Iowa State University Capstones, Theses and Retrospective Theses and Dissertations Dissertations
1983 The effects of osmotically-induced water stress and stripe smut (Ustilago striiformis) or flag smut (Urocystis agropyri) on Kentucky bluegrass (Poa pratensis 'Merion') Jeffery Lynn Nus Iowa State University
Follow this and additional works at: https://lib.dr.iastate.edu/rtd Part of the Agricultural Science Commons, Agriculture Commons, and the Plant Biology Commons
Recommended Citation Nus, Jeffery Lynn, "The effects of osmotically-induced water stress and stripe smut (Ustilago striiformis) or flag smut (Urocystis agropyri) on Kentucky bluegrass (Poa pratensis 'Merion') " (1983). Retrospective Theses and Dissertations. 8950. https://lib.dr.iastate.edu/rtd/8950
This Dissertation is brought to you for free and open access by the Iowa State University Capstones, Theses and Dissertations at Iowa State University Digital Repository. It has been accepted for inclusion in Retrospective Theses and Dissertations by an authorized administrator of Iowa State University Digital Repository. For more information, please contact [email protected]. INFORMATION TO USERS
This reproduction was made from a copy of a document sent to us for microfilming. While the most advanced technology has been used to photograph and reproduce this document, the quality of the reproduction is heavily dependent upon the quality of the material submitted.
The following explanation of techniques is provided to help clarify markings or notations which may appear on this reproduction.
1.The sign or "target" for pages apparently lacking from the document photographed is "Missing Page(s)". If it was possible to obtain the missing page(s) or section, they are spliced into the film along with adjacent pages. This may have necessitated cutting through an image and duplicating adjacent pages to assure complete continuity.
2. When an image on the film is obliterated with a round black mark, it is an indication of either blurred copy because of movement during exposure, duplicate copy, or copyrighted materials that should not have been filmed. For blurred pages, a good image of the page can be found in the adjacent frame. If copyrighted materials were deleted, a target note will appear listing the pages in the adjacent frame.
3. When a map, drawing or chart, etc., is part of the material being photographed, a definite method of "sectioning" the material has been followed. It is customary to begin filming at the upper left hand comer of a large sheet and to continue from left to right in equal sections with small overlaps. If necessary, sectioning is continued again-beginning below the first row and continuing on until complete.
4. For illustrations that cannot be satisfactorily reproduced by xerographic means, photographic prints can be purchased at additional cost and inserted into your xerographic copy. These prints are available upon request from the Dissertations Customer Services Department.
5. Some pages in any document may have indistinct print. In all cases the best available copy has been filmed.
UniversiV Micr^lms International 300 N. Zeeb Road Ann Arbor, Ml 48106
8407114
Nus, Jeffery Lynn
THE EFFECTS OF OSMOTICALLY-INDUCED WATER STRESS AND STRIPE SMUT (USTILAGO STRIIFORMIS) OR FLAG SMUT (UROCYSTIS AGROPYRI) ON KENTUCKY BLUEGRASS (POA PRATENSIS 'MERION')
Iowa State University PH.D. 1983
University Microfilms Intsrnâtional 300 N. zeeb Road, Ann Arbor, Ml 48106
The effects of osmotically-induced water stress and stripe smut (Ustilago striiformis) or flag smut (Urocvstis agropvri) on Kentucky bluegrass (Poa pratensis 'Merion*)
by
Jeffery Lynn Nus
A Dissertation Submitted to the Graduate Faculty in Partial Fulfillment of the Requirements for the Degree of DOCTOR OF PHILOSOPHY
Major: Horticulture Approved:
Signature was redacted for privacy. In Charge^f Majef Wori
Signature was redacted for privacy. For the Major Department
Signature was redacted for privacy. For the Graduate College
Iowa State University Ames, Iowa 1983 il
TABLE OP CONTENTS Page I. INTRODUCTION 1 II. REVIEW OF LITERATURE 6 A. Water Deficits 6 1. Application 6 2. Measurement 9 a. Water content 9 b. Water potential 10 3. Effects on plants 19 a. Growth 19 b. Water relations 24
G. Water loss 25 d. Osmotic adjustment and cell wall 32 elasticity B. Disease 38 1. Effects on plants 38 a. Growth 38 b. Water economy 39 2. Leaf smuts 41 III. GROWTH OF WATER-STRESSED POA PRATENSIS INFECTED 44 BY UROCYSTIS AGROPYRI OR USTILAGO STRIIFORMIS A. Abstract 44 B. Introduction 45 C. Materials and Methods 46 1. Nutrient solution studies 48 2. Water stress studies 49 iii
Page D. Results 50 1. Nutrient solution effects 50 2, Water stress effects 53 E. Discussion 53 IV. EFFECT OF WATER STRESS AND SYSTEMIC INFECTION 59 BY USTILAGO STRIIFORMIS OR UROCYSTIS AGROPYRI ON THE GROWTH OF TILLERS AND RHIZOMES OF POA PRATENSIS A. Abstract 59 B. Introduction 60 C. Materials and Methods 6l 1. Nutrient solution studies 62 2. Water stress studies 63 D. Results 64 1. Nutrient solution effects 64 2. Water stress effects 67 E. Discussion 70 V. EFFECT OF WATER STRESS AND INFECTION BY USTILAGO 72 STRIIFORMIS OR UROCYSTIS AGROPYRI ON LEAF TURGOR AND WATER POTENTIALS OF POA PRATENSIS At Abstract 72 B, Introduction 73 C. Materials and Methods 74 D. Results 78 E, Discussion 87
VI. COMPARATIVE WATER USE BY HEALTHY AND USTILAGO 91 STRIIFORMIS-INFECTED POA PRATENSIS iv
Page A. Abstract 91 B. Introduction 91 C. Materials and Methods 93 1. Water use and efficiency 94 2. Leaf diffusive resistance 95 D. Results 97 E. Discussion 100 VII. CONCLUSIONS 108 VIII. LITERATURE CITED HO IX. ACKNOWLEDGMENTS 1^5 1
I. INTRODUCTION
Water is an essential component of plant life and comprises approximately 85 to ^Ofo of the total fresh weight in physiologically active herbaceous plants. Many physiological functions of the plant are impaired when the water content falls much below this level (Turner and Kramer, 1980). Water shortage is more limiting to crop production in the world than any other single factor (Kozlowski, 1968a). Over one-third of the world's cropland receives less than 30 inches of precipitation per year and crop yields are periodically reduced by drought (Townley-Smith and Kurd, 1979)» Drought is a major limiting factor of yields even in tropical regions (Wein et al., 1979). Kramer (I98O) claims worldwide losses in yield from water stress probably ex ceed the losses from all other causes combined. It is important to differentiate between the terms drought and water stress to avoid confusion when studying the effects of water deficits on plants. Drought is a meterological event defined as the absence of rainfall for a period of time long enough to cause depletion of soil moisture and damage to plants (May and Milthorpe, I962). The length of time without rain that is necessary to cause injury depends on the kind of plants, the water holding characteristics of the soil in which they are growing, and 2
the atmospheric conditions that affect the rate of evaporation and transpiration (Kramer, I98O). Levitt (1972) describes a "biological stress as "any environmental factor capable of inducing a potentially injurious strain in living organisms". Crafts (1939) suggested that the terra water stress be defined as the state a plant enters when the water potential crosses the zero mark and becomes negative. Although plant water stress always accompanies drought, it may occur in the absence of drought, either because of excessive transpiration or be cause water absorption is inhibited by cold soil, an excess of salt in the soil solution, deficient aeration, or injury to root systems (Turner and Kramer, 1980). The degree of dehydration without permanent injury varies widely, depending on the process under consideration, the stage of development, the duration of stress, and the kind of plant (Hsiao, 1973î Begg and Turner, 1976). Because of the overwhelming importance of water for plant growth and development, several books (Kozlowski 1978; 1976a; 1972; 1968a; 1968b; 1964; Kramer 1969? 19^9; Lange et al., 1976; Levitt, 1972; Maximov, 1929î Mussell and Staples, 1979; Paleg and Aspinall, 1982; Rutter and Whitehead, I963; Slatyer, 1967; Slavik, 1974; Turner and Kramer, I98O) and reviews (Boyer and McPherson, 1975» Gates, 1964; Henckel, 1964; Hsiao, 1973; Ilgin, 1957; Kramer, I963; Macklon and Weatherley, 1965; Shaw and Laing, I966; Slatyer, 1957; 3
Vaadia et al., 19^1) have been written. Mechanisms for drought resistance have been reviewed by Ilgin (1957), Levitt (1972), and Paleg and Aspinall (1982). Plant disease is another limiting factor in crop production in addition to shortages of water. Disease is a cause of lowered efficiency or final breakdown in the plants growth and function (Wood, 1953)» The relationship between plant disease and environment is complex. Environ mental conditions may predispose potential hosts to invasion by pathogens (Schoeneweiss, 1975b, 1978), affect the growth, distribution, and survival of pathogens (Cook, 1973î Griffin, 1978; Cook and Duniway, I98I), and modify the expression of the disease syndrome (Talboys, I968). Temperature and moisture are the limiting factors for most plant diseases (Miller, 1953)» and either can be decisive in the initiation, development, and spread of disease. The disease becomes serious if both are constantly favor- able. Symptoms of plant disease may include disease- induced water deficits in the host, i.e., the drought syndrome (Talboys, 1968). Disease-induced host water deficits may arise in several ways. Vascular wilt pathogens induce water deficits by producing vascular occlusions, toxic metabolites, host degradation products, or host reaction mechanisms (Dimond, 1970). Disease-induced water deficits also may result from loss of membrane integrity k
from the action of toxins or enzymes, or the disruption of host hormonal "balance (Ayres, 1978; Talboys, I968). Pathogen-induced reduction in root growth and subsequent decrease in host root-shoot ratio may lead to host water deficits (Murphy, 1935; Sever, 1937). Finally, dramatic increases in water loss may accompany infection. Some pathogens inhibit stomatal function directly (Farrell et al., 1969; Cruikshank and Rider, I96I; Ayres, 1972). Dramatic increases in non-stomatal water loss also may accompany epidermal rupturing caused by sporulation of rust and smut pathogens (Johnston and Miller, 1934; 1940; Ayres, 1978). Ustilago striiformis and Urocvstis agropvri are the causal organisms for stripe smut and flag smut, respectively. They are serious pathogens in several grass species in cluding Poa pratensis (Couch, 1971)• There are no cultivars of P. pratensis that are very resistant to either pathogen (Kreitlow and Juska, 1959)- All P. pratensis cultivars are to some degree tolerant (Wingard, 1953) of U. striiformis and U. agropvri. Under conditions of plentiful moisture and cool temperatures, these pathogens grow systemically throughout the aerial parts of the host (Hodges, 1970b; Hodges and Britton, 1970) and infected tillers and rhizomes are produced from infected plants (Hodges and Britton, I969)* Under conditions of limited moisture or high temperatures 5
(Hodges, 1970c; Kreitlow, 19^3)i however, infected plants exhibit greater mortality rates than healthy plants. 'Merion' P. pratensis exhibits severe damage from in fection by either organism (Kreitlow and Juska, 1959)» This research was initiated to investigate the effects of systemic infection by Ustilago striiformis and Urocystis agropvri on total plant growth, leaf and root growth, growth of tillers and rhizomes, water relations, water use ef ficiency, osmotic adjustment, and cell wall elasticity of 'Merion* P. pratensis under conditions of increasing os- motically-induced water stress. 6
II. REVIEW OF LITERATURE
A. Water Déficits
1. Application The most common method of investigating the effects of drought on plants is to withhold irrigation until plants wilt or until a predetermined soil moisture content is reached (Duniway, 1977; Edmunds, 1964; Moore et al., 1963; Couch and Bloom, I96O; Ghaffar and Erwin, I969; Hobbs et al., 1963; Cykler, 1946). Often these drying cycles range from field capacity to permanent wilting point, or a per centage of field capacity (Cowan, 1965; Ashton, 1956). Field capacity is the water content of a soil profile when, after it has been thoroughly wetted, the rate of drainage from the profile is .negligible '(Baver et al., 1972). Water potentials of a uniform soil profile at field capacity ap proach -0.1 bars for sandy soils and -0.3 bars for loams and silt loams (Papendick and Campbell, I98I). Permanent wilting point is the soil water content at the time leaves of indicator plants become wilted and do not recover turgor overnight (Papendick and Campbell, 1981). Permanent wilting point corresponds to a soil water potential of -15 bars (Crafts, 1968). Problems encountered in controlling soil moisture within field capacity to permanent wilting per centage were summarized by Couch et al. (I967). 7
Interpretation of water deficits based on alternate wetting and drying of soil is subject to criticism since such regular cycles are different than water depletion patterns under field conditions (Schoeneweiss, 1975a). Water stress in nature is more often associated with pro longed drought; where soil water potentials remain low or decrease for a long time and exert more constant stress on plants than wide fluctuations between field capacity and some lower value. Therefore,, techniques in which water stress levels can be held relatively constant should pro vide more valid information on water stress than other methods reported. Much evidence has accumulated showing that the metabolism and growth of plants in drying soil are influenced by even mild water deficits in plant tissues and that such deficits usually occur long before drying soil approaches the permanent wilting percentage (Kozlowski 1958ï 1964). This has led researchers to develop techniques that enabled them to expose plants to relatively constant levels of water stress. Miller and Burke (197^) devised a technique for creating constant water stress which utilized ceramic plates under suction as one side of a soil chamber. They were able to maintain constant matric potentials over a four week period. Several workers have used various osmotica in aqueous environments in efforts to maintain constant levels 8
of water stress (Mizrahi et al., 1970; Darbyshlre, 1971). These osmoticums include NaCl (Itai et al., I968; Vaadia and Itai, I968), mannitol (Itai et al., I968; Vaadia and Itai, 1968; Plaut and Ordin, 1964), and polyethylene gly col (Lawlor, 1979; Williams and Shaykewich, 1969; Michel and Kaufmann, 1973; Steuter et al., 1981; Lagerwerff et al., 1961; Ayres, 1977; Jarvis and Jarvis, I965). However, both NaCl and mannitol can be taken up by the tissue (Trip et al., 1964; Kozinka and Klenovska, 1965; Groenewegen and Milis, I96O; Greenway and Leahy, 1970) and produce some unwanted side effects (Jackson, 1965; Lawlor, 1970). This has led workers to compare the use of mannitol versus polyethylene glycol for use as osmoticums (Williams and Shaykewich, I969; Nechiporenko and Rybalova, 1980a; 1980b; Michel, I97O; 1971)' Most have found that polyethylene glycols of higher molecular weight (greater than 3000 daltons) generally do not penetrate the tissue (Jarvis and Jarvis, 1965; Michel, 1971) nor do they cause injury in most systems (Jarvis and Jarvis, I965). Some workers have found side effects from using polyethylene glycols (Kaufmann and Eckard, 1971; Lawlor, 1970; Michel, 1971; Ruf et al., 1967; Mexal et al., 1975)» so although widely used, the applicability of polyethylene glycols as osmotica depends on the sytems and plants you are using (Hsiao, 1973; Michel and Kaufmann, 1973). 9
2. Measurement The only way to determine whether a plant is being subjected to water stress, according to Kramer (1963)» is to measure the water deficit in the plant itself. Methods for measuring water deficits of plants and soils have been reviewed by Barrs (I968), Slavik (1974), Rawlins (1976), and Boyer (I969). Plant water deficits are primarily described by two basic parameters: water content of the tissue and the energy status of the water contained there in (Slatyer and Taylor, i960). According to Barrs,(I968), determination of both components is necessary to describe any water deficit completely. a. Water content Water content, calculated on either a dry or a fresh weight basis, is an unsatisfactory measure of water stress, but water content as a percentage of water content at saturation is useful (Turner and Kramer, 1980). The two most accepted expressions of water content relative to saturation are relative water content (RWC) and water saturation deficit (WSD) (Barrs, I968). RWC is the water content (as a percentage) relative to the water content of the same tissue at full turgor (Barrs, I968; Turner and Kramer, I98O). Leaf water contents have been calculated for whole leaves (Slavik, 1963b), leaf discs (Begg et al., 1964; Jarvis and Slatyer, I966) or leaf segments of long narrow leaves such as the needles of 10
conifers (Jarvis and Jarvis, 1963) or the blades of grasses (Rychnovska and Bartos, I962). A major shortcoming of RWC or WSD as measurement of plant water status is that they are rather insensitive indicators when water deficits are not severe (Hsiao, 1973). Slavik (1963b) also has shown that leaf water content may vary markedly within the area of one leaf blade. Data such as this have led other workers (Kramer and Brix, 1965; Weatherley, 19^5) to conclude that whole organ or plant water content does not provide a satisfactory measurement of water stress. Because size of plant organs is a function of water content (Anderson and Kerr, 19^3i Kozlowski, 1964), several indirect methods of estimating water content have been used (Barrs, I968). These include leaf thickness (Gardner and Ehlig, 1965; Meidner, 1952), stem and trunk diameters (Kozlowski and Winget, 1964; Kramer and Kozlowski, i960), and size determination by beta ray gauging tYamada et al., 1961; Mederski, I96I; 1964). b. Water potential The physiological response of living cells to water stress is closely related to the free energy of the water in the cells (Gates, 1964; Papendick and Campbell, I98I). Water potential is a fundamental concept widely accepted for quantifying the energy state of water in soil, plants, and microorganisms (Hsiao, 1973)- Water potential is the difference between the chemical poten 11
tial of the water in a particular system and that of pure free water at the same temperature, divided by the partial molal volume of water (Slatyer, I967). Since pure free water is usually assigned zero water potential, the potential energy of water in these systems is almost always negative. Water flow is spontaneous from high to low potentials and the availability of water for physiological processes decreases as the potential is lowered (Papen- dick and Campbell, I98I). According to Kramer (I969), "the degree of water stress (in plants) is probably best expressed in terms of water potential, because this appears to be the most closely related to the physiological and biochemical processes which control growth". The components of plant water potential are osmotic potential arising from the presence of solutes, especially in the vacuole; turgor pressure, the hydrostatic pressure developed in the cell In response to osmotic pressure and the tensile strength of the cell wall; and the matric potential arising from the imbibitional forces of colloids in the cell and forces of capillarity in the cell wall (Slatyer and Taylor, I96O). All components of plant water potential are normally negative except pressure potential (Barrs, I968). Matric potential in many species does not become significant numerically until much of the tissue water 12
(.g., 50fo) is lost (Boyer, 196?; Wiebe, I966). So unless the tissue is badly dehydrated, the components of water potential of concern are pressure and osmotic potentials (Hsiao, 1973). As tissue water decreases, changes in total water potential, pressure potential, and osmotic potential are described by the classical Hofler diagram (Salisbury and Ross, 1978). Water potential is related to RWC of the same tissue, the moisture release curve, though the relationship is dependent on species ans stages of growth (Connor and Tunstall, I968; Millar et al., 1968; Gardner and Ehlig, I965» Knipling, I967; Sanchez-Diaz and Kramer, 1972). Mild stress is considered to represent a lowering of plant water potential by a few bars or a lowering of RWC of 8 to 10 percentage points below values in well-watered plants under mild evaporative demand. Moderate stress represents a lowering of water potential by more than a few bars but less than 12 to 15 bars, or a lowering of RWC by more than 10 but less than 20 percentage points. Stress would be considered severe if water potential is lowered more than 15 bars, or RWC more than 20 percentage points (Hsiao, 1973). It is important to note that water stress is not restricted to times of drought. Even in well-watered soils, regular diurnal development of internal water stresses in plants is shown by afternoon decreases in RWC and water potential of plant tissues (Kozlowski, 1968c). 13
Scholander et al. (I965) noted that xylem streams are usually under stress throughout the major growth period of the plant. Numerous methods are available for measuring water potential (Black et al., 1965» Brown and Van Haveren, 1972; Barrs, I968; Slavik, 1974). The techniques are based on the isopiestic principle which dictates that if a sample has the same chemical potential as a surrounding medium, no net change of water will take place (Barrs, I968). Tech niques based on this principle can be divided into two groups: liquid phase and vapor phase. Liquid phase methods involve immersing parallel samples in a range of concentrations of a non-penetrating solute in order to find the solution in which no net exchange of water occurs between sample and solution (Barrs, I968). This can be detected by measuring properties of the samples such as the size of cells or cell groups (Ernest, 1931), length or thickness of tissue strips (Crafts et al., 1949), or cylinders (Lyon, 1936; 19^0), weight (Ashby and Wolf, 1947), or volume (Currier, 19^3) of size stand ardized samples. Sample water potential has also been
determined by measuring properties of the solution before and after sample immersion. These solution properties include density (Kozlowski, 1964), refractive index (Ashby and Wolf, 1947), and volume (Van Overbeek, 1942). After immersion of samples, solutions which give no change 14
in preceding sample or solution properties are said to be isopiestic with the tissue sample, and the osmotic potential of the isopiestic solution is assumed to be equal to the water potential of the sample (Barrs, I968). Errors may arise in liquid phase isopiestic water potential determination by injection of intercellular spaces (Ashby and Wolf, 194?), infiltration of plasmolyzed cells (Slatyer, 1958), and solute penetration (Slatyer, I966). Vapor phase isopiestic techniques are based on the fact that relative vapor pressure above a solution at temperature equilibrium in a closed container becomes equal to the solution's osmotic potential (Slavik, 1974; Barrs, I968). Samples can be exposed to a range of known water potential values without being immersed in the solutions and iso piestic solutions are found by finding the solution which gives no change in sample weight (Slatyer, 1958) or solution volume (Barrs, I968) or weight (Thut, 1938). Because vapor phase isopiestic water potential determination does not require tissue immersion, it circumvents a major source of error inherent in liquid phase isopiestic techniques, namely, solute penetration (Slatyer, I966). It follows from the isopiestic principle that if the water in a sample is allowed to equilibriate with water vapor in air in a closed system, the water potential in the liquid and vapor phases will be equal (Rawlins, 1976). Water potential of a sample can be computed if one is able 15
