Mechanism of CAP1-mediated apical actinpolymerization in pollen tubesYuxiang Jianga,b,1, Ming Chang (常明)a,1, Yaxian Lana,1, and Shanjin Huanga,2

aCenter for Plant Biology, School of Life Sciences, Tsinghua University, 100084 Beijing, China; and bTsinghua-Peking Center for Life Sciences, 100084 Beijing,China

Edited by Ram Dixit, Washington University, St. Louis, MO, and accepted by Editorial Board Member June B. Nasrallah April 25, 2019 (received for reviewDecember 20, 2018)

Srv2p/CAP1 is an essential regulator of actin turnover, but its exactfunction in regulating actin polymerization, particularly the con-tribution of its actin nucleotide exchange activity, remains in-completely understood. We found that, although ArabidopsisCAP1 is distributed uniformly in the cytoplasm, its loss of functionhas differential effects on the actin cytoskeleton within differentregions of the pollen tube. Specifically, the F-actin level increasesin the shank but decreases in the apical region of cap1 pollentubes. The reduction in apical F-actin results mainly from impairedpolymerization of membrane-originated actin within cap1 pollentubes. The actin nucleotide exchange activity of CAP1 is involvedin apical actin polymerization. CAP1 acts synergistically with pol-len ADF and profilin to promote actin turnover in vitro, and it canovercome the inhibitory effects of ADF and synergize with profilinto promote actin nucleotide exchange. Consistent with its role as ashuttle molecule between ADF and profilin, the cytosolic concen-tration of CAP1 is much lower than that of ADF and profilin in pollen.Thus, CAP1 synergizes with ADF and profilin to drive actin turnoverin pollen and promote apical actin polymerization in pollen tubes ina manner that involves its actin nucleotide exchange activity.

pollen tube | apical actin structure | actin dynamics | CAP1 |actin nucleotide exchange

The actin cytoskeleton has been implicated in numerous fun-damental physiological cellular processes (1). Most actin-

based functions, if not all, are carried out by the polymericform of actin (2). Therefore, a central question in this field ishow actin monomers can rapidly assemble into actin filamentsand organize into distinct structures to meet the demands of var-ious physiological and cellular processes. A pool of polymerization-competent actin monomers must be available to support rapid andsustainable actin polymerization. ADF/cofilin is a central player indriving actin assembly and disassembly, and it preferentially bindsto ADP-G-actin with high affinity and inhibits its nucleotide ex-change (3, 4). Consequently, the dissociated actin monomersshould be in the form of ADP-G-actin-ADF. Therefore, dissoci-ation of G-actin from ADF and reloading of G-actin with ATP isthe key to drive actin assembly.CAP, also known as Srv2p in budding yeast, has been

emerging as an important player in this process. It was originallyidentified as an adenylyl cyclase-associated protein (5, 6). It wasproposed that Srv2p/CAP releases actin monomers from theADF-ADP-G-actin complex by transferring actin monomers toprofilin and subsequently delivering actin subunits to the barbedend of actin filaments (7). To date, there is overwhelming evi-dence supporting the role of Srv2p/CAP in promoting actinturnover both in vitro and in vivo via coordination with ADF/cofilin in different organisms (8–14). The synergistic action be-tween Srv2p/CAP1 and ADF/cofilin enables the release of actinmonomers from ADF, which allows us to speculate that CAP1 willbe required for actin polymerization in cells. Indeed, it was shownthat loss of function of CAS-1, a Caenorhabditis elegans cyclase-associated protein, impaired sarcomeric actin assembly in striatedmuscle (15), but the underlying molecular details remain to be

uncovered. To date, direct evidence supporting the role of CAP1 inpromoting actin polymerization is rather limited in different or-ganisms, which is likely because loss of function of CAP1 causesdramatic defects in actin turnover, which result in increased actinbundling and F-actin levels in cells. To some extent, this explainswhy the actin turnover function of CAP1 has been studied muchmore extensively than its polymerization function. However, someaspects of CAP1 function in actin polymerization—in particular,the contribution of its actin nucleotide exchange activity to thisprocess—remain largely unknown.Pollen tube growth depends on a dynamic actin cytoskeleton

(16–19). Earlier studies suggest that there exists a population ofdynamic actin filaments, and their polymerization is crucial forrapid pollen tube growth (20, 21). Live-cell imaging of actindynamics within the growth domain of pollen tubes revealed thatactin filaments polymerize actively from the apical membrane,and this process is required for and concurrent with pollen tubegrowth (22). Based on the observations that the formins arerequired for apical actin polymerization (22–24) and that loss offunction of profilins impairs apical actin polymerization (25), itwas proposed that formin/profilin module-mediated actin poly-merization is one of the major actin polymerization pathwayswithin the growth domain of pollen tubes. Given that actin ispredicted to be buffered by an equal amount of profilin in pollen(16, 26), continuous generation and maintenance of a pool ofpolymerization-competent ATP-G-actin-profilin complexes withinthe cytoplasm at the pollen tube tip is central to this hypothesis. Incontrast to nonplant profilins, which enhance nucleotide exchange

Significance

Actin polymerization drives rapid polarized pollen tube growth,but the mechanism underlying actin polymerization within thegrowth domain of pollen tubes remains incompletely un-derstood. We here identify CAP1 as a major player in drivingactin polymerization via recharging ADP-G-actin and facilitatingthe formation and maintenance of a pool of polymerization-competent actin monomers in a manner that involves the nu-cleotide exchange activity of CAP1 in pollen tubes. Our studydirectly links the actin nucleotide exchange activity of CAP1 to itsrole in promoting actin polymerization. Our study thus signifi-cantly enhances our understanding of the mechanism of actinpolymerization in pollen tubes.

Author contributions: Y.J., M.C., and S.H. designed research; Y.J., M.C., and Y.L. per-formed research; Y.J., M.C., Y.L., and S.H. analyzed data; and Y.J., M.C., and S.H. wrotethe paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission. R.D. is a guest editor invited by the Editorial Board.

Published under the PNAS license.1Y.J., M.C., and Y.L. contributed equally to this work.2To whom correspondence may be addressed. Email: sjhuang@tsinghua.edu.cn.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1821639116/-/DCSupplemental.

Published online May 23, 2019.

