Plant Cell Physiol. 48(11): 1612–1623 (2007)
doi:10.1093/pcp/pcm133, available online at www.pcp.oxfordjournals.org
ß The Author 2007. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.
All rights reserved. For permissions, please email: [email protected]
Sucrose Synthase Oligomerization and F-actin Association are
Regulated by Sucrose Concentration and Phosphorylation
Kateri A. Duncan
1, 2
and Steven C. Huber
1, 2, 3,
*
1
Department of Plant Biology, University of Illinois at Urbana-Champaign, Urbana, IL 61801, USA
Program in Physiological and Molecular Plant Biology, University of Illinois at Urbana-Champaign, Urbana, IL 61801, USA
3
United States Department of Agriculture-Agricultural Research Service, Photosynthesis Research Unit and Department of Crop Sciences,
University of Illinois at Urbana-Champaign, Urbana, IL 61801, USA
2
Sucrose synthase (SUS) is a key enzyme in plant
metabolism, as it serves to cleave the photosynthetic endproduct sucrose into UDP-glucose and fructose. SUS is
generally assumed to be a tetrameric protein, but results in the
present study suggest that SUS can form dimers as well as
tetramers and that sucrose may be a regulatory factor for the
oligomerization status of SUS. The oligomerization of SUS
may also affect the cellular localization of the protein. We
show that sucrose concentration modulates the ability of
SUS1 to associate with F-actin in vitro and that calciumdependent protein kinase-mediated phosphorylation of recombinant SUS1 at the Ser15 site is a negative regulator of its
association with actin. Although high sucrose concentrations
and hyperphosphorylation have been shown to promote SUS
association with the plasma membrane, we show that the
opposite is true for the SUS–actin association. We also show
that SUS1 has a unique 28 residue coiled-coil domain that
does not appear to play a role in oligomerization, but may
prove to be significant in the future for interactions of SUS
with other proteins. Collectively, these results highlight the
multifaceted nature of SUS association with cellular
structures.
Keywords: Sucrose synthase — F-actin binding — protein
oligomerization — calcium dependent protein kinase —
protein phosphorylation — sugar sensing.
Abbreviations: AEBSF, 4-(aminoethyl)benzenesulfonyl fluoride hydrochloride; CD, circular dichroism; CDPK, calciumdependent protein kinase; DAP, days after pollination; DTT,
dithiothreitol; PBST, phosphate-buffered saline containing 0.1%
(v/v) Tween-20; MBP, maltose-binding protein; MCLR, microcystin-LR; PVPP, polyvinylpolypyrrolidone; SUS, sucrose
synthase; TFE, 2,2,2-trifluoroethanol.
Introduction
Sucrose synthase (SUS) is recognized to be an
important enzyme in sucrose metabolism and is essential
for growth of heterotrophic plant organs (Nguyen-Quoc
et al. 1990, Winter and Huber 2000a, Koch 2004). Although
SUS is a soluble protein, the ability of this enzyme also to
associate with membranes has been well documented
(Winter et al. 1997, Ruan et al. 2003, Hardin et al. 2004,
Duncan et al. 2006, Hardin et al. 2006). Both phosphorylation (Hardin et al. 2004) and high concentrations of sugars
(Hardin et al. 2006) have been shown to promote the
membrane association of SUS, which is postulated to
provide the substrate UDP-glucose to cellulose synthase for
cellulose production (Amor et al. 1995, Carlson and
Chourey 1996, Winter et al. 1997, Hardin et al. 2004).
The ability of SUS to bind to F-actin in vitro and to
associate with the actin cytoskeleton in vivo has also been
documented (Winter et al. 1998, Winter and Huber 2000a,
Winter and Huber 2000b, Azama et al. 2003, Matic et al.
2004). The SUS1–F-actin association has an apparent Kd
value of about 2 mM [calculated from results presented in
Winter et al. (1998)] with a stoichiometry at saturation of
about 0.2 molecules of SUS1 monomer bound per actin
subunit. This is a reasonably high affinity binding; however,
specific conditions mediating the SUS–F-actin association
have not been well studied.
We recently made the unexpected observation that
sucrose can affect the quaternary structure of SUS: in the
absence of sucrose, SUS behaves as a dimer during size
exclusion chromatography and it is only in the presence of
sucrose that the tetrameric form of SUS is observed. We
wanted to characterize this observation further because the
tetrameric form of SUS is thought to represent the active
enzyme (Su and Preiss 1978, Chourey 1981, Echt and
Chourey 1985, Chourey et al. 1986, McElfresh and Chourey
1988, Koch et al. 1992, Chourey and Taliercio 1994, Hardin
and Huber 2004) and, as noted above, sucrose has been
shown to stimulate binding of SUS to membranes in vitro.
We speculated that the effects of sucrose on membrane
association may involve changes in the oligomeric status of
the protein and, if so, that binding of SUS to F-actin might
also be affected.
Another recent observation is that SUS1 is predicted
by in silico analysis to contain a coiled-coil domain. Because
coiled-coils can function in oligomerization of proteins
(Tripet et al. 2000, Burkhard et al. 2001), we speculated that
the coiled-coil in SUS1 may be involved in assembling the
*Corresponding author: E-mail, [email protected]; Fax, þ1-217-244-4419.
1612
SUS oligomerization and actin binding
SEC kDa
tetrameric or dimeric form of the enzyme. However, an
important first step would be to verify that the predicted
coiled-coil can, in fact, function as a protein–protein
interaction domain. Hence, the objectives of the current
study were to (i) characterize the effects of sucrose, pH and
phosphorylation on oligomerization of SUS; (ii) determine
how sucrose and phosphorylation status affect the binding of SUS to F-actin; and (iii) determine whether the
predicted coiled-coil domain in SUS1 functions in oligomerization of SUS1.
