Plant Physiology and Biochemistry 58 (2012) 89e97
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Plant Physiology and Biochemistry
journal homepage: www.elsevier.com/locate/plaphy
Research article
Comprehensive survey of redox sensitive starch metabolising enzymes
in Arabidopsis thaliana
Mikkel A. Glaring a, b, c, Katsiaryna Skryhan a, Oliver Kötting c, Samuel C. Zeeman c, Andreas Blennow a, *
a
VKR Research Centre Pro-Active Plants, Department of Plant Biology and Biotechnology, University of Copenhagen, 40 Thorvaldsensvej, 1871 Frederiksberg C, Denmark
Department of Agriculture and Ecology, University of Copenhagen, 40 Thorvaldsensvej, 1871 Frederiksberg C, Denmark
c
Department of Biology, ETH Zürich, Universitätstrasse 2, 8092 Zürich, Switzerland
b
a r t i c l e i n f o
a b s t r a c t
Article history:
Received 4 April 2012
Accepted 19 June 2012
Available online 28 June 2012
In chloroplasts, the ferredoxin/thioredoxin pathway regulates enzyme activity in response to light by
reduction of regulatory disulfides in target enzymes, ensuring coordination between photosynthesis and
diurnal metabolism. Although earlier studies have suggested that many starch metabolic enzymes are
similarly regulated, redox regulation has only been verified for a few of these in vitro. Using zymograms
and enzyme assays, we performed a comprehensive analysis of the redox sensitivity of known starch
metabolising enzymes in extracts of Arabidopsis thaliana. Manipulation of redox potentials revealed that
several enzymatic activities where activated by reduction at physiologically relevant potentials. Among
these where the isoamylase complex AtISA1/AtISA2, the limit dextrinase AtLDA, starch synthases AtSS1
and AtSS3, and the starch branching enzyme AtBE2. The reversibility of the redox reaction was confirmed
by enzyme assays for AtLDA, AtSS1 and AtSS3. Analysis of an AtBAM1 knock-out mutant identified an
additional redox sensitive b-amylase activity, which was most likely AtBAM3. A similar requirement for
reducing conditions was observed for recombinant chloroplastic a-amylase (AtAMY3) activity. This study
adds further candidates to the list of reductively activated starch metabolising enzymes and supports the
view that redox regulation plays a role in starch metabolism.
Ó 2012 Elsevier Masson SAS. All rights reserved.
Keywords:
Arabidopsis
Starch
Redox regulation
Starch metabolism
Starch synthase
1. Introduction
Starch is a major product of photosynthetic carbon fixation and
serves important functions as a storage carbohydrate. In the chloroplast of leaves, starch accumulates during the day and is
degraded during the night to provide energy for continued growth
in the dark. This repeated cycle of biosynthesis and degradation
requires regulation of key metabolic pathways in order to balance
the available photosynthate with the requirements for growth and
storage [1e3]. Carbon assimilation is limited by the light available
for photosynthesis and light itself acts as a regulatory signal that
allows the plant to switch between light and dark metabolism. This
occurs through the chloroplastic ferredoxin/thioredoxin system,
Abbreviations: AGPase, ADP-glucose pyrophosphorylase; AMY, a-amylase; At,
Arabidopsis thaliana; BAM, b-amylase; BE, starch branching enzyme; DTT, dithiothreitol; DTTox (oxidised DTT), trans-4,5-dihydroxy-1,2-dithiane; Fdx, Ferredoxin;
GWD, a-glucan, water dikinase; ISA, isoamylase; LDA, limit dextrinase; MDH,
NADP-dependent malate dehydrogenase; NTRC, NADP-thioredoxin reductase C;
PHS, starch phosphorylase; SS, starch synthase; Trx, thioredoxin.
* Corresponding author. Tel.: þ45 35333304.
E-mail address: [email protected] (A. Blennow).
0981-9428/$ e see front matter Ó 2012 Elsevier Masson SAS. All rights reserved.
http://dx.doi.org/10.1016/j.plaphy.2012.06.017
which converts a light-activated electron signal into a thiol signal.
Electrons transported through the photosystems during photosynthesis reduce ferredoxin (Fdx), which, in turn, leads to the
sequential reduction of ferredoxin-thioredoxin reductase (FTR) and
a family of thioredoxins (Trx). The Trxs then directly reduce their
target enzymes by catalysing a disulfide to thiol conversion, thus
linking the availability of light to the activity of numerous enzymes
[4]. The elucidation of this process for widespread reversible
posttranslational modification has led to the general view that
redox sensitive biosynthetic enzymes in chloroplasts are mainly
active during the day and inhibited at night. A complete Fdx/Trx
system has also been identified in amyloplasts from wheat endosperm. In this tissue, Fdx is reduced not by light, but by metabolically generated NADPH via ferredoxin-NADP reductase [5].
The building blocks for starch biosynthesis, ADP-glucose, is
produced from glucose 1-phosphate and ATP by the enzyme ADPglucose pyrophosphorylase (AGPase). In the light, glucose 1phosphate is synthesised from Calvin cycle intermediates by
phosphoglucose isomerase and phosphoglucomutase. The activity
of AGPase in leaves is regulated by both the available light and
sugars through the breaking and formation of a disulfide bridge
between the two small subunits of the AGPase heterotetrameric
M.A. Glaring et al. / Plant Physiology and Biochemistry 58 (2012) 89e97
Redu c e d
A
O x i di z ed
ISA1/2
BAM1
LDA
03
17
-3
33
26
-3
-3
38
-3
-3
42
48
-3
55
-3
66
-3
-3
B
-3
80
DPE1
BAM5
mV
SS3
BAM1
AMY3
26
-3
21
-3
17
-3
12
-3
03
-3
30
-3
33
-3
38
-3
42
-3
48
-3
-3
55
SS1
80
enzyme complex [6,7]. This is similar to the light-dependent redox
activation of enzymes of the Calvin cycle and both processes are
activated by reduced Trx, ensuring coordination between photosynthesis and starch biosynthesis in response to light [4]. Recently,
reductive activation of AGPase by the bifunctional NADPthioredoxin reductase C (NTRC) was discovered as an alternative
mechanism of redox regulation in Arabidopsis. NTRC uses NADPH
to reduce AGPase and is linked to photoreduced Fdx by the
ferredoxin-NADP reductase, thus complementing the classical Fdx/
Trx system [8].
