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LSU Historical Dissertations and Theses Graduate School

1992

Selection and Characterization of Populations of Rapid-Cycling Selection and Characterization of Populations of Rapid-Cycling

Brassica Rapa L. Differing in Their Response to Rootzone Brassica Rapa L. Differing in Their Response to Rootzone

Hypoxia. Hypoxia.

Christine Jo Daugherty Louisiana State University and Agricultural & Mechanical College

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Recommended Citation Recommended Citation Daugherty, Christine Jo, "Selection and Characterization of Populations of Rapid-Cycling Brassica Rapa L. Differing in Their Response to Rootzone Hypoxia." (1992). LSU Historical Dissertations and Theses. 5377. https://digitalcommons.lsu.edu/gradschool_disstheses/5377

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S e le c t io n an d c h a r a c te r iza tio n o f p o p u la tio n s o f r a p id -c y c lin g Brassica rapa, L . d ifferin g in th e ir r e sp o n se to r o o tz o n e h y p o x ia

Daugherty, Christine Jo, Ph.D.

T h e Louis iana S t a te Univers i ty and Ag r ic u l t ura l and Mechanical Col., 1992

UMI100 N Z tvb Rd.Ann Arbor. Ml 4X 106

SELECTION AND CHARACTERIZATION OF POPULATIONS OF RAPID-CYCLING BRASSICA RAPA L. DIFFERING IN

THEIR RESPONSE TO ROOTZONE HYPOXIA

A Dissertation

Submitted to the Graduate Faculty of the Louisiana State University and

Agricultural and Mechanical College in partial fulfillment of the requirements for the degree of

Doctor of Philosophyin

The Department of Plant Pathology and Crop Physiology

byChristine Jo Daugherty

B . A. , Central University of Iowa, 1986 M.S., Iowa State University, 1988

August 1992

ACKNOWLEDGEMENTS I would like to sincerely thank Dr. Mary E. Musgrave

for her advice, friendship, and enduring patience throughout my doctoral program. Special thanks to A1 Hopkins and Denyse Cummins for their assistance and camraderie in the lab. I am grateful to Drs. Sharon Matthews, Irv Mendelssohn, and Tao Kong for their assistance in the TEM and energy charge studies. 1 also acknowledge Dr. Ray Schneider, Dr. Marc Cohn and Steven Footitt for providing equipment and technical assistance with the enzyme and isozyme studies. Thanks to Dr. Paul williams and the Crucifer Genetics Cooperative for providing the Brassica rapa seed. I thank Drs. Dave Longstreth and William Blackmon for their contributions to this project. I also acknowledge Sigma Xi Grant-in-Aid Research funding for providing support for this research. Special thanks to my family and friends for supporting me throughout my graduate studies.

ii

TABLE OF CONTENTS gagsACKNOWLEDGMENTS .......................................... iiLIST OF TABLES.......................................... ivLIST OF FIGURES ........................................ VLIST OF ABBREVIATIONS................................. viiABSTRACT.............................................. viiiCHAPTER

1 REVIEW OF LITERATURE .......................... 1Literature Cited .......................... 15

2 SELECTION OF POPULATIONS OF RAPID-CYCLING BRASSICA RAPA L. DIFFERING IN THEIR RESPONSETO ROOTZONE HYPOXIA ......................... 22

Introduction .............................. 23Materials and Methods ..................... 24Results.................................... 29Discussion................................. 39Summary.................................... 4 2Literature Cited .......................... 44

3 CARBOHYDRATE STATUS OF POPULATIONS OF RAPID-CYCLING BRASSICA RAPA L. UNDERROOTZONE HYPOXIA ............................ 46

Introduction .............................. 47Materials and Methods ..................... 48Results.................................... 52Discussion................................. 60Summary.................................... 64Literature Cited .......................... 66

4 EARLY CHANGES IN ROOT METABOLISM OF THREE POPULATIONS OF RAPID-CYCLING BRASSICA RAPA L. DURING WATERLOGGING .......................... 68

Introduction .............................. 69Materials and Methods ..................... 71Results.................................... 75Discussion................................. 86Summary.................................... 94Literature Cited .......................... 96

5 CONCLUSIONS AND PERSPECTIVES ................. 100REFERENCES.............................................. IllV I T A .................................................... 121

iii

LIST OF TABLES

Table

Table

Table

Table

Table

Table

Table

Table

2.1

2 .2 :

3.1

3.2

4.1:

4.2:

4.3:

4.4:

Total chlorophyll content of populations of Brassica rapa responding to waterlogging ............................ 3 3Comparison of Brassica rapa root and shoot weights responding to waterlogging treatment for six and eight d a y s ........ 36Comparison of soluble carbohydrate concentrations in leaves of populations of Brassica rapa responding to waterlogging stress ....................... 54Comparison of leaf starch concentrations in populations of Brassica rapa responding to waterlogging stress .................. 57Activities of six enzymes measured in Brassica rapa roots after 48 hours of waterlogging .......................... 75Alcohol dehydrogenase activities measured in roots of populations of Brassica rapa after 12 and 18 hours of waterlogging ... 78Pyruvate decarboxylase activities measured in roots of populations of Brassica rapa after 48 hours of waterlogging .......... 79Adenylate energy charge ratio and the concentration of ATP measured in roots of populations of Brassica rapa after 18 hours of waterlogging .................... 80

iv

LIST OF FIGURES

Figure 2.1: Experiment design used to obtain distinctpopulations of Brassica rapa from the base B. rapa population which differed in theirresponse to waterlogging stress ........ 27

Figure 2.2: Concentration of total chlorophyll fromthe base population of Brassica rapa under control and waterlogged conditions ..... 31

Figure 2.3: Concentration of total chlorophyll fromthe tolerant and sensitive populations of Brassica rapa under waterlogged conditions 3 2

Figure 2.4: Concentrations of total chlorophyll fromthe tolerant and sensitive populationsof Brassica rapa under controlconditions .............................. 34

Figure 2.5: Root dry weight of populations of Brassicarapa subjected to control or waterlogged conditions for eight days .............. 37

Figure 2.6: Shoot dry weight of populations ofBrassica rapa subjected to controlor waterlogged conditions foreight days .............................. 38

Figure 3.1: Leaf soluble carbohydrates of populationsof Brassica rapa responding towaterlogging stress and concurrentsoil redox values ....................... 53

Figure 3.2: Leaf starch of populations of Brassicarapa responding to waterloggingstress ................................... 56

Figure 3.3: Light micrographs of leaf sectionsfrom leaves of control and waterlogged plants.....................................58

Figure 3.4: Electron micrographs of parenchyma cellsections from leaves of control and waterlogged plants ...................... 59

Figure 4.1: Root alcohol dehydrogenase activities ofpopulations of Brassica rapa responding waterlogging stress ..................... 77

v

LIST OF FIGURES (continued)Figure 4.2:

Figure 4.3:

Composite electrophoretic banding patterns of the four enzymes extracted from Brassica rapa roots that did not show variation in the banding patterns among the three populations regardless of treatment .... 81Electrophoretic banding pattern of alcohol dehydrogenase isozymes extracted from Brassica rapa roots exposed to waterlogged conditions for 2 4 or 48 hours ................................ 82

Figure 4.4 Electrophoretic banding pattern of phosphoglucomutase isozymes extracted from Brassica rapa roots ............ . 84

Figure 4.5 Electron micrographs of Brassica rapa cell sections from waterlogged and control roots showing mitochondrial ultrastructure ....................... 85

Figure 5.1: Physiological response timeline of three populations of Brassica rapa in response to rootzone hypoxia .................. 105

LIST OF ABBREVIATIONS

(ATP) adenosine triphosphate(AEC) adenylate energy charge(ADH) alcohol dehydrogenase(CrGC) Crucifer Genetics Cooperative(G6PDH) glucose 6-phosphate dehydrogenase(MDH) malate dehydrogenase(ME) malic enzyme(PGM) phosphoglucomutase(PDC) pyruvate decarboxylase

vii

ABSTRACTChallenging a heterogenous population of plants (rapid-

cycling Brassica rapa L.) with waterlogging, we selected plants which differed in their response to rootzone hypoxia. Based on visual criteria, individuals from the waterlogged wild-type population were separated into "tolerant" and "sensitive" groups and mass pollinated to produce the next generation. This selection protocol was continued for seven generations. Under waterlogged conditions, chlorophyll concentrations in the tolerant population were significantly greater than in the sensitive population. After eight days of waterlogging, tolerant plants had significantly greater dry root and shoot weights than the sensitive plants. However, when tolerant and sensitive populations were grown under control conditions chlorophyll concentrations and dry weights were not significantly different. Solublecarbohydrates and starch concentrations steadily increased in the leaves of waterlogged plants, but carbohydrate concentrations of control plants maintained a constant level. Examination by transmission electron microscopy showed distorted thylakoid membranes in the chloroplasts of waterlogged sensitive leaves.

Activities and electrophoretic profiles of six root enzymes: alcohol dehydrogenase (ADH), glucose 6-phosphate dehydrogenase (G6PDH), malate dehydrogenase (MDH), malic enzyme (ME), phosphoglucomutase (PGM), and pyruvate

vi ii

decarboxylase (PDC) were examined in the populations subjected to rootzone hypoxia. ADH and PDC activities increased, PGM and MDH activities decreased, and ME and G6PDH activities did not change. There were significant differences in ADH activities in the populations after 18 hours of waterlogging. The isozyme patterns of ADH and PGM were also different among the populations. Adenylate energy charge ratios measured in the populations were similar after 18 hours of waterlogging. There were swollen cristae and electron dense matrices in the mitochondria of roots from all plants grown under waterlogged conditions. Enzyme activities, energy charge, carbohydrate status, and ultrastructure differences were considered to be indicators of rather than causes of differences in population responses to rootzone hypoxia.

ix

CHAPTER ONE

REVIEW OF LITERATURE

I

2We are interested in the physiological mechanisms of

waterlogging tolerance and how tolerance affects short-term phenotypic responses to hypoxic events in the rootzone. The occurrence of morphological and physiological characteristics in species better adapted to waterlogged conditions than others suggests that these characteristics confer a degree of tolerance to the stresses associated with excessive soil moisture. The differential responses by groups of individuals to hypoxic conditions could be used to identify a genetic pool for enhanced tolerance to the waterlogging stress. The following chapters explain the experimental selection of groups of plants that differ in their response to rootzone hypoxia, and the physiological and biochemical mechanisms involved in the differential response.

Many facets of plant responses to waterlogging have been observed, including morphological and ultrastructural changes, differences in metabolic pathways, and alteration of enzyme activity. These responses will be briefly reviewed and discussed in relation to the project.

Gross Morphological Response to HypoxiaSoil waterlogging greatly inhibits gaseous diffusion

from the atmosphere, leading to hypoxia and subsequent anoxia around the rootzone. Gross morphological changes in response to waterlogging include induction of adventitious

3

roots (Kramer, 1951; Wenkert et al. , 1981; Kozlowski andPallardy, 1984) and an increase in aerenchymatous tissue (Drew et al. , 1979; 1981; Kawase and Whitmoyer, 1980;Trought and Drew, 1980). Both mechanisms function to increase oxygen availability to the waterlogged roots. These responses are thought to be mediated via plant growth regulators, especially ethylene (Jackson et al. , 1981;Kawase, 1981) . It has been suggested that spatial concentration of hormones in the shoot near the soil surface may contribute to the initiation of adventitious roots (Jackson, 1984; Reid and Bradford, 1984).

Another morphological change is the yellowing of foliage under prolonged waterlogging stress (Drew and Sisworo, 1977; 1979; Jackson, 1983). Yellowing may be due to changes in nutrient availability (Mendelssohn and Postek, 1982; Drew and Sisworo, 1979), status of nitrogen in the rootzone environment (Trought and Drew, 1980; 1981; Drew and Sisworo, 1979), and the alteration of carbohydrate assimilation and/or partitioning (Laan and Blom, 1990; Pezeshki, 1990; Wample and Davis, 1983).

Foliar Carbohydrate Response to HypoxiaPlants exposed to hypoxic conditions induced by

waterlogging usually have the normal balance between consumption and production of carbohydrates disrupted. Partitioning of carbohydrates may be influenced by

4differences in metabolic pathways, alteration of enzyme activity, and morphological and ultrastructural changes that occur when plants are subjected to waterlogging stress (Crawford, 1978; Drew, 1987). Under hypoxic conditions a continuous supply of soluble carbohydrates to root tissues is needed to preserve and maintain growth of root tips which are a critical component of plant survival (Barta, 1988; Jackson and Drew, 1984).

A few reports have suggested that a decreased carbohydrate supply may occur following waterlogging. The reduction in carbohydrate levels may be caused by stomatal closure induced by reduced water permeability of the roots (Kozlowski and Pallardy, 1984; Save and Serrano, 1986; Trought and Drew, 1980). However, hypoxia also leads to reduced translocation rates, thereby decreasing the movement of assimilates from the leaves (Schumacher and Smucker, 1985) . The basis of carbohydrate accumulation in the leaves has been hypothesized to be due to reduced utilization of carbohydrates by the plant as a whole (Setter et al., 1987; Siji and Swanson, 1973). Subsequently, this may lead to disruption of the photosynthetic apparatus by excessive starch accumulation in the thylakoid membranes (Cave et al. , 1981; Talbot et al. , 1987; Wample and Davis, 1983; Wulff and Strain, 1982).

Root Cell Response to HypoxiaRoots, a major carbohydrate sink, respond at the

cellular level to the hypoxic or anoxic environments imposed by waterlogging conditions. Changes in membrane properties, cellular energy balance, and overall metabolism may limit the demand for assimilates from the leaves (Jackson and Drew, 1984). Longer durations of rootzone hypoxia increase cell membrane permeability as well as distort the ultrastructure of mitochondrial membranes (Drakeford and Reid, 1987; Vartapetian et al. , 1987; Oliveira, 1977;Vartapetian, 1982; 1991). Disruption of the cell membrane and mitochondrial integrity would definitely affect the source/sink balance of foliar carbohydrate supply and root demand. Changes in cellular membrane status of the roots may result in a cascade of events which ultimately affects all metabolic processes in the plant.

Root Metabolism in Response to HypoxiaMetabolic changes in roots include alteration of

respiratory pathways (Opik, 1973; Crawford, 1978; Armstrong, 1978; Smith and ap Rees, 1979; Mendelssohn and Burdick, 1988) and glycolytic pathways (John and Greenway, 1976; Crawford, 1978; Davies, 1980; McKee et al., 1989). Root respiratory processes are immediately affected because of limited oxygen. When roots are under a short duration of waterlogging stress, alcoholic fermentation may be

6accelerated. The response of the accelerated glycolytic cycle (Pasteur effect) is to produce as much ATP as possible during short-term waterlogging. By coupling the glycolysis cycle with the pentose phosphate pathway, another metabolic route for carbohydrate degradation, generation of NADPH is accomplished.

The energy produced by these metabolic pathways can be measured by the adenylate energy charge ratio (AEC). AEC is the ratio of phosphorylated adenine nucleotides to the total adenine nucleotide pool. The AEC ratio will change in response to the concentration of the phosphorylated adenine nucleotides in the cell. The overall energy or metabolic status of a plant can be monitored by determining the AEC ratios. When plants are exposed to a hypoxic stress in the rootzone induced by natural waterlogging or laboratory methods, the AEC ratio can change dramatically. Several reports have shown when plants are exposed to waterlogging stress, the root AEC ratio decreases in comparison to the control level (Mendelssohn et al., 1981; McKee andMendelssohn, 1987; Barta, 1986; Burdick and Mendelssohn, 1987), although in some wetland plants the AEC ratios remain high even under hypoxic stress (Mendelssohn and McKee, 1987). In several waterlogging tolerant species, the AEC ratio initially decreases under hypoxia, but then recovers, possibly due to the increased activity of the glycolysis cycle (Mendelssohn and Burdick, 1988).

