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 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 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 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 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 lea mays 83
Table 6-3 Soluble acid invertase activity in various tissues of Oh 43, an inbred line of lea 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/?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 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 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 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, 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 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: 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. 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 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 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-D-
glucosyltransferase) 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, 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-6-
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, 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 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(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., 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 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 UDP- glucose 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 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 Km 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 K^s for UDPG when compared to other NDPG's have also been
et al. sweet observed for sucrose synthase from pea seedlings (Gabrielyan p 1969), 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 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 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 MgCI2 . In contrast, Pontis (1977) reported that P, inhibited the degradative reaction alone or
2+ in the presence of Mg . 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
2+ 2+ 2+ 2+ Mg , at 5-10 , Mn Ca and Ba 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
2+ inhibition could be reversed by the addition of Mg (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 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 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 + 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 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). 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 .
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 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); 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
4 5 molecular weights ranging from 2.8 x 10 to 2.2 x 10 (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 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
2+ complex could be partially disassociated by low pH or high Mg 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 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-methyl-
D-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%. 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 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 28
Sus1 gene has been described (Chourey et al. but, unlike the Sh1 mutant, p 1988), 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. 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
14 100% activity loss in 20 min for maize and cotton, respectively). Initially C-
14 sucrose cleavage was quantified by recovery of 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 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
1 C with a water potential of -1 .0 MPa (adjusted with PEG 8,000) with 2 g I" 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 5- to old in . 6-day seedlings and quick frozen liquid N2
14 Purification of C-Sucrose
Trace amounts of phosphorylated sugars are common impurities in
14 commercial C-sucrose, and can reduce accuracy of the assay. These were
14 removed by descending paper chromatography of commercially obtained C- 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
14 analysis (data not shown). Molarity and specific activity of purified C-sucrose 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
with mortar pestle. frozen transferred to mortar N2 a and The powder was another 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
14 Cleavage of C-sucrose by sucrose synthase was assayed in a 50 /xl
14 volume consisting of 20 n\ extract, 80 mM Mes (pH 5.5), 5 mM NaF, 100 mM C- sucrose 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
rotary shaker for was rinsed separately, first in 40 ml of H20 at 175 rpm on a 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
14 amount of the 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 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. 35
2
Time after extraction (min)
3-1 Figure . 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. 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 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 hetero- and homotetramers of sucrose synthase). Endosperm tissue contains 39
2.5
^2.0 + Casein .c c1.5 'o 2 1.0 Q_ OXD.5 o i ° B "§2.5 + Casein CO CD ^ ^ o2.0 CD C/) , _ o1.5 o 5 1.0 + Pi 0.5
0 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 P (10 mM) or casein (2% t w:v). 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 homo- and 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. 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., « '
c 1 f co o co ~| co -c E
OJ c H-
(DC 6 « 10 CO CD
^ s S > Q- « o g CD ^ CO c 2 c
co E eg 2 - Q1 E CO t«CO > CD fi c CO o CD LU CO c3 W -C +1 c c >> c Q to CD CD 10 -*— 0 CD OCO 0CO o D W +- 10; O CQ o CD CD CD c N E CD 'co cn c| E 0 — to CD 1c 53 CD to -C CO *r CD O O £ M" aCD cE CD °J « CO o CD ~J c » O Z CO So 81 ~* 5 S to O co o is ^ CO CD CD .£= o -a E CD =3 I C i- 2 CD C D) in O) I <5 ' CD o CO si
CO
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
14 14 also been possible to measure C-sucrose formed from UDP- 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 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
14 14 the incorporation of C-glucose into UDP-glucose from 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
14 separated from the excess 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 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 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
2+ presence of Mg . 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
2+ degradative reaction of sucrose synthase alone or in the presence of Mg .
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, 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. 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 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., 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. 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
1 water potential of -1.0 MPa (adjusted with PEG 8,000) with 2 g I' 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
1 paper in covered glass pans. Continuous airflow was provided (1 liter min" )
final h. throughout the germination period with 40% 02 supplied during the 48 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 side- arm 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
1 agitation airflow of I min" on a rotary shaker (125 rpm) and an 40% 02 (1 ) through an airstone in each flask. Experiments were terminated by twice rinsing in sterile
blotting in liquid . water, excess moisture and freezing them 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
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 53
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
1 1 (/nmol sucrose mg' 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 y immediately after excision. lntact refers to root tips quick frozen in liquid N2 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 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 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 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 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, 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 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 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, 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 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 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 Sh1.
