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Plant Physiol. (1999) 120: 205-216
Identification of the Soluble Starch Synthase Activities of
Maize Endosperm1
Heping Cao,
Jennifer Imparl-Radosevich,
Hanping Guan,
Peter L. Keeling,
Martha G. James, and
Alan M. Myers*
Department of Biochemistry, Biophysics, and Molecular Biology, Iowa
State University, Ames, Iowa 50011 (H.C., M.G.J., A.M.M.); and ExSeed
Genetics, L.L.C., and Department of Agronomy, Food Sciences
Building, Iowa State University, Ames, Iowa 50011 (J.I.-P., H.G.,
P.L.K.)
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ABSTRACT |
This study identified the complement
of soluble starch synthases (SSs) present in developing maize
(Zea mays) endosperm. The product of the
du1 gene, DU1, was shown to be one of the two major soluble SSs. The C-terminal 450 residues of DU1 comprise eight sequence
blocks conserved in 28 known or predicted glucan synthases. This region
of DU1 was expressed in Escherichia coli and shown to
possess SS activity. DU1-specific antisera detected a soluble endosperm
protein of more than 200 kD that was lacking in du1- mutants. These antisera eliminated 20% to 30% of the soluble SS activity from kernel extracts. Antiserum against the isozyme zSSI eliminated approximately 60% of the total soluble SS, and
immunodepletion of du1- mutant extracts with this
antiserum nearly eliminated SS activity. Two soluble SS activities were
identified by electrophoretic fractionation, each of which correlated
specifically with zSSI or DU1. Thus, DU1 and zSSI accounted for the
great majority of soluble SS activity present in developing endosperm.
The relative activity of the two isozymes did not change significantly
during the starch biosynthetic period. DU1 and zSSI may be
interdependent, because mutant extracts lacking DU1 exhibited a
significant stimulation of the remaining SS activity.
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INTRODUCTION |
Plant starches are composed almost entirely of glucosyl units
joined by either  (1 4) or (16) glycoside bonds. In the
amylopectin component of starch, these two types of linkage are located
with a specific spatial distribution, and this defined branching
pattern in turn determines higher-order structures (Gallant et al.,
1997 ). The number of glucosyl units in each (14)-linked chain and
the locations of (16) branch linkages that join the linear chains are expected to be determined by inherent properties of the starch biosynthetic enzymes. The specific mechanisms by which these enzymes determine architectural specificity, however, are not well understood.
Three enzymatic activities are involved in starch synthesis from the
glucosyl donor ADPGlc (for reviews, see Hannah et al., 1993; Nelson and
Pan, 1995; Ball et al., 1996; Nakamura, 1996; Preiss and Sivak, 1996;
Smith et al., 1997). SS enzymes add glucosyl units at the nonreducing
end of linear chains through new (14) linkages. BEs cleave
interior (14) bonds and form a new glycoside bond between the C1
reducing end of the released glucan and a C6 elsewhere within a linear
chain; thus, BEs introduce branch linkages. Debranching enzymes
hydrolyze branch linkages and are needed for determination of the
normal branching pattern in amylopectin. Multiple isoforms exist for
each enzyme, although their specific roles in starch biosynthesis
generally are not known.
In maize (Zea mays), at least five different genes code for
predicted SSs. The wx locus codes for the major
granule-bound SS, GBSSI (Shure et al., 1983; Klösgen et al.,
1986), and the du1 gene codes for a protein similar in
sequence to known SSs (Gao et al., 1998). In addition to these two
genetically defined loci, three different cDNAs are known that code for
the SSs designated zSSI, zSSIIa, and zSSIIb (Harn et al., 1998;
Imparl-Radosevich et al., 1998; Knight et al., 1998). Genetic and
biochemical analyses reveal that individual SS isozymes have specific
effects on starch structure, influencing both chain length and
branch-linkage formation. GBSSI is known to synthesize the amylose
component of starch (Martin and Smith, 1995; Denyer et al., 1996; van
de Wal et al., 1998). Alteration of DU1 causes changes in amylopectin
structure, including increased branching frequency (Shannon and
Garwood, 1984; Wang et al., 1993a, 1993b), and mutations affecting a
Chlamydomonas reinhardtii SS have similar effects
(Fontaine et al., 1993). Mutations of the pea rug5 locus,
which affects SSII, alter starch granule structure and chain-length
distribution (Craig et al., 1998). Maize du1- mutations and
pea rug5 mutations also condition pleiotropic effects on
other starch biosynthetic enzymes (Boyer and Preiss, 1981; Craig et
al., 1998). Thus, various SSs and BEs may act in a concerted manner to
determine starch structure.
An important step in understanding the specific functions of each SS
isozyme in maize starch biosynthesis would be to determine which of the
known cDNAs code for the major soluble enzyme activities present during
endosperm development. Two SS-activity peaks, designated SSI and SSII,
have been fractionated biochemically from soluble extracts of maize
endosperm (Boyer and Preiss, 1981; Preiss and Sivak, 1996).
The enzyme responsible for the SSI-activity peak was purified and
identified as the product of the Ss1 cDNA; this enzyme is designated
zSSI (Mu et al., 1994; Knight et al., 1998). The protein responsible
for the SSII-activity peak has not yet been identified, however,
characterization of the Du1 cDNA provided indirect evidence that DU1 is
this enzyme (Gao et al., 1998). Specifically, the SSII-activity peak is
significantly reduced in du1- mutants (Boyer and Preiss,
1981), and the predicted size of DU1 and the tissue-specific expression
pattern of the Du1 cDNA match the characteristics of this enzyme (Dang
and Boyer, 1988; Mu et al., 1994; Gao et al., 1998). Whether DU1
accounts for a significant portion of soluble SS activity is not clear,
however, because du1- mutations do not cause decreased total
soluble SS activity in endosperm extracts (Singletary et al., 1997).
