Advanced Microscopy Imaging of Phloem Proteins in Arabidopsis Reveal Insights Into Munch’S Pressure Flow Hypothesis

Home , Phloem, Sieve tube element

ADVANCED MICROSCOPY IMAGING OF PHLOEM PROTEINS IN ARABIDOPSIS REVEAL INSIGHTS INTO MUNCH’S PRESSURE FLOW HYPOTHESIS

DANIEL ROBERT FROELICH

A dissertation submitted in partial fulfillment of

the requirements for the degree of

DOCTOR OF PHILOSOPHY

WASHINGTON STATE UNIVERSITY

School of Biological Sciences

MAY 2014

To the faculty of Washington State University:

The members of the Committee appointed to examine the dissertation of DANIEL ROBERT FROELICH find it satisfactory and recommend that it be accepted.

Michael Knoblauch, Ph.D., Chair

Hanjo A. Hellmann, Ph.D.

Winfried S. Peters, Ph.D.

Raymond W. Lee, Ph.D.

Acknowledgments

I would not have been able to complete this Ph.D. without the tremendous support of so many people. Firstly, I want to thank Michael Knoblauch: my committee chair, boss and soccer buddy for coming out to Washington and inviting me into his lab. I thank

Winifried Peters at Indiana Purdue Ft. Wayne for being on my committee despite having to put up all the logistical issues every time we meet or I needed a form signed. Hanjo Hellman has been a great help, always asking the right questions when I have become too myopic, focusing too closely on the details at the expense of the whole. Ray Lee has been invaluable.

He taught me the importance of knowing my audience and how to cater my presentations to them.

In the lab, I especially thank Dan Mullendore. We have worked together from the beginning and having such a talented peer has been invaluable. I hope I have helped him as much as he did for me. Tim Ross Elliott is a great guy to share an office with and he ensures that we will come back from every conference will have some fantastic stories. Hélène

Pellissier and Ray Collier taught me how to clone. Under their guidance, I have created new organisms! Sierra Beecher is always such a sweet person and so nice to talk to. She has helped me both in the lab and out. Other past lab members: Jamie Watts, Adelina Petrova and Hannah Merley made this program a special adventure!

Valerie Lynch-Holm and Christine Davitt are truly microscopy wizards. They have inspired my career in microscopy and taught me so much. Chuck Cody has always kept my plants green despite my best efforts to kill them.

iii

Without my favorite group of runners, the Beer Chasers, I would have surely lost my mind long ago. Wednesday nights are always a highlight of the week. Even the quarterly

Beer Miles, so terrible during the event but amazing shortly after, are a hilarious way to break up the graduate student life. Scott, Annie, Graham, Steffie, Buzz are Aaron are all my closest, greatest friends.

Most of all, I owe all this work to Nicole, my lovely wife. She has put up with me and helped celebrate the successes, and bore more than her fair of the weight from my failures.

Without her, I would have burned out long ago, but with her support, I can flourish.

This research was supported by the NSF. I personally received funding from the

Herbert Eastlick fellowship, Betty Higinbotham travel grants, NASA and the Vincent

Franceschi Graduate Research Fellowship. Science is expensive, but these groups have recognized our merit and paved the way for world class research.

Thanks to everyone I missed and always: Go Cougs!

ADVANCED MICROSCOPY IMAGING OF PHLOEM PROTEINS IN ARABIDOPSIS REVEAL INSIGHTS INTO MUNCH’S PRESSURE FLOW HYPOTHESIS

Abstract

By Daniel Froelich, Ph.D. Washington State University

May 2014

Chair: Michael Knoblauch

Phloem proteins have been widely regarded as a wound response mechanism. All imagery showing apparent occlusions of this protein at a sieve plate have been dismissed as preparation artifact and ignored. Unfortunately, these images only show one still frame of a movie and so all conclusions are susceptible to misinterpretation. Presented here is a combination of high resolution still images with complete context from a dynamic in vivo reference. This new perspective shows that not only are the Sieve Element Occluding

Related (SEOR) phloem protein agglomerations in Arabidopsis common in healthy, translocating, uninjured plants, but that they do not appear to occlude the phloem at all.

The previously known purpose of this very common family of proteins is once again obscure.

Table of Contents Acknowledgments...... iii

Abstract ...... v

Table of Contents ...... vi

List of Figures ...... xi

List of Tables ...... xiii

Chapter 1 - Introduction ...... 1

1.1 Phloem anatomy ...... 2

1.1.1 Phloem fibers ...... 2

1.1.2 Phloem Parenchyma ...... 3

1.1.3 Sieve elements ...... 3

1.1.4 Companion cells ...... 4

1.2 Phloem loading ...... 4

1.3 Phloem Unloading ...... 7

1.4 Phloem Transport ...... 8

1.4.1 Sieve element plastids ...... 10

1.5 Endoplasmic reticulum ...... 11

1.6 Mitochondria ...... 12

1.7 Phloem proteins ...... 12

1.7.1 Dispersive phloem proteins ...... 13

1.7.2 Non-dispersive phloem proteins ...... 13

1.8 Microscopy ...... 14

1.8.1 Optical microscopy ...... 15

1.8.2 Epi-fluorescent microscopy...... 18

1.8.3 Confocal laser scanning microscopy ...... 19

1.8.4 Electron microscopy ...... 20

1.8.5 Scanning electron microscopy ...... 22

1.8.6 Transmission electron microscopy ...... 24

1.9 References ...... 28

Chapter 2 - Phloem Ultrastructure and Pressure Flow: Sieve-Element-Occlusion-Related

Agglomerations Do Not Affect Translocation ...... 32

2.0 Author contributions ...... 32

2.1 Abstract ...... 33

2.2 Introduction ...... 34

2.3 Results ...... 37

2.3.1 Development and Structure of SEOR1 ...... 39

2.3.2 TEM of Sieve Tubes ...... 42

2.3.3 Obstructions in Sieve Tubes ...... 52

2.3.4 Sieve Tube Structure and Its Impact on Phloem Translocation ...... 57

2.3.9 SEOR1 Function ...... 65

2.4 Discussion ...... 68

2.4.1 Sieve Tube Ultrastructure ...... 68

2.4.2 Phloem Translocation ...... 72

2.4.3 SEOR1 Function ...... 75

2.5 Methods ...... 76

2.5.1 Plant Material for Freeze Substitution ...... 76

2.5.2 Micro-ROCs ...... 77

2.5.3 Plunge Freezing and Freeze Substitution ...... 77

vii

2.5.4 Epifluorescence Microscopy ...... 78

2.5.5 Confocal Microscopy ...... 79

2.5.6 FRAP ...... 79

2.5.7 Cloning and Transformation: SEOR1-YFP ...... 80

2.5.8 GFP5-ER ...... 80

2.5.9 SEOR1-GFP ...... 81

2.5.10 T-DNA Insertion Mutants ...... 82

2.5.11 Immunolocalization ...... 82

2.5.12 Accession Numbers ...... 83

2.6 Supplemental Data ...... 83

2.7 Acknowledgments ...... 84

2.8 Author Contributions ...... 84

2.9 References ...... 85

2.10 Appendix A...... 92

2.10.1 Resistance of the sieve tube lumen ...... 93

2.10.2 Resistance of the sieve plate ...... 93

2.10.3 Resistance of the At SEOR 1 agglomeration ...... 94

2.10.4 Resistance of the At SEOR 1 agglomeration opening ...... 94

2.10.5 Resistance of the At SEOR 1 agglomeration fiber network ...... 95

2.10.6 Supplemental References ...... 96

2.10.7 Supplemental Movie 1. SEOR1 in Root Tip ...... 97

2.10.8 Supplemental Movie 2. Real Time Imaging of Phloem Flow ...... 97

2.10.9 Supplemental Movie 3. SEOR1 movement in Injured Sieve Tubes...... 97

viii

Chapter 3 - Arabidopsis P-protein Filament Formation Requires Both AtSEOR1 and AtSEOR2

...... 99

3.0 Author contributions ...... 99

3.1 Abstract ...... 100

3.2 Introduction ...... 101

3.3 Results ...... 104

3.3.1 AtSEOR1 and AtSEOR2 proteins accumulate in Arabidopsis ...... 104

3.3.2 Immunolocalization analysis of AtSEOR mutant lines ...... 105

3.3.3 Formation of the phloem filament matrix requires both SEOR proteins ...... 106

3.3.4 Aphid feeding is not enhanced by the absence of phloem filaments ...... 111

3.4 Discussion ...... 113

3.4.1 Functional redundancy ...... 114

3.4.2 SEOR1/SEOR2 Interactions ...... 115

3.4.3 Plant-insect interactions ...... 117

3.5 Conclusions ...... 118

3.6 Materials and Methods ...... 119

3.6.1 AtSEOR Protein Expression in Arabidopsis...... 119

3.6.2 Arabidopsis T-DNA insertion mutants ...... 120

3.6.3 Immunolocalization of phloem filaments in AtSEOR knockouts ...... 121

3.6.4 Transgenic plants expressing recombinant protein fusions ...... 121

3.6.5 Yeast 2-hybrid analysis of AtSEOR1 and AtSEOR2 interactions ...... 122

3.6.6 Aphid fecundity study ...... 123

3.7 Acknowledgements ...... 124

3.8 References ...... 125

Chapter 4 - SEORious business – structural proteins in sieve tubes and their involvement in sieve element occlusion...... 131

4.0 Author contributions ...... 131

4.1 Abstract ...... 132

4.2 Introduction: struggling with structural sieve tube components ...... 133

4.2.1 Forisome function: seeing is believing—what about knowing? ...... 136

4.2.2 SEO, SEOR, and legume evolution ...... 144

4.2.3 Sieve tube slime: same player shoots again! ...... 146

4.2.4 SEOR proteins: fluid dynamic effects and specific functions ...... 149

4.2.5 Hydraulic effects of SEOR agglomerations in excised organs ...... 152

4.2.6 SEOR interactions with and responses to stress factors ...... 157

4.2.7 Iconoclastic speculations… ...... 162

4.2.8 … on P-proteins and aphids ...... 166

4.2.9 … on phloem exudation and wound sealing ...... 168

4.3 Conclusions ...... 171

4.4 Supplementary data ...... 173

4.5 Acknowledgements ...... 174

4.6 References ...... 174

List of Figures Chapter 1

Figure 1: Schematic diagram labeling the main cell types of the phloem...... 2

Figure 2: Diagram of three different phloem loading strategies. Adapted from (Turgeon,

2010)...... 5

Figure 3: Diagram of phloem transport from source to the sink tissues, facilitated by water cycling via the xylem...... 8

Figure 4: Schematic Reconstruction of an Arabidopsis Sieve Tube...... 10

Chapter 2

Figure 1: Epifluorescence of SEOR1-YFP in Living Roots...... 40

Figure 2: In Vivo Observation of Sieve Tube Structure...... 44

Figure 3: TEM of Sieve Tubes in Arabidopsis...... 47

Figure 4: Fine Structure of Arabidopsis Sieve Tubes...... 49

Figure 5: SEOR1 Mutant-DNA Insertion Line...... 51

Figure 6: Obstructions in Arabidopsis Sieve Tubes...... 56

Figure 7: SEOR1-Like Filaments in Tobacco and Black Cottonwood...... 57

Figure 8: Schematic Reconstruction of an Arabidopsis Sieve Tube...... 58

Figure 9: In Vivo Flow and Injury Experiments...... 67

Chapter 3

Figure 1: Phloem filaments antigenic to RS21 ...... 106

Figure 2: Visualization of GFP-tagged sieve element (SE) occlusion proteins in whole undamaged Arabidopsis roots...... 109

Figure 3: Yeast two-hybrid experiment showing that AtSEOR1 and AtSEOR2 form homo- but not heterodimers...... 111

Figure 4: Mean pre-reproductive period and lifetime fecundity of single Myzus persicae aphids ...... 112

Chapter 4

Fig. 1: Abrupt cold causes stoppage of phloem translocation in the roots of AtSEOR1 knock- outArabidopsis plants...... 161

xii

List of Tables Chapter 2

Table 1. List of Parameters for Flow Calculations ...... 60

Table 2. Parameters Relevant for the Calculation of the Pressure Drop ∆p in Equation

1/(A1) ...... 63

Chapter 3

Table 1: Life history traits of A. gossypii developing wild-type (Columbia) and knockout

Arabidopsis lines ...... 112

xiii

Chapter 1 - Introduction The vascular system of plants is responsible for all intercellular transport between distant regions. It is comprised of the xylem and phloem. Xylem is chiefly responsible for transporting water and nutrients from the site of absorption, the roots, up the shoot to areas of photosynthesis and respiration, the leaves. Aside from a small amount that is produced and consumed within the same tissues, the phloem mainly transports all products of photosynthesis, photosynthates, from sources, sites of sugar production, to sinks, sites of sugar consumption. In addition to this sugary sap, the phloem transports trace amounts of RNA, amino acids, proteins, hormones, ions (Oparka and Cruz, 2000) and conduct action potential-like energy propagation waves (Fromm, 1991; Fromm and

Spanswick, 1993; Fromm and Bauer, 1994).

1.1 Phloem anatomy The phloem is comprised of four main cell types: sieve elements, companion cells, phloem parenchyma and phloem fibers:

Figure 1: Schematic diagram labeling the main cell types of the phloem. Sieve elements (SE) are interconnected at sieve plates via sieve pores. They are closely associated with companion cells (CC). Phloem parenchyma (PP) and phloem fibers (PF) lie adjecent.

1.1.1 Phloem fibers

Phloem fibers are structural elements with thick lignified secondary cell walls which are merely located amongst the other phloem cells. They have little participation in the transport pathway and simply serve as sturdy cells which maintain structural rigidity within the plant.

1.1.2 Phloem Parenchyma

Phloem parenchyma serve two main functions: sugar loading/unloading into the sieve elements and storage (Giaquinta, 1983). At the source tissue, they temporarily store sugars before loading them into the sieve elements. Besides ordinary parenchyma cells, two additional parenchyma cell types called transfer cells and intermediary cells can be structurally discriminated , which appear to have important functions in phloem loading.

They will be discussed in detail later. In the transport phloem, between sources and sinks, there are fewer symplastic connections between the companion cells and phloem parenchyma compared to the loading and unloading zones (van Bel, 2003). Phloem parenchyma cells in the transport phloem may be utilized after a phloem stopping drought stress event to reestablish flow by pushing sugars into the stalled phloem, which is then followed by diffusing water (Cernusak et al., 2003). At the sink tissue, phloem parenchyma is responsible for unloading sugars from the companion cells for dissemination.

1.1.3 Sieve elements

Sieve elements are the translocating cell type which intimately relies on companion cells (van Bel and Knoblauch, 2000). An immature mother cell destined to become a sieve element first divides longitudinally to produce one large sieve element and a smaller companion cell (Fisher and Oparka, 1996). During ontogeny, the sieve element will lose its nucleus, tonoplast, ribosomes, golgi and cytoskeleton (Behnke, 1974; Evert, 1990). It retains mitochondria, plastids, endoplasmic reticulum, specialized proteins and its plasma membrane(van Bel and Knoblauch, 2000). While enucleate, sieve elements are still considered living cells due to the maintenance of a membrane potential between the

3 spacious interior of the cell, the lumen, and outside the plasma membrane. At maturation, they form into a tube system by enlarging plasmodesmata at its distal cell walls (sieve plate), making large sieve pores to another adjacent sieve element (Esau and Vernon, 1961;

Esau et al., 1962). This sieve tube is the primary conduit of translocation and signal transduction throughout the plant.

1.1.4 Companion cells

The neighboring companion cell, which may further divide laterally, is responsible for the production of all maintenance proteins required by the living, enucleate sieve element. Companion cells also provide ATP and phosphorylated glycolytic compounds (van

Bel and Knoblauch, 2000).

1.2 Phloem loading

Loading sugars into the sieve element is fundamentally difficult due to relative amounts of photosynthetic tissue versus transport tissue. All the sugar produced in the voluminous chlorenchyma must be concentrated into the relatively tiny sieve elements, which results in a very large and energetically opposed concentration gradient. Three distinct phloem loading mechanisms have evolved to address this dilemma.

Figure 2: Diagram of three different phloem loading strategies. Adapted from (Turgeon, 2010). Three different sugar loading mechanisms in the phloem are: Passive, Apoplastic and Symplastic. Found more often in woody species, passive phloem loading relies solely on a high concentration gradient between sugar producing mesophyll cells and phloem. Aided, and recognized by a high number of interconnecting plasmodesmata, the sugar simply diffuses into the phloem without any concentration mechanism (Turgeon, 2006). Further, there are two classes of active phloem loaders: apoplastic and symplastic. The most obvious difference between the two is most readily observable by the ultrastructure of their source phloem parenchyma and companion cells. Otherwise, these two mechanisms are distinct by the form of sucrose being transported (Turgeon, 2006). Apoplastic loading species utilize invaginated transfer cells to actively push sugars from the parenchyma cells, through the apoplast (extracellular space) and into the companion cell. This process requires great energy input because it is moving sugar against its concentration gradient and the companion cell is constantly drained by the linked sieve element (Turgeon and Medville,

2004). This sucrose transport pathway includes two steps: efflux from the parenchyma into

5 the apolast and import into the companion cell (Braun, 2012). The SWEET gene family are sucrose efflux proton coupled transporters (Chen et al., 2012). They are responsible for moving sucrose from the mesophyll cell into the apoplastic space at the mesophyll/transfer cell interface. These recently discovered transporters finally solved the mystery of sucrose efflux (Chen et al., 2012). This finding was delayed due to the multiple (17) related and partially redundant genes in Arabidopsis, yielding no noticeable phenotype from a single knockout. Sucrose Transporters (SUC) are responsible for the import of apoplastically located sucrose into the transfer cell. The SUC transporters all share a high affinity for sucrose (Sivitz et al., 2007) while also capable of transporting maltose and other glucosides

(Kühn and Grof, 2010; Gora et al., 2012). The Sucrose Uptake Transporter (SUT) sub-family of the SUC transporters are classified by additionally having a very high specificity to sucrose as a substrate (Reinders et al., 2012). Symplastic loading species utilize intermediary cells which are not invaginated and appear as a normal brick shaped parenchyma cell, but with significantly more vesicles (Oparka and Prior, 1988). The intermediary cell is symplastically linked to the companion cell, and so the produced sugars, disaccharides, are able to freely move into the companion cells. Symplastic loading is therefore a passive loading variant with a specialized concentration mechanism. Once there, they polymerize into raffinose, a trisaccharide, and stachyose, a tetrasaccharide, molecules which serve two purposes (Turgeon, 1991). The plasmodesmata’s small size exclusion limit prohibits movement back into the intermediary cell, and so when the raffinose concentration builds, its only outlet is forward into the sieve element. This is

6 known as the Polymer Trap Hypothesis (Turgeon, 1991). As the polymerization constantly reduces the monomeric sucrose concentration of the sugar, more can diffuse in.

1.3 Phloem Unloading

Primarily at the sink tissues, but also in small quantities along the transport pathway, phloem unloading follows similar mechanisms to loading. Symplastic unloading requires a large concentration of plasodesmata at the companion cell/phloem parenchyma interface so that the sugars are able to passively diffuse out (Turgeon, 1991; Van Bel,

1993). The localized regulation of symplastic unloading is controlled by the plasmodesmata number (long term regulation) and plasmodesmata conductivity by constriction or enlargement (short term regulation). Apoplastic unloading is again under the control of active sucrose transporters. Localized control occurs via up or down regulating the expression of those transporters (Patrick, 1997; Oparka and Cruz, 2000; Williams et al.,

2000). Sucrose diffuses from the sieve element into the symplastically connected companion cell and is transported into the apoplast either by diffusion or potentially the

SWEET transporters (Lalonde et al., 2003; Chen et al., 2012). Sucrose influx into the sink cells is achieved with either facilitated diffusion or proton symport. It is often coupled with hexose invertases, which inhibit backflow into the apoplast and companion cells (Lalonde et al., 2003).

1.4 Phloem Transport

Figure 3: Diagram of phloem transport from source to the sink tissues, facilitated by water cycling via the xylem.

Sugars are transported through the phloem by an osmotically driven pressure pump, hypothesized by Ernst Münch (Münch, 1930). The Münch hypothesis states that the

8 sugars, dissolved at high concentrations in water at the source tissue, form a large osmotic pressure that pushes the solution along the sieve tube to areas of lower concentration, the sinks. This hypothesis assumes the sieve tube acts as a passive tube. In this situation, the sieve tube should adhere to the Hagen-Poiseuille equation of flow through an ideal tube:

This equates the volumetric flow rate (V) from the change in pressure between source and sink (ΔP), radius (r), viscosity of the phloem sap (η) and length of the tube (l). Via the

Hagen-Poiseuille equation, only the radius of the tube has a non-linear effect on the eventual volumetric flow rate. Being raised to the fourth power, the radius will clearly be key factor in considering the suitability of applying this equation to the sieve tube system

(Thompson, 2006; Jensen et al., 2011). As mentioned, the phloem retains several, but not all, cellular organelles. These organelles reduce the effective sieve tube radius and so their retention comes at an exponential cost to the tube’s flow rate and are therefore presumably very important. This drop in flow rate a result in the decrease in the sieve element’s hydraulic conductivity which is calculated both from the impedance due to the remaining organelles as well as other structures, such as phloem proteins and sieve plates (Thompson and Holbrook, 2003). Further, more in depth analysis of sieve plate geometry refined this equation to better compensate for sieve pore number and radii (Mullendore et al., 2010).

While the retained organelles are easily recognized, their specific function in highly specialized cells like sieve elements is not as clear.

Figure 4: Schematic Reconstruction of an Arabidopsis Sieve Tube. Reconstruction of the structure of a sieve element-companion cell complex as found in in vivo confocal studies and after freeze substitution of whole plants. Sieve elements contain ER, mitochondria covered with clamp proteins, and electron-dense vesicles. While those structures are usually embedded in an amorphous ground matrix, SEOR1 filaments and sieve element plastids are always in direct contact with the sieve tube sap. A SEOR1 agglomeration is shown in front of a plate that does not fill the entire lumen of the sieve element. Companion cells contain all organelles typical for a plant cell, but only nucleus, vacuoles, chloroplasts, and mitochondria are shown. Blue lines indicate the location of a cross section for (A) to (C). C, chloroplast; Cl, clamp proteins; EV, electron-dense vesicles; GM, ground matrix; M, mitochondria; N, nucleus; P, plastid; SR, SEOR1 filaments; V, vacuole (Froelich et al., 2011).

1.4.1 Sieve element plastids

Unique sieve element plastids are commonly found containing high concentrations of starch or proteins. Derived from the same proplastids as other common plastids (e.g. chloroplasts, chromoplasts, leukoplasts, etc.) (Behnke, 1974), their function is yet

10 unknown. If they are simply utilized for storage, no regulatory pathway is yet discovered.

Behnke (1972) sorted phloem plastids according to their contents and this classification is consistent within species and used for classifying clades. S-type plastids contain starch and

P-type plastids contain proteins. Numerous, round sieve element plastids are usually found closely associated with the membrane, but with a diameter of about 1µm, they can appear very obstructive in smaller sieve elements. Poor sample preparation for TEM often result in disrupted sieve element plastids (Esau, 1965). The membranes open and spill their contents, which then surge towards the downstream sieve plate. This may lead to the hasty hypothesis that they are injury sensitive and are intended for sieve plate clogging.

Certainly, any organelle that is found at a sieve plate in severely wounded phloem may have a secondary benefit as assisting in isolating wound areas, but it should not be assumed that there is no other purpose, perhaps a much more essential one.

1.5 Endoplasmic reticulum

Sieve element endoplasmic reticulum is commonly found, but never with any associated ribosomes. Usually stacked thick in the corners and thinner along the membrane of the sieve elements, it appears to be arranged specifically to stay out of the translocation stream. Sieve element endoplasmic reticulum is thought to be responsible for the sequestration of calcium (Arsanto, 1986) or aiding the transport from the companion cells

(Sjolund and Shih, 1983). This requires still lacking evidence that it forms a continual link from the companion cell, through the plasmodesmata, into the sieve element (Esau and

Thorsch, 1985). This endoplasmic reticulum is also thought to store and release signaling

11 ions (van Bel et al., 2014) which would help to explain how observed action potentials are able to transmit down the phloem faster than the translocation velocity.

1.6 Mitochondria

Another organelle retained in mature sieve elements are mitochondria. Like the plastids, these are found closely associated to the membrane, anchored by a protein matrix

(Froelich et al., 2011). They are normally less than 1µm in diameter and are consistently spherical (Esau and Cronshaw, 1968), while mitochondria found in other cell types are more often elongated. The mitochondria should remain fully active, as evidenced by their uptake of metabolic indicator dyes (McGivern, 1957; Lee et al., 1971; Moniger et al., 1993).

Sieve elements appear to adapt their organelles in line with the Hagen-Poiseuille equation to optimize flow. Plastids and mitochondria are both rounded and the endoplasmic reticulum is densely stacked where it is out of the translocation stream. Other nonessential organelles have been degraded. During continued investigation of sieve element components, it is important to keep in mind this conserved trend of sieve element evolution: to optimize what is necessary and remove everything else.

1.7 Phloem proteins

The final remaining structure in mature sieve elements are phloem proteins. Phloem proteins are found in nearly all angiosperms and are completely lacking in the poaceae

(Behnke, 1981). They are classified into two groups: dispersive and non-dispersive

(Behnke, 1988). Both are formed early in the sieve elements, and coalesce into phloem protein bodies. Non-dispersive phloem protein bodies remain in this paracrystaline

12 conformation, while dispersive phloem proteins disaggregate into filaments (Cronshaw,

1975).

1.7.1 Dispersive phloem proteins

Ninety percent of all angiosperm families contain examples of dispersive phloem proteins (Behnke, 1991). They have been long suspected as a wound response mechanism

(Johnson et al., 1976; Walz et al., 2004) since they tend to accumulate at the sieve plate and in sieve pores during the surging of sieve element contents during preparation for microscopy. Knock out studies in Arabidopsis and tobacco have shown that this strategy is weak at best (Jekat et al., 2013), but the wound defense theory remains resilient largely because there are no other hypotheses for their function.

The most widely studied filamentous proteins were characterized in pumpkins and were labeled phloem protein 1 and 2 (PP1 and PP2). PP1 is the larger, monomeric 96-kD protein and PP2 is a dimeric 46-kD lectin. Both proteins are synthesized in the companion cells and transport to the sieve elements via plasmodesmata (Bostwick et al., 1992; Clark et al., 1997; Golecki et al., 1999).

1.7.2 Non-dispersive phloem proteins

At least 10% of all dicotyledons contain phloem proteins which do not disperse at maturity, but remain as protein bodies (Behnke, 1991). These are classified according to their shape, which includes: barrel, compound-spherical, spindle, as well as thirteen others

(Behnke, 1991).

The only family of phloem proteins with a known function are the spindle shaped para-crystalline forisomes found in the fabaceae. These proteins lie longitudinally within a sieve element. They are up to 40µm in length and 4µm in diameter and scale along the dimensions of their sieve element (Peters et al., 2006). During wounding, the forisomes undergo a stark confirmation shift from the long and slim low volume state (LVS) to a short, fat and inflated high volume state (HVS). This reaction can be triggered ex planta with as little as 30µM free calcium, strontium and barium and reversed with a chelator, such as EDTA. The reaction does not require any ATP, yet still produces an observable force during contraction (Knoblauch et al., 2003).

Phloem proteins, like all other retained sieve element structures, represent a enigma of significance. They are extremely common, not only across diverse species, but in voluminous abundance within a plant. Their retention is presumably intentional but their purpose is yet undefined. Being a wound sensitive structure, there is inconsistent imaging of healthy in vivo phloem proteins.

1.8 Microscopy

Ultrastructural investigations rely on high resolution imaging and subsequent data analysis. Within light microscopy, there are standard light microscopes which simply magnify reflected (stereo microscope) or transmitted (compound microscope) light to a camera. Fluorescent light microscopes include epi-fluorscent and confocal laser scanning microscopes (CLSM). In lieu of photons, electron microscopes use a focused electron beam to produce images. Scanning (SEM) and transmission (TEM) electron microscopes produce

14 monochromatic images capable of much higher resolution at the detriment of requiring more extensive tissue processing.

Invasive microscopy is largely plagued with artifacts, unnatural features introduced by the microscopist between the living state and imaging. Therefore, all images must be evaluated and ultimately kept or discarded. But what if there are persistent artifacts which occur consistently across various samples and preparation techniques? At a certain point, those artifacts are accepted as a natural, living state. Then, in the future when a better image, free from that artifact is produced, it is quickly discarded as false, perpetuating the misinformation. This occurred in the mid-20th century in sieve element imaging. Phloem proteins and plastid contents were consistently found adjacent to the sieve plate (Hartig,

1854). Also callose, a β-1,3 glucan, was observed within the sieve (Barclay et al., 1977). All this led to the assumption that the sieve elements were persistently occluded, even in a translocating state (Fensom, 1957; Spanner, 1958). Gentler preparation techniques were required, since sieve elements live with such high turgor pressure, and conventional methods resulted in phloem contents surging towards the sieve plate, producing a now well-known artifact (Knoblauch and van Bel, 1998).

1.8.1 Optical microscopy

Optical microscopy often is the best first step in imaging. The samples can be imaged while still living, and with careful technique, without any invasive distress. Static structures, like dead xylem, do not react to plant distress, but dynamic, wound sensitive phloem certainly does. Therefore, optical microscopy can be used for establishing an in vivo

15 reference for more intensive, but higher resolution imaging. An early light microscope investigation of sieve plate pores (Hartig, 1854) was later supplemented by Esau and

Cheadle (1959) when they measured and counted pores in both sieve areas of sieve plates and on lateral walls in 160 species, spanning 60 families. The ability to accurately measure these pores utilized modern optics with oil immersion at 1,350X magnification. Further precision in measuring sieve plates and pores will be discussed in the electron microscopy section.

A standard compound light microscope contains a light source, a condenser lens, a moveable stage, objective lens and a camera/eyepiece. The light is focused and transmits through the sample which is then enlarged and focused to produce a magnified image at the camera. The maximum resolution of a light microscope is dictated by Abbe’s diffraction limit (Abbe, 1873) as:

The resolution or minimal distance between two points that are still discernable as separate points (d) equals the wavelength of the light (λ) divided by twice the numerical aperture ( , where n is the index of refraction and is half of the angle of total collected light, the aperture angle. The numerical aperture is similar to a camera lens’s f- number. It refers to a physical property of the lens – how much light it can accept. A higher numerical aperture accepts more light, which yields better potential resolution. The angle of collected light, , is largely dependent on the refractive indices of everything between

16 the condenser and objective lenses. At every interface (between the lenses, slide, sample, mounting media, coverglass and immersion media), some light is transmitted and some is reflected away. Only the transmitted light will reach the camera and so minimizing light loss due to reflection yields better resolution. Ideally, everything between the two lenses would have identical refractive indices, thus no lost light, but this not practical. Finally, λ, the wavelength of light, has an effect on resolution. Shorter wavelengths have higher potential resolution (e.g., blue light is 800nm; red light is 1400nm). A standard light microscope, under optimal conditions, can resolve up to 230nm. This figure also assumes the sample has sufficient contrast. Contrast is the measure of how much light is refracted while passing through a substance. If all source light passes through unrefracted, there is no image. If light refracts off a surface, then the camera will register varying light intensities forming an image. If a feature is not naturally refractive, because its refractive index is too similar to its surroundings, then stains are used to gain contrast. Stains are colorful dyes that specifically bind to certain substrates. While the substrate may have little contrast, the stain will. They are also useful due to their binding specificity. Multiple stains can be used to specifically color different substrates on one sample.

Some stains and many other endogenous compounds produce images via fluorescence. As certain wavelengths of light strike a sample, electrons are energetically excited to a higher energy shell. This unstable arrangement eventually (within nanoseconds) results in the electron falling back into its original shell. This results in a release of light energy. Since some energy is lost in this process, the light emitted will be at a higher wavelength (lower energy) than the excitation light. All fluorescent compounds

17 have unique excitation and emission spectra. These spectra are both plotted a graph with λ on the x-axis and relative intensity on the y-axis. There is usually a bell shape range of excitation wavelengths that will all be capable of producing emitted fluorescent light.

Multiple fluorescent compounds can be present on a sample just like multiple stains, and they can be used to highlight specific features.

1.8.2 Epi-fluorescent microscopy

An epi-fluorescent microscope looks very similar to a conventional light microscope.

Instead of illuminating the sample with a white, full spectrum light, it uses filters to only shine specific wavelengths of excitation light on the sample. This beam splitter reflects the chosen excitation light towards the sample, and both reflected light and fluorescing light from the sample return along the same path. Reflected excitation light is bounced back towards the light source, while the emission light is transmitted on. There are also further filters to selectively block or transmit desired bands of wavelengths to the camera. One benefit of a fluorescent microscope is that the sample may not need to be injured by cutting. Since the fluorescent light does not need to transmit through the sample, it can shine through the objective and fluoresces back through it to the camera. Therefore, thick and opaque fluorescent samples can be imaged intact. Analine blue is a commonly used phloem indicator dye. It preferentially binds to callose and can be used to quickly identify sieve plates (Currier, 1957; Evert and Derr, 1964).

1.8.3 Confocal laser scanning microscopy

Confocal microscopes are very similar in construction to the light microscopes described above, except they use a focused laser as the light source. It relies on the same fluorescence properties as above. The laser does not illuminate the entire field of view at once. Instead, it sequentially scans the sample, one spot at a time. The light detector is not a normal camera with a large megapixel array of tiny sensors. Instead, it uses a large, single pixel detector to maximize sensitivity. This is fine since only one pixel fluoresces at a time, therefore all the emission light is coming from that single spot. Confocal microscopes have one other large advantage, their pinhole. Light microscopes focus their illumination light at one plane and focus their objective lens to connect light from that same plane. All features above and below that plane will cast a fuzzy blur on the image. A confocal microscope receives emitted light which bounces off the reflective interior walls in a tube after the objective lens, and refocus to a point before spreading back out at the light detector. There is a pinhole aperture at this spot. All the emitted light from a singular focal plane will coalesce at the same plane of the pinhole, while light from out of focus depths in the sample will be spread out. This broad width beam will be largely blocked and will not pass through the pinhole aperture, greatly reducing emitted light from sample depths out of the focal plane. The diameter of the pinhole aperture can be widened to allow imaging of greater sample thickness as a larger range of sample depth’s emitted light is allowed to pass.

Confocal microscopes are therefore always in focus.

Both epi- fluorescent and confocal microscopes mark a huge forward step in capability when paired with fluorescent proteins fused to proteins of interest. They allow

19 the direct imaging of otherwise low contrast structures in vivo. For example, immunological labeling can identify a phloem protein (PP1) using the TEM (Read and Northcote, 1983) of dead fixed tissue, but a fluorescent protein fused to a different phloem protein (At SEOR-1) can be imaged as living and dynamic (Froelich et al., 2011), which provides the full story video instead of possible false assumptions from a single image (Knoblauch et al., 2014).

