Environmental and Genetic Factors Regulating Localization of ......Environmental and Genetic Factors...

Environmental and Genetic Factors RegulatingLocalization of the Plant PlasmaMembrane H+-ATPase1[OPEN]

Miyoshi Haruta,a,b,2 Li Xuan Tan,c Daniel B. Bushey,e Sarah J. Swanson,d and Michael R. Sussmana,b,2

aDepartment of Biochemistry, University of Wisconsin-Madison, Madison, Wisconsin 53706bBiotechnology Center, University of Wisconsin-Madison, Madison, Wisconsin 53706cDepartment of Ophthalmology and Visual Sciences, University of Wisconsin-Madison, Madison, Wisconsin 53706dDepartment of Botany, University of Wisconsin-Madison, Madison, Wisconsin 53706eJanelia Research Campus, HHMI, Ashburn, Virginia 20147

ORCID IDs: 0000-0003-1686-1860 (M.H.), 0000-0001-9374-3353 (M.R.S.).

A P-type H+-ATPase is the primary transporter that converts ATP to electrochemical energy at the plasma membrane of higher plants.Its product, the proton-motive force, is composed of an electrical potential and a pH gradient. Many studies have demonstrated that thisproton-motive force not only drives the secondary transporters required for nutrient uptake, but also plays a direct role in regulating cellexpansion. Here, we have generated a transgenic Arabidopsis (Arabidopsis thaliana) plant expressing H+-ATPase isoform 2 (AHA2) thatis translationally fused with a fluorescent protein and examined its cellular localization by live-cell microscopy. Using a 3D imagingapproach with seedlings grown for various times under a variety of light intensities, we demonstrate that AHA2 localization at theplasma membrane of root cells requires light. In dim light conditions, AHA2 is found in intracellular compartments, in addition to theplasma membrane. This localization profile was age-dependent and specific to cell types found in the transition zone located betweenthe meristem and elongation zones. The accumulation of AHA2 in intracellular compartments is consistent with reduced H+ secretionnear the transition zone and the suppression of root growth. By examining AHA2 localization in a knockout mutant of a receptorprotein kinase, FERONIA, we found that the intracellular accumulation of AHA2 in the transition zone is dependent on a functionalFERONIA-dependent inhibitory response in root elongation. Overall, this study provides a molecular underpinning for understandingthe genetic, environmental, and developmental factors influencing root growth via localization of the plasma membrane H+-ATPase.

A unique feature of the plasmamembrane in plants andfungi compared to animals is the proton electrochemicalgradient produced by a P-typeH+-ATPase (proton pump),

rather than by a sodium pump. Because the proton pumpis moving only one proton per ATP molecule, the protonreversal potential of the enzyme ismuch larger than that ofthe sodium/potassium pump, which moves three sodiumions out of a cell for every two potassium ions moving in.In concert with this intrinsic biochemical difference be-tween the two enzymes, the membrane potential of plantsis routinely found to be much more negative than in ani-mals. It has allowed plants to evolve rapid hydraulic-based methods for organ movement, such as is readilyapparent in touch-sensitive plants, e.g. Mimosa pudica(sensitive plant) and Dionaea muscipula (Venus flytrap).However, all plants, even the slower growthmovementsthat underlie plant specific phenomena such as photoand gravitropism, also rely on the deep membrane po-tential to ensure that the major osmolyte, the potassiumions (K+), is concentrated in the cytoplasm and itsmovement across the membrane mediated by potas-sium channels ultimately induces the changes in cellu-lar turgor pressure. The proton pump is also involvedin tight regulation of the cell-wall pH. This apoplasticpH in turn controls the extensibility of polymers in thewall, and is known to be altered rapidly and to differentextents in different cell typeswithin one organ, such as theroot (Sussman and Harper, 1989; Michelet and Boutry,1995; Palmgren, 2001).

1 The majority of this work on the ATPase is supported by fundingto M.R.S. from the U.S. Department of Energy (Basic Energy Sciences;no. DEFG02-88ER13938), with additional funding for analysis of theFERONIAmutant plant provided toM.R.S. from the National ScienceFoundation (NSF; MCB grants no. 1410164 and 1713899). Assistancein support of software used in data analysis was provided by fundingto the Department of Ophthalmology and Visual Sciences from theNational Institutes of Health (NIH; P30EY016665).

2 Address correspondence to [email protected] [email protected].

The authors responsible for distribution of materials integral to thefindings presented in this article in accordance with the policy de-scribed in the Instructions for Authors (www.plantphysiol.org) are:Miyoshi Haruta ([email protected]) andMichael R. Sussman([email protected]).

M.H. and M.R.S. designed the experiments. M.H. performed all mo-lecular cloning and plant work. M.H. and S.J.S. operated the microscopeand captured images. L.X.T., D.B.B., and M.H. analyzed the image data.D.B.B. coded all of programs. M.H. interpreted biological results withdiscussion with L.X.T., D.B.B., and S.J.S. M.H. wrote the manuscript withM.R.S. L.X.T. andD.B.B. provided the text of themethods used for the dataanalyses.All authors participated in reviewing and editing themanuscript.

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The Arabidopsis (Arabidopsis thaliana) genome con-tains a protein family of 100-kD plasma membraneH+-ATPases (AHAs)with 11members, althoughAHA1and AHA2 are the most predominantly expressed andplay essential roles throughout the lifecycle (Baxteret al., 2003; Haruta et al., 2010). During the seedlingstage, AHA2 plays critical roles in establishing theproton gradient at the root surface as aha2 insertionmutants show impaired proton secretion and rootelongation at high apoplastic potassium concentrations(Fuglsang et al., 2007; Santi and Schmidt, 2009; Harutaand Sussman, 2012). A number of studies indicate thatproton pump function is modulated at many levels,including transcriptional, translational, and posttrans-lational regulation (Duby and Boutry, 2009; Wang et al.,2014; Haruta et al., 2015; Falhof et al., 2016). In terms ofcatalytic activity, phosphorylation of regulatory Ser andThr residues modulate H+-ATPase-mediated protonpumping. Both genetic and biochemical analyses suggestthat phosphorylation/dephosphorylation of the penul-timate Thr residue (i.e. Thr-947 of AHA2 and Thr-948 ofAHA1 and 3) is key for regulating the ATPase by bluelight and auxin (Fuglsang et al., 1999; Kinoshita andShimazaki, 1999; Robertson et al., 2004; Takahashi et al.,2012; Spartz et al., 2014). Additional phosphorylationsites, such as Thr-881 and Ser-899 of AHA2, are alsolikely involved in the activity modulation and these maybe independently regulated by various protein kinasesand phosphatases that function in the signaling path-ways initiated by receptor kinases such as FERONIA(FER) and PSY1R, in response to the cognate peptideligands (Fuglsang et al., 2014; Haruta et al., 2014). Thecomplexity of H+-ATPase regulation is further under-scored by the recent observation that dephosphorylationat Thr-947 of AHA2 by a protein phosphatase, PP2Cplays an important and unique role in growth regulationby proteins encoded by auxin-induced small auxinupregulated genes, suggesting that efforts in character-izing kinase-mediated phosphorylation may only beproviding a partial understanding of the regulation ofthis enzyme (Spartz et al., 2014).Localization of the H+-ATPase protein at the cell

surface has been extensively characterized primarilyusing biochemical approaches. For example, plasmamembrane fractions have been purified by enrichmentsusing two-phase partitioning and/or by buoyant den-sity gradient centrifugation (Scherer and Fischer, 1985;Sze, 1985; Larsson et al., 1987). Using these cellularfractionation methods combined with the enzyme ac-tivity assay, immunoblots, or peptide sequencing viamass spectrometry, the H+-ATPase is often utilized as acommon marker protein for the plasma membrane,although its presence only in the plasmamembrane hasnever been critically tested under all growth conditions(Palmgren, 1990; DeWitt et al., 1996; Nelson et al., 2006).Tissue or cell-type specificity of H+-ATPase localizationwas shown via an immunocytological approach thatallowed microscopic visualization of this protein atthe surface of pollen, roots, and phloem companioncells (Parets-Soler et al., 1990; Obermeyer et al., 1992;

Samuels et al., 1992; Bouche-Pillon et al., 1994; DeWittand Sussman, 1995). In a more recent study, GFP-taggedAHA1 overexpressed in the stomatal guard cells wasvisualized at the cell surface and its localization waspartially dependent on a MUN domain protein, Munc13(Hashimoto-Sugimoto et al., 2013). In another study, thelevel of AHA2-GFP protein expression in the primaryroot was shown to be altered when external nitrogensupplies were limiting (Mlodzi�nska et al., 2015).

