Plant Cell Physiol. 46(8): 1366–1376 (2005)
doi:10.1093/pcp/pci148, available online at www.pcp.oupjournals.org
JSPP © 2005
Sucrose Synthase Controls Both Intracellular ADP Glucose Levels and
Transitory Starch Biosynthesis in Source Leaves
Francisco José Muñoz 1, 3, Edurne Baroja-Fernández 1, 3, María Teresa Morán-Zorzano 1,
Alejandro Miguel Viale 1, Ed Etxeberria 2, Nora Alonso-Casajús 1 and Javier Pozueta-Romero 1, *
1
Agrobioteknologiako Instituta, Nafarroako Unibertsitate Publikoa and Consejo Superior de Investigaciones Científicas, Mutiloako etorbidea
zenbaki gabe, 31192 Mutiloabeti, Nafarroa, Spain
2
University of Florida, IFAS, Department of Horticultural Sciences, Citrus Research and Education Center, 700 Experiment Station Road, Lake
Alfred, FL 33850, USA
;
The prevailing model on transitory starch biosynthesis
in source leaves assumes that the plastidial ADPglucose
(ADPG) pyrophosphorylase (AGP) is the sole enzyme catalyzing the synthesis of the starch precursor molecule,
ADPG. However, recent investigations have shown that
ADPG linked to starch biosynthesis accumulates outside
the chloroplast, presumably in the cytosol. This finding is
consistent with the occurrence of an ‘alternative’ gluconeogenic pathway wherein sucrose synthase (SuSy) is involved
in the production of ADPG in the cytosol, whereas both
plastidial phosphoglucomutase (pPGM) and AGP play a
prime role in the scavenging of starch breakdown products. To test this hypothesis, we have compared the ADPG
content in both Arabidopsis and potato wild-type (WT)
leaves with those of the starch-deficient mutants with
reduced pPGM and AGP. These analyses provided evidence
against the ‘classical’ model of starch biosynthesis, since
ADPG levels in all the starch-deficient lines were normal
compared with WT plants. Whether or not SuSy is involved
in the synthesis of ADPG accumulating in leaves was
tested by characterizing both SuSy-overexpressing and
SuSy-antisensed transgenic leaves. Importantly, SuSy-overexpressing leaves exhibited a large increase of both ADPG
and starch levels compared with WT leaves, whereas SuSyantisensed leaves accumulated low amounts of both ADPG
and starch. These findings show that (i) ADPG produced by
SuSy is linked to starch biosynthesis; (ii) SuSy exerts a
strong control on the starch biosynthetic process; and (iii)
SuSy, but not AGP, controls the production of ADPG accumulating in source leaves.
Keywords: ADPglucose — Arabidopsis thaliana — Solanum
tuberosum —Starch — Sucrose — Sucrose synthase.
Abbreviations: ADPG, ADPglucose; AGP, ADPG pyrophosphorylase; ASPP, adenosine diphosphate sugar pyrophosphatase;
E4P, erythrose-4-phosphate; G1P, glucose-1-phosphate; G6P, glucose6-phosphate; NOS, nopaline synthase; 3PGA, 3-phosphoglycerate;
pPGI, plastid phosphoglucose isomerase; pPGM, plastid phosphoglucomutase; RBCS, ribulose 1,5-bisphosphate carboxylase small subunit; 35S, cauliflower mosaic virus 35S promoter; SPS, sucrose
phosphate synthase; SuSy, sucrose synthase; UGP, UDPglucose pyrophosphorylase; WT, wild type.
3
*
Introduction
Plants accumulate starch in their leaves during the day and
degrade it during the subsequent night. Due to the diurnal rise
and fall cycle of its levels, starch found in photosynthetically
competent cells is termed ‘transitory starch’. Starch biosynthesis in leaves has generally been considered to take place
exclusively in the chloroplast, and segregated from the sucrose
biosynthetic process that takes place in the cytosol (Stitt et al.
1984, Okita 1992, Sonnewald 1992, Baroja-Fernández et al.
2004) (Supplementary Fig. 1). According to this ‘classical’
view, starch is considered to be the end-product of a unidirectional and vectorial pathway linked to the Calvin cycle wherein
ADPglucose (ADPG) pyrophosphorylase (AGP) exclusively
catalyzes the synthesis of the starch precursor molecule,
ADPG, and functions as the major regulatory step in the gluconeogenic process (Caspar et al. 1985, Lin et al. 1988, MüllerRöber et al. 1992, Okita 1992, Stark et al. 1992).
However, recent evidence has exposed inconsistencies in
the ‘classical’ view of transitory starch biosynthesis and previews the occurrence of an ‘alternative’ pathway wherein
ADPG linked to starch biosynthesis is produced de novo in the
cytosol by sucrose synthase (SuSy) (Baroja-Fernández et al.
2001, Baroja-Fernández et al. 2004, Baroja-Fernández et al.
2005) (Supplementary Fig. 1). According to this ‘alternative’
view, both sucrose and starch biosynthetic pathways are tightly
interconnected by means of ADPG producing SuSy activity
(Delmer 1972, Porchia et al. 1999, Baroja-Fernández et al.
2003), and by the action of an ADPG translocator located at the
chloroplast envelope membranes (Pozueta-Romero et al. 1991,
Baroja-Fernández et al. 2004).
Pulse–chase and starch pre-loading experiments using isolated chloroplasts (Stitt and Heldt 1981, Fox and Geiger 1984),
intact leaves (Scott and Kruger 1995, Walters et al. 2004) and
cultured photosynthetic cells (Lozovaya et al. 1996) have provided evidence indicating that chloroplasts can synthesize and
mobilize starch simultaneously. This observation is in agreement with reports dealing with gluconeogenic processes in bacteria (Gaudet et al. 1992, Belanger and Hatfull 1999, Guedon et
al. 2000), animals (David et al. 1990, Massillon et al. 1995),
These authors contributed equally to this work
Corresponding author: E-mail, [email protected]; Fax, +34-948232191.
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SuSy controls starch production in leaves
heterotrophic plant organs (Sweetlove et al. 1996, NguyenQuoc and Foyer 2001) and amyloplasts (Pozueta-Romero and
Akazawa 1993, Neuhaus et al. 1995). Accordingly, the ‘alternative’ view on transitory starch biosynthesis assumes that both
AGP and plastid phosphoglucomutase (pPGM) play vital roles
in the scavenging of glucose units derived from starch breakdown (Pozueta-Romero et al. 1991, Baroja-Fernández et al.
2001, Pozueta-Romero et al. 2003, Baroja-Fernández et al.
2004).
Against the idea that synthesis and breakdown of starch
occur simultaneously in leaves, pulse–chase experiments on
illuminated Arabidopsis leaves have recently shown that none
of the 14C incorporated into starch during a short pulse of 14CO2
was released during a subsequent chase (Zeeman et al. 2002).
Based on these results, and assuming that starch biosynthesis
occurs following a ‘primer non-reducing-end’ mechanism of
polysaccharide chain elongation, Zeeman et al. (2002) concluded that starch synthesis is not accompanied by significant
turnover. This conclusion, however, had an erroneous basis,
since it has been shown recently that starch biosynthesis occurs
following a ‘non-primer reducing-end’ insertion mechanism
(Mukerjea and Robyt 2004). Taking into account that (i) βamylase controls transitory starch degradation (Scheidig et al.
2002, Niittylä et al. 2004, Lloyd et al. 2005) and (ii) this
enzyme liberates maltose from the non-reducing end of starch
chains, it is not surprising that illuminated leaves exposed to
a short pulse of 14CO2 do not release any label during the
subsequent chase.
Under light conditions, sucrose transiently accumulates in
the photosynthetic cell (Stitt et al. 1983, Stitt et al. 1984,
Gerhardt et al. 1987, Draborg et al. 2001, Smith et al. 2004).
According to the ‘alternative’ model of transitory starch biosynthesis, SuSy will produce from sucrose the ADPG necessary for starch biosynthesis. Concurrently, both AGP and
pPGM will scavenge starch breakdown products in the chloroplast. During the night, however, sucrose is depleted, resulting
in considerable decline in ADPG production by SuSy. In
addition, AGP activity will also decline because of both
redox inactivation of the enzyme (Hendriks et al. 2003) and the
low 3-phosphoglycerate (3PGA)/Pi ratio (Kleczkowski 1999,
Kleczkowski 2000, Crevillén et al. 2003).