1.8.4 Electron microscopy

After optical microscopy has been optimized to establish a trusted in vivo reference, higher magnification is necessary to observe the samples ultrastructure.

Electron microscopes use an electron beam in place of light to image a sample. They are therefore fundamentally able to achieve higher resolution since electrons, while physically larger than photons, are able to be focused finer than the wave height of light.

While light is able to pass cleanly through air, electrons tend to scatter if they encounter any mass. Therefore, the electron beam requires a vacuum for highest resolution (Bozzola and Russell, 1999). Vacuums are quite damaging and so it necessitates additional sample preparation. Any water in the sample would immediately begin to draw out in the vacuum.

Due to the bi-polar nature of water, anything with a charge will adhere to it, and as that water evaporates, it will deform. The fundamental purpose of microscopy is to image a sample at its natural state, so this evaporation induced deformation is unacceptable.

Therefore, two different techniques are used to work around this problem: cryogenic imaging and chemical fixatives.

In cryogenic electron microscopy, the fresh sample is rapidly frozen and remains frozen throughout imaging. The sample can then be inserted into the vacuum and imaged since the water (as ice) will not as readily evaporate, causing deformation. The samples are also protected against damage due to the electron beam. Incased in ice, the proteins may otherwise be blasted apart by that high energy beam, but are locked in place and unable to disfigure (Dubochet et al., 1988). Cryogenic microscopes require specialized equipment to ensure the samples remain frozen and are therefore rarer and not utilized in this project.

The more common method to prepare samples for electron microscopy uses chemical fixatives. Like freezing the sample in ice, chemical fixation stabilizes it, via chemical crosslinks. The sample is exposed to aldehyde fixatives, which infiltrate in and form an internal structural architecture that will ultimately retain the living state when all water is removed. A buffered mixture of glutaraldehyde (GA) and paraformaldehyde (PFA) are used to exploit GA’s superior crosslinking strength and PFA’s rapid initial fixation speed

(Karnovsky, 1965; Bozzola and Russell, 1999). Additional fixation with osmium tetroxide

(OsO4) (Claude, 1947) specifically targets double bonds, thus lipids. Unlike the organic and small atoms in GA and PFA, OsO4 is metallic and also stains and makes the sample conductive. After fixation, the sample is dehydrated. The simplest method is freeze drying.

The fixed sample is rapidly frozen in liquid nitrogen and exposed to a vacuum at low temperatures. The ice is then slowly drawn out from the sample. Alternatively, the buffer can be gradually substituted with increasing concentrations of ethanol to slowly remove water. The ethanol can be substituted for liquid carbon dioxide (CO2) in a critical point drier, and that CO2 will be slowly exhausted yielding a dry, dehydrated sample which can be

21 optionally coated for SEM (Bozzola and Russell, 1999). A dehydrated and wet sample can be embedded and polymerized in hydrophobic plastic for sectioning destined for TEM.

Hydrophilic plastic resins may be substituted to avoid dehydration or to allow better post- polymerization infiltration of aqueous stains and immunological markers.

1.8.5 Scanning electron microscopy

Scanning Electron Microscopes (SEM) focus an electron beam similar to a confocal.

They aim the beam at one small area at a time and electron detectors record the number of low energy electrons that are subsequently cast off, due to the interaction of the beam’s high energy electrons with the sample (Bozzola and Russell, 1999). These inelastically scattered electrons are collected by the secondary electron detector, and the differential in detected electrons per pixel forms the final image. Other, elastically scattered electrons bounce off the sample if they strike near the nucleus and have nearly as much energy as the beam electrons. They are not often collected by the weakly positively charged secondary electron detector, but may produce secondary electrons of their own, contributing noise.

Back scattered electrons are collected by another detector, which is mounted surrounding the final gun aperture. These electrons bounce off the sample and reflect back according to the atomic number of the atoms in the sample, and so can be used for elemental analysis

(Goldstein et al., 1981; Bozzola and Russell, 1999). Also similar to a confocal, samples may be thick since SEMs do not require their electrons to pass through to the detector. This allows a three dimensional image of the surface of the sample.

Such an image is ideal for measuring sample features because it does not require careful positioning before imaging. Mentioned above, sieve plates and pores were measured and counted to describe the species phylogenetically, with a future goal of learning their function (Esau and Cheadle, 1959). Unfortunately, the resolution of light microscopy was not adequate to accurately quantify the effect of plates and pores on the tube’s conductivity in species with smaller diameter sieve tubes. There was also an issue of wound response between living tissue and imaging due to surging and callose formation.

Mullendore et al. (2010) pioneered a rapid tissue killing (to stop callose) and clearing (to remove surged proteins) technique to prepare samples for SEM imaging. The resulting measurements of sieve plates and pores revealed a counter intuitive result, that flow velocity increased with decreased conductivity. This finding was only possible due to the

SEM’s higher magnification potential over light microscopes and how it enabled them to look closer at smaller diameter sieve elements.

If the samples are properly fixed, they may be cut to expose interior surfaces for imaging as well. The resolution is dependent on both how precise the electron beam can be focused (how small a spot is energized at once) and how well the sample can cope with that energy. Biological samples naturally do not contain a large quantity of metallic atoms, and therefore behave as an electrical insulator. Under the beam, while many electron bounce off to the detector, some are absorbed into the sample and sporadically discharge after a delay. This will cause an erroneous abundance of recorded electrons which translates into a bright white streak, called charging, on the image (Goldstein et al., 1981). Samples must therefore be prepared to either be internally or externally conductive to ground this

23 retained charge to the sample holder. Internal conductivity described above used OsO4, while external conductivity relies on coating the sample with a thin coating of metal

(Echlin, 1978; Echlin, 1981). A standard coating of gold of about 1 nanometer is usually sufficient to eliminate charging artifacts. This will reduce the imaging resolution as any sub-nanometer features are now obscured. Ideally, the sample is imaged without any coating. A lower power electron beam may still yield high contrast if more low energy electrons interact with the sample. In this situation, there are fewer absorbed electrons, therefore less charging and less of a need to coat the sample. Additionally, a high power beam may simply transmit through the sample, and not bounce enough electrons to the detector (Goldstein et al., 1981). If samples cannot be fixed prior to imaging, and environmental SEM can image them in a hydrated state. While the electron gun is still under vacuum, a differential pumping system allows the sample area to remain near atmospheric pressure. This reduces the rate of evaporation, allowing a period of high resolution imaging of hydrated samples.

1.8.6 Transmission electron microscopy

Transmission Electron Microscopes (TEM) focus their electron beam similar to a standard light microscope. The cone shaped beam focuses to a small point at the sample, and then spreads back out, forming a magnified image. This image is then collected by a camera. In order to transmit the electron beam, the sample must be either very thin, or otherwise electron transparent. The more matter in the sample, the fewer electrons are able to pass through to form the image. Since an atom is largely empty space, the contrast of a TEM image is mostly due to deflection of the negatively charged electrons as they pass

24 near the positively charged nuclei of the sample. This will yield a differential of the amount of electrons at each pixel in the camera, which is translated into an image. Biological samples with smaller weight atoms will struggle to deflect the beam electrons in order to form an image, so thin sections will often be stained with aqueous heavy metal solutions.

The most common TEM stains are lead and uranium based and are usually used together for increased and even contrast across the sample. While a standard thin section for TEM is imaged flat and assumed as two dimensional, there is indeed depth in the sample, usually around 100nm. While imaging protein filament arrangements with diameters under 10nm, there is a considerable amount of three dimensional information that is otherwise discarded. TEM tomography is a technique that takes many images of a sample at various tilt angles, and then separately maps these different views to build a three dimensional volume. This volume can then be digitally rotated and sliced into to produce images that would only otherwise be possible by sectioning along that exact plane.

Normally, TEM sample preparation involves cutting off a chunk of tissue and submerging it in the chemical fixatives described above. The cell layers that immediately bordered the cut will clearly be tore apart and damaged by the razor blade, but hopefully, all cells beyond that cut will be imaged in their natural, living state. Since the phloem is a wound responsive system, all preparation procedures must be carefully designed so that the resulting imaging is not simply all wound artifact (Knoblauch and Oparka, 2012).

Additionally, as a symplastically connected tube system, a very distant cut will cause an immediate drop in pressure everywhere in that tube. Therefore, it is not reasonable to believe that conventionally prepared sieve elements are free from wounding. Sieve element

25 specific TEM procedures can either be especially slow and gentle or extremely rapid. The former will gradually introduce the fixatives into the phloem, so that any wounding response will be minimal and hopefully repaired by the time the fixative kills the plant. The latter strategy involves rapid freezing of the tissue, vitrification, and subsequent substitution of that ice for the chemical fixative at cryogenic temperatures. Normal ice formation distills the cooling water by removing and concentrating all internal solutes into pockets. This would be disastrous for TEM imaging (Froelich et al., 2011). Vitrification is the glassification of liquid water into solid, non-crystalline ice (Bellare et al., 1988). It requires a near instantaneous drop in temperature, quick enough that the individual water molecules are not able to align their dipoles to form crystals, and thus all solutes remain in place. Vitrification requires a decrease in temperature of at least 104 degrees per second

(Bellare et al., 1988). The most widely used procedure is high pressure freezing (Studer et al., 2001). It is unsuitable for phloem imaging because it requires the sample fit into a small copper carrier. The plant would need to be dissected to fit, and so wound response is expected. Another freezing method, plunge freezing, includes quickly submerging the sample in a cryogenic liquid. This method suffers due to the limited depth of vitrified tissue.

Any dermal cells may be perfect glassy ice, but deeper layers will be heat buffered and will thus freeze slower, yielding a gradient towards crystalline ice. The benefit of this technique is that there is no sample size limit, and so entire plants can be submerged, without prior damage. While the sieve elements in roots lie deep, their high concentration of sucrose acts as an intrinsic cryoprotectant. This aides vitrification of the sieve elements even when adjacent cells show complete freezing artifact (Froelich et al., 2011).

By using the best established methods and developing new ones where they are lacking, phloem proteins can be imaged in a natural and living state. Along with knock out studies, the purpose of these seemingly important proteins can begin to be understood.

Other phloem features which also greatly benefited by the advances in microscope include sieve plates, plastids and plasmodesmata. Sieve plates are now known to be almost entirely clear of obstruction of callose, phloem proteins and phloem plastid contents due to careful preparation and non-invasive imaging techniques (Fensom, 1957; Spanner, 1958;

Barclay et al., 1977; Knoblauch and van Bel, 1998). Phloem plastids have been characterized according to their contents by differential staining (Behnke, 1974).

Plasmodesmatal connections between companion cells and sieve elements are still poorly understood, but are currently under investigation regarding their relative numbers in different tissues and the role of sieve element reticulum in regulating their size exclusion limits.

1.9 References

Abbe, E. (1873). Beiträge zur Theorie des Mikroskops und der mikroskopischen Wahrnehmung. Archiv für mikroskopische Anatomie 9: 413-418. Arsanto, J. (1986). Ca2+-binding sites and phosphatase activities in sieve element reticulum and P- protein of chick-pea phloem. A cytochemical and X-ray microanalysis survey. Protoplasma 132: 160-171. Barclay, G.F., Oparka, K.J., and Johnson, R.P.C. (1977). Induced Disruption of Sieve Element Plastids in Heracleum mantegazzianum L. Journal of Experimental Botany 28: 709-717. Behnke, H.-D. (1972). Sieve-tube plastids in relation to angiosperm systematics—an attempt towards a classification by ultrastructural analysis. The Botanical Review 38: 155-197. Behnke, H.-D. (1974). Sieve-element plastids of Gymnospermae: Their ultrastructure in relation to systematics. Plant Systematics and Evolution 123: 1-12. Behnke, H.-D. (1988). Sieve-element plastids, phloem protein, and evolution of flowering plants: III. Magnoliidae. Taxon: 699-732. Behnke, H. (1981). Sieve–element characters. Nordic Journal of Botany 1: 381-400. Behnke, H.D. (1991). Nondispersive protein bodies in sieve elements: a survey and review of their origin, distribution and taxonomic significance. IAWA Bulletin 12: 143-175. Bellare, J.R., Davis, H.T., Scriven, L.E., and Talmon, Y. (1988). Controlled environment vitrification system: an improved sample preparation technique. Journal of electron microscopy technique 10: 87-111. Bostwick, D.E., Dannenhoffer, J.M., Skaggs, M.I., Lister, R.M., Larkins, B.A., and Thompson, G.A. (1992). Pumpkin Phloem Lectin Genes Are Specifically Expressed in Companion Cells. The Plant Cell 4: 1539-1548. Bozzola, J.J., and Russell, L.D. (1999). Electron microscopy: principles and techniques for biologists. (Sudbury, MA: Jones & Bartlett Learning). Braun, D.M. (2012). SWEET! The pathway is complete. Science 335: 173-174. Cernusak, L.A., Arthur, D.J., Pate, J.S., and Farquhar, G.D. (2003). Water Relations Link Carbon and Oxygen Isotope Discrimination to Phloem Sap Sugar Concentration in Eucalyptus globulus. Plant Physiology 131: 1544-1554. Chen, L.-Q., Qu, X.-Q., Hou, B.-H., Sosso, D., Osorio, S., Fernie, A.R., and Frommer, W.B. (2012). Sucrose efflux mediated by SWEET proteins as a key step for phloem transport. Science 335: 207-211. Clark, A.M., Jacobsen, K.R., Bostwick, D.E., Dannenhoffer, J.M., Skaggs, M.I., and Thompson, G.A. (1997). Molecular characterization of a phloem-specific gene encoding the filament protein, Phloem Protein 1 (PP1), from Cucurbita maxima. The Plant Journal 12: 49-61. Claude, A. (1947). Studies on cells: morphology, chemical constitution and distribution of biochemical functions. XLIII. Cronshaw, J. (1975). P-Proteins. In: Phloem Transport, J.D. S. Aronoff, P.R. Gorman, L.M. Srivastava & C.M. Swanson, ed (New York, London: Plenum), pp. 79-115. Currier, H.B. (1957). Callose Substance in Plant Cells. American Journal of Botany 44: 478-488. Dubochet, J., Adrian, M., Chang, J.-J., Homo, J.-C., Lepault, J., McDowall, A.W., and Schultz, P. (1988). Cryo-electron microscopy of vitrified specimens. Quarterly reviews of biophysics 21: 129-228. Echlin, P. (1978). Coating techniques for scanning electron microscopy and x-ray microanalysis. Scanning Electron Microscopy 1: 109-132.

Echlin, P. (1981). Recent advances in specimen coating techniques. SEM SERIES. Esau, K. (1965). Fixation Images of Sieve Element Plastids In Beta. Proceedings of the National Academy of Sciences of the United States of America 54: 429. Esau, K., and Cheadle, V.I. (1959). Size of pores and their contents in sieve elements of dicotyledons. Proceedings of the National Academy of Sciences of the United States of America 45: 156. Esau, K., and Vernon, I.C. (1961). An Evaluation of Studies on Ultrastructure of Sieve Plates. Proceedings of the National Academy of Sciences 47: 1716-1726. Esau, K., and Cronshaw, J. (1968). Plastids and mitochondria in the phloem of Cucurbita. Canadian Journal of Botany 46: 877-880. Esau, K., and Thorsch, J. (1985). Sieve Plate Pores and Plasmodesmata, the Communication Channels of the Symplast: Ultrastructural Aspects and Developmental Relations. American Journal of Botany 72: 1641-1653. Esau, K., Cheadle, V.I., and Risley, E. (1962). Development of sieve-plate pores. Botanical Gazette: 233- 243. Evert, R.F. (1990). Dicotyledons. In: Sieve Elements, H.D.B.R.D. Sjolund, ed (Berlin: Springer), pp. 103- 137. Evert, R.F., and Derr, W.F. (1964). Callose substance in sieve elements. American Journal of Botany: 552- 559. Fensom, D.S. (1957). The bioelectric potentials of plants and their functional significance. The Canadian Journal of Botany 35: 573-582. Fisher, D.B., and Oparka, K.J. (1996). Post-phloem transport: principles and problems. Journal of Experimental Botany 47: 1141-1154. Froelich, D.R., Mullendore, D.L., Jensen, K.H., Ross-Elliott, T.J., Anstead, J.A., Thompson, G.A., Pélissier, H.C., and Knoblauch, M. (2011). Phloem Ultrastructure and Pressure Flow: Sieve- Element-Occlusion-Related Agglomerations Do Not Affect Translocation. The Plant Cell Online 23: 4428-4445. Fromm, J. (1991). Control of phloem unloading by action potentials in Mimosa. Physiologia Plantarum 83: 529-533. Fromm, J., and Spanswick, R. (1993). Characteristics of action potentials in willow (Salix viminalis L.). Journal of Experimental Botany 44: 1119-1125. Fromm, J., and Bauer, T. (1994). Action potentials in maize sieve tubes change phloem translocation. Journal of Experimental Botany 45: 463-469. Giaquinta, R.T. (1983). Phloem loading of sucrose. Annual Review of Plant Physiology 34: 347-387. Goldstein, J.I., Newbury, D.E., Echlin, P., Joy, D.C., Fiori, C., and Lifshin, E. (1981). Electron-Beam- Specimen Interactions. In Scanning Electron Microscopy and X-Ray Microanalysis (Springer), pp. 53-122. Golecki, B., Schulz, A., and Thompson, G.A. (1999). Translocation of Structural P Proteins in the Phloem. The Plant Cell 11: 127-140. Gora, P.J., Reinders, A., and Ward, J.M. (2012). A novel fluorescent assay for sucrose transporters. Plant methods 8: 1-6. Hartig, T. (1854). Über die Querscheidewände zwischen den einzelnen Gliedern der Siebröhren in Cucurbita pepo. Botanischen Zeitschrift 12: 51-54. Jekat, S.B., Ernst, A.M., von Bohl, A., Zielonka, S., Twyman, R.M., Noll, G.A., and Prüfer, D. (2013). P- proteins in Arabidopsis are heteromeric structures involved in rapid sieve tube sealing. Frontiers in Plant Science 4.

Jensen, K.H., Lee, J., Bohr, T., Bruus, H., Holbrook, N.M., and Zwieniecki, M.A. (2011). Optimality of the Münch mechanism for translocation of sugars in plants. Journal of The Royal Society Interface: rsif.2010.0578v2011-rsif20100578. Johnson, R.P., Fruendlich, A., and Barclay, G.F. (1976). Transcellular strands in sieve tubes; what are they? Journal of Experimental Botany 27: 1117-1136. Karnovsky, M.J. (1965). A formaldehyde-glutaraldehyde fixative of high osmolality for use in electron microscopy. J. Cell Biol. 27: 137A-138A. Knoblauch, M., and van Bel, A.J.E. (1998). Sieve Tubes in Action. The Plant Cell 10: 35-50. Knoblauch, M., and Oparka, K. (2012). The structure of the phloem–still more questions than answers. The Plant Journal 70: 147-156. Knoblauch, M., Froelich, D.R., Pickard, W.F., and Peters, W.S. (2014). SEORious business: structural proteins in sieve tubes and their involvement in sieve element occlusion. Journal of experimental botany 65: 1879-1893. Knoblauch, M., Noll, G.A., Müller, T., Prüfer, D., Schneider-Hüther, I., Scharner, D., van Bel, A.J.E., and Peters, W.S. (2003). ATP-independent contractile proteins from plants. Nature Materials 2: 600- 603. Kühn, C., and Grof, C.P. (2010). Sucrose transporters of higher plants. Current opinion in plant biology 13: 287-297. Lalonde, S., Tegeder, M., Throne‐Holst, M., Frommer, W., and Patrick, J. (2003). Phloem loading and unloading of sugars and amino acids. Plant, Cell & Environment 26: 37-56. Lee, D., Arnold, D., and Fensom, D. (1971). Some microscopical observations of functioning sieve tubes of Heracleum using Nomarski optics. Journal of Experimental Botany 22: 25-38. McGivern, M.J. (1957). Mitochondria and plastids in sieve-tube cells. American Journal of Botany: 37-48. Moniger, T., Wang, Q., and Sjolund, R. (1993). Rhodamine 123 labeling of mitochondria in sieve elements. I. Localization with confocal microscopy in plant tissue culture phloem. Acta Micróscop 2: 92-98. Mullendore, D.L., Windt, C.W., Van As, H., and Knoblauch, M. (2010). Sieve Tube Geometry in Relation to Phloem Flow. The Plant Cell 22: 579-593. Münch, E. (1930). Die Stoffbewegungen in der Pﬂanze. (Jena, Germany: Fischer). Oparka, K., and Prior, D. (1988). Movement of Lucifer Yellow CH in potato tuber storage tissues: a comparison of symplastic and apoplastic transport. Planta 176: 533-540. Oparka, K.J., and Cruz, S.S. (2000). The great escape: phloem transport and unloading of macromolecules 1. Annual review of plant biology 51: 323-347. Patrick, J. (1997). Phloem unloading: sieve element unloading and post-sieve element transport. Annual review of plant biology 48: 191-222. Peters, W.S., van Bel, A.J.E., and Knoblauch, M. (2006). The geometry of the forisome-sieve element- sieve plate complex in the phloem of Vicia faba L. leaflets. Journal of Experimental Botany 57: 3091-3098. Read, S., and Northcote, D. (1983). Chemical and immunological similarities between the phloem proteins of three genera of the Cucurbitaceae. Planta 158: 119-127. Reinders, A., Sivitz, A.B., and Ward, J.M. (2012). Evolution of plant sucrose uptake transporters (SUTs). Frontiers in plant science 3: 22. Sivitz, A.B., Reinders, A., Johnson, M.E., Krentz, A.D., Grof, C.P.L., Perroux, J.M., and Ward, J.M. (2007). Arabidopsis sucrose transporter AtSUC9. High-affinity transport activity, intragenic control of expression, and early flowering mutant phenotype. Plant Physiology 143: 188-198.

Sjolund, R.D., and Shih, C.Y. (1983). Freeze-fracture analysis of phloem structure in plant tissue cultures: II. The sieve element plasma membrane. Journal of Ultrastructure and Molecular Structure Research 82: 189-197. Spanner, D.C. (1958). The Translocation of Sugar in Sieve Tubes. Journal of Experimental Botany 9: 332- 342. Studer, D., Graber, W., Al‐Amoudi, A., and Eggli, P. (2001). A new approach for cryofixation by high‐ pressure freezing. Journal of microscopy 203: 285-294. Thompson, M.V. (2006). Phloem: the long and the short of it. Trends in Plant Science 11: 26-32. Thompson, M.V., and Holbrook, N.M. (2003). Application of a Single-solute Non-steady-state Phloem Model to the Study of Long-distance Assimilate Transport. Journal of Theoretical Biology 220: 419-455. Turgeon, R. (1991). Symplastic phloem loading and the sink-source transition in leaves: a model. Recent advances in phloem transport and assimilate compartmentation. Ouest Editions, Nantes, France: 18-22. Turgeon, R. (2006). Phloem loading: how leaves gain their independence. Bioscience 56: 15-24. Turgeon, R. (2010). The Role of Phloem Loading Reconsidered. Plant Physiology 152: 1817-1823. Turgeon, R., and Medville, R. (2004). Phloem Loading. A Reevaluation of the Relationship between Plasmodesmatal Frequencies and Loading Strategies. Plant Physiology 136: 3795-3803. Van Bel, A. (1993). Strategies of phloem loading. Annual review of plant biology 44: 253-281. van Bel, A.J. (2003). Transport phloem: low profile, high impact. Plant Physiology 131: 1509-1510. van Bel, A.J., and Knoblauch, M. (2000). Sieve element and companion cell: the story of the comatose patient and the hyperactive nurse. Functional Plant Biology 27: 477-487. van Bel, A.J., Furch, A.C., Will, T., Buxa, S.V., Musetti, R., and Hafke, J.B. (2014). Spread the news: systemic dissemination and local impact of Ca2+ signals along the phloem pathway. Journal of Experimental Botany: ert425. Walz, C., Giavalisco, P., Schad, M., Juenger, M., Klose, J., and Kehr, J. (2004). Proteomics of curcurbit phloem exudate reveals a network of defence proteins. Phytochemistry 65: 1795-1804. Williams, L.E., Lemoine, R., and Sauer, N. (2000). Sugar transporters in higher plants–a diversity of roles and complex regulation. Trends in plant science 5: 283-290.

Chapter 2 - Phloem Ultrastructure and Pressure Flow: Sieve-Element-

Occlusion-Related Agglomerations Do Not Affect Translocation

Daniel R. Froelich,a Daniel L. Mullendore,a Kåre H. Jensen,b Tim J. Ross-Elliott,a James A.

Anstead,c Gary A. Thompson,c Hélène C. Pélissier,a,d and Michael Knoblaucha a) School of Biological Sciences, Washington State University, Pullman Washington 99164 b) Department of Physics, Technical University of Denmark, 2800 Kongens Lyngby,

Denmark. c) College of Agricultural Sciences, Pennsylvania State University, Pennsylvania 16802 d) Department of Plant Biology and Biotechnology, University of Copenhagen, 1871

Frederiksberg, Denmark.

Published: The Plant Cell Online 23, 4428-4445, 2011.

2.0 Author contributions

This publication is the culmination of a constantly expanding collaboration. It began as an investigation of forisome related phloem proteins in Arabidopsis. Fluorescent fusion constructs were cloned (Froelich and Ross-Elliott, aided by Pélissier), and through extensive confocal (Froelich) and TEM (Froelich and Mullendore) imaging, it became apparent that conventional views regarding phloem translocation blockages were suspect.

Mathematical modeling of these blockages was calculated (Jensen). Through a fortunate encounter at the Plant Vascular Biology conference in 2010, it was discovered that another group was interested in the same proteins (Thompson and Anstead), and so they

It was also featured as a cover image of that issue.

The bulk of the writing (Knoblauch) was supplemented by Froelich and Anstead. All figures were prepared by Froelich, except Figure 5 (Anstead) and Figure 8 (Knoblauch). All authors assisted in editing for publication.

2.1 Abstract

Since the first ultrastructural investigations of sieve tubes in the early 1960s, their structure has been a matter of debate. Because sieve tube structure defines frictional interactions in the tube system, the presence of P protein obstructions shown in many transmission electron micrographs led to a discussion about the mode of phloem transport.

At present, it is generally agreed that P protein agglomerations are preparation artifacts due to injury, the lumen of sieve tubes is free of obstructions, and phloem flow is driven by an osmotically generated pressure differential according to Münch’s classical hypothesis.

Here, we show that the phloem contains a distinctive network of protein filaments. Stable transgenic lines expressing Arabidopsis thaliana Sieve-Element-Occlusion-Related1

(SEOR1)–yellow fluorescent protein fusions show that

At SEOR1 meshworks at the margins and clots in the lumen are a general feature of living sieve tubes. Live imaging of phloem flow and flow velocity measurements in individual tubes indicate that At SEOR1 agglomerations do not markedly affect or alter flow. A transmission electron microscopy preparation protocol has been generated showing sieve tube ultrastructure of unprecedented quality. A reconstruction of sieve tube ultrastructure

33 served as basis for tube resistance calculations. The impact of agglomerations on phloem flow is discussed.

2.2 Introduction

All organisms, in particular multicellular ones, need to maintain functional coherence. They must coordinate activities and processes that occur in their various parts and integrate a variety of stimuli from the outside to produce meaningful responses. In land plants, the phloem tissue is thought to play an essential role in organismal coordination.

The phloem tissue of angiosperms consists of phloem parenchyma cells, sieve elements, and companion cells. Sieve elements assemble into sieve tubes, which form a continuous microfluidics network throughout the plant body. The primary function of the phloem is the long-distance distribution of photoassimilates and signals. For rapid movement of large fluid volumes, tube systems are used in many natural and artificial systems. To support urban centers, we use pipelines for water, oil, sewage, etc. In animals, circulatory tube systems translocate nutrients and waste to be exchanged at dedicated locations. In basically all known cases, the driving force for flow is a pressure differential that may be positive (e.g., garden hose) or negative (e.g., xylem). Thus, it appears intuitive that the driving force to distribute photoassimilates in the phloem would follow similar mechanisms, and it is not surprising that an osmotically generated pressure differential is the central element of Münch’s pressure flow hypothesis (Münch, 1927, 1930).

However, on closer inspection, there are some striking differences between the phloem and other systems. To minimize resistance, the tube should be free of obstructions and the walls should be smooth. This is relatively easy to realize when flow occurs through the extracellular matrix. The phloem, however, is the only long-distance transport system where flow occurs intercellularly in the symplast. Thus, constituents required to maintain tube integrity, such as organelles, are located in the path of flow. Although the cellular infrastructure has been minimized by loss of the nucleus, the vacuole, ribosomes, Golgi, and the cytoskeleton, sieve elements are not empty tubes but contain smooth endoplasmic reticulum (ER), mitochondria, sieve element plastids, and phloem proteins (P proteins;

Knoblauch and Peters, 2010).

Independent of the length of the tube, a single internal obstruction may increase the resistance of the tube to the point of complete flow stoppage. Obstructions can be used for flow control, for example, by a stopcock, but it bears some risks if a clot is formed unintentionally (e.g., stroke and heart attacks). Since the first descriptions of the phloem, clots in the lumen and often on the sieve plate were commonly observed. Initially, these clots were designated as slime (Hartig, 1854). Later, they were renamed P proteins due to their proteinaceous nature (Cronshaw, 1975). When transmission electron microscopy

(TEM) became available, a surprising variety of P proteins were discovered. They were characterized as amorphous, crystalline, filamentous, tubular, and fibrillar (for an overview, see Evert, 1990). The higher resolution, however, did not change the fact that they were most commonly found in the lumen or inside the sieve plate pores, which led to one of the most controversial discussions in plant physiology of the last century. Some

35 investigators believed that electron micrographs represented the in vivo state. Because bulk flow through occluded pores could not be driven by pressure gradients, alternative translocation hypotheses were developed, such as the electroosmotic theory (e.g., Fensom,

1957; Spanner, 1958, 1970; Siddiqui and Spanner, 1970). Other authors, however, believed that P proteins shown in many micrographs were dislocated during tissue preparation.

Sometimes, plates had open pores after gentle preparation (e.g., Fellows and Geiger, 1974;

Fisher, 1975; Russin and Evert, 1985). This led to the conclusion that sieve tubes form a continuous path and that phloem flow can be driven by an osmotically generated pressure differential (Thompson, 2006). However, convincing evidence has not been shown.

The major reason for this controversy is in the nature of phloem anatomy and the resulting difficulties with in vivo observations of sieve tubes. The phloem is generally embedded in layers of ground tissue preventing direct observation of cellular features.

Therefore, some degree of invasive preparation is required. The sieve tube system builds a network in the plant body, and the exceptionally high turgor (Turgeon, 2010) causes an immediate effect over large distances when a tube is severed. This led to an overwhelming amount of ultrastructural data accounting for different degrees of injury, but it is not clear if uninjured sieve tubes have ever been observed in TEM micrographs.

Recently, we isolated three genes expressing phloem-specific P proteins involved in the formation of forisomes (Pélissier et al., 2008). Forisomes are contractile P protein bodies occurring in faboid legumes. They were suggested to reversibly block sieve tubes in case of injury (Knoblauch et al., 2001, 2003). We designated the gene family Sieve-Element-

Occlusion (SEO; Pélissier et al., 2008). We found homologous genes of unknown function in other plant species, including Arabidopsis thaliana (At3g01680; Pélissier et al., 2008), and designated them Sieve- Element-Occlusion-Related (SEOR). Recently, Rüping et al. (2010) suggested calling genes involved in forisome formation SEO-F (for Sieve Element Occlusion by Forisomes). In our opinion, there is neither a reason nor a justification to rename the gene families. The SEO family implies forisome genes and SEOR signifies homologous genes in nonfabaceae families as originally described by Pélissier et al. (2008).

Without a clear understanding of the underlying construction of the sieve tube system, it will be impossible to properly understand its functional principles. Therefore, we intended to elucidate the ultrastructure of uninjured sieve tubes by TEM by comparing our findings to those obtained from in vivo studies by confocal microscopy.

2.3 Results

Because electron microscopy samples are under high vacuum, samples have to be fixed and dehydrated. Since the structure of proteins, membranes, and other cellular components is often defined by their interaction with water molecules, dehydration may lead to artifacts. The degree of artifacts varies with cell and protein type. To draw appropriate conclusions, an in vivo reference is most helpful. Unfortunately, such a reference is lacking for sieve tubes. Sieve tube components are usually invisible in the light microscope because of their size and/or lack of contrast. In addition, it would be important that sieve tubes be observed without preparation, which is unfortunately usually

37 impossible because of the anatomy of the plant. So far, not a single study has shown cellular features of the phloem without preparation of the tissue. Even the method that allowed us to investigate individual uninjured sieve elements in broad bean (Vicia faba) at high resolution requires removal of cortical cell layers (Knoblauch and van Bel, 1998). Our aim for this study, however, was to investigate sieve tubes without any mechanical intervention.

In aboveground organs, in addition to being embedded in a thick layer of ground tissue, the phloem is covered by more or less opaque, pigmented cells, making a direct observation impossible. The cells in roots of many plant species, however, are relatively transparent. To study the phloem, a thin cortical layer is beneficial, since cell wall– cytoplasm interfaces lead to reflection and refraction phenomena. In this regard,

Arabidopsis thaliana appears ideal. The cortical layer in primary roots is just three cell layers thick, and the root cells do not contain significant amounts of polyphenolics and other compounds that would significantly affect optical properties. The small size of

Arabidopsis sieve tubes is a drawback, but with high-end instrumentation, subcellular structures can be visualized.

We expected that the forisome homolog gene SEOR1 in Arabidopsis encodes a specific P protein. We cloned the gene, including its endogenous promoter, fused yellow fluorescent protein (YFP) to its C terminus and generated transgenic Arabidopsis lines. To study roots in vivo, we used microscopy rhizosphere chambers (Micro-ROCs) and grew

Arabidopsis plants expressing SEOR1-YFP for structural studies. The chambers consist of

38 plant pots with a cover glass as one of the side walls, optimized for high resolution. Root growth is funneled along the cover glass by a porous mesh, while root hairs are in direct contact with soil. In contrast with glass-bottom Petri dishes, where plants are grown in an artificial medium under sterile conditions and at 100% humidity, Micro-ROCs allow direct visualization of the root system in a natural soil environment, which also includes symbionts. Maximum resolution without any preparation or manipulation of the tissue is possible (Figure 1A).

2.3.1 Development and Structure of SEOR1

YFP fluorescence was first detectable in the differentiation zone of young roots

(Figure 1B). After elongation, spherical amorphous protein agglomerates were found inside the cells (Figure 1C). Time-lapse movies revealed quick movements of the bodies in actively growing root regions (see Supplemental Movie 1 online). In the course of further development, the protein bodies increased in size and became elongated (Figure 1D).