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on actin (27–29), plant profilins inhibit actin nucleotide exchange(30, 31) or weakly enhance actin nucleotide exchange (12). In thisregard, recharging ADP-G-actin, presumably complexed withADF, before transferring it to profilin to form an ATP-G-actin-profilin complex, is crucial for actin polymerization. ArabidopsisCAP1 is the likely candidate to take on this role since it enhancesnucleotide exchange on actin (12). Several CAP1 proteins fromother organisms also harbor nucleotide exchange activities foractin (32–34). To date, however, the linkage between the activityof CAP1/Srv2p in stimulating actin nucleotide exchange and actinpolymerization is rarely established.By analyzing the function of CAP1 in regulating actin dy-

namics in pollen tubes, we here demonstrate unambiguously thatCAP1 is required for the polymerization of actin filaments fromthe plasma membrane (PM) within the growth domain of pollentubes. Our findings reveal that the reduction in the amount ofapical actin filaments in cap1 pollen tubes results mainly fromimpaired de novo actin polymerization and demonstrate that theactivity of CAP1 in stimulating actin nucleotide exchange is re-quired for its function in driving actin polymerization. We thuspropose that CAP1 acts in concert with ADF and profilin to driveactin turnover in pollen and consequently facilitates the formation ofATP-G-actin-profilin complexes to feed membrane-anchored forminsto drive apical actin polymerization and normal pollen tube growth.

ResultsArabidopsis CAP1 Promotes ADF-Mediated Actin Depolymerizationand Severing via Its N Terminus in Vitro. To understand the mech-anism by which CAP1 regulates actin assembly and disassembly inpollen, we examined its coordination with pollen ADF and pro-filin. Sequence alignment showed that Arabidopsis CAP1 harborsthe conserved N-terminal residues which are crucial for the co-ordination of CAP1/Srv2p with ADF/cofilin (35) (SI Appendix, Fig.S1). Consistent with previous findings that ArabidopsisCAP1 enhances actin nucleotide exchange (12), it also containsthe conserved residues that are crucial for the actin nucleotideexchange activity (36) (SI Appendix, Fig. S1). We therefore gen-erated three recombinant proteins, CAP1, CAP1-90, and CAP1-109 (Fig. 1 A and B), among which CAP1-90 (35) and CAP1-109(36) have mutations in the residues that are crucial for ADF/cofilininteraction and the actin nucleotide exchange activity, respectively.We then examined whether CAP1 and CAP1-90 act synergisticallywith pollen ADFs. Using high-speed F-actin cosedimentation as-says, we found that CAP1 promotes pollen-specific ADF7- andADF10-mediated actin depolymerization whereas CAP1-90 hasreduced activity (Fig. 1 C–E). Taking ADF10 as the representa-tive pollen ADF, we demonstrated by total internal reflectionfluorescence microscopy that Arabidopsis CAP1 promotes ADF10-mediated filament fragmentation; again CAP1-90 has reducedactivity (Fig. 1 F and G). Our data thus suggest that ArabidopsisCAP1 is able to enhance pollen ADF-mediated actin de-polymerization and severing via a conserved mechanism.

Synergy Between CAP1 and ADF Promotes Actin Turnover in Vivo. Toreveal the function of CAP1 in pollen, we characterized two loss-of-function T-DNA insertion mutants of CAP1 (8) SI Appendix,Fig. S2 A and B). Consistent with previous results (8), we foundthat pollen germination and pollen tube growth were signifi-cantly inhibited in cap1 mutants compared with WT (SI Appen-dix, Fig. S2 C–E). In addition, we found that the morphology ofpollen tubes derived from cap1 mutants was dramatically dif-ferent from WT (SI Appendix, Fig. S2C), as evidenced by sig-nificant increases in the width of pollen tubes (SI Appendix, Fig.S2F). The developmental defects and pollen tube growth defectsin Arabidopsis can be rescued by transformation of CAP1 orCAP1-EGFP under the control of its own promoter (SI Appen-dix, Fig. S2 G–K). Interestingly, we found that overexpression ofCAP1 (SI Appendix, Fig. S2L) slightly but significantly inhibited

pollen germination (SI Appendix, Fig. S2M) whereas it enhancedpollen tube growth (SI Appendix, Fig. S2N). Thus, these datatogether suggest that misexpression of CAP1 impairs pollengermination and alters polarized pollen tube growth.To determine the effect of misexpression of CAP1 on the actin

cytoskeleton in vivo, we initially examined the actin cytoskeletonin pollen grains. We found that the amount of actin filamentsand the width of filamentous structures increased in cap1 pollengrains (SI Appendix, Fig. S3 A–C) but decreased in CAP1 over-expression pollen grains (SI Appendix, Fig. S3 D–F). These datasuggested that CAP1 promotes actin turnover in vivo. Strikingly,we found that introduction of CAP1-90 into cap1 mutants (SIAppendix, Fig. S3G) failed to complement the increased amountand bundling of actin filaments in cap1 mutant pollen grains (SIAppendix, Fig. S3 H–J) and the pollen tube growth phenotype incap1 mutant pollen tubes (SI Appendix, Fig. S3K). These datatogether suggest that CAP1 promotes actin turnover in a mannerthat requires its synergy with ADF in vivo.

Loss of Function of CAP1 Has Differential Effects on the F-Actin LevelWithin Different Regions of the Pollen Tube. Next, we examined theeffect of the misexpression of CAP1 on the actin cytoskeleton inpollen tubes. We initially found that actin filaments becamebrighter and more heavily bundled and that the amount of actinfilaments increased in the shank region of cap1 pollen tubes (Fig.2 A and B). Accordingly, we found that overexpression of CAP1causes a reduction in the amount of F-actin throughout theentire pollen tube (SI Appendix, Fig. S4). Surprisingly, we foundthat actin filaments became dimmer and that the amount of actinfilaments decreased within the apical region of cap1 pollen tubescompared with WT pollen tubes (Fig. 2 A and B). The resultssuggest that loss of function of CAP1 has differential effects onactin dynamics within different regions of the pollen tube.

