Comparative phosphoproteome profiling revealsa function of the STN8 kinase in fine-tuningof cyclic electron flow (CEF)Sonja Reilanda,b, Giovanni Finazzic,d,e,f, Anne Endlera,b, Adrian Willigg,h, Katja Baerenfallera,b, Jonas Grossmanna,b,Bertran Gerritsa,b, Dorothea Rutishausera,b, Wilhelm Gruissema,b, Jean-David Rochaixg,h, and Sacha Baginskya,b,1,2
aDepartment of Biology, Eidgenössische Technische Hochschule (ETH) Zurich, 8092 Zurich, Switzerland; bFunctional Genomics Center Zurich, 8057 Zurich,Switzerland; cCentre National Recherche Scientifique, Unité Mixte Recherche 5168, Laboratoire Physiologie Cellulaire et Végétale, F-38054 Grenoble, France;dCommissariat à l’Energie Atomique et Energies Alternatives, l’Institut de Recherches en Technologies et Sciences pour le Vivant, F-38054 Grenoble, France;eUniversité Grenoble 1, F-38041, France; fInstitut National Recherche Agronomique, UMR1200, F-38054 Grenoble, France; and Departments of gMolecularBiology and hPlant Biology, University of Geneva, 1211 Geneva, Switzerland
Edited* by Bob B. Buchanan, University of California, Berkeley, CA, and approved June 24, 2011 (received for review March 24, 2011)
Important aspects of photosynthetic electron transport efficiency inchloroplasts are controlled by protein phosphorylation. Two thyla-koid-associated kinases, STN7 and STN8, have distinct roles in short-and long-term photosynthetic acclimation to changes in light qual-ity and quantity. Although some substrates of STN7 and STN8 areknown, the complexity of this regulatory kinase system implies thatcurrently unknown substrates connect photosynthetic perfor-mance with the regulation of metabolic and regulatory functions.We performed an unbiased phosphoproteome-wide screen withArabidopsis WT and stn8 mutant plants to identify unique STN8targets. The phosphorylation status of STN7 was not affected instn8, indicating that kinases other than STN8 phosphorylate STN7under standard growth conditions. Among several putative STN8substrates, PGRL1-A is of particular importance because of its pos-sible role in the modulation of cyclic electron transfer. The STN8phosphorylation site on PGRL1-A is absent in both monocotyledon-ous plants and algae. In dicots, spectroscopic measurements withArabidopsis WT, stn7, stn8, and stn7/stn8 double-mutant plantsindicate a STN8-mediated slowing down of the transition from cy-clic to linear electron flow at the onset of illumination. This findingsuggests a possible link between protein phosphorylation by STN8and fine-tuning of cyclic electron flow during this critical step ofphotosynthesis, when the carbon assimilation is not commensurateto the electron flow capacity of the chloroplast.
phosphoproteomics | Arabidopsis thaliana
In the field of chloroplast biogenesis, interest in protein phos-phorylation historically focused on photosynthesis-related pro-
teins, with the initial discovery of thylakoid membrane proteinphosphorylation dating back to the late 1970s (1–4). Almosta decade later, AtpB, RNA-binding proteins, and transcriptionfactors were recognized as phosphoproteins in thylakoid mem-branes and stroma fractions (5–7). Because of recent large-scalefunctional genomics and phosphoproteomics approaches, ∼200chloroplast phosphoproteins are known today, and several kinaseshave been identified that are most likely involved in their phos-phorylation (8). However, the exact kinase/substrate relationshipsare not known for most of the proteins, and efforts are underwayto identify in vivo substrates of known kinases. Phosphoproteo-mics data suggest that chloroplast functions are regulated bya highly complex phosphoprotein network in which one kinasephosphorylates several substrates and one substrate is probablyphosphorylated by several kinases at different sites (9, 10).Although we currently do not understand all nodes in the
chloroplast phosphoprotein network, candidate proteins and ex-perimental tools are available to address the above questions.Two of the best-characterized chloroplast kinases are STN7 andSTN8. STN7 is the ortholog of the Stt7 kinase from Chlamydo-monas reinhardtii that was identified in screens for strains with
a defect in state transitions (11). This process balances theabsorbed light excitation energy between the two photosystems.State transitions are regulated by light quality and intensity andmediated by phosphorylation of photosystem II (PSII) light-har-vesting complex (LHCII) proteins (4, 12). It is now well estab-lished that STN7 activity is required for state transitions, althoughit is currently unclear whether STN7 directly phosphorylatesLHCII proteins or triggers their phosphorylation through a cas-cade. STN8 is a paralog of STN7 and is also associated with thethylakoid membrane system. Analyses with phosphothreonine-specific antibodies identified the D1 (PsbA) and D2 (PsbD)proteins of PSII, PsbH, CP43, and a Ca2+-sensitive thylakoidphosphoprotein, calcium-sensing receptor (CaS), as STN8 sub-strates (13–15). However, loss of STN8 function not only affectsthe phosphorylation of thylakoid membrane proteins but also theexpression of nucleus- and plastid-encoded genes for photosyn-thetic proteins (13).These data suggest multiple functional interactions of STN8
