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Title 1 Autophagy promotes organelle clearance and organized cell separation of living root cap 2 cells in Arabidopsis thaliana 3 4 Running title 5 Role of autophagy in root cap 6 7 Authors 8 Tatsuaki Goh1,§,*, Kaoru Sakamoto1,§, Pengfei Wang2, Saki Kozono1, Koki Ueno1, 9 Shunsuke Miyashima1, Koichi Toyokura3, Hidehiro Fukaki3, Byung-Ho Kang2, Keiji 10 Nakajima1,* 11 12 Affiliations 13 1Graduate School of Science and Technology, Nara Institute of Science and Technology, 14 8916-5 Takayama, Ikoma, Nara 630-0192, Japan. 15 2School of Life Sciences, Centre for Cell & Developmental Biology and State Key 16 Laboratory of Agrobiotechnology, The Chinese University of Hong Kong, Shatin, New 17 Territories, Hong Kong, China. 18 3Department of Biology, Graduate School of Science, Kobe University, Rokkodai, Kobe 19 657-8501, Japan 20 §These authors contributed equally. 21 22 1 *Corresponding authors: 23 Tatsuaki Goh <[email protected]> and Keiji Nakajima <[email protected]> 24 25 Keywords 26 Arabidopsis thaliana, amyloplast, autophagy, cell separation, root cap 27 28 Summary statement 29 Time-lapse microscope imaging revealed spatiotemporal dynamics of intracellular 30 reorganization associated with functional transition and cell separation in the Arabidopsis 31 root cap and the roles of autophagy in this process. 32 33 34 2 Abstract 35 The root cap is a multi-layered tissue covering the tip of a plant root that directs root 36 growth through its unique functions such as gravity-sensing and rhizosphere interaction. 37 To prevent damages from the soil environment, cells in the root cap continuously turn 38 over through balanced cell division and cell detachment at the inner and the outer cell 39 layers, respectively. Upon displacement toward the outermost layer, columella cells at 40 the central root cap domain functionally transition from gravity-sensing cells to secretory 41 cells, but the mechanisms underlying this drastic cell fate transition are largely unknown. 42 By using live-cell tracking microscopy, we here show that organelles in the outermost 43 cell layer undergo dramatic rearrangements, and at least a part of this rearrangement 44 depends on spatiotemporally regulated activation of autophagy. Notably, this root cap 45 autophagy does not lead to immediate cell death, but rather is necessary for organized 46 separation of living root cap cells, highlighting a previously undescribed role of 47 developmentally regulated autophagy in plants. 48 3 Introduction 49 50 The root cap is a cap-like tissue covering the tip of a plant root. The root cap protects the 51 root meristem where rapid cell division takes place to promote root elongation (Arnaud 52 et al., 2010; Kumpf and Nowack, 2015). The root cap is also responsible for a number of 53 physiological functions, such as gravity-sensing to redirect the root growth axis (Strohm 54 et al., 2012), and metabolite secretion for lubrication and rhizosphere interaction 55 (Cannesan et al., 2012; Driouich et al., 2013; Hawes et al., 2016; Maeda et al., 2019). In 56 addition to its unique functions, the root cap exhibits a striking developmental feature, 57 namely continuous turnover of its constituent cells (Fig. 1A) (Kamiya et al., 2016). This 58 cell turnover is enabled by concerted production and detachment of cells at the inner stem 59 cells layer and the outer mature cell layer, respectively. Notably, the outermost root cap 60 cells detach from the root tip and disperse into the rhizosphere, creating a unique 61 environment at the border between the root and the soil. For this, detaching root cap cells 62 are called "border cells" (Hawes and Lin, 1990). Cell turnover is commonly seen in 63 animals but rarely found in plants where morphogenesis relies not only on the production 64 of new cells but also on the accumulation of mature and sometimes dead cells. Thus, the 65 root cap serves as a unique experimental material to study how plant cells dynamically 66 change their morphology and functions during tissue maintenance. 67 In the model angiosperm Arabidopsis thaliana (Arabidopsis), the root cap is 68 composed of two radially organized domains, the central columella and the surrounding 69 lateral root cap (LRC) that together constitute five to six cell layers along the root 70 4 proximodistal axis (Fig. 1) (Dolan et al., 1993). In Arabidopsis, the outermost root cap 71 cells do not detach individually, but rather separate as a cell layer (Fig. 1) (Driouich et al., 72 2007; Kamiya et al., 2016; Vicre et al., 2005). Previous studies revealed that detachment 73 of the Arabidopsis root cap cells is initiated by localized activation of programmed cell 74 death (PCD) at the proximal LRC region, and requires the functions of the NAC-type 75 transcription factor SOMBRERO (SMB), a master regulator of root cap cell maturation 76 (Bennett et al., 2010; Fendrych et al., 2014; Willemsen et al., 2008; Xuan et al., 2016). 77 While SMB is expressed in all root cap cells and acts as a master regulator of cell 78 maturation in the root cap, two related NAC-type transcription factors, BEARSKIN1 79 (BRN1) and BRN2, are specifically expressed in the outer two cell layers of the root cap 80 (Bennett et al., 2010; Kamiya et al., 2016). BRN1 and BRN2 share high sequence 81 similarities and redundantly promote the separation of central columella cells. Cell 82 separation in plants requires partial degradation of cell walls. Indeed, ROOT CAP 83 POLYGLACTUROSE (RCPG) gene encoding a putative pectin-degrading enzymes acts 84 downstream of BRN1 and BRN2, and at least BRN1 can directly bind to the RCPG 85 promoter (Kamiya et al., 2016). CELLULASE5 (CEL5) gene encoding a putative 86 cellulose-degrading enzyme is also implicated in cell separation in the root cap (Bennett 87 et al., 2010; del Campillo et al., 2004). 88 Previous electron microscopic studies reported profound differences in the 89 intracellular organization between the inner and the outer root cap cells of Arabidopsis 90 (Maeda et al., 2019; Sack and Kiss, 1989). As expected from their gravity-sensing 91 functions, columella cells in the inner layers accumulate large amyloplasts. Amyloplasts 92 5 are specialized plastids containing starch granules and known to act as statoliths in the 93 gravity-sensing cells (statocytes) in both roots and shoots (Gilroy and Swanson, 2014). 94 In contrast, columella cells constituting the outermost root cap layer do not contain large 95 amyloplasts, and instead accumulate secretory vesicles (Maeda et al., 2019; Poulsen et 96 al., 2008). Thus, the observed difference in subcellular structures correlates well with the 97 functional transition of columella cells from gravity-sensing cells to the secretory cells 98 (Blancaflor et al., 1998; Maeda et al., 2019; Vicre et al., 2005). Before detachment, the 99 outermost root cap cells contain a large central vacuole, likely for the storage of various 100 metabolites (Baetz and Martinoia, 2014). In addition, a novel role of cell death promotion 101 has been proposed for the large central vacuole in the LRC cells (Fendrych et al., 2014). 102 In eukaryotes, dispensable or damaged proteins and organelles are degraded by 103 a self-digestion process called autophagy (Mizushima and Komatsu, 2011). Autophagy 104 initiates with expansion of isolated membranes, which subsequently form spherical 105 structures called the autophagosomes and engulf target components. In later steps, 106 autophagosomes fuse with vacuoles, and the content of autophagosomes degraded by 107 hydrolytic enzymes stored in the vacuole. When eukaryotic cells are subjected to stress 108 conditions such as nutrient starvation, autophagy is activated to recycle nutrients and 109 maintain intracellular environments in order to sustain the life of cells and/or individuals 110 (Mizushima and Komatsu, 2011). Autophagy plays an important role not only in stress 111 response but also in development and differentiation, as autophagy-deficient mutants are 112 lethal in a variety of model organisms including yeast, nematode, fruit fly, and mouse 113 (Mizushima and Levine, 2010). Genes encoding central components of autophagy, the 114 6 core ATG genes, are conserved in the Arabidopsis genome (Hanaoka et al., 2002; Liu and 115 Bassham, 2012). However, under normal growth conditions, autophagy-deficient 116 Arabidopsis mutants grow normally except for early senescence (Hanaoka et al., 2002; 117 Yoshimoto et al., 2009). Thus roles of autophagy in plant growth and development remain 118 largely unknown. 119 In this study, we revealed morphological and temporal dynamics of 120 intracellular rearrangement that enable the functional transition of the root cap cells in 121 Arabidopsis by using motion-tracking time-lapse imaging. We also found that the 122 autophagy-deficient Arabidopsis mutants are defective in cell clearance and vacuolization 123 of the outermost root cap cells. Unexpectedly, the autophagy-deficient mutants are 124 impaired in the organized separation of the outermost root cap layer. Thus our study 125 revealed a novel role of developmentally regulated autophagy in the root cap 126 differentiation and functions. 127 128 129 Results 130 131 Outermost columella cells undergo rapid organelle rearrangement before cell 132 detachment 133 While previous electron microscopic studies have revealed profound differences in 134 intracellular structures between the inner and the outer root cap cells (Maeda et al., 2019; 135 Poulsen et al., 2008; Sack and Kiss, 1989), spatiotemporal dynamics of subcellular 136 7 reorganization in the root cap cells has not been analyzed, due to a difficulty in performing 137 prolonged time-lapse imaging of the root tip that quickly relocates as the root elongates. 138 To overcome this problem, we developed a motion-tracking microscope system with a 139 horizontal optical axis and a spinning disc confocal unit. A similar system has been 140 reported by another group (von Wangenheim et al., 2017). Our microscope system 141 enabled high-magnification time-lapse confocal imaging of the tip of vertically growing 142 roots for up to six days, allowing visualization of cellular and subcellular dynamics of 143 root cap cells during three consecutive detachment events (Supplementary Fig. S1). 144 Under our experimental conditions, the outermost root cap layer of wild-type 145 Arabidopsis sloughed off with a largely fixed interval of about 38 hours (h) 146 (Supplementary Fig. S1F). This periodicity is comparable to that reported for roots 147 growing on agar plates (Shi et al., 2018), indicating that our microscope system does not 148 affect the cell turnover rate of the root cap. Bright-field observation revealed that the cell 149 detachment initiates in the proximal LRC region and extends toward the central columella 150 region (Fig. 1 and Fig. S1A-S1D). In concert with the periodic detachment of the 151 outermost layer, subcellular structures of the neighboring inner cell layer (hereafter called 152 the second outermost layer) rearranged dynamically (Fig. 2A and Supplementary Movie 153 S1). Before the detachment of the outermost layer, columella cells in the inner three to 154 four cell layers contained large amyloplasts that sedimented toward the distal (bottom) 155 side of the cell (Fig. 2A, -4 h, light blue arrowheads), whereas those in the outermost 156 layer were localized in the middle region of the cell (Fig. 2A, -4 h, dark blue arrowhead). 157 A few hours after the outermost layer started to detach at the proximal LRC region, the 158 8 amyloplasts in the second outermost layer relocated toward the middle region of the cell, 159 resulting in a similar localization pattern to those of the outermost layer (Fig. 2A, 0.5 h, 160 dark blue arrowheads). Toward the completion of the cell separation, rapid vacuolization 161 and shrinkage of amyloplasts took place in the outermost layer (Fig. 2A, 18 h, green 162 arrowhead). 163 By using plants expressing nuclear-localized red fluorescent proteins 164 (DR5v2:H2B-tdTomato), we could also visualize dynamic relocation of nuclei, as well as 165 its temporal relationship with amyloplast movement (Fig. 2B and Supplementary Movie 166 S2). In the second outermost layer, nuclei relocated from the proximal (upper) to the 167 middle region of each cell about a few hours before the neighboring outermost layer 168 initiated detachment (Fig. 2B, -8 h, red arrowhead). This nuclear migration was followed 169 by the relocation of amyloplasts around the time when the neighboring outermost layer 170 initiated detachment at the proximal LRC region (Fig. 2B, 0 h, dark blue arrowhead). In 171 later stages, the amyloplasts surrounded the centrally-localized nucleus (Fig. 2B, 13 h, 172 dark blue arrowhead). In the outermost cells, nuclei migrated further to localize to the 173 distal pole of the cell (Fig. 2B, 13 h, purple arrowheads). 