This study used recombinant proteins and antibodies generated through
application of the cloned Du1 and Ss1 cDNAs to investigate in greater
detail the correlation between known cDNAs and the specific soluble SSs
present in maize endosperm during starch biosynthesis. DU1 was shown
directly to possess SS activity and to account for a significant
proportion of the total activity in soluble kernel extracts.
Essentially all of the soluble SS activity was accounted for by the
combination of DU1 and zSSI. The fact that du1- mutations do
not affect total SS activity was explained by the observation that zSSI
is more active in mutant endosperm extracts than in nonmutant extracts.
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MATERIALS AND METHODS |
Biological Materials, Nomenclature, and Media
Maize (Zea mays) nomenclature follows the standard of
Beavis et al. (1995); in addition, cDNA names are not italicized to distinguish them from gene names. Maize inbreds used were W64A and
Oh43. Plants homozygous for du1-Ref (Mangelsdorf,
1947) or du1-R4059 (Gao et al., 1998) were the
progeny of at least four successive backcrosses to W64A.
Escherichia coli strains used for cloning and recombinant
protein expression were XL-1-Blue, DH5, and BL21(DE3) (Ausubel et
al., 1989). Standard Luria-Bertani medium (Ausubel et al., 1989) was
supplemented with 50 µg/mL kanamycin or 100 µg/mL ampicillin to
make Luria-Bertani K or Luria-Bertani A medium, respectively.
Computational Methods
Multiple sequence alignment used the PILEUP program of the GCG
Sequence Analysis Software Package (Genetics Computer Group, Madison,
WI). Conserved sequence motifs were assigned based on the presence of
at least one invariant residue in the 28 polypeptides analyzed and
conservative substitutions of several nearby residues in the same
positions in the alignment among all of the sequences. Consensus
sequences were determined by votes according to the method of Posfai et
al. (1989). Abbreviations as used in Table I and accession numbers are as follows:
Aae (Aquifex aeolicus) GS, AE000704; At (Arabidopsis
thaliana) SS, AL021713; Atu (Agrobacterium tumefaciens)
GS, L24117; Bst (Bacillus stearothermophilus) GS, D87026;
Bsu (Bacillus subtilis) GS, Z25795; Eco (E. coli)
GS, J02616; Hin (Hemophilus influenzae) GS, P45179; Hv
(Hordeum vulgare; barley) GBSSI, X07932; Ib (Ipomoea
batatas; sweet potato) GBSSI, U44126; Me (Manihot
esculenta; cassava) GBSSI, X74160; Os (Oryza sativa;
rice) GBSSI, X53694; SSS, D16202; Ps (Pisum sativum; pea)
GBSSI, X88789; SSII, X88790; Sb (Sorghum bicolor; sorghum)
GBSSI, U23945; St (Solanum tuberosum; potato) GBSSI, X58453;
GBSSII, X87988; SSSI, Y10416; SSSIII, X94400 and X95759; Syn
(Synechocystis sp. PCC6803) GS, D90899; Ta (Triticum
aestivum; wheat) GBSSI, X57233; SS, U66377; SSS, U48227; and Zm
(Z. mays; maize) DU1, AF023159; GBSSI, M24258; SSI,
AF036891; SSIIa, AF019296; SSIIb, AF019297.
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Table I.
Conserved sequence motifs of the ADPGlc-dependent
-1,4-glucosyltransferases from plants and prokaryotes
The plant enzymes are divided arbitrarily into three classes
based on their known subcellular locations or, in instances of
uncharacterized proteins, their high degree of sequence similarity to
such characterized enzymes. Soluble enzymes are those such as DU1 that
are exclusively or nearly exclusively located in the soluble phase.
Granule-bound enzymes are those such as GBSSI, i.e. the product of a
wx gene, that are exclusively found in the granule fraction.
Dual-location enzymes are those such as zSSI that are present in
significant amounts in both the granule-associated and soluble
fractions. Conservative substitutions are noted when they fall into the
functional groups defined by Dayhoff and Orcutt (1979), which are
AGPST, ILMV, HKR, DENQ, FWY, and C. Invariant residues are denoted by
asterisks under the plant SS consensus sequence, and rare exceptions to
the designated consensus are underlined. Numerals refer to amino acid
positions beginning at the first ATG codon of the open reading frame;
exceptions are Ta SSS and Ta SS, for which the complete cDNA sequences
are not available. The number of nonconserved amino acids adjacent to
each conserved motif is indicated. Full references for each sequence
are listed in ``Materials and Methods''.
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Construction of Expression Plasmids
Plasmid pHC1 was constructed as an intermediate in the generation
of the antigens used to raise the antisera anti-DU1N and anti-DU1F.
This plasmid contains the entire Du1 cDNA coding region delineated by
two EcoRI sites, one located immediately upstream of the
presumed initiation codon, and the second 225 bp downstream of the
termination codon. A 1.5-kb fragment was PCR amplified from
the partial Du1 cDNA clone pMgf10 (Gao et al., 1998) using primers HCp1
and M13F. HCp1
(5 -AAACCCGGGAATTCGATGGAGATGGTCCTACG-3) contains SmaI and EcoRI sites located
upstream of the presumed initiation codon (restriction sites and the
initiation codon are underlined), and M13F is located downstream of the
cDNA insert on the noncoding strand. The amplified fragment was cleaved
at the SmaI site of the primer and the unique
AgeI site within the cDNA sequence. This fragment was cloned
in the SmaI and AgeI sites of pMg10-6, which
contains the Du1 cDNA extending from 125 bp upstream of the presumed
initiation codon to the downstream EcoRI site (M. Gao,
unpublished results). The resulting plasmid is pHC1. DNA sequence
analysis indicated that no mutations arose in the Du1 cDNA during
construction of this plasmid.
