suppression of chloroplastic alkenal/one oxidoreductase ......bases (esterbauer et al., 1991; miyata...

Suppression of Chloroplastic Alkenal/OneOxidoreductase Represses the Carbon CatabolicPathway in Arabidopsis Leaves during Night1[OPEN]

Daisuke Takagi2, Kentaro Ifuku2, Ken-ichi Ikeda, Kanako Ikeda Inoue, Pyoyun Park, Masahiro Tamoi,Hironori Inoue, Katsuhiko Sakamoto, Ryota Saito, and Chikahiro Miyake*

Department of Biological and Environmental Science, Faculty of Agriculture, Graduate School of AgriculturalScience (D.T., K.-i.I., K.I.I., P.P., H.I., K.S., R.S., C.M.), and Center for Support to Research and EducationActivities (P.P.), Kobe University, Nada, Kobe 657–8501, Japan; Division of Integrated Life Science, GraduateSchool of Biostudies, Kyoto University, Sakyo-ku, Kyoto 606–8502, Japan (K.I.); and Faculty of Agriculture,Kinki University, Nakamachi, Nara 631–8505, Japan (M.T.)

ORCID IDs: 0000-0003-0880-5877 (D.T.); 0000-0002-2426-2377 (C.M.).

Lipid-derived reactive carbonyl species (RCS) possess electrophilic moieties and cause oxidative stress by reacting with cellularcomponents. Arabidopsis (Arabidopsis thaliana) has a chloroplast-localized alkenal/one oxidoreductase (AtAOR) for thedetoxification of lipid-derived RCS, especially a,b-unsaturated carbonyls. In this study, we aimed to evaluate thephysiological importance of AtAOR and analyzed AtAOR (aor) mutants, including a transfer DNA knockout, aor (T-DNA),and RNA interference knockdown, aor (RNAi), lines. We found that both aor mutants showed smaller plant sizes than wild-typeplants when they were grown under day/night cycle conditions. To elucidate the cause of the aor mutant phenotype, weanalyzed the photosynthetic rate and the respiration rate by gas-exchange analysis. Subsequently, we found that both wild-type and aor (RNAi) plants showed similar CO2 assimilation rates; however, the respiration rate was lower in aor (RNAi) than inwild-type plants. Furthermore, we revealed that phosphoenolpyruvate carboxylase activity decreased and starch degradationduring the night was suppressed in aor (RNAi). In contrast, the phenotype of aor (RNAi) was rescued when aor (RNAi) plantswere grown under constant light conditions. These results indicate that the smaller plant sizes observed in aor mutants grownunder day/night cycle conditions were attributable to the decrease in carbon utilization during the night. Here, we propose thatthe detoxification of lipid-derived RCS by AtAOR in chloroplasts contributes to the protection of dark respiration and supportsplant growth during the night.

Among biomolecules, lipids, especially polyunsatu-rated fatty acids (PUFAs), are easily oxidized by reac-tive oxygen species (ROS). When PUFAs react withROS, sequential lipid peroxidation reactions start tooccur. An allylic/bis-allylic hydrogen atom in PUFAs ishighly reactive against ROS, especially the hydroxylradical (Møller et al., 2007; Poon, 2009). The allylichydrogen atom in PUFAs is extracted by these radicalspecies, and then lipid radical is formed (Vistoli et al.,

2013). Subsequently, the lipid radical reacts with molec-ular oxygen and forms lipid peroxyl radical and lipidalkoxyl radical (Vistoli et al., 2013). Lipid peroxyl radicaland lipid alkoxyl radical cause radical chain oxidationto lipid peroxidation by reacting with neighboring lipidmolecules, as a result of which lipid peroxide andlipid radical accumulate (Catalá, 2010). This sequentiallipid peroxidation cycle disturbs membrane structuresand their fluidity; thus, the physiological functions ofmembranes are inactivated (Pamplona, 2011). Further-more, lipid peroxide produces lipid-derived reactivecarbonyl species (RCS) such as a,b-unsaturated car-bonyls, dicarbonyls, and keto-aldehydes as intermedi-ate products through their breakdown (Vistoli et al.,2013). The electrophilic moieties in lipid-derived RCSare capable of reacting with DNA and proteins thatcontain nucleophilic amino acids (such as Cys, His, Lys,andArg) and form advanced lipoxidation end productsthrough the formation of Michael adducts and Schiffbases (Esterbauer et al., 1991; Miyata et al., 2000; Aldiniet al., 2007). The formation of advanced lipoxidationend products from DNA and proteins is accompaniedby their structural changes or cross-linkages, and DNAand proteins lose their intrinsic physiological functions(Pamplona, 2011).

1 This work was supported by the Japan Society for the Promotionof Science (grant no. 21570041 to C.M.).

2 These authors contributed equally to the article.* Address correspondence to [email protected] author responsible for distribution of materials integral to the

findings presented in this article in accordance with the policy de-scribed in the Instructions for Authors (www.plantphysiol.org) is:Chikahiro Miyake ([email protected]).

D.T., K.I., and C.M. designed the experiments; D.T. performedmost of the work; K.I. generated RNAi mutants (aor-1 and aor-4);K.-i.I., K.I.I., and P.P. performed transmission electron microscopyanalysis; M.T., H.I., K.S., and R.S. supported the experiments andthe interpretation of data; D.T. and C.M. wrote and completed thearticle with the valuable suggestion of all authors.

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Chloroplasts in higher plants are PUFA-rich organ-elles. Monogalactosyldiacylglycerol and digalacto-syldiacylglycerol are major constituents of chloroplasts,and fatty acid moieties, which constitute theselipid molecules, are highly unsaturated (monogalacto-syldiacylglycerol, 18:3 approximately 60%, 16:3 ap-proximately 30%; digalactosyldiacylglycerol, 18:3approximately 70%, 16:3 approximately 2%; Douceet al., 1973; Kelly et al., 2003; Block et al., 2007). Chlo-roplasts are major ROS-producing organelles, andsuperoxide (O2

2), hydroxyl radical, and singlet oxygen(1O2) are inevitably produced by photosynthetic elec-tron transport reactions (Asada and Takahashi, 1987).These facts suggest that chloroplasts are exposed to ahigh risk of lipid peroxidation. Indeed, Mano et al.(2014a) reported that the Arabidopsis (Arabidopsisthaliana) fad7fad8 double mutant contained less lipid-derived RCS (acrolein, crotonaldehyde, and malon-dialdehyde) in its leaves compared with wild-typeleaves. Both Arabidopsis FAD7 and FAD8 encodechloroplastic v-3 fatty acid desaturases that convert16:2 and 18:2 fatty acids into 16:3 and 18:3 fatty acids(Iba et al., 1993; McConn et al., 1994). Hence, the fad7-fad8 double mutant suppressed the production of 16:3and 18:3 fatty acids, and these lipid molecules thatconstitute chloroplasts are less unsaturated (McConnet al., 1994). These reports indicate that chloroplasts arean important source of lipid-derived RCS in plants.Lipid-derived RCS modify various enzymes in plant

cells and inhibit their catalytic activities. For example,the addition of acrolein, crotonaldehyde, or 4-hydroxyl-2-nonenal to isolated chloroplasts inhibits the activitiesof the Calvin cycle enzymes, especially thiol-regulatedphosphoribulokinase (PRK) and glyceraldehyde-3-phosphate dehydrogenase (GAPDH; Mano et al.,2009). The inhibition of enzyme activities reduces theCO2 assimilation rate in chloroplasts (Mano et al., 2009).In addition, Rubisco, the 33-kD oxygen-evolving com-plex, and the light-harvesting complex are modified bymalondialdehyde under heat stress conditions in vivo(Yamauchi et al., 2008; Yamauchi and Sugimoto, 2010).Furthermore, recent studies have revealed that lipid-derived RCS modify not only chloroplastic enzymesbut also cytosolic and mitochondrial enzymes in plantleaves (Fujita and Hossain, 2003; Hoque et al., 2012;Mano et al., 2014b). These facts indicate that the pro-duction and the accumulation of lipid-derived RCSshould be dangerous for plants. On the other hand,recent studies have also shown that lipid-derived RCSplay an important role in the signaling process andmodify the gene expression of several important cellu-lar processes, including detoxification, heat stress, celldivision, auxin signaling, and programmed cell death(Farmer and Mueller, 2013; Biswas and Mano, 2015).Indeed, Yamauchi et al. (2015) reported that exposureto lipid-derived RCS modifies the expression of tran-scriptional factors in plant cells, and the modification ofgene expression networks provides tolerance to theheat stress. On the basis of these reports, although ex-cess amounts of lipid-derived RCS would be toxic for

plant cells, suitable amounts of lipid-derived RCS couldbe beneficial for acclimation to environmental stressconditions. Therefore, the production of lipid-derivedRCS should be strictly regulated in plants cells.