7Although lack of ATP is a significant problem during

waterlogging stress, poisoning of the roots by products of fermentation might also be a potential hazard. Ethanol accumulation which results from increased alcoholic fermentation can lead to membrane destruction and inactivation of mitochondrial activity (Crawford, 1978) . As the cell membrane is destroyed, exudation of cellular contents soon follows. This may facilitate the growth of soil pathogens which invade roots, subsequently leading to plant death. However, roost waterlogging tolerant plants exhibit ethanol diffusion from the roots, resulting in relatively low root concentrations (Drew, 1987; Jackson and Drew, 1984).

Crawford (1978) and his colleagues (McManmon and Crawford 1971; Barclay and Crawford, 1982; Crawford and Zochowski, 1984; Crawford et al. , 1987) reported thatwaterlogging tolerant plants differ from intolerant plants by not accelerating alcoholic fermentation, and subsequently limiting the amount of ethanol produced. These waterlogging tolerant plants respond via an alternative metabolic pathway which increases malate concentrations via the carboxylation of glycolytic phosphoenolpyruvate followed by the reduction of oxaloacetate to malate via malate dehydrogenase (ap Rees et al., 1987). These waterlogging tolerant plants produce malate in the cell, and therefore do not increase ethanol in the rootzone.

8Enzyme Activities in Response to Hypoxia

Roots of plants exposed to hypoxic conditions synthesize a set of approximately 20 soluble proteins, which have been designated as the "anaerobic proteins" (Sachs et al. , 1980). Several of these anaerobic proteins have been identified as enzymes involved in the glycolysis cycle (Ferl et al. , 1979; Kelley and Freeling, 1984a; 1984b;). Anincrease or decrease in enzyme activity in response to waterlogging stress may affect the activities of all other enzymes in a particular pathway. Alteration in activities of several enzymes have been reported to occur under waterlogging stress: cytochrome oxidase (Opik, 1973); lactic dehydrogenase (Sherwin and Simon, 1969; VanToai et al., 1987); malic enzyme (Davies et al., 1974; McManmon andCrawford, 1971; VanToai et al. , 1987); malate dehydrogenase (Crawford, 1978; Avadhani et al. , 1978); alcoholdehydrogenase (McManmon and Crawford, 1971; Wignarajah and Greenway, 1976; Mendelssohn et al. , 1981; VanToai et al. ,1985; Roberts et al., 1989); pyruvate decarboxylase, (John and Greenway, 1976; Wignarajah and Greenway, 1976; VanToai et al. , 1985; 1987) and phosphofructokinase/pyruvate kinase (Vanlerberghe et al., 1990). Changes in enzyme activities in response to waterlogging stress may occur in other metabolic pathways throughout the plant and are not limited to the enzymes involved in the alcoholic fermentation.

9Because changes in enzyme activities are so dramatic

following waterlogging, they have been used as indicators of plant response to waterlogging stress. Often these enzymatic responses to hypoxia are interpreted as potential adaptations to the stress (Hook et al. , 1971; Crawford,1978; Pradet and Bomsel, 1978; Kozlowski and Pallardy, 1984; Burdick and Mendelssohn, 1990). While the validity of this interpretation has been debated (John and Greenway, 1976; Avadhani et al. , 1978; Keeley and Franz, 1979; Drew andLynch, 1980; Smith et al., 1986), these enzyme changes arenevertheless valuable indicators of plant performance when challenged by rootzone hypoxia.

Selection of Populations Tolerant to an Environmental StressMorphological and physiological characteristics

apparent in species better adapted to waterlogged conditions than others suggest that these traits may confer a degree of tolerance to stresses associated with waterlogging. The presence or absence of the various adaptations to waterlogging discussed above have been taken by many authors as evidence of differential tolerance to waterlogging stress among plant species (Crawford, 1978; 1982; Hook andScholtens, 1978; Jackson and Drew, 1984; Kozlowski and Pallardy, 1984; Mendelssohn et al. , 1981). However, if a

10trait is truly of adaptive significance, one would expect it to appear in response to the challenge of environmental stress.

Since differential responses of individuals to an environmental stress ultimately affect fitness of all individuals and exert selective pressure, individual differences can lead to development of tolerant populations. This has been demonstrated in natural populations exposed to heavy metal contamination (Antonovics et al., 1971) andrecently in radish populations challenged with air pollution stress (Gillespie and Winner, 1989).

Thus, another approach to studying waterlogging tolerance would be to challenge an unselected plant population with waterlogging stress, screen multiple generations and observe the traits which distinguish lines of plants which grow well under waterlogged conditions from those which do poorly. Gross morphological changes (adventitious root formation, aerenchyma, yellowing of foliage) could be used to identify desirable traits in a selection program to make plants more tolerant of a hypoxic rootzone. In comparison, other response features such as cellular ultrastructure and metabolic activity are less easily observed and require destructive plant sampling, and therefore would not readily lend themselves to a selection program.

11Benefits of rapid-cycling Brassica rapa

Rapid-cycling Brassica species, relatives of mustard, have the potential for resolving many questions in plant biology. The rapid-cycling Brassica populations were developed through mass pollination and recurrent selection by Dr. Paul Williams and his colleagues at the University of Wisconsin-Madison (Williams and Hill, 1986). Brassica rapa L. populations can produce up to ten generations per year making this plant an excellent tool for research in genetics, population biology, and plant breeding. Their small size (approximately 15 cm in height) , rapid development, and low light requirement make the plants readily usable in a laboratory setting. Rapid-cycling Brassica species respond strongly to their environment, indicating the plants possess reservoirs of genes useful for recombination of favorable genes or genetic resistance to a particular stress (Williams and Hill, 1986). Various mutants have been developed from the base (wild-type) population that respond to physical and chemical stimuli, making these populations excellent for studying physiological responses in plants. The ease of culture, rapid development, and the strong response to environmental factors contributed to the rationale for choosing the rapid- cycling Brassica rapa for these experiments.

12Project Objectives

The purpose of the experiments outlined below is to determine whether a heterogeneous population of plants would display differential tolerance to waterlogging stress and if so, whether differential responses by groups of individuals could serve as a genetic pool to enhance tolerance to environmental stress. if differential tolerance was observed, we would analyze qualitatively differences between selected populations with regard to the putative waterlogging tolerance traits which have been discussed in the literature review. Three issues are addressed:

1. The heterogenous base (wild-type) population of rapid- cycling Brassica rapa L. was chosen as the subject for the experiments. Through recurrent selection techniques and mass pollination, can we obtain populations differing in waterlogging tolerance?

2. Since the populations were separated on the basis of their response to the waterlogging challenge, differences in physiology and morphology observed should be linked to the differential tolerance. Does leaf carbohydrate status relate to the differential tolerance response to waterlogging?

3. Similarly, is root metabolism determining the differential tolerance response to waterlogging?

1 3

By challenging an unselected population of Brassica rapa L. with waterlogging stress and selecting plants using a visual criterion, it was possible to distinguish a population of plants which grew well when waterlogged, from one that did poorly. Characterization of how these populations differ in their response to waterlogging then became the goal of the dissertation study. In Chapter Two, chlorophyll concentrations were used to describe population differences in relation to their waterlogging response. Reductions in growth rate caused by waterlogging stress were compared between the populations using sequential harvesting and growth analysis. Through these procedures it was possible to identify temporal differences in root and shoot responses.

The response of leaf carbohydrates to waterlogging stress in the B . rapa populations is examined in Chapter Three. Soluble carbohydrates, starch levels, andchloroplast ultrastructure of the populations were used to monitor carbohydrate accumulation in the leaves. Onset of the environmental stress was recorded by measuring soil redox values every six hours. Thus it was possible to relate early changes in foliar carbohydrate status to the timecourse of oxygen disappearance in the rootzone.

Root response of the B. rapa populations when exposed to rootzone hypoxia is examined in Chapter Four. Activities and isozyme patterns of six enzymes, and root adenylate

14energy charge were analyzed to observe metabolic changes during the initial hours of waterlogging stress. Mitochondrial ultrastructure was examined to observe subcellular changes that may have occurred during the later period of the waterlogging stress. The timing or onset of alcoholic fermentation was monitored at six-hour intervals to detect differences in populations when exposed to short­term rootzone hypoxia.

The data discussed in Chapters Two through Four are re­examined in Chapter Five to synthesize hypotheses regarding the nature of the tolerant and sensitive responses to waterlogging in the Brassica rapa populations developed for this study. Changes in soluble carbohydrates, starch, and overall dry matter are used as markers for the responses by the shoot, while ADH, PDC, and root dry matter are indicators of responses by the root. This approach allowed construction of a response timeline for shoot and root responses by the populations. Comparison of the timelines elucidates interrelationships between root and shoot metabolism in response to waterlogging stress and also permits development of hypotheses regarding the nature of the tolerance and sensitivity to soil waterlogging.

15Literature Cited

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ap Rees T, Jenkin LET, Smith AH, Wilson PM (1987) The metabolism of flood-tolerant plants. In RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford, pp 227-238

Armstrong W (1978) Root aeration in the wetland condition. Jn DD Hook, RMM Crawford, eds, Plant Life in Anaerobic Environments. Ann Arbor Science, MI, pp 269-297

Avadhani PN, Greenway H, Lefroy R, Prior L (1978) Alcoholic fermentation and malate metabolism in rice germinating at low oxygen concentrations. Aust J Plant Physiol 5: 15-25

Barclay AM, Crawford RMM (1982) Plant growth and survival under strict anaerobiosis. J Exp Bot 33: 541-549

Barta AL (1986) Metabolic response of Medicago sativa L. and Lotus corniculatus L. roots to anoxia. Plant Cell Env 9: 127-131

Barta AL (1988) Response of field grown alfalfa to root waterlogging and shoot removal. I. Plant injury and carbohydrate and mineral content of roots. Agron J 80: 889- 892

Burdick DM, Mendelssohn IA (1987) Waterlogging responses in dune, swale and marsh populations of Spartina patens under field conditions. Oecol 74: 321-329

Burdick DM, Mendelssohn IA (1990) Relationship between anatomical and metabolic responses to soil waterlogging in the coastal grass Spartina patens. 3 Exp Bot 41: 223-228

Cave G, Tolley LC, Strain BR (1981) Effect of carbon dioxide enrichment on chlorophyll content, starch content and starch grain structure in Trifolium subterraneum leaves. Physiol Plant 51: 171-174

Crawford RMM (1978) Metabolic adaptation to anoxia. Tn DD Hook, RMM Crawford, eds. Plant Life in Anaerobic Environments. Ann Arbor Science, MI, pp 119-155

Crawford RMM (1982) Physiological responses to flooding. In OL Lange, PS Nobel, CB Osmond, H Ziegler, eds, Encyclopedia of Plant Physiology vol 12 B, Physiological Plant EcologyII. Springer-Verlag, New York, pp 437-477

1 6

Crawford RMM, Monk LS, Zochowski ZM (1987) Enhancement of anoxia tolerance by removal of volatile products of anaerobiosis. In RMM Crawford, ed. Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford, pp 375-384Crawford RMM, Zochowski ZM (1984) Tolerance of anoxia and ethanol toxicity in chickpea seedlings (Cicer arietinum L.). J Exp Bot 35: 1472-1480

Davies DD (1980) Anaerobic metabolism and the production of organic acids. In DD Davies, ed, The Biochemistry of Plants. Academic Press, New York, pp 581-611

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Drakeford DR, Reid DM (1987) Some rapid responses of sunflower to flooding. In RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford, pp 385-395

Drew MC (1987) Mechanisms of acclimation to flooding and oxygen shortage in non-wetland species. Jn RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford, pp 321-331

Drew MC, Jackson MB, Giffard SC (1979) Ethylene-promoted adventitious rooting and development of cortical air spaces (aerenchyma) in roots may be adaptive responses to flooding in Zea mays L. Planta 147: 83-86

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17Ferl RJ, Dlouhy SR, Schwartz D (1979) Analysis of maize alcohol dehydrogenase by native SDS two-dimensional electrophoresis and autoradiography. Mol Gen Genet 169: 7- 12

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Kawase M, Whitmoyer RE (1980) Aerenchyma development in waterlogged plants. Am J Bot 67: 18-22

Keeley JE, Franz EH (1979) Alcoholic fermentation in swamp and upland populations of Nyssa sylvatica: Temporal changes in adaptive strategy. Am Nat 113: 587-592

18Kelley PM, Freeling M (1984a) Anaerobic expression of maize fructose-1,6-diphosphate aldolase. J Biol Chem 259: 14180- 14183

Kelley PM, Freeling M (1984b) Anaerobic expression of maize glucose phosphate isomerase. J Biol Chem 259: 673-677

Kozlowski TT, Pallardy SG (1984) Effect of flooding on water, carbohydrate, and mineral relations. Jn TT Kozlowski, ed, Flooding and Plant Growth. Academic Press, FL pp 165-193

Kramer PJ (1951) Causes of injury to plants resulting from flooding of the soil. Plant Physiol 26: 722-736

Laan P, Blom, CWPM (1990) Growth and survival responses of Rujnex species to flooded and submerged conditions: The importance of shoot elongation, underwater photosynthesis and reserve carbohydrates. J Exp Bot 411: 77 5-783

McKee KL, Mendelssohn IA (1987) Root metabolism in the black mangrove (Avicennia germinans [L.] L.): response to hypoxia. Env Exp Bot 27: 147-156

McKee KL, Mendelssohn IA, Burdick DM (1989) Effect of long­term flooding on root metabolic response in five freshwater marsh plant species. Can J Bot 67: 3446-3452

McManmon M, Crawford RMM (1971) A metabolic theory of flooding tolerance: The significance of enzyme distribution and behavior. New Phytol 70: 299-306

Mendelssohn IA, Burdick DM (1988) The relationship of soil parameters and root metabolism to primary production in periodically inundated soils. Jn DD Hook, ed, The Ecology and Management of Wetlands Vol. 1: Ecology of Wetlands. Croon Helm Ltd, United Kingdom pp 398-425

Mendelssohn IA, McKee KL (1987) Root metabolic response of Spartina alterniflora to hypoxia. Jn RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford pp 239-253

Mendelssohn IA, McKee KL, Patrick WH Jr (1981) Oxygen deficiency in Spartina alterniflora roots: Metabolic adaptation to anoxia. Science 214: 439-441

Mendelssohn IA, Postek MT (1982) Elemental analysis of deposits on the roots of Spartina alterniflora Loisel. Am J Bot 69: 904-912

19Oliveira L (1977) Changes in the ultrastructure of mitochondria of roots of Trlticala subjected to anaerobiosis. Protoplasma 91: 267-280

Opik H (1973) Effect of anaerobiosis on respiratory rate, cytochrome oxidase activity and coleoptiles of rice (Oryza sativa L.). J Cell Sci 12: 725-739

Pezeshki SR (1990) A comparative study of the response of Taxodium distichum and Nyssa aguatica seedlings to soil anaerobiosis and salinity. For Ecol Manage 34: 531-541

Pradet A, Bomsel JL (1978) Energy metabolism in plants under hypoxia and anoxia. In DO Hook, RMM Crawford, eds, Plant Life in Anaerobic Environments. Ann Arbor Science, Ann Harbor, pp 89-118

Reid DM, Bradford KJ (1984) Effects of flooding on hormone relations. In TT Kozlowski, ed, Flooding and Plant Growth. Academic Press, Orlando, pp 195-219

