The reasonable man attempts to adapt himself to suit the world: the unreasonable one persists in trying to adapt the world to suit himself. Therefore all progress depends upon the unreasonable man.
George Bernard Shaw
UNIVERSITY OF READING
School of Animal and Microbial Sciences
The General Amino Acid Permease of Rhizobium leguminosarum biovar viciae
by
David L. Walshaw
Submitted in partial fulfilment of the requirement for the degree of Docter of Philosophy 1995
I declare that this thesis is my own account of my research and that this work
has not previously been submitted for a degree at any University. However, I
would like to acknowledge the help I received from the undergraduate project
students Adam Wilkinson and Mathias Mondy in restriction mapping the
cosmid pRU3004, under my joint supervision with Dr P.S.Poole.
David Walshaw
TABLE OF CONTENTS
Page
CHAPTER 1 LITERATURE REVIEW 1
1.1 INTRODUCTION 2 1.1.1 Rhizobia 2 1.1.2 Nodule formation and structure 2
1.2 BACTEROID METABOLISM 5 1.2.1 Carbon sources supplied to the bacteroid 5 1.2.2 The TCA cycle in the bacteroid 8 1.2.3 The role of poly-β-hydroxybutyrate biosynthesis 13 1.2.4 The malate-aspartate shuttle 14 1.2.5 Other roles of amino acids in bacteroid metabolism 19
1.3 AMINO ACID TRANSPORT IN BACTERIA 27
1.4 ABC TRANSPORTERS 29 1.4.1 Overall structure of ABC transporters 29 1.4.2 The transmembrane domains 32 1.4.3 The ATP-binding domains 35 1.4.4 Periplasmic binding proteins 40 1.4.5 Mechanism of solute translocation 44 1.4.6 The role of binding protein-dependent transporters 49 1.4.7 Regulation of ABC transporters 49
1.5 AMINO ACID TRANSPORT IN RHIZOBIUM 51
1.6 REGULATION INVOLVING NTRC 53
CHAPTER 2 MATERIALS AND METHODS 58
2.1.1 Bacterial strains 59 2.1.2 Culture conditions 68 2.1.3 DNA and genetic manipulations 68 2.1.4 Mutagenesis 69 2.1.5 Transport Assays 70 2.1.6 Isolation of periplasmic fractions and protein gel
electrophoresis
71 2.1.7 Protein binding assays 71 2.1.8 Enzyme assays 72 2.1.9 Metabolite excretion assays 73 2.1.10 Intracellular concentrations 74
Page
2.1.11 Protein determination 75 2.1.12 Plant Assays 75
CHAPTER 3 THE CLONING AND CHARACTERIZATION OF THE GENERAL AMINO ACID PERMEASE OF RHIZOBIUM LEGUMINOSARUM STRAIN 3841
76
3.1 INTRODUCTION 77
3.2 RESULTS 78 3.2.1 Isolation of cosmid pRU3024 carrying the general
amino acid permease genes of Rhizobium leguminosarum strain 3841
78 3.2.2 Restriction mapping, sub-cloning and mutational
analysis of pRU3024
80 3.2.3 Nucleotide sequence of the 5.4kb MluI-ClaI
fragment of pRU135
85 3.2.4 Coding regions of the nucleotide sequence from
pRU189
97 3.2.5 Other features of the nucleotide sequence from
pRU189
106 3.2.6 Mutation of the general amino acid permease 107 3.2.7 Mapping of promoter sites in the aap operon by
complementation analysis
111 3.2.8 Transcription levels of aap genes 113 3.2.9 Amino acid uptake in strains RU542, RU543, RU634
and RU636
114 3.2.10 Growth of strain RU543 on amino acids as sole
source of carbon and nitrogen
116 3.2.11 Amino acid uptake in strain RU640 118 3.2.12 Expression of the R. leguminosarum general amino
acid permease in E. coli
118 3.2.13 Physical properties of the aapJ gene product 120 3.2.14 Effect of aapJ on amino acid uptake in 3841 122 3.2.15 Effect of aapQMP on amino acid uptake in strains
3841
123 3.2.16 Specificity of the aapJ gene product 124 3.2.17 Substrate-binding activity of AapJ 125
Page
3.2.18 Amino acid exchange 130 3.2.19 Plant properties of strains RU542, RU543, RU634,
and RU636
143 3.2.20 Nucleotide sequence of the 0.8kb BamHI fragment
of pRU3024
144 3.2.21 Amino acid transport in strain RU632 147 3.2.22 Plant properties of RU632 149 3.2.23 Nucleotide sequence adjacent to the transposon in
cosmids pRU3053, pRU3082, pRU3083, pRU3084, pRU3085 and pRU3086
150 3.2.24 Amino acid transport in metC mutants of strain 3841 154
3.3 DISCUSSION 155
CHAPTER 4 NITROGEN REGULATION OF THE GENERAL AMINO ACID PERMEASE OF RHIZOBIUM LEGUMINOSARUM STRAIN 3841
161
4.1 INTRODUCTION 162
4.2 RESULTS 163 4.2.1 Effect of the metC-aapJ intergenic region on growth
of strain 3841
163 4.2.2 Effect of nitrogen supply on the transcription of
aapJQM
164 4.2.3 Amino acid uptake in strain RU929 169 4.2.4 Effect of nitrogen supply on the transcription metC
and cysE
170 4.2.5 Sequence analysis of the metC-aapJ intergenic region 173
4.3 DISCUSSION 175
CHAPTER 5 INTER-REGULATION OF THE TCA CYCLE AND THE GENERAL AMINO ACID PERMEASE OF R. LEGUMINOSARUM STRAIN 3841.
177
5.1 INTRODUCTION 178
5.2 RESULTS 179
Page
5.2.1 Aspartate resistant mutants of R. leguminosarum strain 3841
179
5.2.2 Growth of strains RU116, RU118, RU137 and RU156 on succinate and glucose
183
5.2.3 Amino acid transport in strains RU116 and RU156 184 5.2.4 Transductional analysis of strains RU116, RU118,
RU137 and RU156
186 5.2.5 Nucleotide sequence adjacent to the transposon in
strains RU116, RU137 and RU156
187 5.2.6 Activity of TCA cycle enzymes in strains RU116,
RU118, RU137 and RU156
194 5.2.7 Growth of strains RU116, RU137 and RU156 on
arabinose
195 5.2.8 Complementation of strain RU156 196 5.2.9 Effect of pRU3004 on aspartate transport in strains
RU116, RU137, and RU156
197 5.2.10 Southern blot of pRU3004 against RU116, RU137,
RU156 chromosomal DNA
198 5.2.11 Restriction mapping, sub-cloning and mutation of
pRU3004
199 5.2.12 Genes carried by pRU3004 201 5.2.13 β-galactosidase activities from pRU3004 mutants 210 5.2.14 TCA cycle enzyme activities in sucCDAB mutants of
strain 3841
210 5.2.15 Mapping of promoter sites in pRU3004 211 5.2.16 Amino acid excretion by strains RU116, RU156 and
RU543
215 5.2.17 Intracellular concentrations of α-ketoglutarate and
glutamate in strains RU116 and RU156
220 5.2.18 Transcription of the aap operon in sucDA mutants of
strain 3841
222 5.2.19 Plant properties of strains RU116, RU137 and
RU156
224
5.3 DISCUSSION 226
Page
CHAPTER 6 FINAL DISCUSSION 231
6.1.1 The general amino acid permease of Rhizobium leguminosarum
232
6.1.2 TCA cycle enzymes in Rhizobium leguminosarum 237 6.1.3 Methionine biosynthetic enzymes in Rhizobium
leguminosarum
238 6.1.4 Future work 239
REFERENCES
241
ABSTRACT
The four genes, aapJQMP, encoding the general amino acid permease of R.
leguminosarum strain 3841 have been cloned, sequenced, and shown to be
transcribed as a single operon. Sequence homology data indicate that these
genes encode the components of an ABC transporter. However, the
periplasmic binding protein, AapJ, and the two integral membrane
components, AapQ and AapM, are significantly larger than the equivalent
components of previously described ABC transporters of amino acids. The
strong homology of these proteins to sequence from the Escherichia coli genome
sequencing project suggests that E. coli may possess a previously unreported
general amino acid permease. Transcription of the aap operon has been shown
to be negatively regulated by NtrC in response to nitrogen supply.
Mutation of any one of the aap genes resulted in a reduction in the uptake by
strain 3841 of a range of amino acids, including aliphatic amino acids such as
leucine and alanine, and polar amino acids such as glutamate, aspartate and
histidine. Over expression of the aap operon resulted in a marked increase in
the uptake of all the amino acids tested. The results of experiments to
investigate the effect of mutation and over expression of aap genes on amino
acid exchange by strain 3841, appear to indicate that the general amino acid
permease facilitates both uptake and efflux of amino acids. The involvement of
the general permease in amino acid efflux is also indicated by the reduced
glutamate excretion during growth on glucose/NH4Cl/aspartate, exhibited by
an aapJ mutant of strain 3841.
An attempt to isolate general amino acid permease mutants on the basis of
resistance to a toxic concentration of aspartate, led to the discovery that
mutation of genes encoding the TCA cycle enzymes α-ketoglutarate
dehydrogenase and succinyl-CoA synthetase causes almost total abolition of
uptake by the general amino acid permease of strain 3841. This effect has been
shown not to be due to regulation of aap gene expression at the transcriptional
level. Nor does it appear to be the result of substrate-inhibition of the
transporter, despite the fact that α-ketoglutarate dehydrogenase and succinyl-
CoA synthetase mutants are found to accumulate and excrete glutamate. It is
therefore proposed that the general amino acid permease may be subject to
post-translational modification.
The chromosomal region containing contiguous genes that code for malate
dehydrogenase, succinyl-CoA synthetase and α-ketoglutarate dehydrogenase,
has been cloned and partially sequenced.
Mutants in any one of the components of the general amino acid permease
were found to induce pea root nodules that reduce acetylene as effectively as
those of the wild-type strain. α-Ketoglutarate dehydrogenase and succinyl-
CoA synthetase mutants of strain 3841 formed ineffective nodules.
1
CHAPTER 1 LITERATURE REVIEW
2
1.1 INTRODUCTION
1.1.1 Rhizobia
Bacteria of the genus Rhizobium stimulate leguminous plants to develop root
nodules which the bacteria infect and inhabit. Ultimately, the two organisms
establish metabolic co-operation: The bacteria reduce (fix) molecular nitrogen
to ammonia, which is exported to the plant for assimilation; the plant reduces
carbon dioxide to sugars during photosynthesis and translocates these to the
root where they are used to provide the bacteria with a carbon and energy
source.
This dramatic symbiosis is both ecologically and agronomically significant,
leguminous root nodules being by far the largest single source of organic
nitrogen in the global nitrogen cycle. However, the system has an additional
attraction as an area for study. During a complex series of developmental steps,
the bacteria and the plant each influence in the other such fundamental
activities as cell division, gene expression, metabolic function, and cell
morphogenesis. Analysis of these processes may reveal otherwise elusive
components that are parts of plant and bacterial systems for signal
transduction, gene regulation, cell division, and cell wall formation.
1.1.2 Nodule formation and structure
Individual bacterial species and strains nodulate a particular set of host
plants (Table 1.1), and are characterized as having a host range that is either
broad (nodulating many different plants) or narrow (nodulating one or few
hosts). Nodules formed on different plants by different bacteria nonetheless
display striking developmental similarities (Fig. 1.1). Rhizobia attach to the
roots of their host and cause a characteristic curling of the host's root hairs (Yao
& Vincent, 1969; Dazzo & Gardiol, 1984). As this happens, cells in the root
cortex, under the epidermis, start to divide and form the nodule primordium
(Libbenga & Harkes, 1973; Newcomb, 1981). Bacteria trapped in a curled hair,
3
or between a hair and another cell, proliferate and begin to infect the outer plant
cells; as they do so the plant cells are stimulated to produce infection threads
(Callaham & Torrey, 1981). The infection threads ramify and penetrate
individual primordium cells. Bacteria released from infection threads into the
cytoplasm of target cells are enveloped in plant plasma membrane (Robertson et
al., 1978). The bacteria undergo limited DNA replication and division, then
cease both processes. Finally, the endosymbiotic forms of the bacteria, referred
to as bacteroids, begin to fix nitrogen by the action of the enzyme nitrogenase.
Table 1.1 Some Rhizobium-plant associations Bacterial Species Plant Hosts Rhizobium meliloti alfalfa (Medicago sativa) Rhizobium leguminosarum biovar viciae pea (Pisum sativum), vetch (Vicia sativa) biovar trifolii clovers (Trifolium species) biovar phaseoli Phaseolus bean Bradyrhizobium japonicum soybean (Glycine max) Rhizobium spp. NGR234 broad host range; genera including
Vigna, Macroptilium, Lablab, Glycine
Fig. 1.1 Initial stages in the rhizobia-legume symbiosis (Fisher & Long, 1992). Rhizobia attach to host root hairs (a) and cause root hair deformation (b), branching (c) and curling (d). Concomitant mitosis in the root cortex (e) culminates in the formation of the nodule primordium. Bacteria invade the plant cells through a novel structure termed the infection thread (f), a plant-derived tube which delivers the bacteria into individual cells within the nodule primordium. At this point the bacteria differentiate into bacteroids, which can fix atmospheric nitrogen to ammonia.
4
Nitrogenase is irreversibly inactivated by oxygen. However, the plant
derived "peribacteroid membrane" (PBM) contains leghaemoglobin.
Leghaemoglobin binds oxygen then releases it when the local concentration
drops below a certain level, thus providing a high flux for the bacteroid to use
in respiration, but in an environment with low free oxygen (Appleby, 1984).
The specialization of the PBM may also include specific transport or
permeability functions. Since all metabolite exchange between the host and the
bacteroid has to occur across this membrane, it may play an important role in
the regulation of nitrogen fixation.
5
1.2 BACTEROID METABOLISM
1.2.1 Carbon sources supplied to the bacteroid
The process of nitrogen fixation is energetically very costly with the
nitrogenase reaction utilizing as many as 16 molecules of ATP and reductant
equivalent to 8e- per molecule of N2 reduced.
Bacteroids within plant nodule cells are enclosed within the plant derived
PBM, and are dependent on the plant for provision of carbon/energy
substrates.
Nodule homogenates contain a number of carbohydrates and organic acids
in sufficient quantities to be considered as potential carbon substrates for
bacteroid respiration. To be considered seriously, these candidates must be
taken up and metabolized by isolated bacteroids at rates capable of supporting
nitrogenase, and they must be readily able to cross the PBM.
Although it has been shown that nitrogen fixation is fuelled by recently
synthesized sucrose translocated to the nodule (Reibach & Streeter, 1983;
Gordon et al., 1985; Kouchi & Nakaji, 1985), neither sucrose nor the hexoses
resulting from its hydrolysis are readily accumulated by isolated bacteroids.
This was first established for Rhizobium leguminosarum biovar viciae (Hudman &
Glenn, 1980; Glenn & Dilworth, 1981; de Vries et al., 1982) and was later
reported for other rhizobia (Reibach & Streeter, 1984; Saroso et al., 1984;
Salminen & Streeter, 1987b). In contrast to bacteroids, cultured R.
leguminosarum bv. viciae will take up sugars efficiently, indicating that
fundamental changes in carbon transport and metabolism occur in response to
the special environment in nodules (Hudman & Glenn, 1980; de Vries et al.,
1982; San-Francisco & Jacobson, 1986).
Movement of neutral sugars across the PBM of isolated soybean
symbiosomes is only by slow, passive diffusion with rates being inadequate to
support nitrogenase activity (Day & Udvardi, 1989; Udvardi et al., 1990).
6
The distribution of enzymes of carbohydrate metabolism in bacteroids is also
consistent with a minor role for sugars in bacteroid function. For example,
bacteroids appear to lack invertase, as first demonstrated by Robertson and
Taylor (1973), although they may have low sucrose synthase activity (Salminen
& Streeter, 1987b). Low levels of glycolytic enzymes in bacteroids relative to
host cytoplasm have also been demonstrated by numerous groups (Reibach &
Streeter, 1983; Salminen & Streeter, 1987b; Copeland et al., 1989b). Furthermore,
activity of the Entner-Douderoff pathway, a common mechanism for
conversion of hexose to carboxylic acids in Gram-negative bacteria, is very low
in bacteroids, although it is the main pathway of sugar metabolism in cultured
rhizobia (Salminen & Streeter, 1987b). Saroso et al. (1986) have shown that
snake bean bacteroids of cowpea Rhizobium NGR234 do not induce sugar
catabolic enzymes even though snake bean nodules contain significant
concentrations of these substrates and the enzymes systems are inducible in the
free-living form.
The relative unimportance of sugars in N2 fixation is also indicated by
reports that, with one exception (Duncan, 1981), Rhizobium mutants lacking the
capability to metabolize carbohydrates are still able to form effective symbioses
(Arias et al., 1979; Ronson & Primrose, 1979; Cervenansky & Arias, 1984; Glenn
et al., 1984; Arwas et al., 1985; Dilworth et al., 1986; El-Guezzar et al., 1988;
Lafontaine et al., 1989).
The fact that sugars are found in bacteroids is probably due to high
concentrations effecting passive uptake (Reibach & Streeter, 1984) coupled with
the low oxidation rate of these compounds. However, it should be noted that
glucose has been reported to stimulate nitrogenase activity in French bean
bacteroids (Trinchant et al., 1981) and some glucose uptake by French bean
symbiosomes has also been observed (Herrada et al., 1989). Thus French bean
may be an exception to the above generalization.
7
In contrast to neutral sugars, the importance of one or more of the C4-
dicarboxylate tricarboxylic acid (TCA) cycle intermediates, succinate, fumarate
and L-malate in symbiosis has been demonstrated. Numerous dicarboxylic acid
transport (Dct) mutants have been obtained (Table 1.2).
Most of the work has been done with the fast-growing species, but overall
results are remarkably consistent in showing that bacteroids incapable of
dicarboxylate uptake do not fix nitrogen. Furthermore, in contrast to the
sluggish metabolism of sugars, the metabolism of dicarboxylic acids in the
bacteroid is rapid, as illustrated by the relative rates of conversion of labelled
compounds to CO2 by purified bacteroids incubated under microaerobic
conditions (Salminen & Streeter, 1987a).
Table 1.2 Dicarboxylic acid transport mutants of rhizobia red, reduced uptake of dicarboxylic acids and reduced fixation activity; $, exact genetic lesion unknown. Mutants were generated with transposons apart from: #, spontaneous; *, produced with nitrosoguanidine. Rhizobium
species (biovar) Bacterial
phenotype Phenotype in nodule
Reference
leguminosarum
(viciae) Succinate uptake-# Nod+ Fix- Glenn and Brewin (1981)
Dct- Nod+ Fix- Arwas et al. (1985) Dct- Nod+ Fix- Finan et al. (1983) Dctred Nod+ Fixred Finan et al. (1983) meliloti Dct- Nod+ Fix- Engelke et al. (1987) Dctred Nod+ Fixred Engelke et al. (1987) Dct- Nod+ Fix- Bolton et al. (1986) Dct-* Nod+ Fix- Hornez et al. (1989) Dct- Nod+ Fix- Watson et al. (1988) trifolii Dct-* Nod+ Fix- Ronson et al. (1981) leguminosarum
(phaseoli) Dct- Nod+ Fix- LaFontaine et al. (1989)
japonicum Reduced succinate uptake$
Nod+ Fixred Humbeck and Werner (1989)
8
Since dct mutants nodulate and differentiate into bacteroids, it is clear that
they must be able to obtain sufficient carbon from the host plant to fuel such
processes. Despite attempts to determine the identity of the carbon sources
being used under such circumstances, they remain unknown. Double mutants
of R. leguminosarum lacking the Dct system and the ability to catabolize a range
of other carbon sources including C5-, C6-, and C12-sugars (Arwas et al., 1986)
still nodulate as well as the dct- parent strain.
Thus, while it is apparent that dicarboxylic acids are not the sole source of
reducing equivalents utilized for bacteroid formation, the fact that these TCA
cycle intermediates are required for nitrogen fixation implies an important role
for the TCA cycle in the bacteroid.
The observation that nodules induced by a malic enzyme mutant of R.
meliloti fail to reduce acetylene (Driscoll & Finan, 1993) is consistent with this
suggestion, since malic enzyme, which oxidatively decarboxylates malate to
pyruvate (Fig. 1.2), is required for metabolism of dicarboxylates via the TCA
cycle in Rhizobium (Dilworth et al., 1988). In addition, malic enzyme activity has
been demonstrated in R. leguminosarum (McKay et al., 1988) and B. japonicum
(Copeland et al., 1989a; Kimura & Tajima, 1989) bacteroids.
B. japonicum bacteroids have been found to oxidize 14C-labelled succinate,
pyruvate and acetate in a manner consistent with operation of the TCA cycle
(Stovall & Cole, 1978).
1.2.2 The TCA cycle in the bacteroid
The tricarboxylic acid cycle is the major energy-generating pathway in
aerobic heterotrophs, as well as an important source of intermediates for
cellular biosynthesis. It consists of a series of enzyme-catalyzed reactions which
brings about the total oxidation of acetyl units, derived as acetyl-CoA from
pyruvate and other metabolites (Fig. 1.2).
The demands imposed on cell metabolism vary according to the nutritional
environment. Thus in E. coli, which can grow under aerobic and anaerobic
9
conditions, deriving energy from a variety of respiratory and fermentative
processes, the TCA cycle is an inducible pathway which is fully induced only
under strictly aerobic conditions that make the greatest demands on its dual
anabolic and catabolic functions. Under anaerobic conditions all TCA cycle
enzymes in E. coli are repressed to some extent (Gray et al., 1966; Langley &
Guest, 1978; Buck et al., 1986), however the α-ketoglutarate dehydrogenase
complex is particularly affected (Amarasingham & Davis, 1965; Smith &
Neidhardt, 1983). As a result the TCA cycle becomes a branched non-cyclic
pathway (Miles & Guest, 1987; Guest, 1992; Guest & Russell, 1992), in which
carbon flows at a much reduced rate through two routes, an oxidative route
leading to α-ketoglutarate and a reductive route leading to succinate and
succinyl-CoA. These routes fulfil the biosynthetic functions of the TCA cycle.
10
Fig. 1.2 Some potential carbon metabolic pathways in Rhizobium. TCA cycle: CS, citrate synthase; ACN, aconitase; IDH, isocitrate dehydrogenase; KGDH, α-ketoglutarate dehydrogenase; SCS, succinyl-CoA synthetase; SDH, succinate dehydrogenase; FUM, fumerase; MDH, malate dehydrogenase. PHB biosynthesis: KT, β-ketothiolase; HBDH, acetoacetyl-CoA reductase; PHBP, poly-β-hydroxybutyrate synthase. Glutamate degradation: GOGAT, aspartate aminotransferase; GDH, glutamate dehydrogenase; GPT, glutamate-pyruvate aminotransferase; GDC, glutamate decarboxylase; AGT, γ-aminobutyric-glutamic transaminase; SSDH, succinate semialdehyde dehydrogenase. Other: ME, malic enzyme; PDH, pyruvate dehydrogenase.
11
In the nodule Rhizobium is maintained in an oxygen-limited environment.
Under oxygen-limitation, electron transport in the bacterial cell is saturated
with NAD(P)H, and the NAD(P)H:NAD(P) ratio is increased (Jackson &
Dawes, 1976). α-Ketoglutarate dehydrogenase has been shown to be inhibited
by NADH in the Gram-negative bacteria Acetobacter xylinum (Kornfeld et al.,
1977; DeKok et al., 1980) and Acinetobacter sp. (Weitzman, 1972; Hall &
Weitzman, 1977). Furthermore, when the non-symbiotic N2-fixing bacterium,
Azotobacter beijerinckii is subjected to oxygen limitation, α-ketoglutarate
dehydrogenase activity decreases by approximately 70% (Jackson & Dawes,
1976). Levels of NADH and NAD in anaerobically isolated B. japonicum
bacteroids are compatible with possible inhibition of α-ketoglutarate
dehydrogenase (Tajima & Kouzai, 1989; Salminen & Streeter, 1990), and this
enzyme may be a key point for regulation of TCA cycle function in the
bacteroid. Some potential consequences of the inhibition of α-ketoglutarate
dehydrogenase in the bacteroid are discussed in Section 1.2.5.
Activity of other TCA cycle enzymes such as citrate synthase (Kurz & LaRue,
1977), isocitrate dehydrogenase (Kurz & LaRue, 1977; Karr et al., 1984; Irigoyen
et al., 1990), fumarase (Karr et al., 1984) and malate dehydrogenase (DeVries et
al., 1980; Karr et al., 1984; Waters et al., 1985; Kouchi et al., 1988; Irigoyen et al.,
1990) has been detected in bacteroids of various species of Rhizobium and
Bradyrhizobium. The levels of malate dehydrogenase appear to be particularly
high, with activities in R. leguminosarum strain MNF300 bacteroids being 20-fold
greater than those in free-living cells (McKay et al., 1989).
The genes encoding all of the TCA cycle enzymes of E. coli have been cloned,
sequenced and located in the linkage map (Fig. 1.3). There is a major cluster at
16.3 min encoding citrate synthase, succinate dehydrogenase, the specific
components of the 2-oxoglutarate dehydrogenase complex, and succinyl-CoA
synthetase. There is also a smaller cluster at 2.8 min encoding the specific
components of the pyruvate dehydrogenase complex and lipoamide
12
dehydrogenase, the common component of the pyruvate dehydrogenase and 2-
oxoglutarate dehydrogenase complexes. The significance of this clustering is
unclear. The other genes are scattered about the linkage map.
Fig. 1.3 Linkage map of E. coli showing locations of TCA cycle and related genes (Guest & Russell, 1992). Arrows indicate experimentally identified mRNA transcripts (Miles & Guest, 1987). PDHC, pyruvate dehydrogenase complex; CS, citrate synthase; ACN, aconitase; ICDH, isocitrate dehydrogenase complex; ODHC, 2-oxoglutarate dehydrogenase complex; SCS, succinyl-CoA synthetase; SDH, succinate dehydrogenase; FUM, fumarase; MDH, malate dehydrogenase.
13
In the case of Rhizobium relatively few TCA cycle enzymes have been
characterized. The gene encoding isocitrate dehydrogenase in R. meliloti has
recently been cloned and mutated (McDermott & Kahn, 1992). The isocitrate
dehydrogenase mutants were found to form ineffective nodules. This is
compatible with reports that mutants of R. meliloti lacking α-ketoglutarate
dehydrogenase (Duncan & Fraenkel, 1979) or succinate dehydrogenase (Gardiol
et al., 1987) activity also form ineffective nodules. Interestingly, levels of malate
dehydrogenase and succinyl-CoA synthetase in the α-ketoglutarate
dehydrogenase mutant were found to be approximately 4-fold and 5-fold
higher, respectively than in the wild-type (Duncan & Fraenkel, 1979); while
malate dehydrogenase activity in a succinate dehydrogenase mutant isolated by
Finan et al. (1981) was elevated 5-fold.
Malate dehydrogenase from B. japonicum has been purified from bacteroids
and partially characterized (Waters et al., 1985; Emerich et al., 1988).
1.2.3 The role of poly-β-hydroxybutyrate biosynthesis
Bacteroids in some symbioses accumulate large quantities of poly-β-
hydroxybutyrate (PHB). In B. japonicum bacteroids PHB can account for up to
70% of cell dry mass (Wong & Evans, 1971; Bergersen & Turner, 1990). PHB is
derived from acetyl-CoA and its synthesis requires NAD(P)H (Fig. 1.2). Thus,
assuming the TCA cycle fuels the nitrogenase reaction, it appears that PHB
synthesis and nitrogenase compete for energy and reductants.
In Azotobacter oxygen-limitation brings about accumulation of large
quantities of PHB (Senior et al., 1972). Since very low oxygen tensions result in
an increase in the NAD(P)H:NAD(P) ratio (Jackson & Dawes, 1976), and
NADPH and/or NADH inhibit isocitrate dehydrogenase and citrate synthase
(Senior et al., 1972), it has been proposed that in Azotobacter, the formation of
PHB serves a regulatory role (Senior, 1971; Senior et al., 1972). It is suggested
that under oxygen-limitation, accumulated NAD(P)H is channelled into the
14
synthesis of PHB, thereby reducing the inhibition of citrate synthase and
isocitrate dehydrogenase.
By analogy, it has been proposed that PHB biosynthesis in B. japonicum
bacteroids serves a similar purpose (McDermott et al., 1989). Certainly the
results of measurements of pyridine nucleotide redox state in bacteroids
incubated under conditions mimicking the nodule environment are suggestive.
The NADP pool was found to be even more reduced than the NAD pool
(Tajima et al., 1988; Tajima & Kouzai, 1989). Synthesis of PHB may, in part, be
responsible for the lower NADH/NAD ratio since in B. japonicum acetoacetyl-
CoA reductase, the second enzyme in the biosynthetic pathway from acetyl-
CoA to PHB (Fig. 1.2), has a 20-fold lower Km for NADH than NADPH
(Emerich, 1985).
1.2.4 The malate-aspartate shuttle
Various amino acids have been considered as possible carbon sources for
bacteroids. Kahn et al. (1985) focused on glutamate because it is often needed to
induce nitrogen fixation by ex planta cultures of Rhizobium and because
glutamate catabolism can be accompanied by ammonia excretion (O'Gara &
Shanmugam, 1976; Tubb, 1976). To explain why bacteroids continue to fix
nitrogen whilst showing no other signs of nitrogen limitation, (glutamine
synthetase and glutamate synthase (GOGAT) levels are very low in bacteroids,
and the high-affinity ammonia uptake system that is induced by nitrogen
limitation in free-living R. leguminosarum (O'Hara et al., 1985) in not switched
on), Kahn et al. have proposed the model outlined in Fig. 1.4.
In this scheme glutamate (or another nitrogen-containing compound) is fed
to the bacteroid where it is catabolized to yield ammonia or an amino acid,
energy and carbon. The waste products are returned to the plant with no loss
of fixed nitrogen and the plant uses the nitrogen to regenerate glutamate and
complete the cycle. This cycle benefits only the bacteroid since all of the
nitrogen is being recycled. However, if the plant removes some of the ammonia
15
in order to support its own growth, an "appropriate" bacterial response is to
produce more ammonia in order to replenish the fixed nitrogen that serves as a
carrier of carbon and energy. This model elegantly accounts for the repression
of nitrogen assimilatory enzymes in the bacteroid (nitrogen containing
compounds are available to it) and also explains why the bacteroid exports
fixed nitrogen.
Fig 1.4 Nutrient flow between plant and bacteroid, as proposed by Kahn et al. (1985). PBM, peribacteroid membrane; BM, bacteroid membrane.
16
While glutamate can be catabolized in a variety of ways (Fig. 1.2), Kahn et al.
have suggested the operation of a malate-aspartate shuttle between the plant
cytoplasm and the bacteroid (as an extension of the model in Fig. 1.4). In this
pathway malate and glutamate are imported into the bacteroid in exchange for
α-ketoglutarate and aspartate, respectively. Malate is oxidized within the
bacteroid to give oxaloacetate which is then transaminated by glutamate to give
aspartate. α-Ketoglutarate is transaminated in the plant by aspartate to yield
glutamate. The net result is the transfer of NADH into the bacteroid without
the transfer of carbon (Fig. 1.5).
Fig. 1.5 The malate-aspartate shuttle. PBM, peribacteroid membrane; BM, bacteroid membrane.
17
The malate-aspartate shuttle is used to transfer reducing equivalents into
mitochondria (LaNoue & Tischler, 1974; Meijer & Van Dam, 1974; Indiveri et al.,
1987; Dierks & Krämer, 1988) and therefore argue Kahn et al., should not
require novel behaviour from the plant. The needed transport proteins would
be made by the plant but would be located in the PBM instead of the inner
membrane of the mitochondrion.
There has been much debate over the malate-aspartate shuttle as a possible
alternative or supplementary mechanism for the transfer of reducing
equivalents to bacteroids.
Significant levels of malate dehydrogenase (DeVries et al., 1980; Karr et al.,
1984; Waters et al., 1985; Kouchi et al., 1988; McKay et al., 1989; Irigoyen et al.,
1990; Appels & Haaker, 1991) and aspartate aminotransferase (Ryan et al., 1972;
Ryan & Fottrell, 1974; Reynolds et al., 1981; Kouchi et al., 1988; Appels &
Haaker, 1991) have been found in both cytosol and bacteroids of legume
nodules. Malate dehydrogenase is required for TCA cycle activity and cytosolic
aminotransferase might be required for aspartate and asparagine synthesis, so
the presence of these enzymes does not necessarily indicate the operation of a
malate-aspartate shuttle. However, it is difficult to explain high
aminotransferase activity in bacteroids on the basis of aspartate and asparagine
synthesis.
When nitrogenase activity in cultured Bradyrhizobium japonicum was induced
by lowering O2 concentration, etc., aspartate aminotransferase activity nearly
doubled relative to cells in which nitrogenase was repressed (Werner & Stripf,
1978). Furthermore, a mutant of Rhizobium meliloti lacking aspartate
aminotransferase forms nodules which are Fix- (Rastogi & Watson, 1991).
Appels and Haaker (1991) have recently proposed that the main function of a
nodule stimulated cytoplasmic form of aspartate aminotransferase in Rhizobium
leguminosarum strain PRE is participation in a malate-aspartate shuttle. This
suggestion was based on changes in the concentrations of amino acids and
18
organic acids in the incubation medium of pea bacteroids. However, the
findings of Rosendahl et al. (1992) do not support this hypothesis:
Symbiosomes of R. leguminosarum bv. viciae MNF300 incubated with
[14C]malate did excrete [14C]aspartate, and this excretion was increased 3-fold
when unlabelled glutamate was also added as an "exchange substrate" for
aspartate. However, when symbiosomes were incubated with [14C]glutamate
no radioactive α-ketoglutarate could be detected in the incubation medium,
with or without the addition of unlabelled malate.
This disparity in results is apparently not due to impermeability of the
peribacteroid membrane to α-ketoglutarate, since the bacteroid preparations
used in the earlier study were prepared at a high osmotic value and were
reported to contain a majority of intact symbiosomes. However, it is
conceivable that metabolism of carbon storage materials in the bacteroid
affected the carbon pools recorded by Appels and Haaker.
Streeter and Salminen (1990) concluded that an active malate-aspartate
shuttle is not present in B. japonicum bacteroids. Bacteroids supplied with
[14C]malate released only small amounts of [14C]aspartate, and only when
unlabelled glutamate was also provided. Although α-[14C]ketoglutarate was
released by bacteroids when malate, succinate or α-ketoglutarate was supplied
in addition to [14C]glutamate, the formation of labelled α-ketoglutarate was
attributed to a rapid exchange of label via aspartate aminotransferase in the
periplasm.
While glutamate is readily accumulated by isolated Bradyrhizobium japonicum
bacteroids (Salminen & Streeter, 1987a; Udvardi et al., 1988) and will support
their nitrogenase activity (Bergersen & Turner, 1988; Kouchi & Fukai, 1988;
Appels & Haaker, 1991; Kouchi et al., 1991) the observation that the
peribacteroid membrane in soybean and siratro nodules is virtually
impermeable to glutamate (Udvardi et al., 1988; Udvardi & Day, 1988; Ouyang
& Day, 1992) is not compatible with the operation of a malate-aspartate shuttle.
19
However, comparison of respiratory evolution of 14CO2 from [14C]glutamate by
intact symbiosomes with that from naked bacteroids, indicates that glutamate
uptake at higher concentrations may be significant (Kouchi et al., 1991):
Evolution of 14CO2 from [14C]glutamate by B. japonicum symbiosomes was less
than 15% of that by naked bacteroids at a glutamate concentration of 0.1mM,
but about 45% of that at 2mM. On the other hand, rates of respiratory
utilization of [14C]malate by intact symbiosomes were about 60% of those by
naked bacteroids at both concentrations of 0.1 and 2mM. In soybean nodules
estimates of the in vivo concentration of glutamate in the plant cell cytosol range
from 1 to 10mM (Kouchi & Yoneyama, 1986; Streeter, 1987; Bergersen & Turner,
1988). Therefore, the peribacteroid membrane may not work as a potential
barrier to the supply of glutamate to the bacteroids from plant cytosol within
the range of concentrations expected in vivo in the cytosol in nodules.
Glutamate is neither actively accumulated by, nor does it support
nitrogenase activity of, isolated R. meliloti bacteroids (McRae et al., 1989a;
McRae et al., 1989b), although its respiration following passive diffusion into R.
meliloti bacteroids has been reported (Ta et al., 1988).
1.2.5 Other roles of amino acids in bacteroid metabolism
If the operation of an orthodox malate-aspartate shuttle in many symbioses
now seems unlikely, a variety of evidence suggests that glutamate may still
play a central role in bacteroid metabolism.
B. japonicum bacteroids from soybean nodules contain a large pool of
glutamate (Kouchi & Yoneyama, 1986; Streeter, 1987). This fact is consistent
with several studies which indicate that a significant proportion of
malate/succinate supplied to bacteroids is converted to glutamate: Salminen
and Streeter (1987a) provided anaerobically isolated B. japonicum bacteroids
with 14C labelled succinate or malate under microaerobic conditions. After 1
hour 62% of unrespired succinate (32% of the label taken up) and 75% of
unrespired malate (21% of the label taken up) were recovered as glutamate.
20
Significantly, when bacteroids were supplied with [14C]glutamate 97% of
unrespired substrate (46% of the label taken up) was found as glutamate; i.e.
essentially the only fate of glutamate was conversion to CO2. (This has also
been found with R. leguminosarum symbiosomes (Rosendahl et al., 1992)). In
earlier work with B. japonicum bacteroids under low O2 concentrations (Tajima
et al., 1986) [2,3-14C]succinate was found to be incorporated mainly into TCA
cycle acids, with little or no 14C in amino acids. This discrepancy may be due to
the fact that Tajima et al. incubated bacteroids under an Ar-02 mixture, whereas
the later experiments employed N2-O2, so that the bacteroids were actively
fixing N2 during incubation. Indeed glutamate is the most highly labelled
compound in bacteroids following incubation of intact soybean nodules with 15N2 (Ohyama & Kumazawa, 1980).
Kouchi et al. (1991) have also reported high levels of [14C]glutamate in B.
japonicum bacteroids following 20 minute incubations with either [14C]malate or
[14C]glutamate.
A 3-fold increase in glutamate concentration was observed in anaerobically
isolated R. meliloti bacteroids incubated with succinate or malate for 30 minutes
under 4% oxygen plus either argon or nitrogen (Miller et al., 1991). In this case
the bacteroid pool size of free ammonia was found to be sufficient to account
for the glutamate synthesized under argon.
Experiments with intact nodules, in which [14C]malate was generated by the
action of phosphoenolpyruvate carboxylase on 14CO2, appear to confirm the
results obtained with isolated bacteroids (Salminen & Streeter, 1992): Intact
nodules of soybean inoculated with B. japonicum and pea inoculated with R.
leguminosarum were detached and immediately fed 14CO2 for up to 6 minutes.
Bacteroids rapidly purified from these nodules contained most label in malate.
However, the rate of glutamate labelling was 67% of the rate of malate labelling
in the case of B. japonicum bacteroids, while in R. leguminosarum bacteroids
isolated after 6 minutes incubation, glutamate contained 78% of the amount of
21
label found in malate. Since the radioactivity in glutamate in the cytosolic
fraction of the nodules was found to be low, the labelling of glutamate in the
bacteroids would seem indicative of malate metabolism rather than uptake of
labelled glutamate, especially since the uptake of malate by isolated B.
japonicum bacteroids is 3.4 times faster than that of glutamate (Salminen &
Streeter, 1991).
It has been suggested (McDermott et al., 1989; Salminen & Streeter, 1990) that
glutamate synthesis from malate under the oxygen-limiting conditions of the
nodule is the consequence of diversion of α-ketoglutarate away from the TCA
cycle due to inhibition of α-ketoglutarate dehydrogenase by NADH (Section
1.2.2). The fact that no accumulation of radioactivity in α-ketoglutarate is seen
in bacteroids supplied with [14C]malate (Salminen & Streeter, 1990), probably
reflects the equilibrium constant (1014) of the glutamate dehydrogenase
catalysed reaction for the formation of glutamate, and the high ammonia
concentrations in N2-fixing bacteroids (Klucas, 1974; Streeter, 1989). Indeed
ammonia and α-ketoglutarate are extremely efficient inducers of glutamate
dehydrogenase in B. japonicum (Fottrell & Mooney, 1969).
