Download the fine-tuning of the endomembrane system

Intracellular Accommodation of
Rhizobia in Legume Host Cell: the Finetuning of the Endomembrane System
Aleksandr Gavrin
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Thesis committee
Promotor
Prof. Dr T. Bisseling
Professor of Molecular Biology
Wageningen University
Co-promotor
Dr E. Fedorova
Researcher, Laboratory of Molecular Biology
Wageningen University
Other members
Prof. Dr H.J. Bouwmeester, Wageningen University
Prof. Dr F.P.M. Govers, Wageningen University
Dr M.J. Ketelaar, Wageningen University
Prof. Dr S.C. de Vries, Wageningen University
This research was conducted under the auspices of the graduate School of
Experimental Plant Sciences
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Intracellular accommodation of rhizobia in
legume host cell: the fine-tuning of the
endomembrane system
Aleksandr Gavrin
Thesis
submitted in fulfillment of the requirements for the degree of doctor
at Wageningen University
by the authority of Rector Magnificus
Prof. Dr A.P.J. Mol,
in the presence of the
Thesis Committee appointed by the Academic Board
to be defended in public
on Wednesday 2 September 2015
at 11 a.m. in the Aula.
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Aleksandr Gavrin
Intracellular accommodation of rhizobia in legume host cell: the fine-tuning of the
endomembrane system
160 pages.
PhD thesis, Wageningen University, Wageningen, NL (2015)
With references, with summaries in Dutch and English
ISBN 978-94-6257-418-2
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Contents
OUTLINE7
CHAPTER 1
General introduction
11
CHAPTER 2
VAMP721a and VAMP721d are essential for pectin dynamics during
release of bacteria in soybean nodules
33
CHAPTER 3
Adjustment of host cells for accommodation of symbiotic bacteria:
vacuole defunctionalization, HOPS suppression and TIP1g
retargeting in Medicago59
CHAPTER 4
ARP2/3-mediated actin nucleation associated
with symbiosome membraneis essential for the development
of symbiosomes in infected cells of Medicago truncatula root nodules
95
CHAPTER 5
The establishment of symbiosis: the role of Synaptotagmin 1
119
CHAPTER 6
General Discussion: The adaptation of the endomembrane system
of host cell to intracellular bacteria
139
Summary152
Sumenvatting (summary in Dutch)
154
Acknowledgements156
157
List of publications
158
Curriculum Vitae
Education statement
159
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OUTLINE
Plants are non-motile organisms. Their sessile mode of life restricts access to vital
elements like nitrogen and phosphorus, which are scares in many types of soils. To
get better access to such nutrients plants evolved endosymbiotic interactions with
soil fungi and bacteria. Members of the legume family are able to establish an N2fixing endosymbiosis with certain soil bacteria, collectively called rhizobium. This
involves the formation of a new organ, the root nodule. These root nodules have a
central tissue containing infected cells. Each of the infected cells contains thousands
of rhizobium bacteria. Intracellular rhizobia are surrounded by a membrane of the
host cell and these organelle-like structures are named symbiosomes. Symbiosome
membrane forms a unique symbiotic interface that facilitates exchange of nutrients
between micro- and macrosymbiont. The symbiosome membrane is made by the host
cell and belongs to the plant cell endomembrane system. Therefore, the intracellular
accommodation of rhizobia causes a reorganization of the endomembrane system of
these cells.
In Chapter 1, I give a general introduction and describe the formation of two types
of nodules, which have an indeterminate or determinate type of growth, respectively.
I review the recent studies concerning signaling pathways that control root nodule
development and mechanisms that control the formation of symbiotic interfaces. I
address the recent data concerning adaptation of host cell endomembrane system to
intracellular bacteria and describe membrane proteins of symbiotic interfaces.
In Chapter 2 we study the role of a symbiosis specific exocytotic pathway. This
pathway involves vesicle-associated membrane proteins 721d/e (VAMP721d/e).
Rhizobia enter nodule cells by cell wall bound infection threads and release into the
host cell cytoplasm via cell wall–free regions of infection threads, so called unwalled
droplets. Previously, it was shown that of VAMP721d/e is essential for the formation
of the symbiotic interface in both rhizobial and arbuscular mycorrhizal symbiosis
in Medicago truncatula, however the underlying molecular mechanisms were not
clarified.
We have found that in soybean nodules the silencing of GmVAMP721d blocks the
release of rhizobia and the formation of the symbiosomes. Instead of symbiosomes
the infected cells contained big clusters of bacteria embedded in a matrix of methylesterified and de-methyl-esterified pectin. We have analysed the role of GmVAMP721d
containing vesicles in the transport of pectin and found that these vesicles were
not transporting methyl-esterified pectin. We hypothesized that they may deliver
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enzymes involved in pectin turnover. Subsequently, we found that GmVAMP721d is
partly co-localized with pectate lyase. Therefore the GmVAMP721d may deliver pectin
modifying enzymes to the sites of rhizobial release.
In Chapter 3 of this thesis, we show that vacuole modification plays a crucial role
in the development of infected cells. The maintenance of symbiosomes requires a
major adjustment of the pathway controlling vacuole formation as well as tonoplasttargeted trafficking. We show that the expression of VPS11 and VPS39, two members
of vacuolar tethering complex HOPS, are suppressed in infected cells. This occurs when
the infected cells become mature N2-fixing cells. The vacuoles of the infected cells
contract at this stage of cell development. This contraction permits the expansion
of symbiosomes. Trafficking of tonoplast-targeted proteins in infected cells is
also altered, as shown by retargeting of the aquaporin TIP1g from the tonoplast
membrane to the symbiosome membrane. This retargeting appears to be essential
for the maturation of symbiosomes. Therefore, these alterations in the functionality
of the vacuole are key events in the adaptation of the plant cell to host intracellular
symbiotic bacteria.
In Chapter 4, we have studied the reorganization of the plant cell cytoskeleton
during intracellular accommodation of rhizobia in root nodules of M. truncatula. We
focused on the role of actin networks and nucleating factor ARP3. We have shown
that the actin configuration in infected cells changes markedly during symbiosome
development. When symbiosomes mature, they become surrounded by a dense actin
network. The ARP2/3 complex is operational in the rearrangement of actin around
symbiosomes. We believe that the retargeting of late endosome/tonoplast proteins
to the symbiosome membranes allows the recruitment of actin to the symbiosomes
and that this is necessary for their functional maturation.
Chapter 5 addresses the putative regulatory mechanisms of membrane fusion with
the symbiosome membrane. During the formation of infected cell new protrusions
of the plasma membrane (infection threads, cell wall-free unwalled droplets and
symbiosomes) are formed. These separate the bacteria from the host cytoplasm. The
symbiosomes rapidly increase in number and volume due to the expanding bacteria
and may represent sites with increased membrane tension. We hypothesized that
this membrane tension may create a vector for vesicle fusion to these membranes.
To test this hypothesis, we studied MtSyt2 and MtSyt3, two M. truncatula homologs of
Arabidopsis thaliana synaptotagmin 1 (SYT1) proteins. AtSYT1 is involved in membrane
repair in case of fluctuation of membrane tension or membrane damage. Two M.
truncatula homologs of AtSYT1, MtSyt2 and MtSyt3, are expressed in nodule primordia
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and in the apical part of root nodules. The proteins are localized in plasma membrane
regions of meristematic and elongating cells, showing that they are involved in
housekeeping membrane fusion. However, in the infected cells MtSyt2 and especially
MtSyt3 are very abundant on infection threads and unwalled droplets. The double
silencing of MtSyt2 and MtSyt3 shows that they are operational during growth of
infected cells and intracellular accommodation of rhizobia. The localization of MtSyt2
and MtSyt3 at the site of bacterial release, around infection threads and in the plasma
membrane of growing cells supports the hypothesis that this calcium sensor may be
involved in the creation of a vector for targeted exocytosis in young infected cells.
In Chapter 6 (General discussion) I discuss the results obtained in this thesis
and integrate it with related literature. It focusses on molecular mechanisms that
control the rearrangement of the endomembrane system in infected cells during
accommodation of microsymbionts.
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Chapter 1
General Introduction
Aleksandr Gavrin
Laboratory of Molecular Biology, Graduate School
Wageningen University, Droevendaalsesteeg 1, 6708PB
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Experimental Plant Science,
Wageningen, The Netherlands
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Chapter 1
Nitrogen is often a limiting element for plant growth. Though nitrogen is one
of the most abundant elements in the Earth’s atmosphere, plants can only utilize
reduced forms of it, for example ammonia and nitrate. Biological nitrogen fixation
is an ancient and ecologically friendly nitrogen source for plants. This is carried out
by some groups of prokaryotes, which enzymatically convert atmospheric nitrogen to
ammonia (Wagner, 2012). These prokaryotes include different free-living soil bacteria,
such as Azotobacter, and bacteria that form associative relationships with plants, such
as Azospirillum, and most importantly, bacteria, collectively known as Rhizobia, that
establish a nodule symbioses with legumes (Olivares et al., 2013). Rhizobium-legume
symbiosis is a striking example of an endosymbiotic interaction, which permits the
accommodation of bacteria in host cells of a newly formed organ, a root nodule. In last
20 years the research in the area of root nodule symbiosis were centred on signalling
between symbionts during the initiation of symbiotic relations and triggering the
nodule organogenesis. The analysis of the mutants defected in nodulation helps to
specify the genes involved in Nod factors signalling pathway (Geurts & Bisseling,
2002; Cooper, 2007; Jones et al., 2007; Oldroyd et al., 2011; Bapaume & Reinhardt,
2012; Oldroyd, 2013). However the study of symbiosis is not limited to the tools of
genetics. Symbiosis is based on the fine-tuning of host cell and bacteria that ensure
the co-existence and mutual benefit. Such processes involve the reorganization of
endomembrane system in infected cells that have to be studied by the methods of
plant biology and molecular biology. The morphology and the development of root
nodules are very well studied and reviewed (Day et al., 2001; Brewin, 2004; Kondorosi
et al., 2013; Udvardi & Poole, 2013; Xiao et al., 2014), so I will describe it only briefly. In
this introduction, I focus more on the mechanisms by which symbiotic relationships
are formed and address some of the recent insights obtained in model and crop
legumes.
Morphology of legume root nodules, a newly formed symbiotic organ
Nodules are formed by a new meristem, which is initiated by the dedifferentiation
of root cortical cells. The spatial pattern of meristem initiation and persistence
determines the main differences between the two major types of nodule (Hirsch,
1992) (Figure 1A). The nodules with so-called indeterminate type of growth have the
active meristem at the nodule apex and develop as elongated cylindrical structures.
The nodules of this type are mainly elicited on temperate legumes such as Pisum
sativum (pea), Trifolium pratense (clover) and Medicago truncatula (Medicago) (Hirsch,
1992; Brewin, 2004; Patriarca et al., 2004). Newly produced by the meristem cells are
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General introduction
shifted downwards and get infected thus creating a gradient of cell differentiation
between the meristem and the older cell layers (Xiao et al., 2014). Rhizobia enter and
start to populate host cells symplast in a so-called zone of infection. Infected cells
undergo polyploidization, enlarge and differentiate into fully matured symbiotic
cells (Vinardell et al., 2003)2003. The symbiosomes in indeterminate nodules contain
normally one, rarely two bacteria. The symbiosome differentiation into nitrogen-fixing
stage is accompanied by a significant increase in size due to the growth of bacteria
(bacteroids) inside of symbiosome (Vasse et al., 1990; Gavrin et al., 2014; Chapter 3).
Mature symbiotic cells are partly hypoxic that permits to maintain the environment,
which is favourable for the expression and functioning of rhizobia enzyme nitrogenase
and the fixation of atmospheric nitrogen (Ferguson et al., 2010). The zone, composed
of mature symbiotic cells, is defined as a zone of nitrogen fixation. In most distant
cell layers, in the basal part of the nodule, the nitrogen fixation is ceases and infected
cells get lysed. This zone is defined as a zone of senescence (Van de Velde et al., 2006;
Pérez Guerra et al., 2010). In our opinion it would be more appropriate to name this
last zone “zone of symbiosis termination”, as other nodule tissues as well as cell of the
roots from which nodule is derived are not undergoing senescence simultaneously
with infected cells.
The nodules with determinate type of growth are typical for tropical legumessuch
as Lotus japonicus (Lotus), Glycine max (soybean) or Phaseolus vulgaris (common bean).
These nodules have a spherical form. The meristem is active in primordia and in the
young nodules. The meristematic locuses, composed by several cells, are situated
laterally and remains active approximately for 2-3 weeks. Accordingly, nodule growth
takes place by cell expansion rather than by cell division and shows the spatial
differentiation pattern only in the young nodules (2-3 weeks). More mature nodule
contains a relatively homogenous population of infected cells without clearly visible
zonation (Franssen et al., 1992; Crespi & Gálvez, 2000; Brewin, 2004; Ferguson et al.,
2010; Gossmann et al., 2012). The oldest zone where the symbiosis is terminated is
situated in the central part of determinate nodules.
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Chapter 1
Figure 1. Nodule structure of determinate (A) and indeterminate (B) nodule types. NF, nitrogen fixation tissue; S,
senescence zone; M, meristem; II, infection zone; III fixation zone (adopted from http://commons.wikimedia.
org/wiki/User:Ninjatacoshell).
Initial signal exchange and Nod factor signaling
The establishment of a functional symbiosis requires the coordinated expression
of numerous bacterial and plant genes. Many studies showed that this coordination
of gene expression and other responses is mediated by a sequential exchange of
molecular signals (Cooper, 2007; Jones et al., 2007). The legume-rhizobium interactions
are in general set in motion by the secretion of 2-phenyl-1,4-benzopyrone derivatives
commonly known as flavonoids (Veitch, 2007), which are potent activators of the
transcription of rhizobial nodulation (nod) genes. Proteins encoded by the nod genes
are involved in the synthesis and secretion of Nod factors (NFs) (Geurts & Bisseling,
2002). NFs are lipo-chitooligosaccharides that generally consist of a β-1,4-linked
N-acetyl glucosamine backbone of 4 or 5 residues of which the non-reducing terminal
residue contains an acyl chain (Truchet et al., 1991). Depending on the rhizobial species,
different acyl chains and specific decorations at the reducing and non-reducing
terminal residues can be present. This are major determinants of host specificity the
biological activity of the NFs (Ardourel et al., 1994)1994.