Potential targets of redox regulation by Trxs have previously
been identified by a variety of methods. Initially, many of these
were identified based on in vitro changes in activity observed after
reduction using non-physiological substitutes for Trx, such as
dithiothreitol (DTT) [4,9]. Over the last decade, both proteomic and
in vitro approaches have greatly expanded the number of targets
and among these are several enzymes involved in starch metabolism [9,10]. Apart from AGPase, these include potato a-glucan,
water dikinase (GWD) and Arabidopsis b-amylase 1 (AtBAM1) and
phosphoglucan phosphatase SEX4, all of which are activated by
Trxs in vitro [11e13]. Similarly, a poplar leaf endoamylase was
shown to be reversibly inactivated by oxidation and activated by
DTT and E. coli Trx [14], although it is not clear whether this enzyme
is localised in the chloroplasts. The activity and isoform pattern of
limit dextrinase, also called pullulanase, from wheat and spinach is
influenced by the redox conditions in vitro [15,16], however, activation by Trx has not been demonstrated. Another two enzymes, a1,4-glucan phosphorylase (starch phosphorylase) and starch
branching enzyme IIa, were identified as potential Trx targets in
a proteomic analysis of isolated wheat amyloplasts [5]. The identification of Trx targets among enzymes thought to be active during
starch degradation in the dark has led to speculation about the
significance of these findings. The presence of a redox regulated,
stress-induced starch degradation pathway has been proposed as
an explanation for the Trx induced activation of AtBAM1 [12] and
there is some evidence to support this [17].
The experimental evidence hinting at a potentially extensive
redox regulation of starch metabolism prompted us to perform
a comprehensive survey of the effect of manipulating redox
potentials in vitro on the activity of chloroplastic starch metabolising enzymes in Arabidopsis thaliana. Using native substratecontaining gels (zymograms) and enzyme assays we were able to
identify several enzymatic activities among both the biosynthetic
and degradative enzymes of starch metabolism as potential targets
of redox regulation in planta.
-3
90
mV
Fig. 1. Redox mediated changes in starch metabolising enzyme activities. An extract of
Arabidopsis wild type was treated with different ratios of reduced to oxidised DTT
(DTT/DTTox) in a total concentration of 40 mM. The redox potentials of the mixtures
were calculated using the Nernst equation and a midpoint redox potential of DTT at pH
7.9 of 380 mV. Enzyme activities were detected in zymograms containing amylopectin (A) or glycogen (B). Starch synthase activities were visualised after incubation in
buffer containing ADP-glucose (B). The identified enzyme activities are indicated by
arrows. Representative zymogram shown.
Fig. S1). The identifiable redox insensitive enzymes included the
cytosolic AtBAM5 and disproportionating enzyme 1 (AtDPE1). The
identities of the observed in-gel activities were assigned by
comparison to previously published results [18e21] and, where
required, verified by analysis of Arabidopsis knock-out mutants in
each enzyme (data not shown; Fig. S1). The activity of the
remaining chloroplastic isoforms of BAM, SS, and BE, as well as the
isoamylase AtISA3 could not be visualised by zymogram analysis.
The ability of the observed enzyme activities to respond to
physiological redox potentials was investigated by redox titration of
plant extracts using different ratios of reduced to oxidised DTT.
2. Results
2.1. Zymogram analyses of redox sensitive starch metabolising
enzymes
Potential redox sensitive starch metabolising enzymes were
identified by modulating the redox potential in leaf extracts of
A. thaliana (At) using the reducing redox reagent dithiothreitol
(DTT) and the corresponding oxidised form (trans-4,5-dihydroxy1,2-dithiane; DTTox) followed by zymogram analysis. Among the
starch hydrolysing enzymes active on amylopectin, b-amylase 1
(AtBAM1), limit dextrinase (AtLDA), and the isoamylase complex
AtISA1/AtISA2, were only active under reducing conditions (Fig. 1A).
Analysis of soluble starch synthases (SS) and starch branching
enzymes (BE) in glycogen-containing and substrate-free zymograms, respectively, showed reductive activation of AtSS1, AtSS3
and AtBE2 (Figs. 1B and 2). The glycogen-containing zymograms
also revealed activation of two degrading activities corresponding
to AtBAM1 and the chloroplastic a-amylase AtAMY3 (Fig. 1B;
BE2
1st
2nd
-
DTT
DTT
DTT
DTTox
DTTox
DTTox
DTTox
DTT
Fig. 2. Redox mediated changes in starch branching enzyme activity. Extracts of
Arabidopsis were reduced or oxidised with 20 mM DTT or DTTox, respectively, for 1 h
at 25 C (1st). After gel filtration on Sephadex G-25 columns, extracts were subjected to
the same or the reverse treatment (2nd). Extracts were separated by native PAGE and
SBE activity was visualised using the phosphorylase a stimulation method followed by
iodine staining. Representative zymogram shown.
M.A. Glaring et al. / Plant Physiology and Biochemistry 58 (2012) 89e97
2.2. Quantitative analyses of redox sensitivity
In order to quantitatively analyse the effect of modulating the
redox conditions in vitro, enzyme assays were carried out using
extracts of Arabidopsis wild type and various knock-out mutants
after treatment with reductant or oxidant to obtain a fully reduced
or oxidised extract, respectively. Where possible, the reversibility of
the observed activity-changes was also investigated. For some
enzymes, the direct thiol exchange rate with DTT and DTTox in vitro
is very slow and redox equilibrium is not attained in the lifetime of
the enzyme [25,26]. Thioredoxins can catalyse this exchange and
although plant extracts contain potentially active endogenous Trxs,
recombinant Arabidopsis Trxs were included in the extracts used in
the following analyses to increase the efficiency of the DTT and
DTTox treatments. Two thioredoxins with a ubiquitous expression
pattern in green tissues (www.genevestigator.com) were chosen as
representatives of families f and m of the chloroplastic thioredoxins; TRXf1, which is an activator of most thioredoxin targets [4], and
TRXm4, which is an efficient activator of chloroplastic NADP-malate
dehydrogenase [27]. Since the thiol-disulfide state of enzymes
cannot be reliably preserved during extraction, it was not possible
to directly investigate the changes in activity that might occur as
a response to altered redox conditions in planta, such as during
a diurnal cycle.
