Biochem. J. (2011) 436, 15–34 (Printed in Great Britain)
15
doi:10.1042/BJ20110078
REVIEW ARTICLE
The remarkable diversity of plant PEPC (phosphoenolpyruvate carboxylase):
recent insights into the physiological functions and post-translational
controls of non-photosynthetic PEPCs
Brendan O’LEARY*, Joonho PARK* and William C. PLAXTON*†1
*Department of Biology, Queen’s University, Kingston, ON, Canada, K7L 3N6, and †Department of Biochemistry, Queen’s University, Kingston, ON, Canada, K7L 3N6
PEPC [PEP (phosphoenolpyruvate) carboxylase] is a tightly
controlled enzyme located at the core of plant C-metabolism
that catalyses the irreversible β-carboxylation of PEP to form
oxaloacetate and Pi . The critical role of PEPC in assimilating atmospheric CO2 during C4 and Crassulacean acid
metabolism photosynthesis has been studied extensively. PEPC
also fulfils a broad spectrum of non-photosynthetic functions,
particularly the anaplerotic replenishment of tricarboxylic acid
cycle intermediates consumed during biosynthesis and nitrogen
assimilation. An impressive array of strategies has evolved to
co-ordinate in vivo PEPC activity with cellular demands for
C4 –C6 carboxylic acids. To achieve its diverse roles and
complex regulation, PEPC belongs to a small multigene family
encoding several closely related PTPCs (plant-type PEPCs), along
with a distantly related BTPC (bacterial-type PEPC). PTPC
genes encode ∼110-kDa polypeptides containing conserved
serine-phosphorylation and lysine-mono-ubiquitination sites, and
typically exist as homotetrameric Class-1 PEPCs. In contrast,
BTPC genes encode larger ∼117-kDa polypeptides owing to a
unique intrinsically disordered domain that mediates BTPC’s tight
interaction with co-expressed PTPC subunits. This association
results in the formation of unusual ∼900-kDa Class-2 PEPC
hetero-octameric complexes that are desensitized to allosteric
effectors. BTPC is a catalytic and regulatory subunit of Class2 PEPC that is subject to multi-site regulatory phosphorylation
in vivo. The interaction between divergent PEPC polypeptides
within Class-2 PEPCs adds another layer of complexity to
the evolution, physiological functions and metabolic control
of this essential CO2 -fixing plant enzyme. The present review
summarizes exciting developments concerning the functions,
post-translational controls and subcellular location of plant PTPC
and BTPC isoenzymes.
INTRODUCTION
crystal structures for Zea mays (maize) C4 PTPC and Escherichia
coli PEPC, which, in combination with site-directed mutagenesis,
have uncovered many of the structure–function relationships of
PEPC catalysis, allosteric control and regulatory phosphorylation
[8,9]. Vascular plant BTPC genes encode larger 116–118-kDa
polypeptides exhibiting low (<40 %) sequence identity with
PTPCs [3,4,6,7,11]. BTPCs lack the distinctive N-terminal serinephosphorylation motif of PTPCs, and appear to only exist as
catalytic and regulatory subunits of extraordinary Class-2 PEPC
heteromeric complexes whose novel structure, properties, control,
expression patterns and putative functions are discussed below.
Extensive biochemical and genetic studies of photosynthetic
PTPC isoenzymes of C4 and CAM leaves have elucidated catalytic
and regulatory features that are well suited for their function
as an atmospheric CO2 -concentrating mechanism that reduces
photorespiration to improve overall photosynthetic efficiency up
to 2-fold relative to C3 leaves [10,12–14]. The post-translational
control of photosynthetic PEPCs appears to be largely achieved
by allosteric activation and inhibition by glucose 6-phosphate
and malate respectively, as well as phosphorylation catalysed
PEPC [PEP (phosphoenolpyruvate) carboxylase] (EC 4.1.1.31)
is an important enzyme situated at a crucial branch point
in plant carbohydrate metabolism. It catalyses the irreversible
β-carboxylation of PEP in the presence of HCO3 − to yield
OAA (oxaloacetate) and Pi using Mg2 + as a cofactor. The
enzyme is present in all plants, green algae and cyanobacteria,
most archaea and non-photosynthetic bacteria, but is absent from
animals and fungi [1,2]. Plant PEPCs belong to a small multigene
family encoding several PTPCs (plant-type PEPCs), along with
at least one distantly related BTPC (bacterial-type PEPC) [3–7].
PTPC genes encode closely related 105–110-kDa polypeptides
that have a conserved N-terminal serine-phosphorylation domain
and typically exist as homotetrameric Class-1 PEPCs [8,9].
PTPCs have been categorized as being either photosynthetic
[C4 and CAM (Crassulacean acid metabolism) PEPCs] or nonphotosynthetic (C3 ) isoenzymes. All PTPCs evolved from a
common ancestral gene and display a high degree of conservation
at the genetic level [10]. Katsura Izui and colleagues solved the
Key words: 14-3-3 protein, mono-ubiquitination, phosphoenolpyruvate carboxylase, protein phosphorylation, protein–protein
interaction.
Abbreviations used: BTPC, bacterial-type phosphoenolpyruvate carboxylase; CAM, Crassulacean acid metabolism; co-IP, co-immunopurification; COS,
castor (Ricinus communis ) oil seed; FHA, forkhead-associated; FP, fluorescent protein; GOGAT, glutamate 2-oxoglutarate aminotransferase; GS, glutamine
synthetase; MDH, malate dehydrogenase; ME, malic enzyme; NLS, nuclear localization signal; OAA, oxaloacetate; 2-OG, 2-oxoglutarate; PA, phosphatidic
acid; PDHp , plastidic isoenzyme of the pyruvate dehydrogenase complex; PEP, phosphoenolpyruvate; PEPC, PEP carboxylase; PFK, phosphofructokinase;
PKc , cytosolic pyruvate kinase; PP2A, protein phosphatase 2A; PPCK, PEPC protein kinase; PTM, post-translational modification; PTPC, plant-type PEPC;
RT, reverse transcription; TCA, tricarboxylic acid; USP-2, ubiquitin-specific protease-2.
1
To whom correspondence should be addressed (email [email protected]).
c The Authors Journal compilation c 2011 Biochemical Society
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Figure 1
B. O’Leary, J. Park and W. C. Plaxton
The diverse functions of plant PEPC
by a specific Ca2 + -independent serine/threonine kinase known
as PPCK (PEPC protein kinase), and dephosphorylation by a
PP2A (protein phosphatase 2A) [8,13,15,16]. Phosphorylation at
the conserved N-terminal serine residue activates the enzyme by
decreasing inhibition by malate while increasing activation by glucose 6-phosphate. A number of excellent reviews and book
chapters concerning various aspects of the evolution, molecular
and biochemical characteristics, and physiological functions and
regulation of photosynthetic and non-photosynthetic PEPCs
and their corresponding PPCKs have appeared over the last
15 years [8–10,12,13,15–21]. Development of a more complete
account of PEPC’s diverse and ubiquitous involvement in plant
metabolism has recently made considerable progress owing to the
use of non-photosynthetic model plant systems such as developing
seeds and heterotrophic suspension cell cultures, together
with increased access to state-of-the-art genomic, bioinformatic
and proteomic tools. The first part of the present review
summarizes established and proposed metabolic roles for nonphotosynthetic plant PEPCs. The second section outlines recent
advancements in our understanding of the post-translational
control, protein–protein interactions and subcellular location of
non-photosynthetic PEPCs.
DIVERSE METABOLIC FUNCTIONS FOR NON-PHOTOSYNTHETIC
PLANT PEPC
PEPC and primary carbon metabolism
Besides its fundamental role in the initial fixation of atmospheric
CO2 during C4 and CAM photosynthesis, PEPC has a wide range
of non-photosynthetic roles including supporting carbon–nitrogen
interactions, seed formation and germination, fruit ripening,
guard cell metabolism during stomatal opening and provision
of malate as a respiratory substrate for symbiotic N2 -fixing
bacteroids of legume root nodules (Figure 1). The traditional
view of non-photosynthetic PEPC function is its ubiquitous
anaplerotic carboxylation of PEP needed to replenish TCA
c The Authors Journal compilation c 2011 Biochemical Society
(tricarboxylic acid) cycle carbon skeletons that are withdrawn to
support anabolism. Although certainly valid, this oversimplifies
the contribution of PEPC to primary metabolism for several
reasons. First, the TCA ‘cycle’ does not always operate as a
continuous cycle in certain plant cells. Flux through one part of
the pathway may be negligible or disproportionately low (e.g.
as occurs to various extents in illuminated and senescing leaves
and developing seed embryos) [22–25]. The partial TCA ‘cycle’
thus resembles a linear pathway with PEPC as a component
enzyme, and the notion of anaplerosis is unnecessary as the
maintenance of OAA concentrations is of no more importance
than any other metabolite [22]. Also, when combined with
MDH (malate dehydrogenase) and NAD-ME (NAD + -dependent
malic enzyme), PEPC forms an alternative flux mode which
can function in place of PKc (cytosolic pyruvate kinase) to
generate pyruvate (Figure 2A) [22,26,27]. This endows plants
with the ability to respire OAA generated from PEPC, again
not an anaplerotic function. Secondly, as discussed below, the
organic acids generated by PEPC have diverse roles outside
the TCA cycle [28]. Thirdly, when coupled with the cytosolic
MDH reaction, the metabolism of PEP to malate and other
organic acids has important implications in the redox balance
of NAD(P)H/NAD(P) + [29]. Malate transported from the
cytosol and oxidized to pyruvate within ‘sink’ organelles (e.g.
mitochondria or plastids) by ME isoenzymes functions as both a
NAD(P)H shuttle and a carbon source (Figure 2).
A distinguishing feature of the anaplerotic PEPC reaction is that
it represents a highly flexible aspect of primary plant metabolism.
Several studies have quantified in vivo flux changes through
primary metabolic pathways in isolated plant cells acclimatized
to environmental perturbations such as a change in oxygen
concentrations [30], temperature [31,32], glucose availability
[33] or nitrogen source [24]. As the overall rates of flux and
biosynthesis changed, the ratio of carbon flux through most
nodes within primary metabolism was relatively rigid, whereas
anaplerotic fluxes were relatively plastic. In other words, changes
in carbon partitioning at the PEP branchpoint (Figure 2A) are
Functional control of non-photosynthetic plant PEPC
17
crucial in allowing plant cells to synchronize their metabolism
with changing environmental conditions. Control of PEPC
activity clearly occurs at both the transcriptional/translational and
post-translational levels. Transcript and enzymological studies
have shown that the supply of exogenous sugars, Pi and/or
nitrogen influences both the amount and phosphorylation status of
PEPC in a wide range of tissues, including developing Hordeum
vulgare (barley) seeds and COS [castor oil (Ricinus communis)
seeds] [34–36], Solanum lycopersicum (tomato) cell cultures [33],
Arabidopsis thaliana (thale cress) cell cultures and seedlings
[37,38], Lotus japonicus and Glycine max (soya bean) root
nodules [39,40], Solanum tuberosum (potato) seedlings [41] and
Triticum aestivum (wheat) leaves [42].
