LARGE-SCALE BIOLOGY ARTICLE

Cell Cycle–Dependent Regulation and Function ofARGONAUTE1 in Plants[OPEN]

Adrien Trolet,a,1 Patricia Baldrich,b,1 Marie-Claire Criqui,a Marieke Dubois,a Marion Clavel,a Blake C.Meyers,b,c andPascal Genschika,2

a Institut de Biologie Moléculaire des Plantes (IBMP-CNRS), 12, Rue du Général Zimmer, 67084 Strasbourg Cedex, FrancebDonald Danforth Plant Science Center, 975 N Warson Road, St. Louis, Missouri 63132cDivision of Plant Sciences, 52 Agriculture Lab, University of Missouri, Columbia, Missouri 65211

ORCID IDs: 0000-0003-0710-1034 (A.T.); 0000-0003-4669-6632 (P.B.); 0000-0002-6419-8306 (M.-C.C.); 0000-0002-5190-2130(M.D.); 0000-0003-1322-7362 (M.C.); 0000-0003-3436-6097 (B.C.M.); 0000-0002-4107-5071 (P.G.)

Regulated gene expression is key to the orchestrated progression of the cell cycle. Many genes are expressed at specificpoints in the cell cycle, including important cell cycle regulators, plus factors involved in signal transduction, hormonalregulation, and metabolic control. We demonstrate that post-embryonic depletion of Arabidopsis (Arabidopsis thaliana)ARGONAUTE1 (AGO1), the main effector of plant microRNAs (miRNAs), impairs cell division in the root meristem. We utilizedthe highly synchronizable tobacco (Nicotiana tabacum) Bright yellow 2 (BY2) cell suspension to analyze mRNA, small RNAs,and mRNA cleavage products of synchronized BY2 cells at S, G2, M, and G1 phases of the cell cycle. This revealed that inplants, only a few miRNAs show differential accumulation during the cell cycle, and miRNA-target pairs were only identifiedfor a small proportion of the more than 13,000 differentially expressed genes during the cell cycle. However, this unique set ofmiRNA-target pairs could be key to attenuate the expression of several transcription factors and disease resistance genes.We also demonstrate that AGO1 binds to a set of 19-nucleotide, tRNA-derived fragments during the cell cycle progression.

INTRODUCTION

In all eukaryotes, the basic principles controlling cell divisionappear to be conserved (Nurse, 2000). Thus, the cell cycle iscomposed of four phases: in gap phase 1 (G1), cells increase theirnumber of organelles; during S phase DNA replication occurs; ingap phase 2 (G2), cells still increase their size by extensive proteinsynthesis; and inmitosis (M) phase, chromosomes segregate intotwo nuclei, followed by cytokinesis, duringwhich cells are dividedinto two daughter cells. The orchestration of the cell cycle, andespecially the transition from G1 to S phase as well as the pro-gression andexit fromMphase, requiresmultiple levels of control.Inparticular, cyclin-dependentkinases (CDKs) that arespecificallyactivated by cyclins are key players in this process (MalumbresandBarbacid, 2005;DeVeylder et al., 2007). Several other kinasesand phosphatases, as well as additional regulatory proteins, suchasCDK inhibitors, also regulate progression through the cell cycle(Boutros et al., 2006; Fisher, 2012; Starostina and Kipreos, 2012).In Arabidopsis (Arabidopsis thaliana), and plants in general, mostcore cell cycle genes are conserved (Vandepoele et al., 2002),although several are present in multiple copies. Thus, while in

mammals three CDK inhibitory protein/Kinase inhibitory protein(Cip/Kip) members inhibit a broad spectrum of CDK-cyclincomplexes (Denicourt and Dowdy, 2004), the Arabidopsis ge-nome encodes at least 21 related Cip/Kip proteins (Kumar et al.,2015). Likewise, multigenic gene families also encode severalplant cyclin families, including a unique class of CDKs (B-type) forwhich no counterpart is found in other organisms (Vandepoeleet al., 2002; Boudolf et al., 2004).The control of each step of the cell cycle therefore requires the

dynamic accumulation of specificmRNAs and proteins, achievedthrough transcriptional, posttranscriptional, and posttranslationalcontrols. The overaccumulation of cyclins, the absence of CDKinhibitors, or the misexpression of retinoblastoma proteins iscommonly found in human cancers, indicating that the properprotein level of cell cycle regulators is key for cell division. Theubiquitin proteasome system actively participates in this processby promoting irreversible and timely proteolysis of numerousregulatory proteins absolutely required for transitions in the cellcycle phase (Mocciaro and Rape, 2012; Genschik et al., 2014). Afailure to degrade some cell cycle regulatory proteins leads toabnormal cell division and chromosomal instability.Beside selective proteolysis, microRNAs (miRNAs) have a

demonstrated role in controlling the levels of multiple cell cycleregulators in mammals (Bueno and Malumbres, 2011). miRNAsarederived fromprocessed, single-strandedRNAs, functioning tocontrol the stability and translation of protein-codingmRNAs (Krolet al., 2010; Axtell, 2013). This is achieved through their in-corporation into a protein complex called the RNA-inducedsilencing complex (RISC; Czech and Hannon, 2011). Central

1 These authors contributed equally to this work.2 Address correspondence to pascal.genschik@ibmp-cnrs.unistra.fr.The author responsible for distribution of materials integral to the findingspresented in this article in accordance with the policy described in theInstructions for Authors (www.plantcell.org) is: Pascal Genschik (pascal.genschik@ibmp-cnrs.unistra.fr).[OPEN]Articles can be viewed without a subscription.www.plantcell.org/cgi/doi/10.1105/tpc.19.00069

The Plant Cell, Vol. 31: 1734–1750, August 2019, www.plantcell.org ã 2019 ASPB.

components of RISC are ARGONAUTE (AGO) proteins that bindthe mature miRNA and orient it for interaction with target mRNAs(Meister, 2013; Poulsen et al., 2013). In animal cells, a singlemiRNA can have a large number of different mRNA targets; inhumans, the >1000 annotated miRNAs may control up to 60% ofthegenes (Friedmanet al., 2009; LeungandSharp, 2010).miRNAshave been implicated in numerous developmental and physio-logical processes and are involved in a number of diseases, in-cluding cell proliferative diseases such as cancer (Williams, 2008).

miRNAs target dozens of cell cycle genes inmammals and thusplay functions in the regulation of theG1/S transition and the entryand progression throughM phase (Bueno andMalumbres, 2011).For example, the miRNA cluster miR-15a-16-1 could induce cellcycle arrest at G1 by targeting CDKs and cyclins (Linsley et al.,2007; Liu et al., 2008). miRNAs also target negative regulators ofthe cell cycle such as retinoblastoma proteins (Volinia et al., 2006)and cell cycle inhibitors of the INK4 or Cip/Kip families (Visoneet al., 2007;Kimetal., 2009;Wanget al., 2009), thuspromotingcellcycle entry. AGO proteins also bind other classes of small RNAs(sRNAs), such as small interfering RNAs (siRNAs). In plants,siRNAs thatderive fromadouble-strandedRNAprecursor includeheterochromatic siRNAs (hc-siRNAs) andphasedsiRNAs (BorgesandMartienssen,2015). Inaddition, recentworkshowed thatAGOproteinsbindsRNAs that are tRNA fragments (tRFs),whichact likemiRNAs to regulate cellular functions, including cell proliferationand in genome protection against retrotransposons (Lee et al.,2009; Haussecker et al., 2010; Shigematsu and Kirino, 2015;Martinez et al., 2017).