174 Dynamic change in vacuolar morphology was also visualized using plants 175 expressing a tonoplast marker (VHP1-mGFP) (Segami et al., 2014) (Supplementary Fig. 176 S2 and Supplementary movie S3). Vacuoles in the inner columella cells were smaller and 177 spherical, whereas those in the outer cells were larger and tubular (Supplementary Fig. 178 S2, 5-23 h). Notably, in the outermost layer, vacuoles were dramatically enlarged, and 179 eventually occupied the entire volume of detaching root cap cells (Supplementary Fig. 180 9 S2, 35-47 h). Confocal imaging of plants expressing both tonoplast and nuclear markers 181 (VHP1-mGFP and pRPS5a:H2B-tdTomato) (Adachi et al., 2011; Segami et al., 2014) 182 revealed that both nuclei and amyloplasts were embedded in the meshwork of vacuolar 183 membranes in the outermost cell layer, whereas, in the inner cell layer, amyloplasts were 184 localized in a space devoid of vacuolar membranes (Fig. 2C). Taken together, our time- 185 lapse microscopic imaging revealed a highly organized sequence of organelle 186 rearrangement in the outer root cap cells, as well as its close association with cell position 187 and cell detachment. 188 189 Autophagy is activated in the outermost root cap cells before their detachment 190 Autophagy is an evolutionarily conserved self-digestion system in eukaryotes and 191 operates by transporting cytosolic components and organelles to the vacuole for nutrient 192 recycling and homeostatic control (Mizushima and Komatsu, 2011). The rapid 193 disappearance of amyloplasts and the formation of large vacuoles observed in the 194 outermost root cap cells made us hypothesize that autophagy operates behind their 195 dynamic subcellular rearrangements before the cell detachment. To test this hypothesis, 196 we examined whether autophagosomes, spherical membrane structures characteristics of 197 autophagy, are formed in the root cap cells at the time and space corresponding to the 198 organelle rearrangement. 199 We first observed an autophagosome marker, 35Spro:GFP-ATG8a, which 200 ubiquitously expresses GFP-tagged Arabidopsis ATG8a proteins, one of the nine ATG8 201 proteins encoded in the Arabidopsis genome (Yoshimoto et al., 2004). ATG8 is a 202 10 ubiquitin-like protein, and upon autophagy activation, incorporated into the 203 autophagosome membranes as a conjugate with phosphatidylethanolamine (Liu and 204 Bassham, 2012). Our time-lapse confocal imaging revealed uniform localization of GFP- 205 ATG8a fluorescence in the inner cell layers, suggesting low autophagic activity in these 206 cells (Fig. 3B and Supplementary Movie S4). In contrast, in detaching outermost cells, 207 dot-like signals of GFP-ATG8a became evident and their number and size increased (Fig. 208 3C, -24.0-1.5 h). In later stages, GFP-ATG8a signals largely disappeared in the outermost 209 cells before their detachment (Fig. 3C, 10 h). After the detachment of the outermost cell 210 layer, the inner cells (the new outermost cells) remained showing uniform GFP-ATG8 211 signals (Fig. 3C, 18.5 h). In the later phase of cell detachment, GFP-ATG8a signals 212 exhibited ring-like shapes, a typical image of autophagosomes in confocal microscopy 213 (Fig. 3C, 1.5 h, red arrowhead and a magnified image in the inset). 214 To further confirm whether the GFP-ATG8a-labelled puncta correspond to the 215 typical double membrane-bound autophagosome, we performed correlative light and 216 electron microscopy (CLEM) analysis (Fig. 4) (Wang and Kang, 2020). GFP 217 fluorescence precisely colocalized with spherical structures typical of autophagosomes 218 (Fig. 4C-4F). Together, our observations confirmed that autophagy is activated in the 219 outermost columella cells before their detachment. 220 221 Autophagy promotes organelle rearrangement in the outermost root cap cells 222 To examine whether autophagy plays a role in the maturation of columella cells, we first 223 tested the effect of E-64d, a membrane-permeable protease inhibitor that promotes the 224 11 accumulation of autophagic bodies inside the vacuole (Inoue et al., 2006; Merkulova et 225 al., 2014). In the outermost columella cells of E64d-treated roots, autophagic body-like 226 aggregates accumulated inside the enlarged vacuoles, suggesting the occurrence of active 227 autophagic degradation in these cells (Fig. S3B, compare with S3A). 228 We next carried out the phenotypic characterization of autophagy-deficient 229 mutants. ATG genes encoding autophagy components are known to exist in the genomes 230 of Arabidopsis and other model plant species (Hanaoka et al., 2002; Liu and Bassham, 231 2012). Among them, ATG5 belongs to the core ATG genes and is essential for 232 autophagosome formation as ATG8. In the loss of function atg5-1 mutant (Yoshimoto et 233 al., 2009), GFP-ATG8a signal was uniformly distributed throughout the cytosol both 234 during and after the cell detachment, indicating that autophagosome formation in the 235 detaching columella cells requires functional ATG5 (Fig. S4 and Supplementary movie 236 S5). Furthermore, time-lapse observation revealed a loss of full vacuolation in the 237 detaching outermost cells of atg5-1 (Fig. S5A, Supplementary movie S6). In the 238 detaching outermost cells of wild-type plants, a central vacuole enlarged to occupy the 239 entire cell volume, whereas only a few spherical and small fragmented vacuoles were 240 found in the corresponding cells of atg5-1 (Fig. 5A-5D). Whereas the disappearance of 241 iodine-stained large amyloplasts was not affected in the outer columella cells of atg5-1 242 (Fig. S3C and S3D), plastids in the atg5-1 mutant exhibited abnormal morphologies 243 dominated by tubular structures called stromules (Hanson and Hines, 2018), suggesting 244 a specific role of autophagy in plastid restructuring and/or degradation (Fig. S3E and S3F). 245 We also found that the detaching atg5-1 cells were strongly stained with FDA, a 246 12 compound that emits green fluorescence when hydrolyzed in the cytosol, as compared 247 with the restricted fluorescence in the cortical region of corresponding wild-type cells 248 (Fig. 5E and 5F). Retention of cytosol in detaching columella cells was also observed in 249 FDA-stained roots of additional atg mutants including atg2-1, atg7-2, atg10-1, atg12ab, 250 atg13ab and atg18a (Fig. 5G-5L), as well as in atg5-1 plants expressing GUS-GFP fusion 251 proteins under the outer layer-specific BRN1 promoter (Fig. S5D, compare with S5C). 252 Defects of vacuolization and cytosol digestion in atg5-1 were complemented with an 253 ATG5-GFP transgene, where GFP-tagged GFP5 proteins were expressed under the ATG5 254 promoter (Fig. 5M and 5N). Together, these observations clearly demonstrated a central 255 role of autophagy in cytosol digestion and vacuolization of detaching columella cells. 256 257 Autophagy is required for organized separation of root cap cell layer 258 In the course of time-lapse imaging of atg5-1, we noticed that the autophagy-deficient 259 mutants exhibited a distinct cell detachment behavior as compared with that of wild type. 260 While the outermost root cap cells detach as a cell layer in the wild type (Fig. 6A, white 261 arrowheads, and Supplementary Movie S7) (Kamiya et al., 2016), those of atg5-1 262 detached individually (Fig. 6B, orange arrowheads, and Supplementary Movie S8), 263 indicating that autophagy is required not only for organelle rearrangement but also for the 264 organized separation of root cap cell layers, a behavior typically observed in the root cap 265 of Arabidopsis and related species (Hamamoto et al., 2006; Hawes et al., 2002). The 266 aberrant cell detachment behavior of atg5-1 was complemented by the ATG5-GFP 267 transgene (Fig. 6C, white arrowheads, and Supplementary Movie S9), confirming the 268 13 causal relationship. To clarify whether autophagy activation in the outermost cells is 269 sufficient for organized cell separation, we established atg5-1 plants expressing GFP- 270 tagged ATG5 proteins under the BRN1 and the RCPG promoter, which drive transcription 271 in the outer two cell layers and the outermost root cap layer, respectively (Kamiya et al., 272 2016). Time-lapse imaging revealed that both of the plant lines restored the organized 273 separation of the outermost root cap cell layer (Fig. 7A and 7B, white arrowheads and 274 Supplementary movie S10 and S11). These observations, in particular, restoration of the 275 layered cell separation by the RCPG promoter-driven ATG-GFP, confirmed that 276 autophagy activation in the detaching cells at the timing of active cell wall degradation is 277 sufficient for the organized separation of the outermost root cap layer. 278 279 280 Discussion 281 282 In this study, we revealed spatiotemporal dynamics of the intracellular reorganization and 283 cell detachment in the Arabidopsis root cap, as well as a role of developmentally regulated 284 autophagy in these processes. In the outermost root cap layer, autophagy is activated in a 285 specific cell layer and at the timing closely associated with the functional transition of 286 columella cells and their detachment. This spatiotemporally regulated activation of 287 autophagy is essential not only for cell clearance and vacuolar enlargement but also for 288 the organized separation of the outermost layer of the root cap. 289 290 14 Motion-tracking time-lapse imaging revealed rapid intracellular rearrangement 291 associated with the functional transition of root cap cells 292 Cells constituting the root cap constantly turn over by balanced production and 293 detachment of cells at the innermost and the outermost cell layers, respectively. During 294 their lifetime, columella cells undergo a functional transition from being gravity-sensing 295 statocytes to secretory cells according to their position (Blancaflor et al., 1998; Maeda et 296 al., 2019; Sack and Kiss, 1989; Vicre et al., 2005). While the previous electron 297 microscopic observations revealed a profound difference in the subcellular structures 298 between the inner statocytes and the outer secretory cells of the Arabidopsis root cap 299 (Maeda et al., 2019; Poulsen et al., 2008; Sack and Kiss, 1989), detailed temporal 300 dynamics of organelles rearrangement in relation to the timing of cell displacement and 301 detachment has not been analyzed. 302 Our time-lapse observation using a motion-tracking microscope system with a 303 horizontal optical axis clearly visualized both morphological and temporal details of 304 organelle rearrangement in this transition (Fig. 8). Cells in the inner two to three layers 305 have unique arrangements of organelles, which is likely optimized for their gravity- 306 sensing function (Blancaflor et al., 1998). In these cells, starch granule-containing 307 amyloplasts and nuclei are localized at the distal (lower) and proximal (upper) end of 308 each cell, respectively, whereas small tubular vacuoles preferentially occupy the proximal 309 (upper) half of each cell (Fig. 2) (Leitz et al., 2009; Sack and Kiss, 1989). This organelle 310 arrangement changed dynamically in the outermost cell layer. The first conspicuous sign 311 of rearrangement is relocation of nuclei from the upper to the central region, which 312 15 happens even before the layer containing these columella cells starts to detach at the 313 proximal LRC region (Fig. 2). Around the time of the detachment of this cell layer, 314 amyloplasts 'float up' to the middle region of the cell (Fig. 2). Later, amyloplasts disappear 315 and vacuoles start to expand to occupy the entire cell volume by the time these cells 316 slough off from the root tip (Fig. 2 and Supplementary Fig. S2). The development of large 317 central vacuoles likely constitutes a central component of functional specialization of 318 these cells for storage (Driouich et al., 2013; Hawes et al., 2016; Vicre et al., 2005). A 319 novel role of central vacuoles for cell death promotion has been also proposed for LRC 320 cells (Fendrych et al., 2014). 321 Here, the central question is what controls the spatiotemporal activation of this 322 dramatic rearrangement of organelles in the root cap. The NAC-type transcription factors 323 BRN1 and BRN2 are expressed specifically in the outer two cell layers of the root cap 324 and required for cell detachment (Bennett et al., 2010; Kamiya et al., 2016), seemingly 325 becoming good candidates for the upstream regulators. However, the outermost root cap 326 cells of brn1 brn2 mutants, though defective in cell detachment, were found to be 327 normally vacuolated and lacking amyloplasts as those of wild type, indicating that at least 328 a part of the organelle rearrangement is regulated independently of BRN1 and BRN2 329 (Bennett et al., 2010; Kamiya et al., 2016). On the other hand, our previous study 330 suggested the existence of unknown positional cues that, together with another NAC-type 331 transcription factor SMB, promote the outer layer-specific expression of BRN1 and BRN2 332 (Kamiya et al., 2016). Future identification of factors transmitting such positional 333 16 information will provide a clue to understanding a mechanism underlying position- 334 dependent organelle rearrangement in the root cap. 335 336 Autophagy is activated in the outermost root cap cells to promote cell clearance and 337 vacuolization 338 Our time-lapse imaging revealed specific activation of autophagy in the outermost root 339 cap layer in concert with the progression of the cell separation (Fig. 3). As expected, 340 mutants defective in the canonical autophagy pathway exhibited compromised cell 341 clearance and vacuolization of detaching root cap cells (Fig. 5). Because detached root 342 cap cells are dispersed into the rhizosphere and act in plant defense through their secretory 343 capacity (Driouich et al., 2013; Hawes et al., 2016), degradation of starch-containing 344 amyloplasts and vacuolar expansion appear to be a reasonable differentiation trajectory 345 in view of energy-recycling and storage. 