Fluorescent microscope techniques combined within vivo expression of fluorescently labeled proteins ororganelle stainingdyes have beenwidely used to quantifyprotein localization and cellular dynamics (Lippincott-Schwartz et al., 2001; Hamilton, 2009). By capturing aseries of confocal optical sections and rendering a pro-jection, 3D views of live cells or tissues as well as locali-zation patterns of a protein of interest can be obtained in anoninvasive manner (Masters et al., 1997; Truernit et al.,2008). Imaging developing Arabidopsis organs andquantifying fluorescent signals allow plant biologists totrack cell lineage in addition to observing protein dy-namics, thus enabling computational modeling of bio-logical processes (Roeder et al., 2011). In this study, weestablished 3D live-cell imaging tools and a platform toexamine the localization of AHA2 in an aha2-4 mutantline that stably expresses fluorescent protein (FP)-taggedAHA2 under the native ATPase promoter. We exam-ined how light affects FP-taggedAHA2 localization in theroot and our results demonstrate that, under dim lightconditions, FP-tagged AHA2 proteins accumulate in thecytoplasmic compartments in the cells residing in thetransition zone. This localization change in AHA2 corre-lates with apoplastic pH alkalinization and root growthsuppression. Furthermore, we show that this internal lo-calization of AHA2 in the cytoplasm was dependent onthe normal function of FER receptor protein kinase. Theseresults emphasize the importance of live-cell imagingwith the plasma membrane proton pump, to understandmembrane energetics and growth regulation in planta.

RESULTS

Expression of a mCitrine-AHA2 TranslationalFusion Protein

To perform live-cell imaging of the AHA2 protein inArabidopsis plants, we transformed aha2-4 insertionalknockout mutant plants with an AHA2 genomic con-struct containing mCitrine fluorescent tag fused at theamino acid position, Leu-4 of AHA2 (mCitAHA2; Fig.1A). Two independently transformed mCitrine-AHA2(mCitAHA2) lines, aha2-4;mCitAHA2a and aha2-4;mCitAHA2b that are homozygous for a single insertionof the transgene, were used for further study. Ourprevious results have showed that the aha2-4 insertionmutants display impaired root elongation when grownon media containing high external potassium ion con-centration, presumably due to the stress placed on en-ergetics at the plasmamembrane in the presence of high

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3D Imaging P-type H+-ATPase in Arabidopsis Roots



concentrations of this positively charged current-carrying abundant osmolyte (Haruta and Sussman,2012). A comparison of aha2-4;AHA2 (complementationby wild-type AHA2 sequence) and aha2-4;mCitAHA2lines in the root growth assay indicates that mCitAHA2can partially rescue the shorter root phenotype of aha2-4(Fig. 1B). Together with the carboxyl terminus, the aminoterminus of AHA2 is known to be a regulatory domain ofits enzymatic activity (Ekberg et al., 2010), and thus thereduced ability of mCitAHA2 to rescue the impaired rootgrowth of aha2 mutant is likely caused by misregulationdue to compromising the intact amino terminus. To ex-amine cellular localization of mCitAHA2 protein, weextracted and fractionated seedling tissues from aha2-4;mCitAHA2 line to enrich for plasma membrane pro-teins (Fig. 1C). Immunoblot detection of the soluble,endomembrane (lower phase from the two-phase par-titioning), and the plasma membrane fractions (upperphase from the two-phase partitioning), indicates thatmCitAHA2 is enriched in the plasma membrane frac-tion of seedling extracts. The SDS-gel banding patternsuggests that mCitAHA2 protein is present mainly in amonomeric form at approximately 125 kD, with possi-ble oligomers thatwere visible at higherMr regions. Thelower phase, which contains primarily endomem-branes such as the tonoplast, also containedmCitAHA2protein. This may reflect two possibilities: 1) either thefractionation is not complete and thus PM-localizedmCitAHA2 enters the lower phase in this protocol, or 2)mCitAHA2 protein is normally present in the endo-membrane system, albeit to a smaller extent than foundat the cell surface. Using live cell imaging, we were ableto use an independent orthogonal technology to ex-amine this question.

Correlation of Root Growth Suppression and IntracellularAccumulation of AHA2 under Dim Light

The H+-ATPase plays a critical role in providing theenergy for organ development and elongation pro-cesses during germination. For example, it is well knownthat seedling root and hypocotyl elongation is stronglyinfluenced by the light intensity (Fankhauser and Chory,1997). Light regulation of the H+-ATPase activity isthought to contribute to optimizing growth rates ofseedling organs (Spalding andCosgrove, 1992; Takahashiet al., 2012) although a precise understanding of thisprocess has not yet been achieved. To evaluate AHA2localization at the cellular level, fluorescent signals ofmCitAHA2 were visualized in seedlings grown for 10 dunder low (2.2 mmol m22 s21) or bright light (approxi-mately 10 mmol m22 s21) intensities, using a confocallaser scanning microscopy (CLSM). Consistent withpreviously described typical Arabidopsis growth phe-notypes at various light intensities (Hardtke and Deng,2000; Chory, 2010), aha2-4;mCitAHA2 roots grow longerin the bright light compared with dim light whereashypocotyls grew longer in the dim light conditioncompared with bright light (Fig. 2A). To visualize

mCitAHA2 fluorescent signals in the tissues in whichrapid cell growth is occurring, 3D images were recon-structed from approximately 60 optical slices for theapex of hypocotyls and root tips (Fig. 2, B to E). Underboth the dim and bright light conditions, mCitAHA2signals in hypocotyl cells were detected at the plasmamembrane (Fig. 2, B and C). In roots, mCitAHA2 waslocalized at the cell surface in the seedlings grown un-der the bright light condition (Fig. 2D). The seedlingroots grown under the dim light showed the cell-surfacelocalization of mCitAHA2 signal in the root tip as well aselongation zone but surprisingly, we observed accumu-lation of the signal in the cytoplasm of the cells in thetransition zone (Fig. 2E). Thus, the shorter root length ofthe seedlings grown under the dim light correlates withthe accumulation of mCitAHA2 in the cytoplasm in thisregion.

To further investigate the mCitAHA2 localizationpattern with seedlings grown under the dim lightcondition, root growth was measured daily (Fig. 2F). Acomparison of root growth curves under bright anddim conditions for a 9 d period shows that roots grownunder the bright condition sustain growth whereasroots under the dim condition deviate from this andstop elongation around 8 d old (Supplemental Fig. S1).Because the growth medium contains half-strengthMurashige & Skoog (MS) nutrient salts and 1% (w/v)Suc, the root growth cessation under the dim conditionis likely due to the endogenous developmental pro-gram instead of nutrient depletion. In rapidly growingroots at 3 d old to 5 d old under dim light, mCitAHA2 islocalized at the plasma membrane (Fig. 2G), whereasthe accumulation of mCitAHA2 in the cytoplasm wasvisiblewith 10-d-old seedlings (Fig. 2H). This cytoplasmicaccumulation of mCitAHA2 was specific to cells residingin the transition zone of roots (Supplemental Movie S1).

