Storage compartments for capillary water rarely refill in ... · 1 1 Title: Storage compartments...

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Title: Storage compartments for capillary water rarely refill in an intact woody plant 1

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Authors: Thorsten Knipfer1*, Italo F. Cuneo2, J. Mason Earles1,3, Clarissa Reyes1, Craig R. 3

Brodersen3, Andrew J. McElrone1,4* 4

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Institution Addresses: 6 1Department of Viticulture and Enology, University of California, Davis, CA 95616. USA 7 2School of Agronomy, Pontificia Universidad Católica de Valparaíso, Quillota, Chile. 8 3School of Forestry and Environmental Studies, Yale University, New Haven, CT 06511, USA 9 4USDA-ARS, Crops Pathology and Genetics Research Unit, Davis, CA 95618, USA 10

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Footnotes: 12

*Corresponding authors; e-mail: [email protected], [email protected], 13

T.K. designed and performed most of the experiments, analyzed the data, and wrote the article 14

together with A.J.M. 15

I.F.C., J.M.E., C.R. performed some of the experiments, helped in data analysis, and revised 16

the article. 17

C.R.B. helped in experimental design, performed the ESEM imaging, and revised the article. 18

A.J.M. obtained the grants, helped in experimental design, performed some of the experiments, 19

and wrote the article together with T.K. 20

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This work was supported by a NIFA SCRI grant and funding from the American Vineyard 22

Foundation to A.J.M. and USDA-ARS CRIS funding (grant no. 5306-21220-004-00). The 23

Advanced Light Source is supported by the Director, Office of Science, Office of Basic Energy 24

Science, of the US Department of Energy under contract no. DE-AC02-05CH11231. 25

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Summary: Visualization of refilling dynamics in L. nobilis revealed that both capillary water 27

storage and vessel refilling play a minor role under in-vivo conditions. 28

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Running Head: 34

Plant Physiology Preview. Published on October 17, 2017, as DOI:10.1104/pp.17.01133

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Visualization of water storage and refilling 35

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Corresponding Authors: 38

Thorsten Knipfer 39

Department of Viticulture and Enology, 40

University of California, 41

Davis, CA 95616, USA 42

Phone: 530-752-1762 43

E-mail: [email protected] 44

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Andrew J. McElrone 46

Department of Viticulture and Enology, 47

USDA-ARS, Crops Pathology and Genetics Research Unit, 48

University of California, 49

Davis, CA 95616, USA 50

Phone: 530-754-9763 51

Email: [email protected] 52

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Research Category: 55

Ecophysiology and Sustainability 56

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ABSTRACT 66



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Water storage is thought to play an integral role in the maintenance of whole plant water 67

balance. The contribution of both living and dead cells to water storage can be derived from 68

rehydration and pressure-volume curves on excised plant material, but the underlying tissue-69

specific emptying/refilling dynamics remain unclear. Here, we used X-ray computed micro-70

tomography (microCT) to characterize refilling of xylem fibers, pith cells and vessels under both 71

excised and in-vivo conditions in Laurus nobilis. In excised stems supplied with H2O, water 72

uptake exhibited a biphasic response curve, and microCT images showed that high water 73

storage capacitance was associated with fiber and pith refilling as driven by capillary forces; 74

fibers refilled more rapidly than pith cells while vessel refilling was minimal. In excised stems 75

that were sealed, fiber and pith refilling was associated with vessel emptying, indicating a link 76

between tissue connectivity and water storage. In contrast, refilling of fibers, pith cells and 77

vessels was negligible in intact saplings over two timescales, a period of 24-h and 3-weeks. 78

However, those compartments did refill slowly when the shoot was covered to prevent 79

transpiration. Collectively, our data i) provide direct evidence that storage compartments for 80

capillary water refill in excised stems but rarely under in-vivo conditions, ii) highlight that 81

estimates of capacitance from excised samples should be interpreted with caution as certain 82

storage compartments may not be utilized in the intact plant, and iii) question the paradigm that 83

fibers play a substantial role in daily discharge/recharge of stem capacitance in an intact tree. 84

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Keywords: 87

embolism, fiber, hydraulic capacitance, in-vivo, parenchyma, pith, x-ray micro-tomography, 88

xylem 89

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INTRODUCTION 100



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Stem internal water storage can prolong vessel functionality by sourcing water into the 101

transpiration stream and reducing the risk of gas emboli by buffering xylem tension (Tyree and 102

Sperry, 1989; Holbrook, and Sinclair, 1992; Holbrook, 1995; Cochard et al., 2013). Estimates of 103

water storage are commonly derived from water-release curves measured on excised stems 104

(Tyree and Yang, 1990; Jupa et al., 2016). These data indicate that water stored in dead fibers, 105

non-functional vessels and apoplastic pores provide the largest fraction of stored water in most 106

trees (i.e. capillary water storage). Living cells (e.g. xylem parenchyma) typically provide a 107

relatively small storage volume due to their limited elasticity, and this water is released under 108

more negative xylem pressures (i.e. elastic water storage). Experiments by Borchert and 109

Pockman (2005) indicate that storage compartments for capillary and elastic water refill during 110

stem rehydration, and refilling of capillary water storage requires xylem pressures of >-0.5 MPa. 111

However, the hydraulic methods typically used to measure elastic and capillary water storage 112

require excised plant material released from sustained negative pressures, and the resulting 113

data does not provide detailed information about the temporal, spatial, or tissue-specific 114

emptying or refilling dynamics of an intact plant. 115

116

Measurements on forest trees using sap flow sensors, isotopic tracers, frequency domain 117

reflectometry, and dendrometers indicate that the emptying and refilling of storage 118

compartments occurs on a daily basis (e.g. Goldstein et al., 1998; Cermak et al., 2007; Meinzer 119

et al., 2009; Hao et al., 2013; De Schepper et al., 2012; Carrasco et al., 2014). For example, a 120

release of stored water from the trunk can contribute up to 50% to daily transpiration (Waring et 121

al., 1979; Verbeeck et al., 2007), and tree transpiration can be maintained with stored water for 122

about one week (Cermak et al., 2007). Current data suggests that the volume of stored and 123

discharged can be substantial, which implies that tissue compartments of relatively high storage 124

capacity (such as dead fibers for storage of capillary water) are involved in this process and are 125

able to refill on a daily basis. . Similar to refilling of capillary water storage compartments 126

(Borchert and Pockman, 2005), successful vessel refilling appears to require xylem pressures 127

approaching or exceeding 0 MPa (Hacke and Sperry, 2003; Charrier et al., 2016). Therefore, 128

both refilling processes may be interrelated, and vessel refilling in some species may be limited 129

by a simultaneous recharge of capacitive tissue and competition for ‘free’ water. The link among 130

refilling of water storage compartments and vessel refilling remains to be determined. 131

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A large body of literature has been published over the last few decades concerning the 133

importance of plant water storage (examples cited above), but the tissue compartments that 134



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empty and refill in an intact plant and effectively contribute to maintenance of whole plant water 135

balance remain unclear. Here, our goal was to observe the refilling dynamics of the putative 136

compartments for water storage (fibers, pith) and long-distance, axial transport (vessels) under 137

both excised and in-vivo conditions, and to determine whether the lab-based hydraulics 138

methods used to study capacitance are measuring the same thing that occurs in an intact plant. 139

