similarly hydrated

seeds.

For studies with dormancy-breaking chemicals, manually dehulled seeds (110

seeds used) were incubated as above for 24 h. Germinated seeds were replaced with

similarly treated dormant seeds prior to dormancy-breaking treatments. In the controls

germinated seeds were replaced immediately prior to sampling. Seeds were transferred

to 250 mL wire clasp storage jars (Heritage Industries, Millville, NJ) containing two

sheets of Whatman N°1 filter paper and 10 mL of test solution. Seeds were covered

with tissue paper, and jars were sealed. Dormancy-breaking compounds were applied

as a 24 h/30°C pulse in 25 mM citrate/phosphate buffer at concentrations that broke

dormancy in 90% of the population. Sodium nitrite (4 mM) was applied at pH 3.0 (pK

3.3). Propionic acid (22 mM) was applied at its pK of 4.9. Methyl propionate (32

mM), n-propanol (75 mM), and propionaldehyde (40 mM) were applied at pH 7.0.

At the end of the chemical pulse, seeds were transferred to Petri plates containing H20

(as described above) for 24 h/30°C. Samples were taken at intervals throughout the

chemical pulse and post-pulse phases.

Fru 2,6-P2 extraction

The following extraction and anion exchange steps are after Van Schaftingen

(1984). For 'each determination, 100 embryos were excised and immediately frozen

in liquid nitrogen. Embryos were then ground to a powder in liquid nitrogen. This

powder was homogenized in a cold tissue homogenizer with 1 mL of ice cold, 200

54

mM NaOH. After decanting the homogenate to a cold centrifuge tube, the remaining

material was recovered by rinsing the homogenizer with 1 mL 200 mM NaOH. The

homogenate was then centrifuged at 20,0Q0g for 5 minutes at 1°C. The supernatant

was heated at 80°C for 5 minutes, then cooled on ice. The same procedure was used

for the first forty endosperms, of which the weight was also recorded.

The pericarp of red rice contains anthocyanins (Takahashi et al. , 1989). These

were removed from the extracts by passing each sample through a Pasteur pipette

loaded with Dowex 1x8 (4 x 40 mm, Cl' form, 200 mesh) at 5°C. The loaded column

was washed with 1 mL distilled H20 and 3 mL 0.15 M NaCl. Fru 2,6-P2 was eluted

in 3 mL 0.3 M NaCl. Samples were frozen at -20°C until assayed. Columns were

prepared in advance by washing with 4 mL 1 M NaCl and 10 mL H20 , then stored

hydrated at 0 to 5°C. In each experiment one time point was replicated to test the

recovery of Fru 2,6-P2. An amount double that expected in the tissue was added to

the sample after the first aliquot of NaOH. A recovery control (no tissue included)

was also prepared by the same extraction protocol to confirm the [Fru 2,6-PJ used

in the recovery sample.

Fru 2,6-P2 Assay

Fru 2,6-P2 was assayed after Van Schaftingen (1984). Lyophilized enzymes

were used; aldolase type X (EC 4.1.2.13, rabbit muscle, Sigma), glycerophosphate

dehydrogenase (EC 1.1.1.8, GDH, rabbit muscle, US Biochemical Corp),

pyrophosphate-dependent phosphofructokinase (EC 2.7.1.90, PPi-PFK, potato tuber,

55

Sigma), triose phosphate isomerase type X (EC 5.3.1.1, TPI, rabbit muscle, Sigma).

GDH, PPi-PFK, and TPI were stored in 20 mM TRIS/HC1, pH 8.2, 20 mM KC1, 2

mM dithiothreitol and 60% (w/w) glycerol at -20°C.

Homogenate samples (1 mL) were neutralized with 0.1 mL 1 M HEPES, 5.5%

(w/v) soluble PVP-10 (Polyvinylpyrrolidone), pH 7.8. To remove the effect of

variability in sample composition an internal calibration was performed for each assay;

a calibration solution was prepared by acidifying 0.5 mL of neutralized sample with

0.2 mL 0.4 M HC1 for 20 minutes at room temperature to destroy endogenous Fru

2,6-P2, then neutralizing with 0.2 mL 0.4 M NaOH. This gives a sample:HClrNaOH

ratio of 5:2:2 which is used in all sample assays.

To each of four quartz cuvettes was added: calibration solution, 0.5 mL

reaction buffer (100 mM TRIS/acetate pH 8.0, 4 mM magnesium acetate, 2 mM F6P,

0.3 mM NADH; F6P was first acid treated to destroy contaminating Fru 2,6-P2) and

0.1 mL enzyme solution (25 mM TRIS/acetate pH 8.2, 150 UL'1 PPi-PFK, 20 KUL1

aldolase, 100 KUL'1 TPI, 20 KUL'1 GDH, and 0.2% BSA). Fru 2,6-P2 was added at

0, 0.2, 0.5 and 1.0 pmoles. The volume in each cuvette was made up to 0.95 mL

with distilled H20 . In a fifth cuvette equivalent volumes of 0.4 M HCL, 0.4 M NaOH

(followed by the reaction buffer) and sample were substituted for the calibration

solution. 1'he reaction was started by the addition of 0.05 mL 10 mM pyrophosphate.

The assay-w&s monitored at 339 nm for 10 min/25°C using a temperature controlled

cuvette block in a Gilford Response II spectrophotometer. The Fru 2,6-P2 content of

the sample was determined by linear regression analysis of the internal calibration.

56

Coordinates (0,0) were used to represent the blank. Only data from assays where the

correlation coefficient was > 0.995 were used.

Respirometry

Three replicates of 20 manually dehulled seeds were incubated in 15 mL

respirometer flasks with two sheets of Whatman N°1 filter paper and 2 mL distilled

H20 . The center well was filled with 0.2 mL 20% KOH. A film of lanolin was placed

around the top of the well to stop the KOH from creeping up the glass wall. A fluted

filter paper wick was placed in the well to increase the surface area for absorption of

C 02. An internal control lacking seeds was included with each experiment to indicate

atmospheric pressure fluctuations. The water bath was maintained at 30°C and

monitored periodically as changes in temperature effect the manometer readings.

Respiration was recorded over one hour intervals, every 2 h for 24 h. At the end of

the 24 h period, seeds were transferred to 50 mL Erlenmeyer flasks for the standard

germination test. For dry seeds, respiration was measured over 24 h and the rate

expressed on an hourly basis. Respiration was recorded as /xL 0 2 uptake h'1 20 seeds'1.

Seed incubation for germination tests

Germination tests used five replicates of 20 maually dehulled seeds/treatment

in 50 mL Erlenmeyer flasks with two sheets of Whatman N°1 filter paper and 2 mL

of test solution. Flasks were sealed with rubber septum caps. Nondormant seeds were

incubated for 7 d/30°C. For studies using dormancy-breaking chemicals, seeds were

57

incubated in Erlenmeyer flasks as for [Fru 2,6-PJ determinations and transferred to

distilled H20 for 7 d/30°C, when germination was determined. Germination was

measured as rupture of the pericarp and aleurone over the embryo (criterion used in

time course experiments) or radicle protrusion. Germination >10% was taken as an

indicator of the commencement of germination within the population. Germination and

viability tests were performed with each experiment. Germination data are presented

in the appropriate figures. Seed viability was > 99%, as determined by excising

embryos from ungerminated seeds and incubating on distilled H20 at 30 °C for 7 d.

Results

Embryo [Fru 2,6-P2] during hydration

In dry seeds, embryo [Fru 2,6-P2] was low in both the embryo (Fig. 4. IB) and

endosperm (Appendix E; Table E.3 & E.4). In embryos of both dormant and

nondormant dehulled seeds, the [Fru 2,6-P2] increased during imbibition. In

nondormant embryos, this increase reached a plateau where it remained from 4 to 12

h, when the first visible signs of germination were seen (Fig. 4.1). Between 10 and

12 h desiccation tolerance declined from 94 ± 5% to 11 ± 5 % (Appendix A; Table

A .l). Germination reached 83% by 16 h (Fig. 4.1C). Over this period (12 to 16 <h)

the [Fru 2,6-PJ doubled (Fig. 4. IB). Rapid increases in radicle emergence and

embryo [Fru 2,6-PJ were coincident (18 to 20 h). In dormant seeds, embryo [Fru

2,6-PJ decreased following the plateau associated with imbibition.

58

In endosperm tissue, the [Fru 2,6-P2] increased in both dormant and

nondormant seeds during imbibition, then decreased only to rise at the onset of

germination (Appendix E; Table E.4). These changes may reflect contamination with

embryo tissue. When endosperm was extracted in the absence of adhering embryo

tissue (scutellum), Fru 2,6-P2 was detected at approximately 0.25 ± 0.03 pmoles

endosperm1 over 14 h of hydration.

Seed respiration during hydration

A difference in respiration was seen between dry dormant and nondormant

seeds. During the hydration of dormant and nondormant seeds, respiration rate

increased during the imbibition phase. Following imbibition, the respiration rate of

nondormant seeds remained on a plateau in the germination phase from 8 to 14 h.

From 14 h onwards the respiration rate increased at the point when radicle emergence

commenced. In dormant seeds, respiration rate remained stable (Fig. 4.1). Seed

respiration rate and embryo [Fru 2,6-PJ were highly correlated in dormant and

nondormant seeds during the first 24 h of hydration (Fig. 4.2).

Embryo [Fru 2,6-PJ during and post-chemical contact.

No difference was found between embryo [Fru 2,6-PJ in the controls

conducted at pH 3.0, 4.9, and 7.0, both during and post-chemical contact (pH 4.9

&7.0 only). Hence, these data were combined. Embryo [Fru 2,6-PJ did not change

in the controls over a 48 h period (Fig. 4.3B).

59

-a<uoco

CMO

0- 0- 0 ' °

® - ® - ® - ® - ® - ® - ® - ®

O— O N o n d o r m a n t • — • D o r m a n t

® -® -®

A Pericarp □ Radicle O Shoot

8 12 16 20 24

Hours

Figure 4.1. A) Respiration of dehulled dormant and nondormant seeds over 24 h/30°C as 0 2 uptake /xL1 h'1 seed'1 B) Fru 2,6-P2 content of embryos in dehulled dormant and nondormant seeds during hydration at 30°C. Data are corrected for full recovery based on within-treatment recoveries. Recoveries: dormant, 77%; nondormant, 86%. C) Utilization of pericarp splitting, radicle and shoot emergence as the criteria for % visible germination of nondormant seeds in Fig. IB.

60

Increases in embryo [Fru 2,6-PJ during chemical contact differed depending

on the dormancy-breaking chemicals employed (Fig. 4.3B). In the case of nitrite and

propionaldehyde, embryo [Fru 2,6-PJ, (a) increased after only 2 h of contact, and (b)

increased to plateau levels similar to that of nondormant embryos (Figs. 4. IB &

4.3B). Embryo [Fru 2,6-PJ increased above the plateau level when visible

germination commenced, which was during chemical contact for both nitrite (16 h)

and propionaldehyde (24 h). n-propanol treatment also produced a distinct although

lower embryo [Fru 2,6-PJ plateau level. Propionate and methyl propionate did not

increase embryo [Fru 2,6-PJ above that of the control during contact.

Embryo [Fru 2,6-PJ increased post-chemical contact, prior to the

commencement of germination (Fig. 4.3B & C). When dry dormant seeds were

hydrated on 75 mM n-propanol (Appendix E; Table E.8), embryo [Fru 2,6-PJ

followed the same pattern as when hydrated on H20 .

The mean plateau embryo [Fru 2,6-PJ during chemical contact for all

treatments was significantly correlated with the elapsed time to 30% germination (Fig.

4.4). This relationship also held true for 40% germination (r = -0.9977; P < 0.001).

Following dormancy-breaking, germination percentages were significantly correlated

with embryo [Fru 2,6-PJ in chemically treated seeds (Fig. 4.5).

61

1 1 . 4 -1o ■ ■

1 . 2 -_QECD 1 . 0 -CM

CL1 0 . 8 -

CD ■■CM 0.6 --D , .v _

Li_ 0.4 —COCDO 0 . 2 -E -i-CL 0 .0 - -

y = 0 .1 8(x) - 0 .022; r = 0 .92; P < 0.001 j

O N o n d o r m a n t j iQ i

? b 1

D o r m a n t

- 2 .0 - 1 .0 0.0 1.0 2.0 3.0 4.0 5.0 6.0 7.0

f i L C>2 h — s e e d - ^

Figure 4.2. Correlation between embryo [Fru 2,6-PJ and seed respiration. Data from Fig. 4.1.

62

Figure 4.3. Dormant seeds hydrated for 24 h/30°C, then exposed to each dormancy- breaking chemical. A) Dormancy-breaking after increasing periods of contact. Buffer controls represent 0 hours. B) Fru 2,6-P2 content of embryos in dehulled dormant seeds during and post-chemical contact. Data are corrected for full recovery, based on within-treatment recoveries. Recoveries: control, 76%; methyl propionate, 91%; nitrite, 97%; n-propanol, 97%; propionaldehyde, 90%; propionic acid, 96%. C) Germination (pericarp splitting) in real time of seeds in Fig. 4.2B.