within the chloroplast phosphoprotein network that extendbeyond our current mechanistic knowledge. For example, light-quality–dependent changes of photosystem core protein phos-phorylation mediated by STN8 no longer occur in the stn7 back-ground in Arabidopsis, suggesting functional crosstalk betweenSTN7 and STN8 (16, 17). TheChlamydomonas ortholog of STN8,called Stl1, is a phosphoprotein in vivo whose phosphorylationdepends on Stt7 (18). It is conceivable that a similar crosstalkexists between the correspondingArabidopsis orthologs STN8 andSTN7. However, although STN7 is an abundant phosphoprotein,comprehensive phosphoproteome analyses failed to identify anySTN8 phosphorylation inArabidopsis chloroplasts under differentconditions (9, 10, 19). Interestingly, the sequence of the C-ter-minal region of STN7 containing the four mapped phosphory-lated sites diverges from the corresponding region in Stt7,suggesting a function of STN7 phosphorylation in adaptationprocesses that are specific to higher plants (10). Although it is
Author contributions: S.R., G.F., J.-D.R., and S.B. designed research; S.R., G.F., A.E., andA.W. performed research; S.R., G.F., K.B., J.G., B.G., D.R., W.G., J.-D.R., and S.B. analyzeddata; and G.F. and S.B. wrote the paper.
The authors declare no conflict of interest.
*This Direct Submission article had a prearranged editor.
Data deposition: The MS data and tandem MS spectra have been deposited in thePRoteomics IDEntifications (PRIDE) database, http://www.ebi.ac.uk/pride/ (accessionnos. 13754–13761).1Present address: Institute of Biochemistry and Biotechnology, Martin-Luther-UniversityHalle-Wittenberg, 06120 Halle (Saale), Germany.
2To whom correspondence should be addressed. E-mail: [emailprotected].
This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1104734108/-/DCSupplemental.
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currently unknown which kinase phosphorylates STN7, analysis ofthe phosphorylation motifs has suggested that one of the phos-phorylation sites might be used by casein kinase II (10).Here we report STN8 substrates that we identified in a com-
parative proteome-wide analysis of protein phosphorylation inWT and in STN8-deficient (stn8) plants. We quantified unphos-phorylated proteins in plastids of both genotypes by normalizedspectral counting to distinguish changes in phosphorylation statesfrom changes in protein abundance. Our data show that otherkinase(s) besides STN8 are involved in the phosphorylation ofSTN7 and establish PGRL1-A as a STN8 substrate.
Results and DiscussionPhosphoproteome Profiling from stn8 and WT Arabidopsis LeafTissue. We analyzed the leaf phosphoproteome of WT and stn8plants in three biological replicates by using a combined immo-bilized metal-ion affinity chromatography/titanium dioxide affin-ity chromatography (IMAC/TiO2) phosphopeptide enrichmentstrategy followed by LTQ-Orbitrap mass spectrometry (MS). Intotal, 15,492 spectra were assigned to 3,589 phosphopeptides and1,738 unique phosphoproteins at a false-discovery rate of 0.15%at the spectrum level. All information concerning peptide andprotein identifications are deposited in the PRoteomics IDEnti-fications (PRIDE) database (20). To extract plastid phospho-proteins, we matched this dataset against a chloroplast proteomereference table that was assembled from the overlap of two pre-viously published chloroplast proteome datasets (SI Appendix,Table S1) (10, 21). Altogether, 1,657 spectra, 294 phosphopep-tides, and 149 phosphoproteins matched with chloroplast proteins(SI Appendix, Table S2). Chloroplast phosphopeptide detectionwas similar to previously reported analyses, and 131 of thephosphoproteins of stn8 andWTwere previously identified (9, 10,19), whereas 18 unknown proteins were detected in our analysis.The reproducible detection of these chloroplast phosphoproteinssuggests that we have acquired a robust dataset that reflectsphosphorylation activity in chloroplasts under standard con-ditions. All identified phosphoproteins and peptides are providedin SI Appendix, Table S2.A global qualitative comparison of phosphoprotein identifica-
tion and phosphorylation motif utilization in WT and stn8chloroplasts revealed minor differences at the level of phospho-peptide detection (SI Appendix, Fig. S1). We therefore searchedfor quantitative differences in the phosphorylation state of chlo-roplast proteins by using the workflow presented in Fig. 1.