CAP1 Is Uniformly Distributed in Pollen Tubes. We next askedwhether the differential effect of CAP1 on actin dynamics withindifferent regions of the pollen tube results from the distinct in-tracellular localization of CAP1 in the pollen tube. The abovedata show that introduction of a CAP1-EGFP fusion constructdriven by the native CAP1 promoter into cap1 mutants fullyrescued the developmental defects and pollen tube growth de-fects in cap1-1 mutants (SI Appendix, Fig. S2 I and K), whichsuggests that CAP1-EGFP can act as a faithful probe to indicatethe intracellular localization of CAP1. We found that CAP1 hasa uniform distribution and does not form obvious filamentousstructures in the cytoplasm of pollen grains and pollen tubes (Fig.2 C and D). This is consistent with the fact that CAP1 acts as anactin monomer-binding protein. Thus, our data suggest thatCAP1 is distributed uniformly in the cytoplasm of the pollen tube.

Actin Dynamics Is Reduced in cap1 Pollen Tubes and Increased in CAP1Overexpression Pollen Tubes. To further reveal the defect of actindynamics in cap1 pollen tubes, we performed real-time visuali-zation of the dynamics of actin filaments decorated with Lifeact-eGFP as described previously (37, 38). Similar to the findingsshown above (Fig. 2A), we found that actin filaments becameheavily bundled in the shank region of cap1 pollen tubes com-pared with WT pollen tubes (Fig. 3A). We found that the dy-namics of actin filaments are reduced substantially (SI Appendix,Fig. S5A) and that the filaments are arranged into heavy butcurved bundle structures in the shank region of the cap1 pollentube (SI Appendix, Fig. S5 A and B). This was also supported bymeasurements showing that the filament elongation rate, fila-ment shortening rate, and filament severing frequency are sig-nificantly reduced in the shank region of cap1 mutant pollentubes compared with WT (SI Appendix, Fig. S5 C–E). Next, wevisualized the dynamics of actin filaments within the apical regionof the pollen tube. As reported previously, we found that actin

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filaments continuously polymerize from the apical membrane inWT pollen tubes (Fig. 3 A, Upper, and Movie S1). By comparison,apical membrane-originated actin polymerization was severelycompromised in cap1 pollen tubes, leading to impaired formationof the apical actin structure (Fig. 3 A, Lower, and Movie S2). Aftertracking individual membrane-originated actin filaments (Fig.3B), we found that, consistent with the findings shown for shank-localized actin filaments (SI Appendix, Fig. S5), their depolymer-ization rates and severing frequencies are reduced, which conse-quently leads to an increase in the maximal filament lifetime incap1 pollen tubes compared with WT pollen tubes (Fig. 3C). Thus,we showed that cap1 pollen tubes exhibit a similar reduction inactin dynamics within both apical and shank regions, but theoutcomes are different in the two regions in terms of the amountof actin filaments. This allows us to speculate that the decrease inthe amount of apical actin filaments is due to the defect in de novoactin polymerization within the growth domain of cap1 pollentubes. In support of this speculation, we found that the elongation

rate of actin filaments is reduced in cap1 pollen tubes comparedwith WT (Fig. 3C).Accordingly, we found that the rate of actin elongation in-

creased in CAP1 overexpression pollen tubes compared with WT(SI Appendix, Fig. S6 A and B). However, the amount of actinfilaments polymerized from the apical membrane appears lessoverall in CAP1 overexpression pollen tubes than in WT (SIAppendix, Fig. S6A and Movies S3 and S4). We showed abovethat CAP1 promotes ADF-mediated actin depolymerization invitro (Fig. 1 C–G) and enhances actin turnover in vivo (SI Ap-pendix, Fig. S3 D–F). Therefore, we speculated that the re-duction in the amount of apical membrane-originated actinfilaments in CAP1 overexpression pollen tubes is likely due tothe up-regulation in actin depolymerization. In support of thisspeculation, we found that depolymerization rates and severingfrequencies of membrane-originated actin filaments increased,leading to the reduced maximal filament lifetime and filamentlength in CAP1 overexpression pollen tubes compared with WT(SI Appendix, Fig. S6B). Our data suggest that misexpression of

Fig. 1. Arabidopsis CAP1 enhances pollen ADF-mediated actin depolymerization and severing in vitro via its N terminus. (A) Schematic domain organizationof CAP1 and the mutant constructs used in this study. CC, coiled coil domain; HFD, helically folded domain; P1 and P2, proline-rich region 1 and 2; WH2, WASP-homology 2 domain. The red and blue dots in the HFD domain and β-sheet indicate the amino acid residues that were mutated in CAP1-90 and CAP1-109,respectively. (B) Purified Arabidopsis CAP1 recombinant protein and its variants. (C) SDS/PAGE gels of the high-speed F-actin cosedimentation experiments.F-actin, 3 μM; ADF7, 20 μM; CAP1 or CAP1-90, 3 μM. S, supernatant; P, pellet. Gels in B and C were stained with Coomassie Brilliant Blue. (D and E) Quan-tification of the amount of actin in the supernatant from the high-speed F-actin cosedimentation experiments. F-actin, 3 μM; ADF7 or ADF10, 20 μM; CAP1 orCAP1-90, 3 μM. Data are presented as mean ± SD from three independent experiments. Statistical comparisons were performed using ANOVA with a post hocTukey test; different letters indicate significant differences, P < 0.01. (F) Time-lapse images of actin filaments assembled from Oregon-green (OG) actin. OG-actin, 1 μM, 50% labeled; ADF10, 0.3 μM; CAP1 or CAP1-90, 1 μM. The pH of the buffer is 7.0. Red scissors indicate filament fragmentation events. (Scale bar,5 μm.) (G) Quantification of actin filament-severing frequencies. The data are presented as mean ± SEM. More than 40 actin filaments were counted fromthree independent experiments. Statistical comparisons were performed using ANOVA with a post hoc Tukey test; different letters indicate significantdifferences, P < 0.01. For ADF10 vs. ADF10+CAP1-90, P = 0.85.

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CAP1 affects both actin polymerization and depolymerization,but the final outcome in terms of the amount of actin filamentswithin cells depends on the relative change in actin polymeri-zation and depolymerization.