To avoid the risk of accumulating excess lipid-derived RCS in plant cells and the consequentoxidative modification of biomolecules, plants havedetoxification enzymes like aldo-keto reductase (AKR),aldehyde dehydrogenase (ALDH), aldose/aldehydereductase (ALR), alkenal reductase (AER), andalkenal/one reductase (AOR; Oberschall et al., 2000;Mano et al., 2002; Kirch et al., 2004; Yamauchi et al.,2011; Saito et al., 2013). Both AKR and ALR reduce al-dehyde groups to alcohol groups using NAD(P)H, andALDH oxidizes aldehyde groups to carboxylic acidgroups using NAD(P)+ (Oberschall et al., 2000; Kirchet al., 2004; Saito et al., 2013). Both AER and AOR de-toxify a,b-unsaturated carbonyls through the reductionof a highly electrophilic a,b-unsaturated bond usingNAD(P)H (Mano et al., 2002; Yamauchi et al., 2011).Overexpression of these detoxification enzymes inplants reduces the content of lipid-derived RCS in cellscompared with the wild type. Moreover, these over-expressing plants acquire tolerance to oxidative dam-age under drought and salt stress conditions, whereROS production in chloroplasts is stimulated (Sunkaret al., 2003; Mano et al., 2005; Rodrigues et al., 2006;Turóczy et al., 2011). On the other hand, transgenicplants that have suppressed activities of these detoxi-fication enzymes show higher sensitivity to oxidativestress than wild-type plants (Kotchoni et al., 2006; Shinet al., 2009; Stiti et al., 2011; Yamauchi et al., 2012).These reports indicate that the accumulation of lipid-derived RCS indeed stimulates oxidative stress inplants. However, in spite of these studies on lipid-derived RCS detoxification enzymes in plants, themechanism of how lipid-derived RCS affect plantphysiological reactions such as photosynthesis or res-piration is less clear in vivo. As a result, the physio-logical importance of the detoxification of lipid-derivedRCS in plants is still elusive.

In this study, we aimed to elucidate the physiolog-ical importance of the detoxification of lipid-derivedRCS in chloroplasts that is the source of ROS and lipid-derived RCS in plants. First, we grew the Arabidopsistransfer DNA-tagged knockout mutant aor (T-DNA)and analyzed plant growth. We found that aor(T-DNA) showed growth retardation, compared withthe wild type, under day/night cycle conditions. Thisresult conflicted with previous results (Yamauchiet al., 2012). To confirm whether AOR is related togrowth in Arabidopsis, we constructed RNA interfer-ence mutants of AOR in Arabidopsis, aor (RNAi), andchecked their growth. We observed that aor (RNAi)plants also showed growth retardation, comparedwith the wild type, under day/night cycle conditions.Here, we discuss the physiological importance of lipid-derived RCS detoxification through the investigationof the cause of the growth retardation observed in aormutants.

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RESULTS

aorMutants Show Growth Retardation Compared with theWild Type under Day/Night Cycle Conditions

To study the importance of the detoxification of lipid-derived RCS in higher plants, we grew the transfer DNA-tagged knockout mutant of AtAOR, aor (T-DNA)(Yamauchi et al., 2012), under day/night cycle condi-tions (16 h of light and 8 h of dark) and analyzed thephenotype. First, we checked the mRNA level of AtAORin aor (T-DNA) by reverse transcription (RT)-PCR. Theexpression of AtAOR was successfully inhibited in aor(T-DNA) (Fig. 1A). Subsequently, we evaluated theacrolein-reducing activity in a crude leaf extract of aor(T-DNA). The activity was decreased to about 58% in aor(T-DNA) (2.3 6 0.2 mmol NADPH mg21 protein h21

[n = 3]) compared with the wild type (4 6 0.4 mmolNADPHmg21 protein h21 [n = 3]). Because higher plantshave a cytosolic acrolein detoxification enzyme, aor(T-DNA) would show the residual activity, as has beenmentioned in previous reports (Mano et al., 2005; Papdiet al., 2008; Yamauchi et al., 2012).

Under day/night cycle conditions, aor (T-DNA)showed smaller growth compared with the wild type(Fig. 1B). The growth rate, as evaluated by maximumrosette diameter, was slower in aor (T-DNA) than in thewild type (Fig. 1C). Furthermore, the dry weight of aor(T-DNA) 3 weeks after germination was significantlylower than that of the wild type (Fig. 1D). These results

conflicted with the previous report that the growth ofaor (T-DNA) was the same as that of the wild type(Yamauchi et al., 2012).

To confirm whether the growth retardation in aor(T-DNA) observed in our analysis depends on themalfunction of AtAOR, we generated RNA interference(RNAi) mutants in Arabidopsis, which specificallysuppressed themRNA level ofAtAOR. We isolated twoRNAi lines, aor-1 and aor-4. The mRNA level of AtAORdecreased to approximately 2% and 50% in aor-1 andaor-4, respectively, as comparedwith the wild type (Fig.2A). Subsequently, the accumulation of AtAOR proteinin aor-1 and aor-4 leaves was compared with that of thewild type by western-blot analysis. AtAOR protein inthe wild-type leaves was detected at a molecular massof approximately 38 kD (Fig. 2B). According to ExPASy(http://www.expasy.org/), the molecular mass of full-length AtAOR protein was estimated as 41 kD. How-ever, AtAOR protein has been reported to possess atransit peptide sequence to chloroplasts (Yamauchiet al., 2011). Thus, it was speculated that the true mo-lecular mass of AtAOR protein is smaller than its full-length protein. To clarify whether AtAOR protein islocalized in the chloroplasts of Arabidopsis cells, weexpressed a translational fusion protein betweenAtAOR protein and GFP in protoplasts isolated fromArabidopsis. Green fluorescence from GFP was ob-served in the protoplasts, and the location of emittinggreen fluorescence overlapped with red fluorescence

Figure 1. Phenotype analysis of aor (T-DNA) grown under day/nightcycle growth conditions. A, Confirmation of the expression ofAtAOR inwild-type (WT) and aor (T-DNA) leaves by RT-PCR. As a positive con-trol, the expression of 40S RIBOSOMAL PROTEIN S15A (Rps15aA) isshown. B, Phenotypes of wild-type and aor (T-DNA) plants. These plantsare 3 weeks old after germination. Representative plants are shown.Bar = 1 cm. C, Plant growth evaluated as an increase in maximum ro-sette diameter. Data are expressed as means6 SE (n = 4). Black squaresindicate the wild type, and red circles indicate aor (T-DNA). D, Dryweight of plants compared between the wild type and aor (T-DNA). Theblack bar indicates the wild type, and the red bar indicates aor (T-DNA).Data are expressed as means6 SE (n = 4). Asterisks indicate a significantdifference between the wild type and aor (T-DNA) (Student’s t test: **,P , 0.01).

Figure 2. Expression analysis of AtAOR and the detection of AtAORprotein in wild-type (WT) and aor (RNAi) leaves. A, Relative expressionlevels of AtAOR mRNA in wild-type and aor (RNAi) leaves. Expressionlevel was quantified by real-time PCR. Data are expressed asmeans6 SE

(n = 3). B, AtAOR protein extracted from wild-type and aor (RNAi)leaves was detected by western-blot analysis. M indicates a proteinweight molecular marker. Protein corresponding to 5 mg was loaded ineach lane. C, Protein profile by SDS-PAGE that correlates with thewestern-blot analysis shown in B. M indicates a protein weight mo-lecular marker. Protein corresponding to 5 mg extracted from wild-typeand aor (RNAi) leaves was loaded in each lane.

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emitted from chloroplasts (Supplemental Fig. S1).These results indicated that AtAOR protein was local-ized in chloroplasts. From these results, we concludedthat AtAOR protein exists in the chloroplast as the38-kD protein. In aor-1, no AtAOR protein was detected,whereas a small amount of AtAOR protein was detectedin aor-4 (Fig. 2, B and C). Next, we evaluated the acrolein-reducing activity in crude leaf extracts of aor (RNAi)mutants. The activity decreased to about 60% and 75% inaor-1 and aor-4, respectively, compared with the wildtype (Table I). Like aor (T-DNA), aor-1 showed residualacrolein-reducing activity even though AtAOR proteinwas not detected in western-blot analysis.We evaluated the growth rates in aor-1 and aor-4

under day/night cycle conditions. Like aor (T-DNA),aor-1 and aor-4 showed growth retardation, and dryweights of aor-1 and aor-4 at 3 weeks after germinationdecreased to about 49% and 54%, respectively, com-pared with the wild type (Fig. 3, A and B). From thesefacts, we concluded that AtAOR protein function isrequired for optimal plant growth.

Suppression of AtAOR Decreases Dark Respiration Rate,But Not the Photosynthesis Rate, under Day/NightCycle Conditions

To elucidate the cause of growth retardation in aormutants, we analyzed the photosynthesis activity anddark respiration rate in aor-1 and aor-4 under growthconditions. First, we quantified the nitrogen and chloro-phyll contents in aor-1 and aor-4. Comparedwith the wildtype, the chlorophyll content in aor-1 and aor-4 mutantsdecreased by approximately 14% and 20%, respectively,and the nitrogen content also decreased by approximately18% in both aormutants (Table II). These decreases in thechlorophyll and nitrogen contents also were observed inaor (T-DNA) (Supplemental Table S1).Next,we evaluatedthe photosynthetic activity and thedark respiration rate inwild-type and aor (RNAi)mutant plants. CO2 assimilationrates in aor-1 and aor-4 showed similar values to the wildtype under atmospheric conditions, where actinic lightintensity was the same as growth light intensity (150 mEm22 s21; Table III). Furthermore, internal partialpressure of CO2 (Ci) and stomatal conductance (gs)also were similar among the wild type, aor-1, and aor-4(Table III). Simultaneously, we evaluated themaximum

quantum yield of PSII (Fv/Fm) and the incident quan-tum yield of PSII [Y(II)] under growth light conditions.The values of Fv/Fm and Y(II) were similar amongwild-type, aor-1, and aor-4 plants (Table III). These resultsindicated that aor-1 and aor-4 have similar photosyn-thetic activities compared with the wild type.