Plasmid pHC2 expresses a fusion protein containing the
Schistosoma japonicum glutathione-S-transferase
protein at its N terminus and DU1 residues 1 to 648 at its C terminus;
this polypeptide was used as the DU1N antigen. pHC2 was constructed by
cloning the EcoRI-SalI fragment from pHC1 into
pGEX4T-3 (Pharmacia) digested with the same enzymes. Plasmid pHC4
expresses a fusion protein containing thioredoxin at its N terminus and
full-length DU1 at its C terminus; this protein was used as the DU1F
antigen. pHC4 was constructed by cloning the EcoRI fragment
from pHC1 into pET-32b(+) (Novagen, Madison, WI).
Plasmids pHC5 and pHC6 express the C-terminal region of DU1 (DU1C) in
E. coli. The Du1 cDNA from codon 1226 to termination codon
1675 was PCR amplified using pHC1 as the template. The upstream primer
was HCp2 (5-GCAGAATTCGATGCACATTGTCCAC-3),
which places an EcoRI site adjacent to codon 1226 (the
EcoRI site and codon 1226 are underlined). The downstream
primer was M13F. The amplified fragment digested with EcoRI
was cloned into pET-29b(+) and pET-32b(+) (Novagen) to form pHC5 and
pHC6, respectively. The sequence of the entire DU1C insert and the
junction with the T7 promoter was determined in clones with correct
restriction maps. These data confirmed that no mutations arose in the
Du1 cDNA sequence during construction of the plasmids.
Antigen and Antibody Production
To produce the DU1N antigen, 1-L exponential-phase cultures of
E. coli cells containing pHC2 were grown for 2 h at
37°C in the presence of 0.1 mM IPTG. Cells were
collected by centrifugation, and the pellet (7 g wet weight, from 2 L
of culture) was suspended in 100 mL of 140 mM
NaCl, 2.7 mM KCl, 10 mM
Na2HPO4, 1.8 mM
KH2PO4, 1 mM PMSF, 0.01 mM
trans-epoxysuccinyl-L-leucylamido-(4-guanidino)butane, 10 mM EDTA, 5 mM DTT, and 1 mg/mL lysozyme; all subsequent treatments were at 0°C. Cells were
lysed by sonication. The DU1N fusion protein was affinity purified
using glutathione-agarose beads, essentially as described previously
(Rahman et al., 1998). The fusion protein was eluted in 100 mM Tris-HCl, pH 8.0, 120 mM
NaCl, and 20 mM glutathione.
To produce the DU1F antigen, 0.5-L exponential-phase cultures of
E. coli cells containing pHC4 were grown for 1.5 h at
37°C in the presence of 0.5 mM IPTG. Cells were
collected by centrifugation, suspended in 25 mL of 50 mM Tris-HCl, pH 7.0, 1 mM
PMSF, 10 mM EDTA, 5 mM DTT,
10% glycerol, and 3% 10× proteinase inhibitor cocktail (Sigma no.
P8465), and broken by sonication. Lysates were centrifuged at
10,000g for 10 min, and the pellets were dissolved by
boiling for 10 min in 1× SDS-PAGE sample buffer. A band of more than
200 kD was observed during SDS-PAGE that was specific to cells
containing pHC4 and reacted with anti-DU1N in immunoblot analysis (data
not shown). This protein, therefore, was identified as the DU1F
antigen. The DU1F antigen band was cut out of large-scale 6%
polyacrylamide gels, crushed to a powder, and used for immunization.
Antisera were raised in rabbits by standard procedures (Harlow and
Lane, 1988). For the initial immunization with the DU1N antigen, 300 µg of protein was injected in complete Freund's adjuvant. Booster
immunizations of 200 µg of fusion protein were supplied three times
at 3-week intervals. Immunization with DU1F followed a similar
protocol, except that approximately 50 µg of antigen was supplied in
all four injections.
Expression of DU1C in E. coli
E. coli strain BL21(DE3) containing pHC5 or pHC6 was
grown in Luria-Bertani K or Luria-Bertani A medium, respectively.
Overnight cultures were inoculated into fresh medium at a 1:10 dilution and grown at 37°C until the density was 0.8 A600/mL. IPTG was added to 0.5 mM and the cultures were grown for 5 h at
25°C. Cells were collected by centrifugation, suspended in
one-twentieth culture volume of sonication buffer (50 mM Tris-HCl, pH 7.0, 10% glycerol, 10 mM EDTA, 5 mM DTT, and 3%
10× proteinase inhibitor cocktail [Sigma no. P8465]), and broken by
sonication. Lysates were cleared by centrifugation in a microfuge, and
the supernatants were used for subsequent analyses. The S-tag Rapid
Assay kit (Novagen) was used for detection of S-tag sequences by
measurement of reconstituted RNase A activity.
Zymogram Analysis
Zymogram analysis was performed essentially as described by
Buléon et al. (1997) with a few modifications. Endosperm
from three to four kernels was frozen in liquid
N2, crushed to a fine powder, and suspended by
vortexing in 50 mM Tris acetate, pH 8.0, 10 mM
EDTA, and 5 mM DTT (1 mL/g kernel fresh weight). The crude homogenate was cleared by centrifugation at 10,000g for 10 min at 4°C, and protein concentration in the supernatant was
determined. Protein samples (225 µg) were boiled in SDS-PAGE buffer
(65 mM Tris-HCl, pH 6.8, 2% SDS, 10% glycerol,
and 5% 2-mercaptoethanol) and loaded onto an 8% acrylamide gel
containing 0.1% glycogen. Electrophoresis was performed under
denaturing conditions (25 mM Tris-HCl, pH 8.3, 192 mM Gly, 0.1% SDS, and 5 mM DTT) for 3 h at 4°C at 80 V in a
Bio-Rad Mini-Protean II cell. The gel was washed four times for 30 min
each at room temperature in 40 mM Tris-HCl, pH
7.0, and 5 mM DTT to remove SDS and allow
proteins to renature. The gel was then incubated in reaction buffer
(100 mM Bicine, pH 8.0, 0.5 M citrate, 25 mM potassium
acetate, 0.5 mg/mL BSA, 5 mM ADPGlc, 5 mM 2-mercaptoethanol, and 20 mg/mL glycogen) for
36 h at room temperature. Enzyme activities were detected by the
addition of iodine stain (0.2% iodine and 2% potassium iodide in 10 mM HCl), and the zymograms were photographed
immediately.