CAP1 Is an Abundant Cellular Protein in Pollen That Synergizes withPollen ADF and Profilin to Promote Actin Turnover and Actin NucleotideExchange in Vitro. To gain insights into the biochemical basis of theaction of CAP1 in pollen cells, we sought to determine the cellularconcentration of CAP1 as well as its molar ratio relationship withactin and two other proteins involved in actin turnover, ADF andprofilin. Consistent with previous measurements (20, 39, 40), wefound that the molar ratio of total actin to profilin is roughly 1:1 inArabidopsis pollen (Fig. 4A). The cellular concentration of Ara-bidopsis CAP1 is about 1.2 μM in pollen, which corresponds to amolar ratio of roughly 1:25 to actin or profilin (Fig. 4A). Thissuggests that the actin monomer-sequestering activity is contrib-uted mainly by profilin in pollen. In addition, the concentration ofCAP1 is about one-fourth that of ADF (Fig. 4A), which is actuallyconsistent with the notion that CAP1 acts as an intermediateplayer by transferring actin monomers from ADF to profilin in theactin turnover machinery as proposed previously (7). Indeed, wefound that CAP1 substantially enhances actin turnover in thepresence of pollen-specific ADF10 (41) and PRF5 (25) (Fig. 4B).In addition, we found that CAP1 is able to overcome the in-hibitory effect of ADF7 on ADP-G-actin nucleotide exchange

(Fig. 4C). Strikingly, we found that the role of CAP1 in promotingADP-G-actin nucleotide exchange is much more potent than thatof profilin, as 50 nM CAP1 substantially enhanced the rate ofADP-G-actin nucleotide exchange, whereas 4 μM PRF5 onlyslightly enhanced it (Fig. 4D). Our study, along with previousobservations (12, 30, 31), suggests that the role of rechargingADP-G-actin will be taken over by CAP1 rather than profilin invivo. Interestingly, we found that profilin and CAP1 synergisticallyenhance ADP-G-actin nucleotide exchange (Fig. 4D). Thesynergy between profilin and CAP1 in promoting ADP-G-actinnucleotide exchange is consistent with the fact that CAP1 re-tains the conserved PP1 and PP2 motifs (SI Appendix, Fig. S1),as the PP1 motif of Srv2p was previously demonstrated to in-teract with profilin (42). Therefore, our results showed thatCAP1 is an abundant cytosolic protein that is distributed uni-formly in pollen cells and acts synergistically with pollen ADFand profilin.

Both CAP1-109 and CAP1-90 Fail to Rescue the F-Actin ReductionPhenotype at cap1 Pollen Tube Tips. We next introduced CAP1-109 (Fig. 1A) into cap1 mutants, as CAP1-109 was demonstratedto lack actin nucleotide exchange activity (36). We initiallydemonstrated that CAP1-109 retains the capability to enhanceADF-mediated actin depolymerization in vitro and promoteactin turnover in pollen grains (SI Appendix, Fig. S7 A–F). Wefound that CAP1-109 retains the G-actin–binding activity, as

Fig. 2. CAP1 is distributed throughout the cytoplasm of pollen cells whereas loss of function of CAP1 has differential effects on the actin cytoskeleton withindifferent regions of pollen tubes. (A) Micrographs of actin filaments stained with Alexa-Fluor-488–phalloidin in pollen tubes. The projection image or selectedoptical sections are shown. The dashed red lines indicate the apical region (0–10 μm from the tip). (Lower) Transverse sections at the indicated positions fromthe tip of pollen tubes (the distance from the tip is indicated on each image). (Right) The 3D distribution of fluorescence intensity of actin filaments within theapical region, which was generated by ImageJ software with a 3D interactive “Surface Plot” function. Warm and cold colors indicate higher and lowerfluorescence, respectively. (Scale bar, 5 μm.) (B) Quantification of the fluorescence intensity of Alexa-Fluor-488–phalloidin staining from the tip to the basealong the growth axis of pollen tubes. The dashed red line indicates the apical region as shown in A. Data are presented as mean ± SEM. More than 15 pollentubes were measured. (C) Subcellular localization of CAP1-EGFP in a pollen grain and pollen tubes of different lengths. Pollen derived from cap1 plantsexpressing CAP1pro:CAP1-EGFP was visualized. The projection images and selected optical sections are shown. (Scale bar, 10 μm.) (D) Plot of the fluorescenceintensity of CAP1-EGFP and EGFP (as control) in the pollen tube. More than 15 pollen tubes were measured. Data are presented as mean ± SEM.

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evidenced by data showing that CAP1-109 inhibits spontaneousactin polymerization and binds to ADP-G-actin as effectively asCAP1 and CAP1-90 (SI Appendix, Fig. S8 A–C). However, wefound that the ability of CAP1-109 to promote actin nucleotideexchange is substantially compromised when it is compared withCAP1 and CAP1-90 (SI Appendix, Fig. S8D). Consistent withthis, we found that introduction of both CAP1 and CAP1-109reduced the amount of actin filaments and bundling in the shank

of cap1 pollen tubes (Fig. 5 A and B), which suggests that theyretain the capability to enhance actin turnover in vivo. However,CAP1-109 cannot complement the reduction in the amount ofapical actin filaments in cap1 pollen tubes (Fig. 5 A–C) or thepollen tube growth defect (SI Appendix, Fig. S7G). Further ob-servations revealed that apical actin polymerization is impairedin CAP1-109 pollen tubes compared with WT pollen tubes (Fig.5D and Movies S5 and S6), as the elongation rate of membrane-originated actin filaments is significantly reduced in CAP1-109pollen tubes (Fig. 5E). Surprisingly, the depolymerization rateof apical actin filaments is reduced significantly in CAP1-109pollen tubes compared with WT pollen tubes. This reduceddepolymerization rate leads to an increase in the maximal fila-ment length and lifetime (Fig. 5E). We do not currently know thereason for this. It might be due to a feedback regulatory mech-anism to counteract the reduction in the amount of apical actinfilaments. Nonetheless, our study suggests that the activity ofCAP1 in stimulating actin nucleotide exchange is crucial for itsrole in regulating apical actin polymerization.To determine whether CAP1 mainly promotes the conversion