Quantitative phosphopeptide analysis was performed in threebiological replicates by spectral counting and extracted ionchromatogram quantification (Fig. 1A). To distinguish betweenchanges in protein abundance and changes in phosphorylationstate, we quantified the unphosphorylated proteins from the flow-through fraction of the IMAC. The protein quantification fromthe flow-through enables a quantitative comparison of the plastidproteomes in the two different genetic backgrounds (Fig. 1B andSI Appendix, Table S3). The average Spearman rank correlationcoefficient, ρ, for the spectral count data from 756 identifiedchloroplast proteins is 0.871, suggesting minor quantitativeadaptations of the chloroplast proteome to a loss of STN8 (Fig.1B). The high similarity between the plastid proteomes ofWT andstn8 plants allowed a valid quantitative comparison of proteinphosphorylation in the plastids of the two genotypes.
Quantitative Comparison of Phosphopeptide Detection in WT andSTN8-Deficient Plants. We searched for phosphoproteome differ-ences between WT and stn8 plants by comparing the spectralcount information for individual phosphopeptides in WT andstn8 datasets, which we considered to be different when theywere detected with at least three spectra in total and a twofoldhigher spectral count in WT in at least two biological replicates.We furthermore included relative phosphopeptide quantificationby comparing extracted ion chromatograms using the Progenesissoftware tool (Nonlinear Dynamics). We used both criteria, i.e.,reduced spectral count and reduced relative intensities ofextracted ion chromatograms in stn8, together to assess the effectof loss of STN8 function. This stringent combination of selectioncriteria provides a reliable assessment of those peptides whosephosphorylation is affected in stn8.We first asked whether STN7 phosphorylation was affected in
stn8 plastids. None of the STN7 phosphopeptides fulfilled thecriteria for differentially phosphorylated peptides describedabove, suggesting that STN8 is not required to maintain the STN7phosphorylation state. The extracted ion chromatogram quanti-fication confirms that STN7 phosphopeptides are phosphorylatedto a similar extent in both genotypes (Table 1 and Fig. 2). Thepeptide NALApSALR, with the phosphorylation site at serine,was even increased in abundance in stn8 in all three replicates(Table 1 and Fig. 2, which shows the third replicate). The re-versed-phase chromatography separated two isobaric phospho-peptides that were phosphorylated at different threonine residues(Fig. 2 Center). Although the peptide eluting at 45 min wasidentified as TVTEpTIDEISDGRK by manual spectrum anno-tation, the peptide that eluted at 51 min was pTVTETI-DEISDGRK, TVpTETIDEISDGRK, or a mixture of both (SIAppendix, Fig. S2). The fragmentation pattern does not allow us todistinguish between these two possibilities. The two isobaricpeptides have the same abundance inWT and stn8 plants, and thesame holds true for the doubly phosphorylated peptide pp[TVTETIDEISDGRK] in all three replicates (Table 1 and Fig. 2).Together, our data strongly suggest that STN8 is not responsiblefor STN7 phosphorylation under our experimental conditions.
Differential Quantitative Phosphopeptide Detection in WT and stn8Plants. The previously reported STN8 substrates CaS (At5G-23060) (15), the doubly phosphorylated PsbH peptide ApTQpT-VEDSSR (Table 1) (14), as well as RbcL (ATCG00490), twounknown proteins (AT1G54520 and AT5g08540), the thylakoid-associated proteins CP29 (AT3G08940), and an ATP synthasefamily protein (AT4G32260) (Table 1) were identified with higherspectral counts in WT compared with stn8 and fulfill the criteriafor differential phosphorylation. The two unknown proteins con-tain one and two transmembrane domains, respectively, and bothwere previously identified in the proteome of chloroplast mem-brane preparations (22–24). Although the thylakoid association ofa majority of the above proteins makes their phosphorylation by
1 10 100
1
10
100WT, stn8
3 bi
ol. R
epl.