Roberts JKM, Chang K, Webster C, Callis J, Walbot V (1989) Dependence of ethanolic fermentation, cytoplasmic pH regulation, and viability on the activity of alcohol dehydrogenase in hypoxic maize root tips. Plant Physiol 89: 1275-1278

Sachs MM, Freeling M, Okimoto R (1980) The anaerobic proteins of maize. Cell 20: 761-768

Save R, Serrano L (1986) Some physiological and growth responses of kiwi fruit (Actinidia chinensis) to flooding. Physiol Plant 66: 75-78

Schumacher TE, Smucker AJM (1985) Carbon transport and root respiration of split root systems of Phaseolus vulgaris subjected to short term localized anoxia. Plant Physiol 78: 359-364

Setter TL, Water I, Greenway H, Atwell BJ, Kupkanchanakul T (1987) Carbohydrate status of terrestrial plants during flooding. In RMM Crawford, ed. Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford, pp 411-433

Sherwin T, Simon EW (1969) The appearance of lactic acid in Phasedus seeds germinating under wet conditions. J Exp Bot 20: 776-785

Siji JW, Swanson GC (1973) Effect of petiole anoxia on phloem transport in squash. Plant Physiol 51: 368-371

2 0

Smith AH, ap Rees T (1979) Effects of anaerobiosis on carbohydrate oxidation by roots of Pisum sativum. Phytochem 18: 1453-1458

Smith AM, Hylton CM, Koch L, Woolhouse HW (1986) Alcohol dehydrogenase activity in the roots of marsh plants in naturally waterlogged soils. Planta 168: 13 0-138

Talbot RJ, Etheringtion JR, Bryant JA (1987) Comparative studies of plant growth and distribution in relation to waterlogging. XII. Growth, photosynthetic capacity and metal ion uptake in Salix caprea and S. cinnerea ssp oleifolia. New Phytol 105: 563-574

Trought MCT, Drew MC (1980) The development of waterlogging damage in wheat seedlings (Triticum aestivum L.). I. Shoot and root growth in relation to changes in the concentrations of dissolved gasses and solutes in the soil solution. Plant Soil 54: 77-94

Trought MCT, Drew MC (1981) Alleviation of injury to young wheat plants in anaerobic solution culture in relation to the supply of nitrate and other inorganic nutrients. J Exp Bot 32: 509-522

Vanlerberghe GC, Feil R, Turpin DH (1990) Anaerobic metabolism in the N-limited green alga Selenastrum minutum I. Regulation of carbon metabolism and succinate as a fermentation product. Plant Physiol 94: 1116-1123

VanToai TT, Fausey NR, McDonald MB Jr (198 5) Alcohol dehydrogenase and pyruvate decarboxylase activities in flood-tolerant and susceptible corn seeds during flooding. Agron J 77: 753-757

VanToai TT, Fausey NR, McDonald MB Jr (1987) Anaerobic metabolism enzymes as markers of flooding stress in maize seeds. Plant Soil 102: 33-39

Vartapetian BB (1982) Anaerobiosis and the theory of physiological adaptation of plant to flooding. Soviet Plant Physiol 29: 764-771

Vartapetian BB (1991) Flood-sensitive plants under primary and secondary anoxia: ultrastructure and metabolic responses. Jn MB Jackson, DD Davies, H Lambers eds, Plant Life Under Oxygen Deprivation. SPB Academic Publishing, The Hague, pp 201-216

21Vartapetian BB, Snkhchian HH, Generozova IP (1987) Mitochondrial fine structure in imbibing seeds and seedlings of Zea mays L* under anoxia. Jn RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford pp 205-223

Wample RL, Davis RW (1983) Effect of flooding on starch accumulation in chloroplasts of sunflower (H&lianthus annuus L.). Plant Physiol 73: 195-198

Wenkert W, Fausey NR, Watters HD (1981) Flooding responses in Zea mays L. Plant Soil 62: 351-366

Wignarajah K, Greenway H (1976) Effect of anaerobiosis on activities of alcohol dehydrogenase and pyruvate decarboxylase in roots of Zea mays. New Phytol 77: 575-584

Williams PH, Hill CB (1986) Rapid-cycling populations of Brassica. Science 232: 1385-1389

Wulff RD, Strain BR (1982) Effects of C03 enrichment on growth and photosynthesis in Desmodium paniculatum. Can J Bot 60: 1084-1091

CHAPTER TWO

SELECTION OF POPULATIONS OF RAPID-CYCLING BRASSICA RAPA L. DIFFERING IN THEIR RESPONSE TO ROOTZONE HYPOXIA

22

23Introduction

We are interested in the physiological mechanisms of waterlogging tolerance and how tolerance affects short-term phenotypic responses to hypoxic events in the rootzone. Short-term phenotypic responses to waterlogging may include morphological and ultrastructural changes, as well as alterations in metabolic pathways and enzyme activity (Crawford, 1978; Drew, 1987). Presence of traits in some species that are better adapted to waterlogged conditions than others, suggest that genetically determined characteristics confer a degree of tolerance to the stresses associated with waterlogging (Drew, 1987).

The presence of various physiological traits in plants occupying waterlogged sites has been taken as evidence that these traits are adaptations to hypoxia stress (Crawford, 1978; Hook, 1984; Mendelssohn et al. , 1981). An approach to identifying adaptive traits is to challenge a population with the environmental stress and observe those traits that appear preferentially in the progeny of the stressed plants compared to nonstressed plants. Since differentialresponses of individuals to an environmental stress ultimately affect their fitness, the stress exerts a selective pressure which leads to identification of genotypes that are better adapted to the environment. Stress as a selective pressure has been demonstrated in natural plant populations exposed to heavy metal contamination

24(Antonovics et al., 1971); in the laboratory in rapid-cycling Braseica populations inoculated with plant pathogens (Williams, 1985); and recently in a greenhouse study with radish populations stressed by air pollution (Gillespie and Winner, 1989).

In this study we have challenged a heterogeneous population of plants with waterlogging stress. Using visual criteria to select and interbreeding multiple generations we obtained a population of plants which grew well when waterlogged, from a population which did poorly. The study explored the use of a heterogeneous population of plants displaying differential responses to waterlogging stress as a genetic pool for traits conferring tolerance to the waterlogging stress. We chose rapid-cycling firassica rapa L. as the subject for these experiments because of the short life cycle (40-45 days), compact size, easy culture, and ability to reproduce under waterlogged conditions (Daugherty and Musgrave, 1989).

Materials and MethodsPlant Material

Rapid-cycling Brassica rapa L. CrGC#l-l (identification number of the Crucifer Genetics Cooperative, Madison, Wl) was used as the base (wild-type) population for the selection of two populations differing in their response to waterlogged conditions. The base population of rapid-cycling

25Brassica is genetically diverse and has served as a gene pool for many selection studies (Williams and Hill, 1986). Rapid-cycling B, rapa plants are self-incompatible and mass pollination must be used as the breeding technique. CrGC/l- 33 (marketed commercially by Carolina Biological Supply as MBasic Brassica rapa"), was used as a standard population for comparative studies with the selected populations produced as described below. Rapid-cycling Brassica rapa seeds were obtained from the Crucifer Genetics Cooperative, Madison WI.

Growth and Waterlogging ProceduresPlants were grown at 25°C in plastic multipot packs (55

cmVpack) containing standardized soil mix (Terra-lite RediEarth, W.R. Grace and Co. Cambridge, MA), four Osmocote slow release fertilizer (14:14:14) pellets, and two seeds of CrGC#l-l B. rapa. Cool white fluorescent bulbs provided a continuous light source (260 nmol/m2 sec). Plants were grown for one week after germination and watered with a wick-fed watering system containing deionized water. Thereafter, multipot packs were separated into control and waterlogged treatments. Control packs were maintained with the normal watering regime and the waterlogged packs submerged in water to one cm above soil line. Deionized water was replaced as needed to maintain the necessary water level. Waterlogging intensity (soil redox) was quantified

26using a glass-reference combination electrode (Sargent- Welch, Irving, TX). Redox values (Efc) ranged from 350 mV to 4 00 mV in the control packs and 200 mV to 2 50 mV in the waterlogged packs. Soil redox values were significantly lower in waterlogged treatments when compared to controls.

Selection and BreedingFoliage color was used as the criterion for selecting

individuals for successive generations. Individuals from the waterlogged base population were separated into "tolerant** and "sensitive" populations and mass pollinated to produce the next generation (Figure 2.1). After seven to nine days under waterlogged conditions the green and vigorous plants were selected for the tolerant pool and the yellow, less vigorous plants for the sensitive pool. Recurrent selection was continued for seven generations using the above criterion. After seven generations seed from all plants within a respective population were pooled. Identification numbers CrGC#l-51 (sensitive) and CrGC/1-52 (tolerant) were assigned to these two populations by the Crucifer Genetics Cooperative.

Chlorophyll AnalysisTotal chlorophyll concentration was used to rank plants

relative to their waterlogging responses. After seven days of waterlogging, three 0.25 cm1 leaf disks per plant were

Waterlogged Base Population 27

low vigor yellow follaga

high vigor . grsen fotlago

■aa>c

’oc* 'o

cft)Eo0)

cnc*o>cnOVo

Sensitive Pool Tolerant Pool

IP

1

yellow plants

mass pollination

ysllow plants

mass pollination

yellow plants

continued for seven generations

interbreeding only no further selection

1green plants • —

mass pollination

green plants

mass pollination

green plants

continued for seven generations

interbreeding only no further selection

*o0)co*-»c6

cIDE-*-■oa>

O'ccnC7*_oa)-*-»o

Sensitive Population Tolerant Population

Figure 2.1. Experiment design used to obtain distinctpopulations of Brassica rapa from the base B. rapa population which differed in their response to waterlogging stress.

28used for the chlorophyll determinations. Chlorophyll was extracted in 10 ml boiling methanol and absorbance was read at 650 nm and 665 run using a spectrophotometer (Bausch and Lomb Spectronic 20). Total chlorophyll was calculated with equations specified for extraction in methanol (Holden, 1976; MacKinney, 1941).

Growth AnalysisSequential harvests (Kvet et al. , 1971) were used to

perform a growth analysis on the response of the two selected populations and the CrGC#l-33 population under waterlogged and control conditions. Leaf area (measured with a LiCor Li-3100 leaf area meter), and root, shoot, and leaf dry weight after drying at 65°C to constant weight were recorded for individual plants harvested over an eight day waterlogging period.

Data AnalysisExperimental variables and the calculated ratios were

compared by a randomized block factorial analysis (Cochran and Cox, 1957) in terms of time, treatment, population, time x treatment, time x population, treatment x population, and time x treatment x population interaction effects. Values of the experimental variables were compared by an analysis of variance (ANOVA) and only those variables with significant P-values were compared by Least Significant

29Difference (LSD) analysis (Snedecor and Cochran, 1980). Standard errors of the means were determined for the dry weights of the populations.

Resultschlorophyll

Distribution of total chlorophyll concentrations from the base population under control and waterlogged conditions is presented in Figure 2.2. Waterlogging the base population plants significantly reduced the total chlorophyll concentration. Total chlorophyll concentration from leaves of base population plants subjected to waterlogged conditions ranged from 1 to 68 tig chlorophyll/cm2. However under control conditions, the distribution range of total chlorophyll was 50 to 86 tig chlorophyll/cm2. The tolerant and sensitive populations were selected from the base population based on the foliage color of the plants exposed to waterlogged conditions.

The histogram of total leaf chlorophyll distribution from waterlogged sensitive and tolerant populations is presented in Figure 2.3. Total chlorophyll concentration in the tolerant population exhibited a Gaussian distribution, whereas the sensitive population is skewed toward the left. A large proportion of plants in the waterlogged sensitive population had low concentrations of total chlorophyll in their leaves, with values ranging from 8 to 55 ng

3 0

chlorophyll/cm'1. In contrast, the waterlogged tolerant population had a greater proportion of plants in the upper range of chlorophyll concentrations. Values of total chlorophyll from waterlogged tolerant plants ranged from 13 to 70 jig chlorophyll/cm2.

Distributions of total chlorophyll concentrations from the sensitive and tolerant populations were similar and ranged from 50 to 88 jig chlorophyll/cm7 (Fig 2.4). Comparison of tolerant and sensitive chlorophyll distribution under control conditions indicated that the selection process did not alter the chlorophyll concentration in the leaves when the plants were not stressed. Mean total chlorophyll values from the two selected populations and the base population under control and waterlogged conditions are presented in Table 2.1.

Chlorophyll concentrations were not significantly different among the three populations under control conditions. However, under waterlogged conditions a significant reduction of chlorophyll was observed among all three populations. Chlorophyll concentrations in the waterlogged tolerant population were significantly greater than the concentrations in the waterlogged sensitive and base populations.

31

Total

20Chlorophyll Concentration in Base Population

CO

cO 15 CL

a>cno-*-*c a> ok.a> D_

10

I-. - | Watorioggod

25: O s S

£50

fiq ch forophy ll/cnrV

Control

75

Figure 2.2. Concentration of total chlorophyll from the base population of Brassica rapa under control and waterlogged conditions. Bars represent percentage of plants occurring in 5 nq chlorophyll/cm1 intervals. Distribution is based on 50 plants from the base population.

32

Total Chlorophyll Concentration23

E 2 Tolerant

in 20C_ga.*»—o 15

10CVol_(0CL

12 18 24 30 36 42 48 54 60 666

/zg c h lo ro p h y l l /c m2

Figure 2.3. Concentration of total chlorophyll from the tolerant and sensitive populations of Brassica rapa under waterlogged conditions. Bars represent percentage of plants occurring in 6 nq chlorophyll/cn1 intervals. Distribution is based on 80 plants from each population.

33Table 2.1. Total chlorophyll content of populations of Brassica rapa responding to waterlogging. Data are means from 50 individual plants within each population. Values with the same letter within a column are not significantly different (P > 0.05).

Total chloroohvll (uq/cm1!PoDulation Control WaterloaaedTolerant 69.13 a 44.99 b

Sensitive 73.35 a 23.93 a

Base 68.37 a 31.37 a

Waterlogged sensitive and base population chlorophyll concentrations were not significantly different from each other, however the mean value for the base population had a greater standard deviation (SD 20.31) than either the tolerant (SD 11.38) or sensitive (SD 7.26) population under waterlogged conditions. Plants in the base population varied greatly in their response to waterlogging conditions, therefore it was desirable to use a less variable, standard population for comparison studies. The commercially available CrGC/l-3 3 population, which is homogenous and contains little variation within the population, was used in subsequent analyses to compare with the tolerant and sensitive populations.

34

Total Chlorophyll Concentration

20incoK 15

oCP 10ocQ)O0)CL

50 53 56 59 62 65 66 71 75 78 81 64 87

fig c h lo ro p h y l l /c m2

Figure 2.4. Concentration of total chlorophyll from the tolerant and sensitive populations of Brassica rapa under control conditions. Bars represent percentage of plants occurring in 3 pg chlorophyll/cm1 intervals. Distribution is based on 80 plants from each population.

35Growth Analysis

Root dry weights for the three populations arepresented in Figure 2.5. There was no significant difference in overall root dry weight among the threepopulations under control conditions. Root weight in control plants increased substantially over time in allpopulations. Mean dry root weights of the control plants on Day 8 of treatment were 31.3 mg, 3 6.5 mg, and 31.1 mg for the CrGC#l—33, tolerant, and sensitive populations respectively.

Root weights from the waterlogged treated plants did not have a substantial increase in dry matter over time (Fig 2.5 and Table 2.2). Root weights of waterlogged plants were significantly different from the control plants four, six, and eight days after waterlogging, but not after two days of waterlogging (Fig 2.5). The root dry weights of the sensitive and the crGC/i-33 plants did not significantly increase over time. The tolerant plants did havesignificantly greater dry root weights after eight days of waterlogging than the sensitive or CrGC#l-3 3 plants (Table 2 .2).