of ADP-G-actin bound with ADF into ATP-G-actin to facilitatemembrane-originated actin polymerization, we examined theactin cytoskeleton in pollen tubes derived from CAP1-90 plants.We found that apical actin polymerization is impaired in theCAP1-90 pollen tube, and this causes a reduction in the amountof apical actin filaments compared with WT (Fig. 6 A and B andMovies S7 and S8). This reduction was directly visualized intransverse sections derived from the apical region of the pollentube (Fig. 6C). Surprisingly, we found that the amount of actinfilaments increased in transverse sections derived from the shankregion of the CAP1-90 pollen tube compared with WT (Fig. 6C).Notably, heavy actin cables are more prominent in the shank ofthe CAP1-90 pollen tube compared with WT (Fig. 6C). This isconsistent with the above data showing the reduced rate of actinturnover in CAP1-90 pollen grains (SI Appendix, Fig. S3 H–J).Indeed, we found that actin depolymerization rate and severingfrequency of membrane-originated actin filaments are reducedand that their lifetime is consequently increased in the CAP1-90pollen tube compared with WT (Fig. 6D). These data suggestthat the reduction in the amount of apical actin filaments inCAP1-90 pollen tubes results from the impaired actin poly-merization. In support of this notion, we found that the elon-gation rate of membrane-originated actin filaments is reducedsignificantly in the CAP1-90 pollen tubes compared with WT(Fig. 6D). These data suggest that the coordination ofCAP1 with ADF is crucial for CAP1 in promoting apicalmembrane-originated actin polymerization in pollen tubes,which implies that CAP1 mainly recharges ADP-G-actin boundwith ADF in cells.

DiscussionWe initiated the functional characterization of Arabidopsis CAP1in pollen with the aim of understanding the regulation of actinpolymerization within the growth domain of pollen tubes. Wefound that actin is almost equimolar to profilin in pollen (Fig.4A), consistent with previous measurements (20, 39, 40). Theseprevious findings led to the conclusion that actin monomers existmainly in the form of actin-profilin complexes (16, 26, 43). Inaddition, it was previously shown that the formin/profilin moduleplays an essential role in driving actin polymerization from themembrane within the growth domain of the pollen tube (22–25).This urges us to ask how a pool of polymerization-competentATP-actin-profilin complexes is generated and maintained. Incontrast to the situation in nonplant systems, where profilins canenhance actin nucleotide exchange (27–29), plant profilins lack orinhibit actin nucleotide exchange (30, 31). In this regard,recharging of ADP-G-actin before the formation of ATP-G-actin-profilin complexes should be critical. ADF is a very abundant

Fig. 3. Actin polymerization is impaired within the apical region of cap1pollen tubes. (A) Time-lapse images of actin filaments in growing pollentubes of WT and cap1. Actin filaments were decorated with Lifeact-EGFP.See the entire series in Movies S1 and S2. The red boxes indicate the apicalregion of the pollen tube, and red brackets mark the region with brightapical actin filaments originating from the apical membrane. (Right) Ky-mograph analysis of the Left images: the fluorescence intensity of Lifeact-EGFP indicates the amount of actin filaments in the pollen tube (from tip toshank region along the longitudinal axis) during the growth process. Redarrows indicate the bright apical F-actin polymerized from the membrane.(Scale bars, 10 μm.) (B) Time-lapse images of actin filaments within the apicalregion of pollen tubes. The far Left panels show the pseudotricoloredoverlayered images of WT and cap1 pollen tubes taken at intervals of 6 s.(Right) The time-lapse images of actin filaments within the red-boxed apicalregion (Left) of pollen tubes. Membrane-originated actin filaments are la-beled with different colored dots. The origination sites of actin filaments areindicated by yellow circles. Actin filament elongation, shortening, and sev-ering events are indicated by blue, purple, and green arrowheads, re-spectively. (Scale bars, 5 μm.) (C ) Dynamic parameters of actin filaments inthe apical region (as shown in the red-boxed region of B). Measurementswere taken from more than 15 pollen tubes for each genotype. Numbersof actin filaments measured are indicated in parentheses. Values arepresented as mean ± SD ***P < 0.001; ND, no significant difference (Stu-dent’s t test).

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protein in pollen (Fig. 4A), and pollen ADFs prefer ADP-G-actin, inhibit actin nucleotide exchange, and play an essentialrole in regulating actin turnover in pollen (41, 44). Dissociationof ADP-G-actin from ADF is a prerequisite for the reentryof actin into the actin polymerization cycle. CAP1 will be an ex-tremely relevant player in this process as it synergizes with ADFand profilin to promote actin turnover as well as actin nucleotideexchange activity (12). However, the role of CAP1 in regulatingactin polymerization, particularly the contribution of its actinnucleotide exchange activity, remains largely unknown.By examining the effect of loss of function of CAP1 on the

actin cytoskeleton in pollen tubes, we found that CAP1 hasdifferential effects on the actin cytoskeleton within differentregions of the pollen tube (Figs. 2A and 3A). Loss of function ofCAP1 severely reduces the F-actin level within the growthdomain but increases the F-actin level in the shank region (Figs. 2A and B and 3A). The reduced F-actin level in the apical region ofcap1 pollen tubes is due to impaired actin polymerization, as sev-ering and depolymerization of apical actin filaments are reduced incap1 pollen tubes compared with WT pollen tubes (Fig. 3C). Thissuggests that the formation of the apical actin structure is highlydependent on actin polymerization, as demonstrated recently ingrowing pollen tubes (22). The differential effect of the loss offunction of the diffusely distributed CAP1 (Fig. 2C) on the actincytoskeleton within different regions of the pollen tube is very likelydue to the fact that formins efficiently nucleate actin assembly fromthe PM within apical and subapical regions (24). In addition, thedifferential effect of CAP1 loss of function on the actin cytoskeletonmight also be due to the differential collaboration of CAP1 withADFs within different regions of the pollen tube, as ADFs densely

