Rheology and Conductivity of Phloem

RHEOLOGY AND CONDUCTIVITY OF PHLOEM

SIERRA DAWN BEECHER

A dissertation submitted in partial fulfillment of the requirements for the degree of

DOCTOR OF PHILOSOPHY

WASHINGTON STATE UNIVERSITY Molecular Plant Sciences

JULY 2016

To the Faculty of Washington State University:

The members of the Committee appointed to examine the dissertation of SIERRA DAWN BEECHER find it satisfactory and recommend that it be accepted.

______Michael Knoblauch, Ph.D. Chair

______Asaph Cousins, Ph.D.

______Andrei Smertenko, Ph.D.

______Eric Shelden, Ph.D.

Acknowledgements

I have been very fortunate to have had a lot of excellent support with these projects. My advisor, Michael Knoblauch helped me design experiments to address the questions I was most curious about, and generously made available all tools and technology to carry them out. Dan

Froelich shared many of the techniques he had developed for in vivo confocal imaging, which were the foundation for the methods that I developed to do cold stimulation experiments. Dan

Mullendore graciously shared many insights, and technical tricks about new possibilities presented by the Leica SP8. These were used for fluorescence lifetime imaging to measure the viscosity of translocating phloem sap. Tim Ross-Elliot was always generous with his methods, and helped me develop better molecular biology skills. All members of the Knoblauch Lab, and my committee members, Eric Shelden, Andrei Smertenko, and Asaph Cousins have inspired me with their passion and intelligence. Michael Neff has also been very supportive and helpful. I would like to thank Valerie Lynch-Holm and Chris Davitt, whose kindness and incredible expertise in all aspects of imaging have been very valuable. Chuck Cody’s wealth of taxonomic knowledge and magical ability to keep plants alive has been very much appreciated. Kåre

Hartvig Jensen has been a constant collaborator, and his excellent skills in math and physics were crucial assets for this work. Jessica Savage provided all of the plant materials for analyses in Chapter 4, even climbing trees to do so. My husband, Horst Onken has provided a steady stream of intellectual insight and friendship, which I could not have done without. My parents,

Ron and Cookie Beecher, and their spouses have helped me in more ways than I can count, and my children, Emily and Kevin Beecher have been a wellspring of joy and support.

iii

RHEOLOGY AND CONDUCTIVITY OF PHLOEM

Abstract

by Sierra Dawn Beecher, Ph.D. Washington State University July 2016

Chair: Michael Knoblauch

The Pressure Flow Hypothesis, presented in 1930 by Ernst Münch, is the most accepted model for photoassimilate translocation by the phloem of plants. Net photosynthetic sugar production occurs in source tissues, often mature leaves. These photoassimilates are transported along the sieve tube system to consumptive sink tissues such as roots, meristems, and seeds. Münch’s hypothesis states that a sugar concentration gradient between sources and sinks osmotically generates a hydrostatic pressure differential that drives sap flow. If phloem sap is driven by a pressure differential, geometries of the sieve tube system, and the rheology of phloem sap should scale to equations describing pressure flow in low Reynolds number tubes. Imaging methods developed in this work provide robust geometric and rheological data, expanding the phloem biophysics toolkit. Additionally, insight into a mysterious phenomenon in which transient inhibition of translocation is induced by sudden localized stem chilling is presented. A novel application of fluorescence recovery after photobleaching (FRAP) allowed simultaneous measurement of sap velocity, and diffusivity of a phloem-mobile probe. This technique was used to investigate the cold shock phenomenon directly at the site of cold stimulus at higher resolution than was previously possible. Cold-induced sap velocity declines were not accompanied by similar reductions in sap viscosity. A Fluorescence Lifetime Imaging Microscopy

(FLIM) method was also developed, providing direct in vivo measurements of viscosity in translocating phloem sap. Values obtained indicate phloem sap viscosities of less than 2 mPas, consistent with sugar concentrations estimated by previous time-intensive exudate analyses.

Questions regarding pressure flow in trees, where tube lengths are very long and source phloem turgor pressures have been measured as surprisingly low were also addressed. Sieve element lengths, diameters, and sieve plate angles were measured in 3 positions of 22 different angiosperms. These conductivity factors were large, and scaled to stem length in trees. Because pressure flow equations indicate an inversely proportional relationship between conductivity and pressure, these data are consistent with pressure driven flow as the mechanism for translocation in trees.

TABLE OF CONTENTS

Page

Acknowledgments...... iii

Abstract ...... iv

List of Tables ...... ix

List of Figures ...... x

Chapter 1: Introduction to Phloem ...... 1

Overview ...... 1

Evolution of Translocation ...... 1

Development of Angiosperm Phloem...... 6

Structure of Angiosperm Phloem ...... 9

Sieve Elements ...... 9

Sieve Element Organelles and Subcellular Structures ...... 12

Sieve Element Plastids ...... 12

Sieve Element Mitochondria ...... 12

Sieve Element Endoplasmic Reticulum ...... 12

Sieve Element Proteins ...... 13

Sieve Element Plasmalemma ...... 14

Sieve Element Cell Walls and Sieve Plate Pores ...... 15

Companion Cells ...... 16

Phloem Parenchyma ...... 17

Phloem Fibers ...... 17

Function of the Phloem ...... 18

Phloem Loading...... 21

Passive Symplastic Phloem Loading ...... 23

Active Symplastic Phloem Loading ...... 24

Apoplastic Phloem Loading ...... 24

Transport Phloem ...... 25

Unloading Phloem ...... 27

Rheological Considerations ...... 27

Problems with the Pressure Flow Hypothesis ...... 28

Works Cited ...... 30

Chapter 2: Diffusion and Velocity ...... 36

Abstract ...... 36

Introduction ...... 36

Materials and Methods ...... 38

Results ...... 39

Discussion...... 46

Works Cited ...... 51

Chapter 3: Phloem Sap Viscosity ...... 53

Abstract ...... 53

Introduction ...... 53

Materials and Methods ...... 60

Preparation of Standard Solutions ...... 60

Imaging Standard Solutions ...... 60

Plants ...... 61

Analysis ...... 61

Results ...... 62

Viscosity Effect of Lifetime on 2-NBDG ...... 62

vii

pH and Ionic Strength Effects ...... 63

Effect of soluble protein ...... 66

Intensity and Lifetime Imaging of Living Phloem ...... 66

Discussion...... 68

Works Cited ...... 75

Chapter 4: Angiosperm Sieve Tube Conductivity ...... 79

Abstract ...... 79

Introduction ...... 79

Materials and Methods ...... 83

Sample Collection ...... 83

Sample Preparation ...... 83

Imaging ...... 85

Results ...... 86

Discussion...... 99

Works Cited ...... 104

viii

LIST OF TABLES

Chapter 4.

Table 1: Geometric sieve element data ...... 87

Table 2: Scientific names of species, and total plant height ...... 89

LIST OF FIGURES

Chapter 1.

Figure 1: Phloem evolution ...... 5

Figure 2: Secondary vasculature in the eudicot Catalpa speciosa ...... 7

Figure 3: Leaf vasculature of the monocot Phalaris arundinacea ...... 8

Figure 4: Sieve tube anatomy ...... 11

Figure 5: Protein bodies in sieve elements ...... 14

Figure 6: Sieve plates of Quercus Rubra ...... 16

Figure 7: Arabidopsis thaliana companion cells and sieve elements ...... 17

Figure 8: Schematic of pressure flow mechanism for phloem translocation ...... 20

Figure 9: Schematic of phloem loading strategies ...... 22

Figure 10: H+-ATPase driven phloem loading schematic ...... 25

Chapter 2.

Figure 1. In situ flow velocity measurements ...... 40

Figure 2: In situ phloem sap viscosity analysis ...... 41

Figure 3: A plot of v3 and dkτ−2 indicates a linear relationship ...... 43

Figure 4: Phloem flow velocity change after cold shock ...... 44

Figure 5: v3 versus dkτ−2 ...... 45

Figure 6: Schematic drawing of the development of a bleach profile ...... 47

Chapter 3

Figure 1: Fluorescence lifetime of 2-NBDG is a function of solvent viscosity ...... 63

Figure 2: Low pH of solvent has a slight effect on lifetime of 2-NBDG ...... 64

Figure 3: Lifetimes of 2-NBDG in 0.5 to 3 M of NaCl in PBS ...... 65

Figure 4: Lifetimes of 2-NBDG in sucrose solutions containing 0.1% BSA ...... 66

Figure 5: 2-NBDG intensity and lifetime maps in the sap of living plants ...... 67

Figure 6: Relationship between osmotic potential and viscosity ...... 72

Chapter 4.

Figure 1: Schematic of sample preparation ...... 84

Figure 2: Imaging examples ...... 86

Figure 3: Correlation between sieve element diameter and distance from apex...... 91

Figure 4: Diameter and length of sieve elements increases with length in trees ...... 92

Figure 5: The angle of sieve plates increases with the length of the cell ...... 93

Figure 6: Sieve element measurements from the tree Quercus rubrus ...... 94

Figure 7: Sieve element diameter was not a function of length in vines ...... 95

Figure 8: Diameters and lengths of sieve elements in Solanum lycopersicum...... 96

Figure 9: Sieve element measurements from Solanum lycopersicum ...... 97

Figure 10: Sieve element length to diameter ratios ...... 98

Figure 11: Relationship between stem length and stem diameter ...... 99

Figure 12: Proposed model for sieve tube conductivity in trees ...... 102

Chapter 1: Introduction to Phloem

Overview

The biosphere is almost completely dependent on solar energy. A very significant part of terrestrial biological energy flow involves the vascular system of plants. Photosynthesis fixes atmospheric carbon dioxide into sugars, transforming radiant energy into storable chemical energy, which enters the food web powering biodiversity. The importance of plant physiology for sustaining terrestrial life just begins at photosynthesis, however. Some of the sugars synthesized in source tissues of plants must be transported to non-photosynthetic, energy- demanding plant organs, such as roots, tubers, flowers, and fruits. Many of the plant parts that draw photosynthates from leaves are directly responsible for the ability of other heterotrophic life processes to continue when photosynthetic biomass accumulation is suppressed by seasonality. For example, current human civilizations are completely dependent on storable seeds that include rice, wheat, and corn. The transport process that facilitates the movement of energy from sources to sinks in vascular plants is called translocation, and is accomplished by the phloem.

Evolution of translocation

Oxygenic photosynthesis evolved soon after the cooling of the earth in the Archean. Oxygen began to accumulate in the atmosphere about 3.2 billion years ago, and fossil stromatolites built from cyanobacteria can be dated to this Eon (Blankenship 2010). The tiny cells that formed these structures are independent, and each cell is an autonomous organism with little metabolic need of its neighbors. The evolution of multicellularity eventually resulted in a

1 division of labor, wherein cells became specialized for certain tasks (Nedelcu 2001;

Schirrmeister et al. 2013). This division of labor presented the selective pressure that resulted in the evolution of nutrient transport systems in multicellular plants (Van Bel 2000).

Even before the evolution of green plants, this division of labor was already taking root in photosynthetic bacteria as Earth’s atmosphere changed as a result of their activity. For example, in some freshwater filamentous cyanobacteria, low environmental levels of reduced nitrogen stimulate the formation of heterocysts at regular intervals of about 15 cells. The fixation of nitrogen that occurs in these differentiated cells must be separated from oxygenic photosynthesis, as oxygen irreversibly inhibits the enzyme complex nitrogenase. Also, photosynthesis is suppressed in heterocysts, but ATP is still needed to drive nitrogenase activity and for the cell’s metabolism (Berman-Frank et al. 2003). Anabaena cylindrica, a heterocystic filamentous cyanobacteria exhibits high rates of exchange of molecules between vegetative cells and heterocysts, and exchange rates become more rapid after heterocyst formation. The non-heterocyst forming filamentous Oscillatoria terebriformis does not exhibit this high rate of intercellular molecular exchange, probably because the lack of labor division has not presented a selective pressure for the production of channel proteins that allow it in A. cylindrica

(Mullineaux et al. 2008).

As organisms become larger, simple diffusion, such as described for the filamentous cyanobacteria, is no longer sufficient to maintain communication and nutrient exchange.

Diffusion is not even sufficient in single cells if they become very large. Some of the largest plant cells known belong to the land plant predecessor, Chara. Cytoplasmic streaming through the activity of actin and myosin in the giant cells of Chara improves nutrient exchange both

2 intracellularly, and intercellulary, moving cellular cargo at increased rates, and generating a current. Arabidopsis transformants that express chimeric “high speed” myosins derived from

Chara grow larger, and produce seeds sooner, indicating that this system is directly tied to fitness (Tominaga and Ito 2015). Intercellular translocation in Chara is driven energetically by cytoplasmic streaming, and not bulk flow, as demonstrated by the observations that 14C does not travel cell-to-cell faster than the rate of streaming, and that Cytochalasin B inhibits the movement of radiolabeled carbon both within individual cells, and between them (Box et al.

1984).

Intercellular connectivity similar to that presented by A. cylindrica has convergently evolved in many organisms, but the angiosperms are unique in their possession of diverse and plastic plasmodesmata (Brunkard et al. 2015). Similar, and presumably evolutionarily precursory structures are encountered in Chara (Cook et al. 1997). Bryophytes (Sheirer 1990), pteridophytes (Evert 1990), and conifers (Shulz, 1990) have more developed plasmodesmal structures. Plasmodesmata of terrestrial plants not only connect adjacent cells, rendering their cytoplasm communal, but also present regulated cell-to-cell communication (Lucas and Lee

2004). They are formed during telophase, when parts of the endoplasmic reticulum become trapped in the newly forming cell wall of the daughter cells, or de-novo during cell expansion.

The ability to control the fluxes of molecules through plasmodesmata, and “turn them off” via constriction is a crucial element to plant development and organogenesis (Fitzgibbon et al.

2013). Plasmodesmata are especially important in this work for their roles in long-distance transport, and phloem loading in angiosperms.

Cytoplasmic streaming, while rapid and efficient in aquatic plants, is energy consumptive, and therefore not the best means for long distance transport in organisms that are large, highly differentiated, limited by water, and heavily invested in storage tissue. In land plants, a new transport strategy based on bulk flow begins to emerge.

In the ancient bryophytes, beginnings of a higher level transport system can be seen in the elongated structure of rhizoids. Also, an isolated stele is present in the seta and the stem that contains hydroids for water transport, and leptoids, (elongated, connected cells) that transport sugars (Figure 1A). Leptoids have many morphological similarities to sieve elements of angiosperms, but they differ in development. They do not undergo complete nuclear degeneration, have less developed end wall connections, and lack companion cells. They are very likely the evolutionary precursors to sieve elements (Schierer 1990).

It was not until the evolution of robust vascular systems and the molecule lignin that plants were able to vastly increase their size and range in terrestrial ecosystems. In the late Devonian,

Lycopodium ancestors, the first vascular plants, dominated terrestrial ecosystems.

Lepidodendron trees displayed secondary growth, producing woody water conducting tissue, but had a unifacial cambium (Seward 1902; Eggert and Kanemoto 1977). Therefore,

Lepidodendron did not make secondary phloem, yet still grew very tall. It was hypothesized that they were not deeply rooted, and that photosynthetic shoots supported roots metabolically.

They grew very fast, via short life cycles due in part to the lack of secondary phloem, and the energy storage capacity it imparts. Today, only three clades of Lycopodium persist. All are less than 1m tall, lacking secondary growth.

Ferns have more developed vascular systems. Siphonosteles are organized into strands of amphivasal bundles (Figure 1B). Although still dependent on freshwater ecosystems for reproduction, the evolution of specialized vascular tissues and cell wall materials allowed lycopodium and ferns to gain size and expand their range during the late Devonian and

Carboniferous periods (Kenrick and Crane 1997). This biomass expansion increased terrestrial primary productivity, and had vast impacts on the structure of ecosystems and the composition of the atmosphere. The burial of these massive ferns and lycopods also resulted in the formation of the reserves of coal that we rely so heavily on to power our industrial lifestyles.

A B

E E

P H X L

Figure 1: Phloem evolution. A. Cross section of Polytrichum moss. Haplostele consists of a core of hydroids (H) surrounded by a ring of leptoids (L). Cuticle of epidermis (E) is stained red by

FCA. B. Vascular bundle of dictostele of fern. Amphicribral bundle is ringed with thick endodermis (E). Xylem (X) is in the core, ringed by phloem (P) Sieve areas are steeply angled

5 and labeled with arrows. Stained with calcofluoro white and aniline blue. Xylem, plastids, and lipids autofluoresce in the red. Scale bar is ~ 20 μm.

The cellular structure of fern vasculature is also more similar to angiosperms than that of bryophytes and lycopsids. The phloem is abundant, but differs structurally and developmentally, and is less complex (Evert 1990). During the dry Permian period, gymnosperms came to dominance in ecosystems, because of the evolution of seeds and other special adaptations to aridity. They still dominate in the Taiga, and in some other water-limited ecosystems. Gymnosperm translocation is accomplished by sieve cells with small diameters and large steeply angled sieve areas. The density of venation, and therefore the photosynthetic capacity of angiosperms is far greater than moss, pteridophytes, and gymnosperms, and the steles and bundles are more complex. The expansion of the density of venation in angiosperms during the Cretaceous is correlated with major changes in the earth’s biogeochemistry, including a marked reduction in the atmospheric concentration of CO2 (Field et al. 2011).

Bryophytes and one lineage of ferns were spared extinction by the newfound dominance of seed plants that was imparted by their advanced vasculature, because they possess genes that allow them to thrive in the shade (Li et al. 2014).

Development of angiosperm phloem In the developing seed, ground tissue of the embryo has already differentiated into continuous strands of procambium that extend from the cotyledons to the radicle (Esau 1964). At germination, these strands develop into the primary vasculature of the plant, form patterned venation structures, and begin to produce primary phloem and xylem (Scarpella et al. 2004).

This patterning is distinct in monocots and eudicots. In monocots, venation is often parallel in

6 leaves, and vascular bundles are scattered in stems. In eudicots, the leaf venation is usually reticulate, and in the stem, vascular bundles are organized into a ring. This difference is thought to involve an evolution of different responses to auxin in these two groups of angiosperms

(McSteen 2010). After the primary plant body is complete, in eudicots, the vascular cambium is initiated, and begins to lay down secondary vascular tissues. The secondary phloem usually is positioned external to the xylem in stems and often oriented abaxially in dicot leaves (Evert

2006). The presence of the vascular cambium in eudicots allows for secondary growth, which is not exhibited by monocots. (See Figures 2 and 3).

P X

Toward epidermis

Figure 2: Secondary vasculature in the eudicot Catalpa speciosa. P = phloem, VC = vascular cambium (between dashed white lines), X = xylem. Callose is stained bright red at sieve plates, and cellulose appears blue. Darker red is autofluorescence of lignin and plastids. The xylem is differentiating to the right of the VC, and the phloem to the left. Scale bar is 20 μm.

X P

Figure 3: Leaf vasculature of the monocot Phalaris arundinacea. P = phloem, X = xylem.

Vasculature of monocots lacks vascular cambium.

Angiosperm sieve elements are formed by the unequal anticlinal division of a mother cell that is produced by the procambium, or vascular cambium. The division of the mother cell results in a juvenile sieve element, and a companion cell. The sieve element undergoes nuclear degeneration and loses most of its contents, becoming metabolically dependent on, and forming a unit with its companion cell, to which it is tightly connected by plasmodesmata at

8 maturity. This partial autophagy clears the sieve element for translocation. The sieve plates become perforated by enlargement and modification of plasmodesmata at end walls of the sieve elements, which connect them end to end to each other, forming a microfluidic network throughout the plant (van Bel and Knoblauch 2000).