PEPC plays a central role in the control of plant glycolysis and
respiration
Figure 2
Models illustrating several metabolic functions of plant PEPC
(A) Interactions between carbon and nitrogen metabolism involves three compartments in
plants. This scheme highlights the important role of the two terminal enzymes of plant
cytosolic glycolysis, PEPC and PKc , in controlling the provision of the mitochondria with
respiratory substrates, as well as for generating the 2-OG and OAA respectively required for
NH4 + -assimilation via GS/GOGAT in plastids and aspartate amino transferase in the cytosol. The
co-ordinate control of PEPC and PKc by allosteric effectors, particularly glutamate and aspartate,
provides a mechanism for the regulation of cytosolic glycolytic flux and PEP partitioning during
and following NH4 + -assimilation as discussed in the text. The central role of PEP in providing
‘bottom-up’ control of glycolysis is also illustrated. Broken arrows with a circled plus and minus
sign indicate enzyme activation and inhibition respectively by allosteric effectors. (B) Alternative
metabolic routes for the conversion of sucrose into fatty acids in developing oil seeds. This
model illustrates the role of PEPC in controlling PEP partitioning to malate as a source of carbon
skeletons and reducing power for leucoplast fatty acid synthesis. Abbreviations are as defined
in the text, in addition to the following: AAT, Asp aminotransferase; E.T.C., electron transport
chain; MEm /PDHm and MEp /PDHp , mitochondrial and plastidic isoenzymes of ME and PDH
respectively.
In animals, primary control of glycolytic flux of hexose
phosphates to pyruvate is mediated by ATP-dependent PFK
(phosphofructokinase), with secondary control at PK (pyruvate
kinase). Activation of PFK increases the level of its product,
fructose 1,6-bisphosphate, which is a potent feed-forward
allosteric activator of many non-plant PKs [27]. In contrast,
quantification of changes in levels of glycolytic intermediates
that occur immediately following stimulation of respiration in a
wide variety of plant systems (including green algae, ripening
fruit, aged storage root slices, germinating seeds and suspension
cell cultures) consistently demonstrated that plant glycolysis is
controlled from the ‘bottom up’ with primary and secondary
regulation exerted at the levels of PEP and fructose 6-phosphate
utilization respectively (Figure 2A) [27]. These findings are
compatible with the observation that plant ATP-PFKs demonstrate
potent allosteric feedback inhibition by PEP. It follows that any
enhancement in the activity of PEPC or PKc will relieve the PEP
inhibition of ATP-PFK, thereby elevating glycolytic flux from
hexose phosphate (Figure 2A). Reduced cytosolic PEP levels
also cause elevated levels of the signal metabolite fructose 2,6bisphosphate (and thus activation of the PPi -dependent PFK)
since PEP is a potent inhibitor of plant 6-phosphofructo-2kinase [27]. A possible advantage of bottom-up regulation of
glycolysis is that it permits plants to control net glycolytic flux
independent of related plant-specific metabolic processes such
as the Calvin–Benson cycle and sucrose–starch interconversion.
The hypothesis that plant glycolysis is regulated from the bottom
up was corroborated by the application of metabolic control
analysis to assess the distribution of respiratory flux control in
tubers of transgenic potato plants expressing different amounts
of E. coli ATP-PFK [43]. It was determined that ATP-PFK
exerts a low flux control coefficient over both glycolysis and
respiration, whereas far more flux control was exerted in the
metabolism of PEP. The relatively low flux control coefficient of
ATP-PFK was explained as a consequence of its potent inhibition
by PEP. In this way, PEPC and PKc play a central role in the
overall regulation of plant respiration, since the control of their
activities in vivo will ultimately dictate the rate of mobilization of
sucrose or starch for respiration, while simultaneously controlling
the provision of: (i) respiratory substrates (e.g. pyruvate and/or
malate) for ATP production via oxidative phosphorylation, and
(ii) TCA cycle carbon skeletons needed for nitrogen assimilation
or as biosynthetic precursors. It is obvious that PEPC and
PKc represent promising targets for metabolic engineering of
plant respiration and photosynthate partitioning. For example,
transgenic bean plants were generated that expressed a bacterial
malate-insensitive PEPC in a seed-specific manner [44]. Analysis
c The Authors Journal compilation c 2011 Biochemical Society
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B. O’Leary, J. Park and W. C. Plaxton
of the transgenic seeds indicated enhanced PEP partitioning
through the PEPC anaplerotic pathway, resulting in a shift of
photosynthate partitioning from sugars/starch into organic acids
and amino acids. Consequently, the transgenic seeds accumulated
up to 20 % more storage protein [44].
PEPC produces malate as an alternative respiratory substrate for
plant mitochondria
Malate has been frequently proposed to be an important
respiratory substrate for plant mitochondria, such that a significant
fraction of pyruvate enters the TCA cycle via the combined
reactions of PEPC, MDH and NAD-ME, rather than PKc
(Figure 2A). However, in vivo flux studies of plant cells incubated
with 13 C-labelled substrates demonstrated that the flux of malate to
pyruvate via mitochondrial NAD-ME is usually very low, relative
to the flux of PEP to pyruvate via PKc [22,27]. Thus a marked
reduction in mitochondrial NAD-ME in transgenic potatoes and
Arabidopsis had little impact on respiratory metabolism [45,46].
It was concluded that the import of PEPC-derived malate into
the mitochondria generally serves an anaplerotic role to support
biosynthesis and nitrogen assimilation, whereas pyruvate derived
from the PKc reaction is the most significant substrate for
respiration [45,46]. Nevertheless, there are specific situations,
such as prolonged Pi starvation [47] or the light-enhanced dark
respiration peak that occurs immediately after C3 leaf darkening
[48], when the oxidation of malate as a respiratory substrate
for mitochondrial ATP production appears to be of particular
significance. Similarly, overexpression of PEPC in several plant
species resulted in a stimulation of respiration when illuminated
leaves were darkened, and an increase in extractable NAD-ME
and NADP-ME activities [19,49–51]. Future studies are required
to determine the full suite of physiological and developmental
conditions that stimulate intramitochondrial pyruvate production
via the NAD-ME-dependent oxidative decarboxylation of malate
(Figure 2A).
Carbon–nitrogen interactions
PEPC plays a crucial role regulating respiratory carbon flux in
vascular plant tissues and green algae that are actively assimilating
nitrogen [27,52–54]. The organic acids supplied by PEPC have
several distinct roles within nitrogen metabolism. α-Oxo acids,
especially OAA and 2-OG (2-oxoglutarate), are obligate carbon
skeletons on to which NH4 + is assimilated and exported to other
tissues (Figure 2A). In nodules, malate is also the predominant
respiratory substrate supplied to symbiotic rhizobia bacteria to
support the huge energy demand of reducing N2 into NH4 +
[55,56]. Malate and citrate may also act as counterions to replace
nitrate and maintain cytosolic pH [57]. The tight integration and
critical importance of PEPC at the interface of carbon and nitrogen
metabolism has been continuously elaborated by numerous
studies of green algae and vascular plants. Legume species
express a specific nodule-enhanced PTPC isoenzyme that is upregulated and activated by in vivo phosphorylation during active
nitrogen assimilation by N2 fixing root nodules [39,55,58–60], and
whose suppression results in greatly diminished rates of nitrogen
assimilation [55,56]. In leaves and non-nodular root tissues,
PTPC transcripts, activity and phosphorylation status are also upregulated when metabolism is switched to a nitrogen-assimilating
state following the addition of NO3 − or NH4 + [42,53,61–66].
PEPC is also up-regulated during nitrogen remobilization from
senescing leaves [25].
c The Authors Journal compilation c 2011 Biochemical Society
A key study compared metabolic fluxes of autotrophic
canola (Brassica napus) embryos supplied with organic or
inorganic nitrogen (amino acids compared with NH4 + ) [24]. The
differences in primary metabolism were mainly a restructuring
of flow through the TCA cycle caused by the reciprocal
changes in PEPC compared with mitochondrial NAD-ME
flux. The dynamic response of PEPC to different nitrogen
sources has been particularly well studied in chemostats of
unicellular green algae where nitrogen availability can be
precisely manipulated [52]. NO3 − or NH4 + resupply to N-limited
green algae leads to an immediate and massive stimulation of: (i)
respiratory carbon flux in order to generate TCA carbon skeletons
required by GS (glutamine synthetase)/GOGAT (glutamate 2-OG
aminotransferase), and (ii) dark CO2 fixation catalysed by PEPC
(Figure 2A) [52]. During transient nitrogen assimilation, over
50 % of net plant carbon may be committed to the production
of carbon skeletons and energy (ATP and reductant) required for
the GS/GOGAT system [27,52]. Green algal (Chlamydomonas
reinhardtii) PTPC and BTPC transcript and protein levels are both
up-regulated upon nitrogen starvation, presumably to facilitate
rapid provision of OAA and 2-OG when a nitrogen source
becomes available [5,67]. As discussed below, the control of
green algal and vascular plant PEPCs by allosteric effectors,
particularly glutamate and aspartate, has clearly evolved to be
in tune with carbon–nitrogen interactions [68], and to co-operate
with other enzymes involved in nitrogen assimilation, particularly
PKc , nitrate reductase and GS (Figure 2A) [53,62,69].
Regulatory phosphorylation represents a second level of
post-translational PEPC control in plant cells engaged in
active nitrogen assimilation [27,54]. Nodule-specific PPCKs
have been reported for several legume species whose activity
mirrors PEPC’s phosphorylation status and the presence of
photosynthate supply [6,39,40,70,71]. PEPC phosphorylation
in roots and leaves also responds to the carbon/nitrogen
balance due to the action of PPCK [16,37,42,66,72]. The
importance of post-translational PEPC control in mediating
carbon–nitrogen interactions has been elegantly demonstrated
by a series of metabolic engineering efforts. Various forms of
PEPC with diminished feedback inhibition have been expressed
in transgenic potatoes, Arabidopsis and Vicia narbonsis (vetch)
seeds [44,51,73,74]. These alterations increased carbon flux
into organic acids and amino acids, and boosted the overall
organic nitrogen content at the expense of starch and soluble
sugars. The mutant potato plants had diminished growth rates
and tuber yield, whereas the PEPC-overexpressing Arabidopsis
plants suffered severe stunting owing to reduced PEP levels that
limited aromatic amino acid production via the shikimate pathway
[51]. Conversely, overexpression of PEPC without diminished
feedback inhibition has little impact on overall metabolism, even
when large increases in extractable PEPC specific activity were
achieved [19,49–51]. As a further tier of integration into nitrogen
metabolism, PEPC has been shown to be a major target of the
plant-specific Dof1 transcription factor which is implicated in
organic acid metabolism [75]. Increased expression of Dof1 in
transgenic Arabidopsis caused an increase in nitrogen content
and improved growth under low nitrogen that was correlated with
increased PEPC and PKc expression and activity [75].