In turn, these data would then provide novel insight into the role of different tissue types in plant 140

hydraulic function. Experiments using non-invasive high-resolution X-ray micro-computed 141

tomography (microCT) imaging were complimented with traditional microscopy and 142

physiological measurements. Our study was performed on Laurus nobilis, an evergreen woody 143

plant native to the Mediterranean region that has been documented for its ability to restore plant 144

hydraulic function by vessel refilling (Salleo et al., 1996, 2004; Hacke and Sperry, 2003; Trifilo et 145

al., 2014). 146

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RESULTS 150

Excised stems 151

During rehydration of excised L. nobilis stems, water uptake revealed two distinct phases over a 152

period of 20 hours (Figure 1A). The initial phase of uptake was rapid (<1 h), followed by a 153

second phase that was relatively slow but steady. A biphasic response in water uptake was 154

observed both in stems with and without leaves indicating that this response predominantly 155

reflects stem internal water storage dynamics. During the initial uptake phase, Ψstem recovered 156

to values of close to zero and Ψstem stabilized at >-0.05 MPa during the second phase. The 157

biphasic response curve indicated that different tissue compartments refilled during the 158

rehydration process (Figure 1A), and the relationship of Ψstem and water uptake point to 159

differences in hydraulic capacitance among these compartments (Figure 1B). Capacitance of 160

tissue compartments providing for initial water storage was low (0.02 to 0.06 g MPa-1), whereas 161

capacitance of compartments providing for storage after >1 hour of rehydration was >10-fold 162

higher (Figure 1B). 163

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Tissue-specific refilling during rehydration of excised stems was visualized using microCT 165

imaging. For a representative excised stems supplied with H2O (Figure 2), fibers located in older 166

xylem close to pith and many fibers throughout the second annual ring were initially air-filled 167

(Aair-fibers=0.09 mm2, Figure 2A, t=2.5 h). At the same time, many pith cells were air-filled (Aair-168

pith=0.40 mm2), and the majority of embolized vessels were located in older xylem close to pith 169

(Aair-vessels=0.04 mm2) (Figure 2A, t=2.5 h). A few hours later during stem rehydration, Aair-fibers 170

(=0.05 mm2) was reduced by 45% and Aair-pith (=0.33 mm2) was reduced by 18%; 171

simultaneously, more embolized vessels appeared (Aair-vessels=0.05 mm2) (Figure 2A, t=7.5 h). 172

After 14 hours, Aair-fibers was as low as 0.01 mm2 (Figure 2A). After 18.5 hours, very few air-filled 173

fibers remained in the most recent annual ring, while all fibers in older xylem close to pith had 174

refilled (Aair-fibers=0.003 mm2) (Figure 2A). Remaining air-filled pith cells were concentrated 175

towards the stem center (Aair-pith=0.16 mm2), and many air-filled vessels persisted in an 176

embolized state (Figure 2A, t=18.5 h). These refilling dynamics were confirmed with 3-D 177

observations during a 6.5 hour period (Figure 2B); as visualized for a portion of the stem, the 178

air-filled volume of fibers and pith decreased from 0.006 to 0.003 mm3 and 0.135 to 0.055 mm3, 179

respectively, while changes in air-filled vessel volume were negligible. 180

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Across all excised stem samples, temporal dynamics of refilling differed among tissue 182

compartments and among treatments when stems were either rehydrated (+H2O, Figure 3A to 183



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3C) or entirely sealed (-H2O, Figure 3D to 3F) (see supplemental Table S1 for cross-sectional 184



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areas). In excised stems during rehydration, Aair-fibers declined rapidly (Figure 3A), while Aair-pith 185



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declined at a slower pace compared to fibers (Figure 3B). Simultaneously, Aair-vessels typically 186



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increased in the first couple of hours and decreased thereafter (Figure 3C). Curves fit to the 187

data using nonlinear regressions indicated that 50% of air-filled fibers refilled within around 5 188

hours of rehydration (Figure 3A); in comparison, a 50% reduction in air-filled pith tissue and 189

vessels required more than 15 and 35 hours, respectively (Figure 3B and 3C). Refilling of air-190

filled fibers and pith tissue was also observed for stems that were entirely sealed (Figure 3D and 191

3E), but Aair-fibers and Aair-pith declined more slowly as compared to rehydrated stems; curves fit to 192

the data using nonlinear regressions indicated a 50% reduction of Aair-fibers and Aair-pith after 193

around 30h. Moreover, refilling of fibers and pith tissue was accompanied by a general increase 194

in embolized vessels (Figure 3F), pointing to internal water redistribution from vessels to refilling 195

fibers and pith in stems that were entirely sealed (for example see circle symbols); increases in 196

embolized vessels were least pronounced for sample ‘diamond symbols’ (Figure 3F), which 197

suggest that water from alternative tissue sources was also redistributed towards fiber and pith. 198

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MicroCT images provided visual evidence that tissue-specific refilling in stems was dominated 200

by capillary forces, and as shown for a representative excised stem during rehydration (Figure 201

4): Transverse microCT images showed that the lumen of several adjacent air-filled fibers 202

appeared water-filled three hours later (Figure 4A). Corresponding longitudinal images indicated 203

that while fibers refilled, water columns expanded inside the air-filled lumen from both ends 204

(Figures 4B and 4C), and the liquid/air meniscus with the fiber wall was concave shaped relative 205

to the direction of flow (contact angle ranged from 44 o to 63o, see also supplemental Figure S1 206

for fibers located close and further away from pith). For pith tissue (Figure 4D and 4E), enlarged 207

longitudinal images showed that air-filled cells located in proximity to xylem refilled first (Figure 208

4D). Refilling and water entry into the air-filled pith cell initially resulted in the formation of a 209

concave shaped liquid/air meniscus with the cell wall, and the appearance of a spherical air-void 210

before completion of refilling (Figure 4E). Refilling of the air-filled vessel lumen was related to 211

water droplet formations on the lateral vessel wall and water column expansion (Figure 4F); the 212

liquid/air meniscus with the vessel wall was variable in shape. 213

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Stems in intact plants 215

In contrast to excised conditions, tissue-specific refilling was negligible in the stem of intact 216

saplings (Figures 5 and 6). In a representative L. nobilis sapling, Ψstem recovered from -1.5 MPa 217

to -0.4 MPa after 20 hours of soil rehydration, but Aair-fibers (=0.58 mm2), Aair-vessels (=0.17 mm2) 218

and Aair-pith (=0.54 mm2) remained at similar levels under in-vivo conditions (<6 % change) 219

(Figure 5A and 5B). In line with previous observations, after the stem was excised and 220



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rehydrated for 5 hours, Aair-fibers declined by 43 % to 0.33mm2 which was accompanied by a 13% 221



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and 23% reduction of Aair-pith and Aair-vessels, respectively. 222

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For intact saplings with the shoot not bagged and exposed to ambient conditions (Figure 6A to 224

6C), changes in Aair-fibers, Aair-pith and Aair-vessels by refilling were minimal during recovery in Ψstem 225

from <-1MPa to ~ -0.5 MPa following soil saturation after drought. Similarly, intact saplings that 226

were maintained well-watered and entered the experiment at less negative Ψstem (-0.5 and -0.4 227

MPa), no reductions in Aair-fibers, Aair-pith and Aair-vessels were observed; even after maintaining 228

saplings under well-watered conditions for an additional 3-weeks (Figures 6). However, for well-229

watered saplings where the shoot was bagged during the time period of investigation, there was 230

evidence for a slow but gradual reduction in Aair-fibers, Aair-pith and Aair-vessels over time (Figures 6D 231

to 6F); curves fit to the data using nonlinear regressions indicated that refilling of tissues for the 232

bagged plants resulted in a reduction of Aair-fibers, Aair-pith and Aair-vessels by around 20% after 20 233

hours. 234

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Stem anatomical features 236

Tissue viability staining showed that fibers in stem xylem were not metabolically active and 237

dead, as well as most pith cells towards the stem center (Figure 7A to 7C). Xylem parenchyma 238

cells were relatively inactive in their metabolic activity at the time of analysis as evident from 239

small amounts of ray tissue emitting a green fluorescence signal. Within the pith, only cells 240

located closest to xylem tissue (first 1 to 3 cell layers) were living (Figures 7A -C). Anatomical 241

features of fibers, pith tissue, and vessels were further characterized using ESEM and microCT 242



