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Sucrose synthase and invertase in maize roots differing in carbohydrate status and/or genetic composition

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Sucrose synthase and invertase in maize roots differing in carbohydrate status and/or genetic composition
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Duke, Edwin Ralph, 1960-
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English
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ix, 113 leaves : ill., photos ; 29 cm.

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Cells ( jstor )
Corn ( jstor )
Endosperm ( jstor )
Enzymes ( jstor )
Messenger RNA ( jstor )
Metabolism ( jstor )
Phosphates ( jstor )
Plant roots ( jstor )
Root tips ( jstor )
Sugars ( jstor )
Dissertations, Academic -- Horticultural Science -- UF
Horticultural Science thesis Ph. D
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bibliography ( marcgt )
non-fiction ( marcgt )

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Thesis:
Thesis (Ph. D.)--University of Florida, 1991.
Bibliography:
Includes bibliographical references (leaves 94-112).
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Typescript.
General Note:
Vita.
Statement of Responsibility:
by Edwin Ralph Duke.

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SUCROSE SYNTHASE AND INVERTASE IN MAIZE ROOTS DIFFERING
IN CARBOHYDRATE STATUS AND/OR GENETIC COMPOSITION















By

EDWIN RALPH DUKE


A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY

UNIVERSITY OF FLORIDA


1991














ACKNOWLEDGEMENTS


I would like to extend thanks to the members of my committee, Dr. Karen Koch, Dr. Rebecca Darnell, Dr. Don McCarty, Dr. Curt Hannah and Dr. Alice Harmon, for their assistance and guidance during the completion of this degree. I would also like to thank Dr. Tom Humphreys for his critical review of this dissertation. There are many people in the Fruit Crops Department to whom I owe thanks. I would like to thank the staff and faculty for their help and encouragement during my time here. For their friendship and support, I especially would like to thank Kathy Zimmerman, Teki and Andy Ericson, Dr. Kathy Taylor, Dr. Pat Tomlinson, Wayne Avigne, Kurt Nolte, and Don Merhaut. Thanks are also given to the graduate students, both past and present, of the Fruit Crops Department. Finally, I want to extend my most heartfelt thanks to my parents, Ralph and Mildred Duke; they have stood by me and given me support throughout my education, and I can never thank them enough.














TABLE OF CONTENTS


ACKNOWLEDGEMENTS ................................

LIST OF TABLES .....................................

LIST OF FIGURES ....................................

ABSTRACT .........................................

CHAPTERS

1 INTRODUCTION ............................

2 REVIEW OF THE LITERATURE .................

Sucrose Metabolism .........................
Sucrose Synthase ..........................
Invertases ..............................
Use of Mutants in Physiological Research .........

3 INSTABILITY OF SUCROSE SYNTHASE FROM ROOT
TIPS: CHARACTERIZATION AND STABILIZATION ...

Abstract ..................................
Introduction ...............................
Materials and Methods .......................
Results ...................................
Discussion ................................

4 SUCROSE SYNTHASE ACTIVITY IN WILDTYPE MAIZE
ROOT TIPS RESPONDING TO ALTERED CARBOHYDRATE STATUS ....................

Abstract ..................................
Introduction ...............................
Materials and Methods .......................








Results ......................................... 52
Discussion ...................................... 52

5 SUGAR RESPONSE OF SUCROSE SYNTHASE AT
THE GENE (Susl), PROTEIN AND ENZYME ACTIVITY
LEVELS IN ROOTS OF THE Shl MAIZE MUTANT ......... 57

Abstract ........................................ 57
Introduction ..................................... 58
Materials and Methods ............................. 61
Results ......................................... 63
Discussion ...................................... 68

6 AN ORGAN-SPECIFIC INVERTASE DEFICIENCY IN
THE PRIMARY ROOT OF AN INBRED MAIZE LINE ........ 75

Abstract ........................................ 75
Introduction ..................................... 76
Materials and Methods ............................. 78
Results ......................................... 80
Discussion ...................................... 85

7 SUMMARY AND CONCLUSIONS ..................... 91

LITERATURE CITED ......................................... 94

BIOGRAPHICAL SKETCH ..................................... 113














LIST OF TABLES


Table 3-1 Effect of enzyme protectants on activity of sucrose synthase
from maize root tips assayed five minutes after extraction ........ 36

Table 4-1 Total sucrose synthase activity in wildtype maize root tips
incubated in a range of glucose concentrations for 24 hour ....... 53

Table 5-1 Sucrose synthase activity in mutant maize (W22:shl) root tips
incubated in media containing a range of glucose concentrations for
24 hours ............................................ 66

Table 5-2 Sucrose synthase activity in mutant maize (W22:shl) root tips
incubated in media containing 0 or 2.0% glucose for various time
periods ............................................. 70

Table 6-1 Soluble and insoluble acid invertase activity in sequential 2 mm
segments of primary roots of 5- to 6-day-old seedlings from 1 hybrid
and 2 inbred lines of maize ............................... 81

Table 6-2 Soluble acid invertase and sucrose synthase activity in 0.5 cm
apices of primary and adventitious roots of 5- to 6-day-old seedlings
from 1 hybrid and 2 inbred lines of Zea mays ................. 83

Table 6-3 Soluble acid invertase activity in various tissues of Oh 43, an
inbred line of Zea mays ................................. 84













LIST OF FIGURES


Figure 3-1 Time course of in vitro decrease in sucrose synthase activity in
maize and cotton roots .................................. 35

Figure 3-2 Time course of in vitro decrease in maize root sucrose synthase
activity with and without the serine proteinase inhibitor, PMSF ..... 37

Figure 3-3 Time course of in vitro decrease in maize root sucrose synthase
activity in the presence and absence of either Pi (10 mM) or casein
2% w :v) ............................................. 39

Figure 3-4 Denaturing (A) and native (B,C,D) protein gel blot analysis of
maize root sucrose synthase at various times after extraction ..... 40

Figure 3-5 Time course of in vitro decrease in maize root sucrose synthase
activity from lines with homo- (W22:shl) and heterotetrameric (NK
508) forms of this enzyme ............................... 43

Figure 5-1 RNA gel blot analysis of Susl expression in maize roots
incubated in a range of glucose concentrations for 24 hours ...... 64

Figure 5-2 Protein gel blot of Susl encoded sucrose synthase from maize
roots incubated in a range of glucose concentrations for 24 hours.. 65

Figure 5-3 RNA gel blot analysis of Susl mRNA expression in maize roots
incubated in 0 and 2.0% glucose for various time periods ........ 67

Figure 5-4 Protein gel blot of Susl encoded sucrose synthase from maize
roots incubated in 0 and 2.0% glucose for various time periods .... 69

Figure 6-1 Histochemical localization of invertase activity in free-hand,
cross sections of root apices of 6-day-old maize seedlings ....... 87














Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy
SUCROSE SYNTHASE AND INVERTASE IN MAIZE ROOTS DIFFERING
IN CARBOHYDRATE STATUS AND/OR GENETIC COMPOSITION

By

Edwin Ralph Duke

August, 1991

Chairperson: Dr. Karen E. Koch
Major Department: Horticultural Science

The extent to which sucrose is transported in phloem of higher plant species necessitates its effective metabolism by non-photosynthetic cells. However, only two enzymes can catalyze sucrose breakdown in these instances, invertase and sucrose synthase (a reversible enzyme). Specific molecular, genetic and physiological factors affecting these enzymes were investigated in the root tips of maize. A radiometric assay was first developed for sucrose synthase to circumvent the rapid decline of sucrose synthase activity in vitro. Further, characterization of in vitro instability indicated that activity decline was not associated with any detectable proteolytic degradation, charge alteration, or subunit separation and that inhibition of activity by inorganic phosphate suggested possible phosphorylation of this enzyme.








Message levels of genes encoding sucrose synthase isozymes in maize have been shown to respond to tissue carbohydrate status, thus the effects of such changes were examined at the level of enzyme activity. Total sucrose synthase activity from roots of wildtype plants showed little difference in extracts from root tips incubated for 24 h in a range of glucose levels. This activity, however, is the combined contribution of two isozymes whose genes are responding differentially to experimental conditions.

The shI mutant of maize was used to study expression of the Susl gene for sucrose synthase in response to sugar availability because this mutant has only one gene (Susl) for sucrose synthase and provides a system uncomplicated by the presence of the second isozyme (Shl). Susl mRNA increased 5-fold when incubated for 24 h in 2.0% glucose compared to 0 or 0.2% glucose. Levels of Sus protein were slightly elevated with increasing sugar levels. Enzyme activity was elevated 2-fold under the same conditions. A study of time-course and treatment reversals showed that changes in mRNA or protein were not evident until 24 h and indicated that the response to carbohydrate level had been initiated within 16 h. Overall, enhanced expression of Susl was evident at the mRNA, protein and enzyme levels.

An organ-specific invertase deficiency affecting only the primary root system also was characterized in the Oh 43 maize inbred. Substantial acid invertase activity was evident in extracts of all tissues tested except the primary root system of Oh 43. This deficiency was also evident in lateral roots arising from the primary









root but not in otherwise morphologically identical laterals from adventitious roots. In contrast, sucrose synthase was active in all roots and theoretically provided the only available avenue for sucrose degradation in primary root tips of Oh 43.














CHAPTER 1
INTRODUCTION


Sucrose metabolism is important to the majority of plant species because of the nearly ubiquitous role of this sugar in growth and development. Initial breakdown of sucrose can be catalyzed by either invertase or the reversible enzyme sucrose synthase.

Gene responses to changes in carbohydrate availability have been reported for mammalian cells (Un and Lee, 1984) and yeasts and bacteria (Carlson, 1987; Schuster, 1989). Evidence has also been presented that the transcriptional activity of promoters of photosynthetic genes from maize protoplasts are repressed and coordinated by sugars (Sheen, 1990). Expression of the genes encoding the sucrose synthase isozymes also have been shown to respond to carbohydrate levels (Koch and McCarty, 1988; Koch et al., 1989). Northern blot analysis of mRNA from wildtype maize roots showed levels of one isozyme gene (Shl) was up-regulated in response to carbohydrate depletion whereas the other (Sus) responded positively only when sugar availability was elevated. This sensitivity of genes for a key enzyme in sugar metabolism may prove to be an important control mechanism whereby plant cells are able to react to cellular nutritional conditions.

The importance of invertase, the other enzyme of sucrose catabolism in plants, to root metabolism is unclear. Invertases are especially active in tissues








2

undergoing rapid cell division such as shoot and root apices (Avigad, 1982). Recent evidence, however, indicates that although much hydrolysis is often observed, invertase activity may not be essential for sucrose uptake into either sugar cane stems (Lingle, 1989; Thom and Maretzki, 1990) or maize kernels (Schmalstig and Hitz, 1987). Chapleo and Hall (1989a, b, and c) also have concluded that invertase was not essential to sucrose import into roots of Ricinus. Giaquinta and co-workers (1983) showed that sucrose entering the roots of maize via phloem does not have to pass through the extracellular space. Robbins (1958) first reported that root tips of an inbred line of maize, Oh 43, were unable to retrieve exogenous sucrose. B. Burr (Brookhaven National Laboratory, personal communication) has indicated that the lack of retrieval might possibly be due to an invertase deficiency. The absence of invertase activity could have important implications for sucrose import, not only because of potential effects on the retrieval system, but also because sucrose breakdown in such an instance could theoretically proceed only via action of sucrose synthase. A mutant lacking functional invertase in its roots would also be useful, in combination with nulls for both sucrose synthase isozymes, to elucidate the individual roles for these enzymes.

The purpose of the following research is to further elucidate the roles and regulation of the two sucrose metabolizing enzymes sucrose synthase and invertase in roots of maize. Specific objectives are to:








3

1. Develop a method of accurately assaying sucrose synthase activity

to circumvent the rapid decline in activity exhibited upon extraction

from maize root tips.

2. Clarify the possible causes of the rapid decline of sucrose synthase

activity in extracts from maize root tips.

3. Determine the effects of varying carbohydrate conditions on total

sucrose synthase activity in extracts from wildtype maize root tips. 4. Ascertain the effects of varying carbohydrate conditions on the Susl

gene and its sucrose synthase gene product free from the

confounding effects of Shl.

5. Characterize the extent of invertase activity in various tissues of the

Oh 43 inbred of maize, a putative invertase-deficient mutant.













CHAPTER 2
REVIEW OF THE UTERATURE



Sucrose Metabolism


Sucrose is the major transported sugar in the majority of higher plant species. The major roles of sucrose in higher plants include its function as both a translocatable form of carbon and as a vacuolar storage compound (Hawker, 1985). It is a non-reducing sugar made up of a glucose (a-D-glucopyranose and fructose (pl-D-fructofuranose) joined by an a-1,2 linkage. For metabolic utilization, it must be broken down into its component monosaccharides or their derivatives. Only two enzymatic systems for sucrose breakdown are known in higher plant tissues. Sucrose can be hydrolyzed by the action of invertase or cleaved by sucrose synthase working in the degradative direction. The free energy of sucrose hydrolysis is nearly equal to that of the y phosphoryl group of ATP (AGO= -7.0 kcal/mol and -7.3 kcal/mol, respectively) (Neufeld and Hassid, 1963). This is much greater than the free energy of most other glycosidic bonds (Avigad, 1982). Cleavage by sucrose synthase retains the bond energy in the a-glucosyl bond of UDP-glucose. In contrast, hydrolysis by invertase conserves none of the free energy in the bond.








5

Morell and Copeland (1984, 1985) have investigated the enzymes of sucrose breakdown in soybean nodules and found that both sucrose synthase and alkaline invertase are present. They suggested that sucrose partitioning between the two enzymes could be determined by differences in their affinities for this substrate. The K, of alkaline invertase for sucrose was 10 mM whereas that of sucrose synthase was 31 mM. Given the presence of both enzymes, they proposed the greater affinity of alkaline invertase for sucrose in this system could ensure that most of the sucrose entering the nodule would be converted to hexoses for further catabolism. At the same time some sucrose would be converted to UDP-glucose for subsequent synthesis of nucleotide sugars and polysaccharides. In contrast, Huber and Akazawa (1986) reported essentially an opposite situation in cultured sycamore cells, another system in which both sucrose synthase and neutral (alkaline) invertase were present at the same time and with similar activities. In these cells the sucrose synthase Km for sucrose was substantially lower than that of neutral invertase (15 vs 65 mM). They proposed two pathways of sucrose cleavage, initiated by each of the enzymes, both pathways eventually leading to the production of triose-phosphates. Sucrose concentration was postulated to regulate carbon flow between the two pathways. Sucrose synthase had a lower Km for sucrose and would, therefore, be relatively more important under sucrose limiting conditions (Avigad, 1982). This pathway is more energy efficient and would be more beneficial to the cells when carbon supplies are limited. The work of Morell and Copeland's work (1984, 1985) and








6

Huber and Akazawa (1986) both demonstrate the potential importance of simultaneous sucrose catabolism via two enzyme systems.


Sucrose Synthase

Sucrose synthase (EC 2.4.1.13, UDP-D-glucose: D-fructose 2-a-Dglucosyltransferase) is ubiquitous in higher plants (Keller et al., 1988) and probably occurs in all types of tissues. However, this enzyme is found in greatest abundance in nonphotosynthetic tissues and in developing seeds (Echt and Chourey, 1985). In cell fractionation studies, sucrose synthase was shown to be associated with the soluble fraction (Nishimura and Beevers, 1979; MacDonald and ap Rees, 1983). A cytosolic rather than vacuolar localization for sucrose synthase has been demonstrated in protoplasts isolated from Jerusalem artichoke (Keller et al., 1988).

Although molecular weights as high as one million have been reported for sucrose synthase (Grimes et al., 1970), it is now generally concluded that, in its native state, sucrose synthase has a molecular weight of approximately 36 to 40 kD (Delmer, 1972a; Su and Preiss, 1978; Morell and Copeland, 1985; Moriguchi and Yamaki, 1988) and is composed of four identical subunits. The sucrose synthase subunit from maize has been found to have a molecular weight of 8.8 kD (Su and Preiss, 1978). However, unlike many other plant species, maize has two genes which encode sucrose synthase subunits (Chourey and Nelson, 1976; Echt and Chourey, 1985). The two sucrose synthase subunits of maize, shl and sus, encoded by the Shl and Susl genes, respectively, have similar enzyme kinetics,








7

similar amino acid compositions and share limited structural homologies (Echt and Chourey, 1985). They do differ slightly in their electrophoretic movement during PAGE (Echt and Chourey, 1985). The two subunits are homologous enough to form heterotetrameric structures, apparent as five separate bands on native PAGE (Echt and Chourey, 1985).

Spatial separations of these two sucrose synthase isozymes are sometimes apparent. Shl encoded protein is primarily located in the endosperm (Chourey and Nelson, 1976; Chen and Chourey, 1989), in the root (Springer et al., 1986; Chourey et al., 1986) and in etiolated shoots (Springer et al., 1986). The Susl encoded protein is found throughout the plant (Chourey, 1981; Echt and Chourey, 1985; Chourey et al., 1988). The distribution of both proteins is further distinguished under stress conditions (such as anaerobiosis) where tissue-specific localization in roots is readily apparent (Rowland et al., 1989). Roles of sucrose synthase

Sucrose synthase was initially considered to be a sucrose synthesizing system in plants by Leloir and Cardini (1953, 1955). Sucrose synthase and sucrose phosphate synthase (EC 2.3.1.14, UDP-D-glucose: D-fructose-6phosphate 2-a-D-glucosyltransferase) are the two enzymes that catalyze the transglucosylation reaction from UDP-glucose to fructose and fructose-6phosphate, respectively.









8

The necessity for two systems of sucrose synthesis was puzzling until it was determined that the sucrose synthase reaction is readily reversible (Cardini et al, 1955):

Sucrose + UDP < --------- > UDP-Glucose + Fructose

This reversibility gave rise to the suggestion that sucrose synthase could make UDPG available for utilization as a glucosyl donor in starch synthesis (Turner and Turner, 1957). Other studies of sucrose synthase specificity and kinetics led Avigad and coworkers (Avigad, 1964; Avigad et al., 1964; Milner and Avigad, 1964) to suggest that this enzyme functioned mainly in sucrose cleavage in storage tissues. Sucrose synthase activity typically is highest in tissues during periods of rapid growth and is often not accompanied by high invertase activity (Schaffer et al., 1987).

Several factors lend credence to the view that the role of sucrose synthase is sucrose cleavage in importing cells. First, a substantial level of free fructose is required for sucrose synthase activity in the synthetic direction (Km ca. 2.0-2.5 mM [Avigad, 1982]). Levels of this sugar are low in healthy, intact leaves, but they are higher in storage tissues and roots, where most of the free fructose is in the vacuole or extracellular spaces (Avigad, 1982). Availability of UDPG is also likely to limit the synthetic reaction, because the cellular concentration is typically less than 0.4 mM (Murata, 1975). The Km for UDPG ranges from 0.1 to 8.5 mM (average approximately 2.0 mM)(Avigad, 1982). In contrast, sucrose









9

concentrations are generally elevated in importing areas. Therefore, substrate levels favor the cleavage reaction in the majority of instances.

A second line of evidence for the degradative role of sucrose synthase in importing organs is provided by mutant maize lines lacking a functional sucrose synthase protein. Chourey and Nelson (1976) showed that a deletion of the Sh locus on chromosome 9 of maize (coding for sucrose synthase) led to a 90% deficiency of the respective protein in mutant vs wild-type kernels. Starch formation was also reduced in this line, giving rise to a "shrunken" seed. The association between this shrunken phenotype and a sucrose synthase deficiency was considered evidence that the critical reaction in vivo was that of sucrose cleavage, and that this was essential for conversion of photosynthetically produced sucrose for starch biosynthesis. The residual amount of starch deposited was attributed to the sucrose degrading activity of a second sucrose synthase encoded by another locus (Susl).

Further research also favors the cleavage role of sucrose synthase and implicates its involvement in starch deposition. Dale and Housley (1986), for example, found that developing wheat kernels with the greatest rates of growth and starch deposition had significantly greater sucrose synthase activities. A positive correlation between sucrose synthase activity and starch deposition was also reported in Pisum sativum by Edwards and ap Rees (1986a and b). They proposed that UDP-glucose formed during sucrose cleavage was converted to glucose-i -phosphate by UDP-glucose pyrophosphorylase using pyrophosphate








10

generated by PFK(PP). Morrell and ap Rees (1986) have also suggested that much of the sucrose translocated to developing potato tubers is probably metabolized via the same pathway with the initial step catalyzed by sucrose synthase. Gibson and Shine (1983) have demonstrate that in the presence of inorganic phosphate, UDPG may be hydrolyzed to G-1 -P and UDP by the action of UDPG phosphorylase. Salerno (1986) also has demonstrated the presence a highly nucleotide specific form of UDP-glucose phosphorylase in developing maize endosperms. The activity level of this enzyme followed closely the development of the grain and paralleled that of sucrose synthase. The presence of this enzyme links sucrose cleavage and starch formation via sucrose synthase.

Finally, an additional line of evidence was presented by Cobb and Hannah (1988) that also indicated a primarily degradative function for sucrose synthase in importing organs. They showed that maize kernels from a line deficient in the Sh 1 gene for sucrose synthase still had normal levels of sucrose and normal rates of sucrose synthesis when grown in culture with fructose as the carbon source. If sucrose synthesis had been proceeding via sucrose synthase in wild-type kernels, then the loss of ca. 90% of total sucrose synthase activity in kernels of the mutant line should have affected sucrose formation there. The authors concluded that Shl encoded sucrose synthase was not necessary for sucrose synthesis.

Correlative data suggest that sucrose synthase activity is closely linked with sink strength. The supply and timing of sucrose for export seem to be closely related to the source of photosynthate (Fondy and Geiger, 1982; Servaites et al.,








11

1989) as well as the energy for phloem loading. However, once sucrose is loaded, its eventual fate does not appear to be under the control of the source leaf (Gifford and Evans, 1981) but rather is under the control of the importing sink (Wyse, 1986). Giaquinta (1979) found that young, immature roots of sugar beets had low levels of sucrose synthase, but the onset of rapid sucrose import for storage was accompanied by a significant increase in sucrose synthase activity. Similar correlations were also observed by Silvius and Snyder (1979) and Fieuw and Willenbrink (1987). In sugar beet roots sucrose uptake into parenchyma can proceed without prior hydrolysis in the apoplast or free space. Increases in sucrose synthase activity have also been observed during periods of sucrose import and/or accumulation in sweet melons (Schaffer et al., 1987), netted muskmelon (Lingle and Dunlap, 1987), eggplants (Claussen et al., 1985, 1986), rose flowers (Khayat and Zeslin, 1987), developing chick pea seeds (Setia and Malik, 1985), tomato (Yelle et al., 1988) and Solanum muricatum (Schaffer et al., 1989). tingle (1987), however, found no correlation of sucrose synthase activity with sucrose concentration in sweet sorghum.

Huber and Akazawa (1986) hypothesized that a primary role of sucrose synthase could be to feed glucose-i -phosphate directly into glycolysis. Black and coworkers (Black et al., 1987; Sung et al., 1988; 1989; Xu et al., 1989) also support this hypothesis. This link to glycolysis also involves UDPG-pyrophosphorylase, which converts the UDPG formed by the action of sucrose synthase into G-1-P and UTP. The methods employed to deliver carbohydrates to their respective








12
sinks vary from species to species (ap Rees, 1974, Hawker, 1985). However, sucrose breakdown generally appears to proceed via sucrose synthase in starch and sugar storage sinks (Sung et al., 1988).

Sucrose synthase also may be associated with sink strength through its potential involvement in cell wall synthesis (Hendrix, 1990). Sucrose synthase activity predominates over that of invertase in rapidly expanding cotton ovules, for example where rapid elongation of epidermal hairs (cotton fibers) requires extensive cellulose formation. Hendrix (1990) postulated that sucrose entering the seed coat in the developing cotton boll was cleaved via sucrose synthase, and subsequent carbohydrates went into the rapidly growing epidermal hairs. However, some of the sucrose breakdown products were converted to starch stored in the seed coats. Stepanenko and Morozova (1970) demonstrated cellulose biosynthesis from UDP-glucose in cotton. However, they did not indicate that the source of the UDPG might come from the action of sucrose synthase. Carpita and Delmer (1981) showed that the rate of synthesis and turnover of UDPglucose in developing cotton ovules was more than sufficient to account for rates of biosynthesis for cellulose, -1,3-glucan and sterylglucosides (all cell wall constituents). They found that UDPG levels increased dramatically just prior to the maximum rate of secondary wall cell synthesis and dropped precipitously at the time when cellulose synthesis ceased. Again, sucrose synthase activity was not measured, but could possibly be the source of the increased levels of UDPG. Sucrose synthase activity was measured in cultured cells of Catharanthus roseus








13

(Amino et al., 1985) and found to be elevated during the GI phase when the amount of total cell walls increased significantly. However, UDP-glucose pyrophosphorylase activity (also involved in the formation of UDPG) was greater than sucrose synthase activity at the G1 phase. The former was considered by these authors likely to make a more important contribution to the total UDPG formed. Chourey et al. (1991 a) have reiterated the hypothesis that the resultant shrunken, starch-deficient endosperm of the shl maize mutant may be due to reduced cell wall deposition rather than any direct effect on the starch biosynthetic pathway.


Regulation of Sucrose Synthase

Sucrose synthase, a key enzyme in sucrose metabolism, is subject to a number of complex regulatory factors (Davies, 1974). This enzyme exhibits a wide specificity for the nucleoside base utilized in the reaction. Most enzymes of sugar nucleoside metabolism show a marked specificity for a particular base (Avigad, 1982). Sucrose synthase working in the synthetic direction has been shown to utilize UDPG, ADPG, TDPG, CDPG and GDPG as glucosyl group donors (Avigad, 1982). The Km for UDPG, however, is usually much less than for other NDPG's. Grimes et al. (1970) found that the K, for UDPG was approximately 0.2 mM while ADPG, TDPG, CDPG and GDPG had Km'S of 1.8, 1.7,2.5 and 2.5 mM, respectively. They also found that with UDPG as the nucleoside sugar, the Km for fructose was reduced 10 fold below that apparent when ADPG was utilized. This change was








14

attributed by Grimes et al. (1970) to possibly result from conformational changes in the enzyme.

Reduced Km'S for UDPG when compared to other NDPG's have also been observed for sucrose synthase from pea seedlings (Gabrielyan et al., 1969), sweet potato root (Murata, 1971), potato tubers (Pollock and ap Rees, 1975), sweet corn seeds (de Fekete and Cardini, 1964) and soybean nodules (Morell and Copeland, 1985). In contrast, ADPG and TDPG were reportedly more efficient glucosyl donors than UDPG for sucrose synthase isolated from sorghum seeds (Sharma and Bhatia, 1980) and sugar beet roots (Avigad and Milner, 1966).

No large differences in K,'s for UDP and other nucleoside diphosphates generally are observed when the reverse reaction is analyzed. Delmer (1 972a and b) found minimal or no differences in Km's for NDP's with sucrose synthase from mung bean seedlings. She did, however, find a large difference in the rates of sucrose cleavage with different NDP's. Maxima were observed when UDP was the substrate. The Vmax for UDP in relative terms was 100 compared to 28, 6, 3 and 3 for ADP, TDP, CDP and GDP, respectively. Similar Km'S for various NDP's have also been observed in sweet potato roots (Murata, 1971), potato tubers (Pollock and ap Rees, 1975), Jerusalem artichoke (Pontis et al., 1972) and sugar beet (Avigad and Milner, 1966). Sucrose synthase from sweet corn kernels does, however, exhibit a Km an order of magnitude greater for ADP than UDP (Su and Preiss, 1978; de Fekete and Cardini, 1964). Morell and Copeland (1985) also








15

found that in soybean nodules, the Km of sucrose synthase for UDP (0.5 mM) was lower than that of ADP and CDP (0.13 and 1.1 mM, respectively).

Delmer (1 972a and b) characterized regulation of purified Phaseolus aureus sucrose synthase and found a number of differences in the regulation of the synthetic and degradative reactions. NADP, iodoacetic acid, and gibberellic acid all stimulate sucrose degradation but inhibit sucrose synthesis. Pyrophosphate also enhanced the degradative activity, but only in the presence of MgCl2. In contrast, Pontis (1977) reported that P, inhibited the degradative reaction alone or in the presence of Mg2+. Delmer also tested the effects of intermediates in carbohydrate metabolism and found that G-1 -P, G-6-P, F-6-P, F-1,6-BP, R-5-P, R1,5-BP, PEP and 3-PGA had little or no influence on the sucrose synthase reaction in either direction when present at 2 mM. However, de Fekete (1969) and Pontis (1977) both have reported that G-1 -P, G-6-P and F-1,6-BP were inhibitory to the degradative reaction at 2-5 mM without affecting the synthetic reaction. ATP, ADP and AMP had no inhibitory effect on sucrose synthesis at 4 mM; however, the degradative reaction was inhibited 30% by ADP, and 50% by both ADP and AMP. P-Phenylglucoside has also been shown to inhibit sucrose degradation almost completely and sucrose synthesis by 50% (Wolosiuk and Pontis, 1974b; Lowell, 1986). This has proven useful for distinguishing activities of sucrose phosphate synthase from sucrose synthase.

Pontis and coworkers (Pontis et al., 1972) found that the divalent cations Mg2+, Mn2+, Ca2+ and Ba2+ at 5-10 mM activated sucrose synthase in the









16

synthetic direction but inhibited the cleavage reaction. UDP was found to be a strong inhibitor of the synthetic reaction at 10 mM (70-80% inhibition), but the inhibition could be reversed by the addition of Mg2+ (de Fekete and Cardini, 1964). UDP was also found to inhibit the degradative reaction as a competitive inhibitor for UDPG (Wolosiuk and Pontis, 1974a). Inhibition by other NDP's was very weak. UTP (4 mM) caused a slight inhibition of synthetic activity, but caused an 80% inhibition of the degradative reaction (Tsai, 1974). Echeverria and Humphreys (1985), however, found that UDP and UTP within the cytosolic range (< 4 mM) both had little or no effect on sucrose synthase in the synthetic direction. UDPG was able to inhibit the cleavage reaction by 13% at 10 mM, but tissue concentrations were generally below this level (Echeverria and Humphreys, 1985), with the effect on the synthetic reaction minimal. Wolosiuk and Pontis (1974a) found that UDPG could function as a competitive inhibitor for UDP in the sucrose synthase synthesis reaction.

Carbohydrates also have been found to inhibit sucrose synthase activity in vitro. Fructose was found to function as a competitive inhibitor of sucrose in the cleavage reaction (Pridham et al., 1969; Doehlert, 1987). Sucrose had no inhibitory effects on sucrose synthase activity at saturating levels of fructose and UDPG (Echeverria and Humphreys, 1985), but glucose at 100 mM inhibited sucrose synthesis by 63%-70% and inhibited sucrose cleavage 86%-93%. Expression of sucrose synthase genes also respond to carbohydrate availability. Koch and McCarty (1988, 1990; Koch et al., 1989) have shown that levels of maize








17
root Shl mRNA were elevated when sugar supplies were limited in culture. In contrast, levels of Susl mRNA were elevated in response to increasing glucose concentrations. They speculated that the effect of sugar levels on expression of specific genes could prove to be an important control mechanism whereby plant cells could react to cellular nutritional conditions. The Shl gene is also upregulated under anaerobic conditions (Springer et al., 1986); however, the carbohydrate response of the Shl gene appears to be distinct from its anaerobic regulation (Koch et al., 1989). There is some doubt as to whether anaerobic induction occurs at both the transcriptional and translational levels in maize (McElfresh and Chourey, 1988). Taliercio and Chourey (1989) hypothesized that the expression of anaerobically induced Shl transcripts are blocked at some step beyond polyribosomal loading. However, other researchers have shown that sucrose synthase in maize is anaerobically induced at the protein as well as the gene level (Freeling and Bennett, 1985; Springer et al., 1986). Anaerobic induction of sucrose synthase at both the gene and protein levels also has been demonstrated in rice (Ricard et al, 1991) and Echinochloa phyllopogon (Mujer et al., 1990).


Invertases

Invertases (E.C. 3.2.1.26, 6-D-fructofuranoside fructohydrolases) are widely distributed in the plant kingdom and catalyze the following reaction:

Sucrose + H20 -------- > Glucose + Fructose








18

Invertases are specific for the fructofuranose moiety of sucrose and work by hydrolyzing the glycosidic linkage between the bridge oxygen and the fructose residue (Sum et al., 1980). Up to five different forms of invertase have been reported in plants (Sasaki et al., 1971). However, these enzymes are generally divided into two main types based on the pH at which sucrose hydrolysis is most efficiently accomplished. Acid invertases have pH optima around 4.5 to 5.0; that of alkaline (or neutral invertase) is 7.0 to 7.5.

Most invertases are glycoproteins. Arnold (1966) partially purified an acid invertase from grapes and found that it was approximately 25% carbohydrate. Faye and coworkers (Faye and Berjonneau, 1979; Faye et al., 1981) have shown a 7.7% carbohydrate content of acid invertase from radish seedlings, and date invertase has a carbohydrate content of 8.2% (AI-Bakir and Whitaker, 1978). Invertase preparations from barley (Prentice and Robbins, 1976), sugar cane (del Rosario and Santisopasri, 1977), potato tubers (Anderson and Ewing, 1978; Bracho and Whitaker, 1990b) and banana (Sum et al., 1980) were shown to bind strongly to concanavalin A, a phytagglutinin or lectin isolated from jack bean with a strong binding affinity for carbohydrates. In yeast and Neurospora the carbohydrate content of invertase has been estimated to range from 0 to 50 % (Metzenberg, 1963; Gascon et al., 1968; Holbein et al., 1976). Much of our more detailed knowledge on the molecular structure and mode of action of invertases comes from studies on fungal and yeast enzymes; however, considerable progress has been made in analyses of plant invertases.










Roles of Invertase

Elevated acid invertase activity is characteristic of plant tissues in which there is a need for hexoses produced from stored or recently transported sucrose (ap Rees, 1974; Avigad, 1982). Greater activities of invertase also correlate well with a low content of stored sucrose. In sugar beets, the onset of sucrose storage is accompanied by a decrease in invertase activity (Silvius and Snyder, 1979; Giaquinta, 1979). The same is true for carrot roots (Ricardo and ap Rees, 1970), melon (Hubbard et al., 1989; Lingle and Dunlap, 1987; Schaffer et al., 1987; McCollum et al., 1988), citrus (Kato and Kubota, 1978; Lowell, 1986) and Lycopersicon hirsutum (Miron and Schaffer, 1991). In these systems, invertase was very active prior to sucrose accumulation and dropped significantly upon maturity. Invertase activity is usually greatest in tissues that are at a rapid stage of growth and development (Weil and Rausch, 1990), particularly at the cell division stage (Masuda et al., 1988). Root apices, young leaves and stem internodes fall into this category. Mature leaves, functioning as sources of photosynthates, generally have low levels of apoplastic acid invertase (Dickinson et al., 1991). Transgenic tomato plants expressing yeast invertase in the apoplast of mature leaves had a striking repression of growth (Dickinson et al., 1991). The higher the level of invertase, the greater the inhibition. The general role of acid invertase, therefore, seems to be for the breakdown of sucrose where there is a marked need for hexose (ap Rees, 1974).








20
Previously, the role of invertase in sucrose transfer was considered particularly important in plants such as sugar cane and maize where substantial sugar movement occurred through the cell wall space and was accompanied by action of an extracellular invertase (Glasziou and Gayler, 1972; Hawker and Hatch, 1965). Recent evidence, however, indicates that although much hydrolysis is often observed, invertase activity may not be essential for sucrose uptake into either sugar cane stems (Thom and Maretzki, 1990; Lingle, 1989) or maize kernels (Schmalstig and Hitz, 1987). The role of apoplastic invertase in sucrose import into roots had previously been questioned by Chapleo and Hall (1 989a) who concluded that although present, apoplastic root invertase did not have a direct role in sugar transport. However, substantial activity of invertase has been widely documented in roots of plants such as pea (Lyne and ap Rees, 1971), bean (Robinson and Brown, 1952), tomato (Chin and Weston, 1973), Ricinus (Chapleo and Hall, 1989a, b, and c), oat (Pressy and Avants, 1980), and maize (Hellebust and Forward, 1962; Chang and Bandurski, 1963). Specific tissue localizations also have been described. Peak activity for root invertase is generally 2-3 mm behind the apex and corresponds to the region of expansion and elongation in pea (Robinson and Brown, 1952; Sexton and Sutcliffe, 1969) and maize (Hellebust and Forward, 1962). In Ricinus roots, this activity predominates in the cortex (Chapleo and Hall 1989a).

Although invertase may not have a direct role in sucrose import into roots, it still may be important to two major aspects of root biology. First, invertase is








21
essential to mycorrhizal associations (Purves and Hadley, 1975). Maize (Gerdemann, 1964; Kothari et al., 1990) and 90% of other agriculturally important species form these beneficial symbioses under field conditions (Gerdemann, 1968). However, sucrose must be hydrolyzed for the fungal symbiont (Long et al., 1975), and invertase levels rise at sites of carbon transfer. It is not known whether this is host or fungal invertase.

Another possible role for apoplastic invertase is in the regulation of the intercellular sucrose concentration. Regulation of the free-space sucrose concentration may be important in osmotic relations and in the control of tissue differentiation (ap Rees, 1974). Jeffs and Northcote (1966, 1967) have shown that phloem differentiation in cultures of Phaseolus vulgaris depended on the supply of sucrose; glucose or fructose would not substitute. Wright and Northcote (1972), however, have shown that phloem differentiation in cultures of Acer pseudoplatanus were equally responsive to glucose and sucrose. The results of Jeffs and Northcote show that in certain cases the regulation of the apoplastic sucrose content by acid invertase could be important in differentiation.

A definitive role cannot be assigned to alkaline invertase at present. Studies with sugar cane (Hatch and Glasziou, 1963), carrot roots (Ricardo and ap Rees, 1970), pea roots (Lyne and ap Rees, 1971), melon (Ungle and Dunlap, 1987; McCollum et al., 1988), and Lycopersicon hirsutum (Miron and Schaffer, 1991) indicate an inverse relationship between alkaline and acid invertase and a more positive correlation between alkaline invertase and sucrose concentration. The








22
maximum values for alkaline invertase activity observed to date are consistently less than those of acid invertase (Masuda et al., 1988). The possibility exists that alkaline invertase allows cells that store sucrose in their vacuoles to retain a capacity for breakdown of enough sucrose in the cytoplasm to meet respiratory and metabolic demands for hexoses (ap Rees, 1974). The capacity of a plant to produce two different invertases that are spatially separated may allow the plant cell to regulate sucrose storage independent from sucrose breakdown.


Regulation of Invertase

Acid invertases are generally found in the apoplast and vacuoles of plant tissues. Washed preparations of cell walls contain a large proportion of a plant's acid invertase (Uttle and Edelman, 1973). A portion of the acid invertase can be extracted from the cell wall during grinding, but at least some of the enzyme is considered to be attached to the cell wall in vivo (Edelman and Hall, 1965). The major determinant of how much acid invertase remains bound during extraction is the pH of the buffer used (ap Rees, 1974). Buffers with acidic pH leave most of the activity in the cell wall fraction, whereas neutral or alkaline buffers release the majority into the soluble fraction.

Early evidence suggested that the soluble and insoluble acid invertases were not simply different forms of the same enzyme. The pH optima and the Km of bound acid invertase of mature (Hawker and Hatch, 1965) and immature (Hatch et al., 1963) sugar cane storage tissue differed from those of the soluble fractions. Association with the cell wall may change an enzyme's properties (ap Rees, 1974);









23

however the differences in values, especially those of mature tissues, were considered unlikely to be wholly artifactual. Distinguishing between forms of invertase is further complicated by information obtained from yeast. One gene in yeast, SUC2, has been shown to encode the two forms of invertase in yeast, secreted and intracellular, via two differentially regulated mRNAs (Carlson and Botstein, 1982).

The Km'S of acid invertase for sucrose generally range from 2 to 13 mM. Sucrose is the primary substrate for acid invertase but raffinose also is hydrolyzed, though at a slower rate (10% to 50% the rate of sucrose) (Avigad, 1982). Acid invertase from sugar-cane leaves was inhibited competitively by fructose (K 32 mM) and noncompetitively by glucose (K, 37 mM) (Sampietro et al., 1980). Acid invertases have been partially purified from a number of tissues with apparent molecular weights ranging from 2.8 x 104 to 2.2 x 105 (Roberts, 1973; Ricardo, 1974; Kato and Kubota, 1978; Masuda and Sugawara, 1980; Sum et al., 1980; Faye et al., 1981).

The SUC2 gene of Saccharomyces, encoding invertase, has been shown to be modulated by glucose levels (Carlson et al., 1987). Sucrose or raffinose, substrates of the yeast invertase, have no such effect. Kaufman et al. (1973) found that acid invertase activity rises in Avena stem segments incubated in a sucrosecontaining medium. The response had a lag time of 10-12 hours, suggesting a change in protein levels. Fructose in the incubation medium resulted in a similar response, but glucose caused no change in invertase activity.








24
A naturally occurring acid invertase inhibitor has been detected in a number of plant tissues including beet roots (Burakhanova et al., 1987), potato roots (Pressy, 1967, 1968; Bracho and Whitaker, 1990a and b), maize endosperm (Jaynes and Nelson, 1971), pea pollen (Malik and Sood, 1976) and Ipomea petals (Winkenbach and Matile, 1970). In potato the inhibitor was characterized as a small protein, binding irreversibly to acid invertase (Pressy, 1967; Anderson and Ewing, 1978). Pressy (1967) found that the binding of the potato inhibitor to invertase had a pH optimum of 4.5 (Pressy, 1967), and the enzyme-inhibitor complex could be partially disassociated by low pH or high Mg2+ concentrations. In contrast, Bracho and Whitaker (1 990a) found no effect of pH on inhibitor binding. Sucrose at 2 mM could inhibit binding, but would not dissociate a complex already formed (Pressy, 1967). Neither glucose nor fructose had a similar effect. Matsushita and Uritani (1974) noticed a marked increase of acid invertase activity resulted from wounding of sweet potato roots, but alkaline invertase activity did not change under similar conditions. They also isolated a heat-stable protein component with a molecular weight of approximately 19.5 kD, that fluctuated during the incubation period after the wounding (Matsushita and Uritani, 1976). They found that this putative inhibitor declined with a concomitant rise in invertase activity early in the incubation, but increased in later stages when invertase activity declined (Matsushita and Uritani, 1977). Pressy (1967, 1968) and Matsushita and Uritani (1977) have suggested that the increase in invertase activity caused by cold treatment or by wounding could be explained by a decrease in binding of the








25
inhibitor. Bracho and Whitaker (1 990b) also found a positive correlation between levels of inhibitor and invertase. The possibility therefore exists that this interaction plays a regulatory role in sucrose breakdown (Akazawa and Okamoto, 1980; Avigad, 1982).

Alkaline invertase is generally considered to be cytoplasmic. It is only recovered from the soluble fraction of homogenates and has a pH optimum near neutral. Both findings support its internal localization. Km values of alkaline invertase for sucrose are slightly higher than for acid invertase, generally 9 to 25 mM. Alkaline invertase hydrolyzes raffinose very poorly (< 7% of the rate of sucrose breakdown). Morell and Copeland (1984) found that stachyose (0.1 M) also was hydrolyzed by alkaline invertase but much less efficiently than sucrose (1.5% of the rate of sucrose). Both raffinose and stachyose are polysaccharides containing a fructose moiety. Morell and Copeland (1984) also found that cellobiose, gentiobiose, maltose, turanose, lactose, melezitose, trehalose, a-methylD-glucopyranoside and p-methyl-D-glucopyranoside (all at 0.1 M) were resistant to degradative action by alkaline invertase. None of these sugars contain a fructose moiety, further confirming the specificity of invertase for the fructofuranose moiety of sucrose.

Alkaline invertase from potato tubers was inhibited only slightly by glucose (Matsushita and Uritani, 1974); glucose-6-phosphate also had a slight inhibitory effect. Fructose (15 mM) competitively inhibited soybean nodule alkaline invertase by 50% (Morell and Copeland, 1984); glucose (5 mM) inhibited activity by 7%.








26

Morell and Copeland (1984) also found that the metabolites ATP, ADP, UDP, ADPglucose, UDP-glucose, glucose-i -phosphate, glucose-6-phosphate, and fructose-6phosphate (all at 5 mM) had no inhibitory effects. They also tested the effects of various chloride salts on alkaline invertase activity and found that Na+, K+ or NH4+ at 50 mM had no effects; however, CaCl2 (10 mM) and MgCl2 (10 mM) each inhibited activity by 25%. The anions citrate and inorganic phosphate have been shown to stimulate alkaline invertase from Lupinus luteus nodules (Kidby, 1966); however, Morell and Copeland (1984) found no effect on activity of soybean nodule alkaline invertase. They did find, though, that Tris buffer was a noncompetitive inhibitor of soybean nodule alkaline invertase activity; a 0.7 mM buffer concentration inhibited activity by 50%.


Use of Mutants in Physiological Research

Despite the fact that all mutations have effects on the biochemistry and physiology of the plant, only a small number have been investigated physiologically (Vose, 1981). Advances in knowledge about the molecular bases of cell processes in eukaryotic and prokaryotic microorganisms have been achieved with an array of mutant lines, often induced, that modify or block steps in the processes under study (Nilan et al., 1981). Many mutants will, theoretically, differ in only a single major physiological character. The use of mutants is growing in comparative physiological studies because the alternative is comparison of contrasting genotypes that quite possibly may be altered in undefined characters different from the one of interest.








27
The maize plant (Zea mays L.) has been particularly useful in genetic and cytogenetic studies because of the number of mutants available (Neuffer et al., 1968). Many of the mutants also have proven useful for physiological research. The shrunken-1 mutant of maize was first described by Chourey and Nelson (1976). Less than 10% of the normal sucrose synthase activity in wild-type endosperm was observed. This reduced activity results in a "shrunken" phenotype in the dry kernel. The shl mutant has proven useful in elucidation of the role of sucrose synthase in starch formation. The residual activity of sucrose synthase present in this shl mutant was attributed to the presence of another isozyme encoded by a second gene (Chourey and Nelson, 1976). This second gene, Susl, has been mapped to the same chromosome as Shl (chromosome 9) but is located 32 map units away (McCarty et al., 1986; Gupta et al., 1988). Shl and Susl encode similar proteins. Sucrose synthase is a tetramer in its native form (Su and Preiss, 1978), and the two isozymes encoded by Sh 1 and Sus 1 are able to form heterotetrameric forms of the native protein (Echt and Chourey, 1985).

Shl has been shown to be responsive to anaerobic conditions, with transcript levels increasing 10 to 20 times in shoot and root tissue respectively compared to aerobic controls (Springer et al., 1986). However, Susl exhibits little response to anaerobic stress and seems to be expressed at a relative constant in all tissues (McCarty et al., 1986). Rowland et al (1989), however, found that Susl did show a slight response to anaerobic conditions, decreasing slightly in the lower root, primarily in the pith, root tip and root cap. A maize mutant lacking the








28
Susl gene has been described (Chourey et al., 1988), but, unlike the Shl mutant, the Susl mutant does not have any detectable phenotypic abnormality. A mutation lacking detectable levels of both sucrose synthase isozymes also has been described (Chourey, 1988), but its existence is puzzling considering the expected lethality of a complete sucrose synthase deficiency.














CHAPTER 3
INSTABILITY OF SUCROSE SYNTHASE FROM ROOT TIPS:
CHARACTERIZATION AND STABILIZATION



Abstract


Instability of sucrose synthase from root tips was characterized in maize and an assay developed to circumvent the rapid decline of activity in vitro (35 and 100% activity loss in 20 min for maize and cotton, respectively). Initially 14Csucrose cleavage was quantified by recovery of 14C-UDPG on DEAE ion exchange paper (Delmer, 1972; Su and Preiss, 1978). Subsequently, modifications were made which resulted in increased accuracy, reduced tissue volume required and reduced extraction/assay period. Phenolic protectants did not reduce the activity loss over time. Specific inhibitors for the four classes of proteinases were also tested; only PMSF increased enzyme activity, but did not completely prevent its loss over time. Stabilization and additional elevation of activity were achieved by adding casein. However, western blot analysis indicated that activity decline was not associated with any detectable proteolytic degradation, charge alteration, or subunit separation. In addition, inclusion of 10 mM P, in the extraction medium rapidly reduced activity, indicating the possible involvement of phosphorylation or nucleotide effects.










Introduction

Measurement of the maximum catalytic activities of enzymes in plant tissues can make important contributions to the understanding of metabolic pathways and their mechanisms of control (ap Rees, 1974). Currently available methods of assaying sucrose synthase have proven ineffective for many tissues, particularly those of roots (Duke et al., unpublished data; Ungle, USDA/ARS, Westlaco, TX, personal communication). A precipitous loss of activity follows tissue extraction from root tips of maize and other species (D.L. Hendrix, USDA/ARS, Phoenix, AZ, personal communication). Chan et al. (1990) reported that sucrose synthase activity in roots of rice was detectable in only one stage of growth. However, sucrose synthase protein was present in root tissue at all stages of growth, exceeding that in grain when grain activity was highest among tissues sampled. This report addresses the basis of this instability in maize roots and describes a rapid radiometric assay for sucrose synthase which circumvents this problem and allows assay of small samples. Extraction and assay were optimized for substrate concentration, pH, assay length, and inclusion (or exclusion) of various antioxidants and proteinase inhibitors. The procedure has proven effective for a range of tissues and species examined and provides an accurate measurement of activity, particularly where enzyme stability may be limiting.










Materials and Methods


Plant Material

Maize seed (Zea mays L., NK 508, W22:sh 1) were primed for 6 days at 10 C with a water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I1 captan (Bodsworth and Bewley, 1981). At the end of 6 days, the seeds were rinsed free of PEG, given a 20 min rinse in 1.05% (v/v) sodium hypochlorite and again rinsed in water for 20-30 min. Seeds were germinated in the dark at 18 C on Whatman 3mm filter paper. Moisture level was kept constant throughout. At the end of 7 days, 1 cm primary root tips were excised under a sterile transfer hood. Cotton (Gossypium hirsutum L., Coker 100) root tips were obtained from Dr. D.L. Hendrix (Western Cotton Research Laboratory, USDNARS Phoenix, AZ). One cm root tips were excised from 5- to 6-day old seedlings and quick frozen in liquid N2.


Purification of 14C-Sucrose

Trace amounts of phosphorylated sugars are common impurities in commercial 14C-sucrose, and can reduce accuracy of the assay. These were removed by descending paper chromatography of commercially obtained 14C sucrose in ethanol (NEN, Boston MA) using DEAE cellulose paper. The majority of anion-free sucrose was concentrated into the first 2 to 3 drops eluted from the V-shaped tip of the DEAE paper strip. No impurities were detected using HPLC analysis (data not shown). Molarity and specific activity of purified 14C-sucrose








32
were subsequently adjusted to 1 M and ca. 0.11 LCi per ul. Two and one-half il (ca. 0.27 pCi) were used in each reaction. Tissue Extraction

Weighed tissue (100-200 mg) was frozen and ground to a powder in liquid N2 with a mortar and pestle. The frozen powder was transferred to another mortar containing ice-cold extraction buffer (200 mM HEPES buffer [pH 7.5], with 1 mM DTT, 5 mM MgCI, 1 mM EGTA, 20 mM sodium ascorbate, 1 mM PMSF and 10% [w/w] PVPP) and ground briefly in this medium. One ml of grinding buffer was used for every 100 mg tissue fresh weight. Cysteine (10 mM) was initially but was omitted to prevent non-specific binding of radiolabel to DEAE cellulose paper.

Two-hundred pl of extract were placed on each of 4 to 8 spun columns packed with Sephadex G 50-80 hydrated with extraction buffer. Columns were centrifuged for 1 min at 800 x g. Eluent from each column was pooled with others from the same tissue sample before assay. Ratio of sample to bed volume was maximized at 1:5 (v:v) by HPLC detection of soluble sugar presence in eluent (Yelle, 1991).


Enzyme Assay

Cleavage of 14C-sucrose by sucrose synthase was assayed in a 50 'l volume consisting of 20 pl extract, 80 mM Mes (pH 5.5), 5 mM NaF, 100 mM 14Csucrose and 5 mM UDP. Reactions proceed for 5 minutes at 30 C and were terminated by adding 50 pl of Tris (pH 8.7) and boiling for 1 min. Controls









33

contained all assay components except UDP. The assay was optimized for pH, linearity with time and protein concentration (data not shown). Product Determination

The entire reaction volume was blotted onto a small disk of DEAE ionexchange paper (2.4 cm diameter) and dried completely before rinsing. Each disk was rinsed separately, first in 40 ml of H20 at 175 rpm on a rotary shaker for 2 hours, again for an additional 2 hours and finally rinsed in a gentle stream of DI water for 30 sec. Remaining radiolabel was quantified and compared to total amount of the 14C-sucrose substrate utilized to determine extent of sucrose cleavage.


Protein Gel Blots

Subsamples from protein extracts to be used for enzyme assays were separated on native PAGE using the system of Laemilli (1970) with (denaturing) or without (native) SDS. Polyacrylamide concentrations of the stacking and separating gels were 2.5% and 5%, respectively. Proteins were resolved at 4 C by applying 15 V for 9 h, then 125 V for 11 h (constant current and temperature (4 C). Each lane was loaded with 2 /sg of total protein. Proteins were electroblotted to nitrocellulose membranes and probed with polyclonal antibodies following the procedure of Towbin et al. (1979). Sucrose synthase antisera, obtained from D.R. McCarty, was generated in rabbits using protein purified from maize kernels (W64 x 182E) 22 days after pollination. Antisera was diluted 1:1000








34

and cross reacted strongly to both the Shi and Susl gene products where such were present.


Results

Activity of sucrose synthase from maize and cotton root tips declined rapidly after extraction (Figure 3-1). The greatest decrease in activity occurred between 10 and 15 minutes after extraction from both species. Uttle or no activity was observed after 4 hours (data not shown). After extraction, extracts were maintained at 0 C until used in the radiometric assay.

A wide range of enzyme protectants were examined. No improvement in activity was observed when the polyphenol protectants PVP-40, PEG 20,000 and BSA were utilized (Table 3-1). PVPP, also a phenol absorbent, was utilized in each extraction. In addition, four classes of proteinase inhibitors were tested for their effect on stability of sucrose synthase activity. Addition of leupeptin (1 mM) slightly decreased initial activity (Table 3-1), and pepstatin-A (1 mM) had no effect (Table 3-1). Phenylmethylsulfonyl fluoride (PMSF) (1 mM), a serine proteinase inhibitor, had a substantial positive effect, as did casein (2% w:v), potentially a non-specific proteinase inhibitor.

Further characterization of activity change in the presence of PMSF showed that stabilization was not effective in the first 20 min following extraction (Fig. 3-2). Although total activity prior to this time was elevated by addition of PMSF, a linear decrease was not prevented from occurring.















o1.5
L.

0) E
E 1 -- Maize




0
D0.5 Cotton
Cnn
0



0
0 10 20 30 40 50 Time after extraction (min)


Figure 3-1. Time course of in vitro decrease in sucrose synthase activity in maize
and cotton roots. Bars represent � SE, n=3.










Table 3-1. Effect of enzyme protectants on activity of sucrose synthase from maize
root tips assayed five minutes after extraction.



Protectant (concentration) enhancement of control


PVP-40 (5 %) -6 PEG-20,000 (2% w:v) +6 BSA (2% w:v) +6 Caproic acid (2 mM) 0 Pepstatin A (1 mM) -5 Leupeptin (1 mM) 0 PMSF (1 mM) +46 Casein (2% w:v) + 15


Note: Each protectant was included in the buffer used for extraction and equilibration of desalting columns. PVP-40, PEG-20,000 and BSA were utilized to protect against phenolic compounds; PVPP was included in each extraction. Representative inhibitors of proteinase classes were: pepstatin A (aspartic), leupeptin (cysteine), EGTA (metallo) included in each extraction, caproic acid (serine) and PMSF (serine). Casein was included as a general, non-specific proteinase inhibitor.















--2.5
t-"
C

0
+ PMSF
-2
E







0 (D -5
0
b 0.5
CO

0 p p I
0 10 20 30 40 50 Time after extraction (min)


Figure 3-2. Time course of in vitro decrease in maize sucrose synthase activity with
and without the serine proteinase inhibitor, PMSF. No other class-specific proteinase inhibitors tested affected initial sucrose synthase activity. PMSF (1 mM) was used in extraction buffer and equilibration of desalting columns.
Bars represent � SE, n=3.








38

Addition of casein increased initial enzyme activity compared to controls (Table 3-1) and improved stabilization of sucrose synthase activity with time (Figure 3-3A & B).

In contrast to added protectants, inorganic phosphate (10 mM), a protein regulator through its role in reversible phosphorylation (Bennett, 1984), added to the extraction buffer decreased initial activity of sucrose synthase by ca. 40% (Figure 3-3A & B).

Despite loss of activity in vitro and positive responses to apparent protectants against protease activity, proteolytic degradation of sucrose synthase from maize root tips was not detectable via either denaturing or native (Figure 3-4A & B, respectively) western-blot analysis at various times after enzyme extraction. Further, no change in charge or separation of subunits in native tetramers was evident. Nor was any change evident with time from samples extracted with added casein or phosphate (Figure 3-4C & D, respectively). However, changes in enzyme activity do not necessarily result in changes in electrophoretic mobility. Walker and Huber (1989) demonstrated that activation of sucrose phosphate synthase by light or mannose (a P, sequestering sugar) did not affect immunoprecipitation or mobility of subunit mobility during SDS-PAGE.

The possible involvement of tetramer stability in the loss of activity in vitro was further examined by comparison of the extracts from shi (containing only homotetramers of Susl encoded sucrose synthase) and Shl (containing both hetero- and homotetramers of sucrose synthase). Endosperm tissue contains











+ Casein


Buffer only


2.5

,2.0 c1.5 21.0 CDQ.5


-2.5

E2.0 81.5
_0
C.)
751.0

0.5


+ Casein


Buffer only


+ Pi


0 10 20


30


40


50


60


70


Time after extraction (min)
Figure 3-3. Time course of in vitro decrease in maize root sucrose synthase
activity in the presence and absence of either Pi (10 mM) or casein (2%
w:v).


+ Pi


--MEMO












Time after Extraction

(min)

4 10 20 60

a-, - --


5 15 20 60


B $$NOW



Duin" "U
D



Figure 3-4. Denaturing (A) and native (B,C,D) protein gel blot analysis of maize
root sucrose synthase at various times after extraction. Sub-samples were removed at designated intervals during incubation at 4 C. Proteins were separated by polyacrylamide gel electrophoresis with (A) or without (B,C,D) SDS and sucrose synthase resolved by probing with a polyclonal antibody raised against protein products of both the Shl and Sus genes. The five bands visible in the native gel blot have been described as corresponding to homo- and heterotetrameric forms of sucrose synthase composed entirely of products from the Shl gene (uppermost band), the Sus gene (lowermost band) and combinations of the two (middle three bands).
Extracts for denaturing (A) and native (B) blots were extracted with buffer only. Casein (2% w:v) (C) and Pi (10 mM) (D) were tested as protectants
of enzyme stability.









41
tetramers composed only of Shl encoded protomers (Chourey et al., 1986; Heinlein and Starlinger, 1989; Rowland and Chourey, 1990), and sucrose synthase activity is stable during extraction and dialysis procedures (Chourey and Nelson, 1976; Echt and Chourey, 1985). In extracts from root tissue, the five bands shown by western blot represent the possible combinations of monomers of the two separate isozymes (Shl and Sus) to form the native tetrameric structure (Echt and Chourey, 1985). Heterotetramers could theoretically be more unstable than homotetramers since, although very similar, the two subunits are not identical (Echt and Chourey, 1985). Su and Preiss (1978) found that sucrose synthase tended to polymerize to an inactive polymeric after extraction. Results indicated that formation of the native enzyme from two different isozymes was not a contributing factor in loss of enzyme activity over time (Figure 3-5A & B). Despite differences in the absolute values of sucrose synthase activity from shl vs. Shl, the percent decline in activity was similar for both genotypes.


Discussion

Sucrose synthase activity was stabilized in vitro and an assay developed which enabled accurate measurement of enzyme action in root tips. The described assay allows rapid product recovery in instances where activity is otherwise unstable in vitro, and increases sensitivity to the extent that sample volumes as small as 100 to 200 jg can be used.

Sucrose synthase has previously been assayed in both synthetic and cleavage directions (Avigad and Milner, 1966; Grimes et al., 1970; Pontis et al.,






















Figure 3-5. Time course of in vitro decrease in maize root sucrose synthase activity from lines with homo- (W22:shl) and
heterotetrameric (NK 508) forms of this enzyme. Bars represent � SE, n=3. Data from one replicate using material isogeneic to W22:shl except for the Shl gene (W22) resulted in a curve similar in appearance and of the same
magnitude as that of W22:sh 1.








A B
- :1.8
C 0.8

" 1.6
7. 0.7
1.4 NK 508 W22: shl
-5
E
S1.2 0.6
ai)
0.5 )0.8
0
%- 0.4 " 0.6
CD)
0.4 3 3 . ' U 1 3 ' �5 0.3
40 10 20 30 40 0 10 20 30 40 50
Time after extraction (min)








44
1970; Salerno et al., 1979; Keller et al., 1988; Lowell et al., 1989). Measurements of the synthetic reaction have been based on quantification of either sucrose or UDP production. Sucrose levels can be determined indirectly by using invertase for full conversion to hexoses and measuring glucose colorimetrically (Avigad and Milner, 1966). The latter method, however, is susceptible to interference by substances in the crude enzyme extracts of many plants (Pontis, 1977). It has also been possible to measure 14C-sucrose formed from UDp-14C-glucose by separating labeled product from substrate with anionic resins (Salerno et al., 1979), paper electrophoresis (Grimes et al., 1970) or paper chromatography (Pontis, 1970). Such radioactive assays have proven useful in systems where colorimetric methods have been problematic (Pontis, 1977). In addition, UDP production can be determined spectrophotometrically by coupling its formation to the pyruvate kinase-lactate dehydrogenase reaction and measuring the decrease in absorbance due to oxidation of NADH (Avigad, 1964; Avigad and Milner, 1966; Lowell et al., 1989).

Procedures for assaying the cleavage reaction are based on determination of fructose or UDP-glucose formation. Fructose can be measured colorimetrically by the Nelson reducing sugar assay (1944), or spectrophotometrically by coupling hexokinase, phosphoglucose isomerase and glucose 6-phosphate dehydrogenase for production of NADPH (Avigad, 1964; Keller et al., 1988). Also, UDP-glucose formation can be estimated by coupling its appearance to NAD reduction by UDPdehydrogenase (Avigad, 1964; Lowell, 1986; Lowell et al., 1989). Degradative








45
action of sucrose synthase can also be coupled to that of UDPglucopyrophosphorylase (Xu et al., 1986; Sung et al., 1989).

Radioactive assays of sucrose synthase in the cleavage direction measure the incorporation of 14C-glucose into UDP-glucose from 14C-sucrose (Delmer, 1972; Su and Preiss, 1978). These procedures are among the most sensitive of assays for sucrose synthase (Avigad, 1982). The sugar nucleotide formed can be separated from the excess 14C-sucrose by paper chromatography (Wolosiuk and Pontis, 1974a) or by ion exchange paper (Delmer, 1972a and b; Su and Preiss, 1978). The current procedure utilizes the sensitivity of a radiometric assay along with reduced time from extraction to assay termination and results in a method suitable for time-labile extracts from small tissue samples.

Instability of sucrose synthase was further characterized using this sensitive method in an attempt to better define factors affecting the activity of this key enzyme in vitro. Sucrose synthase is a sulfhydryl enzyme and is sensitive to inhibition by phenolics and oxidized polyphenols (Pontis, 1977). Typical effectors of activity reduction examined during the present study showed that phenolic compounds did not appear to be the primary cause of the sucrose synthase instability observed. No phelolic protectant was able to preserve sucrose synthase activity over time.

Sucrose synthase has been shown to be sensitive to serine proteinases (Wolosiuk and Pontis, 1974b). In their study, trypsin caused a 70% decline in the degradative reaction and a 30% reduction in the synthetic reaction after a 15








46
minute incubation. However, chymotrypsin, also a serine proteinase, and papain, a cysteine proteinase, had no effect. In the current study, PMSF, a serine proteinase inhibitor, gave an increase in initial measurements but not did not prevent the observed short-term loss of activity with time. Echt and Chourey (1985) observed that PMSF did not stop the loss of activity of sucrose synthase from maize endosperm during long term storage. No other specific proteinase inhibitor affected stability of the enzyme from maize root tips. In addition to the specific proteinase inhibitors, casein and BSA were included in some extractions. Due to casein's complex composition and random structure, it undergoes proteolysis with all the known proteolytic enzymes (Reimerdes and Klostermeyer, 1976). Casein increased initial measurements and stabilized activity with time (Figure 3-3A & B). BSA, however, was much less effective. Casein has also been found to preserve the longevity of sucrose synthase extracted from sugar cane (S. Ungle, USDA-ARS, Westlaco, TX, personal communication). Casein (0.75-3.0%) has also been shown to effectively stabilize and increase the initial activity measurements of sucrose phosphate synthase (Raghuveer and Sicher, 1987). Addition of casein is not always feasible, however, especially in instances where accurate quantification of total tissue protein is important.