Materials and Methods
Maize seed (Zea mays L, \N22:sh1) were primed for 6 days at 10 C with a
1 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 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% 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 1 00 ml side- arm 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
1 agitation on a rotary shaker rpm) airflow I" (125 and by an 40% 02 (1 ) through an airstone in each flask. Experiments were terminated by twice rinsing root tips in
sterile water, blotting moisture . excess and freezing them in liquid N2
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. 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 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 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 64
% glucose
Intact 0 0.2 0.5 2.0 4.0
Expt. 1
^^^^
Expt. 2
5-1 Figure . 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
** mrm* ** m ' Expt. 1 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 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
g"1 1 (jumol sucrose protein h" )
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 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. 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 69
1 6h 24h 48h 1 6h + 32h ~ 1 1 Intact + + +~ -/+ +/-
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. 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 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 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 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., 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. 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
1 1 activity (7.3 to 16.1 ^mol 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
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; 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 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
-80 excision. Tissue samples were weighed, frozen in liquid N2 and stored at 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 fine in liquid a powder 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
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, 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 0
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
14 assay quantifying the production of 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
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
1 1 M, pH 6.0) containing 0.24 mg ml" nitroblue tetrazolium, 0.14 mg ml* phenazine
1 1 methosulfate, 25 units ml' glucose oxidase and 5 mg ml" 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 CD
CO in co 1 O -r -H -H c 8 CO N- <2 a "c co cvi CD CO o co CO -t—' .>£6 c may o CD CO *~ g i E CD o d U) CD CO H +1 CD N 00 t- CO H— o z CO CO E n 31~ CO o _3 to - E CD C o c v CM cCO "co "a CD CO c 1— 3CD c -C o CO — LU eq CNJ CO c o CO CO c CO "D CD a +1 c C CO CD o E o o o >, 0) 3 o d CO Q_ CO ctiv rf yb CD +1 CO c o CO CD co E -H -H -H CO a. o CO O co E CM CO * CO CD 15 o 00 LO *- to vert i_ d d 5r CM i- CD CO c. CO Q) CO O CO ~o c f c co CM CM o 3 J3 aci edli O a) 00 cvi in d o -H -H -H CD LO +1 > c CO 3 -Q CO CO CO ^j- CO "D CD o CD 3 "O O CO CD CM d 9 £$ O O CO CO CM CO TO E i CO _ "CD CD ins E 3 CD 2 a. 2 a T3 TS X * CD -4-> C CD CO OT co d O CO to .Q- CD CD +-» H CO .Q LO CM CD 3 O "5 o d f> o Uo a O CD CCD CO CO i i= CO o to £ O o CD CD 1 4-' 3 1 o CD c E £CO CD CD "C o E E CO > CD if *- E E - - o ab O CO o c CO < O CD CM * CO CO i- I I I I I CD h- DC CO O CM CO CO M >.' $ ~ LU 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
g" 1 1 glucose FW h" -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
1 1 from this line. Also, from 83% to 94% (13.3 to 15.1 jumol glucose mg' protein h" ) 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). )
0 1— -o
jimol glucose /^moi giucose
1 1 1 1 mg" protein h" g FW h*
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 W W 1° roots 0.1 ± 0.1 0.2 ± 0.1 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. 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 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.
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 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 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
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 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 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. 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 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. 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 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. 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 DP. 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.
1 1 1 :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. 1 1 1 :381 -390.
Chapleo, S. and J.L Hall. 1989c. Sugar unloading in roots of Ricinus communis
L. III. The extravascular pathway of sugar transport. New Phytol. 111:391- 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. 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. 21 4:300-306.
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. 1 976. 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. 1991 b. 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 LC. 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. 98 de Fekete, M.A.R. and C.E. Cardini. 1964. Mechanism of glucose transfer from sucrose into the starch granules of sweet corn. Arch. Biochem. Biophys. 104:173-184. del Rosario, E.J. and V. Santisopasri. 1977. Characterization and inhibition of invertases in sugar cane juice. Phytochem. 16:443-445.
Delmer, D.P. 1972a. The purification and properties of sucrose synthetase from etiolated Phaseolus aureus seedlings. J. Biol. Chem. 247:3822-3828.
Delmer, D.P. 1972b. The regulatory properties of purified Phaseolus aureus sucrose synthetase. Plant Physiol. 50:469-472.
Delmer, D.P. and P. Albersheim. 1970. The biosynthesis of sucrose and nucleoside diphosphate glucose in Phaseolus aureus. Plant Physiol. 45:782-786.
Dick, P.S. and T. ap Rees. 1 975. The pathway of sugar transport in roots of Pisum sativum. J. Expt. Bot. 26:305-314.
