Regulation of growth response to water stress in the soybean primary root. I. Proteomic analysis...

Regulation of growth response to water stress in thesoybean primary root. I. Proteomic analysis revealsregion-specific regulation of phenylpropanoid metabolismand control of free iron in the elongation zonepce_2073 223..243

MINEO YAMAGUCHI1†, BABU VALLIYODAN1†, JUAN ZHANG2, MARY E. LENOBLE1, OLIVER YU2,ELIZABETH E. ROGERS3*, HENRY T. NGUYEN1 & ROBERT E. SHARP1

Divisions of 1Plant Sciences and 3Biochemistry, University of Missouri, Columbia, MO 65211, USA, and 2Donald DanforthPlant Science Center, St Louis, MO 63132, USA

ABSTRACT

In water-stressed soybean primary roots, elongation wasmaintained at well-watered rates in the apical 4 mm (region1), but was progressively inhibited in the 4–8 mm region(region 2), which exhibits maximum elongation in well-watered roots. These responses are similar to previousresults for the maize primary root. To understand theseresponses in soybean, spatial profiles of soluble proteincomposition were analysed. Among the changes, the resultsindicate that region-specific regulation of phenylpropanoidmetabolism may contribute to the distinct growth responsesin the different regions. Several enzymes related to isofla-vonoid biosynthesis increased in abundance in region 1,correlating with a substantial increase of isoflavonoidcontent in this region which could contribute to growthmaintenance via various potential mechanisms. In contrast,caffeoyl-CoA O-methyltransferase, which is involved inlignin synthesis, was highly up-regulated in region 2. Thisresponse was associated with enhanced accumulation oflignin, which may be related to the inhibition of growth inthis region. Several proteins that increased in abundance inboth regions of water-stressed roots were related to protec-tion from oxidative damage. In particular, an increase in theabundance of ferritin proteins effectively sequestered moreiron and prevented excess free iron in the elongation zoneunder water stress.

Key-words: drought; isoflavonoids; lignin; proteomics; reac-tive oxygen species; root growth.

INTRODUCTION

Drought is the major abiotic stress factor limiting crop pro-ductivity worldwide (Boyer 1982), and understanding the

genetic and biochemical mechanisms that control droughttolerance is a central question in plant biology. One aspectof principal importance in this arena is the response of rootgrowth and development to water stress conditions. Underwater stress, the growth responses of different plant partsare varied and complex, and some types of roots can con-tinue elongation at low water potentials that completelyinhibit shoot growth (Sharp & Davies 1989; Spollen et al.1993). In previous studies of the maize (Zea mays L.)primary root, kinematic principles were applied to charac-terize spatial patterns of expansion within the elongationzone, and revealed that elongation rates were preferentiallymaintained towards the root apex at low water potentials(Sharp, Silk & Hsiao 1988; Liang, Sharp & Baskin 1997).Remarkably, elongation rates were unaffected in the apicalfew millimetres even under severe water stress (waterpotential of -1.6 MPa), but were progressively inhibited atmore basal locations resulting in a shortened elongationzone. This spatial characterization provided an essentialfoundation for extensive research into the physiologicalmechanisms of growth regulation in the maize primary rootat low water potentials (reviewed in Sharp et al. 2004; Ober& Sharp 2007). Recently, these studies were expanded toinclude transcriptomic (Bassani, Neumann & Gepstein2004; Poroyko et al. 2007; Spollen et al. 2008) and cell wallproteomic (Zhu et al. 2007) analyses, which revealed majorand largely region-specific changes in gene expression andprotein composition profiles between well-watered andwater-stressed roots, and provided additional insights intothe processes involved in regulating the differentialresponses of cell elongation to water deficit in the differentregions of the elongation zone.

Although the relative maintenance of overall elongationrate of the primary root compared to the shoot has beenreported for water-stressed plants of several species(Spollen et al. 1993), the associated mechanisms of rootgrowth regulation have not been investigated in detail inspecies other than maize. Accordingly, it is not knownwhether similar or different mechanisms are involved in theadaptation of root growth to water stress in other species.

Correspondence: R. E. Sharp. Fax: +1 573 882 1469; e-mail: [email protected]

*Present address: USDA-ARS, 9611 S. Riverbend Ave., Parlier, CA93648, USA.†These authors contributed equally to the article.

Plant, Cell and Environment (2010) 33, 223–243 doi: 10.1111/j.1365-3040.2009.02073.x

© 2010 Blackwell Publishing Ltd 223

To address this question, we have initiated a comprehen-sive analysis of the molecular and physiological processesinvolved in adaptation to water stress in the primary root ofsoybean [Glycine max (L.) Merr.]. Worldwide, legumes aresecond to grass species in economic importance. Soybean isthe most important leguminous crop plant, and is used as amajor source of protein and oil. In addition, being a dicoty-ledonous plant, soybean provides a contrast with maize incharacteristics such as root system architecture. Soybeanalso has distinct metabolic features including nitrogen fixa-tion processes and isoflavone synthesis.

This study reports the spatial distribution of the responseof elongation rate to low water potential within thesoybean primary root elongation zone, and presents a pro-teomic analysis of associated changes in soluble proteinabundance. Proteomic analyses provide a powerful tool toinvestigate the physiological basis of plant responses tostress. Several proteomic studies have been reported inwater-stressed leaves of various species (Costa et al. 1998;Riccardi et al. 1998; Hajheidari et al. 2005; Vincent et al.2005; Bhushan et al. 2007; Gazanchian et al. 2007; Kottapalliet al. 2009), and the results implicate not only a variety ofstress-related functions of the regulated proteins, but alsodifferent patterns of protein compositional changes in dif-ferent species. However, limited research has been con-ducted on changes in protein composition in the roots ofwater-stressed plants (Bianchi, Damerval & Vartanian2002a; Plomion et al. 2006; Rabello et al. 2008; Yoshimuraet al. 2008), and, to our knowledge, proteomic studies on theroot elongation zone under water stress are limited to thementioned study of cell wall proteins in the maize primaryroot (Zhu et al. 2007). In this study, spatial analysis ofsoluble protein composition within the soybean primaryroot elongation zone was combined with knowledge ofelongation rate patterns to provide insight into novelmechanisms that are involved in soybean primary rootgrowth adaptation to low water potentials.

MATERIALS AND METHODS

Plant material, growth conditions andharvest times

Soybean (cv. Magellan) seeds were surface sterilized in 1%NaClO solution for 2 min, rinsed in deionized water for30 min and germinated between sheets of germinationpaper moistened with a solution containing 5 mm CaCl2

and 5 mm Ca(NO3)2 for 36 h at 29 °C and near-saturationhumidity in the dark. Seedlings with primary roots approxi-mately 15 mm in length were transplanted against the inte-rior surface of Plexiglas cylinders (14.5 cm diameter) filledwith a 1:1 (v/v) mixture of vermiculite (no. 2A, Therm-O-Rock East Inc., New Eagle, PA, USA) and Turface (ProfileProducts LLC, Buffalo Grove, IL, USA) at water potentialsof -0.1 MPa (well-watered treatment, moistened to the drippoint) or -1.6 MPa (water-stressed treatment), which wereobtained by thorough mixing with different volumes of5 mm CaCl2 + 5 mm Ca(NO3)2 solution. Approximately 1.4

or 0.27 L of solution was mixed with 1 kg of vermiculite–Turface to produce high or low water potential media,respectively. Vermiculite–Turface water potentials weremeasured at the beginning of each experiment by isopiesticthermocouple psychrometry (Boyer & Knipling 1965). Theseedlings were then grown under the same conditions untilharvest. Primary root elongation was monitored by periodi-cally marking the position of the root apices on the Plexi-glas. Transplanting, growth measurements and harvestingwere performed using a green ‘safe’ light (Saab et al. 1990).

In the water-stressed treatment, primary roots were har-vested 48 h after transplanting in all experiments except formeasurement of cell length profiles. Root tip water poten-tial (measured by isopiestic thermocouple psychrometry)had decreased to approximately -1.6 MPa by this time(data not shown). Because the water-stressed roots elon-gated more slowly than the well-watered roots, two well-watered controls were collected in most experiments.Firstly, a developmental control was harvested 24 h aftertransplanting (roots of the same length as the waterstressed), and secondly, a temporal control was harvested48 h after transplanting (roots of the same age as the waterstressed) (Fig. 1). The two well-watered controls wereimportant to help identify true responses to water stress,because changes in protein abundance and other param-eters might also vary with root development under

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Figure 1. Representative time courses of primary root lengthincrease after transplanting soybean (cv. Magellan) seedlingsto well-watered (WW, water potential of -0.10 MPa) orwater-stressed (WS, water potential of -1.6 MPa) conditions.Data are means of at least nine seedlings; error bars (�SE) aresmaller than the symbols. As indicated by arrows, root sampleswere collected for proteomic analysis at 48 h after transplantingfor the water-stressed treatment, and at 24 h (WWD,developmental control, roots of the same length as thewater-stressed roots) and 48 h (WWT, temporal control, roots ofthe same age as the water-stressed roots) for the well-wateredtreatment.

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© 2010 Blackwell Publishing Ltd, Plant, Cell and Environment, 33, 223–243

well-watered conditions. Depending on the experiment, theapical 25 mm of each root was immediately sectioned intosome or all of the following regions (distances are from theroot apex including the root cap): region 1, 0–4 mm; region2, 4–8 mm; region 3, 8–15 mm; region 4, 15–25 mm (Fig. 2).

Cell length and relative elongation rate profiles

To define the length of the elongation zone and describe theelongation rate profile in the primary roots of well-wateredand water-stressed seedlings, spatial distributions of relativeelongation rate (h-1) were calculated from root elongationrates and cell length profiles (Silk, Lord & Eckard 1989).Accurate determination of relative elongation rate profilesfrom anatomical records requires conditions of steadygrowth and cell length distribution. Figure 1 illustrates thatroot elongation rates were essentially steady in both well-watered and water-stressed roots after 24 h from trans-planting. However, because the water-stressed roots hadslower elongation rates, the harvest time for cell lengthmeasurements was increased from 48 h in the well-wateredtreatment to 72 h in the water-stressed treatment, to allowgreater time for stabilization of the cell length profile aftertransplanting to the low water potential condition.

In each of four replicate experiments, 10 seedlings weregrown under well-watered and water-stressed conditions,and root elongation rates were measured for 10 h priorto harvest to select one or two roots that were

both straight and had the closest elongation rates to themean. Cell lengths were then measured as a function ofdistance from the root apex by confocal microscopy [Bio-Rad Radiance 2000 (Hercules, CA, USA) coupled to anOlympus IX70 inverted microscope, Olympus America,Valley, PA, USA]. To assist in visualizing the cells, theplasma membranes were labelled with the fluorescentprobe FM 1-43 [N-(3-triethylammoniumpropyl)-4-(4-(dibutylamino)styryl)pyridinium dibromide; MolecularProbes, Eugene, OR, USA] by incubating the root apicesin 80 mm dye solution for 30 min. Optically sectionedroot images were then obtained using 488 nm excitationand emission beyond 580 nm (580 nm long pass), and thelengths of cortical (outer two layers) and epidermal cellswere measured directly from the images (no distinctionwas made between the cortical and epidermal cells,although the measurements were dominated by corticalcells). For each root, lengths of 3–10 cells were measuredat approximately 0.5 mm intervals from the root apexuntil unchanging mean cell lengths were obtained forseveral successive positions.