Another possibility for the regulation of sucrose synthase is through phosphorylation. Many enzymes undergo reversible phosphorylation as a means regulating activity (Bennett, 1984). Increased inorganic phosphate levels added to buffers used for enzymatic extraction provide a substrate for protein kinase








47

activity (Bennett, 1984). Sucrose phosphate synthase (SPS), an enzyme of carbohydrate metabolism, has been shown to be regulated in this manner (Doehlert and Huber, 1983; Walker and Huber, 1989; Huber et al., 1989a). Increases in extractable SPS activity are noted after illumination or inclusion of mannose or glucosamine (phosphate sequestering agents) in darkness (Huber et al., 1989b). Inorganic phosphate (5-10 mM) was found to be a potent inhibitor of SPS (Amir and Preiss, 1982), with the inhibition becoming more sensitive in the presence of Mg2+. Sucrose synthase has also been shown to have exhibit diurnal fluctuations in activity (as does SPS) (Hendrix and Huber, 1986; Vassey, 1989) and be affected by P,. Pontis (1977) reported that Pi (2-5 mM) inhibited the degradative reaction of sucrose synthase alone or in the presence of Mg2+. Delmer (1 972a), however, found that 2 mM Pi had no effect on the rates of either the forward or reverse reactions. Sucrose synthase extracted from maize roots has also been shown to be phosphorylated in vitro (Xu and Koch, University of FL, unpublished data). In the current study, no effect of added phosphate on protein stability was noted (Figure 3-4D). However, initial sucrose synthase activity measurements were less than controls, and a loss of activity with time was observed (Figure 3-3A & B).

The possibility existed that disassociation of subunits from tetramers may have effected activity. Two separate isozymes in maize (Shl and Sus) form subunits which appear to combine randomly into tetramers in root tips and other tissues (Chourey et al., 1986). Endosperm sucrose synthase tetramers, however,









48
are almost entirely composed of subunits encoded by the Shl gene (Chourey and Nelson, 1976; Chourey, 1981; Chourey et al., 1986) and are remain active during extraction (Echt and Chourey, 1985). Five different types result in those tissues which exhibit polymerization of both protomers; two are homotetramers and three are heterotetramers. Heterotetramers could theoretically be more unstable than homotetramers since, although very similar, the two subunits are not identical (Echt and Chourey, 1985). Data (Figure 3-5A & B) indicate that greater instability of heterotetramers relative to homotetramers of sucrose synthase was not the cause of the observed activity loss. The profile of declining activity with time is similar in extracts from root tips of a mutant line having only homotetramers of Shl (SS1) subunits (W22:shl) as it is in extracts from wild type kernels with 5 native tetrameric combinations (NK 508).

Rapid loss of sucrose synthase activity with time in maize and other root tips, as well as small tissue size, necessitated the development of a rapid and sensitive assay. Other assays for sucrose synthase using radiometric techniques have been described (Delmer, 1972a and b; Su and Preiss, 1978; Salerno et al., 1979). However, the procedure described here has proven effective and useful due to reduced time from extraction to assay termination, reduced sample size and increased enzyme stabilization.














CHAPTER 4
SUCROSE SYNTHASE ACTIVITY IN WILD-TYPE MAIZE ROOT TIPS RESPONDING TO ALTERED CARBOHYDRATE STATUS


Abstract


The two genes encoding sucrose synthase isozymes in maize (Shl and Susl) have been shown to respond to altered tissue carbohydrate status in root tips; Shl expression is favored by carbohydrate depletion whereas Susl is upregulated when sugars are plentiful (Koch and McCarty, 1988, 1990; Koch et al., 1989). Response at the level of enzyme activity was tested in the present study by assaying sucrose synthase activity in excised maize root tips after 24 h of incubation in a range of glucose concentrations. Little change was evident at the level of total sucrose synthase activity; however, this represented the collective responses of different isozymes and tissue types.


Introduction

Systems for changes in gene expression in response to altered carbohydrate conditions have been reported for mammalian cells (Lin and Lee, 1984) and in bacteria and yeasts (Carlson, 1987; Schuster, 1989). Recently, seven photosynthetic genes in maize protoplasts have been shown to be repressed and coordinated by sugars (Sheen, 1990). Regulation in higher plants could have









50
important implications for the control of carbohydrate distribution and utilization. Sucrose synthase is considered to have a key function in the allocation of sucrose to various plant organs, and plant carbohydrate status could function as a means of coarse regulation for activity of this enzyme.

Sucrose metabolism is important to the majority of plant species because of the nearly ubiquitous role of this sugar in phloem transport to growing and developing plant parts (Avigad, 1982). Two enzymes can catalyze the initial breakdown of sucrose, invertase or the reversible enzyme sucrose synthase. Recently, expression of the gene encoding the shrunken-1 isozyme of sucrose synthase in maize has been shown to be sensitive to carbohydrate levels (Koch and McCarty, 1988; Koch et al., 1989). Northern blot analysis of shrunken-1 mRNA showed levels were elevated in response to carbohydrate depletion. This regulation is distinct from the previously characterized anaerobic induction (Springer et al., 1986; Koch and McCarty, 1988). Although the anaerobic induction of Shl has received considerable attention in several systems, questions remain regarding the extent to which transcription and translation are synchronized under these conditions. The anaerobic induction in maize has been reported to occur only at the transcriptional level without concomitant changes in protein levels (McElfresh and Chourey, 1988; Taliercio and Chourey, 1989). However, translation of anaerobically induced sucrose synthase mRNA in rice (Ricard et al., 1991) has been demonstrated. The present work examines enzyme-level responses to changes in root carbohydrate status known to alter levels of Shl and Susl mRNA.








51
Regulation by sugar concentration may prove to be an important control mechanism whereby plant cells are able to react to cellular carbohydrate status.


Materials and Methods

Maize seed (Zea mays L, NK 508) were primed for 6 days at 10 C at a water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I"1 captan (Bodsworth and Bewley, 1981). Seeds were subsequently rinsed free of PEG, soaked for 20 min in 1.05% (vlv) sodium hypochlorite and rinsed for 20-30 min with ca. 5 I of water. Seeds were germinated in the dark at 18 C on moist filter paper in covered glass pans. Continuous airflow was provided (1 liter min"') throughout the germination period with 40% 02 supplied during the final 48 h. At the end of 7 days, 1 cm primary root tips were excised under a sterile transfer hood.

Excised root tips (ca. 750 mg per treatment) were incubated in 100 ml sidearm flasks containing 50 ml of sterile White's basal salt mixture (White, 1963) supplemented with a range of glucose (0, 0.2, 0.5, 2.0, and 4.0%). Aerobic conditions were maintained during 24 h incubations in the dark at 18 C by slow agitation on a rotary shaker (125 rpm) and an airflow of 40% 02 (11 min-) through an airstone in each flask. Experiments were terminated by twice rinsing in sterile water, blotting excess moisture and freezing them in liquid N2.

Sucrose synthase activity was determined using a rapid radiometric procedure developed to circumvent enzyme instability previously observed upon extraction from maize root tips (Chapter 3).











Results

Sucrose synthase activity was consistently maximal in root tips supplemented with 0.5% glucose (Table 4-1), a level at which the combined levels of mRNA from the two sucrose synthase genes was also greatest (Koch et al., unpublished data). Overall, however, activity of sucrose synthase in whole root tips was not significantly changed by alteration of carbohydrate status by exogenous sugar supply (Table 4-1). It was not possible to distinguish activities of isozymes encoded by the two sucrose synthase genes. These genes, Sh 1 and Susl, were found to exhibit reciprocal responses at the mRNA level to sugar availability in the same sets of roots used for these experiments (Koch et al., unpublished data). Also, changes in distribution of sucrose synthase protein among tissues within these root tips (Nolte, unpublished data) were not reflected at the level of whole root enzyme activity.


Discussion

Reciprocal regulation of the two isoforms by carbohydrate levels, as has been demonstrated for genes encoding for these isozymes (Koch and McCarty, 1989), could explain the lack of significant differences detected between glucose treatments. The two isozymes of sucrose synthase from maize (encoded by the Shl and Susl genes) are very similar, differing only slightly in their electrophoretic movement during PAGE (Echt and Chourey, 1985). The Shl and Susl encoded proteins are capable of catalyzing the same reaction with little difference in affinities










Table 4-1. Total sucrose synthase activity in wildtype maize root tips incubated in
a range of glucose concentrations for 24 hours.


intacty % glucose


0 0.2 0.5 2.0 4.0



(Mmol sucrose mg"1 protein h-')

Expt. 1 0.9 1 .2 0.9 1.6 0.9 1.5 Expt. 2 0.5 0.5 0.4 0.6 0.6 0.6 Expt. 3 1.0 0.7 0.8 1.3 0.6 0.7 Mean 0.8 0.8 0.7 1.1 0.7 0.9
S.E.M. �0.2 �0.2 �0.2 �0.3 �0.1 �0.3

Ylntact refers to root tips quick frozen in liquid N2 immediately after excision.








54

for substrates (Echt and Chourey, 1985), and both are present in extracts from wildtype maize root tips (Chourey et al., 1986). Therefore the total amount of measured sucrose synthase activity would be due to a combined complement of sucrose synthase protein. Despite close homology, however, they have been shown to be distinctive proteins encoded by separate genes (Chourey, 1981; Echt and Chourey, 1985). The two proteins also are distinct in their localization within the maize plant. The protein encoded by Shl is primarily located in the endosperm (Chourey and Nelson, 1976; Chen and Chourey, 1989), in the root (Springer et al., 1986; Chourey et al., 1986) and in etiolated shoot (Springer et al., 1986). The Susl encoded protein is generally distributed throughout the plant (Chourey, 1981; Echt and Chourey, 1985; Chourey et al., 1988).

Carbohydrate responsive proteins have been identified in roots of pearl millet (Baysdorfer and Van der Woude, 1988). Webster and Henry (1987) have also identified an unknown protein with a molecular weight similar to that of the subunits of sucrose synthase in pea root meristem cells undergoing sugar starvation. This protein, however, has yet to be positively identified. Initial findings by Koch and coworkers (Koch and McCarty, 1988, 1990; Koch et al., 1989) indicated that the Shl gene of maize was stimulated by low carbohydrate conditions and down-regulated under carbohydrate sufficient conditions. The Sus 1 gene responded in an inverse manner. Maas and co-workers (Maas et al., 1990) demonstrated that the promoter from the Shl gene was repressed by high








55

sucrose conditions. However, Salanoubat and Belliard (1989) found that increased sucrose levels promoted genes encoding sucrose synthase in potato.

The possibility also exists that protein fluctuations did not occur after 24 h incubation. However, the shifts in protein localization noted under the same conditions (K Nolte, University of Florida, unpublished data) indicate that some protein level changes did occur. Spatial separation within root tissue also could explain the differential response of Sh 1 and Sus 1 observed at gene level without a concomitant change in total enzyme activity. The distribution of sucrose synthase isozymes has been shown to be developmentally regulated, and changes during kernel development (Heinlein and Starlinger, 1989). Chen and Chourey (1989) have reported that expression of sucrose synthase genes is spatially and/or temporally separated in endosperm cells but not in root cells. However, Rowland et al. (1989) demonstrated tissue specific localization of both sucrose synthase genes and isozymes in roots undergoing anaerobic stress. K. Nolte (University of Florida, unpublished data) has shown that shifts in sucrose synthase protein localization occur in maize root tips under carbohydrate depleted and carbohydrate sufficient conditions. Increases of one isozyme in a particular tissue within the root along with decreases of the other in a different tissue would not be apparent at the level of total root sucrose synthase activity. The lack of significant differences in sucrose synthase activity of wildtype maize roots under carbohydrate sufficient and depleted condition, therefore, does not demonstrate that differences evident at the gene level are not also event at the translational








56
level. Occurrence of maximal enzyme activities in each experiment from samples having the highest levels of both sucrose synthase genes, in fact, tends to indicate that protein changes might be occurring but are not completely detectable under the assay conditions utilized.














CHAPTER 5
SUGAR RESPONSE OF SUCROSE SYNTHASE AT THE GENE (Susl), PROTEIN
AND ENZYME ACTIVITY LEVELS IN ROOTS OF THE Shl MAIZE MUTANT


Abstract


The shl mutant of maize was used to study expression of the Susl gene for sucrose synthase in response to sugar availability because this mutant has only one isozyme gene (Susl) for sucrose synthase and provides a system uncomplicated by the presence of the second gene (Shl). Koch and McCarty (1988, 1990) have previously demonstrated that Susl is up-regulated by plentiful supplies of metabolizable sugars and down-regulated under carbohydrate depletion, whereas Shl responds in an inverse manner. Excised root tips from shl were incubated for 24 h in White's basal salts medium supplemented with different amounts of glucose. Susl mRNA levels were approximately 5-fold greater in treatments with 2.0% vs. 0% or 0.2% glucose. This difference was also reflected in western blot analysis of sus protein. Enzyme activity was elevated 2-fold in root tips from 2% glucose treatments vs. those in 0 or 0.2%. Time-course and switching experiments showed that changes in mRNA or protein were not evident until 24 h and indicated that the response to carbohydrate level had been initiated within 16 h. Roots incubated in 2.0% glucose for 16 h and switched to 0% for 32 h (total of 48 h) responded like those remaining continuously in 2.0% glucose.









58
Overall, enhanced expression of Susl was evident at the mRNA, protein and enzyme levels.


Introduction

Changes in gene expression by carbohydrates have been documented as mechanisms by which bacteria and yeasts respond to changes in their nutrient status (Carlson, 1987; Schuster, 1989). Glucose-responsive genes have also been described in mammalian cells (Un and Lee, 1984). In addition, Sheen (1990) has presented evidence that the transcriptional activity of promoters of seven photosynthetic genes from maize protoplasts are repressed and coordinated by sugars. Regulation of this type in higher plants could have important implications for carbohydrate allocation and utilization.

Sucrose and its metabolic products are important to almost all plant species because of the nearly universal role of this sugar in growth and development (Avigad, 1982). Initial breakdown of sucrose can be catalyzed by either invertase or the reversible enzyme sucrose synthase. Recently, the gene encoding the shrunken-1 isozyme of sucrose synthase in maize has been shown to be responsive to carbohydrate levels (Koch and McCarty, 1988; Koch et al., 1989). Gel blot analysis of shrunken-1 mRNA showed levels were elevated in response to carbohydrate depletion. This may prove to be an important control mechanism whereby plant cells are able to react to cellular nutritional conditions. The Shi gene has also been shown to be regulated by anaerobic conditions (Springer, et al., 1986); however, effects on this gene by altered carbohydrate status are distinct








59
from regulation by anaerobic conditions (Koch and McCarty, 1988). The anaerobic induction has been reported to occur only at the transcriptional level without differences in protein levels (McElfresh and Chourey, 1988; Taliercio and Chourey, 1991). Possible changes in protein levels and enzyme activity of sucrose synthase due to carbohydrate regulation have been difficult to detect because of the nonspecificity of assay methods (Duke and Koch, unpublished).

The two isozymes of sucrose synthase from maize (encoded by the Shl and Susl genes) are very similar, differing only slightly in their electrophoretic movement during PAGE (Echt and Chourey, 1985). Despite close homology, they are distinctive proteins encoded by separate genes (Chourey, 1981; Echt and Chourey, 1985). The two proteins are, however, distinct in their localization within the maize plant. The protein encoded by Shl is primarily located in the endosperm (Chourey and Nelson, 1976; Chen and Chourey, 1989), in the root (Springer et al., 1986; Chourey et al., 1986) and in etiolated shoot (Springer et al., 1986); however Shl mRNA does appear in other tissues such as pollen grains (Hannah and McCarty, 1988). The Susl encoded protein is more widespread in its localization and is found throughout the plant (Chourey, 1981; Echt and Chourey, 1985; Chourey et al., 1988). The distribution of both proteins is further distinguished under stress conditions (such as anaerobiosis) where tissue-specific localization in roots is readily apparent (Rowland et al., 1989). Tissue specific shifts in sucrose synthase have also been noted in wildtype maize root tips incubated in glucose deficient and sufficient media (K Nolte, University of Florida,









60
unpublished data). Mararia and co workers (Mararha et al., 1990) have found that the two genes encoding sucrose synthase in wheat (SsI and Ss2) also show a differential response to stress conditions (anaerobiosis, cold shock and light).

Webster and Henry (1987) reported an unknown protein with a molecular weight similar to that of the subunits of sucrose synthase in pea root meristem cells undergoing sugar starvation. Carbohydrate responsive proteins have also been found in roots of pearl millet (Baysdorfer and VanDerWoude, 1988). These proteins, however, are yet to be definitively identified. Initial findings by Koch and co workers (Koch and McCarty, 1988, 1990; Koch et al., 1989) indicated that the Shl gene of maize was stimulated by low carbohydrate conditions and downregulated under carbohydrate sufficient conditions. The Susl gene responded in an inverse manner. Maas and co-workers (Maas et al., 1990) demonstrated that the promoter from the Shl gene was repressed by high sucrose conditions. However, Salanoubat and Belliard (1989) found that increased sucrose promoted genes encoding sucrose synthase.

The two genes encoding sucrose synthase in maize respond to altered carbohydrate status (Koch and McCarty, 1988, 1990; Koch et al., 1989), and shifts in sucrose synthase protein localization have been observed under the same conditions (K Nolte, University of Florida, unpublished data). However, these studies were carried out using a maize line having both sucrose synthase genes present. The present study utilizes the Shrunken-1 mutant of maize to determine









61
the effects of varying carbohydrate conditions on the Susl gene and its sucrose synthase gene product free from the confounding effects of Shl.


Materials and Methods

Maize seed (Zea mays L., W22:sh1) were primed for 6 days at 10 C with a water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I captan (Bodsworth and Bewley, 1981). Seeds were then rinsed with water, soaked for 20 min in 1.05% (v/v) sodium hypochlorite and rinsed again for 20-30 min with ca. 5 liters of water. Germination took place in the dark at 18 C on moist filter paper in covered glass pans. Continuous airflow was provided (1 liter min-1) throughout the germination period with 40% 02 supplied during the final 48 h. At the end of

7 days, 1 cm primary root tips were excised under a sterile transfer hood.

Excised root tips (ca. 750 mg per treatment) were incubated in 100 ml sidearm flasks containing 50 ml of sterile White's basal salt mixture (White, 1963) supplemented with a range of glucose (0, 0.2, 0.5, 2.0, and 4.0%). Aerobic conditions were maintained during 24 h incubations in the dark at 18 C by slow agitation on a rotary shaker (125 rpm) and by an airflow 40% 02 (1 "') through an airstone in each flask. Experiments were terminated by twice rinsing root tips in sterile water, blotting excess moisture and freezing them in liquid N2.

Responses of root tips to incubation in 0% vs 2.0% glucose were examined after 16, 24, or 48 h. Effects of treatment reversals at 16 h were also studied by switching roots from 0% glucose treatments to 2.0% glucose and vice versa, then continuing incubations for a total of 48 h.











Enzyme Assay

Sucrose synthase activity was determined by a rapid radiometric procedure developed to circumvent enzyme instability previously observed upon extraction from maize root tips (Chapter 3).


RNA Extraction and Northern Blotting

Samples were ground to a fine powder in a mortar and pestle with liquid N2 and RNA extracted according to McCarty (1986). Total RNA was quantified by absorbance at 260 nm.

Total RNA was separated by electrophoresis in 1% agarose gels containing formaldehyde (Thomas, 1980), blotted to a nylon membrane (Hybond-N, Amersham Corporation, Arlington Heights, IL) and probed as per Church and Gilbert (1984) with genomic clones of Sus1 (McCarty et al., 1986) radiolabeled by random primer. Blots were rinsed and placed on X-ray film at -80 C.


Protein Gel Blots

Subsamples from protein extracts to be used for enzyme assays and separated on native PAGE using the system of Laemilli (1970) without SDS. Polyacrylamide concentrations of the stacking and separating gels were 2.5% and 5%, respectively. Proteins were resolved at 4 C by applying 15 V for 9 h, then 125 V for 11 h (constant current) and temperature (4 C).

Proteins were electroblotted to nitrocellulose membranes and probed with polyclonal antibodies following the procedure of Towbin et al. (1979). Sucrose








63

synthase antisera, obtained from D.R. McCarty, was generated in rabbits using protein purified from maize kernels (W64A x 182E) 22 days after pollination. Antisera was diluted 1:1000 and cross reacted strongly to both the Sh 1 and Sus 1 gene products where such were present.


Results

Levels of Susl mRNA in excised maize roots were greater after 24 h of incubation in 2% glucose than in those that had received 0 or 0.2% glucose (Figure 5-1). At the protein level, western blot analysis showed little or no change with increasing carbohydrate concentration (Figure 5-2); however, enzyme activity was elevated in root tips incubated at high vs. low glucose concentrations (Table 5-1). Both lines of evidence indicated the protein level response was less pronounced at 24 h than that of mRNA.

The time-course of changes in Sus 1 message levels in root tips showed that differences between those given 0% vs. 2.0% exogenous glucose became apparent sometime between 16 and 24 h (Figure 5-3). Initial decreases appeared to occur in both treatments, but within 24 h, Susl mRNA levels in glucose supplemented roots had risen well above those with limited sugar supply. The greatest difference between carbohydrate treatments was evident after 48 h of incubation. Treatment reversals indicated that the gene response to carbohydrate level had been initiated within 16 h (Figure 5-3). Roots incubated in 2.0% glucose for 16 h and switched to 0% for 32 h (total of 48 h) responded like those remaining continuously in 2.0% glucose. Roots initially deprived of glucose and then












% glucose


Intact 0 0.2 0.5 2.0 4.0


Expt. 1


I


Expt. 2


Figure 5-1. RNA gel blot analysis of Susl expression in maize roots incubated in
a range of glucose concentrations for 24 hours.













% glucose


0 0.2 0.5 2.0 4.0


Expt. 2


-W -0 OM -PO r0


Figure 5-2. Protein gel blot of Susl encoded sucrose synthase from maize roots
incubated in a range of glucose concentrations for 24 hours. Data from Expts. 1 and 2 were obtained from the same set of roots sampled for RNA
analyses shown in Figure 5-1.


Intact


Expt. 1










Table 5-1. Sucrose synthase activity in mutant maize (W22:shl) root tips
incubated in media containing a range of glucose concentrations for 24
hours.


Sucrose synthase activity % glucose

Intact 0 0.2 0.5 2.0 4.0 (/imol sucrose g-1 protein h-1) Expt. 1 0.22 0.13 0.07 0.13 0.32 0.32 Expt. 2 0.35 0.21 0.13 0.20 0.40 0.39
















16h


Intact


- -+


24h 48h 16h+32h
I+ +


*0


V.o


Figure 5-3. RNA gel blot analysis of Susl mRNA expression in maize roots
incubated in 0% (-) or 2.0% (+) glucose for various time periods. At 16 h switching treatments were conducted by changing roots in 0% glucose to 2.0% glucose (-/+) and vise verse (+/-); incubations were continued for 32
additional hours.








68
switched to 2.0% glucose responded similarly to those remaining in 2.0% glucose for the entire time.

At the protein level,changes in response to altered carbohydrate availability were not apparent at 16 h, remained barely detectable at 24 h, but were clearly evident after 48 h (Figure 5-4). Treatment reversals indicated that a protein-level response occurred only when 16 h of elevated glucose treatment was followed by 32 h of glucose deprivation. The response was similar to that of root tips that had remained continuously in 2.0% glucose. Slight differences in enzyme activity between treatments were evident after 16 h or 24 h (Table 5-2); however activity in glucose supplemented tips had risen to levels two-fold greater than those without exogenous sugars within 48 h.


Discussion

The significance of results described here are two-fold. First, data demonstrate that the differential response to changing carbohydrate availability by the Susl gene for sucrose synthase is apparent at the translational level as well as at the transcriptional level. The two genes encoding sucrose synthase previously have been shown to respond differentially to carbohydrate supply (Koch and McCarty, 1988; 1990; Koch et al., 1989). Differences at the protein and enzyme level, however, have been difficult to detect due to cross reactivity of polyclonal antibodies and the collective contribution of both isozymes to activity measurements. Second, the resulting changes in physiology may allow the cells to adjust their carbohydrate metabolizing capacity to the available supply. Use of

























16h 24h 48h 16h + 32h Intact - + - + - + -/++I-


-M AM


Figure 5-4. Protein gel blot of Sus I encoded sucrose synthase from maize roots
incubated in 0% (-) or 2.0% (+) glucose for various time periods. At 16 h switching treatments were conducted by changing roots in 0% glucose to 2.0% glucose (-/+) and vise verse (+/-); incubations were continued for 32
additional hours.


4OW 4w 4w









Table 5-2. Sucrose synthase activity in mutant maize (W22:shl) root tips
incubated in media containing in 0 or 2.0% glucose for various time periods.


Sucrose synthase activity

16h 24h 48h 16h + 32h Intact - + - + - + -+ +(jimol sucrose g-1 protein h1)

0.12 0.22 0.28 0.22 0.29 0.18 0.40 0.31 0.40 Note: 0 and 2.0% glucose represented by - and +, respectively.








71
the Shl maize mutant has allowed the response to be characterized using a simple, single enzyme system with respect to changing sugar supply. The increased protein and enzyme activity evident at increased exogenous glucose levels indicate that the plant tissue can adjust this first step in their sucrosemetabolizing capacity relative to its carbohydrate status.

After 24 h at a given glucose level, changes in gene expression were more marked than were differences at the protein and enzyme levels (Figure 5-1, Figure 5-2 and Table 5-1). This is not surprising given the probable presence of previously formed RNA and protein (both appear to be relatively long-lived) as well as the comparatively long-term progression of the response to the maximal extent observed at 48 h (Figure 5-3 and Table 5-2). Chourey et al. (1991 b) reported that the sucrose synthase gene in sorghum homologous to Susl gene from maize is anaerobically induced, but levels of the respective protein do not change. Anerobic induction, however, was terminated after only 12 h. Anaerobic induction of Shl in maize becomes apparent between 6 and 12 hours but message levels are not maximal until at least 24 h (Duke and Koch, unpublished data). Data from the present work indicate that like the respiratory drop noted by Brouquisse et al. (1991), at least 20 hours are required before a change in Sus1 is fully apparent at the gene level and even longer at the protein level. Nonetheless, data are presented here at the levels of mRNA, protein and enzyme activity that indicate that expression of the Susl gene for sucrose synthase is responsive to carbohydrate availability to an extent not evident in background levels of total RNA








72

and protein. The duration of time required for this response is consistent with the proposed physiological function of sucrose in coarse adjustment of root growth relative to sugar supply (Farrar and Williams, 1990).

The increased levels of Susl mRNA and subsequent elevation of its respective protein with carbohydrate status may give insight into specific roles for this isozyme as opposed to the Shl gene product. Sucrose synthase activity could be key to the regulation of carbon entry into the respiratory pathway (Huber and Akazawa, 1986; Black et al., 1987). The enhanced expression of Susl under plentiful carbohydrate supplies accompanied probable increases in respiratory activity in the root tips (Saglio and Pradet, 1980; Farrar and Williams, 1990; Brouquisse et al., 1991). In addition, carbohydrate content in many tissues has been correlated with the respiration rate (Penning de Vries et al., 1979; Farrar, 1985). Saglio and Pradet (1980) also found that an exogenous supply of 0.2 M glucose was required to bring the respiration rate of excised maize roots back to the level of intact tissue, indicating that the rate of metabolic activity of the root tips may be closely tied to sugar import. Perhaps another line of evidence supporting control of respiration by carbohydrate status comes from the work of Douce et al. (1990) in which sycamore cells in culture showed loss of mitochondrial function when starved of sucrose; the beginning of the decline coincided with the fall in endogenous sugar concentrations.

Another possible role for the Sus 1 gene product may be in the diversion of carbohydrate to cell wall biosynthesis. Roots in 2.0% glucose medium show








73

marked growth during the 24 hours of incubation (data not shown). During this time, there is a demand for cell wall synthesis by the expanding cells. Sucrose synthase has been implicated in the directing of carbohydrates for polysaccharide biosynthesis (Amino et al.,1985; Hendrix, 1990). The level of involvement for nucleotide-sugars during cell wall polysaccharide biosynthesis has been implicated in a need for greater activity of this enzyme (Maas et al., 1990). Sucrose synthase activity was elevated in cell cultures of Catharanthus roseus during the G1 phase when total amounts of cell wall biosynthesis increased significantly (Amino et al., 1985). Sugar modulation of Sus 1 could convincingly combine production of cell wall precursors with other aspects of increased growth (Farrar and Williams, 1990) likely to accompany an enhanced sugar supply.

The carbohydrate response of sucrose synthase in the present study differs from previously demonstrated regulation in that it occurs in rapidly growing and metabolizing structures. Other studies have involved sucrose synthase regulation in storage tissues where processes such as starch accumulation predominate. Loss of the shrunken-1 gene in maize results in a typical endosperm phenotype where starch deposition is reduced by over 70% (Chourey and Nelson, 1976); however, a mutant lacking a functional Susl gene has no apparent phenotype (Chourey et al., 1988). Starch deposition accompanies protein accumulation in developing potato tubers and levels of mRNA encoding sucrose synthase have been shown to increase in this tissue (Salanoubat and Belliard, 1989). Levels of storage proteins, such as patatin, also accumulate during this time (Paiva et al.,








74

1983). Increased levels of sucrose result in elevated levels of genes for both sucrose synthase (Salanoubat and Belliard, 1989) and patatin (Rocha-Sosa et al., 1989; Wenzler et al., 1989) in tissues where they are not usually found. A rise in sugar availability can also result in increased transcription and translation of a unique storage protein in stem and leaf tissues of sweet potato (Hattori et al., 1990). The diversity of processes operating in the system utilized in this study suggests that sugar-responsive gene expression (ie. Susl) may have broad implications in the formation and function of non-storage plant tissues.














CHAPTER 6
AN ORGAN-SPECIFIC INVERTASE DEFICIENCY IN THE PRIMARY ROOT OF AN INBRED MAIZE LINE



Abstract


An organ-specific invertase deficiency affecting only the primary root system is described in the Oh 43 maize inbred. Invertases (acid and neutral/soluble and insoluble) were assayed in various tissues of hybrid (NK 508) and inbred (Oh 43, W22) maize lines to determine the basis for an early report that Oh 43 root tips were unable to grow on sucrose agar (Robbins, 1958). Substantial acid invertase activity (7.3 to 16.1 imol glucose mg~'protein h) was evident in extracts of all tissues tested except the primary root system of Oh 43. This deficiency was also evident in lateral roots arising from the primary root. In contrast, morphologically identical lateral roots from the adventitous root system had normal invertase levels. These results suggest that ontogenetic origin of root tissues is an important determinant of invertase expression in maize. Adventitious roots (including the seminals) arise above the scutellar node and are, therefore, of shoot origin. The Oh 43 deficiency also demonstrated that invertase activity was not essential for maize root growth. Sucrose synthase was active in extracts from all root apices and theoretically provided the only available avenue for sucrose degradation in









76
primary root tips of Oh 43. The deficiency described here will provide a useful avenue of investigation into the expression and significance of root invertase.


Introduction


Sucrose breakdown is critical to the vast majority of plant species because non-photosynthetic tissues depend on import of this sugar for their growth and development. Initial cleavage of sucrose can be catalyzed by either invertase or the reversible enzyme sucrose synthase. Invertases are especially active in tissues undergoing rapid cell division such as shoot and root apices (Avigad, 1982). Previously, the role of invertases in sucrose transfer was considered particularly important in plants such as sugar cane and maize where substantial sugar movement occurred through the cell wall space and was accompanied by action of an extracellular invertase (Glasziou and Gayler, 1972; Hawker and Hatch, 1965). Recent evidence, however, indicates that although much hydrolysis is often observed, invertase activity may not be essential for sucrose uptake into either sugar cane stems (Ungle, 1989; Thom and Maretzki, 1990) or maize kernels (Schmalstig and Hitz, 1987).

Sucrose generally is believed to enter root tips without traversing the extracellular space (Giaquinta et al., 1983); however growing roots can differ markedly in their capacity to lose (Rovira and Davey, 1974) and retrieve (Robbins, 1958) exogenous sugars. Net losses do occur. The extent of sugar efflux from roots can be affected by irradiance level, nutritional status, moisture availability and








77
temperature (Rovira and Davey, 1974). The composition of root sugars exuded is quite variable but includes both reducing and non-reducing sugars (Rovira and Davey, 1974). Glucose and fructose are often taken up from the extracellular space more rapidly than is sucrose (Humphreys, 1974). Retrieval of solutes from the apoplast and the form in which they are available may thus be a potentially important attribute of root carbon balance.

A deficiency in this retrieval process was first indicted by Robbins' report (1958) that roots of a maize inbred, Oh 43, were unable to grow on sucrose agar medium, yet roots of another line, Hy 2, grew quite well. Only when roots of both were cultured immediately adjacent to one another, were those of Oh 43 able to grow. Growth of excised Oh 43 root tips also occurred when glucose was substituted for the sucrose. Oh 43 was concluded to be "incapable of inverting sucrose" in its root tips. Preliminary investigations by B. Burr (Brookhaven National Laboratory, personal communication) indicated that a lack of invertase may have been the reason for the inability of Oh 43 roots to metabolize sucrose.

The absence of invertase activity could have important implications for sucrose import not only because of potential effects on the retrieval system, but also because sucrose utilization in such an instance could theoretically be initiated only via action of sucrose synthase. In addition, genetic material which lacks activity of a specific enzyme can be useful in investigations of physiological processes normally mediated by these enzymes (Koch et al., 1982). The present report demonstrates that invertase is not essential for primary root growth despite








78

probable advantages of its presence and indicates an unusual organ-specific difference in expression between primary and adventitious roots.


Materials and Methods


Plant Material

Maize seed (Zea mays L NK 508, W22 and Oh 43) were germinated on moist filter paper in petri dishes. Seeds were imbibed for 24 hours and pericarps removed, allowing more uniform germination and more effective surface sterilization (20 min soak in 0.525% sodium hypochlorite).

Five successive 2 mm segments were sampled from the tips of primary roots 4 to 5 days after germination. Intact roots of Oh 43 seedlings grew more slowly than did those of NK 508 or W22, but all roots had reached 2 cm prior to excision. Tissue samples were weighed, frozen in liquid N2 and stored at -80 C until assayed for invertase activity. In subsequent experiments, 5 mm root tips were excised from primary and adventitious roots for invertase and sucrose synthase activity measurements. Plants and tissues were as above. Tissue Extraction

Frozen tissue samples were ground to a fine powder in liquid N2 using a mortar and pestle. Frozen powder was transferred to a second mortar containing ice-cold 200 mM HEPES buffer (pH 7.5) with 1 mM DTT, 5 mM MgCI, 1 mM EGTA, 20 mM sodium ascorbate and 10% (w/w) PVPP. One ml of extraction buffer was used for every 100 mg of tissue fresh weight. Buffered extract was centrifuged at








79
14,000 x g for 1 min to sediment particulate matter. Supernatant was dialyzed (27,000 mw cutoff) at 4 C for 24 h against extraction buffer diluted 1:40. Buffer was changed after 1 h and thereafter, every 4 h. Soluble dialyzed extract was assayed for invertase as described below. Previously separated particulate matter was rinsed with one volume of extraction buffer and assayed for insoluble, cellwall-bound invertase (soluble acid invertase includes both vacuolar and loosely bound extracellular enzyme [Avigad, 1982]).

To test the possibility that the soluble enzyme was present in primary roots of Oh 43 but was being bound or inactivated during the extraction procedure, two additional extraction/assay methods were employed. First, adventitious root extracts, previously shown to contain active invertase activity, were added to those of primary apices. The resulting mixture was dialyzed and assayed for enzyme activity. Second, three cm apices of both primary and adventitious roots Oh 43 roots were excised. Apices of these roots (0.5 cm) were suspended in extraction buffer for three hours at 27 C. Buffer alone was subsequently dialyzed as described above. The portion of each root which had been immersed in the extraction buffer was excised for fresh weight measurement. After dialysis, the buffer-enzyme solution was analyzed for enzyme activity.


Enzyme Assays

Soluble and insoluble forms of acid invertase were assayed as described by Lowell et al. (1989). Reaction media contained 50 mM sucrose, and pH of 4.5 was adjusted with a sodium acetate buffer. Neutral invertase was assayed using








80
the same reaction medium adjusted to pH 7.5 with potassium phosphate buffer. Initial assays were also performed at pH ranges of 4.0 to 5.5 for acid invertase and 7.0 to 8.0 for neutral invertase. After a 15 min incubation at 30 C, glucose production was quantified by the glucose oxidase method (Sigma Chemical Co.). Sucrose synthase was assayed in the degradative direction using a radiometric assay quantifying the production of 14C-UDPG (Chapter 3). Histochemical Staining

Free-hand cross sections from apices of both primary and adventitious roots were fixed in 4% formalin (pH 7.0) for 30 min and rinsed in water at least 10 times over a period of 3 hours to remove endogenous sugars (Doehlert and Felker, 1987). Sections were then incubated in a sodium phosphate buffer (0.38 M, pH 6.0) containing 0.24 mg ml1 nitroblue tetrazolium, 0.14 mg ml1 phenazine methosulfate, 25 units ml1 glucose oxidase and 5 mg ml"1 sucrose (Doehlert and Felker, 1987). Control sections were incubated in the same mixture without sucrose. After rinsing in water, sections were post fixed in 4% formalin (pH 7.0) and photographed under a microscope.


Results

Primary roots of Oh 43 showed little or no acid invertase activity (Table 6-1). In contrast, acid invertase was active in extracts from apical areas of roots from other maize lines examined (NK 508 and W22). Activity, per unit fresh weight, was greatest in root apices, decreasing with distance from the tip until no longer













Table 6-1. Soluble and insoluble acid invertase activity in sequential 2 mm segments of primary
roots of 5 to 6 day-old seedlings from 1 hybrid and 2 inbred lines of Zea mays.


Solubley Insoluble Root
Segment Oh 43 NK 508 W22 Oh 43 NK 508 W22

jimol glucose g'"FW h'

0-2 mm 0.2 *02w 43.8 � 5.2 40.2 � 3.7 ---z 3.8 0.5 3.6 � 0.8
2-4 mm --- 32.8 � 2.2 28.6 � 3.1 --- 3.1 + 1.9 2.7 � 1.5
4-6 mm --- 20.6 � 5.0 15.4 � 4.6 .........
6-8 mm --- 1.4 � 0.9 0.9 � 0.5 ---....
8-10 mm --- --- ---.......
Z-not detectable
Y--Soluble activity is expressed per unit FW to allow comparison with insoluble activity. W--Activity in root tips from axenic culture was 0.0 � 0.0 Amol g1'FW h"; thus microorganisms are a likely source of the residual activity in ca. 1 of 3 assays. Each value represents the mean of 3 separate samples � SEM.








82

detectable farther than 8 mm from the apex. Soluble enzyme accounted for 88 to 92% of the total acid invertase activity, and the remainder was due to action of the insoluble enzyme. Neutral invertase activity was insignificant or absent from all lines (data not shown). Although invertase action was essentially undetectable in the primary roots of Oh 43, adventitious root extracts showed levels of activity as least as great as those from the primary roots of other lines tested (Table 6-2).

Active acid invertase was released into buffer in the extracellular space around adventitious root tips of Oh 43 during 3 hours of emersion (13.4 � 3.5 pmol glucose g' FW h'1--ca. 25% of the activity on a fresh weight basis of N2 and buffer extracted roots). The same was not observed during emersion of primary roots from this line. Also, from 83% to 94% (13.3 to 15.1 Amol glucose mg1 protein h-1) of the invertase activity in adventitious root extracts remained after addition of extracts from primary roots of Oh 43.

Unlike invertase, sucrose synthase was active in extracts of both primary and adventitious roots from Oh 43 seedlings. Activity of this enzyme was approximately similar in both root types, as observed for the other two lines tested (Table 6-2).

Extracts of lateral roots originating from primaries exhibited no detectable acid invertase activity (Table 6-3), in contrast to counterparts derived from adventitious roots. Shoots of 5-day-old Oh 43 seedlings and endosperm or scutellum tissue from developing kernels (23 DAP) also exhibited activity of both soluble and insoluble acid invertase (Table 6-3).













Table 6-2. Soluble acid invertase and sucrose synthase activity in 0.5 cm apices of primary and
adventitious roots of 5-to 6-day-old seedlings from 1 hybrid and 2 inbred lines of Zea mays.

Oh 43 NK 508 W22 primary adventitious primary adventitious primary adventitious Amol glucose mg-'protein h1

Invertase 0.1 � o.1w 16.1 +3.2 16.0 *2.5 14.0 *2.5 12.6 +2.2 11.9 �1.6
Sucrose
Synthase 1.2 � o.1 1.1 �0.1 1.0 �0.1 1.1 �0.2 1.0 �0.1 0.9 �0.1

/hmol glucose g-1 FW h-1

Invertase 0.2 � 0.2w 60.8 � 12.1 56.5 � 8.7 51.6 � 9.2 53.0 + 9.3 48.9 � 6.7
Sucrose
Synthase 4.7 � 0.4 4.2 � 0.1 3.5 � 0.5 4.2 � 0.6 4.0 0 0.4 3.7 � 0.4
w--Activity in root tips from axenic culture was 0.0 � 0.0 /mol g1 FW h'; thus microorganisms are a likely source of the residual activity in ca. 1 of 3 assays. Each value represents the mean of 3 separate samples � SEM.











Table 6-3. Soluble acid invertase activity in various tissues of
Oh 43, an inbred line of Zea mays.


Mmol glucose mg'protein h-'


Shoot
Endosperm
Scutellum Primary roots Lateral roots from 10 roots Adventitious roots Lateral roots from adventitious roots


10.8 � 0.6 7.3 � 1.2 9.0 � 0.8 0.1 � 0.1 0.1 � 0.1W
16.1 �3.2

10.4 � 0.9


gumol glucose g' FW h

25.8 * 1.5 12.6 � 2.1 19.4 * 1.7
0.2 * 0.2

0.2 � 0.1w 60.8 � 12.1

38.7 + 3.2


W--Activity in root tips from axenic culture was 0.0 � 0.0 gmol g1FW h; thus microorganisms are a likely source of the residual activity in ca. 1 of 3 assays. Shoot, primary roots and adventitious roots were samples from 6-day-old seedlings. Endosperm and scutellum were from developing kernels 23 DAP. Lateral roots developed after 2 to 3 weeks of growth and were then excised. Each value represents the mean of 3 separate samples � S.E.M.








85
Histochemical staining indicated that no invertase activity was detectable in the primary roots of Oh 43 (Figure 6-1). Cross sections of primary and adventitious roots of Nk 508 and adventitious roots of Oh 43 all stained positive for invertase activity. Invertase activity was primarily in the cortex and was localized intercellularly.


Discussion

These data confirm that Oh 43 (an inbred line of maize) lacks invertase activity in its primary root tips. Surprisingly, however, a deficiency was not evident in structurally and functionally similar adventitious roots (Table 6-2), other tissues of the same Oh 43 plants (Table 6-3) or in the developing kernels of this line (Doehlert, et al., 1988). The genetic potential for invertase expression is therefore present. The lack of activity may result from altered regulation of gene expression (transcription or translation), the loss of a tissue specific invertase isozyme or the existence of an unidentified effector of enzyme function. The significance of results described here is twofold. First, evidence is presented for differential expression of invertase in morphologically identical organs that differ primarily in point of origin. This is most strikingly illustrated in the apparent distinction between lateral roots arising from primary and adventitious root systems. Differential expression of genes in morphologically identical structures are unusual; however, organ or tissue-specific differences have been well documented (Fluhr et al., 1986; Xie and Wu, 1989). Xie and Wu (1989), for example, found that genes for alcohol dehydrogenase were differentially expressed in root and shoot tissues of rice
























Figure 6-1. Histochemical localization of invertase activity in free-hand, fresh cross
sections of root apices of 6-day-old maize seedlings. Sections are approximately 50 Am in thickness. Primary (A) and adventitious (B) root cross sections from NK 508 incubated in reaction medium without sucrose (controls). Primary (C) and adventitious (D) root cross sections from Oh 43 incubated in reaction medium (note only adventitious root section exhibits blue formazan reaction product [dark areas]). Primary (E) and adventitious (F) root cross sections from NK 508 incubated in reaction medium (note both sections exhibit reaction product [dark areas]). Bar represents 50 jim.








87




B




~
2'.




D










F






V








88
plants. One isozyme predominated in shoot-derived organs (leaves, sheaths, nodes and pollen) and the other isozyme showed highest activity in the roots. It is interesting in this respect that the adventitious roots of maize arise above the scutellar node of the developing seedling and are, therefore, derived from shoot tissue (Hayward, 1938). The difference in invertase expression in primary and adventitious root systems may reflect a similar root/shoot dichotomy. Therefore, the tissue of origin and cell lineage may be more important than organ identity and function in regulating invertase expression. Other variants in invertase expression have been described. Echeverria and Humphreys (1984) reported a hybrid maize line (DKXL80) which exhibited no soluble invertase activity in contrast to previously tested lines. Other tissues of this maize line exhibited normal invertase activity.

Second, data demonstrate that invertase is not essential for primary root growth. The primary roots of Oh 43 exhibited no signs of premature senescence, and, if left intact, continued apparently normal growth for many days (data not shown). Invertase's role in sucrose import into roots has been questioned by Chapleo and Hall (1 989a) who concluded that although present, apoplastic root invertase did not have a direct role in sugar transport in Ricinus. However, substantial activity of invertase has been widely documented in roots of plants such as pea (Lyne and ap Rees, 1971), bean (Robinson and Brown, 1952), tomato (Chin and Weston, 1973), Ricinus (Chapleo and Hall, 1989a, b and c), oat (Pressy and Avants, 1980), and maize (Chang and Bandurski, 1963; Hellebust and Forward, 1962). In maize, the Oh 43 invertase deficiency apparently prevents








89

utilization of exogenous sucrose (Robbins, 1958). Specific tissue localization of invertase also has been described. Peak activity for root invertase is generally 2-3 mm behind the apex and corresponds to the region of expansion and elongation in pea (Robinson and Brown, 1952) and maize (Hellebust and Forward, 1962). In Ricinus this activity predominates in the cortex (Chapleo and Hall, 1989).

Although invertase may not have a direct role in sucrose import in roots, it still may be important to two major aspects of root biology. First, invertase has been implicated in formation of mycorrhizal associations (Purves and Hadley, 1975). Maize (Gerdemann, 1964; Kothari et al., 1990) and 90% of other agriculturally important species form these beneficial symbioses under field conditions (Gerdemann, 1986). Invertase activity typically increases and hexose levels rise at infection sites of biotrophic fungi (Long et al., 1975). Elevated hexose content in roots upon infection by mycorrhizal fungi has been attributed to a rise in invertase levels (Purves and Hadley, 1975). It is not known whether this is host or fungal invertase; however, Oh 43 does not appear to provide the former in its primary root systems.

Second, the purported lack of a sucrose carrier in the plasma membrane of maize root cells (Un et al., 1984) would indicate that if sucrose were released into the apoplast, retrieval might proceed more effectively in the presence of extracellular invertase. Such retrieval could be particularly important during stress or periods when sugar losses from the symplast were elevated. Any physiological consequence of this deficiency would most likely be evident early in seedling








90
development because the root system would consist solely of a primary root at this time. Later in seedling development, adventitious or seminal roots rapidly take over a dominant role, leaving the primary root with little or no essential function (Hayward, 1938). In the field, lines having Oh 43 as a progenitor have been observed to suffer from poor emergence rates in damp soils (B. Martin, Pioneer Seed, personal communication). However, conclusive evidence of a physiological effect will require generation of and analysis of isogeneic lines.

In conclusion, the deficiency described here will be useful for investigation into the regulation and physiological function of root invertase. Because the primary root system in maize is nonessential, invertase deficient roots can be studied without deleterious effects on the overall physiology of the plant. In addition, this deficiency is significant because it reveals an unexpected distinction between primary and adventitious root development in maize. At some level, the mechanisms which regulate invertase in these root systems must differ.














CHAPTER 7
SUMMARY AND CONCLUSIONS


Sucrose metabolism is important to the majority of plant species because of the widespread role of this sugar in growth and development through its function as a phloem transport sugar. Initial breakdown of sucrose can be catalyzed by either sucrose synthase, a reversible enzyme, or invertase.

Expression of genes encoding the sucrose synthase isozymes (Shl and Sus 1) have been found to be sensitive to carbohydrate levels (Koch and McCarty, 1988, Koch et al., 1989). Northern blot analysis of mRNA showed levels showed those of Sh 1 were up-regulated in response to carbohydrate depletion whereas those of Susl were down-regulated under the same conditions. This may prove to be an important control mechanism whereby plant cells are able to react to cellular carbohydrate status.

The first purpose of this work was to determine the extent to which carbohydrate-modulated changes in message levels affected activity of sucrose synthase the level of enzyme activity. However, previously available sucrose synthase assay methods proved ineffective for maize root tips due to a precipitous loss of activity after tissue extraction. A rapid, radiometric assay for sucrose synthase was therefore developed that overcomes these obstacles. Extraction and assay were optimized for factors such as substrate concentration, pH, assay




Full Text
14
attributed by Grimes et al. (1970) to possibly result from conformational changes
in the enzyme.
Reduced K^s for UDPG when compared to other NDPGs have also been
observed for sucrose synthase from pea seedlings (Gabrielyan et al., 1969), sweet
potato root (Murata, 1971), potato tubers (Pollock and ap Rees, 1975), sweet corn
seeds (de Fekete and Cardini, 1964) and soybean nodules (Morell and Copeland,
1985). In contrast, ADPG and TDPG were reportedly more efficient glucosyl
donors than UDPG for sucrose synthase isolated from sorghum seeds (Sharma
and Bhatia, 1980) and sugar beet roots (Avigad and Milner, 1966).
No large differences in Kms for UDP and other nucleoside diphosphates
generally are observed when the reverse reaction is analyzed. Delmer (1972a and
b) found minimal or no differences in Kms for NDPs with sucrose synthase from
mung bean seedlings. She did, however, find a large difference in the rates of
sucrose cleavage with different NDPs. Maxima were observed when UDP was the
substrate. The Vmax for UDP in relative terms was 100 compared to 28, 6, 3 and
3 for ADP, TDP, CDP and GDP, respectively. Similar K^s for various NDPs have
also been observed in sweet potato roots (Murata, 1971), potato tubers (Pollock
and ap Rees, 1975), Jerusalem artichoke (Pontis et al., 1972) and sugar beet
(Avigad and Milner, 1966). Sucrose synthase from sweet corn kernels does,
however, exhibit a Km an order of magnitude greater for ADP than UDP (Su and
Preiss, 1978; de Fekete and Cardini, 1964). Morell and Copeland (1985) also


22
maximum values for alkaline invertase activity observed to date are consistently
less than those of acid invertase (Masuda et al., 1988). The possibility exists that
alkaline invertase allows cells that store sucrose in their vacuoles to retain a
capacity for breakdown of enough sucrose in the cytoplasm to meet respiratory
and metabolic demands for hexoses (ap Rees, 1974). The capacity of a plant to
produce two different invertases that are spatially separated may allow the plant
cell to regulate sucrose storage independent from sucrose breakdown.
Regulation of Invertase
Acid invertases are generally found in the apoplast and vacuoles of plant
tissues. Washed preparations of cell walls contain a large proportion of a plants
acid invertase (Little and Edelman, 1973). A portion of the acid invertase can be
extracted from the cell wall during grinding, but at least some of the enzyme is
considered to be attached to the cell wall in vivo (Edelman and Hall, 1965). The
major determinant of how much acid invertase remains bound during extraction
is the pH of the buffer used (ap Rees, 1974). Buffers with acidic pH leave most
of the activity in the cell wall fraction, whereas neutral or alkaline buffers release the
majority into the soluble fraction.
Early evidence suggested that the soluble and insoluble acid invertases
were not simply different forms of the same enzyme. The pH optima and the
of bound acid invertase of mature (Hawker and Hatch, 1965) and immature (Hatch
et al., 1963) sugar cane storage tissue differed from those of the soluble fractions.
Association with the cell wall may change an enzymes properties (ap Rees, 1974);


95
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41
tetramers composed only of Sh1 encoded protomers (Chourey et al., 1986;
Heinlein and Starlinger, 1989; Rowland and Chourey, 1990), and sucrose synthase
activity is stable during extraction and dialysis procedures (Chourey and Nelson,
1976; Echt and Chourey, 1985). In extracts from root tissue, the five bands shown
by western blot represent the possible combinations of monomers of the two
separate isozymes (Sh1 and Sus) to form the native tetrameric structure (Echt and
Chourey, 1985). Heterotetramers could theoretically be more unstable than
homotetramers since, although very similar, the two subunits are not identical (Echt
and Chourey, 1985). Su and Preiss (1978) found that sucrose synthase tended
to polymerize to an inactive polymeric after extraction. Results indicated that
formation of the native enzyme from two different isozymes was not a contributing
factor in loss of enzyme activity over time (Figure 3-5A & B). Despite differences
in the absolute values of sucrose synthase activity from sh1 vs. Sh1, the percent
decline in activity was similar for both genotypes.
Discussion
Sucrose synthase activity was stabilized in vitro and an assay developed
which enabled accurate measurement of enzyme action in root tips. The
described assay allows rapid product recovery in instances where activity is
otherwise unstable in vitro, and increases sensitivity to the extent that sample
volumes as small as 100 to 200 nQ can be used.
Sucrose synthase has previously been assayed in both synthetic and
cleavage directions (Avigad and Milner, 1966; Grimes et al., 1970; Pontis et al.,


ACKNOWLEDGEMENTS
I would like to extend thanks to the members of my committee, Dr. Karen
Koch, Dr. Rebecca Darnell, Dr. Don McCarty, Dr. Curt Hannah and Dr. Alice
Harmon, for their assistance and guidance during the completion of this degree.
I would also like to thank Dr. Tom Humphreys for his critical review of this
dissertation. There are many people in the Fruit Crops Department to whom I owe
thanks. I would like to thank the staff and faculty for their help and encouragement
during my time here. For their friendship and support, I especially would like to
thank Kathy Zimmerman, Teki and Andy Ericson, Dr. Kathy Taylor, Dr. Pat
Tomlinson, Wayne Avigne, Kurt Nolte, and Don Merhaut. Thanks are also given
to the graduate students, both past and present, of the Fruit Crops Department.
Finally, I want to extend my most heartfelt thanks to my parents, Ralph and Mildred
Duke; they have stood by me and given me support throughout my education,
and I can never thank them enough.
n


107
Paiva, E.p R.M. Lister and W.D. Park. 1983. Induction and accumulation of major
tuber proteins of potato in stems and petioles. Plant Physiol. 71:161-168.
Penning de Vries, F.W.T., J.M. Witlage and D. Kremer. 1979. Rates of respiration
and of increase in structural dry matter in young wheat, ryegrass and maize
plants in relation to temperature, to water stress and to their sugar content.
Ann. Bot. 44:595-609.
Pollock, C.J. and T. ap Rees. 1975. Activities of enzymes of sugar metabolism in
cold-stored tubers of Solarium tuberosum. Phytochem. 14:613-617.
Pontis, H.G. 1977. Riddle of Sucrose, pp. 79-117 in D.H. Northcote, ed. Plant
Biochemistry: Inti. Rev. Biochem. Vol. XIII. Baltimore: University Park Press.
Pontis, H.G., R.A. Wolosiuk, L.M. Fernandez and B. Bettinelli. 1972. The role of
sucrose and sucrose synthase in Helianthus tuberosus. pp. 239-266 in R.
Piras and H.G. Pontis, eds. Biochemistry of the Glycosidic Linkage. New
York: Academic Press.
Prentice, N. and G.S. Robbins. 1976. Composition of invertases from germinated
barley. Cereal Chem. 53:874-880.
Pressy, R. 1967. Invertase inhibitor from potatoes: Purification, characterization and
reactivity with plant invertases. Plant Physiol. 42:1780-1786.
Pressy, R. 1968. Invertase inhibitors from red beet, sugar beet and sweet potato
roots. Plant Physiol. 43:1430-1434.
Pressey, R., J.K. Avants. 1980. Invertases in oat seedlings, separation, properties
and changes in activities in seedling segments. Plant Physiol. 65:136-140.
Pridham, J.B., M.W. Walter and H.G. Worth. 1969. The metabolism of raffinose and
sucrose in germinating broad-bean (Vicia faba) beans. J. Expt. Bot. 20:317-
324.
Purves, S. and G. Hadley. 1975. Movement of carbon compounds between the
partners in orchid mycorrhiza. pp. 119-148 in F.E. Sanders, B. Mosse and
P.B. Tinker, eds. Endomycorrhizas. London: Academic Press.
Raghuveer, P. and R.C. Sicher. 1987. Stabilization of in vitro sucrose phosphate
synthase activity. Ind. J. Plant Physiol. 30:390-392.


26
Morell and Copeland (1984) also found that the metabolites ATP, ADP, UDP, ADP-
glucose, UDP-glucose, glucose-1-phosphate, glucose-6-phosphate, and fructose-6-
phosphate (all at 5 mM) had no inhibitory effects. They also tested the effects of
various chloride salts on alkaline invertase activity and found that Na+, K+ or NH4+
at 50 mM had no effects; however, CaCI2 (10 mM) and MgCI2 (10 mM) each
inhibited activity by 25%. The anions citrate and inorganic phosphate have been
shown to stimulate alkaline invertase from Lupinus luteus nodules (Kidby, 1966);
however, Morell and Copeland (1984) found no effect on activity of soybean
nodule alkaline invertase. They did find, though, that Tris buffer was a
noncompetitive inhibitor of soybean nodule alkaline invertase activity; a 0.7 mM
buffer concentration inhibited activity by 50%.
Use of Mutants in Physiological Research
Despite the fact that all mutations have effects on the biochemistry and
physiology of the plant, only a small number have been investigated
physiologically (Vose, 1981). Advances in knowledge about the molecular bases
of cell processes in eukaryotic and prokaryotic microorganisms have been
achieved with an array of mutant lines, often induced, that modify or block steps
in the processes under study (Nilan et al., 1981). Many mutants will, theoretically,
differ in only a single major physiological character. The use of mutants is growing
in comparative physiological studies because the alternative is comparison of
contrasting genotypes that quite possibly may be altered in undefined characters
different from the one of interest.


7
similar amino acid compositions and share limited structural homologies (Echt and
Chourey, 1985). They do differ slightly in their electrophoretic movement during
PAGE (Echt and Chourey, 1985). The two subunits are homologous enough to
form heterotetrameric structures, apparent as five separate bands on native PAGE
(Echt and Chourey, 1985).
Spatial separations of these two sucrose synthase isozymes are sometimes
apparent. Sh1 encoded protein is primarily located in the endosperm (Chourey
and Nelson, 1976; Chen and Chourey, 1989), in the root (Springer et al., 1986;
Chourey et al., 1986) and in etiolated shoots (Springer et al., 1986). The Sus1
encoded protein is found throughout the plant (Chourey, 1981; Echt and Chourey,
1985; Chourey et al., 1988). The distribution of both proteins is further
distinguished under stress conditions (such as anaerobiosis) where tissue-specific
localization in roots is readily apparent (Rowland et al., 1989).
Roles of sucrose synthase
Sucrose synthase was initially considered to be a sucrose synthesizing
system in plants by Leloir and Cardini (1953, 1955). Sucrose synthase and
sucrose phosphate synthase (EC 2.3.1.14, UDP-D-glucose: D-fructose-6-
phosphate 2-a-D-glucosyltransferase) are the two enzymes that catalyze the
transglucosylation reaction from UDP-glucose to fructose and fructose-e-
phosphate, respectively.


72
and protein. The duration of time required for this response is consistent with the
proposed physiological function of sucrose in coarse adjustment of root growth
relative to sugar supply (Farrar and Williams, 1990).
The increased levels of Sus1 mRNA and subsequent elevation of its
respective protein with carbohydrate status may give insight into specific roles for
this isozyme as opposed to the Sh1 gene product. Sucrose synthase activity
could be key to the regulation of carbon entry into the respiratory pathway (Huber
and Akazawa, 1986; Black et al., 1987). The enhanced expression of Sus1 under
plentiful carbohydrate supplies accompanied probable increases in respiratory
activity in the root tips (Saglio and Pradet, 1980; Farrar and Williams, 1990;
Brouquisse et al., 1991). In addition, carbohydrate content in many tissues has
been correlated with the respiration rate (Penning de Vries et al., 1979; Farrar,
1985). Saglio and Pradet (1980) also found that an exogenous supply of 0.2 M
glucose was required to bring the respiration rate of excised maize roots back to
the level of intact tissue, indicating that the rate of metabolic activity of the root tips
may be closely tied to sugar import. Perhaps another line of evidence supporting
control of respiration by carbohydrate status comes from the work of Douce et al.
(1990) in which sycamore cells in culture showed loss of mitochondrial function
when starved of sucrose; the beginning of the decline coincided with the fall in
endogenous sugar concentrations.
Another possible role for the Sus 1 gene product may be in the diversion of
carbohydrate to cell wall biosynthesis. Roots in 2.0% glucose medium show


27
The maize plant (Zea mays L.) has been particularly useful in genetic and
cytogenetic studies because of the number of mutants available (Neuffer et al.,
1968). Many of the mutants also have proven useful for physiological research.
The shrunken-1 mutant of maize was first described by Chourey and Nelson
(1976). Less than 10% of the normal sucrose synthase activity in wild-type
endosperm was observed. This reduced activity results in a "shrunken" phenotype
in the dry kernel. The sh 1 mutant has proven useful in elucidation of the role of
sucrose synthase in starch formation. The residual activity of sucrose synthase
present in this sh1 mutant was attributed to the presence of another isozyme
encoded by a second gene (Chourey and Nelson, 1976). This second gene,
Sus 1, has been mapped to the same chromosome as Sh 1 (chromosome 9) but
is located 32 map units away (McCarty et al., 1986; Gupta et al., 1988). Sh1 and
Sus 1 encode similar proteins. Sucrose synthase is a tetramer in its native form
(Su and Preiss, 1978), and the two isozymes encoded by Sh1 and Sus 1 are able
to form heterotetrameric forms of the native protein (Echt and Chourey, 1985).
Sh1 has been shown to be responsive to anaerobic conditions, with
transcript levels increasing 10 to 20 times in shoot and root tissue respectively
compared to aerobic controls (Springer et al., 1986). However, Sus1 exhibits little
response to anaerobic stress and seems to be expressed at a relative constant
in all tissues (McCarty et al., 1986). Rowland et al (1989), however, found that
Sus1 did show a slight response to anaerobic conditions, decreasing slightly in the
lower root, primarily in the pith, root tip and root cap. A maize mutant lacking the


Table 6-2. Soluble acid invertase and sucrose synthase activity in 0.5 cm apices of primary and
adventitious roots of 5-to 6-day-old seedlings from 1 hybrid and 2 inbred lines of Zea mays.
Oh 43
NK 508
W22
primary
adventitious
primary
adventitious
primary
adventitious
/mol glucose mg"1 protein h"1
Invertase
0.1 o.iw
16.1 3.2
16.0 2.5
14.0 2.5
12.6 2.2
11.9 1.6
Sucrose
Synthase
1.2 0.1
1.1 0.1
1.0 0.1
1.1 0.2
1.0 0.1
0.9 0.1
/mol glucose g"1 FW h'1
Invertase
Sucrose
0.2 0.2W
60.8 12.1
56.5 8.7
51.6 9.2
53.0 9.3
48.9 6.7
Synthase
4.7 0.4
4.2 0.1
3.5 0.5
4.2 0.6
4.0 0.4
3.7 0.4
W--Activity in root tips from axenic culture was 0.0 0.0 mol g'1FW h'1; thus microorganisms are
a likely source of the residual activity in ca. 1 of 3 assays.
Each value represents the mean of 3 separate samples SEM.