PEPC participates in photosynthate partitioning in developing
seeds and organic acid metabolism of germinating seeds
Developing seeds
Developing seeds are crucial metabolic sinks that import large
amounts of sugars and amino acids from leaves which must be
Functional control of non-photosynthetic plant PEPC
efficiently metabolized into starch, triacylglycerols and storage
proteins. Owing to the agronomic significance of these storage reserves, a detailed understanding of developing seed metabolism
has obvious practical applications in crop breeding and
metabolic engineering. PEP metabolism via PEPC, PKc and PKp
(plastidic pyruvate kinase) plays a prominent role in partitioning
imported photosynthate towards leucoplast fatty acid biosynthesis
compared with the mitochondrial production of carbon skeletons
and ATP needed for amino acid interconversion in support of
storage protein biosynthesis. PEPC may also aid in refixing
respired CO2 to improve overall seed carbon economy [76–80].
Increased PEPC and PPCK activity and PTPC phosphorylation
status during seed development coincides with the main stages
of storage oil and protein biosynthesis [35,78,79,81–84], and
have been shown to be controlled by photosynthate supply in
developing COS and barley seeds [34–36]. Seed storage protein
synthesis has been traditionally linked to PEPC activity owing to
the obvious requirement for OAA and 2-OG needed to assimilate
nitrogen during formation of amino acids (Figure 2A) [44,74,85].
Seeds import nitrogen in the form of amino acids, but still
require additional carbon skeletons because: (i) glutamine, a major
component of the supplied amino acids contains two amino groups
per 2-OG skeleton; and (ii) amino acids such as alanine can be
deaminated and respired through the TCA cycle, yielding NH4 +
that must be rapidly assimilated into glycine and glutamate via
GS/GOGAT (Figure 2A).
PEPC’s role in generating metabolite precursors and reducing
power to support triacylglycerol synthesis in developing oil seeds
has also received considerable attention. De novo fatty acid
synthesis occurs within the leucoplast, is ATP- and NAD(P)Hdependent, and uses acetyl-CoA as an immediate precursor.
However, the pathway leading from sucrose-derived cytosolic
hexose phosphates to plastidial acetyl-CoA is highly flexible
and certainly varies within and between species (Figure 2B)
[86,87]. Studies of isolated leucoplasts from developing COS
and Helianthus annuus (sunflower) seeds showed that, relative
to other precursors, exogenous malate supported maximal rates
of fatty acid synthesis [88,89]. As a consequence of malate’s
oxidation into acetyl-CoA via the plastidial isoenzymes of NADPME and PDH (pyruvate dehydrogenase complex), all of the
reductant required for carbon incorporation into fatty acids is
produced (Figure 2B). Both PEPC and plastidial NADP-ME
are abundant in developing COS endosperm [81,90,91], and a
specific malate/Pi antiporter exists in the COS and maize embryo
leucoplast envelope [92,93]. These results are compatible with
the hypothesis that PEPC has an important role to control sucrose
partitioning at the level of the cytosolic PEP branchpoint for generation of carbon skeletons and reducing power needed for
fatty acid biosynthesis by developing oil seeds such as COS
(Figure 2B). Consistent with this view are the high rates of dark
CO2 fixation exhibited by intact pods of developing COS [94].
Metabolic flux studies of developing oil seeds have reported
a spectrum of values regarding anaplerotic PEP carboxylation
to support biosynthetic pathways [24,26,32,95–99]. The flux
from malate to plastidial acetyl-CoA appears to be negligible in
developing green oil seeds such as canola [97], despite the strong
correlation that exists between PEPC expression and rates of oil
synthesis by developing canola embryos [83]. However, fatty acid
synthesis by green oilseed rape/canola and soya bean embryos is
light-dependent [86,100,101], since most of the reducing power
[NAD(P)H] they need to assemble fatty acids from acetyl-CoA is
generated via the light reactions of leucoplast photosynthesis.
In the non-green developing maize and sunflower embryos,
however, 30 % and 10 % of carbon flux into fatty acids was
derived from PEPC-generated malate respectively [95,96]. One
19
caveat of metabolic flux analyses is that they require artificially
static conditions, such as constant light and sucrose supply, and
non-physiological levels of oxygen [102]. Conversely, the
activities of the enzymes surrounding the PEP node and their
support of fatty acid synthesis are likely to be tightly controlled
to be in tune with changing environmental conditions, including
the diurnal cycle.
Germinating seeds
Research with starch-storing cereal seeds such as barley and
wheat, and oil-storing dicotyledon seeds such as COS have
consistently implied an important anaplerotic function for PEPC
during germination [36,81,103–107]. Cristina Echevarria and
colleagues have performed pioneering studies with PEPC of
germinated cereal seeds such as wheat and barley [105,106], in
which the aleurone layer synthesizes and secretes various acid
hydrolases needed to mobilize starch and protein reserves in
the underlying endosperm. Aleurone PEPC is thought to play
an important role in regulating the production of organic acids,
chiefly malic acid, that are excreted for the acidification of
the starchy endosperm, or transported to the growing embryo
to replenish pools of TCA cycle intermediates consumed
during biosynthesis and transamination reactions. Although
barley and wheat seed PTPC becomes progressively activated
by phosphorylation following imbibition, PPCK was already
present in the dry seeds and was apparently not up-regulated
during germination [36,105]. The phytohormone abscisic acid
appears to play a role in triggering PPCK accumulation during
the desiccation stage of maturing barely seeds [36]. Also, the
protein synthesis inhibitor cycloheximide, as well as other
pharmacological agents known to block the signal-induced
phosphorylation of PEPC in C4 leaf mesophyll cells, had no
influence on PEPC’s phosphorylation in germinated cereal seeds,
suggesting that their PPCK may be subject to a post-translational
control mechanism [36,105].
The carbon metabolism of germinating oil seeds such as COS
is dominated by the mobilization of storage lipid and protein
reserves to support the needs of the growing hypocotyls and
roots. In germinating COS endosperm, this process includes
the massive conversion of reserve triacylglycerols into sucrose,
which is absorbed by the cotyledons of the growing seedling
[108,109]. PEPC has been suggested to fulfil a crucial function
very early in the germination process to build up cellular pools of
C4 acids needed to trigger subsequent TCA and glyoxylate cycle
activity [81]. An ensuing role for PEPC in germinating oil seeds
is to replenish dicarboxylic acids required as substrates for the
substantial transamination reactions that follow storage protein
hydrolysis [108], as well as for GS’s reassimilation of NH4 +
released during the oxidative deamination of glutamate to 2-OG
via glutamate dehydrogenase [110]. The labelling of metabolic
intermediates to isotopic steady state and the modelling of the
TCA cycle in germinating lettuce seeds indicated that 70 % of
glycolytic flux from carbohydrate oxidation enters the TCA cycle
via the PEPC reaction [111]. At the same time, PEP carboxykinase
catalyses the initial step in the gluconeogenic conversion of
storage lipids into sucrose, as it uses ATP to decarboxylate and
phosphorylate OAA derived from the glyoxylate cycle into PEP
[108]. PEP carboxykinase is highly abundant in the cytosol of
germinating COS endosperm [112], but is incompatible with
PEPC, since their combined reactions result in a futile cycle,
i.e. the wasteful hydrolysis of ATP to ADP and Pi . In their
classic study of gluconeogenesis in germinating COS endosperm,
Kobr and Beevers [109] determined that the maximum rate
c The Authors Journal compilation c 2011 Biochemical Society
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B. O’Leary, J. Park and W. C. Plaxton
of glycolysis is about one-tenth that of gluconeogenesis and
that the pools of glycolytic and gluconeogenic intermediates
probably occur in separate intracellular regions and appear to be
regulated independently. Several studies have provided evidence
for glycolytic or gluconeogenic multi-enzyme complexes
(metabolons) in the plant cytosol [27], including the cytosolic
isoenzymes of fructose-1,6-bisphosphate aldolase and fructose1,6-bisphosphatase in germinated COS [113]. It will therefore
be important to establish whether PEPC mono-ubiquitination
(see below) possibly contributes to formation of a glycolytic
metabolon while minimizing its futile cycling with PEP
carboxykinase in the cytosol of the germinating oil seeds such
as COS.
PEPC mediates organic acid accumulation by developing fruit and
expanding cells
Many developing fruits accumulate substantial concentrations of
malate and citrate (>300 mM) in their vacuoles. This stockpile of
organic acids is believed to sustain respiration and act as carbon
source during ripening when sugars are accumulated [114,115].
A fruit-specific PTPC isoenzyme has been identified in tomato
whose transcripts and activity increase throughout development,
simultaneous with enhanced cellular malate and citrate levels
[115,116]. PEPC from tomato and Citrus sinensis (orange) fruits
displayed a significantly decreased sensitivity to malate inhibition
relative to many other PTPCs [116,117]. Whether this is the
result of specific isoenzyme properties or control by a PTM
(post-translational modification) such as phosphorylation remains
unknown. Adding to this uncertainty, a PPCK transcript steadily
increases in tomato fruit from early development until ripening,
but does not correlate with PEPC activity in this tissue [118]. A
Class-1 PEPC has also been purified and characterized from Musa
cavendishii (banana) fruit [119]. The PEPC of ripening bananas is
subject to regulatory phosphorylation, and was suggested to play
an anaplerotic role to replenish carbon skeletons consumed during
nitrogen assimilation and/or transamination reactions [120].
The tomato fruit PTPC is located predominantly in large
or expanding cells of the pericarp, where malate production
via PEPC and MDH has been suggested to sustain osmotic
potential to allow rapid cell expansion [116]. A similar role for
malate has been suggested for other expanding cells as well.
