Amino Acid Transport

US ~ epar~ent of Agriculture-~~S and ~epartment of Plant ~iology, 

~niversi~ of ~llinois at Urb~na-Champai~n, Urbana, Illinois 

* 

of the uni~ue features of lants as multicellular organisms is their ability to synthes 

from simple inorganic salts and the free ener~y made 

tr~sduction reactions of photosyn~esis. 

no acids. ~lthoug~ every pla 

capacity to assi~late inorganic ni 

ly restricted to cells in mature leave 

no acids from sites of pri 

trogen. I ~ aminoacids ~ s ~ e ~ 

esis, they are also the precursors of every ~~ogen 

luding nucleic acids, gro~th regulators, ~hotosynarray 

of other essential compounds. In m ~ y 

of ~~ogen me~bolis~ in the plant and underd 

bet~een organs is a ~ndamental ~uestio~ in 

ids in plants must neces 

from the soil solution by the 

h onium ions 

ssl~lated in the root, or

3 ush 

general patterns have been identified (Pate 1980, 1983; Andrews 1986). Temperate perennials 

and legumes frequently carry out primary assimilation in the roots, whereas 

tropical and subtropical species strongly favor leaf assimilation. Temperate annuals fall 

into both categories. Significantly, if high concentrations of nitrate are available in the 

soil solution (>1 mM), many species carry out primary assimilation in the leaf. Even 

symbiotic nitrogen assimilation is suppressed in the presence of high levels of nitrate 

(Harper 1994; Pate 1980). 

Once assimilated into amino acids, there are several pathways for the long-distance 

transport of reduced nitrogen (Fig. 1). For those plants and conditions in which primary 

assimilation occurs in the leaf, amino acids are transported to the heterotrophic tissues 

by the phloem-translocation stream. In species that use an apoplastic phloem-loading 

mechanism (Bush 1992; Turgeon 1996), this requires the release of amino acids from 

mesophyll cells into the apoplastic space, with subsequent loading into the phloem. A 

few amino acids (generally a combination of two or more of the following: glutamate, 

aspartate, glutamine, asparagine, alanine, and serine) are more concentrated in the 

phloem-translocation stream than in the cytoplasm of the mesophyll (Riens et al. 1991; 

Winter et al. 1992; Lohaus et al. 1994; Lam et al. 1995). These amino acids must be 

actively transported into the phloem (Bush 1993a). In contrast, the measured concentrations 

of other amino acids in the phloem suggests they are at or below ~ermodyn~c 

equilibrium with the mesophyll and passive transport activity has been suggested (Riens 

et al. 199 1; Winter et al. 1992; Lohaus et al. 1994). However, it is important to recognize 

that the phloem represents a dynamic pool of amino acids that is rapidly turning over as 

pressure-driven mass flow continuously removes the loaded metabolites. Thus, although 

snap-shot determinations of amino acid concentrations may implicate passive transport 

mechanisms for some amino acids, the kinetics of amino acid turnover suggests that high 

rates of transport activity must occur, and passive transport activity, for which the rate 

of transport is directly proportional to the concentration difference, may not be rapid 

enough to maintain the phloem pools that are quickly flushed away. Given the dynamics 

of rapid exchange at the phloem interface, it is not surprising that several high-affinity, 

proton-coupled amino acid transporters have been described in plasma membrane vesicles 

isolated from mature leaf tissue (Li and Bush 1990, 1991, 1992; Williams et al. 

1990, 1992, 1996; Weston et al. 1995). These active transporters can maintain high rates 

of flux, even against substantial concentration differences. Nevertheless, evidence for 

low levels of facilitated diffusion is also present in membrane vesicles isolated from leaf 

tissue (Williams et al. 1990; D. R. Bush unpublished data). Although the participation 

of facilitated transporters in phloem loading has yet to be demonstrated, such porters are 

likely candidates to play an important role in releasing amino acids from the surrounding 

mesophyll cells. 

When nitrate or ammonium ions are assimilated in the root, amino acids are transported 

to mature leaf tissue in the xylem by the transpiration stream (Pate 1980, 1983). 

Because the nitrogen requirement of mature leaves is low, a substantial portion of the 

amino acids arriving with the transpiration stream are cycled into the phloem for redistribution 

to heterotrophic sinks (see Fig. 1). Cycling amino acids from the xylem to the 

phloem in mature leaf tissue is also a fundamental process in symbiotic nitrogen fixation. 

For example, atmospheric nitrogen assimilated in the Rhizobium-legume symbiosis is 

exported from the root in the transpiration stream as amide amino acids (examples include, 

clover and alfalfa) or ureides (mung bean and soybean; Schubert 1986; Winkler 

et al, 1988). Even though some of the amide amino acids may be removed from the

Amino Acid Import 

Developing leaves, 


Amino Acid Cycling 

(xylem to phloem) 


Amino Acid Export 

Primary ~ssimilation 

Amino Acid Cycling 

(phloem to xylem) 

/ 


- - = Xylem 

Phloem 

Figure 1 Schematic presentation of amino acid transport pathways in the plant: Amino acids 

synthesized in the root from primary assimilation or symbiotic relations are transported to mature 

leaf tissue in the xylem (----). Amino acids synthesized in the leaf, or arriving from the root in the 

xylem, are transported out of the leaf in the phloem (-) to satisfy the needs of heterotrophic 

sids. Import-dependent tissues include developing leaves, roots, cortical cells in the stem, seed 

and fruits, and apical meristems. 

transpiration stream by stem tissues (McNeil et al. 1979; Pate 1983), much of that nitrogen 

arrives in mature leaf tissue, from whichit is redistributed by the phloem. The 

ureides also arrive in the leaf, but instead of being redistributed in the phloem, they are 

catabolized to C02 and (Winkler et al. 1988). Released ammonia is incorporated 

into amino acids that are used in leaf metabolism or redistributed to heterotrophic tissues 

by the phloem. 

Nitrogen cycling between the vascular systems is not restricted to the leaf tissue. 

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). 

Thus, cycling amino acids between the two vascular systems in mature leaf and root 

tissue (see Fig. 1) permits dynamic adjustments of carbon and nitrogen metabolism in 

response to developmental queues or environmental change. For example, the metabolic 

needs of the mature root system emphasizes catabolic reactions associated with maintenance 

reactions versus the anabolic reactions that drive early growth. Therefore, excess 

amino acids transported to the root under these conditions can be redirected to the shoot 

for subsequent incorporation into developing seed. Significantly, excess amino acid nitrogen 

arriving at the root may also represent a dynamic signal that triggers the downregulation 

of nitrate assimilation (Cooper and Clarkson 1989; Crawford 1995; Padgett 

and Leonard 1996). 

In addition to its role in primary assimilation, amino acid transport is a key process 

in leaf senescence and seed germination. In rice, for example, more than 60% of the 

amino nitrogen delivered to developing leaves and ears is derived from amino acids 

retrieved from senescing older leaves (Mae et al. 1983, 1985; Feller and Fischer 1994). 

Likewise, amino acids released from storage proteins during seed ge~nation are the 

principal nitrogen source in growing seedlings (Schobert and Komor 1989). Taken together, 

it seems clear that amino acids are the currency of nitrogen exchange in the plant. 

Amino acids are transported into plant cell by proton-coupled symporters that link translocation 

across the plasma membrane to the proton-motive force generated by the €I?- 

pumping ATPase (Fig. 2; Bush 1993a). These are widely expressed carriers that appear 

to be the primary pathways for amino acid transport into plant cells. However, there is 

some evidence for facilitated transporters that mediate passive amino acid transport, although 

little biochemical and no molecular characterization of such porters has been 

published. 

The amino acid symporters have been investigated in recent years using purified 

membrane vesicles and imposed proton electrochemical potential differences (Bush 

1993a). Purified membrane vesicles are a very useful experimental system for studying 

membrane transporters. Although membrane transport activity can be examined with 

intact cells, there are many complications associated with metabolism and intracellular 

comp~mentation that limit experimental interpretation. Additionally, it is difficult to 

dissect the bioenergetics of a transport system in intact tissues because of complex interactions 

among the primary pumps, ion channels, and other unrelated porters. Isolated 

membrane vesicles, on the other hand, allow one to focus on specific transport processes 

while minimizing problems associated with the living cell. The major attributes of the 

membrane vesicle approach include (1) unequivocal identification of membrane location 

using purified vesicles, (2) elimination of posttransport metabolism and compartmentation, 

and (3) the ability to control both intra- and extravesiculq solution composition 

(Bush 1992). The ability to control solution co~position on both sides of the membrane 

is particularly useful for investigating the bioenergetics of the transport process. With 

this approach, the basic transport properties of the plant amino acid symporters have 

been described (Bush 1993a). 

Plant amino acid symporters are electrogenic transporters. Electrogenicity was

Model of amino acid symporter-mediated transport into plant cells: The H+-ATPase 

generates a substantial proton electrochemical potential difference (both ApH and AY) that can 

drive transport reactions three or four orders of magnitude away from equilibrium. A diagrammatic 

representation of the 11 transmembrane domains (TMDs) of NAT2/AAPl is also presented. 

demonstrated in isolated membrane vesicles by examining the effect of membrane electrical 

potential (AY) on ApH-dependent transport activity. Potassium gradients and valinomycin 

were used to manipulate AY. In the absence of compensating charge flow, a low 

rate of amino acid flux was observed. When the AY was clamped at zero, transport was 

stimulated fourfold, and when a negative AY was imposed, transport was stimulated 

sixfold. These data are consistent with an electrogenic transport system that results in 

primary charge separation as proton cotransport drives substrate accumulation (Li and 

Bush 1990). Similar results were reported for this porter expressed in Xenopus oocytes 

(Boorer et al. 1996). 

The stoichiometry of H"-amino acid cotransport was recently measured using electrophysiologic~ 

methods to measure conductance through another plant amino acid symporter 

(AAP5) expressed in Xenopus oocytes (Boorer and Fischer 1997). Boorer and 

Fischer (1997) used direct measurements of [3H]amino acid uptake and simultaneous 

measurements of inward current to show that the stoich~o~etry of the amino acid symporters 

is one w' per one amino acid. This held true for the basic, neutral, and acidic 

amino acids examined. Because the acidic amino acids also carried one positive charge, 

these results suggest they are transported in their neutral forms. 

The number of symporters that mediate amino acid transport into higher-plant cells 

has yet to be determined. Transport competition experiments with isolated membr~e 

vesicles provided evidence of at least four amino acid symporters: an acidic amino acid 

symporter, a basic amino acid symporter, and two symporters for the neutral amino acids 

(Li and Bush 1990, 1991). The neutral amino acid porters were resolved in the competition 

experiments because of their differential affinity for isoleucine, valine, threonine, 

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 

(1991) hypothesized that branching at the fharbon in isoleucine, valine, and threonine 

results in steric interactions that block their access to one of the neutral porters. More 

recently, several amino acid symporters have been cloned, and we now know that many 

amino acid symporters are found in the plant (see later discussion). 