346 Autophagosomes are double-membrane vesicles that engulf a wide range of 347 intracellular components and transport them to vacuoles for degradation by lytic enzymes. 348 Rapid reduction of GFP-ATG8a signals and accumulation of autophagic body-like 349 structures inside the vacuoles after the application of the proteinase inhibitor E64d 350 (Supplementary Fig. S3) support occurrence of active autophagic flow and vacuolar 351 degradation in the outermost root cap layer. Such active autophagic transport may act to 352 supply membrane components and to facilitate water influx into the vacuoles by 353 increasing osmotic pressure, leading to enhanced vacuolization of the outermost root cap 354 cells. 355 17 While the autophagy-deficient atg5-1 mutant was capable of eliminating 356 Lugol-stained amyloplasts from mature columella cells as the wild type, morphology of 357 plastids in the detaching root cap cells was abnormal in atg5-1, having tubular structures 358 typical of stromules (Supplementary Fig. S3). Storomules arise from chloroplasts under 359 starvation or senescence conditions. In such stress conditions, chloroplast contents are 360 degraded via piecemeal-type organelle autophagy, in which stromules or chloroplast 361 protrusions are believed to be engulfed by an autophagosome (Ishida et al., 2008), 362 whereas damaged chloroplasts can be engulfed as a whole by an isolated membrane and 363 transported into vacuoles (Izumi et al., 2013). Stromule formation in the autophagy- 364 deficient atg5-1 mutant suggests that amyloplast degradation in the outermost root cap 365 cells proceeds in two steps; first by autophagy-independent degradation of starch granules 366 and stromule formation, followed by the piecemeal chloroplast autophagy. It should be 367 noted, however, that autophagy-dependent amyloplast degradation also occurs as a part 368 of root hydrotropic response, where some starch-containing amyloplasts are engulfed 369 directly by the autophagosome-like structures (Nakayama et al., 2012). Together, these 370 observations suggest that multiple amyloplast degradation pathways exist in the 371 Arabidopsis root cap with different contributions of autophagy. 372 While the present study clearly demonstrated the role of autophagy in the 373 organelle rearrangement in the root cap, spatiotemporal regulation of autophagy 374 activation is yet to be investigated. The root cap autophagy seems to operate via canonical 375 macro-autophagy pathway mediated by the components encoded by the ATG genes (Fig. 376 5) (Liu and Bassham, 2012) (Fig. 5). Autophagy is induced by various stress conditions, 377 18 such as nutrient starvation, as well as abiotic and biotic stresses, where SNF-related 378 kinase 1 (SnRK1) and target of rapamycin (TOR) protein kinase complexes function as 379 key regulators (Liu and Bassham, 2012; Mizushima and Komatsu, 2011). In contrast, the 380 root cap autophagy can occur in plants growing on a sterile nutrient-rich medium in our 381 experiments, suggesting that root cap autophagy is activated independently of nutrient 382 starvation and biotic stress. Instead, activation of the root cap autophagy appears to be 383 closely associated with the process of cell detachment, which in turn is known to be 384 regulated by intrinsic developmental programs (Dubreuil et al., 2018; Shi et al., 2018). 385 Again, BRN1 and BRN2 are unlikely to regulate the root cap autophagy, because cell 386 clearance and vacuolization normally occur in the outermost root cap cells of brn1 brn2 387 mutants. 388 389 Autophagy is required for the organized separation of the Arabidopsis root cap cells 390 Autophagy promotes organelle rearrangement associated with the differentiation of 391 secretory cells that subsequently slough off to disperse into the rhizosphere. Based on this, 392 we expected that the loss of autophagy would inhibit or delay cell detachment in the root 393 cap. Somewhat unexpectedly, however, autophagy-deficient atg5-1 mutants showed a 394 phenotype suggestive of enhanced cell detachment (Fig. 6). In Arabidopsis and related 395 species, the outermost root cap cells separate as a cell layer, rather than as isolated cells 396 (Driouich et al., 2010; Driouich et al., 2007; Kamiya et al., 2016). Although the 397 physiological significance of this detachment behavior has not been demonstrated so far, 398 it has been hypothetically linked with a capacity of secreting mucilage, a mixture of 399 19 polysaccharides implicated in plant defense, aluminum-chelating, and lubrication 400 (Driouich et al., 2010; Maeda et al., 2019). 401 Previous genetic studies suggested a key role of cell wall pectins in the control 402 of root cap cell detachment; when pectin-mediated cell-cell adhesion was compromised 403 by mutations in genes encoding putative pectin-synthesizing enzymes or overexpression 404 of RCPG, a root cap-specific putative pectin-hydrolyzing enzyme, root cap cells slough 405 off as isolated cells (Driouich et al., 2010; Kamiya et al., 2016). Moreover, the 406 morphology of detaching root cap cell layers was altered in the loss-of-function rcpg 407 mutant, likely due to a failure of separating cell-cell adhesion along the lateral cell edge 408 (Kamiya et al., 2016). The similarity between the altered cell detachment behaviors 409 between atg5-1 and pectin-deficient plants suggests a role of autophagy in the control of 410 cell wall integrity during the root cap cell detachment. Both transport and modification 411 of cell wall pectins require Golgi and Golgi-derived vesicles (Driouich et al., 2012; Wang 412 et al., 2017). In outer root cap cells, small vesicles accumulate for their secretory functions 413 (Driouich et al., 2013; Maeda et al., 2019; Wang et al., 2017), and a mutation disrupting 414 this secretory pathway results in the failure of root cap cell detachment (Poulsen et al., 415 2008). If autophagy is required for timely attenuation of such vesicular transport during 416 the cell detachment program, lack of autophagy should lead to prolonged secretion of cell 417 wall modifying enzymes such as RCPG, resulting in enhanced loosening of cell-cell 418 adhesion. Indeed, we could recognize broader gaps at the apoplastic junctions at the distal 419 cell-cell adhesion points in atg5-1 than those in the wild type (Supplementary movie S7 420 and S8). Future studies comparing secretory dynamics of cell wall-modifying enzymes in 421 20 various genetic backgrounds using our live-imaging system will elucidate the molecular 422 mechanism controlling the cell detachment behaviors in the root cap and the role of 423 autophagy. 424 In summary, our study revealed the role of spatiotemporally regulated 425 autophagy in cell clearance and vacuolization in root cap differentiation as well as in cell 426 detachment. While autophagy has been known to promote tracheary element 427 differentiation in Arabidopsis and anther maturation in rice, roles of autophagy in these 428 instances are linked to PCD (Escamez et al., 2016; Kurusu and Kuchitsu, 2017). 429 Considering that autophagy is required for functional transition and detachment of living 430 columella cells, our study revealed a previously undescribed role of developmentally 431 regulated autophagy in plant development. 432 433 21 Materials and Methods 434 435 Plant materials and growth conditions 436 Arabidopsis thaliana L. Heynh (Arabidopsis) accession Col-0 was used as the wild type. 437 The Arabidopsis T-DNA insertional lines, atg5-1 (SAIL_129_B07), atg7-2 (GK- 438 655B06), atg2-1 (SALK_076727), atg10-1 (SALK_084434), atg12a (SAIL_1287_A08), 439 atg12b (SALK_003192), atg13a (GABI_761_A11), atg13b (GK-510F06) and atg18a 440 (GK_651D08) have been described previously (Doelling et al., 2002; Hanaoka et al., 441 2002; Izumi et al., 2013; Thompson et al., 2005; Yoshimoto et al., 2004; Yoshimoto et 442 al., 2009). 35Spro:CT-GFP, RPS5apro:H2B-tdTomato and VHP1-mGFP has been 443 described previously (Adachi et al., 2011; Köhler et al., 1997; Segami et al., 2014). Seeds 444 were grown vertically on Arabidopsis nutrient solution supplemented with 1 % (w/v) 445 sucrose and 1 % (w/v) agar under the 16h light/8h dark condition at 23 ºC. 446 447 Generation of transgenic plants 448 For ATG5pro:ATG5:GFP, a 4.5-kb genomic fragment harboring the ATG5 449 coding region and the 5’-flanking region was amplified by PCR and cloned into 450 pAN19/GFP-NOSt vector, which contained GFP-coding sequence and the 451 Agrobacterium (Rhizobium) nopaline synthase terminator (NOS). The resulting ATG5- 452 GFP fragment was then transferred to pBIN4 to give ATG5pro:ATG5:GFP/pBIN41. 453 22 Layer-specific rescue constructs of ATG5-GFP were constructed by amplifying 454 the ATG5-GFP fragment from ATG5pro:ATG5:GFP/pBIN41, and inserting them to 455 pDONR221 by the GatewayTM technology. The ATG5-GFP fragment was then 456 transferred to pGWB501:BRN1pro and pGWB501:RCPGpro, which respectively 457 contained the BRN1 and RCPG promoter flanking the Gateway cassette in pGWB501 458 (Nakagawa et al., 2007). The cytosolic marker GUS-GFP was similarly constructed by 459 inserting a GUS-GFP fragment into pENTR D-TOPO, and then by transferring the insert 460 to pGWB501:BRN1pro to give BRN1pro:GUS-GFP. 461 For DR5v2:H2B:tdTomato, a DR5v2 promoter fragment was amplified by PCR 462 from the DRv2n3GFP construct (Liao et al., 2015), and inserted into pGWB501 by the 463 In-Fusion technique to give pGWB501:DR5v2. The H2B-tdTomato fragment in pENTR 464 was transferred to the pGWB501:DR5v2. Integrity of the cloned genes was verified by 465 DNA sequencing. Transformation of Arabidopsis plants was performed by the floral dip 466 method using Rhizobium (formerly Agrobacterium) tumefaciens, strain C58MP90. 467 468 Microscopy 469 Time-lapse imaging of the root cap was performed using two microscopic systems 470 developed in the corresponding authors' laboratory, which can automatically track the tip 471 of vertically growing roots. Technical details will be published elsewhere. Briefly, an 472 inverted microscope (ECLIPSE Ti-E and ECLIPSE Ti2-E, Nikon, Tokyo, Japan) was 473 tilted by 90 degrees to vertically orient the sample stage. The motorized stage was 474 controlled by the Nikon NIS-elements software with the “keep object in view” plugin to 475 23 automatically track the tip of growing roots. Three-day-old seedlings were transferred to 476 a chamber slide (Lab-Tek chambered coverglass, Thermofisher, Waltham, MA) and 477 covered with a block of agar medium. 478 Confocal laser scanning microscopy was carried out with a Nikon C2 confocal 479 microscope. Roots were stained with 10 µg/ml of propidium iodide (PI). Fluorescein 480 diacetate (FDA) staining was performed by soaking the roots in a solution containing 2 481 μg/ml of FDA. 482 Iodine staining was performed as described previously (Segami et al., 2018). 483 Root fixed in 4% (w/v) paraformaldehyde in PBS for 30 min under a vacuum at room 484 temperature. The fixed sample was washed twice for 1 min each in PBS and cleared with 485 ClearSee (Kurihara et al., 2015). The samples were transferred to 10% (w/v) xylitol and 486 25% (w/v) urea to remove sodium deoxycholate, and then stained in a solution containing 487 2 mM iodine (Wako), 10 % (w/v) xylitol, and 25 % (w/v) urea. 488 Correlative light and electron microscopy (CLEM) analysis was performed as 489 described previously (Wang and Kang, 2020; Wang et al., 2019). GFP-ATG8a seedlings 490 were grown vertically under 16 h light-8 h dark cycle at 22 °C for seven days. Root tips 491 samples expressing GFP were cryofixed with an EM ICE high-pressure freezer (Leica 492 Microsystems, Austria) and embedded in Lowicryl HM20 resin at -45ºC. TEM sections 493 of 150nm thickness were collected on copper or gold slot grids coated with formvar and 494 examined for GFP after staining the cell wall with Calcofluor White. The grids were post- 495 stained and GFP-positive cells were imaged under an H-7650 TEM (Hitachi High-Tech, 496 24 Japan) operated at 80kV. For electron tomography, tilt series were collected with a TF- 497 20 intermediate voltage TEM (Thermo Fisher Scientific, USA). Tomogram calculation 498 and three-dimensional model preparation were carried out with the 3dmod software 499 package (bio3d.colorado.edu). 500 501 Acknowledgments 502 We thank Masanori Izumi (RIKEN, Japan), Kohki Yoshimoto (Meiji University, Japan), 503 Masayoshi Maeshima (Nagoya University, Japan), Shoji Segami (NIBB, Japan), and 504 Maureen R. Hanson (Cornell University, USA) for providing plant materials, Dolf 505 Weijers (Wageningen University, Netherlands) for providing the DR5v2 construct, and 506 Masako Kanda for technical assistance. 507 508 Competing interests 509 The authors declare no competing interests. 510 511 Funding 512 This work was supported by MEXT/JSPS KAKENHI grants 20H05330 to T.G. and 513 19H05671, 19H05670 and 19H03248 to K.N., and by the Hong Kong Research Grant 514 Council (GRF14121019, 14113921, AoE/M-05/12, C4002-17G) to B.-H. 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Plant Cell 21, 2914-2927. 691 692 31 Figures legends 693 694 Fig. 1. A diagram illustrating structure and cell detachment process of Arabidopsis 695 root cap. 696 Landmark events constituting the cell separation sequence are marked by arrowheads. 