Glutamate may also be formed from α-ketoglutarate via glutamine
synthetase and glutamate synthase, both of which have been detected in R.
leguminosarum and B. japonicum bacteroids (Brown & Dilworth, 1975). In fact,
glutamate synthase activity in B. japonicum bacteroids increases as the symbiosis
progresses, whereas that of glutamate dehydrogenase declines (Stripf &
Werner, 1978); and glutamate synthase activity doubles when B. japonicum is
shifted from nitrogenase repressed to nitrogenase derepressed conditions
(Werner & Stripf, 1978). Furthermore, a NADP-dependent glutamate synthase
deficient mutant of B. japonicum forms ineffective nodules (O'Gara et al., 1984),
while a B. japonicum mutant lacking glutamine synthetase activity fails to
nodulate soybeans (Carlson et al., 1987).
22
However, glutamate synthase does not appear to be significant in R. meliloti
bacteroids, as mutants lacking this enzyme are Fix+ (Kondorosi et al., 1977;
Osburne & Signer, 1980; Lewis et al., 1990). In addition, none of the three forms
of glutamine synthetase, encoded by the genes glnA, glnII and glnT (Section 1.6),
is essential for symbiotic nitrogen fixation in R. meliloti, since strains carrying a
single mutation in any one of these genes are Fix+ (de Bruijn et al., 1989;
Somerville et al., 1989). Furthermore, assuming that glnT is not expressed in
bacteroids, as appears to be the case for R. leguminosarum (Espin et al., 1990), the
Fix+ phenotype of a glnA glnII double mutant (de Bruijn et al., 1989), suggests
that no glutamine synthetase activity is required for nitrogen fixation by R.
meliloti bacteroids.
A possible fate of glutamate derived from α-ketoglutarate is decarboxylation
to form γ-aminobutyrate (GABA), then deamination and oxidation to succinate
(Fig. 1.2). This "GABA-shunt" would provide a detour around α-ketoglutarate
dehydrogenase, removing an equivalent amount of CO2 and yielding the same
end product, succinate, which can re-enter the TCA cycle. While this pathway
appears to operate in R. meliloti bacteroids (Fitzmaurice & O'Gara, 1991; Miller
et al., 1991), the results of tracer experiments and the detection of only extremely
low levels of glutamate decarboxylase activity, suggest that the GABA-shunt is
essentially absent from B. japonicum bacteroids (Salminen & Streeter, 1990;
Kouchi et al., 1991).
There is evidence to suggest that the amino acids aspartate and alanine may
also play a significant part in bacteroid metabolism.
Like glutamate, aspartate is actively accumulated by B. japonicum bacteroids
(Reibach & Streeter, 1984), and at controlled substrate and oxygen
concentrations, stimulates respiration (Salminen & Streeter, 1987a), and
supports nitrogenase activity (Salminen & Streeter, 1987a; Bergersen & Turner,
1988; Kouchi & Fukai, 1988; McRae et al., 1989b; Appels & Haaker, 1991; Kouchi
et al., 1991). B. japonicum bacteroids contain relatively high concentrations of
23
aspartate (Streeter, 1987) but exogenously supplied aspartate appears to be
catabolized without being incorporated into the endogenous aspartate pool
(Salminen & Streeter, 1987a; Tajima & Kouchi, 1990; Kouchi et al., 1991). On
incubation of anaerobically isolated B. japonicum bacteroids with [14C]aspartate
the largest amount of unrespired label is found as glutamate (Salminen &
Streeter, 1987a; Tajima & Kouchi, 1990; Kouchi et al., 1991), and the failure of
amino-oxyacetate, a specific inhibitor of transamination, to affect the utilization
of aspartate (Tajima & Kouchi, 1990; Kouchi et al., 1991), indicates that aspartate
utilization occurs via the TCA cycle after direct deamination by aspartase. In
contrast, glutamate utilization is strongly inhibited by the presence of amino-
oxyacetate, and it is suggested (Tajima & Kouchi, 1990; Kouchi et al., 1991) that
the major fate of exogenously supplied glutamate in B. japonicum bacteroids is
transamination to form aspartate.
While actively accumulated by a low-affinity transport system, aspartate
does not support nitrogenase activity in R. meliloti bacteroids (Miller et al., 1988;
McRae et al., 1989b). However, the observation that a Tn5 induced mutant of R.
meliloti lacking aspartate aminotransferase does not grow on aspartate as a
carbon source and forms Fix- nodules has prompted Rastogi and Watson (1991)
to suggest that aspartate provided by the plant to the bacteria in the nodule is
essential for an effective symbiosis.
Recently isolated isocitrate dehydrogenase mutants of R. meliloti (McDermott
& Kahn, 1992) are found to be strict glutamate auxotrophs. Since these mutants
are Nod+, the host must be directly or indirectly providing enough glutamate
for cell growth and maintenance. However, their Fix- phenotype suggests that
either α-ketoglutarate or glutamate is not provided in large quantities or that
glutamate catabolism is of relatively little importance. McDermott and Kahn
have suggested that glutamate obtained via aspartate aminotransferase may be
significant in R. meliloti (α-ketoglutarate is required for production of glutamate
in this way, and isocitrate dehydrogenase mutants cannot synthesize α-
24
ketoglutarate). This explanation could account for both the Fix- phenotype of R.
meliloti GABA-shunt mutants (Fitzmaurice & O'Gara, 1988) and the Fix+
phenotype of R. meliloti glutamate synthase mutants (Kondorosi et al., 1977;
Osburne & Signer, 1980; Lewis et al., 1990), and is also consistent with aspartate
having an important role in R. meliloti bacteroid metabolism.
Alanine is neither actively accumulated by, nor does it support nitrogenase
activity of, isolated R. meliloti bacteroids (McRae et al., 1989a; McRae et al.,
1989b). However, in anaerobically isolated B. japonicum bacteroids incubated
with [14C]succinate under microaerobic conditions for 1 hour, 11% of the label
taken up was recovered as alanine (Salminen & Streeter, 1987a). Curiously, the
same figure after incubation with [14C]malate was only 3%. In their
experiments involving incubation of pea and soybean nodules with 14CO2, in
which malate was the major form of labelled carbon supplied to the bacteroids
(see above), Salminen and Streeter (1992) found that R. leguminosarum
bacteroids accumulated as much label in alanine as in glutamate (74% of the
amount of label found in malate), while in B. japonicum bacteroids the rates of
labelling of alanine and aspartate were 18 and 24% that of malate, respectively.
Several groups have reported excretion of alanine and/or aspartate by
nitrogen fixing bacteroids. Kretovich et al. (1986) incubated bacteroids of
Rhizobium lupini 359a with succinate, malate or fumarate under nitrogen and 6%
oxygen for up to 30 minutes. Aspartate, and smaller amounts of alanine were
found to be excreted into the incubation medium in each case, with the greatest
excretion (37 nmole of aspartate per gram of bacteroids per minute) being
observed when malate was the substrate. Appels and Haaker (1991) noted the
presence of alanine in the medium after incubation of R. leguminosarum strain
PRE bacteroids with malate plus glutamate or aspartate plus α-ketoglutarate
under nitrogen fixing conditions. They also reported glutamate-pyruvate
aminotransferase activity in the bacteroids and proposed the formation of
alanine through transamination of pyruvate by glutamate. (Pyruvate is formed
25
from malate by the action of malic enzyme, and glutamate and malate would be
formed from aspartate and α-ketoglutarate by a reverse malate-aspartate
shuttle).
Kouchi et al. (1991) found marked accumulation of alanine, but not aspartate,
in the medium when anaerobically isolated bacteroids of B. japonicum A1017
were incubated with malate for 20 minutes under N2 and 1.5% O2. When
bacteroids were incubated with glutamate, aspartate, but not alanine,
accumulated in the medium. However, when bacteroids were incubated with
malate plus glutamate the accumulation in the medium of both aspartate and
alanine, particularly aspartate, increased greatly.
Similar results have been obtained with anaerobically isolated R.
leguminosarum symbiosomes from pea nodules (Rosendahl et al., 1992):
Incubation of symbiosomes with 0.5mM [14C]malate under 4% O2 resulted in
14% of the label recovered after 30 minutes being detected in alanine; 7% inside
the symbiosomes and 7% in the surrounding medium. Aspartate plus
glutamate (indistinguishable by TLC) accounted for 7% of the label: 2%
intracellular, 5% extracellular. When symbiosomes were incubated with 0.5mM
[14C]malate plus 0.5mM unlabelled glutamate under the same conditions the
results were as follows: 9% of the label recovered as alanine inside
symbiosomes; 25% as alanine in the incubation medium; 3% as
aspartate/glutamate inside symbiosomes; 17% as aspartate/glutamate in the
incubation medium.
Rosendahl et al. suggest the following explanation of these results: Labelled
malate is transported across the PBM and bacteroid membrane into the
bacteroids where it is converted to oxaloacetate (malate dehydogenase) and
pyruvate (malic enzyme). Transamination from amino donors such as
glutamate then produces radioactive aspartate and alanine. When exogenous
glutamate is added, the release of alanine and aspartate is accelerated because
glutamate taken into bacteroids is functioning as an additional amino donor in
26
transamination, and/or the rates of amino acid transport across the bacteroid
and/or peribacteroid membrane are increased. The latter suggestion is
prompted by reports of common amino acid transporters capable of exchanging
intra- and extracellular pools of amino acids in R. leguminosarum (Poole et al.,
1985) and cowpea Rhizobium (Glenn et al., 1991).
It has been argued (McDermott et al., 1989) that under the oxygen-limited
conditions of the bacteroid where citrate synthase, isocitrate dehydrogenase
and α-ketoglutarate dehydrogenase may be repressed relative to malate
dehydrogenase (Senior, 1971; Jackson & Dawes, 1976; Karr et al., 1984;
Suganuma & Yamamoto, 1987; McKay et al., 1989), oxaloacetate derived from
actively accumulated malate (or succinate) would build up in the absence of
outlets other than the TCA cycle. Since oxaloacetate competitively inhibits
succinate dehydrogenase, removal or utilization of excess oxaloacetate may be
essential in maintaining the TCA cycle. Excretion of aspartate (derived from
oxaloacetate via aspartate aminotransferase), and/or excretion of alanine
(derived from malate via malic enzyme and glutamate pyruvate transaminase)
by the bacteroid could provide a means of preventing accumulation of
oxaloacetate. If this is the case, amino acid transport across the
bacteroid/peribacteriod membrane, the likely rate determining step in the
oxaloacetate depletion process, could regulate bacteroid metabolism. This idea
is not without precedent since the malate-aspartate shuttle in mitochondria is
regulated by an amino acid exchange transporter (Murphy et al., 1979).
27
1.3 AMINO ACID TRANSPORT IN BACTERIA
The prokaryotic amino acid uptake systems reported to date can be divided
into two categories: (i) periplasmic binding protein-independent, secondary
solute transport systems in which the free energy for accumulation of the amino
acid(s) is supplied by an electrochemical gradient (ii) periplasmic binding
protein-dependent transport systems which utilize ATP hydrolysis to energize
translocation.
Amino acid uptake systems in category (i) are usually symporters, mediating
the coupled movement of two (or more) solutes in the same direction.
Hydropathy analyses and other topographic studies on transport proteins of
this type, point, in most cases, to a secondary structure that includes twelve
hydrophobic domains in α-helical configuration, traversing the membrane in a
zig-zag fashion (Poolman & Konings, 1993). Examples of this type of
transporter are the proton/glutamate symporter (GltP) of Bacillus subtilis
(Tolner et al., 1995), the sodium/proline symporter (PutP) of Salmonella
typhimurium (Miller & Maloy, 1990; Poolman & Konings, 1993), and the
sodium/proton/glutamate symporter (GltT) of Bacillus stearothermophilus and
Bacillus caldotenax (Tolner et al., 1992a). Such transporters are not of direct
relevance to the research reported in this thesis and are further discussed only
briefly in Section 1.4.6. Transporters in category (ii) are described in Section 1.4.
Bacterial amino acid transport systems are usually highly specific for a single
amino acid, or group of related amino acids (Halpern, 1974; Landick et al., 1985;
Antonucci & Oxender, 1986). However, general amino acid permeases, which
transport all amino acids, have been found in the eukaryotic microorganisms
Neurospora and Saccharomyces (Pall, 1969; Rao et al., 1975; DeBusk & DeBusk,
1980; Ogilvie-Villa et al., 1981; Grenson et al., 1970; Grenson & Hon, 1972; Eddy,
1982; Jauniaux & Grenson, 1990).
Transport of one amino acid may be mediated by several different permeases
with overlapping specificities. Thus in E. coli three L-glutamate transport
28
systems have been identified: (i) a binding protein-dependent glutamate-
aspartate system (Schellenberg & Furlong, 1977); (ii) a proton/glutamate-
aspartate system (Wallace et al., 1990; Tolner et al., 1992b); (iii) a
sodium/glutamate system (Deguchi et al., 1989; Deguchi et al., 1990; Kalman et
al., 1991). Some potential reasons for such multiplicity of transport systems are
discussed in Section 1.4.6.
29
1.4 ABC TRANSPORTERS
Prokaryotic periplasmic binding protein-dependent permeases are members
of the ABC superfamily of membrane transporters. Members of this
superfamily have extensive sequence and structural similarity, and the
designation ABC transporters recognizes a highly conserved ATP-binding
cassette, a particularly characteristic feature of family members that is
indicative of the use of ATP hydrolysis in the energization of substrate
translocation. The name traffic ATPases has also been suggested for these
transporters (Mimura et al., 1990).
ABC transporters have been identified in both prokaryotes and eukaryotes.
Among the eukaryotic systems are the medically important cystic fibrosis
transmembrane regulator (CFTR) and multidrug resistance (MDR) protein.
Each ABC transporter is relatively specific for a given substrate. However,
different transporters handle a wide variety of substrates including amino
acids, sugars, inorganic ions, polysaccharides and peptides (Ames, 1986;
Furlong, 1987; Higgins et al., 1990). Some ABC transporters are uptake systems,
while others export substrate from the cell: none has yet been identified that
can pump in both directions (Higgins, 1992).
1.4.1 Overall structure of ABC transporters
ABC transporters require the function of multiple polypeptide/protein
domains, organized in a characteristic fashion (Fig. 1.6). The typical transporter
consists of four membrane-associated domains. Two of these domains are
highly hydrophobic and each usually consists of six membrane-spanning
segments. These domains form the pathway through which substrate crosses
the membrane, and are believed to play a significant part in determining the
substrate specificity of the transporter. The other two domains are peripherally
located at the cytoplasmic face of the membrane, bind ATP and couple ATP
hydrolysis to the transport process.
30
Fig 1.6 Structural organization of a typical ABC transporter (Higgins, 1992). The location of the short sequence motif conserved between the transmembrane domains of many of these transporters is indicated by *.
The individual domains of an ABC transporter are frequently expressed as
separate polypeptides, particularly in prokaryotic species. The oligopeptide
transporter of S. typhimurium (Hiles et al., 1987) is an example of this
arrangement (Fig. 1.7A). However, there are examples in which two or more
domains are fused into larger, multifunctional polypeptides, as in the ribose
transporter (Bell et al., 1986) of E. coli (Fig. 1.7B). Many eukaryotic ABC
transporters, such as the MDR protein and the CFTR gene product, have all four
domains fused into a single protein (Fig. 1.7C).
31
Fig. 1.7 Examples of differing domain organization in ABC transporters (Higgins, 1992).
32
In addition to the four core domains, all bacterial ABC transporters that
mediate solute uptake require a substrate-binding protein located in the
periplasm (Figs. 1.7A, B, D). These periplasmic components are essential for the
function of the transporter with which they are associated, although they are
not integral to the process of transmembrane solute translocation itself. By
contrast, there is no evidence that eukaryotic ABC transporters utilize a soluble
receptor (a periplasmic binding protein equivalent) to transfer substrate to the
membrane-bound components of the transporter.
No ABC transporter has yet been shown to function with fewer than the four
core domains, and in the absence of evidence to the contrary, it is assumed that
the four core domains form the basic unit required to mediate solute
translocation. It is generally assumed that a functional transport complex
consists of one of each of the four core domains rather than a larger oligomeric
assembly. However, at present, this is purely an assumption. A few ABC
transporters appear to lack a full complement of domains. For example, the
operon encoding the histidine transporter of S. typhimurium includes only a
single gene, hisP, encoding an ATP-binding component. However,
coimmunoprecipitation of HisQMP has recently shown that this transport
complex contains HisP in a 2:1 ratio with the other domains (Kerppola et al.,
1991), suggesting that HisP functions as a homodimer (Fig. 1.7D). Similar
results have also been obtained for the maltose system of E. coli (Davidson &
Nikaido, 1991). The core components of minimalist transporters such as the
glutamine uptake system in E. coli, which, in addition to a binding protein gene,
glnH, has only one hydrophobic membrane component gene, glnP, and one
ATP-binding component gene, glnQ, (Nohno et al., 1986), probably function as
homodimers, although this has yet to be tested experimentally.
33
1.4.2 The transmembrane domains
The two transmembrane domains of ABC transporters are highly
hydrophobic. Each is predicted, from its sequence, to consist of multiple α-
helical segments that could span the membrane. The majority of transporters
are predicted to have six membrane-spanning segments per domain (a total of
twelve per transporter), with the N- and C-termini on the cytoplasmic face of
the membrane and three extracellular and two intracellular loops (Fig. 1.6). The
available experimental data are consistent with these predictions. Thus the six
predicted transmembrane segments of each domain of the oligopeptide
permease of S. typhimurium have been identified experimentally, using both
biochemical techniques and β-lactamase fusion analysis of OppB and OppC
(Pearce et al., 1992).
A few ABC transporters apparently do not conform to the two-times-six
transmembrane helix paradigm. For example, TnphoA fusion analysis of MalF
from the maltose transporter of E. coli (Froshauer et al., 1988), suggests a total of
eight membrane-spanning regions for this protein, in agreement with
computer-assisted predictions. Alignment with equivalent components of other
transporters (Overduin et al., 1988) indicates that MalF consists of the six
standard transmembrane segments, but has an N-terminal extension with two
additional transmembrane segments (Fig. 1.8). Furthermore, these two
transmembrane segments can be deleted without loss of MalF function
(Ehrmann et al., 1990). Another potential exception is the histidine transporter
of S. typhimurium, which has two hydrophobic components, HisQ and HisM,
each predicted to have five, rather than six, transmembrane segments (Higgins
et al., 1982b; Ames, 1985). These predictions have recently been confirmed
experimentally through TnphoA fusion analysis of HisQ and HisM, and studies
involving use of proteolytic enzymes and antibodies with oriented membrane
vesicles (Kerppola et al., 1991; Kerppola & Ames, 1992). Alignment with other
transporters (Fig. 1.8) indicates that the usual N-terminal transmembrane
segment of each domain may be absent, which places the N-termini on the
34
exterior of the cell. This orientation may indicate that ten (two-times-five)
transmembrane segments provide the minimal unit required to form the
translocation pathway itself; the additional N-terminal transmembrane
segment(s) of most transporters may simply facilitate correct folding, packing,
and orientation within the membrane.
Fig. 1.8 Alignment of OppC, HisQ and MalF, showing membrane-spanning segments as black boxes (Higgins, 1992). The majority of integral membrane components of ABC transporters are exemplified by OppC, which has six transmembrane segments. HisQ (and HisM) appears to lack transmembrane segment 1, while MalF has two additional N-terminal transmembrane segments. The conserved motif in these proteins (indicated by *) is located between transmembrane segments 4 and 5.
Comparison of the amino acid sequences of the hydrophobic components of
one transporter with those of another reveals little or no similarity. This implies
that the structural constraints required for the function of the hydrophobic
domains can be satisfied by a variety of amino acid combinations. The only
significant sequence conservation between the transmembrane domains of
several different ABC transporters is a short motif identified on many bacterial
transporters (Ames, 1985; Dassa & Hofnung, 1985; Kerppola & Ames, 1992;
Saurin et al., 1994), appropriately positioned on a cytoplasmic loop (Figs. 1.6
and 1.8) to interact with the ATP-binding domains (Pearce et al., 1992). Whether
or not it serves this function has yet to be established.
35
The two integral membrane domains of an individual ABC transporter are
generally more closely related to each other than they are to the equivalent
domains of other transporters in the superfamily. Thus the HisQ and HisM
components of the S. typhimurium histidine transporter are closely related to
each other (Higgins et al., 1982b), as are the OppB and OppC components of the
oligopeptide transporter (Hiles et al., 1987). This similarity implies that the two
domains function symmetrically as a pseudodimer. This would be consistent
with the view that the single hydrophobic domain found in some ABC
transporters, may function as a homodimer.
While, for bacterial uptake systems, the periplasmic binding proteins are
important in determining substrate specificity (Section 1.4.4), it is clear that the
transmembrane components also play a role. Binding protein-independent
mutants still exhibit substrate selectivity (Treptow & Shuman, 1985; Petronilli &
Ames, 1991), and mutations that alter the specificity of a transporter invariably
alter the transmembrane domains. Thus, mutations that alter the selectivity of
the histidine transporter of S. typhimurium from L-histidine to L-histidinol delete
four amino acids from a membrane-spanning segment of the transmembrane
domain HisM (Payne et al., 1985). Similarly, mutations that allow the maltose
transporter of E. coli to transport the analogue p-nitrophenyl-α-maltoside (not
normally a substrate) also alter the transmembrane domains (Reyes et al., 1986).
1.4.3 The ATP-binding domains
The ATP-binding domains of ABC transporters are their most characteristic
feature. Each domain is about 200 amino acids long and the domains from
different transporters share considerable sequence identity. The conserved
sequences include two short motifs, the Walker motifs, associated with many
nucleotide-binding proteins (Walker et al., 1982; Higgins et al., 1985; Higgins et
al., 1986).
The ATP-binding domains are highly hydrophilic, include no potential
membrane-spanning segments, and would not normally be expected to span
36
the membrane. Nevertheless, these components appear to be tightly associated
with the cytoplasmic membrane, although the nature of this association is not
clear. In some cases there is evidence to suggest peripheral attachment to the
cytoplasmic face of the membrane. Thus the MalK component of the maltose
transporter of E. coli is found to be cytoplasmic in the absence of the integral
membrane components (Shuman & Silhavy, 1981), while OppF, the ATP-
binding component of the S. typhimurium oligopeptide transporter, is less
accessible to proteolysis from the exterior of the cell than from the cytoplasm
(Gallagher et al., 1989).
Such evidence contrasts with what has been found for HisP from S.
typhimurium, which behaves neither like a typical integral nor a peripheral
membrane protein when it is in the presence of HisQ and HisM (Kerppola et al.,
1991). Interestingly, HisP prefers to be associated with the membrane even in
the absence of HisQ and HisM, but in such a case it assumes an improper
association, acquiring an unusual level of sensitivity to proteases and
detergents, and in general behaving like a typical peripheral membrane protein.
This observation has led to the suggestion that HisP is deeply embedded in the
membrane (Kerppola et al., 1991; Baichwal et al., 1993). A possible explanation
for the association of HisP with the membrane in the absence of HisQ and HisM
may be found in its predicted tertiary structure, which has been modelled to
include a large loop with some amphipathic characteristics (Mimura et al.,
1991). This loop might allow proper insertion of HisP in the membrane in the
presence, and its improper association in the absence, of the hydrophobic
components. The notion that a segment of the ATP-binding domain protrudes
into or through a pore generated by the transmembrane domains, derives
support from the recent finding that HisP is accessible to proteases and to an
impermeant biotinylating reagent from the exterior surface of the membrane
when the complex has been assembled in the presence of HisQ and HisM, but
not in their absence (Baichwal et al., 1993).
37
While the nature of the interaction between the ATP binding component and
the transmembrane components is uncertain, the interaction must be specific as
the ATP-binding domain from one transporter cannot normally replace that of
another. Indeed, the interaction appears to induce a conformational change in
the ATP-binding domain that alters its biochemical properties (Reyes &
Shuman, 1988; Davidson & Nikaido, 1990). The residues involved in these
interactions are unknown, although gene-fusion studies show that specificity
cannot reside in the N-terminus (Schneider & Walter, 1991).
The two attempts at computer-assisted modelling of ATP-binding
components have generated similar structures by entirely different procedures
(Hyde et al., 1990; Mimura et al., 1991). The core of both models is a nucleotide-
binding pocket which includes five hydrophobic β-sheets and the glycine-rich
Walker motif A, appropriately positioned to interact with ATP and mediate
phosphoryl transfer (Fig. 1.9). Extending from the core nucleotide-binding fold
are loops that have no direct counterpart in adenylate kinase. In one model
(Hyde et al., 1990) these sequences are folded as two separate loops (designated
loops 2 and 3); in the other model (Mimura et al., 1991) they are folded as one
large loop (termed the helical domain). While this difference remains to be
clarified experimentally, it is significant that both models predict that the same
sequences protrude from the nucleotide-binding pocket.
38
Fig. 1.9 Schematic representation of the predicted topology of the ATP-binding component of an ABC transporter (Doige & Ames, 1993). The shaded section corresponds to the Walker motif A. "P-loops" such as this are known to be involved in phosphoryl transfer in many nucleotide-binding proteins. Amino acid residues refer to the HisP sequence. The black box represents a bound mononucleotide.
39
The available data support these model structures. In vitro chemical
modification studies using the photolabelling analogue of ATP, 8-azido ATP are
consistent with the proposed nucleotide-binding fold (Mimura et al., 1990).
Structure function analysis of HisP mutants (Petronilli & Ames, 1991; Shyamala
et al., 1991) showed that the ability of mutant proteins to function in transport
and/or bind ATP was related to the location of the mutation within both the
sequence and the predicted structure. A good correlation was found between
loss of ATP-binding ability (and transport) and location of the mutation within
the predicted ATP-binding pocket, while transport-negative mutants within the
helical domain generally did not affect ATP binding. In addition, a set of
interesting mutations of hisP that enable the mutant HisP proteins to facilitate
transport and ATP hydrolysis even in the total absence of the periplasmic HisJ
protein (Ames & Spudich, 1976; Petronilli & Ames, 1991) also supports the
models because they are all located within the ATP-binding pocket.
Presumably the mutations have changed the binding pocket so that it has
acquired a capacity for unregulated ATP hydrolysis (Petronilli & Ames, 1991;
Shyamala et al., 1991; Speiser & Ames, 1991). Interestingly, although mutations
resulting in unregulated ATP hydrolysis have also been found in the maltose
permease of E. coli (Davidson et al., 1992), these are located in the hydrophobic
domains, indicating that such an effect can arise from a disturbance in the
structure of many regions of the membrane-bound complex.
Each ABC transporter has two ATP-binding domains and both are required
for function. Thus elimination of either one of the two domains of the Opp
oligopeptide transporter of S. typhimurium abolishes function (Hiles et al., 1987).
It is not clear, however, whether the two ATP-binding domains are functionally
equivalent. For transporters such as the histidine permease of S. typhimurium
the two ATP-binding subunits are identical (Kerppola et al., 1991), which
suggests equivalency. The estimated stoichiometry of two ATP molecules
hydrolyzed for each molecule of substrate transported (Mimmack et al., 1989) is
40
also consistent with two equivalent domains, each hydrolyzing one ATP
molecule per transport cycle. However, the observation that genetic analyses of
prokaryotic transport systems in which the ATP-binding components are coded
for by two genes (such as livG/livF in E. coli, braF/braG in Pseudomonas
aeruginosa, and oppD/oppF in both E. coli and S. typhimurium) have usually not
identified the second of the two ATP-binding component genes, has led to the
suggestion (Haney & Oxender, 1992) that most missense mutations and protein
fusions in the second gene are functionally silent. Data for CFTR leads to a
similar conclusion. Many cystic fibrosis mutations fall in the first nucleotide-
binding domain (Cutting et al., 1990; Kerem et al., 1990), yet a mutation in the C-
terminal nucleotide-binding domain that might be expected to disrupt ATP
hydrolysis does not appear to inhibit CFTR function (Anderson et al., 1991a).
In conclusion, while the specific role of each nucleotide-binding domain
remains to be clarified, a working model consistent with the available data
predicts that the ABC proteins have a tightly folded core structure that binds
and hydrolyzes ATP; loops that extend from this core structure interact with the
other components of the transporter and couple, presumably via a
conformational change, the energy of ATP hydroysis to transport.
1.4.4 Periplasmic binding proteins
All known prokaryotic ABC transport systems that facilitate uptake utilize a
periplasmic binding protein. Genetic studies have demonstrated that these
periplasmic proteins are absolutely required for the function of the transport
system with which they are associated. However, they are not integral to the
mechanism of transmembrane translocation itself, since mutants of bacterial
uptake systems can be isolated that function in the absence of the periplasmic
component (Shuman, 1982; Speiser & Ames, 1991).
The periplasmic proteins are relatively easy to purify and many have been
studied in considerable detail. The characterized proteins vary in size from 25
kD for HisJ (Higgins & Ames, 1981) to 59 kD for OppA (Hiles & Higgins, 1986).
41
There is little sequence conservation between binding proteins for different
substrates, the only exceptions being the pairs of periplasmic proteins that
interact with the same core transmembrane complex (e.g. the histidine- and
lysine-arginine-ornithine-binding proteins which deliver substrates to the
HisMQP complex in the membrane of S. typhimurium, and the leucine- and
leucine-isoleucine-valine-binding proteins which deliver substrates to the
LivHMGF complex in the membrane of E. coli). Several binding proteins have
been crystallized and their three dimensional structures determined. These
include the leucine-isoleucine-valine-, leucine-, arabinose-, maltose- and
galactose-binding proteins of E. coli (Adams & Oxender, 1989; Quiocho, 1990;
Vyas et al., 1991). All have a similar structure with two globular domains and a
cleft between that forms the substrate binding site. All substrates appear to be
bound via hydrogen bonds (Quiocho, 1986; Pflugrath & Quiocho, 1988). The
Venus flytrap model in which the proteins undergo a conformational change
upon binding substrate that traps the substrate in the cleft between the two
domains (Mao et al., 1982; Sack et al., 1989), has recently been confirmed for the
lysine-arginine-ornithine-binding protein of S. typhimurium, which has been
purified in both the liganded and unliganded forms (Oh et al., 1993). The
structures determined for the two forms suggest the existence of two states for
the unliganded receptor, open empty and closed empty, that are in dynamic
equilibrium with each other, with the substrate capable of binding to one of the
two globular domains. In the presence of substrate, the closed form is
stabilized by additional interactions between the substrate and the second
globular domain, resulting in a closed liganded form (Oh et al., 1993). In effect,
this model postulates that the act of binding the substrate is responsible for
stabilizing the receptor in its closed conformation.
The periplasmic proteins serve as the initial receptor for transport, delivering
substrate to the membrane-bound components. The in vitro binding
specificities and affinities measured for the purified proteins correspond well
42
with in vivo characteristics of the transport process, which implies that binding
provides the rate-limiting step for transport (Miller et al., 1983). Interaction of
the binding protein-substrate complex with the membrane-associated transport
components has been demonstrated both biochemically and genetically.
Prossnitz et al. (1988) used chemical cross-linking reagents to isolate subunits of
the S. typhimurium histidine transport complex that were cross-linked to each
other in vivo and in vitro. They identified cross-linked products by Western
blot, using antibodies to HisJ and HisQ. Several cross-linked forms were
identified, including one that reacted with antibodies against both HisJ and
HisQ. A previously described mutant of HisJ that is able to bind histidine
normally but does not allow transport (Kustu & Ames, 1974), showed
significantly reduced cross-linking to HisQ relative to the wild-type protein,
supporting the suggestion that the interaction between HisJ and HisQ occurred
during the transport event. That the interaction between binding protein and
transmembrane component is specific was indicated by the finding that MalE
(the binding protein from the maltose transporter of E. coli) could not be cross-
linked to HisQ.
Treptow and Shuman (1988) isolated mutants of malE that restored maltose
uptake in strains of E. coli containing a variety of mutations in malF or malG, the
genes encoding the hydrophobic membrane components of the transporter.
Many of the suppressors were found to be allele specific i.e. a malE mutation
selected in one mutant background was found to be unable to restore transport
in other mutant backgrounds. Mutations mapped widely over the malE gene,
with mutations in the N-terminal end of malE tending to suppress malF alleles
and mutants in the C-terminal end tending to suppress malG alleles. These
results suggest that interaction involves both domains of the periplasmic
binding protein and both hydrophobic transmembrane components.
Interaction of the binding protein with the membrane-associated transport
components does not occur in the absence of bound substrate (Prossnitz et al.,
43
1988; Doige & Ames, 1993), suggesting that the conformational change
undergone by the periplasmic protein upon binding of the substrate enables
interaction with the appropriate complex of membrane proteins.
The precise function of binding proteins in transport is still not fully
understood. Their role has commonly been ascribed to that of increasing the
effective concentration of substrate in the periplasm. However, Hennge and
Boos (1983) have argued that the effect of a high concentration, in the
periplasm, of binding protein with a high affinity for substrate, is to generate a
high concentration of liganded binding protein, not of ligand itself. Estimates
of the concentration of the binding protein in the periplasm vary from 0.5 to
10mM (Silhavy et al., 1975; Hengge & Boos, 1983; Ames, 1986). With
dissociation constants for ligands in the range of 10-5 to 10-7 M, this means that
even when low concentrations of ligand are present in the medium, the
concentration of liganded binding protein in the periplasm can be in the
millimolar range (Hengge & Boos, 1983). The concentration of the free ligand in
the periplasm will be the same as it is outside the periplasm. This by itself
effectively demonstrates that the membrane complex recognizes the liganded
binding protein and not the free ligand, since strains not expressing binding
proteins do not show appreciable transport from the high-affinity system, even
when the ligand is present in millimolar concentrations, whereas wild-type
transport systems show transport affinities in the micromolar range (Hengge &
Boos, 1983; Treptow & Shuman, 1985).
It has been suggested that binding proteins facilitate the movement of
substrate within the periplasm. The periplasm of Gram-negative bacteria is a
gel-like matrix and , at low substrate concentrations, diffusion may be limiting
(Brass et al., 1986). However, the recent identification of binding proteins in
Mycoplasma and other Gram-positive species, which do not have a periplasm
(Dudler et al., 1988; Gilson et al., 1988; Alloing et al., 1990; Perego et al., 1991;
Rudner et al., 1991), argues against this. An adaptation of this hypothesis is that
44
the binding proteins enhance transport by restricting diffusion to two, rather
than three, dimensions. Diffusion of the binding protein-substrate complex in
two dimensions is expected to enhance the efficiency at which substrate is
delivered to the membrane transport complex, compared with three-
dimensional diffusion of the unbound substrate in solution. In Gram-positive
species the binding proteins are anchored to the membrane by a lipid group,
and diffusion is consequently restricted to two dimensions. In Gram-negative
species the dimensions of the periplasm also effectively restrict diffusion of the
binding protein to two dimensions. Indeed, the periplasmic binding protein of
the maltose transporter in E. coli still functions efficiently when anchored to the
membrane via a non-cleavable signal sequence (Fikes & Bassford, 1987).
It is also possible that the periplasmic protein imposes directionality on
transport. No ABC transporter has yet been found that can mediate both
uptake and export of a substrate, yet comparison of the membrane-associated
proteins of uptake and export systems does not identify any feature that allows
the two to be distinguished. Since all known bacterial uptake systems require a
periplasmic component, while no export system does, it is conceivable that the
presence of the binding protein determines the directionality of the transporter.
Consistent with such a suggestion is the fact that interaction of the binding
protein-substrate complex with the membrane-associated domains is required
for ATP hydrolysis (Bishop et al., 1989; Petronilli & Ames, 1991), which implies
an induced conformational change in the membrane-associated components of
the transporter.
1.4.5 Mechanism of solute translocation
The presence of conserved ATP-binding motifs (Walker motifs) in all
characterized ABC proteins indicates a role for ATP in solute translocation by
ABC transporters, and recent studies have provided good evidence that ATP
hydrolysis by the transporters themselves provides the driving force for solute
accumulation by binding protein-dependent systems in Gram-negative bacteria.
45
The OppD and HisP proteins of S. typhimurium, and the MalK protein of E.
coli, have been shown to bind ATP and/or a variety of ATP-affinity analogues
(Hobson et al., 1984; Higgins et al., 1985). The Km of the histidine transporter for
ATP is estimated to be about 200µM, appropriately less than the normal
intracellular ATP concentration (Ames et al., 1989). Furthermore, the ABC
proteins not only bind ATP, but ATP-binding is essential for function. Thus
mutation of the ATP-binding site of several ABC transporters inhibits activity
(Hiles et al., 1987; Joshi et al., 1989; Higgins et al., 1990; Ames & Joshi, 1991;
Shyamala et al., 1991). Additionally, in cell membrane-derived vesicle systems,
histidine and maltose transport by E. coli shows an absolute requirement for
ATP (Dean et al., 1989; Prossnitz et al., 1989).
A requirement for ATP does not necessarily imply that hydrolysis energizes
transport. ATP binding could, potentially, serve a structural or regulatory role.
However, non-hydrolyzable analogues of ATP are unable to support active
transport (Ames et al., 1989), and in vivo studies on several E. coli ABC
transporters demonstrated that ATP hydrolysis is dependent upon, and occurs
concomitantly with, substrate translocation (Mimmack et al., 1989), while no
ATP consumption was observed for non-ABC transporters. Reconstitution of
transporters in vesicles (Ames et al., 1989; Dean et al., 1989; Prossnitz et al., 1989)
and in proteoliposomes (Bishop et al., 1989; Davidson & Nikaido, 1990) also
revealed the complete dependence of transport on ATP hydrolysis, and vice
versa.
There is, therefore, no doubt that ATP hydrolysis is required for transport
and that domains of the transporters themselves can bind ATP. However,
although the simplest interpretation of these facts is that the ABC domains
themselves hydrolyze ATP and directly couple the energy of hydrolysis to the
transport process, this has yet to be formally demonstrated.
Besides ATP, GTP and CTP can also energize histidine transport in vesicles
(Bishop et al., 1989). However, the low cytoplasmic pools of these nucleotides
46
and their poor affinities for the transport proteins (Hobson et al., 1984; Higgins
et al., 1985) make it unlikely that they serve a significant role in energizing
transport in vivo.
The stoichiometry of ATP hydrolysis is not firmly established. In vesicle
systems, stoichiometries of 5 (Bishop et al., 1989) and 1.4 to 17 (Davidson &
Nikaido, 1990) have been reported. These high and variable values may be due
to damage incurred by the complex during reconstitution resulting in the
uncoupling of hydrolysis and transport (Bishop et al., 1989). Estimates using
whole cells suggest a stoichiometry of close to two ATP molecules hydrolysed
per molecule of substrate transported (Mimmack et al., 1989).
While it has been demonstrated that ATP is only hydrolyzed upon
interaction of the periplasmic binding protein with the membrane-bound
complex (Bishop et al., 1989; Petronilli & Ames, 1991), it is not known how the
signal for ATP hydrolysis is transduced. Similarly, the mechanism by which
the energy of ATP hydrolysis is coupled to the transport process is unknown.
There is no evidence that a phosphorylated protein intermediate is involved
(Ames & Nikaido, 1981), and it is generally assumed that ATP binding and
hydrolysis induces a conformational change in the ATP-binding domain, which
is transmitted, via domain-domain interactions, to the transmembrane subunits
that mediate translocation across the membrane. The helical domain (loop 2/3
region) of the ATP-binding components is a good candidate for involvement in
this conformational transduction, particularly since mutations in this loop
region can uncouple ATP hydrolysis from the transport event, without altering
ATP binding or hydrolysis (Petronilli & Ames, 1991).
The nature of the interaction between substrate and transmembrane domains
is also uncertain. At one extreme the transmembrane domains might simply
form a pore in the membrane which the ATP-binding domains serve to open
and close. However, the transmembrane domains do not simply form an open
channel in the absence of the ATP-binding domains (Higgins, 1992).
47
Furthermore, such an arrangement could not account for the directionality, and
ability to concentrate solute against a gradient, of ABC transporters. Classical
kinetic analyses of transporters imply a conformational change exposing
substrate-binding sites to opposing sides of the membrane (Stein, 1990). A
better model may therefore be that substrate interacts with a binding site
located within a pore-like structure generated by the transmembrane domains,
and subsequent ATP hydrolysis reorientates this site to expose it to the opposite
face of the membrane. However, it is equally possible that reorientation of the
binding site is facilitated by substrate-binding and that ATP hydrolysis resets
the orientation in order to impose directionality and the ability to actively
accumulate substrate.
While the great majority of ABC transport systems mediate active transport,
two apparently typical ABC transporters, CFTR and MDR, have recently been
associated with chloride channel activity (Anderson et al., 1991b; Kartner et al.,
1991; Gill et al., 1992; Valverde et al., 1992). This has highlighted the problem of
defining the physical differences between channels and transporters.