localize to the shank-localized actin cables but distribute compar-atively diffusely within the cytoplasm of the pollen tube tip (41, 44).Furthermore, we directly link the actin nucleotide exchange activityof CAP1 to its function in promoting actin polymerization, as wefound that the mutant protein CAP1-109, which binds G-actin andpromotes actin turnover but has reduced actin nucleotide exchangeactivity (SI Appendix, Fig. S8), cannot fully rescue actin polymeri-zation defects within the growth domain of cap1 pollen tubes (Fig.5). Interestingly, we found that CAP1-90 fails to rescue the apicalactin polymerization defect in pollen tubes (Fig. 6), which suggeststhat the conversion of ADP-G-actin bound with ADF is the rate-limiting step for the actin polymerization and depolymerizationcycle in cells. These data together suggest that CAP1 acts totransfer actin monomers from ADF to profilin. It was reportedthat overexpression of profilin can suppress the defects caused byloss of the C-terminal domain of CAP (45), which suggests thatthe actin monomer-sequestering function of CAP might be criticalin vivo. The actin monomer-sequestering function was previouslyproposed for plant CAP1 (46). It was shown that the binding af-finity of CAP1 to plant actin is higher than that of ZmPRO5 (Zeamays profilin5), although both have roughly similar binding af-finity to muscle actin (12). Similar results were obtained in ourresearch for the binding of CAP1 to muscle actin (SI Appendix,Fig. S8C). Our finding that the cellular concentration of CAP1 ismuch lower than that of actin and profilin (Fig. 4A) suggests that theactin monomer sequestering function is achieved mainly by profilinrather than CAP1 in plants. In this regard, actin mainly formscomplexes with profilin rather than CAP1 in plant cells. However,considering that plant profilins inhibit actin nucleotide exchange (30,31), the role in promoting actin nucleotide exchange will be

Fig. 4. CAP1 is an abundant protein in pollen that synergizes with pollen ADF and profilin in promoting actin turnover and stimulating actin nucleotideexchange. (A) Quantification of cytosolic concentrations of Actin, CAP1, ADF, and profilin in mature pollen of Arabidopsis thaliana. Measurements were madeas described in Materials and Methods. Protein abundance (mean ± SD) was measured from at least three experiments. Recombinant Arabidopsis ADF, PRF,CAP1, and ACT1 were used as the loading controls. (B) CAP1 efficiently promotes actin turnover in the presence of ADF and profilin. All reactions contain 4 μMe-ADP-F-actin in Buffer G supplemented with 1×KMEI and 1 μM ATP. ADF10, 4 μM; PRF5, 4 μM; CAP1, 1 μM. (C) CAP1 overcomes the inhibitory effect ofADF7 on actin nucleotide exchange. ADP-actin, 2 μM. The concentrations of actin-binding proteins are indicated. (D) CAP1 synergizes with PRF5 to enhanceactin nucleotide exchange. ADP-actin, 2 μM. The concentrations of actin-binding proteins are indicated.

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taken over by CAP1 in vivo, as we demonstrate in this study. Evenin other systems where both profilin and CAP1 harbor actin nu-cleotide exchange activity, CAP1 appears to be better suited forthis function since it binds to the substrate (ADP-G-actin) withmuch higher affinity than profilin (36, 47). In line with our find-ings, a recent report showed that deletion of four amino acids ofCAP specifically affects its nucleotide exchange activity and im-pairs its role in promoting actin cable formation (34). Therefore,our results provide unambiguous evidence that CAP1 promotesapical actin polymerization in pollen tubes, and this function re-quires the ability of CAP1 to stimulate actin nucleotide exchange.

Previous characterizations support the function of ArabidopsisCAP1 in promoting actin turnover (8, 46). In particular, loss offunction of CAP1 increases the amount and bundling of actinfilaments in Arabidopsis root hairs (8). A similar phenomenonwas noted in the shank region of the pollen tube and pollengrains (Figs. 2A and 3A and SI Appendix, Fig. S5). In support ofthe role of CAP1 in promoting actin turnover in vivo, it waspreviously shown that Arabidopsis CAP1 inhibits spontaneousactin polymerization and is capable of synergizing with ADF topromote actin depolymerization (12). However, the precisemechanism underlying the action of CAP1 is not well understood.

Fig. 5. CAP1-109 cannot fully rescue the defective actin polymerization within the apical region of cap1 pollen tubes. (A) Micrographs of actin filamentsstained with Alexa-Fluor-488–phalloidin in pollen tubes of WT and CAP1-109 transgenic lines. CAP1-109 is CAP1pro:CAP1-109;cap1-1. The Z-projection andselected optical sections are shown. (Scale bar, 5 μm.) The red boxes indicate the apical region (0–10 μm from the tip). (Lower) The distribution of fluorescenceintensity of actin filaments in transverse sections at the indicated distances from the tip of pollen tubes. Pseudocolored transverse images are also shown.Warm and cold colors indicate higher and lower fluorescence, respectively. (Scale bar, 5 μm.) (B) Quantification of the fluorescence intensity of actin filamentsfrom the extreme tip to the base along the growth axis of pollen tubes in WT, cap1-1, COM-1, and CAP1-109 lines. The dashed red line indicates the base ofthe red-boxed regions shown in the Upper of A. More than 20 pollen tubes were measured for each line. Data are presented as mean ± SEM. (C) Quanti-fication of the mean fluorescence intensity of actin filaments within the red-boxed region of pollen tubes. More than 20 pollen tubes were measured for eachgenotype. Data are presented as mean ± SEM. Statistical comparisons were performed using ANOVA with a post hoc Tukey test; different letters indicatesignificant differences, P < 0.01. WT vs. COM-1, P = 0.99; WT or COM-1 vs. cap1-1, P < 0.0001; 1# vs. 2#, P = 0.98; 1# vs. cap1-1 P = 0.021; 2# vs. cap1-1, P = 0.03;WT vs. 1#, P = 0.032; WT vs. 2#, P = 0.02; COM-1 vs. 1#, P = 0.014; COM-1 vs. 2#, P = 0.009. (D) Time-lapse images of actin filaments in growing pollen tubes ofWT and CAP1-109 plants. Actin filaments were revealed by decoration with Lifeact-EGFP. Red brackets indicate the apical actin filaments in the pollen tube.See the entire series in Movies S5 and S6. (Right) Pseudocolored kymograph analysis of apical actin filaments during pollen tube growth. Warm and cold colorsindicate high and low fluorescence intensity, respectively. The fluorescence intensity of Lifeact-EGFP indicates the amount of actin filaments in the pollentube (from tip to shank region along the longitudinal axis) during the growth process. The enlarged kymograph images show the pollen tube tip region(0–10 μm) in the Upper. (Scale bars, 10 μm.) (E) Dynamic parameters of actin filaments in the apical region of pollen tubes. Measurements were taken frommore than 15 pollen tubes for each genotype. The numbers of actin filaments analyzed are shown in parentheses. Values are presented as mean ± SD ***P <0.001; *P < 0.05; ND, no significant difference (Student’s t test).