In contrast to the photosynthetic activity, the darkrespiration rates evaluated from CO2 gas-exchangeanalysis were lower in aor-1 and aor-4 compared withthe wild type (Table III). We also evaluated the darkrespiration rate using an oxygen electrode; aor-1 andaor-4 again showed lower respiration rates comparedwith the wild type (Supplemental Fig. S2). Further-more, we also evaluated the photosynthetic activitiesand the dark respiration rate in aor (T-DNA) grownunder day/night cycle conditions. Both Y(II) and gstended to be lower than in the wild type; however, CO2assimilation rate and Ci were the same between wild-type and aor (T-DNA) plants (Supplemental Table S2).In contrast, the dark respiration rate in aor (T-DNA)waslower compared with the wild type, but it was similarto both aor-1 and aor-4 (Supplemental Table S2). Theseresults suggested that the growth retardation in aormutants was caused by a decrease in the dark respira-tion rate.

Expression Analysis of Respiration-Related Genes byReal-Time PCR

In the previous section, we observed that the aormutants showed lower respiration rates in the dark butnot lower CO2 assimilation rates in the light. These

Table I. Reduction activity of acrolein in crude leaf extract

The acrolein reduction activities in crude protein extracts fromleaves in wild-type and aor (RNAi) mutant plants were determined. Inthis experiment, 3- to 4-week-old plants were used. Data are expressedas means 6 SE (n = 3). Values in parentheses indicate the relative ac-tivity for the wild type.

Line Activity

mmol mg21 protein h21

Wild type 4.5 6 0.6 (100)aor-1 2.7 6 0.1 (60)aor-4 3.0 6 0.3 (75)

Figure 3. Characterization of aor (RNAi) mutants grown under day/night cycle conditions. A, Phenotypes of wild-type (WT), aor-1, and aor-4plants. These plants were 2 weeks old after germination. Representativeplants are shown. Bar = 1 cm. B, Plant growth evaluated as an increase inmaximum rosette diameter. Data are expressed as means 6 SE (n = 6).Black squares indicate the wild type, red circles indicate aor-1, and bluetriangles indicate aor-4. C, Dry weight of plants compared between thewild type and aor (RNAi) mutants. The black bar indicates the wild type,the red bar indicates aor-1, and the blue bar indicates aor-4. Data areexpressed as means 6 SE (n = 16). Different letters indicate a significantdifference between the wild type and aor (RNAi) mutants (Tukey-Kramerhonestly significant difference test: P , 0.05).



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observations indicated that the utilization efficiency ofcarbon acquired during photosynthesis was lower inthe aormutants during the night. Thus, mRNA levels ofgenes related to respiratory metabolism enzymes, suchas those involved in starch degradation, glycolysis, theTCA cycle, and respiratory electron transport reactions,were analyzed. In the following experiments, we usedaor-1 as the aor mutant. For this analysis, mRNA wasextracted from the leaves of both wild-type and aor-1plants at three different times: at the end of the day (1 hbefore the night period started), at midnight (4 h afterthe night period started), and at the end of the night (7 hafter the night period started). As an internal standard,we used Rps15aA (Watanabe et al., 2014), and themRNA level in the wild type at the end of the day wasset to 1 (Fig. 4). First, we analyzed the mRNA levelsinvolved in glycolysis. The mRNA level of FBA1(encoding FRUCTOSE-1,6-BISPHOSPHATE ALDOL-ASE1) decreased from the end of the day to midnightand subsequently increased until the end of the night inboth the wild type and aor-1 (Fig. 4A). For the mRNAlevel of PK (encoding PYRUVATE KINASE), a signifi-cant difference between the wild type and aor-1was notobserved; however, the change in the amount of mRNAwas different, in that the mRNA level tended to de-crease in aor-1 at midnight, compared with the wildtype (Fig. 4B). The change in mRNA level of PDH (en-coding PYRUVATE DEHYDROGENASE E1 a-subunit)increased at midnight and subsequently decreased at theend of the night in the wild type (Fig. 4C). In contrast, themRNA level of PDH did not increase atmidnight in aor-1(Fig. 4C). The mRNA level of PEPC1 (encodingPHOSPHOENOLPYRUVATE CARBOXYLASE1) alsoincreased from the end of the day to midnight and

subsequently decreased to the end of the night in thewild type (Fig. 4D). However, the mRNA level ofPEPC1 in aor-1 decreased linearly from the end of theday to the end of the night (Fig. 4D).

Next, we analyzed the mRNA levels of TCA cycleenzymes and respiratory electron transport chaincomponents. The mRNA levels of ACO3 (encodingACONITASE3) and CSY4 (encoding mitochondrion-targeted CITRATE SYNTHASE4) showed similarchanges to that of PEPC1 from the end of the day to theend of the night, and the mRNA level of ACO3 wassignificantly lower in aor-1 at midnight, as comparedwith the wild type (Fig. 4, E and F). In contrast, themRNA levels of CI76 (encoding COMPLEX I 76-kDsubunit), COX6a (encoding CYTOCHROME C OXI-DASE SUBUNIT 6A) and AOX1a (encoding ALTER-NATIVE OXIDASE1a) showed similar changes in thewild type and aor-1 (Fig. 4, G–I).

In addition, we analyzed the mRNA levels of en-zymes involved in starch degradation: GWD1 (encod-ing GLUCAN WATER DIKINASE1), SEX4 (encodingPHOSPHOGLUCAN PHOSPHATASE), BAM1 (encod-ingb-AMYLASE1), andBAM3 (encodingb-AMYLASE3).In these starch degradation-related genes, no differencesin mRNA levels were observed between the wild typeand aor-1 (Fig. 4, J–M). The expression patterns of thesegenes during the night period were consistent with pre-vious reports (Smith et al., 2004; Santelia et al., 2011). Toconfirm that the mRNA levels of each gene were inde-pendent from the expression levels of an internal stan-dard, we also analyzed ACT2 (encoding ACTIN2)expression each time. In both the wild type and aor-1,ACT2 hardly showed any change in mRNA level duringthe night, and no difference was observed in ACT2 ex-pression between the wild type and aor-1 (Fig. 4N).Therefore, these results indicated that themRNA levels ofeach gene normalized by Rps15Aa showed uniquechanges during the night. These observations indicatedthat the functions of glycolysis and the TCA cycle duringthe night period were modified in aor-1.

Enzyme Activities of PDH, PEPC, ACO, and CSY in Wild-Type and aor-1 Leaves

Based on the gene expression analysis related torespiration, we evaluated the enzyme activities of PDH,PEPC, ACO, and CSY in wild-type and aor-1 leaves,which were sampled during the night period. PDHactivities in the wild type and aor-1 were similar (Fig.

Table II. Chlorophyll and nitrogen contents in wild-type and aor(RNAi) mutant leaves grown under day/night cycle growth conditions

Wild-type and aor (RNAi) mutant plants were grown under day/nightcycle growth conditions. Chlorophyll and nitrogen contents were de-termined on leaves in to 4-week-old plants. Data are expressed asmeans 6 SE (n = 6). Asterisks indicate significant differences betweenthe wild type and aor (RNAi) mutants (Student’s t test: *, P , 0.05 and**, P , 0.01).

Line Chlorophyll Nitrogen

mmol m22 mmol m22

Wild type 219.0 6 7.4 75.6 6 3.0aor-1 189.9 6 6.4** 63.0 6 4.1*aor-4 177.3 6 6.6** 62.4 6 4.2*

Table III. Dark respiration rate and photosynthetic parameters in wild-type and aor (RNAi) plants grown under day/night cycle conditions

Photosynthesis activities were determined under growth light conditions (light intensity of 150 mE m22 s21), and respiration activity was determinedin the dark. Data are expressed as means 6 SE (n = 3). Dagger indicate significant differences between the wild type and aor (RNAi) mutants(Student’s t test: †, P , 0.1).

Line Fv/Fm Respiration CO2 Assimilation Y(II) gs Ci

mmol CO2 m22 s21 mmol CO2 m

22 s21 mmol mol21

Wild type 0.827 6 0.002 1.41 6 0.34 4.72 6 0.20 0.631 6 0.004 0.076 6 0.008 386.1 6 12.4aor-1 0.821 6 0.003 0.54 6 0.14† 4.87 6 0.22 0.625 6 0.016 0.070 6 0.014 374.1 6 9.3aor-4 0.817 6 0.003 0.51 6 0.15† 4.49 6 0.10 0.627 6 0.025 0.086 6 0.003 384.9 6 4.4


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5A). In contrast, PEPC activity in aor-1 decreased signif-icantly to about 75% compared with the wild type (Fig.5B). The activities of ACO and CSY also were similarbetween the wild type and aor-1 (Fig. 5, C and D).

