Proteome Dynamics during Plastid Differentiation in Rice1[W]
Torsten Kleffmann, Anne von Zychlinski, Doris Russenberger, Matthias Hirsch-Hoffmann, Peter Gehrig,
Wilhelm Gruissem, and Sacha Baginsky*
Institute of Plant Sciences, Eidgenössische Technische Hochschule Zurich, 8092 Zurich, Switzerland (T.K., A.v.Z.,
D.R., M.H.-H., W.G., S.B.); and Functional Genomics Center Zurich, 8057 Zurich, Switzerland (P.G., W.G.)
We have analyzed proteome dynamics during light-induced development of rice (Oryza sativa) chloroplasts from etioplasts
using quantitative two-dimensional gel electrophoresis and tandem mass spectrometry protein identification. In the dark, the
etioplast allocates the main proportion of total protein mass to carbohydrate and amino acid metabolism and a surprisingly
high number of proteins to the regulation and expression of plastid genes. Chaperones, proteins for photosynthetic energy
metabolism, and enzymes of the tetrapyrrole pathway were identified among the most abundant etioplast proteins. The
detection of 13 N-terminal acetylated peptides allowed us to map the exact localization of the transit peptide cleavage site,
demonstrating good agreement with the prediction for most proteins. Based on the quantitative etioplast proteome map, we
examined early light-induced changes during chloroplast development. The transition from heterotrophic metabolism to
photosynthesis-supported autotrophic metabolism was already detectable 2 h after illumination and affected most essential
metabolic modules. Enzymes in carbohydrate metabolism, photosynthesis, and gene expression were up-regulated, whereas
enzymes in amino acid and fatty acid metabolism were significantly decreased in relative abundance. Enzymes involved in
nucleotide metabolism, tetrapyrrole biosynthesis, and redox regulation remained unchanged. Phosphoprotein-specific staining
at different time points during chloroplast development revealed light-induced phosphorylation of a nuclear-encoded plastid
RNA-binding protein, consistent with changes in plastid RNA metabolism. Quantitative information about all identified
proteins and their regulation by light is available in plprot, the plastid proteome database (http://www.plprot.ethz.ch).
Plastids perform essential biosynthetic and metabolic functions in plants, including photosynthetic
carbon fixation and synthesis of amino acids, fatty
acids, starch, and secondary metabolites (Neuhaus
and Emes, 2000; Lopez-Juez and Pyke, 2005). In response to tissue-specific and environmental signals,
they differentiate into specialized plastid types that
can be distinguished by their structure, pigment composition (color), and function. Examples of such different plastid types are elaioplasts in seed endosperm,
chromoplasts in fruits and petals, amyloplasts in roots,
etioplasts in dark-grown leaves, and chloroplasts in
photosynthetically active leaf tissues (Neuhaus and
Emes, 2000; Lopez-Juez and Pyke, 2005). Depending
on their specific biosynthetic activity and energy
metabolism, plastids are broadly classified as photosynthetic and nonphotosynthetic plant organelles.
Photosynthetic chloroplasts synthesize sugar phosphates that are catabolized by oxidative metabolism
1
This work was supported by funds from the Eidgenössische
Technische Hochschule Zurich and Strategic Excellence Project Life
Sciences (to W.G. and S.B.) and generous fellowships from the
VELUX foundation (to A.Z.).
* Corresponding author; e-mail [email protected]; fax 41–1–632–
10–79.
The author responsible for distribution of materials integral to the
findings presented in this article in accordance with the policy
described in the Instructions for Authors (www.plantphysiol.org) is:
Sacha Baginsky ([email protected]).
[W]
The online version of this article contains Web-only data.
www.plantphysiol.org/cgi/doi/10.1104/pp.106.090738
912
to produce NADPH and ATP. Nonphotosynthetic
plastid types import cytosolic sugar phosphates and
ATP, which are necessary to sustain their anabolic
metabolism. This difference in energy metabolism is
often used to distinguish the autotrophic chloroplast
from heterotrophic plastids.
Illumination of plant leaves that have developed in
the dark triggers the conversion of etioplasts into
chloroplasts. This plastid differentiation process is
paralleled by the transition from heterotrophic to
autotrophic energy metabolism, which involves massive reorganization of the etioplast proteome to support photosynthesis-dependent autotrophic growth.
General aspects of this transition have been investigated using different plants and were mainly focused
on the accumulation of individual proteins and synthesis of chlorophyll and other pigments (for review,
see Lopez-Juez and Pyke, 2005). A more comprehensive analysis of nonphotosynthetic plastid metabolism
revealed considerable metabolite fluxes between different metabolic pathways, such as carbon flux to
starch, amino acids, lipids, and protein biosynthesis
(Neuhaus and Emes, 2000; Tetlow et al., 2004). It is
therefore conceivable that the transition from heterotrophic to autotrophic plastid metabolism involves
most essential metabolic functions to support the
availability of sufficient energy and reduction equivalents for different metabolic pathways.
Differentiation of chloroplasts from etioplasts in
response to light requires coordination and integration
of plastid and nuclear gene expression. For example,
the nucleus controls plastid gene expression through
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Proteomics of Plastid Differentiation
import of specific nuclear-encoded sigma factors, protein kinases that phosphorylate transcription factors,
or nucleases and RNA-binding proteins (RNPs) that
control plastid RNA stability (for review, see Link,
2003; Bollenbach et al., 2004). Plastids, in turn, regulate
the expression of nuclear genes for photosynthetic
proteins by feedback controls that are not fully understood, but most likely involve specific tetrapyrrole
pathway intermediates (for review, see Surpin et al.,
2002; Strand et al., 2003; Beck, 2005). More recent data
further suggest a signaling function of plastid gene
expression and a role of photosynthetic redox signals
in plastid-to-nucleus signaling (Fey et al., 2005; Pesaresi
et al., 2006). Although global adaptation of plant gene
expression to light requires integrated control of plastid
and nuclear gene expression, very early events must
involve plastid gene expression and the plastid protein
complement. This is important because light-induced
synthesis of chlorophyll generates potentially harmful
molecules that are photoreactive and must be rapidly
scavenged at the site where they are synthesized (i.e.
by the plastid’s intrinsic protection system; Apel and
Hirt, 2004).