For example, malate accumulation in the guard cell vacuole
is involved in stomatal opening. Several studies analysing the
genetic or pharmacological alteration of PEPC activity in guard
cells have correlated stomatal opening with PEPC activity and
PTPC phosphorylation status [19,121,122]. Cotton fibres are hairlike single cells that undergo a stage of rapid elongation to several
centimetres at anthesis. PEPC activity and PTPC transcripts, and
the accumulation of malate were all correlated with the rate, extent
and developmental profile of fibre elongation [123]. Treatment of
fibres with LiCl, an inhibitor of PPCK activity, reduced fibre
length and implicated regulatory PTPC phosphorylation in this
tissue as well. Aside from acting as an osmolyte to increase
turgor pressure, the malate generated by cotton fibre PEPC may
also support fatty acid synthesis (Figure 2B) needed for the rapid
extension of the plasma membrane and tonoplast [123].
PEPC helps plants to acclimatize to abiotic and biotic stresses
PEPC has been widely implicated in plant stress tolerance in two
main ways. First, as discussed above, PEPC provides metabolic
flexibility that allows PEP metabolism to adjust to environmental
conditions. Generally, this involves either increased anaplerosis
c The Authors Journal compilation c 2011 Biochemical Society
to support specifically up-regulated biosynthetic processes and/or
a glycolytic bypass of PKc as an adjustment to differing Pi or
reductant levels. Secondly, some roots excrete huge amounts of
organic acids into the soil (rhizosphere) to acidify the soil and
chelate cations as a response to nutrient deprivation and/or toxic
metal stress. PEPC up-regulation, as documented by increased
transcripts, protein, extractable activity and phosphorylation
status appears to be an integral component of these stress
responses.
Drought, salt and ozone stress
An up-regulated PEPC and PPCK response to salinity or drought
stress has been well documented in C3 , C4 and CAM plants
[11,84,124–133]. Consistent with these reports, RNAi (RNA
interference) knockdown of a canola PTPC isoenzyme resulted
in increased sensitivity to poly(ethylene glycol)-induced osmotic
stress [134]. The adaptive value of increased PEPC activity
upon salt or drought stress requires further clarification, but
could potentially improve carbon metabolism during periods of
reduced stomatal conductance by reassimilating respired CO2
and/or increasing rates of CO2 fixation at night when stomata
are open, as in CAM photosynthesis [126,129,132]. PEPC may
also support the biosynthesis of biocompatible osmolytes such as
proline that rapidly accumulate upon drought or salinity stress in
many plant species [134].
Exposure to elevated ozone causes oxidative stress and leads to
a major rearrangement in primary carbon metabolism involving a
suppression of photosynthesis and an increase in respiration [135].
Specifically, a major up-regulation of PEPC and concomitant
down-regulation of ribulose-1,5-bisphosphate carboxylase is
typical of this response [135,136]. The up-regulation of PEPC
along with ME is hypothesized to contribute to the increased
production of NAD(P)H necessary to detoxify reactive oxygen
species, as well as the increased anaplerotic flux to support protein
synthesis for defence and repair processes [137].
Nutritional phosphate and iron deficiency
Phosphorus is an essential element for growth and metabolism
that plays a central role in virtually all metabolic processes.
Plants preferentially absorb phosphorus from the soil in its fully
oxidized anionic form, Pi . Despite its importance, Pi is one of
the least available macronutrients in many terrestrial and aquatic
environments. In soil, Pi is frequently complexed with Al3 + or
Ca2 + cations and thus exists as insoluble mineral forms that
render it unavailable for plant uptake. PEPC was suggested to
provide a metabolic bypass (with MDH and NAD-ME) to the
ADP-limited PKc to facilitate continued pyruvate supply to
the TCA cycle, while concurrently recycling the PEPC by-product
Pi for its reassimilation into the metabolism of the Pi -deficient
cells [27]. PEPC induction and its regulatory phosphorylation
during Pi stress has also been correlated with the synthesis and
consequent excretion of large amounts of malic and citric acids
by roots [27,38,138,139]. This increases Pi availability to the
roots by acidifying the rhizosphere to: (i) solubilize otherwise
inaccessible sources of mineralized soil Pi [138], while (ii) making
phosphorus esters more accessible to hydrolysis by secreted
acid phosphatases [140]. An excellent correlation of organic
acid excretion and PEPC up-regulation with enhanced rates
of dark CO2 fixation occurs in proteoid (cluster) roots of Pi deficient Lupinus albus (white lupin) [138]. Recent developments
have clarified the molecular basis of the PEPC response to Pi
stress in Arabidopsis cell cultures and seedlings [37,38,141,142],
Functional control of non-photosynthetic plant PEPC
Oryza sativa (rice) leaves [72] and white lupin proteoid roots
[143], which involves the up-regulation of specific PTPC and
PPCK isoenzymes under the control of multiple transcription
factors. The simultaneous induction and in vivo phosphorylation
activation of the PTPC isoenzyme AtPPC1 (Arabidopsis PEPC 1)
at its conserved Ser11 phosphorylation site (Figure 3) contributes
to the metabolic adaptations of Pi -starved Arabidopsis [38]. This
is consistent with the observation that the PPCK-encoding genes
AtPPCK1 and AtPPCK2 are among the most strongly induced
genes when Arabidopsis is subjected to nutritional Pi deprivation
[141,144,145].
Iron deficiency also up-regulates PEPC activity [up to 60fold in Beta vulgaris (sugar beet)], specifically in the cortical
cells of root apical meristem [146]. This was hypothesized to
support organic acid excretion to the soil for increased iron uptake
and organic acid accumulation within the plant for increased
vascular transport [146–150]. The rising intracellular citrate
concentrations that occur may inhibit PKc and thus promote flux
through PEPC, MDH and NAD-ME as a glycolytic bypass [148].
Heavy metal toxicity
The exudation of PEPC-derived organic acids also functions to
chelate metals in the rhizosphere to alleviate heavy metal toxicity
by preventing their uptake into the cell [138,151]. Al3 + toxicity
is the most prevalent form of plant heavy metal stress. Although
elevated soil Al3 + levels result in root efflux of organic acids, there
appears to be no corresponding induction of root PEPC [152,153].
However, a recent study demonstrated that overexpression of a C4
PTPC isoenzyme in rice led to increased oxalate exudation and
Al3 + tolerance [154]. In contrast with Al3 + , Cd2 + toxicity leads
to PEPC up-regulation [155,156]. This is believed to support the
synthesis of the phytochelatins, intracellular metal chelators that
rapidly accumulate during heavy metal stress. Phytochelatins are
polymers of the tripeptide glutathione whose synthesis requires
de novo amino acid synthesis and therefore increased anaplerotic
PEP carboxylation via PEPC.
Biotic stress
PEPC is also up-regulated during infection of tobacco (Nicotiana
tabacum) plants by the potato virus [157,158]. The up-regulation
may be a response to: (i) closed stomata and the ensuing alteration
of metabolism, and/or (ii) increased anaplerotic flux needed to
sustain the rapid synthesis of viral defence proteins.
THE MULTIFACETED POST-TRANSLATIONAL CONTROL OF PLANT
PEPC
21
heteromeric complexes composed of tightly associated PTPC
and BTPC subunits (discussed below) also display distinctive
kinetic properties [116,117,159,160]. Furthermore, aspartate and
glutamate are important feedback effectors of Class-1 PEPC
and PKc isoenzymes in green algae and plant tissues active
in nitrogen assimilation and/or transamination reactions. This
provides a regulatory link between nitrogen metabolism and the
control of respiratory carbon metabolism (Figure 2A). Feedback
inhibition of specific Class-1 PEPC and PKc isoenzymes by
glutamate provides a rationale for the known activation of the
two enzymes that occurs in vivo during periods of enhanced
nitrogen assimilation when intracellular glutamate concentrations
become transiently reduced [27,52]. In contrast with PEPC,
aspartate functions as an allosteric activator of various plant PKc s
by effectively relieving the enzyme’s inhibition by glutamate
(Figure 2A) [27,69,161–164]. Reciprocal control of Class1 PEPC and PKc by aspartate has been documented in a
wide range of plant cells and tissues involved in nitrogen
assimilation and/or transamination reactions including: green
algae, Spinacia oleracea (spinach) and castor oil plant leaves,
ripening banana fruit, canola cell cultures and endosperm of
developing COS [52,69,82,161–164]. This was suggested to
provide a mechanism for decreasing flux from PEP to aspartate
(via PEPC and aspartate aminotransferase) while promoting PKc
activity whenever cytosolic aspartate levels become elevated. This
may occur when the cell’s demands for nitrogen are satisfied, and
the overall rate of protein synthesis becomes more dependent on
ATP availability, rather than the supply of amino acids. In this
instance, respiration may assume a more significant role in terms
of satisfying a large ATP demand, rather than the anaplerotic
generation of biosynthetic precursors.
PA (phosphatidic acid) is a lipid second messenger that
transiently accumulates in plant cells within minutes of applying
a wide array of stress conditions [165]. Several C3 and C4
PTPC isoenzymes were recently identified as PA-binding proteins
[166,167]. Moreover, a low concentration (50 μM) of PA and
other anionic phospholipids resulted in significant inhibition
(>50 %) of C4 Class-1 PEPC activity [167]. Little or no effect
was observed with neutral, zwitterionic or positively charged
phospholipids, ruling out a non-specific effect of the fatty acyl
chains (which activate E. coli PEPC). Interestingly, hypo-osmotic
treatment of Arabidopsis cells increased PEPC’s affinity for PA
through an unknown mechanism [166]. Although most of the C4
PTPC of Sorghum bicolor (sorghum) leaves was in the soluble
fraction, a subfraction was membrane-associated [167]. Anionic
phospholipids represent novel regulators that may inhibit the
activity and/or mediate membrane association of certain PEPCs.
How PEPC activity and/or cellular location is influenced by
dynamic changes in anionic membrane phospholipids such as
PA will be an interesting area for further research.
Allosteric effectors and regulatory phosphorylation: new tricks for
old dogs
Allosteric effectors
Regulatory phosphorylation
The basic features of post-translational control of plant Class-1
PEPCs have been known since the early 1980s. Most
Class-1 PEPCs show allosteric inhibition by malate and activation
by glucose 6-phosphate. However, the enzyme’s specific kinetic
and allosteric properties are quite variable and appear to be
well adapted to the physiological role(s) of specific PEPC
isoenzymes and the nature of their cellular environment. A good
example is the increased K m (PEP) and IC50 (malate) values of
C4 Class-1 PEPC relative to typical C3 Class-1 PEPCs [10].
The C3 Class-1 PEPCs from developing fruit, the chloroplastic
Class-1 PEPC isoenzyme of rice leaves and the Class-2 PEPC
In the early 1980s, Hugh Nimmo (University of Glasgow)
and Raymond Chollet (University of Nebraska) respectively
documented the regulatory phosphorylation of CAM and C4
photosynthetic Class-1 PEPC isoenzymes at their single highly
conserved serine residue close to the N-terminus of the protein.