The mCitAHA2 intensity levels were much greater atthe plasma membrane compared to accumulation incytoplasmic areas. This difference provided a meansto quantify cytoplasmic and membrane accumulation(Fig. 2I, Supplemental Fig. S2). Voxels were dividedby intensity into low and high intensity levels. Thethreshold for low intensity voxels was set above back-ground but below the intensity levels occurring at themembrane. The threshold for high intensity voxels wasset such that these voxels predominately occurred at themembrane. The total number of low (red) and high(yellow) intensity voxels were summed in windows8mmwide along the root length throughout the volumeand divided by the total number of voxels above thebackground within the volume to give the ratio ofvoxels falling into either category. In the roots grownunder the dim light, at 80 mm to 140 mm from the tip,corresponding to the transition zone, a majority ofvoxels contained the low (red) intensity voxels indi-cating that mCitAHA2 was diffused in the cytoplasm.In contrast, in bright conditions, high intensity voxelsoccur muchmore frequently, indicating dense mCitAHA2accumulation at the cytoplasmic compartments. This effectis best summarized by subtracting the number of low

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intensity voxels from the high intensity voxels pro-ducing an accumulation index (blue). In the transitionzone, in dim conditions, .80% of the voxels have lowintensity but in comparison in bright conditions, theaccumulation index reaches almost 0%, indicating thenumber of high and low intensity voxels are equal.Additional experiments testing mCitAHA2 localizationunder various light intensity conditions ranging be-tween 1.2 mmol m22 s21 and 20 mmol m22 s21 found thatthe intensity at approximately 5 mmol m22 s21 is thecritical point at which intracellular accumulation ofmCitAHA1 occurs (Supplemental Fig. S3).

In separate experiments, we have also produceda different set of transgenic lines, aha2;Dendra2AHA2plants that express Dendra2 fluorescent protein-taggedAHA2 in the aha2 mutant background (SupplementalFig. S4, A and B). The roots of aha2;Dendra2AHA2seedlings grown under the bright or dim light condi-tions showed this FP-tagged AHA2 localization profileto be very similar to that seen with the aha2-4;mCitAHA2plants, i.e. cell surface localization under the bright con-dition and intracellular accumulation in the transitionzone of the roots under the dim condition. Those resultssupport our observation that dim light caused the intra-cellular accumulation of AHA2 protein in the transitionzone and that this was not due to a particular reporterprotein used for the detection.

Characterization of Intracellular Accumulation ofmCitAHA2 in the Transition Zone of Roots Grownunder Dim Light

To quantitatively describe the intracellular localiza-tion of mCitAHA2 in the roots grown under the dimlight condition, we computationally segmented the 3Dimage of mCitAHA2 roots (Supplemental Fig. S5) intothree parts: 1) meristem, 2) transition zone, and 3)elongation zone. The number of mCitAHA2-positiveintracellular structures are quantified as detailed in“Materials and Methods” (Fig. 3A). At 4 d old, thenumber of intracellular structures is comparable amongthe three root zones. However, it was evident that at11 d old, themCitAHA2-labeled intracellular structuresspecifically accumulated in the transition zone, butnot in the meristem or elongation zone. Analyses forcolocalization efficiencies of mCitAHA2 signals and thesurface of live cells stained with impermeant propi-dium iodide (PI) revealed that the group of cells resid-ing in the transition zone had significantly lower R2

values than those in the elongation andmeristem regions(Fig. 3D). The relative lower R2 value for the transitionzone agreed with our visual inspection of the mCitAHA2accumulation in the cytoplasmof the cells in the transitionzone because the cell surface was clearly visible in the PIchannel (Fig. 3, E and F).

Examination of the XY section of the 3D image sug-gested that mCitAHA2 signal accumulated in intracel-lular components as large vesicles (Fig. 3, G and H). Inthe cells that are experiencing elongation, mCitAHA2

Figure 1. Expression of fluorescently labeled AHA2 in plants. A,Membrane topology of mCitrineAHA2 (mCitAHA2) fusion protein. ThemCitrine fluorescent protein sequence was inserted at the amino acid posi-tion Leu-4 located in the cytoplasmic face of the AHA2 sequence. B, Rootgrowthmeasurements ofmCitAHA2-expressingplants.Arabidopsis seedlingsof wild type, aha2-4mutant, aha2-4;AHA2 (wild-type complementation) oraha2-4;mCitAHA2weregrown for 3donto1/2MSmedia, then transferred tothemedia supplementedwith 100mMKCl and grown for additional 10 d. C,Immunological detection of mCitAHA2 protein from aha2-4;mCitAHA2plants. Ten-d-old seedlings were extracted to enrich the plasma membranefraction using the two-phase partitionmethod. Fifteenmicrograms of proteinswere loadedontoprotein gel.Blotwasprobedwith anti-GFP (right).Once theblot membranes were scanned for the immunodetection, they were stainedwith Ponceau S tovisualize proteins (left). Sol., the fraction containing solubleproteins; LP, the lower-phase fraction containing endomembrane proteins;UP, the upper-phase fraction containing plasma membrane proteins.









signals were diffused across the cells. At the maturationzone, a majority of mCitAHA2 signals were localized tothe plasma membrane and no detectable signals in theintracellular space (Fig. 3, H to J). Those images may bereflecting the localization of AHA2 protein moleculesthat were in the process of trafficking to or from the PMand were paused in their locations when the cell elon-gation process was inhibited due to the absence ofsufficient light intensity.

Intracellular Accumulation of mCitAHA2 Correlates withpH Alkalinization at the Root Surface of theTransition Zone

Both our previous experiments and other studieshave shown that epidermal cells of the primary rootcontain a high expression level of the AHA2 transcriptand protein, and this expression pattern was supportedby impaired root growth phenotypes of aha2 mu-tants on appropriate growth conditions (Haruta andSussman, 2012; Mlodzi�nska et al., 2015). Thus, protonsecretion by AHA2 activity is expected to play the majorrole in creating proton gradient at the root surface. Toexamine proton concentrations at the surface of rootsgrown under dim light, we surrounded mCitAHA2seedling roots in a solution containing a pH probe,fluorescein-dextran (FITC-dextran), and captured 2Dimages (Fig. 4A). FITC-dextran has been used to visualizepH changes at the root surface in Arabidopsis (Ohkumaand Poole, 1978; Monshausen et al., 2007). Surface pHwascalibrated with standard solutions buffered to pH 5.5, 6.0,or 6.5 (Supplemental Fig. S6). Consistent with the aboveobservation,mCitAHA2protein clearly localizedat thePMin thematuration andmeristematic zones but accumulatedin the cytoplasmic compartments in the transition zone.

Figure 2. Organ length andmCitAHA2 localization in aha2-4;mCitAHA2plants grown under bright or dim light conditions. A, Hypocotyl and rootlength of aha2-4;mCitAHA2 seedlings grown for 10 d under dim (2.2mmolm22 s21) or bright (10 mmol m22 s21) light conditions. B, Hypocotyl ofaha2-4;mCitAHA2 seedling grown for 10 d under the bright light. C, Hy-pocotyl of aha2-4;mCitAHA2 seedling grown for 10 d under the dim light.

D,Root of aha2-4;mCitAHA2 seedling grown for 10dunder the bright light.E, Root of aha2-4;mCitAHA2 seedling grown for 10 d under the dim light. F,Root growth curve of aha2-4;mCitAHA2 seedlings grown under the dimlight. n = 7. Data are shownwithmean6 SE. G, Root of aha2-4;mCitAHA2seedling grown for 4 d under the dim light. H, Root of aha2-4;mCitAHA2seedling grown for 10 d under the dim light. For (B) to (E) and (G) to (H),images are a maximum intensity projection of fluorescent CLSM opticalsections. Scale bars indicate 50 mm for (B) and (C) and 20 mm for (D) to (E)and (G) to (H). I, Quantitation of mCitAHA2 fluorescence intensity. Themeanfluorescent intensity valueswere calculated along the root length bya21-pixel window size corresponding to approximately 8mm from three 3Dimages of aha2-4;mCitAHA2 roots grown the dim or bright conditions(Supplemental Fig. S2). (Top) Heat map for aha2-4;mCitAHA2 intensityscale; white indicates the highest intensity and black indicates the lowestintensity. (Middle) Images of aha2-4;mCitAHA2 seedling roots grownunderthe dim or bright light conditions. The x axis of the root images correspondsto the scale shown in the bottom chart showing the intensity profiling.(Bottom) Quantified intensity profiles for aha2-4;mCitAHA2 seedling rootsgrown under the dim or bright light conditions. Approximately 60 opticalslices were summed at a given pixel position. Voxels were categorized intotwo groups: high intensity (yellow), or medium intensity (red). Subtractingthe number of low-intensity voxels from the high-intensity voxels producedthe accumulation index (shown with blue plot).


