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Studies on the metabolism and accumulation of a, a-trehalose in rhizobia and legume nodules
Hoelzle, Inger Alice, Ph.D. The Ohio State University, 1990
UMI 300 N. Zeeb Rd. Ann Arbor, MI 48106 STUDIES ON THE METABOLISM AND ACCUMULATION OF
a,a-TREHALOSE IN RHIZOBIA AND LEGUME NODULES
DISSERTATION
Presented in Partial Fulfillment of the Requirements for
the Degree Doctor of Philosophy in the Graduate
School of the Ohio State University
By
Inger Alice Hoelzle, B.S., M.S.
*****
The Ohio State University
1990
Dissertation Committee: Approved by A.L. Barta
W.D. Bauer Adviser F.D. Sack department of Agronomy
G.K. Sims ACKNOWLEDGEMENTS
The guidance and advice of Dr. John G. Streeter throughout the course of my research are greatly
appreciated. Further thanks are given to the other members of my committee, Drs. Allen Barta, Dietz Bauer, Fred Sack, and Gerry Sims. Mary Kilpatrick frequently provided me with excellent instruction and invaluable advice that.is sincerely and gratefully acknowledged. The advice and training in molecular biology techniques of Drs. Mike
McMullen and Ron Diebold, and Mark Jones was also very helpful and is appreciated.
My parents, William and Margaret Hoelzle, have been an inspiration since the day I first was old enough to realize how very special they are. Their unwavering support throughout my life has been a source of strength and comfort
I could never describe or repay properly. Ian Lamb, my husband, also deserves far more thanks and appreciation than is possible to put down on paper. Finally, my aunt Jean
Hoelzle has always provided me with understanding and a wonderful perspective on life. Without the support of these loving people I could never have accomplished all that I have. VITA
January 28, 1956...... Born - Flint, Michigan
1981...... B.S. Botany, San Diego State University, San Diego, California 1983...... M.S. Biology, San Diego State University, San Diego, California
1983-1985...... Graduate Assistant, Virginia Polytechnic Institute and State University, Blacksburg, Virginia
1985-Present...... Graduate Research Associate, The Ohio State University, Columbus, Ohio
PUBLICATIONS
Hoelzle, I., and Streeter, J.G. 1989. Higher trehalose accumulation in rhizobia under salt stress. Plant Physiol. 89S:118.
Hoelzle, I., and Streeter, J.G. 1990. Stimulation of a-glucosidases from fast-growing rhizobia and Aqrobacterium tumefaciens by K+, NH+4, and Rb+. Can. J. Microbiol. 36:223-227. FIELDS OF STUDY
Major Field: Agronomy
Studies in Plant Physiology and Plant Biochemistry. Professors Ken D. Johnson, John L. Hess, and Miller B. McDonald.
Studies in Soils. Professors T. Logan, F. Himes, G. Sims, and W.A. Dick.
Studies in Molecular Biology. Professors G. Marzluf, E. Vanin, Lee F. Johnson and Mike D. McMullen. TABLE OF CONTENTS
ACKNOWLEDGEMENTS...... ii
VITA...... iii
LIST OF TABLES...... viii
LIST OF FIGURES...... ix
LIST OF PLATES...... X
INTRODUCTION...... 1
CHAPTER PAGE
I. LITERATURE REVIEW...... *...... 7
General description of trehalose...... 7 Natural occurrence of trehalose...... 10 Factors affecting trehalose concentration in organisms...... 13 Functions of trehalose...... 19 Metabolism of trehalose...... 38 Summary of trehalose occurrence and metabolism in rhizobia...... 52
II. INCREASED ACCUMULATION OF TREHALOSE IN SOME RHIZOBIA WHEN CULTURED UNDER OSMOTIC STRESS...... 55
Introduction...... 55 Materials and Methods...... 57 Results...... 58 Discussion...... 60
III. STIMULATION OF a-GLUCOSIDASES FROM FAST-GROWING RHIZOBIA AND AGROBACTERIUM TUMEFACIENS BY K+, Rb+, AND NH+4...... 67
Introduction...... 67 Materials and Methods...... 68 Results...... 72 Discussion...... 75
v IV. INCREASED ACCUMULATION OF TREHALOSE IN RHIZOBIA CULTURED WITH 1% OXYGEN...... 87
Introduction...... 87 Materials and Methods...... 90 Results...... 94 Discussion...... 98
V. GENETIC STUDIES...... 106
Introduction...... 106 Materials and Methods...... 108 Results...... 116 Discussion...... 119
V. CONCLUSIONS...... 135
APPENDICES
A. CHROMATOGRAPHY TECHNIQUES...... 140
Fractionation of organic acids, amino acids and sugars...... 141 HPLC analysis of organic acids...... 143 Gas chromatographic analysis of carbohydrates..145
B. BACTERIAL TRANSFORMATION WITH PLASMID DNA...... 147
C. ISOLATION AND RESTRICTION DIGESTION OF DNA...... 149
Plasmid mini-prep...... 150 Isolation of genomic DNA from rhizobia...... 152 Restriction digestion of DNA...... 154
D. RUNNING AGAROSE GELS...... 155
E. SOUTHERN TRANSFER OF DNA TO A MEMBRANE...... 158
DNA transfer to a membrane...... 159 Disassembling a DNA transfer...... 161
F. NUCLEIC ACID HYBRIDIZATION...... 162
Aqueous DNA-DNA hybridization...... 163 Formamide DNA-DNA hybridization...... 164 Labeling random primed DNA probe...... 165
G. BLOT WASHING AND STRIPPING...... 167
Low stringency washing...... 168 High stringency washing...... 169 Blot stripping...... 170
vi H. RECIPES FOR MEDIA AND SOLUTIONS...... 171
Recipes for solutions...... 172 Recipes for media...... 176
LIST OF REFERENCES...... 180
vii LIST OF TABLES
TABLE PAGE
1. Trehalose accumulation in R^. lecruminosarum bv. phasedi USDA 2667 in different media and with different salts...... 65
2. Trehalose accumulation by various rhizobia grown in succinate medium with 100 mM excess NaCl...... 66
3. Response to K+ by a-glucosidases from different sources...... 82
4. Effect of different salts on trehalase activity from Ri. lequminosarum bv. phasedi USDA 2667...... 84
5. Kinetic properties of a-glucosidases in crude extracts from Rhizobium lequminosarum bv. phaseoli USDA 2667 grown with a,a-trehalose or glutamate as sole carbon source...... 85
6. Comparison of a-glucosidase activity in crude extracts of two species of rhizobia grown with sucrose or a,a-trehalose as the sole carbon source...86
7. Trehalose accumulation in cells of Rj_ lequminosarum bv. phaseoli USDA 2667 grown with different carbon sources under 21% or 1% oxygen...... 104
8. Accumulation of trehalose in different species of rhizobia grown under 21% or 1% oxygen...... 105
9. Effect of switching from 1% to 21% oxygen on trehalose accumulation in cells of R. lequminosarum bv. phaseoli USDA 2667...... 106
10. Summary of trehalose accumulation patterns in selected Tn5 mutants of Rj. bv. phaseoli USDA 2667...... 135
viii LIST OF FIGURES
FIGURE PAGE
1. Known reactions of trehalose metabolism...... 39
2. Effect of various concentrations of excess NaCl on trehalose accumulation in Ei. lequminosarum bv. Phaseoli USDA 2667...... 64
3. Effect of KC1 concentration on trehalase activity in extracts from R^. lequminosarum bv. phaseoli USDA 2667...... 79
4. pH curves for a-glucosidases from R^. lequminosarum bv. phaseoli USDA 2667...... 81
5. Trehalose and protein levels in cultures of R. lequminosarum bv. Phaseoli USDA 2667 grown in succinate medium under 1% or 21% 02 and harvested at various times during early through late log phases of growth. Increasing protein levels indicate more advanced growth stages within a treatment...... 103
ix LIST OF PLATES
PLATE PAGE
I. Genomic DNA from lequminosarum bv. phaseoli USDA 2667 and CFN42, Rs. meliloti USDA 1021 and E. coli probed with labeled otsA DNA. Films were intentionally overexposed to demonstrate complete lack of binding to rhizobial DNA...... 128
II. Genomic DNA from R^. lequminosarum bv. phaseoli USDA 2667 and CFN42, R^. meliloti USDA 1021 and E. coli probed with labeled treA DNA. Films were intentionally overexposed to demonstrate complete lack of binding to rhizobial DNA...... 130
III. Genomic DNA from Rj_ lequminosarum bv. phaseoli USDA 2667 and CFN42, Rj. meliloti USDA 1021 and E. coli probed with labeled node DNA. The membrane used for Plate I was stripped and Re-probed with the rhizobial probe to demonstrate that the rhizobial DNA could bind DNA of sufficient homology...... 132
IV. Genomic DNA from Rj_ lequminosarum bv. phaseoli USDA 2667 and CFN42, R*. meliloti USDA 1021 and E. coli probed with labeled node DNA. The membrane used for Plate II was stripped and re-probed with the rhizobial probe to demonstrate that the rhizobial DNA could bind DNA of sufficient homology...... 134
x INTRODUCTION
Bacteria in the genera Rhizobium and Bradyrhizobium are unique in their ability to form symbioses with many different species of leguminous plants. These bacteria form colonies in nodules on the roots of receptive legume species, where substantial differentiation of plant tissue occurs and the bacteria establish themselves (as differentiated "bacteroids") in the plant cells. Within the nodules atmospheric nitrogen (N2) is reduced to ammonium
(NH4+) and assimilated into amino acids or ureides before export to the xylem stream of the plant. In "exchange" for this supply of nitrogen the plant provides the fixed carbon necessary for bacteroid growth and metabolism (including N2 fixation energy demands).
The agronomic, economic and ecological importance of biological N2 fixation can hardly be overstated. Many important agricultural crops are legumes, representing an essential source of vegetable protein in both developed and undeveloped countries. The utilization of legume protein for human nutrition is important because of low availability of meat in many countries and because of increased environmental concern about the vast areas of prime
1 agricultural land dedicated to the support of livestock, especially those animals raised purely for human consumption. Use of this land for production of legume crops utilized either for direct human consumption or support of dairy animals would represent both an enormous energy savings and a significant decrease in the amount of land needed to support human populations. Another economic and ecological concern relating to legume-rhizobial symbioses is the amount of fertilizer used for crops that do not fix N2. Nitrogen fertilizers are energetically very costly to produce, and repeated overuse of fertilizers frequently results in detrimental effects on the soil and serious contamination of water systems. Finally, cultivation of legume crops not only decreases the amount of nitrogen fertilizer immediately needed, it also reduces the amount required for non-legume crops grown in rotation with legumes (because of residual nitrogen in the soil).
Both applied and basic research on legume symbioses is vital to insure the continued optimization of this important phenomenon in agricultural systems. A large amount of research has centered on the environmentally important and biologically intriguing nitrogen metabolism associated with
N2-fixing systems. However, nodule carbon metabolism has traditionally received much less attention in spite of the fact it is equally as important as N2 metabolism to the overall function of N2-fixing symbioses. In recent years some of the pathways of energy metabolism and the occurrence
of various carbohydrates have been documented, however, many
questions remain unanswered. For example, a particular
disaccharide, a,a-trehalose, has been identified in every
legume symbiosis examined. In cultured rhizobia and
bradyrhizobia trehalose is the sole mono- or disaccharide
detectable (Streeter, 1985). Nevertheless, in spite of the ubiquitous nature of this sugar, its metabolic function in
rhizobia is not known.
Trehalose is an intriguing sugar and occurs in many other organisms, in some cases playing a very specialized role in the life cycle. In organisms capable of complete dehydration, such as brine shrimp eggs, nematode cysts and
fungal spores, trehalose is vital to survival, apparently because it stabilizes membranes when the water of hydration
is lost (Crowe et al.. 1984a). In most marine algae and some Ei. coli strains, trehalose is utilized as an osmoticum during times of hyperosmotic environmental conditions (Reed et al.. 1984a; Larsen et al.. 1987; Rod et al.. 1988). In this role trehalose is synthesized in response to increasing osmotic concentration of the surrounding sea water, and the tendency to plasmolyze is reduced. A similar role for trehalose has been suggested in some insects and arthropods adapted to extreme cold; trehalose and other small molecular weight carbohydrates accumulate in their hemolymph as a sort of "antifreeze" (Baust, 1983). In other systems trehalose is a source of metabolic energy; it is the blood sugar of
insects (Florkin and Jeuniaux, 1974), and in many fungi it
is stored in reproductive structures and/or utilized as a
specialized carbon source (Thevelein, 1984a). Whether
rhizobial trehalose functions in a role similar to any of those described above is not clear.
Nodules have been documented to contain several other unusual carbon compounds, such as the cyclitols pinitol, ononitol and chiro-inositol, in addition to the disaccharide trehalose (Streeter, 1980). There also are some more common carbohydrates in legume nodules, such as sucrose, dicarboxylic acids, and the carbon storage polymers
(3-hydroxybutyrate and glycogen; these compounds generally occur in far greater amounts than trehalose (Streeter,
1987). In the symbiotic state it is generally believed that bacteroid metabolism and energy for N2 reduction is supported by dicarboxylic acids (Stowers, 1985). These facts suggest that trehalose is not a primary bacteroid energy source or a carbon storage compound in nodules.
Some preliminary studies documenting the occurrence, location, and enzymology of trehalose in rhizobia have been published (Streeter, 1980, 1981, 1982, 1985; Reibach and
Streeter, 1983; Salminen and Streeter, 1986). These papers have established some important points: 5
1) Trehalose has been detected in every strain or
species of Rhizobium and Bradvrhizobium examined.
[Trehalose also occurs in other N2-fixing organisms
such as cyanobacteria (Reed et al.. 1986) and
Frankia sp. (Lopez et al.. 1983)]
2) Trehalose does not appear in the nodules until the
onset of N2 fixation.
3) Trehalose is synthesized by the bacteroids but a
significant portion is transported to host tissues.
4) A greater proportion of total nodule trehalose is
sequestered in bacteroids as the nodules age, and
the concentration of trehalose increases in
senescing nodules.
The above statements suggest a role of trehalose in the life cycle of rhizobia, and possibly in the N2-fixing state.
The following literature review and summary of experiments is the result of efforts directed toward gaining a greater understanding of trehalose and its role in rhizobia and legume nodules. The initial goal of this research was to produce and isolate two types of rhizobial mutants: 1), those that could not synthesize trehalose and 2) those that could not catabolize trehalose. These mutants were to be characterized biochemically and then used in inoculation experiments to determine what effects the mutations would have on the symbiosis, possibly identifying the role of trehalose. When this goal proved untenable (the desired mutants were not obtained) several other experiments were performed to define some factors affecting the occurrence of trehalose, based largely on ideas generated by the literature concerning trehalose in other organisms
(especially other gram- bacteria) or ancillary observations from other experiments. Finally, some experiments were conducted to try to determine the cause(s) for the problems encountered in generating rhizobial trehalose mutants. CHAPTER I
LITERATURE REVIEW
A. General Description of Trehalose
A.1. Physical Properties
Trehalose (1-0-a-o-glucopyranosyl-a-D-glucopyranoside, rarely called mycose or mushroom sugar) occurs naturally as an a-o-glucose disaccharide of approximate molecular weight
342 g/mole and chemical formula A few reports claim to have detected oc-/3- or /3-/3- isomers occurring naturally but these cases have not been well documented (see
Elbein, 1974). All 3 isomers have been chemically synthesized and chromatographically separated (Birch, 1963;
Cook et al.. 1985; Parrish et al.. 1987), and recently a method for extraction of 14C-labeled trehalose from E^ coli culture grown in the presence of 14C-glucose has been published (Brand and Boos, 1989).
Trehalose is a sweet sugar (Birch, 1979), is soluble in dilute ethanol and water, will crystallize in cold 80% ethanol (as a dihydrate), and has a melting point of 96-97°C
(Birch, 1963). Trehalose is alkali stable (non-reducing), and yields a-glucose when hydrolyzed. Thorough characterizations of the crystalline state (Jeffrey and
7 Nanni, 1985) and the chemistry in solution (Bock et al..
1983; Vicent et al.. 1989) of this sugar have been published. The effects of radiolysis of trehalose have also been investigated (Diehl et al.. 1978).
A.2. Naturally Occurring Derivatives
Trehalose is a component of several fatty acid esters found in related bacteria such as Mycobacterium.
Corvnebacterium. Rhodococcus. and Nocardia (Elbein, 1974;
Tomiyasu et al.. 1986) . Trehalose dimycolate, formed by diesterification of trehalose with various mycolic acids, is found in the cell walls of Mycobacterium tuberculosis and other related bacteria. This compound (known as the "cord factor") is part of the antigenic factor that causes tubercles in infected mammalian tissues. Different species of bacteria produce trehalose dimycolate with mycolic acids of differing chain lengths, but all appear to be capable of eliciting the same toxic response in infected mammals (Silva et al.. 1988). Both purified natural and synthetic trehalose dimycolates have been used to inoculate mice and induce some immunity to infection by mycobacteria (Yarkoni and Bekierkunst, 1976; Pimm et al.f 1979). A large body of literature exists concerning trehalose dimycolates and related compounds and is beyond the scope of this review; for a recent review see Brennan (1989).
Some Mycobacterium spp. are classified as "atypical" because they do not produce classical tuberculosis symptoms, although they are generally capable of producing some
symptoms similar to those of true tuberculosis. These
atypical Mycobacteria spp. form two colony types,
categorized as rough and smooth. Only the smooth variants possess unusual trehalose-containing lipooligosaccharides on the cell surface (Hunter et al.. 1983; Belisle and Brennan,
1989). This is in contrast to the true tuberculosis-causing
Mycobacteria spp. (which require the presence of trehalose dimycolates for pathogenicity); among the atypical variants, only those without trehalose-based lipooligosaccharides
(rough colony type) are capable of causing chronic
infections. Various other trehalose-containing glycolipids, oligosaccharides, and sulfolipids have also been isolated
from mycobacteria and related genera (Elbein, 1974; Saadat and Ballou, 1983; Ristau and Wagner, 1983; Tomiyasu et al..
1986; Shimikata et al.. 1985); the role of these compounds
is not well understood.
It is noteworthy that the genus of N2-fixing bacteria,
Frankia. and the mycobacteria discussed above are both included in the actinomycetes (Brock, 1979). Accumulation of an internal pool of free trehalose has been documented in
Frankia HFPArl3 (Lopez et al., 1984), but the cell wall composition has apparently not been investigated.
Furthermore, some species of Mycobacterium can also fix N2
(Brock, 1979); it would be interesting to know if these organisms are similar to Frankia and contain significant 10 amounts of free trehalose in addition to the cell wall trehalose dimycolates already discussed. This may not be the case, because a related bacterium, Brevibacterium flavum, produces free trehalose but does not synthesize mycolic acids (London and Walker, 1985; Jones and Keddie,
1986).
A few other naturally occurring trehalose-containing compounds have been identified; examples include selaginose, a trisaccharide from Selaainella (Fischer and Kandler,
1970) , 3-0-a-D-glucopyranosyl-a,j9-trehalose from
Streptococcus faecalis (Fischer and Krieglstein, 1967), and
2-21-dideoxy-a,a'-trehalose from yeast (Farkas et al..
1969). Some Streptococcus spp. secrete trehaloseamine
(2-amino-2-deoxy-a-D-trehalose), a competitive inhibitor of trehalase, the enzyme that hydrolyzes trehalose (Umezawa,
1967).
B. Natural Occurrence of Trehalose
B.1. Funai
Trehalose was first isolated from rye ergot by Wiggers in 1832 (Birch, 1963). Since then it has been found in virtually all fungi, occurring in varying concentrations and locations depending on the fungal species, developmental stage, and culture or environmental conditions (Elbein,
1974; Thevelein, 1984a; Van Laere, 1989). 11 B.2 Insects
Trehalose has been detected in the hemolymph of nearly all insects, where it is the primary blood sugar (Wyatt and
Kalf, 1957; Barker and Lehner, 1976? Florkin and Jeuniaux,
1974), and it also occurs in other insect tissues (Candy and
Kilby, 1961).
B.3. Bacteria
Trehalose has been found in a wide range of bacteria, including the gram- bacterial genera Rhizobium and
Bradvrhizobium (Streeter, 1985), E_j. coli (Ogino et al..
1982? Larsen et al.. 1987) and Pseudomonas fluorescens
(Smith and Smith, 1989). It has also been documented in 2 genera of archaebacteria, Sulfolobus solfataricus (Nicolaus et al.. 1988) and Methanobacterium thermoautotrophicum
(Evans et al.. 1986), and in the actinomycetes Arthrobacter paraffineus (Suzuki et al.. 1971), Micrococcus spp. (Ahmad et al.. 1980), Streptomyces qriseus (Martin et al.. 1986?
McBride and Ensign, 1987a, b), Propionibacterium
(Stjernholm, 1958), and Frankia HFPArl3 (Lopez et al.. 1983,
1984). Trehalose has been found in the corynebacteria
Brevibacterium flavum (London and Walker, 1985)
Microbacterium ammoniaphilum (Walker et al.. 1982), and
Cellulomonas uda (Schimz et al., 1985; Schimz and Overhoff,
1987), and in the halophilic bacterium Ectothiorhodospira halocloris (Galinski and Triiper, 1982). 12
B.4. Miscellaneous organisms
Trehalose is not so ubiquitous in other taxonomic groups but has been detected in slime molds (Ceccarini and
Filosa, 1965; McBride and Zusman, 1989), liverworts (Matsuo et al.. 1986), many different species of cyanobacteria (Heed et al.. 1986), and the protozoan Euglena gracilis (Dwyer,
1986). It also frequently occurs in helminths (Fairbairn,
1958; Powell et al.. 1986), nematodes (Powell, et al.. 1986;
Madin et al.. 1985), and some red algae, annelids, and crustaceans (see Elbein, 1974). It apparently does not occur in higher plants (Gussin, 1972; Elbein, 1974).
B.5. Svmbioses and Parasitic Associations
It is of some interest that trehalose is found in a great number of symbiotic and parasitic associations.
Trehalose is found in symbiotic N2-fixing legume nodules
(Streeter, 1985), N2-fixing nodules formed by Frankia sp. in association with woody trees such as Casuarina sp. (Lopez et al.. 1984), in mycorrhizae formed by the symbiotic association of fungi with a diverse range of plants (Martin et al.f 1987), in lichens formed by fungi in association with algae (Iacomini et al.. 1987), and in Chlorella sp. in a symbiosis with Paramecium bursaria (Pardy et al.. 1989).
Trehalose is found in many parasitic associations such as protozoan (Undeen et al.. 1987), and nematode infections
(Powell, et al.. 1986), and it also occurs in many fungal 13 infections of plants (Evans et al.. 1981; Edwards and Allen,
1966; Lunderstadt, 1966; Long and Cooke, 1974).