An early indication of branch root development is the appearance of additional small protein bodies beneath the sieve tube (Figure 1E). The bodies increase in size (Figure

1F) until the branch root breaks through the cortical layer and branch root sieve tubes are formed (Figure 1G). At certain locations, a sudden and significant alteration in the shape of the protein bodies can be observed. The oval, amorphous bodies condense and transform into defined filamentous structures (Figure 1H). Both amorphous bodies and filamentous structures are usually aligned in files.

Figure 1: Epifluorescence of SEOR1-YFP in Living Roots. (A) An Arabidopsis plant grown in a Micro-ROC. The root hairs of the plant are in contact with the soil, while the roots are forced to grow along the cover slip. (B) A root tip and a young part of a root as observed by epifluorescence in a Micro-ROC. Cells were stained with synapto-red to visualize cell outline. Bright spots along the root are SEOR1-YFP fusion proteins. The image is a single frame of Supplemental Movie 1 online. (C) and (D) Higher magnification of SEOR1-YFP fusion proteins (C). In young vascular tissue, the proteins appear as round amorphous bodies (arrows), which increase in size and become elongated in consecutive slightly older areas ([D], arrow). (E) Early indication of root branch formation is the abundance of SEOR1-YFP bodies beside the file (arrow). (F) After the root tip broke through the cortical layer, a new vascular file formed. (G) A root containing numerous amorphous bodies in a file (arrows). (H) Ten hours later, the amorphous bodies have developed into more defined structures (arrows). Bars = 150 µm in (B), 25 µm in (C) and (D), 50 µm in (E) and (F), and 100 µm in (G) and (H).

To study subcellular localization, we investigated living root sieve tubes by confocal microscopy. Besides the prominent amorphous bodies, fine strands became visible (Figure

2A). To identify the cell type in which the fine strands occur, we generated a double transgenic line expressing SEOR1-YFP and green fluorescent protein (GFP) tagged to the

ER under control of the Medicago truncatula SEO2 promoter. This promoter is known to be sieve element specific, which allows the unequivocal distinction of sieve elements and companion cells (Knoblauch and Peters, 2010). Filaments are restricted to sieve elements

(Figures 2B and 2C), while amorphous protein may occur outside a file, probably in young developing tubes. To determine the location of actively translocating sieve tubes, we loaded leaves of transgenic Arabidopsis plants grown in Micro-ROCs with carboxyfluorescein diacetate (CFDA; Wright and Oparka, 1997) and observed transport in uninjured roots.

Translocation occurred in cells containing filamentous proteins (Figure 2D).

In older roots, SEOR1-YFP filaments became more prominent (Figures 2E and 2F).

Amorphous protein bodies are located in neighboring and nontranslocating cell files, supporting the notion that these are young developing sieve elements (Figure 2F) and that sieve tubes become active after the proteins transform into filaments.

At highest resolution, a meshwork becomes visible that usually extends throughout the sieve element (Figure 2G). The meshwork and ER cover a significant fraction of the sieve tube membrane (Figure 2H) and are closely associated. At the sieve plate, the meshwork traverses the sieve plate pores, outlining their location (Figures 2I and 2J).

We reinvestigated the literature and our own vast collection of sieve tube micrographs but failed to find any structures in electron micrographs of Arabidopsis and other species that resembled the meshworks found in our confocal images of mature translocating tubes. We therefore decided to reinvestigate sieve tube ultrastructure.

2.3.2 TEM of Sieve Tubes

The formation of artifacts in sieve tubes due to preparation and fixation for electron microscopy has been discussed in numerous publications (e.g., Spanner, 1978; Evert,

1982). A large number of fixation protocols, including chemical and freeze fixation of plant and callus sieve tubes has been tested (e.g., Cronshaw and Esau, 1967; Wooding, 1969;

Sjolund and Shih, 1983). A crucial step, however, is the preparation before fixation. Since electron microscopy generally requires small samples, the tissue is usually sectioned. This procedure induces artifacts before fixation is even initiated. Therefore, we decided to fix entire plants to prevent prefixation artifacts.

Chemical fixation may be suboptimal because of the slow diffusion of the fixative through multiple tissue layers. Ultrafast freezing procedures require vitrification of the tissue; otherwise, water-crystal formation leads to complete distortion of cellular features.

Although high-pressure freezing provides superior vitrification to a depth of up to 500 μm and represents the optimum procedure for tissue cryofixation, the maximum sample size is an area of 1 x 2 mm, too small for any plant (Bozzola and Russell, 1999). Other freezing techniques such as jet freezing or plunge freezing usually vitrify the outer 5 to 40 μm of the

42 tissue at best. Even in very small plants such as Arabidopsis, the phloem is never closer than

50 μm to the surface. However, the phloem has one major advantage over other tissues. It carries a high concentration of an intrinsic cryoprotectant: Suc.

Standard fixation of Arabidopsis leaf and stem segments after excision leads to the typical precipitation of P proteins on the sieve plate (Figure 3A), which has been seen before in many other plant species. Often the pores are filled with protein filaments and lined with a thick layer of callose (e.g., Wooding, 1969; Figure 3B). We compared chemical fixation of excised tissue with chemical fixation of whole young plants. P proteins in uncut sieve tubes were more evenly distributed throughout the lumen, and the organelles were usually intact (Figures 3C and 3D). The appearance resembled tubes after gentle preparation (Ehlers et al., 2000). However, no structure could be found that matched the strands observed by confocal microscopy in living tubes. We then took young Arabidopsis plants in the four to eight leaf state and plunge froze them in slush nitrogen (~63K).

Subsequently, the tissue was freeze-substituted in aldehyde fixative containing acetone and postfixed in osmium tetroxide (see Methods for a detailed protocol). Initially, preservation was poor and it turned out that plants had to be grown at 100% humidity either in soil or on Petri dishes to achieve appropriate preservation. Such growth conditions prevented the formation of a thick cuticle that obviously represents a significant freezing barrier. In some cases, it is beneficial to add 0.1 to 0.5% water to the glutaraldehyde-containing acetone fixative. The water supports preservation and easier sectioning of the tissue.

Figure 2: In Vivo Observation of Sieve Tube Structure. (A) SEOR1-YFP fusion protein distribution within vascular bundles shows files containing amorphous bodies (solid arrows) and files containing fine strands (dashed arrows). (B) and (C) GFP specifically tagged to the sieve tube ER (green) reveals that SEOR1-YFP (cyan) is located in sieve elements. Arrows point toward sieve plates.

(D) Loading of phloem with CFDA (red) shows that fine SEOR1-YFP filaments (cyan) are located within mature, translocating sieve tubes. Amorphous bodies are located outside of translocating files. (E) and (F) In older root tissue, a large amount of SEOR1-YFP is abundant in sieve tubes. Consecutive files lead into branch roots (E). Before dispersion, amorphous SEOR1-YFP bodies (arrow) are indicative of young developing sieve tubes and do not translocate CFDA (red). (G) and (H) At highest resolution, the ER (green) is surrounded by a fine SEOR1-YFP filament meshwork (cyan). (I) SEOR1-YFP filaments cover and/or traverse a sieve plate (arrow), outlining the sieve plate pores. (J) Despite the presence of filaments (cyan) in the pores, sieve tubes are fully functional, as indicated by translocation of CFDA (red). Bars = 25 μm in (A) to (E) and (G), 75 μm in (F), and 5 μm in (H) to (J).

In plunge-frozen and freeze-substituted tissue, parenchyma cells surrounding the sieve elements and companion cells are severely damaged and no subcellular structures are preserved (Figure 3E). However, unprecedented preservation is achieved in sieve elements and companion cells of source leaves. Sieve element plastids and mitochondria are intact. Most importantly, protein filaments, 20 nm (±1.7 nm) in diameter and often forming bundles, are located at the margin of the cells (Figure 3E), while the fine filaments in the lumen, usually found after chemical fixation (cf. Figures 3A and 3C), are absent. In accordance with confocal images (Figures 2G and 2I), filament bundles are preferentially oriented longitudinally to the sieve tube axis (Figures 4A to 4C). Bundles may consist of

<10 to >100 individual filaments (Figure 4A). Tangential and longitudinal sections suggest that the filaments are relatively flexible, may bend backward, and often are not strictly aligned in parallel (Figures 4B and 4C).

Sieve plate pores are often unobstructed (but see below), and there is no indication of callose deposition around the pores (Figure 4D). In accordance with investigations after chemical fixation, the ER is organized in stacks (Figure 4E). Sieve element plastids have a smooth surface and are not in close contact with other structures or organelles (Figures 3E,

4A, and 4F). By contrast, mitochondria are always embedded in a parietal layer (Figure 4G), and they are always surrounded by a “halo” of 34.5 nm (±8 nm; Figures 4H and 4I) to which other structures, such as protein filaments (Figure 4H) or membranes (Figure 4I), are attached. In addition, it appears that there is an amorphous ground matrix of the parietal layer embedding all other structures (Figures 4G to 4I). The nature of this matrix is obscure. In some cases, it looks as though ER membranes disintegrate or transform into this amorphous structure (Figure 4E). The layer, however, could also consist of parietal proteins found in other plant species (e.g., Knoblauch and van Bel, 1998). In addition to mitochondria, smaller, electron-dense vesicles can frequently be found in the parietal ground matrix (Figures 4G to 4I), which seem to bud off of membrane structures. The nature of the membranes is yet unclear. They may be constituents of the ER, but they often appear more electron dense and significantly better preserved than the ER, suggesting a different molecular composition.

Figure 3: TEM of Sieve Tubes in Arabidopsis. (A) and (B) Standard chemical fixation of tissue sections of Arabidopsis shows the typical abundance of P protein filaments (dashed arrow) in front of the sieve plate ([A], solid arrow) or in the sieve plate pores (B). Remnants of sieve element plastids ([B], open arrows) can be found around the sieve plate. (C) and (D) Standard fixation of whole Arabidopsis plants resembles images after gentle preparation. Protein filaments (dashed arrows) are located in the lumen of the sieve

47 element, but a sieve element plastid (asterisk) in front of the sieve plate (solid arrow) is intact. (E) Arabidopsis phloem tissue after plunge freezing of entire plants. Phloem parenchyma cells (PP) are completely destroyed by the freezing procedure, but sieve elements (SE) and companion cells (CC) show unprecedented preservation. Sieve element plastids (asterisk) and mitochondria (solid arrows) are well preserved. Most importantly, protein filaments (dashed arrows) are not randomly located in the lumen but consist of longitudinally aligned filaments at the margins of the cells. Bars = 1000 nm in (A), (B), and (E) and 500 nm in (C) and (D).

To verify that the filaments and bundles are formed by SEOR1, we investigated the

Arabidopsis T-DNA insertion mutant GABI-KAT 609F04. The T-DNA insertion is located in the first exon (Figure 5A). PCR experiments verified that the protein is effectively knocked out, but truncated mRNAs are formed. PlantpromoterDB 2.0 predicted a possible weak promoter in the second intron, which might lead to the formation of the observed truncated mRNAs. However, the mutant did not show antigenicity to the P protein–specific antibody RS21 (Toth and Sjolund, 1994; Toth et al., 1994), while the phloem in wild-type plants was well labeled (Figure 5B).

Figure 4: Fine Structure of Arabidopsis Sieve Tubes. (A) Cross section of an Arabidopsis vascular bundle showing two sieve elements. Large bundles of filaments (solid arrows) are located at the margins of the cells. Filaments and sieve element plastids (dashed arrow) fill a significant portion of the tube lumen. (B) and (C) Tangential section through the marginal layer of a sieve element showing aligned filaments in a bundle (B). While the filaments are usually aligned in parallel to the sieve elements’ (SE) long axis, they appear flexible and may bend backward (C). (D) Sieve plate pores are unobstructed and do not contain any detectable callose. (E) Cross section of a sieve element (SE) showing stacked ER cisternae. The ER is usually not as well preserved as in standard fixed tissue. It appears to descend into a less defined amorphous ground matrix. (F) A sieve element plastid with a smooth surface in direct contact with sieve tube sap. (G) A cross section through a sieve element showing a variety of sieve tube components, such as mitochondria, P protein filaments (solid arrow), ER (open arrow), and electron- dense vesicles (dashed arrow) embedded in an amorphous ground matrix. (H) Two mitochondria (asterisks) covered by a halo of proteins (dashed arrow) that attach them to protein filaments (solid arrow).

(I) In other cases, mitochondria (asterisk) are surrounded by membranes from which electron-dense vesicles (solid arrows) may bud off. Again, membranes are not in direct contact with the mitochondria but are attached by small proteins (dashed arrows). The electron-dense vesicles and mitochondria are usually embedded in the amorphous ground matrix (open arrow), while P protein filaments and sieve element plastids are always in contact with sieve tube sap. Bars = 1000 nm in (A), 500 nm in (B) to (E), (G), and (I), and 250 nm in (F) and (H).

The neighboring gene, At3g01670, which shows high homology to At3g01680, was not affected by the T-DNA insertion (Figure 5B). TEM images confirmed the absence of

SEOR filaments, while all other structures found in wild-type plants were present (Figures

5C to 5E). Complementation by transformation of the mutant with SEOR1-GFP led to recovery of filament generation in the mutant. The complemented line showed reduced fluorescence compared with the SEOR1-YFP line; however, bundles of filaments resembling those of SEOR1- YFP plants could clearly be visualized (Figure 5F).

Figure 5: SEOR1 Mutant-DNA Insertion Line. (A) A representation of the Arabidopsis gene At3g01680 indicating the location of the T- DNA insertion in the GABI-KAT 609F04 line and the location of a possible weak promoter indicated by analysis using PlantpromoterDB 2.0 (http://ppdb.agr.gifu-u.ac.jp/ppdb/cgi- bin/index.cgi). Also shown are three sections amplified by RT-PCR showing that a truncated mRNA product containing sections 2 and 3 is produced in the T-DNA insertion mutant. C, amplification control; KO, GABI-KAT 609f04; WT, wild-type Arabidopsis line Columbia. (B) Immunolocalization using a P protein–specific antibody indicates P proteins are absent in GABI-KAT 609F04 (insets are higher magnification images of single vascular bundles), and RT-PCR analysis shows the expression of the adjacent gene At3g01670 (70) is unaffected in the At3G01680 (80) T-DNA insertion mutant (Actin serves as an amplification control). (C) TEM micrograph of SEOR1 T-DNA insertion mutant after standard chemical fixation. Filaments filling the lumen of the sieve tube as shown in Figure 3 are absent. (D) and (E) TEM micrographs of At SEOR1 T-DNA insertion mutant after freeze substitution of whole plants. At SEOR1 filaments are absent, but all other structures, such as ER, mitochondria, and clamps proteins surrounding the mitochondria, are present.

(F) Transformation of KO:GABI-KAT 609f04 with At SEOR1-GFP leads to filament formation. Bars = 100 μm in (B) (inset = 20 μm), 1 μm in (C), 500 nm in (D) and (E), and 3 μm in (F).

2.3.3 Obstructions in Sieve Tubes

Frequently, we noticed in confocal images that SEOR1-YFP forms agglomerates filling significant portions of the tube diameter at, or close to, the sieve plate. The appearance of these agglomerates is extremely variable. Some sieve tubes may contain large bundles (Figure 6A), while others have agglomerates on both sides of the sieve plate and filaments spanning through the pores (Figure 6B). In many cases, however, multiple large agglomerations fill the entire lumen of the tube (Figures 6C and 6D). We loaded the phloem with CFDA and, surprisingly, independent of the amount of protein in the tube lumen, all sieve tubes were fully functional (Figures 6C and 6D). The structural state of the protein filling the lumen is different in mature and young sieve tubes. Developing sieve tubes contain amorphous protein bodies (Figure 6E, lower body), while the agglomerations in mature sieve tubes consist mainly of filaments and bundles, indicated by their extensions

(Figure 6E, upper body; see also Figures 6H to 6J). Despite the large amount of protein within the flow path, the tube contains CFDA (Figure 6F). The lower tube is still in development with isolated sieve elements and a sieve element in the transition phase

(Figure 6F, lower file, left).

Since CFDA is loaded in leaves and diffuses into source tissue until it reaches the phloem, no distinct front but a gradual increase in fluorescence results in the transport phloem. The problem becomes especially obvious when neighboring companion and

52 parenchyma cells light up almost as fast as the sieve tubes. Since the quality and speed of loading is dependent on multiple factors, it has not yet been possible to standardize the procedure to always obtain the same loading. Therefore, we were not yet able to exclude the possibility that sieve tubes containing large agglomerations did not actually translocate but that the fluorescence diffused from neighboring tubes.

To unequivocally prove that the tubes are actively translocating, we conducted studies on real-time movement of fluorescent dyes within individual sieve tubes. We grew plants in Micro-ROCs, loaded them with CFDA, and photobleached CFDA in the tube to produce a distinct front of fluorescence and imaged refilling at 0.3-s intervals (fluorescence recovery after photobleaching [FRAP]). Since the laser of a confocal microscope can be directed with pixel size accuracy, precise areas can be targeted. Figure 6G shows three frames of a FRAP experiment of the tube shown in Figures 6E and 6F. Refilling occurs at a velocity of ~60 μms-1 downstream of the obstruction (see Supplemental Movie 2 online).

There is no other sieve tube or lateral sieve plate that would allow bypassing the agglomeration. We therefore conclude that transport occurs through agglomerations.

Currently, phloem translocation is thought to be driven by an osmotically generated pressure differential. Sieve tubes supposedly provide a channel of adequately low hydraulic resistance permitting pressure differential driven flow. The presence of agglomerations in the flow path necessitated a reevaluation of the feasibility of a pressure flow. To calculate the increase of resistance by obstructions, the resolution of confocal microscopy is insufficient. Only TEM permits precise measurements. TEM sections, on the other hand, are

53 only in the range of 80-nm thick and the agglomerations are rare in comparison to unobstructed areas. To section through a single sieve element of 120-μm length, 1500 individual cross sections are required, and on average, only every tenth sieve element contains an agglomeration. By conducting serial cross sectioning, we were able to find two sieve plates and one area of obstruction. At the sieve plate, numerous filaments are located that traverse the pores (Figure 6H1). Right behind the sieve plate, ~55% of the lumen is obstructed by filaments (Figure 6H2). With increasing distance, the filaments are located further toward the margins (Figure 6H3) until they form distinct bundles (Figure 6H4).

Sieve plates in Arabidopsis are often not strictly perpendicular to the tube axis (Figure 6I1).

The sieve plate in Figure 6I also contains filaments on the plate. While some pores are open, others contain several filaments that obstruct a certain percentage of the pore’s lumen (Figure 6I3). The only agglomeration we found so far is shown in Figure 6J. The distance between image Figure 6J1 and 6J4 is ~7 μm. Major parts of the lumen are filled with filaments with the exception of an area of ~0.5 x 1 μm. Average filament diameter is

21.8 ± 2.5 nm (n = 100) with 6.1 ± 1.3 nm (n = 43) spacing between filaments within the agglomeration.

To see if the abundance of filaments and bundles is a general feature of dicot sieve tubes, we investigated two nonrelated plant species. While Arabidopsis has developed into an important model plant, ultrastructural studies are very limited. We chose tobacco

(Nicotiana tabacum), since extensive ultrastructural data are available, and black cottonwood (Populus trichocarpa) as model tree species. Preservation of some ultrastructural features was not as good as in Arabidopsis, and modification of the fixation

54 protocol will be required in the future. However, filaments and bundles can be seen in the periphery of sieve elements (Figures 7A to 7C).

Figure 6: Obstructions in Arabidopsis Sieve Tubes. (A) to (D) Protein agglomerations (cyan) in the lumen of sieve tubes are variable. In many sieve elements, filaments are located at the margins of the cells. (A) The presence of filaments on the sieve plate (arrow) outlines their location. (B) A larger agglomeration of P protein (dashed arrows) on both sides of a sieve plate (solid arrow). The P protein agglomerations fray out into filaments. Some of the filaments connect through the sieve plate. (C) and (D) Overview images of CFDA (red) translocating sieve tubes containing massive P protein agglomerations. Sieve plates (solid arrows) are often not directly covered with P protein agglomerations. Some agglomerations appear to completely fill the lumen of the tube (dashed arrows), while others only cover part of it (open arrows). (E) Two P protein agglomerates. The upper agglomeration frays out into filaments. Some darker spots indicate the location of organelles, in this case most likely mitochondria. The lower agglomerate is completely amorphous. (F) The same sieve tubes as shown in (E). The upper file is fully mature and translocates CFDA (red) despite the presence of the large P protein agglomeration (dashed arrow) in front of the sieve plate (solid arrow). The lower tube is not fully mature. The amorphous P protein body (arrowhead) has not transformed into strands and is not translocating CFDA, while the next sieve element on the left in the same file is in the transition phase. (G) Three consecutive images of a FRAP experiment. The dashed arrow indicates the location of the P protein agglomeration shown in (E). The tube has been bleached by the laser and quickly refills after decrease of the laser energy indicating transport. (H) Four TEM images of a serial section of a sieve tube in the area of the sieve plate. (H1) to (H4) A cross section through the plate shows several open pores in the center, while significant portions at the margin of the plate are covered with filaments (H1). In consecutive sections (~1 μm apart from each other), filaments fill >50% of the lumen (H2) and move toward the membrane (H3) until they form discrete bundles (H4). (I) Serial section through an Arabidopsis sieve plate, oriented in a slight angle in relation to the sieve tube. (I1) and (I2) While most pores are open (I1), filaments are present on the plate (I2). (I3) Higher magnification of sieve pores in the sieve plate shown in (I2) (box). SEOR1 filaments can be seen in some pores ([I3], arrows). (I4) A few micrometers behind the plate, filaments move toward the margins. (J) Four images of a serial section through the lumen of a sieve tube containing an agglomeration. Major parts of the lumen are filled with P protein filaments, but a channel is unobstructed. The filaments are mostly oriented in parallel and have a pseudocrystalline appearance. A sieve element plastid is abundant in J4. The distance from (J1) to (J4) is 7 μm. Bars = 10 μm in (A) and (B), 25 μm in (C), (D), (F), and (G), 5 μm in (E), and 500 nm in (H) to (J).

In contrast with Arabidopsis, sieve element plastids of tobacco are decorated with filaments (Figure 7A). The size of the filaments differed slightly from that of Arabidopsis with an average of 18.68 ± 2.1 nm in tobacco and 23.88 ± 2.1 nm in black cottonwood.

2.3.4 Sieve Tube Structure and Its Impact on Phloem Translocation

The structures found in whole-plant freeze substitutions differ significantly from what has been described using other preparation and fixation protocols. The large amount of Arabidopsis SEOR1 filaments in the translocation path inevitably leads to the question of its impact on tube resistance.

Figure 7: SEOR1-Like Filaments in Tobacco and Black Cottonwood. (A) A cross section through a tobacco sieve element (SE) shows several sieve element (SE) plastids covered with Arabidopsis SEOR1 filaments and bundles. (B) A tangential section through a tobacco sieve element along the organelle containing layer close to the plasma membrane. A large SEOR1 bundle of multiple filaments covers the membrane. (C) Longitudinal section through a black cottonwood sieve tube. The preservation is not as good as in Arabidopsis and tobacco, but Arabidopsis SEOR1-like filaments are visible. Bars = 1000 nm (A) and (B) and 150 nm in (C).

The question is: Can a pressure flow, as discussed by Münch (1930), drive phloem translocation? Figure 8 shows a schematic summary of our findings in Arabidopsis sieve

57 tubes. In recent phloem flow calculation models, sieve tubes were expected to be empty tubes and the space occupied by organelles and other structures is considered to be below the error of sieve tube geometry measurements (Thompson and Holbrook, 2003;

Mullendore et al., 2010). In reality, however, even in areas without significant SEOR1 accumulation, a major fraction of the tube lumen turned out to be unavailable for translocation.

Figure 8: Schematic Reconstruction of an Arabidopsis Sieve Tube. Reconstruction of the structure of a sieve element-companion cell complex as found in in vivo confocal studies and after freeze substitution of whole plants. Sieve elements contain ER, mitochondria covered with clamp proteins, and electron-dense vesicles. While those structures are usually embedded in an amorphous ground matrix, SEOR1 filaments and sieve element plastids are always in direct contact with the sieve tube sap. A SEOR1 agglomeration is shown in front of a plate that does not fill the entire lumen of the sieve element. Companion cells contain all organelles typical for a plant cell, but only nucleus, vacuoles, chloroplasts, and mitochondria are shown. Blue lines indicate the location of a

58 cross section for (A) to (C). C, chloroplast; Cl, clamp proteins; EV, electron-dense vesicles; GM, ground matrix; M, mitochondria; N, nucleus; P, plastid; SR, SEOR1 filaments; V, vacuole.

Usually, up to 30% (Figure 4G) are occupied by sieve tube constituents. Even in the wider tubes of tobacco, up to 35% (Figure 7A) of the lumen is filled with sieve element plastids and other organelles. On top of this, SEOR1 agglomerations need to be included in the calculations.

To quantify the effects of the organelles and SEOR1 filaments on the flow, we calculate the influence of these on the hydrostatic pressure difference between source and sink tissues required to drive the observed flow. For simplicity, we consider a single sieve tube as a proxy for the phloem and model the translocation pathway as consisting of a collection of approximately cylindrical sieve tube elements lying end to end separated by sieve plates. This approach has been widely used in previous studies of phloem transport

(for example, see Thompson and Holbrook, 2003, and references therein). The relation between the hydrostatic pressure drop ∆p between source and sink and the volumetric flow rate Q through the sieve tube is the hydraulic equivalent of Ohm’s law (Bruus, 2008)

∆p = RQ. (1)

Here, the volumetric flow rate Q = UA is the product of the flow velocity U and cross- section area A of the sieve element, and R is the hydraulic resistance of the phloem translocation pathway. Due to the abundance of sieve tube constituents at the margins, we estimate that between 65 and 100% of the area is open to flow, such that the cross-section area A lies in the range A ≈ (4.5 – 7.1) μm2. Typical flow speeds observed are of the order U

≈ 100 μm/s, which yields Q ≈ (450 – 710) mm3/s. When calculating the resistance R in

Equation 1, we take into account three major components: (1) the tube lumen including organelles, (2) the sieve plate, and (3) the SEOR1 agglomerations. Assuming that the translocation pathway consists of N ≈ 1250 identical sieve tube elements, M ≈ N/10 = 125 of which contain a SEOR1 agglomeration, we write the resistance of the phloem translocation pathway R as

R = NRlumen + (N - 1)Rplate + MRplug. (2)

Here, Rlumen is the resistance of a single sieve tube element lumen, Rplate is the resistance of a single sieve plate separating adjacent sieve elements, and Rplug is the resistance of a single SEOR1 agglomeration. Please refer to the Supplemental Appendix A for a detailed discussion of how these resistance values are determined and Table 1 for a list of characteristic values of the parameters used. As shown in Table 2, we find typical values of the terms in Equation 2: NRlumen ≈ (0.98 – 2.4) x 1020 Pa s m-3, (N – 1)Rplate ≈ 1.9 x

1020 Pa s m-3, MRplug ≈ 4.4 x 1019 Pa s m-3. While the contribution from the lumen and plate resistances are of comparable magnitude, the contribution from the SEOR1 agglomerations is somewhat smaller, reflecting the fact that these are only found in every ~10 sieve tube elements. We finally have for the total resistance in Equation 2 that R ≈ (3.3 – 4.7) x 1020Pa s m-3 and find from Equation 1 that the pressure drop required to drive the flow over a distance of 15 cm lies in the range ∆p ≈ (0.21 – 0.23) MPa.

Table 1. List of Parameters for Flow Calculations

Parameter Symbol/expression Value, unit, reference

Sieve tube cross section area

Effective Sieve tube cross section area

At SEOR1 agglomeration cross section area

Effective sieve tube radius 1.2 μm

At SEOR1 filament radius a 10 nm f

At SEOR1 agglomeration opening 0.5 µm radius

Average sieve pore radius 156 nm

Sieve tube radius 1.5 µm

Eff. sieve tube diameter 2.4 µm

At SEOR1 filament diameter 20 nm

At SEOR1 agglomeration opening 1 µm diameter

Sieve tube diameter 3.0 µm

Observed flow speed 100 µm s-1

At SEOR1 filament separation b 6 nm distance

Permeability of At SEOR1 agglomeration

Length of plant 15 cm

At SEOR1 agglomeration length 6 µm

Sieve element length 120 µm

Sieve plate thickness 450 nm

Number of sieve elements 1250

Average number of sieve pores 15

Number of At SEOR 1 agglomerations 125

Volume flux m3 s-1

Hydraulic resistance of the phloem Pa s m-3 translocation pathway

Viscosity 1.3 mPa s (Deeken et al.,

2002; Hunt et al., 2009)

Non-dimensional permeability of At

SEOR1 agglomeration

Volume fraction occupied by 0.45 filaments inside agglomeration

Reference is given next to parameter value when not measured by the authors.

Table 2. Parameters Relevant for the Calculation of the Pressure Drop ∆p in Equation 1/(A1)

[MPa

[μm [μm] [Pasm [Pa [Pa [Pa [Pa [Pa [ ]

-3 ] ] s ] s/ ] s/ ] s/ ] s/ ]

[Pa

s ]

(†) (†) (†)

Calculated values of the lumen resistance Rlumen, plate resistance Rplate, agglomeration resistance Rplug, and total resistance R determined from Equations (A3), (A4), (A5), and

(A2) (see Supplemental Appendix 1 and Supplemental References 1 online). The results are given for two values of the effective sieve tube diameter de and three values of the agglomeration opening diameter do. de = 3.0 μm corresponds to a completely empty sieve

64 tube, and de = 2.4 μm corresponds to a sieve tube with only 65% of the area open to flow.

Results marked with an asterisk indicate the measured value of do = 1 μm. Results marked with (†) indicate the case where no At SEOR1 agglomerations are present.

2.3.9 SEOR1 Function

Since SEOR1 is located in the flow path of sieve tubes, we tested a potential influence on translocation. We used the homozygous SEOR1 T-DNA insertion mutant

(GABI-KAT 609F04) and conducted flow velocity studies along intact roots. We studied the flow in eight independent plants of each wild type and T-DNA insertion mutant (Figures 9A to 9D). Velocities in the root system are variable in both lines. So far, no significant difference between insertion mutant and the wild type has been found (Figures 9A and 9B).

We further measured the average sieve tube diameter of the two lines to see if fewer obstructions lead to a change in tube anatomy. Average sieve tube diameters did not differ significantly between the lines.

Our study was initiated because the genomic SEOR1 sequence in Arabidopsis showed homology to genomic sequences of Medicago forisomes (Pélissier et al., 2008) and therefore was likely to be a yet unknown P protein. This relationship also suggested a similar function. Forisomes appear to form reversible agglomerations that temporarily stop sieve tube flow (Knoblauch et al., 2001, 2003; Peters et al., 2006). The transformation from the low volume to the high volume state of a forisome may be completed in 100 ms

(Peters et al., 2008). Thus, we tested SEOR1 for a potential injury reaction. Initially, we observed intact sieve tubes for several hours by epifluorescence and confocal microscopy without any indication of dynamic behavior of SEOR1 bundles and agglomerations. Then, we tested different injury stimuli that are known to trigger the forisome reaction, such as local mechanical injury, distant burning of leaf tips, and local cold shocks (Furch et al.,

2007; Thorpe et al., 2010).

Figure 9: In Vivo Flow and Injury Experiments. (A) and (B) Comparison of phloem flow velocities along a main root of the Arabidopsis wild type (A) and SEOR1 T-DNA insertion mutant (B). The entire root system is visible in MicroROCs after loading with CFDA, permitting flow measurements in individual tubes by FRAP. No significant difference was found between mutant and wild-type plants. (C) and (D) FRAP experiment on an individual tube. Three frames from Supplemental Movie 2 online (C). After bleaching of CFDA, the laser intensity was lowered and refilling of

67 the tube was monitored at subsecond intervals. Regions of interest are marked along the tube (arrows and colors of arrows correspond to colors in graph), and fluorescence intensity is measured and graphed (D), giving a direct reading of flow velocity in the tube. (E) and (E1) to (E4) Four frames of Supplemental Movie 3 online, showing the slow movement (flow is right to left) of SEOR1-YFP filaments through a sieve plate (arrow). Movement does not stop even after 23 min. Bars = 5 μm in (A) and (B), 100 μm in (C), and 10 μm in (E).

None of the treatments triggered any immediate reaction. Even direct application of 1 to 5 mM Ca2+ medium on ruptured sieve tubes or isolated SEOR1 bundles did not result in any structural changes.

Although we were not able to find reactions equivalent to that of forisomes, SEOR1 in Arabidopsis sieve tubes underwent an obvious structural alteration after tissue excision and standard fixation for TEM (Figures 3A and 3B). To understand the development of these structures, we conducted time-lapse movies of injured sieve tubes. A very slow movement of SEOR1 toward the sieve plate was observed in some cases. Surprisingly, the movement did not stop at the sieve plate but agglomerates continued to move through the plate for extended periods of time (Figure 9E; see Supplemental Movie 3 online).

2.4 Discussion

2.4.1 Sieve Tube Ultrastructure

Fluorescent tagging of SEOR1 filaments permitted comparison of in vivo confocal micrographs of sieve tubes with TEM images collected from variably processed tissue.

Freeze substitution of whole plants most accurately resembled the in vivo structure and

68 location of components found in confocal images. Freeze substitution, however, has some limitations. For good preservation, a high sugar content that acts as antifreeze substance is necessary. In addition, a close location to the surface is required. So far, we were only able to preserve sieve tubes for TEM in source leaves. The coverage of root sieve tubes with rhizodermis, large cortical parenchyma cells, endodermis, and pericycle in combination with a lower sugar concentration in sink sieve elements has so far prevented us from studying root sieve tubes. By contrast, in vivo confocal investigations using Micro-ROCs are possible in roots only. Thus, we are currently not able to compare phloem structures in the same organs. On the other hand, removal of some cortical cell layers at the main vein of source leaves exposes uninjured sieve tubes, which show the same fine structure as found in root sieve tubes by confocal microscopy. We conclude that the different location probably has just a minor influence on sieve tube structure. Preservation of phloem tissue in larger plants may become increasingly difficult since sieve tubes are usually covered by a thicker tissue layer, which may increase problems with freezing artifacts. Specific treatment, such as localized chilling, which halts phloem translocation but not loading

(Pickard and Minchin, 1992), might become necessary to increase sieve tube antifreeze concentrations. Specific protocols may have to be developed for different plant species.

Usually, membrane structures were more difficult to preserve. This may be due to the acetone solvent. The addition of water or tannic acid helps to some extent, but in general, standard chemical fixation shows a more pronounced outline of ER stacks. Alterations of the fixation protocol might help solve this problem in the future. However, within those

69 limitations, the method has proven most beneficial, since structures in TEM images match the location and distribution of structures found in translocating sieve tubes.