Shoot dry weights of the three populations were also significantly affected by the waterlogging treatment. Plants maintained under control conditions had an increase in shoot dry weight over time (Fig 2.6) . The mean shoot dry weights of the control plants on Day 8 of treatment were

3682.7 mg, 78.9 mg, 71.8 mg for the tolerant, sensitive, and CrGC/1-33 respectively. These values were not significantly different from each other.

Table 2.2. Comparison of Brassica rapa root and shoot weights responding to waterlogging treatment for six and eight days. Data are means from four replications. Values with the same letter within a column are on a specific day are not significantly different (P > 0.05).

Sample time Population Dry Weight (mg/plant)Day 6 Shoot

Tolerant 10.02 b 39.12 bSensitive 6.77 ab 21.00 aCrGC/1-33 6.37 a 31.07 ab

Day 8Tolerant 18.17 b 66.62 cSensitive 7.27 a 32.25 aCrGC/1-33 9.97 a 44.80 b

Shoot dry weight did not change relative to controls until six days into the waterlogging treatment (Fig 2.6). The sensitive and CrGC/1-33 populations slowed in the amount of shoot dry matter gain, whereas the shoot weights of the tolerant plants maintained a constant increase. The three populations under waterlogged conditions were significantly different from control plants on Day 6, however, the tolerant population was not significantly different from the

R o o t D r y W e i g h t (mg per plant)— - • M W W O t* O « O W O W O t J l O O i O U O O l O O l O W O W O U O O l O U

•KH

Figure 2.5. Root dry weight of populations of Brassica rapa subjected to control (*) or waterlogged (▼) conditions for eight days. Means are from dry weights of four replicates ± s.e. * Represents significant difference fro* control values (P £ 0.05).

S h o o t D r y W e i g h t (mg per plant)

- * M U > u ) 0 ) ^ n e o - » M U * u i a » > j o i e Qo o o o o o o o o o o o o o o o o o o o o o o o o o o o o o o o o

»3inr-t-3"o

(•a<-►3a3aO

W

M

3.

Q .

Figure 2.6. Shoot dry weight of populations of Brassica rapa subjected to control (•) or waterlogged (*) conditions for eight days. Means are froe dry weights of four replicates ± s.e. * Represents significant difference fros control values (P 5 0.05).

39controls on Day B. In addition, tolerant plants maintained under waterlogged conditions had significantly greater shoot weights than the sensitive plants on Day 6 and Day 8 (Table 2.2). Sensitive and CrGC#l-33 waterlogged plants did not show significant differences in the dry shoot weights on Day 6, but were significantly different on Day 8.

DiscussionSelection of plants which differed in their

waterlogging response from the base population of rapid- cycling Brassica rapa L. gave rise to the tolerant and sensitive populations. The base population contained sufficient variability to provide individuals which could be used for this selection procedure (Fig 2.2). Selecting individuals and mass pollinating to produce the next generation was the same procedure used to develop new disease-resistant Brassica populations (Williams, 1985). Like the disease-resistant rapid-cycling Brassica populations, our selected populations were relatively homogenous in plant morphology and flowering time. However, under the waterlogging stress the plants within the two populations did exhibit some variation in morphology and waterlogging response traits, but this variability was confined to a few individuals.

Several screening methods have been attempted to differentiate sensitive and tolerant plants under

4 0

waterlogging stress (McNamara and Mitchell, 1989; Nelson et al. ,1983). Usually the approach focuses on a morphological or biochemical factor that is altered in response to rootzone hypoxia. A visual change that often occurs is the yellowing of foliage with prolonged waterlogging. The yellowing may be due to changes in nutrient availability (Drew and Sisworo, 1979), alteration of carbon assimilation (Laan and Blom, 1990; Pezeshki, 1990), status of nitrogen in the rootzone environment (Drew and Sisworo, 1979; Trought and Drew, 198 0) and in some plants to the physical disruption of chloroplasts by starch grains (Wample and Davis, 1983) .

The selection criterion used to develop two populations distinguished by degree of foliar yellowing under waterlogging stress was quantified by measuring chlorophyll concentrations. When waterlogged, the base population had a large standard deviation in the mean chlorophyll concentration (SD 20.31), whereas under control conditions variability among the base population plants was small (SD 9.09). Taking advantage of this variability in the foliage color we were able to select different populations. The selection process produced populations that had altered chlorophyll concentrations only under the waterlogging stress, whereas in unstressed controls the chlorophyll concentrations in the tolerant and sensitive populations were indistinguishable (Pigs 2.3 and 2.4). Our method of

41monitoring chlorophyll concentrations is a non-lethal screening technique which was found to be an effective method for evaluating the degree of waterlogging tolerance as reflected in production of plant biomass.

Growth analysis from the three populations under waterlogging conditions indicated that overall dry weight was not significantly altered until more than four days after waterlogging. The root dry weight was the first factor to respond to the waterlogging treatment, with a duration of eight days needed to observe significant changes in shoot dry weight (Fig 2.5). This response time agrees with the results of Trought and Drew (1980) and McNamara and Mitchell (1989), who monitored wheat and tomato responses to waterlogging stress. They did not observe any changes in dry weight until after a minimum duration of four days of waterlogging.

Especially striking was the reduction in root and shoot growth with waterlogging in the sensitive population while the tolerant population had continued growth (Figs 2.5 and 2.6). The rate of growth was greatest in the tolerant population followed by the CrCG#l-33 and sensitive populations. Using these populations as a tool we will relate timing of onset of waterlogging damage shown by the growth analysis to timing of potential causative factors.

For example, we observed that root systems from tolerant and CrGC/1-3 3 plants had developed adventitious

42roots by the eighth day of waterlogging, whereas sensitive roots had barely formed any new root growth. Formation of adventitious roots and aerenchyma has been reported to increase the tolerance of plants to a waterlogged environment (Drew et al. , 1979; Gill, 1975; Kawase andWhitmoyer, 1980; Wenkert, et al. , 1981). Similarly,maintenance of continued shoot and root dry matter gain in the tolerant population indicated that these plants were capable of continued use of the available carbohydrates even under waterlogging stress. Further investigations of the carbohydrate status in the three populations may indicate the nature of these tolerant and sensitive responses to waterlogging stress. We will also report on how these populations differ with regard to the behavior of root enzymes associated with hypoxia responses. Development of populations differing in waterlogging tolerance will make possible a better understanding of these environmentally triggered responses.

SummaryChallenging an heterogenous population of plants

(rapid-cycling Brassica rapa L.) with waterlogging stress, we selected plants which differed in their response to rootzone hypoxia. Selected individuals from the waterlogged wild-type population were separated into "tolerant" and "sensitive" populations and mass pollinated to produce the next generation. Under waterlogged conditions the green and

43vigorous plants were selected for the tolerant pool and the yellow, less vigorous plants were selected for the sensitive pool. Recurrent selection was continued for seven generations. Chlorophyll concentrations in the waterlogged tolerant population were significantly greater than the concentrations in the waterlogged sensitive population. Total chlorophyll concentrations were not significantly different between the populations under control conditions. Sequential harvesting was used to perform a growth analysis on the waterlogging response of the two selected populations. There was no significant difference when populations where grown under control conditions, however the tolerant plants had significantly greater root and shoot dry weights after eight days of waterlogging than the sensitive plants. The results demonstrate that stress- specific differences in population performance can be achieved through recurrent selection.

44Literature Cited

Antonovics J, Bradshaw AD, Turner RG (1971) Heavy metal tolerance in plants. Adv Ecol Res 7: 1-85

Cochran WG, Cox GM (1957) Experimental designs Ed 2. John Wiley and Sons Inc, New York, 611 pp

Crawford RMM (1978) Metabolic adaptation to anoxia. Jn DD Hook, RMM Crawford, eds, Plant Life in Anaerobic Environments. Ann Arbor Science, MI, pp 119-155

Daugherty CJ, Musgrave ME (1989) Selection of rapid-cycling Brassica tolerant to hypoxia for use in space biology experiments. Am Soc Grav Space Biol Bull 3: 82

Drew MC (1987) Mechanisms of acclimation to flooding and oxygen shortage in non-wetland species. Jn RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford, pp 321-331

Drew MC, Jackson MB, Giffard SC (1979) Ethylene-promoted adventitious rooting and development of cortical air spaces (aerenchyma) in roots may be adaptive responses to flooding in Zea mays L. Planta 147: 83-86

Drew MC, Sisworo EJ (1979) The development of waterlogging damage in young barley plants in relation to plant nutrient status and changes in soil properties. New Phytol 82: 301- 314

Gill CJ (1975) The ecological significance of adventitious rooting as a response to flooding in woody species with special reference to Alnus glutinosa (L.) Gaertn. Flora 164: 85-97

Gillespie CT, Winner EW (1989) Development of lines of radish differing in resistance to 0, and SO;. New Phytol 112: 353-361

Holden M (1976) Chlorophylls. In TW Goodwin, ed, Chemistry and Biochemistry of Plant Pigments II. Academic Press, New York, p 7

Hook DD (1984) Adaptations to flooding with fresh water. Jn TT Kozlowski, ed, Flooding and Plant Growth. Academic Press, Orlando, pp 265-294

Kawase M, Whitmoyer RE (1980) Aerenchyma development in waterlogged plants. Am J Bot 67: 18-22

45Kvet J, Ondok JP, Necas J, Jarvis PG (1971) Methods of growth analysis. Jn Z Sestak, J Catsky, PG Jarvis, eds, Plant Photosynthetic Production. W. Junk N.V. Publishers, The Hague, pp 343-391

Laan P, Blom CWPM (1990) Growth and survival responses of Rumex species to flooded and submerged conditions: The importance of shoot elongation, underwater photosynthesis and reserve carbohydrates. J Exp Bot 41: 77 5-783

MacKinney B (1941) Absorption of light by chlorophyll solutions, j Biol chem 140: 315-322

McNamara ST, Mitchell CA (1989) Differential flood stress resistance of two tomato genotypes. J Am Soc Hort Sci 114: 976-980

Mendelssohn IA, McKee KL, Patrick WH Jr (1981) Oxygen deficiency in Spartina alterniflora roots: Metabolic adaptation to anoxia. Science 214: 439-441

Nelson RB, Davis DW, Palta JP, Laing DR (1983) Measurement of soil waterlogging tolerance in Phaseolus vulgaris L.: A comparison of screening techniques. Scientia Hortic 20: 303-313

Pezeshki SR (1990) A comparative study of the response of Taxodium distichum and Nyssa aquatica seedlings to soil anaerobiosis and salinity. For Ecol Manage 34: 531-541

Snedecor GW, Cochran WG (1980) Statistical methods 7th ed. Iowa State University Press, Ames, pp 59 3

Trought MCT, Drew MC (1980) The development of waterlogging damage in wheat seedlings (Triticum aestivun L.) I. Shoot and root growth in relation to changes in the concentrations of dissolved gases and solutes in the soil solution. Plant Soil 54: 77-94

Wample RL Davis RW (198 3) Effect of flooding on starch accumulation in chloroplasts of sunflower (Helianthus annuus L.). Plant Physiol 73: 195-198

Wenkert H, Fausey NR, Hatters HD (1981) Flooding responses in Zea nays L. Plant Soil 62: 351-366

Williams PH (1985) Crucifer Genetics Cooperative Resource Book. Department of Plant Pathology, University of Wisconsin-Madison, HI, pp. 100

Williams PH, Hill CB (1986) Rapid-cycling populations of Brassica. Science 232: 1385-1389

CHAPTER THREE

CARBOHYDRATE STATUS OF POPULATIONS OF RAPID-CYCLING BRASSICA RAPA L. DURING WATERLOGGING STRESS

46

47Introduction

Carbohydrate concentrations are determined by the balance between consumption in respiration and growth, and production of assimilates by photosynthesis. Plants exposed to hypoxic conditions induced by waterlogging may have this carbohydrate balance disrupted. The partitioning of carbohydrates may be influenced by differences in metabolic pathways, alteration of enzyme activity, and morphological and ultrastructural changes that occur when plants are subjected to waterlogging stress (Crawford, 1978; Drew, 1987). Soil waterlogging greatly inhibits gaseous diffusion from the atmosphere leading to hypoxia and subsequent anoxia around the root zone. Under hypoxic conditions, a continuous supply of soluble carbohydrates to root tissues is needed to help preserve and maintain growth of the root tips which are a critical component of plant survival (Barta, 1988; Jackson and Drew, 1984).

Several reports suggested that a decreased carbohydrate supply may occur following waterlogging due to stomatal closure induced by reduced water permeability of the roots (Kozlowski and Pallardy, 1984; Save and Serrano, 1986; Trought and Drew, 1980). However, hypoxia also leads to reduced translocation rates thereby decreasing the movement of assimilates from leaves. The basis of increased carbohydrate accumulation in the leaves has been hypothesized to be caused by reduced utilization by the

48plant as a whole (Setter, et al., 1987; Sijl and Swanson,197 3). The overabundance of carbohydrates subsequently leads to disruption of the photosynthetic apparatus by excessive starch accumulation in the thylakoid membranes (Cave et al., 1981; Talbot et al., 1987; Wample and Davis, 1983; Wulff and Strain, 1982). Roots, which are a major sink for carbohydrates, become altered in membrane properties, cellular energy balance, and overall metabolism under waterlogged conditions thereby limiting the demand for assimilates from the leaves (Jackson and Drew, 1984).

Although several workers have described general waterlogging effects on carbohydrates (Setter, et al . , 1987 ; Talbot et al., 1987; Wample and Davis, 1983) to date no one has investigated leaf carbohydrate status of waterlogged stressed plants over a continued period of rootzone hypoxia. We have selected populations of Brassica rapa L. that differ in their response to waterlogged conditions (Chapter Two), and in this study we report leaf carbohydrate status of these tolerant and sensitive populations when exposed to waterlogging.

Materials and MethodsPlant Material

Selected individuals from the base population were separated into "tolerant" and "sensitive" populations and mass pollinated to produce the next generation (Chapter

49Two) . Plants were grown according to the procedure outlined in Chapter Two. Our populations of rapid-cycling Brassica rapa CrGC#l-51 (sensitive) and CrGC#l-52 (tolerant) are available from the Crucifer Genetics Cooperative, Madison WI.

Preparation of Leaf Material for Carbohydrate AnalysisAll leaves except the cotyledons were removed from

individual plants and quick killed in a microwave oven for 15 seconds, dried for 2 4 hours at 60°C and stored desiccated at 4"C. The dried leaves of each plant were ground to a powder in a homogenizer and dispensed into 10 mg samples for analysis.

Carbohydrate AnalysisSamples for carbohydrate determination were extracted

using the method of Sasek et al. , (1985). Soluble sugarswere separated from starch by sequential centrifugations and resuspensions in methanol, chloroform, and water (12:5:3 v/v/v) (Haissig and Dickson, 1979). Soluble carbohydrates were maintained in the roethanol-water fraction. The starch pellets were resuspended in l.o ml of 0.2 M KOH and boiled for 30 minutes. After cooling, pH was adjusted to approximately 5.5 by adding 0.2 ml of 1.0 M acetic acid. Gelatinized starch was digested to glucose units with 34

50units of amyloglucosidase (Sigma Chemical A-3042, St. Louis, MO) which had previously been dialyzed overnight against 50 mM Na-acetate buffer (pH 4.5) (Pharr et al., 1985). After one hour at 55°C, samples were boiled to denature enzyme. Starch and soluble carbohydrate concentrations (expressed as glucose units for both assays) were determined by the glucose oxidase-iodide assay (Eball, 1969) using 1000 units of glucose oxidase (Sigma Chemical G-S135, St. Louis, MO) . Glucose units were calculated from a standard curve prepared with a glucose standard solution (1.0 mg/ml) and absorbance was determined at 3 53 nm.

