Amino Acid Transport
Amino Acid Transport
Amino Acid Transport
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US ~ epar~ent of Agriculture-~~S and ~epartment of Plant ~iology,<br />
~niversi~ of ~llinois at Urb~na-Champai~n, Urbana, Illinois<br />
*<br />
of the uni~ue features of lants as multicellular organisms is their ability to synthes<br />
from simple inorganic salts and the free ener~y made<br />
tr~sduction reactions of photosyn~esis.<br />
no acids. ~lthoug~ every pla<br />
capacity to assi~late inorganic ni<br />
ly restricted to cells in mature leave<br />
no acids from sites of pri<br />
trogen. I ~ aminoacids ~ s ~ e ~<br />
esis, they are also the precursors of every ~~ogen<br />
luding nucleic acids, gro~th regulators, ~hotosynarray<br />
of other essential compounds. In m ~ y<br />
of ~~ogen me~bolis~ in the plant and underd<br />
bet~een organs is a ~ndamental ~uestio~ in<br />
ids in plants must neces<br />
from the soil solution by the<br />
h onium ions<br />
ssl~lated in the root, or
3 ush<br />
general patterns have been identified (Pate 1980, 1983; Andrews 1986). Temperate perennials<br />
and legumes frequently carry out primary assimilation in the roots, whereas<br />
tropical and subtropical species strongly favor leaf assimilation. Temperate annuals fall<br />
into both categories. Significantly, if high concentrations of nitrate are available in the<br />
soil solution (>1 mM), many species carry out primary assimilation in the leaf. Even<br />
symbiotic nitrogen assimilation is suppressed in the presence of high levels of nitrate<br />
(Harper 1994; Pate 1980).<br />
Once assimilated into amino acids, there are several pathways for the long-distance<br />
transport of reduced nitrogen (Fig. 1). For those plants and conditions in which primary<br />
assimilation occurs in the leaf, amino acids are transported to the heterotrophic tissues<br />
by the phloem-translocation stream. In species that use an apoplastic phloem-loading<br />
mechanism (Bush 1992; Turgeon 1996), this requires the release of amino acids from<br />
mesophyll cells into the apoplastic space, with subsequent loading into the phloem. A<br />
few amino acids (generally a combination of two or more of the following: glutamate,<br />
aspartate, glutamine, asparagine, alanine, and serine) are more concentrated in the<br />
phloem-translocation stream than in the cytoplasm of the mesophyll (Riens et al. 1991;<br />
Winter et al. 1992; Lohaus et al. 1994; Lam et al. 1995). These amino acids must be<br />
actively transported into the phloem (Bush 1993a). In contrast, the measured concentrations<br />
of other amino acids in the phloem suggests they are at or below ~ermodyn~c<br />
equilibrium with the mesophyll and passive transport activity has been suggested (Riens<br />
et al. 199 1; Winter et al. 1992; Lohaus et al. 1994). However, it is important to recognize<br />
that the phloem represents a dynamic pool of amino acids that is rapidly turning over as<br />
pressure-driven mass flow continuously removes the loaded metabolites. Thus, although<br />
snap-shot determinations of amino acid concentrations may implicate passive transport<br />
mechanisms for some amino acids, the kinetics of amino acid turnover suggests that high<br />
rates of transport activity must occur, and passive transport activity, for which the rate<br />
of transport is directly proportional to the concentration difference, may not be rapid<br />
enough to maintain the phloem pools that are quickly flushed away. Given the dynamics<br />
of rapid exchange at the phloem interface, it is not surprising that several high-affinity,<br />
proton-coupled amino acid transporters have been described in plasma membrane vesicles<br />
isolated from mature leaf tissue (Li and Bush 1990, 1991, 1992; Williams et al.<br />
1990, 1992, 1996; Weston et al. 1995). These active transporters can maintain high rates<br />
of flux, even against substantial concentration differences. Nevertheless, evidence for<br />
low levels of facilitated diffusion is also present in membrane vesicles isolated from leaf<br />
tissue (Williams et al. 1990; D. R. Bush unpublished data). Although the participation<br />
of facilitated transporters in phloem loading has yet to be demonstrated, such porters are<br />
likely candidates to play an important role in releasing amino acids from the surrounding<br />
mesophyll cells.<br />
When nitrate or ammonium ions are assimilated in the root, amino acids are transported<br />
to mature leaf tissue in the xylem by the transpiration stream (Pate 1980, 1983).<br />
Because the nitrogen requirement of mature leaves is low, a substantial portion of the<br />
amino acids arriving with the transpiration stream are cycled into the phloem for redistribution<br />
to heterotrophic sinks (see Fig. 1). Cycling amino acids from the xylem to the<br />
phloem in mature leaf tissue is also a fundamental process in symbiotic nitrogen fixation.<br />
For example, atmospheric nitrogen assimilated in the Rhizobium-legume symbiosis is<br />
exported from the root in the transpiration stream as amide amino acids (examples include,<br />
clover and alfalfa) or ureides (mung bean and soybean; Schubert 1986; Winkler<br />
et al, 1988). Even though some of the amide amino acids may be removed from the
<strong>Amino</strong> <strong>Acid</strong> Import<br />
Developing leaves,<br />
<strong>Amino</strong> <strong>Acid</strong> Import<br />
<strong>Amino</strong> <strong>Acid</strong> Cycling<br />
(xylem to phloem)<br />
<strong>Amino</strong> <strong>Acid</strong> Import<br />
<strong>Amino</strong> <strong>Acid</strong> Export<br />
Primary ~ssimilation<br />
<strong>Amino</strong> <strong>Acid</strong> Cycling<br />
(phloem to xylem)<br />
/<br />
<strong>Amino</strong> <strong>Acid</strong> Import<br />
- - = Xylem<br />
Phloem<br />
Figure 1 Schematic presentation of amino acid transport pathways in the plant: <strong>Amino</strong> acids<br />
synthesized in the root from primary assimilation or symbiotic relations are transported to mature<br />
leaf tissue in the xylem (----). <strong>Amino</strong> acids synthesized in the leaf, or arriving from the root in the<br />
xylem, are transported out of the leaf in the phloem (-) to satisfy the needs of heterotrophic<br />
sids. Import-dependent tissues include developing leaves, roots, cortical cells in the stem, seed<br />
and fruits, and apical meristems.<br />
transpiration stream by stem tissues (McNeil et al. 1979; Pate 1983), much of that nitrogen<br />
arrives in mature leaf tissue, from whichit is redistributed by the phloem. The<br />
ureides also arrive in the leaf, but instead of being redistributed in the phloem, they are<br />
catabolized to C02 and (Winkler et al. 1988). Released ammonia is incorporated<br />
into amino acids that are used in leaf metabolism or redistributed to heterotrophic tissues<br />
by the phloem.<br />
Nitrogen cycling between the vascular systems is not restricted to the leaf tissue.<br />
Several lines of evidence show that amino acids can also move from the phloem to the
xylem in the root (Pate 1983; Cooper and Clarkson 1989; Schurr and Schulze 1995).<br />
Thus, cycling amino acids between the two vascular systems in mature leaf and root<br />
tissue (see Fig. 1) permits dynamic adjustments of carbon and nitrogen metabolism in<br />
response to developmental queues or environmental change. For example, the metabolic<br />
needs of the mature root system emphasizes catabolic reactions associated with maintenance<br />
reactions versus the anabolic reactions that drive early growth. Therefore, excess<br />
amino acids transported to the root under these conditions can be redirected to the shoot<br />
for subsequent incorporation into developing seed. Significantly, excess amino acid nitrogen<br />
arriving at the root may also represent a dynamic signal that triggers the downregulation<br />
of nitrate assimilation (Cooper and Clarkson 1989; Crawford 1995; Padgett<br />
and Leonard 1996).<br />
In addition to its role in primary assimilation, amino acid transport is a key process<br />
in leaf senescence and seed germination. In rice, for example, more than 60% of the<br />
amino nitrogen delivered to developing leaves and ears is derived from amino acids<br />
retrieved from senescing older leaves (Mae et al. 1983, 1985; Feller and Fischer 1994).<br />
Likewise, amino acids released from storage proteins during seed ge~nation are the<br />
principal nitrogen source in growing seedlings (Schobert and Komor 1989). Taken together,<br />
it seems clear that amino acids are the currency of nitrogen exchange in the plant.<br />
<strong>Amino</strong> acids are transported into plant cell by proton-coupled symporters that link translocation<br />
across the plasma membrane to the proton-motive force generated by the €I?-<br />
pumping ATPase (Fig. 2; Bush 1993a). These are widely expressed carriers that appear<br />
to be the primary pathways for amino acid transport into plant cells. However, there is<br />
some evidence for facilitated transporters that mediate passive amino acid transport, although<br />
little biochemical and no molecular characterization of such porters has been<br />
published.<br />
The amino acid symporters have been investigated in recent years using purified<br />
membrane vesicles and imposed proton electrochemical potential differences (Bush<br />
1993a). Purified membrane vesicles are a very useful experimental system for studying<br />
membrane transporters. Although membrane transport activity can be examined with<br />
intact cells, there are many complications associated with metabolism and intracellular<br />
comp~mentation that limit experimental interpretation. Additionally, it is difficult to<br />
dissect the bioenergetics of a transport system in intact tissues because of complex interactions<br />
among the primary pumps, ion channels, and other unrelated porters. Isolated<br />
membrane vesicles, on the other hand, allow one to focus on specific transport processes<br />
while minimizing problems associated with the living cell. The major attributes of the<br />
membrane vesicle approach include (1) unequivocal identification of membrane location<br />
using purified vesicles, (2) elimination of posttransport metabolism and compartmentation,<br />
and (3) the ability to control both intra- and extravesiculq solution composition<br />
(Bush 1992). The ability to control solution co~position on both sides of the membrane<br />
is particularly useful for investigating the bioenergetics of the transport process. With<br />
this approach, the basic transport properties of the plant amino acid symporters have<br />
been described (Bush 1993a).<br />
Plant amino acid symporters are electrogenic transporters. Electrogenicity was
Model of amino acid symporter-mediated transport into plant cells: The H+-ATPase<br />
generates a substantial proton electrochemical potential difference (both ApH and AY) that can<br />
drive transport reactions three or four orders of magnitude away from equilibrium. A diagrammatic<br />
representation of the 11 transmembrane domains (TMDs) of NAT2/AAPl is also presented.<br />
demonstrated in isolated membrane vesicles by examining the effect of membrane electrical<br />
potential (AY) on ApH-dependent transport activity. Potassium gradients and valinomycin<br />
were used to manipulate AY. In the absence of compensating charge flow, a low<br />
rate of amino acid flux was observed. When the AY was clamped at zero, transport was<br />
stimulated fourfold, and when a negative AY was imposed, transport was stimulated<br />
sixfold. These data are consistent with an electrogenic transport system that results in<br />
primary charge separation as proton cotransport drives substrate accumulation (Li and<br />
Bush 1990). Similar results were reported for this porter expressed in Xenopus oocytes<br />
(Boorer et al. 1996).<br />
The stoichiometry of H"-amino acid cotransport was recently measured using electrophysiologic~<br />
methods to measure conductance through another plant amino acid symporter<br />
(AAP5) expressed in Xenopus oocytes (Boorer and Fischer 1997). Boorer and<br />
Fischer (1997) used direct measurements of [3H]amino acid uptake and simultaneous<br />
measurements of inward current to show that the stoich~o~etry of the amino acid symporters<br />
is one w' per one amino acid. This held true for the basic, neutral, and acidic<br />
amino acids examined. Because the acidic amino acids also carried one positive charge,<br />
these results suggest they are transported in their neutral forms.<br />
The number of symporters that mediate amino acid transport into higher-plant cells<br />
has yet to be determined. <strong>Transport</strong> competition experiments with isolated membr~e<br />
vesicles provided evidence of at least four amino acid symporters: an acidic amino acid<br />
symporter, a basic amino acid symporter, and two symporters for the neutral amino acids<br />
(Li and Bush 1990, 1991). The neutral amino acid porters were resolved in the competition<br />
experiments because of their differential affinity for isoleucine, valine, threonine,<br />
and proline. A more thorough examination of the kinetics of inter-amino acid transport
competition among the neutral amino acids confirmed the initial conclusion. Li and Bush<br />
(1991) hypothesized that branching at the fharbon in isoleucine, valine, and threonine<br />
results in steric interactions that block their access to one of the neutral porters. More<br />
recently, several amino acid symporters have been cloned, and we now know that many<br />
amino acid symporters are found in the plant (see later discussion).<br />
The ability of various amino acid analogues to compete for transport into purified<br />
membrane vesicles was used to identify molecular determinants that are important for<br />