Structure of angiosperm phloem

Sieve elements

The angiosperm sieve tube system is evolved exquisitely for transport. Elongated cells called sieve elements are the building blocks of a continuous syncytium of elongated cells that form a tubular intracellular network between sources and sinks. The sieve elements are connected end to end by perforated cell walls, called sieve plates (Figure 4A and 4B). Because nuclei, tonoplasts, and most other organelles are degraded during sieve element development, they are metabolically supported by their tightly associated companion cells. Sieve elements maintain some mitochondria, unique plastids, and endoplasmic reticulum that is not associated with ribosomes. They often contain fibrous proteins, mostly located on the periphery. These subcellular structures can be observed by transmission electron microscopy (Esau and

Cronshaw 1968; Evert 1990; Mullendore et al. 2010). Sieve elements of all dicots and many monocots also contain various phloem proteins, of which the P-proteins of cucurbits, and the forisomes of faboid legumes are most often discussed (Evert 2006). Sieve elements of angiosperms are distinguishable from other plant sieve tube members, such as the sieve cells of gymnosperms, by the structure of their perforated sieve plates on end walls. In angiosperms,

9 these perforations are highly dilated, reducing resistance of photoassimilate transport flow in this group of plants (Esau and Cheadle 1959).

Lateral sieve pores are also present on parietal sieve element walls. These are fields of plasmodesmata that are not as dilated as the perforations on sieve plates. They allow for solute continuity between adjacent sieve elements and companion cells (Figure 4A).

A SP B

B SE

LSP

SP SP

Figure 4: Sieve tube anatomy. A: longitudinal confocal image of sieve elements of Cucurbita maxima. SP=sieve plate. LSP lateral sieve pores. SE Sieve element. B: Sieve plate of Catalpa

11 speciosa. Counterstained with calcofluoro white for cellulose (blue), and aniline blue for callose (red).

Scale bars are 20 μm.

Sieve element organelles and subcellular structures

Sieve element plastids

The function of sieve element plastids is unknown. Behnke (1981) described two major groups:

S-type plastids that contain starch, and P-type plastids that contain protein. Within these two major groups an array of subgroups can be classified based on their size, shape, and contents.

Sieve element plastids are so distinct that they can even be used for cladistics (Behnke 1981).

Plastids in mature sieve elements contain crystals, starch grains, fibers, or unknown substances enclosed in a double membrane, and positioned laterally near the cell wall (Esau and Cronshaw

1968).

Sieve element mitochondria

Although most organelles are lost in sieve elements, mitochondria persist (Evert 2006). During the course of development, they morph from being very similar to mitochondria in parenchyma cells to taking on a distinct structure, wherein they are smaller, less electron dense, more spherical, and have very stacked and ordered christae (Esau and Cheadle 1962). Organelles, including mitochondria in sieve elements have been observed to be anchored to the sieve element plasmalemma by ~ 15nm branched structures referred to as clamps (Ehlers et al. 2000)

Sieve element endoplasmic reticulum

The endoplasmic reticulum (ER) of sieve elements is smooth, lacking ribosomes, and usually observed in a parietal position. The sieve element ER appears to be continuous with the ER of companion cells, passing through the plasmodesmata that connect them (Martens et al. 2006).

The ER transports lipids between the CC and the SE, and may be involved in controlling the diameter of the plasmodesmata. It is stacked in a very orderly and regular fashion, and is in close proximal association with parietal organelles (Esau and Cronshaw 1968).

Sieve element proteins

Sieve element proteins are found in all dicotyledons observed to date. They are either filamentous, like those in Arabidopsis (Froelich et al. 2011), or crystalline such as those observed in Phaseolus vulgaris and Populus tremuloides (Figure 5A and 5B). The forisomes present in the faboid legumes are crystalline (Knoblauch et al. 2003). Of sieve element proteins, only these forisomes have a known function. Forisomes undergo a reversible ATP-independent change in morphology upon sieve tube wounding, switching from a low volume to a high volume state, occluding sieve plates. It is confounding to consider the roles of fibrous phloem proteins that are present in phloem sap (Froelich et al. 2011). These bodies seem that they would increase resistance in the sieve tube, and reduce flow. Results obtained by Froelich et al.

(2011) do not, however, appear to support this intuition. Arabidopsis plants with reduced sieve element protein expression did not display significantly faster velocity profiles than did the wild-type (Froelich et al. 2011).

B A SE

Figure 5: Protein bodies in sieve elements. (SE) of P. tremuloides (A) and Tailed forisomes in sieve elements of P. vulgaris (B) are indicated with white triangles. Blue is cellulose signal from calcofluoro white. Sieve plates (SP) are stained red by analine blue where callose deposition has occurred. Red autofluorescence in (A) is plastids and protein bodies. Scale bars are 10 μm.

Sieve element plasmalemma

The plasmalemma of SEs lines the cell wall, and intercolates the cell wall at sieve pores, sieve areas, and at plasmodesmata that join it to CCs and phloem parenchyma (Evert 2006). Most of the sparse structural contents of sieve elements are in close proximity to the cell membrane on side walls. It was thought that few energetic pumps would be found in SEs, because low metabolic activity is expected of them. However, H+ ATPase driven solute transporters and tonoplast type pyrophosphatase have been discovered in SEs by immunolocalization. These active transporters may explain the retention of mitochondria in these cells, and probably function to retrieve leaked sucrose in transport phloem (Langhans et al. 2001). Some alkaloid

14 biosynthetic enzymes have been found associated with SE membranes as well (Bird et al. 2003).

Sieve element membranes are also dotted with aquaporins, which are highly conserved proteins that form water specific channels in all plant cells, including bryophytes (Fraysse et al.

2005).

Sieve element cell walls and sieve plate pores

The most striking feature of sieve elements are the previously discussed sieve plates on the end walls, and lateral sieve pore fields. Sieve elements deposit a substance called callose upon injury at these perforations, which can be stained with aniline blue and visualized with confocal microscopy (Figure 2A and 2B). The cell walls of sieve elements are usually not lignified, and must expand much more than the ground parenchyma to allow longitudinal growth. Although lacking secondary cell wall, in many species, sieve elements do deposit nacreous cell wall material, which is a morphologically distinct secondary thickening internal to the primary wall

(Esau and Cheadle 1958). This cell wall is heavily reticulated with tubular strands of plasmalemma. The nacreous layer usually becomes diminished as the SE matures in primary phloem, and does not always do so in secondary sieve elements (Evert and Mierzwa 1989).

The perforated cellulosic cell walls of the sieve plates, and of fields of lateral sieve pores in woody species are crossed by regular bandings of cellulose that appear similar to the perforation plates of xylem vessels (Figure 6)

SP  SE

Figure 6: Sieve plates of Quercus Rubra. Cellulosic banding can be seen across sieve plates. SE: sieve element, SP: sieve plate. Scale bar is ~20 μm.

Companion cells

Companion cells are ontogenetically related to sieve elements in metaphloem. They provide most of the metabolic support for the CC/SE complex. During development, companion cells retain organelles and dense cytoplasm. They maintain a tight connection to their sister sieve elements through plasmodesmata. This can be expoited for microscopic analysis, because the fluorescent dye 5,6 carboxyfluorescein can only leave sieve elements of transport phloem through plasmodesmata. Therefore it also labels companion cells. In Figure 7, this tight association can be observed. The companion cells are wrapped around the sieve elements. The vacuoles exclude CF, and can therefore be visualized, and distinguished from the SEs, as SEs do not have vacuoles and are homogenously labeled with the dye.

V CC

SE V

Figure 7: Arabidopsis thaliana companion cells and sieve elements. Seen here in transport phloem in juvenile roots. Labeled with 5,6 Carboxyfluorescein. CC: companion cell, SE: sieve element, V: vacuole. Scale bars are 100 μm.

Other Phloem Parenchyma Cells

Phloem parenchyma cells are associated with, but not always symplastically linked to sieve elements. Often, their cytoplasm is not as dense as that of companion cells, and their mitochondria are less spherical (Esau 1973). They provide pathogen defense in some species

(Franceschi et al. 1998). In areas of phloem loading and unloading, phloem parenchyma cells may differentiate into transfer cells. These cells take on a convoluted wall and membrane structure, and contain a dense cytoplasm with many mitochondria. It is proposed that these characteristics impart increased active transport capability. Their development can also be induced by environmental factors, including pathogens (Rodiuc et al. 2015). Phloem parenchyma are crucial for phloem loading in rice, Arabidopsis, and maize. Their cell membranes contain SWEET proteins, which actively efflux sucrose into the cell wall, making it available for influx transport to the CC/SE complex by SUC transporters (Chen et al. 2012).

Phloem fibers

Phloem fibers are cells that protect and stabilize the elongated cells of phloem. They form fiber caps that reduce the vulnerability of the sieve tubes. Fibers exhibit tip growth, and become very long with thick secondary walls. Sometimes their inner walls are gelatinous. Secondary phloem fibers form bast fiber strands which occupy the external stem of the plant, just beneath the epidermis. Their tendency to overlap into interconnected strands, and their thick, soft cell walls allow both flexibility and strength (Evert 2006). Aside from their structural and functional roles in plants, these traits present material properties that are excellent for making paper, rope, and textiles (Paridah et al. 2011).

Function of the phloem

The phloem is the route of transport of photoassimilate from the sources to the sinks of plants.

Sources are mature leaves where the rate of photosynthesis is higher than the rate of respiration, generating net production of sugars. Sinks are parts of the plant like flowers, fruit, seed and roots that must be supplied with energy. The phloem also transports amino acids, minerals, proteins, and signaling molecules such as flowering signals, hormones and small RNAs

(Ruiz-Medrano et al. 2001; Zhang et al. 2009). It has also been shown to facilitate conduction of electrical potential waves, similar to action potentials in the nervous system of animals

(Samejima and Sibaoka 1983; Fromm and Bauer 1994).

The flow of sap in the sieve tube system is hypothesized to be driven by an osmotically generated pressure differential constructed by a sugar concentration gradient between sources and sinks (Jensen et al. 2013). First proposed by Ernst Münch in 1930, the mechanism was described as follows: In sources, sugar production rates are higher than respiration. Therefore,

18 osmotically active solutes accumulate. When these energetic molecules enter the phloem in loading zones, the solute potential drops steeply. This results in a water potential gradient that favors the entry of water into the phloem. Cell walls are minimally elastic. Therefore a hydrostatic pressure is built by the osmotically-induced increase in volume that drives sap through the transport phloem towards unloading zones. Altogether, these events result in a pressure-driven flow from sources to sinks (Figure 8). Efforts to examine this hypothesis are becoming more robust as advancements in imaging and pressure sensing are allowing the parameters of equations for pressure driven flow in living plants to be observed (Knoblauch et al. 2016).

X P

CC MC

CC SC

Figure 8: Schematic of pressure flow mechanism for phloem translocation. Adapted from

Freeman (2010). X = xylem, P = phloem, MP = mesophyll cell, CC = companion cell, SE = sieve element, SC = sink cell. Osmotically active photosynthates constructed in MPs is transported into the CC/SE system, creating a hydrostatic pressure that drives sap to lower pressure regions, where photosynthates are delivered to sink cells (SC). Sinks are often storage tissues that heterotrophs are dependent on.

Only recently has it become possible to make measurements allowing the direct testing of the

Münch Hypothesis (Knoblauch et al. 2012; Savage et al. 2013; Windt et al. 2006). If a pressure differential is driving sap flow in plants, measurements should satisfy the Hagen-Poiseuille equation for pressure driven laminar flow in a low Reynolds number tube (Equation 1), and scale accordingly. Equation 2 has been modified for phloem to account for resistance induced by sieve plates and cell walls (Jensen et al. 2013).

ퟖ흁푳푸 ∆푷 = Equation 1 흅풓ퟒ

푼흁풍 ∆푷 = 풌 Equation 2

푈 = sap flow velocity (μm/s), 푘 = sieve tube conductivity (μm2), ∆P = pressure differential

(mPa), µ = sap viscosity (mPa*s), 푙 and 퐿 = length of tube – in the case of plants, length of plant

(μm), Q = volumetric flow rate (μm3/s), r = radius of tube (μm).

Phloem loading

Loading of sugars into the sieve tube system can occur by active or passive mechanisms. Active loading can be symplastic, involving a polymer trap mechanism for building and maintaining a concentration gradient between mesophyll and phloem; or apoplastic, requiring active pumping of solute into the CC/SE complex. Passive loading uses no pumping or polymer formation. Instead, large densities of plasmodesmata allow pure diffusion from sites of photosynthesis to load the phloem (Lische and Shultz 2012; Rennie and Turgeon 2009; Turgeon

2010; see Figure 9).

M CC SE D A

E B

C F

Figure 9: Schematic of phloem loading strategies. Adapted from Turgeon (2010). M = mesophyll, CC = companion cell, SE = Sieve element. In A (passive symplastic loading), a high degree of symplastic connection exists between companion cells and mesophyll. This results in passive diffusion of sugars as the phloem loading mechanism. The radioactive sucrose label is not localized to the vascular tissue, but instead the whole leaf has a similar concentration of sucrose, as depicted in D. B) describes the polymer trap phloem loading strategy. This is also a symplastic mechanism, but it is active. Sucrose enters companion cells by diffusion through plasmodesmata, but is used as substrate to construct oligosaccharides, such as raffinose, which

22 are too large to diffuse back into mesophyll. The radioactive signal is therefore concentrated in the vasculature (E). Active apoplastic loading, diagrammed in figure 9C, involves active transport of sucrose into the CC/SE complex (See Figure 10 for schematic of pump). In this case, radioactive sucrose signal is concentrated in the vasculature (F).

Passive symplastic phloem loading (Figures 9 A and D)

Passive symplastic phloem loading is assumed by the structure of the source phloem tissue, and the compositon of phloem sap. Plasmodesmatal frequencies between mesophyll, phloem parenchyma, and CC/SE complexes are high (Gamalei 1989). This is referred to as an open minor vein anatomy. The sucrose created by photosynthesis in mesophyll cells diffuses into phloem parenchyma, then into the CC/SE complexes. This does not allow a concentration gradient to develop in the minor veins of the leaves and a resultant high pressure in the loading phloem. The lack of sucrose concentration in veins is visualized using radiolabeled leaf discs.

Radiolabel is diffuse, and does not demarcate veins in plants that use this loading strategy.

Plants referred to as passive loaders do not transport large, modified sugars, and have high concentrations of sugars in their mesophyll (Rennie and Turgeon, 2009). Many of the plants that utilize this phloem loading strategy are trees. This has traditionally been puzzling, because the long path length of transport in trees would intuitively lead one to suspect a requirement for high pressure in source phloem. An apoplastic loading strategy would provide a higher pressure in source sieve tubes by pumping osmotically active photoassimilates against a concentration gradient, therefore constructing a higher hydrostatic pressure.

Active symplastic phloem loading (Figures 9 B and E)

Polymer trapping strategies, while also symplastic, are not passive. In this mechanism, diffusion also moves small sugars through plasmodesmata and into companion cells. Once inside the companion cells, ATP is needed for enzymatic synthesis of raffinose family oligosaccharides

(RFOs) from sucrose. These RFOs are too large to diffuse back through the plasmodesmata connecting CCs to mesophyll cells, and therefore are “trapped” in the CC/SE complex. There is a good correlation between plants that transport RFOs and possess an open minor vein configuration. Furthermore, transport is not inhibited by treatment with substances that disrupt active transport proteins (Turgeon 1996).

Apoplastic phloem loading (Figures 9 C and F)

An ATP-dependent concentrating step involving a carrier for phloem loading in many plants was first evidenced by experiments performed by Sovonick et al. (1974). The existence of these transporters was confirmed by Riesmeier et al. (1992) when they successfully cloned a sucrose carrier protein from spinach cDNA into a yeast expression system. The resultant transgenic yeast was able to import sucrose in a proton-dependent manner. Since then, an array of sucrose transporters has been discovered (Peng et al. 2014). In apoplastic phloem loading plants, sucrose is actively concentrated by loading the phloem at sources, using proton pumps and H+/Suc transporters (Figure 10). These transport proteins are concentrated in the source- adjacent membranes of companion cells, further strengthening the argument for their function in phloem loading. The sucrose SUC transporters import to CCs is made available to the apoplast from mesophyll cells by the activity of SWEET proteins that facilitate efflux (Chen et al.

2012). SWEET-mediated efflux may be controlled to limit pathogen access to translocating sucrose. The model plant Arabidopsis thaliana discussed in Chapters 2 and 3 utilizes this phloem loading strategy.

Figure 10: H+-ATPase driven phloem loading schematic. Adapted from Freeman 2010. This is the active step in the phloem sucrose concentrating process of apoplastic loaders. ATP is needed to pump H+ ions out of the cell. This creates an electrochemical gradient that favors re- entry of protons into the cell. They pass passively through symporters, allowing sucrose to travel against a concentration gradient into the CC/SE complex, building an osmotic potential.

Transport Phloem

Not all photosynthate is directly loaded into the phloem. Up to 50% of assimilated carbon is directed to starch storage in chloroplasts, where it is broken down into sugars at night and

25 delivered to the phloem (Zeeman et al. 2004). This aids in the maintenance of a relatively steady sap flow, although sap composition does fluctuate diurnally (Peuke et al. 2001; Windt et al. 2006).

The transport phloem is minimally conductive in the lateral direction. The connection between

CCs and SEs remains tight, but fluorescent dyes loaded into source tissues do not leave the microfluidic track of the sieve tubes past the companion cells, as can be observed in Figure 5. Its major function is to rapidly move solutes from sources to sinks where they are needed.

Mathematical modeling of angiosperm phloem supports the hypothesis that the system has evolved to be optimized for speed (Jensen et al. 2011). This is plausible, considering the dual function of phloem as both an energy delivery, and an information delivery system. Plants can be considered organisms that are both autotrophic, and heterotrophic. Plant parts that are above ground, and actively photosynthesizing must deliver energy to non-photosynthetic plant organs. Similarly, signals and information must be shared between distant parts of the same organism, which may be experiencing very different environmental cues. The phloem also serves as the route of chemical information delivery, and speed is of the essence. Velocity measurements of photosynthates in transport phloem (Minchin 2003), and of phloem bulk sap

(Jensen et al. 2011; Froelich et al. 2011; data presented here) reveal that the velocity of phloem sap in most plants ranges from as low as 20 μm/s (Chapter 2) to over 400 μm/s (Windt et al.

2006). The strong relationship between source length and transport length to the radius of the sieve tube presented by Jensen et al. (2012) is also supportive of the idea that transport phloem construction is geometrically optimized for speed.

The structure of transport phloem and viscosity of sap are crucial to considerations of its capacity to translocate sap over long distances. As can be inferred from Equation 2, sieve tubes with high conductivity and low viscosity will be most effective at rapid translocation that minimizes the need for high pressure gradients.

Unloading phloem

Major phloem unloading zones include vegetative apices such as shoot and root meristems, terminal vegetative storage sinks such as tubers, and terminal reproductive storage sinks such as fleshy fruits. In these sinks, symplastic continuity and observations of the behavior of fluorescent dyes indicate that unloading is symplastic, and accomplished by bulk flow through increasingly porous sieve tube members and companion cells. In developing seeds, symplastic isolation points to an apoplastic mechanism to fill these reproductive structures with energy rich molecules (Patrick 1997). The conversion of sucrose to starch in tubers, storage of small sugars in vacuoles in fruits, and rapid consumption for growth at meristems maintains low concentrations of sucrose in sinks, providing a continual “pulling” force that maintains a concentration and pressure differential, and therefore sap flow.

Rheological considerations

Energy transport from sources to sinks in plants is subject to an elaborate system of trade-offs.

Increasing the concentration of sucrose in the phloem sap will deliver more energy to sinks only at the expense of velocity of transport, and will require more pressure (See equations 1 and 2).