Unlike the situation in animal cells, little is known in plants aboutposttranscriptional regulation of gene expression during the cellcycle and the role and regulation of AGO proteins in this process.Mutations in genes encodingmiRNApathway components causepattern formation defects leading, in the worse cases, to em-bryonic lethality (Nodine and Bartel, 2010; Vashisht and Nodine,2014). Similarly, aberrant patterns of cell division and cell ex-pansion during embryogenesis have been identified inmutants ofArabidopsis SERRATE (SE) and HYPONASTIC LEAVES1 (HYL1);both genes encode proteins that cooperate with DICER-LIKE1(DCL1) formiRNA biogenesis (Grigg et al., 2009; Armenta-Medinaet al., 2017), pointing to an important role of plant miRNAs in theregulation and/or maintenance of cell division. Here, we in-vestigated the regulation of AGO1 during the cell cycle and ex-amined the repertoire of total sRNA and AGO1-bound sRNAs atdifferent phases of the cell cycle.

RESULTS

AGO1 Depletion Affects Arabidopsis Cell Division and RootMeristem Activity

Arabidopsis ago1 null mutants exhibit a severe morphologicalphenotype affecting leaf shape and polarity, along with defects inmeristem identity and function (Bohmert et al., 1998; Morel et al.,2002; Kidner and Martienssen, 2005). Analysis of primary rootgrowth of ago1 mutants revealed a clear reduction in the rootlength of the hypomorphic ago1-27 allele, while this phenotypewas severely compromised in the strong ago1-36 mutant

(Supplemental Figure 1A). Moreover, the highly organizedstructure of root apical meristem was altered in a significantproportionof ago1-27 roots andeven lost in ago1-36mutant roots(Supplemental Figure 1B), precluding the quantification of theirmeristematic cells.Because these defects in root meristem activity might originate

during embryogenesis, we aimed to deplete AGO1 post-embryonically. For this, we used the b-estradiol–inducible(XVE:P0) line to express the F-box protein P0 from Turnip yel-lows virus, which induces the degradation of plant AGO proteins(Derrien et al., 2018). Upon P0 induction, both the root length andthe root apical meristem size were significantly reduced (Figures1A and 1B) and thus recapitulated the phenotype observed instrong ago1mutant alleles. Because P0 triggers the degradationof several, if not all, plant AGOs (Baumberger et al., 2007; Derrienet al., 2018),wealso inducedP0expression in theago1-57geneticbackground, expressing amutated form of AGO1 resistant to P0-mediateddegradation (Derrien et al., 2018). Thestrongeffect ofP0on meristem size and cell division activity was significantly sup-pressed in ago1-57 (Figures 1A and 1B), indicating that thisphenotype is mainly dependent on AGO1. The fact that thephenotype was not fully suppressed might be attributed to theago1-57mutation affecting some siRNA pathways (Derrien et al.,2018) or the involvement, although minor, of some other AGOproteins.Wenextwonderedwhether AGO1degradationwould block cell

division at a specific phase of the cell cycle. To this end, wetransformed the XVE:P0 and XVE:P0 (ago1-57) lines with thepHTR2:CDT1a (C3)-green fluorescent protein (GFP; Yin et al.,2014) and pCYCB1;2:CYCB1;2(Dbox)-GFP constructs as mark-ers of cells in S/G2 and G2/M phases, respectively. Roots ofdouble homozygous plants were visualized by confocal micros-copy before and upon P0 induction. In the wild-type background,the number of cells expressing either the S/G2 or G2/M markersdramatically decreased upon P0-mediated AGO1 degradation,while this was not the case in ago1-57 expressing the stableversion of the AGO1 protein (Figure 2). From these experiments,we conclude that AGO1 activity is required tomaintain normal cellproliferation in theArabidopsis rootmeristembut that its depletiondoes not lead to an arrest specific to the S, G2, or M phases of thecell cycle. Note that we cannot exclude the possibility that AGO1depletionmight alsoaffect the timingof aspecificcell cyclephase.

AGO1 Regulation in Synchronized BY2 Cells

To address more specifically the question of the regulation andfunction of AGO1 during the cell cycle, we moved to the tobacco(Nicotiana tabacum) BY2 cell suspension (Nagata et al., 1992),which offers optimal synchronization efficiencies and also thepossibility to biochemically monitor AGO1 protein steady-statelevels at different cell cycle stages. We established clonal cellsuspension cultures with the pAGO1:GFP-AGO1 construct ex-pressing a functional GFP-AGO1 protein under the control of itsown promoter (Derrien et al., 2012). We observed that at a highexpression level of the GFP-AGO1 transgene, progression of thecell cycle was significantly delayed (Supplemental Figure 2). Wetherefore selected clonal BY2 cell suspensions with moderateGFP-AGO1 protein expression for synchronization experiments.

Cell Cycle–Dependent Accumulation of Small RNAs 1735

Figure 1. AGO1 Is Required for Arabidopsis Cell Division and Root Meristem Activity.

(A)Root lengthmeasurements at 6 and 12 days after stratification (DAS) of the indicated genotypes undermock (2) orb-estradiol (10mM) to induce P0 (1).At least 30 seedlings per line per treatment were measured, and ANOVA was performed to assess significant differences. *** highlights comparisons forwhich P < 0.001.(B) Root-meristem size of wild-type seedlings compared with the indicated genotypes. Cortex meristematic cells showing no sign of differentiation werecounted. Values aremeristem length of 6DASseedlings germinatedwith (1) orwithout (2)b-estradiol (10mM).ANOVAwasperformed toassess significantdifferences. * highlightscomparison forwhich0.01<P<0.05. (Right)Primary root tipsofXVE:P0andXVE:P0 (ago1-57) after 12dofP0 induction.Rootswerecounterstained with propidium iodide. The arrow indicates the length of the meristematic zone (from the quiescent center to the first elongating cell of thecortex). Bar 5 50 mm.

Figure 2. S/G2 and G2/M Markers Dramatically Decreased upon P0-Mediated AGO1 Degradation.

(A)Confocal laser scanning images of primary root tips of XVE:P0 andXVE:P0 (ago1-57) lines expressing the indicated cell cyclemarkers after 6 and 10 d ofP0 induction with b-estradiol (10 mM) or mock treatment. The timing of expression of both cell cycle markers is illustrated on the top graphic. Bar5 50 mm.(B) Immunoblot analysis of AGO1 content in mock (2) or P0 induced (1) 6-d-old seedlings of the same genotypes as in (A), with or without b-estradiol.Probing with the ACTIN antibody and Coomassie blue (CB) staining were used as loading controls. Quantification of AGO1 and AGO2 signal normalized toACTIN is indicated on top of their respective panels.(C) RT-qPCR analysis of P0 (top) and AGO1 and AGO2 (bottom) mRNA levels in the same samples as in (B).

BY2 cells were synchronized by a 24-h treatment with aphidicolin(an inhibitor of DNA polymerase), as reported previously by Criquiet al. (2000), and total RNAsand proteinswere extracted at differenttime points after removal of the drug (Figure 3). Cell cycle pro-gression was monitored by flow cytometry measurements of DNAcontents,byRT-qPCRmonitoringofcellcyclegeneexpression,andby determination of the mitotic index at the different time points(Figures3Aand3B).FromSphase (1 to2hafteraphidicolin removal)toG2 (5 to6hafter aphidicolin removal), theGFP-AGO1proteinwasdetected at all time points of the cell cycle without substantialfluctuations (Figure 3C). By contrast and as expected, cyclin B1protein levels only accumulated during G2/M, rapidly decreasedafter the time of the peak in the mitotic index (6 h), and reachedbackground levels at the exit of mitosis (9 h). Similar to the AGO1protein level, theaccumulationof theAGO1 transcript (Figure3B)didnot dramatically change during the cell cycle. A second synchro-nization experiment is presented in Supplemental Figure 3 showingsimilar results.Thus,weconclude that theAGO1 expression levelremains overall constant throughout the cell cycle.