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imaging (Figure 7D to 7F). ESEM images showed the existence of fiber-to-fiber pits (Figure 7D) 243



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and vessel-vessel bordered pits (Figure 7E). Adjacent fibers were interconnected via pits; the 244

lumen of fibers reconstructed in 3-D (in red color) was clearly visible through pit openings after 245

fibers were sliced open (Figure 7F, panel 1). Images from 3-D volume renderings also indicated 246



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that fibers were interconnected to neighboring vessels (predominantly in tangential direction). 3-247

D volume renderings of the pith showed that the cell wall contained many cell-to-cell 248

connections (Figure 7F, panel 2). 249

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DISCUSSION 253

Water storage and release has been described as an integral physiological process that 254

contributes to whole plant water balance (e.g. Tyree and Sperry, 1989; Holbrook, 1995; Hao et 255

al., 2013). In this study on L. nobilis, we characterized the tissue-specific refilling dynamics 256

related to capillary water storage under both excised and intact (in-vivo) conditions, which 257

revealed significant differences between rehydration processes that take place within an intact 258

plant compared to those that occur in the type of stem material generally used for empirical 259

hydraulics measurements in the lab. Time-series microCT imaging indicated that in intact 260

saplings refilling of fiber and pith tissue was negligible over periods ranging from 20-h to 3-261

weeks unless the shoot of the sapling was bagged and completely covered with petroleum jelly 262

to prevent any cuticular water loss and transpiration. In comparison, fiber and pith tissue refilled 263

within hours in excised stems during rehydration. It can be speculated that L. nobilis, which is a 264

Mediterranean species, would only experience these conditions infrequently during the growing 265

season, if at all, or during rainy winter months that coincide with soil saturation, lower 266

transpiration, and a wet canopy (LoGullo and Salleo 1988; Rhizopoulou and Mitrakos 1990). 267

Contrary to the diurnal discharge and refilling of water from living cells in the bark (de Schepper 268

et al., 2013), our data indicate that for small trees i) refilling of dead tissue compartments with 269

capillary water is an exception and not the rule under in-vivo and transpiring conditions, and ii) 270

dead fiber and pith tissue plays a negligible role in buffering the daily fluctuation in xylem 271

tensions in an intact tree once empty (i.e. ‘single-use water reservoir’). Our conclusions are 272

based on the dynamics of refilling from saplings studied here, but it needs to be tested if these 273

results extend to larger trees in the field, across species that differ in stem anatomical features, 274

and those that grow in different climate regions. 275

276

Several research groups have documented large daily cycles of capacitance discharge and 277

subsequent recharge in trunks of large trees (e.g. Goldstein et al., 1998; James et al., 2003; 278

Cermak et al., 2007; Hao et al., 2013). Using indirect measurement techniques to quantify water 279

storage capacitance, these authors concluded that: i) water storage is important for maintaining 280

short- and long-term plant water balance (e.g. Hao et al., 2013); ii) the diurnal withdrawal of 281

water from, and refill of, internal stores is a dynamic process (e.g. Goldstein et al., 1998); iii) the 282

exchange of water between storage compartments and the transpiration stream has a 283

substantial influence on axial and radial stem water transport (e.g. James et al., 2003); and iv) 284

sapwood is the most important storage site for water (e.g. Cermak et al., 2007). Our study was 285

performed on young intact saplings, and while we cannot comment directly to the contribution of 286



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the trunk of large trees to water storage, we have, for the first time, pinpointed the exact sites of 287

capillary water storage while also characterizing the limitations to their refilling. 288

Since the process of capillary water storage in plants should follow the same biophysical 289

rules independent of stem age, size, or organ type (for fiber refilling see discussion below), our 290

data raise some valid concerns about the current paradigm that capillary water storage (such as 291

in dead fibers) contributes to the daily discharge and recharge of stem capacitance. If this 292

phenomenon extends beyond L. nobilis and is present in other tree species and mature trees, 293

this would require a significant revision of our understanding of xylem structure-function. 294

However, many open questions still remain, such as: Which capacitive tissue compartments 295

provide a means to protect xylem function under drought? How is water re-distributed within 296

stems to buffer daily fluctuations in xylem sap tension? Are there fiber types with specialized 297

anatomical structures that facilitate water storage and release in certain plant species? 298

299

Our current knowledge on tissue-specific water storage is largely based on measurements of 300

water-release and rehydration curves obtained from excised material (e.g. Tyree and Yang, 301

1990; Borchert and Pockman, 2005; Carrasco et al., 2014; Jupa et al. 2016). The advantage of 302

our method is that it allows the researcher to obtain the exact volume of water released/stored 303

for a given change in water potential, and in turn capacitance of different tissue compartments 304

(capillary versus elastic storage) can be derived from the shape of the curve. Water-release 305

curves provide indirect evidence for tissue-specific capacitance under excised conditions, but 306

direct observations of tissue-specific water storage is lacking for most woody species under in-307

vivo conditions. Using microCT imaging we were not able to determine tissue-specific 308

capacitance and volume fractions of water in different tissue regions, but we were able to 309

visualize that substantial temporal differences in fiber refilling can exist under in-vivo and 310

excised conditions, potentially complicating the interpretation of measurements on excised 311

material. For intact L. nobilis saplings, our microCT data show that fibers rarely refill in-vivo 312

after soil saturation suggesting that estimates of capillary water storage obtained on excised 313

woody stems should not be considered as a significant part of the overall stem water storage 314

capacity once these compartments have emptied in the intact plant; otherwise, this may 315

overestimate the ‘real’ water storage capacity that the intact plant utilizes on a regular basis. 316

Nevertheless, the exact implication of our findings for intact, mature trees in the field need to be 317

investigated in more detail in future experiments. 318

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Woody tree species growing in dry environments typically experience low Ψstem values of < -1.5 320

MPa throughout the year, and species with a high abundance of non-living tissue and denser 321

wood usually exhibit the most negative Ψstem (Borchert and Pockman, 2005; Meinzer et al., 322

2009). In L. nobilis, Ψ ranges from -0.4 to -2.5 MPa over the growing season reaching most 323

negative values in late summer (LoGullo and Salleo, 1998; Nardini et al., 1996). Nardini et al. 324

(2016) showed that water is lost from fibers under water stress in L. nobilis, but this observation 325

was not addressed in detail and fiber refilling was not investigated. For L. nobilis and other tree 326

species, this raises the question how and if compartments for capillary water storage refill? To 327

date, microCT imaging data by Suuronen et al. (2013) provides the only visual evidence for 328

refilling of fibers, which was dependent on environmental conditions such as high temperatures 329

and darkness. Similarly, our data highlight that both fiber and pith refilling requires 330

environmental conditions that presumably induce a substantial relaxation of xylem tension. Data 331

collected here during rehydration of excised stems indicated that capillary water storage in dead 332

fibers and pith requires a local Ψstem of >-0.1 MPa in L. nobilis. 333

334

Water transport from vessels into fiber lumen and among fibers is generally considered to occur 335

via pits (Siau, 1984). In Lauraceae species, xylem fibers are typically of the libriform-type, dead, 336

with thick-walls, and simple bordered pits (Esau 1959; Schweingruber et al., 2011). Similarly, 337

fibers studied here were non-septate with scanty pitting to adjacent vessels or fibers. MicroCT 338

data obtained from excised stems that were entirely sealed (no H2O supplied) showed that 339

additional vessels embolized while fibers simultaneously refilled, pointing to water transport via 340

pits during this process. Tyree et al. (1999) showed that vessels in L. nobilis are surrounded in 341

places by paratracheal parenchyma cells that separate the vessel lumen from fibers. Because 342

xylem cavitation in L. nobilis is thought to be nucleated by microbubbles entering the vessel 343

lumen (Salleo et al. 1996), and fibers can function like transport bridges among vessels (Cai et 344

al., 2014), it can be speculated that this layer of paratracheal parenchyma may impose an 345

important barrier restricting the passage of air towards remaining functional vessels. 346