Data AnalysisExperimental variables and the calculated ratios were

compared by a randomized block factorial analysis (Cochran and Cox, 1957) in terms of time, treatment, population, time x treatment, time x population, treatment x population, and time x treatment x population interaction effects. Values of the experimental variables were compared by an analysis of variance (ANOVA) and only those with the significant F- values were compared by Least Significant Difference (LSD) analysis (Snedecor and Cochran, 1980).

Chloroplast UltrastructureLeaves were obtained from control plants and plants

that were waterlogged for four days. Leaf segments were cut

51into 2 x 3 mm strips while immersed in 1.5% glutaraldehyde: 1% Acrolein in 0.1 M Na-Cacodylate (pH 7.2) buffer. After two hours, fixative was removed and samples were rinsed in three 10-minute changes of 0.1 H Na-Cacodylate buffer. Post-fixation was in It 0s04 for two hours followed by a 10 minute distilled water rinse. Samples were initially dehydrated in a 50% and 70% ethanol series. After second 70% ethanol wash, 80% ethanol containing 2% uranyl acetate was added and samples were refrigerated overnight. The dehydration series was continued with 95% and 100% ethanol. Samples were embedded with 100% LR White hard grade epoxy (Ted Pella, Redding CA) and polymerized for 24 hours at 60°C. Sections were cut on a Porter-Blum MT2 ultra­microtome and post-stained with lead citrate and uranyl acetate. Grids were examined on a JEOL 100 CX transmission electron microscope.

Light Microscopy FixationSections were prepared and fixed in same manner as

ultrastructure protocol. Light microscopy sections were post-stained with 1% Toluidine Blue 0 in 2% sodium borate and then cleared with 100% methanol. Sections were examined with a Nikon Microphot-FXA.

52Results

Soluble CarbohydratesWaterlogging produced significantly greater soluble

carbohydrate concentrations in all populations as compared to the control plants (Fig 3.1). Plants subjected to waterlogged conditions showed a significant increase in soluble carbohydrates when compared to the control plants as indicated by asterisks. Overall mean soluble carbohydrate concentrations of leaves in the waterlogged and control treatments were 20.7 nq glucose units/mg dry weight and 7.1 jig glucose units/mg dry weight respectively. Leaf soluble carbohydrates of waterlogged plants were usually three times greater than the concentration in the control leaves.

Soil redox values began to decrease after only 12 hours of waterlogging. Redox values in the control pots were relatively constant with values near 350 mV, but significantly lower in the waterlogged treatments (200 mV) after 48 hours of waterlogging. The decrease in oxygen tension corresponded with a steady increase in soluble carbohydrates in the waterlogged plants throughout the first two to three days of treatment, whereas the control plants maintained a relatively constant concentration of soluble carbohydrates. A decrease in carbohydrates occurred on Day 6 in the waterlogged treated plants coinciding with floral production.

53

Soluble Carbohydrates (glucose units)

y /-

hours

symbols

• CrGCfl —33

■ Tolerant

♦ Sensitive

filled—waterlogged hollow—control

Time

Figure 3.1. Leaf soluble carbohydrates of populations of Brassica rapa responding to waterlogging stress and concurrent soil redox values (inset). Soil redox values are means based on six readings per time interval. Carbohydrate means based on four replications. * Represents significant differences from control values (P £ 0.05).

54Differences In response of soluble carbohydrates were

observed in the waterlogging sensitive and tolerant populations of S. rapa. Soluble carbohydrates increased in leaves of the sensitive plants after only 12 hours of waterlogging and were significantly greater than the control plants. In addition, waterlogged sensitive plants had significantly greater concentrations of solublecarbohydrates than the waterlogged tolerant and CrGC#l-3 3 plants at Hours 12 and 18 (Table 3.1). All waterlogged populations had significantly greater concentrations of soluble carbohydrates than the controls after 24 hours of waterlogging. Elevated concentrations of solublecarbohydrates in the waterlogged populations were maintained through Day 6.

Table 3.1. Comparison of soluble carbohydrateconcentrations in leaves of populations of Brassica rapa responding to waterlogging stress. Waterlogging treatment was maintained for 12, 18, or 2 4 hours. Data are meansbased on four replications, values with the same letter in the same column are not significantly different (P > 0.05).

PoDulation

Lenath of treatment (hours)12

uq

18glucose units/ma drv

24weicrht

Tolerant 2 . 5 a 9.0 a 12.5 a

Sensitive 9.1 b 16.8 b 22. 5 a

CrGC#l-33 2 . 3 a 5.0 a 16. 5 a

55Starch Concentration

Leaf starch concentration of B. rapa plants under waterlogged and control conditions is presented in Figure 3.2. Unlike the soluble carbohydrate concentrations which responded promptly to waterlogging stress, starch concentration was not significantly different between waterlogged and control plants until after 24 hours of waterlogging. Starch concentrations in the waterlogged plants were then significantly different from controls until Day 6. Mean values of waterlogged and control leaves on Day 2 were 313 pg glucose units/mg dry weight and 105 pg glucose units/mg dry weight respectively.

Starch decreased dramatically in the plants after four days of waterlogging, but then increased after six days. Starch concentrations in the control plants did not change significantly after four days, however concentrations increased following Day 6. After six days of waterlogging there was no significant difference in the starch concentration of waterlogged (107 pg glucose units/mg dry weight) and control (119 pg glucose units/mg dry weight) plants.

Starch concentrations in the three B. rapa populations under waterlogging stress are presented in Table 3.2. Throughout the waterlogging treatment, leaves of the tolerant population had lower concentrations of starch than either CrGC#l-33 or the sensitive population. Starch

56

Starch(glucose units)

symbols• CrCCf 1—33 ■ Tolerant♦ Sensitive

300Cn

filled—waterlogged hollow—control•■= 200

o 100

CT>

2 41 6 8Time (days)

Figure 3.2. Loaf starch of populations of Brassica rapa responding to waterlogging stress. Means based on four replications. * Represents significant differences from control values (P s 0.05).

57concentrations of waterlogged tolerant plants were significantly lower on Day 2 when compared to waterlogged sensitive and CrGC#l-33. In all three populations, leaf starch concentration dramatically increased between Day 1 and 2 when waterlogged.

Table 3.2. Comparison of leaf starch concentrations in populations of Brassica rapa responding to waterlogging stress. Waterlogging treatment was maintained for one, two, or four days. Data are means based on four replications. Values with the same letter in the same column are notsignificantly different (P > 0.05) .

Lenoth of treatment (davsl

PoDulation1

UQ2 4

alucose units/mcr dry weiahtTolerant 189 a 243 a 272 a

Sensitive 200 a 377 b 303 a

CrGC#l-3 3 201 a 318 b 328 a

Light MicroscopyLeaf sections of £.

morphological manifestationsrapa populations showed the of the carbohydrate changes in

response to waterlogging (Fig 3.3). The most striking difference between waterlogged and control plants is the size and concentration of starch grains in the palisade cells. Plants waterlogged for four days had extensive starch grains filling most of the chloroplast. The control plants contained starch grains, but the grains were smaller

Figure 3.3. Light micrographs of leaf sections from leaves of A: control plant (magnification 129x), B: waterlogged sensitive plant (magnification 129x), and C: waterlogged tolerant plant (magnification 72x). Duration of waterlogging was four days. Starch grains (SG).

Figure 3.4. Electron Micrographs of parenchyma cell sections fros leaves of A: control tolerant plant, B: waterlogged sensitive plant, and C: waterlogged tolerant plant. Duration of waterlogging was four days. Starch grains (SG), ThylaKoid membranes (TM). m

60and localized near the periphery of the cell. Under waterlogged conditions, the concentration of leaf starch in the three B . rapa populations did not appear different when viewed at the light microscopic level.

Chloroplast UltrastructureExamination of leaf cells of waterlogged and control

plants revealed differences in starch accumulation within the tolerant and sensitive populations (Figure 3.4). Large starch grains were abundant in the chloroplasts of the tolerant and sensitive plants when waterlogged for four days. However, the thylakoid membranes were distorted in the waterlogged sensitive plants. Thylakoid membranes in the waterlogged tolerant plants were still intact despite large starch grains in the chloroplast. The control plants also contained a few starch grains in the chloroplasts but had intact thylakoid membranes.

DiscussionLeaf carbohydrate status appears to be different in the

waterlogging tolerant and sensitive populations of Brassica rapa L. In Chapter Two we reported the maintenance of continued shoot and root dry matter gain in the tolerant population, whereas the sensitive population did not show this increase when the plants were waterlogged. Soluble carbohydrates and starch concentrations were accumulating in

61the leaves of the sensitive plants and possibly not translocated to the roots or other sinks as was the case for the tolerant plants (Figs 3.1 and 3.2).

Increased carbohydrate content in foliage under conditions of rootzone hypoxia has been hypothesized to be related to diminished metabolic sink activity of the root system (Jackson and Drew, 1984). The increased leaf carbohydrate concentration when the B . rapa populations were subjected to rootzone hypoxia is in agreement with previous reports. Wheat and alfalfa grown in waterlogged fields had twice the soluble carbohydrate concentration in the shoots compared to plants grown in well-drained plots (Setter et al. , 1987; Barta, 1988). When subjected to waterloggedconditions, two selections of Apios americana had an increase in specific leaf weight and leaf carbohydrate concentrations and a decrease in tuber yield when compared to control plants (Musgrave et al., 1991).

Carbohydrate accumulation in the leaves has been proposed to be caused by the reduction in translocation of photosynthetic products, specifically, translocating assimilates to roots of waterlogged plants. The reduction in the translocation of carbohydrates to the roots was reported for waterlogged wheat and woody plants (Kozlowski and Pallardy, 1984; Trought and Drew, 1980). This reduction in transported assimilates can occur very quickly under waterlogging stress. Within 30 minutes of waterlogging the

62rootzone, the rate of accumulation of l4C-labeled assimilates in cucumbers decreased by more than half (Siji and Swanson, 1973). The supply of oxygen to the B. rapa roots, as indicated by the decreasing soil redox values, was declining after only 12 hours of waterlogging (Fig 3.1). The lack of oxygen for root metabolic functions could have lessened the demand for assimilates, subsequently causing an increase in leaf carbohydrates within this short period of time.

Observation of palisade cells and chloroplast ultrastructure in the waterlogged sensitive plants indicated that accumulation of starch may have been disruptive to the cells. The partitioning of starch in these cells was not in discrete granules as was the case for starch in the waterlogged tolerant plants (Fig 3.4). We observed a major disruption of the thylakoid membranes possibly by the excessive starch in the waterlogged sensitive population. Loss of thylakoid membrane integrity was recently reported in cucumber leaves (Schaffer et al., 1991). The premature leaf chlorosis was induced by the high starch accumulation in the chloroplasts. Wample and Davis (1983) reported a slight displacement of the thylakoid membranes due to starch accumulation in waterlogged sunflower plants. C02enrichment also affected chloroplast ultrastructure in Trifolium subterraneum and Desmodium pan leu latum by inducing an accumulation of large starch grains which altered the

63normal structure and function of the chloroplasts (Cave et al., 1981; Wulff and Strain, 1982).

The accumulation of starch in the leaves of waterlogged stressed plants may not always be a detrimental factor. Bensari et al. , (1990) proposed that excessive foliar starch content may play a role in resistance to stress. They found the excessive starch in soybean leaves could be immediately utilized by the sinks when the stress was removed thereby avoiding any carbohydrate limitation. The waterlogged tolerant population did show an excess amount of starch in the leaves without disruption of the thylakoid membranes. The excess starch in leaves of waterlogged plants may provide an immediate carbohydrate source for the new adventitious roots or older primary roots once the waterlogged stress is removed.

Carbohydrate accumulation and partitioning may play a role in response of a plant exposed to waterlogging stress. Obviously, a mechanism that allows the plant to continue photosynthesis without disruption by excess storage products would be advantageous. However, excessive accumulation of carbohydrates may be detrimental, as was observed in the sensitive population. If, under waterlogged conditions, carbohydrates are translocated to the roots this must indicate metabolic processes are still active. Examination of the roots of the populations will be the subject of a subsequent investigation.

64Summary

Soil waterlogging greatly inhibits gaseous diffusion from the atmosphere, leading to hypoxia and subsequent anoxia around the root zone. Under hypoxic conditions, a continuous supply of soluble carbohydrates to root tissues is needed to help preserve and maintain growth of the root tips which are a critical component of plant survival. However, rootzone hypoxia usually leads to reduced translocation rates thereby decreasing the movement of assimilates from the leaves to the roots. Leaf carbohydrate status in three populations of Brassica rapa which differed in their response to waterlogging was analyzed during an eight day period of rootzone hypoxia. Soil redox values decreased after 12 hours of waterlogging. The decrease in oxygen tension corresponded with a steady increase in soluble carbohydrates in the leaves of waterlogged plants, whereas the control plants maintained a relatively constant concentration of soluble carbohydrates. Unlike the soluble carbohydrate concentration which responded promptly to waterlogging stress, starch concentration was not significantly different between waterlogged and control plants until 24 hours after soil waterlogging. Light microscopy of leaf sections from waterlogged plants revealed extensive starch grain deposition in most of the cells. Cells in the control plants also contained starch grains, but these grains were smaller and localized near the

6 5

periphery of the cell. Examination of chloroplasts from waterlogged plants in the sensitive population by transmission electron microscopy showed a disruption in the thylakoid membranes, possibly by excessive starch accumulation. Thylakoid membranes in waterlogged plants from the tolerant population were still intact despite large starch grains in the chloroplast. Control plants had small starch grains and intact thylakoid membranes in the chloroplast. Soluble carbohydrates, starch concentrations, and chloroplast ultrastructure of the sensitive population indicated the carbohydrates were accumulating in the leaves and possibly not translocated to the roots or other sinks. The leaf carbohydrate concentrations and chloroplast ultrastructure of the tolerant population during waterlogging stress indicated that these plants are capable of continued use of the available carbohydrates.