To better understand the response of etioplasts to
light-induced signal transduction pathways that initiate chloroplast differentiation processes, it is possible
to analyze rearrangements in the etioplast proteome in
response to light. Several proteomics studies using different plant cell organelles have already been reported
(for review, see Peck, 2005; Agrawal and Rakwal, 2006;
Baginsky and Gruissem, 2006). Although a few studies analyzed chloroplast proteome dynamics (Lonosky
et al., 2004; Giacomelli et al., 2006), most were focused
on a detailed, but static, analysis of the chloroplast
proteome or suborganellar compartments (for review, see van Wijk, 2004; Baginsky and Gruissem,
2004, 2006). This approach has certain limitations
because chloroplasts have highly abundant photosynthetic proteins that impact the dynamic range and
detection of low-abundance proteins (Baginsky et al.,
2005). This becomes apparent when the identical (or
nearly identical) set of chloroplast proteins was repeatedly identified in different studies, whereas the
reported detection rate of new proteins is small
(Kleffmann et al., 2006; Peltier et al., 2006). This is
not unexpected because most abundant chloroplast
proteins are involved in photosynthesis. Moreover,
profiling of static proteomes limits information on proteome dynamics, such as the ordered rearrangement of
the proteome during plastid differentiation. Transcriptional profiling can only be indicative of quantitative
changes at the transcript level and does not provide
information about protein levels or posttranslational
modifications (e.g. Belostotsky and Rose, 2005). Insight
into the regulation of proteome dynamics is essential,
however, to understand how proteome changes affect metabolic activities and regulatory processes during plastid differentiation (Neuhaus and Emes, 2000;
Weber et al., 2005). Plastid protein dynamics most
likely also relate to different protein-targeting routes
that exist in plastids, but comprehensive information
on the regulation of protein targeting during plastid
differentiation is currently not available (Jarvis and
Robinson, 2004).
Here we report a targeted proteomics approach to
explore protein dynamics during the differentiation
of rice (Oryza sativa) etioplasts into chloroplasts. Analysis of the etioplast proteome is not constrained by
abundant photosynthetic proteins, therefore shifting
the dynamic range to facilitate the detection of lowabundance proteins that have regulatory functions
(e.g. transcription, RNA metabolism; Zychlinski et al.,
2005). Using two-dimensional (2-D) gel electrophoresis
coupled with tandem mass spectrometry (MS/MS)
analysis, we have developed a detailed 2-D map of the
etioplast proteome. The etioplast proteome 2-D map
provides insight into protein abundance and the prevalence of different metabolic pathways that constitute
etioplast metabolism. Based on proteome dynamics
during the first 8 h after light induction, we present
a comprehensive model of global reorganization
of heterotrophic etioplast metabolism to autotrophic
chloroplast metabolism. All protein data can be accessed via an interactive 2-D proteome map at plprot,
the plastid proteome database (http://www.plprot.
ethz.ch; Kleffmann et al., 2006).
RESULTS AND DISCUSSION
The Rice Etioplast Proteome
We recently reported a new method to isolate intact and pure etioplasts from etiolated rice leaves
(Zychlinski et al., 2005), which is based on several differential centrifugation steps followed by Nycodenz
density gradient centrifugation. Semiquantitative protein profiling along the Nycodenz gradient established
that isolated etioplasts were free of protein contamination from other cell organelles (Zychlinski et al.,
2005). For the proteome analysis reported here, we
isolated etioplasts from 7-d-old rice seedlings, solubilized the protein in detergent-containing buffer,
and established a quantitative proteome map by 2-D
gel electrophoresis. Proteins from stained spots were
identified by matrix-assisted laser-desorption ionization (MALDI)-tandem time-of-flight (TOF/TOF) analysis using the MASCOT database search algorithm for
unambiguous protein identification (see ‘‘Materials
and Methods’’). For assignment of protein quantities,
we integrated data from four independent experiments (biological replicates) and averaged the proteinstaining intensities (Sypro Ruby) for each protein spot.
Using this approach, we identified 369 proteins with
high confidence. We compared these data with our
shotgun proteome analysis of the etioplast proteome
(Zychlinski et al., 2005) and found that 237 etioplast
proteins were exclusively identified using the 2-D
approach and 118 proteins exclusively by the shotgun
approach.
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Kleffmann et al.
This difference in protein detection can be attributed
to the different physicochemical properties of proteins
that influence their identification in the two different
proteomics strategies. Analysis of membrane proteins
by 2-D gel electrophoresis is seriously limited by
solubility constraints that have a minor impact on
shotgun proteomics. As expected, membrane proteins,
such as transporters and several plastid-encoded proteins, are significantly underrepresented on the 2-D
gel, whereas they were readily detected with the
shotgun approach. On the other hand, we reached
greater overall proteome coverage with the 2-D PAGE
approach reported here because we could detect lowabundance proteins, such as 3-dehydroquinate and
chorismate synthase. These enzymes are active in the
synthesis of aromatic amino acids and represent only
two examples of proteins that were detected on the
2-D map, but not with the shotgun approach. In
general, proteome analyses reach their greatest depth
when they combine different strategies that exploit the
diverse physicochemical properties of proteins. Combining both studies, we have identified 477 unique
proteins from rice etioplasts, which makes the etioplast proteome the best-characterized proteome of all
heterotrophic plastid types analyzed to date.
BLAST searches of etioplast proteins against all plastid proteins that were already identified in large-scale
proteome analyses (Altschul et al., 1997; Kleffmann
et al., 2006) revealed that 88 etioplast proteins were
not previously detected in proteome studies of chloroplasts or undifferentiated Bright-Yellow 2 plastids.
When compared to other plastid types, these proteins
are more prevalent in etioplasts and therefore point to
etioplast-specific metabolic and regulatory functions.
Most of these proteins have no reported function and
therefore may represent new metabolic activities or
regulatory mechanisms (Supplemental Table S1). Additionally, several of the proteins previously unidentified in large-scale proteome analyses are involved
in plastid gene expression comprising ribosomal proteins, an RNA helicase, and the a-subunit of the
plastid-encoded RNA polymerase. This supports our
earlier conclusion that the etioplast gene expression
apparatus appears to be highly tuned to support a
rapid response of plastid gene expression to light
(Zychlinski et al., 2005). The prevalence of proteins
involved in gene expression distinguishes etioplasts
from other fully developed and differentiated plastid
types. Together, our results demonstrate that proteome
analysis of different plastid types can be used as an
efficient strategy to increase the number of newly
identified proteins on the way to establish the complete proteome of higher plant plastids.