IMA
C/ T
iO2
P-pep�des[XIC]
FT proteins[APEX]
A
LOG
nSp
C [F
T (stn8)
]
LOG nSpC [FT (WT)]
B
Fig. 1. Strategy for the quantification of phosphopeptides and unphos-phorylated proteins. (A) WT or stn8 samples were subjected to affinity chro-matography on IMAC or TiO2 as described in Materials and Methods.Phosphopeptides were eluted from the affinity column and identified byMS.The relative quantification of phosphopeptides in the different samples wasbased on their spectral count information and extracted ion chromatograms(XIC). The unphosphorylated peptides were collected in the flow-throughfraction and were used for the quantification of chloroplast proteins bynormalized spectral counting (nSpC) in two biological replicates. Absoluteprotein expression (APEX). (B) Comparison of the abundance of chloroplastproteins in the flow-through fractions from WT and stn8 samples. The dia-gram shows the averaged normalized spectral count information from theIMAC flow-through, plotted on a logarithmic scale.
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Table
1.Protein
andphosp
hopep
tidedetectionan
dquan
tificationin
WTan
dST
N8-defi
cien
tplants
Protein
Spectral
countreplicates
Logab
undan
ceratio(W
T/stn8)
Logratio
protein
expressiona
Iden
tifier
Annotation
Phosphopep
tide
WT(1)
WT(2)
WT(3)
Stn8(1)
Stn8(2)
Stn8(3)
Exp.1
Exp.2
Exp.3
(WT/stn8)
Rep
orted
substrates
ATC
G00
710
PsbH
pp[A
TQTV
EDSS
R(SGPR
)]10
1212
82
5−0.2/only
WTb
5.2
4.6
−0.2
p[A
TQTV
EDSS
R(SGPR
)]18
1914
2230
9−2.3
−0.3
−3.3
p[A
c-ATQ
TVED
SSR(SGPR
)]4
01
12
0−1.0
−1.7
1.3
pp[A
c-ATQ
TVED
SSR(SGPR
)]2
01
00
0ND
9.4
ND
ATC
G00
020
PsbA
p[A
c-TA
ILER
R]
——
——
——
−3.3
ND
1.9
−0.1
p[TAILER
R]
——
——
——
−0.2
ND
ND
ATC
G00
270
PsbD
p[TIALG
KFT
K]
——
——
——
2.1
ND
ND
0.1
p[A
c-TIALG
KFT
K]
30
02
00
0.8
ND
2.8
ATC
G00
280
CP4
3p[A
c-TL
FNGTL
ALA
GR]
——
——
——
−1.7
0.6
−1.0
−0.1
p[TLF
NGTL
ALA
GR]
——
——
——
Only
WT
3.4
−0.7
pp[A
c-TL
FNGTL
ALA
GR]
——
——
——
ND
ND
−1.7
AT5
G23
060
CaS
p[SGTK
FLPS
SD]
22
12
00
0.8
2.7
2.8
0.2
p[IIPAASR
SFGTR
]0
01
00
10.3
ND
ND
pp[IIPAASR
SFGTR
]—
——
——
—−0.7
ND
ND
p[LGTD
SYNFS
FAQVLS
PSR]
21
01
10
−0.5
1.5
ND
p[SFG
TRSG
TK]
10
01
00
−1.3
ND
ND
Substratesin
AT1
G68
830
STN7
p[TVTE
TIDEISD
GRK]
92
75
28
0.8
ND
−0.2
0.1
pp[TVTE
TIDEISD
GRK]
121
59
25
0.1
0.9
−1.0
p[N
ALA
SALR
]1
02
22
4−1.7
−1.0
−1.0
New
substrates
AT4
G04
020
Fibrillin
p[A
TDIDDEW
GQDGVER
]2
02
00
02.0
ND
1.1
0.4
ATC
G00
490
RbcL
p[W
SPEL
AAACEV
WK]
01
20
01
ND
ND
ND
−0.1
At1G54
520
Unkn
own
p[SSS
SSSS
QSY
SVPR
]1
02
00
1−0.3
ND
−0.3
1.6
At5G08
540
Unkn
own
p[KNSS
VEE
ETEE
EVEE
DMPW
IQEK
]3
01
10
00.2
ND
ND
0.7
AT3
G08
940
LHCB4.2
p[N
LYGEV
IGTR
TEAVDPK
]3
30
21
00.8
1.2
−0.2
−0.1
AT4
G32
260
ATP
synthase
p[A
LDSQ
IAALS
EDIVKK]
11
20
00
Only
WT
−0.4
0.5
0.2
AT1
G08
640
Unkn
own
p[G
VTF
GSF
K]
71
11
00
2.1
ND
2.4
0.6
AT4
G22
890
PGRL1
p[A
TTEQ
SGPV
GGDNVDSN
VLP
YCSINK]
23
20
01
2.2
6.0
0.7
0.8
Acetylatedphosphopep
tides
wereassigned
byProgen
esisonthebasisofMASC
OTsearch
results(M
aterialsan
dMethods).L
istedaretheiden
tified
phosphopep
tides,theirnumber
ofspectrain
theindividual
replicates,an
dtheab
undan
ceratiooftheprecu
rsorex
tractedionch
romatogram
ascalculatedbyProgen
esis.T
hequan
tificationwas
donewithoutco
nsideringphosphopep
tides
withoxidized
methioninean
ddid
notdistinguish
thesite
ofphosphorylation
inapep
tidewith
seve
ralhyd
roxy
lated
amino
acids.