Transporters are generally considered to be enzyme-like, interacting
stoichiometrically with their substrate and undergoing defined conformational
changes during each transport cycle. In contrast, channels are viewed as holes
which, when open, allow non-stoichiometric passage of molecules with
appropriate characteristics. This distinction is based on several experimental
observations: (i) Kinetic and biochemical characterization of several
transporters has revealed enzyme-like intermediate states during each transport
cycle, often interpreted as the alternate exposure of a substrate-binding site at
each face of the membrane (Stein, 1990). (ii) Many transporters have been
shown to be stoichiometrically coupled to other events such as ATP hydrolysis
or proton movement. (iii) The turnover number of transporters is restricted by
the enzyme-like conformational changes. In contrast, channels are ultimately
diffusion-limited, and their turnover number can be several orders of
48
magnitude greater than has ever been measured for a transporter. (iv)
Channels simply facilitate equilibration of substrate in response to
concentration and electrochemical gradients. Transporters, in contrast, can
utilize energy to concentrate substrate against a gradient.
In the case of ABC systems the difference between a transporter and a
channel may reside in the organization of the transmembrane domains. At one
extreme , the translocation pathway formed by the transmembrane domains of
ABC transporters and channels may be very similar, with a small structural
change allowing a transporter to function as a channel. At the other extreme,
the transmembrane pathways may be very different. One suggestion (Higgins,
1992) is that transport activity might be associated with the monomeric
membrane-bound complex, while channel function might involve a different
transmembrane translocation pathway generated by the oligomeric association
of complexes (Fig. 1.10).
Fig. 1.10 Schematic representation of transmembrane domains of an ABC transporter viewed from the membrane surface, showing the pathway through the membrane (*) provided by monomeric complexes (A), and a potential alternative pathway (B) provided by oligomerization (Higgins, 1992).
49
1.4.6 The role of binding protein-dependent transporters
In general ABC transporters are low capacity but high affinity (Km 0.01-1µM)
systems and can accumulate substrate against very large concentration
gradients (>10,000-fold). This is in contrast to members of the twelve-
membrane-spanning-helix family of transporters (Section 1.3), which as a
general rule are low affinity but high capacity systems that, for thermodynamic
reasons, cannot accumulate substrate against large concentration gradients
(Hengge & Boos, 1983).
For some substrates, E. coli possesses both an ABC transporter and a twelve-
transmembrane-helix transporter. The reason for this apparent duplication of
effort may be that under certain growth conditions changing energy status
might favour the use of transporters energized by alternate means (twelve-
transmembrane-helix transporters are energized by the electrochemical
gradient or by co- or countertransport). However, it has also been proposed
(Higgins, 1992; Doige & Ames, 1993) that the kinetic properties of twelve-
transmembrane-helix transporters make them suitable for bulk uptake of
carbon and nitrogen sources for growth, while those of ABC transporters
indicate a scavenging role that becomes important in environments where
nutrients are present at extremely low concentrations. In the specific case of
amino acid uptake, it has been suggested that ABC transporters may be
involved in the recapture of biosynthetically produced amino acids that would
otherwise be lost from the cell (Ames, 1972; Antonucci & Oxender, 1986).
1.4.7 Regulation of ABC transporters
As with other bacterial uptake systems, ABC transporters are commonly
regulated at the transcriptional level (Haney & Oxender, 1992), and many are
only expressed in the presence of their specific substrates. However, the
activity of ABC transporters, once expressed, may also be regulated. Uptake of
sugars such as maltose (Nelson & Postma, 1984) is inhibited by direct
50
interaction between a component of glucose metabolism (Enzyme III) and the
ABC transporter, such that glucose is used as the preferred carbon source.
Regulation of transport through alteration of binding protein affinity by
phosphorylation has also been described (Celis, 1984): E. coli was incubated
with 32Pi and an osmotic shock fluid separated by gel chromatography. One
labelled fraction that exhibited binding of arginine was shown to contain
phosphorylated lysine-arginine-ornithine-binding protein. Unphosphorylated
lysine-arginine-ornithine-binding protein was also recovered and was found to
have a 50-fold greater affinity for substrate than the phosphorylated protein
(dissociation constants for arginine of the modified and unmodified proteins
are 5.0µM and 0.1µM, respectively). A mutant has been isolated that does not
express the kinase that phosphorylates the binding protein (Celis, 1990). This
kinase, which has been purified (Celis, 1990), serves to deactivate the binding
protein and limit entry of basic amino acids into the cell.
There is also evidence that the intracellular level of substrate may regulate
uptake. In a study of histidine transport in reconstituted proteoliposomes,
which demonstrates the unidirectional nature of substrate translocation in this
system (Doige & Ames, 1993), it was found that accumulated histidine did not
exit from the proteoliposomes, and that incorporation of ADP, inorganic
phosphate, and histidine inside the proteoliposomes did not result either in the
exit of histidine or the synthesis of ATP. Furthermore, both the internally
accumulated histidine and the hydrolytically produced ADP inhibited
transport, suggesting that the internal pool of histidine (and ADP) regulates
translocation, presumably via a substrate-recognizing site on the cytoplasmic
surface of the membrane-bound complex (Doige & Ames, 1993).
51
1.5 AMINO ACID TRANSPORT IN RHIZOBIUM
Poole et al. (1985) studied the stereospecific and kinetic properties of L-
glutamate transport in R. leguminosarum biovar viciae strain 3841. They found
not only that glutamate uptake is competitively inhibited by a wide range of
other amino acids, but also that intracellular L-leucine can be exchanged out of
strain 3841 by L-glutamate. These results strongly suggest that strain 3841
possesses a general amino acid transport system.
The fact that L-glutamate transport failed to show any substantial deviation
from Michaelis-Menten kinetics suggests that the general permease may be the
only kinetically significant transport system for L-glutamate in strain 3841. The
existence of specific transport systems for glycine and leucine, operating in
addition to the common system in strain 3841, was indicated by the failure of
large excesses of L-glutamate to completely inhibit uptake of these substrates.
Glenn et al. (1991) studied proline transport in cowpea Rhizobium NGR234,
and found that a 5-fold excess of many amino acids (including glutamate)
caused a 70-90% inhibition of L-proline uptake. In addition, [14C]proline taken
up by NGR234 cells over 3 minutes, was found to be exchangeable with
extracellular valine, histidine or isoleucine but not glutamate. This suggests
that proline is carried by a general amino acid transporter. The fact that
intracellular proline is not excreted in the presence of extracellular glutamate
may indicate that glutamate transport is distinct from that of proline, or may
reflect the relative affinities of these substrates for the common carrier.
Glutamate transport in cowpea Rhizobium MNF2030 and R. leguminosarum
biovar trifolii MNF1000 is inhibited 50-100% by the addition of a 10-fold excess
of a variety of other amino acids (Jin et al., 1990). However, kinetic analysis of
glutamate uptake in these strains indicates the presence of two glutamate
transport systems in both cases (a high affinity (low Km) system with a
relatively low capacity, and a low affinity (high Km) system with a greater
capacity).
52
In a recent study (Watson et al., 1993), aspartate transport in R. meliloti was
found to be mediated by at least two systems; the Dct system (apparent Km for
aspartate transport of 10mM) and a second system (apparent Km of 1.5mM)
which was competitively inhibited by glutamate. There was also some kinetic
evidence for the existence of a third, high affinity, system. In dct mutants
growing on aspartate, high initial rates of aspartate uptake preceded the lower
steady-state value. This was taken to be indicative of the presence of an internal
aspartate pool in R. meliloti, and a rapid initial equilibration of aspartate inside
the cell with that outside by an exchange process.
The capacity of the glutamate-aspartate system is sufficient to allow growth
of R. meliloti dct mutants on aspartate as nitrogen source, but not as both carbon
and nitrogen source. Furthermore, this system is not directly induced by
aspartate, suggesting that it may be regulated, at least partly, by nitrogen
requirement. Previously reported experiments with chemostat cultures have
demonstrated that glutamate uptake in R. leguminosarum strain 3841 is
regulated by nitrogen availability (Poole et al., 1987).
53
1.6 REGULATION INVOLVING NTRC
Transcriptional regulation of nitrogen metabolism in enteric bacteria has
been found to involve five genes (Reitzer & Magasanik, 1987; Merrick, 1988):
ntrA (also designated glnF and rpoN), which codes for the alternative sigma
factor, σ54; ntrBC (also designated glnLG), which are located downstream of
glnA (the structural gene for glutamine synthetase) and code for the NtrB (also
called NRII) and NtrC (also called NRI) proteins belonging to the family of
histidine kinase sensors and response regulators (Stock et al., 1989); and finally
glnD and glnB, which code for a uridylyltransferase and PII protein,
respectively. In E. coli, glnB and glnD are expressed independently of the
nitrogen status of the cell and of σ54 (van Heeswijk et al., 1993). The glnA-ntrBC
operon has three promoters; one, ntrBp, located between glnA and ntrB, and
two, glnAp1 and glnAp2, located upstream of glnA. Transcription initiated at
the σ70-dependent promoters glnAp1 and ntrBp serves to maintain a low
intracellular concentration of glutamine synthetase, NtrB and NtrC (Reitzer &
Magasanik, 1986) under conditions of nitrogen-excess. Under nitrogen-
limitation expression of the glnA-ntrBC operon is primarily due to glnAp2.
Initiation of transcription from this promoter requires σ54 and the upstream
binding of the activator protein NtrC.
The cellular nitrogen status, as indicated by the intracellular glutamine:α-
ketoglutarate ratio, is sensed by the uridylyltransferase. It responds to
nitrogen-limitation (low glutamine:α-ketoglutarate ratio) or nitrogen-excess
(high glutamine:α-ketoglutarate ratio) by uridylylating or deuridylylating PII,
respectively (Bueno et al., 1985; Holtel & Merrick, 1989). In its unuridylylated
form, PII interacts with NtrB causing it to be a protein phosphatase (Bueno et al.,
1985; Holtel & Merrick, 1989). However, uridylylation of PII prevents this
interaction allowing NtrB to act as a kinase towards NtrC by transferring
phosphate from ATP, and it is the phosphorylated form of NtrC that acts as an
activator at glnAp2 (Ninfa & Magasanik, 1986; Reitzer & Magasanik, 1987;
54
Holtel & Merrick, 1988; Merrick, 1988). Thus nitrogen-limitation leads to
increased synthesis of glutamine synthetase, NtrB and NtrC.
Glutamine synthetase is the key enzyme in the high-affinity, glutamine
synthetase/glutamate 2-oxoglutarate amino transferase, pathway for ammonia
assimilation, and the same cascade that controls the activity of NtrB as a kinase,
also controls post-translational regulation of glutamine synthetase. Glutamine
synthetase can be converted into a less active form by adenylylation. The
adenylyltransferase (product of the glnE gene) which catalyzes both the
adenylylation and deadenylylation of glutamine synthetase is controlled by PII.
Unuridylylated PII (nitrogen-excess conditions) interacts with the
adenylyltransferase enabling it to catalyze the adenylylation of glutamine
synthetase, while uridylylated PII (nitrogen-limited conditions) causes the
adenylyltransferase to catalyze deadenylylation. Thus the post-translational
regulation of glutamine synthetase mirrors its transcriptional regulation.
In addition to acting as a positive regulator, NtrC can also act as a negative
regulator. Indeed, both glnAp1 and ntrBp are negatively controlled by NtrC. At
ntrBp an NtrC binding site overlaps the transcriptional start and the -10 region
of the promoter, so that binding of NtrC impedes RNA polymerase binding and
ntrBC expression (Reitzer & Magasanik, 1983; Dixon, 1984; Ueno-Nishio et al.,
1984; MacFarlane & Merrick, 1985). In Klebsiella pneumoniae, mutations of NtrB
that result in constitutive phosphorylation of NtrC, are found also to cause
strong repression of ntrBp. This repression is independent of nitrogen status,
and is greater than the repression observed in an ntrB deletion strain
(MacFarlane & Merrick, 1987), suggesting that at ntrBp, phosphorylated NtrC
binds more effectively than the unphosphorylated form.
At glnAp1 in E. coli and S. typhimurium two NtrC binding sites with potential
for inhibiting expression from this promoter have been identified (Ames &
Nikaido, 1985; Hirschman et al., 1985; Reitzer & Magasanik, 1985). Site 1
overlaps the -35 region of the promoter, while site 2 overlaps the transcriptional
55
start site. In K. pneumoniae only site 2 is present (Dixon, 1984; Hawkes et al.,
1985).
In addition to controlling the level of glutamine synthetase, the Ntr system
also regulates the expression of other genes related to the nitrogen status of the
cell, including genes involved in amino acid uptake (Merrick, 1988).
In S. typhimurium uptake of histidine, arginine, lysine and ornithine is
mediated by a single membrane associated complex composed of the proteins
HisQ, HisM and HisP in a 1:1:2 ratio (Section 1.4.1). This complex interacts with
two periplasmic binding proteins; one, HisJ is specific for histidine, while the
other, ArgT, binds arginine, lysine and ornithine (Higgins et al., 1982b). The
argT gene lies directly upstream of the hisJQMP operon, and is transcribed,
under the control of the promoter argTr, as a monocistronic unit. Transcription
of hisJQMP is controlled by the promoter dhuA, which is located between argT
and hisJ. The observation that transcription initiated by dhuA is elevated in an
ntrB mutant (Higgins & Ames, 1982; Stern et al., 1984), has led to the erroneous
suggestion that dhuA is negatively regulated by NtrC (Haney & Oxender, 1992).
In fact, studies of lacZ fusions to argTr and dhuA in ntrA- and ntrC- strains have
shown both these promoters to be σ54-dependent, and activated by NtrC under
nitrogen-limitation (Ames & Nikaido, 1985). Mutational analysis has enabled
the site of action of NtrC at argTr to be described in detail (Schmitz et al., 1988).
Uptake of glutamine, glutamate and aspartate in S. typhimurium is also under
nitrogen control, and expression of genes encoding transporters of these amino
acids is likely to be regulated by the Ntr system (Kustu et al., 1979).
Glutamine transport in E. coli is positively regulated by NtrC, with nitrogen-
limitation resulting in increased expression of the glnHPQ operon (Nohno et al.,
1986; Claverie-Martin & Magasanik, 1991).
The pattern of Ntr control in Rhizobiaceae differs from that found in enteric
bacteria, with glnA and ntrBC being located in separate operons. In R.
leguminosarum biovar phaseoli, ntrBC are found in the operon ORF1-ntrBC which
56
is transcribed from a σ70-dependent promoter independently of nitrogen status
(Patriarca et al., 1993; Amar et al., 1994). The function of ORF1 is currently
unknown. The increase in transcription of ntrC observed in an ntrC mutant has
been taken to indicate that this operon is subject to negative autoregulation by
NtrC, and a putative repressor-binding site for NtrC has been identified in the
promoter region for the operon (Patriarca et al., 1993). These findings indicate
that, unlike the situation in enteric bacteria, changes in the level of NtrC
phosphorylation, and not its intracellular concentration, are sufficient to
activate or repress transcription from its target promoter(s) in R. leguminosarum
biovar phaseoli.
At least two glutamine synthetase isozymes, GSI and GSII, encoded by the
glnA and glnII genes, respectively, are present in Rhizobiaceae (Darrow & Knots,
1977; Fuchs & Keister, 1980; Filser et al., 1986; Colonna-Romano et al., 1987;
Patriarca et al., 1992), with a third isozyme GSIII, encoded by glnT, having been
identified in R. leguminosarum (Chiurazzi et al., 1992) and R. meliloti (de Bruijn et
al., 1989). GSI is similar to the glutamine synthetase of enteric bacteria and is
subject to adenylylation. Transcription of glnA, as part of the glnBA operon, can
be initiated by two promoters, one upstream of glnB and one located between
glnB and glnA (Chiurazzi & Iaccarino, 1990). Although in free living cells, glnA
is mostly transcribed from the promoter upstream of glnB, and the activity of
this promoter is positively regulated by NtrC (Chiurazzi & Iaccarino, 1990),
synthesis of GSI is found not to be greatly affected by nitrogen supply (Carlson
et al., 1987; Szeto et al., 1987; Rossi et al., 1989; Chiurazzi & Iaccarino, 1990; Amar
et al., 1994). In contrast, synthesis of the eukaryote-like GSII is fully dependent
on positive control by NtrC in response to nitrogen source availability, with
glnII being transcribed as a monocistronic unit (Carlson et al., 1987; Martin et al.,
1988; de Bruijn et al., 1989; Rossi et al., 1989; Shatters et al., 1989; Patriarca et al.,
1992). The increase in transcription of glnII and glnB observed in a glnB mutant
of R. leguminosarum biovar phaseoli, has been shown not to result from an
57
increase in the level of NtrC, and is consistent with the suggestion that changes
in the degree of phosphorylation of NtrC are solely responsible for the
regulation of NtrC-controlled promoters in this species (Amar et al., 1994).
It has been suggested that the different patterns of regulation observed for
ntrBC, glnBA and glnII may be the result of differences in the affinity of NtrC for
the upstream activator sequences of glnBA and glnII, and the putative
repressor-binding site for ORF1-ntrBC (Patriarca et al., 1993). Alternatively, it is
also possible that additional, as yet unidentified, regulatory factor(s) could play
a role in the transcriptional regulation of some or all of these genes.
58
CHAPTER 2 MATERIALS AND METHODS
59
2.1.1 Bacterial strains
The bacterial strains, plasmids and bacteriophages used in this work are
listed in Table 2.1.
Table 2.1 Bacterial strains, plasmids and bacteriophages used
Strain, phage or plasmid
Description Source or Reference
Bacterium R. leguminosarum 3841 StrR derivative of strain 300 Johnston and
Beringer (1975) RU116 Strain 3841 sucD::Tn5 This work RU117 Aspartate toxic escape, putative
Tn5 mutant of 3841 This work
RU118 Aspartate toxic escape Tn5 mutant of 3841
This work
RU126 Aspartate toxic escape, putative Tn5 mutant of 3841
This work
RU137 Strain 3841 phbC::Tn5 This work RU140 Aspartate toxic escape, putative
Tn5 mutant of 3841 This work
RU151 Aspartate toxic escape, putative Tn5 mutant of 3841
This work
RU154 Aspartate toxic escape, putative Tn5 mutant of 3841
This work
RU156 Strain 3841 sucA::Tn5 This work RU158 Aspartate toxic escape, putative
Tn5 mutant of 3841 This work
RU216 3841 containing genomic Tn5(KmR) insert introduced by transduction from strain RU116
This work
RU237 3841 containing genomic Tn5(KmR) insert introduced by transduction from strain RU137
This work
RU256 3841 containing genomic Tn5(KmR) insert introduced by transduction from strain RU156
This work
RU368 Strain 3841 containing pMP220 This work RU438 Strain 3841 containing pRU3024 This work RU439 Strain 3841 containing pRU135 This work RU441 Strain 3841 containing pRU3026 This work RU442 Strain 3841 containing pRU3027 This work RU443 Strain 3841 containing pRU3028 This work
60
Strain, phage or plasmid
Description Source or Reference
RU444 Strain RU116 containing
pRU3004 This work
RU449 Strain RU137 containing pRU3004
This work
RU453 Strain RU156 containing pRU3004
This work
RU500 Strain 3841 containing pRU3029 This work RU502 Strain 3841 containing pRU3031 This work RU503 Strain 3841 containing pRU3032 This work RU504 Strain 3841 containing pRU3033 This work RU506 Strain 3841 containing pRU3035 This work RU510 Strain 3841 containing pRU3039 This work RU515 Strain 3841 containing pRU3044 This work RU517 Strain 3841 containing pRU3046 This work RU519 Strain 3841 containing pRU3048 This work RU522 Strain 3841 containing pRU3049 This work RU526 Strain 3841 containing pRU3053 This work RU541 pRU3026 homogenote in strain
3841 This work
RU542 pRU3027 homogenote in strain 3841
This work
RU543 pRU3028 homogenote in strain 3841
This work
RU622 Strain 3841 containing pAR36A This work RU631 pRU3029 homogenote in strain
3841 This work
RU632 pRU3031 homogenote in strain 3841
This work
RU633 pRU3032 homogenote in strain 3841
This work
RU634 pRU3035 homogenote in strain 3841
This work
RU635 pRU3039 homogenote in strain 3841
This work
RU636 pRU3046 homogenote in strain 3841
This work
RU637 pRU3048 homogenote in strain 3841
This work
RU638 pRU3049 homogenote in strain 3841
This work
RU639 pRU3053 homogenote in strain 3841
This work
RU640 Strain 3841 containing pRU191 This work
61
RU641 Strain 3841 containing pRU192 This work
62
Strain, phage or plasmid
Description Source or Reference
RU724 pRU3067 homogenote in strain
3841 This work
RU725 pRU3059 homogenote in strain 3841
This work
RU726 pRU3069 homogenote in strain 3841
This work
RU733 pRU3061 homogenote in strain 3841
This work
RU735 Strain RU542 containing pRU191 This work RU736 Strain RU542 containing pRU307 This work RU737 Strain RU542 containing pRU308 This work RU738 Strain RU542 containing pRU309 This work RU739 Strain RU542 containing pRU310 This work RU740 Strain RU542 containing pRU311 This work RU741 Strain RU542 containing pRU312 This work RU742 Strain RU542 containing pRU313 This work RU743 Strain RU634 containing pRU191 This work RU744 Strain RU634 containing pRU307 This work RU745 Strain RU634 containing pRU308 This work RU746 Strain RU634 containing pRU309 This work RU747 Strain RU634 containing pRU310 This work RU748 Strain RU634 containing pRU311 This work RU749 Strain RU634 containing pRU312 This work RU750 Strain RU634 containing pRU313 This work RU751 Strain RU636 containing pRU191 This work RU752 Strain RU636 containing pRU308 This work RU753 Strain RU636 containing pRU309 This work RU754 Strain RU636 containing pRU310 This work RU755 Strain RU636 containing pRU312 This work RU756 Strain RU636 containing pRU313 This work RU757 Strain RU543 containing pRU191 This work RU758 Strain RU543 containing pRU309 This work RU759 Strain RU543 containing pRU310 This work RU760 Strain RU543 containing pRU313 This work RU889 Strain RU116 containing
pRU3028 This work
RU897 Strain RU156 containing pRU3028
This work
RU891 Strain RU116 containing pAR36A
This work
RU899 Strain RU156 containing pAR36A
This work
RU913 Strain 3841 containing pRK415-1 This work
63
RU914 Strain 3841 containing pIJ1891 This work
64
Strain, phage or plasmid
Description Source or Reference
RU915 Strain 3841 containing pRU309 This work RU916 Strain 3841 containing pRU310 This work RU917 Strain 3841 containing pRU313 This work RU918 Strain 3841 containing pRU388 This work RU919 Strain 3841 containing pRU389 This work RU929 Strain 3841 ntrC::Ω Reid (1995) RU972 Strain 3841 containing pRU3080 This work RU974 Strain 3841 containing pRU3082 This work RU975 Strain 3841 containing pRU3083 This work RU976 Strain 3841 containing pRU3084 This work RU977 Strain 3841 containing pRU3085 This work RU978 Strain 3841 containing pRU3086 This work RU980 Strain RU929 containing
pRU3028 This work
RU981 Strain RU929 containing pRU3031
This work
RU982 Strain RU929 containing pRU3033
This work
RU983 Strain RU929 containing pRU3035
This work
RU984 Strain RU929 containing pRU3046
This work
RU986 Strain RU929 containing pRU3082
This work
RU988 Strain RU929 containing pRU3086
This work
RU990 Strain 3841 containing pRU393 This work RU999 pRU3082 homogenote in strain
3841 This work
RU1000 pRU3084 homogenote in strain 3841
This work
RU1001 pRU3086 homogenote in strain 3841
This work
RU1002 Strain RU116 containing pMP220
This work
RU1003 Strain RU156 containing pMP220
This work
RU1013 Strain RU929 containing pRU3024
This work
RU1017 pRU3028 homogenote in strain RU929
This work
RU1018 pRU3035 homogenote in strain RU929
This work
65
Strain, phage or plasmid
Description Source or Reference
RU1019 pRU3046 homogenote in strain
RU929 This work
RU1024 Strain RU116 containing pRU3024
This work
RU1025 Strain RU156 containing pRU3024
This work
RU1027 pRU3033 homogenote in strain RU929
This work
RU1029 pRU3082 homogenote in strain RU929
This work
RU1030 pRU3086 homogenote in strain RU929
This work
RU1069 Strain 3841 containing pRU3089 This work RU1070 Strain 3841 containing pRU3090 This work RU1071 Strain 3841 containing pRU3091 This work E. coli 803 met- gal-. Wood (1966) DH5α supE44 ∆lacU169 (φ80 lacZ∆M15)
hsdR17 recA1 endA1 gyrA96 thi-1 relA1.
Hanahan (1983); Bethesda Research Laboratories (1986)
JC5412 Low glutamate uptake: does not grow on glutamate as sole carbon and nitrogen source
Willetts and Mount (1969)
MC1061 hsdR mcrB araD139 ∆(araABC-leu)7679 ∆lacX74 galU galK rpsL thi
Meissner et al. (1987)
S17-1 pro hsdR recA [RP4-2(Tc::Mu) (Km::Tn7)]; RP4 integrated into its chromosome.
Simon et al. (1983)
RU1050 Strain JC5412 containing pRU310 This work Plasmid pAR36A R. leguminosarum glnII::lacZ
translational fusion in pMP220 Patriarca et al. (1992)
pBC KS+ Phagemid, pUC19 derivative, f1 origin of replication, ColE1 replicon; CmR
Stratagene Research Laboratories.
pBluescript SK+ Phagemid, pUC19 derivative, f1 origin of replication, ColE1 replicon; AmpR
Stratagene Research Laboratories.
pIJ1891 pLAFR3 containing the pUC118 polylinker; TcR
A. Downie
66
Strain, phage or plasmid
Description Source or Reference
pJQ200 pACYC derivative, P15A origin
of replication; GmR Quandt and Hynes (1993)
pLAFR1 Wide host range P-group cloning vector, mobilizable RK2 cosmids; TcR
Friedman et al. (1982)
pMP220 IncP transcriptional fusion vector; TcR
Spaink et al. (1987)
pPHJI1 P-group chaser plasmid; GmR Hirsch and Beringer (1984)
pRK2013 ColE1 replicon with RK2 tra genes; helper plasmid used for mobilizing P- and Q-group plasmids; NmR KmR.
Figurski and Helinski (1979)
pRK415-1 Broad host range P-group cloning vector; TcR
Keen et al. (1988)
pSP72 Promoterless cloning vector; AmpR
Promega
pSUP202-1::Tn5 mob; KmR Simon et al. (1983) pRU32 pBluescript SK+ carrying 9.6kb
Tn5-bearing EcoRI fragment from strain RU116
This work
pRU34 pBluescript SK+ carrying 10.2kb Tn5-bearing EcoRI fragment from strain RU156
This work
pRU36 Sub-clone of pRU32 generated by BamHI digestion followed by ligation
This work
pRU37 Sub-clone of pRU32 generated by HindIII digestion followed by ligation
This work
pRU40 Sub-clone of pRU34 generated by BamHI digestion followed by ligation
This work
pRU41 Sub-clone of pRU34 generated by HindIII digestion followed by ligation
This work
pRU99 pBluescript SK+ carrying 8.9kb Tn5-bearing EcoRI fragment from strain RU137
This work
pRU100 Sub-clone of pRU99 generated by HindIII digestion followed by ligation
This work
67
pRU101 pBC KS+ carrying 3.2kb KmR BamHI fragment from pRU99
This work
68
Strain, phage or plasmid
Description Source or Reference
pRU133 pRK415-1 carrying 3.2kb EcoRI
fragment from pRU3024
pRU135 pRK415-1 carrying 10.2kb HindIII fragment from pRU3024
This work
pRU149 pRK415-1 carrying 6.0kb HindIII-SstI fragment from pRU135
This work
pRU165 Sub-clone of pRU135 generated by XbaI-MluI digestion followed by blunted-end ligation.
This work
pRU180 pRK415-1 carrying 4.6kb SstI fragment from pRU3004
This work
pRU181 pRK415-1 carrying 2.5kb SstI fragment from pRU3004
This work
pRU182 pRK415-1 carrying 6.3kb PstI fragment from pRU3004
This work
pRU185 Sub-clone of pRU135 generated by XbaI-ClaI digestion followed by blunted-end ligation.
This work
pRU186 pRK415-1 carrying 5.2kb NcoI fragment from pRU135 cloned in BamHI site
This work
pRU189 pBluescript SK+ carrying 5.4kb MluI-ClaI fragment from pRU135, cloned in EcoRV site
This work
pRU190 pBluescript SK+ carrying 5.4kb MluI-ClaI fragment from pRU135, cloned in EcoRV site in opposite orientation to pRU189
This work
pRU191 pRK415-1 carrying 5.4kb XbaI-HindIII fragment from pRU189
This work
pRU192 pRK415-1 carrying 5.4kb XbaI-HindIII fragment from pRU190
This work
pRU194 pJQ200 carrying 10.7kb Tn5-lacZ-bearing salI fragment from pRU3059
This work
pRU222-247 Ordered BstUI deletions of pRU189
This work
pRU248-275 Ordered BstUI deletions of pRU190
This work
pRU276 pRK415-1 carrying 4.4kb EcoRI fragment from pRU3004
This work
69
pRU277 pRK415-1 carrying 3.8kb EcoRI fragment from pRU3004
This work
70
Strain, phage or plasmid
Description Source or Reference
pRU278 pRK415-1 carrying 3.8kb EcoRI
fragment from pRU3004 cloned in opposite orientation to pRU278
This work
pRU307 pRK415-1 carrying 2.0kb XbaI-HindIII fragment from pRU234
This work
pRU308 pRK415-1 carrying 3.1kb XbaI-HindIII fragment from pRU239
This work
pRU309 pRK415-1 carrying 4.4kb XbaI-HindIII fragment from pRU246
This work
pRU310 pIJ1891 carrying 5.4kb XbaI-HindIII fragment from pRU189
This work
pRU311 pIJ1891 carrying 2.0kb XbaI-HindIII fragment from pRU234
This work
pRU312 pIJ1891 carrying 3.1kb XbaI-HindIII fragment from pRU239
This work
pRU313 pIJ1891 carrying 4.4kb XbaI-HindIII fragment from pRU246
This work
pRU383 pBluescript SK+ carrying 8.8kb BamHI Tn5-lacZ-bearing fragment from pRU3031
This work
pRU384 pBluescript SK+ carrying 8.4kb BamHI Tn5-lacZ-bearing fragment from pRU3033
This work
pRU386 pIJ1891 carrying 5.0kb lacZ-bearing PstI fragment from pRU3068
This work
pRU387 pIJ1891 carrying 5.0kb lacZ-bearing PstI fragment from pRU3068 cloned in opposite orientation to pRU386
This work
pRU388 pRK415-1 carrying 1.8kb KpnI-BamHI fragment from pRU256
This work
pRU389 pIJ1891 carrying 1.8kb KpnI-BamHI fragment from pRU256
This work
pRU393 pMP220 carrying 1.0kb EcoRI-PstI fragment from pRU189
This work
pRU394 pSP72 carrying 11.8kb Tn5-lacZ-bearing salI fragment from pRU3061
This work
pRU395 pSP72 carrying 8.9kb Tn5-lacZ-bearing salI fragment from pRU3069
This work
71
Strain, phage or plasmid
Description Source or Reference
pRU396 pSP72 carrying 10.2kb Tn5-lacZ-
bearing salI fragment from pRU3070
This work
pRU397 pRK415-1 carrying 6.3kb PstI fragment from pRU3004 cloned in opposite orientation to pRU182
This work
pRU398 pRK415-1 carrying 4.4kb EcoRI fragment from pRU3004 cloned in opposite orientation to pRU276
This work
pRU3004 pLAFR1 cosmid containing mdh-sucCDAB from strain 3841
This work
pRU3024 pLAFR1 cosmid containing aapJQMP from strain 3841
This work
pRU3026 pRU3024 aapQ::Tn5-lacZ This work pRU3027 pRU3024 aapP::Tn5-lacZ This work pRU3028 pRU3024 aapJ::Tn5-lacZ This work pRU3029 pRU3024 aapQ::Tn5-lacZ This work pRU3031 pRU3024 cysE::Tn5-lacZ This work pRU3032 pRU3024 aapM::Tn5-lacZ This work pRU3033 pRU3024 cysE::Tn5-lacZ This work pRU3035 pRU3024 aapM::Tn5-lacZ This work pRU3039 pRU3024 aapQ::Tn5-lacZ This work pRU3044 pRU3024::Tn5-lacZ This work pRU3046 pRU3024 aapQ::Tn5-lacZ This work pRU3048 pRU3024 aapJ::Tn5-lacZ This work pRU3049 pRU3024::Tn5-lacZ This work pRU3053 pRU3024 metC::Tn5-lacZ This work pRU3059 pRU3004 sucC::Tn5-lacZ This work pRU3061 pRU3004 sucA::Tn5-lacZ This work pRU3067 pRU3004 sucA::Tn5-lacZ This work pRU3068 pRU3004 sucA::Tn5-lacZ This work pRU3069 pRU3004 sucB::Tn5-lacZ This work pRU3070 pRU3004 mdh::Tn5-lacZ This work pRU3075 pRU3004 sucA::Tn5-lacZ This work pRU3076 pRU3004 mdh::Tn5-lacZ This work pRU3080 pRU3024::Tn5-lacZ This work pRU3082 pRU3024 metC::Tn5-lacZ This work pRU3083 pRU3024::Tn5-lacZ This work pRU3084 pRU3024 metC::Tn5-lacZ This work pRU3085 pRU3024::Tn5-lacZ This work pRU3086 pRU3024 metC::Tn5-lacZ This work
72
pRU3089 pRU3024::Tn5-lacZ This work
73
Strain, phage or plasmid
Description Source or Reference
pRU3090 pRU3024::Tn5-lacZ This work pRU3091 pRU3024::Tn5-lacZ This work Bacteriophage λ::Tn5-lacZ λ carrying the Tn5-B20
transposon Simon et al. (1989)
RL38 Generalized transducing phage of R. leguminosarum
Buchanan-Wollaston (1979)
2.1.2 Culture conditions
R. leguminosarum strains were grown at 28°C on either TY (Beringer, 1974) or
acid minimal salts (AMS) medium derived from that of Brown and Dilworth
(1975), the changes being; potassium phosphate (0.5mM), MgSO4 (2mM) and
buffering provided by MOPS (20mM) pH 7.0. All carbon and nitrogen sources
were at 10 mM unless otherwise stated. Y medium used in transductions was
prepared as previously described by Sherwood (1970). Antibiotics were used at
the following concentrations in µg ml-1 unless otherwise stated; gentamicin 20,
kanamycin 40, spectinomycin 100 streptomycin 500, and tetracycline 2 (in
AMS), 5 (in TY).
E. coli strains were grown at 37°C on LB, with antibiotic concentrations in µg
ml-1 as follows; ampicillin 50, chloramphenicol 10 gentamicin 5, kanamycin 25,
and tetracycline 10.
2.1.3 DNA and genetic manipulations
All routine DNA analysis was performed essentially according to Sambrook
et al. (1989). Southern transfer of DNA to positively charged nylon membrane
(Boehringer Mannheim) and hybridisation were done using an Amersham ECL
kit according to the manufacturers instructions. Conjugations were performed
using either Escherichia coli strain S17-1 as the donor strain according to Simon et
74
al. (1983), or as triparental matings according to Figurski et al. (1979) with either
E. coli strain 803 or DH5α as the donor, and strain 803 containing pRK2013
providing the transfer functions. Transductions were performed according to
Buchanan-Wollaston (1979) using the phage RL38. Transductants were selected
for on TY agar containing kanamycin (80 µg ml-1). DNA sequencing was
performed by the cycle sequencing method using a Promega fmol kit according
to the manufacturers instructions. Nucleotide sequences of non-routine
sequencing primers used are as follows:
P0: GTTCAGGACGCTACTTG
P15: GGATCCATAATTTTTTCCTCC
P16: AAGATAAGACAACGGAAAAGG
P21: ATGGGTCAGGCGGGTGTTG
P22: GTCGCAAATGTCACTATGG
Computer-assisted sequence analysis was performed using GCG software.
2.1.4 Mutagenesis
Transposon mutagenesis was carried out on R. leguminosarum bv. viciae 3841
with Tn5 using the suicide vector pSUP202-1 as described (Simon et al., 1983).
Mutations in pRU3024 and pRU3004 were produced by first transforming the
cosmid into E. coli strain MC1061. Transformants were mutagenised with Tn5-
lacZ by using λ containing the transposon derivative B20 essentially as
described (Simon et al., 1989). Kanamycin resistant colonies were pooled, and
the cosmids isolated by the alkaline lysis technique. The cosmids were
transformed into E. coli strain DH5α and kanamycin resistant colonies purified.
Cosmid DNA was isolated from each purified strain and the location and
orientation of transposons determined by restriction mapping.
To create chromosomal mutations, mutated cosmids were conjugated into R.
leguminosarum strain 3841. After purification, the incompatible plasmid pPHJI1
was conjugated into each strain and the homogenotes isolated by the technique
of Ruvkun and Ausubel (1981).
75
2.1.5 Transport Assays
For R. leguminosarum strains, cells were prepared and transport assays
performed as previously described (Poole et al., 1985), using in each case, a total
substrate concentration of 25µM. In exchange experiments, cell suspensions
were incubated in 50µM labelled AIB at 28°C for between 2 and 20 min prior to
the addition of unlabelled AIB or glutamate to a concentration of 4mM, CCCP
to a concentration of 50µM, or KCN to a concentration of 10mM. Subsequent
0.1ml samples for Millipore filtration and counting, were taken at intervals of 1,
2, 5 or 10 min for up to 50 min.
For E. coli strains, cells grown on LB to A660 0.5-0.7 were harvested, washed
twice in 50mM potassium phosphate, pH6.9, containing 0.5mM MgSO4 (buffer
A) and resuspended to a final A660 of approximately 10 in the same buffer. This
cell suspension was stored at 37°C with shaking at 250 rpm. Uptake was
assayed at 37°C, after 10-fold dilution of the cells into buffer A containing
10mM glucose, and incubation for 1 min. Uptake was initiated by addition of
labelled substrate to a final concentration of 25µM. Samples were removed at 1
min intervals for 2 min, and cells collected by Millipore filtration under
vacuum. Filters were washed rapidly with 2 x 3ml ice-cold buffer A, and the
radioactivity counted in Beckman "Ready Safe" scintillation fluid.
The specific activities of labelled substrates in the assays were: L-[U-
14C]aspartic acid (354 MBq mmol-1), L-[U-14C]glutamic acid (357 MBq mmol-1),
L-[U-14C]alanine (348 MBq mmol-1), L-[U-14C]histidine (359 MBq mmol-1), L-[U-
14C]leucine (359 MBq mmol-1), L-[35S]methionine (370 MBq mmol-1) and D-[U-
14C]glucose (358 MBq mmol-1).
Incorporation experiments were performed as previously described (Poole et
al., 1985).
2.1.6 Isolation of periplasmic fractions and protein gel electrophoresis
R. leguminosarum strains were grown in AMS containing 50µM potassium
phosphate and periplasmic proteins released by lysosyme/EDTA treatment as
76
described by Glenn and Dilworth (1979). Crude periplasmic fractions were
concentrated on microconcentrators (Microcon-10, Amicon) as necessary.
Samples were subjected to SDS-PAGE as previously described (Laemmli, 1970),
using a 12.5% gel.
Cell fractions for use in assays of marker enzymes were prepared as follows:
After the removal of periplasmic proteins, cells were resuspended in 30mM
TRIS/HCl buffer pH8.0 containing 20%(w/v) sucrose and 1mM DTT, and
disrupted by two passages through a French press at 69000 kPa. Following
centrifugation at 30000 g for 20 min, the supernatant was used for enzyme
assay.
2.1.7 Protein binding assays
R. leguminosarum strains were grown and periplasmic proteins isolated as for
SDS-PAGE. Each crude extract was dialysed overnight in 3 x 3l of 5mM
HEPES, pH7.2 before being concentrated to approximately 1 mg ml-1 on an
Ultrafree-20 (Millipore) filter (10000 MWCO). Substrate-binding by the
concentrated extracts was assayed by three techniques.
Ammonium sulphate precipitation was performed essentially as described
by Richarme and Kepes (1983). Periplasmic extract (100 µg protein) was
incubated in 10 µM labelled substrate at 28°C for 10 min. Proteins were
precipitated by the rapid addition of 10 volumes of ice-cold saturated
ammonium sulphate, and collected by Millipore filtration under vacuum.
Filters were washed with 2 x 3ml saturated ammonium sulphate, and counted
in Beckman "Ready Safe" scintillation fluid.
Detection of binding activity by direct polyacrylamide gel electrophoresis of
the ligand-protein complex in non-denaturing conditions was carried out as
described by Le Rudulier et al. (1991). Periplasmic protein extract (20 µg) was
incubated in 10 µM labelled substrate in a total volume of 25 µl, for 30 min at 28
°C. Laemmli's sample buffer (Laemmli, 1970) without SDS and β-
mercaptoethanol was added, and the samples subjected to PAGE using a 12.5%
77
gel and omitting SDS. The analyses were performed with a constant voltage of
200V for approximately 55 min. The gels were quickly dried on Whatman 3MM
paper and exposed to X-ray film for 10 days.