The ability of various amino acid analogues to compete for transport into purified 

membrane vesicles was used to identify molecular determinants that are important for 

substrate recognition by the neutral amino acid symporters (Li and Bush 1992). D-ISOmers 

of alanine and isoleucine were not effective transport antagonists of the L-isomers, 

thereby implicating stereospecificity and suggesting that positional relations between the 

a-amino and carboxyl groups is an important parameter in substrate recognition. This 

conclusion was supported by the observation that p-alanine and analogues with methylation 

at the a-carbon, at the carboxyl group, or at the a-amino group, were not effective 

transport antagonists. In contrast, analogues with various substitutions at the distal end 

of the amino acid were less potent antagonists. More recently, el~trophysiological investigations 

of AAPS expressed in Xenopus oocytes also identified the a-amino and carboxyl 

groups, as well as the P-carbon, as important dete~inants in defining substrate 

specificity (Boorer and Fischer 1997). Thus, the binding site for the carboxyl end of the 

amino acid is a well-defined space that is characterized by compact, asymmetric positional 

relations and specific ligand interactions (Li and Bush 1992). 

The amino acid symporters are inhibited by carbonyl cyanide ~-chlorophenyl hydrazone 

(CCCP) and diethylpyrocarbonate (DEPC; Bush and Langston-~nkefer 1988; Li 

and Bush 1990). CCCP is a protonophore that inhibits amino acid transport by dissipating 

the proton-motive force that drives the symport reaction. DEPC is a small molecule that 

forms covalent bonds with the imidazole ring of histidine residues. The time-dependent 

inactivation of alanine transport in the presence and absence of amino acids showed that 

substrate binding protects the symporter from DEPC inactivation (Fig. 3). This observation 

is consistent with DEPC binding at (or at least confo~ationally linked to) the 

substrate-binding site of the carrier (Bush and Li 1991; see Bush 1993b, for discussion). 

Since histidine residues frequently participate in protonation reactions, this observation 

implicates a histidine residue in the reaction mechanism. 

Initial efforts to identify the amino acid syrnporters by using standard biochemical strategieswerelargelyunsuccessful.Althoughthiswasoftenquitefrustrating, 

it was not 

too surprising, given that these are very low-abundance, integral membrane prot6ins. 

Fortunately, a novel molecular approach was successfully used to clone several plant 

amino acid symporters and biochemical analysis has moved forward using these clones 

and a variety of expression systems. 

The first plant amino acid symporter cloned was identified using ~nctional complementation 

of yeast amino acid transport mutants (Frommer et al. 1993; Wsu et ai. 

1993). ~acc~aro~~ces cerevisiae strains that are auxotrophic and transport-deficient for 

a specific amino acid were tr~sfo~ed with plant cDNA libraries that are constructed 

in yeast expression vectors. Those transformants that successfully expressed an appropriate 

plant amino acid symporter were easily identified by screening for growth on low 

concentrations of the limiting amino acid. Because these yeast strains are both auxotrophic 

and transport-deficient, a simple secondary screen in the absence of the limiting

80 

40 

8 20 

0 

0.0 2.0 4.0 6.0 

ure 3 Time- and concentration-dependent inactivation of proton-coupled alanine transport 

into purified plasma membrane vesicles (PMVs) isolated from sugar beet leaf tissue: PMVs were 

pretreated with DEPC plus or minus alanine for the indicated times, at which point the reaction 

was stopped. The vesicles were pelleted and resuspended in inhibitor-~ee buffer, and transport 

activity was measured as described previously (Li and Bush 1990). 

amino acid was used to show that growth was dependent on amino acid transport, rather 

than complementation of the biosynthetic pathway. This is a powerful approach because 

it allows one to clone transporter cDNAs without protein sequence information, and 

because it provides a useful expression system for subsequent investigations of the biochemical 

properties of the trans~ort protein (Bush 1993a; Tanner and Casper 1996). The 

screens used for functional co~ple~entation s~ould also id en ti^ high-affinity facilitated 

diffusion transpo~ers. in spite of numerous screens of different expression 

libraries in a variety of ast strains, no examples of facilitated transporte~ have 

been reported. 

The cDNA clone of the first amino acid symporter identi~ed with functional complementation 

encoded a protein containing 486 amino acids with a calculated molecul~ 

mass (MJ of 52.9 kDa (Prommer et al. 1993; Hsu et al. 1993). Hy~opathy analysis of 

the deduced amino acid sequence suggests this is a typical hydrophobic membrane protein 

with 10 to 12 membrane-spanning domains and three sites of potential N-linked 

glycosylation. The transport kinetics, substrate specificity, and inhibitor sensitivity of the 

heterologously expressed carrier matched those described for one of the neutral amino 

acid symporters ch~acterized in purified membrane vesicles (Li and ush 1991). Hsu et 

al. (1993) designated this gene ~A~ for neutral amino acid transporter TI, and Prommer 

et al. (1993) termed it ~~~. No homologues were identi~ed in the databases, suggesting 

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) 

that are closely related to NAT2/AAPl (Kwart et al, 1993; Fischer et al. 1995; Rentsch 

et al. 1996). They are similar in size and predicted topology, and sequence homologies 

range from 70 to 95% amino acid si~larity, thus providing good evidence that these 

clones represent a family of amino acid translocators (Fischer et al. 1995; Rentsch et al. 

1996). As might be expected for a gene family within a given organism, these porters are 

differentiated by a combination of unique expression patterns and substrate specificities. 

Individually, the AAP transporters have broad substrate specificity, although each exhibits 

some preference (lower Km or higher Yma) for certain classes of amino acids. For 

example, AAP4 is particularly active for valine and proline, whereas AAP5 is an effective 

lysine transporter (Fischer et al, 1995). Within the family, however., there appears to 

be some redundancy, for AAP1, AAP2, AAP4, and Asup6 share similar patterns of substrate 

specificity, whereas AM3 and AN5 are similar (Fischer et al. 1995; Rentsch et 

al. 1996). 

Although similar substrate specificity could lead to widespread functional redundancies 

between the AAPs., differential expression patterns appear to separate the various 

AAI? gene products from one another. For example, AAPS and AAPlNAT2 are widely 

expressed in all tissues, whereas AAP3 expression is restricted to the root (Fischer et al. 

1995). Moreover, even when related AAPs are expressed in the same organ, they can be 

functionally isolated by restricting expression to different cell types within that organ. 

Four other classes of plant amino acid transporters have been cloned that are not 

members of the AAP gene family. Two of them are basic amino acid ~anspo~ers that 

were also cloned by functional complementation: UTI encodes a lysine and arginine 

transpo~er that has 533 amino acids and 14 putative transmembrane domains (Frommer 

et al. 1995). It is a single-copy gene in ~r~bi~o~sis and sequence alignments suggest it 

is closely related to a human basic amino acid transporter (Yoshimoto et al. 1991; Christensen 

1992). The second basic amino acid transporter cloned is LHTl (Chen and Bush, 

1997). It is a lysine and histidine transporter that contains 446 amino acids and 9-10 

transmembrane domains. In contrast to AATl , LHTl does not transport arginine. RNA 

gel blot analysis and whole-mount in situ hybridization indicate that LHTl is most 

strongly expressed in pollen, siliques, and on the root surface. AATl is most strongly 

expressed in vascular tissue and flowers (Frornmer et al. 1995). The expression patterns 

of LHTl and UT1 complement one another, suggesting ~HTl may be involved in 

amino acid acquisition, whereas AATl may contribute to long-distance transpo~. In 

addition to these basic amino acid transporters, an aromatic amino acid transporter has 

also been recently characterized. 

An aromatic amino acid transporter was identified using functional expression of 

an EST cDNA that exhibited low levels of sequence homology to NAT2/AAPl (L. Chen 

and D. R. Bush, unpublished data). The cDNA is 1.6 kb, with an open-reading frame 

that codes for a protein with 432 amino acids and a calculated M, of 50 ma. Hy~opathy 

analysis of the deduced amino acid sequence suggests this is an integral membrane protein 

with 11 putative transmembrane domains. This protein exhibits sequence si~l~ty 

with AUX1 (a putative auxin transporter; Bennett et al. 1996) and to tryptophan and 

tyrosine translocators in E. coli (Wookey and Pittard 1988; Sarsero et al. 1991; Heatwole 

and Somerville 1991). RNA gel blot analysis has shown that this transporter is expressed 

at very low levels and that it is in all tissues. Whole-mount in situ hybridization further 

localized ART1 expression on the surface of roots and leaves and in young seedlings

and in stomata. Overall, ART1 belongs to 

favors aromatic amino acids and, perhaps, 

Chen and D. R. Bush, unpublished data). 

Two proline transporters have also 

a new class of amino acid transporter that 

to other aromatic compounds in plants (L. 

been recently cloned, ProTl and Pro72 

(Rentsch et al. 1996). These transporters were identified by complementing a Shr3- yeast 

strain that is unable to target yeast amino acid transporters to the plasma membrane. 

Significantly, the plant amino acid transporters are not affected by this processing mutation; 

therefore, several amino acid transporters (many of which were previously identified) 

complemented this yeast strain. A survey of different amino acids for evidence of 

transport through these porters showed that they are proline translocators. However, the 

rate of proline transport in yeast cells expressing these porters is 300 times slower than 

proline transport by AAP6 expressed in the same cell line. It is not known if this is a 

result of lower expression levels, or if it is an intrinsic property of these transporters. 

ProTl and ProT2 are closely related genes that encode typical membrane proteins that 

contain 442 and 439 amino acids, and have 10 putative transmembrane domains. Both 

genes are widely expressed in Arabidopsis, with ProTl strongest in roots, flowers and 

stems. Significantly, Pro22 expression was enhanced under conditions of water or salt 

stress, which is consistent with proline’s putative role in water and salt tolerance (Delauney 

and Verma 1993). 

Molecular descriptions of amino acid transport have included the isolation of several 

transport mutants in Arabidupsis. Two mutants that lack a low-affinity basic amino 

acid transport activity were isolated based on their resistance to lysine plus threonine or 

to ~-2-~inoethyl-~-cysteine, respectively (Heremans et al. 1997). The specificity of the 

lost transport activities were resolved based on apparent K, and sensitivity to transport 

inhibitors. Although both mutations have been mapped to chromosome 1, it is not clear 

if they are allelic. In a similar approach, the raz1 mutant of Arabidopsis was selected 

based on its resistance to the toxic proline analogue, azetidine-2-carboxylic acid (Verbruggen 

et al. 1996). Likewise, we have isolated two T-DNA-tagged lines that are resistant 

to high concentrations of valine in their growth medium (Chen and Bush 1994). 

Exogenous valine is a powerful plant growth inhibitor because it is a negative regulator 

of acetohy~ozyacid synthase (also known as acetolactate synthase), an enzyme in the 

valine biosynthetic pathway. Once down-regulated in the presence of excess valine, 

growth is inhibited because the cells starve for other amino acids, the precursors of which 

also pass through acetohy~oxyacid synthase. We were able to show that our lines are 

transport mutants versus regulation mutants because they are also resistant to azaserine, 

a toxic amino acid analogue that targets other enzymes in the cell. Thus, resistance to 

azaserine is consistent with loss of transport activity. We are currently attempting to 

identify the disrupted gene(s). 