Wefurthermore
quan
tified
alliden
tified
chloroplast
proteinsin
theflow-through
fractionsfrom
the
phosphopep
tideaffinitych
romatographybynorm
alized
spectral
counting.ND,notdetected;p,phosphorylation;Ac,
acetylation.
aRatio
ofthemea
nnorm
alized
spectral
countquan
tities
expressed
asLo
gbase2.
bOnly
detectedin
WTin
thestrongcationex
chan
gech
romatographyfraction3from
themem
branepreparation.
Reiland et al. PNAS Early Edition | 3 of 6
PLANTBIOLO
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STN8 possible, the extracted ion chromatogram quantificationdoes not support the conclusion that their phosphorylation is re-duced in stn8 (Table 1). Although this finding does not excludethat the two proteins are STN8 targets, our data suggest that theirphosphorylation is largely independent of STN8 under the ex-perimental conditions and is likely catalyzed by at least one otherchloroplast protein kinase.Nevertheless, three proteins showed consistently reduced
phosphorylation in stn8 based on spectral count and extractedion chromatogram quantification. One of these is fibrillin(AT4G04020), which is phosphorylated at the threonine residuein ATDIDDEWGQDGVER, which represents the most N-ter-minal tryptic peptide detectable in vivo (25). Fibrillin is associ-ated with plastoglobuli and located on the stromal side ofthylakoid membranes, a topology that supports phosphorylationby STN8. The second protein is annotated as “unknown protein”(AT1G08640). This protein was previously identified in chloro-plast proteome analyses as an envelope protein with threetransmembrane domains, and it is thought to be a solute trans-porter of cyanobacterial endosymbiotic origin (26). The phos-phorylation site in the peptide GVTFGSFKVSK is located at theN-terminal side of the three predicted transmembrane domains.Although the consistent trend in spectral count and extracted ionchromatogram quantifications is to argue for the dependence ofthis phosphorylation site on STN8, it is currently unclear how thephosphorylation of an envelope membrane protein is catalyzedby a thylakoid-associated kinase. It is possible that transientinteractions between thylakoid and chloroplast envelope mem-branes could allow for such an STN8-mediated phosphorylation.Alternatively, STN8 could be part of a phosphorylation cascadethat results in the phosphorylation of AT1G08640.The third protein with a reduced phosphorylation state in stn8 is
the PGR5-like protein PGRL1-A (AT4G22890), which wascharacterized further (SI Appendix, Figs. S3–S5). PGRL1-A isa thylakoid membrane protein with two transmembrane domainsand its N-terminal end exposed to the stromal side (27). Thisprotein is most likely phosphorylated at one of the two threonineresidues in ATTEQSGPVGGDNVDSNVLPYCSINK (SI Ap-pendix, Figs. S3 and S4). According to the predicted transit pep-tide cleavage site at position 60 (28) and in vivo large-scaleproteome mapping (25), this phosphopeptide represents the mostN-terminal tryptic peptide of the mature protein. PGRL1-A formsa protein complex with PGR5 that is associated with (but not
bound to) photosystem I (PSI). Plants that lack PGRL1-A showperturbations in their cyclic electron flow (CEF) response, whichappears as an accelerated transition from CEF to linear electronflow (LEF) during the dark-to-light transition, i.e., the activationof photosynthesis (27). We therefore investigated possible changesof CEF in stn8, which would support a functional role for PGRL1-A phosphorylation during this phase. To identify a potential effectof loss of PGRL1-A phosphorylation and to determine its speci-ficity for stn8, we measured parameters related to LEF and CEF instn8, stn7, and stn7/stn8 double mutants.