to measure the vapor pressure of the closed atmosphere surrounding that sample at equilibrium (Brown and Van Haveren, 1972). A psyohrometer is an instrument with wet and dry temperature sensors for measuring the amount of moisture in the air. Thermocouple psychrometry for the measurement of water potentials depends on two thermoelectric effects; the Seebeck and Peltier effects. A thermocouple is formed when wires of two different metals are joined together. If both ends of the wires are joined together to form a closed circuit, an electrical current will flow through the wires if the two junctions are at different tempera tures (Brown and Van Haveren, 1972). This current is called the Seebeck effect. The difference in voltage between the two junctions is a function of their tempera ture difference. By adding a small power source to the circuit, one junction will cool significantly below am bient temperature, while the other junction will become warmer. This is called the Peltier effect. Psychrometric methods are based on measuring the temperature or rate of evaporation from a water droplet or wet surface (Rawlins, 1976). Spanner (1951) was the first to use thermocouple psychrometry to determine water poten tials of plant samples. By using a thermocouple suspended in a closed chamber, Spanner (1951) used Peltier cooling to lower the temperature of the thermocouple below the 16
dewpoint of the atmosphere, and condensation of water resulted on the cooled junction. When the current was shut off, there was an immediate tendency for the con densed water to evaporate back into the enclosed sur roundings. Evaporative cooling depresses the temperature of the thermocouple as a function of evaporation rate, which is a function of the vapor pressure which is a function of the water potential of the sample. By knowing the ambient temperature and temperature depression caused by evaporative cooling. Spanner was able to calculate the vapor pressure of the atmosphere and water potential of the sample. Although it is possible to calculate vapor pressure and hence water potential in this way (Rawlins, 1966; Peck, 1963; I96O; Scotter, 1972), in practice, instruments are calibrated empirically using salt solutions (Rawlins, 1976). Richards and Ogata (1958) built a modified version of Spanner's design in 1958. With this version, water is placed into a loop or ring surrounding the thermocouple to keep the junction wet. That eliminated the need for Peltier cooling to form a drop of condensation and the larger drop of water on the wet loop yields a steady out put for a longer time than Spanner's original psychrometer. Sources of error in psychrometric water potential determination are heat of respiration (Barrs, I968; Boyer, 1969)1 and temperature and vapor pressure gradients (Boyer, 17
1969; Rawlins, 1976). For these reasons, early workers using psychrometry made measurements in carefully con trolled constant temperature baths (Papendick and Camp bell, 1981). It has been found that water baths without temperature control, or enclosures constructed of materials having high thermal conductivities eliminates most error (Campbell and Campbell, 1974; Neuman and Thur- tell, 1972), Since Spanner's psychrometer was built, many variations have appeared including the use of thermistors (Brady et al., 1951; Muller and Stolten, 1953)* Thermistors are more rugged and have greater responses to temperature changes that thermocouples (Rawlins, I976). An important advancement of thermocouple psychrometry has been the use of dew-point depression rather than wet- bulb depression (Neumann and Thurtell, 1972). The dew- point measurement is preferable to wet-bulb because 1) the dew-point temperature measures vapor pressure rather than relative humidity or relative vapor pressure, 2) at the dew point, no net water is exchanged at the wet junction which permits measurements to be made without disturbing the vapor equilibrium in the chamber, 3) the relationship of dew-point temperature to water potential is less dependent upon the ambient temperature than is that of wet-bulb temperature, and 4) psychrometric measurements are influenced by the wetting characteristics of the 18 juncticn, whereas the dew-point technique should be relatively independent of these factors (Rawlins, 1976). Campbell et al. (1973 advanced the dew-point hygrom eter further by utilizing electronic switching that al lowed alternate cooling and measuring of the thermocouple temperature. This circuitry maintains the thermocouple junction precisely at the dew-point. It is this design that many commercial dew-point hygrometers utilize (Wescor, Inc., Logan, Utah). Scholander et aL (1964) used a pressure technique for determination of xylem water potential of excised leafy twigs. This technique involves sealing an excised twig or leaf into a closed chamber and pressurizing it until the xylem sap just begins to exude from the cut surface of the stem or petiole. This technique has certain advantages for measuring plant water deficits. It can be used for field use and since the pressure bomb requires only a short time to obtain a reading, the water status measured is close to that in the intact plant (Tyree and Hammel, 1972). However, because it measures xylem water potential instead of total leaf water potential, relationships between these two values must be established before the pressure bomb technique can be used with meaningful data (Barrs, I968; Boyer, I966; Kaufmann, 1968a). Inmost cases, however, pressure bomb readings compare favorably with those made by thermocouple psychrometers (Boyer, I966; 19
Kaufraann, 1968a). The technique provides accurate, stable estimates of xylem water potential (Schoeneweiss, 1975b) and gives the best field estimates of plant water poten tial (Hodges and Lorio, 1971). 3. Effects on plants a. Growth . Many workers have shown that plant growth is reduced as water stress increases (Gates, I968). This has been well-established for native plants (Petrie and Wood, 1938a; 1938b; Wood, 1932) and cultivated plants such as potato (Cykler, 19^6), wheat (Lehane and Staple, 1962), barley (Williams and Shapter, 1955; Aspinall et al., 1964), rye (Williams and Shapter, 1955)» sorghum (Whiteman and Wilson, 1965), flax (Tiver, 1942), sugar beets (Morton and Watson, 1948), corn (Peters, I96O; Acevedo et al., 1971), tomato (Gates 1955a; 1955b), cotton (Boyer, I965), tobacco (Petrie and Arthur, 19^3), sunflower (Boyer, I968), soybean (Boyer, 1970), and turf- grasses (Burton et al., 1957; Doss et al., 1962; Welton and Wilson, I93I). The level of stress that results in reduction in growth depends on the conditions under which the plants are grown (Begg, I98O). Plants grown in growth chambers are generally more sensitive to water stress than field- grown plants (Begg, I98O). However, response and tolerance of any plant tissue to reduction in leaf water potential is complex because it involves a number of physiological and 20 metabolic processes. As a consequence, growth, as an integrated process, can serve as a measure of plant tolerance under stress (Blum, 1979)» Growth is the irreversible enlargement of cells (Hsiao, 1973)» and involves cell division and enlargement (Salisbury and Ross, 1978). Of these two processes, cell division appears less sensitive to water stress than does cell enlargement (Taylor and Ratliff, 1969a; 1969b; Vaadia et al., I96I; Acevedo et al., 1971; Green and Cummins, 1974; Wenkert et al., 1978a). According to Hsiao (1973)» cell expansion is one of the plant processes most sensitive to water stress, if not the most sensitive of all. Growth of some plants responds to changes in water potential of just fractions of a bar (Cook and Papendick, 1972; 1978). Such sensitive growth retardation responses with increasing water deficits have been cited as a favorable adaptation which increases water use efficiency (Passioura, 1972; Begg and Turner, 1976; Blum and Naveh, 1976). Growth is retarded relative to the degree of tolerance of the tissues or the ability of the whole plant system to avoid drought as moisture stress increases (Blum, 1979)» Tissues from different parts of the plant differ in their sensitivity to water stress (Hsiao, 1973)» Therefore, it may be useful to consider leaf growth, root growth, and growth of the apical meristem separately. 21
Leaf growth is known to be very sensitive to water deficits (Boyer, 1970» Hsiao et al., 1976; Acevedo et al.; I97I; Begg, I98O), Since leaf growth is generally more sensitive to water stress than stomatal conduction and carbon dioxide assimilation, leaf growth can be re duced by water deficits too small to cause a reduction in stomatal aperture and photosynthesis (Acevedo et al., 1971; Boyer, 1970; Fischer and Hagan, I965» Hsiao and Acevedo, 1974; Boyer and McPherson, 1975). In some species, leaf enlargement is so sensitive to water stress that it may be largely confined to the night (Boyer, I968). Low water potentials may cause the loss of existing leaf area if desiccation is prolonged and severe. The loss is caused by a hastened form of leaf senescence, which is particularly rapid in grasses (Boyer, 1976). Also in grasses, leaf rolling or folding is a common response to stress and results in a marked reduction in effective leaf area and a more vertical leaf orientation (Begg, 1980). This is frequently cited in the literature as an adaptive mechanism, which, in addition to reducing the effective leaf area and the energy load on the plant, may markedly reduce transpiration (Parker, I968) and increase water use efficiency (Johns, 1978). Other leaf morphological changes in grasses in response to increasing water deficits include decreased leaf number (Olmsted, 19^1), area (Grant, 1938; 22
Weaver and Hiinmel, 1930), and width (Grant, 1938); leaf thickness generally increases (Roberts and Lage, 1965)'
It is a common observation that the root-shoot ratio of plants increases with water stress (Larcher, 1975» Begg and Turner, 1976; Turner and Kramer, I98O; Beard,
1973; Cheesman et al. , I965» Pearson, I966; Davidson, 1969; El Nadi et al., 1969; Hoffman et al., 1971). In creased root-shoot ratios arise from a greater decrease in growth of shoots compared to the growth of roots (Beard, 1973; Turner, 1979)• In some eases, however, increased root-shoot ratios arise, at least in part, to absolute increases in root growth under moderate water stress (Cheesman et al., 1965; Hsiao and Acevedo, 1974; Bennett and Doss, I96O; Clements, 1964; Doss et al., i960). According to Begg (I98O), the effect is likely to occur at levels of water stress sufficient to reduce shoot growth but not carbon dioxide assimilation. The increase in available assimilates then permits osmotic adjustment and additional root growth. Greater root growth may be reflected by greater root density or rooting depth. Both characteristics are important adaptations in that they allow the plant to extract more soil water and maintain higher plant water potentials (Turner, 1979)» This has important significance in drought resistance because plant water stress generally decreases 23
as the extent of the root system Increases (Donald, 1963; Bertrand, I966; Weatherley, 1965; Wadleigh et al., I966). One selection criterion for crop production under moisture stress is a high root-shoot ratio (Kaul, 196?)« Merion Kentucky bluegrass has good drought resistance that is partially attributed to a deep root system (Funk and Engel, 19635 Hagan, 1954).
Growth of the apical meristem is extremely sensitive to water stress (Husain and Aspinall, 1970; Barlow et al., 1977). However, the apex exhibits drought survivability better than most plant tissues (Barlow et al., 1977). This is especially true for members of the Gramineae for two reasons: 1) the apex is protected from transpirational losses by its protective location within mature leaf sheaths and 2) the apex is not connected to the stem by functional xylem vessels. Consequently, the water content of the apex changes little during stress. During periods of water stress, translocation of solutes continues (Wardlaw, I967). Translocated organic substances are either oxidized to carbon dioxide or accumulate mainly as sugars and free amino acids (Barlow et al., 1980). Accumulation of these substances, along with potassium (Munns et al., 1979) allows the apex to osmotically adjust and maintain turgor at water potentials low enough to kill leaf tissue (Barlow et al., 1977 ; Munns et al., 1979). 24
b. Water relations Water potential is the sum of osmotic, pressure, and raatric potentials. It is well- established that plant tissue water potentials decrease as water deficits become more severe (Kaufinann, 1981). Al though most researchers have disregarded matric potentials in characterizing tissue water relations, there is evidence that matric potentials become increasingly important as water deficits become severe (Boyer, I967; Wiebe, I966; Shepard, 1975)» Although water potential gradients in the soil-plant-atmosphere continuum determine water movement (Salisbury and Ross, I978), osmotic and turgor potentials have more relevance to physiological consequence to increased water deficits (Hsiao, 1973). To fully under stand what is occurring in plants subject to water deficits, it is necessary to measure both water and osmotic potentials and to calculate turgor (Kramer, I98O). The relationship between water, osmotic, and pressure potentials as they change with increasing water deficits is described by the classic Hofler diagram (Salisbury and Ross, 1978). As the tissue loses water from fully hydrated conditions, the most apparent consequence is a loss in turgor until plasmolysis is reached where turgor becomes zero. There have been several reports in the literature of negative turgor, however, these have been discounted in herbaceous plants because the methods used to 25
determine osmotic potential allowed for apoplastic dilution (Tyree, 1976; Kramer, I98O), or matric potentials were disregarded (Shepard, 1975)« c. Water loss Although the major resistance to water flow occurs in the roots (Kramer, 1969; Beyer, 1971; Wind, 1955)* control of water loss from a plant is exercised mainly by regulation of stomatal aperture (Brouwer, 1965; Slatyer and Lake, I966). Several reviews on stomatal control of water loss have been published (Fischer and Turner, 1978; Begg and Turner, 1976; Jones et al., 1982; Burrows and Miltiïorpe, I976; Raschke, 1975» Cowan, 1978; Turner, 1970; Hall et al., 1976) and will not be discussed here. Because plant water loss is so dependent upon stomatal aperture, stomatal aperture can also reflect plant water status (Denmead and Shaw, 1962; Barrs, I968), Stomatal aperture has been used in the field as a criterion for irrigation of crops (Alvim, I966; Shimshi, 1964). Kramer and Brix (I965) have stated that stomatal aperture does not provide a quantitative basis for comparing water deficits in different kinds of plants because stomatal aperture is affected by many other environmental factors. These factors include light glux density (Turner, 1970; 1975; McCree, 1974; Kanemasu and Tanner, 1969b), leaf water status (Troughton, I969; Wilson, 1975; Kanemasu and Tanner, 26
1969a; Biscoei 1972), humidity (Meidner and Mansfield, 1968; Slavik, 1973)» temperature (Goltz and Tanner, 1972; Gifford and Musgrave, 1973; Moss, 1963} Drake et al., 1970), wind (Davies et al., 1974), mineral nutrient supply (Allaway and Milthorpe, 1976), carbon dioxide concen tration (Burrows and Milthorpe, 1976), and other gases such as oxygen (Heath and Orchard, 1956; Zelitch, 1965)» and various air pollutants (Biscoe et al,, 1973; Duggar and Ting, 1970). However, according to Barrs (I968), if the researcher restricts the study to one kind of plant, stomatal aperture can be used to give comparative estimates of water deficits. Most data clearly demonstrate a threshold level of water potential or RWC above which leaf conductance or resistance remains constant (Cowan and Milthorpe, I968; Ehlig and Gardner, 1964; Hsiao, 1973)* When leaf water potentials or RWC fall below a critical value, this leads to a series of partial stomatal closures and reopenings during the day (Ludlow, I98O). The overall effect is a decline in stomatal conductance in a linear or curvilinear manner until it approaches zero or cuticular conductance (Wein et al., 1979). This indicates that stomata are rather insensitive to mild water stress (Hsiao, 1973)* In some species, however, stomata are sensitive to mild water stress showing closure at relatively high water poten 27
tials and no plateau in stomatal response to increasing water stress (El-Sharkawy and Hesketh, 1964; Boyer, 1970; Fischer et al., 1970; Millar et al., 1971; Raschke, 1975; Stalfelt, 1955; 1961). Methods for measuring stomatal apertures and their limitations have been discussed by several authors (Slatyer, 1967; 1958; Barrs, I968; Kramer, I969). There are four main techniques to ascertain stomatal aperture: epidermal peeling and fixing (Lloyd, I9O8; Gale and Poljakoff-Mayber, 1965; MacDowall, 1963)1 surface impressions (Sampson, I96I; Zelitch, I96I; Macklon and Weatherley, 1965), infiltration (Heath, 1959» Dale, 1958; Parker, I963), and porometry (Alvim, I966; Wallihan, 1964). Of these techniques, porometry is the most widelh used for its advantage in expediency and no damage to the leaf being tested (Barrs, 1968). There are basically two types of porometers; pressure- drop viscous flow type (Alvim, I966) and diffusional type (Wallihan, 1964). Diffusional type porometers are most popular because they can estimate stomatal aperture on either side of the leaf and are equally effective with leaves having stomata on one or both sides (Barrs, I968). Diffusion porometers are of two designs: transient and steady state. Transient diffusion porometers depend on measuring the rate of change of water vapor in a small 28
chamber attached temporarily to the leaf (Kanemasu et al., 1969; Van Bavel et al., I965). The air in the chamber is dried and the time taken for the humidity to change over a set range, the transit time, is recorded (Monteith and Bull, 1970)' The diffusion resistance of the sample is determined from a calibration curve relating transit time to known resistance obtained using calibration plates (Monteith and Bull, 1970; Kanemasu et al., I969). In a steady state diffusion porometer, water loss from a leaf is determined by maintaining a constant vapor density in a cuvette that is in contact with the transpiring leaf (Bingham and Coyne, 1977). This is achieved by pumping dry air into the cuvette at an appropriate measured rate to obtain a balance (at a predetermined humidity) between the flux of water transpired by the leaf and the moist air out of the cuvette (Beardsell et al., 1972). Expanding leaves are often accompanied by changing stomatal behavior reaching their highest values for stomatal conductance when the leaves are about 75^ of their full size (Burrows and Milthorpe, I976). Also, there is a rapid increase in the maximum stomatal resistance in the later stages of leaf senescence (Burrows and Milthorpe, I976). Because of this, sampling pre cautions for porometer use include establishing a standard position on the leaf (Alvim, I966), and using leaves of 29 similar age and position (Barrs, I968). In addition, readings should be carried out as quickly as possible with minimal handling and utilizing a large number of samples to overcome variation (Barrs, I968). Stomatal resistances are derived from the difference between the water lost in light and darkness, because it is assumed that stomatal close in the dark and that the cuticular resistance is not affected by illumination (Ayres, 1978). Transpiration is the sum of stomatal and non- stomatal water loss (Crafts, I968). However, stomatal closure is the main cause for transpiration decline as water stress develops (Hsiao, 1973) and, therefore, control of stomatal aperture is very important in drought avoid ance (Townley-Smith and Hurd, 1979)« Plants show a range of stomatal sensitivity to water deficits (Turner, 1979)» as shown by plotting leaf diffusive resistance or con ductance against leaf water potential. The absolute values of leaf water potential that induce stomatal closure depend on many factors including leaf position on the plant (Turner, 1974; Millar and Denmead, 1976), leaf age (Frank et al., 1973)» growth conditions (Jordan and Ritchie, 1971; Beadle et al., 1973), and stress history of the plant (Stocker, I96O; McCree, 1974; Brown et al., 1976; Turner et al., 1978). These factors help to explain the observation that under field conditions, stomata of 30
several plant species remain open at lower leaf water potentials than those found to induce closure under con trolled environment conditions (Davies, 1977» Jordan and Ritchie, 1971). Once threshold water status for stomatal closure is reached, leaf diffusive resitance increases with further drops in leaf water potential (Jordan and Ritchie, 1971; Kanemasu and Tanner, 1969a) or relative water, content (Duniway and Durbin, 1971b; Troughton, I969). Stomates close when bulk leaf turgor pressure is near zero (Turner, 1974; Turner and Jones, 1980). However, changes in stomatal aperture involve changes in the turgor relations of the guard cells and subsidiary cells (Turner, 1974; Meidner and Edwards, 1975)» and several researchers have demonstrated only a loose relationship between bulk leaf turgor and turgor of the guard cells (Turner et al., 1978; Beadle et al., 1978; Turner, 1975» Jones and Rawson, 1979; Brown et al., 1976; Simmelsgaard, 1976). It follows then that if the plant is able to osmotically adjust to increasing water stress and thereby maintain bulk leaf turgor as leaf water potentials de crease, osmotic adjustment can provide an explanation for stomatal adjustment in response to water stress (Turner, 1979). This prompted Kaufmann (1981) to reason that an assessment of drought avoidance capabilities of a plant must include more than a study of the depth and 31
extent of its root systems and the effectiveness of the cuticle and stomata: in restricting water loss. Osmotic adjustment should be evaluated as well. Factors that af fect stomatal adjustment include plant maturity and leaf age (Frank et al., 1973; Morgan, 1977a), rate of stress development (Jones and Rawson, 1979)i and genotype (Blum, 1979; Henzell et al., 1976; Ackerson et al., 1980). Even if the stomatal apparatus is sensitive to water deficits and the leaf water potential at which stomata close is high, full benefit is gained only if the cuticular resistance is also high and stomatal closure is complete (Turner, 1979)* The cuticle of leaves of many crop plants are appreciably permeable, and cuticular transpiration continues after stomatal closure has occurred (Szabo and Buchholtz, I96I; Crafts and Foy, 1962; Darlington and Circulis, I963; Yamada et al., 1964; Bukovac and Norris, 1967). Conversely, it is well-established that when pro longed water deficits prevail, there is an increase in the epidermal waxes of leaves which can decrease cuticular transpiration (Skoss, 1955; Blum, 1975).