66Literature Cited

Barta AL (1988) Response of field grown alfalfa to root waterlogging and shoot removal. I. Plant injury and carbohydrate and mineral content of roots. Agron J 80: 889- 892

Bensari M, Calmes J, Viala G (1990) Deficit hydrigue et distribution du carbone photosynthetique entre saccharose et amidon. Acta Ecol 11: 843-855

Cave G, Tolley LC, Strain BR (1981) Effect of carbon dioxide enrichment on chlorophyll content, starch content and starch grain structure in Trifolium subterraneum leaves. Physiol Plant 51: 171-174

Cochran WG, Cox GM (1957) Experimental designs Ed 2. John Wiley and Sons Inc, New York, 611 pp

Crawford RMM (1978) Metabolic adaptation to anoxia. Jn DD Hook, RMM Crawford, eds, Plant Life in Anaerobic Environments. Ann Arbor Science, MI, pp 119-155

Drew MC (1987) Mechanisms of acclimation to flooding and oxygen shortage in non-wetland species. Xn RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific, Oxford, pp 321-331

Eball LF (1969) Specific total starch determinations in conifer tissues with glucose oxidase. Phytochem 8: 25-36

Haissig BE, Dickson RE (1979) Starch measurement in plant tissue using enzymatic hydrolysis. Physiol Plant 47: 151- 157

Jackson MB, Drew MC (1984) Effects of flooding on growth and metabolism of herbaceous plants. Jn TT Kozlowski ed, Flooding and Plant Growth. Academic Press, Orlando, pp 47- 128

Kozlowski TT, Pallardy SG (1984) Effect of flooding on water, carbohydrate, and mineral relations. Jn TT Kozlowski ed, Flooding and Plant Growth. Academic Press, Orlando, pp 165-193

Musgrave ME, Hopkins AG, Daugherty CJ (1991) Oxygen insensitivity of photosynthesis by waterlogged Apios americana. Env Exp Bot 31: 117-124

67Pharr DM, Huber SC, Sox HN (198 5) Leaf carbohydrate status and enzymes of translocate synthesis in fruiting and vegetative plants of Cucumis sativus L. Plant Physiol 77: 104-108

Sasek TW, DeLucia EH, Strain BR (1985) Reversibility of photosynthetic inhibition in cotton after long-term exposure to elevated C03 concentrations. Plant Physiol 78: 619-622

Save R, Serrano L (1986) Some physiological and growth responses of kiwi fruit (Actinidia chinensis) to flooding. Physiol Plant 66: 75-78

Schaffer AA, Nerson H, Zamski E (1991) Premature leaf chlorosis in cucumber associated with high starch accumulation. J Plant Physiol 138: 186-190

Setter TL, Water I, Greenway H, Atwell BJ, Kupkanchanakul T (1987) Carbohydrate status of terrestrial plants during flooding. In RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford, pp 411-433

Siji JW, Swanson GC (1973) Effect of petiole anoxia on phloem transport in squash. Plant Physiol 51: 368-371

Snedecor GW, Cochran WG (1980) Statistical methods 7th ed. Iowa State University Press, Ames, pp 593

Talbot RJ, Etherington JR, Bryant JA (1987) Comparative studies of plant growth and distribution in relation to waterlogging. XII. Growth, photosynthetic capacity and metal ion uptake in Salix caprea and 5. cinnerea ssp oleifolia. New Phytol 105: 563-574

Trought MCT, Drew MC (1980) The development of waterlogging damage in wheat seedlings (Triticum aestivum L.) I. Shoot and root growth in relation to changes in the concentrations of dissolved gases and solutes in the soil solution. Plant Soil 54: 77-94

Wample RL, Davis RW (1983) Effect of flooding on starch accumulation in chloroplasts of sunflower (Halianthus annuus L.). Plant Physiol 73: 195-198

Wulff RD, Strain BR (1982) Effects of C03 enrichment on growth and photosynthesis in Dasmodium paniculatum. Can J Bot 60: 1084-1091

CHAPTER FOUR

EARLY CHANGES IN ROOT METABOLISM OF THREE POPULATIONS OF RAPID-CYCLING BRASSICA RAPA L. DURING WATERLOGGING

68

69Introduction

Many responses occur in the roots of plants exposed to hypoxic conditions. During the initial hours of hypoxia, metabolic processes in the roots are immediately affected because of lack of sufficient oxygen in the rootzone. Metabolic alterations may include acceleration of the glycolytic cycle (Pasteur effect) to produce excess ATP during short-term waterlogging (John and Greenway, 1976; Crawford, 1978; Pradet and Bomsel, 1978).

Alterations in activities of cytochrome oxidase (Opik, 1973); malate dehydrogenase (Crawford, 1978; Avadhani et al. , 1978), malic enzyme (Davies et al. , 1974; McManmon and Crawford, 1971; VanToai et al . , 1987), alcohol dehydrogenase (McManmon and Crawford, 1971; Wignarajah and Greenway, 1976; Mendelssohn et al. , 1981; VanToai et al. , 1985), andpyruvate decarboxylase, (John and Greenway, 1976; VanToai et al. , 1985; 1987) are so dramatic following waterlogging that they have been used as indicators of plant response to waterlogging stress. However, the role of these enzyme changes in response to the waterlogging stress has been debated (Hook, et al., 1971; Crawford, 1978; Smith et al.,1986; Burdick and Mendelssohn, 1990).

As the duration of hypoxia continues, the metabolic changes in the root ultimately induce distortions in the ultrastructure of the mitochondrial membranes (Vartapetian, 1982; Vartapetian et al. , 1987; Oliveira, 1977). Disruption

70of the respiration and mitochondrial integrity affect the source/sink balance of foliar carbohydrate supply and root demand. The changes in the cellular membrane status of the roots may result in a cascade of events which ultimately affects all metabolic processes in the plant. The final alteration in plants under waterlogged conditions is the formation of adventitious roots and aerenchymatous tissue, gross morphological changes which facilitate oxygen diffusion to the roots.

The purpose of these experiments was to use our selected populations of Brassica rapa L. that differ in their response to waterlogged conditions (Chapter Two) and examine the nature of the tolerant and sensitive responses by analyzing the root metabolic status during the initial hours of hypoxia. We reported differential response by the shoots of these populations as early as 12 hours into the waterlogging period (Chapter Three). In this study we examine the activities and electrophoretic profile of six enzymes, adenylate energy charge ratios, and mitochondrial ultrastructure in the roots of the three populations under rootzone hypoxia. The objective was to describe early changes in root metabolism observed among the populations.

71Material and Methods

Plant MaterialSelected individuals from the base population were

separated into "tolerant" and "sensitive" populations and mass pollinated to produce the next generation (Chapter Two) . Plants were grown according to the procedure outlined in Chapter Two. Our populations of rapid-cycling Brassica rapa CrCC01-51 (sensitive) and CrGC/1-52 (tolerant) are available from the Crucifer Genetics Cooperative, Madison WI.

Root Enzyme AnalysisRoot tissue for enzyme assays was weighed, frozen in

liquid nitrogen, and stored at -80°C until use. All extracts were prepared and maintained at 2-4°C using a modified method of McKee and Mendelssohn (1987). Root tissue was ground in 60mM Tris (TRIZMA, Sigma Chemical Co.) buffer (pH 6.8) which contained 0.015% (w/v) dithiothreitol, and 10% (w/v) of polyvinylpyrrolidone. The plant extracts were centrifuged at 26,000 g for 15 min and the enzymes in the supernatant assayed at immediately.

Assays for alcohol dehydrogenase (ADH; EC 1.1.1.1), NAD+-malate dehydrogenase (MDH; EC 1.1.1.27), malic enzyme (ME; 1.1.1.40), glucose-6 phosphate dehydrogenase (G6PDH; EC 1.1.1.49), pyruvate decarboxylase (PDC; EC 4.1.1.1) and phosphoglucomutase (PGM; EC 2.7.5.1) were performed at 2 5°C

7 2

in a final volume of l ml which contained 100 mM Tris-5 mM HgCl] (pH 7.0) buffer and reagents as follows: ADH: 25 mMNADH, 0.5 mM acetaldehyde, 20 Ml enzyme extract; HOH: 25 mM NADH, 30 mM OAA, 2 nl enzyme extract; ME: 0.125 mM NADP, 30 mM malate, 2 0 /il enzyme extract; G6PDH: 0.12 5 mM NADP, 0.3 mM G6P, 2 0 Ml enzyme extract; PDC: 2 5 mM NADH, 3 0 mMpyruvate, 40 Ml enzyme extract; PGM: 0.125 mM NADP, 0.3 mM G1P, 40 mM Gl,6diP, 12 units G6PDH, 40 m 1 enzyme extract. Enzyme activities were quantified spectrophotometrically by measuring the oxidation of NADH or the reduction of NADP at 339 nm using a Gilford (Response II) spectrophotometer.

Total soluble protein was determined for the root extracts using a modified Lowry et al., (1951) procedure.Each sample was compared against a blank containing the appropriate solutions. Total protein was calculated from a standard curve prepared with a bovine serum albumin solution (10 mg/ml) and the absorbance was determined at 650 nm.

Root Adenylate AnalysisRoot tissue for adenylate assays was prepared according

to the method of Mendelssohn and McKee (1981). Root tissue that had been waterlogged for 18 hours, or maintained under control conditions, was placed in small plastic bags containing approximately 15 ml of deionized water, and frozen in liquid nitrogen. Samples were maintained at -80°C until they could be lyophilized in a Virtis Sentry SSL

7 3

Freeze Dryer. Root tissue was ground, weighed, and then extracted in 1 mM boiling EDTA and 5% (w/v)polyvinylpolypyrrolidone solution. Adenosine mono-, di-, and triphosphates were measured using the ATP-dependent light yielding reaction of the firefly-lantern luciferin luciferase (FLE-50, Sigma Chemical Co., St. Louis) complex with a Model 3000 SAI-Technology Company Integrating Photometer. ATP was determined directly, while ADP and AMP were converted enzymatically to ATP and determined by subtraction.

Isozyme AnalysisAll starch gels (12% w/v) were prepared with

electrophoresis grade starch (Starch-Art, Smithville, TX). The electrode buffer was a LiOH-Borate (0.06 M LiOH, 0.3 M Borate pH 8.1) solution. The gel buffer consisted of a Tris-Citrate (0.03 M Tris, 0.005 M citrate, pH 8.5) solution with the addition of 1% LiOH-Borate electrophoresis buffer.

Root extract was absorbed onto Whatman filter paper wicks Grade-1 (4 mm x 10 mm) and loaded into a horizontal starch gel. All gels were run at 50 mA for approximately four hours. The following enzymes were assayed: ADH, G6PDH, MDH, ME, PGM, and PDC. Banding patterns of each enzyme were analyzed at 6, 12, 18, 24, and 48 hours after waterlogging. Isozymes of PGM and ADH were analyzed further using vertical discontinuous polyacrylamide gel electrophoresis. The two

74enzymes were analyzed on a 7% (w/v) native polyacrylamide system with 0.37 M Tris (pH 8.8) in the analyzing gel and 3.5 % (w/v) polyacrylamide with 0.1 M Tris (pH 6.8) in the stacking gel. Both reservoirs contained a modified Tris/glycine buffer (Guikema and Sherman, 1982).

Root extracts had protein concentrations ranging from 3 to 5 nq/nl and were loaded on an equal volume basis into the separate wells on the gel. A minimum of four gels were run under the same conditions for each time course. Isozymes were stained with specific stains as described by Shaw and Prasad (1970).

Mitochondrial UltrastructureRoots were obtained from control plants and plants that

were waterlogged for four days. Root sections 2 cm in length (cut from the lower third of the root excluding the root tip) were processed in the same manner as described in Chapter Three.

Data AnalysisExperimental variables and the calculated ratios were

compared by a randomized block factorial analysis (Chapter Three). Values of the experimental variables were compared by an analysis of variance (ANOVA) and only those with the significant F-values were compared by Least Significant Difference (LSD) analysis (Snedecor and Cochran, 1980).

75Results

Enzyme ActivitiesWhen plants were subjected to waterlogging stress, a

change in activity was observed in four of the six root enzymes that were measured in the populations of Brassica rapa (Table 4.1). The most dramatic changes were the increases in ADH and PDC activities after 4B hours of waterlogging. Malate dehydrogenase and phosphoglucoroutase both decreased in activity after the roots were exposed to the waterlogging stress. The levels of all six enzymes were not significantly different among the three populations under control conditions.

Table 4.1. Activities {/mol mg protein1 roin1) of six enzymes measured in Brassica rapa roots after 4 8 hours of waterlogging. Data are pooled means of the three populations based on six replications. Values with the same letter in the same row are not significantly different (P > 0.05).

Activity(//mol mg protein1 min'1)

Enzvme Reaction Control WaterloadedME pyruvate <— > malic acid 37 a 46 aMDH OAA <--> malate 2227 b 2058 aADH acetaldehyde <— > ethanol 80 a 672 bG6PDH G6P <— > D-glucono-lactone 6P 52 a 61 aPDC pyruvate <--> acetaldehyde 19 a 90 bPGM glucose-lP <— > glucose-6P 38 b 20 a

7 6

The activity levels of ADH showed a change after only six hours of waterlogging when compared to the control levels (Fig 4.1) . The CrGC#l-3 3 population was the first to show an increase in ADH activity, but the sensitive population had the roost dramatic increase in ADH activity after 18 hours of waterlogging.

There was a significant difference in ADH activities of the three populations after 12 hours of rootzone hypoxia (Table 4.2). The CrGC/1-33 population showed an increase in ADH activity, whereas the ADH activities in the tolerant and sensitive populations did not change between the 6 and 12 hour waterlogging periods. ADH activity in the sensitive population was significantly greater after 18 hours of rootzone hypoxia in comparison to the levels in the tolerant and CrGC#l-33 populations. Levels of ADH in the tolerant and CrGC/1-33 populations remained fairly constant, whereas the ADH activity in the sensitive population was approximately four times greater than the level after 12 hours of waterlogging.

There was an 18 hour lag in ADH response in the waterlogged tolerant population as compared to control levels (Fig 4.1). A 24 hour period of waterlogging was needed to induce a major increase in ADH activity in the tolerant population. After 24 hours of waterlogging stress the populations were not significantly different in ADH activity. However, after only 48 hours of waterlogging

77

O<

7a>-*->D

BOO

700CE

600

Tca>

300-*->oi—CL

400

cr>E

300

oE

200

100

0

Alcohol dehydrogenase

12 24 36

TIME (hours)

symbols

• CrCC#1-33 ■ Tolerant + Sensitive

niled-waterlogged hollow— control

Figure 4.1. Root alcohol dehydrogenase activities of populations of Brassica rapa responding to waterlogging stress. Activity data are means based on six replications. All waterlogging values were significantly different from the controls, except the tolerant population at 12 and 18 hours (P s 0 . 0 5 ) .

78stress, the activities of ADH in the waterlogged plants were approximately seven to eight times greater than the control levels.

Table 4.2. Alcohol dehydrogenase activities (jimol mg protein1 min1) measured in roots of populations of Brassica rapa after 12 and 18 hours of waterlogging. Data are means based on six replications. Values with the same letter in the same column are not significantly different (P > 0.05).

Enzyme Activity Population (Mmol mg protein1 min1)______________________________12 hr_____________19 hrTolerant 90.9 a 100.3 aSensitive 137 . 3 a 557.9 cCrGC#l-3 3 266 . 7 b 267.8 b

After 48 hours of waterlogging stress the activity of PDC increased significantly in the waterlogged roots when compared to the controls (Fig 4.3). Also, after 48 hours of waterlogging, there was a significant difference in PDC activity among the three populations. The sensitive and CrGC#l-33 populations had significantly higher levels of PDC activity when compared to the tolerant population.

79Table 4.3. Pyruvate decarboxylase activities (Mmol mg protein1 min1) measured in roots of populations of Brassica rapa after 48 hours of waterlogging. Data are means based on six replications. Values with the same letter in the same column are not significantly different (P > 0.05).

Population Enayma ActivityControl

(M*ol mg protein1 min'1)Waterloaaed

Tolerant 17.76 a 71.86 a

Sensitive 22.05 a 104.00 b

CrGC#l-3 3 18.90 a 97.11 b

Root Adenylate Energy ChargeAfter 18 hours of waterlogging stress there was no

significant difference in the ATP concentrations or adenylate energy charge (AEC) ratios caused by the waterlogging treatment (Table 4.4). The ATP concentrations in the waterlogged plants were slightly greater than the control values, but the differences were not significantly different.

80Table 4.4. Adenylate energy charge ratio ([ATP] + 0.5 [ADP] / [ ATP] + [ADP] + [AMP]) and the concentration of ATP (nmoles/g dry weight) measured in roots of populations of Brassica rapa after 18 hours of waterlogging. Data are means based on two replications. Values were not significantly different between treatment or population (P > 0.05).

populationControl

AECWaterloaoed

ATP (nmolas/g dvt)Control Waterlogged

Tolerant 0.89 0. 89 995 1020

Sensitive 0.87 0.90 930 1100

CrGC#l-3 3 0. 86 0 . 88 920 990

Isozyme PatternsThe banding patterns of malic enzyme (ME), malate

dehydrogenase (MDH), glucose 6-phosphate dehydrogenase (G6PDH), and pyruvate decarboxylase (PDC) in the waterlogging treatment were not different from the controls or within the three populations (Fig 4.2). Except malate dehydrogenase, all of the enzymes had excellent resolution.