Initial perception of NFs occurs at the epidermis, but NF signaling continues to
be important as the bacteria invade the root cortex (Capoen et al., 2005; Den Herder
et al., 2007), and for bacterial release into cells of the nodule (Limpens et al., 2005;
Heckmann et al., 2006; Moling et al., 2014). NFs are perceived by receptor-like kinases
with N-acetylglucosamine–binding lysin motifs (LysM) in the extracellular domain
such as NFP/LYK3/LYK4 in Medicago and Nod factor receptor NFR5/NFR1 in Lotus
(Limpens et al., 2003; Madsen et al., 2003). These receptors trigger ion fluxes leading
to a depolarization of the plasma membrane and calcium oscillations in and around
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General introduction
the nucleus (Oldroyd & Downie, 2008; Capoen et al., 2011). The activation of this
calcium spiking response also requires a plasma membrane LRR receptor kinase
DMI2/SYMRK, which forms a complex with NFR5 and might acts as a co-receptor
(Ané et al., 2002; Endre et al., 2002; Antolín-Llovera et al., 2014). Though, mechanisms
that can connect ligand perception at the plasma membrane with calcium changes in
the nucleus remain unclear. Probably, the receptors activate secondary messengers
that can regulate DMI1 ion channels. Pharmacological approach and measurements
of phospholipids have indicated the importance of phospholipase C (PLC) and
phospholipase D (PLD) in NF signaling (Charron et al., 2004), but there is no genetic
evidence supporting this. By contrast, interaction studies have shown that DMI2/
SYMRK associate with 3-hydroxy-3methylglutaryl-CoA reductase (HMGR). HMGRs
regulate the mevalonate biosynthetic pathway leading to the production of different
isoprenoid compounds, precursor to many secondary products. Recruitment of
HMGR1 by DMI2/SYMRK could be required for production of specific isoprenoid
compounds that could provide the signaling between the plasma membrane–
localized NF receptors and the nuclear-localized proteins (Kevei et al., 2007).
NF induced calcium changes occur in the nucleoplasm and the perinuclear
plasm of root hairs. The nature of the calcium channels remains unknown, but a
nuclear pore complex (Groth et al., 2010)2010 and a putative cation channel (DMI1
in Medicago, Castor and Pollux in Lotus) have been identified as essential for NF
induced calcium oscillation (Edwards et al., 2007; Charpentier et al., 2008). These
channels are preferentially permeable to potassium and located on the inner nuclear
membrane. Furthermore, a calcium ATPase MCA8 essential for symbiotic calcium
oscillations is also targeted to the inner nuclear membrane and can pump the
calcium back into the nuclear envelope (Capoen et al., 2011). Mathematical model
based on these components and a presumed voltage-activated calcium channel
describes a mechanism of calcium spiking where DMI1 regulates the calcium channel
by driving depolarization of the membrane through regulating potassium flow, and
it compensates the charge produced by the movement of calcium (Granqvist et al.,
2012).
Calcium spiking signal is decoded by a nuclear calcium/calmodulin-dependent
kinase CCaMK (Tirichine et al., 2006). CCaMK phosphorylates CYCLOPS, a DNAbinding transcriptional activator of the NODULE INCEPTION (NIN) gene (Limpens &
Bisseling, 2014; Singh et al., 2014). NIN is a nodulation-specific gene that encodes a
putative transcription factor, which regulates NF-YA1 and NF-YB1 expression in Lotus.
Two genes encode different subunits of a NF-Y CCAAT box binding protein complex.
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Chapter 1
NF-Y complex triggers entry of the cell into division cycle (Laloum et al., 2013).
Therefore, activation of the NF-Y subunit genes induces cortical cell division, which is
an initial step in root nodule organogenesis (Soyano et al., 2013).
Nodulation signalling pathway 1 (NSP1) and NSP2 are GRAS domain transcription
factors. NSP1 and NSP2 form a heterocomplex that associates with the promoters
of Nod factor-inducible genes (Hirsch et al., 2009). Previously, it was thought that
this transcription complex promotes the expression of NIN gene (Oldroyd, 2013).
However, findings of Singh et al. (2014) shows that CYCLOPS induces NIN expression
in an NSPs independent way. It remains unclear how the action of NSP1 and NSP2
is associated with the NF signaling. Perhaps NSPs acts downstream of NIN or in an
alternative pathway (Limpens & Bisseling, 2014).
Several studies indicate that rhizobia continue to produce NFs inside root nodules
(Marie et al., 1994; Schlaman et al., 1998) and components of NF signaling pathway are
also expressed in nodule cells, especially in the infection zone of the nodule (Bersoult
et al., 2005; Capoen et al., 2005; Limpens et al., 2005; Riely et al., 2006). Silencing of
them, driven by nodule specific promoters, does not affect the nodulation but leads
to aberration of microsymbiont differentiation. For instance, knockdown of SymRK/
DMI2 by RNA interference causes a block of the release of rhizobia into host cells
while nodule formation and infection thread growth still occurred (Capoen et al.,
2005; Limpens et al., 2005). Specific silencing of NFP in the infection zone blocks
rhizobial release in cells derived from the meristem (Moling et al., 2014). Functional
characterization of IPD3 and DMI3 genes also clearly demonstrates an essential role
of the NF signalling pathway in intracellular accommodation of bacteria (Morzhina
et al., 2000; Voroshilova et al., 2009; Ovchinnikova et al., 2011). These data as well as
the upregulation of several NF signalling components in the nodule suggest that
NF signalling is required to allow bacterial release and symbiosome development
(Kereszt et al., 2011).
Nodule organogenesis and the infection
The interaction of rhizobia and legumes results in the formation of root nodules.
The formation of this organ involves the coordinated processes. On one hand, the
mitotic reactivation of differentiated root cells (Xiao et al., 2014) from which a nodule
primordium is formed and second the infection process. Rhizobia penetrate into
nodule cells via special tubular structures called infection threads (ITs) or by “crack
entry” from the colonies situated in intercellular spaces between epidermis/cortical
cells. ITs are transcellular, cell wall-bound tubular structures, covered by membrane
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General introduction
and populated by rhizobia embedded in a luminal matrix (VandenBosch et al., 1989;
Rae et al., 1992; Brewin, 2004). It is worth to mention that in relatively primitive
legumes, as Caesalpinioideae, bacteria are not released from ITs but retained in
modified ITs, known as fixation threads (Sprent & Embrapa, 1980; Masson-Boivin et
al., 2009; Op den Camp et al., 2012).
The IT mediated colonization of legume roots is probably the best studied of all
the rhizobial infection processes, requires host-specific NFs (Gage, 2004; Den Herder
et al., 2007). In general, ITs originate in root hairs that curl around attached bacteria
through a continued reorientation of their growth direction by which rhizobia become
entrapped in a closed cavity and form a microcolony (Jones et al., 2007). Inside the
curl, the plant cell wall is locally degraded and new membrane and cell wall material
is deposited forming a tubular IT. Bacteria growing within ITs remain topologically
outside of root hair cells, and are separated from the plant cytoplasm by the IT wall and
membrane. IT grows through the root hair into the apoplast of cortical cells. The path
of ITs is determined by formation of so-called pre-infection threads. Activated cortical
cells form phragmoplast-like cytoplasmic bridges that traverse the central vacuole
and facilitate passing of ITs through cortical cell layers (Yang et al., 1994; Timmers et
al., 1998). In this way ITs reach nodule primordium cells, the precursors of nodule cells,
which formed through a series of reprogrammed cortical cells divisions (Oldroyd et al.,
2011). Branching of ITs as it grows through the root and enters the nodule primordium
increases the number of sites from which bacteria can enter primordium cells. Then,
bacteria are released into developing cells. In determinate nodules, release takes
place only at the primordium stage. Release of bacteria in indeterminate nodules is
restricted to a very specific area, just 1-2 cell layers bellow the meristem, and does
not occur anymore in next cell layers despite the presence of the network of infection
threads through all nodule zones (Vasse et al., 1990; Monahan-Giovanelli et al., 2006).
During “crack entry”, rhizobia take advantage of local disruptions in the epidermal
cell layers, for example during lateral root formation. Bacteria enter, multiply and
form extracellular infection pockets, an example is nodule formation in Sesbania and
Aeschynomene (Den Herder et al., 2007). Infection of the nodule primordium cells
occurs by a direct uptake from these infection pockets (Chandler et al., 1982; Boogerd
& van Rossum, 1997). Colonization of Sesbania nodules requires NFs, whereas the
infection of Aeschynomene plants is a NF-independent process (D’Haeze et al., 2003;
Goormachtig et al., 2004; Giraud et al., 2007).
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Chapter 1
The release of rhizobia into the host cell and symbiosomes development
Rhizobia are released into the host cells by a process resembling endocytosis,
but which is controlled by a specific exocytotic pathway (Figure 2). Bacterial release
is mediated by formation of unwalled infection droplets, regions of ITs with locally
degraded cell wall and a slightly outbulging membrane (Figure 2). Creation of such
a cell-wall free interface allows rhizobia to come into close contact with the host
membrane, and be engulfed into the cytoplasm surrounded by a host membrane. As
a result, rhizobial cells becomes a cellular compartments defined as the symbiosomes
- bacteroids enclosed by a plant-derived symbiosome membrane (Brewin et al., 1994;
Bolaños et al., 2004). After bacteria have been released, they divide, differentiate and
fill the cells.
Figure 2. Schematic representation of bacterial release. Symbiosome formation starts with the formation of
unwalled infection droplet, where bacteria come into close contact with the host plasma membrane.
Subsequently bacteria are individually “pinched off” into the cytoplasm and become surrounded by symbiosome
membrane. IT, infection thread; UD, unwalled infection droplet; B, bacteroid; V, vacuole; N, nucleus; ER,
endoplasmic reticulum; G, Golgi complex.
In determinate nodules division and fusion of young single-bacteroid
symbiosomes form symbiosomes with multiple bacteroids (Fedorova et al., 1999;
Brewin, 2004). Bacteroids are morphologically not very different from newly released
rod-shaped bacteria.
Differentiation of symbiosomes is significantly diverse in indeterminate
and determinate nodules. In indeterminate nodules, division of bacteroids and
symbiosomes is strictly coupled and leads to the formation of symbiosomes with
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General introduction
a single-bacteroid. After the release young symbiosomes of developmental stage 1
(Vasse et al., 1990) are able to divide, at this stage bacteroids are similar in size to freeliving bacteria. After several rounds of division, bacteroids gradually differentiate into
mature bacteroids capable of reducing atmospheric nitrogen. They become much
larger, radially arranged and completely fill the cytoplasm of infected cells in the
fixation zone (Vasse et al., 1990; Timmers et al., 1998).
Differentiation of bacteroids, in indeterminate nodules, is an irreversible
process. At the endpoint of it the bacteroids lose the ability to divide. This stage is
called terminal differentiation. Terminal differentiation depends on the host plant
(Mergaert et al., 2006) and triggered by a family of nodule-specific cysteine-rich
(NCR) peptides, which resemble defensin-like antimicrobial peptides (Van de Velde
et al., 2010; Maróti et al., 2011). In Medicago, they represent a huge gene family, most
of which are expressed specifically in infected cells (Mergaert et al., 2003). Targeting
of NCR peptides in infected cells is provided by a nodule-specific protein secretory
pathway. Specific signal peptidase SPC22 cleaves off the NCR signal peptide and
controls targeting of NCR peptides to symbiosomes (Wang et al., 2010)2010.
Despite knowledge of the critical role of Medicago NCR in bacteroid development,
the exact bacterial targets are unknown. BacA protein was discovered to be essential
for bacteroid development within Medicago legumes (Glazebrook et al., 1993). BacA is
critical to reduce the level of NCR-induced membrane permeabilization and bacterial
killing in vitro. Therefore, BacA protein is essential for protection of Sinorhizobium
meliloti against the antimicrobial activities of host NCR peptides (Haag et al., 2011)
changing the host cell defense reaction. There is a growing realization now that one
of the crucial components in the establishment and maintenance of symbiosis is a
modulation of plant innate immunity processes (Bourcy et al., 2013; Berrabah et al.,
2014).
Though terminal differentiation is mainly a characteristic of indeterminate
nodules; it does not really depend on the nodule type. For example, several
mimosoid (Mimosoideae) legumes form indeterminate nodules with non-terminal
differentiated symbiosomes (Ishihara et al., 2011; Kereszt et al., 2011; Marchetti et
al., 2011). Instead, terminal differentiation of bacteroids seems to be a characteristic
for legumes of the so-called inverted-repeat-lacking (IRLC) clade of Papillionoideae
(Wojciechowski et al., 2004; Mergaert et al., 2006). The NCR peptide genes are
specific for the IRLC legumes and do not occur in the genomes of sequenced non-IRLC
legumes such as soybean and Lotus (Alunni et al., 2007). The distribution of terminal
bacteroid differentiation in the legume family indicates that the non-terminal
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Chapter 1
differentiated state is ancestral, and terminal differentiation likely evolved several
times independently in the Papilionoidea (Oono et al., 2010).
Intracellular lifestyle of rhizobia
Specialized symbiotic (infected) cells of root nodules contain thousands of living
bacteria. These bacteria are kept in host cell cytoplasm separated by the individual
membrane provided by the host (Oldroyd & Downie, 2008; Kereszt et al., 2011;
Kondorosi et al., 2013). Morphologically symbiosomes have something in common
with vacuoles housing pathogen microbes of mammals (Brumell & Scidmore, 2007;
Isberg et al., 2009; von Bargen et al., 2009). Though, only legumes and insects have
specialized cells, which are permissive for a long maintenance of intracellular bacteria.
After bacterial release, symbiosomes become part of the host cell endomembrane
system. Eukaryotic cells have internal membrane organelles. Vesicle transport
between them is highly dependent on the identity of the membranes. Main two
groups of the proteins, which mediate the membrane identity, are regulatory small
GTPases of the Rab family and soluble N-ethylmaleimide sensitive factor attachment
protein receptors (SNAREs). Rab GTPases control the transport and docking of
vesicles to their target membrane compartment. Subsequently, a vesicle-associated
SNARE (v-SNARE) proteins forms a complex with cognate SNARE proteins in the
target compartment (t-SNARE) which drives the membrane fusion (Sanderfoot et al.,
2000; Sanderfoot et al., 2001; Pfeffer & Aivazian, 2004; Behnia & Munro, 2005; Lipka
et al., 2007; Pfeffer, 2007; Bassham & Blatt, 2008; Nielsen et al., 2008).
Release of bacteria and symbiosome formation involves a major reorganization
of the host endomembrane system and massive production of new membrane
material to accommodate bacteria in symbiosome compartments. It is controlled by a
symbiosis specific exocytotic process, involving specific vesicle-associated membrane
proteins VAMP72s (Ivanov et al., 2012). Further, another member of the same
family, VAMP721a, was recently shown to be a target of regulator of symbiosome
differentiation (RSD). RSD is a member of the Cysteine-2/Histidine-2 (C2H2) family
of plant transcription factors and is required for normal symbiosome differentiation
of M. truncatula nodules (Sinharoy et al., 2013). Moreover, plasma membrane t-SNARE
SYP132 is present on symbiosome membranes from the start of symbiosome
formation throughout its development (Catalano et al., 2007; Limpens et al., 2009).
For instance, above described NCR peptides may also be transported by specific
exocytotic vesicles to differentiating symbiosomes (Wang et al., 2010)2010.
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General introduction
It has been generally thought that rhizobia enter host cells via an endocytosislike process similar to the phagocytic uptake of bacteria in animal cells (Knodler et
al., 2002; Brumell & Grinstein, 2004). However, these suggestions were merely based
on morphological observations. Recent progress in membrane biology allows now
the use of membrane identity markers and shows that symbiosomes do not accept
the molecular markers of early endosome, a trans-Golgi marker SYP4 or the late
endosome/prevacuolar marker Rab5, that means that symbiosomes are not following
the default endocytotic pathway. The late endosomal/vacuolar marker Rab7, however,
is recruited to symbiosome membrane at later stages of development and is required
for symbiosomes maturation. During the senescence, which may be also called the
termination of symbiosis, the symbiosomes start to fuse and form lytic vacuole-like
compartments; at this stage proteins of vacuolar SNAREs complex are accumulated
on the symbiosome membranes (Limpens et al., 2009).
