When families in the math group used the appthe least (Bin 0), children with high-math-anxiousparents grew significantly less in math achieve-ment by the end of the school year relative tochildren with low-math-anxious parents ½^b20 ¼−7:94; t ¼ −2:42;P ¼ 0:02� (Fig. 2 andModel S5).Strikingly, using the math appmitigated this neg-ative relation between parents’ math anxiety andchildren’smath achievement. When families usedthe app on average once a week or more, chil-dren with high-math-anxious parents made gainsinmath achievement by the end of the school yearthat didnot significantly differ from thosemadebychildren with low-math-anxious parents {Bin 1:½^b20 ¼ 3:44; t ¼ 1:53;P ¼ 0:13�; Bin 2+: ½^b20 ¼−3:60; t ¼ −1:21;P ¼ 0:23�} (Model S5).Thus,whenparents and children interact about

math story problems—even as little as once aweek—children show increased math achieve-ment by the end of the school year. The benefitsof occasional math-related interactions are es-pecially apparent for children whose parentsare anxious about math. By providing an en-gaging way for math-anxious parents to sharemath with their children, the math app may helpcut the link between parents’ high math anxietyand children’s low math achievement (6).The current findings are of particular relevance

in view of the multibillion-dollar educational appmarket (19). Scant research exists on the effective-ness of apps marketed as educational (20), andthe research that has been done does not alwaysfind benefits for children’s learning. Use of en-hanced e-books that target literacy can actually bedetrimental to children’s basic reading compre-hension when they contain distracting soundsand animations (21). The math app used herehas several specific features that may have con-tributed to its effectiveness. First, it was basic innature (very few sounds, animations, or videos)to avoid distracting elements. Second, it was de-signed to align with the goals of the CommonCore Standards at varying grade levels. Third, itwas designed to be used by parents and childrentogether, based on the known importance of earlyparental input, and specifically parent math talk,for children’s achievement. The app may giveparents—especially high-math-anxious parents oreven parents with less skill or interest in engagingin math—more and better ways to talk to theirchildren about math not only during app usagebut also in other everyday interactions. We haveshown that using this math app enhances thelikelihood that childrenwill succeed inmath, whichis essential for academic success and for the robust-ness of the science, technology, engineering, andmath pipeline.

REFERENCES AND NOTES

1. J. Cannon, H. Ginsburg, Early Educ. Dev. 19, 238–260(2008).

2. E. Duursma, M. Augustyn, B. Zuckerman, Arch. Dis. Child. 93,554–557 (2008).

3. S. C. Levine, L. W. Suriyakham, M. L. Rowe, J. Huttenlocher,E. A. Gunderson, Dev. Psychol. 46, 1309–1319 (2010).

4. A. K. Tekin, Int. J. App. Educ. Studies 11, 1–13 (2011).5. K. Zickuhr, Tablet Ownership 2013 (Pew Research Center,

Washington, DC, 2013); http://pewinternet.org/2013/06/10/tablet-ownership-2013.

6. E. A. Maloney, G. Ramirez, E. A. Gunderson, S. C. Levine,S. L. Beilock, Psychol. Sci. 26, 1480–1488 (2015).

7. B. Butterworth, S. Varma, D. Laurillard, Science 332,1049–1053 (2011).

8. E. A. Gunderson, S. C. Levine, Dev. Sci. 14, 1021–1032(2011).

9. S. C. Levine, K. Ratliff, J. Huttenlocher, J. Cannon, Dev. Psychol.48, 530–542 (2012).

10. S. M. Pruden, S. C. Levine, J. Huttenlocher, Dev. Sci. 14,1417–1430 (2011).

11. B. N. Verdine, C. M. Irwin, R. M. Golinkoff, K. Hirsh-Pasek,J. Exp. Child Psychol. 126, 37–51 (2014).

12. R. Hembree, J. Res. Math. Educ. 21, 33–46 (1990).13. NRC, Mathematics Learning in Early Childhood (National

Academy Press, Washington, DC, 2009).14. R. Woodcock, K. McGrew, N. Mather, Woodcock-Johnson-III

Tests of Achievement (Riverside Publishing, Itasca, IL, 2001).15. S. W. Raudenbush, A. S. Bryk, R. Congdon, HLM 6-Windows

[Computer software] (Scientific Software International, Inc,Skokie, IL, 2004).