Starch Degradation in Wild-Type and aor-1 Leaves duringthe Night Period

During the night period, plants degrade starchthrough respiratory metabolism (Zeeman et al., 2010).Accordingly, we quantified the amount of starch inchloroplasts in the wild type and aor-1. For this mea-surement, leaves from wild-type and aor-1 plants wereharvested at the end of the day and night. The starch inchloroplasts was observed by transmission electronmicroscopy, and the amount of starch in chloroplastswas quantified by a point-counting method (Weibel,1979). In wild-type leaves, starch accumulated in chlo-roplasts at the end of the day was hardly detected at theend of the night (Fig. 6A). This result showed that about98% of the starch accumulated during the day wasconsumed during the night in chloroplasts of the wild

type (Fig. 6B). In contrast, starch remained in chloro-plasts of aor-1 at the end of the night, although thestarch content decreased significantly at the end of thenight, compared with the end of the day (Fig. 6). Thisresult showed that about 62% of the starch accumulatedin the day was consumed in aor-1 chloroplasts duringthe night period. Compared with the wild type, starchcontent in aor-1 was significantly higher at the end ofboth day and night (Fig. 6B). These results indicatedthat starch degradation was suppressed in aor-1 duringthe night, resulting in the accumulation of starch inchloroplasts. We also quantified the amount of starch inwild-type and aor-1 leaves biochemically, as describedby Sawicki et al. (2012). In the wild type, starch accu-mulated during the day was degraded significantlyduring the night (Supplemental Fig. S3). In contrast,starch degradation was suppressed in aor-1 during thenight, and the starch content was not decreased sig-nificantly at the end the night, compared with the endof the day (Supplemental Fig. S3). This result supportsthe idea that starch degradation is suppressed in aor-1during the night.

Figure 4. Gene expression analysis related torespiration and starch degradation in leaves.mRNAwas extracted from wild-type (WT) andaor-1 leaves at end of day (1 h before the nightperiod started), midnight (4 h after the nightperiod started), and end of night (7 h after thenight period started). In each gene, the mRNAlevel in the wild type at the end of day was setto 1, and relative expression change is shown.Black squares indicate the wild type, and redcircles indicate aor-1. Data are expressed asmeans 6 SE (n = 3–6). Asterisks indicate sig-nificant differences between the wild type andaor-1 (Student’s t test: *, P , 0.05).



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Growth of Wild-Type and aor-1 Plants under ConstantLight Conditions

We grew both wild-type and aor-1 plants underconstant light conditions and analyzed their growth.We found that wild-type and aor-1 plants grew simi-larly (Fig. 7A). Dry weight at 3 weeks after germinationalso was similar between the wild type and aor-1 undercontinuous light conditions (Fig. 7B). Furthermore, thenitrogen and chlorophyll contents also were at similarvalues for the wild type and the aor-1mutant (Table IV).These results indicate that constant light conditionsrescued the growth in aor-1.

Next, we analyzed the photosynthetic activities inwild-type and aor-1 plants grown under constant lightgrowth conditions. CO2 assimilation rate and Y(II) werenot different between the wild type and aor-1 (Table V).The values of Fv/Fm, Ci, and gs also were not differentbetween the wild type and aor-1 (Table V). These resultsshowed that wild-type and aor-1 plants have similarphotosynthetic activities, similar to the day/night cyclegrowth conditions. In contrast, the dark respiration ratein aor-1 was lower compared with the wild type (TableV). We also checked the phenotype of aor (T-DNA)grown under continuous light conditions. Like aor-1,

aor (T-DNA) grew similar to thewild type (SupplementalFig. S4). Chlorophyll and nitrogen contents were similarbetween thewild type and aor (T-DNA); furthermore, thephotosynthetic activities estimated by CO2 assimilationrate and Y(II) were not different between the wild typeand aor (T-DNA) (Supplemental Tables S3 and S4).However, the dark respiration rate was lower in aor(T-DNA) than in the wild type, as well as in aor-1(Supplemental Table S4).

Figure 5. Evaluation of enzyme activities of PDH (A), PEPC (B), ACO(C), and CSY (D) in wild-type (WT) and aor-1 leaves. Leaves were har-vested from wild-type and aor-1 plants grown under day/night cyclegrowth conditions, and these enzyme activities were evaluated inleaves harvested at midnight. Black bars indicate the wild type, and redbars indicate aor-1. Data are expressed as means 6 SE (n = 4). The as-terisk indicates a significant difference between the wild type andaor-1 (Student’s t test: *, P , 0.05).

Figure 6. Evaluation of starch metabolism in wild-type (WT) andaor-1 leaves during the night. A, Ultrastructural observation ofchloroplasts in wild-type and aor-1 leaves. Leaveswere harvested at endof day (1 h before the night period started) and end of night (7 h after thenight period started) for this analysis. Representative images are shown.Bars = 2 mm. B, Starch occupancy ratio in chloroplasts in wild-type andaor-1 leaves evaluated by a point-counting method (see “Materials andMethods”). Data are expressed as means 6 SE. More than 16 chloro-plasts were observed in each treatment. Different letters indicate sig-nificant differences between the wild type and aor-1 (Tukey-Kramerhonestly significant difference test: P , 0.05).


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Next, we analyzed the mRNA levels of PDH, PEPC1,ACO3, and CSY4 in both wild-type and aor-1 plantsgrown under constant light growth conditions. ThemRNA levels of these genes were not different betweenthe wild type and aor-1 (Supplemental Fig. S5).We also evaluated the enzyme activities of PDH,

PEPC, ACO, and CSY in both wild-type and aor-1plants grown under constant light conditions. Wefound that PEPC activity in aor-1 decreased comparedwith the wild type (Supplemental Fig. S6B). In contrast,the activities of PDH, ACO, and CSY were similar be-tween the wild type and aor-1 (Supplemental Fig. S6, A,C, and D). These results indicated that PEPC activitywas lower in aor-1, although the growth in aor-1 wassimilar to that of the wild type.Finally, the amount of starch in chloroplasts was

observed in both wild-type and aor-1 plants grownunder constant light conditions (Supplemental Fig. S7).Although the amount of starch tended to be higher inaor-1 than in the wild type, no significant difference wasobserved between the wild type and aor-1.

DISCUSSION

In this study, we analyzed the physiological impor-tance of the detoxification of lipid-derived RCS inchloroplasts using Arabidopsis mutants that have alower amount ofAtAOR [aor (RNAi); aor-1 and aor-4] ordo not express a functional AtAOR [aor (T-DNA)]. Werevealed that the suppression of AtAOR functioninhibited carbon metabolism initiated by starch

degradation during the night, leading to a lowering ofdark respiration activity and suppressed plant growth(Figs. 3 and 6; Table III; Supplemental Fig. S3). These re-sults conflict with a previous report inwhich aor (T-DNA)was analyzed (Yamauchi et al., 2012).However,we foundthat the same aor (T-DNA) also showed smaller growthsize than the wild type when they were grown underday/night cycle conditions (Fig. 1). These results corre-sponded to those of aor (RNAi). Therefore, our differentresults for the growth of aor (T-DNA)would be due to thedifference in the growth conditions, including nutrition,growingmedium, light intensity, or growth stage. In fact,the sizes of the aor (T-DNA) plants reported by Yamauchiet al. (2012) were smaller than those reported here (Fig. 1).

We revealed that the growth retardation in aor mu-tants under day/night cycle conditions was not causedby carbon acquisition capacity during the day. Underthe growth light intensity, CO2 assimilation and Y(II) inaormutants were similar to wild-type values (Table III;Supplemental Table S2). Based on these results, thegrowth retardation in aor mutants was not accountedfor by their photsynthetic ability. However, these re-sults did not mean that the suppression of AtAORhardly affects photosynthetic ability. In aor-1, the Calvincycle enzymes FBPase and PRKhad significantly loweredactivities comparedwith thewild type (Supplemental Fig.S8). FBPase and PRK have thioredoxin-regulated Cysin their structures, and the reduction of the disulfidebridge between two Cys residues activates their activities(Martin et al., 2000). These enzymes are susceptible tooxidation by ROS; furthermore, these enzymes are mod-ified by lipid-derived RCS, which inhibit their functions(Asada and Takahashi, 1987; Tamoi et al., 1996a; Manoet al., 2009, 2014b). Based on these previous reports, thedecrease in FBPase and PRK activities in aor-1 could bedue to oxidative stress induced by the suppression ofAtAOR in chloroplasts. In fact, we observed that oxida-tive stress evaluted by the 3,3-diaminobenzidine stainingmethod was stimulated in aor-1, compared with the wildtype, under their growth conditions (Supplemental Fig.S9). FBPase and PRK activities were lower in aor-1; nev-ertheless, the CO2 assimilation rate and Y(II) were similarbetween wild-type and aor-1 plants under their growthlight conditions (Table III).