Fractionation of Maize Kernel Extracts and Glucan Synthase Assays
Kernels were collected from developing ears, immediately frozen in
liquid N2, and stored at  80°C. Frozen kernels
were ground on ice with a mortar and pestle in homogenization buffer
(50 mM Tris-HCl, pH 7.0, 10% glycerol, 10 mM
EDTA, 5 mM DTT, 1 mM PMSF, and 50 µL/g tissue
10× proteinase inhibitor cocktail [Sigma no. P2714]; total,
2.5 mL/g tissue). The homogenate was centrifuged at 10,000g
for 10 min, and the supernatant was used for SS assays and
determination of protein concentration. To obtain starch granules, the
10,000g pellet was vortexed vigorously in homogenization
buffer and centrifuged again. The pellet from the third wash was
suspended in homogenization buffer and used as the starch granule
fraction.
Glucan synthase assays were performed in microfuge tubes in a total
volume of 0.1 mL. The standard reactions contained 100 mM
Bicine-NaOH, pH 8.0, 5 mM EDTA, 0.5 M sodium
citrate, 0.5 mg/mL BSA, 10 mg/mL glycogen, 1 mM
ADP-[14C]Glc (150 cpm/nmol; catalog no. CFB144,
Amersham), and various amounts of total soluble extract. Reactions were
initiated by the addition of the labeled ADPGlc, incubated for 30 min
at 30°C, and terminated by the addition of 1 mL of 75% methanol/1%
KCl. Incorporation of radioactive label into methanol-insoluble glucan was determined according to the method of Cao and Preiss (1996). All
assays were performed in duplicate or triplicate, and the maximal
observed variation was approximately 10%. Preliminary experiments
demonstrated that the amount of 14C incorporated
into methanol-precipitable glucan is linear with the amount of protein
in the assay. Furthermore, approximately 10% of the
14C in the assay was recovered in insoluble
glucan. Thus, the assays were performed in conditions of substrate
excess.
Some assays varied from the standard procedure by the omission of
glycogen and/or sodium citrate. When glycogen was omitted from the
assay, it was added to the standard concentration after the reaction
was stopped by the addition of methanol.
Immunoblot and Immunodepletion Methods
Protein concentrations were determined according to the method of
Bradford (1976). SDS-PAGE and transfer of protein from the gels to
nitrocellulose filters followed standard methods (Sambrook et al.,
1989). The primary antisera were anti-SSI (Mu et al., 1994) diluted
1:1,000 or 1:3,000, anti-DU1N diluted 1:10,000 or 1:75,000, and
anti-DU1F diluted 1:2,000. The secondary antibody was goat anti-rabbit
IgG-alkaline phosphatase conjugate (Bio-Rad) diluted 1:3,000, which was
detected using the 5-bromo-4-chloro-3-indolyl phosphate/nitroblue
tetrazolium reagent system (Bio-Rad). Fusion proteins containing the
S-tag amino acid sequence were detected by the same procedure, except
that S-protein-alkaline phosphatase conjugate (Novagen) diluted 1:5,000
was used instead of a primary antibody.
Immunodepletion experiments were performed as follows. Total soluble
kernel extracts (50 µL) were mixed with an equal volume of serum. The
solutions were incubated on ice for 90 min with gentle mixing every 10 to 15 min, after which 10% (w/v) protein A-Sepharose CL-4B (Sigma) was
added. The mixtures were then gently shaken continuously for 30 min and
centrifuged for 10 min at 10,000g, and the supernatants were
assayed for SS activity. The pellets were washed with buffer three
times before immunoblot analysis of the precipitated proteins.
Preliminary experiments in which the volume of immune serum added to
the assays was varied were used to ensure that these conditions were
saturating for the amount of antibody present (data not shown).
SS-activity values obtained after treatment with preimmune serum were
taken as 100%.
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RESULTS |
Sequence Motifs Conserved in DU1 and SSs
Three conserved sequence blocks identified previously in
comparisons of various GBSSI proteins and E. coli GS (van
der Leij et al., 1991; Preiss and Sivak, 1996) were present in the DU1 C-terminal region (Gao et al., 1998). This comparative analysis was
extended to include 28 SS or GS sequences from 17 species (Table I).
Thirty-three residues are conserved in all 28 enzymes. Five conserved
sequence motifs were identified in addition to the three noted
previously. The eight conserved sequence blocks are designated motifs I
to VIII, in order from the N to the C termini; according to this
notation, motifs I, VII, and VIII correspond to regions I, II, and III,
respectively, as designated previously (Preiss and Sivak, 1996). The
conserved sequences are distributed in the 359 residues of DU1 between
positions 1237 and 1595.
Recombinant DU1 Exhibits SS Activity
The 449 C-terminal residues of DU1 (positions 1226-1674;
designated DU1C) were expressed in E. coli from plasmids
pHC5 or pHC6. These plasmids are based in the expression vector
pET-29b(+) or pET-32b(+), respectively, and thus produce DU1C fusion
proteins containing either 35 or 167 plasmid-derived residues at their N termini. Expression of the fusion proteins was monitored by enzymatic
and immunoblotting analyses that detected the S-tag sequence present in
these N-terminal extensions. Proteins of the expected sizes were
expressed specifically when the DU1C coding region was present (Fig.
1).
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| Figure 1.
Expression of DU1C in E. coli. Gene
expression from the T7 promoter of the indicated plasmid was induced in
exponential-phase E. coli cells. Total soluble lysates
were fractionated by SDS-PAGE, and specific proteins containing the
S-tag sequence (specified by the pET plasmid) were detected by
S-protein-alkaline phosphatase conjugate. Lane 1, pET-32b; lane 2, pHC6
(DU1C in pET-32b); lane 3, pET-29b; and lane 4, pHC5 (DU1C in pET-29b).