In the third technique, 420µl of protein extract was incubated in 7µM labelled
amino acid in a total assay volume of 450µl, for 30 min at 28°C. Following
incubation, 50µl of the assay was removed and scintillation counted. The
remaining 400µl was spun through a Microcon-10 (Amicon) microconcentrator
at 5°C and the radioactivity in 50µl of the filtrate counted. The amount of
substrate bound to protein was calculated by comparison of the amount of
radioactivity present in the filtrate to that in the incubation mixture prior to
spinning. Controls containing no protein were performed to determine any
background binding of substrate by the filter in the microconcentrator.
Specific activities of substrates in the assays were: L-[U-14C]aspartic acid (676
MBq mmol-1), L-[U-14C]glutamic acid (688 MBq mmol-1), L-[U-14C]alanine (655
MBq mmol-1), L-[U-14C]histidine (695 MBq mmol-1), L-[U-14C]leucine (695 MBq
mmol-1), and L-[35S]methionine (3.01 GBq mmol-1)).
2.1.8 Enzyme assays
Cultures of R. leguminosarum strains were harvested at a cell density of
approximately 5 x 108 cells ml-1, washed and resuspended in 40 mM HEPES,
pH7.0, containing 1 mM DTT. Cells were disrupted by two passages through a
French press at 69000 kPa. Following centrifugation at 30000 g for 20 min, the
supernatant was used for enzyme assay. Citrate synthase, α-ketoglutarate
dehydrogenase, isocitrate dehydrogenase and succinyl-CoA synthetase were
assayed according to Reeves et al. (1971). Malate dehydrogenase was assayed
by the technique of Saroso et al. (1986).
β-Galactosidase fusions were assayed according to Miller (1972) with the
modifications described by Poole et al. (1994a). Alkaline phosphatase activity
was measured as described by de Maagd and Lugtenberg (1986).
78
2.1.9 Metabolite excretion assays
Cells of R. leguminosarum strains grown to A600 0.5-0.7 on AMS containing the
pertinent carbon and nitrogen sources, were harvested aseptically, washed once
in AMS and resuspended to an A600 of approximately 1 in AMS containing the
carbon and nitrogen sources used for initial growth. Resupended cells were
incubated at 28°C and samples removed at 0, 60, 150 and 270 min. Samples
were centrifuged at 11000 g for 15 min and the supernatants assayed for one or
more metabolites.
Glutamate was determined by a technique derived from Bernt and
Bergmeyer (1974). Each assay contained 75 µmol glycine, 60 µmol hydrazine
monohydrate, 4 µmol NAD+, 3 units of glutamate dehydogenase and 250 µl of
sample, in a total volume of 1.5 ml. Assays were incubated for 90 min at 37°C
prior to reading of the absorbance at 340 nm and determining the glutamate
concentrations from a standard curve prepared between 20 and 100 nmol.
Minus enzyme controls were run as blanks.
Aspartate was measured by a procedure adapted from Bergmeyer et al.
(1974). Each assay contained 150 µmol potassium phosphate (pH 7.2), 0.3 µmol
NADH, 15 µmol 2-oxoglutarate, 2 units of aspartate aminotransferase, 2 units of
malate dehydrogenase and 250 µl of sample, in a total volume of 1.5 ml. Assays
were incubated and read as for the glutamate determination.
Alanine was determined in a final volume of 1.5 ml containing 75 µmol
glycine, 60 µmol hydrazine monohydrate, 4 µmol NAD+, 0.5 unit of alanine
dehydrogenase and 250 µl of sample. Assays were incubated and read as for
the glutamate determination.
α-Ketoglutarate assays contained 150 µmol potassium phosphate (pH 7.2),
0.3 µmol NADH, 75 µmol aspartate, 2 units of aspartate aminotransferase, 2
units of malate dehydrogenase and 250 µl of sample, in a total volume of 1.5 ml.
Assays were incubated and read as for the glutamate determination.
79
2.1.10 Intracellular concentrations
500 ml cultures of R. leguminosarum strains were grown on glucose/NH4Cl,
harvested by centrifugation at a cell density of approximately 5 x 108 cells ml-1,
washed three times in AMS, and resuspended in 15 ml 100 mM HEPES, pH7.2,
all operations being carried out at 5°C. Cells were disrupted by two passages
through a French press at 69000 kPa, centrifuged at 30000 g, 5°C, for 20 min,
and the supernatant assayed for protein content. 2.5 ml of ice-cold 10% TCA
was added to 5 ml of supernatant. After centrifugation at 3500 g, 5°C, for 15
min, the pH of the supernatant was adjusted to 7.0 and the volume made up to
8 ml. Samples of the neutralized supernatant were assayed for glutamate and
α-ketoglutarate as described in Section 2.1.9.
2.1.11 Protein determination
The protein concentration of whole cells was determined by the method of
Lowry et al. (1951), using bovine serum albumin as standard. Protein
concentrations of extracts used in enzyme assays and PAGE were determined
by the method of Bradford (1976).
2.1.12 Plant Assays
Seeds of Pisum sativum c.v. meteor were surface sterilised, germinated,
inoculated with R. leguminosarum strains, and grown as described by Poole et al.
(1994b). Four weeks after inoculation, plants were harvested and acetylene
reduction carried out on whole plants as described by Trinick et al. (1976).
Sample nodules were removed and surface sterilised by immersion in calcium
hypochlorite (0.7%) for 10 min. Nodules were then washed three times in
sterile distilled water, crushed and bacteria streaked on TY agar. Isolated
bacteria were subsequently replica plated and screened for appropriate
antibiotic markers.
80
CHAPTER 3 THE CLONING AND CHARACTERIZATION OF THE GENERAL AMINO ACID PERMEASE OF RHIZOBIUM LEGUMINOSARUM STRAIN 3841
81
3.1 INTRODUCTION
Observations that Rhizobium bacteroids under nitrogen-fixing conditions
excrete alanine and/or aspartate (Kretovich et al., 1986; Appels & Haaker, 1991;
Kouchi et al., 1991; Rosendahl et al., 1992), are compatible with the proposal that
oxidation of the dicarboxylate malate by the bacteroid is coupled to
transamination and excretion of aspartate, forming a malate-aspartate shuttle in
the nodule (Kahn et al., 1985). Alternatively, amino acids may be excreted in
order to alleviate inhibition of the TCA cycle by preventing accumulation of
ketoacids and reducing equivalents in the bacteroid. Whichever pathway
operates, amino acid transport across the bacteroid/peribacteroid membrane is
likely to be crucial to nitrogen fixation, and may regulate bacteroid metabolism
(Murphy et al., 1979).
The unusual properties of the high affinity glutamate uptake system of R.
leguminosarum strain 3841 - a broad specificity for structurally unrelated amino
acid side chains, and an apparent ability to mediate exchange between
intracellular and extracellular amino acid pools - are compatible with a role in
nutrient exchange in the nodule, and make this transporter an interesting
subject for study in its own right.
In this chapter the cloning, sequencing and mutation of the genes encoding
the general amino acid permease of R. leguminosarum strain 3841 are described.
The results of experiments to investigate the involvement of this transporter in
amino acid exchange are discussed with regard to both the structure of the
transporter, and models of nutrient exchange in the nodule.
82
3.2 RESULTS
3.2.1 Isolation of cosmid pRU3024 carrying the general amino acid permease genes of Rhizobium leguminosarum strain 3841
The strategy employed in cloning the general amino acid permease genes of
Rhizobium leguminosarum strain 3841 is based on the potential for a strain
carrying additional copies of these genes to show increased labelling when
grown on [14C]glutamate.
Strain 3841 containing a strain 3841 chromosomal library (as EcoRI fragments
in pLAFR1 (Downie et al., 1983)) was grown on acid minimal salts agar
containing glucose (carbon source), NH4Cl (nitrogen source), [14C]glutamate,
and aspartate. Labelling by growth on [14C]glutamate is dependent on
glutamate incorporation and will only be indicative of glutamate uptake [by the
general amino acid permease] if glutamate uptake is the rate limiting-step in
incorporation. Hence, aspartate, which has been shown to cause a severe
reduction in uptake by the general amino acid permease (Reid, 1995), was
included in the growth media. After 2-3 days growth, colonies were lifted onto
nitro-cellulose filters, which were dried and exposed to X-ray film. On visual
inspection of the autoradiograph of approximately 3000 colonies, one colony
that apparently exhibited increased labelling was discerned with difficulty.
Following its purification from the selection media, some of the uptake
properties of this strain, RU438, were investigated (Table 3.1). It can be seen
that the uptake of all the amino acids tested is elevated in strain RU438 in
comparison to the wild-type. The increase in the transport rate of the non-
metabolizable amino acid α-aminoisobutyrate (AIB) is particularly significant
as it indicates that the increase in amino acid uptake is not the result of
increased metabolic drag. (AIB is not incorporated into TCA-precipitated
material from either strain 3841 or strain RU438 following uptake of AIB (Table
3.2)). The fact that glucose uptake is not increased in RU438 suggests that this
strain is affected in amino acid transport specifically.
83
Table 3.1 Rates of amino acid transport in R. leguminosarum strains grown on glucose/NH4Cl. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. ND, not determined.
Substrate Strain 3841 RU438 RU439 RU640 RU641 RU913 L-Glutamate 5.6±0.4 19.3±1.1 38.1±4.0 43.9±4.3 38.1±2.6 4.9±0.2 L-Aspartate 3.9±0.1 13.8±0.1 ND 43.9±3.3 ND 3.0±0.8 L-Alanine 6.4±0.7 18.7±1.2 ND 63.7±2.7 ND 6.6±0.8 AIB 4.3±0.6 21.2±2.7 40.4±6.5 70.3±5.3 ND 5.0±0.3 L-Histidine 5.4±0.5 15.7±2.0 ND 40.0±1.2 ND 5.5±0.7 L-Leucine 5.3±0.2 10.8±1.3 ND 54.5±4.8 ND 5.6±0.2 L-Methionine 9.0±0.1 ND ND 37.7±2.8 ND ND D-Glucose 39.5±3.2 29.1±2.3 38.7±2.2 46.2±6.9 ND 44.5±2.1
Table 3.2 Labelling of TCA precipitated material by [14C]AIB and [14C]Alanine in R. leguminosarum strains 3841 and RU438. Cells grown on glucose/NH4Cl/aspartate were washed and resuspended in minimal salts in the presence of either [14C]AIB or [14C]alanine. After the time intervals shown, two identical samples were taken. One of these was added to ice-cold 10%(w/v) TCA and the amount of label in the precipitate determined. The label in whole cells was obtained from the second sample. Values are the result from a single experiment and are expressed as percentage of added label. Substrate Assay Time (min) and strain 1 5 10 AIB 3841 Whole cells 2.0 7.9 14.2 RU438 Whole cells 7.0 28.1 41.8 3841 TCA precipitate <0.1 <0.1 <0.1 RU438 TCA precipitate <0.1 <0.1 <0.1 L-Alanine 3841 Whole cells 2.0 10.0 16.6 RU438 Whole cells 6.3 21.0 27.4 3841 TCA precipitate 0.3 3.5 10.3 RU438 TCA precipitate 0.4 2.5 6.6
84
In order to confirm that the observed transport phenotype of RU438 was
caused by the cosmid, pRU3024, carried by the strain, rather than a
chromosomal mutation in the host, pRU3024 DNA was isolated from strain
RU438 and used to transform E. coli strain DH5α. Cosmid pRU3024 was then
conjugated from the resulting E. coli strain back into strain 3841. The transport
phenotype of the resulting R. leguminosarum strain with respect to the
previously tested amino acids and glucose was found to be the same as that of
strain RU438 (data not shown).
3.2.2 Restriction mapping, sub-cloning and mutational analysis of pRU3024
Restriction analysis of pRU3024 was carried out using BamHI, EcoRI, and
HindIII. The resulting map of strain 3841 DNA is shown in Fig. 3.1. This map
does not show all of the 17.1kb insert DNA. Two small EcoRI fragments (2.1kb
and 0.7kb) which lie outside the region shown were not mapped.
The 6.6kb and 3.2kb BamHI fragments, the 6.3kb, 4.8kb and 3.2kb EcoRI
fragments, and the 10.2kb and 2.9kb HindIII fragments of pRU3024 were cloned
in the broad host range, medium copy number vector pRK415-1. These sub-
clones were then conjugated into strain 3841 and the rate of glutamate uptake
by the resulting strains investigated. For the purposes of comparison glutamate
uptake is taken to be representative of general amino acid uptake. All strains
showed glutamate transport similar to the wild-type (data not shown) except
strain RU439, which contains the clone of the 10.2kb HindIII fragment
(pRU135). Strain RU439 exhibits similar rates of glutamate, AIB and glucose
uptake to those of strain RU438 (Table 3.1).
85
Fig. 3.1 Restriction map of pRU3024 with sub-clones below. The shaded arrows indicate the direction of transcription initiation from the lac promoter in the vector of the sub-clones. Restriction sites are: B, BamHI; C, ClaI; D, DraI; E, EcoRI; H, HindIII; M, MluI; N, NcoI; Nt, NotI; S, SstI; Sp, SspI.
86
Plasmid pRU135 was subjected to further restriction analysis (Fig. 3.1) and a
variety of sub-clones in pRK415-1 created. The effect of these sub-clones on
glutamate uptake in strain 3841 was investigated. Two sub-clones of pRU135
capable of producing the same effect on amino acid transport in strain 3841 as
pRU135 itself (data not shown) were found to be pRU165, a clone of the 8.7kb
HindIII-MluI fragment, and pRU185, a clone of the 6.9kb HindIII-ClaI fragment
(Fig. 3.1). By contrast, pRU149 and pRU186, clones of the 6.0kb HindIII-SstI and
5.2kb NcoI fragments of pRU135, respectively (Fig. 3.1), have no effect on amino
acid transport. From these results it was concluded that the DNA responsible
for increasing amino acid uptake in strain 3841 is contained within the 5.4kb
region of pRU135 that is bounded by the unique MluI and ClaI sites (Fig. 3.1).
Concomitant to the above restriction analysis and sub-cloning, pRU3024 was
subjected to saturation Tn5-lacZ mutagenesis. Mutated cosmids were screened
for their effect on amino acid transport in strain 3841 (Table 3.3). Nine cosmids
(pRU3026, pRU3027, pRU3028, pRU3029, pRU3032, pRU3035, pRU3039,
pRU3046 and pRU3048) were isolated that produce no effect on amino acid
uptake. Restriction analysis of these cosmids revealed that in all cases the
transposon had inserted in the 5.4kb MluI-ClaI region found in pRU135 (Fig.
3.2). It was therefore decided to sequence this 5.4kb of DNA.
87
Table 3.3 Rates of glutamate uptake by R. leguminosarum strains grown on glucose/NH4Cl/aspartate. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the result from a single experiment except those for strains 3841 and RU438 which are the mean±SEM of determinations from three or more independent cultures. Strain Uptake 3841 3.2±0.1 RU438 (3841/pRU3024) 19.9±1.2 RU441 (3841/pRU3026) 3.6 RU442 (3841/pRU3027) 5.7 RU443 (3841/pRU3028) 3.1 RU500 (3841/pRU3029) 4.5 RU502 (3841/pRU3031) 8.0 RU503 (3841/pRU3032) 5.3 RU504 (3841/pRU3033) 8.4 RU506 (3841/pRU3035) 4.9 RU510 (3841/pRU3039) 3.7 RU515 (3841/pRU3044) 20.8 RU517 (3841/pRU3046) 3.0 RU519 (3841/pRU3048) 3.0 RU522 (3841/pRU3049) 9.4 RU526 (3841/pRU3053) 15.7 RU972 (3841/pRU3080) 18.8 RU974 (3841/pRU3082) 21.7 RU975 (3841/pRU3083) 19.7 RU976 (3841/pRU3084) 20.5 RU977 (3841/pRU3085) 19.0 RU978 (3841/pRU3086) 19.7 RU1069 (3841/pRU3089) 19.8 RU1070 (3841/pRU3090) 22.7 RU1071 (3841/pRU3091) 21.1
88
Fig. 3.2 Restriction map showing location of transposons in mutants of cosmid pRU3024. The locations of the Tn5-lacZ insertions are flagged with the number of the cosmid in which they occur. Each flag points in the direction of transcription of the lacZ gene in the transposon. Filled flags represent active fusions. Restriction sites are: B, BamHI; C, ClaI; E, EcoRI; H, HindIII; M, MluI.
89
Three mutated pRU3024 cosmids were isolated that do not cause the increase
in glutamate uptake in strain 3841 that is associated with pRU3024 itself, but do
increase uptake to a lesser degree (Table 3.3). In one of these cosmids,
pRU3049, the insert lies close to one end of the previously described 5.4kb MluI-
ClaI region (Fig. 3.2), and the phenotype produced by this cosmid is likely to be
due to the proximity of the transposon to the promoter region for the genes
lying between these restriction sites (Section 3.2.6). Intriguingly however, the
transposons in the other two cosmids in this class, pRU3031 and pRU3033, are
both located in a 0.8kb BamHI fragment approximately 3.5kb away from the
5.4kb MluI-ClaI region (Fig. 3.2). Since cosmids containing mutations in the
intervening 3.5kb region, for example pRU3083, pRU3085, and pRU3086 (Fig.
3.2), produce the same increase in glutamate uptake in 3841 as pRU3024 (Table
3.3), it appears that pRU3024 contains two distinct regions of DNA that affect
amino acid uptake in strain 3841.
3.2.3 Nucleotide sequence of the 5.4kb MluI-ClaI fragment of pRU135
A variation of the ordered deletion strategy described by Robson et al. (1986)
was employed in determining the nucleotide sequences of both strands of the
5.4kb MluI-ClaI fragment from pRU135:
The 5.4kb fragment was cloned, in both orientations, into the EcoRV site of
pBluescript SK+, creating plasmids pRU189 and pRU190. Each plasmid was cut
at its unique SmaI site within the polylinker, partially digested with BstUI and
ligated (Fig. 3.3). E. coli strain DH5α was transformed with the resulting DNA
and AmpR transformants were screened for a range of plasmids between 3.0kb
and 8.4kb that retained the XbaI site of the pBluescript SK+ polylinker.
90
Fig. 3.3 Schematic representation of the strategy employed to obtain ordered deletions of pRU189 and pRU190 for sequencing. The unfilled arrow indicates the direction of sequencing from primers Reverse and SK. Restriction sites are: X, XbaI; S, SmaI.
91
Sequencing of the insert DNA in these plasmids (which are illustrated in Fig.
3.4) using Reverse and SK primers to the pBluescript SK+ polylinker, provided
the nucleotide sequence shown in Fig. 3.5. (In the case of pRU247 an additional
primer (P16), to the end of the sequence provided by the SK primer, was
needed to obtain sequence overlapping that from the next deletion, pRU246).
In the course of this sequencing it was found that sequence obtained from
pRU190 lacked the G at position 438 of the sequence in Fig. 3.5. That this was
due to the deletion of a base during the construction of pRU190 rather than the
presence of an erroneous additional base in pRU189, was demonstrated by
sequencing cosmid pRU3024 itself in the vicinity of the ambiguity, using two
primers (P21 and P22), one to each strand approximately 100bp upstream of the
problematic base.
In order to be certain that the sequenced DNA was capable of enhancing
amino acid uptake in strain 3841, the insert DNA from both pRU189 and
pRU190 was excised as a 5.4kb HindIII-XbaI fragment and in each case cloned
into pRK415-1. The two resulting plasmids, pRU191 and pRU192, respectively,
were then conjugated into strain 3841 and glutamate uptake in the resulting
strains (RU640 and RU641, respectively) measured (Table 3.1). Both strains
show a greater than six-fold increase in glutamate transport in comparison to
the wild-type.
92
Fig. 3.4 Schematic representation of ordered deletions of pRU189 and pRU190. Figures in parentheses indicate the position of the relevant BstUI site for each deletion with reference to the sequence in Fig. 3.5. Arrows indicate the direction of sequencing from primers Reverse and SK.
93
94
95
96
97
98
99
100
101
3.2.4 Coding regions of the nucleotide sequence from pRU189
The deduced amino acid sequences from four open reading frames (ORFs)
found in the nucleotide sequence from pRU189 are shown in Fig. 3.5. All of
these ORFs are transcribed in the same direction. Proposed protein starts are
based on the presence of methionine codons which are preceded by sequence
resembling a ribosome binding site, and on comparisons of the proteins to
similar proteins in other bacteria.
Sequence comparisons suggest that the polypeptides encoded by the four
ORFs constitute the components of an ABC transporter, with particular
homology being found to carriers of amino acids and opines. Consequently, we
propose the gene designation aap (for amino acid permease). The letter
assigned to an individual gene (Fig. 3.5) is based on that of the corresponding
gene in the histidine transport operon of Salmonella typhimurium.
The deduced amino acid sequence of the aapP gene product (molecular
weight 29kD) exhibits extensive homology to the ATP-binding proteins of other
bacterial amino acid transporters. The strongest identities are to GlnQ from
Bacillus stearothermophilus (61.4%), GlnQ, HisP and ArtP from E. coli (52.1%,
50.2% and 43.9%, respectively), GluA from Cornyebacterium glutamicum (61.8%),
HisP from S. typhimurium (50.0%), and OccP and NocP from Agrobacterium
tumefaciens (46.4% and 49.0%, respectively). AapP also contains the ATP/GTP-
binding site motif A and the ABC transporters family signature (Fig. 3.5) that
are characteristic of the ATP-binding proteins of ABC transporters (Higgins et
al., 1986; Higgins, 1992).
The C-terminal ends of AapQ and AapM show significant homology to
integral membrane proteins from other ABC transporters. In particular, both
proteins contain the recently determined consensus sequence (Saurin et al.,
1994) characteristic of this type of protein (Fig. 3.5). AapQ exhibits 30.6%
identity over 121 amino acids to OccM from A. tumefaciens and 33.0% identity
over 115 amino acids to HisM from E. coli, while AapM is 28.4% identical over
221 residues to HisM from S. typhimurium and 27.7% identical over 223 amino
102
acids to GlnP from E. coli. However, both AapM and AapQ are larger by 110 to
180 amino acids than other previously described amino acid transport proteins
of this type, with their predicted molecular weights being 42kD and 43kD,
respectively.
AapJ exhibits similarities to binding proteins. The homology to the
corresponding component of the glutamine transport system of B.
stearothermophilus (29.5% identity over 245 residues) is greatest (although AapJ
possesses an additional 96 amino acids at the C-terminal end). At the N-
terminal end of the protein a putative signal sequence (Fig. 3.5) is identified by
the GCG program SIGCLEAVE, indicating translocation of AapJ through the
cytoplasmic membrane.
Interestingly, the most significant homology to each of the Aap proteins is
provided by the deduced polypeptide from one of four adjacent unidentified
ORFs in the 67.4 to 76.0 minute region of the E. coli K12 chromosome (EMBL
accession number U18997). Each aap gene product shows identity to a different
E. coli ORF as follows: AapJ, 58.4%; AapQ, 48.9%; AapM, 50.7%; AapP, 71.0%.
Indeed, the genes occur in the same order in E. coli as they do in R.
leguminosarum. The homology of the E. coli ORFs to AapJ, AapQ and AapM is
particularly striking since these proteins are significantly bigger, and only
approximately 30% identical to the corresponding components of previously
reported ABC transporters of amino acids.
AapJ is also 57.6% identical to the translation of an incomplete unidentified
ORF from Pseudomomas fluorescens (EMBL accession number D00852; (Hong et
al., 1991) ). An alignment of AapJ and the AapJ-like proteins from E. coli and P.
fluorescens is shown in Fig. 3.6.
103
Rl MKNKLLSAAI--GAAVLAVGASAASATTLSDVKAKGFVQCGVNTGLTGFAAPDASGNWAGFD
Ec EKDDDSHTGC--RQRAACRCKSGAAGATLDAVQKKGFVQCGISDGLPGFSYADADGKFSGID
Pf MKVLKSTLAIFTAAAVLGVSGFAQAGATLDAVQKKGFVQCGVSDGLPGFSVPDASGKILGID
Pf2 MKVLKSTLAIVCAAAVLGVSGFAQAGATLDAVQKKGFVQCGVSDGLPGFSV-----------
# ~ ~ ~~~## #~ #######~~ ## ##~ ~## #~ #~#
Rl VDFCKAVASAVFGDPTKVKYTPTNAKERFTALQSGEIDVLSRNTTWTINRDTALGFNFRPVT
Ec VDICRGVAAAVFGDDTKVKYTPLTAKERFTALQSGEVDLLSRNTTWTSSRDAGMGMAFTGVT
Pf ADYCRAVAAAVFGDATKVKFSQLNAKERFTALQSGEVDILSRNTTMTSSRDAGMGLKF----
Pf2 --------------------------------------------------------------
# #~~##~##### ####~~ ############~#~###### # ~##~~~# # ##
Rl YYDGQGFMVRKGLNVKSALELSGAAICVQSGTTTELNLADYFKTNNLQYNPVVFENLPEVNA
Ec YYDGIGFLTHDKAGLKSAKELDGATVCIQAGTDTELNVADYFKANNMKYTPVTFDRSDESAK
Pf --------------------------------------------------------------
Pf2 --------------------------------------------------------------
#### ##~ ~ ~### ## ##~~#~#~## ####~#####~##~~# ## #~ #
Rl AYDAGRCDVYTTDQSGLYSLRLTLKNPDEHIILPEIISKEPLGPAVRQGDDQWFDIVSWTAY
Ec ALESGRCDTLASDQSQLYALRIKLSNPAEWIVLPEVISKEPLGPVVRRGDDEWFSIVRWTLF
Pf --------------------------------------------------------------
Pf2 --------------------------------------------------------------
# ~~#### ~~### ##~##~ # ## # #~###~######## ##~###~## ## ## ~
Rl ALINAEEFGITQANVDEMKNSPN-PDIKRFLGSETDTKIGTDLGLTNDWAANVIKGVGNYGE
Ec AMLNAEEMGINSQNVDEKAANPATPDMAHLLGKEGDY--GKDLKLDNKWAYNIIKQVGNYSE
Pf --------------------------------------------------------------
Pf2 --------------------------------------------------------------
#~~#### ## #### ~# ##~ ~~## # # # ## # # ## #~## ####~#
Rl IFERNIGQGSPLKIARGLNALWNKGGIQYAPPVR
Ec IFERNVGSESPLKIKRGQNNLWNNGGIQYAPPVR
Pf ----------------------------------
Pf2 ----------------------------------
#####~# ##### ## # ###~##########
Fig. 3.6 Alignment of AapJ from R. leguminosarum (Rl) with similar proteins from E. coli (Ec, EMBL accession number U18997) and P. fluorescens (Pf, EMBL accession number D00852; Pf2, (Hong et al., 1991)). Alignment was performed using the program ALIEN. #, identical amino acids; ~, conservative substitutions
104
The EMBL entry U18997 indicates a possible frame shift in the region of the
sequence corresponding to aapJ, and an arbitrary deletion of the two nucleotides
at positions 199844-5 in this sequence yields an E. coli protein which now also
exhibits significant homology to the other two AapJ proteins over the first
twenty residues, including a methionine start at the same position as that
suggested for the R. leguminosarum protein.
Hydropathy analysis of the deduced amino acid sequences of AapQ and
AapM by the method of Engelman et al. (1986), incorporating the "positive-
inside rule" of von Heijne (1986; 1992), predicts eight membrane-spanning
regions for each protein (Figs. 3.7 and 3.8). Kyte and Doolittle (1982) plots
predict nine membrane-spanning regions for each protein, with the additional
transmembrane segment occurring between the sixth and seventh membrane-
spanning regions of the Engelman plot, in each case. This is a greater number
of transmembrane segments than has been reported for the corresponding
components of other ABC transporters of amino acids. Indeed, six membrane-
spanning regions has been suggested as the paradigm for each integral
membrane component of an ABC transporter (Higgins, 1992).
105
Fig. 3.7 Topology prediction for AapM generated by the program TOP-PRED (Claros & von Heijne, 1994). The hydrophobicity profile (top) calculated by the method of Engelman et al. (1986) with a full window of 21 amino acids and a core window of 11 amino acids, indicates 6 certain and 3 putative membrane-spanning regions. The most likely topology (bottom), selected from the various possible topologies that include all the certain transmembrane segments, but either include or exclude each of the putative ones, is that which has the greatest degree of bias in the distribution of positively charged residues. LL, loop length; KR, number of lysine and arginine residues; KR Diff, positive charge difference.
106
Fig. 3.8 Topology prediction for AapQ generated by the program TOP-PRED (Claros & von Heijne, 1994). The hydrophobicity profile (top) calculated by the method of Engelman et al. (1986) with a full window of 21 amino acids and a core window of 11 amino acids, indicates 8 certain membrane-spanning regions. The most likely topology, on the basis of the distribution of positively charged residues, is illustrated (bottom). LL, loop length; KR, number of lysine and arginine residues; KR Diff, positive charge difference.
107
Sequence alignment of AapM and AapQ with each other, and the integral
membrane components of ABC transporters that mediate uptake of polar amino
acids or glutamine, reveals a region of 63 amino acids that contains 18 well
conserved residues (Fig. 3.9). On the basis of the topologies illustrated in Figs.
3.7 and 3.8, the conserved residues in both AapM and AapQ lie within two
membrane-spanning regions and the cytoplasmic loop that connects them. In
the case of AapM it is transmembrane segments 4 and 5 (Fig. 3.7) that are
involved, while for AapQ segments 2 and 3 (Fig. 3.8) contain the conserved
amino acids. The reason for this difference is not clear, however in the case of
AapQ, the location of the conserved residues may indicate a significant
difference to MalF, the integral membrane component of the ABC transporter of
maltose in E. coli. In MalF the presence of eight transmembrane segments has
been demonstrated experimentally (Froshauer et al., 1988). However, the two
N-terminal transmembrane segments can be deleted without loss of protein
function (Ehrmann et al., 1990). This has given credence to the suggestion that
MalF consists of the six standard membrane-spanning regions, with an
extension of two transmembrane segments at the N-terminus (Overduin et al.,
1988; Higgins, 1992). The presence of conserved residues in transmembrane
segment 2 (Fig. 3.8) of AapQ, suggests that deletion of the two N-terminal
membrane-spanning regions of this protein is likely to impair function.
In all the proteins in Fig. 3.9 except those from general permeases, the
conserved region is located very close to the N-terminus. Since in AapM and
AapQ from R. leguminosarum, these conserved residues apparently constitute
two transmembrane segments and a cytoplasmic loop, it might be predicted
that the first two membrane-spanning regions of the non-general permeases in
Fig. 3.9 are joined by a cytoplasmic loop, and that therefore the N-terminus of
these proteins is periplasmic. Hydropathy analysis of each of these proteins by
both the method of Engelman et al. (1986), and that of Kyte and Doolittle (1982),
taking account of the "positive-inside rule" of von Heijne (1986; 1992), predicts
108
five membrane-spanning regions, with a periplasmic N-terminus and a
cytoplasmic C-terminus in the majority of cases. Most exceptions arise from use
of the Engelman scale, which leads to the prediction of four transmembrane
segments for OccQ, YckJ, ArtQ, GltJ, HisM, and GlnP from E. coli and R.
prowazekii, while a Kyte and Doolittle plot results in a prediction of six
transmembrane segments for GltK, and GlnP from R. prowazekii. However, a
five-transmembrane-segment topology has been demonstrated experimentally
for HisM and HisQ (Kerppola et al., 1991; Kerppola & Ames, 1992), and in
addition to producing a consistent location for the N-terminal conserved region,
the five-transmembrane-segment topology places the "integral membrane
component signature" sequence (Saurin et al., 1994) in the cytoplasm in each of
the proteins analyzed. This is consistent with the known location of this
conserved sequence in other ABC transporters (Higgins, 1992). It may therefore
be the case that five, rather than six, membrane-spanning regions is the norm
for the integral membrane components of ABC transporters of this group of
amino acids.
Some of the residues in the region illustrated in Fig. 3.9 may be involved in
substrate specificity, since spontaneous mutants of S. typhimurium in which the
specificity of the histidine transporter is altered from L-histidine to L-histidinol,
are found to contain a deletion from HisM of four amino acids, corresponding
to positions 25-28 of this region (Payne et al., 1985).
In addition, there is a correlation between the nature of the amino acid that
replaces the leucine at position 53 of the conserved region (Fig. 3.9), and the
nature of the substrate transported by the proteins in which this leucine is not
conserved. In NocQ, OccQ, ArtQ and HisQ, each of which constitutes one half
of the integral membrane complex of a transporter of (substituted) basic amino
acid(s), the leucine at position 53 is replaced by an acidic amino acid, either
glutamate or aspartate (Fig. 3.9). Similarly, in one of the two integral membrane
components of the general amino acid permease of both E. coli (see below) and
109
R. leguminosarum, this leucine is replaced by proline (Fig. 3.9). In both the
integral membrane components (or the sole integral membrane component) of
the remaining transporters in Fig. 3.9, all of which transport either glutamate or
glutamine, the leucine at position 53 is either conserved or replaced by a similar
neutral amino acid.
For each of the integral membrane proteins in Fig. 3.9, position 53 of the
conserved region is predicted to lie within a transmembrane segment, as are
residues 25-28 of the conserved sequence in HisM. Thus this data appears to be
compatible with substrate specificity being determined within the pore formed
by the integral membrane components of the transporter.
110
111
As well as the four complete ORFs in the sequence in Fig. 3.5, another
incomplete ORF which starts at base 351 and is transcribed in the opposite
direction to the aap operon was investigated. The deduced amino acid sequence
from this ORF (Fig. 3.5) shows significant homology to the N-terminus of MetC
from Bordetella avium and E. coli (39.6% and 33.6% identity, respectively).
Furthermore, translations of sequence obtained using a primer to the 5' end of
the Tn5-lacZ in mutated pRU3024 cosmids in which the insertion lies
downstream of the start of this ORF , also show homology to MetC (Section
3.2.23). It therefore seems likely that this ORF encodes beta-cystathionase in R.
leguminosarum.
3.2.5 Other features of the nucleotide sequence from pRU189
Downstream of the stop codon at the end of aapP there is an inverted repeat,
centred at positions 5134-5, with potential for forming a G/C-rich stem-loop
structure (∆G = -156.5 kJ mol-1, calculated according to Tinoco et al. (1973))
followed by at least four T residues. These are the characteristics of a rho-
independent terminator (Rosenberg & Court, 1979; Platt & Bear, 1983).
The intergenic region between aapJ and aapQ also contains an inverted repeat
(centred at a position 1773) with potential for forming a stem-loop structure (∆G
= -128.9 kJ mol-1, calculated according to Tinoco et al. (1973)). Such stem-loop
structures are probably involved in differential gene expression in polycistronic
operons (Higgins et al., 1988), and this role has been suggested for extragenic
palindromic sequences following the coding region for the periplasmic protein
HisJ in the histidine transport operon of S. typhimurium (Higgins et al., 1982a;
Higgins et al., 1982b; Stern et al., 1988).
112
3.2.6 Mutation of the general amino acid permease
Tn5-lacZ mutants of strain 3841 were generated by homogenotization
(Ruvkun & Ausubel, 1981) of mutated pRU3024 cosmids that showed impaired
ability to increase amino acid uptake. The location of the mutation in each of
the resulting strains, which was confirmed by Southern blotting (Fig. 3.10), is
illustrated in Fig. 3.11. In the case of mutations lying within the sequenced
5.4kb MluI-ClaI region, the position of the transposon was mapped precisely by
sequencing the corresponding cosmid DNA adjacent to the insert, using a
primer (P15) to the 5' end of Tn5-B20 (Fig. 3.5).
113
Fig. 3.10 Southern blots of Tn5-B20 mutants of strain 3841. A: HindIII digested chromosomal DNA probed with the 10.2kb insert from pRU135. The signal above the 10.2kb band in the 3841 lane is due to incomplete digestion. In each of the mutants the 10.2kb band is replaced by two bands (or a presumed doublet in the case of strain RU636), of the expected size. (Tn5-B20 is ~8.3kb and contains an internal HindIII fragment of ~2.8kb). B: BamHI digested chromosomal DNA probed with the 3.2kb insert from pRU133 (Fig. 3.1). Of the two bands expected to replace the ~0.8kb band of the wild-type in the strain RU632 lane, one is calculated to be only ~0.2kb, with approximately half this DNA being derived from the transposon. This fragment presumably contains insufficient DNA homologous to the probe to be detected.
114
Fig. 3.11 Restriction map of pRU3024 and mutants derived from it. Boxes represent coding regions, the unfilled arrows indicating direction of transcription. The locations of Tn5-lacZ insertions are flagged with the number of the mutant strain (and cosmid) in which they occur. Each flag points in the direction of transcription of the lacZ gene in the transposon. Filled flags represent active fusions. Restriction sites are: B, BamHI; C, ClaI; E, EcoRI; H, HindIII; M, MluI.
115
The uptake rates of glutamate and glucose for the mutants were determined
(Table 3.4).
Table 3.4 Rates of glutamate and glucose uptake by R. leguminosarum strains grown on glucose/NH4Cl. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. ND, not determined. Strain Relevant Substrate genotype L-Glutamate D-Glucose 3841 Wild-type 5.6±0.4 39.5±3.2 RU541 aapQ::Tn5-lacZ 1.7±0.1 40.0±1.6 RU542 aapP::Tn5-lacZ 1.6±0.3 36.9±1.9 RU543 aapJ::Tn5-lacZ 1.2±0.6 33.3±1.8 RU631 aapQ::Tn5-lacZ 2.0±0.1 ND RU632 cysE::Tn5-lacZ 4.6±0.3 39.5±3.2 RU633 aapM::Tn5-lacZ 1.8±0.5 ND RU634 aapM::Tn5-lacZ 1.8±0.8 44.7±3.2 RU635 aapQ::Tn5-lacZ 2.0±0.2 ND RU636 aapQ::Tn5-lacZ 1.9±0.6 37.4±0.7 RU637 aapJ::Tn5-lacZ 1.5±0.1 ND RU638 See text 2.8±0.5 ND
With the exception of RU632 and RU638, all the strains show an
approximately 3-fold reduction in glutamate uptake in comparison to the wild-
type, while glucose uptake, in the strains for which it was measured, is
unimpaired. This phenotype is consistent with mutation of the general amino
acid permease, and four of these mutants RU542, RU543, RU634 and RU636,
having mutations in aapP, aapJ, aapM and aapQ, respectively, where chosen for
further study.
Glutamate uptake in strain RU638 is impaired in comparison to the wild-type
strain, but is not as severely affected as that in strain RU543. This is consistent
with the location of the start of aapJ being as proposed in Fig. 3.5, with the
116
mutation in RU638 lying upstream of aapJ and potentially affecting the
promoter region. (Alternative ATG starts for aapJ result in the transposon in
RU638 lying within aapJ. If this were the case, glutamate uptake in strains
RU543 and RU638 would be expected to be identical).
The mutation in strain RU632 and its effects are discussed elsewhere
(Sections 3.2.20 and 3.2.21).
3.2.7 Mapping of promoter sites in the aap operon by complementation analysis
In order to determine the location of promoters for the aap genes, the ability
of plasmid-borne copies of aapJQMP, aapQMP, aapMP and aapP to complement
chromosomal mutations in aapJQMP was investigated.
While strain 3841 will grow on minimal salts agar containing glutamate as
the sole source of carbon and nitrogen, strains RU542, RU543, RU634 and
RU636 were found to be unable to grow on this medium. It was therefore
possible to use growth on glutamate as a test for complementation of mutations
in the aap operon.
Clones in both pRK415-1 and pIJ1891 of aapJQMP, aapQMP, aapMP and aapP
were each constructed from an appropriate deletion of pRU189 previously
generated for sequencing, the insert 3841 DNA being excised from pBluescript
SK+ as a HindIII-XbaI fragment (Fig. 3.12). In plasmid pRK415-1 the cloned
genes are in the opposite orientation to that of the lac promoter in the vector,
whereas in pIJ1891 transcription of the genes carried by the inserts can be
initiated by this promoter. (The lac promoter is constitutive in R. leguminosarum
(Labes et al., 1990)). The cosmid vector pIJ1891 was employed in cloning aap
genes downstream of the lac promoter, since its low copy number minimises the
possibility of over expression of these genes from the foreign promoter.
Each of the clones was conjugated into RU542, RU543, RU634 and RU636 and
the resulting strains tested for growth on minimal salts agar containing
glutamate as the sole carbon and nitrogen source (Fig. 3.12).
