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Louisiana State UniversityLSU Digital Commons
LSU Historical Dissertations and Theses Graduate School
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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Recommended CitationFootitt, Steven, "Seed Dormancy in Red Rice (Oryza Sativa): Changes in Embryo pH and Metabolism During the Dormancy-Breaking Process." (1993). LSU Historical Dissertations and Theses. 5501.https://digitalcommons.lsu.edu/gradschool_disstheses/5501
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3 0 0 North Z e e b R oad . Ann Arbor. Ml 4 8 1 0 6 -1 3 4 6 USA 3 1 3 /7 6 1 -4 7 0 0 8 0 0 /5 2 1 -0 6 0 0
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**
ww F|*V«
-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
b £ 100ro t
£ 8 0
p 6 0
a> o
® 3 —Hydroxypropionate—2 — 13O P r o p io n a t e —2 — C
A C i t r a t e - 2 , 4 - 1 3 C
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
O O Pr o p a n o l— 1 —5 0 - T
■+->
‘coca;
■+->c
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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Glass capillary
4*A3Has
5E
r . ,
■ 1
A
f t
Circulatory flow
Lift tube
Embryos
Perforated rubber support
Reference capillary (TEMDP)
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