Asterisks indicate polypeptides of approximately the size predicted
from the plasmid and Du1 cDNA sequences, which are present only when
the DU1C coding region is contained within the plasmid.
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Increased glucan synthase activity was observed in total soluble
extracts of E. coli cells expressing DU1C. Cells containing pHC5 or pHC6 were exposed to IPTG to induce expression of the DU1C
proteins, and total soluble extracts were tested for glucan synthase
activity. DU1C expression resulted in approximately 5-fold increased SS
activity compared with control cells lacking the maize coding region
(Table II). A similar increase also
occurred in the reconstituted RNase A activity conferred by the S-tag
sequence of the N-terminal extension (data not shown). Nearly identical results were obtained when DU1C was expressed in pET-29b(+) or pET-32b(+). The activity increase relative to the endogenous level was
relatively modest, although similar levels were detected also for zSSI
expressed in E. coli (Knight et al., 1998). In addition, the
level of recombinant enzyme activity observed for DU1C was comparable
to that of potato GBSSII expressed in a similar system (Edwards et al.,
1995). These data provide direct evidence that DU1 is a SS and that its
C-terminal 449 residues are sufficient to provide this enzymatic
activity.
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Table II.
Glucan synthase activity in E. coli soluble
extracts
Gene expression was induced for 5 h in the exponential phase of
E. coli cells transformed with the indicated plasmid. Total
soluble extracts were assayed for SS activity in the presence of
citrate and glycogen primer. Values indicate means ± SE (n = 4).
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Immunological Detection of DU1 in Kernel Extracts
Detection of DU1 in kernel extracts revealed the apparent size of
this SS, its temporal expression pattern, and its lack of association
with starch granules. The polyclonal antiserum anti-DU1N was raised in
rabbits against the N-terminal 648 residues of DU1. This region of DU1
is unique among known protein sequences (Gao et al., 1998), so
anti-DU1N is expected to react specifically with DU1 and not with other
SSs. Figure 2a shows that in immunoblot analysis of total soluble kernel extracts (i.e. the 10,000g
supernatant) from nonmutant kernels, anti-DU1N detected a protein that
migrated at an apparent molecular mass of more than 200 kD. This
protein was missing in two different du1- mutants. In
kernels homozygous for the reference mutation
du1-Ref, a smaller immunoreactive protein was
detected, whereas in kernels homozygous for the presumed
transposon-induced allele du1-R4059 (Gao et al.,
1998), the protein was completely eliminated (Fig. 2a). Identical
results were obtained using a different antiserum, anti-DU1F, which was
raised against full-length DU1 (data not shown). Thus, both anti-DU1N
and anti-DU1F recognized DU1, the product of the du1 gene.
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| Figure 2.
Immunological detection of DU1 and SSI in kernel
extracts. a, Total soluble extracts from 20-DAP kernels of the W64A
genetic background homozygous for the indicated allele were
fractionated by SDS-PAGE and probed with anti-DU1N or anti-SSI. An
equal amount of protein was loaded in each lane. du1-M5 indicates the
allele du1-R4059. The asterisk indicates
full-length DU1. b, Extracts of nonmutant W64A kernels and congenic
du1-Ref mutant kernels collected 20 DAP
were separated into granule (i.e. 10,000g pellet) and
total soluble fractions (i.e. 10,000g supernatant). An
equal volume of each fraction was separated by SDS-PAGE;
therefore, each pair of lanes is standardized to kernel fresh weight.
The samples were probed with anti-DU1N or anti-SSI, as indicated. c,
Total soluble extracts of W64A kernels harvested at various times after
pollination (as indicated) were analyzed by SDS-PAGE and immunoblot
analysis using anti-DU1N or anti-SSI.
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The zSSI protein of apparently 76 kD also was identified in immunoblot
analysis of these same kernel extracts, using anti-SSI antiserum (Fig.
2a). Anti-DU1N did not recognize SSI, and anti-SSI did not recognize
DU1. Therefore, in this assay, both antisera reacted specifically with
a distinct isozyme.
DU1 was found to be located primarily in the soluble fraction of kernel
extracts, as opposed to being associated with starch granules. Kernels
harvested 20 DAP were fractionated into soluble and granule fractions.
The identity of the granule fraction was verified by enrichment for
zSSI (Fig. 2b), which is known to be both granule associated and
soluble (Mu et al., 1994). The amount of DU1 present in the granule and
soluble fractions was determined by immunoblot analysis of protein
samples standardized based on kernel fresh weight. In contrast to zSSI,
the anti-DU1N signal was found almost exclusively in the soluble
fraction (Fig. 2b), indicating that DU1 is not stably associated with
starch granules in 20-DAP endosperm.
The temporal expression pattern of DU1 and SSI in kernels at various
times after pollination was monitored. DU1 was detected first at 12 DAP
and was maintained at a nearly constant level throughout the period of
starch biosynthesis for up to at least 32 DAP (Fig. 2c). Anti-SSI
produced a signal in the 8-DAP kernel extract (Fig. 2c), indicating
that in these tissue samples zSSI was expressed earlier than DU1.
Immunodepletion of SS Activity in Kernel Extracts
Immunodepletion experiments investigated the amount of SS activity
in endosperm provided by DU1 and zSSI. Total soluble extracts of
kernels harvested 20 DAP were treated with anti-DU1N, anti-DU1F, anti-SSI, or preimmune serum. Immune complexes were then removed from
solution after binding to protein A-Sepharose beads. Residual SS
activity remaining in the supernatant was determined in the presence of
citrate and exogenous primer, conditions known to yield maximal
activity of these enzymes (Preiss and Sivak, 1996). Preliminary
experiments titrated the amount of serum; the following data were
obtained in conditions of antibody excess. Nonmutant extracts of either
the W64A or Oh43 background were depleted of approximately 28% of
their total SS activity by either of the two anti-DU1 sera (Fig.