2.3. b-Amylases
A b-amylase (BAM) was first picked out as a target of Trx in
spinach chloroplasts [28]. Subsequently, AtBAM1 was shown to be
redox regulated in vitro and directly activated by DTT and chloroplast Trxs, with TRXf1 being the most efficient redox mediator
[12,17]. In an attempt to identify additional redox regulated bamylases we analysed the effect of modulating the redox conditions
in extracts of an AtBAM1 mutant. A small, but significant (p < 0.01,
paired t-test), increase in total b-amylase activity was observed
after treatment with DTT suggesting that at least one other bamylase is activated by reduction (Fig. 3). This is comparable to
results obtained earlier [17]. Experiments with an AtBAM3 mutant
showed a similar activation by DTT, presumably of endogenous
AtBAM1. Interestingly, analysis of an AtBAM1/3 double mutant did
not reveal any significant changes in activity in response to DTT or
DTTox, indicating that AtBAM3 could be responsible for the
observed response (Fig. 3).
There is no conservation of the two redox-active cysteine residues between AtBAM1 and any other BAM in Arabidopsis (C32 and
C470 in the mature peptide) [12]. Three of the seven cysteines in
AtBAM3 are located in two flexible loops involved in substrate
binding in soybean b-amylase and are adjacent to, or part of, the
active site (C169, C177 and C257) [29]. Homology modelling the
AtBAM3 structure on soybean b-amylase showed that the three
cysteines are close in the tertiary structure and could potentially
form disulfide bridges that would block access to the active site
(Fig. S2).
120
115
% of water control
Redox potentials covering the range of midpoint potentials for
known redox regulated chloroplastic enzymes were investigated
(380 to 300 mV at pH 7.9). A clear change in activity was
observed at potentials around 340 to 320 mV (Fig. 1A and B). This
is within the range reported for other redox regulated chloroplast
enzymes such as Arabidopsis phosphoribulokinase (PRK, 330 mV)
[22], spinach NADP glyceraldehyde-3-phosphate dehydrogenase
(GAPDH, 353 mV) [23], and tomato fructose-1,6-bisphosphatase
(FBPase), PRK, and ATP synthase (348 mV, -315 mV and
335 mV, respectively) [24], as well as the starch metabolising
enzymes BAM1 (350 mV) [12] and GWD (310 mV) [11].
91
110
105
100
95
90
85
DTT
DTTox
Fig. 3. Total b-amylase activity in reduced and oxidised extracts of Arabidopsis.
Extracts of wild type Col-0 and BAM knock-out mutants were treated with 20 mM
DTT/DTTox and 2 mM each of recombinant TRXf1 and TRXm4 for 1 h at 25 C. Total bamylase activity was assayed on p-nitrophenol maltopentaoside (PNPG5). Measurements are the means SE of four independent samples. The activity of a water-treated
control was set to 100% for each sample.
2.4. a-Amylases
An endoamylase was previously isolated from poplar leaves and
shown to be activated by reductants and E. coli Trx [14]. However,
given the reported size (44 kD) this is unlikely to be an orthologue
of the large chloroplastic a-amylases [30]. We initially observed
a reductive activation of a degrading activity in glycogencontaining zymograms (Fig. 1B). This activity was absent in
knock-out mutants of the chloroplastic a-amylase AtAMY3 suggesting that this enzyme could be redox regulated (Fig. S1). This
prompted us to study the effect of DTT on recombinant, purified
AtAMY3, which demonstrated a clear dependency on DTT for
activity on amylopectin. The oxidised inactive form had only 1.6%
(0.7%) activity compared to the active form obtained after treatment with DTT (100 1.3%).
2.5. Starch phosphorylases
An a-1,4-glucan phosphorylase (PHS) was previously identified
as a putative Trx target in wheat amyloplasts [5]. Using enzyme
assays of plant extracts, we tested the effect of DTT and DTTox on
total PHS activity on amylopectin and maltoheptaose. Neither
treatment led to a significant change in activity (p > 0.05, paired ttest) when compared to a water-treated control (Table 1). Oxidation
by 100 mM CuCl2 did lead to a substantial loss of activity, however,
this could not be efficiently recovered by subsequent desalting and
treatment with DTT (Table 2), indicating an irreversible inactivation
of PHS activity.
2.6. Isoamylases
The Arabidopsis genome encodes three isoamylases. AtISA1 and
AtISA2 form a multimeric enzyme complex involved in starch
biosynthesis and AtISA3 is involved in starch degradation [31].
Visualisation of AtISA1/AtISA2 in zymograms showed that DTT is
required for full activity of this enzyme complex (Fig. 1A). It cannot
be ruled out that this is a consequence of a redox mediated effect on
the enzyme complex, leading to an altered position in the gel, and
not a change in total activity. However, as it is not possible to
specifically assay AtISA1/AtISA2 activity in plant extracts, this could
92
M.A. Glaring et al. / Plant Physiology and Biochemistry 58 (2012) 89e97
Table 1
Redox sensitivity of starch metabolising enzymes. Extracts of Arabidopsis were
treated with reductant (Red; 20 mM DTT) or oxidant (Ox; 20 mM DTTox for PHS,
ISA3 and MDH; 250 mM CuCl2 for DPE1) supplemented with 2 mM each of
recombinant TRXf1 and TRXm4 for 1 h at 25 C and immediately assayed. PHS
activity was assayed on both amylopectin and maltoheptaose. ISA3 activity was
measured in extracts of a LDA mutant on b-limit dextrin in the presence of saturating amounts of barley b-amylase and the results are given as production of
maltose and maltotriose. Chloroplastic NADP-dependent malate dehydrogenase
(MDH) was used as a redox control. Measurements are the means SE from three
independent samples. The activity is given as percentage of a water-treated control.