63

P u l s e P o s t —p u l s eooOfO\x>h-

l_V

c_ooc

Q>O

I0JDEa>CN

CL1

CD<NI

Li_mooEa .

Q>E

oa>

co'- 4->Oc

oo

• Control

A Methyl propionate

V Nitrite

□ n—Propanol

O Propionaldehyde

0 Propionic acid

8 0 - - Y

.2

100 - -

0 4 8 12 16 2 0 2 4 2 8 3 2 3 6 4 0 4 4 48

Hours

64

13OCD

_oQ_C0

T_ Q

ECD

CNQ.1

CD

CN

D LL.C/3CD

C L

0.6-

0.5-

0.4-

0.3-

0.2- AV□

0.1 -O

0 0- ----- i

t y = -0.01 4(x) + 0.752 v r = -0 .984; P < 0.001

H h0 4 8 1 2 1 6 20 24 28 32 36 40 44 48

Hours to 30% germinat ionFigure 4.4. Correlation between mean plateau [Fru 2,6-PJ and the time to 30% germination. Data are from Fig. 4.3B and 4.3C.

Ger

min

atio

n

65

100.0

80.0

60.0

40.0

^ 20.0

0.00.0 0.2 0.4 0.6 0.8 1.0 1.2 1.4

-i

p m o le s Fru 2 , 6 —Po e m b r y o

y = 8 6 . 1 ( x ) - 2 1 . 5 ; r = 0 . 9 0 5 ; P < 0 . 0 0 1

A Methyl propionate v Nitri te □ n —Propanol O Propionaldehyde

HO Propionic acid

O —«

H 1 H H 1 H

Figure 4.5. Correlation between % germination and [Fru 2,6-PJ. Data are from Fig. 4.3B and 4.3C.

66

Discussion

In this study, the levels of embryo Fru 2,6-P2 during dormancy-breaking

treatments were determined and evaluated as a potential marker for delineating the

transition from the dormant to the nondormant state. Fru 2,6-P2 determinations were

highly reproducible with consistent recoveries in this high phenolic-containing tissue.

Fru 2,6-P2 levels in red rice were consistent with previous studies of Oryza species

(Mertens et al., 1990).

Dormancy-breaking chemicals and embryo [Fru 2,6-P2]

Although the chemicals employed produce a saturating dormancy-breaking

response, their effect on embryo [Fru 2,6-P2] varied with regard to speed and

magnitude of response. Dormant embryo [Fru 2,6-PJ dramatically increased in the

presence of nitrite and propionaldehyde after only two hours contact. Only nitrite

broke dormancy after 2 h contact. The propionaldehyde data demonstrate that a

dormancy-breaking chemical can rapidly induce a physiological response without

breaking dormancy. Furthermore, methyl propionate and propionate readily break

dormancy, yet embryo [Fru 2,6-PJ remained at control levels during chemical

contact. Hence, embryo [Fru 2,6-PJ is not directly related to the onset of dormancy-

breaking. This suggests that the chemicals employed are acting on two levels (a)

dormancy-breaking, and (b) direct or indirect control of PFK2 and FBPase-2 activity.

In the case of dormancy-breaking, these chemicals may act via embryo acidification

(Chapter 3).

67

Contact with nitrite, propionaldehyde, and //-propanol resulted in an increase

in embryo Fru 2,6-P2 to distinct plateau levels. The higher the mean plateau embryo

[Fru 2,6-PJ, the faster the subsequent rate of germination. This suggests that the

higher the mean plateau embryo [Fru 2,6-PJ, the faster the embryo can traverse the

germination phase.

Fru 2,6-P2 stimulates glycolysis (Van Schaftingen, 1987; Stitt, 1990). Thus,

high plateau levels of embryo Fru 2,6-P2 may reflect increased glycolytic activity. The

different plateau levels seen may result from differing effects of the chemicals on

glycolysis, despite their uniform effect on dormancy-breaking.

Chemicals capable of breaking dormancy (Table 1.1) affect glycolysis and [Fru

2,6-PJ. Concentrations of free acid in the /xM range stimulated glycolysis (benzoic

acid) in yeast (Warth, 1991) and increased the [Fru 2,6-PJ in Neurospora (acetic and

salicylic) (Dumbrava and Pall, 1987). Concentrations of free acid in the mM range

inhibited glycolysis (acetic and benzoic acids) (Pampulha and Loureiro-Dias, 1990;

Warth, 1991), decreased the [Fru 2,6-PJ, inhibited PFK2, and stimulated FBPase-2

activity in yeast (benzoic and salicylic acids) (Francois et al., 1986; 1988a). Various

nitrogen sources, including the uncouplers azide and dinitrophenol, stimulated

glycolysis with a correlative increase in [Fru 2,6-PJ (Dumbrava and Pall, 1987;

Francois et al.,. 1988b). Nitrite inhibits oxygen uptake and oxidative phosphorylation

in bacteria (Rowe et al., 1979) -conditions that would lead to increased glycolysis and

[Fru 2,6-PJ (Mertens, 1991). Acetaldehyde, acetate and ethanol stimulated respiration

in potato tubers (Rychter et al., 1979). Conversely, in gluconeogenic tissues,

68

acetaldehyde, ethanol and organic weak acids, inhibited glycolysis and decreased the

[Fru 2,6-PJ in fed hepatocytes (Van Schaftingen et al., 1984; Hue et al., 1988) and

ripening tomatoes (acetaldehyde) (Halinska and Frenkel, 1991).

In other model systems, dormancy-breaking chemical treatments resulted in

transient increases in [Fru 2,6-PJ in both dormant spores of Phycornyces

blakesleeanus and Avena sativa seeds (Van Laere et a l , 1983; Van Larondelle et al.,

1987). However, no transient increase was seen in red rice during chemical contact.

In Avena sativa [Fru 2,6-P2] did not increase with germination but did in red rice.

Differences between red rice and Avena sativa may be due to the use of dehulled,

hydrated red rice vs. intact, dry caryopses of Avena sativa. During the dormancy-

breaking chemical treatment, oxygen uptake by the glumes (Lenoir et al., 1986) may

induce anaerobic conditions in the Avena embryo leading to increased [Fru 2,6-PJ

(Mertens et al., 1990). The glumes also delay the time to visible germination.

Contrary to the effect of ethanol on Avena seeds, hydration of dry, dehulled, dormant

red rice in the presence of n-propanol had no effect on embryo [Fru 2,6-PJ above that

of the control. The [Fru 2,6-PJ found in Avena sativa was an order of magnitude

greater than in red rice and may reflect differing sensitivities of PPi-PFK to Fru 2,6-

P2, with the red rice enzyme being more sensitive. This would be consistent with

comparisons between potato and Jerusalem artichoke tubers. In potatoes, PPi-PFK is

highly sensitive to the low [Fru 2,6-PJ present, the reverse being true in artichoke

tubers (Van Schaftingen and Hers, 1983).

69

Embryo [Fru 2,6-PJ and respiration during germination in real time

The time course of embryo [Fru 2,6-P2] and respiration in nondormant seeds

followed a triphasic response as shown for embryo moisture content and embryo pH

(Chapter 3). The main component of the increase in embryo [Fru 2,6-PJ is a result

of growth subsequent to pericarp splitting. Loss of desiccation tolerance and the onset

of germination occurred at 12 h, immediately prior to the increases in embryo [Fru

2,6-P2] and respiration. Therefore, careful consideration of the visual cue used to

mark the onset of germination (post-germinative growth) is important. The use of

pericarp splitting identified germination four hours prior to radicle emergence (the

traditional germination marker).

In dormant red rice seeds, embryo [Fru 2,6-PJ increased during imbibition,

confirming a previous report (Van Larondelle et al., 1987). This indicated that PFK2

is initially active in imbibing, dormant seeds. Embryo [Fru 2,6-PJ then decreased

following imbibition possibly due to an increase in the PFK2 inhibitors, glycerate 3-

phosphate and phosphoenolpyruvate (Van Larondelle et al., 1987). Respiration

remained constant following imbibition.

After dormancy was broken, percent visible germination in real time and

embryo [Fru 2,6-PJ were significantly correlated, as was visible germination and

embryo pH (Chapter 3). Such correlations indicate that biochemical changes during

visible germination are independent of the chemicals employed to break dormancy.

Similar positive correlations were seen during dormancy-breaking in Phycomyces

blakesleeanus spores for germination capacity vs. both maximal [Fru 2,6-PJ and

70

[glucose 6-phosphate] (Van Laere et al., 1983). The implication is that [Fru 2,6-P2]

vs. physiological response correlations reflect glycolytic activity. This is, to some

extent, borne out by the correlation between embryo [Fru 2,6-PJ and respiration in

hydrating dormant and nondormant red rice seeds. Attempts to measure respiration

during dormancy-breaking chemical treatments were unsuccessful due to absorption

of chemicals by the KOH trap.

Control of Fru 2,6-P2 synthesis

The mechanism by which dormancy-breaking chemicals effect the activity of

PFK2 and FBPase-2 is unclear. In both potato tubers and dormant Avena sativa seeds,

ethanol treatments decreased the levels of 3-phosphoglycerate and

phosphoenolpyruvate; both metabolites inhibit PFK2 and stimulate FBPase-2 (Rychter

et al., 1979; Van Larondelle et al., 1987). This indicates that a potential block on

glycolysis lies between phosphoenolpyruvate and pyruvate. Activation of these

enzymes may be via phosphorylation state, enzyme binding (Moorhead and Plaxton,

1988; Storey, 1990) or sulfhydryl status. Reduction of enzyme sulfhydryls is required

for activity in PFK2 (El-Magrabi et al., 1990; Espinet et al., 1990; Van Larondelle

et al., 1986). In PPi-PFK, the sulfhydryl groups required for Fru-2,6-P2 activation are

protected by phosphate metabolites. Reduced thioredoxin h will restore PPi-PFK

activity (Kiss et al., 1991). Thioredoxin activity increased during the first 24 h of

wheat germination (Kobrehel et al., 1992). It is unclear when this activity increased

71

since germination was 70% at the first time point (24 h) (Buchanan, personal

communication).

In summary, it was found that (a) seed respiration increased during hydration;

and in dormant seeds respiration remained constant following imbibition; (b) embryo

[Fru 2,6-PJ increased during hydration, but in dormant seeds this was transient; (c)

embryo [Fru 2,6-PJ reached a plateau during the germination phase; (d) embryo [Fru

2,6-P2] increased following germination; (e) embryo [Fru 2,6-PJ and respiration

during hydration were highly correlated. When dormant seeds are subjected to

dormancy-breaking chemical treatments, (f) dormancy-breaking chemicals could be

rapidly perceived by dormant embryos; (g) a dormancy-breaking chemical

(propionaldehyde) can induce a physiological response without breaking dormancy;

(h) embryo Fru 2,6-P2 levels were not related to dormancy-breaking; and (i) the

magnitude of the embryo [Fru 2,6-PJ plateau was highly correlated with the speed of

subsequent germination. The data suggest glycolysis may be activated before

dormancy is broken.

C H APTER 5

A 13C NMR STUDY OF THE METABOLIC FATE OF

DORMANCY-BREAKING CHEMICALS IN DORMANT RED

RICE EMBRYOS

Introduction

In a previous study, embryo acidification during dormancy-breaking chemical

treatments was proposed as a marker for the termination of developmental arrest. It

was also, suggested that embryo acidification resulted from metabolism of acid

analogues to the parent acid (Chapter 3). The metabolic fate of dormancy-breaking

chemicals is a neglected area with few available studies (Bradbeer and Colman, 1967;

Eilers and Sussman, 1970; Rast and Stauble, 1970).

I4C-labelled acetate was metabolized in excised tissues from stratifying dormant

hazel nuts (Bradbeer and Colman, 1967). This may reflect increased metabolic activity

as a result of the dormancy-breaking effects of stratification, but wounding will also

contribute to increased metabolism (Van Schaftingen and Hers, 1983; Moorhead and

Plaxton, 1988). When dormancy was broken by furfural (Neurospora ascospores) or

isovaleric acid (Agaricus bisporus (Lange) Sing, spores), these chemicals were

metabolized prior to visible germination (Eilers and Sussman, 1970; Rast and Stauble,

1970).

72

73

While these metabolic events were considered to be important in the dormancy-

breaking process (Rast and Stauble, 1970), they were dismissed in Neurospora (Eilers

and Sussman, 1970) even though the furfural metabolite furfuryl alcohol also broke

dormancy (Sussman and Halvorson, 1966). None of these studies attempted to relate

metabolism of dormancy-breaking chemicals to the dormant/nondormant transition.

Propionate and its analogues have previously been shown to alter both embryo

pH and fructose 2,6-bisphosphate (Chapters 3 & 4). It was considered important to

ascertain whether these changes are related to the metabolism of these dormancy-

breaking chemicals during contact. To this end the metabolism of propionate-2-13C and

n-propanol-l-13C was investigated during dormancy-breaking chemical treatments.