C. Factors Affecting Trehalose Concentration in Organisms
C.1. Microorganisms in Culture
C.l.a Media
Although most fungi accumulate trehalose to some extent, the concentration can be altered by culture conditions (see Thevelein, 1984a; Van Laere, 1989); many researchers have documented changes in trehalose levels in fungi cultured under various conditions. Maltose or galactose media will stimulate trehalose accumulation in yeast (Panek and Matoon, 1977), and acetate-sucrose media under limited aeration or gluconeogenic carbon substrates will induce trehalose accumulation in Aspergillus nidulans
(Dijkema et al.. 1985). The addition of nitrogen to yeast culture media can affect the glucose-stimulated breakdown of
\ _ . trehalose during germination (Thevelein and Jones, 1983). A mannitol-arabinose based medium stimulated trehalose accumulation in Rhizobium spp., although the nitrogen source also affected trehalose concentration (Streeter, 1985).
When glutamate was supplied as both carbon and nitrogen source the bacteria had a lower concentration of trehalose than with mannitol-arabinose media, but essentially no other mono- or disaccharides were detected. However, because trehalose accumulates during log phase of growth in rhizobia 14 and is broken down in stationary phase (Streeter, 1985), and some cultures were harvested at different time points during this study, it is possible that some differences observed were due to growth stage and not to nutritional conditions.
Another bacterium, Cellulomonas sp. DSM20108, accumulated trehalose when cultured with several different carbon sources, including those which required cellulase for utilization (Schimz et al.. 1985).
Starvation and reduced growth rate (due to environment or stage of development) will induce trehalose synthesis and accumulation in fungi (see Thevelein, 1984a), but trehalose was degraded during starvation of the bacterium Cellulomonas sp. (Schimz et al.. 1985; Schimz and Overhoff, 1987). In the latter studies trehalose accumulated during logarithmic growth and disappeared during stationary phase. Rhizobia show the same pattern of accumulation during log phase and degradation during stationary phase (Streeter, 1985).
Starvation for glucose, nitrogen, phosphate, or sulfur will induce trehalose synthesis and accumulation in yeast (Kuenzi and Fiechter, 1972? Lillie and Pringle, 1980).
Oxine (8-hydroxyquinoline), an inhibitor of bacterial and fungal growth, stimulates trehalose accumulation in
Allomvces macrogynus, as does cycloheximide; the mechanism of this effect is not clear but may be due to disruption of membrane transport systems (Youatt, 1982). 15
C.l.b Uptake from Culture Media
Obviously the internal concentration of trehalose can be affected by the external presence of the sugar, and the ability of the microorganism to absorb it. Some organisms are incapable of utilizing external trehalose, for example, members of the bacterial genus Bradvrhizobium synthesize trehalose but cannot use it as a carbon source. Little is known about trehalose transport in most microorganisms, however, some information is available concerning yeasts and a few bacteria.
Trehalose is transported by a phosphotransferase system in ELs_ coli. Salmonella typhimurium (Marechal, 1984) and
Vibrio parahaemolvticus (Kubota et al.. 1982). Postma et al. (1986) found that trehalose was transported in S. tvphimurium either by the mannose phosphotransferase system, or by a galactose permease if the former transport system was unavailable, somewhat in contradiction to the results of
Marechal (1984). There is some evidence that the transport of trehalose via the mannose phosphotransferase system does not involve phosphorylation, in contrast to the usual mechanism of sugar uptake by microorganisms (Postma et al..
1986). This is also the case in yeast; both the soil yeast
Trichosporon cutaneum (Mortberg and Neujahr, 1986) and S. cerevisiae (Koytk and Michaljanicova, 1979) have been reported to transport trehalose without phosphorylation.
Mandels and Vitols (1967) found inducible and constitutive 16
trehalose transport mechanisms in the fungus Mvrothecium verrucaria.
C.l.c Environmental Conditions
High temperature incubation of yeast cells on glucose media produces cells with higher levels of trehalose than
those grown at lower temperatures (Grba et al.. 1975; Hottiger et al.. 1987a, b). Many bacteria and cyanobacteria
and some rhizobia increase their internal trehalose
concentration in response to osmotic stress (see section
D.3.). Some insects increase the trehalose level in their hemolymph in response to falling temperatures in the autumn
(see section D.4.a), and hemolymph trehalose concentration
falls during flight (Jutsum and Goldsworthy, 1976; King et al.. 1986). C.2. Developmental Effects on Trehalose Concentration
C.2.a. Fungi
Different growth stages can affect the concentration of trehalose in fungi. During fruiting trehalose may become the predominant carbohydrate in reproductive structures
(Thevelein, 1984a). In yeast, trehalose accumulated in the maturation phase of the cell cycle and, in several other species of fungi, trehalose synthesis and storage was correlated with formation of reproductive structures (Ng et al. . 1974; Catley and Kelly, 1975; Kiienzi and Fiechter,
1969). Yeast cells metabolize trehalose during budding, and accumulate it during the maturation phase of the cell cycle 17
(Kiienzi and Fiechter, 1969; Van Doorn et al.. 1988), with essentially none present in actively growing cells
(Thevelein, 1984a). In Dictvostelium sp. (a slime mold) there is a gradient of increasing trehalose concentration from the fruiting body to the stalk cells (Wright et al..
1982) .
C.2.b. Insects
Trehalose accumulates with age in beetles, representing a larger proportion of the total stored carbohydrates in older individuals (Rastogi and Dhand, 1985). Changes in metabolic demand, such as flight, diapause, and progression to more advanced growth stages can alter the level of free trehalose in the insect hemolymph and muscles, presumably as different demands are placed on stored carbohydrate (Islam and Roy, 1982). Various insect larvae accumulate trehalose in preparation for overwintering (Goto et al.. 1986; Tsutsui et al.. 1988; Lee and Lewis, 1985). Hemolymph trehalose levels in some insects can be elevated by certain regulatory neuropeptides (Gade, 1989). The trehalose concentration in hemolymph changes during flight, however the degree and length of time before recovery to non-flight levels varies with the type of insect. It appears that insects adapted to prolonged flight experience a sharp drop in hemolymph trehalose concentration when flying (Clegg and Evans, 1961;
Jutsum and Goldsworthy, 1976; Vaandrager et al.. 1988), but those not adapted to flight experience a slight increase in 18 hemolymph trehalose concentration (King et al.. 1986).
Trehalose synthesis and secretion by the insect fat body is apparently regulated by the corpus cardiacum (a hormone- secreting gland in the head); the effect may be mediated via trehalose-6-phosphate phosphatase (Steele, 1963? Steele and
Hall, 1985; Steele et al.. 1988).
C.2.c. Bacteria
Cultures of the bacterium Streptomyces antibioticus increased trehalose synthesis as sporulation began (Brana et al.. 1986), and high concentrations were documented in spores and aerial hyphae. Cultured rhizobia accumulate trehalose during the log phase of growth and break it down rapidly upon reaching stationary phase (Streeter, 1985).
Bacteroids (differentiated rhizobia or bradyrhizobia) generally have concentrations of trehalose than when cultured in liquid media (Streeter, 1985).
C.2.d. Legume nodules
Trehalose is not detected in legume nodules until the onset of N2 fixation (Streeter, 1980). The concentration increases during vegetative growth, flowering, and seed set, and all trehalose is lost as the plant becomes senescent
(Streeter, 1981). In young soybean nodules most of the trehalose present is localized in the cytosol (plant tissue) but in older nodules a greater proportion of the total nodules trehalose is found in the bacteroids (Streeter, 1985). 19 D. Functions of Trehalose
It is important to realize that there may be more than one function for trehalose in any given organism. For example, Van Laere (1989) suggests that trehalose may often function as a reserve carbon compound in fungi, but that additional roles for this sugar (such as desiccation or heat tolerance) are also possible.
D.1. Energy Supply
Trehalose is an important source of metabolic energy in nearly all insects. It is used for energy supply during flight, during chitin synthesis, and for cellular energy in many tissues. In the insect fat body (a tissue similar in function to the mammalian liver) the trehalose concentration stays relatively constant during molt but the glycogen concentration falls drastically? apparently glycogen is catabolized to supply trehalose to the hemolymph for chitin synthesis (Florkin and Jeuniaux, 1974).
In general, trehalose is thought to serve mainly as a specialized storage carbohydrate in fungi, utilized during periods of reduced growth (whether in response to some environmental insult or due to the developmental stage of the fungus) and in certain stages of germination or reproduction (Van Laere, 1989; Thevelein, 1984a; Donnini et al., 1988). There is some disagreement in the literature regarding the specific function and metabolism of trehalose in fungi, probably due to the use of different genera and 20 species for experiments, and to different culture conditions. Slow-growing, synchronous fungal cultures accumulate more trehalose than faster growing cultures, and this accumulation correlates positively with the onset of differentiation (Thevelein, 1984a). Yeast spores often have high levels of trehalose, which is usually metabolized during germination and considered essential for energy supply in germinating spores (Panek and Bernardes, 1983), although Van Laere et al. (1987) suggest it is broken down to glycerol during germination of Phvcomvces spores to enhance hydration of cellular components via osmotic effects. However, some results suggest that trehalose is not required for germination and may instead be important in dormancy (Donnini et al.. 1988; Lingappa and Sussman, 1959;
Inoue and Shimoda, 1981b). In some fungal spores, trehalose may be the source of energy during both dormancy and early germination stages. Dormant yeast spore survival depends on sufficient trehalose supply, stored during differentiation
(Lillie and Pringle, 1980).
The role of trehalose in bacterial metabolism has not been clarified. It is fairly common in occurrence (section
B.3), and has been suggested to be a storage form of carbon
(Schimz et al.. 1987), an energy source (Lopez et al..
1983), and an agent in desiccation tolerance (Martin et al.. 1986). 21
D.2. Structural Role
The presence of trehalose-containing compounds in some bacterial cell walls has already been discussed (section
A.2). Trehalose is esterified to fatty acids, forming trehalose dimycolate, and is incorporated into the cell walls of species of mycobacteria and corynebacteria. This glycolipid has been implicated in the antigenic reactions of infected tissues which result in tuberculosis; an additional function of trehalose dimycolate in strengthening and/or maintaining cell wall integrity of these bacteria is not clear. However, only strains of Mycobacterium kansaii that can synthesize trehalose-containing lipooligosaccharides have the "rough" colony morphology; this suggests that these glycolipids are responsible for some structural characteristics (Belisle and Brennan, 1989).
D.3. Osmoregulation
Marine fungi, a few bacteria, and cyanobacteria are examples of organisms that have the ability to osmoregulate in response to increased solute concentration of the surrounding (usually liquid) habitat (Jennings, 1983;
Borowitzka et al.. 1980; Measures, 1975; Csonka, 1989).
These organisms increase their internal osmotic pressure in response to increased external solute concentration, which allows them to maintain turgor. Several different types of compounds have been identified as osmotica under these conditions; they generally are low molecular weight 22 molecules that do not interfere with other physiological processes (Yancey et al.. 1982; Warr et al.. 1984; Somero,
1986). Examples of common osmoregulatory solutes are trehalose, sucrose, glycine, glutamate, quaternary ammonium compounds (such as glycine betaine), glycerol, glucosylglycerol (0-a-D-glucopyranosyl-(l-+2) -glycerol), and ions such as K+ and Na+.
D.3.a. Cvanobacteria
All species of cyanobacteria (previously called blue-green algae) tested have the ability to osmoregulate
(Reed et al.. 1984a; Stal and Reed, 1987); many species of cyanobacteria accumulate trehalose as a primary osmoticum.
Other frequently identified osmoregulatory solutes in cyanobacteria are sucrose, glucosylglycerol, and the quaternary ammonium compound glycine betaine. These may occur alone or in combination in osmoregulating cyanobacteria (Mackay et al.. 1983, 1984).
It has been suggested that in general, cyanobacteria can be divided into categories of salt tolerance by the type of osmoticum they accumulate; species from soil and freshwater tend to produce saccharides (such as trehalose), and more salt tolerant species accumulate quaternary ammonium compounds and glucosylglycerol, or a combination of these (Mackay et al.. 1983), although there are some exceptions to these trends (Mackay et al.. 1984; Reed et al., 1984b; Stal and Reed, 1987). Further support for the correlation between salinity tolerance and type of osmoticum was provided by Warr et al. (1987), who found that strains synthesizing disaccharides in response to osmotic stress
(generally those from intertidal zones) responded more rapidly to osmotic "upshock" than organisms synthesizing glucosylglycerol (which are more likely to be in deep ocean habitats). They suggest that this is correlated with the environment: intertidal and brackish waters are more subject to fluctuations in salinity than deep ocean water, so the ability to osmo-adapt rapidly is more important to organisms in the former habitat, which apparently are more likely to rapidly synthesize or degrade disaccharides in response to external osmotic changes.
Soirulina olatensis accumulates trehalose and glucosylglycerol in response to salt stress, but temperature can alter the ratio of these compounds (Warr et al.. 1985a).
While the total amount of osmoticum did not change significantly, at 37°C 31% of the total trehalose + glucosylglycerol was trehalose, but at 20°C trehalose represented only 9% of the total osmoticum present. The highest trehalose concentration noted in this study was at high temperature and low salt concentration. Muller and
Wegman (1978) noted an additional osmoregulatory solute accumulated at higher temperatures in Dunaliella tertiolecta that was not present at lower temperatures, and Warr et al.
(1985b) noted the same response in Svnechocvstis. It 24 appears that high temperatures alone or in concert with high osmotic pressures may induce osmotic adjustment in cyanobacteria.
Warr et al. (1985a) noted that glycogen concentration in cyanobacteria was inversely correlated with glucosylglycerol concentration: when these algae were transferred from high to low percent sea water the glycogen concentration increased and glucosylglycerol levels decreased. This is in contrast to the reaction of Rivularia atra to osmotic "downshock": trehalose accumulates as an osmoticum with incubation in high percent sea water, but is wastefully excreted when the alga is transferred to fresh water (Reed and Stewart, 1983), rather than being deposited as some non-osmotically active storage carbohydrate polymer.
The same study documented a positive correlation between low salinity and high nitrogenase activity in this alga.
D.3.b. Bacteria
Several studies have documented various E±. coli strains with higher levels of trehalose when under salt stress
(Strom et al., 1986; Larsen et al.. 1987; Rod et al.. 1988;
Giaever et al.. 1988; Munro et al. f 1989). Mutants lacking an enzyme necessary to synthesize UDP-glucose were consequently unable to synthesize trehalose and were osmo- sensitive (Strom et al.. 1986); this phenomenon was successfully used to screen for other trehalose-negative mutants (Giaever et al. . 1988). These authors also showed 25
increased activity of EL. coli trehalose phosphate synthase when assayed in vitro under high osmotic pressure.
Rod et al. (1988) screened over 20 strains of EL. coli
for increased trehalose accumulation under osmotic stress
and showed that some strains that had been in culture for many years had mutations that left them unable to respond to
osmotic stress by increasing their trehalose concentrations,
and also found that these strains were osmosensitive. They were able to trace the mutations through the ancestry of the
strains, and showed that the wild type EL_ coli strains are those that can accumulate trehalose in response to osmotic
stress, even though the strain EL. coli K-12 (which has been
in culture since 1922, longer than nearly all other strains) does not have this ability. Several strains that were descendants of EL. coli K-12 had regained the ability to
synthesize trehalose under osmotic stress because they had amber suppressor mutations in the affected genes. They suggest that amber mutations have occurred in cultures of descendants of Ej. coli K-12 (unable to synthesize trehalose) which were maintained in laboratories where the media was allowed to desiccate; the resulting osmotic stress led to selection for restoration of the ability to synthesize trehalose, via the amber suppressor mutations.
Other studies (Larsen et al.. 1987; Dinnbier et al..
1988) have found that EL. coli accumulates glutamate in addition to trehalose when under osmotic stress. Glutamate 26
accumulates rapidly after initiation of osmotic stress but
is then replaced by trehalose over a period of one or two
hours.
D.3.c. Osmoregulation in Rhizobia
There is one report showing an increase in trehalose
concentration in rhizobia under osmotic stress (Smith and
Smith, 1989). They surveyed several strains of R^. meliloti.
and found that four strains synthesized essentially no
trehalose when under osmotic stress and one accumulated a
significant amount of the sugar. They also were able to
identify a dipeptide that was present only in osmotically
stressed cells. Based on these observations it appears that
trehalose accumulation is not a universal mechanism of
osmotic stress tolerance among rhizobia.
Several other studies have examined rhizobia cultured
under osmotic stress, however, trehalose has not been
assayed (Hua et al.. 1982; Yap and Lim, 1983; Botsford,
1984; Bernard et al.. 1986). Some of these reports have
documented increased glutamate levels in salt stressed
rhizobia, and may have actually been observing a response
similar to that previously discussed in Ej. coli (section
D.3.b), where glutamate is synthesized initially and is
subsequently replaced by trehalose. Hua et al. (1982) found
that a Rhizobium species isolated from the Sonoran desert
accumulated glutamate in response to NaCl stress, as did R. meliloti (Botsford, 1984) and Rhizobium sp. UMKL 20, a 27 tropical fast-growing species (Yap and Lim, 1983). Upchurch and Elkan (1977) noted that some isolates of B^. ianonicum strains 61A76 and USDA 110 differed in colony morphology and that the more slimy, large colony types were more salt tolerant but less efficient at N2 fixation. D.3.d. Miscellaneous Organisms
There are a few scattered reports of increased trehalose accumulation in other organisms under osmotic stress. Euqlena gracilis (a protozoan) accumulates trehalose and mannitol in response to osmotic stress; the trehalose;mannitol ratio increases with increasing osmolarity of the growth medium (Dwyer, 1986). There is also one report of osmoregulation via trehalose accumulation in mosquitoes (Garrett and Bradley, 1987); larvae of Culex tarsalis in media of high osmolarity had increased levels of trehalose, proline and serine in their hemolymph. Meikle et al. (1988) documented increased intracellular trehalose concentration in stationary phase yeast cells under osmotic stress. In contrast, Bellinger and Larher (1988) found increases in glycerol and arabitol concentration in saline- grown yeasts, but no change in trehalose level. However, the latter study compared cultures at different points during exponential growth, not during stationary phase, so the results of these two studies are not necessarily conflicting. Van Laere et al. (1987) suggest that trehalose is degraded to glycerol in germinating spores of Phvcomvces 28 blakesleeanus as a mechanism of increasing internal osmolarity and consequently increasing H20 uptake.
D.4. Crvoprotection
D.4.a. Supercooling
Some cold blooded organisms accumulate compounds that function as "anti-freeze" and allow the organism to withstand temperatures below 0°C without freezing
(supercooling). A variety of beneficial effects of these protective agents have been documented, including enhancement of water binding, decreased ice content, maintenance of cell volume, reduction of water flux, and greater membrane stability (Baust, 1983). Examples of protective compounds are sorbitol, glycerol, glucose, glycogen, myo-inositol, erythritol, and trehalose (Block,
1984; Lee and Lewis, 1985; Baust and Lee, 1983).
The Antarctic midge Belaia antarctica does not freeze until daily average temperatures drop below -10°C (Baust and
Lee, 1983), and the cryoprotectant concentration in this organism increases as the average daily temperature falls in the autumn. Four species of terrestrial arthropods from
Antarctica can withstand temperatures of -20°C without freezing, and were shown to contain high levels of trehalose and other cryoprotectants (Block, 1984). Bakken (1985) documented supercooling in alpine beetles which had accumulated myo-inositol or trehalose in their hemolymph during winter acclimation. Hemolymph osmolality increased 29
in these beetles; in Calanthus melanocephalus maximum
trehalose concentration was 40-60mM.
The phenomenon of supercooling is not limited to
organisms adapted to extreme environments: gall fly (Eurpsta
solidaainis) larvae from the eastern US can withstand -25°C
due to accumulation of a three component solute profile
which includes trehalose (Lee and Lewis, 1985), and larvae
of the spotted cutworm Xestia c-niqrum accumulate trehalose
when subjected to low temperatures and short days (Goto et
al. 1986). Chrvsooerla carnea. the common lacewing,
decreases its supercooling point to -17°C while
overwintering, apparently by accumulating glycerol and
trehalose (Vannier, 1986). Increased levels of trehalose in
the sunflower moth, Homoeosoma electellum. correlated well
with decreases in glycogen content in cooled larvae (Rojas
et al.. 1989). This insect is common in the warmer climate
of Texas and farther south, but has a facultative ability to
enter diapause and consequently may be able to overwinter in more severe climates, apparently aided by accumulating
trehalose.
Hayakawa and Chino (1981) were able to induce trehalose
accumulation in silkworm larvae by cooling them for several weeks; this increase was accompanied by a decrease in glycogen content, and was observed repeatedly during warming and cooling cycles. Similarly, the lipid reserves of nematode eggs are converted to trehalose and glycogen when 30 they are chilled (Ash and Atkinson, 1983). However, in both these studies increased freezing tolerance was demonstrated but not decreased freezing point. It is possible that there are roles for trehalose in overwintering organisms other than supercooling, such as membrane stabilization or protection of macromolecules during desiccation or freezing.
It should be mentioned that a decrease in supercooling point is not correlated with increased trehalose concentration in some insects. In the army cabbageworm
Memestra brassicae, different aged larvae with differing trehalose concentrations had similar supercooling points
(Tsutsui et al.. 1988), and increased trehalose content in overwintering larvae of the soybean pod borer leauminovora glvcinivorella did not result in decreased supercooling points (Shimada et al.. 1984). However, in both these studies no other compounds were measured, so increased or decreased levels of some other osmoticum may have confounded interpretation of the results. Furthermore, Baust (1983) has suggested that optimal cooling rates vary widely between species and has also emphasized the importance of controlling thawing procedures; careless experimental procedures could also lead to conflicting results. Rojas et al. (1989) suggested that insect larvae that had been in diapause longer were able to withstand lower temperatures than those larvae recently entering diapause. 31
D.5. Freezing Preservation
Trehalose has been used as a protectant in the freezing and freeze-drying preservation of some organisms and macromolecules. Suspension cultures of carrot and tobacco cells were pretreated in 10% trehalose and then frozen with
40% trehalose as a cryoprotectant (Bhandal et al.. 1985).
Post-thaw viability was greater in trehalose-preserved samples, and was slightly greater in samples treated with both trehalose and dimethylsulfoxide. Lactobacillus bulaaricus and Saccharomvces cerevisiae cells also have been frozen in the presence of trehalose as a method of preservation of stock cultures (De Antoni et al.. 1989;
Coutinho et al.. 1988) , as have mouse embryos (Honadel and
Killian, 1988). Yeast strains that accumulate greater amounts of trehalose have been used in commercially produced raw dough breads (sold frozen, before proofing) because they withstand freezing better (Oda et al.. 1986). ATPase activity of mitochondrial inner membranes was preserved when mitochondria were frozen and thawed in the presence of trehalose (Tsvetkov et al.. 1985). Finally, isolated spinach thylakoid membranes have been found to be less susceptible to osmotic rupture if frozen in the presence of trehalose rather than sucrose (Hincha, 1989).
Some proteins, most notably phosphofructokinase, have been stabilized during freezing by the addition of trehalose
(Carpenter et al.. 1987? Carpenter et al.P 1986). Many 32 other organic solutes also have been shown to exert a protective effect on phosphofructokinase during freezing
(Carpenter and Crowe, 1988), and the inclusion of heavy metal ions along with trehalose or other osmotica may have a synergistic effect both on short term freezing (Carpenter et al.. 1986) and long term freezing and freeze-drying
(Carpenter et al.. 1988). Trehalose in combination with divalent cations has also been used to stabilize antibodies during freezing (Hazen et al.. 1988).