substrate recognition by the neutral amino acid symporters (Li and Bush 1992). D-ISOmers<br />
of alanine and isoleucine were not effective transport antagonists of the L-isomers,<br />
thereby implicating stereospecificity and suggesting that positional relations between the<br />
a-amino and carboxyl groups is an important parameter in substrate recognition. This<br />
conclusion was supported by the observation that p-alanine and analogues with methylation<br />
at the a-carbon, at the carboxyl group, or at the a-amino group, were not effective<br />
transport antagonists. In contrast, analogues with various substitutions at the distal end<br />
of the amino acid were less potent antagonists. More recently, el~trophysiological investigations<br />
of AAPS expressed in Xenopus oocytes also identified the a-amino and carboxyl<br />
groups, as well as the P-carbon, as important dete~inants in defining substrate<br />
specificity (Boorer and Fischer 1997). Thus, the binding site for the carboxyl end of the<br />
amino acid is a well-defined space that is characterized by compact, asymmetric positional<br />
relations and specific ligand interactions (Li and Bush 1992).<br />
The amino acid symporters are inhibited by carbonyl cyanide ~-chlorophenyl hydrazone<br />
(CCCP) and diethylpyrocarbonate (DEPC; Bush and Langston-~nkefer 1988; Li<br />
and Bush 1990). CCCP is a protonophore that inhibits amino acid transport by dissipating<br />
the proton-motive force that drives the symport reaction. DEPC is a small molecule that<br />
forms covalent bonds with the imidazole ring of histidine residues. The time-dependent<br />
inactivation of alanine transport in the presence and absence of amino acids showed that<br />
substrate binding protects the symporter from DEPC inactivation (Fig. 3). This observation<br />
is consistent with DEPC binding at (or at least confo~ationally linked to) the<br />
substrate-binding site of the carrier (Bush and Li 1991; see Bush 1993b, for discussion).<br />
Since histidine residues frequently participate in protonation reactions, this observation<br />
implicates a histidine residue in the reaction mechanism.<br />
Initial efforts to identify the amino acid syrnporters by using standard biochemical strategieswerelargelyunsuccessful.Althoughthiswasoftenquitefrustrating,<br />
it was not<br />
too surprising, given that these are very low-abundance, integral membrane prot6ins.<br />
Fortunately, a novel molecular approach was successfully used to clone several plant<br />
amino acid symporters and biochemical analysis has moved forward using these clones<br />
and a variety of expression systems.<br />
The first plant amino acid symporter cloned was identified using ~nctional complementation<br />
of yeast amino acid transport mutants (Frommer et al. 1993; Wsu et ai.<br />
1993). ~acc~aro~~ces cerevisiae strains that are auxotrophic and transport-deficient for<br />
a specific amino acid were tr~sfo~ed with plant cDNA libraries that are constructed<br />
in yeast expression vectors. Those transformants that successfully expressed an appropriate<br />
plant amino acid symporter were easily identified by screening for growth on low<br />
concentrations of the limiting amino acid. Because these yeast strains are both auxotrophic<br />
and transport-deficient, a simple secondary screen in the absence of the limiting
80<br />
40<br />
8 20<br />
0<br />
0.0 2.0 4.0 6.0<br />
ure 3 Time- and concentration-dependent inactivation of proton-coupled alanine transport<br />
into purified plasma membrane vesicles (PMVs) isolated from sugar beet leaf tissue: PMVs were<br />
pretreated with DEPC plus or minus alanine for the indicated times, at which point the reaction<br />
was stopped. The vesicles were pelleted and resuspended in inhibitor-~ee buffer, and transport<br />
activity was measured as described previously (Li and Bush 1990).<br />
amino acid was used to show that growth was dependent on amino acid transport, rather<br />
than complementation of the biosynthetic pathway. This is a powerful approach because<br />
it allows one to clone transporter cDNAs without protein sequence information, and<br />
because it provides a useful expression system for subsequent investigations of the biochemical<br />
properties of the trans~ort protein (Bush 1993a; Tanner and Casper 1996). The<br />
screens used for functional co~ple~entation s~ould also id en ti^ high-affinity facilitated<br />
diffusion transpo~ers. in spite of numerous screens of different expression<br />
libraries in a variety of ast strains, no examples of facilitated transporte~ have<br />
been reported.<br />
The cDNA clone of the first amino acid symporter identi~ed with functional complementation<br />
encoded a protein containing 486 amino acids with a calculated molecul~<br />
mass (MJ of 52.9 kDa (Prommer et al. 1993; Hsu et al. 1993). Hy~opathy analysis of<br />
the deduced amino acid sequence suggests this is a typical hydrophobic membrane protein<br />
with 10 to 12 membrane-spanning domains and three sites of potential N-linked<br />
glycosylation. The transport kinetics, substrate specificity, and inhibitor sensitivity of the<br />
heterologously expressed carrier matched those described for one of the neutral amino<br />
acid symporters ch~acterized in purified membrane vesicles (Li and ush 1991). Hsu et<br />
al. (1993) designated this gene ~A~ for neutral amino acid transporter TI, and Prommer<br />
et al. (1993) termed it ~~~. No homologues were identi~ed in the databases, suggesting<br />
that this plant amino acid sympo~er was the first example of a new class of tr~s~rters.
Fromer’s group identified five additional amino acid transporters (AAP2-AAP6)<br />
that are closely related to NAT2/AAPl (Kwart et al, 1993; Fischer et al. 1995; Rentsch<br />
et al. 1996). They are similar in size and predicted topology, and sequence homologies<br />
range from 70 to 95% amino acid si~larity, thus providing good evidence that these<br />
clones represent a family of amino acid translocators (Fischer et al. 1995; Rentsch et al.<br />
1996). As might be expected for a gene family within a given organism, these porters are<br />
differentiated by a combination of unique expression patterns and substrate specificities.<br />
Individually, the AAP transporters have broad substrate specificity, although each exhibits<br />
some preference (lower Km or higher Yma) for certain classes of amino acids. For<br />
example, AAP4 is particularly active for valine and proline, whereas AAP5 is an effective<br />
lysine transporter (Fischer et al, 1995). Within the family, however., there appears to<br />
be some redundancy, for AAP1, AAP2, AAP4, and Asup6 share similar patterns of substrate<br />
specificity, whereas AM3 and AN5 are similar (Fischer et al. 1995; Rentsch et<br />
al. 1996).<br />
Although similar substrate specificity could lead to widespread functional redundancies<br />
between the AAPs., differential expression patterns appear to separate the various<br />
AAI? gene products from one another. For example, AAPS and AAPlNAT2 are widely<br />
expressed in all tissues, whereas AAP3 expression is restricted to the root (Fischer et al.<br />
1995). Moreover, even when related AAPs are expressed in the same organ, they can be<br />
functionally isolated by restricting expression to different cell types within that organ.<br />
Four other classes of plant amino acid transporters have been cloned that are not<br />
members of the AAP gene family. Two of them are basic amino acid ~anspo~ers that<br />
were also cloned by functional complementation: UTI encodes a lysine and arginine<br />
transpo~er that has 533 amino acids and 14 putative transmembrane domains (Frommer<br />
et al. 1995). It is a single-copy gene in ~r~bi~o~sis and sequence alignments suggest it<br />
is closely related to a human basic amino acid transporter (Yoshimoto et al. 1991; Christensen<br />
1992). The second basic amino acid transporter cloned is LHTl (Chen and Bush,<br />
1997). It is a lysine and histidine transporter that contains 446 amino acids and 9-10<br />
transmembrane domains. In contrast to AATl , LHTl does not transport arginine. RNA<br />
gel blot analysis and whole-mount in situ hybridization indicate that LHTl is most<br />
strongly expressed in pollen, siliques, and on the root surface. AATl is most strongly<br />
expressed in vascular tissue and flowers (Frornmer et al. 1995). The expression patterns<br />
of LHTl and UT1 complement one another, suggesting ~HTl may be involved in<br />
amino acid acquisition, whereas AATl may contribute to long-distance transpo~. In<br />
addition to these basic amino acid transporters, an aromatic amino acid transporter has<br />
also been recently characterized.<br />
An aromatic amino acid transporter was identified using functional expression of<br />
an EST cDNA that exhibited low levels of sequence homology to NAT2/AAPl (L. Chen<br />
and D. R. Bush, unpublished data). The cDNA is 1.6 kb, with an open-reading frame<br />
that codes for a protein with 432 amino acids and a calculated M, of 50 ma. Hy~opathy<br />
analysis of the deduced amino acid sequence suggests this is an integral membrane protein<br />
with 11 putative transmembrane domains. This protein exhibits sequence si~l~ty<br />
with AUX1 (a putative auxin transporter; Bennett et al. 1996) and to tryptophan and<br />
tyrosine translocators in E. coli (Wookey and Pittard 1988; Sarsero et al. 1991; Heatwole<br />
and Somerville 1991). RNA gel blot analysis has shown that this transporter is expressed<br />
at very low levels and that it is in all tissues. Whole-mount in situ hybridization further<br />
localized ART1 expression on the surface of roots and leaves and in young seedlings
and in stomata. Overall, ART1 belongs to<br />
favors aromatic amino acids and, perhaps,<br />
Chen and D. R. Bush, unpublished data).<br />
Two proline transporters have also<br />
a new class of amino acid transporter that<br />
to other aromatic compounds in plants (L.<br />
been recently cloned, ProTl and Pro72<br />
(Rentsch et al. 1996). These transporters were identified by complementing a Shr3- yeast<br />
strain that is unable to target yeast amino acid transporters to the plasma membrane.<br />
Significantly, the plant amino acid transporters are not affected by this processing mutation;<br />
therefore, several amino acid transporters (many of which were previously identified)<br />
complemented this yeast strain. A survey of different amino acids for evidence of<br />
transport through these porters showed that they are proline translocators. However, the<br />
rate of proline transport in yeast cells expressing these porters is 300 times slower than<br />
proline transport by AAP6 expressed in the same cell line. It is not known if this is a<br />
result of lower expression levels, or if it is an intrinsic property of these transporters.<br />
ProTl and ProT2 are closely related genes that encode typical membrane proteins that<br />
contain 442 and 439 amino acids, and have 10 putative transmembrane domains. Both<br />
genes are widely expressed in Arabidopsis, with ProTl strongest in roots, flowers and<br />
stems. Significantly, Pro22 expression was enhanced under conditions of water or salt<br />
stress, which is consistent with proline’s putative role in water and salt tolerance (Delauney<br />
and Verma 1993).<br />
Molecular descriptions of amino acid transport have included the isolation of several<br />
transport mutants in Arabidupsis. Two mutants that lack a low-affinity basic amino<br />
acid transport activity were isolated based on their resistance to lysine plus threonine or<br />
to ~-2-~inoethyl-~-cysteine, respectively (Heremans et al. 1997). The specificity of the<br />
lost transport activities were resolved based on apparent K, and sensitivity to transport<br />
inhibitors. Although both mutations have been mapped to chromosome 1, it is not clear<br />
if they are allelic. In a similar approach, the raz1 mutant of Arabidopsis was selected<br />
based on its resistance to the toxic proline analogue, azetidine-2-carboxylic acid (Verbruggen<br />
et al. 1996). Likewise, we have isolated two T-DNA-tagged lines that are resistant<br />
to high concentrations of valine in their growth medium (Chen and Bush 1994).<br />
Exogenous valine is a powerful plant growth inhibitor because it is a negative regulator<br />
of acetohy~ozyacid synthase (also known as acetolactate synthase), an enzyme in the<br />
valine biosynthetic pathway. Once down-regulated in the presence of excess valine,<br />
growth is inhibited because the cells starve for other amino acids, the precursors of which<br />
also pass through acetohy~oxyacid synthase. We were able to show that our lines are<br />
transport mutants versus regulation mutants because they are also resistant to azaserine,<br />
a toxic amino acid analogue that targets other enzymes in the cell. Thus, resistance to<br />
azaserine is consistent with loss of transport activity. We are currently attempting to<br />
identify the disrupted gene(s).<br />
One of the unexpected observations emerging from recent molecular descriptions<br />
of plant transporter genes is that there are large families of related porters that function<br />
in assimilate partitioning (Bush et al. 1996). Examples include the AHA family of protonpumping<br />
ATPases (Harper et al. 1990), the major facilitator superfamily of sugar transporters<br />
(Griffith et al. 1992; Bush et al. 1996), and the amino acid transporter families<br />
reviewed here. Although it is easy to imagine how these porters fulfill complementary<br />
functions in different organs and cells, it is clear that a major challenge in this field is<br />
to determine the unique contributions of the various transporters and, ultimately, to integrate<br />
their combined activities across the plant as a multicellular organism.