These results showed that the lack of AtAOR inchloroplasts induces oxidative stress, which affects the

Figure 7. Characterization of aor-1 plants grown under constant lightgrowth conditions. A, Phenotypes of wild-type (WT) and aor-1 plants.These plants are 3 weeks old after germination. Representative plantsare shown. B, Plant growth evaluated as an increase inmaximum rosettediameter. Black squares indicate the wild type, and red circles indicateaor-1. Data are expressed as means 6 SE (n = 9–10). C, Dry weight ofplants compared between the wild type and aor-1. The black bar in-dicates the wild type, and the red bar indicates aor-1. Data areexpressed as means 6 SE (n = 9–10).

Table IV. Chlorophyll and nitrogen contents in wild-type and aor(RNAi) mutant leaves grown under continuous light conditions

Wild-type and aor (RNAi) mutant plants were grown under constantlight growth conditions. Chlorophyll and nitrogen contents were de-termined on leaves in 3- to 4-week-old plants. Data are expressed asmeans 6 SE (n = 6). The dagger indicates a significant difference be-tween the wild type and aor-1 (Student’s t test: †, P , 0.1).

Line Chlorophyll Nitrogen

mg m22 mmol m22

Wild type 261.3 6 9.7 75.1 6 4.8aor-1 288.3 6 8.4† 83.5 6 6.5aor-4 290.2 6 20.0 78.8 6 3.5



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Calvin cycle enzymes, although it did not cause the sup-pression of photosynthesis under our growth conditions.This would be because photosynthesis was not limited bythe regeneration of ribulose 1,5-bisphosphate (RuBP) un-der our growth conditions (Farquhar et al., 1980). Underour growth conditions, the light intensity did not saturateagainst photosynthesis; that is, photosynthesis was lim-ited by the supply of photon energy to the photosystemsin the thylakoid membranes. Wild-type and aor-1 plantsshowed the same Fv/Fm values; therefore, there was nodifference in photosynthetic activity between wild-typeand aor-1 plants grown under the growth light condi-tions (Table III). On the other hand, under high-light andhigh-CO2 conditions, where photosynthesis is limited bythe regeneration of RuBP, we observed that aor-1 showedlower CO2 assimilation rate and lower Y(II) comparedwith the wild type; otherwise, Ci was similar between thewild type and aor-1 (Supplemental Fig. S10). In contrast,under high-light and ambient CO2 conditions, rangingfrom20 to 30 PaCi, where photosynthesis is limited by thecarboxylation reaction of RuBPbyRubisco,wild-type andaor-1 plants showed the same CO2 assimilation rate and Y(II) (Supplemental Fig. S10). Therefore, aor mutants mayshowamore severe phenotype under high-CO2 andhigh-light conditions.

Carbon utilization during the night period is sup-pressed in aor mutants. Although aor mutants grownunder the day/night cycle showed growth retardation,the growth retardation was alleviated under constantlight conditions (Figs. 1, 3, and 7; Supplemental Fig. S4).These results indicated that the growth retardationobserved in aor mutants grown under the day/nightcycle might be due to the dark respiration during thenight. In fact, aor mutants showed a decreased rate ofdark respiration comparedwith thewild type (Table III;Supplemental Table S2; Supplemental Fig. S2). In thedark, chlororespiration is active, as is mitochondrialrespiration (Nawrocki et al., 2015). Chlororespirationconsists of two reactions: (1) first, the reduction of plas-toquinone (PQ) by the NAD(P)H dehydrogenase-likecomplex (NDH) embedded in the thylakoid mem-branes; and (2) the oxidation of reduced plastoquinone(PQH2) by PLASTID TERMINAL OXIDASE (PTOX).The oxygen consumption by PTOX affects to the oxygenabsorption rate in leaves in the dark (Häusler et al., 2009).Thus, there is a possibility that chlororespiration activitywas lowered in the aor mutants. To determine whetherthis was the case, we conducted three experiments.

First, we analyzed initial (minimum) PSII fluores-cence in the dark-adapted state (Fo) and maximum PSII

fluorescence in the dark-adapted state (Fm) levels indark-adapted wild-type and aor-1 leaves. If oxygenconsumption by PTOX was suppressed in the aor-1mutant, PQH2 would accumulate in the dark as a resultof NDH activity and Fo would increase (Häusler et al.,2009). However, we found that the Fo levels were sim-ilar between wild-type and aor-1 plants (SupplementalFig. S11A). Second, we analyzed NDH-dependent PQreduction activity in the dark. After illumination (150mE m22 s21) for 10 min, we monitored the kinetics ofchlorophyll fluorescence, which represents NDH-dependent PQ reduction, in the dark (Hashimoto et al.,2003; Yamori et al., 2015). We observed similar kineticsin both wild-type and aor-1 plants (Supplemental Fig.S11B). These results indicate that chlororespiration ac-tivity, which consisted of PQ reduction by NDH andPQH2 oxidation by PTOX, was not impaired in aor-1.Furthermore, we analyzed the time course of change inthe PQ redox state (qL and Fs/Fm) after the start of il-lumination in dark-adapted wild-type and aor-1 leaves,because the function of chlororespiration could be em-phasized in dark/light transient conditions (Casanoet al., 2000; Joët et al., 2002; Kramer et al., 2004; Miyakeet al., 2009; Suorsa et al., 2012). After illuminationat growth light intensity, qL and Fs/Fm showed the samekinetics in bothwild-type and aor-1 plants (SupplementalFig. S11, C and D). On the basis of these results, wesuggest that aor-1 maintains chlororespiration activitysimilar to the wild type. Therefore, the decrease in oxy-gen consumption rate in the aormutants in the dark couldbe attributed to the suppression of mitochondrial respi-ratory reactions.

Photosynthesis assimilates CO2 to accumulate carbonas a starch in chloroplasts during the day period; in con-trast, respiration degrades starch to acquire energy tosupply carbon to the catabolic metabolism. Under day/night cycle growth conditions, relative growth rate showsa linear correlation with starch degradation rate duringthe night period, rather than starch synthesis rate duringthe day (Gibon et al., 2009). Furthermore, the starchdegradation rate during the night period also shows alinear correlation with the utilization efficiency of organicacid metabolism (Gibon et al., 2009). This means thatcarbonflow from starch in chloroplasts to glycolysis in thecytosol and the TCA cycle in mitochondria is importantfor plant growth. Indeed, mutant analysis showed thatthe suppression of starch degradation caused a lowerrespiration rate and severe growth retardation (Zeemanand Rees, 1999; Yu et al., 2001; Chia et al., 2004; Niittyläet al., 2004; Kötting et al., 2005; Fulton et al., 2008).

Table V. Dark respiration rate and photosynthetic parameters in wild-type and aor-1 plants grown under constant light conditions

Photosynthesis activities were determined under growth light conditions (light intensity of 150 mE m22 s21), and the respiration activity wasdetermined in the dark. Data are expressed as means 6 SE (n = 3).

Line Fv/Fm Respiration CO2 Assimilation Y(II) gs Ci

mmol CO2 m22 s21 mmol CO2 m

22 s21 mmol mol21

Wild type 0.822 6 0.003 0.46 6 0.18 4.69 6 0.44 0.685 6 0.016 0.097 6 0.008 378.8 6 9.1aor-1 0.818 6 0.003 0.25 6 0.07 4.40 6 0.19 0.682 6 0.020 0.092 6 0.008 369.7 6 7.4


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In this study, we found that aor mutants showed alower dark respiration rate (Table III; Supplemental Fig.S2), where PEPC expression level during the night pe-riod and its enzyme activity decreased significantly,compared with the wild type (Figs. 4 and 5). PEPCconverts phosphoenolpyruvate and HCO3

2 to oxalo-acetic acid (OAA) and inorganic phosphate (Pi) byb-carboxylation of phosphoenolpyruvate in the pres-ence ofMg2+, and this reaction contributes to the supplyof carbon skeletons to the TCA cycle and amino acidsynthesis by the production of OAA (Lepiniec et al.,1994; Izui et al., 2004). PEPC activity greatly affectscarbon and nitrogen metabolism. For example, anArabidopsis knockout mutant of PEPC showed growthretardation (Shi et al., 2015). Furthermore, the decreaseof PEPC activity in plants caused a decrease in starchdegradation during the night period (Häusler et al.,1999; Shi et al., 2015). In addition, nitrogen assimilationalso was suppressed and amino acid content producedfrom OAA (Asp and Asn) was lowered compared withthe wild type (Häusler et al., 1999; Shi et al., 2015). Incontrast, plants overexpressing PEPC showed highernitrogen content compared with wild-type plants aswell as the stimulation of starch degradation (Häusleret al., 1999; Agarie et al., 2002; Rademacher et al., 2002;Chen et al., 2004). Furthermore, the change in PEPCactivity is related to that in the dark respiration rate inleaves. The PEPC suppression mutant showed a lowerdark respiration rate, and plants overexpressing PEPCshowed a higher respiration rate, compared with thewild type (Häusler et al., 1999; Agarie et al., 2002;Rademacher et al., 2002). These changes accompaniedthe change of carbon supply from starch to respiratorymetabolism. The phenotype of the PEPC suppressionmutant reported in these previous studies is similar tothe phenotype of aor-1 observed in this study (Figs. 3and 6; Table III; Supplemental Fig. S3).Based on these observations, one of the reasons why