Because an apoplastic loader like Arabidopsis thaliana uses ATP to load the phloem and build turgor pressure, this comes also at an energy expenditure cost. Furthermore, sucrose presents

27 exponential increases in viscosity with incremental gains in solute potential drops. Compared to salts as osmolites, these viscosity increases are extreme. It is not surprising that most measurements of phloem sap viscosity correspond to a phloem sap concentration of sucrose that is just under 20%, as this is the point where further increases in sucrose concentration and solute potential drops begin to deliver higher viscosities at a steeply increasing rate (Swindells

1958). This means that this sucrose concentration in phloem sap provides high energy delivery with minimal viscosity, optimizing pressure driven flow. Although a wide range of sucrose concentrations have been observed in phloem (Jensen et al. 2013), the mean and the mode for all species surveyed in the above work converge near 20%.

Problems with the Pressure Flow Hypothesis

If pressure flow is driving sap, length would directly scale to pressure (Equation 2) in phloem.

The pressure measured in source phloem of trees is often lower than in herbaceous plants

(Turgeon 2010). Furthermore, trees often display an open minor vein configuration consistent with passive symplastic loading. It is surprising that a plant with such a long path length would not take advantage of an active loading strategy (Turgeon 2010). When the long path length, passive loading mechanism, and low phloem pressure measurements of trees are considered, it seems unlikely that Pressure Flow could achieve translocation in trees (Thompson and Holbrook

2003). It appears that only an especially high conductivity in the transport phloem of trees could explain why these plants can satisfy their transport requirements at low pressure gradients. Therefore, the morphology of several species of vines and trees was investigated in the present study in order to determine if there is indeed a correlation between long path lengths and increased conductivity.

The viscosity of phloem sap is also a limiting factor of flux as can be seen in Equations 1 and 2.

Sieve elements are deep in the tissue, small, and their contents tend to gel upon exposure to oxygen (Lang and Minchin 1986). Therefore, all previous reports of phloem sap viscosity have been estimated from analysis of exudates and aphid stylectomy collections. Here, for the first time, direct in-vivo measurements of translocating phloem sap viscosity are presented using the molecular rotor 2-(N-(7-Nitrobenz-2-oxa-1,3-diazol-4-yl)Amino)-2-Deoxyglucose (2-NBDG). Also presented are diffusivity measurements of living phloem using fluorescence recovery after photobleaching analysis in transporting phloem sap providing direct rheological measurements useful for rapid analysis of phloem sap solute concentrations, and for testing the Münch

Hypothesis.

WORKS CITED:

Behnke, Heinz-Dietmar. "Sieve–element characters." Nordic Journal of Botany 1, no. 3 (1981): 381-400. Berman-Frank, Ilana, Pernilla Lundgren, and Paul Falkowski. "Nitrogen fixation and photosynthetic oxygen evolution in cyanobacteria." Research in Microbiology 154, no. 3 (2003): 157-164. Bird, David A., Vincent R. Franceschi, and Peter J. Facchini. "A tale of three cell types: alkaloid biosynthesis is localized to sieve elements in opium poppy." The Plant Cell 15, no. 11 (2003): 2626-2635. Blankenship, Robert E. "Early evolution of photosynthesis." Plant physiology 154, no. 2 (2010): 434-438. Box, Raineer, M. Andrews, and J. A. Raven. "Intercellular transport and cytoplasmic streaming in Chara hispida." Journal of experimental botany 35, no. 7 (1984): 1016-1021. Brunkard, Jacob O., Anne M. Runkel, and Patricia C. Zambryski. “The cytosol must flow: intercellular transport through plasmodesmata.” Current opinion in cell biology 35, no. 1 (2015): 13-20. Chen, Li-Qing, Xiao-Qing Qu, Bi-Huei Hou, Davide Sosso, Sonia Osorio, Alisdair R. Fernie, and Wolf B. Frommer. "Sucrose efflux mediated by SWEET proteins as a key step for phloem transport." Science 335, no. 6065 (2012): 207-211. Cook, Martha E., Linda E. Graham, C. Botha, and Colleen Lavin. “Comparative ultrastructure of plasmodesmata of Chara and selected bryophytes: toward an elucidation of the evolutionary origin of plant plasmodesmata.” American Journal of Botany 84, no 9 (1997): 1169-1169. Eggert, Donald A., and N. Y. Kanemoto. "Stem phloem of a middle Pennsylvanian Lepidodendron." Botanical Gazette 138, no. 1 (1977): 102-111. Ehlers, Katrin, Michael. Knoblauch, and Aart J. E. Van Bel. "Ultrastructural features of well- preserved and injured sieve elements: minute clamps keep the phloem transport conduits free for mass flow." Protoplasma 214, no. 1-2 (2000): 80-92. Esau, Katherine. "Comparative structure of companion cells and phloem parenchyma cells in Mimosa pudica L." Annals of Botany 37, no. 3 (1973): 625-632. Esau, Katherine. "Structure and development of the bark in Dicotyledons."From: Formation of wood in forest trees (1964): 37-50. Esau, Katherine, and James Cronshaw. "Endoplasmic reticulum in the sieve element of Cucurbita." Journal of ultrastructure research 23, no. 1 (1968): 1-14. Esau, Katherine, and James Cronshaw. "Plastids and mitochondria in the phloem of Cucurbita." Canadian Journal of Botany 46, no. 7 (1968): 877-880.

Esau, Katherine, and Vernon I. Cheadle. "Mitochondria in the phloem of Cucurbita." Botanical Gazette 124, no. 2 (1962): 79-85. Esau, Katherine, and Vernon I. Cheadle. "Wall thickening in sieve elements."Proceedings of the National Academy of Sciences 44, no. 6 (1958): 546-553. Esau, Katherine, and Vernon I. Cheadle. "Size of pores and their contents in sieve elements of dicotyledons." Proceedings of the National Academy of Sciences 45, no. 2 (1959): 156-162. Evert, Ray F. "Seedless vascular plants." In Sieve Elements, pp. 35-62. Springer Berlin Heidelberg, 1990. Evert, Ray F., and Robert J. Mierzwa. "The cell wall-plasmalemma interface in sieve tubes of barley." Planta 177, no. 1 (1989): 24-34. Evert, Ray Franklin, and Katherine Esau. Esau's Plant Anatomy: Meristems, Cells, and Tissues of the Plant Body: Their Structure, Function, and Development. Hoboken, NJ: Wiley-Interscience, 2006. Feild, Taylor S., Timothy J. Brodribb, Ari Iglesias, David S. Chatelet, Andres Baresch, Garland R. Upchurch, Bernard Gomez et al. "Fossil evidence for Cretaceous escalation in angiosperm leaf vein evolution." Proceedings of the National Academy of Sciences 108, no. 20 (2011): 8363- 8366. Fitzgibbon, Jessica, Martina Beck, Ji Zhou, Christine Faulkner, Silke Robatzek, and Karl Oparka. "A developmental framework for complex plasmodesmata formation revealed by large-scale imaging of the Arabidopsis leaf epidermis." The Plant Cell 25, no. 1 (2013): 57-70. Franceschi, Vincent, Trygve Krekling, A. Berryman, and Erik Christiansen. "Specialized phloem parenchyma cells in Norway spruce (Pinaceae) bark are an important site of defense reactions." American Journal of Botany 85, no. 5 (1998): 601-601. Fraysse, Laure C., Brian Wells, Maureen C. McCann, and Per Kjellbom. "Specific plasma membrane aquaporins of the PIP1 subfamily are expressed in sieve elements and guard cells." Biology of the Cell 97, no. 7 (2005): 519-534. Freeman, Scott, Joan Catherine Sharp, and Mike Harrington. Biological science. Benjamin Cummings, 2010. Froelich, Daniel R., Daniel L. Mullendore, Kaare. H. Jensen, Timothy J. Ross-Elliott, James A. Anstead, Gary A. Thompson, Helene C. Pelissier, and Michael Knoblauch. "Phloem Ultrastructure and Pressure Flow: Sieve-Element-Occlusion-Related Agglomerations Do Not Affect Translocation." The Plant Cell 23, no. 12 (2011): 4428-445. Fromm, Jörg, and Tillmann Bauer. "Action potentials in maize sieve tubes change phloem translocation." Journal of Experimental Botany 45, no. 4 (1994): 463-469. Gamalei, Yuri. "Structure and function of leaf minor veins in trees and herbs." Trees 3, no. 2 (1989): 96-110.

Jensen, Kåre H., Jessica A. Savage, and Noel M. Holbrook. 2013. “Optimal concentration for sugar transport in plants.” Journal of the Royal Society Interface 10, no. 83 (2013): 0055. Jensen, Kåre Hartvig, Johannes Liesche, Tomas Bohr, and Alexander Schulz. "Universality of phloem transport in seed plants." Plant, cell & environment 35, no. 6 (2012): 1065-1076. Jensen, Kåre H., J. Lee, Tomas Bohr, Henrik Bruus, Noel M. Holbrook, and Maciej Zwieniecki. "Optimality of the Münch mechanism for translocation of sugars in plants." Journal of the Royal Society Interface 8, no. 61 (2011): 1155-1165. Kenrick, Paul, and Peter R. Crane. "The origin and early evolution of plants on land." Nature 389, no. 6646 (1997): 33-39. Kenrick, Paul, and Peter R. Crane. "The origin and early evolution of plants on land." Nature 389, no. 6646 (1997): 33-39. Knoblauch, Michael, Jan Knoblauch, Daniel L. Mullendore, Jessica A. Savage, Benjamin A. Babst, Sierra D. Beecher, Adam C. Dodgen, Kaare H. Jensen, and Noel Michele Holbrook. "Testing the Münch hypothesis of long distance phloem transport in plants." eLife 5 (2016): e15341. Knoblauch, Michael, Gundula A. Noll, Torsten Müller, Dirk Prüfer, Ingrid Schneider-Hüther, Dörte Scharner, Aart JE van Bel, and Winfried S. Peters. "ATP-independent contractile proteins from plants." Nature materials 2, no. 9 (2003): 600-603. Knoblauch, Michael, and Karl Oparka. "The Structure of the Phloem - Still More Questions than Answers." The Plant Journal 70, no. 1 (2012): 147-56. Knoblauch, Michael, Mike Stubenrauch, Aart J. E. Van Bel, and Winfried S. Peters. "Forisome Performance in Artificial Sieve Tubes." Plant, Cell & Environment 35, no. 8 (2012): 1419-427. Lang, Alexander, and Peter E. H. Minchin. "Phylogenetic distribution and mechanism of translocation inhibition by chilling." Journal of Experimental Botany (1986): 389-398. Langhans, Markus, Rafael Ratajczak, Martin Lützelschwab, Wolfgang Michalke, Rebecca Wächter, Elke Fischer-Schliebs, and Cornelia I. Ullrich. "Immunolocalization of plasma- membrane H+-ATPase and tonoplast-type pyrophosphatase in the plasma membrane of the sieve element-companion cell complex in the stem of Ricinus communis L." Planta 213, no. 1 (2001): 11-19. Li, Fay-Wei, Juan Carlos Villarreal, Steven Kelly, Carl J. Rothfels, Michael Melkonian, Eftychios Frangedakis, Markus Ruhsam et al. "Horizontal transfer of an adaptive chimeric photoreceptor from bryophytes to ferns."Proceedings of the National Academy of Sciences 111, no. 18 (2014): 6672-6677. Liesche, Johannes, and Alexander Schulz. "In vivo quantification of cell coupling in plants with different phloem-loading strategies." Plant physiology 159, no. 1 (2012): 355-365. Lucas, William J., and Jung-Youn Lee. "Plasmodesmata as a supracellular control network in plants." Nature Reviews Molecular Cell Biology 5, no. 9 (2004): 712-726.

Martens, Helle J., Alison G. Roberts, Karl J. Oparka, and Alexander Schulz. "Quantification of plasmodesmatal endoplasmic reticulum coupling between sieve elements and companion cells using fluorescence redistribution after photobleaching." Plant Physiology 142, no. 2 (2006): 471-480. McSteen, Paula. "Auxin and monocot development." Cold Spring Harbor Perspectives in Biology 2, no. 3 (2010): 1479. Minchin, Peter EH, and Michael R. Thorpe. "Using the short-lived isotope 11C in mechanistic studies of photosynthate transport." Functional Plant Biology 30, no. 8 (2003): 831-841. Mullendore, Daniel. L., Carel. W. Windt, Henk Van As, and Michael Knoblauch. "Sieve Tube Geometry in Relation to Phloem Flow." The Plant Cell 22, no. 3 (2010): 579-593. Mullineaux, Conrad W., Vicente Mariscal, Anja Nenninger, Hajara Khanum, Antonia Herrero, Enrique Flores, and David G. Adams. "Mechanism of intercellular molecular exchange in heterocyst‐forming cyanobacteria." The EMBO journal 27, no. 9 (2008): 1299-1308. Münch, Ernst. "Stoffbewegungen in der Pflanze." (1930). Nedelcu, Aurora M. "Evolution of Multicellularity." eLS (2001). Patrick, John W. "Phloem unloading: sieve element unloading and post-sieve element transport." Annual review of plant biology 48, no. 1 (1997): 191-222. Paridah, Md Tahir, Amel B. Basher, Syeed SaifulAzry, and Zakiah Ahmed. "Retting process of some bast plant fibres and its effect on fibre quality: A review." BioResources 6, no. 4 (2011): 5260-5281. Peuke, Andreas D., Marcus Rokitta, Ulrich Zimmermann, L. Schreiber, and Axel Haase. "Simultaneous Measurement of Water Flow Velocity and Solute Transport in Xylem and Phloem of Adult Plants of Ricinus Communis over a Daily Time Course by Nuclear Magnetic Resonance Spectrometry." Plant, Cell and Environment Plant Cell Environ 24, no. 5 (2001): 491-503. Peng, Duo, Xi Gu, Liang-Jiao Xue, James H. Leebens-Mack, and Chung-Jui Tsai. "Bayesian phylogeny of sucrose transporters: ancient origins, differential expansion and convergent evolution in monocots and dicots."Frontiers in plant science 5 (2014): 615. Rennie, Emilie A., and Robert Turgeon. "A comprehensive picture of phloem loading strategies." Proceedings of the National Academy of Sciences 106, no. 33 (2009): 14162-14167. Riesmeier, Jorg W., Lothar Willmitzer, and Wolf B. Frommer. "Isolation and characterization of a sucrose carrier cDNA from spinach by functional expression in yeast." The EMBO Journal 11, no. 13 (1992): 4705. Rodiuc, Natalia, Paulo Vieira, Mohamed Youssef Banora, and Janice de Almeida Engler. "On the track of transfer cell formation by specialized plant-parasitic nematodes." Transfer Cells (2015): 81.

Ruiz-Medrano, Roberto, Beatriz Xoconostle-Cázares, and William J. Lucas. "The phloem as a conduit for inter-organ communication." Current opinion in plant biology 4, no. 3 (2001): 202- 209. Samejima, Michikazu, and Takao Sibaoka. "Identification of the excitable cells in the petiole of Mimosa pudica by intracellular injection of procion yellow." Plant and Cell Physiology 24, no. 1 (1983): 33-39. Savage, Jessica A., Maciej A. Zwieniecki, and N. Michele Holbrook. "Phloem transport velocity varies over time and among vascular bundles during early cucumber seedling development." Plant physiology 163, no. 3 (2013): 1409-1418. Scarpella, Enrico, Philip Francis, and Thomas Berleth. "Stage-specific markers define early steps of procambium development in Arabidopsis leaves and correlate termination of vein formation with mesophyll differentiation." Development 131, no. 14 (2004): 3445-3455. Scheirer, Daniel C. "Mosses." In Sieve Elements, pp. 19-33. Springer Berlin Heidelberg, 1990. Schirrmeister, Bettina E., Jurriaan M. de Vos, Alexandre Antonelli, and Homayoun C. Bagheri. "Evolution of multicellularity coincided with increased diversification of cyanobacteria and the Great Oxidation Event." Proceedings of the National Academy of Sciences 110, no. 5 (2013): 1791-1796. Seward, Albert Charles. "On the so-called phloem of lepidoendron." New Phytologist 1, no. 2 (1902): 38-46. Shulz, Alexander. “Conifers.” In Sieve Elements, pp. 63-88. Springer Berlin Heidelberg, 1990. Sovonick, Susan A., Donald R. Geiger, and Robert J. Fellows. "Evidence for active phloem loading in the minor veins of sugar beet." Plant Physiology 54, no. 6 (1974): 886-891. Thompson, Matthew V., and N. Michele Holbrook. "Application of a Single-solute Non-steady- state Phloem Model to the Study of Long-distance Assimilate Transport." Journal of Theoretical Biology 220, no. 4 (2003): 419-55. Tominaga, Motoki, and Kohji Ito. "The molecular mechanism and physiological role of cytoplasmic streaming." Current opinion in plant biology, 27 (2015): 104-110. Turgeon, Robert. "Phloem loading and plasmodesmata." Trends in Plant Science 1, no. 12 (1996): 418-423. Turgeon, Robert. "The Puzzle of Phloem Pressure." Plant Physiology 154, no. 2 (2010): 578-81. Turgeon, Robert. "The role of phloem loading reconsidered." Plant physiology152, no. 4 (2010): 1817-1823. Van Bel, Aart J.E., and Michael Knoblauch. "Sieve element and companion cell: the story of the comatose patient and the hyperactive nurse. "Functional Plant Biology 27, no. 6 (2000): 477- 487.

Van Bel, Aart J.E. "Evolution, polymorphology and multifunctionality of the phloem system." Perspectives in Plant Ecology, Evolution and Systematics 2, no. 2 (1999): 163-184. Windt, Carel W., Frank J. Vergeldt, P. Adrie De Jager, and Henk Van As. "MRI of Long-distance Water Transport: A Comparison of the Phloem and Xylem Flow Characteristics and Dynamics in Poplar, Castor Bean, Tomato and Tobacco." Plant, Cell and Environment Plant Cell Environ 29, no. 9 (2006): 1715-729. Zeeman, Samuel C., Steven M. Smith, and Alison M. Smith. "The breakdown of starch in leaves." New Phytologist 163, no. 2 (2004): 247-261. Zhang, Shoudong, Li Sun, and Friedrich Kragler. "The phloem-delivered RNA pool contains small noncoding RNAs and interferes with translation." Plant physiology 150, no. 1 (2009): 378-387.

Chapter 2: Diffusion and Velocity FRAP imaging in living sieve tubes of Arabidopsis thaliana confirms that cold shock velocity response does not scale to changes in sap rheology.

Sierra D. Beecher1, Kåre H Jensen2, Michael Knoblauch1 1: School of Biological Sciences, Washington State University 2: Department of Physics, Technical University of Denmark Abstract: Here we present a new method to simultaneously measure in vivo phloem sap translocation velocity and diffusivity of carboxyfluorescein in living, translocating sieve tubes of Arabidopsis thaliana using fluorescence recovery after photobleaching (FRAP). Experimental results indicate that cold shock phenomenon is not caused by changes in the viscosity of the phloem sap in these plants.

Introduction:

The transport of photoassimiliates form source leaves to distant heterotrophic sinks through the sieve tube network of the phloem is a critical step to maintain functional integrity of vascular plants. It has long been recognized that rapid local chilling of petioles or stem sections causes immediate halt of phloem flow (Swanson and Geiger 1967; Lang 1974; Pickard and

Minchin 1990; Pickard and Minchin 1992). Over the last decades a comprehensive set of information on the phenomenology of shock induced flow stoppage has been collected which include: a) Stimuli that cause sudden halt are rapid cooling (but not warming), electro shock, strong vibration (Pickard and Minchin 1990); b) fatigue occurs when shocks of the same kind are applied within short time, but fatigue may not occur when different stimuli are applied within the same time frame (Pickard and Minchin 1992); c) the duration of fatigue is dependent on the strength of the stimulus (Minchin and Thorpe 1983); d) in cold shocked plants, flow

36 resumes after a while even if the tissue is kept at low temperatures (Swanson and Geiger 1967); e) flow stoppage is only local at the stimulated region, but flow continues apical or basal of the stimulated region although at a lower rate (Pickard et al. 1978; Pickard and Minchin 1992).