D.6. Desiccation Tolerance
D.6.a. Organisms
Trehalose has been detected at high concentrations (up to 20% dry weight) in anhydrobiotic organisms (those capable of withstanding extreme drying). Dry active bakers yeast, brine shrimp cysts, slime mold macrocysts, some nematodes
(both larvae and adults), and some fungal spores can exist completely dehydrated in a metabolic "limbo" for indefinite lengths of time, when they come into contact with water they rehydrate and become fully functional (see J.H. Crowe et al.. 1984a). This ability to dehydrate completely has been positively correlated with the presence of trehalose in many organisms (J.H. Crowe et al.. 1984a; Womersley and
Smith, 1981).
The interaction of trehalose with the phospholipid bilayer is considered to be important in maintenance of membrane integrity during desiccation (L.M. Crowe et al. 1984a). The mode of action is thought to be hydrogen bonding of the phosphate head groups with the hydroxyl hydrogens of trehalose, which effectively replaces
stabilizing water molecules normally associated with membranes (Crowe et al. 1985a, Rudolph et al. 1986). This may prevent the polar phospholipid head groups from
interacting with each other during dehydration by keeping them spaced far enough apart. It has been suggested that trehalose in dormant organisms helps protect cellular components by substituting for water molecules that normally stabilize membranes (J.H. Crowe et al. 1984a).
Spores of Streptomvces griseus normally contain approximately 25% trehalose (dry weight), but spores produced from cultures grown on glucose media have only about 1% trehalose and also have reduced viability after desiccation or heat stress (McBride and Ensign, 1987a), and
Martin et al. (1986) found similar results with S. antibioticus. The ability of yeast to withstand desiccation has also been correlated with internal trehalose concentration; cultures of Saccharomvces cerevisiae manipulated so that they had reduced levels of trehalose also have reduced ability to withstand drying (Keller et al.. 1982: Zikmanis et al.. 1985; Mackenzie et al.. 1988;
Gadd et al.. 1987). A minimal internal trehalose concentration of 120 mM was necessary to achieve a significant increase in dehydration resistance by Saccharomyces cerevisiae. and external trehalose was able to
increase desiccation resistance (Gadd et al.. 1987).
Coutinho et al. (1988) have shown increased viability of
yeast cells desiccated in the presence of trehalose,
relative to cells dried in phosphate buffer. The changes in
activity of the enzymes of trehalose metabolism during
desiccation and rehydration of baker's yeast cells has been documented (Marino et al.. 1989); as has trehalose retention
and distribution to the culture medium during drying and
rehydration (Zikmanis et al.. 1988).
D.6.b. Membranes
Numerous studies have been documented the stabilizing
effect of trehalose on dehydrated membranes, both artificial
and those isolated from living organisms (see Crowe et al..
1987a; Crowe et al.. 1988; Chandrasekhar and Gaber, 1988).
Many other sugars have similar effects, but in general trehalose is the best stabilizing agent known for dry membranes and phospholipid bilayers.
The presence of trehalose in artificial membranes
significantly depressed the transition (melting) temperature
(J.H. Crowe et al.. 1984b, L.M. Crowe, 1985b), and decreased the degree of fusion of vesicles during freeze-drying
(Rudolph and Crowe, 1985; Womersley et al.. 1986), apparently by preventing phase separation (Crowe et al..
1986). Lee et al. (1986) suggest that trehalose in contact with artificial membranes forms a novel liquid crystalline- like phase that is responsible for the stabilizing effects observed with trehalose. These experimental artificial membranes are formed by lipids that are derivatives of phosphatidylcholine, and it has been shown that they can be completely dehydrated and retain their integrity (Crowe et al.. 1987b). The functional integrity of biological membranes during long-term storage increased in the presence of high levels of trehalose, low 02 tension, low humidity and darkness (Mouradian et al.. 1985). The potential application of trehalose-stabilized artificial vesicles in the administration of drugs has been investigated (Madden et al. 1985). The stability of some dry liposomes can be affected by the size, surface charge, and the dry-mass ratio between the amount of sugar and the amount of lipid (Crowe and Crowe, 1988).
Mitochondrial membranes have been lyophilized in the presence of trehalose and rehydrated with marginal loss of enzyme activity (Tsvetkov et al.. 1985), and lobster sarcoplasmic reticulum vesicles have also been successfully freeze dried and rehydrated after trehalose treatment without substantial loss of transport activity (Rudolph and
Crowe, 1985).
D.7. Heat Tolerance
There are a few reports showing a correlation between trehalose concentration and heat resistance in some microorganisms. Hecker and Sussman (1973b) found that trehalase extracted from mycelia and spores of Neurosoora was more stable when heated in solutions containing trehalose than when heated in other solutions. Other sugars and polyols were not effective, and invertase was not protected by trehalose (Yu et al.. 1967), suggesting the mode of protection was by trehalose binding to the active site. McBride and Ensign (1987a) manipulated the internal trehalose content of spores of Streptomvces ariseus and showed that those with higher trehalose levels were more resistant to heating. Hottiger et al. (1987a, b) found increased trehalose accumulation (and heat tolerance) in Saccharomyces cerevisiae incubated at high temperatures, apparently due to increased activity of trehalose-6- phosphate synthase. However, these cells also had increased activity of trehalase, the enzyme necessary for trehalose breakdown, which apparently lead to a futile cycle of trehalose synthesis and breakdown. In a later study they found the same stresses that induce synthesis of heat shock proteins also induced trehalose synthesis in Saccharomyces
(Hottinger et al.. 1989). Finally, Dictvostelium discoideum spores with increased internal trehalose content had higher heat tolerance than spores with low trehalose levels
(Emyantioff and Wright, 1979).
D.8. Trehalose as a Carbon Sink
Trehalose is not synthesized by higher plants, although in some parasitic and symbiotic associations (notably ectomycorrhizae) trehalose is synthesized from photosynthate
(glucose or sucrose) received by the fungus and thus forms a carbon "sink” (see Smith et al. 1969, Lewis and Harley 1965;
Martin et al.. 1987). A continuous flow of photosynthate to the fungus is maintained because feed-back inhibition mechanisms are not activated as they would be if a plant-synthesized compound, such as sucrose, was stored by the fungus.
It is interesting that many N2-fixing root nodule symbionts synthesize trehalose, both in culture and in the symbiotic state (Streeter, 1985; Lopez et al.. 1983, 1984).
Lichens are a similar type of symbiosis, because photosynthate and fixed atmospheric N2 are transported between partners. However, lichens are an association between a fungus and one or more algae, and the algae are responsible for both N2 fixation and photosynthesis (see
Smith 1980; Englund 1977). This carbon transported is usually in the form of glucose, but can be sugar alcohols.
The flow of carbon is larger than necessary to support the very low growth rate of lichen-associated fungi, and is proposed to be necessary for maintenance of a high level of soluble carbohydrate (Smith 1980). This would help protect the organism from environmental extremes (and lichens are commonly found in extremely harsh environments), and also would require the synthesis of a "sink" compound. The composition of soluble carbohydrates in lichens is not well 38 documented, but trehalose is present at least in some species (Iacomini et al.. 1987).
E. Metabolism of Trehalose
E.1. Synthesis
E.1.a Biosynthetic Pathway
The pathway of trehalose synthesis was first elucidated from yeast extracts (Cabib and Leloir, 1958); it is a relatively simple two-step process: uridine diphosphoglucose
(UDP) and glucose-6-phosphate combine to yield trehalose-6-phosphate (T-6-P) and UDP, then T-6-P is cleaved to form trehalose and inorganic phosphate (Fig. 1) (see Elbein 1974). The enzymes involved are T-6-P synthetase (EC
2.4.1.15) and T-6-P phosphatase (EC 3.1.2.12) respectively.
Relatively few reports on the isolation and purification of these two enzymes have been published: T-6-P synthetase was first isolated by Leloir and Cabib (1953) and since then it has been partially purified and characterized from the slime mold Dictvostelium discoideum (Killick, 1979) and from Mycobacterium smeqmatis (Lapp et al.. 1971). Both
T-6-P synthetase and T-6-P phosphatase have been purified from Saccharomyces cerevisiae (Vandercammen et al.. 1989).
There is some evidence that the synthetase and the phosphatase form an enzyme complex in yeast (Vandercammen et al.. 1989? Panek and Panek, 1989). The synthetase requires
Mg++ for activity, and is usually thought to utilize uridine 39 diphosphoglucose (see Fig. 1.), but Paschoalin et al. (1989) reported a trehalose synthase from yeast that was adenine diphosphoglucose dependent. This enzyme also required Mg++.
Piper and Lockheart (1988) isolated a mutant of
Saccharomvces cerevisiae that was deficient in T-6-P phosphatase. This strain accumulated T-6-P at low temperatures but was unable to grow at 37°C, apparently because of some toxic effect of increased pools of T-6-P.
GLUCOSE GLUCOSE + + GLUC0SE-6-P fi-GLUCOSE-l-P
UT
GLUCOSE-I- TREHAL0SE-6-P TREHALOSE
GLUCOSE
© UDPG PYROPHOSPHCiRYLASE GLUCOSE © TREHALOSE-6*PHOSPHATE SYNTHETASE © TREHALOSE-6-PHOSPHATE PHOSPHATASE
© PHOSPHOTREHALASE
© TREHALASE
© TREHALOSE PHOSPHORYLASE
Fig. 1. Known reactions of trehalose metabolism. From
Salminen and Streeter (1986).
E.l.b Regulation of Trehalose Synthesis
The synthesis of trehalose may be regulated by end-product inhibition from either T-6-P or trehalose itself
(see Elbein 1974). In Mycobacterium sp. trehalose synthesis may be controlled indirectly by regulation of sugar-6-P and nucleotide diphosphate levels (Roth and Sussman, 1968).
Panek et al. (1987) found that T-6-P synthetase from yeast was regulated by phosphorylation; the dephosphorylated form was inactive and could be generated by Sephadex G-25 filtration of the active form (Panek and Panek, 1989).
Similarly, phosphofructokinase activity may be less active in insects that accumulate trehalose instead of glycerol, causing glycogen breakdown products to be preferentially incorporated into trehalose rather than glycerol (Hayakawa and Chino, 1982). Killick and Wright (1972) found an inactivated form of T-6-P synthetase in differentiating slime molds that was activated during later development.
Literature detailing various external stimuli required to induce trehalose synthesis in different organisms was discussed in section III. There is some evidence of a link between trehalose synthesis and maltose utilization in
Saccharomvces cerevisiae (Panek et al.. 1980; Oliveira et al.. 1986; Paschoalin et al.. 1986).
E.2. Trehalose Catabolism
E.2.a Enzymes
The chief enzyme responsible for the degradation of trehalose is trehalase (EC 3.2.1.28, a,a-trehalose-l-D- glucohydrolase), first demonstrated by Bourquelot (1893).
The products of hydrolysis have been shown to be one a-glucose and one /3-glucose molecule (Clifford, 1979; Hehre et al.. 1977; Defaye et al.. 1983; Nakano et al., 1989), although other catalytic reactions of trehalase have been documented under certain (non-physiological) conditions
(Hehre et al.. 1982; Kasumi et al.. 1986? Weiser and
Lehmann, 1988). The specificities of some different trehalases have been reported (Labat-Robert et al.. 1978;
Fleischmacher et al.. 1980), and a comparison of different trehalases has been published (Alabran et al.. 1983). The latter authors concluded trehalase was more similar to a - D -
(1-+4)-glucan glucohydrolases (EC 3.2.1.3) than it is to the other a-glucosidases. Killick (1980) documented some very sensitive fluorometric assays for trehalase activity.
Trehalose phosphorylase (EC 2.4.1.1) also catalyses the breakdown of trehalose, although in this case the products are a-glucose and /3-glucose-l-phosphate (Marechal and
Belocopitow, 1972). The latter enzyme is apparently much more rare in nature than trehalase, but has also been detected at low levels of activity in Bradvrhizobium bacteroids (Salminen and Streeter, 1986). A similar trehalose phosphorylase from the basidiomycete fungus
Flammulina velutioes has been described, however, the products of trehalose phosphorolysis are glucose and a- glucose-1-phosphate (rather than glucose and /3-glucose-l- phosphate) (Kitamoto et al.. 1988). The specific activity of the phosphorylase was about double that of hydrolytic trehalase activity in mycelia, and fruiting bodies had approximately 10-fold greater phosphorylase specific 42 activity than trehalase activity. The phosphorylase also had the ability to synthesize trehalose from the catabolic products.
Another enzyme capable of breaking down trehalose is phosphotrehalase, which catalyses the breakdown of T-6-P to glucose and a-glucose-l-phosphate. This enzyme was originally found in Bacillus popilliae (Bhumiratana et al..
1974) and the only other report of its occurrence is in extracts of soybean bacteroids (Salminen and Streeter,
1986). The same products as those from phosphotrehalase- mediated T-6-P hydrolysis were detected in E_s_ coli and
Salmonella tvphimurium extracts and were attributed to a
"trehalose hydrolase" (Marechal, 1984).
E.2.b Distribution of Trehalase
Trehalase has been isolated from many animals, including both those that can make trehalose and those that cannot. The presence of this enzyme in non-trehalose synthesizing organisms is probably a defense mechanism for protection against trehalose toxicity in the event of consumption or uptake (via symbiotic or parasitic associations) of trehalose. Intestinal forms of this enzyme have been isolated from several non-trehalose synthesizing animals and probably function in absorption (Azuma and
Yamashita 1985, Ogiso et al. 1985, Yokota et al. 1986).
There are reports of significant trehalase activity in some higher plants (Fleischmacher et al.f 1980; Gussin et al.. 43
1969; Gussin and McCormack, 1970; Hisajima et al.. 1981), however, plants low in trehalase activity produce a black phenolic substance and die quickly if trehalose is added to their nutrient media (Veluthambi et al.. 1981, 1982 a, b).
There are a few interesting facts concerning trehalase in humans. It has been suggested that trehalose was more important than sucrose in the diet of ancient man, and intestinal trehalase activity is common in humans (see
Semenza, 1968). Trehalase deficiency has been documented in the population of Greenland, and is associated with intestinal discomfort after consumption of trehalose- containing foods similar to that seen in (more commonly occurring) lactose-intolerant individuals (Gudman-Hoyer et al.. 1988). These authors felt avoiding trehalose would only require elimination of mushrooms from the diets of sensitive people, apparently they were not aware of the significant presence of trehalose in wine and sherry (Olano,
1982; Santa-Maria et al.. 1985)! Elevated levels of plasma trehalase have been correlated with an increased chance of developing diabetes mellitus (Eze, 1988), and low levels of trehalase activity in human amniotic fluid is considered indicative of cystic fibrosis in the fetus (Szabo et al..
1985).
E.2.c. Trehalase purification
Trehalase has been purified from many diverse sources: rabbit intestine and kidney (Yokota et al. 1986, Takesue et al. 1986? Nakano and Sacktor, 1985), pig kidney (Yoneyama,
1987), rat intestine (Riby and Galand 1985), Neurospora
(Hill and Sussman, 1963? White et al. 1985), Frankia (Lopez and Torrey, 1985), yeast (Kelly and Catley, 1976? Panek and
Souza, 1964? Londesborough and Varimo, 1984? Dellamora-Ortiz et al.. 1986? Mittenbiihler and Holzer, 1988? App and Holzer,
1989), silkworms (Sumida and Yamashita 1983, Azuma and
Yamashita 1985), cockroaches (Osigo et al. 1985), grasshoppers (Teo and Heng, 1987) , mycobacteria (Patterson et al.. 1972), slime molds (Killick 1983a, 1985), and soil
(Smith and Rodriguez-Kabana, 1982). The results of these studies vary somewhat but in general the enzyme is reported to be a glycoprotein, has an average molecular weight of
100,000 daltons, and usually has pH optimum of about 5.5, although some sources have trehalases with two pH optima, approximately 4.0 and 6.5, and still others are near neutrality. Some organisms have more than one type of trehalase (Arguelles and Gacto, 1985? Takesue et al.. 1989), which may vary in activation properties and membrane association. Reports of subunit composition of the functional enzyme vary from one polypeptide chain to four.
Isozymes have been documented in developing slime molds
(Killick, 1983b), rabbit kidney (Nakano and Sactor, 1985), and silkworm (Bombvx mori) intestines, where they differ in solubility because one form has an "anchor" segment binding it to a membrane (Takesue et al.. 1989). Competitive 45 inhibition studies of rabbit renal trehalase suggest there are two binding sites for trehalose at the active site
(Nakano and Sactor, 1984). A variety of methods are available for assay of trehalase (Killick, 1980; Alabran et al.. 1983).
Many reports of trehalase inhibitors are in the literature. Trehalases from various sources are inhibited by the following; Tris buffer, p-aminophenylglucoside, sucrose, and maltose (Riby and Galand, 1985), f3- methylglucoside, phlorizin, and phloritin (Nakano and
Sacktor, 1984), validoxylamines (Kameda et al.. 1987), p-nitrophenyl-£-D-glucoside (Sumida and Yamashita, 1983) ,
1,5-dideoxy-l, 5-imino-D-mannitol (Evans et al.. 1983) and dioxane and 5-gluconolactone (Terra et al.. 1983).
Non-competitive inhibition is reported to occur in the presence of HgCl2, Tris buffer and ATP (Panek, 1969; Sumida and Yamashita; 1983; Nakano and Sactor; 1984). A peptide inhibitor of trehalase was isolated from the hemolymph of
Triatoma vitticeps (Conter and Veiga, 1984), and a protein inhibitor of trehalase was isolated from Periplaneta americana hemolymph (Hayakawa et al.. 1989). The trehalase inhibitor trehaloseamine is secreted by Streptococcus spp.
(Umezawa et al.. 1967).
E.2.d. Localization of Trehalase
Two reports have used immunochemistry to localize trehalase. Immunofluorescent and enzyme-labeled antibody techniques were used to identify trehalase in the intestinal epithelial cells of rabbit kidney proximal tubule brush borders (Nakano, 1982), and similar results were obtained by
Azuma and Yamashita (1986) with silkworm midgut, using peroxidase-linked antibodies to localize trehalase in the plasma membrane on the basal surface of the epithelium.
They verified the membrane location of the enzyme with sucrose density gradient centrifugation and analysis of marker enzymes along with trehalase. Both of these studies suggest the role of absorption for trehalase, although the latter indicated that trehalase could also aid in uptake of trehalose from insect hemolymph during starvation.
In yeast, activated trehalase (see section E.2.e) is found in the vacuoles, but inactive trehalase and trehalose are located in the cytosol (Londesborough and Varimo, 1984;
Wiemken and Schellenberg, 1982, Keller et al.. 1982; Harris and Cotter, 1988). In ascospores of Neurospora and
Schizosaccharomvces trehalase is associated with the cell wall (Inoue and Shimoda, 1981a; Chang and Trevithick, 1972;
Hecker and Sussman, 1973a), and Ram et al. (1984) found the same location for trehalase in budding yeast cells. Another study indicated a peripheral location in yeast cells, although not necessarily in the cell wall (Mandels, 1981).
Van Assche et al. (1978) and Arnold (1979) both found two locations of trehalase, in Phycomyces spores and cells of the yeast Torulopsis qlabrata. respectively. They detected 47 cytosolic and periplasmic trehalases, and suggested the internal enzyme was active during breakdown of endogenous trehalose, and that the periplasmic trehalase was used for catabolism of exogenous trehalose.
Trehalase is associated with the soluble fraction in extracts of Dictvostelium. Pullularia. and ascospores of
Pichia (Killick 1983a, Merdinger et al. 1971, Thevelein
1984a). However, Yokota et al. (1986) documented an
"anchor" segment attaching the enzyme to membranes in mammalian kidneys, so it is possible that these studies inadvertently cleaved this anchor segment and incorrectly concluded that the enzyme was soluble. Killick (1985) found that trehalase was compartmentalized in slime molds, and that the locality shifted with development. In spores nearly all the enzyme was soluble, while in stalk cells up to 50% insoluble. An extracellular trehalase has also been documented in slime molds (Killick 1983a).
E.2,e. Regulation of Trehalase
In natural systems there appears to be a variety of mechanisms for the control of trehalase activity, including activation, compartmentalization, cleavage of proenzymes, subunit polymerization, and regulation of transcription.
Considerable effort has been directed at elucidating the mechanism of cyclic adenosine monophosphate (cAMP) activation of trehalase in fungi. The addition of glucose to yeast cell cultures stimulates cAMP production (via 48 adenylate cyclase), which activates a kinase that phosphorylates the inactive form of trehalase; the phosphorylated trehalase becomes active (Ortiz et al.. 1983,
Uno et al.. 1983, van der Plaat, 1974; Thevelein, 1988).
This phenomenon can make purification of this enzyme complicated because accidental dephosphorylation of the enzyme will appear to be enzyme loss. Recently a technique to avoid this problem has been published (Araujo et al..
1989).
This cAMP mediated activation of trehalase can be induced by glucose (see Thevelein, 1984a), heat shock
(Thevelein, 1984b), and agents causing internal acidification (Valle et al.. 1986). Reports of activation by membrane depolarization (Thevelein, 1984c) have not distinguished between activation by internal acidification resulting from membrane depolarizing agents and true activation by depolarization. Valle et al. (1986) investigated the mode of cAMP activation to determine if acidification or depolarization or both effects were needed to induce trehalase activation. Depolarization of membranes did not result in trehalase activation unless acidification also occurred, so the authors concluded that internal acidification was the actual mechanism of activation.
Arguelles and Gacto (1985) found evidence of both regulatory (cAMP activated) and nonregulatory trehalases in
Candida utilis. Nonregulatory trehalases are thought to be 49 controlled by compartmentalization of trehalase and trehalose. Hecker and Sussman (1973a) first proposed the concept of physical separation of trehalase and trehalose in
Neurospora. and this has also been suggested to occur in
Mvrothecium (Mandels, 1981), Dictvostelium (Jackson et al..
1982), dormant Pilobolus lonaipjpes spores (Bourret et al..
1989), and Bradvrhizobium bacteroids (Salminen and Streeter,
1986). It is thought that membrane integrity is lost under certain conditions and trehalose is allowed to come into contact with the enzyme, although the specifics of this mechanism are not clear (see Thevelein, 1984a). A different mechanism of trehalase regulation was proposed by Harris and
Cotter (1987); they obtained evidence that a vacuolar proenzyme of trehalase in Saccharomvces cerevisiae is cleaved by protease A for activation.
The effect of changes in the internal pH of brine shrimp embryos has also been investigated (Hand and
Carpenter, 1986). The internal pH of dormant and hatching brine shrimp embryos was manipulated with C02 exposure, and the effects on trehalase activity monitored. When the pH was lowered (due to increased respiration at hatching or to incubation with elevated C02 concentration), the enzyme depolymerized and was much less active than at pH 7.4 or higher. When isolated and subjected to gel electrophoresis the degree of subunit polymerization and specific activity could be changed by changing the pH of the gel. The enzyme was not polymerized at pH 8.3 and showed high catalytic activity, but if electrophoresed at pH 7.0 the enzyme polymerized and lost much of the activity seen at pH 8.3.