Both were found to be metabolized prior to dormancy-breaking and visible

germination.

Materials and Methods

Mature, dormant red rice (Oryza sativa) seeds (straw-hulled, awnless) were

obtained from the South Farm, Rice Research Station, Crowley, LA in 1990. Seeds

were harvested by hand-shattering of individual plants. Moisture content at harvest

was 24.2%. After drying for four days in open trays at 22°C, the moisture content

was 13.8%. Seeds were stored in Mason jars at -15°C. Propionate-2-13C and n-

propanol-l-13C (both 99% isotopic enrichment) were obtained from Isotec

(Miamisburg, OH) and 3-hydroxypropionate from Chemical Service Inc. (West

74

Chester, PA). Nystatin, rifampicin and deuterium oxide were from Sigma (St. Louis,

MO). All other chemicals were of reagent grade. Freshly prepared solutions of

dormancy-breaking chemicals were used for each experiment. Experiments were

repeated three times. All data are presented as the mean ± the standard error of the

mean.

Seed incubation for 13C labelling studies

Prior to exposure to 13C labelled dormancy-breaking chemicals, manually

dehulled seeds (110 seeds used) were incubated on 9 cm Petri plates containing three

sheets of germination paper, 10 mL distilled H20 and covered with a double layer of

tissue paper (Kimwipe) and incubated for 24 h/30°C in darkness. Seeds were then

surface sterilized based on the recommendations of Abdul-Baki (1974), and Abdul-

Baki and Moore (1979). Seeds were placed in 25 mL 5% sodium hypochlorite solution

(available chlorine 5%) at pH 7.0 for 15 min (the hypochlorite solution was adjusted

to pH 7.0 using 1 M citrate/phosphate). Seeds were then transferred to 25 mL 0.1 N

HC1 for 10 min then rinsed 8 times with 25 mL aliquots of sterile HzO. Germinated

seeds were then removed to leave 100 dehulled dormant seeds. Seeds were transferred

to 250 mL wire clasp storage jars (Heritage Industries, Millville, NJ) containing two

sheets of Whatman N°1 filter paper and 10 mL of test solution. Seeds were covered

with tissue paper, and jars were sealed. Dormancy-breaking compounds were applied

as a 24 h/30°C pulse at concentrations previously shown to break dormancy in 90%

of the population (Chapter 4). Propionate-2-13C (22 mM) was buffered at its pK of 4.9

75

with 25 mM citrate. n-Propanol-l-13C (75 mM) was buffered at pH 7.0 with 25 mM

citrate/phosphate. Each dormancy-breaking chemical was applied in presence of the

antibiotics, nystatin (60 fig mL'1) and rifampicin (25 fig mL"1) in 10 mM

dimethylsulfoxide (DMSO). DMSO was required to dissolve the rifampicin. At the

end of the chemical pulse (propionate treatment only), seeds were transferred to Petri

plates containing distilled H20 (as described above) for 12 h/30°C. Samples were

taken at intervals throughout the chemical pulse and post-pulse phases. Germination

was recorded for each sample. Germination was measured as rupture of the pericarp

and aleurone over the embryo (criterion used in time course experiments) or radicle

protrusion. Germination data are presented in the appropriate figures.

Labelled metabolite extraction

For each determination, 100 embryos were excised and immediately frozen in

liquid nitrogen. Embryos were then ground to a powder in liquid nitrogen. This

powder was homogenized in a cold tissue homogenizer with 0.75 mL of ice cold, 2

M perchloric acid and 0.25 mL deuterium oxide. The homogenate was decanted to an

Eppendorf tube and centrifuged for 2 min at 20,000g/4°C. The supernatant was

removed, frozen in liquid N2, and 120 fiL of 8 N KOH (-20°C) was added. Freezing

prevented the decomposition or volatilization of metabolites during the highly

exothermic ''chlorate precipitation. The KC104 precipitate was removed by

centrifugation. The supernatant was removed, and 0.2 mL 2M phosphate at pH 6.8

was added as buffer. The volume was then made up to 1 mL with distilled H20 .

76

Samples were stored at -20°C. The same procedure was used for the first forty

endosperms; care was taken to ensure no embryo tissue was attached.

13C NMR of extracts

Spectra were recorded on a Bruker AM 400 spectrometer operating at 100.62

MHz with a 5 mm broad band probe and 2H lock signal. Spectra were acquired using

a 106° pulse angle of 10 jxsec, an acquisition time of 0.655 s, and recycle delay of

2.0 s. High power broadband proton decoupling was used during acquisition. Unless

otherwise stated each spectrum consists of 1024 scans obtained with 32K data points.

The resulting free induction decays were Fourier transformed after the application of

exponential multiplication with line broadening of 2 Hz. The signal of the methylene

carbon of propionate-2-13C [31.7 5(ppm)] was used as an internal standard. Signal

identification was obtained by comparison with 13C NMR spectra of standards and

addition of different standards to PC A extracts. Methylene groups were identified

using proton coupled spectra to determine carbon-proton couplings ( / C - h ) - In proton

coupled spectra, signal:noise ratio was improved using nuclear Overhauser

enhancement achieved by proton decoupling during the recycle delay (gated

decoupling) (Fig. 5.1) (Breitmaier and Voelter, 1987). Signal intensities in the

propionate and n-zi-propanol labelling studies are expressed relative to a 3-

hydroxypropionate-2-13C signal after 24 h contact with propionate-2-13C. The signal

was set at 100 arbitrary intensity units. This enabled direct comparisons to be made

between the two studies.

77

A) Broad band decouplingON

H (decoupler)o f F

PW AQ

C RD

B) Gated decouplingON

(decoupler)OFF

PW AQ

13C RD

Figure 5.1. Pulse sequences used for acquisition of 13C NMR spectra. (A) Broad band proton decoupling, and (B) gated proton decoupling. Abbreviations: AQ, acquisition; PW, pulse width; RD, recycle delay.

Results

Metabolism of propionate-2-13C by dormant red rice

Metabolism of propionate-2-l3C in the incubating solution was not detected

after 24 h/30°C contact with dormant red rice seeds. This indicated that propionate-2-

13C metabolism occurred within the seeds. On transfer of hydrated, dehulled, dormant

seeds to propionate-2-13C, metabolism to 3-hydroxypropionate-2-13C was detected in

the embryo after only 2 h (Fig. 5.2 & 5.3B). Penetration of embryo tissue by

propionate was linear. Its conversion to the hydroxy acid proceeded at the same rate

as uptake from 0 to 8 h (Fig. 5.3B). The propionate signal reaches a plateau after 8

h before declining when seeds are transferred to H2Q at 24 h. The intensity of the 3-

hydroxypropionate signal continued to increase up to 24 h. Total intensity of the

propionate plus the 3-hydroxypropionate signals was linear over the first 24 h and then

declined (Fig. 5.4). Low level metabolism of propionate-2-!3C to the hydroxy acid was

detected in the endosperm during chemical contact (Appendix F.3). The onset of

dormancy-breaking occurs between 8 to 12 h in the presence of unlabelled propionate;

at this point signal intensities had diverged (Fig. 5.3 A & B). Visible germination

increased following transfer to H20 . The 3-hydroxypropionate signal shows a rapid

decline at 36 h as radicle (4%) and shoot (4%) emergence commence.

In the post-pulse phase a signal identified as that of the C-2 and C-4 carbons

of citrate appeared at 46 ppm. An unidentified signal also appeared at 32 ppm.

Throughout these experiments, weak, unidentified signals that possibly correspond to

the CHOH groups of carbohydrates appeared in the region 50 to 80 ppm.

79

b

m/ti &f*ts3 6 h

3 2 h

2 8 h

*WvTivw i \iKw*Vr**

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-I—I—I—I—I—I—I—I—I—I—I—I—I—I—I—I—I—I I I I I I- 1 I 1 1 1 r4 5 4 0 3 5 3 0 2 5

PPM

Figure 5.2. 13C NMR spectra of perchloric acid extracts. Time course of propionate-2-13C metabolism in red rice embryo during (0 to 24 h) and post-chemical contact (28 to 36 h). Chemical shift assignments, (ppm); (a) C-2,4 of citrate-2,4-13C, 46.6; (b) C- 2 of 3-hydroxypropionate-2-13C, 41; (c) unidentified, 32.1; (d) C-2 of propionate-2- 13C, 31.7,- ;

P u l s e80

P o s t —pu l se

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® 3 —Hydroxypropionate—2 — 13O P r o p io n a t e —2 — C

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A 3 2 ppm

-CL'O -

'0 „

'O -- A - -Ar

8 1 2 1 6 2 0 2 4 2 8 3 2 3 6

Hours

Figure 5.3. (A) Visible germination percentages after 7 d/30°C following increasing periods of contact with propionate and visible germination percentages in real time. (B) Time course of signal intensities (arbitrary units) of 13C-resonances from citrate- 2,4-13C, 3-hydroxypropionate-2-13C, propionate-2-13C, and an unidentified resonance at 32.1 ppm.

81

Pulse Post—pulse

>N H—* 1 2 0 -' ( f )c0-4-» 1 0 0 -c

"5 8 0 -cCD

' ( f )6 0 -

0> • ■

■+->4 0 -

o0

O d2 0 -

8 12 16 20 24 28 32 36

Hour s

Figure 5.4. Total relative intensity of the signals from C2 of both 3- hydroxypropionate-2-,3C and propionate-2-,3C during and post chemical pulse. Data from Fig. 5.3B.

82

V V V W t . 2 4 h

2 0 h

H * H # t f 1 6 h

y ' W w W f l 1 2 h

W V i U 8 h

kW 4 h

2 h

o h

1 8 0PPM

- i — i— i— |— i— i— i r

6 5PPM

Figure 5 .5 .13C NMR spectra of perchloric acid extracts. Time course of zi-propanol- 1-13C metabolism in red rice embryos during chemical contact (0 to 24 h). Chemical shift assignments (ppm): (a) C-l of 3-hydroxypropionate-1-13C, 181.6; (b) C-l of n- propanol-l-I3C, 64.6.

83

E 1 0 0 -

Da>L_L_ 8 0 -cco

O '-P P o D C

6 0 -EL.

fc o 4 0 -

* ♦ '

2 0 -

♦

6 0 — • -— 9 3 —Hydroxypropionate—1 — C

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■+->

‘coca;

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4 0 -

3 0 -

2 0 -

_oCD 1 0 -

80 4 12 1 6 20 24

H o u r s

Figure 5.6. (A) Visible germination percentages after 7 d/30°C following increasing periods of contact with n-propanol and visible germination percentages in real time. (B) Time course of relative signal intensities (arbitrary units) of 13C-resonances from 3-hydroxypropionate-l-,3C and n-propanol-l-13C.

84

Table 5.1. 13C NMR assignments (ppm) of standards and metabolite signals (100.6 MHz in 85% H20/15% D20)

Citrate 3-Hydroxypropionate

C2 & C4 C2

5(ppm) 7 c.„(H z)‘ 5(ppm) J c-h(H z)

standard 4 4 .1 5 2 131.65 4 0 .6 0 6 126 .10

metabolite 46 .731 126 .69 40 .9 8 5 127 .04

m etabolite + standard 46 .655 128 .37 40 .9 6 7 126 .72

Vc.H = coupling constant

Table 5.2. Effect of pH on 3-hydroxypropionate assignments

3-Hydroxypropionate

Cl

S(ppm)

C2

5(ppm)

C3

5(ppm)

pH 3.0 177.537 37.792 58.342

pH 5.3 180.890 40.483 59.701

pH 9.8 181.628 41.068 60.000

/t-propanol-1-13C labelled sample

pH 7.7 181.641

/z-propanol-l-13C labelled sample + standard

pH 5.7 181.452

85

Metabolism of n-propanol-l-13C by dormant red rice

Embryo tissue is rapidly penetrated by n-propanol-l-I3C;

a steady state level is achieved after only 2 h (Fig. 5.5 & 5.6B). A signal at 181 ppm

appears after 4 h and corresponds to 3-hydroxypropionate-l-13C. Metabolism of n-

propanol clearly commences prior to the onset of dormancy-breaking (16 h) (Fig.

5.6A & B). Visible germination is below 10% at the end of the chemical contact

period.

Identification of metabolite signals

Metabolite signals where identified as originating from Cl and C3 of 3-

hydroxypropionate, and C2 and C4 of citrate. Methylene groups were identified by

comparison of signal multiplicities (Table 5.1). The Cl signal of 3-hydroxypropionate

is pH sensitive. This explains the chemical shift seen upon spiking of samples (Table

5.2). Spectra were recorded using a 106° pulse angle rather than a 90° pulse angle

of 8.5 /xs. This reduces signal intensity but does not effect signal assignments or

metabolic trends observed. No signals were identified as originating from the

methylenes of malate, 40.3 ppm; succinate, 35.1 ppm; or N-carbamyl-DL-aspartic

acid, 41.3 ppm. Coenzyme A derivatives were not detected.