Transition from CEF to LEF Is Altered in Plants Deficient in the STN8Kinase. In steady-state photosynthesis, LEF involves both PSIIand PSI activity, producing ATP and NADPH and finally leadingto CO2 assimilation. Conversely, a significant fraction of thephoto-generated electrons can be recycled around PSI (CEF) atthe onset of illumination, i.e., when the Calvin cycle is still largelyinactive while the electron flow capacity is already at maximum(29). The transition from CEF to LEF was first assayed bymeasuring the kinetics of P700 oxidation upon illumination withfar-red light, i.e., under conditions where PSI is preferentiallyexcited. In dark-adapted conditions, slow P700 oxidation rates areobserved, whereas P700 oxidation is accelerated upon light ex-posure for 10 min (SI Appendix, Fig. S6), in agreement withprevious findings (27, 30). This period corresponds to the timerequired to activate carbon assimilation (31). Starting with dark-adapted leaves, similar kinetics was seen in all of the genotypeswhen activity was probed at the beginning of illumination (30 s;SI Appendix, Fig. S6), suggesting a similar CEF capacity. Simi-larly, no differences were measured at the end of illumination(10 min), indicating the same rate of LEF, consistent with pre-vious data (11, 13). In contrast, a faster transition from slow tofast P700 oxidation was seen in stn8 and the stn7/stn8 doublemutant compared with WT or stn7 (SI Appendix, Fig. S6 and Fig.3), suggesting a faster transition from CEF to LEF in the absenceof STN8. To confirm this conclusion, PSII activity was estimatedfrom fluorescence parameters (32) (SI Appendix, Fig. S7A) andcompared with the overall rate of electron flow [from the elec-trochromic shift (ECS); SI Appendix, Fig. S7B and Table S4] to
NALApSALR p[TVTETIDEISDGRK] pp[TVTETIDEISDGRK]W
Tstn8
Fig. 2. Phosphopeptide quantification of phosphorylation sites in STN7.Displayed is the intensity (y axis) over time during chromatography (re-tention time, x axis) for the mass window [calculated phosphopeptidemass ± 5 ppm]. (Upper) Phosphopeptide intensity in WT. (Lower) Phospho-peptide intensity in the stn8 mutant. The intensity of the precursor withhigher intensity was set to 100%, and the same y axis scale was used for theother precursor to allow a direct comparison. Presented are representativedata for the STN7 phosphopeptides from the third replicate.
Fig. 3. Transition from CEF to LEF during a dark-to-light shift is faster instn8 and stn7/stn8 than in WT. Dark-adapted leaves from WT, stn7, stn8, andthe stn7/stn8 double mutant were exposed to red light (37 μE·m−2·s−1). Atany indicated time, PSII activity (A), the sum of PSII plus PSI activity (B), therate of CEF (C), and the fraction of PSI involved in CEF (D) were measured.Data refer to the average of six samples from two independent experiments(± SE). Parameters were derived from fluorescence, P700, and ECS measure-ments as detailed in SI Appendix, SI Materials and Methods.
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derive changes in the rate of CEF. The ECS is a light-inducedshift in the absorption spectrum of some photosynthetic pig-ments, which is caused by charge separation within the two pho-tosystems (33). During illumination, PSII activity increased in thesame way in all genotypes (Fig. 3A), and the PSI + PSII electronflow rate was higher in the STN8-containing lines (Fig. 3B). Al-though the estimated rate of cyclic PSI activity (Fig. 3C) (34) wassimilar in dark- and light-adapted leaves, a faster deactivation wasobserved in stn8 and stn7/stn8 compared with the WT and stn7, inagreement with the P700-derived data (Fig. 3D), suggesting thatthe loss of PGRL1 phosphorylation does not impair CEF or LEFactivity but does impact the plant’s capacity to maintain an activeCEF during the onset of carbon assimilation. This phenotype isvery similar to the one observed in pgr5 (35) and pgrl1 (27), twomutants lacking a protein complex (PGR5/PGRL1) likely in-volved in the regulation of CEF. In both mutants, maximum CEFcapacity is unchanged relative to WT when tested under the sameconditions as here (27, 30), but a faster activation of LEF occursupon light exposure. This similarity between pgr5, pgrl1, and stn8and stn7/stn8 supports our proposal for a link between PGRL1-Aphosphorylation and CEF stability.