117
Fig. 3.12 Complementation of aap mutants for growth on 10mM glutamate as sole source of carbon and nitrogen. Growth of strains (right) containing plasmids (left) is scored: +, good growth; +/-, poor growth; -, no growth. * On incubation for a further 4-5 days these strains show good (+) growth. The shaded arrows indicate the direction of transcription initiation from the lac promoter in the vector of the plasmids. Restriction sites are: Bs, BstUI; C, ClaI; M, MluI.
118
The complete aap operon carried by either of the vectors complements a
chromosomal mutation in any of the aap genes, whilst plasmids lacking a
particular gene fail to complement a mutation in that gene. Mutations in aapQ
and aapM are not complemented by any of the clones in pRK415-1 other than
that carrying the complete aap operon, suggesting that the first three genes in
the operon are under the control of a single promoter at the start of the operon.
This is confirmed by the ability of aapQMP cloned in pIJ1891, under the control
of the lac promoter, to complement mutations in both aapQ and aapM.
Likewise, aapMP complements a mutation in aapM. Since a mutation in aapP is
eventually complemented by aapQMP, aapMP or aapP cloned in pRK415-1, it
would appear that there may be some weak promoter activity directly
upstream of aapP. Enhanced transcription of aapP relative to that of aapQM is
consistent with the expected association of two copies of AapP with one each of
AapQ and AapM within the transporter (Kerppola et al., 1991; Davidson &
Nikaido, 1991).
3.2.8 Transcription levels of aap genes
β-Galactosidase assays were carried out on the reverse mutants previously
described in Section 3.2.6, with cells being grown under both nitrogen-excess
(glucose/NH4Cl) and nitrogen-limited (glucose/glutamate) conditions.
The results (Table 3.5) for the aap mutants are consistent with the direction of
transcription for the aap operon proposed in Section 3.2.4. Only those aap
mutants in which the lacZ gene in the transposon is in the same orientation as
that proposed for the mutated aap gene show appreciable β-galactosidase
activity. Active fusions of aap genes show the expected repression of
transcription under nitrogen-excess conditions (Poole et al., 1985; Poole et al.,
1987), though whether strains mutated in their major glutamate uptake system
are likely to be simply nitrogen-limited when grown on glutamate as the
nitrogen source, may need to be taken into account when considering absolute
values (Section 4.2.2).
119
Table 3.5 β-galactosidase activities in aap mutants of R. leguminosarum strain 3841. Activities are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. Strain Relevant Growth Conditions genotype Glucose/NH4Cl Glucose/Glutamate 3841 Wild-type 15±2 22±3 RU541 aapQ::Tn5-lacZ 23±5 30±2 RU542 aapP::Tn5-lacZ 15±1 27±1 RU543 aapJ::Tn5-lacZ 155±23 690±39 RU631 aapQ::Tn5-lacZ 14±1 21±1 RU632 cysE::Tn5-lacZ 139±10 269±23 RU633 aapM::Tn5-lacZ 16±5 14±1 RU634 aapM::Tn5-lacZ 50±5 309±46 RU635 aapQ::Tn5-lacZ 38±3 123±6 RU636 aapQ::Tn5-lacZ 75±9 325±50 RU637 aapJ::Tn5-lacZ 164±41 568±37 RU638 See text 30±2 35±1
There is a considerable difference in activity between the fusions in strains
RU543/RU637 (aapJ) and those in strains RU635/RU636 (aapQ) and RU634
(aapM), both under nitrogen-excess and nitrogen-limited conditions. This result
is in accordance with the expectation that the periplasmic binding protein will
be expressed at a greater level than the membrane components of the
transporter (Higgins et al., 1982b). Since aapJ, aapQ and aapM are under the
control of a single promoter (Section 3.2.7), the difference in the expression of
these genes is presumably due to transcriptional attenuation by the putative
stem-loop located between aapJ and aapQ (Section 3.2.5).
3.2.9 Amino acid uptake in strains RU542, RU543, RU634 and RU636
The rates of uptake of a range of amino acids by strains RU542, RU543,
RU634 and RU636 grown on both glucose/NH4Cl and glucose/glutamate were
measured (Tables 3.6 and 3.7).
120
Table 3.6 Rates of amino acid transport in R. leguminosarum strains grown on glucose/NH4Cl. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures.
Substrate Strain 3841 RU542 RU543 RU634 RU636 L-Glutamate 5.6±0.4 1.6±0.3 1.2±0.6 1.8±0.8 1.9±0.6 L-Aspartate 3.9±0.1 0.7±0.2 0.8±0.2 0.9±0.3 1.0±0.3 L-Alanine 6.4±0.7 2.6±0.4 2.8±0.5 3.8±0.5 3.5±0.8 AIB 4.3±0.6 2.0±0.6 2.0±0.4 2.7±0.6 3.1±1.0 L-Histidine 5.4±0.5 1.9±0.4 2.0±0.3 2.5±0.3 2.0±0.3 L-Leucine 5.3±0.2 3.5±0.4 3.5±0.2 3.7±0.7 3.6±0.2 L-Methionine 9.0±0.1 5.3±0.9 4.6±0.2 4.2±0.1 5.0±1.0 D-Glucose 39.5±3.2 36.9±1.9 33.3±1.8 44.7±3.2 37.4±0.7
Table 3.7 Rates of amino acid transport in R. leguminosarum strains grown on glucose/glutamate. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures.
Substrate Strain 3841 RU542 RU543 RU634 RU636 L-Glutamate 26.9±2.9 5.1±1.0 5.4±1.1 3.8±0.4 3.9±2.0 L-Aspartate 18.9±0.8 2.1±0.4 2.0±0.3 1.7±0.2 2.1±0.1 L-Alanine 28.8±1.7 6.0±1.2 4.9±1.2 5.3±1.3 6.3±1.1 AIB 37.8±4.3 12.0±2.3 9.3±0.7 10.0±2.4 11.5±0.7 L-Histidine 23.4±2.0 6.3±0.2 6.0±0.8 5.1±0.3 5.5±0.3 L-Leucine 27.2±6.5 10.1±1.1 8.6±0.8 8.3±0.5 9.8±0.4 L-Methionine 15.9±0.6 6.7±0.1 6.1±0.1 6.2±0.2 6.8±0.3 D-Glucose 42.4±0.6 22.8±2.1 17.8±0.8 16.3±1.6 18.3±0.7
121
Uptake of glutamate, aspartate and histidine by the mutants is severely
affected. This suggests that the general permease is the main high affinity
transport system for these amino acids in strain 3841. Alanine, AIB, leucine and
methionine uptake is much less significantly affected and the existence of
additional, specific systems for transporting these substrates seems likely from
previous kinetic studies (Poole et al., 1985).
Initially, the uptake results from cells grown with glutamate as the nitrogen
source (i.e. nitrogen-limited) seem to suggest a greater contribution by the
general permease to the transport of alanine, AIB, leucine and methionine.
However, glucose uptake in the mutants is also substantially reduced under
these conditions, presumably because cells lacking the major glutamate uptake
system struggle to grow on glutamate. It is therefore uncertain whether the low
amino acid uptake rates in mutants grown on glutamate are due directly to the
loss of the general amino acid uptake system, or whether a significant
proportion of the reduction in uptake is attributable to a global decrease in
transport.
3.2.10 Growth of strain RU543 on amino acids as sole source of carbon and nitrogen
Strain 3841 will grow on minimal salts agar containing alanine, histidine or
proline as the sole source of carbon and nitrogen. Growth of strain RU543 on
minimal salts agar containing each of these amino acids as sole carbon and
nitrogen source was therefore investigated, in an attempt to assess the
contribution of the general amino acid permease to the uptake of these
substrates in strain 3841. The results are shown in Table 3.8. Growth of strain
RU543 on the carbon/nitrogen sources glucose/NH4Cl and glutamate was
tested as a positive and a negative control, respectively.
122
Table 3.8 Growth of R. leguminosarum strain RU543 on individual amino acids as sole source of carbon and nitrogen. Growth medium was AMS agar with C/N sources added at 20mM except for glucose and NH4Cl which were at 10mM. +++, good growth; ++, moderate growth; +, poor growth; -, no growth. C/N Source Strain 3841 RU543 Glucose/NH4Cl +++ +++ Glutamate +++ - Alanine ++ ++ Histidine + + Proline +++ +
The ability to grow on proline is significantly reduced in strain RU543
compared to that in strain 3841. This is clearly not due to the mutation in strain
RU543 causing a general impairment of growth, since growth on
glucose/NH4Cl is unaffected. It therefore seems likely that the retardation in
growth of strain RU543 is due to reduced proline uptake, suggesting that the
general amino acid permease is responsible for a significant proportion of
proline uptake in strain 3841 under these conditions.
Growth of strain RU543 on both alanine and histidine is unimpaired. In the
case of alanine this is consistent with the earlier evidence (Section 3.2.9; (Poole et
al., 1985)) for the existence of at least one alanine carrier in strain 3841 besides
the general amino acid permease. However, for histidine the result is
somewhat surprising considering the extent of the reduction in transport of this
substrate in strain RU543 (Section 3.2.9). It is noticeable that the wild-type
strain itself grows only poorly on histidine (giving growth comparable to that of
strain RU543 on proline), so one explanation could be that uptake of nutrient is
not the rate determining step in growth on histidine, and the reduced histidine
uptake in the mutant is still sufficient to support growth at the wild-type rate.
Another possibility is that in addition to the general amino acid permease strain
123
3841 possesses another histidine carrier that is either of low affinity (and hence
insignificant under transport assay conditions where the substrate is only
25µM), or only induced under certain metabolic conditions, such as those
arising from growth on histidine as sole carbon and nitrogen source.
3.2.11 Amino acid uptake in strain RU640
In order to further investigate the broad specificity of the cloned amino acid
permease, the amino acid transport properties of strain RU640 (which contains
additional copies of the aap operon carried by pRU191) grown on
glucose/NH4Cl were studied (Table 3.1).
For the amino acids tested, strain RU640 shows a 4- to 16-fold increase in
uptake compared to the wild-type. This includes aliphatic amino acids such as
leucine and alanine, in addition to polar amino acids such as glutamate,
aspartate and histidine. That this increase is not caused by the presence of the
vector pRK415-1 is demonstrated by the uptake values for strain RU913 (strain
3841/pRK415-1) which are very similar to those of strain 3841. In addition,
glucose transport in strain RU640 shows no significant increase, indicating that
enhancement of uptake is restricted to amino acids.
Strain RU640 grows extremely poorly on glucose/glutamate and uptake
rates for both glutamate and glucose are found to be lower than wild-type
under these conditions (data not shown). This apparent unhealthiness of strain
RU640 is thought to be due to the intolerably high number of copies of the
amino acid transporter that are likely to be present in cells carrying additional
copies of the aap operon under non-repressing conditions.
3.2.12 Expression of the R. leguminosarum general amino acid permease in E. coli
Following the discovery that the four proteins that comprise the general
amino acid permease in R. leguminosarum show high homology to four
124
unidentified proteins from E. coli, it was decided to investigate the effect of the
aap operon, expressed via the lac promoter, on amino acid transport in E. coli.
E. coli strain JC5412, which lacks appreciable glutamate uptake, was
transformed with pRU310 DNA and a TetR transformant purified, to generate
strain RU1050. Glutamate, Histidine and AIB uptake by strain RU1050 grown
in both LB, and LB containing IPTG, was measured (Table 3.9).
Table 3.9 Rates of amino acid transport in E. coli strains JC5412 and RU1050 grown on LB. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. Substrate Strain JC5412 RU1050 RU1050+IPTG L-Glutamate 0.74±0.02 1.06±0.21 1.22±0.10 L-Histidine 0.64±0.24 0.91±0.02 1.16±0.13 AIB 0.02±0.01 0.80±0.41 1.61±0.81
Uptake of all three amino acids is elevated in strain RU1050 in comparison to
strain JC5412, with further elevation of uptake apparent in strain RU1050 grown
in the presence of IPTG. Transport of AIB is particularly informative since in
the absence of the general amino acid permease uptake of this substrate is
barely detectable, while in its presence AIB uptake is similar to that of both
glutamate and histidine. This is consistent with functional expression of the
general amino acid permease from R. leguminosarum strain 3841 in E. coli strain
JC5412.
125
3.2.13 Physical properties of the aapJ gene product
Periplasmic proteins from strains 3841, RU542, RU543, RU640, RU913 and
RU918 grown phosphate-limited on glucose/NH4Cl were isolated by
lysosyme/EDTA treatment and subjected to SDS-PAGE (Fig. 3.13).
Fig. 3.13 SDS-PAGE gel of periplasmic proteins from strains 3841, RU542, RU543, RU640, RU913 and RU918.
The gel shows a band at approximately 33kD in the 3841, RU542 and RU913
lanes, that is greatly enhanced in the RU640 and RU918 lanes and is apparently
missing from the RU543 lane. Since strains RU640 and RU918 (Section 3.2.14)
carry additional copies of aapJ, while strains 3841, RU542 and RU913 carry only
the chromosomal copy of aapJ, and strain RU543 is an aapJ mutant, this is
compatible with the band at ~33kD being AapJ. The apparent size of this
126
presumed aapJ gene product is slightly smaller than the 34.5kD that is expected
from the sequence, however, a discrepancy between the calculated molecular
weight and the molecular weight determined on an SDS gel is not unusual
(Noel et al., 1979; Higgins et al., 1982b).
In order to confirm that the protein extracts run on the gel in Fig. 3.13
contained only periplasmic proteins, the activity of the cytoplasmic marker
enzyme malate dehydrogenase was measured in these extracts and compared
to the activity found in the cell fractions remaining after removal of the
periplasmic extracts. Activities of the periplasmic marker enzyme alkaline
phosphatase were also measured (Table 3.10).
Table 3.10 Activities of cytoplasmic and periplasmic marker enzymes in periplasmic protein fractions of R. leguminosarum strains prepared for SDS-PAGE. Activities are expressed as nmol min-1 for the total sample of cells, equivalent to 20ml of culture A600 1.0. Values are the result from a single experiment. UD, undetectable. Strain and Malate dehydrogenase Alkaline phosphatase fraction assayed Activity % of total Activity % of total 3841 Periplasm UD UD 1141 53.7 French-pressed cells 862 100.0 984 46.3 Total 862 100.0 2125 100.0 RU542 Periplasm UD UD 353 18.3 French-pressed cells 823 100.0 1579 81.7 Total 823 100.0 1932 100.0 RU543 Periplasm UD UD 302 17.5 French-pressed cells 753 100.0 1421 82.5 Total 753 100.0 1723 100.0 RU640 Periplasm UD UD 1071 51.3 French-pressed cells 997 100.0 1018 48.7
127
Total 997 100.0 2089 100.0
128
Since no malate dehydrogenase activity was observed in the periplasmic
fractions, whereas this activity was high in the French pressed samples, it can
be concluded that the samples run on the gel in Fig. 3.13 do not contain
intracellular proteins. The alkaline phosphatase data indicate that periplasmic
proteins were released from 17-54% of cells in the preparation of these samples.
The RU640 lane of the gel in Fig. 3.13 was scanned using a LKB Ultroscan XL
densitometer. The optical density of the aapJ band relative to the total optical
density of the lane, indicated that AapJ constitutes ~15% of the total periplasmic
protein in strain RU640.
3.2.14 Effect of aapJ on amino acid uptake in 3841
Since the increase in amino acid uptake by 3841 strains carrying additional
copies of the aap operon might be due solely to the binding of increased
quantities of substrate by the additional periplasmic binding protein without
attendant transport, the effect of aapJ on amino acid uptake was studied.
The 1.8kb insert in pRU256, which contains aapJ and its upstream promoter
region, was excised as a BamHI-KpnI fragment and cloned in both pRK415-1
and pIJ1891, creating pRU388 and pRU389, respectively. (Plasmid pRU256 is a
deletion of pRU190 generated for sequencing). In pRU389, but not pRU388,
transcription of aapJ can be initiated by the lac promoter in the vector. Both
pRU388 and pRU389 were conjugated into strain 3841, creating strains RU918
and RU919, respectively, and glutamate uptake by these strains was measured
(Table 3.11). (That expression of aapJ carried by pRU388 is not affected by the
single base pair deletion in this plasmid that will have been inherited from
pRU190 (Section 3.2.3), is demonstrated by the enhanced AapJ band in the
RU918 lane of the SDS-PAGE gel in Fig. 3.13, and by the ability of pRU192 to
enhance amino acid uptake in strain 3841 (Section 3.2.3)).
129
Table 3.11 Rates of L-glutamate transport in R. leguminosarum strains grown on glucose/NH4Cl. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. Strain Relevant genotype Uptake Host Plasmid 3841 Wild-type None 5.6±0.4 RU913 Wild-type Vector only 4.9±0.2 RU640 Wild-type aapJQMP 43.9±4.3 RU918 Wild-type aapJ 4.4±0.4 RU915 Wild-type aapQMP 4.5±0.1 RU914 Wild-type Vector only 5.3±0.7 RU916 Wild-type aapJQMP 23.0±1.2 RU919 Wild-type aapJ 2.6±0.3 RU917 Wild-type aapQMP 5.1±0.5
In contrast to strains RU640 and RU916 (strain 3841/pRU310), neither RU918
or RU919 shows an increase in glutamate uptake in comparison to strains
RU913 and RU914 (strain 3841/pIJ1891), respectively. This indicates that in the
wild-type the rate of substrate transport by the membrane components of the
carrier is not limited by the amount of periplasmic binding protein available for
ferrying substrate. Crucially, the additional substrate binding capacity
provided by increased amounts of binding protein does not produce an
apparent increase in uptake. The latter conclusion is confirmed by the
observation that apparent glutamate uptake is not increased in either of the
strains RU543 or RU542 carrying either pRU388 or pRU389 (data not shown).
3.2.15 Effect of aapQMP on amino acid uptake in strains 3841
In order to determine whether additional copies of the membrane
components of the general amino acid permease alone are capable of increasing
amino acid uptake in strain 3841, pRU309 and pRU313 were conjugated into
3841, creating strains RU915 and RU917, respectively.
130
Neither strain shows an increase in glutamate uptake in comparison to the
strain 3841 containing the corresponding vector alone (Table 3.11). That
pRU309 produces no effect is unsurprising since in this plasmid aapQ and aapM
are promoterless (Fig. 3.12). However, in pRU313 aapQMP are expressed via
the lac promoter in the vector (as demonstrated in the complementation
experiments described in Section 3.2.7), so a wild-type value for glutamate
uptake by strain RU917 indicates that the presence of additional copies of the
membrane components of the carrier, in the absence of increased amounts of
binding protein, does not lead to an increase in amino acid uptake.
Taken in conjunction with the results from Section 3.2.14, this result
demonstrates that the amino acid uptake rates observed in strain RU640 are due
to the intracellular accumulation of substrate rather than merely the binding of
substrate to individual components of the transporter. It also suggests that in
the wild-type cell the relative quantities of the periplasmic binding protein and
the membrane components of the amino acid permease are optimal.
3.2.16 Specificity of the aapJ gene product
The high amino acid uptake rates for strain RU640, and the wild-type level of
glutamate transport observed for strain RU917, are consistent with the idea that
the functioning of the general amino acid permease is dependent on the ability
of a single periplasmic binding protein (the aapJ gene product) to bind the
complete range of amino acids, rather than there being a number of binding
proteins interacting with the membrane component of the transporter, each of
which is specific to one, or a small number of related substrates. The latter
possibility is well documented for the histidine/LAO system from Salmonella
typhimurium and the LS/LIV-I system from E. coli (Higgins et al., 1982b; Adams
et al., 1990).
To investigate this issue further the uptake of a range of amino acids by
strain RU760 (Fig. 3.12) was measured (Table 3.12).
131
Table 3.12 Rates of amino acid transport in R. leguminosarum strains grown on glucose/NH4Cl. Rates of uptake are expressed as nmol min-1 (mg protein)-1. Values are the mean±SEM of determinations from three or more independent cultures. Substrate Strain 3841 RU543 RU760 L-Glutamate 5.6±0.4 1.2±0.6 1.2±0.6 L-Aspartate 3.9±0.1 0.8±0.2 0.4±0.2 L-Alanine 6.4±0.7 2.8±0.5 3.3±0.2 AIB 4.3±0.6 2.0±0.4 1.6±0.7 L-Histidine 5.4±0.5 2.0±0.3 2.4±0.1 L-Leucine 5.3±0.2 3.5±0.2 2.1±0.5
In strain RU760 the chromosomal copy of the aap operon is not functional
because of the transposon in aapJ, but functional copies of aapQMP are provided
by pRU313. If binding proteins other than the aapJ gene product are supporting
amino acid transport by the aapQMP gene products, strain RU760 would be
expected to exhibit increased uptake of a sub-set of amino acids in comparison
to strain RU543. This assumes that the genes encoding other binding proteins
do not lie immediately downstream of the aap operon where they could
potentially be affected by the transposon in aapJ. However, no significant
difference is observed between strains RU760 and RU543 in the uptake of any of
the amino acids tested.
3.2.17 Substrate-binding activity of AapJ
In an endeavour to demonstrate directly the broad specificity of the
periplasmic binding protein component of the general amino acid permease,
amino acid-binding activity in the periplasmic protein fraction from strains
3841, RU640 and RU543 was studied.
As an initial experiment an attempt was made to assay the substrate-binding
affinity for glutamate using ammonium sulphate precipitation (Richarme &
132
Kepes, 1983; Dahl & Manson, 1985; Higgins et al., 1987). Periplasmic proteins
from each of the strains 3841, RU640 and RU543 were isolated as described in
Section 2.1.6. The crude extracts were dialysed overnight in 5mM HEPES,
pH7.2 and concentrated, before being assayed as described in Section 2.1.7.
However, no glutamate-binding activity was detected by this method in any of
the extracts. This was presumed to be due to the glutamate-binding protein
complex failing to survive the precipitation step.
The second approach to the problem employed non-denaturing
polyacrylamide gel electrophoresis, a technique which has been used
successfully in the demonstration of substrate binding by other periplasmic
proteins (Nobile & Deshusses, 1988; Le Rudulier et al., 1991; Talibart et al., 1994).
Dialysed and concentrated periplasmic protein extracts from strains 3841,
RU640 and RU543 were prepared as above. Each of these extracts was then
incubated with each of the substrates [14C]glutamate, [14C]aspartate,
[14C]alanine, [14C]AIB, [14C]histidine and [14C]leucine, before being subjected to
non-denaturing PAGE and autoradiography as described in Section 2.1.7.
In a trial non-denaturing PAGE gel of crude periplasmic protein extracts, the
presumed Aap binding protein band is clearly visible in strain 3841, absent
from strain RU543, and greatly enhanced in strain RU640 (Fig. 3.14).
133
Fig. 3.14 Non-denaturing PAGE gel of periplasmic proteins from strains 3841, RU543 and RU640.
However, the autoradiographs of the gels run following incubation of
protein extracts with radioactive amino acids indicate no labelling of this band
by any of the substrates tested. Nor was labelling of any other periplasmic
protein apparent except in the case of leucine. In the autoradiograph of the gel
loaded with proteins incubated with leucine, each of the three lanes
corresponding to the different strains exhibits an identical single band,
produced by a protein that runs further on the gel than the Aap binding protein
(Fig. 3.15).
134
Fig. 3.15 Autoradiograph of non-denaturing PAGE gel of periplasmic proteins from strains 3841, RU543 and RU640 pre-incubated with [14C]leucine. The control contained no protein.
It is probable that this band corresponds to the binding protein from a
leucine specific transporter and that the highly specific interaction between this
protein and its substrate is strong enough to enable the protein-substrate
complex to survive the electrophoresis conditions. The same electrophoresis
conditions are apparently too harsh for the Aap binding protein-substrate
complex, which is perhaps not surprising given that a binding protein with a
very broad specificity may have a relatively "loose" binding site.
It seemed possible that the use of less stringent assay conditions than had
been previously employed might allow substrate-binding by the Aap binding
protein to be observed. Unfortunately the equilibrium dialysis method (Argast
& Boos, 1979; May et al., 1986) was not financially feasible if a number of
potential substrates were to be screened. The following approach was therefore
adopted: Periplasmic protein extracts from strains 3841, RU640 and RU543
were each dialysed, concentrated, and incubated with substrates exactly as
described for the non-denaturing gel experiment. The radioactivity in each
135
incubation mixture was determined by the scintillation counting of a small
sample. The incubation mixtures were then each spun through a
microconcentrator with a molecular weight cut-off below the size of AapJ, and
the radioactivity of the filtrates determined. Comparison of the radioactivity
present in the filtrate to that in the incubation mixture prior to spinning,
enabled the amount of substrate bound to protein to be calculated (Table 3.13).
Table 3.13 Binding of amino acids by periplasmic fractions from cells of R. leguminosarum strains grown on glucose/NH4Cl. Binding is expressed as pmol [mg protein]-1. Values are the mean±SEM of three or more replicate assays. UD, undetectable. Substrate Strain 3841 RU543 RU640 L-Glutamate 265±6 UD 1824±100 L-Aspartate 208±93 UD 1315±37 L-Alanine 534±37 535±67 594±35 AIB UD UD UD L-Histidine 357±86 UD 1470±242 L-Leucine 74±58 36±4 173±103 L-Methionine 62±54 22±13 122±28
The results for glutamate, aspartate and histidine demonstrate binding of
each of these substrates by the Aap binding protein. In each case there was
binding of the substrate by the periplasmic protein extract from the wild-type
strain 3841, substantially increased binding in strain RU640 (which carries
additional copies of aapJ), but no binding in strain RU543 (an aapJ mutant).
Leucine- and methionine-binding appear to follow a similar pattern, but the
level of binding is significantly lower. The binding of leucine by the extract
from strain RU543 is consistent with the existence of a leucine-specific binding
protein in strain 3841, as indicated by the non-denaturing PAGE gel of
136
periplasmic proteins incubated with this amino acid. Binding of alanine is
evident for each of the strains 3841, RU640 and RU543. The fact that the level of
alanine binding is the same in each of the three strains indicates that rather than
being due to the Aap binding protein, this binding is presumably caused by one
or more alanine-specific proteins. There was no detectable binding of AIB by
the periplasmic protein extracts from any of the three strains.
These results are in line with the amino acid uptake data for the aap mutants
RU542, RU543, RU634 and RU636 (Section 3.2.9). The level of binding by AapJ
of ligands such as leucine, alanine and AIB, for which the general amino acid
permease is apparently not the only significant carrier, is found to be lower than
that for substrates such as glutamate, aspartate and histidine for which this is
the predominant uptake system. That no binding at all could be detected for
alanine and AIB is perhaps not surprising as kinetic evidence suggests that the
general permease has a lower affinity for these substrates (Poole et al., 1985),
and assay conditions such as pH and ionic strength may not have been optimal
for the binding of these amino acids.
3.2.18 Amino acid exchange
Cells of R. leguminosarum strain 3841 have previously been found to exhibit
high rates of exchange of pre-loaded intracellular amino acids with amino acids
in the surrounding medium (Poole et al., 1985). In an attempt to determine the
role of the general amino acid permease in this process, it was decided to
investigate the effect of both mutation and over expression of the aap operon on
amino acid exchange by strain 3841.
Having been allowed to accumulate [14C]AIB, cells of each of the strains 3841,
RU543 and RU640 were incubated in an excess of AIB or glutamate, or had
carbonyl cyanide m-chlorophenylhydrazone (CCCP) added, and the
intracellular concentrations of [14C]AIB monitored over time. The duration of
pre-incubation with [14C]AIB was varied between strains so that the cells of
each strain contained approximately equal concentrations of the radioactive
137
substrate immediately prior to exchange, thus allowing unambiguous
comparison of efflux rates. AIB was chosen as the exchange substrate in these
experiments because it is not metabolized by R. leguminosarum strain 3841
(Section 3.2.1). Consequently the results are not complicated by incorporation
of radioactivity into cell proteins or loss of label as 14CO2.
In the wild-type strain 3841, the intracellular concentration of [14C]AIB
rapidly decreases on addition of either extracellular AIB or extracellular
glutamate (Fig. 3.16). This decrease is presumed to be the result of exchange
between the intracellular and extracellular amino acid pools (Poole et al., 1985).
In the case of exchange with glutamate, some recovery in the internal
[14C]AIB concentration is observed at longer incubation times. This may be due
to uptake of [14C]AIB by a carrier with affinity for AIB but not glutamate,
probably an alanine transporter, after glutamate has saturated the exchange
system on both sides of the membrane. This effect has been seen before in the
exchange of intracellular leucine with extracellular glutamate (Poole et al., 1985).
In strain RU543 the rate of exchange of internal [14C]AIB with external AIB
(homologous exchange) is lower than that in strain 3841, while loss of internal
[14C]AIB due to the presence of external glutamate (heterologous exchange) is
very slight in comparison to that in the wild-type (Fig. 3.17). By contrast, in
strain RU640 the rates of both homologous and heterologous exchange are
increased relative to those in strain 3841 (Fig. 3.18). The fact that in strain
RU543 homologous exchange is not as significantly affected as heterologous
exchange, points to the existence of at least one additional exchange system
specific to AIB. This is likely to be an alanine transporter.
138
Time (min)
0 10 20 30 40 50 60 70
Intra
cellu
lar l
abel
led
AIB
(nm
ol [m
g pr
otei
n]-1
)
0
10
20
30
40
50
60
70
Fig. 3.16 Efflux of AIB from cells of R. leguminosarum strain 3841. The data shown is the result from a single representative experiment. , No addition; , AIB addition; , L-glutamate addition; , CCCP addition. The time of addition is indicated by the arrow.
139
Time (min)
0 10 20 30 40 50 60 70
Intra
cellu
lar l
abel
led
AIB
(nm
ol [m
g pr
otei
n]-1
)
0
10
20
30
40
50
60
70
Fig. 3.17 Efflux of AIB from cells of R. leguminosarum strain RU543. The data shown is the result from a single representative experiment. , No addition; , AIB addition; , L-glutamate addition; , CCCP addition. The time of addition is indicated by the arrow.
140
Time (min)
0 10 20 30 40 50 60 70
Intra
cellu
lar l
abel
led
AIB
(nm
ol [m
g pr
otei
n]-1
)
0
10
20
30
40
50
60
70
Fig. 3.18 Efflux of AIB from cells of R. leguminosarum strain RU640. The data shown is the result from a single representative experiment. , No addition; , AIB addition; , L-glutamate addition; , CCCP addition. The time of addition is indicated by the arrow.
141
The proton ionophore CCCP has a profound effect on the level of
intracellular [14C]AIB in all three strains (Figs. 3.16, 3.17 and 3.18). The rapid
efflux of AIB from CCCP-poisoned cells suggests that maintaining the internal
concentration of AIB against a concentration gradient is energy dependent.
However, nothing can be deduced about the energy requirements of AIB efflux.
Curtailing energy dependent processes may result in active transport systems
becoming simply "open pores" in the cytoplasmic membrane, through which
substrates are free to diffuse. This idea could explain the observation that
CCCP treatment results in similar very rapid rates of AIB efflux from all three
strains, if it is assumed that the differences in the number of copies of the amino
acid transporter between strains, are insignificant in comparison to the total
number of transporters in each strain. That the cytoplasmic membrane has not
simply fallen apart upon treatment with CCCP is shown by the fact that the
level of intracellular [14C]AIB is not reduced to zero. The residual intracellular
[14C]AIB is lost when cells are treated with CCCP together with an excess of
unlabelled AIB (Fig. 3.19).
That the effect of CCCP is due to the loss of energy dependent processes
rather than some specific property of CCCP itself, is demonstrated by the effect
of the cytochrome oxidase inhibitor CN- on cells loaded with [14C]AIB (Fig.
3.20). It can be seen that although CN- poisoning takes longer to affect cells, it
also results in the efflux of AIB.
142
Time (min)
0 10 20 30 40 50 60 70
Intra
cellu
lar l
abel
led
AIB
(nm
ol [m
g pr
otei
n]-1
)
0
10
20
30
40
50
60
70
Fig. 3.19 Effect of excess extracellular AIB on CCCP-induced efflux of AIB from cells of R. leguminosarum strain RU640. The data shown is the result from a single representative experiment. , No addition; , AIB addition; , L-glutamate addition; , CCCP + AIB addition. The time of addition is indicated by the arrow.
143
Time (min)
0 10 20 30 40 50 60 70
Intra
cellu
lar l
abel
led
AIB
(nm
ol [m
g pr
otei
n]-1
)
0
20
40
60
80
Fig. 3.20 Efflux of AIB from cells of R. leguminosarum strain RU640. The data shown is the result from a single representative experiment. , No addition; , AIB addition; , L-glutamate addition; , KCN addition. The time of addition is indicated by the arrow.
144
The observation that amino acid exchange is reduced in strain RU543 and
increased in strain RU640 relative to the wild-type, suggests that the general
amino acid permease is involved in the exchange process. However, it is not
clear whether the efflux, as well as the uptake of amino acids, is facilitated by
the general amino acid permease itself.
Two possible models of the exchange process are illustrated in Fig 3.21.
Fig. 3.21 Schematic representation of two possible mechanisms for exchange of amino acids by R. leguminosarum: A, uptake via the general amino acid permease (AAP) and efflux via a second carrier; B, uptake and efflux via the general amino acid permease.
145
In A (Fig. 3.21), amino acid uptake is catalyzed by the general amino acid
permease, with amino acid efflux being facilitated by an entirely separate
carrier. In B, the general amino acid permease is responsible for both the
uptake and the efflux of amino acids. In both models it is suggested that uptake
and efflux are continuous competing processes, with a net build up of substrate
in the cytoplasm resulting from a greater rate of uptake than efflux. Exchange
is explained by assuming that the added extracellular excess of unlabelled
substrate effectively monopolizes uptake, while labelled substrate continues to
leave the cytoplasm via the efflux system. In the case of heterologous exchange,
the final, steady state intracellular concentrations will be a function of the
relative affinities of the two substrates for the uptake and efflux systems.
It is hard to explain the observed effects on amino acid exchange of varying
the copy number of the general amino acid permease, on the basis of model A
in Fig. 3.21. For example, if this model were correct, then the additional copies
of the general amino acid permease carried by strain RU640, would result in
increased uptake capacity without a concomitant rise in the potential for efflux.
It would therefore be expected that in strain RU640 the intracellular
concentration of the chaser substrate would rise more rapidly than in the wild-
type strain, resulting, in the case of homologous exchange at least, in a faster
rise in competition for the efflux system from the chaser, and hence a lower rate
of exchange. In fact, precisely the opposite is observed in practice (compare
Figs. 3.16 and 3.18), which suggests that the general amino acid permease is
intimately involved in the efflux process.
The mechanism by which exchange might occur in the case where the
general amino acid permease provides both uptake and efflux facilities (B, Fig.
3.21) is not known. It seems clear that the general amino acid permease actively
imports amino acids. However, since previously studied ABC transporters are
reported to be uniporters, catalysing transport of substrates in one direction
only (Higgins, 1992), it seems unlikely that this transporter also energizes amino
146
acid efflux. One alternative possibility is that amino acids simply leak out of
the cytoplasm through the general amino acid permease. This may be
particularly feasible for a relatively small amino acid such as AIB, since the
broad specificity of the amino acid permease presumably necessitates it
providing a relatively "wide" pore in the cytoplasmic membrane.
If efflux of intracellular AIB is the result of diffusion through the general
permease, it might be anticipated that the periplasmic binding protein of the
transporter would be unnecessary for such efflux. In order to investigate this
possibility, exchange of AIB in strain RU760 was examined (Fig. 3.22) and
compared to that in strain RU543.
Strain RU760 carries functional copies of aapQMP but not aapJ (Section 3.2.7),
whereas in strain RU543 aapJQM are not transcribed. Therefore, if the three
proteins AapQ, AapM and AapP alone permit efflux of AIB, the rates of
exchange in strain RU760 are expected to be greater than those in strain RU543.
This does not appear to be the case (compare Figs. 3.17 and 3.22), suggesting
that AapJ is essential for efflux via the general amino acid permease.
AapP also appears to be essential to exchange, as loss of pre-loaded [14C]AIB
from strain RU542 on addition of external glutamate or AIB (Fig. 3.23), occurs at
a similar rate to that observed for strain RU543 (Fig. 3.17).
147
Time (min)
0 10 20 30 40 50 60 70
Intra
cellu
lar l
abel
led
AIB
(nm
ol [m
g pr
otei
n]-1
)
0
10
20
30
40
50
60
70
Fig. 3.22 Efflux of AIB from cells of R. leguminosarum strain RU760. The data shown is the result from a single representative experiment. , No addition; , AIB addition; , L-glutamate addition. The time of addition is indicated by the arrow.
148
Time (min)
0 10 20 30 40 50 60 70
Intra
cellu
lar l
abel
led
AIB
(nm
ol [m
g pr
otei
n]-1
)
0
10
20
30
40
50
60
70
Fig. 3.23 Efflux of AIB from cells of R. leguminosarum strain RU542. The data shown is the result from a single representative experiment. , No addition; , AIB addition; , L-glutamate addition. The time of addition is indicated by the arrow.
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3.2.19 Plant properties of strains RU542, RU543, RU634, and RU636
Strains RU542, RU543, RU634, and RU636 all nodulated peas, with 25 nodule
isolates of each strain all retaining kanamycin resistance. Acetylene reduction
rates (Table 3.14) show that each of these strains reduce acetylene, and hence
presumably fix nitrogen, at least as well as the wild-type strain 3841.
Table 3.14 Rates of acetylene reduction by pea nodules harbouring aap and cysE mutants of R. leguminosarum strain 3841 Reduction rates are expressed as µmol h-1 per plant. Values are the mean±SEM of three or more independent determinations. Strain Relevant Acetylene reduction genotype 3841 Wild-type 0.45±0.02 RU542 aapP::Tn5-lacZ 0.63±0.03 RU543 aapJ::Tn5-lacZ 0.56±0.05 RU632 cysE::Tn5-lacZ 0.83±0.01 RU634 aapM::Tn5-lacZ 0.56±0.05 RU636 aapQ::Tn5-lacZ 0.66±0.01
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3.2.20 Nucleotide sequence of the 0.8kb BamHI fragment of pRU3024
In an attempt to determine the nature of the gene (or genes) mutated in
pRU3031 (and hence strain RU632) and pRU3033 it was decided to sequence as
much of the DNA adjacent to the transposons in these cosmids as was
conveniently possible. The aim being to compare the sequence obtained with
sequence databases.
To this end, BamHI restriction fragments from each of pRU3031 and
pRU3033 were randomly cloned into the BamHI site of pBluescript SK+. In each
case the resulting DNA was used to transform E. coli strain DH5α, and KmR,
AmpR transformants were selected. In this way, one clone containing the
majority of the transposon plus the cosmid DNA from the end of the
transposon to the next BamHI site, was isolated from each of pRU3031 and
pRU3033. These clones are pRU383 and pRU384, respectively (Fig. 3.24).
The insert DNA in each of pRU383 and pRU384 was sequenced using the KS
primer to the pBluescript SK+ polylinker and a primer (P0) to the insertion
sequence of the transposon (Fig. 3.24). Additional sequence was obtained by
sequencing pRU3033 DNA directly using a primer (P15) to the 5' end of Tn5-
B20. The total sequence obtained, much of which is from one strand only, is
shown in Fig. 3.25.
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Fig. 3.24 Restriction map of the cysE region of pRU3024 and mutants derived from it. Tn5-B20 insertions are labelled with the designation of the mutated cosmid in which they occur. Sub-clones of the two mutated cosmids in pBluescript SK+ are shown below. The unfilled arrow indicates the direction of transcription of cysE. The labelled arrows indicate the primers used to obtain the sequence in Fig. 3.25. Restriction site is: B, BamHI.
152
153
The direction of transcription of the DNA in Fig. 3.25 was determined from
the observation that the fusions in both pRU3031 and pRU3033 are active (Fig.
3.2). Of the three possible reading frames for the sequence, that leading to the
deduced amino acid sequence given in Fig. 3.25 appeared the most likely on the
basis of the occurrence of stop codons. Screening of the GenEmbl sequence
database with this amino acid sequence reveals 60.9%, 60.3% and 40.1% identity
to CysE from S. typhimurium, E. coli and Bacillus subtilis, and 39.7% identity to
NifP from Azotobacter chroococcum. (Screening of the database with the deduced
amino acid sequences from the two other possible reading frames reveals no
significant homologies). Both cysE and nifP encode serine acetyltransferase.
Thus the sequence homology data suggest that the mutated gene in pRU3031
and pRU3033 encodes serine acetyltransferase in R. leguminosarum, particularly
since the gene directly upstream of, and transcribed divergently from, the aap
operon apparently also encodes an enzyme on the methionine biosynthetic
pathway (Sections 3.2.4 and 3.2.23).
3.2.21 Amino acid transport in strain RU632
Both pRU3031 and pRU3033 are substantially impaired in their ability to
increase amino acid uptake in strain 3841 as compared to the parent cosmid
pRU3024 (Table 3.3). However, strain RU632, the chromosomal mutant of
strain 3841 derived from pRU3031 (Section 3.2.6), grown on glucose/NH4Cl,
shows no reduction in the uptake of any of the amino acids tested in
comparison to strain 3841 (Table 3.15).