3). Anti-SSI depleted 66% and 61% of
the total SS activity in the two genotypes, respectively. Treatment of
a du1- mutant extract with either of the two anti-DU1 sera had virtually no effect on the total SS activity, suggesting that the
particular enzyme affected by these antibodies is specifically that
coded for by Du1. In contrast, anti-SSI treatment depleted virtually all of the SS activity present in the du1- mutant
extract, which suggests that the great majority of SS activity in the
soluble fraction of 20-DAP endosperm is provided by a combination of
zSSI and DU1.
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| Figure 3.
Immunodepletion of SS activity. Total soluble
extracts from kernels of the indicated genotype collected 20 DAP were
treated with preimmune serum or saturating amounts of the indicated
antiserum, and residual SS activity was assayed after removal of the
immune complexes. The du1-Ref mutant was
in the W64A genetic background. SS activity remaining after treatment
with preimmune serum was defined as 100%. These values were 7.0 nmol
min 1 mg1 for W64A, 12.9 nmol
min1 mg1 for the
du1-Ref mutant, and 16.4 nmol
min1 mg1 for Oh43.
|
|
This analysis was applied to kernel extracts from nonmutant W64A plants
collected 12, 18, or 32 DAP and also to an independently prepared
20-DAP extract. In these experiments, the range of total SS activity
neutralized by anti-DU1N ranged from 17% to 24%, and anti-SSI
treatment depleted 47% to 66% of the activity (data not shown). The
relative abundance of the DU1 and zSSI activities, therefore, did not
change significantly during the period of starch biosynthesis.
Fractionation of SS Activities in Total Endosperm Extracts
The SS activities present in 20-DAP endosperm also were correlated
with particular cloned cDNAs by a combination of zymogram, immunoblot,
and mutational analyses. These SSs were fractionated by SDS-PAGE and
detected by their activity in gels after protein renaturation. Two
activity bands were observed, one of more than 200 kD and the other of
approximately 76 kD (Fig. 4a). The sizes of these isozymes correlate roughly with those predicted by the Du1
cDNA and the Ss1 cDNA, respectively (Gao et al., 1998; Knight et al.,
1998). Immunoblot analysis of the same protein samples revealed that
the >200-kD isozyme reacted with anti-DU1N, whereas the 76-kD isozyme
reacted with anti-zSSI (Fig. 4b). Extracts from du1- mutant
endosperm entirely lacked activity of the >200-kD isozyme. These
results support the conclusions of the previous section: (a) that there
are two major soluble SSs present in developing endosperm cells, and
(b) that one of these is DU1, the product of the du1 gene,
and the other is zSSI, the product of the Ss1 cDNA.
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| Figure 4.
Specific identification of SS isozymes. a, SS
activity zymogram. Proteins in total soluble endosperm extracts were
separated based on molecular mass by SDS-PAGE and then allowed to
renature in the gel. SS substrates were provided to the entire gel, and
positions of glucan synthesis were detected by staining with iodine.
Two congenic strains in the W64A genetic background were analyzed, one
bearing the nonmutant allele Du1 and the other
containing du1-Ref (indicated as
du1-). Two SS activities are evident in the nonmutant
endosperm, one of which is missing from the
du1-Ref extract. b, Immunoblot analysis.
Proteins in duplicates of the gel shown in a were transferred to
nitrocellulose paper and probed with the indicated antiserum. A
polypeptide of the same mobility and genetic specificity as the larger
SS activity is recognized by anti-DU1N, whereas a protein of the same
mobility as the smaller SS activity is recognized by anti-SSI.
|
|
Increased Total SS Activity in du1- Mutant Extracts
The conclusion that du1 specifies a SS appears to be
inconsistent with a previous report indicating that soluble SS activity is not decreased in a du1- mutant. To the contrary, the
total soluble SS activity was found to be increased approximately
2-fold in du1-Ref mutant extracts (Singletary et
al., 1997); this observation was confirmed independently in the current
study (Fig. 5). Congenic strains were
analyzed, ruling out genetic background differences as the explanation
for the different total SS levels. A possible explanation for this
phenomenon is that a SS other than DU1 is hyperactive in
du1- mutants. To test this possibility, SS activity in total
soluble kernel extracts was assayed in the presence or absence of
citrate and/or exogenous glucan primer. These experiments were intended
to differentiate between zSSI, which is known to be stimulated
significantly by citrate and to be independent of exogenous primer, and
the enzyme accounting for the SSII-activity peak, which is primer
dependent and largely citrate independent (Boyer and Preiss, 1981).
View larger version (30K):
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| Figure 5.
SS activity in total soluble kernel extracts.
Total soluble extracts from kernels of the indicated genotype collected
20 DAP were assayed for SS activity in the presence or absence of
exogenous primer (10 mg/mL glycogen) and 0.5 M citrate, as
indicated. The du1-Ref mutant was in the
W64A genetic background.
|
|
Citrate-stimulated, primer-independent SS activity was increased
approximately 8-fold in du1-Ref mutant extracts
compared with nonmutant extracts (Fig. 5). Similar results were
obtained for six other independent du1- alleles (data not
shown). The immunodepletion data described above indicated that the
only SS remaining in du1- mutants was zSSI (Fig. 3). Thus,
it appears that the activity of zSSI was increased in du1-
mutants. The level of zSSI protein detected by immunoblot analysis
appeared to be slightly increased in du1- mutant kernels
relative to nonmutants, either when the extracts were compared on a
total protein basis (Fig. 2a) or when they were standardized by the
fresh weight of the tissue used to prepare the extracts (Fig. 2b). This
apparent increase, however, is significantly less than the 8-fold
increase in enzyme activity. Thus, the elevated zSSI activity in
du1- mutant extracts most likely results in part from the
increased specific activity of the enzyme.
| |
DISCUSSION |
Important goals in the understanding of the mechanisms of starch
biosynthesis include identification of the particular SS isozymes
active during endosperm development and determination of the specific
functions of each enzyme. Multiple soluble SSs are present in
endosperm, as was shown initially by biochemical fractionation. Two
activity peaks were observed, designated SSI, which does not require
exogenous glucan primer and is stimulated by citrate, and SSII, which
is dependent on exogenous primer and is largely insensitive to citrate
(enzyme designations according to Boyer and Preiss, 1981). Five
different cDNA clones are known that code for SSs, however, so it is
necessary to correlate each enzymatic activity with a particular
genetic element.