100
443
200
150
A
SUS1
80
60
40
20
Results
0
10
100
% Maximum SUS Protein
Native maize leaf SUS oligomerization is affected by sucrose
levels
SUS was partially purified from the maize leaf
elongation zone, and size exclusion chromatography was
performed in the presence or absence of 0.1 M sucrose. We
typically include sucrose in buffers used to extract and
purify SUS because in our experience the sugar tends to
stabilize SUS protein and activity (data not shown). In the
presence of sucrose, SUS eluted from the column at a
position consistent with the tetrameric form of the protein
(360 kDa), while, in the absence of sucrose, SUS protein
eluted at a position expected for the dimeric form of the
protein (180 kDa). All three of the SUS isoforms were
affected similarly by sucrose (Fig. 1A, B, C). It was verified
that sucrose in the elution buffer had no effect on the
elution volume of standard proteins used to calibrate the
column (data not shown).
1613
11
12
13
14
11
12
13
14
12
13
14
B
SUS-SH1
80
60
40
20
0
10
100
C
SUS2
Recombinant MBP–SUS1 oligomerization is affected by
sucrose and phosphorylation
Size exclusion experiments were performed with the
recombinant maltose-binding protein (MBP)–SUS1 fusion
protein in the presence or absence of sucrose to determine if
the apparent oligomerization state of the recombinant
protein would behave similarly to the native protein. As
shown in Fig. 2A, the recombinant protein clearly eluted as
a dimer in the absence of sucrose, but as a tetramer in the
presence of sucrose. Free MBP eluted as a monomer in both
the presence and absence of sucrose (data not shown). Thus,
recombinant SUS1 protein behaves similarly to the native
protein in terms of the effect of sucrose on oligomerization.
To test whether phosphorylation affects oligomerization, the recombinant SUS1 protein (in these experiments
with the MBP tag removed) was run on a size exclusion
column before or after phosphorylation by recombinant
soybean calcium-dependent protein kinase b (CDPKb),
which readily phosphorylates the protein at the Ser15 site
(Hardin et al. 2004). When not phosphorylated and in
the absence of sucrose, recombinant SUS1 was a dimer,
but when the protein was phosphorylated it formed
80
60
40
20
0
10
11
Elution Volume (mL)
Fig. 1 Sucrose affects the elution of native maize SUS isoforms
during size exclusion chromatography. Elution of (A) SUS1, (B)
SUS-SH1 and (C) SUS2 isoforms as determined by immunoblotting
with isoform-specific antibodies (Duncan et al. 2006). SUS was
partially purified from the leaf elongation zone (basal 8 cm) and
size exclusion chromatography was performed in the presence
(filled circles) or absence (open circles) of 0.1 M sucrose.
Densitometry of immunoblots was performed and results are
plotted as a percentage of the maximum SUS protein measured.
Vertical dashed lines indicate the elution volumes of molecular
weight standards (SEC kDa).
1614
SUS oligomerization and actin binding
SEC kDa
21
21
250
− Sucrose 150
100
250
+ Sucrose 150
100
Fraction #
IB: MBP
IB: MBP
% Maximum SUS Protein
0
SEC kDa
15
0
44
20
13
14
15
16
17
18
19
11
12
66
6
7
8
9
10
kDa
3
B
9
A
443
200
150
100
80
− CDPK
+ CDPK
60
40
20
0
9
10
11
12
13
14
Elution Volume (mL)
15
Fig. 2 Sucrose affects the elution of recombinant maize SUS1 during size exclusion chromatography. (A) Immunoblots (IB) of
chromatography fractions of recombinant MBP–SUS1 fusion protein probed with the anti-MBP antibody. Elution positions of size exclusion
standard proteins are indicated at the top of the panel (SEC kDa). Size exclusion was performed in the presence or absence of 0.1 M
sucrose, as indicated to the left of each panel. (B) Size exclusion chromatography elution profiles of recombinant SUS1 (MBP tag cleaved
off) fractions in the presence of CDPK (phosphorylated, filled diamonds) or absence (non-phosphorylated, open diamonds).
a tetramer (Fig. 2B). Previous attempts to identify a
docking site on SUS1 for CDPKs were not successful
(S. C. Hardin and S. C. Huber, unpublished) and, thus,
formation of a higher molecular weight SUS1–CDPK
complex could not readily explain the shift in elution
volume observed. Moreover, CDPK was not present at
equimolar amounts with the SUS1 protein, which would
also reduce the likelihood of complex formation. These
results suggest that phosphorylation of SUS1 can also
influence the oligomerization state of the protein.
At pH 8.5, SUS1 protein had the broadest profile in the
absence of sucrose and apparently existed in various states
of aggregation, ranging from tetramers to monomers
(Fig. 3C). In the presence of sucrose at pH 8.5, SUS1 was
predominantly found as a tetramer. It is important to note
that the total amount of extract added to the columns was
equal and that the broad profile at pH 8.5 in the absence
of sucrose is not a reflection of differences in protein
concentration. Thus, pH affected the elution behavior of
SUS1 in both the presence and absence of sucrose.
pH affects SUS oligomerization
It is well recognized that pH is one of the factors that
has an impact on the enzymatic properties of SUS. In
general, sucrose cleavage activity usually has an acid pH
optimum while sucrose synthetic activity has an alkaline pH
optimum (Pontis et al. 1981, Morell and Copeland 1985,
Klotz et al. 2003). We wanted to determine whether pH
would affect the distribution of SUS protein between the
dimeric and tetrameric forms. To do this, SUS protein was
partially purified from developing kernels and equilibrated
with buffers at pH values ranging from pH 6.0 (optimum
for cleavage activity) to pH 8.5 (optimum for synthetic
activity), and the pH of the gel filtration chromatography
buffer was varied accordingly. The chromatographic
behavior of SUS1 was monitored using isoform-specific
antibodies (Duncan et al. 2006). When the pH was low
(pH 6.0), SUS1 protein eluted as a broad peak that
appeared to include both dimeric and tetrameric forms,
and the presence of sucrose had little effect (Fig. 3A). At pH
7.5, SUS1 had more distinct elution profiles, with the
tetramer predominant in the presence of sucrose and the
dimer predominant in the absence of sucrose (Fig. 3B).