Plastid Protein Targeting
Most of the nuclear-encoded plastid proteins contain N-terminal transit peptides (TPs). On the basis of
TP predictions from genome sequences, the approximate number of expected plastid proteins has been
calculated (Abdallah et al., 2000). The robustness of
this approach depends on the sensitivity and specificity value of the target prediction algorithm. The
specificity value indicates how many proteins predicted to localize to plastids are true plastid proteins
(Emanuelsson et al., 2000). The proteomics approach
we have used does not allow conclusions about specificity because lack of protein detection does not necessarily suggest a false-positive prediction. However,
proteomics can provide information about the sensitivity of target prediction algorithms. Provided that
the set of etioplast proteins reported here comprises
only true plastid proteins, the sensitivity of correct
target prediction is 68% (Table I). This value is lower
than reported (Emanuelsson et al., 2000), but consistent with previously reported plastid proteome and
bioinformatics analyses (Kleffmann et al., 2004; Richly
and Leister, 2004; Nair and Rost, 2005; Zychlinski et al.,
2005; Peltier et al., 2006). Of all etioplast proteins we
have identified, TargetP predicts that 68% are localized
to plastids, 14% to mitochondria, 14% to any other
location, and 3% to the secretory pathway (Table I).
Although we identified a small number of putative
contaminants, most of the proteins not predicted by
TargetP are true plastid proteins. Examples include the
putative shikimate/quinate 5-dehydrogenase and
stroma ascorbate peroxidase, both well-known plastid
proteins that are predicted by TargetP for the secretory
pathway and a plastid diaminopimelate decarboxylase that is predicted for any other location.
False-negative prediction of true plastid proteins
can have several reasons. TPs differ in amino acid
composition, which makes reliable prediction per se
difficult (Bruce, 2001). To identify differences in TP
composition between identified rice etioplast proteins
(this study) and Arabidopsis (Arabidopsis thaliana)
Table I. Identified proteins and prediction of subcellular localization
a
TargetP Prediction
Shotguna
2-D PAGE
In Both
Nonredundant
Plastid
Mitochondria
Secretory pathway
Any other location
Plastid genome
Total
168
22
10
24
16
240
245
55
11
50
8
369
97
11
5
11
8
132
316
66
16
63
16
477
As reported by Zychlinski et al. (2005).
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Proteomics of Plastid Differentiation
chloroplast proteins (Kleffmann et al., 2004), we
aligned 39 N-terminal sequences (50 amino acids) of
those rice proteins without a predictable plastid TP
that have a homolog in Arabidopsis with a predictable
plastid TP. The alignment supports the view that the
difference in targeting prediction of evolutionary related proteins stems from different TP compositions
because the 39 N termini from rice are significantly
enriched in the unpolar/aliphatic amino acids Ala and
Leu (Supplemental Fig. S1A). A more general alignment of 100 randomly chosen Arabidopsis and rice
proteins with a predicted TP supports the view that
the occurrence of Ala and Leu distinguishes rice TPs
from those of Arabidopsis. The latter are enriched
in the polar amino acid Ser (Supplemental Fig. S1, B
and C).
Furthermore, different TP structures could have
evolved to maintain efficient import of low-abundance
proteins in the presence of highly abundant photosynthetic proteins (Kubis et al., 2003, 2004; Ivanova et al.,
2004; Baldwin et al., 2005). We have recently shown
that TargetP prediction sensitivity decreases for nuclearencoded plastid proteins with lower transcript levels
(Baginsky et al., 2005). This could suggest that lowabundance proteins use TPs with modified amino acid
composition that cannot be recognized by TargetP
(Jarvis and Robinson, 2004). Another reason for the
false-negative plastid-targeting prediction is the existence of alternative import routes that differ from the
common inner translocation chloroplast/outer translocation chloroplast complex protein import pathway.
Recent reports on chloroplast protein import via the
secretory pathway (Chen et al., 2004; Villarejo et al.,
2005) and the existence of internal targeting information (Miras et al., 2002) support this view. Furthermore, dual targeting has already been reported for a
substantial number of plastid proteins that likely contribute significantly to intracellular protein trafficking
(Silva-Filho, 2003).
Despite the exceptions of plastid protein import
discussed above, primary import of plastid proteins
occurs via translocation by the outer translocation
chloroplast and inner translocation chloroplast complexes (for review, see Soll and Schleiff, 2004; Bedard
and Jarvis, 2005; Kessler and Schnell, 2006). Following
protein translocation, the N-terminal TP is recognized
and cleaved by a stromal-processing peptidase (SPP;
Richter and Lamppa, 2003). TPs have no significant
sequence or structure conservation, suggesting that
the substrate specificity of SPP may be rather low. It
was suggested, however, that cysteins in the TP are
involved in the cleavage reaction by coordinating the
zinc ion in the SPP catalytic site (Richter and Lamppa,
2003). We analyzed the MS spectra of the rice etioplast
proteins for acetylated N termini to obtain new insights into TP length and exact localization of the
cleavage site. N-terminal acetylation is a common
posttranslational modification of proteins in eukaryotes and has also been reported for a number of
chloroplast proteins now (Gomez et al., 2002; Ferro
et al., 2003). Chloroplasts contain at least one putative
N-acetyltransferase that was recently identified in
Arabidopsis chloroplasts (Kleffmann et al., 2004) supporting enzyme-catalyzed N-terminal acetylation. We
Table II. Acetylated N-terminal peptides that were identified by MALDI-TOF/TOF analyses
Some peptides were identified several times; in these cases, only the highest Mascot Score is provided.