The ‘alternative’ view is highly compatible with most, if
not all, starch mutant and transgenic plants described to date.
For example, the ‘alternative view’ predicts that in the ‘low
starch’ phenotype resulting from deficiencies in either pPGM
or AGP (Caspar et al. 1985, Lin et al. 1988, Müller-Röber et al.
1992), the recovery towards starch biosynthesis of the glucose
units derived from the starch breakdown will also be deficient,
resulting in a parallel decline of starch accumulation (BarojaFernández et al. 2001, Baroja-Fernández et al. 2004, BarojaFernández et al. 2005). In this respect, it is worth mentioning
that both leaves and heterotrophic organs totally lacking pPGM
accumulate readily detectable amounts of starch (Saether and
1367
Iversen 1991, Harrison et al. 1998), which indicates that ADPG
sources other than AGP occur in plants.
Leaves with a ‘high starch/sucrose’ balance resulting
when (i) export of triose phosphates from the chloroplast is
constrained (Riesmeier et al. 1993, Heineke et al. 1994); (ii)
either cytosolic phosphoglucose isomerase or fructose bisphosphatase activities are low (Neuhaus et al. 1989, Zrenner et al.
1996, Strand et al. 2000); and (iii) fructose-2,6-bisphosphate
levels are high (Scott and Kruger 1995), are characterized by a
high 3PGA/Pi ratio as compared with wild-type (WT) leaves.
A high 3PGA/Pi ratio activates AGP. Under those circumstances, the ‘alternative’ view predicts that the recovery
towards starch biosynthesis of the glucose units derived from
the starch breakdown will also be enhanced, resulting in a parallel increase of the starch/sucrose balance. We must emphasize
that a high 3PGA/Pi ratio has been shown to stimulate leaf
starch synthesis in the dark (Scott and Kruger 1995).
The ‘alternative’ model of starch biosynthesis in leaves
predicts that both sucrose and starch biosynthetic pathways are
tightly connected by means of ADPG-producing SuSy. In this
respect, the occurrence of UDP-glucose pyrophosphorylase
(UGP)-, cytosolic PGM- and sucrose phosphate synthase
(SPS)-deficient plants is consistent with the ‘alternative’ view
(but not with the ‘classical’ one), since these plants are characterized by containing low levels of both sucrose and starch in
their leaves compared with WT leaves (Strand et al. 2000,
Fernie et al. 2002, Kleczkowski et al. 2004). It is worth noting
that leaves from the latter two types of plants have normal
3PGA/Pi ratios compared with WT leaves.
The ‘alternative’ view postulates that the Calvin cycle is
not directly connected to the starch biosynthetic pathway by
means of plastid PGI (pPGI). In fact, Calvin cycle intermediates such as erythrose-4-phosphate (E4P) and 3PGA are
potent inhibitors of pPGI (Kelly and Latzko 1980, Dietz 1985,
Backhausen et al. 1997), the stromal concentrations of these
compounds in the illuminated chloroplast being much higher
than their Ki values for pPGI (Bassham and Krause 1969, Dietz
1985). Furthermore, the stromal glucose-6-phosphate (G6P)/
fructose-6-phosphate ratio is quite low and displaced from
equilibrium in the illuminated leaf (Dietz 1985), indicating that
pPGI is strongly inhibited under conditions of active CO2 fixation and starch production. The presumption that the Calvin
cycle is not connected to starch biosynthesis by means of pPGI
does not conflict with the occurrence of starch-deficient pPGI
mutants (Jones et al. 1986, Kruckeberg et al. 1989, Yu et al.
2000) since leaves from these plants have a reduced photosynthetic capacity that, in addition to reducing starch biosynthesis, causes a reduced growth phenotype.
Using transgenic plants expressing ADP sugar pyrophosphatase (ASPP) from Escherichia coli (Moreno-Bruna et
al. 2001) in either the cytosol or chloroplast, we recently provided strong evidence indicating that most of ADPG linked to
starch biosynthesis accumulates outside the chloroplast, presumably in the cytosol (Baroja-Fernández et al. 2004). Further-
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SuSy controls starch production in leaves
more, we also demonstrated that extraplastidial ADPG is
intimately linked to starch biosynthesis. Our results strongly
indicated that ADPG cannot accumulate in the chloroplast,
most probably because it is transferred rapidly to starch, and/or
because it spontaneously hydrolyzes under pH conditions
occurring in the illuminated chloroplast. The implications of
these findings evidently raised the question as to which of
the two enzymes known to synthesize ADPG, i.e. AGP and
SuSy, is the predominant source of cytosolic ADPG accumulating in leaves.
To address this question, in this work we measured the
ADPG content in starch-deficient Arabidopsis thaliana and
potato (Solanum tuberosum L.) plants with reduced pPGM,
AGP or SuSy (Caspar et al. 1985, Lin et al. 1988, MüllerRöber et al. 1992, Zrenner et al. 1995). Furthermore, we characterized both SuSy-overexpressing and -antisensed leaves, but
which contain normal pPGM and AGP activities.
The rationale behind this approach was that if ADPG
accumulating in the photosynthetically active cell is exclusively produced in the chloroplast, plants with reduced activities of either pPGM or AGP will not accumulate ADPG. In
addition, ADPG and starch levels in both SuSy-overexpressing
and -antisensed leaves will be normal compared with WT
leaves. Conversely, if ADPG accumulating in the cytosol is
produced by SuSy, ADPG levels in starch-deficient plants with
reduced plastid gluconeogenic enzymes will be normal compared with WT leaves. Furthermore, both ADPG and starch
levels in SuSy-overexpressing leaves will be higher than in WT
leaves, whereas SuSy-antisensed leaves will accumulate low
amounts of both ADPG and transitory starch.
The results presented in this communication undoubtedly
recognize the essential participation of SuSy in the production
of ADPG destined for the transitory starch biosynthetic process in leaves.
Results and Discussion
ADPG extraction and measurement
ADPG was extracted and measured by HPLC as described
in Materials and Methods. To ensure the effectiveness of the
ADPG extraction method, we added known amounts of commercially available ADPG to leaves immediately before
homogenization. The added amounts of ADPG were comparable with values previously reported for leaves (Smith et al.
1990, Baroja-Fernández et al. 2004), and their recoveries were
98 and 95% for potato and Arabidopsis leaves, respectively.
We subsequently checked the reliability of the two distinct chromatographic methods of ADPG detection and measurement employed in this work by adding known amounts of
ADPG to leaf extracts previously digested with purified ASPP
(Moreno-Bruna et al. 2001). ASPP is a nucleotide-sugar hydrolase that specifically cleaves ADP-sugars (Bessman et al. 1996,
Moreno-Bruna et al. 2001). Fig. 1A and B demonstrates that
Fig. 1 ADPG measurement in 100 µl potato leaf extracts (A) without and (B–H) with an ASPP digestion step prior to analysis by HPLC
with a Waters and Associates system fitted to a Partisil-10-SAX column. In (B–H), once the ASPP reactions were stopped, the indicated
amounts of commercially available ADPG were added to the extracts
and then analyzed by HPLC. Nucleotide elution times are: AMP,
3.2 min; UMP, 8.9 min; ADPG, 15.5 min; ADPribose, 18.3 min;
UDPglucose, 21.8 min; ADP, 31.8 min; ATP, 35 min; UTP, 36.1 min.
AU, absorbance units of A260. An equivalent type of analysis was performed using HPLC with pulsed amperometric detection on a DX-500
system fitted to a CarboPac PA10 column (not shown).
ASPP digestion exclusively and totally removed a substance
that eluted at the level of ADPG. This substance was collected
and hydrolyzed with ASPP. The products of the enzymatic
reaction were subjected to ion chromatography and proved to
SuSy controls starch production in leaves
1369
Fig. 2 ADPG calibration curve for potato leaf extracts. The curve
was obtained from chromatographic analyses illustrated in Fig. 1 using
100 µl of leaf extracts. An equivalent curve was also obtained for
Arabidopsis leaves (not shown). AU, absorbance units of A260
be AMP and glucose-1-phosphate (G1P) (not shown), the overall results confirming the correct identification of ADPG.