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We here demonstrate that Arabidopsis CAP1 enhances ADF-mediated actin severing and depolymerization via its N terminus(Fig. 1 C–G), which suggests that Arabidopsis CAP1 promotesactin turnover using the same mechanism as reported for CAPprotein from other organisms (14). We previously demonstratedthat ADF is an essential regulator of actin turnover in pollen (41,44). Accordingly, we found here that ADF is very abundant, as itscytosolic concentration reaches about 5 μM in pollen (Fig. 4A).Although the molar ratio of ADF to actin is roughly 1:6 in Ara-bidopsis pollen (Fig. 4A), the percentage of F-actin is less than10% of total actin in pollen (20, 39). Therefore, we speculate thatADF is sufficient to saturate F-actin in pollen cells. In partialsupport of this speculation, we found that both ADF7 andADF10 form prominent filamentous actin structures in pollencells (41, 44). Strikingly, we found that disruption of the in-teraction between the N terminus of CAP1 and ADF impairs itsfunction in promoting actin turnover in pollen cells (Fig. 6D andSI Appendix, Fig. S3 H–J). This suggests that the role of CAP1 indriving actin turnover in vivo is exclusively through its interactionwith ADF.Based on the in vitro and in vivo data presented here, we

propose that CAP1 synergizes with pollen ADF to enhance actinmonomer dissociation and filament severing, which subsequentlyreleases ADP-G-actin from ADF. After stimulating the nucleo-tide exchange of ADP-G-actin to convert it into ATP-G-actin,CAP1 subsequently transfers ATP-G-actin to profilin toform ATP-G-actin-profilin complexes to maintain a pool ofpolymerization-competent actin monomers. This will be utilized

by membrane-localized formins to support actin polymerizationfrom the PM within the growth domain of pollen tubes (Fig. 7).We thus identify CAP1 as a major player in driving actin poly-merization via recharging ADP-G-actin and facilitating the for-mation and maintenance of a pool of polymerization-competentactin monomers. Our study significantly enhances our un-derstanding of the mechanism of actin polymerization in pollentubes as well as the mechanism of action of Srv2p/CAP1 inregulating actin assembly and disassembly in general.

Materials and MethodsPlant Materials and Growth Conditions. The information about two CAP1T-DNA insertion lines, cap1-1 (Salk_112802) and cap1-2 (GK_453G08), hadbeen described previously (8). Arabidopsis CAP1 overexpression and com-plementation plants were generated as described below. ArabidopsisColumbia-0 ecotype was used as WT in this study. Arabidopsis plants weregrown in a culture room at 22 °C under a 16-h-light/8-h-dark photoperiod.

Complementation of cap1 Mutants and Visualization of the IntracellularLocalization of CAP1. To generate the CAP1 complementation construct, inwhich expression of CAP1 was driven by the CAP1 promoter, the genomicsequence of CAP1 was amplified with primers pgCAP1For and pgCAP1Rev+TAA(SI Appendix, Table S1) using Arabidopsis genomic DNA as the template andsubsequently moved into pCAMBIA1301-NOS to generate pCAMBIA1301-CAP1pro:CAP1g-NOS plasmid. To determine the function of the interactionbetween CAP1 and ADF and the nucleotide exchange activity of CAP1 in vivo,we generated pCAMBIA1301-CAP1pro:CAP1-90g-NOS and pCAMBIA1301-CAP1pro:CAP1-109g-NOS plasmids using pCAMBIA1301-CAP1pro:CAP1g-NOSplasmid as the template with primer pairs pgCAP1-90For/pgCAP1-90Revand CAP1-109For/CAP1-109Rev (SI Appendix, Table S1), respectively. They were

Fig. 6. CAP1-90 cannot fully rescue the defective membrane-originated actin polymerization within the apical region of cap1 pollen tubes. (A) Time-lapseimages of actin filaments in growing pollen tubes of WT and CAP1-90 plants. Actin filaments were revealed by decoration with Lifeact-EGFP. Red bracketsindicate the apical actin filaments in the pollen tube. See the entire series in Movies S7 and S8. (Scale bar, 10 μm.) (B) Pseudocolored kymograph analysis ofapical actin filaments during pollen tube growth. The fluorescence intensity of Lifeact-EGFP indicates the amount of actin filaments in the pollen tube (fromthe tip to shank region along the longitudinal axis) during the growth process. The green box indicates the red-bracketed region shown in A. Warm and coldcolors indicate high and low fluorescence intensity, respectively. (Scale bar, 10 μm.) (C) Projection images showing the organization of actin filaments in WTand CAP1-90 pollen tubes. Pseudocolored images are shown at the Right. The (Lower) The distribution of fluorescence intensity of actin filaments inpseudocolored transverse sections at the indicated distances from the tip of pollen tubes. Warm and cold colors indicate higher and lower fluorescence,respectively. White arrowheads indicate heavy actin cables. (Scale bar, 10 μm.) (D) Dynamic parameters of actin filaments in the apical region of pollen tubes.Measurements were taken from more than 15 pollen tubes for each genotype. The numbers of actin filaments analyzed are shown in parentheses. Values arepresented as mean ± SD ***P < 0.001; *P < 0.05; ND, no significant difference (Student’s t test).

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transformed into cap1-1 to generate transgenic plants pCAMBIA1301-CAP1pro:CAP1g-NOS;cap1-1, pCAMBIA1301-CAP1pro:CAP1-90g-NOS; cap1-1, andpCAMBIA1301-CAP1pro:CAP1-109g-NOS; cap1-1, respectively. To generate theCAP1-EGFP fusion construct driven by the CAP1 promoter, the genomic sequenceof CAP1 was initially amplified with primers pgCAP1For and pgCAP1Rev-TAA (SIAppendix, Table S1). EGFP was amplified with primers EGFPFor and EGFPRev (SIAppendix, Table S1). Both fragments were digested and subsequently movedinto pCAMBIA1301-NOS to generate the final pCAMBIA1301-CAP1pro:CAP1g-EGFP-NOS plasmid. pCAMBIA1301-CAP1pro:CAP1g-EGFP-NOS construct wastransformed into cap1-1 to generate the transgenic plant pCAMBIA1301-CAP1pro:CAP1g-EGFP-NOS;cap1. To visualize the intracellular localization of CAP1, pollenderived from pCAMBIA1301-CAP1pro:CAP1g-EGFP-NOS;cap1 plants was visualizedunder a fluorescence light microscope equipped with a 100× oil objective (1.46numerical aperture HC PLAN), and images were captured by confocal laser scan-ning microscopy excited with the 488-nm line of an argon laser. Optical sectionswere collected with a step size of 0.5 μm for both pollen grains and pollen tubes.