Therefore, mature symbiosomes appear to have a unique mosaic identity,
combining plasma membrane and late endosomal markers. This dual character of
symbiosome membranes suggests that symbiosomes are able to intercept plasma
membrane- and vacuole-targeted vesicle traffic. This could explain presence of
several plasma membrane and vacuolar transporters on symbiosome membrane
(Whitehead & Day, 1997; Guenther & Roberts, 2000; Wienkoop & Saalbach, 2003;
Jones et al., 2007). Furthermore, delayed acquisition of complete vacuolar identity
by symbiosomes seems a regulatory mechanism of intracellular maintenance of
microsymbiont in the host cell (Limpens et al., 2009).
1
Membrane transporters of symbiotic interfaces
At the “heart” of symbiosis is an exchange of nutrients for the purpose of mutual
benefits for both symbionts. In case of Rhizobium–legumes symbiosis, plants receive
fixed nitrogen in the form of ammonium and provide rhizobia with whatever is
required for nitrogen fixation in return. Symbiosome membrane is operational in
this exchange. Proteomic and genomic studies have shown presence of a range of
membrane transporters on symbiosome membrane (Panter et al., 2000; Wienkoop
& Saalbach, 2003; Catalano et al., 2004; Colebatch et al., 2004; Høgslund et al., 2009;
Benedito et al., 2010; Clarke et al., 2015).
Intensive investigation of symbiosome membrane permeability to various
metabolites began with appearance of methods of symbiosomes isolation from
the nodules (Price et al., 1987). It was found that symbiosomes from soybean root
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Chapter 1
nodules possesses a dicarboxylate transporter capable of mediating a rapid flux of
dicarboxylate anions, such as succinate and especially malate, to the bacteroids inside
the nodule (Udvardi et al., 1988; Day et al., 1989; Ou Yang et al., 1990). The same approach
revealed that symbiosome membrane is weakly permeable for sugars and apparently
lacks carriers for these compounds (Udvardi et al., 1990). Another strong evidence
that dicarboxylic acids are the primary source of carbon provided by the plant for
nitrogen fixing bacteroids, comes from bacterial genetic investigations. Bacteroid’s
genes for the transport and catabolism of dicarboxylic acids are strongly expressed
and indispensable for symbiosis (Terpolilli et al., 2012). However, dicarboxylate
transporters of symbiosome membrane have not been found.
Metabolism of bacteroids, besides carbon, also requires various elements. For
instance, bacterial nitrogenase is a sophisticated enzymatic complex, which needs
molybdenum, iron and sulfur-containing cofactors. All these elements are essential
for symbiosis and supposed to be provided by the host cell. It requires appropriate
transporters on the symbiosome membrane. Sulfate from the plant cell cytoplasm to
the bacteroids in Lotus nodules is transported by SST1 (Krusell et al., 2005). Expression
of LjSEN1, the member of vacuolar iron/manganese transporter family, was detected
exclusively in nodule-infected cells and may have a role in the transport of iron to
bacteroids (Hakoyama et al., 2012)2012. A second iron transporter of symbiosome
membrane is a soybean divalent metal transporter (GmDmt1). GmDmt1 is a noduleenhanced transporter capable of iron and zinc transport across the symbiosome
membrane in soybean root nodules (Kaiser et al., 2003). GmZIP1 is another zinc
transporter of soybean. GmZIP1 is also specifically expressed in nodules and localized
to the symbiosome membrane (Moreau et al., 2002). Besides cation transportation
symbiosome membrane also controls transport of anions. GmN70 and LjN70
are inorganic anion transporters of the symbiosome membrane with enhanced
preference for nitrate. Their transport activities may aid in regulation of ion and
membrane potential homeostasis (Vincill et al., 2005).
The water is transported by aquaporins, and in infected cells of the nodules
were identified several aquaporins: PIP1, PIP2 and LIMP1 in Lotus nodules and
TIP1g in Medicago. PIP1 and PIP2 are related to plasma membrane intrinsic proteins
(Wienkoop & Saalbach, 2003). LIMP1 and TIP1g belong to the tonoplast intrinsic
protein (TIP) subfamily of plant membrane intrinsic proteins. Both water channels
are expressed at high levels in nodule and root tissues. Localization of LIMP1 was not
determined. MtTIP1g occurs on tonoplast membrane in root cells and cells of the
apical part of nodules, but it is retargeted to symbiosome membrane in infected cells
22
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General introduction
of the fixation zone. Functional analysis of these transporters by expression in Xenopus
laevis oocytes shows that it is a water-specific aquaporins (Guenther & Roberts, 2000;
Wienkoop & Saalbach, 2003; Gavrin et al., 2014; Chapter 3).
It is remarkable that mechanisms of transportation of fixed nitrogen into the
host cell are not completely clear. One of candidates is Nodulin 26 (NOD26). NOD26,
the most studied transporter of symbiosome membrane, was identified in soybean
nodules by Fortin et al. (1987). Later, it was shown that NOD26 is a member of
aquaporin water channel family, which is able to transport water and glycerol (Rivers
et al., 1997; Guenther & Roberts, 2000). Further studies revealed a range of these
aquaglyceroporins, which were combined in nodulin26-like intrinsic proteins (NIP)
subfamily (Wallace et al., 2006). Relatively recent work of Hwang et al. (2010) showed
ability of NOD26 to facilitate the transport of ammonia (NH3) in an Hg2+-sensitive
manner. An interesting mechanism of assimilation of NH3, after transportation across
symbiosome membrane, was proposed by Masalkar et al. (2010). Authors showed that
soybean glutamine synthetase GS1β1, the principal ammonia assimilatory enzyme,
binds the NOD26 C-terminal domain and therefore promotes efficient assimilation of
fixed nitrogen, as well as prevents potential ammonia toxicity. The use of patch-clamp
techniques, in isolated symbiosome membranes of soybean and L. japonicus nodules,
has revealed the permeability for NH4+ by means of channel-like activity (Roberts &
Tyerman, 2002; Obermeyer & Tyerman, 2005). Nonetheless, a particular transporter,
providing this activity, is still unknown. LjAMT1;1 and LjAMT2 are plant ammonium
transporters of the AMT1 family which expressed in the infected cells of Lotus nodules,
however the biochemical properties of LjAMT1;1 is not compatible with such a role, on
the basis of the ammonium concentrations estimated within the symbiosome space
(Streeter, 1989; Salvemini et al., 2001; D’Apuzzo et al., 2004; Rogato et al., 2008). LjAMT2
are plasma membrane located and has a possible role in ammonium recovery from
the apoplast and potassium homeostasis in plant cells, respectively, rather than roles
in ammonium transport across the symbiosome membrane (Simon-Rosin et al., 2003;
Udvardi & Poole, 2013). Transcription factor GmbHLHm1 was proposed to be linked to
a novel mechanism for NH4+ transport common to both yeast and plants (Chiasson et
al., 2014).
1
Concluding remarks
The host cell undergoes significant changes during the intracellular stage of
symbiosis. Genetic studies in the area of root nodule symbiosis are mainly focused
on NF perception and signaling, nodule organogenesis, and rhizobial infection in the
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Chapter 1
epidermis (Geurts & Bisseling, 2002; Ferguson et al., 2010; Popp & Ott, 2011; Oldroyd,
2013; Maróti & Kondorosi, 2014; Moling et al., 2014; Xiao et al., 2014). However, the
membrane mechanisms controlling intracellular stage of bacterial accommodation
and the development of symbiosomes are crucial part to the symbiotic process. In the
last couple of years, information concerning transcriptomics of infected cells (Limpens
et al., 2013; Roux et al., 2014) and mechanisms controlling symbiosome formation and
development (Van de Velde et al., 2010; Wang et al., 2010; Ivanov et al., 2012) become
available. However, the mechanisms by which architecture of infected cells is created
and endomembrane system is adjusted for the symbiotic accommodation of rhizobia
remain unclear. To get insight into these processes we studied the modifications of
endo- and exocytotic pathways and key regulators of vesicle traffic induced by rhizobia
in host cells of Medicago truncatula and Glycine max. The results of these studies are
presented in this thesis.
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General introduction
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Chapter 2
VAMP721a and VAMP721d are
essential for pectin dynamics
during release of bacteria
in soybean nodules
Aleksandr Gavrin1, David Chiasson2, Evgenia Ovchinnikova3, Brent N. Kaiser3,
Ton Bisseling1,4, Elena E. Fedorova1
Laboratory of Molecular Biology, Graduate School Experimental Plant Sciences, Wageningen
University, 6708 PB, The Netherlands; 2University of Munich (LMU) Institute of Genetics,
Grosshaderner Str. 2-4 D-82152 Martinsried, Germany; 3Centre for Carbon, Water and
Food Faculty of Agriculture and Environment, The University of Sydney, Brownlow Hill,
NSW 2570, Australia; 4College of Science, King Saud University, Riyadh 11451, Saudi Arabia
1
Accepted in The New Phytologist 2015
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Abstract
In the root nodules rhizobia enter host cells via infection threads, the tubular
invaginations of host cell wall and plasma membrane. The release of bacteria to
host cell is possible only from cell wall-free regions of the infection thread. We
hypothesized that the VAMP721d and VAMP721e exocytotic pathway, identified
before in Medicago truncatula (Ivanov et al., 2012), has a role in the local modification of
cell wall during the release of rhizobia. To address this question we use Glycine max, the
plant with a determinate type of nodules. The localization of several polysaccharide
compounds of cell walls was analysed in control versus nodules with partially silenced
GmVAMP72d. The silencing of GmVAMP72d blocks the release of rhizobia. Instead of
rhizobia-containing membrane compartments, symbiosomes, infected cells were
filed by clusters of bacteria embedded in a matrix of methyl-esterified and demethyl-esterified pectin. These clusters were surrounded by membrane. Surprisingly,
we found that GmVAMP72d-positive vesicles do not transport methyl-esterified
pectin. Therefore, we hypothesized that GmVAMP721d-positive vesicles may deliver
the enzymes involved in pectin turnover. Subsequently, we found that GmVAMP72dpositive vesicles partly co-localize with pectate lyase GmNPL1. Therefore, the
biological role of VAMP72d may be explained by its action in delivering pectin
modifying enzymes to the site of release.
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VAMP721d/a are essential for pectin dynamics during release of bacteria
Introduction
The legume symbiosis with soil bacteria (rhizobia) enables the fixation of
atmospheric dinitrogen gas (N2) into ammonium. N2-fixation provides legumes with
a competitive advantage in nitrogen-deficient soils. N2-fixation takes place in root
nodules, which originate from dedifferentiated root cells. Legume nodules exhibit
either indeterminate (with an apical meristem; crops such as pea, alfalfa, clover) or
determinate growth (meristem active only during initial nodule formation; soybean,
birdsfoot trefoil, common bean) (Brewin, 2004; Ferguson et al., 2010).
Rhizobia enter nodules via infection threads, tubular structures, directed from
the apoplast invaginations of host cell wall and plasma membrane. Rhizobia are
released into the host cell cytoplasm via unwalled droplets, which are cell wall–free
regions of infection threads. After gaining entry, rhizobia (now called bacteroids)
subsist in plasma membrane–derived organelles called symbiosomes (Haag et al.,
2013; Kondorosi et al., 2013). In soybean, each symbiosome contains several (4-12)
bacteroids, resulting from the division of bacteria within symbiosomes as well as
continuous fusion of symbiosomes (Fedorova et al., 1999; Brewin, 2004). Development
of the symbiotic interface (i.e., cell wall-free membrane) between the host cell and
intracellular bacteria is crucial for the symbiosis. However, the mechanisms of plant
cell, which control its formation, are largely unresolved.
In our previous work, we have identified the Medicago truncatula exocytotic
pathway crucial for the formation of the symbiotic interface in both rhizobial and
arbuscular mycorrhizal symbiosis. Specifically, members of the vesicle-associated
membrane protein 721 family, named VAMP721d/e, are indispensable for proper
interface development (Genre et al., 2012; Ivanov et al., 2012). However, the biological
mechanisms of their role need to be clarified. The silencing of VAMP721d/e (Ivanov
et al., 2012) in M. truncatula nodules resulted in the formation of abnormal unwalled
droplets. These droplets aberrantly develop a layer of cell wall material near their
membrane forming a physical barrier for rhizobia, preventing their release into
host cells. We hypothesised that VAMP721d/e may be involved in the redirection of
transported cell wall materials to the symbiotic interface region.
Our objective in the present study was to clarify the role of the VAMP721d
exocytotic pathway during the formation of symbiosis in determinant nodule of
Glycine max. We analysed the localization of GmVAMP721d and its functional role in
infected cells. To elucidate the effect of VAMP721d silencing on cell wall modification
during the release of rhizobia we analysed the primary cell wall components: methyl-
2
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Chapter 2
esterified pectin, de-methyl-esterified pectin, cellulose and callose in infected cells of
control nodules versus the transgenic nodules with partial silencing of GmVAMP72d.
The partial silencing of GmVAMP721d has hampered the bacteria release and the
formation of symbiosomes. Host cells were filled with clusters of bacteria embedded
in a pectin matrix. Surprisingly, we found that GmVAMP721d-positive vesicles were
not involved in the delivery of methyl-esterified pectin from Golgi. We therefore
hypothesized that GmVAMP721d-positive vesicles may be transporting some cell wall
remodelling enzymes. Subsequently, we found that GmVAMP721d-positive vesicles
partly co-localize with pectate lyase GmNPL1, an ortholog of the nodule pectate
lyase (LjNPL) from Lotus japonicus (Xie et al., 2012). Therefore the biological role of
VAMP721d in part is explained by its action in delivering pectin modifying enzymes to
the site of release of rhizobia.
Results
GmVAMP72d localized to the site of rhizobial release
To study the role of GmVAMPs in the soybean/Bradyrhizobium japonicum
symbiosis, we targeted the orthologs of Meducago truncatula VAMP721d/e (Ivanov et
al., 2012). GmVAMP721d and GmVAMP721h are orthologous to MtVAMP721d, whereas
GmVAMP721a and GmVAMP721m are orthologous to MtVAMP721e. This shows that
the duplication, which resulted in two related Medicago VAMPs, has occurred in a
common ancestor of Medicago and soybean. Soybean obtained two additional copies
as a result of its genome being a diploidized tetraploid (Shultz et al., 2006). Soybean
therefore has four VAMP721 genes that can play role in the symbiotic exocytotic
pathway. According to SoyBase gene expression data (http://soybase.org/soyseq/),
GmVAMP721d and its closest paralog GmVAMP721a are expressed in roots and nodules,
while the level of GmVAMP721d expression is four times higher in nodules relative to
roots (Supplemental Figure 1). Therefore, GmVAMP721d represents a good candidate
to study exocytotic mechanisms involved in symbiosome interface formation in
soybean nodules.
