The Plant Journal (2005) 41, 815–830
doi: 10.1111/j.1365-313X.2005.02348.x
Arabidopsis mutants Atisa1 and Atisa2 have identical
phenotypes and lack the same multimeric isoamylase,
which influences the branch point distribution of
amylopectin during starch synthesis
Thierry Delatte1, Martine Trevisan1, Mary L. Parker2 and Samuel C. Zeeman1,*
Institute of Plant Sciences, University of Bern, CH-3013, Switzerland, and
2
Institute of Food Research, Norwich Research Park, Norwich NR4 7UA, UK
1
Received 23 November 2004; accepted 9 December 2004.
*
For correspondence (fax þ41 31 631 4942; e-mail [email protected]).
Summary
The aim of this work was to evaluate the function of isoamylase in starch granule biosynthesis in Arabidopsis
leaves. A reverse-genetic approach was used to knockout AtISA1, one of three genes in Arabidopsis encoding
isoamylase-type debranching enzymes. The mutant (Atisa1-1) lacks functional AtISA1 transcript and the major
isoamylase activity (detected by native gels) in crude extracts of leaves. The same activity is abolished by
mutation at the DBE1 locus, which encodes a second isoamylase-type protein, AtISA2. This is consistent with
the idea that ISA1 and ISA2 proteins are subunits of the same enzyme in vivo. Atisa1-1, Atisa2-1 (dbe1), and the
Atisa1-1/Atisa2-1 double mutant all have identical phenotypes. Starch content is reduced compared with
the wild type but substantial quantities of the soluble glucan phytoglycogen are produced. The amylopectin of
the remaining starch and the phytoglycogen in the mutants are structurally related to each other and differ
from wild-type amylopectin. Electron micrographs reveal that the phytoglycogen-accumulating phenotype is
highly tissue-specific. Phytoglycogen accumulates primarily in the plastids of the palisade and spongy
mesophyll cells. Remarkably, other cell types appear to accumulate only starch, which is normal in appearance
but is altered in structure. As phytoglycogen accumulates during the day, its rate of accumulation decreases,
its structure changes and intermediates of glucan breakdown accumulate, suggesting that degradation occurs
simultaneously with synthesis. We conclude that the AtISA1/AtISA2 isoamylase influences glucan branching
pattern, but that this may not be the primary determinant of partitioning between crystalline starch and
soluble phytoglycogen.
Keywords: Arabidopsis, starch synthesis, debranching enzyme, isoamylase, amylopectin structure, glucan.
Introduction
The major component of starch is the branched glucan,
amylopectin. It is responsible for the semi-crystalline nature
of starch granules and consists of chains of a-1,4-linked
glucose residues connected to one another by a-1,6-linkages
(branch points). The distribution of branch points within the
amylopectin molecule gives rise to clusters of adjacent unbranched chains, which are viewed as crucial in allowing
amylopectin to form a semi-crystalline matrix. Precisely how
the starch-synthesizing enzymes determine the molecular
architecture of amylopectin is an open question.
ª 2005 Blackwell Publishing Ltd
Isoamylase-type debranching enzymes (ISA, EC: 3.2.1.68)
appear to play a major role in amylopectin synthesis.
Mutations affecting isoamylase have been described in
maize, rice and barley (Burton et al., 2002; James et al.,
1995; Kubo et al., 1999), in Arabidopsis (Zeeman et al.,
1998a), and in the unicellular green alga Chlamydomonas
reinhardtii (Dauvillée et al., 2000, 2001a; Mouille et al., 1996).
All result in striking phenotypes, reducing or abolishing
starch synthesis and causing the accumulation of a soluble,
more highly branched glucan (phytoglycogen). The residual
815
816 Thierry Delatte et al.
starch is usually present as small granules, often with an
aberrant morphology and often containing altered amylopectin. In some cases, however, amylopectin appears to be
unaltered (Burton et al., 2002).
In addition to isoamylase, plants also contain a second
type of debranching enzyme, limit-dextrinase (LDA or pullulanase-type, EC: 3.2.1.142). In maize and Arabidopsis,
mutations that abolish activity of LDA do not cause phytoglycogen accumulation (Dinges et al., 2003; T. Delatte and
S.C. Zeeman, unpublished data). However, evidence suggests that LDA may determine the extent of phytoglycogen
accumulation in isoamylase-deficient mutants (Dinges et al.,
2003; Kubo et al., 1999).
Models have been proposed that suggest either a direct or
an indirect involvement of isoamylase in the synthesis of
amylopectin (Mouille et al., 1996; Myers et al., 2000; Zeeman
et al., 1998a). The direct model suggests that the synthesis
of the correct branching pattern of amylopectin requires
debranching enzyme activity and that, in its absence,
glucans are elongated and branched to form phytoglycogen.
The indirect model suggests that phytoglycogen and amylopectin are distinct products and that soluble phytoglycogen precursors, normally degraded by isoamylase activity,
are elaborated into phytoglycogen when isoamylase is
lacking. Both models address primarily the existence of
phytoglycogen in the isoamylase mutants. More recently it
has been suggested that isoamylase might also influence
starch granule initiation because the starch remaining in
isoamylase-deficient plants is present as numerous small
granules (Burton et al., 2002; Bustos et al., 2004). However,
the way in which isoamylase deficiency causes the complex
phenotypes observed has yet to be fully explained.
Where studied, isoamylase has a native molecular mass
of between 350 and 500 kDa (Dauvillée et al., 2001b; Fujita
et al., 1999; Ishizaki et al., 1983), indicating a multimeric
enzyme. In rice, isoamylase purified from developing endosperm was found to contain a single 83 kDa polypeptide, on
the basis of which the enzyme was proposed to be a
homomultimer (Fujita et al., 1999). In Chlamydomonas,
however, mutations at two loci (STA7 and STA8) affect
isoamylase activity and cause phytoglycogen accumulation
(Dauvillée et al., 2001a). Mutations at the STA7 locus result
in the loss of measurable isoamylase activity, whereas
mutations at the STA8 locus reduce isoamylase activity and
alter its native molecular mass. Recent evidence from potato
also suggests that isoamylase is heteromultimeric and that
two isoamylase proteins, ISA1 and ISA2, associate with each
other to form a single isoamylase enzyme in vivo (Hussain
et al., 2003).
The Arabidopsis genome encodes three isoamylase-like
proteins (AtISA1, AtISA2 and AtISA3) together with one LDA
protein (AtLDA). All four genes are widely conserved in both
monocots and dicots (Hussain et al., 2003). Interestingly,
amino acids thought to be required for catalysis are not
conserved in the ISA2 proteins, suggesting that they may
not be active debranching enzymes (Hussain et al., 2003). In
the isoamylase-deficient maize, barley and rice, the ISA1
gene is affected (Fujita et al., 2003; James et al., 1995; Kubo
et al., 1999), whereas in the Arabidopsis mutant dbe1-1, the
expression of the ISA2 gene is affected (Zeeman et al.,
1998a). Here we report a genetic approach to investigate the
nature of isoamylase activity and the function of the ISA1
protein in Arabidopsis, and examine the role of this enzyme
in starch metabolism.
Results
Mutations in AtISA1 and AtISA2
The three genes encoding isoamylase-like proteins annotated in the Arabidopsis genome are AtISA1 (At2g39930,
GenBank accession ATAF2109), AtISA2 (At1g03310, GenBank accession AC005278) and AtISA3 (At4g09020, GenBank
accession AC005359). Full-length cDNAs from these genes
have been isolated (AY139980, BT000443 and AY091058
respectively). All three proteins contain predicted N-terminal
chloroplast transit peptides (ChloroP; Emanuelsson et al.,
1999). We obtained a line (SALK_029442) containing a
putative T-DNA insertion in the coding sequence of AtISA1
and confirmed the location of the T-DNA within exon 13
using PCR (Figure 1a). Plants homozygous for the T-DNA
insert (Atisa1-1) were identified from the segregating population obtained from Salk.