The electrophoretic banding pattern of ADH in the three populations after 24 and 48 hours of waterlogging stress is presented in Figure 4.3. At both the 24 and 48 hours after waterlogging, two regions of activity were present when stained for ADH. Two distinct bands, present in all three populations, were noted at the 24 hour sampling (Fig 4.3A). The band at the anodal portion of the gel had a much greater staining intensity than the slower migrating band. After 48

+ ME MDH G-6PDH PDC

Figure 4.2. Composite electrophoretic banding patterns of the four enzymes extracted from Brasetlca rapa roots that did not show variation in the banding patterns among the three populations, regardless of treatment. Isozymes were analyzed on a 12% (w/v) starch gel. Malic enzyme (ME); Malatedehydrogenase (MDH); Glucose 6-phosphate dehydrogenase (G- 6PDH); Pyruvate decarboxylase (PDC).

+

CrGC#1-33 Tolerant Sensitive CrGC#1-33 Tolerant Sensitive

Figure 4.3. Electrophoretic banding pattern of alcohol dehydrogenase isozymes extracted from Brassica rapa roots exposed to waterlogged conditions for 24 hours (A) or 48 hours (B). Isozymes were analyzed on a 7% (w/v)polyacrylamide gel.

83hours of waterlogging the slower migrating ADH band was not apparent in the tolerant and sensitive populations but was observed in the CrGC/1-3 3 population. The faster migrating ADH band stained intensely in all three populations.

The electrophoretic banding pattern of phosphoglucomutase (PGM) from the roots of three populations of fi. rapa is presented in Figure 4.4. The CrGC/1-3 3 population showed four distinct regions of activity when stained for PGM, whereas the sensitive and tolerant populations had only three and two regions, respectively. The staining intensity of the PGM isozymes also was dissimilar. The fastest migrating zone was apparent in all three populations, but had low staining intensity, whereas at least two isozymes of PGM were intensely stained. The isozyme banding pattern of PGM within the three populations did not change regardless of treatment.

Mitochondrial UltrastructureTEM examination of B. rapa root cells revealed

differences in the mitochondrial ultrastructure between the waterlogged and control treatments. After four days of waterlogging the mitochondria exhibited structures as shown in Figure 4.5. Mitochondria ranged from severely distorted (Fig 4.5A and C) to mildly distorted (Fig 4.5B) in comparison to the controls (Fig 4.5D). The control mitochondria were round and oval shaped with randomly

84

CrGC#1-33 Tolerant Sensitive

Figure 4.4. Electrophoretic banding pattern ofphosphoglucomutase isozymes extracted from Brassica rapa roots. The banding pattern was identical under control or waterlogged conditions. Isozymes were analyzed on a 7% (w/v) polyacrylamide gel.

Figure 4.5. Electron micrographs of Brassica rapa cell sections from waterlogged and control roots showing mitochondrial ultrastructure. A: B: C: waterlogged roots, D: control roots. Duration of waterlogging was four days, (m) Mitochondria.

86arranged cristae (Fig 4.5D). Many of the waterlogged roots had mitochondria with electron dense matrices and cristae that were organized in a linear fashion. A few of the mitochondria in roots exposed to hypoxic conditions had irregular shapes, but were similar in cristae and matrix organization to the control mitochondria (Fig 4.5B). The mitochondrial ultrastructure in the populations did not appear to be different among the three populations.

DiscussionEnzyme Activities

For many plant species under hypoxic stress, metabolic alterations may include changes in specific enzyme activity, or increases in the concentrations and types of enzymes produced (Crawford, 1978; Davies, 1980; VanToai et al., 1987; Crawford et ai. , 1987; Mendelssohn and Burdick,1988;). One of the main alterations occurs through the increased activity of alcoholic fermentation (Wignarajah and Greenway, 1976; Mendelssohn et al. , 1981; VanToai et ai. ,1985; John and Greenway, 1976; McKee et al. , 1989). Alcohol dehydrogenase activity has been shown to increase in hypoxic roots and the extent of increase has been correlated directly with the severity of oxygen deprivation (Bertani and Brambilla, 1982). This indicates that partial alcoholic fermentation and ethanol accumulation can take place in

87roots at low oxygen concentrations as well as under totally anoxic conditions.

Crawford (1978) and his colleagues (McManmon and Crawford 1971; Crawford and Zochowski, 1984; Crawford et al. , 1987) reported that waterlogging tolerant plants differ from intolerant species by not accelerating alcoholic fermentation, but by employing an alternative metabolic pathway. These waterlogging-tolerant plants respond by producing malate and therefore do not increase the levels of ethanol in the rootzone. However several reports provide contradictory evidence to this hypothesis (Hook, 1971; John and Greenway, 197 6; ap Rees et al., 1987). Manyinvestigators reported an increase in malic enzyme activity in plants exposed to waterlogged conditions, but the malate concentrations also accumulated under aerobic conditions, or in conjunction with increases in alcohol dehydrogenase activity (Davies et al., 1974; Keeley, 1978).

The three populations of Brassica rapa exhibited an increase in ADH activity, with no change in the activity level of malic enzyme. Malic enzyme was not altered in activity or isozyme banding patterns, regardless of treatment or population (Table 4.2 and Fig 4.2). The suggestion of waterlogging-tolerance in plants by accumulation of malate rather than ethanol was not supported by these experiments.

88The populations of S. rapa responded to the

waterlogging stress by altering ADH activity, which increased more than seven fold the control levels after 48 hours (Fig 4.1). The activity level of ADH in the B . rapa roots is comparable to the activities reported for rice, Zea mays, and Vicia faba that were exposed to anaerobic conditions (John and Greenway, 1976; Hignarajah and Greenway, 1976; Thynn and Werner, 1990). It seems likely that all three populations were, at some point during the waterlogging stress, relying on the increased activity of alcoholic fermentation to provide the energy needed for cellular function. In addition, the activity of pyruvate decarboxylase also increased in the three populations, but to a lesser extent than ADH activity levels (Table 4.3). Low PDC activity levels have been reported for several species exposed to waterlogged conditions (Wignarajah and Greenway, 1976; Kimmer, 1987). Activities of PDC were much lower than those of ADH, favoring control of alcoholic fermentation via control of PDC, which prevents accumulation of phytotoxic acetaldehyde (John and Greenway, 1976). Pyruvate decarboxylase has been suggested to be the regulatory step in ethanol biosynthesis (Davies, 1980).

Root Adenylate Energy ChargeGiven the differences in ADH activity among the three

populations by 18 hours, we investigated whether population

89differences in root metabolism would be reflected in energy charge. Adenylate energy charge ratios and ATPconcentrations in the roots were not significantly different in the three populations (Table 4.4). The ATP levels, although similar in the three populations, may have been produced from different metabolic pathways within the cell. Alcohol dehydrogenase activity was relatively high in the sensitive population early in the waterlogging treatment, which may indicate increased alcoholic fermentation. Increases in ADH activity corresponding with high adenylate energy charge ratios were reported in rice exposed to hypoxic conditions (Mocquot et al. , 1981; Rumpho et al. ,1984) . The adenylate levels in the sensitive population may have been produced from the acceleration of the glycolytic cycle, whereas the tolerant population was able to maintain ATP concentrations without an increase in glycolytic activity. The tolerant population may have been able to maintain normal cellular functions through regular respiration or pathways that lie outside fermentation, such as the pentose phosphate shunt.

Isozyme PatternsThe banding patterns of the six enzymes were evaluated

to determine if the differential tolerance to rootzone hypoxia in the three populations of 5. rapa was influenced by alterations in isozyme patterns (Fig 4.2). Two enzymes,

90alcohol dehydrogenase and phosphoglucomutase, had electrophoretic profiles that showed variations among the populations (Figs 4.3 and 4.4). The alteration in PGM isozyme banding patterns in the populations of B. rapa did not appear to affect overall enzyme activity, which decreased under waterlogging stress. The difference in isozyme banding patterns may have been one of several alterations that occurred during the selection processes of the two populations of S. rapa (Chapter Two) because the banding pattern is identical under waterlogged and control conditions.

Mutants of maize (Roberts et al. , 1984) and barley(Harberd and Edwards, 1982) which lacked the major isozyme of alcohol dehydrogenase were markedly less capable of withstanding hypoxic conditions than those with the normal complement of enzyme. The sensitive population did not appear to be lacking in a major ADH isozyme. Therefore the sensitivity of this population was probably not caused by a lack of a particular ADH isozyme.

The isozyme banding pattern of alcohol dehydrogenase was similar in the three populations, except for the slower migrating band. This band which had a low staining intensity was lacking in both the tolerant and sensitive populations. After 48 hours of waterlogging, the tolerant and sensitive populations may have produced only one ADH

91isozyme for use in alcoholic fermentation rather than two isozymes as was apparent in the CrCG/1-33 population.

The fastest migrating ADH isozyme increased in staining intensity from the 24 to 48 hour time period in all three populations. This increased staining intensity also corresponded to the increased alcohol dehydrogenase activity. Staining intensity and subsequent activity of the isozymes may have resulted from the specific activity of the enzyme or a greater quantity of total enzyme.

Mitochondrial ultrastructureMany reports have examined mitochondrial ultrastructure

when roots have been exposed to sudden anoxic conditions, resulting from purging the system with nitrogen (Opik, 1973; Oliveira, 1977; Vartapetian et al., 1987; Vartapetian,1991). Vartapetian and colleagues reported a variety of mitochondrial morphologies in roots exposed to anoxic conditions and in roots which were maintained under anoxic conditions supplemented with 3% glucose in the medium (Vartapetian et al., 1977; 1987; Vartapetian and Andreeva,1986). Mitochondria from completely anoxic conditions increased in size, contained fewer cristae, and had less electron-dense matrices. However, the mitochondria in the anoxic treatment supplemented with glucose were oval shaped, had cristae arranged in a parallel order and were electron dense (Vartapetian and Andreeva, 1986).

92The mitochondria in the waterlogged B . rapa roots were

exposed to a gradual decline in oxygen availability as opposed to sudden rootzone anoxia when purged with nitrogen. After four days of hypoxic treatment, the mitochondria exhibited features similar to those exposed to the 24 hour anoxia supplemented with glucose treatment (Fig 4.5). Mitochondria of B. rapa exhibited elongated and branched structure similar to the ultrastructural arrangement observed in the roots exposed to anaerobic conditions (Oliveira, 1977; Vartapetian, 1982). Elongation has been proposed to be an adaptive rearrangement of the mitochondria under conditions of hypoxic stress. Branched mitochondria would ensure closer contact of the mitochondrial membranes with the cytoplasm and the endoplasmic reticulum, thus aiding in rapid transport of ATP along the membranes to protein synthesis sites on the endoplasmic reticulum (Vartapetian et al. , 1977). Mitochondria from B . rapa roots grown under hypoxic conditions varied greatly in their appearance and were not distinctly different among the populations.

Root enzyme activities, energy charge, and ultrastructure, were considered to be indicators of rather than causes of differences in the response of populations of Brassica rapa L. to rootzone hypoxia. None of the populations had complete inhibition of a metabolic process, suggesting that the lack of a particular enzyme was not the

93determining factor for the cause of the differential response observed under rootzone hypoxia.

In a previous paper we reported on the leaf carbohydrate accumulation and partitioning in the three populations in response to waterlogging stress (Chapter Three). Leaf carbohydrate concentrations increased very quickly in the sensitive population under hypoxic stress, whereas carbohydrate accumulation was not as great in the tolerant population. From these studies it is apparent that the tolerant population had active root metabolism and was able to utilize the foliar carbohydrates. The roots could obtain the needed ATP without increasing the glycolytic cycle at least during the first 24 hours of waterlogging. The maintenance of the normal metabolic pathways, even for a short duration, may allow the plant to begin initiation processes for gross morphological changes (adventitious roots, aerenchymatous tissue) that would aid in oxygen diffusion to the roots.

Our results indicate that the initial hours of waterlogging are the most critical in terms of identifying waterlogging tolerant plants, because this is when metabolic processes must be altered in order to ensure survival. Waterlogging tolerant plants may use a combination of root and foliar metabolic adaptations to withstand the effects of short-term hypoxic conditions. A closer examination of the metabolic relationship between the foliage and the roots,

94rather than identification of a particular root metabolic process as the determining factor, may provide additional information to determine what constitutes a waterlogging tolerant species.

SummaryRoots exposed to waterlogging stress may develop

altered metabolism and morphology. Changes in the glycolytic pathway, enzyme activities and forms, and mitochondrial ultrastructure may help maintain the plant through the initial period of hypoxic stress. The root metabolic status of three selected populations of Brassica rapa L. that differ in their response to rootzone hypoxia was analyzed. Activities and electrophoretic profile of six enzymes: alcohol dehydrogenase (ADH), pyruvate decarboxylase (PDC), malate dehydrogenase (MDH), malic enzyme (ME), glucose 6-phosphate dehydrogenase (G6PDH), and phosphoglucomutase (PGM) were examined. Under waterlogged conditions, activities of ADH and PDC increased, PGM and MDH decreased, and ME and G6PDH did not change. After 18 hours of waterlogging, activity levels of ADH were significantly different in the three populations. The isozyme patterns of ADH and PGM also were different among the populations. No significant difference in the adenylate energy charge or ATP concentrations were observed after 18 hours of waterlogging. Mitochondrial ultrastructure in the roots of the three

95populations under rootzone hypoxia revealed swollen cristae and electron dense matrices. Enzyme activities, energy charge, and ultrastructure, were considered to be indicators of rather than causes of differences in population responses to rootzone hypoxia.