Since their seminal discoveries, this PTM has been implicated in
the in vivo control of a wide array of non-photosynthetic PTPCs
[34,36,38,42,106,120,168–172]. This research area has been
elaborated in several excellent reviews that have appeared within
the last decade [8,16,17]. In brief, Class-1 PEPC phosphorylation
uniformly results in enzyme activation at physiological pH
c The Authors Journal compilation c 2011 Biochemical Society
22
Figure 3
B. O’Leary, J. Park and W. C. Plaxton
Amino acid sequence alignment of Arabidopsis and castor oil plant PEPC isoenzymes
Experimentally verified in vivo phosphorylation sites of castor oil plant and Arabidopsis PTPCs (RcPPC3 and AtPPC1 respectively) and castor oil plant BTPC (RcPPC4), are highlighted in green
[34,38,191,201], whereas RcPPC3’s conserved Lys628 mono-ubiquitination site is marked with a red font [103]. BTPC’s highly divergent ∼10-kDa domain that was predicted to exist largely as an
intrinsically disordered region [191] is enclosed in a red rectangle. The proteolytically susceptible peptide bond (Lys446 –Ile447 ) within RcPPC4’s disordered region [4] is marked with an X. Boxes
I–III denote conserved subdomains essential for PEPC catalysis [9]. The predicted pI, molecular mass and sequence identity (%) of the various PEPCs are shown. The deduced PEPC sequences
were aligned using ClustalW software (http://www.ebi.ac.uk/Tools/clustalw2/index.html). Semi-colons and asterisks indicate identical and conserved amino acids respectively.
c The Authors Journal compilation c 2011 Biochemical Society
Functional control of non-photosynthetic plant PEPC
by decreasing PEPC’s sensitivity to allosteric inhibitors while
increasing its affinity for PEP and sensitivity to allosteric
activators. Hence, much attention has also been directed to the
Ca2 + -independent PPCK and, to a far lesser extent, PP2A,
that mediate Class-1 PEPC’s reversible phosphorylation in vivo.
Detailed PPCK studies, including its cloning from CAM, C4
and C3 plants, continues to provide one of the best understood
examples of regulatory enzyme phosphorylation in the plant
kingdom [16,17]. There is also compelling evidence from
several systems that Class-1 PEPC phosphorylation correlates
with increased flux through the enzyme in vivo [16]. With
molecular masses of 31–33 kDa, the PPCKs are the smallest
protein kinases yet reported, consisting of only a core kinase
domain [8,17]. Class-1 PEPC phosphorylation status appears to
be generally controlled by changes in rates of PPCK synthesis
compared with degradation (with the notable exception of
germinating cereal seeds; see above) [8,16]. This is a novel means
of control, since most protein kinases are post-translationally
controlled by second messengers such as Ca2 + ions, intracellular
signals such as AMP, and/or phosphorylation by other protein
kinases.
PPCK is up-regulated in response to a range of different
signals, including light in C4 leaves, a circadian rhythm in CAM
leaves, light and nitrogen supply in C3 leaves, nutritional Pi
deprivation in C3 plants and cell cultures, and photosynthate
supply in legume root nodules and developing seeds [16,17,35–
37,72,127,173,174]. Endogenous PPCK activity and PTPC
subunit phosphorylation of developing COS and soya bean root
nodules were both eliminated following prolonged darkness of
intact plants [6,34,35,40,70]. Both effects were fully reversed
following reillumination of darkened plants. These results imply
a direct relationship between the up-regulation of COS or
soya bean nodule PPCK and PTPC phosphorylation during the
recommencement of photosynthate delivery from illuminated
leaves to these non-photosynthetic sink tissues. The ability
of plant cells to sense sugars plays a crucial role in carbon
partitioning and allocation in source and sink tissues. These
processes are modulated as a consequence of the plant’s sugar
status, and sugar signals function both at the transcriptional and
translational levels in tight co-ordination with light and other
environmental stimuli [84]. PPCK of non-green sink tissues such
as soya bean root nodules or developing oil seeds appears to be
one of the many proteins whose expression is markedly influenced
by sucrose supply. Developing seed and legume nodule gene
expression, respiration, storage product synthesis and carbon–
nitrogen interactions are highly dependent upon the supply of
recent photosynthate [85]. A key area for future studies will
be to establish signalling pathway(s) that link sucrose supply
to enhanced PPCK expression and PTPC phosphorylation in
heterotrophic sinks such as legume root nodules or developing
seeds.
A variety of post-translational PPCK controls have also been
proposed including: (i) malate inhibition of PPCK activity,
possibly by interacting with PEPC [35,120,175], (ii) thiol–
disulfide interconversion [35,176,177], and (iii) a proteinaceous
inhibitor [178]. The mechanisms and in vivo relevance of these
controls requires further evaluation. Functional genomic studies
have evaluated the impact of in vivo PTPC phosphorylation on
the growth and development of transgenic C4 (Flaveria bidentis)
and C3 (Arabidopsis) plants in which PPCK expression and thus
in vivo PEPC phosphorylation was disrupted [171,179]. Although
perturbation of growth and/or metabolite levels were observed,
the results of both studies led to the surprising conclusion
that inhibition of PTPC phosphorylation has little impact on
metabolic flux through PEPC (albeit, in non-stressed plants).
23
In Arabidopsis, however, several pathways were perturbed,
and it was hypothesized that leaf PPCK may have several
targets in addition to PEPC [171]. The physiological roles of
PTPC phosphorylation, and the substrate specificity of PPCK
require further clarification. A PP2A appears to dephosphorylate
photosynthetic and non-photosynthetic PTPCs [8,15,16,20].
Additional research is needed to assess the possibility that the
activity of PP2A catalytic subunits are themselves controlled by
regulatory subunits with which they might be associated.
Protein phosphorylation can not only control enzymatic activity
directly, but also generate specific docking sites for other
proteins, particularly the 14-3-3 proteins, a family of highly conserved and abundant proteins that play a central regulatory
role in eukaryotic cells [27]. The 14-3-3 homodimers bind to
specific phospho-sites on diverse target proteins, thereby forcing
conformational changes or influencing interactions between
their targets and other molecules. In these ways, 14-3-3s
‘finish the job’ when phosphorylation alone lacks the power
to drive changes in the activities of intracellular enzymes. For
example, the phosphorylated forms of plant nitrate reductase and
sucrose-phosphate synthase are both potently inhibited by 14-33 binding [27]. Proteomic profiling of tandem-affinity-purified
14-3-3 protein complexes from transgenic Arabidopsis seedlings
recently identified the PTPC isoenzyme AtPPC1 as being a
14-3-3-binding protein [180]. However, plant PEPCs, including
AtPPC1, do not contain any established 14-3-3-binding motifs,
and recent attempts to demonstrate 14-3-3 binding by the purified
phosphorylated or dephosphorylated forms of native AtPPC1 [38]
using 14-3-3 far-Western overlay assays have proven unsuccessful
(C. MacKintosh and W.C. Plaxton, unpublished work). It is
possible that AtPPC1 binds to (i) 14-3-3 proteins, but does
not perform well in the overlay assay which requires protein
refolding, or (ii) another 14-3-3-interacting phosphoprotein and
thus appeared in the tandem-affinity-purified 14-3-3 complexes.
It would be of interest to employ 14-3-3 pull-down assays coupled
with PEPC immunoblotting to further assess putative 14-3-3
binding by PTPCs, and/or whether 14-3-3s exert any influence
on the kinetic and regulatory properties of AtPPC1 or other plant
PEPCs.
PEPC mono-ubiquitination: the new kid in town
Ubiquitin is a highly conserved globular protein of eukaryotic
cells that modifies target proteins via its covalent attachment
through an isopeptide bond between the C-terminal glycine
residue of ubiquitin and the ε-amino group of a lysine residue on
a target protein. A multi-enzyme system consisting of activating
(E1), conjugating (E2) and ligating (E3) enzymes attach ubiquitin
to cellular proteins. Polyubiquitination is a well-known PTM
that tags many proteins for their proteolytic elimination by
the 26S proteasome. Indeed, degradation of both the PTPC
and PPCK by the polyubiquitin–proteasome pathway has been
reported [181,182]. However, protein mono-ubiquitination has
been demonstrated recently to play a variety of crucial nondestructive functions in yeast and mammalian cells [183,184].
Mono-ubiquitination is a reversible PTM that mediates protein–
protein interactions (by recruiting ubiquitin-binding domain
client proteins) and localization to help control processes such
as endocytosis, DNA repair, transcription and translation, and
signal transduction [183,184]. Ubiquitin-related pathways are
believed to be of widespread importance in the plant kingdom.
Genomic analyses indicated that the ubiquitin-related pathway
alone comprises over 6 % of the Arabidopsis or rice proteomes
with thousands of different proteins being probable targets [185].
c The Authors Journal compilation c 2011 Biochemical Society
24
B. O’Leary, J. Park and W. C. Plaxton
Recent studies on Class-1 PEPC from germinated COS
endosperm provided the first example of regulatory monoubiquitination of a metabolic enzyme [103]. Extracts of fully mature and germinating COS endosperm had been shown to contain
an immunoreactive PTPC polypeptide that co-migrated with the
107-kDa subunit of the developing COS Class-1 PEPC
homotetramer [4,77,81]. However, an additional immunoreactive
PTPC polypeptide of approximately 110 kDa appeared
immediately following COS imbibition and persisted throughout
germination at an equivalent ratio with the 107-kDa subunit. In
order to clarify the molecular basis for this observation, a 440kDa Class-1 PEPC heterotetramer composed of an equivalent
ratio of non-phosphorylated 110- and 107-kDa subunits was
purified from 3-day-old germinated COS endosperm [103]. Nterminal microsequencing, MS and immunoblotting established
that both subunits arise from the same PTPC gene (RcPpc3)
(Figure 3) encoding the phosphorylated 410-kDa Class-1
PEPC homotetramer of developing COS, but that the 110-kDa
subunit is a mono-ubiquitinated form of the 107-kDa subunit.