As illustrated in Fig. 1B–H, this approach allowed us to
produce accurate calibration curves for ADPG content in both
Arabidopsis and potato leaves (Fig. 2). Using these calibration
curves, we observed that ADPG levels in both Arabidopsis and
potato leaves ranged between 0.25 and 0.35 nmol g–1 FW
(0.35–0.40 nmol mg–1 Chl) (Fig. 1A). These values were comparable with those previously reported for WT pea leaves
(0.6 nmol mg–1 Chl) (Smith et al. 1990).
AGP- and pPGM-deficient leaves have low levels of starch but
normal levels of ADPG
The ‘classical’ view of starch biosynthesis, according to
which AGP is the sole enzyme producing ADPG linked to
starch biosynthesis, predicts that ADPG levels in starch-deficient leaves with reduced activities of either AGP or pPGM
will be low when compared with WT leaves. Conversely, the
‘alternative’ starch biosynthetic model, according to which
SuSy catalyzes the de novo production of ADPG and AGP
plays a role in the scavenging of the glucose units derived
from the starch breakdown, predicts that intracellular levels of
ADPG in leaves with low activities of chloroplast gluconeogenic enzymes will be normal when compared with those of
WT plants (Baroja-Fernández et al. 2001, Pozueta-Romero et
al. 2003, Baroja-Fernández et al. 2004, Baroja-Fernández et al.
2005).
As a first step to distinguish which of these two metabolic
models may entail a predominant role in starch biosynthesis in
source leaves, we measured the ADPG content in leaves of
TC7 and TL25 Arabidopsis plants lacking pPGM and AGP,
respectively (Caspar et al. 1985, Lin et al. 1988). The starchdeficient phenotype of these plants has been used as a crucially
important argument to support the ‘classical’ starch biosynthetic model (Okita 1992). As illustrated in Fig. 3A, meas-
Fig. 3 Starch-deficient AGP and pPGM mutant Arabidopsis leaves
contain normal ADPG levels. Starch (A) and ADPG (B) levels in fully
expanded source leaves of WT, TL25 (Lin et al. 1988) and TC7
(Caspar et al. 1985) plants. Analyses were conducted using fully
expanded leaves after 9 h illumination (300 µmol photons s–1 m–2).
ADPG levels were measured by the two chromatographic methods
described in Materials and Methods.
urements of starch content in Arabidopsis plants confirmed
previous studies showing that both TL25 and TC7 plants contain low starch levels when compared with WT plants (Caspar
et al. 1985, Lin et al. 1988). Most importantly however, this
reduction in starch content was not accompanied by any measurable reduction in intracellular ADPG levels, which ranged
between 0.25 and 0.35 nmol g–1 FW (Fig. 3B).
Normal levels of ADPG in starch-deficient plants with
low AGP activity were not limited to Arabidopsis plants. As
illustrated in Fig. 4A, and in conformity with the results of
Müller-Röber et al. (1992), leaves of transgenic potato plants
having low AGP activity accumulate less starch than leaves of
WT plants. In clear contrast, however, ADPG levels in the
transgenic lines were shown to be nearly identical to those
found in WT plants (Fig. 4B).
Collectively, these results strongly indicate that AGP is
not the predominant source of ADPG accumulating in photosynthetic cells.
Changes in SuSy expression are accompanied by changes in
both ADPG content and rate of transitory starch biosynthesis
Compared with reserve starch-storing organs, SuSy activity and starch content are considerably lower in source leaves.
This enzyme is highly regulated (Huber et al. 1996, Nakai et al.
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SuSy controls starch production in leaves
Fig. 5 ADPG-producing SuSy activities in WT and both SuSy-antisensed and -overexpressing source leaves (both 35S-SuSy-NOS and
RBCS-SuSy-NOS). The results are the mean ± SE of 10 independent
plants per line.
Fig. 4 Starch-deficient AGP-antisensed potato leaves contain normal ADPG levels. Starch (A) and ADPG (B) levels in fully expanded
(fifth) source leaves from 5-week-old WT plants and plants with
reduced AGP activity (AGP62 and AGP85, Müller-Röber et al. 1992).
Analyses were conducted using fully expanded leaves after 9 h illumination (300 µmol photons s–1 m–2). ADPG levels were measured by the
two chromatographic methods described in Materials and Methods.
1998, Asano et al. 2002) and is involved in the sucrose–starch
conversion process in sink organs (Zrenner et al. 1995, BarojaFernández et al. 2003). SuSy is expressed in the mesophyll
cells (Fu et al. 1995, Wang et al. 1999), though its physiological role still remains unknown.
The ‘alternative’ model of starch biosynthesis assumes
that SuSy catalyzes the production of a sizable pool of ADPG
linked to starch biosynthesis. Whether or not SuSy is responsible for the ADPG accumulating in leaves was tested by characterizing both SuSy-overexpressing and -antisensed leaves. The
rationale behind this approach was that if this sucrolytic
enzyme catalyzes the production of cytosolic ADPG linked to
transitory starch biosynthesis in mesophyll cells, varying SuSy
activities will be accompanied by changes in both ADPG and
starch levels. We therefore proceeded to generate and characterize genetically modified potato plants expressing SuSy
either constitutively or in the illuminated leaves [caulifower
mosaic virus 35S promoter (35S)-SuSy-nopaline synthase
(NOS) and ribulose 1,5-bisphosphate carboxylase small subunit (RBCS)-SuSy-NOS, respectively]. In addition, we characterized the leaves of S-112 and S-129 SuSy-antisensed potato
plants (Zrenner et al. 1995). As illustrated in Fig. 5, ADPGproducing SuSy activities in the WT leaves were approxi-
Fig. 6 ADPG levels in fully expanded (fifth) source leaves from 6week-old WT and both SuSy-antisensed and -overexpressing plants
(both 35S-SuSy-NOS and RBCS-SuSy-NOS). Leaf samples were
taken and quenched in liquid nitrogen 9 h after the beginning of the
light period. The results are the mean ± SE of 10 independent plants
per line. ADPG levels were measured by the two chromatographic
methods described in Materials and Methods.
mately 15 mU g–1 FW, whereas SuSy activities in the leaves of
the SuSy-overexpressing and -antisensed transgenic lines were
35–90 and 8–10 mU g–1 FW, respectively.
Leaves of both SuSy-overexpressing and -antisensed
plants were analyzed further for their ADPG and starch contents. As it is our experience that biochemical analyses are subject to considerable variation, we analyzed 10 plants per line to
ensure the attainment of reliable data. Importantly, analyses of
the ADPG content in leaves of both WT and transgenic plants
(Fig. 6, Supplementary Fig. 2) revealed a pattern similar to that
of ADPG-producing SuSy activities shown in Fig. 5. Most
important was the fact that starch content patterns in both WT
and transgenic leaves paralleled that of ADPG-producing SuSy
activities shown in Fig. 5 (Fig. 7).
During the day, sucrose accumulates in the source leaf
(Stitt et al. 1983, Stitt et al. 1984, Gerhardt et al. 1987,
SuSy controls starch production in leaves
1371
Fig. 7 Starch levels in fully expanded (fifth) source leaves from 6week-old WT and both SuSy-antisensed and -overexpressing plants
(both 35S-SuSy-NOS and RBCS-SuSy-NOS). Leaf samples were
taken and quenched in liquid nitrogen 9 h after the beginning of the
light period. The results are the mean ± SE of 10 independent plants
per line.
Draborg et al. 2001, Smith et al. 2004) followed by a decline
during the night. If SuSy plays an important role in the starch
biosynthetic process by channeling ADPG from sucrose to
starch, it can be presumed that both ADPG and starch accumulation patterns will be determined by both sucrose availability
and SuSy activity. We therefore compared the ADPG and
starch accumulation patterns between WT, 35S-SuSy-NOS
(line 5) and RBCS-SuSy-NOS (line 4) leaves throughout the
photoperiod.