ConclusionThe large-scale analysis reported here demonstrates that phos-phoproteome profiling is a powerful tool for kinase characterizationand for the detection of unique and unexpected kinase targets.Because our identification of unknown STN8 substrates was basedon stringent selection criteria, we identified a restricted set of STN8targets, and our analysis most likely underestimates their number.Among them, PGRL1-A is particularly interesting because it sug-gests a link between protein phosphorylation and modulation ofelectron flow in plants. This possibility is supported by the modifiedCEF capacity observed in stn8 and the stn7/stn8 double mutant(which both lack STN8 activity) in the experimental conditionsreported above. Because the observed effect on CEF is transitory, itcould be relevant under rapidly changing light conditions, such asshading by light flecks, which is consistent with a fast-operatingcontrol mechanism such as phosphorylation. However, the fasterdeactivation of CEF observed in plants lacking STN8 has no ap-parent effect on the overall rate of carbon assimilation, which issimilar in WT, stn8, stn7, and the double mutant (Fig. 3A). Thisfinding rules out the possibility of a direct effect of STN8 on thecarbon assimilatory process, at least under the conditions exploredin this work. Recently, a protein supercomplex has been identifiedin Chlamydomonas that is capable of performing CEF and containsstoichiometric amounts of PSI, the cytochrome b6f complex (in-cluding the small subunit PetO), ferredoxin:NADPH oxidoreduc-tase (FNR), LHCI, LHCII, and PGRL1 (36). Its accumulation inthylakoids is related to the activation of the Stt7 kinase during statetransitions. The complex does not contain PGR5, and it was pro-posed that this protein would be replaced in Chlamydomonas byPetO, which also undergoes phosphorylation under state 2 con-ditions (37). Based on these findings, it is therefore tempting tospeculate that modulation of CEF by protein phosphorylation couldbe a possible leitmotif in electron flow regulation in Viridiplantae.In Chlamydomonas, this process would operate through sequester-ing of diffusing carriers within the PetO-driven cyclic supercomplex.In plants, where PetO is absent and no CEF supercomplex has beenfound so far (31), the STN8/PGRL1-A system could directly orindirectly control the transition between CEF and LEF in a freelydiffusing system through a still-unknown mechanism. Alternatively,phosphorylation of PGRL1 could stabilize a supercomplex involvedin CEF, similar to the one found in Chlamydomonas (36). Consis-tent with the specificity of the STN8/PGRL1-A regulatory pathway,a comparison of the PGRL1-A sequences in different photosyn-thetic organisms reveals that the phosphorylation site identified inArabidopsis is absent in mosses, green algae, and other marinephotosynthetic organisms (SI Appendix, Fig. S8). Intriguingly, this
site is also absent in monocots, suggesting that the N-terminalSTN8-mediated phosphorylation of PGRL1-A originated after theseparation of dicots from monocots. At present, the exact mecha-nism how PGRL1-A phosphorylation modulates the transition ki-netics between CEF and LEF remains to be explored.
Materials and MethodsPlant Material, Growth Conditions, and Protein Extraction. Arabidopsis thali-ana Col0 and stn8 (SALK 060869) seedlings were grown on soil under short-day conditions in a controlled environment chamber (8 h light/16 h dark, 100μE·m−2·s−1). Plants were harvested after 6 wk, 3 h after the start of the light,and immediately frozen in liquid nitrogen, ground, and subsequently storedat −80 °C until further analyses. Proteins were extracted exactly as describedpreviously (10) (SI Appendix, SI Materials and Methods).
In-Solution Tryptic Protein Digest. Before tryptic digestion, cysteine residueswere reduced with 10 mM DTT for 45 min at 50 °C and alkylated by 50 mMiodoacetamide for 1 h at room temperature in the dark. Trypsin (sequencinggrade; Promega) was added in a ratio of 1:20 and incubated over night at 37 °C.
Fractionation of Peptides by Strong Cation Exchange Chromatography. Pep-tides were desalted by using Sep-Pak reverse-phase cartridges (Waters),dissolved in buffer A [10 mMKH2PO4 (pH 2.6) in 25% acetonitrile] and loadedonto a 4.6 × 200 mm PolySULFOETHYL Aspartamide A column (PolyLC) on anAgilent HP1100 binary HPLC system. Peptides were eluted with an increasingKCl gradient [10-40 min at 0–30% buffer B then 40–60 min at 30–100%buffer B; buffer B: 10 mM KH2PO4 (pH 2.6) and 350 mM KCl in 25% aceto-nitrile]. The eluate was fractionated into four fractions and desalted withSep-Pak reverse-phase cartridges (Waters).