CHAPTER 3
INSTABILITY OF SUCROSE SYNTHASE FROM ROOT TIPS:
CHARACTERIZATION AND STABILIZATION
Abstract
Instability of sucrose synthase from root tips was characterized in maize and
an assay developed to circumvent the rapid decline of activity in vitro (35 and
100% activity loss in 20 min for maize and cotton, respectively). Initially de
suerse cleavage was quantified by recovery of 14C-UDPG on DEAE ion exchange
paper (Delmer, 1972; Su and Preiss, 1978). Subsequently, modifications were
made which resulted in increased accuracy, reduced tissue volume required and
reduced extraction/assay period. Phenolic protectants did not reduce the activity
loss over time. Specific inhibitors for the four classes of proteinases were also
tested; only PMSF increased enzyme activity, but did not completely prevent its
loss over time. Stabilization and additional elevation of activity were achieved by
adding casein. However, western blot analysis indicated that activity decline was
not associated with any detectable proteolytic degradation, charge alteration, or
subunit separation. In addition, inclusion of 10 mM P¡ in the extraction medium
rapidly reduced activity, indicating the possible involvement of phosphorylation or
nucleotide effects.
29


97
Chourey, P.S., G.A. DeRobertis and P.E. Still. 1988. Altered tissue specificity of the
revertant shrunken allele upon Dissociation (Ds) excision is associated with
loss of expression and molecular rearrangement at the corresponding non
allelic isozyme locus in maize. Mol. Gen. Genet. 214:300-306.
Chourey, P.S., M.D. Latham and P.E. Still. 1986. Expression of two sucrose
synthetase genes in endosperm and seedling cells of maize: evidence of
tissue specific polymerization of protomers. Mol. Gen. Genet. 203:251-255.
Chourey, P.S. and O.E. Nelson. 1976. The enzymatic deficiency conditioned by the
shrunken-1 mutations in maize. Biochem. Gen. 14:1041-1055.
Chourey, P.S., E.W. Taliercio and E.J. Kane. 1991b. Tissue-specific expression and
anaerobically induced posttranscriptional modulation of sucrose synthase
genes in Sorghum bicolor M. Plant Phyisol. 96:485-490.
Church, G.M. and W. Gilbert. 1984. Genomic sequencing. Proc. Natl. Acad. Sci.
(USA) 81:1991-1995.
Claussen, W. B.R. Loveys and J.S. Hawker. 1985. Comparative investigations on
the distribution of sucrose synthase activity and invertase activity within
growing, mature and old leaves of some C3 and C4 plant species. Physiol.
Plant. 65:275-280.
Claussen, W. B.R. Loveys and J.S. Hawker. 1986. Influence of sucrose and
hormones on the activity of sucrose synthase and invertase in detached
leaves and leaf sections of eggplants (Solanum melongena). J. Plant
Physiol. 124:345-358.
Cobb, B.G. and L.C. Hannah. 1988. Shrunken-1 encoded sucrose synthase is not
required for sucrose synthesis in the maize endosperm. Plant Physiol.
88:1219-1221.
Dale, E.M. and T.L. Housley. 1986. Sucrose synthase activity in developing wheat
[Triticum aestivum) endosperms differing in maximum weight. Plant Physiol.
82:7-10.
Davies, D.R. 1974. Some aspects of sucrose metabolism, pp. 61-81 in J.B.
Pridham, ed. Plant Carbohydrate Biochemistry. New York: Academic Press.
de Fekete, M.A.R. 1969. Zum Stoffwechsel der Starke I. Die Umwandlung von
Saccharose in Starke in den Kotyledon von Vicia faba. Planta 87:311-323.


59
from regulation by anaerobic conditions (Koch and McCarty, 1988). The anaerobic
induction has been reported to occur only at the transcriptional level without
differences in protein levels (McElfresh and Chourey, 1988; Taliercio and Chourey,
1991). Possible changes in protein levels and enzyme activity of sucrose synthase
due to carbohydrate regulation have been difficult to detect because of the non
specificity of assay methods (Duke and Koch, unpublished).
The two isozymes of sucrose synthase from maize (encoded by the Sh1
and Sus1 genes) are very similar, differing only slightly in their electrophoretic
movement during PAGE (Echt and Chourey, 1985). Despite close homology, they
are distinctive proteins encoded by separate genes (Chourey, 1981; Echt and
Chourey, 1985). The two proteins are, however, distinct in their localization within
the maize plant. The protein encoded by Sh 1 is primarily located in the
endosperm (Chourey and Nelson, 1976; Chen and Chourey, 1989), in the root
(Springer et al., 1986; Chourey et al., 1986) and in etiolated shoot (Springer et al.,
1986); however Sh 1 mRNA does appear in other tissues such as pollen grains
(Hannah and McCarty, 1988). The Sus 1 encoded protein is more widespread in
its localization and is found throughout the plant (Chourey, 1981; Echt and
Chourey, 1985; Chourey et al., 1988). The distribution of both proteins is further
distinguished under stress conditions (such as anaerobiosis) where tissue-specific
localization in roots is readily apparent (Rowland et al., 1989). Tissue specific
shifts in sucrose synthase have also been noted in wildtype maize root tips
incubated in glucose deficient and sufficient media (K. Nolte, University of Florida,


15
found that in soybean nodules, the Km of sucrose synthase for UDP (0.5 mM) was
lower than that of ADP and CDP (0.13 and 1.1 mM, respectively).
Delmer (1972a and b) characterized regulation of purified Phaseolus aureus
sucrose synthase and found a number of differences in the regulation of the
synthetic and degradative reactions. NADP, iodoacetic acid, and gibberellic acid
all stimulate sucrose degradation but inhibit sucrose synthesis. Pyrophosphate
also enhanced the degradative activity, but only in the presence of MgCI2. In
contrast, Pontis (1977) reported that P¡ inhibited the degradative reaction alone or
in the presence of Mg2+. Delmer also tested the effects of intermediates in
carbohydrate metabolism and found that G-1-P, G-6-P, F-6-P, F-1,6-BP, R-5-P, R-
1,5-BP, PEP and 3-PGA had little or no influence on the sucrose synthase reaction
in either direction when present at 2 mM. However, de Fekete (1969) and Pontis
(1977) both have reported that G-1-P, G-6-P and F-1,6-BP were inhibitory to the
degradative reaction at 2-5 mM without affecting the synthetic reaction. ATP, ADP
and AMP had no inhibitory effect on sucrose synthesis at 4 mM; however, the
degradative reaction was inhibited 30% by ADP, and 50% by both ADP and AMP.
/3-Phenylglucoside has also been shown to inhibit sucrose degradation almost
completely and sucrose synthesis by 50% (Wolosiuk and Pontis, 1974b; Lowell,
1986). This has proven useful for distinguishing activities of sucrose phosphate
synthase from sucrose synthase.
Pontis and coworkers (Pontis et al., 1972) found that the divalent cations
Mg2+, Mn2+, Ca2+ and Ba2+ at 5-10 mM activated sucrose synthase in the


38
Addition of casein increased initial enzyme activity compared to controls
(Table 3-1) and improved stabilization of sucrose synthase activity with time (Figure
3-3A & B).
In contrast to added protectants, inorganic phosphate (10 mM), a protein
regulator through its role in reversible phosphorylation (Bennett, 1984), added to
the extraction buffer decreased initial activity of sucrose synthase by ca. 40%
(Figure 3-3A & B).
Despite loss of activity in vitro and positive responses to apparent
protectants against protease activity, proteolytic degradation of sucrose synthase
from maize root tips was not detectable via either denaturing or native (Figure 3-4A
& B, respectively) western-blot analysis at various times after enzyme extraction.
Further, no change in charge or separation of subunits in native tetramers was
evident. Nor was any change evident with time from samples extracted with
added casein or phosphate (Figure 3-4C & D, respectively). However, changes
in enzyme activity do not necessarily result in changes in electrophoretic mobility.
Walker and Huber (1989) demonstrated that activation of sucrose phosphate
synthase by light or mannose (a P¡ sequestering sugar) did not affect
immunoprecipitation or mobility of subunit mobility during SDS-PAGE.
The possible involvement of tetramer stability in the loss of activity in vitro
was further examined by comparison of the extracts from sh1 (containing only
homotetramers of Sus1 encoded sucrose synthase) and Sh1 (containing both
hetero- and homotetramers of sucrose synthase). Endosperm tissue contains


2
undergoing rapid cell division such as shoot and root apices (Avigad, 1982).
Recent evidence, however, indicates that although much hydrolysis is often
observed, invertase activity may not be essential for sucrose uptake into either
sugar cane stems (Lingle, 1989; Thom and Maretzki, 1990) or maize kernels
(Schmalstig and Hitz, 1987). Chapleo and Hall (1989a, b, and c) also have
concluded that invertase was not essential to sucrose import into roots of Ricinus.
Giaquinta and co-workers (1983) showed that sucrose entering the roots of maize
via phloem does not have to pass through the extracellular space. Robbins (1958)
first reported that root tips of an inbred line of maize, Oh 43, were unable to
retrieve exogenous sucrose. B. Burr (Brookhaven National Laboratory, personal
communication) has indicated that the lack of retrieval might possibly be due to
an invertase deficiency. The absence of invertase activity could have important
implications for sucrose import, not only because of potential effects on the
retrieval system, but also because sucrose breakdown in such an instance could
theoretically proceed only via action of sucrose synthase. A mutant lacking
functional invertase in its roots would also be useful, in combination with nulls for
both sucrose synthase isozymes, to elucidate the individual roles for these
enzymes.
The purpose of the following research is to further elucidate the roles and
regulation of the two sucrose metabolizing enzymes sucrose synthase and
invertase in roots of maize. Specific objectives are to:


24
A naturally occurring acid invertase inhibitor has been detected in a number
of plant tissues including beet roots (Burakhanova et al., 1987), potato roots
(Pressy, 1967, 1968; Bracho and Whitaker, 1990a and b), maize endosperm
(Jaynes and Nelson, 1971), pea pollen (Malik and Sood, 1976) and Ipomea petals
(Winkenbach and Matile, 1970). In potato the inhibitor was characterized as a
small protein, binding irreversibly to acid invertase (Pressy, 1967; Anderson and
Ewing, 1978). Pressy (1967) found that the binding of the potato inhibitor to
invertase had a pH optimum of 4.5 (Pressy, 1967), and the enzyme-inhibitor
complex could be partially disassociated by low pH or high Mg2+ concentrations.
In contrast, Bracho and Whitaker (1990a) found no effect of pH on inhibitor
binding. Sucrose at 2 mM could inhibit binding, but would not dissociate a
complex already formed (Pressy, 1967). Neither glucose nor fructose had a similar
effect. Matsushita and Uritani (1974) noticed a marked increase of acid invertase
activity resulted from wounding of sweet potato roots, but alkaline invertase activity
did not change under similar conditions. They also isolated a heat-stable protein
component with a molecular weight of approximately 19.5 kD, that fluctuated
during the incubation period after the wounding (Matsushita and Uritani, 1976).
They found that this putative inhibitor declined with a concomitant rise in invertase
activity early in the incubation, but increased in later stages when invertase activity
declined (Matsushita and Uritani, 1977). Pressy (1967, 1968) and Matsushita and
Uritani (1977) have suggested that the increase in invertase activity caused by cold
treatment or by wounding could be explained by a decrease in binding of the


6
Huber and Akazawa (1986) both demonstrate the potential importance of
simultaneous sucrose catabolism via two enzyme systems.
Sucrose Synthase
Sucrose synthase (EC 2.4.1.13, UDP-D-glucose: D-fructose 2-a-D-
glucosyltransferase) is ubiquitous in higher plants (Keller et al., 1988) and probably
occurs in all types of tissues. However, this enzyme is found in greatest
abundance in nonphotosynthetic tissues and in developing seeds (Echt and
Chourey, 1985). In cell fractionation studies, sucrose synthase was shown to be
associated with the soluble fraction (Nishimura and Beevers, 1979; MacDonald and
ap Rees, 1983). A cytosolic rather than vacuolar localization for sucrose synthase
has been demonstrated in protoplasts isolated from Jerusalem artichoke (Keller
et al., 1988).
Although molecular weights as high as one million have been reported for
sucrose synthase (Grimes et al., 1970), it is now generally concluded that, in its
native state, sucrose synthase has a molecular weight of approximately 36 to 40
kD (Delmer, 1972a; Su and Preiss, 1978; Morell and Copeland, 1985; Moriguchi
and Yamaki, 1988) and is composed of four identical subunits. The sucrose
synthase subunit from maize has been found to have a molecular weight of 8.8 kD
(Su and Preiss, 1978). However, unlike many other plant species, maize has two
genes which encode sucrose synthase subunits (Chourey and Nelson, 1976; Echt
and Chourey, 1985). The two sucrose synthase subunits of maize, sh1 and sus,
encoded by the Sh1 and Sus1 genes, respectively, have similar enzyme kinetics,


69
16h 24h 48h 16h + 32h
Intact + ~ + + "/+ +/-
Figure 5-4. Protein gel blot of Sus 1 encoded sucrose synthase from maize roots
incubated in 0% (-) or 2.0% (+) glucose for various time periods. At 16 h
switching treatments were conducted by changing roots in 0% glucose to
2.0% glucose (-/+) and vise verse (+/-); incubations were continued for 32
additional hours.


Sucrose cleaved (pmol g_1 protein h'1)


56
level. Occurrence of maximal enzyme activities in each experiment from samples
having the highest levels of both sucrose synthase genes, in fact, tends to indicate
that protein changes might be occurring but are not completely detectable under
the assay conditions utilized.


This dissertation was submitted to the Graduate Faculty of the College of
Agriculture and to the Graduate School and was accepted as partial fulfillment of
the requirements for the degree of Doctor of Philosophy.
August, 1991
cuA
Dean, College of AgrioliKure
Dean, Graduate School


82
detectable farther than 8 mm from the apex. Soluble enzyme accounted for 88 to
92% of the total acid invertase activity, and the remainder was due to action of the
insoluble enzyme. Neutral invertase activity was insignificant or absent from all
lines (data not shown). Although invertase action was essentially undetectable in
the primary roots of Oh 43, adventitious root extracts showed levels of activity as
least as great as those from the primary roots of other lines tested (Table 6-2).
Active acid invertase was released into buffer in the extracellular space
around adventitious root tips of Oh 43 during 3 hours of emersion (13.4 3.5 /imol
glucose g'1 FW h'1-ca. 25% of the activity on a fresh weight basis of N2 and buffer
extracted roots). The same was not observed during emersion of primary roots
from this line. Also, from 83% to 94% (13.3 to 15.1 Mmol glucose mg1 protein h"1)
of the invertase activity in adventitious root extracts remained after addition of
extracts from primary roots of Oh 43.
Unlike invertase, sucrose synthase was active in extracts of both primary
and adventitious roots from Oh 43 seedlings. Activity of this enzyme was
approximately similar in both root types, as observed for the other two lines tested
(Table 6-2).
Extracts of lateral roots originating from primaries exhibited no detectable
acid invertase activity (Table 6-3), in contrast to counterparts derived from
adventitious roots. Shoots of 5-day-old Oh 43 seedlings and endosperm or
scutellum tissue from developing kernels (23 DAP) also exhibited activity of both
soluble and insoluble acid invertase (Table 6-3).


96
Carlson, M., L. Neigeborn and L. Sarokin. 1987. Genetics of carbon catabolite
repression of the SUC2 gene in Saccharomyces cervisiae. pp. 25-32 in G.G.
Stewart, I. Russell, R.D. Klein and R.R. Heibsch, eds. Biological Research
on Industrial Yeasts. Vol. III. Boca Raton: CRC Press.
Carpita, N.C. and D.P. Delmer. 1981. Concentration and metabolic turnover of
UDP-glucose in developing cotton fibers. J. Biol. Chem. 256:308-315.
Chan, H-Y., T-Y. Ling, R-H. Juang, l-N. Ting, H-Y Sung and J-C. Su. 1990. Sucrose
synthase in rice plants. Growth-associated changes in tissue specific
distributions. Plant Physiol. 94:1456-1461.
Chang, C.W. and R.S. Bandurski. 1963. Exocellular enzymes of corn roots. Plant
Physiol. 38:60-64.
Chapleo, S. and J.L. Hall. 1989a. Sugar unloading into roots of Ricinus communis
L. I. The characteristics of enzymes concerned with sucrose catabolism and
a comparison of their distribution in roots and shoot tissues. New Phytol.
111:369-379.
Chapleo, S. and J.L. Hall. 1989b. Sugar unloading into roots of Ricinus communis
L. II. Characteristics of the extravascular apoplast. New Phytol. 111:381 -390.
Chapleo, S. and J.L. Hall. 1989c. Sugar unloading in roots of Ricinus communis
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396.
Chen, Y-C. and P.S. Chourey. 1989. Spatial and temporal expression of the two
sucrose synthase genes in maize: immunohistological evidence. Theor.
Appl. Genet. 78:553-559.
Chin, C.K. and G.D. Weston. 1973. Distribution in excised Lycopersicum
esculentum roots of the principal enzymes involved in sucrose metabolism.
Phytochem. 12:1229-1235.
Chourey, P.S. 1981. Genetic control of sucrose synthetase in maize endosperm.
Mol. Gen. Genet. 184:372-376.
Chourey, P.S. 1988. Recombinants lacking in detectable levels of both sucrose
synthases are functionally normal. Maize Genet. Coop. Newsletter 62:62-63.
Chourey, P.S., Y-C. Chen and M.E. Miller. 1991a. Early cell degeneration in
developing endosperm is unique to the shrunken mutation in maize.
Maydica 36:141-146.


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Morell, M. and L Copeland. 1984. Enzymes of sucrose breakdown in soybean
nodules: Alkaline invertase. Plant Physiol. 74:1030-1034.
Morell, M. and L. Copeland. 1985. Sucrose synthase of soybean (Glycine max cv.
Williams) nodules. Plant Physiol. 78: 149-154.
Moriguchi, T. and S. Yamaki. 1988. Purification and characterization of sucrose
synthase from peach (Prunus prsica) fruit. Plant and Cell Physiol. 29:1361-
1366.
Morrell, S. and T. ap Rees. 1986. Sugar metabolism in developing tubers of
Solarium tuberosum. Phytochem. 25:1579-1586.
Mujer, C.V., M.E. Rumpho, T.C. Fox and R.A. Kennedy. 1990. Developmental
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of Echinochloa phyllopogon. Plant Physiol. 93S:101.
Murata, T. 1971. Enzymic mechanisms of starch synthesis in sweet potato roots.
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Murata, T. 1975. Soluble nucleotides in Konjak corn. Agrie. Biol. Chem 39:1401-
1406.
Nelson, N. 1944. A photometric adaptation of the Somogyi method for the
determination of glucose. J. Biol. Chem. 153:357-380.
Neufeld, E.F. and W.Z. Hassid. 1963. Biosynthesis of saccharides from
glucopyranosyl esters of nucleotides (sugar nucleotides). Adv.
Carbohydrate Chem. 18:309-356.
Neuffer, M.G., L. Jones and M.S. Zuber. 1968. The Mutants of Maize. Madison, Wl:
Crop Science Society of America.
Nilan, R.A., A. Kleinhofs and R.L. Warner. 1981. Use of induced mutants of genes
controlling nitrate reductase, starch deposition, and anthocyanin synthesis
in barley, pp. 183-200 in Induced Mutations-A Tool in Plant Research.
Vienna, Austria: International Atomic Energy Agency.
Nishimura, M. and H. Beevers. 1979. Subcellular distribution of gluconeogenetic
enzymes in germinating castor bean endosperm. Plant Physiol. 64:31-37.


89
utilization of exogenous sucrose (Robbins, 1958). Specific tissue localization of
invertase also has been described. Peak activity for root invertase is generally 2-3
mm behind the apex and corresponds to the region of expansion and elongation
in pea (Robinson and Brown, 1952) and maize (Hellebust and Forward, 1962). In
Ricinus this activity predominates in the cortex (Chapleo and Hall, 1989).
Although invertase may not have a direct role in sucrose import in roots, it
still may be important to two major aspects of root biology. First, invertase has
been implicated in formation of mycorrhizal associations (Purves and Hadley,
1975). Maize (Gerdemann, 1964; Kothari et al., 1990) and 90% of other
agriculturally important species form these beneficial symbioses under field
conditions (Gerdemann, 1986). Invertase activity typically increases and hexose
levels rise at infection sites of biotrophic fungi (Long et al., 1975). Elevated hexose
content in roots upon infection by mycorrhizal fungi has been attributed to a rise
in invertase levels (Purves and Hadley, 1975). It is not known whether this is host
or fungal invertase; however, Oh 43 does not appear to provide the former in its
primary root systems.
Second, the purported lack of a sucrose carrier in the plasma membrane
of maize root cells (Lin et al., 1984) would indicate that if sucrose were released
into the apoplast, retrieval might proceed more effectively in the presence of
extracellular invertase. Such retrieval could be particularly important during stress
or periods when sugar losses from the symplast were elevated. Any physiological
consequence of this deficiency would most likely be evident early in seedling


Sucrose cleaved (jumol mg 1protein h'1)
37
Time after extraction (min)
Figure 3-2. Time course of /'/7 vitro decrease in maize sucrose synthase activity with
and without the serine proteinase inhibitor, PMSF. No other class-specific
proteinase inhibitors tested affected initial sucrose synthase activity. PMSF
(1 mM) was used in extraction buffer and equilibration of desalting columns.
Bars represent SE, n=3.


40
Time after Extraction
(min)
4 10 20 60
A
5 15 20 60
b
c
D nnnn
Figure 3-4. Denaturing (A) and native (B,C,D) protein gel blot analysis of maize
root sucrose synthase at various times after extraction. Sub-samples were
removed at designated intervals during incubation at 4 C. Proteins were
separated by polyacrylamide gel electrophoresis with (A) or without (B.C.D)
SDS and sucrose synthase resolved by probing with a polyclonal antibody
raised against protein products of both the Sh1 and Sus genes. The five
bands visible in the native gel blot have been described as corresponding
to homo- and heterotetrameric forms of sucrose synthase composed
entirely of products from the Sh 1 gene (uppermost band), the Sus gene
(lowermost band) and combinations of the two (middle three bands).
Extracts for denaturing (A) and native (B) blots were extracted with buffer
only. Casein (2% w:v) (C) and Pi (10 mM) (D) were tested as protectants
of enzyme stability.


92
length, and inclusion (or exclusion) of various anti-oxidants and protease inhibitors.
The assay has proven effective for a range of tissues and species examined and
provides a particularly effective measurement for sucrose synthase action in root
tips. Characterization of the in vitro instability also provided evidence suggestive
of possible phosphorylation effects on activity.
Activity of sucrose synthase was then assayed in extracts from intact
wildtype maize root tips and from those that had been excised and incubated for
24 h in a basal salts medium with varying levels of glucose. Enzyme activity
showed little or no significant differences between treatments. However, results
represented contributions by two isozymes (encoded by genes exhibiting
reciprocal responses under the same conditions [Koch et al., 1989]), and were the
collective sum of different tissues (that showed specific alterations in sucrose
synthase distribution [K. Nolte, University of Florida, unpublished data]).
A different approach was thus utilized to examine the relationship between
sugar-modulated changes in message levels and extent of enzyme level effects.
Use of the shrunken-1 maize mutant (deficient in a functional Sh 1 gene) showed
that carbohydrate-responsive gene expression was evident for Sus1 at the levels
of mRNA, protein and enzyme activity. Time-course and treatment reversals also
indicated greater responses between glucose sufficient and deficient treatments
were observed after 48 h of incubation, indicating a possible coarse control
mechanism.
The second purpose of this work was to explore a possible line of


32
were subsequently adjusted to 1 M and ca. 0.11 /Ci per /I. Two and one-half /I
(ca. 0.27 /iCi) were used in each reaction.
Tissue Extraction
Weighed tissue (100-200 mg) was frozen and ground to a powder in liquid
N2 with a mortar and pestle. The frozen powder was transferred to another mortar
containing ice-cold extraction buffer (200 mM HEPES buffer [pH 7.5], with 1 mM
DTT, 5 mM MgCI, 1 mM EGTA, 20 mM sodium ascorbate, 1 mM PMSF and 10%
[w/w] PVPP) and ground briefly in this medium. One ml of grinding buffer was
used for every 100 mg tissue fresh weight. Cysteine (10 mM) was initially but was
omitted to prevent non-specific binding of radiolabel to DEAE cellulose paper.
Two-hundred il of extract were placed on each of 4 to 8 spun columns
packed with Sephadex G 50-80 hydrated with extraction buffer. Columns were
centrifuged for 1 min at 800 x g. Eluent from each column was pooled with others
from the same tissue sample before assay. Ratio of sample to bed volume was
maximized at 1:5 (v:v) by HPLC detection of soluble sugar presence in eluent
(Yelle, 1991).
Enzyme Assay
Cleavage of 14C-sucrose by sucrose synthase was assayed in a 50 /I
volume consisting of 20 /I extract, 80 mM Mes (pH 5.5), 5 mM NaF, 100 mM de
suerse and 5 mM UDP. Reactions proceed for 5 minutes at 30 C and were
terminated by adding 50 /xl of Tris (pH 8.7) and boiling for 1 min. Controls


9
concentrations are generally elevated in importing areas. Therefore, substrate
levels favor the cleavage reaction in the majority of instances.
A second line of evidence for the degradative role of sucrose synthase in
importing organs is provided by mutant maize lines lacking a functional sucrose
synthase protein. Chourey and Nelson (1976) showed that a deletion of the Sh
locus on chromosome 9 of maize (coding for sucrose synthase) led to a 90%
deficiency of the respective protein in mutant vs wild-type kernels. Starch
formation was also reduced in this line, giving rise to a "shrunken" seed. The
association between this shrunken phenotype and a sucrose synthase deficiency
was considered evidence that the critical reaction in vivo was that of sucrose
cleavage, and that this was essential for conversion of photosynthetically produced
sucrose for starch biosynthesis. The residual amount of starch deposited was
attributed to the sucrose degrading activity of a second sucrose synthase encoded
by another locus (Sus1).
Further research also favors the cleavage role of sucrose synthase and
implicates its involvement in starch deposition. Dale and Housley (1986), for
example, found that developing wheat kernels with the greatest rates of growth
and starch deposition had significantly greater sucrose synthase activities. A
positive correlation between sucrose synthase activity and starch deposition was
also reported in Pisum sativum by Edwards and ap Rees (1986a and b). They
proposed that UDP-glucose formed during sucrose cleavage was converted to
glucose-1-phosphate by UDP-glucose pyrophosphorylase using pyrophosphate


12
sinks vary from species to species (ap Rees, 1974, Hawker, 1985). However,
sucrose breakdown generally appears to proceed via sucrose synthase in starch
and sugar storage sinks (Sung et al., 1988).
Sucrose synthase also may be associated with sink strength through its
potential involvement in cell wall synthesis (Hendrix, 1990). Sucrose synthase
activity predominates over that of invertase in rapidly expanding cotton ovules, for
example where rapid elongation of epidermal hairs (cotton fibers) requires
extensive cellulose formation. Hendrix (1990) postulated that sucrose entering the
seed coat in the developing cotton boll was cleaved via sucrose synthase, and
subsequent carbohydrates went into the rapidly growing epidermal hairs.
However, some of the sucrose breakdown products were converted to starch
stored in the seed coats. Stepanenko and Morozova (1970) demonstrated
cellulose biosynthesis from UDP-glucose in cotton. However, they did not indicate
that the source of the UDPG might come from the action of sucrose synthase.
Carpita and Delmer (1981) showed that the rate of synthesis and turnover of UDP-
glucose in developing cotton ovules was more than sufficient to account for rates
of biosynthesis for cellulose, 0-1,3-glucan and sterylglucosides (all cell wall
constituents). They found that UDPG levels increased dramatically just prior to the
maximum rate of secondary wall cell synthesis and dropped precipitously at the
time when cellulose synthesis ceased. Again, sucrose synthase activity was not
measured, but could possibly be the source of the increased levels of UDPG.
Sucrose synthase activity was measured in cultured cells of Catharanthus roseus


28
Sus 1 gene has been described (Chourey et al., 1988), but, unlike the Sh 1 mutant,
the Sus 1 mutant does not have any detectable phenotypic abnormality. A
mutation lacking detectable levels of both sucrose synthase isozymes also has
been described (Chourey, 1988), but its existence is puzzling considering the
expected lethality of a complete sucrose synthase deficiency.


93
investigation into the physiological significance of invertase by investigating a
putative deficiency in sucrose-metabolizing capacity in the Oh 43 line of maize
(Robbins, 1958). Complete absence of invertase in this line was not likely due to
its demonstrated activity in the scutellum (Doehlert et al., 1988). However, the
hypothesis tested here was that an organ-specific deficiency or invertase
suppression was responsible for the metabolic anomalies in Oh 43. The primary
root of Oh 43 indeed was shown to lack invertase activity. In contrast, adventitious
roots of the same plants exhibited wildtype levels of invertase activity. Initial
characterization of this mutation will provide an effective tool for future
investigations into the physiological role(s) of this enzyme in roots. Use of this
mutation and isogeneic wildtype counterparts in combination with the nulls for
each sucrose synthase gene may allow a better understanding of the biological
functions of each.


8
The necessity for two systems of sucrose synthesis was puzzling until it was
determined that the sucrose synthase reaction is readily reversible (Cardini et al,
1955):
Sucrose + UDP < > UDP-Glucose + Fructose
This reversibility gave rise to the suggestion that sucrose synthase could make
UDPG available for utilization as a glucosyl donor in starch synthesis (Turner and
Turner, 1957). Other studies of sucrose synthase specificity and kinetics led
Avigad and coworkers (Avigad, 1964; Avigad et al., 1964; Milner and Avigad, 1964)
to suggest that this enzyme functioned mainly in sucrose cleavage in storage
tissues. Sucrose synthase activity typically is highest in tissues during periods of
rapid growth and is often not accompanied by high invertase activity (Schaffer et
al., 1987).
Several factors lend credence to the view that the role of sucrose synthase
is sucrose cleavage in importing cells. First, a substantial level of free fructose is
required for sucrose synthase activity in the synthetic direction (Km ca. 2.0-2.5 mM
[Avigad, 1982]). Levels of this sugar are low in healthy, intact leaves, but they are
higher in storage tissues and roots, where most of the free fructose is in the
vacuole or extracellular spaces (Avigad, 1982). Availability of UDPG is also likely
to limit the synthetic reaction, because the cellular concentration is typically less
than 0.4 mM (Murata, 1975). The Km for UDPG ranges from 0.1 to 8.5 mM
(average approximately 2.0 mM)(Avigad, 1982). In contrast, sucrose


64
% glucose
Intact 0 0.2 0.5 2.0 4.0
Expt. 1
Expt. 2
Figure 5-1. RNA gel blot analysis of Sus1 expression in maize roots incubated in
a range of glucose concentrations for 24 hours.


104
Lingle, S.E. 1989. Evidence for the uptake of sucrose intact into sugarcane
internodes. Plant Physiol. 90:6-8.
Lingle, S.E. and J.R. Dunlap. 1987. Sucrose metabolism in netted muskmelon fruit
during development. Plant Physiol. 84:386-389.
Little, B. and J. Edelman, 1973. Solubility of plant invertases. Phytochem. 12:67-71.
Long, D.E., A.K. Fung, E.E.M. McGee, R.C. Cooke and D.H. Lewis. 1975. The
activity of invertase and its relevance to the accumulation of storage
polysaccharides in leaves infected by biotrophic fungi. New Phytol. 74:173-
182.
Loomis, W.D. 1969. Removal of phenolic compounds during the isolation of plant
enzymes, pp. 555-563 in J.M. Lowenstein, ed. Methods in Enzymology. Vol.
XIII. Citric Acid Cycle. New York: Academic Press.
Lowell, C.A. 1986. Structure and sucrose metabolizing enzymes of the transport
path: Implications for assimilate translocation in grapefruit. Ph.D.
Dissertation. Gainesville: University of Florida.
Lowell, C.A., P.T. Tomlinson and K.E. Koch. 1989. Sucrose-metabolizing enzymes
in transport tissues and adjacent sink structures in developing citrus fruit.
Plant Physiol. 90:1394-1402.
Lyne, R.L. and T. ap Rees. 1971. Invertase and sugar content during differentiation
of roots of Pisum sativum. Phytochem. 10:2593-2599.
Maas, C., S. Schaal and W. Werr. 1990. A feedback control element near the
transcription start site of the maize Shrunken gene determines promoter
activity. EMBO J. 9:3447-3452.
MacDonald, F.D. and T. ap Rees. 1983. Enzymic properties of amyloplasts from
suspension cultures of soybean. Biochim. Biophys. Acta 755:81-89.
Malik, C.P. and R. Sood. 1976. Ecophysiological regulation of invertase activity in
pea pollen. Ind. J. Ecol. 3:44-48.
Maraa, C. F. Garcia-Olmedo and P. Carbonero. 1990. Differential expression of
two types of sucrose synthase-encoding genes in wheat in response to
anaerobiosis, cold shock and light. Gene 88:167-172.


102
Holbein, B.E., C.W. Forsberg and D.K. Kidby. 1976. A modified procedure for
studying enzyme secretion in yeast sphaeroplasts: Subcellular distribution
of invertase. Can. J. Microbiol. 22:989-995.
Hubbard, N.L., S.C. Huber and D.M. Pharr. 1989. Sucrose phosphate synthase
and acid invertase as determinants of sucrose concentration in developing
muskmelon (Cucumis melo L.) fruits. Plant Physiol. 42:1527-1534.
Huber, J.L.A., S.C. Huber and T.H. Nielsen. 1989a. Protein phosphorylation as a
mechanism for regulation of spinach leaf sucrose phosphate synthase
activity. Arch. Biochem. Biophys. 270:681-690.
Huber, S.C. and T. Akazawa. 1986. A novel sucrose synthase pathway for sucrose
degradation in cultured sycamore cells. Plant Physiol. 81:1008-1013.
Huber, S.C., T.H. Nielsen, J.L.A. Huber and D.M. Pharr. 1989b. Variation among
species in light activation of sucrose phosphate synthase. Plant and Cell
Physiol. 30:277-285.
Humphreys, T.E. 1974. Sucrose transport and hexose release in the maize
scutellum. Phytochem. 13:2387-2396.
Jaynes, T.A. and O.E. Nelson. 1971. An invertase inactivator in maize endosperm
and factors affecting inactivation. Plant Physiol. 47:629-634.
Jeffs, R.A. and D.H. Northcote. 1966. Experimental induction of vascular tissue in
an undifferentiated plant callus. Biochem. J. 101:146-152.
Jeffs, R.A. and D.H. Northcote. 1967. The influence of indol-2yl acetic acid and
sugar on the pattern of induced differentiation in plant tissue culture. J. Cell
Sci. 2:77-88.
Kato, T. and S. Kubota. 1978. Properties of invertases in sugar storage tissues of
citrus fruit and changes in their activities during maturation. Physiol. Plant.
42:283-297.
Kaufman, P.B., N.S. Ghosheh, J.D. LaCroix, S.L. Soni and H. Ikuma. 1973.
Regulation of invertase levels in Avena stem segments by gibberellic acid,
sucrose, glucose, and fructose. Plant Physiol. 52:221-228.
Keller, F., M. Frehner and A. Wiemken. 1988. Sucrose synthase, a cytosolic
enzyme in protoplasts of Jerusalem artichoke tubers (Helianthus tuberosus
L.) Plant Physiol. 88:239-241.


44
1970; Salerno et al., 1979; Keller et al., 1988; Lowell et al., 1989). Measurements
of the synthetic reaction have been based on quantification of either sucrose or
UDP production. Sucrose levels can be determined indirectly by using invertase
for full conversion to hexoses and measuring glucose colorimetrically (Avigad and
Milner, 1966). The latter method, however, is susceptible to interference by
substances in the crude enzyme extracts of many plants (Pontis, 1977). It has
also been possible to measure 14C-sucrose formed from UDP-14C-glucose by
separating labeled product from substrate with anionic resins (Salerno et al., 1979),
paper electrophoresis (Grimes et al., 1970) or paper chromatography (Pontis,
1970). Such radioactive assays have proven useful in systems where colorimetric
methods have been problematic (Pontis, 1977). In addition, UDP production can
be determined spectrophotometrically by coupling its formation to the pyruvate
kinase-lactate dehydrogenase reaction and measuring the decrease in absorbance
due to oxidation of NADH (Avigad, 1964; Avigad and Milner, 1966; Lowell et al.,
1989).
Procedures for assaying the cleavage reaction are based on determination
of fructose or UDP-glucose formation. Fructose can be measured colorimetrically
by the Nelson reducing sugar assay (1944), or spectrophotometrically by coupling
hexokinase, phosphoglucose isomerase and glucose 6-phosphate dehydrogenase
for production of NADPH (Avigad, 1964; Keller et al., 1988). Also, UDP-glucose
formation can be estimated by coupling its appearance to NAD reduction by UDP-
dehydrogenase (Avigad, 1964; Lowell, 1986; Lowell et al., 1989). Degradative


112
Wolosiuk, R.A. and H.G. Pontis. 1974a. Studies on sucrose synthetase kinetic
mechanism. Arch. Biochem. Biophys. 165:140-145.
Wolosiuk, R.A. and H.G. Pontis. 1974b. The role of sucrose and sucrose
synthetase in carbohydrate plant metabolism. Mol. Cell Biochem. 4:115-123.
Wright, K. and D.H. Northcote. 1972. Induced root differentiation on sycamore
callus. J. Cell Sci. 11:319-337.
Wyse, R. 1986. Sinks as determinants of assimilate partitioning: Possible sites for
regulation, pp. 197-210 in J. Cronshaw, W.J. Lucus, R.T. Giaquinta, eds.
Phloem Transport. New York: Alan R. Liss, Inc.
Xie, Y. and R. Wu. 1989. Rice alcohol dehydrogenase genes: anaerobic induction,
organ specific expression and characterization of cDNA clones. Plant Mol.
Biol. 13:53-68.
Xu, D-P., S.S. Sung, C.A. Alvarez and C.C. Black. 1986. Pyrophosphate-dependent
sucrose metabolism and its activation by fructose 2,6-bisphosphate in
sucrose importing plant tissues. Biochem. Biophys. Res. Comm. 141:440-
445.
Xu, D-P., S.J.S. Sung, T. Loboda, P.P. Kormanik and C.C. Black. 1989.
Characterization of sucrolysis via the uridine diphosphate and
pyrophosphate-dependent sucrose synthase pathway. Plant Physiol.
90:635-642.
Yelle, S., R.T. Chetelat, M. Doris, J.W. DeVerna and A.B. Bennett. 1990. Sink
metabolism in tomato fruit. IV. Genetic and biochemical analysis of sucrose
accumulation. Plant Physiol. 95:1026-1035.
Yelle, S., J.D. Hewitt, N.L. Robinson, S. Damon and A.B. Bennett. 1988. Sink
metabolism in tomato fruit III. Analysis of carbohydrate assimilation in a wild
species. Plant Physiol. 87:737-740.


10
generated by PFK(PP¡). Morrell and ap Rees (1986) have also suggested that
much of the sucrose translocated to developing potato tubers is probably
metabolized via the same pathway with the initial step catalyzed by sucrose
synthase. Gibson and Shine (1983) have demonstrate that in the presence of
inorganic phosphate, UDPG may be hydrolyzed to G-1-P and UDP by the action
of UDPG phosphorylase. Salerno (1986) also has demonstrated the presence a
highly nucleotide specific form of UDP-glucose phosphorylase in developing maize
endosperms. The activity level of this enzyme followed closely the development
of the grain and paralleled that of sucrose synthase. The presence of this enzyme
links sucrose cleavage and starch formation via sucrose synthase.
Finally, an additional line of evidence was presented by Cobb and Hannah
(1988) that also indicated a primarily degradative function for sucrose synthase in
importing organs. They showed that maize kernels from a line deficient in the Sh 1
gene for sucrose synthase still had normal levels of sucrose and normal rates of
sucrose synthesis when grown in culture with fructose as the carbon source. If
sucrose synthesis had been proceeding via sucrose synthase in wild-type kernels,
then the loss of ca. 90% of total sucrose synthase activity in kernels of the mutant
line should have affected sucrose formation there. The authors concluded that
Sh1 encoded sucrose synthase was not necessary for sucrose synthesis.
Correlative data suggest that sucrose synthase activity is closely linked with
sink strength. The supply and timing of sucrose for export seem to be closely
related to the source of photosynthate (Fondy and Geiger, 1982; Servaites et al.,


Figure 3-5. Time course of in vitro decrease in maize root sucrose synthase activity from lines with homo- (W22:s/77) and
heterotetrameric (NK 508) forms of this enzyme. Bars represent SE, n=3. Data from one replicate using material
isogeneic to \N22:sh1 except for the Sh1 gene (W22) resulted in a curve similar in appearance and of the same
magnitude as that of W22:sh1.


66
Table 5-1. Sucrose synthase activity in mutant maize (W22:s/77) root tips
incubated in media containing a range of glucose concentrations for 24
hours.
Sucrose synthase activity
% glucose
Intact
0
0.2
0.5 2.0
4.0
(/xmol sucrose g1
protein h"1)
Expt. 1
0.22
0.13
0.07
0.13 0.32
0.32
Expt. 2
0.35
0.21
0.13
0.20 0.40
0.39


63
synthase antisera, obtained from D.R. McCarty, was generated in rabbits using
protein purified from maize kernels (W64A x 182E) 22 days after pollination.
Antisera was diluted 1:1000 and cross reacted strongly to both the Sh 1 and Sus 1
gene products where such were present.
Results
Levels of Sus 1 mRNA in excised maize roots were greater after 24 h of
incubation in 2% glucose than in those that had received 0 or 0.2% glucose
(Figure 5-1). At the protein level, western blot analysis showed little or no change
with increasing carbohydrate concentration (Figure 5-2); however, enzyme activity
was elevated in root tips incubated at high vs. low glucose concentrations (Table
5-1). Both lines of evidence indicated the protein level response was less
pronounced at 24 h than that of mRNA.
The time-course of changes in Sus1 message levels in root tips showed that
differences between those given 0% vs. 2.0% exogenous glucose became
apparent sometime between 16 and 24 h (Figure 5-3). Initial decreases appeared
to occur in both treatments, but within 24 h, Sus 1 mRNA levels in glucose
supplemented roots had risen well above those with limited sugar supply. The
greatest difference between carbohydrate treatments was evident after 48 h of
incubation. Treatment reversals indicated that the gene response to carbohydrate
level had been initiated within 16 h (Figure 5-3). Roots incubated in 2.0% glucose
for 16 h and switched to 0% for 32 h (total of 48 h) responded like those remaining
continuously in 2.0% glucose. Roots initially deprived of glucose and then


13
(Amino et al., 1985) and found to be elevated during the G1 phase when the
amount of total cell walls increased significantly. However, UDP-glucose
pyrophosphorylase activity (also involved in the formation of UDPG) was greater
than sucrose synthase activity at the G1 phase. The former was considered by
these authors likely to make a more important contribution to the total UDPG
formed. Chourey et al. (1991a) have reiterated the hypothesis that the resultant
shrunken, starch-deficient endosperm of the sh1 maize mutant may be due to
reduced cell wall deposition rather than any direct effect on the starch biosynthetic
pathway.
Regulation of Sucrose Svnthase
Sucrose synthase, a key enzyme in sucrose metabolism, is subject to a
number of complex regulatory factors (Davies, 1974). This enzyme exhibits a wide
specificity for the nucleoside base utilized in the reaction. Most enzymes of sugar
nucleoside metabolism show a marked specificity for a particular base (Avigad,
1982). Sucrose synthase working in the synthetic direction has been shown to
utilize UDPG, ADPG, TDPG, CDPG and GDPG as glucosyl group donors (Avigad,
1982). The Km for UDPG, however, is usually much less than for other NDPGs.
Grimes et al. (1970) found that the Km for UDPG was approximately 0.2 mM while
ADPG, TDPG, CDPG and GDPG had Kms of 1.8,1.7, 2.5 and 2.5 mM, respectively.
They also found that with UDPG as the nucleoside sugar, the Km for fructose was
reduced 10 fold below that apparent when ADPG was utilized. This change was


23
however the differences in values, especially those of mature tissues, were
considered unlikely to be wholly artifactual. Distinguishing between forms of
invertase is further complicated by information obtained from yeast. One gene in
yeast, SUC2, has been shown to encode the two forms of invertase in yeast,
secreted and intracellular, via two differentially regulated mRNAs (Carlson and
Botstein, 1982).
The Kms of acid invertase for sucrose generally range from 2 to 13 mM.
Sucrose is the primary substrate for acid invertase but raffinose also is hydrolyzed,
though at a slower rate (10% to 50% the rate of sucrose) (Avigad, 1982). Acid
invertase from sugar-cane leaves was inhibited competitively by fructose (K¡ 32
mM) and noncompetitively by glucose (K¡ 37 mM) (Sampietro et al., 1980). Acid
invertases have been partially purified from a number of tissues with apparent
molecular weights ranging from 2.8 x 104 to 2.2 x 105 (Roberts, 1973; Ricardo,
1974; Kato and Kubota, 1978; Masuda and Sugawara, 1980; Sum et al., 1980;
Faye et al., 1981).
The SUC2 gene of Saccharomyces, encoding invertase, has been shown
to be modulated by glucose levels (Carlson et al., 1987). Sucrose or raffinose,
substrates of the yeast invertase, have no such effect. Kaufman et al. (1973) found
that acid invertase activity rises in Avena stem segments incubated in a sucrose-
containing medium. The response had a lag time of 10-12 hours, suggesting a
change in protein levels. Fructose in the incubation medium resulted in a similar
response, but glucose caused no change in invertase activity.


18
Invertases are specific for the fructofuranose moiety of sucrose and work by
hydrolyzing the glycosidic linkage between the bridge oxygen and the fructose
residue (Sum et al., 1980). Up to five different forms of invertase have been
reported in plants (Sasaki et al., 1971). However, these enzymes are generally
divided into two main types based on the pH at which sucrose hydrolysis is most
efficiently accomplished. Acid invertases have pH optima around 4.5 to 5.0; that
of alkaline (or neutral invertase) is 7.0 to 7.5.
Most invertases are glycoproteins. Arnold (1966) partially purified an acid
invertase from grapes and found that it was approximately 25% carbohydrate.
Faye and coworkers (Faye and Berjonneau, 1979; Faye et al., 1981) have shown
a 7.7% carbohydrate content of acid invertase from radish seedlings, and date
invertase has a carbohydrate content of 8.2% (Al-Bakir and Whitaker, 1978).
Invertase preparations from barley (Prentice and Robbins, 1976), sugar cane (del
Rosario and Santisopasri, 1977), potato tubers (Anderson and Ewing, 1978;
Bracho and Whitaker, 1990b) and banana (Sum et al., 1980) were shown to bind
strongly to concanavalin A, a phytagglutinin or lectin isolated from jack bean with
a strong binding affinity for carbohydrates. In yeast and Neurospora the
carbohydrate content of invertase has been estimated to range from 0 to 50 %
(Metzenberg, 1963; Gascon et al., 1968; Holbein et al., 1976). Much of our more
detailed knowledge on the molecular structure and mode of action of invertases
comes from studies on fungal and yeast enzymes; however, considerable
progress has been made in analyses of plant invertases.


79
14,000 x g for 1 min to sediment particulate matter. Supernatant was dialyzed
(27,000 mw cutoff) at 4 C for 24 h against extraction buffer diluted 1:40. Buffer
was changed after 1 h and thereafter, every 4 h. Soluble dialyzed extract was
assayed for invertase as described below. Previously separated particulate matter
was rinsed with one volume of extraction buffer and assayed for insoluble, cell-
wall-bound invertase (soluble acid invertase includes both vacuolar and loosely
bound extracellular enzyme [Avigad, 1982]).
To test the possibility that the soluble enzyme was present in primary roots
of Oh 43 but was being bound or inactivated during the extraction procedure, two
additional extraction/assay methods were employed. First, adventitious root
extracts, previously shown to contain active invertase activity, were added to those
of primary apices. The resulting mixture was dialyzed and assayed for enzyme
activity. Second, three cm apices of both primary and adventitious roots Oh 43
roots were excised. Apices of these roots (0.5 cm) were suspended in extraction
buffer for three hours at 27 C. Buffer alone was subsequently dialyzed as
described above. The portion of each root which had been immersed in the
extraction buffer was excised for fresh weight measurement. After dialysis, the
buffer-enzyme solution was analyzed for enzyme activity.
Enzyme Assays
Soluble and insoluble forms of acid invertase were assayed as described
by Lowell et al. (1989). Reaction media contained 50 mM sucrose, and pH of 4.5
was adjusted with a sodium acetate buffer. Neutral invertase was assayed using


77
temperature (Rovira and Davey, 1974). The composition of root sugars exuded
is quite variable but includes both reducing and non-reducing sugars (Rovira and
Davey, 1974). Glucose and fructose are often taken up from the extracellular
space more rapidly than is sucrose (Humphreys, 1974). Retrieval of solutes from
the apoplast and the form in which they are available may thus be a potentially
important attribute of root carbon balance.
A deficiency in this retrieval process was first indicted by Robbins report
(1958) that roots of a maize inbred, Oh 43, were unable to grow on sucrose agar
medium, yet roots of another line, Hy 2, grew quite well. Only when roots of both
were cultured immediately adjacent to one another, were those of Oh 43 able to
grow. Growth of excised Oh 43 root tips also occurred when glucose was
substituted for the sucrose. Oh 43 was concluded to be "incapable of inverting
sucrose" in its root tips. Preliminary investigations by B. Burr (Brookhaven National
Laboratory, personal communication) indicated that a lack of invertase may have
been the reason for the inability of Oh 43 roots to metabolize sucrose.
The absence of invertase activity could have important implications for
sucrose import not only because of potential effects on the retrieval system, but
also because sucrose utilization in such an instance could theoretically be initiated
only via action of sucrose synthase. In addition, genetic material which lacks
activity of a specific enzyme can be useful in investigations of physiological
processes normally mediated by these enzymes (Koch et al., 1982). The present
report demonstrates that invertase is not essential for primary root growth despite



PAGE 1

SUCROSE SYNTHASE AND INVERTASE IN MAIZE ROOTS DIFFERING IN CARBOHYDRATE STATUS AND/OR GENETIC COMPOSITION By EDWIN RALPH DUKE A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT OF THE REQUIREMENTS FOR THE DEGREE OF DOCTOR OF PHILOSOPHY UNIVERSITY OF FLORIDA 1991

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ACKNOWLEDGEMENTS I would like to extend thanks to the members of my committee, Dr. Karen Koch, Dr. Rebecca Darnell, Dr. Don McCarty, Dr. Curt Hannah and Dr. Alice Harmon, for their assistance and guidance during the completion of this degree. I would also like to thank Dr. Tom Humphreys for his critical review of this dissertation. There are many people in the Fruit Crops Department to whom I owe thanks. I would like to thank the staff and faculty for their help and encouragement during my time here. For their friendship and support, I especially would like to thank Kathy Zimmerman, Teki and Andy Ericson, Dr. Kathy Taylor, Dr. Pat Tomlinson, Wayne Avigne, Kurt Nolte, and Don Merhaut. Thanks are also given to the graduate students, both past and present, of the Fruit Crops Department. Finally, I want to extend my most heartfelt thanks to my parents, Ralph and Mildred Duke; they have stood by me and given me support throughout my education, and I can never thank them enough.

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TABLE OF CONTENTS ACKNOWLEDGEMENTS ii LIST OF TABLES v LIST OF FIGURES vi ABSTRACT vii CHAPTERS 1 INTRODUCTION 1 2 REVIEW OF THE LITERATURE 4 Sucrose Metabolism 4 Sucrose Synthase 6 Invertases 17 Use of Mutants in Physiological Research 26 3 INSTABILITY OF SUCROSE SYNTHASE FROM ROOT TIPS: CHARACTERIZATION AND STABILIZATION 29 Abstract 29 Introduction 30 Materials and Methods 31 Results 34 Discussion 41 4 SUCROSE SYNTHASE ACTIVITY IN WILDTYPE MAIZE ROOT TIPS RESPONDING TO ALTERED CARBOHYDRATE STATUS 49 Abstract 49 Introduction 49 Materials and Methods 51 iii

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Results 52 Discussion 52 5 SUGAR RESPONSE OF SUCROSE SYNTHASE AT THE GENE {Sus1), PROTEIN AND ENZYME ACTIVITY LEVELS IN ROOTS OF THE Sh1 MAIZE MUTANT 57 Abstract 57 Introduction 58 Materials and Methods 61 Results 63 Discussion 68 6 AN ORGAN-SPECIFIC INVERTASE DEFICIENCY IN THE PRIMARY ROOT OF AN INBRED MAIZE LINE 75 Abstract 75 Introduction 76 Materials and Methods 78 Results 80 Discussion 85 7 SUMMARY AND CONCLUSIONS 91 LITERATURE CITED 94 BIOGRAPHICAL SKETCH 113 iv

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LIST OF TABLES Table 3-1 Effect of enzyme protectants on activity of sucrose synthase from maize root tips assayed five minutes after extraction 36 Table 4-1 Total sucrose synthase activity in wildtype maize root tips incubated in a range of glucose concentrations for 24 hour 53 Table 5-1 Sucrose synthase activity in mutant maize (W22:s/7)) root tips incubated in media containing a range of glucose concentrations for 24 hours 66 Table 5-2 Sucrose synthase activity in mutant maize (W22:sfr7) root tips incubated in media containing 0 or 2.0% glucose for various time periods 70 Table 6-1 Soluble and insoluble acid invertase activity in sequential 2 mm segments of primary roots of 5to 6-day-old seedlings from 1 hybrid and 2 inbred lines of maize 81 Table 6-2 Soluble acid invertase and sucrose synthase activity in 0.5 cm apices of primary and adventitious roots of 5to 6-day-old seedlings from 1 hybrid and 2 inbred lines of lea mays 83 Table 6-3 Soluble acid invertase activity in various tissues of Oh 43, an inbred line of lea mays 84 v

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LIST OF FIGURES Figure 3-1 Time course of in vitro decrease in sucrose synthase activity in maize and cotton roots 35 Figure 3-2 Time course of in vitro decrease in maize root sucrose synthase activity with and without the serine proteinase inhibitor, PMSF 37 Figure 3-3 Time course of in vitro decrease in maize root sucrose synthase activity in the presence and absence of either Pi (10 mM) or casein 2%w:v) 39 Figure 3-4 Denaturing (A) and native (B,C,D) protein gel blot analysis of maize root sucrose synthase at various times after extraction 40 Figure 3-5 Time course of in vitro decrease in maize root sucrose synthase activity from lines with homo(W22:s/?7) and heterotetrameric (NK 508) forms of this enzyme 43 Figure 5-1 RNA gel blot analysis of Sus1 expression in maize roots incubated in a range of glucose concentrations for 24 hours 64 Figure 5-2 Protein gel blot of Sus1 encoded sucrose synthase from maize roots incubated in a range of glucose concentrations for 24 hours . . 65 Figure 5-3 RNA gel blot analysis of Sus1 mRNA expression in maize roots incubated in 0 and 2.0% glucose for various time periods 67 Figure 5-4 Protein gel blot of Sus1 encoded sucrose synthase from maize roots incubated in 0 and 2.0% glucose for various time periods .... 69 Figure 6-1 Histochemical localization of invertase activity in free-hand, cross sections of root apices of 6-day-old maize seedlings 87 vi

PAGE 7

Abstract of Dissertation Presented to the Graduate School of the University of Florida in Partial Fulfillment of the Requirements for the Degree of Doctor of Philosophy SUCROSE SYNTHASE AND INVERTASE IN MAIZE ROOTS DIFFERING IN CARBOHYDRATE STATUS AND/OR GENETIC COMPOSITION By Edwin Ralph Duke August, 1991 Chairperson: Dr. Karen E. Koch Major Department: Horticultural Science The extent to which sucrose is transported in phloem of higher plant species necessitates its effective metabolism by non-photosynthetic cells. However, only two enzymes can catalyze sucrose breakdown in these instances, invertase and sucrose synthase (a reversible enzyme). Specific molecular, genetic and physiological factors affecting these enzymes were investigated in the root tips of maize. A radiometric assay was first developed for sucrose synthase to circumvent the rapid decline of sucrose synthase activity in vitro. Further, characterization of in vitro instability indicated that activity decline was not associated with any detectable proteolytic degradation, charge alteration, or subunit separation and that inhibition of activity by inorganic phosphate suggested possible phosphorylation of this enzyme. vii

PAGE 8

Message levels of genes encoding sucrose synthase isozymes in maize have been shown to respond to tissue carbohydrate status, thus the effects of such changes were examined at the level of enzyme activity. Total sucrose synthase activity from roots of wildtype plants showed little difference in extracts from root tips incubated for 24 h in a range of glucose levels. This activity, however, is the combined contribution of two isozymes whose genes are responding differentially to experimental conditions. The sh1 mutant of maize was used to study expression of the Sus1 gene for sucrose synthase in response to sugar availability because this mutant has only one gene (Sus1) for sucrose synthase and provides a system uncomplicated by the presence of the second isozyme (Sh1). Sus1 mRNA increased 5-fold when incubated for 24 h in 2.0% glucose compared to 0 or 0.2% glucose. Levels of Sus protein were slightly elevated with increasing sugar levels. Enzyme activity was elevated 2-fold under the same conditions. A study of time-course and treatment reversals showed that changes in mRNA or protein were not evident until 24 h and indicated that the response to carbohydrate level had been initiated within 16 h. Overall, enhanced expression of Sus 7 was evident at the mRNA, protein and enzyme levels. An organ-specific invertase deficiency affecting only the primary root system also was characterized in the Oh 43 maize inbred. Substantial acid invertase activity was evident in extracts of all tissues tested except the primary root system of Oh 43. This deficiency was also evident in lateral roots arising from the primary viii

PAGE 9

root but not in otherwise morphologically identical laterals from adventitious roots. In contrast, sucrose synthase was active in all roots and theoretically provided the only available avenue for sucrose degradation in primary root tips of Oh 43. ix

PAGE 10

CHAPTER 1 INTRODUCTION Sucrose metabolism is important to the majority of plant species because of the nearly ubiquitous role of this sugar in growth and development. Initial breakdown of sucrose can be catalyzed by either invertase or the reversible enzyme sucrose synthase. Gene responses to changes in carbohydrate availability have been reported for mammalian cells (Lin and Lee, 1 984) and yeasts and bacteria (Carlson, 1 987; Schuster, 1 989). Evidence has also been presented that the transcriptional activity of promoters of photosynthetic genes from maize protoplasts are repressed and coordinated by sugars (Sheen, 1990). Expression of the genes encoding the sucrose synthase isozymes also have been shown to respond to carbohydrate levels (Koch and McCarty, 1988; Koch et al., 1989). Northern blot analysis of mRNA from wildtype maize roots showed levels of one isozyme gene (Sh1) was up-regulated in response to carbohydrate depletion whereas the other (Sus) responded positively only when sugar availability was elevated. This sensitivity of genes for a key enzyme in sugar metabolism may prove to be an important control mechanism whereby plant cells are able to react to cellular nutritional conditions. The importance of invertase, the other enzyme of sucrose catabolism in plants, to root metabolism is unclear. Invertases are especially active in tissues 1

PAGE 11

undergoing rapid cell division such as shoot and root apices (Avigad, 1982). Recent evidence, however, indicates that although much hydrolysis is often observed, invertase activity may not be essential for sucrose uptake into either sugar cane stems (Lingle, 1989; Thorn and Maretzki, 1990) or maize kernels (Schmalstig and Hitz, 1987). Chapleo and Hall (1989a, b, and c) also have concluded that invertase was not essential to sucrose import into roots oiRicinus. Giaquinta and co-workers (1983) showed that sucrose entering the roots of maize via phloem does not have to pass through the extracellular space. Robbins (1 958) first reported that root tips of an inbred line of maize, Oh 43, were unable to retrieve exogenous sucrose. B. Burr (Brookhaven National Laboratory, personal communication) has indicated that the lack of retrieval might possibly be due to an invertase deficiency. The absence of invertase activity could have important implications for sucrose import, not only because of potential effects on the retrieval system, but also because sucrose breakdown in such an instance could theoretically proceed only via action of sucrose synthase. A mutant lacking functional invertase in its roots would also be useful, in combination with nulls for both sucrose synthase isozymes, to elucidate the individual roles for these enzymes. The purpose of the following research is to further elucidate the roles and regulation of the two sucrose metabolizing enzymes sucrose synthase and invertase in roots of maize. Specific objectives are to:

PAGE 12

3 1 . Develop a method of accurately assaying sucrose synthase activity to circumvent the rapid decline in activity exhibited upon extraction from maize root tips. 2. Clarify the possible causes of the rapid decline of sucrose synthase activity in extracts from maize root tips. 3. Determine the effects of varying carbohydrate conditions on total sucrose synthase activity in extracts from wildtype maize root tips. 4. Ascertain the effects of varying carbohydrate conditions on the Sus1 gene and its sucrose synthase gene product free from the confounding effects of Sh1. 5. Characterize the extent of invertase activity in various tissues of the Oh 43 inbred of maize, a putative invertase-deficient mutant.

PAGE 13

CHAPTER 2 REVIEW OF THE LITERATURE Sucrose Metabolism Sucrose is the major transported sugar in the majority of higher plant species. The major roles of sucrose in higher plants include its function as both a translocatable form of carbon and as a vacuolar storage compound (Hawker, 1985). It is a non-reducing sugar made up of a glucose (a-D-glucopyranose and fructose 03-D-fructofuranose) joined by an a-1 ,2 linkage. For metabolic utilization, it must be broken down into its component monosaccharides or their derivatives. Only two enzymatic systems for sucrose breakdown are known in higher plant tissues. Sucrose can be hydrolyzed by the action of invertase or cleaved by sucrose synthase working in the degradative direction. The free energy of sucrose hydrolysis is nearly equal to that of the y phosphoryl group of ATP (AG°= -7.0 kcal/mol and -7.3 kcal/mol, respectively) (Neufeld and Hassid, 1963). This is much greater than the free energy of most other glycosidic bonds (Avigad, 1982). Cleavage by sucrose synthase retains the bond energy in the a-glucosyl bond of UDP-glucose. In contrast, hydrolysis by invertase conserves none of the free energy in the bond. 4

PAGE 14

5 Morell and Copeland (1984, 1985) have investigated the enzymes of sucrose breakdown in soybean nodules and found that both sucrose synthase and alkaline invertase are present. They suggested that sucrose partitioning between the two enzymes could be determined by differences in their affinities for this substrate. The ^ of alkaline invertase for sucrose was 10 mM whereas that of sucrose synthase was 31 mM. Given the presence of both enzymes, they proposed the greater affinity of alkaline invertase for sucrose in this system could ensure that most of the sucrose entering the nodule would be converted to hexoses for further catabolism. At the same time some sucrose would be converted to UDP-glucose for subsequent synthesis of nucleotide sugars and polysaccharides. In contrast, Huber and Akazawa (1986) reported essentially an opposite situation in cultured sycamore cells, another system in which both sucrose synthase and neutral (alkaline) invertase were present at the same time and with similar activities. In these cells the sucrose synthase ^ for sucrose was substantially lower than that of neutral invertase (15 vs 65 mM). They proposed two pathways of sucrose cleavage, initiated by each of the enzymes, both pathways eventually leading to the production of triose-phosphates. Sucrose concentration was postulated to regulate carbon flow between the two pathways. Sucrose synthase had a lower K m for sucrose and would, therefore, be relatively more important under sucrose limiting conditions (Avigad, 1982). This pathway is more energy efficient and would be more beneficial to the cells when carbon supplies are limited. The work of Morell and Copeland's work (1984, 1985) and

PAGE 15

6 Huber and Akazawa (1986) both demonstrate the potential importance of simultaneous sucrose catabolism via two enzyme systems. Sucrose Synthase Sucrose synthase (EC 2.4.1.13, UDP-D-glucose: D-fructose 2-a-Dglucosyltransferase) is ubiquitous in higher plants (Keller et al., 1 988) and probably occurs in all types of tissues. However, this enzyme is found in greatest abundance in nonphotosynthetic tissues and in developing seeds (Echt and Chourey, 1985). In cell fractionation studies, sucrose synthase was shown to be associated with the soluble fraction (Nishimura and Beevers, 1 979; MacDonald and ap Rees, 1983). A cytosolic rather than vacuolar localization for sucrose synthase has been demonstrated in protoplasts isolated from Jerusalem artichoke (Keller et al., 1988). Although molecular weights as high as one million have been reported for sucrose synthase (Grimes et al., 1970), it is now generally concluded that, in its native state, sucrose synthase has a molecular weight of approximately 36 to 40 kD (Delmer, 1972a; Su and Preiss, 1978; Morell and Copeland, 1985; Moriguchi and Yamaki, 1988) and is composed of four identical subunits. The sucrose synthase subunit from maize has been found to have a molecular weight of 8.8 kD (Su and Preiss, 1978). However, unlike many other plant species, maize has two genes which encode sucrose synthase subunits (Chourey and Nelson, 1 976; Echt and Chourey, 1985). The two sucrose synthase subunits of maize, sh1 and sus, encoded by the Sh1 and Sus 7 genes, respectively, have similar enzyme kinetics,

PAGE 16

7 similar amino acid compositions and share limited structural homologies (Echt and Chourey, 1985). They do differ slightly in their electrophoretic movement during PAGE (Echt and Chourey, 1985). The two subunits are homologous enough to form heterotetrameric structures, apparent as five separate bands on native PAGE (Echt and Chourey, 1985). Spatial separations of these two sucrose synthase isozymes are sometimes apparent. Sh1 encoded protein is primarily located in the endosperm (Chourey and Nelson, 1976; Chen and Chourey, 1989), in the root (Springer et al., 1986; Chourey et al., 1986) and in etiolated shoots (Springer et al., 1986). The Sus1 encoded protein is found throughout the plant (Chourey, 1981 ; Echt and Chourey, 1985; Chourey et al., 1988). The distribution of both proteins is further distinguished under stress conditions (such as anaerobiosis) where tissue-specific localization in roots is readily apparent (Rowland et al., 1989). Roles of sucrose synthase Sucrose synthase was initially considered to be a sucrose synthesizing system in plants by Leloir and Cardini (1953, 1955). Sucrose synthase and sucrose phosphate synthase (EC 2.3.1.14, UDP-D-glucose: D-fructose-6phosphate 2-a-D-glucosyltransferase) are the two enzymes that catalyze the transglucosylation reaction from UDP-glucose to fructose and fructose-6phosphate, respectively.