16. OECD, PISA 2012 Results: Ready to Learn (Volume III):Students' Engagement, Drive and Self-Beliefs (OECD Publishing,Paris, 2013);

17. C. Eden, A. Heine, A. Jacobs, Psychology 4, 27–35(2013).

18. L. Alexander, C. Martray, Meas. Eval. Couns. Dev. 22, 143–150(1989).

19. Apple Press info, App store sales top $10 billion in 2013(7 January 2014); www.apple.com/pr/library/2014/01/07App-Store-Sales-Top-10-Billion-in-2013.html.

20. K. Hirsh-Pasek et al., Psychol. Sci. Public Interest 16, 3–34(2015).

21. J. Parish-Morris, N. Mahajan, K. Hirsh-Pasek, R. M. Golinkoff,M. F. Collins, Mind Brain Educ. 7, 200–211 (2013).

ACKNOWLEDGMENTS

This work was funded by an Overdeck Family Foundation grant toS.L.B. and S.C.L. The chair, Laura Overdeck, established theBedtime Math Foundation—a nonprofit, 501(c)(3) organization thatproduces the Bedtime Math App. None of the authors have afinancial interest in Bedtime Math. The BLT app, written for AppleiOS8, is freely available on request to S.L.B. The supplementarymaterials contain additional data. The BLT reading and math appcan be accessed from an Apple iPad, iPod, or iPhone at URL appitms-services://?action=download-manifest&url=https://app-builds.uchicago.edu/ios/bedtimemathstudy/BLT.plist; Readingcode: 55dc86e7c9; Math code: 55dc8695c7. We thankS. Raudenbush for providing HLM consultation.

SUPPLEMENTARY MATERIALS

www.sciencemag.org/content/350/6257/196/suppl/DC1Materials and MethodsSupplementary TextFig. S1Table S1Models S1 to S8Accessory Data File S1

5 June 2015; accepted 3 September 201510.1126/science.aac7427

PLANT SCIENCE

Visualization of cellulose synthases inArabidopsis secondary cell wallsY. Watanabe,1,2 M. J. Meents,1,2 L. M. McDonnell,2* S. Barkwill,2 A. Sampathkumar,3

H. N. Cartwright,4 T. Demura,5 D. W. Ehrhardt,4,6 A. L. Samuels,1† S. D. Mansfield2†

Cellulose biosynthesis in plant secondary cell walls forms the basis of vasculardevelopment in land plants, with xylem tissues constituting the vast majority of terrestrialbiomass. We used plant lines that contained an inducible master transcription factorcontrolling xylem cell fate to quantitatively image fluorescently tagged cellulosesynthase enzymes during cellulose deposition in living protoxylem cells. The formation ofsecondary cell wall thickenings was associated with a redistribution and enrichmentof CESA7-containing cellulose synthase complexes (CSCs) into narrow membranedomains. The velocities of secondary cell wall–specific CSCs were faster than those ofprimary cell wall CSCs during abundant cellulose production. Dynamic intracellulartrafficking of endomembranes in combination with increased velocity and high densityof CSCs, enables cellulose to be synthesized rapidly in secondary cell walls.

Cellulose, themost abundant biopolymer onEarth, is a key biomechanical componentof land plants and a valuable natural re-source. Cellulose in the primary cell wall,which is laid down during plant growth,

determines plant shape (1). However, the bulk ofterrestrial biomass is composed of the cellulosein secondary cell walls, which are laid down afterthe cell has stopped growing to strengthen plantvasculature and structure (2). The strength ofthese walls is derived from the organization ofcellulose microfibrils, which, relative to primarycell walls, possess cellulose with a higher degreeof polymerization, increased microfibril crystal-linity, and a higher degree of microfibril organi-zation (2, 3).Cellulose is synthesized at the plasma mem-

brane by cellulose synthase (CESA) enzymes thatare organized in multiprotein cellulose synthasecomplexes (CSCs) (4). In Arabidopsis thaliana,10 CESA isoforms exist, with CESA1, CESA3, andCESA6 involved in primary cell wall synthesis

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1Department of Botany, University of British Columbia,Vancouver, BC, Canada. 2Department of Wood Science,University of British Columbia, Vancouver, BC, Canada. 3Divisionof Biology and Biological Engineering, California Institute ofTechnology, Pasadena, CA, USA. 4Department of Plant Biology,Carnegie Institution for Science, Stanford, CA, USA. 5GraduateSchool of Biological Sciences, Nara Institute of Science andTechnology, Nara, Japan. 6Department of Biological Sciences,Stanford University, Stanford, CA, USA.*Present address: Division of Biological Sciences, University ofCalifornia, San Diego, CA, USA. †Corresponding author. E-mail:annelacey.samuels@botany.ubc.ca (L.S.);shawn.mansfield@ubc.ca (S.D.M.)