697 Definition of the proximodistal polarity used in this study is shown on the left. 698 699 Fig. 2. Organelle rearrangement takes place in the outer root cap layers 700 (A) Time-lapse images visualizing the sequences of root cap cell detachment and 701 relocation of amyloplasts. Representative images before (left panel), at the beginning 702 (central panel), and around the end (right panel) of cell layer detachment are shown. Light 703 blue and dark blue arrowheads indicate sedimenting and floating amyloplasts, 704 respectively. Green arrowhead points to a highly vacuolated cell. Corresponding video is 705 available as Supplementary movie S1. 706 (B) Time-lapse images showing intracellular relocation of nuclei (red fluorescence of 707 DR5v2:H2B-tdTomato) and amyloplasts (gray particles in the bright field). Orange and 708 red arrowheads point to the nuclei localized in the proximal (upper) and the middle 709 regions of the cell, respectively. Light blue and dark blue arrowheads point to the 710 amyloplasts in the distal (bottom) and the middle regions of the cell, respectively. Purple 711 arrowheads point to the nuclei localized at the distal pole of the cells. Corresponding 712 video is available as Supplementary movie S2. 713 (C) Confocal images visualizing differential localization of organelles between the inner 714 32 and the outermost cell layers. Orange and red arrowheads point to red-fluorescent nuclei 715 in the proximal (upper) and the middle regions in the cell, respectively. Light blue and 716 dark blue arrowheads point to the amyloplasts in the distal (bottom) and the middle 717 regions in the cell, respectively. Green color indicates vacuolar membranes. 718 Time tables shown in (A) and (B) represent durations of the cell detachment process (gray 719 box). Timing of image capturing is indicated at the upper right corner of each image 720 where the origin (0 h) is set at the time when the outermost layer started detachment in 721 the proximal LRC region. Cell outlines are delineated by white dotted lines. Scale bar, 20 722 µm. 723 724 Fig. 3. Autophagosomes are formed specifically in the outermost root cap layer 725 Representative confocal time-lapse images of the 35Spro:GFP-ATG8a root. Bright-field 726 (A) and GFP-ATG8a fluorescence (B, C) images are shown. Images in (C) are magnified 727 images of the boxed regions in (B). White arrowheads in (C) indicate autophagosomes 728 marked by GFP-ATG8a. They showed the typical donut-shaped autophagosome images 729 in the later phase of detachment (red arrowhead at 1.5h, inset: enlarged view). Timing of 730 image capturing is indicated at the upper right corner of each image where the origin (0 731 h) is set at the time when the outermost layer started detachment in the proximal LRC 732 region. Scale bar, 50 µm (A, B), 20 µm (C), 2 µm (C, inset). A corresponding video is 733 available as Supplementary movie S4. 734 735 Fig. 4. CLEM imaging revealed localization of GFP-ATG8a in autophagosomes 736 33 (A, B) GFP fluorescence (A) and TEM (B) images of a section from a GFP-ATG8a root 737 cap. 738 (C-E) Magnification of the region boxed in (A) and (B). GFP-ATG8a (C), TEM (D), and 739 their merged image (E) are shown. Red arrowhead in (E) indicates an autophagosome 740 with GFP-ATG8a fluorescence. 741 (F) A 3D electron tomographic model built for an amyloplast (blue), two mitochondria 742 (brown,) and an autophagic compartment (magenta) overlaid with the TEM image. 743 Scale bar, 10 µm (A, B); 500 nm (C-F). 744 745 Fig. 5. Vacuolization and cytosol digestion were inhibited in detaching columella 746 cells in atg mutants 747 (A-D) Vacuolar morphologies in wild-type (A, B) and atg5-1 (C, D) columella cells. (A, 748 C) VHP1-mGFP fluorescence (green). (B, D) Merged images with PI-stained cell walls 749 (red). 750 (E-L) Retention of cytosol in the detaching root cap cells of various atg mutants (F-L) as 751 compared with wild type (E). Cytosol and cell walls were stained with FDA (green) and 752 PI (red), respectively. 753 (M, N) Vacuolization and cytosol digestion defects of detaching atg5-1 root cap cells 754 were complemented by the ATG5-GFP transgene (white arrowheads). Note the uniform 755 ATG5:GFP expression by the ATG5 promoter. 756 Scale bar, 10 µm (A-D); 50 µm (E-N). 757 758 34 Fig. 6. Autophagy activation is required for organized separation of the outermost 759 root cap cell layer 760 (A-C) Time-lapse images of root cap detachment processes in wild-type (A), atg5-1 (B), 761 and ATG5pro:ATG5:GFP atg5-1 (C) plants at the time points indicated at the top. Note 762 that the outermost root cap cells detach as a layer (white arrowheads) in wild type (A) 763 and ATG5:GFP atg5-1 (C), whereas they detach individually in atg5-1 (B, orange 764 arrowheads). Scale bar, 50 µm. Corresponding videos are available as Supplementary 765 movie S7-S9. 766 767 Fig. 7. Autophagy activation at the timing of cell wall degradation is sufficient for 768 organized cell separation 769 (A-D) Time-lapse images of root cap detachment processes in BRN1pro:ATG5-GFP 770 atg5-1 (A, B) and RCPGpro:ATG5:GFP atg5-1 (C, D) at the time points indicated at the 771 top right corner of each panel. Note that the outermost root cap cells detach as a cell layer 772 in both genotypes (white arrowheads), as compared with individual detachment in atg5- 773 1 (Fig. 6B). Bright-field (A, C) and GFP fluorescence (B, D) images were shown. Scale 774 bar, 50 µm. Corresponding videos are available as Supplementary movies S10 and S11. 775 776 Fig. 8. Schematic illustration of the sequence of organelle rearrangement and 777 autophagy activation during maturation and detachment of columella cells. 778 779 Fig. S1. Arabidopsis root cap cells detach at fixed intervals 780 35 (A-D) Time-lapse images showing periodic detachment of Arabidopsis root cap cells. 781 Detachment of the outermost root cap layer initiates at the proximal LRC region and 782 progressively extends toward the central columella region (B, black arrowheads). 783 Detached root cap cells adhere together to keep a cell layer morphology (C, red 784 arrowhead). Detachment of the next cell layer initiates in the same manner as the previous 785 one (D). Elapsed time after the start of culture is indicated in each panel. Scale bar, 100 786 µm. 787 (E) A time table showing periodic detachment of root cap cell layers in five (#1-5) root 788 samples each experiencing three rounds of root cap detachment. Gray, blue, and orange 789 boxes indicate the duration from the start (initial detachment at the proximal LRC region) 790 and the end (complete detachment at the columella region) of the first, second, and third 791 cell layer, respectively. The x-axis indicates elapsed time (h) from the start of culture. 792 Red lines indicate average time points of the start of detachment. 793 (F) Intervals between the start of detachment between the first and second cell layers 794 (gray bar), and between the second and third cell layer (black bar). Mean and SE are 795 shown (n = 5). 796 797 Fig. S2. Morphological transition of vacuoles during the detachment of root cap cells 798 (A, B) Time-lapse images showing vacuolar morphology by the tonoplast-localized 799 VHP1-mGFP fluorescence (A) and bright-field images (B). In the outermost cells, 800 vacuoles are initially small and fragmented (up to 17 h), and gradually expand to form 801 large central vacuoles before the cell detachment (41 h). Elapsed time after the start of 802 36 observation is indicated in each panel. A corresponding video is available as 803 Supplementary movie S3. 804 (C-E) The entire cell volume was occupied by a large central vacuole in detaching root 805 cap cells. Images of VHP1-mGFP fluorescence (C) and its overlay with a DIC image (D) 806 were shown. (F) is a Z-stack projection encompassing 50-µm depth. Note that cells at the 807 center of the detached cell layer possess large central vacuoles as visualized by VHP1- 808 mGFP (white arrowheads), whereas those at the periphery do not show fluorescence 809 (orange arrowheads) likely due to the loss of cell viability. 810 Scale bar, 20 µm. 811 812 Fig. S3. Accumulation of autophagic body-like structures in the E64d-treated wild- 813 type root cap cells and abnormal plastid morphology in atg5-1 814 (A, B) Accumulation of autophagic body-like structures inside the vacuoles of the wild- 815 type outermost root cap cells after E-64d treatment (B, orange arrowheads), as compared 816 with the translucence vacuolar images of a non-treated control (A, white arrowheads). 5- 817 day-old seedlings grown on the medium with or without 10 µM E-64d were observed. 818 Scale bar, 20 µm. 819 (C, D) Amyloplasts in the outermost root cap cells lost starch granules in both wild type 820 and atg5-1. Black arrowheads indicate the detaching outermost cell layers. Scale bar, 50 821 µm. 822 (E, F) Amyloplasts exhibit abnormal morphologies in the outermost root cap cells of 823 atg5-1 (F) as compared with those in the wild type (E). Plastids are visualized by the CT- 824 37 GFP fluorescence marker line. Note that small spherical plastids accumulate in the wild- 825 type cells (white arrowheads), whereas those with tubular morphologies dominate in 826 atg5-1 cells (orange arrowheads). Scale bar, 20 µm. 827 828 Fig. S4. Autophagosomes do not form in the detaching root cap cells of atg5-1 829 Time-lapse images of the 35Spro:GFP-ATG8a atg5-1 root tip. Bright-field (A) and GFP- 830 ATG8a fluorescence images (B, C) are shown. Images in (C) are magnified views of 831 boxed regions in (B) of respective time points. Note that the GFP-ATG8a signals were 832 uniformly distributed throughout the cytosol. Occasionally observed punctate signals did 833 not form a donut-shape typical of an autophagosome (D, E). Elapsed time after the start 834 of observation is indicated at the top. Scale bar, 50 µm (A, B); 20 µm (C); 10 µm (D, E). 835 A corresponding video is available as Supplementary movie S5. 836 837 Fig. S5. Vacuolization and cytosol digestion do not occur in detaching atg5-1 cells 838 (A, B) Time-lapse images showing vacuolar morphology by the tonoplast-localized 839 VHP1-mGFP fluorescence (A), and corresponding bright-field images (B) in atg5-1. In 840 the outermost cells, vacuoles are initially small and fragmented and gradually expand as 841 those in wild type, but fail to expand fully (43 h). Elapsed time after the start of 842 observation is indicated at the upper right corner of each panel. Corresponding video is 843 available as Supplementary movie S6. 844 (C, D) Cytosolic GUS-GFP proteins expressed under the outer layer-specific BRN1 845 promoter revealed cytosol digestion in the detaching root cap cells of wild type, as 846 38 compared with its retention in atg5-1 (white arrowheads). 847 Scale bar, 20 µm (A, B); 50 µm (C, D). 848 849 Supplementary Movie S1. Time-lapse movie showing root cap cell detachment and 850 organelle rearrangement in wild-type root cap cells 851 Scale bar, 20 µm. 852 853 Supplementary Movie S2. Time-lapse movie showing intracellular relocation of 854 nuclei (red, DR5v2:H2B-tdTomato) and amyloplasts (gray particles in the bright 855 field) in the root cap cells 856 Scale bar, 20 µm. 857 858 Supplementary Movie S3. Time-lapse movie showing morphological transition of 859 vacuoles during cell detachment 860 Scale bar, 20 µm. 861 862 Supplementary Movie S4. Time-lapse movie showing autophagosome formation in 863 the outermost root cap cells visualized by 35Spro:GFP-ATG8a 864 Scale bar, 20 µm. 865 866 Supplementary Movie S5. Time-lapse movie showing the absence of autophagosome 867 formation in 35Spro:GFP-ATG8a in atg5-1. 868 39 Scale bar, 20 µm. 869 870 Supplementary Movie S6. Time-lapse movie showing morphological transition of 871 vacuoles during cell detachment in atg5-1. 