Enzyme
Red (%)
PHS on amylopectin
PHS on maltoheptaose
ISA3, maltose
ISA3, maltotriose
MDH
DPE1
108
115
111
114
2130
100
Ox (%)
6.0
6.2
3.3
4.1
59
5.7
101
100
91.6
96.7
96.9
98.9
2.8
0.8
2.0
2.2
0.5
5.9
not be verified by other methods. The redox sensitivity of AtISA3
was determined by a specific assay on b-limit dextrin in extracts of
an AtLDA mutant following treatment with DTT or DTTox.
Measurements of maltose and maltotriose release from b-limit
dextrin showed a significant difference between the reduced and
oxidised samples for both products (p < 0.05, paired t-test; Table 1),
although this only represented a total difference in activity of less
than 20%.
2.7. Disproportionating enzyme
The chloroplastic disproportionating enzyme AtDPE1 was
visible as a faint red band in amylopectin-containing gels and did
not appear to respond to changes in the redox potential (Fig. 1A).
This was confirmed by enzyme assays in extracts of Arabidopsis
using the DPE1 specific substrate maltotriose. No change in AtDPE1
activity was observed after treatment with DTT or high concentrations of CuCl2 (250 mM; Table 1).
2.8. Limit dextrinase
Limit dextrinase, also called pullulanase, has previously been
identified as redox sensitive in vitro in spinach and wheat [15,16] and
Table 2
Inactivation and reactivation of starch metabolising enzymes. Extracts of Arabidopsis wild type Col-0 were treated with reductant (1st; 20 mM DTT) or oxidant
(2nd; 100 mM CuCl2 or 20 mM DTTox) for 1 h at 25 C, desalted on Sephadex G-25 gel
filtration columns, and subsequently treated with 20 mM DTT or 20 mM DTTox/
water for 1 h at 25 C before assaying for enzyme activity. MDH was used as a redox
control. Measurements are the means SE from three (LDA, PHS, MDH) or five (SS,
total soluble starch synthase activity) independent samples. Activity is given as
a percentage of the fully reduced samples.
1st
2nd
Enzyme
Activity (%)
DTT
DTT
DTT
Water
CuCl2
DTT
CuCl2
Water
DTT
DTT
DTTox
DTTox
Water
DTTox
Water
DTT
LDA
PHS
MDH
LDA
PHS
MDH
LDA
PHS
MDH
LDA
PHS
MDH
SS
SS
SS
SS
100
100
100
89.9
98.1
5.07
106
84.0
46.3
6.73
79.5
3.79
100
51.2
14.9
71.4
7.7
5.2
3.9
8.7
4.6
0.1
6.5
4.7
7.3
2.6
3.8
0.2
9.6
4.1
1.5
5.2
AtLDA showed a clear dependency on DTT for full activity when
examined in amylopectin-containing gels (Fig. 1A). This observation
was confirmed in gels containing red-pullulan, which is a substrate
exclusively degraded by LDA (Fig. S3). Using CuCl2 as an oxidising
agent it was possible to decrease the activity of the fully reduced
AtLDA to less than 10% of maximum activity. This loss of activity could
be completely recovered by subsequent treatment with DTT,
demonstrating the reversibility of the oxidationereduction reaction
(Table 2).
The three cysteines tentatively proposed to be involved in redox
sensitivity and isoform microheterogeneity of spinach pullulanase
(C390, C452 and C677) [15] are conserved in AtLDA (C456, C518 and
C743). Modelling the structure of AtLDA on barley LDA indicated
that, although these three cysteines are located on the same side of
the central a/b-barrel, as suggested earlier [15], there is considerable distance between them, making direct disulfide bridge
formation dependent on a significant conformational change
(Fig. S4). Only one cysteine is conserved between Arabidopsis,
spinach and wheat LDA (Arabidopsis C743), hinting at a different
mechanism of redox sensitivity in these enzymes.
2.9. Soluble starch synthases
Arabidopsis contains four soluble starch synthases (SS) which
serve distinct functions in amylopectin biosynthesis [21]. Two
major SS activities can be visualised in glycogen-containing
zymograms (SS1 and SS3) [21] and an analysis of Arabidopsis
extracts showed that both of these SSs were activated by
reduction (Fig. 1B). Following separation under oxidising conditions, both enzymes could be activated in the gel by incubation
with DTT, indicating that the lack of visible activity of the oxidised enzymes was not caused by an altered position in the gel
(Fig. S5). The observed redox sensitivity was corroborated by
enzyme assay of both wild type and AtSS1 and AtSS3 mutant
extracts after treatment with DTT or DTTox, clearly showing that
DTT is required for full SS activity (Table 3). Gel filtration of
oxidised and reduced extracts followed by incubation with DTT
or DTTox, respectively, demonstrated that the redox mediated
changes in activity were mostly reversible, suggesting that AtSS1
and AtSS3 could be targets of redox regulation in Arabidopsis
(Table 2).
2.10. Starch branching enzymes
The Arabidopsis genome encodes three starch branching
enzymes (BE) of which one (AtBE2) is responsible for the majority
of the measurable activity in plant extracts [20]. The closest
homologue of AtBE2 in wheat is SBEIIa (77% identity), which has
been identified as a potential Trx target in wheat amyloplasts [5].
Zymograms of BE activities in Arabidopsis extracts revealed one
major activity corresponding to AtBE2, that was only active under
reducing conditions. Similar to the observation made for SS, this
redox mediated change was reversible (Fig. 2).
Table 3
Redox sensitivity of soluble starch synthases. Extracts of wild type (wt) and SS1 and
SS3 mutants were treated with 20 mM DTT or DTTox, or water (control) supplemented with 2 mM each of TRXf1 and TRXm4 for 1 h at 25 C and immediately
assayed for soluble starch synthase activity. Activity is given as a percentage of the
DTT-treated wild type sample. Measurements are the means SE from five samples.