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(5, 6), and CESA4, CESA7, and CESA8 requiredfor secondary cell wall production (7). At leastthree distinct isoforms are required for normalcellulose production (5–7). Live-cell imaging ofprimary cell wall CSCs clearly showsCESA velocityand distribution at the plasma membrane as wellas intracellular trafficking (8–10). Secondary cellwalls are produced in vasculature and fibersdeep within plant tissues, limiting the resolutionof live-cell imaging (11, 12). Here, we visualizedcellulose synthesis in secondary cell walls usingfluorescently tagged CESA7 in a unique systemwhere live epidermal cells are induced to formsecondary cell walls ectopically (13).Arabidopsisplantswere engineered to constitu-

tively express a transcription factor controllingxylem tracheary element cell fate, VASCULARNAC-DOMAIN7, fused to a glucocorticoid receptor(VND7::GR). When these plants are exposed to aglucocorticoid hormone such as dexametha-

sone, cells are induced to become protoxylem-like tracheary elements (13). After induction, thesecondary cell wall cellulose synthase genes weretranscriptionally up-regulated by a factor of 1.5 to3.0, whereas primary cell wall–related cellulosesynthase genes were transcriptionally unaffected(table S1). Although it is not possible to knowwhether the VND7::GR levels in these plants arecomparable to endogenous VND7 in protoxylemtracheary elements, the induced cells have featurestypical of developing protoxylem, such as second-ary cell wall thickenings alternating with primarycell wall domains, in spiral or annular patterns.To investigate cellulose deposition, we crossedplants carrying the VND7::GR induction systemwith cesa7/irx3-4 null mutants complementedwith a functional fluorescently taggedCESA7 (YFP::CESA7) driven by its native promoter (fig. S1).Fluorescent signal from CSCs was detectable ininduced epidermal cells, although identification

of individual plasma membrane–localized CSCswas possible only with the use of an optimizedspinning disk confocal imaging system (optimi-zation parameters are described in fig. S2).Use of this optimized system permitted the

visualization of discrete YFP::CESA7 particles inthe plasma membrane of induced protoxylemtracheary elements (Fig. 1). Their linear move-ment at slow and steady velocities meets thecriterion of actively synthesizing CSCs (movie S1)(8–10, 14). CSCs moved in a bidirectional fashionalong plasmamembrane domainsunderlying sec-ondary cell wall thickenings. Relative to labeledprimary cell wall CSCs (8–10), secondary cell wallCSCs showed a higher density at the plasmamem-brane, with overlapping signals from individualparticles largely indistinguishable over extendedareas (Fig. 1, B and C, and fig. S3). Moreover, CSCdistribution changed over the course of second-ary cell wall development. Early in development,

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Fig. 1. CESA7 and microtubules are constrained to secondary cell walldomains during protoxylem development. (A) Early development: YFP::CESA7-labeled CSCs and RFP::TUB6-labeled microtubules localize diffuselyacross the plasma membrane. (B) Mid-development: Narrow microtubulebundles lie underneath tight tracks of membrane-localized CSCs. (C) Latedevelopment: Apparent concentrations of YFP::CESA7 and RFP::TUB6 at

the edges of forming secondary cell walls are revealed as uniform signal insubcortical optical sections. YFP::CESA7-labeled Golgi (arrowhead) are visiblebecause of the increased depth of imaging through secondary cell wall thick-enings. (D) TEM micrographs highlight plasma membrane (PM) curvatureover secondary cell walls (2CW), lined by cortical microtubule (MT) bundles.Scale bars, 10 mm [(A) to (C)], 500 nm (D).