The spatial distribution of displacement velocity(mm h-1) away from the root apex was calculated from theroot elongation rate and cell length profile using the rela-tionship LA/LF = VA/VF, where LA is the mean cell length atposition A, LF is final cell length, VA is displacement velocityat position A, and VF is the final displacement velocity(equal to the root elongation rate). Final cell lengths weredetermined by averaging cell lengths at the four most distalmeasurement positions. It should be noted that displace-ment velocities cannot be accurately derived from celllength measurements in the meristematic region (Silk et al.1989); therefore, velocities were calculated starting at thedistal end of the meristem, which was estimated to occur ata cell length of 2.5 times the length of the shortest cell(Erickson 1961). The relative elongation rate profile wasobtained from the derivative of a fifth-order polynomialcurve fitted to the values of displacement velocity plottedagainst position (Fig. 2, inset), including the origin at theroot–root cap junction.

Root harvest and extraction of totalsoluble proteins

In each of four replicate experiments, approximately 200roots were harvested from the water-stressed treatment andfrom the well-watered developmental and temporal con-trols. The root tips were sectioned into regions 1–2 (waterstressed) and 1–3 (controls), giving a total of eight samplesper experiment. The root sections were immediatelyimmersed in liquid nitrogen and then stored at -80 °C.Totalsoluble proteins were extracted with phenol followed bymethanolic ammonium acetate precipitation (Hurkman &Tanaka 1986; Mooney, Krishnan & Thelen 2004). Briefly,root segments (0.2–0.3 g fresh weight) were ground to apowder with liquid nitrogen, and suspended directly withhomogenization media containing 50% (v/v) phenol, 0.45 msucrose, 5 mm EDTA, 0.2% (v/v) 2-mercaptoethanol, and

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Figure 2. Relative elongation rate and displacement velocity(inset) as a function of distance from the apex (including the rootcap) of primary roots of well-watered (WW) and water-stressed(WS) soybean seedlings. Relative elongation rates were obtainedas the derivative of displacement velocity with respect to positionusing curves fitted by a fifth-order polynomial. Displacementvelocities were calculated from root elongation rates and celllength profiles at 48 h (well watered) or 72 h (water stressed)after transplanting; values are means � SE of four to five rootsfrom four independent experiments. Region 1 (R1, 0–4 mm),region 2 (R2, 4–8 mm), region 3 (R3, 8–15 mm) and region 4(R4, 15–25 mm), as harvested for measurements in this study, areindicated. Root cap lengths (�SE) were 0.48 � 0.01 mm and0.41 � 0.02 mm in well-watered and water-stressed roots,respectively.

Proteomic analysis of soybean root growth response to water stress 225


50 mm Tris–HCl (pH 8.8). The homogenate was agitated for30 min at 4 °C, and centrifuged for 30 min at 5000 g. Theupper phenol phase was collected and proteins were pre-cipitated by adding six volumes of 0.1 m ammonium acetatein 100% methanol, and incubating at -20 °C for a minimumof 1 h. Precipitated proteins were collected by centrifuga-tion (20 min at 5000 g) and washed three times with0.1 m ammonium acetate in 100% methanol, three timeswith 80% (v/v) acetone, and once with 70% (v/v) ethanol.Washed proteins were stored in 80% acetone at -20 °C.

Protein separation by two-dimensionalelectrophoresis (2-DE) and gel image analysis

For each of the four experiments, the eight proteinsamples were separated by 2-DE in two sets as follows.For set 1, three gels were run for the region 1 samples(water-stressed, well-watered developmental and temporalcontrols). For set 2, five gels were run for the region 2samples (water-stressed, well-watered developmental andtemporal controls) and region 3 samples (well-wateredcontrols only). 2-DE was carried out using 24 cm pH 4.0–7.0 immobiline pH gradient (IPG) strips (Bio-Rad) and12–18% acrylamide separating gels. (In preliminaryexperiments with region 1 samples, pH 3.0–10.0 stripswere used. Because most of the differentially expressedproteins were detected within pH range 4.0–7.0, thisrange was selected for detailed analysis to obtainmaximum separation of protein spots and improvedconfidence of protein identifications.) Protein pellets weredissolved in isoelectric focusing resuspension media[8 m urea, 2 m thiourea, 4% (w/v) 3-[(3-cholamidopropyl)dimethylammonio]-1-propanesulphonate, 100 mm DTT,2% Triton X-100, 1% (v/v) IPG buffer (GE Healthcare,Piscataway, NJ, USA)], and proteins were quantified usingthe EZQ protein assay kit (Molecular Probes). The IPGstrips were passively rehydrated with 450 mL samples(500 mg protein) for 15 h (preliminary trials with differentprotein amounts determined that 500 mg was optimal forgel resolution). Isoelectric focusing was carried out usingProtean IEF Cell (Bio-Rad) with the following program:1 h at 250 V, 0.5 h at 1000 V, 3 h to increase from 1000 to8000 V, and 11 h at 8000 V (total of 94 750 Vh). Afterfocusing, the IPG strips were incubated for 20 min in8.5 mL of equilibration buffer containing 50 mm Tris–HCl(pH 8.8), 6 m urea, 20% (v/v) glycerol and 2% (w/v)sodium dodecyl sulphate (SDS) supplemented with 2%(w/v) DTT, and then further incubated in equilibrationbuffer supplemented with 2.5% iodoacetamide for 20 min.The IPG strips were loaded onto second-dimension gelsand sealed with 0.5% (w/v) agarose (Fisher Biotech, WestPerth, Western Australia, Australia) containing a trace ofthe tracking dye bromophenol blue (Fisher Scientific,Waltham, MA, USA). Sodium dodecyl sulphate–polyacrylamide gel electrophoresis (SDS–PAGE) was runat 3 W gel-1 for 1 h, and then at 2 W gel-1 at 20 °C until thebromophenol blue dye front reached the bottom of thegels using the Ettan DALTsix Electrophoresis system (GE

Healthcare). The running buffer composition was 25 mmTris, 0.192 m glycine and 0.1% (w/v) SDS; 1¥ concentrationwas used as the upper tank buffer, and 2¥ as the lowertank buffer. Gels were then stained with a modified col-loidal Coomassie staining method (Neuhoff et al. 1988),and gel images were taken using a FLA5000 gel scanner(Fujifilm, Tokyo, Japan).

Spot detection, quantification and matching were per-formed for each set of gels.Thus, for each of the four experi-ments, the results provided within-region comparisons forregion 1 (set 1) and region 2 (set 2), and comparison ofwater-stressed region 2 with well-watered region 3 (set 2;see Results for further details of this analysis). Gels wereanalysed with Bio-Rad PDQuest version 7.3.1 software;within each region, the well-watered temporal control wasused as the reference (master) gel. After background sub-traction and spot detection, spots were matched and nor-malized using the method of total density in the gel image.

To determine water stress-regulated proteins, spotintensities in regions 1 and 2 were compared between thewater-stressed sample and each of the well-watered controlsamples. For differentially expressed proteins within region2, spot intensities of the water-stressed samples were furthercompared with region 3 of the well-watered controls.Restricted maximum likelihood analysis (Imin et al. 2005)was used to test for statistical significance of treatmenteffects using PROC MIXED in SAS version 9.1 (SAS Insti-tute Inc., Cary, NC, USA). The water stress treatment andthe two well-watered controls were considered as fixedeffect factors, and replicates were treated as random factors(Imin et al. 2005). Proteins were considered to be differen-tially expressed when the spot intensity of the water-stressed sample was statistically different (P < 0.05) fromboth of the control samples in pairwise comparisons usingthe Tukey–Kramer adjustment method. If spot fusion pre-vented spot quantification in one replicate, the other threereplicates were used for statistical analysis and determina-tion of induction level. To describe the level of proteinregulation under water stress, protein abundance ratioswere calculated between the water-stressed sample andeach of the well-watered controls, and the two abundanceratios were averaged.

In-gel tryptic digestion of proteins

Thirty-five water stress-responsive proteins were excisedmanually from each of the four replicate gels of region 1and/or 2 samples (region 3 samples were not used forprotein identifications) using a 1.5 mm 2D gel spot picker(The Gel Company, San Francisco, CA, USA). In general,spots were excised from gels of water-stressed samples forproteins that increased in abundance under water stress,and from gels of well-watered samples for proteins thatdecreased in abundance under water stress; final selectionwas based on quality of spot resolution on the gels. In-geltryptic digestion was conducted according to Mooney et al.(2004, 2006). Gel plugs were washed four times for 15 minin acetonitrile/50 mm ammonium bicarbonate (50, 50%,



v/v), incubated for 20 min in 100% acetonitrile, and com-pletely air dried. The plugs were then re-hydrated with20–40 mL trypsin solution (20 mg mL-1; Promega, Madison,WI, USA), and incubated for 1 h at 4 °C. The trypsin solu-tion was then removed and replaced with 30 mL of 50 mmammonium bicarbonate and 10% acetonitrile to keep thegel plugs hydrated throughout the digest. Trypsin digestionwas performed overnight at 37 °C. After centrifugation, thesupernatants containing tryptic peptides were transferredto new tubes. Gel plugs were washed twice with acetonitrile/MilliQ water/trifluoroacetic acid (60, 39, 1%, v/v) solution,and peptides were collected and pooled. The pooled digestswere lyophilized and reconstituted in 5 mL of acetonitrile/MilliQ water/88% formic acid (50, 49, 1%, v/v).