PAGE 17

8 The necessity for two systems of sucrose synthesis was puzzling until it was determined that the sucrose synthase reaction is readily reversible (Cardini et al, 1955): Sucrose + UDP < > UDP-Glucose + Fructose This reversibility gave rise to the suggestion that sucrose synthase could make UDPG available for utilization as a glucosyl donor in starch synthesis (Turner and Turner, 1957). Other studies of sucrose synthase specificity and kinetics led Avigad and coworkers (Avigad, 1 964; Avigad et al., 1 964; Milner and Avigad, 1 964) to suggest that this enzyme functioned mainly in sucrose cleavage in storage tissues. Sucrose synthase activity typically is highest in tissues during periods of rapid growth and is often not accompanied by high invertase activity (Schaffer et al., 1987). Several factors lend credence to the view that the role of sucrose synthase is sucrose cleavage in importing cells. First, a substantial level of free fructose is required for sucrose synthase activity in the synthetic direction (K,,, ca. 2.0-2.5 mM [Avigad, 1982]). Levels of this sugar are low in healthy, intact leaves, but they are higher in storage tissues and roots, where most of the free fructose is in the vacuole or extracellular spaces (Avigad, 1982). Availability of UDPG is also likely to limit the synthetic reaction, because the cellular concentration is typically less than 0.4 mM (Murata, 1975). The ^ for UDPG ranges from 0.1 to 8.5 mM (average approximately 2.0 mM) (Avigad, 1982). In contrast, sucrose

PAGE 18

9 concentrations are generally elevated in importing areas. Therefore, substrate levels favor the cleavage reaction in the majority of instances. A second line of evidence for the degradative role of sucrose synthase in importing organs is provided by mutant maize lines lacking a functional sucrose synthase protein. Chourey and Nelson (1976) showed that a deletion of the Sh locus on chromosome 9 of maize (coding for sucrose synthase) led to a 90% deficiency of the respective protein in mutant vs wild-type kernels. Starch formation was also reduced in this line, giving rise to a "shrunken" seed. The association between this shrunken phenotype and a sucrose synthase deficiency was considered evidence that the critical reaction in vivo was that of sucrose cleavage, and that this was essential for conversion of photosynthetically produced sucrose for starch biosynthesis. The residual amount of starch deposited was attributed to the sucrose degrading activity of a second sucrose synthase encoded by another locus (Sus1). Further research also favors the cleavage role of sucrose synthase and implicates its involvement in starch deposition. Dale and Housley (1986), for example, found that developing wheat kernels with the greatest rates of growth and starch deposition had significantly greater sucrose synthase activities. A positive correlation between sucrose synthase activity and starch deposition was also reported in Pisum sativum by Edwards and ap Rees (1986a and b). They proposed that UDP-glucose formed during sucrose cleavage was converted to glucose-1 -phosphate by UDP-glucose pyrophosphorylase using pyrophosphate

PAGE 19

10 generated by PFK(PPj). Morrell and ap Rees (1986) have also suggested that much of the sucrose translocated to developing potato tubers is probably metabolized via the same pathway with the initial step catalyzed by sucrose synthase. Gibson and Shine (1983) have demonstrate that in the presence of inorganic phosphate, UDPG may be hydrolyzed to G-1-P and UDP by the action of UDPG phosphorylase. Salerno (1986) also has demonstrated the presence a highly nucleotide specific form of UDP-glucose phosphorylase in developing maize endosperms. The activity level of this enzyme followed closely the development of the grain and paralleled that of sucrose synthase. The presence of this enzyme links sucrose cleavage and starch formation via sucrose synthase. Finally, an additional line of evidence was presented by Cobb and Hannah (1988) that also indicated a primarily degradative function for sucrose synthase in importing organs. They showed that maize kernels from a line deficient in the Sh1 gene for sucrose synthase still had normal levels of sucrose and normal rates of sucrose synthesis when grown in culture with fructose as the carbon source. If sucrose synthesis had been proceeding via sucrose synthase in wild-type kernels, then the loss of ca. 90% of total sucrose synthase activity in kernels of the mutant line should have affected sucrose formation there. The authors concluded that Sh1 encoded sucrose synthase was not necessary for sucrose synthesis. Correlative data suggest that sucrose synthase activity is closely linked with sink strength. The supply and timing of sucrose for export seem to be closely related to the source of photosynthate (Fondy and Geiger, 1982; Servaites et al.,

PAGE 20

11 1 989) as well as the energy for phloem loading. However, once sucrose is loaded, its eventual fate does not appear to be under the control of the source leaf (Gifford and Evans, 1981) but rather is under the control of the importing sink (Wyse, 1986). Giaquinta (1979) found that young, immature roots of sugar beets had low levels of sucrose synthase, but the onset of rapid sucrose import for storage was accompanied by a significant increase in sucrose synthase activity. Similar correlations were also observed by Silvius and Snyder (1979) and Fieuw and Willenbrink (1987). In sugar beet roots sucrose uptake into parenchyma can proceed without prior hydrolysis in the apoplast or free space. Increases in sucrose synthase activity have also been observed during periods of sucrose import and/or accumulation in sweet melons (Schaffer et al., 1987), netted muskmelon (Lingle and Dunlap, 1987), eggplants (Claussen et al., 1985, 1986), rose flowers (Khayat and Zeslin, 1987), developing chick pea seeds (Setia and Malik, 1985), tomato (Yelle et al., 1988) and Solanum muricatum (Schaffer et al., 1989). Lingle (1987), however, found no correlation of sucrose synthase activity with sucrose concentration in sweet sorghum. Huber and Akazawa (1986) hypothesized that a primary role of sucrose synthase could be to feed glucose-1 -phosphate directly into glycolysis. Black and coworkers (Black et al., 1987; Sung et al., 1988; 1989; Xu et al., 1989) also support this hypothesis. This link to glycolysis also involves UDPG-pyrophosphorylase, which converts the UDPG formed by the action of sucrose synthase into G-1-P and UTP. The methods employed to deliver carbohydrates to their respective

PAGE 21

12 sinks vary from species to species (ap Rees, 1974, Hawker, 1985). However, sucrose breakdown generally appears to proceed via sucrose synthase in starch and sugar storage sinks (Sung et al., 1988). Sucrose synthase also may be associated with sink strength through its potential involvement in cell wall synthesis (Hendrix, 1990). Sucrose synthase activity predominates over that of invertase in rapidly expanding cotton ovules, for example where rapid elongation of epidermal hairs (cotton fibers) requires extensive cellulose formation. Hendrix (1 990) postulated that sucrose entering the seed coat in the developing cotton boll was cleaved via sucrose synthase, and subsequent carbohydrates went into the rapidly growing epidermal hairs. However, some of the sucrose breakdown products were converted to starch stored in the seed coats. Stepanenko and Morozova (1970) demonstrated cellulose biosynthesis from UDP-glucose in cotton. However, they did not indicate that the source of the UDPG might come from the action of sucrose synthase. Carpita and Delmer (1981) showed that the rate of synthesis and turnover of UDPglucose in developing cotton ovules was more than sufficient to account for rates of biosynthesis for cellulose, /3-1 ,3-glucan and sterylglucosides (all cell wall constituents). They found that UDPG levels increased dramatically just prior to the maximum rate of secondary wall cell synthesis and dropped precipitously at the time when cellulose synthesis ceased. Again, sucrose synthase activity was not measured, but could possibly be the source of the increased levels of UDPG. Sucrose synthase activity was measured in cultured cells of Catharanthus roseus

PAGE 22

13 (Amino et al., 1985) and found to be elevated during the G1 phase when the amount of total cell walls increased significantly. However, UDP-glucose pyrophosphorylase activity (also involved in the formation of UDPG) was greater than sucrose synthase activity at the G1 phase. The former was considered by these authors likely to make a more important contribution to the total UDPG formed. Chourey et al. (1991a) have reiterated the hypothesis that the resultant shrunken, starch-deficient endosperm of the sh1 maize mutant may be due to reduced cell wall deposition rather than any direct effect on the starch biosynthetic pathway. Regulation of Sucrose Synthase Sucrose synthase, a key enzyme in sucrose metabolism, is subject to a number of complex regulatory factors (Davies, 1974). This enzyme exhibits a wide specificity for the nucleoside base utilized in the reaction. Most enzymes of sugar nucleoside metabolism show a marked specificity for a particular base (Avigad, 1982). Sucrose synthase working in the synthetic direction has been shown to utilize UDPG, ADPG, TDPG, CDPG and GDPG as glucosyl group donors (Avigad, 1982). The K,,, for UDPG, however, is usually much less than for other NDPG's. Grimes et al. (1970) found that the K m for UDPG was approximately 0.2 mM while ADPG, TDPG, CDPG and GDPG had K m 's of 1 .8, 1 .7, 2.5 and 2.5 mM, respectively. They also found that with UDPG as the nucleoside sugar, the K m for fructose was reduced 10 fold below that apparent when ADPG was utilized. This change was

PAGE 23

14 attributed by Grimes et al. (1970) to possibly result from conformational changes in the enzyme. Reduced K^s for UDPG when compared to other NDPG's have also been observed for sucrose synthase from pea seedlings (Gabrielyan et al. p 1969), sweet potato root (Murata, 1971), potato tubers (Pollock and ap Rees, 1975), sweet corn seeds (de Fekete and Cardini, 1964) and soybean nodules (Morell and Copeland, 1985). In contrast, ADPG and TDPG were reportedly more efficient glucosyl donors than UDPG for sucrose synthase isolated from sorghum seeds (Sharma and Bhatia, 1980) and sugar beet roots (Avigad and Milner, 1966). No large differences in K^s for UDP and other nucleoside diphosphates generally are observed when the reverse reaction is analyzed. Delmer (1 972a and b) found minimal or no differences in K m 's for NDP's with sucrose synthase from mung bean seedlings. She did, however, find a large difference in the rates of sucrose cleavage with different NDP's. Maxima were observed when UDP was the substrate. The V max for UDP in relative terms was 100 compared to 28, 6, 3 and 3 for ADP, TDP, CDP and GDP, respectively. Similar K^/s for various NDP's have also been observed in sweet potato roots (Murata, 1971), potato tubers (Pollock and ap Rees, 1975), Jerusalem artichoke (Pontis et al., 1972) and sugar beet (Avigad and Milner, 1966). Sucrose synthase from sweet corn kernels does, however, exhibit a K,,, an order of magnitude greater for ADP than UDP (Su and Preiss, 1978; de Fekete and Cardini, 1964). Morell and Copeland (1985) also

PAGE 24

15 found that in soybean nodules, the K m of sucrose synthase for UDP (0.5 mM) was lower than that of ADP and CDP (0.13 and 1.1 mM, respectively). Delmer (1 972a and b) characterized regulation of purified Phaseolus aureus sucrose synthase and found a number of differences in the regulation of the synthetic and degradative reactions. NADP, iodoacetic acid, and gibberellic acid all stimulate sucrose degradation but inhibit sucrose synthesis. Pyrophosphate also enhanced the degradative activity, but only in the presence of MgCI 2 . In contrast, Pontis (1977) reported that P, inhibited the degradative reaction alone or in the presence of Mg 2+ . Delmer also tested the effects of intermediates in carbohydrate metabolism and found that G-1-P, G-6-P, F-6-P, F-1.6-BP, R-5-P, R1 ,5-BP, PEP and 3-PGA had little or no influence on the sucrose synthase reaction in either direction when present at 2 mM. However, de Fekete (1969) and Pontis (1977) both have reported that G-1-P, G-6-P and F-1.6-BP were inhibitory to the degradative reaction at 2-5 mM without affecting the synthetic reaction. ATP, ADP and AMP had no inhibitory effect on sucrose synthesis at 4 mM; however, the degradative reaction was inhibited 30% by ADP, and 50% by both ADP and AMP. /3-Phenylglucoside has also been shown to inhibit sucrose degradation almost completely and sucrose synthesis by 50% (Wolosiuk and Pontis, 1974b; Lowell, 1986). This has proven useful for distinguishing activities of sucrose phosphate synthase from sucrose synthase. Pontis and coworkers (Pontis et al., 1972) found that the divalent cations Mg 2+ , Mn 2+ , Ca 2+ and Ba 2+ at 5-10 mM activated sucrose synthase in the

PAGE 25

16 synthetic direction but inhibited the cleavage reaction. UDP was found to be a strong inhibitor of the synthetic reaction at 10 mM (70-80% inhibition), but the inhibition could be reversed by the addition of Mg 2+ (de Fekete and Cardini, 1964). UDP was also found to inhibit the degradative reaction as a competitive inhibitor for UDPG (Wolosiuk and Pontis, 1974a). Inhibition by other NDP's was very weak. UTP (4 mM) caused a slight inhibition of synthetic activity, but caused an 80% inhibition of the degradative reaction (Tsai, 1974). Echeverria and Humphreys (1985), however, found that UDP and UTP within the cytosolic range (< 4 mM) both had little or no effect on sucrose synthase in the synthetic direction. UDPG was able to inhibit the cleavage reaction by 13% at 10 mM, but tissue concentrations were generally below this level (Echeverria and Humphreys, 1985), with the effect on the synthetic reaction minimal. Wolosiuk and Pontis (1974a) found that UDPG could function as a competitive inhibitor for UDP in the sucrose synthase synthesis reaction. Carbohydrates also have been found to inhibit sucrose synthase activity in vitro. Fructose was found to function as a competitive inhibitor of sucrose in the cleavage reaction (Pridham et al., 1969; Doehlert, 1987). Sucrose had no inhibitory effects on sucrose synthase activity at saturating levels of fructose and UDPG (Echeverria and Humphreys, 1985), but glucose at 100 mM inhibited sucrose synthesis by 63%-70% and inhibited sucrose cleavage 86%-93%. Expression of sucrose synthase genes also respond to carbohydrate availability. Koch and McCarty (1988, 1990; Koch et al., 1989) have shown that levels of maize

PAGE 26

17 root Sh1 mRNA were elevated when sugar supplies were limited in culture. In contrast, levels of Sus1 mRNA were elevated in response to increasing glucose concentrations. They speculated that the effect of sugar levels on expression of specific genes could prove to be an important control mechanism whereby plant cells could react to cellular nutritional conditions. The Sh1 gene is also upregulated under anaerobic conditions (Springer et al., 1986); however, the carbohydrate response of the Sh1 gene appears to be distinct from its anaerobic regulation (Koch et al., 1989). There is some doubt as to whether anaerobic induction occurs at both the transcriptional and translational levels in maize (McElfresh and Chourey, 1988). Taliercio and Chourey (1989) hypothesized that the expression of anaerobically induced Sh1 transcripts are blocked at some step beyond polyribosomal loading. However, other researchers have shown that sucrose synthase in maize is anaerobically induced at the protein as well as the gene level (Freeling and Bennett, 1985; Springer et al., 1986). Anaerobic induction of sucrose synthase at both the gene and protein levels also has been demonstrated in rice (Ricard et al, 1991) and Echinochloa phyllopogon (Mujer et al., 1990). Invertases Invertases (E.C. 3.2.1.26, /3-D-fructofuranoside fructohydrolases) are widely distributed in the plant kingdom and catalyze the following reaction: Sucrose + H 2 0 > Glucose + Fructose

PAGE 27

18 Invertases are specific for the fructofuranose moiety of sucrose and work by hydrolyzing the glycosidic linkage between the bridge oxygen and the fructose residue (Sum et al., 1980). Up to five different forms of invertase have been reported in plants (Sasaki et al., 1971). However, these enzymes are generally divided into two main types based on the pH at which sucrose hydrolysis is most efficiently accomplished. Acid invertases have pH optima around 4.5 to 5.0; that of alkaline (or neutral invertase) is 7.0 to 7.5. Most invertases are glycoproteins. Arnold (1966) partially purified an acid invertase from grapes and found that it was approximately 25% carbohydrate. Faye and coworkers (Faye and Berjonneau, 1979; Faye et al., 1981) have shown a 7.7% carbohydrate content of acid invertase from radish seedlings, and date invertase has a carbohydrate content of 8.2% (Al-Bakir and Whitaker, 1978). Invertase preparations from barley (Prentice and Robbins, 1976), sugar cane (del Rosario and Santisopasri, 1977), potato tubers (Anderson and Ewing, 1978; Bracho and Whitaker, 1990b) and banana (Sum et al., 1980) were shown to bind strongly to concanavalin A, a phytagglutinin or lectin isolated from jack bean with a strong binding affinity for carbohydrates. In yeast and Neurospora the carbohydrate content of invertase has been estimated to range from 0 to 50 % (Metzenberg, 1963; Gascon et al., 1968; Holbein et al., 1976). Much of our more detailed knowledge on the molecular structure and mode of action of invertases comes from studies on fungal and yeast enzymes; however, considerable progress has been made in analyses of plant invertases.

PAGE 28

19 Roles of Invertase Elevated acid invertase activity is characteristic of plant tissues in which there is a need for hexoses produced from stored or recently transported sucrose (ap Rees, 1974; Avigad, 1982). Greater activities of invertase also correlate well with a low content of stored sucrose. In sugar beets, the onset of sucrose storage is accompanied by a decrease in invertase activity (Silvius and Snyder, 1979; Giaquinta, 1979). The same is true for carrot roots (Ricardo and ap Rees, 1970), melon (Hubbard et al., 1989; Lingle and Dunlap, 1987; Schaffer et aJ., 1987; McCollum et al., 1988), citrus (Kato and Kubota, 1978; Lowell, 1986) and Lycopersicon hirsutum (Miron and Schaffer, 1991). In these systems, invertase was very active prior to sucrose accumulation and dropped significantly upon maturity. Invertase activity is usually greatest in tissues that are at a rapid stage of growth and development (Weil and Rausch, 1990), particularly at the cell division stage (Masuda et al., 1988). Root apices, young leaves and stem internodes fall into this category. Mature leaves, functioning as sources of photosynthates, generally have low levels of apoplastic acid invertase (Dickinson et al., 1991). Transgenic tomato plants expressing yeast invertase in the apoplast of mature leaves had a striking repression of growth (Dickinson et al., 1991). The higher the level of invertase, the greater the inhibition. The general role of acid invertase, therefore, seems to be for the breakdown of sucrose where there is a marked need for hexose (ap Rees, 1974).

PAGE 29

20 Previously, the role of invertase in sucrose transfer was considered particularly important in plants such as sugar cane and maize where substantial sugar movement occurred through the cell wall space and was accompanied by action of an extracellular invertase (Glasziou and Gayler, 1 972; Hawker and Hatch, 1 965). Recent evidence, however, indicates that although much hydrolysis is often observed, invertase activity may not be essential for sucrose uptake into either sugar cane stems (Thorn and Maretzki, 1990; Lingle, 1989) or maize kernels (Schmalstig and Hitz, 1987). The role of apoplastic invertase in sucrose import into roots had previously been questioned by Chapleo and Hall (1989a) who concluded that although present, apoplastic root invertase did not have a direct role in sugar transport. However, substantial activity of invertase has been widely documented in roots of plants such as pea (Lyne and ap Rees, 1971), bean (Robinson and Brown, 1952), tomato (Chin and Weston, 1973), Ricinus (Chapleo and Hall, 1989a, b, and c), oat (Pressy and Avants, 1980), and maize (Hellebust and Forward, 1962; Chang and Bandurski, 1963). Specific tissue localizations also have been described. Peak activity for root invertase is generally 2-3 mm behind the apex and corresponds to the region of expansion and elongation in pea (Robinson and Brown, 1 952; Sexton and Sutcliffe, 1 969) and maize (Hellebust and Forward, 1962). In Ricinus roots, this activity predominates in the cortex (Chapleo and Hall 1989a). Although invertase may not have a direct role in sucrose import into roots, it still may be important to two major aspects of root biology. First, invertase is

PAGE 30

21 essential to mycorrhizal associations (Purves and Hadley, 1975). Maize (Gerdemann, 1964; Kothari et al., 1990) and 90% of other agriculturally important species form these beneficial symbioses under field conditions (Gerdemann, 1 968) . However, sucrose must be hydrolyzed for the fungal symbiont (Long et al., 1975), and invertase levels rise at sites of carbon transfer. It is not known whether this is host or fungal invertase. Another possible role for apoplastic invertase is in the regulation of the intercellular sucrose concentration. Regulation of the free-space sucrose concentration may be important in osmotic relations and in the control of tissue differentiation (ap Rees, 1974). Jeffs and Northcote (1966, 1967) have shown that phloem differentiation in cultures of Phaseolus vulgaris depended on the supply of sucrose; glucose or fructose would not substitute. Wright and Northcote (1972), however, have shown that phloem differentiation in cultures of Acer pseudoplatanus were equally responsive to glucose and sucrose. The results of Jeffs and Northcote show that in certain cases the regulation of the apoplastic sucrose content by acid invertase could be important in differentiation. A definitive role cannot be assigned to alkaline invertase at present. Studies with sugar cane (Hatch and Glasziou, 1963), carrot roots (Ricardo and ap Rees, 1970), pea roots (Lyne and ap Rees, 1971), melon (Lingle and Dunlap, 1987; McCollum et al., 1988), and Lycopersicon hirsutum (Miron and Schaffer, 1991) indicate an inverse relationship between alkaline and acid invertase and a more positive correlation between alkaline invertase and sucrose concentration. The

PAGE 31

22 maximum values for alkaline invertase activity observed to date are consistently less than those of acid invertase (Masuda et al., 1988). The possibility exists that alkaline invertase allows cells that store sucrose in their vacuoles to retain a capacity for breakdown of enough sucrose in the cytoplasm to meet respiratory and metabolic demands for hexoses (ap Rees, 1974). The capacity of a plant to produce two different invertases that are spatially separated may allow the plant cell to regulate sucrose storage independent from sucrose breakdown. Regulation of Invertase Acid invertases are generally found in the apoplast and vacuoles of plant tissues. Washed preparations of cell walls contain a large proportion of a plant's acid invertase (Little and Edelman, 1973). A portion of the acid invertase can be extracted from the cell wall during grinding, but at least some of the enzyme is considered to be attached to the cell wall in vivo (Edelman and Hall, 1965). The major determinant of how much acid invertase remains bound during extraction is the pH of the buffer used (ap Rees, 1974). Buffers with acidic pH leave most of the activity in the cell wall fraction, whereas neutral or alkaline buffers release the majority into the soluble fraction. Early evidence suggested that the soluble and insoluble acid invertases were not simply different forms of the same enzyme. The pH optima and the K,,, of bound acid invertase of mature (Hawker and Hatch, 1 965) and immature (Hatch et al., 1963) sugar cane storage tissue differed from those of the soluble fractions. Association with the cell wall may change an enzyme's properties (ap Rees, 1974);

PAGE 32

23 however the differences in values, especially those of mature tissues, were considered unlikely to be wholly artifactual. Distinguishing between forms of invertase is further complicated by information obtained from yeast. One gene in yeast, SUC2, has been shown to encode the two forms of invertase in yeast, secreted and intracellular, via two differentially regulated mRNAs (Carlson and Botstein, 1982). The K^s of acid invertase for sucrose generally range from 2 to 13 mM. Sucrose is the primary substrate for acid invertase but raffinose also is hydrolyzed, though at a slower rate (10% to 50% the rate of sucrose) (Avigad, 1982). Acid invertase from sugar-cane leaves was inhibited competitively by fructose (Kj 32 mM) and noncompetitively by glucose (Kj 37 mM) (Sampietro et al., 1980). Acid invertases have been partially purified from a number of tissues with apparent molecular weights ranging from 2.8 x 10 4 to 2.2 x 10 5 (Roberts, 1973; Ricardo, 1974; Kato and Kubota, 1978; Masuda and Sugawara, 1980; Sum et al., 1980; Faye et al., 1981). The SUC2 gene of Saccharomyces, encoding invertase, has been shown to be modulated by glucose levels (Carlson et al., 1987). Sucrose or raffinose, substrates of the yeast invertase, have no such effect. Kaufman et al. (1 973) found that acid invertase activity rises in Avena stem segments incubated in a sucrosecontaining medium. The response had a lag time of 10-12 hours, suggesting a change in protein levels. Fructose in the incubation medium resulted in a similar response, but glucose caused no change in invertase activity.

PAGE 33

24 A naturally occurring acid invertase inhibitor has been detected in a number of plant tissues including beet roots (Burakhanova et al., 1987), potato roots (Pressy, 1967, 1968; Bracho and Whitaker, 1990a and b), maize endosperm (Jaynes and Nelson, 1971), pea pollen (Malik and Sood, 1976) and Ipomea petals (Winkenbach and Matile, 1970). In potato the inhibitor was characterized as a small protein, binding irreversibly to acid invertase (Pressy, 1967; Anderson and Ewing, 1978). Pressy (1967) found that the binding of the potato inhibitor to invertase had a pH optimum of 4.5 (Pressy, 1967), and the enzyme-inhibitor complex could be partially disassociated by low pH or high Mg 2+ concentrations. In contrast, Bracho and Whitaker (1990a) found no effect of pH on inhibitor binding. Sucrose at 2 mM could inhibit binding, but would not dissociate a complex already formed (Pressy, 1 967). Neither glucose nor fructose had a similar effect. Matsushita and Uritani (1 974) noticed a marked increase of acid invertase activity resulted from wounding of sweet potato roots, but alkaline invertase activity did not change under similar conditions. They also isolated a heat-stable protein component with a molecular weight of approximately 19.5 kD, that fluctuated during the incubation period after the wounding (Matsushita and Uritani, 1976). They found that this putative inhibitor declined with a concomitant rise in invertase activity early in the incubation, but increased in later stages when invertase activity declined (Matsushita and Uritani, 1977). Pressy (1967, 1968) and Matsushita and Uritani (1 977) have suggested that the increase in invertase activity caused by cold treatment or by wounding could be explained by a decrease in binding of the

PAGE 34

25 inhibitor. Bracho and Whitaker (1 990b) also found a positive correlation between levels of inhibitor and invertase. The possibility therefore exists that this interaction plays a regulatory role in sucrose breakdown (Akazawa and Okamoto, 1980; Avigad, 1982). Alkaline invertase is generally considered to be cytoplasmic. It is only recovered from the soluble fraction of homogenates and has a pH optimum near neutral. Both findings support its internal localization. K,^ values of alkaline invertase for sucrose are slightly higher than for acid invertase, generally 9 to 25 mM. Alkaline invertase hydrolyzes raffinose very poorly (< 7% of the rate of sucrose breakdown). Morell and Copeland (1984) found that stachyose (0.1 M) also was hydrolyzed by alkaline invertase but much less efficiently than sucrose (1 .5% of the rate of sucrose). Both raffinose and stachyose are polysaccharides containing a fructose moiety. Morell and Copeland (1984) also found that cellobiose, gentiobiose, maltose, turanose, lactose, melezitose, trehalose, a-methylD-glucopyranoside and 0-methyl-D-glucopyranoside (all at 0.1 M) were resistant to degradative action by alkaline invertase. None of these sugars contain a fructose moiety, further confirming the specificity of invertase for the fructofuranose moiety of sucrose. Alkaline invertase from potato tubers was inhibited only slightly by glucose (Matsushita and Uritani, 1974); glucose-6-phosphate also had a slight inhibitory effect. Fructose (15 mM) competitively inhibited soybean nodule alkaline invertase by 50% (Morell and Copeland, 1984); glucose (5 mM) inhibited activity by 7%.

PAGE 35

26 Morell and Copeland (1984) also found that the metabolites ATP, ADP, UDP, ADPglucose, UDP-glucose, glucose-1 -phosphate, glucose-6-phosphate, and fructose-6phosphate (all at 5 mM) had no inhibitory effects. They also tested the effects of various chloride salts on alkaline invertase activity and found that Na + , K + or NH 4 + at 50 mM had no effects; however, CaCI 2 (10 mM) and MgCI 2 (10 mM) each inhibited activity by 25%. The anions citrate and inorganic phosphate have been shown to stimulate alkaline invertase from Lupinus luteus nodules (Kidby, 1966); however, Morell and Copeland (1984) found no effect on activity of soybean nodule alkaline invertase. They did find, though, that Tris buffer was a noncompetitive inhibitor of soybean nodule alkaline invertase activity; a 0.7 mM buffer concentration inhibited activity by 50%. Use of Mutants in Physiological Research Despite the fact that all mutations have effects on the biochemistry and physiology of the plant, only a small number have been investigated physiologically (Vose, 1981). Advances in knowledge about the molecular bases of cell processes in eukaryotic and prokaryotic microorganisms have been achieved with an array of mutant lines, often induced, that modify or block steps in the processes under study (Nilan et al., 1981). Many mutants will, theoretically, differ in only a single major physiological character. The use of mutants is growing in comparative physiological studies because the alternative is comparison of contrasting genotypes that quite possibly may be altered in undefined characters different from the one of interest.

PAGE 36

27 The maize plant (Zea mays L.) has been particularly useful in genetic and cytogenetic studies because of the number of mutants available (Neuffer et al., 1968). Many of the mutants also have proven useful for physiological research. The shrunken-1 mutant of maize was first described by Chourey and Nelson (1976). Less than 10% of the normal sucrose synthase activity in wild-type endosperm was observed. This reduced activity results in a "shrunken" phenotype in the dry kernel. The sh1 mutant has proven useful in elucidation of the role of sucrose synthase in starch formation. The residual activity of sucrose synthase present in this sh1 mutant was attributed to the presence of another isozyme encoded by a second gene (Chourey and Nelson, 1976). This second gene, Sus1, has been mapped to the same chromosome as Sh1 (chromosome 9) but is located 32 map units away (McCarty et al., 1986; Gupta et al., 1988). Sh1 and Sus1 encode similar proteins. Sucrose synthase is a tetramer in its native form (Su and Preiss, 1978), and the two isozymes encoded by Sh1 and Sus1 are able to form heterotetrameric forms of the native protein (Echt and Chourey, 1985). Sh1 has been shown to be responsive to anaerobic conditions, with transcript levels increasing 1 0 to 20 times in shoot and root tissue respectively compared to aerobic controls (Springer et al., 1986). However, Sus1 exhibits little response to anaerobic stress and seems to be expressed at a relative constant in all tissues (McCarty et al., 1986). Rowland et al (1989), however, found that Sus1 did show a slight response to anaerobic conditions, decreasing slightly in the lower root, primarily in the pith, root tip and root cap. A maize mutant lacking the

PAGE 37

28 Sus1 gene has been described (Chourey et al. p 1988), but, unlike the Sh1 mutant, the Sus1 mutant does not have any detectable phenotypic abnormality. A mutation lacking detectable levels of both sucrose synthase isozymes also has been described (Chourey, 1988), but its existence is puzzling considering the expected lethality of a complete sucrose synthase deficiency.

PAGE 38

CHAPTER 3 INSTABILITY OF SUCROSE SYNTHASE FROM ROOT TIPS: CHARACTERIZATION AND STABILIZATION Abstract Instability of sucrose synthase from root tips was characterized in maize and an assay developed to circumvent the rapid decline of activity in vitro (35 and 100% activity loss in 20 min for maize and cotton, respectively). Initially 14 Csucrose cleavage was quantified by recovery of 14 C-UDPG on DEAE ion exchange paper (Delmer, 1972; Su and Preiss, 1978). Subsequently, modifications were made which resulted in increased accuracy, reduced tissue volume required and reduced extraction/assay period. Phenolic protectants did not reduce the activity loss over time. Specific inhibitors for the four classes of proteinases were also tested; only PMSF increased enzyme activity, but did not completely prevent its loss over time. Stabilization and additional elevation of activity were achieved by adding casein. However, western blot analysis indicated that activity decline was not associated with any detectable proteolytic degradation, charge alteration, or subunit separation. In addition, inclusion of 10 mM P, in the extraction medium rapidly reduced activity, indicating the possible involvement of phosphorylation or nucleotide effects. 29

PAGE 39

30 Introduction Measurement of the maximum catalytic activities of enzymes in plant tissues can make important contributions to the understanding of metabolic pathways and their mechanisms of control (ap Rees, 1974). Currently available methods of assaying sucrose synthase have proven ineffective for many tissues, particularly those of roots (Duke et al., unpublished data; Lingle, USDA/ARS, Westlaco, TX, personal communication). A precipitous loss of activity follows tissue extraction from root tips of maize and other species (D.L Hendrix, USDA/ARS, Phoenix, AZ, personal communication). Chan et al. (1990) reported that sucrose synthase activity in roots of rice was detectable in only one stage of growth. However, sucrose synthase protein was present in root tissue at all stages of growth, exceeding that in grain when grain activity was highest among tissues sampled. This report addresses the basis of this instability in maize roots and describes a rapid radiometric assay for sucrose synthase which circumvents this problem and allows assay of small samples. Extraction and assay were optimized for substrate concentration, pH, assay length, and inclusion (or exclusion) of various antioxidants and proteinase inhibitors. The procedure has proven effective for a range of tissues and species examined and provides an accurate measurement of activity, particularly where enzyme stability may be limiting.

PAGE 40

Materials and Methods 31 Plant Material Maize seed (Zea mays L, NK 508, VJ22:sh1) were primed for 6 days at 10 C with a water potential of -1 .0 MPa (adjusted with PEG 8,000) with 2 g I" 1 captan (Bodsworth and Bewley, 1981). At the end of 6 days, the seeds were rinsed free of PEG, given a 20 min rinse in 1 .05% (v/v) sodium hypochlorite and again rinsed in water for 20-30 min. Seeds were germinated in the dark at 1 8 C on Whatman 3mm filter paper. Moisture level was kept constant throughout. At the end of 7 days, 1 cm primary root tips were excised under a sterile transfer hood. Cotton (Gossypium hirsutum L, Coker 100) root tips were obtained from Dr. D.L Hendrix (Western Cotton Research Laboratory, USDA/ARS Phoenix, AZ). One cm root tips were excised from 5to 6-day old seedlings and quick frozen in liquid N 2 . Purification of 14 C-Sucrose Trace amounts of phosphorylated sugars are common impurities in commercial 14 C-sucrose, and can reduce accuracy of the assay. These were removed by descending paper chromatography of commercially obtained 14 Csucrose in ethanol (NEN, Boston MA) using DEAE cellulose paper. The majority of anion-free sucrose was concentrated into the first 2 to 3 drops eluted from the V-shaped tip of the DEAE paper strip. No impurities were detected using HPLC analysis (data not shown). Molarity and specific activity of purified 14 C-sucrose

PAGE 41

32 were subsequently adjusted to 1 M and ca. 0.11 /xCi per pi. Two and one-half /xl (ca. 0.27 /xCi) were used in each reaction. Tissue Extraction Weighed tissue (1 00-200 mg) was frozen and ground to a powder in liquid N 2 with a mortar and pestle. The frozen powder was transferred to another mortar containing ice-cold extraction buffer (200 mM HEPES buffer [pH 7.5], with 1 mM DTT, 5 mM MgCI, 1 mM EGTA, 20 mM sodium ascorbate, 1 mM PMSF and 10% [w/w] PVPP) and ground briefly in this medium. One ml of grinding buffer was used for every 100 mg tissue fresh weight. Cysteine (10 mM) was initially but was omitted to prevent non-specific binding of radiolabel to DEAE cellulose paper. Two-hundred n\ of extract were placed on each of 4 to 8 spun columns packed with Sephadex G 50-80 hydrated with extraction buffer. Columns were centrifuged for 1 min at 800 x g. Eluent from each column was pooled with others from the same tissue sample before assay. Ratio of sample to bed volume was maximized at 1 :5 (v:v) by HPLC detection of soluble sugar presence in eluent (Yelle, 1991). Enzyme Assay Cleavage of 14 C-sucrose by sucrose synthase was assayed in a 50 /xl volume consisting of 20 n\ extract, 80 mM Mes (pH 5.5), 5 mM NaF, 100 mM 14 Csucrose and 5 mM UDP. Reactions proceed for 5 minutes at 30 C and were terminated by adding 50 /xl of Tris (pH 8.7) and boiling for 1 min. Controls

PAGE 42

33 contained all assay components except UDP. The assay was optimized for pH, linearity with time and protein concentration (data not shown). Product Determination The entire reaction volume was blotted onto a small disk of DEAE ionexchange paper (2.4 cm diameter) and dried completely before rinsing. Each disk was rinsed separately, first in 40 ml of H 2 0 at 175 rpm on a rotary shaker for 2 hours, again for an additional 2 hours and finally rinsed in a gentle stream of Dl water for 30 sec. Remaining radiolabel was quantified and compared to total amount of the 14 C-sucrose substrate utilized to determine extent of sucrose cleavage. Protein Gel Blots Subsamples from protein extracts to be used for enzyme assays were separated on native PAGE using the system of Laemilli (1970) with (denaturing) or without (native) SDS. Polyacrylamide concentrations of the stacking and separating gels were 2.5% and 5%, respectively. Proteins were resolved at 4 C by applying 15 V for 9 h, then 125 V for 11 h (constant current and temperature (4 C). Each lane was loaded with 2 /xg of total protein. Proteins were electroblotted to nitrocellulose membranes and probed with polyclonal antibodies following the procedure of Towbin et al. (1979). Sucrose synthase antisera, obtained from D.R. McCarty, was generated in rabbits using protein purified from maize kernels (W64 x 182E) 22 days after pollination. Antisera was diluted 1 :1000

PAGE 43

34 and cross reacted strongly to both the Sh1 and Sus1 gene products where such were present. Results Activity of sucrose synthase from maize and cotton root tips declined rapidly after extraction (Figure 3-1). The greatest decrease in activity occurred between 10 and 15 minutes after extraction from both species. Little or no activity was observed after 4 hours (data not shown). After extraction, extracts were maintained at 0 C until used in the radiometric assay. A wide range of enzyme protectants were examined. No improvement in activity was observed when the polyphenol protectants PVP-40, PEG 20,000 and BSA were utilized (Table 3-1 ). PVPP, also a phenol absorbent, was utilized in each extraction. In addition, four classes of proteinase inhibitors were tested for their effect on stability of sucrose synthase activity. Addition of leupeptin (1 mM) slightly decreased initial activity (Table 3-1), and pepstatin-A (1 mM) had no effect (Table 3-1). Phenylmethylsulfonyl fluoride (PMSF) (1 mM), a serine proteinase inhibitor, had a substantial positive effect, as did casein (2% w:v), potentially a non-specific proteinase inhibitor. Further characterization of activity change in the presence of PMSF showed that stabilization was not effective in the first 20 min following extraction (Fig. 3-2). Although total activity prior to this time was elevated by addition of PMSF, a linear decrease was not prevented from occurring.

PAGE 44

35 2 Time after extraction (min) Figure 3-1 . Time course of in vitro decrease in sucrose synthase activity in maize and cotton roots. Bars represent ± SE, n=3.

PAGE 45

36 Table 3-1 . Effect of enzyme protectants on activity of sucrose synthase from maize root tips assayed five minutes after extraction. Note: Each protectant was included in the buffer used for extraction and equilibration of desalting columns. PVP-40, PEG-20,000 and BSA were utilized to protect against phenolic compounds; PVPP was included in each extraction. Representative inhibitors of proteinase classes were: pepstatin A (aspartic), leupeptin (cysteine), EGTA (metallo) included in each extraction, caproic acid (serine) and PMSF (serine). Casein was included as a general, non-specific proteinase inhibitor. Protectant (concentration) enhancement of control PVP-40 (5 %) PEG-20,000 (2% w:v) BSA (2% w:v) Caproic acid (2 mM) Pepstatin A (1 mM) Leupeptin (1 mM) PMSF (1 mM) Casein (2% w:v) % -6 +6 +6 0 -5 0 +46 + 15

PAGE 46

Figure 3-2. Time course of in vitro decrease in maize sucrose synthase activity with and without the serine proteinase inhibitor, PMSF. No other class-specific proteinase inhibitors tested affected initial sucrose synthase activity. PMSF (1 mM) was used in extraction buffer and equilibration of desalting columns. Bars represent ± SE, n=3.

PAGE 47

38 Addition of casein increased initial enzyme activity compared to controls (Table 3-1 ) and improved stabilization of sucrose synthase activity with time (Figure 3-3A & B). In contrast to added protectants, inorganic phosphate (10 mM), a protein regulator through its role in reversible phosphorylation (Bennett, 1984), added to the extraction buffer decreased initial activity of sucrose synthase by ca. 40% (Figure 3-3A & B). Despite loss of activity in vitro and positive responses to apparent protectants against protease activity, proteolytic degradation of sucrose synthase from maize root tips was not detectable via either denaturing or native (Figure 3-4A & B, respectively) western-blot analysis at various times after enzyme extraction. Further, no change in charge or separation of subunits in native tetramers was evident. Nor was any change evident with time from samples extracted with added casein or phosphate (Figure 3-4C & D, respectively). However, changes in enzyme activity do not necessarily result in changes in electrophoretic mobility. Walker and Huber (1989) demonstrated that activation of sucrose phosphate synthase by light or mannose (a Pj sequestering sugar) did not affect immunoprecipitation or mobility of subunit mobility during SDS-PAGE. The possible involvement of tetramer stability in the loss of activity in vitro was further examined by comparison of the extracts from sh1 (containing only homotetramers of Sus1 encoded sucrose synthase) and Sh1 (containing both heteroand homotetramers of sucrose synthase). Endosperm tissue contains

PAGE 48

39 2.5 ^2.0 .c c1.5 'o 2 1.0 Q_ OXD.5 o i ° "§2.5 CO CD ^ ^ o2.0 CD C/) , _ o1.5 o 5 1.0 0.5 0 B + Casein + Casein + Pi 0 10 20 30 40 50 60 Time after extraction (min) 70 Figure 3-3. Time course of in vitro decrease in maize root sucrose synthase activity in the presence and absence of either P t (10 mM) or casein (2% w:v).

PAGE 49

40 Time after Extraction (min) 4 10 20 60 A 5 15 20 60 b 0BBH c mmmo ° n mi ft Figure 3-4. Denaturing (A) and native (B.C.D) protein gel blot analysis of maize root sucrose synthase at various times after extraction. Sub-samples were removed at designated intervals during incubation at 4 C. Proteins were separated by polyacrylamide gel electrophoresis with (A) or without (B.C.D) SDS and sucrose synthase resolved by probing with a polyclonal antibody raised against protein products of both the Sh1 and Sus genes. The five bands visible in the native gel blot have been described as corresponding to homoand heterotetrameric forms of sucrose synthase composed entirely of products from the Sh1 gene (uppermost band), the Sus gene (lowermost band) and combinations of the two (middle three bands). Extracts for denaturing (A) and native (B) blots were extracted with buffer only. Casein (2% w:v) (C) and Pi (10 mM) (D) were tested as protectants of enzyme stability.

PAGE 50

41 tetramers composed only of Sh1 encoded protomers (Chourey et al., 1986; Heinlein and Starlinger, 1989; Rowland and Chourey, 1990), and sucrose synthase activity is stable during extraction and dialysis procedures (Chourey and Nelson, 1 976; Echt and Chourey, 1 985). In extracts from root tissue, the five bands shown by western blot represent the possible combinations of monomers of the two separate isozymes (Sh1 and Sus) to form the native tetrameric structure (Echt and Chourey, 1985). Heterotetramers could theoretically be more unstable than homotetramers since, although very similar, the two subunits are not identical (Echt and Chourey, 1985). Su and Preiss (1978) found that sucrose synthase tended to polymerize to an inactive polymeric after extraction. Results indicated that formation of the native enzyme from two different isozymes was not a contributing factor in loss of enzyme activity over time (Figure 3-5A & B). Despite differences in the absolute values of sucrose synthase activity from sh1 vs. Sh1, the percent decline in activity was similar for both genotypes. Discussion Sucrose synthase activity was stabilized in vitro and an assay developed which enabled accurate measurement of enzyme action in root tips. The described assay allows rapid product recovery in instances where activity is otherwise unstable in vitro, and increases sensitivity to the extent that sample volumes as small as 1 00 to 200 /xg can be used. Sucrose synthase has previously been assayed in both synthetic and cleavage directions (Avigad and Milner, 1966; Grimes et al., 1970; Pontis et al.,

PAGE 51

c 1 f co o co ~| co -c E OJ c H(DC 6 « 10 CO CD ^ s S > o Q« ^ g CD c CO 2 c E co 2 1 Q t« > CO fi « CO c CD LU c3 W -C +1 c >> to c CD 10 0 CD CO O o D W ; +10 O CQ o eg E CO CD o CO c Q CD -*— ' CO 0 CD N 'co E CD E | c 0 — to CD 1c to 53 CO *r CD O O £ CD a CD c CD cn CD -C E M " c o is ^ « CO o c » O Z So 5 S CO CD .£= E I i2 in I ' CD CO si a> CO CD °J CD ~J 81 CO ~* to O co CD o -a CD =3 C CD C O) D) o <5
PAGE 53

44 1970; Salerno et al., 1979; Keller et al., 1988; Lowell et al., 1989). Measurements of the synthetic reaction have been based on quantification of either sucrose or UDP production. Sucrose levels can be determined indirectly by using invertase for full conversion to hexoses and measuring glucose colorimetrically (Avigad and Milner, 1966). The latter method, however, is susceptible to interference by substances in the crude enzyme extracts of many plants (Pontis, 1977). It has also been possible to measure 14 C-sucrose formed from UDP14 C-glucose by separating labeled product from substrate with anionic resins (Salerno et al., 1 979), paper electrophoresis (Grimes et al., 1970) or paper chromatography (Pontis, 1970). Such radioactive assays have proven useful in systems where colorimetric methods have been problematic (Pontis, 1977). In addition, UDP production can be determined spectrophotometrically by coupling its formation to the pyruvate kinase-lactate dehydrogenase reaction and measuring the decrease in absorbance due to oxidation of NADH (Avigad, 1964; Avigad and Milner, 1966; Lowell et al., 1989). Procedures for assaying the cleavage reaction are based on determination of fructose or UDP-glucose formation. Fructose can be measured colorimetrically by the Nelson reducing sugar assay (1 944), or spectrophotometrically by coupling hexokinase, phosphoglucose isomerase and glucose 6-phosphate dehydrogenase for production of NADPH (Avigad, 1964; Keller et al., 1988). Also, UDP-glucose formation can be estimated by coupling its appearance to NAD reduction by UDPdehydrogenase (Avigad, 1964; Lowell, 1986; Lowell et al., 1989). Degradative

PAGE 54

45 action of sucrose synthase can also be coupled to that of UDPglucopyrophosphorylase (Xu et al., 1986; Sung et al., 1989). Radioactive assays of sucrose synthase in the cleavage direction measure the incorporation of 14 C-glucose into UDP-glucose from 14 C-sucrose (Delmer, 1 972; Su and Preiss, 1978). These procedures are among the most sensitive of assays for sucrose synthase (Avigad, 1982). The sugar nucleotide formed can be separated from the excess 14 C-sucrose by paper chromatography (Wolosiuk and Pontis, 1974a) or by ion exchange paper (Delmer, 1972a and b; Su and Preiss, 1978). The current procedure utilizes the sensitivity of a radiometric assay along with reduced time from extraction to assay termination and results in a method suitable for time-labile extracts from small tissue samples. Instability of sucrose synthase was further characterized using this sensitive method in an attempt to better define factors affecting the activity of this key enzyme in vitro. Sucrose synthase is a sulfhydryl enzyme and is sensitive to inhibition by phenolics and oxidized polyphenols (Pontis, 1977). Typical effectors of activity reduction examined during the present study showed that phenolic compounds did not appear to be the primary cause of the sucrose synthase instability observed. No phelolic protectant was able to preserve sucrose synthase activity over time. Sucrose synthase has been shown to be sensitive to serine proteinases (Wolosiuk and Pontis, 1974b). In their study, trypsin caused a 70% decline in the degradative reaction and a 30% reduction in the synthetic reaction after a 15

PAGE 55

46 minute incubation. However, chymotrypsin, also a serine proteinase, and papain, a cysteine proteinase, had no effect. In the current study, PMSF, a serine proteinase inhibitor, gave an increase in initial measurements but not did not prevent the observed short-term loss of activity with time. Echt and Chourey (1985) observed that PMSF did not stop the loss of activity of sucrose synthase from maize endosperm during long term storage. No other specific proteinase inhibitor affected stability of the enzyme from maize root tips. In addition to the specific proteinase inhibitors, casein and BSA were included in some extractions. Due to casein's complex composition and random structure, it undergoes proteolysis with all the known proteolytic enzymes (Reimerdes and Klostermeyer, 1976). Casein increased initial measurements and stabilized activity with time (Figure 3-3A & B). BSA, however, was much less effective. Casein has also been found to preserve the longevity of sucrose synthase extracted from sugar cane (S. Lingle, USDA-ARS, Westlaco, TX, personal communication). Casein (0.75-3.0%) has also been shown to effectively stabilize and increase the initial activity measurements of sucrose phosphate synthase (Raghuveer and Sicher, 1987). Addition of casein is not always feasible, however, especially in instances where accurate quantification of total tissue protein is important. Another possibility for the regulation of sucrose synthase is through phosphorylation. Many enzymes undergo reversible phosphorylation as a means regulating activity (Bennett, 1984). Increased inorganic phosphate levels added to buffers used for enzymatic extraction provide a substrate for protein kinase

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47 activity (Bennett, 1984). Sucrose phosphate synthase (SPS), an enzyme of carbohydrate metabolism, has been shown to be regulated in this manner (Doehlert and Huber, 1983; Walker and Huber, 1989; Huber et al., 1989a). Increases in extractable SPS activity are noted after illumination or inclusion of mannose or glucosamine (phosphate sequestering agents) in darkness (Huber et al., 1989b). Inorganic phosphate (5-10 mM) was found to be a potent inhibitor of SPS (Amir and Preiss, 1982), with the inhibition becoming more sensitive in the presence of Mg 2+ . Sucrose synthase has also been shown to have exhibit diurnal fluctuations in activity (as does SPS) (Hendrix and Huber, 1986; Vassey, 1989) and be affected by Pj. Pontis (1977) reported that Pj (2-5 mM) inhibited the degradative reaction of sucrose synthase alone or in the presence of Mg 2+ . Delmer (1972a), however, found that 2 mM Pj had no effect on the rates of either the forward or reverse reactions. Sucrose synthase extracted from maize roots has also been shown to be phosphorylated in vitro (Xu and Koch, University of FL, unpublished data). In the current study, no effect of added phosphate on protein stability was noted (Figure 3-4D). However, initial sucrose synthase activity measurements were less than controls, and a loss of activity with time was observed (Figure 3-3A & B). The possibility existed that disassociation of subunits from tetramers may have effected activity. Two separate isozymes in maize (Sh1 and Sus) form subunits which appear to combine randomly into tetramers in root tips and other tissues (Chourey et al., 1986). Endosperm sucrose synthase tetramers, however,

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48 are almost entirely composed of subunits encoded by the Sh1 gene (Chourey and Nelson, 1976; Chourey, 1981; Chourey et al., 1986) and are remain active during extraction (Echt and Chourey, 1985). Five different types result in those tissues which exhibit polymerization of both protomers; two are homotetramers and three are heterotetramers. Heterotetramers could theoretically be more unstable than homotetramers since, although very similar, the two subunits are not identical (Echt and Chourey, 1985). Data (Figure 3-5A & B) indicate that greater instability of heterotetramers relative to homotetramers of sucrose synthase was not the cause of the observed activity loss. The profile of declining activity with time is similar in extracts from root tips of a mutant line having only homotetramers of Sh1 (SS1) subunits (W22:s/7?) as it is in extracts from wild type kernels with 5 native tetrameric combinations (NK 508). Rapid loss of sucrose synthase activity with time in maize and other root tips, as well as small tissue size, necessitated the development of a rapid and sensitive assay. Other assays for sucrose synthase using radiometric techniques have been described (Delmer, 1972a and b; Su and Preiss, 1978; Salerno et al., 1979). However, the procedure described here has proven effective and useful due to reduced time from extraction to assay termination, reduced sample size and increased enzyme stabilization.

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CHAPTER 4 SUCROSE SYNTHASE ACTIVITY IN WILD-TYPE MAIZE ROOT TIPS RESPONDING TO ALTERED CARBOHYDRATE STATUS Abstract The two genes encoding sucrose synthase isozymes in maize (Sh1 and Sus1) have been shown to respond to altered tissue carbohydrate status in root tips; Sh1 expression is favored by carbohydrate depletion whereas Sus1 is upregulated when sugars are plentiful (Koch and McCarty, 1988, 1990; Koch et al., 1989). Response at the level of enzyme activity was tested in the present study by assaying sucrose synthase activity in excised maize root tips after 24 h of incubation in a range of glucose concentrations. Little change was evident at the level of total sucrose synthase activity; however, this represented the collective responses of different isozymes and tissue types. Introduction Systems for changes in gene expression in response to altered carbohydrate conditions have been reported for mammalian cells (Lin and Lee, 1984) and in bacteria and yeasts (Carlson, 1987; Schuster, 1989). Recently, seven photosynthetic genes in maize protoplasts have been shown to be repressed and coordinated by sugars (Sheen, 1990). Regulation in higher plants could have 49

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50 important implications for the control of carbohydrate distribution and utilization. Sucrose synthase is considered to have a key function in the allocation of sucrose to various plant organs, and plant carbohydrate status could function as a means of coarse regulation for activity of this enzyme. Sucrose metabolism is important to the majority of plant species because of the nearly ubiquitous role of this sugar in phloem transport to growing and developing plant parts (Avigad, 1982). Two enzymes can catalyze the initial breakdown of sucrose, invertase or the reversible enzyme sucrose synthase. Recently, expression of the gene encoding the shrunken-1 isozyme of sucrose synthase in maize has been shown to be sensitive to carbohydrate levels (Koch and McCarty, 1988; Koch et al., 1989). Northern blot analysis of shrunken-1 mRNA showed levels were elevated in response to carbohydrate depletion. This regulation is distinct from the previously characterized anaerobic induction (Springer et al., 1 986; Koch and McCarty, 1 988). Although the anaerobic induction of Sh1 has received considerable attention in several systems, questions remain regarding the extent to which transcription and translation are synchronized under these conditions. The anaerobic induction in maize has been reported to occur only at the transcriptional level without concomitant changes in protein levels (McElfresh and Chourey, 1988; Taliercio and Chourey, 1989). However, translation of anaerobically induced sucrose synthase mRNA in rice (Ricard et al., 1991) has been demonstrated. The present work examines enzyme-level responses to changes in root carbohydrate status known to alter levels of Sh1 and Sus1 mRNA.

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51 Regulation by sugar concentration may prove to be an important control mechanism whereby plant cells are able to react to cellular carbohydrate status. Materials and Methods Maize seed (Zea mays L, NK 508) were primed for 6 days at 10 C at a water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I' 1 captan (Bodsworth and Bewley, 1981). Seeds were subsequently rinsed free of PEG, soaked for 20 min in 1 .05% (v/v) sodium hypochlorite and rinsed for 20-30 min with ca. 5 I of water. Seeds were germinated in the dark at 1 8 C on moist filter paper in covered glass pans. Continuous airflow was provided (1 liter min" 1 ) throughout the germination period with 40% 0 2 supplied during the final 48 h. At the end of 7 days, 1 cm primary root tips were excised under a sterile transfer hood. Excised root tips (ca. 750 mg per treatment) were incubated in 1 00 ml sidearm flasks containing 50 ml of sterile White's basal salt mixture (White, 1963) supplemented with a range of glucose (0, 0.2, 0.5, 2.0, and 4.0%). Aerobic conditions were maintained during 24 h incubations in the dark at 1 8 C by slow agitation on a rotary shaker (125 rpm) and an airflow of 40% 0 2 (1 I min" 1 ) through an airstone in each flask. Experiments were terminated by twice rinsing in sterile water, blotting excess moisture and freezing them in liquid N 2 . Sucrose synthase activity was determined using a rapid radiometric procedure developed to circumvent enzyme instability previously observed upon extraction from maize root tips (Chapter 3).

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52 Results Sucrose synthase activity was consistently maximal in root tips supplemented with 0.5% glucose (Table 4-1), a level at which the combined levels of mRNA from the two sucrose synthase genes was also greatest (Koch et al., unpublished data). Overall, however, activity of sucrose synthase in whole root tips was not significantly changed by alteration of carbohydrate status by exogenous sugar supply (Table 4-1). It was not possible to distinguish activities of isozymes encoded by the two sucrose synthase genes. These genes, Sh1 and Sus1, were found to exhibit reciprocal responses at the mRNA level to sugar availability in the same sets of roots used for these experiments (Koch et al., unpublished data). Also, changes in distribution of sucrose synthase protein among tissues within these root tips (Nolte, unpublished data) were not reflected at the level of whole root enzyme activity. Discussion Reciprocal regulation of the two isoforms by carbohydrate levels, as has been demonstrated for genes encoding for these isozymes (Koch and McCarty, 1989), could explain the lack of significant differences detected between glucose treatments. The two isozymes of sucrose synthase from maize (encoded by the Sh1 and Sus1 genes) are very similar, differing only slightly in their electrophoretic movement during PAGE (Echt and Chourey, 1985). The Sh1 and Sus1 encoded proteins are capable of catalyzing the same reaction with little difference in affinities

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53 Table 4-1 . Total sucrose synthase activity in wildtype maize root tips incubated in a range of glucose concentrations for 24 hours. intact y % glucose 0 0.2 0.5 2.0 4.0 (/nmol sucrose mg' 1 protein h' 1 ) Expt. 1 0.9 1.2 0.9 1.6 0.9 1.5 Expt. 2 0.5 0.5 0.4 0.6 0.6 0.6 Expt. 3 1.0 0.7 0.8 1.3 0.6 0.7 Mean 0.8 0.8 0.7 1.1 0.7 0.9 S.E.M. ±0.2 ±0.2 ±0.2 ±0.3 ±0.1 ±0.3 y lntact refers to root tips quick frozen in liquid N 2 immediately after excision.

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54 for substrates (Echt and Chourey, 1985), and both are present in extracts from wildtype maize root tips (Chourey et al., 1986). Therefore the total amount of measured sucrose synthase activity would be due to a combined complement of sucrose synthase protein. Despite close homology, however, they have been shown to be distinctive proteins encoded by separate genes (Chourey, 1 981 ; Echt and Chourey, 1985). The two proteins also are distinct in their localization within the maize plant. The protein encoded by Sh1 is primarily located in the endosperm (Chourey and Nelson, 1976; Chen and Chourey, 1989), in the root (Springer et al., 1986; Chourey et al., 1986) and in etiolated shoot (Springer et al., 1986). The Sus1 encoded protein is generally distributed throughout the plant (Chourey, 1981; Echt and Chourey, 1985; Chourey et al., 1988). Carbohydrate responsive proteins have been identified in roots of pearl millet (Baysdorfer and Van der Woude, 1988). Webster and Henry (1987) have also identified an unknown protein with a molecular weight similar to that of the subunits of sucrose synthase in pea root meristem cells undergoing sugar starvation. This protein, however, has yet to be positively identified. Initial findings by Koch and coworkers (Koch and McCarty, 1988, 1990; Koch et al., 1989) indicated that the Sh1 gene of maize was stimulated by low carbohydrate conditions and down-regulated under carbohydrate sufficient conditions. The Sus1 gene responded in an inverse manner. Maas and co-workers (Maas et al., 1990) demonstrated that the promoter from the Sh1 gene was repressed by high

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55 sucrose conditions. However, Salanoubat and Belliard (1 989) found that increased sucrose levels promoted genes encoding sucrose synthase in potato. The possibility also exists that protein fluctuations did not occur after 24 h incubation. However, the shifts in protein localization noted under the same conditions (K. Nolte, University of Florida, unpublished data) indicate that some protein level changes did occur. Spatial separation within root tissue also could explain the differential response of Sh1 and Sus1 observed at gene level without a concomitant change in total enzyme activity. The distribution of sucrose synthase isozymes has been shown to be developmentally regulated, and changes during kernel development (Heinlein and Starlinger, 1989). Chen and Chourey (1989) have reported that expression of sucrose synthase genes is spatially and/or temporally separated in endosperm cells but not in root cells. However, Rowland et al. (1989) demonstrated tissue specific localization of both sucrose synthase genes and isozymes in roots undergoing anaerobic stress. K. Nolte (University of Florida, unpublished data) has shown that shifts in sucrose synthase protein localization occur in maize root tips under carbohydrate depleted and carbohydrate sufficient conditions. Increases of one isozyme in a particular tissue within the root along with decreases of the other in a different tissue would not be apparent at the level of total root sucrose synthase activity. The lack of significant differences in sucrose synthase activity of wildtype maize roots under carbohydrate sufficient and depleted condition, therefore, does not demonstrate that differences evident at the gene level are not also event at the translational

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56 level. Occurrence of maximal enzyme activities in each experiment from samples having the highest levels of both sucrose synthase genes, in fact, tends to indicate that protein changes might be occurring but are not completely detectable under the assay conditions utilized.