Dick, P.S. and T. ap Rees. 1 976. Sucrose metabolism by roots of Pisum sativum Phytochem. 15:255-259.
Dickinson, CD., T. Altabella and M.J. Chrispeels. 1 991 . Slow-growth phenotype of transgenic tomato expressing apoplastic invertase. Plant Physiol. 95:420- 425.
Dixon, M. and E.C. Webb. 1979. Enzymes, 3rd ed. New York: Academic Press.
Doehlert, D.C. 1 987. Substrate inhibition of maize endosperm sucrose synthase by fructose and its interaction with glucose inhibition. Plant Sci. 52:153-158.
Doehlert, D.C. and F.C. Felker. 1987. Characterization and distribution of invertase in developing maize (Zea mays) kernels. Physiol. Plant. 70:51-57.
Doehlert, D.C. and S.C. Huber. 1983. Regulation of spinach leaf sucrose phosphate synthase by glucose-6-phosphate, inorganic phosphate, and pH. Plant Physiol. 73:989-994.
Doehlert, D.C, T.M. Kuo and F.C. Felker. 1988. Enzymes of sucrose and hexose metabolism in developing kernels of two inbreds of maize. Plant Physiol. 86:1013-1019. .
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.
Echeverria, E. and T. Humphreys. 1 984. In vitro and in vivo inhibition of sucrose synthesis by sucrose. Phytochem. 23:2727-2731
Echeverria, E. and T. Humphreys. 1 985. 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. 1 965. 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. 1 986a. 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 of . Plant, Cell source C02 Env. 8:427-438.
Faye, L and C. Berjonneau. 1979. Evidence for the glycoprotein nature of radish jS-fructosidase. Biochemie 61 :330-332.
Faye, L. C. Berjonneau and P. Rollin. 1981. Studies on 0-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 (Sefa vulgaris ssp. altissima). J. Plant Physiol. 131:153-162.
Ruhr, R., C. Kuhlemeier, F. Nagy and N-H. Chua. 1986. Organ-specific and light- induced expression of plant genes. Science 232:1106-1112. 100
Freeling, M. and D.C. Bennett. 1985. Maize Adh1. Ann. Rev. Genet. 19:297-323.
Fondy, B.R. and D.R. Geiger. 1982. Diurnal pattern of translocation and carbohydrate metabolism in source leaves of Beta vulgaris L. Plant Physiol. 70:671-676.
Gabrielyan, N.D., E.B. Lapina, S.M. Spiridonova, V.N. Shibaev, E.I. Budowsky and K.N. Kochetkov. 1969. The interaction of adenosine diphosphate glucose
and its synthetic analogs with uridine diphosphate glucose-fructose
glucosyltransferase from pea seedlings. Biochem. Biophys. Acta. 1 85:478- 481.
Gascon, S., N.P. Neumann and J.O. Lampen. 1968. Comparative study of the properties of the purified internal and external invertases from yeast. J. Biol Chem. 7:1573-1577.
Gerdemann, J.W. 1964. The effect of mycorrhizas on the growth of maize. Mycologia 56:342-349.
Gerdemann, J.W. 1968. Vesicular-arbuscular mycorrhiza and plant growth. Ann. Rev. Phytopath. 6:397-418.
Giaquinta, R.T. 1979. Sucrose translocation and storage in the sugar beet. Plant Physiol. 63:828-832.
Giaquinta, R.T., W. Lin, N.L Sadler and V.R. Franceschi. 1983. Pathway of phloem unloading of sucrose in corn roots. Plant Physiol. 72:362-267.
Gibson, D.M. and W.E. Shine. 1983. Uridine diphosphate glucose breakdown is
mediated by a unique enzyme activated by fructose 2,6-bisphosphate in Solanum tuberosum. Proc. Natl. Acad. Sci. (USA) 80:2491-2494.
Gifford, R.M. and L.T. Evans. 1981. Photosynthesis, carbon partitioning and yield. Ann. Rev. Plant Physiol. 32:485-509.
Glasziou, K.T. and K.R. Gayler. 1972. Storage of sugars in stalks of sugar cane. Bot. Rev. 38:471-490.
Grimes, W.J., B.L Jones and P. Albersheim. 1970. Sucrose synthetase from Phaseolus aureus seedlings. J. Biol. Chem. 245:188-197. 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. 1 0:21 5- 224.
Hannah, LC. 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.
Hattori, 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. 1 8:444-453.
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. 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. 1 978. 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. 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, LE. 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, LF. 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. .
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, 1 973. Solubility of plant invertases. Phytochem. 1 2:67-71
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. 1 986. 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, CP. and R. Sood. 1976. Ecophysiological regulation of invertase activity in pea pollen. Ind. J. Ecol. 3:44-48.