Binding of native SUS to F-actin in vitro
F-actin association experiments were performed in the
presence of various sucrose concentrations with SUS
partially purified from developing kernels. As expected,
the ability of SUS1 to associate with F-actin was strongly
stimulated by sucrose (Winter and Huber 2000b). However,
as shown in Fig. 4, SUS1 binding to F-actin displayed a
distinct sucrose optimum at concentrations of 20–60 mM
sucrose (Fig. 4A). Concentrations of sucrose below 23 mM
were tested, which confirmed that maximum binding
required at least 23 mM sucrose (Fig. 4B). The densitometry
of the immunoblots in Fig. 4A and B indicated that binding
of SUS1 to F-actin was essentially dependent on sucrose,
but at concentrations above approximately 60 mM, binding
was strongly inhibited (Fig. 4C). Polymerization of actin
and sedimentation of F-actin were not affected by sucrose
concentration; rather, the binding of SUS was specifically
modulated.
Recombinant SUS1 actin binding
Recombinant MBP–SUS1 association with F-actin was
also tested to determine its ability to associate under various
SUS oligomerization and actin binding
200
150
kDa
100
A
+ Actin
pH 6.0
80
Pellets
Supernatants
A
3
23
53
10
153
3
20
3
443
3
23
53
10
153
203
3
SEC kDa
1615
mM Sucrose
100
75
IB: SUS1
50
Actin Pellets: CBB
60
− Actin
B
3
8
13
18
Supernatants
kDa
0
10
11
12
13
14
15
16
100
50
C
20
0
10
11
12
13
14
15
16
C
pH 8.5
60
% SUS1 associated with actin pellet
− Actin
40
80
Actin Pellets: CBB
pH 7.5
60
100
IB: SUS1
75
30
25
20
15
10
5
0
0
40
mM Sucrose
IB: SUS1
75
B
80
100
Pellets
100
+ Actin
% Maximum SUS Protein
IB: SUS1
3
8
13
18
21
23
20
100
75
21
23
40
20
40
60
80
100 120 140 160 180 200
mM Sucrose
20
0
10
11
12
13
14
15
16
Elution Volume (mL)
Fig. 3 Effect of pH and sucrose on the elution of SUS1 protein
during size exclusion chromatography. SUS was partially purified
from developing maize kernels, and the SUS1 isoform was
specifically monitored using isoform-specific antibodies.
Chromatography was performed at (A) pH 6.0, (B) pH 7.5 and
(C) pH 8.5 in the presence (filled circles) or absence (open circles)
of 0.1 M sucrose. Vertical dashed lines indicate the elution volumes
of molecular weight standard proteins (SEC kDa).
sucrose concentrations. We found that the recombinant
protein (Fig. 5A) had a similar response to that observed
with native SUS1, in that binding was minimal at the lowest
concentration of sucrose (3 mM), and there was an
optimum concentration around 50 mM above which significant inhibition of binding was observed (Fig. 5C).
Interestingly,
CDPK-mediated
phosphorylation
of
Fig. 4 Sucrose dependence of native SUS1 binding to F-actin
in vitro. SUS protein was partially purified from 20 DAP maize
kernels, and the SUS1 isoform was monitored using SUS1-specific
antibodies in the supernatants (left panels) and pellets (right panels)
obtained after sedimentation of F-actin. Coomassie blue (CBB)
staining of F-actin pellets is shown, and concentrations of sucrose
are indicated at the top of the lane. Immunoblots (IB) showing cosedimentation of SUS1 with F-actin in the presence of (A) high and
(B) low sucrose concentrations. In both experiments, equivalent
reactions without exogenous actin (‘–actin’) verified that SUS1
protein did not sediment in the absence of F-actin.
(C) Densitometry of immunoblots in (A) and (B) showing sucrose
dependence of SUS1 protein binding to F-actin, expressed as a
percentage of the total SUS1 protein supplied.
recombinant SUS1 at the Ser15 site was a negative regulator
of F-actin binding in vitro especially at higher concentrations of sucrose (Fig. 5B, C).
Manipulation of sugar levels in vivo
We wanted to determine whether sucrose concentration in vivo would affect the binding of endogenous SUS1
SUS oligomerization and actin binding
Pellets
A
10
3
15
3
20
3
3
23
53
10
3
15
3
20
3
mM Sucrose
100
+ Actin 75
IB: SUS1
50
Actin Pellets: CBB
100
75
− Actin
CBB
Supernatants
B
kDa
100
wash
− +
−
+
sucrose
75
IB: SUS1
50
37
IB: Actin
Pellets
3
3
20
3
15
3
23
53
10
10
3
15
3
20
3
3
23
53
mM Sucrose
Dark
+ Actin
150
IB: pS15
100
50
B
AU/mg total protein
IB: pS15
100
35
30
Dephosphorylated SUS1
Phosphorylated SUS1
25
Dark
3
2
15
0
Sucrose:
0
0
50
100
150
200
250
mM sucrose
Fig. 5 Sucrose dependence of recombinant SUS1 protein binding
to F-actin in vitro. (A) Unphosphorylated SUS1, as monitored with
SUS1-specific antibodies, and (B) SUS1 phosphorylated at the
Ser15 site, as monitored with phospho-specific antibodies (pS15).
The experimental design is as described in legend of Fig. 4.
(C) Densitometry of immunoblots in (A) and (B) showing the effect
of Ser15 phosphorylation on sucrose dependence of SUS1 protein
binding to F-actin, expressed as a percentage of the total SUS1
protein supplied.
to the actin cytoskeleton or to membranes. To test this,
4-week-old maize seedlings were kept in normal light/dark
conditions or transferred to darkness for 36 h to deplete
the plant of non-structural carbohydrates, including
sucrose (Brouquisse et al. 1998). The leaf elongation zone
was harvested from these plants and tissue was extracted
with and without addition of 0.15 M sucrose to buffers.