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CHAPTER 5 SUGAR RESPONSE OP SUCROSE SYNTHASE AT THE GENE {Sus1), PROTEIN AND ENZYME ACTIVITY LEVELS IN ROOTS OF THE Sh1 MAIZE MUTANT Abstract The sh1 mutant of maize was used to study expression of the Sus1 gene for sucrose synthase in response to sugar availability because this mutant has only one isozyme gene (Sus1) for sucrose synthase and provides a system uncomplicated by the presence of the second gene (Sh1). Koch and McCarty (1988, 1990) have previously demonstrated that Sus1 is up-regulated by plentiful supplies of metabolizable sugars and down-regulated under carbohydrate depletion, whereas Sh1 responds in an inverse manner. Excised root tips from sh1 were incubated for 24 h in White's basal salts medium supplemented with different amounts of glucose. Sus1 mRNA levels were approximately 5-fold greater in treatments with 2.0% vs. 0% or 0.2% glucose. This difference was also reflected in western blot analysis of sus protein. Enzyme activity was elevated 2-fold in root tips from 2% glucose treatments vs. those in 0 or 0.2%. Time-course and switching experiments showed that changes in mRNA or protein were not evident until 24 h and indicated that the response to carbohydrate level had been initiated within 16 h. Roots incubated in 2.0% glucose for 16 h and switched to 0% for 32 h (total of 48 h) responded like those remaining continuously in 2.0% glucose. 57

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58 Overall, enhanced expression of Sus1 was evident at the mRNA, protein and enzyme levels. Introduction Changes in gene expression by carbohydrates have been documented as mechanisms by which bacteria and yeasts respond to changes in their nutrient status (Carlson, 1987; Schuster, 1989). Glucose-responsive genes have also been described in mammalian cells (Lin and Lee, 1984). In addition, Sheen (1990) has presented evidence that the transcriptional activity of promoters of seven photosynthetic genes from maize protoplasts are repressed and coordinated by sugars. Regulation of this type in higher plants could have important implications for carbohydrate allocation and utilization. Sucrose and its metabolic products are important to almost all plant species because of the nearly universal role of this sugar in growth and development (Avigad, 1982). Initial breakdown of sucrose can be catalyzed by either invertase or the reversible enzyme sucrose synthase. Recently, the gene encoding the shrunken-1 isozyme of sucrose synthase in maize has been shown to be responsive to carbohydrate levels (Koch and McCarty, 1988; Koch et al., 1989). Gel blot analysis of shrunken-1 mRNA showed levels were elevated in response to carbohydrate depletion. This may prove to be an important control mechanism whereby plant cells are able to react to cellular nutritional conditions. The Sh1 gene has also been shown to be regulated by anaerobic conditions (Springer, et al., 1986); however, effects on this gene by altered carbohydrate status are distinct

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59 from regulation by anaerobic conditions (Koch and McCarty, 1988). The anaerobic induction has been reported to occur only at the transcriptional level without differences in protein levels (McElfresh and Chourey, 1 988; Taliercio and Chourey, 1 991 ). Possible changes in protein levels and enzyme activity of sucrose synthase due to carbohydrate regulation have been difficult to detect because of the nonspecificity of assay methods (Duke and Koch, unpublished). The two isozymes of sucrose synthase from maize (encoded by the Sh1 and Sus1 genes) are very similar, differing only slightly in their electrophoretic movement during PAGE (Echt and Chourey, 1985). Despite close homology, they are distinctive proteins encoded by separate genes (Chourey, 1981; Echt and Chourey, 1985). The two proteins are, however, distinct in their localization within the maize plant. The protein encoded by Sh1 is primarily located in the endosperm (Chourey and Nelson, 1976; Chen and Chourey, 1989), in the root (Springer et al., 1986; Chourey et al., 1986) and in etiolated shoot (Springer et al., 1986); however Sh1 mRNA does appear in other tissues such as pollen grains (Hannah and McCarty, 1988). The Sus1 encoded protein is more widespread in its localization and is found throughout the plant (Chourey, 1981; Echt and Chourey, 1985; Chourey et al., 1988). The distribution of both proteins is further distinguished under stress conditions (such as anaerobiosis) where tissue-specific localization in roots is readily apparent (Rowland et al., 1989). Tissue specific shifts in sucrose synthase have also been noted in wildtype maize root tips incubated in glucose deficient and sufficient media (K. Nolte, University of Florida,

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60 unpublished data). Marana and co workers (Marana et al., 1990) have found that the two genes encoding sucrose synthase in wheat {Ss1 and Ss2) also show a differential response to stress conditions (anaerobiosis, cold shock and light). Webster and Henry (1987) reported an unknown protein with a molecular weight similar to that of the subunits of sucrose synthase in pea root meristem cells undergoing sugar starvation. Carbohydrate responsive proteins have also been found in roots of pearl millet (Baysdorfer and VanDerWoude, 1988). These proteins, however, are yet to be definitively identified. Initial findings by Koch and co workers (Koch and McCarty, 1988, 1990; Koch et al., 1989) indicated that the Sh1 gene of maize was stimulated by low carbohydrate conditions and downregulated under carbohydrate sufficient conditions. The Sus1 gene responded in an inverse manner. Maas and co-workers (Maas et al., 1990) demonstrated that the promoter from the Sh1 gene was repressed by high sucrose conditions. However, Salanoubat and Belliard (1989) found that increased sucrose promoted genes encoding sucrose synthase. The two genes encoding sucrose synthase in maize respond to altered carbohydrate status (Koch and McCarty, 1988, 1990; Koch et al., 1989), and shifts in sucrose synthase protein localization have been observed under the same conditions (K. Nolte, University of Florida, unpublished data). However, these studies were carried out using a maize line having both sucrose synthase genes present. The present study utilizes the Shrunken-1 mutant of maize to determine

PAGE 70

61 the effects of varying carbohydrate conditions on the Sus1 gene and its sucrose synthase gene product free from the confounding effects of Sh1. Materials and Methods Maize seed (Zea mays L, \N22:sh1) were primed for 6 days at 10 C with a water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I" 1 captan (Bodsworth and Bewley, 1981). Seeds were then rinsed with water, soaked for 20 min in 1 .05% (v/v) sodium hypochlorite and rinsed again for 20-30 min with ca. 5 liters of water. Germination took place in the dark at 1 8 C on moist filter paper in covered glass pans. Continuous airflow was provided (1 liter min1) throughout the germination period with 40% 0 2 supplied during the final 48 h. At the end of 7 days, 1 cm primary root tips were excised under a sterile transfer hood. Excised root tips (ca. 750 mg per treatment) were incubated in 1 00 ml sidearm flasks containing 50 ml of sterile White's basal salt mixture (White, 1963) supplemented with a range of glucose (0, 0.2, 0.5, 2.0, and 4.0%). Aerobic conditions were maintained during 24 h incubations in the dark at 1 8 C by slow agitation on a rotary shaker (125 rpm) and by an airflow 40% 0 2 (1 I" 1 ) through an airstone in each flask. Experiments were terminated by twice rinsing root tips in sterile water, blotting excess moisture and freezing them in liquid N 2 . Responses of root tips to incubation in 0% vs 2.0% glucose were examined after 1 6, 24, or 48 h. Effects of treatment reversals at 1 6 h were also studied by switching roots from 0% glucose treatments to 2.0% glucose and vice versa, then continuing incubations for a total of 48 h.

PAGE 71

62 Enzyme Assay Sucrose synthase activity was determined by a rapid radiometric procedure developed to circumvent enzyme instability previously observed upon extraction from maize root tips (Chapter 3). RNA Extraction and Northern Blotting Samples were ground to a fine powder in a mortar and pestle with liquid N 2 and RNA extracted according to McCarty (1986). Total RNA was quantified by absorbance at 260 nm. Total RNA was separated by electrophoresis in 1 % agarose gels containing formaldehyde (Thomas, 1980), blotted to a nylon membrane (Hybond-N, Amersham Corporation, Arlington Heights, IL) and probed as per Church and Gilbert (1984) with genomic clones of Sus1 (McCarty et al., 1986) radiolabeled by random primer. Blots were rinsed and placed on X-ray film at -80 C. Protein Gel Blots Subsamples from protein extracts to be used for enzyme assays and separated on native PAGE using the system of Laemilli (1970) without SDS. Polyacrylamide concentrations of the stacking and separating gels were 2.5% and 5%, respectively. Proteins were resolved at 4 C by applying 1 5 V for 9 h, then 1 25 V for 11 h (constant current) and temperature (4 C). Proteins were electroblotted to nitrocellulose membranes and probed with polyclonal antibodies following the procedure of Towbin et al. (1979). Sucrose

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63 synthase antisera, obtained from D.R. McCarty, was generated in rabbits using protein purified from maize kernels (W64A x 182E) 22 days after pollination. Antisera was diluted 1 :1000 and cross reacted strongly to both the Sh1 and Sus1 gene products where such were present. Results Levels of Sus1 mRNA in excised maize roots were greater after 24 h of incubation in 2% glucose than in those that had received 0 or 0.2% glucose (Figure 5-1). At the protein level, western blot analysis showed little or no change with increasing carbohydrate concentration (Figure 5-2); however, enzyme activity was elevated in root tips incubated at high vs. low glucose concentrations (Table 5-1). Both lines of evidence indicated the protein level response was less pronounced at 24 h than that of mRNA. The time-course of changes in Sus1 message levels in root tips showed that differences between those given 0% vs. 2.0% exogenous glucose became apparent sometime between 16 and 24 h (Figure 5-3). Initial decreases appeared to occur in both treatments, but within 24 h, Sus1 mRNA levels in glucose supplemented roots had risen well above those with limited sugar supply. The greatest difference between carbohydrate treatments was evident after 48 h of incubation. Treatment reversals indicated that the gene response to carbohydrate level had been initiated within 16 h (Figure 5-3). Roots incubated in 2.0% glucose for 16 h and switched to 0% for 32 h (total of 48 h) responded like those remaining continuously in 2.0% glucose. Roots initially deprived of glucose and then

PAGE 73

64 % glucose Intact 0 0.2 0.5 2.0 4.0 Expt. 1 ^^^^ Expt. 2 Figure 5-1 . RNA gel blot analysis of Sus1 expression in maize roots incubated in a range of glucose concentrations for 24 hours.

PAGE 74

65 % glucose Intact 0 0.2 0.5 2.0 4.0 Expt. 1 ** m mrm * ** m ' Expt. 2 Figure 5-2. Protein gel blot of Sus1 encoded sucrose synthase from maize roots incubated in a range of glucose concentrations for 24 hours. Data from Expts. 1 and 2 were obtained from the same set of roots sampled for RNA analyses shown in Figure 5-1 .

PAGE 75

66 Table 5-1. Sucrose synthase activity in mutant maize (W22:s/77) root tips incubated in media containing a range of glucose concentrations for 24 hours. Sucrose synthase activity % glucose Intact 0 0.2 0.5 2.0 4.0 (jumol sucrose g" 1 protein h" 1 ) Expt. 1 0.22 0.13 0.07 0.13 0.32 0.32 Expt. 2 0.35 0.21 0.13 0.20 0.40 0.39

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67 16h 24h 48h 16h+32h Intact + + +-/++/Figure 5-3. RNA gel blot analysis of Sus1 mRNA expression in maize roots incubated in 0% (-) or 2.0% (+) glucose for various time periods. At 16 h switching treatments were conducted by changing roots in 0% glucose to 2.0% glucose (-/+) and vise verse (+/-); incubations were continued for 32 additional hours.

PAGE 77

68 switched to 2.0% glucose responded similarly to those remaining in 2.0% glucose for the entire time. At the protein level.changes in response to altered carbohydrate availability were not apparent at 16 h, remained barely detectable at 24 h, but were clearly evident after 48 h (Figure 5-4). Treatment reversals indicated that a protein-level response occurred only when 1 6 h of elevated glucose treatment was followed by 32 h of glucose deprivation. The response was similar to that of root tips that had remained continuously in 2.0% glucose. Slight differences in enzyme activity between treatments were evident after 16 h or 24 h (Table 5-2); however activity in glucose supplemented tips had risen to levels two-fold greater than those without exogenous sugars within 48 h. Discussion The significance of results described here are two-fold. First, data demonstrate that the differential response to changing carbohydrate availability by the Sus1 gene for sucrose synthase is apparent at the translational level as well as at the transcriptional level. The two genes encoding sucrose synthase previously have been shown to respond differentially to carbohydrate supply (Koch and McCarty, 1988; 1990; Koch et al., 1989). Differences at the protein and enzyme level, however, have been difficult to detect due to cross reactivity of polyclonal antibodies and the collective contribution of both isozymes to activity measurements. Second, the resulting changes in physiology may allow the cells to adjust their carbohydrate metabolizing capacity to the available supply. Use of

PAGE 78

69 1 6h 24h 48h 1 6h + 32h Intact ~ + 1 + 1 +~ -/+ +/Figure 5-4. Protein gel blot of Sus1 encoded sucrose synthase from maize roots incubated in 0% (-) or 2.0% (+) glucose for various time periods. At 16 h switching treatments were conducted by changing roots in 0% glucose to 2.0% glucose (-/+) and vise verse (+/-); incubations were continued for 32 additional hours.

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71 the Sh1 maize mutant has allowed the response to be characterized using a simple, single enzyme system with respect to changing sugar supply. The increased protein and enzyme activity evident at increased exogenous glucose levels indicate that the plant tissue can adjust this first step in their sucrosemetabolizing capacity relative to its carbohydrate status. After 24 h at a given glucose level, changes in gene expression were more marked than were differences at the protein and enzyme levels (Figure 5-1 , Figure 5-2 and Table 5-1). This is not surprising given the probable presence of previously formed RNA and protein (both appear to be relatively long-lived) as well as the comparatively long-term progression of the response to the maximal extent observed at 48 h (Figure 5-3 and Table 5-2). Chourey et al. (1991 b) reported that the sucrose synthase gene in sorghum homologous to Sus1 gene from maize is anaerobically induced, but levels of the respective protein do not change. Anerobic induction, however, was terminated after only 12 h. Anaerobic induction of Sh1 in maize becomes apparent between 6 and 12 hours but message levels are not maximal until at least 24 h (Duke and Koch, unpublished data). Data from the present work indicate that like the respiratory drop noted by Brouquisse et al. (1991), at least 20 hours are required before a change in Sus1 is fully apparent at the gene level and even longer at the protein level. Nonetheless, data are presented here at the levels of mRNA, protein and enzyme activity that indicate that expression of the Sus1 gene for sucrose synthase is responsive to carbohydrate availability to an extent not evident in background levels of total RNA

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72 and protein. The duration of time required for this response is consistent with the proposed physiological function of sucrose in coarse adjustment of root growth relative to sugar supply (Farrar and Williams, 1990). The increased levels of Sus1 mRNA and subsequent elevation of its respective protein with carbohydrate status may give insight into specific roles for this isozyme as opposed to the Sh1 gene product. Sucrose synthase activity could be key to the regulation of carbon entry into the respiratory pathway (Huber and Akazawa, 1986; Black et al., 1987). The enhanced expression of Sus1 under plentiful carbohydrate supplies accompanied probable increases in respiratory activity in the root tips (Saglio and Pradet, 1980; Farrar and Williams, 1990; Brouquisse et al., 1991). In addition, carbohydrate content in many tissues has been correlated with the respiration rate (Penning de Vries et al., 1979; Farrar, 1985). Saglio and Pradet (1980) also found that an exogenous supply of 0.2 M glucose was required to bring the respiration rate of excised maize roots back to the level of intact tissue, indicating that the rate of metabolic activity of the root tips may be closely tied to sugar import. Perhaps another line of evidence supporting control of respiration by carbohydrate status comes from the work of Douce et al. (1 990) in which sycamore cells in culture showed loss of mitochondrial function when starved of sucrose; the beginning of the decline coincided with the fall in endogenous sugar concentrations. Another possible role for the Sus1 gene product may be in the diversion of carbohydrate to cell wall biosynthesis. Roots in 2.0% glucose medium show

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73 marked growth during the 24 hours of incubation (data not shown). During this time, there is a demand for cell wall synthesis by the expanding cells. Sucrose synthase has been implicated in the directing of carbohydrates for polysaccharide biosynthesis (Amino et al.,1985; Hendrix, 1990). The level of involvement for nucleotide-sugars during cell wall polysaccharide biosynthesis has been implicated in a need for greater activity of this enzyme (Maas et al., 1990). Sucrose synthase activity was elevated in cell cultures of Catharanthus roseus during the G1 phase when total amounts of cell wall biosynthesis increased significantly (Amino et al., 1985). Sugar modulation of Sus1 could convincingly combine production of cell wall precursors with other aspects of increased growth (Farrar and Williams, 1 990) likely to accompany an enhanced sugar supply. The carbohydrate response of sucrose synthase in the present study differs from previously demonstrated regulation in that it occurs in rapidly growing and metabolizing structures. Other studies have involved sucrose synthase regulation in storage tissues where processes such as starch accumulation predominate. Loss of the shrunken-1 gene in maize results in a typical endosperm phenotype where starch deposition is reduced by over 70% (Chourey and Nelson, 1976); however, a mutant lacking a functional Sus1 gene has no apparent phenotype (Chourey et al., 1988). Starch deposition accompanies protein accumulation in developing potato tubers and levels of mRNA encoding sucrose synthase have been shown to increase in this tissue (Salanoubat and Belliard, 1989). Levels of storage proteins, such as patatin, also accumulate during this time (Paiva et al.,

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74 1983). Increased levels of sucrose result in elevated levels of genes for both sucrose synthase (Salanoubat and Belliard, 1989) and patatin (Rocha-Sosa et al., 1989; Wenzler et al., 1989) in tissues where they are not usually found. A rise in sugar availability can also result in increased transcription and translation of a unique storage protein in stem and leaf tissues of sweet potato (Hattori et al., 1990). The diversity of processes operating in the system utilized in this study suggests that sugar-responsive gene expression (ie. Sus1) may have broad implications in the formation and function of non-storage plant tissues.

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CHAPTER 6 AN ORGAN-SPECIFIC INVERTASE DEFICIENCY IN THE PRIMARY ROOT OF AN INBRED MAIZE LINE Abstract An organ-specific invertase deficiency affecting only the primary root system is described in the Oh 43 maize inbred. Invertases (acid and neutral/soluble and insoluble) were assayed in various tissues of hybrid (NK 508) and inbred (Oh 43, W22) maize lines to determine the basis for an early report that Oh 43 root tips were unable to grow on sucrose agar (Robbins, 1958). Substantial acid invertase activity (7.3 to 16.1 ^mol glucose mg' 1 protein h' 1 ) was evident in extracts of all tissues tested except the primary root system of Oh 43. This deficiency was also evident in lateral roots arising from the primary root. In contrast, morphologically identical lateral roots from the adventitous root system had normal invertase levels. These results suggest that ontogenetic origin of root tissues is an important determinant of invertase expression in maize. Adventitious roots (including the seminals) arise above the scutellar node and are, therefore, of shoot origin. The Oh 43 deficiency also demonstrated that invertase activity was not essential for maize root growth. Sucrose synthase was active in extracts from all root apices and theoretically provided the only available avenue for sucrose degradation in 75

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76 primary root tips of Oh 43. The deficiency described here will provide a useful avenue of investigation into the expression and significance of root invertase. Introduction Sucrose breakdown is critical to the vast majority of plant species because non-photosynthetic tissues depend on import of this sugar for their growth and development. Initial cleavage of sucrose can be catalyzed by either invertase or the reversible enzyme sucrose synthase. Invertases are especially active in tissues undergoing rapid cell division such as shoot and root apices (Avigad, 1982). Previously, the role of invertases in sucrose transfer was considered particularly important in plants such as sugar cane and maize where substantial sugar movement occurred through the cell wall space and was accompanied by action of an extracellular invertase (Glasziou and Gayler, 1972; Hawker and Hatch, 1965). Recent evidence, however, indicates that although much hydrolysis is often observed, invertase activity may not be essential for sucrose uptake into either sugar cane stems (Lingle, 1989; Thorn and Maretzki, 1990) or maize kernels (Schmalstig and Hitz, 1987). Sucrose generally is believed to enter root tips without traversing the extracellular space (Giaquinta et al., 1983); however growing roots can differ markedly in their capacity to lose (Rovira and Davey, 1974) and retrieve (Robbins, 1 958) exogenous sugars. Net losses do occur. The extent of sugar efflux from roots can be affected by irradiance level, nutritional status, moisture availability and

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77 temperature (Rovira and Davey, 1974). The composition of root sugars exuded is quite variable but includes both reducing and non-reducing sugars (Rovira and Davey, 1974). Glucose and fructose are often taken up from the extracellular space more rapidly than is sucrose (Humphreys, 1974). Retrieval of solutes from the apoplast and the form in which they are available may thus be a potentially important attribute of root carbon balance. A deficiency in this retrieval process was first indicted by Robbins' report (1958) that roots of a maize inbred, Oh 43, were unable to grow on sucrose agar medium, yet roots of another line, Hy 2, grew quite well. Only when roots of both were cultured immediately adjacent to one another, were those of Oh 43 able to grow. Growth of excised Oh 43 root tips also occurred when glucose was substituted for the sucrose. Oh 43 was concluded to be "incapable of inverting sucrose" in its root tips. Preliminary investigations by B. Burr (Brookhaven National Laboratory, personal communication) indicated that a lack of invertase may have been the reason for the inability of Oh 43 roots to metabolize sucrose. The absence of invertase activity could have important implications for sucrose import not only because of potential effects on the retrieval system, but also because sucrose utilization in such an instance could theoretically be initiated only via action of sucrose synthase. In addition, genetic material which lacks activity of a specific enzyme can be useful in investigations of physiological processes normally mediated by these enzymes (Koch et al., 1982). The present report demonstrates that invertase is not essential for primary root growth despite

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78 probable advantages of its presence and indicates an unusual organ-specific difference in expression between primary and adventitious roots. Materials and Methods Plant Material Maize seed (Zea mays L. NK 508, W22 and Oh 43) were germinated on moist filter paper in petri dishes. Seeds were imbibed for 24 hours and pericarps removed, allowing more uniform germination and more effective surface sterilization (20 min soak in 0.525% sodium hypochlorite). Five successive 2 mm segments were sampled from the tips of primary roots 4 to 5 days after germination. Intact roots of Oh 43 seedlings grew more slowly than did those of NK 508 or W22, but all roots had reached 2 cm prior to excision. Tissue samples were weighed, frozen in liquid N 2 and stored at -80 C until assayed for invertase activity. In subsequent experiments, 5 mm root tips were excised from primary and adventitious roots for invertase and sucrose synthase activity measurements. Plants and tissues were as above. Tissue Extraction Frozen tissue samples were ground to a fine powder in liquid N 2 using a mortar and pestle. Frozen powder was transferred to a second mortar containing ice-cold 200 mM HEPES buffer (pH 7.5) with 1 mM DTT, 5 mM MgCI, 1 mM EGTA, 20 mM sodium ascorbate and 10% (w/w) PVPP. One ml of extraction buffer was used for every 100 mg of tissue fresh weight. Buffered extract was centrifuged at

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79 1 4,000 x g for 1 min to sediment particulate matter. Supernatant was dialyzed (27,000 mw cutoff) at 4 C for 24 h against extraction buffer diluted 1 :40. Buffer was changed after 1 h and thereafter, every 4 h. Soluble dialyzed extract was assayed for invertase as described below. Previously separated particulate matter was rinsed with one volume of extraction buffer and assayed for insoluble, cellwall-bound invertase (soluble acid invertase includes both vacuolar and loosely bound extracellular enzyme [Avigad, 1982]). To test the possibility that the soluble enzyme was present in primary roots of Oh 43 but was being bound or inactivated during the extraction procedure, two additional extraction/assay methods were employed. First, adventitious root extracts, previously shown to contain active invertase activity, were added to those of primary apices. The resulting mixture was dialyzed and assayed for enzyme activity. Second, three cm apices of both primary and adventitious roots Oh 43 roots were excised. Apices of these roots (0.5 cm) were suspended in extraction buffer for three hours at 27 C. Buffer alone was subsequently dialyzed as described above. The portion of each root which had been immersed in the extraction buffer was excised for fresh weight measurement. After dialysis, the buffer-enzyme solution was analyzed for enzyme activity. Enzyme Assays Soluble and insoluble forms of acid invertase were assayed as described by Lowell et al. (1989). Reaction media contained 50 mM sucrose, and pH of 4.5 was adjusted with a sodium acetate buffer. Neutral invertase was assayed using

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80 the same reaction medium adjusted to pH 7.5 with potassium phosphate buffer. Initial assays were also performed at pH ranges of 4.0 to 5.5 for acid invertase and 7.0 to 8.0 for neutral invertase. After a 15 min incubation at 30 C, glucose production was quantified by the glucose oxidase method (Sigma Chemical Co.). Sucrose synthase was assayed in the degradative direction using a radiometric assay quantifying the production of 14 C-UDPG (Chapter 3). Histochemical Staining Free-hand cross sections from apices of both primary and adventitious roots were fixed in 4% formalin (pH 7.0) for 30 min and rinsed in water at least 1 0 times over a period of 3 hours to remove endogenous sugars (Doehlert and Felker, 1987). Sections were then incubated in a sodium phosphate buffer (0.38 M, pH 6.0) containing 0.24 mg ml" 1 nitroblue tetrazolium, 0.14 mg ml* 1 phenazine methosulfate, 25 units ml' 1 glucose oxidase and 5 mg ml" 1 sucrose (Doehlert and Felker, 1987). Control sections were incubated in the same mixture without sucrose. After rinsing in water, sections were post fixed in 4% formalin (pH 7.0) and photographed under a microscope. Results Primary roots of Oh 43 showed little or no acid invertase activity (Table 6-1). In contrast, acid invertase was active in extracts from apical areas of roots from other maize lines examined (NK 508 and W22). Activity, per unit fresh weight, was greatest in root apices, decreasing with distance from the tip until no longer

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82 detectable farther than 8 mm from the apex. Soluble enzyme accounted for 88 to 92% of the total acid invertase activity, and the remainder was due to action of the insoluble enzyme. Neutral invertase activity was insignificant or absent from all lines (data not shown). Although invertase action was essentially undetectable in the primary roots of Oh 43, adventitious root extracts showed levels of activity as least as great as those from the primary roots of other lines tested (Table 6-2). Active acid invertase was released into buffer in the extracellular space around adventitious root tips of Oh 43 during 3 hours of emersion (1 3.4 ± 3.5 yimol glucose g" 1 FW h" 1 -ca. 25% of the activity on a fresh weight basis of N 2 and buffer extracted roots). The same was not observed during emersion of primary roots from this line. Also, from 83% to 94% (13.3 to 15.1 jumol glucose mg' 1 protein h" 1 ) of the invertase activity in adventitious root extracts remained after addition of extracts from primary roots of Oh 43. Unlike invertase, sucrose synthase was active in extracts of both primary and adventitious roots from Oh 43 seedlings. Activity of this enzyme was approximately similar in both root types, as observed for the other two lines tested (Table 6-2). Extracts of lateral roots originating from primaries exhibited no detectable acid invertase activity (Table 6-3), in contrast to counterparts derived from adventitious roots. Shoots of 5-day-old Oh 43 seedlings and endosperm or scutellum tissue from developing kernels (23 DAP) also exhibited activity of both soluble and insoluble acid invertase (Table 6-3).

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Table 6-3. Soluble acid invertase activity in various tissues of Oh 43, an inbred line of lea mays. jimol glucose /^moi giucose mg" 1 protein h" 1 g 1 FW h* 1 Shoot 10.8 t 0.6 25.8 ± 1.5 Endosperm 7.3 ± 1.2 12.6 ± 2.1 Scutellum 9.0 ± 0.8 19.4 ± 1.7 Primary roots 0.1 ±0.1 0.2 ± 0.2 Lateral roots from 1° roots 0.1 ± 0.1 W 0.2 ± 0.1 W Adventitious roots 16.1 ±3.2 60.8 ± 12.1 Lateral roots from adventitious roots 10.4 ± 0.9 38.7 ± 3.2 W --Activity in root tips from axenic culture was 0.0 ± 0.0 /xmol g' 1 FW h ; thus microorganisms are a likely source of the residual activity in ca. 1 of 3 assays. Shoot, primary roots and adventitious roots were samples from 6-day-old seedlings. Endosperm and scutellum were from developing kernels 23 DAP. Lateral roots developed after 2 to 3 weeks of growth and were then excised. Each value represents the mean of 3 separate samples ± S.E.M.

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85 Histochemical staining indicated that no invertase activity was detectable in the primary roots of Oh 43 (Figure 6-1). Cross sections of primary and adventitious roots of Nk 508 and adventitious roots of Oh 43 all stained positive for invertase activity. Invertase activity was primarily in the cortex and was localized intercellularly. Discussion These data confirm that Oh 43 (an inbred line of maize) lacks invertase activity in its primary root tips. Surprisingly, however, a deficiency was not evident in structurally and functionally similar adventitious roots (Table 6-2), other tissues of the same Oh 43 plants (Table 6-3) or in the developing kernels of this line (Doehlert, et al., 1988). The genetic potential for invertase expression is therefore present. The lack of activity may result from altered regulation of gene expression (transcription or translation), the loss of a tissue specific invertase isozyme or the existence of an unidentified effector of enzyme function. The significance of results described here is twofold. First, evidence is presented for differential expression of invertase in morphologically identical organs that differ primarily in point of origin. This is most strikingly illustrated in the apparent distinction between lateral roots arising from primary and adventitious root systems. Differential expression of genes in morphologically identical structures are unusual; however, organ or tissue-specific differences have been well documented (Fluhr et al., 1986; Xie and Wu, 1989). Xie and Wu (1989), for example, found that genes for alcohol dehydrogenase were differentially expressed in root and shoot tissues of rice

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ure 6-1 . Histochemical localization of invertase activity in free-hand, fresh cross sections of root apices of 6-day-old maize seedlings. Sections are approximately 50 nm in thickness. Primary (A) and adventitious (B) root cross sections from NK 508 incubated in reaction medium without sucrose (controls). Primary (C) and adventitious (D) root cross sections from Oh 43 incubated in reaction medium (note only adventitious root section exhibits blue formazan reaction product [dark areas]). Primary (E) and adventitious (F) root cross sections from NK 508 incubated in reaction medium (note both sections exhibit reaction product [dark areas]). Bar represents 50 nm.

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88 plants. One isozyme predominated in shoot-derived organs (leaves, sheaths, nodes and pollen) and the other isozyme showed highest activity in the roots. It is interesting in this respect that the adventitious roots of maize arise above the scutellar node of the developing seedling and are, therefore, derived from shoot tissue (Hayward, 1938). The difference in invertase expression in primary and adventitious root systems may reflect a similar root/shoot dichotomy. Therefore, the tissue of origin and cell lineage may be more important than organ identity and function in regulating invertase expression. Other variants in invertase expression have been described. Echeverria and Humphreys (1984) reported a hybrid maize line (DKXL80) which exhibited no soluble invertase activity in contrast to previously tested lines. Other tissues of this maize line exhibited normal invertase activity. Second, data demonstrate that invertase is not essential for primary root growth. The primary roots of Oh 43 exhibited no signs of premature senescence, and, if left intact, continued apparently normal growth for many days (data not shown). Invertase's role in sucrose import into roots has been questioned by Chapleo and Hall (1989a) who concluded that although present, apoplastic root invertase did not have a direct role in sugar transport in Ricinus. However, substantial activity of invertase has been widely documented in roots of plants such as pea (Lyne and ap Rees, 1971), bean (Robinson and Brown, 1952), tomato (Chin and Weston, 1973), Ricinus (Chapleo and Hall, 1989a, b and c), oat (Pressy and Avants, 1980), and maize (Chang and Bandurski, 1963; Hellebust and Forward, 1962). In maize, the Oh 43 invertase deficiency apparently prevents

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89 utilization of exogenous sucrose (Robbins, 1958). Specific tissue localization of invertase also has been described. Peak activity for root invertase is generally 2-3 mm behind the apex and corresponds to the region of expansion and elongation in pea (Robinson and Brown, 1952) and maize (Hellebust and Forward, 1962). In Ricinus this activity predominates in the cortex (Chapleo and Hall, 1989). Although invertase may not have a direct role in sucrose import in roots, it still may be important to two major aspects of root biology. First, invertase has been implicated in formation of mycorrhizal associations (Purves and Hadley, 1975). Maize (Gerdemann, 1964; Kothari et aJ., 1990) and 90% of other agriculturally important species form these beneficial symbioses under field conditions (Gerdemann, 1986). Invertase activity typically increases and hexose levels rise at infection sites of biotrophic fungi (Long et al., 1 975). Elevated hexose content in roots upon infection by mycorrhizal fungi has been attributed to a rise in invertase levels (Purves and Hadley, 1975). It is not known whether this is host or fungal invertase; however, Oh 43 does not appear to provide the former in its primary root systems. Second, the purported lack of a sucrose carrier in the plasma membrane of maize root cells (Lin et al., 1984) would indicate that if sucrose were released into the apoplast, retrieval might proceed more effectively in the presence of extracellular invertase. Such retrieval could be particularly important during stress or periods when sugar losses from the symplast were elevated. Any physiological consequence of this deficiency would most likely be evident early in seedling

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90 development because the root system would consist solely of a primary root at this time. Later in seedling development, adventitious or seminal roots rapidly take over a dominant role, leaving the primary root with little or no essential function (Hayward, 1938). In the field, lines having Oh 43 as a progenitor have been observed to suffer from poor emergence rates in damp soils (B. Martin, Pioneer Seed, personal communication). However, conclusive evidence of a physiological effect will require generation of and analysis of isogeneic lines. In conclusion, the deficiency described here will be useful for investigation into the regulation and physiological function of root invertase. Because the primary root system in maize is nonessential, invertase deficient roots can be studied without deleterious effects on the overall physiology of the plant. In addition, this deficiency is significant because it reveals an unexpected distinction between primary and adventitious root development in maize. At some level, the mechanisms which regulate invertase in these root systems must differ.

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CHAPTER 7 SUMMARY AND CONCLUSIONS Sucrose metabolism is important to the majority of plant species because of the widespread role of this sugar in growth and development through its function as a phloem transport sugar. Initial breakdown of sucrose can be catalyzed by either sucrose synthase, a reversible enzyme, or invertase. Expression of genes encoding the sucrose synthase isozymes (Sh1 and Sus1) have been found to be sensitive to carbohydrate levels (Koch and McCarty, 1988, Koch et al., 1989). Northern blot analysis of mRNA showed levels showed those of Sh1 were up-regulated in response to carbohydrate depletion whereas those of Sus1 were down-regulated under the same conditions. This may prove to be an important control mechanism whereby plant cells are able to react to cellular carbohydrate status. The first purpose of this work was to determine the extent to which carbohydrate-modulated changes in message levels affected activity of sucrose synthase the level of enzyme activity. However, previously available sucrose synthase assay methods proved ineffective for maize root tips due to a precipitous loss of activity after tissue extraction. A rapid, radiometric assay for sucrose synthase was therefore developed that overcomes these obstacles. Extraction and assay were optimized for factors such as substrate concentration, pH, assay 91

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92 length, and inclusion (or exclusion) of various anti-oxidants and protease inhibitors. The assay has proven effective for a range of tissues and species examined and provides a particularly effective measurement for sucrose synthase action in root tips. Characterization of the in vitro instability also provided evidence suggestive of possible phosphorylation effects on activity. Activity of sucrose synthase was then assayed in extracts from intact wildtype maize root tips and from those that had been excised and incubated for 24 h in a basal salts medium with varying levels of glucose. Enzyme activity showed little or no significant differences between treatments. However, results represented contributions by two isozymes (encoded by genes exhibiting reciprocal responses under the same conditions [Koch et al., 1 989]), and were the collective sum of different tissues (that showed specific alterations in sucrose synthase distribution [K. Nolte, University of Florida, unpublished data]). A different approach was thus utilized to examine the relationship between sugar-modulated changes in message levels and extent of enzyme level effects. Use of the shrunken-1 maize mutant (deficient in a functional Sh1 gene) showed that carbohydrate-responsive gene expression was evident for Sus1 at the levels of mRNA, protein and enzyme activity. Time-course and treatment reversals also indicated greater responses between glucose sufficient and deficient treatments were observed after 48 h of incubation, indicating a possible coarse control mechanism. The second purpose of this work was to explore a possible line of

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93 investigation into the physiological significance of invertase by investigating a putative deficiency in sucrose-metabolizing capacity in the Oh 43 line of maize (Robbins, 1958). Complete absence of invertase in this line was not likely due to its demonstrated activity in the scutellum (Doehlert et at., 1988). However, the hypothesis tested here was that an organ-specific deficiency or invertase suppression was responsible for the metabolic anomalies in Oh 43. The primary root of Oh 43 indeed was shown to lack invertase activity. In contrast, adventitious roots of the same plants exhibited wildtype levels of invertase activity. Initial characterization of this mutation will provide an effective tool for future investigations into the physiological role(s) of this enzyme in roots. Use of this mutation and isogeneic wildtype counterparts in combination with the nulls for each sucrose synthase gene may allow a better understanding of the biological functions of each.

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LITERATURE CITED Akazawa, T. and K. Okamoto. 1 980. Biosynthesis and Metabolism of Sucrose, pp. 1 99-220 in The Biochemistry of Plants. New York: Academic Press. Al-Bakir, A.Y. and J.R. Whitaker. 1978. Purification and properties of invertase from dates (Phoenix dactylifera L. var. Zahdi) J. Food Biochem. 2:32-160. Amino, S., Y. Takeuchi and A. Komamine. 1985. Changes in enzyme activities involved in formation and interconversion of UDP-sugars during the cell cycle in a synchronous culture of Catharanthus roseus. Physiol. Plant. 64:111-117. Amir, J. and J. Preiss. 1982. Kinetic characterization of spinach leaf sucrosephosphate synthase. Plant Physiol. 69:1027-1030. Anderson, R.S. and E.E. Ewing. 1978. Partial purification of potato tuber invertase and its proteinaceous inhibitor. Phytochem. 17:1077-1081. ap Rees, T. 1974. Sucrose metabolism, pp. 53-73 in D.H. Lewis, ed. Storage Carbohydrates in Vascular Plants. Cambridge, U.K.: Cambridge University Press. Arnold, W.N. 1966. A column method for enzymic characterization of coarse cellular fractions: application to insoluble /3-fructofuranosidase from grape. Arch. Biochem. Biophys. 113:451-456. Avigad, G. 1 964. Sucrose-uridine diphosphate glucosyl-transferase from Jerusalem artichoke tubers. J. Biol. Chem. 239:3613-3618. Avigad, G. 1982. Sucrose and other disaccharides. pp. 217-347 in F.A. Loewus and W. Tanner, eds. Encyclopedia of Plant Physiology, New Series. Vol. 1 3A. New York: Springer-Verlag. Avigad, G., N. Levin and Y. Milner. 1964. Sucrose metabolism in plant storage tissues. 6th Intl. Congr. Biolchem. Abst. p. 502. 94

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95 Avigad, G. and Y. Milner. 1966. UDP-Glucose: fructose transglucosylose from sugar beet root. Methods Enzymol. 8:341-345. Baysdorfer, C. and W.J. VanDerWoude. 1988. Carbohydrate responsive proteins in the roots of Pennisetum americanum. Plant Physiol. 87:566-570. Bennett, J. 1984. Regulation of photosynthesis by protein phosphorylation, pp. 227-247 in P. Cohen, ed. Enzyme Regulation by Reversible Phosphorylation-Further advances. New York: Elsevier. Black, C.C., L Mustardy, S.S. Sung, P.P. Kormanik, D-P. Xu and N. Paz. 1987. Regulation and roles for alternative pathways of hexose metabolism in plants. Physiol. Plant. 69:387-394. Bodsworth, S. and J.D. Bewley. 1981. Osmotic priming of seeds of crop species with polyethylene glycol as a means of enhancing early and synchronous germination at cool temperatures. Can. J. Bot. 59:672-676. Bracho, G.E. and J.R. Whitaker. 1990a. Characteristics of inhibition of potato (Solanum tuberosum) invertase by an endogenous proteinaceous inhibitor in potatoes. Plant Physiol. 92:381-385. Bracho, G.E. and J.R. Whitaker. 1990b. Purification and partial characterization of potato (Solanum tuberosum) invertase and its endogenous proteinaceous inhibitor. Plant Physiol. 92:386-394. Brouquisse, R., F. James, P. Raymond and A. Pradet. 1991. Study of glucose starvation in excised maize root tips. Plant Physiol. 96:619-626. Burakhanova, E.A., I.M. Dubinina and LF. Kudryartseva. 1987. Endogenous invertase inhibitor in sugar beet root ontogenesis. Soviet Plant Physiol. 34:235-239. Cardini, C.E., LF. Leloir and J. Chiriboga. 1955. The biosynthesis of sucrose. J. Biol. Chem. 214:149-155. Carlson, M. 1987. Regulation of sugar utilization in Saccharomyces species. J. Bacterid. 169:4873-4877. Carlson, M. and D. Botstein. 1982. Two differentially regulated mRNAs with different 5' ends encode secreted and intracellular forms of yeast invertase. Cell 28:145-151.

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109 Salanoubat, M. and G. Belliard. 1989. The steady-state level of potato sucrose synthase mRNA is dependent on wounding anaerobiosis and sucrose concentration. Gene 84: 181-1 85. Salerno, G.L 1986. Relationship between sucrose and starch in developing maize endosperm UDP-glucose phosphorylase activity. Plant Sci. 44:111-118. Salerno, G.L, S.S. Gamundi and H.G. Pontis. 1979. A procedure for the assay of sucrose synthetase and sucrose phosphate synthetase in plant homogenates. Anal. Biochem. 93:196-199. Sampietro, A.R., M.A. Vattuone and F.E. Prado. 1980. A regulatory invertase from sugar cane leaf-sheaths. Phytochem. 19:1637-1642. Sasaki, T., K. Tadokoro and S. Suzuki. 1971. Multiple forms of invertase of potato tuber stored at low temperature. Phytochem. 1 0:2047-2050. Schaffer, A.A., B. Aloni and E. Fogelman. 1987. Sucrose metabolism and accumulation in developing fruit of Cucumis. Phytochem. 26:1883-1887. Schaffer, A.A., I. Rylski and M. Fogelman. 1989. Carbohydrate content and sucrose metabolism in developing Solanum muricatum fruits. Phytochem. 28:737-739. Schmalstig J.G. and W.D. Hitz. 1987. Transport and metabolism of a sucrose analog (1 'fluorosucrose) into Zea mays L. endosperm without invertase hydrolysis. Plant Physiol. 85:902-905. Schuster, J.R. 1989. Regulated transcriptional systems for the production of proteins in yeast: Regulation by carbon source, pp. 83-108 in P.J. Barr, A.J. Brike and P. Valenzuela, eds. Yeast Genetic Engineering. London: Butterworths. Servaites, J.C., B.R. Fondy, B. Li and D.R. Geiger. 1989. Sources of carbon for export from spinach leaves throughout the day. Plant Physiol. 90: 1 1 68-1 1 74. Setia, N. and CD. Malik. 1985. Changes in some enzymes of carbohydrate metabolism in developing pod and seed of chick pea (C/cer arietinum) Phyton 25:93-99. Sexton R. and J.F. Sutcliffe. 1969. The distribution of 0-glycerophosphatase in young roots of Pisum sativum. L. Ann. Bot. 33:407-419.

PAGE 118

110 Sharma, K.P. and I.S. Bhatia. 1980. Sucrose metabolism in Sorghum vulgare at ripening. Physiol. Plant. 48:470-476. Sheen, J. 1990. Metabolic repression of transcription in higher plants. Plant Cell 2:1027-1038. Silvius, J.E. and F.W. Snyder. 1979. Comparative enzymatic studies of sucrose metabolism in the taproots and fibrous roots of Beta vulgaris L Plant Physiol. 54:1070-1073. Springer, B., W. Werr, P. Starlinger, D.C. Bennett, M. Zokolica and M. Freeling. 1986. The shrunken gene on chromosome 9 of Zea mays L. is expressed in various plant tissues and encodes an anaerobic protein. Mol. Gen. Genet. 205:461-468. Stepanenko, B.N. and A.V. Morozova. 1970. Possibility of cellulose biosynthesis from UDP-glucose in the cotton plant. Soviet Plant Physiology 1 7:250-255. Su, J-C. and J. Preiss. 1 978. Purification and properties of sucrose synthase from maize kernels. Plant Physiol. 61 :389-393. Sum, W.F. P.J. Rogers, I.D. Jenkins and R.D. Guthrie. 1980. Isolation of invertase from banana fruit (Musa cavendishii). Phytochem. 19:399-401. Sung, S.S., D-P. Xu, and C.C. Black. 1989. Identification of actively filling sucrose sinks. Plant Physiol. 89:1117-1121. Sung, S.S., D-P. Xu, CM. Galloway and C.C. Black. 1988. A reassessment of glycolysis and gluconeogenesis in higher plants. Physiol Plant. 72:650-654. Taliercio, E.W. and P.S. Chourey. 1989. Post-transcriptional control of sucrose synthase expression in anaerobic seedlings of maize. Plant Physiol. 90:1359-1364. Thorn, M. and A. Maretzki. 1990. Sugar transport in stalk tissue of sugarcane. Proc. Intl. Conf. Phloem Transport and Assimilate Compartmentation. Cognac, France, Aug. 19-24, 1990. p. 29. Thomas, P.S. 1980. Hybridization of denatured RNA and small DNA fragments transferred to nitrocellulose paper. Proc. Natl. Acad. Sci. (USA) 77 52015205.

PAGE 119

111 Towbin, H., T. Staehelin and J. Gerdon. 1979. Electrophoretic transfer of proteins from polyacrylamide gels to nitrocellulose sheets: Procedure and some applications. Proc. Natl. Acad. Sci. (USA) 76:4350-4354. Tsai, C.Y. 1974. Sucrose UDP glucosyltransferase of Zea mays endosperm. Phytochem. 13:885-891. Turner, J.F. and D.H. Turner. 1957. Physiology of pea fruits. Ill Changes in starch and starch phosphorylase in the developing seed. Aust. J. Biol. Sci. 10:302309. Vassey, T.L 1989. Light/dark profiles of sucrose phosphate synthase, sucrose synthase, and acid invertase in leaves of sugar beets. Plant Physiol. 89:347351. Vose, P.B. 1981. Potential use of induced mutants in crop plant physiology studies, pp. 159-181 in Induced Mutations-A Tool in Plant Research. Vienna, Austria: International Atomic Energy Agency. Walker, J.L and S.C. Huber. 1989. Regulation of sucrose phosphate synthase activity in spinach leaves by protein level and covalent modification. Planta 177:116-120. Webster, P.L and M. Henry. 1987. Sucrose regulation of protein synthesis in pea root meristem cells. Env. and Exp. Bot. 27:255-262. Weil, M. and T. Rausch. 1990. Cell wall invertase in tobacco crown gall cells. Plant Physiol. 94:1575-1581. Wenzler, H.C., G.A. Mignery, LM. Fisher and W.D. Park. 1989. Analysis of a chimeric class-l patatin-GUS gene in transgenic potato plants: High-level expression in tubers an sucrose-inducible expression in cultured leaf and stem explants. Plant Mol. Biol. 12:41-50. White, P.R. 1963. The Cultivation of Animal and Plant Cells. New York: Ronald Press. Williams, J.H.H. and J.F. Farrar. 1990. Control of barley root respiration by sucrose. Physiol. Plant. 79:259-266. Winkenbach, F. and P. Matile. 1970. Evidence for de novo synthesis of an invertase inhibitor protein in senescing petals of Ipomoea. Z. Pflanzenphysiol. 63:292-295.

PAGE 120

112 Wolosiuk, R.A. and H.G. Pontis. 1974a. Studies on sucrose synthetase kinetic mechanism. Arch. Biochem. Biophys. 165:140-145. Wolosiuk, R.A. and H.G. Pontis. 1974b. The role of sucrose and sucrose synthetase in carbohydrate plant metabolism. Mol. Cell Biochem. 4:1 1 5-1 23. Wright, K. and D.H. Northcote. 1972. Induced root differentiation on sycamore callus. J. Cell Sci. 11:319-337. Wyse, R. 1986. Sinks as determinants of assimilate partitioning: Possible sites for regulation, pp. 197-210 in J. Cronshaw, W.J. Lucus, R.T. Giaquinta, eds. Phloem Transport. New York: Alan R. Liss, Inc. Xie, Y. and R. Wu. 1989. Rice alcohol dehydrogenase genes: anaerobic induction, organ specific expression and characterization of cDNA clones. Plant Mol. Biol. 13:53-68. Xu, D-P., S.S. Sung, C.A. Alvarez and C.C. Black. 1986. Pyrophosphate-dependent sucrose metabolism and its activation by fructose 2,6-bisphosphate in sucrose importing plant tissues. Biochem. Biophys. Res. Comm. 141:440445. Xu, D-P., S.J.S. Sung, T. Loboda, P.P. Kormanik and C.C. Black. 1989. Characterization of sucrolysis via the uridine diphosphate and pyrophosphate-dependent sucrose synthase pathway. Plant Physiol. 90:635-642. Yelle, S., R.T. Chetelat, M. Dorais, J.W. DeVerna and A.B. Bennett. 1990. Sink metabolism in tomato fruit. IV. Genetic and biochemical analysis of sucrose accumulation. Plant Physiol. 95:1026-1035. Yelle, S., J.D. Hewitt, N.L Robinson, S. Damon and A.B. Bennett. 1988. Sink metabolism in tomato fruit III. Analysis of carbohydrate assimilation in a wild species. Plant Physiol. 87:737-740.

PAGE 121

BIOGRAPHICAL SKETCH Edwin Ralph Duke was bom in Bainbridge, Georgia on February 22, 1 960. He attended public school in Rossville, Georgia, and graduated from Macon Christian Academy, Macon, Georgia, in May 1978. In June 1980 he received his Associate of Science degree from Macon Junior College, Macon, Georgia, majoring in agriculture. He graduated from the University of Georgia in June 1982 with a Bachelor of Science in Agriculture degree, majoring in horticultural science. Edwin entered the University of Florida in August 1982 in the Department of Ornamental Horticulture and received his Master of Science degree from this department in May 1985. He transferred to the Department of Fruit Crops to pursue the Doctor of Philosophy degree. Edwin plans to receive his doctorate in August 1 991 . He will spend a short time as a postdoctoral research associate in the laboratory of Dr. Karen Koch at the University of Florida and will then move to Peoria, Illinois where he will be a postdoctoral research associate for the United States Department of Agriculture in the laboratory of Dr. Douglas Doehlert. He is a member of Phi Theta Kappa, Phi Kappa Phi and Gamma Sigma Delta honor societies and a member of the American Society for Horticultural Science and the American Society of Plant Physiologists. 113

PAGE 122

I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Caren E. Koch, Chair Professor of Horticultural Science I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Rebecca L. Darnell Assistant Professor of Horticultural Science I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Donald ft. Associate Professor of "Horticultural Science I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. QOuA (L c. L Curtis Hannah Professor of Horticultural Science I certify that I have read this study and that in my opinion it conforms to acceptable standards of scholarly presentation and is fully adequate, in scope and quality, as a dissertation for the degree of Doctor of Philosophy. Alice C. Harmon Assistant Professor of Botany

PAGE 123

This dissertation was submitted to the Graduate Faculty of the College of Agriculture and to the Graduate School and was accepted as partial fulfillment of the requirements for the degree of Doctor of Philosophy. August, 1991 )J^cuA Dean, Allege of Agria£jKure Dean, Graduate School


17
root Sh1 mRNA were elevated when sugar supplies were limited in culture. In
contrast, levels of Sus1 mRNA were elevated in response to increasing glucose
concentrations. They speculated that the effect of sugar levels on expression of
specific genes could prove to be an important control mechanism whereby plant
cells could react to cellular nutritional conditions. The Sh1 gene is also up-
regulated under anaerobic conditions (Springer et al., 1986); however, the
carbohydrate response of the Sh 1 gene appears to be distinct from its anaerobic
regulation (Koch et al., 1989). There is some doubt as to whether anaerobic
induction occurs at both the transcriptional and translational levels in maize
(McElfresh and Chourey, 1988). Taliercio and Chourey (1989) hypothesized that
the expression of anaerobically induced Sh1 transcripts are blocked at some step
beyond polyribosomal loading. However, other researchers have shown that
sucrose synthase in maize is anaerobically induced at the protein as well as the
gene level (Freeling and Bennett, 1985; Springer et al., 1986). Anaerobic induction
of sucrose synthase at both the gene and protein levels also has been
demonstrated in rice (Ricard et al, 1991) and Echinochloa phyllopogon (Mujer et
al., 1990).
Invertases
Invertases (E.C. 3.2.1.26, /3-D-fructofuranoside fructohydrolases) are widely
distributed in the plant kingdom and catalyze the following reaction:
Sucrose + H20
> Glucose + Fructose


16
synthetic direction but inhibited the cleavage reaction. UDP was found to be a
strong inhibitor of the synthetic reaction at 10 mM (70-80% inhibition), but the
inhibition could be reversed by the addition of Mg2+ (de Fekete and Cardini, 1964).
UDP was also found to inhibit the degradative reaction as a competitive inhibitor
for UDPG (Wolosiuk and Pontis, 1974a). Inhibition by other NDPs was very weak.
UTP (4 mM) caused a slight inhibition of synthetic activity, but caused an 80%
inhibition of the degradative reaction (Tsai, 1974). Echeverra and Humphreys
(1985), however, found that UDP and UTP within the cytosolic range (< 4 mM)
both had little or no effect on sucrose synthase in the synthetic direction. UDPG
was able to inhibit the cleavage reaction by 13% at 10 mM, but tissue
concentrations were generally below this level (Echeverra and Humphreys, 1985),
with the effect on the synthetic reaction minimal. Wolosiuk and Pontis (1974a)
found that UDPG could function as a competitive inhibitor for UDP in the sucrose
synthase synthesis reaction.
Carbohydrates also have been found to inhibit sucrose synthase activity in
vitro. Fructose was found to function as a competitive inhibitor of sucrose in the
cleavage reaction (Pridham et al., 1969; Doehlert, 1987). Sucrose had no
inhibitory effects on sucrose synthase activity at saturating levels of fructose and
UDPG (Echeverra and Humphreys, 1985), but glucose at 100 mM inhibited
sucrose synthesis by 63%-70% and inhibited sucrose cleavage 86%-93%.
Expression of sucrose synthase genes also respond to carbohydrate availability.
Koch and McCarty (1988,1990; Koch et al., 1989) have shown that levels of maize


61
the effects of varying carbohydrate conditions on the Sus1 gene and its sucrose
synthase gene product free from the confounding effects of Sh 1.
Materials and Methods
Maize seed (Zea mays L, \N22.sh1) were primed for 6 days at 10 C with a
water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I'1 captan
(Bodsworth and Bewley, 1981). Seeds were then rinsed with water, soaked for 20
min in 1.05% (v/v) sodium hypochlorite and rinsed again for 20-30 min with ca. 5
liters of water. Germination took place in the dark at 18 C on moist filter paper in
covered glass pans. Continuous airflow was provided (1 liter min-7) throughout
the germination period with 40% 02 supplied during the final 48 h. At the end of
7 days, 1 cm primary root tips were excised under a sterile transfer hood.
Excised root tips (ca. 750 mg per treatment) were incubated in 100 ml side-
arm flasks containing 50 ml of sterile Whites basal salt mixture (White, 1963)
supplemented with a range of glucose (0, 0.2, 0.5, 2.0, and 4.0%). Aerobic
conditions were maintained during 24 h incubations in the dark at 18 C by slow
agitation on a rotary shaker (125 rpm) and by an airflow 40% 02 (1 I'1) through an
airstone in each flask. Experiments were terminated by twice rinsing root tips in
sterile water, blotting excess moisture and freezing them in liquid N2.
Responses of root tips to incubation in 0% vs 2.0% glucose were examined
after 16, 24, or 48 h. Effects of treatment reversals at 16 h were also studied by
switching roots from 0% glucose treatments to 2.0% glucose and vice versa, then
continuing incubations for a total of 48 h.


99
Douce, R., R. Bligny, D. Brown and A-J. Dome. 1990. Biochemical changes during
sucrose deprivation in higher plants, pp. 167-188 in M.J. Ernes, ed.
Compartmentation of Plant Metabolism in Non-photosynthetic Tissues.
Cambridge, U.K.: Cambridge University Press.
Duke, E.R., D.R. McCarty, L.C. Hannah and K.E. Koch. 1990. A rapid radiometric
assay for sucrose synthase. Plant Physiol. 93S:117.
Echeverra, E. and T. Humphreys. 1984. In vitro and in vivo inhibition of sucrose
synthesis by sucrose. Phytochem. 23:2727-2731.
Echeverra, E. and T. Humphreys. 1985. Glucose control of sucrose synthase in
the maize scutellum. Phytochem. 24:2851-2855.
Echt, C.S. and P.S. Chourey. 1985. A comparison of two sucrose synthetase
isozymes from normal and shrunken-1 maize. Plant Physiol. 79:530-536.
Edelman, J. and M.A. Hall. 1965. Development of invertase and ascorbate oxidase
activities in mature storage tissue of Helianthus tuberosus. Biochem. J.
95:403-410.
Edwards, J. and T. ap Rees. 1986a. Sucrose partitioning in developing embryos
of round and wrinkled varieties of Pisum sativum. Phytochem. 25:2027-
2032.
Edwards, J. and T. ap Rees. 1986b. Metabolism of UDP-glucose by developing
embryos of round and wrinkled varieties of Pisum sativum. Phytochem.
25:2033-2040.
Farrar, J.F. 1985. The respiratory source of C02. Plant, Cell Env. 8:427-438.
Faye, L. and C. Berjonneau. 1979. Evidence for the glycoprotein nature of radish
/3-fructosidase. Biochemie 61:330-332.
Faye, L. C. Berjonneau and P. Rollin. 1981. Studies on /3-fructosidase from radish
seedlings. I. Purification and partial characterization. Plant Sci. Letters 22:77-
87.
Fieuw, S. and J. Willenbrink. 1987. Sucrose synthase and sucrose phosphate
synthase in sugar beet plants (Beta vulgaris ssp. altissima). J. Plant Physiol.
131:153-162.
Fluhr, R., C. Kuhlemeier, F. Nagy and N-H. Chua. 1986. Organ-specific and light-
induced expression of plant genes. Science 232:1106-1112.


111
Towbin, H.t T. Staehelin and J. Gerdon. 1979. Electrophoretic transfer of proteins
from polyacrylamide gels to nitrocellulose sheets: Procedure and some
applications. Proc. Natl. Acad. Sci. (USA) 76:4350-4354.
Tsai, C.Y. 1974. Sucrose UDP glucosyltransferase of Zea mays endosperm.
Phytochem. 13:885-891.
Turner, J.F. and D.H. Turner. 1957. Physiology of pea fruits. Ill Changes in starch
and starch phosphorylase in the developing seed. Aust. J. Biol. Sci. 10:302-
309.
Vassey, T.L. 1989. Light/dark profiles of sucrose phosphate synthase, sucrose
synthase, and acid invertase in leaves of sugar beets. Plant Physiol. 89:347-
351.
Vose, P.B. 1981. Potential use of induced mutants in crop plant physiology
studies, pp. 159-181 in Induced Mutations-ATool in Plant Research. Vienna,
Austria: International Atomic Energy Agency.
Walker, J.L. and S.C. Huber. 1989. Regulation of sucrose phosphate synthase
activity in spinach leaves by protein level and covalent modification. Planta
177:116-120.
Webster, P.L. and M. Henry. 1987. Sucrose regulation of protein synthesis in pea
root meristem cells. Env. and Exp. Bot. 27:255-262.
Weil, M. and T. Rausch. 1990. Cell wall invertase in tobacco crown gall cells. Plant
Physiol. 94:1575-1581.
Wenzler, H.C., G.A. Mignery, L.M. Fisher and W.D. Park. 1989. Analysis of a
chimeric class-l patatin-GUS gene in transgenic potato plants: High-level
expression in tubers an sucrose-inducible expression in cultured leaf and
stem explants. Plant Mol. Biol. 12:41-50.
White, P.R. 1963. The Cultivation of Animal and Plant Cells. New York: Ronald
Press.
Williams, J.H.H. and J.F. Farrar. 1990. Control of barley root respiration by
sucrose. Physiol. Plant. 79:259-266.
Winkenbach, F. and P. Matile. 1970. Evidence for de novo synthesis of an
invertase inhibitor protein in senescing petals of Ipomoea. Z.
Pflanzenphysiol. 63:292-295.


110
Sharma, K.P. and I.S. Bhatia. 1980. Sucrose metabolism in Sorghum vulgare at
ripening. Physiol. Plant. 48:470-476.
Sheen, J. 1990. Metabolic repression of transcription in higher plants. Plant Cell
2:1027-1038.
Silvius, J.E. and F.W. Snyder. 1979. Comparative enzymatic studies of sucrose
metabolism in the taproots and fibrous roots of Beta vulgaris L. Plant
Physiol. 54:1070-1073.
Springer, B., W. Werr, P. Starlinger, D.C. Bennett, M. Zokolica and M. Freeling.
1986. The shrunken gene on chromosome 9 of Zea mays L. is expressed
in various plant tissues and encodes an anaerobic protein. Mol. Gen. Genet.
205:461-468.
Stepanenko, B.N. and A.V. Morozova. 1970. Possibility of cellulose biosynthesis
from UDP-glucose in the cotton plant. Soviet Plant Physiology 17:250-255.
Su, J-C. and J. Preiss. 1978. Purification and properties of sucrose synthase from
maize kernels. Plant Physiol. 61:389-393.
Sum, W.F. P.J. Rogers, I.D. Jenkins and R.D. Guthrie. 1980. Isolation of invertase
from banana fruit (Musa cavendishii). Phytochem. 19:399-401.
Sung, S.S., D-P. Xu, and C.C. Black. 1989. Identification of actively filling sucrose
sinks. Plant Physiol. 89:1117-1121.
Sung, S.S., D-P. Xu, C.M. Galloway and C.C. Black. 1988. A reassessment of
glycolysis and gluconeogenesis in higher plants. Physiol Plant. 72:650-654.
Taliercio, E.W. and P.S. Chourey. 1989. Post-transcriptional control of sucrose
synthase expression in anaerobic seedlings of maize. Plant Physiol.
90:1359-1364.
Thom, M. and A. Maretzki. 1990. Sugar transport in stalk tissue of sugarcane.
Proc. Inti. Conf. Phloem Transport and Assimilate Compartmentation.
Cognac, France, Aug. 19-24, 1990. p. 29.
Thomas, P.S. 1980. Hybridization of denatured RNA and small DNA fragments
transferred to nitrocellulose paper. Proc. Natl. Acad. Sci. (USA) 77:5201-
5205.


46
minute incubation. However, chymotrypsin, also a serine proteinase, and papain,
a cysteine proteinase, had no effect. In the current study, PMSF, a serine
proteinase inhibitor, gave an increase in initial measurements but not did not
prevent the observed short-term loss of activity with time. Echt and Chourey
(1985) observed that PMSF did not stop the loss of activity of sucrose synthase
from maize endosperm during long term storage. No other specific proteinase
inhibitor affected stability of the enzyme from maize root tips. In addition to the
specific proteinase inhibitors, casein and BSA were included in some extractions.
Due to caseins complex composition and random structure, it undergoes
proteolysis with all the known proteolytic enzymes (Reimerdes and Klostermeyer,
1976). Casein increased initial measurements and stabilized activity with time
(Figure 3-3A & B). BSA, however, was much less effective. Casein has also been
found to preserve the longevity of sucrose synthase extracted from sugar cane (S.
Lingle, USDA-ARS, Westlaco, TX, personal communication). Casein (0.75-3.0%)
has also been shown to effectively stabilize and increase the initial activity
measurements of sucrose phosphate synthase (Raghuveer and Sicher, 1987).
Addition of casein is not always feasible, however, especially in instances where
accurate quantification of total tissue protein is important.
Another possibility for the regulation of sucrose synthase is through
phosphorylation. Many enzymes undergo reversible phosphorylation as a means
regulating activity (Bennett, 1984). Increased inorganic phosphate levels added
to buffers used for enzymatic extraction provide a substrate for protein kinase


51
Regulation by sugar concentration may prove to be an important control
mechanism whereby plant cells are able to react to cellular carbohydrate status.
Materials and Methods
Maize seed (Zea mays L, NK 508) were primed for 6 days at 10 C at a
water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I'1 captan
(Bodsworth and Bewley, 1981). Seeds were subsequently rinsed free of PEG,
soaked for 20 min in 1.05% (v/v) sodium hypochlorite and rinsed for 20-30 min
with ca. 5 I of water. Seeds were germinated in the dark at 18 C on moist filter
paper in covered glass pans. Continuous airflow was provided (1 liter min"1)
throughout the germination period with 40% 02 supplied during the final 48 h. At
the end of 7 days, 1 cm primary root tips were excised under a sterile transfer
hood.
Excised root tips (ca. 750 mg per treatment) were incubated in 100 ml side-
arm flasks containing 50 ml of sterile Whites basal salt mixture (White, 1963)
supplemented with a range of glucose (0, 0.2, 0.5, 2.0, and 4.0%). Aerobic
conditions were maintained during 24 h incubations in the dark at 18 C by slow
agitation on a rotary shaker (125 rpm) and an airflow of 40% 02 (1 I min'1) through
an airstone in each flask. Experiments were terminated by twice rinsing in sterile
water, blotting excess moisture and freezing them in liquid N2.
Sucrose synthase activity was determined using a rapid radiometric
procedure developed to circumvent enzyme instability previously observed upon
extraction from maize root tips (Chapter 3).


36
Table 3-1. Effect of enzyme protectants on activity of sucrose synthase from maize
root tips assayed five minutes after extraction.
Protectant (concentration)enhancement of control
PVP-40 (5 %)
%
-6
PEG-20,000 (2% w:v)
+6
BSA (2% w:v)
+6
Caproic acid (2 mM)
0
Pepstatin A (1 mM)
-5
Leupeptin (1 mM)
0
PMSF (1 mM)
+46
Casein (2% w:v)
+ 15
Note: Each protectant was included in the buffer used for extraction and
equilibration of desalting columns. PVP-40, PEG-20,000 and BSA were utilized to
protect against phenolic compounds; PVPP was included in each extraction.
Representative inhibitors of proteinase classes were: pepstatin A (aspartic),
leupeptin (cysteine), EGTA (metallo) included in each extraction, caproic acid
(serine) and PMSF (serine). Casein was included as a general, non-specific
proteinase inhibitor.


Table 6-1. Soluble and insoluble acid invertase activity in sequential 2 mm segments of primary
roots of 5 to 6 day-old seedlings from 1 hybrid and 2 inbred lines of Zea mays.
Root
Solubley
Insoluble
Segment
Oh 43
NK 508
W22 Oh 43
NK 508 W22
/mol glucose g'1FW h'1
0-2 mm
2-4 mm
4-6 mm
6-8 mm
8-10 mm
0.2 0.2W
43.8 5.2
32.8 2.2
20.6 5.0
1.4 0.9
40.2 3.7 z
28.6 3.1
15.4 4.6
0.9 0.5
3.8 0.5 3.6 0.8
3.1 1.9 2.7 1.5
z-not detectable
y-Soluble activity is expressed per unit FW to allow comparison with insoluble activity.
"--Activity in root tips from axenic culture was 0.0 0.0 /mol g'1FW h'1; thus microorganisms are a
likely source of the residual activity in ca. 1 of 3 assays.
Each value represents the mean of 3 separate samples SEM.