To understand the mechanism of, for example, long-distance transport, the interactions of sieve tubes with pathogens such as aphids or viruses and the interactions of sieve elements and companion cells, a good understanding of the cellular equipment available for those interactions is fundamental. Besides the well known previously described sieve tube components, mitochondria, sieve element plastids, and ER, some new, frequently found components have to be added to our picture of sieve tube infrastructure. Mitochondria in

Arabidopsis are always surrounded by a halo of small protein spikes that attach them to membranes and/or SEOR1 filaments (Figure 4). Similar clamps have been found in one earlier study in tomato (Solanum lycopersicum) and fava bean (Ehlers et al., 2000). Clamp proteins do not attach to all organelles in all species. In Arabidopsis, mitochondria are completely covered, while sieve element plastids lack clamps (Figure 4F). In Vicia and

Solanum, clamps are present on all organelles, including the ER. In contrast with other organelles, mitochondria in Arabidopsis have a layer of clamp proteins each, doubling the distance between the organelles (Figure 4H).

In addition, we found electron-dense vesicles of various sizes. The vesicles seem to bud off of membranes. The nature of the membranes and vesicles has yet to be established.

Vesicles are always embedded in the ground matrix of the parietal layer.

The structures that clearly stand out are SEOR1 filaments. Sieve plate pores are mostly unobstructed, but large SEOR1 agglomerations exist in the lumen of some

70 translocating sieve tubes. Agglomerations have frequently been observed previously.

However, the ultrastructure and location of the agglomeration differs depending on the preparation used. After standard fixation of sectioned tissue, Arabidopsis sieve plate precipitates consist of fine 5- to 10-nm-thick filaments (Figures 3A and 3B). The precipitates are located on the plate or in the lumen, but no filaments are found at the margins that would match the location of in vivo confocal images. In tobacco, different forms of P proteins have been described to occur after standard preparation and fixation, including 23-nm tubules designated as P1 protein, 15-nm striated filaments designated as

P2 protein, very fine filaments (Cronshaw and Esau, 1967; Gilder and Cronshaw,

1973), and crystalline filaments of ~100 nm diameter (Johnson, 1969). The diameters reported may vary depending on the study. All filaments are located in the lumen or the pores but do not form a distinct meshwork at the margins. In Cucurbita maxima, phloem protein1 (PP1) is a 96-kD protein that forms filaments, and PP2 is a 25-kD dimeric lectin that binds covalently to PP1 (Bostwick et al., 1992; Golecki et al., 1999). The structural component PP1 belongs to a gene family found only in cucurbits (Clark et al., 1997;

Beneteau et al., 2010). By contrast, genes encoding SEOR proteins have been reported in many dicot families (Pélissier et al., 2008; Huang et al., 2009, Rüping et al., 2010).

In Arabidopsis and tobacco, freeze substitution of whole plants results in only one morphological form of P protein: filaments of ~20 nm diameter, which are absent in SEOR1

T-DNA insertion mutants. Our data suggest that many P protein structures described are

71 alterations of SEOR proteins due to preparation and that P proteins usually exist in the form of SEOR filaments in active sieve tubes.

2.4.2 Phloem Translocation

The controversy about the mode of phloem translocation in the last century mainly revolved around the question of the abundance of P protein agglomerations inside the sieve tube lumen and inside sieve plate pores and has split phloem researchers into two groups. While investigators believing in occluded plates favored the electroosmotic theory

(Fensom, 1957; Spanner, 1958, 1970), the pressure flow hypothesis was supported by the group believing in open pores and that occlusion is due to preparation artifacts (for an overview, see Knoblauch and Peters, 2010). Over the years, gentle preparation methods for

TEM (e.g., Fisher, 1975; Turgeon et al., 1975; Lawton and Newman, 1979) and in vivo studies on translocating sieve tubes (Knoblauch and van Bel, 1998) supported an unobstructed sieve tube path. To date, an osmotically generated pressure flow is generally accepted as the mode of action of long-distance translocation in sieve tubes. In this context, our finding that massive SEOR1 agglomerates are a standard feature in the lumen of translocating sieve tubes in Arabidopsis is most surprising. In the end, both groups of investigators were right. Most of the pores are usually unobstructed, but massive agglomerates exist in the lumen. The resulting question is: Is a pressure differential driven flow possible?

We calculated the pressure differential required to drive flow through a 15-cm-long

Arabidopsis sieve tube from a source leaf to the root at a velocity of 100 μm s-1 to be 0.2

MPa. The osmotic concentration of Arabidopsis sieve tube sap in source tissue can be taken from measurements of sap collected by stylectomy to be 0.7 M (Deeken et al., 2002; Hunt et al., 2009), which can generate a pressure of ~1.7 MPa or less, depending on the osmolarity of the apoplastic solution. Not all of this pressure is available for transport, since sink cells possess a turgor of ~0.7 MPa (Pritchard, 1996; Turgeon, 2010). This leaves a maximum of

1 MPa pressure differential for flow. For our calculations, we assumed all parameters to be at the lower end of observation and to favor pressure flow. All pores were assumed to be unobstructed, the average occupation of cross-sectional area with organelles was assumed to be only 20%, the surface of the parietal layer was assumed to be smooth, the channel in the SEOR1 agglomeration was assumed to be 1 μm in diameter, and the tubes were assumed to be circular. The calculated pressure differential required is ~0.2 MPa. Even if we assume less favorable conditions, the potential 1 MPa pressure differential leaves plenty of margin in comparison to the calculated 0.2 MPa required pressure, since the agglomeration opening would have to be smaller than 500 nm in diameter in every agglomeration to increase the required pressure to more than 1 MPa.

The situation, however, changes if we assume that SEOR1 agglomerations do not contain open channels. Unfortunately, serial sectioning for TEM is so labor intensive that we were only able to find a single SEOR1 agglomeration. This agglomeration had a channel of ~0.5 x 1 μm. In vivo confocal images, on the other hand, show a variety of agglomerations. Confocal resolution is ~230 nm and would have allowed us to identify the

73 opening in the agglomeration shown in Figure 6J. In many confocal images, however, agglomerations appear to fill the entire lumen (Figures 6C to 6E), which would increase the required pressure significantly.

To calculate the impact of agglomerations on flow, the porosity of the material is critical. For our calculations, we assumed filaments in the agglomeration to be straight rods, with a smooth surface and without major interaction with the surrounding medium, similar to glass filaments. Proteins, especially if they contain a high percentage of charged amino acids, form a hydration shell and turn the surrounding water into a viscous layer, which increases the effective filament diameter significantly (Bánó and Marek, 2006). In this case, a single agglomeration without an opening would add considerable resistance to the flow and would most likely be sufficient to block the flow entirely. Assuming flow favoring conditions with filaments lacking hydration shells, ~10 agglomerations would be needed to increase the resistance to the point that the calculated pressure differential of 1

MPa would not be sufficient to drive flow at the measured velocities (Table 2).

In summary, despite the existence of large SEOR1 agglomerations in the lumen of sieve tubes, a pressure differential–driven flow appears feasible, given that the porosity of

SEOR1 agglomerations is high. It is, however, surprising that there is no significant difference in transport velocity between the mutant and wild type. The existence of agglomerations necessarily has an influence on tube resistance and must result in a higher pressure differential in wild-type plants to maintain constant flow velocities.

The pressure flow hypothesis remains an issue of debate. While the tube anatomy does appear to scale with plant size (Jensen et al., 2011), pressure does not (Turgeon,

2010). Also, larger tubes with significantly lower resistance translocate at slower velocities than tubes with higher resistance (Mullendore et al., 2010). It appears necessary to conduct correlated determinations of translocation velocity, pressure differential, and sieve tube structure.

2.4.3 SEOR1 Function

It has been repeatedly suggested that P proteins are involved in sieve tube occlusion, and the discovery of forisome function supported this notion (Pélissier et al.,

2008). On the other hand, it had also been questioned whether occlusion is the (only) function of P proteins (Sabnis and Sabnis, 1995).

The reaction of SEOR1 to injury is not comparable to the reaction found in forisomes. SEOR1 filaments do not show a detectable structural reaction to Ca2+ ions, nor do they react within milliseconds. TEM snapshots of injured sieve tubes gave the impression that SEOR1 occluded pores (e.g., Figure 3B). SEOR1 filaments were supposed to be compressed by callose formation to form a tight seal (Mullendore et al., 2010). In reality, there is a slow movement of SEOR1 through the pores, which can continue for at least 45 min. SEOR1 often moves out of the wound site and disappears in the surrounding medium.

This explains why high concentrations of P protein filaments can be found in phloem exudates (Cronshaw et al., 1973) in different plant species, including tobacco. This argues

75 against a targeted sealing mechanism and also suggests that callose occlusion is relatively inefficient, at least in Arabidopsis.

The structure of sieve tubes is probably the least understood of all major plant cell types. Although we have known of the existence of sieve element plastids for almost a century, and meanwhile have studied the function of all other plastid types in detail, we still have no indication of sieve element plastid function. The function of many other structures, including SEOR1, is also obscure. Because the phloem is a key player to maintain plant integrity, it will be crucial to obtain more detailed insights into the functions of its components. Sieve elements are far from being empty tubes. The existence of a protein filament meshwork that structurally resembles a cytoskeleton may lead to new insights in short- and long-distance signaling, plant–pathogen interaction, such as viral movement, and, among others, sieve element–companion cell interactions. Some of the sieve tube components might have a direct effect on translocation and/or flow control. The tools to investigate these open questions by in vivo studies on a cellular basis are now available.

2.5 Methods

2.5.1 Plant Material for Freeze Substitution

Arabidopsis thaliana ecotype Columbia, SEOR1 T-DNA insertion mutant GABI-KAT

609F04, and tobacco (Nicotiana tabacum) were grown on 0.44% (w/v) Murashige and

Skoog (MS) medium containing 87.6 mM Suc, 2.56 mM MES buffer, pH 5.8, and 0.8% (w/v) agar. Seeds were surface sterilized with 70%ethanol, plated, and cold treated overnight at

4°C before being placed into the growth chamber. The plants were grown at 25°C with a

16/8-h light/dark period. Black cottonwood (Populus trichocarpa) was grown in pots in a greenhouse at 23°C, with 60 to 70% relative humidity, and a 14/10-h light/dark period

(daylight plus additional lamp light [model PL 90; PL Lighting Systems]) with a minimum irradiance of 150 μE m-2 s-1.

2.5.2 Micro-ROCs

Plants were grown in Micro-ROCs (Advanced Science Tools) in the greenhouse at a

14-h photoperiod, 300 to 400 μE m-2 s-1, at 20°C day and 15°C night. Plants were grown to the six- to eight-leaf stage for SEOR1-YFP imaging. FRAP plants were grown until the first true leaves matched the diameter of the cotyledons.

2.5.3 Plunge Freezing and Freeze Substitution

Liquid nitrogen was placed into a shallow, thick-sided polystyrene container and placed under vacuum for ~7min until the nitrogen became slushy. Whole Arabidopsis plants, in the four-leaf state, were gently teased from the MS agar and rapidly plunged into the slush nitrogen. The frozen plants were then transferred to 2% glutaraldehyde in acetone with 0.1% water, 0.1% tannic acid, or 4% tannic acid in scintillation vials on dry ice. The plants in solution were transferred to -80°C for 24 h and then placed into a -20°C

77 freezer while removing most of the dry ice. The solution was allowed to ramp up to -20°C over a period of at least 8 h. The plants were rinsed twice for 30 min with cooled (-20°C) acetone. Postfixation was achieved in cooled (-20°C) 2% OsO4 in acetone overnight. The material was ramped to 20°C over a period of 6 h. The OsO4 was rinsed with acetone two times for 30 min and exchanged for propylene oxide (PO). The plants were infiltrated with a soft recipe Spurr’s resin (SR; Bozzola and Russell, 1999) as follows: 3:1 PO:SR, 48 h; 2:1

PO:SR, 24 h; 1:1 PO:SR, 24 h; 1:2 PO:SR, 24 h; 1:3 PO:SR, 24 h; 100% SR, 24 h; 100% SR, 48 h; 100% SR, 24 h. Before each exchange, the samples were cycled three times in vacuum for

5 min each cycle. The samples were embedded in fresh SR and cured for 2 d at 60°C.

Ultrathin sections (70 to 100 nm) were taken with an ultramicrotome (Reichert Ultracut R;

Leica) and placed on formvar-coated slot grids. They were stained with a solution of 1% uranyl acetate and 0.01% potassium permanganate for 10 min and poststained for 6 min in

Reynolds lead citrate (Reynolds, 1963). The sections were imaged on a FEI Tecnai G2 TEM

(FEI Company) or a Philips CM200 UT Intermediate Voltage TEM (FEI Company).

2.5.4 Epifluorescence Microscopy

Epifluorescence microscopy was performed with a Leica DM LFSA microscope or with a Leica MZ8 stereomicroscope. Images and movies were recorded with a Leica DFC

300FX-cooled charge-coupled device camera. To show the outline of root cells, 0.1 mg/mL synapto-red (EMD Chemicals) was added to agar plates. For synapto red and YFP double

78 labeled tissue, a Leica filter cube I3 was used, and for YFP or CFDA detection, a GFP filter cube was used.

2.5.5 Confocal Microscopy

All confocal laser scanning microscope images were obtained with a Leica TCS SP5.

Respective excitation and emission for YFP, GFP, GFP5, and CFDA were 514 argon/520 to

550, 488 argon/500 to 600, 405 diode/ 475 to 530, and 488 argon/490 to 515. Subsequent processing used ImageJ for time series and Leica LAS AF Lite software for images. For flow velocity, measurements were conducted with plants in the four-leaf state grown in micro-

ROCs.

2.5.6 FRAP

CFDA was loaded into the first true leaves and cotyledons by half clipping and applying 20 mL 1:5 (v/v) 50 mg mL-1 CFDA in acetone to water. Loaded sieve elements in the primary root were manually photobleached at 488 nm at maximum laser intensity, pinhole at Airy 3, and at x8 zoom, starting apically and moving toward the hypocotyl. A 3- frames per second time series, to record refilling of the sieve element, immediately followed the reduction of the laser power to 15% and zoom to x1. Region-of-interest intensities were generated using Leica LAS AF Lite software.

2.5.7 Cloning and Transformation: SEOR1-YFP

The Modular Binary Construct System (gift from Christopher G. Taylor) was used for all constructs. The K4 adapter made from 5’-TTCGGATCCACTAGTTCTGCTGCTGGTTCTG-

CTGCTGGTTCTGGGGGATCCCTT-3’ and 5’-AAGGGATCCCCCAGAACCAGCAGCAG-

AACCAGCAGCAGAACTAGTGGATCCGAA-3’, which contains a unique SpeI restriction site was cloned into the BamHI site of a modified AKK 1435 vector containing the YFP gene and sequenced for directionality. SEOR1, minus the stop codon, and its 1500-bp promoter region was amplified from BAC clone F4P13 with 5’-

TCGGTACCGAACTAATACACAAGTAACACA-AGT-3’ and 5’-

TTCACTAGTGAAGTTGTAGTTCTCGTCTT-3’. This was ligated into the AKK 1435 shuttle vector at the KpnI and SpeI sites. The PacI promoter-gene fusion cassette was then ligated into the AKK 1426b binary vector containing in planta glufosinate resistance (Thompson et al., 1987). The construct was used to transform Arabidopsis ecotype Columbia via

Agrobacterium rhizogenes 18r12v using the floral dip method (Clough and Bent, 1998), and the transformed seeds were screened with daily spraying of 0.003% glufosinate ammonium (Sigma-Aldrich) and 0.05% Silwet L-77. T2 generations were screened by epifluorescence microscopy to identify homozygous lines.

2.5.8 GFP5-ER

GFP5-ER was amplified from pBINmGFP5ER (Haseloff et al., 1997) with primers 5’-

TTCAA-GCTTAAGGAGATATAACAATGAAGACTA-3’ and 5’-TTCGGATCCGATCTAGTAACA-

TAGATGACACC-3’ and subsequently cloned into AKK 1408 at the 3’ end of the 2047-bp

Medicago truncatula SEO2 promoter (Pélissier et al., 2008). The Pro-Mt-SEO2-GFP5-ER cassette was then cloned into binary vector AKK 1426b via SdaI. Arabidopsis expressing At

SEOR1-YFP was transformed with Pro-Mt-SEO2-GFP5-ER by A. rhizogenes 18r12v using the floral dip method (Clough and Bent, 1998) with seeds screened on MS plates containing

50 mg mL-1 kanamycin.

2.5.9 SEOR1-GFP

Approximately 1000 bp of promoter sequences extending 5’ from, but not including, the translation start codon of SEOR1 were PCR amplified from the bacterial artificial chromosome F4P13. The amplicons were initially cloned into the pGEM-T easy vector, and specific primers were designed to subclone the amplicons into SalI and XbaI restriction sites located 3 bp 5’ of the translation initiation codon of the GUS reporter gene (uidA) in the pGPTV-Kan binary vector (Becker et al., 1992). The enhanced GFP gene was PCR amplified and subcloned into the pGPTV-Kan binary vector in place of the uidA gene using the SmaI andKpnI restriction sites, and these primers also created a multiple cloning site at the 3’ end of the enhanced GFP gene. Subsequently, the SEOR1 open reading frame was PCR amplified and subcloned into this multiple cloning site (KpnI and ApaI). The binary vectors were transformed into Agrobacterium tumefaciens strain GV3101 and used to transform

Arabidopsis by the floral dip method (Clough and Bent, 1998). Transgenic plants were then screened on kanamycin-supplemented media.

2.5.10 T-DNA Insertion Mutants

T-DNA insertions in SEOR1 were identified using T-DNA Express

(http://signal.salk.edu/cgi-bin/tdnaexpress). Seeds for GABI-KAT 609F04 (SEOR1 knockout) were obtained from the Genomanalyse im Biologischen System Pflanze. Plants from the original seed stocks or one generation later were screened to identify individual homozygous plants using the PCR based screening technique according to the method of

Siebert et al. (1995). The GABI-KAT 609F04 mutant contained a second T-DNA insertion so plants were allowed to self-fertilize and plants homozygous for the SEOR1 insertion alone were identified.

Successful knockout of the gene was confirmed using RT-PCR. In brief, total RNA was extracted using the Trizol method, and total RNA was reverse transcribed using

SuperScript II according to the manufacturer’s instructions. Partial, intron-spanning sections of the gene were amplified using gene-specific primers, including section 1 (1 to

351 bp) 5’-ATGGAGTCGCT-GATCAAGTC-3’ and 5’-TATCTCGCAGGCAACACG AT-3’, section 2

(860 to 988) 5’-ACC-ATCTCGCTGAGACCTTGAGG-3’ and 5’-

GGCCGTGAGAATCTTCATGTTATCA-3’, section 3 (1494 to 1659) 5’-GAGAGAGACCT-

TTTCCCTTAACCTCA-3’ and 5’-TTCACGT-TGGAATCTTTGGCC-3’, and subsequently visualized on a 1.6% agarose gel containing ethidium bromide.

2.5.11 Immunolocalization

Cross sections of unfixed floral stems from Arabidopsis Columbia plants and GABI-

KAT 609F04 were cut with a vibrating microtome (Vibratome) at 50 μm and collected in

PBS. Sections were washed twice in 10 mM PBS and then incubated for 30 min in blocking buffer (PBS with 3% nonfat dry milk). Sections were washed twice more with PBS and incubated for 45 min with the RS21 primary monoclonal antibody in blocking buffer

(1:100). After incubation with primary antibody, the sections were washed three times with PBS and then incubated in PBS with ALEXA 488-nm fluorescently tagged secondary goat anti-mouse antibody (Molecular Probes) (1:250). Finally, the labeled sections were washed twice with PBS and once with nanopure water and observed under a Nikon E600 epifluorescence microscope, with an excitation wavelength of 490 nm and an emission wavelength of 512 nm.

2.5.12 Accession Numbers

Sequence data from this article can be found in the Arabidopsis Genome Initiative or

GenBank/EMBL databases under the following accession numbers: At3g01680 (SEOR1) and GK-609F04-021864 (GABI-KAT 609F04).

2.6 Supplemental Data

The following materials are available in the online version of this article.

Supplemental Movie 1. SEOR1 in Root Tip.

Supplemental Movie 2. Real-Time Imaging of Phloem Flow.

Supplemental Movie 3. SEOR1 Movement in Injured Sieve Tubes.

Supplemental Movie Legends. Legends for Supplemental Movies1 to 3.

Supplemental Appendix 1. Mathematical Derivation of Expressions for the Three

Resistances given in Equation (2).

Supplemental References 1. Supplemental References for Supplemental Appendix 1.

2.7 Acknowledgments

We thank Karl J. Oparka (University of Edinburgh) and Winfried S. Peters (Indiana

University–Purdue University Fort Wayne) for helpful discussions and critical reading of the manuscript. We acknowledge technical support from Washington State University’s

Franceschi Microscopy and Imaging Center and thank Valerie Lynch-Holm, Christine Davitt, and Chuck Cody (Washington State University) for technical assistance. We thank four anonymous reviewers for helpful comments. This work was supported by National Science

Foundation Integrated Organismal Systems Grants 0818182 and 1022106.

2.8 Author Contributions

D.R.F., D.L.M., and M.K. designed and conducted confocal and electron microscopy experiments. D.R.F., T.J.R.-E., and H.C.P. performed cloning and transformation. J.A.A. and

G.A.T. analyzed the T-DNA insertion mutant GABI-KAT 609F04 and complementation plants. K.H.J. analyzed results and calculated pressures based on microscopy data. M.K. wrote the article with participation of all the authors.

Received October 26, 2011; revised November 21, 2011; accepted December 7, 2011; published December 23, 2011.

2.9 References

Bánó , M., and Marek, J. (2006). How thick is the layer of thermal volume surrounding the protein? Biophys. Chem. 120: 44–54.

Becker, D., Kemper, E., Schell, J., and Masterson, R. (1992). New plant binary vectors with selectable markers located proximal to the left T-DNA border. Plant Mol. Biol. 20:

1195–1197.

Beneteau, J., Renard, D., Marché, L., Douville, E., Lavenant, L., Rahbé , Y., Dupont, D.,

Vilaine, F., and Dinant, S. (2010). Binding properties of the N-acetylglucosamine and high- mannose N-glycan PP2-A1 phloem lectin in Arabidopsis. Plant Physiol. 153: 1345–1361.

Bostwick, D.E., Dannenhoffer, J.M., Skaggs, M.I., Lister, R.M., Larkins, B.A., and

Thompson, G.A. (1992). Pumpkin phloem lectin genes are specifically expressed in companion cells. Plant Cell 4: 1539–1548.

Bozzola, J.J., and Russell, L.D. (1999). Electron Microscopy: Principles and Techniques for

Biologists. (Sudbury, MA: Jones & Bartlett Learning).

Bruus, H. (2008). Theoretical Microfluidics. (Cambridge, UK: Cambridge University Press).

Clark, A.M., Jacobsen, K.R., Bostwick, D.E., Dannenhoffer, J.M., Skaggs, M.I., and

Thompson, G.A. (1997). Molecular characterization of a phloem-specific gene encoding the filament protein, phloem protein 1 (PP1), from Cucurbita maxima. Plant J. 12: 49–61.

Clough, S.J., and Bent, A.F. (1998). Floral dip: A simplified method for Agrobacterium- mediated transformation of Arabidopsis thaliana. Plant J. 16: 735–743.

Cronshaw, J. (1975). P-proteins. In Phloem Transport, J.D.S. Aronoff, P.R. Gorman, L.M.

Srivastava, and C.M. Swanson, eds (New York, London: Plenum), pp. 79–115.

Cronshaw, J., and Esau, K. (1967). Tubular and fibrillar components of mature and differentiating sieve elements. J. Cell Biol. 34: 801–815.

Cronshaw, J., Gilder, J., and Stone, D. (1973). Fine structural studies of P-proteins in

Cucurbita, Cucumis, and Nicotiana. J. Ultrastruct. Res. 45: 192–205.

Deeken, R., Geiger, D., Fromm, J., Koroleva, O., Ache, P., Langenfeld-Heyser, R., Sauer,

N., May, S.T., and Hedrich, R. (2002). Loss of the AKT2/3 potassium channel affects sugar loading into the phloem of Arabidopsis. Planta 216: 334–344.

Ehlers, K., Knoblauch, M., and van Bel, A.J.E. (2000). Ultrastructural features of well- preserved and injured sieve elements: Minute clamps keep the phloem transport conduits free for mass flow. Protoplasma 214: 80–92.

Evert, R.F. (1982). Sieve-tube structure in relation to function. Bioscience 32: 789–795.

Evert, R.F. (1990). Dicotyledons. In Sieve Elements, H.D.B.R.D. Sjolund, ed (Berlin:

Springer), pp. 103–137.

Fellows, R.J., and Geiger, D.R. (1974). Structural and physiological changes in sugar beet leaves during sink to source conversion. Plant Physiol. 54: 877–885.

Fensom, D.S. (1957). The bioelectric potentials of plants and their functional significance.

Can. J. Bot. 35: 573–582.

Fisher, D.B. (1975). Structure of functional soybean sieve elements. Plant Physiol. 56: 555–

569.

Furch, A.C.U., Hafke, J.B., Schulz, A., and van Bel, A.J.E. (2007). Ca2+-mediated remote control of reversible sieve tube occlusion in Vicia faba. J. Exp. Bot. 58: 2827–2838.

Gilder, J., and Cronshaw, J. (1973). Adenosine triphosphatase in the phloem of Cucurbita.

Planta 110: 189–204.

Golecki, B., Schulz, A., and Thompson, G.A. (1999). Translocation of structural P proteins in the phloem. Plant Cell 11: 127–140.

Hartig, T. (1854). Über die Querscheidewände zwischen den einzelnen Gliedern der

Siebröhren in Cucurbita pepo. Bot. Z. 12: 51–54.

Haseloff, J., Siemering, K.R., Prasher, D.C., and Hodge, S. (1997). Removal of a cryptic intron and subcellular localization of green fluorescent protein are required to mark transgenic Arabidopsis plants brightly. Proc. Natl. Acad. Sci. USA 94: 2122–2127.

Huang, S., et al. (2009). The genome of the cucumber, Cucumis sativus L. Nat. Genet. 41:

1275–1281.

Hunt, E., Gattolin, S., Newbury, H.J., Bale, J.S., Tseng, H.-M., Barrett, D.A., and Pritchard,

J. (2009). A mutation in amino acid permease AAP6 reduces the amino acid content of the

Arabidopsis sieve elements but leaves aphid herbivores unaffected. J. Exp. Bot. 61: 55–64.

Jensen, K.H., Lee, J., Bohr, T., Bruus, H., Holbrook, N.M., and Zwieniecki, M.A. (2011).

Optimality of the Münch mechanism for translocation of sugars in plants. J. R. Soc. Interface

8: 1155–1165.

Johnson, R.P.C. (1969). Crystalline fibrils and complexes of membranes in the parietal layer in sieve elements. Planta 84: 68–80.

Knoblauch, M., Noll, G.A., Müller, T., Prüfer, D., Schneider-Hüther, I., Scharner, D., van

Bel, A.J.E., and Peters, W.S. (2003). ATP-independent contractile proteins from plants.

Nat. Mater. 2: 600–603.

Knoblauch, M., and Peters, W.S. (2010). Münch, morphology, microfluidics- Our structural problem with the phloem. Plant Cell Environ. 33: 1439–1452.

Knoblauch, M., Peters, W.S., Ehlers, K., and van Bel, A.J.E. (2001). Reversible calcium- regulated stopcocks in legume sieve tubes. Plant Cell 13: 1221–1230.

Knoblauch, M., and van Bel, A.J.E. (1998). Sieve tubes in action. Plant Cell 10: 35–50.

Lawton, D.M., and Newman, Y.M. (1979). Ultrastructure of phloem in young runner-bean stem: Discovery, in old sieve elements on the brink of collapse, of parietal bundles of P protein tubules linked to the plasmalemma. New Phytol. 82: 213–222.

Mullendore, D.L., Windt, C.W., Van As, H., and Knoblauch, M. (2010). Sieve tube geometry in relation to phloem flow. Plant Cell 22: 579–593.

Münch, E. (1927). Versuche über den Saftkreislauf. Ber. Deut. Bot. Ges. 45: 340–356.

Münch, E. (1930). Die Stoffbewegungen in der Pflanze. (Jena, Germany: Fischer).

Pélissier, H.C., Peters, W.S., Collier, R., van Bel, A.J.E., and Knoblauch, M. (2008). GFP tagging of sieve element occlusion (SEO) proteins results in green fluorescent forisomes.

Plant Cell Physiol. 49: 1699–1710.

Peters, W.S., Knoblauch, M., Warmann, S.A., Pickard, W.F., and Shen, A.Q. (2008).

Anisotropic contraction in forisomes: Simple models won’t fit. Cell Motil. Cytoskeleton 65:

368–378.

Peters, W.S., van Bel, A.J.E., and Knoblauch, M. (2006). The geometry of theforisome- sieve element-sieve plate complex in the phloem of Vicia faba L. leaflets. J. Exp. Bot. 57:

3091–3098.

Pickard, W.F., and Minchin, P.E. (1992). The nature of the short-term inhibition of stem translocation produced by abrupt stimuli. Funct. Plant Biol. 19: 471–480.

Pritchard, J. (1996). Aphid stylectomy reveals an osmotic step between sieve tube and cortical cells in barley roots. J. Exp. Bot. 47: 1519–1524.

Reynolds, E.S. (1963). The use of lead citrate at high pH as an electronopaque stain in electron microscopy. J. Cell Biol. 17: 208–212.

Rüping, B., Ernst, A.M., Jekat, S.B., Nordzieke, S., Reineke, A.R., Müller, B., Bornberg-

Bauer, E., Prüfer, D., and Noll, G.A. (2010). Molecular and phylogenetic characterization of the sieve element occlusion gene family in Fabaceae and non-Fabaceae plants. BMC Plant

Biol. 10: 219.

Russin, W.A., and Evert, R.F. (1985). Studies on the leaf of Populus deltoides (Salicaceae):

Ultrastructure, plasmodesmatal frequency, and solute concentrations. Am. J. Bot. 72: 1232–

1247.

Sabnis, D.D., and Sabnis, H.M. (1995). Phloem proteins: Structure, biochemistry and function. In The Cambial Derivatives (Encyclopedia of Plant Anatomy, Vol. 9), M. Iqbal, ed

(Berlin: Borntraeger), pp. 271–292.

Siddiqui, A.W., and Spanner, D.C. (1970). The state of the pores in functioning sieve plates. Planta 91: 181–189.

Siebert, P.D., Chenchik, A., Kellogg, D.E., Lukyanov, K.A., and Lukyanov, S.A. (1995). An improved PCR method for walking in uncloned genomic DNA. Nucleic Acids Res. 23: 1087–

1088.

Sjolund, R.D., and Shih, C.Y. (1983). Freeze-fracture analysis of phloem structure in plant tissue cultures. II. The sieve element plasma membrane. J. Ultrastruct. Res. 82: 189–197.

Spanner, D.C. (1958). The translocation of sugar in sieve tubes. J. Exp. Bot. 9: 332–342.

Spanner, D.C. (1970). The electro-osmotic theory of phloem transport in the light of recent measurements on Heracleum phloem. J. Exp. Bot. 21: 325–334.

Spanner, D.C. (1978). Sieve-plate pores, open or occluded. A critical review. Plant Cell

Environ. 1: 7–20.

Thompson, C.J., Movva, N.R., Tizard, R., Crameri, R., Davies, J.E., Lauwereys, M., and

Botterman, J. (1987). Characterization of the herbicide-resistance gene bar from

Streptomyces hygroscopicus. EMBO J. 6: 2519–2523.

Thompson, M.V. (2006). Phloem: the long and the short of it. Trends Plant Sci. 11: 26–32.

Thompson, M.V., and Holbrook, N.M. (2003). Application of a single solute non-steady- state phloem model to the study of long-distance assimilate transport. J. Theor. Biol. 220:

419-455.

Thorpe, M.R., Furch, A.C.U., Minchin, P.E.H., Föller, J., Van Bel, A.J.E., and Hafke, J.B.

(2010). Rapid cooling triggers forisome dispersion just before phloem transport stops.

Plant Cell Environ. 33: 259–271.

Toth, K.F., and Sjolund, R.D. (1994). Monoclonal antibodies against plant P-protein from plant tissue cultures. II. Taxonomic distribution of cross reactivity. Am. J. Bot. 81: 1378–

1383.

Toth, K.F., Wang, Q., and Sjolund, R.D. (1994). Monoclonal antibodies against plant

Pprotein from plant tissue cultures. I. Microscopy and biochemical analysis. Am. J. Bot. 81:

1370–1377.

Turgeon, R. (2010). The puzzle of phloem pressure. Plant Physiol. 154: 578–581.

Turgeon, R., Webb, J.A., and Evert, R.F. (1975). Ultrastructure of minor veins in Cucurbita pepo leaves. Protoplasma 83: 217–232.

Wooding, F.B.P. (1969). P protein and microtubular systems in Nicotiana callus phloem.

Planta 85: 284–298.

Wright, K.M., and Oparka, K.J. (1997). Metabolic inhibitors induce symplastic movement of solutes from the transport phloem of Arabidopsis roots. J. Exp. Bot. 48: 1807–1814.

2.10 Appendix A.

In this appendix, we derive expressions for the three resistances given in Equation

(2). With measured values of the phloem flow speed U, this allows us to determine the hydrostatic pressure difference ∆p required to drive the flow given in Eq. (1). Characteristic values of the parameters used in the calculations can be found in Table 1 while the calculated values of the hydrostatic pressure are given in Table 2.

Our starting point is the relation between the hydrostatic pressure drop ∆p between source and sink and the volumetric flow rate Q given in Eq. (1):

∆p = RQ (A1)

Here, the volume volumetric flux Q = UA is the product of the flow velocity U and cross section area A and R is the hydraulic resistance of the phloem translocation pathway.

Assuming that the translocation pathway consists of N identical sieve tube elements, M of which contain an At SEOR1 agglomeration, we write the resistance as

R = NRlumen + (N-1)Rplate +MRplug. (A2)

Here, we take into account three major components: a) the sieve tube lumen including organelles, , b) the sieve plate, , and c) the At SEOR1 agglomerations, . An expression for each of the terms in Equation (A2) is derived in the following sections, and numerical values are given in Table 2.

2.10.1 Resistance of the sieve tube lumen

Assuming that the cell lumen is well approximated by a cylindrical tube, we have for the resistance of the lumen (Bruus, 2008)

8nLt Rlumen  4 (A3) ae

Here, η is the viscosity, Lt is the length of the sieve tube element and ae is the radius of the part of the tube which is open to flow. Due to the abundance of sieve tube constituents at the margins, we estimate that the effective radius ae is between 80% and 100% of the total sieve tube element radius at .