Given the central role played by plant amino acid symporters in nitrogen p~itioning<br />
into cells and between organs, and the complexity presented by the many transporters<br />
cloned to date, it is necessw to obtain basic information about their structure and regulation<br />
as the foundation for understanding their contributions to plant growth. Unfortunately,<br />
these kinds of investigations have only recently become possible with the cloning<br />
of the various transporters; therefore, very little has been published in this area. However,<br />
several laboratories have initiated projects investigating st~cture and function, and we<br />
anticipate significant progress in the near future.<br />
itiated a detailed investigation of N<br />
transporter family. We chose NAT<br />
sed, it transports many amino acid e co~on in the<br />
phloem-translocation stream, and because at least one member of this gene fdly appears<br />
to be expressed in every tissue of the plant. We are using an integrated approach<br />
with both biochemical and molecular strategies to learn more about the structure and<br />
function of this protein.<br />
The initial focus of our §tructural analysis has been establishing the membrane<br />
topology of ~ A This ~ is an , important question because our initial analysis suggested<br />
it does not contain 12 membrane-spanning helices as is commonly described for other<br />
metabolite translocators. We started by engineering a c-myc epitope on either the NH2-<br />
or COOH-termini of the expressed protein. We then used in vitro trans~ation, partial<br />
digestion with proteinase , and immuno~recipitation to determine the location of the<br />
NH2- and COOH-termini and to identify a family of oriented peptide fragments that we<br />
resolved on sodium dodecyl sulfate-polyacryldde gel electrophoresis (SDS-PAGE).<br />
These results showed that the ~H2-terminus is located inside the cell and the COOHterminus<br />
is outside. In addition, the lengths of the peptide fragments recovered after<br />
partial proteolysis allowed us to predict the location of protease-accessible cleavage sites<br />
between putative ~ansmembrane domains. We independently identified the location of<br />
the NH2- and COOH-termini using immuno~uorescence microscopy of NAT2 expressed<br />
in COS-1 cells. From the combined data, we proposed an 11 ~ansmembrane domain<br />
model, with the NH2-terminus in the cytoplasm and CO~H-terminus facing outside of<br />
cell (see Fig. 2; Chang and Bush 1997). This model of protein topology anchors our<br />
co~plement~ investigations of porter structure and function using site-directed and<br />
random mutagenesis.<br />
Previous biochemical investigations of ~E~C-dependent inactivation of the amino<br />
acid sympo~ers implicated a histidine residue in the reaction ~echanism (Li and Bush<br />
1990; see Fig. 3). An inspection of the ~ A gene family ~ identi~ed / two ~ histidine ~<br />
is-337, that are conserved in every transporter. Therefore, we used<br />
site-directed mutagenesis to change se residues and then we determined the effect of<br />
those changes on transport activity 0th residues are essential because the modified<br />
proteins were no longer active, owever, the change in is-337 destabiliz~ the protein,<br />
thereby complicating our inte~re~tion of that result. Several amino acid substitutions of<br />
His-47, on the other hand, blocked transport activity in the stable protein, thus demonstrating<br />
the significa~ce of this residue in the transport reaction (L. Chen and D. R. Bush,<br />
un~ublished data).<br />
A second approach we have employed to identify impo~nt &no acid residues<br />
and protein domains uses random mutagenesis and highly selective screens. The advan-
tage of this approach is that it does not require prior biochemical knowledge to identify<br />
residues for potential modification. ~ A ~ was randomly / ~ mutagenized ~ l in a mutator<br />
strain of E. coli and then thousands of modified versions of ~ A were trans- ~ /<br />
formed back into the yeast transport mutants. The yeast were then screened under new<br />
growth conditions that wild-type ~ A ~ does / not permit ~ ~ growth. l For example, we<br />
identified cells that acquired the ability to grow in the presence of transport competitors<br />
and others that grew on very low concentrations of histidine. This kind of positive selection<br />
identified mutant forms of the porter that exhibit new transport properties while<br />
eliminating those that simply lost transport activity. Although loss of function mutants<br />
might be revealing, far too many would inhibit transport activity for unspecific reasons,<br />
such as premature te~ination of the peptide. Thus far, we have identified important<br />
residues in TMDs 1,5,6,8, and 11. ~reliminary analysis of these results suggests TMDs<br />
1, 5, 6, and 8 may be involved in defining the substrate translocation pathway through<br />
this transporter (L. Chen and D. R. Bush, unpublished data).<br />
The amino acid symporters described here are excellent candidates for using biotechnological<br />
methods to modify the nutritional value of harvested tissues. For example, many<br />
cereals are poor sources of protein for animals because they are low t~ptophan, in lysine,<br />
and threonine (Bright and Shewry 1983; z and Larkins 1991 ; Galili 1995), and legumes<br />
are low in cysteine and methionine (Shewry et al. 1995). One strategy for modifying<br />
the nutrition^ value of these deficient crops would be to use targeted expression of<br />
high-affinity amino acid transporters to enhance the content of specific amino acids in<br />
harvested seed. For instance, targeted expression MTl in the phloem of mature leaf<br />
tissue may increase the amount of lysine and histidine transported per unit carbon. This<br />
would increase the lysine and histidine content in developing seeds. The success of this<br />
approach is dependent on several unknown variables, such as reciprocal increases in<br />
mesophyll synthesis and release as these amino acids are actively transported into the<br />
phloem. There is precedence for increased rates of synthesis under increased demand,<br />
and recent work in Heldt’s laboratory suggests the mesophyll pool may be dynamically<br />
linked to phloem loading (Mens et al. 1991 ; Winter et al. 1992; Lohaus et al. 1994) and,<br />
thus, active loading by a high-affinity transporter may shift the equation to increased<br />
synthesis. Other examples of potential engineering include targeted expression in stem<br />
tissue to capture amino acids passing by in the phloem, combining ove~roduction and<br />
targeted expression, or targeted expression to engineer uptake of novel xenobiotics.<br />
Our understanding of amino acid transport has progressed si~nificantly in recent<br />
years. The challenges that lay ahead focus on defining the coarse and fine-con~ol mechanisms<br />
that regulate that function of these amino acid transporters. This knowledge will<br />
be the fo~ndation of our understanding of resource allocation across the plant as a multicellular<br />
organism, and for developing novel strategies for improving crop yields and<br />
nu~tion~ value.<br />
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regulator of root gravitropism. Science, 273: 948-950.<br />
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by functional complementation of a yeast amino acid transport mutant. Proc, Natl, Acad.<br />
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related amino-acid transporters with differing substrate specificity in Arabidopsis thaliana.<br />
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3<br />
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Verbruggen, N., Borstlap, A. C., Jacobs, M., Van Montagu, M., and Messens, E. (1996). The razl<br />
mutant of Ara~idopsis thaliana lacks the activity of a high affinity amino acid transporter.<br />
Planta, 200: 247-253.<br />
Weston, K., Hall, J. L., and williams, L. E. (1995). Characte~zation of amino acid transport in<br />
Ricinus communis roots using isolated membrane vesicles. Planta, 196 166-173.<br />
Williams, L. E., Nelson, S, J., and Hall, J, L. (1990). Characte~zation of solute transport in plasma<br />
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Planta, 186 541-550.<br />
Williams, L. E., Bick, J. A., Neelam, A., Weston, K. N., and Hall, J, L, (1996). Biochemical and<br />
molecular characte~zation of sucrose and amino acid carriers in ~icinus communis. J. Exp.<br />
Bot., 47: 1211-1216.<br />
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a novel human gene homologous to the murine ecotropic retroviral receptor. Virology, 185:<br />
10-17.
David Rhodes<br />
Purdue Unjversj~, West ~fa~ette, Indiana<br />
~n~ve~si~ of M~sso~~i, Colu~bia, Missouri<br />
1. INT TI<br />
This chapter will consider the central role of amino acid metabolism in abiotic stress<br />
resistance of plants. We will attempt to ~ighlight progress made since the last comprehensive<br />
review of this field by Stewart and Larher (1980). Major emphasis will be placed<br />
on the role of amino acid metabolism in the synthesis of compatible osmolytes. This<br />
class of molecules includes certain amino acids (notably proline), quaternary ammonium<br />
compounds (e.g., glycinebetaine, prolinebetaine, P-alaninebetaine, and choline-0-sulfate),<br />
and the tertiary sulfonium compound 3-dim~thylsulfoniopropionate (DMSP), The<br />
quaternary ammonium compounds and DMSP are derived from amino acid precursors.<br />
These compounds share the property of being uncharged at neutral pH and are of high<br />
solubility in water (Ballantyne and Chamberlin 1994). Moreover, at high concen~ations<br />
they have little or no perturbing effect on macromolecule-solvent interactions (Yancey<br />
et al. 1982; Low 1985; Somero 1986; Timasheff 1993; Yancey 1994). Unlike perturbing<br />
solutes (such as inorganic ions) that readily enter the hydration sphere of proteins, favoring<br />
unfolding, compatible osmolytes tend to be excluded from the hydration sphere of<br />
proteins and stabilize folded protein structures (Low 1985). These compounds are<br />
thought to play a pivotal role in plant cytoplasmic osmotic adjustment in response to<br />
osmotic stresses (Wyn Jones et al. 1977).<br />
The synthesis of quate~ary ammonium and tertiary sulfonium compounds requires<br />
participation of the “activated methyl cycle’’ to provide methyl groups for the onium<br />
moieties of these compounds (Hanson et al. 1995; Bohnert and Jenson 1996). Therefore,<br />
attention is given to the role of sulfur amino acid metabolism, and in particular, the role<br />
of methionine and ~-adenosylmethionine (SAM) in the biosynthesis of these methylated<br />
onium compounds. The central position of SAM in stress metabolism in plants is further<br />
illustrated by briefly considering its role as a precursor of the hormone ethylene (Kende<br />
1983) and of the polyamines spermine and spermidine (Flores et ai. 1989). The role of<br />
sulfur amino acid metabolism in plant stress-resistance mechanisms is further explored<br />
in relation to the p~icipation of cysteine as a precursor of peptides (glutathione and<br />
319
phytochelatins) that chelate heavy metals (Rauser 1990, 1995) and facilitate detoxification<br />
of herbicides (Li et al. 1995; Kreuz et al. 1996) and active oxygen species (Alscher<br />
1989; Foyer et al. 1995; Smirnoff 1995).<br />
The amino acids alanine and y-aminobutyrate (CABA) accumulate markedly in<br />
response to anaerobic stress in plants (Streeter and Thompson 1972); these amino acids<br />
are specifically discussed in relation to anaerobic carbon metabolism, and intracellular<br />
pH regulation. Because it is now recognized that CABA synthesis is catalyzed by a<br />
C~-calmodulin-activated enzyme, glutamate decarboxylase (Snedden et al. 1995), we<br />
will consider the specific role of GfU3A connecting intermediary amino acid metabolism<br />
to perturbations of cytoplasmic Ca" concentrations induced by stress.<br />
Because proline is treated in a separate chapter in this volume (see Chap. 8), and has<br />
been the subject of numerous reviews over the last 20 years (see, e.g., Stewart and Larher<br />
1980; Thompson 1980; Stewart 1981; Hanson and Hitz 1982; Csonka and Baich 1983;<br />
Rhodes 1987; Delauney and Verma 1993; Samaras et al. 1995), including a comprehensive<br />
recent review (Hare and Cress 1997), our treatment of this amino acid will be brief.<br />
Proline accumulation represents a common metabolic responses of higher plants to water<br />
deficits and salinity stress. This highly water-soluble imino acid is accumulated to osmotically<br />
significant levels by leaves of many halophytic higher-plant species grown in saline<br />
environments (Stewart and Lee 1974; Treichel 1975; Briens and Larher 1982), in leaf<br />
tissues and shoot apical meristems of plants experiencing water stress (Barnett and<br />
Naylor 1966; Boggess et al. 1976; Jones et al. 1980), in desiccating pollen (Hong-qi et<br />
al. 1982; Lansac et al. 1996), and in cultured plant cells adapted to water stress (Handa<br />
et al. 1986; Rhodes et al. 1986), or NaCl stress (Katz and Tal 1980; Treichel 1986;<br />
Binzel et al. 1987; Rhodes and Handa 1989; Thomas et al. 1992). In the apical millimeter<br />
of maize roots, proline represents a major solute, reaching concentrations of 120 mM in<br />
roots growing at a water potential of -1.6 MPa (Voetberg and Sharp, 1991). The accurnulated<br />
proline accounts for a significant fraction ("50%) of the osmotic adjustment in this<br />
region (Voetberg and Sharp 1991). Proline accumulation in maize root apical meristems<br />
in response to water deficits involves increased proline deposition to the growing region<br />