The soluble protein fraction (100 K supernatant) and
Light
4
1
5
IB: Actin
5
20
10
IB: SUS1
75
50
37
Actin Pellets: CBB
150
− Actin
% SUS asscociated with actin pellet
mem
+
100
kDa
C
actin
−
Light
kDa
3
23
53
Supernatants
A
100K Super
1616
actin
actin
mem
mem
wash
wash
−
+
−
+
−
+
Fig. 6 Extended dark treatment (36 h) of maize seedlings reduces
SUS1 association with microsomal membranes and the actin
cytoskeleton. (A) Immunoblots (IB) using SUS1- specific antibodies
of the soluble protein fraction (100 K Super), detergent-insoluble
fraction (actin), detergent-soluble (mem) fraction and wash fraction
(wash), produced after inversion of plasma membrane vesicles with
Brij-58. Sucrose (0.15 M) was added to the extraction buffer as
indicated at the top of the figure, and light and dark treatments are
indicated to the left of the IB panels. (B) Densitometry of the
immunoblots in (A); values plotted are means SEs from two
independent experiments.
microsomal membrane and crude actin cytoskeleton fractions were prepared and analyzed for actin and SUS1
protein by immunoblotting. As expected, the majority of
the SUS1 protein was present in the 100 K supernatant
(soluble protein) fraction, and the content of SUS1 protein
in the light- and dark-treated samples was very similar
(Fig. 6A). Much of the actin protein was also found in the
soluble fraction, which reflects either depolymerization
during extraction or the presence of monomeric G-actin
in vivo. However, actin protein was clearly present in
the crude cytoskeleton fraction (labeled ‘actin’ fraction
SUS oligomerization and actin binding
in Fig. 6A) and the recovery of actin protein was similar for
the light- and dark-treated samples. The most important
result is that the recovery of SUS1 protein in the actin
cytoskeleton and membrane fractions was substantially
lower in the plants subjected to 36 h of darkness compared
with the control plants maintained on a normal day/night
cycle (Fig. 6B). It is well known that soluble sugars are
rapidly depleted by short-term extended darkness in maize
(Kalt-Torres and Huber 1987, Brouquisse et al. 1998) and
thus these results are consistent with the notion that sugars
promote the actin and membrane association of SUS1 in
planta. Interestingly, the only effect of adding 0.15 M
sucrose to the extraction buffer was to decrease recovery of
SUS1 protein in the actin cytoskeleton fractions (Fig. 6B).
This is consistent with the notion that high concentrations
of sucrose reduce actin binding in vitro (Figs. 4C, 5C)
whereas membrane association is not inhibited (Hardin
et al. 2006).
Analysis of a predicted coiled-coil in SUS1
Algorithms are available for the analysis of the primary
structure of proteins to predict the occurrence of coiledcoils, which can function broadly in protein–protein
interactions, including oligomerization (Burkhard et al.
2001). Prediction programs such as Coils and MultiCoil
(Wolf et al. 1997) that are available on the ExPASy
Proteomics Server (http://us.expasy.org/) predicted that
SUS1 contains a coiled-coil whereas the probability of a
coiled-coil domain in SUS-SH1 and SUS2 is very low
(Table 1). The Paircoil2 program (McDonnell et al. 2006),
which predicts coiled-coils based on a pairwise residue
correlation, is the most recently updated and is considered
the most stringent analysis and also predicted a four-heptad
coiled-coil in SUS1 but not in the other two isoforms.
We wanted to test the prediction that SUS1 contains a
coiled-coil. To do this, we produced a synthetic peptide
(referred to as SS26) that is based on residues 199–226 of
SUS1 (Fig. 7A). Two experimental approaches were taken
to test the ability of SS26 to form a coiled-coil. First, we
attempted to cross-link the SS26 peptide chemically with a
low concentration of glutaraldehyde in the presence of the
helix-stabilizing agent 2,2,2-trifluoroethanol (TFE; Luo and
Baldwin 1997, and references within) and compared the
results with a control peptide (SS4) not predicted to form a
coiled-coil. As shown in Fig. 7B, the SS26 peptide formed
higher order structures (dimer, trimer and tetramer),
indicating that it has the ability to form a coiled-coil
in vitro, while the SS4 control peptide did not form higher
order structures (Fig. 7B). The second approach involved
circular dichroism (CD) analysis of the SS26 peptide in the
presence of varying concentrations of TFE to stabilize
helical structures. The CD spectrum of the SS26 peptide
in the presence of 10–30% TFE was characteristic of an
Table 1
1617
Prediction of a Coiled-coil domain in maize SUS1
Program
Coils
Paircoil2
Multicoil
Parameter
3 Heptad
4 Heptad
--Overall
probability
Dimer
Trimer
Probabilities
SUS1
SUS-SH1
SUS2
0.939
0.556
0.022
0.849
0.096
0.022
No coil
0.106
0.175
0.003
No coil
0.520
0.188
0.660
0.008
0.098
0.002
0.049
Output from the indicated programs are shown (programs
available on the ExPASy Proteomics server (http://ca.expasy.org/).
In the Coils and Muticoil programs values 4 0.8 indicate a high
probability of containing a coiled-coil; Paircoil2 values 5 0.025 are
indicative of a coiled-coil.
a-helix with a trough in the spectrum between 205 and
225 nm (Yang et al. 1986). The ratio of ellipticities measured
at 222 and 208 nm was calculated because ratios above
0.80 are indicative of a coiled-coil region (Lau et al. 1984).
At concentrations of 20% TFE and above, the 222 nm/
208 nm ratio was above 0.80 (Fig. 7C inset), indicating that
the SS26 peptide has the ability to form a coiled-coil. Thus,
two lines of evidence confirm the prediction that SUS1
contains a coiled-coil region involving residues between
Leu199 and Asp226.