Protein
Identifier
Identified Acetylated Peptide
Peptide
Start-End
TP-Length
(Cleavage Site)a
Mascot
Scoreb
Expressed protein
Expressed protein
29-kD ribonucleoprotein A,
chloroplast precursor
Plastid-specific ribosomal
protein 2
Expressed protein
RNA recognition motif family
protein
Chloroplast inositol phosphatase
Glucose-1-P adenylyltransferase
large subunit 1
3-Oxoacyl-(acyl-carrier-protein)
synthase III
Expressed protein
Isocitrate dehydrogenase, NADP
dependent
Putative enoyl-ACP reductase
Expressed protein
LOC_Os03g21370
LOC_Os03g01030
LOC_Os07g43810
Ac-AVAVDSDQQGSPEPPDQEAKPK
Ac-ADLLGDFGAR
Ac-VAVSSEVEEEEGGAESEGEFAEDLK
48-69
44-53
57-81
47 (C/RC4)
43 (C/RC3)
38 (C/RC2)
112
41
210
LOC_Os09g10760
Ac-SSSVLEAPEEVAAR
57-70
53 (C/RC1)
127
LOC_Os05g01970
LOC_Os03g25960
38-55
58-87
37 (C/RC5)
56 (C/RC1)
129
101
LOC_Os07g37250
LOC_Os05g50380
Ac-AAAGGPLPTVLVTGAGGR
Ac-VAVSEEVETEEDEEEEEEGSGGEEFSDDLR
Ac-VATAGDVPPTVAETK
Ac-VLTSDAGPDTLHVR
51-65
61-74
49 (C/RC5)
74 (C/RC5)
27
106
LOC_Os04g55060
Ac-ASTVDDGVVSAAAAPKPR
52-69
51 (M/RC4)
142
LOC_Os08g15500
LOC_Os04g42920
Ac-(A)AAASAPAPATPVQAQQR
Ac-AAAAAAVAEQHR
46/47-62
48-59
67 (C/RC2)
47 (C/RC4)
80/97
61
LOC_Os08g23810
LOC_Os10g30870
Ac-AM(ox)SSESGPQGLPIDLR
Ac-VILGPDGRPIGGGPR
138-153
48-62
– (AO/RC2)
46 (C/RC2)
84
49
a
Prediction of localization and cleavage site was performed with TargetP; in parentheses, prediction of localization (C, chloroplast; M,
b
mitochondria; S, secretory pathway; AO, any other) and reliability class of prediction (1–5).
MASCOT scores above 34 (P , 0.05) were
considered significant; all spectra were manually examined for correct assignment.
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Kleffmann et al.
detected 13 acetylated N termini (Table II), which
closely match the cleavage site predicted by TargetP
and suggest an average TP length of 50 amino acids
(Table II). Although this number of acetylated N
termini is still small, the data are nevertheless useful
to obtain additional information about the chemical
properties of the actual cleavage site. Statistical analysis of multiple sequence alignments around the
cleavage site points to several Args located at various
distances upstream of the cleavage site and an enrichment of aliphatic amino acids immediately proximal to
the cleavage site (Supplemental Fig. S2).
Etioplast Pathways and Metabolic Activities
The rice etioplast proteome map provides quantitative information about the identified proteins that can
be used to estimate relative pathway prevalence. This
is feasible because Sypro Ruby is a proportional stain
that provides direct correlation between staining intensity and protein abundance. Because membrane
proteins are not completely soluble in mild detergent,
they are not included in the quantitative analysis.
Figure 1 shows a comprehensive presentation of total
protein mass distribution relative to specific metabolic
functions. The largest group of identified proteins (diamonds, top graph) and 38% of the total assigned protein masses (gray bars, top graph) represent amino acid
(16%) and carbohydrate (22%) metabolism pathways.
In addition, approximately 28% of the total assigned
protein masses represent proteins involved in gene
expression, secondary metabolism, and tetrapyrrole
biosynthesis (Fig. 1). Proteins that participate in nucleotide and fatty acid metabolism are represented only by
a small number of total assigned protein masses.
Figure 1 also shows individual protein abundance in
each functional category as the median value of spot
intensity. Interestingly, based on this analysis, chaperones are among the most abundant etioplast proteins,
especially the 60-kD chaperonin (CPN60), the heat
shock 70-kD protein (HSP70), and a 10-kD chaperonin
(Supplemental Table S2). Proteins that function in
photosynthesis and the subunits of the ATP synthase
complex are already moderately abundant in the dark.
An important biosynthetic function of etioplasts is the
synthesis of chlorophyll via the tetrapyrrole pathway.
In the dark, this pathway terminates at protochlorophyllide because subsequent reactions depend on the
presence of light (for review, see Grimm, 1998). As
expected, Figure 2 shows that enzymes involved in
chlorophyll biosynthesis belong to the most abundant
proteins in etioplasts, consistent with rapid production of chlorophyll after illumination. All other metabolic pathways are represented by individual proteins
of average abundance.
Etioplasts rely on the import of cytosolic sugar
phosphates and ATP for their energy metabolism
(Neuhaus and Emes, 2000; Fischer and Weber, 2002;
Weber et al., 2005). Our recent proteomics results
confirm biochemical data that etioplasts import triose
phosphates via a triose phosphate translocator, but not
hexose phosphates (Neuhaus and Emes, 2000; Tetlow
et al., 2004; Zychlinski et al., 2005). The conversion of
triose phosphates into hexose phosphates (e.g. to feed
the oxidative pentose phosphate pathway [OPPP])
requires, among other enzymes, Fru 1,6-bisphosphatase, which is activated by reduced thioredoxin
(Scheibe, 1994; Scheibe et al., 2005). This type of redox
regulation was thought to be restricted to photosynthetic chloroplasts, which reduce ferredoxin and thioredoxin via their photosynthetic electron transport. We
identified low concentrations of Fru 1,6-bisphosphatase (0.162 staining intensity), suggesting that etioplasts
may convert imported triose phosphates into hexose
phosphates. Similar to nonphotosynthetic amyloplasts
(Balmer et al., 2006), etioplasts also contain thioredoxin
(m type, 0.3 staining intensity), ferredoxin-NADP reductase (0.717 staining intensity), ferredoxin (Zychlinski et al.,
2005), and a putative thioredoxin reductase (0.257
staining intensity), suggesting that thioredoxin reduction could also occur in nonphotosynthetic etioplasts
Figure 1. Quantitation of etioplast proteins based on 2-D gel electrophoresis and Sypro Ruby staining. Identified proteins (in total 369) were
classified by their molecular function. The top graph shows the number
of identified proteins in each functional category (black diamonds) and
the sum of all individual staining intensities of the proteins in each
functional category (gray bars). Percentage of total mass was calculated
by dividing the summed staining intensities of the proteins in each
category by the total staining intensity of all identified proteins on the
gel. The bottom graph shows the distribution of staining intensities of
the proteins in each functional category as box plots. Median values
(line in the box), the 25% and 75% percentile (bottom and top end of
the box), the 5% and 95% percentile (bottom and top bars), and the
outlier (single dots) are indicated.