Plant
DTT (%)
DTTox (%)
Water (%)
wt
SS1 mutant
SS3 mutant
100 7.8
49.0 2.5
49.5 3.8
10.1 1.3
1.63 1.4
13.7 2.7
13.0 2.8
0.94 2.1
24.6 4.6
M.A. Glaring et al. / Plant Physiology and Biochemistry 58 (2012) 89e97
3. Discussion
It has been known for some time that the rate of starch
biosynthesis is controlled at least partially at the level of ADPglucose production by redox regulation of the AGPase enzyme.
Production of ADP-glucose by AGPase is generally viewed as the
first committed step in starch biosynthesis and the enzyme makes
a considerable contribution to the control of flux through this
pathway [32,33]. Despite increasing amounts of data from
numerous in vitro studies pointing to a broader role of redox
regulation in starch metabolism, the subject has never been
comprehensively investigated. In this study we have surveyed the
majority of chloroplastic enzymes known to be involved in starch
metabolism in A. thaliana and have identified several redox sensitive enzyme activities.
3.1. Redox regulatory mechanisms
A thiol-disulfide exchange can lead to subtle changes in the
properties of an enzyme such as pH optimum and range, substrate
affinity, and sensitivity to allosteric regulators. For example,
reduction of spinach LDA broadens the pH range of activity by one
unit in the alkaline range, making the reduced enzyme active at
chloroplastic pH values (above pH 7.0) [15]. Reduction of AGPase
changes the kinetic parameters of the enzyme, leading to increased
sensitivity to the allosteric activator glycerate 3-phosphate and
a decreased sensitivity to inhibition by Pi, thus complementing
other regulatory mechanisms [7,34].
A thiol-disulfide exchange can potentially influence the formation of protein complexes. The Trx-regulated Calvin cycle enzymes
GAPDH and PRK form a supramolecular complex with the protein
CP12 under oxidising conditions [35] and starch biosynthetic
enzymes can associate to form large enzyme complexes in amyloplasts [36]. Consequently, it is possible that the redox effects
observed in the zymograms are caused by changes in protein
complex formation leading to an altered position in the zymogram.
At least for AtSS1 and AtSS3 this was not the case, as both enzymes
could be reactivated at the same position in the gel after separation
under oxidising conditions. In addition, it cannot be ruled out that
a perceived redox sensitivity is a consequence of a thiol-disulfide
exchange in another protein, which leads to an indirect effect on
the activity of the analysed enzyme. For example, post-translational
modification by phosphorylation has been suggested as a regulatory mechanism for many starch metabolising enzymes [10] and
the controlling factors themselves (e.g. kinases and phosphatases)
could be subject to redox regulation.
Glutathionylation of proteins, and the reverse process catalysed
by the glutaredoxins (Grx), has been proposed as an alternative
mechanism of redox regulation. There is considerable overlap
between the Grx- and Trx-interacting targets in vitro, including
AGPase and several Calvin cycle enzymes, and the Grx and Trx
pathways could serve complementary functions in redox regulation in vivo [37,38]. There is no evidence in the literature for a direct
role of glutathionylation in leaf starch metabolism, however, based
on the current study, such an effect cannot be ruled out and an
inhibition of activity caused by glutathionylation in oxidized
extracts could potentially be reversed by DTT treatment.
Multiple factors might contribute to the regulation of an
enzyme’s activity. Furthermore, the regulatory mechanisms might
influence one another. Hence, manipulation of the redox conditions
may not necessarily lead to a large change in enzyme activity if,
under a given set of conditions, its activity and/or redox sensitivity
is simultaneously affected by other factors. Since some factors, such
as the presence of allosteric regulators or proteinaceous inhibitors/
activators, cannot be accurately controlled in plant extracts, any
93
verification and detailed analyses of redox sensitivity would
require the recombinant enzyme for in vitro studies.
3.2. Identification of redox sensitive starch metabolising enzymes
Starch biosynthesis in Arabidopsis requires the combined action
of SSs and BEs, as well as a debranching activity provided by ISA
[39]. Using zymograms, we observed an effect of modulating the
redox conditions on all three classes of enzymes suggesting that
starch biosynthesis is under redox control. Due to lack of a specific
assay for AtISA1/AtISA2 activity in plant extracts [40], the redox
sensitivity of this enzyme could not be verified outside the gelbased system. Similarly, we were unable to determine the activity
of AtBE2 in plant extracts using the phosphorylase a stimulation
method, possibly due to interference from endogenous degrading
enzymes. For the SSs, the observation was confirmed by enzyme
assays of wild type and SS mutants indicating that AtSS1 and AtSS3
are potential targets of redox regulation. The notion that SS activity
is under redox control adds a significant layer of complexity to the
control of starch biosynthesis, as it suggests that both the supply of
ADP-glucose and its utilisation is co-ordinately regulated and
coupled to light capture by the photosynthetic electron transport
chain.
The precise molecular mechanisms determining starch structure are not fully understood, but is a product of the combined
activities of SS, BE and ISA isoforms [21,41,42]. Our findings raise
the possibility that the structure of starch might be influenced by
the redox status of the chloroplast by changes in the relative
contributions of the different redox regulated isoforms. It has also
been proposed from work in maize and wheat endosperm that
both protein phosphorylation and protein complex formation
involving SS and BE isoforms may play roles in controlling starch
biosynthesis [36,42,43]. Redox regulation could act in parallel with
such mechanisms to ensure correct synthesis of the starch granule,
although there is as yet no evidence for such protein complex
formation in leaves. Redox regulation of starch biosynthetic
enzymes could also serve to ensure that the pathway of starch
synthesis is not active during the dark period, preventing energy
wastage in a futile cycle.