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CESA7 signal was observed across the plane of theplasma membrane on tracks defined by bands ofmicrotubules forming in the cell cortex (Fig. 1A,merged image, and table S2). As secondary cellwall synthesis progressed (Fig. 1B), CESA7 signalwas enriched in regions of the plasmamembraneassociated with tight bundles of microtubules(Fig. 1B and table S2) (15). In late secondary cellwall development (Fig. 1C), the YFP::CESA7 signalwas apparent as U-shaped plasma membranefurrows curving around the secondary cell wallthickenings. The majority of the CESA7 signalwas evenly distributed and restricted to thesecurved domains (Fig. 1C, inset). When VND7::GR-induced cells were cryofixed and examined bytransmission electron microscopy (TEM), tan-gential sections along the cell surface revealedthe curved domain of plasma membrane aroundsecondary cell walls, as well as their associatedmicrotubules in the cell cortex (Fig. 1D).The rate of cell wall deposition is the conse-

quence of both the concentration of CSCs andthe rate at which each CSC produces cellulose.The importance of the restriction of the CSCs tothe curved membrane domains is that cellulosedeposition is concentrated in a discrete area of

intense production. Such concentration may helpto explain why secondary cell walls are synthe-sized more quickly than primary cell walls (16).With live-cell imaging, we measured depositionrates of individual CSCs. We assumed that CSCmovement through the plasma membrane is afunction of glucan chain biosynthesis and itssubsequent crystallization into cellulose micro-fibrils, such that faster CSC velocities would re-flect faster rates of synthesis (3, 17). To facilitatethis comparison, we quantified the velocities ofYFP::CESA7–labeled CSCs at the cell membraneand primary cell wall CSCs (GFP::CESA3) (9, 10)under identical growth and imaging conditions,alternating data collection between plant lines inan imaging session to control for possible en-vironmental effects (Fig. 2). Time projections overa 5-min data collection period showed CSC tracksacross the plasma membrane (Fig. 2, A and B).These tracks defined lines for kymograph analysisof particle velocity. In primary cell wall CSC ky-mographs, the tracks of each GFP::CESA3 weredistinct (Fig. 2A). The tracks of CESA7 were moredifficult to discern because of their higher density(Fig. 2B and fig. S3). The slope of the lines in thekymograph (Fig. 2, A and B, arrows) represents

the CSC velocities, with steeper slopes indicatingfaster CSCmovement (more displacement on thespatial axis). The average velocity of CESA7-containing CSCs producing secondary cell wallswas 265 ± 75 nm/min [mean ± SD,n= 36 cells (40CSCs per cell) from 12 plants] (Fig. 2D). Velocitychanged over the course of protoxylem differen-tiation, which was divided into stages accordingtomicrotubule banding and secondary cell wall fea-tures. AverageCESA7 velocitywas293±29nm/minduring early development, 327 ± 37 nm/min dur-ing mid-development, and 187 ± 38 nm/minduring late development (Fig. 2E). In contrast,primary cell wall CESA3-containingCSCs displayedan average velocity of 231 ± 34 nm/min (n = 12;4 cells from 4 plants; Fig. 2C). Analysis of var-iance (ANOVA) identified significant variationamong conditions (F3,44 = 38.7, P = 2.133 × 10−12).Post hoc Tukey’s pairwise comparison tests in-dicated that primary cell wall CSC velocity wassignificantly different from all stages of sec-ondary cell wall development. Secondary cellwall CSC velocity did not differ significantlybetween early and mid-development, but bothwere significantly different from late develop-ment (P < 0.01; Fig. 2E). This illustrates that

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Fig. 2. Secondary cell wall CSCs have a higher velocity than primary cell wallCSCs during peak cellulose deposition. (A and B) CSC tracks in single frames andtime projections for primary cell wall (A) and secondary cell wall (B) visualized usingGFP::CESA3 and YFP::CESA7, respectively. Kymographs sampled along the yellow linesshow CSC trajectories over time, from which CSC velocity was calculated. Scale bars,10 mm. (C and D) Histograms of GFP::CESA3 (C) and YFP::CESA7 (D) velocities cal-culated from kymograph analysis. (E) Box plot of CSC velocities across stages ofxylem cell development. Means with different letters represent statistically significantdifferences (Tukey’s pairwise comparison, P < 0.01). For each developmental stage, 480CSC velocities were measured from 12 cells in four plants. In (D), 1440 velocities werepooled from all developmental stages. In (E), velocities were averaged for each cellbefore analysis.