We investigated the impact of the T-DNA insertion on
expression of the AtISA1 gene using RT-PCR. Primers pairs
were designed to amplify mRNA sequences corresponding
to positions upstream, downstream and spanning the
T-DNA insertion site. The correct mRNA sequences were
amplified from the wild type. For Atisa1-1, the upstream and
downstream primer combinations also amplified the correct
sequences, but no product was obtained for the primer
combination spanning the insertion site (Figure 1b). Using a
gene-specific primer (5¢ of the insertion site) and a T-DNAspecific primer, we amplified a hybrid fragment containing
both AtISA1 and T-DNA sequences from mutant but not
wild-type RNA (not shown). The hybrid mRNA contains a
predicted stop codon 18 bp into the T-DNA-derived
sequence, which would result in a truncated protein ending
in loop 7 of the conserved (ab)8 barrel structure. The
RT-PCR product from Atisa1-1 RNA with downstream primers probably represents a hybrid transcript initiated within
the T-DNA itself.
The DBE1 locus was mapped to the same chromosomal
location as the AtISA2 gene (Zeeman et al., 1998a). RNA gel
blot analysis revealed that AtISA2 expression was reduced
or abolished in this line (Zeeman et al., 1998a). To determine
the nature of the mutation, we amplified the AtISA2 coding
sequence and flanking regions (2 kb upstream and 1 kb
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Isoamylase function in Arabidopsis 817
known that non-sense mutations can destabilize transcripts,
possibly explaining why no transcript was detected previously (Zeeman et al., 1998a). Hereafter we refer to dbe1-1 as
Atisa2-1.
(a)
AATGGAGAAGA atatattgt
ATG
LB
(b)
1
2
3
4
(d)
T-DNA
Mutations in AtISA1 and AtISA2 abolish the same
isoamylase enzyme activity
1
2
3
4
M
1
2
3
D1
A1
A2
D2
5
6
7
8
A3
(e)
(c)
CACGG
ATG
A
AACCT
100
75
Figure 1. Identification and analysis of mutants at the AtISA1 and AtISA2
loci.
(a) Structure of the AtISA1 gene; exons are depicted as closed boxes.
Sequence flanking the T-DNA left border (hatched background) indicates that
T-DNA insertion in Atisa1-1 occurred in exon 13.
(b) RT-PCR on RNA isolated from the wild type (odd-numbered lanes) and
Atisa1-1 (even-numbered lanes) using AtISA1-specific primer combinations
upstream (lanes 1 and 2), spanning (lanes 3 and 4) and downstream (lanes 5
and 6) of the T-DNA insertion site (shown in a). Lanes 7 and 8 contain control
reactions using actin2-specific primers.
(c) Structure of the AtISA2 gene (DBE1 locus); the single exon is depicted as a
closed box. The single base pair deletion in Atisa2-1 (dbe1-1) is shown.
(d) Proteins in crude extracts of leaves from the wild type (lane 1), Atisa1-1
(lane 2), Atisa2-1 (dbe1-1; lane 3) and the Atisa1-1/Atisa2-1 double mutant
(lane 4) were separated by native PAGE in gels containing amylopectin.
Starch-hydrolysing activities were detected by staining the gel with iodine
solution after incubation for 16 h at 20C in 100 mM Tris-HCl, pH 7.0, 1 mM
DTT, 1 mM CaCl2, 1 mM MgCl2. D1 and D2 are isoamylase and limit dextrinase
respectively, A1–A3 are a- or b-amylases.
(e) Proteins in crude extracts of leaves from the wild type (lane 1), Atisa1-1
(lane 2), Atisa2-1 (dbe1-1; lane 3) were separated by SDS-PAGE, blotted onto
PVDF membranes and probed with anti-AtISA1 antibodies. M, molecular
weight markers, given as kDa on the left. Arrow indicates the position of the
ISA1 protein.
downstream) from dbe1-1 genomic DNA, using PCR.
Sequencing of the amplified fragments revealed a single
base pair deletion in the coding sequence of AtISA2 in dbe11, compared with amplified wild-type DNA and with the
published genomic sequence (Figure 1c). No other differences between dbe1-1 and wild-type sequences were found.
The resulting frame shift introduces a stop codon, truncating
the C-terminal part of the protein [in loop 8 of the (ab)8
barrel]. It seemed unlikely that this mutation would abolish
transcription of the gene and we were able to amplify AtISA2
mRNA from dbe1-1 using RT-PCR (not shown). However, it is
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Measurement of isoamylase activity in crude extracts of
higher plants is problematic due to the interference of
other glucan hydrolysing enzymes. Therefore we used
native PAGE to investigate whether the loss of AtISA1
expression resulted in a loss of isoamylase activity. We
found that a major debranching enzyme activity present in
crude extracts of wild-type leaves was missing in Atisa1-1
(Figure 1d, band D1). This enzyme is a chloroplastic isoamylase (Zeeman et al., 1998a,b). Mutations at the DBE1
locus (encoding AtISA2) also result in the loss of D1
activity (Figure 1d; Zeeman et al., 1998a). One explanation
for this result is that both AtISA1 and AtISA2 contribute to
the same isoamylase activity, as reported for potato
(Hussain et al., 2003).
Given that the AtISA2 protein is likely to be non-catalytic
(Hussain et al., 2003; also see Introduction), we investigated
the expression of AtISA1 in the Atisa2-1 mutant using RTPCR. No difference in AtISA1 mRNA abundance was
observed between Atisa2-1 and wild-type plants (not
shown). However, protein gel blots probed with an antibody
raised against an AtISA1-specific peptide revealed that the
AtISA1 protein was appreciably reduced in Atisa2-1 (Figure 1e). This suggests that in the absence of wild-type
AtISA2, AtISA1 may be unstable. No AtISA1 protein was
detected in Atisa1-1, as expected.
We investigated the native molecular weight of the D1
isoamylase using gel permeation chromatography. The D1
activity co-eluted with ribulose 1,5-bisphosphate carboxylase, which is known to have a native molecular mass of
around 500 kDa (Figure S1). This is much larger than the
predicted size of AtISA1 or AtISA2 (85 and 96 kDa respectively) and is consistent with the idea that the isoamylase D1
is a multimeric protein in vivo. Using protein gel blots
probed with the AtISA1-specific antibodies, we confirmed
that AtISA1 co-eluted with D1 activity in wild-type extracts
(Figure S2a). We also investigated the native molecular
weight of the residual AtISA1 protein in Atisa2-1. The very
low levels of AtISA1 in this line made its detection impossible with the same conditions used for the wild-type extracts.
However, by increasing fivefold the protein loaded the gel
and using sensitive chemiluminescent methods, a protein of
the correct molecular weight was just detectable (Figure S2b). This protein eluted late from the gel permeation
column suggesting that, in addition to being reduced in
abundance, AtISA1 has a lower native molecular weight in
the absence of AtISA2.
818 Thierry Delatte et al.
Atisa1-1 and Atisa2-1 have identical phenotypes and
accumulate altered glucans
Loss of D1 isoamylase caused by mutations of the AtISA2
gene results in a reduction in starch accumulation and the
simultaneous production of soluble phytoglycogen in the
chloroplasts of leaf mesophyll cells (Zeeman et al., 1998a).