aor mutants showed a suppression of the dark respira-tion rate, a lower nitrogen content, and growth retar-dation would be the lower activity of PEPC (Table II;Supplemental Table S1; Reich et al., 1998; Agarie et al.,2002; Foyer et al., 2011). However, the decrease in PEPCactivity cannot be wholly responsible for the growthretardation observed in the aormutants. This is becauseShi et al. (2015) reported that a moderate decrease inPEPC activity did not result in growth inhibition com-pared with the wild type, although the growth wasevaluated at the early growth phase in that study. Here,we found that the decrease in the detoxification activityof lipid-derived RCS in chloroplasts affects the cytosolicenzyme and nucleus-encoded gene expression (Figs. 4and 5). That is, our results suggest that lipid-derivedRCS produced in chloroplasts spread widely through-out plant cells. Therefore, lipid-derived RCS also couldaffect various molecular targets, which were not ex-amined in this study in plant cells because of their re-activities, and aor mutants might cause the decrease ofcarbon utilization during the night and the growth re-tardation as a result of the modification of plant cell

components. Moreover, lipid-derived RCS have beenreported to be involved in signaling processes such asauxin signaling and cell division (Farmer and Mueller,2013; Biswas and Mano, 2015; Yamauchi et al., 2015).Based on these reports, lipid-derived RCS diffused inplant cells might modify the gene expression network,which is not analyzed in this study, and cause the de-crease of carbon utilization during the night and thegrowth retardation in aor mutants. Combining the re-sults of Mano et al. (2014a) and our study, we con-cluded that chloroplasts are a major production site oflipid-derived RCS and might be an initiator for lipid-derived RCS-dependent signaling cascades in plantcells. Accordingly, further intensive studies are re-quired to elucidate the molecular targets of lipid-derived RCS, especially the receptors of lipid-derivedRCS triggering the signal cascade in plant cells (Farmerand Mueller, 2013; Biswas and Mano, 2015).

PEPC is one of the targeted molecules that is oxida-tively modified by lipid-derived RCS in vivo. Therefore,the decrease in PEPC activity might not be limited to thesuppression of PEPC gene expression in the aor mutant.Mano et al. (2014b) reported that the production of lipid-derived RCS is stimulated in plants under severe saltstress conditions, and they showed that several candidateproteins are carbonylated by lipid-derived RCS and ox-idative stress. Among the candidate proteins, PEPC wasdetected as a target for protein carbonylation (Mano et al.,2014b). PEPC has nucleophilic amino acids as a prereq-uisite for its activity. For example, a His residue is locatedin the hydrophobic pocket of PEPC; this His residuestabilizes an intermediate of carboxylation products andextracts H+ from the carboxyl group in phosphoenol-pyruvate (Lepiniec et al., 1994; Izui et al., 2004). Fur-thermore, PEPC has a Lys residue in its active site for thecarboxylation reaction (Podesta et al., 1986; Jiao et al.,1990). Interestingly, this Lys residue reacts with an al-dehyde group in pyridoxal 59-phosphate, whereby PEPCactivity is reduced by the formation of a Schiff base be-tween Lys and the aldehyde group (Podesta et al., 1986;Jiao et al., 1990). In addition, plant PEPC isozymes con-serve seven Cys residues in their amino acid alignments,and their oxidation of thiol groups suppresses PEPC ac-tivity (Chardot and Wedding, 1992). Based on these re-ports, PEPC is structurally highly susceptible to oxidativemodification by reactive aldehyde groups. Therefore, wesuggest that oxidative modification of these nucleophilicamino acid residues in PEPC might have occurred, andthis might be one of the reasons for a decrease in PEPCactivity in aor-1, because the detoxification activity oflipid-derived RCS is suppressed and oxidative stress isstimulated in aor mutants (Supplemental Fig. S9).

Under constant light conditions, the change of carbonsupply from starch degradation to glycolysis and theTCA cyle does not affect plant growth (Izumi et al.,2013). Furthermore, the dark respiration rate is largelysuppressed (about 80%) under illumination comparedwith under darkness, because TCA cycle activity andthe supply of hexose molecules like Glc and Suc toglycolysis are inhibited under illumination (Brooks and



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Farquhar, 1985; Villar et al., 1995; Shapiro et al., 2004;Tcherkez et al., 2005, 2008; Yin et al., 2011). These re-ports indicate that the carbon flow from photosynthesisto glycolysis and the TCA cycle is severely restrictedunder illumination, compared with that occurring un-der darkness. In fact, we observed that the amount ofstarch in chloroplasts was similar between wild-typeand aor-1 plants grown under constant light condi-tions (Supplemental Fig. S7). Therefore, we suggest thatthe decrease of carbon utilization in the aor mutantswould be masked by lowering the carbon flow in gly-colysis under constant light conditions and the growthretardation in aor mutants would be alleviated (Fig. 7;Supplemental Fig. S4).

In natural conditions, but not in the Arctic and Ant-arctica, the sun rises and sets and plants experience nightlife. Furthermore, global warming is predicted to ac-company the increases in both earth temperature and thepartial pressure of CO2 in the atmosphere (Stocker et al.,2013). The elevated temperature increases the dark res-piration rate in plants (Atkin and Tjoelker, 2003; Noguchiet al., 2015). The increased partial pressure of CO2 in theatmosphere shifts the rate-limiting step of photosyntheticCO2 assimilation rate from Rubisco limiting to RuBP re-generation (Farquhar et al., 1980). From our findings, wesuggest that the function of AOR would be important inthe future environment, because the lack of AtAORsuppressed dark respiration and the regeneration effi-ciency of RuBP in theCalvin cycle. In addition, heat stressstimulates the production of lipid-derived RCS in higherplants (Yamauchi et al., 2008; Yamauchi and Sugimoto,2010). Therefore, future studies should be conducted toinvestigate the detoxification and signaling processes oflipid-derived RCS in order to better understand the re-sponses of plants to future environments.

MATERIALS AND METHODS

Plant Growth Conditions

Arabidopsis (Arabidopsis thaliana) wild-type (ecotype Columbia-0) and aormutant plants were grown for 3 to 4 weeks under day/night cycle growthconditions (16 h of light [23°C, 150 mE m22 s21] and 8 h of dark [20°C]) orconstant light conditions (24 h of light [23°C, 150 mEm22 s21]) in a plant growthchamber. Relative humidity was kept at 50% to 60%. Seeds were sown in soil(commercial peat-based compost) and kept at 4°C for 3 d to stimulate theirgermination, before seeds were transferred to the plant growth chamber. A0.1% (v/v) Hyponex solution (nitrogen:phosphorus:potassium, 5:10:5) wasused to irrigate plants every other day.

Production of the RNAi Transgenic Lines for AtAOR

The complementary DNA (cDNA) region of AtAOR (At1g23740) that trig-gers silencing was selected using the dsCheck program (Naito et al., 2005) toavoid off-target effects. The PCR primers used to amplify the selected 387-bpfragment of the AtAOR cDNA were 59-caccgagcgagaaagcattggaag-39 and59-gcgtcaaagacaacatcgta-39. The PCR products were cloned into the pENTR/D-TOPO vector and then transferred into the pHellsgate8 vector (Helliwell andWaterhouse, 2003) by an LR recombination reaction (Gateway; Invitrogen).Arabidopsis Columbia-0 was transformed with the pHellsgate8-At1g23740plasmid using Agrobacterium tumefaciens strain GV3101 and the floral dipmethod. Seeds were collected, and transformants were selected on mediumcontaining Murashige and Skoog salt mix, 50 mg mL21 kanamycin, and0.8% agar. The selected transgenic lines (T1 generation) were transferred to soil,

and T1 lines showing a strong silencing phenotype were selected by RT-PCR,cultivated until they flowered, and self-fertilized. The lines that showed stablesilencing effects in T2 were analyzed further.

Transient Expression in ArabidopsisMesophyll Protoplasts

The preparation of protoplasts from Arabidopsis rosette leaves and poly-ethylene glycol-calcium transfection were performed according to a methoddescribed previously (Yoo et al., 2007). The coding region of AtAOR in bothtransit and nontransit forms obtained by PCR using KOD-FXNeo (Toyobo)wassubcloned into SalI andNcoI sites of a pCaMV35S_GFP vector (Chiu et al., 1996)using the In Fusion HD Cloning Kit (Takara). The primers used in this exper-iment were 59-gtcgacatgaacgcagcgcttgcaac-39 and 59-ccatggcaggaatgggataaa-caacgacc-39. For transformation, 10 mg of each plasmid DNA was transfectedinto 2 3 104 protoplasts, and GFP and chlorophyll fluorescence were observedusing a fluorescence microscope (BZ-8000; Keyence). The field of cells was ex-cited at 480 nm, and fluorescence emission was detected at 510 nm.

Measurements of Chlorophyll Content in Leaves

The chlorophyll content was measured by the method of Porra et al. (1989).Leaf segments were incubated in N,N-dimethylformamide at 4°C overnight.Absorbances at 750, 663.8, and 646.8 nm were measured to calculate chloro-phyll content. Results were expressed as chlorophyll content per unit of leafarea.