347

Tradeoffs between xylem efficiency and embolism safety have been linked to water storage and 348

fiber traits in angiosperms (see review of Pratt and Jacobsen, 2016). For L. nobilis saplings as 349

used here, microCT data combined with additional transpiration measurements indicated that 350

water lost from fibers would only contribute a negligible amount of water to the transpiration 351

stream. Calculations showed that a volume of water equivalent to the maximum volume of air-352

filled fibers in the stem (approximated to be on average 6% =Aair-fibers/Astem in saplings analyzed, 353



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see supplemental table S1), would be transpired in only 2 min during the daytime (transpiration 354

rate of 4.0±0.4 x10-6 m3 h-1, measured gravimetrically from water loss of potted saplings where 355

the soil was covered with plastic foil) and 30 min during the nighttime (0.3±0.01 x 10-6 m3 h-1); 356

for calculations the estimated stem volume and maximum air-filled fiber volume of saplings (leaf 357

area was 25693±1211 mm2) was 2596±73 mm3 (as derived from stem height and diameter) 358

and 159±4 mm3, respectively. Since the amount of water released from fibers was relatively 359

small compared to the volume of water transpired and fibers rarely refilled in-vivo, it can be 360

inferred that fibers are rather ineffective in discharging water into the transpiration stream to 361

buffer daily fluctuations in xylem tensions in L. nobilis saplings. However, water stored in fibers 362

may have been large enough to be of relevance on a localized tissue level by providing water 363

into xylem parenchyma cells, thereby allowing for turgor maintenance and cell function when 364

xylem tensions fluctuated, but this topic needs more in-depth experimental testing. 365

366

In theory, fibers can only rehydrate if the liquid/air meniscus inside the lumen is able to 367

overcome the negative pressure in neighboring xylem vessels (Px). The negative pressure (P, 368

relative to atmospheric pressure) of a water column inside a cylindrical tube can be estimated 369

according to the Young-Laplace equation: 370 = 2 cos /

where γ is the surface tension of water (0.072 N m-1), Θ is the liquid/wall contact angle of the 371

meniscus, and r is the radius. MicroCT images collected in this study showed that fibers had a 372

radius ranging from 2.5 to 9 μm. For a concave-shaped meniscus with a contact angle of around 373

60o, we estimated a P of -0.001 to -0.005 MPa, respectively, which suggests that only under 374

conditions when Px is >-0.005 MPa will water enter the air-filled lumen via capillary forces. Given 375

that vessels and pith cells were of larger lumen diameter than fibers, it would require a Px of >>-376

0.005 MPa for those compartments to refill solely by capillarity. Such values were not obtained 377

from indirect measurements of xylem pressure by Ψstem, but it can be speculated that such 378

pressures exist locally where refilling of these tissue compartments was observed. 379

380

Pith tissue commonly suffers mechanical damage during development, and cells lose their 381

ability to store carbohydrates and become devoid of content as stems mature (Esau 1959). 382

Under drought and stem elongation, autolysis of pith cells is a common phenomenon leading to 383

air-filled cell cavities (for review see Beers et al. 1997). Together, microCT and fluorescent light 384

microscopy images provided visual evidence for the spatial distribution of dead and living pith 385

cells in young stems of L. nobilis, and data indicate that most pith cells are air-filled, dead and 386



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devoid of starch granules, while the remaining living pith cells are located at the periphery close 387

to xylem. Our data suggest that during early stages of stem development xylem parenchyma 388

may be the only effective site for carbohydrate storage in L. nobilis (Plavcova and Jansen, 389

2015). Furthermore, in woody stems with extensive secondary growth, the pith tissue typically 390

becomes ‘crushed’ and disappears. Our microCT data on L. nobilis indicated that before the pith 391

becomes ‘crushed’, air-filled cavities of dead pith cells can fill with water especially for excised 392

samples. In turn, water released from dead pith cells may contribute significantly to measured 393

water volume obtained from water-release curves; especially when dead pith tissue occupies a 394

large fraction of the entire organ volume. 395

396

Recent literature using microCT imaging indicates that most woody plant species studied lack 397

an efficient mechanism for short-term (days) embolism repair in-vivo (Brodersen and McElrone 398

2013; Cochard and Delzon, 2013; walnut, Knipfer et al., 2015b; redwood, Choat et al., 2015), 399

with grapevine as the exception (Brodersen et al., 2010; Knipfer et al., 2015a, 2016; Charrier et 400

al., 2016). However, CryoSEM data collected by Tyree et al. (1999) indicate that embolism 401

repair in L. nobilis is associated with water droplets emerging from lateral walls similar to in-vivo 402

observations on grapevine (Brodersen et al., 2010; Knipfer et al., 2015a, 2016). In line with 403

these findings, the present microCT images emphasize that L. nobilis stems have the ability to 404

form water droplets on lateral vessel walls and refill by water column expansion, but over the 405

time course of investigation vessel refilling was insignificant. Hacke and Sperry (2003) reported 406

that vessel refilling in L. nobilis required maintenance of plants at Ψstem >-0.3 MPa for 1-h by 407

pressurizing the root system. Together with our data, it can be followed that a mechanism for 408

short-term embolism repair by water droplet growth is mostly inactive in L. nobilis and water 409

column expansion inside the vessel lumen may also be related to capillary action, similar to the 410

refilling mechanism of dead fibers and pith cells at Ψstem of close to zero. 411

412

413 MATERIAL AND METHODS 414

Plant Material 415

Experiments were performed on excised stem material as obtained from terminal branches of L. 416

nobilis trees growing at the Arboretum, University of California Davis. Terminal branches were 417

harvested with pruning shears about 1 m behind the shoot tip, and maintained in a sealed 418

plastic bag containing a moist paper towel prior analysis. In addition, in-vivo experiments were 419

performed on the main stem of intact L. nobilis saplings (ca. 30 cm in height) that were obtained 420



21

from ArtForm Nurseries (Chagrin Falls, Ohio, USA). Saplings (n=7 in total) were grown in 4-inch 421

diameter plastic pots filled with soil mix (equals parts of peat moss, composted bark, sand and 422

perlite) and maintained for 4 weeks under greenhouse conditions (approximated day/night 423

temperature of 8 / 25 ˚C, photoperiod of 15 / 9 h, relative humidity of 35 %) at University of 424

California, Davis. Saplings were irrigated daily with water supplemented with macro and micro-425

nutrients (similar to Knipfer et al., 2015a,b). Prior to analysis, some saplings were subjected to 426

drought by not watering for 3-7 days while others were maintained under well-watered 427

conditions. Because we were not able to monitor the entire growth period of L nobilis branches 428

or saplings, plant material used may have experienced some level of stress resulting in air 429

embolism prior investigation. 430

431

Stem water potential 432

Water status of intact saplings and harvested branch material was measured with a Scholander 433

Pressure Chamber (Plant Moisture Stress Model 1505D, PMS Instrument Company, Albany, 434

OR, USA) on mature leaves that were covered and sealed with a foiled plastic bag for >30 min 435

(Knipfer et al., 2015a,b). The measured parameter was defined as stem water potential (Ψstem). 436

437

Water uptake curves 438

Terminal branches were harvested and transported to the laboratory as described in section 439

‘Plant Material’. In the laboratory, the branch was maintained for 1-2 hours in the sealed plastic 440

bag to allow for Ψstem-equilibration. Subsequently, initial Ψstem was measured on an apical and 441

basal leaf, which were located at opposite sides of the stem portion of interested (length of 7 to 442

13 cm containing six leaves). When corresponding Ψstem values differed by less than 0.05 MPa, 443

stem surface and leaves were covered with petroleum jelly and plastic foil to prevent 444

evaporation, and the portion of interest was excised with a fresh razorblade from the branch. 445