To visualize GmVAMP721d in infected cells, we used GFP translational fusion
of GmVAMP721d driven by leghemoglobin (LBc3) promoter of soybean. Transgenic
root nodules expressing pLBc3:GFP-GmVAMP721d were obtained via Agrobacterium
rhizogenes-mediated hairy root transformation. In contrast to Medicago, nodules of
soybean have short-lived infection threads and rhizobia are more often released via
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VAMP721d/a are essential for pectin dynamics during release of bacteria
a local opening of the infection thread cell wall (Brewin, 2004). GFP-GmVAMP721d
signal was observed on the membrane of infection threads during rhizobial release
(Figure 1A). To verify the localization of GmVAMP72d, we used M. truncatula antiVAMP721d antibodies (Ivanov et al., 2012) and performed immunolocalization of
GmVAMP721 in wild-type soybean root nodules. Western blot assay showed that this
antibody specifically bound to a protein of an expected size (24.94 kD) in soybean
nodule extract (Supplemental Figure 2). Confocal microscopy showed signal over the
infection threads (Figure 1B), the pattern was identical to one obtained in pLBc3:GFPGmVAMP721d transgenic nodules.
Silencing of GmVAMP721d/a leads to the defect in bacteria release and the formation
of aberrant rhizobial clusters
To investigate the functional role of GmVAMP721d, we used RNA interference
(RNAi) to knockdown its transcript levels. The nodules from pLBc3:RNAi-GmVAMP721d
transgenic roots were analysed 28 days post inoculation (dpi). The expression of
VAMP721d and VAMP721a was approximately 10% and 30% of the control (Supplemental
Figure 3). It is similar to the previous study on Medicago where it was shown that only
knock-down of both Medicago VAMPs (MtVAMP721d and MtVAMP721e) affects the
bacteria release and formation of symbiosomes.
2
Figure 1. GmVAMP721d labels membrane of infection threads during rhizobia release in infected cells of soybean
root nodules. (A) Confocal image of pLBc3:GFP-GmVAMP721d transgenic nodule. (B) Immunolocalization of
GmVAMP721d by using anti-VAMP721d antibody in wild type soybean root nodules. It, infection thread.
Counterstained by FM4-64 (A) and by propidium iodide (B). Bars: 1µm.
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Chapter 2
Light microscopy analysis of GmVAMP721d-silenced nodules (28 dpi) revealed an
altered infection phenotype in comparison to control nodules on roots transformed
with the empty vector (Figure 2A-D). Control nodules show the typical pattern of
developing infected cells. After release from infection threads rhizobia colonize
the host cell forming symbiosomes with several bacteroids inside. During this
process the host cells lose the vacuoles and expand (Figure 2A, B). In GmVAMP721dsilenced nodules the infected cells were filled by large intracellular bacterial clusters
containing 20-40 bacteria embedded in a dense matrix in contrast to relatively small
symbiosomes with transparent peribacteroid space in control (Figure 2C, D). These
bacteria clusters were partly interlinked and formed a net-like structure. Further,
silenced nodules at 28 dpi contained a large number of dead host cells without
cytoplasm packed with bacterial clusters (Figure 2D). The phenotype was observed in
20 nodules (n=20) from 2 separate experiments.
Figure 2. Silencing of GmVAMP72d blocks the release of bacteria and the formation of symbiosomes. (A) Section
of a 28 dpi empty vector control nodule. (B) Magnification of (A) shows the gradient of infected cell development:
young cells with infection threads, freshly colonized infected cells with small vacuoles and symbiosomes. (C)
Mature infected cells are filled with symbiosomes containing 2-6 bacteria. (D) Section of RNAi-GmVAMP721d
nodule. (E) Magnification of (D) shows infected cells filled by large clusters of bacteria embedded in dense
matrix. (F) Infected cell with clusters containing 20-50 bacteria. IC, infected cell; It, infection thread (arrowheads);
BCl, bacteria clusters; (*) dead cell with disintegrating cytoplasm; V, vacuoles (arrows); S, symbiosomes. Bars: a, d
75mm; b, e 25mm; c, f 5mm.
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VAMP721d/a are essential for pectin dynamics during release of bacteria
To describe the phenotype in more detail we performed electron microscopy
(EM) analysis of transgenic nodules versus control nodules. Aberration of root nodule
structure was already observed at the stage of bacterial release. In control nodules,
the release of bacteria from cell wall-free regions of infected threads is preceded by
the formation of a low electron-dense rim of infection thread matrix around bacteria
that are soon to be released (Figure 3A).
After release, bacteria normally divide within symbiosomes, and symbiosomes
fuse forming multi-bacteroids units (Figure 3B, C). In GmVAMP721d-silenced nodules,
the bacterial colonies, which have entered the host cell from the apoplast, were not
able to form a tubular infection thread, which normally contains the single file of
rhizobia surrounded by a plant-derived cell wall. Instead, broad clusters of bacteria
embedded in electron-dense material, structurally similar to cell wall, were formed
(Figure 3D). Further, these clusters grew in volume and bacterial division inside the
cluster was frequently observed. Finally, the intracellular clusters occupied almost
the entire volume of a host cell; the bacteria inside of the clusters showed the signs
of degradation (Figure 3E). We also observed, albeit rarely, normal bacterial release.
Infected cells filled by the bacterial cluster seemed to be losing their integrity, as
organelles were not clearly visible, cytoplasm disintegrated and the loss of contact
between cytoplasm and the cell wall indicates that the cell was also losing turgor
(Figure 3F). The results therefore show that silencing of VAMP721d and VAMP721a
blocks normal bacterial release and symbiosome formation in soybean nodules,
demonstrating that the VAMP721 exocytotic pathway is operational and required for
bacterial release in this symbiosis, as previously observed in Medicago.
2
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Figure 3. Electron microscopy analysis of empty vector control (A, B, C) and RNAi-GmVAMP721d transgenic
nodules (D, E, F). (A) Bacteria release in control nodules. (B) Rhizobia propagation in a young infected cell. (C)
Maturing multi-bacteroids symbiosomes. (D) Young bacteria cluster with dividing bacteria. (E) Mature cluster
containing bacteria with partly degraded cytoplasm. (F) Dead host cells completely filled with bacterial clusters.
S, symbiosomes; YBCl, young bacteria cluster; (*) protruding bacteria is still connected with the cluster; MBCl,
mature bacteria cluster; CDH, colonized dead host cell; Rb, released bacteria; It, infection thread; arrowhead
indicates low electron-dense rim of infection thread matrix around bacteria that are soon to be released; arrow
indicates fusing young symbiosomes.
Finally, to assess the N2-fixing capacity of of pLBc3:RNAi-GmVAMP721d transgenic
nodules, we performed acetylene-reduction analyses (ARA; Supplemental Figure 4).
The ARA of transgenic nodules estimated as µmol C2H4 h-1 g-1 fw was only 10% of the
activity of control nodules. Therefore, silencing of GmVAMP721d results in nearly fixphenotype.
Partial silencing of GmVAMP721d/a impairs the distribution of methyl-esterified
and de-methyl-esterified forms of pectin in infected cells.
To gain insight into the cell wall deposition phenotype observed after silencing
of GmVAMP721d, we analysed the localization of cell wall polysaccharide compounds
(cellulose, callose, methyl-esterified and de-methyl-esterified pectin) in nodules.
The distribution of methyl-esterified and de-methyl-esterified forms of pectin was
analysed by immunostaining with monoclonal antibodies which recognize methyl-
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VAMP721d/a are essential for pectin dynamics during release of bacteria
esterified (LM20) and de-methyl-esterified (LM19) forms of pectin (Verhertbruggen
et al., 2009). The methyl-esterified form of pectin is assembled in the Golgi and
transported to the cell wall in secretory vesicles. After secretion, it is de-methylated by
pectin methylesterases in the cell wall. In soybean control nodules, immunostaining
revealed both epitopes in host cell walls and infection threads (Figure 4A, B).
Further, dot-like structures labelled for methyl-esterified pectin were detected in the
cytoplasm of young cells where bacteria were released. This may reflect the transport
of this form of pectin from Golgi to the plasma membrane. The dot-like structures,
which contain de-methyl-esterified pectin, were also detected in young infected
cells filled with symbiosomes (Figure 4D). The signal of de-methyl-esterified pectin
in infected cells was stronger than the signal for methyl-esterified pectin. In the
transgenic nodules with partial silencing of GmVAMP721d/a, immunostaining of both
methyl-esterified and de-methyl-esterified forms of pectin revealed strong labelling
in the large bacterial clusters (Figure 4E, F, G) described above (Figure 3D). The signal
of these structures was as strong as the signal over the cell walls.
To determine cellulose distribution in infected cells we stained sections with
calcofluor white. Confocal microscopy of control nodules showed localization of
cellulose in host cell walls and weakly in infection threads, however staining was
not observed at the site of release (Figure 5A). pLBc3:RNAi-GmVAMP721d/a nodules
showed normal cellulose staining during the stage of infection and the clusters of
bacteria were not surrounded by cellulose (Figure 5B). Immunolocalization of callose
with anti-(1→3)-β-D-glucan antibodies in control nodules revealed a strong signal
in cell walls of uninfected cells, while staining in walls of infected cell was much
weaker, callose was also found on infection threads walls (Figure 5C). In pLBc3:RNAiGmVAMP721d/a transgenic nodules callose distribution was similar to control during
the stage of release and in young living infected cells. However, we observed numerous
early senescent cells showing the callose deposition, where bacterial clusters also
showed the callose signal (Figure 5D, E).
2
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Figure 4. Imunolocalization of methyl-esterified (MEP) (A, C, E) and de-methyl-esterified (DMEP) pectin (B, D, F,
G) in control (A, B, C, D) and RNAi-GmVAMP721d/a (E, F, G) nodules. (A) Signal indicates MEP in the cell wall of
infected threads and host cells. Image is a 3D reconstruction of the Z-stack showing the cell wall continuity. (B)
DMEP in the cell walls of infection threads and host cell walls. (C) In the young infected cells, filled with
symbiosomes, MEP is mainly present in the cell wall. (D) In contrast, DMEP in the same host cells appears also as
dots in the cytoplasm. (E) MEP and (F) DMEP are present in the wall and the lumen of bacterial clusters in RNAiGmVAMP721d nodules. Image (F) is a 3D reconstruction from z-stack. (G) Early senescent infected cells filled with
bacterial clusters show the strong presence of DMEP in RNAi-GmVAMP721dnodules. Bacteria and nuclei are
contrasted with propidium iodide (red). It, infection thread; CW, cell wall; BCl, bacteria clusters; arrow, bacterial
cluster is attached to the host cell wall; IC, infected cell filled with symbiosomes. Bars: a-d 10μm; e 5μm; f, g 20
μm.
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VAMP721d/a are essential for pectin dynamics during release of bacteria
2
Figure 5. Staining of cellulose (A, B) and callose (C, D, E) in control (A, C) and GmVAMP72d-RNAi nodules (B, D, E).
To spotlight the difference in the labelling of cell walls, infection threads and bacteria clusters in control and
GmVAMP721d-RNAi nodules the staining for cellulose (blue) was coupled with the immunolabelling for methylesterified pectin (red) (A, B). Infection threads in control nodules do not show strong cellulose-positive staining
in comparison with cell wall (A). In GmVAMP721d-RNAi nodules the bacteria clusters did not show the cellulose
labelling, however they are labelled for methyl-esterified pectin. Callose deposition (green) is stronger in the cell
walls of non-infected cells in control nodules (C). In GmVAMP721d-RNAi young nodules the cell wall staining is
more equal in infected and non-infected cells (D). In dead cells the increased callose deposition is observed
around the bacterial clusters (arrowhead). Bacteria are counterstained by propidium iodide (red). It, Infection
thread; IC, infected cell; NI, non-infected cell; BCl, bacterial clusters; CW, cell wall. Bars: a, b, c 10µm; d, e 5µm.
Hence, the rhizobia in silenced nodules were imbedded in a matrix of both
methyl-esterified and de-methyl-esterified pectin, in contrast with infection threads
or symbiosomes from control plants. These observations show that the transport,
targeting or turnover of pectins in the nodules with partially silenced GmVAMP721d/a
was impaired.
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Role of the GmVAMP721d pathway in pectin transport
To clarify the possible role of GmVAMP721d vesicles in the transport of
methyl-esterified pectin from Golgi to the infected cell wall, we performed coimmunolocalization of GmVAMP721d and methyl-esterified pectin using the
antibodies LM20 and anti-VAMP721d. Both antibodies displayed dot-like patterning;
however the co-localization of the dots was not observed in infected cell from soybean
nodule (Figure 6A). We additionally have tested the co-localization in the Medicago
infected cell (Supplemental Figure 5) with the same negative result. This suggests that
GmVAMP721d-positive vesicles do not transport methyl-esterified pectin.
To clarify the nature of the dot-like structures containing de-methyl-esterified
pectin in infected cells, we performed electron microscopy immunogold detection
using antibody LM19. In infected cells of control nodules, the epitopes of de-methylesterified pectin were observed in the cell walls and in the walls of infection threads
(Figure 7A, B). The signal was also observed in the lumen of vesicles with a diameter
of 50-200 µm. These vesicles are similar in both size and structure to endosomes and
young vacuoles (Figure 7B). Further, the signal of de-methyl-esterified pectin was
observed in the lumen of vacuoles (Figure 7C), implying that the labelled endosomes
may be targeted to the vacuole. Endocytosis of de-methyl-esterified pectin, which
implies substantial level of pectin turnover in young infected cells, may explain our
observations. In control nodules, the labelling of de-methyl-esterified pectin in
symbiosomes was weak; however some gold particles were visible in peribacteroid
space of mature symbiosomes (Figure 7D). The de-methyl-esterified pectin epitopes
in the peribacteroid space of mature symbiosomes may derive from the fusion
with endosomes carrying pectin, possibly endocytosed from the cell wall (Figure
7B). The retargeting of the endosomes to symbiosomes instead of the vacuole may
be explained by the fact that the vacuole in soybean infected cells is a short-lived
transitory organelle (Figure 2 B, C). The collapse ofthe vacuole in infected cells may
result in retargeting of tonoplast-bound traffic towards symbiosomes, as occurs in
Medicago nodules (Gavrin et al., 2014; Chapter 3).
Hence, in GmVAMP721d/a-silenced nodules we observed strong labelling for demethyl-esterified and methyl-esterified pectin (Figure 7E, F) in the matrix of enlarged
bacterial clusters comparable with the level of labelling of the cell wall. This supports
a role of GmVAMP721d in cell wall remodelling during symbiosis formation.
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VAMP721d/a are essential for pectin dynamics during release of bacteria
2
Figure 6. Co-localization of VAMP721d-positive vesic-les with MEP and GmNPL1. (a) VAMP72d-positive vesic-les
(red) are not co-localized MEP-carrying vesicles (green). (b) The colocaliza-tion of GFP-GmNPL1 (green) with
VAMP72d-positive vesicles (red) over the infection thread gives a combined yellow signal. (c) anti-VAMP72d
labelling (red) in infected cells shows partial co-localisation with GFP-GmNPL1 (arrowheads). (d) Signal of GFPtagged GmNPL1 detected using anti-GFP antibody. Bars: a, b 5µm; c, d 20µm.