More recent investigations on cold shock induced signaling indicate that action potentials are generated that travel at 3 cm /s along the phloem, while wounding induces variation potentials traveling at 0.5 cm/s (Fromm et al. 2013).

It is yet unclear what downstream events lead to a halt of phloem flow. In order to stop flow, at least one of the flow defining parameters has to change dramatically. If we assume that phloem flow follows an osmotically generated pressure gradient, then the defining parameters are described in the Hagen-Poiseuille equation

푘 훥푝 푈 = , (1) 휂 퐿 where U is the flow velocity, Δ푝 is the pressure differential, L is the tube length, ɳ is the sap viscosity, and k is the conductivity of the tube.

A drastic change in the length of the tube to account for flow stoppage can be excluded. A pressure drop in the stimulated area would generate a local sink, while an increase would generate a local source. In both cases this would result in a partial reversal of flow towards the new sink or source respectively, which is not supported by the available data. Therefore, only a change in conductivity by physical barriers like callose formation or p-protein plug formation, or a drastic increase in viscosity can be the reason for flow stoppage.

The discovery of a) the reversible calcium induced volume change of specific p-proteins called forisomes in the family fabaceae (Knoblauch and Peters 2004), b) evidence that forisomes can occlude microchannels efficiently (Shen et al. 2006), and c) the fact that action potentials lead to intracellular calcium increases, prompted studies on the correlation of the phenomena and indeed simultaneous reactions have been demonstrated (Thorpe et al. 2010). Calcium induced forisome reaction roughly correlated with flow behavior.

Investigations on p-proteins in non-fabaceae plants, however, did not provide evidence that p- proteins in other species have similar functions. In situ observation of forisome homologue p- proteins in Arabidopsis did not show a visible reaction to either cold shock or electro shock.

Most importantly, Arabidopsis knockout mutants which lack structural p-proteins show a cold shock induced flow stoppage (Knoblauch et al. 2014). Thus p-protein independent control occurs and the question arises if an overarching p-protein independent mechanism exists.

An increase in phloem sap viscosity could account for a flow stoppage, and phloem sap gelling in phloem sap exudates is a well-known phenomenon in many plant species (Lang and Minchin

1986; Dinant et al. 2003). Therefore we aimed to investigate if there is a correlation between cold shock and intracellular viscosity changes. We developed a confocal microscope based method that provides values on relative phloem sap viscosity changes in situ.

Materials and Methods

Plants were grown in microscopy Rhizosphere chambers (micro-ROCs, Advanced Science Tools,

Pullman, WA) in a 14/10 hour light/dark period in a greenhouse at (night 20-24deg ~ day 24-

27deg) with natural light supplemented with a 400W high pressure sodium lamp one meter

38 above the plants that maintained a light intensity of 250 μmol/m2/s, and in a growth chamber with a 16/8hr light/dark photoperiod at 20 °C with 190 μmol/ m2/s intensity provided by T5 fluorescent lamps containing 6500k and 3000k bulbs. Plants were investigated 8-15 days after germination. Stock 5(6)-Carboxyfluorescein diacetate (CFDA) from Sigma-Aldrich solution is prepared by dissolving 100mg of CFDA into 5mL acetone. This stock was mixed 1:5 with deionized water and applied as described in Knoblauch et al. (2015). Imaging was done with a

Leica SP5 confocal microscope. CF was excited with the 488nm laser line of a supercontinuum

Argon laser with a Leica SP5 confocal microscope using a 20X Leica water immersion lens. The

520 – 560 band of fluorescence emission was detected with a HyD detector. After the sieve tube system was well labeled with the dye, a straight section of transport phloem was photo- bleached and phloem flow was monitored as described in Froelich et al. (2011). Analysis of transport velocity and viscosity was performed as described below. Cold shock stimulation was achieved by replacing the immersion water with 1 ml of ice-cold water.

Results

In situ phloem flow velocity measurements in individual tubes was established recently

(Froelich et al. 2011). The methodology employs distant application of phloem mobile fluorochromes to a leaf and consecutive observation of tube filling in the root system by confocal microscopy. Several appropriate fluorochromes are now available (Knoblauch et al.

2015). Once the tube is filled, high energy laser light is applied to bleach the fluorochrome and to generate a distinct dye front. After decrease of the laser intensity, refilling of the tube is monitored (Figure 1).

Figure 1. In situ flow velocity measurements. Three frames extracted from supplemental movie 1 showing the progression of the dye front (arrows) over time. The spread of the front due to diffusion provides a reading of sap viscosity. Example is a cold-stimulated plant, exhibiting a slow velocity of about 20 μm/s.

Flow velocity can be analyzed by defining regions of interest with known distance along the tube, and measuring a fluorescence recovery curve for each ROI (Figure 2).

Figure 2: In situ phloem sap viscosity analysis.

To analyze phloem sap viscosity data, ROIs are defined along the sieve tube and the distance between them is measured. The fluorescence intensity of each frame of the movie is then measured and plotted for each ROI individually. The distance between ROIs and the difference in time it takes to reach the 50% fluorescence intensity recovery point between ROIs provides the flow velocity. The change in slope of the recovery curve can be used to calculate the viscosity using equation (5).

Depending on the flow velocity, refilling of the tube within the visible area of the microscope requires several seconds. Within this time period, the bleach front will spread out due to diffusion which is directly correlated to tube sap viscosity and is manifested in a change of the slope in the recovery curve (Figure 2). The lower the viscosity, the shallower the slope will become within a given time frame. This follows the relation:

The width w of the front grows with the square root of time t

푤 = (푘퐷푡)1/2 (2) where k = 4π (Carroll et al. 2014) and D is the diffusivity. The front moves with speed v, and the time it takes to reach a specific ROI is

푑 푡 = (3) 푣 where d is the distance from the bleaching point to the ROI position. The transition time τ is related to the speed and width by

푤 휏 = (4) 푣

This leads to a relation between speed v, transition time τ, and distance d

푘푑 푣3 = 퐷 (5) 휏2

We conducted in situ flow velocity measurements at ambient temperature in micro-ROCs, as described above and analyzed the spread of the dye front according to equation (5). The results show a linear relationship between v3 and dkτ−2 over 3 orders of magnitude (Figure 3; n=24).

Figure 3: A plot of v3 and dkτ−2 indicates a linear relationship.

After the initial flow velocity and viscosity analysis, we replaced the immersion water of the microscope objective with ice water to induce a cold shock. Temperature measurements in micro-ROCs where the root was replaced by a thermocouple indicated an instant temperature drop of 8.2 °C (n=6, +/- 0.44 [stdev]) when applying 1ml of ice water. Cold shocked roots showed an immediate flow velocity reduction (Figure 4). The flow, however, did not stop

43 entirely but continued at a slower speed (Figure 4) with an average reduction of 4.9 times

(n=7).

Figure 4: Phloem flow velocity change after cold shock. Seven individual measurements of the phloem flow velocity before and after cold shock. In average, phloem flow was reduced 4.9 times at a temperature reduction of 8.5 °C.

Analysis of the spread of the dye front according to equation (5) to evaluate viscosity changes, showed that the data points from cold shock experiments follow the linear relationship which indicates that the increase in viscosity is low and only due to the temperature induced reduction. No additional significant viscosity increase that could account for the observed 4.9 times flow velocity reduction was measured (Figure 5).

Figure 5: v3 versus dkτ−2 The phloem sap viscosity after cold shock (open circles) decreases as expected for temperature reduction, but there is no additional viscosity increase that could explain the 4.9 times reduction in flow velocity. For this the viscosity values would have to cluster around the indicated black line.

The magnitude of increase in viscosity of a 20% sucrose solution caused by this temperature drop (from 18C to 9.8C) is 1.21 fold/8.2 C, (equation derived from Table 130, Swindells 1958) compared to the 4.9 times reduction in phloem flow velocity observed in in-vivo plants subjected to the same temperature treatment.

Discussion

Our aim was to develop a method to evaluate phloem sap viscosity in situ. The method provides relative viscosity changes. Although absolute viscosity and diffusivity values could in principle be extracted, it is in practice not possible in living plant tissue, since the laser energy to generate a distinct bleach front would be too high. Because of the continuous flow, new fluorochromes arrive constantly in the bleach area and it takes a certain time until all fluorochromes are bleached. The time is dependent on the applied laser energy and the flow velocity. In any case a bleach profile instead of a distinct straight front develops because of the laminar flow profile with faster flow towards the center of the tube (Figure 6 A). This bleach profile remains constant at specific flow velocity and laser intensity. Increasing the laser intensity can reduce the bleach profile to a negligible size (Figure 6 B), which is feasible in an artificial microfluidic system (Carroll et al. 2014). In living plants, however, it turned out that the required energy input is too high and phloem flow stops instantly due to artifact generation.

Since a straight bleach front is assumed in equation (5), the parabolic nature of the bleach front leads to a calculated diffusivity of 703 ± 49 µm2 / s which is less than the viscosity of water.

Figure 6: Schematic drawing of the development of a bleach profile. Depending on tube diameter and flow velocity, a specific flow profile will develop (indicated by black arrows). The laser in the application area will photobleach the dye. Due to differential velocities in the tube and dependent on the laser intensity, a laser bleach profile develops (upper image). Increase of the laser intensity leads to faster bleaching and a more distinct bleach profile.

When considering effects of the initial shape of the front on the dispersion timescales we have to alter the equation accordingly. We assume that the width w of the front grows with the square root of time t

1/2 푤 = 푤0 + (푘퐷푡) (6)

where w0 is the initial front width. The front moves with speed v, and the time it takes to reach the observer is

푑 푡 = (7) 푣

where d is the distance from the bleaching point to the observation position. The transition time τ is related to the speed and width by

푤 푤 +(푘퐷푡)1/2 휏 = = 0 (8) 푣 푣

This leads to a relation between speed v, transition time τ, and distance d:

2 (푣휏 − 푤0) = 퐷푑푘 (9)

Expanding the parenthesis gives

푘푑 (푣3휏2 + 푤2푣 − 2푣2휏푤 )/휏2 = 퐷 (10) 0 0 휏2

This allows us to estimate the diffusivity D. With an initial front width of w0 = 50 μm we find the value D = 215 ± 39 μm2/s, close to reported values for 20% sugar solution.

Because the flow velocity, dye concentration, and tube geometry vary between specimens, it is basically impossible to estimate a front width in in situ experiments to receive absolute values.

But it should be noted that the methods and mathematical principles can be applied to other

(e.g. artificial) systems.

To answer the question if a viscosity increase is a general mechanism for the cold shock induced phloem flow stoppage, the measurement of relative viscosity changes is of high value. Our data show two important findings. A) Measured sap viscosity values are - as expected for

48 temperature reduction - due to the natural increase in molecular interaction and friction at lower temperatures, but no additional viscosity increases that could account for the observed flow reduction were measured. Thus, only a mechanism that leads to a conductivity reduction appears plausible to explain the cold shock reaction. B) In contrast to other flow velocity measurement techniques, the spatial and temporal resolution of direct flow observation using fluorochromes is significantly better. The cold shock induced flow response is not an all or nothing response, but the flow may only slow down. This points towards a gating mechanism rather than an all or nothing blockage. Minchin (1983) mentioned that some of his plants showed reduced velocity instead of completely stopping, and Lang and Minchin (1986) noted partial sensitivity (1/2 velocity reduction) in radish.

The two sieve tube structures that have been implicated with phloem occlusion are P-proteins and callose. Forisomes are the classical p-protein example for tube occlusion, but they follow an all or nothing scheme rather than a graded swelling. A lag time between forisome reaction and cold stimulated flow stoppage in Vicia faba has been noticed (Thorpe et al. 2010). In addition, the mechanism of cold shock induced flow reduction has been observed in all dicots and several monocots. The phylogenetic distribution of p-proteins in monocots does not coincide with the observed distribution of cold shock reaction. Poaceae are the only angiosperms without p- proteins, but the lack of cold shock reaction has been reported for monocots such as onions, yucca etc. Furthermore, Arabidopsis p-protein knock-outs maintain cold shock sensitivity.

Callose on the other hand is deposited on sieve plate pores to reduce their diameter which is a graded event and a step wise gating would certainly be possible. Reported callose deposition

49 rates are, however, in the range of 1nm per second (Mullendore et al. 2010) which would not be sufficient to reduce flow in the observed time frame, even in plants with very small sieve plate pores of 100-250 nm like in Arabidopsis. In plants with large pores such as Cucurbita, full pore occlusion did not occur even after 24 hours (Mullendore et al. 2010), but rapid, cold stimulated flow stoppage did occur (Lang and Minchin 1986).

Cold shock induced action potentials traveling at speeds of 3cm per second (Fromm et al 2013) would be sufficiently fast to induce source or sink alterations. But even shutting down loading or unloading mechanisms would not result in an instant halt of flow. Specifically the observation that flow stoppage is local and flow continues up- and downstream of the stimulated area (although at a lower rate) speaks against distantly induced mechanisms.

In essence, none of the known or suggested mechanisms of sieve tube occlusion can be brought in line with all observations. In conclusion, there may be species specific differences in employing callose, p-proteins, and maybe other, yet to identify processes.

WORKS CITED:

Carroll, Nick J., Kaare H. Jensen, Shima Parsa, N. Michele Holbrook, and David A. Weitz. "Measurement of flow velocity and inference of liquid viscosity in a microfluidic channel by fluorescence photobleaching."Langmuir 30, no. 16 (2014): 4868-4874. Dinant, Sylvie, Anna M. Clark, Yanmin Zhu, Françoise Vilaine, Jean-Christophe Palauqui, Chantal Kusiak, and Gary A. Thompson. "Diversity of the superfamily of phloem lectins (phloem protein 2) in angiosperms." Plant Physiology 131, no. 1 (2003): 114-128. Froelich, Daniel R., Daniel L. Mullendore, Kåre H. Jensen, Tim J. Ross-Elliott, James A. Anstead, Gary A. Thompson, Hélène C. Pélissier, and Michael Knoblauch. "Phloem ultrastructure and pressure flow: sieve-element-occlusion-related agglomerations do not affect translocation." The Plant Cell 23, no. 12 (2011): 4428-4445. Fromm, Jörg, Mohammad-Reza Hajirezaei, Verena K. Becker, and Silke Lautner. "Electrical signaling along the phloem and its physiological responses in the maize leaf." Frontiers in plant science 4, no. (2013): 239. Knoblauch, Michael, and Winfried S. Peters. "Forisomes, a novel type of Ca2+‐dependent contractile protein motor." Cell motility and the cytoskeleton 58, no. 3 (2004): 137-142. Knoblauch, Michael, Daniel R. Froelich, William F. Pickard, and Winfried S. Peters. "SEORious business: structural proteins in sieve tubes and their involvement in sieve element occlusion." Journal of experimental botany 65, no. 7 (2014): 1879-1893. Knoblauch, Michael, Marc Vendrell, Erica de Leau, Andrea Paterlini, Kirsten Knox, Tim Ross- Elliot, Anke Reinders, Stephen A. Brockman, John Ward, and Karl Oparka. "Multispectral phloem-mobile probes: Properties and applications." Plant physiology 167, no. 4 (2015): 1211- 1220. Lang, Alexander, and Peter E. H. Minchin. "Phylogenetic distribution and mechanism of translocation inhibition by chilling." Journal of Experimental Botany (1986): 389-398. Lang, Alexander. "The effect of petiolar temperature upon the translocation rate of 137Cs in the phloem of Nymphoides peltata." Journal of experimental botany 25, no. 1 (1974): 71-80. Minchin, Peter, and Michael R. Thorpe. “A Rate of Cooling Response in Phloem Translocation.” Journal of Experimental Botany 34, no. 142 (1983): 529-536. Mullendore, Daniel L., Carel W. Windt, Henk Van As, and Michael Knoblauch. "Sieve Tube Geometry in Relation to Phloem Flow." The Plant Cell 22, no. 3 (2010): 579-593. Peng, Duo, Xi Gu, Liang-Jiao Xue, James H. Leebens-Mack, and Chung-Jui Tsai. "Bayesian phylogeny of sucrose transporters: ancient origins, differential expansion and convergent evolution in monocots and dicots." Frontiers in plant science 5 (2014): 615.

Pickard, William F., and Peter E.H. Minchin. “The Transient Inhibition of Phloem Translocation in Phaseolus vulgaris by Abrupt Temperature Drops, Vibration, and Electric Shock." Journal of Experimental Botany 41, no. 232 (1990): 1361-1369. Pickard, William F., Peter E. H. Minchin, and John H. Troughton. "Transient inhibition of translocation in Ipomoea alba L. by small temperature reductions."Functional Plant Biology 5, no. 2 (1978): 127-130. Pickard, William F., and Peter E.H. Minchin. “The Nature of the Short Term Inhibition of Stem Translocation Produced by Abrupt Stimuli.” Australian Journal of Plant Physiology 19 (1992): 471-480. Shen, Amy Q., Benjamin D. Hamlington, Michael Knoblauch, Winfried S. Peters, and William F. Pickard. "Forisome based biomimetic smart materials." Smart Structures and Systems 2, no. 3 (2006): 225-235. Swanson, Carroll A., and Donald R. Geiger. "Time course of low temperature inhibition of sucrose translocation in sugar beets." Plant Physiology 42, no. 6 (1967): 751-756. Swindells, James F. “Viscosities of sucrose solutions at various temperatures: tables of recalculated values.” USGPO. Vol. 440. (1958). Thorpe, Michael R., Alexandra CU Furch, Peter EH Minchin, Jens Foeller, Aart JE Van Bel, and Jens B. Hafke. "Rapid cooling triggers forisome dispersion just before phloem transport stops." Plant, cell & environment 33, no. 2 (2010): 259-271.

Chapter 3: Phloem sap Viscosity

The fluorescent glucose analog 2-(N-(7-Nitrobenz-2-oxa-1,3-diazol-4-yl)Amino)-2- Deoxyglucose (2-NBDG) is also a phloem mobile molecular rotor, useful for the direct measurement of the viscosity of translocating phloem sap.

Sierra Beecher, Michael Knoblauch and Daniel Mullendore Washington State University, School of Biological Sciences

Abstract:

We established the cell-permeant, phloem-mobile vital stain 2-(N-(7-Nitrobenz-2-oxa-1,3- diazol-4-yl)Amino)-2-Deoxyglucose (2-NBDG) as a molecular rotor, that responds to higher viscosities induced by sucrose and glycerol by exhibiting longer fluorescence lifetimes.

Fluorescence lifetime was shown to dominantly be affected by viscosity, as even high concentrations of NaCl did not appreciably alter the response, and low pH had only a minimal effect. We used this method to measure the viscosity of in-vivo transporting phloem sap in the roots of juvenile Arabidopsis thaliana and leaf major veins of Ipomoea nil. Results are in good agreement with previously obtained estimations of phloem sap viscosity based on exudate analysis.

Introduction:

The viscosity of biofluids is of interest in many lines of inquiry in life sciences. Diffusion of small molecules through cytoplasm is a function of viscosity (Kao et al. 1993), and this affects all aspects of cell biology. The viscosity of extracellular fluids is also determinant to physiology, as demonstrated by the findings that blood plasma viscosity is a biomarker for Alzheimer’s disease in humans (Aras et al. 2012), and that the viscosity of saliva can be an indication of oral health

(Animireddy et al. 2014). It can also indicate oncological illness. For example, benign ovarian cells are more viscous than late stage Mouse Ovarian Surface Epithelial cells (Ketene et al.

2012), and cell death is accompanied by an increase in cell viscosity (Kuimova et al. 2009).

Plants are no exception. The rheological nature of biofluids is also very important for many aspects of plant physiology. Water relations in plants are a dominant part of their evolutionary strategies. The vacuoles of plant cells maintain turgor with low-viscosity solutes (Evert 2006).