2.10.2 Resistance of the sieve plate

For the resistance of the sieve plate we follow (Mullendore et al., 2010) and take into account the contribution to the resistance from each individual pore. The resistance of a sieve plate of thickness l consisting of Np pores of (generally different) radii ap,n has two contributions. One due to the finite length of the pore and one due to the flow near the orifice [Weissberg 1962, Dagan 1982]. We thus have for the plate resistance Rplate that

(∑ ( ) ) (A4)

where we have assumed that the sieve plates are unobstructed. Individual pore radii ap,n and average plate thickness from 22 sieve plates was determined as described in

(Mullendore et al., 2010). The plate resistance Rplate was subsequently calculated from Eq.

(A4). The value given in Table 2 is the average of the values obtained from 22 sieve plates.

Average plate thickness, pore diameter, and number of pores are given in Table 1.

2.10.3 Resistance of the At SEOR 1 agglomeration

As shown in Figures 5J and 6, the At SEOR 1 agglomerations usually has a roughly circular opening of diameter do ~ 1.0 μm. The fibrous part of the agglomeration can thus be thought of as acting in series with a cylindrical tube, such that the total resistance of the agglomeration is given by

( ) (A5)

2.10.4 Resistance of the At SEOR 1 agglomeration opening

The hydraulic resistance of the opening is completely analogous to that of a single sieve pore, Eq. (A4) (Weissberg, 1962; Dagan, 1982)

(A6)

where is the radius of the openingn and Lp is the length of the agglomeration.

2.10.5 Resistance of the At SEOR 1 agglomeration fiber network

To calculated the resistance of the At SEOR 1 fiber agglomeration Rfibers we think of the fibers as a porous medium consisting of a large number of parallel solid cylindrical rods of uniform diameter df. Analogous to Eq. (A1), we write the hydraulic resistance of the fiber network as

(A7)

where ∆pfibers is the pressure drop across the agglomeration and Qfibers is the volume flux through the fibers. To determine Rfibers we follow Jackson and James [Jackson 1986] and consider Darcy's law for the volumetric flow rate Qfiber

(A8)

where is the cross section area of the fiborous part of the agglomeration, Lp is the length of the agglomeration, and K is the permeability of the

agglomeration. The non-dimensional permeability depends on the volume fraction

of solid material and on the arrangement of the fibers. It has been determined experimentally and theoretically for several different classes of cylinder arrangements

(Jackson, 1986). For flow parallel to an array of parallel cylindrical rods, the non- dimensional permeability K is given by

( ) (A9)

95 where depends on the arrangement of the cylinders (Jackson, 1986). A comparison with experiments suggest that gives the best fit to a large collection of data, including flow through polymer gels, glass fibers and collagen, materials with dimensions similar to that of At SEOR1 (Jackson, 1986).

The arrangement of cylinders is not know in detail. We therefore approximate the solid volume fraction by the mean value obtained in three simple geometries

(A10)

{

such that and where is the distance between adjacent fiber centers

(Tamayol, 2011). From Equations (A7) and (A8) we finally have for the agglomeration resistance

. (A11)

2.10.6 Supplemental References

Dagan, Z., Weinbaum, S., and Pfeffer, R. (1982). An infinite-series solution for the creeping motion through an orifice of finite length. J. Fluid Mech. 115: 505–523. Jackson, G.W., and James, D.F. (1986). The permeability of fibrous porous media. Can. J. Chem. Eng. 64: 364–374.

Tamayol, A., and Bahrami, M. (2011). Transverse permeability of fibrous porous media. Phys. Rev. E Stat. Nonlin. Soft Matter Phys. 83: 046314. Weissberg, H.L. (1962). End correction for slow viscous flow through long tubes. Phys. Fluids 5: 1033.

2.10.7 Supplemental Movie 1. SEOR1 in Root Tip

Movement of AtSEORI-eYFP protein bodies (green) is apparent in the root tip and vascular tissue in the elongation zone of a growing root. The root is stained with

Synaptored (EMD Chem., San Diego) to highlight the cell membranes. Images were taken at

10 s interval. Total running time is 16:40 minutes.

2.10.8 Supplemental Movie 2. Real Time Imaging of Phloem Flow

After loading the phloem of a plant with carboxyfluorescein diacetate, the entire sieve tube system is fluorescent (not shown). Photo-bleaching a sieve tube by high laser power produces a distinct front of florescent label, and yellow Carboxyfluoresceindiacetate refills a sieve element towards the root tip. The first 10 frames are the final seconds of photobleaching before reducing the laser intensity to observe refilling. Images were taken at 0.355 s interval.

2.10.9 Supplemental Movie 3. SEOR1 movement in Injured Sieve Tubes

A sieve tube of a transgenic Arabidopsis line carrying SEOR1-eYFP fusion proteins.

The tube is initially slightly out of focus. The location of the sieve plate is indicated by an

97 arrow. After cutting the root tip, large yellow AtSEORI-eYFP protein agglomerates pass through a sieve plate towards the severed edge of the injured root. New agglomerates appear in the field of view from upstream sieve elements. Images were taken at 10 s interval.

Chapter 3 - Arabidopsis P-protein Filament Formation Requires Both

AtSEOR1 and AtSEOR2

James A. Ansteada, Daniel R. Froelichb, Michael Knoblauchb and Gary A. Thompsona a) College of Agricultural Sciences, The Pennsylvania State University, University Park

PA16802, USA b) School of Biological Sciences, Washington State University, Pullman WA 99164-4236,

USA

Published: Plant and Cell Physiology, 2012.

3.0 Author contributions

This publication investigated a second Aradibopsis SEOR phloem protein. It complemented Froelich 2011 by demonstrating the interaction between the two proteins in both its native plant, Arabidopsis, and in a yeast two-hybrid system.

Anstead was responsible for the majority of the publication. Figure 2 was supplied by Froelich, which imaged six different GFP-fusion constructs of the two SEOR proteins.

Additions (wildtype plus each gene with GFP), reliefs (single knockouts with the gene replaced with GFP) and complements (single knockouts with the other gene tagged with

GFP) revealed that both genes are necessary for proper filament formation. All authors assisted in editing for publication.

3.1 Abstract

The structure-function relationship of proteinaceous filaments in sieve elements has long been a source of inquiry in order to understand their role in the biology of the phloem.

Two phloem filament proteins AtSEOR1 (At3g01680.1) and AtSEOR2 (At3g01670.1) in

Arabidopsis have been identified that are required for filament formation.

Immunolocalization experiments using a phloem filament-specific monoclonal antibody in respective T-DNA insertion mutants provided an initial indication that both proteins are necessary to form phloem filaments. To further investigate the relationship between these two proteins, green fluorescent protein (GFP)-AtSEO fusion proteins were expressed in

Columbia wild-type and T-DNA insertion mutants. Analysis of these mutants by confocal microscopy confirmed that phloem filaments could only be detected in the presence of both proteins, indicating that despite significant sequence homology the proteins are not functionally redundant. Individual phloem filament protein subunits of AtSEOR1 and

AtSEOR2 were capable of forming homodimers, but not heterodimers in a yeast 2-hybrid system. The absence of phloem filaments in phloem sieve elements did not result in gross alterations of plant phenotype or affect basal resistance to green peach aphid (Myzus persicae).

Keywords: Arabidopsis, AtSEOR, Myzus persicae, phloem, P-protein, sieve element

100

3.2 Introduction

Historically, P-protein (phloem protein) is an all-inclusive term used to describe a group of ultrastructurally distinct components of sieve elements in the phloem of angiosperms (Cronshaw 1981; Esau and Cronshaw 1967). P-proteins accumulate as non- membrane bounded aggregates (P-protein bodies) in differentiating nucleate sieve elements (SEs) that either disperse forming filamentous or tubular structures (Esau and

Cronshaw 1967; Kollmann et al. 1970) or remain as non-dispersive bodies in mature SEs

(Johnson 1969). Ultrastructural studies of well preserved SEs indicate that the filamentous

P-proteins, plastids, mitochondria and smooth endoplasmic reticulum (ER) are associated together and firmly attached to the plasma membrane through clamp-like structures, resulting in a low resistance lumen free of occlusions (Ehlers et al., 2000). The presence of large accumulations of P-protein at the sieve plates of damaged SEs has led to the generalized belief that these structural proteins primarily function in sieve element occlusion (Eschrich 1970; Sjolund et al. 1983; Will and van Bel 2006) and secondarily, as a physical barrier to phloem feeding insects or microbes (Read and Northcote 1983; Will and van Bel 2006).

The dispersive P-proteins of cucurbits have been widely studied because of the ease of acquiring sieve-tube exudates. Two abundant exudate proteins, the phloem filament protein or phloem protein 1 (PP1) and the phloem lectin or phloem protein 2 (PP2), undergo reversible, oxidative cross-linkage forming high molecular weight polymers in dilute sieve-tube exudate samples (Read and Northcote 1983). PP1 monomeric subunits have a predicted molecular mass of 95.4 kDa; however, the apparent molecular size is

101 dependent on pH and oxidation state as conformational isoforms exist that appear to be related to either the polymerized or un-polymerized, translocated forms of the protein

(Clark et al. 1997; Leineweber et al. 2000). Cucurbits have an unusual phloem anatomy that consists of two ontogenetically distinct phloem systems; the fascicular or phloem of the vascular bundle, which might be considered homologous to the phloem in other plant families and the extra-fascicular phloem located at the periphery of the vascular bundles and scattered throughout the stem. Recent work has highlighted the functional differences of these two phloem systems (Zhang et al. 2010) and the soluble, translocated PP1 studied primarily in phloem exudates could be derived from the extra-fascicular phloem (Petersen et al. 2005). PP2 does not appear to be an essential component since purified PP1 subunits can form filaments/aggregations in its absence (Kleinig et al. 1975). PP1 is, however, not phylogenetically related to more recently characterized Sieve Element Occlusion proteins

(SEOs) (Pelissier et al. 2008), leading to the speculation that PP1-type filaments are an unique structural adaptation of the unusually large SEs of cucurbits (Lin et al. 2009).

Forisomes are another unique structural adaptation that are limited to sieve elements in members of the Fabaceae (Knoblauch et al. 2003) although not present in all tribes (Peters et al. 2010). Forisomes, classified as “non-dispersive,“ crystalline P-protein bodies, undergo reversible conformational changes from crystalline to dispersed states that plug sieve elements in response to local changes in calcium concentration (Knoblauch et al. 2003; Knoblauch et al. 2001). Forisomes appear to be composed of multiple sieve element occlusion (SEO) proteins, although the level of redundancy is unclear (Pelissier et al. 2008). Phylogenetic analysis has shown that forisomes are encoded by a large gene

102 family that includes filamentous SEOs from other species including Arabidopsis (Pelissier et al. 2008; Ruping et al. 2010).

Two contiguous Arabidopsis thaliana genes, At3g01670 and At3g01680, located on chromosome 3 encode putative SEO proteins that have been assigned alternative nomenclatures by different authors; AtSEOa or AtSEOR2 are encoded by At3g01670 and

AtSEOb, AtSEO1 or AtSEOR1 are encoded by At3g01680 (Froelich et al. 2011; Pelissier et al.

2008; Ruping et al. 2010). For the benefit of clarity this paper will refer to the respective proteins as AtSEOR1 and AtSEOR2 and use the same nomenclature for the genes (AtSEOR1 and AtSEOR2) that encode these proteins. Generation of transgenic lines expressing

AtSEOR1–YFP fusions revealed a previously unseen network of protein filaments in the sieve tube lumen (Froelich et al. 2011). TEM investigations unveiled a matrix of 20 nm thick filaments that often form bundles of ten to 100 individual filaments. In some cases the bundles form agglomerations that appear to fill the lumen of the sieve tube without impeding phloem flow even when large accumulation occurred at the sieve plate (Froelich et al. 2011). Whether AtSEOR2 forms part of these networks is not known, but

AtSEOR2promoter-GFPER analysis showed a phloem specific expression pattern and a higher titer of its mRNA was found in phloem enriched tissue (Ruping et al. 2010). A third related Arabidopsis gene, (At1g67790) is reported as a likely pseudogene on the basis of its failure to amplify in RT-PCR experiments. Sieve element occlusion and sieve element occlusion related proteins (SEOs and SEORs) are found in gene families ranging in size from two (and one probable pseudogene) in Arabidopsis to 26 in soybean (Glycine max).

While recent work has highlighted the structures formed by AtSEOR1 in Arabidopsis,

103 revealing a complex network of filaments within sieve elements, the role of the other phloem filament protein has not been elucidated. This study was designed to determine whether AtSEOR2 is an integral component of the filamentous network and whether the

AtSEOR1 and AtSEOR2 genes are functionally redundant.

3.3 Results

3.3.1 AtSEOR1 and AtSEOR2 proteins accumulate in Arabidopsis

A number of methods have been used to demonstrate that the genes encoding the

AtSEOR proteins are expressed in Arabidopsis; however, direct evidence for the accumulation of native AtSEOR1 and AtSEOR2 proteins has not been given in sieve elements or in sieve element exudates (Batailler et al. 2012). Analysis of MALDI-TOF MS peptide mass fingerprints showed AtSEOR1 and AtSEOR2 were both present in a total protein extraction and an immunoreactive band from Arabidopsis floral stems (Figure S1).

Five different peptide sequences from the total protein extraction and seven from the immunoreactive band were matched to the deduced AtSEOR1 amino acid sequence and two from the total protein extraction and eight from the immunoreactive band were matched to the deduced AtSEOR2 amino acid sequence. The absence of peptide sequences matching the deduced protein sequence encoded by At1g67790 combined with the inability to detect At1g67790 mRNA previously reported by Ruping and coworkers (2010) strongly indicates that it is a pseudogene.

104

3.3.2 Immunolocalization analysis of AtSEOR mutant lines

To elucidate the role of AtSEOR proteins in phloem filament formation, two independent T-DNA insertion mutants were examined. GABI-KAT 609F04 is an At3g01680

T-DNA insertion mutant (Atseor1-1), previously characterized by Froelich et al (2011).

SALK 148614C, is an At3g01670 T-DNA insertion mutant (Atseor2-1) with a T-DNA insertion in the third exon of the gene. PCR experiments confirmed that mRNA accumulation is effectively eliminated in the respective mutant lines (Figure 1) and that the expression of the adjacent, non-mutated AtSEOR gene was unaffected. The presence or absence of phloem filaments was initially examined by immunofluorescent localization in mutant and wild-type plants using the phloem filament-specific monoclonal antibody RS21.

RS21 is an antibody identified from a monoclonal antibody library created by injecting mice with phloem-enriched fraction of Streptanthus turtuosus callus culture, and subsequently screened against Arabidopsis and found to be specific for P-proteins in this and a number of other plant species although the gene encoding the target protein was not identified (Toth and Sjolund 1994; Toth et al. 1994). Figure 1c shows fluorescence in the vasculature of Arabidopsis floral stems and fluorescently-labeled filamentous exudates at higher magnification (inset) when both AtSEOR1 and AtSEOR2 are expressed. Similar filamentous exudates were present in roots and leaf veins (not shown). When either gene is mutated (Figure 1a and 1b) fluorescence was not detected in the vasculature and no phloem filaments were observed (inset). These results indicate that both AtSEORs are required for the formation of antigenic phloem filaments; however, it is possible that the

105

RS21 antibody recognizes a unique feature of the heteropolymer comprised of both proteins, while non-antigenic phloem filaments could be formed by homopolymerization of a single protein species. High resolution electron microscopy of Atseor1-1 suggests that is unlikely as no filament proteins are visible (Froelich et al., 2011). It is also possible that

RS21 recognizes a component of the phloem filament complex that is degraded or highly soluble in the absence of either AtSEOR1 or AtSEOR2 and is therefore not detected.

Figure 1: Phloem filaments antigenic to RS21 are not visible in the Atseor2-1 (A) and Atseor1-1 (B) T-DNA insertion mutants. They are, however, clearly labeled in the wild-type line Col-0 (C). Large pictures show 50 µM transverse sections of the inflorescence stem; insets are higher magnification images of single vascular bundles from 50 µM sections of the root transition zone. RT–PCR of total RNA isolated from whole seedling tissues from each line shows which gene products are present; actin was used as a positive control.

3.3.3 Formation of the phloem filament matrix requires both SEOR proteins

To establish the role of each AtSEOR protein in the formation of the phloem filament matrix, living root sieve tubes expressing GFP tagged AtSEOR proteins were visualized by

106 confocal microscopy. Micro-ROC chambers were used to allow structures in live

Arabidopsis root SEs to be visualized. All fusion proteins were created with amino-terminal

GFP tags and the recombinant genes were expressed in transgenic Arabidopsis plants under the control of their respective promoters. Transgenic plants expressing GFP-tagged proteins were created in a wild-type Columbia background and in each of the T-DNA insertion mutants (i.e. in Atseor2-1, GFP-AtSEOR1 was expressed and in Atseor1-1, GFP-

AtSEOR2 was expressed). Complementation mutants were also generated where GFP- tagged protein was expressed in the respective mutants (i.e. GFP-AtSEOR1 was expressed in Atseor1-1, and GFP-AtSEOR2 was expressed in Atseor2-1). Either GFP-tagged AtSEOR1 or

AtSEOR2 expressed in a wild-type background labeled a complex meshwork of phloem filaments inside the sieve element with some protein accumulating at the sieve plate

(Figure 3 A & B). Both lines showed the same pattern, indicating both proteins form part of the filament matrix. These patterns matched those previously found for AtSEOR1 using a carboxy-terminal YFP tag including the presence and pattern of filament structure and the presence of accumulations (plugs) at the sieve plate (Froelich et al. 2011). These data also show that the GFP tag has no apparent effect on P-protein formation.

107

108

Figure 2: Visualization of GFP-tagged sieve element (SE) occlusion proteins in whole undamaged Arabidopsis roots. Wild-type Columbia roots with GFP-tagged AtSEOR1 (A) and wild-type Columbia roots with GFP-tagged AtSEOR2 (B) show fluorescence in the SE with some build-up at the sieve plate (arrow). Atseor1-1 complemented with GFP-tagged AtSEOR1 (C) shows fluorescence in filamentous strands with similar build-up at the sieve plate to Atseor2-1complemented with GFP-tagged AtSEOR2 (D). However, in the Atseor2- 1 line expressing GFP-tagged AtSEOR1 (E), fluorescence in present in small (^) globular bodies as well as in a diffuse and amorphous pattern in both the SE and companion cell (CC). A sieve plate (arrow) interrupts uniform fluorescence intensity. Atseor1-1 expressing GFP-tagged AtSEOR2 (F) shows florescence only in globular bodies (^). Scale bars indicate 100 µm.

In contrast, when GFP-tagged AtSEOR1 was expressed in Atseor2-1 (plant expressing AtSEOR1 and GFP-AtSEOR1) the normal meshwork of filaments was absent and the fluorescence was uniformly distributed throughout the lumen of the sieve element and companion cell. Filaments were not observed in either the sieve element or companion cell and labeled protein failed to accumulate at the sieve plate, which was clearly visible as an interruption in fluorescence (Figure 3E). In addition, small fluorescent globular bodies were visible in both the sieve element and companion cell. This phenotype was rescued and the phloem filaments restored when the Atseor2-1 was complemented with GFP-

AtSEOR2 (Figure 3C). Similarly, filaments were absent when GFP-tagged AtSEOR2 was expressed in Atseor1-1 (plant expressing AtSEOR2 and GFP-AtSEOR2). While numerous globular bodies were observed in what appear to be both the sieve element and companion cell (Figure 3F) the distribution of fluorescence throughout the sieve element lumen was less apparent. This phenotype is also rescued when the mutant was complemented with

GFP-AtSEOR1 (Figure 3D). The globular bodies in both mutant lines are <500nm in size and

109 are similar in appearance to the “amorphous bodies” observed using YFP labeled AtSEOR1 in Froelich et al (2011). There was some localized movement of fluorescent bodies in

Atseor1-1 expressing GFP tagged AtSEOR2 (Figure 3F). These experiments strongly indicated that interactions between the two AtSEOR protein subunits are necessary to form phloem filaments. Yeast 2-hybrid experiments were conducted to determine whether the phloem filament subunits can directly interact. The respective coding sequences for

AtSEOR1 or AtSEOR2 were both inserted in BD- or AD-plasmids and used to transform

MATa PJ69-4A (yRM1757) and MATa PJ69-4a (yRM1756) reporter strains. The following matings were conducted: AtSEOR1 x AtSEOR1; AtSEOR2 x AtSEOR2; AtSEOR1 x AtSEOR2;

AtSEOR2 x AtSEOR1. Growth of the diploid colonies on the -leucine/-uracil media (Figure

4A) demonstrated successful mating between all of the constructs made in the BD and AD plasmids. Colony growth on both the –histidine (Figure 4B) and –adenine (Figure 4C) media of the positive control and the lack of growth of the empty vector negative controls on the respective selective media demonstrated that the experimental system was functioning correctly. Both AtSEOR1 and AtSEOR2 showed strong protein-protein interactions as homodimers evidenced by the growth of the diploid colonies on both the – adenine and –histidine media (Figure 4B and C). In contrast, there was no evidence of protein-protein interaction between AtSEOR1 and AtSEOR2. The strength of the self- interactions could have prevented the detection of weaker interactions between heterodimers; however, it is possible that AtSEOR1 and AtSEOR2 protein interactions occur at a higher structural level or require additional assembly or component proteins.

110

Figure 3: Yeast two-hybrid experiment showing that AtSEOR1 and AtSEOR2 form homo- but not heterodimers. Diploid two-hybrid reporter strains were generated by crossing yRM1757/PJ69-4A containing KAR9-BD (+ve control), AtSEOR1, AtSEOR2 or empty BD (−ve control) with yRM1756/PJ69-4a containing BIM1-BD (+ve control), AtSEOR1, AtSEOR2 or empty AD (−ve control). Diploids were selected on SD medium without uracil or leucine (A) and tested for interaction by growth on SD medium without adenine (B) and SD medium without histidine (C) at 30°C for 3 d. 3.3.4 Aphid feeding is not enhanced by the absence of phloem filaments

Phloem filaments have been hypothesized to have a negative effect on aphid feeding by blocking aphid stylets or SEs. Aphids appear to be able to overcome this by ejecting saliva that modifies the environment of the sieve element to prevent the stylet plugging during feeding (Tjallingii 2006). If phloem filaments actually present a physiological or structural barrier to feeding, a significant increase in aphid fitness would be expected in their absence as the result of removing the fitness cost associated with overcoming their effect during feeding. There was no statistical difference between the mutant lines and the control in the length of the pre-reproductive period, lifespan or nymphs laid per day during the reproductive period (Table 1). The total reproductive fitness was higher and the reproductive period was longer in aphids feeding on the wild-type line than in either of the

111 mutant lines (Figure 5 and Table 1). The total number of nymphs produced on the wild- type plants was 24% higher than on Atseor1-1plants and 15% higher than on Atseor2-1.

This decrease in fitness could be due to a nutritional effect caused by a reduction in phloem protein, as phloem filament proteins are present at high concentrations (Malter and Wolf

2011; Zhang et al. 2010) and aphid reproductive rate is often limited by amino-acid availability (Sandstrom and Moran 1999; Sandstrom and Pettersson 1994).

Figure 4: Mean pre-reproductive period and lifetime fecundity of single Myzus persicae aphids reared on homozygous Atseor2-1,Atseor1-1 and wild-type (Col-0) plants grown at 21°C under(14 h/10 h light/dark at 40.0 µmol m−2 s−1 in a randomized block design. Different letters indicate statistically significant lifetime fecundity (Student’s t- test, P < 0.05).

Table 1: Life history traits of A. gossypii developing wild-type (Columbia) and knockout Arabidopsis lines Trait Columbia Atseor1-1 Atseor2-1 6.7 ± 6.5 ± 0.42 (P = Pre-reproductive period (d) 0.18 7.2 ± 0.34 (P = 0.32) 0.43) 18.4 ± 15.7 ± 1.5 (P = Reproductive period (d) 0.84 15.8 ± 1.3 (P = 0.05) 0.03) 25.1 ± 22.2 ± 1.0 (P = Lifespan (d) 0.77 23 ± 1.3 (P = 0.07) 0.06) 31.3 ± 25.2 ± 0.87 (P = 27.2 ± 2.3 (P = Total nymphs deposited 1.2 0.002) 0.04)

112

Daily reproduction (nymphs 1.73 ± 1.64 ± 0.13 (P = 1.75 ± 0.13 (P = d-1) 0.13 0.63) 0.92)

3.4 Discussion

Phloem filament proteins are encoded by multiple members of the sieve element occlusion (SEO) and sieve element occlusion related (SEOR) gene family. The most comprehensively studied members of this family are the SEOs that encode subunits of forisomes (Peters et al. 2010). Forisomes are crystalline protein bodies specific to SEs in many species within the Fabaceae that function to reversibly block sieve tubes after injury

(Knoblauch et al. 2003; Knoblauch et al. 2001). Forisomes respond to changes in calcium concentration independently of ATP, changing from a contracted “spindle” shape at low

Ca2+ concentration to an expanded state at high Ca2+ concentrations blocking sieve tube transport (Knoblauch et al. 2005; Peters et al. 2008; Peters et al. 2007). In Medicago truncatula forisomes are composed of at least three proteins (Pelissier et al. 2008) although there are other members of the gene family in this species (Ruping et al. 2010).

Proteins encoded by this gene family share conserved domains, including a thioredoxin fold potentially involved in calcium binding, the M1 motif with its four spatially conserved cysteines residues, and a number of conserved motifs of unknown function (Ruping et al.

2010) . In Arabidopsis, this gene family is composed of two actively transcribed genes

(AtSEOR1 and AtSEOR2) and one pseudogene; however, other plant species have much larger SEO/SEOR gene families (Ruping et al. 2010; Zhang et al. 2010). AtSEOR1 is known to be a component of a complex network of phloem filaments found within the sieve elements that includes a mesh-like matrix as well as globular agglomerations (Froelich et al. 2011).

113

Observations of GFP-tagged AtSEOR2 along with GFP-AtSEOR1 clearly showed that

AtSEOR2 is a second structural component of the complex filament matrix (Figure 3B) with an expression pattern similar to GFP-AtSEOR1 (Figure 3A). In wild-type Arabidopsis plants, both GFP-labeled AtSEOR1 and AtSEOR2 label a network of filaments with comparable intensity that are distributed throughout the sieve tube.

3.4.1 Functional redundancy

The overall homology of the sequence and intron/exon structure of these genes as well as their similarity in expression and localization patterns could suggest that AtSEOR1 and AtSEOR2 genes are functionally redundant. However, phloem filaments were not detected in either of the T-DNA insertion mutants of the respective genes when probed with a phloem filament-specific monoclonal antibody (Figure 1). Analysis of mutant lines expressing GFP tagged AtSEORs also revealed that both proteins must be present to form phloem filaments (Figure 3E & F). In both mutants, the phloem filament phenotype was rescued in transgenic plants expressing a functional version of the mutated gene (Figure 3C

& D); thus, the genes do not appear to be functionally redundant.

Gene families encoding sieve element occlusion proteins have been identified in seven plant species, with members ranging from the two functional genes in Arabidopsis to the 26 putative SEO(R) genes identified in soybean (Huang et al. 2009; Ruping et al. 2010).

Phylogenetic analysis revealed that AtSEOR1 is also somewhat divergent from other SEO sequences and is the sole member of subgroup 6 (Ruping et al. 2010), although that branch is not strongly supported by the analysis. This divergence in sequence could be an indicator

114 of the functional divergence noted in the mutant analysis. Pelissier and co-workers (2008) examined GFP-tagged MtSEOF1, MtSEOF2, and MtSEOF3 proteins in composite Medicago truncatula plants and found that all three proteins were components of forisomes. Based on homology and expression patterns they concluded that the proteins are functionally redundant isoforms; however, MtSEO1 and MtSEO2 are members of a phylogenetic subgroup that does not include MtSEO3 (Ruping et al. 2010), raising the possibility for a more complex functional relationship among these proteins. It is unknown whether other, more divergent MtSEO genes encode forisome components, are subunits of phloem filaments, or have other unrelated functions. Very little is known about the function or redundancy of SEO proteins in other plant species.

3.4.2 SEOR1/SEOR2 Interactions

While it is clear that both AtSEOR1 and AtSEOR2 are required to establish phloem filaments, the formation of different structures in the T-DNA insertion mutants could indicate different roles for each gene in the assembly of the phloem filaments. In Atseor1-1 plants, GFP-AtSEOR2 accumulated predominantly in undispersed globular bodies (Figure

3F), whereas, GFP-AtSEOR1 expressed in the absence of AtSEOR2 primarily accumulated as diffuse, amorphous protein that filled the lumen of the sieve element with only a few globular bodies (Figure 3E). Given that previous reports indicate that globular bodies condense and transform into filaments (Froelich et al 2011), the absence of filaments and presence of large numbers of globular bodies in Atseor1-1expressing GFP tagged AtSEOR2 indicate that AtSEOR1 plays an essential role in the process of filament formation. In the

115 absence of AtSEOR1, AtSEOR2 remains in globular bodies and filaments are never formed.

The presence of diffuse GFP tagged AtSEOR1 in both sieve elements and companion cells in

Atseor2-1 raises several questions. AtSEOR derived phloem filaments have only been detected in sieve elements to date and promoter-reporter analysis has shown that MtSEO1-

3, VfSEO1 and AtSEOR2 promoters are only active in immature sieve elements (Noll et al.

2007; Noll et al. 2009; Ruping et al. 2010). Co-localization of YFP-tagged AtSEOR1 with a

SE-specific marker confirmed AtSEOR1 accumulation is also limited to the SE (Froelich et al. 2011). The simplest explanation is that in the absence of AtSEOR2, the unpolymerized, soluble form of AtSEOR1 readily traffics between the sieve element and companion cell through pore-plasmodesmata. This 112kDa fusion protein is considerably larger than GFP- fusion constructs previously shown to readily traffic through pore-plasmodesmata between phloem SEs and CCs, the largest of which was 67kDa (Stadler et al. 2005).

However, heterografting experiments clearly demonstrated that the unpolymerized, soluble 96kDa Cucurbita maxima phloem protein 1 (CmPP1) translocated from C. maxima to Cucumis sativus, accumulating in both C. sativus sieve elements and companion cells

(Golecki et al. 1999; Petersen et al. 2005). In contrast, the polymerized phloem filament proteins are too large to be trafficked through the pore-plasmodesmata.

The mutant analysis demonstrated that AtSEOR1 and AtSEOR2 interact in SEs forming nm scale structures visible with fluorescence microscopy. Yeast 2-hybrid experiments were conducted to gain further insight into the interactions between these two proteins. Both SEOR1 and SEOR2 proteins exhibit strong self-interactions (Figure 4), but did not appear to have detectable interactions between the two proteins. Given the

116 experimental evidence of their in vivo interactions, there are several possible explanations for the absence of detectable interactions in the yeast 2-hybrid experiments. The strong homodimeric interactions could be inhibiting weaker interactions between the two proteins as each of the individual components preferentially self-aggregated. Alternatively, multimers of one or both protein could be necessary for interactions to occur at a higher structural order. This would be consistent with the detection of non-filament structures in

Atseor1-1 expressing AtSEOR2 and the Atseor2-1expressing AtSEOR1.

3.4.3 Plant-insect interactions

On the basis of both the forisome and cucurbit phloem filament models it has been widely assumed that the major role of SEO(R) proteins in phloem is the formation of proteinaceous occlusions as the first line of defense to prevent the loss of both photoassimilates and turgor pressure after SE injury. Similar proteinaceous occlusions have been observed in aphid stylets following stylectomy (severing the aphid stylets while they are embedded in the phloem SE to collect phloem exudate) (Tjallingii and Esch 1993).

Aphids are believed to prevent both sieve element and stylet occlusion by secreting watery saliva containing calcium-chelating proteins into the sieve tube to scavenge free calcium ions released in response to the disruption of the sieve element plasma membrane, preventing or reversing sieve pore occlusions (Furch et al. 2010). This effect has been partially demonstrated with forisomes where the addition of concentrated aphid saliva in vitro reverses their dispersal (Will et al. 2007). Aphids also exhibit increased salivation following sieve element blockage (Will et al. 2009). Increased salivation has an energetic

117 cost to the aphid, both in terms of energy expenditure (production of salivary components and ATP expended) and in delayed feeding, that should be reflected by improved aphid performance when feeding on the phloem filament mutants. Removal of this structural feeding barrier should result in a corresponding reduction of these energetic costs and increased aphid fitness. However, no statistically significant fitness advantage was gained by aphids feeding on mutant plants lacking phloem filaments (Figure 5). This result fails to support the hypothesis that phloem filament proteins provide a significant barrier to aphid feeding by blocking either sieve pores or aphid stylets.

3.5 Conclusions

Analysis of GFP-tagged mutants showed that both Arabidopsis Sieve Element

Occlusion Related proteins (AtSEOR1 and AtSEOR2) are important scaffold proteins that are required to form the phloem filament matrix. Analysis of GFP-tagged SEOR proteins expressed in T-DNA insertion mutant lines showed that both proteins are required to form the characteristic phloem filament matrix in sieve elements. AtSEOR1 and AtSEOR2 T-DNA insertion mutants have different SEOR expression phenotypes that can be rescued when complemented with the appropriate GFP-tagged protein. These data show that despite their sequence homology these proteins do not have redundant functions. The differences in the protein accumulation patterns in the mutant plants suggest they have different roles in the formation of phloem filament proteins. Both genes readily form homodimers in yeast

2-hybrid experiments, but no evidence of heterodimerization was found. M. persicae

118 feeding experiments indicate that the presence of phloem filaments does not impose a fitness cost during aphid feeding.

3.6 Materials and Methods

All oligonucleotide primers were designed using Vector NTI Advance 11

(Invitrogen) and synthesized by Integrated DNA technologies (Coralville, Iowa) and are listed in Table S1.

3.6.1 AtSEOR Protein Expression in Arabidopsis

The RS21 monoclonal antibody (mab) recognizes phloem filament proteins in A. thaliana (Toth and Sjolund 1994; Toth et al. 1994). The RS21 mab was produced by hybridomas grown in a bioreactor at the Iowa State University Hybridoma Facility and the mab was concentrated by ammonium sulfate precipitation. Two grams of floral stem tissue was frozen in liquid nitrogen and homogenized in 2ml of purification buffer (10mM Tris,

10mM EGTA, 150mM NaCl, 10mM KCl, 1% Sigma protease inhibitor cocktail, 20mM DTT).