Recombinant truncation analysis of SUS1
Truncation mutants of SUS1 were made to determine
which regions of the SUS1 polypeptide are required for
oligomerization. The truncation mutants, which are shown
schematically in Fig. 8A, contained only the N-terminal
portions of SUS1 (up to Asn191, Leu271 or Thr362) and
were compared with full-length MBP–SUS1. The truncation at Asn191 terminates the polypeptide before the coiledcoil, while the truncations at Leu271 and Thr362 terminate
after the coiled-coil region. The elution profiles of the
truncations and full-length MBP–SUS1 during size exclusion chromatography are shown in Fig. 8B. Densitometry
analysis of the immunoblots shown in Fig. 8B was used to
calculate the peak elution volumes for the SUS1 polypeptides. Comparison with the elution volumes for the standard
proteins used to calibrate the column indicated that the
truncation before the coiled-coil (Asn191) formed a monomer, as did the truncation after the coiled-coil (Leu271; see
Fig. 8C). The truncation at Thr362 formed a trimer/
tetramer while the full-length MBP–SUS1 formed a
tetramer (Fig. 8C). Thus, formation of the tetrameric
form of SUS1 required the catalytic region of the protein,
whereas formation of dimeric molecules appears to
require residues located between Leu271 and Thr362.
1618
SUS oligomerization and actin binding
A
Heptad 1
Heptad 2
Heptad 3
Heptad 4
N- L N D R I R S | L S A L Q G A | L R K A E E H | L S T L Q A D -C
a b c d e f g | a b c d e f g |a b c d e f g | a b c d e f g
B
% TFE
C
Ellipticity (deg*cm2*dmol−1)
SS26
SS4
Da
17300
Da
Tr
T
D
M
17300
8200
3700
1704
M
TFE (%) 0
Gld.(0.001%) −
0
+
8200
3700
15
+
0
+
15
+
30
+
θ 222/208
0.32
0.41
0.52
0.85
0.87
0
5
10
20
30
8000
6000
4000
2000
0
−2000
−4000
−6000
−8000
194
204
214
224
234
244
Wavelength (nm)
Fig. 7 Confirmation of the predicted coiled-coil in maize SUS1. (A) Sequence of the four- heptad repeat in SUS1 that is predicted to be a
coiled-coil region (residues 199–226). (B) Tris-Tricine gels of a control synthetic peptide (SS4), not predicted to form a coiled-coil, and the
SS26 synthetic peptide based on the sequence shown in (A). Peptides were incubated with the indicated concentration of TFE, a helixstabilizing agent, and the chemical cross-linker glutaraldehyde (Gld) at 0.001%. Oliogomeric structures are listed to the right of the panels
indicating monomer (M), dimer (D), trimer (T) and tetramer (Tr). (C) CD spectrum of the SS26 synthetic peptide in the presence of 30% TFE.
Ratios of the ellipticities at 222 and 208 nm at varying concentrations of TFE are presented in the inset.
A
Catalytic
N
MBP
CC
MBP
CC
MBP
Monomer
L271
Monomer
N191
SEC kDa
fraction #
IB: MBP
16
17
18
19
20
21
22
23
24
25
26
27
28
15
0
50
75
50
IB: MBP
13
4
14 43
15
16
17
18
2
19 00
20
21
22
23
73
L271
100
T362
75
IB: MBP
132
Full
150
Length 100
6
5.8
5.6
5.4
5.2
5
4.8
4.6
4.4
4.2
4
9.5
T362 (Trimer)
Full length
(Tetramer)
N191
(monomer)
L271
(monomer)
10.5
11.5
12.5
13.5
14.5
15.5
16.5
Elution volume (mL)
6
7
6
8 69
9
10
11
12
13 443
14
15
16
2
17 00
18
83
C
log (Molecular weight)
kDa
75
N191
66
Estimated
Monomer Size
(kDa)
64
Trimer/ Tetramer
T362
18
19
20
21
22
23 66
24
25
26
27
28
29
30 2
9
B
C Tetramer
Linker
CC
IB: MBP
Fig. 8 Effect of C-terminal truncation on the oligomerization status of MBP–SUS1 fusion proteins. (A) Schematic representation of SUS1
showing the predicted catalytic domain (MacGregor 2002), predicted coiled-coil, and location of stop codons introduced to produce the
truncation mutants T362, L271 and N191. (B) Size exclusion chromatography of recombinant proteins depicted in (A) detected by
immunoblotting (IB) with anti-MBP antibodies. Chromatography fraction numbers are indicated above each lane at the top of the IB panels.
The elution positions of molecular weight standard proteins are indicated above the fraction numbers for each panel (SEC kDa).
(C) Calibration curve for the Superdex 200 HR size exclusion column showing the estimated molecular weight for the full-length
MBP–SUS1 and truncation mutants. Comparison with the predicted monomer size of each protein, shown at the left side of (B), resulted in
the aggregation state noted alongside each recombinant protein.
SUS oligomerization and actin binding
Hence, the coiled-coil region, which is located between
residues 199 and 226, is not sufficient for dimerization.
Discussion
The fundamental observation made in the present
study is that SUS, which is generally regarded as a
tetrameric protein, appears to readily form dimers and,
under some conditions, monomers as well. One of the major
factors affecting oligomerization is the sucrose concentration, which may represent a new component of sucrose
‘sensing’ in heterotrophic plant cells. In addition, a
correlation that is beginning to emerge is that factors that
promote SUS oligomerization also affect binding of the
protein to membranes and F-actin. This has potential
implications for the nature of the binding site(s) involved
and how the intracellular distribution of SUS in vivo may
be controlled by various metabolic factors.
SUS oligomerization is affected by sucrose concentration,
phosphorylation and pH
Both native SUS, partially purified from developing
kernels, and recombinant MBP–SUS1 were affected similarly by sucrose levels (Figs. 1, 2A). Previous studies with
developing maize kernels confirmed the presence of all three
SUS isoforms (SUS1, SUS-SH1 and SUS2) and suggested
that SUS2 existed primarily as a hetero-oligomer with
SUS1, and that SUS-SH1 formed primarily homooligomers (Duncan et al. 2006). In the present study, we
monitored all three isoforms separately using isoformspecific antibodies and could demonstrate the sucrosemediated conversion of dimers to tetramers in all cases
(Fig. 1). These results indicate that sucrose might act as
a metabolic signal that controls the ability of SUS to
form tetramers. Recently, sucrose was shown to be essential
for the binding of SUS to membranes in vitro (Hardin et al.
2006), which now leads us to speculate that the tetrameric
form of SUS may be essential for membrane binding.