Protein identification by matrix-assisted laserdesorption/ionization-time-of-flight tandemmass spectrometry (MALDI-TOF MS/MS) anddatabase searching

MALDI-TOF MS/MS analyses were conducted accordingto Medzihradszky et al. (2000). A 0.5 mL fraction of eachpeptide sample was mixed with an equal volume of5 mg mL-1 a-cyano-4-hydroxycinnamic acid (MS grade,Fluka, St Louis, MO, USA) in acetonitrile/MilliQ water/10% TFA/100 mm ammonium dihydrogen phosphate (50,38, 2, 10%, v/v). The sample/matrix mix (0.3 mL) was depos-ited on a polished stainless steel target plate (ABI01-192-6-AB, Applied Biosystems, Foster City, CA, USA).Crystallization of the mixture proceeded under ambientconditions, and the crystals were not washed (Smirnov et al.2004). Analysis of the tryptic peptides was conducted usinga MALDI-TOF/TOF MS (model 4700, Applied Biosys-tems) with a 355 nm Nd : YAG laser (200 Hz) in the posi-tive ion delayed extraction reflector MS or MS/MS mode.Peptide standards (4700 Mass Standards Kit; Applied Bio-systems) were used for calibration. Parent ion spectra (400laser shots summed/averaged) were obtained over the massrange 700–4000 Da. The eight most intense parent ionswere automatically selected for MS/MS analysis. Threecommon trypsin autolysis peptides, 842.51, 870.5412 and2211.1046 Da, were excluded from the MS/MS analysis.MS/MS spectra (3000 laser shots) were obtained with themetastable ion suppressor on and using a 1 kV potentialdifference. The peak list was generated using GPS Explorersoftware (version 3.0; Applied Biosystems). Peak listsobtained from the MALDI-TOF MS, MS/MS mass spectrawere deisotoped, and following baseline correction andnoise reduction, were submitted online to Matrix Science’ssearch engines (www.matrixscience.com) against theNCBInr ‘viridiplantae’ protein database. Protein identifica-tions were considered significant only if the Mascot scorewas confident (>95%), at least three peaks matched theprotein (except in the case of spot 16, sc32f08.y1, forwhich only two peaks were matched) and a percentage ofcoverage higher than 10% was obtained. Recentlyavailable soybean genome sequence information (http://www.phytozome.net) also allowed us to search for the

respective soybean gene identification of differentiallyexpressed proteins through basic local alignment searchtool (BLAST) analysis.

Isoflavonoid analysis

In each of three experiments, approximately 100 roots wereharvested from the water-stressed and well-watered (devel-opmental and temporal controls) treatments, and the roottips were sectioned into regions 1–3, immediately immersedin liquid nitrogen and stored at -80 °C. The segments (100–200 mg fresh weight per region) were ground to a powderunder liquid nitrogen, and extracted with 500 mL of 80%methanol. The extracts were concentrated using an Eppen-dorf vacuum concentrator at 30 °C, and then dissolved in80% methanol for high-performance liquid chromatogra-phy (HPLC) analysis. Samples were analysed for isoflavonecomposition using an Agilent 1100 series HPLC system(Agilent Technologies, Santa Clara, CA, USA) with aSpherisorb ODS-2 reversed-phase C-18 column (5 mm;250 ¥ 4.6 mm). HPLC samples were separated using an18 min linear gradient from 20% methanol, 80% 10 mmammonium acetate (pH 5.6), to 100% methanol at a flowrate of 1 mL min-1. Elution of metabolites was monitoredby a photodiode array. Retention time and UV spectra werecompared to those of authentic standards.

For histochemical analysis of isoflavonoids and fla-vonoids, roots were stained with 0.25% (w/v) diphenyl boricacid-2-aminoethyl ester (DPBA) in 0.02% (v/v) TritonX-100 for 30 min (Sheahan & Rechnitz 1993). Stained rootswere observed under a stereomicroscope (MZFLIII, Leica,Wetzlar, Germany) with UV excitation (maximum at350 nm) and emission beyond 420 nm.

Lignin detection

Histochemical analysis of lignin was performed for water-stressed and well-watered (temporal control) roots accord-ing to Fan et al. (2006) with slight modifications. Rootelongation rates of 10 seedlings were measured during the10 h prior to harvest to select one or two roots which hadthe closest elongation rates to the mean. One-millimetre-long segments were sectioned from the middle of regions1–4 (1.5–2.5 mm, 5.5–6.5 mm, 11–12 mm and 19.5–20.5 mmfrom the root apex). Cell wall ghosts were prepared bytreatment with hot methanol (65 °C) for 5 min, and werethen washed with methanol and phosphate-buffered saline(pH 7.4). Lignification was examined by autofluorescenceof wall phenolics and by Mäule staining (Higuchi 1998). Forautofluorescence, the methanol-treated root segments werefixed with 3.7% formaldehyde for 48 h. Transverse sections(35 mm in thickness) were made from the middle of eachsegment using a cryostat microtome (CM 1850; Leica).Autofluorescence was observed under UV excitation(maximum at 359 nm) and emission between 460 and500 nm using a fluorescence microscope (Olympus IX70).The fluorescence intensity of the xylem and epidermis wasquantified using Metamorph software (MDS Inc., Toronto,



ON, Canada). For Mäule staining, transverse sections(0.15 mm in thickness) were made from the middle ofeach methanol-treated segment using a Lancer series1000 Vibratome (Vibratome, St Louis, MO, USA). The sec-tions were treated with 0.5% KMnO4 for 5 min, rinsed withwater, destained with 10% HCl for 5 min, rinsed with waterand then mounted in 28% NH4OH. The stained sectionswere observed under a stereomicroscope (MZFLIII,Leica).

Ferric iron staining

Ferric iron in root tips (apical 5 mm) harvested from thewater-stressed and well-watered (developmental and tem-poral controls) treatments was visualized using the Perls’staining method, as described by Green & Rogers (2004).Briefly, equal amounts of solutions of 4% (v/v) HCl and 4%(w/v) potassium ferrocyanide were mixed immediatelyprior to use. The staining solution was vacuum infiltratedinto the root tips for approximately 3 h at room tempera-ture. The root tips were then rinsed in water, and stainingwas observed immediately using a stereomicroscope. Toexpose iron staining inside the root, stained root tips weresquashed between a glass slide and cover glass by hand. Tomake transverse or longitudinal sections, stained root tipswere fixed in 4% (w/v) formaldehyde and embedded in 4%(w/v) low-melt agarose. Sections (0.15 mm in thickness)were cut with a Vibratome, and observed under a stereo-scope or inverted microscope.

Native PAGE, staining for protein-bound ironand ferritin Western blot analysis

Total soluble proteins were extracted from water-stressedand well-watered (temporal control) roots. The root tipswere sectioned into regions 1 and 2, and region 1 was sub-divided into the 0–1 mm and 1–4 mm regions. The root seg-ments were homogenized in buffer containing 0.25 msorbitol, 10 mm 3-morpholinopropanesulphonic acid–Tris(pH 7.3) and 0.01% proteinase inhibitor cocktail (Sigma).The slurry was centrifuged at 15 000 g for 30 min, and thesupernatant was used for protein analysis. One-dimensionalnative PAGE was performed using 4–20% acrylamide gra-dient gels according to Laemmli (1970) except that SDSwas omitted from the buffer system. Approximately 70 mgof protein was loaded per lane. For detection of iron, thegels were stained with reagent solution containing 0.75 mmFerene S [3-(2-pyridyl)-5,6-di(2-furyl)-1,2,4-triazine-5′,5″-disulfonic acid; Sigma], 15 mm thioglycolic acid and 2%(v/v) acetic acid (Chung 1985). Bands, indicating presenceof iron, developed in 5 min. Gels were further stainedaccording to Kuo & Fridovich (1988) and Topham, Eads &Butler (1992) except that diaminobenzoic acid was replacedwith diaminobenzidine (DAB) which proved to be moresensitive for iron detection.

Western blots were performed using standard protocols.Separated proteins were transferred to polyvinylidene

difluoride membranes, and the membrane was probed withantiserum raised against pea ferritin (which recognizessoybean ferritins, as well as those from other plants;Laulhere, Lescure & Briat (1988). The second antibodywas a goat anti-rabbit IgG conjugated to alkaline phos-phatase, and the band was visualized by reaction withnitro-blue tetrazolium chloride and 5-bromo-4-chloro-3′-indolylphosphate.The intensity of each band was quantifiedby G : BOX (Syngene, Cambridge, UK).

RESULTS

Growth response of the soybean primary rootto water stress

To study the response of soybean primary root growth towater stress, low water potentials were imposed using amodified version of the seedling culture system that hasbeen used extensively by Sharp and co-workers to studymaize primary root growth under water stress (Sharp et al.2004; Ober & Sharp 2007). Preliminary experiments estab-lished that for studies of soybean, a 50/50 (v/v) mixture ofvermiculite and Turface (a fired clay substrate) resulted ingreater root elongation rates (possibly because of superiorbuffering characteristics) under both well-watered andwater-stressed conditions compared to growth in 100% ver-miculite as has been used for studies of maize. In addition,provision of 10 mm [Ca2 +] [supplied as 5 mm CaCl2 + 5 mmCa(NO3)2] during germination and after transplanting (asrecommended for soybean seedling studies; Boyer, Univer-sity of Delaware, personal communication) was found toprevent the seed-borne fungal disease anthracnose, whichotherwise caused progressive inhibition of growth rates insome lines.

Initial studies were conducted with a genetically diversecollection of 19 soybean lines, which were examined forprimary root elongation rates under well-watered and mild(-0.25 MPa) and severe (-1.6 MPa) water stress conditions(data not shown). Both natively and non-natively adaptedgenotypes including elite as well as Plant Introduction lineswere tested, which were chosen because of known variabil-ity in agronomic characteristics including yield, drought tol-erance and deep and/or prolific rooting. From these results,cv. Magellan was selected as the focus of this study. Rootelongation rates of this line were among the most rapid(approximately 3 mm h-1) under well-watered conditions,and among the least inhibited under water-stressed condi-tions (approximately 1.3 mm h-1, or 43% of well-watered, ata water potential of -1.6 MPa) (Fig. 1). (In the lines tested,root elongation rate at -1.6 MPa ranged from 27 to 50% ofwell-watered values.) Magellan (maturity group IV, indeter-minate growth habit) has also been reported to have highyield potential and drought tolerance characteristics(Schapaugh et al. 1998; Sleper, University of Missouri,personal communication).

By growing the seedlings at near-saturation humidity inthe dark, evaporative water loss after transplanting wasminimized, and, therefore, the seedlings were exposed to



essentially constant conditions of media water potential(measurements showed that the water potential of thevermiculite–Turface media decreased by less than 0.06 MPaover the course of the water stress experiments).As a result,root elongation rates were steady from 24 to 72 h (themaximum duration of experiments in this study) after trans-planting to both well-watered and water-stressed conditions(Fig. 1). Shoot growth was completely inhibited under thiswater stress condition (data not shown).

Analysis of the spatial distribution of relative elongationrate showed that in well-watered roots, longitudinal expan-sion occurred throughout the apical 16 mm (relative elon-gation rate fell to 0 at approximately 16 mm from the rootapex) and exhibited a maximum rate at approximately5 mm from the apex (Fig. 2). In the water-stressed roots,relative elongation rates were the same as in well-wateredroots until almost 4 mm from the apex, but were then pro-gressively inhibited and fell to zero at about 8 mm from theapex. The maximum relative elongation rate also decreasedfrom 0.37 to 0.30 h-1, and shifted apically compared to thewell-watered roots. These results show that the spatial dis-tribution of the response of elongation rate to water stressin the soybean primary root is very similar to that previ-ously reported for the maize primary root (Sharp et al. 1988;Liang et al. 1997); longitudinal expansion is fully main-tained under quite severe water stress conditions in theearly ontogenetic phases of growth, but deceleration andcessation of expansion occur closer to the apex than inwell-watered roots, resulting in a shorter elongation zone.