The apical cut of the excised stems was sealed with petroleum jelly and a 2-cm piece of PVC 446

tubing was placed over this stem end to hold the petroleum jelly in place. The distal cut was 447

connected to water-filled PVC tubing that was inserted into a water-filled cylinder that was sitting 448

on an electronic balance (Mettler, Toledo). Stem water uptake was recorded continuously in 30 449

sec intervals by weight change of the cylinder. During stem rehydration, Ψstem was measured on 450

leaves harvested from the excised stem. Water uptake was also measured for excised stems for 451

which all leaves were removed during sample preparation and prior analysis. Stem hydraulic 452

capacitance was determined from the slope (in g/MPa) of the linear portions of the relationship 453

of Ψstem versus water uptake (see also Borchert and Pockman, 2005). 454



22

455

X-ray micro-tomography 456

Plant material was scanned at the X-ray micro-tomography facility (Beamline 8.3.2) at the 457

Lawrence Berkley National Laboratory. Saplings were transported by car to the Advanced Light 458

Source (ALS, Lawrence Berkeley National Lab,Berkeley, CA) less than 4 h prior to analysis. To 459

assess the impact of initial plant water status on tissue-specific dynamics of refilling, saplings 460

subjected to microCT analysis covered a range of initial Ψstem (-1.5 to -0.2 MPa) as measured at 461

ALS after arrival. During the time period of microCT investigation, the soil of saplings was fully 462

saturated with water and Ψstem was measured periodically; for some saplings, the entire shoot 463

was coated with petroleum jelly and covered with a sealed plastic bag containing a wet paper 464

towel to test if minimizing transpiration affects tissue-specific refilling dynamics. For visualization 465

of stem tissue, the potted sapling was placed in an aluminum cage and the same stem portion 466

located 2 to 3 cm above the soil was scanned repeatedly over a period of 24 h. Some saplings 467

were transported back to the greenhouse and maintained under well-watered conditions for an 468

additional 3 weeks before the stem portion was subjected to a re-scan. 469

470

Excised stem samples were prepared within 5 to 8 hours following branch harvest, and were 471

either rehydrated by supplying water to cut ends or entirely sealed. For both types of 472

experiments, excised stem samples containing no leaves were prepared as follows: Ψstem was 473

measured on a bagged leaf of the branch that was located less than 2 cm away from the stem 474

portion of interest. Immediately after, the branch was submerged in water, and a stem portion of 475

5 to 10 cm in length was cut under water with pruning shears. (i) For rehydrated excised stems, 476

stem ends were recut using a fresh razor blade to remove air trapped at distal ends during the 477

initial cut and in turn ensure maximal connectivity to externally supplied H2O. Following the 478

procedure by Knipfer et al. (2016), the trimmed stem (length 3 to 8 cm) was connected to a 2 479

cm piece of PVC tubing on the top end and to a 2 cm piece of PVC-tubing attached to a valve 480

(i.e. open position) on the bottom end. Tubing was sealed with the stem using high vacuum 481

grease (976V, Dow Corning Company). PVC tubing was entirely filled with H2O, and the valve at 482

the bottom end was closed. The stem with attached tubing was removed from the water bath 483

and its entire surface was covered with vacuum grease to prevent surface evaporation. (ii) For 484

excised stems that were not supplied with H2O, stems were excised under water as described 485

above, removed from the water bath, and the entire stem surface including the cut stem ends 486

were coated with vacuum grease and wrapped with parafilm to prevent evaporation. Following 487



23

sample preparation, excised stems were placed in a sample holder and the same stem portion 488

about midway along the sample was scanned repeatedly over time. 489

490

Stems were scanned in a 21 keV synchrotron X-ray beam using a continuous tomography 491

setting yielding 1025 two-dimensional longitudinal images (resolution of 3.22 μm/pixel) that were 492

captured on a CMOS camera (PCO.edge; PCO, Kehlheim, Germany) at 350 ms exposure time. 493

Acquired raw images were reconstructed into transverse images using a custom software plugin 494

for Fiji image-processing software (www.fiji.sc, ImageJ) that used Octopus software (ver. 8.3; 495

National Institutes for Nuclear Science, Ghent University,Ghent, Belgium) in the background 496

(Knipfer et al., 2016). Longitudinal images were generated using the slice tool in the software 497

AVIZO (ver. 6,2; Visualization Sciences Group / FEI, Hillsboro, OR). 498

499

The cross-sectional area of air-filled fibers, pith tissue, and vessels was quantified from binary 500

images as generated from representative transverse microCT images using a semi-automated 501

routine in Fiji software: For a time-series of microCT images, the contrast and brightness was 502

adjusted so that air-filled tissue was clearly visible at comparable intensity (‘Image-Adjust’ tool). 503

Subsequently, the xylem including pith was extracted manually using the ‘Image-Crop’ tool, and 504

the ‘Image-Threshold’ tool was used to label exclusively air-filled tissue; a noise filter was 505

applied (‘Process –Noise –Despeckle’ tool) to remove black outlier pixels. Subsequently, 506

labelled air-filled pith tissue was erased manually (‘Paint brush’ tool), and the remaining air-filled 507

cross-sectional area of fibers and vessels (Aair-xylem) was measured (‘Analyze Particle’ tool). 508

Following this step, labelled air-filled vessels were erased manually (‘Paint brush’ tool) from 509

images and the cross-sectional area of remaining air-filled fibers (Aair-fibers) was measured 510

(‘Analyze Particle’ tool); air-filled cross-sectional area of vessels (Aair-vessels) was determined by 511

Aair-xylem – Aair-fibers. By using the original binary image again, Aair-xylem was erased and the 512

remaining cross-sectional area of air-filled pith (Aair-pith) was measured (‘Analyze Particle’ tool). 513

The percentage changes of air-filled tissue was determined by (Aair-x (Scan2,3,..., n) / Aair-x (Scan1)) x 514

100% (subscript ‘x’ = fibers, pith tissue, or vessels). Based on these data, the rate of %-515

reduction in Aair-fibers, Aair-pith, and Aair-vessels over time was estimated from the slope (% h-1) 516

following linear regression analyses. In addition, contact angles of liquid/air menisci within the 517

lumen of air-filled tissues were measured on longitudinal microCT images using the ‘Angle tool’ 518

in Fiji image-processing software (www.fiji.sc, ImageJ). 519

520



24

For 3-D visualization, stems were imaged at higher resolution (1.27 μm/pixel). The stack of 521

microCT images was uploaded into Fiji image-processing software (www.fiji.sc, ImageJ), and a 522

semi-automated routine was used to segment air-filled portions of each tissue type: Pith tissue 523

was separated manually from xylem using the ‘Polygon’ tool, and both images stacks were 524

saved separately. The ‘‘Image-Threshold’ tool was used to label air-filled fibers and vessels in 525

one image stack and air-filled pith in the other image stack. For the binary image stack of air-526

filled fibers and vessels, the ‘size criterion’ feature as part of the ‘Analyze particle’ tool was used 527

to separate both air-filled tissue types, and image stacks were saved separately. Image stacks, 528

were inverted using Fiji software, uploaded into AVIZO software, and air-filled fibers, pith tissue 529

and vessels were visualized in 3-D using the ‘volume rendering’ tool. 530

531

Stem anatomy and tissue connectivity was studied on dry stem samples scanned at very high 532

resolution (0.96 μm/pixel). For imaging, stem samples (around 3 cm in length) were prepared 533

from branches harvested in the Arboretum at UC Davis, and dehydrated slowly at around 30 ˚C 534

for 5 days prior scanning. Dimensions of fibers, vessels, pith cells were determined from 535

microCT images using the ‘Line’ and ‘Polygon’ tools in Fiji image-processing software 536