When grown on glucose/glutamate however, strain RU632 shows
considerably reduced rates of uptake for all the amino acids tested except
methionine, for which uptake is only slightly below the wild-type level (Table
3.15). The relatively high rates of methionine transport in strain RU632 under
these conditions may be due to increased expression of methionine-specific
uptake systems initiated by the low intracellular methionine levels that
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presumably arise in a methionine biosynthetic mutant under nitrogen-
limitation.
Table 3.15 Rates of amino acid transport in R. leguminosarum strain RU632 grown on glucose/NH4Cl and glucose/glutamate. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. Substrate Glucose/NH4Cl Glucose/glutamate 3841 RU632 3841 RU632 L-Glutamate 5.6±0.4 4.6±0.3 26.9±2.9 10.8±2.7 L-Aspartate 3.9±0.1 4.3±1.5 18.9±0.8 5.6±0.9 L-Alanine 6.4±0.7 6.7±0.6 28.8±1.7 12.2±1.1 AIB 4.3±0.6 6.6±0.4 37.8±4.3 16.5±2.1 L-Histidine 5.4±0.5 6.6±2.1 23.4±2.0 10.4±1.7 L-Leucine 5.3±0.2 5.1±1.1 27.2±6.5 8.6±2.1 L-Methionine 9.0±0.1 9.0±0.5 15.9±0.6 11.7±0.5 D-Glucose 39.5±3.2 44.1±1.8 42.4±0.6 25.2±2.6
Transport of glucose is also impaired in strain RU632 grown on
glucose/glutamate, but not to the same degree as that of amino acids. It
therefore appears that although some of the reduction in amino acid transport
in strain RU632 under nitrogen-limited conditions can be attributed to a global
effect of the mutation, there is also a specific effect on the general amino acid
permease. This is consistent with the observation that pRU3031 and pRU3033
do not increase amino acid transport in strain 3841 to the same degree as
pRU3024 (Table 3.3). However, it is not clear why a chromosomal mutation of
cysE has no effect on amino acid uptake under nitrogen-excess conditions when,
under the same conditions, mutation of this gene in pRU3024 reduces the ability
of this cosmid to increase amino acid uptake.
The role of serine acetyltransferase in amino acid uptake in strain 3841 is
obscure. It seems unlikely that a reduction in methionine biosynthesis per se is
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responsible for the effect on the general amino acid permease, since loss of beta-
cystathionase is expected to have at least as much of an effect on methionine
production as inactivation of serine acetyltransferase (Fig. 3.26), and metC
mutants of strain 3841 exhibit wild-type amino acid transport (Section 3.2.24).
Unlike a metC mutation, a cysE mutation is expected to be detrimental to
cysteine biosynthesis (Fig. 3.26), and it is plausible that a change in the
intracellular cysteine concentration has an effect on the nitrogen regulation of
amino acid transport. However, neither the cysE or metC mutants are
methionine or cysteine auxotrophs. Indeed, it is interesting that CysE shows
homology to NifP, and it is possible that R. leguminosarum, like Azotobacter
(Evans et al., 1991), possesses two genes encoding serine acetyltransferase. Thus
it may be the case in strain 3841, that one gene encoding serine acetyltransferase
enables sufficient cysteine biosynthesis for normal amino acid transport under
nitrogen-excess, but that expression of two serine acetyltransferase genes is
required to allow a wild-type response of amino acid transport to nitrogen-
limitation.
3.2.22 Plant properties of RU632
Strain RU632 nodulated peas, and all of 25 nodule isolates retained
kanamycin resistance. The acetylene reduction rate for this strain is shown in
Table 3.14, and from comparison of this to the wild-type rate, it is apparent that
the mutation in RU632 does not cause a decrease in acetylene reduction, and
hence presumably nitrogen fixation.
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Fig. 3.26 Methionine biosynthesis in microorganisms (Bender, 1978).
3.2.23 Nucleotide sequence adjacent to the transposon in cosmids pRU3053, pRU3082, pRU3083, pRU3084, pRU3085 and pRU3086
In the hope of identifying R. leguminosarum strain 3841 genes lying between
aapJ and cysE, nucleotide sequence adjacent to the transposon in pRU3053,
pRU3082, pRU3083, pRU3084, pRU3085 and pRU3086 was obtained by
sequencing cosmid DNA directly using a primer (P15) to the 5' end of Tn5-B20.
Translations of these sequences were screened for homology to the deduced
proteins from the GenBank and EMBL sequence databases.
Sequences obtained from pRU3053, pRU3082, pRU3084 and pRU3086 were
found to overlap, and were combined to produce one continuous length of
sequence. In addition, bases 1-351 of the sequence from pRU189 (Fig. 3.5), a
translation of which shows homology to MetC (Section 3.2.4), overlap the
sequence from pRU3084. Combination of the cosmid sequence with the 351 bp
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from pRU189 yields the nucleotide sequence in Fig. 3.27. The deduced
polypeptide from this sequence (Fig. 3.27) shows 41.5% and 37.7% identity to
beta-cystathionase from E. coli and Bordetella avium, respectively. It therefore
seems likely that the gene directly upstream of aapJ in strain 3841 is the R.
leguminosarum equivalent of metC. This gene is transcribed divergently from
aapJ.
The sequences from pRU3083 and pRU3085 did not show significant
homology to any of the sequences in the databases searched.
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159
160
3.2.24 Amino acid transport in metC mutants of strain 3841
In contrast to the cysE mutant RU632, the metC mutants RU639, RU999,
RU1000 and RU1001 generated by homogenotization of cosmids pRU3053,
pRU3082, pRU3084 and pRU3086, respectively, in strain 3841, exhibit no
significant reduction in glutamate transport under either nitrogen-excess or
nitrogen-limited conditions (Table 3.16).
Table 3.16 Rates of glutamate uptake by metC mutants of R. leguminosarum strain 3841 grown on glucose/NH4Cl and glucose/glutamate. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the result from a single experiment except those for strains 3841 and RU639 which are the mean±SEM of determinations from three or more independent cultures. Strain Growth conditions Glucose/NH4Cl Glucose/glutamate 3841 5.6±0.4 26.9±2.9 RU639 4.5±0.1 21.0±0.6 RU999 4.7 30.5 RU1000 4.8 29.5 RU1001 4.4 27.8
It is interesting that attempts to make mutants of strain 3841 by
homogenotization of pRU3083 and pRU3085 were unsuccessful, suggesting that
mutation of the gene(s) carrying the transposon in these cosmids may be lethal.
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3.3 DISCUSSION
The sequence homology data strongly suggests that the products of the four
complete R. leguminosarum strain 3841 genes carried by pRU189 comprise the
components of an ABC transporter of amino acids. A mutation in any one of
these four genes leads to a decrease in the transport of all of the amino acids
tested, and the ability of this transporter to carry a broad range of amino acids is
further demonstrated by the uptake rates for strain RU640. It is therefore
reasonable to conclude that the genes carried by pRU189 encode a general
amino acid permease, and the gene designation aap is suggested.
The general amino acid permease appears to be the main high-affinity uptake
system for glutamate, aspartate, histidine and proline in free-living cells of R.
leguminosarum strain 3841. This transporter also accounts for a significant
percentage of alanine, leucine and methionine uptake, although there are
clearly other specific uptake systems for these substrates, as indicated by earlier
kinetic studies (Poole et al., 1985).
The uptake of amino acids by strain RU760 and the results of substrate
binding assays on periplasmic fractions from cells of strains 3841, RU543 and
RU640, suggest that the general amino acid permease utilizes a single
periplasmic binding protein of broad specificity (the aapJ gene product). Such
broad specificity is unusual, and the protein, of apparent molecular weight
~33kD, shows little overall homology to any previously reported periplasmic
binding protein. In particular, AapJ shows no homology to OppA, the
periplasmic binding protein from the oligopeptide transporter of S. typhimurium
(Hiles and Higgins, 1986; Hiles et al., 1987), which also shows broad substrate
specificity, but is considerably larger (52kD) than AapJ.
The rate of exchange by strain 3841 of intracellular AIB with extracellular
AIB or glutamate appears to be directly related to the number of copies of the
aap operon present, and it is hard to envisage a mechanism for exchange in
which the general amino acid permease provides only uptake functions, that
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can account for this observation. Since ABC transporters are reported to be
uniporters (Higgins, 1992), the most likely possibility seems to be that the
general amino acid permease actively facilitates amino acid uptake but also
allows passive efflux. If this is the case, then the pore formed by the transporter
must be accessible to intracellular substrate. Whether access to the pore can be
gained from the cytoplasm at all times, or is dependent upon the binding
and/or uptake of extracellular substrate, is a critical question for the
functioning of this and possibly other ABC transporters.
The requirement of AapJ for efflux suggests that binding of external
substrate is necessary for this process. An alternative possibility is that in the
wild-type unliganded binding protein interacts with the membrane complex,
and the absence of AapJ results in a change in the conformation of the
membrane components that prevents efflux. However, this seem unlikely since
the binding protein of the histidine uptake system of S. typhimurium appears
only to interact with the membrane components of the transporter in its
liganded form (Prossnitz et al., 1988; Prossnitz et al., 1989).
While the reduction in amino acid exchange in an aapP mutant of strain 3841
may indicate that ATP hydrolysis is required to fuel efflux, it seems more likely
that the energy is needed to open a membrane pore upon binding of a liganded
binding protein complex. It is also possible that the loss of AapP alters the
conformation of the remaining components of the transporter so that efflux can
no longer occur, particularly since it is generally assumed that AapP induces a
conformational change in the transmembrane subunits upon ATP hydrolysis
(Higgins, 1992; Doige & Ames, 1993).
The physiological significance of the exchange capability of the general
amino acid permease is unknown. Clearly the conditions employed in the
exchange experiments described here are unlikely to be encountered in nature,
however, relatively high intracellular concentrations of substrates may
ordinarily be present in cells as a result of active accumulation or endogenous
163
synthesis. Consequently, intracellular substrate will flow out of the cell down a
concentration gradient if suitable pores are present in the cytoplasmic
membrane. If such pores are provided by an uptake system upon
binding/uptake of an external substrate, then there is potential for regulated
exchange of substrates, in which the passage of substrates across the membrane
is determined by the differential affinities of the substrates for a common
carrier.
Certainly the exchange capacity of the general amino acid permease is not
required for nitrogen fixation by strain 3841, as aap mutants induce pea nodules
that reduce acetylene as effectively as those induced by the wild-type. Since the
general amino acid permease appears to be the major high affinity glutamate
transporter in strain 3841, this observation suggests that nitrogen fixation in
Rhizobium leguminosarum is not fuelled via a malate-aspartate shuttle in the
nodule as proposed by Kahn et al. (1985). However, the possibility that
alternative glutamate/aspartate transporters are induced under symbiotic
conditions means that the operation of this shuttle can not be completely
discounted.
In the case of Rhizobium leguminosarum it is alanine rather than aspartate that
is found to be excreted from bacteroids and symbiosomes under nitrogen-fixing
conditions (Appels & Haaker, 1991; Rosendahl et al., 1992). If this excretion of
alanine is the primary mechanism for maintaining sufficient flux through the
TCA cycle to fuel nitrogen fixation in the bacteroid, then the fact that the
general amino acid permease is not required for an effective symbiosis is
unsurprising: Alanine uptake by aap mutants indicates the existence of one or
more alternative alanine carriers, and the results of exchange experiments on
strain RU543, using the alanine analogue AIB, suggest that such a carrier (or
carriers) allows efflux of substrate.
The striking homology between the deduced amino acid sequences of the aap
genes and those of four unidentified ORFs from the 67.4 to 74.0 minute region
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of the E. coli K12 chromosome, suggests that E. coli possesses a general amino
acid permease. Similarly, the fact that the aapJ gene product shows extremely
high homology to the polypeptide deduced from what is available of the
sequence of an unidentified ORF in P. fluorescens, but shows little homology to
other known binding proteins, makes it likely that a homologue of this
transporter is also found in Pseudomonas.
E. coli strain D2W is reported to have five separate systems for dicarboxylic
amino acid transport (Schellenberg & Furlong, 1977). These are: (i) a binding-
protein-independent, Na+-dependent, glutamate specific system; (ii) a binding-
protein-dependent, Na+-independent system for transport of glutamate and
aspartate; (iii) a binding-protein-independent, Na+-independent glutamate-
aspartate system; (iv) a binding-protein-independent, aspartate-specific system;
and (v) a dicarboxylic acid transport system that carries aspartate in addition to
malate, fumarate and succinate. Genes corresponding to systems (i) and (iii)
from E. coli K12 and/or E. coli B have been cloned and sequenced (Deguchi et
al., 1989; Deguchi et al., 1990; Kalman et al., 1991; Wallace et al., 1990; Tolner et
al., 1992b), and the nucleotide sequence of three genes encoding the membrane
components of a binding-protein-dependent glutamate-aspartate transporter
(corresponding to (ii) above) from E. coli K12 has been entered in the EMBL
database (accession number U10981, B. Wallace (1994)). However, the latter
sequence is not the same as that found in the 67.4 to 74.0 minute region of the
K12 chromosome, and although the deduced ATP-binding protein from this
sequence shows 60.2% identity to AapP, the other two membrane components
show less than 30% identity to either AapQ or AapM. It therefore seems
possible that the general amino acid permease-like transporter is a previously
uncharacterized system for glutamate and aspartate uptake in E. coli.
Since Aap-like proteins are so highly conserved between Escherichia coli,
Pseudomonas fluorescens and Rhizobium leguminosarum it seems reasonable to
conclude that this transporter has an important physiological role, and may be
165
common to a wide range of Gram-negative bacteria. However, the fact that
mutation of aap genes is not lethal to R. leguminosarum, and that the Aap-like
system in E. coli has apparently gone undetected, suggests that this system is
not prominent under laboratory culture conditions.
TFASTA and BLAST searches indicate homology of AapQ and AapM to the
integral membrane components of ABC transporters of polar amino acids or
glutamine. In addition to the "integral membrane component signature"
(Saurin et al., 1994), a sequence alignment of AapQ and AapM with these
proteins, reveals a region of 63 amino acids containing 18 highly conserved
residues, that is located, in the majority of cases, at the N-terminal end of the
protein. This suggests that amino acid transporters of this type may constitute a
sub-family of the ABC superfamily.
The presence of two conserved regions in proteins belonging to this sub-
family, provides a constraint for topological models, since such regions are
expected to occur in similar locations (i.e. periplasmic, transmembrane, or
cytoplasmic) in each protein. Predicted topologies containing eight
transmembrane segments in the case of the general permeases, and five
membrane-spanning regions in the case of the other known members of the
sub-family, are consistent with this constraint. Indeed, a five-transmembrane-
segment topology for HisM and HisQ from S. typhimurium has been confirmed
experimentally (Kerppola et al., 1991; Kerppola & Ames, 1992). In these
topologies the N-terminal conserved region spans two transmembrane
segments and a connecting cytoplasmic loop, while the integral membrane
component signature is located in a cytoplasmic loop.
The N-terminal conserved region may be involved in substrate specificity,
since for one position in this sequence there appears to be a correlation between
the nature of the substrate translocated, and the nature of the amino acid found
at that position; while deletion of four other amino acids from this N-terminal
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region in HisM of S. typhimurium, results in a change of substrate specificity
from L-histidine to L-histidinol (Payne et al., 1985).
On the basis of sequence homology, the gene lying immediately upstream of
the aap operon in R. leguminosarum strain 3841, is likely to be metC, encoding
beta-cystathionase. This gene is transcribed divergently from the aap operon,
and assuming it is of a similar size to its E. coli counterpart, its 3' end lies
approximately 2.4kb upstream of the probable start of a gene, designated cysE,
which yields a deduced polypeptide with significant homology to serine
acetyltransferase. Both these enzymes are involved in methionine biosynthesis.
However, the β-galactosidase activities produced by Tn5-lacZ mutants of
pRU3024 in which the transposon has inserted between the 3' end of metC and
the start of cysE (pRU3083 and pRU3085), are consistent with transcription of a
gene(s) in the opposite orientation to metC and cysE (Fig. 3.2), suggesting that
these genes are not contained in one operon.
Mutation of cysE, but not metC, results in a reduction in uptake by the
general amino acid permease under nitrogen-limitation. Such a reduction in
transport might occur if the aap operon is regulated in response to a metabolite
level(s) that is altered as a consequence of the mutation of cysE, but not metC.
The fact that a mutation in cysE only affects amino acid uptake under nitrogen-
limitation, is compatible with the possible existence of two genes encoding
serine acetyl transferase in R. leguminosarum, as suggested by the homology of
CysE to NifP from Azotobacter: The expression of one gene encoding serine
acetyltransferase may be sufficient to allow wild-type amino acid transport
under nitrogen excess, however the level of serine acetyltransferase resulting
from expression of two genes, may be required to enable a normal response of
amino acid transport to nitrogen-limitation.
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CHAPTER 4 NITROGEN REGULATION OF THE GENERAL AMINO ACID PERMEASE OF RHIZOBIUM LEGUMINOSARUM STRAIN 3841
168
4.1 INTRODUCTION
It has previously been reported that glutamate uptake by R. leguminosarum
strain 3841 is significantly lower under conditions of nitrogen-excess than it is
under nitrogen-limitation (Poole et al., 1985; Poole et al., 1987). The cloning and
mutation of the genes encoding the general amino acid permease of strain 3841,
described in Chapter 3, has shown that this transporter is the major high-
affinity uptake system for glutamate in this strain.
It was therefore decided to investigate the regulation of the aap operon in
response to nitrogen supply. In particular, the role of the nitrogen response
regulator NtrC was examined. The results are discussed in this chapter.
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4.2 RESULTS
4.2.1 Effect of the metC-aapJ intergenic region on growth of strain 3841
For the purpose of monitoring transcription of the aap operon in R.
leguminosarum strain 3841, a lacZ-fusion to the N-terminus of aapJ and its
upstream promoter region was created by cloning a 1.0kb EcoRI-PstI fragment
from pRU189 into the corresponding sites in the polylinker of the expression
vector pMP220.
The resulting plasmid, pRU393 (Fig. 4.1), was conjugated into strain 3841,
creating strain RU990. The intention was to measure β-galactosidase activity in
strain RU990 grown on both glucose/NH4Cl and glucose/glutamate. However,
the presence of pRU393 appears to be extremely detrimental to strain 3841, and
growth of strain RU990 on minimal media was deemed too poor to yield any
meaningful results.
Fig. 4.1 Map of pRU383 showing the location and orientation of the promoterless lacZ gene in the vector.
Since strain RU368 (3841/pMP220) exhibits no difference in growth to that of
the wild-type, it can be concluded that it is the additional copies of the
intergenic region between metC and aapJ, carried by pRU393, that are
deleterious to the growth of strain RU990. A possible reason for this effect is
that one or more transcriptional regulators that normally bind to the native
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metC-aapJ intergenic region, and perhaps other promoter regions in the
chromosome, are bound to plasmid-borne intergenic regions in RU990, leading
to potentially harmful changes in the regulation of certain genes in this strain.
Indeed, it has previously been observed that over expression of the aap operon
can cause poor growth of strain 3841 (Section 3.2.11).
4.2.2 Effect of nitrogen supply on the transcription of aapJQM
Since the aapJ::lacZ reporter fusion, pRU393, could not be used, it was
decided to use the aapJ::lacZ, aapQ::lacZ and aapM::lacZ fusions in cosmids
pRU3028, pRU3046 and pRU3035, respectively, to study transcription of the aap
operon. The disadvantage to the use of these fusions is the potentially
consequential introduction into the host strain of additional copies of the other
genes carried by the cosmid. Indeed, all these cosmids necessarily contain the
metC-aapJ intergenic region. However, unlike pRU393, none of these cosmids
appears to affect the growth of strain 3841, suggesting that the smaller numbers
of the metC-aapJ intergenic region introduced into the host by the cosmids, in
comparison to pRU393 (due to the lower copy number of the cosmids), do not
have such a significant impact on the cell.
Strains RU443, RU506 and RU517, generated by conjugating cosmids
pRU3028, pRU3035 and pRU3046, respectively, into strain 3841, were grown
under both nitrogen-excess (glucose/NH4Cl) conditions and nitrogen-limited
(glucose/glutamate) conditions, and β-galactosidase activities measured in each
case (Table 4.1).
The transcription-attenuating effect of the putative stem-loop located
between aapJ and aapQ is evident in the relative level of β-galactosidase activity
produced by pRU3028 (aapJ::lacZ) compared to that from pRU3035 (aapM::lacZ)
and pRU3046 (aapQ::lacZ) under the same growth conditions. However, all
three fusions exhibit a similar degree of repression under nitrogen-excess
conditions.
171
Table 4.1 Effect of ntrC on plasmid-borne aap::lacZ fusion activity in R. leguminosarum strain 3841. Strains were grown on AMS containing either glucose/NH4Cl or glucose/glutamate at 10mM. Activities are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. Strain Relevant genotype Growth conditions Host Cosmid Glucose/NH4Cl Glucose/glutamate RU438 (3841/pRU3024) Wild-type aapJQMP 40±5 43±1 RU443 (3841/pRU3028) Wild-type aapJ::Tn5-lacZ 3325±279 7923±666 RU506 (3841/pRU3035) Wild-type aapM::Tn5-lacZ 752±208 2103±184 RU517 (3841/pRU3046) Wild-type aapQ::Tn5-lacZ 1221±265 2853±331 RU1013 (RU929/pRU3024) ntrC::Ω aapJQMP 34±1 34±2 RU980 (RU929/pRU3028) ntrC::Ω aapJ::Tn5-lacZ 7278±546 8256±1738 RU983 (RU929/pRU3035) ntrC::Ω aapM::Tn5-lacZ 2113±527 1484±309 RU984 (RU929/pRU3046) ntrC::Ω aapQ::Tn5-lacZ 2096±517 2121±350
172
Lower rates of uptake by the general amino acid permease of strain 3841
grown on glucose/NH4Cl compared to those of the same strain grown on
glucose/glutamate, have been previously reported for both batch cultures
(Poole et al., 1985) and chemostat cultures (Poole et al., 1987). However, the
reduction in transcription (~3-fold) between glutamate-grown and NH4Cl-
grown cells indicated by the results in Table 4.1, is significantly less than the
reduction in glutamate transport observed between corresponding batch
cultures (~5-fold) and chemostat cultures (~17-fold). This discrepancy is
discussed later in this Section.
In order to investigate whether the repression of the aap operon under
nitrogen-excess conditions is mediated by ntrC, cosmids pRU3028, pRU3035
and pRU3046 were conjugated into strain RU929, an ntrC interposon mutant of
strain 3841. The resulting strains, RU980, RU983 and RU984, respectively, were
grown on both glucose/NH4Cl and glucose/glutamate, and β-galactosidase
activities measured in each case (Table 4.1).
After nitrogen-limited growth, the activities of all three fusions is similar in
the strain RU929 background to that in the strain 3841 background. However,
under nitrogen-excess conditions, transcription of aapJQM in strain RU929
occurs at a similar level to that obtaining under nitrogen-limitation. This
suggests that NtrC may negatively regulate the aap operon in R. leguminosarum
strain 3841. Certainly the data in Table 4.1 are indicative of the involvement in
some way of NtrC in the regulation of the aap operon, since both nitrogen-
limitation, which results in the phosphorylation of NtrC, and mutation of NtrC,
result in a similar increase in the transcription of aapJQM from that found in a
nitrogen-excess wild-type background.
In view of the effect of pRU393 on the growth of strain 3841, it was
considered that the presence of additional copies of the metC-aapJ intergenic
region in the strains in Table 4.1 may have affected the β-galactosidase activities
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observed in these strains. It was therefore decided to study the activities of a
lacZ fusion to the chromosomal copy of each of aapJQM.
To this end, strains RU1017, RU1018 and RU1019, containing mutations in
both ntrC and aapJ, aapM or aapQ, respectively, were generated by
homogenotization of pRU3028, pRU3035 and pRU3046 in strain RU929. The β-
galactosidase activity resulting from the Tn5-lacZ mutations in aapJQM in these
strains, and in strains RU543, RU634 and RU636 (which contain only the
corresponding single mutations of aapJ, aapM and aapQ, respectively), was
investigated after growth on both glucose/NH4Cl and glucose/glutamate
(Table 4.2).
Table 4.2 Effect of ntrC on chromosomal aap::lacZ fusion activity in R. leguminosarum strain 3841. Strains were grown on AMS containing either glucose/NH4Cl or glucose/glutamate at 10mM. Activities are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. Strain Relevant genotype Growth conditions Glucose/NH4Cl Glucose/glutamate 3841 Wild-type 15±2 22±3 RU543 aapJ::Tn5-lacZ 155±23 690±39 RU634 aapM::Tn5-lacZ 50±5 309±46 RU636 aapQ::Tn5-lacZ 75±9 325±50 RU929 ntrC::Ω 27±1 23±1 RU1017 ntrC::Ω aapJ::Tn5-lacZ 603±106 678±49 RU1018 ntrC::Ω aapM::Tn5-lacZ 250±38 278±40 RU1019 ntrC::Ω aapQ::Tn5-lacZ 352±49 344±52
β-galactosidase activities in strains RU543, RU634 and RU636 exhibit the
expected repression of the aap operon under nitrogen-excess. However, in
contrast to the results obtained from cosmid-borne fusions, the approximately
5-fold difference in activity between glucose/NH4Cl and glucose/glutamate
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cultures of these strains, is consistent with the scale of repression demonstrated
by amino acid uptake in batch cultures.
One potential problem in the interpretation of the data in Table 4.2 is the
possibility that strains mutated in their major high-affinity glutamate uptake
system may not be simply nitrogen-limited when grown on glutamate as the
sole nitrogen source. However, the observation that glucose/glutamate
cultures of the mutant strains in Table 4.2 show no significant difference in
growth to those of their parental strains, suggests that mutation of aap genes has
little effect on the utilization of glutamate as the sole nitrogen source in strain
3841. Furthermore, addition of glutamate to chemostat cultures of strain 3841
growing nitrogen-limited on NH4Cl has been previously shown to result in a
slight increase in glutamate uptake (Poole et al., 1987). It therefore seems
unlikely that the loss of glutamate uptake in aap mutants leads to an increase in
the transcription of the aap operon.
The ntrC-aapJQM double mutants exhibit the complete loss of repression of
the aap operon under nitrogen-excess previously observed for cosmid-borne
fusions in a strain RU929 background. The similarity of the β-galactosidase
activities in these mutants grown on glucose/NH4Cl to the corresponding
activities following glucose/glutamate growth, would be hard to explain if the
activity observed in glucose/glutamate cultures was affected by the reduced
glutamate uptake in these strains.
The ratio between the nitrogen-limited and nitrogen-excess levels of
transcription of the aap operon as indicated by the activity of chromosomal
fusions (Table 4.2), is more compatible with the observed rates of glutamate
transport under the two conditions (Poole et al., 1985), than that obtained from
cosmid fusions (Table 4.1). In the case of the cosmid fusions, the ratio of
nitrogen-limited to nitrogen-excess values is too low. This leads to the
conclusion that either the β-galactosidase activities in strains RU443, RU506 and
RU517 grown on glucose/NH4Cl are artificially high or the activities in
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glucose/glutamate grown RU443, RU506, RU517, RU980, RU983 and RU984,
and glucose/NH4Cl grown RU980, RU983 and RU984 are artificially low. (β-
galactosidase activities from the cosmid fusions are expected to be greater than
those from the chromosomal fusions due to their greater copy number). If the
aap operon is negatively regulated by the binding of NtrC to the upstream
intergenic region, then the binding of the finite pool of NtrC in the cell to an
increased number of metC-aapJ intergenic regions carried by the cosmids in
RU443, RU506 and RU517, might be expected to result in the partial
derepression of the amino acid permease in these strains under nitrogen-excess.
This would account for relatively high β-galactosidase activities in strains
RU443, RU506 and RU517 grown on glucose/NH4Cl.
4.2.3 Amino acid uptake in strain RU929
As an alternative measure of the effect of the mutation of ntrC on the
transcription of the aap operon in R. leguminosarum strain 3841, glutamate
uptake in strain RU929 was determined under both nitrogen-excess and
nitrogen-limited conditions (Table 4.3).
Table 4.3 Rates of glutamate uptake R. leguminosarum strain RU929 grown on glucose/NH4Cl and glucose/glutamate. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. Strain Relevant Growth conditions genotype Glucose/NH4Cl Glucose/glutamate 3841 Wild-type 5.6±0.4 26.9±2.9 RU929 ntrC::Ω 21.4±1.3 22.4±3.7
The elevation of glutamate transport in strain RU929 grown on
glucose/NH4Cl to a similar level to that found in strain 3841 grown on
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glucose/glutamate, is consistent with the aapJQM::lacZ fusion data discussed in
Section 4.2.2.
4.2.4 Effect of nitrogen supply on the transcription metC and cysE
Since mutation of cysE has an effect on amino acid uptake in strain 3841
(Section 3.2.21), it was decided to investigate the nitrogen regulation of this
gene. The nitrogen regulation of metC, which is transcribed divergently to the
aap operon, and like cysE, encodes an enzyme involved in methionine
biosynthesis, was also investigated.
Cosmids pRU3031 and pRU3033, carrying Tn5-lacZ mutated copies of cysE,
and cosmids pRU3082 and pRU3086, carrying Tn5-lacZ mutated copies of metC,
were conjugated into both strain 3841 and strain RU929. β-galactosidase
activities were measured in the resulting strains grown on both glucose/NH4Cl
and glucose/glutamate (Table 4.4).
β-galactosidase activities from corresponding chromosomal fusions were
also studied. Strains RU632, RU999 and RU1001 containing Tn5-lacZ mutations
in cysE, metC and metC, respectively, were generated by homogenotization of
pRU3031, pRU3082 and pRU3086 in strain 3841. (The cysE mutant derived
from pRU3033 could not be isolated). The ntrC-metC double mutants RU1029
and RU1030 were generated by the homogenotization in strain RU929 of
pRU3082 and pRU3086, respectively; while homogenotization of pRU3033 in
strain RU929 generated the ntrC-cysE double mutant RU1027 (homogenotes of
pRU3031 in RU929 could not be isolated). β-galactosidase activities in these
strains were measured under both nitrogen-excess and nitrogen-limited
conditions (Table 4.5).
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Table 4.4 β-galactosidase activities in R. leguminosarum strains 3841 and RU929 containing pRU3031, pRU3033, pRU3080 and pRU3086. Strains were grown on AMS containing either glucose/NH4Cl or glucose/glutamate at 10mM. Activities are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. Strain Relevant genotype Growth conditions Host Cosmid Glucose/NH4Cl Glucose/glutamate RU438 (3841/pRU3024) Wild-type metC cysE 40±5 43±1 RU502 (3841/pRU3031) Wild-type cysE::Tn5-lacZ 878±189 1313±156 RU504 (3841/pRU3033) Wild-type cysE::Tn5-lacZ 1324±103 2096±103 RU974 (3841/pRU3082) Wild-type metC::Tn5-lacZ 967±183 1188±64 RU978 (3841/pRU3086) Wild-type metC::Tn5-lacZ 1010±140 1133±107 RU1013 (RU929/pRU3024) ntrC::Ω metC cysE 34±1 34±2 RU981 (RU929/pRU3031) ntrC::Ω cysE::Tn5-lacZ 1213±150 1285±134 RU982 (RU929/pRU3033) ntrC::Ω cysE::Tn5-lacZ 2118±168 2162±267 RU986 (RU929/pRU3082) ntrC::Ω metC::Tn5-lacZ 1197±129 1256±409 RU988 (RU929/pRU3086) ntrC::Ω metC::Tn5-lacZ 1055±47 1378±34
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Table 4.5 β-galactosidase activities in R. leguminosarum strains RU632, RU999, RU1001, RU1026, RU1027, RU1029 and RU1030. Strains were grown on AMS containing either glucose/NH4Cl or glucose/glutamate at 10mM. Activities are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. Strain Relevant genotype Growth conditions Glucose/NH4Cl Glucose/glutamate 3841 Wild-type 15±2 22±3 RU632 cysE::Tn5-lacZ 139±10 269±23 RU999 metC::Tn5-lacZ 91±4 193±10 RU1001 metC::Tn5-lacZ 78±9 172±31 RU929 ntrC::Ω 27±1 23±1 RU1027 ntrC::Ω cysE::Tn5-lacZ 289±28 309±44 RU1029 ntrC::Ω metC::Tn5-lacZ 190±32 152±9 RU1030 ntrC::Ω metC::Tn5-lacZ 181±19 201±51
The regulation of cysE and metC in response to nitrogen supply results in a
similar pattern for the transcription of these genes to that observed for the aap
operon (Section 4.2.2). The data from the chromosomal fusions indicate
repression of both cysE and metC under nitrogen-excess, though the magnitude
of this effect (an approximately 2-fold reduction in transcription) is less than
that observed for the aap operon. However, as was the case for the aap operon,
mutation of ntrC results in the loss of this repression, suggesting that NtrC
negatively regulates transcription of cysE and metC.
The difference between the nitrogen-excess and nitrogen-limited levels of
transcription of cysE and metC as indicated by the β-galactosidase activities
produced by the cosmid-borne fusions to these genes in the strain 3841
background (Table 4.4), is not as great as that indicated by the chromosomal
fusions. This may be explained by the presence of additional copies of the
metC-aapJ intergenic region in strains containing the cosmids, leading to
derepression of genes negatively regulated by NtrC, as was suggested to
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account for the discrepancy between the chromosomal and cosmid fusion data
for transcription of the aap operon (Section 4.2.2).
The difference in β-galactosidase activities produced by pRU3031 and
pRU3033 under the same conditions (Table 4.4) appears to reflect the relative
distances of the fusions in these cosmids from the N-terminus of cysE (Fig. 3.24),
and may indicate occasional incomplete transcription of this gene.
4.2.5 Sequence analysis of the metC-aapJ intergenic region
The nucleotide sequence between the methionine start of metC and that of
aapJ, contains a putative NtrC binding site (Fig 4.2). This binding site could
potentially regulate both promoters.
180
181
4.3 DISCUSSION
The data presented in this chapter suggest that the aap operon of R.
leguminosarum strain 3841 is negatively regulated by NtrC. The results from
both glutamate uptake assays and aapJQM::lacZ transcriptional fusion studies
show that the aap operon is fully derepressed in an ntrC mutant of strain 3841
under both nitrogen-excess and nitrogen-limited conditions. This is in contrast
to the wild-type strain in which nitrogen-excess leads to an approximately 5-
fold repression of the aap operon relative to the nitrogen-limited state. In
addition, a putative NtrC-binding site can be identified in the metC-aapJ
intergenic region.
While transcriptional activation by NtrC, under nitrogen-limitation, of genes
encoding amino acid transporters in enteric bacteria has been observed (Ames
& Nikaido, 1985; Nohno et al., 1986; Schmitz et al., 1988; Claverie-Martin &
Magasanik, 1991), negative regulation of an amino acid uptake system by NtrC
has not been reported previously. However, negative regulation by NtrC in R.
leguminosarum is not unprecedented, as NtrC has been found to repress
transcription of the operon containing ntrBC in this bacterium (Patriarca et al.,
1993).
Details of the mechanism by which NtrC regulates aap gene expression in
response to nitrogen supply have not been investigated. Any model has to
account for the fact that both loss of NtrC (through mutation), and
phosphorylation of NtrC (the presumed result of nitrogen-limitation), lead to
the derepression of the aap operon.
One potential explanation is that phosphorylated NtrC (NtrC-P) has a
significantly lower affinity for the binding sites that impair transcription of
aapJQMP than the unphosporylated form. Thus the phosphorylation of NtrC
under nitrogen-limiting conditions would lead to increased expression of these
genes. However, this seems unlikely, particularly since in K. pneumoniae, NtrC-
P is reported to bind more effectively as a negative regulator of ntrBC
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expression, than NtrC (MacFarlane & Merrick, 1987). An alternative possibility
is that NtrC-P, but not NtrC, has a lower affinity for the binding sites upstream
of aapJQMP, than it does for those involved in controlling the expression of
other genes, such as glnII, which has been shown to be positively regulated by
NtrC (Carlson et al., 1987; Martin et al., 1988; de Bruijn et al., 1989; Rossi et al.,
1989; Shatters et al., 1989; Patriarca et al., 1992). In this case, if the total
concentration of NtrC (phosphorylated and unphosphorylated) in the cell is
largely unaffected by the nitrogen supply, then under nitrogen-limitation, the
amount of NtrC (in the form of NtrC-P) available to bind to the promoter region
of aapJQMP will be reduced, and transcription of these genes consequently
increased. The observation that, in contrast to enteric bacteria where NtrC
activates its own transcription under nitrogen-limited conditions (Reitzer &
Magasanik, 1987; Merrick, 1988), transcription of ntrC in R. leguminosarum is
essentially independent of nitrogen status (Patriarca et al., 1993; Amar et al.,
1994), is therefore consistent with this model.
The presence of additional, plasmid-borne, copies of the metC-aapJ intergenic
region in strain 3841, results in severe retardation of growth. This effect might
be caused by the binding of one or more regulators to the extra binding sites
provided by the additional intergenic regions. However, the binding of NtrC to
such additional binding sites is unlikely to be solely responsible for the effect,
since the introduction into strain 3841, of pAR36A, which carries an NtrC-
binding site upstream of the glnII promoter of R. leguminosarum in the same
plasmid as carried the metC-aapJ intergenic region, is not detrimental to growth.
This suggests that at least one other regulator may bind to the metC-aapJ
intergenic region.
The cysE::lacZ and metC::lacZ fusion data suggest that both cysE and metC are
negatively regulated by NtrC in response to nitrogen supply, presumably in a
similar fashion to the aap operon.
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CHAPTER 5 INTER-REGULATION OF THE TCA CYCLE AND THE GENERAL AMINO ACID PERMEASE OF R. LEGUMINOSARUM STRAIN 3841.
184
5.1 INTRODUCTION
The excretion of the amino acids alanine and aspartate by bacteroids under
nitrogen-fixing conditions has been widely reported (Kretovich et al., 1986;
Appels & Haaker, 1991; Kouchi et al., 1991; Rosendahl et al., 1992), and a
possible role for this excretion in the regulation of the TCA cycle in the
bacteroid has been discussed (Section 1.2.5).
The results from amino acid exchange experiments presented in chapter 3,
demonstrated the potential for the involvement of the general amino acid
permease from R. leguminosarum strain 3841 in the excretion of amino acids.
However, the Fix+ phenotype of aap mutants suggests either that the general
amino acid permease is not significantly involved in excretion by the bacteroid
or that such excretion is not necessary for bacteroid function.
In this chapter the isolation of α-ketoglutarate dehydrogenase and succinyl-
CoA synthetase mutants of R. leguminosarum strain 3841, which exhibit
dramatically reduced rates of uptake by the general amino acid permease, is
described. The results of experiments performed to investigate this apparent
inter-regulation of the TCA cycle and the general amino acid permease are
discussed.
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5.2 RESULTS
5.2.1 Aspartate resistant mutants of R. leguminosarum strain 3841
Prior to the isolation of cosmid pRU3024, one strategy employed to obtain
amino acid transport mutants of R. leguminosarum was the screening of
transposon mutants of strain 3841 for resistance to a lethal concentration of a
toxic amino acid analogue. The logic being that at least some of the mutants
able to survive such toxic conditions might derive their immunity from an
inability to take up the poison. This approach has been used successfully to
obtain amino acid transport mutants in other gram-negative bacteria (Ames et
al., 1977; Kay, 1971; Weiner & Heppel, 1971; Oxender, 1972; Halpern, 1974;
Schellenberg & Furlong, 1977; Masters & Hong, 1981; Payne et al., 1985; Dila &
Maloy, 1986; Yamato et al., 1990).
In the case of R. leguminosarum however, common toxic amino acid
analogues were ineffective: The aspartate analogue β-hydroxyaspartate which
is toxic to E. coli and has been used to obtain mutants deficient in aspartate
transport in this organism (Kay, 1971; Schellenberg & Furlong, 1977) was found
not to be toxic to strain 3841 at financially viable concentrations. γ-
Glutamylhydrazide, an analogue of glutamine which strongly inhibits 5-amino
imidazole ribonucleotide synthetase (Schroeder et al., 1969), an enzyme of the
purine biosynthetic pathway, has been used at concentrations of 190-500µM to
select for glutamine transport mutants in E. coli (Weiner & Heppel, 1971;
Masters & Hong, 1981). Unfortunately, this analogue appears to be unstable in
acid minimal salts and is unable to survive in this media long enough to do
more than retard the growth of R. leguminosarum over the first 1-2 days, even
when added at concentrations in excess of 10mM. Some of the larger colonies
that grew on plates containing 10mM γ-glutamylhydrazide, which had been
inoculated with Tn5 mutants of strain 3841, were picked and the strains
purified. However, none of these strains exhibited a phenotype suggestive of
mutation of the general amino acid permease, and were not studied further.