AGO1 Localizes to Different Cellular Bodies in SynchronizedBY2 Cells

Wenext imaged the localization of GFP-AGO1 in our transgenic celllines. In asynchronizedBY2cells, theGFP-AGO1 fusionproteinwasfound predominantly in the cytosol, as reported previously in Ara-bidopsis root cells (Derrien et al., 2012), but it was also present in thenuclear compartment (Figure 4A). Note that the nuclear localizationof AGO1 has been documented in Arabidopsis previously (Dolataet al., 2016; Bologna et al., 2018; Liu et al., 2018). We next syn-chronizedBY2cellsandimagedGFP-AGO1atthedifferentcellcyclephases (Figures 4B to 4H). During S phase, the GFP-AGO1 signalwasmainlyenriched in thecytosol (Figures4Cand4D). Interestingly,duringG2, we observed theGFP-AGO1 signal also in larger nuclearbodies (of;2 to6mm)distinct fromnucleoli (Figure4E).Quantitativeanalysis showed that the number of cells that contain these nuclearbodies increased from S to G2 and dropped significantly when themitotic index started to increase, suggesting a cell cycle–dependentperiodic accumulation of GFP-AGO1 nuclear foci (Figure 4B). Inmitotic cells, the fluorescence signal was excluded fromcondensedchromosomes (Figures 4F and 4G).

To confirm this observation, we transformed the GFP-AGO1clonal cell suspensionswith aconstruct expressing thehistone2B(H2B) fused to tandem dimer tomato (tdTomato), which onceincorporated intochromatin allowsvisualizationof nuclear events.Hence, the dual localization of both fluorescent proteins inmitoticcells indicated no substantial overlap in condensed chromatin(Figure 4I). Finally, after mitotic exit, cells in G1 showed againa predominant cytosolic distribution of the GFP-AGO1 protein(Figure 4H). From these observations, we conclude that AGO1exhibits a complex subcellular localizationpatternwith foci in boththe nucleus and cytosol, supporting the existence of pools ofRISCs that may play distinct functions along the cell cycle.

Differential Gene Expression in Synchronized BY2 Cells

To achieve a global view of gene expression during the cell cycle,we performed five independent synchronization experiments

(called replicates 1 to 5) using a clonal BY2 cell suspension (131)expressingGFP-AGO1; the aimwas to analyze total RNA, sRNAs,and AGO1-associated sRNAs at key points of the cell cycle.According to flow cytometry measurements of DNA contents andthe monitoring of mitotic indexes, all five synchronization ex-periments were highly reproducible in the timing of the differentcell cycle phases and their level of synchrony (SupplementalFigure 4). For each experiment, we collected RNA and proteinsamples for S, G2, M, and G1 phases (at 1, 3, 7, and 10 h afteraphidicolin removal, respectively).To focusongenes regulatedduring thecell cycle,weperformed

RNA sequencing (RNA-seq) with three biological replicates toidentify differentially expressed (DE) genes, comparing S, G2, M,and G1 phases (Supplemental Figure 5A; Supplemental Table 1).Of the 69,500 genes identified in the tobacco genome (Edwardset al., 2017), we were able to detect 66,752 (96%). Among thosegenes, considering all time points together, we identified a total of23,060 DE transcripts corresponding to 13,623 genes witha functional annotation (Supplemental Data Sets 1A and 1B).However, consideringeachcomparisonseparately, thenumberofDE genes was variable, ranging from 6737 to 14,771 DE genes inM/G1 and S/M comparisons, respectively (Figure 5A). However,the ratio of down- and upregulated genes wasmaintained in eachphase of the cell cycle and was always ;50%. Considering onlythe DE genes with a fold change of 5 or higher, we observeda change in the expression trend, that is, a high percentage of theDE genes were upregulated in M and G1 phases and down-regulated in S and G2 phases (Figure 5B and SupplementalFigure 5B). The genes highly expressed (minimal counts permillion > 10) and showing high fold change (>5) in expressionmainly encode histone-related proteins and proteins involved incytoskeleton and cytokinesis (Supplemental Figure 5C, greendots; Supplemental Data Sets 2A and 2B). As expected, the ex-pression of the histone-related genes peaked in Sphase, and theywere weakly expressed in M phase, while cytokinesis-relatedgenes peaked in M phase. In the group of genes showing highfold changes, we identified multiple cell cycle regulators such ascyclins. These genes are not highly abundant in all phases, butinstead have expression levels dropping to values close to 0 in atleast one cell cycle phase (Supplemental Figure 5C, blue dots;Supplemental Data Sets 2C and 2D).For a better understanding of the RNA-seq data structure, we

performed a soft clustering analysis usingMfuzz package (Kumarand Futschik, 2007). Soft clustering groups genes that havea similar expression pattern, providing insight in gene functionsand networks. We divided our DE data set, containing more than23,000genes (q-value<0.05), into 16different clusters (Figure5C;Supplemental Figure 6). We observed distinct expression patters,mostly due to the highly dynamic transcription. The two mostabundant clusters were clusters 11 and 6, representing a singlepeak of expression during phases M and S, respectively(Figure 5C). Genes included in cluster 6 are involved in chromatinassembly and organization, receptor-mediated endocytosis, andphosphorylation,while genes included in cluster 11were enrichedin cellular component movement, cytoskeleton organization, andintercellular protein transport (Supplemental Data Set 1).Next, we conducted a Gene Ontology (GO) analysis on the DE

genes using PANTHER software (see “Methods”). As expected,

1738 The Plant Cell

Figure 3. Accumulation Pattern of AGO1 mRNA and Protein in Synchronized BY2 Cells.

(A) DNA content measurements of synchronized cell nuclei by flow cytometry at different time points. Arrows represent 2C and 4C DNA content.(B) Transcript analysis of gene expression by RT-qPCR in aphidicolin-synchronized cells. (Left) Transcript accumulation of the cell cycle–regulated geneshistoneH4andCyclinB1used tomonitorSandG2/Mphase, respectively.Thedashed line represents themitotic indexdeterminedbycounting thepercentageofdividing cells. (Right) Transcript level of endogenousAGO1 (Nta AGO1) andGFP-AGO1. RT-qPCRvalues represent themean6 SE of three technical replicates.(C)EndogenousAGO1andGFP-AGO1protein levelsatdifferent timepointsof thesynchronizationexperiment. Total proteins (30mg)were separatedby4 to16% gradient SDS-PAGE for GFP-AGO1 (Coomassie blue stain CB1) and for endogenous AGO1 (Coomassie blue stain CB2). The respective antibodiesused are indicated (@). Antibodies against Nta CYCLIN B1were used as G2/M phasemarker. ACTIN immunodetection and Coomassie blue staining wereused as loading controls. The graph (bottom) shows GFP-AGO1 and endogenous Nta AGO1 quantifications normalized to CB. The orange dashed linerepresents endogenous AGO1 signal quantification normalized to ACTIN.

Figure 4. Subcellular Localization of GFP-AGO1 during the Cell Cycle in Transgenic BY2 Cells.