(www.fiji.sc, ImageJ). 537

538

Environmental-SEM (ESEM) imaging 539

ESEM experiments were performed to validate anatomical observations from microCT images. 540

Stem samples (around 5 cm in length) were excised from branches collected at the UC Davis 541

Arboretum, placed in a plastic bag containing a wet paper towel, sent overnight to the 542

Environmental-SEM (ESEM) facility at Yale University, stored at 4 ˚C, and imaged <48 h after 543

harvest. Samples were dissected with a razor blade to expose the xylem, and fresh tissue 544

fragments ca. 2 mm2 were placed on a Peltier-cooled stage and maintained at 0.5 ˚C during 545

ESEM imaging. Samples were observed at 10.0 kV with a FEI/Philips Field Emission XL-30 546

ESEM under true environmental mode. Water vapor was injected into the sample chamber at 547

4.0 Torr water vapor pressure, thereby establishing a 95 % relative humidity to prevent 548

desiccation. With this sample preparation no sputter coating was required. 549

550

FDA-PI viability staining 551

Stem tissue viability was analyzed using a fluorescence-based staining assay (Krasnow et al., 552

2008; Knipfer et al. 2016). For analysis, two fluorescent dyes (fluorescein-diacetate ‘FDA’ and 553

propidium iodide ‘PI’) were used simultaneously that allow a two-color discrimination between 554



25

living and dead tissue compartments (FDA can permeate through the intact cell membrane and 555

non-fluorescence FDA is converted into the green fluorescence metabolite fluorescein if cells 556

are living and the esterase enzyme is active; PI cannot permeate through the intact cell 557

membrane, and stains the cell wall or the cell nuclei if the membrane is disrupted/leaky). The 558

FDA-PI staining solution was prepared by adding 8 µl of FDA and 50 µl of PI to 5 ml of water. 559

For analysis, stem samples were obtained from branches harvested at Arboretum, University of 560

California Davis. Transverse stem sections were cut free-handed using a fresh razor blade, and 561

were immediately submerged in the staining solution for 30 min and incubated in dark at ~23 ˚C. 562

Subsequently, samples were mounted on a glass slide and observed under fluorescent light 563

(excitation filter 490 nm and 575 nm, dichromatic mirror 505 nm, barrier filter 525 nm and 625 564

nm) using a Leica DM4000 B LED microscope equipped with a Leica DFC7000 T 2.8 MP 565

camera. Images were captured in <4 h following sample preparation. 566 567 Supplemental Data 568

Table S1. Summary of cross-sectional areas and Ψstem of samples used in microCT 569

experiments. 570

Figure S1. Visualization of fiber refilling in an excised stem during rehydration for fibers that 571

were located either close or further away from pith. 572

573

574

ACKNOWELEDGEMENTS 575

The authors kindly thank D. Parkinson and A. MacDowell for their assistance at the Lawrence 576

Berkeley National Laboratory Advanced Light Source Beamline 8.3.2 microtomography facility. 577

The Advanced Light Source is supported by the Director, Office of Science, Office of Basic 578

Energy Science, of the US Department of Energy under contract no. DE-AC02-05CH11231. 579

580

581

582

FIGURE LEGENDS 583

Figure 1: (A) Time course of water uptake (indicated by solid line) and corresponding Ψstem 584

(indicated by symbols) during rehydration of excised L. nobilis stems. Following measurement of 585

initial Ψstem (t=0h), the stem was connected to a water source and stem surface and leaves were 586

sealed to prevent evaporation during rehydration. Values in legend are length and diameter of 587



26

excised stems. Stems indicated in blue color (with leaves to measure Ψstem) and red color (no 588

leaves) were obtained from the same branch and analyzed simultaneously. (B) Relationship of 589

Ψstem and corresponding amount of water uptake. For analysis of the stem indicated in red color 590

(no leaves), Ψstem values of the stem indicated in blue color were used. Linear regression 591

analysis allowed to determine capacitance (values in g MPa-1) of elastic (dashed line fitted 592

across data point 1 to 3 at most negative Ψstem) and capillary (solid line fitted across data point 3 593

to 6) storage compartments. 594

595

Figure 2: Visualization of tissue-specific refilling dynamics during rehydration (at t=0h) of an 596

excised L. nobilis stem (length ~5cm containing no leaves; indicated by circle symbols in 597

Figures 4A to 4C). (A) Binary images were generated from transverse microCT images and 598

show air-filled fibers (top row), pith tissue (middle row) and vessels (bottom row) in black color. 599

Values in mm2 are air-filled cross-sectional areas; lines indicate the estimated boundaries 600

between xylem annual rings (dashed line) and xylem to pith (solid bold line). (B) Corresponding 601

3-D visualizations of air-filled fibers (green color), pith tissue (white color) and vessels (gold 602

color) for a portion of the stem; values in mm3 are air volumes (some of the air-filled protoxylem 603

vessels were excluded to allow for a better view of pith). 604

605

Figure 3: Temporal refilling dynamics of air-filled fibers, pith tissue and vessels in excised L. 606

nobilis stems that were either (A-C) supplied with H2O at cut ends or (D-E) entirely sealed. 607

Values in MPa are initial stem water potentials measured immediately prior sample preparation; 608

all excised stems contained no leaves. Dashed lines provide an estimate of tissue-specific 609

refilling dynamics and were obtained from nonlinear regression analysis across data points of all 610

samples (y=a∙e(b/(x+c); A, a=0.004, b=582, c=57, R2=0.92, P<0.0001; B, a=0.005, b=2279, c=229, 611

R2=0.73, P<0.0001; D, a=46, b=7, c=8, R2=0.63, P=0.018; E, a=25, b=56, c=40, R2=0.60, 612

P=0.02; lines were omitted for panels C and F were a continuous trend of refilling was lacking). 613

614

Figure 4: Visualization of the refilling process of air-filled (A-C) fibers, (D-E) pith, and (F) 615

vessels in an excised L. nobilis stem during rehydration (at t =0h, circle symbols in Figures 4A 616

to 4C). Representative microCT images show water- and air-filled tissue in light and dark gray 617

color, respectively. (A) Enlarged transverse images show adjacent air-filled fibers, and many of 618

these fibers refilled over time (examples indicated by black arrows). (B, C) Corresponding 619

longitudinal images show the expansion of water columns (direction of movement indicated by 620

white arrows) inside the air-filled lumen; fibers labelled ‘a-d’ and ‘e-j’ were positioned along 621



27

dashed lines ‘B’ and ‘C ’in panel A, respectively. Liquid/air menisci with the fiber wall were 622

concave-shaped. (D) Longitudinal images show the progression of pith refilling from periphery 623

towards stem center (black arrows indicate examples of refilled cells). (E) Corresponding 3-D 624

volume rendering for an enlarged portion of the pith (position indicated by white box in panel D) 625

visualizing cell ‘a’ during refilling (dashed line indicates the cell wall). The liquid/air meniscus 626

with the cell wall was concave-shaped. (F) Longitudinal images show refilling of an air-filled 627

vessel. Water columns formed and expanded inside the air-filled lumen (direction of movement 628

indicated by white arrows); black triangles indicate droplets forming on the lateral vessel wall. 629

(f=fiber, Pi=pith, V=vessels,) 630

631

Figure 5: (A, B) Transverse microCT images visualizing air-filled tissues in the stem of an intact 632

L. nobilis sapling following soil saturation (at t=0.h) and (C) following stem excision (length 633

~5cm) and rehydration for 6 hours. For better orientation, examples of the same air-filled fibers 634

and vessels are labelled in red and yellow color, respectively, and air-filled pith tissue is labelled 635

in orange color. Values in hours (h) is time following soil saturation; values in mm2 are air-filled 636

cross-sectional areas. 637

638

Figure 6: Temporal refilling dynamics of air-filled fibers, pith tissue and vessels in the stem of 639

intact L. nobilis saplings. During the time period of investigation, the soil was fully saturated for 640

all saplings. The shoot of saplings was either exposed to (A-C) ambient conditions or (D-F) 641

covered in petroleum jelly and a humid plastic bag. Stem water potential (open symbols in panel 642