The cDNA and protein that account for the SSI-activity peak were
identified recently; however, the protein(s) responsible for the second
SS had not been clearly assigned before this study. An apparent 76-kD
protein copurified with the SSI activity (Mu et al., 1994). Sequence
information indicated that the Ss1 cDNA codes for this polypeptide, and
this cDNA directed expression of an active SS that is immunologically
cross-reactive with the purified enzyme (Imparl-Radosevich et al.,
1998; Knight et al., 1998). Thus, the genetic element responsible for
synthesis of zSSI has now been identified. Presumably, at least one
other protein provides additional SS activity in the soluble fraction,
because of the distinct enzymatic characteristics and apparent
Mr of the enzyme responsible for the
SSII-activity peak. Detailed characterization of this second enzyme is
lacking because it has proven difficult to purify.
The gene du1 was proposed to code for a soluble SS activity
based in part on the facts that du1- mutants lack the SSII
activity (Boyer and Preiss, 1981) and that Du1 codes for a
protein similar in sequence to known SSs (Table I). This study confirms
the identification of DU1 as an SS active in developing endosperm.
Expression of the DU1 C terminus correlated with induction of SS
activity, and DU1-specific antibodies immunodepleted a significant
portion of the enzyme present in kernel extracts. Furthermore, a
specific SS enzyme activity identified by zymogram analysis migrated in SDS-PAGE at the same rate as DU1 and was missing in a du1-
mutant. The SS activity of DU1 resides within the C-terminal 450 residues; the function(s) of the remaining 1224 residues remains to be
determined.
Inferences drawn from the immunodepletion data presume that anti-DU1N
is specific for DU1. Immunological specificity was indicated by three
observations. First, in immunoblot analysis, anti-DU1N failed to detect
zSSI (and anti-SSI failed to detect DU1). Second, when du1-
mutant extracts were treated with anti-DU1N, there was no decrease in
residual SS activity, even though anti-SSI treatment of the same
extracts reduced the activity almost completely. Thus, anti-DU1N did
not neutralize zSSI. Third, the anti-DU1 and anti-SSI immunoprecipitates were analyzed by immunoblotting using both antisera;
the anti-DU1N complexes did not contain zSSI, and visa versa (H. Cao,
unpublished results).
DU1 and zSSI most likely account for all of the soluble SS activity in
developing kernels. Only two enzymes were observed in zymograms, and
each of these could be correlated with either DU1 or zSSI. The fact
that the combined effects of a du1- mutation and anti-zSSI
immunodepletion caused nearly complete loss of SS activity suggests
that these are the only two isozymes present in the soluble fraction.
Consistent with this conclusion is the fact that antibodies reactive
with either of the remaining known SSs, zSSIIa or zSSIIb (Harn et al.,
1998), failed to detect polypeptides in soluble extracts of 20-DAP
kernels (J. Imparl-Radosavich and H. Cao, unpublished results). There
is one inconsistency with the idea that DU1 and zSSI account for all
soluble SS activity, which is the fact that the amount of enzyme
neutralized by anti-DU1 and anti-SSI sera together is about 90% of the
total (Fig. 3). A possible explanation for this observation is that the
anti-DU1 sera are not capable of binding all of the DU1 present,
perhaps as a result of partial degradation of the protein.
Alternatively, one or more additional soluble SSs might exist that are
not detected in the zymograms, either because they fail to recover
activity after denaturation or they comigrate with zSSI. We consider
the latter possibility unlikely, because if an additional enzyme
exists, it should be evident after treatment of the du1-
mutant extract with anti-SSI. Despite these arguments, the data in this
study certainly do not exclude the possibility that SSs in addition to
DU1 and zSSI exist as minor activities in the soluble fraction of maize
endosperm.
This study supports the hypothesis that DU1 accounts for the
SSII-activity peak (Boyer and Preiss, 1981). The fact that there are
two SS peaks in anion-exchange chromatography and two enzymes in the
zymograms is most simply explained by a direct correspondence. Such a
correspondence is further indicated by molecular mass comparisons: immunoblots indicated that DU1 has a mass of more than 200 kD, and the
native size of the protein responsible for the SSII-activity peak was
estimated to be 180 kD (Mu et al., 1994). The >200-kD protein detected
by anti-DU1N was not present in du1- mutants. Most
tellingly, zymogram analysis revealed the existence of a >200-kD SS
that is missing in du1- mutants, as is the case also for the
SSII-activity peak (Boyer and Preiss, 1981). All of these diverse
observations would be explained if DU1 were the active SS enzyme
present in the SSII-activity peak. The correspondence between the
molecular mass determined for the native enzyme in the SSII-activity
peak, the size of the SS identified as DU1 separated in denaturing
zymograms, and the size of DU1 predicted by cDNA cloning indicates that
this enzyme functions as a monomer. The proteolytically labile nature
of DU1 may explain the facts that purification of the native SS present
in the SSII-activity peak has been problematic and that different
molecular masses (180 and 95 kD) have been reported (Mu et al., 1994;
Preiss and Sivak, 1996).
Assignment of DU1 as a soluble protein of more than 200 kD was
supported by an independent study (Yu et al., 1998). A protein of this
size was present in the purified amyloplast stromal fraction from
nonmutant plants but was lacking in a du1- mutant. This
protein most likely is the same as the >200-kD SS and the >200-kD
anti-DU1N reactive protein shown here to be absent in du1-
kernels. Taken together, these data indicate that DU1, as expected, is
located within plastids.