The tissue was incubated for one hour at 4°C with rocking, then centrifuged and the supernatant removed. The pellet was then washed once with 10ml of the purification buffer and centrifuged. The supernatant was discarded and 2ml of SDS extraction buffer was added (4% SDS, 125mM Tris-HCl, 20mM DTT, 1% Sigma protease inhibitor cocktail) to the pellet and incubated at RT for one hour with rocking. The protein extraction was centrifuged and the supernatant decanted and half saved for analysis. The other half was

119 boiled and separated in duplicate 8-16% gradient SDS-polyacrylamide gels. Duplicate gels were either stained with Coomassie blue or the proteins transferred to nitrocellulose membrane using the semi-dry method, blocked with 2.5% dry milk in TBST and incubated with purified RS21 overnight at 4°C. A gel slice was cut from the stained gel at the site of the immunoreactive band and proteins were identified from the total protein extraction and the immunoreactive band from MALDI-TOF MS peptide mass fingerprints that were obtained by the Oklahoma State Recombinant DNA/Protein Core Facility.

3.6.2 Arabidopsis T-DNA insertion mutants

T-DNA Express (http://signal.salk.edu/cgi-bin/tdnaexpress) was used to identify T-

DNA insertions in the genes At3g01670 (AtSEOR2) and At3g01680 (AtSEOR1). Seeds for the mutant lines SALK 148614C (AtSEOR2 knock-out, Atseor2-1), obtained from the

Arabidopsis Biological Resource Center (Columbus, OH). Seeds for GABI-KAT 609F04

(AtSEOR1 knock-out, Atseor1-1), were obtained from the Genomanalyse im Biologischen

System Pflanze (Bielefeld, Germany) (both mutants are in a Columbia background).

Homozygous plants were identified using PCR-based screening according to the method of

Siebert et al. (1995). The GABI-KAT 609F04 mutant contained an additional T-DNA insertion in a different gene so plants were allowed to self-fertilize and plants homozygous for the At3g01680 insertion alone were further analyzed. Successful knockout of each gene was also confirmed using RT-PCR. In brief total RNA was extracted using the Trizol method and total RNA was reverse transcribed using SuperScript II according to the manufacturer’s

120 instructions. Partial, intron spanning sections of each gene were amplified using gene specific primers (Table S1) and visualized on an Ethidium bromide stained agarose gel.

3.6.3 Immunolocalization of phloem filaments in AtSEOR knockouts

Living tissue sections of A. thaliana Col-0 plants, Atseor1-1, and Atseor2-1floral stems were cut with a vibrating microtome (Vibratome, Bannockburn, IL 60015) and collected in phosphate buffered saline (PBS). The 50 μm cross sections were washed twice in 10 mM

PBS and incubated for 30 minutes in PBS with 3% non-fat dry milk (blocking buffer).

Sections were then washed twice more with PBS and incubated for 45 minutes with the

RS21 primary monoclonal antibody in blocking buffer (1:100). After incubation with primary antibody the sections were washed three times with PBS and then incubated in

PBS with ALEXA 488nm fluorescently tagged secondary goat anti-mouse antibody

(Invitrogen, Carlsbad, CA) (1:250). Finally, the labeled sections were washed twice with

PBS and once with nanopure water and observed under a Nikon E600 epifluorescence microscope with an excitation wavelength of 490 nm and an emission wavelength of 512 nm.

3.6.4 Transgenic plants expressing recombinant protein fusions

The eGFP gene was PCR amplified using primers designed to subclone the gene into the pGPTV-Kan binary vectors generated previously in place of the uidA gene using the

SmaI and KpnI restriction sites, these primers also created a multiple cloning site at the 3’

121 end of the eGFP gene. Subsequently the AtSEOR1 and AtSEOR2 ORFs were PCR amplified using specific primers designed to subclone the ORF’s into the multiple cloning site (KpnI and ApaI). The binary vectors were transformed into the A. tumefaciens strain GV3101 and used to transform Arabidopsis lines Col-0, Atseor1-1 and Atseor2-1by the floral dip technique (Clough and Bent 1998). Transgenic plants were selected on kanamycin supplemented media, transplanted and then grown in a Percival growth chamber (14:10

L:D 21°C). The presence of the correct GFP fusion construct was confirmed using primers that overlapped the eGFP-ORF boundary. The presence of the T-DNA insertion and its location were also re-confirmed by PCR. In total six transgenic genotypes were created.

Two wild type (Col-o) lines expressing GFP tagged AtSEOR1 and GFP tagged AtSEOR2 respectively. Two Atseor1-1 lines expressing GFP tagged AtSEOR1 and GFP tagged AtSEOR2 respectively and two Atseor2-1 lines expressing GFP tagged AtSEOR1 and GFP tagged

AtSEOR2 respectively. For analysis plants were grown in Microscopy Rhizosphere

Chambers (micro-ROCs; Advanced Science Tools, Pullman, WA) in the greenhouse (14:10

L:D, 20°C:15°C) to the six-eight leaf stage. Micro-ROCs allow live root cells to be visualized without preparation in a natural soil environment. Confocal laser scanning microscopy images were obtained with a Leica TCS SP5 with 488nm argon laser excitation and 500-

555nm emission. Image processing was performed with Leica LAS AF lite software.

3.6.5 Yeast 2-hybrid analysis of AtSEOR1 and AtSEOR2 interactions

The mating strains used for the yeast 2-hybrid experiment were MATa PJ69-4A

(yRM1757) and MATa PJ69-4a (yRM1756) reporter strains (generously provided by Dr.

122

Rita Miller, Oklahoma State University, Stillwater, OK). To serve as a positive control for the analysis, pRM1154-BD plasmid containing Kar9 protein sequence and pRM1151-AD plasmid containing Bim1 protein sequence were transformed into Saccharomyces cerevisiae yRM1757 and yRM1756 strains, respectively (Meednu et al. 2008). As negative controls, strains containing empty AD plasmid (E-AD) were mated with strains containing empty BD plasmid (E-BD). The ORFs of AtSEOR1 and AtSEOR2 were each cloned into pRM1151-AD using specific primers designed to subclone the ORF’s into the multiple cloning sites BamHI/XhoI and EcoRI/XhoI respectively and transformed into yRM1757 strains. Likewise both ORFs were cloned into pRM1154-BD using the same restriction sites and transformed into yRM1756 strains. Haploid yeast cells were mated by crossing the two strains containing corresponding plasmids to generate multiple combinations that were replicate-plated onto SC plates lacking uracil and leucine (only diploid cells will grow) and grown for 48-72 hours at 30°C. These diploid cells were then replica plated onto –adenine and –histidine plates to determine which proteins showed protein-protein interactions.

They were then grown for a further 72 hours at 30°C.

3.6.6 Aphid fecundity study

Six replicates of homozygous T-DNA insertion mutants from the Atseor1-1 and

Atseor2-1 lines, and wild-type (col-0) plants were grown at 21 °C under 14:10 L:D at 40.0

μmol m-2 s-1 for three weeks (just beginning to bolt) in a randomized block design.

Individual adults from a clonal M. persicae colony were placed on each plant and allowed to deposit a single nymph, at which point the adult was removed. Each plant was covered with

123 a cage made from an adapted Aracon (Betatech, Gent Belgium) tube. Aphids were then allowed to feed on the plants and were examined daily, when reproductive age was reached newly deposited nymphs were removed each day and their numbers recorded.

The experiment was terminated when all adult aphids had died.

3.7 Acknowledgements

We would like to thank Rita Miller for kindly providing yeast strains and advice for the yeast 2-hybrid experiments and Steve Hartson and the Oklahoma State Recombinant

DNA/Protein Core Facility for MALDI-TOF MS peptide mass fingerprint analyses.

124

3.8 References

Batailler, B., Lemaitre, T., Vilaine, F., Sanchez, C., Renard, D., Cayla, T., Beneteau, J. and

Dinant, S. (2012) Soluble and Filamentous proteins in Arabidopsis sieve elements. Plant

Cell Environ. Accepted Article: doi: 10.1111/j.1365-3040.2012.02487.x.

Becker, D., Kemper, E., Schell, J. and Masterson, R. (1992) New plant binary vectors with selectable markers located proximal to the left T-DNA border. Plant Mol. Biol. 20: 1195-

1197.

Clark, A.M., Jacobsen, K.R., Bostwick, D.E., Dannenhoffer, J.M., Skaggs, M.I. and

Thompson, G.A. (1997) Molecular characterization of a phloem-specific gene encoding the filament protein, phloem protein 1 (PP1), from Cucurbita maxima. Plant J. 12: 49-61.

Clough, S.J. and Bent, A.F. (1998) Floral dip: a simplified method for Agrobacterium- mediated transformation of Arabidopsis thaliana. Plant J. 16: 735-743.

Cronshaw, J. (1981) Phloem Structure and Function. Annu. Rev. Plant Phys.32: 465-484.

Esau, K. and Cronshaw, J. (1967) Tubular components in cells of healthy and tobacco mosiac virus infected Nicotiana. Virology 33: 26-35.

Eschrich, W. (1970) Biochemistry and Fine Structure of Phloem in Relation to Transport.

Annu. Rev. Plant Physio. 21: 193-214.

Froelich, D.R., Mullendore, D.L., Jensen, K.H., Ross-Elliott, T.J., Anstead, J.A.,

Thompson, G., Pelissier, H.C. and Knoblauch, M. (2011) Phloem Ultrastructure and

Pressure Flow: Sieve-Element-Occlusion-Related Agglomerations Do Not Affect

Translocation. Plant Cell 23: 4428-4445.

125

Furch, A.C.U., Zimmermann, M.R., Will, T., Hafke, J.B. and van Bel, A.J.E. (2010) Remote- controlled stop of phloem mass flow by biphasic occlusion in Cucurbita maxima. J. Exp. Bot.

61: 3697-3708.

Golecki, B., Schulz, A. and Thompson, G.A. (1999) Translocation of structural P proteins in the phloem. Plant Cell 11: 127-140.

Huang, S., Li, R., Zhang, Z., Li, L., Gu, X., et al. (2009) The genome of the cucumber,

Cucumis sativus L. Nat. Genet. 41: 1275-U1229.

Johnson, R.P.C. (1969) Crystalline Fibrils and Complexes of Membranes in Parietal Layer in Sieve Elements. Planta 84: 68-&.

Kleinig, H., Thones, J., Dorr, I. and Kollmann, R. (1975) Filament formation in vitro of a sieve tube protein from Cucurbita maxima and Cucurbita pepo. Planta 127: 163-170.

Knoblauch, M., Noll, G.A., Muller, T., Prufer, D., Schneider-Huther, I., Scharner, D., Van

Bel, A.J.E. and Peters, W.S. (2005) ATP-independent contractile proteins from plants (vol

2, pg 600, 2003). Nat. Mater. 4: 353-353.

Knoblauch, M., Noll, G.A., Van Bel, A.J.E., Prufer, D. and Peters, W.S. (2003) Forisomes consist of a novel class of contractile proteins. Eur. J. Cell Biol. 82: 70-71.

Knoblauch, M., Peters, W.S., Ehlers, K. and van Bel, A.J.E. (2001) Reversible calcium- regulated stopcocks in legume sieve tubes. Plant Cell 13: 1221-1230.

Kollmann, R., Dorr, I. and Kleinig, H. (1970) Protein Filaments - Structural Components of Phloem Exudate .1. Observations with Cucurbita and Nicotiana. Planta 95: 86-94.

Leineweber, K., Schulz, A. and Thompson, G.A. (2000) Dynamic transitions in the translocated phloem filament protein. Aust. J. Plant Physiol. 27: 733-741.

126

Lin, M.-K., Lee, Y.-J., Lough, T.J., Phinney, B.S. and Lucas, W.J. (2009) Analysis of the

Pumpkin Phloem Proteome Provides Insights into Angiosperm Sieve Tube Function. Mol.

Cell. Prot. 8: 343-356.

Malter, D. and Wolf, S. (2011) Melon phloem-sap proteome: developmental control and response to viral infection. Protoplasma 248: 217-224.

Meednu, N., Hoops, H., D'Silva, S., Pogorzala, L., Wood, S., Farkas, D., Sorrentino, M.,

Sia, E., Meluh, P. and Miller, R.K. (2008) The Spindle Positioning Protein Kar9p Interacts

With the Sumoylation Machinery in Saccharomyces cerevisiae. Genetics 180: 2033-2055.

Noll, G.A., Fontanellaz, M.E., Rueping, B., Ashoub, A., van Bel, A.J.E., Fischer, R.,

Knoblauch, M. and Pruefer, D. (2007) Spatial and temporal regulation of the forisome gene for1 in the phloem during plant development. Plant Mol. Biol. 65: 285-294.

Noll, G.A., Ruping, B., Ernst, A.M., Bucsenez, M., Twyman, R.M., Fischer, R. and Prufer,

D. (2009) The Promoters of Forisome Genes MtSEO2 and MtSEO3 Direct Gene Expression to Immature Sieve Elements in Medicago truncatula and Nicotiana tabacum. Plant Mol. Biol.

Rep. 27: 526-533.

Pelissier, H.C., Peters, W.S., Collier, R., van Bel, A.J.E. and Knoblauch, M. (2008) GFP

Tagging of Sieve Element Occlusion (SEO) Proteins Results in Green Fluorescent Forisomes.

Plant Cell Physiol. 49: 1699-1710.

Peters, W.S., Haffer, D., Hanakam, C.B., van Bel, A.J.E. and Knoblauch, M. (2010)

Legume phylogeny and the evolution of a unique contractile apparatus that regulates phloem transport. Am. J. Bot. 97: 797-808.

127

Peters, W.S., Knoblauch, M., Warmann, S.A., Pickard, W.F. and Shen, A.Q. (2008)

Anisotropic contraction in forisomes: Simple models won't fit. Cell Mot. Cyto. 65: 368-378.

Peters, W.S., Knoblauch, M., Warmann, S.A., Schnetter, R., Shen, A.Q. and Pickard, W.F.

(2007) Tailed forisomes of Canavalia gladiata: A new model to study Ca2+-driven protein contractility. Ann. Bot. 100: 101-109.

Petersen, M.L., Hejgaard, J., Thompson, G.A. and Schulz, A. (2005) Cucurbit phloem serpins are graft-transmissible and appear to be resistant to turnover in the sieve element- companion cell complex. J. Exp. Bot. 56: 3111-3120.

Read, S.M. and Northcote, D.H. (1983) Subunit structure and interactions of the phloem proteins of Cucurbita maxima (pumpkin). Eur. J. Biochem. 134: 561-569.

Ruping, B., Ernst, A.M., Jekat, S.B., Nordzieke, S., Reineke, A.R., Mueller, B., Bornberg-

Bauer, E., Pruefer, D. and Noll, G.A. (2010) Molecular and phylogenetic characterization of the sieve element occlusion gene family in Fabaceae and non-Fabaceae plants. BMC Plant

Biol. 10: 219

Sandstrom, J. and Moran, N. (1999) How nutritionally imbalanced is phloem sap for aphids? Entomol. Exp. Appl. 91: 203-210.

Sandstrom, J. and Pettersson, J. (1994) Amino-Acid composition of phloem sap and the relation to intraspecific variation in pea-aphid (Acyrtosiphon pisum) performance. J. Insect

Physiol. 40: 947-955.

Siebert, P.D., Chenchik, A., Kellogg, D.E., Lukyanov, K.A. and Lukyanov, S.A. (1995) An improved PCR method for walking in uncloned genomic DNA. Nucleic Acids Res. 23: 1087-

1088.

128

Sjolund, R.D., Shih, C.Y. and Jensen, K.G. (1983) Freezer-Fracture Analysis of Phloem

Structure in Plant-Tissue Cultures .3. P-Protein, Sieve Area Pores, and Wounding. J. Ultras.

Research 82: 198-211.

Stadler, R., Wright, K.M., Lauterbach, C., Amon, G., Gahrtz, M., Feuerstein, A., Oparka,

K.J. and Sauer, N. (2005) Expression of GFP-fusions in Arabidopsis companion cells reveals non-specific protein trafficking into sieve elements and identifies a novel post- phloem domain in roots. Plant J. 41: 319-331.

Tjallingii, W.F. (2006) Salivary secretions by aphids interacting with proteins of phloem wound responses. J. Exp. Bot. 57: 739-745.

Tjallingii, W.F. and Esch, T.H. (1993) Fine-structure of aphid stylet routes in plant-tissues in correlation with EPG signals. Physiol. Entomol. 18: 317-328.

Toth, K.F. and Sjolund, R.D. (1994) Monoclonal-antibodies against phloem P-protein from plant-tissue cultures. 2. Taxanomic distribution of cross reactivity. American Journal of

Botany 81: 1378-1383.

Toth, K.F., Wang, Q. and Sjolund, R.D. (1994) Monoclonal-Antibodies Against Phloem P-

Protien From Plant -Tissue Cultures .1. Microscopy and Biochemical-Analysis. Am. J. Bot.

81: 1370-1377.

Will, T., Kornemann, S.R., Furch, A.C.U., Tjallingii, W.F. and van Bel, A.J.E. (2009) Aphid watery saliva counteracts sieve-tube occlusion: a universal phenomenon? J. Exp. Bot. 212:

3305-3312.

Will, T., Tjallingii, W.F., Thonnessen, A. and van Bel, A.J.E. (2007) Molecular sabotage of plant defense by aphid saliva. P. Natl. Acad. Sci. USA 104: 10536-10541.

129

Will, T. and van Bel, A.J.E. (2006) Physical and chemical interactions between aphids and plants. J. Exp. Bot. 57: 729-737.

Zhang, B., Tolstikov, V., Turnbull, C., Hicks, L.M. and Fiehn, O. (2010) Divergent metabolome and proteome suggest functional independence of dual phloem transport systems in cucurbits. P. Natl. Acad. Sci. USA 107: 13532-13537.

130

Chapter 4 - SEORious business – structural proteins in sieve tubes and

their involvement in sieve element occlusion

Michael Knoblaucha, Daniel R. Froelicha, William F. Pickardb and Winfried S. Petersc a) School of Biological Sciences, Washington State University, Pullman WA 99164, USA b) Department of Electrical and Systems Engineering, Washington Univ., St. Louis, Missouri

63130, USA. c) School of Biological Sciences, Washington State University, Pullman WA 99164, USA; on

Sabbatical Leave from Indiana/Purdue University Fort Wayne, 2101 East Coliseum

Boulevard, Fort Wayne, IN 46805-1499, USA,

Published: Journal of Experimental Botany, 2014.

4.0 Author contributions

This review was published in a special edition of the Journal of Experimental Botany following the Plant Vascular Biology 2013 conference. It intends to review and ultimately clarify what is currently known about the role of phloem proteins, based on actual observation and experimentation, not conclusions of conjecture.

Knoblauch and Peters contributed most of the review writing with sections contributed by Froelich and Pickard. The cold shock experiment showing Arabidopsis is capable of halting phloem flow without At-SEOR-1 was performed by Froelich, using a novel technique adapted from (Froelich, 2011). All authors assisted in editing for publication.

131

4.1 Abstract

The phloem provides a network of sieve tubes for long-distance translocation of photosynthates. For over a century, structural proteins in sieve tubes have presented a conundrum since they presumably increase the hydraulic resistance of the tubes while no potential function other than sieve tube or wound sealing in the case of injury has been suggested. Here we summarize and critically evaluate current speculations regarding the roles of these proteins. Our understanding suffers from the suggestive power of images; what looks like a sieve tube plug on micrographs may not actually impede translocation very much. Recent reports of an involvement of SEOR (sieve element occlusion-related) proteins, a class of P-proteins, in the sealing of injured sieve tubes are inconclusive; various lines of evidence suggest that, in neither intact nor injured plants, are SEORs determinative of translocation stoppage. Similarly, the popular notion that P-proteins serve in the defence against phloem sap-feeding insects is unsupported by empirical facts; it is conceivable that in functional sieve tubes, aphids actually could benefit from inducing a plug. The idea that rising cytosolic Ca2+ generally triggers sieve tube blockage by P-proteins appears widely accepted, despite lacking experimental support. Even in forisomes, P-protein assemblages restricted to one single plant family and the only Ca2+-responsive P-proteins known, the available evidence does not unequivocally suggest that plug formation is the cause rather than a consequence of translocation stoppage. We conclude that the physiological roles of structural P-proteins remain elusive, and that in vivo studies of their dynamics in continuous sieve tube networks combined with flow velocity measurements will be required to (hopefully) resolve this scientific roadblock.

132

4.2 Introduction: struggling with structural sieve tube components

The practical investigation of biochemical and molecular properties of cell components usually starts from an initial isolation and purification. The isolation of a specific structure or substance may be extremely complicated if it is present in low quantities. This is a common problem when working with phloem components. Sieve elements, companion cells, and phloem parenchyma are very different cell types, but form a functional unit of structurally and functionally interconnected elements (Esau,

1969; Evert, 1982; Knoblauch and Peters, 2010). The separation of one cell type from the others is practically impossible. In addition, the phloem forms a network, embedded in other tissues. The phloem contributes <1% of the plant body in most species, which complicates isolation and purification even more. Historically, biochemical and molecular investigations into phloem composition were mostly restricted to soluble substances that could be found in phloem sap exudates (Atkinset al., 2010).

It is no surprise that the first thorough biochemical studies of phloem components were conducted on cucurbits (Cucurbitaceae) which exude phloem sap over prolonged periods when treated properly, enabling the collection of millilitre volumes of sap (Crafts,

1932). Components such as the phloem filament protein (PP1) and phloem lectin (PP2) were isolated in large quantities, allowing thorough biochemical analysis (Lin et al., 2009).

The phloem of many cucurbits is unusual not only because of the bicollateral vascular bundles in which two sets of sieve tubes are located externally and internally of the xylem, but also because of an extrafascicular sieve tube network that is scattered throughout the non-vascular parenchyma (Fischer, 1884; Crafts, 1932). Unfortunately, most of the exudate

133 that can be easily collected in cucurbits does not seem to originate from the phloem at all

(Zhang et al., 2012). Furthermore, it is not really clear how much exudate is contributed from the extrafascicular phloem whose contents and function differ significantly from those of the vascular phloem (Zhang et al., 2010; Gaupels et al., 2012; Zhang et al., 2012).

This makes comparisons with non-cucurbit species difficult, and suggests that results obtained with cucurbits should not be rashly generalized (Turgeon and Oparka,

2010; Slewinski et al., 2013).

Exudates—although usually in lower quantities compared with cucurbits—can also be collected from other plant species, and a variety of soluble substances including proteins, RNAs, amino acids, and sugars have been detected (Marentes and Grusak,

1998; Schobert et al., 2000; Lough and Lucas, 2006). Structural components, however, are usually absent from exudates due to their size and insolubility. Until recently it was believed that these structural components comprise endoplasmic reticulum (ER), mitochondria, sieve element plastids, and phloem-specific proteins (P-proteins; for a review, see Knoblauch and Peters, 2010). However, investigations by confocal laser- scanning microscopy (CLSM) of living plants in microscopy rhizosphere chambers (Micro-

ROCs), and by electron microscopy after freeze substitution indicated that the peripheral cytoplasmic layer in sieve tubes may contain previously unknown elements (Froelich et al.,

2011).

The absence of structural components from exudates has prevented biochemical and molecular studies. The alternative isolation method, extraction from homogenates, is

134 difficult as well, since sieve tube components are attached to the plasma membrane via small protein linkers (Ehlers et al., 2000; Froelich et al., 2011). When the tissue is homogenized, these linkers lead to mixtures of different quantities of the various sieve tube components that have different densities, impeding the formation of specific bands in density gradients. The surprising exclusion of sieve element plastids from textbooks as a plastid type deriving from proplastids exemplifies the dilemma. It is comparatively easy to purify the large quantities of chloroplasts, chromoplasts, and leucoplasts that are floating in the cytoplasm of numerous cells, restricted only by transient connections to the cytoskeleton via motor proteins (Vick and Nebenführ, 2012). Isolating the small numbers of sieve element plastids that are attached rigidly to the plasma membrane is a different ball game.

The situation is less difficult for non-dispersive P-protein bodies (NDPPBs; for a review, see Behnke, 1991), which are visible in the light microscope and can be found in

~10% of the angiosperm families. At least in some cases, NDPPBs seem to move freely in the sieve tube lumen, as indicated by their preferential localization at the downstream end of the sieve element (Peters et al., 2006). Nonetheless they are absent from exudates, since their size exceeds the sieve plate pore diameter. On the other hand, their size allows them to be isolated and analysed individually. The analysis of one particular type of NDPPBs, the contractile forisomes, has not only elucidated forisome evolution (Peters et al., 2010) but also led to the molecular identification of a family of dispersive P-proteins (Pélissier et al.,

2008). Starting with NDPPBs and forisomes, and proceeding to the related dispersive P- proteins, we will critically discuss current ideas about the function of these phloem

135 components. Because we believe that there are valid alternatives to currently popular interpretations of several key experiments, we shall add some iconoclastic speculations in our final section.

4.2.1 Forisome function: seeing is believing—what about knowing?

Form and shape of NDPPBs vary and often are specific for certain taxa (Behnke,

1991). Some NDPPBs are capable of rapidly switching between a low-volume state at the low Ca2+ levels that are typical of transporting sieve elements, and a high-volume state at the increased Ca2+ levels of stressed or injured sieve tubes (Knoblauch et al.,

2001; Pickard et al., 2006; Peters et al., 2007). This peculiar, Ca2+-dependent but ATP- independent contractility of NDPPBs is known only from the papilionoid legumes (the

Fabaceae sensu stricto); in fact, it appears to be one of the synapomorphies that define this huge taxon as a monophyletic clade (Peters et al., 2010).

From principles of fluid dynamics alone, it is clear that NDPPBs must affect fluid flow in sieve tubes. Just like sieve plates and the lateral borders of the sieve elements,

NDPPBs contribute to the total hydraulic resistance in the system. The contractile NDPPBs of the papilionoids, however, are unique as their shape and size, two factors that control the hydrodynamic properties of an object, change dependent on the cytosolic Ca2+ level which can be regulated by the cell (Knoblauch et al., 2001; Pickard et al., 2006; Furch et al.,

2009). The active regulation of hydraulic resistance and the passive, merely structural contribution to total hydraulic resistance are fundamentally different phenomena. For

136 these reasons, papilionoid NDPPBs were re-named gate bodies, or forisomes (Knoblauch et al., 2003). Their postulated function, however, proved hard to demonstrate in situ.

Micrographs produced by CLSM and transmission electron microscopy (TEM) of forisomes in the high-volume state in situ were highly suggestive of a structural block (Knoblauch et al., 2001). However, if based on the visual appearance of forisome plugs alone, the conclusion that forisomes actually are blocking phloem flow will remain problematic at best, for several reasons. First, what appears like a block on a 2D picture does not necessarily block fluid flow in 3D reality, since open passages may exist outside of the 2D plane. Secondly, some materials that appear just as dense as forisome plugs on electron micrographs allow fluids to permeate at significant rates. Cell walls, for example, look quite solid, but aqueous solutions readily pass through them; otherwise common phenomena such as plasmolysis would be inexplicable, as botanists realized more than a century ago

(de Vries, 1877; Pfeffer, 1877). Apoplasmic transport (i.e. fluid flow in the cell wall space) has been monitored using non-membrane-permeant dyes (Hanson et al., 1985; Moon et al.,

1986). Unfortunately, the apoplasmic movement of dyes does not necessarily provide a quantitative measure for concurrent water fluxes since hydrophobic wall components found, for example, in Casparian strips inhibit the apoplasmic movement of water and solutes selectively (Zimmermann and Steudle, 1998). Generally, the identification of such barriers requires functional tests and cannot be achieved by simply looking at micrographs

(Schreiber et al., 1999; Hose et al., 2001;Ranathunge and Schreiber, 2011). We see no reason to assume that the hydrodynamic behaviour of forisome plugs and other protein agglomerations in sieve tubes necessarily is less complex than that of cell wall materials.

137

Thirdly, the 3D geometry of the sieve tubes containing forisomes cannot be ignored if we are to evaluate the efficiency of forisome plugs. An analysis of anatomical data available at the time indicated that forisomes were incapable of occluding sieve tubes for geometric reasons in Vicia faba (Peters et al., 2006), but the popularity of the idea that forisomes could block sieve tubes apparently remained unaffected. A causal relationship between forisome activity and phloem flow stoppage was implied by Thorpe et al. (2010) who reported that the transition of forisomes into the high-volume state correlated with the stoppage of phloem transport following rapid cooling, but, as always, correlation does not imply causation. Many plants exhibit cold shock-inducible transient stoppages of phloem translocation (Lang and Minchin, 1986), but only papilionoid legumes possess forisomes.

Therefore, the reported temporal correlation of forisome phase change and flow stoppage in a papilionoid species (Thorpe et al., 2010) does not imply a causal relationship between the two phenomena, which might well be separate effects of a common cause. On the other hand, forisomes can be isolated by pre-purifying phloem tissue before extraction and gradient centrifugation (Knoblauch et al., 2003), opening up the possibility to study their proposed function in vitro. The first published attempt to regulate fluid flow in channels on microfluidics chips using isolated forisomes failed: the movement of suspended particles, but not that of the fluid, stopped when the forisomes switched into their high-volume state

(Uhliget al., 2008). Apparently, all these problems were no match for the suggestive power of micrographs that showed occlusion of sieve elements or artificial microchannels by forisomes, and occasionally wishful thinking took over. Groscurth et al. (2012, p. 3077), for example, celebrated ‘the technological potential of forisomes, as recently demonstrated by

138 their incorporation as smart materials into a prototype microfluidic system to control fluid flow (Uhlig et al., 2008)’. Ironically, controlling fluid flow is exactly what Uhlig and colleagues had not accomplished, as mentioned above.

As it turned out, the main problem working with isolated forisomes is that their reactivity sharply deteriorates as soon as they are released from their cells. Only after isolation procedures had been optimized, and after the incubation conditions had been re- designed to mimic closely the reducing milieu in the phloem, did it become possible actually to demonstrate the occlusion of artificial sieve elements by forisomes

(Knoblauch et al., 2012). On the basis of this prima facie evidence generated by direct functional tests, it would appear most unreasonable to doubt that forisomes are capable in principle of controlling fluid flow in natural sieve tubes. However, there is to date still no direct demonstration of such flow controlin vivo. Assuming that forisomes actually do occlude sieve tubes when prompted by a rise in cytosolic Ca2+, what could be a biological context in which such a reaction would be adaptive?

The plant phloem is attacked by various specialized consumers that extract phloem sap from more or less intact sieve tubes (Dixon, 1975;Douglas, 2006; Walling, 2008).

Therefore, the possibility that forisomes might be involved in the defence against aphids and other phloem sap thieves is obvious (Knoblauch et al., 2001). Aphids secrete gelling saliva that hardens rapidly to form a stylet sheath as they penetrate the plant tissue with their stylets (Miles, 1999). They also intermittently discharge watery saliva while probing as well as during phloem sap feeding (Miles, 1999; Tjallingii, 2006; Moreno et al., 2011),

139 suggesting that watery saliva may have a dual function in target as well as non-target tissues. Watery saliva contains proteins including a variety of enzymes (Miles,

1999;Harmel et al., 2008; Carolan et al., 2009; Rao et al., 2013) and factors thought to induce or suppress plant defence responses (Hilker and Meiners, 2010; Hogenhout and

Bos, 2011; Consales et al., 2012; Coppolaet al., 2013; Elzinga and Jander, 2013). An essential role in phloem sap feeding has been demonstrated for Protein C002 from the pea aphid (Acyrthosiphon pisum; Mutti et al., 2008). Putative calcium-binding proteins have been found in the watery saliva of a leafhopper (a non-aphid phloem feeder; Hattori et al.,

2012), and in those of several aphids (Carolan et al., 2011; Nicholson et al., 2012; Rao et al.,

2013). In an in vitro assay, calcium-binding proteins from the saliva of the aphidMegoura viciae competed for Ca2+ with forisomes isolated from V. faba.This interference resulted in an inhibition of the forisomes’ transition into the Ca2+-induced high-volume state (Will et al., 2007). In this experiment, protein concentrates derived from artificial media on which aphids had fed were used; it remained unexplored how the concentrations of saliva protein in these artificial concentrates compared with those that could realistically be expected to build up in functional sieve elements if delivered into the flowing sieve tube sap by an aphid. Another problem is that according to Miles (1999, p. 49), the validity of saliva analyses based on secretions into non-natural food sources is generally questionable, because of the excretory function of the glands from which the watery saliva is derived. It should be stressed also that any pair of arbitrarily chosen calcium-binding proteins will show competition for Ca2+ in tests of this type, so that observed interferences do not necessarily indicate physiological relevance. Notably, watery saliva is secreted right from

140 the start of tissue penetration, long before a sieve tube is impaled (Moreno et al., 2011).

This opens up the possibility that the physiological target of the Ca2+-binding saliva proteins is not located in the phloem at all. Despite these caveats, Will and colleagues

(2007)definitely have identified a candidate saliva protein that might interfere with forisome function in vivo.

Will et al. (2007) also documented a sudden shift in the electrical penetration graph

(EPG) pattern of aphids feeding on Vicia leaves that occurred shortly after the leaf had been burned 5cm from the aphid, in the upstream direction of phloem flow (Will et al., 2007).

This EPG pattern shift was interpreted as a switch from phloem sap ingestion (E2 pattern) to salivation (E1 pattern) behaviour, which supposedly coincided with the plausible but undocumented stoppage of phloem flow following burning.Will et al. (2007) suggested that the aphids reacted to the postulated burning-induced sieve tube occlusion by secreting watery saliva into the sieve element in order to unplug the tube. It is worth noting that the aphid saliva could not possibly have prevented the assumed forisome-dependent stoppage of phloem flow that had been triggered by burning the leaf several centimetres upstream

(source-ward) of the aphid (Will et al., 2007). Phloem transport velocities measured in intact plants ranged from 0.25mm s–1 to 0.4mm s–1 (Windt et al., 2006), implying that the entire contents of a large V. faba sieve element of 250 μm length (Peterset al., 2006) are completely replaced every 0.6–1 s. Thus, in an operating sieve tube, watery saliva will be strongly diluted and carried away immediately in the downstream direction, ruling out the possibility that saliva components could interact with P-proteins upstream of the inserted aphid stylet (the preferential translocation of salivary components towards sinks has been

141 demonstrated in principle, but the temporal resolution of those experiments—24 h—did not allow for conclusions concerning fast processes on the cellular scale; Madhusudhan and

Miles, 1998). Similarly, it seems practically impossible that the saliva was responsible for the assumed reopening of the phloem in the experiments of Will and colleagues. In a blocked sieve tube with stagnant contents, injected saliva components can travel by diffusion only. Therefore, it is conceivable that a significant concentration of saliva components could build up in the sieve element into which they are secreted, maybe also in the adjacent sieve elements on both sides, but certainly not all the way up to the wounded tissues several centimetres away. Thus, it is unclear how the secretion of watery saliva could provide a continuous flow of phloem sap which obviously requires certain lengths of tubes.