A and D) of saplings was monitored periodically; values in legend are initial stem water 643

potentials of saplings. As for Figure 3, dashed lines provide an estimate of tissue-specific 644

refilling dynamics and were obtained from nonlinear regression analysis across data points of all 645

samples (y=a∙e(b/(x+c); A, a=98, b=4E-11, c=1, R2=-1.64E-11, P=1; B, a=47, b=2156, c=2953, 646

R2=0.58, P=0.005; C, a=0.73, b=1002, c=201, R2=0.63, P=0.01; D, a=72, b=8, c=24, R2=0.63, 647

P=0.02; E, a=80, b=1, c=5, R2=0.80, P=0.001; F, a=12, b=421, c=195, R2=0.66, P=0.01). 648

649

Figure 7: Characterization of tissue viability and anatomical features in L. nobilis stems. (A-C) 650

Fluorescence light microscopy images stained with FDA (green signal inside viable tissue) and 651

PI (red signal in non-viable apoplast) solutions. Corresponding enlarged images (location 652

indicated by white box in panel A) show that (B) fibers surrounding vessels were dead and that 653

only xylem ray parenchyma and (C) pith cells located in close proximity to xylem were living; 654

pith cells located further inward were dead but maintained an intact cell wall. (D-E) ESEM 655



28

images show pit connections (indicated by triangles) between fibers and between vessels. (F) 656

MicroCT longitudinal images show that adjacent fibers were arranged in radial rows; examples 657

of fiber lumen are reconstructed in 3-D and labelled in red color. Panels 1 and 2 (position 658

indicated by yellow box in panel F) show enlarged images of 3-D volume renderings. Fiber-to-659

fiber, vessel-to-fiber and vessel-to-vessel pits in xylem (panel 1) and cell-to-cell connections in 660

pith (panel 2) are clearly visible; examples of pits and cell-to-cell connections are indicated by 661

black triangles. Examples of granules in pith cells are indicated by arrow (f=fiber, RP=ray 662

parenchyma, pi=pith, V=vessel). 663

664

665

666



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http://www.ncbi.nlm.nih.gov/pubmed?term=Brodersen CR%2C McElrone AJ %282013%29 Maintenance of xylem network transport capacity%3A a review of embolism repair in vascular plants%2E Frontiers in Plant Science 4%2C 1%2D11%2E&dopt=abstract

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http://www.ncbi.nlm.nih.gov/pubmed?term=Krasnow M%2C Matthews M%2C Shackel K %282008%29 Evidence for substantial maintenance of membrane integrity and cell viability in normally developing grape %28Vitis vinifera L%2E%29 berries throughout development%2E Journal of Experimental Botany 59%2C 849%2D859%2E&dopt=abstract

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Meinzer FC, Johnson DM, Lachenbruch B, McCulloh KA, Woodruff DR (2009) Xylem hydraulic safety margins in woody plants:coordination of stomatal control of xylem tension with hydraulic capacitance. Functional Ecology 23, 922-930.


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Nardini A, Savi T, Losso A, Petit G, Pacile S, Tromba G, Mayr S, Trifilo P, Lo Gullo MA, Salleo S (2016). X-ray microtomographyobservations of xylem embolism in stems of Laurus nobilis are consistent with hydraulic measurements of percentage loss ofconductance. New Phytologist 213, 1068-1075.


Plavcová L, Jansen S (2015) The Role of Xylem Parenchyma in the Storage and Utilization of Nonstructural Carbohydrates. In: Hacke U.(eds) Functional and Ecological Xylem Anatomy. Springer, Cham


Pratt and Jacobsen (2016) Conflicting demands on angiosperm xylem: Tradeoffs among storage, transport and biomechanics. PlantCell and Environment 40, 897-913.


Rhizopoulou S, Mitrakos K (1990) Water relations of evergreen sclerophylls. I. seasonal changes in the water relations of elevenspecies from the same environment. Annals of Botany 65, 171-178.


Salleo S, Lo Gullo MA, De Paoli D, Zippo M (1996) Xylem recovery from cavitation-induced embolism in young plants of Laurus nobilis: apossible mechanism. New Phytologist 132, 47-56.


Schweingruber FH, Boerner A, Schulze ED (2011) Atlas of stem anatomy in herbs, shrubs and trees. Volume 1. Heidelberg DordrechtLondon New York. Springer Verlag.


Siau JF (1984) Transport processes in wood. Springer-Verlag. Berlin Heidelberg New York Tokyo.Pubmed: Author and TitleCrossRef: Author and TitleGoogle Scholar: Author Only Title Only Author and Title

Suuronen J-P, Peura M, Fagerstedt K, Serimaa R (2013) Visualizing water-filled versus embolized status of xylem conduits by desktopx-ray microtomography. Plant Methods 9, 1-13.


Trifilo P, Raimondo F, Lo Gullo MA, Barbera PM, Salleo S, Nardini A (2014) Relax and refill: xylem rehydration prior to hydraulicmeasurements favours embolism repair in stems and generates artificially low PLC values. Plant, Cell and Environment 37, 2491-2499.


Tyree MT, Yang S (1990) Water-storage capacity of Thuja, Tsuga and Acer stems measured by dehydration isotherms. Planta 182, 420-426.


Tyree MT, Salleo S, Nardini A, Mosca R, Lo Gullo M, Mosca R (1999) Refilling of embolized vessels in young stems of laurel: do weneed a new paradigm? Plant Physiology 120, 11-22.

Pubmed: Author and Title www.plantphysiol.orgon August 13, 2019 - Published by Downloaded from Copyright © 2017 American Society of Plant Biologists. All rights reserved.

http://www.ncbi.nlm.nih.gov/pubmed?term=Meinzer FC%2C Johnson DM%2C Lachenbruch B%2C McCulloh KA%2C Woodruff DR %282009%29 Xylem hydraulic safety margins in woody plants%3A coordination of stomatal control of xylem tension with hydraulic capacitance%2E Functional Ecology 23%2C 922%2D930%2E&dopt=abstract

http://search.crossref.org/?page=1&rows=2&q=Meinzer FC, Johnson DM, Lachenbruch B, McCulloh KA, Woodruff DR (2009) Xylem hydraulic safety margins in woody plants: coordination of stomatal control of xylem tension with hydraulic capacitance. Functional Ecology 23, 922-930.

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http://search.crossref.org/?page=1&rows=2&q=Nardini A, Lo Gullo MA, Tracanelli S (1996) Water relations of six scerophylls growing near Trieste (Northeastern Italy): has scelrophylly a univocal functional significance? Giornale botanica intaliano 130, 811-828.

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http://search.crossref.org/?page=1&rows=2&q=Nardini A, Savi T, Losso A, Petit G, Pacile S, Tromba G, Mayr S, Trifilo P, Lo Gullo MA, Salleo S (2016). X-ray microtomography observations of xylem embolism in stems of Laurus nobilis are consistent with hydraulic measurements of percentage loss of conductance. New Phytologist 213, 1068-1075.

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http://search.crossref.org/?page=1&rows=2&q=Plavcov� L, Jansen S (2015) The Role of Xylem Parenchyma in the Storage and Utilization of Nonstructural Carbohydrates. In: Hacke U. (eds) Functional and Ecological Xylem Anatomy. Springer, Cham

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http://search.crossref.org/?page=1&rows=2&q=Pratt and Jacobsen (2016) Conflicting demands on angiosperm xylem: Tradeoffs among storage, transport and biomechanics. Plant Cell and Environment 40, 897-913.