Marana, C. F. Garcia-Olmedo and P. Carbonero. 1 990. Differential expression of two types of sucrose synthase-encoding genes in wheat in response to anaerobiosis, cold shock and light. Gene 88:167-172. .
105
Masuda, H. and S. Sugawara. 1980. Purification and some properties of cell wall bound invertases from sugar beet seedlings and aged slices of sugar beet. Plant Physiol. 66:93-96.
Masuda, H., T. Takahashi and S. Sugarwara. 1988. Acid and alkaline invertases in suspension cultures of sugar beet cells. Plant Physiol. 86:312-317.
Matsushita, K. and I. Uritani. 1974. Change in invertase activity of sweet potato in
response to wounding and purification and properties of its invertases. Plant Physiol. 54:60-66.
Matsushita, K. and I. Uritani. 1 976. Isolation and characterization of acid invertase inhibitor from sweet potato. J. Biochem. 79:633-639.
Matsushita, K. and I. Uritani. 1977. Synthesis and apparent turnover of acid invertase in relation to invertase inhibitor in wounded sweet potato root tissue. Plant Physiol. 59:879-883.
McCarty, D.R. 1986. A rapid and simple method for extracting RNA from maize tissues. Maize Gen. Coop. Newsletter 60:61
McCarty, D.R., J.R. Shaw and LC. Hannah. 1986. The cloning, genetic mapping, and expression of the constituitive sucrose synthase locus of maize. Proc. Natl. Acad. Sci. (USA) 83:9099-9103.
McCollum, T.G., D.J. Huber and D.J. Cantliffe. 1988. Soluble sugar accumulation
and activity of related enzymes during muskmelon fruit development. J. Amer. Soc. Hort. Sci. 113:399-403.
McElfresh, K.C. and P.S. Chourey. 1988. Anaerobiosis induces transcription but not translation of sucrose synthase in maize. Plant Physiol. 87:542-546.
Metzenberg, R.L 1963. The purification and properties of invertase of Neurospora. Arch. Biochem. Biophys. 100:503-511.
Milner, Y. and G. Avigad. 1964. The UDP-glucose: D-fructose glucosyltransferase system from sugar beet roots. Isr. J. Chem. 2:316-320.
Miron, D. and A.A. Schaffer. 1991 . Sucrose phosphate synthase, sucrose synthase
and invertase activities in developing fruit of Lycopersicon esculentum Mill, and the sucrose accumulating Lycopersicon hirsutum Humb. and Bonpl. Plant Physiol. 95:623-627. 106
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: 1 49-1 54.
Moriguchi, T. and S. Yamaki. 1988. Purification and characterization of sucrose
synthase from peach (Prunus persica) fruit. Plant and Cell Physiol. 29:1361- 1366.
Morrell, S. and T. ap Rees. 1986. Sugar metabolism in developing tubers of Solanum tuberosum. Phytochem. 25:1579-1586.
Mujer, C.V., M.E. Rumpho, T.C. Fox and R.A. Kennedy. 1990. Developmental expression of two sucrose synthase isozymes in anerobically-grown seeds of Echinochloa phyllopogon. Plant Physiol. 93S:101.
Murata, T. 1 971 . Enzymic mechanisms of starch synthesis in sweet potato roots. V. Sucrose synthetase of sweet potato roots. Agric. Biol. Chem. 35:1441- 1448.
Murata, T. 1975. Soluble nucleotides in Konjak corn. Agric. 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. 1 8:309-356.
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. 1 979. Subcellular distribution of gluconeogenetic enzymes in germinating castor bean endosperm. Plant Physiol. 64:31 -37. 107
Paiva, E., 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. 1 975. Activities of enzymes of sugar metabolism in cold-stored tubers ol Solarium tuberosum. Phytochem. 14:613-617.
Pontis, H.G. 1977. Riddle of Sucrose, pp. 79-117 in D.H. Northcote, ed. Plant Biochemistry: Intl. Rev. Biochem. Vol. XIII. Baltimore: University Park Press.
Pontis, H.G., R.A. Wolosiuk, LM. 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. 1 975. 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. 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 /3-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. 1 958. Sucrose and growth of excised roots of an inbred lea mays. Proc. Natl. Acad. Sci. (USA) 44:1210-1212.
Roberts, D.W.A. 1 973. 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. 1 952. 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, LJ. 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, LJ. 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 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. 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 5201- 5205. 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. 10:302- 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-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. 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. 141:440- 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. 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. 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 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 (Lc. 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 )J^cuA Dean, Allege of Agria£jKure
Dean, Graduate School