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Chapter 2
Figure 7. Immunogold labelling of DMEP in infected cells of empty vector (control) nodules (A-D); DMEP and
MEP in pLBc3:RNAi-VAMP721d/a transgenic nodules (E, F). (A) In the site of bacteria release the labelling for DMEP
is reduced in comparison with the labelling over the wall of infection thread. (B) DMEP is endocytosed in infected
cells: the strong labelling for DMEP is visible over the cell walls and endosomes, and over the (C) infection
threads and vacuoles. (D) The fusion of a symbiosome with DMEP-labelled endosomes. Arrows indicate
endosomes. Asterisk indicates weak immunogold labelling. Arrowhead indicates a symbiosome membrane
fusion. (E) Immunogold labelling of MEP in VAMP721d/a transgenic nodules. (F) Labelling of DMEP (E) over the
filamentous material inside of the clusters of bacteria. Rel, bacteria release; B, bacteroid; BCl, clusters of bacteria;
En, endosome; It, infection thread; S, symbiosome; Vac, vacuole. Bars: b, e, f 200 nm; a, c, d 500 nm.
Co-localization of pectate lyase with GmVAMP721d
Silencing of GmVAMP721d/a results in the colonization of infected cells by clusters
of bacteria embedded in a matrix containing two forms of pectin. We therefore
hypothesized that GMVAMP721d may be involved in the delivery of cell wall
remodelling enzymes and can be operational on the site of bacteria release. To test
this, we have selected the enzyme pectate lyase (GmNPL1), the soybean orthologue
of Lotus japonicas nodulation pectate lyase LjNPL (Xie et al., 2012). The pectate lyase
LjNPL plays a critical role in symbiosis development and is involved in the infection
threads progression and bacteria release. We have studied the localization and
putative relations of GmNPL1 with VAMP721d pathway.
We generated transgenic root nodules expressing GFP-tagged GmNPL1. The
sections of pLBc3:GFP-GmNPL1 transgenic nodules were used for immuno colocalization with the specific antibody against VAMP721d and observed by confocal
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VAMP721d/a are essential for pectin dynamics during release of bacteria
microscopy. The co-localization of GmNPL1 (green) with VAMP721d-positive vesicles
(red) over the infection thread has been shown by the strong combined yellow signal
(Figure 6B, C).
The quantitative estimation of co-localized dots across 12 infected cells from
3 transgenic nodules showed that 39±12% of dots positive for VAMP721d were colocalized with dots carrying the GFP-GmNPL1 (Figure 6B, C). The dots positive for
VAMP721d were used as a reference because they were more abundant. This suggests
that VAMP721d may be involved in the targeting of GmNPL1 to the sites of release as
well as to the plasma membrane of young expanding cells.
Discussion
2
The development of infected nodule cells results in the coordinated expansion
of symbiotic bacteria in number and volume, the enlargement of the host cell and
the formation of the cell wall-free symbiotic membrane interface between the host
cytoplasm and bacteria. To accommodate the microsymbionts the infected cell
requires a high level of cell wall expansibility that is coordinated with bacterial release
from unwalled regions of infected threads and with the bacteria expansion. This
necessitates localised modification of the host cell wall for the rhizobial symbiosis to
occur (Giordano & Hirsch, 2004).
The comprehensive review concerning the modification of cell walls during the
nodule initiation and infection threads growth was given by Brewin (2004). However,
the molecular mechanisms of host cell to form the symbiotic interface have to
be clarified. Our objective was to define the role of GmVAMP721d in the symbiosis
between soybean and B. japonicum. The function of the symbiotic VAMP721d
exocytotic pathway has previously been studied in Medicago (Ivanov et al., 2012).
Further, another member of the same family, VAMP721a, was recently shown to be
a target of regulator of symbiosome differentiation (RSD). RSD is a member of the
Cysteine-2/Histidine-2 (C2H2) family of plant transcription factors and is required for
normal symbiosome differentiation of M. truncatula (Sinharoy et al., 2013).
Silencing of VAMP721d exocytotic pathway had no effect on infection thread
formation in Medicago nodules, but bacteria release was blocked (Ivanov et al., 2012).
In this case the formation of unwalled bacterial containing droplets was initiated but
a wall, albeit markedly thinner than a normal cell wall, enclosed the membrane and
prevented bacterial release. Hence, we hypothesized that the VAMP721d controlled
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Chapter 2
exocytotic pathway may have a role in the local modification of infected cell wall
structure and formation of symbiotic interface.
To clarify the role of GmVAMP721d we analysed the location of this protein in
soybean nodules relative to the distribution of key cell wall components, including
methyl-esterified and de-methyl-esterified pectins, cellulose and callose in infected
cells after silencing of GmVAMP721d. The soybean ortholog of the VAMP721d displayed
similar nodule localization patterns to that observed in Medicago, accumulating at
the site of bacterial release from infection threads and amongst membranes of young
symbiosomes. The partial silencing of GmVAMP721d also blocked rhizobia release but
caused a structurally different phenotype to that in Medicago. In Medicago, loss of
MtVAMP721d caused a failure to remove cell wall material from unwalled droplets
that has prevented the bacterial release. In soybean nodule, however, the formation of
tubular infection threads was inhibited creating baggy clusters of bacteria embedded
in a pectin matrix. The reason for the differences in phenotypes between soybean and
Medicago is not clear but supports the idea that membrane mechanisms by which the
symbiotic interface is formed may be different between determinate (soybean) and
indeterminate (Medicago) nodules.
According to the previous works the glycoproteins and glycolipids but not
polysaccharides or cell wall material were found in symbiosomes from pea and
soybean root nodules (Roth & Stacey, 1989; Perotto et al., 1991). That also support our
conclusion that bacteria clusters that we have observed in GmVAMP721d/a-silenced
nodules are not symbiosomes. Though, it is worth to mention that the symbiosomes
from the caesalpinioid legume Dimorphandra wilsonii are embedded in de-methylesterified pectin, that structural arrangement may represent the ancient state in the
evolution of symbiotic compartments (Fonseca et al., 2012).
The analysis of primary cell wall components in VAMP721d-silenced nodules
shows that there is no influence on the deposition of cellulose or callose on host cell
wall development; however there was an impact on pectin accumulation. This was
surprising as we found that the vesicles labeled by GmVAMP721d are not involved in
the transport of methyl-esterified pectin from the Golgi. The second unusual feature
was the high level of pectin endocytosis manifested by endocytosis of de-methylesterified pectin in young infected cells. Pectin endocytosis has been observed in
Arabidopsis thaliana root meristem cells and found important for the maintenance
of plant cell wall integrity and flexibility during cell division (Baluška et al., 2005).
However, this feature is rather unusual for differentiated young infected cells and
represents an interesting biological phenomenon, which may be important for host
cell extensibility during infected cell expansion.
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VAMP721d/a are essential for pectin dynamics during release of bacteria
Flexible cell walls in growing regions like pollen tube tips are enriched in methylesterified pectin delivered from the Golgi (Bárány et al., 2010). In differentiated cells
de-methyl-esterfied pectin is more abundant (Bosch et al., 2005). This implies that
high levels of methyl-esterified pectin in infected cell walls would favor cell wall
expansion (Peaucelle et al.; Bárány et al., 2010). Cell wall pectin composition also
affects interactions with other cell walls components (Palin & Geitmann, 2012; Wolf
et al., 2012; Bashline et al., 2014). For example, the formation of a pectin-cellulose
complex is highly dependent on the degree of pectin methylation. It was shown that
highly methyl-esterified pectin has a low binding affinity for cellulose microfibrils,
resulting in a more flexible cell wall (Yoneda et al., 2010). De-methylated pectin is less
flexible and more adhesive (Wolf et al., 2009). The defect in bacteria release, which
we have observed in GmVAMP721d-silenced nodules partly, may be explained by a
defect in pectin turnover that leads to excess deposition of both forms of pectin in the
lumen of bacterial clusters. The physical constraints and compromised rheological
characteristics of the matrix of bacterial clusters, which become too adhesive, may
explain the observed block of bacterial release. Another component of a fluid-to-solid
transition might involve the enzymatic removal of the arabinogalactan side chains
from root nodule extensin in the luminal matrix (Brewin, 2004). The importance of
pectin turnover in symbiosis has been demonstrated in M. truncatula (RodríguezLlorente et al., 2004) and in L. japonicas (Xie et al., 2012). A mutation in L. japonicas
pectate lyase LjNPL affects infection thread progression, rhizobial release and nodule
development. De-methyl-esterification facilitates degradation by pectate lyases and
polygalacturonases and promotes pectin turnover (Lionetti et al., 2012).
With the aim to clarify the link between VAMP721d and pectin turnover we
performed co-localization of soybean pectate lyase GmNPL1 and VAMP721d. From
this analysis we found that a part of the VAMP721d-positive vesicles contained
GmNPL1. This suggests that VAMP721d is involved in the delivery of pectate lyase and,
possibly, other cell wall modifying enzymes during symbiosis. We cannot exclude
that other enzymes involved in increasing pectin elasticity, for example inhibitors
of demethylation or inhibitors of pectin polymerization (Rodríguez-Llorente et al.,
2004; Wolf et al., 2012; Sénéchal et al., 2014), could also be delivered to the cell wall via
VAMP721d vesicles. Hence, the phenotype we observed as a result of partial silencing
of VAMP721d may reflect a more complex situation than only the shortage of the
vesicles carrying the pectate lyase.
In conclusion, this study shows that soybean has a GmVAMP721d-controlled
exocytotic pathway that is essential for release of rhizobia from infection threads.
2
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Chapter 2
This pathway is crucial in creating unwalled regions of the infected thread. The
partial association of GmNPL1 with VAMP721d vesicles suggests a possible role of the
VAMP721 exocytosis pathway in delivering pectin modifying enzymes to the site of
release and to host cell wall.
Materials and Methods
Plant Materials, Transformation and Inoculation.
Glycine max L. cv. Djakal seeds were sown in Waikerie river sand, inoculated with
Bradyrhizobium japonicum USDA 110 four days after planting. Plants were fertilized
twice weekly with nutrient solution lacking nitrogen (Herridge et al., 1990). Plants
were grown in a glasshouse (24–28º C) under long day conditions (16/8 h day/
night). The protocol of plant transformation is described previously (MohammadiDehcheshmeh et al., 2013).
Cloning
The open reading frames of GmVamp721d (Glyma09g02310) and GmNPL1
(Glyma11g37620) were amplified using PCR from 28-day-old nodule cDNA template
using Phusion high fidelity polymerase (Finnzymes). Entry clones were generated
by TOPO cloning (Invitrogen). The Gateway cloning system (Invitrogen) was used
to create genetic constructs for RNA interference (RNAi) and GFP-fusion. pENTR
clones were recombined into the following destination vectors using LR Clonase
(Invitrogen): pGmLBc3-pK7GWIWG2 for RNAi and pGmLBc3-pK7WGF2-R for
N-terminal GFP-X fusions. G. max leghemoglobin (LBc3) promoter (2000-bp region
upstream of the ATG start codon of Glyma10g34260; X15061) was directionally cloned
into pGEM-T (Promega) for modifications of destination vectors. A GFP fusion vector
containing the GmLBc3 promoter, was created by the exchange of the 35S promoter in
35S-pK7WGF2-R for GmLBc3 promoter using Hind III-Spe I restriction sites (Smit et al.,
2005). Modification of the RNAi vector was not possible by direct exchange of promoters
as there were several Spe I restriction sites in pK7GWIWG2 (Karimi et al., 2002). To
create an RNAi vector driven by the GmLBc3 promoter, a region of pK7GWIWG2, which
is flanked by Mlu I and Sac I, was PCR-amplified and cloned into pCR8 (Invitrogen). This
vector was further modified by replacing the 35S promoter for the GmLBc3 promoter by
Sac I – Spe I restriction sites. After the modified region of pK7GWIWG2 was introduced
into pK7GWIWG2 by Mlu I – Sac I, replacing indigenous sequence.
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VAMP721d/a are essential for pectin dynamics during release of bacteria
Gene expression.
Total RNA was extracted from roots and 28 days post inoculation (dpi) root
nodules using E.Z.N.A. Plant RNA Mini Kit (Omega Bio-Tek) and transcribed into
cDNA using the iScript cDNA synthesis kit (Bio-Rad). Real-time PCR was set up in
a 20-μL reaction system using iQ SYBR Green Supermix (Bio-Rad). Gene-specific
primers were designed with Primer-3-Plus software (Untergasser et al., 2007). Geneexpression profiles were normalized against the transcription level of reference gene
cons6(Libault et al., 2008).
Confocal microscopy.
Confocal imaging of GFP-fused proteins was done on transgenic hand sectioned
nodules using a Zeiss LSM 5 Pascal confocal laser-scanning microscope (Carl
Zeiss). For the localization study of GmVamp721d, we used polyclonal rabbit antiVAMP721d/VAMP721e (Ivanov et al., 2012) at a dilution of 1:50–100 and secondary
anti-rabbit Alexa 488 antibodies (Molecular Probes) at a dilution 1:200. For callose
immunolocalization, a monoclonal anti-callose antibody (Biosupplies, Australia)
was used (1:20 dilution). The secondary antibody was anti-mouse-Alexa Fluor 488
(Molecular Probes) diluted 1:50. Cellulose was analyzed with Calcofluor White
(Sigma-Aldrich) (1g/l) mixed with Evans blue (0.5 g/l).
Monoclonal rat antibodies LM19 and LM20 were used for the localization of
pectins (PlantProbes). Nodule sections were blocked in 1% (w/v) BSA. Sections were
counterstained either by FM4-64 (30 μg/mL) or by propidium iodide (PI) 0.01%.
2
Sample preparation for light and electron microscopy (EM) and EM
immunodetection.
The protocol of tissue processing has been described previously (Limpens et al.,
2009). Semi-thin sections (0.6 μm) for light microscopy and thin sections (60 nm)
for EM of transgenic nodules were cut by using a Leica Ultracut microtome. Nickel
grids with the sections were blocked in normal goat serum diluted in PBS and then
incubated with the primary antibody according to dilutions described above. Goat
anti-rat antibody coupled with 10-nm gold (BioCell) (1:50 dilution) were used as the
secondary antibody. Sections were examined using a JEOL JEM 2100 transmission
electron microscope equipped with a Gatan US4000 4K×4K camera.
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Acknowledgements
A.G. received a PhD fellowship from EPS School of Biological Sciences (Wageningen
University). E.F. and T.B. are supported by the European Research Council (ERC-2011AdG294790).
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P, Palomares AJ. 2004. From pollen tubes to infection threads: recruitment of Medicago floral pectic genes
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E, Udvardi MK. 2013. The C2H2 Transcription Factor REGULATOR OF SYMBIOSOME DIFFERENTIATION
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Is Essential for Rhizobial Nod Factor-Induced Transcription. Science 308(5729): 1789-1791.
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Verhertbruggen Y, Marcus SE, Haeger A, Ordaz-Ortiz JJ, Knox JP. 2009. An extended set of monoclonal
antibodies to pectic homogalacturonan. Carbohydrate Research 344(14): 1858-1862.