One of the unexpected observations emerging from recent molecular descriptions 

of plant transporter genes is that there are large families of related porters that function 

in assimilate partitioning (Bush et al. 1996). Examples include the AHA family of protonpumping 

ATPases (Harper et al. 1990), the major facilitator superfamily of sugar transporters 

(Griffith et al. 1992; Bush et al. 1996), and the amino acid transporter families 

reviewed here. Although it is easy to imagine how these porters fulfill complementary 

functions in different organs and cells, it is clear that a major challenge in this field is 

to determine the unique contributions of the various transporters and, ultimately, to integrate 

their combined activities across the plant as a multicellular organism.

Given the central role played by plant amino acid symporters in nitrogen p~itioning 

into cells and between organs, and the complexity presented by the many transporters 

cloned to date, it is necessw to obtain basic information about their structure and regulation 

as the foundation for understanding their contributions to plant growth. Unfortunately, 

these kinds of investigations have only recently become possible with the cloning 

of the various transporters; therefore, very little has been published in this area. However, 

several laboratories have initiated projects investigating st~cture and function, and we 

anticipate significant progress in the near future. 

itiated a detailed investigation of N 

transporter family. We chose NAT 

sed, it transports many amino acid e co~on in the 

phloem-translocation stream, and because at least one member of this gene fdly appears 

to be expressed in every tissue of the plant. We are using an integrated approach 

with both biochemical and molecular strategies to learn more about the structure and 

function of this protein. 

The initial focus of our §tructural analysis has been establishing the membrane 

topology of ~ A This ~ is an , important question because our initial analysis suggested 

it does not contain 12 membrane-spanning helices as is commonly described for other 

metabolite translocators. We started by engineering a c-myc epitope on either the NH2- 

or COOH-termini of the expressed protein. We then used in vitro trans~ation, partial 

digestion with proteinase , and immuno~recipitation to determine the location of the 

NH2- and COOH-termini and to identify a family of oriented peptide fragments that we 

resolved on sodium dodecyl sulfate-polyacryldde gel electrophoresis (SDS-PAGE). 

These results showed that the ~H2-terminus is located inside the cell and the COOHterminus 

is outside. In addition, the lengths of the peptide fragments recovered after 

partial proteolysis allowed us to predict the location of protease-accessible cleavage sites 

between putative ~ansmembrane domains. We independently identified the location of 

the NH2- and COOH-termini using immuno~uorescence microscopy of NAT2 expressed 

in COS-1 cells. From the combined data, we proposed an 11 ~ansmembrane domain 

model, with the NH2-terminus in the cytoplasm and CO~H-terminus facing outside of 

cell (see Fig. 2; Chang and Bush 1997). This model of protein topology anchors our 

co~plement~ investigations of porter structure and function using site-directed and 

random mutagenesis. 

Previous biochemical investigations of ~E~C-dependent inactivation of the amino 

acid sympo~ers implicated a histidine residue in the reaction ~echanism (Li and Bush 

1990; see Fig. 3). An inspection of the ~ A gene family ~ identi~ed / two ~ histidine ~ 

is-337, that are conserved in every transporter. Therefore, we used 

site-directed mutagenesis to change se residues and then we determined the effect of 

those changes on transport activity 0th residues are essential because the modified 

proteins were no longer active, owever, the change in is-337 destabiliz~ the protein, 

thereby complicating our inte~re~tion of that result. Several amino acid substitutions of 

His-47, on the other hand, blocked transport activity in the stable protein, thus demonstrating 

the significa~ce of this residue in the transport reaction (L. Chen and D. R. Bush, 

un~ublished data). 

A second approach we have employed to identify impo~nt &no acid residues 

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 

residues for potential modification. ~ A ~ was randomly / ~ mutagenized ~ l in a mutator 

strain of E. coli and then thousands of modified versions of ~ A were trans- ~ / 

formed back into the yeast transport mutants. The yeast were then screened under new 

growth conditions that wild-type ~ A ~ does / not permit ~ ~ growth. l For example, we 

identified cells that acquired the ability to grow in the presence of transport competitors 

and others that grew on very low concentrations of histidine. This kind of positive selection 

identified mutant forms of the porter that exhibit new transport properties while 

eliminating those that simply lost transport activity. Although loss of function mutants 

might be revealing, far too many would inhibit transport activity for unspecific reasons, 

such as premature te~ination of the peptide. Thus far, we have identified important 

residues in TMDs 1,5,6,8, and 11. ~reliminary analysis of these results suggests TMDs 

1, 5, 6, and 8 may be involved in defining the substrate translocation pathway through 

this transporter (L. Chen and D. R. Bush, unpublished data). 

The amino acid symporters described here are excellent candidates for using biotechnological 

methods to modify the nutritional value of harvested tissues. For example, many 

cereals are poor sources of protein for animals because they are low t~ptophan, in lysine, 

and threonine (Bright and Shewry 1983; z and Larkins 1991 ; Galili 1995), and legumes 

are low in cysteine and methionine (Shewry et al. 1995). One strategy for modifying 

the nutrition^ value of these deficient crops would be to use targeted expression of 

high-affinity amino acid transporters to enhance the content of specific amino acids in 

harvested seed. For instance, targeted expression MTl in the phloem of mature leaf 

tissue may increase the amount of lysine and histidine transported per unit carbon. This 

would increase the lysine and histidine content in developing seeds. The success of this 

approach is dependent on several unknown variables, such as reciprocal increases in 

mesophyll synthesis and release as these amino acids are actively transported into the 

phloem. There is precedence for increased rates of synthesis under increased demand, 

and recent work in Heldt’s laboratory suggests the mesophyll pool may be dynamically 

linked to phloem loading (Mens et al. 1991 ; Winter et al. 1992; Lohaus et al. 1994) and, 

thus, active loading by a high-affinity transporter may shift the equation to increased 

synthesis. Other examples of potential engineering include targeted expression in stem 

tissue to capture amino acids passing by in the phloem, combining ove~roduction and 

targeted expression, or targeted expression to engineer uptake of novel xenobiotics. 

Our understanding of amino acid transport has progressed si~nificantly in recent 

years. The challenges that lay ahead focus on defining the coarse and fine-con~ol mechanisms 

that regulate that function of these amino acid transporters. This knowledge will 

be the fo~ndation of our understanding of resource allocation across the plant as a multicellular 

organism, and for developing novel strategies for improving crop yields and 

nu~tion~ value. 

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R., Schulz,B.,andFeldmann,K.A. (1996). Arabidopsis AUX1 gene:Apermease-like 

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10-17.

David Rhodes 

Purdue Unjversj~, West ~fa~ette, Indiana 

~n~ve~si~ of M~sso~~i, Colu~bia, Missouri 

1. INT TI 

This chapter will consider the central role of amino acid metabolism in abiotic stress 

resistance of plants. We will attempt to ~ighlight progress made since the last comprehensive 

review of this field by Stewart and Larher (1980). Major emphasis will be placed 

on the role of amino acid metabolism in the synthesis of compatible osmolytes. This 

class of molecules includes certain amino acids (notably proline), quaternary ammonium 

compounds (e.g., glycinebetaine, prolinebetaine, P-alaninebetaine, and choline-0-sulfate), 

and the tertiary sulfonium compound 3-dim~thylsulfoniopropionate (DMSP), The 

quaternary ammonium compounds and DMSP are derived from amino acid precursors. 

These compounds share the property of being uncharged at neutral pH and are of high 

solubility in water (Ballantyne and Chamberlin 1994). Moreover, at high concen~ations 

they have little or no perturbing effect on macromolecule-solvent interactions (Yancey 

et al. 1982; Low 1985; Somero 1986; Timasheff 1993; Yancey 1994). Unlike perturbing 

solutes (such as inorganic ions) that readily enter the hydration sphere of proteins, favoring 

unfolding, compatible osmolytes tend to be excluded from the hydration sphere of 

proteins and stabilize folded protein structures (Low 1985). These compounds are 

thought to play a pivotal role in plant cytoplasmic osmotic adjustment in response to 

osmotic stresses (Wyn Jones et al. 1977). 

The synthesis of quate~ary ammonium and tertiary sulfonium compounds requires 

participation of the “activated methyl cycle’’ to provide methyl groups for the onium 

moieties of these compounds (Hanson et al. 1995; Bohnert and Jenson 1996). Therefore, 

attention is given to the role of sulfur amino acid metabolism, and in particular, the role 

of methionine and ~-adenosylmethionine (SAM) in the biosynthesis of these methylated 

onium compounds. The central position of SAM in stress metabolism in plants is further 

illustrated by briefly considering its role as a precursor of the hormone ethylene (Kende 

1983) and of the polyamines spermine and spermidine (Flores et ai. 1989). The role of 

sulfur amino acid metabolism in plant stress-resistance mechanisms is further explored 

in relation to the p~icipation of cysteine as a precursor of peptides (glutathione and 

319

phytochelatins) that chelate heavy metals (Rauser 1990, 1995) and facilitate detoxification 

of herbicides (Li et al. 1995; Kreuz et al. 1996) and active oxygen species (Alscher 

1989; Foyer et al. 1995; Smirnoff 1995). 

The amino acids alanine and y-aminobutyrate (CABA) accumulate markedly in 

response to anaerobic stress in plants (Streeter and Thompson 1972); these amino acids 

are specifically discussed in relation to anaerobic carbon metabolism, and intracellular 

pH regulation. Because it is now recognized that CABA synthesis is catalyzed by a 

C~-calmodulin-activated enzyme, glutamate decarboxylase (Snedden et al. 1995), we 

will consider the specific role of GfU3A connecting intermediary amino acid metabolism 

to perturbations of cytoplasmic Ca" concentrations induced by stress. 

Because proline is treated in a separate chapter in this volume (see Chap. 8), and has 

been the subject of numerous reviews over the last 20 years (see, e.g., Stewart and Larher 

1980; Thompson 1980; Stewart 1981; Hanson and Hitz 1982; Csonka and Baich 1983; 

Rhodes 1987; Delauney and Verma 1993; Samaras et al. 1995), including a comprehensive 

recent review (Hare and Cress 1997), our treatment of this amino acid will be brief. 

Proline accumulation represents a common metabolic responses of higher plants to water 

deficits and salinity stress. This highly water-soluble imino acid is accumulated to osmotically 

significant levels by leaves of many halophytic higher-plant species grown in saline 

environments (Stewart and Lee 1974; Treichel 1975; Briens and Larher 1982), in leaf 

tissues and shoot apical meristems of plants experiencing water stress (Barnett and 

Naylor 1966; Boggess et al. 1976; Jones et al. 1980), in desiccating pollen (Hong-qi et 

al. 1982; Lansac et al. 1996), and in cultured plant cells adapted to water stress (Handa 

et al. 1986; Rhodes et al. 1986), or NaCl stress (Katz and Tal 1980; Treichel 1986; 

Binzel et al. 1987; Rhodes and Handa 1989; Thomas et al. 1992). In the apical millimeter 

of maize roots, proline represents a major solute, reaching concentrations of 120 mM in 

roots growing at a water potential of -1.6 MPa (Voetberg and Sharp, 1991). The accurnulated 

proline accounts for a significant fraction ("50%) of the osmotic adjustment in this 

region (Voetberg and Sharp 1991). Proline accumulation in maize root apical meristems 

in response to water deficits involves increased proline deposition to the growing region 

(Voetberg and Sharp 1991) and requires abscisic acid (ABA; Ober and Sharp 1994; 

Sharp et al. 1994). 