We compared the phenotypes of Atisa1-1, Atisa2-1, the
Atisa1-1/Atisa2-1 double mutant and the wild type. First,
plants were stained with iodine at the end of the day (Figure 2a). Wild-type plants stained a dark brown colour, typical
of amylopectin. Atisa1-1 and Atisa2-1 stained a more
(a)
(b)
Starch
10
Glucan content (mg g–1 FW)
8
6
4
2
0
Soluble glucan
4
2
n.d.
0
WT
Atisa1-1 Atisa2-1
DM
Figure 2. Starch and soluble glucan content of wild-type, Atisa1-1, Atisa2-1
and Atisa1-1/Atisa2-1 double mutant plants.
(a) Wild-type (middle), Atisa1-1 (left) and Atisa2-1 (right) plants were
harvested at the end of a 12-h photoperiod, decolourized with hot ethanol
and stained for starch with iodine solution.
(b) All the leaves of individual wild-type (WT), Atisa-1, Atisa2-1 and Atisa1-1/
Atisa2-1 double mutant (DM) plants were harvested and immediately frozen
in liquid N2. Insoluble (upper graph) and soluble (lower graph) glucans were
extracted using perchloric acid and measured as described in Experimental
procedures. Each bar is the mean standard error from five replicate
samples. n.d., not detected.
reddish-brown (Figure 2a), as did the double mutant (not
shown). This is indicative of more highly branched glucan
polymers in the mutants than in the wild type. Second, we
measured the starch and soluble-glucan contents of the
leaves at the end of the photoperiod. We found that Atisa1-1,
Atisa2-1 and the double mutant accumulated reduced
amounts of starch relative to the wild type, together with
large amounts of soluble glucan (Figure 2b).
We compared the structures of the starches and phytoglycogens from the single mutants and double mutant with
the starch from wild-type plants using a combination of
enzymatic digestions and high-performance anion exchange chromatography with pulsed amperometric detection (HPAEC-PAD). The analyses were repeated on several
batches of plants, in each of which wild-type and mutant
plants were grown side by side in the growth chamber.
There were small variations in the glucan structures
between batches of plants, but the differences observed
between wild-type and mutant plants were highly consistent. We used two different extraction methods; the first
involved extracting the glucans in ice-cold buffered medium
and the second involved extraction with perchloric acid (see
Experimental procedures). Both types of extraction have
been used in earlier studies, but the two have not been
previously compared. The results from the two methods
were similar for starch, but for phytoglycogen the two
extraction methods yielded qualitatively different results,
which affects the interpretation of the mutant phenotypes in
both this and other studies.
Using ice-cold buffered extraction medium, starch and
soluble glucans were extracted from leaves of the wild type,
the two single mutants and the double mutant. The chain
length distributions of the amylopectin from all three mutant
lines were identical to each other, but differed from that of
the wild type (Figure 3a,b; Figure S3). The amylopectin from
the mutants had a small increase in the relative number of
very short glucan chains with a degree of polymerization
(dp) between 3 and 8, and a decrease in slightly longer
chains (dp 9–16) compared with wild-type amylopectin. The
chain length distributions of the phytoglycogen from all
three mutant lines were also very similar to each other
(Figure 3c; Figure S3). Compared with wild-type amylopectin, the phytoglycogen contained far more very short chains
(dp 3–9) and fewer slightly longer chains (dp 11–35)
(Figure 3d; Figure S3).
Figure 4 (and Figure S4) shows similar chain-length analyses performed on starch and soluble glucans extracted
using perchloric acid. The results for the amylopectin from
the mutants and wild type were similar to those obtained
with the buffered extraction medium (compare Figures 3b
and 4b; see also Figures S3 and S4). The amylopectins from
all three mutant lines were similar to each other but differed
from that of the wild type. However, the phytoglycogen
extracted using perchloric acid contained relatively fewer
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Isoamylase function in Arabidopsis 819
10
WT amylopectin
8
6
4
2
0
3
8
13
18
23
28
33
38
43
10
Relative percentage
Relative percentage
(a)
Atisa1-1 amylopectin
8
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3
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(b)
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–2
–4
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Chain length
12
Relative percentage
(c)
Atisa1-1 phytoglycogen
10
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4
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0
3
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18
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28
33
38
43
48
Chain length
8
(d)
Relative percentage
difference
Figure 3. Comparisons of the chain length distributions
of glucans extracted from leaves of wild-type, Atisa1-1,
Atisa2-1 and the Atisa1-1/Atisa2-1 double mutant plants
using ice-cold buffered extraction medium.
Glucans were extracted from batches of approximately 50
plants, debranched with Pseudomonas isoamylase, and
analysed by HPAEC-PAD. Peak areas were summed and
the areas of individual peaks expressed as a percentage of
the total. The mean standard errors of three independent isoamylase digests are shown. For difference plots,
standard errors of the compared data sets were added
together. For clarity, only the results of a representative
experiment for Atisa1-1 are shown in full. The complete
data set is presented in Figure S3.
(a) Chain length distributions of amylopectins from the
wild type and Atisa1-1.
(b) Difference plots derived by subtracting the relative
percentage values of wild-type amylopectin from those of
Atisa1-1 (black), Atisa2-1 (red) and the Atisa1-1/Atisa2-1
double mutant (yellow) amylopectins.
(c) Chain length distribution of phytoglycogen from
Atisa1-1.
(d) Difference plots derived by subtracting the relative
percentage values of wild-type amylopectin from those of
phytoglycogen from Atisa1-1 (black), Atisa2-1 (red) and the
Atisa1-1/Atisa2-1 double mutant (yellow).
23
Chain length
Relative percentage
difference
Chain length
6
4
2
0
–2
–4
3
8
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Chain length
very short chains and more longer chains compared with the
phytoglycogen extracted in the buffered extraction medium.
The likely explanation for this difference is that the phytoglycogen is susceptible to some enzymatic degradation
during extraction in the buffered medium. This would not
occur during extraction in perchloric acid, as enzymes would
very rapidly be inactivated. Thus, we suggest that the results
obtained using this latter method reflect more closely the
chain length distribution of the phytoglycogen in vivo. As
mentioned previously, the phytoglycogens from the single
mutants and the double mutant were all very similar to one
another (Figure 4d; Figure S4). However, it is striking that
the magnitude of the alterations in the chain length distribution when the phytoglycogen is compared with wild-type
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
amylopectin is quite small (Figure 4d). When compared with
the amylopectin from the mutants, the difference is smaller
still (Figure 4e).
Analysis of the chain length distribution of branched
glucans fails to address whether the distribution of branch
points within the molecule is altered (Thompson, 2000).
Therefore, we analysed the glucans from wild-type and
mutant plants after first digesting the external segments of
each polymer using b-amylase. b-Amylase removes maltosyl units from the non-reducing end of glucan chains to
within two or three glucosyl positions of a branch point. The
resulting b-limit glucans were debranched and their chain
length distributions determined. This technique allows the
comparison of the relative proportions of A chains (chains
820 Thierry Delatte et al.
(a)
Relative percentage
Relative percentage
10
WT amylopectin
8
6
4
2
0
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10
Atisa1-1 amylopectin
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0
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18
Relative percentage
difference
(b)
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33
38
43
48
38
43
48
4
2
0
–2
–4
3
8
13
18
23
28
33
Chain length
(c)
Relative percentage
Figure 4. Comparisons of the chain length distributions of
glucans extracted from leaves of wild-type, Atisa1-1,
Atisa2-1 and the Atisa1-1/Atisa2-1 double mutant plants
using perchloric acid.
Glucans were extracted from four to six individual plants.