186
Following the lack of success with toxic analogues, the possibility of using an
amino acid itself as the toxic agent was investigated. This type of approach has
been employed successfully in the isolation of dicarboxylate transport (dct)
mutants of R. leguminosarum by selecting for growth on 100mM succinate
(Glenn & Brewin, 1981).
The toxicity of the amino acids L-alanine, L-aspartate, L-glutamate and L-
serine towards R. leguminosarum strain 3841 was tested (Table 5.1).
Table 5.1 Growth of R. leguminosarum strain 3841 in the presence of increasing concentrations of four different amino acids. Growth medium was AMS agar with glucose/NH4Cl added at 10mM as the C/N source. +++, good growth; ++, moderate growth; +, poor growth; -, no growth.
Concentration Amino acid L-Alanine L-Aspartate L-Glutamate L-Serine
10mM +++ +++ +++ +++ 25mM + ++ +++ +++ 50mM - ++ +++ ++ 100mM - - ++ -
Although alanine shows the greatest toxicity, kinetic evidence suggests that
the general amino acid permease is not the only significant high affinity
transporter for this substrate in strain 3841 (Poole et al., 1985). Mutation of the
general amino acid permease is therefore less likely to allow escape from
alanine toxicity than it is to allow escape from a toxic level of a substrate for
which the general amino acid permease is the major transporter. Consequently,
it was decided to attempt to isolate amino acid permease mutants by selecting
for growth on 100mM aspartate.
Subsequent work in this laboratory (P.S.Poole, unpublished data) has shown
that at concentrations of 10mM or above, aspartate is transported at appreciable
187
rates by the dct system of R. leguminosarum. This suggests that mutation of an
aspartate-carrying amino acid transporter is unlikely to allow escape from
aspartate toxicity, and in fact strain RU543 exhibits the same susceptibility to
aspartate as the wild-type strain (Table 5.2).
Table 5.2 Growth of R. leguminosarum strains RU543 and 3841 in the presence of increasing concentrations of L-aspartate. Growth medium was AMS agar with glucose/NH4Cl added at 10mM as the C/N source. +++, good growth; ++, moderate growth; +, poor growth; -, no growth. Aspartate Strain concentration 3841 RU543 0mM +++ +++ 50mM ++ ++ 100mM - -
However, at the time, the use of aspartate as a toxic agent for selecting amino
acid permease mutants seemed reasonable. Tn5 mutants of strain 3841 were
grown on acid minimal salts agar containing glucose/NH4Cl as the
carbon/nitrogen source, 100mM aspartate, and kanamycin at 80 µg ml-1. The
increased concentration of kanamycin in this medium was found to be
necessary as 100mM aspartate appears to have an inhibitory effect on the action
of this antibiotic. Fifty colonies able to grow on this medium were purified and
the uptake of aspartate and glucose by the resulting strains was investigated.
The results for a representative sample are shown in Table 5.3.
The selection criteria used to isolate these mutants is not expected to yield
only mutants deficient in amino acid uptake. There will presumably be other
ways in which the cell can overcome the toxic effect of aspartate, such as having
reduced aspartate catabolism (if it is a product of this catabolism that causes
death) or increased metabolism (to use up excess aspartate and/or products of
188
its catabolism). Indeed, a proportion of the mutants in Table 5.3 show no
difference in aspartate uptake to the wild-type. Nevertheless, in many of the
mutants in Table 5.3 aspartate uptake is significantly impaired. However, in
every case where there is a reduction in aspartate transport there is a
concomitant, though not so severe, reduction in glucose transport. This is not
the phenotype expected to result from a mutation of the general amino acid
permease and suggests that these mutants are altered in a gene (or genes) with
a more global impact.
Table 5.3 Rates of aspartate and glucose transport in glucose/NH4Cl grown aspartate toxic escape Tn5 mutants of R. leguminosarum strain 3841. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the result from a single experiment except those for strains 3841, RU116 and RU156 which are the mean±SEM of determinations from three or more independent cultures. Strain Substrate L-Aspartate D-Glucose 3841 3.9±0.1 39.5±3.2 RU116 0.6±0.1 19.5±0.6 RU117 4.3 40.3 RU118 4.3 39.6 RU126 2.3 26.0 RU137 1.5 26.2 RU140 3.5 37.5 RU151 2.6 31.7 RU154 0.1 14.1 RU156 0.2±0.1 12.4±1.1 RU158 0.1 11.1
However, since the effect of the mutation on amino acid uptake in some of
these strains is so severe (in strain RU156 for example, aspartate transport is less
than 5% of that in the wild-type strain), and the effect appears to be particularly
marked for amino acid transport (glucose transport in strain RU156 is more
189
than 30% of that in strain 3841), it was decided that some of these mutants
warranted further investigation. As a representative sample of those mutants in
which amino acid uptake is impaired, strains RU116, RU137 and RU156 were
chosen for further study. Some properties of strain RU118, which overcomes
aspartate toxicity without a reduction in aspartate transport were also
investigated.
5.2.2 Growth of strains RU116, RU118, RU137 and RU156 on succinate and glucose
Since strains RU116 and RU156 were found to grow slowly on plates relative
to the wild-type, strains RU116, RU118, RU137 and RU156 were further
characterized by measuring their growth rates on glucose/NH4Cl and
succinate/NH4Cl (Table 5.4).
Table 5.4 Growth rates of aspartate toxic escape Tn5 mutants of R. leguminosarum strain 3841 on glucose/NH4Cl and succinate/NH4Cl. Growth rates are expressed as mean generation times in min. Values are the result from a single experiment. Carbon source Strain 3841 RU116 RU118 RU137 RU156 Glucose 270 405 230 255 990 Succinate 220 315 210 205 305
Growth of strains RU116 and RU156 on glucose is significantly slower than
that of the wild-type strain 3841. However, both mutants are partially rescued
by growth on succinate This suggests that mutations in strains RU116 and
RU156 are more deleterious to glucose catabolism than they are to that of
succinate.
190
The increased growth rate of strain RU118 on glucose relative to that of strain
3841 suggests that the mutation in this strain may result in an enhanced
metabolic rate under these conditions.
5.2.3 Amino acid transport in strains RU116 and RU156
In order to further characterize the effect of the mutation in each of RU116
and RU156 on amino acid transport, uptake of AIB, alanine and glutamate by
these strains grown on glucose/NH4Cl was investigated. Since a significant
difference is apparent in the growth of these strains on glucose compared to
that on succinate (Section 5.2.2), uptake of glutamate and aspartate was also
measured in cells grown on succinate/NH4Cl (Table 5.5).
191
Table 5.5 Rates of amino acid transport in R. leguminosarum strains RU116 and RU156 grown on glucose/NH4Cl and succinate/NH4Cl. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. ND, not determined. Growth conditions Substrate and strain L-Aspartate L-Glutamate L-Alanine AIB D-Glucose Succinate Glucose/NH4Cl 3841 3.9±0.1 5.6±0.4 6.4±0.7 4.3±0.6 39.5±3.2 ND RU116 0.6±0.1 0.8±0.3 2.0±0.2 1.2±0.1 19.5±0.6 ND RU156 0.2±0.1 0.3±0.1 1.2±0.1 0.1±0.0 12.4±1.1 ND Succinate/NH4Cl 3841 3.1±0.1 3.6±0.4 ND ND ND 46.7±0.2 RU116 1.3±0.1 1.4±0.4 ND ND ND 31.2±0.6 RU156 0.6±0.1 0.5±0.1 ND ND ND 22.7±3.9
192
The uptake of all the amino acids tested is significantly reduced in strains
RU116 and RU156. Since AIB transport is affected, it appears that this is not a
metabolic effect. Although the reductions in glutamate and aspartate transport
in RU116 and RU156 cells grown on succinate are less than those found in
glucose grown cultures of these strains, the impairment of amino acid uptake is
significantly greater than that of succinate in each case. Furthermore, the effect
on succinate uptake in the succinate grown mutants is less than that on glucose
uptake in the same strains grown on glucose. This suggests that the partial
rescuing of amino acid uptake observed in strains RU116 and RU156 grown on
succinate is the result of a global effect within the cell rather than a specific
effect on amino acid uptake.
5.2.4 Transductional analysis of strains RU116, RU118, RU137 and RU156
Mutations caused by the insertion of Tn5 are stable, and strongly polar (Berg,
1977). If a single copy of Tn5 is present in the genome of a mutant strain, and if
the insertion of this element is solely responsible for the mutant phenotype,
then the mutant phenotype is expected to be 100% cotransducible with the Tn5
kanamycin resistance marker (Beringer et al., 1978). This expectation applies
not only in cases where Tn5 inactivates a single structural gene, but also in
those cases where a single insertion of Tn5 in a polycistronic operon exerts a
polar effect on a larger number of genes.
Therefore, in order to confirm that the insertion of Tn5 was responsible for
the phenotype of the chosen aspartate toxic escape mutants (rather than a
concurrent spontaneous mutation), the kanamycin resistance marker of each of
strains RU116, RU118, RU137 and RU156 was transduced to strain 3841. The
purified transductants were tested for growth on 100mM aspartate and for the
presence of the high-level streptomycin resistance allele of strain 3841. This can
be distinguished from the low-level streptomycin resistance which is encoded
by Tn5 (Putnoky et al., 1983) by plating on medium containing streptomycin at
500 µg ml-1. Five transductants from each mutant strain were each found to be
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resistant to 100mM aspartate. In addition, aspartate and glucose uptake in the
transductants RU216, RU237 and RU256 from strains RU116, RU137 and
RU156, respectively, were measured. All were found to have transport rates for
these substrates similar to those of their respective transductional donor strains
(Table 5.6).
Table 5.6 Rates of aspartate and glucose transport in transductants from R. leguminosarum strains RU116, RU137 and RU156, grown on glucose/NH4Cl. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the result from a single experiment except those for strains 3841, RU116 and RU156 which are the mean±SEM of determinations from three or more independent cultures. ND, not determined. Strain Substrate L-Aspartate D-Glucose 3841 3.9±0.1 39.5±3.2 RU116 0.6±0.1 19.5±0.6 RU216 0.9 17.0 RU137 1.5 26.2 RU237 1.5 ND RU156 0.2±0.1 12.4±1.1 RU256 0.2 11.8
From these results it was concluded that the phenotypes of strains RU116,
RU137 and RU156 are each tightly linked to a single Tn5 insertion.
5.2.5 Nucleotide sequence adjacent to the transposon in strains RU116, RU137 and RU156
In an attempt to determine the identity of the gene(s) mutated in strains
RU116, RU137 and RU156 it was decided to sequence the chromosomal DNA
either side of the Tn5 insert in each case, and screen the resulting sequences
against sequence databases.
Chromosomal DNA from each strain was isolated, restricted with EcoRI and
the resulting restriction fragments randomly cloned into the EcoRI site of
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pBluescript SK+. In each case the resulting DNA was used to transform E. coli
strain MC1061, and KmR, AmpR transformants were selected. In this way
clones pRU32, pRU99 and pRU34, containing the approximately 10kb, 9kb and
10kb Tn5-bearing EcoRI fragment from the chromosome of strains RU116,
RU137 and RU156, respectively, were isolated.
Since a primer to the insertion sequence of Tn5 was to be used for sequencing
the DNA adjacent to the transposon, it was necessary to create sub-clones of
pRU32, pRU99 and pRU34 containing only one end of Tn5 (and hence only one
copy of the insertion sequence). Tn5 possesses internal BamHI and HindIII sites,
while in the pBluescript polylinker the BamHI and HindIII sites lie either side of
the EcoRI site. Since the DNA inserts in pRU32 and pRU34 contain either no
BamHI or HindIII sites other than those located within Tn5 (pRU34) or only one
additional BamHI site located between the transposon and the BamHI site in the
polylinker (pRU32), BamHI and HindIII digestion of these plasmids followed by
ligation yielded in each case, two sub-clones, each containing a different end of
Tn5 together with the adjacent chromosomal DNA (Fig. 5.1). The existence of
BamHI sites in the insert DNA between the transposon and the HindIII site in
the polylinker in pRU99 means that BamHI digestion and ligation will not yield
a sub-clone containing a Tn5 insertion sequence. In this case the required sub-
clone, pRU101, was obtained by randomly cloning BamHI restriction fragments
from pRU99 into pBC SK+, transforming E. coli strain DH5α with the resulting
DNA and selecting a KmR, CmR transformant. (This strategy makes use of the
fact that the required BamHI fragment carries the complete kanamycin
resistance gene of Tn5). Plasmid pRU100, the other sub-clone of pRU99 used
for sequencing, was provided by HindIII digestion and ligation.
195
Fig. 5.1 Creation of sub-clones of transposon clones from strains RU116 and RU156, for use in sequencing. Restriction sites are: B, BamHI; E, EcoRI; H, HindIII. *, Site present only in pRU32 and pRU37.
196
Sequencing pRU36/pRU37, pRU101/pRU100 and pRU40/pRU41 using a
primer (P0) to the insertion sequence of Tn5 provided the nucleotide sequence
either side of the transposon in strains RU116, RU137 and RU156, respectively
(Figs. 5.2-5.4).
The GenBank and EMBL databases were searched for sequences showing
homology to each of the six possible translations of each sequence. The
translation shown in Fig. 5.2 of the nucleotide sequence from strain RU116
shows 53.7%, 51.6% and 45.5% identity to SucD, the alpha subunit of succinyl-
CoA synthetase, from E. coli, Coxiella burnetti and Thermus aquaticus,
respectively. In addition, the translation shown in Fig. 5.4 of the nucleotide
sequence from strain RU156 shows 76.5%, 71.4%, 69.4%, 69.4% and 58.1%
identity to α-ketoglutarate dehydrogenase (encoded by sucA or an equivalent
gene) from E. coli, Bacillus subtilis, C. burnetti, Azotobacter vinelandii and Homo
sapiens, respectively. These data suggest that both RU116 and RU156 are TCA
cycle mutants.
The translation shown in Fig. 5.3 of the nucleotide sequence from strain
RU137 shows 60.9% identity to poly-beta-hydroxybutyrate synthase (encoded
by phbC) from R. meliloti. This suggests that biosynthesis of poly-beta-
hydroxybutyrate (PHB) is blocked in strain RU137.
197
198
199
200
5.2.6 Activity of TCA cycle enzymes in strains RU116, RU118, RU137 and RU156
In order to obtain experimental evidence for the nature of the mutation in
strains RU116 and RU156, the activities of a selection of TCA cycle enzymes in
these strains were assayed (Table 5.7). The activity of these enzymes in strains
RU118 and RU137 was also measured (Table 5.7).
Table 5.7 TCA cycle enzyme activities in glucose/NH4Cl grown aspartate toxic escape Tn5 mutants of R. leguminosarum strain 3841. Activities are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of three or more independent determinations, except the succinyl-CoA synthetase values for strains RU116 and RU156 which are the average of two independent determinations. αKDH, α-ketoglutarate dehydrogenase; SCS, succinyl-CoA synthetase; MDH, malate dehydrogenase; ICDH, isocitrate dehydrogenase; CS, citrate synthase; UD, undetectable; ND, not determined.
Enzyme Strain 3841 RU116 RU118 RU137 RU156 αKDH 109±10 6±2 86±3 42±12 UD SCS 55±21 20 ND ND 252 MDH 4978±833 18830±2524 3838±505 3893±1835 17435±1622 ICDH 864±164 904±94 886±147 890±61 957±66 CS 166±26 227±31 155±24 100±23 123±37
No α-ketoglutarate dehydrogenase activity can be detected in strain RU156,
while strain RU116 lacks succinyl-CoA synthetase activity. This is consistent
with mutation of sucA and sucD, respectively in these strains. The lack of α-
ketoglutarate dehydrogenase activity in strain RU116 is discussed elsewhere
(Section 5.2.14). Interestingly, malate dehydrogenase levels in strains RU116
and RU156 are greatly elevated in comparison to strain 3841. This effect is not
observed for citrate synthase or isocitrate dehydrogenase. One potential
explanation, which is discussed in Section 5.2.15, is that the increase in malate
dehydrogenase activity is a consequence of mdh lying upstream in the same
201
operon as sucA and sucD. The increased activity of succinyl-CoA synthetase in
strain RU156 can be explained similarly (Section 5.2.15).
The activities of α-ketoglutarate dehydrogenase, and to a lesser degree,
citrate synthase are reduced in strain RU137 in comparison to strain 3841, while
malate dehydrogenase and isocitrate dehydrogenase are unaffected. This
suggests that PHB biosynthesis may be involved in the regulation of certain
TCA cycle enzymes. This possibility is discussed elsewhere (Section 5.3).
The activities of all the enzymes assayed in strain RU118 are comparable to
those in strain 3841, suggesting that the mutation in RU118 is unrelated to those
in strains RU116, RU137 and RU156.
5.2.7 Growth of strains RU116, RU137 and RU156 on arabinose
If strains RU116 and RU156 are mutated in succinyl-CoA synthetase and α-
ketoglutarate dehydrogenase, respectively, it is expected that they will be
unable to grow on arabinose as the carbon source, since in R. leguminosarum the
catabolic pathway for this substrate (Fig. 5.5) enters the TCA cycle at α-
ketoglutarate (Dilworth et al., 1986). Consequently, these strains were tested for
growth on acid minimal salts containing arabinose and NH4Cl. Growth of
strain RU137 on this medium was also investigated. Neither strain RU116 nor
strain RU156 was able to grow, whereas strains 3841 and RU137 grew normally.
202
Fig. 5.5 Pathway of catabolism of L-arabinose in R. leguminosarum (Dilworth et al., 1986).
5.2.8 Complementation of strain RU156
Concomitant to the isolation and sequencing of the transposon clones of
strains RU116, RU137 and RU156, the mutation in strain RU156 was
complemented from a strain 3841 chromosomal library.
203
Strain RU156 was found to be unable to grow on glutamate as sole source of
carbon and nitrogen, presumably because glutamate uptake is insufficient
and/or α-ketoglutarate dehydrogenase is one of the principle catabolic
enzymes for glutamate. It was therefore possible to look for complementation
of the mutation in strain RU156 by testing for growth on glutamate. (Growth
on arabinose can be used to test for complementation of the mutation in RU156
(Section 5.2.15), however the inability of this strain to grow on arabinose as the
carbon source was not known when these experiments were performed).
A strain 3841 chromosomal library (as EcoRI fragments in pLAFR1) was
conjugated from E. coli strain 803 into strain RU156. Five transconjugants able
to grow on AMS agar containing glutamate as sole source of carbon and
nitrogen were isolated. Purified cosmids from these transconjugants were used
to transform E. coli strain S17-1 and thence reconjugated into strain RU156. This
was done in order to distinguish between Glu+ revertants and true
transconjugants. On reconjugation, four of the five cosmids were found to
complement strain RU156 for growth on glutamate. Since all four of these
cosmids show identical restriction patterns, only one, pRU3004, was employed
in further experiments.
5.2.9 Effect of pRU3004 on aspartate transport in strains RU116, RU137, and RU156
Cosmid pRU3004 was conjugated from E. coli strain S17-1 into RU116, RU137
and RU156, creating strains RU444, RU449 and RU453, respectively. Rates of
aspartate uptake in these strains indicate that pRU3004 complements strains
RU116 and RU156, but not strain RU137, for aspartate transport (Table 5.8).
204
Table 5.8 Rates of aspartate transport in R. leguminosarum strains RU444, RU449 and RU453 grown on glucose/NH4Cl. Rates of uptake are expressed as nmol min-1 [mg protein]-1. Values are the result from a single experiment except those for strains 3841, RU116 and RU156 which are the mean±SEM of determinations from three or more independent cultures. Strain Uptake 3841 3.9±0.1 RU116 0.6±0.1 RU444 2.7 RU137 1.5 RU449 1.6 RU156 0.2±0.1 RU453 3.5
5.2.10 Southern blot of pRU3004 against RU116, RU137, RU156 chromosomal DNA
Southern blotting of EcoRI digested chromosomal DNA from each of strains
3841, RU116, RU137 and RU156 with pRU3004, demonstrated that pRU3004
contains DNA homologous to the Tn5-bearing EcoRI fragment in both RU116
and RU156, but not to that in RU137.
Although the inserts in pRU32 and pRU34 both appear to be approximately
10kb in size, the fact that pRU32 contains a BamHI site which is absent from
pRU34 indicates that the transposon in strains RU116 and RU156 has inserted
into a different approximately 4kb EcoRI fragment in each case. Both these
fragments must be present in pRU3004 (because pRU3004 carries an EcoRI
fragment of the strain 3841 chromosome).
In other bacteria, sucA and sucD are clustered with genes encoding other
TCA cycle enzymes (Miles & Guest, 1987; Nicholls et al., 1990; Nishiyama et al.,
1991; Guest, 1992; Guest & Russell, 1992). It was therefore anticipated that
several R. leguminosarum TCA cycle genes might be carried by pRU3004, in
addition to sucA and sucD. Consequently, pRU3004 was subjected to further
205
study, in an attempt to establish whether the effect on amino acid transport in
strain 3841 was specific to mutation of α-ketoglutarate dehydrogenase and
succinyl-CoA synthetase, or whether the TCA cycle in general was implicated.
5.2.11 Restriction mapping, sub-cloning and mutation of pRU3004
A combination of sub-cloning and Southern blotting was used to produce the
restriction map of the 13.4kb of insert DNA from pRU3004 shown in Fig. 5.6.
Cosmid pRU3004 was subjected to saturation Tn5-lacZ mutagenesis. The
position of the transposon in mutated cosmids was determined by restriction
analysis. Cosmids in which the mutation is located within the mapped 13.4kb
region of pRU3004 are illustrated in Fig. 5.6.
Southern blotting of pRU3004 with pRU32 and pRU34 demonstrated that the
transposon in each of strains RU116 and RU156 lies within the region of the
strain 3841 chromosome corresponding to the mapped 13.4kb of pRU3004
DNA. The position of these mutations (determined by restriction analysis of
pRU32 and pRU34) is illustrated in Fig. 5.6.
206
Fig. 5.6 Map of pRU3004 and mutants derived from it, with sub-clones of mutants below. The locations of Tn5 and Tn5-lacZ insertions are flagged with the number of the cosmid and/or mutant strain in which they occur. Flags representing Tn5-lacZ insertions point in the direction of transcription of the lacZ gene in the transposon. Filled flags represent active fusions. Restriction sites are: B, BamHI; E, EcoRI; P, PstI; S, SalI; Ss, SstI.
207
5.2.12 Genes carried by pRU3004
The Tn5-B20-bearing salI fragments from pRU3059, pRU3061, pRU3069 and
pRU3070 were each cloned into the salI site of either pJQ200 or pSP72, creating
plasmids pRU194, pRU394, pRU395 and pRU396, respectively (Fig. 5.6).
Sequencing of the insert DNA in these plasmids using a primer (P15) to the 5'
end of Tn5-B20, and either Reverse, SK and KS primers to the pJQ200
polylinker, or T7 primer to the pSP72 polylinker, provided the nucleotide
sequences shown in Figs. 5.7-5.12. Sequencing of pRU41 using Reverse and SK
primers, provided the sequence shown in Fig. 5.13.
One potential translation of each of these sequences (Figs. 5.7-5.13) exhibits
significant homology to one of the TCA cycle enzymes α-ketoglutarate
dehydrogenase, succinyl-CoA synthetase or malate dehydrogenase from other
organisms as indicated in Table 5.9.
The locations of these homologies within the various genes (Table 5.9) are
consistent with the arrangement of Rhizobium genes proposed in Fig. 5.14.
208
1 ACGTAACAAGATCGCACTCATTGGTTCTGGCATGATTGGTGGCACGCTGGCGCACCTCGC 60
R N K I A L I G S G M I G G T L A H L A
61 CGGCCTGAAGGAACTGGGCGACATCGTTCTCTTCGACATCGCGGACGGCATTCCCCAGGG 120
G L K E L G D I V L F D I A D G I P Q G
121 CAAGGGTCTCGATATTTCCCAGTCGTCGCCGGTCGAAGGCTTCGACGTCAATCTGACGGG 180
K G L D I S Q S S P V E G F D V N L T G
181 CGCCAGCGACTATTCCGCGATCGAAGGCGCTGACGTCTGCATCGTCACGCGCGTCGCCCG 240
A S D Y S A I E G A D V C I V T R V A R
241 CAAGCCCGGCATGAGCCGCGATGACCTTCTCGGC 274
K P G M S R D D L L G
Fig. 5.7 Nucleotide sequence from pRU396 obtained using primer P15, and putative translation.
209
1 CCAAGTCGATCGACGAAGTCGTCGCCCATGCCAAGGAAATGCTGGGCAACACGCTGGTGA 60
K S I D E V V A H A K E M L G N T L V T
61 CGGNGCAGACCGGCGAAGCCGGCAAGCAGGTCAACCGCCTGTACATCGAAGACGGCGCCA 120
X Q T G E A G K Q V N R L Y I E D G A N
121 ACATCGCTCGCGAGCTCTATTGCTCGCTGCTGGTCGACCGTTCGGTCGGTCGCGTGGCTT 180
I A R E L Y C S L L V D R S V G R V A X
181 TNGTGGTCTCCACCGAAGGCGGCATGGACATCGAAGCTGTCGCCCACGACACGCCTGAGA 240
V V S T E G G M D I E A V A H D T P E K
241 AGATCCAGACGATCGCCATCGATCCGGAAGCCGGCGTGACGGCTGCCGACGTTGCTGCGA 300
I Q T I A I D P E A G V T A A D V A A I
301 TCTCCAAGGCTTTTGAGCTCGAGGGTGCTGCCGCCGAAGACGCCAAGACGCTTTTT 356
S K A F E L E G A A A E D A K T L F
Fig. 5.8 Combined nucleotide sequence from pRU396 and pRU194 obtained using primers T7 and Universal respectively, and putative translation.
210
1 ATCAAGCTCTACGGCAAGGAGCCGGCTAACTTCTGCGACGTCGGCGGTGGCNCCGGCAAG 60
I K L Y G K E P A N F C D V G G G X G K
61 GAGAAGGTTGCTGCGGCTTTCAAGATCATCACCGCTGTCCCCANGGNCGATGGCATTCTC 120
E K V A A A F K I I T A V P X X D G I L
121 GTCATCATCTTCGGCGGCATCATGAAGTGTCTGTGCAGGGCCTGTGGGCGCNTTGCTGCG 180
V I I F G G I M K C L C R A C G R X A A
181 GTCAAGGAAGTCGGTCTCAAGGTTCCGCTCGTCGTGCGCCTTGAAGGCACCAATGTCGAG 240
V K E V G L K V P L V V R L E G T N V E
241 CTCGGCAAGAAGATCCTGAACGAGTCGGGTNTGGCGATCACGGCGGCTGACGACTTGGAC 300
L G K K I L N E S G X A I T A A D D L D
301 GATGCGGCCAAGAAGATCGTCGCGGCGATCAACGGCTGAGAATGATCATGG 351
D A A K K I V A A I N G * E * S W
Fig. 5.9 Nucleotide sequence from pRU194 obtained using primer P15, and putative translation.
211
1 ACGGCATCTGCGCCTTCCGCCCGGCCTCGACAACACGAGACCAGNAGATGCGCNGCGGAC 60
G I C A F R P A S T T R D Q X M R X G
61 AGGCGGCAATCGGAATGCGGCAGCCGGCCATCAAGCCGGTAAATTCATCAAAGTCAGGAG 120
Q A A I G M R Q P A I K P V N S S K S G
121 GCGGACGGAAGCGTCCGCATAACACCATGGCACGGCAAGAAGCCAACGAGCAGTTTCAGA 180
G G R K R P H N T M A R Q E A N E Q F Q
181 TCACCTCGTTTCTGGATGGCGCCAACGCTGCCTATATCGAGCAGCTCTACGCGC 234
I T S F L D G A N A A Y I E Q L Y A
Fig. 5.10 Nucleotide sequence from pRU194 obtained using primers SK and Reverse, and putative translation.
212
213
1 GCCTNGNGNGAAGAGNGNGTGAAGATGANNTGCCTTGCNAGACGATNGCNAAGNGNCCTC 60
A X X E E X V K M X C L A R R X X X X L
61 AAGGATGTGCAGAACACNGCCGCCATGCTGACCACCTACAATGAGGTGGACATGAAGGCG 120
K D V Q N T A A M L T T Y N E V D M K A
Fig. 5.12 Nucleotide sequence from pRU395 obtained using primer P15, and putative translation.
1 AATTCCGCATGAAGTTCCACAAGCCTGTTGTGCTCGACCTGTTCTGCTACCGTCGCTACG 60
F R M K F H K P V V L D L F C Y R R Y
61 GCCACAATGAAGGCGACGAACCGTCCTTCACGCAGCCGAAGATGTACAAGGTGATCCGCG 120
G H N E G D E P S F T Q P K M Y K V I R
121 CCCACAAGACCGT 133
A H K T
Fig. 5.13 Nucleotide sequence from pRU41 obtained using primers SK and Reverse, and putative translation.
214
Table 5.9 Homology of deduced polypeptides from pRU3004 to known sequences of TCA cycle enzymes. The number in the left hand column refers to the corresponding shaded area in Fig. 5.14. mdh encodes malate dehydrogenase; sucC and scsB encode the beta subunit of succinyl-CoA synthetase; sucD encodes the alpha subunit of succinyl-CoA synthetase; sucA encodes α-ketoglutarate dehydrogenase; sucB encodes dihydrolipoamide succinyltransferase. Number Sequence Identities 1 Fig. 5.7 40.4% over amino acids 1 to 92 of Mdh in Photobacterium
36.2% over amino acids 1 to 92 of Mdh in E.coli 2 Fig. 5.8 38.1% over amino acids 63 to 180 of SucC in E.coli
44.6% over amino acids 64 to 151 of SucC in C. burnetti 36.3% over amino acids 81 to 171 of ScsB in T. aquaticus
3 Fig. 5.9 54.3% over amino acids 275 to 389 of SucC in E. coli
55.5% over amino acids 273 to 382 of SucC in C. burnetti 38.9% over amino acids 270 to 379 of ScsB in T. aquaticus
4 Fig. 5.2 53.7% over amino acids 223 to 289 of SucD in E. coli
51.6% over amino acids 223 to 286 of SucD in C. burnetti 45.5% over amino acids 222 to 287 of SucD in T. aquaticus
5 Fig. 5.10 57.1% over amino acids 7 to 28 of SucA in C. burnetti 6 Fig. 5.11 62.8% over amino acids 438 to 480 of SucA in C. burnetii
60.5% over amino acids 445 to 487 of SucA in A. vinelandii 58.1% over amino acids 439 to 481 of SucA in E. coli 55.8% over amino acids 443 to 485 of SucA in B. subtilis
7 Fig. 5.12 68.1% over amino acids 679 to 797 of SucA in E. coli
63.7% over amino acids 688 to 810 of SucA in A. vinelandii 63.5% over amino acids 681 to 795 of SucA in C. burnetti 60.3% over amino acids 681 to 803 of SucA in B. subtilis
8 Fig. 5.13 50.0% over amino acids 187 to 222 of SucB in B. subtilis
45.0% over amino acids 172 to 211 of SucB in E. coli 43.2% over amino acids 168 to 204 of SucB in A. vinelandii
215
Fig. 5.14 Putative arrangement of genes in mapped region of pRU3004. Gene locations and orientations are based on homologies of sequenced regions (shaded) to genes in other bacteria as detailed in Table 5.9. Numbering of shaded areas refers to Table 5.9.
216
5.2.13 β-galactosidase activities from pRU3004 mutants
As experimental confirmation of the proposed directions of transcription for
mdh and sucCDAB, β-galactosidase activities produced in a strain 3841
background by corresponding pRU3004 mutants were investigated. Those
cosmids in which lacZ in the transposon is aligned with the proposed direction
of transcription for the corresponding gene show significantly higher activity
than those in which lacZ is in the opposite orientation (Fig. 5.6).
5.2.14 TCA cycle enzyme activities in sucCDAB mutants of strain 3841
In order to investigate directly the nature of the partially sequenced genes
carried by pRU3004, activities of the corresponding TCA cycle enzymes were
assayed in chromosomal mutants of these genes.
The Tn5-lacZ mutants of strain 3841, RU725, RU733, RU724 and RU726 were
generated by homogenotization (Ruvkun & Ausubel, 1981) of cosmids
pRU3059, pRU3061, pRU3067 and pRU3069, respectively. No homogenotes of
cosmids pRU3070 or pRU3076 could be isolated, presumably because mutation
of mdh is lethal.
α-Ketoglutarate dehydrogenase and succinyl-CoA synthetase levels in the
mutant strains were measured (Table 5.10).
Table 5.10 TCA cycle enzyme activities in R. leguminosarum strains RU724, RU725, RU726 and RU733 grown on glucose/NH4Cl. Activities are expressed as nmol min-1 (mg protein)-1. Values are the mean of two independent determinations, except the 3841 values which are the mean±SEM of four independent determinations. αKDH, α-ketoglutarate dehydrogenase; SCS, succinyl-CoA synthetase; UD, undetectable; ND, not determined.
Enzyme Strain 3841 RU724 RU725 RU726 RU733 αKDH 109±10 UD UD UD UD SCS 55±21 ND 22 98 ND
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The loss of α-ketoglutarate dehydrogenase activity in the sucA (RU724,
RU733) and sucB (RU726) mutants is expected, as is the reduction in succinyl-
CoA synthetase activity in the sucC mutant (RU725). The observation that
strain RU725 exhibits negligible α-ketoglutarate dehydrogenase activity, as
does strain RU116 (Section 5.2.6), can be understood if sucAB are under the
control of the same promoter as sucCD, or if high levels of succinyl-CoA,
accumulated as a result of the loss of succinyl-CoA synthetase, cause feed-back
inhibition of α-ketoglutarate dehydrogenase (Williamson & Cooper, 1980). In
order to distinguish between these two possibilities, the location of promoters
in the mapped 13.4kb region of pRU3004 was investigated (Section 5.2.15).
The elevated succinyl-CoA synthetase level in strain RU726 is discussed
elsewhere (Section 5.2.15).
5.2.15 Mapping of promoter sites in pRU3004
Promoter activity in the region of the mdh and sucCDAB genes of R.
leguminosarum strain 3841 was investigated by testing the ability of various sub-
clones of pRU3004 to complement chromosomal mutations in these genes.
None of the strains RU725, RU733, RU724 and RU726 is able to grow on
arabinose as the carbon source, a phenotype consistent with mutation of α-
ketoglutarate dehydrogenase or succinyl-CoA synthetase (Section 5.2.7). It was
therefore possible to use growth on arabinose as a test for complementation of
the mutation in these strains.
A selection of EcoRI, PstI and SstI fragments of pRU3004 were cloned in the
corresponding site in pRK415-1, creating plasmids pRU180, pRU181, pRU182,
pRU276, pRU277, pRU278, pRU397 and pRU398 (Fig. 5.15). Each of these
plasmids was conjugated into RU116, RU156, RU724, RU725, RU276 and RU733
and the resulting strains tested for growth on arabinose (Table 5.11).
218
Fig 5.15 Sub-clones of pRU3004 used in complementation analysis. Two sub-clones of pRU3068 are also shown. The shaded arrows indicate the direction of transcription initiation from the lac promoter in the vector of each sub-clone. Restriction sites are: E, EcoRI; P, PstI; Ss, SstI.
219
Table 5.11 Growth of sucCDAB mutants of R. leguminosarum strain 3841, containing sub-clones of pRU3004, on arabinose/NH4Cl. Growth medium was AMS agar with the C/N source added at 10mM. +, good growth; -, no growth; ND, not determined. Plasmid Strain RU116 RU156 RU724 RU725 RU726 RU733 None - - - - - - pRU277 - ND ND - ND ND pRU278 - ND ND - ND ND pRU181 - ND ND - ND ND pRU180 ND - - ND - - pRU276 ND - - ND + - pRU398 ND - - ND - - pRU397 ND - - ND + - pRU182 ND ND ND ND - ND
Transcription of the strain 3841 genes carried by pRU181, pRU276, pRU277
and pRU397 can be initiated by the lac promoter in the vector. Therefore the
observation that mutations in strains RU725 and RU116 are not complemented
by pRU277, which carries sucCD, indicates that sucCDA are under the control of
a single promoter upstream of sucC (because a chromosomal insertion in either
sucC or sucD appears to prevent transcription of at least one gene downstream
of sucD). This result is consistent with the observation that strains RU116 and
RU725 exhibit negligible α-ketoglutarate dehydrogenase activity (Sections 5.2.6
and 5.2.14).
The absence of a promoter between the transposon in pRU3068 and the PstI
site upstream of it was confirmed by cloning, in both orientations in pIJ1891, the
lacZ-bearing PstI fragment from pRU3068, and measuring the β-galactosidase
activity produced by the resulting clones in a strain 3841 background. In
pRU386 (Fig. 5.15) transcription of the lacZ fusion can be initiated by the lac
promoter in the vector, and β-galactosidase activity in strain 3841 containing
this plasmid is high (1336±211 nmol min-1 [mg protein]-1). In pRU387 (Fig. 5.15)
220
the direction of transcription of the insert DNA is opposite to the lac promoter
in the vector, so the low level of β-galactosidase activity (95±4 nmol min-1 [mg
protein]-1) observed in strain 3841 containing this plasmid is indicative of a lack
of native promoter activity in the insert. (Cosmid pRU3068, which contains the
same replicon as pRU386 and pRU387, produces β-galactosidase activity of 3119
±195 nmol min-1 [mg protein]-1 in strain 3841).
Since the mutation in strain RU726 is not complemented by pRU398, either
there is no promoter in the intergenic region between sucA and sucB, or a sucB
mutation is polar on a downstream gene required for growth on arabinose.
However, since the mutation in strain RU726 is complemented by pRU276, it
can be concluded that pRU398 does not complement strain RU726 because sucB
lacks a promoter of its own. The fact that pRU397 complements the mutation in
RU726 demonstrates that the gene mutated in this strain lies within the 2.4kb
EcoRI-PstI fragment containing the transposon.
It is not possible to deduce from these complementation data whether the
promoter controlling sucCDAB lies in the intergenic region between mdh and
sucC, or upstream of mdh. However, the latter arrangement might provide an
explanation for the increased malate dehydrogenase activity observed in sucDA
mutants (RU116 and RU156; Table 5.7), and the elevated levels of succinyl-CoA
synthetase in sucAB mutants (RU156 and RU726; Table 5.7 and 5.10). A
mutation in sucCDAB may lead to an increase in the expression of the mutated
sucCDAB in response to a change in the intracellular concentration of one or
more metabolites, such as α-ketoglutarate, in the mutant. If this is the case, then
the location of mdh upstream of sucCDAB, and under the control of the same
promoter, would result in the increased expression of mdh in a sucCDAB
mutant. Similarly, mutation of sucAB would lead to increased sucCD
expression.
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5.2.16 Amino acid excretion by strains RU116, RU156 and RU543
If, as has been suggested (Section 1.2.5), it is inhibition and/or repression of
α-ketoglutarate dehydrogenase that results in the excretion of amino acids by
bacteroids, it might be expected that mutants of R. leguminosarum lacking α-
ketoglutarate dehydrogenase activity will also excrete amino acids.
Consequently, strains RU116 and RU156 were tested for amino acid excretion.
Supernatants from cultures growing on L-malate/NH4Cl were assayed over
time for alanine, aspartate, glutamate and α-ketoglutarate (Table 5.12). L-
malate was provided as the carbon source because this is the most abundant C4-
dicarboxylate in the nodule (Streeter, 1987), and consequently the most likely
carbon source for the bacteroid (Section 1.2.1).
Although no excretion of alanine or aspartate is apparent, significant
concentrations of glutamate and α-ketoglutarate were detected in the
supernatants from cultures of both strain RU116 and strain RU156. This is in
contrast to the wild-type, for which no excretion of any of the four metabolites
assayed was detected.
The excretion of certain metabolites, including α-ketoglutarate, by bacteria
under particular metabolic conditions is well documented (Tempest & Neijssel,
1992; Kramer, 1994), and the excretion of α-ketoglutarate by R. leguminosarum
strains containing mutations that block the major catabolic pathway for this
substrate is not surprising. However, the excretion of glutamate by strains
RU116 and RU156 suggests that a proportion of the excess α-ketoglutarate is
converted to glutamate.