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secondary cell wall CSCsmovemore rapidly in theplane of the membrane during peak secondarycell wall synthesis, then slow as the cell nearsmaturity, prior to programmed cell death. Thus,both the high density and high velocity of syn-thesis by individual CSCs contribute to rapidcell wall synthesis during secondary cell wallformation.Previous studies examining fluorescently tagged

secondary cell wall CESAs, visualized throughlayers of tissue to the center of the root, could notresolve CSCs at the plasma membrane, althoughthese authorsmade contributions to understandingthe trafficking of CSCs in intracellular endomem-branes (11, 12). The authors described “CESA7-containing organelles” actively streaming through

the cytoplasm and pausing at domains of sec-ondary cell wall deposition (12). However, becauseof technical limitations, the nature of the organ-elles was not clear. In the induced protoxylemtracheary element system of the VND7::GR lines,we found that the CESA7-containing organelleswere both small CESA-containing compartments(SmaCCs) (10) and Golgi (Fig. 3; red arrows inmovie S2). SmaCCs were both Golgi-independentSmaCCs (blue arrows in movie S3) and Golgi-associated SmaCCs (yellow arrows), and theirbehaviors were most easily identified using fluo-rescence recovery after photobleaching (FRAP)(Fig. 3, A and B, and movies S3 and S4). Afterbleaching the bright plasma membrane–localizedCSC signal, both SmaCCs and Golgi repopulated

the underlying cytoplasm (Fig. 3A). Golgi andSmaCCs often moved in a coordinated fashion(movie S3; 40 ± 15% of SmaCCs were associatedwith Golgi, n = 3 cells), and frequently Golgi andassociated SmaCCs would pause (ranging from15 s to 3 min) at secondary cell wall domains,consistent with previous observations (12).Whenthe Golgi moved on, a CSC signal would persistand split within 70 ± 20 s (n = 8 events; Fig. 3Bandmovie S4). This stationary signal followed byslow steadymovement (280 ± 30 nm/min, n = 16particles) fits the criterion for an insertion eventof multiple CSCs into the plasma membrane, asdefined for primary cell wall CSCs (10). We wereunable to quantify the number of insertion eventsat secondary cell wall domains. However, it was

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Fig. 3. Golgi and SmaCCs densely populate and rapidly deliver CSCsto domains of secondary cell wall formation. (A) Fluorescence recoveryafter photobleaching (FRAP) of YFP::CESA7 in the boxed area, overlying asecondary cell wall thickening. Abundant Golgi-independent SmaCCs (redand blue arrows) rapidly repopulate bleached regions. Additionally, Golgi(arrowhead) and closely associated SmaCCs (yellow arrow) can be seenmoving from one secondary cell wall band and pausing at another. (B)FRAP and kymograph analyses demonstrating insertion at the plasma

membrane of at least two YFP::CESA7-labeled CSCs from a SmaCC (yel-low arrow). After the Golgi (arrowhead) moves away, the signal from theSmaCC splits into two distinct punctae with steady velocities (red andorange arrows). (C) TEM micrographs of cytoplasm around secondary cellwalls showing a diversity of closely associated vesicles. Trans-Golginetworks (arrows), secretory vesicle clusters (arrowheads), and electron-lucent vesicles (asterisks) are indicated. Scale bars, 5 mm (A), 2.5 mm (B),500 nm (C).

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apparent that CSC insertion events were restrictedto thickenings (in 12 photobleached regions of10 mm2 in six cells, no CSC inserted in plasmamembrane regions between thickenings), hencethe targeting of these events is tightly limitedto secondary cell wall domains. TEM of induceddevelopingprotoxylemcells revealed several vesiclepopulations associated with Golgi that may carryCSCs, including trans-Golgi networks, secretoryvesicle clusters, and larger vesicles lacking in-ternal content (Fig. 3C). High-resolution live-cellimaging of CESA7-containing organelles demon-strates the dynamic exchange of CSCs among theGolgi, SmaCCs, and the plasma membrane asso-ciated with microtubule bands.Previous studies of primary cell wall CSCs