Measurements of Nitrogen Content in Leaves

Total leaf nitrogen content was determined with Nessler’s reagent in a di-gestion solution after the addition of sodium-potassium tartrate (Makino andOsmond, 1991). Detached leaves were kept in a drying machine (60°C) anddehydrated overnight. Sulfuric acid (60% [v/v]; 100 mL) and dehydrated leaveswere mixed in the glass tube and incubated at 150°C in a heating block ther-mostat bath for 40 min. After cooling the glass tube in the air, 30% (v/v)hydrogen peroxide (H2O2; 50 mL) was added to the mixture. The mixturewas incubated at 180°C for 40 min in the heating block; subsequently, H2O2 (50mL) was added after the mixture was cooled. The incubation in the heatingblock and the addition of H2O2 were repeated another two times, but the in-cubation temperature was changed to 220°C and 260°C in these two incuba-tions. The incubation at 260°C and the addition of H2O2 were continued untilthe color of the mixture turned clear from brown. When the color turned clear,distilled water (4.95 mL) was added to the mixture and mixed vigorously. Themixture (500 mL), distilled water (4.25 mL), 10% (w/v) potassium sodium tar-trate solution (100mL), and 2.5 N sodium hydroxide (NaOH; 50mL)weremixed,andNessler’s reagent (100mL)was immediately added to themixture. Nitrogencontent was determined by the absorbance change at 420 nm.

SDS-PAGE

For protein analysis, approximately 1.5 g fresh weight of leaf tissue washomogenizedwith a pestle in 3mLof extraction buffer (50mMHEPES-KOH, pH7.6, 1 mM dithiothreitol [DTT], 2% [w/v] polyvinylpolypyrrolidone, 1 mM

phenylmethylsulfonyl fluoride, and 10 mM leupeptin). The homogenate wascentrifuged at 15,000g for 15 min at 4°C. The supernatant was treated as thesoluble fraction. The proteins were electrophoresed on 12.5% (w/v) SDS-polyacrylamide gels, as described by Laemmli (1970). The proteins (5 mg)were loaded on each lane, and the gels were stained with Coomassie BrilliantBlue R-250.

Western Blotting

The proteins were separated by SDS-PAGE, transferred onto a poly-vinylidene difluoride (PVDF) membrane (Millipore), and then blocked withBlocking One reagent (Nakalai Tesque) for 30 min at room temperature (25°C).The PVDF membrane was subsequently incubated with AtAOR-specific pep-tide antibody purchased from Hokkaido System Science for 1 h at room tem-perature. The PVDFmembranewaswashed three times with TBS-Tween buffer(10mMTris-HCl [pH 7.4], 0.14 MNaCl, and 0.1% [v/v] Tween 20) and incubatedwith enhanced chemiluminescence peroxidase-labeled anti-rabbit antibody(GEHealthcare) for 1 h at room temperature. The PVDFmembranewaswashed


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three times with TBS-Tween buffer. AtAOR protein was detected with Chemi-Lumi One Super (Nakarai Tesque) and scanned with an Ez-Capture MG (ATTO).

Gas-Exchange and Chlorophyll Fluorescence Analyses

The rates of CO2 andwater vapor exchangeweremeasuredwithDual-PAM-100 (Heinz Walz) and infrared gas analyzer (LI-7000; LI-COR) measuring sys-tems equipped with the 3010-DUAL gas-exchange chamber (Heinz Walz). Thesystems have been detailed by Makino et al. (1988). Gas with the indicatedmixture of pure oxygen and CO2 was prepared by mixing 20.1% (v/v) oxygenin 79.9% (v/v) N2 and 1% (v/v) CO2 in 99% (v/v) N2 using a mass-flow con-troller (Kofloc model 1203; Kojima Instruments). The mixture of gases wassaturated with water vapor at 13.5°C 6 0.1°C. The leaf temperature was con-trolled at 25°C. In the experiments that analyze the photosynthetic activity inArabidopsis leaves, chlorophyll fluorescence analysis was conducted simulta-neously by the Dual-PAM-101 (Walz). Gas-exchange parameters were calcu-lated according to the equations of von Caemmerer and Farquhar (1981), andchlorophyll fluorescence parameters (Fv/Fm and quantum yield of PSII) werecalculated according to the equations of Butler and Kitajima (1975) and Gentyet al. (1989). Leaves were illuminated (150 mE m22 s21) for 10 min, and photo-synthetic parameters were determined. For the calculation of chlorophyll flu-orescence parameters, measured light intensity (0.1 mE m22 s21) and saturatingpulse (300 ms, 10,000 mE m22 s21) were applied to determine Fo, maximum PSIIfluorescence in the dark-adapted state (Fm), steady-state fluorescence (Fs), andmaximum PSII fluorescence in the light-adapted state (Fm’).

Oxygen-Exchange Analysis

Leaf segments were placed in a cuvette (LD2/2; Hansatech). Oxygen uptakewas measured in the dark according to the method of Deliu andWalker (1981).Temperature-controlled water was circulated through the water jacket, and leaftemperature was maintained at 25°C.

Enzyme Activity Assays

For the evaluation of AtAOR activity, leaf protein was extracted in buffer(80 mM Tris-HCl, 1 mM EDTA, and 5% glycerol) and leaf protein solution wasobtained by centrifugation (15,000g, 2 min, and 4°C). AtAOR activity was de-termined in a reaction mixture (1 mL) that contained 50 mM potassium phos-phate (pH 7), 0.2 mM NADPH, and 20 mM acrolein. The acrolein-dependentoxidation of NADPH was followed by monitoring the decrease in A340, as-suming an absorption coefficient of 6.2 mM cm21.

For the evaluation of Rubisco activity, leaf protein was extracted in buffer(100 mM Tris-HCl [pH 7.8], 5 mM MgCl2, 2 mM EDTA, 5 mM DTT, 1 mM reducedglutathione, 0.5% [v/v] Triton X-100, and 5% [w/v] polyvinylpyrrolidone) andleaf protein solution was obtained by centrifugation (15,000g, 2 min, and 4°C).Rubisco activitywasmeasured as the decrease inA340 by coupling the activity ofRubisco to NADH oxidation using phosphoglycerate kinase and NAD+-GAPDH (Sawada et al., 1990). To measure total Rubisco activity, 1 mL of theleaf protein solution was transferred to a new tube, the MgCl2 concentrationwas brought to 20 mM, and the NaHCO3 concentration was brought to 10 mM.This mixture was kept on ice for 10 min before assaying for total Rubisco ac-tivity. The reaction mixture contained 50 mM HEPES-KOH (pH 8), 15 mM

MgCl2, 20 mM NaCl, 10 mM DTT, 0.5 mM ATP, 0.2 mM NADH, 5 mM phos-phoenolpyruvate, 5 mM creatine phosphate, 10 mM NaHCO3, 20 units of pyru-vate kinase, 2 units of phosphocreatine kinase, 18 units of NAD-GAPDH,18 units of phosphoglycerate kinase, 1 mM RuBP, and leaf protein solution.The reaction was started by adding 200 mM RuBP (10 mL) after preincubationfor 2 min and monitored by following the A340.

For the evaluation of FBPase, PRK, and NADP+-GAPDH activities, leafprotein was extracted in buffer (100 mM Tris-HCl [pH 7.8], 10 mM MgCl2, 1 mM

EDTA, 2.5 mM DTT, 1 mM reduced glutathione, 0.5% [v/v] Triton X-100, and5% [w/v] polyvinylpyrrolidone) and leaf protein solution was obtained bycentrifugation (15,000g, 2 min, and 4°C). FBPase activity was determined in areaction mixture (1 mL) that contained 100 mM Tris-HCl (pH 8), 10 mM MgCl2,0.5 mM EDTA, 0.4 mM NADP+, 0.1 mM Fru-1,6-bisP, 0.5 units of Glc-6-P dehy-drogenase, 1.5 units of phosphoglucose isomerase, and the leaf protein solution.The reaction was started by adding 20 mM Fru-1,6-bisP (10 mL) after pre-incubation for 2 min, and A340 was monitored (Tamoi et al., 1996a).

PRK activity was determined in a reaction mixture (1 mL) that contained100 mM Tris-HCl (pH 8), 100 mM KCl, 10 mM MgCl2, 0.2 mM NADH, 2 mM ATP,2.5 mM phosphoenolpyruvate, 2 mM ribose-5-phosphate, 5 units of lactate

dehydrogenase, 2 units of pyruvate kinase, 1 unit of phosphoriboisomerase,and the leaf protein solution. The reactionwas started by adding 200mM ribose-5-phosphate (10 mL) after preincubation for 2 min and was followed by mon-itoring A340 (Kobayashi et al., 2003).

NADP+-GAPDH activity was determined in a reaction mixture (1 mL) thatcontained 100 mM Tris-HCl buffer (pH 8), 10 mM MgCl2, 5 mM ATP, 0.2 mM

NADPH, 2 units of phosphoglycerate kinase, 3mM 3-phosphoglycerate, and the leafprotein solution. The reaction was started by adding 300 mM 3-phosphoglycerate(10 mL) after preincubation for 2 min, andA340 was monitored (Tamoi et al., 1996b).

For the evaluation of PEPC, CSY, ACO, and PDH activities, leaf protein wasextracted in buffer (50 mM KH2PO4-K2HPO4 [pH 7.6], 10 mM MgSO4, 1 mM

EDTA, 5 mM DTT, 0.05% [v/v] Triton X-100, 5% [w/v] polyvinylpyrrolidone,and one protein cocktail tablet [Roche]), as described byWatanabe et al. (2014), andleaf protein solution was obtained by centrifugation (15,000g, 2 min, and 4°C).