Tandem MS sequencing of tryptic peptides identified Lys628
as PEPC’s mono-ubiquitination site [103]. Lys628 is absolutely
conserved in all PTPCs and BTPCs, and is proximal to a PEPbinding/catalytic domain (Figure 3). Incubation of the purified
germinated COS Class-1 PEPC with a recombinant human
deubiquitinating enzyme [USP-2 (ubiquitin-specific protease-2)
core] cleaved ubiquitin from its 110-kDa subunits (Figure 4) while
significantly reducing the enzyme’s K m (PEP) and sensitivity to
allosteric activators (hexose phosphates, glycerol 3-phosphate)
and inhibitors (malate, aspartate) [103]. It is also notable that
elimination of photosynthate supply to developing COS by
detaching intact developing COS pods from the plant caused
the 107-kDa PTPC subunits of the endosperm and cotyledon
to become dephosphorylated [34,35], and then subsequently
mono-ubiquitinated in vivo (B. O’Leary and W. C. Plaxton,
unpublished work), concomitant with the disappearance of
PPCK activity. Because depodding abolishes photosynthate
delivery to COS, it is conceivable that the metabolism of
the depodded developing endosperm is rearranged to mimic
that of the gluconeogenic germinating COS. PTPC monoubiquitination has since been demonstrated in Lilium longiflorum
(lily) pollen [7] and Arabidopsis seedlings (B. O’Leary and
W. C. Plaxton, unpublished work). An immunoreactive PTPC
‘doublet’ highly reminiscent of the mono-ubiquitinated Class-1
PEPC of germinated COS has been frequently observed on PTPC
immunoblots of clarified extracts from a broad variety of plants,
including Hydrilla verticillata (hydrilla) leaves [186], Vicia
faba (broad bean) stomata [187], Cucumis sativus (cucumber)
roots [188], germinating seeds of wheat, barley and sorghum
[104,105,107], and ripening banana fruit [119]. In contrast with
the COS system, the Class-1 PEPC of cereal seeds such as
barley and wheat is phosphorylated during germination, whereas
that of fully mature maize seeds appears to be maximally
phosphorylated before germination [36,106,168]. We recently
completed a survey of tissue-specific PTMs of PTPC polypeptides
in various photosynthetic and non-photosynthetic tissues of the
castor oil plant. Immunoblotting using PTPC (anti-RcPPC3)
and phospho-site-specific PTPC (anti-pS11) antibodies following
pre-incubation of clarified extracts with USP-2 demonstrated
that PTPC mono-ubiquitination rather than phosphorylation
appears to be the predominant PTM of Class-1 PEPCs that
occurs in non-stressed castor oil plants (B. O’Leary and
W. C. Plaxton, unpublished work). Moreover, the distinctive
developmental patterns of PTPC phosphorylation compared with
mono-ubiquitination in the castor oil plant indicated that these
PTMs may be mutually exclusive. As discussed above, Class-1
c The Authors Journal compilation c 2011 Biochemical Society
Figure 4 A human deubiquitinating enzyme (USP-2 core) cleaved
ubiquitin from the 110-kDa subunits of the germinating castor bean PEPC
heterotetramer
Native PEPC purified from endosperm of 4-day-old germinated COS endosperm was incubated
with or without 1 μM USP-2 core. Aliquots were removed at various times and subjected
to immunoblot analysis using (A) anti-(bovine ubiquitin IgG) (1 μg of PEPC/lane), or (B)
anti-[developing COS Class-1 PEPC (RcPPC3) IgG] (0.1 μg of PEPC/lane). Figure modified
from [103] with permission. Molecular masses are indicated in kDa.
PEPC phosphorylation activation was mainly observed in plant
tissues in which a high and tightly controlled flux of PEP to
malate has an obvious metabolic role; e.g. as occurs during
atmospheric CO2 assimilation in C4 or CAM leaves, photosynthate
partitioning to storage end-products by developing COS, rapid
nitrogen assimilation following NH4 + or NO3 − resupply
to nitrogen-limited cells, or during nutritional Pi starvation
[16,17,34,38,42,54,169]. Clearly, additional research is warranted
to assess the interplay between and metabolic functions of
PTPC phosphorylation and mono-ubiquitination. Future research
also needs to characterize the: (i) ubiquitin-binding domain
proteins that might interact with the ubiquitin-‘docking site’ of
mono-ubiquitinated PEPCs, as well as the possible influence
of this PTM on their subcellular location, and (ii) signalling
pathways and specific E3 ligase that mediate tissue-specific
PTPC mono-ubiquitination. High-throughput proteomic screens
have identified numerous ubiquitinated metabolic enzymes
in Arabidopsis, including PTPC [189,190]. However, it was
not determined whether the various targets were poly- or
mono-ubiquitinated. This discrimination is crucial for future
studies of the plant ubiquitome since polyubiquitination and
mono-ubiquitination are destructive and non-destructive PTMs
respectively that mediate entirely different effects on target protein
function [183,184].
BTPC functions as a catalytic and regulatory subunit of Class-2
PEPC complexes of vascular plants and green algae
BTPC genes are unique and specifically expressed
In 2003, interrogation of Arabidopsis and rice PEPC gene families
led to the surprising discovery that, alongside their various PTPC
genes, both genomes also encode and express an enigmatic BTPC
gene whose deduced amino acid sequence shares a slightly higher
similarity with PEPCs from proteobacteria than with other plant
PEPCs (Figure 5) [3]. These novel PEPC isoenzymes were
thus named BTPCs, whereas all remaining C3 , C4 and CAM
PEPCs were classified as PTPCs. All plant genomes sequenced
to date, including that of ancestral green algae, contain at least
one BTPC gene. The BTPCs constitute a monophyletic group,
separate from either the PTPCs or bacterial and archaeal PEPCs,
and appear to have evolved in green algae (Figure 5). PTPC
genes have a highly conserved genomic structure composed of
approximately ten exons, whereas BTPC genes have a very
different and more complex structure with approximately 20
exons [8]. Deduced BTPC polypeptides of vascular plants range
from ∼116 to 118 kDa (or approximately 130 kDa in green algae)
Functional control of non-photosynthetic plant PEPC
Figure 5
25
Phylogenetic analysis of vascular plant, green algal, bacterial and archaeal PEPCs using a neighbour-joining consensus tree
Protein accession numbers from GenBank® , Phytozome and Maizesequence databases were used to construct the phylogenetic tree and are shown in square brackets. Bootstrap analysis was carried
out with 100 replicates. Numbers at the branches correspond to percentage bootstrap frequencies for each branch. Only values >50 are shown.
c The Authors Journal compilation c 2011 Biochemical Society
26
B. O’Leary, J. Park and W. C. Plaxton
compared with ∼105–110 kDa for the corresponding PTPCs [8].
All BTPC sequences deduced contain residues critical for PEPC
catalysis (Figure 3), and heterologous expression of green algal
(C. reinhardtii) and castor oil plant BTPCs yielded active PEPCs
[5,160,191]. Deduced PEPC polypeptides are readily classified
as a BTPC or PTPC by three main criteria: (i) their C-terminal
tetrapeptide is either (R/K)NTG for BTPCs or QNTG for PTPCs,
(ii) BTPCs lack the distinctive N-terminal serine phosphorylation
motif (acid-base-XXSIDAQLR) characteristic of PTPCs [3–5],
and (iii) all BTPCs contain a unique and highly divergent insertion
of approximately 10 kDa (corresponding to residues 325–467
of the COS BTPC, RcPPC4) that was predicted to exist in a
largely unstructured and highly flexible conformation, known as
an intrinsically disordered region (Figure 3) [191]. Disorderedregion-containing proteins are ubiquitous in all organisms. Their
disordered region typically exists as a flexible linker that connects
two globular domains to mediate protein–protein interactions.
The flexible linker allows the connected domains to freely twist
and rotate through space to recruit their binding partners. Linker
sequences can vary greatly in length and amino acid sequence,
but tend to be susceptible to proteolytic cleavage while exhibiting
a low content of bulky hydrophobic amino acids and a higher
proportion of polar and electrically charged amino acids [192].
Indeed, a recent imaging study of transiently expressed wildtype and truncated mutants of castor oil plant BTPC (RcPPC4)
and PTPC (RcPPC3) fluorescent fusion proteins in tobacco BY2
suspension cells demonstrated that RcPPC4’s disordered region
mediates its in vivo interaction with RcPPC3 (e.g. as a Class-2
PEPC complex, see below) [193].
The tissue-specific expression of BTPC transcripts has been
investigated in several species. Although expression of BTPC
transcripts tends to be quite low relative to PTPC transcripts, and
is dependent upon the developmental stage or metabolic status
of the tissue being examined, no obvious pattern of expression
has emerged [3,4,6,7,11,194]. Arabidopsis BTPC transcripts
were initially detected at low levels in siliques and flowers
[3], but were shown subsequently to be specifically expressed
in the late stages of pollen development [7] and induced in
roots by salt and drought stress [11]. BTPC transcripts and
polypeptides are co-ordinately and highly expressed during COS
development; their developmental profile coincided with stages of
rapid endosperm growth and oil accumulation [4]. Nevertheless,
our current understanding of plant/algal BTPC biochemistry
was initially built upon a foundation of integrating classical
enzyme biochemistry with modern tools of MS and associated
bioinformatic databases.
Native BTPCs interact with PTPCs as subunits of heteromeric Class-2 PEPC
complexes
The study of plant BTPC biochemistry began unknowingly in
the 1990s with a series of papers describing the purification and
characterization of two distinct PEPC classes from unicellular
green algae (Selenastrum minutum and C. reinhardtii) that
exhibited highly dissimilar physical and kinetic properties, but
that shared an identical PTPC subunit [195–199]. From these
preparations, typical 400-kDa PEPC homotetramers composed
solely of PTPC subunits were classified as a Class-1 PEPC,
whereas high-molecular-mass complexes of PTPC and an
immunologically unrelated 130-kDa subunit (subsequently shown
to be a BTPC) were termed Class-2 PEPCs. Relative to the
Class-1 PEPC, the algal Class-2 PEPCs displayed enhanced
thermal stability, a broader pH-activity profile, biphasic PEP
saturation kinetics and a markedly reduced sensitivity to allosteric
c The Authors Journal compilation c 2011 Biochemical Society
effectors. This work was corroborated and significantly extended
by Raymond Chollet and co-workers who cloned and analysed
the expression of C. reinhardtii PTPC and BTPC genes [5,67].
The PTPC cDNA expressed as an active ∼110-kDa subunit
of both Class-1 PEPC and Class-2 PEPC, whereas the BTPC
cDNA expressed as an active ∼130-kDa subunit found only
in Class-2 PEPC. At the level of gene and protein expression,
both algal PEPC subunits responded to inorganic nitrogen levels.
However, the effect was more pronounced with the PTPC
subunits, consistent with their proposed anaplerotic function
during transient nitrogen assimilation [5,67].