When considering sites for resistance to transpiration, possibilities have been raised that non-stomatal factors other than cuticular resistance also play a role in cur tailing plant water loss (Crafts, I968), The terms "mesophyll" or "wall resistance" have been used to describe 32
this type of diffusion resistance. However, it is generally agreed that stomatal resistance is by far the greatest resistance to water loss (Hsiao, 1973)» and mesophyll or wall resistance is negligible until water deficits become quite severe (Fischer, I968; Slatyer, I967). Expressing dry matter or crop yield as a function of water use has proven to be a useful selection criterion for drought tolerant germplasm (Townley-Smith and Kurd, 1979)•
This is called water use efficiency, and is a direct measure of the plants water economy. Various strategies for maximizing water use efficiency have been discussed (Stone, 1975; Pierre,et al., I966). d. Osmotic adjustment and cell wall elasticity Lawlor (1979) states that although the terms water potential has been commonly associated with water stress effects on metabolism, the controlling factors may not be tissue water potential per se, but rather osmotic or turgor potential. Turgor pressure is the driving force for cell expansion (Hsiao, 1973)» Therefore, in order for growth and turgor-mediated processes such as stomatal conductance to be maintained as water deficits increase and plant water potentials decrease, the plant must be able to lower its osmotic potential to maintain turgor (Turner and Jones, 1980). A lowering of osmotic potential in response to water 33
deficits can arise from the concentration of osmotic solutes as water is withdrawn from the vacuole and the cell volume decreases, or additionally, from the accumulation of solutes in the cell (Turner and Jones, I98O). Osmotic adjustment référés to the lowering of osmotic potential arising from the net accumulation of solutes in response to water deficits. By actively accumulating organic and inorganic solutes, plants lower their osmotic potentials more than can be attributed to solute concentration due to a decrease in cell volume (Hsiao et al., 1976; Turner and Jones, 198O), Organic compounds that are utilized for osmotic adjustment include sugars, sugar alcohols, free amino acids (especially proline), and carboxylic acids, while inorganic ions include potassiuiji, sodium, and chloride (Borowitzka, I98I). Earlier literature has demonstrated the ability of plants growing in saline environments to accumulate inorganic ions and thus maintain turgor pressure by lowering their osmotic potentials (Berstein, I96I; Meiri and Poljakoff-Mayber, 1969)» There are now several reports in the literature indicating that leaves of higher plants lower their osmotic potentials in response to slowly developing water deficits (Hsiao et al., 1976; Morgan, 1977%; Fereres et al., 1978; Jones and Turner, 1978; Turner et al., 1978; Jones and Rawson, 1979)• In addition, 34
turgor maintenance by osmotic adjustment has been shown to occur in response to the daily changes in water status of leaf tissue (Turner, 1975; Wehkert et al., 1978b). The degree of osmotic adjustment is measured as the change in osmotic potential at a particular water potential or water content, usually full or zero turgor (Turner and Jones, 1980), However, the degree of osmotic adjustment is limited and depends on the rate of water stress develop ment (Turner and Jones, 1980; Hsiao et al., 1976; Turner et al., 1978; Jones and Turner, 1978) and the plant organ in question (Turner and Jones, 1980). Osmotic adjustment takes place in leaves, hypocotyls, roots, and reproductive organs of several plant species (Turner and Jones, 1980). A greater degree of osmotic adjustment is found in apices compared to expanded leaves. Several species have been shown to osmotically adjust in response to water deficits. They include sorghum (Fereres et al., 1978), corn (Fereres et al., 1978), sunflower (Jones and Turner, 1980), soybean hypocotyls (Meyer and Boyer, 1972), pea (Greacen and Oh, 1972), wheat (Morgan, 1977b), and rice (Cutler et al., 1980). From these studies, it has become clear that considerable differences exist between species in their capacity for osmotic adjustment. However, even though osmotic adjustment is an important mechanism in the drought tolerance of plants 35
(Turner, 1979; Jones et al., 1981), not all species or cultivars show evidence of osmotic adjustment (Turner and Jones, 1980). The pressure-volume technique, originally introduced by Scholander et al., (1964), has been frequently used to measure water, solute, and turgor potentials of a wide variety of tissues (Tyree and Hararael, 1972; Goode and Higgs, 1973; Boyer, 1969; Hellkvist et al., 1974; Richter, 1978; Wenkert et al., 1978b. Pressure-volume curves are plotted as the inverse of the applied pressure (leaf water potential as determined with a pressure bomb) versus the cumulative volume or weight of the expressed sap or residual weight of the tissue (Melkonian et al., 1982). Estimates of osmotic potential are made from the linear region of the curve which is described by the Boyle- Van't Hoff relation, i.e., volume expressed varies linearly with the inverse of the applied pressure (Tyree and Hammel, 1972). Data can also be plotted as water potential versus relative water content of the tissue (Brown and Tanner, 1983). Plots of such data represent moisture release curves (Jarvis and Jarvis, I963). Important parameters affecting drought resistance can be derived from water release curves including relative water content at zero turgor, and osmotic potential at full and zero turgor 36
(Brown and Tanner, 1983î Townley-Smith and Kurd, 1979). In tissue with a high osmotic potential, the osmotic potential decreases with tissue dehydration and solute concentration, and the turgor pressure reaches zero at a relative water content of about 92^. In tissue with a low osmotic potential due to osmotic adjustment, the turgor pressure is much higher at all relative water contents and zero turgor is not reached until the relative water content is about 80^ (Turner, 1979)» Numerous studies indicate that species differences exist, both in the moisture release curve and in relative water content at zero turgor (Turner and Jones, 1980), Age has an effect on this relationship. As the tissue matures, the relative water content becomes higher at given leaf water potentials or at zero turgor (Kassam and Elston, 1974; Knipling, 1967). The moisture release curve depends not only on leaf osmotic potential, but also on cell wall elasticity (Jones and Turner, 1978; Jones et al., 1981; Weatherley, 1970; Turner, 1974; 1979)» Therefore, estimates of cell wall elasticity are important because elasticity influences the relation between water content and water potential (Dainty, 1972; Cheung et al., 1975; 1976; Zimmermann, 1978). Cell wall elasticity is related to cell wall thickness (Turner, 1979; Turner and Jones, 1980). Comparative estimates of cell wall thickness have been made using 37
turgid weight/dry weight ratios of leaves (Wilson et al., 1980). Cell wall elasticity has been characterized by volumetric elastic modulus (Jones and Turner, 1980; Gardner and Ehlig, I965). Estimates of the elastic modulus can be derived from the curvilinear region of the pressure- volume curve where positive turgor is maintained (Melkonian et al., 1982; Tyree and Hammel, 1972; Cutler et al., I98O), or from the relationship between leaf turgor potential and relative water content (Campbell et al., 1979; Melkonian et al., 1982). Although some workers have suggested that dependence of the modulus of elasticity on turgor pressure is inferred from such pressure-volume relations (Hellkvist et al., 1974; Cutler et al., I98O; Steudle et al., 1977; Jones and Turner, 1980), others have shown if estimates are derived from data from a pressure-volume curve of high turgor, then the modulus of elasticity does not depend on turgor or volume (Melkonian et al., 1982; Campbell et al., 1979). The effect of water deficits on cell wall elasticity are unclear. While water deficits reportedly increase cell wall elasticity in Vicia faba (Kassam and Elston, 1974; Elston et al., 1976), decreases in cell wall elas ticity were reported in drought-conditioned sorghum (Jones and Turner, 1978; Jones et al., 1981). No signif- 38 leant changes in elasticity as a result of drought conditioning was observed in either cotton (Hsiao et al,, 1978), rice (Cutler et al., 1980), sunflower (Jones and Turner, I98O), or wheat (Campbell et al., 1979).
B. Disease
1. Effects on plants a. Growth It is axiomatic that pathogens affect normal growth and function of their hosts (Wood, 1953)- Pathogens may affect growth of their .hosts in several ways. Through alterations of their host's hormone metabolism (Sequiera, 1973)» various morphological and developmental changes become evident as a result of in fection. Growth of galls, tumors, and nodules are a direct result of pathogen infection (Agrios, 1969)» In addition, certain pathogens can induce dwarfing of their hosts (Bewley, 1922; Talboys et al., I96I). One pathogen-induced growth response of special interest when investigating the combined effects of infection and water stress is that of reduced root growth. Several pathogens have been shown to reduce root growth of their hosts (Johnston and Miller, 193^» 1940; Duniway and Durbin, 1971a; Amatya and Jones, I966; Van der Wal and
Cowan, 1974; Van der Wal et al., 1975; Gerwitz and Durbin, 39
1965). Hodges (1977) has shown that infection by Ustilàgo striiformis and, to a greater degree, Urocystis agropyri reduce root growth in Poa pratensis 'Merlon'. Root growth reduction is more pronounced than the reduction in shoot growth (Hodges, 1977)» The result is that infected plants exhibit lower root-shoot ratios. Root-shoot ratios have been used as a selection criterion in breeding programs to improve drought re sistance of several crops including wheat (Townley-Smith and Kurd, 1979) and sorghum (Blum, 1979)» Plants with higher root-shoot ratios most often are able to maintain higher water status during periods of drought than plants with low root-shoot ratios. Pathogen-induced reductions in root-shoot ratios would likely predispose infected plants to desiccation during periods of limited water supply or high water loss. Hodges (1977) has shown this to be true for Poa pratensis 'Merion' infected with U. striiformis and U. agropyri. Infected plants exhibited
greater mortality when irrigation was withheld compared to non-infected, healthy plants. b. Water economy Plant pathogens have been shown to adversely affect the water economy of their hosts (Ayres, 1978; Talboys, I968), which can lead to disease-induced water deficits reflected by lower leaf water potentials (Duniway and Durbin, 1971b; Ayres, 1977» 40
Threlfall, 1959? Beckman et al., 1962). Disease-induced water deficits occur because of decreased stomatal resistance (Duniway and Durbin, 1971a, Ayres, 1976), decreased cuticular resistance (Sempio et al., 1966; Amatya and Jones, I966; Gerwitz and Durbin, I965), reduced hydraulic conductivity (Dimond, 1970; Talboys, I968), and reduced water uptake due to pathogen-induced root growth inhibition (Murphy, 1935» Hodges, 1969; 1970a). Duniway and Durbin (1971b) found that bean rust totally inhibited the movement of stomata. Other pathogens have been shown to exhibit similar inhibition of stomatal function in potatoes (Farrell et al., I969), tomato (Rubin and Artsikhovskaya, I963)» barley (Ayres, 1972;
Ayres and Jones, 1975)» and tobacco (Cruickshank and Rider, I96I). In addition to stomatal inhibition, the pathogen may affect the cuticular resistance of diseased plants by invasion through host epidermis, or rupture of the epidermis to sporulate (Ayres, 1978). This is true for the rust diseases where transpiration is often reduced early in the infection cycle, but in creases after the pathogen ruptures the epidermis to re lease its spores. Rust diseases also reduce leaf area development and dry matter accumulation (Ayres, 1978). This reduction in growth often causes the total amount of water taken up or lost by rusted plants to be less than 41
that taken up or lost by healthy plants (Johnston and Miller, 193^? 19^0; Murphy, 1935; Sever, 1937). Although smut fungi are closely related to the rust, little is known about their effects on host water economy except that they increase transpiration (Ayres, 1978). Transpiration stream interruption can lead to water deficits in aerial parts of the plant. Various vascular pathogens are known to lower the hydraulic conductivity of their hosts by the production of toxins or gums. Infec tion also stimulates the hosts to produce occlusions and tyloses (Ayres, 1978). Transpiration stream interruptions may lead to symptom development on leaves which can parallel a drought syndrome (Talboys, I968). It is well-documented that many pathogens inhibit root growth of their hosts (Ayres, 1978; Talboys, I968; Gerwitz and Durbin, 1965; Duniway and Durbin, 1971a, Johnston and Miller, 1934; 1940), Hodges (1970a) has demonstrated decreased root growth of P. pratensis infected with U. striiformis or U,. agropvri which contributed to increased mortality of infected plants during periods of water stress compared to healthy control plants. 2. Leaf smuts
Urocvstis agro-pvri (flag smut) and Ustilago striifor mis (stripe smut) are pathogens of a number of grass species including Poa pratensis (Kentucky bluegrass) 42
(Couch, 1971). 'Merion' Kentucky bluegrass is one of the most widely planted turfgrasses in the United States, and is very susceptible to both smut organisms (Gaskin, 1965; Kreitlow and Juska, 1959)• Primary infection sites include coleoptiles on seedlings and axillary buds on crowns and rhizome nodes of mature plants (Hodges, 1976). Mycelia systemically infects leaf, sheath, and floral parts of the plant (Couch, 1971)» and becomes perennially established for the life of the host (Hodges and Britton, 1970)• The disease spreads in established grasses by systemic growth within tillers and rhizomes produced by infected plants, and by new primary infections of axillary buds (Hodges and Britton, I969; 1970)* Invading fungal hyphae cause striking morphological changes in the host (Hodges, 1970a). Infection by U. agropvri results in overall stunting, reduced root growth, and increased intravaginal branching (Hodges, 1970a). Infection by U. striiformis has been reported to cause an increase in plant height, increased lamina width, wider lamina to sheath angles, as well as a striping symptom on leaf sheaths and laminas (Hodges., 1970c). Both pathogens produce sori in leaf mesophyll that may extend the entire length of the leaf (Davies, 1922; Kreitlow, 1948; Thirumalaohar and Dickson, 1953)» As sori mature, sporulation results in epidermal rupturing as teliospores ^3
are released• Epidermal rupturing results in loss of non- stomatal control of water loss and plants become more susceptible to periods of water stress. Hodges (1977) has demonstrated increased mortality rates of P. pratensis infected by U, striiformis or U. agropyri when irrigation was withheld compared to healthy plants. Other researchers have established that infected plants are readily killed by high temperatures and drought (Fischer, 19^0; Kreitlow, 19^3; Leach et al., 19^6), 44
III. GROWTH OF WATER-STRESSED POA PRATENSIS INFECTED BY UROCYSTIS AGROPYRI OR USTILAGO STRIIFORMIS
A. Abstract
Growth of healthy Poa pratensis 'Merion' or systemically infected by Ustilago striiformis (stripe smut) or Urocystis agropvri (flag smut) was evaluated for total, leaf, and root weights, and root-shoot ratioç in nutrient solution and in nutrient solution amended
with polyethylene glycol to provide increasing levels of osmotically-induced water stress. Infection by U. striiformis stimulated total and leaf weight, but de creased root weight and root-shoot ratios of P. pratensis grown in nutrient solution. P. pratensis infected by U. agropvri exhibited decreased total, leaf, and root weights and root-shoot ratios.
Total, leaf, and root weights of all three health groups decreased, and root-shoot ratios increased with decreasing osmotic potentials of nutrient solutions. The increases in total and leaf weights of U. striiformis- infected plants compared to healthy plants grown in nutrient solution were lost as nutrient solution osmotic potentials decreased. Healthy plants maintained higher root-shoot ratios than infected plants at all stress levels. 45
B. Introduction
Urocystis agropvri (flag smut) and Ustilago striiformis (stripe smut) are pathogens of a number of grass species including Poa pratensis (Kentucky bluegrass) (Couch, 1971), 'Merion' Kentucky bluegrass is very susceptible to both pathogens (Gaskin, 1965; Kreitlow and Juska, 1959). Primary infection sites include coleoptiles on seedlings and axillary buds on crown and rhizome nodes of established plants (Hodges, 1976). Mycelium systemically infects leaf, sheath, and floral parts of the plant (Couch, 1971), and becomes perennially established for the life of the host (Hodges and Britton, 1970). The disease spreads in established grasses by systemic growth within tillers and rhizomes produced by infected plants, and by new primary infections of axillary buds (Hodges and Britton, I969; 1970), Leaf-smutted P. pratensis has higher mortality rates than healthy plants when subjected to water stress (Hodges, 1977). This suggests that the imposition of water stress on leaf-smutted plants enhances the biotic stress induced by the pathogen and increases the mortality rate of diseased plants. It has been proposed that the process of growth may be the most sensitive of all plant processes to water stress (Hsiao, 1973)» and that growth, as an 46 integrated process, can serve as a measure of plant tolerance under stress (Blum, 1979)• This study was initiated to investigate the effects of progressively lower nutrient solution osmotic potentials on the growth of P. pratensis 'Merion* systemically infected by U. striiformis or U. agropvri.
C. Materials and Methods
A single clone of healthy, U. striiformis-, or U. agropyri-infected Poa pratensis 'Merion' was vegetatively- propagated from individual shoots and grown in a loams peatsperlite (1:1:1) mix under greenhouse conditions. Individual shoots selected for experiments were removed from the soil mix, roots were washed, and all but the two youngest fully-expanded leaves were removed from the shoot. Individual shoots were placed in specially designed nutrient culture jars (Pig. 3.1) to provide continuous aeration of the nutrient solution (Table '3.1). Nutrient solution was similar to that used by Pellet and Roberts (1963) except higher rates of nitrogen were used because Hull et al. (1979) have shown that it favors symptom development. Experiments were initiated to determine the effect of infection on growth in nutrient solution per se, and in nutrient solution with progressively lower 47
- POA PRATENSIS SHOOT SILICONE GREASE AIR INLET SEAL
AIR EXHAUST INVERTED #4 SINGLE HOLE RUBBER STOPPER 100ml VOLUME MARK SILVER PAINTED NUTRIENT CULTURE JAR
Figure 3.1. Nutrient culture jar used to grow Poa pratensis 48
Table 3.1. List of nutrients, their sources and con centrations used in nutrient solution
Nutrient Source Concentration (ppm)
N Ca(NO^)2 ' 156 ^
P 25 K KOH 50 ^
Mg MgSO^ • 7H2O 19 Fe FegfSO^ 1.2 ^3 Mn MnSO^ • HgO 0.25 B H^BOj 0.1 Zn ZnSO^ • yHgO 0.1 Cu CUSO4 ' 0.01 Mo MoO- ^ 0.01
^Different from Pellet and Roberts (1963)» osmotic potentials induced by polyethylene glycol. Healthy (non-infected) control plants were grown under the same conditions. 1. Nutrient solution studies Seventy-five two-leaf plants were selected for size uniformity from each health groups (i.e., stripe- smutted, flag-smutted, and healthy) for evaluation of 49 the effect of infection on growth of P. pratensis in nutrient solution. Plants were randomized in the growth chamber using a randomized complete block design utilizing 3 blocks and 3 treatments (health groups). Growth chamber conditions included 12-hour photoperiods, 50^ relative humidity, 65 (- 5) nE cm~^ sec~^ light intensity, 23 C (- 2) constant temperature. Plants were grown in nutrient solution for 4 weeks and the solution was flushed twice per week. Fifteen plants from each treatment (5 from each block) were sampled for total dry weight (dried at 60 C, 48 hours), leaf dry weight, and root dry weight at 0, 1, 2, 3, and 4 weeks and presented as weekly means for each health group. Root-shoot ratios were calculated from dry weights. Means were bracketed by their standard errors• 2. Water stress studies Plants evaluated for growth in response to in fection and progressively lower osmotic potentials of nutrient solution were preconditioned by growing them in nutrient solution alone for 10 days before applying water stress treatments. Plants were then placed in nutrient solution containing polyethylene glycol (PEGqqqq) at concentrations of 0, 168, 242, and 292 g/L. These concentrations provided initial osmotic potentials of -0,2, -0.4, -0.6, and -0.8 MPa as measured by a Wescor 50
HR-33T dewpoint microvoltmeter using calibrated C-52 sample chambers and the dewpoint mode. Nutrient solutions were flushed twice per week and growth chamber conditions were as previously stated. Plants were grown for 4 weeks under applied water stress after which 12 plants from each treatment combination were sampled for total, leaf, and root dry weights. Root-shoot ratios were calculated. The experiment utilized a 3 by 4 factorial in a randomized complete block design,. Treatment factors included 3 health groups (healthy, U. striiformis-. and U. agropyri-infected) and 4 levels of osmotically- induced water stress. Data were presented as means of healthy, U. striiformis-. and U. agropyri-infected plants for total, leaf, and root weight, and root-shoot ratios for -0.2, -0.4, -0.6, and -0.8 MPa initial nutrient solution osmotic potentials. Means were bracketed by their standard errors.
D, Results
1, Nutrient solution effects Poa pratensis infected by U, striiformis and grown in nutrient solution increased in total and leaf dry weight, and decreased in root growth and root-shoot ratios compared to healthy plants (Fig. 3.2). Plants Figure 3.2. Total (A), leaf (B), and root (C) dry- weights and root-shoot ratios (D) of healthy (H), Ustilago striiformis- infected (stripe smut) (S), and Uro- ovstis agropyri-infeeted (flag smut) (F) Poa pratensis 'Merion* shoots grown four weeks in nutrient solution ROOT/SHOOI RATIO (mg/mg) ROOT WEIGHT (mg) lEAF WEIGHT (mg) lOTAl WEIGHT (mg) 53 infected by U. agropyri decreased in total, leaf, and root weight, and in root-shoot ratios compared to healthy plants (Fig. 3»3A-D). No differences were found, however, between root-shoot ratios between flag-smutted and stripe-smutted plants (Pig. 3»2D). 2. Water stress effects Total, leaf, and root dry weights of all three health groups decreased with decreasing osmotic potentials of nutrient solution (Fig, 3.3A-D). Root-shoot ratios in creased as the osmotic potential of the nutrient solution decreased for all health groups (Fig, 3»3D). Total and leaf weight of stripe-smutted plants increased compared to healthy plants in -0.2 MPa nutrient solution, and then decreased as osmotic potential of the nutrient solution decreased (Fig, 3.3A-C), Root dry weights and root-shoot ratios of infected plants were less than that of healthy plants at all nutrient solution osmotic potentials (Fig, 3.3C and 3.3D).