At high concentrations, proline protects membranes and proteins against the adverse 

effects of inorganic ions and temperature extremes (reviewed in Samaras et al. 

1995). Proline may also function as a hydroxyl radical scavenger (Smirnoff and Cumbes 

1989). Its synthesis is implicated as a mechanism of alleviating cytoplasmic acidosis, 

and it may maintain NADP+/NAPH ratios at values compatible with metabolism (Hare 

and Cress, 1997). Rapid catabolism of proline after relief of stress may provide reducing 

equivalentsthatsupportmitochondrialoxidativephosphorylationandthegeneration of 

ATP for recovery from stress and repair of stress-induced damage (Hare and Cress, 1997). 

In vivo-labeling studies with 14C-labeled precursors (Moms et al. 1969; Oaks et al. 

1970; Boggess et al. 1976), ["CC]glutamate (Hayser et al. 1989a,b) or ''NH~/''NO~ 

(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 

increase of proline biosynthesis rate (Boggess et al. 1976; Stewart, 1981; Rhodes et al. 

1986; Rhodes and Handa 1989). Proline accumulation involves induction of enzymes of 

proline biosynthesis (Peng et al. 1996; and see Chap. 8), possibly coupled with a relaxation 

of proline feedback-inhibition control of the pathway (Boggess et al. 1976; Stewart 

1981), decreased proline oxidation to glutamate (Stewart et al. 1977; Stewart and Boggess 

1978; Wuang and Cavalieri 1979; Sells and Koeppe 1981; Elthon and Stewart 1982), 

mediated at least partly by down-regulation of proline dehydrogenase (Kiyosue et al. 

1996, Peng et al. 1996), decreased utilization of proline in protein synthesis (Boggess 

and Stewart 1980; Stewart 1981), and enhanced protein turnover (Fukutoku and Yamada 

1984). Water deficits induce dramatic increases in the proline concentration of phloem 

sap in alfalfa (Girousse et al. 1996), suggesting that increased deposition of proline at 

the root apex in water-stressed plants (e,g., Voetberg and Sharp 1991) could occur partly 

by phloem transport of proline (Girousse et al. 1996). A proline transporter gene ProT2 

is strongly induced by water and salt stress in Arnbidopsis t~~Ziann (Rentsch et al. 1996). 

In bacteria, proline synthesis from glutamate is catalyzed by three enzymes: y- 

glutamyl kinase (GK; EC 2.7.2.1 I), y-glutamyl phosphate reductase (GPR; glutamic semialdehyde 

dehydrogenase; EC 1.2.1 Al), and A1-pyrroline-5-carboxylate reductase (PSCR; 

EC 1.5,1.2), encoded by genes pro& proA, and yroC, respectively (Adams and Frank 

1980; Csonka and Baich 1983). In higher plants it is now clear that a similar pathway 

operates, with the exception that GK and GPR are embodied in one bifunctional polypeptide, 

A1-py~oline-5-carboxylate synthetase (PSCS), encoded by a single gene (Hu et al. 

1992; Fig. 1). The prevailing evidence is that both PSCS and PSCR are localized in the 

cytoplasm (Hare and Cress, 1997). The GK activity of the bifunctional PSCS is inhibited 

by proline (Nu et al. 1992). PSCS gene expression is induced by desiccation, salinity 

stress, and abscisic acid (ABA; Hu et al. 1992; Yoshiba et al. 1995; Peng et ai. 1996; 

Savourk et al. 1997). P5CR transcripts also increase in abundance in response to osmotic 

stress (Delauney and Verma 1990; Williamson and Slocum, 1992; Verbruggen et al. 

1993). Savourd et al. (1997) suggest that the expression of the proline biosynthetic genes 

are dependent on at least two signal transduction cascades: one triggered by exogenously 

applied ABA in the absence of stress, and the other triggered by cold and osmotic stress 

independently of exogenously applied ,ABA. 

Recently, transgenic tobacco plants overexpressing PSCS have been obtained, and 

these appear to have increased resistance to both water deficits and salinity stress (Kishor 

et al. 1995). However, insufficient data are available to conclude with certainty that 

proline accumulation in these transgenic plants contributes to their enhanced stress resistance 

by osmotic adjustment, or by some other mechanisms (Sharp et al. 1996). As discussed 

by Hare and Cress (1997), the controversial data reported for the water relations 

of these transgenic plants emphasize the need to investigate nonosmotic explanations for 

the phenotypes observed. 

A decrease in proline oxidation rate can contribute to net proline accumulation 

during drought and salinity stress. Proline is oxidized to PSC by proline dehydrogenase 

(EC 1.5.99.8) and the resulting P5C is further oxidized to glutamate by P5C dehydrogenase 

(EC 1.5.1.12; see Fig. 1). Both enzymes are localized in the mitochondrion (reviewed 

in Hare and Cress, 1997). A cDNA-encoding proline dehydrogenase (PDH) has 

recently been cloned from Arn~i~opsis and is induced by proline under nonstressed conditions, 

but strongly repressed in response to osmotic stress (Kiyosue et al. 1996). It 

appears that osmotic stress overrides proline induction of proline dehydrogenase gene

OH glutamate + 

8 

0 0 

*. 

I 

* 

*. 

NADH 

+ H+ 

NADS 

* * 

.I 

0 . 

.*.* 

PSCR 

H O 

proline - 

P 

proline 

FADH2 

+ H20 

FAD + 

c 

+ 02 

iigure 1 Pathways of synthesis and oxidation of proline in higher plants: GK, y-glutamyl kinase; 

GPR, y-glutamylphosphate reductase; PSC, A1-pyrroline-5-~arboxylate; PSCS, A'-pyrroline-S-carboxylate 

synthetase; PSCR, A'-pyrroline-5-carboxylate reductase; PDH, proline dehydrogenase; 

PSCDH, A1-pyrroline-S-c~boxylate dehydrogenase. The dotted line denotes feedback inhibition of 

the GK activity of PSCS by the end product, proline,

expression (Kiyosue et al. 1996). The reciprocal regulation of PSCS and PD 

count for the rapid accumulation of proline in response to osmotic stress and its rapid 

catabolism after stress relief (Peng et al. 1996). To our knowledge the gene encoding 

P5C dehydrogenase has not yet been cloned from higher plants, but the enzyme has 

recently been extensively purified and characterized (Forlani et al. 1997). 

Hare and Cress (1997) argue that the different subcellular localizations of proline 

biosynthesis (cytoplasm) and oxidation (mitochondrion), the NADPH cofactor preference 

of the biosynthetic enzymes, and NADH cofactor preference for the proline oxidation 

pathway, would enable proline biosynthesis to enhance activity of the cytoplasmic oxidative 

pentose phosphate pathway and provide a mechanism of interconversion of the phosphorylated 

and nonphosphorylated pools of pyridine nucleotide cofactors. They suggest 

that the osmoprotective effects of proline accumulation may be of secondary impo~ance 

to the associated metabolic implications of proline synthesis and degradation. Further 

investigations of transgenic plants, engineered for altered proline synthesis or catabolism, 

clearly need to consider not only the consequences on absolute proline level and the 

biophysical effects exerted by elevated cytoplasmic proline concentrations, but also the 

fluxes to and from the proline pool (Hare and Cress 1997). 

In growing tissue, the proline deposition rate will be equal to the combined rates 

of proline synthesis, proline release from protein, and proline import, minus the combined 

rates of proline catabolism, proline export, proline utilization in protein synthesis, 

and the rate of pool dilution caused by water uptake during growth (Rhodes and Handa 

1989; Voetberg and Sharp 1991). The latter, in turn, will be determined by fundamental 

plant-water relations of the growing region, including water potential, solute potential, 

yield threshold, turgor, and potential difference between the xylem and growing cells 

(Nonami et al. 1997), as well as the levels of growth inhibitory metabolites such ABA, as 

for which synthesis or compartmentation may respond rapidly to changes in turgor or 

metabolism (e.g., intracellular pH; Rhodes 1987). The feedback loops in this system are 

clearly complex, with ABA and osmotic signals altering expression of genes encoding 

proline biosynthesis enzymes, proline and osmotic signals altering expression of PD 

proline feedback inhibiting its own synthesis, and proline itself (if accumulated to sufficiently 

high levels) contributing to solute potential and, hence, turgor and growth maintenance. 

The full promise of transgenic plants engineered for proline metabolism toward 

an understanding of proline’s role(s) in stress resistance will likely not be realized until 

attention is given to the growing regions as well as mature tissues (cf., Kishor et al. 

1995). The consideration of growing and nongrowing tissues and the fluxes between 

them will be essential in testing the intriguing scheme, recently proposed by Hare and 

Cress (1997), in which proline and PSC might act as an intercellu1~-signaling system. 

In this scheme, proline produced from P5C in an “effector” tissue is transported by the 

phloem to a “target” tissue, characterized by a high-energy requirement, where proline 

degradation generates reducing equivalents needed to drive the tricarboxylic acid (TCA) 

cycle activity. The PSC or glutamate generated may then be translocated back to the 

effector tissue where conversion to proline regenerates the NADP’ needed to prime the 

oxidative pentose phosphate pathway. 

Stress-hypersensitive mutants of higher plants that exhibit disturbed proline metabolism 

(e.g., the sosl mutant of Ar~~i~o~sis; Liu and Zhu 1997) can contribute significantly 

to the elucidation of thesignalsto which proline accumulation may respond. 

However, we urge caution in concluding from such mutants that “Pro accumulation is a 

symptom of stress injury rather than an indicator of stress tolerance” (Liu and Zhu 1997).

324 ~ho~es et a!. 

111. IN0 ACIDS AS PREC 

Y AMMONIUM COMP 

OMPATIBLE OSMOLYTES 

The role of quaternary ammonium compounds as compatible solutes has been extensively 

reviewed (Wyn Jones and Storey 1981 ; Anthoni et al. 199 1 ; Rhodes and Hanson 1993; 

Gorham 1995). These zwitterionic compounds include glycinebetaine, choline-0-sulfate, 

P-alaninebetaine, prolinebetaine, and hydroxyprolinebetaine. They possess a fully 

methyl-substituted nitrogen atom (creating a permanent positive charge on the N moiety) 

and a negatively charged carboxyl group (in the case of betaines) or sulfate group (in 

the case of choline-0-sulfate). The function of these compounds as osmoprotectants is 

illustrated by their stimulation of the growth of Escherichia coli or ~aZ~oneZZa ~ ~ ~ i ~ 

rium in saline medium (Strom et al. 1983; Hanson et al. 1991, 1994a). In these assays, 

glycinebetaine, P-alaninebetaine, prolinebetaine, and choline-0-sulfate at 1-mM concentrations 

are more effective than 1 mM of proline (Hanson et al. 1991, 1994a). Choline, 

an alcohol without a negatively charged group, is not an osmoprotectant per se. Although 

it stimulates the growth of wild-type E. coli in a saline medium, it must be oxidized to 

glycinebetaine to function as an osmoprotectant. Thus, betA mutants of E. coli, defective 

in choline oxidation, are unable to use choline as an osmoprotectant (Hanson et al. 1991). 