As illustrated in Fig. 8A, these analyses revealed that the
highest rate of transient starch accumulation in WT plants
(7 nmol of glucose transferred to starch g–1 FW min) can be
well accounted for by the ADPG-producing SuSy activity
measured in vitro (∼15 mU g–1 FW, cf. Fig. 5). Furthermore,
these analyses revealed that, essentially similar to the case of
starch and sucrose, ADPG transiently accumulates in the photosynthetic cell during the light period (Fig. 8B) followed by a
decline during the subsequent night. Most important, however,
was the fact that the rates of both ADPG and transitory starch
accumulation in both 35S-SuSy-NOS and RBCS-SuSy-NOS
leaves were considerably higher than that of WT leaves. The
increase in the rates of both ADPG and transitory starch accumulation in SuSy-enhanced lines during the day was mirrored
by a similar increase in the rates of both ADPG and starch
depletion during the night.
Collectively, these results are a strong indication that
SuSy plays a dominant role in the transitory starch biosynthetic process in source leaves. Furthermore, these results
also indicate that (i) sucrose availability is a limiting step in the
transitory starch biosynthetic process and (ii) SuSy catalyzes
the production of cytosolic ADPG accumulating in the leaf.
Fig. 8 Time course analysis of (A) starch and (B) ADPG content in
leaves of 5-week-old WT (circle), 35S-SuSy (line 5, square) and RBCSSuSy-NOS (line 4, triangle) plants grown in chambers under an 8 h/
16 h light (300 µmol photons s–1 m–2, 20°C)/dark (20°C) regime. At
each time point, samples were taken from mature source leaves and the
data represent the means ± SE of measurements on 10 plants per line.
SuSy overexpression is not accompanied by pleiotropic changes
in enzymes directly connected to starch and sucrose metabolism
In order to verify that increases in ADPG and starch contents in SuSy-overexpressing leaves were not due to changes in
the activities of enzymes other than SuSy, we measured the
maximum catalytic activities of a range of enzymes closely
connected to starch and sucrose metabolism. These analyses
revealed no significant changes in AGP, alkaline pyrophosphatase, UGP, SPS, acid invertase, total starch synthase,
starch phosphorylase and total amylolytic activity (Table 1,
Supplementary Table 1).
SuSy overexpression does not affect the photosynthetic capacity of the plant
Given that both ADPG and starch levels are higher in the
leaves of SuSy-overexpressing plants than in the WT plants,
we proceeded to investigate whether transgenic plants dis-
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Table 1
SuSy controls starch production in leaves
Enzyme activities (mU g–1 FW) in WT and 35S-SuSy-NOS potato leaves
WT
6
35S-SuSy-NOS
12
3
5
4
7
AGP
144 ± 6 149 ± 5 140 ± 3
148 ± 7 124 ± 6 122 ± 7 137 ± 7
UGP
107 ± 3 113 ± 5 108 ± 5
116 ± 4 125 ± 5 103 ± 4 116 ± 8
Acid invertase
118 ± 9 137 ± 11 104 ± 8
155 ± 9 125 ± 7 120 ± 9 146 ± 12
SPS
2,855 ± 220 3,279 ± 245 3,235 ± 362 3,211 ± 264 2,875 ± 303 2,542 ± 231 3,611 ± 316
Alkaline
756 ± 61 879 ± 80 880 ± 70
717 ± 65 786 ± 72 762 ± 74 762 ± 75
pyrophosphatase
Amylolytic activity
114 ± 9 135 ± 10 119 ± 5
124 ± 6 125 ± 6 143 ± 8 133 ± 10
Total starch synthase
6.1 ± 1
6.6 ± 0.5
5.9 ± 0.9
7.0 ± 1
6.7 ± 0.7
5.9 ± 0.8
7.9 ± 0.9
Starch phosphorylase 39 ± 6
39 ± 7
33 ± 6
49 ± 12
35 ± 7
43 ± 9
44 ± 8
The activities were determined in samples from source leaves of 6-week-old plants grown in chambers at ambient CO2 conditions, 20°C and at an
irradiance of 300 µmol photons s–1 m–2. Leaf samples were taken and quenched in liquid nitrogen 7 h after the beginning of the light period. The
results are the mean ± SE of extracts from 10 independent plants per line.
Table 2 Photosynthetic parameters of whole leaves attached to 6-week-old WT and SuSy-expressing plants grown in the same
conditions as in Table 1
WT
6
35S-SuSy-NOS
12
3
5
4
7
O2 evolution
9.1 ± 1.1 8.5 ± 1.2 9.1 ± 1.1 9.0 ± 1.0 9.6 ± 0.5 8.9 ± 1.3 9.0 ± 1.1
(µmol m–2 s–1)
Transpiration
1.8 ± 0.4 2.3 ± 0.3 2.0 ± 0.3 1.9 ± 0.5 2.2 ± 0.4 1.8 ± 0.2 1.8 ± 0.3
(mmol m–2 s–1)
Stomatal conductance of
170 ± 41 200
± 25 180
± 20 190
± 35 180 ± 30 130 ± 20 150 ± 40
CO2 (mmol m–2 s–1)
Substomatal CO2 (ppm) 372
± 53 374
± 19 379
± 18 362
± 39 355 ± 39 342 ± 21 366 ± 30
Chlorophyll (mg/g FW)
0.80 ± 0.06 0.88 ± 0.07 0.80 ± 0.06 0.77 ± 0.01 0.82 ± 0.01 0.75 ± 0.08 0.80 ± 0.02
The results are the mean ± SE of extracts from 10 independent plants per line.
played increased photosynthetic activity. Towards this end,
photosynthetic parameters of whole attached leaves in SuSyexpressing and control plants were compared at light intensities in which the plants were grown (300 µmol photon m–2 s–1)
and at ambient CO2 (350 ppm). Our results revealed no significant changes in the rates of O2 production, transpiration, CO2
stomatal conductance and substomatal CO2 concentration
(Table 2, Supplementary Table 2). Furthermore, there were no
significant differences between chlorophyll contents of the
SuSy lines and WT plants. Therefore, increase of both ADPG
and starch contents in the leaves of SuSy-overexpressing plants
cannot be ascribed to increased photosynthetic capacities.
Metabolic profile of SuSy-overexpressing leaves
Leaves from both 35S-SuSy-NOS and RBCS-SuSy-NOS
plants were shown to accumulate normal levels of glucose,
fructose, 3PGA and Pi when compared with control plants
(Table 3, Supplementary Table 3). In contrast, G1P levels in the
SuSy-overexpressing leaves decreased up to 2-fold when compared with control plants. This observation is highly significant given previous observations that reducing the levels of
cytosolic ADPG is accompanied by an increase of total G1P
levels (Baroja-Fernández et al. 2004), strongly indicating that
G1P and cytosolic ADPG pools are closely connected. Levels
of G6P, an allosteric effector of SPS (Doehlert and Huber 1983,
Huber and Huber 1996), were 30–40% higher in the SuSyoverexpressing leaves than in the WT leaves. As a possible
consequence of the stimulatory effect of high G6P levels on
SPS, SuSy-overexpressing leaves were shown to accumulate
30–40% more sucrose than WT leaves.
Additional remarks
In source leaves, SuSy plays a pivotal role in basic aspects
of plant survival such as supplying energy for the sucrose loading and unloading process (Martin et al. 1993, Nolte and Koch
1993). Taking into account all the limitations that are inherent
in basing conclusions on genetically engineered and mutant
plants, results from the work presented in this communication
show that SuSy exerts a strong control on the starch biosynthetic process in source leaves. Furthermore, evidence is
provided that SuSy, but not AGP, catalyzes the production of
ADPG accumulating in source leaves (Fig. 3, 4).