This mode of enzyme regulation is termed hysteresis, and the two enzyme states were found to be differently sensitive to
ATP and other inhibitors. The ability to regulate enzyme activity by changing the internal pH was suggested to be a possible means of survival in the rapidly changing environment encountered by hatching brine shrimp. A decrease in respiration (due to adverse conditions) would lower the pH sufficiently to slow the breakdown of trehalose and arrest the hatching process.
E.2.f. Effects of Fructose-2.6-Bisphosphate
As mentioned earlier, trehalose phosphorylase cleaves trehalose to j8-D-glucosyl phosphate and glucose, and this enzyme has been found in Eualena gracilis and Bradvrhizobium bacteroids. In Euqlena fructose-2,6-bisphosphate (F26BP) has an inhibitory effect on trehalose phosphorylase
(Miyatake et al.. 1984). Fructose-1,6-bisphosphatase is also inhibited by F26BP, but phosphofructokinase is stimulated. This combination of regulatory actions is suggested to be a means of control of trehalose and paramylon (a glycogen-like carbohydrate) synthesis. The mechanism of regulation of trehalose phosphorylase in bacteroids is not known. Panek et al. (1989) investigated the effect of F26BP on fructose 1,6 bisphosphatase and phosphoglucose isomerase from yeast, in an attempt to demonstrate a regulatory role for F26BP in yeast. They concluded that cAMP activation was responsible for regulation of trehalose metabolic enzymes, and that F26BP was not involved in regulation of these enzymes. Van Laere et al. (1983) and Vandercammen et al.
(1990) have examined the metabolism of F26BP during germination of Phvcomvces blakesleeanus and Blastocladiella emersonii. respectively. The latter study found some distinct differences in carbohydrate metabolism in
Blastocladiella cells (class oomycetes) relative to higher fungi. Reserve carbohydrates like glycogen and trehalose were degraded rather than synthesized during sporulation, however, the mechanism of regulation of trehalose metabolism was not clear because cAMP levels remained constant.
Because the effects of F26BP were similar to that reported in Euglena (Miyatake et al.. 1984), i.e. stimulation of phosphofructokinase and inhibition of fructose bisphosphatase, it appears possible that F26BP has a similar effect in Blastocladiella.
E.3. Genes Related To Trehalose Metabolism
A few reports have been published concerning the genes for trehalose metabolism in different organisms. St. Pierre
(1968) isolated mutants of Salmonella typhimurium defective in trehalose utilization and mapped the chromosomal lesion 52 with respect to other known genes, and Becerra de Lares et al. (1977), White et al. (1985) and Burton and La Spada
(1986) have done the same with E_s_ coli. Neurospora crassa. and Drosophila melanoaaster. respectively. The gene for an osmoregulated periplasmic trehalase from ILt. coli has been cloned and sequenced (Boos et al.. 1987; Gutierrez et al..
1989) . Two Ei. coli genes involved with osmotic induction of trehalose synthesis have been cloned, but it is not clear whether they represent.structural or regulatory genes
(Giaever et al. . 1988).
F. Summary of Trehalose Occurrence and Metabolism in
Rhizobia
Trehalose has been identified in every legume symbiosis tested, and is produced by the bacteroids (Reibach and
Streeter, 1983; Streeter, 1985). Different strains of the same species of rhizobia accumulate different amounts of trehalose, and in general the amount found in cultured bradyrhizobia and rhizobia is proportional to the amount found in bacteroids formed by the same strain. Trehalose does not appear in the nodules until the onset of nitrogen fixation (Streeter and Bosler, 1976; Streeter, 1980), it then increases in concentration until the nodules become senescent, and then decreases (Streeter, 1981). A curious aspect of the trehalose found in legume nodules is the large proportion of (bacteroid-synthesized) trehalose found in the 53 plant fraction; however, the proportion of total nodule trehalose in the bacteroids increases as the nodule ages
(Streeter, 1985? 1987).
Bradyrhizobia (slow-growing rhizobia) accumulate trehalose in culture, but cannot use it for a carbon source
(Stowers, 1985), and uptake by isolated bacteroids is small relative to uptake of succinate (Salminen and Streeter,
1987). This is a major difference in the cultural characteristics of the fast and slow growing rhizobia, as
Rhizobium spp. (fast-growing rhizobia) can use trehalose as a carbon source, including the (fast-growing) soybean- infecting species fredii. Different strains and species of rhizobia accumulate different amounts of trehalose in culture (Streeter, 1985).
The occurrence and activity of trehalase in soybean bacteroids and nodules has been analyzed. Trehalase was found in the nodule cytosol (plant tissue) but not in the bacteroids in two studies (Streeter, 1982; Mellor, 1988), and in both the cytosol and bacteroid fractions in a third report (Salminen and Streeter, 1986). Differences in centrifugation protocols are probably responsible for the discrepancies. It apparently is a coincidence that soybean trehalase activity has the same pH optimum (approximately
6.6) as rhizobial trehalase (Streeter, 1982; Hisajima et al.. 1981; Bassarab et al.. 1984)? this has contributed to the confusion concerning bacteroid derived trehalase. Other enzymes of trehalose metabolism (UDPGlc phosphorylase, T-6-P phosphatase, trehalose phosphorylase and phosphotrehalase) have been localized in the bacteroids and cytosol, but T-6-P synthetase, the key synthetic enzyme, was found only in the bacteroid fraction (Salminen and Streeter, 1986). CHAPTER II
INCREASED ACCUMULATION OF TREHALOSE IN SOME RHIZOBIA
WHEN CULTURED UNDER OSMOTIC STRESS
Introduction
Several types of micro-organisms are known to increase their internal solute concentration in response to hyperosmotic stress, as discussed in the literature review
(section IV.C). The increased internal solute concentration means the internal osmotic pressure has increased, allowing these organisms to prevent dehydration and consequently maintain turgor. The compounds that accumulate under osmotic stress are generally sugars or other low molecular weight compounds. Trehalose is one of these compounds, frequently increasing in concentration in both cyanobacteria and Ei coli when under osmotic stress (Larsen et al.. 1987;
Giaever et al. , 1988; Rod et al.. 1988; Reed et al.. 1986).
Escherichia coli is a gram negative bacterium, as are rhizobia and bradyrhizobia. Cyanobacteria frequently fix atmospheric N2, another distinctive characteristic of the rhizobia and bradyrhizobia. The combination of these two facts led to the suggestion that osmotic stress might play a role in induction of trehalose accumulation in rhizobia
55 56 and/or bradyrhizobia, either in the free-living state or in the nodule.
It is not clear whether the nodule environment is osmotically stressful for the bacteroids, however, some estimates of nodule solute concentration are higher than many common rhizobial media. For example, YEMG (Vincent,
1970; Appendix H), a commonly used rhizobial medium, has an osmotic potential of about 100 mOsmol (Hoelzle, unpublished data), but well-watered soybean nodules have been documented with water potentials of 150 mOsmol (Fellows et al.. 1987) and cowpea (Vigna unouiculata) nodules at about 220 mOsmol
(Khanna-Chopra et al.. 1984). These same two studies measured water potentials in water-stressed nodules of approximately 580 mOsmol, considerably more than YEMG.
Furthermore, the drying conditions of soils certainly must subject free-living rhizobia to far more stressful osmotic potentials that those found in culture media or in the nodule environment.
Because of the similarity to rhizobia of some organisms that accumulate trehalose under hyperosmotic stress, and because it appears that both the nodule environment and the soil may be osmotically stressful for rhizobia, a survey of rhizobia and bradyrhizobia was conducted to determine if there was a correlation between trehalose accumulation and salt (osmotic) stress. 57 Materials and Methods
The arctic rhizobial isolate was obtained from Dr. H.M.
Schulman, Lady Davis Institute, Montreal, Canada. All other
rhizobia were provided by Dr. H. Keyser, USDA Nitrogen
Fixation Laboratory, Beltsville, MD. Rhizobia were grown at
29°C in 50 ml of liquid culture medium in 125 ml Erlenmeyer
flasks on a rotary shaker at 125 RPM.
The succinate medium was a modification of the defined succinate medium of Lepo and McAllister (1983) (Appendix H).
Malate medium was the same as succinate medium with 3.0 g sodium malate/1 substituted for sodium succinate. The arabinose/mannitol medium and the micronutrients were as described by Manhart and Wong (1979) (Appendix H). Vitamins were filter sterilized before addition to autoclaved media in the following quantities (in mg/1): thiamin HC1 (0.4), calcium pantothenate (0.5) and biotin (0.1). Excess salts were added as indicated in the text. All rhizobia and bradyrhizobia were initially cultured in YEMG (Vincent,
1970; Appendix H) and a small amount of this starter culture used to inoculate experimental flasks.
Cells were harvested by centrifugation at 12,000 x g for 5 minutes. The pellet was resuspended in 10 ml H20 and an aliquot taken for dry weight measurement, and the remainder re-centrifuged for 5 minutes at 12,000 x g. This pellet was suspended in 5 ml 80% ethanol and a few drops of chloroform, and refrigerated for at least 3 hours. After 58 refrigeration the extracts were centrifuged at 12,000 x g for 10 minutes and the supernatant fraction decanted into 10 ml drying bottles. The liquid was evaporated under an air stream, the extraction process repeated two more times, and the extracts from each sample were pooled. The dried extracts were dissolved in 2 ml H20 and 1 ml chloroform.
After the phases had completely separated an aliquot of the aqueous fraction was taken and dried down in a 4 ml glass vial.
The samples were then derivatized using reagents from
Pierce Chemicals, Rockford, IL (Appendix A). The oxime- trimethylsilyl derivatives were analyzed using a Varian 3700 gas chromatograph fitted with a glass column packed with 3%
OV-17 (Appendix A). Standards were run daily.
Results
Rhizobium leauminosarum bv. phaseoli USDA 2667 was grown in succinate medium with 0, 50, 100, or 150 mM excess
NaCl. The water potentials of these media ranged from about
125 mOsmol (0 excess salt) to approximately 350 mOsmol (150 mM excess salt), which is in the range of osmotic potentials previously measured in legume nodules (Khanna-Chopra et al.f
1984; Fellows et al.. 1987). One flask of each treatment was harvested at five different times after inoculation
(Fig. 2). Trehalose concentration increased with increasing osmolarity up to 100 mM excess NaCl. Salt additions greater than 100 mM led to decreased growth rate and decreased trehalose concentration in the cells (Fig. 2; data for 250 mM excess NaCl not shown). At stationary phase
(approximately 70 hours after inoculation) the internal reserves of trehalose rapidly disappeared from the cells.
This experiment was conducted only once, however, the increased trehalose accumulation response of R. leauminosarum bv. phaseoli USDA 2667 to 100 mM excess NaCl was observed numerous times during the course of other experiments (see Chapter IV).
Cells of R±. 1^ bv. phaseoli USDA 2667 harvested from either malate or arabinose/mannitol media with 100 mM extra
NaCl demonstrated the same response to salt stress as those grown in succinate medium (Table 1). Furthermore, substitution of 100 mM KC1 for 100 mM NaCl in the succinate medium also did not alter the response to osmotic stress in this strain (Table 1). Similar results were observed at least one additional time with this rhizobial strain cultured in each of these media.
A total of seventeen isolates of rhizobia were screened for increased trehalose accumulation when cultured with excess NaCl (Table 2). Of the nine types of R. 1. bv. phaseoli tested, three (including strain USDA 2667 used for the experiments in Fig. 2 and Table 1) had trehalose concentrations under salt stress that were at least double that observed when they were cultured with no salt stress. 60
The remaining six iL*. bv. phaseoli strains either showed no difference in trehalose concentration or did not grow at all when under salt stress, even if the salt concentration was reduced to 25 mM over that already in the medium (Table
2).
Among the other strains and species tested, only R. loti USDA 3071, the symbiont for Lotus corniculatus. had increased trehalose concentration under osmotic stress.
Several other species tested did not show any change in trehalose content under osmotic stress, but were nevertheless capable of growing quite well. Two strains of
Bradvrhizobium iaponicum did not grow under salt stress
(Table 2). All strains indicated in Table 2 with increased trehalose accumulation under osmotic stress were tested at least twice; some other strains were tested twice.
Discussion
Rod et al. (1988) described a similar pattern of trehalose accumulation in EL. coli under osmotic stress; some strains accumulated more trehalose under osmotic stress than when not stressed, but in other strains the trehalose pool size was not affected by osmotic conditions. However, their system differed from that found in rhizobia in several ways;
1) All rhizobia make some trehalose regardless of the osmotic conditions (Streeter, 1985), but Ejs. coli synthesize it only when under osmotic stress (Rod et al.. 1988; Larsen et al.. 1987), 2) Some rhizobial strains tested in this study grew well under osmotic stress even though they did not accumulate more trehalose; E^. coli strains that are unable to synthesize trehalose are osmo-sensitive (Larsen et al.. 1987; Giaever et al.. 1988), 3) Rod et al. (1988) were able to show a close genetic relatedness of coli strains lacking trehalose synthetic capability and also among those that are osmo-tolerant; the various rhizobia examined here are of very diverse origins. Even the three R. 1. bv. phaseoli strains that did accumulate more trehalose under osmotic stress are probably quite unrelated; the origin of only two are known, strain USDA 2667 was isolated in
Washington and strain USDA 2670 was isolated from Wisconsin
(Keyser and Griffin, 1987). While it is possible that they could be related if they were introduced species, they were isolated in 1948 (Keyser and Griffin, 1987), before extensive use of Rhizobium spp. inoculants was common. Any
Rhizobium which nodulates Phaseolus vulgaris is considered a member of the R^. leauminosarum bv. phaseoli taxonomic group.
This group has previously been documented to be genetically diverse (Pinero et al., 1988), and this diversity can now be extended to differences in osmotic stress response.
Smith and Smith (1989) documented a phenomenon in rhizobia similar to that described here; they found increased trehalose accumulation in some rhizobia when grown under osmotic stress but not in other strains. However, they identified a dipeptide that was present during osmotic stress in the strains of rhizobia that did not alter their trehalose pool size (when under osmotic stress). Other studies (Hua et al.. 1982; Yap and Lim, 1983; Botsford,
1984) have found increased glutamate accumulation in some osmotically stressed rhizobia. One of these mechanisms
(glutamate or dipeptide accumulation) may be the mode of salt tolerance by the rhizobia in this study that grew well under salt stress but did not show increased trehalose pool size.
Accumulation of trehalose does not appear to be a universal mechanism among rhizobia for adaptation to osmotic stress. Some strains (e.g. Rj.lj.bv. phaseoli USDA 2667) accumulated high trehalose concentrations in response to salt stress and the sugar may play a protective role in these strains. Some of the other strains of rhizobia tested apparently have alternate methods of surviving osmotically stressful environmental conditions. Differences in response to salt stress may be useful in determining genetic relationships among some rhizobia. S3
Figure 2. Effect of various concentrations of excess NaCl on trehalose accumulation in Rhizobium lequminosarum bv. phaseoli USDA 2667. Each point represents the trehalose concentration detected in one culture. ♦ , 0 excess NaCl ;
T / 50 mM excess NaCl; # , 100 mM excess NaCl;
B , 150 mM excess NaCl ^ 20 (1) O +-> si O) *
-b
20 40 60 80 100 hours after inoculation
Figure 2
o> 4^ 65
TABLE 1. Trehalose accumulation in
R. leauminosarum bv. ohaseoli USDA 2667 cultured
in different media and with different salts.
Medium/ mM mg Dry Weight Cells/ mg Trehalose/ Salt added Salt ml Culture1'2 g Dry Weight Cells
Succinate/ 0 1.2 4.8 NaCl 100 1.2 14.9
Succinate/ 0 0.7 2.4 KC1 100 1.1 14.2
Malate/ 0 1.4 3.9 NaCl 100 1.4 8.3
Arabinose + 0 0.7 1.0 Mannitol/ NaCl 100 1.0 10.9
1. Cultures were harvested at mid-log phase of growth.
2. Data are representative of at least two experiments. TABLE 2. Trehalose accumulation by various rhizobia
grown in succinate medium with 100 mM excess NaCl1.
Soecies Resoonse Rhizobium leauminosarum bv. phaseoli USDA 2667
R. 1. bv. ohaseoli USDA 2669 +
R. 1. bv. ohaseoli USDA 2670 +
R. 1. bv. Dhaseoli USDA 2674 _3
R. 1. bv. ohaseoli USDA 2668 -
R. 1. bv. ohaseoli USDA 2681 -
R. 1. bv. Dhaseoli USDA 2672 -
R. 1. bv. ohaseoli USDA 3319 NG4
R. 1. bv. ohaseoli USDA 3251 NG
R. 1. bv. trifolii USDA 2066 -
R. 1. bv. viceae USDA 2370 -
R. loti USDA 3071 +
R. meliloti USDA 1020a -
R. meliloti USDA 1082 -
Arctic rhizobial isolate N35 -
Bradvrhizobium iaoonicum 61A76 NG
B. iaoonicum USDA 110 NG
1. Data are from a single experiment with duplicate flasks.
2. +, trehalose concentration at least double relative to 0 excess NaCl treatment.
3. no difference in trehalose concentration between salt treatments.
4. NG, no growth with 25 or 100 mM excess NaCl. CHAPTER III
STIMULATION OF a-GLUCOSIDASES FROM FAST-GROWING RHIZOBIA
AND AGROBACTERIUM TUMEFACIENS BY K+, Rb+, AND NH+A
Introduction
Disaccharides such as maltose, sucrose, and trehalose are a-glucosides; hydrolysis of these sugars to monosaccharides is the first step of their enzymatic breakdown. This can be accomplished by different enzymes, such as maltase (EC 3.2.1.20, a-glucosidase), invertase (EC
3.2.1.26, /3-fructofuranosidase) and a,a-trehalase (EC
3.2.1.28, a,a-trehalose glucohydrolase). Properties of these enzymes such as pH, substrate affinity, and substrate specificity vary among organisms, and the enzymes may react with a variety of a-glucosides and other substrates (Webb,
1984; Kelly and Fogarty, 1984).
In the course of assaying disaccharidases in Rhizobium leauminosarum biovar phaseoli USDA 2667 we observed that the a-glucosidase activities were stimulated by K+ ions.
Numerous enzymes have previously been documented to be activated by K+ and other monovalent ions, especially Rb+ and NH4+ (Suelter, 1970), but to our knowledge rhizobial a-glucosidases have not been reported to have this property.
67 Singh and Singh (1985) surveyed 45 strains of rhizobia
(covering all species and genera) and Agrobacterium tumefaciens for a-glucosidase, /3-glucosidase, and /?- galactosidase activity, but they used sodium phosphate buffer in their assays and did not report any effect of K+ on enzyme activity.
In this chapter I present data on the range of or ganisms having a-glucosidases that are activated by K+, the degree of activation, the effect of other ions, and some kinetic data for the enzymes from Rj_ leauminosarum bv. ohaseoli USDA 2667.
Materials and Methods
Bacterial strains and culture
Agrobacterium tumefaciens A281 was provided by Dr. J.
Finer (Ohio State University, Wooster, OH), Bradvrhizobium sp. Rp501 by Dr. J. Torrey (Harvard Forest, Petersham, MA), all other rhizobia by Dr. H. Keyser (USDA Nitrogen Fixation
Laboratory, Beltsville, MD), and Escherichia coli S17-1 by
Dr. R. Simon (Universitat Bielefeld, Federal Republic of
Germany). All chemicals and enzymes were from Sigma
Chemical Company, St. Louis, MO, unless otherwise noted.
Bacteria were cultured in liquid media at 29°C on a rotary shaker set at 150 rpm. Agrobacterium tumefaciens was grown in YEB medium supplemented with 0.2% sucrose (Hood et al..
1984). E. coli S17-1 was grown in LB broth (Miller, 1972; Appendix H) supplemented with 0.1% sucrose. Rhizobia were cultured at pH 7.0 with either trehalose, sucrose or glutamate (0.2%) as a carbon source and 0.8 g KN03/1 in the trehalose and sucrose media. Rhizobial media also contained micronutrients (Manhart & Wong, 1979; Appendix H), mineral salts (Appendix H), and (in mg/1) thiamin HC1 (0.4), Ca- pantothenate (0.5), biotin (0.1). Vitamins and carbon sources were filter sterilized before addition to autoclaved media.
Preparation of crude bacterial extracts
Bacterial cultures (250 to 1000 ml, absorbance Aaoo 0.5 to 1.5) were harvested by centrifugation for 10 minutes at
15,000 x g, resuspended in 20 ml cold 0.01 M sodium phosphate buffer (pH 7.0), recentrifuged at 12,000 x g, and the pellet resuspended in a small volume of the same buffer.
Three drops of 10% Triton X-100 were added and the cells were sonicated 3 minutes with a Branson model 350 sonifier
(Branson Ultrasonics Corp., Danbury, CT), set at an output of 3 on 50% pulsed duty cycle. This solution was then centrifuged at 27,000 x g for 10 minutes and the supernatant fraction desalted by passage through a 12 cm X 1 cm column of Sephadex G-25, using 10 mM sodium phosphate buffer (pH
7.0) as eluant. Sufficient blue dextran was added to give a visible color and was used to monitor the location of the protein front; blue effluent was collected and used for all enzyme assays. For the Saccharomyces cerevisiae extract,
1.0 g of dried baker's yeast (Red Star, Milwaukee, WI) was dissolved in 10 ml 0.01 M sodium phosphate buffer (pH 7.0) and processed in the same manner as bacteria.
Disaccharidase assays
Assays were run in duplicate with boiled enzyme controls
(zero substrate controls also gave no activity but boiled substrate controls were preferred because any glucose contamination in the substrates could be detected). Each assay contained crude enzyme extract (approximately 100 /zg protein) or an appropriate amount of commercial enzyme, and
12.5 /zmole sucrose or trehalose or 5.0 /zmole maltose, dissolved in 80 mM buffer at a final volume of 0.5 ml.
Extracts from rhizobia, A^. tumefaciens. legume nodules, and
E. coli were assayed at pH 6.6, yeast extract at pH 5.0, rice a-glucosidase at pH 4.0, and porcine trehalase at pH
5.7. Na- or K-Pipes buffer [piperazine-N,N'-bis(2- ethanesulfonic acid)] was used for the pH 6.6 assays, sodium phosphate/citrate (with KCl or NaCl added to a final concentration of 0.1 M) for all others, including the assays used for determination of pH curves. Na+ (as Na-Pipes buffer or NaCl) was used as a control for assays using K-
Pipes buffer or added KCl, thus maintaining ionic charge and osmotic concentration across treatments. In some cases, other salts were added as indicated in the text. After incubation at 37°C, all reactions were terminated by boiling for 5 minutes and glucose production was measured by the glucose oxidase method (Lloyd and Whelan, 1969).