DU1 and zSSI share the property that their mobility in SDS-PAGE is
slower than predicted from their cDNA sequences. The Ss1 cDNA predicts
a 64-kD protein, whereas zSSI runs in gels at 76 kD (Knight et al.,
1998). The Du1 cDNA predicts a 188-kD protein; however, DU1 in kernel
extracts runs significantly slower than the 200-kD marker. Anomalous
migration in SDS-PAGE is thought to be an intrinsic property of zSSI
(Knight et al., 1998) and other SSs (Edwards et al., 1995, 1996). The
same phenomenon may apply to DU1, or it could be posttranslationally
modified.
Removal of DU1 from the soluble endosperm fraction apparently causes
some change that results in increased activity of zSSI. A possible
explanation is that DU1 deficiency causes accumulation of a glucan not
present normally, and this provides an efficient primer for zSSI. This
observation explains the fact that total SS activity is not reduced in
du1- mutant extracts (Singletary et al., 1997), even though
a specific SS isozyme is lacking.
The emerging characterization of the complement of maize SSs warrants
some discussion here of their nomenclature in relation to that of the
starch biosynthesis enzymes of other higher plants. The situation is
straightforward with regard to GBSSI, which is the product of the
wx locus and is highly conserved in all plant tissues that
produce storage starch (for review, see Ball et al., 1998). The
original nomenclature for the maize soluble SSs was based on DEAE
chromatography fractions (Boyer and Preiss, 1981; Preiss and
Sivak, 1996), and in this paper we refer accordingly to the SSI- and
SSII-activity peaks. The enzyme responsible for the SSI-activity peak,
termed zSSI, has been characterized by both biochemical purification
and cDNA cloning (Mu et al., 1994; Harn et al., 1998; Imparl-Radosevich
et al., 1998; Knight et al., 1998). zSSI is very similar in primary
sequence to a soluble SS of rice (Baba et al., 1993). It exists both
bound to starch granules and in a soluble form in the amyloplast
stroma, and in this regard it is similar to the enzymes of pea embryos
and potato tubers designated as SSII. However, in terms of amino acid
sequence, zSSI is not the counterpart of pea SSII or potato SSII. The
sequences of the maize enzymes coded for by the zSSIIa and zSSIIb cDNAs (Harn et al., 1998; Knight et al., 1998) are significantly closer to
those of pea or potato SSII (Dry et al., 1992; Edwards et al., 1995)
than is the zSSI sequence. Thus, zSSI-type enzymes have been reported
so far only in monocots, whereas zSSIIa and zSSIIb are members of a
more generally conserved class, along with pea SSII and potato SSII. It
is not known whether this sequence conservation extends to functional
conservation. zSSIIa and zSSIIb are expressed in developing endosperm
not at all or only at very low levels, whereas the SSII enzymes of pea
and potato clearly are present at significant levels during
storage-starch biosynthesis.
We argue above that the SS designated here by virtue of its genetic
identification as DU1 is the enzyme responsible for the SSII-activity
peak. This enzyme is the evolutionary counterpart of potato SSIII (Abel
et al., 1996; Marshall et al., 1996) based on both the very high
sequence identity between these two enzymes and the fact that both
enzymes are exclusively soluble. We propose that the most logical
nomenclature for these enzymes is one based on evolutionary sequence
conservation and species; therefore, we suggest that DU1 should also be
known as zSSIII. In summary, the enzyme coded for by the du1
locus, zSSIII/DU1, is most likely responsible for the maize
SSII-activity peak and is the apparent evolutionary counterpart of
potato SSIII. Based on sequence identity, zSSIIa and zSSIIb appear to
be the evolutionary counterparts of pea SSII and potato SSII, although
they are not highly expressed in developing maize endosperm. zSSI
accounts for the SSI-activity peak, and as yet a distinct evolutionary
counterpart from pea or potato has not been reported. zSSI is distinct
from GBSSI, the product of the wx locus. Finally, the
lowercase "z" in the names designated here indicates that the
species of origin of each enzyme is maize.
The reason that multiple soluble SSs are used in storage-starch
biosynthesis is not known at present. DU1 clearly is distinct from zSSI
in that it is located almost entirely within the soluble phase of
endosperm cells, whereas zSSI is abundant in both the granule and the
soluble fractions (Fig. 2b). The fact that du1- mutations
alter starch structure indicates that DU1 provides a specific
function(s) that cannot be compensated for by zSSI. Similarly, severe
reduction of potato SSIII by antisense RNA expression causes significant changes in granule structure that cannot be compensated for
by the remaining soluble SS activity (Abel et al., 1996; Marshall et
al., 1996). Although the specific functions of each soluble SS remain
to be determined, identification of the genetic sources of the two
major isoforms in maize will provide significant new tools for such
investigations.
| |
FOOTNOTES |
1
This work was supported by the U.S. Department
of Agriculture (grant no. 96-35300-3779 to A.M.M. and M.G.J.) and the
National Science Foundation (grant no. DIR-9113593 to the Iowa State
University Signal Transduction Training Group). This is journal paper
no. J-18330 of project no. 3197 of the Iowa Agriculture and Home
Economics Experiment Station (Ames). This paper is dedicated to the
memory of Dr. Bruce Wasserman.
*
Corresponding author; e-mail ammyers{at}iastate.edu; fax
1-515-294-0453.
Received October 6, 1998;
accepted January 23, 1999.
| |
ABBREVIATIONS |
Abbreviations:
BE, branching enzyme.
DAP, days after
pollination.
GBSS, granule-bound starch synthase.
GS, glycogen
synthase.
IPTG, isopropyl-1-thio- -D-galactopyranoside.
SS, starch synthase.
| |
ACKNOWLEDGMENTS |
We thank Afroza Rahman and Tracie Bierwagen for technical advice
and assistance.
| |
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Abstract
Introduction