(A) Confocal laser imaging of GFP-AGO1–expressing BY2 cells (left) and wild-type (WT) BY2 cells (middle) with its corresponding differential interferencecontrast (DIC) reference image to highlight the background signal. The fluorescence signal was acquired in the 500 to 550 nm band (excitation, 480 nm) for GFPdetection.(B) to (H)Subcellular localization of GFP-AGO1 in synchronized cell culture. (B)Mitotic index (MI) of the synchronization experiment used for imaging. Thegreen line shows thepercentageof cells exhibitingGFP-AGO1nuclear foci. (C) to (H)Confocal acquisition imagesofGFP-AGO1–expressing synchronizedBY2 cells at the indicated time points (after aphidicolin removal). Images correspond to cells in S phase (see [C] and [D]), G2 phase (E), metaphase andanaphase (see [F] and [G], respectively), and G1 (H). Arrows highlight the presence of nuclear foci in G2 cells (E), quantified in (B).(I)Confocal laser imaging of asynchronous cells expressingGFP-AGO1 and the chromatinmarker pRPS5:H2B-tdTomato. The two panels on the left showthe GFP-AGO1 signal only or both the GFP-AGO1 and H2B-tdTomato signals. The two panels on the right are close-up views of the area delimited by thedashed squares on the left panels. Arrows highlight the presence of nuclear foci.

gene categories related to cell cycle functions such as DNAreplication, chromosome segregation, and mitotic processesincluding cytokinesis were well represented (Figure 5D). In ad-dition, genes involved in DNA repair and epigenetic regulation ofgene expression were also greatly represented. Finally, wesearchedspecifically for genes known toplay regulatory functionsduring thecell cycleand found thatmostof themareDE inourdataset (Supplemental Data Set 2E). Moreover, for many of them, wecould identify their homologs inArabidopsis, forwhich expressionor functional data are already available.

Expression and Targets of miRNAs in SynchronizedBY2 Cells

As hundreds of different transcripts showed highly dynamicpatternsofaccumulation,wequestionedwhethersRNAsmightbeinvolved in their regulation. We performed deep-sequencinganalyses on total sRNAs and GFP-AGO1–associated sRNAs inthe different cell cycle phases (Supplemental Table 2). AGO1-loaded sRNAs were isolated by immunoprecipitation (IP) witha GFP antibody (Supplemental Figure 7). As expected, we ob-served a size distribution of sRNAs in total RNA samples typical ofSolanaceae, that is, one in which 24-mers are the most abundantclass of sRNA, followed by 22- and 21-oligomers (SupplementalFigure 8A). By contrast, the AGO1-associated sRNAs werestrongly enriched in 21- and 22-nucleotide sRNAs (SupplementalFigure 8B), consistent with published AGO1 IP data (Mallory andVaucheret, 2010), and thespecific22-nucleotidepeakobserved inSolanaceaespecies, namely, tomato (Solanum lycopersicum; TheTomato Gene Consortium., 2012).

To investigate the distribution of sRNAs, we quantified theaccumulation of miRNAs, trans-acting siRNAs, hc-siRNAs, andtRFs along the cell cycle (Figure 6A). hc-siRNAswere consistentlythe most abundant category in total RNA samples, while miRNAsrepresented themostabundantcategory inAGO1-IPsamples.Weobserved no differences in trans-acting siRNAs or hc-siRNAs inour data, nor were there significant differences between the cellcycle phases for the proportion of different sRNA classes(SupplementalDataSet3). Therefore,we focusedonmiRNAs (thissection) and tRFs (the subsequent section).

For miRNA analysis, we used the workflow described inSupplemental Figure 9. A total of 312 different mature sequences(including those from 39 and 59 arms, i.e., 3p and 5p miRNAs),belonging to 192 miRNA precursors and 103 different miRNAfamilies (Supplemental Data Sets 4A to 4C), were identified in thisstudy, including the already-annotated miRNAs deposited inmiRBase as well as newly predicted miRNAs. This combined settotaled 221 distinct sequences. Most miRNAs (215 of 221) werepresent in both samples, total RNA and AGO1-IP. We found 10miRNAs differentially accumulated (DA) in total RNA and three inAGO1-IP (Figure6B).Mostof theDAmiRNAshadaphase-specificaccumulation. Six miRNAs showed a higher accumulation duringthe G2 phase, while only a single miRNA, miR168a-5p, wasspecific to M phase (Figure 6C). Two newly identified miRNAs(miR-33-3p and miR-12-5p) and two known miRNAs (miR6147a-3pandmiR479)weremoreabundant in theG1phase.Additionally,miR390-3p and miR482-5p levels peaked during S phase. In theAGO1-IP samples, we only found DA miRNAs when comparing

opposite phases of the cell cycle (S/M and G2/G1). Thus, onlya fewmiRNAs showed DAwhen comparing contiguous cell cyclephases.We next asked whether somemiRNAs identified in our analysis

could target cell cycle–regulated transcripts. To address thisquestion, we generated parallel analysis of RNA ends (PARE) li-braries for threebiological replicatesofeachphaseof thecell cycle(Supplemental Table 3). To minimize sample variability, we fo-cused on miRNA-target pairs present in all biological replicatesand supported with robust evidence of cleavage (i.e., a sRNA-PARE target analyzer–defined score # 3, see “Methods”;Supplemental Data Set 4D). Thus, we found 769 unique miRNA-target pairs considering all samples, corresponding to 124 dif-ferent miRNAs and 168 different genes (Figure 6D). Among those,we found a core subset of miRNAs and their respective targetgenes that were present in all phases of the cell cycle. This in-cludes 179 miRNA-target pairs, involving 51 miRNAs from 24families, and 31 target genes (Figure 6D). When considering onlygenes that are DE during the cell cycle, we identified 48 targetgenes, representing ;30% of the total miRNA-target pairs, butonly a small proportion of all the DE genes (Figure 6E;Supplemental Figure 10; Supplemental Data Sets 4E to 4H).The majority of the miRNA targets encode transcription factors

(TFs) from APETALA2/ETHYLENE RESPONSIVE FACTOR, MYB,SCARECROW, and GROWTH-REGULATING FACTOR TF fami-lies (Supplemental Data Set 4D). The elevated number of TFstargeted by miRNAs in all phases of the cell cycle suggests twolayers of transcriptional control during the cell cycle: TFs andmiRNAs. The second biggest group of target genes were diseaseresistance proteins (Supplemental Data Set 4D), suggesting thatmiRNAs may dampen their expression in proliferating cells. Fi-nally, we observed several phase-specific miRNA-target pairs,that is,miRNA-targetpairs thatwereonly found inonephaseof thecell cycle (Figure 6D; Supplemental Data Set 4).

Differential Accumulation of tRFs in Synchronized BY2 Cells

Knowing that tRFs can form a complex with AGO proteins withimportant cellular functions (Shigematsu and Kirino, 2015;Martinez et al., 2017), we investigated whether there were any DAtRFs in our samples. None of the tRFs derived from the 1529tRNAs analyzed showedDA in total RNA samples. However, tRFsoriginating from 67 different tRNAs were DA in the AGO1-IPsamples (Supplemental Data Sets 5A to 5C). These tRFs werederived from 14 different types of tRNAs (Figure 7A). Interestingly,each DA tRF-producing tRNA was associated with a differentphase of the cell cycle, with the exception of G1, which did notaccumulate high levels of DA tRFs (Figure 7B). To further char-acterize the role of the DA tRFs during the cell cycle, we analyzedtheir size, sequence, and targets. These DA tRFs had a sizedistributionpatterndifferent fromall tRFs (Figure 7C),with ahigherabundance at 19 nucleotides. We observed a conserved se-quence among the DA 19-mers (Figure 7D), and they lacked theCCA or poly(U) signatures typical of the 39 end of the tRNA. Weconfirmed this 59originationbyanalysis of their positionof origin intRNAs; they were thus classified as 59-end tRFs.Taking advantage of our PARE data, we looked for putative

targets for the identified tRFs. As described previously (Martinez

Cell Cycle–Dependent Accumulation of Small RNAs 1741

Figure 5. RNA-Seq Analysis of DE Genes during Cell Cycle.