Root tips secrete viscous mucins to lubricate their path through the soil (Raven and Edwards

2001). Nectar with regulated sugar concentrations are crucial for pollination. Some plants even use viscous secretions as insect traps (Ellison and Gotelli 2001). Importantly for this inquiry, the viscosity of phloem sap is essential information to understand photoassimilate transport in vascular plants (Münch 1930, Knoblauch and Peters 2013, Mullendore et al. 2010).

Fluorescence microscopy methods have previously been used to investigate viscosity in living cells. Spot bleaching and subsequent measurement of fluorescence recovery after photobleaching (FRAP) was used by Verkman (2002) to show that the mitochondrial matrix, and the ER lumen have lower diffusion coefficients than cytoplasm, and that diffusion in cytosol is only about 30-40% slower than it is in water. Total Internal Reflectance-Fluorescence Recovery

After Photobleaching was used in Madin-Darby Canine Kidney Epithelial Cells by Swaminathan et al. in 1996 to show that the cytoplasm near the cell membrane was more viscous than bulk cytoplasm, functioning as a selective sieve. Ratiometric fluorescence is a method that uses two fluorescent probes. When microinjected into cells, the ratio of their fluorescent signal can indicate important viscometric information about cytoplasm, and cytosol. Luby-Phelps et al.

(1993) used this method to show that the fluid phase of cytosol in female rat kidney epithelial

54 cells of the line PtK1, and CV1 cells, a line from the kidney of an adult male African green monkey, had a very low viscosity, and their results suggest that cytosolic water was probably in a free, not organized form. This use of fluorescent viscosity measurement settled a long debate about the microenvironment of cell cytoplasm.

More recently, the use of molecular rotors has been employed to analyze the viscosity of living cells (Kuimova et al. 2009; Levitt et al. 2011). Molecular rotors are fluorescent molecules that, when excited with light, return to their ground state energy level not only by emitting fluorescence, but also by rotoation. In more viscous mediums, more energy is lost as fluorescence, as rotation is inhibited. This increases the fluorescence intensity output, and also the fluorescence lifetime (Suhling et al. 2012).

In all of these previous efforts, the presence of cytoskeleton and membranes incurred issues that required correction factors to solve for the fluid phase cytosol, because these structures reduce movement of probes physically when they interact with them. Diffusion is impeded, and molecular rotation is slowed or stopped. These physical obstructions can be misinterpreted as a high degree of solute in the cytosol, rather than the effect of presence of cytoskeletal elements.

2-(N-(7-nitrobenz-2-oxa-1,3-diazol-4- yl)amino)-2-deoxyglucose (2-NBDG) is synthesized by reacting D-glucosamine with NBD-Cl. This molecule shows spectra with maxima of 475 nm excitation and 550nm emission. This fluorescent glucose analog is demonstrated to be useful as a fluorescent probe for analyzing viability in E. coli cells by monitoring glucose uptake. Living cells accumulate the probe, and dead ones do not. Accumulation of fluorescence is inhibited by

D-glucose, but not L-glucose. Sucrose does not present inhibition (Yoshioka et al. 1996).

O’Neill et al. (2005) discovered that breast and liver tumorogenic cells take up 2-NBDG at a rate four times that of non tumorogenic cells, and retain it in cytoplasm in phosphorylated form.

The fluorescent subunit is attached at the 2 position on the glucose molecule; therefore, metabolism of 2-NBDG is stalled at the isomerase step of glycolysis, and fluorescence accumulates. Because the glucose transporter GLUT1 is expressed to a higher degree in tumor cells, increased glucose uptake is expected. 2-NBDG is presented as a probe for cancer cells, and is implicated as an index for higher rates of metabolism in tumorogenic cells.

Work by Etxeberria et al. (2005) demonstrates that 2-NBDG can also enter the cytoplasm, vacuoles, and organelles of protoplasts and cytoplasts prepared by cell wall removal and centrifugation of Acer pseudoplatanus L cell cultures. Hexoses are transported apoplastically; either selectively, by H+ transporters into the cytosol, or non-selectively, and straight to vacuoles, by endocytosis through vesicles. Import of 2-NBDG was stunted into the cytoplasm in the presence of Phloridzin, which inhibits transporters. However, transport into vacuoles by endocytosis was not disturbed, and fluorescent vesicles were observed. Cells incubated with wortmannin-A, an endocytosis disruptor, showed low-level fluorescence in the cytosol, but inhibited fluorescence accumulation in the vacuole, even after 24 h incubation. Therefore, 2-

NBDG may enter plant cells through two apoplastic pathways - endocytosis, or H+/Hexose mediated membrane symport. Knoblauch et al. (2015) reported that the probe is not transported by SUC transporters.

2-NBDG is also phloem-mobile (Hofmann et al. 2009; Knoblauch et al. 2015), and its structure is favorable to rotational energy loss. This led to the current efforts to investigate the fluorophore

56 as a molecular rotor, and therefore as a means to measure viscosity using FLIM in the phloem sap of living plants (Knoblauch et al. 2016; This work).

Molecular rotors exhibit both fluorescence intensity and fluorescence lifetime increases as a function of viscosity. Fluorescence intensity will also increase as a function of the concentration of the dye. Therefore, fluorescence lifetime is a more useful parameter to measure viscosity in biological tissue, where the relative concentrations in different domains are not known

(Kuimova 2012).

Translocating sieve elements do not contain cytoskeleton, and have few organelles, which are dominantly located in the periphery (Evert 2006; Mullendore et al. 2010). Fluorescent techniques to measure in-vivo cellular viscosity are therefore optimized in this tissue. This is especially useful information, as phloem sap viscosity is an index of sucrose concentration, and viscosity is a needed parameter to evaluate the hypothesis that pressure driven flow is responsible for translocation, and to test models about optimization.

The Pressure Flow Hypothesis is the model of phloem transport that most plant science researchers work under. It states that sugars are transported from photosynthetic sources such as leaves in plants, to heterotrophic sinks such as roots or fruits by a hydrostatic pressure differential. This pressure differential is generated by a high concentration of osmotically active sugars in the sources pulling water into the phloem down a water potential gradient. Because cell walls in plants prevent the rupture seen in animal cells exposed to hypotonic media, a hydrostatic pressure is built, which pushes sap to low-sugar concentration sinks of the plant through the interconnected, enucleate, syncytious cellular sieve tube system of the plant.

This hypothesis can be tested by comparing measured parameters in plants to the Haagen-

Poiseulle equation describing pressure flow in a low Reynolds number tube that has been recently modified for phloem by Jensen et al. (2011). The Jensen equation considers the periodicity of sieve plates, and introduces the value of conductivity (k). Viscosity of the phloem sap is inversely proportional to conductivity, the pressure differential, and velocity in these equations. Previous to this investigation, phloem sap viscosity has never been directly measured in vivo, but rather viscosity has been estimated by chemical analysis of contents of exudates. The accuracy of these previous estimations is unclear, due to contamination from proximal cells, and potential gelling of the sap when it is exposed to ambient oxygen.

Exudate measurements return varying results for sucrose concentration. Ricinus displays superior phloem exudation rates with minimal gelling. Hall and Baker (1972) made some measurements of Ricinus phloem sap, using this collection method. Although his observations that 80-90% of Ricinus phloem sap dry matter is sucrose, and that phloem sap has a low viscosity, and an alkaline pH (8.03) are in agreement with newer analyses by Gessler et al.

(2008), their measurements of total sucrose concentration are widely divergent, with Hall reporting a 10-12% w/w concentration of sucrose in exudate, and Gessler finding over 3.5M total carbon, with 91% of that being from sucrose.

Compared to incision and exudation, aphid stylectomy is a superior method for phloem sap collection. In 1953, Kennedy and Mittler reported that sleeving the part of an aphid’s rostrum that remains in the plant after severing it from the insect during feeding with a capillary offers the ability to collect phloem sap from willow at the rate of about 1 mm3/h. It was inferred from this that aphids rely on the turgor of the phloem rather than “sucking” to obtain sap. The

58 observation that wilted leaves are less preferred by aphids was supportive of this inference.

Although developed to study aphid physiology, this method presented to plant physiologists a new method for phloem sap collection and analysis superior to whole tissue phloem sap extraction methods in use at the time of publication. Since then, optimized means have been developed to use multiple cutting and collection techniques for aphid stylectomy, possibly allowing more precise analysis of phloem sap (Fisher and Frame 1984).

Measurements of sugar w/w % concentrations in phloem sap of many plants by many workers using both methods range from 4.3% in cabbage to over 50% in potato (compiled in Jensen et al. 2013). It is difficult to know if these varying measurements indicate interesting phyisiological aspects of the plants in question, or whether they are artefacts of collection. Additionally, sap collection methods have been shown to yield varying measurements of sugar type and concentration when applied to the same species and cultivar of lettuce. Finally, some plants, such as those that are resistant to aphids, are not possible to gain measurements from via stylectomy (Helden et al. 1994).

Recently, Fluorescence Recovery After Photobleaching (FRAP) has been utilized to measure the diffusivity of in-vivo, translocating phloem sap (See Chapter 2). Though this is not a direct measurement of viscosity, diffusivity and viscosity are inversely related by the Einstein-Stokes equation, and values for diffusivity allow calculated estimates of viscosity.

Because sieve elements are optimal for using molecular rotors to measure in-vivo viscosity due to the minimal obstructions present in these cells, and because 2-NBDG is phloem mobile, non-

59 toxic, and behaves as a molecular rotor, this system presents an excellent opportunity to contribute to growth in understanding of energy transport in vascular plants.

Materials and Methods:

Preparation of standard solutions:

A Series of w/w sucrose solutions corresponding to viscosities of 1, 2.03, 3.02, 5.30, and 10.30 mPas were prepared using PBS as solvent at 0%, 21%, 29%, 38%, and 46% at ambient temperature (20°C) as per Swindells (1958) using PBS at pH 7.2 as solvent. All concentrations were prepared at least three separate times for precision. Glycerol viscosity series were prepared in PBS solvent similarly with 10% (1.3 mPas), 50% (5.98 mPas), 80% (59.9 mPas), and

100% (1400 mPas) at ambient temperature, according to tables from Segur and Oberstar

(1951). Sodium chloride solutions were prepared to molarity with PBS solution. pH series were prepared thusly:

Solution A: 6.995 g NaH2PO4 in 250 mL deionized water

Solution B: 7.09 g Na2HPO4 in 250 mL deionized water

A pH 5.5 buffer was prepared by mixing 93.5 mL of Solution A with 6.5 mL Solution B. A pH 8 buffer was prepared by mixing 94.7 mL of Solution B with 5.3 mL of Solution A. pH was re- adjusted by titration with NaOH or HCl just prior to measurement.

Imaging of standard solutions:

1μL of a 5mg/mL solution of 2-NBDG was added to each 1g solution just prior to imaging.

Solutions were promptly recapped, gently inverted a few times, transferred to depression slides

60 and immediately covered with a cover slip to prevent evaporation. Confocal imaging with a

Leica SP8 was performed utilizing a white light laser set to 470 nm excitation. Fluorescence was collected in the range of 490-550, with a 1-3 Airy Unit pinhole. FLIM imaging was performed at

80MHz utilizing a Photomultiplier tube (PMT) internal spectral detector channel for 200-1000 counts. Five measurements were made from each solution at two different regions of interest.

Plants:

Arabidopsis plants were grown and imaged in Microscopy Rhizoshpere Chamber (Advanced

Science Tools, Pullman, WA) as described by Froelich et al. (2011). To image phloem, 5-10 μl of a 1:5 mixture of stock 2-NBDG in acetone to deionized water was applied to a slit in the leaf also as described by Froelich et al. (2011). Fluorescence was apparent in the phloem of roots in less than 30 min. The same microscope and settings were used as for the standard curve, with a larger pinhole (3-5 AU). Ipomoea nil plants were prepared and imaged as described by

Knoblauch et al. (2016). To quantify the relationship between lifetime and viscosity, regions of interest (ROIs) in the sieve tube are selected that do not contain peripheral sieve tube contents.

These are analyzed and averaged with SymPhoTime PicoQuant software (Rudower Chaussee 29

(IGZ) 12489 Berlin, Germany). ROIs that provide a single exponential fit, with Chi-squared values less than 1.4 were considered reliable.

Analysis:

The lifetimes of the 2-NBDG in solvents were measured by tail-fitting, or deconvoluted measurements. Both methods provided strong (X2 < 1.4) values with only one 2-NBDG lifetime.

Deconvoluted analysis was performed utilizing the instrument response function-included curve as outlined by Suhling et al. (2012).

Results:

Viscosity effect on lifetime of 2-NBDG

A molecular rotor will rotate less in a more viscous solution, resulting in a longer fluorescence lifetime. The fluorescence lifetime of 2-NBDG increases both as a function of sucrose-induced viscosity, and glycerol-induced viscosity (Figure 1). At viscosities between 1 - 5 mPas, glycerol caused similar 2-NBDG lifetime effects as sucrose, but the curves of these two solutes’ effect on lifetime diverged at higher viscosities. At viscosities higher than 10mPas, sucrose solutions’ effects on lifetime of NBDG begin to saturate, differing less linearly as a function of viscosity, but glycerol solutions were able to induce increasing lifetimes at viscosities up to 1,000 mPas, although with broader deviation, and may be useful for analyzing higher-viscosity fluids. In both solvents, the measured lifetimes of phloem sap fell within the linear range, differing by only about 20% if calculated by separated curves. Viscosity of a 1 - 10.3 mPas viscosity solution was described by the equation 2-NBDG lifetime = 0.318ln(viscosity) + 1.2059, as derived from the standard curve (Figure 1).

All measurements were made at ambient temperature (20+ °C) and the standard solutions were mixed at concentrations that correspond to viscosities at 20°C presented in tables of

Swindells (1958).

Sucrose (•) Glycerol (Δ) 4

3.5

y = 0.318ln(x) + 1.2059 2.5 Phloem sap R² = 0.9847 lifetime ~1.37 ns

NBDG Lifetime NBDG Lifetime (ns)

- 2 2 Calculated phloem sap viscosity ~ 1.67 mPas 1.5

1 0.9 9 90 900 Viscosity (mPa s)

Fig 1. Fluorescence lifetime of 2-NBDG is a function of solvent viscosity. Lifetime values for 2-

NBDG as a function of viscosity imposed by glycerol were higher than those obtained from sucrose solvents at increasing viscosities, but were similar at low viscosities, such as those expected in phloem. For all points, n ≥10, (bars represent standard deviation). The viscosity of the phloem sap of both in vivo juvenile A. thaliana, and I. nil was determined to be in the range of 1.7 mPas utilizing this equation (n=8, stdev=0.003). pH and ionic strength effects on behavior of 2-NBDG lifetime

The pH of phloem sap has been reported as 8.02 in Ricinus (Hall and Baker 1972), and is presented as ranging in various plants from 7.3-8.5 by Dinant et al. (2010). Because Arabidopsis uses proton symport to load sucrose into the phloem, a gradient of protons and charge favoring

63 the extracellular environment is actively maintained. It is therefore likely that the pH of

Arabidopsis phloem sap is slightly alkaline. However, in citrus, where the phloem sap contains organic acids, an acidic pH of 6.03 phloem sap has been reported (Hijaz and Killiny 2014). It is therefore important to understand how the behavior or 2-NBDG is affected by pH. At pH 5.5, lifetime values were higher than they were in PBS standard (pH 7.2) in both 0% and 21% sucrose solutions. High pH (8) slightly decreased lifetime measurements at 0% sucrose, compared to pH 7.2 PBS standards, and did not significantly affect lifetime measurements in

21% sucrose solutions. The Hedges’ effect size is a method that quantifies the comparative strength of relationships. In these experiments, Hedges’ effect size of lifetime increase at 21% sucrose compared to 0% at pH 7.2 was 6.533. At pH 5.5, the Hedges’ effect value comparing the two concentrations was 6.73. The effect of viscosity is more than 4 times greater than the effect of reduced pH (1.367 at 0%, and 1.567 21% sucrose).

1.6

1.5

1.4

1.3 0% suc

21% suc

NBDG lifetime NBDG lifetime (ns) - 2 1.2

1.1

1 pH 5.5 pH 7.2 pH8 Figure 2: Low pH of solvent has a slight effect on lifetime of 2-NBDG. The effect of low pH on fluorescence lifetime of 2-NBDG was much smaller in magnitude than the effect of solute concentration. For all means, n ≥ 20. Bars represent standard deviation.

The effect on 2-NBDG lifetime of 0.5 M NaCl PBS solution was not significant compared to that of lifetimes obtained in PBS alone. The effect of 1.0 M NaCl solution was significant at 95% confidence level compared to the dye in PBS alone. Lifetime measurements of 2-NBDG in obtained from the dye in a 3.0 M solution of NaCl in PBS that would have an expected viscosity of 1.34 mPas at 20°C (Kestin et al. 1981) were 10% lower than expected, however the lifetime values at this concentration were higher than PBS alone (p=0.03) It is possible that high ion concentrations affect lifetime values of 2-NBDG in a non-linear manner (Figure 3).

NaCl effect on 2-NBDG lifetime 2

1.9

1.8

1.7

1.6

1.5 y = 0.2798ln(x) + 1.199

NBDG lifetime (ns) 1.4 R² = 0.9915 -

2 0.5M NaCl 1.3 3M NaCl 1.2 1M NaCl 1.1 0.9 1.34 2 3 5 9 calculated viscosity (mPas)

Figure 3: Lifetimes of 2-NBDG in 0.5 to 3M of NaCl in PBS. 2-NBDG in ionic solutions returned lifetime values (red markers) in the range of standard deviation for PBS alone compared to a 1-

10 mPas sucrose-induced viscosity gradient. Ionic strength has only a minimal effect on lifetime of 2-NBDG. All data points represent a mean from ≥10 measurements. Bars are standard deviation.

Effect of soluble protein on behavior of 2-NBDG

Phloem sap has been measured to contain 0.1mg/ml of soluble proteins by Fisher and Frame

(1984). The presence of this concentration of BSA increased lifetime measurements by 3.56% when added to a 21% sucrose solution, and did not significantly affect lifetime measurements when added to 29% (w/w) sucrose solutions.

1.55 BSA effect on NBDG lifetime (ns)

1.5

1.45

1.4

1.35

1.3 2.031 mPas 3.015 mPas w/bsa no bsa Figure 4: Lifetimes of 2-NBDG in sucrose solutions containing 0.1% BSA. The presence of similar concentrations of BSA as soluble protein expected in phloem sap had a small effect on fluorescence lifetime of 2-NBDG in a 2 mPas sucrose solution, and no effect in a 29% sucrose solution. Grey bars represent lifetimes measured in sucrose solutions containing 0.1% BSA. n=4 for both with bsa measurements, and n ≥ 30 for without BSA measurements. Bars are standard deviation.

Intensity and lifetime imaging of living phloem

Phloem in the roots of juvenile Arabidopsis thaliana was labeled with 2-NBDG by loading through the leaves. Ipomoea nil source leaf phloem was loaded by upstream labeling of the phloem by pressure injection of dye through the stomata into the leaf. The average

66 fluorescence lifetime of 2-NBDG in source phloem in morning glory was 1.366 ns (± 0.037 ns, n

= 8), which corresponds to a viscosity of 1.7 mPas or a sucrose concentration of about 18%

(Knoblauch et al. 2016). Results were very similar for Arabidopsis root phloem, where measured lifetimes at four positions in each of 2 different plants corresponded to viscosity values of 1.677 mPas (n = 8, ster = 0.003). Measurements obtained by tail-fitting single exponential curves with

Chi-squared values less than 1.4. The equation listed on the calibration curve presented in

Figure 1 was used to calculate viscosity from lifetime data.