(Voetberg and Sharp 1991) and requires abscisic acid (ABA; Ober and Sharp 1994;<br />
Sharp et al. 1994).<br />
At high concentrations, proline protects membranes and proteins against the adverse<br />
effects of inorganic ions and temperature extremes (reviewed in Samaras et al.<br />
1995). Proline may also function as a hydroxyl radical scavenger (Smirnoff and Cumbes<br />
1989). Its synthesis is implicated as a mechanism of alleviating cytoplasmic acidosis,<br />
and it may maintain NADP+/NAPH ratios at values compatible with metabolism (Hare<br />
and Cress, 1997). Rapid catabolism of proline after relief of stress may provide reducing<br />
equivalentsthatsupportmitochondrialoxidativephosphorylationandthegeneration of<br />
ATP for recovery from stress and repair of stress-induced damage (Hare and Cress, 1997).<br />
In vivo-labeling studies with 14C-labeled precursors (Moms et al. 1969; Oaks et al.<br />
1970; Boggess et al. 1976), ["CC]glutamate (Hayser et al. 1989a,b) or ''NH~/''NO~<br />
(Rhodes et al. 1986; Rhodes and Handa 1989) suggest that glutamate is a major precursor
of osmotic stress-induced proline accumulation in plants. Osmotic stress results in an<br />
increase of proline biosynthesis rate (Boggess et al. 1976; Stewart, 1981; Rhodes et al.<br />
1986; Rhodes and Handa 1989). Proline accumulation involves induction of enzymes of<br />
proline biosynthesis (Peng et al. 1996; and see Chap. 8), possibly coupled with a relaxation<br />
of proline feedback-inhibition control of the pathway (Boggess et al. 1976; Stewart<br />
1981), decreased proline oxidation to glutamate (Stewart et al. 1977; Stewart and Boggess<br />
1978; Wuang and Cavalieri 1979; Sells and Koeppe 1981; Elthon and Stewart 1982),<br />
mediated at least partly by down-regulation of proline dehydrogenase (Kiyosue et al.<br />
1996, Peng et al. 1996), decreased utilization of proline in protein synthesis (Boggess<br />
and Stewart 1980; Stewart 1981), and enhanced protein turnover (Fukutoku and Yamada<br />
1984). Water deficits induce dramatic increases in the proline concentration of phloem<br />
sap in alfalfa (Girousse et al. 1996), suggesting that increased deposition of proline at<br />
the root apex in water-stressed plants (e,g., Voetberg and Sharp 1991) could occur partly<br />
by phloem transport of proline (Girousse et al. 1996). A proline transporter gene ProT2<br />
is strongly induced by water and salt stress in Arnbidopsis t~~Ziann (Rentsch et al. 1996).<br />
In bacteria, proline synthesis from glutamate is catalyzed by three enzymes: y-<br />
glutamyl kinase (GK; EC 2.7.2.1 I), y-glutamyl phosphate reductase (GPR; glutamic semialdehyde<br />
dehydrogenase; EC 1.2.1 Al), and A1-pyrroline-5-carboxylate reductase (PSCR;<br />
EC 1.5,1.2), encoded by genes pro& proA, and yroC, respectively (Adams and Frank<br />
1980; Csonka and Baich 1983). In higher plants it is now clear that a similar pathway<br />
operates, with the exception that GK and GPR are embodied in one bifunctional polypeptide,<br />
A1-py~oline-5-carboxylate synthetase (PSCS), encoded by a single gene (Hu et al.<br />
1992; Fig. 1). The prevailing evidence is that both PSCS and PSCR are localized in the<br />
cytoplasm (Hare and Cress, 1997). The GK activity of the bifunctional PSCS is inhibited<br />
by proline (Nu et al. 1992). PSCS gene expression is induced by desiccation, salinity<br />
stress, and abscisic acid (ABA; Hu et al. 1992; Yoshiba et al. 1995; Peng et ai. 1996;<br />
Savourk et al. 1997). P5CR transcripts also increase in abundance in response to osmotic<br />
stress (Delauney and Verma 1990; Williamson and Slocum, 1992; Verbruggen et al.<br />
1993). Savourd et al. (1997) suggest that the expression of the proline biosynthetic genes<br />
are dependent on at least two signal transduction cascades: one triggered by exogenously<br />
applied ABA in the absence of stress, and the other triggered by cold and osmotic stress<br />
independently of exogenously applied ,ABA.<br />
Recently, transgenic tobacco plants overexpressing PSCS have been obtained, and<br />
these appear to have increased resistance to both water deficits and salinity stress (Kishor<br />
et al. 1995). However, insufficient data are available to conclude with certainty that<br />
proline accumulation in these transgenic plants contributes to their enhanced stress resistance<br />
by osmotic adjustment, or by some other mechanisms (Sharp et al. 1996). As discussed<br />
by Hare and Cress (1997), the controversial data reported for the water relations<br />
of these transgenic plants emphasize the need to investigate nonosmotic explanations for<br />
the phenotypes observed.<br />
A decrease in proline oxidation rate can contribute to net proline accumulation<br />
during drought and salinity stress. Proline is oxidized to PSC by proline dehydrogenase<br />
(EC 1.5.99.8) and the resulting P5C is further oxidized to glutamate by P5C dehydrogenase<br />
(EC 1.5.1.12; see Fig. 1). Both enzymes are localized in the mitochondrion (reviewed<br />
in Hare and Cress, 1997). A cDNA-encoding proline dehydrogenase (PDH) has<br />
recently been cloned from Arn~i~opsis and is induced by proline under nonstressed conditions,<br />
but strongly repressed in response to osmotic stress (Kiyosue et al. 1996). It<br />
appears that osmotic stress overrides proline induction of proline dehydrogenase gene
OH glutamate +<br />
8<br />
0 0<br />
*.<br />
I<br />
*<br />
*.<br />
NADH<br />
+ H+<br />
NADS<br />
* *<br />
.I<br />
0 .<br />
.*.*<br />
PSCR<br />
H O<br />
proline -<br />
P<br />
proline<br />
FADH2<br />
+ H20<br />
FAD +<br />
c<br />
+ 02<br />
iigure 1 Pathways of synthesis and oxidation of proline in higher plants: GK, y-glutamyl kinase;<br />
GPR, y-glutamylphosphate reductase; PSC, A1-pyrroline-5-~arboxylate; PSCS, A'-pyrroline-S-carboxylate<br />
synthetase; PSCR, A'-pyrroline-5-carboxylate reductase; PDH, proline dehydrogenase;<br />
PSCDH, A1-pyrroline-S-c~boxylate dehydrogenase. The dotted line denotes feedback inhibition of<br />
the GK activity of PSCS by the end product, proline,
expression (Kiyosue et al. 1996). The reciprocal regulation of PSCS and PD<br />
count for the rapid accumulation of proline in response to osmotic stress and its rapid<br />
catabolism after stress relief (Peng et al. 1996). To our knowledge the gene encoding<br />
P5C dehydrogenase has not yet been cloned from higher plants, but the enzyme has<br />
recently been extensively purified and characterized (Forlani et al. 1997).<br />
Hare and Cress (1997) argue that the different subcellular localizations of proline<br />
biosynthesis (cytoplasm) and oxidation (mitochondrion), the NADPH cofactor preference<br />
of the biosynthetic enzymes, and NADH cofactor preference for the proline oxidation<br />
pathway, would enable proline biosynthesis to enhance activity of the cytoplasmic oxidative<br />
pentose phosphate pathway and provide a mechanism of interconversion of the phosphorylated<br />
and nonphosphorylated pools of pyridine nucleotide cofactors. They suggest<br />
that the osmoprotective effects of proline accumulation may be of secondary impo~ance<br />
to the associated metabolic implications of proline synthesis and degradation. Further<br />
investigations of transgenic plants, engineered for altered proline synthesis or catabolism,<br />
clearly need to consider not only the consequences on absolute proline level and the<br />
biophysical effects exerted by elevated cytoplasmic proline concentrations, but also the<br />
fluxes to and from the proline pool (Hare and Cress 1997).<br />
In growing tissue, the proline deposition rate will be equal to the combined rates<br />
of proline synthesis, proline release from protein, and proline import, minus the combined<br />
rates of proline catabolism, proline export, proline utilization in protein synthesis,<br />
and the rate of pool dilution caused by water uptake during growth (Rhodes and Handa<br />
1989; Voetberg and Sharp 1991). The latter, in turn, will be determined by fundamental<br />
plant-water relations of the growing region, including water potential, solute potential,<br />
yield threshold, turgor, and potential difference between the xylem and growing cells<br />
(Nonami et al. 1997), as well as the levels of growth inhibitory metabolites such ABA, as<br />
for which synthesis or compartmentation may respond rapidly to changes in turgor or<br />
metabolism (e.g., intracellular pH; Rhodes 1987). The feedback loops in this system are<br />
clearly complex, with ABA and osmotic signals altering expression of genes encoding<br />
proline biosynthesis enzymes, proline and osmotic signals altering expression of PD<br />
proline feedback inhibiting its own synthesis, and proline itself (if accumulated to sufficiently<br />
high levels) contributing to solute potential and, hence, turgor and growth maintenance.<br />
The full promise of transgenic plants engineered for proline metabolism toward<br />
an understanding of proline’s role(s) in stress resistance will likely not be realized until<br />
attention is given to the growing regions as well as mature tissues (cf., Kishor et al.<br />
1995). The consideration of growing and nongrowing tissues and the fluxes between<br />
them will be essential in testing the intriguing scheme, recently proposed by Hare and<br />
Cress (1997), in which proline and PSC might act as an intercellu1~-signaling system.<br />
In this scheme, proline produced from P5C in an “effector” tissue is transported by the<br />
phloem to a “target” tissue, characterized by a high-energy requirement, where proline<br />
degradation generates reducing equivalents needed to drive the tricarboxylic acid (TCA)<br />
cycle activity. The PSC or glutamate generated may then be translocated back to the<br />
effector tissue where conversion to proline regenerates the NADP’ needed to prime the<br />
oxidative pentose phosphate pathway.<br />
Stress-hypersensitive mutants of higher plants that exhibit disturbed proline metabolism<br />
(e.g., the sosl mutant of Ar~~i~o~sis; Liu and Zhu 1997) can contribute significantly<br />
to the elucidation of thesignalsto which proline accumulation may respond.<br />
However, we urge caution in concluding from such mutants that “Pro accumulation is a<br />
symptom of stress injury rather than an indicator of stress tolerance” (Liu and Zhu 1997).
324 ~ho~es et a!.<br />
111. IN0 ACIDS AS PREC<br />
Y AMMONIUM COMP<br />
OMPATIBLE OSMOLYTES<br />
The role of quaternary ammonium compounds as compatible solutes has been extensively<br />
reviewed (Wyn Jones and Storey 1981 ; Anthoni et al. 199 1 ; Rhodes and Hanson 1993;<br />
Gorham 1995). These zwitterionic compounds include glycinebetaine, choline-0-sulfate,<br />
P-alaninebetaine, prolinebetaine, and hydroxyprolinebetaine. They possess a fully<br />
methyl-substituted nitrogen atom (creating a permanent positive charge on the N moiety)<br />
and a negatively charged carboxyl group (in the case of betaines) or sulfate group (in<br />
the case of choline-0-sulfate). The function of these compounds as osmoprotectants is<br />
illustrated by their stimulation of the growth of Escherichia coli or ~aZ~oneZZa ~ ~ ~ i ~<br />
rium in saline medium (Strom et al. 1983; Hanson et al. 1991, 1994a). In these assays,<br />
glycinebetaine, P-alaninebetaine, prolinebetaine, and choline-0-sulfate at 1-mM concentrations<br />
are more effective than 1 mM of proline (Hanson et al. 1991, 1994a). Choline,<br />
an alcohol without a negatively charged group, is not an osmoprotectant per se. Although<br />
it stimulates the growth of wild-type E. coli in a saline medium, it must be oxidized to<br />
glycinebetaine to function as an osmoprotectant. Thus, betA mutants of E. coli, defective<br />
in choline oxidation, are unable to use choline as an osmoprotectant (Hanson et al. 1991).<br />
Similar to proline, glycinebetaine and structurally related compounds have stabilizing<br />
effects on proteins and membranes (Rhodes and Hanson 1993), and are implicated in<br />
frost protection (Kishitani et al. 1994; Nomura et al. 199Sb).<br />
In higher plants, choline is derived from the amino acid serine, presumably by decarboxylation<br />
of serine to ethanolamine (Fig. 2). The NH?-moiety of ethanolamine is then N-<br />
methylated to (CH&-P- at the level of either free ethanolamine bases, O-phosphorylethanolamine<br />
bases, or 0-phosphatidylethanolamine bases, catalyzed by S-adenosylmethionine<br />
(SAM)-dependent N-methyltransferases (Rhodes and Hanson 1993; Corham<br />
1995; Hanson et .al. 1995). Datko and Mudd (1988a,b) have proposed phospho~lethanolamine<br />
(PE) as a common co~itting step in the synthesis of choline moieties in plants,<br />
implicating ethanolamine kinase (EC 2.7.1.82) as a key enzyme in this pathway (see<br />
Fig. 2).<br />
In glycinebetaine-accumulating chenopods, the main pathway of choline synthesis<br />
appears to be by the phosphoryl-bases: PE, phospho~lmonomethylethanolamine (PMME),<br />
phosphoryldimethylethanolamine (PDME), and phosphorylcholine (PC) (Hanson and<br />
Rhodes 1983; Hanson et al. 1995; see Fig. 2). Choline is then liberated from PC by a<br />
phosphorylcholine phosphatase (Rhodes and Hanson 1993; Hanson et al. 1995; see<br />
Fig. 2).<br />
In grasses, PC is incorporated into phosphatidylcholine from which choline is then<br />
released (Giddings and Hanson 1982; Hitz et al.1.991),presumably by the action of<br />
phospholipase-D (Rhodes and Hanson 1993).<br />
Choline down-regulates its own synthesis at the level of the PMME and PDME N-<br />
methyltransferases (Mudd and Datko 1989a,b). These enzymes are induced by salinity<br />
stress in spinach (Weretilnyk and Summers 1992; Hanson et al. 1995; Weretilnyk et al.<br />
1995). PC may feedback-inhibit its own synthesis in sugar beet (Hanson and Rhodes<br />
1983).