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Proteomics of Plastid Differentiation
Figure 2. Sugar phosphate import, glycolysis, and OPPP. Depicted are the identified proteins and their abundance (staining
intensity in the 2-D map [squares] and number of peptides in shotgun proteomics [circles]) together with potential
interconnections between these two pathways (glycolysis and OPPP).
(Schürmann, 2003; Balmer et al., 2006; Fig. 2). Detection
of a set of enzymes that enable etioplasts to catalyze
the conversion of triose phosphates into hexose phosphates suggests that etioplasts feed Glc 6-P into the
OPPP (Fig. 2).
Early Events in Light-Induced Differentiation of
Etioplasts to Chloroplasts
Single quantitative measurements of plastid proteomes as discussed above and reported in the literature to date do not capture proteome dynamics and
therefore do not reveal regulatory aspects of plastid
differentiation. Taking advantage of the 2-D map, we
explored the response of the etioplast proteome to
illumination to address two questions. First, what are
early regulatory processes that integrate the light
signal with control of plastid gene expression; and
second, what is the sequence of events in the transition
of heterotrophic etioplast metabolism to autotrophic
chloroplast metabolism? To answer these questions,
we analyzed changes in the etioplast proteome at 2,
4, and 8 h after illumination. Figure 3A documents
the light-induced changes in plastid morphology and
chlorophyll accumulation. The etioplast-specific prolamellar body, which contains a ternary complex of
protochlorophyllide, protochlorophyllide oxidoreductase, and NADPH, is disassembled during the first 2 h
in the light as the thylakoid membrane system develops (Apel et al., 1980; Vothknecht and Westhoff, 2001;
Fig. 3A). In parallel, protochlorophyllide is photo-
reduced to chlorophyllide and further converted into
chlorophyll as indicated by red chlorophyll fluorescence (Fig. 3A).
We evaluated the suitability of our experimental
approach to reveal light-dependent regulation of proteins by following the accumulation of known lightinduced proteins during the early phase of chloroplast
development. As expected, thylakoid membrane proteins, such as the chlorophyll-binding proteins, accumulated rapidly after illumination (Fig. 3B). In parallel,
enzymes of the Calvin cycle accumulated (Fig. 4), including Fru 1,6-bisphosphatase and Rubisco (Fig. 4,
inset). Interestingly, Rubisco is one of the most abundant etioplast proteins and its light induction was
strongest among the Calvin cycle enzymes. Together,
the results confirmed that 2-D PAGE was suitable to
reveal significant increases in protein accumulation
as early as hours after illumination. We therefore chose
2-D PAGE analysis to identify molecular processes
that may be involved in the regulation of plastid and
nuclear gene expression during the early illumination
phase.
Regulatory Modules That Adjust Gene Expression
in Response to Illumination
Developmental adaptations to light depend on retrograde communication between the plastid and
nucleus that may involve intermediates of the tetrapyrrole pathway (for review, see Nott et al., 2006). We
therefore analyzed whether the abundance of enzymes
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Kleffmann et al.
Figure 3. Light-induced development of chloroplasts from etioplasts.
A, Plastids that were isolated from etiolated rice plants after the
indicated illumination times. The left image shows light microscopy
images of the plastids and the right image shows the corresponding red
chlorophyll fluorescence. B, Regulation of chlorophyll-binding proteins
by light. We followed the spot intensity of chlorophyll-binding proteins
on the different 2-D gels after the indicated illumination times. All
chlorophyll-binding proteins were up-regulated by light. Protein spots
are: 1 and 2, LHC II type I (CAB21); 3 to 5, LHC II type I (CAB 26); 6,
LHC II type III.
active in this pathway was regulated by illumination.
With the exception of glutamyl-tRNA reductase
(GluTR), all known enzymes of the tetrapyrrole pathway were detected, confirming that they accumulate
to moderately high levels in the dark (Fig. 5; Supplemental Table S2). No significant changes of enzyme
abundance were visible during the first 4 h of illumination (Fig. 5), although the flux through this pathway
is significantly increased by light (Grimm, 1998). Tetrapyrrole pathway flux is regulated by feedback inhibition of GluTR by aminolaevulinic acid (Goslings
et al., 2004). It is therefore possible that the increase in
tetrapyrrole pathway flux is regulated by the abundance of GluTR or modification of its catalytic activity.
Because we have not detected GluTR in our proteomics study, it is difficult to distinguish between these
two possibilities. We therefore compared our results
with light induction of the tetrapyrrole pathway at the
transcriptional level. To this end, we used data obtained from Arabidopsis because transcript data of the
early light induction phase are not available for rice
(Zimmermann et al., 2004). Such a comparison is valid
because recent data confirmed that transcriptional
regulation is very similar for Arabidopsis and rice at
later phases of light induction (Jiao et al., 2005). The
transcript data from Arabidopsis support our finding
that enzyme abundance is not significantly changed
during the first 2 h of illumination (Supplemental Fig.
S3). After 5 h of illumination, expression of all genes of
known enzymes involved in the tetrapyrrol pathway
was up-regulated at the transcriptional level, except
for the GluTR genes. These were expressed at very low
levels throughout the illumination period, which is
most likely the reason why GluTR was not detected at
the protein level. The proteomics and transcriptomics
data together suggest that the accumulation of enzymes of the tetrapyrrole pathway in the dark is
sufficient to support a rapid increase in pathway flux
in response to light (Fig. 5) and that tetrapyrrole
pathway flux is likely increased by modification of
the GluTR catalytic activity rather than by increased
GluTR abundance.
Light also controls changes in plastid gene expression. For example, control of plastid mRNA stability is
an important mechanism in the regulation of plastid
gene expression (for review, see Hayes et al., 1999;
Bollenbach et al., 2004). Transcription rates of many
chloroplast DNA-encoded genes do not change significantly during chloroplast development, whereas
the relative half-lives of plastid mRNAs increase (Klaff
and Gruissem, 1991; Stern et al., 1997). These observations suggest that genes encoding proteins for photosynthetic functions are transcribed, but their mRNAs
are rapidly degraded in etioplasts. This is consistent
with the presence of enzymes for the efficient degradation of structured mRNAs in etioplasts, including
helicases, endonucleases, and exonucleases (Zychlinski
et al., 2005). Thus, RNA degradation enzymes must be
controlled by light to increase the half-lives of mRNAs
from photosynthetic genes.