IMAC. Chelating Sepharose Fast Flow beads (GEHealthcare) were charged fourtimes with 0.1 M FeCl3 freshly prepared solution and washed four times withwashingbuffer (74:25:1water:acetonitrile:acetic acid). Desalted peptideswereacidified with 0.1% TFA in 25% acetonitrile, applied to 40 μL of 25% beadslurry, and incubated for 30 min at room temperature. Samples were washedfive times with washing buffer and once with water. Phosphopeptides wereelutedby adding30 μL of 100mMsodiumphosphate buffer (pH8.9). The pHofall samples was adjusted to 3 by adding drops of 10% TFA followed bydesalting and concentrating samples with ZipTips (μC18; Millipore).
TiO2 Affinity Chromatography. Phosphopeptides were enriched using TiO2
affinity chromatography as described by Bodenmiller et al. (38) with minormodifications. Peptides were desalted and dissolved in phthalic acid solution(80% acetonitrile, 2.5% TFA, and 0.13 M phthalic acid). The peptide mixturewas incubated with 0.3 mg of TiO2 (GL Science) for 30 min in closed Mobicolspin columns. After different washing steps (SI Appendix, SI Materials andMethods), peptides were eluted with 0.3 M NH4OH and dried in a speed vac.Before MS analysis, samples were desalted with ZipTips (μC18; Millipore).
Analysis by Liquid Chromatography/Electrospray Ionization/Tandem MS andInterpretation of MS Data. Phosphopeptide analysis was performed with anLTQ-Orbitrap as described previously (10) (SI Appendix, SI Materials andMethods). Up to five data-dependent tandemMS spectra were acquired in thelinear ion trap for each Fourier transform MS spectral acquisition range, thelatter acquired at 60,000 FWHM nominal resolution settings with an overallcycle time of ∼1 s. The samples were acquired by using internal lock mass cali-bration onm/z 429.088735 and 445.120025. TandemMS spectra were searchedwith Mascot (Matrix Science) version 2.2.04 against The Arabidopsis In-formation Resource (TAIR9) protein database (downloaded on June 29, 2009)with concatenated decoy database supplemented with contaminants (67,079entries) as described previously (10) (SI Appendix, SI Materials and Methods)and integrated into the pep2pro database (39). Identifications with a MASCOTion score >30 and a MASCOT expected value of <0.015 were accepted. Phos-phorylation site assignment was based on normalized delta ion score (ΔI) thatwas calculated for phosphopeptides for which the only difference between therank 1 and the rank 2 hit was the phosphorylation position. Phosphorylationsite assignmentswithΔI≥ 0.4were accepted (10, 40). From thefinal data, PRIDE2.1 XML files were created and exported to the PRIDE database (20) (accessionnos. 13754–13761). Data are also available in the pep2pro database (www.pep2pro.ethz.ch) (39). Relative quantification by extracted ion chromatogramswas achieved by commercial Progenesis software from Nonlinear Dynamics.Analysis was performed in pairs of liquid chromatography/MS runs for WT andthe corresponding stn8 experiment, according to the manufacturer’s instruc-tions. The quantitative analysis was done in three biological replicates.
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Spectroscopic Analysis. Spectroscopic changes were followed on intact leaveswith a flash spectrophotometer (JTS 10; Biologic). P700 oxidation kinetics wasassessed at 820–870 nm, and fluorescence was probed in the near infraredupon excitation with blue light. The ECS was measured at 520–545 nm toavoid interference by redox-related signals (SI Appendix, SI Materials andMethods). Actinic light was provided by a red LED peaking at 620 nm, whichwas transiently switched off to allow for the measurements of the P700 andECS changes (SI Appendix, SI Materials and Methods).
ACKNOWLEDGMENTS. We are grateful for support from the FunctionalGenomics Center Zurich, especially for the advice of Drs. Bernd Roschitzkiand Peter Gehrig concerning the interpretation of MS data. This work wassupported by funds from ETH Zurich and Swiss SystemsX.ch Project PlantGrowth Regulation (to W.G.), by CNRS funds (to G.F.), and by Swiss NationalFoundation Grant 31003A_133089 (to J.-D.R.). S.R. was supported by MarieCurie Early Stage Training Fellowship ADONIS MEST-CT-2005-020232 (to W.G. and S.B.). A.E. was supported by a Zurich-Basel Plant Science Center Grad-uate Research Fellowship (to S.B.).
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