Low- and high-molecular-mass PEPC isoforms were
subsequently purified and characterized from the triacylglycerolrich endosperm of developing COS whose respective physical
and kinetic/regulatory properties were remarkably analogous to
the highly distinctive Class-1 and Class-2 PEPCs of unicellular
green algae [82]. Thus native COS Class-1 PEPC is a classic
410-kDa homotetramer of 107-kDa PTPC subunits. In contrast,
the allosterically desensitized COS Class-2 PEPC 910-kDa
hetero-octameric complex consists of the same Class-1 PEPC
homotetrameric core tightly associated with four 118-kDa BTPC
subunits (Figures 6 and 7) [4,82]. The COS Class-2 PEPC
complex has been documented by additional in vitro techniques
including co-IP (co-immunopurification) and non-denaturing
PAGE of clarified extracts coupled with in-gel PEPC activity
staining and parallel immunoblotting using BTPC- and PTPCspecific antibodies [4,34,77]. Both approaches were recently
employed to demonstrate comparable Class-1 and Class-2 PEPC
isoforms in extracts of developing lily pollen [7].
Upon extraction, the BTPC subunits of green algal and vascular
plant Class-2 PEPCs are extremely susceptible to rapid in vitro
proteolytic cleavage at a specific site within their disordered
region (e.g. see Figure 3) by an endogenous thiol endopeptidase
[4,196,197]. This in vitro proteolysis has prevented purification
of native COS Class-2 PEPC containing non-truncated BTPC
subunits, even in the presence of a wide array of protease
inhibitors and cocktails. However, limited protection, suitable
for BTPC immunoblotting and co-IP from clarified extracts,
was afforded by a combination of PMSF and the ProteCEASE
100 Cocktail marketed by G-Biosciences [4,77]. Two additional
techniques were central in allowing further characterization of
non-proteolytically degraded native BTPC from developing COS.
First, co-IP of highly enriched native BTPC from developing
COS endosperm extracts was achieved using an anti-(castor oil
plant PTPC IgG) immunoaffinity column [77]. Relatively large
quantities (>5 mg) of purified non-proteolysed 118-kDa BTPC
subunits were eluted from this column using Pierce’s Gentle
Ag/Ab Elution Buffer and used for detailed MS characterization
of its in vivo phosphorylation sites [77,191]. Secondly, E. coli
lysates containing heterologously expressed wild-type and mutant
versions of castor oil plant BTPC (RcPPC4) were mixed with a
recombinant Arabidopsis PTPC (AtPPC3) to create intact stable
chimaeric Class-2 PEPCs in vitro, which were then purified and
characterized [160,191]. Both PTPC and BTPC subunits were
catalytically active and consequently the chimaeric Class-2 PEPC
displayed biphasic PEP saturation kinetics [160], as documented
previously for the purified non-proteolysed native Class-2 PEPC
from the green alga S. minutum [197]. BTPC is a low-affinity
allosterically desensitized Class-2 PEPC subunit that exhibits K m
(PEP) and IC50 (malate) values approximately 10- and 15-fold
higher respectively than those of the PTPC subunits [160,191].
The BTPC subunits appear to have an additional regulatory role
because the PTPC subunits of Class-2 PEPC were far less sensitive
to allosteric inhibitors compared with the same subunits within a
Class-1 PEPC (Figure 7) [160,191]. It is also notable that the
Functional control of non-photosynthetic plant PEPC
Figure 6
27
Model illustrating the biochemical complexity of castor bean PEPC
In developing COS endosperm, the PTPC RcPPC3 exists: (i) as a typical Class-1 PEPC homotetramer (PEPC1) which is activated in vivo by sucrose-dependent phosphorylation of its 107-kDa
subunit (p107) at Ser11 [34,35], and (ii) tightly associated with 118-kDa BTPC (RcPPC4) subunits (p118) to form the allosterically desensitized Class-2 PEPC hetero-octameric complex (PEPC2).
The RcPPC4 subunits are subject to in vivo multi-site phosphorylation at Thr4 , Ser425 and Ser451 [77,191,201]. COS maturation is accompanied by disappearance of the Class-2 PEPC complex and
RcPPC4 polypeptides and transcripts, a marked reduction in the amount of Class-1 PEPC coupled with dephosphorylation of its p107-PTPC subunits [4,34,82]. COS imbibition and germination
is accompanied by increased RcPpc3 gene expression, PEPC activity and amount, and ubiquitination of 50 % of RcPPC3’s subunits at Lys628 to form the mono-ubiquitinated Class-1 PEPC
heterotetramer. UB, ubiquitin. Figure modified from [103] with permission.
Figure 7 Differential influence of glucose 6-phosphate and malate on the
activity of native Class-1 and Class-2 PEPC (PEPC1 and PEPC2 respectively)
isoforms purified from developing castor beans
A 0.5 and IC50 denote the concentration of glucose 6-phosphate (Glc-6-P/G6P) and malate
required for half maximal activation and inhibition respectively. Assays were conducted at pH 7.3
with subsaturating PEP concentrations (0.2 mM). All values represent the means ( +
− S.E.M.) of
three separate determinations. Figure modified from [82] with permission.
recombinant COS BTPC readily formed insoluble aggregates
when extracted from E. coli cells in which it was overexpressed.
However, when mixed with native or recombinant Class-1 PEPCs
from various plant sources, the recombinant COS BTPC subunits
spontaneously rearranged to form stable soluble Class-2 PEPC
complexes [160]. Although plant and algal BTPCs exhibit PEPC
activity, several additional lines of evidence strongly suggest that
PTPC subunits are essential binding partners for BTPC subunits:
(i) native BTPC subunits have only been observed in association
with PTPC subunits as a Class-2 PEPC [4,67,82,196,197], (ii)
the BTPC subunits tightly associate with the PTPC subunits
during purification [77,82], and (iii) FP (fluorescent protein)tagged castor oil plant PTPC and BTPC subunits interact in
vivo during their transient co-expression in tobacco BY2 cells
(see below) [193]. PTPCs have thus been hypothesized to be
compulsory BTPC-binding partners that play an indispensable
role in maintaining BTPCs in their proper structural and functional
state in Class-2 PEPCs [160].
Significant levels of Class-2 PEPC have been observed in
developing COS endosperm and lily pollen. Both are reproductive
sink tissues in which mitosis has been completed and the
cells are rapidly expanding while simultaneously metabolizing
large amounts of imported photosynthate into storage lipids and
proteins [7,82,160]. The unique kinetic and regulatory properties
of Class-2 PEPC have been hypothesized to function as a
‘metabolic overflow’ mechanism capable of sustaining significant
flux from PEP to malate under in vivo conditions where the
corresponding Class-1 PEPC activity would become largely
suppressed by feedback allosteric inhibitors [7,160]. For example,
malate levels in developing COS endosperm have been measured
at 5 mM, a value ∼80-fold higher than the IC50 (malate) of COS
Class-1 PEPC at physiological pH (Figure 7), but within the range
of the IC50 (malate) of the BTPC subunit of Class-2 PEPC [82,89].
BTPC of developing castor beans is subject to multi-site
phosphorylation in vivo
As plant BTPCs lack the conserved N-terminal serine phosphorylation motif characteristic of PTPCs (Figure 3), they were
suggested to be non-phosphorylatable [11,37]. However, the use
of Pro-Q Diamond phosphoprotein staining, immunoblotting with
commercially available phospho-(serine/threonine) Akt substratespecific antibodies, and Pi -affinity PAGE using Phos-TAG
acrylamide [200] demonstrated that co-immunopurified native
c The Authors Journal compilation c 2011 Biochemical Society
28
B. O’Leary, J. Park and W. C. Plaxton
BTPC from developing COS endosperm was phosphorylated
at multiple sites in vivo [77]. Detailed analyses of coimmunopurified COS BTPC using Fourier-transform MS (>93 %
sequence coverage) identified three novel phosphorylation sites:
Thr4 at the N-terminus, and Ser425 and Ser451 within its
disordered region (respectively corresponding to acidophilic,
proline-directed and basophilic protein kinase recognition motifs)
(Figure 3) [77,201]. The Thr4 and Ser451 phosphorylation sites
are conserved in other BTPC orthologues, but the Ser425 site
is only partially conserved (e.g. see Figure 3) [191,201].
Phosphomimetic mutagensis of the Ser425 and Ser451 sites have
shown them to be regulatory in nature, with both causing marked
inhibition of the BTPC subunits within a Class-2 PEPC by
increasing their K m (PEP) and sensitivity to feedback inhibition
by malate and aspartate [191,201]. The developmental patterns
of BTPC phosphorylation at Ser425 and Ser451 were examined
by immunoblotting clarified COS extracts with the respective
phosphosite-specific antibodies, and shown to be very distinct
from the in vivo phosphorylation activation of COS Class-1
PEPC’s PTPC subunits at Ser11 , implying control by separate
protein kinases and signalling pathways [191,201]. In green
algae, BTPC phosphorylation: (i) also appears to be inhibitory
in nature, and (ii) may be involved in mediating BTPC/PTPC
subunit stoichiometry within algal Class-2 PEPC complexes
[198]. The function of the N-terminal phosphorylation site of
developing COS BTPC (Thr4 ) is unknown. Kinetic analysis of
a T4D phosphomimetic RcPPC4 mutant indicated that it is not
regulatory in nature [201]. However, it is intriguing that Thr4 of
plant BTPC exists in a conserved FHA (forkhead-associated)binding domain (pTXXD) (Figure 3) and also corresponds to an
acidophilic protein kinase CK2 motif. As FHA domains have
gained considerable prominence as phosphothreonine-dependent
protein interaction modules [202], it will be necessary to establish
the role that BTPC phosphorylation at Thr4 might play in
mediating the interaction of Class-2 PEPCs with FHA domaincontaining proteins. Characterization of BTPC phosphorylation
from additional vascular plant and algal sources, alongside
identification of the responsible protein kinases and related
signalling pathways, will also be essential to further validate and
extend the role of BTPC phosphorylation in Class-2 PEPCs.
SUBCELLULAR LOCALIZATION AND PROTEIN–PROTEIN
INTERACTIONS OF PLANT PEPC
Class-2 PEPC appears to associate with the mitochondrial outer
envelope
The subcellular location and in vivo interaction of COS PTPC
(RcPPC3) and BTPC (RcPPC4) were assessed recently by
imaging FP–PEPC fusion proteins that had been transiently
expressed in heterotrophic tobacco suspension cells [193]. FP–
PTPC sorted to the diffuse cytosol, whereas NLS (nuclear
localization signal)–FP–PTPC sorted to the nucleus [193]. In
contrast, BTPC-FP localized to discrete punctate structures,
tentatively identified as mitochondria by immunostaining of
endogenous mitochondrial cytochrome oxidase. However, when
NLS–FP–PTPC or FP–PTPC was co-expressed with BTPC–
FP, fluorescent PTPC signals co-localized with BTPC–FP.