(A) Summary of DE genes upon pairwise comparison of two cell cycle phases. Down- and upregulated genes are represented in purple and orange,respectively.(B) Heatmap representation of differentially regulated genes with a fold change greater than or equal to 5.(C)Heatmap representing the expression pattern of all clusters. Clusters are divided into six groups (I to VI) following their expression in each phase of thecell cycle.(D) GO Slim categories representing the biological function of DE genes. Here, only represented GO categories with more than 1.53 representation foldchange.

Figure 6. sRNAs and the Cell Cycle.

(A) sRNAabundanceper category. Five biological replicates of total RNAsamples (left) and threebiological replicates ofAGO1-IP samples (right). The xaxisrepresents the different sRNA categories included in this study, and the y axis represents their abundance (reads per million [RPM]).(B) miRNA fold change (FC) accumulation (in log2 scale) for the differentially accumulated (DA) miRNAs (*q-value # 0.05 and **q-value # 0.01).(C)Highly abundantmiRNAs in each cell cycle phase. In black are the DA genes identified in total RNA samples, and in blue are those identified in AGO1-IPsamples.(D)miRNA-target pairs in each phase of the cell cycle. Colors are indicative of the abundance, with dark gray representing themost abundant andwhite theleast abundant.(E) Overlap of the miRNA targets with genes DE in the RNA-seq. DE, differential expression.

Figure 7. tRFs and the Cell Cycle.

(A) tRF fold change (FC) accumulation (in log2 scale) for the DE tRNA-derived sRNAs (*q-value # 0.05, **q-value # 0.01).(B) High abundant tRNAs with DA fragments in each phase of the cell cycle from the AGO1-IP fractions.(C)Sizedistributionof tRFabundance indifferentsamples. Inblackare totalRNAsamples, in redareallAGO1-IPsamples,and ingreenonly theDAtRFs fromAGO1-IP samples. RPM, reads per million.(D) Logo representation of the most abundant size category (19-mers) of the differentially accumulated tRFs of AGO1-IP samples.(E) TE distribution of the tRF targets identified using PARE data. The two main categories are represented here: DNA and retrotransposons.(F) Retrotransposable elements targeted by tRFs identified using PARE data, divided into superfamilies. Represented here are the percentages of totalnumber of targeted retrotransposons (left) and the normalized number of retrotransposons (right).

et al., 2017), tRFs have the ability to target transposable elements(TEs). Most of the targets of the tRFs that we could validate byPARE analysis were retrotransposons (Figure 7E; SupplementalData Set 5D). Among these TEs, the majority was from the LongTerminal Repeat-Gypsy superfamily (Figure 7F, left). However,when normalizing to the total number of TEs of each superfamilypresent in the tobacco genome, short interspersed nuclear ele-ments (a category of non-LTR retrotransposons with an internalregion that originated from tRNAs) were enriched among the tRFtargets (Figure 7F, right). We next analyzed the expression of theTEs identifiedas tRF targets inourRNA-seqdataset.We identified381,452 expressed TEs in our RNA-seq data of 1,980,189 (;20%expressed). Of these, 18 (;0.005% of the identified TEs) were DE(Supplemental Data Set 5) in one or more cell cycle comparisons,and only one of these was targeted by a tRF (;0.0003% ofidentified TEs).

DISCUSSION

AGO1 Is Required for Cell Division Activity but RegulatesOnly a Subset of DE Cell Cycle Genes

In mammalian cells, the role of AGO proteins and miRNAs in theregulation of gene expression during the cell cycle has been wellestablished (Bueno and Malumbres, 2011). However, in plants,this question remained unclear despite the fact that defects inmiRNAbiogenesisor inpost-transcriptionalgenesilencing impactcell proliferation during embryonic and post-embryonic de-velopment. For example, mutation ofDCL1 alters embryogenesisat the globular stage, showing abnormal cell divisions throughoutthe suspensor and hypophysis (Schwartz et al., 1994; Nodine andBartel, 2010). Similarly, aberrant patterns of cell division and cellexpansion during embryogenesis have also been identified inmutants of Arabidopsis proteins that act together with DCL1,SERRATE, and HYPONASTIC LEAVES1 (Grigg et al., 2009;Armenta-Medina et al., 2017). The molecular basis of alteredembryogenesis, pattern formation, and abnormal cell divisions ishowever not yet well understood. Although dcl1 mutants exhibitaltered auxin responses (Grigg et al., 2009; Nodine and Bartel,2010; Seefried et al., 2014) and several miRNAs target AUXINRESPONSE FACTORs (ARFs; Jones-Rhoades and Bartel, 2004;Armenta-Medina et al., 2017), a causal link between auxin sig-naling and defects of embryogenesis in these mutants has notbeen established. To investigate whether miRNAs are required tomaintain cell proliferations in meristems at a post-embryonicstage, we depleted AGO1 by targeted degradation using the vi-ral encoded F-box protein P0. Under these conditions, depletionof AGO1 impaired meristem size and cell division activity in theroot apical meristem. According to our cell cycle reporters, cellsdid however not arrest in the S or G2/M phases of the cell cycle,suggesting that they remained in G1.

To investigate the repertoire of miRNAs that are expressed atspecific cell cycle phases and identify their targets, we chose towork with the tobacco BY2 cell suspension culture (Nagata et al.,1992), offering optimal synchronization efficiencies. The analysisof total and AGO1-associated miRNAs revealed that only a fewmiRNAsshowedadifferential accumulationpatternduring thecell

cycle. This situation is very different from mammals for which thetranscriptionof somemiRNAs is cell cycle regulated. For example,some miRNAs are DE through direct activation by E2F and otherTFs (Sylvestre et al., 2007; Woods et al., 2007; Bueno et al., 2010)and degraded in a cell cycle–dependent manner by specific en-donucleases (Elbarbary et al., 2017). In addition,whilemammalianmiRNAs target numerous cell cycle transcripts at different cellcycle stages, this is not the case for plant miRNAs. Our analysisshows that miRNAs target only a subset of genes DE during thecell cycle and none of them correspond to core cell cycle genes(Supplemental Figure 10; Supplemental Data Set 4). A likely ex-planation of such a difference is that themode of action of RISC ingenesilencingdiffersbetweenplantandanimal cells. Even thoughplant AGO1 is able to perform translational repression (Brodersenet al., 2008; Li et al., 2013), most of its activity is in slicing targettranscripts. As this process requires a perfect or near-perfectcomplementarity between a miRNA and its mRNA target, itmight preclude the possibility that a unique miRNA targets a widespectrum of genes, as is the case in mammals (Friedman et al.,2009). Nonetheless, according to our PARE data requiringcleavage to be detected, we presently cannot exclude the pos-sibility that some plant miRNAs might also control the translationof DE genes during the cell cycle.Yet, our data support that plant miRNAs indirectly contribute to

the control of cell proliferation by targeting some important reg-ulators. Indeed, we found that most of miRNA targets are tran-scription factors, somebeingexpressed in acell cycle–dependentmanner (Supplemental Data Set 4; Supplemental Figure 10). Forexample, we found that miR164 and miR160 target the NAC TFCUP-SHAPED COTYLEDON2 and ARFs, respectively, in a cellcycle–dependent manner. CUP-SHAPED COTYLEDON2 pro-motes the generation of auxin response maxima and thereforecould release ARF repression, triggering downstream auxin sig-naling components suchascyclins andCDKs.Also interesting aresome miRNAs that target F-box domain containing transcripts,including AUXIN SIGNALING F-BOX 2. Two F-box encodinggenes are targeted by members of the miR6149 family and willdeserve particular attention as their expression is enriched in Sphase. Although they belong to a different subclade than theArabidopsis F-box FBL17, a key player of theG1/S transition (Noiret al., 2015), theycouldbe involved in the turnoverof important cellcycle regulatory proteins.