Haruta et al.





Signals from the FITC pH reporter at the root surface andits profiling reveal higher pH values in the apoplast of thetransition zone comparedwith that in thematuration zone(Fig. 4, B to D), correlating with reduced abundance ofmCitAHA2protein in the plasmamembrane in this region.Unexpectedly, we also noticed that this higher pH bandspans above the transition zone to the elongation zone,suggesting that the H+-ATPase in this region is less activepossibly caused by less 14-3-3 binding, altered phospho-rylation, or low cytosolic pH. Alternatively, it may becaused by changes in other proton transporters (e.g.proton-coupled carriers) or other pH-regulating mecha-nisms in the elongation zone (see “Discussion”).

As a control, we tested the effect of buffered solutions(pH 5.5) containing the pHprobe to ensure that changesin the FITC fluorescent intensity are responding to pHvalues and not to changes in other ion concentrations(Fig. 4, E to H). In these experiments, the bright greensignals seen at the transition zone of the root sub-merged in the unbuffered solution disappeared in thebuffered medium, supporting the idea that our obser-vations, using unbuffered media, indeed report changesin pH values.

FERONIA Receptor Kinase Is Required for Root GrowthSuppression and mCitAHA2 Intracellular Accumulation inSeedling Roots Grown under Dim Light

During seedling development, root growth is pre-sumably optimized by the fine adjustment of positiveand negative regulatory mechanisms. Our model pre-dicts that insufficient light intensities will suppress rootelongation, and this is in part mediated by an intracel-lular signaling cascade initiated by the FERONIA recep-tor kinase (Haruta et al., 2014). This idea is supported by

Figure 3. Visualization and quantitation of intracellular localization ofmCitAHA2 fluorescent signals in aha2-4;mCitAHA2 roots grown underdim light for 11 d. A, Quantitation of intracellularly localized mCitAHA2structures (i.e. blobs). Root region212mmfrom the tip in the3D imageswas

segmented into the meristem (61.28 mm from the tip), the transition zone(63.77mmabove themeristem), and the elongation zone (86.95mmabovethe transition zone). For each region, surface rendering and identification ofintracellularly localized structures was performed as detailed in “Materialsand Methods” (an example of the surfaces generated is shown inSupplemental Fig. S5). Graph shows mean number of intracellular struc-tures 6 SD (n = 3). B, 3D image of root tip of aha2-4;mCitAHA2 seedlinggrown for 10 d under the dim light. Image was detected by mCitrineemission. Bar, 20 mm. C, PI staining of the identical aha2-4;mCitAHA2seedling shown in (B). Image was detected by PI emission. D, Two-channelmerge of (B) and (C). Regression values for colocalization ofmCitAHA2andPI were shown at the right of the corresponding regions. E, Magnified viewof the transition zone that was visualized in the mCitAHA2 channel shownin (B). Bar, 10 mm. F, Magnified view of PI-stained cells in the same regionshown in (E).G,Orthogonal viewofmCitAHA2 root. (Top)XZ section of theroot at the position indicated with the green line shown in the bottomimage. (Bottom) XY section of the root at the position indicated with theblue line shown in the top image. Bar, 15mm.H,Magnifiedviewof the rootregions showing the transition of mCitAHA2 localization from the intra-cellular compartments to the plasma membrane. Bar, 15 mm. I, Diagramshowing the interpretation of the fluorescent raw image shown in (H). Toassist the interpretation of changes in mCitAHA2 localization, the imagewas reproduced by drawing. J, Annotation of cell types and mCitAHA2cellular localization.






loss-of-function mutants of FERONIA that show fasterroot elongation under a dim light condition, whereaswild-type root growth is suppressed in the samedim lightcondition (Haruta et al., 2014). To further compare spatialregulation of mCitAHA2 protein between wild type andfer mutant plants, we produced fer-4;aha2-4 doublemutants carrying the mCitAHA2 fluorescent reporterby genetic crossing of fer-4 and aha2-4;mCitAHA2aor aha2-4;mCitAHAb. Under the bright light condition(10 mmol/m2/s), root length measurements of aha2-4;mCitAHA2 and fer-4;aha2-4;mCitAHA2 are comparable(Fig. 5, A andC). As the light intensity decreases, aha2-4;mCitAHA2 root length was dramatically decreasedwhenmeasured at 10 d old (Fig. 5, A and B). Root growthmeasurements of fer-4;aha2-4;mCitAHA2 seedlings werealso reduced as the light intensity decreases and the effect

of fer-4 knockout mutation was clearly visible becausefer-4;aha2-4;mCitAHA2 roots grow faster than aha2-4;mCitAHA2 roots.

The roots of seedlings carrying a wild-type FER alleleshow the accumulation of mCitAHA2 fluorescentsignals in cytoplasmic compartments of specific cellsunder the dim light conditions (2.2 mmol/m2/s). Fluo-rescent intensity profiling of mCitAHA2 in the aha2-4;mCitAHA2 seedling grown under the dim light wasperformedwith amethod similar to that shown in Figure2I, and these displayed a distinct profile from that in thesame genotype grown under bright light (Fig. 5, D andE). In contrast, under dim light conditions, mCitAHA2localization profiles in the fer-4;aha2-4;mCitAHA2 rootsappeared to resemble those observed with the samegenotype grown under the bright light (Fig. 5, F and G).

Figure 4. Measurement of root surfacepH. A, Image of aha2-4;mCitAHA2 rootgrown under dim light for 10 d. A, Rootimage was visualized with mCitrinechannel detection. The yellow color inthe bathing media is due to overlappingemission from FITC-dextran. A root wassubmerged in an unbuffered solutioncontaining 1 mM KCl, 1 mM CaCl2, and3 mg/mL FITC-dextran. Bar, 20 mm. B,Visualization of FITC-dextran as a pHprobe for measuring the root surface pHof the identical plant. Brighter green in-dicates higher pH values. C, Intensityprofiling the root surface pH. (Left) Colorscale for pH. (Right) Heat map of pHvalues at the root surface. Region sur-rounded by the white line was used tomeasure root surface pH. D, Quantita-tion of the mean pH values at the rootsurface. Data are shown as the mean6 SE.n = 4. E, Image of aha2-4;mCitAHA2root grown under dim light for 10 d andvisualized with mCitrine channel de-tection. A root was submerged in abuffered solution containing 10 mM MES(pH5.5), 1 mM KCl, 1 mM CaCl2, and3 mg/mL FITC-dextran. F, Visualizationof FITC-dextran as a pH probe for mea-suring the root surface pH. G, Profilingthe root surface pH. (Left) Color scale forpH. (Right) Heat map of pH values at theroot surface. H, Quantitation of pHvalues at the root surface. Data are shownas the mean 6 SE. n = 4.


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Because mCitAHA2 signals in the plants with fer-4 ge-netic background appear to be lower than those withFER wild-type plants (Fig. 5, F and G), we extractedtotal protein from seedlings grown under bright or dimconditions and examined mCitAHA2 protein abun-dance with GFP immunoblot (Fig. 5, H and I). WhereasmCitAHA2 level is comparable between the two geno-types when plants were grown under bright light, themCitAHA2 protein level in plants with fer-4 allele issomehow higher than in those with the FER wild-typeallele when plants were grown under dim light (Fig. 5I).This observation seems to be in contradiction with theresults seenwithmCitAHA2fluorescent imaging (Fig. 5,F and G); however, it is likely that the results fromthe immunoblot assay reflect true mCitAHA2 proteinabundance as fer-4 roots secretes more protons to theapoplast and elongates faster than wild type (Harutaet al., 2014; Du et al., 2016; Yeats et al., 2016; see “Dis-cussion” for further description). Regardless of weakmCitrine fluorescent signals in the fer-4 genetic back-ground, overall these results are consistent with thefaster growth of fer-4 mutant roots being caused by theabsence of FERONIA kinase suppressing the intracellu-lar accumulation of mCitAHA2 protein under the dimlight condition, leading to increased proton pumpfunction and an increased rate of cell expansion.