E, Discussion
Systemic infection of 'Merion' P. pratensis by U. striiformis increased total and leaf dry weight of plants grown in nutrient solution without polyethylene glycol compared to healthy plants, but root growth Figure 3«3* Total (A), leaf (B), and root (C) dry- weights androot-shoot ratios (D) of healthy (H), Ustilago striiformis- infeoted (stripe smut) (S), and Uro- cvstis agropyri-infe cted (flag smut) (F) Poa pratensis 'Merion' shoots after four weeks in PEGgQQQ-amended nutrient solution ROOT/SHOOT RATIO (mg/mg) ROOT WEIGHT Imgl LEAF WEIGHT img) TOTAL WEIGHT (mg) 56
is inhibited. Infection by U. agropyri inhibits all growth of the central crown. Related morphological changes in P. pratensis associated with these pathogens include reduced extravaginal branching and inflorescence production, and increased intravaginal branching and lamina to sheath angles (Hodges, 1969» 1970a). These observations are suggestive of metabolic imbalances induced by the pathogens. The increase in total and leaf weight of U. striiformis-infected plants compared to healthy plants at -0.2 MPa may be related to hormonal imbalances (Sequeira, 1973). Both pathogens decrease the total sugar and amino acid content of infected plants (Hodges and Robinson, 1977; Madsen et al., 1983)*. but these effects cannot explain the increase in total and dry leaf weights under minimal stress. The decrease in all growth parameters of U. agropvri-infected plants, however, may be due to the severe decrease in sugars and amino acids alone. Pathogen-induced dwarfing of hosts has been observed in other plant diseases (Bewley, 1922; Talboys et al., I96I). Decreased root growth of infected plants may be in part due to hormone imbalances that affect assimilate partitioning (Zeevart, 1979), however, one cannot rule out that root growth inhibition is due to assimilate depletion, per se. 57
The decrease in leaf weight in response to in creasing water deficits was disproportionately greater than that of the roots and is in agreement with similar observations (Larcher, 1975; Begg and Turner, 1976; Turner and Begg, 1978; Beard, 1973). The resulting increase in root-shoot ratios is attributed mainly to this differential growth response (Turner, 1979» Beard, 1973)» and not to an increase, in root growth as has been reported in some cases (Cheesman et al., 1965; Hsiao and Acevedo, 1974; Bennett and Doss, 1969). Infection of P. pratensis by U. striiformis or U. agropvri sharply reduces root-shoot ratios (Fig, 3.2D) and the effect is persistent as water stress is increased (Fig. 3.3D). High root-shoot ratios are indicative of greater drought resistance and are used as selection criteria for drought resistance (0'Toole and Chang, 1979). The lower root-shoot ratio suggests that drought resistance is decreased by the pathogens and supports the observation that mortality of infected plants increases without irrigation (Hodges, 1977). If overall growth, as an integrated process, can serve as a measure of plant tolerance under stress (Blum, 1979)1 then U. striiformis-infected plants grown in nutrient solution can tolerate the amount of stress induced by the pathogen (Fig, 3.3A). Total weight was 58 not reduced by infection by U. striiformis in P. pratensis growing in nutrient solution. The detri mental combination of the stress imposed by the presence of the pathogen and osmotically-induced water stress, however, becomes quickly evident as water stress in creases (Fig, 3,3A). Decreases in total weight as a response to increasing water stress is more severe in infected plants and may partially explain the ob servation that infection by either U, agropvri or U, striiformis results in growth reduction of field-grown grasses (Couch, 1971). This suggests that the pathogens have decreased the stress tolerance of their hosts, which when water deficits increase may lead to increased rates of mortality of leaf-smutted Poa pratensis demon strated by Hodges (1977), 59
IV. EFFECT OF WATER STRESS AND SYSTEMIC INFECTION BY USTILAGO STRIIFORMIS OR UROCYSTIS AGROPYRI ON THE GROWTH OF TILLERS AND RHIZOMES OF POA PRATENSIS
A. Abstract
Water stress and infection of Poa pratensis 'Marion* by Ustilago striiformis (stripe smut) or or Urocystis agropvri (flag smut) were evaluated for their effects on tiller and rhizome growth. Plants were grown in nutrient solution and in nutrient solution with increasing amounts of polyethylene glycol to provide increasing levels of osmotically-induced water stress. Stripe smut had no affect on mean total dry weight of rhizomes per shoot, but the mean percentage of rhizome dry weight as a portion of total shoot dry weight decreased compared to healthy plants. Dry weight of tillers produced by stripe-smutted plants in nutrient solution increased and were a greater mean percentage of total shoot dry weight compared to healthy plants. Flag-smutted plants in nutrient solution showed decreased rhizome growth and increased tiller growth compared to healthy plants. The stimulation of tiller growth by stripe-smutted plants was negated with increasing water stress. Flag-smutted plants, however, continued 60
to show increased tillering at the most severe levels of water stress. Implications of altered hormonal balance in U. agropyri-infected plants are discussed.
B, Introduction
Poa pratensis (Kentucky bluegrass) vegetatively propagates itself by producing tillers and rhizomes. Rhizome and tiller production is at a maximum in spring, late summer, and fall, and is minimal during periods of environmental stresses including heat, cold, and drought (Beard, 1973). Ustilago striiformis and Urocvstis agropyri are the causal organisms of stripe and flag smut, respectively. They are serious pathogens of several grass species in cluding P. pratensis (Couch, I97I). Invading fungal hyphae can infect leaf, sheath, and floral parts of the host and cause striking morphological changes in the
host (Hodges, 1970a; I969). Because of the directional growth and systemic nature of the pathogens (Hodges and Britten, 1970), tillers and rhizomes from infected plants are almost always infected as they develop from axillary buds of infected plants (Hodges and Britten, 1969)» Therefore, such factors as irrigation and nitrogen fertilization that increase the survival of infected 61
plants and favor tiller and rhizome production can enhance the spread of the disease (Hodges, 1977; Hull
et al., 1979)' Physiological control of tillering is subject to auxin and carbohydrate concentrations in the grass plant (Leopold, 1949). Infection by U, striiformis or U. agropyri may decrease the soluble sugar content in leaves and roots of P. pratensis (Hodges and Robinson, 1977; Madsen et al., I983). Growth responses of P. pratensis infected with U. agropvri or U. striiformis also are suggestive of growth hormone imbalances (Hodges, Hodges, 1970a; 1969). This study was initiated to determine the effect of infection by U. striiformis or U. agropvri on tiller and rhizome growth of P. pratensis grown in nutrient solution, and to evaluate the combined effects of sys temic infection and osmotically-induced water stress on tiller and rhizome growth.
C. Materials and Methods
A single clone of healthy, U. striiformis-, or U. agropvri-infected Poa pratensis was vegetatively- propagated from individual shoots and grown in a loam: peat:perlite (1:1:1) mix under greenhouse conditions. 62
Individual shoots selected for experiments were removed from the soil mix, roots were washed, and all but the two youngest fully-expanded leaves were stripped from the shoot. Individual shoots were selected for size uniformity for growth chamber experiments. 1. Nutrient solution studies Seventy-five shoots with two leaves each were taken from each health group (i.e., stripe-smutted, flag-smutted, and healthy) for evaluation of tiller and rhizome growth in nutrient solution. Plants were randomized in the growth chamber using a randomized complete block design utilizing 3 blocks and 3 treat ments (health groups). Growth chamber conditions included 12-hour photoperiods, 65 (- 5) nE cm"^ sec~^ light intensity, 23 C (- 2) constant temperature. Plants were placed individually in specially designed nutrient culture jars (See Chapter III) and grown for 4 weeks. Nutrient solutions were flushed twice per week. At 0, 1, 2, 3, and 4 weeks, 15 plants from each treatment (5 from each block) were sampled for their respective total dry weight (dried at 60 C, 48 hours) of rhizomes and tillers per shoot. Tiller and rhizome percentages were calculated by expressing the tiller and rhizome dry weights as a percentage of the total dry weight of each shoot. Data were plotted as weekly means of healthy, U. 63
striiformis-, and U. agropvri-infected plants for rhizome dry weight per shoot, rhizome percentage of total shoot dry weight, tiller dry weight per shoot, and tiller percentage of total shoot dry weight. Each mean was bracketed by its standard error. 2. liJater stress studies Plants evaluated for growth in response to infection by U. striiformis and U. agropyri and to osmotically- induced water stress were preconditioned by growing them in nutrient solution without polyethylene glycol
(PEGqqqq) for 10 days before water stress treatments were applied. Plants were selected for size uniformity and placed in specially designed nutrient solution jars (See Chapter III) containing 0, 168, 242, or 292 grams
of PEGqqqq per liter nutrient solution. This provided initial nutrient solution osmotic potentials of -0,2, -0.4, -0.6, and -0.8 MPa as measured by a Wescor HR- 33T dewpoint microvoltmeter using calibrated C-52 sample chambers and the dewpoint mode. Nutrient solutions were flushed twice per week and growth chamber conditions were as previously stated. Plants were grown for four weeks under applied water stress after which 12 plants from each of 12 treatment combinations were sampled for
total dry weight per shoot and dry weight of rhizomes and tillers per shoot. Rhizome and tiller percentages were 64
calculated by dividing rhizome and tiller dry weight per shoot by total dry weight per shoot, respectively, and expressing ratios as percentages. The experiment utilized a 3 by 4 factorial in a randomized complete block design. Treatment factors included 3 health groups (healthy, stripe-smutted, and flag-smutted plants) and 4 levels of osmotically-induced water stress (-0.2, -0.4, -0.6, and -0.8 MPa). Data were plotted as the means of healthy, U. striiformis-. and U. agropyri-infected plants for rhizome and tiller dry weight per shoot and for rhizome and tiller percentage of total shoot dry weight. Each mean was bracketed by its standard error.
D. Results
1. Nutrient solution effects Infection by U. striif ormis had no effect on total dry weight of rhizomes per shoot on P. pratensis grown in nutrient solution (Fig. 4.1A). Rhizome growth of stripe-smutted plants expressed as a mean percentage of total shoot dry weight, however, decreased between the second and third weeks of the study (Fig. 4.IB). Infection by U. striiformis increased the total dry weight of tillers per shoot and their mean percentage of the total dry weight of the shoot (Fig. 4.ID) compared to healthy plants grown Figure 4.1. Weekly mean dry weights of rhizomes (A) and tillers (C) and their respective mean percentages of the total shoot dry weight (B and D) for healthy (H), Ustilago stri- iformis-infected (stripe smutted) (S), and Urocystis agropvri-infected (flag smutted) (F) Poa pratensis 'Merion' shoots grown four weeks in nutrient solution HUER PERCENTAGE TILLER WEIGHT (mg) RHIZOME PERCENTAGE S S 67
in nutrient solution. Infection by U. agropvri sharply decreased rhizome growth on plants grown in nutrient solution (Fig. 4.1A and 4.IB), Conversely, flag-smutted plants exhibited a sharp increase in tiller growth compared to healthy plants (Fig. 4.1C and 4.ID). 2. Water stress effects Progressively decreasing the osmotic potential of the nutrient solution in which healthy, U. striiformis-, and U. agropvri-infected plants were growing generally decreased the mean dry weight of rhizomes and tillers and their respective percentages of total plant dry weight (Fig. 4.2A-D). Tiller growth associated with both pathogens was greater than that of healthy controls under minimal stress (Fig, 4.20). Increasing water stress de creased tillering by plants infected by either pathogen, but tillering by flag-smutted plants remained greater than that of controls. Rhizome growth was decreased by both pathogens under minimal water stress, but more severely by flag smut (Fig. 4.2A). Increasing water stress reduced the differential decrease in rhizome growth between healthy and stripe-smutted plants, but flag-smutted plants showed severe decreases in rhizome growth at all levels of water stress (Fig. 4.2A and 4.2B). Figure 4.2. Mean dry weight of rhizomes (A) and tillers (C) and their respective mean percentages of the total shoot dry weight (B and D) for healthy (H), Ustilago striiformis- infected (stripe-smutted) (S), and Urocystis agropyri-infected (flag-smutted) (F) Poa pratensis 'Merion' shoots after four weeks growth in nutrient solution amended with
^^^8000 69
250
= 100
30 B
- 200 S 150
60
-0.8 -0,6 -0.2 NUTRIENT SOLUTION OSMOTIC POTENTIAL (MPa) 70
E. Discussion
This study has shovm that systemic infection by U. agropyri greatly stimulates the growth of tillers of Poa pratensis, but decreases the grwoth of rhizomes. The same general trend appears for U, striiformis- infected plants, but the effect is much less dramatic (Fig, 4.1A-D). This agrees with the observation that U. agropyri severely decreases rhizome production of P. pratensis grown in pot culture (Hodges, 1970a, I969). Growth of tillers in grasses uses large reserves of stored carbohydrates (Younger and Nudge, I968; Sachs and Thimann, 196^). The stimulation of tilering in U. striiformis and U. agropyri-infected plants may account for the re duction in carbohydrates found in infected plants (Hodges and Robinson, 1977» Madsen et al., I983). Tiller and rhizome growth in P. pratensis are sensitive indicators of water stress (Fig. 4.2), and decreases in their growth in response to water deficits has been previously observed (Olmsted, 1941; Quarrie and Jones, 1977). Decreased tiller and rhizome growth in response to water stress may be due to the extreme sensitivity of apical meristems (Hsiao, 1973)» or environmentally- induced hormonal disturbances. Hormonal disturbances in water stressed plants include increases in abscisic acid 71
and decreases in cytokinins (Livne and Vaadia, 1972). Increased lAA oxidase activity has also "been observed in response to water deficits (Darbyshire, 1971). Tiller and rhizome grwoth in grasses depends on development of lateral buds (Beard, 1973)» the growth of which may be inhibited by auxin produced in the apical meristem (Snow, 1925; Leopold, 19^9; Thimann and Skoog, 193^)» or require growth promoters such as gibberellins (Eagles and Wareing, 1964) to stimulate their development. Plant pathogens of the Basidiomycetes often induce hormonal imbalances in their hosts (Brian et al., 1954; Sequiera, 1973). Auxin imbalances induced by U. agropvri might promote lateral bud break (Leopold, 1949); and increase tillering, but does not explain the near absence of rhizome production. 72
V. EFFECT OF WATER STRESS AND INFECTION BY USTILAGO STRIIFORMIS OR UROCYSTIS AGROPYRI ON LEAF TURGOR AND WATER POTENTIALS OF POA PRATENSIS
A. Abstract
Systemic infection by Ustilago striiformis or Urocystis agropvri and water stress were evaluated for their effects on leaf turgor and water potentials of Poa pratensis 'Merion* when grown in nutrient solution. Infection by either pathogen decreased leaf turgor and water potentials during light and dark periods compared to healthy controls. Healthy plants maintained higher leaf turgor and water potentials than infected plants as the nutrient solution osmotic potential was lowered with polyethylene glycol. Moisture release curves for single leaves from healthy, U, striiformis-, and U. agropvri-infected plants were established to evaluate plants for osmotic adjust ment by estimating leaf osmotic potentials and relative water contents at zero turgor as plants were grown in nutrient solution and nutrient solution amended with polyethylene glycol to increase water stress. Turgid weight/dry weight ratios also were calculated for healthy, U. striiformis-, and U. agropyri-infected leaves before 73 and after water stress treatments. Water stress lowered the leaf osmotic potentials and relative water contents at zero turgor and decreased the turgid weight/dry weight ratios of healthy and infected leaves. Healthy plants exhibited lower osmotic potentials at zero turgor, however, and smaller turgid I weight/dry weight ratios after water stress than infected plants suggesting that infection by either pathogen inhibited osmotic adjustment and an increase in cell wall thickness of P. pratensis leaves in response to water stress.
B. Introduction
Several pathogens cause water deficits in their hosts by altering the host's water relations (Duniway, 1973; Ayres, 1977; 1978; Duniway and Durbin, 1971a; Talboys, I968). Host water deficits can be brought about in a number of ways including accelerated water loss, reduced water uptake, and loss of membrane in tegrity which may result in the loss of the cell's ability to retain solutes and water (Ayres, 1978). Few studies have considered the combined effects of environmentally-imposed and disease-imposed water stress on plants (Ayres, 1978). 74
Urocystis agropvri and Ustilago striiformis are pathogens of a number of grass species including Poa pratensis (Couch, 1971). During periods of water stress, mortality of leaf-smutted P. pratensis is higher than that of healthy plants (Hodges, 1977; Kreitlow, 1943). This suggests that infection by these pathogens may impose a direct water stress on the host and/or alter drought resistance of the host. To determine the effects of either environmentally-imposed and/or disease- imposed water stress, it is necessary to measure both water potential and osmotic potential and to calculate turgor potential (Kramer, 1980). This study was initiated to determine the effect of U. striiformis or U. agropyri on leaf turgor and water potential of P. pratensis grown in nutrient solution and to determine if infection by either organism affects osmotic adjustment or turgid weight/dry weight ratios of leaves as healthy and host plants are subjected to increasing water stress.
C. Materials and Methods
A single clone of healthy, U. striiformis-, or U. agropvri-infected Poa pratensis 'Merion' was vegetatively propagated from individual shoots and grown in a loam: 75 peat:perlite (1:1:1) mix under greenhouse conditions. Individual shoots selected for experiments were removed from the soil mix, roots were washed, and all but the two youngest fully-expanded leaves were stripped from the shoot. Individual shoots of healthy, U. striiformis-, and U. agropvri-infected plants were grown in the growth chamber in nutrient solution (see Chapter III) for three weeks, then all but the youngest three fully- expanded leaves were removed and the plants were placed back into the growth chamber in aerated nutrient culture jars (see Chapter III) and held for osmotic and water potential measurements. Growth chamber conditions included a 12-hour photoperiod, 65 (- 5) nE cm" sec ~ light intensity, 23 (- 2) C, and 50^ relative humidity. Leaf water potentials were determined over a period of 10 days utilizing a PMS pressure bomb. Only the second fully- expanded leaf was used for water potential determinations. Prior to leaf excision, leaf blades were sealed with petroleum jelly to avoid rapid water loss before and during pressurization. A slow pressurization rate (0.01 MPa sec"^) was used to insure accurate endpoint deter mination. Osmotic potentials were determined from expressed sap from the same leaf using a Wescor HR-33T 76
dewpoint microvoltmeter in the dewpoint mode. Cali brated C-52 sample chambers were used with a vapor equilibrium time of 5 minutes. Leaf turgor potentials were calculated as the difference between water and osmotic potentials. Data were plotted as scatter diagrams of water and turgor potentials versus time for healthy, U. striiformis-. and U. agropyri-infected plants. Each data point represents the mean of three leaves. Healthy, U. striiformis-, and U. agropvri-infected plants were measured for prelight (synonymous with predawn) leaf water and turgor potentials under conditions of increasing water stress. Prelight values were taken from 3 hours before to onset of light period. Polyethylene glycol (PEGgogg) was used to lower the osmotic potential of the nutrient solution at the rate of 0,1 MPa day"^ until -0.8 MPa was reached. Solutions were flushed at the end of the light period daily and replaced with pro gressively more concentrated solutions. Prelight leaf water and osmotic potentials were determined as pre viously described at nutrient solution osmotic potentials of -0.2, -0.4, -0.6, and -0.8 MPa. Prelight turgor potentials were calculated as the difference between leaf water potentials and osmotic potentials. Data were expressed as prelight leaf water and turgor potentials versus nutrient colution osmotic potential. Each data 77
point represents the mean of 8 leaves and has been bracketed by its standard error. Evidence for osmotic adjustment was evaluated by establishing water release curves using a method similar to that described by Brown and Tanner (1983) for the second fully-expanded leaf of healthy, U. striiformis-, and U. agropvri-infected plants. Curves were established for plants growing in nutrient solution before and after plants were water stressed using PEGggoO "the manner previously described. After nutrient solution osmotic potential reached -0.8 MPa (6 days), nutrient culture jars (See Chapter III) were flushed with distilled , water and plants were allowed to réhydraté in the dark for at least 6 hours. Before leaf excision for water potential determinations, leaves were sealed with a very thin coat of petroleum jelly. After excision, the leaf was weighed immediately and placed in a PMS pressure bomb and pressurized at the rate of 0,01 MPa sec"^. After endpoints were attained, pressure was relieved slowly (approximately 0,03 MPa sec"^). The leaf was weighed again, water potential determined, and the process was repeated until leaf water potentials were below -3«5 MPa. Data were plotted as leaf water potential versus leaf weight, linear regression was used on the linear 78 portion of the curve to estimate leaf weight at full turgor {Fig. 5.1A). values for these regressions were consistently near O.99. After curves were established, leaves were oven-dried (60 C, 48 hours) and dry weights recorded. Relative water contents (Barrs» I968) for each data point were calculated and data were replotted as water potential versus relative water content (the moisture release curve) (Fig. 5*1B). The point of inter section between the linear and curvilinear phases of the moisture release curve was used as the estimate of zero turgor. Values for leaf osmotic potential and relative water content at zero turgor were recorded (Fig. 5»1B). Eight leaves of healthy, U. striiformis-. and U. agropyri-infected plants were evaluated before and after stress treatments. Turgid weight/dry weight ratios were also calculated for those healthy, U. striiformis-, and U. agropyri-infected leaves of P. pratensis before and after water stress. The ratios were used as comparative estimates of cell wall thickness which affects cell wall elasticity (Wilson et al., I98O).