Figure 2 illustrates the four contiguous regions within theapical 25 mm of the roots that were harvested for thevarious measurements in this study. Region 1 (0–4 mm fromthe apex, including the root cap) encompassed the region inwhich elongation rates were maintained under water stress.Region 2 (4–8 mm) exhibited maximum elongation rates inwell-watered roots, but progressive deceleration underwater stress. In region 3 (8–15 mm), elongation deceleratedin well-watered roots and was completely inhibited inwater-stressed roots. Region 4 (15–25 mm) was non-elongating in both well-watered and water-stressed roots.

In addition to within-region comparisons, the spatialgrowth analysis also allowed consideration of effects thatmay have been attributable to differences in stage ofdevelopment between treatments rather than to specificresponses to water stress (Zhu et al. 2007). Thus, responsesto water stress in region 2 were also compared to region 3 inwell-watered roots, which exhibited a comparable profileand rates of elongation. Changes in protein abundance andother responses under water stress that were consistentwhen compared to both regions 2 and 3 in well-wateredroots were considered to be independent of developmentalchanges associated with the stress-induced shortening ofthe elongation zone and, therefore, likely to be specificresponses to water stress. Because relative elongation rateswere the same in region 1 under well-watered and water-stressed conditions, all significant changes of protein abun-dance and other responses in region 1 were considered tobe responses to water stress.

The final cell length at the end of the elongation zone wasreduced by 39% in the water-stressed roots (mean valueswere 257 mm in well-watered roots, and 157 mm in water-stressed roots). Because the roots were growing understeady conditions, rates of cell flux [root elongation ratedivided by final cell length; Silk et al. (1989)] allowed anestimate of rates of cell production. This analysis indicatedthat the rate of cell production decreased by 29% underwater stress, from 11.1 to 7.9 cells h-1 in the well-wateredand water-stressed roots, respectively. Thus, although localelongation rates were maintained under water stress in theapical 4 mm (which encompassed the meristematic region),processes related to cell division were substantially inhib-ited. Nevertheless, the overall inhibition of root elongationunder water stress was more attributable to the inhibitionof cell elongation in the basal region of the growth zonethan to the inhibition of cell production.

2-DE gel analysis and identification of waterstress-responsive proteins

Total soluble proteins were analysed by 2-DE in regions 1and 2 of water-stressed (48 h after transplanting) comparedto well-watered roots (both developmental and temporalcontrols), and, as detailed above, in region 2 of water-stressed roots compared with region 3 of well-wateredroots. Figure 3 shows a representative gel image for region2 proteins from water-stressed roots; representative gelimages from all sampled regions of well-watered and water-stressed roots are shown in Supporting Information Fig. S1.Gel images of replicate samples were closely comparable inspot patterns and intensity. Approximately 1000 proteinspots were detected in each gel, and most of the spots(97–99% in the different replicate samples) were matched

Figure 3. Representative two-dimensional electrophoresis(2-DE) gel image of total soluble proteins extracted from region2 of water-stressed roots. Proteins which were differentiallyexpressed compared to both the well-watered developmental andtemporal controls are indicated, and were picked for MS analysis.



among gels from all regions and treatments. In total, 35proteins exhibited reproducible and significant changes inabundance in one or more regions of water-stressed rootscompared to both of the well-watered controls (SupportingInformation Fig. S2; spatial patterns of protein response aredescribed in more detail below).

In most cases, protein identifications were made indepen-dently for regions 1 and 2 (Table 1), and were accepted ifidentifications were reproduced in at least two of the fourreplicate samples from each region; peptide sequences,peptide masses and MS/MS spectral details are provided inSupporting Information Table S1. In total, as many as sevenreproduced identifications were obtained for proteins thatwere analysed in both the region 1 and 2 samples. Amongthe accepted protein identifications, a different proteinidentification was obtained in one of the other replicatesonly for spot 23 (which was identified as isoflavone reduc-tase homolog 2 in four region 1 and two region 2 samples).Three of the proteins that were stress responsive in bothregions 1 and 2 (spots 3, 14, 20) were identified from oneregion only because no peptides were detected in thesamples from the other region. Eight protein spots couldnot be identified either because there were no detectablepeptides (spots 2, 8, 11, 18, 27, 32) or because of inconsistentresults among replicate samples (spots 6, 29), possiblybecause of the presence of multiple proteins in one spot.

The protein identifications, together with their predictedfunctions, are shown in Table 1.Twenty-seven proteins wereidentified from the 35 stress-responsive proteins (77% iden-tification rate), 21 and six being identified from proteinand soybean EST databases, respectively. Experimentallyobserved and theoretical values of molecular mass and pIwere generally close (Table 1). In three cases, the identifica-tion of the same protein from different spots may indi-cate different isoforms or post-translational modifications:aldose reductase, spots 3 and 5; trypsin inhibitor, spots 4 and11; isoflavone reductase, spots 10 and 23. For the proteinsidentified in the soybean EST database, a homology searchwas used to find similar proteins using BLAST analysis(NCBI), and the matched proteins are shown in paren-theses after the EST identifications in Table 1. The peptidesequences of the differentially expressed proteins were alsoused for BLAST analysis to obtain gene identifications fromrecently available soybean genome sequence information(http://www.phytozome.net) (Table 1). It should be notedthat spot 13 consisted of two proteins (identified as ferritinlight chain and ferritin in region 2), which migrated sepa-rately in three of the four replicates of region 2 samples, butwere fused in all gels of region 1 samples. Accordingly, theresponse to water stress of spot 13 in region 1 was calculatedfrom the combined spot intensity of the fused spots, and isreferred to as spot ‘13L + 13U’ (lower plus upper spots).

Spatial patterns of protein responses towater stress

Figure 4 illustrates the abundance ratios of all of thewater stress-responsive proteins in regions 1 and 2 of

water-stressed compared to well-watered roots, andbetween region 2 of water-stressed roots compared toregion 3 of well-watered roots; all of the regulated proteinspots in regions 1 and 2 are labelled in Fig. 3. The numbersof up- and down-regulated proteins are summarized in Sup-porting Information Fig. S2, and mean spot intensities areprovided in Supporting Information Fig. S3. In region 1, theabundance of 23 proteins significantly increased, and oneprotein significantly decreased under water stress. In region2, the abundance of 29 proteins significantly increased, andtwo proteins significantly decreased under water stress.Twenty out of the total of 35 water stress-responsive pro-teins were common between regions 1 and 2, and in all casesthe direction (although not the magnitude) of the changesin abundance was consistent in the two regions. Four of thestress-responsive proteins, all of which showed increases inabundance, were specific to region 1, while 10 others, nine ofwhich showed increases in abundance, were specific toregion 2. Six of the proteins which showed region 2-specificresponses were not detected in the region 1 samples fromany of the treatments, including spot 29 which was detectedonly in water-stressed roots. Only three of the total of 31stress-induced changes in protein abundance in region 2 didnot exhibit similar and significant responses when region 2of water-stressed roots was compared with region 3 of well-watered roots. Accordingly, the majority of the changes inprotein abundance in region 2 were likely to be specificresponses to water stress rather than being attributable togrowth deceleration/tissue maturation.

Eighteen proteins (spots 1–10, 13–17, 19, 21, 23) exhibitedincreases in abundance in both regions 1 and 2 under waterstress, as well as the more common situation where thestress response in region 2 was similar when compared withregion 3 of well-watered roots (Figs 4 & S3). Accordingly,this category of proteins is considered to exhibit trueresponses to water stress in both regions 1 and 2. Caffeoyl-CoA O-methyltransferase (CCoAOMT, spot 20) alsoexhibited stress-induced increases in both regions 1 and 2(Fig. 4). However, although the response in region 2 wasmuch more pronounced than in region 1, this protein alsoincreased in abundance in region 3 of well-watered roots(Supporting Information Fig. S3), such that the comparisonof water-stressed region 2 with well-watered region 3 wasnot significant. The other two proteins that exhibitedresponses in region 2 which were not significant whencompared with well-watered region 3 were spot 25, putativequinone oxidoreductase, and spot 31, sg29g03.x1 (ripening-related protein), neither of which showed a significantresponse to water stress in region 1. The results suggest thatthe effects of water stress on the abundance profiles of thesethree proteins (spots 20, 25, 31) may have been largelyattributable to the developmental shift in deceleration/maturation towards the root apex rather than specificresponses to water stress.

Four proteins (spots 11, 12, 22, 24) exhibited a region1-specific increase in abundance under water stress (Fig. 4).Among them, chalcone synthase 7 (spot 22) also showeda greater abundance in region 1 than in region 2 in



Tab

le1.

Iden

titi

esof

wat

erst

ress

-res

pons

ive

prot

eins

inth

eel

onga

tion

zone

ofth

eso

ybea

npr

imar

yro

ot

Spot

no.

Pro

tein

orE

STid

enti

ficat

ion

GI

num

ber

Exp

erim

enta

lm

ass/

pIT

heor

etic

alm

ass/

pIM

asco

tsc

ore

Org

anis

mm

atch

edId

enti

fied

from

regi

on(s

)P

redi

cted

func

tion

Gen

eid

enti

ficat

ion

inP

hyto

zom

e

1C

yste

ine

prot

eina

sein

hibi

tor

1944

319

26.5

/6.6

27.6

/7.3

223

Gly

cine

max

R1,

R2

End

opep

tida

sein

hibi

tor

Gly

ma1

5g36

180.

13

Ald

ose

redu

ctas

e13

1603

9935

.3/6

.336

.5/5

.699

Dig

italis

purp

urea

R1

Ald

ehyd

ere

duct

ion

Gly

ma0

1g25

000.

14

Tryp

sin

inhi

bito

r93

6704

220

.3/5

.118

.0/6

.140

7G

.max

R1,

R2

End

opep

tida

sein

hibi

tor

Gly

ma0

9g28

310.

15

Ald

ose

redu

ctas

e13

1603

9935

.9/6

.536

.5/5

.613

2D

.pur

pure

aR

1,R

2A

ldeh

yde

redu

ctio

nG

lym

a01g

2500

0.1

7Fe

rrit

in-4

,chl

orop

last

prec

urso

r29

8393

8625

.7/5

.623

.6/5

.137

1G

.max

R1,

R2

Iron

hom

eost

asis

Gly

ma1

4g06

160.

19

Glu

tath

ione

pero

xida

se92

8942

2320

.0/6

.218

.3/6

.618

6M

edic

ago

trun

catu

laR

1,R

2D

etox

ifica

tion

ofR

OS

Gly

ma0

8g01

700.

110

Isofl

avon

ere

duct

ase

hom

olog

265

7317

136

.5/6

.033

.9/5

.669

2G

.max

R1,

R2

Isofl

avon

oid

bios

ynth

esis

Gly

ma0

4g01

380.

111

Tryp

sin

inhi

bito

r93

6704

220

.6/5

.018

.0/6

.136

0G

.max

R1,

R2

End

opep

tida

sein

hibi

tor

Gly

ma0

9g28

310.