It was previously proposed that plastid RNPs are
involved in the regulation of mRNA stability (Hayes
et al., 1999). We have identified RNP29 in two protein spots on the 2-D map, one of which is shifted
toward a higher molecular mass and a more acidic pI
Figure 4. Quantification of Calvin cycle enzymes. We followed the
spot intensity of Calvin cycle enzymes after illumination and plotted
the staining intensity against the illumination time. Large subunit of
Rubisco, whose light induction is the strongest among all Calvin cycle
enzymes, is shown in the inset.
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Proteomics of Plastid Differentiation
Figure 5. Regulation of the tetrapyrrole biosynthesis
pathway by illumination. Spot intensities of individual enzymes of the tetrapyrrole biosynthesis pathway
after 0-, 2-, and 4-h illumination are presented.
(Fig. 6), which is characteristic of phosphorylation. ProQ-Diamond specifically stains the protein with the
higher molecular mass and more acidic pI, suggesting
that this protein represents the phosphorylated form
of RNP29 (data not shown). Full staining of both protein
spots with Sypro Ruby showed that the ratio of phosphorylated and unphosphorylated RNP29 increased
from 0.41 at 0 h to 0.88 at 2 h of illumination (Fig. 6).
Phosphorylation of RNPs alters their affinity for RNA
and has been proposed to increase the stability of
plastid mRNAs in response to external signals (Lisitsky
and Schuster, 1995; Bollenbach et al., 2004; LozaTavera et al., 2006). Light-induced RNP phosphorylation shown here could therefore protect mRNAs for
photosynthetic proteins from degradation and could
be the primary regulatory event during light-dependent
chloroplast development. Other RNA degradation enzymes, such as polyribonucleotide phosphorylase (Hayes
et al., 1996; Baginsky et al., 2001), CSP41 (Yang et al.,
1996), and an RNA helicase (Zychlinski et al., 2005) remain unchanged after 2 and 4 h of illumination (Supplemental Table S2).
translation in plastids and their molecular function
suggests that higher in vivo concentrations of these
proteins allow for higher translation rates. Efficient
translation machinery is fundamental for reorganization of proteomes during developmental processes
and necessary for fast accumulation of enzymes that
are needed in photosynthetic chloroplasts. An additional reason for the need for an efficient translation
system in developing chloroplasts is the damage of
proteins by photooxidative processes that accompany
the onset of photosynthetic electron transport. Damaged proteins must be rapidly degraded and replaced
by newly synthesized proteins (Aro et al., 1993).
The reorganization of plastid proteomes and the
removal of damaged proteins not only require protein
synthesis, but also machinery for their degradation.
We detected strong light-induced accumulation of
plastid proteases, especially of those proteases that
belong to the Clp protease system (Fig. 7; Supplemental Table S2). Clp proteases are important for plastid
viability and constitute an efficient protein degradation system (Fig. 7; for review, see Sakamoto, 2006).
Global Rearrangement of Metabolic Modules
toward Autotrophy
We next addressed the transition of heterotrophic
etioplast metabolism to the autotrophic metabolism of
developing chloroplasts. One adjustment to light
within the etioplast proteome is the accumulation of
proteins with a function in translation (Table III; Fig. 7;
Supplemental Table S2). These proteins comprise ribosomal proteins and the translation elongation factors P and Tu. Both elongation factors are of pivotal
functional importance for the translation process.
Elongation factor P stimulates the peptidyltransferase
function of ribosomes and is essential for viability in
Escherichia coli (Aoki et al., 1997). Elongation factor Tu
delivers aminoacyl tRNAs to the acceptor site of
ribosomes and is thought to be rate limiting for the
speed of translation elongation (Pape et al., 1998). It is
therefore conceivable that the availability of these
translation factors is crucial for rapid and efficient
Figure 6. Light-induced phosphorylation of a plastid RNP. The graph
shows the ratio of phosphorylated to unphosphorylated RNP29
(5850.m00157) on the basis of the Sypro Ruby staining intensity
(shown as inset). The top-left spot was identified as the phosphorylated version of RNP29 by a positive phosphoprotein-specific ProQ-Diamond stain (data not shown).
Plant Physiol. Vol. 143, 2007
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Kleffmann et al.
Table III. Light-regulated reorganization of the etioplast’s nonphotosynthetic metabolism toward
photosynthesis-dependent autotrophy
Total protein mass is shown as the sum of staining intensities of proteins in each functional category after
0-, 2-, and 4-h illumination. PS, Photosynthesis.
Amino acid
Carbohydrate
PS and energy
Gene expression
Fatty acid
Nucleotide
Redox
Chaperones
Hypothetical/unknown
Secondary
Tetrapyrrole
Proteases
Other
Dark
2-h Light
4-h Light
57.5
76.8
30.0
34.8
13.7
6.3
16.3
34.1
31.4
6.7
20.0
13.5
15.1
44.7
86.8
32.6
39.7
10.6
4.5
17.0
36.2
29.0
5.3
20.7
15.6
13.5
37.3
112.3
37.8
42.7
9.4
5.3
16.0
38.1
27.6
6.2
17.1
17.5
13.5
It can be speculated that degradation of etioplast
proteins fills the plastid amino acid reservoir, which
makes de novo synthesis of amino acids during a
phase of plastid-type transition unnecessary. This
could explain why the relative abundance of amino
acid-synthesizing enzymes decreases in the early illumination phase (Fig. 7), although most of their genes
are induced at the transcriptional level in Arabidopsis
(data acquired from Genevestigator; Zimmermann
et al., 2004; Supplemental Fig. S3). In contrast to the
enzymes of amino acid metabolism, Calvin cycle enzymes and thylakoid membrane proteins accumulate
to higher levels in the light because they are immediately needed for construction of photosynthetic
machinery. Their accumulation at the protein level
correlates with induction at the transcript level and
suggests a positive correlation between transcript and
protein levels for these proteins during the deetiolation process (Supplemental Fig. S3).
The examples reported above illustrate that data at
different levels of gene expression are necessary and
must be integrated for full comprehension of the
numerous molecular processes that constitute the development of chloroplasts from etioplasts. Quantitative information about proteins as reported here is
necessary to infer regulatory events that take place
between the expression of a gene and the metabolite
that is synthesized by the gene product. Recent analyses with Arabidopsis confirmed that, in many cases,
no apparent correlations between transcript levels,
protein activity, and metabolite accumulation exist,
suggesting that regulation occurs at different nodes in
the network (Gibon et al., 2004, 2006). Knowledge
about the concentration of an enzyme, however, may
not be sufficient to infer metabolite turnover rates
because it is often unfeasible to correlate enzyme
abundance with enzyme activity. This can be attributed to posttranslational regulation of enzyme activity.
