ADVANCED MICROSCOPY IMAGING OF PHLOEM PROTEINS IN ARABIDOPSIS
REVEAL INSIGHTS INTO MUNCH’S PRESSURE FLOW HYPOTHESIS
By
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
© Copyright by DANIEL ROBERT FROELICH, 2014
All Rights Reserved
© Copyright by Daniel Robert Froelich, 2014
All Rights Reserved
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.
ii
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!
iv
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.
v
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
vi
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
ix
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
x
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
xi
Figure 3: Yeast two-hybrid experiment showing that AtSEOR1 and AtSEOR2 form homobut 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 knockoutArabidopsis 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
1.1 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.
2
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.
4
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).
7
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.
9
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).
13
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 fnumber. 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).
18
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.
20
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 postpolymerization 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.
22
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).
26
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.
27
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Chapter 2 - Phloem Ultrastructure and Pressure Flow: Sieve-ElementOcclusion-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
32
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).
34
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-
36
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.
39
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).
40
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).
41
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.
43
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.
44
(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).
45
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.
46
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).
48
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 electrondense 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).
49
(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).
50
Figure 5: SEOR1 Mutant-DNA Insertion Line.
(A) A representation of the Arabidopsis gene At3g01680 indicating the location of the TDNA 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/cgibin/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.
51
(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).
55
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).
56
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 crosssection 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
59
≈ 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
60
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
At SEOR1 filament radius
1.2 μm
10 nm
af
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
61
Sieve tube diameter
3.0 µm
Observed flow speed
100 µm s-1
At SEOR1 filament separation
6 nm
b
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
62
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
-3]
s
]
[Pa
[Pa
[Pa
[Pa
s/ ]
s/ ]
s/ ]
s/ ]
[Pa
s
]
63
[
]
(†)
(†)
(†)
(†)
(†)
(†)
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
65
(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).
66
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?
72
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
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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.
74
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
76
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 microROCs.
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 3frames 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.
79
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’-TTCGGATCCACTAGTTCTGCTGCTGGTTCTGCTGCTGGTTCTGGGGGATCCCTT-3’ and 5’-AAGGGATCCCCCAGAACCAGCAGCAGAACCAGCAGCAGAACTAGTGGATCCGAA-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-
80
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.
81
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’-GAGAGAGACCTTTTCCCTTAACCTCA-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 GABIKAT 609F04 were cut with a vibrating microtome (Vibratome) at 50 μm and collected in
82
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.
83
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.
84
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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.
92
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
Rlumen 
(Bruus, 2008)
8nLt
ae4
(A3)
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
93
(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.
94
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
of solid material
depends on the volume fraction
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 nondimensional permeability K is given by
(
)
95
(A9)
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.
96
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.
98
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.
99
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 nonmembrane 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.
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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, GFPAtSEOR2 was expressed). Complementation mutants were also generated where GFPtagged 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 Atseor21 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 GFPAtSEOR2 (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 selfinteractions 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 homobut 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 wildtype 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 ttest, 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
d-1)
1.73 ±
0.13
1.64 ± 0.13 (P =
0.63)
1.75 ± 0.13 (P =
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
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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
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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 MtSEO13, 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 GFPfusion 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
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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
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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
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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
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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 TDNA 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
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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’
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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 500555nm 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
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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.
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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.
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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.
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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
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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 laserscanning microscopy (CLSM) of living plants in microscopy rhizosphere chambers (MicroROCs), 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
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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 Pproteins, we will critically discuss current ideas about the function of these phloem
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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 ATPindependent 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
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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.
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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
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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 redesigned 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),
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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
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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
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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
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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
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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 salivasecreting 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,
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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
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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,
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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 longdistance 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
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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.
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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)
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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)
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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
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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
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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-
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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
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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
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concentration in the exudate from SEOR knock-out leaves. Since the amount of phloemderived 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.
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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
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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
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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.
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Fig. 1: Abrupt cold causes stoppage of phloem translocation in the roots of AtSEOR1 knockout 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
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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 Pproteins 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
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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-
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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
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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 coworkers 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.
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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 sapfeeding 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
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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 Pproteins 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
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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 Pproteins 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
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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
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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 preparationinduced 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 icecold 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.
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