86

Discussion

The ability of red rice embryos to metabolize dormancy-breaking chemicals

was observed and compared to the onset of dormancy-breaking and visible

germination within the population. The metabolic activity detected in the endosperm

is considered to reside in the aleurone. Catabolism of these dormancy-breaking

chemicals to a common metabolite occurred prior to the onset of dormancy-breaking.

This indicates that the metabolism of dormancy-breaking chemicals to a weak acid

may indeed be required for dormancy-breaking activity.

In red rice the metabolism of -propanol to propionaldehyde is assumed to be

via alcohol dehydrogenase as in Avena sativa (Corbineau et al., 1991).

Propionaldehyde-1-13C was not detected in red rice. Exogenous propionaldehyde

acidifies embryos rapidly upon contact (Chapter 3); this is taken to indicate rapid

oxidation to propionic acid by cytosolic aldehyde dehydrogenase (Oppenheim and

Castelfranco, 1967). Propionic acid in turn is metabolized to 3-hydroxypropionate, an

intermediate of modified j8-oxidation (Hatch and Stumpf, 1962) located in the

peroxisome (glyoxysome in seeds) (Gerhardt, 1986; Gerbling and Gerhardt, 1989).

Oxidation finally yields C 02 and acetyl CoA which then enters the glyoxylate cycle

as citrate (Stumpf, 1970).

Embryo metabolism of n-propanol and propionate to 3-hydroxypropionate was

evident prior to the contact interval required for the onset of dormancy-breaking. In

87

each case embryo acidification occurred prior to or coincident (^-propanol) with the

onset of dormancy-breaking (Chapter 3). Metabolism of n-propanol to 3-

hydroxypropionate was detected after 4 h contact, at the same time embryo pH

decreased and embryo [Fru 2,6-P2] increased compared to the controls (Chapter 3 &

4). If further metabolism of the hydroxy acid is rate limiting, the resulting slow

accumulation of C 02 (Stumpf, 1970) may explain late embryo acidification coincident

with the onset of dormancy-breaking (Chapter 3). The requirement for metabolism to

a weak acid and the resulting acidification is also suggested by another study;

bamyardgrass dormancy is maintained in wounded seeds by removal of C 02 generated

by the wounding response (Leather et ah, 1992).

3-Hydroxypropionate does not appreciably accumulate in n-propanol-treated

embryos. It does, however, accumulate in a linear fashion during contact with

propionate. The rate of synthesis is linear over 24 h of chemical contact. Further

metabolism appears to be limited until the onset of visible germination. Overall, the

data from red rice suggest the following: (1) oxidation of n-propanol to

propionaldehyde is rate limiting; (2) propionaldehyde is rapidly oxidized to

propionate, as indicated by rapid embryo acidification and the inability to detect

propionaldehyde-1-13C. (3) /3-oxidation rapidly oxidizes propionate to 3-

hydroxypropionate. The following steps are rate-limiting as indicated by the late

appearance of citrate. Similar results were obtained for propionate metabolism in

Phaseolus limensis L. stems (Halamkar et al., 1988). As the extraction conditions

88

lead to the conversion of CoA derivatives to the free acid form, the 3-

hydroxypropionate may be derived in part from the CoA derivative.

Both dormancy-breaking chemicals penetrated the embryo rapidly. Penetration

by n-propanol was saturating after only 2 h. Penetration by propionate appeared to be

saturating by 8 h but the total intensity of the propionate and 3-hydroxypropionate

signals shows that uptake was linear over the chemical contact period.

In summary, it was found that (a) dormant embryos rapidly take up n-propanol

and propionate; (b) w-propanol and propionate are metabolized by dormant red rice

embryos prior to the onset of dormancy-breaking; (c) a dormancy-breaking chemical,

n-propanol, that lacks a dissociable proton is metabolized to an acid form (3-

hydroxypropionate and possibly C 02) leading to embryo acidification.

CHAPTER 6

31P NMR OF DORMANT RED RICE EMBRYOS: THE

OBSERVATION OF INTERNAL 31P USING A SPIN ECHO

SEQUENCE

Introduction

Embryo acidification has been measured in filtered homogenates during

dormancy-breaking chemical treatments and germination (Chapter 3). It would be

preferable to measure embryo pH in vivo in order to avoid the mixing of subcellular

compartments with differing pH values (e.g., cytoplasmic and vacuolar). Intracellular

pH (pH;) of developing seed embryos was measured in vivo using distribution of the

weak acid DMO (5,5-dimethyloxazolidine-2,4-dione) (Barthe et a l., 1986). Weak base

distribution can also be used (Roos and Boron, 1981). Unfortunately, weak acids

(DMO included) and weak bases break dormancy (Cohn et al., 1989; Chapter 1).

Fluorescent indicators (e.g., fluorescein and pyranin) used to determine pH; are weak

acids (Roos and Boron, 1981; Wolfbeis et al., 1983) which themselves perturb pH*.

A non-invasive technique that does not in itself perturb pH; is 31P NMR. First

used to measure the pH; of erythrocytes (Moon and Richards, 1973), this technique

has been widely used on both animal and plant model systems. The chemical shift

[5(ppm)] of the internal inorganic phosphate signal is pH-sensitive. In solution

89

90

inorganic phosphate (Pj) undergoes rapid exchange between HP042 and H2P 04. The

chemical shifts of these two species are separated approximately 2.4 ppm, but due to

rapid exchange only one signal is seen. As their equilibrium changes with pH, so does

the observed chemical shift which becomes more negative (moves upfield) as pH

decreases (Gadian, 1982; Pfeffer and Gerasimowicz, 1989). Measurement of pH; with

31P NMR has been achieved in several developmentally arrested systems (Nuccitelli

et al., 1981; Morrill et al., 1984; Drinkwater and Crowe, 1987; Candelier et al.,

1989; Goudeau et al., 1989; Hervd et al., 1989). This chapter describes the

development of a 31P NMR protocol designed to enable in vivo measurement of pH;

in embryos of red rice.

Materials and Methods

Mature, dormant red rice (Oryza saliva) seeds (straw-hulled, awnless) were

obtained from the South Farm, Rice Research Station, Crowley, LA in 1987. Seeds

were harvested by hand shattering of individual plants. Moisture content at harvest

was 17.8%. After drying for two days in open trays at 22°C, the moisture content

was 12.6%. Seeds were stored in Mason jars at -15°C. Freshly prepared solutions

were used for each experiment. All chemicals were of reagent grade.

91

Seed incubation and embryo preparation

For each 31P NMR run, five replicates of 110 manually dehulled dormant seeds

were incubated on 9 cm Petri plates containing three sheets of germination paper, 10

mL distilled H20 and covered with a double layer of tissue paper (Kimwipe) and

incubated for 24 h/30°C in darkness.

For each determination, visibly germinated seeds were discarded. Embryos

from 500 seeds were excised and placed on moistened filter paper. Loose endosperm

tissue was gently washed away. Embryos were then loaded into a 10 mm NMR tube.

Embryos were perfused with aerated distilled HzO by means of an airlift system

(Kramer and Bailey, 1991) (Fig. 6.1). The liquid height was 12 cm. A polyethylene

tube (cocktail straw) with dimensions of 10 x 0.3 cm was used to provide the lift

column, around which the embryos were packed. Air was introduced via a 5 fiL glass

capillary that terminated 6 cm from the base of the NMR tube [2 cm above the radio

frequency (RF) coils]. Air was introduced to the system at a flow rate of 50 mL min'1.

Flow rate was controlled by a Matheson 602 flowmeter. Air bubbles did not enter the

region of the RF coils. A reference capillary containing 2 M tetraethyl methylene

diphosphonate (TEMDP) (Aldrich) was inserted in the lift column.

To replace the incubation solution, fluid above the embryos was drawn off and

replaced with the next incubation solution, which was then circulated. This procedure

was carried out twice to ensure replacement of the old incubation solution. The tube

was then refilled a third time. Embryos were incubated in 90% H2Q/10% D20 , and

single pulse and spin echo spectra were recorded. The incubating solution was then

O tn

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Figure 6.1. Airlift perfusion system used to acquire 3,P NMR spectra of red embryos.

93

replaced with 20 mM NaNO2/10% DzO at pH 2.2 (pK 3.3). Spin echo spectra were

then recorded after 5, 30, and 60 min of incubation. After 100 min the perfusate was

alkalinized to pH 8.3 with 6 N NaOH. Spin echo spectra were then recorded after 5,

15, and 30 min of incubation. The perfusate was then replaced with 20 mM

phosphate/10% D20 at pH 7.9, and a spin echo spectrum recorded.

Preparation of embryo sap and standard solutions

Dormant manually dehulled seeds were hydrated for 6 h/30°C as described

above. The embryos of 3,500 seeds were excised, and the sap was expressed under

a 20,000 lb load using a Carver Laboratory Press, Model B (FS Carver Inc., Summit,

NJ). The sap was centrifuged to remove particulate matter. The supernatant (0.55 mL)

was frozen at -20°C until required. Embryo sap pH was 6.667 after thawing. The

divalent metal ion chelator CDTA (rra/w-1,2-diaminocyclohexane-N,N,N’ ,N’-

tetraacetic acid) (0.1 M solution/pH 6.6) was added to a final concentration of 7.4

mM. The solution standard of 10 mM phosphate (dibasic sodium salt)/5 mM phytic

acid (dodecasodium salt)/40 mM MOPS (3-[N-Morpholino]propanesulfonic acid) at

pH 7.0 was made up in 90% H2O/10% D20 .

3,P NMR

Spectra were recorded on a Bruker AM 400 spectrometer operating at 162

MHz with high power broadband proton decoupling. A 2H lock signal was used.

Single pulse and spin echo experiments were performed. Spin echo experiments used

94

the Carr-Purcell-Meiboom-Gill (CPMG) sequence (90°-[D2-180o-D2]n-Acq-D,)

(Meiboom and Gill, 1958; Derome, 1987). The spin echo sequence was required to

resolve the inorganic phosphate peak from the broad phytate peaks seen in red rice

embryos. Due to their shorter transverse relaxation times (T2), the broad phytate peaks

are allowed to decay during the echo or refocusing cycle [D2-180°-DJ.

A 5 mm broad band probe was used to acquire 31P NMR spectra of embryo

sap and phosphate/phytate standards. The 5mm probe acquired spectra using a 90°

pulse angle of 10 usee, an acquisition time of 1.44 s, and recycle delay of 1.0 s. Spin

echo experiments used a D t of 1 s, D2 varied, and only one refocusing cycle (n = 2)

was used. A 10 mm probe was used to acquire spectra of perfused embryos using the

following acquisition parameters: a 90° pulse angle of 19 nsec, an acquisition time

of 1.016 s, and recycle delay of 2.0 s. Spin echo experiments (10 mm probe) used a

B 1 of 1 s, D2 of 4 ms. Two refocusing cycles (n = 4) were used to allow decay of

residual signals. Unless otherwise stated each spectrum consists of 256 scans obtained

with 32K data points. The resulting free induction decays were Fourier transformed

after the application of exponential multiplication with line broadening of 20 Hz.

Signals were referenced to the TEMDP resonance at 22.49 ppm relative to 85%

phosphoric acid at 0 ppm.

Results

31P NMR of phosphate solutions and embryo sap

The feasibility of using the CPMG spin echo sequence to eliminate the phytate

signals was tested using a phosphate/phytate solution. Single pulse spectra show the

four phytate resonances and that of inorganic phosphate (Fig. 6.2A). When the spin

echo sequence was employed, the phytate resonances were seen to decay more rapidly

than that of inorganic phosphate. A D2 of 55 ms resulted in the almost total

disappearance of the phytate resonances (Fig. 6.2B).

Single pulse spectra of embryo sap revealed four peaks (Fig. 6.3A). The

application of the spin echo sequence with a D2 of 8 ms resulted in the collapse of the

phytate signals leaving the Pj peak at 2.1 ppm and a low intensity phytate signal at 2.6

ppm (Fig. 6.3B). Peaks were identified by comparison of proton coupled and

decoupled spectra. The proton coupled signals of the phosphate groups of phytate

exhibit signal splitting (3/ c.H = 5 to 10 Hz). The signal at 2.1 ppm did not exhibit

signal splitting in coupled spectra, and its identity as P; was confirmed by the addition

of P;.

3IP NMR of dormant red rice embryos

Single pulse spectra of dormant embryos produced a broad peak yielding little

detail. Application of the spin echo sequence revealed a narrow peak at 1.99 ppm

(Fig. 6.4). Subjecting embryos to acid loading by the permeant weak acid nitrite

produced changes in the spectra (Fig. 6.5). After 5 min exposure to nitrite (pH 2.2),

96

B

»--1—' ........... i 16 . 5 6 . 0

1— I— 1— l _

5 . 0 PPM

- "■»■ ■ I ■ 4 . 5 4 . 0 3 . 55 . 5

I i i i i (■

Figure 6.2. (A) Single pulse spectra (64 scans) and (B) spin echo spectra (64 scans) of phosphate/phytate solution at pH 7.0. The inorganic phosphate peak is (a), all other peaks are phytate signals.