Plant miRNAs Repress Defense Genes during the Cell Cycle

Themolecular links between the cell cycle anddefense responsesin plants are still not well understood, but surprising connectionshavealreadybeenpointedout (Zebell andDong,2015).Microarraydata fromsynchronizedArabidopsis cell suspension revealed thatsome resistance genes are DE during the cell cycle, with a peak ofexpression either in theSorMphase (Menges et al., 2005), andwealso identified100of thoseDEresistancegenes fromourRNA-seqlibraries. Interestingly, it has been shown that cell cycle mis-regulation can lead to the activation of disease resistancegenes (Bao et al., 2013). For example, overexpression of OMIS-SION OF SECOND DIVISION1 or UV-B-INSENSITIVE4, twonegative regulators of the anaphase-promoting complex or cy-closome, results in theexpressionof thedefensemarkergenePR1

Cell Cycle–Dependent Accumulation of Small RNAs 1745

and confers resistance to virulent pathogens. Conversely, lossof these two proteins, as well as the APC10 subunit or theCCS52 anaphase-promoting complex or cyclosome coactivator,led to enhanced susceptibility to bacterial infection (Bao et al.,2013). In addition, the loss of the resistance gene SUPPRESSOROF npr1-1 CONSTITUTIVE1 abolished pathogen resistanceconferred by the osd1 mutation. On the other hand, plantpathogens and beneficial symbionts can also modulate the cellcycle activity by inducing or suppressing cell cycle gene ex-pression (Favery et al., 2002). Taken together, these data suggestthat the balance between cell proliferation and defense againstpathogens implies complex regulatory circuits.

What our study showed is that genes encoding disease re-sistance proteins represent the second main group of miRNAtargets. While many of these miRNA targets are constitutivelyexpressed, some are DE during the cell cycle (SupplementalFigure10;SupplementalDataSet4). Therefore, onecouldassumethat miRNAs in proliferating cells or even at specific cell cyclestages repress the expression of numerous resistance genes andare thus key for the antagonistic regulation of cell cycle and de-fense gene expression programs.

A Subset of tRFs Differentially Associate to AGO1 during theCell Cycle

In contrast to other sRNAs, tRF abundances were surprisinglydifferent between total sRNAs and AGO1-associated data sets.This suggests that tRF biogenesis is most likely constant duringcell cycle,while loading inAGO1of at least some tRFsmust be cellcycle regulated. It is already known that some plant tRFs accu-mulate in a wide range of stresses including oxidative stresses(Thompson et al., 2008) and UV stresses (Gebetsberger et al.,2012) that induce deleterious effects such as DNA damage. In-terestingly, some tRFs that differentially accumulated in our li-braries originate from the tRNAs that are also processed uponoxidative stresses, particularly tRNA-Arg (CCT) and tRNA-Glu(CTC; Figure 7; Supplemental Data Set 5). In addition, 11 tRFsamong the 14 DE tRFs appear to be enriched in S and G2, sug-gesting that they could be linked to DNA replication and/or G2checkpoint processes.

Notably, most of the 19-mer tRFs associated with AGO1 havea G or a C at the 59 position (Figure 7; Supplemental Data Set 5),while it is established thatAGO1hasapreference for a59UorA (Miet al., 2008). This might explain why tRFs represent such a smallsubset of AGO1-associated sRNAs. Moreover, the presence ofG/C at the tRFs 59 end might also affect the stability of the RISCcomplex and therefore further explain whywe have detected highfoldchanges forAGO1-associated tRFswhileothersRNAsremainoverall unchangedduring cell cycle. This raises thequestion of thebiological significanceof these tRFsdifferentially accumulated, aswell as their mode of action.

To examinewhether tRFs are able to triggermRNA slicingwhileassociated with RISC complex, we investigated the PARE sig-natures that match with tRF sequences. We found that Gypsyelements are the most abundant target for tRFs (Figure 7;Supplemental Data Set 5); this corresponds to what was found inpreviousstudies inArabidopsis (Martinezetal., 2017).On theotherhand, short interspersed nuclear elements were also abundant

targetsof tRFs. Thismightbedue to their highlyconserved internalregions that originated from tRNAs.Besides slicing functions, some tRFs have also been shown to

induce translation inhibitionof some transcripts. For example, Val-tRF from the archaea Haloferax volcanii associates to the smallribosomal subunit and is able to inhibit protein synthesis by in-terfering with peptide bond formation, presumably to fine-tuneoverall protein synthesis in response to several stresses(Gebetsberger et al., 2012). A similar mechanism of tRF-mediatedtranslational inhibition in HeLa cells has also been describedpreviously (Sobala and Hutvagner, 2013). The 39GG dinucleotideof these tRFs is essential for their function. Interestingly, thismotifis also represented in our AGO1-associated tRF sequences(Figure 7D), but whether they are involved in translational re-pression in plants requires further investigation.

METHODS

Constructs

For the pAGO1:GFP-AGO1 construct, the pAGO1:GFP-AGO1 describedinDerrien et al. (2012)was transferred from thepGREEN II to abinary vectorwith plant kanamycin resistance for transformant selection. The sequencewas amplified using primers FOR-attB1 (GGGGACAAGTTTGTACAAAAAAGCAGGCTTAACGACGGCCAGTGAATTGTAATACGA) and REV-attB2(GGGGACCACTTTGTACAAGAAAGCTGGGTAGCTATGACCATGATTACGCCAAGCT) and used for recombination in the pDONR221 vector(Invitrogen). The LR clonase reaction was performed to transfer the con-struct in the pKGW vector backbone (VIB).

For the pCYCB1;2:CYCB1.2(Dbox)-GFP (AT5G06150) construct,a genomic fragment of 1929 bp containing 1090 bp of promoter sequence,the 59 untranslated region, and 839 bp from the coding sequence includingthe first intron was amplified from wild-type Columbia-0 genomic DNAusing primers FOR-attB1 (GGGGACAAGTTTGTACAAAAAAGCAGGCTTAAGACTTACTCTGATCTTCAACGCCAA) and REV-attB2 (GGGGACCACTTTGTACAAGAAAGCTGGGTACTTAGGTGACATCGCTACTTCCTT)and used for recombination in the pDONR221 vector (Invitrogen).pCYCB1;2:CYCB1;2(Dbox) and pHTR2:CDT1a (C3) (Yin et al., 2014) entryclones were recombined with pGWB650 binary vector (RIKEN) to createa C-terminal fusion to the enhanced GFP.