D. Results
Plants infected with either pathogen and grown in nutrient solution exhibited lower leaf water and turgor Figure 5*1* Leaf weight versus leaf water potential (A) for a single leaf of healthy P. pratensis 'Merion' grown in nutrient solution to determine leaf weight at full turgor. Data are replotted as relative water content versus leaf water potential to yield the moisture release curve (B) which yields estimates of os motic potential (TTp) and relative water content (RWC^) at zero turgor (plasmolysis) 80
LEW WATER POTENTIAL (MPj)
LEAF WEIGHT AT FULL TURGOR
POINT OF INTERSECTION USED AS PIASMOIVSIS
RWC AT ZERO TURGOR (RWCp) 81
potentials than healthy plants during both light and dark periods (Fig. 5.2). Healthy plants regained turgor during the dark period rapidly to prelight values of approximately 1.5 MPa. U. agropvri- and U. striiformis- infected plants regained turgor less rapidly to prelight values of approximately 0.8 MPa and 0.5 MPa, respectively. Healthy plants subjected to progressively greater water stress were able to regain higher prelight leaf water and turgor potentials than U. striiformis- or U. agropyri-infected plants (Fig. 5*3)• Systemic infection by either pathogen resulted in near equal reductions in prelight leaf water and turgor potentials over the entire range of water stress (Fig. 5,3). Plants grown in nutrient solution and infected by U. striiformis exhibited lower mean osmotic potentials at zero turgor than leaves from healthy plants (Table 5.1). Plants infected by U. agropyri exhibited decreased mean relative water content at zero turgor and increased turgid weight/dry weight ratios compared to leaves from healthy plants (Table 5.1). Water stress treatments induced osmotic adjustment of healthy and infected leaves (Table 5.1). Leaves for healthy plants, however, exhibited lower osmotic potentials at zero turgor and lower turgid weight/dry weight ratios than infected plants after water stress treatments. Mean relative water contents of leaves Figure 5»2. Leaf water and turgor potentials of healthy {•), U. striiformis-infected (stripe-smutted) (o), and U. agropvri- infected (flag-smutted) (a) leaves of Pea pratensis over a 24-hour cycle from plants growing in a growth chamber in nutrient solution. Each point represents the mean of 3 leaves 83
1.8 1.7 DARK LIGHT 1.6 15 1.4 • • • • . • •• U • • 1.2 • • • 1.1 1.0 • ^ AO ^ s 0.9 0.8 0.7 0.6 05 o* o o 0.4 #0 cf 0.3 A A o 02 QAO^O cf A 0.1 ,0 A, T
-0.1 -0.2 • HEALTHY -0.3 •• • O STRIPE-SMUTTEO -0.4 A RAG-SMUTTED -0.5 -0.6 -0.7 -0.8 -0.9 -1.0 -1.1 -1.2 ^ o .o 8 -1.3 A • -1.4 -15 o A° -1.6 A" O -1.7 - o -1.8 -1.9 A A % A A -2.0 O O -2.1 ' A O -22 -2.3 -2.4 _L I I I I L 2100 2300 0100 0^ OMO 0700 0900 1100 1300 1500 1700 1900 2100 • TIME Figure 5«3« Prelight leaf water and turgor potentials versus nutrient solution osmotic potential of the second fully-expanded leaf from healthy (H), U# striiformis-infected (stripe-smutted) (S), and U. agropvri- infected (flag-smutted) (F) Poa pratensis grown in nutrient solution amended with
PEGqooo* Each point represents the mean of 8 leaves and has been bracketed by its standard error. PREDAWN LEAF WATER POTENTIAL (MPa) PREDAWN LEAF TURGOR (MPa)
roiliiiliiiiil ro ro • i—' >—• •—* ;—' CD CD CZ) o c—CD CD CD I—» i—• i—» #—< i—> r>o (=) bo (3") 4=» ro o bo ctî Ks o ro ^ cr> bo o ro 4s. ct> bo
OO
C/3 <=> o
iI—
ro C/3 Table 5.1. Mean osmotic potentials (MPa) and relative water contents (%) at zero turgor and mean turgid weight/dry weight ratios of leaves from healthy, U. striiformis-. and U. agropyri-infected Poa pratensis 'Merion' before and after water stress treatment
Health group Before Stress Treatment After Stress Treatment TTp KWCp TW/DW° -JXp RWCp TW/DW
Healthy -1.33 93.1 3.39 -2.72 78.9 2.87
Infected by U. striiformis -1.54 91.0 3.36 -2.21 78.1 3.26
uT^I2roDV^l -1.40 89.3 3-85 -2.10 77.6 3.69
LSD(O^05) 0.15 2.4 0.30 0.42 3.1 0.46
^Osmotic potentials at zero turgor (plasmolysis). ^Relative water content at zero turgor (plasmolysis), °Turgid weight/dry weight ratios. 87
at zero turgor after water stress treatment were not significantly different between healthy and infected plants (Table 5*1)•
E, Discussion
Infection of P. pratensis by U. agropyri or U. striiformis resulted in pathogen-induced water stress. Infected plants exhibited lower leaf water and turgor potentials during the light and less rapid recovery during the dark to prelight levels that were substantially below healthy plants (Fig. 5«2), Pathogen-induced water stress has been demonstrated for other host-pathogen systems (Ayres, 1978» Talboys, I968; Kozlowski, 1976b; Van der Wal et alt, 1975)» and can arise from reduced water absorption due to root growth inhibition, reduced water transport due to vascular blockages, loss of cell membrane integrity, or accelerated water loss due to disruption of normal stomatal function or epidermal rupturing (Ayres, 1978; Dimond, 1970; Kozlowski, 1976). Infection by U. agropyri and U. striiformis de creases root growth of P. pratensis (see Chapter III). Infected plants are subsequently limited in the volume of soil from which they can extract moisture compared to healthy plants. In addition, the hyphae of both 88
pathogens grow between mesophyll cells and, with the formation of teliospores, rupture mesophyll and epidermal cells (Hodges.and Britton, 1970), which may lead to accelerated water loss. The combination of root growth inhibition and epidermal rupturing leading to accelerated water loss may explain the lower leaf water and turgor potentials seen in Figure 5«2. The ability of cell membranes to retain solutes is reflected, in part, by the degree of osmotic adjust ment for turgor maintenance in response to water stress. As plants were subjected to progressively more severe water stress, healthy plants regained higher prelight leaf water and turgor potentials than plants infected with either U. striiformis and U. agropyri (Fig. 5*3)• Turgor potential at a given water potential is dependent on the osmotic potential and elasticity of the tissue (Turner, 1979). It seems, therefore, that leaf-smutted plants are less able to adjust osmotically or that tissue elasticity may be adversely affected by infection. Quantitative assessment concerning the degree of osmotic adjustment in healthy versus infected plants, however, cannot be made from data presented in Figure 5«3 because methods for turgor potential determination involving expressed sap are subject to errors by symplastic dilution by apoplastic water and possible differences in 89
tissue hydration (Brown and Tanner, 1983). Infection by either U. striiformis or U. agropvri results in lower leaf water and turgor potentials of P. pratensis (Fig. 5*2).. This suggests that P. pratensis is capable of osmotic adjustment in response to water stress induced by either pathogen as reflected by the lower values for osmotic potential at zero turgor (Table 5» 1). Osmotic adjustment of the host also would explain the lower relative water contents at zero turgor exhibited by infected plants. Healthy and infected P. pratensis showed evidence for osmotic adjustment of leaves in response to osmotically- induced water stress (Table 5*1)• Infection by either pathogen, however, decreased the degree of osmotic adjustment by P. pratensis leaves. Leaves from healthy plants were able to osmotically adjust to lower values than leaves from flag or stripe smutted plants (Table 5« 1). Although plants use a number of osmotica to lower their osmotic potentials in response to water stress (Hsiao, 1973î Borowitzka, 1981), soluble sugars are a main component, Hodges and Robinson (1977) and Madsen et al. (1983) have shown that infection by either pathogen results in decreased levels of soluble sugars in P. pratensis leaves. This may partially account for the decrease in degree of osmotic adjustment exhibited by 90 infected plants. Infection by U. agropyri results in a sharp decrease in leaf dry weight compared to healthy or U. striiformis-infected plants (see Chapter III). Higher turgid weight/dry weight ratios may be a result of this, and provide evidence for thinner cell walls compared to those of healthy or U. striiformis-infected plants (Wilson et al., 1980). Thicker cell walls are positively correlated with greater drought resistance (Turner, I979). 91
VI. COMPARATIVE WATER USE BY HEALTHY AND USTILAGO STRIIFORMIS-INFECTED POA PRATENSIS
A. Abstract
Pea pratensis 'Merion' systemically infected by Ustilago striiformis was evaluated for water economy and stomatal action. Infected plants with leaves having little or no visible sporulation of teliospores and healthy plants did not differ in rate of water use (ml day"^) or water use efficiency (ml mg~^). Infected plants with leaves having moderate to heavy sporulation exhibited a sharp decrease in water use efficiency and per day use of water lower than healthy plants or infected plants with leaves having non-sporulating sori. Stomatal closure on infected leaves with non- sporulating sori occurred at lower leaf water potentials than on healthy leaves suggestive of a degree of osmotic adjustment.
B. Introduction
Plant pathogens have been shown to interfere with the water status of their hosts (Ayres, 1978; Talboys, 1968; See Chapter V). Pathogen-induced inhibition of 92
root growth (Murphy, 1935; Johnston and Miller, 193^; 1940; Hodges, 1969; 1970a; See Chapter III) and vascular blockages (Dimond, 1970) can limit water uptake. Water loss may be accelerated by decreased cuticular resistance (Sempio et al., I966; Amatya and Jones, I966; Gerwitz and Durbin, 19^5) or inhibition of stomatal movement (Duniway and Durbin, 1971b, Farrell et al., 1969; Ayres and Jones, 1975)* Transpiration is the sum of stomatal and non- stomatal water loss (Crafts, I968). Stomatal resis tances are derived from differences between water lost in light and darkness because it is assumed that stomata close in the dark and that cuticular resistance is not affected by illumination (Ayres, 1978). Because stomatal closure is the main cause for transpiration decline as water stress develops, control of stomatal aperture is very important in drought resistance (Hsiao, 1973; Townley-Smith and Hurd, 1979). However, even if the stomatal apparatus is sensitive to water deficits, and the leaf water potential at which stomata close is high, full benefit is gained only if the cuticular resistance is also high and stomatal closure is com plete (Turner, 1979)' Ustilago striiformis is the causal organism for stripe smut and is a pathogen of several grass species 93
including Poa pratensis (Couch, 1971). It is a member of the Ustilaginales and produces thick-walled telio- spores in sori on leaves, sheaths, and floral parts of infected plants (Couch, 1971). Sporulation of telio- spores ruptures the epidermis of the host and may result in rapid water loss in the area of the sorus. In closely related rust diseases, transpiration by in fected plants is often reduced early in the infection cycle because of inhibition of stomatal activity (Duniway and Durbin, 1971b; Farrell et al., 1969; Ayres, and Jones, 1975). After sporulation, however, water economy is reduced due to epidermal rupturing and a decrease in cuticular resistance (Ayres, 1978). A direct measure of the plants water economy is water use efficiency and has been used to evaluate drought resistance (Stone, 1975, Pierre et al., 1966} Townley- Smith and Kurd, 1979). This study was initiated to test the effects of systemic infection by Ustilago striiformis on the water use, water use efficiency, leaf diffusive resistance, and stomatal activity of Poa pratensis 'Merion'.
C. Materials and Methods
A single clone of healthy or U, striiformis-infected P. pratensis 'Merion' was vegetatively-propagated from 94
individual shoots and grown in a loam :peat:perlite (1:1:1) mix under greenhouse conditions. Individual shoots selected for experiments were removed from the soil mix, roots were washed, and all but the two youngest fully-expanded leaves were stripped from the shoot. Individual shoots were then placed in nutrient culture jars (see Chapter III) and grown in a growth chamber + —2 with a 12-hour photoperiod (65 - 5 nE cm" sec ), $0% relative humidity, and 23 (-2) C. 1, Water use and efficiency Plants were stripped of all but the three youngest fully-expanded leaves and carefully selected for size uniformity after three weeks growth in the growth chamber. U. striiformis-infected plants were separated into two groups: (1) plants with leaves with immature sori with little or no sporulation of teliospores and without epidermal rupturing, and (2) plant with leaves with mature sori and teliospore sporulation that had ruptured the epidermis. Fifteen healthy plants and U. striiformis- infected plants were chosen at random and oven-dried (60 C, 48 hours). Initial mean dry weights (IMDW) were recorded for both healthy and infected plants. An additional fifteen healthy and infected plants were placed individually in aerated nutrient culture jars with 100 ml of nutrient solution (see Chapter III), 95
and sealed with silicone grease (Dow Corning, Midland, Mich.). Plants were grown two weeks and nutrient solution was flushed twice per week. Total water loss was recorded at each nutrient solution flush which included both water usage by the plant and evaporation from the solution aeration exhaust port (see Chapter III). Evaporation from the aeration exhaust port was determined by running the same experiment at equal aeration rates without plants and the rubber stoppers sealed with sili cone grease. Evaporation was recorded, and plant water usage was calculated as total water loss minus evaporation from the aeration exhaust port. All plants were oven-dried (60 C, 48 hours) at the end of the two week growth period. Initial mean dry weights (IMDW) for healthy and infected plants were subtracted from each healthy or infected plant, re spectively. Estimates for dry matter produced during the two week period were thus obtained. Water use efficiency was derived by dividing dry matter produced by plant water usage for each plant (Kramer, 1980). 2. Leaf diffusive resistance A Li-Cor leaf diffusive resistance meter (Model LI-60) was used to measure diffusive resistance of healthy leaves and leaves with non-sporulating and sporulating sori of plants grown in nutrient solution. 96
Measurements were taken after greenhouse-grown plants were selected for size uniformity at a fully-expanded two leaf stage and grown in a growth chamber for three weeks. Growth chamber conditions were as pre viously stated. Data were plotted as diffusive re sistance of healthy leaves and leaves with non-sporulating and sporulating sori over a 24-hour period. Each point represents the mean of three measurements. Leaf diffusive resistance also was recorded as a function of leaf water potential for healthy and in- fected plants. Twenty 6-cm plastic pots of healthy and infected plants growing in a 1:1:1 (loam:peat: perlite) mix were taken from the greenhouse and placed in the growth chamber. Plants were watered daily with distilled water and fertilized with nutrient solution at 5 day intervals. After two weeks, pots were allowed to dry and leaf diffusive resistance and water potential measurements were initiated. Leaf diffusive resistance measurements were taken only during the light cycle and only on the second fully-expanded leaf blade. Humidity sensor placement on U. striiformis-infected leaves was restricted to non-sporulated areas of the sori. Leaf blades were then sealed with petroleum jelly and excised. Leaf water potential measurements were obtained using a
PMS pressure bomb at a pressurization rate of 0,01 MPa sec~\ 97
Data were plotted as leaf diffusive resistance versus leaf water potential for healthy leaves and infected leaves with non-sporulated areas of the sori.
D. Results
Leaves of plants infected with U. striiformis showing little or no sporulation or epidermal rupturing of sori did not differ in water use from healthy plants (Fig. 6.1A). Infected leaves of plants with moderate to severe epidermal rupturing of sori used less water than leaves of healthy plants or infected leaves with non- sporuxating sori (Fig. 6.1A). Dry matter accumulation (mg day"^) of P. pratensis decreased as leaf sori matured and epidermal cell rupturing increased (Fig. 6.IB). Leaves of infected plants with immature sori and little or no epidermal damage did not differ in dry matter accumulation from healthy plants, but leaves of infected plants with mature sori and moderate to severe epidermal damage decreased in dry matter accumulation compared to healthy or U. striiformis- infected plants with immature sori. No differences were found in the water use ef ficiency between healthy plants and infected plants with immature leaf sori (Fig. 6.1C). Infected plants with Figure 6.1. Water use (A), dry matter production (B), and water use efficiency (C) of healthy and Ustilago striiformis-infected Poa pratensis 'Merion* leaves with immature (non-sporulating) and mature (sporulating) sori WATER USE EFFICIENCY (mg/ml) DRY MATTER PRODUCTION (mg/day) WATER USE (mis/day)
1 1 1 1 1 1 i 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1 1
h -4 t- -4 so\o
9 t- 4 t- H h H
l-H t-4 ^ 4
o OD > 100
mature leaf sori and moderate to severe epidermal damage were less water efficient that healthy plants or infected plants with immature leaf sori. No differences were observed in leaf diffusive resistance between healthy plants and infected plants with mature or immature leaf sori during the 12-hour photo-period (Fig. 6.2). Leaf diffusive resistance in the dark was affected by infection on leaves with both im mature and mature sori (Fig. 6.2). Infected P. pratensis with leaves having mature sori exhibited little resis tance to water loss in the dark (Fig. 6.2). However, infected leaves with immature sori (non-sporulating) also exhibited slightly less leaf diffusive resistance in the dark than leaves of healthy plants (Fig. 6.2). Leaf diffusive resistances of healthy leaves ex hibited rapid increases at leaf water potentials between -1.2 and -1.7 MPa (Fig, 6.3). Immature portions of sori on infected leaves exhibited sharp increases in leaf diffusive resistances at leaf water potentials of -2.1 to -2.4 MPa.