Based on the fact that aphids secret watery saliva while penetrating sieve tubes (Prado and

Tjallingii, 1994), various authors have asserted that aphids ‘release Ca2+-binding proteins in the phloem sieve cells preventing occlusion of these cells upon mechanical damage by the aphid stylets’ (Hougenhout and Bos, 2011, p. 424; compare Will et al., 2009;Hilker and

Meiners, 2010). This interpretation pre-supposes that stylet insertion triggers a release of

Ca2+ into the sieve element, an idea that seems intuitive for two reasons. First, Ca2+ ions are involved in numerous cellular signal transduction processes in plants including the interaction with herbivorous arthropods (Maffei et al., 2007a, b) where cellular Ca2+levels rise in the immediate vicinity of bite-induced injuries (Maffei et al., 2004). It must be cautioned, though, that biting herbivores and probing aphids inflict distinct types of wounds in different kinds of cells that do not necessarily launch similar responses. The

142 notion also seems plausible because of the assumed analogy between aphid stylets and microelectrodes, which may trigger sieve tube occlusion when inserted into a sieve element [Will et al. (2007, (2013) refer to microelectrode experiments reported by Knoblauch and van Bel (1998) to support this analogy]. However, microelectrodes actually can be inserted into sieve elements without causing damage (Knoblauch and van

Bel, 1998), and electrophysiological studies of sieve elements using intracellular microelectrodes are feasible (Hafke et al., 2003; Furch et al., 2009), demonstrating that the analogy does not hold. Moreover, the general facts should be stressed that in contrast to aphid stylets, microelectrodes have not been reported to produce a protective sheath around themselves as they penetrate the tissue, do not bend around cells when their tips proceed through multiple cell layers, and have a tapering shape that causes destruction in overlying tissue when deeply embedded cells are impaled. So one may ask: what is the empirical evidence supporting the idea of a stylet insertion-induced Ca2+ rise in sieve elements? Astonishingly, there does not seem to be any. Quite the contrary—the first published investigation into the behaviour of Ca2+-regulated phloem proteins during the initial phase of aphid attack reports that forisomes did not respond to stylet insertion even before E1 salivation started (Walker and Medina-Ortega, 2012). As a result, the authors found it ‘difficult to envision a potential role of E1 salivation immediately after sieve element penetration in preventing sieve element occlusion in the pea aphid–faba bean interaction. The possibility cannot be ruled out that E1 salivation at the onset of phloem phase [i.e. the period just after sieve element penetration] serves a function totally unrelated to phloem-sealing responses’ (Walker and Medina-Ortega, 2012, p. 333). In a

143 subsequent in vivo study, the same authors tested the hypothesis that apparent sieve element occlusions by high-volume forisomes are removed through interactions of the forisome with the saliva an aphid secretes into the sieve element. They found no differences in the behaviour of forisomes in sieve elements with and without saliva- secreting aphids (Medina-Ortega and Walker, 2013).

One has to conclude that the idea of an involvement of forisomes in the response to phloem sap-feeding insects is not supported by the empirical data available at this time. As a consequence, the interaction of concentrated aphid saliva proteins with forisomes in vitro (Will et al., 2007) is intriguing but of unclear physiological significance.

4.2.2 SEO, SEOR, and legume evolution

As mentioned above, forisomes can be isolated in large numbers (Knoblauch et al.,

2003). This facilitated the identification of forisome proteins and candidate genes. Tagging of the gene products with green fluorescent protein (GFP) resulted in fluorescent, reactive forisomes (Pélissier et al., 2008). The gene family was named sieve element occlusion

(SEO; Pélissier et al., 2008)—which was bold, as no efficient sieve element occlusion by the corresponding proteins had been demonstrated. Intriguingly, the same authors found similar genes in published sequences of non-papilionoids in which forisomes have never been reported, and these sieve element occlusion-related (SEOR) genes had a homologue in the papilionoids themselves. The gene products of both groups—SEO and SEOR—could be distinguished unambiguously on the amino acid sequence level: the papilionid SEOR protein was significantly more similar to non-papilionid SEORs than to papilionid SEOs,

144 and both groups were defined by unique sets of conserved motifs (Pélissier et al., 2008).

Taken together, these findings prompted the hypothesis that ‘a previously not characterized, well-defined group of proteins [i.e. SEOR] exists in higher plants including the Fabaceae, from which the evolution of SEO proteins in the Fabaceae originated’

(Supplementary Data 3 of Pélissier et al., 2008). Supposedly, the SEO gene family had branched from the widely distributed SEOR gene family in that lineage that gave rise to the last common ancestor of the papilionoid legumes (Peters et al., 2010). The idea is in agreement with the fact that no P-proteins other than forisome-forming SEOs have been shown to respond to Ca2+ (for reports of unsuccessful attempts, see Knoblauch et al.,

2001; Froelich et al., 2011). Available evidence thus suggests that Ca2+ responsiveness evolved in the ancestral protein at the root of the SEO protein family (Peters et al., 2010). It is worth emphasizing that this interpretation is in line with current views of legume evolution (for an overview, see Legume Phylogeny Working Group, 2013).

In the following year, Lin et al. (2009) detected a protein homologous to the one now called

AtSEOR1 (compare Froelich et al., 2011) in the phloem proteome of Cucurbita maxima. At the same time, the cucumber (Cucumis sativus) genome was published by Huang et al. (2009), leading to the identification of a cucumber homologue of the Arabidopsis gene that encodes AtSEOR1. Huang and collaborators concluded that ‘sieve element occlusion proteins (gene cluster 4754), present in all eudicots but absent from mosses and monocots, represent a novel mechanism that evolved for sealing the sieve tube system after wounding

(Pélissier et al., 2008)’ (Huang et al., 2009, p. 1280; our emphasis). In this statement, Huang and co-workers confused SEOR and SEO proteins as originally defined, and jumped to a

145 conclusion regarding SEOR function and evolution that lacked an empirical basis, and that certainly was not supported by the reference cited. However, the presence of SEO-related genes in non-papilionoids could hardly be considered surprising from here on.

Obviously unaware of the earlier discoveries, Rüping et al. (2010, p. 1) announced the ‘unexpected occurrence’ of SEO-related genes in non-papilionoids. These authors expanded the analysis of SEO/SEOR sequence similarities, and also identified possible orthologue and paralogue relationships between SEO as well as between SEOR genes, both within and between species. Unfortunately, they ignored the sequence-based distinction between SEOR and SEO gene products although their data were in line with this interpretation. Like Huang et al. (2009) before them, they used the term ‘SEO’ in an inclusive sense that comprised SEOs and SEORs, only to define a subgroup, ‘SEO-F’, that included all proteins for which an involvement in forisome formation had been demonstrated experimentally (Rüping et al., 2010). We consider this nomenclature unnecessarily confusing because it bases the definition of groups of genes partly on sequence data and partly on the gene product’s function. On the other hand, Rüping et al. (2010) never provided a rationale for rejecting Pélissier et al.’s (2008) distinction between SEOs and SEORs. Therefore, we will retain the original definitions, first and foremost because they integrate the molecular facts into the wider evolutionary picture.

4.2.3 Sieve tube slime: same player shoots again!

The existence of SEO-related genes in plants not shown to generate forisomes raised questions. Are the gene products located in sieve elements? If so, what is their structure,

146 and do they possess phloem-specific functions? Tagging of Arabidopsis (Froelich et al.,

2011) and tobacco (Ernst et al., 2012) SEOR gene products with fluorescent proteins revealed meshworks of SEOR filaments within sieve tubes and dense slime agglomerations that occluded the sieve elements—or so it appeared from the micrographs. Evidently, SEOR proteins represent or are at least part of the sieve tube slime of the older literature. It has long been discussed why multiple occlusion mechanisms including callose de novo synthesis and slime plug formation by P-proteins (SEO and SEOR) appear to exist

(Sabnis and Sabnis, 1995). The model plant Arabidopsis possesses twoSEOR genes; both

AtSEOR1 and AtSEOR2 must be present for SEOR filaments and agglomerations to form

(Anstead et al., 2012). Soon a debate started about the possible physiological function(s) of

SEOR agglomerations that presents yet another chapter of the sieve tube slime controversy that traces its origins to the middle of the 19th century (Sabnis and Sabnis,

1995).

By 1860, light microscopy had revealed the basic structural principles of elongated phloem cells with perforated end walls (Hartig, 1837) whose function appeared to be long- distance transport (Hartig, 1860). Dense proteinaceous slime masses that consistently were found on the perforated walls which separated these sieve elements were in line with the contemporary notion that sieve tubes represented a storage and transport tissue for nitrogen-rich compounds, but not for carbohydrates (Strasburger, 1891). In those days, the translocation of sugar solutions in sieve tubes seemed unlikely for a variety of reasons, and the apparent occlusion of sieve plates by protein agglomerations was one of them. It took an outsider who struggled to establish a career, Alfred Fischer, to demonstrate that the

147 slime masses consistently observed were artefacts caused by common but inappropriate preparation protocols, and that the contents of live sieve elements were more or less homogeneous and apparently fluid (Fischer, 1885). Bulk fluid flow in sieve tubes thus became a plausible concept. Before long, turgor-driven bulk translocation was discussed in textbooks (e.g. Haberlandt, 1896), ultimately leading toMünch’s (1926, 1927, 1930) presentation of the conceptual framework of an osmotically generated, pressure-driven flow that dominates current thinking about the mechanisms of phloem transport

(Knoblauch and Peters, 2013).

With the advent of electron microscopy in the 1930s, investigators realized that sieve elements contained structural components that had remained invisible in the light microscope. Proteinaceous slime in the lumen of sieve elements and in sieve pores, now called P-protein, made a reappearance and created a problem for Münch’s pressure flow hypothesis. The hydraulic resistance to bulk flow offered by dense protein agglomerations in sieve pores appeared too high to be overcome by pressure gradients of plausible magnitudes. Alternative explanations for phloem translocation were developed

(MacRobbie, 1971; Wardlaw, 1974; see also the four review articles by Canny, Spanner,

Milburn, and Fensom in Zimmermann and Milburn, 1975) yet the Münch hypothesis still prevails as the leading hypothesis. One reason was that numerous workers in the field never stopped believing that occluded sieve plates represented preparation artefacts rather than the functional state. A number of novel preparation methods were devised, and some indeed showed open pores. However, the debate remained unresolved for decades.

148

The digital age provided new tools such as CLSM, which enabled the live imaging of functional sieve tubes. Important findings produced with the new tool included the direct confirmation of bulk flow in the phloem, and the visualization of the formation of P-protein agglomerations on sieve plates in response to injuries (Knoblauch and van Bel, 1998). In this context, the observation of dense SEOR agglomerations in apparently functional, uninjured Arabidopsis sieve elements mentioned above came as a surprise. What was even more surprising was the fact that the apparent sieve element occlusions by SEOR agglomerations seemed to have little effect on flow velocity, as the comparison of functional sieve tubes in roots of wild-type plants and AtSEOR1 knock-out mutants showed

(Froelich et al., 2011).

4.2.4 SEOR proteins: fluid dynamic effects and specific functions

Hydraulic effects of SEOR agglomerations in intact plants

P-protein agglomerations in sieve tubes that appear to occlude the tube have been reported to allow the passage of fluid and dissolved macromolecules (Kempers et al., 1993) before the recent studies on AtSEOR1 agglomerations (Froelich et al., 2011). To understand the counterintuitive ineffectualness of such apparent sieve tube occlusions,Froelich et al. (2011) studied AtSEOR1 sieve tube agglomerations in the roots of intact Arabidopsis in depth. Based on sieve element structure and the ultrastructure of AtSEOR1 agglomerations, the authors evaluated the contribution of the flow resistance offered by the SEOR agglomerations to the total hydraulic resistance (R total) in the sieve tube:

Rtotal=n Rlumen+(n−1) Rplate+m Raggl (1)

149 where R lumen is the resistance of the lateral walls of one sieve element and n is the number of sieve elements in a tube, R plate is the resistance of a sieve plate, and R aggl is the resistance of one of the m SEOR agglomerations present in the tube. The authors concluded that for a typical Arabidopsis sieve tube, n R lumen and (n−1) R plate are about equal, whereas m R aggl is somewhat smaller, owing to the comparatively low frequency of agglomerations (~1 per

10 sieve elements). While the value of R aggl can only be estimated as it depends on the porosity of the SEOR protein material which is not quantitatively known, calculations based on a range of plausible assumptions suggested that bulk flow driven by turgor pressure differentials of the expected magnitudes should be possible with the observed frequency of

SEOR agglomerations (Froelich et al., 2011).

There are several important conclusions to be drawn from these findings. First, a protein agglomeration in a sieve tube does not necessarily produce total occlusion, no matter how dense and tight it may look on a micrograph. Secondly, the contribution of

AtSEOR agglomerations to total flow resistance is probably smaller than that of open sieve plates and that of the tube itself. Thirdly, despite its relatively small contribution to total flow resistance, the resistance offered by AtSEOR agglomerations is a significant biophysical factor; Froelich et al. (2011) estimate R aggl to be ~20% of R total. Since the volumetric flow rate, Q, in a sieve tube relates to its driving force given by the pressure differential, Δp, and the total hydraulic resistance, R total, according to

Q=Δp/Rtotal (2)

150 our conclusions imply that a plant can maintain a given flow rate under increasing numbers of SEOR agglomerations as long as its phloem loading and unloading machineries are capable of increasing Δpcommensurately. Such functional adjustment does not necessarily require complex regulation (which might be hard for the plant to accomplish anyway; Thompson and Holbrook, 2003b; Thompson, 2006). Phloem flow according to

Münch’s hypothesis is driven by loading and unloading in sources and sinks, respectively; the osmotically induced Δp is generated by the loading and unloading processes, and also links them mechanistically like a transmission belt. If R total along the pathway rises and loading and/or unloading continues, Δp will increase automatically, either until a new equilibrium according to Equation (2) is established, or until the loading/unloading machinery reaches maximum capacity (this effect has been measured in vivo by Gould et al.,

2004).

To appreciate fully our proposed explanation of why Froelich et al. (2011)did not detect any differences in the phloem flow velocities between wild-type plants and AtSEOR1 knock-out mutants, it is important to realize that they made their observations in intact plants growing in the newly developed Micro-ROCs. In these miniature rhizotrons, roots remain in contact with natural soil at all times, even while being observed under the microscope. Such a nearly natural environment obviously is preferable over the artificial environment provided by the usual agar plate cultivation methods when a systemic phenomenon such as phloem translocation is studied in vivo. In the whole-plant physiology approach which Froelich and co-workers took, the plants studied were intact except for a small leaf incision for fluorescent dye loading, and did not

151 need to be removed for experimentation from the natural soil in which they grew. There is no evidence and plausible reason why one should assume that the phloem loading and unloading machineries in these plants were not fully operational. Consequently, phloem flow proceeded at similar velocities with and without SEOR proteins.

4.2.5 Hydraulic effects of SEOR agglomerations in excised organs

In intact plants, flow rates in the phloem (Q) can be maintained as long as shifts in R total are balanced by changes in Δp [see Equation (2)], which requires full functionality of the loading/unloading machineries. The latter include a potent water source—the xylem—to fuel the osmotic generation of hydrostatic pressure, especially in source organs.

Therefore, the influence of R aggl on phloem flow might become detectable in excised organs in which the ability to modulate Δp is impaired due to the disconnection from the physiological water source. When a petiole is cut, export of fluid from the leaf through the phloem must slow down, because the refilling of the sieve elements becomes more difficult in a leaf that is disconnected from the plant’s xylem. Consequently, we may expect to see a correlation between the rate of phloem exudation and the amount of SEOR proteins in excised leaves. Such a correlation has been demonstrated for excised leaves of tobacco

(Ernst et al., 2012) andArabidopsis (Jekat et al., 2013). In both cases, the contribution of the phloem to the total exudate secreted from the excised leaves over a period of 10min was estimated, in both wild-type plants and genetically modified plants lacking SEORs, by measuring the amount of glucose exuded in the presence and absence of β-fructosidase.

Under the assumption that none of the glucose but all of the sucrose present in the original exudates originated exclusively from sieve elements (for possible problems with this

152 assumption, see van Bel and Hess, 2008; Liu et al., 2012), it was inferred that the presence of SEOR proteins reduced phloem exudation from excised leaves by factors of nine in tobacco (Ernst et al., 2012) and two in Arabidopsis (Jekat et al., 2013). It should be emphasized that SEOR proteins did not occlude (in the common sense of block) or seal the sieve tubes, but only reduced flow rate Q under conditions where the capability to maintain turgor and thus Δp in the phloem was disturbed compared with the intact plant. These findings are in full agreement with the notion that SEOR agglomerations add a summand

(m R aggl) to the total hydraulic resistance of the sieve tube (R total), according to Equation

(1). A specific wound response is not required to explain the observations.

To obtain a more intuitive picture, imagine a garden hose of 1cm inner diameter and

1 km length; this roughly equals the length-to-diameter ratio of a sieve tube extending from the inflorescence of an Arabidopsisplant to a root tip. Clearly, one will have to apply pressure to drive water through this hose, and even more pressure will be required to drive flow through a similar hose in which solid dirt deposits increase the total hydraulic resistance by a quarter. Consequently, if we cut the clean and the dirty hose in the middle, we expect that the water will flow from the clean halves faster than from the dirty halves, and this is what Ernst et al.(2012) and Jekat et al. (2013) have shown.

The original authors seem to disagree. In the title of their paper, Jekat et al. (2013, p. 1) announced that Arabidopsis P-proteins (AtSEORs) are ‘involved in rapid sieve tube sealing’, which somewhat overstates the matter—a reduction in phloem exudation by half over

10min. Similarly,Ernst et al. (2012, p. E1987) claimed to ‘have demonstrated clearly that P-

153 protein accumulations block translocation following injury’. This, however, is misleading since what they showed was reduced, not blocked translocation, and because no comparison between the injured and the non-injured state was presented. Taken together, neither Ernst et al.(2012) nor Jekat et al. (2013) provided evidence to support the idea that

SEOR agglomerations affect sieve tube flow through mechanisms other than a merely structural contribution to total hydraulic resistance. Their conclusion that the demonstration of such a structural contribution establishes a role for SEORs in injury responses and sieve tube sealing is logically flawed. To see why, consider viscosity, a parameter that so far we had excluded from our discussion (and from Equation 1) to keep things simple. Viscosity is the internal resistance of a medium to being sheared as in pipe flow, which the driving force of flow, in our case Δp, must overcome to initiate and maintain flow; each of the several resistive terms in R total [compare Equation (1)] is commonly thought to be linearly proportional to it. In transporting sieve tubes, sucrose usually is the most important solute. The viscosity of sucrose solutions depends on various physical factors, but under physiologically relevant conditions there is a straight-forward, positive relationship between sucrose concentration and viscosity (Longinotti and Corti, 2008).

Now let us set, for argument’s sake, all parameters other than the sucrose concentration constant. In this case, a decreased sucrose level will result in a decreased bulk viscosity and thus in an increased flow rate (Thompson and Holbrook, 2003a; Hölttä et al., 2006).

However, this is analogous to what can be said about a decreased number of SEOR agglomerations, which, if all other parameters remain unchanged, will also result in an

154 increased flow rate. To be sure, no plant physiologist will conclude that sucrose functions in sieve tube sealing following injury.

Among plant physiologists, the claim that sieve tube occlusion by P-proteins following injury prevents the loss of energetically expensive photoassimilates would hardly meet resistance. However, whether photoassimilates flowing from severed sieve tubes represent a significant contribution to injury-induced losses is not clear. In the frequently studied cucurbits, the fluid material lost from open wounds, for example after leaf excision, originates mostly from non-phloem cells (Zhang et al., 2012), but this does not necessarily tell us what proportion of the soluble carbohydrates lost exudes from the phloem.

Analysing the exudation data presented as fig. 4B by Jekat et al. (2013) for Arabidopsis leaves, we find that phloem-derived sugars contributed a mere

16% of the total sugar loss through exudation in the wild type. This proportion roughly doubled in the two AtSEOR knock-out mutants, while the amounts of sugars apparently originating from non-phloem sources were slightly decreased in the mutants. Taking both trends into account, we find that the presence of AtSEORs in wild-type plants prevented about one-fifth of the total sugar loss from excised leaves that Jekat et al. (2013) had observed in the AtSEOR knock-out plants. While this estimate is not impressive, it still may be too high. Given the stochastic variation in the data, the AtSEOR effect is too weak to show in the total budget: we detect no statistically significant difference in the total amounts of sugars lost between the wild-type and knock-out plants. In the data from an analogous experiment using tobacco (fig. S6 in Ernst et al., 2012), we find a relative proportion of apparently phloem-derived sugars of about three-quarters of the total

155 concentration in the exudate from SEOR knock-out leaves. Since the amount of phloem- derived sugars was nine times higher in SEOR knock-out than in wild-type leaves while the levels of non-phloem-derived sugars were about equal (Ernst et al., 2012), we arrive at the conclusion that SEOR proteins reduced the amount of carbohydrates exuding from excised leaves by approximately two-thirds in tobacco. This estimate is much higher than that for Arabidopsis, indicating large differences between species in the same experiment.

Nonetheless, the general conclusion remains: photosynthate leakage due to continuing phloem translocation is not always the most dramatic loss a wounded plant experiences.

This fact as such does not speak against a specific function for SEOR proteins in sieve tube sealing, but it questions the relative importance of any such function, should one exist, for the prevention of sugar loss following injury.

Last but not least, an obvious fact deserves to be highlighted in this context: the fitness of a plant cannot possibly increase from the reduction of sugar loss from a peripheral organ after excision of that organ. To generalize conclusions from studies on excised leaves (as done by Ernst et al., 2012; Jekat et al., 2013), one has to assume that the isolated organs are valid partes pro toto and functionally represent intact plants. However, as we have argued above, phloem transport is a systemic phenomenon that is lost when the system is cut into pieces, and, even if residual phloem functionalities remain, not all pieces are equal after cutting. Results obtained with excised leaves cannot simply be extrapolated to the intact or remaining plant; whether mechanisms that apparently reduce sugar loss where it is irrelevant (excised leaves) are valid models for mechanisms that may prevent sugar loss where it may count (petiole stumps on the plant) is not guaranteed.

156

4.2.6 SEOR interactions with and responses to stress factors

On the conceptual level, it is essential to distinguish between specific biological processes on one hand and mere physical necessity on the other. Forisome action provides an example of a specific biological process. Forisomes change the hydraulic resistance they offer to sieve tube flow through a process—interaction with Ca2+—that is under the control of the live sieve element, which regulates cytosolic Ca2+ in response to external stimuli.

Despite the open questions discussed above, these facts very strongly support the idea that forisomes function in the regulation of phloem translocation. On the other hand, any object in the path of the flowing sieve tube contents will add to overall hydraulic resistance, tending to slow the flow. Therefore, the fact that SEOR agglomerations in sieve tubes reduce flow rates does not prove anything. It strongly suggests, though, that SEOR proteins have a beneficial function or functions that outweigh the disadvantage the plant incurs; first, by the cost of synthesis of the SEOR proteins, and; secondly, by the increased sieve tube hydraulic resistance. What do we know about potential functions of SEOR proteins in the regulation of phloem activity?

To test the responsiveness of AtSEOR agglomerations to various treatments known to induce rapid stress responses in functional sieve tubes, Froelich et al. (2011) mechanically injured sieve tubes, applied distant wounds by burning and local cold shock, and added Ca2+ to open sieve tubes and SEOR agglomerations. No immediate reactions were observed. In a few cases, the protein agglomerations started to move slowly towards the downstream sieve plate but did not occlude it; the protein rather continued its movement through the pores. This process could be followed for at least 45min, suggesting

157 that electron micrographs previously thought to show occluded sieve pores may in fact represent snapshots of ongoing translocation. We here report an extension of the experiments of Froelich et al. (2011). Electroshocks are known to stop phloem flow rapidly

(Pickard and Minchin, 1990, 1992a, b), and we used pAtSEOR1:AtSEOR1:YFP (yellow fluorescent protein)Arabidopsis plants growing in Micro-ROCs as detailed before

(Froelich et al., 2011) to investigate the possible involvement of SEOR proteins. Through small holes made in the walls of the Micro-ROCs, we placed electrodes in the soil next to a root ~1cm apart from each other and applied voltages of 10–120V at pulses of 1–5 s. Even at a field strength of 8kV m−1, there was no visible reaction of SEOR proteins. However, at field strengths >10kV m−1, SEOR agglomerations disappeared. However, this could hardly be considered a specific response because at the same time irreversible distortions of the entire root system occurred.

Taken together, the idea of an involvement of AtSEORs in targeted sealing mechanisms finds no support in the lack of AtSEOR responses to stimuli known to affect sieve tube transport. But do Arabidopsis plants without SEORs respond to such stimuli as their wild-type conspecifics do? We studied wild-type plants (which obviously produced

SEOR proteins), pAtSEOR1:AtSEOR1:YFP transgenic plants (in which fluorescent AtSEOR1 could be observed microscopically), and SEOR knock-out plants (which lacked SEOR agglomerations). In all three plant types, phloem translocation in roots was monitored by

FRAP (fluorescence recovery after photo-bleaching) after the feeding of CFDA

(carboxyfluorescein diacetate) into the phloem, employing the methods we have described before (Froelich et al. 2011). The plants were kept in Micro-ROCs during the experiment

158 and were cold shocked by applying ice water to the hypocotyl and most proximal part of the root system. As expected, the AtSEOR1 agglomerations visible in the pAtSEOR1:AtSEOR1:YFP plants showed no reaction. Phloem flow, however, stopped within seconds in all three plant lines (Fig. 1; Supplementary Movie S1 available at JXB online), even in SEOR knock-out lines. These results indicated that Arabidopsis does not require

SEOR proteins to halt phloem translocation in response to cold shock.

159

160

Fig. 1: Abrupt cold causes stoppage of phloem translocation in the roots of AtSEOR1 knock- out Arabidopsis plants. After the phloem had been loaded with the fluorescent dye carboxyfluorescein diacetate (CFDA), the dye was photobleached and the effect of cold shock on the movement of the dye front was observed. (A) The dye front (arrow) is moving into the field of view just before the cold shock is applied. (B) The plant is cold-shocked; the dye front slows down (C) and comes to a halt 15 s after the shock (D). (E–G) Dye movement did not occur over the next 30 s, and did not resume for at least another minute (see Supplementary Movie S1available at JXB online). For comparison, in control experiments without cold shock, the dye front moved through the entire horizontal length of the images in <4 s. The times on each image refer to Supplementary Movie S1 available at JXB online from which the images were taken. Plant cultivation, CFDA feeding to the phloem, confocal laser-scanning microscopy, and photobleaching were performed as described in detail byFroelich et al. (2011).

It has been speculated that P-proteins block sieve pores in Arabidopsis in response to the insertion of an aphid stylet into the sieve tube (Kuśnierczyk et al., 2008). If the idea is correct and applicable to SEOR proteins, we should expect that aphids exploiting Arabidopsis greatly benefit from living on AtSEOR knock-out mutants rather than on wild-type plants. However, the opposite seems to be true: aphids exhibited decreased fitness (in terms of life time offspring production) when grown on plants lacking SEOR proteins (Anstead et al., 2012), indicating that the presence of SEOR proteins may even be beneficial to the insects at least in the compatible interaction of Myzus persicae and Arabidopsis. This finding is not totally unexpected, as amino acid availability often limits aphid growth and reproductive success (Sandström and Moran, 2001;Douglas,

2006), and aphids seem to prefer host plants growing on N-rich substrates (Nowak and

Komor, 2010). Moreover, at least some aphid species possess proteolytic enzymes to break down ingested phloem proteins (Pyatti et al., 2011), an ability that previously had appeared doubtful (for a review, see Kehr, 2006). Therefore, it is not entirely implausible

161 that SEOR proteins and their building blocks might actually increase the nutritional value of the phloem sap for aphids. Plants may benefit from the presence of P-proteins due to increased resistance in other plant–insect interactions and P-proteins may be one reason for incompatibilities. However, only direct experimental evidence can validate this idea.

In summary, currently available information suggests that (i) SEOR proteins do not seal the phloem efficiently in the case of injury; (ii) SEOR agglomerations show no visible responses to selected stimuli known to slow or halt phloem translocation; (iii) SEOR proteins show no visible responses to Ca2+, an effector widely thought to control sieve tube occlusion; (iv)

SEOR knock-out plants display qualitatively unchanged cold-shock-induced stoppage of phloem transport; and (v) SEOR proteins promote the well-being of phloem sap-feeding aphids, at least in compatible interactions of M. persicae and Arabidopsis. These findings do not support currently popular notions concerning the physiological roles of dispersive P- proteins in general and SEORs in particular. We conclude that the biological function of

SEOR proteins remains obscure at this time.

4.2.7 Iconoclastic speculations…

We believe that valid interpretations of several key experiments are possible that go against the grain of currently popular notions. In this section, we present some of these iconoclastic ideas. We do not necessarily think that they are all correct; however, we do feel that the current debate might benefit from considering alternative views.

… on forisomes

162

According to a current model, forisomes occlude sieve elements in response to a transient membrane depolarization (sometimes referred to as a ‘plant action potential’) that originates from sites of injury (leaf burning, in particular) and travels from there along the vasculature at velocities in the range of 1mm s−1 (Furch et al., 2007). The membrane depolarization coincides with Ca2+ influx into the sieve element; thus, the travelling ‘plant action potential’ is accompanied by a travelling cytosolic Ca2+ wave. However, the rise in cytosolic Ca2+ brought about by the ‘action potential’ has been reported to be too weak to trigger the transition of forisomes into the high-volume state (Furch et al., 2009).

Supposedly, an amplification of the Ca2+ signal by Ca2+ released from the ER is required to produce local Ca2+ hotspots, and only forisomes located at such a hotspot appear likely to respond by switching into the high-volume state (Furch et al., 2009). A problem rarely discussed in this context is flow. As mentioned above, we have to assume that the entire fluid content of a typical, transporting Vicia sieve tube is replaced in less than a second, which is in the range of the typical reaction time of Viciaforisomes that we observe in situ.

This flow vector has to be added to any diffusional movement of dissolved particles, making it hard to imagine how stable, local Ca2+ hotspots could develop in a transporting tube. If the hotspot scenario described by Furch and co-workers (2009) is valid, the system would appear to work optimally when flow has come to a halt already. In other words, forisomes would expand as an indirect consequence of flow stoppage, as it were.

The idea that forisome transition into the high-volume state is a result rather than the cause of a stoppage of phloem flow runs against deeply engrained convictions. For example, in their seminal paper on aphid interference with forisome function, Will and co-

163 workers (2007) did not monitor phloem flow but assumed that the burning of their experimental leaves induced a remote sieve tube occlusion to which the aphid would react.

The reason that apparently justified the assumption was that in a similar set-up ‘burning of V. faba leaf tips results in remote dispersion of forisomes, followed by a stoppage of mass flow (Furch et al., 2007)’ (Willet al., 2007, p. 10537). However, Furch et al. (2007) did not measure mass flow; thus there is no way of knowing whether there was a stoppage and, if there was, whether flow stoppage preceded, followed, or coincided with the forisome response. However, if we accept the explanation Furch et al.(2007) offered for their results, then we will have to conclude that the transition of forisomes into the high-volume state actually followed the stoppage of flow rather than vice versa in the experiments by Furch and colleagues as well as in those by Will and colleagues. Why?

Imagine the depolarization and cytosolic Ca2+ wave described by Furch et al. (2007, 2009) running along a sieve tube. Because the Ca2+ stimulus that triggers the forisome response travels along the sieve tube network, a zone of high-volume forisomes will expand like a wake behind that travelling trigger. In other words, just before a given forisome reacts, the forisomes in the upstream sieve elements (those located towards the origin of the moving cytosolic Ca2+ wave) must already have reacted. If forisomes in the high-volume state actually occlude (in the sense of block flow) sieve elements, flow stoppage thus must precede the forisome response. In this scenario, the idea that the transition of forisomes into the high-volume state depends on the build-up of local

Ca2+ hotspots (Furch et al., 2009) is more easily digestible than in a transporting sieve tube

164 in which a concentration hotspot might be carried past a typically sized Vicia forisome in

<100ms.

One may say that our argument creates a chicken or egg problem; if the forisome response requires prior flow stoppage, why does flow stop in the first place? Furch and co- workers do not give details on the destruction caused by the burning injury that they applied to initiate the flow stoppage/forisome transformation cascade, but remarked that tissue movements under the microscope were inevitable consequences of the pressure waves induced by burning (Furch et al., 2009, p. 2121). It is inconceivable that the sieve tube network at the burning site survived such treatment without structural damage, implying that sieve tubes were opened, resulting in a pressure drop and a stoppage of translocation in the vicinity of the burned tissue. This forisome-independent, initial stoppage of translocation at the wounding site may have allowed forisomes to switch into the high-volume state, thereby triggering the expanding flow stoppage/forisome transformation cascade that Furch and colleagues analysed.

We speculate that under the conditions described above, forisomes might not function in stopping flow, but rather in locking an idling sieve tube network in its physiologically passive state. In this interpretation, forisomes could be viewed as analogues of the plaster cast around a broken ankle, providing stability to the system by preventing any attempts to perform normal function, thus enabling undisturbed repair activities.

165

4.2.8 … on P-proteins and aphids

Screening the recent literature, one can hardly escape the conclusion that the role of

Ca2+-induced sieve tube occlusion in defending the plant against attacks by phloem sap- feeding insects is firmly established (e.g.Goggin, 2007; Kuśnierczyk et al., 2008; Hilker and

Meiners, 2010; Consales et al., 2011; Hogenhout and Bos, 2011; Kamphuis et al.,

2013;Rodriguez and Bos, 2013; Will et al., 2013). As the references usually given in this context show, the notion rests exclusively on the finding that Ca2+-binding proteins from concentrated aphid saliva can inhibit the Ca2+-dependent transition of isolated forisomes into the high-volume state in an in vitro assay (Will et al., 2007). It is worth stressing once again that neither the prevention of sieve tube occlusion by Ca2+-binding saliva components nor the removal of existing occlusions by such components have been demonstrated experimentally (cf. Medina-Ortega and Walker, 2013).