Thus, the presence of abundant sucrose in vivo might be
expected to promote SUS association with the plasma
membrane, where it could provide substrates for cell
wall glucan synthesis (Amor et al. 1995). This may be
one of the cellular mechanisms that control allocation of
carbon to structural carbohydrates in relation to sugar
availability. Indeed, depletion of sucrose in vivo by shortterm extended darkness reduced the association of SUS
with microsomal membranes (Fig. 6), which is consistent
with this notion.
Use of a recombinant SUS1 protein also allowed us to
study the effect of phosphorylation on oligomerization.
As purified from Escherichia coli, the recombinant protein
is completely unphosphorylated (Hardin et al. 2004)
and behaved as a dimeric protein in the absence of
1619
sucrose (Fig. 2B). However, following phosphorylation by
CDPKb, the recombinant SUS1 exhibited a marked shift in
elution volume, suggesting an increase in oligomerization
(Fig. 2B). Thus, phosphorylation of SUS1 at the Ser15 site
(Huber et al. 1996) promotes maintenance of the tetrameric
structure even in the absence of sucrose. Phosphorylation,
specifically at the N-terminal Ser15 site, has also been
shown to promote the membrane association of SUS
(Huber et al. 1996, Hardin et al. 2004). Hence, we are
speculating that the effect of phosphorylation may be
mediated, in part at least, by promotion of tetramer
formation. Differences in the phosphorylation status of
native SUS purified from different sources may explain why
tetramers are observed in some studies even in the absence
of sucrose (Morell and Copeland 1985, Klotz et al. 2003).
Previous studies with maize kernel SUS reported the
occurrence of more highly aggregated forms, including
tetramers, hexamers and octamers (Su and Preiss 1978);
interestingly, ionic strength was reported to be a major
factor affecting oligomerization, with low ionic strength
promoting aggregation. Ionic strength is not likely to be a
factor in our experiments, as gel filtration buffers contained
0.1 M NaCl and addition of sucrose would not, of course,
affect ionic strength. We do, however, recognize that
sucrose will affect the osmotic strength of the medium and
that this may play a role in SUS oligomerization, but we
have not explored this aspect further.
In the absence of sucrose, low pH (6.0) promoted
tetramer formation whereas high pH (8.5) produced
a mixture of forms from monomers to tetramers (Fig. 3).
The ability of SUS1 to form different oligomeric structures
as a function of pH has potential implications for
mechanisms regulating catalytic activity and also membrane
binding activity. Low pH has been shown to stimulate
membrane binding of SUS1 strongly in vitro (Hardin et al.
2006), and we speculate that promotion of tetramer
formation by low pH may be partially responsible.
In vivo, there is evidence for pH variant microdomains
(Schwiening and Willoughby 2002), and it is possible that
these regions within cells may affect oligomerization of
SUS. If SUS were to encounter such a microdomain within
the plant cell, such as in the vicinity of the plasma
membrane Hþ-ATPase where pH gradients exist (Young
et al. 1998), it is possible that SUS oligomerization would
occur and would promote association with the membrane.
In addition, pH is known to have a profound effect on the
catalytic activities of SUS, with acidic conditions promoting
sucrose cleavage while basic conditions promote sucrose
synthetic activity (Tsai 1974, Pontis et al. 1981, Morell and
Copeland 1985). Thus, acidic microdomains in the vicinity
of the plasma membrane would enhance the sucrose
cleavage activity of membrane-bound SUS. One further
implication of our results is that the oligomeric status of
1620
SUS oligomerization and actin binding
SUS may directly control the balance between sucrose
cleavage and synthetic activities. The alkaline pH optimum
of sucrose synthetic activity implies that SUS monomers or
dimers may have increased synthetic activity, whereas SUS
tetramers would be primarily involved in sucrose cleavage.
Independent regulation of SUS binding to F-actin and
membranes
The stimulation of SUS binding to F-actin by sucrose
(Figs. 4, 5) indicates that SUS may bind to the cytoskeleton
in vivo only when the cell has a sufficient supply of sucrose.
Typical physiological concentrations of sucrose range from
20 to 100 mM (Gerhard et al. 1987, Winter et al. 1994),
which are comparable with the sucrose concentrations that
produce maximum binding in vitro (Figs. 4, 5). Whether
SUS oligomerization affects F-actin binding cannot be
readily discerned at the present time. However, the
observation that high sucrose concentrations inhibit binding of SUS1 to F-actin but do not inhibit membrane
binding (Hardin et al. 2006) indicates that the site(s)
involved in the two binding activities are not identical.
This is significant because many known actin-binding
proteins also bind lipids apparently at the same site
(Isenberg and Goldmann 1995, Meerschaert et al. 1998).
Another difference between the two binding activities
involves the effect of SUS phosphorylation, which promotes
membrane binding (Hardin et al. 2004) but inhibits F-actin
binding (Fig. 5). Thus, the binding sites are probably not
identical, which would allow for independent regulation of
these two activities. The physiological significance of SUS
binding to F-actin is not clear. However, actin mediates
various functions in plant cells including directing the plane
of cell division and cell wall synthesis, positioning organelles, and allowing a means for cytoplasmic streaming
(McCurdy et al. 2001, and references within). The ability of
SUS1 to associate with actin could be either a transport
mechanism within cells to shuttle SUS from one area of the
cell to another or a mechanism to localize SUS at specific
positions in the cytoplasm of cells to cleave sucrose and
produce metabolites for specific cellular fucntions.
Identifying the factors that differentially regulate membrane
and F-actin binding may help elucidate the physiological
significance of these associations.
SUS1 contains a coiled-coil region
The predicted coiled-coil domain in SUS1 was confirmed by both chemical cross-linking and CD analysis
(Fig. 7). Because coiled-coils can function in oligomerization of proteins (Tripet et al. 2000, Burkhard et al. 2001), we
wanted to determine whether the coiled-coil region in SUS1
played such a role. However, analysis of MBP–SUS1
truncation mutants indicated that the coiled-coil domain
is not sufficient to confer oligomerization of the protein.