Plant Material

Arabidopsis (Arabidopsis thaliana) missense mutant alleles ago1-36(Baumberger and Baulcombe, 2005) and ago1-27 (Morel et al., 2002) havebeen described previously. The XVE:P0 and the XVE:P0 (ago1-57) transgeniclines are described by Derrien et al. (2018). Both lines were transformed withthe pHTR2:CDT1a (C3)-GFP and pCYCB1;2:CYCB1.2(Dbox)-GFP con-structs, and double homozygous plants for both transgenes were obtainedand used for further experiments.

For in vitro culture conditions, Arabidopsis seedswere surface sterilizedusingethanol andplatedongrowthmedium (MES-bufferedMurashigeandSkoog saltsmedium [Duchefa], 1% (w/v) Suc, and 0.8 or 1% (w/v) agar, pH5.7) in the presence of selective agent when appropriate. Seeds werestratified for 2 d at 4°C in the dark and then transferred in 16-h-light/8-h-dark (at 20.5/17°C, 70%humidity) growth chamber, under fluorescent light(Osram Biolux 58W/965). For P0 induction, seedlings were grown onmedium supplemented with 10 mM b-estradiol or supplemented with anequal volume of DMSO for the mock treatment.

1746 The Plant Cell

BY2 Cell Culture, Transformation, and Synchronization

A rapidly growing suspension culture of tobacco BY2 cells (Nicotianatabacum cv Bright Yellow 2) was maintained by weekly dilution (1.5:80) ofcells into fresh medium according to Nagata et al. (1992) and cultured at27°C with shaking at 130 rpm in the dark. The pAGO1:GFP-AGO1 (Derrienet al., 2012) and H2B-tdTomato constructs (Adachi et al., 2011) were in-troducedby electroporation into thedisarmedAgrobacterium tumefaciensstrain LBA4404, and the resulting Agrobacterium strains were used totransform BY2 cells. Four milliliters of a 3-d-old BY2 culture was co-cultivated with 100 mL of an overnight culture of Agrobacterium in Petridishes in the dark for 2 d at 27°C. Cells were then collected and washedthree times with BY2 culture medium by centrifugation and were plated onsolid medium supplemented with carbenicillin (500 mg mL21), cefotaxime(500 mg mL21), and hygromycin (30 mg mL21). In order to establish clonalcell cultures for both pAGO1:GFP-AGO1 and pAGO1:GFP-AGO1;pRPS5a:H2B-tdTomato, at least 4 to 6 weeks was required before callicould be recovered. Individual calli were subcultured into liquid mediumcontaining carbenicillin (500 mg mL21), cefotaxime (500 mg mL21), andhygromycin (30 mg mL21).

TobaccoBY2cellswere synchronizedaccording toNagataet al. (1992).Clonal BY2 cell suspensions 131 and 408 were used for Figure 3 andSupplemental Figure 3, respectively. Note that the cell suspension 408expresses slightlymoreGFP-AGO1protein thancell suspension131, likelyexplaining the delay in synchronization. For determination of the mitoticindex, cells were stained with 0.2mgmL21 49,6-diamidino-2-phenylindole(Sigma-Aldrich) in the presence of 0.2% Triton X-100; interphase, pro-phase, metaphase, anaphase, and telophase figures were determined forat least 600 cells by using UV light microscopy. Flow cytometry analysis ofthe tobacco BY2 cell suspension was performed as described previously(Noir et al., 2015). Measurements with at least two biological replicateswere performed.

Root Length Analysis and Confocal Imaging

After 6 to 12 d of growth in vitro in the vertical orientation, the plates werescanned and the root length of Arabidopsis wild type, mutants, andtransgenic lines was measured with ImageJ. For root meristem meas-urements and the imaging of the root tip, roots were mounted on a mi-croscopic slide in a 75 mg/mL propidium iodide solution. Root tips wereimaged under the confocal microscope (Leica TCSSP8) in the plane of thequiescent center.GFPwasdetectedusing argon laser excitation at 488nmand through 505 to 550 nm emission filter-set. Propidium iodide wasdetected using diode-pumped solid state laser excitation at 561 nm andthrough 600 to 630 nm emission filter-set. Imaging was performed usingHCXAPOCS (203 -0.7) objective lens. Themeristemsizewasmeasured inImageJ as the distance between the quiescent center and the last dividingcell of the cortex. To assess the statistical significance, analysis of variance(ANOVA) tests were performed using the R software.

To image GFP-AGO1 in transformed BY2 cells (clonal cell suspensions304), confocal imageswereobtainedusingargon laser excitationat488nmand through 505 to 550 nm emission filter-set. H2B-tdTomato was de-tected with DSSP laser excitation at 561 nm and through 575 to 650 nmemission filter-set. Imaging was performed using HCX APOCS (203 - 0.7)air, HC PL APO CS2 (403 - 1.1) water, and HC PL APO CS2 (633 - 1.4) oilobjective lens. Images are presented as single sections. Differential in-terference contrast (Nomarski) was used for transmission light images.

RNA Analyses by RT-qPCR

RNAextractionwasperformedondriedmaterial vacuum filtered from2mLof BY2 cell suspension using Tri-Reagent (Sigma-Aldrich) according to themanufacturer’s instructions. For quantitative RT-PCR, 2 mg of total RNAtreated with DNase I (Thermo Fisher Scientific) were reverse transcribed

using a High-Capacity cDNA reverse transcription kit (Thermo FisherScientific). qPCR reactions were performed in a total volume of 10 mL ofLightCycler 480 SYBR GREEN 1 mix (Roche) on a LightCycler LC480apparatus (Roche) according to themanufacturer’s instructions. Themeanvalues of at least three biological replicates were normalized using N.tabacum ubiquitin-conjugating enzyme E2 (AB026056), protein phos-phatase 2A (X97913), andL25 ribosomal protein (L18908) genes as internalcontrols (Schmidt and Delaney, 2010).

Protein Analysis and Immunoblotting

Total proteins were extracted from tobacco BY2 cells and Arabidopsisseedlings using a phenol extraction protocol. Frozen material was groundand resuspended in extraction buffer (0.7Msaccharose, 500mMTris-HCl,pH 8.5, 5 mM EDTA, pH 8, 100 mM NaCl, 2% b-mercaptoethanol, andcOmplete mini protease inhibitor [Roche]). The lysate was incubated withan equal volume of biophenol, pH 8.0, under agitation. Proteins wereprecipitated from the phenol phase with 5 volumes of cold methanol and100mMammoniumacetate.Pelletswerewashed twicewithcoldmethanoland resuspended in a buffer (10% glycerol, 3%SDS, and 62mM Tris-HCl,pH 8). Prior to loading, protein samples were supplemented with 43loading buffer (40% glycerol, 16% SDS, 250 mM Tris-HCl, pH 8, and 20%b-mercaptoethanol) to 13 final concentration and incubated at 95°C for2 min. Ten to 30 mg of total protein extracts was separated on SDS-PAGEgels and blotted onto Immobilon-Pmembrane (Millipore). GFP-AGO1 wasdetected using anti-GFP antibody (mouse monoclonal JL-8, Clontech)diluted to 1:2000; Nicotiana tabacum AGO1 (Nta AGO1) was detectedusing anti-N.benthamianaAGO1antibody (rabbit polyclonal; Csorba et al.,2010) diluted to 1:10,000 (v/v); Nta CYCLIN B1was detected using anti-N.tabacumCyclinB1 antibody (rabbit polyclonal; Criqui et al., 2000) diluted to1:1000 (v/v); and ACTIN protein was detected using anti-ACTIN antibody(mouse monoclonal AS163141, Agrisera) diluted 1:20,000 (v/v). Arabi-dopsis AGO1andAGO2were detected using anti-AGO1 (rabbit polyclonalAS09527, Agrisera) diluted to 1:10,000 (v/v) and anti-AGO2 (rabbit poly-clonal AS132682, Agrisera) diluted to 1:2000 (v/v), respectively.