55
sucrose conditions. However, Salanoubat and Belliard (1989) found that increased
sucrose levels promoted genes encoding sucrose synthase in potato.
The possibility also exists that protein fluctuations did not occur after 24 h
incubation. However, the shifts in protein localization noted under the same
conditions (K. Nolte, University of Florida, unpublished data) indicate that some
protein level changes did occur. Spatial separation within root tissue also could
explain the differential response of Sh1 and Sus 1 observed at gene level without
a concomitant change in total enzyme activity. The distribution of sucrose
synthase isozymes has been shown to be developmentally regulated, and
changes during kernel development (Heinlein and Starlinger, 1989). Chen and
Chourey (1989) have reported that expression of sucrose synthase genes is
spatially and/or temporally separated in endosperm cells but not in root cells.
However, Rowland et al. (1989) demonstrated tissue specific localization of both
sucrose synthase genes and isozymes in roots undergoing anaerobic stress. K.
Nolte (University of Florida, unpublished data) has shown that shifts in sucrose
synthase protein localization occur in maize root tips under carbohydrate depleted
and carbohydrate sufficient conditions. Increases of one isozyme in a particular
tissue within the root along with decreases of the other in a different tissue would
not be apparent at the level of total root sucrose synthase activity. The lack of
significant differences in sucrose synthase activity of wildtype maize roots under
carbohydrate sufficient and depleted condition, therefore, does not demonstrate
that differences evident at the gene level are not also event at the translational


root but not in otherwise morphologically identical laterals from adventitious roots.
In contrast, sucrose synthase was active in all roots and theoretically provided the
only available avenue for sucrose degradation in primary root tips of Oh 43.
IX


CHAPTER 1
INTRODUCTION
Sucrose metabolism is important to the majority of plant species because
of the nearly ubiquitous role of this sugar in growth and development. Initial
breakdown of sucrose can be catalyzed by either invertase or the reversible
enzyme sucrose synthase.
Gene responses to changes in carbohydrate availability have been reported
for mammalian cells (Lin and Lee, 1984) and yeasts and bacteria (Carlson, 1987;
Schuster, 1989). Evidence has also been presented that the transcriptional activity
of promoters of photosynthetic genes from maize protoplasts are repressed and
coordinated by sugars (Sheen, 1990). Expression of the genes encoding the
sucrose synthase isozymes also have been shown to respond to carbohydrate
levels (Koch and McCarty, 1988; Koch et al., 1989). Northern blot analysis of
mRNA from wildtype maize roots showed levels of one isozyme gene (Sh 1) was
up-regulated in response to carbohydrate depletion whereas the other (Sus)
responded positively only when sugar availability was elevated. This sensitivity of
genes for a key enzyme in sugar metabolism may prove to be an important control
mechanism whereby plant cells are able to react to cellular nutritional conditions.
The importance of invertase, the other enzyme of sucrose catabolism in
plants, to root metabolism is unclear. Invertases are especially active in tissues
1


SUCROSE SYNTHASE AND INVERTASE IN MAIZE ROOTS DIFFERING
IN CARBOHYDRATE STATUS AND/OR GENETIC COMPOSITION
By
EDWIN RALPH DUKE
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
1991


45
action of sucrose synthase can also be coupled to that of UDP-
glucopyrophosphorylase (Xu et al., 1986; Sung et al., 1989).
Radioactive assays of sucrose synthase in the cleavage direction measure
the incorporation of 14C-glucose into UDP-glucose from 14C-sucrose (Delmer, 1972;
Su and Preiss, 1978). These procedures are among the most sensitive of assays
for sucrose synthase (Avigad, 1982). The sugar nucleotide formed can be
separated from the excess 14C-sucrose by paper chromatography (Wolosiuk and
Pontis, 1974a) or by ion exchange paper (Delmer, 1972a and b; Su and Preiss,
1978). The current procedure utilizes the sensitivity of a radiometric assay along
with reduced time from extraction to assay termination and results in a method
suitable for time-labile extracts from small tissue samples.
Instability of sucrose synthase was further characterized using this sensitive
method in an attempt to better define factors affecting the activity of this key
enzyme in vitro. Sucrose synthase is a sulfhydryl enzyme and is sensitive to
inhibition by phenolics and oxidized polyphenols (Pontis, 1977). Typical effectors
of activity reduction examined during the present study showed that phenolic
compounds did not appear to be the primary cause of the sucrose synthase
instability observed. No phelolic protectant was able to preserve sucrose synthase
activity over time.
Sucrose synthase has been shown to be sensitive to serine proteinases
(Wolosiuk and Pontis, 1974b). In their study, trypsin caused a 70% decline in the
degradative reaction and a 30% reduction in the synthetic reaction after a 15


25
inhibitor. Bracho and Whitaker (1990b) also found a positive correlation between
levels of inhibitor and invertase. The possibility therefore exists that this interaction
plays a regulatory role in sucrose breakdown (Akazawa and Okamoto, 1980;
Avigad, 1982).
Alkaline invertase is generally considered to be cytoplasmic. It is only
recovered from the soluble fraction of homogenates and has a pH optimum near
neutral. Both findings support its internal localization. Km values of alkaline
invertase for sucrose are slightly higher than for acid invertase, generally 9 to 25
mM. Alkaline invertase hydrolyzes raffinose very poorly (< 7% of the rate of
sucrose breakdown). Morell and Copeland (1984) found that stachyose (0.1 M)
also was hydrolyzed by alkaline invertase but much less efficiently than sucrose
(1.5% of the rate of sucrose). Both raffinose and stachyose are polysaccharides
containing a fructose moiety. Morell and Copeland (1984) also found that
cellobiose, gentiobiose, maltose, turanose, lactose, melezitose, trehalose, a-methyl-
D-glucopyranoside and /3-methyl-D-glucopyranoside (all at 0.1 M) were resistant
to degradative action by alkaline invertase. None of these sugars contain a
fructose moiety, further confirming the specificity of invertase for the fructofuranose
moiety of sucrose.
Alkaline invertase from potato tubers was inhibited only slightly by glucose
(Matsushita and Uritani, 1974); glucose-6-phosphate also had a slight inhibitory
effect. Fructose (15 mM) competitively inhibited soybean nodule alkaline invertase
by 50% (Morell and Copeland, 1984); glucose (5 mM) inhibited activity by 7%.


58
Overall, enhanced expression of Sus1 was evident at the mRNA, protein and
enzyme levels.
Introduction
Changes in gene expression by carbohydrates have been documented as
mechanisms by which bacteria and yeasts respond to changes in their nutrient
status (Carlson, 1987; Schuster, 1989). Glucose-responsive genes have also been
described in mammalian cells (Lin and Lee, 1984). In addition, Sheen (1990) has
presented evidence that the transcriptional activity of promoters of seven
photosynthetic genes from maize protoplasts are repressed and coordinated by
sugars. Regulation of this type in higher plants could have important implications
for carbohydrate allocation and utilization.
Sucrose and its metabolic products are important to almost all plant species
because of the nearly universal role of this sugar in growth and development
(Avigad, 1982). Initial breakdown of sucrose can be catalyzed by either invertase
or the reversible enzyme sucrose synthase. Recently, the gene encoding the
shrunken-1 isozyme of sucrose synthase in maize has been shown to be
responsive to carbohydrate levels (Koch and McCarty, 1988; Koch et al., 1989).
Gel blot analysis of shrunken-1 mRNA showed levels were elevated in response
to carbohydrate depletion. This may prove to be an important control mechanism
whereby plant cells are able to react to cellular nutritional conditions. The Sh1
gene has also been shown to be regulated by anaerobic conditions (Springer, et
al., 1986); however, effects on this gene by altered carbohydrate status are distinct


101
Gupta, M., P.S. Chourey, B. Burr and P.E. Still. 1988. cDNAs of two non-allelic
sucrose synthase genes in maize: cloning, expression, characterization and
molecular mapping of the sucrose synthase-2 gene. Plant Mol. Biol. 10:215-
224.
Hannah, L.C. and D.R. McCarty. 1988. Mature pollen contains transcripts of the
constitutive sucrose synthase (Css) gene. Maize Gen. Coop. Newsletter
62:59-60.
Hatch, M.D. and K.T. Glasziou. 1963. Sugar accumulation cycle in sugar cane II.
Relationship of invertase activity to sugar content and growth rate in storage
tissue of plants grown in controlled environments. Plant Physiol. 38:344-348.
Hatch, M.D., J.A. Sacher and K.T. Glasziou. 1963. Sugar accumulation cycle in
sugar cane I. Studies on enzymes of the cycle. Plant Physiol. 38:338-343.
Hatton, T., S. Nakagawa and K. Nakamura. 1990. High-level expression of
tuberous root storage protein genes of sweet potato in stems of plantlets
grown in vitro on sucrose medium. Plant Mol. Biol. 14:595-604.
Hawker, J.S. 1985. Sucrose, pp. 1-51 in P.M. Dey, ed. Biochemistry of Storage
Carbohydrates in Green Plants. London: Academic Press.
Hawker, J.S. and M.D. Hatch. 1965. Mechanism of sugar storage by mature stem
tissue of sugar cane. Physiol Plant. 18:444-453.
Hayward, H.E. 1938. The Structure of Economic Plants. New York: The Macmillan
Co.
Heinlein, M. and P. Starlinger. 1989. Tissue- and cell-specific expression of the two
sucrose synthase isoenzymes in developing maize kernels. Mol. Gen.
Genet. 215:441-446.
Hellebust, J.A. and D.F. Forward. 1962. The invertase of the corn radicle and its
activity in successive stages of growth. Can. J. Bot. 40:113-126.
Hendrix, D.L. 1990. Carbohydrates and carbohydrate enzymes in developing
cotton ovules. Physiol. Plant. 78:85-92.
Hendrix, D.L. and S.C. Huber. 1986. Diurnal fluctuations in cotton leaf carbon
export, carbohydrate content, and sucrose synthesizing enzymes. Plant
Physiol. 81:584-586.


54
for substrates (Echt and Chourey, 1985), and both are present in extracts from
wildtype maize root tips (Chourey et al., 1986). Therefore the total amount of
measured sucrose synthase activity would be due to a combined complement of
sucrose synthase protein. Despite close homology, however, they have been
shown to be distinctive proteins encoded by separate genes (Chourey, 1981; Echt
and Chourey, 1985). The two proteins also are distinct in their localization within
the maize plant. The protein encoded by Sh 1 is primarily located in the
endosperm (Chourey and Nelson, 1976; Chen and Chourey, 1989), in the root
(Springer et al., 1986; Chourey et al., 1986) and in etiolated shoot (Springer et al.,
1986). The Sus 1 encoded protein is generally distributed throughout the plant
(Chourey, 1981; Echt and Chourey, 1985; Chourey et al., 1988).
Carbohydrate responsive proteins have been identified in roots of pearl
millet (Baysdorfer and Van der Woude, 1988). Webster and Henry (1987) have
also identified an unknown protein with a molecular weight similar to that of the
subunits of sucrose synthase in pea root meristem cells undergoing sugar
starvation. This protein, however, has yet to be positively identified. Initial findings
by Koch and coworkers (Koch and McCarty, 1988, 1990; Koch et al., 1989)
indicated that the Sh 1 gene of maize was stimulated by low carbohydrate
conditions and down-regulated under carbohydrate sufficient conditions. The Sus 1
gene responded in an inverse manner. Maas and co-workers (Maas et al., 1990)
demonstrated that the promoter from the Sh 1 gene was repressed by high


50
important implications for the control of carbohydrate distribution and utilization.
Sucrose synthase is considered to have a key function in the allocation of sucrose
to various plant organs, and plant carbohydrate status could function as a means
of coarse regulation for activity of this enzyme.
Sucrose metabolism is important to the majority of plant species because
of the nearly ubiquitous role of this sugar in phloem transport to growing and
developing plant parts (Avigad, 1982). Two enzymes can catalyze the initial
breakdown of sucrose, invertase or the reversible enzyme sucrose synthase.
Recently, expression of the gene encoding the shrunken-1 isozyme of sucrose
synthase in maize has been shown to be sensitive to carbohydrate levels (Koch
and McCarty, 1988; Koch et al., 1989). Northern blot analysis of shrunken-1
mRNA showed levels were elevated in response to carbohydrate depletion. This
regulation is distinct from the previously characterized anaerobic induction
(Springer et al., 1986; Koch and McCarty, 1988). Although the anaerobic induction
of Sh1 has received considerable attention in several systems, questions remain
regarding the extent to which transcription and translation are synchronized under
these conditions. The anaerobic induction in maize has been reported to occur
only at the transcriptional level without concomitant changes in protein levels
(McElfresh and Chourey, 1988; Taliercio and Chourey, 1989). However, translation
of anaerobically induced sucrose synthase mRNA in rice (Ricard et al., 1991) has
been demonstrated. The present work examines enzyme-level responses to
changes in root carbohydrate status known to alter levels of Sh1 and Sus1 mRNA.


TABLE OF CONTENTS
ACKNOWLEDGEMENTS ii
LIST OF TABLES v
LIST OF FIGURES vi
ABSTRACT vii
CHAPTERS
1 INTRODUCTION 1
2 REVIEW OF THE LITERATURE 4
Sucrose Metabolism 4
Sucrose Synthase 6
Invertases 17
Use of Mutants in Physiological Research 26
3 INSTABILITY OF SUCROSE SYNTHASE FROM ROOT
TIPS: CHARACTERIZATION AND STABILIZATION 29
Abstract 29
Introduction 30
Materials and Methods 31
Results 34
Discussion 41
4 SUCROSE SYNTHASE ACTIVITY IN WILDTYPE MAIZE
ROOT TIPS RESPONDING TO ALTERED
CARBOHYDRATE STATUS 49
Abstract 49
Introduction 49
Materials and Methods 51
iii


70
Table 5-2. Sucrose synthase activity in mutant maize (W22:s/7i) root tips
incubated in media containing in 0 or 2.0% glucose for various time periods.
Sucrose synthase activity
16h 24h
48h
16h + 32h
Intact
+ +
+
-/+
+/-
0.12
(/mol sucrose g'1
0.22 0.28 0.22 0.29
protein h1)
0.18 0.40
0.31
0.40
Note: 0 and 2.0% glucose represented by and +, respectively.


65
% glucose
Intact 0 0.2 0.5 2.0 4.0
Expt. 1
Expt. 2
Figure 5-2. Protein gel blot of Sus 1 encoded sucrose synthase from maize roots
incubated in a range of glucose concentrations for 24 hours. Data from
Expts. 1 and 2 were obtained from the same set of roots sampled for RNA
analyses shown in Figure 5-1.


Message levels of genes encoding sucrose synthase isozymes in maize
have been shown to respond to tissue carbohydrate status, thus the effects of
such changes were examined at the level of enzyme activity. Total sucrose
synthase activity from roots of wildtype plants showed little difference in extracts
from root tips incubated for 24 h in a range of glucose levels. This activity,
however, is the combined contribution of two isozymes whose genes are
responding differentially to experimental conditions.
The sh 1 mutant of maize was used to study expression of the Sus1 gene
for sucrose synthase in response to sugar availability because this mutant has only
one gene (Sus1) for sucrose synthase and provides a system uncomplicated by
the presence of the second isozyme (Sh1). Sus1 mRNA increased 5-fold when
incubated for 24 h in 2.0% glucose compared to 0 or 0.2% glucose. Levels of Sus
protein were slightly elevated with increasing sugar levels. Enzyme activity was
elevated 2-fold under the same conditions. A study of time-course and treatment
reversals showed that changes in mRNA or protein were not evident until 24 h and
indicated that the response to carbohydrate level had been initiated within 16 h.
Overall, enhanced expression of Sus 1 was evident at the mRNA, protein and
enzyme levels.
An organ-specific invertase deficiency affecting only the primary root system
also was characterized in the Oh 43 maize inbred. Substantial acid invertase
activity was evident in extracts of all tissues tested except the primary root system
of Oh 43. This deficiency was also evident in lateral roots arising from the primary
VIII


Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
SUCROSE SYNTHASE AND INVERTASE IN MAIZE ROOTS DIFFERING
IN CARBOHYDRATE STATUS AND/OR GENETIC COMPOSITION
By
Edwin Ralph Duke
August, 1991
Chairperson: Dr. Karen E. Koch
Major Department: Horticultural Science
The extent to which sucrose is transported in phloem of higher plant
species necessitates its effective metabolism by non-photosynthetic cells.
However, only two enzymes can catalyze sucrose breakdown in these instances,
invertase and sucrose synthase (a reversible enzyme). Specific molecular, genetic
and physiological factors affecting these enzymes were investigated in the root tips
of maize. A radiometric assay was first developed for sucrose synthase to
circumvent the rapid decline of sucrose synthase activity in vitro. Further,
characterization of in vitro instability indicated that activity decline was not
associated with any detectable proteolytic degradation, charge alteration, or
subunit separation and that inhibition of activity by inorganic phosphate suggested
possible phosphorylation of this enzyme.
VII


CHAPTER 4
SUCROSE SYNTHASE ACTIVITY IN WILD-TYPE MAIZE ROOT TIPS RESPONDING
TO ALTERED CARBOHYDRATE STATUS
Abstract
The two genes encoding sucrose synthase isozymes in maize (Sh1 and
Sus1) have been shown to respond to altered tissue carbohydrate status in root
tips; Sh 1 expression is favored by carbohydrate depletion whereas Sus 1 is up-
regulated when sugars are plentiful (Koch and McCarty, 1988, 1990; Koch et al.,
1989). Response at the level of enzyme activity was tested in the present study
by assaying sucrose synthase activity in excised maize root tips after 24 h of
incubation in a range of glucose concentrations. Little change was evident at the
level of total sucrose synthase activity; however, this represented the collective
responses of different isozymes and tissue types.
Introduction
Systems for changes in gene expression in response to altered
carbohydrate conditions have been reported for mammalian cells (Lin and Lee,
1984) and in bacteria and yeasts (Carlson, 1987; Schuster, 1989). Recently, seven
photosynthetic genes in maize protoplasts have been shown to be repressed and
coordinated by sugars (Sheen, 1990). Regulation in higher plants could have
49


103
Khayat, E. and N. Zieslin. 1987. Effect of night temperature on the activity of
sucrose phosphate synthase, acid invertase and sucrose synthase in
source and sink tissues of Rosa hybrida cv Golden Times. Plant Physiol.
84:447-449.
Kidby, D.K. 1966. Activation of plant invertase by inorganic phosphate. Plant
Physiol. 41:1139-1144.
Koch, K.E., W.T. Avigne and D.R. McCarty. 1989. Differential response of 2
sucrose synthase genes to decreased sugar supply. Plant Physiol. 89S:7.
Koch, K.E. and D.R. McCarty. 1988. Induction of sucrose synthase by sucrose
depletion in maize root tips. Plant Physiol. 86S:35.
Koch, K.E. and D.R. McCarty, 1990. Optimum conditions for sugar-modulated
expression of sucrose synthase genes in maize root tips. Plant Physiol.
93:8.
Koch, K.E., C-L. Tsui, L.E. Schrader and O.E. Nelson. 1982. Source-sink relations
in maize mutants with starch-deficient endosperms. Plant Physiol. 70:322-
325.
Kothari, S.K., H. Marschner and E. George. 1990. Effect of VA mycorrhizal fungi
and rhizosphere microorganisms on root and shoot morphology, growth
and water relations in maize. New Phytol. 116:303-312.
Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the
head of the bacteriophage T4. Nature 227:680-685.
Leloir, L.F. and C.E. Cardini. 1953. The biosynthesis of sucrose. J. Amer. Chem.
Soc. 75:4118-4123.
Leloir, L.F. and C.E. Cardini. 1955. The biosynthesis of sucrose phosphate. J. Biol.
Chem 214:157-165.
Lin, A.Y. and A.S. Lee. 1984. Induction of two genes by glucose starvation in
hamster fibroblasts. Proc. Natl. Acad. Sci. (USA) 81:988-992.
Lin, W., M.R. Schmitt, W.D. Hitz and R.T. Giaquinta. 1984. Sugar transport in
isolated corn root protoplasts. Plant Physiol. 76:894-897.
Lingle, S.E. 1987. Aspects of sucrose transport in stem parenchyma of sweet
sorghum. Plant Physiol. 83S:142.


53
Table 4-1. Total sucrose synthase activity In wlldtype maize root tips incubated in
a range of glucose concentrations for 24 hours.
intacty
% glucose
0
0.2 0.5 2.0 4.0
(/mol sucrose mg'1 protein h'1)
Expt. 1
0.9
1.2
0.9
1.6
0.9
1.5
Expt. 2
0.5
0.5
0.4
0.6
0.6
0.6
Expt. 3
1.0
0.7
0.8
1.3
0.6
0.7
Mean
0.8
0.8
0.7
1.1
0.7
0.9
S.E.M.
0.2
0.2
0.2
0.3
0.1
0.
ylntact refers to root tips quick frozen in liquid N2 immediately after excision.


CHAPTER 2
REVIEW OF THE LITERATURE
Sucrose Metabolism
Sucrose is the major transported sugar in the majority of higher plant
species. The major roles of sucrose in higher plants include its function as both
a translocatable form of carbon and as a vacuolar storage compound (Hawker,
1985). It is a non-reducing sugar made up of a glucose (a-D-glucopyranose and
fructose (/?-D-fructofuranose) joined by an a-1,2 linkage. For metabolic utilization,
it must be broken down into its component monosaccharides or their derivatives.
Only two enzymatic systems for sucrose breakdown are known in higher plant
tissues. Sucrose can be hydrolyzed by the action of invertase or cleaved by
sucrose synthase working in the degradative direction. The free energy of sucrose
hydrolysis is nearly equal to that of the y phosphoryl group of ATP (AG= -7.0
kcal/mol and -7.3 kcal/mol, respectively) (Neufeld and Hassid, 1963). This is much
greater than the free energy of most other glycosidic bonds (Avigad, 1982).
Cleavage by sucrose synthase retains the bond energy in the a-glucosyl bond of
UDP-glucose. In contrast, hydrolysis by invertase conserves none of the free
energy in the bond.
4


90
development because the root system would consist solely of a primary root at
this time. Later in seedling development, adventitious or seminal roots rapidly take
over a dominant role, leaving the primary root with little or no essential function
(Hayward, 1938). In the field, lines having Oh 43 as a progenitor have been
observed to suffer from poor emergence rates in damp soils (B. Martin, Pioneer
Seed, personal communication). However, conclusive evidence of a physiological
effect will require generation of and analysis of isogeneic lines.
In conclusion, the deficiency described here will be useful for investigation
into the regulation and physiological function of root invertase. Because the
primary root system in maize is nonessential, invertase deficient roots can be
studied without deleterious effects on the overall physiology of the plant. In
addition, this deficiency is significant because it reveals an unexpected distinction
between primary and adventitious root development in maize. At some level, the
mechanisms which regulate invertase in these root systems must differ.


CHAPTER 5
SUGAR RESPONSE OF SUCROSE SYNTHASE AT THE GENE (Sus 7), PROTEIN
AND ENZYME ACTIVITY LEVELS IN ROOTS OF THE Sh1 MAIZE MUTANT
Abstract
The sh1 mutant of maize was used to study expression of the Sus 7 gene
for sucrose synthase in response to sugar availability because this mutant has only
one isozyme gene (Sus 7) for sucrose synthase and provides a system
uncomplicated by the presence of the second gene (Sh1). Koch and McCarty
(1988, 1990) have previously demonstrated that Sus 7 is up-regulated by plentiful
supplies of metabolizable sugars and down-regulated under carbohydrate
depletion, whereas Sh 1 responds in an inverse manner. Excised root tips from
sh 1 were incubated for 24 h in Whites basal salts medium supplemented with
different amounts of glucose. Sus 7 mRNA levels were approximately 5-fold greater
in treatments with 2.0% vs. 0% or 0.2% glucose. This difference was also reflected
in western blot analysis of sus protein. Enzyme activity was elevated 2-fold in root
tips from 2% glucose treatments vs. those in 0 or 0.2%. Time-course and
switching experiments showed that changes in mRNA or protein were not evident
until 24 h and indicated that the response to carbohydrate level had been initiated
within 16 h. Roots incubated in 2.0% glucose for 16 h and switched to 0% for 32
h (total of 48 h) responded like those remaining continuously in 2.0% glucose.
57


67
16h
Intact +
24h
+
48h 16h+32h
- + -/+ +/-
Figure 5-3. RNA gel blot analysis of Sus1 mRNA expression in maize roots
incubated in 0% (-) or 2.0% (+) glucose for various time periods. At 16 h
switching treatments were conducted by changing roots in 0% glucose to
2.0% glucose (-/+) and vise verse (+/-); incubations were continued for 32
additional hours.


84
Table 6-3. Soluble add invertase activity in various tissues of
Oh 43, an inbred line of Zea mays.
/imol glucose
nmol glucose
g'1FW h'1
mg'1 protein h'1
Shoot
10.8 0.6
25.8 1.5
Endosperm
7.3 1.2
12.6 2.1
Scutellum
9.0 0.8
19.4 1.7
Primary roots
Lateral roots from
0.1 0.1
0.2 0.2
10 roots
0.1 0.1w
0.2 0.1W
Adventitious roots
Lateral roots from
16.1 3.2
60.8 12.1
adventitious roots
10.4 0.9
38.7 3.2
W-Activity in root tips from axenic culture was 0.0 0.0 /mol
g'1FW h'1; thus microorganisms are a likely source of the
residual activity in ca. 1 of 3 assays.
Shoot, primary roots and adventitious roots were
samples from 6-day-old seedlings. Endosperm and scutellum
were from developing kernels 23 DAP. Lateral roots
developed after 2 to 3 weeks of growth and were then
excised. Each value represents the mean of 3 separate
samples S.E.M.


21
essential to mycorrhizal associations (Purves and Hadley, 1975). Maize
(Gerdemann, 1964; Kothari et al., 1990) and 90% of other agriculturally important
species form these beneficial symbioses under field conditions (Gerdemann, 1968).
However, sucrose must be hydrolyzed for the fungal symbiont (Long et al., 1975),
and invertase levels rise at sites of carbon transfer. It is not known whether this
is host or fungal invertase.
Another possible role for apoplastic invertase is in the regulation of the
intercellular sucrose concentration. Regulation of the free-space sucrose
concentration may be important in osmotic relations and in the control of tissue
differentiation (ap Rees, 1974). Jeffs and Northcote (1966,1967) have shown that
phloem differentiation in cultures of Phaseolus vulgaris depended on the supply
of sucrose; glucose or fructose would not substitute. Wright and Northcote
(1972), however, have shown that phloem differentiation in cultures of Acer
pseudoplatanus were equally responsive to glucose and sucrose. The results of
Jeffs and Northcote show that in certain cases the regulation of the apoplastic
sucrose content by acid invertase could be important in differentiation.
A definitive role cannot be assigned to alkaline invertase at present. Studies
with sugar cane (Hatch and Glasziou, 1963), carrot roots (Ricardo and ap Rees,
1970), pea roots (Lyne and ap Rees, 1971), melon (Lingle and Dunlap, 1987;
McCollum et al., 1988), and Lycopersicon hirsutum (Miron and Schaffer, 1991)
indicate an inverse relationship between alkaline and acid invertase and a more
positive correlation between alkaline invertase and sucrose concentration. The


LIST OF FIGURES
Figure 3-1 Time course of in vitro decrease in sucrose synthase activity in
maize and cotton roots 35
Figure 3-2 Time course of in vitro decrease in maize root sucrose synthase
activity with and without the serine proteinase inhibitor, PMSF 37
Figure 3-3 Time course of in vitro decrease in maize root sucrose synthase
activity in the presence and absence of either Pi (10 mM) or casein
2% w:v) 39
Figure 3-4 Denaturing (A) and native (B.C.D) protein gel blot analysis of
maize root sucrose synthase at various times after extraction 40
Figure 3-5 Time course of in vitro decrease in maize root sucrose synthase
activity from lines with homo- (W22:s/77) and heterotetrameric (NK
508) forms of this enzyme 43
Figure 5-1 RNA gel blot analysis of Sus 1 expression in maize roots
incubated in a range of glucose concentrations for 24 hours 64
Figure 5-2 Protein gel blot of Sus1 encoded sucrose synthase from maize
roots incubated in a range of glucose concentrations for 24 hours . 65
Figure 5-3 RNA gel blot analysis of Sus 1 mRNA expression in maize roots
incubated in 0 and 2.0% glucose for various time periods 67
Figure 5-4 Protein gel blot of Sus1 encoded sucrose synthase from maize
roots incubated in 0 and 2.0% glucose for various time periods .... 69
Figure 6-1 Histochemical localization of invertase activity in free-hand,
cross sections of root apices of 6-day-old maize seedlings 87
VI


SUCROSE SYNTHASE AND INVERTASE IN MAIZE ROOTS DIFFERING
IN CARBOHYDRATE STATUS AND/OR GENETIC COMPOSITION
By
EDWIN RALPH DUKE
A DISSERTATION PRESENTED TO THE GRADUATE SCHOOL
OF THE UNIVERSITY OF FLORIDA IN PARTIAL FULFILLMENT
OF THE REQUIREMENTS FOR THE DEGREE OF
DOCTOR OF PHILOSOPHY
UNIVERSITY OF FLORIDA
1991

ACKNOWLEDGEMENTS
I would like to extend thanks to the members of my committee, Dr. Karen
Koch, Dr. Rebecca Darnell, Dr. Don McCarty, Dr. Curt Hannah and Dr. Alice
Harmon, for their assistance and guidance during the completion of this degree.
I would also like to thank Dr. Tom Humphreys for his critical review of this
dissertation. There are many people in the Fruit Crops Department to whom I owe
thanks. I would like to thank the staff and faculty for their help and encouragement
during my time here. For their friendship and support, I especially would like to
thank Kathy Zimmerman, Teki and Andy Ericson, Dr. Kathy Taylor, Dr. Pat
Tomlinson, Wayne Avigne, Kurt Nolte, and Don Merhaut. Thanks are also given
to the graduate students, both past and present, of the Fruit Crops Department.
Finally, I want to extend my most heartfelt thanks to my parents, Ralph and Mildred
Duke; they have stood by me and given me support throughout my education,
and I can never thank them enough.
n

TABLE OF CONTENTS
ACKNOWLEDGEMENTS ii
LIST OF TABLES v
LIST OF FIGURES vi
ABSTRACT vii
CHAPTERS
1 INTRODUCTION 1
2 REVIEW OF THE LITERATURE 4
Sucrose Metabolism 4
Sucrose Synthase 6
Invertases 17
Use of Mutants in Physiological Research 26
3 INSTABILITY OF SUCROSE SYNTHASE FROM ROOT
TIPS: CHARACTERIZATION AND STABILIZATION 29
Abstract 29
Introduction 30
Materials and Methods 31
Results 34
Discussion 41
4 SUCROSE SYNTHASE ACTIVITY IN WILDTYPE MAIZE
ROOT TIPS RESPONDING TO ALTERED
CARBOHYDRATE STATUS 49
Abstract 49
Introduction 49
Materials and Methods 51
iii

Results 52
Discussion 52
5 SUGAR RESPONSE OF SUCROSE SYNTHASE AT
THE GENE {Sus1), PROTEIN AND ENZYME ACTIVITY
LEVELS IN ROOTS OF THE SM MAIZE MUTANT 57
Abstract 57
Introduction 58
Materials and Methods 61
Results 63
Discussion 68
6 AN ORGAN-SPECIFIC INVERTASE DEFICIENCY IN
THE PRIMARY ROOT OF AN INBRED MAIZE LINE 75
Abstract 75
Introduction 76
Materials and Methods 78
Results 80
Discussion 85
7 SUMMARY AND CONCLUSIONS 91
LITERATURE CITED 94
BIOGRAPHICAL SKETCH 113
iv

LIST OF TABLES
Table 3-1 Effect of enzyme protectants on activity of sucrose synthase
from maize root tips assayed five minutes after extraction 36
Table 4-1 Total sucrose synthase activity in wildtype maize root tips
incubated in a range of glucose concentrations for 24 hour 53
Table 5-1 Sucrose synthase activity in mutant maize ^N22:sh1) root tips
incubated in media containing a range of glucose concentrations for
24 hours 66
Table 5-2 Sucrose synthase activity in mutant maize (W22:s/77) root tips
incubated in media containing 0 or 2.0% glucose for various time
periods 70
Table 6-1 Soluble and insoluble acid invertase activity in sequential 2 mm
segments of primary roots of 5- to 6-day-old seedlings from 1 hybrid
and 2 inbred lines of maize 81
Table 6-2 Soluble acid invertase and sucrose synthase activity in 0.5 cm
apices of primary and adventitious roots of 5- to 6-day-old seedlings
from 1 hybrid and 2 inbred lines of Zea mays 83
Table 6-3 Soluble acid invertase activity in various tissues of Oh 43, an
inbred line of Zea mays 84
v

LIST OF FIGURES
Figure 3-1 Time course of in vitro decrease in sucrose synthase activity in
maize and cotton roots 35
Figure 3-2 Time course of in vitro decrease in maize root sucrose synthase
activity with and without the serine proteinase inhibitor, PMSF 37
Figure 3-3 Time course of in vitro decrease in maize root sucrose synthase
activity in the presence and absence of either Pi (10 mM) or casein
2% w:v) 39
Figure 3-4 Denaturing (A) and native (B.C.D) protein gel blot analysis of
maize root sucrose synthase at various times after extraction 40
Figure 3-5 Time course of in vitro decrease in maize root sucrose synthase
activity from lines with homo- (W22:s/77) and heterotetrameric (NK
508) forms of this enzyme 43
Figure 5-1 RNA gel blot analysis of Sus 1 expression in maize roots
incubated in a range of glucose concentrations for 24 hours 64
Figure 5-2 Protein gel blot of Sus1 encoded sucrose synthase from maize
roots incubated in a range of glucose concentrations for 24 hours . 65
Figure 5-3 RNA gel blot analysis of Sus 1 mRNA expression in maize roots
incubated in 0 and 2.0% glucose for various time periods 67
Figure 5-4 Protein gel blot of Sus1 encoded sucrose synthase from maize
roots incubated in 0 and 2.0% glucose for various time periods .... 69
Figure 6-1 Histochemical localization of invertase activity in free-hand,
cross sections of root apices of 6-day-old maize seedlings 87
VI

Abstract of Dissertation Presented to the Graduate School
of the University of Florida in Partial Fulfillment of the
Requirements for the Degree of Doctor of Philosophy
SUCROSE SYNTHASE AND INVERTASE IN MAIZE ROOTS DIFFERING
IN CARBOHYDRATE STATUS AND/OR GENETIC COMPOSITION
By
Edwin Ralph Duke
August, 1991
Chairperson: Dr. Karen E. Koch
Major Department: Horticultural Science
The extent to which sucrose is transported in phloem of higher plant
species necessitates its effective metabolism by non-photosynthetic cells.
However, only two enzymes can catalyze sucrose breakdown in these instances,
invertase and sucrose synthase (a reversible enzyme). Specific molecular, genetic
and physiological factors affecting these enzymes were investigated in the root tips
of maize. A radiometric assay was first developed for sucrose synthase to
circumvent the rapid decline of sucrose synthase activity in vitro. Further,
characterization of in vitro instability indicated that activity decline was not
associated with any detectable proteolytic degradation, charge alteration, or
subunit separation and that inhibition of activity by inorganic phosphate suggested
possible phosphorylation of this enzyme.
VII

Message levels of genes encoding sucrose synthase isozymes in maize
have been shown to respond to tissue carbohydrate status, thus the effects of
such changes were examined at the level of enzyme activity. Total sucrose
synthase activity from roots of wildtype plants showed little difference in extracts
from root tips incubated for 24 h in a range of glucose levels. This activity,
however, is the combined contribution of two isozymes whose genes are
responding differentially to experimental conditions.
The sh 1 mutant of maize was used to study expression of the Sus1 gene
for sucrose synthase in response to sugar availability because this mutant has only
one gene (Sus1) for sucrose synthase and provides a system uncomplicated by
the presence of the second isozyme (Sh1). Sus1 mRNA increased 5-fold when
incubated for 24 h in 2.0% glucose compared to 0 or 0.2% glucose. Levels of Sus
protein were slightly elevated with increasing sugar levels. Enzyme activity was
elevated 2-fold under the same conditions. A study of time-course and treatment
reversals showed that changes in mRNA or protein were not evident until 24 h and
indicated that the response to carbohydrate level had been initiated within 16 h.
Overall, enhanced expression of Sus 1 was evident at the mRNA, protein and
enzyme levels.
An organ-specific invertase deficiency affecting only the primary root system
also was characterized in the Oh 43 maize inbred. Substantial acid invertase
activity was evident in extracts of all tissues tested except the primary root system
of Oh 43. This deficiency was also evident in lateral roots arising from the primary
VIII

root but not in otherwise morphologically identical laterals from adventitious roots.
In contrast, sucrose synthase was active in all roots and theoretically provided the
only available avenue for sucrose degradation in primary root tips of Oh 43.
IX

CHAPTER 1
INTRODUCTION
Sucrose metabolism is important to the majority of plant species because
of the nearly ubiquitous role of this sugar in growth and development. Initial
breakdown of sucrose can be catalyzed by either invertase or the reversible
enzyme sucrose synthase.
Gene responses to changes in carbohydrate availability have been reported
for mammalian cells (Lin and Lee, 1984) and yeasts and bacteria (Carlson, 1987;
Schuster, 1989). Evidence has also been presented that the transcriptional activity
of promoters of photosynthetic genes from maize protoplasts are repressed and
coordinated by sugars (Sheen, 1990). Expression of the genes encoding the
sucrose synthase isozymes also have been shown to respond to carbohydrate
levels (Koch and McCarty, 1988; Koch et al., 1989). Northern blot analysis of
mRNA from wildtype maize roots showed levels of one isozyme gene (Sh 1) was
up-regulated in response to carbohydrate depletion whereas the other (Sus)
responded positively only when sugar availability was elevated. This sensitivity of
genes for a key enzyme in sugar metabolism may prove to be an important control
mechanism whereby plant cells are able to react to cellular nutritional conditions.
The importance of invertase, the other enzyme of sucrose catabolism in
plants, to root metabolism is unclear. Invertases are especially active in tissues
1

2
undergoing rapid cell division such as shoot and root apices (Avigad, 1982).
Recent evidence, however, indicates that although much hydrolysis is often
observed, invertase activity may not be essential for sucrose uptake into either
sugar cane stems (Lingle, 1989; Thom and Maretzki, 1990) or maize kernels
(Schmalstig and Hitz, 1987). Chapleo and Hall (1989a, b, and c) also have
concluded that invertase was not essential to sucrose import into roots of Ricinus.
Giaquinta and co-workers (1983) showed that sucrose entering the roots of maize
via phloem does not have to pass through the extracellular space. Robbins (1958)
first reported that root tips of an inbred line of maize, Oh 43, were unable to
retrieve exogenous sucrose. B. Burr (Brookhaven National Laboratory, personal
communication) has indicated that the lack of retrieval might possibly be due to
an invertase deficiency. The absence of invertase activity could have important
implications for sucrose import, not only because of potential effects on the
retrieval system, but also because sucrose breakdown in such an instance could
theoretically proceed only via action of sucrose synthase. A mutant lacking
functional invertase in its roots would also be useful, in combination with nulls for
both sucrose synthase isozymes, to elucidate the individual roles for these
enzymes.
The purpose of the following research is to further elucidate the roles and
regulation of the two sucrose metabolizing enzymes sucrose synthase and
invertase in roots of maize. Specific objectives are to:

3
1. Develop a method of accurately assaying sucrose synthase activity
to circumvent the rapid decline in activity exhibited upon extraction
from maize root tips.
2. Clarify the possible causes of the rapid decline of sucrose synthase
activity in extracts from maize root tips.
3. Determine the effects of varying carbohydrate conditions on total
sucrose synthase activity in extracts from wildtype maize root tips.
4. Ascertain the effects of varying carbohydrate conditions on the Sus 1
gene and its sucrose synthase gene product free from the
confounding effects of Sh 1.
Characterize the extent of invertase activity in various tissues of the
Oh 43 inbred of maize, a putative invertase-deficient mutant.
5.

CHAPTER 2
REVIEW OF THE LITERATURE
Sucrose Metabolism
Sucrose is the major transported sugar in the majority of higher plant
species. The major roles of sucrose in higher plants include its function as both
a translocatable form of carbon and as a vacuolar storage compound (Hawker,
1985). It is a non-reducing sugar made up of a glucose (a-D-glucopyranose and
fructose (/?-D-fructofuranose) joined by an a-1,2 linkage. For metabolic utilization,
it must be broken down into its component monosaccharides or their derivatives.
Only two enzymatic systems for sucrose breakdown are known in higher plant
tissues. Sucrose can be hydrolyzed by the action of invertase or cleaved by
sucrose synthase working in the degradative direction. The free energy of sucrose
hydrolysis is nearly equal to that of the y phosphoryl group of ATP (AG= -7.0
kcal/mol and -7.3 kcal/mol, respectively) (Neufeld and Hassid, 1963). This is much
greater than the free energy of most other glycosidic bonds (Avigad, 1982).
Cleavage by sucrose synthase retains the bond energy in the a-glucosyl bond of
UDP-glucose. In contrast, hydrolysis by invertase conserves none of the free
energy in the bond.
4

5
Morell and Copeland (1984, 1985) have investigated the enzymes of
sucrose breakdown in soybean nodules and found that both sucrose synthase
and alkaline invertase are present. They suggested that sucrose partitioning
between the two enzymes could be determined by differences in their affinities for
this substrate. The Km of alkaline invertase for sucrose was 10 mM whereas that
of sucrose synthase was 31 mM. Given the presence of both enzymes, they
proposed the greater affinity of alkaline invertase for sucrose in this system could
ensure that most of the sucrose entering the nodule would be converted to
hexoses for further catabolism. At the same time some sucrose would be
converted to UDP-glucose for subsequent synthesis of nucleotide sugars and
polysaccharides. In contrast, Huber and Akazawa (1986) reported essentially an
opposite situation in cultured sycamore cells, another system in which both
sucrose synthase and neutral (alkaline) invertase were present at the same time
and with similar activities. In these cells the sucrose synthase for sucrose was
substantially lower than that of neutral invertase (15 vs 65 mM). They proposed
two pathways of sucrose cleavage, initiated by each of the enzymes, both
pathways eventually leading to the production of triose-phosphates. Sucrose
concentration was postulated to regulate carbon flow between the two pathways.
Sucrose synthase had a lower Km for sucrose and would, therefore, be relatively
more important under sucrose limiting conditions (Avigad, 1982). This pathway is
more energy efficient and would be more beneficial to the cells when carbon
supplies are limited. The work of Morell and Copelands work (1984, 1985) and

6
Huber and Akazawa (1986) both demonstrate the potential importance of
simultaneous sucrose catabolism via two enzyme systems.
Sucrose Synthase
Sucrose synthase (EC 2.4.1.13, UDP-D-glucose: D-fructose 2-a-D-
glucosyltransferase) is ubiquitous in higher plants (Keller et al., 1988) and probably
occurs in all types of tissues. However, this enzyme is found in greatest
abundance in nonphotosynthetic tissues and in developing seeds (Echt and
Chourey, 1985). In cell fractionation studies, sucrose synthase was shown to be
associated with the soluble fraction (Nishimura and Beevers, 1979; MacDonald and
ap Rees, 1983). A cytosolic rather than vacuolar localization for sucrose synthase
has been demonstrated in protoplasts isolated from Jerusalem artichoke (Keller
et al., 1988).
Although molecular weights as high as one million have been reported for
sucrose synthase (Grimes et al., 1970), it is now generally concluded that, in its
native state, sucrose synthase has a molecular weight of approximately 36 to 40
kD (Delmer, 1972a; Su and Preiss, 1978; Morell and Copeland, 1985; Moriguchi
and Yamaki, 1988) and is composed of four identical subunits. The sucrose
synthase subunit from maize has been found to have a molecular weight of 8.8 kD
(Su and Preiss, 1978). However, unlike many other plant species, maize has two
genes which encode sucrose synthase subunits (Chourey and Nelson, 1976; Echt
and Chourey, 1985). The two sucrose synthase subunits of maize, sh1 and sus,
encoded by the Sh1 and Sus1 genes, respectively, have similar enzyme kinetics,

7
similar amino acid compositions and share limited structural homologies (Echt and
Chourey, 1985). They do differ slightly in their electrophoretic movement during
PAGE (Echt and Chourey, 1985). The two subunits are homologous enough to
form heterotetrameric structures, apparent as five separate bands on native PAGE
(Echt and Chourey, 1985).
Spatial separations of these two sucrose synthase isozymes are sometimes
apparent. Sh1 encoded protein is primarily located in the endosperm (Chourey
and Nelson, 1976; Chen and Chourey, 1989), in the root (Springer et al., 1986;
Chourey et al., 1986) and in etiolated shoots (Springer et al., 1986). The Sus1
encoded protein is found throughout the plant (Chourey, 1981; Echt and Chourey,
1985; Chourey et al., 1988). The distribution of both proteins is further
distinguished under stress conditions (such as anaerobiosis) where tissue-specific
localization in roots is readily apparent (Rowland et al., 1989).
Roles of sucrose synthase
Sucrose synthase was initially considered to be a sucrose synthesizing
system in plants by Leloir and Cardini (1953, 1955). Sucrose synthase and
sucrose phosphate synthase (EC 2.3.1.14, UDP-D-glucose: D-fructose-6-
phosphate 2-a-D-glucosyltransferase) are the two enzymes that catalyze the
transglucosylation reaction from UDP-glucose to fructose and fructose-e-
phosphate, respectively.

8
The necessity for two systems of sucrose synthesis was puzzling until it was
determined that the sucrose synthase reaction is readily reversible (Cardini et al,
1955):
Sucrose + UDP < > UDP-Glucose + Fructose
This reversibility gave rise to the suggestion that sucrose synthase could make
UDPG available for utilization as a glucosyl donor in starch synthesis (Turner and
Turner, 1957). Other studies of sucrose synthase specificity and kinetics led
Avigad and coworkers (Avigad, 1964; Avigad et al., 1964; Milner and Avigad, 1964)
to suggest that this enzyme functioned mainly in sucrose cleavage in storage
tissues. Sucrose synthase activity typically is highest in tissues during periods of
rapid growth and is often not accompanied by high invertase activity (Schaffer et
al., 1987).
Several factors lend credence to the view that the role of sucrose synthase
is sucrose cleavage in importing cells. First, a substantial level of free fructose is
required for sucrose synthase activity in the synthetic direction (Km ca. 2.0-2.5 mM
[Avigad, 1982]). Levels of this sugar are low in healthy, intact leaves, but they are
higher in storage tissues and roots, where most of the free fructose is in the
vacuole or extracellular spaces (Avigad, 1982). Availability of UDPG is also likely
to limit the synthetic reaction, because the cellular concentration is typically less
than 0.4 mM (Murata, 1975). The Km for UDPG ranges from 0.1 to 8.5 mM
(average approximately 2.0 mM)(Avigad, 1982). In contrast, sucrose

9
concentrations are generally elevated in importing areas. Therefore, substrate
levels favor the cleavage reaction in the majority of instances.
A second line of evidence for the degradative role of sucrose synthase in
importing organs is provided by mutant maize lines lacking a functional sucrose
synthase protein. Chourey and Nelson (1976) showed that a deletion of the Sh
locus on chromosome 9 of maize (coding for sucrose synthase) led to a 90%
deficiency of the respective protein in mutant vs wild-type kernels. Starch
formation was also reduced in this line, giving rise to a "shrunken" seed. The
association between this shrunken phenotype and a sucrose synthase deficiency
was considered evidence that the critical reaction in vivo was that of sucrose
cleavage, and that this was essential for conversion of photosynthetically produced
sucrose for starch biosynthesis. The residual amount of starch deposited was
attributed to the sucrose degrading activity of a second sucrose synthase encoded
by another locus (Sus1).
Further research also favors the cleavage role of sucrose synthase and
implicates its involvement in starch deposition. Dale and Housley (1986), for
example, found that developing wheat kernels with the greatest rates of growth
and starch deposition had significantly greater sucrose synthase activities. A
positive correlation between sucrose synthase activity and starch deposition was
also reported in Pisum sativum by Edwards and ap Rees (1986a and b). They
proposed that UDP-glucose formed during sucrose cleavage was converted to
glucose-1-phosphate by UDP-glucose pyrophosphorylase using pyrophosphate

10
generated by PFK(PP¡). Morrell and ap Rees (1986) have also suggested that
much of the sucrose translocated to developing potato tubers is probably
metabolized via the same pathway with the initial step catalyzed by sucrose
synthase. Gibson and Shine (1983) have demonstrate that in the presence of
inorganic phosphate, UDPG may be hydrolyzed to G-1-P and UDP by the action
of UDPG phosphorylase. Salerno (1986) also has demonstrated the presence a
highly nucleotide specific form of UDP-glucose phosphorylase in developing maize
endosperms. The activity level of this enzyme followed closely the development
of the grain and paralleled that of sucrose synthase. The presence of this enzyme
links sucrose cleavage and starch formation via sucrose synthase.
Finally, an additional line of evidence was presented by Cobb and Hannah
(1988) that also indicated a primarily degradative function for sucrose synthase in
importing organs. They showed that maize kernels from a line deficient in the Sh 1
gene for sucrose synthase still had normal levels of sucrose and normal rates of
sucrose synthesis when grown in culture with fructose as the carbon source. If
sucrose synthesis had been proceeding via sucrose synthase in wild-type kernels,
then the loss of ca. 90% of total sucrose synthase activity in kernels of the mutant
line should have affected sucrose formation there. The authors concluded that
Sh1 encoded sucrose synthase was not necessary for sucrose synthesis.
Correlative data suggest that sucrose synthase activity is closely linked with
sink strength. The supply and timing of sucrose for export seem to be closely
related to the source of photosynthate (Fondy and Geiger, 1982; Servaites et al.,

11
1989) as well as the energy for phloem loading. However, once sucrose is loaded,
its eventual fate does not appear to be under the control of the source leaf (Gifford
and Evans, 1981) but rather is under the control of the importing sink (Wyse,
1986). Giaquinta (1979) found that young, immature roots of sugar beets had low
levels of sucrose synthase, but the onset of rapid sucrose import for storage was
accompanied by a significant increase in sucrose synthase activity. Similar
correlations were also observed by Silvius and Snyder (1979) and Fieuw and
Willenbrink (1987). In sugar beet roots sucrose uptake into parenchyma can
proceed without prior hydrolysis in the apoplast or free space. Increases in
sucrose synthase activity have also been observed during periods of sucrose
import and/or accumulation in sweet melons (Schaffer et al., 1987), netted
muskmelon (Lingle and Dunlap, 1987), eggplants (Claussen et al., 1985, 1986),
rose flowers (Khayat and Zeslin, 1987), developing chick pea seeds (Setia and
Malik, 1985), tomato (Yelle et al., 1988) and Solanum muricatum (Schaffer et al.,
1989). Lingle (1987), however, found no correlation of sucrose synthase activity
with sucrose concentration in sweet sorghum.
Huber and Akazawa (1986) hypothesized that a primary role of sucrose
synthase could be to feed glucose-1 -phosphate directly into glycolysis. Black and
coworkers (Black et al., 1987; Sung et al., 1988; 1989; Xu et al., 1989) also support
this hypothesis. This link to glycolysis also involves UDPG-pyrophosphorylase,
which converts the UDPG formed by the action of sucrose synthase into G-1-P
and UTP. The methods employed to deliver carbohydrates to their respective

12
sinks vary from species to species (ap Rees, 1974, Hawker, 1985). However,
sucrose breakdown generally appears to proceed via sucrose synthase in starch
and sugar storage sinks (Sung et al., 1988).
Sucrose synthase also may be associated with sink strength through its
potential involvement in cell wall synthesis (Hendrix, 1990). Sucrose synthase
activity predominates over that of invertase in rapidly expanding cotton ovules, for
example where rapid elongation of epidermal hairs (cotton fibers) requires
extensive cellulose formation. Hendrix (1990) postulated that sucrose entering the
seed coat in the developing cotton boll was cleaved via sucrose synthase, and
subsequent carbohydrates went into the rapidly growing epidermal hairs.
However, some of the sucrose breakdown products were converted to starch
stored in the seed coats. Stepanenko and Morozova (1970) demonstrated
cellulose biosynthesis from UDP-glucose in cotton. However, they did not indicate
that the source of the UDPG might come from the action of sucrose synthase.
Carpita and Delmer (1981) showed that the rate of synthesis and turnover of UDP-
glucose in developing cotton ovules was more than sufficient to account for rates
of biosynthesis for cellulose, 0-1,3-glucan and sterylglucosides (all cell wall
constituents). They found that UDPG levels increased dramatically just prior to the
maximum rate of secondary wall cell synthesis and dropped precipitously at the
time when cellulose synthesis ceased. Again, sucrose synthase activity was not
measured, but could possibly be the source of the increased levels of UDPG.
Sucrose synthase activity was measured in cultured cells of Catharanthus roseus

13
(Amino et al., 1985) and found to be elevated during the G1 phase when the
amount of total cell walls increased significantly. However, UDP-glucose
pyrophosphorylase activity (also involved in the formation of UDPG) was greater
than sucrose synthase activity at the G1 phase. The former was considered by
these authors likely to make a more important contribution to the total UDPG
formed. Chourey et al. (1991a) have reiterated the hypothesis that the resultant
shrunken, starch-deficient endosperm of the sh1 maize mutant may be due to
reduced cell wall deposition rather than any direct effect on the starch biosynthetic
pathway.
Regulation of Sucrose Svnthase
Sucrose synthase, a key enzyme in sucrose metabolism, is subject to a
number of complex regulatory factors (Davies, 1974). This enzyme exhibits a wide
specificity for the nucleoside base utilized in the reaction. Most enzymes of sugar
nucleoside metabolism show a marked specificity for a particular base (Avigad,
1982). Sucrose synthase working in the synthetic direction has been shown to
utilize UDPG, ADPG, TDPG, CDPG and GDPG as glucosyl group donors (Avigad,
1982). The Km for UDPG, however, is usually much less than for other NDPGs.
Grimes et al. (1970) found that the Km for UDPG was approximately 0.2 mM while
ADPG, TDPG, CDPG and GDPG had Kms of 1.8,1.7, 2.5 and 2.5 mM, respectively.
They also found that with UDPG as the nucleoside sugar, the Km for fructose was
reduced 10 fold below that apparent when ADPG was utilized. This change was

14
attributed by Grimes et al. (1970) to possibly result from conformational changes
in the enzyme.
Reduced K^s for UDPG when compared to other NDPGs have also been
observed for sucrose synthase from pea seedlings (Gabrielyan et al., 1969), sweet
potato root (Murata, 1971), potato tubers (Pollock and ap Rees, 1975), sweet corn
seeds (de Fekete and Cardini, 1964) and soybean nodules (Morell and Copeland,
1985). In contrast, ADPG and TDPG were reportedly more efficient glucosyl
donors than UDPG for sucrose synthase isolated from sorghum seeds (Sharma
and Bhatia, 1980) and sugar beet roots (Avigad and Milner, 1966).
No large differences in Kms for UDP and other nucleoside diphosphates
generally are observed when the reverse reaction is analyzed. Delmer (1972a and
b) found minimal or no differences in Kms for NDPs with sucrose synthase from
mung bean seedlings. She did, however, find a large difference in the rates of
sucrose cleavage with different NDPs. Maxima were observed when UDP was the
substrate. The Vmax for UDP in relative terms was 100 compared to 28, 6, 3 and
3 for ADP, TDP, CDP and GDP, respectively. Similar K^s for various NDPs have
also been observed in sweet potato roots (Murata, 1971), potato tubers (Pollock
and ap Rees, 1975), Jerusalem artichoke (Pontis et al., 1972) and sugar beet
(Avigad and Milner, 1966). Sucrose synthase from sweet corn kernels does,
however, exhibit a Km an order of magnitude greater for ADP than UDP (Su and
Preiss, 1978; de Fekete and Cardini, 1964). Morell and Copeland (1985) also

15
found that in soybean nodules, the Km of sucrose synthase for UDP (0.5 mM) was
lower than that of ADP and CDP (0.13 and 1.1 mM, respectively).
Delmer (1972a and b) characterized regulation of purified Phaseolus aureus
sucrose synthase and found a number of differences in the regulation of the
synthetic and degradative reactions. NADP, iodoacetic acid, and gibberellic acid
all stimulate sucrose degradation but inhibit sucrose synthesis. Pyrophosphate
also enhanced the degradative activity, but only in the presence of MgCI2. In
contrast, Pontis (1977) reported that P¡ inhibited the degradative reaction alone or
in the presence of Mg2+. Delmer also tested the effects of intermediates in
carbohydrate metabolism and found that G-1-P, G-6-P, F-6-P, F-1,6-BP, R-5-P, R-
1,5-BP, PEP and 3-PGA had little or no influence on the sucrose synthase reaction
in either direction when present at 2 mM. However, de Fekete (1969) and Pontis
(1977) both have reported that G-1-P, G-6-P and F-1,6-BP were inhibitory to the
degradative reaction at 2-5 mM without affecting the synthetic reaction. ATP, ADP
and AMP had no inhibitory effect on sucrose synthesis at 4 mM; however, the
degradative reaction was inhibited 30% by ADP, and 50% by both ADP and AMP.
/3-Phenylglucoside has also been shown to inhibit sucrose degradation almost
completely and sucrose synthesis by 50% (Wolosiuk and Pontis, 1974b; Lowell,
1986). This has proven useful for distinguishing activities of sucrose phosphate
synthase from sucrose synthase.
Pontis and coworkers (Pontis et al., 1972) found that the divalent cations
Mg2+, Mn2+, Ca2+ and Ba2+ at 5-10 mM activated sucrose synthase in the

16
synthetic direction but inhibited the cleavage reaction. UDP was found to be a
strong inhibitor of the synthetic reaction at 10 mM (70-80% inhibition), but the
inhibition could be reversed by the addition of Mg2+ (de Fekete and Cardini, 1964).
UDP was also found to inhibit the degradative reaction as a competitive inhibitor
for UDPG (Wolosiuk and Pontis, 1974a). Inhibition by other NDPs was very weak.
UTP (4 mM) caused a slight inhibition of synthetic activity, but caused an 80%
inhibition of the degradative reaction (Tsai, 1974). Echeverra and Humphreys
(1985), however, found that UDP and UTP within the cytosolic range (< 4 mM)
both had little or no effect on sucrose synthase in the synthetic direction. UDPG
was able to inhibit the cleavage reaction by 13% at 10 mM, but tissue
concentrations were generally below this level (Echeverra and Humphreys, 1985),
with the effect on the synthetic reaction minimal. Wolosiuk and Pontis (1974a)
found that UDPG could function as a competitive inhibitor for UDP in the sucrose
synthase synthesis reaction.
Carbohydrates also have been found to inhibit sucrose synthase activity in
vitro. Fructose was found to function as a competitive inhibitor of sucrose in the
cleavage reaction (Pridham et al., 1969; Doehlert, 1987). Sucrose had no
inhibitory effects on sucrose synthase activity at saturating levels of fructose and
UDPG (Echeverra and Humphreys, 1985), but glucose at 100 mM inhibited
sucrose synthesis by 63%-70% and inhibited sucrose cleavage 86%-93%.
Expression of sucrose synthase genes also respond to carbohydrate availability.
Koch and McCarty (1988,1990; Koch et al., 1989) have shown that levels of maize

17
root Sh1 mRNA were elevated when sugar supplies were limited in culture. In
contrast, levels of Sus1 mRNA were elevated in response to increasing glucose
concentrations. They speculated that the effect of sugar levels on expression of
specific genes could prove to be an important control mechanism whereby plant
cells could react to cellular nutritional conditions. The Sh1 gene is also up-
regulated under anaerobic conditions (Springer et al., 1986); however, the
carbohydrate response of the Sh 1 gene appears to be distinct from its anaerobic
regulation (Koch et al., 1989). There is some doubt as to whether anaerobic
induction occurs at both the transcriptional and translational levels in maize
(McElfresh and Chourey, 1988). Taliercio and Chourey (1989) hypothesized that
the expression of anaerobically induced Sh1 transcripts are blocked at some step
beyond polyribosomal loading. However, other researchers have shown that
sucrose synthase in maize is anaerobically induced at the protein as well as the
gene level (Freeling and Bennett, 1985; Springer et al., 1986). Anaerobic induction
of sucrose synthase at both the gene and protein levels also has been
demonstrated in rice (Ricard et al, 1991) and Echinochloa phyllopogon (Mujer et
al., 1990).
Invertases
Invertases (E.C. 3.2.1.26, /3-D-fructofuranoside fructohydrolases) are widely
distributed in the plant kingdom and catalyze the following reaction:
Sucrose + H20
> Glucose + Fructose

18
Invertases are specific for the fructofuranose moiety of sucrose and work by
hydrolyzing the glycosidic linkage between the bridge oxygen and the fructose
residue (Sum et al., 1980). Up to five different forms of invertase have been
reported in plants (Sasaki et al., 1971). However, these enzymes are generally
divided into two main types based on the pH at which sucrose hydrolysis is most
efficiently accomplished. Acid invertases have pH optima around 4.5 to 5.0; that
of alkaline (or neutral invertase) is 7.0 to 7.5.
Most invertases are glycoproteins. Arnold (1966) partially purified an acid
invertase from grapes and found that it was approximately 25% carbohydrate.
Faye and coworkers (Faye and Berjonneau, 1979; Faye et al., 1981) have shown
a 7.7% carbohydrate content of acid invertase from radish seedlings, and date
invertase has a carbohydrate content of 8.2% (Al-Bakir and Whitaker, 1978).
Invertase preparations from barley (Prentice and Robbins, 1976), sugar cane (del
Rosario and Santisopasri, 1977), potato tubers (Anderson and Ewing, 1978;
Bracho and Whitaker, 1990b) and banana (Sum et al., 1980) were shown to bind
strongly to concanavalin A, a phytagglutinin or lectin isolated from jack bean with
a strong binding affinity for carbohydrates. In yeast and Neurospora the
carbohydrate content of invertase has been estimated to range from 0 to 50 %
(Metzenberg, 1963; Gascon et al., 1968; Holbein et al., 1976). Much of our more
detailed knowledge on the molecular structure and mode of action of invertases
comes from studies on fungal and yeast enzymes; however, considerable
progress has been made in analyses of plant invertases.

19
Roles of Invertase
Elevated acid invertase activity is characteristic of plant tissues in which
there is a need for hexoses produced from stored or recently transported sucrose
(ap Rees, 1974; Avigad, 1982). Greater activities of invertase also correlate well
with a low content of stored sucrose. In sugar beets, the onset of sucrose storage
is accompanied by a decrease in invertase activity (Silvius and Snyder, 1979;
Giaquinta, 1979). The same is true for carrot roots (Ricardo and ap Rees, 1970),
melon (Hubbard et alM 1989; Lingle and Dunlap, 1987; Schaffer et al., 1987;
McCollum et al., 1988), citrus (Kato and Kubota, 1978; Lowell, 1986) and
Lycopersicon hirsutum (Miron and Schaffer, 1991). In these systems, invertase
was very active prior to sucrose accumulation and dropped significantly upon
maturity. Invertase activity is usually greatest in tissues that are at a rapid stage
of growth and development (Weil and Rausch, 1990), particularly at the cell
division stage (Masuda et al., 1988). Root apices, young leaves and stem
internodes fall into this category. Mature leaves, functioning as sources of
photosynthates, generally have low levels of apoplastic acid invertase (Dickinson
et al., 1991). Transgenic tomato plants expressing yeast invertase in the apoplast
of mature leaves had a striking repression of growth (Dickinson et al., 1991). The
higher the level of invertase, the greater the inhibition. The general role of acid
invertase, therefore, seems to be for the breakdown of sucrose where there is a
marked need for hexose (ap Rees, 1974).