113

LL

ower

spot

:fer

riti

nlig

htch

ain

1700

7825

.0/5

.323

.0/5

.249

3G

.max

R2

Iron

hom

eost

asis

Gly

ma1

8g43

650.

113

UU

pper

spot

:fer

riti

n96

8987

25.1

/5.3

23.0

/5.3

430

G.m

axR

2Ir

onho

meo

stas

isG

lym

a07g

1906

0.1

14sd

58e0

8.y1

(In2

-1pr

otei

n)60

7128

627

.1/5

.5N

D16

4G

.max

R1

Det

hiol

atio

nof

glut

athi

onat

edpr

otei

nN

A15

In2-

1pr

otei

n11

3855

7925

.3/5

.127

.0/5

.243

6G

.max

R1,

R2

Det

hiol

atio

nof

glut

athi

onat

edpr

otei

nG

lym

a13g

1983

0.1

16sc

32f0

8.y1

(Dis

ease

resi

stan

ce-r

espo

nsiv

efa

mily

prot

ein)

5509

112

19.3

/5.7

ND

135

G.m

axR

1,R

2U

nkno

wn

NA

17C

halc

one

redu

ctas

e45

8657

033

.4/6

.5N

D96

Cic

erar

ietin

umR

1,R

2Is

oflav

onoi

dbi

osyn

thes

isG

lym

a18g

5225

0.1

19G

luta

thio

neS-

tran

sfer

ase

811

3854

3126

.6/5

.825

.9/5

.736

2G

.max

R1,

R2

Det

oxifi

cati

onof

reac

tive

com

poun

dsG

lym

a07g

1691

0.1

20C

affe

oyl-

CoA

O-m

ethy

ltra

nsfe

rase

1934

859

29.1

/5.5

27.9

/5.0

180

Euc

alyp

tus

gunn

iiR

2L

igni

nbi

osyn

thes

isG

lym

a02g

1153

0.1

21D

ehyd

roas

corb

ate

redu

ctas

e28

1924

2725

.0/5

.923

.6/7

.712

7N

icot

iana

taba

cum

R1,

R2

Asc

orba

tere

cycl

ing

Gly

ma2

0g38

440.

122

Cha

lcon

esy

ntha

se7

2317

9942

.5/6

.642

.9/6

.012

3G

.max

R1

Isofl

avon

oid

bios

ynth

esis

Gly

ma0

1g43

880.

123

Isofl

avon

ere

duct

ase

hom

olog

265

7317

136

.6/5

.833

.9/5

.651

8G

.max

R1,

R2

Isofl

avon

oid

bios

ynth

esis

Gly

ma0

4g01

380.

124

Cyt

osol

icph

osph

oglu

com

utas

e62

7228

178

.9/5

.563

.3/5

.525

5P

isum

sativ

umR

1,R

2G

lyco

lysi

sG

lym

a05g

3479

0.1

25P

utat

ive

quin

one

oxid

ored

ucta

se21

0686

6424

.3/6

.521

.7/6

.518

2C

.ari

etin

umR

2R

educ

tion

ofce

rtai

nqu

inon

esG

lym

a08g

0657

0.1

26si

51d0

8.y1

(Qui

none

oxid

ored

ucta

se)

7146

301

37.7

/6.4

ND

273

G.m

axR

1,R

2R

educ

tion

ofce

rtai

nqu

inon

esG

lym

a16g

0804

0.1

28sa

34f0

1.y1

(Euk

aryo

tic

tran

slat

ion

init

iati

onfa

ctor

5A)

4289

908

19.4

/5.9

ND

323

G.m

axR

1,R

2m

RN

Ash

uttl

ing

for

cell

cycl

eG

lym

a04g

3295

0.1

30P

utat

ive

amin

otra

nsfe

rase

5235

3685

48.8

/6.4

42.2

/6.5

117

Ory

zasa

tiva

R2

Am

ino

acid

met

abol

ism

Gly

ma0

1g32

090.

131

sg29

g03.

x1(R

ipen

ing-

rela

ted

prot

ein)

6726

549

17.4

/6.3

ND

189

G.m

axR

2U

nkno

wn

NA

33gm

rtD

rNS0

139

-CM

13R

E03

023.

s4W

ater

stre

ssed

48h

segm

ent

2(p

utat

ive

C2

dom

ain-

cont

aini

ngpr

otei

n)

5802

1988

41.6

/4.9

ND

105

G.m

axR

2Si

gnal

tran

sduc

tion

Gly

ma0

9g24

890.

1

34P

lasm

am

embr

ane

intr

insi

cpo

lype

ptid

e64

6912

131

.7/5

.123

.3/5

.077

C.a

riet

inum

R2

Unk

now

nG

lym

a09g

0523

0.1

Pro

tein

spot

sw

ere

iden

tifie

dby

mat

rix-

assi

sted

lase

rde

sorp

tion

/ioni

zati

on-t

ime-

of-fl

ight

tand

emm

ass

spec

trom

etry

(MA

LD

I-T

OF

MS/

MS)

anal

ysis

;pep

tide

sequ

ence

sar

elis

ted

inSu

ppor

ting

Info

rmat

ion

Tabl

eS1

.Id

enti

ficat

ion

and

acce

ssio

nnu

mbe

rs(p

rote

inG

Inu

mbe

r)ar

efr

omth

eN

atio

nalC

ente

rfo

rB

iote

chno

logy

Info

rmat

ion

(NC

BI)

data

base

.For

prot

eins

wit

hout

anno

tati

onin

the

soyb

ean

data

base

,pro

babl

ean

nota

tion

sob

tain

edfr

omba

sic

loca

lalig

nmen

tse

arch

tool

(BL

AST

)an

alys

isar

esh

own

inpa

rent

hese

s.E

xper

imen

tala

ndth

eore

tica

lmol

ecul

arm

ass

are

show

nin

kDa.

Cor

resp

ondi

ngso

ybea

nge

nes

wer

ede

term

ined

byB

LA

STan

alys

isfr

omth

eP

hyto

zom

eso

ybea

nda

taba

se(h

ttp:

//ww

w.p

hyto

zom

e.ne

t).N

D,n

otde

term

ined

;the

oret

ical

mas

san

dpI

are

notc

ompu

ted

beca

use

the

enti

rese

quen

ceis

nota

vaila

ble

inth

eda

taba

se.N

A,n

otav

aila

ble;

Phy

tozo

me

gene

iden

tific

atio

nis

not

avai

labl

e,po

ssib

lybe

caus

ean

nota

tion

ofth

eso

ybea

nge

nest

ruct

ures

isin

com

plet

e.



0

1

2

3

4

5

6

7

1. C

yste

ine

prote

inas

e inhib

itor 2

3. A

ldos

e re

ductas

e

4. T

ryps

in in

hibito

r

5. A

ldos

e re

ductas

e 6

7. F

errit

in-4

, chlor

oplast p

recu

rsor 8

9. G

luta

thione

per

oxid

ase

10. I

sofla

vone

redu

ctas

e

11. T

ryps

in in

hibito

r12

13L. F

errit

in ligh

t cha

in

13U. F

errit

in

14. In2-

1 pr

otein

15. In2-

1 pr

otein

16. D

iseas

e resistanc

e pro

tein

17. C

halco

ne re

ductas

e18

19. G

luta

thione

S-tr

ansfe

rase

8

20. C

affeoy

l-CoA O

-met

hyltr

ansfer

ase

21. D

ehyd

roas

corb

ate

redu

ctas

e

22. C

halco

ne sy

nthas

e 7

23. I

sofla

vone

redu

ctas

e

24. C

ytoso

lic p

hosp

hogluc

omuta

se

25. Q

uino

ne o

xidor

educ

tase

26. Q

uino

ne o

xidor

educ

tase 27

28. T

rans

latio

n in

itiat

ion

factor

5A

29

30. A

minot

ransf

erase

31. R

ipen

ing-

relate

d pro

tein 32

33. C

2 do

main

-con

taining

protein

34. P

lasm

a mem

bran

e intri

nsic p

olyp

eptid

e

Pro

tein

ab

un

da

nce

ra

tio

R2WS R2WW

R2WS R3WW

**

** **

**

**

**

** ****

****

**

****

**

**

** **

*

****

*

**

**

**

**

*

*

***

**

*

*

0

1

2

3

4

5

6

7

1. C

yste

ine

prote

inas

e inhib

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Spot number and protein identif ication

Region 1

Region 2 Detected

only in WS

R1WS R1WW

Figure 4. The abundance ratio of differentially expressed proteins in region 1 (R1, upper panel) and region 2 (R2, lower panel) ofwater-stressed (WS) compared to well-watered (WW) roots. Because of the use of well-watered developmental (WWD) and temporal(WWT) controls (see Fig. 1), two abundance ratios (WS/WWD, WS/WWT) were calculated for each protein in each region, and the averagewas used to describe the level of regulation under water stress (WS/WW). The protein abundance ratio in water-stressed region 2compared with well-watered region 3 (R3) is also shown in the lower panel. Changes in protein abundance which were significant in boththe R2WS/R2WW and R2WS/R3WW comparisons are considered to be independent of developmental changes associated with thestress-induced shortening of the elongation zone (see text for details of this analysis). Values are means � SE of analyses from three tofour sets of gels from independent experiments. Asterisks denote differentially expressed proteins for which spot intensities weresignificantly different between the water-stressed and well-watered treatments (P < 0.05; mean spot intensities are provided in SupportingInformation Fig. S3). Spots 13 lower (L) and upper (U) were fused in all gels of region 1 samples. Spots 29–34 were not detected (ND) inregion 1 in any treatment. Spot 29 was detected only in region 2 of water-stressed roots (note that the scale of protein abundance ratiodoes not apply for this spot). Unidentified proteins are shown by spot number only.



well-watered roots (Supporting Information Fig. S3). Fiveproteins (spots 29–33) were not detected in any treatmentin region 1, and exhibited increases in abundance in region2 that were also significant when compared to well-wateredregion 3 (Fig. 4). Accordingly, this category of proteins isconsidered to exhibit true responses to water stress specifi-cally in region 2.

Only one protein, sa34f01.y1 (eukaryotic translation ini-tiation factor 5A, spot 28), was down-regulated under waterstress in both regions 1 and 2. The other down-regulatedprotein was plasma membrane intrinsic polypeptide (spot34), which was not detected in any treatment in region 1,and down-regulated in region 2 under water stress. Both ofthese proteins had similar abundances in regions 2 and 3 ofwell-watered roots (Supporting Information Fig. S3), andthus the decreases in abundance in region 2 under waterstress were also significant when compared to well-wateredregion 3. Accordingly, the down-regulation of both proteinsis considered to represent a true response to water stress inboth regions 1 and 2.