AGO1 Immuno-precipitation

Onegramof frozenBY2cellswasground and resuspended in 3 volumes ofextraction buffer (50 mM Tris-HCl, pH 7.5, 300 mM NaCl, 10% glycerol,5 mM MgCl2, 0.5% Triton X-100, 5 mM DTT, and cOmpete mini proteaseinhibitor [Roche]). The lysate was incubated with 50 mL of GFP-Trapmagnetic beads (Chromotek) for 1 h at 4°C on a rotary shaker. Beadswere washed four times with extraction buffer on a magnetic stand. RNAwas recovered from IP fractions using Tri-Reagent (according to themanufacturer’s instructions) and resuspended in ultrapure water (Sigma-Aldrich). For proteins, one tenth of themagnetic beadswas resuspended in13 loading buffer (10% glycerol, 4% SDS, 50 mM Tris-HCl, pH 8, and 5%b-mercaptoethanol) and incubated at 95°C for 2 min.

Library Preparation and Sequencing

For this study, a total of 40 sRNA libraries (only the 32 with at least 10 Mmapped reads were ultimately used), 12 RNA libraries, and 12 PARE li-brarieswere constructed. sRNA librarieswere constructed using total RNAand AGO1-IP RNAs, using TruSeq sRNA Sample Preparation kit (Illumina)according tomanufacturer’smanual. RNA librarieswereconstructedusingTruSeq Stranded mRNA Preparation kit (Illumina). These libraries weresequenced using Illumina HiSeq 2500 system and Illumina HiSeq 4000system, respectively, by Fasteris SA. PARE libraries were constructedusing total RNA following the protocol described in Zhai et al. (2014) andsequenced using Illumina HiSeq 2500 system at the Delaware Bio-technology Institute.

Cell Cycle–Dependent Accumulation of Small RNAs 1747

Data Analysis

Sequencing data quality was assessed using FastQC. Reads were pro-cessed by removing the adaptor sequences using Trimmomatic (Bolgeret al., 2014) and mapped to the N. tabacum genome version 4 (https://solgenomics.net/organism/Nicotiana_tabacum/genome; Edwards et al.,2017) using Bowtie (Langmead and Salzberg, 2012).

For sRNA analysis, a modified version of miREAP was used as de-scribed in Arikit et al. (2014) for miRNA prediction. Only miRNA precursorswith reads present in both arms and mature miRNAs 21 nucleotides or 22nucleotides longwereselected.NewmiRNAcandidateswere thenused forhomology search using miRBase v22 (Kozomara and Griffiths-Jones,2014), with a maximum difference of four nucleotides. To analyzehc-siRNAs, we first ran RepeatMasker (http://www.repeatmasker.org)using the Viridiplantae RepBase database (http://www.girinst.org/repbase)as library. For RNA-seq, analyses were performed using the HISAT2 andStrigtTie as described in Pertea et al. (2016). Clustering analysis was doneusingMFuzz v3.6. GOanalyseswere done using the Arabidopsis orthologsof tobacco genes and PANTHER v11 software (Mi et al., 2017) with defaultparameters. PARE libraries were analyzed using sRNA-PARE target ana-lyzer (Kakrana et al., 2014) and CleaveLand v4 (Addo-Quaye et al., 2009).Three biological replicates of each cell cycle phase were used as startingmaterial. Only miRNA-target pairs present in all biological replicates, witha score of 3 or lower, were considered for further analysis. All statisticalanalyses were performed with DESeq2 (Love et al., 2014) using defaultparameters and raw counts, establishing the significant threshold ata q-value < 0.05.

Accession Numbers

The data discussed in this article have been deposited in the NationalCenter for Biotechnology Information’s Gene Expression Omnibus (Edgaret al., 2002) and are accessible through Gene Expression Omnibus SeriesaccessionnumberGSE102344 (https://www.ncbi.nlm.nih.gov/geo/query/acc.cgi?acc5GSE102344).

Supplemental Data

Supplemental Figure 1. Loss of Arabidopsis AGO1 affects rootdevelopment.

Supplemental Figure 2. Selection of clonal BY2 cell suspensions withmoderate GFP-AGO1 expression.

Supplemental Figure 3. Accumulation pattern of AGO1 mRNA andprotein in synchronized BY2 cells.

Supplemental Figure 4. Quality control of five independent synchro-nization experiments conducted with the GFP-AGO1 clonal BY2 cellsuspension used for high-throughput sequencing.

Supplemental Figure 5. Analysis of the RNA-seq data.

Supplemental Figure 6. Cluster analysis of RNA expression data.

Supplemental Figure 7. RNA-immunoprecipitation (RIP) of GFP-AGO1 loaded small RNAs in BY2 tobacco cell suspension.

Supplemental Figure 8. Size distribution of small RNAs.

Supplemental Figure 9. Workflow including all steps followed formiRNA identification, as well as the numbers of known and newmiRNAs identified.

Supplemental Figure 10. List of miRNA-target pairs only for DE genesduring the cell cycle and for which homologs have been identified inArabidopsis.

Supplemental Table 1. RNA-seq libraries mapping statistics.

Supplemental Table 2. Small RNA libraries mapping statistics.

Supplemental Table 3. PARE libraries mapping statistics.

Supplemental Data Set 1. Gene expression during cell cycleprogression.

Supplemental Data Set 2. List of transcripts indicated in green andblue in Supplemental Figure 5C.

Supplemental Data Set 3. List of the identified small RNAs mappingon transposable elements and TAS loci, including their expression foldchange between phases.

Supplemental Data Set 4. List of the identified miRNAs in total RNAand AGO1-IP samples and list of miRNA-target pairs.

Supplemental Data Set 5. List of the DE tRFs in AGO1-IP samplesand list of targets of the tRFs validated by PARE analysis.

ACKNOWLEDGMENTS

We thank Masaaki Umeda for the gift of the pHTR2:CDT1a (C3) plasmid,Sachihiro Matsunaga for the gift of the pMDC99-proRPS5A-H2B-tdTomato plasmid, and József Burgyán for the N. benthamiana AGO1antibody. Support was provided by the European Research Council underthe European Union’s Seventh Framework Programme (grant FP7/2007-2013 to P.G.), the European Research Council (advanced grant 338904 toP.G.), and from LABEX (grant ANR-10-LABX-0036_NETRNA). Supportwas also provided by the National Science Foundation Integrative Organ-ismal Systems (award 1842698 toB.C.M. andP.B.) from thePlantGenomeResearch Program.

AUTHOR CONTRIBUTIONS

A.T., P.B., B.C.M., and P.G. designed research; A.T., P.B., M.-C.C., M.D.,and M.C. performed research; A.T., P.B., B.C.M., and P.G. analyzed thedata; A.T., P.B., and P.G. wrote the article, with help fromM.D. and B.C.M.

Received January 31, 2019; revised May 7, 2019; accepted June 7, 2019;published June 12, 2019.

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1750 The Plant Cell

DOI 10.1105/tpc.19.00069; originally published online June 12, 2019; 2019;31;1734-1750Plant Cell

and Pascal GenschikAdrien Trolet, Patricia Baldrich, Marie-Claire Criqui, Marieke Dubois, Marion Clavel, Blake C. Meyers

Dependent Regulation and Function of ARGONAUTE1 in Plants−Cell Cycle

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