Global analysis of all measurable posttranslational
modifications, paralleled by detailed analysis of their
effect on enzyme activity and kinetics, are additional
sources of information that, when integrated with
transcript- and protein-profiling data, provide deeper
insights into the functioning of a cell.
CONCLUSION
Our comprehensive proteome analysis of chloroplast development from etioplasts shows that this
transition is associated with a significant increase of
proteins for photosynthesis and enzymes involved in
the Calvin cycle. At the gene regulatory level, proteins
that control translation of plastid mRNAs are upregulated and mRNAs are stabilized by light-induced
phosphorylation of a plastid RNP. Because most plastid
genes encode proteins that function in photosynthesis,
Figure 7. Reorganization of etioplast metabolism after illumination.
The total protein-staining intensity for proteins in each functional
category as identified from etioplasts (dark) was set to 100%. The
absolute changes of protein mass in each functional category after
2- and 4-h illumination are provided as the difference from the dark
value (2/4-h light value minus 100%). PS, Photosynthesis.
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Proteomics of Plastid Differentiation
the increased translation rates of their mRNAs support
the rapid assembly of a functional thylakoid membrane
system. We suggest that most of the energy required
for the early phase of chloroplast development is generated by the OPPP and a partial glycolysis module. Further experiments, especially analysis of metabolite and
pathway fluxes, will be necessary to expand this model.
MATERIALS AND METHODS
Isolation of Rice Plastids
For plastid isolation, 200 g of rice (Oryza sativa cv japonica) seeds were
washed in 5% sodium hydrochloride solution for 10 min, rinsed four times
with deionized water, and swollen overnight at 29°C. Seeds were transferred
to wet Vermiculite supplemented with one-half-strength concentrated Murashige and Skoog medium (3:2 [v/v] vermiculite:medium). Seedlings were
grown in the dark at constant 29°C for 10 d. Prior to isolation of plastids, the
plants were illuminated for 0, 2, 4, and 8 h at 29°C. Plastid isolation was
performed as described before (Zychlinski et al., 2005). In brief, plastids were
purified by a combination of two consecutive density gradient centrifugation
steps using Nycodenz as the density gradient medium and several differential
centrifugation steps. Each step of the isolation procedure was carried out at
4°C. Batches of 50-g plant material, excluding the roots, were homogenized in
500 mL of etioplast isolation solution (EIS; 10 mM HEPES/KOH, pH 7.8, 2 mM
EDTA, 2 mM MgCl2, 1 mM tetrasodiumpyrophosphate, 600 mM sorbitol)
supplemented with 0.2% (w/v) bovine serum albumin using a blender. The
homogenate was filtered through two layers of Miracloth. The homogenization step and filtration were repeated once. The two filtrates were pooled and
refiltered through four layers of Miracloth. The filtrate was subsequently
centrifuged for 4 min at 200g to remove cellular debris. The supernatant was
recentrifuged for 10 min at 8,000g. Pellets containing plastids were carefully
resuspended in EIS supplemented with 0.2% (w/v) bovine serum albumin
and subjected to Nycodenz density gradient centrifugation. To this end, the
plastid suspension was adjusted with 50% Nycodenz stock solution (50%
[w/v] Nycodenz, 10 mM HEPES/KOH, pH 7.8, 2 mM EDTA, 2 mM MgCl2,
1 mM tetrasodiumpyrophosphate, 5 mM dithiothreitol [DTT]) to a final
Nycodenz concentration of 30% and a maximal volume of 20 mL. Five milliliters of plastid suspension in 30% Nycodenz were loaded into a 30-mL Corex
tube (no. 8445) and finally overlaid with a Nycodenz step gradient of 6 mL
25%, 8 mL 20%, 6 mL 15%, and 3 mL 10% Nycodenz in ESI. The gradient was
centrifuged for 45 min at 4°C and 8,000g. The two yellowish bands (bands 2
and 3) at the interface of 20% to 15% and 25% to 20% Nycodenz were
harvested from the gradient, pooled, and diluted 3-fold (v/v) with EIS plus
5 mM DTT. The organelle suspension was than centrifuged for 5 min at 8,000g
to remove the remaining Nycodenz. The pellet was resuspended in maximal
20-mL EIS per centrifugal tube and used for the second centrifugation step
at 500g for 10 min. The pellet was resuspended as before and centrifuged
again for 15 min at 500g. The resulting first plastid pellet was subjected to a
second Nycodenz gradient centrifugation using only one gradient. From the
second gradient, the two yellowish bands were harvested separately, subjected to differential centrifugation, and further processed as described before
(Zychlinski et al., 2005). Although both bands contain intact etioplasts that
do not differ in their protein pattern as judged by 2-D gel electrophoresis
(Zychlinski et al., 2005) they were further analyzed separately as time point
0 h A (top band 2) and 0 h B (bottom band 3). In the gradients of time points 2, 4,
and 8 h, only band 2 plastids were visible. Pellets containing the isolated plastids
were stored at 280°C.
2-D PAGE
Plastids isolated from plant material grown from 200g seeds were
resuspended in approximately 100-mL solubilization buffer (40 mM Trisbase, 7 M urea, 2 M thiourea, 2% CHAPS, 0.5% Brij 35, 0.4% carrier ampholites,
2 mM tributyl phosphine, 20 mM DTT, protease inhibitor cocktail EDTA-free
[Roche]) and adjusted to a protein concentration of at least 1 mg/mL. Insoluble
material was pelleted for 30 min at 40,000g. For the first dimension, 100 mg of
protein were loaded in a total volume of 420-mL solubilization buffer without
Tris-base by in-gel rehydration onto 24-cm-long strips with an immobilized
linear pH gradient from 4 to 7 (Bio-Rad). Rehydration was performed
overnight. Proteins were separated using the IPGphor (GE Healthcare, formerly Amersham Biosciences) for a total of approximately 80 kVh by the
following voltage gradient: 3 h from 500 to 2,000 V, 1.5 h from 2,000 to 4,000 V,
15 h at 4,000 V, 0.5 h from 4,000 to 8,000 V, and up to 80 kVh at 8,000 V. Prior to
the second dimension, the immobilized pH gradient strips were equilibrated
for the reduction/alkylation steps, first for 10 min in equilibration buffer (6 M
urea, 2% SDS, 50 mM Tris/HCl, pH 8.8, 20% glycerol) containing 2% DTT and
next for 10 min in equilibration buffer containing 2.5% iodacetamide. The
second dimension was performed in laboratory-made homogeneous 12%
polyacrylamide gels using the Ettan Dalt II unit (GE Healthcare). From time
point 0 h A, four different plastid isolations were analyzed in four different
electrophoretic separations to include four biological and technical replicates
in this experiment. For time points 0 h B, 2 and 4 h of illumination, two
biological and technical replicates were analyzed. Altogether, this adds up to
six biological replicates for the dark control (0 h A and 0 h B) and four
biological replicates for illumination (2 and 4 h light). Only one gel was
prepared for the time point of 8 h of illumination, which was not integrated
into the quantitative time course experiment.