Equal quantities of starch or phytoglycogen from each
plant were pooled, debranched with Pseudomonas isoamylase, and analysed by HPAEC-PAD. Peak areas were
summed and the areas of individual peaks expressed as a
percentage of the total. The mean standard errors of
three independent isoamylase digests are shown. For
difference plots, standard errors of the compared data sets
were added together. Only the results for Atisa1-1 are
shown in full. The complete data set is presented in
Figure S4.
(a–d) As described in Figure 3.
(e) Difference plots derived by subtracting the relative
percentage values of Atisa1-1 (black), Atisa2-1 (gold) and
the Atisa1-1/Atisa2-1 double mutant (cyan) amylopectin
from the respective values for phytoglycogen from the
same lines.
23
Chain length
Chain length
10
Atisa1-1 phytoglycogen
8
6
4
2
0
3
8
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(d)
Relative percentage
difference
Chain length
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2
0
–2
–4
3
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(e)
Relative percentage
difference
Chain length
4
2
0
–2
–4
3
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28
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Chain length
that carry no other chains) and B chains (chains that carry
one or more other chains via a branch point) and of the
lengths of internal B-chain segments (Thompson, 2000).
Glucans extracted using both extraction methods (buffered
medium and perchloric acid) were analysed in this way and
yielded comparable results, both for the amylopectins and
the phytoglycogens (Figure 5 compare a and b; compare
Figures S5 and S6). This indicates that the modification to
phytoglycogen structure during extraction in buffered
extraction medium, detected when comparing the chain
length profiles (see above; Figures 3 and 4), affects primarily
the linear outer chains and not the internal branch point
distribution.
The results from the single mutants and the double
mutant were very similar to one another and reveal that both
amylopectin and phytoglycogen synthesized in the absence
of isoamylase are different in branch point distribution from
wild-type amylopectin. First, maltose and maltotriose, representing residual stubs of A-chains, dominated all the
distributions but were relatively more abundant in wild-type
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Isoamylase function in Arabidopsis 821
(a) 60
50
40
Relative percentage
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20
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Chain length
Chain length
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Chain length
(b)
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Chain length
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Chain length
60
50
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Relative percentage
30
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Chain length
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Chain length
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Chain length
Figure 5. Comparisons of the chain length distributions of wild-type and Atisa1-1 glucans after b-amylolysis of the external chains.
Glucans were extracted as described in Figures 3 and 4, treated with b-amylase and debranched with both Pseudomonas isoamylase and Klebsiella pullulanase.
Results obtained for Atisa1-1, Atisa2-1 and the Atisa1-1/Atisa2-1 double mutant were very similar. Note the differences and changes in scale of the y-axis. For clarity
only the Atisa1-1 results are displayed. The complete data sets are presented in Figures S5 and S6.
(a) b-Limit chain length profiles of glucans extracted using ice-cold buffered medium. The mean standard errors of three independent isoamylase digests are
shown.
(b) b-Limit chain length profiles of glucans extracted using perchloric acid. Values for a single set of digests is shown. Replicate digests gave very similar results. The
abundance of dp 6 varied considerably between individual samples. However, this variability was not consistent between individual samples from a single
genotype.
b-limit amylopectin than in b-limit phytoglycogen from the
mutants (Figure 5). This reflects a relative reduction in
A-chains in phytoglycogen due to its increased branch
frequency compared with amylopectin. Chains longer than
maltotriose, representing internal B-chain segments, had
the opposite relative abundances, as would be expected.
However, the b-limit phytoglycogen was greatly enriched in
the shorter residual B-chains, indicating that branch points
in phytoglycogen are closer together relative to wild-type
amylopectin. The chain length profiles of b-limit amylopectin from the mutants were intermediate between those of
wild-type b-limit amylopectin and of b-limit phytoglycogen,
However, in qualitative terms the b-limit profiles of amylopectin from the mutants resembled those of phytoglycogen
more closely than of wild-type amylopectin. This shows that,
in addition to an altered overall chain length distribution, the
amylopectin from the mutants also has an increased
branching frequency, with branches located closer together
than in wild-type amylopectin.
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Phytoglycogen structure is modified via degradation in vivo
Analysing the structure of the glucans present at a single
time point cannot give a full picture of how these glucans are
produced. Therefore, we analysed the pattern of starch and
phytoglycogen accumulation throughout the day and night
and investigated whether there were any differences in
glucan structure at different times. All extracts for these
experiments were made using the perchloric acid extraction
method described above.
We determined the starch and phytoglycogen content of
leaves of Atisa1-1 and Atisa2-1 and the wild type during the
diurnal cycle. The results for the two single mutants were
identical to each other and similar to those previously
reported for Atisa2-1 (Figure 6a; Zeeman et al., 1998a). In the
wild type, starch accumulated at a fairly constant rate during
the day, reaching a peak value of 8 mg g)1 FW and, after a
brief lag, was degraded at a fairly constant rate during the
night. In the mutants, starch also accumulated and was
822 Thierry Delatte et al.
degraded at constant rates during the day and night,
respectively, but the amount synthesized was less than in
the wild type (2 mg g)1 FW; Figure 6a). Phytoglycogen
accumulation, however, occurred most rapidly in the first
8 h of the day, after which time the amount of phytoglycogen increased more slowly, if at all. The rate of phytoglycogen breakdown was fastest during the early hours of the
night and after 8 h of darkness, most of the phytoglycogen
10
(a)
Starch (mg g–1 FW)
8
6
had been metabolized. The total glucan accumulation in the
isoamylase mutants was less than that in the wild type.
We investigated the cause of the reduced rate of phytoglycogen accumulation late in the day in the isoamylase
mutants. First, we measured the rate of photosynthesis in
the wild type and Atisa2-1 to see if the rate of carbon
assimilation was altered in the mutant. This was not the
case: the carbon assimilation rate was the same in both lines
throughout the day (not shown). This suggests that the
accumulation of phytoglycogen does not generally inhibit
metabolic processes in the chloroplast. Second, we illuminated wild type and Atisa1 and Atisa2 mutant plants in a
range of different light intensities for the duration of a single
12 h photoperiod to determine whether increasing the
rate of carbon assimilation would increase the amount of
glucan synthesized. As expected, increasing the light intensity led to increased starch accumulation in the wild type
(Figure 7). In the mutants, the amount of starch synthesized also appeared to increase, whereas the amount of
4
25
20
0
Starch (mg g–1 FW)
Soluble glucan (mg g–1 FW)
2
4
3
2
1
15
10
0
0
6
12
18
5
24
(b)
0.1
0
Soluble glucan
(mg g–1 FW)
Maltose (mg g–1 FW)
0.12
0.08
0.06
0.04
0.02
9
6
3
0
0
0
0
4
8
12
16
20
24
200
400
600
–1
800
–2
Light intensity (µmol photons s m )
Time (h)
Figure 6. Starch and soluble glucan contents of wild-type, Atisa1-1 and
Atisa2-1 plants.
All the leaves of individual wild-type (black symbols), Atisa-1 (grey symbols)
and Atisa2-1 (white symbols, no line) plants were harvested and immediately
frozen in liquid N2.
(a) Starch (upper graph) and soluble glucans (lower graph) were extracted
using perchloric acid and measured as described in Experimental procedures.
Each point is the mean standard error from six replicate samples.
(b) Soluble glucans were extracted from a replicate batch of plants using
perchloric acid and maltose in the neutral fraction measured by HPAEC-PAD.
Figure 7. Starch and phytoglycogen accumulation in leaves of wild-type and
isoamylase-deficient mutants illuminated at different light intensities.
Plants were grown as described in Experimental procedures until 6 weeks old.