Similar to proline, glycinebetaine and structurally related compounds have stabilizing 

effects on proteins and membranes (Rhodes and Hanson 1993), and are implicated in 

frost protection (Kishitani et al. 1994; Nomura et al. 199Sb). 

In higher plants, choline is derived from the amino acid serine, presumably by decarboxylation 

of serine to ethanolamine (Fig. 2). The NH?-moiety of ethanolamine is then N- 

methylated to (CH&-P- at the level of either free ethanolamine bases, O-phosphorylethanolamine 

bases, or 0-phosphatidylethanolamine bases, catalyzed by S-adenosylmethionine 

(SAM)-dependent N-methyltransferases (Rhodes and Hanson 1993; Corham 

1995; Hanson et .al. 1995). Datko and Mudd (1988a,b) have proposed phospho~lethanolamine 

(PE) as a common co~itting step in the synthesis of choline moieties in plants, 

implicating ethanolamine kinase (EC 2.7.1.82) as a key enzyme in this pathway (see 

Fig. 2). 

In glycinebetaine-accumulating chenopods, the main pathway of choline synthesis 

appears to be by the phosphoryl-bases: PE, phospho~lmonomethylethanolamine (PMME), 

phosphoryldimethylethanolamine (PDME), and phosphorylcholine (PC) (Hanson and 

Rhodes 1983; Hanson et al. 1995; see Fig. 2). Choline is then liberated from PC by a 

phosphorylcholine phosphatase (Rhodes and Hanson 1993; Hanson et al. 1995; see 

Fig. 2). 

In grasses, PC is incorporated into phosphatidylcholine from which choline is then 

released (Giddings and Hanson 1982; Hitz et al.1.991),presumably by the action of 

phospholipase-D (Rhodes and Hanson 1993). 

Choline down-regulates its own synthesis at the level of the PMME and PDME N- 

methyltransferases (Mudd and Datko 1989a,b). These enzymes are induced by salinity 

stress in spinach (Weretilnyk and Summers 1992; Hanson et al. 1995; Weretilnyk et al. 

1995). PC may feedback-inhibit its own synthesis in sugar beet (Hanson and Rhodes 

1983).

6iotic Stress ~esista~ce 

ethanolamine @A) 

. 

e 

. 

0 

transferase 

0 SAHC 

0 

* 

. -0 

e N- 

H 

O 

phospho~lmonometh~lethanolamine 

(PIMME) 

phospho~ldimethylethanolamine 

(PDME) 

phospho~lcholine (PC) 

+- 

choline 

Figure 2 Pathway of choline synthesis in the Chenopodiaceae: The dotted line denotes feedback 

regulation of the first N-methyltransferase by choline. 

Glycinebetaine is synthesized from choline in a two-step oxidation catalyzed by a ferredoxin 

(Fd)-dependent choline rnonooxygenase (CMO) and a betaine aldehyde dehydrogenase 

(BADH; EC 1.2.1.8) with a strong preference for NAD' (Rhodes and Hanson 1993; 

Fig. 3).

,-,. betaine ~dehyde 

+ H20 - H20 

0 - betaine aldehyde 

NAD+ 

NADH + Elt 

~~ycinebetai~~ 

Pathway of glycinebetaine synthesis in the Chenopo~iacea~: Fd, fe~e~oxin. 

th enzymes are predominantly localized in the chloroplast stroma of chenopods 

et al. 1986, 1988; Brouquisse et al. 1989), although in spinach leaves approximately 

10% of the BADH may exist as a cytosolic isofom (~eretilnyk and Hanson 

1988). ~lycinebetaine is also localized predominantly in the chloroplasts of salinized 

spinach leaves, where it provides osmotic adjustment (Robinson and Jones 1986). Oxygen-1 

8 tracer studies confirm that CMO uses molecular oxygen as a substrate (Lema et 

al. 1988). 

Both CMO (Burnet et al. 1995) and BADH (~eretilnyk and Hanson 1989) have 

been purified to homogeneity from spinach. Complementary DNAs encoding BADH 

n isolated from spinach (Weretilnyk and Hanson 1990) and sugar beet (McCue 

son 1992). Salinity stress leads to a two- to threefold increase of CMO and 

ene expression, and concomitant increases 

enzyme level (Hanson et al. 1995). 

ADH has an unusual chloroplast-transit target sequence (Rathinasabapathi 

ansgenic tobacco plants overexpressing chloroplast-localized BADH have 

been obtained (Rathinasabapathi et al. 1994). However, these are unable to accumulate 

glycinebetaine in the absence of exogenously supplied betaine aldehyde because tobacco 

lacks C ~ (~athinasabapathi 

O 

et al. 1994). A cDNA encoding CMO has recently been 

cloned from spinach (Rathinasabapathi et al. 1997). The cDNA sequence confirms that 

is an Fe-S protein, and that CMO has a typical chloroplast-transit peptide sequence 

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 

glycinebetaine in salinity and drought stress resistance. 

It should be cautioned, however, that BADH may be a general aldehyde dehydrogenase, 

acting on other aldehyde substrates in addition to betaine aldehyde (Trossat et al. 

1997).BADHwilluse 3-dimethylsulfoniopropionaldehyde, an intermediate in DMSP 

synthesis (see Sec. 1V.B) and certain aldehydes (3-aminopropionaldehyde and 4-aminobutyraldehyde) 

involved in polya~ne metabolism (Trossat et al, 1997). Transgenic 

plants engineered for BADH expression (Holmstrom et al. 1994; ~athinasabapathi et al. 

1994) may possibly exhibit phenotypes related to perturbed polyamine metabolism, in 

addition to phenotypes attributable to alterations in the glycinebetaine synthesis pathway 

(Trossat et al. 1997). The multiple substrate specificities of BADH may explain the 

occurrence of BADH in plants that do not accumulate glycinebetaine (Ishitani et al. 

1993; Weretilnyk et al. 1989), and in organs of glycinebetaine-accumulating plants that 

do not contain glycinebetaine (e.g., roots of cereals; Ishitani et al. 1995). 

In monocotylednous plants, recent studies suggest that BADH is localized in peroxisomes 

(Nakamura et al. 1997). All monocotyledenous BADHs have a COOH-terminal 

tripeptide (SKI.,) that is a signal for targeting preproteins to microbodies (Nakamura et 

al. 1997). This raises the question of whether the subcellular localization of the choline 

oxidation pathway is the same in grasses as in chenopods. 

In bacteria, such as E. coli, choline is an osmoprotectant only because it is oxidized 

to glycinebetaine. Mutants in which choline fails to serve as an osmoprotectant prove to 

have defects in choline transport (betT), choline oxidation (betA), betaine aldehyde oxidation 

(betB), or a regulatory locus (betl) encoding a repressor protein (Lamark et al, 1991, 

1996). The entire E. coli bet gene cluster has been introduced into the freshwater cyanobacterium 

~y~ec~ococc~s and has conferred glycinebetaine accumulation and increased 

salt tolerance, partly owing to stabilization of photosystem I1 (PS-11; Nomura et al. 

1995a). However, this increased salt tolerance appears to depend on the presence of an 

exogenous supply of choline. Transgenic plants expressing the E. coli betA gene encoding 

choline dehydrogenase putatively capable of oxidizing both choline and betaine aldehyde,arereportedtohaveincreasedsalttolerance 

(Liliusetal.1996).However,no 

evidence was presented that these transgenic plants actually accumulated glycinebetaine. 

In maize, several naturally occurring glycinebetaine-de~cient inbred lines have 

been identified (Brunk et al. 1989). These lack the ability to accumulate glycinebetaine 

in leaf tissue in response to either salinity stress or water deficits owing to a single 

recessive gene (betl), which has been mapped to the short arm of chromosome 3 near 

the centromere (Rhodes et al. 1993; rang et al. 1995). Glycinebetaine deficiency in 

maize is associated with an inability to oxidize ['4C]choline to ~'4C]glycinebetaine, but 

an unimpaired ability to oxidize [2H3]betaine aldehyde to [2H3]glycinebetaine, suggesting 

a lesion at the CMO step in the biosynthetic pathway (Lema et al. 1991). Homozygous 

glycinebetaine-de~cient maize lines appear to be more salt-sensitive than near-isogenic 

homozygous glycinebetaine-containing lines (Saneoka et al. 1995), and they exhibit 

greater membrane injury and damage to PS-I1 in response to heat stress (Yang et al. 

1996). ~lycinebetaine-deficient maize lines exhibit a signi~cantly elevated pool of free 

choline; however, choline does not accumulate to levels equal to the glycinebetaine level 

of glycinebetaine-containing lines, suggesting that choline must down-regulate its own 

synthesis (Yang et al. 1995). Consistent with this, glycinebeta~ne-deficient lines of maize 

exhibit an elevated level of serine in comparison with homozygous glycinebetaine-containing 

lines (Yang et al. 1995).

Clycinebetaine is thought to be the archetypal betaine in the plant kingdom (Weretilnyk 

et al. 1989; Hanson and Burnet 1994; Hanson et al. 1994a). Some species have apparently 

lost the capacity to accumulate large quantities of this compound, and others have 

evolved the capacity to synthesize alternative quaternary a~onium compounds, including 

choline- sulfate, p-alaninebetaine, or prolinebetaine. This is particularly well 

illustrated in a consideration of the quaternary ammonium compounds accumulated by 

membersofthePlumbaginaceae (Hansonetal.1994a).Ancestralsubfamilies of the 

Plumbaginaceae are glycinebetaine-accumulators (Hanson et al. 1994a). In more advanced 

subfamilies, however, glycinebetaine has been replaced in part or in whole by p- 

alaninebetaine or prolinebet~ne (Hanson et al. 1994a). In addition, all members of the 

Plumba~inaceae accumulate choline-0-sulfate. This compound is synthesized from choline 

by a salt stress-inducible cholinesulfotransferase (EC 2.8.2.6; Rivoal and Hanson 

1994b; Fig. 4). 

The adaptive significance of choline-0-sulfate in the Plumbaginac~ae is thought to 

reside in the fact that salt glands of these species can excrete chloride, but not sulfate; 

thus, conjugation of sulfate with choline is proposed to be a mechanism of sulfate detoxification, 

converting a normally deleterious anion to a compatible osmolyte (Hanson et 

al. 1994a). Because choline- sulfate synthesis competes with glycinebetaine synthesis 

for available choline, this may have cont~buted to the evolution of alternative betaine 

biosynthetic pathways (p-alaninebetaine or prolinebetaine) that draw on substrates other 

than choline (Hanson et al. 1994a). 

p-Alaninebetaine is confined to species of the Plurnbaginaceae (Hanson et ai. 1994a). It 

is synthesized by direct ~-methylation of the amino acid p-alanine, presumably catalyzed 

by SAM-dependent ~-methyltransferases (Hanson et al. 1991, 1994a; Fig. 5). In contrast 

with glycinebetaine synthesis, p-alaninebetaine does not require oxygen (Hanson et al. 