SuSy controls starch production in leaves
Table 3
1373
Metabolite levels (given in nmol g–1 FW) in control (WT) and 35S-SuSy-NOS source leaves
Control WT
6
5
35S-SuSy-NOS
12
3
4
7
Glucose
848 ± 31
922 ± 29
860 ± 30
933 ± 29
881 ± 56
895 ± 32
871 ± 60
Fructose
996 ± 43
1,035 ± 67
1,094 ± 17
1,022 ± 10
1,067 ± 58
1,078 ± 63
817 ± 41
Sucrose
1,012 ± 27
1,529 ± 48
1,402 ± 68
1,642 ± 58
1,307 ± 35
1,317 ± 35
1,391 ± 70
G6P
244 ± 28
329 ± 15
320 ± 25
291 ± 27
355 ± 23
298 ± 12
315 ± 9.8
G1P
22.7 ± 1.9
15.5 ± 2.1
10.3 ± 1.1
9.9 ± 1.2
9.5 ± 1.5
15.2 ± 1.9
11.4 ± 1.8
3PGA
324 ± 42
354 ± 40
332 ± 21
362 ± 12
386 ± 27
311 ± 47
359 ± 27
Pi
40.6 ± 2.8
36.9 ± 1.2
39.9 ± 2.1
43.7 ± 3.1
39.8 ± 0.5
41.6 ± 1.7
42.3 ± 0.9
3PGA/Pi ratio
8.1 ± 1.0
9.5 ± 1.3
8.3 ± 1.0
8.2 ± 1.0
9.6 ± 0.9
7.5 ± 1.1
8.3 ± 0.7
Leaf samples were taken from 6-week-old plants grown in chambers at ambient CO2 conditions, 20°C and at an irradiance of 300 µmol photons s–1
m–2. Leaves were taken and quenched in liquid nitrogen 7 h after the beginning of the light period. The results are the mean ± SE of extracts from
10 independent plants per line. Values that are significantly different from the control plants are marked in bold.
SuSy has the capacity of producing ADPG from sucrose
(Delmer 1972, Porchia et al. 1999, Baroja-Fernández et al.
2003). It is therefore highly conceivable that the high levels of
ADPG occurring in SuSy-overexpressing leaves (Fig. 6) are
due to the direct conversion of sucrose into ADPG by means of
SuSy. An alternative explanation for our results is that, essentially identical to the ‘classical’ view on sucrose–starch conversion in heterotrophic tissues (Neuhaus and Emes 2000), SuSy
converts sucrose into UDPG, which is then stepwise transformed to G1P and ADPG by the coupled reactions of UGP
and AGP. This possibility, however, is highly unlikely because
(i) UGP in leaves functions primarily in the production of
UDPG rather than in its degradation and (ii) G1P levels in
SuSy-overexpressing leaves are lower than in WT leaves
(Table 3, Supplementary Table 3). Furthermore, this presumption indirectly implies the operation of a G1P transport machinery which, as shown by Quick et al. (1995), does not occur in
the envelope membranes of chloroplasts. Also, and contrary to
our recent findings showing the extraplastidial localization of
ADPG (Baroja-Fernández et al. 2004), this possibility would
imply that ADPG in the SuSy-overexpressing leaves would
accumulate in the chloroplast.
A second alternative explanation for the occurrence of
high levels of both ADPG and starch in the SuSy-overexpressing leaves is that high SuSy activity alters the exchange
of Pi and triose phosphates between the cytosol and the chloroplast. This situation will lead to a high 3PGA to Pi ratio in the
chloroplast, thus activating AGP. This possibility, however, is
highly unlikely since the 3PGA to Pi ratio in SuSy-overexpressing leaves is normal when compared with WT leaves
(Table 3, Supplementary Table 3).
The overall data thus strongly indicate that the starch biosynthetic process occurring in the chloroplast is tightly linked
to the cytosolic sucrose metabolism by means of ADPG-producing SuSy activity.
Materials and Methods
Plants, bacterial strains and media
This work was carried out using plants regenerated from WT
potato plants (S. tuberosum L. cv Desirée), and plants transformed
with either anti-AGP-1 (Müller-Röber et al. 1992, lines 62 and 85),
anti-SuSy (Zrenner et al. 1995, lines S-112 and S-129), 35S-SuSyNOS terminator or RBCS-SuSy-NOS (see below). Columbia WT,
TL25 and TC7 Arabidopsis plants were obtained from the Nottingham
Arabidopsis Stock Centre.
Unless otherwise indicated, plants were grown at ambient CO2
(350 ppm) for 4–6 weeks in growth chambers with a 16 h light
(300 µmol photons s–1 m–2) photoperiod and at a constant temperature
of 20°C. For biochemical analyses, fully expanded source leaves were
harvested at the indicated times of illumination, immediately quenched
in liquid nitrogen and stored at –80°C before use.
Agrobacterium tumefaciens cells, strain C58C1:GV2260, transformed with either pBIN35S-SuSy-NOS or pBINRBCS-SuSy-NOS
(see below) were grown in yeast extract broth medium, with the appropriate selection using standard techniques.
Production of SuSy-overexpressing potato plants
For the production of 35S-SuSy-NOS plants, a complete Sus4
cDNA (EMBL accession no. AJ537575) was amplified by PCR using
the primers 5′-GTCTCGATAAGCTTCTGCCATGGCTGAACGTGTTTTGACTCG-3′ (forward) and 5′-CCGGTGGATCCCCCCTTCATTCACTCAGCAGCCAATGGAAC-3′ (reverse) and a potato cDNA
library. The approximately 2,400 bp PCR product obtained was cloned
into pBluescript SK- to produce pSK-SuSy (Supplementary Fig. 3).
The NcoI–BamHI fragment of pSK-SuSy was ligated with the corresponding restriction sites of p35S-NOS (Baroja-Fernández et al. 2004)
to produce p35S-SuSy-NOS. This construct was digested successively
with HindIII, T4 DNA polymerase and NotI. The fragment thus released
was cloned into the pBIN20 plant expression vector (Hennegan and
Danna 1998) previously digested successively with the enzymes HindIII, T4 DNA polymerase and EcoRI to produce pBIN35S-SuSy-NOS.
For production of SuSy-overexpressing leaves, approximately
1,000 bp of the 5′-upstream region of the tobacco RBCS gene (EMBL
accession no. X02353) was amplified by PCR that confers expression
to illuminated chloroplast-containing cells (Barnes et al. 1994).
Towards this end, we used tobacco genomic DNA and the following
primers: (forward) 5′-AAGCTTGTGGGAACGAGATAAGGGC-3′;
(reverse) 5′-CCATGGTTAATTACACTTAGACAGAAAGATTTCTC-
1374
SuSy controls starch production in leaves
TTTCTCTCTTTTCCTTTAACTTCC-3′. The fragment thus obtained
was cloned into pGEMT-easy vector (Promega) to produce pGEMTRBCSprom (Supplementary Fig. 4). This construct was digested with
HindIII and NcoI and the fragment released was ligated with the corresponding restriction sites of p35S-SuSy-NOS to produce pRBCS-SuSyNOS. This final construct was digested successively with HindIII, T4
DNA polymerase and NotI. The resulting fragment was cloned into
pBIN20 previously digested successively with the enzymes HindIII,
T4 DNA polymerase and EcoRI to produce pBINRBCS-SuSy-NOS.
Transfer of constructs into A. tumefaciens was carried out by
electroporation. Subsequent transformation of potato plants was conducted as described by Baroja-Fernández et al. (2004). Transgenic
plants were selected on kanamycin-containing medium. The presence
of the recombinant Sus4 was confirmed by Southern blot analyses
using radiolabeled Sus4 cDNA as probe. A second screening of the
transgenic lines was performed by SuSy activity. Six independent lines
per construct were selected and used for all subsequent analyses
described in this report.
Enzyme assays
All enzymatic reactions were carried out at 37°C. Harvested
leaves were immediately freeze-clamped and ground to a fine powder
in liquid nitrogen with a pestle and mortar. To assay enzyme activity, 1
g of the frozen powder was resuspended at 4°C in 5 ml of 100 mM
HEPES (pH 7.5), 2 mM EDTA and 5 mM dithiothreitol. The suspension was desalted and assayed for enzymatic activities. We checked
that this procedure did not result in loss of enzymatic activity as evidenced by comparing the activity in extracts prepared from the frozen
powder and extracts prepared by homogenizing fresh tissue in extraction medium. AGP, acid invertase, UGP, alkaline pyrophosphatase,
total starch synthase, starch phosphorylase, total amylolytic and SPS
activities were assayed as described by Baroja-Fernández et al. (2004).