6iotic Stress ~esista~ce<br />
ethanolamine @A)<br />
.<br />
e<br />
.<br />
0<br />
transferase<br />
0 SAHC<br />
0<br />
*<br />
. -0<br />
e N-<br />
H<br />
O<br />
phospho~lmonometh~lethanolamine<br />
(PIMME)<br />
phospho~ldimethylethanolamine<br />
(PDME)<br />
phospho~lcholine (PC)<br />
+-<br />
choline<br />
Figure 2 Pathway of choline synthesis in the Chenopodiaceae: The dotted line denotes feedback<br />
regulation of the first N-methyltransferase by choline.<br />
Glycinebetaine is synthesized from choline in a two-step oxidation catalyzed by a ferredoxin<br />
(Fd)-dependent choline rnonooxygenase (CMO) and a betaine aldehyde dehydrogenase<br />
(BADH; EC 1.2.1.8) with a strong preference for NAD' (Rhodes and Hanson 1993;<br />
Fig. 3).
,-,. betaine ~dehyde<br />
+ H20 - H20<br />
0 - betaine aldehyde<br />
NAD+<br />
NADH + Elt<br />
~~ycinebetai~~<br />
Pathway of glycinebetaine synthesis in the Chenopo~iacea~: Fd, fe~e~oxin.<br />
th enzymes are predominantly localized in the chloroplast stroma of chenopods<br />
et al. 1986, 1988; Brouquisse et al. 1989), although in spinach leaves approximately<br />
10% of the BADH may exist as a cytosolic isofom (~eretilnyk and Hanson<br />
1988). ~lycinebetaine is also localized predominantly in the chloroplasts of salinized<br />
spinach leaves, where it provides osmotic adjustment (Robinson and Jones 1986). Oxygen-1<br />
8 tracer studies confirm that CMO uses molecular oxygen as a substrate (Lema et<br />
al. 1988).<br />
Both CMO (Burnet et al. 1995) and BADH (~eretilnyk and Hanson 1989) have<br />
been purified to homogeneity from spinach. Complementary DNAs encoding BADH<br />
n isolated from spinach (Weretilnyk and Hanson 1990) and sugar beet (McCue<br />
son 1992). Salinity stress leads to a two- to threefold increase of CMO and<br />
ene expression, and concomitant increases<br />
enzyme level (Hanson et al. 1995).<br />
ADH has an unusual chloroplast-transit target sequence (Rathinasabapathi<br />
ansgenic tobacco plants overexpressing chloroplast-localized BADH have<br />
been obtained (Rathinasabapathi et al. 1994). However, these are unable to accumulate<br />
glycinebetaine in the absence of exogenously supplied betaine aldehyde because tobacco<br />
lacks C ~ (~athinasabapathi<br />
O<br />
et al. 1994). A cDNA encoding CMO has recently been<br />
cloned from spinach (Rathinasabapathi et al. 1997). The cDNA sequence confirms that<br />
is an Fe-S protein, and that CMO has a typical chloroplast-transit peptide sequence<br />
inasabapathi et al, 1997). Expression of chloropIast-localized CMO in BADH-con-
taining transgenic tobacco plants will provide a critical test of the adaptive value of<br />
glycinebetaine in salinity and drought stress resistance.<br />
It should be cautioned, however, that BADH may be a general aldehyde dehydrogenase,<br />
acting on other aldehyde substrates in addition to betaine aldehyde (Trossat et al.<br />
1997).BADHwilluse 3-dimethylsulfoniopropionaldehyde, an intermediate in DMSP<br />
synthesis (see Sec. 1V.B) and certain aldehydes (3-aminopropionaldehyde and 4-aminobutyraldehyde)<br />
involved in polya~ne metabolism (Trossat et al, 1997). Transgenic<br />
plants engineered for BADH expression (Holmstrom et al. 1994; ~athinasabapathi et al.<br />
1994) may possibly exhibit phenotypes related to perturbed polyamine metabolism, in<br />
addition to phenotypes attributable to alterations in the glycinebetaine synthesis pathway<br />
(Trossat et al. 1997). The multiple substrate specificities of BADH may explain the<br />
occurrence of BADH in plants that do not accumulate glycinebetaine (Ishitani et al.<br />
1993; Weretilnyk et al. 1989), and in organs of glycinebetaine-accumulating plants that<br />
do not contain glycinebetaine (e.g., roots of cereals; Ishitani et al. 1995).<br />
In monocotylednous plants, recent studies suggest that BADH is localized in peroxisomes<br />
(Nakamura et al. 1997). All monocotyledenous BADHs have a COOH-terminal<br />
tripeptide (SKI.,) that is a signal for targeting preproteins to microbodies (Nakamura et<br />
al. 1997). This raises the question of whether the subcellular localization of the choline<br />
oxidation pathway is the same in grasses as in chenopods.<br />
In bacteria, such as E. coli, choline is an osmoprotectant only because it is oxidized<br />
to glycinebetaine. Mutants in which choline fails to serve as an osmoprotectant prove to<br />
have defects in choline transport (betT), choline oxidation (betA), betaine aldehyde oxidation<br />
(betB), or a regulatory locus (betl) encoding a repressor protein (Lamark et al, 1991,<br />
1996). The entire E. coli bet gene cluster has been introduced into the freshwater cyanobacterium<br />
~y~ec~ococc~s and has conferred glycinebetaine accumulation and increased<br />
salt tolerance, partly owing to stabilization of photosystem I1 (PS-11; Nomura et al.<br />
1995a). However, this increased salt tolerance appears to depend on the presence of an<br />
exogenous supply of choline. Transgenic plants expressing the E. coli betA gene encoding<br />
choline dehydrogenase putatively capable of oxidizing both choline and betaine aldehyde,arereportedtohaveincreasedsalttolerance<br />
(Liliusetal.1996).However,no<br />
evidence was presented that these transgenic plants actually accumulated glycinebetaine.<br />
In maize, several naturally occurring glycinebetaine-de~cient inbred lines have<br />
been identified (Brunk et al. 1989). These lack the ability to accumulate glycinebetaine<br />
in leaf tissue in response to either salinity stress or water deficits owing to a single<br />
recessive gene (betl), which has been mapped to the short arm of chromosome 3 near<br />
the centromere (Rhodes et al. 1993; rang et al. 1995). Glycinebetaine deficiency in<br />
maize is associated with an inability to oxidize ['4C]choline to ~'4C]glycinebetaine, but<br />
an unimpaired ability to oxidize [2H3]betaine aldehyde to [2H3]glycinebetaine, suggesting<br />
a lesion at the CMO step in the biosynthetic pathway (Lema et al. 1991). Homozygous<br />
glycinebetaine-de~cient maize lines appear to be more salt-sensitive than near-isogenic<br />
homozygous glycinebetaine-containing lines (Saneoka et al. 1995), and they exhibit<br />
greater membrane injury and damage to PS-I1 in response to heat stress (Yang et al.<br />
1996). ~lycinebetaine-deficient maize lines exhibit a signi~cantly elevated pool of free<br />
choline; however, choline does not accumulate to levels equal to the glycinebetaine level<br />
of glycinebetaine-containing lines, suggesting that choline must down-regulate its own<br />
synthesis (Yang et al. 1995). Consistent with this, glycinebeta~ne-deficient lines of maize<br />
exhibit an elevated level of serine in comparison with homozygous glycinebetaine-containing<br />
lines (Yang et al. 1995).
Clycinebetaine is thought to be the archetypal betaine in the plant kingdom (Weretilnyk<br />
et al. 1989; Hanson and Burnet 1994; Hanson et al. 1994a). Some species have apparently<br />
lost the capacity to accumulate large quantities of this compound, and others have<br />
evolved the capacity to synthesize alternative quaternary a~onium compounds, including<br />
choline- sulfate, p-alaninebetaine, or prolinebetaine. This is particularly well<br />
illustrated in a consideration of the quaternary ammonium compounds accumulated by<br />
membersofthePlumbaginaceae (Hansonetal.1994a).Ancestralsubfamilies of the<br />
Plumbaginaceae are glycinebetaine-accumulators (Hanson et al. 1994a). In more advanced<br />
subfamilies, however, glycinebetaine has been replaced in part or in whole by p-<br />
alaninebetaine or prolinebet~ne (Hanson et al. 1994a). In addition, all members of the<br />
Plumba~inaceae accumulate choline-0-sulfate. This compound is synthesized from choline<br />
by a salt stress-inducible cholinesulfotransferase (EC 2.8.2.6; Rivoal and Hanson<br />
1994b; Fig. 4).<br />
The adaptive significance of choline-0-sulfate in the Plumbaginac~ae is thought to<br />
reside in the fact that salt glands of these species can excrete chloride, but not sulfate;<br />
thus, conjugation of sulfate with choline is proposed to be a mechanism of sulfate detoxification,<br />
converting a normally deleterious anion to a compatible osmolyte (Hanson et<br />
al. 1994a). Because choline- sulfate synthesis competes with glycinebetaine synthesis<br />
for available choline, this may have cont~buted to the evolution of alternative betaine<br />
biosynthetic pathways (p-alaninebetaine or prolinebetaine) that draw on substrates other<br />
than choline (Hanson et al. 1994a).<br />
p-Alaninebetaine is confined to species of the Plurnbaginaceae (Hanson et ai. 1994a). It<br />
is synthesized by direct ~-methylation of the amino acid p-alanine, presumably catalyzed<br />
by SAM-dependent ~-methyltransferases (Hanson et al. 1991, 1994a; Fig. 5). In contrast<br />
with glycinebetaine synthesis, p-alaninebetaine does not require oxygen (Hanson et al.<br />
1994a). It is possible, therefore, that this pathway represents an adaptation to anoxic<br />
saline environments (Hanson et al. 1994a). The ~-alaninebetaine biosynthetic pathway is<br />
HO<br />
I<br />
3-phosp~oadenosi~~-<br />
5'phosphosulfate (PAPS)<br />
3-phosphoadenosine-<br />
0 5 'phosphate (PAP)<br />
- G~oli~e-0-sulfate<br />
Pathway of choline-0-sulfate synthesis in the Plumbaginaceae.