The major starch degradation pathway in Arabidopsis involves
the interdependent actions of the chloroplastic b-amylases AtBAM1
and AtBAM3 and the debranching enzyme AtISA3 [10,29,39]. A
small but significant difference in AtISA3 activity was observed
between reduced and oxidised extracts. If this is indicative of true
redox sensitivity, it is possible that oxidation by DTTox is inefficient,
so that the enzyme remains in a stable reduced form in the oxidized
extracts. The previously reported redox sensitivity of AtBAM1 [12]
was confirmed in zymograms of Arabidopsis leaf extracts. A
second visible b-amylase activity corresponded to the cytosolic
AtBAM5 and this enzyme was unaffected by DTT treatment and
high concentrations of the oxidiser CuCl2 (data not shown). Enzyme
assays of an AtBAM1 knock-out mutant suggested the presence of
an additional redox sensitive b-amylase activity. AtBAM3 was
previously identified as a major contributor to starch degradation
[29] and total b-amylase activity in the double mutant AtBAM1/
AtBAM3 was insensitive to redox changes, suggesting that AtBAM3
itself, or a component influencing AtBAM3 activity, is redox sensitive. Due to the relatively small changes in total b-amylase activity
measured in the AtBAM1 mutant and the extra steps required to
sequentially oxidise and reduce the plant extract, it was not
possible to reliable determine the reversibility of this redox reaction. Although a homology model of AtBAM3 identified three
cysteine residues in close proximity to both each other and the
active site, further experiments will be required to clarify the
potential redox sensitivity of AtBAM3.
94
M.A. Glaring et al. / Plant Physiology and Biochemistry 58 (2012) 89e97
AtAMY3 and AtLDA are part of a minor, alternative pathway that
can support starch degradation in Arabidopsis, although the
physiological significance of this pathway is unclear [39,40]. Both
enzymes were sensitive to redox conditions and clearly activated by
reduction in zymograms and enzyme assays. Furthermore, inactivation of AtLDA by oxidation was almost completely reversible by
subsequent reduction. Interestingly, reduction of spinach LDA
broadens the pH range of the enzyme, making it active at nighttime chloroplastic pH values (above pH 7.0) [15]. Spinach and
wheat LDAs are unusual in that they exist as multiple interconvertible isomeric forms [15,16]. Different redox states of three
cysteine residues, as either thiols or disulfides, were proposed as an
explanation for the coexistence of multiple forms [15]. Given that
spinach LDA does not appear to respond to Trx in vitro [15] and that
only one cysteine is conserved between the redox sensitive LDAs
from spinach, wheat and Arabidopsis, the redox sensitivity of this
enzyme may represent an unusual adaptation to diurnal
metabolism.
3.3. The role of redox regulation in starch metabolism
Starch degradation in leaves includes an initial cycle of reversible glucan phosphorylation involving GWD and SEX4 [39,44]. Both
of these enzymes are redox regulated and activated by Trx in vitro
[11,13], suggesting that the initial attack on the starch granule is
under redox control. The results presented here point towards
a similar regulatory mechanism of the enzymes acting downstream
of GWD/SEX4 and opens up the possibility that both of the established pathways of starch degradation in Arabidopsis are redox
regulated and, surprisingly, activated by reduction. While reductive
activation of enzymes involved in starch biosynthesis is intuitively
relevant for starch deposition during active photosynthesis, the
significance of reductive activation of starch degrading enzymes is
less obvious. At least two explanations can be imagined. First, that
redox regulation represents an adaptation to special conditions,
such as abiotic or biotic stresses, which may require a change in the
normal diurnal metabolism of starch to fulfil the altered needs of
the plant and secondly, that some degrading enzymes carry out
specialized functions in specific tissues (e.g. in amyloplasts of
heterotrophic tissues) or at certain developmental stages. There is
some evidence to support the idea of such specialised roles. Potato
GWD has the most positive midpoint redox potential among known
redox regulated enzymes (310 mV at pH 7.9) [11] suggesting that
GWD exists primarily in the reduced, active form during a normal
diurnal cycle as proposed by Edner et al. [45]. Any condition that
alters the redox balance and results in a more oxidising environment could thus affect the activity of GWD and hence starch
degradation. AtBAM1 is not required for normal starch degradation
in leaves, but contributes to starch breakdown in the absence of the
major b-amylase AtBAM3 [29]. A recent study suggested that
AtBAM1 is required for starch degradation in guard cells during the
day [17]. Furthermore, AtBAM1 expression and activity is induced
by osmotic stress in mesophyll cells and evidence was provided for
a role of AtBAM1 in a stress-induced pathway leading to starch
degradation in the light [17]. Starch degradation in the light
involving b-amylase activity has also been demonstrated under
photorespiratory conditions in Arabidopsis [46] and b-amylase
expression and activity is up-regulated and involved in production
of maltose as a compatible solute in response to abiotic stress [47].
While reductive activation clearly requires reducing power, this
does not necessarily come directly from photosynthesis. Reductive
activation is possible when photoreduced Fdx is not available. In
wheat amyloplasts, Fdx can be reduced by NADPH via ferredoxinNADP reductase and this metabolically reduced Fdx can provide
reducing equivalents for downstream Trxs [5]. The NADPH itself is
produced by the respiration of carbohydrates via the oxidative
pentose phosphate pathway. This mechanism was suggested to
couple photosynthesis in leaves to biosynthetic pathways in the
amyloplasts of storage tissues via the availability of translocated
sugars. However, reductive activation of enzymes can also occur in
chloroplasts in the absence of light provided that NADPH is present.
External feeding of sugars to Arabidopsis leaves in the dark
increases the pool of reduced AGPase and the rate of starch
biosynthesis, linking sugar availability and redox regulation [6,8].
This activation of AGPase is dependent on the plastidial NTRC,
which uses NADPH as a source of reducing power [8,17]. Despite
these observations, it is still unclear what mechanism would trigger
the reductive activation of starch degrading enzymes, as starch
breakdown is a process which supplies sugars in the dark, rather
than being triggered by the presence of sugars. Simultaneous
activation of both biosynthetic and degradative enzymes seems
unlikely, so some as yet unidentified specificity in the activation
pathways is to be expected. This could either be in the interaction
between the reducing activator (e.g. Trx/NTRC) and the target
enzyme or in the interaction with potential redox-regulated
protein factors controlling activity by other post-translational
mechanisms (see above). At the plant level, this could imply
separation in terms of the signals (e.g. light or sugars) that lead to
activation or the conditions under which such a regulation is
required.