20
Previously, the role of invertase in sucrose transfer was considered
particularly important in plants such as sugar cane and maize where substantial
sugar movement occurred through the cell wall space and was accompanied by
action of an extracellular invertase (Glasziou and Gayler, 1972; Hawker and Hatch,
1965). Recent evidence, however, indicates that although much hydrolysis is often
observed, invertase activity may not be essential for sucrose uptake into either
sugar cane stems (Thom and Maretzki, 1990; Lingle, 1989) or maize kernels
(Schmalstig and Hitz, 1987). The role of apoplastic invertase in sucrose import
into roots had previously been questioned by Chapleo and Hall (1989a) who
concluded that although present, apoplastic root invertase did not have a direct
role in sugar transport. However, substantial activity of invertase has been widely
documented in roots of plants such as pea (Lyne and ap Rees, 1971), bean
(Robinson and Brown, 1952), tomato (Chin and Weston, 1973), Ricinus (Chapleo
and Hall, 1989a, b, and c), oat (Pressy and Avants, 1980), and maize (Hellebust
and Forward, 1962; Chang and Bandurski, 1963). Specific tissue localizations also
have been described. Peak activity for root invertase is generally 2-3 mm behind
the apex and corresponds to the region of expansion and elongation in pea
(Robinson and Brown, 1952; Sexton and Sutcliffe, 1969) and maize (Hellebust and
Forward, 1962). In Ricinus roots, this activity predominates in the cortex (Chapleo
and Hall 1989a).
Although invertase may not have a direct role in sucrose import into roots,
it still may be important to two major aspects of root biology. First, invertase is

21
essential to mycorrhizal associations (Purves and Hadley, 1975). Maize
(Gerdemann, 1964; Kothari et al., 1990) and 90% of other agriculturally important
species form these beneficial symbioses under field conditions (Gerdemann, 1968).
However, sucrose must be hydrolyzed for the fungal symbiont (Long et al., 1975),
and invertase levels rise at sites of carbon transfer. It is not known whether this
is host or fungal invertase.
Another possible role for apoplastic invertase is in the regulation of the
intercellular sucrose concentration. Regulation of the free-space sucrose
concentration may be important in osmotic relations and in the control of tissue
differentiation (ap Rees, 1974). Jeffs and Northcote (1966,1967) have shown that
phloem differentiation in cultures of Phaseolus vulgaris depended on the supply
of sucrose; glucose or fructose would not substitute. Wright and Northcote
(1972), however, have shown that phloem differentiation in cultures of Acer
pseudoplatanus were equally responsive to glucose and sucrose. The results of
Jeffs and Northcote show that in certain cases the regulation of the apoplastic
sucrose content by acid invertase could be important in differentiation.
A definitive role cannot be assigned to alkaline invertase at present. Studies
with sugar cane (Hatch and Glasziou, 1963), carrot roots (Ricardo and ap Rees,
1970), pea roots (Lyne and ap Rees, 1971), melon (Lingle and Dunlap, 1987;
McCollum et al., 1988), and Lycopersicon hirsutum (Miron and Schaffer, 1991)
indicate an inverse relationship between alkaline and acid invertase and a more
positive correlation between alkaline invertase and sucrose concentration. The

22
maximum values for alkaline invertase activity observed to date are consistently
less than those of acid invertase (Masuda et al., 1988). The possibility exists that
alkaline invertase allows cells that store sucrose in their vacuoles to retain a
capacity for breakdown of enough sucrose in the cytoplasm to meet respiratory
and metabolic demands for hexoses (ap Rees, 1974). The capacity of a plant to
produce two different invertases that are spatially separated may allow the plant
cell to regulate sucrose storage independent from sucrose breakdown.
Regulation of Invertase
Acid invertases are generally found in the apoplast and vacuoles of plant
tissues. Washed preparations of cell walls contain a large proportion of a plants
acid invertase (Little and Edelman, 1973). A portion of the acid invertase can be
extracted from the cell wall during grinding, but at least some of the enzyme is
considered to be attached to the cell wall in vivo (Edelman and Hall, 1965). The
major determinant of how much acid invertase remains bound during extraction
is the pH of the buffer used (ap Rees, 1974). Buffers with acidic pH leave most
of the activity in the cell wall fraction, whereas neutral or alkaline buffers release the
majority into the soluble fraction.
Early evidence suggested that the soluble and insoluble acid invertases
were not simply different forms of the same enzyme. The pH optima and the
of bound acid invertase of mature (Hawker and Hatch, 1965) and immature (Hatch
et al., 1963) sugar cane storage tissue differed from those of the soluble fractions.
Association with the cell wall may change an enzymes properties (ap Rees, 1974);

23
however the differences in values, especially those of mature tissues, were
considered unlikely to be wholly artifactual. Distinguishing between forms of
invertase is further complicated by information obtained from yeast. One gene in
yeast, SUC2, has been shown to encode the two forms of invertase in yeast,
secreted and intracellular, via two differentially regulated mRNAs (Carlson and
Botstein, 1982).
The Kms of acid invertase for sucrose generally range from 2 to 13 mM.
Sucrose is the primary substrate for acid invertase but raffinose also is hydrolyzed,
though at a slower rate (10% to 50% the rate of sucrose) (Avigad, 1982). Acid
invertase from sugar-cane leaves was inhibited competitively by fructose (K¡ 32
mM) and noncompetitively by glucose (K¡ 37 mM) (Sampietro et al., 1980). Acid
invertases have been partially purified from a number of tissues with apparent
molecular weights ranging from 2.8 x 104 to 2.2 x 105 (Roberts, 1973; Ricardo,
1974; Kato and Kubota, 1978; Masuda and Sugawara, 1980; Sum et al., 1980;
Faye et al., 1981).
The SUC2 gene of Saccharomyces, encoding invertase, has been shown
to be modulated by glucose levels (Carlson et al., 1987). Sucrose or raffinose,
substrates of the yeast invertase, have no such effect. Kaufman et al. (1973) found
that acid invertase activity rises in Avena stem segments incubated in a sucrose-
containing medium. The response had a lag time of 10-12 hours, suggesting a
change in protein levels. Fructose in the incubation medium resulted in a similar
response, but glucose caused no change in invertase activity.

24
A naturally occurring acid invertase inhibitor has been detected in a number
of plant tissues including beet roots (Burakhanova et al., 1987), potato roots
(Pressy, 1967, 1968; Bracho and Whitaker, 1990a and b), maize endosperm
(Jaynes and Nelson, 1971), pea pollen (Malik and Sood, 1976) and Ipomea petals
(Winkenbach and Matile, 1970). In potato the inhibitor was characterized as a
small protein, binding irreversibly to acid invertase (Pressy, 1967; Anderson and
Ewing, 1978). Pressy (1967) found that the binding of the potato inhibitor to
invertase had a pH optimum of 4.5 (Pressy, 1967), and the enzyme-inhibitor
complex could be partially disassociated by low pH or high Mg2+ concentrations.
In contrast, Bracho and Whitaker (1990a) found no effect of pH on inhibitor
binding. Sucrose at 2 mM could inhibit binding, but would not dissociate a
complex already formed (Pressy, 1967). Neither glucose nor fructose had a similar
effect. Matsushita and Uritani (1974) noticed a marked increase of acid invertase
activity resulted from wounding of sweet potato roots, but alkaline invertase activity
did not change under similar conditions. They also isolated a heat-stable protein
component with a molecular weight of approximately 19.5 kD, that fluctuated
during the incubation period after the wounding (Matsushita and Uritani, 1976).
They found that this putative inhibitor declined with a concomitant rise in invertase
activity early in the incubation, but increased in later stages when invertase activity
declined (Matsushita and Uritani, 1977). Pressy (1967, 1968) and Matsushita and
Uritani (1977) have suggested that the increase in invertase activity caused by cold
treatment or by wounding could be explained by a decrease in binding of the

25
inhibitor. Bracho and Whitaker (1990b) also found a positive correlation between
levels of inhibitor and invertase. The possibility therefore exists that this interaction
plays a regulatory role in sucrose breakdown (Akazawa and Okamoto, 1980;
Avigad, 1982).
Alkaline invertase is generally considered to be cytoplasmic. It is only
recovered from the soluble fraction of homogenates and has a pH optimum near
neutral. Both findings support its internal localization. Km values of alkaline
invertase for sucrose are slightly higher than for acid invertase, generally 9 to 25
mM. Alkaline invertase hydrolyzes raffinose very poorly (< 7% of the rate of
sucrose breakdown). Morell and Copeland (1984) found that stachyose (0.1 M)
also was hydrolyzed by alkaline invertase but much less efficiently than sucrose
(1.5% of the rate of sucrose). Both raffinose and stachyose are polysaccharides
containing a fructose moiety. Morell and Copeland (1984) also found that
cellobiose, gentiobiose, maltose, turanose, lactose, melezitose, trehalose, a-methyl-
D-glucopyranoside and /3-methyl-D-glucopyranoside (all at 0.1 M) were resistant
to degradative action by alkaline invertase. None of these sugars contain a
fructose moiety, further confirming the specificity of invertase for the fructofuranose
moiety of sucrose.
Alkaline invertase from potato tubers was inhibited only slightly by glucose
(Matsushita and Uritani, 1974); glucose-6-phosphate also had a slight inhibitory
effect. Fructose (15 mM) competitively inhibited soybean nodule alkaline invertase
by 50% (Morell and Copeland, 1984); glucose (5 mM) inhibited activity by 7%.

26
Morell and Copeland (1984) also found that the metabolites ATP, ADP, UDP, ADP-
glucose, UDP-glucose, glucose-1-phosphate, glucose-6-phosphate, and fructose-6-
phosphate (all at 5 mM) had no inhibitory effects. They also tested the effects of
various chloride salts on alkaline invertase activity and found that Na+, K+ or NH4+
at 50 mM had no effects; however, CaCI2 (10 mM) and MgCI2 (10 mM) each
inhibited activity by 25%. The anions citrate and inorganic phosphate have been
shown to stimulate alkaline invertase from Lupinus luteus nodules (Kidby, 1966);
however, Morell and Copeland (1984) found no effect on activity of soybean
nodule alkaline invertase. They did find, though, that Tris buffer was a
noncompetitive inhibitor of soybean nodule alkaline invertase activity; a 0.7 mM
buffer concentration inhibited activity by 50%.
Use of Mutants in Physiological Research
Despite the fact that all mutations have effects on the biochemistry and
physiology of the plant, only a small number have been investigated
physiologically (Vose, 1981). Advances in knowledge about the molecular bases
of cell processes in eukaryotic and prokaryotic microorganisms have been
achieved with an array of mutant lines, often induced, that modify or block steps
in the processes under study (Nilan et al., 1981). Many mutants will, theoretically,
differ in only a single major physiological character. The use of mutants is growing
in comparative physiological studies because the alternative is comparison of
contrasting genotypes that quite possibly may be altered in undefined characters
different from the one of interest.

27
The maize plant (Zea mays L.) has been particularly useful in genetic and
cytogenetic studies because of the number of mutants available (Neuffer et al.,
1968). Many of the mutants also have proven useful for physiological research.
The shrunken-1 mutant of maize was first described by Chourey and Nelson
(1976). Less than 10% of the normal sucrose synthase activity in wild-type
endosperm was observed. This reduced activity results in a "shrunken" phenotype
in the dry kernel. The sh 1 mutant has proven useful in elucidation of the role of
sucrose synthase in starch formation. The residual activity of sucrose synthase
present in this sh1 mutant was attributed to the presence of another isozyme
encoded by a second gene (Chourey and Nelson, 1976). This second gene,
Sus 1, has been mapped to the same chromosome as Sh 1 (chromosome 9) but
is located 32 map units away (McCarty et al., 1986; Gupta et al., 1988). Sh1 and
Sus 1 encode similar proteins. Sucrose synthase is a tetramer in its native form
(Su and Preiss, 1978), and the two isozymes encoded by Sh1 and Sus 1 are able
to form heterotetrameric forms of the native protein (Echt and Chourey, 1985).
Sh1 has been shown to be responsive to anaerobic conditions, with
transcript levels increasing 10 to 20 times in shoot and root tissue respectively
compared to aerobic controls (Springer et al., 1986). However, Sus1 exhibits little
response to anaerobic stress and seems to be expressed at a relative constant
in all tissues (McCarty et al., 1986). Rowland et al (1989), however, found that
Sus1 did show a slight response to anaerobic conditions, decreasing slightly in the
lower root, primarily in the pith, root tip and root cap. A maize mutant lacking the

28
Sus 1 gene has been described (Chourey et al., 1988), but, unlike the Sh 1 mutant,
the Sus 1 mutant does not have any detectable phenotypic abnormality. A
mutation lacking detectable levels of both sucrose synthase isozymes also has
been described (Chourey, 1988), but its existence is puzzling considering the
expected lethality of a complete sucrose synthase deficiency.

CHAPTER 3
INSTABILITY OF SUCROSE SYNTHASE FROM ROOT TIPS:
CHARACTERIZATION AND STABILIZATION
Abstract
Instability of sucrose synthase from root tips was characterized in maize and
an assay developed to circumvent the rapid decline of activity in vitro (35 and
100% activity loss in 20 min for maize and cotton, respectively). Initially de
suerse cleavage was quantified by recovery of 14C-UDPG on DEAE ion exchange
paper (Delmer, 1972; Su and Preiss, 1978). Subsequently, modifications were
made which resulted in increased accuracy, reduced tissue volume required and
reduced extraction/assay period. Phenolic protectants did not reduce the activity
loss over time. Specific inhibitors for the four classes of proteinases were also
tested; only PMSF increased enzyme activity, but did not completely prevent its
loss over time. Stabilization and additional elevation of activity were achieved by
adding casein. However, western blot analysis indicated that activity decline was
not associated with any detectable proteolytic degradation, charge alteration, or
subunit separation. In addition, inclusion of 10 mM P¡ in the extraction medium
rapidly reduced activity, indicating the possible involvement of phosphorylation or
nucleotide effects.
29

30
Introduction
Measurement of the maximum catalytic activities of enzymes in plant tissues
can make important contributions to the understanding of metabolic pathways and
their mechanisms of control (ap Rees, 1974). Currently available methods of
assaying sucrose synthase have proven ineffective for many tissues, particularly
those of roots (Duke et al., unpublished data; Lingle, USDA/ARS, Westlaco, TX,
personal communication). A precipitous loss of activity follows tissue extraction
from root tips of maize and other species (D.L. Hendrix, USDA/ARS, Phoenix, AZ,
personal communication). Chan et al. (1990) reported that sucrose synthase
activity in roots of rice was detectable in only one stage of growth. However,
sucrose synthase protein was present in root tissue at all stages of growth,
exceeding that in grain when grain activity was highest among tissues sampled.
This report addresses the basis of this instability in maize roots and describes a
rapid radiometric assay for sucrose synthase which circumvents this problem and
allows assay of small samples. Extraction and assay were optimized for substrate
concentration, pH, assay length, and inclusion (or exclusion) of various anti
oxidants and proteinase inhibitors. The procedure has proven effective for a range
of tissues and species examined and provides an accurate measurement of
activity, particularly where enzyme stability may be limiting.

31
Materials and Methods
Plant Material
Maize seed (Zea mays L, NK 508, VJ22:sh1) were primed for 6 days at 10
C with a water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I*1 captan
(Bodsworth and Bewley, 1981). At the end of 6 days, the seeds were rinsed free
of PEG, given a 20 min rinse in 1.05% (v/v) sodium hypochlorite and again rinsed
in water for 20-30 min. Seeds were germinated in the dark at 18 C on Whatman
3mm filter paper. Moisture level was kept constant throughout. At the end of 7
days, 1 cm primary root tips were excised under a sterile transfer hood. Cotton
(Gossypium hirsutum L, Coker 100) root tips were obtained from Dr. D.L. Hendrix
(Western Cotton Research Laboratory, USDA/ARS Phoenix, AZ). One cm root tips
were excised from 5- to 6-day old seedlings and quick frozen in liquid N2.
Purification of 14C-Sucrose
Trace amounts of phosphorylated sugars are common impurities in
commercial 14C-sucrose, and can reduce accuracy of the assay. These were
removed by descending paper chromatography of commercially obtained de
suerse in ethanol (NEN, Boston MA) using DEAE cellulose paper. The majority
of anion-free sucrose was concentrated into the first 2 to 3 drops eluted from the
V-shaped tip of the DEAE paper strip. No impurities were detected using HPLC
analysis (data not shown). Molarity and specific activity of purified 14C-Sucrose

32
were subsequently adjusted to 1 M and ca. 0.11 /Ci per /I. Two and one-half /I
(ca. 0.27 /iCi) were used in each reaction.
Tissue Extraction
Weighed tissue (100-200 mg) was frozen and ground to a powder in liquid
N2 with a mortar and pestle. The frozen powder was transferred to another mortar
containing ice-cold extraction buffer (200 mM HEPES buffer [pH 7.5], with 1 mM
DTT, 5 mM MgCI, 1 mM EGTA, 20 mM sodium ascorbate, 1 mM PMSF and 10%
[w/w] PVPP) and ground briefly in this medium. One ml of grinding buffer was
used for every 100 mg tissue fresh weight. Cysteine (10 mM) was initially but was
omitted to prevent non-specific binding of radiolabel to DEAE cellulose paper.
Two-hundred il of extract were placed on each of 4 to 8 spun columns
packed with Sephadex G 50-80 hydrated with extraction buffer. Columns were
centrifuged for 1 min at 800 x g. Eluent from each column was pooled with others
from the same tissue sample before assay. Ratio of sample to bed volume was
maximized at 1:5 (v:v) by HPLC detection of soluble sugar presence in eluent
(Yelle, 1991).
Enzyme Assay
Cleavage of 14C-sucrose by sucrose synthase was assayed in a 50 /I
volume consisting of 20 /I extract, 80 mM Mes (pH 5.5), 5 mM NaF, 100 mM de
suerse and 5 mM UDP. Reactions proceed for 5 minutes at 30 C and were
terminated by adding 50 /xl of Tris (pH 8.7) and boiling for 1 min. Controls

33
contained all assay components except UDP. The assay was optimized for pH,
linearity with time and protein concentration (data not shown).
Product Determination
The entire reaction volume was blotted onto a small disk of DEAE ion-
exchange paper (2.4 cm diameter) and dried completely before rinsing. Each disk
was rinsed separately, first in 40 ml of H20 at 175 rpm on a rotary shaker for 2
hours, again for an additional 2 hours and finally rinsed in a gentle stream of Dl
water for 30 sec. Remaining radiolabel was quantified and compared to total
amount of the 14C-sucrose substrate utilized to determine extent of sucrose
cleavage.
Protein Gel Blots
Subsamples from protein extracts to be used for enzyme assays were
separated on native PAGE using the system of Laemilli (1970) with (denaturing)
or without (native) SDS. Polyacrylamide concentrations of the stacking and
separating gels were 2.5% and 5%, respectively. Proteins were resolved at 4 C
by applying 15 V for 9 h, then 125 V for 11 h (constant current and temperature
(4 C). Each lane was loaded with 2 /g of total protein. Proteins were
electroblotted to nitrocellulose membranes and probed with polyclonal antibodies
following the procedure of Towbin et al. (1979). Sucrose synthase antisera,
obtained from D.R. McCarty, was generated in rabbits using protein purified from
maize kernels (W64 x 182E) 22 days after pollination. Antisera was diluted 1:1000

34
and cross reacted strongly to both the Sh1 and Sus 1 gene products where such
were present.
Results
Activity of sucrose synthase from maize and cotton root tips declined rapidly
after extraction (Figure 3-1). The greatest decrease in activity occurred between
10 and 15 minutes after extraction from both species. Little or no activity was
observed after 4 hours (data not shown). After extraction, extracts were
maintained at 0 C until used in the radiometric assay.
A wide range of enzyme protectants were examined. No improvement in
activity was observed when the polyphenol protectants PVP-40, PEG 20,000 and
BSA were utilized (Table 3-1). PVPP, also a phenol absorbent, was utilized in each
extraction. In addition, four classes of proteinase inhibitors were tested for their
effect on stability of sucrose synthase activity. Addition of leupeptin (1 mM) slightly
decreased initial activity (Table 3-1), and pepstatin-A (1 mM) had no effect (Table
3-1). Phenylmethylsulfonyl fluoride (PMSF) (1 mM), a serine proteinase inhibitor,
had a substantial positive effect, as did casein (2% w:v), potentially a non-specific
proteinase inhibitor.
Further characterization of activity change in the presence of PMSF showed
that stabilization was not effective in the first 20 min following extraction (Fig. 3-2).
Although total activity prior to this time was elevated by addition of PMSF, a linear
decrease was not prevented from occurring.

Sucrose cleaved (urnol mg protein h 1)
35
0 10 20 30 40 50
Time after extraction (min)
Figure 3-1. Time course of in vitro decrease in sucrose synthase activity in maize
and cotton roots. Bars represent SE, n=3.

36
Table 3-1. Effect of enzyme protectants on activity of sucrose synthase from maize
root tips assayed five minutes after extraction.
Protectant (concentration)enhancement of control
PVP-40 (5 %)
%
-6
PEG-20,000 (2% w:v)
+6
BSA (2% w:v)
+6
Caproic acid (2 mM)
0
Pepstatin A (1 mM)
-5
Leupeptin (1 mM)
0
PMSF (1 mM)
+46
Casein (2% w:v)
+ 15
Note: Each protectant was included in the buffer used for extraction and
equilibration of desalting columns. PVP-40, PEG-20,000 and BSA were utilized to
protect against phenolic compounds; PVPP was included in each extraction.
Representative inhibitors of proteinase classes were: pepstatin A (aspartic),
leupeptin (cysteine), EGTA (metallo) included in each extraction, caproic acid
(serine) and PMSF (serine). Casein was included as a general, non-specific
proteinase inhibitor.

Sucrose cleaved (jumol mg 1protein h'1)
37
Time after extraction (min)
Figure 3-2. Time course of /'/7 vitro decrease in maize sucrose synthase activity with
and without the serine proteinase inhibitor, PMSF. No other class-specific
proteinase inhibitors tested affected initial sucrose synthase activity. PMSF
(1 mM) was used in extraction buffer and equilibration of desalting columns.
Bars represent SE, n=3.

38
Addition of casein increased initial enzyme activity compared to controls
(Table 3-1) and improved stabilization of sucrose synthase activity with time (Figure
3-3A & B).
In contrast to added protectants, inorganic phosphate (10 mM), a protein
regulator through its role in reversible phosphorylation (Bennett, 1984), added to
the extraction buffer decreased initial activity of sucrose synthase by ca. 40%
(Figure 3-3A & B).
Despite loss of activity in vitro and positive responses to apparent
protectants against protease activity, proteolytic degradation of sucrose synthase
from maize root tips was not detectable via either denaturing or native (Figure 3-4A
& B, respectively) western-blot analysis at various times after enzyme extraction.
Further, no change in charge or separation of subunits in native tetramers was
evident. Nor was any change evident with time from samples extracted with
added casein or phosphate (Figure 3-4C & D, respectively). However, changes
in enzyme activity do not necessarily result in changes in electrophoretic mobility.
Walker and Huber (1989) demonstrated that activation of sucrose phosphate
synthase by light or mannose (a P¡ sequestering sugar) did not affect
immunoprecipitation or mobility of subunit mobility during SDS-PAGE.
The possible involvement of tetramer stability in the loss of activity in vitro
was further examined by comparison of the extracts from sh1 (containing only
homotetramers of Sus1 encoded sucrose synthase) and Sh1 (containing both
hetero- and homotetramers of sucrose synthase). Endosperm tissue contains

39
O 10 20 30 40 50 60 70
Time after extraction (min)
Figure 3-3. Time course of in vitro decrease in maize root sucrose synthase
activity in the presence and absence of either P. (10 mM) or casein (2%
w:v).

40
Time after Extraction
(min)
4 10 20 60
A
5 15 20 60
b
c
D nnnn
Figure 3-4. Denaturing (A) and native (B,C,D) protein gel blot analysis of maize
root sucrose synthase at various times after extraction. Sub-samples were
removed at designated intervals during incubation at 4 C. Proteins were
separated by polyacrylamide gel electrophoresis with (A) or without (B.C.D)
SDS and sucrose synthase resolved by probing with a polyclonal antibody
raised against protein products of both the Sh1 and Sus genes. The five
bands visible in the native gel blot have been described as corresponding
to homo- and heterotetrameric forms of sucrose synthase composed
entirely of products from the Sh 1 gene (uppermost band), the Sus gene
(lowermost band) and combinations of the two (middle three bands).
Extracts for denaturing (A) and native (B) blots were extracted with buffer
only. Casein (2% w:v) (C) and Pi (10 mM) (D) were tested as protectants
of enzyme stability.

41
tetramers composed only of Sh1 encoded protomers (Chourey et al., 1986;
Heinlein and Starlinger, 1989; Rowland and Chourey, 1990), and sucrose synthase
activity is stable during extraction and dialysis procedures (Chourey and Nelson,
1976; Echt and Chourey, 1985). In extracts from root tissue, the five bands shown
by western blot represent the possible combinations of monomers of the two
separate isozymes (Sh1 and Sus) to form the native tetrameric structure (Echt and
Chourey, 1985). Heterotetramers could theoretically be more unstable than
homotetramers since, although very similar, the two subunits are not identical (Echt
and Chourey, 1985). Su and Preiss (1978) found that sucrose synthase tended
to polymerize to an inactive polymeric after extraction. Results indicated that
formation of the native enzyme from two different isozymes was not a contributing
factor in loss of enzyme activity over time (Figure 3-5A & B). Despite differences
in the absolute values of sucrose synthase activity from sh1 vs. Sh1, the percent
decline in activity was similar for both genotypes.
Discussion
Sucrose synthase activity was stabilized in vitro and an assay developed
which enabled accurate measurement of enzyme action in root tips. The
described assay allows rapid product recovery in instances where activity is
otherwise unstable in vitro, and increases sensitivity to the extent that sample
volumes as small as 100 to 200 nQ can be used.
Sucrose synthase has previously been assayed in both synthetic and
cleavage directions (Avigad and Milner, 1966; Grimes et al., 1970; Pontis et al.,

Figure 3-5. Time course of in vitro decrease in maize root sucrose synthase activity from lines with homo- (W22:s/77) and
heterotetrameric (NK 508) forms of this enzyme. Bars represent SE, n=3. Data from one replicate using material
isogeneic to \N22:sh1 except for the Sh1 gene (W22) resulted in a curve similar in appearance and of the same
magnitude as that of W22:sh1.

Sucrose cleaved (pmol g_1 protein h'1)

44
1970; Salerno et al., 1979; Keller et al., 1988; Lowell et al., 1989). Measurements
of the synthetic reaction have been based on quantification of either sucrose or
UDP production. Sucrose levels can be determined indirectly by using invertase
for full conversion to hexoses and measuring glucose colorimetrically (Avigad and
Milner, 1966). The latter method, however, is susceptible to interference by
substances in the crude enzyme extracts of many plants (Pontis, 1977). It has
also been possible to measure 14C-sucrose formed from UDP-14C-glucose by
separating labeled product from substrate with anionic resins (Salerno et al., 1979),
paper electrophoresis (Grimes et al., 1970) or paper chromatography (Pontis,
1970). Such radioactive assays have proven useful in systems where colorimetric
methods have been problematic (Pontis, 1977). In addition, UDP production can
be determined spectrophotometrically by coupling its formation to the pyruvate
kinase-lactate dehydrogenase reaction and measuring the decrease in absorbance
due to oxidation of NADH (Avigad, 1964; Avigad and Milner, 1966; Lowell et al.,
1989).
Procedures for assaying the cleavage reaction are based on determination
of fructose or UDP-glucose formation. Fructose can be measured colorimetrically
by the Nelson reducing sugar assay (1944), or spectrophotometrically by coupling
hexokinase, phosphoglucose isomerase and glucose 6-phosphate dehydrogenase
for production of NADPH (Avigad, 1964; Keller et al., 1988). Also, UDP-glucose
formation can be estimated by coupling its appearance to NAD reduction by UDP-
dehydrogenase (Avigad, 1964; Lowell, 1986; Lowell et al., 1989). Degradative

45
action of sucrose synthase can also be coupled to that of UDP-
glucopyrophosphorylase (Xu et al., 1986; Sung et al., 1989).
Radioactive assays of sucrose synthase in the cleavage direction measure
the incorporation of 14C-glucose into UDP-glucose from 14C-sucrose (Delmer, 1972;
Su and Preiss, 1978). These procedures are among the most sensitive of assays
for sucrose synthase (Avigad, 1982). The sugar nucleotide formed can be
separated from the excess 14C-sucrose by paper chromatography (Wolosiuk and
Pontis, 1974a) or by ion exchange paper (Delmer, 1972a and b; Su and Preiss,
1978). The current procedure utilizes the sensitivity of a radiometric assay along
with reduced time from extraction to assay termination and results in a method
suitable for time-labile extracts from small tissue samples.
Instability of sucrose synthase was further characterized using this sensitive
method in an attempt to better define factors affecting the activity of this key
enzyme in vitro. Sucrose synthase is a sulfhydryl enzyme and is sensitive to
inhibition by phenolics and oxidized polyphenols (Pontis, 1977). Typical effectors
of activity reduction examined during the present study showed that phenolic
compounds did not appear to be the primary cause of the sucrose synthase
instability observed. No phelolic protectant was able to preserve sucrose synthase
activity over time.
Sucrose synthase has been shown to be sensitive to serine proteinases
(Wolosiuk and Pontis, 1974b). In their study, trypsin caused a 70% decline in the
degradative reaction and a 30% reduction in the synthetic reaction after a 15

46
minute incubation. However, chymotrypsin, also a serine proteinase, and papain,
a cysteine proteinase, had no effect. In the current study, PMSF, a serine
proteinase inhibitor, gave an increase in initial measurements but not did not
prevent the observed short-term loss of activity with time. Echt and Chourey
(1985) observed that PMSF did not stop the loss of activity of sucrose synthase
from maize endosperm during long term storage. No other specific proteinase
inhibitor affected stability of the enzyme from maize root tips. In addition to the
specific proteinase inhibitors, casein and BSA were included in some extractions.
Due to caseins complex composition and random structure, it undergoes
proteolysis with all the known proteolytic enzymes (Reimerdes and Klostermeyer,
1976). Casein increased initial measurements and stabilized activity with time
(Figure 3-3A & B). BSA, however, was much less effective. Casein has also been
found to preserve the longevity of sucrose synthase extracted from sugar cane (S.
Lingle, USDA-ARS, Westlaco, TX, personal communication). Casein (0.75-3.0%)
has also been shown to effectively stabilize and increase the initial activity
measurements of sucrose phosphate synthase (Raghuveer and Sicher, 1987).
Addition of casein is not always feasible, however, especially in instances where
accurate quantification of total tissue protein is important.
Another possibility for the regulation of sucrose synthase is through
phosphorylation. Many enzymes undergo reversible phosphorylation as a means
regulating activity (Bennett, 1984). Increased inorganic phosphate levels added
to buffers used for enzymatic extraction provide a substrate for protein kinase

47
activity (Bennett, 1984). Sucrose phosphate synthase (SPS), an enzyme of
carbohydrate metabolism, has been shown to be regulated in this manner
(Doehlert and Huber, 1983; Walker and Huber, 1989; Huber et al., 1989a).
Increases in extractable SPS activity are noted after illumination or inclusion of
mannose or glucosamine (phosphate sequestering agents) in darkness (Huber et
al., 1989b). Inorganic phosphate (5-10 mM) was found to be a potent inhibitor of
SPS (Amir and Preiss, 1982), with the inhibition becoming more sensitive in the
presence of Mg2+. Sucrose synthase has also been shown to have exhibit diurnal
fluctuations in activity (as does SPS) (Hendrix and Huber, 1986; Vassey, 1989) and
be affected by P¡. Pontis (1977) reported that P¡ (2-5 mM) inhibited the
degradative reaction of sucrose synthase alone or in the presence of Mg2+.
Delmer (1972a), however, found that 2 mM P¡ had no effect on the rates of either
the forward or reverse reactions. Sucrose synthase extracted from maize roots
has also been shown to be phosphorylated in vitro (Xu and Koch, University of FL,
unpublished data). In the current study, no effect of added phosphate on protein
stability was noted (Figure 3-4D). However, initial sucrose synthase activity
measurements were less than controls, and a loss of activity with time was
observed (Figure 3-3A & B).
The possibility existed that disassociation of subunits from tetramers may
have effected activity. Two separate isozymes in maize (Sh1 and Sus) form
subunits which appear to combine randomly into tetramers in root tips and other
tissues (Chourey et al., 1986). Endosperm sucrose synthase tetramers, however,

48
are almost entirely composed of subunits encoded by the Sh1 gene (Chourey and
Nelson, 1976; Chourey, 1981; Chourey et al., 1986) and are remain active during
extraction (Echt and Chourey, 1985). Five different types result in those tissues
which exhibit polymerization of both protomers; two are homotetramers and three
are heterotetramers. Heterotetramers could theoretically be more unstable than
homotetramers since, although very similar, the two subunits are not identical (Echt
and Chourey, 1985). Data (Figure 3-5A & B) indicate that greater instability of
heterotetramers relative to homotetramers of sucrose synthase was not the cause
of the observed activity loss. The profile of declining activity with time is similar in
extracts from root tips of a mutant line having only homotetramers of Sh1 (SS1)
subunits (y\/22:sh1) as it is in extracts from wild type kernels with 5 native
tetrameric combinations (NK 508).
Rapid loss of sucrose synthase activity with time in maize and other root
tips, as well as small tissue size, necessitated the development of a rapid and
sensitive assay. Other assays for sucrose synthase using radiometric techniques
have been described (Delmer, 1972a and b; Su and Preiss, 1978; Salerno et al.,
1979). However, the procedure described here has proven effective and useful
due to reduced time from extraction to assay termination, reduced sample size
and increased enzyme stabilization.

CHAPTER 4
SUCROSE SYNTHASE ACTIVITY IN WILD-TYPE MAIZE ROOT TIPS RESPONDING
TO ALTERED CARBOHYDRATE STATUS
Abstract
The two genes encoding sucrose synthase isozymes in maize (Sh1 and
Sus1) have been shown to respond to altered tissue carbohydrate status in root
tips; Sh 1 expression is favored by carbohydrate depletion whereas Sus 1 is up-
regulated when sugars are plentiful (Koch and McCarty, 1988, 1990; Koch et al.,
1989). Response at the level of enzyme activity was tested in the present study
by assaying sucrose synthase activity in excised maize root tips after 24 h of
incubation in a range of glucose concentrations. Little change was evident at the
level of total sucrose synthase activity; however, this represented the collective
responses of different isozymes and tissue types.
Introduction
Systems for changes in gene expression in response to altered
carbohydrate conditions have been reported for mammalian cells (Lin and Lee,
1984) and in bacteria and yeasts (Carlson, 1987; Schuster, 1989). Recently, seven
photosynthetic genes in maize protoplasts have been shown to be repressed and
coordinated by sugars (Sheen, 1990). Regulation in higher plants could have
49

50
important implications for the control of carbohydrate distribution and utilization.
Sucrose synthase is considered to have a key function in the allocation of sucrose
to various plant organs, and plant carbohydrate status could function as a means
of coarse regulation for activity of this enzyme.
Sucrose metabolism is important to the majority of plant species because
of the nearly ubiquitous role of this sugar in phloem transport to growing and
developing plant parts (Avigad, 1982). Two enzymes can catalyze the initial
breakdown of sucrose, invertase or the reversible enzyme sucrose synthase.
Recently, expression of the gene encoding the shrunken-1 isozyme of sucrose
synthase in maize has been shown to be sensitive to carbohydrate levels (Koch
and McCarty, 1988; Koch et al., 1989). Northern blot analysis of shrunken-1
mRNA showed levels were elevated in response to carbohydrate depletion. This
regulation is distinct from the previously characterized anaerobic induction
(Springer et al., 1986; Koch and McCarty, 1988). Although the anaerobic induction
of Sh1 has received considerable attention in several systems, questions remain
regarding the extent to which transcription and translation are synchronized under
these conditions. The anaerobic induction in maize has been reported to occur
only at the transcriptional level without concomitant changes in protein levels
(McElfresh and Chourey, 1988; Taliercio and Chourey, 1989). However, translation
of anaerobically induced sucrose synthase mRNA in rice (Ricard et al., 1991) has
been demonstrated. The present work examines enzyme-level responses to
changes in root carbohydrate status known to alter levels of Sh1 and Sus1 mRNA.

51
Regulation by sugar concentration may prove to be an important control
mechanism whereby plant cells are able to react to cellular carbohydrate status.
Materials and Methods
Maize seed (Zea mays L, NK 508) were primed for 6 days at 10 C at a
water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I'1 captan
(Bodsworth and Bewley, 1981). Seeds were subsequently rinsed free of PEG,
soaked for 20 min in 1.05% (v/v) sodium hypochlorite and rinsed for 20-30 min
with ca. 5 I of water. Seeds were germinated in the dark at 18 C on moist filter
paper in covered glass pans. Continuous airflow was provided (1 liter min"1)
throughout the germination period with 40% 02 supplied during the final 48 h. At
the end of 7 days, 1 cm primary root tips were excised under a sterile transfer
hood.
Excised root tips (ca. 750 mg per treatment) were incubated in 100 ml side-
arm flasks containing 50 ml of sterile Whites basal salt mixture (White, 1963)
supplemented with a range of glucose (0, 0.2, 0.5, 2.0, and 4.0%). Aerobic
conditions were maintained during 24 h incubations in the dark at 18 C by slow
agitation on a rotary shaker (125 rpm) and an airflow of 40% 02 (1 I min'1) through
an airstone in each flask. Experiments were terminated by twice rinsing in sterile
water, blotting excess moisture and freezing them in liquid N2.
Sucrose synthase activity was determined using a rapid radiometric
procedure developed to circumvent enzyme instability previously observed upon
extraction from maize root tips (Chapter 3).

52
Results
Sucrose synthase activity was consistently maximal in root tips
supplemented with 0.5% glucose (Table 4-1), a level at which the combined levels
of mRNA from the two sucrose synthase genes was also greatest (Koch et al.,
unpublished data). Overall, however, activity of sucrose synthase in whole root
tips was not significantly changed by alteration of carbohydrate status by
exogenous sugar supply (Table 4-1). It was not possible to distinguish activities
of isozymes encoded by the two sucrose synthase genes. These genes, Sh1 and
Sus 1, were found to exhibit reciprocal responses at the mRNA level to sugar
availability in the same sets of roots used for these experiments (Koch et al.,
unpublished data). Also, changes in distribution of sucrose synthase protein
among tissues within these root tips (Nolte, unpublished data) were not reflected
at the level of whole root enzyme activity.
Discussion
Reciprocal regulation of the two isoforms by carbohydrate levels, as has
been demonstrated for genes encoding for these isozymes (Koch and McCarty,
1989), could explain the lack of significant differences detected between glucose
treatments. The two isozymes of sucrose synthase from maize (encoded by the
Sh1 and Sus1 genes) are very similar, differing only slightly in their electrophoretic
movement during PAGE (Echt and Chourey, 1985). The Sh1 and Sus1 encoded
proteins are capable of catalyzing the same reaction with little difference in affinities

53
Table 4-1. Total sucrose synthase activity In wlldtype maize root tips incubated in
a range of glucose concentrations for 24 hours.
intacty
% glucose
0
0.2 0.5 2.0 4.0
(/mol sucrose mg'1 protein h'1)
Expt. 1
0.9
1.2
0.9
1.6
0.9
1.5
Expt. 2
0.5
0.5
0.4
0.6
0.6
0.6
Expt. 3
1.0
0.7
0.8
1.3
0.6
0.7
Mean
0.8
0.8
0.7
1.1
0.7
0.9
S.E.M.
0.2
0.2
0.2
0.3
0.1
0.
ylntact refers to root tips quick frozen in liquid N2 immediately after excision.

54
for substrates (Echt and Chourey, 1985), and both are present in extracts from
wildtype maize root tips (Chourey et al., 1986). Therefore the total amount of
measured sucrose synthase activity would be due to a combined complement of
sucrose synthase protein. Despite close homology, however, they have been
shown to be distinctive proteins encoded by separate genes (Chourey, 1981; Echt
and Chourey, 1985). The two proteins also are distinct in their localization within
the maize plant. The protein encoded by Sh 1 is primarily located in the
endosperm (Chourey and Nelson, 1976; Chen and Chourey, 1989), in the root
(Springer et al., 1986; Chourey et al., 1986) and in etiolated shoot (Springer et al.,
1986). The Sus 1 encoded protein is generally distributed throughout the plant
(Chourey, 1981; Echt and Chourey, 1985; Chourey et al., 1988).
Carbohydrate responsive proteins have been identified in roots of pearl
millet (Baysdorfer and Van der Woude, 1988). Webster and Henry (1987) have
also identified an unknown protein with a molecular weight similar to that of the
subunits of sucrose synthase in pea root meristem cells undergoing sugar
starvation. This protein, however, has yet to be positively identified. Initial findings
by Koch and coworkers (Koch and McCarty, 1988, 1990; Koch et al., 1989)
indicated that the Sh 1 gene of maize was stimulated by low carbohydrate
conditions and down-regulated under carbohydrate sufficient conditions. The Sus 1
gene responded in an inverse manner. Maas and co-workers (Maas et al., 1990)
demonstrated that the promoter from the Sh 1 gene was repressed by high

55
sucrose conditions. However, Salanoubat and Belliard (1989) found that increased
sucrose levels promoted genes encoding sucrose synthase in potato.
The possibility also exists that protein fluctuations did not occur after 24 h
incubation. However, the shifts in protein localization noted under the same
conditions (K. Nolte, University of Florida, unpublished data) indicate that some
protein level changes did occur. Spatial separation within root tissue also could
explain the differential response of Sh1 and Sus 1 observed at gene level without
a concomitant change in total enzyme activity. The distribution of sucrose
synthase isozymes has been shown to be developmentally regulated, and
changes during kernel development (Heinlein and Starlinger, 1989). Chen and
Chourey (1989) have reported that expression of sucrose synthase genes is
spatially and/or temporally separated in endosperm cells but not in root cells.
However, Rowland et al. (1989) demonstrated tissue specific localization of both
sucrose synthase genes and isozymes in roots undergoing anaerobic stress. K.
Nolte (University of Florida, unpublished data) has shown that shifts in sucrose
synthase protein localization occur in maize root tips under carbohydrate depleted
and carbohydrate sufficient conditions. Increases of one isozyme in a particular
tissue within the root along with decreases of the other in a different tissue would
not be apparent at the level of total root sucrose synthase activity. The lack of
significant differences in sucrose synthase activity of wildtype maize roots under
carbohydrate sufficient and depleted condition, therefore, does not demonstrate
that differences evident at the gene level are not also event at the translational

56
level. Occurrence of maximal enzyme activities in each experiment from samples
having the highest levels of both sucrose synthase genes, in fact, tends to indicate
that protein changes might be occurring but are not completely detectable under
the assay conditions utilized.

CHAPTER 5
SUGAR RESPONSE OF SUCROSE SYNTHASE AT THE GENE (Sus 7), PROTEIN
AND ENZYME ACTIVITY LEVELS IN ROOTS OF THE Sh1 MAIZE MUTANT
Abstract
The sh1 mutant of maize was used to study expression of the Sus 7 gene
for sucrose synthase in response to sugar availability because this mutant has only
one isozyme gene (Sus 7) for sucrose synthase and provides a system
uncomplicated by the presence of the second gene (Sh1). Koch and McCarty
(1988, 1990) have previously demonstrated that Sus 7 is up-regulated by plentiful
supplies of metabolizable sugars and down-regulated under carbohydrate
depletion, whereas Sh 1 responds in an inverse manner. Excised root tips from
sh 1 were incubated for 24 h in Whites basal salts medium supplemented with
different amounts of glucose. Sus 7 mRNA levels were approximately 5-fold greater
in treatments with 2.0% vs. 0% or 0.2% glucose. This difference was also reflected
in western blot analysis of sus protein. Enzyme activity was elevated 2-fold in root
tips from 2% glucose treatments vs. those in 0 or 0.2%. Time-course and
switching experiments showed that changes in mRNA or protein were not evident
until 24 h and indicated that the response to carbohydrate level had been initiated
within 16 h. Roots incubated in 2.0% glucose for 16 h and switched to 0% for 32
h (total of 48 h) responded like those remaining continuously in 2.0% glucose.
57

58
Overall, enhanced expression of Sus1 was evident at the mRNA, protein and
enzyme levels.
Introduction
Changes in gene expression by carbohydrates have been documented as
mechanisms by which bacteria and yeasts respond to changes in their nutrient
status (Carlson, 1987; Schuster, 1989). Glucose-responsive genes have also been
described in mammalian cells (Lin and Lee, 1984). In addition, Sheen (1990) has
presented evidence that the transcriptional activity of promoters of seven
photosynthetic genes from maize protoplasts are repressed and coordinated by
sugars. Regulation of this type in higher plants could have important implications
for carbohydrate allocation and utilization.
Sucrose and its metabolic products are important to almost all plant species
because of the nearly universal role of this sugar in growth and development
(Avigad, 1982). Initial breakdown of sucrose can be catalyzed by either invertase
or the reversible enzyme sucrose synthase. Recently, the gene encoding the
shrunken-1 isozyme of sucrose synthase in maize has been shown to be
responsive to carbohydrate levels (Koch and McCarty, 1988; Koch et al., 1989).
Gel blot analysis of shrunken-1 mRNA showed levels were elevated in response
to carbohydrate depletion. This may prove to be an important control mechanism
whereby plant cells are able to react to cellular nutritional conditions. The Sh1
gene has also been shown to be regulated by anaerobic conditions (Springer, et
al., 1986); however, effects on this gene by altered carbohydrate status are distinct

59
from regulation by anaerobic conditions (Koch and McCarty, 1988). The anaerobic
induction has been reported to occur only at the transcriptional level without
differences in protein levels (McElfresh and Chourey, 1988; Taliercio and Chourey,
1991). Possible changes in protein levels and enzyme activity of sucrose synthase
due to carbohydrate regulation have been difficult to detect because of the non
specificity of assay methods (Duke and Koch, unpublished).
The two isozymes of sucrose synthase from maize (encoded by the Sh1
and Sus1 genes) are very similar, differing only slightly in their electrophoretic
movement during PAGE (Echt and Chourey, 1985). Despite close homology, they
are distinctive proteins encoded by separate genes (Chourey, 1981; Echt and
Chourey, 1985). The two proteins are, however, distinct in their localization within
the maize plant. The protein encoded by Sh 1 is primarily located in the
endosperm (Chourey and Nelson, 1976; Chen and Chourey, 1989), in the root
(Springer et al., 1986; Chourey et al., 1986) and in etiolated shoot (Springer et al.,
1986); however Sh 1 mRNA does appear in other tissues such as pollen grains
(Hannah and McCarty, 1988). The Sus 1 encoded protein is more widespread in
its localization and is found throughout the plant (Chourey, 1981; Echt and
Chourey, 1985; Chourey et al., 1988). The distribution of both proteins is further
distinguished under stress conditions (such as anaerobiosis) where tissue-specific
localization in roots is readily apparent (Rowland et al., 1989). Tissue specific
shifts in sucrose synthase have also been noted in wildtype maize root tips
incubated in glucose deficient and sufficient media (K. Nolte, University of Florida,

60
unpublished data). Maraa and co workers (Maraa et al., 1990) have found that
the two genes encoding sucrose synthase in wheat (Ss1 and Ss2) also show a
differential response to stress conditions (anaerobiosis, cold shock and light).
Webster and Henry (1987) reported an unknown protein with a molecular
weight similar to that of the subunits of sucrose synthase in pea root meristem
cells undergoing sugar starvation. Carbohydrate responsive proteins have also
been found in roots of pearl millet (Baysdorfer and VanDerWoude, 1988). These
proteins, however, are yet to be definitively identified. Initial findings by Koch and
co workers (Koch and McCarty, 1988, 1990; Koch et al., 1989) indicated that the
Sh 1 gene of maize was stimulated by low carbohydrate conditions and down-
regulated under carbohydrate sufficient conditions. The Sus1 gene responded in
an inverse manner. Maas and co-workers (Maas et al., 1990) demonstrated that
the promoter from the Sh 1 gene was repressed by high sucrose conditions.
However, Salanoubat and Belliard (1989) found that increased sucrose promoted
genes encoding sucrose synthase.
The two genes encoding sucrose synthase in maize respond to altered
carbohydrate status (Koch and McCarty, 1988,1990; Koch et al., 1989), and shifts
in sucrose synthase protein localization have been observed under the same
conditions (K. Nolte, University of Florida, unpublished data). However, these
studies were carried out using a maize line having both sucrose synthase genes
present. The present study utilizes the Shrunken-1 mutant of maize to determine

61
the effects of varying carbohydrate conditions on the Sus1 gene and its sucrose
synthase gene product free from the confounding effects of Sh 1.
Materials and Methods
Maize seed (Zea mays L, \N22.sh1) were primed for 6 days at 10 C with a
water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I'1 captan
(Bodsworth and Bewley, 1981). Seeds were then rinsed with water, soaked for 20
min in 1.05% (v/v) sodium hypochlorite and rinsed again for 20-30 min with ca. 5
liters of water. Germination took place in the dark at 18 C on moist filter paper in
covered glass pans. Continuous airflow was provided (1 liter min-7) throughout
the germination period with 40% 02 supplied during the final 48 h. At the end of
7 days, 1 cm primary root tips were excised under a sterile transfer hood.
Excised root tips (ca. 750 mg per treatment) were incubated in 100 ml side-
arm flasks containing 50 ml of sterile Whites basal salt mixture (White, 1963)
supplemented with a range of glucose (0, 0.2, 0.5, 2.0, and 4.0%). Aerobic
conditions were maintained during 24 h incubations in the dark at 18 C by slow
agitation on a rotary shaker (125 rpm) and by an airflow 40% 02 (1 I'1) through an
airstone in each flask. Experiments were terminated by twice rinsing root tips in
sterile water, blotting excess moisture and freezing them in liquid N2.
Responses of root tips to incubation in 0% vs 2.0% glucose were examined
after 16, 24, or 48 h. Effects of treatment reversals at 16 h were also studied by
switching roots from 0% glucose treatments to 2.0% glucose and vice versa, then
continuing incubations for a total of 48 h.

62
Enzyme Assay
Sucrose synthase activity was determined by a rapid radiometric procedure
developed to circumvent enzyme instability previously observed upon extraction
from maize root tips (Chapter 3).
RNA Extraction and Northern Blotting
Samples were ground to a fine powder in a mortar and pestle with liquid N2
and RNA extracted according to McCarty (1986). Total RNA was quantified by
absorbance at 260 nm.
Total RNA was separated by electrophoresis in 1% agarose gels containing
formaldehyde (Thomas, 1980), blotted to a nylon membrane (Hybond-N,
Amersham Corporation, Arlington Heights, IL) and probed as per Church and
Gilbert (1984) with genomic clones of Sus1 (McCarty et al., 1986) radiolabeled by
random primer. Blots were rinsed and placed on X-ray film at -80 C.
Protein Gel Blots
Subsamples from protein extracts to be used for enzyme assays and
separated on native PAGE using the system of Laemilli (1970) without SDS.
Polyacrylamide concentrations of the stacking and separating gels were 2.5% and
5%, respectively. Proteins were resolved at 4 C by applying 15 V for 9 h, then 125
V for 11 h (constant current) and temperature (4 C).
Proteins were electroblotted to nitrocellulose membranes and probed with
polyclonal antibodies following the procedure of Towbin et al. (1979). Sucrose

63
synthase antisera, obtained from D.R. McCarty, was generated in rabbits using
protein purified from maize kernels (W64A x 182E) 22 days after pollination.
Antisera was diluted 1:1000 and cross reacted strongly to both the Sh 1 and Sus 1
gene products where such were present.
Results
Levels of Sus 1 mRNA in excised maize roots were greater after 24 h of
incubation in 2% glucose than in those that had received 0 or 0.2% glucose
(Figure 5-1). At the protein level, western blot analysis showed little or no change
with increasing carbohydrate concentration (Figure 5-2); however, enzyme activity
was elevated in root tips incubated at high vs. low glucose concentrations (Table
5-1). Both lines of evidence indicated the protein level response was less
pronounced at 24 h than that of mRNA.
The time-course of changes in Sus1 message levels in root tips showed that
differences between those given 0% vs. 2.0% exogenous glucose became
apparent sometime between 16 and 24 h (Figure 5-3). Initial decreases appeared
to occur in both treatments, but within 24 h, Sus 1 mRNA levels in glucose
supplemented roots had risen well above those with limited sugar supply. The
greatest difference between carbohydrate treatments was evident after 48 h of
incubation. Treatment reversals indicated that the gene response to carbohydrate
level had been initiated within 16 h (Figure 5-3). Roots incubated in 2.0% glucose
for 16 h and switched to 0% for 32 h (total of 48 h) responded like those remaining
continuously in 2.0% glucose. Roots initially deprived of glucose and then

64
% glucose
Intact 0 0.2 0.5 2.0 4.0
Expt. 1
Expt. 2
Figure 5-1. RNA gel blot analysis of Sus1 expression in maize roots incubated in
a range of glucose concentrations for 24 hours.

65
% glucose
Intact 0 0.2 0.5 2.0 4.0
Expt. 1
Expt. 2
Figure 5-2. Protein gel blot of Sus 1 encoded sucrose synthase from maize roots
incubated in a range of glucose concentrations for 24 hours. Data from
Expts. 1 and 2 were obtained from the same set of roots sampled for RNA
analyses shown in Figure 5-1.

66
Table 5-1. Sucrose synthase activity in mutant maize (W22:s/77) root tips
incubated in media containing a range of glucose concentrations for 24
hours.
Sucrose synthase activity
% glucose
Intact
0
0.2
0.5 2.0
4.0
(/xmol sucrose g1
protein h"1)
Expt. 1
0.22
0.13
0.07
0.13 0.32
0.32
Expt. 2
0.35
0.21
0.13
0.20 0.40
0.39

67
16h
Intact +
24h
+
48h 16h+32h
- + -/+ +/-
Figure 5-3. RNA gel blot analysis of Sus1 mRNA expression in maize roots
incubated in 0% (-) or 2.0% (+) glucose for various time periods. At 16 h
switching treatments were conducted by changing roots in 0% glucose to
2.0% glucose (-/+) and vise verse (+/-); incubations were continued for 32
additional hours.

68
switched to 2.0% glucose responded similarly to those remaining in 2.0% glucose
for the entire time.
At the protein level,changes in response to altered carbohydrate availability
were not apparent at 16 h, remained barely detectable at 24 h, but were clearly
evident after 48 h (Figure 5-4). Treatment reversals indicated that a protein-level
response occurred only when 16 h of elevated glucose treatment was followed by
32 h of glucose deprivation. The response was similar to that of root tips that had
remained continuously in 2.0% glucose. Slight differences in enzyme activity
between treatments were evident after 16 h or 24 h (Table 5-2); however activity
in glucose supplemented tips had risen to levels two-fold greater than those
without exogenous sugars within 48 h.
Discussion
The significance of results described here are two-fold. First, data
demonstrate that the differential response to changing carbohydrate availability by
the Sus1 gene for sucrose synthase is apparent at the translational level as well
as at the transcriptional level. The two genes encoding sucrose synthase
previously have been shown to respond differentially to carbohydrate supply (Koch
and McCarty, 1988; 1990; Koch et al., 1989). Differences at the protein and
enzyme level, however, have been difficult to detect due to cross reactivity of
polyclonal antibodies and the collective contribution of both isozymes to activity
measurements. Second, the resulting changes in physiology may allow the cells
to adjust their carbohydrate metabolizing capacity to the available supply. Use of

69
16h 24h 48h 16h + 32h
Intact + ~ + + "/+ +/-
Figure 5-4. Protein gel blot of Sus 1 encoded sucrose synthase from maize roots
incubated in 0% (-) or 2.0% (+) glucose for various time periods. At 16 h
switching treatments were conducted by changing roots in 0% glucose to
2.0% glucose (-/+) and vise verse (+/-); incubations were continued for 32
additional hours.

70
Table 5-2. Sucrose synthase activity in mutant maize (W22:s/7i) root tips
incubated in media containing in 0 or 2.0% glucose for various time periods.
Sucrose synthase activity
16h 24h
48h
16h + 32h
Intact
+ +
+
-/+
+/-
0.12
(/mol sucrose g'1
0.22 0.28 0.22 0.29
protein h1)
0.18 0.40
0.31
0.40
Note: 0 and 2.0% glucose represented by and +, respectively.

71
the Sh1 maize mutant has allowed the response to be characterized using a
simple, single enzyme system with respect to changing sugar supply. The
increased protein and enzyme activity evident at increased exogenous glucose
levels indicate that the plant tissue can adjust this first step in their sucrose-
metabolizing capacity relative to its carbohydrate status.
After 24 h at a given glucose level, changes in gene expression were more
marked than were differences at the protein and enzyme levels (Figure 5-1, Figure
5-2 and Table 5-1). This is not surprising given the probable presence of
previously formed RNA and protein (both appear to be relatively long-lived) as well
as the comparatively long-term progression of the response to the maximal extent
observed at 48 h (Figure 5-3 and Table 5-2). Chourey et al. (1991 b) reported that
the sucrose synthase gene in sorghum homologous to Sus 1 gene from maize is
anaerobically induced, but levels of the respective protein do not change.
Anerobic induction, however, was terminated after only 12 h. Anaerobic induction
of Sh1 in maize becomes apparent between 6 and 12 hours but message levels
are not maximal until at least 24 h (Duke and Koch, unpublished data). Data from
the present work indicate that like the respiratory drop noted by Brouquisse et al.
(1991), at least 20 hours are required before a change in Sus1 is fully apparent at
the gene level and even longer at the protein level. Nonetheless, data are
presented here at the levels of mRNA, protein and enzyme activity that indicate
that expression of the Sus 1 gene for sucrose synthase is responsive to
carbohydrate availability to an extent not evident in background levels of total RNA

72
and protein. The duration of time required for this response is consistent with the
proposed physiological function of sucrose in coarse adjustment of root growth
relative to sugar supply (Farrar and Williams, 1990).
The increased levels of Sus1 mRNA and subsequent elevation of its
respective protein with carbohydrate status may give insight into specific roles for
this isozyme as opposed to the Sh1 gene product. Sucrose synthase activity
could be key to the regulation of carbon entry into the respiratory pathway (Huber
and Akazawa, 1986; Black et al., 1987). The enhanced expression of Sus1 under
plentiful carbohydrate supplies accompanied probable increases in respiratory
activity in the root tips (Saglio and Pradet, 1980; Farrar and Williams, 1990;
Brouquisse et al., 1991). In addition, carbohydrate content in many tissues has
been correlated with the respiration rate (Penning de Vries et al., 1979; Farrar,
1985). Saglio and Pradet (1980) also found that an exogenous supply of 0.2 M
glucose was required to bring the respiration rate of excised maize roots back to
the level of intact tissue, indicating that the rate of metabolic activity of the root tips
may be closely tied to sugar import. Perhaps another line of evidence supporting
control of respiration by carbohydrate status comes from the work of Douce et al.
(1990) in which sycamore cells in culture showed loss of mitochondrial function
when starved of sucrose; the beginning of the decline coincided with the fall in
endogenous sugar concentrations.
Another possible role for the Sus 1 gene product may be in the diversion of
carbohydrate to cell wall biosynthesis. Roots in 2.0% glucose medium show

73
marked growth during the 24 hours of incubation (data not shown). During this
time, there is a demand for cell wall synthesis by the expanding cells. Sucrose
synthase has been implicated in the directing of carbohydrates for polysaccharide
biosynthesis (Amino et al.,1985; Hendrix, 1990). The level of involvement for
nucleotide-sugars during cell wall polysaccharide biosynthesis has been implicated
in a need for greater activity of this enzyme (Maas et al., 1990). Sucrose synthase
activity was elevated in cell cultures of Catharanthus roseus during the G1 phase
when total amounts of cell wall biosynthesis increased significantly (Amino et al.,
1985). Sugar modulation of Sus1 could convincingly combine production of cell
wall precursors with other aspects of increased growth (Farrar and Williams, 1990)
likely to accompany an enhanced sugar supply.
The carbohydrate response of sucrose synthase in the present study differs
from previously demonstrated regulation in that it occurs in rapidly growing and
metabolizing structures. Other studies have involved sucrose synthase regulation
in storage tissues where processes such as starch accumulation predominate.
Loss of the shrunken-1 gene in maize results in a typical endosperm phenotype
where starch deposition is reduced by over 70% (Chourey and Nelson, 1976);
however, a mutant lacking a functional Sus 1 gene has no apparent phenotype
(Chourey et al., 1988). Starch deposition accompanies protein accumulation in
developing potato tubers and levels of mRNA encoding sucrose synthase have
been shown to increase in this tissue (Salanoubat and Belliard, 1989). Levels of
storage proteins, such as patatin, also accumulate during this time (Paiva et al.,

74
1983). Increased levels of sucrose result in elevated levels of genes for both
sucrose synthase (Salanoubat and Belliard, 1989) and patatin (Rocha-Sosa et al.,
1989; Wenzler et al., 1989) in tissues where they are not usually found. A rise in
sugar availability can also result in increased transcription and translation of a
unique storage protein in stem and leaf tissues of sweet potato (Hattori et al.,
1990). The diversity of processes operating in the system utilized in this study
suggests that sugar-responsive gene expression (ie. Sus 1) may have broad
implications in the formation and function of non-storage plant tissues.

CHAPTER 6
AN ORGAN-SPECIFIC INVERTASE DEFICIENCY IN THE PRIMARY ROOT OF AN
INBRED MAIZE LINE
Abstract
An organ-specific invertase deficiency affecting only the primary root system
is described in the Oh 43 maize inbred. Invertases (acid and neutral/soluble and
insoluble) were assayed in various tissues of hybrid (NK 508) and inbred (Oh 43,
W22) maize lines to determine the basis for an early report that Oh 43 root tips
were unable to grow on sucrose agar (Robbins, 1958). Substantial acid invertase
activity (7.3 to 16.1 ^mol glucose mg'1protein h'1) was evident in extracts of all
tissues tested except the primary root system of Oh 43. This deficiency was also
evident in lateral roots arising from the primary root. In contrast, morphologically
identical lateral roots from the adventitous root system had normal invertase levels.
These results suggest that ontogenetic origin of root tissues is an important
determinant of invertase expression in maize. Adventitious roots (including the
seminis) arise above the scutellar node and are, therefore, of shoot origin. The
Oh 43 deficiency also demonstrated that invertase activity was not essential for
maize root growth. Sucrose synthase was active in extracts from all root apices
and theoretically provided the only available avenue for sucrose degradation in
75

76
primary root tips of Oh 43. The deficiency described here will provide a useful
avenue of investigation into the expression and significance of root invertase.
Introduction
Sucrose breakdown is critical to the vast majority of plant species because
non-photosynthetic tissues depend on import of this sugar for their growth and
development. Initial cleavage of sucrose can be catalyzed by either invertase or
the reversible enzyme sucrose synthase. Invertases are especially active in tissues
undergoing rapid cell division such as shoot and root apices (Avigad, 1982).
Previously, the role of invertases in sucrose transfer was considered particularly
important in plants such as sugar cane and maize where substantial sugar
movement occurred through the cell wall space and was accompanied by action
of an extracellular invertase (Glasziou and Gayler, 1972; Hawker and Hatch, 1965).
Recent evidence, however, indicates that although much hydrolysis is often
observed, invertase activity may not be essential for sucrose uptake into either
sugar cane stems (Lingle, 1989; Thom and Maretzki, 1990) or maize kernels
(Schmalstig and Hitz, 1987).
Sucrose generally is believed to enter root tips without traversing the
extracellular space (Giaquinta et al., 1983); however growing roots can differ
markedly in their capacity to lose (Rovira and Davey, 1974) and retrieve (Robbins,
1958) exogenous sugars. Net losses do occur. The extent of sugar efflux from
roots can be affected by irradiance level, nutritional status, moisture availability and

77
temperature (Rovira and Davey, 1974). The composition of root sugars exuded
is quite variable but includes both reducing and non-reducing sugars (Rovira and
Davey, 1974). Glucose and fructose are often taken up from the extracellular
space more rapidly than is sucrose (Humphreys, 1974). Retrieval of solutes from
the apoplast and the form in which they are available may thus be a potentially
important attribute of root carbon balance.
A deficiency in this retrieval process was first indicted by Robbins report
(1958) that roots of a maize inbred, Oh 43, were unable to grow on sucrose agar
medium, yet roots of another line, Hy 2, grew quite well. Only when roots of both
were cultured immediately adjacent to one another, were those of Oh 43 able to
grow. Growth of excised Oh 43 root tips also occurred when glucose was
substituted for the sucrose. Oh 43 was concluded to be "incapable of inverting
sucrose" in its root tips. Preliminary investigations by B. Burr (Brookhaven National
Laboratory, personal communication) indicated that a lack of invertase may have
been the reason for the inability of Oh 43 roots to metabolize sucrose.
The absence of invertase activity could have important implications for
sucrose import not only because of potential effects on the retrieval system, but
also because sucrose utilization in such an instance could theoretically be initiated
only via action of sucrose synthase. In addition, genetic material which lacks
activity of a specific enzyme can be useful in investigations of physiological
processes normally mediated by these enzymes (Koch et al., 1982). The present
report demonstrates that invertase is not essential for primary root growth despite

78
probable advantages of its presence and indicates an unusual organ-specific
difference in expression between primary and adventitious roots.
Materials and Methods
Plant Material
Maize seed (Zea mays L. NK 508, W22 and Oh 43) were germinated on
moist filter paper in petri dishes. Seeds were imbibed for 24 hours and pericarps
removed, allowing more uniform germination and more effective surface
sterilization (20 min soak in 0.525% sodium hypochlorite).
Five successive 2 mm segments were sampled from the tips of primary
roots 4 to 5 days after germination. Intact roots of Oh 43 seedlings grew more
slowly than did those of NK 508 or W22, but all roots had reached 2 cm prior to
excision. Tissue samples were weighed, frozen in liquid N2 and stored at -80 C
until assayed for invertase activity. In subsequent experiments, 5 mm root tips
were excised from primary and adventitious roots for invertase and sucrose
synthase activity measurements. Plants and tissues were as above.
Tissue Extraction
Frozen tissue samples were ground to a fine powder in liquid N2 using a
mortar and pestle. Frozen powder was transferred to a second mortar containing
ice-cold 200 mM HEPES buffer (pH 7.5) with 1 mM DTT, 5 mM MgCI, 1 mM EGTA,
20 mM sodium ascorbate and 10% (w/w) PVPP. One ml of extraction buffer was
used for every 100 mg of tissue fresh weight. Buffered extract was centrifuged at

79
14,000 x g for 1 min to sediment particulate matter. Supernatant was dialyzed
(27,000 mw cutoff) at 4 C for 24 h against extraction buffer diluted 1:40. Buffer
was changed after 1 h and thereafter, every 4 h. Soluble dialyzed extract was
assayed for invertase as described below. Previously separated particulate matter
was rinsed with one volume of extraction buffer and assayed for insoluble, cell-
wall-bound invertase (soluble acid invertase includes both vacuolar and loosely
bound extracellular enzyme [Avigad, 1982]).
To test the possibility that the soluble enzyme was present in primary roots
of Oh 43 but was being bound or inactivated during the extraction procedure, two
additional extraction/assay methods were employed. First, adventitious root
extracts, previously shown to contain active invertase activity, were added to those
of primary apices. The resulting mixture was dialyzed and assayed for enzyme
activity. Second, three cm apices of both primary and adventitious roots Oh 43
roots were excised. Apices of these roots (0.5 cm) were suspended in extraction
buffer for three hours at 27 C. Buffer alone was subsequently dialyzed as
described above. The portion of each root which had been immersed in the
extraction buffer was excised for fresh weight measurement. After dialysis, the
buffer-enzyme solution was analyzed for enzyme activity.
Enzyme Assays
Soluble and insoluble forms of acid invertase were assayed as described
by Lowell et al. (1989). Reaction media contained 50 mM sucrose, and pH of 4.5
was adjusted with a sodium acetate buffer. Neutral invertase was assayed using

80
the same reaction medium adjusted to pH 7.5 with potassium phosphate buffer.
Initial assays were also performed at pH ranges of 4.0 to 5.5 for acid invertase and
7.0 to 8.0 for neutral invertase. After a 15 min incubation at 30 C, glucose
production was quantified by the glucose oxidase method (Sigma Chemical Co.).
Sucrose synthase was assayed in the degradative direction using a radiometric
assay quantifying the production of 14C-UDPG (Chapter 3).
Histochemical Staining
Free-hand cross sections from apices of both primary and adventitious
roots were fixed in 4% formalin (pH 7.0) for 30 min and rinsed in water at least 10
times over a period of 3 hours to remove endogenous sugars (Doehlert and
Felker, 1987). Sections were then incubated in a sodium phosphate buffer (0.38
M, pH 6.0) containing 0.24 mg ml'1 nitroblue tetrazolium, 0.14 mg ml'1 phenazine
methosulfate, 25 units ml'1 glucose oxidase and 5 mg ml'1 sucrose (Doehlert and
Felker, 1987). Control sections were incubated in the same mixture without
sucrose. After rinsing in water, sections were post fixed in 4% formalin (pH 7.0)
and photographed under a microscope.
Results
Primary roots of Oh 43 showed little or no acid invertase activity (Table 6-1).
In contrast, acid invertase was active in extracts from apical areas of roots from
other maize lines examined (NK 508 and W22). Activity, per unit fresh weight, was
greatest in root apices, decreasing with distance from the tip until no longer

Table 6-1. Soluble and insoluble acid invertase activity in sequential 2 mm segments of primary
roots of 5 to 6 day-old seedlings from 1 hybrid and 2 inbred lines of Zea mays.
Root
Solubley
Insoluble
Segment
Oh 43
NK 508
W22 Oh 43
NK 508 W22
/mol glucose g'1FW h'1
0-2 mm
2-4 mm
4-6 mm
6-8 mm
8-10 mm
0.2 0.2W
43.8 5.2
32.8 2.2
20.6 5.0
1.4 0.9
40.2 3.7 z
28.6 3.1
15.4 4.6
0.9 0.5
3.8 0.5 3.6 0.8
3.1 1.9 2.7 1.5
z-not detectable
y-Soluble activity is expressed per unit FW to allow comparison with insoluble activity.
"--Activity in root tips from axenic culture was 0.0 0.0 /mol g'1FW h'1; thus microorganisms are a
likely source of the residual activity in ca. 1 of 3 assays.
Each value represents the mean of 3 separate samples SEM.