Wolf S, Hématy K, Höfte H. 2012. Growth Control and Cell Wall Signaling in Plants. Annual Review of Plant Biology
63(1): 381-407.
Wolf S, Mouille G, Pelloux J. 2009. Homogalacturonan Methyl-Esterification and Plant Development. Molecular
Plant 2(5): 851-860.
Xie F, Murray JD, Kim J, Heckmann AB, Edwards A, Oldroyd GED, Downie JA. 2012. Legume pectate lyase
required for root infection by rhizobia. Proceedings of the National Academy of Sciences 109(2): 633-638.
Yoneda A, Ito T, Higaki T, Kutsuna N, Saito T, Ishimizu T, Osada H, Hasezawa S, Matsui M, Demura T. 2010.
Cobtorin target analysis reveals that pectin functions in the deposition of cellulose microfibrils in parallel
with cortical microtubules. The Plant Journal 64(4): 657-667.
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VAMP721d/a are essential for pectin dynamics during release of bacteria
Supplementary materials
root nodule 1800 1600 1400 1200 1000 800 600 400 200 0 2
Supplemental Figure 1. Expression profile of GmVAMP72 genes based on RNA-Seq Atlas of Glycine max data
(http://soybase.org/soyseq/) in roots and nodules.
Supplemental Figure 2. Western blot analysis of the anti-VAMP72d antibodies specificity on extracts from (A)
soybean and (B) M. truncatula nodules.
Supplemental Figure 3. The level of gene silencing of VAMP72d and VAMP72a in pLBc3:RNAi-GmVAMP72d
transgenic nodules.
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1,2 1 0,8 0,6 0,4 0,2 0 control
RNAi-VAMP72d
Supplemental Figure 4. N2-fixing capacity of pLBc3:RNAi-GmVAMP72d transgenic nodules in comparison with
empty vector control nodules estimated by acetylene-reduction analysis (µmol C2H4/h/g).
Supplemental Figure 5. VAMP72-positive vesicles are not co-localized with methyl-esterified pectin in M.
truncatula infected cell
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VAMP721d/a are essential for pectin dynamics during release of bacteria
Table S1. Primers used for cloning, vector modifications and qRT-PCR analysis.
Cloning primers
Forward
Reverse
GmVAMP72d
CACCATGGGGCAGAAGTCTCTGAT
CTATTTTCCACAGTTGAACCCA
GmNPL1
CACCATGGCATTTTCATTCACATTT
CTAGTGGCATAGATTGCCCTT
pLBc3
CCGTATCGATCGCACAAAA
ATTTCTTTTCTACTTTTCTGTTATTTCTATT
Primers for vector modifications
pK7GWIWG2
Forward
Reverse
ACGCGTGGATCAGCTTAATA
GAGCTCTCCCATATGGTCGA
2
qRT-PCR primers
Forward
Reverse
GmVAMP72d/h
CGCACACACCTTCAATTACC
AGCCATGGGTAACTGTCTTCC
GmVAMP72a/m
ACACAATCGCGTTCCAGTG
TCGACGAGGTAGTTGAAGGTG
GmCons6
CTAATGGCAATTGCAGCTCTC
TAGATAGGGAAATTGTGCAGGTC
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Chapter 3
Adjustment of host cells
for accommodation of
symbiotic bacteria: vacuole
defunctionalization, HOPS
suppression and TIP1g retargeting
in Medicago.
Aleksandr Gavrin1,Brent N. Kaiser2, Dietmar Geiger3, Stephen D. Tyerman4,
Zhenqyu Wen2, Ton Bisseling1,5, and Elena. E. Fedorova1
1
Laboratory of Molecular Biology, Graduate School Experimental Plant Science, Wageningen University,
Droevendaalsesteeg 1, 6708PB Wageningen, The Netherlands; 2Centre for Carbon, Water and Food Faculty
of Agriculture and Environment, The University of Sydney, Brownlow Hill, NSW 2570, Australia; 3Institute
for Molecular Plant Physiology and Biophysics, University Würzburg, Julius-von-Sachs Platz 2, D-97082,
Germany; 4School of Agriculture, Food and Wine, Waite Research Institute, The University of Adelaide,
SA5064, Australia; 5College of Science, King Saud University, Post Office Box 2455, Riyadh 11451, Saudi Arabia
Published in The Plant Cell 2014 26: 3809–3822
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Chapter 3
Abstract
In legume-rhizobial symbioses, the bacteria in infected cells are enclosed in
a plant membrane, forming organelle-like compartments called symbiosomes.
Symbiosomes remain as individual units and avoid fusion with lytic vacuoles of host
cells. We observed changes in vacuole volume of infected cells, and thus hypothesized
that microsymbionts may cause modifications in vacuole formation or function. To
examine this, we quantified the volumes and surface areas of plant cells, vacuoles,
and symbiosomes in root nodules of Medicago truncatula and analysed the expression
and localization of VPS11 and VPS39, members of the HOPS vacuolar-tethering
complex. During the maturation of symbiosomes to become N2-fixing organelles,
a developmental switch occurs and changes in vacuole features are induced. For
example, we found that expression of VPS11 and VPS39 in infected cells is suppressed
and host cell vacuoles contract, permitting the expansion of symbiosomes.
Trafficking of tonoplast-targeted proteins in infected symbiotic cells is also altered,
as shown by retargeting of the aquaporin TIP1g from the tonoplast membrane to the
symbiosome membrane. This retargeting appears to be essential for the maturation
of symbiosomes. We propose that these alterations in the function of the vacuole are
key events in the adaptation of the plant cell to host intracellular symbiotic bacteria.
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Adjustment of host cells vacuoles for accommodation of symbiotic bacteria
Introduction
Legumes can establish symbioses with the N2-fixing bacteria that are collectively
named rhizobia. The symbiosis leads to the formation of a new organ, the root nodule.
Unique in higher plants, the nodule cells contain thousands of bacteria, which are kept
in individual membrane compartments provided by the host. The membrane-bound
bacterial units are called symbiosomes and show structural similarities to microbes
housed in mammalian pathogenic vacuoles (Brumell & Scidmore, 2007; Isberg et al.,
2009; von Bargen et al., 2009). However, unlike mammals, legumes have specialized
cells that promote intracellular bacteria accommodation whereas in mammalian
tissues such cells do not exist.
In nitrogen-fixing infected cells, symbiosomes do not fuse with the lytic vacuole
and remain as individual units within the cytosol. The mechanisms that inhibit
this fusion and subsequently enhance lytic clearance in senescing infected cells are
unknown. To clarify the mechanisms of symbiotic cell adaptation to intracellular
bacteria, we first quantified cell, vacuole, and microsymbiont surface/volume
dynamics during nodule development. This showed that vacuole modification plays
a crucial role in symbiotic cell progression. We hypothesized that maintenance of
symbiosomes requires a major adjustment of the vacuole formation pathway and
tonoplast-targeted trafficking. Therefore, we characterized the vacuoles of host cells
during intracellular bacterial accommodation.
We selected for our studies the model legume Medicago truncatula. M. truncatula
nodules have a persistent meristem; as a result, the nodule is composed of zones
representing subsequent stages of development. The apical part of the nodule
consists of the meristem and the infection zone. At this site, bacteria are released from
infection threads into the host cell cytoplasm. Upon release, bacteria are surrounded
by a host cell–derived membrane to form symbiosomes. The release requires a specific
exocytotic pathway (Ivanov et al., 2012) and the symbiosomes continue to share some
properties of the plasma membrane during their lifespan (Catalano et al., 2007). After
release, rhizobia grow, divide, and gradually colonize the entire host cell.
Next, mature infected cells form in the so-called fixation zone. In these cells, the
rhizobial enzyme nitrogenaseis induced, allowing the bacteria to reduce atmospheric
nitrogen to ammonia, and the bacterial differentiation process is terminated (Vasse
et al., 1990; Maagd et al., 1994; Farkas et al., 2014). At later stages of maturation, the
symbiosome membrane acquires tonoplast and late endosomal identity markers
(Behnia & Munro, 2005), including small GTPase Rab7 and vacuolar SNAREs (Limpens
3
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Chapter 3
et al., 2009). Symbiosomes have some vacuolar properties, but they do not fuse with
the vacuole in nitrogen-fixing infected cells. To test our hypothesis that the pathway
of vacuole formation in infected cells is impaired, we examined the expression and
localization of proteins belonging to the tethering complex HOPS (homotypic
fusion and vacuole protein sorting complex). HOPS is the key regulator involved in
formation of the vacuole (Nickerson et al., 2009; Balderhaar & Ungermann, 2013).
In yeast, the HOPS complex consists of six vacuolar sorting proteins (VPS): VPS11,
VPS16, VPS18, VPS33, VPS39 and VPS41. The HOPS complex ensures specificity during
fusion of membranes with the vacuole (Balderhaar & Ungermann, 2013). In plants,
HOPS proteins also function in vacuole formation and localize to the tonoplast and
prevacuolar compartments. A null mutation of VPS16 causes embryonic lethality in
Arabidopsis thaliana (Rojo et al., 2001; Rojo et al., 2003).
To test whether default tonoplast-targeted trafficking is compromised in infected
cells, we investigated the localization of the vacuolar aquaporin TIP1g. Aquaporins
are membrane proteins that facilitate the transport of small molecules such as water,
glycerol and ammonia (Chaumont & Tyerman, 2014). Higher plant aquaporins are
subdivided into five main subfamilies: the plasma membrane intrinsic proteins (PIPs),
the tonoplast intrinsic proteins (TIPs), the Nodulin26-like intrinsic proteins (NIPs),
the small basic intrinsic proteins (SIPs), and the X intrinsic proteins (XIPs) (Forrest &
Bhave, 2007; Maurel et al., 2009; Wudick et al., 2009; Hwang et al., 2010).
The transition from infection to nitrogen fixation requires a developmental switch
that controls the adaptation of host cells to the accommodation of nitrogen-fixing
rhizobia. At this transition, rhizobial nif genes are expressed and the HOPS complex
is turned off, a process strictly correlated with the collapse and defunctionalization
of vacuoles in the infected cells. As a consequence, the tonoplast aquaporin TIP1g
is retargeted toward symbiosomes. This retargeting is in line with the loss of fusion
specificity of tonoplast-targeted vesicles due to temporary suppression of HOPS.
Retargeting appears to be essential for functional maturation of symbiosomes
and the maintenance of turgor pressure in the infected cells. Our study shows that
a major adjustment in vacuole formation and tonoplast targeting occurs during
the development of the infected cells and we discuss how this contributes to the
mechanisms by which symbiosomes are maintained in infected cells of root nodules.
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Adjustment of host cells vacuoles for accommodation of symbiotic bacteria
Results
Host cell architecture during development of infected cells
To determine the adaptation of infected cells to accommodate intracellular
rhizobia, we quantified the volume and surface area of cells, symbiosomes and vacuoles.
For this, we used confocal microscopy and 3D reconstruction in combination with
Imaris 7.5 software, to image transgenic roots expressing GFP-tagged SYP22 (vacuolar
SNARE), which specifically labels the vacuoles (Limpens et al., 2009). Nodules were
stained with propidium iodide (PI) to contrast the cell wall, nuclei and symbiosomes.
Z-stacks obtained by confocal microscopy were used for 3D reconstruction and
quantification of the volumes occupied by vacuoles and symbiosomes. Infected and
non-infected cells were analyzed in subsequent developmental stages. An overview
of nodule zones and the developmental stages of symbiosomes is provided in Figure 1
as a cartoon (Figure 1A) and light microscopy images (Figure 1B-E). Infected and noninfected cells were analyzed at four stages of development: (1) young (distal) cells of
the infection zone (Figure 1A, B, C, F); (2) the most proximal cell layer of the infection
zone (3); the (adjacent) first cell layer of the fixation zone (interzone 2/3 according to
Vasse et al., 1990) (Figure 1A, B, D, G) and more mature infected cells of the fixation
zone (Figure 1A, B, E, H). For 3D reconstruction, Z-stacks covering whole cells were
obtained for at least eight infected/non-infected cells at each selected developmental
stage (Figure 1F-H). Figure 1I shows an example of 3D reconstructions of infected cells.
Young infected cells of the infection zone are relatively small, compared to mature
cells, and about 65% of their cell volume is occupied by vacuoles. The vacuole size is
comparable with that of non-infected cells (Figure 1C, F, and Figure2A). In the most
proximal layer (=oldest) of the infection zone, cells have increased in size with about
60% of the cell volume occupied by the vacuole and 30% by the symbiosomes. In
the adjacent cell layer, the first cell layer of the nitrogen-fixing zone, the vacuoles of
infected cells collapse as their volume decreases (Figure 1D, G and Figure 2A). The
infected cells have only slightly increased in size in this transition layer. In contrast,
the absolute volume of the vacuoles was reduced 4-fold and the absolute volume
of symbiosomes increased 2.5-fold to occupy about 65% of the cell volume (Figure
2A). Hence, the absolute volume of thevacuoles in this zone decreased and the total
volume of microsymbiont increased.
3
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Chapter 3
Volume, µm3
Figure 1. Nodule zonation, stages of symbiosome development, and 3D reconstructed cells of M. truncatula
nodules. (A) Scheme of the nodule zones: M, meristem; zII, infection zone; zIII, zone of nitrogen fixation. The
stages of symbiotic cells in development are shown next to each other (1 to 4). Zones described here correspond
to the light microscopy image of the nodule (B). The arrow points to the zone of transition between the infection
and fixation zones. The vacuoles of infected cells in this zone are highlighted by red. Here, the arrows point to the
cells in consequent stages of development given in high-magnification images ([C] to [E]) and as examples of 3D
reconstruction ([F] to [H]). (C) And (F) Cells from the zone of
A bacteria release. (D) And (G) The transition zone
**
(interzone 2/3). It is recognized by the appearance of starch grains. (E) And (H) Mature infected cells.
Flaccid
vacuoles of a mature symbiotic cell have a “scalloped” surface, whereas vacuoles of an uninfected cell are
roundish and turgid. (I) An example of Imaris rendering based on a confocal Z-stack of a*GFP-SYP22-expressing
nodule. Frames show the reconstructed images of vacuoles from the neighboring cells. Samples were contrasted
by PI (red). IC, infected cell; It, infection thread; s, symbiosomes; SG, starch grains; UC, uninfected cell; Uv, vacuole
of uninfected cell; v, vacuole. Bar in (B) = 100 μm; bar in (C) = 25 μm; bars in (D) and (E) = 75 μm; bar in (H) = 12.5
μm; bar in (F) = 5 μm; bar in (G) = 10 μm; bar in (I) = 50 μm.
A
B
**
Area, µm2
Volume, µm3
*
.
Area, µm2
2. The quantification
of cell, vacuole
and SBs
volumefrom
and vacuole
surface area
B
Figure
2. Quantification of cell, vacuole, and symbiosomeFigure
volume
and vacuole
surface
area
Imaris
3Dbased on
Imaris 3-D reconstructed Z-stacks of infected cells.
(A) Dynamics
of vacuole, SBs and
cell volume.
*P<0.001,
**P<0.05. *P < 0.001,
Reconstructed Z-stacks of infected cells. (A) Dynamics of vacuole,
symbiosome
(SBs),
and
cell
volume.