872 Scale bar, 20 µm. 873 874 Supplementary Movie S7. Time-lapse movie showing root cap cell detachment in the 875 wild type 876 Scale bar, 50 µm. 877 878 Supplementary Movie S8. Time-lapse movie showing root cap cell detachment in 879 atg5-1 880 Scale bar, 50 µm. 881 882 Supplementary Movie S9. Time-lapse movie showing root cap cell detachment in 883 atg5-1 complemented with ATG5pro:ATG-GFP 884 Scale bar, 50 µm. 885 886 Supplementary Movie S10. Time-lapse movie showing root cap cell detachment in 887 atg5-1 complemented with BRN1pro:ATG-GFP 888 Scale bar, 50 µm. 889 890 40 Supplementary Movie S11. Time-lapse movie showing root cap cell detachment in 891 atg5-1 complemented with RCPG1pro:ATG5-GFP 892 Scale bar, 50 µm. 893 894 41 Separation of living cells by cell-wall degradation Cleavage of LRC layer Division of initial cells Proximal Distal Columella Lateral root cap (LRC) Programmed cell death (PCD) of proximal LRC cells Fig. 1. A diagram illustrating structure and cell detachment process of Arabidopsis root cap. Landmark events constituting the cell separation sequence are marked by arrowheads. Definition of the proximodistal polarity used in this study is shown on the left. 42 DR5v2:H2B-tdTomato (nucleus) B pRPS5a:H2B:tdTomato pRPS5a:H2 (nucleus) Merge VHP1 P - P1 mGFP VHPP1- GFP mG m (vacuole) C 0 5 10 -5 -10 -15 -20 -5 10 20 -10 0 5 15 25 30 -4 h 0.5 h 18 h end (h) (h) A start -12 h -8 h 0 h 13 h 5 0 start Fig. 2. Organelle rearrangement takes place in the outer root cap layers 43 Fig. 2. Organelle rearrangement takes place in the outer root cap layers (A) Time-lapse images visualizing the sequences of root cap cell detachment and relocation of amyloplasts. Representative images before (left panel), at the beginning (central panel), and around the end (right panel) of cell layer detachment are shown. Light blue and dark blue arrowheads indicate sedimenting and floating amyloplasts, respectively. Green arrowhead points to a highly vacuolated cell. Corresponding video is available as Supplementary movie S1. (B) Time-lapse images showing intracellular relocation of nuclei (red fluorescence of DR5v2:H2B-tdTomato) and amyloplasts (gray particles in the bright field). Orange and red arrowheads point to the nuclei localized in the proximal (upper) and the middle regions of the cell, respectively. Light blue and dark blue arrowheads point to the amyloplasts in the distal (bottom) and the middle regions of the cell, respectively. Purple arrowheads point to the nuclei localized at the distal pole of the cells. Corresponding video is available as Supplementary movie S2. (C) Confocal images visualizing differential localization of organelles between the inner and the outermost cell layers. Orange and red arrowheads point to red-fluorescent nuclei in the proximal (upper) and the middle regions in the cell, respectively. Light blue and dark blue arrowheads point to the amyloplasts in the distal (bottom) and the middle regions in the cell, respectively. Green color indicates vacuolar membranes. Time tables shown in (A) and (B) represent durations of the cell detachment process (gray box). Timing of image capturing is indicated at the upper right corner of each image where the origin (0 h) is set at the time when the outermost layer started detachment in the proximal LRC region. Cell outlines are delineated by white dotted lines. Scale bar, 20 µm. 44 Bright field GFP-ATG8 A B C -24.0 h -15.5 h -7.0 h 1.5 h 10.0 h 18.5 h Fig. 3. Autophagosomes are formed specifically in the outermost root cap layer Representative confocal time-lapse images of the 35Spro:GFP-ATG8a root. Bright-field (A) and GFP-ATG8a fluorescence (B, C) images are shown. Images in (C) are magnified images of the boxed regions in (B). White arrowheads in (C) indicate autophagosomes marked by GFP-ATG8a. They showed the typical donut-shaped autophagosome images in the later phase of detachment (red arrowhead at 1.5h, inset: enlarged view). Timing of image capturing is indicated at the upper right corner of each image where the origin (0 h) is set at the time when the outermost layer started detachment in the proximal LRC region. Scale bar, 50 µm (A, B), 20 µm (C), 2 µm (C, inset). A corresponding video is available as Supplementary movie S4. 45 A B C D E F GFP-ATG8a GFP-ATG8a TEM TEM GFP-ATG8a + TEM Fig. 4. CLEM imaging revealed localization of GFP-ATG8a in autophagosomes (A, B) GFP fluorescence (A) and TEM (B) images of a section from a GFP-ATG8a root cap. (C-E) Magnification of the region boxed in (A) and (B). GFP-ATG8a (C), TEM (D), and their merged image (E) are shown. Red arrowhead in (E) indicates an autophagosome with GFP-ATG8a fluorescence. (F) A 3D electron tomographic model built for an amyloplast (blue), two mitochondria (brown,) and an autophagic compartment (magenta) overlaid with the TEM image. Scale bar, 10 µm (A, B); 500 nm (C-F). 46 VHP1-mGFP VHP1-mGFP + PI WT atg5-1 FDA (green) / PI (red) A B C D E F G H I J K L WT atg5-1 atg2-1 atg7-2 atg10-1 atg12ab atg13ab atg18a ATG5 G - 5-GFP ATGG5- FP GF G Cell wall M N ATG5pro:ATG5:GFP (atg5-1) Fig. 5. Vacuolization and cytosol digestion were inhibited in detaching columella cells in atg mutants (A-D) Vacuolar morphologies in wild-type (A, B) and atg5-1 (C, D) columella cells. (A, C) VHP1-mGFP fluorescence (green). (B, D) Merged images with PI-stained cell walls (red). (E-L) Retention of cytosol in the detaching root cap cells of various atg mutants (F-L) as compared with wild type (E). Cytosol and cell walls were stained with FDA (green) and PI (red), respectively. (M, N) Vacuolization and cytosol digestion defects of detaching atg5-1 root cap cells were complemented by the ATG5-GFP transgene (white arrowheads). Note the uniform ATG5:GFP expression by the ATG5 promoter. Scale bar, 10 µm (A-D); 50 µm (E-N). 47 WT atg5-1 atg5-1 with ATG5pro:ATG5:GFP A B C Fig. 6. Autophagy activation is required for organized separation of the outermost root cap cell layer (A-C) Time-lapse images of root cap detachment processes in wild-type (A), atg5-1 (B), and ATG5pro:ATG5:GFP atg5-1 (C) plants at the time points indicated at the top. Note that the outermost root cap cells detach as a layer (white arrowheads) in wild type (A) and ATG5:GFP atg5-1 (C), whereas they detach individually in atg5-1 (B, orange arrowheads). Scale bar, 50 µm. Corresponding videos are available as Supplementary movie S7-S9. 48 RCPGpro:ATG5-GFP (atg5-1) BRN1pro:ATG5-GFP (atg5-1) 0.0 h 7.0 h 19.5 h 20.5 h 30.0 h 20.5 h 34.0 h 43.5 h 48.0 h 50.0 h A B C D Fig. 7. Autophagy activation at the timing of cell wall degradation is sufficient for organized cell separation (A-D) Time-lapse images of root cap detachment processes in BRN1pro:ATG5-GFP atg5-1 (A, B) and RCPGpro:ATG5:GFP atg5-1 (C, D) at the time points indicated at the top right corner of each panel. Note that the outermost root cap cells detach as a cell layer in both genotypes (white arrowheads), as compared with individual detachment in atg5-1 (Fig. 6B). Bright-field (A, C) and GFP fluorescence (B, D) images were shown. Scale bar, 50 µm. Corresponding videos are available as Supplementary movies S10 and S11. 49 autophagosomes vacuolization cytosol digestion nuclei translocation cell wall degradation detachment second outermost layer outermost layer : Nucleus : Amyloplast with starch granules (statolith) : Shrinking amyloplast : Vacuole autophagy amyloplast floating-up Fig. 8. Schematic illustration of the sequence of organelle rearrangement and autophagy activation during maturation and detachment of columella cells. 50 1st to 2nd 2nd to 3rd 0 10 20 30 40 Interval (h) 37.3 h (±2.3) 39.3 h (±4.4) Start of observation End of observation 1st detachment 2nd detachment 3rd detachment 0 50 100 150 200 250 Time of culture (h) #5 #4 #3 #2 #1 79.2 h (±4.8) 116.5 h (±3.5) 155.8 h (±6.7) 3rd detachment A B C D E F 2nd detachment Fig. S1. Arabidopsis root cap cells detach at fixed intervals (A-D) Time-lapse images showing periodic detachment of Arabidopsis root cap cells. Detachment of the outermost root cap layer initiates at the proximal LRC region and progressively extends toward the central columella region (B, black arrowheads). Detached root cap cells adhere together to keep a cell layer morphology (C, red arrowhead). Detachment of the next cell layer initiates in the same manner as the previous one (D). Elapsed time after the start of culture is indicated in each panel. Scale bar, 100 µm. (E) A time table showing periodic detachment of root cap cell layers in five (#1-5) root samples each experiencing three rounds of root cap detachment. Gray, blue, and orange boxes indicate the duration from the start (initial detachment at the proximal LRC region) and the end (complete detachment at the columella region) of the first, second, and third cell layer, respectively. The x-axis indicates elapsed time (h) from the start of culture. Red lines indicate average time points of the start of detachment. (F) Intervals between the start of detachment between the first and second cell layers (gray bar), and between the second and third cell layer (black bar). Mean and SE are shown (n = 5). 51 VHP1-mGFP VHP1-mGFP + DIC VHP1-mGFP (Z-projection) C D E VHP1-mGFP Bright field 5 h 11 h 17 h 23 h 29 h 35 h 41 h 47 h A B 5 h 11 h 17 h 23 h 29 h 35 h 41 h 47 h Fig. S2. Morphological transition of vacuoles during the detachment of root cap cells (A, B) Time-lapse images showing vacuolar morphology by the tonoplast-localized VHP1-mGFP fluorescence (A) and bright-field images (B). In the outermost cells, vacuoles are initially small and fragmented (up to 17 h), and gradually expand to form large central vacuoles before the cell detachment (41 h). Elapsed time after the start of observation is indicated in each panel. A corresponding video is available as Supplementary movie S3. (C-E) The entire cell volume was occupied by a large central vacuole in detaching root cap cells. Images of VHP1-mGFP fluorescence (C) and its overlay with a DIC image (D) were shown. (F) is a Z-stack projection encompassing 50-µm depth. Note that cells at the center of the detached cell layer possess large central vacuoles as visualized by VHP1-mGFP (white arrowheads), whereas those at the periphery do not show fluorescence (orange arrowheads) likely due to the loss of cell viability. Scale bar, 20 µm. 52 Control E-64d (proteinase inhibitor) E WT 35Spro:CT-GFP (plastid) Z-projection Starch granule (iodine staining) A B E F F WT atg5-1 C D Fig. S3. Accumulation of autophagic body-like structures in the E64d-treated wild-type root cap cells and abnormal plastid morphology in atg5-1 (A, B) Accumulation of autophagic body-like structures inside the vacuoles of the wild-type outermost root cap cells after E-64d treatment (B, orange arrowheads), as compared with the translucence vacuolar images of a non-treated control (A, white arrowheads). 5-day-old seedlings grown on the medium with or without 10 µM E-64d were observed. Scale bar, 20 µm. (C, D) Amyloplasts in the outermost root cap cells lost starch granules in both wild type and atg5-1. Black arrowheads indicate the detaching outermost cell layers. Scale bar, 50 µm. (E, F) Amyloplasts exhibit abnormal morphologies in the outermost root cap cells of atg5-1 (F) as compared with those in the wild type (E). Plastids are visualized by the CT-GFP fluorescence marker line. Note that small spherical plastids accumulate in the wild-type cells (white arrowheads), whereas those with tubular morphologies dominate in atg5-1 cells (orange arrowheads). Scale bar, 20 µm. 53 0 h 9 h 18 h 27 h 36 h 45 h Bright field GFP-ATG8 A B C D E 2 h 19.5 h Fig. S4. Autophagosomes do not form in the detaching root cap cells of atg5-1 Time-lapse images of the 35Spro:GFP-ATG8a atg5-1 root tip. Bright-field (A) and GFP-ATG8a fluorescence images (B, C) are shown. Images in (C) are magnified views of boxed regions in (B) of respective time points. Note that the GFP-ATG8a signals were uniformly distributed throughout the cytosol. Occasionally observed punctate signals did not form a donut-shape typical of an autophagosome (D, E). Elapsed time after the start of observation is indicated at the top. Scale bar, 50 µm (A, B); 20 µm (C); 10 µm (D, E). A corresponding video is available as Supplementary movie S5. 54 15 h 19 h 23 h 27 h 31 h 35 h 39 h 43 h 15 h 19 h 23 h 27 h 31 h 35 h 39 h 43 h Bright field VHP1-mGFP atg5-1 A B C D BRN1pro:GUS-GFP WT atg5-1 Fig. S5. Vacuolization and cytosol digestion do not occur in detaching atg5-1 cells (A, B) Time-lapse images showing vacuolar morphology by the tonoplast- localized VHP1-mGFP fluorescence (A), and corresponding bright-field images (B) in atg5-1. In the outermost cells, vacuoles are initially small and fragmented and gradually expand as those in wild type, but fail to expand fully (43 h). Elapsed time after the start of observation is indicated at the upper right corner of each panel. Corresponding video is available as Supplementary movie S6. (C, D) Cytosolic GUS-GFP proteins expressed under the outer layer-specific BRN1 promoter revealed cytosol digestion in the detaching root cap cells of wild type, as compared with its retention in atg5-1 (white arrowheads). Scale bar, 20 µm (A, B); 50 µm (C, D). 55 Supplementary Movie S1. Time-lapse movie showing root cap cell detachment and organelle rearrangement in wild-type root cap cells Scale bar, 20 µm. 56 Supplementary Movie S2. Time-lapse movie showing intracellular relocation of nuclei (red, DR5v2:H2B-tdTomato) and amyloplasts (gray particles in the bright field) in the root cap cells Scale bar, 20 µm. 57 Supplementary Movie S3. Time-lapse movie showing morphological transition of vacuoles during cell detachment Scale bar, 20 µm. 58 Supplementary Movie S4. Time-lapse movie showing autophagosome formation in the outermost root cap cells visualized by 35Spro:GFP-ATG8a Scale bar, 20 µm. 59 Supplementary Movie S5. Time-lapse movie showing the absence of autophagosome formation in 35Spro:GFP-ATG8a in atg5-1. Scale bar, 20 µm. 60 Supplementary Movie S6. Time-lapse movie showing morphological transition of vacuoles during cell detachment in atg5-1. Scale bar, 20 µm. 61 Supplementary Movie S7. Time-lapse movie showing root cap cell detachment in the wild type Scale bar, 50 µm. 62 Supplementary Movie S8. Time-lapse movie showing root cap cell detachment in atg5-1 Scale bar, 50 µm. 63 Supplementary Movie S9. Time-lapse movie showing root cap cell detachment in atg5-1 complemented with ATG5pro:ATG-GFP Scale bar, 50 µm. 64 Supplementary Movie S10. Time-lapse movie showing root cap cell detachment in atg5-1 complemented with BRN1pro:ATG-GFP Scale bar, 50 µm. 65 Supplementary Movie S11. Time-lapse movie showing root cap cell detachment in atg5-1 complemented with RCPG1pro:ATG5-GFP Scale bar, 50 µm. 66

2022

Autophagy promotes organelle clearance and organized cell separation of living root cap cells in

10.1101/2022.02.16.480624

creative-commons