Isoflavonoid biosynthesis

Several proteins that exhibited increases in abundanceunder water stress are enzymes involved in isoflavonoidbiosynthesis (regions 1 and 2, two isoflavone reductasehomologs, and chalcone reductase; region 1 only, chalconesynthase 7) (Fig. 4). Chalcone synthase 7 has been reportedto play a critical role in isoflavonoid synthesis in soybeanseeds (Dhaubhadel et al. 2007).These results suggested thatisoflavonoid biosynthesis may have increased in the rootelongation zone under water stress, perhaps especially inregion 1. To test this prediction, the isoflavone compositionwas quantified in regions 1–3 of well-watered and water-stressed roots. In well-watered roots (developmental andtemporal controls), total isoflavone content was highest inregion 1 and decreased with increasing distance from theapex (Fig. 5). In the water-stressed roots, the isoflavonecontent doubled in region 1 compared to the well-wateredvalue, and then decreased to a similar level to that of well-watered roots in region 2. The isoflavone content was alsosignificantly higher under water stress in region 3, althoughthe levels were relatively low in this region. The effect ofwater stress on the composition of isoflavones and theirderivatives in each region is shown in Table 2. Glucosyldaidzein was the major component in all regions of alltreatments. Region 1 of water-stressed roots had the highestlevel of every isoflavone component, especially in glucosyldaidzein and malonyl glucosyl daidzein. Accumulation ofisoflavonoids/flavonoids in the apical region under waterstress was also examined by staining with DPBA (Fig. 5inset).The level of DPBA fluorescence was enhanced in theapical 2.5 mm of water-stressed roots, in agreement with thequantitative measurements of isoflavone content.

Lignin accumulation

CCoAOMT (spot 20), which is involved in lignin biosynthe-sis, exhibited one of the largest increases in abundance in

region 2 of water-stressed roots (Fig. 4), and also exhibiteda large increase in abundance between regions 2 and 3 inwell-watered roots (Supporting Information Fig. S3). Theseresults suggested that water stress may have enhanced thedegree of lignification in the basal region of the elongationzone. This prediction was tested by histochemical localiza-tion of lignin staining patterns in regions 1–4 (Fig. 6).Lignification was examined by autofluorescence of wallphenolics (Fig. 6a, c, d; blue autofluorescence is emittedfrom lignin and other phenolic compounds under UV exci-tation) and Mäule staining (Fig. 6b); the two methodsshowed similar patterns among the regions and treatments.In well-watered roots, weak fluorescence and Mäule stain-ing were detected in the epidermis in region 3, and in boththe xylem and epidermis in region 4. Under water stress,the onset of lignification shifted apically; lignification wasdetected in the epidermis and xylem in region 2 and wasenhanced in both tissues in regions 3 and 4 compared withwell-watered roots. The results support the suggestionfrom the kinematic growth analysis that the large increasein CCoAOMT abundance in region 2 under water stress(Fig. 4) was partly attributable to the maturation of tissuecloser to the root apex, although comparisons of regions 2and 3 in water-stressed roots with regions 3 and 4 in well-watered roots show that lignification was much greaterunder water stress (Fig. 6), suggesting that lignin synthesiswas also specifically enhanced in response to water stress inthe basal region of the elongation zone.

0

3

6

9

12

15

WWT

WWD

WS

Region 1 Region 2 Region 3

*

*

Tota

l is

oflavone c

onte

nt (n

mol m

g–

1 F

W)

WWT WS

Figure 5. Total isoflavone content in regions 1–3 ofwater-stressed (WS) and well-watered developmental (WWD)and temporal (WWT) control roots. Values are means � SE ofthree samples from independent experiments. Asterisks denotesignificant differences between water-stressed values whencompared to both well-watered controls (Tukey–Kramer test,P < 0.05). The isoflavone composition of the same samples isshown in Table 2. FW, Fresh weight. Inset: representative imagesshowing diphenyl boric acid-2-aminoethyl ester (DPBA) stainingfor isoflavonoids/flavonoids in the apical region of water-stressedand well-watered temporal control roots. The bar indicates 1 mm.



The Mäule staining indicated that the type of lignin wasdifferent between the xylem and epidermis (Fig. 6b). TheMäule reagent stains guaiacyl–syringyl lignin or guaiacyllignin red-purple or brown, respectively (Higuchi 1998). Itshould also be noted that suberin accumulation might havecontributed to the increase of autofluorescence under waterstress (Fig. 6a), because CCoAOMT may also participate insuberin biosynthesis (Bernards & Lewis 1998).

Ferritin and iron localization in the rootelongation zone

Ferritin proteins consist of 24 homologous or heterogonoussubunits that form a cavity that sequesters iron. In regions 1and 2 of the root elongation zone, two or three ferritinsubunits exhibited increases in abundance under waterstress (Fig. 4). Therefore, it was hypothesized that theincrease in ferritin abundance served to tightly control andprevent free iron levels from increasing in the root elonga-tion zone under water stress, probably to limit the genera-tion of hydroxyl radicals via the Fenton reaction (seeDiscussion). To test this hypothesis, we studied the effect ofwater stress on free and protein-bound iron levels.

Perls’s iron staining was used to detect free iron; thismethod preferentially stains free (or loosely chelated) ironrather than iron packed in ferritin (Richter 1978; Harrison& Arosio 1996; Ghio, Churg & Roggli 2004). Figure 7 illus-trates Perls’s iron staining in the apical 5 mm of the roots. Inboth water-stressed and well-watered roots, blue ferric ironstaining was detected only in the apical millimetre; stainingappeared slightly higher in water-stressed than in well-watered roots (Fig. 7a). Squashed root images (Fig. 7b) sug-gested that the slight increase in free iron under water stressoccurred primarily in the most apical region of the stele,which was confirmed in transverse (Fig. 7c) and longitudi-nal (Fig. 7d) sections. Potentially, the slight increase in freeiron level represented excess iron that was not capturedby ferritin.

Because staining for free iron was not present as the cellswere displaced away from the root apex, it was postulatedthat the increase in ferritin abundance under water stressserved to sequester more iron in this region. Native PAGEprotein separation followed by iron staining was conductedto determine the amount of iron captured in the ferritincore (Fig. 8). For this analysis, region 1 was subdivided intotwo segments from 0 to 1 mm and 1 to 4 mm from the apexbecause of the slight increase in free iron observed in theapical millimetre (Fig. 7). Protein-bound iron was detectedusing two staining methods, Ferene S and DAB. Ferene Sforms a blue complex by binding iron stoichiometrically,whereas the DAB method uses the catalytic activity of ironto oxidize DAB and form an insoluble brown pigment inthe presence of H2O2.

Figure 8 shows iron staining with Ferene S and DAB,and immunodetection of ferritins in regions 1 and 2 ofwell-watered and water-stressed roots. Ferene S and DABreagent, and the anti-ferritin antibody detected a singleband at 530 kD (Fig. 8a), indicating that ferritin was theTa

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prominent iron-binding protein in the root elongationzone. The molecular mass of ferritin was in good agreementwith previous studies (Laulhere et al. 1988). DAB stainingwas not decreased when azide was added to the stainingsolution to inhibit peroxidase activity (peroxidase can alsooxidize DAB), indicating that peroxidase activity did notinfluence the results (data not shown). Consistent with theproteomic results (Fig. 4), ferritin protein level was higherin the water-stressed roots in all regions (Fig. 8c). Iron stain-ing with DAB was more sensitive than with Ferene S, butboth methods revealed that the amount of bound ironincreased with increasing distance from the apex in bothwell-watered and water-stressed roots (Fig. 8a,b). In all

regions, however, the levels of bound iron were substan-tially higher under water stress, demonstrating that ferritinproteins sequestered more iron as the cells were displacedthrough the elongation zone of water-stressed compared towell-watered roots.

DISCUSSION

Functional classification of the waterstress-responsive proteins

Most of the water stress-responsive proteins that were iden-tified in the elongation zone of the soybean primary root

Figure 6. Autofluorescence of cell wallphenolics under UV excitation (a) andlignin staining (Mäule stain, b) intransverse sections from the center ofregions 1–4 of water-stressed (WS) andwell-watered temporal control (WWT)roots. Lignins are stained red-purple(guaiacyl–syringyl lignin) or brown(guaiacyl lignin). The bars in both panelsindicate 0.1 mm. The experiments wererepeated with similar results. Relativefluorescence intensity in the xylem andepidermis from the autofluorescenceimages is shown in (c) and (d),respectively; values are means � SE ofthree roots from one experiment.Asterisks denote significant differencesbetween water-stressed values whencompared to the well-watered control(t-test, P < 0.05).

0

10

20

30

0 5 10 15 20

Region 1 Region 2 Region 3 Region 4

WS

WWT

WS

WWT

0

6

12

18

0 5 10 15 20

(c) Fluorescence in xylem

(d) Fluorescence in epidermis

WS

WWT

WS

WWT

Re

lative

in

ten

sity

Re

lative

in

ten

sity

Distance from apex (mm)

Distance from apex (mm)

(a) Wall phenolics autofluorescence

(b) Lignin stain

Region 1 Region 2 Region 3 Region 4

*

*

*

*

*

*

Figure 7. Localization of ferric iron inthe apical 5 mm region of water-stressed(WS) and well-watered developmental(WWD) and temporal (WWT) controlroots. Ferric iron was visualized as a bluecomplex using the Perls’s stainingmethod, and the stained roots wereobserved under a stereoscope (a–c) orinverted microscope (d). (a) Intact rootapices, showing that staining was localizedto the apical millimetre. The bar indicates0.5 mm. (b) Stained roots in panel a weresquashed to expose the inside of theroots. (c) Sequential transverse sections0.15 mm in thickness from the stainedregion at the root apex. The diagramindicates the positions where each sectionwas made. (d) Longitudinal section(0.15 mm thickness) of stained roots. Thebar indicates 0.1 mm.

(a) (b)

(c) (d)



can be postulated to have roles in water stress tolerancemechanisms. The majority of the proteins can be classifiedinto four functional categories: control of reactive oxygenspecies (ROS) metabolism, isoflavonoid biosynthesis,control of apoptosis-like cell death, and control of proteindegradation. These four overall functions are closely inter-related in their potential roles in stress tolerance, and aschematic representation of their possible synergistic inter-actions is shown in Fig. 9. Most of the proteins assigned tothese categories exhibited increases in abundance in bothregions 1 and 2, indicating that these overall functions are ofgeneral importance in stress adaptation rather than specifi-cally associated with growth maintenance in region 1 orgrowth inhibition in region 2. The largest of the categorieswas the control of ROS, which included 14 regulated pro-teins, indicating that this function is probably of particularimportance in adaptation of the root elongation zone towater stress conditions. Abiotic stresses including waterstress often result in increased ROS production, which maycause damage including oxidation of proteins (Apel & Hirt2004; Moller, Jensen & Hansson 2007; Cruz de Carvalho2008; Bian & Jiang 2009). Several lines of evidence alsoshow that increased ROS can trigger apoptosis-like celldeath in plants (Solomon et al. 1999; Gechev et al. 2006).Isoflavonoids have several functions in plant cells, includingserving as ROS scavengers (Nerya et al. 2004; Kruk et al.2005). Accordingly, several enzymes for isoflavonoid bio-synthesis that exhibited increases in abundance in regions 1and 2 are included in Fig. 9 as a subgroup in the category of

control of ROS; other potential functions of isoflavonoidsare considered below.