PEPC activity was determined in a reaction mixture (1 mL) that contained100 mMHEPES-NaOH buffer (pH 7.5), 10 mM MgCl2, 1 mM NaHCO3, 5 mMGlc-6-P, 0.2 mMNADH, and 12 units of malate dehydrogenase at 35°C. The reactionwas started by adding 400 mM phosphoenolpyruvate to 10 mL after pre-incubation for 2 min, and A340 was monitored (Fukayama et al., 2003).

CSY activity was determined in a reaction mixture (1 mL) that contained82mM Tris-HCl buffer (pH 7.9), 0.05% (v/v) Triton X-100, 2.8 mMmalate, 20 mM

3-acetylpyridine adeninedinucleotide, and 12 units of malate dehydrogenase at30°C. The reaction was started by adding 17 mM acetyl-CoA for 10 mL afterpreincubation for 2 min, and A365 was monitored (Jenner et al., 2001).

ACO activity was determined in a reaction mixture (1 mL) that contained80 mM HEPES-NaOH buffer (pH 7.5), 0.42 mM MnCl2, 0.05% Triton X-100,0.5 mM NADP+, and 1 unit of NADP isocitrate dehydrogenase at 30°C. Thereaction was started by adding 800 mM aconitate to a 10-mL reaction mixtureafter preincubation for 2 min, and A340 was monitored (Jenner et al., 2001).

PDH activity was determined in a reaction mixture (1 mL) that contained75 mM TES-NaOH buffer (pH 7.5), 0.5 mM MgCl2, 2 mM NAD+, 0.2 mM CoA,0.2 mM thiamine diphosphate, and 2.5 mM Cys-HCl at 25°C. The reaction wasstarted by adding 100 mM pyruvate to a 10-L reaction mixture after pre-incubation for 2 min, and A340 was monitored (Millar et al., 1999).

Isolation of RNA, RT-PCR, and Real-Time RT-PCR

Total RNA was isolated from plants using the RNeasy Plant Mini Kit(Qiagen). For the detection of mRNAs, first-strand cDNA was synthesizedfrom 1 mg of total RNA using a primer mix [random primer and oligo(dT)primer] and an enzyme mix (Rever Tra Ace and RNase inhibitor; Toyobo).PCR was performed in a 50-mL reaction solution containing 1 mL of cDNAmixture, 0.4 mM of each primer, and TaKaRa Ex Taq (Takara). The PCR cyclesconsisted of denaturation at 95°C for 10 s, annealing at 55°C for 30 s, and ex-tension at 72°C for 15 s. Quantitative real-time PCR was performed with SYBRPremix Ex Taq (Takara) using a LightCycler 1.5 (Roche), and the comparativethreshold cycle method was applied to determine the relative levels of mRNAs.The primers used in RT-PCR and real-time PCR are listed in SupplementalTable S5.

Detection of H2O2 in Leaves

H2O2 was detected visually in Arabidopsis leaves using 3,3-diaminobenzidine(Ren et al., 2002). Leaves were infiltrated with 10 mM MES (pH 5.5) containing0.1% (w/v) 3,3-diaminobenzidine for 3 h. Then, the leaveswere transferred to darkor light conditions (150 mE m22 s21, 23°C) and incubated for 3 h. After the incu-bation, the leaves were boiled in 96% ethanol until a brown color developed.

Transmission Electron Microscopy

Rosette leaves harvested at the end of the day and the end of the night weretrimmed into small pieces (2 3 3 mm2) with a razor blade; these pieces wereprefixed with 2.5% glutaraldehyde (Nisshin EM) in 0.1 M phosphate-bufferedsaline (PBS; pH 7.4) at 4°C overnight. The leaf pieces were then rinsed with PBSthree times at intervals of 10 min and postfixed with 1% PBS-buffered osmiumtetroxide (Nisshin EM) at room temperature (25°C) for 1 h. After the sampleshad been rinsed brieflywith distilledwater, theywere dehydrated immediatelyin an ethanol series (50%, 70%, 90%, and 100%). They were then immersed in anintermediate solvent (propylene oxide; Nisshin EM) for 10min and in amixture(1:1, v/v) of propylene oxide and Spurr’s resin (Spurr, 1969; Polysciences) for6 h at room temperature. Following this, the leaf pieces were placed in pureSpurr’s resin at 4°C for 3 d, embedded on plain embedding plates, and



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polymerized at 70°C for 24 h. Blocks of leaveswere cutwith a Porter-BlumMT-1ultramicrotome (Ivan Sorvall) and a diamond knife (Diatome). Ultrathin sec-tions of about 80 nmwere prepared; thesewere stainedwith 4% aqueous uranylacetate for 10 min and Sato’s lead for 10 min (Sato, 1968) at room temperature.Every staining step was succeeded by a step of washing with water three timesfor 3 min. The stained sections were observed with a JEM-1400 electron mi-croscope (JEOL) at an accelerating voltage of 75 kV.

Point-Counting Method

Chloroplasts in leaves were photographed at random at a magnification of4,0003 using a CCD camera. The photographs were printed and examinedusing point lattice-counting methods (Weibel, 1979; Ylä-Anttila et al., 2009). Asquare lattice was placed over the photographs, and the lattice line intersectionsthat were placed on the chloroplasts and starch granules were counted.

Starch Quantification

The amount of starch in wild-type and aor-1 leaves was quantified bio-chemically, as described by Sawicki et al. (2012) andMorita et al. (2015). Rosetteleaves were harvested at the end of the day and the end of the night (approx-imately 0.5 g fresh weight) and ground in liquid nitrogen, and 80% (v/v) eth-anol was added in the leaf powder. The solutionwas centrifuged at 10,000g for 5min at 4°C. The pellet was dried by vacuum centrifugation at 500g for 30 min at25°C. The dried pellet was suspended in 400 mL of dimethyl sulfoxide and 100mL of HCl, and this solution was incubated at 60°C for 30 min. After the in-cubation, the solution was centrifuged at 5,000g for 5 min at 25°C, and an ali-quot was used for starch quantification. A total of 400 mL of starch solution wasmixed with 400 mL of Lugol’s indole solution (24 mM KI, 9.5 mM I2, and 10 mM

HCl), and A600 was monitored.

Statistical Analysis

All measurement data were expressed as means 6 SE of at least three in-dependent measurements. We used Student’s t test and the Tukey-Kramerhonestly significant difference test to detect differences between wild-typeand aor mutant plants. All statistical analyses were performed using Micro-soft Excel 2010 and JMP8 (SAS Institute).

Supplemental Data

The following supplemental materials are available.

Supplemental Figure S1. Subcellular localization of AtAOR in a plant cell.

Supplemental Figure S2. Comparison of respiration rates in wild-type andaor (RNAi) mutant plants evaluated by oxygen electrode.

Supplemental Figure S3. Quantification of the starch content in wild-typeand aor-1 leaves harvested at the end of the day and the end of the night.

Supplemental Figure S4. Characterization of aor (T-DNA) plants grownunder constant light growth conditions.

Supplemental Figure S5. Gene expression analysis related to respirationand ACT2 in wild-type and aor-1 leaves grown under constant lightconditions.

Supplemental Figure S6. Evaluation of enzyme activities of PDH, PEPC,ACO, and CSY in wild-type and aor-1 leaves grown under constant lightconditions.

Supplemental Figure S7. Evaluation of starch content in wild-type andaor-1 leaves grown under constant light conditions.

Supplemental Figure S8. Evaluation of enzyme activities of Rubisco,GAPDH, FBPase, and PRK in wild-type and aor-1 leaves grown underday/night cycle conditions.

Supplemental Figure S9. Detection of H2O2 accumulation in wild-type andaor-1 leaves grown under day/night cycle growth conditions.

Supplemental Figure S10. Photosynthetic activity in wild-type and aor-1leaves grown under day/night cycle conditions.

Supplemental Figure S11. Evaluation of chlororespiration activity in wild-type and aor-1 leaves grown under day/night cycle conditions.

Supplemental Table S1. Chlorophyll and nitrogen contents in wild-typeand aor (T-DNA) plants grown under day/night cycle conditions.

Supplemental Table S2. Dark respiration rate and photosynthetic param-eters in wild-type and aor (T-DNA) plants grown under day/night cycleconditions.

Supplemental Table S3. Chlorophyll and nitrogen contents in wild-typeand aor (T-DNA) plants grown under continuous light conditions.

Supplemental Table S4. Dark respiration rate and photosynthetic param-eters in wild-type and aor (T-DNA) plants grown under constant lightconditions.

Supplemental Table S5. Genes and primers used for RT-PCR and real-time PCR.

ACKNOWLEDGMENTS

We thank Dr. Yasuo Yamauchi (Kobe University) for giving us aor (T-DNA)mutant seeds and Editage (Cactus Communications; http://www.editage.jp/)for editing the article.

Received October 6, 2015; accepted February 13, 2016; published February 16,2016.

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suppression of chloroplastic alkenal/one oxidoreductase ......bases (esterbauer et al., 1991; miyata...

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