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Table 5.12 Excretion of metabolites by R. leguminosarum strains RU116 and RU156 grown on L-malate/NH4Cl. Cells grown on L-malate/NH4Cl were washed and resuspended in AMS containing 10mM L-malate/NH4Cl and incubated at 28°C. After the time intervals shown, the concentrations of each of the four metabolites in samples of supernatants was determined. At the 0 min and 150 min time points, samples of culture were taken and assayed for protein content. This enabled an approximate calculation of the rate of excretion, in nmol min-1 [mg cell dry weight]-1, assuming 0.2 [mg protein] ml-1 ≡ 0.4 mg dry weight of cells (P.S.Poole, unpublished data). Values are the result from a single representative experiment, with the concentration at different times given in µM. Metabolite Time (min) Excretion and strain 0 60 150 270 rate L-Alanine 3841 <10 <10 <10 <10 <1 RU116 <10 <10 <10 <10 <1 RU156 <10 <10 <10 <10 <1 L-Aspartate 3841 <10 <10 <10 <10 <1 RU116 <10 <10 <10 <10 <1 RU156 <10 <10 <10 <10 <1 L-Glutamate 3841 <10 <10 <10 <10 <1 RU116 <10 81 158 477 15 RU156 <10 52 98 138 11 α-Ketoglutarate 3841 <10 <10 <10 <10 <1 RU116 <10 218 805 >1000 79 RU156 <10 404 >1000 >1000 >105
Since aspartate is a potential amino-donor in the production of glutamate
from α-ketoglutarate, it was thought possible that the escape of strains RU116
and RU156 from aspartate toxicity might be due to transamination of aspartate
to glutamate, and subsequent excretion of the glutamate. To investigate this
possibility, excretion of glutamate by strains 3841, RU116 and RU156 grown on
both glucose/NH4Cl and glucose/NH4Cl/aspartate was measured. For the
purpose of comparison to the results from cultures containing malate as the
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carbon source, supernatants from the glucose/NH4Cl cultures were also
assayed for α-ketoglutarate (Table 5.13).
Table 5.13 Excretion of glutamate and α-ketoglutarate by R. leguminosarum strains 3841, RU116, RU156 and RU543 grown on glucose/NH4Cl and glucose/NH4Cl/aspartate. Cells grown on glucose/NH4Cl or glucose/NH4Cl/(20mM)aspartate were washed and resuspended in AMS containing 10mM glucose/NH4Cl or glucose/NH4Cl/(20mM)aspartate respectively, and incubated at 28°C. After the time intervals shown, the concentrations of the α-ketoglutarate and/or glutamate in samples of supernatants was determined. At the 0 min and 150 min time points, samples of culture were taken and assayed for protein content. This enabled an approximate calculation of the rate of excretion, in nmol min-1 [mg cell dry weight]-
1, assuming 0.2 [mg protein] ml-1 ≡ 0.4 mg dry weight of cells (P.S.Poole, unpublished data). Values are the result from a single representative experiment, with the concentration at different times given in µM. Glc, glucose; NH4, NH4Cl; Asp, aspartate. Metabolite Time (min) Excretion and culture 0 60 150 270 rate L-Glutamate 3841/Glc/NH4 <10 <10 <10 <10 <1 RU116/Glc/NH4 <10 46 179 333 18 RU156/Glc/NH4 <10 <10 65 120 8 RU543/Glc/NH4 <10 <10 <10 <10 <1 3841/Glc/NH4/Asp <10 34 106 225 10 RU116/Glc/NH4/Asp <10 195 602 >1000 59 RU156/Glc/NH4/Asp <10 170 443 863 49 RU543/Glc/NH4/Asp <10 <10 18 55 2 α-Ketoglutarate 3841/Glc/NH4 <10 <10 <10 <10 <1 RU116/Glc/NH4 <10 38 49 55 5 RU156/Glc/NH4 <10 95 195 311 24
Excretion of glutamate by strains RU116 and RU156 is substantially increased
by the presence of aspartate in the medium, consistent with increased
transamination of α-ketoglutarate. This result supports the notion that these
strains are able to survive high levels of aspartate because the increased
availability of intracellular α-ketoglutarate enables the toxic aspartate to be
224
removed by transamination. The increased rate of glutamate excretion by
strains RU116 and RU156 grown on glucose/NH4Cl/aspartate appears to be
greater than can be accounted for by the rate of α-ketoglutarate excretion by
these strains grown on glucose/NH4Cl, given the small apparent size of the
intracellular pool of α-ketoglutarate (Section 5.2.17). However, metabolism, via
the TCA cycle, of oxaloacetate generated from the transamination of α-
ketoglutarate by aspartate, will presumably lead to the generation of additional
α-ketoglutarate in cells grown on glucose/NH4Cl/aspartate. The rate of
excretion of α-ketoglutarate by strains RU116 and RU156 grown on malate is
significantly greater than that found when glucose is the carbon source. This is
consistent with greater carbon flux through the TCA cycle during growth on the
TCA cycle intermediate, malate.
Interestingly, the presence of aspartate in the medium results in excretion of
glutamate, but not α-ketoglutarate, by strain 3841. It seems likely that the
excreted glutamate is derived from α-ketoglutarate, and its synthesis is
presumably the result of an increased supply of amino-donor (aspartate) for the
transamination of α-ketoglutarate, and/or increased production (via the TCA
cycle) of α-ketoglutarate from oxaloacetate generated from aspartate by
transamination. It is also possible that the presence of aspartate in the medium
allows intracellular glutamate to be exchanged out of the cell (Section 3.2.18).
The observation that α-ketoglutarate is not excreted by 3841 grown on
glucose/NH4Cl/aspartate suggests that, in this case, the rate of glutamate
synthesis equals the rate of α-ketoglutarate production. In strains RU116 and
RU156 grown on glucose/NH4Cl and malate/NH4Cl, the rate of α-
ketoglutarate generation presumably exceeds the maximum rate of
transamination to glutamate, and consequently α-ketoglutarate is excreted. The
fact that, while the rates of α-ketoglutarate excretion by these strains differ
substantially between growth on malate and growth on glucose, the rates of
225
glutamate excretion are very similar for both carbon sources, is consistent with
this suggestion.
Since strain 3841 excretes glutamate when grown on
glucose/NH4Cl/aspartate, it was possible to investigate the role of the general
amino acid permease in the excretion of glutamate, by assaying glutamate
excretion by strain RU543 (an aapJ mutant) grown on glucose/NH4Cl/aspartate
(Table 5.13).
Glutamate excretion by strain RU543 is significantly reduced in comparison
to strain 3841. It is unlikely that this result is a consequence of an effect on
aspartate uptake in strain RU543, since aspartate uptake in R. leguminosarum is
mediated by the dct system at the concentration of aspartate employed in this
experiment (Section 5.2.1). Strain RU543 grows more slowly on
glucose/NH4Cl/aspartate than strain 3841 (mean generation times are 1095 min
and 660 min, respectively), and a proportion of the reduction in glutamate
excretion observed in RU543 may be attributable to the reduced metabolic rate
in this strain. However, a potential explanation of the poor growth of RU543 on
glucose/NH4Cl/aspartate, is that this strain can not easily excrete the glutamate
it synthesizes under these conditions. These data therefore appear to indicate
the involvement of the general amino acid permease in the excretion of
glutamate from strain 3841 grown on glucose/NH4Cl/aspartate. In view of the
results of amino acid exchange experiments on strain 3841 (Section 3.2.18), and
the presence of aspartate in the medium, glutamate may be exported via an
exchange mechanism.
The protocol used in this research to prepare cells for transport assays
involves the final resuspension of cells in minimal salts medium lacking carbon
and nitrogen sources, and incubation for between one and six hours prior to the
assay(s). It therefore seemed possible (though unlikely, given the lack of carbon
and nitrogen sources) that the low amino acid transport values obtained for
strains RU116 and RU156 might be the result of competitive inhibition of the
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general permease by glutamate excreted from these strains into the medium
during the incubation period before the assays. However, no glutamate
excretion could be detected from cells of strains RU116 and RU156 resuspended
and incubated for five hours in minimal salts containing no carbon or nitrogen
source (data not shown). In addition, glutamate transport values for cells of
either strain RU116 or strain RU156, spun down and resuspended in fresh salts
immediately prior to assaying, were found to be identical to those of cells
treated in the usual way before being assayed (data not shown). It is evident
therefore, that the impaired amino acid transport in strains RU116 and RU156 is
not due to the excretion of glutamate by these strains.
5.2.17 Intracellular concentrations of α-ketoglutarate and glutamate in strains RU116 and RU156
The fact that α-ketoglutarate and glutamate are excreted by strains RU116
and RU156 suggests that the intracellular concentrations of these substrates
may be elevated in these mutants. It seems improbable that an elevated
intracellular concentration of any metabolite will be maintained during an
incubation without carbon or nitrogen of up to six hours, so it is unlikely that
direct inhibition of the general amino acid permease by, for example,
intracellular glutamate, is the reason for the reduced amino acid uptake in
sucDA mutants of strain 3841. However, an elevated level of intracellular
glutamate or α-ketoglutarate may be the signal that initiates down-regulation of
the amino acid permease.
It was therefore decided to ascertain whether intracellular glutamate or α-
ketoglutarate concentrations are in fact elevated in strains RU116 and RU156.
The assays were carried out on cells grown on glucose/NH4Cl (Table 5.14).
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Table 5.14 Intracellular concentrations of glutamate and α-ketoglutarate in R. leguminosarum strains 3841, RU116 and RU156 grown on glucose/NH4Cl. Concentrations are calculated assuming: 1g dry weight of cells ≡ 1.45ml (Dilworth & Glenn, 1982); 0.2 [mg protein] ml-1 ≡ 0.4 mg dry weight of cells (P.S.Poole, unpublished data); the periplasmic space constitutes 20% total cell volume (Dilworth & Glenn, 1982). Values are the result from a single experiment and are given in mM. Metabolite Strain 3841 RU116 RU156 L-Glutamate 4 12 52 α-Ketoglutarate <1 <1 <1
Since glutamine is likely to have been hydrolysed to glutamate during the
preparation of the samples in which intracellular substrate concentrations were
assayed (Section 2.1.10), the figures given in Table 5.14 strictly represent the
combined intracellular concentrations of glutamate and glutamine. However,
since glutamate is known to be excreted by strains RU116 and RU156 under the
growth conditions employed in this experiment, it is probable that a significant
proportion of the elevated figure for intracellular glutamate in each of these
strains (Table 5.14) is attributable to glutamate rather than glutamine. If this is
the case, the involvement of the intracellular glutamate level in the regulation of
the general amino acid permease seems likely.
Very little intracellular α-ketoglutarate was detected for all three strains
RU116, RU156 and 3841. This seems unlikely to be an experimental artefact
since α-ketoglutarate was shown to be stable under the conditions used to
prepare samples (Section 2.1.10), and the results for intracellular glutamate are
as anticipated. It therefore appears that the intracellular pool of α-ketoglutarate
in the wild-type is low under these conditions, and this is reflected in the ready
excretion of this metabolite by the mutants.
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5.2.18 Transcription of the aap operon in sucDA mutants of strain 3841
In order to determine whether transcriptional regulation mediates the
reduction in amino acid transport observed in succinyl-CoA synthetase and α-
ketoglutarate dehydrogenase mutants of strain 3841, the activity of an aapJ::lacZ
transcriptional fusion in strain RU116 and RU156 backgrounds was
investigated.
Cosmid pRU3028, a Tn5-lacZ mutant of pRU3024 in which the transposon
has inserted in aapJ such that lacZ is in the same orientation as the mutated gene
(Fig. 3.2), was conjugated into strains RU116 and RU156. β-Galactosidase
activity in the resulting strains, RU889 and RU897, respectively, was measured
under both nitrogen-excess and nitrogen-limited conditions (Table 5.15). The β-
galactosidase activity produced by pRU3024 in strain RU116 and RU156
backgrounds (strains RU1024 and RU1025, respectively) was also measured as a
control.
β-Galactosidase activities in strains RU889 and RU897 are generally very
similar to those in strain RU443 (3841/pRU3028), although activity in strain
RU897 grown on glucose/glutamate is somewhat reduced, perhaps because
this strain grows poorly under these conditions. Certainly, the activity of the
fusion in strains RU889 and RU897 does not indicate a decrease in the
transcription of the aap operon sufficient to account for the severe reduction in
amino acid transport observed in strains RU116 and RU156 grown on
glucose/NH4Cl (Section 5.2.3).
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Table 5.15 β-galactosidase activities in R. leguminosarum strains RU116 and RU156 containing pRU3028 and pAR36A. Strains were grown on AMS containing either glucose/NH4Cl or glucose/glutamate at 10mM. Activities are expressed as nmol min-1 [mg protein]-1. Values are the mean±SEM of determinations from three or more independent cultures. Strain Relevant genotype Growth conditions Host Cosmid/plasmid Glucose/NH4Cl Glucose/glutamate RU438 (3841/pRU3024) Wild-type aapJQMP 40±5 43±1 RU1024 (RU116/pRU3024) sucD::Tn5 aapJQMP 32±2 40±3 RU1025 (RU156/pRU3024) sucA::Tn5 aapJQMP 41±4 34±4 RU443 (3841/pRU3028) Wild-type aapJ::Tn5-lacZ 3325±279 7923±666 RU889 (RU116/pRU3028) sucD::Tn5 aapJ::Tn5-lacZ 3225±209 6484±118 RU897 (RU156/pRU3028) sucA::Tn5 aapJ::Tn5-lacZ 3853±234 4726±259 RU368 (3841/pMP220) Wild-type Promoterless lacZ 62±3 105±1 RU1002 (RU116/pMP220) sucD::Tn5 Promoterless lacZ 111±21 50±7 RU1003 (RU156/pMP220) sucA::Tn5 Promoterless lacZ 37±11 40±1 RU622 (3841/pAR36A) Wild-type glnII:lacZ 101±14 531±79 RU891 (RU116/pAR36A) sucD::Tn5 glnII:lacZ 270±6 879±66 RU899 (RU156/pAR36A) sucA::Tn5 glnII:lacZ 53±4 513±42
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It is interesting that, like the wild-type strain, both mutants exhibit repression
of the aap operon when grown on NH4Cl as the nitrogen source. This result is
consistent with the finding that in strains RU116 and RU156, as in strain 3841,
the intracellular α-ketoglutarate concentration is low (Section 5.2.17). Strains
lacking α-ketoglutarate dehydrogenase activity might have been expected to
have an elevated intracellular concentration of α-ketoglutarate and hence an
elevated α-ketoglutarate to glutamine ratio, resulting in a phenotype
compatible with apparent nitrogen-limitation (Section 1.6), even under
conditions of nitrogen-excess. This is apparently not the case.
As confirmation of this observation β-galactosidase activities in strains
RU116 and RU156 carrying the R. leguminosarum glnII::lacZ reporter fusion
pAR36A (Patriarca et al., 1992) were measured (Table 5.15). pAR36A activity in
the RU116 and RU156 backgrounds (strains RU891 and RU899, respectively)
responds to a change from nitrogen-limited to nitrogen-excess conditions as it
does in the 3841 background (strain RU622).
While it is apparent that the aap operon is not affected at the transcriptional
level by a mutation in sucDA, inhibition of the transporter by intracellular
glutamate (or indeed any substrate) also appears to be unlikely (Section 5.2.16).
It seems reasonable to propose that mutation of α-ketoglutarate dehydrogenase
or succinyl-CoA synthetase may lead to post-translational modification of the
amino acid permease, and it is plausible that this modification is initiated by an
increase in intracellular glutamate. Post-translational regulation, in the form of
phosphorylation of the binding protein, has previously been reported for the
lysine-arginine-ornithine uptake system of E. coli (Celis, 1984; Celis, 1990).
5.2.19 Plant properties of strains RU116, RU137 and RU156
Although both strain RU116 and strain RU156 nodulate peas, the nodules are
extremely small and green. No kanamycin resistant bacteria could be recovered
from nodules produced by these strains, and acetylene reduction was not
tested. These results are consistent with an earlier report that an α-
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ketoglutarate dehydrogenase mutant of R. meliloti formed ineffective nodules
(Duncan & Fraenkel, 1979).
The pea nodules produced by strain RU137 appeared on average to be larger
than those produced by strain 3841, however acetylene reduction by RU137
nodules was not investigated. All 25 nodule isolates of this strain were found to
have retained kanamycin resistance. Bean nodules induced by a PHB synthase
mutant of Rhizobium etli have been found to exhibit increased nitrogenase
activity (M.A.Cevallos, personal communication). In contrast, PHB
biosynthesis mutants of R. meliloti have been reported to have similar symbiotic
traits to their parental strain (Povolo et al., 1994).
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5.3 DISCUSSION
Mutation of the TCA cycle enzymes α-ketoglutarate dehydrogenase and
succinyl-CoA synthetase in R. leguminosarum strain 3841 leads to a severe
reduction in amino acid uptake by the mutants. Uptake of other substrates
such as glucose or succinate is also impaired in such mutants, probably as a
result of general dehabilitation produced by the disruption of central
metabolism, however the particularly marked reduction in amino acid transport
suggests that there is a specific effect on this system. This is in accordance with
the suggestion that it is the co-ordinated operation of the TCA cycle and amino
acid transport that regulates the flow of carbon and energy through the
bacteroid (Section 3.1).
The lacZ fusion data indicate that there is no change in the transcription
levels of the aap operon in sucDA mutants, and the conditions encountered by
cells prior to transport assays effectively eliminate the possibility of intracellular
substrate inhibition of the general amino acid permease. This suggests that the
general amino acid permease may be subject to post-translational modification.
The intracellular glutamate concentration was found to be elevated in sucDA
mutants grown on glucose/NH4Cl, and it is possible that this elevation is the
signal for the down-regulation of the general amino acid permease. In addition,
glutamate is excreted by sucDA mutants grown on both glucose/NH4Cl and
malate/NH4Cl. These results are consistent with several studies which indicate
that a significant proportion of malate supplied to bacteroids under nitrogen-
fixing conditions, is converted to glutamate (Salminen & Streeter, 1987a; Kouchi
et al., 1991; Miller et al., 1991; Salminen & Streeter, 1992). Such studies have led
to the suggestion that glutamate synthesis in the bacteroid is a consequence of
inhibition of α-ketoglutarate dehydrogenase by NADH (McDermott et al., 1989;
Salminen & Streeter, 1990). Although complete loss of α-ketoglutarate
dehydrogenase activity is artificial, the accumulation of glutamate by free-living
sucDA mutants appears to support this hypothesis.
233
Although no excretion of glutamate by bacteroids has been reported, such
excretion does add credence to the suggestion that inhibition and/or repression
of central metabolic enzymes in the bacteroid may lead to excretion of amino
acids (McDermott et al., 1989; Rosendahl et al., 1992). Indeed, inhibition of
pyruvate dehydrogenase, an enzyme with strong structural and functional
similarities to α-ketoglutarate dehydrogenase (Miles & Guest, 1987; Guest &
Russell, 1992), would lead to the accumulation of pyruvate. This might explain
the observed excretion of alanine by bacteroids: Transamination of pyruvate
followed by excretion of the resulting alanine, could relieve pyruvate
accumulation. This hypothesis is supported by the observation that alanine
dehydrogenase and alanine aminotransferase activity has been detected in
bacteroids but not free-living cells of R. leguminosarum strain 3841 (P.S.Poole,
unpublished data).
A further indication of the potential importance of transamination in
determining the fate of metabolites and hence regulating growth of R.
leguminosarum, is provided by the finding that addition of aspartate to the
growth medium leads to an increase in glutamate excretion by sucDA mutants.
This is consistent with an increase in transamination of α-ketoglutarate due to
the presence of additional amino-donor, although increased synthesis of α-
ketoglutarate from oxaloacetate generated by transamination of aspartate, is
also likely to be a contributory factor.
Although α-ketoglutarate is excreted by sucDA mutants, there is apparently
no significant increase in the intracellular concentration of this metabolite in
these mutants. This is consistent with a report that radioactivity failed to
accumulate in α-ketoglutarate in bacteroids supplied with labelled malate, an
effect that was ascribed to rapid conversion of α-ketoglutarate to glutamate
(Salminen & Streeter, 1990).
The significantly reduced rate of glutamate excretion by the aapJ mutant
RU543 grown on glucose/NH4Cl/aspartate, in comparison to that of strain
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3841, suggests that the general amino acid permease is involved in glutamate
excretion. This is consistent with results from amino acid exchange
experiments which indicate that efflux of intracellular AIB in the presence of an
excess of extracellular amino acid is dependent on the general permease
(Section 3.2.18). It is possible that glutamate excretion occurs via exchange with
aspartate.
In view of the apparent involvement of the general amino acid permease in
the excretion of glutamate, it is interesting that strains RU116 and RU156 are
able to excrete glutamate despite exhibiting severely reduced glutamate uptake
(Table 5.5). The low rate of glutamate uptake in these strains does not appear to
be due to repression of the aap operon, or substrate inhibition, and one
possibility is that the amino acid permease is covalently modified so as to
reduce uptake but maintain efflux; although this would argue against an
exchange mechanism. Modification of the periplasmic binding protein could
potentially alter uptake without affecting efflux, and the regulation of amino
acid uptake via post-translational modification of a binding protein has been
reported previously (Celis, 1984; Celis, 1990). Furthermore, it has been
suggested that a function of high affinity ABC transporters of amino acids may
be to recapture biosynthetically produced amino acids as they escape from the
cell (Ames, 1972; Antonucci & Oxender, 1986). If the bacteroid is required to
excrete amino acids, this may provide a rationale for reducing uptake, but
maintaining efflux, via the general amino acid permease of R. leguminosarum, in
circumstances which result in the accumulation of intracellular amino acid.
In strain 3841, the gene encoding the TCA cycle enzyme, malate
dehydrogenase, is found to be located immediately upstream of the genes
encoding succinyl-CoA synthetase, which in turn lie immediately upstream of
the α-ketoglutarate dehydrogenase genes. This arrangement of genes is
different from that found in E. coli, where mdh is located in an entirely different
part of the chromosome to sucABCD (Guest, 1992; Guest & Russell, 1992), but
235
appears to be similar to that in T. aquaticus where sucCD are clustered with mdh
(Nicholls et al., 1990; Nishiyama et al., 1991), although the locations of sucAB are
not known in this case. The occurrence of the genes for the three enzymes in a
single operon in R. leguminosarum could account for the elevated malate
dehydrogenase levels observed in succinyl-CoA synthetase and α-ketoglutarate
dehydrogenase mutants, and the increased succinyl-CoA synthetase activity
found in α-ketoglutarate dehydrogenase mutants, if a mutation in sucCDAB
leads to an increase in the expression of the mdh-sucCDAB operon. One
potential reason for such an increase in expression, is that the mdh-sucCDAB
operon may be regulated in response to metabolite level(s) that are altered as a
result of the mutation.
A similar pattern of enzyme activities has been observed previously (but not
explained) for an α-ketoglutarate dehydrogenase mutant of R. meliloti (Duncan
& Fraenkel, 1979). Interestingly, a succinate dehydrogenase mutant of R.
leguminosarum is reported to exhibit a 5-fold increase in malate dehydrogenase
activity (Finan et al., 1981). This suggests that a mutation in sdhABCD has a
similar effect on the regulation of mdh expression to a sucCDAB mutation.
Attempts to isolate malate dehydrogenase mutants of strain 3841 were
unsuccessful, probably because loss of this enzyme from central metabolism is
lethal. However, if, as suspected, the promoter that controls sucCDAB
expression lies upstream of mdh, then an mdh mutant would not provide insight
into the question of whether it is the overall disruption of the TCA cycle, or the
specific loss of α-ketoglutarate dehydrogenase, that leads to severely reduced
amino acid uptake.
It seems likely that it is the lowered α-ketoglutarate dehydrogenase activity
in strain RU137, leading to some accumulation of glutamate, that is the cause of
the impaired amino acid transport in this strain. Indeed, across the strains
RU116, RU137 and RU156, there is a good correlation between the degree of
reduction in amino acid transport and that in α-ketoglutarate dehydrogenase
236
activity. However, it is not clear why α-ketoglutarate dehydrogenase activity is
reduced in strain RU137. If mdh and sucAB are indeed in the same operon, then
any regulation of α-ketoglutarate dehydrogenase must be post-transcriptional,
since malate dehydrogenase activity in strain RU137 is unaffected (Table 5.7).
It has been proposed that biosynthesis of PHB in the oxygen-limited
environment of Bradyrhizobium japonicum bacteroids serves to lower
NAD(P)H/NAD(P) ratios, thereby reducing the inhibition of certain TCA cycle
enzymes including α-ketoglutarate dehydrogenase (McDermott et al., 1989).
Under such circumstances a mutation that prevents PHB biosynthesis may lead
to increased inhibition of α-ketoglutarate dehydrogenase as a result of
increased NAD(P)H levels. If PHB production occurs in free-living cells of R.
leguminosarum under aerobic conditions, then this may provide an explanation
of the phenotype of strain RU137 with regard to α-ketoglutarate activity.
Another possibility that also implies a role for PHB biosynthesis in free-living
cells, is that mutation of phbC affects acetyl-CoA concentrations in the cell.
Acetyl-CoA plays a significant role in determining the level of TCA cycle
intermediates due to its allosteric activation of the enzyme pyruvate
carboxylase, which catalyzes the conversion of pyruvate to oxaloacetate. It is
therefore conceivable that alteration of the acetyl-CoA level may lead to
reduced α-ketoglutarate dehydrogenase activity.
Reduction in α-ketoglutarate dehydrogenase activity is the probable reason
for the escape from aspartate toxicity of strains RU116, RU137 and RU156. A
plausible escape mechanism is the removal of toxic aspartate through the
transamination of α-ketoglutarate to glutamate by aspartate aminotransferase.
Glutamate is less toxic to strain 3841 than aspartate (Section 5.2.1), and is also
excreted.
237
CHAPTER 6 FINAL DISCUSSION
238
6.1.1 The general amino acid permease of Rhizobium leguminosarum
The general amino acid permease of Rhizobium leguminosarum is encoded by
four genes, aapJQMP. Mutation of any one of these genes results in a decrease
in the uptake of amino acids as diverse as leucine, aspartate and histidine, while
over expression of all four genes together leads to a substantial increase in
transport of all the amino acids tested.
The results from amino acid uptake assays on mutants indicate that the
general amino acid permease is the main high-affinity uptake system for
glutamate, aspartate and histidine in free-living cells of R. leguminosarum strain
3841. While this transporter also accounts for a significant percentage of
alanine, leucine and methionine uptake, the residual transport rates in aap
mutants indicate that there are other specific uptake systems for these
substrates. The failure of an aapJQM mutant to grow on proline, suggests that
the general permease is the major uptake system for this amino acid.
Complementation studies indicate that the genes aapJQMP are transcribed
from a single promoter upstream of aapJ, although there may be some
additional weak promoter activity upstream of aapP. The nucleotide sequence
contains a region of dyad symmetry downstream of aapP, that has the
characteristics of a rho-independent transcriptional terminator. It is therefore
likely that these four genes comprise an operon.
The sequence homology data, and in particular the presence of known
signature sequences, suggest that aapJQMP encode the components of an ABC
transporter. The periplasmic binding protein, AapJ, and the two integral
membrane components, AapQ and AapM, are significantly larger than the
equivalent components of previously described ABC transporters of amino
acids. This increased size may be a function of the broad specificity of this
transporter. The deduced polypeptides from the aapJQMP genes exhibit strong
homology to sequence from the Escherichia coli genome sequencing project,
239
suggesting that E. coli may possess a previously unreported general amino acid
permease.
TFASTA and BLAST searches indicate homology of AapQ and AapM to the
integral membrane components of ABC transporters of polar amino acids or
glutamine. In addition to the "integral membrane component signature"
(Saurin et al., 1994), a sequence alignment of AapQ and AapM with these
proteins, reveals a region of 63 amino acids containing 18 highly conserved
residues, that is located, in the majority of cases, at the N-terminal end of the
protein. This suggests that amino acid transporters of this type may constitute a
sub-family of the ABC superfamily.
Predicted topologies containing eight transmembrane segments in the case of
the general permeases, and five membrane-spanning regions in the case of the
other known members of this sub-family, suggest that the N-terminal
conserved region spans two transmembrane segments and a connecting
cytoplasmic loop, while the integral membrane component signature is located
in a cytoplasmic loop. This is consistent with the experimentally determined
topologies of HisM and HisQ of the histidine transporter from S. typhimurium
(Kerppola et al., 1991; Kerppola & Ames, 1992).
For one of the conserved leucines in the N-terminal region, there is a
correlation between the nature of the substrate translocated by proteins in
which it is substituted, and the nature of the amino acid that substitutes. It is
therefore possible that the residue at this position, which is predicted to lie in a
transmembrane segment, is involved in determining substrate specificity.
Furthermore, the deletion from HisM of four amino acids lying in the other
predicted transmembrane segment in this N-terminal region, has been found to
alter the specificity of the histidine transporter in S. typhimurium, from L-
histidine to L-histidinol (Payne et al., 1985).
The rate of exchange of intracellular and extracellular amino acids by strain
3841 is dependent upon the number of copies of the aap operon present,
240
suggesting that the general permease facilitates efflux of amino acids, in
addition to uptake. Translocation of substrate in both directions has not been
previously reported for an ABC transporter, and the most likely explanation is
that the general amino acid permease actively imports amino acids but also
allows outward diffusion. Thus, while the reduction in amino acid exchange in
an aapP mutant of strain 3841 may indicate that ATP hydrolysis is required to
fuel efflux, it seems more likely that the energy is needed to open a membrane
pore upon binding of a liganded binding protein complex. Whether
intracellular substrate has access to such a pore at all times, or is dependent
upon the binding and/or uptake of extracellular substrate for efflux, is an
important mechanistic question, of potential relevance to other ABC
transporters. Since amino acid exchange is reduced in a strain lacking only
AapJ, and assuming that, as in the histidine transporter of S. typhimurium
(Prossnitz et al., 1988; Prossnitz et al., 1989), unliganded binding protein does
not interact with the membrane-bound complex, it is reasonable to conclude
that binding of extracellular substrate, at least, is required for efflux of
intracellular substrate.
The physiological significance of the exchange capability of the general
amino acid permease is unknown. Certainly this capability is not required for
nitrogen fixation by strain 3841, as aap mutants induce pea nodules that reduce
acetylene as effectively as those induced by the wild type. Since the general
amino acid permease appears to be the major high affinity glutamate
transporter in strain 3841, this observation suggests that nitrogen fixation in
Rhizobium leguminosarum is not fuelled via a malate-aspartate shuttle in the
nodule as proposed by Kahn et al. (1985). However, the possibility that
alternative glutamate/aspartate transporters are induced under symbiotic
conditions means that the operation of this shuttle can not be completely
discounted.
241
The results from both aapJQM::lacZ transcriptional fusion studies and
glutamate uptake assays on a ntrC mutant, suggest that the aap operon of R.
leguminosarum strain 3841 is negatively regulated by NtrC in response to
nitrogen supply. In batch culture, nitrogen-excess leads to an approximately 5-
fold repression of the aap operon relative to the nitrogen-limitation.
One potential explanation of this repression that fits the available data, is
that, in contrast to NtrC, phosphorylated NtrC has a lower affinity for the
binding sites upstream of aapJQMP, than it does for those involved in
controlling the expression of other genes, such as glnII, which has been shown
to be positively regulated by NtrC (Carlson et al., 1987; Martin et al., 1988; de
Bruijn et al., 1989; Rossi et al., 1989; Shatters et al., 1989; Patriarca et al., 1992). If
this is the case, then because in R. leguminosarum transcription of ntrC is
essentially independent of the nitrogen status of the cell (Patriarca et al., 1993;
Amar et al., 1994), under nitrogen-limitation, the amount of NtrC (in the form of
NtrC-phosphate) available to bind to the promoter region of the aap operon will
be reduced, and transcription of aapJQMP consequently increased.
Uptake by the general amino acid permease of R. leguminosarum strain 3841
may also be regulated in response to activity of the TCA cycle enzyme α-
ketoglutarate dehydrogenase. Abolition of α-ketoglutarate dehydrogenase
activity through mutation of sucA, results in the almost total loss of uptake by
the general amino acid permease. Mutation of sucD, a gene located upstream of
sucA in the same operon and encoding the alpha subunit of succinyl-CoA
synthetase, reduces α-ketoglutarate dehydrogenase activity by 95% and amino
acid uptake via the general permease by 86%. Mutation of phbC, the gene
encoding poly-beta-hydroxybutyrate synthase, reduces α-ketoglutarate
dehydrogenase activity by 61%, possibly as a result of altering intracellular
NAD(P)H or acetyl-CoA levels, and also causes a 61% reduction in aspartate
uptake.
242
Data from an aapJ::lacZ fusion indicate that there is no change in the
transcription levels of the aap operon in sucDA mutants, and the conditions
encountered by cells prior to transport assays effectively eliminate the
possibility of intracellular substrate inhibition of the general amino acid
permease. This suggests that the general amino acid permease may be subject
to post-translational modification in these mutants.
The intracellular glutamate concentration is elevated in sucDA mutants
grown on glucose/NH4Cl, and it is possible that this elevation is the signal for
the down-regulation of the general amino acid permease. In addition,
glutamate is excreted by sucDA mutants grown on both glucose/NH4Cl and
malate/NH4Cl. These results are consistent with suggestions that the observed
synthesis of glutamate in bacteroids is a consequence of inhibition of α-
ketoglutarate dehydrogenase by NADH (McDermott et al., 1989; Salminen &
Streeter, 1990).
Indeed the inhibition and/or repression of central metabolic enzymes under
the oxygen-limited conditions of the nodule, may account for the observed
excretion of the amino acids alanine and aspartate by bacteroids under nitrogen
fixing conditions (Kretovich et al., 1986; Appels & Haaker, 1991; Kouchi et al.,
1991; Rosendahl et al., 1992). Such excretion may alleviate inhibition of the TCA
cycle by ketoacids and/or reducing equivalents, which would otherwise
accumulate in the non-growing bacteroid cell.
The potential for the involvement of the general amino acid permease in
amino acid excretion by the bacteroid is indicated by the fact that while strain
3841 is found to excrete glutamate when grown on glucose/NH4Cl/aspartate,
an aapJQM mutant exhibits only a relatively low rate of glutamate excretion
under these conditions. This is compatible with the results from amino acid
exchange experiments which indicate that efflux of intracellular amino acid can
be mediated by the general permease.
243
In addition, sucDA mutants of strain 3841 are able to excrete glutamate
despite the fact that glutamate uptake by these mutants is extremely low. Since
glutamate transport in these mutants is apparently not repressed, or subject to
substrate inhibition, this suggests that the general amino acid permease may be
covalently modified so as to reduce uptake but maintain efflux. Modification of
the periplasmic binding protein could potentially alter uptake without affecting
efflux, and the regulation of amino acid uptake via post-translational
modification of a binding protein has been reported previously (Celis, 1984;
Celis, 1990). If the bacteroid is required to excrete amino acids, such
modification may be required to prevent recapture of excreted amino acids by
this high affinity system, while maintaining its ability to mediate efflux.
However, since aap mutants form effective nodules, it is clear that the amino
acid excretion capacity of the general permease is not required for nitrogen
fixation.
In the case of Rhizobium leguminosarum alanine is the predominant amino
acid found to be excreted from bacteroids and symbiosomes under nitrogen-
fixing conditions (Appels & Haaker, 1991; Rosendahl et al., 1992). If this
excretion of alanine is the primary mechanism for maintaining sufficient flux
through the TCA cycle to fuel nitrogen fixation in the bacteroid, then the fact
that the general amino acid permease is not required for an effective symbiosis
is unsurprising: Alanine uptake by aap mutants indicates the existence of one
or more alternative alanine carriers, and the results of exchange experiments on
an aapJQM mutant, using the alanine analogue AIB, suggest that such a carrier
(or carriers) allows efflux of substrate.
6.1.2 TCA cycle enzymes in Rhizobium leguminosarum
Partial sequencing of a cosmid that complements a mutation in sucA,
suggests that in R. leguminosarum strain 3841 the genes encoding the TCA cycle
enzymes malate dehydrogenase, succinyl-CoA synthetase and α-ketoglutarate
dehydrogenase are clustered, and occur in the order mdh-sucCD-sucAB. The
244
results of complementation studies suggest that sucCDAB are under the control
of a single promoter, and it is possible that the same promoter also initiates
transcription of mdh, although this was not verified.
The presence of mdh-sucCDAB in a single operon could account for the
elevated malate dehydrogenase levels observed in succinyl-CoA synthetase and
α-ketoglutarate dehydrogenase mutants, and the increased succinyl-CoA
synthetase activity found in α-ketoglutarate dehydrogenase mutants, if a
mutation in sucCDAB leads to an increase in the expression of the mdh-sucCDAB
operon. One potential reason for such an increase in expression, is that the mdh-
sucCDAB operon may be regulated in response to metabolite level(s) that are
altered as a result of the mutation.
Attempts to isolate malate dehydrogenase mutants of strain 3841 by
homogenotization of cosmids mutated in mdh, were unsuccessful, suggesting
that loss of this enzyme may be lethal to R. leguminosarum.
6.1.3 Methionine biosynthetic enzymes in Rhizobium leguminosarum
On the basis of sequence homology, the gene lying immediately upstream of
the aap operon in R. leguminosarum strain 3841, is likely to be metC, encoding
beta-cystathionase. Similarly, the gene lying ~3.9kb upstream of the aap operon
is likely to be cysE, encoding serine acetyltransferase. Both these enzymes are
involved in methionine biosynthesis and the genes are transcribed in the
opposite direction to the aap operon. However, metC and cysE are apparently
not contained in the same operon, since activities of lacZ fusions to the metC-
cysE intergenic region of ~2.4kb, are indicative of transcription in the opposite
direction to metC and cysE. Data from cysE::lacZ and metC::lacZ fusions suggest
that both cysE and metC are negatively regulated by NtrC.
Mutation of cysE causes a reduction of amino acid transport in strain 3841
grown on glutamate as the nitrogen source. This indicates that the ability of
amino acid transport in R. leguminosarum to respond normally to nitrogen-
limitation is dependent upon the presence of serine acetyltransferase.
245
However, the homology of CysE to NifP from Azotobacter, and the observation
that a cysE mutant is not a cysteine auxotroph, suggest that R. leguminosarum
may possess two genes encoding serine acetyltransferase. This may account for
the wild-type amino acid transport rates exhibited by a cysE mutant of strain
3841 under nitrogen-excess.
6.1.4 Future work
Because of the unusual characteristics of the R. leguminosarum general amino
acid permease, the general amino acid permeases of other bacteria, particularly
E. coli, probably warrant further investigation.
Since the nucleotide sequence of the DNA flanking the aapJQMP operon of E.
coli is known, the DNA containing these genes could be amplified from
chromosomal DNA by PCR and hence cloned. This would allow investigation
of the effects of mutation and over expression of these genes on amino acid
transport in E. coli. The nucleotide sequence in the two areas of the E. coli
sequence where frame shifts are apparent from the R. leguminosarum sequence,
could also be checked.
It may be possible to investigate the extent of distribution of this transporter
among the bacterial species through a combination of PCR, using degenerate
primers designed from the known aapJ sequences of R. leguminosarum, E. coli
and P. fluorescens, to amplify a band of known size from chromosomal DNA
containing an aapJ gene; and Southern blot, using R. leguminosarum aapJ DNA as
the probe, to confirm the nature of the amplified band.
Further study of the R. leguminosarum general amino acid permease could
yield structural and mechanistic details of general applicability to ABC
transporters. In order to investigate whether the general permease actively
exports amino acids, or simply enables diffusion, the transporter could be
reconstituted in E. coli inside-out membrane vesicles, and amino acid uptake by
these vesicles measured in both the presence and absence of ATP. The topology
of the integral membrane components AapM and AapQ could be investigated
246
by TnphoA fusion analysis, while the possible involvement of particular
residues in these proteins in substrate specificity could be tested through site-
directed mutagenesis.
Since alanine excretion may be important to bacteroid function, and this
excretion may be mediated by an alanine-specific transporter(s) in R.
leguminosarum, the cloning of the gene(s) encoding alanine-specific uptake
system(s) could be attempted. Cloning of this gene(s) would enable the
symbiotic effectiveness of strains mutated in both the general amino acid
permease and alanine-specific transport to be investigated. Strategies for
cloning alanine-specific transporters include looking for increased labelling by
[14C]alanine in an aap mutant carrying an aap mutant chromosomal library; or
obtaining alanine transport mutants by screening for loss of growth on alanine
in transposon mutants of an aap-deleted strain, and then complementing the
mutation from an aap mutant chromosomal library.
The question of whether mdh lies in the same operon as sucCDAB in R.
leguminosarum could be resolved by attempting to complement a sucCDAB
mutation with a clone of sucCDAB that includes the complete mdh-sucC
intergenic region, or by cloning the mdh-sucC intergenic region into a promoter
probe vector. Both these options would require further restriction mapping of
the mdh-sucCDAB region, in order to reveal suitable fragments for cloning.
Investigation of the activity of lacZ fusions to mdh and sucCD, in sucAB mutant
backgrounds, would reveal whether elevated malate dehydrogenase and
succinyl-CoA synthetase activities in α-ketoglutarate dehydrogenase mutants
are the result of increased transcription.
247
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