Isoflavonoid and lignin accumulation patterns,and region-specific regulation ofphenylpropanoid metabolism

The apical localization of isoflavones in roots under well-watered conditions (Fig. 5) is consistent with previousstudies of the soybean primary root (Graham 1991). Inaddition, in transgenic tobacco plants expressing GUSunder the promoter of bean chalcone synthase 8, GUSactivity was detected at the root meristem (Schmid et al.1990), consistent with the protein expression pattern ofchalcone synthase 7 in well-watered roots (SupportingInformation Fig. S3). To our knowledge, however, the dra-matic increase of isoflavone content in region 1 under waterstress (Fig. 5) is the first report of an increase in isoflavonoidaccumulation in the root elongation zone in response towater stress. In leaves of hawthorn and Cistus clusii, fla-vonoids, compounds with similar structure to isoflavones,were increased under water stress, and their potential func-tion against oxidative damage was discussed (Kirakosyanet al. 2003; Hernández, Alegre & Munné-Bosch 2004).

The spatial pattern of response of proteins involved inthe phenylpropanoid biosynthetic pathway (Fig. 4),together with the differential accumulation profiles ofisoflavonoids and lignin (Figs 5 & 6), suggests that metabo-lite flow in this pathway may respond differently to water

0–1 mm 4–8 mm

WS WWT WS WWT WS WWT

1–4 mm

Region 1 Region 2

669-

kD

440-

669-440-

669-440-

Iron stain, Ferene S

Iron stain, DAB

Ferritin, immunoblot

0

10

20

30

Rela

tive inte

nsity

(b) Iron stain, DAB

WWT

0

2

4

6

8

0–1 mm 1–4 mm 4–8 mm

WWTWS WS WWTWS

Rela

tive inte

nsity

(c) Ferritin, immunoblot

7.0

7.1

3.1

4.3

4.4

3.8

2.0

1.9

5.9

5.0

3.3

2.1

(a)

*

*

*

Figure 8. Evidence for the accumulation of ferritin-bound iron in the growth zone of water-stressed (WS) compared to well-watered(temporal control, WWT) roots. Total soluble proteins were extracted from root segments and separated on native PAGE gels.(a) Detection of protein-bound iron in root segments from 0 to 1 mm, 1–4 mm and 4–8 mm from the apex of water-stressed andwell-watered roots; 70 mg of proteins was loaded in each lane. Note that region 1 was divided into two segments because of thelocalization of Perls’s iron staining in the apical millimetre (see Fig. 7). The gels were stained with Ferene S and DAB for iron, andthe ferritin band was detected by Western blot. The position of ferritin (530 kD) is indicated by the arrows on the left side of the gels.(b) Relative intensity of the ferritin-bound iron band in the DAB-stained gels. Values are means � SE of three independent experiments.Asterisks denote significant differences between water-stressed values when compared to the well-watered control (t-test, P < 0.05).(c) Relative intensity of native ferritins in the immunoblot analysis. The bars represent the means of two independent experiments; thenumbers above the bars are the values in each of the two experiments.



stress in regions 1 and 2. Chalcone reductase, with co-actionof chalcone synthase, produces trihydroxychalcone. Impor-tantly, this is a precursor only for the biosynthesis of isofla-vonoids, whereas tetrahydroxychalcone, which is producedwithout the action of chalcone reductase, is involved inboth isoflavonoid and flavonoid metabolic pathways (Yu &McGonigle 2005). Accordingly, the increase in abundanceof chalcone synthase specifically in region 1 of water-stressed roots (Fig. 4), together with the increase in chal-cone reductase in both regions, was likely to have increasedmetabolite flow into the isoflavonoid branch pathwayand could have been a determining factor for the majoraccumulation of isoflavones in region 1 under waterstress. Isoflavone reductase catalyses the reduction of2′-hydroxydaidzein to form 7,2′-dihydrodaidzein, one of thesteps of isoflavonoid biosynthesis, and the up-regulation oftwo proteins for this enzyme in both regions 1 and 2 pro-vides further support for enhanced isoflavonoid biosynthe-sis in the root elongation zone under water stress.

The shift in initial development of lignification towardsthe apex of the soybean primary root under water stress(Fig. 6) is similar to the response previously reported inmaize primary roots by Fan et al. (2006). CCoAOMT cataly-ses the fourth step in lignin biosynthesis from p-coumarate-CoA (Boerjan, Ralph & Baucher 2003), and the majorincrease in abundance of this enzyme in region 2 of water-stressed roots supports the conclusion that lignin synthesiswas enhanced in response to water stress in the basal regionof the elongation zone. In water-stressed maize leaves, simi-larly, both the protein level of this enzyme and lignification

were reported to shift towards the base (i.e. the origin) ofthe elongation zone, in association with a shortened elon-gation zone (Vincent et al. 2005).

Importantly, the metabolic pathways of isoflavonoid andlignin biosynthesis share the same precursor, p-coumarate-CoA (Yu & McGonigle 2005). Therefore, up-regulation ofCCoAOMT activity could increase the metabolite flow tolignin synthesis, and limit substrate availability for isofla-vonoid synthesis. Accordingly, the large increase in abun-dance of CCoAOMT in region 2 of water-stressed rootsmay have contributed to the lack of isoflavone accumula-tion in this region (Fig. 5). Figure 10 illustrates the possibledifferences in phenylpropanoid metabolite flux pathwaysbetween regions 1 and 2 under water stress. A recent studyof Arabidopsis (Besseau et al. 2007) supports the concept ofsubstrate competition between lignin synthesis and (iso)fla-vonoid synthesis. It was shown that silencing of an enzymefor lignin biosynthesis, hydroxycinnamoyl-CoA shikimate/quinate hydroxycinnamoyl transferase, induced the accu-mulation of flavonoids in the leaves because of increasedmetabolite flow into flavonoid synthesis [Arabidopsis doesnot have the isoflavonoid synthesis pathway (Yu &McGonigle 2005)].

Possible functions of isoflavonoids and ligninin root growth regulation under water stress

As mentioned earlier, isoflavonoids and also chalcone arepotent antioxidants (Nerya et al. 2004; Kruk et al. 2005), andthus may play a role as scavengers of ROS in the root

Figure 9. Schematic representation ofthe potential roles of water stress-regulated proteins in water stresstolerance mechanisms in the root growthzone. Stress-responsive proteins fromboth regions 1 and 2 are included. Notethat cysteine proteinase inhibitor andglutathione peroxidase could be involvedin two of the overall functional categories[control of apoptosis-like cell death andprotein degradation for cysteineproteinase inhibitor (Solomon et al. 1999;Belenghi et al. 2003), control of reactiveoxygen species (ROS) and control ofapoptosis-like cell death for glutathioneperoxidase (Chen et al. 2004)].

Water stress ROS≠ Apoptosis-like cell death

Control of ROS

Aldose reductase (2)

Ferritin subunits (3)

Glutathione peroxidase

Glutathione transferase 8

Dehydroascorbate reductase

Quinone oxidoreductase (2)

Isoflavone biosynthesis

Isoflavone reductase (2)

Chalcone reductase

Chalcone synthase

Oxidation of proteinShortened lifetime of

proteins

Control of protein degradation

Cysteine proteinase inhibitor

Trypsin inhibitor (2)

In2-1 protein (2)

Control of apoptosis-like

cell death

Cysteine proteinase inhibitor

Glutathione peroxidase



elongation zone under water stress. There is also evidencethat isoflavones can regulate auxin transport in a mannersimilar to flavonoids (Subramanian, Stacey & Yu 2006).Accordingly, the accumulation of isoflavones in the apicalregion of the elongation zone in water-stressed roots mighthave influenced the level of auxin, which in turn could playa role in regulating the growth response. This is an intrigu-ing possibility, because water stress was reported to increasethe auxin content in the elongation zone of maize primaryroots (Ribaut & Pilet 1994), and applied auxin at certainconcentrations has been shown to cause shortening of theelongation zone in primary roots of wheat (Hejnovicz1961), timothy (Goodwin 1972) and maize (Ishikawa &Evans 1993) in a manner similar to the effect of water stressin maize (Sharp et al. 1988) and soybean (Fig. 2). However,the role(s) of auxin in regulating primary root growthresponses to water stress has yet to be determined in anyspecies. Interestingly, CCoAOMT (spot 20) and quinonereductase (spot 25), which showed similar patterns ofresponse to water stress in the root elongation zone (Fig. 4),are both known to be induced by auxin (Bianchi et al.2002a; Laskowski et al. 2002), and thus their responsesmight also reflect an increase in auxin levels.

Lignin confers mechanical strength and stiffness to cellwalls (Jones, Ennos & Turner 2001). Accordingly, theincrease in lignification in regions 2 and 3 of the elongationzone in water-stressed roots (Fig. 6) may have been associ-ated with decreased cell wall extensibility in this region,which has been shown to occur in the equivalent region ofthe maize primary root (Wu et al. 1996; Fan et al. 2006). Theincreased lignification of the epidermis and xylem may alsorestrict water loss from the root, and facilitate longitudinalwater transport to the elongating tissues. It is also interest-ing to note the relationship between lignification andoxidative stress; lignification and secondary cell wall depo-sition are induced in response to ozone and hydrogenperoxide treatments (Potikha et al. 1999; Cabane et al.2004). As shown in Fig. 9, the likely function of many of thewater stress-responsive proteins in the control of ROS

suggests that ROS generation might have been increasedunder water stress, which in turn could have triggered theonset of lignification closer to the root apex.

Ferritin and iron localization in the rootelongation zone

Ferritin sequesters intracellular iron (approximately 4500atoms of Fe per ferritin molecule), and thereby can limitthe generation of hydroxyl radicals (•OH) via the Fentonreaction (Fe2+ + H2O2→Fe3+ + OH- + •OH) (Deak et al.1999; Briat et al. 2009; Ravet et al. 2009). The control of freeiron in plant cells is also important because Fe3+ resultingfrom the Fenton reaction can be reduced back to Fe2+ byO2- radicals or by ascorbate, thus sustaining the reaction.The increased abundance of three ferritin subunits (spots 7,13L, 13U) under water stress indicates that regulation offree iron levels is an important component of adaptation tolow water potentials in the elongation zone of the soybeanprimary root (Figs 4 & 9). Ferritin has also been reported toaccumulate in response to water stress in the primary rootof maize (discussed further below; Spollen et al. 2008), andin leaves of several species including maize (Riccardi et al.1998) and chickpea (Bhushan et al. 2007).

Taken together, the patterns of localization of free andprotein-bound iron (Figs 7 & 8) support the conclusion thatthe increase in ferritin abundance under water stress effec-tively minimized free iron levels in the root elongationzone. The increase in bound iron under water stress indi-cates that increased ferritin levels were required because ofan apparently greater total amount of iron in the elongationzone than in well-watered roots. It is possible that iron thatwas transported to the root tip in the phloem accumulatedin the elongation zone because of the decrease in overallroot elongation rate (i.e. the rate of iron dilution by volumeexpansion was decreased). Interestingly, a study of wheatalso reported that the total iron content of the roots wasincreased during water stress (Price & Hendry 1991).