Measurements of SuSy were performed in the direction of ADPG synthesis. The assay mixture contained 50 mM HEPES (pH 7.0), 1 mM
MgCl2, 15 mM KCl, 250 mM sucrose and 2 mM ADP. After 5 min at
37°C, reactions were stopped by boiling the assay mixture for 1 min.
ADPG was measured by HPLC on a Waters Associate’s system fitted
with a Partisil-10-SAX column as described by Sweetlove et al.
(1996). We define 1 U of enzyme activity as the amount of enzyme
that catalyzes the production of 1 µmol of product min–1.
Determination of soluble sugars
Fully expanded leaves were harvested and immediately ground
to a fine powder in liquid nitrogen with a pestle and mortar. A 0.5 g
aliquot of the frozen powdered tissue was resuspended in 0.4 ml of
1.4 M HClO4, left at 4°C for 2 h and centrifuged at 10,000×g for
5 min. The supernatant was neutralized with K2CO3 and centrifuged at
10,000×g. ADPG content in the supernatant was determined by using
either one of the following methods: (i) by HPLC on a system obtained
from P.E. Waters and Associates fitted with a Partisil-10-SAX column
as described by Sweetlove et al. (1996); or (ii) by HPLC with pulsed
amperometric detection on a DX-500 system (Dionex) fitted to a CarboPac PA10 column.
To confirm further that measurements of ADPG were correct,
ADPG eluted from either the Partisil-10-SAX or CarboPac PA10 columns was enzymatically hydrolyzed with purified E. coli ASPP (see
below) and assayed for conversion into AMP and G1P.
Glucose, sucrose, fructose, G1P and G6P were determined by
HPLC with pulsed amperometric detection on a DX-500 system as
described by Baroja-Fernández et al. (2003). 3PGA and Pi were determined as described by Lytovchenko et al. (2002)
Production and purification of recombinant ASPP
pET-ASPP cells (Moreno-Bruna et al. 2001) were grown in
100 ml of liquid LB medium to an absorbance at 600 nm of about 0.5
and then 0.3 mM isopropyl-β-D-thiogalactopyranoside (IPTG) was
added. After 3 h, cells were centrifuged at 6,000×g for 10 min. The
pelleted bacteria were resuspended in 4 ml of His-bind binding buffer
(Novagen), sonicated and centrifuged at 10,000×g for 10 min. The
supernatant thus obtained was subjected to His-bind chromatography
(Novagen). The eluted His-tagged ASPP was then rapidly desalted by
ultrafiltration on Centricon YM-10 (Amicon, Beford, MA, USA).
SDS–PAGE of the purified His-tagged ASPP preparation and subsequent stain revealed a single approximately 30 kDa band (not shown).
Analytical procedures
Starch in desalted plant extracts obtained by precipitation with
70% ethanol was measured by using an amyloglucosidase-based test
kit (Sigma-Aldrich Chemical Co, St Louis, MO, USA). Chlorophyll
was quantified according to the method of Wintermans and de Mots
(1965). Photosynthetic parameters of the first fully expanded (fifth)
leaves attached to the potato plants were determined under growth
conditions by means of an Lci portable photosynthesis system (ADC
BioScientfic Ltd., Hoddesdon, Herts, UK) in the same chamber where
the leaves were grown.
Supplementary material
Supplementary material mentioned in the article is available to
online subscribers at the journal website www.pcp.oupjournals.org.
Acknowledgments
This research was partially supported by the grants BIO200–
1080 and BIO2004–01922 from the Comisión Interministerial de
Ciencia y Tecnología and Fondo Europeo de Desarrollo Regional
(Spain). M.T.M.-Z. acknowledges the Spanish Ministry of Culture and
Education for a pre-doctoral felloship. A.M.V. expresses his gratitude
to the Spanish Ministry of Culture and Education for financial support. E.E. expresses his gratitude to the Spanish Ministry of Culture
and Education, to the Consejo Superior de Investigaciones Científicas
and to the Public University of Nafarroa for their support. We thank
Vanessa Rubio, Maria José Villafranca and Sorospen Barón Lacunza
(I.A.R.N., Spain) for expert technical support. We are indebted to Dr.
R. N. Trethewey (Metanomics GmbH, Berlin) who kindly made us a
gift of the anti-AGP1 and anti-SuSy plants. We are also indebted to
Axel Tiessen (Max Planck Institut, Golm, Germany), Eckehard
Neuhaus (University of Kaiserslautern, Germany) and Kay Denyer
(John Innes Centre, Norwich, UK) for critical discussions and readings of the manuscript.
References
Asano, T., Kunieda, N., Omura, Y., Ibe, H., Kawasaki, T., et al. (2002) Rice
SPK, a calmodulin-like domain protein kinase, is required for storage product accumulation during seed development: phosphorylation of sucrose
synthase is a possible factor. Plant Cell 14: 619–628.
Backhausen, J.E., Jöstingmeyer, P. and Scheibe, R. (1997) Competitive inhibition of spinach leaf phosphoglucose isomerase isoenzymes by erythrose 4phosphate. Plant Sci. 130: 121–131.
Barnes, S.A., Knight, J.S. and Gray, J.C. (1994) Alteration of the amount of the
chloroplast phosphate translocator in transgenic tobacco affects the distribution of assimilate between starch and sugar. Plant Physiol. 106: 1123–1129.
Baroja-Fernández, E., Muñoz, F.J., Akazawa, T. and Pozueta-Romero, J. (2001)
Reappraisal of the currently prevailing model of starch biosynthesis in photosynthetic tissues: A proposal involving the cytosolic production of ADP-glu-
SuSy controls starch production in leaves
cose by sucrose synthase and occurrence of cyclic turnover of starch in
chloroplast. Plant Cell Physiol. 42: 1311–1320.
Baroja-Fernández, E., Muñoz, F.J. and Pozueta-Romero, J. (2005) A response to
Neuhaus et al. Trends Plant Sci. 10: 156–159.
Baroja-Fernández, E., Muñoz, F.J., Saikusa, T., Rodríguez-López, M., Akazawa,
T. and Pozueta-Romero, J. (2003) Sucrose synthase catalyzes the de novo
production of ADPglucose linked to starch biosynthesis in heterotrophic tissues of plants. Plant Cell Physiol. 44: 500–509.
Baroja-Fernández, E., Muñoz, F.J., Zandueta-Criado, A., Morán-Zorzano, M.T.,
Viale, A.M., Alonso-Casajús, N. and Pozueta-Romero, J. (2004) Most of
ADPglucose linked to starch biosynthesis occurs outside the chloroplast in
source leaves. Proc. Natl Acad. Sci. USA 101, 13080–13085.
Bassham, J.A. and Krause, G.H. (1969) Free energy changes and metabolic regulation in steady-state photosynthetic carbon reduction. Biochim. Biophys.
Acta 189: 207–221.
Belanger, A.E. and Hatfull, G.F. (1999) Exponential-phase glycogen recycling
is essential for growth of Mycobacterium smegmatis. J. Bacteriol. 181: 6670–
6678.
Bessman, M.J., Fricks, D.N. and O’Handley, S.F. (1996) The MutT proteins or
‘Nudix’ hydrolases, a family of versatile, widely distributed, housecleaning
enzymes. J. Biol. Chem. 271: 25059–25062.
Caspar, T., Huber, S.C. and Somerville, C. (1985) Alterations in growth, photosynthesis, and respiration in a starchless mutant of Arabidopsis thaliana (L.)
deficient in chloroplast phosphoglucomutase activity. Plant Physiol. 79: 11–
17.
Crevillén, P., Ballicora, M.A., Mérida, A., Preiss, J. and Romero, J.M. (2003)
The different large subunit isoforms of Arabidopsis thaliana ADP-glucose
pyrophosphorylase confer distinct kinetic and regulatory properties to the heterotetrameric enzyme. J. Biol. Chem. 278: 28508–28515.