A ST CC 10 μm 10 um B

Arabidopsis root phloem

ST CC

Ipomoea leaf phloem

Figure 5: 2-NBDG intensity and lifetime maps in the sap of living plants. Arabidopsis (top) and

Ipomoea (bottom). Confocal intensity mapping images are (A), and lifetime map of the same viewing frames are (B). Cytoplasm in companion cells (CC) and peripheral cell organelles and proteins in sieve elements (SE) show higher fluorescence intensity and longer lifetimes than sieve tubes (ST). In the lifetime map (B), blue indicates low relative viscosity, red and yellow high viscosity, and green is intermediate. Solid arrows point to sieve plates. Dashed arrows indicate plastids. I. nil image from Knoblauch et al. 2016

DISCUSSION:

2-NBDG probably enters the root phloem symplastically, starting in the sieve elements of the minor veins that are disrupted at the cut site where it is applied, and not via transporters, since sieve elements contain very little hexose, sieve elements have no vacuoles, and SUC transporters do not transport 2-NBDG (Knoblauch et al. 2015). The higher intensity fluorescence in the companion cells’ cytoplasm (Figure 5) could be caused by the higher concentration of the dye in this compartment, due to movement from the sieve tube through glucose transporters, a higher viscosity in cytoplasm, or cytoskeletal elements in the cytoplasm resulting in inhibited rotation, and therefore higher intensity fluorescence than in the vacuoles and sieve tubes, or a combination of factors. Peripheral organelles and proteins in sieve elements also display higher fluorescence. This is likely due to hindered rotational energy loss in these domains. Vacuoles of CCs and bulk contents of SEs present lower intensity (Figure 5).

Lifetime measurements are more useful to map viscosity than intensity, because they are independent of dye concentration (Kuimova 2012). Lifetime analysis of phloem sap returns only

68 one exponential decay rate, indicating a homogenous fluid. This single exponential function correlates to a viscosity comparable to a sucrose concentration of about 18% w/w.

Imprecision of measurements for the standard curves, and in vivo phloem is presented by temperature. All measurements were obtained at ambient temperature, which was measured between 20-22°C. These small temperature fluctuations can affect viscosity of standards and therefore the curve utilized to calculate viscosity of phloem sap. The difference between calculated viscosities of phloem sap at 20, and 22°C is about 7% (1.67 mPas at 20°C, and 1.56 at

22°C). Exposure to the laser can also cause temperature fluctuations.

Lifetime measurements of 2-NBDG in phloem sap correlate with viscosities that are 30% higher than can be explained by the 11% sucrose concentration reported by Deeken et al. (2002).

These concentrations may change as a function of changing photosynthetic rates during growth of the plant, and other factors. It has been reported that Arabidopsis thaliana phloem sap also contains free amino acids (Zhang et al. 2010), and phloem sap of multiple plant species contains protein concentrations in the range of 100-400 ng/μL (Hayashi et al. 2000). The average sucrose:amino acid ratio in phloem sap exudate in petioles of potato was measured to be 4.28

(Karley et al. 2002). Hijaz and Killiny (2014) measured up to 32% of phloem sap solutes as amino acids in Citrus sinensis. In phloem sap of Brassica carinata, 500mM concentrations of amino acids were measured by Lohaus and Moellers (2000), and a threefold variation in amino acid concentrations across 3 species was reported by Lohaus and Schwerdtfeger (2014), while carbohydrate phloem sap concentrations only varied by about 50%. Previously measured values for solute concentrations and ratios have varied by species and developmental stage. Variation is also demonstrated by different methods of collection and analysis. Because it has not been

69 possible to distinguish whether these variations are caused by physiology or methodology, this new method, which rapidly and directly measures phloem sap viscosity in vivo is a valuable addition to the study of phloem physiology.

It is interesting also to note that the measurements obtained for viscosity in roots of A. thaliana had little variability. While pressure in phloem was shown to be a function of position, viscosity measurements in source leaves of Morning glory (Knoblauch et al. 2016) obtained by FLIM were very similar to the measurements obtained from roots of A. thaliana. Additionally, FRAP analysis of multiple plants at multiple positions (Chapter 2) suggest that diffusivity is not a function of position.

Münch flow requires an osmotic potential differential to drive transport. It is often posited that this potential differential is a function of sucrose concentration differences between source and sink. By eliminating oxygen supply from source leaves, and thereby reducing the capacity of active sucrose loading, Gould et al. (2005) were able to stimulate both a reduced osmotic potential, and a reduced sucrose concentration in sieve tubes of mature barley leaves by similar magnitudes. Because the decrease in sucrose concentration and measured velocity reduction were concurrent, they conclude that the most important osmolyte in phloem sap is sucrose.

This may however not be so straightforward. Sucrose loading is tightly tied to the expression of potassium channels, and a change of sucrose concentration in phloem affects this ion’s concentration (Deeken et al. 2002; Dreyer and Uozumi 2011). Results from transport in plants with diminished expression of AtSUC transporters show that although sucrose loading is significantly reduced, translocation continues at a lower velocity (Srivistana et al. 2008).

Additionally, phloem sap transport velocities in Maize SUT2 mutant plants are only halved,

70 while phoem loading and transport of sucrose is steeply diminished (Babst 2016, pc). These findings are interesting to consider in tandem with the results of this study. While it is likely that a concentration differential driving flow in roots of such small and young plants as those used in this study would not be detectable by this method, it is unlikely that source leaves of a 3 meter plant would have the same sucrose concentration as phloem of roots, if indeed a sucrose concentration differential is driving flow. Because they are strong osmolytes, and do not present high viscosity, the role of salts in maintaining osmotic potential differentials in plant phloem may be very important for maintenance of a pressure differential (Lang 1983).

Knoblauch et al. (2016) measured a viscosity in source leaf phloem of 1.7. This viscosity corresponds to a sucrose concentration of about 20%. This concentration of sucrose would only provide about 1.22 MPa of osmotic pressure, but pressure in this part of the sieve tube system was measured as 2.2 MPa. If the rest of the pressure were supplied by ions, it would require

200mM of a monovalent cation to achieve this osmotic pressure. Other small solutes, such as amino acids may contribute to osmotic potential in phloem as well. Ionic concentrations in phloem sap have been measured at values in the range of 40-150mM, and occur in a top-to- bottom, source-to-sink gradient in Ricinus (Vreugdenhil and Koot-Gronsveld 1989). 2-NBDG in

1M and 3M solutions of NaCl returned lifetime values that were lower than expected by the equation of the standard curve, but still in the range of standard deviation of PBS alone.

Solutions of 500mM NaCl in PBS were not significantly different from PBS alone. The presence of small ions is not accurately detectable by this method, presumably because they do not have a large enough effect on viscosity.

There is a tradeoff when considering energy delivery to sinks. Increasing sucrose concentrations will deliver more energy per unit volume, but will come at a cost of increased viscosity. More viscosity will increase the needed pressure differential, which costs energy in apoplastic loaders and reduces the speed of transport. Incremental sucrose concentrations begin to increase viscosity more steeply after about 20% (w/w). Simulations conducted by Hölttä et al. (2005) suggest that too much sucrose loading at sources can even result in viscosity increases that would severely impede flow. According to Van ’t Hoff calculations, similar osmotic potential could be delivered to source phloem by loading salt ions with very little viscosity cost, thereby causing high hydrostatic pressure without reducing flow velocity. (Equation 1 and Figure 6)

Ψs = -i*c*R*T Equation 1

Where i is the Van’t Hoff factor, c is the molar concentration, R is the pressure constant, and T is the temperature in Kelvins.

6 Solute potential/viscosity

3 μ suc μ NaCl 2

Viscosity Viscosity (mPas) 1

0 0 2 4 6 8 10 12 |Ψs| (MPa)

Figure 6: Relationship between osmotic potential and viscosity. Sucrose (•) and NaCl (Δ) at

20°C. Viscosity values from Swindells (1958), and Kestin et al. (1981). The exponential increase of viscosity becomes steeper at about 1.7 mPas, providing little additional solute potential.

Solute potential continues to increase with higher concentrations of salts, while imparting only small, linear increases in viscosity.

It is important to consider that the phloem is not only tasked with transport of energetic molecules, but also serves as a signaling pathway, delivering hormones, RNA molecules, and proteins to sinks (Lough and Lucas 2006). Therefore, this microfluidic biological system may also be optimized for speed. Modeling results utilizing a compilation of plant data deliver a mathematical scaling law between radius, and path lengths of source and sink phloem that strongly support this hypothesis in a broad range of plant species (Jensen et al. 2011).

A compilation of phloem sap data, when utilized in mathematical pressure flow models support a pressure flow mechanism that is driven under constant pressure, rather than one that is driven by a pressure that is proportional to the sucrose concentration (Jensen et al. 2013).

Constant pressure in plant phloem, under this perspective, would be able to be maintained by modulation of potassium ion concentration in the sieve tubes. This would also explain the maintenance of velocity in the sieve tubes, even under the conditions of impaired sucrose loading. Results from this study, indicating a low viscosity of phloem sap in sieve tubes, would support high velocity translocation in sieve tubes.

2-NBDG functions as a molecular rotor, presenting a clear relationship between its lifetime, and the viscosity of the solvent it exists in. Its phloem-mobility allows direct measurement in-vivo of

73 translocating phloem sap. Future efforts may include comparing different plant species, different treatment regimes, and different genetic constructs. This new method adds a valuable tool to the pursuit of understanding phloem translocation in plants, and may prove highly useful for the rapid measurement of other biofluids as well.

Works Cited: Animireddy, Dwitha, Venkata Thimma Reddy Bekkem, Pranitha Vallala, Sunil Babu Kotha, Swetha Ankireddy, and Noorjahan Mohammad. "Evaluation of pH, buffering capacity, viscosity and flow rate levels of saliva in caries-free, minimal caries and nursing caries children: An in vivo study." Contemporary clinical dentistry 5, no. 3 (2014): 324. Aras, S., I. Tek, M. Varli, A. Yalcin, O. K. Cengiz, V. Atmis, and T. Atli. "Plasma Viscosity: Is a Biomarker for the Differential Diagnosis of Alzheimer's Disease and Vascular Dementia?" American Journal of Alzheimer's Disease and Other Dementias 28, no. 1 (2012): 62-68. Babst, Benjamin. “Plant Resource utilization is crucial during growth and stress responses: How is it controlled?” p.c. Invited talk for Molecular Plant Science Program. Jan 27th, 2016. Washington State University. Deeken, Rosalia, Dietmar Geiger, Jörg Fromm, Olga Koroleva, Peter Ache, Rosemarie Langenfeld-Heyser, Norbert Sauer, Sean T. May, and Rainer Hedrich. "Loss of the AKT2/3 potassium channel affects sugar loading into the phloem of Arabidopsis." Planta 216, no. 2 (2002): 334-344. Dinant, Sylvie, Jean-Louis Bonnemain, Christine Girousse, and Julia Kehr. "Phloem Sap Intricacy and Interplay with Aphid Feeding." Comptes Rendus Biologies 333, no. 6-7 (2010): 504-15. Dreyer, Ingo, and Nobuyuki Uozumi. "Potassium Channels in Plant Cells." FEBS Journal 278, no. 22 (2011): 4293-303. Ellison, Aaron M., and Nicholas J. Gotelli. "Evolutionary ecology of carnivorous plants." Trends in ecology & evolution 16, no. 11 (2001): 623-629. Etxeberria, Ed, Pedro González, Patricia Tomlinson, and Javier Pozueta-Romero. "Existence of two parallel mechanisms for glucose uptake in heterotrophic plant cells." Journal of Experimental Botany 56, no. 417 (2005): 1905-1912. Evert, Ray Franklin., Katherine Esau, and Katherine Esau. Esau's Plant Anatomy: Meristems, Cells, and Tissues of the Plant Body: Their Structure, Function, and Development. Hoboken, NJ: Wiley-Interscience, 2006. Fisher, Donald B., and John M. Frame. "A Guide to the Use of the Exuding-stylet Technique in Phloem Physiology." Planta 161, no. 5 (1984): 385-393. Froelich, Daniel R., Daniel L. Mullendore, Kåre H. Jensen, Tim J. Ross-Elliott, James A. Anstead, Gary A. Thompson, Hélène C. Pélissier, and Michael Knoblauch. "Phloem ultrastructure and pressure flow: sieve-element-occlusion-related agglomerations do not affect translocation." The Plant Cell23, no. 12 (2011): 4428-4445. Gessler, Arthur, Guillaume Tcherkez, Andreas D. Peuke, Jaleh Ghashghaie, and Graham D. Farquhar. "Experimental Evidence for Diel Variations of the Carbon Isotope Composition in Leaf, Stem and Phloem Sap Organic Matter in Ricinus Communis." Plant, Cell & Environment 31, no. 7 (2008): 941-953.

Giavalisco, Patrick, Kristin Kapitza, Anna Kolasa, Anja Buhtz, and Julia Kehr. "Towards the Proteome Of Brassica Napus Phloem Sap." Proteomics 6, no. 3 (2006): 896-909. Gould, Nick, Michael R. Thorpe, Olga Koroleva, and Peter E. H. Minchin. "Phloem Hydrostatic Pressure Relates to Solute Loading Rate: A Direct Test of the Münch Hypothesis." Functional Plant Biology 32, no. 11 (2005): 1019. Hall, Shelagh M., and D. A. Baker. "The Chemical Composition of Ricinus Phloem Exudate." Planta 106, no. 2 (1972): 131-40. Hayashi, Hiroaki, Akari Fukuda, Nobuo Suzui, and Shu Fujimaki. "Proteins in the Sieve Element- companion Cell Complexes: Their Detection, Localization and Possible Functions." Functional Plant Biology 27, no. 6 (2000): 489. Helden, Maarten Van, W. Freddy Tjallingh, and Teris A. Van Beek. "Phloem Sap Collection from Lettuce (Lactuca Sativa L.): Chemical Comparison among Collection Methods." Journal of Chemical Ecology 20, no. 12 (1994): 3191-206. Hijaz, Faraj, and Nabil Killiny. "Collection and chemical composition of phloem sap from Citrus sinensis L. Osbeck (sweet orange)." PloS one 9, no. 7 (2014): e101830. Hofmann, Julia, Paul H. Hess, Dagmar Szakasits, Andreas Blöchl, Krzysztof Wieczorek, Sabine Daxböck-Horvath, Holger Bohlmann, Aart JE van Bel, and Florian MW Grundler. "Diversity and activity of sugar transporters in nematode-induced root syncytia." Journal of experimental botany 60, no. 11 (2009): 3085-3095. Hölttä, Teemu, Timo Vesala, Sanna Sevanto, Martti Perämäki, and Eero Nikinmaa. "Modeling Xylem and Phloem Water Flows in Trees According to Cohesion Theory and Münch Hypothesis." Trees 20, no. 1 (2005): 67-78. Jensen, Kaare H., Jessica A. Savage, and N. Michele Holbrook. "Optimal concentration for sugar transport in plants." Journal of The Royal Society Interface 10, no. 83 (2013): 20130055. Jensen, Kåre H., J. Lee, Tomas Bohr, Henrik Bruus, Noel M. Holbrook, and Maciej Zwieniecki. "Optimality of the Münch mechanism for translocation of sugars in plants." Journal of the Royal Society Interface 8, no. 61 (2011): 1155-1165. Kao, H. Pin, James R. Abney, and Alan S. Verkman. "Determinants of the translational mobility of a small solute in cell cytoplasm." The Journal of cell biology 120, no. 1 (1993): 175-184. Karley, Alison J., Angela E. Douglas, and William E. Parker. "Amino acid composition and nutritional quality of potato leaf phloem sap for aphids." Journal of Experimental Biology 205, no. 19 (2002): 3009-3018. Kehr, Julia. "Phloem sap proteins: their identities and potential roles in the interaction between plants and phloem-feeding insects." Journal of Experimental Botany 57, no. 4 (2006): 767-774. Kennedy, John S., and Thomas E. Mittler. "A Method of Obtaining Phloem Sap via the Mouth- parts of Aphids." Nature 171, no. 4351 (1953): 528.

Kestin, Joseph, H. Ezzat Khalifa, and Robert J. Correia. "Tables of the Dynamic and Kinematic Viscosity of Aqueous NaCl Solutions in the Temperature Range 20–150 °C and the Pressure Range 0.1–35 MPa." Journal of Physical and Chemical Reference Data 10, no. 1 (1981): 71. Ketene, Alperen N., Eva M. Schmelz, Paul C. Roberts, and Masoud Agah. "The effects of cancer progression on the viscoelasticity of ovarian cell cytoskeleton structures." Nanomedicine: Nanotechnology, Biology and Medicine 8, no. 1 (2012): 93-102. Knoblauch, Michael, Jan Knoblauch, Daniel L. Mullendore, Jessica A. Savage, Benjamin A. Babst, Sierra D. Beecher, Adam C. Dodgen, Kaare H. Jensen, and Noel Michele Holbrook. "Testing the Münch hypothesis of long distance phloem transport in plants." eLife 5 (2016): e15341. Knoblauch, Michael, and Winfried S. Peters. "Long-distance Translocation of Photosynthates: A Primer." Photosynth Res Photosynthesis Research 117, no. 1-3 (2013): 189-196. Knoblauch, Michael, Marc Vendrell, Erica De Leau, Andrea Paterlini, Kirsten Knox, Tim Ross- Elliot, Anke Reinders, Stephen A. Brockman, John Ward, and Karl Oparka. "Multispectral Phloem-Mobile Probes: Properties and Applications." Plant Physiology 167, no. 4 (2015): 1211- 1220. Kuimova, Marina K. "Mapping Viscosity in Cells Using Molecular Rotors." Physical Chemistry Chemical Physics 14, no. 37 (2012): 12671-12686. Kuimova, Marina K., Stanley W. Botchway, Anthony W. Parker, Milan Balaz, Hazel A. Collins, Harry L. Anderson, Klaus Suhling, and Peter R. Ogilby. "Imaging Intracellular Viscosity of a Single Cell during Photoinduced Cell Death." Nature Chemistry 1, no. 1 (2009): 69-73. Lang, Alexander. “Turgor-related translocation.” Plant, Cell, and Environment 6, no. 9 (1983): 683-689. Levitt, James A., Pei‐Hua Chung, Marina K. Kuimova, Gokhan Yahioglu, Yan Wang, Junle Qu, and Klaus Suhling. "Fluorescence anisotropy of molecular rotors." ChemPhysChem 12, no. 3 (2011): 662-672. Lohaus, Gertrud, and Christian Moellers. "Phloem Transport of Amino Acids in Two Brassica Napus L. Genotypes and One B. Carinata Genotype in Relation to Their Seed Protein Content." Planta 211, no. 6 (2000): 833-840. Lohaus, Gertrud, and Michael Schwerdtfeger. "Comparison of Sugars, Iridoid Glycosides and Amino Acids in Nectar and Phloem Sap of Maurandya Barclayana, Lophospermum Erubescens, and Brassica Napus." PLoS ONE 9, no. 1 (2014). Lough, Tony J., and William J. Lucas. "Integrative Plant Biology: Role of Phloem Long-Distance Macromolecular Trafficking." Annual Review of Plant Biology 57, no. 1 (2006): 203-232. Luby-Phelps, Katherine, Swati Mujumdar, Ratnakar B. Mujumdar, Lauren A. Ernst, William Galbraith, and Alan S. Waggoner. "A Novel Fluorescence Ratiometric Method Confirms the Low Solvent Viscosity of the Cytoplasm." Biophysical Journal 65, no. 1 (1993): 236-242. Münch, Ernst. "Stoffbewegungen in der Pflanze." (1930).