E. Discussion
The decreased water use per day exhibited by U. striiformis-infected plants with mature leaf sori seems Figure 6.2, Leaf diffusive resistance of healthy (#), and immature (non-sporulated) (Q) and mature (sporulated) (A) sori of Ustilago striif ormis - infected Poa pratensis 'Merion' grovmin nutrient solution DARK LIGHT 100
90 • • 80 o •o ° . • 70 o • HEALTHY o * • o o STRIPE (NON-SPORULATED) o 60 o o A STRIPE (SPORULATED) o o 50 o 40
30
20 8 4 # A A A'A A o •A 8 ^o# J L J I L I I I ± _L JL 00 2300 0100 0300 0500 0700 0900 1100 1300 1500 1700 1900 2100 TIME Figure 6.3» Leaf diffusive resistance versus leaf water potential of healthy Poa pratensis 'Merlon' leaves and Ustilago striiformis- infected leaves with immature, non- spcrulating sori 104
56- 54 • 52 • 50- 48 - 46 - • HEALTHY 44 - o STRIPE-SMUTTED (NON-SPORULATED) 42 - o o 40 - 38 - 35- o 34 - o o 32 - • • 30 - o 28- 26 - 24 - o 22 - o o 20 - 18 - o o 16 - o 14 - 12 - 0 o ° • °
10 - oO °*o.o * • 8 - o '.o. . •. 6 - 4 -
2 -
_J I I I I I I I I I I L -2.6 -2.4 -2.2 -2.0 -1,8 -1.6 -1.4 -1.2 -1.0 -08 -06 -0.4 -0.2 LEAF WATER POTENTIAL (MPa) 105
inconsistent with the ruptured epidermal cells associated with mature sori (Fig. 6.1A and 6.2). A decrease
in water use also has been observed in rusted wheat (Johnston and Miller, 193^; 1940), oats (Murphy, 1935)» and barley (Sever, 1937)» The dry matter accumulation of infected plants with mature leaf sori was decreased reflecting less cell division and expansion; these responses would decrease water use and partially explain the decrease in water use per day. Infection by U. striiformis also results in decreases in leaf water potential in P. pratensis leaves (see Chapter V). Such pathogen-induced water stress in U. striiformis- infected plants may result in stomatal closure on leaves of infected plants with mature sori. This also could contribute to a decrease in dry matter accumulation due to limited carbon dioxide fixation (Burrows and Milthorpe, 1976). Water use efficiency is a direct measure of the water economy of the plant (Kramer, 1980), Water use (ml day""^) of infected plants with mature leaf sori decreased compared to healthy plants, but, because dry matter production decreased even more, water use efficiency (mg ml"^) also decreased (Pig. 6.IB and 6,1C). Water use efficiency reflects drought resistance (Townley-Smith and Kurd, 1979)* The sharp decreases in 106
water use efficiency of infected plants with mature leaf sori may partly explain the increase in mortality of diseased plants during periods of limited soil moisture (Hodges, 1977). Leaf diffusive resistance of infected plants with immature leaf sori and a non-ruptured epidermis was not as great as that of healthy leaves in the dark. Decreased leaf diffusive resistance in the dark may arise from decreased cuticular resistance or incomplete stomatal closure (Sempio et al., I966; Duniway and Durbin» 1971b). Other pathogens have been shown to directly interfere with stomatal action (Farrell et al., 1969; Ayres, 1972; Cruickshank and Rider, I96I). This also may apply to U, striiformis in P. pratensis. The presence of the pathogen's hyphae or teliospores within the mesophyll and beneath the non-ruptured epidermis may affect normal cuticle formation or function in infected leaves and cause a decrease in cuticular resistance to water loss. The comparative leaf diffusive resistance of healthy leaves versus immature portions (non-sporulated) of leaf sori on infected plants suggests the stomates associated with immature leaf sori of infected leaves close at lower leaf water potentials than those on healthy leaves. Stomatal conductance is turgor dependent (Hsiao, 1973)» and closure occurs when bulk leaf turgor is near zero 107
(Burrows and Milthorpe, I976). Stomatal closure at lower leaf water potentials is evidence for pathogen- induced water stress, P. pratensis can osmotically adjust to the water stress imposed by U, striiformis and maintain turgor and stomatal aperture (See Chapter V) at lower leaf water potentials than healthy plants. Rapid increases in leaf diffusive resistances at lower leaf water potentials in infected plants compared to healthy plants is not evidence for direct pathogen inhibition of stomatal movement. Rather, decreased stomatal sensitivity in U, striiformis-infected plants seems to suggest that the host reacts to pathogen- induced water stress by osmotic adjustment. 108
VII. CONCLUSIONS
Growth responses of Poa pratensis suggest that infection by either Ustilago striiformis or Urocystis agropvri disrupts normal hormonal balance of the host, but in different ways. Growth responses of stripe- smutted plants are suggestive of growth promoting ac tions, whereas, flag smut results in dwarfing of P. pratensis. a proliferation of tillering, and virtual elimination of rhizome growth. Because so little is
known about hormonal control of tillering versus rhizome
growth, the Poa pratensis-Urocvstis agropyri complex may prove to be an excellent system to study such dif ferential control. Overall growth as an integrated process can also serve as a measure of the plant * s tolerance during water stress. Healthy plants are better able to main tain overall growth as water stress increases which sug gests healthy plants possess greater drought tolerance than infected plants. Turgor pressure is the driving force for growth, therefore better maintenance of growth during periods of water stress implies healthy plants have a higher water status than infected plants. This is true for P. pratensis infected with U. agropyri or U. striiformis. 109
There are several reasons why healthy plants are able to maintain a higher plant water status and tolerate water stress better than infected plants. Infection by either pathogen greatly decreases root growth of P. pratensis which leads to lower root-shoot ratios of in fected plants. Infection by either pathogen inhibits the ability of P. pratensis leaves to osmotically adjust to water stress. The increase in cell wall thickness in response to water stress of P. pratensis leaves as measured by turgid weight/dry weight ratios is inhibited by infection by either pathogen. Thinner cell walls limit the amount of leaf turgor for a given leaf water potential. Lastly, as sori from infected leaves mature and leaf epidermal cells are ruptured, non-stomatal water loss is greatly increased, and water use efficiency is subsequently reduced sharply. The combination of these actions predispose infected plants to desiccation during periods of water stress and results in increased mortality of diseased plants. 110
VIII. LITERATURE CITED
Acevedo, E., T. C. Hsiao, and D. W. Henderson. 1971. Immediate and subsequent growth responses of maize leaves to changes in water status. Plant Physiol. 48:631-636. Aokerson, R. G., D. R, Krieg,: and F. J. M. Sung. I98O. Leaf conductance and osmoregulation of field- grown Sorghum genotypes. Crop Sci. 20:10-14. Ackerson, R. C., D, R. Krieg, T. D. Milley, and R. E. Zartman. 1977t Water relations of field grown cotton and sorghum: Temporal and diurnal changes in leaf water, osmotic, and turgor potentials. Crop Sci. 17:76-80. Agrios, G. N. I969. Plant Pathology. Academic Press, New York. Allaway, W. G., and F, L. Milthorpe. 1976. Structure and functioning of stomata. Pages 57-102 in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 4. Academic Press, New York. Alvim, P. de T. I966. Stomatal opening as a practical indicator of moisture stress in cotton. Physiol. Plant. 19008-312. Amatya, P., and J. P. Jones. I966, Transpiration changes in oat plants infected with crown rust. Proc. Ark. Acad. Sci. 20:59-67. Anderson, D. B., and T. Kerr. 1943. A note on the growth behaviour of cotton bolls. Plant Physiol. 18:261-262. Ashby, E., and R. Wolf. 1947. A critical examination of the gravimetric method of determining suction force. Ann. Bot. 11:261-270. Ashton, P. 1956. Effects of a series of cycles of alternating low and high soil water contents on the rate of apparent photosynthesis in sugar cane. Plant Physiol. 31:266-270. Ill
Aspinall, D,, P. B. Nicholls, and I, H. May. 1964. The effects of soil moisture stress on the growth of barley. I. Vegetative development and grain yield. Aust, J, Agric. Res. 15:729-737. Ayres, P. G. 1978. Water relations of diseased plants. Pages l-60 in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 5« Academic Press, New York. Ayres, P. G. 1977' Effects of leaf water potential on sporulation of Ervsiphe pisi (pea mildew). Trans. Br. Mycol. Soc. 68:97-100. Ayres, P. G. 1976. Patterns of stomatal behaviour, transpiration, and GG2 exchange in pea following in fection by powdery mildew (Ervsiphe pisi). J. Exp. Bet. 27Î35^-363. Ayres, P. G. 1972. Abnormal behaviour of stomata in barley leaves infected with Rhvnchosporium sec'alis. J. Exp. Bot. 23s683-691. Ayres, P. G., and P. Jones. 1975. Increased traspiration and the accumulation of root absorbed 86r6 in barley leaves infected by Rhvnchosporium secalis (leaf blotch). Physiol. Plant Pathol. 7:49-58. Barlow, E. W. R., R. E. Munns, and 0. J. Brady. 1980. Drought responses of apical meristems. Pages I9I- 206 in N. G. Turner and P. J. Kramer, editors. Adaptation of Plants to Water and High Temperature Stress. John Wiley and Sons, New York. Barlow, E. W. R., R. E. Munns, N. S. Scott, and A. H. Reisner. 1977» Water potential, growth, and poly ribosome content of the stressed wheat apex. J. Exp. Bot. 28:909-925* Barrs, H. D. I968. Determination of water deficits in plant tissues. Pages 235-368 in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 1. Academic Press, New York. Barrs, H. D. 1964. Heat of respiration as a possible source of error in the estimation by psychrometric methods of water potential in plant tissue. Nature 203,1136-1142. 112
Baver, L. D., W. H. Gardner, and W, R. Gardner. 1972. Soil Physics. Fourth Edition. John Wiley and Sons, New York. Beadle, 0. L., K.-R. Stevenson, H. H. Neumann, G. W, Thurtell, and K. M. King. 1973. Diffusive resistance, transpiration, and photosynthesis in single leaves of corn and sorghum in relation to leaf water potential. Can. J. Plant Soi. 53:537-544. Beard, J. B. 1973* Turfgrass: Science and Culture, Prentice-Hall, Englewood Cliffs, N. J. Beardsell, M. F., P. G. Jarvis, and B. Davidson. 1972. A null-balance diffusion porometer suitable for use with leaves of many shapes. J. Appl. Ecol. 9:677-690. Beckman, C. H., W. A. Brun, and I. W. Buddenhagen. I962. Water relations in banana plants infected with Pseudomonas solanacearum. Phytopathology 52:1144-1147. Begg, J. E. 1980. Morphological adaptations of leaves to water stress. Pages 33-43 in N. C. Turner and P. J. Kramer, editors. Adaptation of Plants to Water and High Temperature Stress. John Wiley and Sons, New York. Begg, J. E., J. P. Bierhuizen, E, R. Lemon, D. K. Misra, R. 0. Slatyer, and W. Stern. 1964. Diurnal energy and water exchanges in bullrush millet in an area of high solar radiation. Agric. Meteorol. 1:294-310. Begg, J. E., and N. C. Turner. 1976. Crop water deficits. Advan. Agron. 28:161-217. Bennett, 0. 1., and B. D. Doss. i960. Effect of soil moisture level on root distribution of cool-season forage species. Agron. J. 52:204-207. Bernstein, L. I96I. Osmotic adjustment of plants to saline media. I. Steady state. Am. J. Bot. 48:909- 918. Bertrand, A. R. I966. Water conservation through improved practices. Pages 207-235 in W. Pierre, D. Kirkham, J. Pesek, and R. Shaw, editors. Plant Environment and Efficient Water Use. Am. Soc. Agron., Madison, Wise. 113
Sever, W. M. 1937. Influence of stripe rust on growth, water economy and yield of wheat and barley» J. Agric. Res. 5^075-385. Bewley, W, F. 1922. Sleepy disease of tomato. Ann. Appl. Biol. 9s116-126. Bingham, G. E., and P. I. Coyne. 1977* A portable temperature controlled steady-state porometer for field measurements of transpiration and photosyn thesis. Photosynthetica 11:148-160. Biscoe, P. V, 1972. The diffusion resistance and water status of leaves of Beta vulgaris, J. Exp. Bot. 23:930-940. Biscoe, P. v., M. H. Unsworth, and H. R. Pinckney. 1973- The effects of low concentrations of sulphur dioxide on stomatal behaviour in Vicia faba. New Phytol. 72:1299-1306. Black, C, A., D. D. Evans, J. L. White, L. E. Ensminger, and F. E. Clark (eds.). I965. Methods of Soil Analysis. Am. Soc. Agron., Madison, Wise. Blum, A. 1979» Genetic improvement of drought resistance in crop plants: A case for Sorghum. Page 429-446 in H. Mussell and R. C. Staples, editors. Stress Physiology in Crop Plants. John Wiley and Sons, New York. Blum, A. 1975' Effect of the Bm gene on epicuticular wax and the water relations of Sorghum bicolor. Isr, J. Bot. 24:50 (Abstr.). Blum, A., and M. Naveh. 1976. Improved water use efficiency in dryland grain sorghum by promoted plant competition. Agron. J. 68:111-116. Borowitzka, L. I98I. Solute accumulation and regulation of cell water activity. Pages 97-130 in L. G. Paleg and D. Aspinall, editors. The Physiology and Biochemistry and Drought Resistance in Plants. Academic Press, New York.
Boyer, J« S. 1976. Water deficits and photosynthesis. Pages I53-I9O in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 4. Academic Press, New York. 114
Boyer, J. S. 1971» Resistances to water transport in soybeans, bean, and sunflower. Crop Soi. 11:403- 407. Boyer, J. S. 1970. Leaf enlargement and metabolic rates of corn, soybean, and sunflower at various leaf water potentials. Plant Physiol. 46Î233-235. Boyer, J. S. 1969. Measurement of the water status of plants. Ann. Rev. Plant Physiol. 20:3^1-364. Boyer, J. S. I968. Relationship of water potential to growth of leaves. Plant Physiol. 43:10^6-1062. Boyer, J. S. I967. Matric potential of leaves. Plant Physiol. 42:213-217. Boyer, J. S. I966. Leaf water potentials measured with a pressure chamber. Plant Physiol. 42:133-137. Boyer, J. S. I965. Effects of osmotic water stress on metabolic rates of cotton plants with open stomates. Plant Physiol. 40:229-234. Boyer, J. S., and H. G. McPherson. 1975* Physiology of water deficits in cereal crops. Adv. Agron. 27:1-23. Brady, A, P., H. Huff, and J. W. McBain. 1951» Measure ment of vapor pressures by means of matched thermis tors. J. Phys. Colloid Chem. 55s304-311. Brian, P. W., C. W. Elson, H. G. Hemming, and M. Radley. 1954. The plant growth promoting properties of gibberellic acid, a metabolic product of fungi Gibberella fu.iikaroi. J. Soi. Food Agric. <^602-6l2. Brouwer, R. I965. Water movement across the root. Symp. Soc. Exp. Biol. 19:131-152. Brown, K. W., W. R. Jordan, and J. C. Thomas. 1976. Water stress induced alterations of the stomatal response to decreases in leaf water potential. Physiol. Plant. 37:1-5. Brown, P. W,, and C. B. Tanner. 1983. Alfalfa osmotic potential: A comparison of the water-release curve and frozen-tissue methods. Agron, J. 75«91-93» 115
Brown, R. W., and B. P. Van Haveren (eds.). 1972. Psychrometry in Water Relations Research. Utah Agric. Ejcp. Stn. , Logan, Utah. Bukovac, M. J., and R. F, Norris. I967. Foliar penetration of plant growth substances with special reference to binding by cuticular surfaces of pear leaves. Agrochimica 12:296-300.
Burrows, F, J., and F. L. Milthorpe. 1976. Stomatal conductance in the control of gas exchange. Pages IO3-I53 in T. r. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 4. Academic Press, New York. Burton, G. W., G. M. Prine, and J. E. Jackson. 1957- Studies of drought tolerance and water use of several southern grasses, Agron. J. 49:498-503. Campbell, E. C., G. S. Campbell, and W. K. Barlow. 1973« A dewpoint hygrometer for water potential measurement. Agric. Meteorol. 12:113-121. Campbell, G. S«, and M. D. Campbell. 1974. Evaluation of a thermocouple hygrometer for measuring leaf water potential.in situ. .Agrom. J. 66:24-27. Campbell, G. S., R. I. Papendick, E. Rabie, and A. J. Shayo-Ngowi. 1979. A comparison of osmotic potential, elastic modulus, and apoplastic water in leaves of dryland winter wheat, Agron. J. 71:31-36. Chessman, J. H., E. 0. Roberts, and L. H. Tiffany. 1965» Effects of nitrogen level and osmotic pressure of the nutrient solution on incidence of Puccinia graminis and Helminthosporium sativm infection in Merion Kentucky bluegrass, Agron. J. 57:599-602. Cheung, Y. N. S., M. T. Tyree, and J. Dainty. 1976. Some possible sources of error in determining bulk elastic moduli and other parameters from pressure volume curves of shoots and leaves. Can. J. Bot. 54:758-765»
Cheung, Y. N. S., M. T. Tyree, and J. Dainty. 1975» _ Water relations parameters on single leaves obtained in a pressure bomb and some ecological interpretations. Can. J. Bot. 53:1342-1346. 116
Chu, A. C. P., and H. G. McPherson. 1977. Sensitivity to desiccation of leaf extension in prairie grass. Aust. J. Plant Physiol. 4:381-387. Clements, H. F. 1964. Interaction of factors affecting yield. Annu, Rev. Plant Physiol. 1^:409-442. Clough, B. P., and F. L. Milthorpe. 1975* Effects of water deficit on leaf development in tobacco. Aust. J. Plant Physiol. 2:291-300. Connor, D. J., and B. R. Tunstall. 1968. Tissue water relations for brigalow and mulga. Aust. J. Bot. 16:487-490. Cook, R. J. 1973' Influence of low plant and soil water potentials on diseases caused by soilborne fungi. Phytopathology 63«451-458. Cook, R. J., and J. M. Duniway. I98I. Water relations in the life-cycles of soilborne plant pathogens. Pages 119-140 in J. F. Parr, W. R. Gardner, and L. F. Elliott, editors. Water Potential Relations in Soil Microbiology. Soil Soi. Soc. Am.,.Madison, Wise. Cook, R. J., and R. I. Papendick. 1978. Role of water potential in microbial growth and development of plant disease with special reference to post harvest pathology. HortScience 13!559-564. Cook, R. J., and R. I. Papendick. 1972. Influence of water potential of soils and plants on root disease. Annu. Rev. Phytopathol. 10:349-374. Couch, H. B. 1971. Disease of Turfgrasses. 2nd Edition. Reinhold Pub. Co., New York, Couch, H. B., and J. R. Bloom. I96O, Influence of en- fironment on diseases of turfgrasses. II. Effect of nutrition, pH, and soil moisture on Sclerotinia dollar spot. Phytopathology 50'761-763. Couch, H. B,, L. H. Purdy, and D. W. Henderson. I967. Application of soil moisture principles to the study of plant disease. Va. Polytech. Inst. State Univ., Dept. Plant Pathol., Bull. No. 4. Cowan, I. R. 1978. Stomatal behaviour and environment. Adv. Bot. Res. 4:117-228. 117
Cowan, I. R. I965. Transport of water in the soil- plant-atmosphere system. J. Appl. Ecol. 2:221-226. Cowan, I. R., and F. L. Milthorpe, I968. Plant factors influencing the water status of plant tissues. Pages 137-194' in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 1. Academic Press, New York. Crafts, A. S, I968. Water deficits and physiological processes. Pages 85-133 in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 2. Academic Press, New York. Crafts, A. S. 1939. Solute transport in plants. Science 90:337-340. Crafts, A. S., and C. L. Foy. I962. The chemical and physical nature of plant surfaces in relation to the use of pesticides and their residues. Residue Rev. 1. Crafts, A. S., H. B. Currier, and C. R. Stocking. 1949. Water in the Physiology of Plants. Chronica Botanica, Waltham, Mass. Cruickshank, I. A. M., and N. E. Rider. I96I. Peronospora tabacina in tobacco: Transpiration, growth, and related energy considerations. Aust. J. Biol. Sci. 14:45-57. Currier, H. B. 1943» Water relations of root cells of Beta vulgaris. Ph. D. Thesis, Univ. of Calif., Berkeley, Calif. Cutler, J. M., K. W. Shahan., and P. L. Steponkus. 1980. Dynamics of osmotic adjustment in rice. Crop Sci. 20:310-314. Cykler, J. F. 1946. Effect of variations in available soil water on yield and quality of potatoes. Agric. Eng. 27:363-368. Dainty, J. 1972. Plant-cell water relations: The elasticity of the cell wall. Proc. R. Soc. Edinburgh 70:89- 93. Dale, J. E. 1958. The use of the infiltration method in the study of the stomata of upland cotton. Empire Cotton Growing Rev. 35:254-261. 118
Darbyshire, B. 1971. Changes in indoleacetic acid oxidase activity associated with plant water poten tial. Physiol. Plant. 25î80-88. Darlington, W. A., and N. Cirulis. 1963. Permeability of apricot leaf cuticle. Plant Physiol. 38:462-46^. Davidson, R. L. 19^9 « Effects of soil nutrients and moisture on root/shoot ratios in Lolium perenne and Trifolium repens. Ann. Bot. 33'571-589.
Davies, W. H. 1922. Urocystis agropyri on redtop. Mycologia 14:279-281. Davies, W. J. 1977» Stomatal responses to water stress and light in plants grown in controlled environments and in the field. Crop Sci. 17:735-740. Davies, W. J., T. T. Kozlowski, and J. Pereira. 1974. Effect of wind on transpiration and stomatal aperture of woody plants. Roy. Soc. N. Z., Bull. 12:433-438. Denmead, 0. T., and R. H. Shaw. I962, Availability of soil water to plants as affected by soil moisture content and meteorological conditions. Agron. J. 54:385-361. Dimond, A. E. 1970. Biophysics and biochemistry of vascular wilt syndrome. Annu. Rev, Phytopathol. 8: 301-322. Donald, C. M. I963. Competition among crop and pasture plants. Adv. Agron. 15s1-118. Doss, B. D., D. A. Ashley, and 0. L. Bennett, i960. Effect of soil moisture regime on root distribution of warm season forage species. Agron. J. 52:569-572. Doss, B. D., 0. L. Bennett, D. A. Ashley, and H. A. Weaver. 1962. Soil moisture regime effect on yield and évapotranspiration from warm season perennial forage species. Agron. J. 54:239-242. Drake, B. G., K. Raschke, and F. B. Salisbury. 1970. Temperature and transpiration resistances of Xanthium leaves as affected by air temperature, humidity, and wind speed. Plant Physiol. 46:324-330» 119
Duggar, W. M., and I. P. Ting. 1970. Air pollution oxidants—their effect on metabolic processes in plants. Annu. Rev. Plant Physiol. 21:21^-224.
Duniway, J. M. 1977» Predisposing effect of water stress on the severity of Phvtophthora root rot in safflower. Phytopathology 67:884-889. Duniway, J. M. 1973« Pathogen-induced changes in host water relations. Phytopathology 63:458-466. Duniway, J. M., and R. D. Durbin. 1971a. Detrimental effect of rust infection on the water relations of bean. Plant Physiol. 48:69-72. Duniway, J. M., and R. D. Durbin. 1971b. Some effects of Uromvces phaseoli on the transpiration rate and stomatal response of bean leaves. Phytopathology 61:114-119. Eagles, C. F., and P. F. Wareing. 1964. The role of growth substances in the regulation of bud dormancy. Physiol. Plant. 17:697-709. Eagles, C. F., and P. F. Wareing. I963. Dormancy regulators in woody plants. Nature 199:874-875* Edmunds, L. K. 1964. Combined relation of plant maturity, temperature, and soil moisture to charcoal stalk rot development in grain sorghum. Phytopathology 54: 514-517. Ehlig, C. P., and W. R. Gardner. 1964. Relationship between transpiration and internal water relations of plants. Agron. J. 56:127-133. El Nadi, A. H., R. Brouwer, and J. T. Locher. I969. Some responses of the root and the shoot of Vicia faba plants to water stress. Neth. J. Agric. Sci. 17:133-152. El-Sharkawy, M. A., and J. D. Hesketh. 1964. Effects of temperature and water deficits on leaf photosynthetic rates of differnt species. Crop Sci, 4:514-518.