At the beginning of a photoperiod, plants were shifted to a range of different
light intensities. The exact light intensity at the surface of individual mature
leaves was measured and these leaves (one per plant) were harvested at the
end of the 12-h photoperiod and immediately frozen in liquid N2. Starch
(upper graph) and soluble glucans (lower graph) were extracted with
perchloric acid and measured as described in Experimental procedures. Wild
type, grey circles; Atisa1-1, open circles; Atisa2-1, open squares; Atisa2-2
(dbe1-2) open triangles. Lines were fitted by eye.
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Isoamylase function in Arabidopsis 823
the end of the day (12 h; Figure 8c). Together, these data
suggest that some external chains are being degraded (e.g.
by b-amylase), resulting both in the production of the very
short chains of phytoglycogen and the observed maltose. In
contrast, the chain length profiles of the amylopectin
present in the wild type after 4 h and at the end of the day
(12 h) were similar to one another (Figure 8a). Intriguingly,
the chain length profiles of the amylopectin from the
mutant exhibited small changes similar in nature to that of
the phytoglycogen (Figure 8b). The results from both
experiments support the idea that glucans in the mutant,
especially the phytoglycogen, are subject to the action of
degradative enzymes during the day, whereas in the wild
type relatively little such daytime glucan degradation
occurs.
We also compared the chain length profiles of glucans
extracted from the leaves harvested after 4 h of darkness
(16 h) with those extracted at the end of the day. Phytoglycogen contained many more very short chains less than
phytoglycogen accumulated was restricted and increased
marginally, if at all, at higher levels of illumination (Figure 7).
We reasoned that during the period of glucan synthesis,
glucan-degrading enzymes could be acting on phytoglycogen, limiting its accumulation through turnover. To test this
hypothesis, we first measured the levels of maltose, a known
intermediate of starch breakdown in Arabidopsis leaves
(Zeeman et al., 2004). The maltose contents increased
throughout the day in the isoamylase mutants, whereas in
the wild type the levels remained low until the onset of
starch degradation at night (Figure 6b). During the night, the
amount of maltose in the mutant declined faster than in the
wild type, presumably because the glucan reserves are
exhausted more rapidly (Figure 6a). Next, we examined the
structure of phytoglycogen at different times during the day
and night in wild-type and Atisa1-1 plants. The phytoglycogen structure synthesized early in the day (4 h time point)
had fewer chains less than dp 11 (and notably very few
chains smaller than dp 6) than the phytoglycogen present at
10
8
4h
8
12 h
8
6
6
6
4
4
4
2
2
2
0
8
13
18
23
28
33
38
43
48
16 h
0
0
3
3
8
13
Chain length
18
23
28
33
38
43
8
13
18
23
28
33
38
43
8
12 h
6
6
4
4
4
2
2
2
0
0
0
8
13
18
23
28
33
38
43
48
3
8
13
Chain length
18
23
28
33
8
13
38
43
16 h
23
28
33
38
43
48
38
43
48
38
43
48
3
1
-1
-3
-5
3
48
18
5
8
13
Chain length
12
3
Chain length
8
6
3
-3
48
10
10
4h
1
-1
Chain length
(b)
8
3
-5
3
48
Chain length
10
Relaitve percenatge
5
Relative percentage
difference
10
(a)
Relative percentage
difference
Relaitve percenatge
10
18
23
28
33
38
43
48
3
8
13
Chain length
18
23
28
33
Chain length
12
12
7
10
10
5
Relaitve percenatge
10
8
4h
8
12 h
6
6
4
4
4
2
2
2
0
0
3
8
13
18
23
28
33
Chain length
38
43
48
16 h
8
6
8
13
18
23
28
33
38
43
48
3
1
-1
-3
-5
0
3
Relative percentage
difference
(c)
3
Chain length
8
13
18
23
28
33
Chain length
38
43
48
3
8
13
18
23
28
33
Chain length
Figure 8. Comparisons of the chain length distributions of glucans extracted from leaves of the wild type and Atisa1-1 at different times of the diurnal cycle.
Glucans were extracted using perchloric acid after 4 h of light (4 h), at the end of the photoperiod (12 h) and after 4 h of darkness (16 h). Values for a single
experiment are shown. A second experiment with another batch of plants yielded similar results.
(a) Chain length distributions of wild-type amylopectin at 4, 12 and 16 h time points. The difference plots are derived by subtracting the relative percentage values of
4 h from 12 h (filled circles), and of 12 h from 16 h (open circles).
(b) Chain length distributions of Atisa1-1 amylopectin at 4, 12 and 16 h time points, as described for (a).
(c) Chain length distributions of Atisa1-1 phytoglycogen at 4, 12 and 16 h time points, as described for (a).
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
824 Thierry Delatte et al.
dp 8, compared with the phytoglycogen present at the end of
the day, indicating that the phytoglycogen pool as a whole is
subject to gradual, progressive degradation. The amylopectin from the wild type, however, showed no such differences
suggesting that, during degradation, amylopectin at the
surface of the granule is completely broken down, whilst
that within the granule remains untouched. Again, results
intermediate between wild-type amylopectin and phytoglycogen were obtained with the amylopectin from Atisa1-1,
which exhibited a small increase in chains less than dp 8
between the 12-h and 16-h time points.
Tissue-specific patterns of glucan accumulation in Atisa1-1
and Atisa2-1
Leaf starch is synthesized primarily from photoassimilates
in the mesophyll cells and previous investigation of Atisa21 focussed on this tissue (Zeeman et al., 1998a). We used
transmission electron microscopy (TEM) to survey the
different cell types in the mutants because other cells, such
as the bundle sheath cells surrounding the vasculature
(starch sheath; Müller-Röber et al., 1994; Rook et al., 2001),
also contain starch. The results for Atisa1-1 and Atisa2-1
were identical. Many glucan structures were visible in the
mutants, ranging from starch granules with a normal
appearance through to phytoglycogen, including many
aberrant granules and, in some cases, material that was
difficult to define either as insoluble starch or soluble
phytoglycogen (Figure 9). However, a striking, tissue-specific pattern was observed. The majority of mesophyll
chloroplasts in the mutants contained very small amounts
of starch in the form of tiny granules, often irregular in
outline (Figure 10, compare a with b and c). Most of the
space was occupied by phytoglycogen, causing the chloroplasts to be distended compared with those of the wild
type. In guard-cell chloroplasts of the mutants, more starch
and less phytoglycogen were present compared with the
mesophyll cells (e.g. Figure 9f). Remarkably, the plastids of
epidermal cells in the mutants (both upper and lower
epidermis) contained starch but no apparent phytoglycogen and were indistinguishable from the corresponding
cells of the wild type (Figure 10f,g). Chloroplasts in the
subtending palisade cells of the mutant were typical of the
mesophyll. Sieve element companion cells and the adjacent bundle sheath cells also contained starch but no
apparent phytoglycogen (Figure 10h,i) and chloroplasts in
some of the outermost bundle sheath cells contained
predominantly starch, but in the form of many irregular
granules (Figure 10i).
We examined the chain length profiles of starch preparations from epidermal peels of wild-type and Atisa2-1 plants
to determine whether the apparently normal starch granules
in the epidermal cells had amylopectin similar to the wild
type. Lower epidermal peels were prepared from plants at
the end of the day and the ratio of starch and phytoglycogen
determined. The preparation from Atisa2-1 predominantly
(a)
(c)
(e)
(g)
(i)
(b)
(d)
(f)
(h)
(j)
Starch
Phytoglycogen
Figure 9. Transmission electron micrographs of a range of glucan structures from plastids of Atisa2-1 leaf cells.