F-Actin Staining and Quantification in Fixed Pollen Grains and Pollen Tubes.Actin filaments in pollen grains and pollen tubes were stained with Alexa-488 phalloidin as previously described (48, 49). To reveal actin filaments inpollen grains and pollen tubes, pollen grains were subjected to staining with

Alexa488 phalloidin after culturing for 10 min and 2 h at 28 °C. Briefly,300 μM 3-maleimidobenzoic acid N-hydroxysuccinimide ester in liquid ger-mination medium (GM) was added onto the surface of solid GM. After in-cubation for 1 h, the GM plate was washed three times with TBSS buffer[50 mM Tris·HCl, 200 mM NaCl, 400 mM sucrose and 0.05% (vol/vol) NonidetP-40, pH 7.5] in GM. Finally, pollen grains or pollen tubes were incubatedwith 150 nM Alexa-488 phalloidin in TBSS buffer overnight at 4 °C. The sampleswere observed under a laser scanning confocal microscope (OlympusFV1000MPE) equipped with a 100× oil objective (numerical aperture of 1.4). Thesample was excited with the 488-nm line of an argon laser, the emission was setin a range of 505–525 nm, and the Z-series images were collected with the Z-step set at 0.5 μm. Maximum intensity projection of Z-series and the opticalsection of both pollen grains and pollen tubes are displayed in the figures. Theamount of actin filaments was quantified by determining the fluorescence in-tensity of Alexa-Fluor-488–phalloidin staining with ImageJ software.

Direct Visualization and Quantification of Actin Filament Dynamics in LivingPollen Tubes. Actin filaments in pollen tubes were revealed by decorationwith Lifeact-EGFP as described previously (37, 38). To reveal actin filaments inpollen tubes of cap1, CAP1 overexpression, and CAP1-90 and CAP1-109plants, the marker Lifeact-EGFP was introduced by crossing them withWT Arabidopsis plants harboring Lat52:Lifeact-EGFP. After segregation,T3 homozygous plants harboring Lat52:Lifeact-EGFP were used for theanalysis of actin filament dynamics in pollen tubes, and pollen tubes derivedfrom sibling Arabidopsis plants harboring Lat52:Lifeact-EGFP were used asthe control for comparison. Actin filaments were observed under a spinningdisk confocal microscope equipped with a Yokogawa CSUX1 scanning head.The time-lapse Z-series images were acquired with an Andor iXon3 DU888EMCCD camera at 2-s intervals, and the Z-step size was set at 0.7 μm. Toreveal the overall dynamics of apical actin filaments, three consecutive im-ages, pseudocolored differently, were merged as described previously (50). Akymograph along the growing direction at the center of the growing pollentube was generated to analyze the intensity of F-actin. First, a 5-μm wideband was drawn along the growing axis of a pollen tube in the time-lapsestacks with the built-in tool “segmented line” in ImageJ. Then the plotprofiles were analyzed with the macro “Stack profile data,” and the resultswere further imported into ImageJ to produce the kymograph. Each line alongthe growth axis represents the average plot profile along the 5-μm band in thepollen tube at one time point. To clearly observe the apical actin filaments, agrayscale kymograph image was applied to the color lookup tables in thesubmenu of ImageJ. The kymograph images of the pollen tube tip were dis-played after pseudocolor processing. Warm and cold colors indicate high andlow fluorescence intensity, respectively. The dynamic parameters of individualactin filaments were analyzed as described previously (38). Two additional pa-rameters, bundling and debundling frequencies, were measured to reveal actindynamics within the shank region of pollen tubes as described previously (44).

Generation of CAP1 overexpression transgenic plants, qRT-PCR analysis,observation and quantification of pollen germination and pollen tubegrowth, sequence alignment, protein production, production of polyclonalantisera, determination of cytosolic protein concentration by quantitativeWestern blotting assay, in vitro actin assembly and disassembly assays, actinmonomer-binding assay, and nucleotide exchange assay are described in SIAppendix, Materials and Methods.

ACKNOWLEDGMENTS. We thank the Nottingham Arabidopsis Stock Centrefor the sequence-indexed T-DNA insertion lines and Dr. Xin Liang (School ofLife Sciences, Tsinghua University) for help with quantifying the proteinconcentration in pollen. This work was supported by a grant from the Na-tional Natural Science Foundation of China (31671390). The research in theS.H. laboratory is also supported by funding from Tsinghua-Peking JointCenter for Life Sciences. Y.J. is supported by postdoctoral fellowships fromTsinghua-Peking Center for Life Sciences and the China Postdoctoral ScienceFoundation (Grant 2017M620755).

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Fig. 7. Schematic model illustrating the action of CAP1 in promoting apicalactin polymerization in the pollen tube. This simple schematic model showsCAP1 as an important player in coordinating with ADF and profilin to pro-mote the turnover of membrane-originated actin filaments within thegrowth domain of the pollen tube. The diagram on the Left reflects themolar ratio of CAP1 with actin and several other actin associated proteinsand their intracellular localization patterns, as shown by refs. 25, 41, 44, 51.(Right) An enlarged picture of the boxed apical region. The relative amountof CAP1 is increased to highlight the function of CAP1. Briefly, within thecytoplasm of the pollen tube, CAP1 coordinates via its N terminus with ADFto promote actin turnover. After releasing ADP-G-actin from ADF,CAP1 enhances actin nucleotide exchange via its C terminus to generateATP-G-actin and subsequently transfers ATP-G-actin to profilin to form ATP-G-actin–profilin complexes. In this regard, CAP1 plays an important role inmaintaining the pool of polymerization-competent actin monomers, whichcan be utilized by the membrane-anchored formins within the apical andsubapical regions of pollen tubes as we demonstrated recently (24).

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