97

a

T i T i T n - i T p i m i t t i i j ' i n r i i m n 11 r i T i m | -> m n 1 1 1 n 1 1 n i 1 1 1 1 1 i rm i n » T i i r » r u t

4 .0 2 .0 0 .0PPM

Figure 6.3. (A) Single pulse spectra (127 scans) and (B) spin echo spectra (352 scans) of CDTA amended embryo sap. The inorganic phosphate peak is (a).

98

V'/V^A/VvAx/ ^ / v

\\

B iH/ y

5 . 0 0 . 0PPM

- 5 . 0

Figure 6.4. (A) Single pulse spectra and (B) spin echo spectra of perfused red rice embryos.

99

Figure 6.5. Spin echo spectra of perfused red rice embryos following increasing contact with 20 mM NaN02 at pH 2.2. Spectra represent, (A) control, (B) 5 min contact, (C) 30 min contact, (D) 60 min contact. Control peak (a); new emerging peak (b).

100

5 . 0 0 . 0PPM

- 5 . 0

101

A

B

C

5 . 0 0 . 0PPM

- 5 . 0

Figure 6 .t>/Spin echo spectra of perfused red rice embryos following increasing periods at pH 8.3. Spectra represent, (A) 5 min contact, (B) 15 min contact, (C) 30 min contact. Peak produced by acidification in figure 6.5 is (a); peak produced by treatment at pH 8.3 is (b).

PP

M

102

j ' - — I ■ I | . . .

8 . 0 6 . 0 4 . 0 2 . 0 1................. 1— " ' • I .............................. ....

- 4 . 0 - 5 . 0 - 8 . 0- 2 . 0

Figure 6.7. Spin echo spectra of perfused red rice embryos following the addition of 20 mM inorganic phosphate (a) at pH 7.9.

103

a new peak appeared upfield at 1.25 ppm (Fig. 6 .6B). Over time, the intensity of this

peak increased whilst that at 1.99 ppm decreased. After 100 min the pH of the

perfusate was adjusted to 8.3 to stop acid loading and test the effect of high external

pH on the signals observed. After 5 min at pH 8.3, a peak started to build downfield

at 1.71 ppm. Over the next 30 min this peak continued to increase in intensity and

migrated further downfield (Fig. 6.7). The addition of 20 mM phosphate (pH 7.9)

produced a large external phosphate peak (Fig. 6.7), demonstrating that the pH

sensitive signal assumed to be inorganic phosphate (1.99 ppm in Fig. 6.3A) was

internal to the tissue.

Discussion

Single pulse spectra of seeds and embryos typically show a single broad peak

consisting of the overlapping signals from phytate and inorganic phosphate (Kime et

al., 1982; Delfini et al., 1985). Spectra of red rice embryos are no exception. Single

pulse 31P NMR has been used to measure the pH; of phytate rich tissues by observing

the chemical shift of the internal Pj (Martin et al., 1987; Candelier et al., 1989; Pdtel

et al., 1992). However, phytate signals can mask the position of the internal Pj peak

(Delfini et al., 1985; Martin et al., 1987; Candelier et al., 1989). This was not

considered to be a factor in the broad spectra of cotton embryos as phytate signals

were not detected in embryo homogenates (Hendrix et al., 1987), despite the likely

presence of phytate in the adhering aleurone (Lui and Altschul, 1967). However,

104

phytate in embryo homogenates readily precipitates in the presence of Ca2+ and Mg2+,

especially above pH 7.5. In the solid state it will not be detected by NMR. Phytate

will also be hydrolyzed if phosphatases are not denatured (personal observations;

Hayakawa et al., 1990). Therefore, the cotton embryo spectra probably represent an

amalgam of inorganic phosphate and phytate signals.

Similar problems were encountered when 31P NMR was used to measure

changes in pHj upon fertilization and artificial activation of Xenopus laevis (Nuccitelli

et al., 1981), Rampipiens Schreber (Morrill et al., 1984), and Carcinus maenas L.

eggs (Goudeau et al., 1989; Hervd et al., 1989). The broad signals of yolk phospho-

proteins necessitated the use of spin echo sequences in order to resolve the internal

Pi peak. In red rice, application of the CPMG spin echo sequence reduced the

intensity of the phytate signals in vitro enabling the Pj signal to be identified (Fig.

6.3). In vivo a single narrow peak emerged above the residual signals of the

previously broad peak seen in the single pulse spectra (Fig. 6.4). This peak is pH

sensitive, moving upfield when the tissue is acidified with nitrite. The appearance of

two peaks during embryo acidification is assumed to represent two separate cell

populations -those yet to be acidified and those that have been acidified. The effect

of embryo acidification was slowly reversed by increasing the external pH. Permeant

weak acids and rapid changes in external pH have previously been shown to effect

internal pH (Guem et al., 1986; Fox and Ratcliffe, 1990).

The proposed internal Pj signal was not conclusively identified as internal P;.

However, it is (a) internal to the tissue as shown by the addition of external Pb and

105

(b) pH sensitive. It is unlikely to be a composite of the two most intense phytate peaks

as they would split into separate peaks as the internal pH changed (Costello et al.,

1976; Zuiderweg et al., 1979). If the peak in question is internal P; it would not

exhibit signal splitting in a proton coupled spectrum as found for embryo sap. Further

confirmation of the internal peak as P; would result from (a) the absence of signal

splitting in a proton coupled spectrum; and (b) a decrease in signal intensity during

choline uptake as internal Pj is sequestered as phosphocholine (Bligny et al., 1990).

In spin echo experiments it was necessary to decrease the value of Dz as

samples progressed from an aqueous solution of phosphate/phytate (55 ms), to embryo

sap (8 ms), to embryos (4 ms). This indicated increasing transverse relaxation due to

increased interaction between excited nuclei (Gadian, 1982) as solute concentration

increased. The implication is that the environment experienced by Pj in the embryo

sap is close to that in vivo.

It was not possible to complete this study due to unresolved technical

difficulties in acquiring a signal in spin echo experiments with embryos subsequent to

those shown here. However, spin echo experiments were successfully used on several

occasions to remove the phytate signals from embryo spectra. The CPMG spin echo

sequence clearly offers an opportunity to detect the internal P; signal in red rice

embryos. Once the present technical difficulties are overcome, this protocol will

enable the in vivo measurement of pH; in the dense tissues of the red rice embryos

during dormancy-breaking chemical treatments and subsequent germination. The

technique also has potential for long term monitoring of metabolism and pH; in

106

somatic seed embryos. In summary, this is the first report of a spin echo sequence

being used to visualize the P; signal in phytate rich plant tissues.

CHAPTER 7

CONCLUSION

The aim of this research was to identify physiological and biochemical

markers, prior to radicle emergence, that could be used to (a) differentiate between

hydrating dormant and nondormant seeds; (b) identify the transition from the dormant

to nondormant state during a dormancy-breaking treatment; and (c) identify the onset

of the germination phase. This research provided detailed information as to changes

in physiological and biochemical markers pre-, during, and post-contact with

dormancy-breaking chemicals.

Differentiating between hydrating dormant and nondormant seeds

The classical triphasic pattern of hydration (Fig. 2.1 & 3.1 A) has been used

to great effect in red rice. In nondormant seeds, the markers studied conformed to the

temporal phases of the hydration pattern (seed respiration, embryo pH and [Fru 2,6-

P2]). Desiccation tolerance was lost when visible germination was first seen (i.e., at

the transition from phase 2 to 3). In dormant seeds, seed respiration, embryo pH and

[Fru 2,6-PJ enter phase 2 similar to nondormant seeds, with embryo [Fru 2,6-PJ

showing a transient peak. In the phase 2 of dormant seeds, these markers reflect the

basal metabolic activity required to maintain viability. The divergence of markers in

dormant vs. nondormant seeds during phase 2 is associated with the onset of the

107

108

germination phase. In nondormant seeds, seed respiration increased, and embryo pH

decreased, while in dormant seeds they remained constant in phase 2. Embryo [Fru

2,6-P2] was sustained in nondormant but was transient in dormant seeds at this time.

Dormancy-breaking

Embryo acidification during chemical contact is proposed as a marker for the

termination of dormancy. A second acidification event occurs upon visible germination

(Fig. 7.1). Embiyo [Fru 2,6-PJ does not emulate this pattern. Increases in [Fru 2,6-

P2] during chemical contact were not related to dormancy-breaking, but to the

subsequent length of the germination phase and rate of germination (Fig. 7.1). The

pH and Fru 2,6-P2 markers seem to be independent of one another as dormancy is

broken in the absence of elevated embryo [Fru 2,6-P2] (compare propionaldehyde and

propionate). The chemicals employed in these studies exhibit dual activity: acting as

(a) dormancy-breaking agents, and (b) germination stimulants.

Dormancy-breaking chemicals without a dissociable proton acidify embryos to

a similar extent (Table 3.2). One explanation for this is their metabolism to a weak

acid. This was shown to be so, with metabolism of n-propanol to 3-hydroxypropionate

by dormant embryos. The inability of a compound to break dormancy has been

equated with an inability to be metabolized based on structure activity studies (Cohn

et al., 1991). All told, these data indicate the activation of processes that respond

independently of one another but which act coordinately i.e., those related to

dormancy-breaking (pH) and the germination phase (Fru 2,6-PJ.

109

Embrvo pH

(0o

o

IB

D/ND transition

N

Embrvo Fructose 2.6-bisphosohate

co(0

cf l>ocoO

T i m e

Length of germination phase

Germinationr a t e

Figure 7.1. Paths of embryo pH and fructose 2,6-bisphosphate during dormancy- breaking and germination. Hydrated dormant seeds are exposed to a dormancy- breaking chemical pulse (A). Following dormancy-breaking visible germination is seen (B). Embryos are acidified during chemical contact (C) and during visible germination (D). Embryo Fru 2,6-P2 levels increase in response to some chemicals (E) and with visible germination (F). The higher the embryo [Fru 2,6-PJ the shorter the germination phase and the greater the subsequent rate of germination.

110

Future research

This research has identified several areas worthy of further research.

(1) The Fru 2,6-P2 transient seen in hydrating dormant seeds indicates that the

germination phase is blocked. The activities of PFK2 and PPr PFK are regulated by

sulfhydryl status (El-Magrabi et al., 1990; Kiss et al., 1991). The thioredoxin system

is important in regulating enzyme sulfhydryl status (Buchanan, 1991). Therefore,

changes in the activity of this system during hydration and dormancy-breaking may

be early events in metabolic activation.

(2) Short chemical contact periods can (a) break dormancy and (b) induce

physiological responses not associated with dormancy-breaking. The rapidity of these

responses provides an opportunity to probe for changes in gene expression associated

with both dormancy-breaking and the germination phase within a narrow time frame

(Cohn and Footitt, 1992).

(3) 13C NMR was successfully used to study the metabolic fate of dormancy-

breaking chemicals. This demonstrates the potential of tracer methods for studying the

fate of dormancy-breaking chemicals, such as ketones and secondary alcohols. The

location of the metabolic blocks imposed on dormant seeds during hydration may be

identified using tracer methods.

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APPENDIX A

ACCOUNTING FOR DIFFERENCES BETWEEN 24H PULSE

AND OH POST-PULSE EMBRYO pH

128

129

When measuring embryo homogenate pH, a discrepancy was found between

the 2 4h/30°C time point at the end of the chemical pulse in experiments separated

by several months (Figs. 3.3 & 3.5; Append. D). The cause was traced to a faulty

pipette, which resulted in a reduction in the pH measurements in the post pulse

period.

Eppendorf 100-1,000 juL pipette used for all tissue pH determinations.

Pipette delivered 916.8 + 4.4 nL when set at 1,000 /uL.

If 100 embryo extracted into 1 mL gives a pH of 7.0, then

pH 7.0 = 1 x 10'10 moles mL'1

If only 916.8 /ul is used to extract 1 x 10"10 moles mL'1, then

1000/916.8 /;L x (1 x lO'^mL'1) = 1.09075 x 10'10 moles mL'1

= 1.09075 x 10'7 moles L '1

= pH 6.962

This accounts for most of the difference observed between these time points. Since the

exact time of occurrence of the pipette problem is unknown and the problem was

recognized after all data were collected, the uncorrected values are presented in the

figures.

130

Table A.I. Embryo homogenate pH as measured in 1989, recalculated for extraction in 916.8 nL, and determined again in 1990 using the same harvest (SH 87-1)

Embryo pH after 24 h/30°C chemical contact

1989 1989 recalculated 1990

Treatment pH +SE -SE pH +SE -SE pH +SE -SE

Control pH 4.87 7.192.01 .01 7.154.009 .01 7.140.031 .029

Propionate 6.859 .047 .043 6.822 .048 .043 6.878 .003 .003

Propanol 7.144 .004 .004 7.106 .004 .003 7.047 .009 .008

Control pH 7.0 7.387 .027 .025 7.349 .027 .026 7.271 .017 .017

Propanol 7.239 .016 .015 7.201 .015 .015 7.177 .016 .015

Propionaldehyde 7.184 .03 .028 7.146 .03 .028 7.096 .011 .011

Methyl propionate 7.161 .028 .027 7.123.029 .027 7.048.023 .022

APPENDIX B

EMBRYO pH DURING DRY AFTERRIPENING

131

132

Embryo pH differed between dry dormant and nondormant seeds and in the

initial stages of hydration. It was decided to see whether this change could be caused

by dry afterripening. Four samples of dormant harvest 1990-1 were placed in 1 L

Mason jars at room temperature. Every five days 110 seeds were taken, manually

dehulled and hydrated for 6 h/30°C in 9 cm Petri plates on three sheets of

germination paper with 10 mL distilled H20 . Seeds were covered with a sheet of

tissue. At the same time standard germination tests were set up. After 6 h, 100 seeds

were taken, and embryo and endosperm homogenate pH were determined. The mean

afterripening temperature over 85 days was 22.7 ± 0.5°C. The mean viability for all

germination tests was 99.9 ± 0.1% (n=72).