DISCUSSION

In this study, we investigated when and whereplasma membrane H+-ATPase function is regulated atthe cellular level in Arabidopsis roots. The H+-ATPaseis long known to be localized at the plasma membrane,as this enzyme fulfills the essential role in secretingprotons into the apoplast to generate both the chargeand pH gradients necessary to energize nutrient up-take. However, the dynamic regulation of pump lo-calization at the cellular and subcellular levels is poorlyunderstood. By visualizing fluorescently labeled AHA2protein with a 3D live-cell imaging approach, we have

Figure 5. FERONIA-dependent root growth suppression and mCitAHA2intracellular accumulation. A, Effect of fer-4 mutation on dim-light-dependent root growth suppression. The aha2-4;mCitAHA2 or fer-4;aha2-4;mCitAHA2 seedlings were grown under various light intensityconditions for 10 d. n = 7. Data are shown as themean6 SD. B, Images of

aha2-4;mCitAHA2 and fer-4; aha2-4;mCitAHA2 seedlings grown underthe dim light for 10 d. Bar, 1 cm. C, Images of aha2-4;mCitAHA2 andfer-4; aha2-4;mCitAHA2 seedlings grownunder the bright light for 10 d.D,Fluorescent intensity profiling of aha2-4;mCitAHA2 root grown under thedim light for 10 d. E, Fluorescent intensity profiling of aha2-4;mCitAHA2root grown under the bright light for 10 d. F, Fluorescent intensity profilingof fer-4;aha2-4;mCitAHA2 root grown under the dim light for 10 d. G,Fluorescent intensity profiling of fer-4;aha2-4;mCitAHA2 root grown un-der the bright light for 10 d. Scale bar indicates 20 mm for (D) to (G). H,Comparison of mCitAHA2 protein abundance between FER wild type andfer-4 knockout plants grown under bright or dim light conditions. Crudeprotein extracts (6.5 mg) from seedlings grown for 10 d under bright or dimlight were analyzed. (Top) Immunoblot detected with GFP antibody.(Bottom) Ponceau S staining of the identical blot is shown as a loadingcontrol. I, Quantitation of GFP positive signals shows that fer-4;aha2-4;mCitAHA2 plants grown under dim light contain moremCitAHA2 proteinthan aha2-4;mCitAHA2 plants. n = 3. Data are shown as the mean 6 SE

values.





found that AHA2 localization in roots is influenced bylight during elongating growth, which is specificallyobserved in a particular region of the root, the transitionzone.We also demonstrate that this light- and cell-type-dependent localization of AHA2 requires the functionof the FERONIA receptor kinase localized at the plasmamembrane.

Fluorescent Fusion Protein of AHA2 Reports Its CellularLocalization in Live Cells

Our challenge in the understanding H+-ATPase reg-ulation at cellular localization level has been, in part,due to the absence of an appropriate genetic resourcethat can report on membrane trafficking processes ofthe H+-ATPase in vivo. We have attempted to produceArabidopsis lines expressing a genetically encoded fu-sion protein of AHA2 with fluorescent proteins, bycomplementing the aha2 insertion mutant phenotypeusing the native promotor. Although not all the fluo-rophores tested here provided robust and reliable sig-nals, we confirmed that mCitrine, which is a yellowderivative of a jelly fish GFP, and Dendra2, which is aderivative of a coral fluorescent protein (FP), reportedsimilar AHA2 localization patterns in the roots of de-veloping seedlings. In our root growth phenotypingassay, these fluorescently tagged AHA2 constructs canrescue the aha2-4 mutant phenotype only partially;however, FP-tagged AHA2 localization changes in re-sponse to the growth environment, and genetic ablationindicates that the signals from the FP-tagged AHA2proteins likely reflect in vivo behavior of theH+-ATPasemolecule. The importance of choosing appropriatefluorophores and protein-tagging sites within the pro-tein sequence was also suggested from an FP-taggingstudy of vacuolar H+-pyrophosphatase, which is also apolytopic transmembrane protein (Segami et al., 2014).

In our control experiment, we grew an Arabidopsisline expressing GFP-tagged ATP-binding cassettetransporter 36, ABCG36 (PEN3), known to be localizedat the plasma membrane and proposed to transporta wide range of solutes to the apoplast, under iden-tical conditions to those of the FP-tagged AHA2(Supplemental Fig. S6; Stein et al., 2006; Kim et al.,2007). This protein, however, was not accumulated inintracellular compartments, unlike the mCitAHA2 andDendra2AHA2, thus indicating that not all of theplasma membrane proteins undergo intracellular ac-cumulation under a dim-light-growth condition. AHA2is preferentially localized at the top and bottom of ep-idermal cells in the transition to elongation zones,whereas consistent with the previous study, ABCG36/PEN3 is localized at the outer edge of the epidermalcells in those regions (Supplemental Fig. S7, A and B;Strader and Bartel, 2009). Thus, it is possible that themCitAHA2 molecules that were normally targeted tothe PM at the top or bottom of cells under the bright-light conditions were internalized to down-regulateAHA2 function in the absence of sufficient light inten-sities. This effect of light could be to either increase the

movement of vesicles to, or decrease the movementfrom, the plasma membrane, because our experimentscannot distinguish between these two processes.

A previous study of the plasmamembraneH+-ATPasemovement to and from the plasma membrane hasreported that indole acetic acid treatment enhancedmembrane flow from the endoplasmic reticulum to theplasmamembrane (Hager et al., 1991). The indole aceticacid-induced increase in the H+-ATPase protein level atthe plasma membrane was observed within 10 min,which correlated with a decrease in the pH value ofbathing medium and the initiation of the elongationgrowth of maize coleoptiles. This study utilized cole-optiles of maize and thus, together with this study us-ing Arabidopsis, our collective results support a criticalrole of theH+-ATPase trafficking dynamics between theendomembrane and plasma membrane in the regula-tion of elongation growth of plant organs.

Seedling Roots Adapt to Dim Light Conditions byIntracellular Accumulation of AHA2, Cell Surface pHAlkalinization in the Transition Zone, andGrowth Suppression

The transition zone has also been referred to previ-ously as the distal elongation zone, in which roots ini-tiate responses to various stimuli such as the gravityandmechanical force (Ishikawa and Evans, 1995). Morerecent studies suggest that the cells in this region aremore active in ion transport processes compared to thecells in the regions above and below (Masi et al., 2009).Our observations of the transition zone acting as theimportant growth regulatory region for localizing theAHA2 protein at the PM coincide with the previouslynoted distinctive roles of the cells in this region in de-termining root physiology; however, there are nocausal links to specifically relate the down-regulation ofAHA2 function in this region and the previous obser-vations reporting on the role of the transition zone inenvironmental sensing. Interestingly, Wan et al. (2012)showed that another plasma membrane transporter,PIN2, known to function in auxin transport processes,was accumulated in intracellular compartments in thearea covering the transition zone in dark-grown roots.The authors of this study and another earlier studyproposed vacuole-like compartments or prevacuolarcompartment as possible organelles in which PIN2 ac-cumulates when seedlings were grown under thedark condition (Kleine-Vehn et al., 2008). BecauseAHA2 is the major cation-transporting protein at theplasmamembrane during longitudinal growth of roots,it would not be surprising if other anionic solutetransporters such as PIN2 are also subject to the re-duced accumulationwithin the same PM compartment.Although it remains to be determined whether AHA2and PIN2 are trafficked in the same or different types ofvesicles/organelles, one may speculate that AHA2 alsoaccumulates or is subject to proteolysis in the vacuole-like compartments or prevacuolar compartment in the