David, M., Petit, W.A., Laughlin, M.R., Shulman, R.G., King, J.E. and Barrett,
E.J. (1990) Simultaneous synthesis and degradation of rat liver glycogen. J.
Clin. Invest. 86: 612–617.
Delmer, D.P. (1972) The biosynthesis of sucrose and nucleoside diphosphate
glucoses in Phaseolus vulgaris. J. Biol. Chem. 247: 3822–3828.
Dietz, K.-J. (1985) A possible rate-limiting function of chloroplast hexosemonophosphate isomerase in starch synthesis of leaves. Biochim. Biophys. Acta
839: 240–248.
Doehlert, D.C. and Huber, S.C. (1983) Regulation of spinach leaf sucrose phosphate synthase by glucose-6-phosphate, inorganic phosphate, and pH. Plant
Physiol. 73: 989–994
Draborg, H., Villadsen, D. and Nielsen, T.H. (2001) Transgenic Arabidopsis
plants with decreased activity of fructose-6-phosphate, 2-kinase/fructose-2, 6bisphosphatase have altered carbon partitioning. Plant Physiol. 126: 750–758.
Fernie, A.R., Tauberger, E., Lytovchenko, A., Roessner, U., Willmitzer, L. and
Trethewey, R.N. (2002) Antisense repression of cytosolic phosphoglucomutase in potato (Solanum tuberosum) results in severe growth retardation,
reduction in tuber number and altered carbon metabolism. Planta 214: 510–
520.
Fox, T.C. and Geiger, D.R. (1984) Effects of decreased net carbon exchange on
carbohydrate metabolism in sugar beet source leaves. Plant Physiol. 76: 763–
768.
Fu, H., Kim, S.Y. and Park, W.D. (1995) A potato Sus3 sucrose synthase gene
contains a context-dependent 3′ element and a leader intron with both positive and negative tissue-specific effects. Plant Cell 7: 1395–1403.
Gaudet, G., Forano, E., Dauphin, G. and Delort, A.-M. (1992) Futile cycling of
glycogen in Fibrobacter succinogenes as shown by in situ 1H-NMR and 13CNMR investigation. Eur. J. Biochem. 207: 155–162.
Gerhardt, R., Stitt, M. and Heldt, H.W. (1987) Subcellular metabolite levels in
spinach leaves. Plant Physiol. 83: 399–407.
Guedon, E., Desvaux, M. and Petitdemange, H. (2000) Kinetic analysis of
Clostridium cellulolyticum carbohydrate metabolism: importance of glucose
1-phosphate and glucose 6-phosphate branch points for distribution of carbon fluxes inside and outside cells as revealed by steady-state continuous culture. J. Bacteriol. 182: 2010–2017.
Harrison, C.J., Hedley, C.L. and Wang, T.L. (1998) Evidence that the rug3 locus
of pea (Pisum sativum L.) encodes plastidial phosphoglucomutase confirms
that the imported substrate for starch synthesis in pea amyloplasts is glucose6-phosphate. Plant J. 13: 753–762.
Heineke, D., Kruse, A., Flügge, U.I., Frommer, W.B., Riesmeier, J.W.,
Willmitzer, L. and Heldt, H.W. (1994) Effect of antisense repression of the
1375
chloroplast triose-phosphate translocator on photosynthetic metabolism in
transgenic potato plants. Planta 193: 174–180.
Hendriks, J.H.M., Kolbe, A., Gibon, Y., Stitt, M. and Geigenberger, P. (2003)
ADP-glucose pyrophosphorylase is activated by posttranslational redox-modification in response to light and to sugars in leaves of Arabidopsis and other
plant species. Plant Physiol. 133: 838–849
Hennegan, K.P. and Danna, K.J. (1998) PBIN20: an improved binary vector for
Agrobacterium-mediated transformation. Plant Mol. Biol. Rep. 16: 129–131.
Huber, S.C. and Huber, J.L. (1996) Role and regulation of sucrose-phosphate
synthase in higher plants. Annu. Rev. Plant Physiol. Plant Mol. Biol. 47: 431–
443.
Huber, S.C., Huber, J.L., Liao, P.-C., Gage, D.A., McMichael, R.W., Chourey,
P.S., Hannah, L.C. and Koch, K. (1996) Phosphorylation of serine-15 of
maize leaf sucrose synthase. Plant Physiol. 112: 793–802.
Jones, T.W.A., Gottlieb, L.D. and Pichersky, E. (1986) Reduced enzyme activity and starch level in an induced mutant of chloroplast phosphoglucose
isomerase. Plant Physiol. 81: 367–371.
Kelly, G.J. and Latzko, E. (1980) Oat leaf phosphoglucose isomerase: competitive inhibition by erythrose-4-phosphate. Photosynth. Res. 1: 181–187.
Kleczkowski, L.A. (1999) A phosphoglycerate to inorganic phosphate ratio is
the major factor in controlling starch levels in chloroplasts via ADP-glucose
pyrophosphorylase regulation. FEBS Lett. 448: 153–156.
Kleczkowski, L.A. (2000) Is leaf ADP-glucose pyrophosphorylase an allosteric
enzyme. Biochim. Biophys. Acta 1476: 103–108.
Kleczkowski, L.A., Geisler, M., Ciereszko, I. and Johansson, H. (2004) UDPglucose pyrophosphorylase. An old protein with new tricks. Plant Physiol.
134: 912–918
Kruckeberg, L., Neuhaus, H.E., Feil, R., Gottlieb, L.D. and Stitt, M. (1989)
Decreased-activity mutants of phosphoglucose isomerase in the cytosol and
chloroplast of Clarkia xantiana. Biochem. J. 261: 457–467.
Lin, T.-P., Caspar, T., Somerville, C.R. and Preiss, J. (1988) Isolation and characterization of a starchless mutant of Arabidopsis thaliana (L.) Heynh lacking
ADPglucose pyrophosphorylase activity. Plant Physiol. 88: 1131–1135.
Lloyd, J.R., Kossmann, J. and Ritte, G. (2005) Leaf starch degradation comes
out of the shadows. Trends Plant Sci. 10: 130–137.
Lozovaya, V.V., Zabotina, O.A. and Widholm, J.M. (1996) Synthesis and turnover of cell-wall polysaccharides and starch in photosynthetic soybean suspension cultures. Plant Physiol. 111: 921–929.
Lytovchenko, A., Bieberich, K., Willmitzer, L. and Fernie, A.R. (2002) Carbon
assimilation and metabolism in potato leaves deficient in plastidial phosphoglucomutase. Planta 215: 802–811.
Martin, T., Frommer, W.B., Salanoubat, M. and Willmitzer, L. (1993) Expression of an Arabidopsis sucrose synthase gene indicates a role in metabolization of sucrose both during phloem loading and in sink organs. Plant J. 4:
367–377.
Massillon, D., Bollen, M., de Wulf, H., Overloop, K., Vanstapel, F., van Hecke,
P. and Stalmans, W. (1995) Demonstration of a glycogen/glucose 1-phosphate cycle in hepatocytes from fasted rats. J. Biol. Chem. 270: 19351–
19356.
Moreno-Bruna, B., Baroja-Fernández, E., Muñoz, F.J., Bastarrica-Berasategui,
A., Zandueta-Criado, A., Rodríguez-López, M., Lasa, I., Akazawa, T. and
Pozueta-Romero, J. (2001) Adenosine diphosphate sugar pyrophosphatase
prevents glycogen biosynthesis in Escherichia coli. Proc. Natl Acad. Sci.
USA 98: 8128–8132.
Mukerjea, R. and Robyt, J.F. (2004) Starch biosynthesis: the primer nonreducing-end mechanism versus the nonprimer reducing-end two-site insertion
mechanism. Carbohydr. Res. 340: 245–255.
Müller-Röber, B., Sonnewald, U. and Willmitzer, L. (1992) Inhibition of the
ADP-glucose pyrophosphorylase in transgenic potatoes leads to sugar-storing tubers and influences tuber formation and expression of tuber storage protein genes. EMBO J. 11: 1229–1238.