Embryo pH was stable for 55 days at which time dormancy had decayed.

After this time embryo pH started to decline, presumably due to the increasing

kinetics of germination (data not collected).

It was concluded that differences in embryo pH resulting from dry

afterripening were too small to detect in a consistant manner during the early stages

of hydration. Embryo pH follows a downward trend but this may not be significant.

Figure B .l. Embryo and endosperm pH of dehulled seeds hydrated for 6 h/30°C, and germination (7 d/30°C) after increasing periods of dry afterripening at room temperature of intact dormant seeds.

APPENDIX C

HARVEST COMPARISON

134

135

7 .0

Figure C .l. Comparison of embryo and endosperm pH of dehulled seeds from four separate harvests of dormant red rice, after hydration for 24 h/30°C (all escapes were replaced). Homogenate pH was determined as described in materials and methods. After 24 h/30°C on H20 there was no significant difference between the four harvests.

APPENDIX D

DATA TABLES FOR CHAPTER 3

136

Table D .l . Embryo data for hydrating dehulled seeds of dormant red rice at 30°C.

Hours pH -SE +SE Fr Wt +SE Dr Wt +SE Fresh Wt Dry Wt %Germ SE

pg emb'1 pg emb4 %MC ±SE %MC +SE

0 7.368 0.024 0.025 461.0 18.6 412.0 19.0 10.9 1.0 12.1 1.2 0 0

1 7.352 0.018 0.018 496.0 8.0 366.0 4.0 26.5 0.7 35.8 1.4 0 0

2 7.281 0.017 0.017 616.0 24.0 416.0 22.0 32.8 0.7 48.6 2.0 0 0

4 7.242 0.012 0.013 704.0 10.0 410.0 8.0 41.6 0.5 71.8 1.0 0 0

6 7.283 0.008 0.008 752.0 20.0 416.0 12.0 44.4 0.1 80.3 0.4 0 0

8 7.258 0.024 0.027 768.0 14.0 418.0 10.0 45.4 0.3 83.6 1.2 0 0

10 7.306 0.011 0.011 756.0 20.0 406.0 14.0 46.2 0.3 86.3 1.0 0 0

12 7.276 0.015 0.016 760.0 4.0 398.0 2.0 47.3 0.3 90.6 1.0 0 0

Table D .2. Embryo data for hydrating dehulled seeds o f nondormant red rice at 30°C.

Hours pH -SE +SE Fr Wt +SE Dr Wt ±SE Fresh Wt Dry Wt %Germ SE

fig emb'1 fig emb'1 %MC ±SE %MC +SE

0 7.255 0.026 0.027 0 0

1 7.252 0.025 0.026 502.0 10.0 368.0 8.0 26.5 0.7 36.3 1.3 0 0

2 7.227 0.011 0.011 574.0 6.0 388.0 6,0 32.2 0.8 47.9 1.6 0 0

4 7.161 0.011 0.011 622.0 34.0 426.0 40.0 41.3 0.3 71.0 0.9 0 0

6 7.164 0.007 0.007 732.0 22.0 410.0 14.0 44.0 0.4 78.7 1.3 0 0

8 7.065 0.015 0.015 768.0 4.0 412.0 2.0 46.2 0.4 86.3 1.4 0 0

10 7.050 0.019 0.020 846.0 40.0 452.0 38.0 47.6 1.2 88.8 6.8 0 0

12 7.011 0.013 0.014 802.0 14.0 376.0 8.0 52.9 0.6 113.0 2.7 5 1

14 6.932 0.016 0.016 874.0 28.0 358.0 2.0 57.0 0.8 143.3 6.1 29 8

Table D .3. Endosperm data for hydrating dehulled seeds o f dormant red rice at 30°C.

Hours pH -SE +SE Fr Wt ±SE Dr Wt +SE Fresh Wt Dry Wt

mg end1 mg end'1 %MC ±SE %MC ±SE

0 7.656 0.020 0.021 17.4 0.1 15.0 0.1 13.7 0.1 15.9 0.1

1 7.554 0.038 0.042 18.6 0.1 15.2 0.1 18.3 0.1 22.6 0.2

2 7.538 0.035 0.038 19.3 0.1 15.3 0.1 21.0 0.2 26.7 0.1

4 7.495 0.009 0.009 20.1 0.0 15.3 0.0 24.0 0.2 31.5 0.3

6 7.425 0.079 0.097 20.3 0.1 15.1 0.0 25.5 0.0 34.3 0.1

8 7.510 0.024 0.026 20.5 0.1 15.1 0.0 26.3 0.1 35.7 0.1

10 7.512 0.025 0.027 20.5 0.1 14.9 0.2 27.1 0.5 37.2 0.9

12 7.529 0.024 0.026 20.7 0.2 15.1 0.1 27.0 0.0 37.0 0.0

Table D .4. Endosperm data for hydrating dehulled seeds o f nondormant red rice at 30°C.

Hours pH -SE +SE

0 7.496 0.051 0.058

1 7.483 0.042 0.047

2 7.435 0.062 0.073

4 7.384 0.022 0.023

6 7.390 0.019 0.019

8 7.451 0.015 0.016

10 7.365 0.024 0.025

12 7.349 0.019 0.020

14 7.324 0.061 0.071

Fr Wt ±SE Dr Wt ±SE

mg end1 mg end"1

18.6 0.2 15.2 0.1

19.0 0.1 15.1 0.1

19.6 0.9 15.0 0.1

20.3 0.1 15.2 0.1

20.3 0.1 15.1 0.1

20.5 0.2 15.1 0.1

20.7 0.1 15.2 0.1

20.6 0.1 15.1 0.1

Fresh Wt Dry Wt

%MC ±SE %MC ±SE

18.1 0.1 22.1 0.2

20.4 0.2 25.7 0.4

23.6 0.1 30.9 0.2

25.1 0.1 33.5 0.1

25.9 0.1 34.9 0.2

26.4 0.1 35.8 0.1

26.5 0.1 36.1 0.2

27.0 0.2 36.9 0.3

141

Table D.5. Effect of nitrite at pH 3.0 on embryo pH and germination, as in figure 3.2.

pH 3.0 Control 10 mM Nitrite

Germination Germination

Hours pH -SE +SE % ±SE pH -SE +SE % ±

-4 7.199 0.057 0.065 0 0 7.260 0.054 0.062 0 0

-3 7.099 0.049 0.055 0 0 6.894 0.023 0.025 0 0

-2 7.071 0.024 0.025 0 0 6.814 0.022 0.024 0 0

-1 7.075 0.012 0.013 0 0 6.803 0.024 0.025 0 0

0 7.077 0.040 0.044 0 0 6.787 0.037 0.040 0 0

1 7.249 0.038 0.042 0 0 6.885 0.019 0.020 0 0

2 7.199 0.015 0.015 2 1 6.896 0.009 0.010 0 0

4 7.250 0.028 0.030 1 0 6.948 0.018 0.019 0 0

6 7.229 0.019 0.019 3 1 6.941 0.011 0.012 0 0

8 7.229 0.009 0.009 4 2 6.972 0.013 0.014 1 0

10 7.213 0.018 0.018 6 2 6.931 0.019 0.019 3 1

12 7.202 0.019 0.020 6 1 6.904 0.015 0.016 11 2

142

Table D.6. Effect of nitrite at pH 7.0 on embryo pH as in figure 3.2.

pH 7.0 Control 10 mM Nitrite

Hours pH -SE +SE pH -SE +SE

-4 7.274 0.006 0.006 7.274 0.017 0.017

-3 7.328 0.031 0.033 7.324 0.007 0.007

0 7.300 0.026 0.027 7.372 0.010 0.010

Table D.7. Effect of 10 mM nitrite at pH 3.0 on endosperm pH as in figure 3.2.

pH 3.0 Control 10 mM Nitrite

Hours pH -SE +SE pH -SE +SE

-4 7.527 0.036 0.040 7.524 0.040 0.044

-3 7.432 0.055 0.063 7.286 0.040 0.044

-2 7.354 0.041 0.045 7.209 0.023 0.025

-1 7.348 0.034 0.036 7.237 0.017 0.017

0 7.380 0.028 0.030 7.209 0.026 0.028

1 7.469 0.041 0.045 7.215 0.043 0.047

2 7.509 0.021 0.022 7.192 0.032 0.035

4 7.477 0.018 0.019 7.250 0.017 0.017

6 7.479 0.002 0.002 7.248 0.010 0.011

8 7.485 0.019 0.020 7.298 0.030 0.032

10 7.465 0.014 0.015 7.276 0.059 0.069

12 7.463 0.024 0.026 7.326 0.031 0.033

144

Table D.8. Effect of nitrite at pH 7.0 on endosperm pH as in figure 3.2.

pH 7.0 Control 10 mM Nitrite

Hours pH -SE +SE pH -SE +SE

-4 7.485 0.019 0.020 7.470 0.027 0.029

-3 7.469 0.033 0.035 7.528 0.026 0.027

0 7.502 0.047 0.053 7.548 0.016 0.017

145

Table D.9. Effect of 25 mM citrate/phosphate buffer at pH 4.9 on embryo pH and germination as in figure 3.3.

Pulse Post-pulse

Germination Germination

in real time in real time

Hours pH -SE +SE % ±SE pH -SE +SE % ±

0 7.293 0.009 0.009 0 0 7.140 0.030 0.032 - -

1 7.229 0.020 0.021 0 0 7.135 0.030 0.032 4 1

2 7.214 0.013 0.014 0 0 7.147 0.007 0.007 1 0

4 7.213 0.015 0.016 1 0 7.099 0.018 0.018 3 1

8 7.202 0.011 0.012 1 0 7.108 0.018 0.019 4 0

12 7.188 0.021 0.022 1 1 7.117 0.022 0.024 3 1

16 7.217 0.017 0.018 1 1 7.137 0.014 0.015 9 2

20 7.209 0.021 0.022 2 1 - - - - -

24 7.192 0.010 0.010 2 1 7.160 0.027 0.028 0 0

Table D .10. Effect o f 20 mM propionate at pH 4.9 on embryo pH and germination as in figure 3.3 & 3.5.

Pulse Post-pulse

Germination Dormancy-breaking Germination

in real time with contact in real time

Hours pH -SE +SE % ±SE % ±SE pH -SE +SE % ±SE

0 7.292 0.028 0.030 0 0 4 0 6.879 0.003 0.003 0 0

1 7.211 0.007 0.007 0 0 6.855 0.022 0.023 2 0

2 7.145 0.011 0.011 0 0 3 1 6.889 0.024 0.026 2 1

4 7.090 0.013 0.013 1 1 4 1 6.922 0.020 0.021 2 1

8 7.045 0.014 0.015 0 0 7 1 7.020 0.006 0.006 6 3

12 6.913 0.025 0.026 1 0 36 8 7.046 0.011 0.011 4 1

16 6.902 0.017 0.018 0 0 74 10 7.013 0.014 0.014 9 1

20 6.917 0.043 0.047 1 0 88 3 6.948 0.014 0.015 12 1

24 6.860 0.043 0.048 2 1 94 2 6.911 0.018 0.019 21 3

40 \

Table D . l l . Effect o f 70 mM n-propanol at pH 4.9 on embryo pH and germination.

Pulse Post-pulse

Germination Dormancy-breaking Germination

in real time with contact in real time

Hours pH -SE +SE % +SE % +SE pH -SE +SE % +SE

0 7.324 0.019 0.020 0 0 7.047 0.009 0.009 3 1

1 7.249 0.022 0.024 0 0 7.059 0.003 0.003 0 0

2 7.231 0.011 0.012 0 0 2 0 7.104 0.007 0.007 0 0

4 7.191 0.017 0.017 0 0 3 1 7.053 0.005 0.005 1 0

8 7.154 0.005 0.005 0 0 5 1 7.054 0.011 0.012 3 1

12 7.137 0.023 0.024 0 0 11 2 7.020 0.021 0.022 6 2

16 7.179 0.025 0.027 0 0 47 6 6.982 0.019 0.019 10 2

20 7.141 0.009 0.009 1 0 72 4 6.906 0.025 0.026 14 2

24 7.144 0.004 0.004 0 0 93 2 6.830 0.017 0.018 28 2

148

Table D.12. Effect of 25 mM citrate/phosphate buffer at pH 7.0 on embryo pH and germination as in figure 3.3 & 3.5.