The above examples and the results obtained in this study
support the idea that redox regulation of enzyme activity plays
a role in starch metabolism. This does not necessarily only include
regulation of the normal diurnal cycle in leaves, but could be
a mechanism for regulating starch metabolic pathways operating
outside the normal diurnal cycle in heterotrophic or specialised
tissues or in response to environmental signals. Further work will
be needed to clarify the role and significance of redox regulation for
both individual enzymes and the pathways in which they operate
in planta.
4. Materials and methods
4.1. Plant material
A. thaliana wild type Col-0 and mutants were grown from seed
in potting compost in growth chambers at 20 C and 70% relative
humidity with a 12/12 h photoperiod at a photon flux density of
120e150 mmol m2 s1. Leaves were harvested directly into liquid
nitrogen 2 h before the end of the light period and kept at 80 C
until analysis.
4.2. Zymograms
Soluble proteins were extracted in buffer using a ground-glass
homogeniser (200e400 mg fresh-weight (FW) per ml). Insoluble
material was pelleted at 14,000 g for 10 min at 4 C and the supernatant was used immediately in zymogram experiments. Gels were
prepared as previously described [48] using 0.2% (w/v) potato
amylopectin (SigmaeAldrich 10118, Copenhagen, Denmark),
0.8e1.0% (w/v) glycogen (SigmaeAldrich G1508), or 1.0% (w/v) red
pullulan (Megazyme, Bray, Ireland) as substrate in the separating gel.
After electrophoresis at 4 C, gels were washed twice in incubation
buffer for 15 min and subsequently incubated in the same buffer for
3e5 h at 37 C or up to 16 h at 25 C. Activities were revealed after
staining with an iodine solution (0.34% I2, 0.68% KI (w/v)).
Starch hydrolysing activities were extracted in 100 mM MOPS
pH 7.2, 10% (v/v) glycerol and separated on 1% (w/v) amylopectin
gels. Gels were washed and incubated in 100 mM Tris pH 7.0, 1 mM
CaCl2, 1 mM MgCl2.
M.A. Glaring et al. / Plant Physiology and Biochemistry 58 (2012) 89e97
Starch synthases were extracted in 100 mM Bicine pH 8.3, 1 mM
EDTA, 10% (v/v) glycerol and separated on gels containing 0.8% (w/
v) glycogen. Gels were washed in 100 mM Bicine pH 8.3, 0.5 M
sodium citrate, 0.5 mM EDTA, 10% (v/v) glycerol and incubated in
the same buffer supplemented with 2 mM ADP-glucose.
Starch branching enzymes were extracted in 50 mM HEPES pH
7.0, 1 mM EDTA, 10% (v/v) glycerol and separated on a native gel
with no substrate. Gels were incubated in 50 mM HEPES pH 7.0, 10%
(v/v) glycerol, 50 mM glucose-1-phosphate, 2.5 mM AMP, 1 U/ml
phosphorylase a from rabbit muscle (SigmaeAldrich P1261).
4.3. Enzyme assays
Soluble proteins were extracted as described for the zymograms. Based on previous Trx activation experiments with potato
GWD and AtBAM1 [11,12] extracts were supplemented with 2 mM
each of recombinant Arabidopsis thioredoxins TRXf1 and TRXm4.
Assay conditions were essentially as previously described [48]
unless otherwise mentioned.
Recombinant AtAMY3: Recombinant, purified AtAMY3 was
obtained from a previous study [30]. Activity of a-amylase was
assayed on amylopectin (5 mg/ml) in 20 mM HEPES pH 7.5, 0.5 mM
CaCl2, 0.05 mg/ml BSA at 37 C. The assay was stopped by adding an
equal volume of 0.5 M NaOH, and liberated reducing ends were
determined using the MBTH method as described [49].
b-amylase: Plant material (200 mg FW/ml) was extracted in
50 mM MES pH 6.2, 10% (v/v) glycerol, and b-amylase activity was
measured using a p-nitrophenol maltopentaoside substrate
(PNPG5) according to manufacturer instructions (Betamyl Reagent,
Megazyme). Activity was measured for 1 h at 30 C in a 300 ml total
volume containing 50 mM MES pH 6.2, 1 mM EDTA and 25 ml
Betamyl substrate.
Limit dextrinase: Plant material (400 mg FW/ml) was extracted
in 100 mM MOPS pH 7.2, 10% (v/v) glycerol, 2 mM MgCl2, 50 mg/ml
polyvinylpolypyrrolidone (PVPP). The 200 ml assay mixture contained 50 mM MOPS pH 7.2 and 10 mg/ml pullulan (Megazyme).
After a 4 h incubation at 30 C, the assay was stopped by adding
200 ml 0.5 M NaOH and reducing ends were determined using the
MBTH method as described [49].
Disproportionating enzyme: Plant material (200 mg FW/ml)
was extracted in 100 mM MOPS pH 7.2, 1 mM EDTA, 10% (v/v)
glycerol. The 250 ml assay mixture contained 50 mM MOPS pH 6.8
and 60 mM maltotriose. Assays were stopped by boiling after 1 h at
25 C and liberated glucose was determined.
Starch phosphorylase: Plant material (200 mg FW/ml) was
extracted in 100 mM MOPS pH 7.0, 10% (v/v) glycerol. The 250 ml
assay mixture contained 20 mM MOPS pH 7.0, 10 mM MgCl2,
20 mM NaH2PO4/Na2HPO4, 3.4 mM NAD, 2.5 mM glucose-1,6bisphosphate, 1 U/ml phosphoglucomutase, 1 U/ml glucose-6phosphate dehydrogenase (Roche Applied Science, Rotkreuz,
Switzerland) and the assay was started by adding 2.5 mg/ml
amylopectin or 1 mM maltoheptaose as substrate. Activity,
expressed as the production of NADH, was measured by monitoring
the absorbance at 340 nm in a microplate reader.