Elston, J., A. J. Karmanos, A. H. Kassam, and R. M. Wadsworth. 1976. The water relations of the field bean crop. Philos. Tran. R. Soc. Lond. Ser. B 293«581-591. 120
Ernest, E. C. M. 1931. Suction pressure gradients and the measurement of suction pressure. Ann. Bot, 45:717-725. Farrell, G. M., T. F. Preece, and M. J. Wren. I969. Effects of infection by Phytophthora infestans on stomatal activity of potato leaves. Ann. Appl. Biol. 63:265-275. Pereres, E., E. Acevedo, D. W. Henderson, and T. C. Hsiao. 1978. Seasonal changes in water potential and turgor maintenance in sorghum and maize under water stress. Physiol. Plant. 44:261-267. Fischer, G. W. I940. Fundamental studies of the stripe smut of grasses (Ustilago striifomris) in the Pacific Northwest. Phytopathology 30:93-118. Fischer, R. A. I968. Resistance to water loss in the mesophyll of leek (Allium porrum). J. Exp. Bot. 19:135-145. Fischer, R. A., and N. C. Turner. 1978. Plant productivity in the arid and semi-arid zones. Annu. Rev. Plant Physiol. 29:277-317. Fischer, R. A., and R. M. Hagan. I965* Plant water re lations, irrigation management, and crop yield. Exp. Agric. 1:161-177. Fischer, R. A., T. G. Hsiao, and R. M. Hagan. 1970. After-effect of water stress on stomatal opening potential. J. Exp. Bot. 21:171-185. Frank, A. B., J. P. Power, and W. 0. Willis. 1973. Effect of temperature and plant water stress on photosynthesis, diffusion resistance, and leaf water potential in spring wheat. Agron. J. 65:777-780. Funk, C. R., and R. E. Engel. I963. Effect of cutting height and fertilizer on species composition and turf quality ratings of various turfgrass mixtures. Rutger Univ. Short Course on Turfgrass Management. pp. 47-56. Gale, J., and A. Poljakoff-Mayber. I965. Antitranspirants as a research tool for the study of the effects of water stress on plant behaviour. UNESCO Arid Zone Res. 25:269-280. 121
Gardner, J. L. 1942. Studies in tillering. Ecology 23:162-174. Gardner, W. R., and C. F. Ehlig. 1965- Physical aspects of the internal water relations of plant leaves. Plant Phsyiol. 40:705-710» Gaskin, T. A. 19^5• Varietal resistance to flag smut in Kentucky bluegrass. Plant Dis. Rep. 49:1017» Gates, C. T. I968. Water deficits and growth of herbaceous plants. Pages 145-19® in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 2. Academic Press, New York. Gates, C. T. 1964. The effect of water stress on plant growth. J. Aust. Inst, Agric. Sci. 30:3-22. Gates, C, T. 1955a. The response of the young tomato plant to a brief period of water shortage. I. The whole plant and its principal parts. Aust. J. Biol. Sci. 8:196-214. Gates, C. T. 1955b. The response of the young tomato: plant to a brief period of water shortage. II. The individual leaves. J. Biol. Sci. 215-232. Gerwitz, D. L., and R. D. Durbin. 1965» The influence of rust on the distribution of P32 in the bean plant. Phytopathology 55'57-64. Ghaffar, A., and D. 0. Erwin. 1969» Effect of soil water stress on root rot of cotton caused by Macrophomina phasioli. Phytopathology 59:795-797» Gifford, R. M., and R. B. Musgrave. 1973» Stomatal role in the variability of net 002 exchange rates by two maize inbreds. Aust. J. Biol. Sci. 26:35-^^» Goltz, S. M., and 0. B. Tanner. 1972. Seed onion temperatures and their effects on stomata. HortScience 7:180-181. Goode, J. E., and K. H. Higgs. 1973* Water, osmotic, and pressure potential relationships in apple leaves. J. Am. Soc. Hort. Sci, 48:203-215. Grant, C. L. 1938. Plant structure as influenced by soil moisture. Proc. Ind. Acad. Soi. 48:67-70. 122
Greacen, E. L., and J. S. Oh, 1972. Physics of root growth. Nature (London) New Biol. 235î24-25. Green, P. B., and W. R. Cummins. 1974. Growth rate and turgor pressure. Plant Physiol. 54:863-867. Greenway, H., and M. Leahy. 1970. Effects of rapidly and slowly permeating osmotica on metabolism. Plant Physiol. 46:259-262. Griffin, D. M. 1978. Effect of soil moisture on survival and spread of pathogens. Pages 175-198 in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 5» Academic Press, New York. Grimes, D. W., and H. Yamada. 1982. Relation of cotton growth and yield to minimum'leaf water potential. Crop Sci. 22:134-139. Groenewegen, H., and J. A. Milis. I96O. Uptake of mannitol into the shoots of intact barley plants. Aust. J. Biol. Sci. 13:1-11. Hagan, R. M. 1954. Water requirements of turfgrasses. Proc. 8th Annu. Texas Turfgrass Conf. pp» 9-20, Hall, A. E., E. D, Schulze, and 0. L. Lange. 1976. Current perspectives of steady-state stomatal re sponses to environment. Pages I69-I88 in 0. L, Lange, L, Kappen, and E. D. Schulze, editors. Water and Plant Life: Problems and Modern Approaches, Springer-Verlag, New York. Heath, 0. V, S. 1959* The water relations of stomatal cells and the mechanism of stomatal movement. Pages 193-250 in F, C, Steward, editor. Plant Physiology. Vol. II. Academic Press, New York, Heath, 0. V. S., and B. Orchard. 1956. Studies in stomatal behaviour. VII. Effects of anaerobic conditions upon stomatal movement—a test of William's hypothesis of stomatal mechanism. J. Exp. Bot, 7:313-325. Hellkvist, J,, G. P. Richards, and P, G. Jarvis. 1974. Vertical gradients of water potential and tissue water relations in Sitka spruce trees measured with the pressure chamber. J, Appl. Ecol. 11:637-667. 123
Member, T. 1949. The occurrence of acid inhibitors in resting terminal buds of Fraxinus. Physiol. Plant. 11 s 610-614. Henckel, P. A. 1964. Physiology of plants under drought. Annu. Rev. Plant Physiol. 15:363-386. Henzell, R. G., K. J. McCree, C. H. M. Van Bavel, and K. F. Shertz. 1976. Sorghum genotype variation in stomatal sensitivity to leaf water deficit. Crop Soi. 16:660-662. Hobbs, E. H., K. K. Krogman, and L. G. Sonmor. I963. Effects of levels of minimum available soil moisture on crop yields. Can. J. Plant Sci. 43:441-44$. Hodges, C. F. 1977« Influence of irrigation on survival of Poa pratensis infected by Ustilago striiformis stripe smut and Uroevstis agropvri flag smut. Can. J. Bot. 55:216-218. Hodges, C. F. 1976. Comparative primary infection characteristics of Ustilago striiformis and Uroevstis agropvri on cultivars of Poa pratensis. Phytopathology 66:1111-1115. Hodges, C. F. 1970a. Comparative morphology and develop ment of Poa pratensis infected by Ustilago striiformis and Uroevstis agropvri. Phytopathology 60:1794-1797. Hodges, C. F. 1970b. Directional growth and the perennial characteristic of Ustilago striiformis in Foa pratensis. Phytopathology 60:849-851. Hodges, C. F. 1970c. Influence of temperature on growth of stripe-smutted creeping bentgrass and on sorus development of Ustilago striiformis. Phytopathology 60:665-668, Hodges, C. F. i969. Morphological differences in Kentucky bluegrass infected by stripe smut and flag smut. Plant Dis. Rep. 53!967-968. Hodges, C. F., and M. P. Britton. 1970. Directional growth and the perennial characteristic of Ustilago striiformis in Poa pratensis. Phytopathology 60: 849-851. 124
Hodges, C. F., and M. P. Britton. 1969* Infection of 'Merion' bluegrass (Poa pratensis) by stripe smut (Us til ago striiformis). Phytopathology 59001-304. Hodges, C, F., and P. W. Robinson. 1977. Sugar and amino acid content of Poa pratensis infected with Ustilago striiformis and Urocystis agropyri. Physiol. Plant. 41«25-28. Hodges, J, D., and P. L. Lorio, Jr. 1971. Comparison of field techniques for measuring moisture stress in large loblolly pines. For. Sci. 17:220-223. Hoffman, G. F., S, L. Rawlins, M. J. Garber, and E. M. Cullen. 1971. Water relations and growth of cotton as influenced by salinity and relative humidity. Agron. J. 63:822-826. Hsiao, T. C. 1973. Plant responses to water stress. Annu. Rev. Plant Physiol. 24:519-570. Hsiao, T. C., and E. Acevedo. 1974. Plant responses to water deficits, water use efficiency, and drought resistance. Agric. Meteorol. 14:59-66. Hsiao, T. C., E. Acevedo, E. Fereres, and D, W. Henderson. 1976. Stress metabolism: water stress, growth, and osmotic adjustment. Philos. Trans. R. Soc.(London Ser. B 273:479-500.
Hsiao, Tb C., E. C. Oliveira, and W. M. Hall. 1978» Adaptation of cotton leaves to water stress. Plant Physiol. 61:80(Suppl.). Hull, R. J., N. Jackson, and C. R» Skogley. 1979» Influence of nutrition on stripe smut severity in Kentucky bluegrass turf. Agron. J. 71 s553-555. Husain, I., and D. Aspinall. 1970. Water stress and apical morphogenesis in barley. Ann. Bot. 34:393-407. Iljin, W. S. 1957. Drought resistance in plants and physiological processes. Annu. Rev. Plant Physiol. 8:257-274. Itai, C., A. E. Richmond, and Y. Vaadia. I968. The role of root cyiokinins during water and salinity stress. Isr. J. Bot. 17: I87-I93. 125
Jackson, W. T. I965. Mannitol-induced stimulation of elongation of root hairs of Agrostis alba. Physiol. Plant. 18:24-30. Jarvis, P. G., and M. S. Jarvis. 19^5» The water relations of tree seedlings. V. Growth and root respiration in relation to the osmotic potential of the root medium. Pages I67-I92 B. Slavik, editor. Water Stress In Plants. Czech. Acad. Scil, Prague. Jarvis, P. G., and M. S. Jarvis. I963. The water relations of tree seedlings. IV. Some aspects of the tissue water relations and drought re sistance. Physiol. Plant, 16:501-511. Jarvis, P. G., and R. 0. Slatyer. I966. Calibration of beta gauges for determining water status. Science 153:78-81. Johns, G. G. 1978. Transpirational, leaf area, stomatal, and photosynthetic responses to gradually induced water stress in four warm temperature herbage species. Aust. J. Plant Physiol. 5:113-125. Johnston, C. 0., and E. C. Miller. 1940. Modification of diurnal transpiration in wheat by infections of Puccinia triticina. J. Agric. Res. 61:427-444. Johnston, C. 0., and E. C. Miller. 1934. Relation of leaf rust infection to yield, growth, and water economy of two varieties of wheat. J. Agric. Res. 49 s 955-981. Jones, H. G. 1979- Stomatal behaviour and breeding for drought resistance. Pages 407-428 in H. Mussells and R. C. Staples, editors. Stress Physiology in Crop Plants. John Wiley and Sons, New York, Jones, M. M., and H. M. Rawson. 1979* Influences of rate of development of leaf water deficits upon photosynthesis, leaf conductance, water use efficiency, and osmotic potential in sorghum. Physiol. Plant. 45:103-111. Jones, M. M., and N. C. Turner. I98O. Osmotic adjustment in expanding and fully expanded leaves of sunflower in response to water deficits. Aust. J. Plant Physiol. 7:181-192. 126
Jones, M, M., and N« C. Turner. 1978. Osmotic adjustment in leaves of sorghum in response to water deficits. Plant Physiol. 61:122-126. Jones, M. M., N. C, Turner, and C. B. Osmond. 1981. Mechanisms of drought resistance. Pages 15-38 in L. G. Paleg and D. Aspinall, editors. The Physiology and Biochemistry of Drought Resistance in Plants. Academic Press, Sydney, Australia. Jordan, W. R. 1970. Growth of cotton seedlings in relation to maximum daily plant-water potential. Agron. J. 62:699-701• Jordan, W. R., and J. T. Ritchie. 1971» Influence of soil water stress on evaporation, root absorption, and in ternal water status of cotton. Plant Physiol. 48: 783-788. Kanemasu, E. T., and C. B. Tanner. 1969a. Stomatal diffusion resistance of snap beans. I. Influence of leaf-water potential. Plant Physiol. 44^1^47-1552. Kanemasu, E. T., and C. B. Tanner. 1969b. Stomatal diffusion resistance of snap beans. II. Effect of light. Plant Physiol. 44:1542-1546. Kanemasu, E. T., G. W. Thurtell, and C. B, Tanner. 1969» Design, calibration, and field use of a stomatal diffusion porometer. Plant Physiol. 44:881-885. Kassam, H. H., and J. F. Elston. 1974. Seasonal changes in the status of water and tissue characteristics of leaves of Vicia faba. Ann. Bot. 38:419-429. Kaufmann, M. R. I98I. Water relations during drought. Pages 55-70 in L, G. Paleg and D. Aspinall, editors. The Physiology and Biochemistry of Drought Resistance in Plants. Academic Press, Sydney, Australia. Kaufmann, M. R. 1968a. Evaluation of the pressure chamber technique for estimating plant water potential of forest tree species. For. Soi. 14:369-374. Kaufmann, M. R. 1968b. Water relations of pine seedlings in relation to root and shoot growth. Plant Physiol. 43:281-288. 127
Kaufmann, M. R., and A. N. Eckard, 1971» Evaluation of water stress control with polyethylene glycols by analysis of guttation. Plant Physiol. 47:453- 456. Kaul, R. 1967, A survey of water suction forces in some prairie wheat varieties. Can. J. Plant Sci. 47:323-327. Knipling, E. B. I967. Effect of leaf aging on water deficit-water potential relationships of dogwood leaves growing in two environments. Physiol. Plant. 20:65-72. Kozinka, V., and S. Klenovska. 1965. The uptake of mannitol "by higher plants. Biol. Plant. 7:285-294. Kozlowski, T. T. (ed.). 1978. Water Deficits and Plant Growth. Vol. 5. Academic Press, New York. Kozlowski, T. T. (ed.). 1976a. Water Deficits and Plant Growth. Vol. 4. Academic Press, New York. Kozlowski, T. T, 1976b. Water supply and leaf shedding. Pages 191-232 in T. T, Kozlowski, editor. Water Deficits and Plant Growth. Vol. 4. Academic Press, New York. Kozlowski, T. T. (ed.). 1972. Water Deficits and Plant Growth. Vol. 3. Academic Press, New York. Kozlowski, T, T. 1968a. Introduction. Pages 1-21 T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 1. Academic Press, New York. Kozlowski, T. T. (ed.). 1968b. Water Deficits and Plant Growth. Vol. 2. Academic Press, New York. Kozlowski, T. T. (ed.). 1968c. Water Deficits and Plant Growth. Vol. 1. Academic Press, New York. Kozlowski, T. T. 1964. Water Metabolism in Plants. Harper, New York. Kozlowski, T. T. 1958. Water relations and growth of trees. J. For. 56:498-503.
Kozlowski, T. T., and C. H. Winget. 1964. Diurnal and seasonal variation in radii of tree stems. Ecology 45:149-156. 128
Kramer, P. J. 1980. Drought stress and the origin of adaptations. Pages 7-20 N. C. Turner and P. J. Krmaer, editors. Adaptation of Plants to Water and High Temperature Stress. John Wiley and Sons.,. New York. Kramer, P. J. 1969* Plant and Soil Water Relationships: A Modern Synthesis. McGraw-Hill, New York. Kramer, P. J. I963. Water stress and plant growth. Agron. J. 55i31-37• Kramer, P. J. 1949. Plant and Soil Water Relationships. McGraw-Hill, New York. Kramer, P. J., and H. Brix. 1965. Measurement of water deficits in plants. UNESCO Arid Zone Res, 25043-360. Kramer, P. J., and T. T. Kozlowski. i960. Physiology of Trees. McGraw-Hill, New York. Kreitlow, K. W. 1948. Urocvstis agropvri on Phleum pratense. Phytopathology 38:158-159» Kreitlow, K. W. 1943. Ustilago striiformis II. Temperature as a factor influencing development of smutted plants of Poa pratensis and germination of of fresh chlamydospores. Phytopathology 33:1055- 1063. Kreitlow, K. W., and P. V. Juska. 1959» Susceptibility of Merion and other Kentucky bluegrass varieties to stripe smut (Ustilago striiformis). Agron. J. 51 s 596-597. Lagerwerff, J. V,, G. Ogata, and H. E. Eagle. i96i. Control of osmotic pressure in culture solution with polyethylene glycol. Science 133:1486-1487. Lange, 0. L., L. Kappen, and E. D. Schulze (eds.). 1976. Water and Plant Life: Problems and Modern Approaches. Springer-Verlag, New York. Langer, R. H. M. i963. Tillering in herbage grasses. Herb. Abstr. 33:141-148. Larcher, W. 1975. Physiological Plant Ecology. Springer- Verlag, New York. 129
Lawlor, D. W. 1979- Effects of water and heat stress on carbon metabolism of plants with C3 and 04 photosynthesis. Pages 303-326 ûi H. Mussell and R. C. Staples, editors. Stress Physiology in Crop Plants. John Wiley and Sons, New York. Lawlor, D. W. 1973» Growth and water absorption of wheat with parts of the roots at different water potentials. New Phytol. 72:297-305. Lawlor, D. W. 1970. Absorption of polyethylene glycols by plants and their effects on plant growth. New Phytol. 69:501-513» Leach, J. G., C. V. Lowther, and M. A. Ryan. 19^6. Stripe smut (Ustilago striiformis) in relation to bluegrass improvement. Phytopathology 36:57-72. Lehane, J. J., and W. J. Staple. i962. Effect of soil moisture tension on growth of wheat. Can. J. Soil Sci. 42:180-184. Leopold, A. C. 1949. The control of tillering in grasses by auxin. Am. J. Bot. 36:437-440. Levitt, J. 1972. Responses of Plants to Environmental Stresses. Academic Press, New York. Livne, A., and Y. Vaadia. 1972. Water deficits and hormone relations. Pages 241-275 in T. T, Kozlowski, editor. Water Deficits and Plant Growth. Vol. 3- Academic Press, New York. Lloyd, F. E. i9o8. The physiology of stomata. Carnegie Inst. Wash. Pub. 82. Ludlow, M. M. 1980. Adaptive significance of stomatal responses to water stress. Pages 123-138 in N. C. Turner and P. J. Kramer, editors. Adaptation of Plants to Water and High Temperature Stress. John Wiley and Sons, New York. Lyon, C. J. 1940. Improvements in the simplified method for osmotic measurements. Plant Physiol. 15:561-563* Lyon, C. J. 1936. Analysis of osmotic relations by extending the simplified method. Plant Physiol. 11:167-172. 130
MacDowall, F, D. H. I963. Midday closure of stomata In agin tobacco leaves. Can. J. Bot. 41:1289-1293. Macklon, A. E. S,, and P. E. Weatherley. 1965» Controlled environment studies of nature and origins of water deficits in plants. New Phytol. 64:414-419. Madsen, J. P., C. F. Hodges, and J. L. Nus. i983. Temperature and the soluble sugar content of Poa pratensis infected by Ustllago strilformis or Urocystis agropvri. Bot. Gaz. (in print). Maximov, N. A. I929. The Plant in Relation to Water. Allen and Unwln, London. May, L. H., and F. 1. Milthorpe. I962. Drought resistance in crop plants. Field Crop Abstr. 15:171-191* McCree, K. J. 1974. Changes in the stomatal response characteristics of grain sorghum produced by water stress during growth. Crop Scl. 14:273-278. Mederski, H. J, 1964. Plant water balance determination by beta gauging technique. Agron. J. 56:126-132. Mederski, H. J. i96i. Determination of internal water status of plants by beta ray gauging. Soil Scl. 92:143-147. Meidner, H. 1952. An instrument for the continuous determination of leaf thickness changes in the field. J. Exp. Bot. 3:319-322. Meidner, H., and M. Edwards. 1975» Direct measurements of turgor pressure potentials of guard cells. J. Exp. Bot. 26:319-330. Meidner, H., and T. A. Mansfield. i968. Physiology of Stomata. McGraw-Hill, New York. Meiri, A., and A. Poljakoffr-Mayber. i969. Effects of variations in substrate salinity on the water balance and ionic composition of bean leaves. Isr. J. Bot. 18:99-112. Melkonlan, I. I., J. Wolfe, and P. L. Steponkus. 1982. Determination of the volumetric modulus of elasticity of wheat leaves by pressure-volume relations and the effect of drought conditioning. Crop Scl. 22:116-123. 131
Mexal, J., J. I. Fisher, J. Osteryoung, and C. P. Reid. 1975- Oxygen availability in polyethylene glycol solutions and its implications in plant- water relations. Plant Physiol. 55^20-2^. Meyer, R. F., and J. S, Boyer, 1972. Sensitivity of cell division and cell elongation to low water potentials in soybean hypocotyles. Planta 108:77-87. Michel, B. E. 1971» Further comparisons between Carbowax 6000 and mannitol as suppressants of cucumber hypocotyl elongation. Plant Physiol. 48:513-516. Michel, B. E. 1970. Carbowax 6000 compared with mannitol as a suppressant of cucumber hypocotyl elongation. Plant Physiol. 45:507-509' Michel, B. E., and M. R. Kaufmann. 1973» The osmotic potential of polyethylene glycol 6000. Plant Physiol. 51 s914-916. Miller, D. E., and D. W, Burke. 1974. Influence of soil bulk density and water potential on Fusarium root rot of beans. Phytopathology 64:526-529» Miller, P. R. 1953» The effect of weather on disease. Pages 83-93 in A. Steffereud, editor. Plant Diseases. 1953 Yearbook of Agric. US Govt. Printing Office,. Washington, D. C. Millar, A. A., Mo E. Duysen, and G. E. Wilkinson. I968. Internal water balance of barley under soil moisture stress. Plant Physiol. 43:968-972. Millar, A. A., W. R. Gardner, ans S, M. Goltz. 1971» Internal water status and water transport in seed onion plants. Agron. J. 63:779-784. Millar, B, D., and 0. T, Denmead. 1976. Water relations of wheat leaves in the field. Agron. J. 68:303-307» Mizrahi, Y», A. Blumenfeld, and A. E. Richmond. 1970. Abscisic acid and transpiration in leaves in relation to osmotic root stress. Plant Physiol. 46:169-175» Monteith, J. L., and T. A. Bull. 1970. A diffusion porometer for field use. II. Theory, calibration, and performance. J. Appl. Ecol. 7s623-638. 132
Moore, L. D., H. B. Couch, and J, R= Bloom. I963. In fluence of environment on diseases of turfgrasses. III. Effect of nutrition, pH, soil temperature, air temperature I and soil moisture on Pythium blight of Highland bentgrass. Phytopathology 53 s53-57• Morgan, J. M. 1977a. Changes in the diffusive con ductance and water potential of wheat plants before and after anthesis. Aust. J. Plant Physiol. 4:75-86. Morgan, J. M. 1977b. Differences in osmoregulation betwen wheat genotypes. Nature 270:234-235. Morton, A. G., and D. j, Watson. 1948. A phsyiological study of leaf growth. Ann. Bot. 12:281-289. Moss, D. N. 1963. The effect of environment on gas exchange in leaves. Conn. Agric. Exp. Stn. Bull. 664. Muller, R. H., and H. J. Stolten. 1953» Use of thermis tors in precise measurement of small temperature differences. Thermometric determination of molecular weights. Anal. Chem. 25:1103-1106. Munns, R. E., C. J. Brady, and E. W. R. Barlow. 1979» Solute accumulation in the apex and leaves of wheat during water stress. Aust. J. Plant Physiol. 6: 379-389. Murphy, H. C. 1935» Effect of crown rust infection on yield and water requirement of oats. J. Agric. Res. 50:387-411. Mussell, H., and R. C. Staples (eds.). 1979» Stress Physiology in Crop Plants. John Wiley and Sons, New York, Nechiporenko, G. A., and B, A. Rybalova. 1980a. Ap plicability of various osmotic agents for studying water metabolism in plant tissues. Fiziol. Rast 27:203-208. Nechiporenko, G. A., and B. A. Rybalova. 1980b. Use of different osmotic agents to investigate water exchange of plant tissues. Fiziol. Rast 27:165- 169. 133