Pulse Post-pulse

Germination

in real time

Hours pH -SE +SE % ±SE pH -SE +SE %

0 7.308 0.015 0.016 0 0 7.271 0.017 0.018 1

1 7.362 0.003 0.003 0 0 7.237 0.019 0.020 0

2 7.359 0.017 0.018 0 0 7.203 0.021 0.022 1

4 7.372 0.008 0.008 0 0 7.206 0.021 0.023 1

8 7.356 0.019 0.020 1 0 7.200 0.024 0.025 2

12 7.357 0.011 0.012 2 1 7.199 0.012 0.013 1

16 7.384 0.024 0.026 1 0 7.143 0.032 0.035 3

20 7.387 0.009 0.009 1 1

24 7.387 0.026 0.027 1 1 7.188 0.016 0.017 3

Germination

in real time

+SE

0

0

Table D .13. Effect of 70 mM n-propanol at pH 7.0 on embryo pH and germination as in figure 3.3 & 3.5.

Pulse Post-pulse

Germination Dormancy-breaking Germination

in real time with contact in real time

Hours pH -SE +SE % ±SE % ±SE pH -SE +SE % ±SE

0 7.298 0.005 0.005 0 0 2 1 7.177 0.015 0.016 0 0

1 7.334 0.016 0.016 0 0 - 7.036 0.043 0.048 0 0

2 7.324 0.005 0.005 0 0 3 0 7.132 0.013 0.014 0 0

4 7.301 0.029 0.031 0 0 3 0 7.105 0.016 0.016 1 0

8 7.303 0.003 0.003 0 0 5 1 7.123 0.028 0.030 0 0

12 7.307 0.013 0.013 0 0 12 3 7.077 0.009 0.010 2 1

16 7.252 0.014 0.014 0 0 40 5 7.025 0.021 0.022 6 1

20 7.237 0.020 0.021 0 0 62 3 6.984 0.017 0.017 10 2

24 7.239 0.016 0.016 1 1 88 2 6.908 0.043 0.047 24 3

M3

Table D .14. Effect o f 40 mM propionaldehyde at pH 7.0 on embryo pH and germination as in figure 3.3 & 3.5.

Pulse Post-pulse

Germination Dormancy-breaking Germination

in real time with contact in real time

Hours pH -SE +SE % ±SE % ±SE pH -SE +SE % ±SE

0 7.287 0.048 0.054 0 0 2 0

1 7.305 0.013 0.014 0 0

2 7.243 0.056 0.064 0 0 4 0

4 7.304 0.018 0.018 0 0 6 1

8 7.215 0.012 0.012 0 0 19 4

12 7.197 0.009 0.009 1 0 51 13

16 7.209 0.021 0.022 1 0 78 2

20 7.189 0.015 0.015 5 1 97 1

24 7.184 0.028 0.030 8 2 97 1

7.096 0.011 0.012 9 1

6.968 0.020 0.020 13 2

6.994 0.006 0.006 10 1

6.998 0.014 0.014 17 3

6.880 0.030 0.033 31 4

6.771 0.015 0.016 49 3

O

Table D .15. Effect o f 30 mM methyl propionate at pH 7.0 on embryo pH and germination as in figure 3.3 & 3.5.

Pulse Post-pulse

Germination Dormancy-breaking Germination

in real time with contact in real time

Hours pH -SE +SE % +SE % +SE pH -SE +SE % +SE

0 7.290 0.011 0.012 0 0 2 1 7.048 0.022 0.023 0 0

1 7.308 0.017 0.018 0 0 6.994 0.018 0.019 1 0

2 7.263 0.028 0.030 0 0 2 1 7.035 0.031 0.033 0 0

4 7.236 0.029 0.031 1 0 4 1 6.985 0.009 0.009 1 0

8 7.179 0.022 0.023 0 0 5 0 7.032 0.017 0.018 1 0

12 7.168 0.016 0.017 1 0 12 3 7.072 0.015 0.015 3 1

16 7.191 0.012 0.012 0 0 50 4 7.076 0.019 0.020 2 1

20 7.148 0.010 0.010 1 0 85 3 6.984 0.037 0.040 8 2

24 7.161 0.027 0.029 1 0 92 1 6.940 0.004 0.004 12 2

152

Table D .16. Effect o f 25 mM citrate/phosphate buffer at pH 4.9 on endosperm pHas in figure 3.4 & 3.6.

Pulse Post-pulse

Hours pH -SE +SE pH -SE +SE

0 7.521 0.041 0.045 7.320 0.038 0.042

1 7.519 0.035 0.039 7.337 0.023 0.024

2 7.437 0.025 0.026 7.322 0.024 0.026

4 7.461 0.034 0.037 7.395 0.023 0.024

8 7.490 0.016 0.017 7.294 0.053 0.060

12 7.474 0.036 0.040 7.296 0.023 0.025

16 7.488 0.029 0.031 7.337 0.024 0.025

20 7.426 0.028 0.030 - - -

24 7.448 0.019 0.020 7.322 0.037 0.041

153

Table D .17. Effect o f 20 mM propionate at pH 4.9 on endosperm pH as in figure 3.4& 3.6.

Pulse Post-pulse

Hours pH -SE +SE pH -SE +SE

0 7.545 0.023 0.024 7.094 0.008 0.008

1 7.504 0.013 0.014 7.055 0.025 0.027

2 7.474 0.011 0.011 7.072 0.016 0.016

4 7.415 0.023 0.024 7.053 0.044 0.049

8 7.359 0.022 0.024 7.113 0.025 0.027

12 7.277 0.014 0.015 7.136 0.016 0.017

16 7.216 0.025 0.026 7.167 0.014 0.015

20 7.190 0.029 0.031 7.173 0.015 0.016

24 7.128 0.021 0.022 7.199 0.022 0.023

154

Table D.18. Effect of 70 mM n-propanol at pH 4.9 on endosperm pH.

Pulse Post-pulse

Hours pH -SE +SE PH -SE +SE

0 7.567 0.011 0.011 7.310 0.020 0.020

1 7.538 0.032 0.035 7.260 0.025 0.026

2 7.520 0.031 0.034 7.251 0.038 0.042

4 7.483 0.024 0.025 7.193 0.029 0.031

8 7.463 0.046 0.052 7.239 0.021 0.022

12 7.464 0.013 0.014 7.168 0.035 0.037

16 7.418 0.027 0.029 7.194 0.024 0.026

20 7.399 0.015 0.016 7.156 0.017 0.018

24 7.396 0.018 0.019 7.148 0.022 0.023

155

Table D .19. Effect o f 25 mM citrate/phosphate at pH 7 .0 on endosperm pH as infigure 3.4 & 3.6.

Pulse Post-pulse

Hours PH -SE +SE pH -SE +SE

0 7.539 0.020 0.021 7.410 0.034 0.037

1 7.564 0.030 0.032 7.427 0.023 0.024

2 7.601 0.035 0.039 7.334 0.042 0.047

4 7.633 0.024 0.026 7.339 0.033 0.036

8 7.588 0.025 0.027 7.346 0.033 0.036

12 7.598 0.039 0.043 7.419 0.030 0.033

16 7.668 0.017 0.018 7.366 0.022 0.024

20 7.656 0.024 0.026

24 7.609 0.019 0.020 7.322 0.034 0.037

Table D .20. Effect o f 70 mM /7-propanol at pH 7.0 on endosperm pH as in figure 3.4& 3.6.

Pulse Post-pulse

Hours pH -SE +SE PH -SE +SE

0 7.550 0.019 0.019 7.351 0.033 0.036

1 7.573 0.016 0.017 7.307 0.047 0.052

2 7.632 0.018 0.019 7.285 0.040 0.044

4 7.574 0.011 0.011 7.282 0.030 0.032

8 7.583 0.016 0.017 7.272 0.026 0.027

12 7.576 0.007 0.008 7.258 0.021 0.022

16 7.563 0.011 0.011 7.279 0.015 0.015

20 7.521 0.018 0.019 7.256 0.021 0.022

24 7.496 0.007 0.007 7.233 0.048 0.053

157

Table D .21. Effect o f 40 mM propionaldehyde at pH 7.0 on endosperm pH as infigure 3.4 & 3.6.

Pulse Post-pulse

Hours pH -SE +SE pH -SE +SE

0 7.526 0.019 0.020 7.329 0.047 0.053

1 7.559 0.017 0.018 7.250 0.030 0.032

2 7.586 0.043 0.048 7.249 0.017 0.017

4 7.578 0.031 0.033 7.264 0.010 0.011

8 7.512 0.037 0.035 7.213 0.030 0.032

12 7.485 0.013 0.013 7.178 0.027 0.028

16 7.472 0.021 0.022

20 7.412 0.015 0.016

24 7.429 0.027 0.029

158

Table D .22. Effect o f 30 mM methyl propionate at pH 7.0 on endosperm pH as infigure 3.4 & 3.6.

Pulse Post-pulse

Hours pH -SE +SE pH -SE +SE

0 7.510 0.020 0.021 7.171 0.027 0.029

1 7.547 0.024 0.026 7.166 0.035 0.038

2 7.520 0.014 0.014 7.169 0.038 0.042

4 7.509 0.016 0.017 7.130 0.031 0.034

8 7.394 0.015 0.015 7.205 0.022 0.023

12 7.348 0.022 0.023 7.165 0.030 0.032

16 7.352 0.022 0.024 7.214 0.027 0.029

20 7.322 0.004 0.004 7.181 0.022 0.023

24 7.263 0.017 0.018 7.150 0.024 0.025

159

Table D.23. Embryo pH in the dormant 1990-1 harvest following 24 h/30°C exposure to 22 mM propionate or 75 mM w-propanol in 25 mM citrate/phosphate buffer at pH 4.9.

Treatment hours pH +SE -SE % ±

Control 0 7.155 0.012 0.011 0 0

pH 4.9 24 7.024 0.032 0.030 5 2

Propionate 24 6.882 0.015 0.014 2 1

Propanol 24 6.960 0.025 0.024 4 0

APPENDIX E

DATA TABLES FOR CHAPTER 4

160

161

Loss of desiccation tolerance in nondormant red rice

Desiccation tolerance was tested for the 1990 harvest. Five replicates of 20

manually dehulled nondormant seeds were used for each time point. Seeds were sown

in 50 mL Erlenmeyer flasks with two sheets of Whatman No. 1 filter paper and 2 mL

of distilled H20 . Flasks were sealed with rubber septum caps. Seeds were incubated

at 30°C. Seeds were transferred at two-hour intervals to 9 cm Petri plates containing

three sheets of germination paper and dried for 7 d/30°C. After 7 days, seeds were

transferred to Erlenmeyer flasks as above and incubated for 7 d/30°C, at which time

germination was assessed. Loss of desiccation tolerance was taken as radicle death.

Viability was 100%.

Table E .l. Radicle and shoot emergence after hydration of nondormant seeds for 8 to 12 h/30°C followed by drying for 7 d/30° then rehydration for 7 d/30°C.

% Radicle +SE % Shoot +SE

Hours Emergence Emergence

8 99 0 99 1

10 94 5 100 0.3

12 11 5 97 1

162

Table E.2. Seed respiration and embryo [Fru 2,6-PJ data for hydrating dormant seeds in figure 4.1 and 4.2.

Respiration +SE Fru 2,6-P2 ±SE

Hours fiL 0 2 h-1 seed'1 pmoles embryo"1

0 -1.5 0.3 0.10 0.01

2 1.0 0.0 0.23 0.01

4 1.6 0.0 0.26 0.04

6 1.9 0.1 0.33 0.03

8 1.9 0.1 0.29 0.02

10 1.6 0.0 0.21 0.01

12 1.5 0.0 0.13 0.02

14 1.5 0.0 0.11 0.01

16 1.5 0.0 0.13 0.01

18 1.3 0.0

20 1.4 0.0 0.10 0.01

22 1.5 0.0

24 1.4 0.1 0.09 0.01

Table E.3. Seed respiration, embryo [Fru 2,6-PJ and germination data for hydrating nondormant seeds in figure 4.1 and 4.2.

Hours Respiration ±SE Fru 2,6-P2 +SE %Pericarp +SE %Radicle +SE %Shoot +SE

_______ f i t 0 2 h-1 seed-1________pmoles embryo1__________ Splitting_____________ Emergence__________ Emergence

0 -0.1 0.2 0.03 0.02 0 0 0 0 0 02 1.5 0.0 0.14 0.01 0 0 0 0 0 04 2.0 0.0 0.38 0.01 0 0 0 0 0 06 2.2 0.1 0.38 0.01 0 0 0 0 0 08 2.9 0.1 0.34 0.03 0 0 0 0 0 0

10 2.9 0.1 0.36 0.02 1 0 0 0 0 012 2.9 0.1 0.40 0.01 6 1 0 0 0 014 3.1 0.1 0.54 0.03 54 5 0 0 0 016 3.7 0.1 0.76 0.06 83 2 15 4 0 018 4.1 0.2 0.71 0.06 90 5 28 4 3 220 4.9 0.2 1.15 0.12 95 1 66 4 8 022 6.0 0.4 1.05 0.11 98 1 76 2 15 324 6.9 0.3 1.26 0.16 97 1 85 6 28 2

asu>

Table E.4. Endosperm [Fru 2,6-PJ data for hydrating dormant and nondormant seeds.