Calculations for invertase activity were doubled because sucrose, unlike maltose and trehalose, contains fructose which is produced in amounts equal to glucose during hydrolysis, but is not detected in the glucose oxidase assay. /3-glucosidase and 0-galactosidase were assayed with the crude extracts and analyses used for a-glucosidases, using cellobiose and lactose for the respective substrates.
Protein was measured by the bicinchoninic assay method
(Smith et al.. 1985) with reagents from Pierce Chemicals
(Rockford, IL). Units of enzyme activity are nmole glucose produced/mg protein/minute.
Nodule extracts
Soybean, pole bean (Phaseolus vulgaris) and alfalfa plants were grown as previously described (Streeter, 1985).
The soybeans were inoculated with Bradvrhizobium iaponicum
USDA 110, pole beans with R. 1. bv. ohaseoli USDA 2667 and alfalfa with R^. meliloti USDA 1021. Nodules were frac tionated into plant and bacteroid fractions by the method of
Salminen and Streeter (1986). Sephadex-filtered cytosol
(plant fraction) and bacteroid soluble protein (rhizobial fraction) were assayed. The assay mixture was the same as that described for bacterial extracts. 72 Results
All a-glucosidase activities (maltase, invertase, and trehalase) from fast-growing rhizobia and A^_ tumefaciens
A281 were stimulated by K+ ions (Table 3, Fig. 3) .
Increases in activity were less than 2-fold to greater than
12-fold, depending on enzyme and species of bacterium. It should be noted that no attempt was made to harvest cultures at identical growth stages, so variations in specific activity or degree of activation may reflect culture age as well as genetic differences.
Of 10 other ions tested, only Rb+ and NH4+ had the same effect as K+ on a-glucosidase activity in R. !_• hv. ohaseoli extracts; most other ions had only a slight effect on enzyme activity (in the absence of K+; Na-Pipes assay) or were slightly inhibitory when added to K-Pipes assays (Table 4).
Mn+2 was slightly stimulatory in the presence of K+ (K-pipes assay); this was observed twice but was not pursued further
(Table 4) . AgN03, CdCl2, CuS04, HgCl2, NiCl2, and ZnCl2 were not tested because they interfered with the glucose oxidase assay; all salts listed in Table 4 did not interfere with glucose determination. R. 1. bv. ohaseoli extracts also contained /?-glucosidase and /3-galactosidase activity but no stimulation of these enzymes was found by any salts tested
(data not shown).
Increased trehalase activity could be detected in R. 1. bv. lequminosarum USDA 2667 extracts at KCl concentrations 73 as low as 0.5 mM. Increases in activity with additional KCl began leveling off at about 30 mM K+, but increased slightly even at 100 mM KCl (Fig. 3) . The Ka for K+ was calculated to be 1.8 mM from a Lineweaver-Burk plot of the data shown in Fig. 3.
All other a-glucosidases tested (from yeast, rice, pig,
E. coli. and soybean and alfalfa nodule cytosol) were not stimulated by the presence of K+ (Table 3). It should be noted that rice a-glucosidase had activity with maltose only, and the porcine enzyme had activity only with trehalose. Bacteroid soluble protein from P. vulgaris and G. max nodules did not contain detectable a-glucosidase. In addition, three species of cultured bradyrhizobia (B. iaponicum 61A76, Bradvrhizobium sp. Rp501, and
Bradvrhizobium sp. USDA 3045) did not have detectable a-glucosidase activities (K+ assay), even after being grown in the presence of trehalose (data not shown).
Specific activity of a-glucosidases in extracts from fast-growing rhizobia was greatly increased by culturing cells with an a-glucoside as the carbon source (Tables 3 and
4). a-Glucosidase activities and the degree of K+ stimulation were similar whether trehalose or sucrose was used for the carbon source (Table 6 ). Extracts of cells grown with glutamate contained very low levels of trehalase activity, although sucrose and maltose hydrolysis were still easily measured (Table 5). Kinetic data for maltose, sucrose, and trehalose hydrolysis by R. 1. bv. ohaseoli USDA
2667 crude extracts showed that all three a-glucosidases had
increased affinity for a-glucosides (indicated by the values) and increased catalytic rate (approximated by Vm) after bacteria were grown with trehalose (Table 5). The Vm values calculated from crude extracts do not reflect the true maximum catalytic rate of these enzymes because they were assayed in the presence of many other proteins.
However, differences in catalytic rate (represented by Vm) in the presence or absence of K+ do reflect changes in enzymatic properties of the a-glucosidase being assayed.
This is especially true with enzymes from the same culture because each assay contained an identical aliquot of protein. All K+-stimulated enzymes except for those from A. tumefaciens A281 had consistently higher specific activities for maltose and sucrose than for trehalose hydrolysis (Table
3) . The pH curves for the three a-glucosidase activities from rhizobia were similar with an optimum of about 6 . 6
(Fig. 4). In contrast, the three a-glucosidases from E. coli S17-1 had distinctly different pH optima (data not shown), and the same three disaccharidase activities in soybean nodule cytosol (Streeter, 1982) had pH optima which differ from those in Fig. 4. 75 Discussion
Among the organisms surveyed, only Rhizobium spp.
(sometimes categorized as "fast-growing" rhizobia) and the related genus Agrobacterium possess a-glucosidases that are stimulated by K+, Rb+, and NH4+. These two genera and the genus Bradvrhizobium (also known as "slow-growing" rhizobia) belong to the bacterial family Rhizobiaceae (Jordan, 1984).
Members of the genus Bradyrhizobium have been previously documented to lack disaccharidase enzymes in general and are unable to grow on disaccharides (Stowers, 1985; Singh and
Singh, 1985), so our inability to demonstrate a-glucosidase activity in this genus was expected and confirms previous observations.
Suelter (1970) observed that K+, Rb+ or NH^+ frequently can activate the same enzyme. It is intriguing that an enzyme from one organism may respond to an ion activator while taxonomically identical enzymes (those with the same
E.C. number) from other organisms do not have the same property. This is not always the case with monovalent cation-activated enzymes; pyruvate kinases from evolutionarily diverse organisms have all retained the same ion activation property (Suelter, 1970).
Enzymes reported to be activated by monovalent cations are generally phosphorylases, dehydrogenases or lyases
(Suelter, 1970); to our knowledge the activation of a- glucosidases by monovalent cations has not been previously 76 reported. Few other hydrolases have been reported to possess this property; Suelter (1970) listed a few phosphohydrolases in a review of enzymes activated by monovalent cations, and plant asparaginase (an amidohydrolase) has been reported to be stimulated by K+ (Sodek et al.. 1980).
The significance of the stimulation of this enzyme by
K+ and NH4+ is not clear. The effect is quite interesting in light of data indicating that K+ concentration influences the metabolism of free-living bradyrhizobia (Gober and
Kashket, 1987). They reported that combined high K+ and low
0 2 induced several metabolic changes in cultured
Bradvrhizobium sp. 32H1, all of which are associated with differentiation of free-living rhizobia into symbiotic bacteroids. However, because there was no detectable a-glucosidase activity in the two species of bacteroids we tested and no stimulation of nodule cytosol a-glucosidase activity by K+ (Table 3), the enzyme activation which we report is probably only relevant to the free-living state.
The fact that nodule cytosol and rhizobial trehalase activities both have the same pH optima is apparently a coincidence. Questions concerning the location and origin of nodule trehalase (Mellor, 1988; Salminen and Streeter,
1986) may be resolved by taking advantage of the fact that any rhizobial a-glucosidase should be activated by K+. Based on the data presented here it is possible that the three a-glucosidase activities of free living rhizobia and Ai tumefaciens A281 are all carried out by the same enzyme, with a higher specificity for maltose and sucrose than for trehalose. This idea is supported by several lines of evidence: all three enzyme activities are stimulated by
K+, NH4+, and Rb+ (Table 3) ; all exhibit the same pH optima
(Fig. 4), in contrast to the situation in E^. coli and nodule extracts (Streeter, 1982); all specific activities increased after culture on sucrose or trehalose (Tables 3 and 4); and all enzyme activities responded to different buffers in the same manner (data not shown). It would be interesting if all three hydrolytic activities were accomplished by the same enzyme.
Although the physiological implications of the stimulation of a-glucosidases by certain monovalent cations is not clear, the observations reported here have obvious practical value. In view of the large stimulation of enzyme activity, it is essential that monovalent cations be included in assays of these enzymes in extracts from members of the Rhizobiaceae in order to provide an accurate estimate of catalytic activity. Furthermore, although our survey was only superficial, the stimulation of these enzymes by monovalent cations might provide a convenient and valuable taxonomic tool for the delineation of the limits of this bacterial family. 78
Figure 3. Effect of KCl concentration on trehalase activity in extracts from IL. leauminosarum bv. ohaseoli USDA 2667. c E c 0) o h. cs E E s K a for K E 1.8 m M q> (/) o o 3 0 > (I) 100
KCl in reaction mixture (mM) Figure 3
vo so
Figure 4. pH curves for a-glucosidases from SU. leauminosarum bv. phaseoli USDA 26611 All assays were in 0.08 M sodium phosphate/citrate buffer with 0.2 M KCl. Maltase, o; Invertase, □ ; Trehalase, a . relative activity
01
18 82
TABLE 3. Response to K+ by a-glucosidases
from different sources1.
Source of Enzyme Maltase Invertase Trehalase
R. lecruminosarum bv.
Dhaseoli USDA 2667 + + ‘ ++
R. leauminosarum bv.
trifolii USDA 2066 +++ ++ ++
R. leauminosarum bv.
viceae USDA 2370 ++ ++ ++++++
R. fredii USDA 191 +++++
R. meliloti USDA 1082 ++ ++ +++++
R. loti USDA 3071 ++++ +++ +++++
Agrobacterium
tumefaciens A281 ++ ++ +++
Escherichia coli S17-1
Baker's yeast ND
Rice (G-9259) ND ND
Porcine Kidney (T-8778) ND ND 83
TABLE 3. (continued)
Glvcine max nodule
cytosol NA6 NA
Medicaqo sativa
nodule cytosol NA NA
1. All rhizobia were cultured with trehalose as sole carbon source. Data, in general, are for single observations.
2. +, Enzyme activity stimulated by K+. More pluses indicate a relatively greater degree of stimulation, a single plus indicates at least a 2 0 % stimulation.
3. -, No enzyme stimulation by K+.
4. ND, Not detected.
5. Purified commercial enzymes (Sigma Chemical Co. stock numbers).
6 . NA, Not assayed. 84
TABLE 4. Effect of different salts on trehalase activity
from R*. leguminosarum bv. phaseoli USDA 2667
% maximum activity1
Salt2 Na-Pipes3 K-Pipes3
NH4 C1 1 0 0 4 96
RbCl 98 95
KCl 87 94
KNO3 83 94
CsCl2 46 8 8 LiCl 32 90
MnCl2 31 1 0 0 5
BaCl2 31 89 NaCl 29 96
CaCl2 28 91
MgCl2 25 93
CoCl2 1 1 71
1. Values are the average of two experiments.
2. Salts were present at a concentration of 10 mM in final reaction mixture.
3. Buffer used for assay.
4. Maximum value obtained within Na-Pipes assays set to
100%, actual value 53.6 nmole glucose/minute/mg protein.
5. Maximum value obtained within K-Pipes assays set to
100%, actual value 74.1 nmole glucose/minute/mg protein. 85
TABLE 5. Kinetic properties of a-glucosidases in crude extracts from Rhizobium leauminosarum bv. ohaseoli USDA 2667 grown with a ,a-trehalose or glutamate as sole carbon source.
Assay Conditions and Km Vm Carbon Source (mM) (nmole alucose/min/ma orotein)
TREHALASE
Trehalose Culture:
K-Pipes assay 3.6 73.1
Na-Pipes assay 1 1 . 2 44.4
Glutamate Culture:
K-Pipes assay 17.3 4.6
Na-Pipes assay ND2 0.9 INVERTASE
Trehalose Culture:
K-Pipes assay 2 . 6 152.6
Glutamate Culture:
K-pipes assay 35.8 58.3 ■u- Na-Pipes assay 44.4 29.6
MALTASE
Trehalose Culture:
K-Pipes assay 1.3 71.2 Glutamate Culture:
K-Pipes assay 2 . 2 35.2
Na-Pioes assav 3.3 17.6
1. All values were calculated from Lineweaver-Burk plots with regression coefficients >0.99
2. ND, not determined due to low activity. 86
TABLE 6 . Comparison of a-glucosidase activity in crude
extracts of two species of rhizobia grown with sucrose or
a,a-trehalose as the sole carbon source.
Species Maltase Invertase Trehalase
R. leauminosarum bv.
phaseoli USDA 2667
carbon source: *4* o | VO o sucrose 127.4 (2.9) 3 150.1 (3.7) •
a,a-trehalose 114.1 (1.5) 171.5 (2.6) 77.3 (2.6)
R. fredii USDA 191
carbon source:
sucrose 6 6 . 8 (1 .2 ) 141.9 (1.4) 67.1 (5.5)
a,a-trehalose 80.7 (1 .2 ) 173.1 (2.1) 85.1 (5.9)
1. Units are nmole glucose produced/minute/mg protein and are the average of two replicate assays.
2. Activity with K-Pipes used as buffer and source of
K+.
3. Numbers in parentheses are the fold-stimulation in activity of the enzyme assayed in K-Pipes vs. Na-Pipes (no
K+) buffer.
4. No activity was detected in the Na-Pipes assay. CHAPTER IV
INCREASED ACCUMULATION OF TREHALOSE IN RHIZOBIA
CULTURED UNDER 1% OXYGEN
Introduction
It is well established that N2-fixing nodules have greatly reduced internal 0 2 tensions, with most estimates in the range of 10-20 nM free 02 in the bacteroids (Appleby,
1969; Tjepkema and Yocum, 1974). This low level of 02 is essential for the functioning of nitrogenase (for reviews see Appleby, 1984; Hill, 1988). Many studies have been concerned with the effects of low 0 2 levels on various growth parameters of cultured rhizobia, and a wide range of physiological changes under low 0 2 tension has been documented. These include; reduced exopolysaccharide synthesis (Agarwal and Keister, 1983; Pankhurst and Craig,
1978; Tulley and Terry, 1985), increased nitrogenase synthesis (Scott et al.. 1979), increased levels of mRNA transcripts for genes involved with N2 fixation such as
NifB, NifH, NifDK, FixA and Glnll (Adams and Chelm, 1988), induction of NifA (Ditta et al.. 1987), increased levels of
87 88 high affinity terminal oxidases (Bergersen and Turner,
1980), induction of a methylammonium transport system and a
K+/H+ antiporter (Gober and Kashket, 1987), synthesis of heme (Avissar and Nadler, 1978), and induction of N2-fixing activity (Bergersen et al.. 1976? Keister, 1975; Tjepkema and Evans, 1975).
The changes which occur in rhizobia cultured under low
0 2 tension are similar or identical to those observed in differentiated rhizobia (bacteroids) found within legume nodules, and are associated with the development of a successful symbiotic relationship. Many of the rhizobial responses to low 0 2 also have been shown to be sensitive to increases in 0 2; rapid reversals of the effects are associated with increased 0 2 levels once the response to reduced 02 has been established (Adams and Chelm, 1988;
Bergersen and Turner, 1980; Bergersen et al.. 1976? Hill,
1988; Scott et al.. 1979).
Consideration of the low levels of free 02 in infected legume nodule cells, and the accumulation of trehalose in nodules established by rhizobia known to accumulate essentially no trehalose when cultured aerobically, led to the hypothesis that low 0 2 levels in the nodule environment may induce the accumulation of trehalose. This chapter is a report of the effects of reduced Oz tension on trehalose accumulation in rhizobia cultured under a variety of conditions. 89
Materials and Methods
All rhizobia were obtained from the USDA Nitrogen
Fixation Laboratory, Beltsville, MD. Cultures were maintained at 29°C on a rotary shaker at 125 RPMf and supplied with either 21% or 1% 02 in the gas phase. Gases were vented into the headspace of 125 ml Erlenmeyer flasks
(containing 50 ml liquid culture media) via a sterile filtration system. The 1% 02 (remainder N2) source was a commercial mixture from a compressed gas cylinder, with
99.6% pure 02 and a concentration tolerance of 1.00 + 0.05%, and 99.999% pure N2 (Liquid Carbonic Corp., Harrison, NJ).
The 21% 02 source was ambient air from pressurized air lines in the building. All gases were filtered through a Na0H/H20 trap and a 0.3 fm pore size bacterial air vent (Gelman
Sciences Inc., Ann Arbor, MI) before being vented into the flasks.
Media were placed in the flasks in 50 ml aliquots prior to autoclaving the entire gas filtration and distribution system. The succinate medium was based on McAllister and
Lepo (1983; Appendix G), with 25 mM succinate, micronutrients as in Manhart and Wong (1979; Appendix G), and vitamins as indicated below. Both the glucose and glycerol media contained 0.2% C source and mineral salts
(Appendix G). vitamins were filter sterilized and added to autoclaved media for Ri. 1. bv. phaseoli USDA 2667 (except where noted) in the following quantities (in mg/1 ): biotin 90
(0.1), thiamin HC1 (0.4) and Ca-pantothenate (0.5). All
cultures for the experiment described in Table 8 were
supplemented with 0 . 1 g yeast extract/ 1 in place of vitamins. Yeast extract mannitol gluconate (YEM6 ) medium was the YM broth described in Vincent (1970) with 2 g sodium gluconate/1 added. All media contained a trace (2 mg/1) of phenol red as a pH indicator. Bacteria were initially cultured aerobically in liquid YEMG and 100 fil of these start cultures were used to inoculate the experimental flasks.
The pH of the cultures was monitored by the color of the phenol red, and after harvesting cells the pH of the remaining liquid was determined using a pH meter. This information was useful as a rough indicator of the growth phase of the cells, especially in the succinate medium which routinely attained a pH of 8.5 by the time stationary phase was reached. Absorbance measurement, a commonly used technique to monitor the growth of liquid cultures, was not practical with these experiments because many of the 1 % 0 2 cultures produced small clumps of cells. Final comparisons of trehalose accumulation between cultures were based on total protein content. Monitoring the pH of media to follow the growth of bacterial cultures has been previously documented (Pirt, 1975? Rice and Hempfling, 1978).
Cells were collected by centrifugation at 12,000 x g for 5 minutes and were resuspended in 10 ml H2 0. An aliquot was taken for protein determination and the remaining cells
, centrifuged again at 12,000 x g for 5 minutes. The
pellet was then suspended in 80% ethanol for extraction of
carbohydrates. After refrigeration for several hours or
overnight the samples were centrifuged for 5 minutes at
27,000 x g, and the supernatant fraction was dried down
under an air stream. Each sample was extracted 3 times and
the extracts were combined and evaporated to dryness. The
dried samples were dissolved in H2 0 and lipids were removed
with CHC13. Portions of the extracts were derivatized using
the STOX-HMDS method with reagents from Pierce Chemical
Company (Rockford, IL) (Appendix A), and analyzed for
carbohydrate content using gas chromatography (Streeter and
Bosler, 1976; Appendix A). The trimethylsilyl ethers of the
carbohydrates were injected into a Varian 3700 gas
chromatograph (Varian Associates, Palo Alto, CA) fitted with
a column packed with 3% OV-17 on Chromosorb W HP (Supelco
Inc, Beliefonte, PA). Standards were analyzed frequently to
assure proper quantitation of sample peaks.
For organic acid analysis of culture media, aliquots of
the liquid left after the cells were removed were
concentrated, filtered and applied to ion exchange columns
to separate organic acid, amino acid and neutral (sugars and
sugar alcohols) fractions (Streeter, 1987; Appendix A). QAE
and SP Sephadex (Pharmacia Inc., Piscataway, NJ) columns
were connected in tandem, and the neutral fraction was 92 collected by rinsing the columns with water after the samples were applied. The organic acids were collected by eluting the QAE Sephadex with 4% formic acid, and the amino acids by eluting the SP Sephadex with 0.2 M NH4 OH. Neutral sugars were processed as described above for the ethanolic extracts. The amino acid fractions were discarded.
Organic acid fractions were dried, the solids dissolved in H2 0, titrated with KOH to pH 7.0-7.5, and re-dried. The
K+ salts of the organic acids were then derivatized by boiling them in phosphate buffer with phenacyl bromide and dicyclohexane-18-crown-6 dissolved in acetone (Streeter,
1987; Appendix A). The phenacyl esters were then analyzed with a Waters (Milford Associates, MA) high pressure liquid chromatograph fitted with a Novapak C 18 steel column.
Samples were eluted with a methanol gradient and monitored at 245 nm with a Waters variable wavelength detector.
Tartrate was used as an internal standard, and standards were used daily to calibrate the instrument.
Aliquots of cultures were kept frozen in H20 at -20 °C until analyzed for protein concentration using the bicinchoninic assay method (Smith et al.. 1985) using reagents from the Pierce Chemical Company. Samples were thawed and portions added directly to assay tubes. The freeze/thaw cycle in H20 coupled with the very basic assay conditions apparently was sufficient to lyse the cells? boiling the samples to lyse cells further before assaying them did not lead to higher protein values. All reaction mixtures were centrifuged to remove cellular debris before determining the absorbance. All other chemicals were from
Sigma Chemical Co., St. Louis, HO.
Results
Cells of Ri L bv. phasedi USDA 2667 were grown in succinate medium and harvested at different times from early log phase through late log phase (Fig. 5). Cells grown under 1% 02 reached a maximum of about 5 mg protein per culture in approximately mid-log phase and ceased to accumulate more protein, although trehalose levels continued to increase slightly. In contrast, cultures maintained at
2 1 % 0 2 increased in protein content until late log phase (as indicated by maximal protein concentration), reaching levels of protein/culture about 5 times greater than those for the low 02 treatment. Thus, cultures grown with 21% 02 accumulated trehalose at a much lower ratio of trehalose to protein; three to four-fold greater amounts of protein were present in 2 1 % 0 2-treated cultures with the same amount of trehalose as the 1% 02-treated cultures (Fig. 5). In some cases the total amount of trehalose per culture was greater in the 2 1 % 0 2 cultures than in the 1 % 0 2 cultures; however, the higher levels were probably achieved by a much greater number of cells as indicated by the significantly higher protein concentrations obtained in the 2 1 % 0 2 treatments. Information on the amount of trehalose present per cell would verify that the increases in trehalose concentration
(expressed on a per mg protein basis) under 1 % 0 2 are indeed due to elevated levels in individual cells. Unfortunately, this data is not available for these experiments. Any future work with this phenomenon should include plate counts at the time of harvest in addition to protein determination so the concentration of trehalose per cell can be evaluated.
The increased accumulation of trehalose in R. 1. bv. phaseoli USDA 2667 under 02 stress was not dependent on C source (Table 7). Some differences between media in both total trehalose content and trehalose concentration (on a per mg protein basis) were observed. However, these may be due to harvesting the cultures at slightly different times during the log phase of growth and therefore do not necessarily represent a differential response to C source.