1994a). It is possible, therefore, that this pathway represents an adaptation to anoxic 

saline environments (Hanson et al. 1994a). The ~-alaninebetaine biosynthetic pathway is 

HO 

I 

3-phosp~oadenosi~~- 

5'phosphosulfate (PAPS) 

3-phosphoadenosine- 

0 5 'phosphate (PAP) 

- G~oli~e-0-sulfate 

Pathway of choline-0-sulfate synthesis in the Plumbaginaceae.

0 

ll 

HO NW, @-alanine 

SAM 

SAHG 

0 

"0 - @-~~inebetaine 

ure 5 Putative pathway of P-alaninebetaine synthesis in the Plumbaginaceae. 

active in both leaves and roots of p-alaninebetaine-accumulating Li~oniu~ species 

(Hanson et al. 1991 ). The precise origin of p-alanine remains obscure. This amino acid 

apparently is not derived from decarboxylation of aspartate in Li~o~2i~~ 

species (Hanson 

and Burnet 1994). 

In some higher-plant species (notably members of the Labiateae, Asteraceae, Fabaceae, 

Bataceae, Rutaceae, Plumbaginaceae, and Capparaceae), proline can be metabolized to 

prolinebetaine (stachydrine; Naidu et al. 1987, 1992; Wood et al. 1991; Hanson et al. 

1994a; Bonham et al. 1995; Gorham 1995; Blunden et ai. 1996; McLean et al. 1996; 

Nolte et al. 1997). The synthesis of prolinebetaine is thought to involve N-methylation of 

proline by SAM[-dependent methyltransferases through the intermediate N-methylproline 

(hygric acid; Robertson and Marion, 1960; Essery et al. 1962), but the enzyme(s) have 

not yet been characterized (Fig. 6). Hydroxyprolinebetaine often (but not always) accumulates 

with prolinebetaine in these prolinebetaine-accu~ulating species (Wood et al. 

1991; Hanson et al. 1994a; Noltet al. 1997). The proline and N-methylproline hydroxylase(s) 

putatively involved in hydroxyproIinebetaine synthesis have not yet been characterized. 

Comparisons of related species with different capacities for accumulation of prolinebetaine 

and hydroxyprolinebetaine suggest that the free-proline pool size is inversely 

related to the total prolinebetaine plus hydroxyprolinebetaine pool size (Hanson et al. 

1994a; Nolte et al. 1997). Prolinebetaine is a more potent osmoprotectant than proline 

(Hanson et al. 1994a). Conversion of proline to prolinebetaine, therefore, may enhance 

osmotic stress resistance. In principle, increased flux of proline to N-methylated derivatives 

should alleviate P5CS from feedback regulation by proline by decreasing the pool

33 o~es et a/. 

HO 

. P5C P5C 

. 

. 

0 

sA.Hc 

SAM 

SAHC 

6 Putative pathway of prolinebetaine biosynthesis. 

size of free proline (See Sec. 11; and also Chap. 8), simultaneously restricting proline 

oxidation and, hence, osmolyte catabolism (Samaras et al. 1995). One isomer of hydroxyprolinebetaine 

(~~~~~-4-hydroxy-~-prolinebetaine) is a potent inhibitor of animal acetylcholinesterase 

(Friess et al. 1957), raising the possibility that the ac~umulation of this 

compound may serve a dual function: as a compatible osmolyte and an antifeedant, 

affording protection against herbivores (Hanson and Burnet 1994). In contrast, proline is 

a stimulant of insect herbivory (Haglund 1980). 

Pipecolatebetaine is accumulated together with prolinebetaine in certain species; notably 

~e~icago and Ac~i~Zea (Wood et al. 1991; Bonham et al. 1995). Pipecolic acid is often 

accumulated to high levels by members of the Fabaceae (Stewart and Larher 1980; Rosenthal 

1982), and indeed, free pipecolic acid is a major constituent of the free amino 

acid pool of ~e~icugo (Wood et al. 1991). The putative pathway of pipecolatebetaine

SAM 

SAHC 

SAM 

SAHC 

ure 7 Putative pathway of pip~colatebetaine biosynthesis. 

synthesis is illustrated in Figure 7. It is unknown if the proline and N-methylproline N- 

methyltransferases have dual substrate specificity, and if they are also capable of N- 

methylating pipecolic acid and N-methylpipecol~c acid, respectively. Pipecolic acid is 

derived from lysine catabolism by ~-aminoadipic semialdehyde (Gon~alves-Butruille et 

al. 1996), which spontaneously cyclizes to ~'-pipe~dine-6-c~boxylate. The latter can 

then be reduced to pipecolic acid in a reaction analogous with that catalyzed by P5CR 

(Stewart and Larher 1980; Rosenthal 1982; Galili 1995). 

tiv 

The synthesis of the foregoing ~uatern~y a~monium compounds is dependent on the 

activity of SAM-dependent ~-methyltransferases for N-methylation of ethanolamine 

bases, p-alanine, proline, and pipecolic acid. This implies that osmolyte accumulation is 

critically dependent on the supply of SAM as a methyl donor (Hanson et al. 1995; 

Bohnert and Jenson 1996). When SAM is used as a methyl donor in ~-~ethyl~ansferase 

reaction, the resulting ~-adenosylhomocysteine (SAHC) is recycled to homocysteine 

(HC) by adenosylhomoc~steinase (EC 3.3.1.1). HC can then accept a methyl group from 

the ~5-~ethy1-tetrahy~ofolate (~~-m~thyl-THF) pool to regenerate methionine by the 

action of methionine synthase (EC 2.1.1.14) and, thereby, SAM by the action of S 

synthetase (methionine adenosyltransferase; EC 2.5.1.6) (Giovanelli et al. 1980; Fig. 8). 

The foregoing cycle is known as the act~v~te~ ~et~yl cycle (Bohnert and Jenson

T€€F 

ethionine 

TEE 

~o~ocysteine 

Ns-me~yl-THF (CH,-THF) 

FAD 

FADH, 

glycine NH, + CO, glycine serine 

N5,Nl0-~ethylene-T~F 

.. 

N5,~l0-methe~yl-~HF 

formate 

+ THE: 

Alternative pathways of synthesis of ~5-methyl-tetr~ydrofolate as methyl donor for 

methionine synthesis from homocysteine: GDC, glycine decarboxylase; SHMT, serine hydroxymethyltransferase. 

hydrolase (EC 3.5.4.9), and ~5,~10-methenyl-THF dehydrogenase (EC 1.5.1 S ) occur on 

a single polypeptide called C1-THF synthase (Prabhu et al. 1996). In plants, however, 

the synthetase activity occurs on a protein moiety separate from that of the cyclohydrolase 

and reductase, which occur on a bifunctional protein (Prabhu et al. 1996). Because 

the latter reactions are reversible, this allows an equilibrium between the pools of Cl 

units at the formyl and methylene levels of oxidation (Cossins 1987). Therefore, serine, 

glycine, and formate can serve as the precursors for methionine synthesis (Cossins 1987; 

see Fig. 9). [13CG]Formate is metabolized to p-['3C]serine in Ar~bi~opsis in a manner 

consistent with C1-THF synthase and SHMT activities interacting with a common THF 

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 

Bregoff 1958; Hitz et al. 1981). Formate can be derived from nonenzymatic Hz02-mediated 

decarboxylation of glyoxylate, although Kleczkowski and Givan (1988) question 

whether this pathway is of physiological significance. 

The sulfonium analogue of P-alaninebetaine, 3-dimethylsulfoniopropionate (DMSP), is 

synthesized and accumulated to osmotically significant levels in a few higher-p~ant species 

from diverse families; certain species of the Asteraceae (most notably WoZZasto~ia 

b$ora), and Poaceae (some [but not all] species of S~arti~ and Sacc~ar~~) 

(Paquet et 

al. 1994; J.ames et al. 1995b), and by many marine algae (see, e.g., Blunden and Gordon 

1986; Dickson and Kirst 1986; Gage et al. 1997). DMSP is as effective as glycinebetaine 

as an osmoprotectant for E. coli (Paquet et al. 1994), and is noted to be a potent cryoprotectant 

(~ishiguchi and Somero 1992; Karsten et al. 1996). 

In ~oZZaston~ff, DMSP is synthesized by methylation of methionine to S-methylmethionine 

(SMM) in the cytosol (Hanson et al. 1994b; James et al, 1995a,b), transport of 

SMM into the chloroplast, dec~boxylation and deamination of SMM to DMSP-aldehyde 

(James et al. 1995b), and oxidation of the aldehyde to DMSP in the chloroplast (Trossat 

et al. 1996; Fig. 30). The first enzyme of the pathway, SA~~L-methionine methylt transferase 

(EC 2.1.1.12) has been purified from WoZlastoni~ leaves (James et al. 1995a). 

The final step in the pathway is catalyzed by a chloroplast-localized DMSP-aldehyde 

dehydrogenase, which may be identical with the BADH involved in glycinebetaine synthesis 

(James et al. 1995b; Trossat et al. 1996, 1997; Vojtechovd et al. 1997). The enzymes 

responsible for metabolism of SMM to DMSP-aldehyde are not yet known. 

In marine algae, DMSP synthesis occurs by a route that is entirely different from 

that in higher plants (Gage et al. 1997; Fig. 11). Methionine is first transaminated to 4- 

methylthio-2-oxobutyrate (MTOB), which is then reduced to 4-methylthio-~-hy~oxybutyrate 

(MTHB). S-Methylation of MTHB then yields 4-dimethylsulfonio-2-hydroxybutyrate 

(DMSWB), which is subs~uently oxidatively decarboxylated to DMSP (Gage et al. 

1997; see Fig. 11). 

Because DMSP does not contain nitrogen, the accumulation of DMSP could offer 

advantages over accumulation of nitrogenous onium compounds for osmotic adjustment 

in N-li~ting environments; N-deficiency increases DMSP levels ~oZZasto~i~ 

in (Wanson 

et al. 1994b) and marine algae (Gage et al. 1997). The enzyme DMSP lyase generates 

the gas dimethyl sulfide (DMS) and the potentially toxic compound, acrylate, from 

DMSP. Acrylate generated from DMSP may play a role in defense against herbivores 

(Wolfe et al. 1997). 

S-Adenosylmethionine serves as a precursor of the plant hormone ethylene, implicated 

in the control of numerous developmental processes (Lieberman 1979; Yang and Hoffman 

1984; Yang 1989; Kende 1993). SAM is converted to the amino acid 1 -aminocyclopropane-1-carboxylic 

acid (ACC) by ACC synthase (EC 4.4.1.14), which is then oxidized 

to ethylene by ACC oxidase (Kende 1993; Fig. 12). 