0<br />
ll<br />
HO NW, @-alanine<br />
SAM<br />
SAHG<br />
0<br />
"0 - @-~~inebetaine<br />
ure 5 Putative pathway of P-alaninebetaine synthesis in the Plumbaginaceae.<br />
active in both leaves and roots of p-alaninebetaine-accumulating Li~oniu~ species<br />
(Hanson et al. 1991 ). The precise origin of p-alanine remains obscure. This amino acid<br />
apparently is not derived from decarboxylation of aspartate in Li~o~2i~~<br />
species (Hanson<br />
and Burnet 1994).<br />
In some higher-plant species (notably members of the Labiateae, Asteraceae, Fabaceae,<br />
Bataceae, Rutaceae, Plumbaginaceae, and Capparaceae), proline can be metabolized to<br />
prolinebetaine (stachydrine; Naidu et al. 1987, 1992; Wood et al. 1991; Hanson et al.<br />
1994a; Bonham et al. 1995; Gorham 1995; Blunden et ai. 1996; McLean et al. 1996;<br />
Nolte et al. 1997). The synthesis of prolinebetaine is thought to involve N-methylation of<br />
proline by SAM[-dependent methyltransferases through the intermediate N-methylproline<br />
(hygric acid; Robertson and Marion, 1960; Essery et al. 1962), but the enzyme(s) have<br />
not yet been characterized (Fig. 6). Hydroxyprolinebetaine often (but not always) accumulates<br />
with prolinebetaine in these prolinebetaine-accu~ulating species (Wood et al.<br />
1991; Hanson et al. 1994a; Noltet al. 1997). The proline and N-methylproline hydroxylase(s)<br />
putatively involved in hydroxyproIinebetaine synthesis have not yet been characterized.<br />
Comparisons of related species with different capacities for accumulation of prolinebetaine<br />
and hydroxyprolinebetaine suggest that the free-proline pool size is inversely<br />
related to the total prolinebetaine plus hydroxyprolinebetaine pool size (Hanson et al.<br />
1994a; Nolte et al. 1997). Prolinebetaine is a more potent osmoprotectant than proline<br />
(Hanson et al. 1994a). Conversion of proline to prolinebetaine, therefore, may enhance<br />
osmotic stress resistance. In principle, increased flux of proline to N-methylated derivatives<br />
should alleviate P5CS from feedback regulation by proline by decreasing the pool
33 o~es et a/.<br />
HO<br />
. P5C P5C<br />
.<br />
.<br />
0<br />
sA.Hc<br />
SAM<br />
SAHC<br />
6 Putative pathway of prolinebetaine biosynthesis.<br />
size of free proline (See Sec. 11; and also Chap. 8), simultaneously restricting proline<br />
oxidation and, hence, osmolyte catabolism (Samaras et al. 1995). One isomer of hydroxyprolinebetaine<br />
(~~~~~-4-hydroxy-~-prolinebetaine) is a potent inhibitor of animal acetylcholinesterase<br />
(Friess et al. 1957), raising the possibility that the ac~umulation of this<br />
compound may serve a dual function: as a compatible osmolyte and an antifeedant,<br />
affording protection against herbivores (Hanson and Burnet 1994). In contrast, proline is<br />
a stimulant of insect herbivory (Haglund 1980).<br />
Pipecolatebetaine is accumulated together with prolinebetaine in certain species; notably<br />
~e~icago and Ac~i~Zea (Wood et al. 1991; Bonham et al. 1995). Pipecolic acid is often<br />
accumulated to high levels by members of the Fabaceae (Stewart and Larher 1980; Rosenthal<br />
1982), and indeed, free pipecolic acid is a major constituent of the free amino<br />
acid pool of ~e~icugo (Wood et al. 1991). The putative pathway of pipecolatebetaine
SAM<br />
SAHC<br />
SAM<br />
SAHC<br />
ure 7 Putative pathway of pip~colatebetaine biosynthesis.<br />
synthesis is illustrated in Figure 7. It is unknown if the proline and N-methylproline N-<br />
methyltransferases have dual substrate specificity, and if they are also capable of N-<br />
methylating pipecolic acid and N-methylpipecol~c acid, respectively. Pipecolic acid is<br />
derived from lysine catabolism by ~-aminoadipic semialdehyde (Gon~alves-Butruille et<br />
al. 1996), which spontaneously cyclizes to ~'-pipe~dine-6-c~boxylate. The latter can<br />
then be reduced to pipecolic acid in a reaction analogous with that catalyzed by P5CR<br />
(Stewart and Larher 1980; Rosenthal 1982; Galili 1995).<br />
tiv<br />
The synthesis of the foregoing ~uatern~y a~monium compounds is dependent on the<br />
activity of SAM-dependent ~-methyltransferases for N-methylation of ethanolamine<br />
bases, p-alanine, proline, and pipecolic acid. This implies that osmolyte accumulation is<br />
critically dependent on the supply of SAM as a methyl donor (Hanson et al. 1995;<br />
Bohnert and Jenson 1996). When SAM is used as a methyl donor in ~-~ethyl~ansferase<br />
reaction, the resulting ~-adenosylhomocysteine (SAHC) is recycled to homocysteine<br />
(HC) by adenosylhomoc~steinase (EC 3.3.1.1). HC can then accept a methyl group from<br />
the ~5-~ethy1-tetrahy~ofolate (~~-m~thyl-THF) pool to regenerate methionine by the<br />
action of methionine synthase (EC 2.1.1.14) and, thereby, SAM by the action of S<br />
synthetase (methionine adenosyltransferase; EC 2.5.1.6) (Giovanelli et al. 1980; Fig. 8).<br />
The foregoing cycle is known as the act~v~te~ ~et~yl cycle (Bohnert and Jenson
T€€F<br />
ethionine<br />
TEE<br />
~o~ocysteine<br />
Ns-me~yl-THF (CH,-THF)<br />
FAD<br />
FADH,<br />
glycine NH, + CO, glycine serine<br />
N5,Nl0-~ethylene-T~F<br />
..<br />
N5,~l0-methe~yl-~HF<br />
formate<br />
+ THE:<br />
Alternative pathways of synthesis of ~5-methyl-tetr~ydrofolate as methyl donor for<br />
methionine synthesis from homocysteine: GDC, glycine decarboxylase; SHMT, serine hydroxymethyltransferase.<br />
hydrolase (EC 3.5.4.9), and ~5,~10-methenyl-THF dehydrogenase (EC 1.5.1 S ) occur on<br />
a single polypeptide called C1-THF synthase (Prabhu et al. 1996). In plants, however,<br />
the synthetase activity occurs on a protein moiety separate from that of the cyclohydrolase<br />
and reductase, which occur on a bifunctional protein (Prabhu et al. 1996). Because<br />
the latter reactions are reversible, this allows an equilibrium between the pools of Cl<br />
units at the formyl and methylene levels of oxidation (Cossins 1987). Therefore, serine,<br />
glycine, and formate can serve as the precursors for methionine synthesis (Cossins 1987;<br />
see Fig. 9). [13CG]Formate is metabolized to p-['3C]serine in Ar~bi~opsis in a manner<br />
consistent with C1-THF synthase and SHMT activities interacting with a common THF<br />
pool (Prabhu et al. 1996). Formate is a very effective precursor of the methyl groups of
choline and glycinebetaine in glycinebetaine-accumulating plant species (Delwiche and<br />
Bregoff 1958; Hitz et al. 1981). Formate can be derived from nonenzymatic Hz02-mediated<br />
decarboxylation of glyoxylate, although Kleczkowski and Givan (1988) question<br />
whether this pathway is of physiological significance.<br />
The sulfonium analogue of P-alaninebetaine, 3-dimethylsulfoniopropionate (DMSP), is<br />
synthesized and accumulated to osmotically significant levels in a few higher-p~ant species<br />
from diverse families; certain species of the Asteraceae (most notably WoZZasto~ia<br />
b$ora), and Poaceae (some [but not all] species of S~arti~ and Sacc~ar~~)<br />
(Paquet et<br />
al. 1994; J.ames et al. 1995b), and by many marine algae (see, e.g., Blunden and Gordon<br />
1986; Dickson and Kirst 1986; Gage et al. 1997). DMSP is as effective as glycinebetaine<br />
as an osmoprotectant for E. coli (Paquet et al. 1994), and is noted to be a potent cryoprotectant<br />
(~ishiguchi and Somero 1992; Karsten et al. 1996).<br />
In ~oZZaston~ff, DMSP is synthesized by methylation of methionine to S-methylmethionine<br />
(SMM) in the cytosol (Hanson et al. 1994b; James et al, 1995a,b), transport of<br />
SMM into the chloroplast, dec~boxylation and deamination of SMM to DMSP-aldehyde<br />
(James et al. 1995b), and oxidation of the aldehyde to DMSP in the chloroplast (Trossat<br />
et al. 1996; Fig. 30). The first enzyme of the pathway, SA~~L-methionine methylt transferase<br />
(EC 2.1.1.12) has been purified from WoZlastoni~ leaves (James et al. 1995a).<br />
The final step in the pathway is catalyzed by a chloroplast-localized DMSP-aldehyde<br />
dehydrogenase, which may be identical with the BADH involved in glycinebetaine synthesis<br />
(James et al. 1995b; Trossat et al. 1996, 1997; Vojtechovd et al. 1997). The enzymes<br />
responsible for metabolism of SMM to DMSP-aldehyde are not yet known.<br />
In marine algae, DMSP synthesis occurs by a route that is entirely different from<br />
that in higher plants (Gage et al. 1997; Fig. 11). Methionine is first transaminated to 4-<br />
methylthio-2-oxobutyrate (MTOB), which is then reduced to 4-methylthio-~-hy~oxybutyrate<br />
(MTHB). S-Methylation of MTHB then yields 4-dimethylsulfonio-2-hydroxybutyrate<br />
(DMSWB), which is subs~uently oxidatively decarboxylated to DMSP (Gage et al.<br />
1997; see Fig. 11).<br />
Because DMSP does not contain nitrogen, the accumulation of DMSP could offer<br />
advantages over accumulation of nitrogenous onium compounds for osmotic adjustment<br />
in N-li~ting environments; N-deficiency increases DMSP levels ~oZZasto~i~<br />
in (Wanson<br />
et al. 1994b) and marine algae (Gage et al. 1997). The enzyme DMSP lyase generates<br />
the gas dimethyl sulfide (DMS) and the potentially toxic compound, acrylate, from<br />
DMSP. Acrylate generated from DMSP may play a role in defense against herbivores<br />
(Wolfe et al. 1997).<br />
S-Adenosylmethionine serves as a precursor of the plant hormone ethylene, implicated<br />
in the control of numerous developmental processes (Lieberman 1979; Yang and Hoffman<br />
1984; Yang 1989; Kende 1993). SAM is converted to the amino acid 1 -aminocyclopropane-1-carboxylic<br />
acid (ACC) by ACC synthase (EC 4.4.1.14), which is then oxidized<br />
to ethylene by ACC oxidase (Kende 1993; Fig. 12).<br />
The HCN generated from the catalytic action of ACC oxidase is detoxified by rapid
‘S<br />
~~~:L-methionine<br />
~-methyltransf~ras~<br />
OH c ethionine<br />
SAM:<br />
SAHC<br />
tr~s~inatio~<br />
& dec~boxylation<br />
3-dime~ylsulfonio-<br />
(~~SP-alde~yde)<br />
Figure 10 Pathway of synthesis of dimethylsulfoniopropionate (DMSP) in ~ollffstonjff ~~orff:<br />
SAM, ~-adenosyl~ethionine; SAHC, S-adenosylhomocysteine.<br />
metabolism to the amino acid P-cyanoalanine that can then be hydrated to asparagine or<br />
conjugated to y-glut~yl-P-cyanoalanine (Yang 1989). 5’-~ethylthioadenosine is metabolized<br />
back to methionine by 5’-methyl~hio~bose and MTOB by the methionine salvage<br />
pathway (Miyazaki and Yang 198’7; see Fig. 12). ACC synthase may be regulated by its<br />
substrate SAM through S~-induced inactivation. This mechanism may be responsible<br />
for the short half-life of this enzyme in vivo (Yang 1989). ACC oxidase requires Fe2”<br />
for catalytic activity and, in some systems, is activated by C02 (Kende 1993). The plant<br />
species ~ ot~~ogeto~ ~ecti~at~s is unusual in that it lacks ACC oxidase and ethylene<br />
biosynthesis; this species is consequently rich in ACC (Summers et al. 1996).<br />
1 -<strong>Amino</strong>cyclopropane- 1 -carboxylate synthase is induced by various environmental<br />
stresses, including mechanical stress (wounding or mechanical impedance), drought,<br />
chilling, anoxia, and hypoxia (Yang 1989; Kende 1993; He et al. 1996a). ACC oxidase<br />
isalsoinduced by hypoxia and by mechanical impedance in maize roots (He et al.<br />
1996a); however, with the combination of hypoxia plus mechanical impedance, ethylene<br />
formation and ACC synthase induction exhibit a greater synergism than ACC oxidase<br />
induction, cellulase induction, and aerenchyma formation (He et al. 1996a). ~erenchyma<br />