Transmission electron microscopy of immunogold-labelled
developing COS endosperm and cotyledon using monospecific
COS BTPC and PTPC antibodies indicated that both proteins
tended to cluster together into discrete regions of the
mitochondrial outer envelope. The overall results indicate that: (i)
COS BTPC and PTPC interact in vivo as a Class-2 PEPC complex,
c The Authors Journal compilation c 2011 Biochemical Society
(ii) BTPC’s unique and divergent intrinsically disordered region
mediates its tight interaction with PTPC, (iii) the BTPCcontaining Class-2 PEPC may be located on the mitochondrial
outer membrane, whereas (iv) the PTPC-containing Class-1 PEPC
is uniformly distributed throughout the cytosol. Mitochondrialassociated cytosolic glycolytic isoenzymes have also been
reported in several studies. Lee Sweetlove and colleagues showed
that seven different glycolytic enzymes formed a metabolon
on the mitochondrial surface of Arabidopsis suspension cells
during periods of increased respiration so as to channel carbon
from cytosolic metabolite pools into the mitochondria while
restricting substrate use by competing metabolic pathways
[203,204]. Furthermore, FP-tagged aldolase, enolase, ATPPFK and hexokinase have shown a dual mitochondrial (outer
envelope) and cytosolic localization [203]. We therefore need to
determine the prevalence, mechanism and metabolic role(s) of
mitochondrial-associated Class-2 PEPC complexes. In addition,
it will be important to identify any Class-2 PEPC-interacting
proteins that may form a metabolon to facilitate respiratory
CO2 refixation (e.g. carbonic anhydrase) and/or anaplerotic PEP
partitioning to metabolic end-products such as storage lipids and
proteins.
Class-1 PEPC may interact with a cytosolic-targeted PDH
Biochemical analyses of several purified plant PTPC isoenzymes
indicated that they form a complex with the metabolically
sequential MDH and ME in C4 leaves [9], its own PPCK in
ripening banana fruit [120], as well as the BTPC leading to
the formation of Class-2 PEPC hetero-oligomeric complexes of
green algae and vascular plants [4,5,67,82,197]. Multienzyme
complexes containing PEPC, MDH and acetyl-CoA carboxylase
were also isolated from the unicellular photosynthetic protist
Euglena gracilis [205]. A surprising observation of the recent
co-IP proteomics study of the Class-1 PEPC interactome of
developing COS endosperm was the identification of the PDHp
(plastidial PDH) as a putative PTPC interactor [77]. PDH catalyses
the irreversible oxidative decarboxylation of pyruvate into acetylCoA (Figure 2). Immunoblotting using monospecific antibodies
raised against E1α, E1β, E2 and E3 subunits of Arabidopsis
PDHp verified the presence of all four PDHp subunits in eluates
from the anti-[COS Class-1 PEPC (RcPPC3)] immunoaffinity
column [77]. PDHp appears to specifically associate with the
PTPC subunits of COS Class-1 PEPC, since a parallel co-IP study
of the PEPC interactome of germinating COS endosperm (which
contains no detectable BTPC or Class-2 PEPC) also identified
PDHp as a PTPC-interacting protein [193]. Although COS Class-1
PEPC and PDHp are believed to be localized in different metabolic
compartments, there are only two enzymatic steps between them
(MDH and ME), and PEPC and PDHp have both been implicated
in playing an important role in supporting fatty acid synthesis in
developing COS [81,89,206]. Plant cells are unique in containing
two different PDH isoenzymes: mitochondrial PDH which links
cytosolic glycolysis with the TCA cycle, and PDHp which
produces acetyl-CoA and NADH for plastidic fatty acid synthesis
(Figure 2) [206,207]. The structure and post-translational control
of PDHp is very different from that of mitochondrial PDH, and
both isoenzymes can be readily discriminated by immunoblotting.
Immunoreactive PDHp subunits were markedly enriched when
clarified COS extracts were subjected to co-IP on the anti(COS Class-1 PEPC) immunoaffinity column. In contrast, no
immunoreactive bands were evident when parallel immunoblots
of the same co-IP column eluates were probed with anti-(plant
mitochondrial PDH) antibodies [77]. A previous report indicated
that PDHp may not be exclusively plastidic in developing COS
Functional control of non-photosynthetic plant PEPC
endosperm [206]. In this study, the plastid marker enzyme acetylCoA carboxylase demonstrated 98 % of its total activity to be
leucoplast-localized, whereas only 62 % of total PDH activity was
in the same fraction. Conversely, 2 % and 38 % of total acetylCoA carboxylase and PDH activity were respectively measured
in the corresponding cytosolic fraction [206]. Further studies are
required to establish PDHp ’s distribution in the plastid compared
with cytosol of non-green plant cells, such as developing and
germinating COS endosperm. Nevertheless, the aforementioned
findings support the hypothesis that a specific PEPC–PDHp
interaction might exist in developing COS. This interaction could
facilitate CO2 recycling from PDH to PEPC and/or occur as part of
a metabolon that channels PEP to cytosolic acetyl-CoA required
for the biosynthesis of isoprenoids, flavonoids and malonated
derivatives, in addition to the elongation of C16 and C18 fatty acids.
Cytosolic acetyl-CoA is believed to be generated from citrate
and CoA by ATP-citrate lyase [208]. However, ATP-citrate lyase
activity was barely detectable in developing COS extracts [208].
PDHp could provide an alternative metabolic route for acetylCoA production within the COS cytosol. It will be interesting
to determine whether the observed in vitro interaction between
a Class-1 PEPC and PDHp exists in vivo, and, if so, the role
that it plays in carbohydrate partitioning and CO2 recycling in
developing seeds.
The chloroplast Class-1 PEPC isoenzyme of rice leaves
Until recently, PEPC had been described as being exclusively
cytosolic, so it was a major surprise when a chloroplastic
PTPC isoenzyme was identified in rice leaves [159]. This PTPC
isoenzyme contains a transit peptide and its expression was
largely confined to leaf mesophyll cells where it was estimated to
account for a third of the total PEPC protein. It exhibited a lower
affinity for both PEP and allosteric effectors compared with
typical C3 PTPC isoenzymes, and gene knockouts produced
a phenotype which implied its involvement in leaf nitrogen
assimilation [159]. Earlier immunogold imaging studies reported
that PTPC may associate with the chloroplast outer envelope
in C3 , C4 and CAM leaves [209,210], whereas PTPC partially
pelleted with the total membrane fraction from sorghum leaves
[167]. However, this is the first well-documented instance of a
plant PEPC being targeted to the plastid stroma. It is unknown
whether plastidic PEPC isoenzymes exist in other genera, as
orthologous genes were not detected. Nevertheless, the existence
of a chloroplastic PTPC isoenzyme endows rice mesophyll cells
with an alternative route of organic acid metabolism to support
nitrogen assimilation that had not previously been considered in
plant primary metabolism [159].
CONCLUDING REMARKS
Although the central importance of PEPC in plant metabolism and
physiology has been appreciated for many years, recent research
has provided numerous insights into the complex biochemical
and molecular mechanisms that underpin the control of PEPC
activity, and the influence that PEPC and PPCK exert on a wide
array of cellular processes. Integrative analyses at the genomic,
transcriptomic, enzymological/biochemical and cellular levels
have revealed PEPC’s biochemical complexity (e.g. see Figure 6),
whereas the application of metabolic flux analysis and functional
genomic tools have consistently indicated the importance of posttranslational PEPC control in ensuring the optimal regulation and
plasticity of anaplerotic PEP metabolism. Owing to its multiple
functions and location at a pivotal branchpoint of plant primary
29
metabolism (Figures 1 and 2A), an impressive array of strategies
has evolved to post-translationally control the activity of plant
PEPCs. Regulatory phosphorylation and mono-ubiquitination,
14-3-3 proteins, as well as changes in intracellular pH and
allosteric effector levels have all been described as mechanisms
to control Class-1 PEPC activity in different plant tissues under
various physiological conditions. The discovery of allosterically
desensitized Class-2 PEPC hetero-octameric complexes in green
algae and vascular plants that arise from a tight physical
association between a Class-1 PEPC with co-expressed BTPC
subunits adds another layer of complexity to plant PEPC functions
and control. A major challenge will be to link the multifaceted
post-translational controls and associated PTMs of Class-1 and
Class-2 PEPCs with the cell-specific partitioning of PEP into
discrete metabolic pathways.
Many metabolic functions have been attributed to malate
and other TCA cycle intermediates within different cell types
[28], and the versatility of these metabolites is undoubtedly
linked to the evolved diversity of plant PEPC (Figure 1).
A comprehensive understanding of the linkages between the
expression of individual PEPC isoenzymes, their specific
physiological/metabolic functions, and the synchronous regulation of PEPC with other key enzymes of the PEP branchpoint
(e.g. PKc , PEP carboxykinase, pyruvate orthophosphate dikinase,
the shikimate pathway) represents a major challenge. Only with
the concerted efforts of plant physiologists, biochemists and
enzymologists and molecular biologists, will the cell-specific
functions and control of plant PEPC be fully comprehended.
It is anticipated that the further characterization of green algal
and vascular plant Class-1 and Class-2 PEPC, their in vivo
PTMs (including the requisite interconverting enzymes such as
PPCK or E3 ubiquitin ligases, and relevant signalling pathways),
subcellular localization and protein–protein interactions will
remain a fruitful research area for the foreseeable future. It
will also be important to establish a structural model for plant
and algal Class-2 PEPC subunit architecture. Lastly, patterns
of Class-1 compared with Class-2 PEPC expression need to
be re-evaluated together with the continued application of
functional genomic tools to assess the impact of altered PTPC and
BTPC expression on anaplerotic PEP-partitioning to storage endproducts in developing seeds, as well as plant carbon–nitrogen
interactions and acclimatization to environmental stress.
ACKNOWLEDGEMENTS
W.C.P. is greatly indebted to past and present members of his laboratory who made
interesting discoveries concerning the molecular and biochemical characteristics of nonphotosynthetic plant PEPCs. W.C.P. is also very grateful to collaborators who have
contributed to our PEPC research, particularly Dr David Turpin, Dr Florencio Podestá,
Dr Jean Rivoal, Dr Wayne Snedden, Dr Chao-fu Lu, Dr Yi-min She, Dr Craig Leach,
Dr Alexander Valentine and Dr Robert Mullen.
FUNDING
Research in this laboratory is generously supported by grants from the Natural Sciences
and Engineering Research Council of Canada (NSERC) and the Queen’s Research Chair’s
program (to W.C.P.), an NSERC post-graduate scholarship (to B.O.’L.) and a Queen’s
University-Province of Ontario postdoctoral fellowship (to J.P.).
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