Rather, sequences located between residues 272 and 362
were required for dimer formation. The basis for this is not
clear, but indicates that the identified coiled-coil cannot be
assigned a function at present. Further demonstration that
the coiled-coil is not required for oligomerization of the
SUS tetramer or heterotetramer is provided by the
observation that SUS-SH1 (which is not predicted to
contain a coiled-coil) forms homo-oligomers in vivo
(Duncan et al. 2006) and occurs as a tetramer (Fig. 1B).
The possibility that the confirmed coiled-coil in SUS1
mediates interactions with other cellular proteins will be
interesting to explore in the future.
Concluding remarks
SUS was previously thought to be only a tetrameric
protein but we now show that SUS can also form dimers.
The oligomerization of SUS is probably regulated by
several factors, but sucrose concentration may be one of
the most important in vivo. Sucrose depletion experiments
are consistent with the notion that binding of SUS to
F-actin and membranes in vivo is enhanced by sucrose
(Fig. 6B). This may be an important mechanism that
controls allocation of carbon among competing pathways.
Regulation of SUS binding to membranes and F-actin may
be mediated, at least in part, by changes in protein
oligomerization. Much remains to be done to confirm and
extend the new working model that is presented here.
Materials and Methods
Chemical cross-linking and Tris-tricine gels
All chemicals were purchased from Sigma-Aldrich (St Louis,
MO, USA) unless otherwise stated. Synthetic peptides were
produced by United Biochemical Research (Seattle, WA, USA).
A synthetic peptide (SS26) based on the predicted coiled-coil
region of SUS1 (residues 199–226: LNDRIRSLSALQGALRK
AEEHLSTLQAD) and a control peptide (SS4) that is not
predicted to form a coiled-coil (VLARLHSVRERIKK) were
prepared as 1 mg ml1 stocks in H2O and the pH was adjusted to
7.0 with 0.5 M Tris base. Cross-linking was performed with 0.001%
glutaraldehyde in a reaction containing 200 mM peptide and 5 mM
KH2PO4. The helix-stabilizing agent TFE was added at varying
concentrations (0–30%) to the reaction as indicated. The crosslinking reactions were incubated at room temperature for 3 h and
reactions were stopped by addition of 200 mM NaBH4 at a final
concentration of 200 mM. Reaction mixtures were taken to dryness
in a Heto speed vac (Heto Lab Equipment, Allerod, Denmark).
The dried samples were resuspended in 15 ml of H2O, an equal
volume of Tris-Tricine loading buffer (200 mM Tris pH 6.8, 2%
SDS, 40% glycerol, 0.04% Coomassie blue G-250) was added,
and the mixtures were heated at 958C for 5 min. The samples were
then run on pre-cast 16.5% 1 mm Tris-Tricine gels (Bio-Rad,
Hercules, CA, USA) at constant current (20 mA) in a buffer
containing 100 mM Tris, pH 8.3, 100 mM Tricine and 0.1% SDS.
Gels were silver stained according to protocol, using the Bio-Rad
Silver Stain Plus kit (Hercules, CA, USA), and scanned using a flat
bed scanner.
SUS oligomerization and actin binding
Circular dichroism (CD)
Samples of the SS26 peptide were analyzed on a JASCO J-600
CD spectropolarimeter. The samples analyzed contained 50 mM
peptide, 5 mM KH2PO4 and varying concentrations of TFE
(0–30%). Spectra were collected from 190 to 250 nm in a 1 mm
cuvette. The measurements for ellipticity (y) of a coiled-coil were
calculated by taking the ratio of values obtained at wavelengths of
222 and 208 nm (Lau et al. 1984).
Preparation of SUS extracts
Maize tissue used in these studies was from the hybrid
B73 Mo17; tissues were harvested and frozen in liquid nitrogen
and then stored at 808C until use. SUS was extracted from
kernels at 20 days after pollination (DAP), the 0–8 cm basal region
of the maize leaf elongation zone, or seedlings grown for light/dark
treatments. Frozen tissue was ground in an extraction buffer
containing 100 mM MOPS pH 7.5, 10 mM dithiothreitol (DTT),
5 mM EDTA, 1 mM EGTA, 20 mM NaF, 5 mM Na2MoO4, 1 mM
Na3VO4, 0.1 mM microcystin-LR (MCLR), 1 mM 4-(aminoethyl)benzenesulfonyl fluoride hydrochloride (AEBSF), 5 mM caproic
acid, 1 mM benzamidine, 2 mM leupeptin, 5 mg ml1 soybean
trypsin inhibitor, 10 mM MG132, 1% (w/v) polyvinylpolypyrrolidone (PVPP), 2% (v/v) polyethyleneglycol (PEG) and (for plus
sucrose treatments) 0.25 M sucrose at a ratio of 1 : 4 (g ml1) in a
mortar with a pestle. The samples were then centrifuged at
9,000 g for 20 min and the supernatant was filtered through one
layer of Miracloth and then centrifuged again at the same speed for
clarification. The supernatants were removed and then centrifuged
at 100,000 g for 1 h. The resulting supernatants were recovered
and are referred to as the 100 K supernatants. The samples were
then dialyzed into a buffer containing 20 mM MOPS pH 7.5, 1 mM
DTT, 1 mM EDTA, 1 mM EGTA, 1 mM NaF and 1 mM
phenylmethylsulfonyl flouride (PMSF).
Phosphorylation of recombinant SUS1
Recombinant SUS1 was produced using the MBP tag (New
England Biolabs, Beverly, MA, USA) and purified according to
the protocol specified below. The purified MBP–SUS1 was
phosphorylated using a recombinant soybean His6-CDPKb that
was purified by using Qiagen nickel affinity agarose (Valencia, CA,
USA); the protein was purified according to Qiagen protocols and
was tested for activity by methods described in Sebastia et al.
(2004). Phosphorylation reactions were carried out in a batch
reaction by incubating MBP–SUS1 with CDPKb in a 100 ml
volume containing 20 mM MOPS, 2 mM ATP, 10 mM MgCl2,
0.1 mM CaCl2 and 5 mM NaF. Phosphorylation reactions were
incubated at room temperature for 1 h before further experimentation was initiated.