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transition zone of roots grown under dim light. Thisnotion is also supported by a potential truncation ofmCitrine tag in the seedling grown under dim light(Fig. 5H).Our pH measurement assay shows an overall cor-

relative trend between higher surface pH values andintracellular accumulation of mCitAHA2 protein in thetransition zone of the roots grown under dim light. Thissupports our model that AHA2 proton secretion ac-tivity in this region is a major contributor to the regu-lation of the proton-motive force and root growth. Thisidea is also supported by a recent observation thatacidic apoplastic pH contributes to cell expansion inroots (Barbez et al., 2017). We did not identify possiblecauses of a stretch of higher pH region also observedabove the transition zone. One possibility is that thealkaline pH observed here was caused by a diffusion ofthe anionic form of the pH reporter, FITC, from theadjacent transition zone during the experiment, thusresulting in bright fluorescent signals near the elonga-tion zone. The use of an apoplastic pH reporter systemthat is based on a genetically encoded pH-sensitivefluorophoremay also help clarify our observation of thealkaline pH above the transition zone (Gjetting et al.,2012). Alternatively, the use of another pH reporter dyefor ratio measurements helps correct the potential in-homogeneous concentration errors observed with theFITC reporter. Other plausible explanations are that thetransport of ions other than protonsmay also contributeto the alkalinization seen at the cell surface. Alterna-tively, proton-coupled solute transporters could bemore active at the region and cause a higher rate ofproton import into these cells. If the latter is the case, inthe rapidly expanding cells, nutrients are likely activelytaken up into the cells together with protons (Yeatset al., 2016), thus depleting apoplastic proton concen-tration and inducing the external pH alkalinization inthe absence of AHA2 function or in conjunction withthe reduction of proton extrusion by AHA2.

FERONIA Receptor Kinase Acts Upstream AHA2 in aSignal Pathway that Restricts Seedling Root Elongationunder a Dim Light Condition

During germination and seedling stages, the pre-formed cotyledon and root stored in seeds contributesto provide nutrients and energy to drive expansiongrowth. The reserved nutrients would last until trueleaves emerged during postembryonic development,which occurs at approximately 10 d after germination(Boyes et al., 2001). During this period, the rates of seedgermination and seedling growth are highly influencedby light intensity. Our observation that the intracellularaccumulation of mCitAHA2 became visible at approx-imately 10 d old when grown under dim light, corre-lates with the timing of the onset of this postembryonicdevelopment. We predict that dim-light grown seed-lings that did not establish photosynthetic capabilitycease root-growth at this age, which coincides with the

removal of the H+-ATPase from the plasma membranein the cells of the transition zone. Although our resultsdo not provide any experimental evidence as to howseedling roots sense light intensity or transmit lightsignaling, previous studies showed that blue lightreceptors, cryptochrome and phototropin are bothexpressed in seedling roots (Sakamoto and Briggs, 2002;Endo et al., 2007). Thus, it is possible that seedling rootscan directly sense light intensity and quality. We didnot test whether the intracellular accumulation ofmCitAHA2 occurred in the seeds germinating on thesoil surface; however, one could speculate a similarsituationwould occur when roots are crawling just nearon the soil surface for the first several days after thegermination, and then seedling roots can directly senselight intensity to adapt to the light environment. De-spite this speculation, it remains to be studied whethersoil-grown adult plants respond like the seedlings didwith the intracellular accumulation of mCitAHA2. Inthe adult plants grown in soil, there are at least severaldifferent light-sensing mechanisms by which leavesand stems sense light and transmit signals to roots (Leeet al., 2017; van Gelderen et al., 2017).

In Arabidopsis roots, AHA2 activity appears to beregulated by protein phosphorylation pathways in-volving multiple protein kinases and phosphatases(Fuglsang et al., 2007; Nühse et al., 2007; Spartz et al.,2014). FERONIA receptor kinase is one of the proteinsregulating AHA2 through a transient phosphorylationat the Ser-899 site (Haruta et al., 2014). To support agenetic basis for FERONIA kinase as a regulator ofAHA2, we examined mCitAHA2 fluorescent signals ina fer-4;aha2-4 double mutant background and com-pared that in an aha2-4 single mutant. In agreementwith the fer-4 root elongation phenotype, mCitAHA2 inseedling roots carrying the fer-4 allele remained local-ized at the PM in the seedling roots grown under thedim-light growth condition, whereas in roots carryingthe FER wild-type allele, the pump is internalized (Fig.5). This supports our model that FERONIA is actingupstream of AHA2 in the signaling pathway of RALF-and FERONIA-regulated root cell growth expansion.We have also noticed that mCitAHA2 fluorescent sig-nals from fer-4;aha2-4 seedling roots were weaker thanthose from aha2-4 seedling roots. This was observed inthe two independently transformed aha2-4;mCitAHA2lines that are independently crossed to the fer-4mutant.Because fer-4 roots secrete more protons and elongatefaster than wild type (Haruta et al., 2014; Du et al.,2016), it is possible that fer-4 root cells accumulate moreor less of other solutes, such as chloride anions, in thecytoplasm. Indirect measurements of the plasmamembrane electric potential suggested that fer-4 rootcells display a deeper electric potential (Yeats et al.,2016). Fluorescent intensity emitted by many fluo-rophores, including mCitrine, is influenced by micro-environments including pH and ion concentrationssurrounding the FP molecules (Griesbeck et al., 2001).Thus, if cytoplasmic ion(s) concentrations are alteredin the fer-4mutant,wemay observe increases or reduction





of mCitAHA2 signals caused by this change. Regardlessof the cause, mCitAHA2 localization profiles in fer-4 rootsgrown under the two light conditions are very distinctfrom those in FER wild-type genetic backgrounds, sup-porting our model that FERONIA action is located at theupstream of AHA2 function and influences its localiza-tion at the PM.

CONCLUSIONS

Altogether, we have produced imaging tools andmethods to analyze spatial regulation of the plasmamembrane H+-ATPase molecule localization in vivo.The use of 3D imaging and intensity profiling approachesprovide an unbiased view of AHA2 localization.Manyprevious studies have provided observations that sup-port an acid growth model in which there is a positivecorrelation between H+-ATPase activity and cell ex-pansion (Rayle and Cleland, 1992; Hager, 2003). Al-though the model has been extensively examined anddiscussed, no single experimental system has shownlive-cell measurement of apoplastic pH and H+-ATPaselocalization in vivo. Our experiment uncovered dif-ferential localization of the H+-ATPase protein withinroot regions, in which this protein is subject to accu-mulation in intracellular vesicles that cannot con-tribute to events at the plasma membrane surface.Together with the previous discussion from a study ofhigh-resolution analysis of cell elongation control byauxin and the H+-ATPase in hypocotyls (Fendrychet al., 2016), our result highlights the importance of theexperimental approach for testing a causal relationshipbetween H+-ATPase activation/inactivation and cellgrowth in cell-type specific or single cell levels. Futuredevelopment of a higher resolution imaging assay willallow us to directly examine how individual H+-ATPaseproteins move within the plane of the membrane andoverall, how their function influences cell elongation inplanta.

MATERIALS AND METHODS

Plant Materials

Arabidopsis (Arabidopsis thaliana) mutant plants, aha2-4 and feronia-4 (eco-type Columbia), were previously isolated and published (Haruta and Sussman,2012; Haruta et al., 2014). PEN3-GFP seeds (CS67802) were obtained fromArabidopsis Biological Resource Center (Stein et al., 2006). Arabidopsis plantswere grown at 22°C with constant light and transformed by the floral dipmethod using the Agrobacterium strain, AGL1(Clough and Bent, 1998).