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http://search.crossref.org/?page=1&rows=2&q=Rhizopoulou S, Mitrakos K (1990) Water relations of evergreen sclerophylls. I. seasonal changes in the water relations of eleven species from the same environment. Annals of Botany 65, 171-178.

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http://www.ncbi.nlm.nih.gov/pubmed?term=Salleo S%2C Lo Gullo MA%2C De Paoli D%2C Zippo M %281996%29 Xylem recovery from cavitation%2Dinduced embolism in young plants of Laurus nobilis%3A a possible mechanism%2E New Phytologist 132%2C 47%2D56%2E&dopt=abstract

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http://www.ncbi.nlm.nih.gov/pubmed?term=Siau JF %281984%29 Transport processes in wood%2E Springer%2DVerlag%2E Berlin Heidelberg New York Tokyo%2E&dopt=abstract

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http://www.ncbi.nlm.nih.gov/pubmed?term=Suuronen J%2DP%2C Peura M%2C Fagerstedt K%2C Serimaa R %282013%29 Visualizing water%2Dfilled versus embolized status of xylem conduits by desktop x%2Dray microtomography%2E Plant Methods 9%2C 1%2D13%2E&dopt=abstract

http://search.crossref.org/?page=1&rows=2&q=Suuronen J-P, Peura M, Fagerstedt K, Serimaa R (2013) Visualizing water-filled versus embolized status of xylem conduits by desktop x-ray microtomography. Plant Methods 9, 1-13.

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http://scholar.google.com/scholar?as_q=Visualizing water-filled versus embolized status of xylem conduits by desktop x-ray microtomography&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

http://scholar.google.com/scholar?as_q=Visualizing water-filled versus embolized status of xylem conduits by desktop x-ray microtomography&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Suuronen&as_ylo=2013&as_allsubj=all&hl=en&c2coff=1

http://www.ncbi.nlm.nih.gov/pubmed?term=Trifilo P%2C Raimondo F%2C Lo Gullo MA%2C Barbera PM%2C Salleo S%2C Nardini A %282014%29 Relax and refill%3A xylem rehydration prior to hydraulic measurements favours embolism repair in stems and generates artificially low PLC values%2E Plant%2C Cell and Environment 37%2C 2491%2D2499%2E&dopt=abstract

http://search.crossref.org/?page=1&rows=2&q=Trifilo P, Raimondo F, Lo Gullo MA, Barbera PM, Salleo S, Nardini A (2014) Relax and refill: xylem rehydration prior to hydraulic measurements favours embolism repair in stems and generates artificially low PLC values. Plant, Cell and Environment 37, 2491-2499.

http://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Trifilo&hl=en&c2coff=1

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http://www.ncbi.nlm.nih.gov/pubmed?term=Tyree MT%2C Yang S %281990%29 Water%2Dstorage capacity of Thuja%2C Tsuga and Acer stems measured by dehydration isotherms%2E Planta 182%2C 420%2D426%2E&dopt=abstract

http://search.crossref.org/?page=1&rows=2&q=Tyree MT, Yang S (1990) Water-storage capacity of Thuja, Tsuga and Acer stems measured by dehydration isotherms. Planta 182, 420-426.

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http://scholar.google.com/scholar?as_q=Water-storage capacity of Thuja, Tsuga and Acer stems measured by dehydration isotherms&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

http://scholar.google.com/scholar?as_q=Water-storage capacity of Thuja, Tsuga and Acer stems measured by dehydration isotherms&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Tyree&as_ylo=1990&as_allsubj=all&hl=en&c2coff=1

http://www.ncbi.nlm.nih.gov/pubmed?term=Tyree MT%2C Salleo S%2C Nardini A%2C Mosca R%2C Lo Gullo M%2C Mosca R %281999%29 Refilling of embolized vessels in young stems of laurel%3A do we need a new paradigm%3F Plant Physiology 120%2C 11%2D22%2E&dopt=abstract


CrossRef: Author and TitleGoogle Scholar: Author Only Title Only Author and Title

Tyree MT, Sperry JS (1989) Vulnerability of xylem to cavitation and embolism. Annual Review of Plant Physiology and Plant MolecularBiology 40, 19-38.


Verbeeck H, Steppe K, Nadezhdina N, Op de Beeck M, Deckmyn G, Meiresonne L, Lemeur R, Cermak J, Ceulemans R, Janssens IA(2007). Stored water use and transpiration in Scots pine: a modeling analysis with ANAFORE. Tree Physiology 27, 1671-1685.


Waring RH, Whitehead D, Jarvis PG (1979) The contribution of stored water to transpiration in Scots pine. Plant Cell and Environment2, 309-317.



http://search.crossref.org/?page=1&rows=2&q=Tyree MT, Salleo S, Nardini A, Mosca R, Lo Gullo M, Mosca R (1999) Refilling of embolized vessels in young stems of laurel: do we need a new paradigm? Plant Physiology 120, 11-22.


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http://www.ncbi.nlm.nih.gov/pubmed?term=Tyree MT%2C Sperry JS %281989%29 Vulnerability of xylem to cavitation and embolism%2E Annual Review of Plant Physiology and Plant Molecular Biology 40%2C 19%2D38%2E&dopt=abstract

http://search.crossref.org/?page=1&rows=2&q=Tyree MT, Sperry JS (1989) Vulnerability of xylem to cavitation and embolism. Annual Review of Plant Physiology and Plant Molecular Biology 40, 19-38.


http://scholar.google.com/scholar?as_q=Vulnerability of xylem to cavitation and embolism&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

http://scholar.google.com/scholar?as_q=Vulnerability of xylem to cavitation and embolism&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Tyree&as_ylo=1989&as_allsubj=all&hl=en&c2coff=1

http://www.ncbi.nlm.nih.gov/pubmed?term=Verbeeck H%2C Steppe K%2C Nadezhdina N%2C Op de Beeck M%2C Deckmyn G%2C Meiresonne L%2C Lemeur R%2C Cermak J%2C Ceulemans R%2C Janssens IA %282007%29%2E Stored water use and transpiration in Scots pine%3A a modeling analysis with ANAFORE%2E Tree Physiology 27%2C 1671%2D1685%2E&dopt=abstract

http://search.crossref.org/?page=1&rows=2&q=Verbeeck H, Steppe K, Nadezhdina N, Op de Beeck M, Deckmyn G, Meiresonne L, Lemeur R, Cermak J, Ceulemans R, Janssens IA (2007). Stored water use and transpiration in Scots pine: a modeling analysis with ANAFORE. Tree Physiology 27, 1671-1685.

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http://scholar.google.com/scholar?as_q=Stored water use and transpiration in Scots pine: a modeling analysis with ANAFORE&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Verbeeck&as_ylo=2007&as_allsubj=all&hl=en&c2coff=1

http://www.ncbi.nlm.nih.gov/pubmed?term=Waring RH%2C Whitehead D%2C Jarvis PG %281979%29 The contribution of stored water to transpiration in Scots pine%2E Plant Cell and Environment 2%2C 309%2D317%2E&dopt=abstract

http://search.crossref.org/?page=1&rows=2&q=Waring RH, Whitehead D, Jarvis PG (1979) The contribution of stored water to transpiration in Scots pine. Plant Cell and Environment 2, 309-317.

http://scholar.google.com/scholar?as_q=&num=10&as_occt=any&as_sauthors=Waring&hl=en&c2coff=1

http://scholar.google.com/scholar?as_q=The contribution of stored water to transpiration in Scots pine&num=10&btnG=Search+Scholar&as_epq=&as_oq=&as_eq=&as_occt=any&as_publication=&as_yhi=&as_allsubj=all&hl=en&lr=&c2coff=1

http://scholar.google.com/scholar?as_q=The contribution of stored water to transpiration in Scots pine&num=10&btnG=Search+Scholar&as_occt=any&as_sauthors=Waring&as_ylo=1979&as_allsubj=all&hl=en&c2coff=1


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