Isoamylase: Plant material (400 mg FW/ml) was extracted in
50 mM MOPS pH 7.2, 10% (v/v) glycerol and desalted on Sephadex
G-25 gel filtration columns (NAP-5, GE Healthcare, Glattbrugg,
Switzerland) equilibrated with extraction buffer. Debranching
activity by AtISA3 was determined on b-limit dextrin essentially as
described [40] except that commercial b-limit dextrin (Megazyme)
was used as a starting point for the second round of b-amylase
treatment. The 100 ml assay mixture contained 50 mM MOPS pH
7.2, 1 mM EDTA and 2 mg/ml b-limit dextrin. Assays were stopped
by boiling after 1 h at 30 C. Maltose and maltotriose release was
quantified by high-performance anion-exchange chromatography.
95
Soluble starch synthase: Plant material (200 mg FW/ml) was
extracted in 100 mM Bicine pH 8.3, 1 mM EDTA, 10% (v/v) glycerol.
Assays were performed in a final volume of 100 ml containing SS
buffer (100 mM Bicine pH 8.3, 25 mM potassium acetate, 5 mM
EDTA), 18 mg/ml amylopectin, 740 Bq (20 nCi) [14C]ADP-glucose,
1 mM ADP-glucose, and incubated for 30 min at 30 C. The reactions were stopped by heating at 90 C for 3 min. Excess [14C]ADPglucose was removed by anion-exchange on Dowex 1 8 200e400
Mesh (SigmaeAldrich) columns. Radioactivity incorporated into
amylopectin was determined by scintillation counting after addition of 3 ml scintillation fluid.
NADP-dependent malate dehydrogenase: Assays were carried
out in microtiter plates at 25 C in a 250 ml volume containing
100 mM Tris pH 8.0, 0.2 mM NADPH, 1 mM oxaloacetic acid. The
rate of NADPH consumption was followed spectrophotometrically
at 340 nm.
4.4. Redox manipulations
Redox titration of Arabidopsis extracts was carried out under
aerobic conditions at 25 C by incubation with different ratios of
reduced to oxidised DTT in a total concentration of 40 mM for 1 h.
The redox potentials were calculated using the Nernst equation and
a midpoint redox potential of DTT at pH 7.9 of 380 mV. For
demonstrations of redox reversibility, extracts were treated with
reductant (20 mM DTT) or oxidant (20 mM DTTox or 100 mM CuCl2)
for 1 h at 25 C, desalted on Sephadex G-25 gel filtration columns
(NAP-5, GE Healthcare), and then subjected to the reverse treatment for 1 h. The oxidant CuCl2 was used when DTTox had no
apparent effect on enzyme activity and it has previously been
successfully used as an inactivator of potato GWD [11]. Watertreated controls were used as a baseline for activity in the
untreated extracts and were treated exactly as the other samples
except water replaced the added reductant or oxidant. Recombinant AtAMY3 was stored as the inactive form in the absence of
reductant and activated by treatment with 20 mM DTT for 20 min
at 25 C.
4.5. Production of recombinant thioredoxins
The open reading frames of A. thaliana thioredoxins f1
(AT3G02730) and m4 (AT3G15360), excluding the predicted chloroplast transit peptide (amino acids 1e59 and 1e76 for f1 and m4,
respectively), were synthesised as codon-optimised constructs for
expression in E. coli. The ORFs were cloned directly into the
expression vector pET15b and expressed as N-terminally 6xHistagged proteins in E. coli BL21-CodonPlus (Stratagene, Agilent
Technologies, Basel, Switzerland). Expression was induced by 1 mM
IPTG and cells were harvested after 4 h of induction at 30 C. Cells
were disrupted by sonication in lysis buffer (50 mM Tris pH 8.0,
300 mM NaCl, 20 mM imidazole, 0.05% (v/v) b-mercaptoethanol).
His-tagged proteins were purified on a 1 ml nickel-chelating resin
column using the ProBond Purification System (Invitrogen, Zug,
Switzerland). The bound thioredoxins were eluted with a stepwise
gradient of imidazole in elution buffer (50 mM Tris pH 8.0, 50 mM
NaCl, 100e500 mM imidazole) and stored at 80 C in 50 mM Tris
pH 8.0, 10% (v/v) glycerol.
4.6. Computational analyses
The AGI gene codes and any alternative names for the proteins
examined in this study were as follows: AMY3 (AT1G69830); BAM1
(BMY7, AT3G23920); BAM3 (BMY8/ctBMY, AT4G17090); BAM5
(BMY1/RAM1, AT4G15210); BE2 (SBE2.2, AT5G03650); DPE1 (AT5G
64860); ISA1 (AT2G39930); ISA2 (DBE1, AT1G03310); ISA3 (AT4G
96
M.A. Glaring et al. / Plant Physiology and Biochemistry 58 (2012) 89e97
09020); LDA (PUL, AT5G04360); PHS1 (PHO1, AT3G29320); PHS2
(PHO2, AT3G46970); SS1 (AT5G24300); SS3 (AT1G11720). Amino acid
alignments were produced using the ClustalW2 tool at EBI (www.ebi.
ac.uk/Tools/msa/clustalw2) with default settings and visualised using
Jalview (www.jalview.org). All amino acid numberings include the
putative chloroplast transit peptides. Structure modelling was performed using the automated mode in Swiss-Model (www.swissmodel.
expasy.org) [50]. The final models were based on the following
templates (PDB ID, www.pdb.org); BAM3, 1BYB; LDA, 2Y4S.
Role of the funding sources
The funding sources have had no role in any stages of this study.
Acknowledgements
We thank Professor Alison M. Smith (John Innes Centre, Norwich, UK) for her donation of starch synthase mutants. The financial support from The Danish Research Council for Technology and
Production Sciences (grant no. 274-06-0312), The Villum Kann
Rasmussen Foundation (to the VKR Research Centre Pro-Active
Plants), and ETH Zürich is gratefully acknowledged.
Appendix A. Supplementary material
Supplementary data associated with this article can be found, in
the online version, at http://dx.doi.org/10.1016/j.plaphy.2012.06.017.
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