formation accelerates the transfer of O2 from the aerial tissues to the ~2-de~cient tissues
ho~es et al.<br />
4-methylthio-2-<br />
OH hydro~bu~rate (MTHB)<br />
OH<br />
4-dime~ylsulfonio-2-<br />
hydro~ybu~rate ( ~ ~ S ~ )<br />
ure 11 Proposed pathway of synthesis of di~ethylsulfoniopropionate (DMSP) in marine<br />
algae.<br />
of the stem base and root of flooded plants (He et al. 3996b). Ethylene production in<br />
response to hypoxia is implicated in the signal transduction pathway leading to cell death<br />
and lysis in the root cortex resulting in aerenchyma formation in maize roots (He et al.<br />
1996b; Drew 1997). Because ACC oxidase is oxygen-dependent, ACC can accumulate<br />
markedly in roots exposed to anoxia. In flooded tomato plants, the accumulated ACC is<br />
transported from the root to the shoot (through the xylem) where it is then rapidly oxidized<br />
to ethylene, inducing epinasty (Bradford and Yang 1980; English et al, 1995) and<br />
adventitious root formation (Visser et al. 1996). The latter may partly involve an ethylene-mediated<br />
increase in the sensitivity of root-forming tissue to endogenous indoleacetic<br />
acid (Visser et al. 1996).<br />
ACC can be conjugated to malonyl-ACC (MACC). MACC synthesis is catalyzed<br />
by ACC ~-~alonyltransferase (Yang1989;MartinandSaftner1995). An alternative<br />
ACC conjugate has recently been identified in tomato fruits: (y-gluta~yla~ino)-cyclo-
337<br />
ATP<br />
I<br />
\<br />
methio~ine<br />
salvage<br />
pathway<br />
PPi + Pi<br />
'S<br />
I<br />
Ad<br />
Smethyl-<br />
~hioadenosin~<br />
(rnA.1<br />
OH<br />
C,Hd + C02 + HCN + 2H20<br />
Figure 12 Pathway of ethylene (C21-&) biosynthesis in plants, and pathway of recycling of 5'-<br />
methylthioadenosine (MTA) to methionine (the methionine salvage cycle) in plants: ACC, 1-<br />
aminocyclopropane-1-carboxylate; MTR, ~ethylthio~bose; MTOB, 4-methylthio-2-oxobutyrate;<br />
SAMS, ~-adenosylmethionine synthetase.<br />
propane-1-carboxylic acid (GACC; Martin et ai. 1995). MACC and GACC formation<br />
may contribute to the regulation of ACC levels and, thereby, of ethylene formation (Yang<br />
1989; Kende 1993; Banga et al. 1996).<br />
The diamine putrescine, the trimine spermidine, and the tetramine spermine are ubiquitous<br />
in plant cells (Smith et al. 1979; Bagni and Pistocchi 1992). They occur as free<br />
cations and as conjugates with phenolic acids and macromolecules (Galston and Sawnhey<br />
1990). Their levels increase greatly in response to environmental stresses, most notably<br />
under conditions of potassium deficiency, water deficits, salinity stress, anaerobiosis,<br />
and acid stress (Flores et al. 1989). Because polyamines are synthesized by amino acid<br />
decarboxylation reactions that consume €I?, polyamine accumulation may function as<br />
part of a homeostatic mechanism to keep intracellular pH at a constant value (Flores et<br />
al. 1985). Polyamines may also play a role in the regulation of DNA replication and cell<br />
division, and they are implicated in the control of senescence and morphogenesis (Evans<br />
and Malrnberg 1989; Galston and Sawnhey 1990).<br />
Arginine decarboxylase (ADC; EC 4.1.1.19)-(a chloroplast-localized enzyme;<br />
Borrell et al. 1995)"is induced by a variety of stresses (most notably potassium deficiency;<br />
Watson and Malmberg 1996) and is thought to be the enzyme primarily responsible<br />
for environmental stress-induced putrescine accumulation (Galston and Sawnhey
199~; Fig. 13). ADC of oats is clipped from a precursor into two polypeptides found in<br />
the soluble enzyme (Malmberg et al. 1992). The two polypeptides are linked by disul~de<br />
bonds to form the active enzyme (Watson and Malmberg 1996). The oat (Bell and Malmberg<br />
1990), tomato (Rastogi et al. 1993), and Ar~bi~o~si~ atso son and Malmberg 1996).<br />
C genes have been cloned. In Ar~bi~o~sis, ADC enzyme levels do not correspond<br />
to ADC protein amounts, suggesting post~anslational modification of the enzyme<br />
in response to potassium deficiency (Watson and Malmberg 1996). In osmotically<br />
stressed oat leaves, spermine inhibits posttranslational processing of the ADC precursor,<br />
with subsequent decreases in mature ADC (Borrell et al. 1996).<br />
a~~atine<br />
~athways of synthesis and metabolism of putrescine.
The alternative, more direct, pathway of synthesis of putresine through ornithine<br />
decarboxylation catalyzed by cytosolic ornithine decarboxylase (ODC; EC 4" 1.1.17; see<br />
Fig. 13) is proposed to be of little importance in stress-induced putrescine accumulation,<br />
but may be critical in regulation of developmental processes (Galston and Sawhney 1990;<br />
Walden et al. 1997). Increased putrescine biosynthesis, mediated by transgenic ODC,<br />
promotes somatic embryogenesis in carrots (Bastola and Minocha 1995). The diamine<br />
cadaverine, derived from the amino acid lysine by decarboxylation, may play an important<br />
role in root development (Gamarnik and Frydrnan 1991).<br />
The triamine spermidine is synthesized by condensation of putrescine with decarboxylated<br />
SAM, synthesized by SAM decarboxylase (EC 4.1.1.50; Flores et al. 1989):<br />
SAM + Ht SAM decarboxylase<br />
> decarboxylated SAM + C02<br />
The condensation of decarboxylated SAM and putrescine is catalyzed by spermine<br />
synthase (putrescine aminopropyltransferase; EC 2.5.1.16). Further condensation of spermidine<br />
with decarboxylated SAM, catalyzed by spermine synthase (EC 2.5.1.22), produces<br />
the tetramine, spermine (Flores et al. 1989; see Fig. 13). Recently, the assumption<br />
that spermidine synthase and spermine synthase are two separate enzymes in plants, has<br />
been questioned (Bagga et al. 1997). The putrescine aminopropyltransferase of alfalfa<br />
may be responsible not only for spermidine and spermine synthesis, but also the synthesis<br />
of the uncommon polyamines, thermospermine, homocaldopentamine, and homocaldohexamine<br />
(Bagga et al. 1997).<br />
Antisense SAM decarboxylase potato plants show markedly altered phenotypes,<br />
including stunted growth; short internodes; stem branching and small leaves; decreased<br />
SAM decarboxylase transcripts; a reduction in SAM decarboxylase activity; reductions<br />
in the levels of putrescine, Spermidine, and spermine; and a marked increase in ethylene<br />
production (Kumar et al. 1996). This supports the idea that there is a competitive interaction<br />
between polyamine and ethylene biosynthesis (Walden et al. 1997).<br />
In addition to serving as a precursor of spermidine and spermine, putrescine has<br />
three other metabolic fates: conjugation with hydroxycinnamic acids, metaboli~m to y-<br />
aminobutyrate (GABA), and utilization in alkaloid biosynthesis, as indicated in Figure<br />
13. Hydroxycinnamic amide (HCA) conjugates of polyamines accumulate markedly in<br />
the floral apex during flower development and are implicated in the development of<br />
competence to flower; certain mutants of tobacco that are deficient in HCAs are unable<br />
to flower, and male sterile mutants of maize lack HCA accumulation in anthers (Flores<br />
et al. 1989). A novel polyamine conjugate, ~-hexanoylspermidine, has recently been<br />
identified in senescing pea ovaries and petals (Perez-Amador et al. 1996). GABA can be<br />
derived from putrescine (through y-aminobutyraldehyde) through the reactions catalyzed<br />
by diamine oxidase and y-aminobutyraldehyde dehydrogenase (Flores et al, 1989). As<br />
noted in Section III.B, the latter enzyme may be the same as BADH involved in glycinebetaine<br />
synthesis (Trossat et al. 1997). Putrescine serves as precursor of the nicotine and<br />
tropane alkaloids (Smith et al. 1979; Flores et al. 1989; Hashimoto and Yamada 1994),<br />
which may play important roles in plant defense against herbivores (see Fig. 13).<br />
The tripeptide glutathione (GSH), is synthesized from the amino acids glutamate, cysteine,<br />
and glycine by the enzymes y-glut~ylcysteine synthetase (EC 6.3.2.2) and glutathione<br />
synthetase (EC 6.3.2.3; Meister 1983):
Glutamate + cysteine + ATP<br />
~Glutamylcysteine<br />
synthetase<br />
> y-glutamylcysteine + ADP + Pi<br />
y-glutamylcysteine + glycine + ATP<br />
synthetase > glutathione + ADP + Pi<br />
Glutathione (GSH)playsa key role in detoxification of certainxenobioticsin<br />
higher plants through the action of glutathione S-transferases (EC 2.5.1 I) 18; Breaux 1986;<br />
Pickett and Lu 1989; Rossini et al. 1996). The glutathione conjugate of a xenobiotic can<br />
then be transported into and sequestered by the vacuole (Li et al. 1995; Kreuz et al.<br />
1996; Marrs 1996). Glutathione S-transferases are induced by a wide range of chemical<br />
agents (Ulmasov et al. 1995), wounding, heavy metals, ethylene, and ozone (Marrs<br />
1996). These enzymes may have initially evolved to conjugate naturally occurring compounds,<br />
such as phenylpropanoids (Dean et al. 1995), and naturally occurring endogenous<br />
reactive electrophilic compounds (Marrs 1996), but may have then been recruited<br />
to detoxify anthropogenic exogenous xenobiotics (Kreuz et al, 1996; Mans 1996).<br />
Under a variety of environmental stresses (e.g., high light and low temperature,<br />
water deficits, or others that disrupt membrane-bound electron transport systems), plants<br />
produce a range of active oxygen species, including superoxide, hydrogen peroxide, and<br />
the hydroxyl radical (Smirnoff 1993). Superoxide is converted to hydrogen peroxide by<br />
the action of superoxide dismutase. The hydroxyl radical can then form from the reaction<br />
of hydrogen peroxide with superoxide, particularly in the presence of iron or other transition<br />
state metals (Srnirnoff 1993). These toxic species can also form when plants are<br />
exposed to ozone, SO2, NO, and NO2. They react with membranes, proteins, and nucleic<br />
acids (Smirnoff 1993). Glutathione is involved in quenching these free radicals through<br />
the ascorbate-GSH cycle (Alscher 1989; Smirnoff 1993, 1995). Thus, hydrogen peroxide<br />
can be detoxified by the action of ascorbate peroxidase (EC 1.11.1.11), dehydroascorbate<br />
reductase (EC 1.8.5.1), and glutathione reductase (EC 1.6.4.2):<br />
Ascorbate + H202 Ascorbate peroxidase > H20 + dehydroascorbate<br />
~ehydroascorbate<br />
Dehy~oascorbate<br />
reductase<br />
+ GSH ascorbate + oxidized glut at hi on^ (GSSG)<br />
GSSG + NADPH + H+ Glutathione reductase<br />
> GSH + NADP'<br />
Strictly speaking, the initial product of the ascorbate peroxidase reaction is monodehydroascorbate,<br />
which can then either be reduced back to ascorbate by the action of an<br />
NAD(P)H-dependent monodehydroascorbate reductase (EC 1.6.5,4), or directly reduced<br />
back to ascorbate by an electron from PS-I (Smirnoff 1995). Any monodehydroascorbate<br />
not so reduced disproportionates to ascorbate plus dehydroascorbate (Smirnoff 1995); it<br />
is only the latter that is reduced by glutathione in the reactions depicted in the foregoing.<br />
Specific isoforms of the enzymes of this cycle are chloroplast-localized and are induced<br />
by an array of environmental factors (Alscher 1989; Pastori and Trippi 1992; Smirnoff<br />
1995). Overexpression of glutathione reductase leads to an increase in antioxidant capacity<br />
and resistance to photoinhibition in poplar (Foyer et al. 1995). However, overproduction<br />
of ascorbate peroxidase in the tobacco chloroplast does not appear to provide protection<br />
against ozone (Torsethau~en et al. 199'7).