showed that disruption of cortical microtubulesdid not affect the rate of insertion of CSCs intothe plasma membrane (10), although CSC distri-bution over the membrane was transiently dis-organized (9, 10). To test the effect of loss ofmicrotubule bundles on secondary cell wall CSCinsertion events, wemeasured the velocities andtrajectories of CESA7-containing CSCs in theVND7::GR induction system after oryzalin treat-ment. In contrast to the dimethyl sulfoxide (DMSO)control (Fig. 4A), where the tracks of CSCs wererestricted to the secondary cell wall bands, theplasma membrane–localized CSCs in oryzalin-treated cells were disorganized (Fig. 4A andmovie S5). These CSC clusters were similar to the“swarms” of primary cell wall CSCs described aftertreatment with an intermediate (8) but not higher(18) concentration of oryzalin. In the absence ofmicrotubule bands, the velocity of secondary cellwall CSCs at the plasmamembranewas unaffected(313 ± 41 nm/min for control, 324 ± 33 nm/min fororyzalin-treated;n= 12 cells from4plants for each;Fig. 4B) while CSC insertion events continued(Fig. 4C and movie S6). Oryzalin treatment alsodid not affect the total cellulose content pro-duced by the differentiating cells (fig. S4). Incontrast, inhibiting cellulose biosynthesis with2,6-dichlorobenzonitrile (DCB) treatment influ-enced only cellulose deposition, while microtu-bule formationwas unaffected (fig. S5). Therefore,the banded microtubule pattern is independentfrom secondary cell wall formation, and althoughmicrotubules are important for the overall sec-ondary cell wall banding pattern, they are notnecessary for CSC delivery from endomembranesto the plasma membrane, nor for reaching peakCSC velocity.We attribute the rapid formation of the sec-

ondary cell wall to both increased concentrationand velocity of CSCs at the plasma membrane.Secondary cell wall CSCs are delivered to thesedomains by populations of Golgi-associated andindependent SmaCCs. These insertion eventsare associated with, but do not require, micro-tubule bundles underlying secondary cell walldomains. These secondary cell walls provide struc-tural support and the water-conducting functionsnecessary for plants to inhabit land. These dataprovide important insights into how land plantsproduce secondary cell walls with the ultrastruc-tural features required for upright growth.

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Fig. 4. Secondary cell wall CSC distribution at the plasma membrane is disorganized by the lossof microtubules after oryzalin treatment while CSC delivery and motility are unaffected. (A)VND7::GR-induced cells expressing YFP::CESA7 and RFP::TUB6 after oryzalin treatment. Plasmamembrane–localized secondary cell wall CSCs follow aberrant tracks in the absence of microtubules. (B) CSCvelocities were not significantly different between oryzalin- and DMSO-treated cells (Student’s t test, P =0.49). For each condition, 40 CSC velocities were averaged for each of 12 cells in four plants. (C) FRAPofYFP::CESA7 signal revealed insertion of CSCs at the plasma membrane after microtubule loss (arrows).Scale bars, 10 mm [(A) and (B)].

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REFERENCES AND NOTES

1. G. O. Wasteneys, Curr. Opin. Plant Biol. 7, 651–660 (2004).2. C. P. Joshi, S. D. Mansfield, Curr. Opin. Plant Biol. 10, 220–226 (2007).3. I. M. Saxena, R. M. Brown Jr., Ann. Bot. 96, 9–21 (2005).4. S. C. Mueller, R. M. Brown Jr., J. Cell Biol. 84, 315–326 (1980).5. T. Desprez et al., Proc. Natl. Acad. Sci. U.S.A. 104,

15572–15577 (2007).6. S. Persson et al., Proc. Natl. Acad. Sci. U.S.A. 104,

15566–15571 (2007).7. N. G. Taylor, R. M. Howells, A. K. Huttly, K. Vickers, S. R. Turner,

Proc. Natl. Acad. Sci. U.S.A. 100, 1450–1455 (2003).8. A. R. Paradez, C. R. Somerville, D. W. Ehrhardt, Science 312,

1491–1495 (2006).9. E. F. Crowell et al., Plant Cell 21, 1141–1154 (2009).10. R. Gutierrez, J. J. Lindeboom, A. R. Paredez, A. M. C. Emons,

D. W. Ehrhardt, Nat. Cell Biol. 11, 797–806 (2009).11. J. C. Gardiner, N. G. Taylor, S. R. Turner, Plant Cell 15,

1740–1748 (2003).12. R. Wightman, S. R. Turner, Plant J. 54, 794–805 (2008).

13. M. Yamaguchi et al., Plant Physiol. 153, 906–914 (2010).14. A. Sampathkumar et al., Plant Physiol. 162, 675–688

(2013).15. C. Ambrose, J. F. Allard, E. N. Cytrynbaum, G. O. Wasteneys,

Nat. Commun. 2, 430 (2011).16. B. Schneider, W. Herth, Protoplasma 131, 142–152 (1986).17. H. E. McFarlane, A. Döring, S. Persson, Annu. Rev. Plant Biol.