Dormant Nondormant

Fru 2,6-P2 ±SE Fru 2,6-P2 ±SE

Hours pmoles endosperm'1 pmoles endosperm

0 0.04 0.04 0.04 0.04

2 0.03 0.07 0.24 0.06

4 0.37 0.03 0.60 0.03

6 0.38 0.01 0.64 0.06

8 0.40 0.09 0.44 0.09

10 0.33 0.03 0.41 0.04

12 0.25 0.03 0.37 0.06

14 0.38 0.03 0.21 0.03

16 0.29 0.04 0.57 0.04

18 0.45 0.03

20 0.25 0.02 0.41 0.05

22 0.34 0.06

24 0.21 0.07 0.59 0.03

165

Table E.5. Embryo [Fru 2,6-PJ, real time germination, and contact time germination for the combined controls during and post contact as in figure 4.3.

Fru 2,6-P2 ±SE %Germ +SE %Germ ±SE

Hours pmoles embryo'1 Real Time Contact Time

0 0.08 0.01 0 0

4 0.10 0.02 0 0

8 0.10 0.01 0 0

12 0.10 0.01 1 0

16 0.10 0.01 1 0

20 0.10 0.01 1 0

24 0.11 0.01 2 0

28 0.10 0.01 0 0

32 0.10 0.01 1 0

36 0.11 0.03 1 0

40 0.11 0.01 1 0

44 0.10 0.02 1 0

48 0.10 0.01 0 0

166

Table E.6. Embryo [Fru 2,6-PJ, real time germination, and contact time germination resulting from a 24 h/30°C pulse of 32 mM methyl propionate in 25 mM citrate/ phosphate buffer at pH 7.0 as in figures 4.3 and 4.5.

Fru 2,6-P2 ±SE %Germ +SE %Germ ±SE

Hours pmoles embryo'1 Real Time Contact Time

0 0.08 0.01 0 0 6 0

2 3 2

4 0.08 0.01 0 0 3 1

8 0.08 0.01 0 0 10 0

12 0.09 0.02 0 0 38 2

16 0.10 0.03 1 1 54 1

20 0.09 0.03 0 0 73 3

24 0.13 0.00 2 1 96 2

28 0.22 0.04 2 1

32 0.27 0.04 1 0

36 0.38 0.04 3 1

40 0.42 0.04 12 1

44 0.54 0.05 25 3

48 0.65 0.05 40 3

167

Table E.7. Embryo [Fru 2,6-PJ, real time germination, and contact time germination during a 24 h/30°C pulse of 4 mM sodium nitrite/ 25 mM citrate/phosphate buffer at pH 3.0 as in figures 4.3 and 4.5.

Fru 2,6-P2 +SE %Germ +SE %Germ ±SE

Hours pmoles embryo'1 Real Time Contact Time

0 0.08 0.01 0 0 15 1

2 0.33 0.04 0 0 82 4

4 0.45 0.06 0 0 92 3

8 0.61 0.12 1 0 89 1

12 0.50 0.06 5 0 76 2

16 0.66 0.04 11 2 93 1

20 0.82 0.07 46 1 87 5

24 0.94 0.13 68 4 88 2

168

Table E.8. Embryo [Fru 2,6-PJ, real time germination, and contact time germination when dry dehulled dormant seeds are exposed 75 mM n-propanol in 25 mM citrate/ phosphate buffer pH 7.0.

Fru 2,6-P2 ±SE %Germ ±SE %Germ ±SE

Hours pmoles embryo'1 Real Time Contact Time

0 0 0

4 0.32 0.03 0 0

8 0.30 0.04 0 0

12 0.19 0.04 0 0

24 0.16 0.02 0 0

169

Table E.9. Embryo [Fru 2,6-PJ, real time germination, and contact time germination resulting from a 24 h/30°C pulse of 75 mM //-propanol in 25 mM citrate/phosphate buffer at pH 7.0 as in figure 4.3 and 4.5.

Fru 2,6-P2 ±SE %Germ +SE %Germ +SE

Hours pmoles embryo'1 Real Time Contact Time

0 0.08 0.01 0 0 6 0

2 3 1

4 0.15 0.01 0 0 4 1

8 0.17 0.02 0 0 5 2

12 0.16 0.02 0 0 20 2

16 0.15 0.03 1 0 41 3

20 0.18 0.01 1 0 74 2

24 0.20 0.02 1 1 91 1

28 0.22 0.03 4 1

32 0.23 0.05 4 1

36 0.27 0.01 8 0

40 0.48 0.08 22 3

44 0.54 0.02 35 5

48 0.61 0.10 48 4

170

Table E.10. Embryo [Fru 2,6-PJ, real time germination, and contact time germination as a result of a 24 h/30°C pulse of 40 mM propionaldehyde in 25 mM citrate/phosphate buffer at pH 7.0 as in figure 4.3 and 4.5.

Fru 2,6-P2 +SE %Germ +SE %Germ +SE

Hours pmoles embryo'1 Real Time Contact Time

0 0.10 0.01 0 0 4 1

2 0.33 0.01 0 0 8 3

4 0.36 0.03 0 0 12 2

8 0.33 0.04 0 0 35 5

12 0.30 0.06 0 0 68 4

16 0.34 0.01 1 1 82 4

20 0.39 0.07 7 1 89 2

24 0.61 0.05 14 3 94 1

28 1.00 0.10 56 3

32 1.05 0.03 78 5

36 1.32 0.23 91 3

171

Table E .l l . Embryo [Fru 2,6-PJ, real time germination, and contact time germination resulting from a 24 h/30°C pulse of 22 mM propionate in 25 mM citrate/phosphate buffer at pH 4.9 as in figure 4.3 and 4.5.

Fru 2,6-P2 ±SE %Germ ±SE %Germ ±SE

Hours pmoles embryo"1 Real Time Contact Time

0 0.10 0.01 0 0 9 3

2 4 1

4 0.06 0.01 0 0 3 1

8 0.08 0.00 1 1 8 2

12 0.09 0.02 1 0 31 10

16 0.09 0.01 1 0 65 15

20 0.10 0.01 0 0 86 10

24 0.12 0.03 4 1 94 1

28 0.18 0.03 3 1

32 0.33 0.01 8 1

36 0.40 0.03 7 1

40 0.47 0.04 17 3

44 0.49 0.06 29 3

48 0.67 0.08 35 5

172

Table E.12. Embryo [Fru 2,6-PJ on plateau during chemical contact vs. time to 30 and 40% germination as in figure 4.4.

Time on Fru 2,6-P2 Time to Time to

Compound Plateau pmoles embryo"1 30% Germ 40%

Sodium nitrite 4 - 12 h 0.52 ± 0.05 18 h 19 h

Propionaldehyde 2 - 20 h 0.34 ± 0.01 25 h 26 h

^-Propanol 4 - 24 h 0.17 ± 0.01 42 h 46 h

Propionic acid 4 - 24 h 0.09 ± 0.01 44 h 48 h

Methylpropionate 4 - 24 h 0.10 ± 0.01 46 h 48 h

Controls 4 - 24 h 0.08 ± 0.003 not applicable

APPENDIX F

DATA TABLES FOR CHAPTER 5

173

Table F .l. Relative signal intensities (RSI) and carbon number of metabolite signals in perchloric acid extracts and germination in real time of propionate-2-13C labelled embryos (100.6 MHz in 85% H20/15% D20) in figure 5.2 and 5.3.

Hours

propionate

C2RSI +SE

3-hydroxypropionateC2RSI ±SE

citrate

C 2& C 4RSI ±SE

32.1 ppma

RSI +SE %GERM +SE

0 0 0 0 0 0 0 0 0 0 02 7.4 2.4 3.1 0.9 0 0 0 0 0 04 13.0 0.9 8.7 0.9 0 0 0 0 0 08 28.9 1.3 27.2 1.1 0 0 0 0 1 1

12 32.3 2.6 40.5 3.5 0 0 0 0 0 016 37.4 3.1 51.1 1.5 0 0 0 0 0 020 29.7 2.1 65.6 3.9 0 0 0 0 4 224 25.2 2.8 87.3 14.8 0 0 0 0 6 128 13.6 1.5 89.0 21.2 5.9 0.6 0 0 12 132 8.1 0.5 91.1 7.2 6.7 6.7 5.6 5.6 16 336 3.2 0.4 49.6 5.5 7.8 1.4 5.5 1.3 20 1

a unidentified signalFor contact time germination see appendix Table E .l l .Signal intensities set relative to a 3-hydroxypropionate-2-13C signal at 24 h set at 100 arbitrary units.

175

Table F.2. Total relative signal intensities (TRSI) of the combined relative signal intensities of the signals from C2 of both 3-hydroxypropionate-2-13C and propionate-2- 13C in figure 5.3B. Data from Table F .l.

Hours TRSI ±SE

0 0 0

2 10.5 3.2

4 22.0 1.6

8 56.1 2.4

12 72.8 3.4

16 88.5 4.7

20 95.3 2.1

24 112.5 12.9

28 102.6 22.6

32 99.3 7.4

36 52.8 5.2

176

Table F.3. Relative signal intensities (RSI) and carbon number of metabolite signals in perchloric acid extracts and germination in real time of «-propanol-l-13C labelled embryos (100.6 MHz in 85% H20/15% D20) in figure 5.5 and 5.6.

n-Propanol 3-Hydroxy

propionate

Cl

Hours RSI ±SE

Cl

RSI ±SE

Real Time

% Germ +SE

0 0 0 0 0 0 0

2 38.6 7.7 1.7 1.7 0 0

4 42.6 8.4 4.9 4.9 0 0

8 42.1 5.7 6.2 0.5 0 0

12 43.1 4.8 6.5 0.9 1 1

16 47.2 10.4 8.0 0.5 2 1

20 42.0 8.2 7.1 0.3 5 1

24 36.9 4.7 9.8 0.9 7 0

For contact time germination see appendix Table E.9.

Signal intensities set relative to a 3-hydroxypropionate-2-13C signal at 24 h set at 100

arbitrary units.

177

Table F.4. Relative signal intensities (RSI) and carbon number of metabolite signals in perchloric acid extracts propionate-2-13C labelled endosperms (100.6 MHz in 85% H20/15% D20).

Propionate 3-Hydroxy

propionate

C2 C2

Hours RSI ±SE RSI ±SE

0 0 0 0 0

2 3.3 1.0 1.7 0.8

4 5.4 0.6 4.3 0.6

8 7.8 1.2 9.0 1.5

12 8.8 0.5 10.2 1.2

16 10.8 1.3 14.5 0.5

20 8.9 1.5 12.9 2.0

24 10.1 1.2 18.6 1.1

APPENDIX G

DATA TABLE FOR CHAPTER 6

178

179

Table G .l. 31P NMR assignments (ppm) of the presumed P; signal in perfused red rice embryos in a single pulse spectra (Fig. 6.4) and in spin echo spectra during exposure to nitrite (Fig. 6.5), alkaline pH (Fig. 6.6).

Treatment 5(ppm)

Single pulse

Control 2.071

Spin echo

Control 1.991

Nitrite pH 2.2

5 min 1.978 1.271

30 min 1.857 1.254

60 min 1.820 1.253

pH 8.3

5 min 1.709 1.182

15 min 1.983 1.305

30 min 2.000 1.197

VITA

Steven Footitt was bom on November 19th 1958, Stockport, England, and was

raised in the village of Furness Vale in the Peak District of Derbyshire. He worked

in manufacturing industry before attending North East London Polytechnic in 1982,

attaining a B.Sc. honours degree in Applied Biology in 1985. While attending college,

a one year internship was spent at the Royal Botanic Gardens, Kew, working on seed

dormancy in wild grasses under the direction of Dr. R Probert. In 1986, he came to

Louisiana State University to join the research program of Dr. MA Cohn in the

Department of Plant Pathology and Crop Physiology. His graduate research was on

the mechanism of dormancy-breaking in red rice. This research focused on the

changes in embryo pH and metabolism in dormant seeds when exposed to dormancy-

breaking chemicals. He is now a candidate for the degree of Doctor of Philosophy in

Plant Health.

180

DOCTORAL EXAMINATION AND DISSERTATION REPORT

Candidate: Steven Footitt

Major Field: Plant Health

Title of Dissertation: S eed Dormancy in Red Rice (O ryza sa t i v a ) : Changesin Embryo pH and Metabolism During the Dormancy- Breaking Process

Approved:

' 4 m-/ /

Major Professor and Chairman

0 A<11 P' A f ' y vDean of the Gradate School

EXAMINING COMMITTEE:■! -< n ,

- / / / ) .■ /

/ C rri\ { Cl. J - ( . i l r V Y i L

C cO

fh.

Date of Examination:

December 3, 1992