82
detectable farther than 8 mm from the apex. Soluble enzyme accounted for 88 to
92% of the total acid invertase activity, and the remainder was due to action of the
insoluble enzyme. Neutral invertase activity was insignificant or absent from all
lines (data not shown). Although invertase action was essentially undetectable in
the primary roots of Oh 43, adventitious root extracts showed levels of activity as
least as great as those from the primary roots of other lines tested (Table 6-2).
Active acid invertase was released into buffer in the extracellular space
around adventitious root tips of Oh 43 during 3 hours of emersion (13.4 3.5 /imol
glucose g'1 FW h'1-ca. 25% of the activity on a fresh weight basis of N2 and buffer
extracted roots). The same was not observed during emersion of primary roots
from this line. Also, from 83% to 94% (13.3 to 15.1 Mmol glucose mg1 protein h"1)
of the invertase activity in adventitious root extracts remained after addition of
extracts from primary roots of Oh 43.
Unlike invertase, sucrose synthase was active in extracts of both primary
and adventitious roots from Oh 43 seedlings. Activity of this enzyme was
approximately similar in both root types, as observed for the other two lines tested
(Table 6-2).
Extracts of lateral roots originating from primaries exhibited no detectable
acid invertase activity (Table 6-3), in contrast to counterparts derived from
adventitious roots. Shoots of 5-day-old Oh 43 seedlings and endosperm or
scutellum tissue from developing kernels (23 DAP) also exhibited activity of both
soluble and insoluble acid invertase (Table 6-3).

Table 6-2. Soluble acid invertase and sucrose synthase activity in 0.5 cm apices of primary and
adventitious roots of 5-to 6-day-old seedlings from 1 hybrid and 2 inbred lines of Zea mays.
Oh 43
NK 508
W22
primary
adventitious
primary
adventitious
primary
adventitious
/mol glucose mg"1 protein h"1
Invertase
0.1 o.iw
16.1 3.2
16.0 2.5
14.0 2.5
12.6 2.2
11.9 1.6
Sucrose
Synthase
1.2 0.1
1.1 0.1
1.0 0.1
1.1 0.2
1.0 0.1
0.9 0.1
/mol glucose g"1 FW h'1
Invertase
Sucrose
0.2 0.2W
60.8 12.1
56.5 8.7
51.6 9.2
53.0 9.3
48.9 6.7
Synthase
4.7 0.4
4.2 0.1
3.5 0.5
4.2 0.6
4.0 0.4
3.7 0.4
W--Activity in root tips from axenic culture was 0.0 0.0 mol g'1FW h'1; thus microorganisms are
a likely source of the residual activity in ca. 1 of 3 assays.
Each value represents the mean of 3 separate samples SEM.

84
Table 6-3. Soluble add invertase activity in various tissues of
Oh 43, an inbred line of Zea mays.
/imol glucose
nmol glucose
g'1FW h'1
mg'1 protein h'1
Shoot
10.8 0.6
25.8 1.5
Endosperm
7.3 1.2
12.6 2.1
Scutellum
9.0 0.8
19.4 1.7
Primary roots
Lateral roots from
0.1 0.1
0.2 0.2
10 roots
0.1 0.1w
0.2 0.1W
Adventitious roots
Lateral roots from
16.1 3.2
60.8 12.1
adventitious roots
10.4 0.9
38.7 3.2
W-Activity in root tips from axenic culture was 0.0 0.0 /mol
g'1FW h'1; thus microorganisms are a likely source of the
residual activity in ca. 1 of 3 assays.
Shoot, primary roots and adventitious roots were
samples from 6-day-old seedlings. Endosperm and scutellum
were from developing kernels 23 DAP. Lateral roots
developed after 2 to 3 weeks of growth and were then
excised. Each value represents the mean of 3 separate
samples S.E.M.

85
Histochemical staining indicated that no invertase activity was detectable in
the primary roots of Oh 43 (Figure 6-1). Cross sections of primary and
adventitious roots of Nk 508 and adventitious roots of Oh 43 all stained positive
for invertase activity. Invertase activity was primarily in the cortex and was
localized intercellularly.
Discussion
These data confirm that Oh 43 (an inbred line of maize) lacks invertase
activity in its primary root tips. Surprisingly, however, a deficiency was not evident
in structurally and functionally similar adventitious roots (Table 6-2), other tissues
of the same Oh 43 plants (Table 6-3) or in the developing kernels of this line
(Doehlert, et al., 1988). The genetic potential for invertase expression is therefore
present. The lack of activity may result from altered regulation of gene expression
(transcription or translation), the loss of a tissue specific invertase isozyme or the
existence of an unidentified effector of enzyme function. The significance of results
described here is twofold. First, evidence is presented for differential expression
of invertase in morphologically identical organs that differ primarily in point of
origin. This is most strikingly illustrated in the apparent distinction between lateral
roots arising from primary and adventitious root systems. Differential expression
of genes in morphologically identical structures are unusual; however, organ or
tissue-specific differences have been well documented (Fluhr et al., 1986; Xie and
Wu, 1989). Xie and Wu (1989), for example, found that genes for alcohol
dehydrogenase were differentially expressed in root and shoot tissues of rice

Figure 6-1. Histochemical localization of ¡nvertase activity in free-hand, fresh cross
sections of root apices of 6-day-old maize seedlings. Sections are
approximately 50 /xm in thickness. Primary (A) and adventitious (B) root
cross sections from NK 508 incubated in reaction medium without sucrose
(controls). Primary (C) and adventitious (D) root cross sections from Oh 43
incubated in reaction medium (note only adventitious root section exhibits
blue formazan reaction product [dark areas]). Primary (E) and adventitious
(F) root cross sections from NK 508 incubated in reaction medium (note
both sections exhibit reaction product [dark areas]). Bar represents 50 /xm.

87

88
plants. One isozyme predominated in shoot-derived organs (leaves, sheaths,
nodes and pollen) and the other isozyme showed highest activity in the roots. It
is interesting in this respect that the adventitious roots of maize arise above the
scutellar node of the developing seedling and are, therefore, derived from shoot
tissue (Hayward, 1938). The difference in invertase expression in primary and
adventitious root systems may reflect a similar root/shoot dichotomy. Therefore,
the tissue of origin and cell lineage may be more important than organ identity and
function in regulating invertase expression. Other variants in invertase expression
have been described. Echeverra and Humphreys (1984) reported a hybrid maize
line (DKXL80) which exhibited no soluble invertase activity in contrast to previously
tested lines. Other tissues of this maize line exhibited normal invertase activity.
Second, data demonstrate that invertase is not essential for primary root
growth. The primary roots of Oh 43 exhibited no signs of premature senescence,
and, if left intact, continued apparently normal growth for many days (data not
shown). Invertases role in sucrose import into roots has been questioned by
Chapleo and Hall (1989a) who concluded that although present, apoplastic root
invertase did not have a direct role in sugar transport in Ricinus. However,
substantial activity of invertase has been widely documented in roots of plants
such as pea (Lyne and ap Rees, 1971), bean (Robinson and Brown, 1952), tomato
(Chin and Weston, 1973), Ricinus (Chapleo and Hall, 1989a, b and c), oat (Pressy
and Avants, 1980), and maize (Chang and Bandurski, 1963; Hellebust and
Forward, 1962). In maize, the Oh 43 invertase deficiency apparently prevents

89
utilization of exogenous sucrose (Robbins, 1958). Specific tissue localization of
invertase also has been described. Peak activity for root invertase is generally 2-3
mm behind the apex and corresponds to the region of expansion and elongation
in pea (Robinson and Brown, 1952) and maize (Hellebust and Forward, 1962). In
Ricinus this activity predominates in the cortex (Chapleo and Hall, 1989).
Although invertase may not have a direct role in sucrose import in roots, it
still may be important to two major aspects of root biology. First, invertase has
been implicated in formation of mycorrhizal associations (Purves and Hadley,
1975). Maize (Gerdemann, 1964; Kothari et al., 1990) and 90% of other
agriculturally important species form these beneficial symbioses under field
conditions (Gerdemann, 1986). Invertase activity typically increases and hexose
levels rise at infection sites of biotrophic fungi (Long et al., 1975). Elevated hexose
content in roots upon infection by mycorrhizal fungi has been attributed to a rise
in invertase levels (Purves and Hadley, 1975). It is not known whether this is host
or fungal invertase; however, Oh 43 does not appear to provide the former in its
primary root systems.
Second, the purported lack of a sucrose carrier in the plasma membrane
of maize root cells (Lin et al., 1984) would indicate that if sucrose were released
into the apoplast, retrieval might proceed more effectively in the presence of
extracellular invertase. Such retrieval could be particularly important during stress
or periods when sugar losses from the symplast were elevated. Any physiological
consequence of this deficiency would most likely be evident early in seedling

90
development because the root system would consist solely of a primary root at
this time. Later in seedling development, adventitious or seminal roots rapidly take
over a dominant role, leaving the primary root with little or no essential function
(Hayward, 1938). In the field, lines having Oh 43 as a progenitor have been
observed to suffer from poor emergence rates in damp soils (B. Martin, Pioneer
Seed, personal communication). However, conclusive evidence of a physiological
effect will require generation of and analysis of isogeneic lines.
In conclusion, the deficiency described here will be useful for investigation
into the regulation and physiological function of root invertase. Because the
primary root system in maize is nonessential, invertase deficient roots can be
studied without deleterious effects on the overall physiology of the plant. In
addition, this deficiency is significant because it reveals an unexpected distinction
between primary and adventitious root development in maize. At some level, the
mechanisms which regulate invertase in these root systems must differ.

CHAPTER 7
SUMMARY AND CONCLUSIONS
Sucrose metabolism is important to the majority of plant species because
of the widespread role of this sugar in growth and development through its
function as a phloem transport sugar. Initial breakdown of sucrose can be
catalyzed by either sucrose synthase, a reversible enzyme, or invertase.
Expression of genes encoding the sucrose synthase isozymes (Sh1 and
Sus 1) have been found to be sensitive to carbohydrate levels (Koch and McCarty,
1988, Koch et al., 1989). Northern blot analysis of mRNA showed levels showed
those of Sh1 were up-regulated in response to carbohydrate depletion whereas
those of Sus 1 were down-regulated under the same conditions. This may prove
to be an important control mechanism whereby plant cells are able to react to
cellular carbohydrate status.
The first purpose of this work was to determine the extent to which
carbohydrate-modulated changes in message levels affected activity of sucrose
synthase the level of enzyme activity. However, previously available sucrose
synthase assay methods proved ineffective for maize root tips due to a precipitous
loss of activity after tissue extraction. A rapid, radiometric assay for sucrose
synthase was therefore developed that overcomes these obstacles. Extraction and
assay were optimized for factors such as substrate concentration, pH, assay
91

92
length, and inclusion (or exclusion) of various anti-oxidants and protease inhibitors.
The assay has proven effective for a range of tissues and species examined and
provides a particularly effective measurement for sucrose synthase action in root
tips. Characterization of the in vitro instability also provided evidence suggestive
of possible phosphorylation effects on activity.
Activity of sucrose synthase was then assayed in extracts from intact
wildtype maize root tips and from those that had been excised and incubated for
24 h in a basal salts medium with varying levels of glucose. Enzyme activity
showed little or no significant differences between treatments. However, results
represented contributions by two isozymes (encoded by genes exhibiting
reciprocal responses under the same conditions [Koch et al., 1989]), and were the
collective sum of different tissues (that showed specific alterations in sucrose
synthase distribution [K. Nolte, University of Florida, unpublished data]).
A different approach was thus utilized to examine the relationship between
sugar-modulated changes in message levels and extent of enzyme level effects.
Use of the shrunken-1 maize mutant (deficient in a functional Sh 1 gene) showed
that carbohydrate-responsive gene expression was evident for Sus1 at the levels
of mRNA, protein and enzyme activity. Time-course and treatment reversals also
indicated greater responses between glucose sufficient and deficient treatments
were observed after 48 h of incubation, indicating a possible coarse control
mechanism.
The second purpose of this work was to explore a possible line of

93
investigation into the physiological significance of invertase by investigating a
putative deficiency in sucrose-metabolizing capacity in the Oh 43 line of maize
(Robbins, 1958). Complete absence of invertase in this line was not likely due to
its demonstrated activity in the scutellum (Doehlert et al., 1988). However, the
hypothesis tested here was that an organ-specific deficiency or invertase
suppression was responsible for the metabolic anomalies in Oh 43. The primary
root of Oh 43 indeed was shown to lack invertase activity. In contrast, adventitious
roots of the same plants exhibited wildtype levels of invertase activity. Initial
characterization of this mutation will provide an effective tool for future
investigations into the physiological role(s) of this enzyme in roots. Use of this
mutation and isogeneic wildtype counterparts in combination with the nulls for
each sucrose synthase gene may allow a better understanding of the biological
functions of each.

LITERATURE CITED
Akazawa, T. and K. Okamoto. 1980. Biosynthesis and Metabolism of Sucrose, pp.
199-220 in The Biochemistry of Plants. New York: Academic Press.
Al-Bakir, A.Y. and J.R. Whitaker. 1978. Purification and properties of invertase from
dates (Phoenix dactylifera L. var. Zahdi) J. Food Biochem. 2:32-160.
Amino, S., Y. Takeuchi and A. Komamine. 1985. Changes in enzyme activities
involved in formation and interconversion of UDP-sugars during the cell
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95:669-674.
Ricardo, C.P.P. 1974. Alkaline /S-fructofuranosidases of tuberose roots: Possible
physiological function. Planta 118:333-343.
Ricardo, C.P.P. and T. ap Rees. 1970. Invertase activity during the development
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cell specific expression of Adh-1, Sh and Sus genes in roots of maize
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Rowland, L.J. and P.S. Chourey. 1990. In situ hybridization analysis of sucrose
synthase expression in developing kernels of maize. Maydica 35:373-382.
Saglio, P.H. and A. Pradet. 1980. Soluble sugars, respiration, and energy charge
during aging of excised maize root tips. Plant Physiol. 66:516-519.

109
Salanoubat, M. and G. Belliard. 1989. The steady-state level of potato sucrose
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endosperm UDP-glucose phosphorylase activity. Plant Sci. 44:111-118.
Salerno, G.L., S.S. Gamundi and H.G. Pontis. 1979. A procedure for the assay of
sucrose synthetase and sucrose phosphate synthetase in plant
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Sampietro, A.R., M.A. Vattuone and F.E. Prado. 1980. A regulatory invertase from
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BIOGRAPHICAL SKETCH
Edwin Ralph Duke was born in Bainbridge, Georgia on February 22,1960.
He attended public school in Rossville, Georgia, and graduated from Macon
Christian Academy, Macon, Georgia, in May 1978. In June 1980 he received his
Associate of Science degree from Macon Junior College, Macon, Georgia,
majoring in agriculture. He graduated from the University of Georgia in June 1982
with a Bachelor of Science in Agriculture degree, majoring in horticultural science.
Edwin entered the University of Florida in August 1982 in the Department
of Ornamental Horticulture and received his Master of Science degree from this
department in May 1985. He transferred to the Department of Fruit Crops to
pursue the Doctor of Philosophy degree. Edwin plans to receive his doctorate in
August 1991. He will spend a short time as a postdoctoral research associate in
the laboratory of Dr. Karen Koch at the University of Florida and will then move to
Peoria, Illinois where he will be a postdoctoral research associate for the United
States Department of Agriculture in the laboratory of Dr. Douglas Doehlert.
He is a member of Phi Theta Kappa, Phi Kappa Phi and Gamma Sigma
Delta honor societies and a member of the American Society for Horticultural
Science and the American Society of Plant Physiologists.
113

I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
Professor of Horticultural Science
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
Rebecca L. Darnell
Assistant Professor of Horticultural Science
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
Associate Professor of Horticultural
Science
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
C? 0 (cas
L. Curtis Hannah
Professor of Horticultural Science
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
Alice C. Harmon
Assistant Professor of Botany

This dissertation was submitted to the Graduate Faculty of the College of
Agriculture and to the Graduate School and was accepted as partial fulfillment of
the requirements for the degree of Doctor of Philosophy.
August, 1991
cuA
Dean, College of AgrioliKure
Dean, Graduate School



109
Salanoubat, M. and G. Belliard. 1989. The steady-state level of potato sucrose
synthase mRNA is dependent on wounding anaerobiosis and sucrose
concentration. Gene 84:181-185.
Salerno, G.L. 1986. Relationship between sucrose and starch in developing maize
endosperm UDP-glucose phosphorylase activity. Plant Sci. 44:111-118.
Salerno, G.L., S.S. Gamundi and H.G. Pontis. 1979. A procedure for the assay of
sucrose synthetase and sucrose phosphate synthetase in plant
homogenates. Anal. Biochem. 93:196-199.
Sampietro, A.R., M.A. Vattuone and F.E. Prado. 1980. A regulatory invertase from
sugar cane leaf-sheaths. Phytochem. 19:1637-1642.
Sasaki, T., K. Tadokoro and S. Suzuki. 1971. Multiple forms of invertase of potato
tuber stored at low temperature. Phytochem. 10:2047-2050.
Schaffer, A.A., B. Aloni and E. Fogelman. 1987. Sucrose metabolism and
accumulation in developing fruit of Cucumis. Phytochem. 26:1883-1887.
Schaffer, A.A., I. Rylski and M. Fogelman. 1989. Carbohydrate content and
sucrose metabolism in developing Solanum muricatum fruits. Phytochem.
28:737-739.
Schmalstig J.G. and W.D. Hitz. 1987. Transport and metabolism of a sucrose
analog (1 fluorosucrose) into Zea mays L. endosperm without invertase
hydrolysis. Plant Physiol. 85:902-905.
Schuster, J.R. 1989. Regulated transcriptional systems for the production of
proteins in yeast: Regulation by carbon source, pp. 83-108 in P.J. Barr, A.J.
Brike and P. Valenzuela, eds. Yeast Genetic Engineering. London:
Butterworths.
Servaites, J.C., B.R. Fondy, B. Li and D.R. Geiger. 1989. Sources of carbon for
export from spinach leaves throughout the day. Plant Physiol. 90:1168-1174.
Setia, N. and C.D. Malik. 1985. Changes in some enzymes of carbohydrate
metabolism in developing pod and seed of chick pea (Cicer arietinum)
Phyton 25:93-99.
Sexton R. and J.F. Sutcliffe. 1969. The distribution of jS-glycerophosphatase in
young roots of Pisum sativum. L. Ann. Bot. 33:407-419.


60
unpublished data). Maraa and co workers (Maraa et al., 1990) have found that
the two genes encoding sucrose synthase in wheat (Ss1 and Ss2) also show a
differential response to stress conditions (anaerobiosis, cold shock and light).
Webster and Henry (1987) reported an unknown protein with a molecular
weight similar to that of the subunits of sucrose synthase in pea root meristem
cells undergoing sugar starvation. Carbohydrate responsive proteins have also
been found in roots of pearl millet (Baysdorfer and VanDerWoude, 1988). These
proteins, however, are yet to be definitively identified. Initial findings by Koch and
co workers (Koch and McCarty, 1988, 1990; Koch et al., 1989) indicated that the
Sh 1 gene of maize was stimulated by low carbohydrate conditions and down-
regulated under carbohydrate sufficient conditions. The Sus1 gene responded in
an inverse manner. Maas and co-workers (Maas et al., 1990) demonstrated that
the promoter from the Sh 1 gene was repressed by high sucrose conditions.
However, Salanoubat and Belliard (1989) found that increased sucrose promoted
genes encoding sucrose synthase.
The two genes encoding sucrose synthase in maize respond to altered
carbohydrate status (Koch and McCarty, 1988,1990; Koch et al., 1989), and shifts
in sucrose synthase protein localization have been observed under the same
conditions (K. Nolte, University of Florida, unpublished data). However, these
studies were carried out using a maize line having both sucrose synthase genes
present. The present study utilizes the Shrunken-1 mutant of maize to determine


33
contained all assay components except UDP. The assay was optimized for pH,
linearity with time and protein concentration (data not shown).
Product Determination
The entire reaction volume was blotted onto a small disk of DEAE ion-
exchange paper (2.4 cm diameter) and dried completely before rinsing. Each disk
was rinsed separately, first in 40 ml of H20 at 175 rpm on a rotary shaker for 2
hours, again for an additional 2 hours and finally rinsed in a gentle stream of Dl
water for 30 sec. Remaining radiolabel was quantified and compared to total
amount of the 14C-sucrose substrate utilized to determine extent of sucrose
cleavage.
Protein Gel Blots
Subsamples from protein extracts to be used for enzyme assays were
separated on native PAGE using the system of Laemilli (1970) with (denaturing)
or without (native) SDS. Polyacrylamide concentrations of the stacking and
separating gels were 2.5% and 5%, respectively. Proteins were resolved at 4 C
by applying 15 V for 9 h, then 125 V for 11 h (constant current and temperature
(4 C). Each lane was loaded with 2 /g of total protein. Proteins were
electroblotted to nitrocellulose membranes and probed with polyclonal antibodies
following the procedure of Towbin et al. (1979). Sucrose synthase antisera,
obtained from D.R. McCarty, was generated in rabbits using protein purified from
maize kernels (W64 x 182E) 22 days after pollination. Antisera was diluted 1:1000


Sucrose cleaved (urnol mg protein h 1)
35
0 10 20 30 40 50
Time after extraction (min)
Figure 3-1. Time course of in vitro decrease in sucrose synthase activity in maize
and cotton roots. Bars represent SE, n=3.


108
Reimerdes, E.H. and H. Klostermeyer. 1976. Determination of proteolytic activities
on casein substrates, pp. 26-28 in L. Lorand, ed. Methods in Enzymology.
Vol. XLV. Proteolytic Enzymes. New York: Academic Press.
Ricard, B., J. Rivoal, A. Spiteri and A. Pradet. 1991. Anaerobic stress induces the
transcription and translation of sucrose synthase in rice. Plant Physiol.
95:669-674.
Ricardo, C.P.P. 1974. Alkaline /S-fructofuranosidases of tuberose roots: Possible
physiological function. Planta 118:333-343.
Ricardo, C.P.P. and T. ap Rees. 1970. Invertase activity during the development
of carrot roots. Phytochem. 9:239-247.
Robbins W.J. 1958. Sucrose and growth of excised roots of an inbred Zea mays.
Proc. Natl. Acad. Sci. (USA) 44:1210-1212.
Roberts, D.W.A. 1973. A survey of the multiple forms of invertase in the leaves of
winter wheat, Triticum aestivum L. Emend Thell ssp. vulgare. Biochem.
Biophys. Acta. 321:220-227.
Robinson E. and R. Brown. 1952. The development of the enzyme complement of
growing root cells. J. Expt. Bot. 3:356-374.
Rocha-Sosa, M., U. Sonnewald, W. Frommer, M. Stratmann, J. Schell and L.
Willmitzer. 1989. Both developmental and metabolic signals activate the
promoter of a class I patatin gene. The EMBO J. 8:23-29.
Rovira, A.D. and C.B. Davey. 1974. Biology of the Rhizosphere. pp. 153-204 in
E.W. Carson, ed. The Plant Root and its Environment. Charlottesville:
University Press of Virginia.
Rowland, L.J. Y-C. Chen and P.S. Chourey. 1989. Anaerobic treatment alters the
cell specific expression of Adh-1, Sh and Sus genes in roots of maize
seedlings. Mol. Gen. Genet. 218:33-40.
Rowland, L.J. and P.S. Chourey. 1990. In situ hybridization analysis of sucrose
synthase expression in developing kernels of maize. Maydica 35:373-382.
Saglio, P.H. and A. Pradet. 1980. Soluble sugars, respiration, and energy charge
during aging of excised maize root tips. Plant Physiol. 66:516-519.


LIST OF TABLES
Table 3-1 Effect of enzyme protectants on activity of sucrose synthase
from maize root tips assayed five minutes after extraction 36
Table 4-1 Total sucrose synthase activity in wildtype maize root tips
incubated in a range of glucose concentrations for 24 hour 53
Table 5-1 Sucrose synthase activity in mutant maize ^N22:sh1) root tips
incubated in media containing a range of glucose concentrations for
24 hours 66
Table 5-2 Sucrose synthase activity in mutant maize (W22:s/77) root tips
incubated in media containing 0 or 2.0% glucose for various time
periods 70
Table 6-1 Soluble and insoluble acid invertase activity in sequential 2 mm
segments of primary roots of 5- to 6-day-old seedlings from 1 hybrid
and 2 inbred lines of maize 81
Table 6-2 Soluble acid invertase and sucrose synthase activity in 0.5 cm
apices of primary and adventitious roots of 5- to 6-day-old seedlings
from 1 hybrid and 2 inbred lines of Zea mays 83
Table 6-3 Soluble acid invertase activity in various tissues of Oh 43, an
inbred line of Zea mays 84
v


71
the Sh1 maize mutant has allowed the response to be characterized using a
simple, single enzyme system with respect to changing sugar supply. The
increased protein and enzyme activity evident at increased exogenous glucose
levels indicate that the plant tissue can adjust this first step in their sucrose-
metabolizing capacity relative to its carbohydrate status.
After 24 h at a given glucose level, changes in gene expression were more
marked than were differences at the protein and enzyme levels (Figure 5-1, Figure
5-2 and Table 5-1). This is not surprising given the probable presence of
previously formed RNA and protein (both appear to be relatively long-lived) as well
as the comparatively long-term progression of the response to the maximal extent
observed at 48 h (Figure 5-3 and Table 5-2). Chourey et al. (1991 b) reported that
the sucrose synthase gene in sorghum homologous to Sus 1 gene from maize is
anaerobically induced, but levels of the respective protein do not change.
Anerobic induction, however, was terminated after only 12 h. Anaerobic induction
of Sh1 in maize becomes apparent between 6 and 12 hours but message levels
are not maximal until at least 24 h (Duke and Koch, unpublished data). Data from
the present work indicate that like the respiratory drop noted by Brouquisse et al.
(1991), at least 20 hours are required before a change in Sus1 is fully apparent at
the gene level and even longer at the protein level. Nonetheless, data are
presented here at the levels of mRNA, protein and enzyme activity that indicate
that expression of the Sus 1 gene for sucrose synthase is responsive to
carbohydrate availability to an extent not evident in background levels of total RNA


47
activity (Bennett, 1984). Sucrose phosphate synthase (SPS), an enzyme of
carbohydrate metabolism, has been shown to be regulated in this manner
(Doehlert and Huber, 1983; Walker and Huber, 1989; Huber et al., 1989a).
Increases in extractable SPS activity are noted after illumination or inclusion of
mannose or glucosamine (phosphate sequestering agents) in darkness (Huber et
al., 1989b). Inorganic phosphate (5-10 mM) was found to be a potent inhibitor of
SPS (Amir and Preiss, 1982), with the inhibition becoming more sensitive in the
presence of Mg2+. Sucrose synthase has also been shown to have exhibit diurnal
fluctuations in activity (as does SPS) (Hendrix and Huber, 1986; Vassey, 1989) and
be affected by P¡. Pontis (1977) reported that P¡ (2-5 mM) inhibited the
degradative reaction of sucrose synthase alone or in the presence of Mg2+.
Delmer (1972a), however, found that 2 mM P¡ had no effect on the rates of either
the forward or reverse reactions. Sucrose synthase extracted from maize roots
has also been shown to be phosphorylated in vitro (Xu and Koch, University of FL,
unpublished data). In the current study, no effect of added phosphate on protein
stability was noted (Figure 3-4D). However, initial sucrose synthase activity
measurements were less than controls, and a loss of activity with time was
observed (Figure 3-3A & B).
The possibility existed that disassociation of subunits from tetramers may
have effected activity. Two separate isozymes in maize (Sh1 and Sus) form
subunits which appear to combine randomly into tetramers in root tips and other
tissues (Chourey et al., 1986). Endosperm sucrose synthase tetramers, however,


20
Previously, the role of invertase in sucrose transfer was considered
particularly important in plants such as sugar cane and maize where substantial
sugar movement occurred through the cell wall space and was accompanied by
action of an extracellular invertase (Glasziou and Gayler, 1972; Hawker and Hatch,
1965). Recent evidence, however, indicates that although much hydrolysis is often
observed, invertase activity may not be essential for sucrose uptake into either
sugar cane stems (Thom and Maretzki, 1990; Lingle, 1989) or maize kernels
(Schmalstig and Hitz, 1987). The role of apoplastic invertase in sucrose import
into roots had previously been questioned by Chapleo and Hall (1989a) who
concluded that although present, apoplastic root invertase did not have a direct
role in sugar transport. However, substantial activity of invertase has been widely
documented in roots of plants such as pea (Lyne and ap Rees, 1971), bean
(Robinson and Brown, 1952), tomato (Chin and Weston, 1973), Ricinus (Chapleo
and Hall, 1989a, b, and c), oat (Pressy and Avants, 1980), and maize (Hellebust
and Forward, 1962; Chang and Bandurski, 1963). Specific tissue localizations also
have been described. Peak activity for root invertase is generally 2-3 mm behind
the apex and corresponds to the region of expansion and elongation in pea
(Robinson and Brown, 1952; Sexton and Sutcliffe, 1969) and maize (Hellebust and
Forward, 1962). In Ricinus roots, this activity predominates in the cortex (Chapleo
and Hall 1989a).
Although invertase may not have a direct role in sucrose import into roots,
it still may be important to two major aspects of root biology. First, invertase is


CHAPTER 6
AN ORGAN-SPECIFIC INVERTASE DEFICIENCY IN THE PRIMARY ROOT OF AN
INBRED MAIZE LINE
Abstract
An organ-specific invertase deficiency affecting only the primary root system
is described in the Oh 43 maize inbred. Invertases (acid and neutral/soluble and
insoluble) were assayed in various tissues of hybrid (NK 508) and inbred (Oh 43,
W22) maize lines to determine the basis for an early report that Oh 43 root tips
were unable to grow on sucrose agar (Robbins, 1958). Substantial acid invertase
activity (7.3 to 16.1 ^mol glucose mg'1protein h'1) was evident in extracts of all
tissues tested except the primary root system of Oh 43. This deficiency was also
evident in lateral roots arising from the primary root. In contrast, morphologically
identical lateral roots from the adventitous root system had normal invertase levels.
These results suggest that ontogenetic origin of root tissues is an important
determinant of invertase expression in maize. Adventitious roots (including the
seminis) arise above the scutellar node and are, therefore, of shoot origin. The
Oh 43 deficiency also demonstrated that invertase activity was not essential for
maize root growth. Sucrose synthase was active in extracts from all root apices
and theoretically provided the only available avenue for sucrose degradation in
75


78
probable advantages of its presence and indicates an unusual organ-specific
difference in expression between primary and adventitious roots.
Materials and Methods
Plant Material
Maize seed (Zea mays L. NK 508, W22 and Oh 43) were germinated on
moist filter paper in petri dishes. Seeds were imbibed for 24 hours and pericarps
removed, allowing more uniform germination and more effective surface
sterilization (20 min soak in 0.525% sodium hypochlorite).
Five successive 2 mm segments were sampled from the tips of primary
roots 4 to 5 days after germination. Intact roots of Oh 43 seedlings grew more
slowly than did those of NK 508 or W22, but all roots had reached 2 cm prior to
excision. Tissue samples were weighed, frozen in liquid N2 and stored at -80 C
until assayed for invertase activity. In subsequent experiments, 5 mm root tips
were excised from primary and adventitious roots for invertase and sucrose
synthase activity measurements. Plants and tissues were as above.
Tissue Extraction
Frozen tissue samples were ground to a fine powder in liquid N2 using a
mortar and pestle. Frozen powder was transferred to a second mortar containing
ice-cold 200 mM HEPES buffer (pH 7.5) with 1 mM DTT, 5 mM MgCI, 1 mM EGTA,
20 mM sodium ascorbate and 10% (w/w) PVPP. One ml of extraction buffer was
used for every 100 mg of tissue fresh weight. Buffered extract was centrifuged at


76
primary root tips of Oh 43. The deficiency described here will provide a useful
avenue of investigation into the expression and significance of root invertase.
Introduction
Sucrose breakdown is critical to the vast majority of plant species because
non-photosynthetic tissues depend on import of this sugar for their growth and
development. Initial cleavage of sucrose can be catalyzed by either invertase or
the reversible enzyme sucrose synthase. Invertases are especially active in tissues
undergoing rapid cell division such as shoot and root apices (Avigad, 1982).
Previously, the role of invertases in sucrose transfer was considered particularly
important in plants such as sugar cane and maize where substantial sugar
movement occurred through the cell wall space and was accompanied by action
of an extracellular invertase (Glasziou and Gayler, 1972; Hawker and Hatch, 1965).
Recent evidence, however, indicates that although much hydrolysis is often
observed, invertase activity may not be essential for sucrose uptake into either
sugar cane stems (Lingle, 1989; Thom and Maretzki, 1990) or maize kernels
(Schmalstig and Hitz, 1987).
Sucrose generally is believed to enter root tips without traversing the
extracellular space (Giaquinta et al., 1983); however growing roots can differ
markedly in their capacity to lose (Rovira and Davey, 1974) and retrieve (Robbins,
1958) exogenous sugars. Net losses do occur. The extent of sugar efflux from
roots can be affected by irradiance level, nutritional status, moisture availability and


BIOGRAPHICAL SKETCH
Edwin Ralph Duke was born in Bainbridge, Georgia on February 22,1960.
He attended public school in Rossville, Georgia, and graduated from Macon
Christian Academy, Macon, Georgia, in May 1978. In June 1980 he received his
Associate of Science degree from Macon Junior College, Macon, Georgia,
majoring in agriculture. He graduated from the University of Georgia in June 1982
with a Bachelor of Science in Agriculture degree, majoring in horticultural science.
Edwin entered the University of Florida in August 1982 in the Department
of Ornamental Horticulture and received his Master of Science degree from this
department in May 1985. He transferred to the Department of Fruit Crops to
pursue the Doctor of Philosophy degree. Edwin plans to receive his doctorate in
August 1991. He will spend a short time as a postdoctoral research associate in
the laboratory of Dr. Karen Koch at the University of Florida and will then move to
Peoria, Illinois where he will be a postdoctoral research associate for the United
States Department of Agriculture in the laboratory of Dr. Douglas Doehlert.
He is a member of Phi Theta Kappa, Phi Kappa Phi and Gamma Sigma
Delta honor societies and a member of the American Society for Horticultural
Science and the American Society of Plant Physiologists.
113


74
1983). Increased levels of sucrose result in elevated levels of genes for both
sucrose synthase (Salanoubat and Belliard, 1989) and patatin (Rocha-Sosa et al.,
1989; Wenzler et al., 1989) in tissues where they are not usually found. A rise in
sugar availability can also result in increased transcription and translation of a
unique storage protein in stem and leaf tissues of sweet potato (Hattori et al.,
1990). The diversity of processes operating in the system utilized in this study
suggests that sugar-responsive gene expression (ie. Sus 1) may have broad
implications in the formation and function of non-storage plant tissues.


31
Materials and Methods
Plant Material
Maize seed (Zea mays L, NK 508, VJ22:sh1) were primed for 6 days at 10
C with a water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I*1 captan
(Bodsworth and Bewley, 1981). At the end of 6 days, the seeds were rinsed free
of PEG, given a 20 min rinse in 1.05% (v/v) sodium hypochlorite and again rinsed
in water for 20-30 min. Seeds were germinated in the dark at 18 C on Whatman
3mm filter paper. Moisture level was kept constant throughout. At the end of 7
days, 1 cm primary root tips were excised under a sterile transfer hood. Cotton
(Gossypium hirsutum L, Coker 100) root tips were obtained from Dr. D.L. Hendrix
(Western Cotton Research Laboratory, USDA/ARS Phoenix, AZ). One cm root tips
were excised from 5- to 6-day old seedlings and quick frozen in liquid N2.
Purification of 14C-Sucrose
Trace amounts of phosphorylated sugars are common impurities in
commercial 14C-sucrose, and can reduce accuracy of the assay. These were
removed by descending paper chromatography of commercially obtained de
suerse in ethanol (NEN, Boston MA) using DEAE cellulose paper. The majority
of anion-free sucrose was concentrated into the first 2 to 3 drops eluted from the
V-shaped tip of the DEAE paper strip. No impurities were detected using HPLC
analysis (data not shown). Molarity and specific activity of purified 14C-Sucrose


I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
Professor of Horticultural Science
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
Rebecca L. Darnell
Assistant Professor of Horticultural Science
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
Associate Professor of Horticultural
Science
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
C? 0 (cas
L. Curtis Hannah
Professor of Horticultural Science
I certify that I have read this study and that in my opinion it conforms to
acceptable standards of scholarly presentation and is fully adequate, in scope and
quality, as a dissertation for the degree of Doctor of Philosophy.
Alice C. Harmon
Assistant Professor of Botany


88
plants. One isozyme predominated in shoot-derived organs (leaves, sheaths,
nodes and pollen) and the other isozyme showed highest activity in the roots. It
is interesting in this respect that the adventitious roots of maize arise above the
scutellar node of the developing seedling and are, therefore, derived from shoot
tissue (Hayward, 1938). The difference in invertase expression in primary and
adventitious root systems may reflect a similar root/shoot dichotomy. Therefore,
the tissue of origin and cell lineage may be more important than organ identity and
function in regulating invertase expression. Other variants in invertase expression
have been described. Echeverra and Humphreys (1984) reported a hybrid maize
line (DKXL80) which exhibited no soluble invertase activity in contrast to previously
tested lines. Other tissues of this maize line exhibited normal invertase activity.
Second, data demonstrate that invertase is not essential for primary root
growth. The primary roots of Oh 43 exhibited no signs of premature senescence,
and, if left intact, continued apparently normal growth for many days (data not
shown). Invertases role in sucrose import into roots has been questioned by
Chapleo and Hall (1989a) who concluded that although present, apoplastic root
invertase did not have a direct role in sugar transport in Ricinus. However,
substantial activity of invertase has been widely documented in roots of plants
such as pea (Lyne and ap Rees, 1971), bean (Robinson and Brown, 1952), tomato
(Chin and Weston, 1973), Ricinus (Chapleo and Hall, 1989a, b and c), oat (Pressy
and Avants, 1980), and maize (Chang and Bandurski, 1963; Hellebust and
Forward, 1962). In maize, the Oh 43 invertase deficiency apparently prevents


30
Introduction
Measurement of the maximum catalytic activities of enzymes in plant tissues
can make important contributions to the understanding of metabolic pathways and
their mechanisms of control (ap Rees, 1974). Currently available methods of
assaying sucrose synthase have proven ineffective for many tissues, particularly
those of roots (Duke et al., unpublished data; Lingle, USDA/ARS, Westlaco, TX,
personal communication). A precipitous loss of activity follows tissue extraction
from root tips of maize and other species (D.L. Hendrix, USDA/ARS, Phoenix, AZ,
personal communication). Chan et al. (1990) reported that sucrose synthase
activity in roots of rice was detectable in only one stage of growth. However,
sucrose synthase protein was present in root tissue at all stages of growth,
exceeding that in grain when grain activity was highest among tissues sampled.
This report addresses the basis of this instability in maize roots and describes a
rapid radiometric assay for sucrose synthase which circumvents this problem and
allows assay of small samples. Extraction and assay were optimized for substrate
concentration, pH, assay length, and inclusion (or exclusion) of various anti
oxidants and proteinase inhibitors. The procedure has proven effective for a range
of tissues and species examined and provides an accurate measurement of
activity, particularly where enzyme stability may be limiting.


87


11
1989) as well as the energy for phloem loading. However, once sucrose is loaded,
its eventual fate does not appear to be under the control of the source leaf (Gifford
and Evans, 1981) but rather is under the control of the importing sink (Wyse,
1986). Giaquinta (1979) found that young, immature roots of sugar beets had low
levels of sucrose synthase, but the onset of rapid sucrose import for storage was
accompanied by a significant increase in sucrose synthase activity. Similar
correlations were also observed by Silvius and Snyder (1979) and Fieuw and
Willenbrink (1987). In sugar beet roots sucrose uptake into parenchyma can
proceed without prior hydrolysis in the apoplast or free space. Increases in
sucrose synthase activity have also been observed during periods of sucrose
import and/or accumulation in sweet melons (Schaffer et al., 1987), netted
muskmelon (Lingle and Dunlap, 1987), eggplants (Claussen et al., 1985, 1986),
rose flowers (Khayat and Zeslin, 1987), developing chick pea seeds (Setia and
Malik, 1985), tomato (Yelle et al., 1988) and Solanum muricatum (Schaffer et al.,
1989). Lingle (1987), however, found no correlation of sucrose synthase activity
with sucrose concentration in sweet sorghum.
Huber and Akazawa (1986) hypothesized that a primary role of sucrose
synthase could be to feed glucose-1 -phosphate directly into glycolysis. Black and
coworkers (Black et al., 1987; Sung et al., 1988; 1989; Xu et al., 1989) also support
this hypothesis. This link to glycolysis also involves UDPG-pyrophosphorylase,
which converts the UDPG formed by the action of sucrose synthase into G-1-P
and UTP. The methods employed to deliver carbohydrates to their respective


68
switched to 2.0% glucose responded similarly to those remaining in 2.0% glucose
for the entire time.
At the protein level,changes in response to altered carbohydrate availability
were not apparent at 16 h, remained barely detectable at 24 h, but were clearly
evident after 48 h (Figure 5-4). Treatment reversals indicated that a protein-level
response occurred only when 16 h of elevated glucose treatment was followed by
32 h of glucose deprivation. The response was similar to that of root tips that had
remained continuously in 2.0% glucose. Slight differences in enzyme activity
between treatments were evident after 16 h or 24 h (Table 5-2); however activity
in glucose supplemented tips had risen to levels two-fold greater than those
without exogenous sugars within 48 h.
Discussion
The significance of results described here are two-fold. First, data
demonstrate that the differential response to changing carbohydrate availability by
the Sus1 gene for sucrose synthase is apparent at the translational level as well
as at the transcriptional level. The two genes encoding sucrose synthase
previously have been shown to respond differentially to carbohydrate supply (Koch
and McCarty, 1988; 1990; Koch et al., 1989). Differences at the protein and
enzyme level, however, have been difficult to detect due to cross reactivity of
polyclonal antibodies and the collective contribution of both isozymes to activity
measurements. Second, the resulting changes in physiology may allow the cells
to adjust their carbohydrate metabolizing capacity to the available supply. Use of


52
Results
Sucrose synthase activity was consistently maximal in root tips
supplemented with 0.5% glucose (Table 4-1), a level at which the combined levels
of mRNA from the two sucrose synthase genes was also greatest (Koch et al.,
unpublished data). Overall, however, activity of sucrose synthase in whole root
tips was not significantly changed by alteration of carbohydrate status by
exogenous sugar supply (Table 4-1). It was not possible to distinguish activities
of isozymes encoded by the two sucrose synthase genes. These genes, Sh1 and
Sus 1, were found to exhibit reciprocal responses at the mRNA level to sugar
availability in the same sets of roots used for these experiments (Koch et al.,
unpublished data). Also, changes in distribution of sucrose synthase protein
among tissues within these root tips (Nolte, unpublished data) were not reflected
at the level of whole root enzyme activity.
Discussion
Reciprocal regulation of the two isoforms by carbohydrate levels, as has
been demonstrated for genes encoding for these isozymes (Koch and McCarty,
1989), could explain the lack of significant differences detected between glucose
treatments. The two isozymes of sucrose synthase from maize (encoded by the
Sh1 and Sus1 genes) are very similar, differing only slightly in their electrophoretic
movement during PAGE (Echt and Chourey, 1985). The Sh1 and Sus1 encoded
proteins are capable of catalyzing the same reaction with little difference in affinities


34
and cross reacted strongly to both the Sh1 and Sus 1 gene products where such
were present.
Results
Activity of sucrose synthase from maize and cotton root tips declined rapidly
after extraction (Figure 3-1). The greatest decrease in activity occurred between
10 and 15 minutes after extraction from both species. Little or no activity was
observed after 4 hours (data not shown). After extraction, extracts were
maintained at 0 C until used in the radiometric assay.
A wide range of enzyme protectants were examined. No improvement in
activity was observed when the polyphenol protectants PVP-40, PEG 20,000 and
BSA were utilized (Table 3-1). PVPP, also a phenol absorbent, was utilized in each
extraction. In addition, four classes of proteinase inhibitors were tested for their
effect on stability of sucrose synthase activity. Addition of leupeptin (1 mM) slightly
decreased initial activity (Table 3-1), and pepstatin-A (1 mM) had no effect (Table
3-1). Phenylmethylsulfonyl fluoride (PMSF) (1 mM), a serine proteinase inhibitor,
had a substantial positive effect, as did casein (2% w:v), potentially a non-specific
proteinase inhibitor.
Further characterization of activity change in the presence of PMSF showed
that stabilization was not effective in the first 20 min following extraction (Fig. 3-2).
Although total activity prior to this time was elevated by addition of PMSF, a linear
decrease was not prevented from occurring.


Figure 6-1. Histochemical localization of ¡nvertase activity in free-hand, fresh cross
sections of root apices of 6-day-old maize seedlings. Sections are
approximately 50 /xm in thickness. Primary (A) and adventitious (B) root
cross sections from NK 508 incubated in reaction medium without sucrose
(controls). Primary (C) and adventitious (D) root cross sections from Oh 43
incubated in reaction medium (note only adventitious root section exhibits
blue formazan reaction product [dark areas]). Primary (E) and adventitious
(F) root cross sections from NK 508 incubated in reaction medium (note
both sections exhibit reaction product [dark areas]). Bar represents 50 /xm.


CHAPTER 7
SUMMARY AND CONCLUSIONS
Sucrose metabolism is important to the majority of plant species because
of the widespread role of this sugar in growth and development through its
function as a phloem transport sugar. Initial breakdown of sucrose can be
catalyzed by either sucrose synthase, a reversible enzyme, or invertase.
Expression of genes encoding the sucrose synthase isozymes (Sh1 and
Sus 1) have been found to be sensitive to carbohydrate levels (Koch and McCarty,
1988, Koch et al., 1989). Northern blot analysis of mRNA showed levels showed
those of Sh1 were up-regulated in response to carbohydrate depletion whereas
those of Sus 1 were down-regulated under the same conditions. This may prove
to be an important control mechanism whereby plant cells are able to react to
cellular carbohydrate status.
The first purpose of this work was to determine the extent to which
carbohydrate-modulated changes in message levels affected activity of sucrose
synthase the level of enzyme activity. However, previously available sucrose
synthase assay methods proved ineffective for maize root tips due to a precipitous
loss of activity after tissue extraction. A rapid, radiometric assay for sucrose
synthase was therefore developed that overcomes these obstacles. Extraction and
assay were optimized for factors such as substrate concentration, pH, assay
91


85
Histochemical staining indicated that no invertase activity was detectable in
the primary roots of Oh 43 (Figure 6-1). Cross sections of primary and
adventitious roots of Nk 508 and adventitious roots of Oh 43 all stained positive
for invertase activity. Invertase activity was primarily in the cortex and was
localized intercellularly.
Discussion
These data confirm that Oh 43 (an inbred line of maize) lacks invertase
activity in its primary root tips. Surprisingly, however, a deficiency was not evident
in structurally and functionally similar adventitious roots (Table 6-2), other tissues
of the same Oh 43 plants (Table 6-3) or in the developing kernels of this line
(Doehlert, et al., 1988). The genetic potential for invertase expression is therefore
present. The lack of activity may result from altered regulation of gene expression
(transcription or translation), the loss of a tissue specific invertase isozyme or the
existence of an unidentified effector of enzyme function. The significance of results
described here is twofold. First, evidence is presented for differential expression
of invertase in morphologically identical organs that differ primarily in point of
origin. This is most strikingly illustrated in the apparent distinction between lateral
roots arising from primary and adventitious root systems. Differential expression
of genes in morphologically identical structures are unusual; however, organ or
tissue-specific differences have been well documented (Fluhr et al., 1986; Xie and
Wu, 1989). Xie and Wu (1989), for example, found that genes for alcohol
dehydrogenase were differentially expressed in root and shoot tissues of rice


80
the same reaction medium adjusted to pH 7.5 with potassium phosphate buffer.
Initial assays were also performed at pH ranges of 4.0 to 5.5 for acid invertase and
7.0 to 8.0 for neutral invertase. After a 15 min incubation at 30 C, glucose
production was quantified by the glucose oxidase method (Sigma Chemical Co.).
Sucrose synthase was assayed in the degradative direction using a radiometric
assay quantifying the production of 14C-UDPG (Chapter 3).
Histochemical Staining
Free-hand cross sections from apices of both primary and adventitious
roots were fixed in 4% formalin (pH 7.0) for 30 min and rinsed in water at least 10
times over a period of 3 hours to remove endogenous sugars (Doehlert and
Felker, 1987). Sections were then incubated in a sodium phosphate buffer (0.38
M, pH 6.0) containing 0.24 mg ml'1 nitroblue tetrazolium, 0.14 mg ml'1 phenazine
methosulfate, 25 units ml'1 glucose oxidase and 5 mg ml'1 sucrose (Doehlert and
Felker, 1987). Control sections were incubated in the same mixture without
sucrose. After rinsing in water, sections were post fixed in 4% formalin (pH 7.0)
and photographed under a microscope.
Results
Primary roots of Oh 43 showed little or no acid invertase activity (Table 6-1).
In contrast, acid invertase was active in extracts from apical areas of roots from
other maize lines examined (NK 508 and W22). Activity, per unit fresh weight, was
greatest in root apices, decreasing with distance from the tip until no longer


98
de Fekete, MAR. and C.E. Cardini. 1964. Mechanism of glucose transfer from
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del Rosario, E.J. and V. Santisopasri. 1977. Characterization and inhibition of
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Delmer, D.P. 1972a. The purification and properties of sucrose synthetase from
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Dick, P.S. and T. ap Rees. 1975. The pathway of sugar transport in roots of Pisum
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Dickinson, C.D., T. Altabella and M.J. Chrispeels. 1991. Slow-growth phenotype of
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Masuda, H. and S. Sugawara. 1980. Purification and some properties of cell wall
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Masuda, H., T. Takahashi and S. Sugarwara. 1988. Acid and alkaline invertases in
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Matsushita, K. and I. Uritani. 1976. Isolation and characterization of acid invertase
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Matsushita, K. and I. Uritani. 1977. Synthesis and apparent turnover of acid
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McCarty, D.R. 1986. A rapid and simple method for extracting RNA from maize
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McCollum, T.G., D.J. Huber and D.J. Cantliffe. 1988. Soluble sugar accumulation
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McElfresh, K.C. and P.S. Chourey. 1988. Anaerobiosis induces transcription but
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Miron, D. and A.A. Schaffer. 1991. Sucrose phosphate synthase, sucrose synthase
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Plant Physiol. 95:623-627.


Results 52
Discussion 52
5 SUGAR RESPONSE OF SUCROSE SYNTHASE AT
THE GENE {Sus1), PROTEIN AND ENZYME ACTIVITY
LEVELS IN ROOTS OF THE SM MAIZE MUTANT 57
Abstract 57
Introduction 58
Materials and Methods 61
Results 63
Discussion 68
6 AN ORGAN-SPECIFIC INVERTASE DEFICIENCY IN
THE PRIMARY ROOT OF AN INBRED MAIZE LINE 75
Abstract 75
Introduction 76
Materials and Methods 78
Results 80
Discussion 85
7 SUMMARY AND CONCLUSIONS 91
LITERATURE CITED 94
BIOGRAPHICAL SKETCH 113
iv


73
marked growth during the 24 hours of incubation (data not shown). During this
time, there is a demand for cell wall synthesis by the expanding cells. Sucrose
synthase has been implicated in the directing of carbohydrates for polysaccharide
biosynthesis (Amino et al.,1985; Hendrix, 1990). The level of involvement for
nucleotide-sugars during cell wall polysaccharide biosynthesis has been implicated
in a need for greater activity of this enzyme (Maas et al., 1990). Sucrose synthase
activity was elevated in cell cultures of Catharanthus roseus during the G1 phase
when total amounts of cell wall biosynthesis increased significantly (Amino et al.,
1985). Sugar modulation of Sus1 could convincingly combine production of cell
wall precursors with other aspects of increased growth (Farrar and Williams, 1990)
likely to accompany an enhanced sugar supply.
The carbohydrate response of sucrose synthase in the present study differs
from previously demonstrated regulation in that it occurs in rapidly growing and
metabolizing structures. Other studies have involved sucrose synthase regulation
in storage tissues where processes such as starch accumulation predominate.
Loss of the shrunken-1 gene in maize results in a typical endosperm phenotype
where starch deposition is reduced by over 70% (Chourey and Nelson, 1976);
however, a mutant lacking a functional Sus 1 gene has no apparent phenotype
(Chourey et al., 1988). Starch deposition accompanies protein accumulation in
developing potato tubers and levels of mRNA encoding sucrose synthase have
been shown to increase in this tissue (Salanoubat and Belliard, 1989). Levels of
storage proteins, such as patatin, also accumulate during this time (Paiva et al.,


LITERATURE CITED
Akazawa, T. and K. Okamoto. 1980. Biosynthesis and Metabolism of Sucrose, pp.
199-220 in The Biochemistry of Plants. New York: Academic Press.
Al-Bakir, A.Y. and J.R. Whitaker. 1978. Purification and properties of invertase from
dates (Phoenix dactylifera L. var. Zahdi) J. Food Biochem. 2:32-160.
Amino, S., Y. Takeuchi and A. Komamine. 1985. Changes in enzyme activities
involved in formation and interconversion of UDP-sugars during the cell
cycle in a synchronous culture of Catharanthus roseus. Physiol. Plant.
64:111-117.
Amir, J. and J. Preiss. 1982. Kinetic characterization of spinach leaf sucrose-
phosphate synthase. Plant Physiol. 69:1027-1030.
Anderson, R.S. and E.E. Ewing. 1978. Partial purification of potato tuber invertase
and its proteinaceous inhibitor. Phytochem. 17:1077-1081.
ap Rees, T. 1974. Sucrose metabolism, pp. 53-73 in D.H. Lewis, ed. Storage
Carbohydrates in Vascular Plants. Cambridge, U.K.: Cambridge University
Press.
Arnold, W.N. 1966. A column method for enzymic characterization of coarse
cellular fractions: application to insoluble /3-fructofuranosidase from grape.
Arch. Biochem. Biophys. 113:451-456.
Avigad, G. 1964. Sucrose-uridine diphosphate glucosyl-transferase from Jerusalem
artichoke tubers. J. Biol. Chem. 239:3613-3618.
Avigad, G. 1982. Sucrose and other disaccharides, pp. 217-347 in F.A. Loewus
and W. Tanner, eds. Encyclopedia of Plant Physiology, New Series. Vol.
13A. New York: Springer-Verlag.
Avigad, G., N. Levin and Y. Milner. 1964. Sucrose metabolism in plant storage
tissues. 6th Inti. Congr. Biolchem. Abst. p. 502.
94


19
Roles of Invertase
Elevated acid invertase activity is characteristic of plant tissues in which
there is a need for hexoses produced from stored or recently transported sucrose
(ap Rees, 1974; Avigad, 1982). Greater activities of invertase also correlate well
with a low content of stored sucrose. In sugar beets, the onset of sucrose storage
is accompanied by a decrease in invertase activity (Silvius and Snyder, 1979;
Giaquinta, 1979). The same is true for carrot roots (Ricardo and ap Rees, 1970),
melon (Hubbard et alM 1989; Lingle and Dunlap, 1987; Schaffer et al., 1987;
McCollum et al., 1988), citrus (Kato and Kubota, 1978; Lowell, 1986) and
Lycopersicon hirsutum (Miron and Schaffer, 1991). In these systems, invertase
was very active prior to sucrose accumulation and dropped significantly upon
maturity. Invertase activity is usually greatest in tissues that are at a rapid stage
of growth and development (Weil and Rausch, 1990), particularly at the cell
division stage (Masuda et al., 1988). Root apices, young leaves and stem
internodes fall into this category. Mature leaves, functioning as sources of
photosynthates, generally have low levels of apoplastic acid invertase (Dickinson
et al., 1991). Transgenic tomato plants expressing yeast invertase in the apoplast
of mature leaves had a striking repression of growth (Dickinson et al., 1991). The
higher the level of invertase, the greater the inhibition. The general role of acid
invertase, therefore, seems to be for the breakdown of sucrose where there is a
marked need for hexose (ap Rees, 1974).


39
O 10 20 30 40 50 60 70
Time after extraction (min)
Figure 3-3. Time course of in vitro decrease in maize root sucrose synthase
activity in the presence and absence of either P. (10 mM) or casein (2%
w:v).


3
1. Develop a method of accurately assaying sucrose synthase activity
to circumvent the rapid decline in activity exhibited upon extraction
from maize root tips.
2. Clarify the possible causes of the rapid decline of sucrose synthase
activity in extracts from maize root tips.
3. Determine the effects of varying carbohydrate conditions on total
sucrose synthase activity in extracts from wildtype maize root tips.
4. Ascertain the effects of varying carbohydrate conditions on the Sus 1
gene and its sucrose synthase gene product free from the
confounding effects of Sh 1.
Characterize the extent of invertase activity in various tissues of the
Oh 43 inbred of maize, a putative invertase-deficient mutant.
5.


48
are almost entirely composed of subunits encoded by the Sh1 gene (Chourey and
Nelson, 1976; Chourey, 1981; Chourey et al., 1986) and are remain active during
extraction (Echt and Chourey, 1985). Five different types result in those tissues
which exhibit polymerization of both protomers; two are homotetramers and three
are heterotetramers. Heterotetramers could theoretically be more unstable than
homotetramers since, although very similar, the two subunits are not identical (Echt
and Chourey, 1985). Data (Figure 3-5A & B) indicate that greater instability of
heterotetramers relative to homotetramers of sucrose synthase was not the cause
of the observed activity loss. The profile of declining activity with time is similar in
extracts from root tips of a mutant line having only homotetramers of Sh1 (SS1)
subunits (y\/22:sh1) as it is in extracts from wild type kernels with 5 native
tetrameric combinations (NK 508).
Rapid loss of sucrose synthase activity with time in maize and other root
tips, as well as small tissue size, necessitated the development of a rapid and
sensitive assay. Other assays for sucrose synthase using radiometric techniques
have been described (Delmer, 1972a and b; Su and Preiss, 1978; Salerno et al.,
1979). However, the procedure described here has proven effective and useful
due to reduced time from extraction to assay termination, reduced sample size
and increased enzyme stabilization.


5
Morell and Copeland (1984, 1985) have investigated the enzymes of
sucrose breakdown in soybean nodules and found that both sucrose synthase
and alkaline invertase are present. They suggested that sucrose partitioning
between the two enzymes could be determined by differences in their affinities for
this substrate. The Km of alkaline invertase for sucrose was 10 mM whereas that
of sucrose synthase was 31 mM. Given the presence of both enzymes, they
proposed the greater affinity of alkaline invertase for sucrose in this system could
ensure that most of the sucrose entering the nodule would be converted to
hexoses for further catabolism. At the same time some sucrose would be
converted to UDP-glucose for subsequent synthesis of nucleotide sugars and
polysaccharides. In contrast, Huber and Akazawa (1986) reported essentially an
opposite situation in cultured sycamore cells, another system in which both
sucrose synthase and neutral (alkaline) invertase were present at the same time
and with similar activities. In these cells the sucrose synthase for sucrose was
substantially lower than that of neutral invertase (15 vs 65 mM). They proposed
two pathways of sucrose cleavage, initiated by each of the enzymes, both
pathways eventually leading to the production of triose-phosphates. Sucrose
concentration was postulated to regulate carbon flow between the two pathways.
Sucrose synthase had a lower Km for sucrose and would, therefore, be relatively
more important under sucrose limiting conditions (Avigad, 1982). This pathway is
more energy efficient and would be more beneficial to the cells when carbon
supplies are limited. The work of Morell and Copelands work (1984, 1985) and


62
Enzyme Assay
Sucrose synthase activity was determined by a rapid radiometric procedure
developed to circumvent enzyme instability previously observed upon extraction
from maize root tips (Chapter 3).
RNA Extraction and Northern Blotting
Samples were ground to a fine powder in a mortar and pestle with liquid N2
and RNA extracted according to McCarty (1986). Total RNA was quantified by
absorbance at 260 nm.
Total RNA was separated by electrophoresis in 1% agarose gels containing
formaldehyde (Thomas, 1980), blotted to a nylon membrane (Hybond-N,
Amersham Corporation, Arlington Heights, IL) and probed as per Church and
Gilbert (1984) with genomic clones of Sus1 (McCarty et al., 1986) radiolabeled by
random primer. Blots were rinsed and placed on X-ray film at -80 C.
Protein Gel Blots
Subsamples from protein extracts to be used for enzyme assays and
separated on native PAGE using the system of Laemilli (1970) without SDS.
Polyacrylamide concentrations of the stacking and separating gels were 2.5% and
5%, respectively. Proteins were resolved at 4 C by applying 15 V for 9 h, then 125
V for 11 h (constant current) and temperature (4 C).
Proteins were electroblotted to nitrocellulose membranes and probed with
polyclonal antibodies following the procedure of Towbin et al. (1979). Sucrose