Figure 8 shows that the ferritin protein level detected byimmunoblot was higher in water-stressed than well-wateredroots in both regions 1 and 2 of the elongation zone.Accordingly, the increase in protein level could partlyaccount for the increase in bound iron in the water-stressedroots. However, whereas the amount of ferritin was as highor higher in the apical millimetre than at greater distancesfrom the apex in both treatments, the amount of bound ironwas least in this region. Thus, the small increase in free ironthat was observed in the apical region of the water-stressedroots occurred despite the presence of ferritin protein, pos-sibly indicating differential compartmentation within theroot tissues.Whether this localized increase in free iron wasof physiological significance for the root response to waterstress is unknown.

Comparison between soybean and maizeprimary root responses to water stress

Because of post-transcriptional regulation and turnover ofproteins, the total soluble protein analysis of the soybean

Figure 10. Hypothesis for differential patterns of metaboliteflux within the phenylpropanoid pathway in regions 1 (R1) and 2(R2) of water-stressed roots. Relative magnitudes ofstress-induced increases in protein abundance between theregions are indicated by the upward arrows. The thickness of theblock arrows in each branch pathway indicates relative rates ofmetabolite flow.



primary root reported in this study is not directly compa-rable to the recent transcriptomic study of the maizeprimary root (Spollen et al. 2008). However, the cultureconditions for seedling growth were similar (although notidentical; see Methods), and the water stress treatmentwas the same in the two experimental systems. Moreover,the spatial patterns of the growth response to water stresswithin the elongation zone were very similar in the twospecies (Fig. 2; Sharp et al. 1988). Therefore, it was of inter-est to compare and contrast the responses to water stress ofthe proteins identified in this study with the stress-responsive maize genes identified by Spollen et al. (2008).

As listed in Supporting Information Table S2, severalof the water-stress responsive proteins in soybean havesimilar functions to the regulated genes in maize, whileother responses appeared to differ between the twosystems. In the maize root, proteinase inhibitors, metal-binding proteins (including ferritin), glutathione trans-ferase and O-methyltransferase were up-regulated underwater stress, and their expression profiles between regions 1and 2 were generally similar to the responses of functionallysimilar proteins in soybean. Maize does not have theenzymes for isoflavonoid synthesis that were up-regulatedin soybean, but flavanone 3-hydroxylase, an enzyme forflavonoid synthesis, was up-regulated in region 1, and chal-cone isomerase was up-regulated in region 2. Alanine glut-arate aminotransferase and a putative aminotransferasewere highly up-regulated in region 2 under water stress inmaize and soybean roots, respectively, indicating activeamino acid metabolism.

In contrast, several distinct responses are also evident.For example, aldo/keto-reductase was down-regulatedunder water stress in region 2 of maize roots, whereassoybean aldose reductases were highly up-regulated in bothregions 1 and 2. Glutathione peroxidase, dehydroascorbatereductase and quinone oxidoreductases were not regulatedin water-stressed maize roots, in contrast to the results insoybean. In addition, sa34f01.y1 (eukaryotic translation ini-tiation factor 5A) was down-regulated in both regions 1 and2 in soybean, but was not regulated in maize. Dresselhaus,Cordts & Lorz (1999) showed that eukaryotic translationinitiation factor 5A was expressed during the G1 phaseof cell division using partially synchronized suspension-cultured cells of rice. Accordingly, the down-regulationof sa34f01.y1 in water-stressed soybean roots was possiblyrelated to the apparent decrease in cell division rate inthis system.

It is noteworthy that cysteine proteinase inhibitors wereup-regulated in region 1 in both the soybean and maizestudies. Cysteine proteinase inhibitor has been shown toinhibit apoptosis in soybean and Arabidopsis cultured cells(Solomon et al. 1999; Belenghi et al. 2003), and it has beensuggested that plant apoptosis is regulated by the balancebetween cysteine proteinase and cysteine proteinase inhibi-tor activities. Several other proteins that were up-regulatedin region 1 (two In 2–1 proteins and two trypsin inhibitors insoybean, and subtilisin–chymotrypsin inhibitor in maize)may also function in control of protein degradation under

water stress (Moons 2005; Huang, Xiao & Xiong 2007).Up-regulation of trypsin inhibitors under water stress wasshown to have a possible protective role against proteina-ses during leaf senescence (Reviron et al. 1992). Takentogether, the up-regulation of these proteinase inhibitors inregion 1 could prevent water stress-induced protein degra-dation by inhibiting proteinases, thereby contributing tothe maintenance of elongation in this region under waterstress (Fig. 9).

Comparison to other studies

Thirteen of the stress-responsive proteins identified in thisstudy (cysteine proteinase inhibitor, two trypsin inhibitors,two aldose reductases, three ferritin subunits, glutathioneperoxidase, glutathione S-transferase, CCoAOMT, dehy-droascorbate reductase, plasma membrane intrinsicpolypeptide) have been shown to be regulated under waterstress in various organs (Reviron et al. 1992; Riccardi et al.1998, 2004; Karuna Sree, Rajendrakumar & Reddy 2000;Bianchi, Roux & Vartanian 2002b; Vincent et al. 2005;Plomion et al. 2006; Bhushan et al. 2007; Gazanchian et al.2007; Yoshimura et al. 2008). To our knowledge, however,this study provides the first analysis of the spatial patternsof changes in abundance of these proteins in the elongationzone of water-stressed roots. Notably, the list of target pro-teins for thioredoxin reviewed by Buchanan & Balmer(2005) includes several of the stress-responsive proteinsidentified in this study (aldose reductases, trypsin inhibitors,glutathione peroxidase, glutathione S-transferase, dehy-droascorbate reductase and phosphoglucomutase), indicat-ing a correlation between the effects of water stress andoxidative stress (Hajheidari et al. 2007).

It is interesting to note that nine of the identified proteins(two trypsin inhibitors, two aldose reductases, threeferritin subunits, glutathione peroxidase, glutathioneS-transferase), all of which were up-regulated in region 1,have been reported to be regulated by abscisic acid (ABA)in other systems (Kiyosue, Yamaguchi-Shinozaki & Shi-nozaki 1993; Lobreaux, Hardy & Briat 1993; Oberschallet al. 2000; Seki et al. 2002; Rodriguez Milla et al. 2003).Accumulation of ABA is essential for the maintenance ofelongation rates in the apical region of the maize primaryroot at low water potentials (Saab et al. 1990; Saab, Sharp &Pritchard 1992; Sharp et al. 1994); whether ABA is similarlyrequired for the adaptation of soybean primary roots towater stress has not been investigated.

CONCLUSIONS

Under water stress, relative elongation rates are maintainedin the apical region, but inhibited at more basal locations ofthe elongation zone in the soybean primary root. Theseresponses are similar to previous results for the maizeprimary root. Proteomic analysis revealed several novelfeatures that are involved in the adaptation of soybeanprimary root growth to low water potentials. The spatialpattern of regulation of enzymes for phenylpropanoid



biosynthesis is consistent with a substantial increase ofisoflavone content in the apical region, and an increaseof wall phenolics including lignin in the basal region ofthe elongation zone under low water potential conditions.These responses may be causally related to the mainte-nance and inhibition of elongation, respectively, in theseregions. Increased abundance of several proteins for pro-tection against ROS is pronounced throughout the elonga-tion zone of water-stressed roots. In particular, localizationof free and bound iron indicates that the increased abun-dance of ferritin proteins effectively prevents excess freeiron in the elongation zone of water-stressed roots, andthereby is likely of importance to prevent excess ROS pro-duction. The next paper in this series will provide a com-prehensive analysis of transcriptome changes to providefurther insight into the metabolic networks involved inregulation of soybean root growth under water stress.

ACKNOWLEDGMENTS

We thank Lindsey Hejlek for assistance with the genotypescreening experiments; Drs Mark Ellersieck and WilliamMcClain for advice in statistical analysis; Drs Dale Blevins,Anand Chandrasekhar and Qisheng Song for equipmentuse; and Dr David Sleper for supplying soybean seeds. Gelelectrophoresis and mass spectrometry were performed atthe MU Proteomics Center, and microscopic analyses wereconducted at the MU Molecular Cytology core facility. Theferritin antibody was a gift from Dr Jean-François Briat,Institut de Biologie Intégrative des Plantes, Montpellier,France. Root growth analysis and proteomics studies weresupported by a grant from the MU/Monsanto Plant BiologyResearch Program. Initial development of the soybeanseedling system and screening of soybean cultivars weresupported by a grant from the Missouri Soybean Merchan-dising Council. Isoflavone analysis was supported by grantsfrom NSF (MCB0519634) and USDA (NRI2005-05190)to O.Y.

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Received 30 July 2009; received in revised form 23 October 2009;accepted for publication 25 October 2009

SUPPORTING INFORMATION

Additional Supporting Information may be found in theonline version of this article:

Figure S1. Representative two-dimensional electrophoresis(2-DE) gel images of total soluble proteins extracted fromregions 1 and 2 of water-stressed (WS) roots, and regions1–3 of well-watered developmental (WWD) and temporal(WWT) control roots. Four replicate samples were analysedfor all regions.Figure S2. Numbers of up-regulated and down-regulatedwater stress-responsive proteins in region 1 (R1) and region2 (R2). The numbers of responsive proteins in region 2which exhibited similar and significant responses whenregion 2 of water-stressed roots was compared with region 3(R3) of well-watered roots are indicated in parentheses andmarked with asterisks; these changes are considered to beindependent of developmental changes associated with thestress-induced shortening of the elongation zone (see textfor details of this analysis).

Figure S3. Abundance of differentially expressed proteinsin regions (R) 1 and 2 of water-stressed (WS) roots (solidbars), and in regions 1–3 of well-watered developmental(WWD) and temporal (WWT) controls (open bars). Thehistograms show mean spot intensities (�SE) of three tofour replicate samples from independent experiments; thecolumns from left to right represent region 1 (WS, WWD,WWT), region 2 (WS, WWD, WWT) and region 3 (WWD,WWT). Asterisks denote differentially expressed proteinsfor which spot intensities were significantly differentbetween the water-stressed treatment and both of the well-watered controls within region 1 and/or region 2. Trianglesdenote that spot intensities for the water-stressed treatmentin region 2 were also significantly different to region 3 of thewell-watered controls.Table S1. Peptide sequences determined by matrix-assistedlaser desorption/ionization-time-of-flight tandem massspectrometry (MALDI-TOF MS/MS) for identified waterstress-responsive proteins in the elongation zone of thesoybean primary root.Table S2. Comparison between responses to water stress ofregulated proteins in the soybean primary root elongationzone (present study) and genes for proteins with similarfunctions in the maize primary root elongation zone (fromSpollen et al. 2008).

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Regulation of growth response to water stress in the soybean primary root. I. Proteomic analysis...

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