Mullendore, Daniel L., Carel W. Windt, Henk Van As, and Michael Knoblauch. "Sieve Tube Geometry in Relation to Phloem Flow." The Plant Cell 22, no. 3 (2010): 579-593. O’Neil, Roger G., Ling Wu, and Nizar Mullani. "Uptake of a Fluorescent Deoxyglucose Analog (2- NBDG) in Tumor Cells." Molecular Imaging and Biology 7, no. 6 (2005): 388-392. Raven, J. A., and Dianne Edwards. "Roots: evolutionary origins and biogeochemical significance." Journal of Experimental Botany 52, no. suppl 1 (2001): 381-401. Segur, John Bartlett, and Helen E. Oberstar. "Viscosity of glycerol and its aqueous solutions." Industrial & Engineering Chemistry 43, no. 9 (1951): 2117-2120. Srivastava, Avinash C., Savita Ganesan, Ihab O. Ismail, and Brian G. Ayre. "Functional characterization of the Arabidopsis AtSUC2 sucrose/H+ symporter by tissue-specific complementation reveals an essential role in phloem loading but not in long-distance transport." Plant Physiology 148, no. 1 (2008): 200-211. Suhling, Klaus, James A. Levitt, Pei-Hua Chung, Marina K. Kuimova, and Gokhan Yahioglu. "Fluorescence lifetime imaging of molecular rotors in living cells." Journal of Visualized Experiments 1, no. 60 (2012): e2925. Swaminathan, Rajaram, S. Bicknese, N. Periasamy, and Alan Verkman. "Cytoplasmic Viscosity near the Cell Plasma Membrane: Translational Diffusion of a Small Fluorescent Solute Measured by Total Internal Reflection-fluorescence Photobleaching Recovery." Biophysical Journal 71, no. 2 (1996): 1140-151. Swindells, James F. “Viscosities of sucrose solutions at various temperatures: tables of recalculated values.” USGPO. Vol. 440. (1958). Verkman, Alan S. "Solute and Macromolecule Diffusion in Cellular Aqueous Compartments." Trends in Biochemical Sciences 27, no. 1 (2002): 27-33. Vreugdenhil, Dick, and Elly A.M. Koot‐Gronsveld. "Measurements of pH, sucrose and potassium ions in the phloem sap of castor bean (Ricinus communis) plants." Physiologia Plantarum 77, no. 3 (1989): 385-388. Yoshioka, Kazuaki, Hirokazu Takahashi, Tomoo Homma, Mikako Saito, Ki-Bong Oh, Yasushi Nemoto, and Hideaki Matsuoka. "A Novel Fluorescent Derivative of Glucose Applicable to the Assessment of Glucose Uptake Activity of Escherichia Coli." Biochimica Et Biophysica Acta 1289, no. 1 (1996): 5-9. Zhang, Lizhi, Qiumin Tan, Raymond Lee, Alexander Trethewy, Yong-Hwa Lee, and Mechthild Tegeder. "Altered xylem-phloem transfer of amino acids affects metabolism and leads to increased seed yield and oil content in Arabidopsis." The Plant Cell 22, no. 11 (2010): 3603- 3620.

Chapter 4: Angiosperm Sieve Tube Conductivity

Sierra Beecher1, Jessica Savage2 and Michael Knoblauch1 1. Washington State University, School of Biological Sciences 2. Harvard University, Department of Organismic and Evolutionary Biology

Abstract:

Calculating the conductivity of sieve tubes requires knowledge of the tube diameter, periodicity and architecture of sieve plates, and length of the sieve elements and the entire sieve tube.

Here we present data that provides sieve tube diameters, sieve element lengths, and sieve plate angles at three different in-planta locations for 22 plant species; 8 trees, 9 vines, 3 herbaceous plants, and 2 shrubs. In trees, the long tube lengths and low source pressures measured in leaves have cast doubt on pressure flow as a possible mechanism of transport.

Rather than a single number, conductivity factors measured in this study are arranged in a gradient, increasing significantly in the trunk. These data indicate that conductivity has been underestimated in trees, and that Münch pressure flow is sufficient to drive sap through even tall trees. Further analysis of sieve plates will be needed to strengthen these conclusions.

Introduction:

A strong understanding of phloem physiology is an important part of the toolbox needed to take adaptive and mitigative measures in the face of the myriad of global changes we face.

Drought conditions increase herbivory on trees by phloem-feeding insects, resulting in whole community structure changes (Kozlov et al. 2015). Forest losses increase carbon flux into the atmosphere (DeFries et al. 2002), reduce soil moisture, disturb whole watersheds, impact the

79 water cycle (Vörösmarty and Sahagian 2000), and are associated with large-scale habitat shrinkage and biodiversity reduction.

Although understanding the phloem of trees is critical for successful climate change action, even the physics of tree phloem is in question. While data available for the phloem of small herbaceous plants largely support the dominant Münch Hypothesis of pressure driven flow as a means to move photoassimilate from sources to sinks (Magnuson et al. 1986), the question is not settled in trees (Thompson and Holbrook 2003; Turgeon 2010). Furthermore, it is not understood how changes in plant anatomy may affect phloem conductivity in herbaceous plants, many of which provide food for our growing population.

Pressure driven flow is the overarching hypothesis about how plants move photoassimilates from sources, such as leaves where they are produced, to sinks, such as roots where they are stored. The hypothesis states that in source regions, the phloem contains a large concentration of sugars. These sugars pull water into sieve tubes by osmosis, building a hydrostatic pressure that drives sap to low pressure regions of the plant where solute concentrations are low. It is a very logical and attractive idea, with experimental support in herbs (Magnuson et al. 1986), but in trees, not all data has seemed to support it (Turgeon 2010; Ryan and Asao 2014).

Mathematical analysis of pressure driven flow in a tube demands that the length of the tube scales linearly with the pressure differential (Eq. 2, Chapter 1). Conductivity has been modeled to decrease with the square of the plant length (Thompson and Holbrook 2003). Therefore, in trees, where length is at least an order of magnitude longer than in herbaceous plants, one would expect to measure higher pressure in source phloem of trees than in herbs, and pressure

80 should scale to length. Although pressure has been shown to scale with length in Morning glory

(Knoblauch et al. 2016), the opposite has been shown to be the case in trees (Hammel 1968;

Wright and Fisher 1980). Furthermore, many trees examined have proven to be passive loaders, with extensive symplastic connections to mesophyll cells, rather than utilizing energetic transporters, which could isolate sap, and maintain a stronger uphill concentration gradient (Fu et al. 2011). Some measurements for sieve tube pressure even put phloem in the range of osmotic equilibrium with nearby cells (Pritchard 1996), calling to question the ability of any pressure being available to drive flow through sieve tubes (Turgeon 2010). Sugar concentration gradients have also not been measured to be sufficient to support a pressure gradient in trees that could drive flow over such a long distance (Zimmerman 1957).

This has led to a re-evaluation of sugar transport in trees by some workers. Lang (1979), suggested that a relay mechanism would resolve these questions. This mechanism assumes that the sieve tube length is not defined by the length of the plant, but rather that several overlapping sieve tubes exist in tandem. At each overlap, loading occurs into the next tube, giving each new section a boost of pressure. The discovery of leakage and retrieval mechanisms in transport phloem led some workers to consider that relays may exist even in each sieve element, with pressure building up at each adjoining plate in sieve tubes (De Schepper et al.

2013). Thompson and Holbrook have demonstrated that this would be mathematically favorable (2003), however, anatomical verification has not been presented. A pressure- concentration wave mechanism, rather than pressure driven flow has also been proposed

(Thompson 2006). This model states that rather than a strong concentration gradient building a hydrostatic pressure differential that “pushes” sap, the phloem may maintain a fairly even

81 concentration of sugars, with sink demand ultimately “pulling” resources, and controlling transport, and even photosynthesis (Thompson 2006). This situation, in which energetic molecules are similarly available at all sinks is compared to a delivery manifold, and requires the pressure differential to be small (Fisher and Cash-Clark 2000). This would explain the relatively low pressures measured in source phloem of trees.

Although pressure has not been demonstrated to scale to length in angiosperm tree leaves, other models, based largely on phloem anatomy provide scaling support for traditional Münch pressure flow in trees (Jensen et al. 2012). A high degree of conductivity in transport phloem is estimated based on large sieve element diameters in angiosperm transport phloem. Because larger diameters and overall phloem area scale to higher velocities compared to gymnosperms, which have higher resistance imposed by their smaller sieve element cross sectional areas, and cuboid geometries, the Munch Hypothesis is supported by this analysis. Sieve element cross sectional area measurements scale to stem length in a manner that is predicted by the Jensen et al. (2012) model.

It is important when assessing sieve tube conductivity to consider that geometries are not anatomically similar throughout the plant. Auxin gradients result in cell differentiation patterns that yield increasingly wider xylem cells at increasing distances from new leaves (Aloni 1987).

Additionally, phytochrome activity is sensitive to light and its effects on cell growth are modulated by brassinosteroids (Azpiroz et al. 1998). These effects have not been demonstrated specifically for phloem, but have presented in xylem, which is also constructed from the vascular cambium, and are therefore relevant. Genetic constructs have been developed that modulate cell elongation based on this system (Nomura et al. 1997).

The dataset presented here will be enhanced greatly when integrated with the upcoming collaborative dataset from Jessica Savage et al. that will provide other conductivity factors including sieve plate porosity from SEM analysis, and will complete this anatomical analysis of comparative phloem conductivity. Savage has designed a large scale phylogenetic and ecotope study of phloem and xylem anatomy, of which the results presented here are a part of.

Materials and Methods:

Sample collection: Samples were collected from Harvard Forest and other American Northeast sites by Jessica

Savage. For each plant, vascular tissue samples were collected from the top, middle, and bottom. The height of the whole plant was recorded, as well as the distance from the ground at each sample collection locale. These were stored in 70% ethanol and refrigerated at 4°C.

Sample preparation: For vines and herbs, all of the samples were whole stem sections of 2-10 cm. In trees, the top samples were ~ 10 cm sections of branches, and these were also stored as stem samples. For middle and bottom sections of trees, cores of up to 5cm in diameter, and 5cm deep that contained outer bark, cork, phloem, cambium, and some xylem were obtained and analyzed. In order to make hand sections, the samples were manipulated as described in Figure 1.

CC CC C X

VC CC

C B

Outer bark Cork

Vascular Plane of first cut. Sectioning proceeded upward to phloem. Cambium Phloem The cambium is softer, indicating proximity to phloem. Here thinner sections were Xylem made and analyzed until aniline blue signature of sieve plates was detected.

Figure 1: Schematic of sample preparation. Stems were cut roughly in half as shown in (A), and hand sectioned longitudinally with a razor. Cores were sectioned in the manner described in

(B).

Hand sections were placed on a slide, and immersed in a solution of aniline blue and calcofluor white, with a cover slip placed on top. The counterstain solution was mixed such that it contained 100 mM KCl, 10mM CHES buffer, and 0.1% Aniline blue in double deionized water. To this, 20mL of a 3.5mg/ml stock solution of calcofluor white in deionized water was added, and the pH was adjusted to 10.

Imaging: The greater part of imaging was performed on a Zeiss 510 META confocal laser scanning microscope. Excitation of both fluorophores was achieved with a 405nm UV laser. Collection of aniline blue (which stains callose) emission was set at 480nm with a longpass filter, and calcofluor white, (which stains cellulose), at 420nm with a bandpass. False-color was employed such that emission from aniline blue appears red, and emission from calcofluor white appears blue. Magnification was varied to optimize imaging, with higher magnification used for smaller sieve elements. Autofluorescence in the red is associated with plastids and lignin, but based on anatomy and to some degree hue, sieve plates are easily identifiable. Some of the images for S. lypercum and C. maxima were obtained from a Leica TCS SP8, employing the microscope’s internal Mosaicking stitching program.

High magnification images of sieve elements were obtained with a Leica LCI Plan-Neofluor

25X/10.8 water immersion lens, and stitched together with Adobe Photoshop CC 2014 software to make visualization of multiple sieve elements possible. Example in Figure 2B.

Low magnification imaging with a Plan-Neofluor 10X/0.3 Zeiss lens allowed more sieve elements to be captured in one frame such as in Figure 2A. This method was employed when sectioning was found to be especially straight through the longitudinal plane. Measurements of sieve element length, diameter, and sieve plate angle were done in imageJ in a manner outlined in Figures 6 and 9.

A B

RP SE RP

SP SP

Figure 2: Imaging examples. Calcofluor white stains cellulose blue, and callose, stained by aniline blue appears red. Middle sample of L. Tulipiflora (A). Middle sample of C. speciosa (B).

Sieve plates (SP) Sieve elements (SE) and Ray parenchyma (RP) are indicated. Scale bars in both images are 50μm.

Results: Here we present data from different positions in plants with long phloem path lengths. In trees, factors that define conductivity are increasing as a function of length, not only across species, but in the individual plants. Twigs and branches near the sources have relatively small secondary phloem diameters, but in the trunk, conductivity factors are much higher (Figures 3 and 4). In some trees, they saturate near the bottom of the tree. A relationship between distance and the conductivity factors these methods provide was not apparent in vines observed in this study (See Figure 7). Although the few herbs and shrubs presented show a similar in planta trend as trees, the n values are too low to make a definitive analysis.

SE L SE D SE V angle STER STER SE L/D Stem D D from top Spp Abbv STER D (μm) (μm) (μm^3) (deg) L ang ratio (cm) (m) TIAM/tree T 219.9 15.8 43046.7 63.7 9.4 0.6 3.4 13.9 0.5 0.15 M 268.6 29.4 181740.9 62.8 9.4 1.5 2.0 9.2 51.7 5.7 B 350.7 28.3 219846.4 69.5 9.3 1.1 1.6 12.4 81.2 13.95

CASP/tree T 137.0 13.5 19533.2 15.2 14.7 0.7 1.6 10.2 0.6 0.13

M 195.2 41.6 265593.1 32.1 9.2 1.4 3.2 4.7 NA 12.5 B 145.8 43.9 220161.3 32.6 8.3 2.1 3.1 3.3 69.7 24.25

POTR/tree T 312.4 20.4 102175.2 61.0 14.7 0.8 4.5 15.3 0.9 0.9 M 389.4 33.6 344448.4 72.9 20.9 1.2 2.3 11.6 10.2 5.7 B 388.9 35.7 390217.6 73.5 19.7 2.8 1.8 10.9 19.4 10.6

NYSY/tree T 262.1 12.4 31658.1 62.9 12.3 0.5 5.3 21.1 0.7 0.5 M 429.7 34.4 400018.4 57.8 14.4 1.4 3.2 12.5 11.7 3.4 B 441.3 29.6 303242.6 60.5 23.2 1.0 3.5 14.9 24.2 7.1

LITU/tree T 381.3 16.2 78513.0 NA 21.6 0.7 NA 23.6 0.7 0.6 M 476.7 33.3 415385.1 73.6 15.1 1 3.1 14.3 26.4 10.8 B 563.7 50.6 1131609.3 74.6 37.3 6.5 2.7 11.2 28.7 21.5 ACSA/tree T 95.9 8.2 5077.7 43.3 7.2 0.5 2.6 11.7 0.3 0.05 M 220.6 19.0 62663.4 75.9 10.9 0.6 1.7 11.6 53.8 11 B 269.2 39.1 322386.7 34.3 13.9 1.5 2.9 6.9 70.0 21.55

QURU/tree T 261.7 19.9 81315.1 51.8 15.4 0.6 2.3 13.2 0.7 0.195 M 345.5 48.1 628529.0 58.5 22.9 1.8 1.6 7.2 38.2 14 B 438.2 55.9 1076925.9 58.4 28.6 1.3 2.3 7.8 57.3 26.9

PRSE/tree T 212.0 16.7 46510.4 64.9 9.5 0.5 3.3 12.7 0.4 0.19 M 328.8 37.9 370449.7 54.7 14.0 1.9 2.0 8.7 23.2 11.8 B 312.7 35.3 305187.6 63.0 20.0 0.8 2.1 8.9 30.6 23.4

FAAU/vine T 179.0 23.0 74023.8 20.2 34.9 0.6 1.3 7.8 0.1 1.2 M 238.4 17.3 55807.6 31.0 22.4 0.8 2.7 13.81 0.6 1.5 B 154.4 18.0 39200.0 27.7 27.3 1.1 2.7 8.59 4.0 2.95 HULU/vine T 96.8 15.3 17881.0 31.2 6.0 0.8 1.6 6.3 0.3 0.43 M 137.4 17.5 33168.7 24.4 3.4 0.5 2.1 7.8 0.3 1 B 133.3 15.4 24841.8 29.2 4.9 0.7 1.7 8.7 0.5 1.95 AKQU/vine T 269.5 15.4 50110.2 58.7 18.3 1.0 3.7 17.5 0.4 1.07 M 266.6 28.9 175042.5 57.5 4.6 1.0 2.3 9.2 0.6 1.5 B 290.0 29.7 200660.1 43.7 9.0 1.3 2.4 9.8 0.7 2.98 CLVI/vine T 422.1 20.8 143745.1 46.3 42.8 0.9 3.0 20.3 0.2 0.25 M 365.9 19.4 107856.1 50.5 50.7 0.8 4.7 18.9 0.2 0.95 B 112.9 15.7 21882.7 19.6 4.3 0.4 4.7 7.2 0.4 1.88 CASE/vine T 216.9 15.8 42252.5 0 11.8 0.5 0 13.8 0.1 0.41 M 171.9 23.5 74462.6 8.5 14.7 1.2 2.2 7.3 0.1 1.14 B 106.6 25.1 52538.1 0 6.5 1.2 0 4.3 0.3 2.54 CUMA/vine T 412.9 39.2 498902.4 10.9 19.9 2.1 1.7 10.5 0.5 0.11 M 176.2 14.0 26991.0 14.8 16.4 1.9 3.3 12.6 0.6 0.65 B 184.4 18.8 51104.0 8.8 14.3 1.9 1.6 9.8 0.6 1.28 PHVU/vine T 194.4 19.1 55449.1 14.8 15.1 0.8 2.7 10.2 0.15 0.23 M 262.6 19.0 74508.8 21.4 10.1 0.7 2.5 13.8 0.16 0.7 B 117.0 15.2 21241.9 16.9 2.2 0.6 1.8 7.7 0.59 1.38 SOLY/vine T 206.9 17.8 51213.5 11.0 15.8 0.7 3.2 11.7 0.42 0.22 M 196.1 12.6 24339.2 6.0 21.1 0.6 1.7 15.6 0.54 0.65 B 154.3 21.2 54278.9 11.9 16.0 1.3 2.6 7.3 0.83 1.23 PASP/vine T 274.9 19.3 80343.9 44.4 3.9 0.8 18.7 14.3 0.2 0.35 M 333.7 21.7 123617.5 55.0 2.3 0.9 12.4 15.4 0.19 0.89 B 242.2 17.7 59514.5 66.5 3.1 0.9 12.8 13.7 0.62 1.81 COJA/shrub T 149.9 11.9 16782.6 35.9 8.5 0.5 3.7 12.6 0.46 0.47 M 176.6 13.9 26644.1 24.7 11.0 0.6 2.7 12.8 6 1.12 B 171.6 18.5 46282.0 31.2 4.3 0.6 3.8 9.3 13.1 2.15 DEVE/shrub T 167.5 16.8 37258.3 0.0 11.0 0.8 0.0 10.0 0.25 0.27 M 181.2 19.1 51815.7 0.0 15.7 0.4 0.0 9.5 0.39 0.7 B 145.7 14.5 23883.1 0.0 11.8 0.6 0.0 10.1 0.85 1.35 OEBI/herb T 117.1 9.3 7961.1 17.6 6.3 0.4 1.9 12.6 0.26 0.1 M 156.9 10.4 13200.6 23.5 7.9 0.4 3.9 15.2 0.38 0.36 B 183.7 10.9 17126.6 16.1 19.8 0.4 3.7 16.9 0.62 0.57 DEBI/herb T 135.9 6.0 3794.4 5.8 7.1 0.4 2.3 22.79 0.41 0.12 M 90.8 5.8 2405.4 14.8 2.9 0.4 1.4 15.63 0.38 0.155 B 93.6 10.1 7427.1 0.0 7.1 0.3 0 9.32 0.45 0.51 DACA/herb T 109.8 6.3 3403.3 0.0 9.2 0.3 0 17.48 0.2 0.07 M 190.0 7.2 7748.2 0.0 16.5 0.6 0 26.38 0.25 0.33 B 119.9 7.8 5683.5 20.4 5.5 0.5 2.8 15.43 0.76 0.63

Table 1: Geometric sieve element data. Values for length, diameter, volume and angle of sieve elements from top (T) middle (M) and bottom (B) of plants, stem diameters, and distance from top of plant. All values are a mean of ten measurements of at least ten different sieve elements. SE L = sieve element length in μm. SE D = sieve element diameter in μm. SE V = sieve element volume (assumes cylindrical shape) in μm3. SP ang = sieve plate angle in degrees. SE

L/D ratio = sieve element length to diameter ratio. Stem D = diameter of stem in cm. D from top

= distance from top of plant in m. See Table 2 for species abbreviation list, and total height of each plant.