The YEMG medium initially contained approximately 100 nq trehalose (from the yeast extract) per 50 ml culture, so trehalose synthesis would not have been required in this medium. However, the results with YEMG-grown cells were essentially the same as those grown in the three defined media. Succinate was the only C source that promoted trehalose accumulation in rhizobia grown under 2 1 % 0 2; the induction of trehalose accumulation by dicarboxylic acids had been previously observed with this strain. An ancillary observation in this experiment was the change in pH in the 95 glycerol medium: the pH increased in the 21% 02 treatment and decreased in the 1% 02 treatment (data not shown). This suggests induction of different metabolic pathways for glycerol metabolism in the two 0 2 treatments.
Three different species (R^. meliloti. R. leauminosarum. and Rj_ fredii), including two biovars of R^. lequminosarum
("phaseoli'1 and "viceae") of rhizobia were tested for their response to reduced 02 tension (Table 8 ). This selection of rhizobia includes strains (R±. meliloti USDA 1021, EL*. fredii
USDA 191) previously documented to produce very low levels of trehalose in liquid media under ambient atmospheric 0 2
(Streeter, 1985; Hoelzle, unpublished observations). The strains Rj. fredii USDA 191, R*. JL-. bv. viceae USDA 2391 and
R. meliloti USDA 1021 were also chosen because they do not respond to osmotic stress by accumulating trehalose, which is a previously documented property of R^. 1 ^ bv. phaseoli
USDA 2667 and a few other rhizobia (Hoelzle and Streeter,
1989; Smith and Smith, 1989). In all four types of rhizobia the concentration of trehalose was considerably greater in cells grown under low 0 2 tension relative to those grown with 21% 02. The strains that generally accumulate nil trehalose under atmospheric 02 (R^ fredii USDA 191 and R. meliloti USDA 1021) had lower levels of trehalose in the 1% treatment than the 2 strains that normally accumulate some trehalose under 2 1 % 0 2. In a final experiment, FU li. bv. phaseoli USDA 2667 was grown in defined succinate medium under 1% 02 for 48 hr, then some flasks were aseptically switched to 21% 02 (Table
9). After 7.5 hours the amount of trehalose present in the cells of the 21% 02 treatment had decreased approximately 7- fold and the protein levels had increased more than 2 -fold, relative to cultures that had remained at 1% 02. The trehalose was apparently lost via respiration or converted to some storage polymer, because there were no detectable sugars in the neutral fraction of the culture supernatant, nor was there any significant change in the organic acid concentration in the medium (data not shown). Increased concentration of poly-/?-hydroxybutyrate in cultured
Rhizobium sp. ORS571 has been documented in response to increases in free 0 2 level, so it is possible that the trehalose was converted to poly-/3-hydroxybutyrate (De Vries et al.. 1986). Trehalose breakdown could not be explained by culture senescence, because succinate concentration remained high in the medium and the pH had not reached maximum levels (data not shown). Significant levels of malate (265.5+13.8 #xg/culture) and fumarate (90.8+4.8
/ig/culture) were found in the culture media from this experiment. However, there were no significant differences in the amounts of organic acids between 0 2 treatments and the amount per culture did not change following the switch from 1 % to 2 1 % 0 2. 97
One complication with these experiments is the fact that trehalose accumulates in rhizobial cultures until
stationary phase and then is rapidly broken down (Streeter,
1985? Hoelzle and Streeter, 1989). If the cultures grown with 2 1 % 0 2 were to reach stationary phase it would be possible to observe low levels of trehalose in the cells.
However, we have been careful to insure this is not the case by monitoring the culture pH (all experiments) and the succinate concentration remaining in the culture medium
(percent 0 2 switching experiment).
Discussion
We have found that several species of rhizobia accumulate much more trehalose when cultured with 1 % 0 2 than with 2 1 % 0 2 in the gas phase (theoretically 1 0 and 2 2 0 nM 0 2 in the media, respectively). Furthermore, we observed this phenomenon in rhizobia cultured in media with different C sources requiring distinct metabolic pathways for both synthesis of trehalose and for cell growth and maintenance: glycerol, glucose, succinate, and a complex medium. These results indicate that the response to low 0 2 is not dependent on nutritional status or species of rhizobia.
The biochemical mechanism for trehalose accumulation under conditions of 0 2 stress is not clear? the accumulation could be due to increased synthesis, decreased breakdown or some combination of both. We have not conducted any assays of the trehalose metabolic enzymes in cells from our
oxygenation system; however, previous data from legume nodules suggests that bacteroid disaccharidases are not
functional (Hoelzle and Streeter, 1990), although it has been shown that isolated bacteroids can take up and metabolize trehalose at a low rate (relative to dicarboxylic acids) (Salminen and Streeter, 1987). The lack of significant trehalase activity in the nodule bacteroids leads us to the suggestion that increased trehalose pool size in cultured rhizobia under low 0 2 tension is due to decreased breakdown of the disaccharide via some type of inactivation of trehalase. However, McKay et al. (1989) have recently shown that glucose-6 -phosphate dehydrogenase activity is reduced in 0 2-limited rhizobia; this might lead to a buildup of glucose-6 -phosphate, which is a precursor of trehalose (Elbein, 1974) and consequently could be utilized for trehalose synthesis.
It is intriguing that under 02 stress the cells converted a dicarboxylic acid (succinate) to a disaccharide
(Fig. 5, Tables 7, 8 and 9). Low 02 tension has been previously documented to lead to reduced activity of TCA cycle enzymes in bacteria (Gray et al.. 1965; Jackson and
Dawes, 1976), so under these conditions (low 02 tension combined with a succinate medium) the bacteria may not be able to utilize dicarboxylic acids in the same manner as 99 when cultured aerobically. Trehalose synthesis may
represent a response to this dilemma.
It is not clear why the accumulation of trehalose by bacteroids in nodules or cultured rhizobia under 0 2 stress would be a useful adaptation. Trehalose is found in a wide variety of organisms (Elbein, 1974), and many different
functions for trehalose have been suggested (see Chapter I).
At this point I am not able to identify any of these roles as the function of trehalose in rhizobia. The possibility of a role in osmotic stress response can be eliminated because some rhizobia have been found to accumulate trehalose in response to salt stress but others do not
(Hoelzle and Streeter, 1989? Smith and Smith, 1989). Legume nodules contain many other compounds in much greater abundance (Streeter, 1987), so the possibility of a role for trehalose as an energy storage compound can also probably be eliminated.
It appears that increased trehalose accumulation is among the responses of rhizobia to low 0 2 levels; this concept can now be added to several other observations concerning trehalose in legume nodules: the appearance of trehalose at the onset of N2 fixation (Streeter, 1980), the synthesis of nodule trehalose by bacteroids (Reibach and
Streeter, 1983; Salminen and Streeter, 1986), the transfer of a significant proportion of the trehalose in nodules from the bacteroids to the plant fraction (Streeter, 1985), and 100 the presence of trehalose in all legume nodules examined
(Streeter, 1985). Although the function of trehalose in N2- fixing nodules is not clear from the present results, the possibility remains that this unusual carbohydrate has a role in the symbiotic state. 101
Figure 5. a,a-Trehalose and protein levels in cultures of R. lecruminosarum bv. phaseoli USDA 2667 grown in succinate medium under 1 % or 2 1 % 0 2 and harvested at various times during early through late log phases of growth. Increasing protein levels indicate more advanced growth stages within a treatment. O , 1 % 0 2; □ 2 1 % 0 2. Trehalose/Culture (/jq) 200 300 100 0 5 oenCutr (mg) ulture rotein/C P 10 iue 5 Figure 15
20 25 103
Table 7. Trehalose accumulation in cells of
R. leouminosarum bv. Phaseoli USDA 2667
grown with different carbon sources under 2 1 % or 1 % oxygen.
Percent jug Trehalose/ ng Trehalose/ Medium1 Oxygen Culture mg Protein
Glucose 21 ND2 <0. 9±0.13'4
1 84.1±18.8 36.4+5.7
Glycerol 21 ND <0. 9+0. 23
1 61.1±4.7 29.2+1.9
Succinate 21 53.7+3.4 8.5±0.1
1 57.8+4.8 25.1±0.1
YEMG 21 ND <1.4+0.23
1 25.9+3.8 12.2+0.4
1. Glucose, glycerol, and succinate media contained a defined vitamin mixture, YEMG contained yeast extract as a vitamin source.
2. ND, not detected; less than 4 ng trehalose present in total culture extract.
3. 4 ng trehalose/total culture extract used for calculations; SE represents variability of protein concentration per culture.
4. Values represent the mean ± SE of 3 or 4 replicate samples. 104
Table 8 . Accumulation of trehalose in different species of
rhizobia grown under 2 1 % or 1 % 0 21'2.
% Hg Trehalose/ Hg Trehalose/ Species Oxygen culture mg Protein
R. 1. bv. nhaseoli 2 1 39.3+1.23 4.7+0.1
USDA 2667 1 89.4+4.5 57.8±2.4
R. 1. bv. viceae 2 1 1 1 .0+3. 8 4.8 + 0 .9
USDA 2391 1 115.8+57.3 8 5.6+41.8
R. fredii 2 1 9.5±0.5 0 .8 + 0 .6
USDA 191 1 1 2 .7+1.5 1 0 .9+2.5
R. meliloti 2 1 ND4 <0.5+0.25
USDA 1021 1 16.9+4.0 4.3+1.0
1. All cells were cultured in succinate medium supplemented with 0 .1 g yeast extract/ 1 as a vitamin source.
2. All cultures were harvested during log phase of growth.
3. Values represent the mean + SE of 3 replicate samples.
4. ND, not detected, less than 4 ng trehalose present in total cell extract.
5. 4 fig trehalose used for calculations; SE represents variation in total protein per culture. Table 9. Effect of switching from 1% to 21% oxygen
on trehalose accumulation in cells of
R. lecruminosarum bv. phaseoli USDA 26671
fig Trehalose/ ng Trehalose/ Oxygen Treatment Culture mg protein
1 %, time 0 45.1±6.02 '3 27.9+1.3
1 % -* 1 % 4 52.7+8.4 27.6±4.2
1 % -*• 2 1 % 5 6 .7+0.1 1 . 2 ± 0 . 1
1. All cells were cultured in succinate medium supplemented with a defined vitamin mixture.
2. Values represent the mean + SE of 3 replicate samples.
3. No trehalose was detected in culture filtrates.
4. Cultures incubated for 7.5 hours after time 0.
5. Cultures switched to 21% 02 at time 0 and incubated 7.5 hours. CHAPTER V
GENETIC STUDIES
Introduction
The focus of the research for this dissertation was to determine the role of trehalose in legume nodules. It has been established that the source of trehalose in nodules is the bacteroids in infected cells (Reibach and Streeter,
1983; Streeter, 1985). An obvious technique to determine the effects of trehalose (and possibly the role and/or importance) in the symbiotic state would be to generate trehalose-negative and trehalase-negative mutants of rhizobia. By using such mutants to establish symbioses, any differences in physiology or morphology between plants inoculated with trehalose-metabolism mutants and plants inoculated with wild-type rhizobia would reflect differences in trehalose metabolism, with the desired mutants, critical experiments could be done to investigate the role of trehalose in legume nodules.
In an attempt to acquire trehalose metabolism mutants of rhizobia, Tn5 mutants were generated and screened for the necessary phenotypes. Rhizobium leauminosarum bv. phaseoli
USDA 2667 was selected for mutagenesis. This strain was
106 107 chosen for 3 reasons: 1) It could be grown with trehalose as the sole carbon source (in contrast to members of the genus
Bradvrhizobium\, which would allow selection for lack of trehalase production by inability to grow on trehalose media, 2) It accumulates an easily detectable amount of trehalose in culture, so any mutant not synthesizing trehalose would be obvious and 3) It responds to osmotic stress by accumulating excess trehalose (Hoelzle and
Streeter, 1989; Chapter II), so mutants lacking the ability to synthesize trehalose could be identified in a preliminary screen by impaired growth under hyperosmotic stress (Strom et al. . 1986; Giaever et al.. 1988).
A possible problem with generation of rhizobial Tn5 mutants could be the presence of multiple copies of the genes of interest. If this was the case, one gene copy might be successfully interrupted by transposon insertion but other(s) would still be functional. The phenomenon of multiple gene copies has been documented in several species of rhizobia, and more than one gene copy may be functional simultaneously (Quinto et al.. 1982; Flores, et al.. 1987;
Appelbaum et al.f 1988). During the mutant screening time period two papers documenting cloning of genes of trehalose metabolism from ELs. coli were published (Boos et al.. 1987;
Giaever et al.. 1988) . Clones of periplasmic trehalase (Boos et al. f 1987) and a gene involved in osmotic induction of trehalose synthesis (Giaever et al.. 1988) were obtained from 108 those authors. These genes were used as probes of rhizobial
DNA, in an effort to determine if there were multiple copies of these genes in rhizobia. Finally, a complementation experiment was done, using a rhizobial gene library to replace the otsAB genes in an EL coli deletion mutant. This was an effort to obtain the rhizobial equivalent(s) of the osmotic trehalose synthesis gene(s) from EL. coli.
Materials and Methods
Tn5 mutagenesis
Rhizobium 1. bv. phaseoli USDA 2667 was obtained from
Dr. H. Keyser, Nitrogen Fixation Laboratory, Beltsville, MD.
The Tn5 vector was EL coli S17-l/pSUP2021, provided by Dr.
R. Simon, Universitat Bielefeld, Bielefeld, FRG.
Tn5 mutants were generated according to the method of
Dr. D. Noel (personal communication; modified from Simon et al.. 1983). Prior to mating, rhizobia were grown at 29°C in liquid low sugar medium (Appendix H) and EL coli were grown at 37°C in LB broth (Appendix H). Best results were obtained when the rhizobial culture used for mating had reached an Aaoo of about 0.3 and the EL coli culture about
2.0. Initiation of transfer of the Tn5 insert to the rhizobia was accomplished by pipeting 0 . 2 ml of the EL coli culture and 0.5 ml of the rhizobial culture onto a plate of solid low sugar medium. The bacteria were mixing by gently swirling the plate, and incubated at 29#C for 2 days. 109
Bacteria were then resuspended from the agar in 3.0 ml of
0.1 M MgS04, vcrtexed, and diluted 10:1 with 0.1 M MgS04.
Mutant Screening
Rhizobia containing Tn5 insertions were selected by plating 0.1 ml of the diluted mating mixture onto solid TY agar plates (Beringer, 1984; Appendix H) containing 200
Mg/ml kanamycin sulfate and 2 0 nq/ml nalidixic acid.
Colonies formed after approximately 1 week incubation at
29'C, and were transferred with sterile toothpicks to grids stamped onto fresh TY plates containing antibiotics. Grid marking and all further transfers for secondary screening were accomplished using a plastic carpet protector (American
Hardware Supply Co., Butler, PA), manufactured to be placed under the legs of heavy furniture. These are square, about
3 X 3 inches, and have short pegs on the bottom surface that served as prongs to transfer colonies. Use of this tool, which could be repeatedly sterilized in 70% ethanol, allowed replicate transfers of 40 colonies per plate, providing a convenient method for mutant organization, identification and storage.
A primary screen of all Tn5 mutant colonies was done by replica plating each mutant onto three types of media: one with a high salt concentration, one with trehalose as the sole carbohydrate source, and a defined minimal medium
(Appendix H). The high salt medium was used to screen for 110 inability to produce trehalose as an osmoticum under salt stress, the trehalose medium for mutants lacking trehalase, and the minimal medium to identify auxotrophs (which might give a false negative reaction on either of the first two media). All media contained 1.0% agar, 200 jug/ml kanamycin sulfate, and 2 0 jug/ml nalidixic acid, and in addition the trehalose and the minimal media contained the following vitamins (mg/1): biotin (0.5), calcium pantothenate (1.0), and thiamin HC1 (0.4).
All Tn5 mutants were replica plated onto all three selection media and incubated at 29°C for about 3 days.
Sets of replicate plates were compared, and any colonies growing poorly on the high salt or trehalose media but showing normal growth on minimal media were noted. The mutants that grew poorly on the high salt medium were subcultured and analyzed for trehalose accumulation. The mutants that did not grow on the trehalose medium were re streaked onto the same medium, and then subcultured and also analyzed to determine their trehalose accumulation patterns. In a few cases mutant strains with superior growth on the test media were also analyzed for trehalose accumulation.
Evaluation of trehalose accumulation in Tn5 mutants
A modified succinate medium was used for subculture, both because it induces trehalose synthesis in rhizobia, even under aerobic conditions (Hoelzle and Streeter, 1989; Ill Chapter II), and because the organic acid succinate does not interfere with carbohydrate analysis. This medium is based on McAllister & Lepo (1984; Appendix H) with 100 mM excess
NaCl, 200 ng/ml kanamycin, 20 /xg/ml nalidixic acid, and a trace amount of phenol red as a pH indicator.
Cultures were allowed to grow until the medium had obtained a pH of approximately 8.0, and harvested by centrifugation at 12,000 x g for 5 minutes. The cells were brought up in 10 ml H20 and an aliquot was taken for dry weight measurement. The remaining cells were extracted 3 times in 80% ethanol and the extracts pooled and dried under and air stream. Dried extracts were dissolved in 2 ml H2 0, extracted with 1 ml CHC13, and portions taken for sugar analysis. Samples were derivitized using the STOX-HMDS method of the Pierce Chemical Co. (Rockford, IL), and analyzed as silylated oxime derivatives with a Varian 3700 gas chromatograph fitted with a i m glass column packed with
3.0% OV-17 (Supelco, Inc., Bellefonte, PA) (Appendix A).
Any mutant strain with a low trehalose accumulation pattern was analyzed in the same manner a second time to insure no error in trehalose measurement had occurred.
Some mutants were used to inoculate seeds of Phaseolus vulgaris cv. "Kentucky Wonder". Cultures were grown in YEMG
(Appendix H) and used for inoculum at an A^q of approximately 0.3 units. Three ml of inoculum was applied to each seed at the time of planting. Plants were grown in 112 sterilized silica sand, irrigated daily with a nutrient solution lacking combined nitrogen, and kept in a greenhouse environment supplemented with artificial illumination during the fall and winter months (Streeter, 1985). Nodules were harvested and carbohydrates extracted in some cases.
Molecular biology techniques:
Source of DNA
The gene for EL. coli periplasmic trehalase (treA) was provided by Dr. E. Bremer (Universitat Konstanz, GDR), and a gene for osmotic trehalose synthesis (otsA) in EL. coli was provided by Dr. A. Strom (University of Tromso, Norway).
The plasmids carrying the genes are referred to as pTrell and pFF47, respectively. The plasmid pRmJ30, which contains primarily node DNA from IL. meliloti USDA 1021 (Jacobs et al.. 1985) was provided by Dr. Kent Peters (Ohio State
University, Columbus, OH). Rhizobium 1. bv. phaseoli USDA
2667 and IL. meliloti USDA 1021 were obtained from Dr. H.
Keyser (Nitrogen Fixation Laboratory, Beltsville MD), and Rt.
1. bv. phaseoli CFN42 was provided by Dr. L. Flores
(University of Mexico, Cuernavaca, Mexico).
Bacterial transformation
The treA gene was provided as an isolated plasmid rather than as a plasmid in a bacterial host, so it was necessary to transform EL. coli strain MV1190 [A(srlR- 113 recA)306::TnlO, A(lac-pro), thi, sup E, (F', ProAB+, lac Iqf lacZ aM15, trad 36, provided by Dr. M. McMullen] to amplify the amount of DNA available for hybridization experiments
(Sambrook et al.. 1989? Appendix B).
Isolation of DNA for labeling
Single colonies from pTrell transformation selection plates or from plates streaked with EL. coli 6169/pFF47 were used to inoculate 5 ml LB broth (Appendix H) containing 15
Hi ampicillin/ml and incubated overnight at 378C and 250
RPM. Plasmids were isolated using a modified plasmid mini- prep (Birnboim and Doly, 1979? McMullen and Louie, 1989?
Appendix B). Small amounts of the isolated plasmids were linearized with EcoRI restriction enzyme (Appendix C) and run out on a mini-gel (Appendix D) with bacteriophage lambda
HindiII markers to check that the plasmids were the expected size. The inserts were then cut out of the plasmid vectors with appropriate restriction enzymes (EcoRI for pRmJ30,
EcoRV and BamHl for pFF47, and EcoRV and Hindlll for pTrell?
Appendix C). The digestions were run on a 10 X 14 cm agarose gel with markers, photographed, and the insert DNA cut out of the gel with a small spatula. The DNA was purified from the agarose by using a "Geneclean" kit (Bio
101, La Jolla, CA) and dissolved in 40 ill TE (Appendix H) .
Five nl of each insert was run on a mini-gel with markers to verify the size and estimate the concentration of the DNA. 114
Immobilization of Bacterial Genomic DNA
Genomic DNA was isolated from E*. 1. bv. phaseoli USDA
2667 and CFN42 and R*. meliloti USDA 1021 using a modification of the technique of Meade et al. (1982)
(Appendix C) . Genomic E_s_ coli DNA was purchased from Sigma
Chemical Co., St Louis, MO. The DNA was digested with ecoRI for 5 hours (Appendix C) and run with markers on two 10 X 14 cm gels. The gels were photographed, then denatured using an acid depurination protocol (McMullen and Louie, 1989;
Appendix E). The DNA was transferred to a nylon membrane
(Zetaprobe, Bio-Rad Laboratories, Richmond, CA) using a modified Southern capillary transfer (Southern, 1975;
McMullen and Louie, 1989; Appendix E).
The baked membranes were pre-hybridized in Seal-a-Meal bags (Dazey Corp., Industrial Airport, KS) for at least 4 hours, then transferred to fresh bags and hybridization solutions added (Appendix F). The DNA inserts from pFF47 and pTrell were used for probes after they were labeled using a random primer technique (Feinberg and Vogelstein,
1983; Appendix F). After 10-48 hours incubation at 37°C or
65°C the membranes were removed from the bags and washed at low stringency (Appendix G). The membranes were wrapped in
Saran Wrap (Dow Chemical Co., Midland, MI) and placed in an autoradiography cassette with Kodak X-OMAT AR film (Eastman
Kodak Co., Rochester, NY) and two intensifying screens. 115
After 1-24 hours exposure at -80°C the film was developed using a Kodak MR100 automatic developer. After autoradiography the blots were usually washed at slightly higher stringency and re-autoradiographed. The probes were then stripped off the membranes (Appendix G) and both membranes reprobed with labeled pRmJ30 insert in the same manner as the two E_j_ coli genes.
Complementation of otsA deletion mutant
The E_s_ coli otsAB deletion mutant, FF4031 (Giaever et al.. 1988) was provided by Dr. A. Strflm, University of Tromso, Norway. A cosmid library of L. bv. phaseoli
CFN42 DNA ligated into pLAFRl and maintained in E_s_ coli
HB101 was provided by Dr. Ron Diebold, Ohio State
University, Columbus, Ohio. Escherichia coli containing the helper plasmid pRK2013 (Figurski and Helinski, 1979) was obtained from Dr. D. Coplin, Ohio State University.