(B) Dynamics of vacuole surface area.
Error bars represent
standard deviations
(n = 8).the data sets.
**P < 0.05. (B) Dynamics of vacuole surface area. Error bars indicate
SD calculated
from
64Figure 2. The quantification of cell, vacuole and SBs volume and vacuole surface area based on
.
Imaris 3-D reconstructed Z-stacks of infected cells.
(A) Dynamics of vacuole, SBs and cell volume. *P<0.001, **P<0.05.
(B) Dynamics of vacuole surface area.
Error bars represent standard deviations (n = 8).
35351_Gavrin.indd 64
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Adjustment of host cells vacuoles for accommodation of symbiotic bacteria
In the cell layers after the termination of microsymbiont expansion, infected cells
have doubled in size. The vacuoles are again clearly present; hence the vacuole volume
returns to pre-infection level. However, vacuoles remain flaccid in the mature infected
cells of the fixation zone (Figure 1E, H). The change in vacuole volume suggests that
during the transition from infection to fixation zone the vacuoles are maintained and
only their volume has been reduced (Figure 1D, E). To determine whether this is the
case, we quantified the surface of the vacuoles using the Imaris 7.5 software (Reisen
et al., 2005). This showed that the surface of the tonoplast indeed remained similar,
whereas the volume of vacuoles decreased 4-fold (Figure2B). This experiment was
repeated twice and both experiments showed a similar change in the vacuole volume
in infected cells. The total vacuole volume in mature infected cells is about 30% of
total cell volume, whereas for plant cells it is normally about 80-90% (Reisen et al.,
2005). This change in vacuole morphology prompted us to determine whether other
vacuolar properties are also modified.
Vacuoles of infected cells are not acidic
Functional lytic vacuoles have a characteristic acidic pH. To determine the pH
in infected cells we stained them with Neutral Red (NR), a supravital stain and
pH marker. In its unprotonated form, NR penetrates the plasma membrane and
tonoplast and accumulates in acidic compartments, resulting in red staining of
these compartments. However, the detection of NR staining in root nodules by light
microscopy (Fedorova & Brown, 2007; Sujkowska et al., 2011) has disadvantages, due
to the multilayer structure of nodule sections. Therefore, hand sections of the nodules
were analyzed by confocal microscopy, which permits the observation of a single cell
layer and shows the cell zones with different pH. NR accumulated in the vacuoles of
non-infected cells, but not in vacuoles of infected cells (Figure3A, B). To verify this, we
stained two different sets of nodules (n=10-12 nodules per experiment), obtaining
the same result for each. Our results indicate that the pH of infected cell vacuoles
is not acidic. We suggest this non-acidic vacuolar pH is already established in cells
where bacteria are released from the infection thread and is maintained throughout
the development of infected cells, whereas vacuoles of neighboring uninfected cells
remain acidic. Vacuoles in infected cells often contained some rhizobia. In nodules
stained with NR, the bacteria had red fluorescence most likely due to the dye sticking
to dead bacteria (see also Discussion). This strongly suggests that NR enters the
vacuoles of infected cells, but does not change color because the pH of these vacuoles
is not acidic. The difference in pH status between non-infected and infected cells was
further verified by using the ratiometric probe LysoSensor Yellow/Blue DND-160 (Han
3
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Chapter 3
& Burgess, 2009). In its acidic form, the dye fluoresces brightly in yellow and in neutral
pH the fluorescence becomes blue. The nodules were hand sectioned, immediately
loaded with LysoSensoYellow/Blue and observed. Vacuoles of non-infected cells
emitted fluorescence corresponding to an acidic pH 5-5.5 according to the calibration
(Supplemental Figure 1A, B). In contrast, vacuoles of infected cells emitted fluorescence
corresponding to neutral pH (Supplemental Figure 1A, B). This analysis was repeated
twice with 10-12 nodules each time. 25 infected and 25 uninfected cells from zone III of
wild type nodules were analyzed for each experiment.
Figure 3. NR Staining to determine vacuolar pH in nodule cells. (A) Infection zone. (B) Fixation zone. Confocal
images display the red color of the acidotrophic dye showing acid compartments–vacuoles. Symbiosomes were
counterstained by Sytox Green. inf, infected cells; R, bacteria release; UC uninfected cells. Asterisks indicate dead
bacteria stained by NR inside the vacuole lumen. Bar in (A) =20 μm; bar in (B) = 50 μm.
The HOPS complex is temporary repressed in young nodule infected cells of the
fixation zone
The HOPS complex is essential for tethering and fusion of vesicles to vacuoles,
a process which assures the specificity of fusion (Balderhaar & Ungermann, 2013).
To determine the role of the HOPS complex in symbiotic cell development, two M.
truncatula genes encoding subunits of the HOPS complex, VPS11 and VPS39, were
analyzed.
The promoter regions (2.5kbupstream of the translation start) of VPS11 and
VPS39 were fused to β-glucuronidase (GUS) and introduced into M. truncatula roots
by Agrobacterium rhizogenes-mediated transformation. Analysis of GUS staining in
transgenic root nodules expressing ProVPS11:GUS (Figure 4A, B) or ProVPS39:GUS
(Figure 4E, F) showed that these promoters were active in the meristem, in cells of
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Adjustment of host cells vacuoles for accommodation of symbiotic bacteria
the peripheral tissues and in the infection zone. Surprisingly, the promoters were not
expressed in the infected cells at the transition of the infection to the fixation zone
(interzone 2/3 according to Vasse et al., 1990), but were expressed in the neighboring
non-infected cells (Figure 4A, E).
Figure 4. Expression of ProVPS11:GUS and ProVPS39:GUS in young (14dpi) nodules and the localization of GFPVPS11 and GFP-VPS39 in mature (35 dpi) and senescent infected cells undergoing lysis. (A) And (B) GUS staining
shows the expression of VPS11 in all cells of the apical part of nodules and only in uninfected cells in the zone of
nitrogen fixation. (C) And (G) GFP-VPS labels young infected cells during bacteria release and only uninfected
cells in the fixation zone. (D) And (H) VPS11 appears on symbiosomes in senescent infected cells where
symbiosomes are scattered in a disintegrated cytoplasm, a bacteria in a vacuole-like structure formed by the
fusion of senescent symbiosomes. VPS signal is marked by 15-nm gold particles in (H). (E) And (F) GUS staining
shows the expression of VPS39 in all cells of the apical part of nodules and only in uninfected cells in the zone of
nitrogen fixation. Membranes are contrasted by staining with FM4-64 in (C), (D), and (G). B, bacteroid; IC,
infected cell; It, infection thread; UC, uninfected cell; zII, infection zone; zIII, fixation zone. Arrowheads indicate
starch grains of infected cells in the first cell layer of the fixation zone. Bars in (A) and (D) = 75 μm; bars in (B) and
(E) = 25 μm; bars in (C), (D), and (G) = 5 μm; bar in (H) = 200 nm.
3
To check whether the repression of the HOPS complex is specific to the legumerhizobial symbiosis, we inoculated the transgenic roots expressing ProVPS11:GUS
or ProVPS39:GUS, with Glomus intraradices, fungi that form arbuscular mycorrhizal
symbiosis in the roots. VPS11 and VPS39 promoters were active in the inner root
cortical cells where the fungus forms the arbuscules (Supplemental Figure 2A,
B). The vacuoles in the cells, containing arbuscules, also retained their acidic pH
(Supplemental Figure 2C). Hence, we conclude that the suppression of HOPS does not
occur in the mycorrhizal interaction.
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The repression of VPS genes is rather unusual, as the absence of HOPS is generally
lethal (Rojo et al., 2001; Rojo et al., 2003). Therefore, we analyzed the distribution of
VPS11 andVPS39 proteins in nodules. We used nodules formed on transgenic roots
expressing ProVPS11:GFP-VPS11 and ProVPS39:GFP-VPS39 harvested 14 and 35 days
postinoculation (dpi). As VPS proteins are not abundant in the cells, the analysis
required enhancement of the GFP signal with anti-GFP antiserum. Hand sections of
transgenic nodules were analyzed by confocal microscopy. 14 dpi nodules showed
GFP-VPS signal in endosomes and the tonoplast of cells in the zone where bacteria are
released. However, the labeling was not detected in infected cells of the fixation zone,
while neighboring uninfected cells displayed the signal. VPS proteins were absent, or
their level was drastically reduced in all infected cells of the fixation zone (Figure 4C,
G). This suggests that maintenance of symbiosomes in the fixation zone correlates
with the temporary repression of the HOPS tethering complex in infected cells.
In senescing nodule cells, lytic compartments form de novo (Dupont et al., 2012).
This likely requires a HOPS complex. Confocal and immunogold labeling electron
microscopy (EM) of senescing nodules 35 dpi showed that the GFP-VPS proteins indeed
accumulate in infected cells undergoing lysis (Figure 4D, H). Reappearance of the VPS
proteins in infected cells during senescence implies that fusion of symbiosomes and
formation of lytic vacuole-like structures, typical for nodule senescence, requires the
presence of a functional HOPS tethering complex.
To verify the correct localization of GFP-VPS proteins we examined their colocalization with VTI11, a member of the vacuolar SNAREpin complex, in ProVPS11:GFPVPS11 and ProVPS39:GFP-VPS39 transgenic roots and nodules. VTI11 is involved
in membrane fusion to the tonoplast and interaction with the HOPS complex
(Balderhaar & Ungermann, 2013). Using a M. truncatula-specific anti-VTI11 antibody
(Limpens et al., 2009), we showed that VTI11 was generally co-localized with GFPtagged VPS proteins (Supplemental Figure 3A, B), confirming that GFP-VPS tagged
proteins maintain their correct localization in ProVPS11:GFP-VPS11 and ProVPS39:GFPVPS39 transgenic roots and nodules.
To validate the localization of VPS11 on the tonoplast we used a construct
whereGFP-VPS11 was expressed under control of the Ubiquitin3 promoter (ProUBQ3).
The strong expression under the UBQ3 promoter permitted clear visualization of GFPVPS11 on the tonoplast membrane (Supplemental Figure 3C).
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Functional analysis of VPS proteins
RNA interference (RNAi) was used to analyze the function of VPS. Initially the
cauliflower mosaic virus 35S promoter (Pro35S) was used to direct expression of the
constructs, but transgenic roots expressing either Pro35S:RNAi-VPS39 or Pro35S:RNAiVPS11 were not obtained. This suggested that the VPS proteins are essential for root
growth and development, confirming previous studies (Rojo et al., 2001)2001. We
therefore used the promoter of Nodulin E12, (ProE12) which is active only in nodule
primordia and in the youngest part of the infection zone of the nodule (Vijn et al., 1995),
to express the RNAi constructs. The level of silencing, as measured by quantitative RTPCR (qRT-PCR), is shown on Supplemental Figure 4A and B.
Nodules from ProE12:RNAi-VPS11 and ProE12:RNAi-VPS39 transgenic roots showed
a higher number of small non-fused vacuoles (8.12 ± 3.8 across 3 sections from
individual ProE12:RNAi-VPS39 nodules; Figure 5A, B) relative to empty vector controls
(2.40 ± 1.21 [P<0.005]; Figure 5C, D). In two separate experiments, where a total of 20
random nodules were examined, the vacuole phenotype was detected in 9 nodules.
The significant increase of non-fused vacuole numbers supports the hypothesis that
the identified VPS genes encode a part of a HOPS tethering complex that controls
homo- and heterotypic fusion of vacuoles. This raised the question whether vesicles
normally targeted to the tonoplast are mistargeted in infected cells or alternatively,
the specificity of their fusion is lost. To test this, we selected and used a tonoplast
aquaporin as a marker for vesicle targeting.
3
MtTIP1g as a membrane marker for vacuolar protein targeting in infected cells
TIPs (tonoplast intrinsic proteins) have been used as vacuolar markers in a variety
of plant species (Gattolin et al., 2009). M. truncatula TIPs were identified based on
homology with A. thaliana TIPs. Seven homologs were found in the available genomic
and cDNA sequences. As shown in Supplemental Figure 5A, all of them cluster with A.
thaliana TIPs in a phylogenetic analysis.
The predicted amino acid sequences of M. truncatula TIPs were aligned and
compared with aquaporins that previously had been functionally analyzed. The M.
truncatula TIPs share with these aquaporins six alpha-helical transmembrane domains
and five inter-helical loops (predicted by TMHMM, http://www.cbs.dtu.dk/services/
TMHMM/) and two highly conserved NPA (Asn-Pro-Ala) motifs (Forrest & Bhave,
2007) (Supplemental Dataset 1, Supplemental Figure 5B). To select a M.truncatula
TIP for the study, we first determined the expression of TIP genes in nodules (14 dpi)
compared to roots, using qRT-PCR analysis (Supplemental Figure 6).
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Figure 5. Silencing of VPS39 disturbs the formation of the central vacuole in the infected cell. (A) In the zone of
infection, infected cells have numerous small vac-uoles. (B) Magnifi- cation of (A). (C) And (D) Empty vector
control. V, vacuole. Bars in (A) and (C) = 25 μm; bars in (B) and (D) = 5 μm.
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Most examined TIP genes had low expression in nodules with the exception of
TIP1g, which displayed a high level of expression in nodules. Clustal W alignments
against other TIP aquaporins suggest that TIP1g is most likely a tonoplast intrinsic
protein. This conclusion was supported by the functional characterization of TIP1g in
Xenopus laevis oocytes.
Water permeability by TIP1g was measured using a X. laevis oocyte heterologous
expression system. We confirmed localization of TIP1g to the oocyte plasma membrane
by injecting the oocytes with YFP-tagged version of TIP1g (Supplemental Figure
7). Water permeability was measured by incubating oocytes in a five-fold diluted
ND96 solution (osmolarity 47 mOsmol kg-1). Oocyte volume change was derived
from images captured at 3 sec intervals for 2 min under a dissecting microscope.
The value of the water permeability coefficient for TIP1g expressing oocytes was
0.00413±0.00125 cms-1 while water-injected and AQP1 expressing oocytes exhibited
water permeability coefficients of 0.00096±0.00093 cms-1 and 0.00878±0.0023 cms-1,
respectively. On average 10 oocytes were tested for each trial (Supplemental Figure
8). These results suggest that TIP1g is a functional water transporter. Whole oocyte
current measurements revealed that TIP1g does not transport malate or ammonia
(Supplemental Figure 9A, B).
To determine whether TIP1g is targeted to the tonoplast we analyzed its subcellular
localization in young nodule cells. TIP1g was fused to the N-terminus of GFP and
expressed under control of either the ProUBQ3 or Leghemoglobin (ProLB) promoters.
Confocal microscopy of ProUBQ3:GFP-TIP1g expressing nodules showed that it is
located on the tonoplast (Figure 6A).Under the ProLB promoter, GFP-TIP1g signal
was found on the tonoplast and symbiosomes of infected nodule cells (Figure 6B).
Collectively these data show that TIP1g can be used as a marker to study targeting of
tonoplast-residing proteins as well as those localized to the symbiosome membrane.