Nakai, T., Konishi, T., Zhang, X.-Q., Chollet, R., Tonouchi, N., Tsuchida, T.,
Yoshinaga, F., Mori, H., Sakai, F. and Hayashi, T. (1998) An increase in
apparent affinity for sucrose of mung bean sucrose synthase is caused by in
vitro phosphorylation or directed mutagenesis of Ser11. Plant Cell Physiol.
39: 1337–1341.
Neuhaus, H.E. and Emes, M.J. (2000) Nonphotosynthetic metabolism in
plastids. Annu. Rev. Plant Physiol. Plant Mol. Biol. 51: 111–140.
1376
SuSy controls starch production in leaves
Neuhaus, H.E., Henrichs, G. and Scheibe, R. (1995) Starch degradation in intact
amyloplasts isolated from cauliflower floral buds (Brassica oleracea L.).
Planta 195: 496–504.
Neuhaus, H.E., Kruckeberg, A.L., Feil, R. and Stitt, M. (1989) Reduced-activity mutants of phosphoglucose isomerase in the cytosol and chloroplast of
Clarkia xantiana. Planta 178: 110–122.
Nguyen-Quoc, B. and Foyer, C.H. (2001) A role for ‘futile cycles’ involving
invertase and sucrose synthase in sucrose metabolism of tomato fruit. J. Exp.
Bot. 52: 881–889.
Niittylä, T., Messerli, G., Trevisan, M., Chen, J., Smith, A.M. and Zeeman, S.C.
(2004) A novel maltose transporter essential for starch degradation in leaves.
Science 303: 87–89.
Nolte, K.D. and Koch, K.E. (1993) Companion-cell specific localization of
sucrose synthase in zones of phloem loading and unloading. Plant Physiol.
101: 899–905.
Okita, T.W. (1992) Is there an alternative pathway for starch synthesis? Plant
Physiol. 100: 560–564.
Porchia, A.C., Curatti, L. and Salerno, G.L. (1999) Sucrose biosynthesis in a
prokaryotic organism: presence of two sucrose-phosphate synthases in
Anabaena with remarkable differences compared with the plant enzymes.
Planta 210: 34–40.
Pozueta-Romero, J. and Akazawa, T. (1993) Biochemical mechanism of starch
biosynthesis in amyloplasts from cultured cells of sycamore (Acer pseudoplatanus). J. Exp. Bot. 44 Suppl.: 297–306.
Pozueta-Romero, J., Ardila, F. and Akazawa, T. (1991) ADP-glucose transport
by the chloroplast adenylate translocator is linked to starch biosynthesis.
Plant Physiol. 97: 1565–1572.
Pozueta-Romero, J., Muñoz, F.J., Rodríguez-López, M., Baroja-Fernández, E.
and Akazawa, T. (2003) New waves in the starch field. JSPP Lett (August)
24–31.
Quick, W.P., Scheibe, R. and Neuhaus, E. (1995) Induction of hexose-phosphate translocator activity in spinach chloroplasts. Plant Physiol. 109: 113–
121.
Riesmeier, J.W., Flügge, U.I., Schulz, B., Heineke, D., Heldt, H.W., Willmitzer,
L. and Frommer, W.B. (1993) Antisense repression of the chloroplast triose
phosphate translocator affects carbon partitioning in transgenic potato plants.
Proc. Natl Acad. Sci. USA 90: 6160–6164.
Saether, N. and Iversen, T.-H. (1991) Gravitropism and starch statoliths in an
Arabidopsis mutant. Planta 184: 491–497.
Scheidig, A., Fröhlich, A., Schulze, S., Lloyd, J.R. and Kossmann, J. (2002)
Downregulation of a chloroplast-targeted β-amylase leads to a starch-excess
phenotype in leaves. Plant J. 30: 581–591.
Scott, P. and Kruger, N. (1995) Influence of elevated fructose-2, 6-bisphosphate
levels on starch mobilization in transgenic tobacco leaves in the dark. Plant
Physiol. 108: 1569–1577.
Smith, S.M., Fulton, D.C., Chia, T., Thorneycroft, D., Chapple, A., Dunstan, H.,
Hylton, C., Zeeman, S.C. and Smith, A. (2004) Diurnal changes in the transcriptome encoding enzymes of starch metabolism provide evidence for both
transcriptional and posttranscriptional regulation of starch metabolism in
Arabidopsis leaves. Plant Physiol. 136: 2687–2699.
Smith, A.M., Neuhaus, H.E. and Stitt, M. (1990) The impact of decreased activity of starch-branching enzyme on photosynthetic starch synthesis in leaves
of wrinkled-seeded peas. Planta 181: 310–315.
Sonnewald, U. (1992) Expression of E. coli inorganic pyrophosphatase in transgenic plants alters photoassimilate partitioning. Plant J. 2: 571–581.
Stark, D.M., Timmerman, K.P., Barry, G.F., Preiss, J. and Kishore, G.M. (1992)
Regulation of the amount of starch in plant tissues by ADPglucose pyrophosphorylase. Science 258: 287–292.
Stitt, M., Gerhardt, R., Kürzel, B. and Heldt, H. (1983) A role for fructose 2, 6bisphosphate in the regulation of sucrose synthesis in spinach leaves. Plant
Physiol. 72: 1139–1141
Stitt, M. and Heldt, H.W. (1981) Simultaneous synthesis and degradation of
starch in spinach chloroplasts in the light. Biochim. Biophys. Acta 638: 1–11.
Stitt, M., Kürzel, B. and Heldt, H.W. (1984) Control of photosynthetic sucrose
synthesis by fructose 2, 6-bisphosphate: partitioning between sucrose and
starch. Plant Physiol. 75: 554–560.
Strand, A., Zrenner, R., Trevanion, S., Stitt, M., Gustafsson, P. and Gardeström,
P. (2000) Decreased expression of two key enzymes in the sucrose biosynthesis pathway, cytosolic fructose-1, 6-bisphosphatase and sucrose phosphate synthase, has remarkably different consequences for photosynthetic
carbon metabolism in transgenic Arabidopsis thaliana. Plant J. 23: 759–770.
Sweetlove, L.J., Burrell, M.M. and ap Rees, T. (1996) Characterization of transgenic potato (Solanum tuberosum) tubers with increased ADPglucose pyrophosphorylase Biochem. J. 320: 493–498.
Walters, R.G., Ibrahim, D.G., Horton, P. and Kruger, N.J. (2004) A mutant of
Arabidopsis lacking the triose-phosphate/phosphate translocator reveals metabolic regulation of starch breakdown in the light. Plant Physiol. 135: 891–
906.
Wang, A.-Y., Kao, M.-H., Yang, W.-H., Sayion, Y., Liu, L.-F., Lee, P.-D. and Su,
J.-C. (1999) Differentially and developmentally regulated expression of three
rice sucrose synthase genes. Plant Cell Physiol. 40: 800–807.
Wintermans, J.F.G.M. and de Mots, A. (1965) Spectrophotometric characteristics of chlorophylls a and b and their pheophytins in ethanol. Biochim.
Biophys. Acta 109: 448–453.
Yu, T.-S., Lue, W.-L., Wang, S.-M. and Chen, J. (2000) Mutation of Arabidopsis
plastid phosphoglucose isomerase affects leaf starch synthesis and floral initiation. Plant Physiol. 123: 319–325.
Zeeman, S.C., Tiessen, A., Pilling, E., Kato, K.L., Donald, A.M. and Smith,
A.M. (2002) Starch synthesis in Arabidopsis. Granule synthesis, composition, and structure. Plant Physiol. 129: 516–529.
Zrenner, R., Krause, K.P., Apel, P. and Sonnewald, U. (1996) Reduction of the
cytosolic fructose-1, 6-bisphosphatase in transgenic potato plants limits photosynthetic sucrose biosynthesis with no impact on plant growth and tuber
yield. Plant J. 9: 671–681.
Zrenner, R., Salanoubat, M., Willmitzer, L. and Sonnewald, U. (1995) Evidence
of the crucial role of sucrose synthase for sink strength using transgenic
potato plants (Solanum tuberosum L.). Plant J. 7: 97–107.
(Received April 27, 2005; Accepted June 3, 2005)