Sp. Code Scientific name Height (m) COJA Coriaria japonica 2.2 FAAU Fallopia aubertii 3.0 HULU Humulus lupulus 2.0 AKQU Akebia quinata 3.0 DEVE Decodon verticillatus 1.4 TIAM Tilia americana 14.0 CASP Catalpa speciosa 24.5 CASE Calstegia sepium 2.9 OEBI Oenothera biennis 0.7 POTR Populus tremuloides 10.7 NYSY Nyssa Sylvatica 7.2 CUMA Cucurbita maxima 1.3 PHVU Phaseolus vulgare 1.4 SOLY Solanum lycopersicum 1.3 DEBI Deinanthe bifida 0.5 PASP Passiflora sp. 1.9 LITU Liriodendron tulipifera 21.6 ACSA Acer saccharum 21.7 QURU Quercus rubra 27.0 PRSE Prunus serotina 23.5 DACA Daucus carota 0.7 CLVI Clematis viticella 1.9

Table 2: Scientific names of species, and total plant height. For plants utilized in this work, trees had transport lengths of an order of magnitude greater than vines, and vine path lengths were at least double that of herbs.

In trees, there was a strong correlation between distance from the apex of the plant and the diameter of sieve elements overall. Diameter of sieve elements increased by 1.2 μm per meter of stem length, with an R2 fit of more than 0.7, indicating that taller trees have higher stem conductivities (Figure 3). The correlation becomes even stronger when considering the gradient in individual trees, and is also apparent for sieve element length, such as is shown in Figure 4B.

These factors increase conductivity (Equation 1) in more distal parts of the tree indicating that conductivity is an in planta gradient in trees.

Trees 65

m) μ 45

25 Sieve Element Diameter ( Diameter Element Sieve 15 y = 1.2422x + 18.2 R² = 0.7066

5 0 5 10 15 20 25 30 Distance from top of plant (m)

Figure 3: Correlation between Sieve element (SE) diameter and distance from plant apex.

Data from all positions of all trees. N = 10 for all measurements. Bars are standard error.

Equation 1

2 Sieve element specific conductivity (Equation 1). k = conductivity in μm , Np = number of pores, rp = radius of pores, l = length of sieve element, lp = length of pores, r = radius of sieve element.

QURU QURU B 65 A 500 60 55 450

m) 50

μ m)

45 μ 400 40 35 350

30 y = 0.0136x + 22.715 SElength ( y = 0.066x + 258.05 SE diameter ( diameter SE R² = 0.9146 R² = 0.9977 25 300 20 15 250 0 500 1000 1500 2000 2500 3000 0 500 1000 1500 2000 2500 3000 Distance from top (cm) Distance from top (cm)

QURU QURU 14 D 1200000 C 1000000 12

m^3) y = -6E-06x + 12.697 μ 800000 R² = 0.7124 600000 10 y = 373.09x + 84514 400000 R² = 0.9986 8 SE volume ( volume SE 200000 0 6 0 500 1000 1500 2000 2500 3000 ratio diameter to length SE 0 500000 1000000 1500000 Distance from the top (cm) SE volume (μm^3)

Figure 4: Diameter (A) and length (B) of sieve elements increases with stem length in trees.

This results in an increasing sieve element volume gradient (C) which is more strongly influenced by the increasing diameter (D). There is more variation in length than diameter.

Example shown here is Quercus rubrus. Volume is calculated using the equation for a cylinder.

The angle of the sieve plates also affects the sieve tube conductivity by increasing the area of the plates, resulting in more area for sieve pores. Longer sieve element in trees had more steeply angled plates (Figure 5). Sieve elements also became longer in trees the further they were from the top (See Figure 4B). Therefore, sieve plate area is another geometric property that increases phloem conductivity in a top-to-bottom gradient in trees.

Trees 90.00 80.00 70.00 60.00 50.00 40.00 y = 0.0841x + 31.427

30.00 R² = 0.3884 Sieve (deg) angle Sieve plate 20.00 10.00 0.00 100.00 200.00 300.00 400.00 500.00 600.00 Sieve element length (μm)

Figure 5: The angle of sieve plates increases with the length of the cell. Bars are standard error. N=10 for all measurements of length and angle.

The images in Figure 6 depict an example of measurements obtained from confocal imaging using image J. In this example, conductivity provided by radius increases by an order of magnitude, and sieve element length provides another 60% increase in the trunk. Sieve plate area is also increasing, which likely increases the number of pores.

B M

52°

53° T Figure 6: Sieve element measurements from the tree Quercus rubrus. T = 19.5 cm from top, M

= 1400 cm from top, and B = 2690 cm from top. The length and diameter, important factors of

94 sieve tube conductivity, increase as a function of distance from the top of the tree. Scale bars are 20 μm.

In the vines, phloem anatomy displayed much variation, but geometric parameters did not appear to be a linear function of distance from the apex of the plant either as a general trend

(Figure 7), or as an in planta gradient (Figure 8). This may be related to irregular internode length, and lack of variation in stem diameter. Previous work on the vine Morning Glory has detected a gradient in sieve tube conductivity (Knoblauch 2016), but this did not present in the factors measured in these plants.

Vines 45

40 CUMA T

SE diameter (um) diameter SE 20

0 0 50 100 150 200 250 300 350 Distance from top of plant (cm)

Figure 7: Sieve element diameter was not a function of length in vines. The total lengths of the vines were an order of magnitude smaller than those in trees. Cucurbita maxima (CUMA) sieve elements have previously been noted for having high conductivity by Mullendore et al. (2010).

In S. lycopersicum, a vine, sieve element diameter decreased in the middle of the plant, and then increased near the bottom, while sieve element length decreased (Figure 8 A and B).

Similar to other vines examined in this work, conductivity factors do not seem to be positively correlated with distance from the apex in-planta, unlike the strong linear trends in conductivity factors observed in trees. Much variation in sieve element length is apparent by the large error bars.

A SOLY B SOLY 24 240

22 220 m)

20 m)

μ 200 μ 18 180 16 160

14 ( length SE SE diameter ( diameter SE 12 140 10 120 0 50 100 150 0 50 100 150 Distance from tom (cm) Distance from top (cm)

Figure 8: Diameters (A) and lengths (B) of sieve elements in Solanum lycopersicum. The trends in conductivity factors apparent in individual trees are not apparent in individual vines. For all measurements, n = 10.

Figure 9 displays imaging and measurements from S. lycopersicum. The sieve plates are not steeply angled as is the case in Q. rubrus, and even the largest sieve elements in the vine had a smaller volume than the smallest sieve elements in the tree. The largest diameters and lengths of sieve elements observed in Q. rubrus were more than twice that of the largest diameters and lengths of sieve elements in S. lycopersicum.

75um 75 um

20 μm

75μm 18 μm

Figure 9: Sieve element measurements from Solanum lycopersicum. While there was variation in length and diameter of sieve elements at different positions in vines, it was not patterned as a function of distance from apex, as is the case in trees. Note scale bars of T and B are 75 μm, and M is 20 μm.

Although there was much variability in the sieve element dimensions measured in this study, the average ratio between the length and the diameter of the sieve elements in all plants was remarkably similar. On average sieve elements are about twelve times longer than they are wide, with no significant difference between groups (Figure 10).

μ ( 10

5 Sieve element length/diameter ratio ratio length/diameter element Sieve

0 trees vines shrubs herbs Figure 10: Sieve element length to diameter ratios. This ratio was similar in all plants. Trees n =

24, vines n = 27, shrubs n = 9, herbs n = 6. For all plants, average L/D ratio is 12.1, Standard

Deviation = +/- 4.6, n = 66.

As can be seen in Figure 11, the diameter of the whole stem also changed more in trees (Figure

11A) as a general rule than in vines (Figure 11B) as a function of distance from the top of the plant. In trees, the rate of increase was 20 times greater than in vines, and correlated 5 times more strongly. This was also apparent when comparing individual plants. For example the stem diameter of P. tremuloides (Figure 11 C) increased at a rate 5 times that of P. speciosa (Figure

11 D). This implies that the whole phloem area also increases more in tree stems than vines as a function of distance from the top. Trees were also an order of magnitude longer than vines.

Trees Vines

90 A 0.9 B 80 0.8 70 0.7 60 0.6 50 0.5 40 0.4 30 0.3

20 0.2 Stem diameter Stem diameter (cm) Stem diameter Stem diameter (cm) y = 0.0226x + 5.0628 y = 0.0011x + 0.2709 10 R² = 0.6058 0.1 R² = 0.1467 0 0 0 500 1000 1500 2000 2500 3000 0 50 100 150 200 250 300 350 Distance from top of plant (cm) Distance from top of plant (cm)

POTR PASP D 25 C 0.9

20 0.8 0.7 15 0.6 10

0.5 Stem diameter (cm) Stemdiameter

Stem diameter (cm) Stemdiameter y = 0.0191x - 0.7954 y = 0.0041x + 0.3042 5 R² = 0.9999 0.4 R² = 0.9807

0 0.3 0 200 400 600 800 1000 1200 0 20 40 60 80 100 120 140 Distance from top of plant (cm) Distance from top of plant (cm)

Figure 11: Relationship between stem length and stem diameter. This relationship is stronger and steeper than in trees than in vines. All tree data (A), All vine data (B), Individual tree P. tremuloides (C), Individual vine P. speciose (D).

Discussion: As noted by Thompson (2006), conductivity has been the least understood variable for mathematical consideration of translocation in plants. Here this is addressed by delivery of some important factors for calculating conductivity.

Inspection of the equation presented by Thompson and Holbrook (2003) and again by

Mullendore et al. (2010) (Equation 1), reveals that sieve element diameter is a very important variable, as the radius is squared. Sieve element length is also important, as long lengths

99 present lower periodicities of resistance imposed by plates. Large sieve plate angles will supply a greater area of plates, and therefore either more or larger pores, also increasing conductivity.

In Ipomoea nil, sieve plates with a larger area were found to have sieve plate pores of larger diameter by Knoblauch et al. (2016), and the degree of conductivity increased in defoliated plants.

The diameter of sieve tubes in trees increases by roughly 1.2 μm per meter from the apex, meaning that the radius increases by a factor of 0.6. Because the radius is squared, this increases conductivity by a factor of 3.6 each meter. So a 30 meter tree such as Q. rubrus with sieve elements in the ends of branches with a radius of ~ 8 μm will see phloem conductivity increase by at least an order of magnitude in its trunk. These rough calculations assume that the pores are all the same size in the top, middle, and bottom. This is most likely not the case. If anything, previous investigation (Knoblauch et al. 2016) indicate that pore size increases with increasing sieve tube diameter in defoliated vines, and therefore, these may even be underestimations.

The length of the sieve tubes also scaled linearly with conductivity in trees, and this positive correlation supplied nearly an additional 60% more conductivity to the phloem of trees in the trunk compared to branches. In vines, the stems did not vary in diameter as much as in trees, and the total plant length in the species analyzed in this work is an order of magnitude smaller

(Table 2). Trees live for many years, while many vines are annuals. Because the trunk of the tree functions as a sink in summer by storing starch in the plastids of rays, and as a source in spring, supporting new growth, this relationship is probably not incidental.

100

High turgor pressures have been measured in the source phloem of the vine I. nil whose stem leaves had been stripped, leaving only one set of apical source leaves (Knoblauch et al. 2016). If these high pressures are typical for vines, they may reduce the need for higher conductivity. It is also interesting to consider the growth habit of vines. Their internodes vary in length, responding to environmental factors and their genetics. Previous workers have shown an axial increase in top-to-bottom diameters of xylem vessels in Phaseolus vulgaris. This is also ascribed to an auxin gradient (Aloni and Zimmerman 1983). The lack of linear geometric gradients found in sieve tube conductivity factors the phloem of vines studied in this work may indicate that secondary phloem growth is different than secondary xylem growth, or that the species studied here have different growth patterns than P. vulgaris. It could also be related to variable internode length. Internode length in Chenopodium polyspermum has been shown to correlate with cell length, and cell elongation rates (Lecharney 1979).

Nonlinear correlations of internode lengths with respect to distance from apical meristems have been reported by Reid et al. (1983) in Pisum. Increasing internode length was correlated strongly with both the number of cells, and increasing cell lengths by these workers. The relative influence of these factors depended on tissue type, and genetics. If this also applies to phloem, it could explain the nonlinear geometric variability presented by the sieve elements in this study, and also could mean that changing growth conditions could influence sink filling in important food crops.

For vessel elements, an increasing diameter of cells has been reported to continue to increase into the roots (Aloni 1987). The six point hypothesis based on decreasing auxin concentrations, and therefore longer differentiation times, resulting in increasing cell diameter as a function of

101 distance from the apical meristem (Aloni and Zimmerman 1983) appears to also be supported in phloem of trees in the data presented here. The strength of the trunk as a seasonal sink presents a growing “pull” force of carbohydrates.

The low pressures, and symplastic loading strategies presented in tall trees can be explained in the framework of a pressure flow mechanism by their high conductivity. Results presented here support a model of conductivity (Figure 12) that depicts all three factors of phloem conductivity examined here as an increasing top-to-bottom gradient in trees. This model is in harmony with the idea that Münch’s original hypothesis of pressure driven flow in sieve tubes could also be sufficient for transport in trees.

Top branches

Middle trunk

Base of trunk

Figure 12: Proposed model for sieve tube conductivity in trees. In trees, sieve tube conductivity factors were present in a gradient. All factors increased from the top and ends of branches to the bottom of the trunk. Although there was considerable variation in geometric properties in all of the plants’ phloem cells, the trees presented the strongest correlation between conductivity and distance from the apex.

102

Because trees are often dominant in ecosystems, and their ranges are being effected by global warming, especially due to threats to the phloem from invasive pests, much more knowledge of phloem anatomy and physiology is critical for adapting to a changing biome.

The conductivity data presented for vines are also of high value, because many vines are food crops. These plants present excellent models to investigate the relationships between environmental factors and sink filling. The methods presented here offer the possibility to study comparisons between sieve element lengths and diameters to internode length that have previously only been studied in xylem. Many vines are also leguminous, and water efficient, and therefore a deeper understanding of sink filling in this group is very useful.

103

Works Cited:

Aloni, Roni, and Martin H. Zimmermann. "The control of vessel size and density along the plant axis." Differentiation 24, no. 1‐3 (1983): 203-208. Aloni, Roni. "Differentiation of vascular tissues." Annual review of plant physiology 38, no. 1 (1987): 179-204. Azpiroz, Ricardo, Yewen Wu, Jeffrey C. LoCascio, and Kenneth A. Feldmann. "An Arabidopsis brassinosteroid-dependent mutant is blocked in cell elongation." The Plant Cell 10, no. 2 (1998): 219-230. DeFries, Ruth S., Richard A. Houghton, Matthew C. Hansen, Christopher B. Field, David Skole, and John Townshend. "Carbon emissions from tropical deforestation and regrowth based on satellite observations for the 1980s and 1990s." Proceedings of the National Academy of Sciences 99, no. 22 (2002): 14256-14261. De Schepper, Veerle, Tom De Swaef, Ingvar Bauweraerts, and Kathy Steppe. "Phloem transport: a review of mechanisms and controls." Journal of experimental botany 64, no. 16 (2013): 4839- 4850. Fisher, Donald B., and Cora E. Cash-Clark. "Gradients in water potential and turgor pressure along the translocation pathway during grain filling in normally watered and water-stressed wheat plants." Plant physiology 123, no. 1 (2000): 139-148. Fu, Qiushi, Lailiang Cheng, Yangdong Guo, and Robert Turgeon. "Phloem loading strategies and water relations in trees and herbaceous plants." Plant Physiology 157, no. 3 (2011): 1518-1527. Hammel, Harold T. "Measurement of turgor pressure and its gradient in the phloem of oak." Plant Physiology 43, no. 7 (1968): 1042-1048. Jensen, Kåre H., Johannes Liesche, Tomas Bohr, and Alexander Schulz. "Universality of phloem transport in seed plants." Plant, cell & environment 35, no. 6 (2012): 1065-1076. Knoblauch, Michael, Jan Knoblauch, Daniel L. Mullendore, Jessica A. Savage, Benjamin A. Babst, Sierra D. Beecher, Adam C. Dodgen, Kaare H. Jensen, and Noel Michele Holbrook. "Testing the Münch hypothesis of long distance phloem transport in plants." eLife 5 (2016): e15341. Kozlov, Mikhail V., Andrey V. Stekolshchikov, Guy Söderman, Eugenia S. Labina, Vitali Zverev, and Elena L. Zvereva. "Sap‐feeding insects on forest trees along latitudinal gradients in northern Europe: a climate‐driven patterns." Global change biology 21, no. 1 (2015): 106-116. Lang, Alexander. "A relay mechanism for phloem translocation." Annals of Botany 44, no. 2 (1979): 141-145. Lecharny, A. "Phytochrome and internode elongation in Chenopodium polyspermum L. sites of photoreception." Planta 145, no. 5 (1979): 405-409.

104

Magnuson, C. E., J. D. Goeschl, and Y. Fares. "Experimental tests of the Münch–Horwitz theory of phloem transport: effects of loading rates." Plant, Cell & Environment 9, no. 2 (1986): 103- 109. Mullendore, Daniel L., Carel W. Windt, Henk Van As, and Michael Knoblauch. "Sieve tube geometry in relation to phloem flow." The Plant Cell 22, no. 3 (2010): 579-593. Münch, Ernst. "Stoffbewegungen in der Pflanze." (1930). Nomura, Takahito, Masayoshi Nakayama, James B. Reid, Yasutomo Takeuchi, and Takao Yokota. "Blockage of brassinosteroid biosynthesis and sensitivity causes dwarfism in garden pea." Plant Physiology 113, no. 1 (1997): 31-37. Pritchard, Jeremy. "Aphid stylectomy reveals an osmotic step between sieve tube and cortical cells in barley roots." Journal of Experimental Botany 47, no. 10 (1996): 1519-1524. Reid, James B., Ian C. Murfet, and William C. Potts. "Internode length in Pisum. II. Additional information on the relationship and action of loci Le, La, Cry, Na and Lm." Journal of Experimental Botany 34, no. 3 (1983): 349-364. Ryan, Michael G., and Shinichi Asao. "Phloem transport in trees." Tree physiology 34, no. 1 (2014): 1-4. Thompson, Matthew V. "Phloem: the long and the short of it." Trends in plant science 11, no. 1 (2006): 26-32. Thompson, Matthew V., and N. Michele Holbrook. "Application of a single-solute non-steady- state phloem model to the study of long-distance assimilate transport." Journal of Theoretical Biology 220, no. 4 (2003): 419-455. Turgeon, Robert. "The puzzle of phloem pressure." Plant physiology 154, no. 2 (2010): 578-581. Vörösmarty, Charles J., and Dork Sahagian. "Anthropogenic disturbance of the terrestrial water cycle." BioScience 50, no. 9 (2000): 753-765. Wright, John P., and Donald B. Fisher. "Direct measurement of sieve tube turgor pressure using severed aphid stylets." Plant physiology 65, no. 6 (1980): 1133-1135. Zimmermann, Martin H. "Translocation of Organic Substances in Trees. II. On the Translocation Mechanism in the Phloem of White Ash (Fraxinus Americana L.)." Plant physiology 32, no. 5 (1957): 399.

105