Root Growth Assay

Arabidopsis seeds were plated at approximately 3 mm apart onto a half-strength MS medium (pH 5.7) containing 1% Suc (w/v) and 0.8% agar (w/v).Seeds were stratified at 4°C for 48 h. Plates containing seeds were placed at thevertical orientation in a plastic box with blue translucent color. The entire boxcontaining plates and seeds were placed onto the shelves in a growth incubator(model no. I36LLVL; Percival) at 22°C and illuminated with cool-white light(model no. F20T12 CW, Cool-white, 20W; General Electric). Detailed descrip-tion of the experimental setting for bright (10 mmol m22 s21) or dim (2.2 mmolm22 s21) light conditions were provided in Supplemental Fig. S8A. Light

wavelengths transmitted through the blue colored boxes were measured with aspectrophotometer (SpectroSuite; Ocean Optics), resulting in a spectrum thatwas enriched in blue light ranges over red lights compared with the white light(Supplemental Fig. S8B). The use of the light condition that was enriched withblue light ranges was justified as seedlings grown under this dim blue-enrichedlight condition grew more uniformly compared with those under dim whitelight, which was critical to minimize the variability in root length measure-ments and observe the consistent mCitAHA2 localization patterns.

For a root growth assay with high potassium concentration, seeds weregrown for 3 d at the vertical orientation under the constantwhite light. Seedlingswere aseptically transferred to the control media or the media supplementedwith 100 mM KCl. Plants were grown for an additional 10 d. To quantify rootlength, plates containing plants were scanned with an office scanner and themeasurements were carried out with the software ImageJ (National Institutes ofHealth; Schneider et al., 2012).

Expression Plasmid DNA Construct

For expression of mCitrine-AHA2 translational fusion protein in planta, anAHA2 genomic DNA fragment containing the native promoter, the entire gene,and 39 regulatory sequences was used (Haruta et al., 2010). To produce themCitrine tag insertion site, a restriction site for XbaI was introduced at theamino acid position, Leu-4, using the site-directed mutagenesis technique. Dueto the large size of the plasmid-containing genomic sequence of AHA2 (a totalof approximately 20 kb), to facilitate the molecular cloning procedure, a smallportion of the AHA2 genomic sequence (2602 kb) flanked by two restrictionenzyme sites, StuI and PacI, was subcloned in pCR2.1 (Invitrogen). This pCR-AHA2sp plasmid containing the Leu-4 region was used for introducingthe restriction site XbaI via site-directed mutagenesis using a Change-ITMutagenesis Kit (Affymetrix) and a primer, 59-AGTGGTGAGAGATGTCGAGTCTAGAAGATATCAAGAACGA-39. The resulting AHA2 sequence contain-ing the restriction site XbaI was excised from pCR2.1-AHA2sp plasmid bydigesting with StuI and PacI and used to ligate into the plant expression vectorcarrying the genomic AHA2 sequence, pCAMBIA1200-gAHA2 (Haruta et al.,2010). Thus, the wild-type DNA fragment containing the Leu-4 position wasreplaced with the DNA fragment containing the XbaI restriction site. DNAsequence-encoding mCitrine (a variant of yellow fluorescent protein, YFP) wasproduced by mutagenizing the YFP sequence of pEarleyGate104 [CD3-686;Arabidopsis Biological Resource Center; (Earley et al., 2006)]. To improve thefunction of YFP, two mutations, A206K (monomeric fluorescent protein;von Stetten et al., 2012) and Q69M (photostability; Griesbeck et al., 2001), wereintroduced by site-directed mutagenesis using two primers, 59-/5Phos/CTGAGCTACCAGTCCAAACTGAGCAAAGACCCC-39 and 59-/5Phos/TTCGGCTACGGCCTGATGTGCTTCGCCCGCTA-39. The mCitrine sequenceflanked by XbaI was PCR-amplified using a pair of primers, 59-TCTAGAAATGGTGAGCAAGGGCGA GGAG-39and 59-TCTAGACTCTTGTACAGCTCGTCCATGCC-39, and used to ligate into the pCAMBIA1200-gAHA2 XbaI to produce a construct for expressing mCitrine-AHA2 translationalfusion under the native promoter in planta. Transgenic lines were selected onthe media containing 25 mg/mL hygromycin, and the transformation wasvalidated by PCR-based genotyping with a set of primers amplifying a regioncovering a junction of AHA2 and mCitrine sequences, 59-GAACTTAAAGAGGCAGCAA-39 and 59-TCTAGACTCTTGTACAGCTCGTCCATGCC-39.

A plasmid-carryingDendra2 codingDNAwas obtained fromAddGene (no.57701;McKinney et al., 2009). The Dendra2 sequencewas amplified using a pairof primers: 59-TCTAGAAAACACCCCGGGAATTAACCTG ATCAA-39 and59-TTCTAGACTCCACACCTGGCTGGGCAGGGGGCTGT-39. The amplifiedPCR fragment was cloned into pCAMBIA1200-gAHA2 XbaI. Transgenic lineswere genotyped with a pair of primers, 59-GAACTTAAAGAGGCAGCAA-39and 59-TTCTAGACTCCACACCTGGCTGGGCAGGGGGCTGT-39.

Plasma Membrane Protein Extraction and Immunoblotting

Arabidopsis seeds (10 mg) were grown in a magenta box containing 75 mLMS liquidmedia for 14dunder constant light. Tissues (approximately 45 g)wereground in ahomogenizerwith 150mLextractionbuffer (100mMTris-HClpH7.5,300 mM Suc, 25 mM EDTA, 25 mMNaF, 1 mMNa2MoO4, 1 mM PMSF, 1 mMDTT,100 mM 1,10-phenanthroline) for 90 s at 4°C. The homogenate was filteredthrough two layers of miracloth and spun at 1,000g for 10 min at 4°C. Super-natant was centrifuged at 100,000g for 35 min at 4°C to obtain the microsomalpellet. The pellets were resuspended in suspension buffer (10 mM Tris-HCl,pH7.5, 300 mM Suc, 1 mM EDTA) and subjected to the two-phase partition


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protocol to obtain the plasmamembrane-enriched fraction (Larsson et al., 1987).The upper (PM) and lower (endomembrane) phases were diluted 2-fold with theresuspension buffer and spun at 100,000g for 50 min for 4°C. The pellet wasresuspended in the suspension buffer and stored at -80°C. Protein concentrationwas determined by BCA assay (Pierce).

Soluble or membrane protein samples were adjusted to 15 mg in 30 mL andmixed with 0.1% Triton X-100. Protein samples were mixed with LDS samplebuffer (Novex; Life Technologies) and Sample ReducingAgent (Novex). Proteinsolution was heated at 70°C for 10 min. The protein sample was loaded onto aprotein gel and resolved by running at approximately 125 V for 45 min. Proteinwas transferred to amembrane filter (Immobilon FL;MerckMillipore) using theXCell II Blot module (Invitrogen). Membrane was blocked with an Odysseyblocking buffer (LI-COR) and probedwith primary antibodies. A GFP antibody(Genscript) was diluted to 1:1000 with the blocking buffer and used to reactwith the epitopes. A secondary antibody, goat anti-mouse IgG labeled withIRDye 680RD (LI-COR) was reacted with the blot. Signal was visualized andrecorded using the Odyssey Imaging System (LI-COR).

The identical blot membranes were stained with Ponceau S to visualize theproteins loaded after the detection of signals from the secondary antibody.

Analysis of AHA2 Protein Abundance in Total ProteinExtract from Seedlings

Protein was extracted from seven 10-d-old seedlings grown under bright ordim light conditions. Seedlings were ground in 140 mL 50 mM HEPES buffer(pH7.5) containing 13 cOmplete Protease Inhibitor Cocktail (Roche) and 0.5%Triton X-100. The homogenate was spun at 12,000 rpm for 10min at 4°C and thesupernatant was recovered. Protein concentration was determined by BCAassay. Protein samples (7.5 mg/30 mL) were mixed with the LDS buffer and thereducing agent before heating at 70°C. Proteins were resolved by protein geland mCitAHA2 was detected with a GFP immunoblot, as described above.