1 Profiles of secoiridoids and alkaloids in tissue of susceptible and resistant green ash progeny reveal patterns of induced responses to emerald ash borer in Fraxinus pennsylvanica Robert K. Stanley1, David W. Carey2, Mary E. Mason2, Therese M. Poland3, Jennifer L. Koch2, A. Daniel Jones4, Jeanne Romero-Severson1* 1Department of Biological Sciences, University of Notre Dame, Notre Dame, IN 46556, USA e-mail: [email protected] e-mail: [email protected] 2Northern Research Station, Forest Service, U.S. Department of Agriculture, Delaware, OH 43015, USA e-mail: [email protected] e-mail: [email protected] e-mail: [email protected] 3Northern Research Station, Forest Service, U.S. Department of Agriculture, Lansing, MI 48910, USA e-mail: [email protected] 4Department of Biochemistry and Molecular Biology, Michigan State University, East Lansing, MI 49503, USA 2 e-mail: [email protected] *Corresponding author Classification: Plant Biology, Chemistry Keywords: Emerald Ash Borer, Fraxinus pennsylvanica, Invasive Species, Plant Defenses, Untargeted Metabolomics, Preprint Server: Biorxiv This PDF file includes Main Text Figures 1 to 5 Table 1 3 Abstract The emerald ash borer (Agrilus planipennis, EAB) invasion in North America threatens most North American Fraxinus species, including green ash (F. pennsylvanica), the mostly widely distributed species (1, 2). A small number of green ash (“lingering ash”, 0.1-1%) survive years of heavy EAB attack (3) and kill more EAB larvae when challenged in greenhouse studies than susceptible controls (4). We combined untargeted metabolomics with intensive phenotyping of segregating F1 progeny from susceptible or lingering ash parents to detect chemotypes associated with defensive responses to EAB. We examined three contrasting groups: low larval kill (0-25% of larvae killed), high larval kill (55-95% of larvae killed) and uninfested. Contrasting the chemotypes of these groups revealed evidence of an induced response to EAB. Infested trees deployed significantly higher levels of select secoiridoids than uninfested trees. Within the infested group, the low larval kill (LLK) individuals deployed significantly higher levels of select secoiridoids than the high larval kill (HLK) individuals. The HLK individuals deployed significantly higher concentrations of three metabolites annotated as aromatic alkaloids compared to the LLK and uninfested individuals. We propose a two-part model for the North American Fraxinus response to EAB wherein every individual has the capacity to detect and respond to EAB, but only certain trees mount an effective defense, killing enough EAB larvae to prevent or minimize lethal damage to the vascular system. Integration of intensive phenotyping of structured populations with metabolomics reveals the multi-faceted nature of the defenses deployed in naïve host populations against invasive species. Significance 4 Long-lived forest trees employ evolutionarily conserved templates to synthesize an array of defensive metabolites. The regulation of these metabolites, honed against native pests and pathogens, may be ineffective against novel species, as illustrated by the high mortality (>99%) in green ash infested by the invasive emerald ash borer (EAB). However, high standing genetic variation may produce a few individuals capable of an effective defense, as seen in the rare surviving green ash. In an investigation of this plant-insect interaction, we annotated metabolites associated with generalized but ineffective responses to EAB, and others associated with successful defensive responses. Untargeted metabolomics combined with intensive phenotyping of structured populations provides a framework for understanding resistance to invasive species in naïve host populations. INTRODUCTION Invasive pests and pathogens, now widely dispersed through globalization, threaten nearly two thirds of North American forests (1). Exacerbated by climate change, these increasingly severe infections and infestations destabilize forested ecosystems and inflict billions of dollars in direct costs to individuals and local communities (5-14). Two of these pathogens and pests, chestnut blight (Cryphonectria parasitica) and emerald ash borer (Agrilus planipennis, EAB) have had a profound impact on public awareness: chestnut blight because this disease caused the ecological extinction of the iconic American chestnut (Castanea dentata) and EAB because of the widespread, rapid and continuing loss of ash trees from streets, parks, and forests (14). The severe impact of the loss of chestnut, pales in comparison 5 to the economic and ecological damage already inflicted by EAB, the most destructive and economically devastating invasive insect pest of forest trees in North American history (15). EAB, a beetle native to Asia, was discovered in Michigan, United States and in Ontario, Canada in 2002 (16). EAB attacks ash (Fraxinus) species; larvae hatch from eggs laid in bark furrows and burrow into the living tissue directly beneath the bark (2). The larvae feed on the vascular cambium, cork cambium, phloem, and xylem inflicting severe vascular damage that ultimately kills the tree. Larvae feed during the summer and the fall and may take one or two years to complete development. The larvae pass through four developmental instars and then chew a pupal chamber in the outer sapwood or inner bark in which they overwinter as mature larvae folded over in a J-shape. During the spring, they enter the pupal stage and transform into adults that emerge in late spring and early summer through characteristic D-shaped exit holes (17, 18). EAB infestation has resulted in the rapid loss of hundreds of millions of green ash, not only in forests and rural areas but also in cities, where green ash was once one of the most widely planted street and park trees in the United States (19, 20). Green ash, the most widely distributed Fraxinus species in North America, is a dioecious, diploid, and deciduous tree species native to the eastern and central United States and eastern Canada (16, 18, 20). Mortality in green ash from EAB infestation can approach 100% within six years of the local detection of EAB (20). EAB invasion threatens not only green ash, but survival of the majority of native North American Fraxinus species including white (F. americana), pumpkin (F. profunda), Carolina (F. caroliniana) and black ash (F. nigra) (20, 21). Fraxinus are ecologically important in a wide range of forested ecosystems and are also extensively utilized for soil conservation, rural 6 water management, riparian zone stabilization, flood control, and urban green spaces in North America (19). Long term forest plot monitoring initiated in 2005, two years (3, 20, 22) after the initial detection of EAB in North America, revealed a small number of green ash (0.1-1%) that survive for years after all other surrounding green ash have died (3). These “lingering ash” (L) have been and continue to be propagated as potential sources of genetic resistance for a breeding program (23, 24). Eleven years of replicated egg bioassay tests, conducted by placing controlled densities of EAB eggs on test trees and monitoring larval development and survival, revealed reproducible larval kill capabilities with phenotypic distributions among trees that suggest quantitative inheritance (24, 25). Durable genetic resistance in the host, the most effective control measure for any pest or pathogen (26, 27), was not initially considered a strategic goal for saving North American Fraxinus species from EAB. The assumption was that a species cannot have any resistance to a pest with which it has not coevolved (28, 29). However, many studies have shown that in many cases native species do marshal heritable defensive responses to non-native invaders (30, 31). Successful breeding programs have produced American beech (Fagus grandifolia) resistant to beech bark disease (Neonectria spp transmitted by Cryptococcus fagisuga) (32), eastern white pine (Pinus strobus) resistant to white pine blister rust (Cronartium ribicola) (33) and Port Orford cedar (Chamaecyparis lawsoniana) (22) resistant to the root rot pathogen Phytophthora lateralis. The success of these and other programs demonstrates that heritable resistance exists in wild populations and can be used to develop resistant populations for species restoration through breeding (30). Once a genetic component is confirmed, a detailed study of the 7 mechanisms of the response can contribute to a body of knowledge on the omics of heritable defensive responses. Previous investigations on the role of secondary metabolites as defenses against EAB have focused on comparing small numbers of cultivars from susceptible Fraxinus spp. to the naturally resistant F. mandshurica cultivar ‘mancana’(34). Application of methyl jasmonate in infested susceptible F. americana individuals induced production of verbascoside and suppressed EAB larval development (35).These studies collectively proposed a positive association between lignan glycosides and host resistance, particularly pinoresinol and verbascoside, as well as suggesting a role for secoiridoid glycosides (36). Secoiridoids are also implicated in the response of European Ash (F. excelsior) to ash dieback disease (ADB) caused by Hymenoscyphus fraxineus. Ash dieback ultimately infects the woody stem tissue, killing the tree (37, 38). High concentrations of specific secoiridoids were identified with tolerant genotypes in one study, and with susceptible genotypes in another. Both groups of investigators proposed that the different levels of secoiridoids are the result of differential transcriptional regulation (39, 40). Investigations of ash dieback phenotypes, in these and other studies suggest that susceptibility to ADB in F. excelsior, is a quantitative trait (41). Other recent investigations of resistance to wood-boring insects have shown that some trees utilize secondary metabolite-based constitutive and induced defensive responses against specific insect pests (42-44). The concentration and profiles of certain plant secondary metabolites strongly predict resistance in maritime pine (Pinus pinaster) to the pine weevil (Hylobius abietis), after accounting for genetic relatedness among the host trees (42). Other 8 investigations have shown that the response consists of altered rates of synthesis for existing metabolites, rather than the synthesis of unique compounds (42, 45). Here we combine untargeted metabolomics and intensive phenotyping on structured populations using an experimental design that accounts for the confounding effect of genetics and environment to detect chemotypes associated with defensive responses to EAB. We hypothesized that the full sibling progeny of Susceptible x Susceptible (SxS), and Lingering x Lingering (LxL) parents would produce a wide range of larval kill phenotypes and that the family means of the progeny from LxL parents would be significantly higher than the family means of the progeny of SxS parents. If both these hypotheses are correct, and the defense is associated with secondary metabolites, we expect a contrast in chemotypes between the high larval kill (HLK, tree defenses killed 55-95% of larvae) and low larval kill (LLK, tree defenses killed 0-25% of larvae) phenotypes. If infestation induces a response, we expect that the chemotypes of infested individuals will be distinct from uninfested individuals within families. Our data showed that some secondary metabolites including select secoiridoids occurred at higher concentrations in infested individuals regardless of larval kill phenotype, while a smaller number of compounds, annotated as aromatic alkaloids were found in higher concentrations in high percent larval kill individuals. Our work will spur future investigations for the molecular basis of durable genetic resistance to EAB in green ash and provide a framework for discovering resistance to invasive species in naïve host populations. RESULTS 9 Analysis of EAB-resistance in full-sibling families of reveals that resistance to EAB in green ash is a multigenic quantitative trait Seedlings (2-3 years old) from two green ash F1 families produced through crosses between lingering parents (LxL) and one family produced by a cross between susceptible parents (SxS) were infested with EAB to confirm the genetic basis of the larval kill phenotype (Figure 1a).. One-way ANOVA and Tukey-Kramer multiple comparison tests revealed that the mean percent larval kill of (LxL) families Pe-Y and Pe-Z were significantly different from the mean percent larval kill of the (SxS) family Pe-C (p < 0.01), but there was no significant difference among the L x L families’ means (Figure. 1b). The shape and range of the larval kill distributions strongly suggests that the phenotype is a quantitative trait and provides support for the hypothesis of complex inheritance (Figure. 1b). Each family produced a range of larval kill phenotypes, (Pe-C: 0-44%, Pe-Y: 8-95%, Pe-Z: 0-75%). Based on the distribution of phenotypes across families (Figure. 1b), we classified individual trees with larval kill values of 25% or lower as LLK and those with larval kill values greater than 55% as HLK. The value of 55% is higher than the highest larval kill value for the collection of lingering ash parents described in a previous report, and the value of 25% is higher than the parents of family Pe-C, and most of the progeny (88%) in the susceptible family Pe-C (Figure 1) (24). As a comparison, the resistant Asian ash F. mandshurica typically kills 80-90% of EAB larvae when tested with the egg bioassay (24). The lingering families in this study included some progeny that performed similarly to resistant Manchurian ash individuals. Generation of untargeted metabolomic profiles. 