Leaf samples were harvested 9 h into a 12-h photoperiod and prepared for TEM as described in Experimental procedures. Bars ¼ 200 nm.
(a) Epidermal cell plastid.
(b, d, e) Chloroplasts from bundle sheath cells.
(c, g, h, i, j) Chloroplasts from mesophyll cells.
(f) Chloroplast from stomatal guard cell.
All the corresponding cell types of the wild type accumulated regular starch granules similar in appearance to those in (a) and (b) and none of the structures shown in
(c–j) (see also Figure 10).
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Isoamylase function in Arabidopsis 825
(a) WT
(d) WT
epi
(b)
Atisa1-1
(e)
Atisa1-1
epi
(c)
Atisa2-1
(f)
Atisa2-1
epi
epi
pal
pal
(g)
WT
bs
sm
pal
(i)
Atisa2-1
sm
bs
(h)
cc
bs
Atisa1-1
bs
sm
bs
bs
Figure 10. Starch- and phytoglycogen-accumulating phenotypes of different cell types in wild-type and Atisa1-1 and Atisa2-1 (dbe1-1) leaves.
Leaf samples were harvested 9 h into a 12-h photoperiod and prepared for TEM as described in Experimental procedures. The pattern of starch and phytoglycogen
accumulation was the same in Atisa1-1 and Atisa2-1. Bars ¼ 2 lm.
(a–c) Chloroplasts from palisade cells. Note the increased size of the mutant chloroplasts.
(d–f) Epidermal cell plastids (above) and the chloroplasts from subtending palisade cells (below). pal, palisade cell; epi, epidermal cell.
(g–i) Cells surrounding the vasculature. Arrows indicate the chloroplasts of the mutants that contain predominantly starch. sm, spongy mesophyll cell; bs, bundle
sheath cell; cc, sieve element companion cell.
contained starch (not shown), consistent with the TEM
observations. Comparison of the amylopectin chain length
distribution of these preparations revealed a difference
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
pattern that was very similar to that obtained for the starch
derived from the whole tissue (Figure 11; compare with
Figure 3a,b). This suggests that the epidermal starch in
826 Thierry Delatte et al.
10
Relative percentage
WT
Atisa2-1
8
6
4
2
Figure 11. Comparisons of the chain length distributions of starch purified from leaf epidermal
peels of wild-type and Atisa2-1 plants.
Starch in the insoluble fraction obtained from
peels of the lower epidermis was debranched
and analysed as described in Figure 3. The
difference plot was obtained by subtracting the
relative percentage values of wild type amylopectin from that of Atisa2-1.
0
3
8
13
18
23
28
33
38
43
48
Relative percentage
dfiferecne
Chain length
3
8
13
18
23
28
33
38
43
48
38
43
48
Chain length
4
2
0
-2
-4
3
8
13
18
23
28
33
Chain length
Atisa2-1 has an altered amylopectin structure, even though
no phytoglycogen is apparent in this tissue.
Discussion
AtISA1 and AtISA2 are both required for a multimeric
isoamylase
The insertion of a T-DNA in SALK_029442 effectively eliminates the expression of the AtISA1 gene and is reflected in
the loss of D1 isoamylase activity in crude extracts of leaves
(Figure 1b,d). The fact that D1 is also absent in AtISA2 mutants indicates that both AtISA1 and AtISA2 are needed for
this activity. The most likely explanation is that AtISA1 and
AtISA2 are both components of a heteromultimeric enzyme,
as is the case for the orthologous proteins in potato (Bustos
et al., 2004; Hussain et al., 2003). This is consistent with the
high native molecular mass of D1 (Figure S1). Hussain et al.
(2003) argued that, as a result of amino acid substitutions,
the Arabidopsis and potato ISA2 proteins were unlikely to
retain hydrolytic activity. This conclusion was supported by
in vitro analysis of the StISA2 protein, which was inactive
when expressed in Escherichia coli. In contrast, the ISA1
proteins retain the consensus sequences for active hydrolases and StISA1 was active when expressed in E. coli. This
suggests that in a heteromultimeric enzyme of ISA1 and
ISA2, ISA1 would be the catalytic subunit.
In the absence of AtISA2, AtISA1 may or may not be active
in vivo. However, our results suggest that AtISA1 is unstable
in Atisa2-1, as the protein is dramatically reduced in
abundance, even though its transcript is still detected.
Thus, we cannot determine if the phenotype of Atisa2-1
is the same as Atisa1-1 because of the reduction of AtISA1
protein or because AtISA1 is inactive (or has altered activity/
specificity) in the absence of AtISA2, or both. If AtISA1
were stable and active, then the phenotype of Atisa2-1
might be distinct from that of Atisa1-1. It is possible that
the phenotype of the sta8 mutant of Chlamydomonas
reflects such a situation. Chlamydomonas possesses a
multimeric isoamylase enzyme and mutations at STA8
locus affect its integrity and properties, but do not abolish
its activity. Interestingly, the sta8 mutant displays a mild
phytoglycogen-accumulating phenotype (Dauvillée et al.,
2001a), whereas the sta7 mutant, which completely lacks
isoamylase activity, has a severe phenotype (Mouille et al.,
1996). However, the gene encoded at the STA8 locus has
yet to be identified.
The role of AtISA1 and AtISA2 in starch biosynthesis
The data presented here shed new light on the function of
isoamylase and, on balance, favour a direct involvement for
the enzyme in determining amylopectin structure. The
starch synthesized in the absence of the AtISA1/AtISA2 isoamylase is altered in structure compared with wild-type
starch. These differences were not detected in our previous
study of the Atisa2-1 mutant due to the relative insensitivity
of the methods used at that time (Zeeman et al., 1998a). The
overall chain length distribution of amylopectin differs from
that of the wild type in essentially the same way as in the
isoamylase mutants of maize and rice (Dinges et al., 2001;
Wong et al., 2003). Importantly, we find that the branching
pattern of the amylopectin in the mutants is also altered
compared with wild-type amylopectin, but resembles that of
phytoglycogen. In particular, the residual B-chains of the
b-limit amylopectin and b-limit phytoglycogen from the
mutant have a high proportion of short chains compared
with wild-type amylopectin, indicating branch points that
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Isoamylase function in Arabidopsis 827
are very close together. Thus, the function of the AtISA1/
AtISA2 isoamylase may be to hydrolyse preferentially
branch points that are very close to other branch points,
thereby facilitating the crystallization of amylopectin. If this
is the case, the proposed heteromultimeric nature of the
ISA1/ISA2 isoamylase may be significant in conferring such
structural specificity. It is possible to speculate that the noncatalytic ISA2 subunit could recognize one branch point and
facilitate the removal of a nearby branch by the catalytic
ISA1 subunit. It is important to note that alterations in the
chain length profiles of the residual starch are not always
observed in isoamylase-deficient plants (e.g. barley; Burton
et al., 2002). In such cases, it would be interesting to determine whether there is any change in the branch point
distribution.
Our data suggest that the structural differences between
granular starch and soluble phytoglycogen may have been
overestimated previously because phytoglycogen is prone
to structural alterations both during extraction and in vivo.