The HCN generated from the catalytic action of ACC oxidase is detoxified by rapid

‘S 

~~~:L-methionine 

~-methyltransf~ras~ 

OH c ethionine 

SAM: 

SAHC 

tr~s~inatio~ 

& dec~boxylation 

3-dime~ylsulfonio- 

(~~SP-alde~yde) 

Figure 10 Pathway of synthesis of dimethylsulfoniopropionate (DMSP) in ~ollffstonjff ~~orff: 

SAM, ~-adenosyl~ethionine; SAHC, S-adenosylhomocysteine. 

metabolism to the amino acid P-cyanoalanine that can then be hydrated to asparagine or 

conjugated to y-glut~yl-P-cyanoalanine (Yang 1989). 5’-~ethylthioadenosine is metabolized 

back to methionine by 5’-methyl~hio~bose and MTOB by the methionine salvage 

pathway (Miyazaki and Yang 198’7; see Fig. 12). ACC synthase may be regulated by its 

substrate SAM through S~-induced inactivation. This mechanism may be responsible 

for the short half-life of this enzyme in vivo (Yang 1989). ACC oxidase requires Fe2” 

for catalytic activity and, in some systems, is activated by C02 (Kende 1993). The plant 

species ~ ot~~ogeto~ ~ecti~at~s is unusual in that it lacks ACC oxidase and ethylene 

biosynthesis; this species is consequently rich in ACC (Summers et al. 1996). 

1 -Aminocyclopropane- 1 -carboxylate synthase is induced by various environmental 

stresses, including mechanical stress (wounding or mechanical impedance), drought, 

chilling, anoxia, and hypoxia (Yang 1989; Kende 1993; He et al. 1996a). ACC oxidase 

isalsoinduced by hypoxia and by mechanical impedance in maize roots (He et al. 

1996a); however, with the combination of hypoxia plus mechanical impedance, ethylene 

formation and ACC synthase induction exhibit a greater synergism than ACC oxidase 

induction, cellulase induction, and aerenchyma formation (He et al. 1996a). ~erenchyma 

formation accelerates the transfer of O2 from the aerial tissues to the ~2-de~cient tissues

ho~es et al. 

4-methylthio-2- 

OH hydro~bu~rate (MTHB) 

OH 

4-dime~ylsulfonio-2- 

hydro~ybu~rate ( ~ ~ S ~ ) 

ure 11 Proposed pathway of synthesis of di~ethylsulfoniopropionate (DMSP) in marine 

algae. 

of the stem base and root of flooded plants (He et al. 3996b). Ethylene production in 

response to hypoxia is implicated in the signal transduction pathway leading to cell death 

and lysis in the root cortex resulting in aerenchyma formation in maize roots (He et al. 

1996b; Drew 1997). Because ACC oxidase is oxygen-dependent, ACC can accumulate 

markedly in roots exposed to anoxia. In flooded tomato plants, the accumulated ACC is 

transported from the root to the shoot (through the xylem) where it is then rapidly oxidized 

to ethylene, inducing epinasty (Bradford and Yang 1980; English et al, 1995) and 

adventitious root formation (Visser et al. 1996). The latter may partly involve an ethylene-mediated 

increase in the sensitivity of root-forming tissue to endogenous indoleacetic 

acid (Visser et al. 1996). 

ACC can be conjugated to malonyl-ACC (MACC). MACC synthesis is catalyzed 

by ACC ~-~alonyltransferase (Yang1989;MartinandSaftner1995). An alternative 

ACC conjugate has recently been identified in tomato fruits: (y-gluta~yla~ino)-cyclo-

337 

ATP 

I 

\ 

methio~ine 

salvage 

pathway 

PPi + Pi 

'S 

I 

Ad 

Smethyl- 

~hioadenosin~ 

(rnA.1 

OH 

C,Hd + C02 + HCN + 2H20 

Figure 12 Pathway of ethylene (C21-&) biosynthesis in plants, and pathway of recycling of 5'- 

methylthioadenosine (MTA) to methionine (the methionine salvage cycle) in plants: ACC, 1- 

aminocyclopropane-1-carboxylate; MTR, ~ethylthio~bose; MTOB, 4-methylthio-2-oxobutyrate; 

SAMS, ~-adenosylmethionine synthetase. 

propane-1-carboxylic acid (GACC; Martin et ai. 1995). MACC and GACC formation 

may contribute to the regulation of ACC levels and, thereby, of ethylene formation (Yang 

1989; Kende 1993; Banga et al. 1996). 

The diamine putrescine, the trimine spermidine, and the tetramine spermine are ubiquitous 

in plant cells (Smith et al. 1979; Bagni and Pistocchi 1992). They occur as free 

cations and as conjugates with phenolic acids and macromolecules (Galston and Sawnhey 

1990). Their levels increase greatly in response to environmental stresses, most notably 

under conditions of potassium deficiency, water deficits, salinity stress, anaerobiosis, 

and acid stress (Flores et al. 1989). Because polyamines are synthesized by amino acid 

decarboxylation reactions that consume €I?, polyamine accumulation may function as 

part of a homeostatic mechanism to keep intracellular pH at a constant value (Flores et 

al. 1985). Polyamines may also play a role in the regulation of DNA replication and cell 

division, and they are implicated in the control of senescence and morphogenesis (Evans 

and Malrnberg 1989; Galston and Sawnhey 1990). 

Arginine decarboxylase (ADC; EC 4.1.1.19)-(a chloroplast-localized enzyme; 

Borrell et al. 1995)"is induced by a variety of stresses (most notably potassium deficiency; 

Watson and Malmberg 1996) and is thought to be the enzyme primarily responsible 

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 

the soluble enzyme (Malmberg et al. 1992). The two polypeptides are linked by disul~de 

bonds to form the active enzyme (Watson and Malmberg 1996). The oat (Bell and Malmberg 

1990), tomato (Rastogi et al. 1993), and Ar~bi~o~si~ atso son and Malmberg 1996). 

C genes have been cloned. In Ar~bi~o~sis, ADC enzyme levels do not correspond 

to ADC protein amounts, suggesting post~anslational modification of the enzyme 

in response to potassium deficiency (Watson and Malmberg 1996). In osmotically 

stressed oat leaves, spermine inhibits posttranslational processing of the ADC precursor, 

with subsequent decreases in mature ADC (Borrell et al. 1996). 

a~~atine 

~athways of synthesis and metabolism of putrescine.

The alternative, more direct, pathway of synthesis of putresine through ornithine 

decarboxylation catalyzed by cytosolic ornithine decarboxylase (ODC; EC 4" 1.1.17; see 

Fig. 13) is proposed to be of little importance in stress-induced putrescine accumulation, 

but may be critical in regulation of developmental processes (Galston and Sawhney 1990; 

Walden et al. 1997). Increased putrescine biosynthesis, mediated by transgenic ODC, 

promotes somatic embryogenesis in carrots (Bastola and Minocha 1995). The diamine 

cadaverine, derived from the amino acid lysine by decarboxylation, may play an important 

role in root development (Gamarnik and Frydrnan 1991). 

The triamine spermidine is synthesized by condensation of putrescine with decarboxylated 

SAM, synthesized by SAM decarboxylase (EC 4.1.1.50; Flores et al. 1989): 

SAM + Ht SAM decarboxylase 

> decarboxylated SAM + C02 

The condensation of decarboxylated SAM and putrescine is catalyzed by spermine 

synthase (putrescine aminopropyltransferase; EC 2.5.1.16). Further condensation of spermidine 

with decarboxylated SAM, catalyzed by spermine synthase (EC 2.5.1.22), produces 

the tetramine, spermine (Flores et al. 1989; see Fig. 13). Recently, the assumption 

that spermidine synthase and spermine synthase are two separate enzymes in plants, has 

been questioned (Bagga et al. 1997). The putrescine aminopropyltransferase of alfalfa 

may be responsible not only for spermidine and spermine synthesis, but also the synthesis 

of the uncommon polyamines, thermospermine, homocaldopentamine, and homocaldohexamine 

(Bagga et al. 1997). 

Antisense SAM decarboxylase potato plants show markedly altered phenotypes, 

including stunted growth; short internodes; stem branching and small leaves; decreased 

SAM decarboxylase transcripts; a reduction in SAM decarboxylase activity; reductions 

in the levels of putrescine, Spermidine, and spermine; and a marked increase in ethylene 

production (Kumar et al. 1996). This supports the idea that there is a competitive interaction 

between polyamine and ethylene biosynthesis (Walden et al. 1997). 

In addition to serving as a precursor of spermidine and spermine, putrescine has 

three other metabolic fates: conjugation with hydroxycinnamic acids, metaboli~m to y- 

aminobutyrate (GABA), and utilization in alkaloid biosynthesis, as indicated in Figure 

13. Hydroxycinnamic amide (HCA) conjugates of polyamines accumulate markedly in 

the floral apex during flower development and are implicated in the development of 

competence to flower; certain mutants of tobacco that are deficient in HCAs are unable 

to flower, and male sterile mutants of maize lack HCA accumulation in anthers (Flores 

et al. 1989). A novel polyamine conjugate, ~-hexanoylspermidine, has recently been 

identified in senescing pea ovaries and petals (Perez-Amador et al. 1996). GABA can be 

derived from putrescine (through y-aminobutyraldehyde) through the reactions catalyzed 

by diamine oxidase and y-aminobutyraldehyde dehydrogenase (Flores et al, 1989). As 

noted in Section III.B, the latter enzyme may be the same as BADH involved in glycinebetaine 

synthesis (Trossat et al. 1997). Putrescine serves as precursor of the nicotine and 

tropane alkaloids (Smith et al. 1979; Flores et al. 1989; Hashimoto and Yamada 1994), 

which may play important roles in plant defense against herbivores (see Fig. 13). 

The tripeptide glutathione (GSH), is synthesized from the amino acids glutamate, cysteine, 

and glycine by the enzymes y-glut~ylcysteine synthetase (EC 6.3.2.2) and glutathione 

synthetase (EC 6.3.2.3; Meister 1983):

Glutamate + cysteine + ATP 

~Glutamylcysteine 

synthetase 

> y-glutamylcysteine + ADP + Pi 

y-glutamylcysteine + glycine + ATP 

synthetase > glutathione + ADP + Pi 

Glutathione (GSH)playsa key role in detoxification of certainxenobioticsin 

higher plants through the action of glutathione S-transferases (EC 2.5.1 I) 18; Breaux 1986; 

Pickett and Lu 1989; Rossini et al. 1996). The glutathione conjugate of a xenobiotic can 

then be transported into and sequestered by the vacuole (Li et al. 1995; Kreuz et al. 

1996; Marrs 1996). Glutathione S-transferases are induced by a wide range of chemical 

agents (Ulmasov et al. 1995), wounding, heavy metals, ethylene, and ozone (Marrs 

1996). These enzymes may have initially evolved to conjugate naturally occurring compounds, 

such as phenylpropanoids (Dean et al. 1995), and naturally occurring endogenous 

reactive electrophilic compounds (Marrs 1996), but may have then been recruited 

to detoxify anthropogenic exogenous xenobiotics (Kreuz et al, 1996; Mans 1996). 

Under a variety of environmental stresses (e.g., high light and low temperature, 

water deficits, or others that disrupt membrane-bound electron transport systems), plants 

produce a range of active oxygen species, including superoxide, hydrogen peroxide, and 

the hydroxyl radical (Smirnoff 1993). Superoxide is converted to hydrogen peroxide by 

the action of superoxide dismutase. The hydroxyl radical can then form from the reaction 

of hydrogen peroxide with superoxide, particularly in the presence of iron or other transition 

state metals (Srnirnoff 1993). These toxic species can also form when plants are 