96Literature Cited

ap Rees T, Jenkin LET, Smith AM, Wilson PM (1987) The metabolism of flood-tolerant plants. Jn RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford, pp 227-23B

Avadhani PN, Greenway H, Lefroy R, Prior L (1978) Alcoholic fermentation and malate metabolism in rice germinating at low oxygen concentrations. Aust J Plant Physiol 5: 15-25

Bertani A, Brambilla I (1982) Effect of decreasing oxygen concentration on wheat roots: growth and induction of anaerobic metabolism. Z Pflanzenphysiol 108: 283-288

Burdick DM, Mendelssohn IA (1990) Relationship between anatomical and metabolic responses to soil waterlogging in the coastal grass Spartina patens. J Exp Bot 41: 223-228

Crawford RMM (1978) Metabolic adaptation to anoxia. Jn DD Hook, RMM Crawford, eds, Plant Life in Anaerobic Environments. Ann Arbor Science, MI, pp 119-155

Crawford RMM, Monk LS, Zochowski ZM (1987) Enhancement of anoxia tolerance by removal of volatile products of anaerobiosis. Jn RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford, pp 375-384

Crawford RMM, Zochowski ZM (1984) Tolerance of anoxia and ethanol toxicity in chickpea seedlings (Cicer arietinum L.). J Exp Bot 35: 1472-1480

Davies DD (1980) Anaerobic metabolism and the production of organic acids. Jn DD Davies, ed, The Biochemistry of Plants. Academic Press, New York, pp 581-611

Davies DD, Nascimento KH, Patil KD (1974) The distribution and properties of NADP malic enzyme in flowering plants. Phytochem 13: 2417-2425

Guikema ja, Sherman LA (1982) Protein composition and architecture of the photosynthetic membranes from the cyanobacterium, Anacystis nidulans R2. Biochem Biophys Acta 681: 440-450

Harberd NP, Edwards KJR (1982) The effect of a mutation causing alcohol dehydrogenase deficiency on flooding tolerance in barley. New Phytol 90: 631-644

97Hook. DD, Brown CL, Kormanik PP (1971) Inductive flood tolerance in swamp tupelo (Nyssa sylvatica var. biflora [Walt.] Sarg.). J Exp Bot 22: 78-89

John CD, Greenway H (1976) Alcoholic fermentation and activity of some enzymes in rice roots under anaerobiosis. Aust J Plant Physiol 3: 325-336

Keeley JE (1978) Malic acid accumulation in roots in response to flooding: Evidence contrary to its role as an alternative to ethanol. J Exp Bot 29: 1345-1349

Kimmer TW (1987) Alcohol dehydrogenase and pyruvate decarboxylase activity in leaves and roots of eastern cottonwood (Populus deltoides Bartr.) and soybean (Glycine max L.). Plant Physiol 84: 1210-1213

Lowry OH, Rosebrough NJ, Farr AL, Randall RJ (1951) Protein measurement with the folin phenol reagent. J Biol Chem 193: 265-275

McKee KL, Mendelssohn IA (1987) Root metabolism in the black mangrove (Avicennia germinans (L.) L): response to hypoxia. Env Exp Bot 27: 147-156

McKee KL, Mendelssohn IA, Burdick DM (1989) Effect of long­term flooding on root metabolic response in five freshwater marsh plant species. Can J Bot 67: 3446-3452

McManmon M, Crawford RMM (1971) A metabolic theory of flooding tolerance: The significance of enzyme distribution and behavior. New Phytol 70: 299-306

Mendelssohn IA, Burdick DM (1988) The relationship of soil parameters and root metabolism to primary production in periodically inundated soils. Jn DD Hook, ed, The Ecology and Management of Wetlands Vol. 1: Ecology of Wetlands. Croom Helm Ltd, United Kingdom pp 398-42 5

Mendelssohn IA, McKee KL (1981) Determination of adenine nucleotide levels and adenylate energy charge ratio in two Spartina species. Aguat Bot 11: 37-55

Mendelssohn IA, McKee KL, Patrick WH Jr (1981) Oxygen deficiency in Spartina alterniflora roots: Metabolic adaptation to anoxia. Science 214: 439-441

Mocquot BI, Mouches C, Pradet A (1981) Effect of anoxia on energy charge and protein synthesis in rice embryo. Plant Physiol 68: 636-640

98Oliveira L (1977) Changes in the ultrastructure of mitochondria of roots of Triticale subjected to anaerobiosis. Protoplasms 91: 267-280

Opik H (1973) Effect of anaerobiosis on respiratory rate, cytochrome oxidase activity and coleoptiles of rice (Oryza satlva L.). J Cell Sci 12: 725-739

Pradet A, Bomsel JL (1978) Energy metabolism in plants under hypoxia and anoxia. Jn DD Hook, RMM Crawford, eds, Plant Life in Anaerobic Environments. Ann Arbor Science, Ann Harbor, pp 89-118

Roberts JKM, Callis J, Wemmer D, Walbot V, Jardetzy O (1984) Mechanism of cytoplasmic pH regulation in hypoxic maize root tips and its role in survival under hypoxia. Proc Nat Acad sci 81: 3379-3383

Rumpho ME, Pradet A, Khalik A, Kennedy RA (1984) Energy charge and emergence of the coleoptile and radicle at varying oxygen levels in Echinochloa crus-galli. Physiol Plant 62: 133-138

Shaw CR, Prasad R (1970) Starch gel electrophoresis of enzymes-A compilation of recipes. Biochem Gen 4: 297-320

Smith AM, Hylton CM, Koch L, Woolhouse HW (1986) Alcohol dehydrogenase activity in the roots of marsh plants in naturally waterlogged soils. Planta 168: 130-138

Snedecor GW, Cochran WG (1980) Statistical methods 7th ed. Iowa State University Press, Ames, pp 593

Thynn M, Werner D (1990) A more rapid increase of alcohol dehydrogenase activity in seedling roots of Vicia faba L. by addition of ethylene compared to anaerobiosis. Angew Botanik 64: 123-131

VanToai TT, Fausey NR, McDonald MB Jr (1985) Alcohol dehydrogenase and pyruvate decarboxylase activities in flood-tolerant and susceptible corn seeds during flooding. Agron J 77: 753-757

VanToai TT, Fausey NR, McDonald MB Jr (1987) Anaerobic metabolism enzymes as markers of flooding stress in maize seeds. Plant Soil 102: 33-39

Vartapetian BB (1982) Anaerobiosis and the theory of physiological adaptation of plant to flooding. Soviet Plant Physiol 29: 764-771

99Vartapetian BB (1991) Flood-sensitive plants under primary and secondary anoxia: ultrastructure and metabolic responses. In MB Jackson, DD Davies, H Lambers eds, Plant Life Under Oxygen Deprivation. SPB Academic Publishing, The Hague, pp 201-216

Vartapetian BB, Andreeva IN (1986) Mitochondrial ultrastructure of three hydrophyte species at anoxia and in anoxic glucose-supplemented medium. J Exp Bot 37: 685- 692

Vartapetian BB, Andreeva IN, Kozlova GI, Agapova LP (197"M Mitochondrial ultrastructure in roots of mesophyte and hydrophyte at anoxia and after glucose feeding. Protoplasma 91: 243-256

Vartapetian BB, Snkhchian HH, Generozova IP (1987) Mitochondrial fine structure in imbibing seeds and seedlings of Zea mays L. under anoxia. In RMM Crawford, ed, Plant Life in Aquatic and Amphibious Habitats. Blackwell Scientific Publications, Oxford pp 205-223

Wignarajah K, Greenway H (1976) Effect of anaerobiosis on activities of alcohol dehydrogenase and pyruvate decarboxylase in roots of Zea mays. New Phytol 77: 575-584

Williams PH (1985) Crucifer Genetics Cooperative Resource Book. Department of Plant Pathology, University of Wisconsin-Madison, WI, pp. 100

CHAPTER FIVE

CONCLUSIONS AND PERSPECTIVES

1 0 0

1 0 1

The purpose of this chapter is to present the conclusions of this work in relation to our knowledge of the physiological and biochemical mechanisms of waterlogging tolerance.

One of the key features of this waterlogging study involved mimicking the stress that might occur in plants exposed to a natural waterlogging event. The Brassica rapa plants were grown in a soil-type matrix which was then waterlogged causing a slow decline in oxygen availability to the roots. In contrast, many studies have utilized nitrogen purging of the root environment to suddenly reduce oxygen concentration (Oliveira, 1977; Roberts et al., 1989; Sachset al., 1980; Vartapetian, 1991). In a natural waterlogged environment it is very unlikely that complete and immediate removal of oxygen in the rootzone will occur. An experimental system that is aerobic and then completely anaerobic within several minutes may produce artificial responses in the plants that may not occur under natural soil waterlogging. Our studies mimicked hypoxic conditions in the rootzone and probably induced responses in the plants that realistically occur under natural waterlogged conditions.

By using mass pollination and recurrent selection we were able to successfully select populations of rapid- cycling Brassica rapa that differed in their response to rootzone hypoxia. The development of populations differing

1 0 2

in waterlogging stress supports the rapid selection of tolerance to severe environmental stress found by others (Antonovics et al., 1971; Gillespie and winner, 1989). The different populations of Brassica rapa demonstrate that stress-specific differences in population performance can be achieved through recurrent selection.

Our waterlogging study is unique in creating a realistic hypoxic environment in the rootzone, using three different populations of Brassica rapa, and evaluating many characteristics in the entire plant over an extended period of waterlogging. Relatively few studies have evaluated the whole plant status over an extended duration of waterlogging stress and these studies investigated responses which occurred at a longer interval from the initiation of the waterlogging treatment (Trought and Drew, 1980; Talbot et al., 1987). In our study we have investigated thecombination of foliar and root responses during the initial hours of waterlogging. By assaying foliar and root responses at six hour intervals in different populations, we were able to provide detailed information on early manifestations of waterlogging tolerance.

Responses of the populations of rapid-cycling Brassica rapa to a hypoxic environment is not likely to be caused by a single cellular or morphological characteristic. Crawford et al. (1987) suggested that waterlogging tolerant plantsdiffer from intolerant species by not accelerating alcoholic

103fermentation, but respond by shunting carbohydrates into nontoxic organic acids, specifically malate. our results showed no changes in the activities or isozyme patterns of malate dehydrogenase or malic enzyme, two key enzymes involved in the pathways described by Crawford et al., (1987) . The suggestion of waterlogging tolerance in plants by accumulation of malate rather than ethanol was not supported by these experiments.

Enzyme activities, energy charge, carbohydrate status, and ultrastructure differences were considered to be indicators of rather than causes of differences in population responses to rootzone hypoxia. All populations responded very quickly to the decreasing levels of oxygen in the rootzone. After only 12 hours of waterlogging several metabolic processes throughout the plant were accelerated. Even though the redox value of the soil did not indicate hypoxia at 12 hours, the oxygen levels in the roots may have been low enough to trigger activation of these metabolic pathways.

All three populations had slightly elevated ADH activities in comparison to the controls six hours after soil waterlogging. However, the sensitive population had a dramatic increase in ADH activity 18 hours after waterlogging, whereas the tolerant and the CrGC/l-33 populations did not show this jump in activity until approximately 48 hours after waterlogging. Even though the

104sensitive population had the initial increase in ADH activity, apparently this did not result in a favorable overall response to the hypoxic conditions. The slightest decrease in rootzone oxygen triggered an increase in ADH and probably activation of alcoholic fermentation. Alternatively, the delay in ADH activation in the tolerant population may indicate that this population can rely on the normal respiratory pathways until oxygen is severely limited in the roots.

The tolerant population may have avoided or delayed activating alcoholic fermentation during the initial hours of rootzone hypoxia. Maintaining regular metabolicfunctions during the first hours of waterlogging may allow the roots to initiate development of adventitious roots and/or aerenchyma that would ultimately facilitate oxygen diffusion to the roots. As more oxygen is delivered to the root cells, normal respiratory processes could be further utilized and therefore rely less on the inefficient glycolytic cycle for the energy needs.

One of the interesting points presented in Figure 5.1 is the response of foliar soluble carbohydrates in relation to alcohol dehydrogenase activities. In all threepopulations leaf soluble carbohydrates accumulated prior to increased levels of ADH activity. The increases in foliar soluble carbohydrates within 12 hours after waterlogging indicate that the foliage was responding to the lack of

105Figure 5.1. Physiological response timeline of three populations of Brassica rapa in response to rootzone hypoxia. Direction and length of arrows indicate an increase or decrease of the response in comparison to the control values. (SC) leaf soluble carbohydrates, (St) leaf starch, (ADH) alcohol dehydrogenase, (PDC) pyruvate decarboxylase, (DW) dry weight.

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107oxygen within a similar time frame as the roots in the sensitive population.

Carbohydrate accumulation was significantly greater in the sensitive as compared to the tolerant population during the initial hours of hypoxia. One possibility was a feedback signal from the waterlogged roots. If normal root metabolic functions decreased, fewer carbohydrates would be needed in the roots and therefore an accumulation may occur in the leaves. Was there a "signal" transported from the roots to the foliage which indicated metabolic stress, and subsequently were carbohydrates not transported to the roots? Possibly product accumulation or lack of substrates were regulating key enzymes in the metabolic pathways ultimately affecting photosynthesis regulation. However, the foliar carbohydrate response was initiated prior to full activation of ADH. Soluble carbohydrates may not have been readily transported to the roots of the sensitive population during the initial hours of waterlogging. Obviously lack of oxygen is detrimental, but the lack of carbohydrates for catabolism would be just as critical.

Many reports have demonstrated that carbohydrates accumulate in the foliage of waterlogged plants, possibly because of reduced utilization by the roots (Siji and Swanson, 1973; Setter, et al. , 1987; Wample and Davis, 1983; Barta, 1988; Musgrave et al. , 1991). Employing alcoholicfermentation for the cell's energy needs requires more

108carbohydrates than aerobic respiratory pathways. However, if there was a decrease in growth and only the critical cellular functions were maintained, fewer carbohydrates would be utilized. The waterlogged roots in the tolerant population were evidently able to maintain cellular activities and continue growth as indicated by root dry weights. The sensitive population could only maintain critical cellular functions and was not able to implement new growth. The roots in the tolerant population continued to be a sink for carbohydrates, and subsequently the foliar carbohydrate accumulation was delayed.

The dry weight accumulation in the tolerant population in comparison to the CrGC#l-33 population was very different, and yet early metabolic responses are similar in the two populations. The metabolic responses of the foliage in the tolerant and CrGC#l-33 populations are similar in timing but not magnitude. These responses may also indicate that the shoot or the root/shoot relationship is the factor determining the differences observed in the three populations.

The sensitive population accumulated carbohydrates in the foliage, may have had a carbohydrate deficiency in the roots, and was activating an inefficient metabolic pathway. The population could not effectively withstand the waterlogging stress. The metabolic responses in the sensitive population were unable to maintain the plant as

109indicated by the decreased dry weights. The tolerant population did show increases in soluble carbohydrates and ADH activity, but this was not as detrimental to the plantas indicated by our data on the minimal loss in dry weights.A closer examination of the foliar metabolic status, specifically enzymes that regulate carbohydratepartitioning, and their relationship to root metabolism is needed for plants exposed to rootzone hypoxia.

From these studies it is apparent that plants respond very quickly to hypoxic conditions in the roots. The metabolic changes in the rapid-cycling B. rapa were taking place within 12-24 hours after waterlogging. These metabolic changes ultimately affect the long term performance of a plant under waterlogged conditions.Tolerance under hypoxic conditions is probably not determined by a single metabolic factor, but rather the combination of several responses.

However, there may be a trait in the foliage or roots which was not investigated and led to the differential responses observed in the populations. Foliar enzymes and their activities may have influenced the carbohydrate status of the populations under waterlogged conditions. In the root system, physical differences in morphology could have aided in maintaining oxygen to the cells during the initial hours of waterlogging. Obviously, many characters could have been altered to create the differential responses

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observed in the populations of 3. rapa. Waterlogging tolerant plants nay use a combination of root and foliar metabolic adaptations to withstand the effects of short-term hypoxic conditions. The populations of rapid-cycling Brask "a rapa L. could be used as a tool for further intensive and detailed studies of the adaptive responses to hypoxic stress in the rootzone.

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VITAChristine Jo Daugherty was born on April 28, 1964 in

Glenwood, Iowa. Christine completed her undergraduate study in biology at Central College in Pella, Iowa. After receiving her B.A. degree in 1986, she entered Iowa State University where she received her M.S. degree in Plant Pathology under the direction of Dr. Charlie Martinson. She continued her graduate studies by joining the Department of Plant Pathology & Crop Physiology in 1988 and is currently under the direction of Dr. Mary Musgrave. Christine has received several distinguished awards during her graduate study at L.S.U. She was the recipient of a Sigma Xi Research grant, recognized for two Outstanding Graduate Student paper presentations (by the American Society for Space & Gravitational Biology in 1989, and by the American Society of Plant Physiologists in 1992), and was recently selected as the C.w. Edgerton Outstanding Graduate Student Award recipient. She is now a candidate for the Doctor of Philosophy degree in the Interdepartmental Plant Physiology Program.

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DOCTORAL EXAMINATION AND DISSERTATION REPORT

C a n d i d a t e : Christine Jo Daugherty

M a j o r F i e l d : Plant Health

T i t l e o f D i a a e r t a t i o n :

Selection and characterization of populations of rapid-cycling Hr ass lea rapa I„. differing in their response to rootzone hypoxia.

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D a t e o f B x a m i n a t i o n :

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