First, by extracting glucans in perchloric acid to rapidly
inactivate enzymes, we obtained chain length profiles for
phytoglycogen that were quite different to those from
buffer-extracted phytoglycogen. The phytoglycogen extracted with perchloric acid had longer chains on average,
indicating that during extraction in buffered medium degradation of the external chains may be occurring. Second,
comparison of perchloric acid-extracted phytoglycogen
from leaves of the Atisa1-1 harvested at different times of
the day reveals that the phytoglycogen present early in the
day has longer chains on average than phytoglycogen
present later in the day. This could be caused by an
increased degree of branching or by shortening of the
chains through degradation. The evidence points towards
the latter explanation because maltose, a known intermediate of starch breakdown (Chia et al., 2004; Niittylä et al.,
2004), accumulates in the leaves of the mutant in a pattern
that is reciprocal to the rate of phytoglycogen accumulation,
but does not accumulate in the wild type until the onset of
starch breakdown. Furthermore, the phytoglycogen accumulated early in the day has few chains shorter than dp 6,
consistent with action of branching enzymes, which are not
thought to transfer chains shorter than dp 6 (Nielsen et al.,
2002; Rydberg et al., 2001). Thus, the relative increase in
chains shorter than dp 6 in the phytoglycogen present later
in the day is caused by degradation of longer chains.
We suggest that a balance between synthesis and degradation of the outer chains of phytoglycogen is achieved in
the mutants during the latter part of the day. The maltose
produced by degradation is presumably exported to the
cytosol and metabolized (Niittylä et al., 2004). Previous
experiments using a 14C pulse-labelling approach failed to
detect phytoglycogen turnover in Atisa2-1 (Zeeman et al.,
2002). However, these experiments were performed early in
the day when phytoglycogen was still accumulating rapidly.
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Interestingly, we also detected small changes in the chain
length profile of the mutant amylopectin at different times of
the day, indicating that at least some of the starch of the
mutant may be more susceptible to modification than wildtype starch (see below). During the night, glucan synthesis
stops and degradation commences. The abundance of very
short chains in the phytoglycogen from the perchloric acidextracted night-time samples was even greater than in
samples extracted at the end of the day. In fact, the 16-h
chain length profile resembled most closely the phytoglycogen extracted using buffered extraction medium, supporting our earlier conclusions about the unsuitability of this
extraction method.
A range of glucans accumulates in tissue-specific patterns
Our ultrastructural observations provide further new
insights into the phenotype caused by isoamylase deficiency. First, specific cell types in the mutants displayed
strikingly different phenotypes. Second, many different
glucan structures were apparent within the mutant tissues
as a whole.
Localized phytoglycogen and starch accumulation has
been observed in the developing endosperm of isoamylase
mutant of cereals (Boyer et al., 1977; Burton et al., 2002;
Nakamura et al., 1997) but it was not apparent from these
studies that particular cell types were behaving differently.
In Arabidopsis, some cell types appear to accumulate only
starch with no apparent phytoglycogen, some primarily
phytoglycogen with small amounts of starch, while others
appear to have an intermediate phenotype. An obvious
explanation for the cell-type-specific phenotype is that other
debranching enzyme isoforms (i.e. ISA3 or LDA) are highly
expressed in epidermal and bundle sheath cells and can, to
some extent, fulfil the same role as ISA1/ISA2. We are
pursuing research to evaluate the function of these proteins
in Arabidopsis and thereby test this possibility. However, it
is also reasonable to suppose that other aspects of starch
metabolism may be different between palisade cells (primary photosynthetic cells), guard cells (photosynthetic, but
which synthesize starch at night and metabolize it during the
day) and epidermal cells (largely non-photosynthetic). The
complement of enzyme isoforms, both biosynthetic and
degradative, may differ between cell types, as could the way
in which these enzymes are regulated.
The interpretation of the chain length profiles of the
glucans must be considered in the light of the range of
apparent glucan structures visible by TEM (Figures 9 and
10). The results from whole-leaf preparations may represent
an average of many glucan structures. However, our analysis of the starch in preparations of epidermal peels suggests
that the granules in epidermal cells, although normal in
appearance, are still made of altered amylopectin (Figure 11). This is important because it raises the possibility
828 Thierry Delatte et al.
that factors other than AtISA1/AtISA2 determine whether or
to what extent glucan is accumulated in an insoluble form
(starch granules) or a soluble form (phytoglycogen).
Partitioning between starch and phytoglycogen
We consider it likely that other enzymes of starch metabolism influence the partitioning between starch and phytoglycogen when the ISA1/ISA2 isoamylase is lacking. Myers
et al. (2000) suggested that during the synthesis of starch,
crystallization prevents continued modification of the nascent, soluble amylopectin molecules by glucan-modifying
enzymes. In this context, it has been suggested that disproportionating enzyme (D-enzyme) may participate in the
synthesis of amylopectin, based on the altered starch-phenotype of a D-enzyme-deficient mutant of Chlamydomonas
(Colleoni et al., 1999). Although there is no evidence that
D-enzyme is involved in normal amylopectin synthesis in
Arabidopsis (Critchley et al., 2001), the evidence presented
here indicates that the glucans produced in the AtISA1/
AtISA2 isoamylase mutants are subjected to appreciable
secondary modification.
We suggest that the AtISA1/AtISA2 isoamylase mutants
produce glucans with an altered branching pattern which
delays, but does not necessarily prevent crystallization. This
would render these glucans susceptible to more modification by enzymes that may or may not normally feature in the
starch biosynthetic process. The proportion of glucan that
remains soluble or crystallizes into granules could depend
on the extent of secondary modifications and therefore on
which other glucan-metabolizing enzymes are present. Such
a process could explain the variation from cell type to cell
type and the range of glucan structures we observe. It is also
possible that crystallization of the altered amylopectin in the
mutants is incomplete, compared with wild-type amylopectin, with a fraction of the glucan still accessible for modification. This might explain why the structure of the
amylopectin in Atisa1-1 changes slightly, both as the day
progresses and during degradation, whereas that of the wild
type does not. Further analyses are underway to evaluate
these hypotheses.
Experimental procedures
Plants and growth conditions
T-DNA insertion mutant lines from the Salk Institute (San Diego, CA,
USA) were obtained via the Nottingham Arabidopsis Stock Centre
(Nottingham, UK). Plants were grown in a controlled environment
chamber. Unless otherwise stated the conditions were as follows:
constant 20C, 75% relative humidity and a 12-h/12-h light/dark cycle, with a uniform illumination of 175 lmol photons m)2 sec)1.
Sown seeds were covered with a clear plastic lid and stratified at 4C
for 2 days. Lids were removed after the cotyledons fully emerged
(approximately 10 days after sowing). Seeds were sown either
directly onto a peat-based potting compost, or germinated first on
fine-grade seed compost then transplanted into potting compost
after 2–3 weeks.
Molecular methods
DNA was extracted using the GenEluteTM Plant genomic DNA
miniprep kit (Sigma, Buchs, Switzerland) and RNA using RNeasy
plant mini kit (Qiagen, Hombrechtikon, Switzerland), according to
the manufacturer’s instructions. RNA preparations were DNAsetreated to reduce the possibility of contamination. RT-PCR was
performed using cMasterTM RTplus PCR system kit (Eppendorf,
Basal, Switzerland) according to the manufacturer’s instructions.
The actin2 gene (At3g18780) was used as a constitutively expressed
control. Primer sequences for PCR and RT-PCR are given in
Table S1.
Native PAGE and size exclusion chromatography
Native PAGE of crude extracts of leaves was performed as described
previously (Zeeman et al., 1998b). Separation of native proteins by
size exclusion chromatography was carried out at 4C using a 50-ml
Sepharose CL-6B column (1.3 cm diameter · 38 cm length), equilibrated with extraction buffer (Zeeman et al., 1998b). The flow rate
was 0.3 ml min)1 and fractions were collected at a rate of 1 every
3 min.