Although the chloroplast-localized ascorbate-glutathione cycle is well characterized,<br />
it is now becoming clear that the enzymes of this cycle are also found in mitochondria<br />
and peroxisomes, and they may represent an important antioxidant protection system<br />
against H203 generated in these organelles (Jimhez et al. 1997).<br />
Glutathione transferases may also be involved in conjugating electrophiles generated<br />
from the reaction of hydroxyl radicals with lipids and DNA and, thus, contribute to<br />
oxidative stress resistance (Marrs 1996). Apoplastic GST induction may be mediated by<br />
an oxidative signal in soybean (Flury et al. 1996).<br />
F. P ~ y ~ o c ~ ~<br />
of Heavy<br />
Glutathione serves as a precursor of peptides known as' phytochelatins9 which are composed<br />
of two or more repeating y-glutamylcysteine units, with a terminal glycine residue;<br />
(~-glutamylcysteine)~-gly, where n = 2-11 (Steffens et al, 1986; Reese and ~agner<br />
1987). The enzyme responsible for the synthesis of these peptides is known as phytochelatin<br />
synthase (Rauser 1990, 1995). Phytochelatins (PCs) play an important role in the<br />
detoxification of certain heavy metals (p~ticularly cad~ium) in plants (Rauser 1990,<br />
1995; Howden et al. 1995a,b). Phytochelatin synthase is activated by cadmium (Rauser<br />
1995). In certain plants (notably legumes) that can synthesize homoglutathione, in which<br />
g-alanine is substituted for glycine as the terminal amino acid, homophytochelatins are<br />
synthesized along with PCs in response to Cd (Klapheck et al, 1995).Inmaizeand<br />
certain other species of the Poaceae, a third family of phytochelatins has been found in<br />
which serine is the COOH-te~inal amino acid (Rauser and ~euwly 1995). ~admium<br />
detoxification may require transport of the Cd-phytochelatin complexes into the vacuole;<br />
a transport system has been recently described for these complexes (Salt and Rauser<br />
1995).<br />
In Rubia tinctoru~, phytochelatins (class I11 metallothionein) are induced by many<br />
metal ions, but only a few (Ag, Cd, and Cu) are bound to the PCs that they induce<br />
(Maitani et al. 1996).<br />
v. A<br />
A<br />
During anaerobic stress, pyruvate must be metabolized to either lactate (by lactate dehydrogenase;<br />
LDH) or ethanol (by pyruvate decarboxylase [PDC] and alcohol dehydrogenase<br />
[ADH]) to regenerate the NAD" required to sustain glycolysis and associated ATP<br />
production (Davies 1980). Lactate fermentation may cont~bute to the acidification of the<br />
cytoplasm (Roberts et al, 1984; Ratcliffe 1995), which may progressively inhibit LDH<br />
and activate PDC, promoting alcohol fe~entation (Rivoal and Hanson 1994a). Accumulation<br />
of pyruvate under anaerobic conditions may shift the equilibrium of alanine aminotransferases<br />
toward alanine accumulation; a rapid response of plants to anaerobic stress<br />
(Streeter and Thompson 1972; Stewart and Larher 1980):<br />
Pyruvate + amino acid 3 alanine + 2-keto acid
An alanine a~notransferase (g1utamate:pyruvate a~notransferase; EC 2.6.1.2) is induced<br />
under anaerobic conditions in barley roots (Good and Crosby 1989; Good and<br />
uench 1992). The precise function of alanine accumulation under anoxia is unknown,<br />
butitisspeculatedthatit may serve as a storage form of pyruvate (perhaps in the<br />
vacuole), controlling supply of pyruvate to LDH and PIX and, thereby, the flux to<br />
lactate and ethanol (Good and Crosby 1989; Good and Muench 1992). The synthesis of<br />
alanine may occur at the expense of the acidic amino acids, glutamate and aspartate<br />
(Streeter and Thompson 1972; Stewart and Larher 1980), and occurs concomitantly with<br />
the accumulation of GABA (see following Section VI).<br />
.<br />
The accumulation of the amino acid y-~nobutyrate (GABA) is markedly stimulated<br />
under anaerobic conditions (Streeter and Thompson, 1972; Stewart and Larher 1980;<br />
Aurisano et al. 1995; Ratcliffe 1995; Drew 1997). Although G A can be derived from<br />
putrescine (see Sec. IV.D), the anaerobic stress-induced accumulation of GABA is<br />
thought to result primarily from the catalytic action of glutamate decarboxylase (EC<br />
4.1.1.15). In part, the stimulation of glutamate decarboxylase may be the consequence of<br />
cytoplas~c acidification; glutamate decarboxylase has an acid pH optimum (-5.8; Patterson<br />
and Graham 1987):<br />
Glutamate + H+ ‘lutarnate<br />
> GABA + CQ2<br />
ause this reaction is proton-consuming, it could represent an adaptive response conuting<br />
to regulation of cytoplasmic pH (Patterson and Graham 1987; Crawford et al,<br />
atcliffe 1995). Crawford et al. (1994) have shown that weak acids causing cytoplasmic<br />
acidification also induce GABA accumulation, and they conclude that this response<br />
is consistent with a role for GABA synthesis in active pH regulation.<br />
y-A~nobutyrate can be transaminated with pyruvate to yield alanine (or with 2-<br />
oxoglutarate to yield glutamate), generating succinic semialdehyde, which can then be<br />
metabolized to succinate by the action of succinate semialdehyde dehydrogenase (Vandewalle<br />
and Qlsson 1983; Patterson and Graham 1987; Shelp et al. 1995; Fig. 14). The<br />
latter enzymes are mitochondrial (Hear1 and Churchich 1984; Breitkreuz and Shelp 1995)<br />
and have alkaline pH optima (Patterson and Graham 1987). GABA accumulation promoted<br />
by cytosolic acidi~cation may partly result from inhibition of GABA-transaminases.<br />
The conversion of glutamate to succinate by the action of glutamate decarboxylase,<br />
GABA ~ansa~nases, and succinic semialdehyde dehydrogenase is known as the<br />
GABA shunt (Vandewalle and Olsson 1983; Patterson and Graham 1987; Breitkreuz and<br />
Shelp 1995; Shelp et al, 1995), affording an alternative pathway for glutamate entry into<br />
the TCA cycle (see Fig. 14).<br />
The accumulation of GABA is induced<br />
response to a sudden decrease in temperature<br />
(Wallace et al. 1984; Patterson and Graham 1987), heat shock (Mayer et al. 1990),<br />
mechanical manipulation (Wallace et al. 1984), and water stress (Rhodes et al. 1986).<br />
Rapid GABA accumulation in response to wounding may play a role in plant defense<br />
against insects (Ramputh and Brown 1996).<br />
Glutamate decarboxylase is a cytosol-localized enzyme (Breitkreuz and Shelp
glutamate<br />
NH,<br />
2-oxo<br />
HO<br />
succinate<br />
Figure 14 Pathway of synthesis and metabolism of GABA.<br />
1995) and has recently been shown to be a calmodulin-binding protein that is<br />
C~~al~odulin activated (Baum et al. 1993; Ling et al. 1994; Baum and Fridmann<br />
1996; Arazi et al. 1995; Snedden et al. 1995). Crawford et al. (1994) note that reduced<br />
cytosolic pH values increase Ca”* levels, and rapid and transient increases in Ca”’ levels<br />
occur in response to mechanical stress and cold stress; conditions that elicit rapid GABA<br />
accumulation (Wallace et al. 1984). The Ca”+-calmodulin activation of glutamate decarboxylase<br />
provides a link between intermedi~ amino acid metabolism and perturbations<br />
of cytosolic ea2+ (Ling et al. 1994; Baum and Fridmann 1996; Arazi et al. 1995; Snedden<br />
et al. 1995) that regulate a host of other metabolic activities (Allan and Trewavas 1985;<br />
Bush 1995).<br />
The only other enzyme of glutamate metabolism currently known to be stimulated<br />
by Ca” in plants in glutamate dehydrogenase (GDH; EC 1.4.1.2), a ~itochondrial enzyme<br />
(Turano et al. 1997):<br />
NH~ + 2-oxoglutarate + NADH + H+ < dehydrogenase > glutamate + NAD+<br />
A calcium-binding domain has been identified in the f3-subunit of GDH encoded<br />
by GDH2, but not in the a-subunit of GDH encoded by GDHI in Arabidopsis, suggesting<br />
that the different isoforms of the hexameric GDH composed of different combinations<br />
of subunits, may be differentially regulated by Ca” (Turano et al. 1997).
We have attempted to highlight areas of amino acid metabolism that may play an important<br />
role in plant “stress” resistance, by osmotic adjustment and the accumulation of<br />
compatible osmolytes; the detoxi~cation of active oxygen sp<br />
heavy metals; and intracellular pH regulation. The advances since 1980 have been primarily<br />
in the cloning of the genes encoding key enzymes of these biosynthetic pathways<br />
and the characterization of their regulation in relation to environmental stimuli. As noted<br />
by Bohnert et al. (1993, the use of metabolic engineering holds great promise for testing<br />
hypotheses conce~ing the role of radical scavenging and osmolyte accumulation as<br />
stress-protection mechanisms (e.g., Foyer et al. 1995; Kishor et al. 1995). These traits<br />
are not restricted to amino acid metabolism; significant advances have also been achieved<br />
in engineering of superoxide dismutase, and the accumulation of polyols and fructans as<br />
compatible solutes or radical scavengers that may limit injury (Bohnert et al. 1995; Shen<br />
et al. 1997, and references cited therein), These traits may eventually need to be combined<br />
with modified ion uptake, exclusion or co~partmentation, facilitated water permeability,<br />
molecular chaperones, modified membrane properties, or stress-signaling pathways<br />
to realize their full potential (Bohnert et al. 1995; Bohnert and Jensen 1996). The<br />
next decade is likely to witness further exciting progress in these areas. In the testing of<br />
transgene effects on plant resistance to low water potentials and salinity stress, we urge<br />
(1) careful measurements of growth and water relations to quantify the degree of stress<br />
and to assess potential growth penalties of transgene expression, (2) consideration of<br />
relevant fluxes in addition to pool sizes, and (3) consideration of the metabolism of<br />
growing regions as well as mature tissues.<br />
Adams, E. and Frank, L. (1980). Metabolism of proline and the hydroxyprolines. Annu. Rev. Biochern.,<br />
49: 1005-1061.<br />
Allan, E. E;. and Trewavas, A. J. (1987). The role of calcium in metabolic control. The ~joche~jstry<br />
of Plants, Vol. 12 (D. D. Davies, ed.), Academic Press, New York, pp. 117-149.<br />
Alscher, R. G. (1989). Biosynthesis and antioxidant function of glutathione in plants. P~ysjoZ<br />
Plant., 77 457-464,<br />
Anthoni, U., Christophersen, C., Hougaard, L., and Nielsen, P. H. (1991). Quate~ary ammonium<br />
compounds in the biosphere-an example of a versatile adaptive strategy. Cornp. Bioche~z.<br />
Bi~~hys~, 99B: 1-18.<br />
Arazi, T., Baum, G,, Snedden, W. A,, Shelp, B. J., and Fromm, H. (1995). Molecular and biochemical<br />
analysis of calmodulin interactions with the calmodulin-binding domain of plant glutamate<br />
decarboxylase. Plant Physiol., 108: 551-561.<br />
Aurisano, N., Bertani, A,, and Reggiani, R. (1995). Anaerobic accumulation of 4-ami~obutyrate in<br />
rice seedlings; causes and significance. P~ytoche~istry, 38: 1147-1 150.<br />
Bagga, S,, Rochford, J., Klaene, Z,, Kuehn, G, D., and Phillips, 6. C. (1997). Putrescine aminopropyltransferase<br />
is responsible for biosynthesis of spermidine, spermine, and multiple uncommon<br />
polyamines in osmotic stress-tolerant alfalfa. Plant P~ysjol., 114: 445454,.<br />
Bagni, N. and Pistocchi, R. (1992). Polyamine metabolism and compa~mentation in plant cells.<br />
~jt~ogen ~etaboljsrn of Plants (K. Mengelm and D. J. Pilbeam, eds.), Clarendon Press,<br />
Oxford, pp. 229-248,<br />
~allantyne, J. S, and Chamberlin, M. E. (1994). Regulation of cellular amino acid levels. Cellular
and Molecular Physiology of Cell Volume Regulation (K. Strange, ed.), CRC Press, Boca<br />
Raton, FL, pp. 11 1-122.<br />
Banga, M., Slaa, E. J., Blom, C. W, P. M,, and Voesenek, L. A. C. J. (1996). Ethylene biosynthesis<br />
and accumulation under drained and submerged conditions: A comparative study of two<br />
Rumex species, Plant Physiol., 112: 229-237.<br />
Barnett, N. M, and Naylor, A. W. (1966). <strong>Amino</strong> acid and protein metabolism in Bermuda grass<br />
during water stress. Plant Physiol., 41: 1222-1230.<br />
Bastola, D. R. and Minocha, S. C. (1995). Increased putrescine biosynthesis through transfer of<br />
mouse ornithine decarboxylase cDNA in carrot promotes somatic embryogenesis. Plant<br />
Physiol., 109: 63-7 1.<br />
Baum, G., Chen, Y., Arazi, T., Takatsuji, H., and Fromm, H, (1993). A plant glutamate decarboxylase<br />
containing a calmodulin binding domain. Cloning, sequence, and functional analysis.<br />
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