Nonetheless, the results Will et al. (2007) produced with V. faba were generalized by several authors to cover angiosperms in general, despite the facts that forisomes are specific to the papilionoids and that no Ca2+responsiveness has ever been reported from phloem proteins other than forisomes. For example, Kuśnierczyk et al. (2008, p. 1109) presented a model of defence mechanisms in Arabidopsis in which a ‘rising concentration of

Ca2+ in sieve elements initializes protein clogging’. The incorrect notion that P-proteins other than forisomes respond to Ca2+ in such a manner has been promoted by claims such as: ‘Occlusion is triggered by Ca2+ influx induced by damage (Knoblauch and van Bel, 1998)’

(cited from Will et al., 2009, p. 3305). However, while Knoblauch and van Bel

(1998) certainly documented the formation of supposedly irreversible depositions of cell

166 components on sieve plates following severe injury, they did not mention, let alone demonstrate, a role for Ca2+in the process. Will et al. (2009) expanded their original aphid behavioural experiment (Will et al., 2007) to four plant species including three dicotyledons and Hordeum vulgare, a member of the monocotyledonous Poaceae, or grass family. They also determined the stoppage of bulk flow, and found no significant differences between the four plant species regarding flow stoppage and aphid saliva secretion as induced by leaf burning. The authors concluded that sieve plate plugging by phloem proteins is a universal phenomenon occurring in all species, even the grass H. vulgare (Will et al., 2009). However, H. vulgare lacks P-proteins (Evert et al., 1971) as grasses do in general (Eleftheriou, 1990). Therefore, the conclusion that the presence of P- proteins is entirely unrelated to flow stoppage and aphid behaviour in these experiments is at least equally plausible.

For argument’s sake, let us ignore the empirical evidence (Froelich et al.,

2011; Walker and Medina-Ortega, 2012; Medina-Ortega and Walker, 2013) for a moment and assume that the insertion of an aphid stylet into a sieve tube triggers sieve plate occlusion by Ca2+-responsive P-proteins.Will et al. (2007) had interpreted the switch from

E2 to E1 EPG patterns after a burn stimulus as indicative of occlusions of the sieve tubes on which their aphids were feeding. Later they could induce similar EPG switches by reducing the hydrostatic pressure in an artificial feeding system (Will et al., 2008). This seemed to make good sense to Will et al.(2009, p. 3305) who maintained that ‘sieve tube occlusion is accompanied by a decrease of sieve tube pressure (Gould et al., 2004)’ (see also Will et al.,

2013, p. 6). However, Gould et al. (2004) had demonstrated a decrease in turgor only

167 downstream (sink-ward) of the sieve tube block; on the upstream (source-ward) side, turgor actually increased—exactly as we would expect if the loading/unloading machinery remained operational while the sieve tubes were blocked. On which side of a stylet insertion-induced sieve plate occlusion would we find the aphid? On the upstream or source-ward side of course, because any Ca2+ entering the sieve element at the penetration site will promptly be carried away in the downstream direction, and only there, downstream or sink-ward of the aphid, could it induce a protein plug. Thanks to the occlusion of the tube just downstream of the penetration site, the aphid would find itself at the downstream terminus of a continuous pipe connecting it directly to the source tissues.

Any volume of phloem sap the aphid may remove would immediately be replaced, especially (but not only) if turgor increases.

We cannot think of any reason why the aphid would want to release a sieve tube occlusion of this kind, and therefore speculate that sieve tube occlusions are not generally a bad thing for phloem-sap thieves. Our chances of elucidating the enigmatic biological function of P- proteins would probably not suffer if the consistently reiterated dogma that aphids need to prevent sieve tube plugging in order to enjoy a continuous flow of nutrients were to be carefully re-evaluated in a fluid dynamics context.

4.2.9 … on phloem exudation and wound sealing

In the late 19th century, Alfred Fischer (1885) demonstrated that the slimyocclusions, which at the time were interpreted as functionally essential components of sieve elements, were in fact artefacts caused by tissue preparation and fixation for light

168 microscopy. Fischer had discovered that by cutting through the phloem, he could induce the formation of slime agglomerations on that side of sieve plates that was facing away from the cut. He concluded that the slime had been carried to its position by the surging of the phloem sap towards the open cut, and hypothesized that the wounding-induced artefacts ‘served, so to speak, as provisional seals of the sieve tube system’ (Fischer, 1885, p. 236). More than a century later, we still have not identified possible physiological functions in the intact plant of the proteins Fischer called sieve tube slime. However, since

Fischer’s artefacts consistently occur when we prepare a plant for experimentation a little too clumsily, we have come to see the biological function in the artefact, assuming or rather implying implicitly that the evolution and phylogenetic conservation of energetically costly

P-proteins was and is driven only by the adaptive benefits of a protein-based emergency shut-down system that works in parallel with an already existing callose synthesis machinery (for a review, see Eschrich, 1975; a critical view is offered by Sabnis and Sabnis,

1995). We cannot exclude that the idea is correct; but neither can we exclude that our position is analogous to that of an extraterrestrial observer who, after having witnessed a few traffic accidents from his remote vantage point, concludes that the main function of automobiles is the prevention of direct contact between fast moving humans and obstacles in their path.

The provisional seal hypothesis of P-protein function appears intuitively plausible; plants must shut down injured sieve tubes promptly to avoid losing expensive photosynthates. But is that so always? Zhang and co-workers recently suggested ‘that the role of P-proteins in the cucurbits may be to prevent excessive water loss from wounded

169 xylem as much as it is to seal wounded phloem’ (Zhang et al., 2012, p. 1881). This suggestion is based on the observation of P-proteins that exude from severed sieve tubes rapidly in large amounts to form plugs that cover the entire cut surface of the vascular bundles. An argument following the same logics had been put forward by Read and

Northcote (1983) who suggested that lectins arriving at wound sites by phloem exudation carry out an anti-invasive role. These postulated functions obviously require the exact opposite of what usually is assumed: in order to deliver functionally important substances to wound areas, sieve tubes need to remain unoccluded to enable the loss of sufficiently large amounts of P-proteins and other factors, together with expensive photoassimilates.

Apparently, the assumption that plants must rapidly seal injured sieve tubes to prevent losing expensive materials is not quite as self-evidently true as it sometimes sounds. At least occasionally, the phloem seems to function like lactifers and secretory ducts, the defensive tube networks present in many tracheophytes that fulfil their ecological functions by extensive secretion (Franceschi et al., 2005; Pickard, 2008; Agrawal and

Konno, 2009); the extrafascicular phloem of the cucurbits may even be specialized for such a role in defence (Turgeon and Oparka, 2010; Zhanget al., 2010; Gaupels and Ghirardo,

2013). In this context, we are intrigued by the following thought. When a small herbivore chews away on a leaf, why should the plant allow sieve tubes to occlude? Photoassimilates lost through severed sieve tubes at the site of biting cannot be saved by sealing the tubes as they will be lost anyway with the herbivore’s next bite; would it not seem beneficial to crank up phloem loading in the leaf to export as much photoassimilate as possible in the remaining time, rather than locking transportable goodies in a doomed organ? Plants

170 respond differently to feeding herbivores and mechanical injury (Baldwin, 1988;Korth and

Dixon,1997; Reymond et al., 2000; Bricchi et al., 2010), so differential responses by the phloem to continuing biting as opposed to single wounding events are not implausible.

However, we do not intend to speculate about herbivore–plant interactions; what we are suggesting is that plugging sieve tubes in response to injury is not obviously and always a good idea. Whether a plant benefits from injury-induced sieve tube occlusions depends on the nature of the agent that inflicted the injury, the nature of the injury, and its position. If cases could be identified in which injury-induced sieve tube plugging by P-proteins evidently harms the plant—if, in other words, sieve tube plugging could be shown to be maladaptive—a strong argument against sieve tube plugging as the primary biological function of P-proteins could be made.

4.3 Conclusions

Proteinaceous sieve tube slime, aka P-proteins, has bamboozled plant physiologists for more than a century. We think that there are two main reasons. First, the rapid reaction of some types of P-proteins to injuries makes it difficult to distinguish unambiguously between their state in the functional, transporting sieve element on one hand and preparation-induced artefacts on the other. Secondly, some of the assumed preparation- induced artefacts actually may represent the functional state of P-proteins. We came to realize that the problem is aggravated by the linguistic sloppiness in many publications including some of our own. To say that a sieve tube is occluded, sealed, clogged, or plugged is not (and should not be meant as) a statement about how the tube looks, but about its functional state. If we would use these terms only in cases in which microscopically

171 visualized putative occlusions, seals, or plugs actually had been demonstrated to be temporally associated with stoppage of phloem translocation, the terms would become rare in our literature while we would be forced to take the fluid dynamics of the phloem seriously and analyse hydraulic resistances quantitatively. Another essential point in the elucidation of P-protein function is the apparent reversibility of any observed responses, which provides prima facie evidence for biological regulation. We consider it less than helpful when reversible processes such as forisome responses and callose deposition

(Knoblauch et al., 2001; Furch et al., 2007) are compounded with the irreversible effects of catastrophic structural failure (demonstrated, for example, by Knoblauch and van Bel,

1998) into all-embracing, overly generalized hypotheses, especially when evident functional differences between taxa are ignored. This seems to be the case with some current notions about aphid–plant interactions (cf. Smith and Boyko, 2007; Cooper et al.,

2011).

The direct observation of fully operational sieve tubes harboring SEOR proteins in Arabidopsis plants that were growing in an almost natural environment produced intriguing results (Froelich et al., 2011). AtSEOR agglomerations showed no visible reactions to various stimuli known to induce a slowing of phloem flow. Under certain circumstances, AtSEOR filaments and agglomerations moved slowly through sieve plates, providing an exemplary justification for our above argument: describing P-proteins visible within sieve plate pores on static micrographs asocclusions certainly is misleading, at least in Arabidopsis. Most importantly, the presence of AtSEOR proteins did not seem to inhibit phloem flow in vivo, leading Froelich et al. (2011, p. 4435) to conclude that ‘transport

172 occurs through agglomerations’. In other words, SEOR agglomerations need not always associate with an infinite hydraulic resistance in intact plants; thus the idea of their involvement in wound sealing appears questionable.

This of course leaves us with a conundrum. If, as it now seems plausible, SEOR agglomerations do not associate with infinite hydraulic resistance in intact plants, then how are we to explain the rapid cessation of label movement down the stems of plants in response to sudden chilling, drastic intracellular pH change, audio frequency vibration, and electroshock (cf. Pickard and Minchin, 1992b)? Moreover, the chilling sensitivity is very widely distributed in the dicots (Lang and Minchin, 1986).

It can hardly be doubted that SEORs and other structural P-proteins contribute to the hydraulic resistance of sieve tubes. Their physiological functions, however, still remain elusive. We think that in vivo studies of P-protein dynamics in combination with flow velocity measurements, although methodologically demanding, represent the most promising approach to overcome this scientific roadblock, especially if methodologies can be developed that enable the monitoring of continuous sieve tubes and networks.

4.4 Supplementary data

Supplementary data are available at JXB online.

Movie S1. Cold shock experiment in a root of an AtSEOR1 knockout plant. When the ice- cold water is applied, the root moves slightly and the second, unbleached phloem file enters the plane of focus. However, refocusing occurs within a second and slowing as well as halt of phloem transport can be seen. This movie corresponds to Fig. 1.

173

4.5 Acknowledgements

We are grateful for help and assistance from the Franceschi Microscopy and Imaging

Center at Washington State University, Pullman. This work was supported by NSF IOS #

1146500 and NSF IOS # 1022106, and by a Sabbatical Leave granted to WSP by

Indiana/Purdue University Fort Wayne. Any opinions, findings, and conclusions or recommendations expressed in this material are those of the author(s) and do not necessarily reflect the views of the National Science Foundation.

4.6 References

Agrawal AA, Konno K. 2009. Latex: a model for understanding mechanisms, ecology, and evolution of plant defense against herbivory. Annual Review of Ecology, Evolution and

Systematics 40, 311–331.

Anstead JA, Froelich DR, Knoblauch M, Thompson GA. 2012. Arabidopsis P-protein filament formation requires both AtSEOR1 and AtSEOR2. Plant & Cell Physiology 53, 1033–

1042.

Atkins C, Smith P, Rodriguez-Medina C. 2010. Macromolecules in phloem exudates – a review. Protoplasma 248, 165–172.

Baldwin IT. 1988. The alkaloidal response of wild tobacco to real and simulated herbivory.

Oecologia 77, 378–381.

Behnke HD. 1991. Nondispersive protein bodies in sieve elements: a survey and review of their origin, distribution and taxonomic significance. International Association of Wood

Anatomists Bulletin 12, 143–175.

174

Bricchi I, Leitner M, Foti M, Mithöfer A, Boland W, Maffei ME. 2010. Robotic mechanical wounding (MecWorm) versus herbivore-induced responses: early signaling and volatile emission in Lima bean (Phaeolus lunatus L.). Planta 232, 719–729.

Carolan JC, Fitzroy CIJ, Ashton PD, Douglas AE, Wilkinson TL. 2009. The secreted salivary proteome of the pea aphid Acyrthosiphon pisum characterised by mass spectrometry. Proteomics 9, 2457–2467.

Carolan JC, Caragea D, Reardon KT, et al. 2011. Predicted effector molecules in the salivary secretome of the pea aphid (Acyrthosiphon pisum): a dual transcriptomic/proteomic approach. Journal of Proteome Research 10, 1505–518.

Consales F, Schweizer F, Erb M, Gouhier-Darimont C, Bodenhausen N, Bruessow F,

Sobhy I, Reymond P. 2012. Insect oral secretions suppress wound-induced responses in

Arabidopsis. Journal of Experimental Botany 63, 727–737.

Cooper WR, Dillwith JW, Puterka GJ. 2011. Comparisons of salivary proteins from five aphid (Hemiptera: Aphididae) species. Environmental Entomology 40, 151–156.

Coppola V, Coppola M, Rocco M, et al. 2013. Transcriptomic and proteomic analysis of a compatible tomato-aphid interaction reveals a predominant salicylic acid-dependent plant response. BMC Genomics 14, 515.

Crafts AS. 1932. Phloem anatomy, exudation, and transport of organic nutrients in cucurbits. Plant Physiology 7, 183–225. de Vries H. 1877. Untersuchungen über die mechanischen Ursachen der Zellstreckung.

Leipzig: Engelmann.

175

Dixon AFG. 1975. Aphids and translocation. In: Zimmermannn MH, Milburn JA, eds.

Encyclopedia of plant physiology. New series, vol. 1. Transport in plants. 1. Phloem transport.

Berlin: Springer, 154–170.

Douglas AE. 2006. Phloem-sap feeding by animals: problems and solutions. Journal of

Experimental Botany 57, 747–754.

Ehlers K, Knoblauch M, van Bel AJE. 2000. Ultrastructural features of well-preserved and injured sieve elements: minute clamps keep the phloem conduits free for mass flow.

Protoplasma 214, 80–92.

Eleftheriou EP. 1990. Monocotyledons. In: Behnke HD, Sjolund RD, eds. Sieve elements.

Berlin: Springer, 139–159.

Elzinga DA, Jander G. 2013. The role of protein effectors in plant-aphid interactions.

Current Opinion in Plant Biology 16, 451–456.

Ernst AM, Jekat SB, Zielonka S, Müller B, Neumann U, Rüping B, Krzyzanek V, Prüfer

D, Noll GA. 2012. Sieve Element Occlusion (SEO) genes encode structural phloem proteins involved in wound sealing of the phloem. Proceedings of the National Academy of Science

USA 109, E1980–E1989.

Esau K. 1969.The phloem. Berlin, Gebrueder Borntraeger.

Eschrich W. 1975. Sealing systems in phloem. In: Zimmermannn MH, Milburn JA, eds.

Encyclopedia of plant physiology. New series, vol. 1. Transport in plants. 1. Phloem transport.

Berlin: Springer, 39–56.

Evert RF. 1982. Sieve-tube structure in relation to function. Bio-Science 32, 789–795.

176

Evert RF, Eschrich W, Eichhorn SE. 1971. Sieve-plate pores in leaf veins of Hordeum vulgare. Planta 100, 262–267.

Fischer A. 1884. Untersuchungen über das Siebröhren-System der Cucurbitaceen. Berlin:

Gebrüder Borntraeger.

Fischer A. 1885. Ueber den Inhalt der Siebröhren in der unverletzten Pflanze. Berichte der

Deutschen Botanischen Gesellschaft 3, 230–239.

Franceschi VR, Krokene P, Christiansen E, Krekling T. 2005. Anatomical and chemical defenses of conifer bark against bark beetles and other pests. New Phytologist 167, 353–

376.

Froelich DR, Mullendore DL, Jensen KH, Ross-Elliott TJ, Anstead JA, Thompson GA,

Pélissier HC, Knoblauch M. 2011. Phloem ultrastructure and pressure flow: sieve- element-occlusion-related agglomerations do not affect translocation. Plant Cell 23, 4428–

4445.

Furch ACU, Hafke JB, Schulz A, van Bel AJE. 2007. Ca2+-mediated remote control of reversible sieve tube occlusion in Vicia faba. Journal of Experimental Botany 58, 2827–

2838.

Furch ACU, van Bel AJE, Fricker MD, Felle HH, Fuchs M, Hafke JB. 2009. Sieve element

Ca2+ channels as relay stations between remote stimuli and sieve tube occlusion in Vicia faba. Plant Cell 21, 2118–2132.

Gaupels F, Ghirardo A. 2013. The extrafascicular phloem is made for fighting. Frontiers in

Plant Science 4, 187.

177

Gaupels F, Sarioglu H, Beckmann M, Hause B, Spannagl M, Draper J, Lindermayr C,

Durner J. 2012. Deciphering systemic wound responses of the pumpkin extrafascicular phloem by metabolomics and stable isotope-coded protein labeling. Plant Physiology 160,

2285–2299.

Goggin FL. 2007. Plant-aphid interactions: molecular and ecological perspectives. Current

Opinion in Plant Biology 10, 399–408.

Gould N, Minchin PEH, Thorpe MR. 2004 Direct measurements of sieve element hydrostatic pressure reveal strong regulation after pathway blockage. Functional Plant

Biology 31, 987–993.

Groscurth S, Müller B, Schwan S, et al. 2012. Artificial forisomes are ideal models of forisome assembly and activity that allow the development of technical devices.

Biomacromolecules 13, 3076–3086.

Haberlandt G. 1896. Physiologische Planzenanatomie. 2nd ed. Leipzig: Wilhelm Engelmann.

Hafke JB, Hafke Y, Smith JAC, Lüttge U, Thiel G. 2003. Vacuolar malate uptake is mediated by an anion-selective inward rectifier. The Plant Journal 35, 116-128.

Hanson PJ, Sucoff EI, Markhart AH. 1985. Quantifying apoplastic flux through Red Pine root systems using trisodium,3-hydroxy-5,8,10-pyrenetrisulfonate. Plant Physiology 77,

21–24.

Harmel N, Létocart E, Cherqui A, Giordanengo P, Mazzucchelli G, Guilloneau F, de

Pauw E, Haubruge E, Francis F. 2008. Identification of aphid salivary proteins: a proteomic investigation of Myzus persicae. Insect Molecular Biology 17, 165–174.

178

Hartig T. 1837. Vergleichende Untersuchungen über die Organization des Stammes der einheimischen Waldbäume. Jahresberichte über die Fortschritte der Forstwissenschaften und der Forstlichen Naturkunde 1, 125-168.

Hartig T. 1860. Beiträge zur physiologischen Forstbotanik. Allgemeine Forst- und

Jagdzeitschrift 36, 257-261

Hattori M, Nakamura M, Komatsu S, Tsuchihara K, Tamura Y, Hasegawa T. 2012.

Molecular cloning of a novel calcium-binding protein in the secreted saliva of the green leafhopper Nephotettix cincticeps. Insect Biochemistry and Molecular Biology 42, 1–9.

Hilker M, Meiners T. 2010. How do plants “notice” attack by herbivorous arthropods?

Biological Reviews 85, 267–280.

Hölttä T, Vesala T, Sevanto S, Perämäki M, Nikinmaa E. 2006. Modeling xylem and phloem water flows in trees according to cohesion theory and Münch hypothesis. Trees 20,

67–78.

Hogenhout SA, Bos JIB. 2011. Effector proteins that modulate plant-insect interactions.

Current Opinion in Plant Biology 14, 422–428.

Hose E, Clarkson DT, Steudle E, Schreiber L, Hartung W. 2001. The exodermis: avatiable apoplastic barrier. Journal of Experimental Botany 52, 2245–2264.

Huang S, Li R, Zhang Z, et al. 2009. The genome of the cucumber, Cucumis sativus L.

Nature Genetics 41, 1275–1281.

Jekat SB, Ernst AM, von Bohl A, Zielonka S, Noll GA, Prüfer D. 2013. P-proteins in

Arabidopsis are heteromeric structures involved in rapid sieve tube sealing. Frontiers in

Plant Science 4, 255.

179

Kamphuis LG, Zulak K, Gao LL, Anderson J, Singh KB. 2013. Plant-aphid interactions with a focus on legumes. Functional Plant Biology 40, 1271–1284.

Kehr J. 2006. Phloem sap proteins: their identities and potential roles in the interaction between plants and phloem-feeding insects. Journal of Experimental Botany 57, 767–774.

Kempers R, Prior DAM, van Bel AJE, Oparka KJ. 1993. Plasmodesmata between sieve element and companion cell of extrafascicular stem phloem of Cucurbita maxima permit passage of 3 kDa fluorescent probes. Plant Journal 4, 567–575.

Knoblauch M, van Bel AJE. 1998. Sieve tubes in action. Plant Cell 10, 35–50.

Knoblauch M, Peters WS. 2010. Münch, morphology, microfluidics – our structural problem with the phloem. Plant, Cell and Environment 33, 149–1452.

Knoblauch M, Peters WS. 2013. Long-distance translocation of photosynthates: a primer.

Photosynthesis Research 117, 189–196.

Knoblauch M, Peters WS, Ehlers K, van Bel AJE. 2001. Reversible calcium-regulated stopcocks in sieve tubes. Plant Cell 13, 1221–1230.

Knoblauch M, Noll GA, Müller T, Prüfer D, Schneider-Hüther I, Scharner D, van Bel

AJE, Peters WS. 2003. ATP-independent contractile proteins from plants. Nature Materials

2, 600–603.Knoblauch M, Stubenrauch M, van Bel AJE, Peters WS. 2012. Forisome performance in artificial sieve tubes. Plant, Cell and Environment 35, 1419–1427.

Korth KL, Dixon RA. 1997. Evidence for chewing insect-specific molecular events distinct from a general wound response in leaves. Plant Physiology 115, 1299–1305.

Kuśnierczyk A, Winge P, Jørstad TS, Troczyńska J, Rossiter JT, Bones AM. 2008.

Towards global understanding of plant defence against aphids – timing and dynamics of

180 early Arabidopsis defence responses to cabbage aphid (Brevicoryne brasicae) attack. Plant,

Cell and Environment 31, 1097–1115.

Lang A, Minchin PEH. 1986. Phylogenetic distribution and mechanism of translocation inhibition by chilling. Journal of Experimental Botany 37, 389–398.

Legume Phylogeny Working Group. 2013. Legume phylogeny and classification in the

21st century: progress, prospects and lessons for other species-rich clades. Taxon 62, 217–

248.

Lin M-K, Lee Y-J, Lough TJ, Phinney BS, Lucas WJ. 2009. Analysis of the pumpkin phloem proteome provides insights into angiosperm sieve tube function. Molecular & Cellular

Proteomics 8, 343-356.

Liu DD, Chao WM, Turgeon R. 2012. Transport of sucrose, not hexose, in the phloem.

Journal of Experimental Botany 63, 4315–4320.

Longinotti MP, Corti HR. 2008. Viscosity of concentrated sucrose and trehalose aqueous solutions including the supercooled regime. Journal of Physical and Chemical Reference Data

37, 1503–1515.

Lough TJ, Lucas WJ. 2006. Integrative plant biology: role of phloem long-distance macromolecular trafficking. Annual Review of Plant Biology 57, 203-232.

MacRobbie EAC. 1971. Facts and mechanisms: a comparative survey. Biological Reviews

46, 429–481.

Madhusudhan VV, Miles PW. 1998. Mobility of salivary components as a possible reason for differences in the responses of alfalfa to the spotted alfalfa aphid and pea aphid.

Entomologia Experimentalis et Applicata 86, 25–39.

181

Maffei M, Bossi S, Spiteller D, Mithöfer A, Boland W. 2004. Effects of feeding Spodoptera littoralis on lima bean leaves. I. Membrane potentials, intracellular calcium variations, oral secretions, and regurgitate components. Plant Physiology 134, 1752–1762.

Maffei ME, Mithöfer A, Boland W. 2007a. Insects feeding on plants: rapid signals and responses preceding the induction of phytochemical release. Phytochemistry 68, 2946–

2959.

Maffei ME, Mithöfer A, Boland W. 2007b. Before gene expression: early events in plant– insect interaction. Trends in Plant Science 12, 310–316.

Marentes E, Grusak MA. 1998. Mass determination of low-molecular-weight proteins in phloem sap using matrix-assisted laser desorption/ ionization time-of-flight mass spectrometry. Journal of Experimental Botany 49, 903-911.

Medina-Ortega KJ, Walker GP. 2013. Does aphid salivation affect phloem sieve element occlusion in vivo? Journal of Experimental Botany in press, Advanced Access doi:

10.1093/jxb/ert325.

Miles PW. 1999. Aphid saliva. Biological Reviews 74, 41–85.

Moon GJ, Clough BF, Peterson CA, Allaway WG. 1986. Apoplastic and symplastic pathways in Avicennia marina (Forsk.) Vierh. roots revealed by fluorescent tracer dyes.

Australian Journal of Plant Physiology 13, 637–648.

Moreno A, Garzo E, Fernandez-Mata G, Kassem M, Aranda MA, Fereres A. 2011. Aphids secrete watery saliva into plant tissues from the onset of stylet penetration. Entomologia

Experimentalis et Applicata 139, 145–153.

182

Münch E. 1926. Dynamik der Saftströmungen. Berichte der Deutschen Botanischen

Gesellschaft 44, 68–71.

Münch E. 1927. Versuche über den Saftkreislauf. Berichte der Deutschen Botanischen

Gesellschaft 45, 340–356.

Münch E. 1930. Die Stoffbewegungen in der Pflanze. Jena: Gustav Fischer.

Mutti NS, Louis J, Papan LK, et al. 2008. A protein from the salivary glands of the pea aphid, Acyrthosiphon pisum, is essential in feeding on a host plant. Proceedings of the

National Academy of Science USA 105, 9965–9969.

Nicholson SJ, Hartson SD, Puterka GJ. 2012. Proteomic analysis of secreted saliva from

Russion wheat aphid (Diurapis noxia Kurd.) biotypes that differ in virulence to wheat.

Journal of Proteomics 75, 252–2268.

Nowak H, Komor E. 2010. How aphids decide what is good for them: experiments to test aphid feeding behaviour on Tanacetum vulgare (L.) using different nitrogen regimes.

Oecologia 163, 973–984.

Pélissier HC, Peters WS, Collier R, van Bel AJE, Knoblauch M. 2008. GFP tagging of sieve element occlusion (SEO) proteins results in green fluorescent forisomes. Plant & Cell

Physiology 49, 1699–1710.

Peters WS, van Bel AJE, Knoblauch M. 2006. The geometry of the forisome–sieve element–sieve plate complex in the phloem of Vicia faba L. leaflets. Journal of Experimental

Botany 57, 3091–3098.

183

Peters WS, Knoblauch M, Warmann SA, Schnetter R, Shen AQ, Pickard WF. 2007.

Tailed forisomes of Canavalia gladiata: a new model to study Ca2+-driven protein contractility. Annals of Botany 100, 101–109.

Peters WS, Haffer D, Hanakam CB, van Bel AJE, Knoblauch M. 2010. Legume phylogeny and the evolution of a unique contractile apparatus that regulates phloem transport.

American Journal of Botany 97, 797–808.Pfeffer W. 1877. Osmotische Untersuchungen.

Studien zur Zellmechanik. Leipzig: Wilhelm Engelmann.

Pickard WF. 2008. Lactifers and secretory ducts: two other tube systems in plants. New

Phytologist 177, 877–888.

Pickard WF, Minchin PEH. 1990. The transient inhibition of phloem translocation in

Phaseolus vulgaris by abrupt temperature drops, vibration, and electric shock. Journal of

Experimental Botany 41, 1361–1369.

Pickard WF, Minchin PEH. 1992a. The electroshock-induced inhibition of phloem translocation. Journal of Experimental Botany 43, 409–417.

Pickard WF, Minchin PEH. 1992b. The nature of the short-term inhibition of stem translocation produced by abrupt stimuli. Australian Journal of Plant Physiology 19, 471–

480.

Pickard WF, Knoblauch M, Peters WS, Shen AQ. 2006. Prospective energy densities in the forisome, a new smart material. Materials Science and Engineering C 26, 471–

480.Prado E, Tjallingii WF. 1994. Aphid activities during sieve element punctures.

Entomologia Experimentalis et Applicata 72, 157–165.

184

Pyatti P, Bandani AR, Fitches E, Gatehouse JA. 2011. Protein digestion in cereal aphids

(Sitobion avenae) as a target for plant defence by endogenous proteinase inhibitors. Journal of Insect Physiology 57, 881–891.

Ranathunge K, Schreiber L. 2011. Water and solute permeabilities of Arabidopsis roots in relation to the amount and composition of aliphatic suberin. Journal of Experimental Botany

62, 1961–1974.

Rao SAK, Carolan JC, Wilkinson TL. 2013. Proteomic profiling of cereal aphid saliva reveals both ubiquitous and adaptive secreted proteins. PLOS ONE 8, e57413.

Read SM, Northcote DH. 1983. Subunit structure and interactions of the phloem proteins of Cucurbita maxima (Pumpkin). European Journal of Biochemistry 134, 561–569.

Reymond P, Weber H, Damond M, Farmer EE. 2000. Differential gene expression in response to mechanical wounding and insect feeding in Arabidopsis. Plant Cell 12, 707–719.

Rodriguez PA, Bos JIB. 2013. Toward understanding the role of aphid effectors in plant infestation. Molecular Plant-Microbe Interactions 26, 25–30.

Rüping B, Ernst AM, Jekat SB, Nordzieke S, Reineke AR, Müller B, Bornberg-Bauer E,

Prüfer D, Noll GA. 2010. Molecular and phylogenetic characterization of the sieve element occlusion gene family in Fabaceae and non-Fabaceae plants. BMC Plant Biology 10, 219.

Sabnis DD, Sabnis HM. 1995. Phloem proteins: structure, biochemistry and function. In:

Iqbal M, ed. The cambial derivatives (Encyclopedia of Plant Anatomy, Vol. 9 Part 4). Berlin:

Gebrüder Borntraeger, 271–292.

Sandström JP, Moran NA. 2001. Amino acid budgets in three aphid species using the same host plants. Physiological Entomology 26, 202–211.

185

Schobert C, Gottschalk M, Kovar D, Staiger C, Yoo BC, Lucas W. 2000. Characterization of Ricinus communis phloem profilin, RcPRO1. Plant Molecular Biology 42, 719-730.

Slewinski TL, Zhang C, Turgeon R. 2013. Structural and functional heterogeneity in phloem loading and transport. Frontiers in Plant Science 4, 244.

Schreiber L, Hartmann K, Skrabs M, Zeier J. 1999. Apoplastic barriers in roots: chemical composition of endodermal and hypodermal cell walls. Journal of Experimental Botany 50,

1267–1280.

Smith CM, Boyko EV. 2007. The molecular bases of plant resistance and defense responses to aphid feeding: current status. Entomologia Experimentalis et Applicata 122, 1–16.

Strasburger E. 1891. Ueber den Bau und die Verrichtungen der Leitungsbahnen in den

Pflanzen. Jena: Gustav Fischer.

Thompson MV. 2006. Phloem: the long and the short of it. Trends in Plant Science 11, 26–

32.

Thompson MV, Holbrook NM. 2003a. Application of a single-solute non-steady-state phloem model to the study of long-distance assimilate transport. Journal of Theoretical

Biology 220, 419–455.

Thompson MV, Holbrook NM. 2003b. Scaling phloem transport: water potential equilibrium and osmoregulatory flow. Plant, Cell and Environment 26, 1561–1577.

Thorpe MR, Furch ACU, Minchin PEH, Föller J, van Bel AJE, Hafke JB. 2010. Rapid cooling triggers forisome dispersion just before phloem flow stops. Plant, Cell &

Environment 33, 259–271.

186

Tjallingii FW. 2006. Salivary secretions by aphids interacting with proteins of phloem wound responses. Journal of Experimental Botany 57, 739–745.

Turgeon R, Oparka K. 2010. The secret phloem of pumpkins. Proceedings of the National

Academy of Science USA 107, 13201–13202.

Uhlig K, Jaeger MS, Lisdat F, Duschl C. 2008. A biohybrid microfluidic valve based on forisome protein complexes. Journal of Microelectromechanical Systems 17, 1322–1328. van Bel AJE, Hess PH. 2008. Hexoses as phloem transport sugars: the end of a dogma?

Journal of Experimental Botany 59, 261–272.

Vick JK, Nebenführ A. 2012. Putting on the breaks: regulating organelle movements in plant cells. Journal of Integrative Plant Biology 54, 868–874.

Walker GP, Medina-Ortega KJ. 2012. Penetration of faba bean sieve elements by pea aphid does not trigger forisome dispersal. Entomologia Experimentalis et Applicata 144,

326–335.

Walling LL. 2008. Avoiding effective defenses: strategies employed by phloem-feeding insects. Plant Physiology 146, 859–866.

Wardlaw IF. 1974. Phloem transport: physical chemical or impossible. Annual Review of

Plant Physiology 25, 515–539.

Will T, Tjallingii WF, Thönnessen A, van Bel AJE. 2007. Molecular sabotage of plant defense by aphid saliva. Proceedings of the National Academy of Science USA 104, 10536–

10541.

187

Will T, Hewer A, van Bel AJE. 2008. A novel perfusion system shows that aphid feeding behaviour is altered by decrease of sieve-tube pressure. Entomologia Experimentalis et

Applicata 127, 237–245.

Will T, Kornemann SR, Furch ACU, Tjallingii WF, van Bel AJE. 2009. Aphid saliva counteracts sieve-tube occlusion: a universal phenomenon? Journal of Experimental Biology

212, 3305–3312.

Will T, Furch ACU, Zimmermann MR. 2013. How phloem-feeding insects face the challenge of phloem-located defenses. Frontiers in Plant Science 4, 336.Windt CW,

Vergeldt FJ, de Jager PA, van As H. 2006. 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 29, 1715–1729.

Zhang B, Tolstikov V, Turnbull C, Hicks LM, Fiehn O. 2010. Divergent metabolome and proteome suggest functional independence of dual phloem transport systems in cucurbits.

Proceedings of the National Academy of Science USA 107, 13532–13537.

Zhang C, Yu X, Ayre BG, Turgeon R. 2012. The origin and composition of cucurbit

“phloem” exudate. Plant Physiology 158, 1873–1882.

Zimmermannn MH, Milburn JA, eds. 1975. Encyclopedia of plant physiology. New series, vol. 1. Transport in plants. 1. Phloem transport. Berlin: Springer.

Zimmermann HM, Steudle E. 1998. Apoplastic transport across young maize roots: effect of theexodermis. Planta 206, 7–19.

188