65, 69–94 (2014).18. S. DeBolt et al., Proc. Natl. Acad. Sci. U.S.A. 104, 5854–5859

(2007).

ACKNOWLEDGMENTS

Arabidopsis plants containing RFP::TUB6 were the kind gift ofC. Ambrose and G. O. Wasteneys. Supported by CREATE andDiscovery grants (L.S. and S.D.M.); Canadian Natural Sciences andEngineering Research Council doctoral postgraduate scholarships(Y.W., M.J.M., and L.M.M.); NSF grant MCB-1158372 (D.W.E. andH.N.C.); and Japan Society for the Promotion of Science KAKENHIgrants 24114002 and 25291062 (T.D.). We thank the technical

support staff of the UBC Bioimaging Facility and R. White forstatistical advice. The pTA7001 plasmid (VP16-GR vector) isavailable from N.-H. Chua of The Rockefeller University under amaterial transfer agreement; the VND7-GR line is available fromT.D.; and the GFP::CESA3 and YFP::CESA7 VND7-GR lines arefreely available from L.S. or S.D.M. for noncommercial purposesunder a materials transfer agreement on the GVG system withN.-H. Chua. Further data are reported in the supplementarymaterials.

SUPPLEMENTARY MATERIALS

www.sciencemag.org/content/350/6257/198/suppl/DC1Materials and MethodsFigs. S1 to S5Tables S1 and S2Movies S1 to S6References (19–22)

5 June 2015; accepted 26 August 201510.1126/science.aac7446

PLANT SCIENCE

Structure-function analysis identifieshighly sensitive strigolactonereceptors in StrigaShigeo Toh,1 Duncan Holbrook-Smith,1 Peter J. Stogios,2,3 Olena Onopriyenko,2

Shelley Lumba,1 Yuichiro Tsuchiya,4 Alexei Savchenko,2 Peter McCourt1*

Strigolactones are naturally occurring signaling molecules that affect plant development,fungi-plant interactions, and parasitic plant infestations. We characterized the functionof 11 strigolactone receptors from the parasitic plant Striga hermonthica usingchemical and structural biology. We found a clade of polyspecific receptors, includingone that is sensitive to picomolar concentrations of strigolactone. A crystal structureof a highly sensitive strigolactone receptor from Striga revealed a larger binding pocketthan that of the Arabidopsis receptor, which could explain the increased range ofstrigolactone sensitivity.Thus, the sensitivity of Striga to strigolactones from host plants isdriven by receptor sensitivity. By expressing strigolactone receptors in Arabidopsis, wedeveloped a bioassay that can be used to identify chemicals and crops with alteredstrigolactone levels.

Strigolactones are small molecules that actas endogenous plant hormones to influenceplant growth and development (1, 2). Unlikeother plant hormones, multiple forms ofbioactive strigolactones exist and vary in

moieties attached to the A and B rings as wellas the orientation of the D ring (fig. S1) (2). Asidefrom their hormonal role, strigolactones are alsoexuded through roots into the soil, where they actas exogenous signals to attract mycorrhizal fungifor a symbiotic interaction that brings nutrientsto the plant (3). Unfortunately, parasitic plants of

the Striga, Phelipanche, and Orobanche genera alsouse root-emitted strigolactones to locate a nearbyhost (2). Upon perception of strigolactones pro-duced by host plants, seeds of parasitic plantsgerminate. Resulting seedlings attach themselvesto the host to deplete them of nutrients with dev-astating consequences. In sub-Saharan Africaalone, Striga has infested up to two thirds of thearable land and represents the largest challengeto food security on that continent, affecting morethan 100 million people in 25 countries (4). Amechanistic understanding of strigolactone syn-thesis and signaling particularly in Striga couldopen avenues for pest control.Studies from model plants suggest that a fam-

ily of related a/b hydrolases is central to strigo-lactone perception. For example, D14 (DWARF14)is a strigolactone receptor important for shootbranching in rice (5–7). A related a/b hydrolasegene, HYPOSENSITIVE TO LIGHT/KARRIKININSENSITIVE2 (HTL/KAI2), herein designatedHTL, has been shown to play a role in Arabidopsisgermination (8). Because Striga requires strigolac-