Neuman, H. H., and B. W. Thurtell. 1972. A Peltier cooled thermocouple dewpoint hygrometer for in situ measurement of water potentials. Pages 103-112 in R. W, Brown and B. P. Van Haveren, editors. Psychrometry in Water Relations Research. Utah Agric. Exp. Stn., Logan, Utah. Olmsted, C. E, 1941. Growth and development in range grasses. I. Early development of Bouteloua curtipendula in relation to water supply. Bot. Gaz. 102:499-519. O'Toole, J. C., and T. T, Chang. 1979» Drought resistance in cereals—rice: A case study. Pages 373-406 in H. Mussell and R. C. Staples, editors. Stress Physiology in Crop Plants. John Wiley and Sons, New York. Paleg, L. G., and D. Aspinall (eds.). 1982. Physiology and Biochemistry of Drought Resistance in Plants. Academic Press, Sydney, Australia. Papendick, R. I., and G. S. Campbell. 1981. Theory and measurement of water potential. Pages 1-22 in J. F. Parr, W. R. Gardner, and L. F. Elliot, editors. Water Potential Relations in Soil Microbiology. Soil Sci. Soc. Am., Madison, Wise. Parker, J. I968. Drought resistance mechanisms. Pages 195-235 in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 1. Academic Press, New York. Parker, J. I963. Comparison of some methods of deter mining stomatal opening. Plant Physiol. 38 (Suppl.): xlviii. Passioura, J. B. 1972. The effect of root geometry on the yield of wheat growing on stored water. Aust. J. Agric. Res. 23:745-752. Paulech, C., and A» Haspelova-Horvatovicova. 1970. Photosynthesis, plant pigments, and transpiration in healthy barley and barley infected by powdery mildew. Biologia 25:477-487. Pearson, R. W. I966. Soil environment and root development. Pages 95-126 in W. H. Pierre, D. Kirkham, J. Pesek, and R.. Shaw, editors. Plant Environment and Efficient Water Use. Am. Soc. Agron., Madison, Wise. 134
Peck, A, J. 1963. Theory of the Spanner psyohrometer, II. The thermocouple. Agric. Meteorol. 5^433-447. Peck, A. J. i960. Change of moisture tension with temperature and air pressure: Theoretical. Soil Sol. 89:303-310. Pellet, H. M,, and E. C, Roberts. 1963* Effects of mineral nutrition on high temperature growth re tardation of Kentucky bluegrass. Agron. J. S5- 473-476. Peters, D. B. i960. Growth and water absorption by corn as influenced by soil moisture tension, moisture content, and relative humidity. Soil Sci. 24:523-527. Petrie, A. H. K., and J. G. Wood. 1938a. Studies on the nitrogen metabolism of plants. I. The relation between the content of proteins, amino acids, and water in the leaves. Ann. Bot. 2:33-41. Petrie, A. H. K., and J. G. Wood. 1938b. Studies of the nitrogen metabolism of plants. III. On the effect of water content on the relationship between proteins and amino acids. Ann. Bot. 2:881-888. Petrie, A. H. K., and J, I. Arthur. 1943. Physiological onogeny in the tobacco plants: The effect of varying water supply on the drifts in dry weight and leaf area and on various components of the leaves. Aust. J. Exp. Biol. 21:191-209. Pierre, W. H., D. Kirkham, J. Pesek, and R. Shaw (eds.). 1966, Plant Environment and Efficient Water Use. Am. Soc. Agron., Madison, Wise. Plaut, A., and L. Ordin. 1964. The effect of moisture tension and nitrogen supply on cell wall metabolism of sunflower leaves. Physiol. Plant. 17:279-286. Quarrie, S. A., and H. G. Jones. 1977. Effects of abscisic acid and water stress on development and morphology of wheat. J. Exp. Bot. 28:192-203. Raschke, K, 1975. Stomatal action. Annu. Rev. Plant Physiol. 26:309-340. 135
Raschke, K. 19^9• Stomatal responses to pressure changes and interruptions in the water supply of detached leaves of Zea mays. Plant Physiol. 45: 415-423. Rawlins, S, L. 1976. Measurement of water content and the state of water in soils. Pages 1-55 in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 4. Academic Press, New York. Rawlins, S, L. I966. Theory for thermocouple psychrometers used to measure water potential in soil and plant samples. Agric. Meteorol. 3:293-310. Richter, H. 1978. Water relations of single drying leaves: Evaluation with a dewpoint hygrometer. J. Exp. Bot. 29:277-280. Richards, L. A., and G. Ogata. 1958. Thermocouple for vapor pressure measurement in biological and soil systems at hi^ humidity. Science 128:1089-1090. Roberts, E, C., and D. P. Lage. 1965* Effects of an evaporation retardant, a surfactant, and an osmotic agent on foliar and root development of Kentucky bluegrass. Agron. J. 57:71-74. Rubin, B. A., and Y. V. Artsikhovskaya. I963. Biochemistry and Physiology of Plant Immunity. Permagon Press, Oxford. Ruf, R. H., R. E. Eckert, and R. 0. Gifford. 19^7. Components of osmotic adjustment of plants to rapid changes in root medium osmotic pressure. Soil Sci. 104:159-162. Rutter, A. J., and F. H. Whitehead (eds.). I963. The Water Relations of Plants. John Wiley and Sons, New York. Rychnovska, M., and J. Artos. I962. Measurement of photosynthesis by the air dry weight increment of sample composed of leaf segments. Biol. Plant, 4:91-99. Sachs, T,, and K. V. Thimann. 1964. Release of lateral buds from apical dominance. Nature 201:939-940. Salisbury, F. B., and C. W. ^^ss. 1978. Plant Physiology. 2nd Edition. Wadsworth Pub. Co., Belmont, OA. 136
Sampson, J. I96I. A method of replicating dry or moist surfaces for examination "by light microsopy. Nature 191:932-935. Sanchez-Diaz, M. P., and P. J. Kramer. 1972. Behaviour of corn and sorghum under water stress and during recovery. Plant Physiol. 48:613-161. Schoeneweiss, D. F, 1978. Water stress as a predisposing factor in plant disease. Pages 6I-IOO in T. T. Kozlov/ski, editor. Water Deficits and Plant Growth. Vol. 5« Academic Press, New York. Schoeneweiss, D. F. 1975a. A method for controlling plant water potentials for studies on the influence of water stress on disease susceptibility. Can. J. Bot. 53:647-652. Schoeneweiss, D. F. 1975%» Predisposition, stress, and plant disease. Annu. Rev. Phytopathol. 13:193-211. Scholander, P. F., H. T. Hammel, E. D. Bradstreet, and F, A. Hemmingspen. I965. Sap pressure in vascular plants. Science 148:339-346. Scholander, P. F., H. T. Hammel, F, A. Hemmingspen, and E. D. Bradstreet. 1964. Hydrostatic pressure and osmotic potential in leaves of mangroves and {.some., other plants. Proc. Nat. Acad. Soi. 52:119-125» Scotter, D. R. 1972. The theoretical and experimental behaviour of a Spanner psychrometer. Agric. Meteorol. 70:125-136. Sempio, C., 0. Majernik, and V. Raggi. I966. Water loss and stomatal behaviour of bean (Phaseolus vulgaris) infected by Uromvces appendiculatus. Biologia 21:99-104. Sepaskhah, A. R., and L. Boersma. 1979a. Elongation of wheat leaves exposed to several levels of raatric potential and NaCl-induced osmotic potential of soil water. Agron. J. 848;852. Sepaskhah, A. R., and L. Boersma. 1979b. Shoot and root growth of wheat seedlings exposed to several levels of raatric potential and NaCl-induced osmotic potential of soil water. Agron. J. 71:746-752. 137
Sequeira, L, 1973- Hormone metabolism in diseased plants. Annu. Rev. Plant Physiol. 24:353-380. Shaw, M., and D. J. Samborski. 1957» The physiology of host-parasite relations. III. The pattern of respiration in rusted and mildewed cereal leaves. Can. J. Bot. 35:389-395. Shaw, R. H., and D. R. Laing. I966. Moisture stress and plant response. Pages 73-9^ in W. H. Pierre, D. Kirkham, J. Pesek, and R. Shaw, editors. Plant Environment and Efficient Water Use. Am. Soc. Agron., Madison, Wise. Shepard, W. 1975* Matric water potential of leaf tissue- measurement and significance. J. Exp. Bot. 26:465- 468. Shimshi, D. 1964. The use of a field porometer for the study of water stress in plants. Isr. J. Agric. Res. 14:137-145. Simmelsgaard, S. E. 1976. Adaptation to water stress in wheat, Physiol. Plant. 37:167-174. Skoss, J. D. 1955. Structure and composition of plant cuticle in relation to environmental factors and permeability. Bot. Gaz. 117:55-72. Slatyer, R. 0. I967. Plant-Water Relationships. Academic Press, New York. Slatyer, R. 0. I966. An underlying cause of measurement discrepancies in determination of osmotic characteris tics in plant cells and tissues. Protoplasma 62: 34-38. Slatyer, R. 0. 1958. The measurement of diffusion pressure deficit in plants by a method of vapour equilibration. Aust. J. Biol. Sci. 11:349-358. Slatyer, R. 0, 1957. Plant response to soil moisture stress. Aust. J. Biol. Sci. 10:320-331. Slatyer, R. 0., and J. V. Lake. I966. Resistance to water transport in plants—whose misconception? Nature 212:1585-1591.
Slatyer, R. 0., and S. A. Taylor, i960. Terminology in plant and soil-water relations. Nature 187:922-924. 138
Slavik, B. 197^* Methods of Studying Plant-Water Relations. Springer-Verlag, New York. Slavik, B. 1973» Transpiration resistance in leaves of maize grown in humid and dry air. Pages 267-269 in R. 0. Slatyer, editor. Plant Responses to Climatic Factors. UNESCO, Paris. Slavik, B. 1963a. Relationship between the osmotic potential of cell sap and the water saturation deficit during the wilting of leaf tissue. Biol. Plant. 5s258-265. Slavik, B. 1963b. The distribution pattern of transpiration rate, water saturation deficit, stomata number and size, photosynthetic and respiration rate in the area of the tobacco leaf blade. Biol. Plant. 5:143-152. Snow, R. 1925» The correlative inhibition of the growth of axillary buds. Ann. Bot. 39:841-856. Spanner, D. D. 1951» The Peltier effect and its use in the measurement of suction pressure. J. Exp. Bot. 2:145-168. Stalfelt, M. G. I96I. The effect of the water deficit on the stomatal movements in a carbon dioxide-free atmosphere. Physiol. Plant. 14:826-843. Stalfelt, M. G. 1955» The stomata as a hydrophotoic regulator of the water deficit of the plants. Physiol. Plant. 8:572-593. Steudle, E., U. Zimmermann, and U. Luttge. 1977' Effect of turgor pressure and cell size on the wall elasticity of plant cells. Plant Physiol. 59:285-289. Steuter, A., A. Mozafar, and J. R. Goodwin. I98I. Water potential of aqueous polyethylene glycol. Plant Physiol, 67:64-67. Stocker, 0. i960. Physiological and morphological changes in plants due to water deficiency. Pages 63-104 in Plant Water Relationships in Arid and Semi-Arid Conditions—Review of Research. UNESCO, Paris. Stone, J. F. 1975. Plant Modification for More Efficient Water Use. Elsevier, New York. 139
Szabo, S. S., and K. P. Buchholtz. I96I. Penetration of living and non-living surfaces by 2, 4-D as influenced by ionic additives. Weeds 9s177-179* Talboys, P. W. I968. Water deficits in vascular disease. Pages 255-311 in T. T. Kozlowski, editor. Water Deficits and Plant Growth. Vol. 2, Academic Press, New York. Talboys, P. W., M. Bennett, and J. F. Wilson. I96I. Tolerance to Verticillium wilt diseases in the strawberry. Ann. Rept. East Mailing Res. Stn., Kent. Taylor, H. M., and L. F. Ratliff. 1969a. Root elongation rates of cotton and peanuts as a function of soil strength and soil water content. Soil Sci. 108: 113-119. Taylor, H. M., and L. F. Ratliff. 1969b. Root growth pressures of cotton, pears, and peanuts. Agron. J. 61:398-402. Thiitiann, K. V., and F. Skoog. 193^. Inhibition of bud development and other functions of growth substances in Vicia faba. Proc, Roy. Soc. London Ser. B 114: 317-331. Thirumalachar, M. J., and J. G. Dickson. 1953» Spore germination, cultural character, and cytology of varieties of Ustilago striiformis and the reaction of hosts. Phytopathology 43î527-534. Threlfall, R. J. 1959. Physiological studies on the Verticillium wilt disease of tomato. Ann. Appl. Biol. 47:57-60. Thut, H. F. 1938. Relative humidity variations affecting transpiration. Am. J. Bot. 25:589-594. Tiver, N. S. 1942. Studies of the flax plant. I. Physiology of growth, stem anatomy, and fiber development in fiber flax. Aust. J. Exp. Biol. 20:149-153. Townley-Smith, T. F,, and E. A. Kurd. 1979. Testing and selecting for drought resistance in wheat. Pages 447- 465 in H. Mussell and R. C. Staples, editors. John Wiley and Sons, New York. 140
Trip, P., G. Krotkov, and C. D. Nelson. 1964. Metabolism of mannitol in higher plants. Am. J. Bot, 51'828-832. Troughton, J. H. I969. Plant water status and carbon dioxide exchange of cotton leaves. Aust. J, Biol. Sci. 22:289-302. Turner, N. C. 1979• Drought resistance and adaptation to water deficits in crop plants. Pages 343-372 in H. Mussell and R. C. Staples, editors. Stress Physiology in Crop Plants. John Wiley and Sons, New York. Turner, N. C. 1975* Concurrent comparisons of stomatal behaviour, water status, and evaporation of maize in soil at high or low water potential. Plant Physiol. 55:932-937. Turner, N. C. 1974. Stomatal behavior and water status of maize, sorghum, and tobacco under field conditions. II. At low soil water status. Plant Physiol. 53O60- 365. Turner, N. C. 1970. Response of adaxial and abaxial stomata to light. New Phytol. 69s647-653• Turner, N. C., J. E. Begg, and M. L. Tonnet. 1978. Osmotic adjustment of sorghum and sunflower crops in response to water deficits and its influence on the water potential at which stomata close. Aust, J. Plant Physiol. 5:597-608. Turner, N. C., and M. M. Jones. I98O. Turgor maintenance by osmotic adjustment. A review and evaluation. Pages 87-104 in N. C. Turner and P. J. Kramer, editors. Adaptation of Plants to Water and High Temperature Stress. John Wiley and Sons, New York. Turner, N. C., and P. J. Kramer (eds.). I98O. Adaptation of Plants to Water and High Temperature Stress. John Wiley and Sons, New York. Tyree, M. T. 1976. Negative turgor pressure in plant cells—fact or fallacy? Can. J. Bot. 54:2738-2746. Tyree, M. T., and H. T. Hammel. 1972. The measurement of the turgor pressure and the water relations of plants by the pressure bomb technique. J. Exp. Bot. 23:267-282. 141
Vaadia, Y., and G. Itai. I968. Interrelationships of growth with reference to the distribution of growth substances. Pages 65-99 in W. J. Whittington, editor. Root Growth. Butterworth, London. Vaadia, Y., F. C. Raney, and R. M. Hagan. I96I. Plant water deficits and physiological processes. Annu. Rev. Plant Physiol. 12s265-292. Van Bavel, C. H. M., F. S. Nakayama, and W. L. Ehrler. 1965. Measuring transpiration resistance of leaves. Plant Physiol. Î0;535-5^0. Van der Wal, A. P., H. Smeitink, and G. C. Maan. 19?5« An ecophysiological approach to crop losses exemplified in the system wheat, leaf rust, and glume blotch. III. Effects of the soil-water potential on development, growth, transpiration, symptoms, and spore production of leaf rust-infected wheat. Neth. J. Plant Pathol, 81:1-13. Van der Wal, A. F., and M. C. Cowan. 1974. An eco physiological approach to crop losses exemplified in the system wheat, leaf rust, and glume blotch. II. Development, growth, and transpiration of unin fected plants and plants infected with Puccinia recondita f. sp. triticina and/or Septoria nodorum in a climate chamber experiment. Neth. J. Plant Pathol. 80:192-214. Van Overbeek, J. 1942. Water uptake by excised root systems of the tomato due to osmotic forces. Am. J. Bot. 31:265-271. Wadleigh, C. H., W. A. Raney, and D. m. Herschfield. 1966. The moisture problem. Pages I-I9 in W, H. Pierre, D. Kirkham, J. Pesek, and R. Shaw, editors. Plant Environment and Efficient Water Use. Am. Soc. Agron., Madison, Wise. Wallihan, E. F. 1964. Modification and use of an electric hygrometer for estimating relative stomatal apertures. Plant Physiol. 39s86-89. Wardlaw, I. F. I967. The effect of water stress on translocation in relation to photosynthesis and growth. I. Effect during grain development in wheat. Aust. J. Biol. Soi. 20:25-39. 142
Weatherley, P. E. 1970. Some aspects of water relations. Adv. Bot. Res. 3sl71-206. Weatherley, P. E. 1965. The state and movement of water in the leaf. Symp. Soc. Exp. Biol. 19s157-161. Weatherley, P. E. 1950- Studies on the water relations of the cotton plants. I. The field measurement of water deficits in leaves. New Phytol. 49:81-97" Weaver, J. E., and W. J. Hiramel. 1930» Relation of increased water content and decreased aeration to root development in hydrophytes. Plant Physiol. 5Î69-92.
Wein, H. C., E. J. Littleton, and A. Ayanaba. 1979* Drought stress of cowpea and soybean under tropical conditions. Pages 283-302 in H. Mussell and R. C. Staples, editors. Stress Physiology of Crop Plants. John Wiley and Sons, New York. Welton, F. A., and J. D. Wilson. 1931» Water-supplying power of the soil under different species of grass and with different rates of water application. Plant Physiol. 6:485-493» Wenkert, W., E. R. Lemon, and T. R. Sinclair. 1978a. Leaf elongation and turgor pressure in field-grown soybean. Agron. J. 70:761-767. Wenkert, W., E. R. Lemon, and T. R. Sinclair. 1978b. Water content-potential relationship in Soya bean; Changes in component potentials for mature and immature leaves under field conditions. Ann. Bot. 42:295-308. Whiteman, P. C., and G. L. Wilson. I965. Effects of water stress on the reproductive development of Sorghum vulgare. Univ. Queensland Papers, Dept. Bot. 4* 233-239. Wickson, A., and K. V. Thimann. 1958. The antagonism of auxin and kinetin in apical dominance. Physiol. Plant. 11:597-618. Wiebe, H. H. I966. Matric potential of several plant tissues and biocolloids. Plant Physiol. 41:1439-1^^2. 143
Williams, J., and C. F, Shaykevirich. I969. An evaluation of PEG 6000 and PEG 20,000 in the osmotic control of soil water matric potential. Can. J. Soil Sci. 49:397-401. Williams, R. F., and R. E. Shapter. 1955* A comparative study of growth and nutrition in barley and rye as affected by low water treatment. Aust. J. Biol. Sci. 8:435-441. Wilson, D. 1975» Stomatal diffusion resistances and leaf growth during draughting of Lolium perenne plants selected for contrasting epidermaJ. ridging. Ann. Appl. Biol. 79:83-94. Wilson, J. R., M. J. Fisher, E. D. Schulze, G. R. Dolby, and M. M. Ludlow. 1979» Comparison between pressure volume and dewpoint hygrometry techniques for de termining the water relations characteristics of grass and legume leaves. Oecologia (Berlin) 41:77-88. Wilson, J. R., M. M. Ludlow, M. J. Fisher, and E. D. Schulze. 1980. Adaptation to water stress of the leaf water relations of four tropical forage species. Aust. J. Plant Physiol. 7:207-220. Wind, G. P. 1955* Flow of water through plant roots. Neth. J. Agric. Sci. 3:259-264. Wingard, S. A. 1953» The nature of resistance to disease. Pages 165-173 in A. Steffereud, editor. Plant Diseases. 1953 Yearbook of Agric. US Govt. Printing Office, Washington, D. C. Wold, D. D., and D. J. Parrish. I98I. Short-term growth responses of tall fescue to changes in soil water potential and to defoliation. Crop Sci. 22:996-999» Wood, J. G. 1932. The physiology of xerophytism in Australian plants. The carbohydrate metabolism of plants with tomentose succulent leaves. Aust. J. Exp. Biol. 10:89-95. Wood, J. I. 1953" Three billion dollars a year. Pages 1-9 in A. Steffereud, editor. Plant Diseases. 1953 Yearbook of Agric. US Govt. Printing Office, Washington, D. C. 1#
Yamda, Y., S. H. Wittwer, and M. J. Bukovac. 1964. Penetration of ions through isolated cuticles. Plant Physiol. 39:28-34. Yamada, Y., S. Tamia, and T. Miyaguchi. I96I. The measure ment of the thickness of leaves using S35. Proc. 2nd Japan. Conf. Radioisotopes, 1958. p. I692. Yeh, R. H., A. G. Matches, and R. L. Larson. 1976. Endogenous growth regulators and summer tillering of tall fescue. Crop Soi. 16:409-413. Younger, V. B., and F. J. Nudge. I968. Growth and carbohydrate storage of three Poa pratensis L. strains as influenced by temperature. Crop Soi. 8:455-457. Zeevart, J. A. D. 1979» Regulation of assimilate partitioning. Summary Reports from Workshop held at Michigan State University, East Lansing, Mich, pp. 14-17, Am. Soc. Plant Physiol., Rockville, MD. Zelitch, I. 1965. Environmental and biochemical control of stomatal movement in leaves. Biol. Rev. Cambridge Phil. Soc. 40:463-482. Zelitch, I. 1961. Biochemical control of stomatal opening in leaves. Proc. Nat. Acad. Soi. 47:1423-1440. Zimmermann, U. 1978. Physics of turgor and osmoregulation. Ann. Rev. Plant Physiol. 29:121-168. 1^5
IX. ACKNOWLEDGMENTS
I wish to thank my major professor, Dr. Clinton Hodges, for his outstanding help in preparation of this manuscript and advice during the study. His professional abilities are the major strength of the department.
I am also indebted to the members of my graduate committee, Dr. Don Nevins, Dr. Cecil Stewart, Dr. Nick Christians, and Dr. Richard Gladon, for their assistance and comments regarding this study. A special thanks goes to Dr. Cliff LaMotte for filling in for Dr. Don Nevins during the final exam. His suggestions and comments were valuable. Last, but not least, I would like to thank my wife, Ann, and my sons, Shawn and Casey, for being patient and understanding during a crucial part of my life.