10 We produced untargeted metabolomic profiles from acetonitrile:isopropanol:water extractions using ultra-high performance liquid chromatography/ high resolution mass spectrometry (UHPLC/MS). The levels of metabolites were normalized to a constant internal standard and replicated, with a constant mass of tissue extracted. An analysis of the relative standard deviation (RSD) of pooled controls had a median of 29.8% for all features considered in downstream analyses (Figure S1) Metabolite based OPLS-DA models correctly identify progeny classes. We conducted pairwise comparisons of the metabolite profiles of HLK, LLK, and uninfested (UNI) individuals within families to determine if metabolites were associated with infestation status or the larval kill phenotype. We assessed 194 metabolite features (Figure 2) with pairwise one-way analysis of variance (ANOVA) tests for the following contrasts: Family C: UNI vs LLK; Family Y: UNI vs LLK, UNI vs HLK, LLK vs HLK; Family Z: UNI vs LLK, UNI vs HLK, LLK vs HLK. Between 9 and 49 features were significant (p < 0.05) in each comparison (Figure 2, Table S2). We then took these features and used orthogonal partial least squares-discriminant analyses (OPLS-DAs) to examine their ability to accurately identify the correct progeny classification (Figure 2). To prevent overfitting of our model we performed a three-fold cross validation on our data and report the average prediction accuracies as the performance of our model. Overall, our model performed quite well, with over 70 % of individuals correctly assigned to progeny larval-kill phenotype class across all models, with the majority of other individuals being unclassified, not incorrectly assigned (Figure 3a, Table S2). Our confidence in these models was supported by principal component analyses (PCAs) yielding similar 11 separations(Figures 3b, 3c), suggesting that OPLS-DAs are producing statistically meaningful group separations (46). This workflow can serve as a template for assessing the relationship of chemotypes and complex phenotypes in a non-model system. Chemotypes across families distinguish a general defense response from a successful defense response. We focused our attention on the 32 features that had a significant p-value in more than one family’s comparisons or were suggested as important for EAB defense in previous investigations. This latter category included verbascoside (35) and salidroside (47). We generated electrospray ionization tandem mass spectra (ESI-MS/MS) for the features detected in our analysis and annotated them based on comparisons with MS/MS databases including the Massbank of North America, along with published literature and purchased standards. The annotation confidence is labeled according to the recommendations of the Metabolomics Standards Initiative (MSI) (48). Our annotations revealed compounds from a wide variety of chemical families, including the first record of specific alkaloids present in green ash tissue (Table 1, Figure 4, Figure 5). We annotated 12 secoiridoids with similar structures to secoiridoids previously hypothesized to be indicative of a resistance mechanism (36). We found that these secoiridoids were elevated in both low and high larval kill phenotypes compared to uninfested controls (Figure S2). However, five secoiridoids had significantly higher concentrations in low larval kill phenotypes compared to high larval kill phenotypes with no significant difference in concentration in the other seven secoiridoids (Figure 5b-d). 12 One secoiridoid (m/z 569.23) was elevated in all infested comparisons (LLK v UNI, HLK v UNI) across all families, and may have some value as an indicator of infestation across many ash genotypes. Another two secoiridoids, nueznehide (m/z 704.2781) and GL5 (m/z 928.3429), found in higher concentrations in trees that are highly susceptible to ash dieback in previous investigations (39, 49), were significantly higher in low larval kill individuals compared to uninfested individuals across families. Additionally, we found that concentrations of verbascoside, a phenylethanoid glycoside also proposed as a component of the resistance response (36), were highest in low larval kill individuals. Overall, our data suggests that these specific secoiridoids and verbascoside may be indicative of a general wound response, but do not appear to be responsible for the high larval kill phenotype. The only compounds that were higher in high larval kill individuals compared to low larval kill individuals were three compounds annotated as aromatic alkaloids and the phenylethanoid glycoside salidroside (Table 1, Figure 5, Figure S3). These alkaloids are the first reported in green ash and suggest a novel role of alkaloid in defense against herbivory in forest trees. DISCUSSION We investigated the ability of select green ash individuals to respond to EAB using structured populations, a reproducible phenotyping method, and an untargeted metabolomics approach. We found that all green ash seedlings analyzed displayed metabolic changes in response to infestation, but in most individuals, this response was ineffective to kill many EAB larvae. OPLS-DA and multivariate analyses showed that high and low performing individuals had chemotypes distinct from each other and from uninfested individuals. These chemotypes are distinguishable based on the relative concentrations of select metabolites (Table 2, Figure 5, 13 Table S1), not their presence or absence, suggesting genetic regulation of multiple synthesis pathways may be responsible for the high larval kill phenotype. We provide an initial annotation of metabolites for further study, including secoiridoids that may prove to be reliable indicators of infestation across all genetic backgrounds, and three aromatic alkaloids that may be part of an effective defensive response. Defensive responses based on multigenic mechanisms confer durable genetic resistance, the most effective control measure for any pest or pathogen. In our study, full sibling F1 progeny of lingering ash parents performed better on average than the F1 progeny of susceptible parents and produced progeny with phenotypes ranging from 0 % larvae killed to 95% larvae killed. This is the result expected when a phenotype is the result of complex genetic mechanisms involving multiple loci (50). A multigenic mechanism for the lingering ash phenotype is also consistent with two recent candidate gene studies, one on the pan-genome of EAB resistant Fraxinus species, and the other which utilized the 2021 release of the green ash genome (37, 51). Additional studies will be necessary to fully elucidate the genetic architecture of these defensive responses. Although we did not examine other components of the lingering ash phenotype, including adult feeding preferences or attractiveness of egg-laying sites to female EAB, our controlled greenhouse experiments did allow us to examine the defensive mechanisms deployed in the woody tissues, where the primary host insect interaction occurs. The chemotypes of high larval kill, low larval kill, and uninfested individuals from the same parents could be distinguished based on relative concentrations of groups of metabolites. Comparisons of the same contrast across multiple families reveals that secoiridoids are associated with a generalized infestation response that does not predict effective defensive 14 responses. This association is consistent with previous studies that suggested that infested trees, or trees artificially stressed with methyl jasmonate produced higher amounts of these metabolites (52). High concentrations of specific secoiridoids in F. excelsior are proposed to be indicative of tolerance (40) or susceptibility to ash dieback (37), and were predicted to provide a future robust reservoir of anti-feeding deterrents to EAB (49). Our data suggests that these specific secoiridoids function best as indicators of a generalized stress response and not necessarily of resistance to EAB. The study design allowed us to disentangle the generalized stress response from an effective defense response as indicated by percent larval kill. Our results suggest that part of the effective defensive response may consist of four metabolites, annotated as three aromatic alkaloids and salidroside, that were significantly elevated in high larval kill individuals compared to low larval kill or uninfested individuals. Overall, our study has distinguished, for the first time, between an effective defensive response and a generalized defensive response to EAB. Based on our results, we propose a two-part model for the North American Fraxinus response to EAB wherein every individual has the biochemical capacity to synthesize chemical defenses as a response to EAB, but only certain trees deploy an effective induced defense response that kills enough EAB larvae to prevent or minimize lethal damage to the vascular system. This model is consistent with forest observations and controlled studies that show most individuals in North American ash species can kill a few larvae, but cannot withstand a heavy infestation (24, 25). The high concentrations of secoiridoid glycosides in infested individuals, especially those with the most live larvae, suggests that even susceptible ash trees detect that they have been wounded by EAB larvae, and attempt to respond but are unable to do so in a 15 manner that results in effectively killing the larvae. A previous study demonstrated that application of methyl jasmonate induced a defensive response and suppressed EAB larval development or killed larvae in susceptible Fraxinus individuals (35), supporting the hypothesis that even susceptible trees have the necessary synthetic machinery, but lack the ability to conduct a tailored reconfiguration of their metabolism, as outlined by Schuman and Baldwin (53), to kill the EAB larvae. This study provides a list of metabolites that could be targeted in future work focusing on the response of green ash to EAB. Key questions for future experiments include determining if the compounds identified extend to additional lingering ash families and gaining a better understanding of the timing and spatial distribution of effective defense responses. Additional phenotypic, genomic, proteomic, transcriptomic, and metabolomic analyses will benefit from the recent release of the green ash genome (51). This future work on the interaction of green ash and EAB will contribute to our understanding of how forest trees recognize and defend themselves against stem-boring insects. In summary, our data supports the hypothesis that the high larval kill phenotype is a multi-genic and heritable trait. We have also shown that green ash responds to EAB infestation with increased concentrations of secoiridoids, regardless of the larval kill phenotype. While infestation with EAB induces a response in all green ash tested, the induced response is ineffective in most cases. In the individuals that mount a successful response, we found higher concentrations of three aromatic alkaloids and salidroside, a result that merits further investigation. Similar metabolites were seen across all phenotypes, but the concentrations varied, suggesting that the high larval kill phenotype is based on complex regulatory 16 mechanisms. Elucidation of the genetic mechanisms driving defensive responses to EAB in green ash will be an essential part of a multidisciplinary effort for saving North American Fraxinus species and guide future investigations of resistance in native species to invasive threats. Materials & Methods Study System and Phenotyping Green ash were selected in the forest based on two criteria: 1) a healthy canopy at least two years after the mortality rate of the stand exceeded 95 percent, and 2) a minimum diameter at breast height (DBH, 1.37 m from the ground) of 26 cm, indicating they were over the minimum size preferred by EAB when the infestation was at peak levels (24). These ‘lingering ash’ trees show evidence of less severe emerald ash borer infestation compared to susceptible phenotypes in the forest, often accompanied by evidence of vigorous wound healing, and maintain a healthy crown for years after local conspecifics have died(3, 54). Over the last 14 years, individuals meeting these criteria have been clonally propagated through grafting and subjected to greenhouse bioassays that provided evidence of the ability of some selected lingering ash trees to mount defensive responses against EAB (24). Although there is evidence of multiple types of defenses, this work is focused on EAB egg bioassays (described below) to assess host defenses that result in larval mortality. Clonal replicates of lingering green ash genotypes, some used as parents in this study, consistently kill more early instar larvae (35 to 50 percent) than the susceptible green ash controls (0 to 10 percent) (24). Plant Material 17 Plant material was comprised of 97 two-year-old potted F. pennsylvanica seedlings reared in an outdoor growing area, then transferred into an environmentally controlled greenhouse in the spring of