Design of AtISA1 antigen and immunoblot analysis
Antibodies specific to the AtISA1 protein were purchased from
Sigma Genosys (Haverhill, Suffolk, UK). A 16-amino acid peptide
specific to AtISA1 (CFDWEGDMHLKLPQKD) was synthesized, conjugated to keyhole limpet hemacyanin and used for rabbit immunization. Anti-AtISA1 antibodies were purified by the manufacturer
via immunoaffinity purification, using a column containing the
covalently bound peptide antigen. The purified antibodies were
used to probe protein gel blots of crude extracts or concentrated
fractions eluted from the size exclusion chromatograms. Blots were
developed using Sigma-FastTM (Sigma) or Immun-StarTM (Bio-Rad,
Reinach, Switzerland) alkaline phosphatase reagents.
Carbohydrate measurements and structural analyses
Leaves were stained for starch with Lugol solution (Sigma) after
clearing in hot 80% (v/v) ethanol. To measure starch and phytoglycogen, samples (all the leaves of individual plants, unless otherwise
stated) were harvested and immediately frozen in liquid N2. Subsequent steps were conducted at 0–4C. Samples were homogenized using an all-glass homogenizer in 0.7 M perchloric acid. Soluble
and insoluble fractions were separated by centrifugation (3000 g,
15 min, 4C). Insoluble material including starch was resuspended
once in water to remove residual soluble glucans, and at least three
times in 80% (v/v) ethanol (20C), then finally resuspended in water
and stored at )20C. The soluble fraction, including phytoglycogen
was adjusted to pH 5 by adding 2 M KOH, 0.4 M MES, 0.4 M KCl.
Precipitated potassium perchlorate was removed by centrifugation
(20 000 g, 15 min, 4C) and extracts were stored at )20C. Total
glucan in each fraction was determined by measuring the amount of
glucose released by treatment with a-amylase and amyloglucosidase as described previously (Hargreaves and ap Rees, 1988).
Maltose in the soluble fraction was determined by HPAEC-PAD as
described previously (Critchley et al., 2001).
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Isoamylase function in Arabidopsis 829
For the extraction of starch and phytoglycogen for structural
analyses, two different methods were used. The first method was
that described above for starch and phytoglycogen measurement.
For each sample, equal quantities of glucan from four or more
individually extracted plants were pooled to yield sufficient
material for structural analyses. The second extraction method
was similar to that described in Zeeman et al. (1998a). Briefly,
leaves (5–10 g) were homogenized using an electric blender in
ice-cold buffered extraction medium (100 mM Tris, pH 7.0, 5 mM
EDTA). Leaf debris was removed by filtration. Starch was
removed from the filtrate by centrifugation and contamination
removed by several washes with 1% (w/v) SDS. The cleaned
starch was rinsed five times with water. The supernatant of the
filtrate was heated rapidly to 95C for 30 min and precipitated
proteins removed by centrifugation (3000 g, 15 min, 4C). Phytoglycogen in the supernatant was precipitated by adjusting to 75%
(v/v) methanol, 1% (w/v) KCl and collected by centrifugation
(3000 g, 15 min, 4C).
Glucan samples (0.1 or 0.2 mg) were boiled for 15 min in water.
After isoamylase treatment to debranching glucans (Zeeman et al.,
1998a), all phytoglycogen samples and the starch samples extracted using the perchloric acid method were passed through
sequential Dowex 50 and Dowex 1 mini-columns (Harley and
Beevers, 1963) to remove contaminating proteins and charged
compounds. The neutral glucan chains were eluted with four
column volumes of water, lyophilized, and re-dissolved in a small
volume of water. Starch samples extracted using buffered medium
were clean and did not require this treatment. Using these samples
we compared untreated with Dowex-treated samples to confirm
that the Dowex procedure did not alter the results of the subsequent HPAEC-PAD analyses. For the analysis of starch in the
epidermis, peels were immediately frozen in liquid N2. Samples
were homogenized in a ground glass homogenizer in buffered
extraction medium (as above). In this case, starch was not
separated from other insoluble material, but instead washed once
in water, three times in 80% (v/v) ethanol and finally resuspended
in water. Starch in the insoluble material was debranched and
passed through Dowex columns as described above. The HPAEC
gradient used for the separation of chain lengths was as described
in Zeeman et al. (1998a).
b-Limit glucans were prepared by incubating glucan samples (0.2
or 0.4 mg) with 100 units of b-amylase (Megazyme, Bray, Ireland) for
2 h at 37C in 10 mM MES, pH 6.0. The b-limit glucans were
precipitated by the addition of 3 volumes of methanol, redissolved
in water and then debranched by sequential incubations with
isoamylase (Zeeman et al., 1998a) and pullulanase (Megazyme; 1
unit of enzyme for 2 h at 37C in 10 mM Na acetate, pH 5.2).
Subsequent sample treatment and HPAEC-PAD analyses were
performed as described above.
Transmission electron microscopy
Transverse sections of leaves were cut with a razor blade and
fixed in 3% glutaraldehyde in 0.1 M cacodylate buffer, pH 7.4 for
4 h. Tissue was washed three times in buffer, post-fixed overnight in 1% (w/v) aqueous osmium tetroxide, then dehydrated in
an ethanol series of increasing 10% steps and transferred to
acetone. The sections were then infiltrated and embedded in
epoxy resin (Spurr’s, Agar Scientific, Stansted, UK). Ultra-thin
sections were cut with a diamond knife, collected on copper
grids and stained sequentially with uranyl acetate and Reynold’s
lead citrate. Stained sections were examined and photographed
in a JEOL 1200EX/B electron microscope (JEOL, Welwyn Garden
City, UK).
ª Blackwell Publishing Ltd, The Plant Journal, (2005), 41, 815–830
Gas exchange measurements
Gas exchange measurements were made using a CIRAS 1 infra-red
gas analyser (PP Systems, Hitchin, UK). Fully expanded, attached
leaves of mature plants were carefully placed in the cuvette at the
start of the photoperiod and CO2 uptake rate was recorded at 15-min
intervals.
Acknowledgements
We thank Cris Kuhlemeier for his support, Alison Smith and Cathie
Martin for valuable discussions, Christopher Ball, Rebecca Alder
and Andrew Chapple for assistance in growing the plants and Pierre
Haldimann for help in making the measurements of photosynthetic
rate. We also thank the Salk Institute Genomic Analysis Laboratory
for providing the sequence-indexed Arabidopsis T-DNA insertion
mutants. The work was funded by the Swiss National Science
Foundation (Grant 3100-067312.01/1) and the Swiss National Centre
for Competence in Research (Plant Survival).
Supplementary Material
The following material is available from http://www.
blackwellpublishing.com/products/journals/suppmat/TPJ/TPJ2348/
TPJ2348sm.htm
Figure S1. Estimation of the native molecular weight of Arabidopsis
isoamylase D1.
Figure S2. Estimation of the native molecular weight of the AtISA1
protein in extracts of wild-type and Atisa2-1 plants.
Figure S3. Comparisons of the chain length distributions of glucans
extracted from leaves of wild-type, Atisa1-1, Atisa2-1 and the Atisa11/Atisa2-1 double mutant plants using ice-cold buffered extraction
medium.
Figure S4. Comparisons of the chain length distributions of glucans
extracted from leaves of wild-type, Atisa1-1, Atisa2-1 and the Atisa11/Atisa2-1 double mutant plants using perchloric acid.
Figure S5. Comparisons of the chain length distributions of wildtype and Atisa1-1 glucans after b-amylolysis of the external chains.
Figure S6. Comparisons of the chain length distributions of wildtype and Atisa-1 glucans after b-amylolysis of the external chains.
Table S1 Primer sequences for PCR and RT-PCR amplifications
Supplementary references
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