All 3 coli strains were grown to an A^jo of approximately 1.0 in liquid LB (Appendix H) medium; 15 nq tetracycline HCl/ml was added to the medium for the library and 50 nq kanamycin sulfate/ml to the pRK2013 medium. Tri- parental matings were performed by mixing 0 . 2 ml of each strain together on LB plates and incubating them overnight at 37°C. The cells were then re-suspended from the mating medium in 3 ml 0.1 M MgS04. Selection for transfer of genes involved in trehalose synthesis was done by plating diluted 116 samples of the mating mixture on M63 plates containing 0.45 M excess NaCl and 15 fig tetracycline HCl/ml (Giaever et al..
1988; Appendix H). Plates were incubated for 3-7 days at
37°C and any colonies picked and streaked onto LB plated supplemented with 15 fig/ml tetracycline HC1. Complemented cells were tested for the ability to synthesize trehalose by culturing them in 250 ml liquid M63 and analyzing them for trehalose accumulation in the same manner as the rhizobial
Tn5 mutants.
Results
Tn5 Mutagenesis
Over 6000 Tn5 mutants were generated and screened with the preliminary three-medium growth assay. About 200 mutants were identified as unusual in their growth on high salt or trehalose media and subcultured for further analysis
(Table 10). This technique identified a variety of mutants with altered trehalose accumulation patterns, but none that could not synthesize trehalose at all, and all those that grew poorly or not at all on the trehalose medium could synthesize and degrade trehalose internally in a wild type manner. Several mutants were tested that had the opposite phenotype than originally searched for; this also identified some strains with altered trehalose accumulation patterns (Table 10). 117
Several mutants were isolated that produced lower than usual amounts of trehalose; some of these were used in inoculation experiments and consistently induced a prolonged yellowing period (3-6 days) during establishment of the symbiosis (a short yellowing phase of 1-3 days is common with this variety of Phaseolus vulgaris when inoculated with the wild-type L, 1^. bv. phaseoli USDA 2667). However, because trehalose was still present in the nodules, and the plants eventually recovered from the extended yellowing period, no further experiments were conducted with these mutants.
The complementation experiment was not successful in isolating a rhizobial gene for trehalose synthesis (data not shown). The matings were performed twice, and all colonies that formed on the M63 medium analyzed for trehalose production. A total of 25 different complemented colonies were tested, and even when the quantity of cells extracted represented 1 0 - 2 0 times the amount needed to detect trehalose in the wild-type cells, no trehalose was found.
The rhizobial plasmids were easily lost by dilution from cultures on LB plates if the tetracycline HC1 was old or omitted; the complemented EL. coli 4031 cells apparently required the transferred plasmids for tolerance to high salt concentration because they did not grow if tetracycline was omitted from the medium. The library used for the complementations has been successfully used for 118 complementation of rhizobial mutants (R. Diebold, personal communication), so the failure to obtain complemented E. coli cells was probably not due to some flaw in the library.
Furthermore, the cultures that gained the ability to grow in osmotically stressful media apparently were synthesizing some (unidentified) compound that conferred osmotolerance, in addition to the utilizing the genes conferring tetracycline resistance, indicating that the transferred plasmids were successfully transcribed and translated by the
E. coli cells.
The attempt to hybridize cloned trehalose metabolism genes from EL. coli to rhizobial DNA was also unsuccessful.
Originally the hybridizations were performed at 65°C in an aqueous hybridization solution, which are conditions more appropriate for generating DNA duplexes of a high degree of homology (data not shown). A second set of conditions was also employed, using a 50% formamide hybridization solution,
37"C incubation and low stringency washing. These conditions are more likely to allow formation of a DNA heteroduplex of low homology, however, no hybridization between the EL. coli genes and rhizobial DNA was detected even under these conditions (Plates I and II). The homologous reactions produced strong binding signals (Plates
I and II for otsA/E. coli and treA/E. coli. respectively), however, even when the films were intentionally overexposed no binding of EL. coli DNA to rhizobial DNA was detectable. 119
The immobilized rhizobial genomic DNA was capable of binding a probe, even at low homology, as demonstrated by the binding of the pRmJ30 insert (from Rs_ meliloti USDA 1021) to
DNA from both Rs_ meliloti USDA 1021 (high homology) and R-. li. bv. phaseoli (low homology) (Plates III and IV).
Discussion
The mutants selected for no or greatly reduced growth on trehalose media apparently were uptake mutants, rather than lacking in the ability to breakdown trehalose. This conclusion is based on the trehalose accumulation patterns of these mutants; there was no buildup of trehalose as might be expected if it was synthesized and not broken down (Table
1 0 ), and all stored trehalose disappeared at stationary phase as in the wild-type cells (data not shown). These mutants nodulated Phaseolus vulgaris with no apparent effect on phenotype, however, since the trehalose in the nodule originates in the bacteroids, the ability to take up trehalose may not be of any consequence. Furthermore, there is some evidence that nodules formed by wild-type rhizobia do not have disaccharidase activity (Hoelzle and Streeter,
1990a; Chapter III), so even if these were bonafide trehalase mutants it is possible that lack of rhizobial trehalase activity due to a mutation would have no effect on the symbiosis. 120
It is somewhat intriguing that it was not difficult to obtain mutants of Rj. L. bv. nhaseoli USDA 2667 that accumulated reduced amounts of trehalose, but that no completely trehalose-negative strains were obtained. It is possible that trehalose production is essential for some aspect of rhizobial metabolism, so that any mutants that could not produce trehalose were non-viable. However, it is also possible that there are multiple copies of the trehalose metabolic genes. Other strains of Rj. jU. bv. phaseoli and other rhizobia have been identified with multiple copies of various genes and unidentified DNA
(Quinto et al.. 1982; Prakash and Atherly, 1984; Martinez et al.. 1985; Flores et al.f 1987; Rodriguez-Quinones et al..
1987; Appelbaum et al.. 1988), and more than one copy may function simultaneously (Romero et al.. 1988; Morett et al..
1988).
It is also possible that the osmotic stress screen was not appropriate for selection of trehalose-negative mutants.
One alternative outcome of the osmotic stress screen could have been the inadvertent selection of mutants that could not synthesize or accumulate some osmoticum other than trehalose; K+, glutamate and proline all have been implicated in osmotic tolerance in rhizobia and other bacteria (Hua et al.. 1982; Yap and Lim, 1983; Botsford,
1984; Somero, 1986; Larsen et al.. 1987; Dinnbier et al..
1988). In this scenario the mutants that grew poorly on the 121 high salt medium but nevertheless had normal trehalose accumulation patterns (93/110 tested; Table 10) could have been deficient in some osmotic tolerance mechanism other than trehalose accumulation. Nevertheless, because some mutants were successfully identified with lowered trehalose accumulation ability by using the salt stress screening procedure, it seems to have been potentially capable of identifying a trehalose-less mutant. Furthermore, this technique has been successfully used for selection of trehalose-negative mutants in E^. coli (Giaever et al. .
1988), a species which also has several different osmotic tolerance mechanisms.
The complementation experiment is difficult to interpret: only a few cells gained the ability to grow on osmotically stressful media containing tetracycline HC1, yet they did not produce detectable amounts of trehalose. They did not grow as well as the wild type cells, but they clearly had acquired a far higher tolerance to hyperosmotic stress than they had before complementation. Furthermore, they were unable to grow in hyperosmotic media if not kept under antibiotic pressure (presumably due to loss of the recombinant plasmid), indicating that the transferred rhizobial DNA cosmids did indeed confer the newly acquired salt tolerance. It is possible that a rhizobial gene for glutamate or proline synthesis under osmotic stress had been acquired, and was sufficient to confer osmotolerance. A similar problem was encountered by de Bruijn et al.
(1989)? using a rhizobial library and the same promoter
(pLAFRl) as in the current study, they successfully restored the wild-type phenotype to an EL. coli alnA (glutamine synthesis) mutant by complementation, but could not detect glutamine synthetase (GS) activity. They determined that all the complemented cells contained the same cosmid, cloned this gene (glnT), and found that it was different from the two previously known genes for glutamine synthesis in rhizobia (glnA and glnll). They attributed their inability to isolate glnA by complementation to the fact that the rhizobial gene apparently has an ecoRI site within the coding region and consequently was inactivated in the process of constructing the library cosmids. Apparently only the EL. coli mutants receiving cosmids carrying glnT were able to grow without added glutamine, however, they do not address why no GS activity was detected in these cells or why glnll was not functional. The current study could have been thwarted by the same type of problem: either the gene of interest was disrupted in the process of constructing the library or genes other than those for trehalose synthesis were capable of restoring the osmo- tolerant phenotype.
To pursue the possibility that multiple copies of trehalose-metabolisra genes might be present in rhizobia, two genes of trehalose metabolism in EL. coli were used to probe the genomes of two species of rhizobia. However, no binding occurred, even when hybridization was performed in 50% formamide at 37°C. Guerinot and Chelm (1984) also were unable to get binding of rhizobial DNA to E^. coli DNA that coded for the same enzyme function, and they concluded the two genes were very divergent. They isolated the adenylate cyclase genes from each species and attempted to hybridize them, although they had an advantage over the current study in that they had demonstrated adenylate cyclase activity in the cloned DNA from each species prior to binding experiments. In the case at hand, in addition to the possible lack of homology between the probes and the rhizobial genes, it is also possible that the E^. coli genes used for probes do not exist in rhizobia. However, Streeter
(1989) found some periplasmic trehalase activity in R. 1. bv. phaseoli USDA 2667, and it also has been shown that this strain has increased trehalose concentration when under osmotic stress (Hoelzle and Streeter, 1989; Chapter II), so the same metabolic capabilities apparently do exist in both
L. coli and the rhizobia used for these experiments.
As mentioned above, it is possible that the genes for periplasmic trehalase and osmotic trehalose synthesis do exist in rhizobia but are divergent enough in homology from those in E^. coli that no DNA heteroduplex could form during hybridization. According to an equation for estimating the melting temperature of a DNA duplex, in a 50% formamide 124 hybridization solution the melting temperature for the binding of the JjL. coli probes to E^. coli DNA would be 79° and the rhizobial probe/rhizobial DNA melting temperature would be 84°C (Sambrook, 1989; equation 1).
equation 1 :
Tm = 81.5 - 16.6 (log1 0 [Na+]) + 0.41 (%G+C) - 0.63 F - 600/L where [Na+] = sodium ion concentration in hybridization
solution, 0.3 M in Appendix F protocol
% G + C = percent guanosine plus cytosine in the DNA,
51% in E_s_ coli. 61% in Rj. meliloti JJ1
(Caudry-Reznick et al.. 1986)
F = percent formamide in hybridization solution
L = length of hybrid in base pairs, 2 2 0 0 for
otsA. 2500 for treA, and 8700 for node
Thus the temperature for the formamide hybridizations was well below the melting point of a homologous hybrid, however, the DNA binding of interest was the annealing of the Ei. coli probes to rhizobial genomic DNA, which would require non-homologous association. It has been estimated that the melting temperature of a DNA/DNA heteroduplex decreases about 1°C for every percent decrease in homology
(Lewin, 1980), so a hybridization temperature of 37°C should allow for 42% - 47% non-homology (using equation 1 for rhizobial or E^. coli DNA melting temperatures) between the two genes before binding would be prohibited. This may seem like quite a wide margin, however, many genes considered evolutionarily related are less than 50% homologous. A family of transcriptional activators which is found in both gram- and gram+ bacteria (and includes the rhizobial nodD gene) share only about 45% DNA homology (Schell and
Sukordhaman, 1989; Renault et al.. 1989), and several rhizobial regulatory proteins (considered to be related) such as NtrC, NifA and DctD, and other similar gene products from E_s_ coli. Klebsiella pneumoniae and Pseudomonas svrinaae share only about 40% protein homology (Drummond et al..
1986; Grimm and Panopoulos, 1989). In these studies there is a conspicuous lack of Southern blots showing DNA between these functionally related genes; it seems probable that there was insufficient homology to demonstrate such binding.
Based on these considerations it appears that the trehalose metabolism genes cloned from Ej. coli either do not exist in rhizobia or are too divergent to produce a stable DNA het erodup1 ex. 126
Plate I. Genomic DNA from Rj_ JL-. bv. phaseoli USDA 2667 and CFN42, Ri. meliloti USDA 1021 and Ej. coli probed with
labeled E_j. coli otsA DNA. Films were intentionally
overexposed to demonstrate lack of binding to
rhizobial DNA. Plate I
IL:. lj. bv. phaseoli CFN42
R. meliloti USDA 1021
R-_ 1^ bv. phaseoli USDA 2667
E. coli
markers 128
Plate II. Genomic DNA from Rj, L bv. phaseoli USDA 2667 and
CFN42, Rj. meliloti USDA 1021 and EL. coli probed with
labeled EL. coli treA DNA. Films were intentionally
overexposed to demonstrate lack of binding to rhizobial DNA. Plate II
R. 1. bv. Phaseoli CFN42
R. meliloti USDA 1021
R. 1. bv. Phaseoli USDA 2667
E. coli
markers 129 130
Plate III. Genomic DNA from R^. bv. phaseoli USDA 2667 and
CFN42, Ri meliloti USDA 1021 and E^. coli probed with
labeled R^. meliloti USDA 1021 node DNA. The membrane
used for Plate I was stripped and re-probed with the
rhizobial probe to demonstrate that the rhizobial DNA
could bind DNA of sufficient homology. Plate III
R;. 1^ bv. phaseoli CFN42
L. meliloti USDA 1021
E l I*, bv. phaseoli USDA 2667
E. coli
markers m 132
Plate IV. Genomic DNA from R±. bv. phaseoli USDA 2667 and
CFN42, R^. meliloti USDA 1021 and EL. coli probed with
labeled R^. meliloti USDA 1021 node DNA. The membrane
used for Plate II was stripped and re-probed with the
rhizobial probe to demonstrate that the rhizobial DNA
could bind DNA of sufficient homology. Plate IV
I R. 1. bv. Phaseoli CFN42
R. meliloti USDA 1021
R. 1. bv. Phaseoli USDA 2667
E. coli
markers
H* f-5 W 134
TABLE 10. Summary of trehalose accumulation patterns in
selected1 Tn5 mutants2 of Ri. L. bv. phaseoli USDA 2667.
Selection Number Number with Number With Medium: Growth Tested Low Trehalose High Trehalose
High Salt:
Poor Growth 1 1 0 17 2
Superior Growth 18 1 3
Trehalose:
Poor Growth 40 1 3
Superior Growth 34 1 0 2
1. Approximately 6000 mutants were screened using these media.
2 . 1 -2 % auxotrophs (poor or no growth on minimal media).
3. Mutants were grown in succinate medium and analyzed for trehalose accumulation relative to wild-type cells. CHAPTER VI
GENERAL DISCUSSION
Several conclusions can be drawn from the experiments documented in this dissertation:
1) Osmotic stress induces some, but not all, rhizobia
to accumulate more trehalose than is present under non-
osmotically stressful conditions.
2) The ions K+, Rb+, and Ntf1* stimulate the
a-glucosidases from members of the genus Rhizobium and
in Aarobacterium tumefaciens. but not from other
sources.
3) The a-glucosidase activity present in legume nodules
is probably of plant origin.
4) Low 02 tension induces increased trehalose
accumulation in rhizobia.
The most interesting conclusion from these facts is the combination of the lack of rhizobial. a-glucosidase activity
135 136 in the nodule in concert with increased rhizobial trehalose accumulation in low 02 environments. This suggests that the marked increase in trehalose concentration in nodules is due to the inhibition of rhizobial trehalase activity under the low 02 conditions found in infected cells. It also suggests that under conditions of high 0 2 availability there is a constant turnover of trehalose, because this would be necessary for the buildup of trehalose upon trehalase inhibition. The reason for this turnover is not clear.
Alternatively, trehalose synthesis could also increase during periods of low 0 2, in addition to inhibition of a- glucosidase activity. The lack of rhizobial a-glucosidases in the nodule also implies that all the sucrose and maltose
(in addition to trehalose) are unavailable to the bacteroids until hydrolyzed by plant enzymes. This may be a mechanism for control (by the plant) of carbon flow to the bacteroids.
It is unfortunate that no trehalose-negative mutants were obtained. The data available (inability to obtain a trehalose-negative mutant and the numerous reports in the literature documenting gene reiteration in various rhizobia) suggest that there are multiple copies of the genes necessary for trehalose metabolism but this has not been conclusively demonstrated at this point.
The role of trehalose in legume nodules remains unclear. The response to 02 status suggests that trehalose may function in adjustment to the low 0 2 environment found in the nodule, perhaps in a manner similar to its role as a membrane stabilizer in desiccated anhydrobiotic organisms.
The precise function of trehalose in this model is not clear. However, the adaptation of previously aerobic enzyme and membrane systems to microaerobic conditions most likely involves at least some degree of molecular (re)stabilization along with greatly altered physiology. Alternatively, trehalose may interfere with plant cell wall synthesis (as suggested by Veluthambi et al.. 1981, 1982a, 1982b) within the nodule. In this scenario the plant's tendency to wall off intruding organisms (more or less the "hypersensitive response") could be circumvented by altering the structure of the developing cell walls such that the nodule environment could be maintained in a condition favorable to the bacteroids. Another idea that draws from the same "cell wall interference" hypothesis is the possible secretion of trehalose immediately after binding of a free-living rhizobium to a receptive host root hair, causing root hair curling by stopping cell wall deposition in the locale of the bacterium but not on the distal side of the root hair.
The "beneficial" effects of trehalose on plant cell wall deposition could also aid the rhizobia as they penetrate the cell wall and travel through the developing infection thread, which also is bounded by a cell wall. Still another timepoint in the life cycle of rhizobia during which trehalose might be important is the free-living phase. 138
Perhaps the strains identified in Chapter II that accumulated extra trehalose under osmotic stress require this ability for survival in the drying conditions of the soil, but other strains and species of rhizobia have other osmotic tolerance mechanisms. This idea does not address the occurrence of trehalose in the strains that did not utilize it for an osmoticum. However, it is possible that there is more than one role for trehalose in these organisms. It also is possible that there are uses for trehalose in the free-living state other than osmotic tolerance.
While many interesting questions remain about the occurrence of trehalose in rhizobia, unfortunately many of the most direct experiments are currently unfeasible. If new data concerning the trehalose metabolism genes of E. coli are published, new experiments (such as complementing a trehalase mutant of EL*. coli) may become possible.
Alternatively, use of a different rhizobial library (perhaps from Ri. meliloti. or constructed using a different restriction enzyme) to complement the E^. coli otsAB deletion mutant might prove successful. Further work with the 02 phenomenon is another attractive avenue: perhaps the genes that are sensitive to 0 2 stress could be identified by use of lac fusions or some similarity to other (already identified) 0 2-sensitive rhizobial genes, and the genes associated with trehalose metabolism identified among them. Further work with the strains that accumulate extra trehalose under osmotic stress might identify which genes
are switched on (or off) under those conditions, and an
analysis similar to Rod et al. (1988) might reveal a similar system of mutations with amber supressors in the strains that do synthesize trehalose under osmotic stress. Finally, the occurrence of trehalose dimycolates in the cell walls of many mycobacteria and other actinomycetes suggests that the
N2-fixing actinomycete Frankia sp. may contain the same compounds. Because these mycolates are responsible for antigenic responses in some diseases, it seems possible that they might have a role in recognition or establishment of Frankia nodules. APPENDIX A
CHROMATOGRAPHY TECHNIQUES
140 141
FRACTIONATION OF ORGANIC ACIDS, AMINO ACIDS AND SUGARS
1. Soak 10 g Sephadex QAE-A25 in 500 ml 0.5 M Na-formate for
48 hours, change solution once and stir occasionally.
Collect Sephadex by vacuum filtration.
2. Store treated Sephadex QAE-A25 in 0.05 M Na-formate.
3. Soak 10 g Sephadex SP-C25 in 500 ml 0.5 M (NH4 )2 S04 for
48 hours. Change solution once and repeat soaking.
Collect Sephadex by vacuum infiltration.
4. Resuspend soaked Sephadex SP-C25 in 7% (v/v) formic acid,
set 2 hours, and decant. Repeat 3-4 times.
5. Store treated Sephadex SP-C25 in 1% formic acid (v/v).
6 . Fill small columns with 1.75 ml of treated Sephadex.
7. Rinse Sephadex QAE column with 5 ml 4% formic acid (v/v),
then with 7 ml H2 0.
8 . Rinse Sephadex SP column with 7 ml 7% formic acid (v/v),
then with 7 ml H2 0.
9. Arrange the Sephadex SP column so it is above the
Sephadex QAE column and will drain into it.
10. Add 100 /zl tartaric acid (331.6 ng/100 fil) for organic
acid internal standard, and 1 0 0 / * 1 norleucine
(41 ng/100 nl) for amino acid standard, to top column.
11. Place bottle for sugar (neutral) fraction below columns.
13. Add sample to top column.
14. Add 0.5 ml H2 0. Repeat. Add 7 ml H2 0.
15. When liquid stops flowing separate columns.
16. Place vials under the columns for collection of amino 142 and organic acid fractions.
17. Elute amino acids from the Sephadex SP column by adding
10 ml 0.2 M NH4 0H.
18. Elute organic acids from the Sephadex QAE column by
adding 15 ml 4% formic acid (v/v).
19. Dry all fractions under an air stream.
20. Rinse columns with storage solutions, return to bottles
for re-use. 143 HPLC ANALYSIS OF ORGANIC ACIDS
1. Dissolve an aliquot of the organic acid fraction from
Sephadex column chromatography in 2.5 ml H2 0.
2. Titrate the solution to pH 7.0-7.5 with 0.15 or 0.30 N
KOH (not NaOH).
3. Add a few drops of chloroform and air dry sample.
4. Add 0.5 ml H20 to dissolve sample. Transfer 250 /tl to a
10 cm screw-cap tube. Re-dry remaining sample.
5. Add 250 /il phosphate buffer and 1.5 ml esterification
reagent to the sample and to 250 /il standard (Appendix
H).
6 . Close tube very tightly and set in boiling H20 bath for 40 minutes.
7. Cool tube on bench. Do not use ice bath.
8 . Filter reaction mixture through 0.45 /im syringe filter
into 4 ml vial and seal with septum cap.
9. Prepare Waters HPLC for analysis of organic acids:
WISP settings:
injection volume = 15 /il
run time = 2 7 minutes
equilibration delay = 8 minutes
Variable wavelength detector settings:
wavelength = 245 nm
absorbance units on full scale = 0.5 channel = l
Gradient controller settings; time flow %A %B curve
initial 1.0 ml/min 90 10 none
26 min 1.0 ml/min 30 70 6
27 min 1.0 ml/min 30 70 6
28 min 1.0 ml/min 90 10 6 Eluants:
A = 35% HPLC methanol/65% HPLC H20 (degassed)
B = 100% HPLC methanol (degassed)
Column:
8 mm X 10 mm Waters Novapak C-18 steel
temperature = 40°C
Waters 740 Data Module:
attenuation = 1
Load samples and standard into WISP, run standard
first and last to be sure times haven't changed. GAS CHROMATOGRAPHIC ANALYSIS OF CARBOHYDRATES
Dry an aliquot of an ethanolic extract or of the neutral
fraction from Sephadex column chromatography. Use a
4 ml glass vial. Dry a standard also (Appendix H).
Add 200 /il "Stox" reagent to the sample vial and to a
standard. Cap with a teflon septum. Vortex thoroughly.