tones to germinate, by inference HTL homologsmay have similar roles in parasitic plant species.ArabidopsisHTL, however, binds a smoke-derivedgermination stimulant, karrikin (9). In contrast,Striga does not germinate in response to karrikinbut is sensitive to picomolar concentrations ofstrigolactones (10). Moreover, Striga appears to dif-ferentiate among hosts by sensing different com-binations of strigolactones (10). Because Strigahermonthica (Striga) is an obligate and outcrossinghemiparasite, the functional roles of Striga HTL/KAI2homologs (ShHTLs) in strigolactone perceptionare difficult to elucidate. Here, we describe theroles of ShHTLs in strigolactone perception.Phylogenetic analysis of public databases led

us to identify a nested clade of 11 HTL homologsin Striga (Fig. 1A). To analyze the function ofthese ShHTL genes, we expressed each of the11 ShHTLs individually in an Arabidopsis loss-of-functionHTL/KAI2mutant (htl-3) backgroundand assayed for germination phenotypes. Seeds ofthe Arabidopsis ecotype Columbia germinatepoorly after exposure to high temperature, butthis thermoinhibition is alleviated by strigolac-tone addition (11). The strigolactone-dependentrescue of thermoinhibited seeds requires a func-tional AtHTL gene, which provided the basisfor introducing ShHTL genes into an htl-3 ge-netic background. We monitored the functionalactivity of these genes by assaying for suppres-sion of thermoinhibited seeds. Thermoinhibitedhtl-3 seed constitutively overexpressing AtHTLdriven by the 35S promoter germinated uponaddition of a synthetic strigolactone, GR24 (fig.S2). Similarly, thermoinhibited htl-3 seed con-taining different ShHTL transgenes germinatedon GR24 to varying degrees, depending on thetransgene (fig. S2). To compare lines, we de-termined the effective concentration of GR24required for 50% germination (EC50) of transgen-ic seed (fig. S3). The EC50 of a transgenic lineoverexpressing AtHTL is lower than that of thewild type, indicating increased strigolactone re-sponsiveness (Fig. 1B). In contrast, ShHTL1,ShHTL2, and ShHTL3 expression lines were lessresponsive to GR24 compared with either wildtype or misexpressed AtHTL seed (Fig. 1B).ShHTL8 and ShHTL9 transgenics were less

SCIENCE sciencemag.org 9 OCTOBER 2015 • VOL 350 ISSUE 6257 203

1Cell and Systems Biology, University of Toronto, 25Willcocks Street, Toronto M5S 3B2, Canada. 2Department ofChemical Engineering and Applied Chemistry, Banting andBest Department of Medical Research, University of Toronto,200 College Street, Toronto M5S 3E5, Canada. 3Center forStructural Genomics of Infectious Diseases, contracted byNational Institute of Allergy and Infectious Diseases, NationalInstitutes of Health, Bethesda, MD, USA. 4Institute ofTransformative Bio-Molecules, Nagoya University, Japan,Furo-cho, Chikusa-ku, Nagoya, Aichi 464-8602, Japan.*Corresponding author. E-mail: peter.mccourt@utoronto.ca

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secondary cell wallsArabidopsisVisualization of cellulose synthases in

Samuels and S. D. MansfieldY. Watanabe, M. J. Meents, L. M. McDonnell, S. Barkwill, A. Sampathkumar, H. N. Cartwright, T. Demura, D. W. Ehrhardt, A.L.

DOI: 10.1126/science.aac7446 (6257), 198-203.350Science 

, this issue p. 198, see also p. 156Scienceare built faster than primary cell walls, perhaps due to increased velocity and density of cellulose synthase complexes.Schneider and Persson). Combining this improved accessibility with fluorescent tagging showed that secondary cell wallsexpression to bring xylem-style cellulose synthase activity to the epidermal surface of the plant (see the Perspective by

leveraged ectopicet al.the sturdiness of wood, is formed by xylem cells embedded in the core of the plant. Watanabe synthesized outside the cell membrane by cellulose synthase enzymes. Much of the secondary cell wall, responsible for

Plant cell walls provide the cellulose that is integral for wood, cotton fiber, and many biofuels. Cellulose isSecondary cell walls built with speed

ARTICLE TOOLS http://science.sciencemag.org/content/350/6257/198

MATERIALSSUPPLEMENTARY http://science.sciencemag.org/content/suppl/2015/10/07/350.6257.198.DC1

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