Protein Detection and Gel Image Analysis
Proteins were detected by in-gel staining with Sypro Ruby (Steinberg et al.,
2000) according to the manufacturer’s instructions. For detection of phosphoproteins, some gels were stained with the phosphoproptein-specific Pro-QDiamond stain (Invitrogen, formerly Molecular Probes) prior to the total
protein stain according to the manufacturer’s protocol. All gels were scanned
at a resolution of 100 mm/pixel using a Typhoon 9400 scanner (GE Healthcare). The electropherograms were analyzed with ProteomWeaver software
(Bio-Rad). Therefore, gels of each time point (0 h A, 0 h B, 2 and 4 h) were
assigned to different groups. After automated spot detection and spot
matching, replicates of one group (one time point) were averaged into an
average gel. All gels were included in the statistics. The time point 0 h A was
used as the reference group. We considered a relative staining intensity of 0.1
for statistics and protein identification. Proteins that are absent from gels of
time point 0 h and those that are strongly up-regulated at time points 2, 4, and
8 h were compared in an additional experiment. In the latter case, spot
detection and matching were manually verified.
Spot Excision and In-Gel Protein Digest
Spots were excised from gels of time point 0 h B. Only spots that were also
present in the average gel of group 0 h A and having a relative staining
intensity of at least 0.1 in the average gel of group 0 h B were considered for the
spot-picking process. Proteins that were highly up-regulated during the time
course were picked from gels of time points 4 and 8 h. Gel spots were excised
using the GelPix (Genetix) robot. Because no carryover was detected in
preliminary experiments, we excised the spots by their staining intensity in
descending order. Altogether 945 different protein spots were excised from gel
1 of group 0 h B and further processed. Protein spots that were not identified
were picked again from gel 2 of group 0 h B. Digestion, Zip tipping, and
MALDI target spotting were carried out with a Tecan liquid-handling robot
(Genesis ProTeam 150; Tecan AG) using standard protocols according to the
manufacturer’s instructions. In brief, gel slices were washed in 30% acetonitrile in 50 mM ammonium bicarbonate for 5 min at 37°C and shrunken in 80%
acetonitrile for 10 min at room temperature prior to the addition of 25 to 50 ng
trypsin in 4 mL 5 mM Tris/HCl, pH 8.2, per spot (Shevchenko et al., 1996). After
5 min, the reswollen gel pieces were overlaid with an additional 5 mL 5 mM
Tris/HCl, pH 8, and incubated for 3 h at 37°C. Digestion was stopped with
12 mL of 1% trifluoroacetic acid (TFA). Subsequently, tryptic peptides were
extracted and desalted with prewetted (with 80% acetonitril in 0.1% TFA) and
washed (with 0.1% TFA) m-C18 ZipTips (Millipore Corporation) by five
binding cycles and two washing steps in 0.1% TFA. Bound peptides were
eluted with 0.8 mL MALDI matrix solution (3 mg a-cyano-4-hydroxycinnamic
acids in 65% acetonitrile containing 0.1% TFA) and directly spotted onto the
MALDI target.
MALDI-TOF/TOF Analysis and Protein Identification
Samples were analyzed on a 4700 proteomics analyzer MALDI-TOF/TOF
system (Applied Biosystems), which is equipped with a Nd:YAG laser
operating at 200 Hz. All mass spectra were recorded in positive reflector
mode and they were generated by accumulating data from 5,000 laser pulses.
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Kleffmann et al.
First, MS spectra were recorded from the standard peptides on each of the six
calibration spots and the default calibration parameters of the instrument
were updated. Subsequently, MS spectra were recorded for all sample spots
on the plate and internally calibrated using signals from autoproteolytic
fragments of trypsin if these signals were detectable. Up to five spectral peaks
per spot that met the threshold criteria were included in the acquisition list for
MS/MS spectra. Peptide fragmentation was performed at a collision energy of
1 kV and a collision gas pressure of approximately 2 3 1027 Torr. Data from
6,000 laser pulses were summed up for each fragment ion spectrum. Global
Proteome Server explorer software (Applied Biosystems) was used for submitting data acquired with the mass spectrometer for database searching. MS
and MS/MS data were searched using Mascot version 1.9.05 (Matrix Science)
as the search engine (www.matrixscience.com) against the The Institute for
Genomic Research rice protein database (downloaded January, 2006). Typically, the following search settings were used: maximal number of missed
cleavages, 1; peptide tolerance, 25 to 50 ppm; MS/MS tolerance, 0.2 amu.
Carboxyamidomethylation of Cys was set as fixed modification and oxidation
of Met was selected as variable modification. In additional searches, acetylation of the N terminus and oxidation of Met were set as variable modifications
and semitrypsin was selected as protease specificity.
Supplemental Data
The following materials are available in the online version of this article.
Supplemental Figure S1. Comparison of the amino acid composition of
rice and Arabidopsis plastid transit peptides.
Supplemental Figure S2. Amino acid composition around the transit
peptide cleavage site.
Supplemental Figure S3. Light induction for selected proteins at the
transcriptional level.
Supplemental Table S1. Proteins only found in etioplasts.
Supplemental Table S2. List of all identified proteins with staining
intensity following illumination.
ACKNOWLEDGMENTS
We would like to thank the staff of the Functional Genomics Center Zurich
for their support and Dr. Johannes Fütterer for critical reading of the
manuscript.
Received October 2, 2006; accepted December 7, 2006; published December 22,
2006.
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