exposed to ozone, SO2, NO, and NO2. They react with membranes, proteins, and nucleic 

acids (Smirnoff 1993). Glutathione is involved in quenching these free radicals through 

the ascorbate-GSH cycle (Alscher 1989; Smirnoff 1993, 1995). Thus, hydrogen peroxide 

can be detoxified by the action of ascorbate peroxidase (EC 1.11.1.11), dehydroascorbate 

reductase (EC 1.8.5.1), and glutathione reductase (EC 1.6.4.2): 

Ascorbate + H202 Ascorbate peroxidase > H20 + dehydroascorbate 

~ehydroascorbate 

Dehy~oascorbate 

reductase 

+ GSH ascorbate + oxidized glut at hi on^ (GSSG) 

GSSG + NADPH + H+ Glutathione reductase 

> GSH + NADP' 

Strictly speaking, the initial product of the ascorbate peroxidase reaction is monodehydroascorbate, 

which can then either be reduced back to ascorbate by the action of an 

NAD(P)H-dependent monodehydroascorbate reductase (EC 1.6.5,4), or directly reduced 

back to ascorbate by an electron from PS-I (Smirnoff 1995). Any monodehydroascorbate 

not so reduced disproportionates to ascorbate plus dehydroascorbate (Smirnoff 1995); it 

is only the latter that is reduced by glutathione in the reactions depicted in the foregoing. 

Specific isoforms of the enzymes of this cycle are chloroplast-localized and are induced 

by an array of environmental factors (Alscher 1989; Pastori and Trippi 1992; Smirnoff 

1995). Overexpression of glutathione reductase leads to an increase in antioxidant capacity 

and resistance to photoinhibition in poplar (Foyer et al. 1995). However, overproduction 

of ascorbate peroxidase in the tobacco chloroplast does not appear to provide protection 

against ozone (Torsethau~en et al. 199'7).

Although the chloroplast-localized ascorbate-glutathione cycle is well characterized, 

it is now becoming clear that the enzymes of this cycle are also found in mitochondria 

and peroxisomes, and they may represent an important antioxidant protection system 

against H203 generated in these organelles (Jimhez et al. 1997). 

Glutathione transferases may also be involved in conjugating electrophiles generated 

from the reaction of hydroxyl radicals with lipids and DNA and, thus, contribute to 

oxidative stress resistance (Marrs 1996). Apoplastic GST induction may be mediated by 

an oxidative signal in soybean (Flury et al. 1996). 

F. P ~ y ~ o c ~ ~ 

of Heavy 

Glutathione serves as a precursor of peptides known as' phytochelatins9 which are composed 

of two or more repeating y-glutamylcysteine units, with a terminal glycine residue; 

(~-glutamylcysteine)~-gly, where n = 2-11 (Steffens et al, 1986; Reese and ~agner 

1987). The enzyme responsible for the synthesis of these peptides is known as phytochelatin 

synthase (Rauser 1990, 1995). Phytochelatins (PCs) play an important role in the 

detoxification of certain heavy metals (p~ticularly cad~ium) in plants (Rauser 1990, 

1995; Howden et al. 1995a,b). Phytochelatin synthase is activated by cadmium (Rauser 

1995). In certain plants (notably legumes) that can synthesize homoglutathione, in which 

g-alanine is substituted for glycine as the terminal amino acid, homophytochelatins are 

synthesized along with PCs in response to Cd (Klapheck et al, 1995).Inmaizeand 

certain other species of the Poaceae, a third family of phytochelatins has been found in 

which serine is the COOH-te~inal amino acid (Rauser and ~euwly 1995). ~admium 

detoxification may require transport of the Cd-phytochelatin complexes into the vacuole; 

a transport system has been recently described for these complexes (Salt and Rauser 

1995). 

In Rubia tinctoru~, phytochelatins (class I11 metallothionein) are induced by many 

metal ions, but only a few (Ag, Cd, and Cu) are bound to the PCs that they induce 

(Maitani et al. 1996). 

v. A 

A 

During anaerobic stress, pyruvate must be metabolized to either lactate (by lactate dehydrogenase; 

LDH) or ethanol (by pyruvate decarboxylase [PDC] and alcohol dehydrogenase 

[ADH]) to regenerate the NAD" required to sustain glycolysis and associated ATP 

production (Davies 1980). Lactate fermentation may cont~bute to the acidification of the 

cytoplasm (Roberts et al, 1984; Ratcliffe 1995), which may progressively inhibit LDH 

and activate PDC, promoting alcohol fe~entation (Rivoal and Hanson 1994a). Accumulation 

of pyruvate under anaerobic conditions may shift the equilibrium of alanine aminotransferases 

toward alanine accumulation; a rapid response of plants to anaerobic stress 

(Streeter and Thompson 1972; Stewart and Larher 1980): 

Pyruvate + amino acid 3 alanine + 2-keto acid

An alanine a~notransferase (g1utamate:pyruvate a~notransferase; EC 2.6.1.2) is induced 

under anaerobic conditions in barley roots (Good and Crosby 1989; Good and 

uench 1992). The precise function of alanine accumulation under anoxia is unknown, 

butitisspeculatedthatit may serve as a storage form of pyruvate (perhaps in the 

vacuole), controlling supply of pyruvate to LDH and PIX and, thereby, the flux to 

lactate and ethanol (Good and Crosby 1989; Good and Muench 1992). The synthesis of 

alanine may occur at the expense of the acidic amino acids, glutamate and aspartate 

(Streeter and Thompson 1972; Stewart and Larher 1980), and occurs concomitantly with 

the accumulation of GABA (see following Section VI). 

. 

The accumulation of the amino acid y-~nobutyrate (GABA) is markedly stimulated 

under anaerobic conditions (Streeter and Thompson, 1972; Stewart and Larher 1980; 

Aurisano et al. 1995; Ratcliffe 1995; Drew 1997). Although G A can be derived from 

putrescine (see Sec. IV.D), the anaerobic stress-induced accumulation of GABA is 

thought to result primarily from the catalytic action of glutamate decarboxylase (EC 

4.1.1.15). In part, the stimulation of glutamate decarboxylase may be the consequence of 

cytoplas~c acidification; glutamate decarboxylase has an acid pH optimum (-5.8; Patterson 

and Graham 1987): 

Glutamate + H+ ‘lutarnate 

> GABA + CQ2 

ause this reaction is proton-consuming, it could represent an adaptive response conuting 

to regulation of cytoplasmic pH (Patterson and Graham 1987; Crawford et al, 

atcliffe 1995). Crawford et al. (1994) have shown that weak acids causing cytoplasmic 

acidification also induce GABA accumulation, and they conclude that this response 

is consistent with a role for GABA synthesis in active pH regulation. 

y-A~nobutyrate can be transaminated with pyruvate to yield alanine (or with 2- 

oxoglutarate to yield glutamate), generating succinic semialdehyde, which can then be 

metabolized to succinate by the action of succinate semialdehyde dehydrogenase (Vandewalle 

and Qlsson 1983; Patterson and Graham 1987; Shelp et al. 1995; Fig. 14). The 

latter enzymes are mitochondrial (Hear1 and Churchich 1984; Breitkreuz and Shelp 1995) 

and have alkaline pH optima (Patterson and Graham 1987). GABA accumulation promoted 

by cytosolic acidi~cation may partly result from inhibition of GABA-transaminases. 

The conversion of glutamate to succinate by the action of glutamate decarboxylase, 

GABA ~ansa~nases, and succinic semialdehyde dehydrogenase is known as the 

GABA shunt (Vandewalle and Olsson 1983; Patterson and Graham 1987; Breitkreuz and 

Shelp 1995; Shelp et al, 1995), affording an alternative pathway for glutamate entry into 

the TCA cycle (see Fig. 14). 

The accumulation of GABA is induced 

response to a sudden decrease in temperature 

(Wallace et al. 1984; Patterson and Graham 1987), heat shock (Mayer et al. 1990), 

mechanical manipulation (Wallace et al. 1984), and water stress (Rhodes et al. 1986). 

Rapid GABA accumulation in response to wounding may play a role in plant defense 

against insects (Ramputh and Brown 1996). 

Glutamate decarboxylase is a cytosol-localized enzyme (Breitkreuz and Shelp

glutamate 

NH, 

2-oxo 

HO 

succinate 

Figure 14 Pathway of synthesis and metabolism of GABA. 

1995) and has recently been shown to be a calmodulin-binding protein that is 

C~~al~odulin activated (Baum et al. 1993; Ling et al. 1994; Baum and Fridmann 

1996; Arazi et al. 1995; Snedden et al. 1995). Crawford et al. (1994) note that reduced 

cytosolic pH values increase Ca”* levels, and rapid and transient increases in Ca”’ levels 

occur in response to mechanical stress and cold stress; conditions that elicit rapid GABA 

accumulation (Wallace et al. 1984). The Ca”+-calmodulin activation of glutamate decarboxylase 

provides a link between intermedi~ amino acid metabolism and perturbations 

of cytosolic ea2+ (Ling et al. 1994; Baum and Fridmann 1996; Arazi et al. 1995; Snedden 

et al. 1995) that regulate a host of other metabolic activities (Allan and Trewavas 1985; 

Bush 1995). 

The only other enzyme of glutamate metabolism currently known to be stimulated 

by Ca” in plants in glutamate dehydrogenase (GDH; EC 1.4.1.2), a ~itochondrial enzyme 

(Turano et al. 1997): 

NH~ + 2-oxoglutarate + NADH + H+ < dehydrogenase > glutamate + NAD+ 

A calcium-binding domain has been identified in the f3-subunit of GDH encoded 

by GDH2, but not in the a-subunit of GDH encoded by GDHI in Arabidopsis, suggesting 

that the different isoforms of the hexameric GDH composed of different combinations 

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 

role in plant “stress” resistance, by osmotic adjustment and the accumulation of 

compatible osmolytes; the detoxi~cation of active oxygen sp 

heavy metals; and intracellular pH regulation. The advances since 1980 have been primarily 

in the cloning of the genes encoding key enzymes of these biosynthetic pathways 

and the characterization of their regulation in relation to environmental stimuli. As noted 

by Bohnert et al. (1993, the use of metabolic engineering holds great promise for testing 

hypotheses conce~ing the role of radical scavenging and osmolyte accumulation as 

stress-protection mechanisms (e.g., Foyer et al. 1995; Kishor et al. 1995). These traits 

are not restricted to amino acid metabolism; significant advances have also been achieved 

in engineering of superoxide dismutase, and the accumulation of polyols and fructans as 

compatible solutes or radical scavengers that may limit injury (Bohnert et al. 1995; Shen 

et al. 1997, and references cited therein), These traits may eventually need to be combined 

with modified ion uptake, exclusion or co~partmentation, facilitated water permeability, 

molecular chaperones, modified membrane properties, or stress-signaling pathways 

to realize their full potential (Bohnert et al. 1995; Bohnert and Jensen 1996). The 

next decade is likely to witness further exciting progress in these areas. In the testing of 

transgene effects on plant resistance to low water potentials and salinity stress, we urge 

(1) careful measurements of growth and water relations to quantify the degree of stress 

and to assess potential growth penalties of transgene expression, (2) consideration of 

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Amino Acid Transport

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