the treatment year to allow acclimatization prior to the start of the EAB treatment. The individuals tested were generated by controlled cross-pollinations of lingering or susceptible green ash to produce full sibling families of known parentage. Individuals belonged to one of three families: Pe-C (21 individuals, susceptible parentage Pe-97 x “Summit”), Pe-Y (42 individuals, lingering parentage, Pe-53 x Pe-56), or Pe-Z (35 individuals, lingering parentage Pe-53 x Pe-59). Both susceptible parents, (Pe-97) and the cultivar “Summit”, had susceptible phenotypes in egg bioassays and did not persist on the landscape after the arrival of EAB. “Summit”, in particular, has been proven susceptible in our egg bioassay (16 replications), in common garden studies (55), and by its rapid demise under natural EAB infestation in city streets and parks (16). Emerald Ash Borer Resistance Bioassays EAB eggs were raised and prepared as described in Koch et al 2015 (24). Twelve eggs were applied to each tree at a density of 400 eggs per square meter, as previously described (24). Eight weeks after eggs were applied, larval galleries were carefully dissected, starting at the entry hole, and followed to determine the outcome of each larva that successfully hatched and entered the tree. Larvae were designated as alive, tree-killed (killed by a host defense response), or dead by other means such as parasitism, cannibalism, or fungal infection. The proportion of tree-killed larvae was calculated based on the total number of larvae that hatched and entered the tree. One-way ANOVA and Tukey-Kramer multiple comparison tests were used to analyze the performance of families Pe-Y, Pe-Z and Pe-C. 18 Metabolite analyses of F. pennsylvanica woody tissue by UHPLC-MS. Trees were destructively sampled to collect tissue for metabolite analyses eight weeks after eggs were placed, during phenotyping. The entire stem, 2.5 cm above the highest EAB larval galleries, was collected and stored immediately on dry ice, before being transferred to - 80°C storage. This ensured the collection of the vascular cambium, the cork cambium, the phloem, and the ray parenchyma. All samples were ground under liquid nitrogen in a Spex Sample Prep freezer mill and stored at -80°C prior to extraction For each sample, 1 g of frozen powdered plant tissues was extracted in 10 ml of acetonitrile/isopropanol/water (3:3:2) containing 1.00 mM telmisartan (internal standard) and 0.01% formic acid and incubated in the dark at 4°C for 24 h. samples were then centrifuged at 4°C and 10,000g for 10 minutes, supernatants were transferred to fresh tubes, and 50:1 diluted aliquots were prepared by adding deionized water. An additional aliquot of undiluted extracted sample has been archived at -80 ˚C. UHPLC/MS analyses were performed using a Shimadzu LC-20AD ternary pump coupled to a SIL-5000 autosampler, column oven, and Waters Xevo G2-XS QTof mass spectrometer equipped with an electrospray ionization source. The operation parameters for the positive-ion mode analyses are as previously detailed(56). A 10- µL volume of each diluted extract was analyzed using a 20-minute gradient method on an Ascentis Express C18UHPLC column (2.1x100mm, 2.7µm) with mobile phases consisting of 10 mM ammonium formate in water, adjusted to pH 2.8 with formic acid (solvent A) and acetonitrile (solvent B). The 20-min method gradient was as follows: 1% B at 0.00 to 1.00 min, then step to 5% B at 1.01 min, linear gradient to 25% B at 8.00 min, then a linear gradient to 75% B at 12.50 min, another linear gradient to 19 98% B at 15.00 min, and a hold at 98% B until 18.00 min, a step to 1% B at 18.01 min, and a hold at 1% B until 20.00 min. Analyte samples were injected in a randomized order while process blank and quality control samples were injected at regular intervals. All calculated peak areas were normalized to the peak area for the internal standard telmisartan utilizing Progenesis QI v2.4software (Nonlinear Dynamics Ltd., Newcastle, UK). Standards of oleuropein, apigenin and salidroside were purchased from Sigma Aldrich, prepared in the extraction solvent, and run at 5 µg/mL. Untargeted Metabolomics Data Processing For untargeted metabolomic analysis, data were initially processed using Progenesis software. Leucine enkephalin lockmass correction (m/z 556.2766) was applied during run importation and all runs were aligned to retention times of a bulk pool run automatically selected by the software from a selection of QC samples. Peak picking and deconvolution was conducted as previously described (57). After deconvolution, 1,278 compound ions remained. To remove features from the dataset introduced by solvents, glassware, or instrumentation and to remove lipids, several filters were applied to the 1,278 compound ions remaining after deconvolution. Concentrations of each feature were normalized to the internal telmisartan standard (m/z = 515.2448). Compounds with the highest mean abundance in process blank samples, maximum abundance less than 0.1% of the most abundant compound in the dataset, or retention times greater than 16 minutes were excluded from the dataset. This reduced the total number of metabolic features to 323. Further analysis and statistical comparisons of compound signals extracted by Progenesis QI software was executed using EZinfo v3.0.2 software (Umetrics, Umeå, Sweden). 20 One way analysis of variance (ANOVA) tests were used to assess significance between each pairwise comparison for individual metabolic features in the seven following contrasts: Family C: UNI vs LLK ; Family Y: UNI vs LLK, UNI vs HLK, LLK vs HLK; Family Z: UNI vs LLK, UNI vs HLK, LLK vs HLK. Features that were significant (p < 0.05) were included in pairwise orthogonal partial least squares discriminant analysis (OPLS-DA) and principal component analysis (PCA) analyses (Table S1). OPLS-DAs and PCAs were run using pareto scaling. To prevent overfitting of our model we performed a threefold cross validation on our data and took the averages as the performance of our model. For all metabolic features extracted with Progenesis QI and used in downstream analyses with EZinfo, spectra were processed using MassLynx v4.2 software (Waters Corporation, Milford, MA,USA) as previously detailed (57) (Table S2). Of the metabolites considered, thirty-two had significance in more than one family, or had a previously proposed purpose and were annotated. Annotation of the electrospray ionization tandem mass spectrometry (ESI-MS/MS) data relied on comparisons with MS/MS databases such as the Massbank of North America as well as previous studies and purchased standards. The confidence levels in the metabolite annotation were following recommendations of the Metabolomics Standards Initiative (48). The quantities present in individual tissue extracts were too small for complete structure elucidation. Acknowledgements Acknowledgments: The authors thank Warren Chatwin and Christina Murray for their helpful comments on the manuscript. The authors thank Aletta Doran, Julia Wolf, Gavin Nupp, Miranda McKibben, and Jarod Sanchez for their work propagating and maintaining the study trees and 21 their assistance conducting the EAB resistance bioassays. The authors also thank Patrick Cunniff, Brandon Chou, Kingsley Owusu Otoo and Julie Huston for assistance in collecting and organizing tissue samples and managing logistics. J.R-S acknowledges support from USDA-USFS APHIS grants 18-IA-11242316-105 and 20-JV-11242303-050. J.R-S also acknowledges support from the Tree Fund Foundation, Tree Fund grant 18-JD-01. R.K.S. acknowledges support from NIH training grant T32GM075762. JK acknowledges support from USDA APHIS 18-IA-11242316- 105, Michigan Invasive Species Grant Program grant IS18-119, the Commonwealth of Pennsylvania Department of Conservation and Natural Resources Bureau of Forestry 18-CO- 11242316-014, and the U.S Forest Service Special Technology Development Program grant NA- 2017-01. A.D.J. acknowledges support from Michigan AgBioResearch through the USDA National Institute of Food and Agriculture, Hatch project number MICL02474, and USDA-USFS grant 20-JV-11242303-050. Competing Interest Statement All the authors declare that they have no competing interests. Data Availability The data that support the findings of this study are available upon request from the corresponding author. The raw data will also be submitted to MetaboLights or similar repository. References 1. K. M. Potter, M. E. Escanferla, R. M. Jetton, G. 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Last, An integrated analytical approach reveals trichome acylsugar metabolite diversity in the wild tomato Solanum pennellii. Metabolites 10, 401 (2020). 57. R. Sadre et al., Metabolite diversity in alkaloid biosynthesis: a multilane (diastereomer) highway for camptothecin synthesis in Camptotheca acuminata. The Plant Cell 28, 1926- 1944 (2016). Figure Legends Figure 1: Egg Bioassay Protocol & Phenotypic Distributions. (a) EAB greenhouse bioassay protocol. Two year old green ash individuals are artifcially dissected at eight weeks to ascertain larval fate. (b) Percentage of EAB larvae kill by infested individuals in families Pe-C, Pe-Y and Pe-Z when sampled 8 weeks post infestation. Family C contained 17 F1 infested progeny from two susceptible parents. Families Pe-Y and Pe-Z both contained 30 F1 full sib progeny of two lingering ash parents. Family means of Pe-Y and Pe-Z were significantly different from the family mean of susceptible family C (p<0.0001) Figure 2: Data Processing Schematic. Flowthrough of the untargeted metabolomics workflow beginning with data generation, and highlighting the number of features at each stage of the analysis. 26 Figure 3: Classification summary. (a) Orthogonal partial least squares projection to latent square discriminate analysis (OPLS-DA) model performance, averaged across triplicate prediction models. The graph indicates that percentage that each model classified correctly, incorrection, or was unable to classify. (b) Principal component analysis plot comparing high and low larval kill (LK) in family Pe-Y, utilizing 43 features. (c) OPLS-DA model utilizing all test samples in a comparison of high vs low larval kill using 43 features. Figure 4: Metabolite Annotations. MS/MS spectra in positive ion mode support annotations of metabolite structures: (a) product ions of m/z 642.24 ([M+NH4]+) for verbascoside, (product ions of m/z 271.06 ([M+H]+) for apigenin, (c) product ions of m/z 584.21 ([M+NH4]+) for Excelside A. Figure 5: Chemical Families of Annotated Compounds. (a) proportions of the chemical families in the annotated metabolites. (b) Pairwise comparisons for specific compounds. ‘Number’ is metabolite number (table 1). LLK v UNI, HLK v UNI, HLK v LLK indicates pairwise comparisons between larval kill phenotypes or uninfested individuals. Pe-C, Pe-Y, Pe-Z refer to full sibling families (figure 1). Box with letter indicates the phenotypic category that had significantly higher concentration of the indicated metabolite (p < 0.05, L in red LLK, H in gray HLK, U in blue UNI). Annotated metabolites 1-6 are alkaloids, 9-17 are secoiridoid glycosides, 18-20 are secoiridoids, 24 is salidroside and 25 is verbascoside. ���������� ���������� ���������� � �� �� �� ��� ��������������������� � � � EAB eggs applied to two year old gra�ed trees EAB egg on filter taped to stem Healthy larva Host-killed larva Data collected 8 weeks later Egg hatched Y/N Larva dead/alive Larval instar: 1-4 Larval weight EAB Larvae hatches, chews through filter into tree * p<0.0001 a b Alkaloid Ammonium Flavone Lignan Lignan glycoside Phenolic glycoside Secoiridoid Secoiridoid glycoside Sugar Unknown 1278 Features 32 Features Collected MS/MS and annotated features 323 Features 194 Features Removed - Contaminants - Lipids - Low Abundance OPLS-DA generation with three fold cross validation, averages reported Selected all features that were signifcant in more than 1 family or had a previous proposed role 32 Features correct unkown incorrect Pe-C UNIvLLK Pe-Y UNIvLLK Pe-Y UNIvHLK Pe-Y HLKvLLK Pe-Z UNIvLLK Pe-Z UNIvHLK Pe-Z HLKvLLK percent assigned Calculated RSD and tested for signifcant relationships in pairwise comparisons Removed - Adducts - Multiple Fragments Pe-C UNI v LLK 34 features Pe-Y UNI v LLK 49 features Pe-Y UNI v HLK 44 features Pe-Y HLK v LLK 43 features Pe-Z UNI v LLK 25 features Pe-Z UNI v HLK 35 features Pe-Z HLK v LLK 9 features Average Model Performance 0 50 100 79 % 83 % 83 % 73 % 87 % 71 % 76 % HLK LLK LLK HLK � � � Unknown Correct Incorrect PC2:13% PC1:56% -80 60 30 0 -40 -140 0 -70 140 70 -70 70 35 0 -35 -120 0 -60 120 60 R2Y: 70% Q2: 59% Pe-C UNI v LLK Pe-Y UNI v LLK Pe-Z UNI v LLK Pe-Y UNI v HLK Pe-Y HLK v LLK Pe-Z UNI v HLK Pe-Z HLK v LLK percent assigned 0 20 40 60 80 100 Average Model Performance � �� ��� ����������������� ������������������ ������ ������ ��� �� � ��� �� � � � � �������� ���������������� ������������������ ������� 100 300 200 400 500 600 [M+H]+ [M+NH4]+ 642.24 325.09 163.04 O OH HO O OH ���������� HO HO O O O OH O O OH OH OH O HO OH H HO ���������������� ������������������� [M+NH4]+ ������ ������ ������ O O O O O O O HO HO OH OH HO OH OH O O 100 300 200 100 300 200 400 500 Alkaloid Ammonium Flavone Lignan Lignan glycoside Phenolic glycoside Secoiridoid Secoiridoid glycoside Sugar Unknown b a

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