Incubate in a heating block for 30 minutes at 70°C.
Vortex occasionally if solids are not completely dissolved.
Cool to room temperature. Add 200 /il "HMDS" reagent and
25 /il trifluoro-acetic acid. Vortex 30 seconds.
Let mixture stand for 30 minutes.
Prepare Varian 3700 Gas Chromatograph:
Gas flow rates: Helium 60 psi = 30 ml/minute
Hydrogen 30 psi = 30 ml/minute
Air 40 psi = 300 ml/minute
Column: 6 ft. X 2.0 mm I.D. glass
Support: 3.0% OV-17 on 80/100 Chromosorb W HP
Injection temperature: 280°C
Detector temperature: 330°C
Temperature program for nodule carbohydrates:
initial temperature 150°C for 8 minutes,
increase to 240"C at 15°C/minute,
hold at 240°C for 6 minutes
Temperature for trehalose detection only:
isothermal at 240#C 146 7. Inject 1 n1 of sample. Hit start button on GC if using
programmed temperature protocol.
8 . Start HP 3392A integrator. Attenuation set at 2.
9. Samples should be run within 2 or 3 hours of reacting. APPENDIX B
BACTERIAL TRANSFORMATION WITH PLASMID DNA
147 148
BACTERIAL TRANSFORMATION
1. Inoculate 50 ml LB broth (Appendix H) with Ea. coli
MV1190. Grow at 37°C with shaking to A5 5 0 0.4-0.5.
2. Centrifuge at 6000 RPM at 4°C in cold 30 ml centrifuge
tubes for 1 0 minutes.
3. Resuspend in 25 ml cold 0.1 M MgCl2. Set on ice for 20
minutes.
4. Centrifuge as in step 2.
5. Resuspend in 5 ml cold 0.1 M CaCl2. At this stage cells
are competent for up to 24 hours if kept on ice.
6 . Set 13 X 100 mm borosilicate tube in ice and add 20 fil
T 1/10 E (Appendix H) and approximately 2 ng plasmid.
Also prepare control tube without DNA.
7. Add 200 fil of competent cells in CaCl2 to borosilicate
tube. Set on ice for 40 minutes.
8 . Heat shock cells at 42°C for 120 seconds (use timer).
9. Transfer to 15 ml tube and add 2 ml LB broth. Incubate
at 37"C for 1 hour without shaking.
10. Centrifuge 10 minutes at 2000 RPM in table top
centrifuge.
11. Resuspend pellet in 200 fil LB broth.
12. Spread 100 fil of cells on antibiotic plate. Also spread
some plates with 1 0 0 fil diluted cells.
13. Invert plates and incubate overnight at 37°C. 150
PLASMID DNA MINI-PREP
1. All solutions, tubes, and pipet tips should be sterile.
2. Culture plasmid-containing bacterium overnight in 5 ml
liquid LB medium (Appendix H) with appropriate
antibiotics.
3. Transfer 1.5 ml to Eppendorf tube, spin in tabletop
centrifuge for 1 minute at top speed. Remove all
supernatant fraction with a pipet.
4. Suspend pellet in 180 /il ice cold solution 1 (Appendix
H). Incubate 5 minutes at room temperature.
5. Add 20 /tl lysozyme (10 mg lysozyme/ml in cold solution 1
from step 4). Vortex gently, incubate 5 minutes at
room temperature.
6 . Add 400 /il solution of 0.2 N NaOH/1% SDS. Mix by
inverting, do not vortex. Incubate 5 min on ice.
7. Add 300 /tl cold 3 M sodium acetate. Vortex gently,
incubate on ice 5 min.
8 . Spin 3 min in microcentrifuge at top speed, transfer
supernatant fraction to fresh tube, re-spin and
again transfer supernate to a fresh tube.
9. Add 500 /il cold isopropanol. Set in -20°C freezer 20
minutes (or longer).
10. Centrifuge in cold microcentrifuge 15 min at top speed.
Discard supernate.
11. Wash pellet with 100 /il cold 70% ethanol. Spin 3 min
at top speed, remove ethanol with pipet. 151 12. Dry in vacuum centrifuge for 10 minutes or air dry 1
hour.
13. Resuspend pellet in 50 ill TE pH 8.0 (Appendix H) . ISOLATION OF GENOMIC DNA FROM RHIZOBIA
Grow rhizobia in YEMG or TY medium (Appendix H) to A^oo
0.5-1.0. Cultures with lower A^oo maY be difficult to lyse.
Centrifuge 15 ml of culture in a refrigerated centrifuge
for 5 minutes at 10,000 RPM (12,000 X g). Use sterile
30 ml tube. Discard supernatant fraction.
Wash cells with 5 ml ice cold TE/0.1 M NaCl (Appendix H).
Set on ice 5 minutes, centrifuge 5 minutes at 10,000
RPM.
Repeat step 3.
Suspend in 375 /il TE in microcentrifuge tube.
Add: 50 /il of 20 mg/ml fresh lysozyme solution
50 /tl of 20 mg/ml proteinase K stock solution
25 /il O f 20% SDS
Vortex very gently. Incubate 3 hours at 37°C.
Shear DNA by passing mixture in and out of a syringe
fitted with a 22 gauge needle. Repeat 2 or 3 times.
Add 500 /il Tris-saturated phenol, shake. Centrifuge in
microcentrifuge 2 minutes at top speed.
Transfer the top layer to a fresh microcentrifuge tube
using an automatic pipet. Do not transfer any of the
white interface. It helps to cut the end of a pipet
tip off at an angle; the opening can be held against
the edge of the tube while collecting the upper layer
and decreases the tendency for the interface to enter 153 the pipet tip.
10. Repeat steps 8 and 9 at least 2 times. The white
interface should become negligible.
11. Add an equal volume of chloroform and extract as with
phenol in steps 8 and 9.
12. Precipitate DNA by adding an equal volume of cold
isopropanol. Set in freezer for 20 minutes (or
longer).
13. Spin in microcentrifuge 10 minutes at top speed.
14. Add 100 nl cold 70% ethanol. Do not resuspend pellet.
15. Centrifuge 3 minutes at top speed.
16. Pour off ethanol, let drain thoroughly. Air dry 1 hour. 17. Suspend pellet in 50 /il TE. RESTRICTION DIGESTION OF DNA
Select restriction enzyme. For genomic digests ecoRI is
usually used.
Put 0.5-16 /il DNA solution in microcentrifuge tube.
Amount varies depending on concentration of DNA and
type of gel to be run after digestion.
Add H20 so that DNA + H20 = 16 /il.
Add: 2 /il 10X digestion buffer appropriate for enzyme.
1 /il of 1 mg/ml RNase
1 /il restriction enzyme ( 1 0 units)
Incubate at 37°C for 2-6 hours. APPENDIX D RUNNING AGAROSE GELS
155 RUNNING AGAROSE GELS
Dissolve agarose in tris-acetate buffer (Appendix H),
0 .8 -1 .2 % w/v.
Pour into gel tray with well-former. Amount varies,
mini-gels are 3-4 mm deep, 10 X 14 cm gels are 5-10 mm
deep- if the gel is to be used for a Southern blot be
careful not to add so much agarose that a ridge forms
by capillary action under the well former.
Remove well-former when gel is solid. Place in gel in
electrophoresis box.
Add tris-acetate buffer until gel is submerged.
Dilute hindlll-digested lambda phage DNA in lambda
buffer (Appendix H). Incubate at 65°C for 5-10 minutes
to separate fragments.
Add 1/10 volume gel-loading buffer (Appendix H) to DNA
samples. Mix and spin briefly to be sure all solution
is in bottom of tube.
Load samples into wells with automatic pipet. If gel is
to be used for a Southern blot leave empty lanes
between loaded lanes. Record lane order of samples.
Connect electrodes and turn on voltage. Remember wells
should be oriented so the DNA can "run to red"
electrode! 10 X 14 cm gels can run at 22 volts for
about 14 hours, mini-gels at 45 volts for 1.5-2.0
hours.
Stain gel with ethidium bromide (25 p1 of 10 mg/ml stock APPENDIX C
ISOLATION AND RESTRICTION DIGESTION OF DNA
149 157
solution/1) for 10-20 minutes. Always wear gloves when
handling ethidium bromide-it is toxic and mutagenic.
10. Examine gel under UV light. Wear protective glasses and gloves.
11. Photograph gel using UV light and a red filter. APPENDIX E
SOUTHERN TRANSFER OF DNA TO A MEMBRANE
158 DNA TRANSFER TO A MEMBRANE
(Acid Depurination Protocol for Gel Denaturation)
1. Run 10 x 14 cm gel in tris-acetate buffer (Appendix H),
5-10 mg DNA per lane. Run markers and leave empty
lanes between loaded lanes.
2. Prepare 1 1 of each soaking solution (steps 5, 7 & 9).
3. Stain gel in ethidium bromide (25 /il of 10 mg/ml stock
EtBr solution/1 tris-acetate buffer) for 20-30 min.
Photograph gel. EtBr is toxic and mutagenic-always wear
gloves when handling.
4. Rinse gel with H20.
5. Soak 15 min in 500 ml 0.2 N HC1 in cake pan on rotator
table. Only gentle rotation is needed. Rinse with H20.
6 . Repeat step 5.
7. Soak 30 min in 500 ml 0.5 M NaOH/1.5 M NaCl in cake pan
on rotator table. Rinse with H20.
8 . Repeat step 7.
9. Soak 40 min in 1 M NH^OAc/O.l M NaOH in cake pan on
rotator table in fume hood, rinse with H20.
10. Blot by capillary transfer overnight in fume hood in
1 M NfyOAc/O.l NaOH as follows:
11. Place four 15 x 22 cm strips of filter paper over 15 x
15 cm platform in cakepan. Strips should criss-cross
so all 4 sides of platform have paper hanging into the
transfer solution. Wet filter papers with transfer
solution and smooth out any bubbles with test tube. 160
12. Place two 15 x 15 cm pieces of filter paper on platform.
Wet with transfer solution and smooth out bubbles.
13. Invert denatured gel onto filter paper. Remove bubbles.
14. Wet one 10 x 14 cm piece of Zeta-probe membrane with
H20, then with transfer solution. Place on gel.
Membrane should never be touched with bare hands-
including when cutting on paper cutter.
15. Wet two 10 x 14 cm filter papers with H20. Place on
membrane. Put approximately 10 cm stack of pre-cut
brown paper towels on filter paper.
16. Place weight (unopened Sigma catalog is ideal) on top of
paper towel stack.
17. Approximately 500 ml of transfer solution should be in
bottom of cake pan.
18. Let stand undisturbed overnight. DISASSEMBLING A DNA TRANSFER
Remove and discard paper towels and top 2 pieces of
filter paper.
Lift gel and membrane in one piece. Place them gel side
up (inverted from position during transfer) on Saran Wrap.
Mark gel edges and position of wells on membrane with
black ball-point pen. Also include date, your initials
and any other useful information. Other colors and
types of ink do not work well.
Remove and discard gel. Place membrane in 2X SSC
(Appendix H) and rub gently with fingers (wearing
gloves) to remove any remaining agarose.
Air dry membrane on kim-wipe.
Bake membrane 2 hours at 80°C under vacuum. APPENDIX F
NUCLEIC ACID HYBRIDIZATION
162 AQUEOUS DNA/DNA HYBRIDIZATION
Place baked membrane in a Seal-a-Meal freezer bag.
Add 30 ml pre-hybridization mixture:
18.5 ml H20
5X SSC (7.5 ml 2OX) (Appendix H)
2X Denhardt's reagent (0.6 ml 100X) (Appendix H)
0.2% SDS (0.3 ml 20%)
0.05 M Tris pH 8.0 (1.5 ml 1 M)
0.01 M Na2EDTA (0.6 ml 0.5 M)
1 ml salmon sperm (10 mg/ml, boil 5 min, set on ice)
Remove bubbles and seal bag with heat sealer.
Incubate at 65°C 4-8 hours.
Add boiled, labeled probe and boiled salmon sperm DNA to
29 ml hybridization solution:
labeled probe
6.0 ml H20
10% dextran sulfate (12.5 ml 25%)
Other ingredients same as in pre-hybridization mixture.
Open pre-hybridization bag and remove membrane. Place
membrane in fresh Seal-a-Meal bag.
Add 30 ml hybridization solution to bag. Heat seal
immediately, seal again about 1 cm behind first seal.
Push any bubbles to top of bag, make new heat seal
separating bubbles from the membrane.
Incubate at 65°C 8-24 hours. 164
FORMAMIDE DNA/DNA HYBRIDIZATION:
1. Protocol is the same as for an aqueous Southern blot
except the solutions are 50% formamide and the
incubations are done at 37°C or 42°C instead of 65°C,
and for longer times- up to 2 days.
Pre-hybridization Mixture:
3.5 ml H20
15.0 ml deionized formamide
other ingredients as in aqueous pre-hybridization
Hybridization Mixture:
3.0 g dextran sulfate (dissolve in H20 and formamide) 3.5 ml H20
15.0 ml deionized formamide
other ingredients as in aqueous hybridization. LABELING RANDOM PRIMED DNA PROBE
Always wear double gloves when handling radioactivity!
Isolate insert from plasmid using "Geneclean".
Boil approximately 25 ng of the DNA to be labeled for 5
minutes, in small microcentrifuge tube. Set on ice.
Spin quenched, denatured DNA briefly in microcentrifuge.
Add to denatured DNA (in this order):
11.5 /il low salt buffer (Appendix H)
1.0 /i 1 BSA (10 mg/ml)
3.0 /il 32P-dCTP (60 /iCi)
0.5 /il Klenow fragment of DNA polymerase I
note: DNA + dCTP + H20 = 12.0 /nl, total reaction
mixture volume must be 25 /tl (or a multiple of 25) .
Incubate 2 hours at room temperature.
Rinse Sephadex G50 column NICK Column (Pharmacia Inc.,
Piscataway, NJ) with several volumes TE (Appendix H).
Stop labeling reaction with 5 /il 0.5 M Na2EDTA. Add to
top of column. Rinse reaction tube with approximately
50 /il TE and add to column. Maximum volume of reaction
mixture + TE rinse is 125 /il.
Add 400 /il TE to column. Collect and save effluent in
microcentrifuge tube until it is clear that no label
has accidentally come through, then discard.
Add another 400 /il TE to column, collect effluent in
large microcentrifuge tube. This fraction is the
radioactive probe. 166 10. Check how radioactive the probe is with geiger counter.
11. Boil the probe and 1 ml of 10 mg/ml salmon sperm DNA
for 5 minutes. Quench on ice.
12. Add probe to hybridization mixture. Use salmon sperm
DNA to rinse the tube that contained the probe and add
to hybridization mixture.
11. Rinse column with 4-5 volumes of TE and discard in
radioactive waste. Rinsings will contain
unincorporated nucleotides. APPENDIX G
BLOT WASHING AND STRIPPING
167 168
BLOT WASH
Use the same protocol for washing blots from aqueous or formamide hybridizations but heat the low stringency wash solution to the same temperature as the hybridization. High stringency wash solutions may be hotter than the hybridization temperature if desired.
LOW STRINGENCY WASH:
1. Heat 2 1 2X SSC/0.1% SDS (Appendix H) to desired
temperature. Set on hot plate with magnetic stirrer.
2. Cut one corner of Seal-a-Meal bag and drain off
hybridization solution into liquid radioactive waste or
into 50 ml sterile plastic tube if it is to be reused.
3. Cut bag open and remove blot. Place blot in cake pan and
add 400 ml heated wash solution.
4. Let set behind 32P shield for 2-3 minutes. Discard
solution into radioactive waste.
5. Add 400 ml more wash solution. Place in H20 bath or
incubator at appropriate temperature for 1 0 minutes.
6 . Pour rinse into radioactive beaker. Check for
radioactivity with geiger counter. If none is detected
the solution may be put in hot sink, otherwise discard
in radioactive liquid waste.
7. Repeat steps 5 and 6 three times.
8 . If no high stringency wash is to be performed, air dry
filter. Wrap with Saran Wrap. 169 HIGH STRINGENCY WASH:
1. Complete steps 1-7 in low stringency wash protocol. Do
not dry blotI
2. Heat 2 1 0.1X SSC/0.1% SDS (Appendix H) to appropriate
temperature.
3. Rinse blot with this solution as in steps 5 and 6 of low
stringency wash. This will be 5 washes.
4. Air dry filter. Wrap with Saran Wrap. 170
BLOT STRIPPING
1. Remove any Saran Wrap or tape from blot.
2. Place in 0.2 N NaOH for 30 minutes at room temperature
with gentle shaking on rotator table.
3. Soak in 0.1% SDS/0.1X SSC (Appendix H) for 30 minutes
at room temperature with gentle shaking.
4. Dry blot, wrap in Saran Wrap until ready to re-probe. APPENDIX H
RECIPES FOR MEDIA AND SOLUTIONS
171 172
RECIPES FOR SOLUTIONS
PHOSPHATE BUFFER FOR ORGANIC ACID DERIVITIZATION:
1.01 g KH2P04
2.86 g Na2P04 «12H20
H20 to 125 ml, pH 6 . 8
ESTERIFICATION REAGENT FOR ORGANIC ACID DERIVITIZATION:
7.50 g phenacyl bromide
0.37 g dicyclohexane-18-crown-6
acetone to 250 ml
ORGANIC ACID STANDARD (mg/ml) (store frozen):
1.250 K-malate
1.750 K-malonate
0.500 K-succinate
0.500 K-citrate
0.300 K-fumarate
1.000 K-transaconitate
0.375 K-benzoate
0.250 K-lactate
1.250 K-tartrate (internal standard)
use 200 til standard + 50 nl H20 (or other
standard) per derivitization 173
STANDARD FOR NODULE CARBOHYDRATE ANALYSIS (mg/ml):
1 . 8 sucrose
1 . 0 glucose
1 . 0 mvo-inositol
0 .4 fructose
0.4 trehalose
0.4 maltose
store frozen and use 1 0 0 /il per derivitization
CARBOHYDRATE STANDARD FOR CULTURED RHIZOBIA:
1 . 0 mg/ml trehalose
store frozen and use 1 0 0 /tl per derivitization
SOLUTION 1 FOR PLASMID MINI-PREP:
3.94 g Tris-Cl
9.0 g glucose
3.72 g Na2EDTA*2H20
HzO to 1 1, pH 8.0
50X TRIS-ACETATE ELECTROPHORESIS BUFFER:
242.00 g Tris-Cl
0.37 g Na2EDTA*2H20
57.10 ml glacial acetic acid HzO to l 1 174
LAMBDA BUFFER:
0.37 g Na2EDTA-2H20 (1 mM)
14.61 g NaCl (25 mM)
1.21 g Tris base (10 mM)
HzO to 1 1
GEL LOADING BUFFER:
IX Tris-acetate electrophoresis buffer
25% glycerol
100X DENHARDT'S REAGENT (Denhardt, 1966):
10 g Ficoll, type 400
1 0 g polyvinylpyrrolidone
5 g bovine serum albumin (fraction V) H20 to 500 ml
freeze at -20°C in 1 ml aliquots
LOW SALT BUFFER:
Solution 1: Solution 2: 100 /tM dATP 500 /il Solution 1
100 /iM dGTP 252 /xl random primer (Pharmacia)
100 /iM dTTP 250 /il 2M hepes buffer
250 mM Tris pH 8.0 118 /tl H20
25 mM MgCl2 store at -20°C
50 mM /3-mercaptoethanol 175
TE:
15.76 g Tris-Cl (10 mM)
0.37 g Na2EDTA*2H20 (1 mM)
H20 to 1 1, pH to 8.0
T 1/10 E:
15.76 g Tris-CL (10 mM)
0.037 g Na2EDTA«2H20 (0.1 mM) H20 to 1 1
SSPE (20X):
175.3 g NaCl
27.6 g NaH2P04 *H20
7.4 g Na2EDTA
H20 to 1 1, pH to 7.0 with NaOH
SSC (20X):
175.3 g NaCl
88.2 g NaCitrate
H20 to 1 1, pH 7.0 176
RECIPES FOR MEDIA
MINIMAL MEDIUM:
5.0 g gluconic acid
1 . 0 g sodium glutamate
0.8 g KN03
mineral salts (Appendix H)
micronutrients (Appendix H)
H20 to 1 1, pH 7.0
M63 MEDIUM:
13.6 g KH2P04
2.0 g (NH4 )2S04
0.5 mg FeS04• 7H20
26.3 g NaCl
H20 to 1 1, pH 7.0
Autoclave and cool, then add 1 ml each of sterilized
1 M MgS04 and CaCl2, and filter sterilize and add 40 mg arginine, and 15 mg tetracycline HC1, and 10 ml of 20% mannitol.
LURIA-BERTANI MEDIUM (LB):
1 0 . 0 g bacto-tryptone
5.0 g yeast extract
10.0 g NaCl
HjO to 1 1, pH 7.5 177 SUCCINATE MEDIUM:
Solution 1:
0.75 g CaCl2 «2H20
0.02 g Na2EDTA-2H20
1.00 g (NH4 )2S04
1.00 g NaCl
3.00 g sodium succinate
H20 to 960 ml, pH 7.0
Solution 2:
1.0 g MgS04 • 7H20 in 20 ml H20
Solution 3:
1.06 g K2HP04
0.53 g KH2 P04 in 20 ml H20
Autoclave and cool all solutions, then add 0.5 ml each of
solutions 2 and 3 per 50 ml solution 1, and add filter-
sterilized vitamins if needed. Some precipitate may form.
YEMG MEDIUM:
0.35 g K2HP04
0.35 g KH2P04
0.20 g MgS04 *7H20
0.10 g NaCl
0.50 g yeast extract
2 . 0 0 g sodium gluconate
1 0 . 0 0 g mannitol
H20 to 1 1, pH 7.0 178
TY MEDIUM:
5.0 g tryptone
3.0 g yeast extract
1.5 g CaCl2 *2H20
H20 to 1 1, pH 7.0
LOW SUGAR MEDIUM:
1 . 0 g yeast extract
mineral salts
micronutrients
H20 to 1 1, pH 7.0
HIGH SALT MEDIUM:
5.0 g arabinose
2 . 0 g sodium gluconate
0.5 g yeast extract
10.0 g NaCl
micronutrients
H20 to 1 1, pH 7.0
TREHALOSE MEDIUM:
2 . 0 g trehalose
0.8 g KN03
mineral salts
micronutrients
HzO to 1 1, pH 7.0 MINERAL SALTS FOR MEDIA:
g/ 1 0 0 ml stock ml stock/ 1 media
CaCl2 ‘2H20 0.066 10
MgS04 *7H20 1.0 10
K2HP04 1.8 10
FeS04 '7H20 0.013 1
1000X MICRONUTRIENTS:
1.45 g H3 BO3
0.05 g CuS04• 7H20
0.05 g MrC12• 4H20
1.08 g ZnS04 «7H20
2.50 g Na2Mo04 «2H20
0.10 g CoCl2 *6H20
4.00 g FeCl3• 6H20
5.50 g Na2EDTA*2H20
H20 to 1 1, use 1 ml/1 media LITERATURE CITED
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