Size exclusion chromatography
An aliquot of the 100 K supernatant (200 ml, equivalent to
700 mg of total protein) or recombinant SUS1 protein was loaded
onto a Superdex 200 HR 10/30 size exclusion column (Amersham
Biosciences, GE Healthcare, Piscataway, NJ, USA) that had been
pre-equilibrated with a buffer containing 20 mM MOPS, pH 7.5,
1 mM DTT, 100 mM NaCl and 100 mM sucrose as indicated. The
column was run at 0.5 ml min1 and 250 ml fractions were collected.
Thyroglobulin (669 kDa), apoferritin (443 kDa), b-amylase
(200 kDa), alcohol dehydrogenase (150 kDa) and carbonic anhydrase (29 kDa) were used as standards to calibrate the column.
1621
Electrophoresis/immunoblotting
Samples were separated on 7 or 10% polyacrylamide–0.1%
SDS gels and transferred to polyvinylidene fluoride fluorescencespecific membranes (Millipore, Bedford, MA, USA). Membranes
were blocked in a 2% fish gelatin solution in phosphate-buffered
saline (PBS: 5 mM NaH2PO4, 150 mM NaCl, pH 7.4) before
being incubated with primary antibodies at 1 : 10,000–1 : 25,000
dilution in PBS containing 0.1% (v/v) Tween-20 (PBST).
Washes were performed in PBST, and an Alexa- Fluor 680- or
IR 800-conjugated secondary antibody was used (Rockland
Innumochemicals Gillbertsville, PA, USA) diluted at 1 : 20,000 in
PBST. Densitometry of immunoblots was performed using a
Li-Cor Odyssey (Lincoln, NE, USA).
Preparation of recombinant protein
The MBP–SUS1 fusion protein was expressed in E. coli and
purified as specified by the protocol provided by New England
BioLabs (Beverly, MA, USA). The MBP–SUS1 was dialyzed into a
buffer containing 20 mM MOPS pH 7.4 and 1 mM DTT before
experiments were carried out.
F-actin binding
Rabbit muscle actin was purchased from Cytoskeleton Inc.
(Denver, CO, USA) and resuspended in 500 ml of actin resuspension buffer (5 mM Tris, pH 8, 0.2 mM ATP, 0.2 mM CaCl2 and
0.5 mM DTT). SUS–F-actin reactions were run at fixed molar
ratios as follows. Either 100 K supernatant (15 mg of total protein,
equivalent to 10 pmol SUS) from 20 DAP kernels or recombinant
MBP–SUS1 (30 pmol, pre-spun at 100,000 g) were incubated
with 300 pmol actin. The actin polymerization reaction was
initiated by adding to a final concentration of 50 mM KCl, 5 mM
MgCl2, 1 mM ATP and 1 mM DTT. For experiments with varying
sucrose concentrations, the amount of sucrose added to the actin
polymerization reaction took into account that the actin contributed 1% (w/v) sucrose to the final reaction mixture. The
complete reactions were incubated at room temperature for 1 h to
allow the actin to polymerize. Reactions were then spun at
100,000 g for 30 min and the supernatant was denatured directly
in SDS–PAGE loading buffer. Reaction pellets were washed once
with actin resuspension buffer, and then resuspended in SDS–
PAGE loading buffer.
Light/dark treatment of maize seedlings
Maize seedlings were grown for 4 weeks in soil in a growth
chamber at 28/208C day/night temperatures, under 14 h days with
a light intensity of 250 mE m2 s1 and 70% humidity. After 4
weeks, a subset of the seedlings was placed in complete darkness
for 36 h at 258C. Samples were collected by harvesting the first
10 cm (from the soil surface) of leaf sheath material from plants in
either the light or dark treatment. Actin (along with microsomes)
was purified by grinding in an extraction buffer (described above)
in either the presence or absence of 150 mM sucrose. Samples were
centrifuged twice at 9,000 g for 20 min and the final supernatant
was collected. Samples were then centrifuged at 100,000 g and an
aliquot of the supernatant was collected as the soluble fraction.
The 100,000 g pellet was treated with 0.1% (w/v) Brij-58 to invert
the vesicles (as done in Duncan et al. 2006) and then centrifuged at
100,000 g to re-pellet the microsome/cytoskeleton fractions. The
Brij-58-treated pellet was then resuspended in a treatment buffer
containing 100 mM MOPS pH 7.5, 10 mM DTT, 5 mM EDTA,
1 mM EGTA, 20 mM NaF, 5 mM Na2MoO4, 1 mM Na3VO4,
0.1 mM MCLR, 1 mM AEBSF, 5 mM caproic acid, 1 mM
benzamidine, 2 mM leupeptin, 5 mg ml1 soybean trypsin inhibitor
1622
SUS oligomerization and actin binding
and 10 mM MG132 with or without 150 mM sucrose. The sample
was allowed to sit on ice for 30 min and was then centrifuged again
at 100,000 g. The supernatant was taken as the ‘wash’ sample.
The pellet was resuspended in the above treatment buffer (again
with or without sucrose) with the addition of 1% (v/v) Triton
X-100 and 0.25% (w/v) CHAPS to solubilize membranes. The
sample was then centrifuged at 40,000 g for 30 min; the pellet
(detergent insoluble) was taken as the cytoskeleton (actin) fraction
and the supernatant was taken as the membrane or microsomal
fraction (detergent soluble).
Acknowledgments
This research was supported in part by funds from the US
Department of Energy (Grant DE-AI05-2003ER to S.C.H.). The
circular dichroism experiments reported here were performed at
the Laboratory for Fluoresence Dynamics (LFD) at the University
of Illinois at Urbana-Champaign (UIUC), which is supported
jointly by the National Center for Research Resources of the
National Institutes of Health (PHS 5 P41-RR03155) and UIUC.
Any opinions, findings and conclusions or recommendations
expressed in this publication are those of the author(s) and do
not necessarily reflect the views of the US Department of Energy or
the National Institutes of Health.
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(Received September 2, 2007; Accepted October 8, 2007)