CLSM for 3D Imaging of Roots

Confocal microscope imaging was performedwith the LSM780 ELYRA PS.1(Carl Zeiss). Seedlings were submerged in water at the surface of a coverslip(483 60 mm, no. 1 cover glass; Gold Seal). To visualize mCitAHA2 protein, thefluorophore was excited at 514 nm with 2.5% to 10% laser power, and theemission signal was detected at 520 nm to 620 nm. The image was viewed andcaptured, using an objective lens, with an LD C-Apochromat 403/1.1 W KorrM27, unless otherwise noted. 3D imaging was performed by scanning ap-proximately 60 optical slices of an Arabidopsis root, with an interval of ap-proximately 1.06 mm for z-depth range of approximately 68 mm. Detector gainwas typically set as 750 to 800. Before recording images, signal intensity acrossthe entire view was manually inspected to prevent signal saturation. Imageswere acquired using the line sequential mode and an averaging of 4. A stack ofoptically sliced images was compiled into single 3D images using the softwaresZEN or ZEN Lite 2012 (Carl Zeiss) and presented with the maximum intensityprojection protocol. Image was recorded with pinhole size 1.01 Airy Units(1.2 mm section), 512 3 512-pixel sizes, 8-bit depth, 3.15-ms pixel time, 0.03-msline time, and 7.75-s frame time. For the movie, 3D images (.czi file) of Arabi-dopsis roots obtained with the confocal microscope were animated with thesoftware ZEN, exported to a QuickTime movie format (.mov), and edited withthe software Premiere Pro (Adobe). For GFP or Dendra2 detection, the fluo-rophore was excited at 488 nm with a 2.0% laser power and detected at 493 nmto 550 nm. For both mCitrine and GFP detection, wild-type nontransgenicseedlings were also viewed with the identical conditions to ensure that anyfluorescent signals detected with the mCitrine- or GFP plants are not due toendogenous autofluorescence.

Quantitation of Intracellular Components andColocalization Analysis

Images were analyzed with the software Imaris 7.6.1 (Bitplane). All imageswere subjected to background subtraction and Gaussian filtering before surfacerendering with the “Surfaces” tool. Each image was divided into three regionsof interest: meristem (61.28 mm from tip), transition zone (63.77 mm abovemeristem), and elongation zone (86.95 mm above transition zone). Each of theseregions was then subjected to surface rendering using histogram-based auto-matic thresholding. Because intracellular components appear vesicular, thesphericity filterwas applied to the rendered surfaces to select and quantify these

structures. A manually selected sphericity threshold of 0.67 (where a value of1 is a perfect sphere) was applied and kept constant between images. The fol-lowing data were extracted from the statistics tab of Imaris: number of sphericalstructures, fluorescence intensity, and number of disconnected components.

For colocalization analyses, images were similarly subjected to backgroundsubtraction and Gaussian filtering. The image was then segmented into fourregions of interest. “Pearson’s coefficient in dataset volume” for each region wasobtained from the “Coloc” tab of the software Imaris (Bitplane). R2 values werecalculated by obtaining the square of Pearson’s coefficient values (Dunn et al., 2011).

Fluorescent Intensity Profiling of mCitAHA2

Image files (.czi) were processed using the software Fiji (Schindelin et al.,2012). The image files were converted to the .tif format and used for intensityprofiling using custom scripts written in the software language Python. Whencomparing mCitAHA2 levels, the images were normalized by subtracting theminimum intensity from all pixels and then dividing by the maximum pixelintensity. Intensity values range from 0 to 1, with 1 being the maximum in-tensity corresponding to maximummCitAHA2 concentration. The voxels weredivided into two distinct groups by intensity. The root regions that had intra-cellular accumulation of mCitAHA2 signals had reduced intensity because theH+-ATPase was spread over a larger volume. In contrast, mCitAHA2 locali-zation to the membrane concentrated the protein into smaller volume pro-ducing high intensity voxels. Within the images, an intensity ranging from 0.1to 0.35 occurred predominately within the intracellular space, whereas inten-sities greater than 0.35 occurred predominately at the membrane. When sum-ming up the number of voxels along the length of the root, the volume includes8 mm below and above the given location. The number of voxels at a givenintensity level were divided by the total number of voxels above background tocontrol for differences in root size. For the data shown in Figure 2I, quantitationwas performed using three images per condition and presented with themean6 SE. Because the triplicated image analysis produced an intensity profilethat is very similar to the individual images (Fig. 5, D to G; Supplemental Figs.S1 and S3), intensity profiling was performed from one imaging data type percondition as a representation of multiple images (n . 3 per condition) thatshowed similar localization patterns. All the programs developed in this studyare available at theGitHub code repository site (https://github.com/DanBushey/Image-analysis-H- -ATPase-in-Arabidopsis.git).

Root Surface pH Measurement

A 10-d-old seedling grown on a half-strength MS media at vertical positionwas submerged in a 20-mL solution containing 1 mM CaCl2, 1 mM KCl, and3 mg/mL FITC-dextran (average Mr 10,000, FD10s; Sigma-Aldrich) on thesurface of a coverslip. Image was acquired with a 203 objective (Plan-Apochromat 203/0.8 M27), image size 1024 3 1024 pixels, 16-bit depth, av-eraging 16. The FITC-dextran was excited at 488 nmwith 1.8% laser power anddetected at emission wavelengths of 493 nm to 597 nm. Detector gain was set to400. Imageswere recordedwith 0.64-ms pixel time, 0.35-ms line time, and 25.02-sframe time. The pH standard solutions were composed of 10 mM MES, 1 mM

CaCl2, and 1 mM KCl, and adjusted to pH values of 5.0, 5.5, 6.0, or 6.5. FITC-dextran were added to the pH standard solutions at 3 mg/mL immediatelybefore imaging. Images for the pH standard solutions were acquired using theidentical method with the apoplastic pH measurement described above.

The regression analysis was performed to examine and generate a pHstandard curve. A liner fit was obtained with the data points of pH 5.5, 6.0, and6.5. The slope was used to determine the pH of the root surface. A region ofinterest (ROI) including the periphery of the root was generated through anautomated script. Thedyedidnot penetrate the root,which allowed amaximumthreshold to be set that separated the root from the surrounding fluorescentmedium.Themask including the rootwasdilatedbyanarea of 50pixels.AnROIincluding only the periphery was generated by subtracting the dilated maskfrom the original rootmask.Mean intensitywas calculated along the root lengthin an 8-mm-wide area. ROI generation and calculations were performed usingscripts written in the software Python (https://www.python.org/).

Supplemental Data

The following supplemental materials are available.

Supplemental Figure S1. Root growth curve of aha2-4;mCitAHA2 rootsgrown under bright or dim light conditions.







https://www.python.org/



Supplemental Figure S2. Raw root images that are used for mCitAHA2fluorescent intensity profiling.

Supplemental Figure S3. Effect of a series of light intensities on mCitAHA2localization.

Supplemental Figure S4. Localization and intensity profiling ofDendra2AHA2 protein.

Supplemental Figure S5. Segmentation of 3D image obtained with aha2-4;mCitAHA2 root.

Supplemental Figure S6. Calibration of a pH probe, FITC-dextran.

Supplemental Figure S7. Localization of mCitAHA2 and PEN3-GFP in theseedlings grown under dim light.

Supplemental Figure S8. Description of light conditions for seedlinggrowth assay.

Supplemental Movie S1. 3D image of the transition zone of aha2-4;mCitAHA2 root grown under dim light.

ACKNOWLEDGMENTS

We thank Marisa Otegui and Simon Gilroy for microscope instrumentation,Marisa Otegui for assistance in interpreting SDS-PAGE-based observations ofAHA2, and Heather Burch and Shanthi Cambala for technical assistance.Images were recorded at the Newcomb Imaging Center, University ofWisconsin-Madison. M.H thanks Kimberly Toops andMatt Curtis for technicaladvice on image acquisition and data analyses, Robin Davies at MediaLab,UW-Madison for technical assistance with video processing, Brian Thompsonfor assistance with the imaging software, and Edgar Spalding and BrettUnks for technical assistance with the photospectrometer and constructivediscussion, and two anonymous reviewers for constructive feedback to themanuscript.

Received August 14, 2017; accepted October 14, 2017; published November 17,2017.

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