3
TIP1g is retargeted to the symbiosome membrane
To examine the targeting of TIP1g in infected cells of M. truncatula nodules, a new
construct was made with the GFP-TIP1g fusion driven by the TIP1g promoter (ProTIP1g).
ProTIP1g:GUS expression analysis showed that this promoter is active in all zones of
the nodule including the fixation zone (Figure 6C). Anti-GFP antibodies were used to
enhance the GFP-TIP1g signal. Confocal microscopy of transgenic nodules expressing
ProTIP1g:GFP-TIP1g showed that TIP1g is located in the tonoplast of infected and noninfected cells in the infection zone, although the level of signal was not high (Figure
6D). The signal of GFP-TIP1g was abundant in infected cells of the fixation zone (Figure
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6D, E). In these cells it appeared that TIP1g is localized on the symbiosome membrane,
indicating this protein might be retargeted toward the symbiosomes from its default
route to the tonoplast. Electron microscopy and immunogold labeling of ProTIP1g:GFPTIP1g expressing nodules confirmed that GFP-TIP1g is present on the symbiosome
membrane (Figure 6F). According to the estimation from 5 EM frames, 80% of the
symbiosomes showed GFP labelling.
Figure 6. Localization of TIP1g. (A) Confocal image of ProUBQ3:GFP-TIP1g expressing nodules. GFP-TIP1g labels
the tonoplast of the apical nodule cells. (B) Confocal image of ProLB:GFP-TIP1g–expressing nodules. GFP-TIP1g
labels the tonoplast and symbiosome membrane in infected cells of the fixation zone. (C) TIP1g promoter
activity. Histochemical GUS staining is observed throughout the developing and fixing nodule zones on young 14
dpi nodules. (D) Confocal image of the apical part of a ProTIP1g:GFP-TIP1g–expressing nodule. GFP-TIP1g labels
the symbiosome membrane from the first cell layer of the fixation zone. (E) Magnification of the first cell layer of
the fixation zone. (F) EM immunogold labeling with anti-GFP antibody. Signal (arrowhead) is present over the
symbiosome membrane. The immunogold labeling has been quantified from five different frames, each
containing 6 to 8 symbiosomes (37 symbiosomes in total), and 85% of the symbiosomes showed the labeling. Inf,
infected cell; It, infection thread; zII, infection zone; zIII, fixation zone. Bars in (A), (B), and (D) = 20 μm; bar in (C)
= 75 μm; bar in (E) = 5 μm; bar in (F) = 200 nm.
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Silencing of TIP1g affects symbiosome development
Since TIP1g is targeted to the symbiosome, we investigated whether this is
essential for symbiosome maturation. RNAi constructs under the control of either
Pro35S or ProLB promoters were introduced into M. truncatula hairy roots. These
promoters are active in different zones of the nodule. Pro35S is active in the meristem
and in the proximal infection zone and ProLB is most active in the fixation zone, which
contains mature, N2-fixing symbiosomes, although expression already begins in the
distal part of the infection zone (Yang et al., 1991).
To check whether partial silencing of TIP1g affects nitrogen fixation in
symbiosomes, transgenic roots were inoculated with a Sinorhizobium meliloti line
expressing GFP under the control of the nifH promoter (nifH:GFP). In nodules that are
not fixing nitrogen, the rhizobial nifH gene is not induced, allowing discrimination
between Fix+ and Fix- nodules on ProLB:RNAi-TIP1g transgenic roots by analysis of GFP
fluorescence in infected cells (Supplemental Figure 10). The Fix+ and Fix- nodules (14
dpi) from transgenic ProLB:RNAi-TIP1g roots were harvested separately and analyzed
by confocal and light microscopy. In control nodules, the expression of nifH:GFP was
clearly detectable (Figure 7A) at the transition between the zone of infection and the
zone of nitrogen fixation (interzone 2/3) (Vasse et al., 1990). The first cell layer of the
fixation zone can be distinguished by the appearance of starch grains on the periphery
of the infected cells (Figure 7B). Previously this zone was named interzone 2/3 as it was
presumed that symbiosomes of this cell layers were not able to fix nitrogen (Vasse
et al., 1990). However as nifH is specifically induced at this transition, this shows
that functionally, it belongs to the fixation zone (Figure 7A, B). Fix+ ProLB:RNAi-TIP1g
transgenic nodules showed an identical pattern to the control nodules. However in
Fix- transgenic nodules, the zone of nitrogen fixation did not develop, nifH:GFP was
not detectable, and the zone of transition, which contains undeveloped symbiosomes,
expanded to 4-6 cell layers (Figure 7C). The Fix+ and Fix- transgenic nodules were
harvested separately and the expression level ofTIP1g was estimated by qRT-PCR
(Supplemental Figure 11). In Fix- nodules TIP1g expression was 5% of that in control
nodules while Fix+ nodules have expression similar to control nodules (Supplemental
Figure 11).
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Figure 7. nifH Expression is not induced in ProLB:RNAi-TIP1g transgenic nodules. (A) And (B) nifH expression is
switched on in the first cell layer of the fixation zone in control nodules. (B) is a magnification of (A); the rectangle
highlights starch grains in the infected cell of the first cell layer of the fixation zone. (C) In ProLB:RNAi-TIP1g–
expressing nodules, the zone of nitrogen fixation is not developed, such that the zone of transition, which
contains small undeveloped symbiosomes, is expanded to four to six cell layers. Hand sections of M. truncatula
nodules formed by S. meliloti (nifH:GFP) were counterstained by FM4-64. zII, infection zone; zIII, fixation zone.
Bar in= 100 μm; bars in (B) and (C) = 20 μm.
RNAi-TIP1g transgenic nodules (14 dpi) were analyzed by light microscopy. No
effect on nodule development was observed in Fix+ ProLB:RNAi-TIP1g and Pro35S:RNAiTIP1g transgenic root nodules. However, each Fix- nodule (12 nodules analyzed) from
ProLB:RNAi-TIP1g expressing roots, displayed an interruption in the development of
symbiosomes: maturation of symbiosomes from elongation stage 3 to nitrogen-fixing
stage 4 (classification of Vasse et al., 1990) did not occur (Figure 8A, B, E, F). This suggests
that TIP1g is required specifically for symbiosome maturation between stages 3 and 4.
In some nodules, small vacuoles in the nitrogen fixation zone were observed (Figure
8F). Sections of the nodules also displayed more pronounced defects, including the
distortion of plant cell turgor and the detachment of the plasma membrane from the
cell wall of symbiotic cells. These differences in the phenotype may reflect the level of
silencing of TIP1g (Figure 8B).
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Figure 8. TIP1g is required for symbiosome development and infected cell turgidity. (A), (B), (E), and (F) Light
microscopy of ProLB:RNAi-TIP1g 14 dpi nodules. Symbiosome development in transgenic nodules did not
proceed furtherthan stage 3. Detachment of the plasma membrane from the cell wall in infected cells is indicated
by arrows. Secondary release of bacteria in infected cells is indicated by arrowheads. (C), (D), (G), and (H) Electron
microscopy of ProLB:RNAi-TIP1g 14 dpi nodules. Symbiosomes reached only developmental stage 2 or 3, and
cellsshowed extreme ER proliferation (C). Detachment of plasma membrane formed a gap between the plasma
membrane and the cell wall (D). The asterisk indicates autophagic bodies containing small parts of the cytoplasm
(G). Secondary release of rhizobia into already populated cells is indicated in (H). (I) and (J) Light microscopy of
empty vector control 14 dpi nodules. (K) and (L) EM of empty vector control 14 dpi nodules. A young infected cell
populated by symbiosomes in stage 2 to 3 is shown in (K), and mature radially aligned symbiosomes are shown
in (L). The asterisk indicates starch grains. Bars in (A), (E), and (I) = 75 μm; bars in (B), (F), and (J) = 25 μm; bar in (C)
= 1 μm; bars in (D) and (G) = 0.5 μm; bars in (H), (K), and (L) = 2 μm.
3
Electron microscopy analysis of the effects of TIP1g silencing on nodule
ultrastructure showed that symbiosomes from Fix- ProLB:RNAi-TIP1g transgenic
root nodules reached only stage 2 or 3 in their maturation (Figure 8C, D), even in
nodules where plasma membrane-cell wall detachments were not observed by
light microscopy (Figure 8G, H). In most symbiotic cells proliferation of endoplasmic
reticulum (ER) occurred and the ER lumen widened. Structures similar to
autophagosomes were present in symbiotic cells (Figure 8G). These autophagosomes
were situated peripherally in the host cell cytoplasm, forming a vacuole-like structure
adjacent to the cell wall. ER membranes encased symbiosomes and parts of host
cell cytoplasm. We consider that in these cells the ER is involved in the formation
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of membranes to isolate the autophagic bodies, as was recently shown for animal
cells omegasomes (Li et al., 2012). Some infected cells displayed an electron-dense
cytoplasm that indicates premature senescence. In some cells “secondary” bacterial
release occurred, with rhizobia entering already populated cells, a process that may
be facilitated by low turgor pressure and gaps between the cell wall and the plasma
membrane. Bacteria released into these cells have to be considered saprophytic and
not symbiotic, as the cell was already undergoing lytic clearance (Figure 8H). Since
symbiosomes in the RNAi nodules did not reach functional maturity as in control
nodules (Figure 8I-L), they were not able to fix atmospheric nitrogen and the symbiosis
appears to be terminated prematurely.
Discussion
We report here that host cell architecture and vacuole formation are modified in
nodule infected cells. In root nodules in the transition from the zone of infection to the
zone of nitrogen fixation several vacuolar changes occur. The host cell vacuole volume
is reduced by 75% and this is accompanied by tonoplast folding. The HOPS complex
in infected cells is temporarily repressed and the integral tonoplast aquaporin TIP1g
accumulates at relatively high levels and is retargeted to the symbiosomes. The
collapse of the vacuoles in this transition may permit the expansion of bacteria in the
host cell cytoplasm at the expense of vacuole volume.
The cell volume change related to the modulation of the vacuole volume is not
uncommon for plant cells. This process is best studied in guard cells. Stomatal opening
is partly caused by vacuolar convolution (Zouhar & Rojo, 2009; Bak et al., 2013). The
conditions that induce stomatal closing result in fragmentation of the central vacuole
and loss of guard cell turgor. Loss of vacuole volume and tonoplast folding have also
been described in cells exposed to hyper-osmotic stress (Reisen et al., 2005; Brett
& Merz, 2008). Hence, the cell architecture remodeling, which is mediated by the
change in volume, may be a common reaction during plant development. Vermeer et
al. (2014) showed that volume loss in endodermal cells, and vacuole remodeling via
fragmentation, take place during lateral root formation in A. thaliana. They considered
this to be a coordinated response reflecting the mechanical stresses at neighboring
cell layers, which might affect tissue and organ patterning.
The collapse of the vacuole in the transition zone of M. truncatula root nodules
is most likely due to the combination of suppression of the HOPS complex, high
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water demand from proliferating bacteria and a functionally compromised nonacidic vacuole. It may also reflect the reaction of nodule cells to mechanical stress
due to rhizobial expansion. We assume that the pathway of vacuole formation in
infected nodule cells is modified and traffic to the tonoplast is impaired. This is
confirmed by the accumulation of TIP1g and its retargeting to the symbiosome
membrane in the transition between the infection and fixation zones. It is interesting
to note that increase of TIP protein content and the retargeting of TIP aquaporin
to other endomembranes from the tonoplast was previously reported as a result
of osmotic stress in leaves of Mesembryanthemum crystallinum (Vera-Estrella et al.,
2004). In this case, it was shown to be a developmental adaptation by which cells
are preprogrammed for such a response. By analogy, symbiotic cells may also have
evolved such reprogramming mechanisms.
To clarify the role of the HOPS tethering complex during the development of the
symbiosis we studied VPS11 and VPS39, which encode putative HOPS subunits. The
expression of both VPS11 and VPS39 was temporary switched off at the transition of the
infection to the fixation zone and the proteins were not detected in infected cells of the
fixation zone, while neighboring uninfected cells display the signal. The mechanism
for this quite specific block of expression remains to be elucidated, but the analysis
of the expression of VPS11 and VPS39 in arbuscular mycorrhizal symbiosis shows that
this block is nodule specific. Functional silencing of VPS11 and VPS39 in the infection
zone results in infected cells that have an increased number of small non-fused
vacuoles instead of a large central vacuole. Further, both VPS proteins are localized on
endosomes and tonoplast membranes. We considered that the localization of VPS11
and VPS39 is similar to localization reported for VPS proteins in yeast and A. thaliana
(Rojo et al., 2003; Balderhaar & Ungermann, 2013). We did not find GFP-tagged VPS
proteins on symbiosomes during the infection stage, despite the presence of the
protein in the cell. The explanation is that the identity of young symbiosomes is
similar to that of the plasma membrane (Ivanov et al., 2012) and the vacuolar identity
is acquired at later stages (Limpens et al., 2009). In the zone of fixation, VPS proteins
are not detected. However, during the termination of symbiosis and the senescence of
infected cells VPS proteins accumulate. In these cells symbiosomes lose the ability to
fix nitrogen and are transformed into lytic compartments that fuse and form vacuolelike units (Van de Velde et al., 2006; Dupont et al., 2012). These results show that the
vacuolar fusion machinery controls symbiosome lysis.
The retargeting of TIP1g indicates a loss of specificity of tonoplast-targeted
vesicle fusion, probably due to the absence of the HOPS complex. The TIP1g RNAi
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experiments indicate that this retargeting is essential for symbiosome expansion and
maturation to the nitrogen-fixing stage, as partial silencing of TIP1g causes a block in
symbiosome maturation. The presence of TIP1g on symbiosomes might have a direct
function in symbiosome maturation, for example by enhancing the availability of
water. A similar block in symbiosome development was observed after silencing of
the small GTPase Rab7, a non-integral tonoplast protein involved in membrane fusion
(Limpens et al., 2009). This underlines the importance of retargeting of tonoplast
proteins to symbiosomes.
In TIP1g RNAi experiments, the nodules with high levels of silencing (95%) are Fix-.
In these nodules the symbiosis is not functional, and the symbiosomes do not mature
to reach the nitrogen-fixing stage. In the majority of infected cells of TIP1g:RNAi,
nodules in which nifH expression is blocked, a massive burst of autophagy occurs.
Macroautophagy involves the formation of autophagosomes, double-membrane
structures that sequester part of the cytoplasm or organelles (Bassham et al., 2006;
Avila-Ospina et al., 2014). The mechanisms for the formation of the autophagosome
isolation membrane are not completely understood, but its formation always
begins with the ER surrounding the part of the cytoplasm that later appears inside
the autophagosome. Recently, Uemura et al. (2014) suggested the mechanisms for
transformation of ER to autophagic isolation membranes. In TIP1g:RNAi expressing
infected cells we have observed the formation of autophagic bodies, and massive
proliferation of ER, surrounding the symbiosomesand part of the cytoplasm. Therefore
it is quite well possible, that formation of the isolation membrane of autophagic
bodies from ER in the nodules is similar to the process described for animal cells (Li et
al., 2012; Uemura et al., 2014).
The spatial and temporal pattern of autophagic body formation in TIP1g:RNAi
nodules is quite different from macroautophagy events in wild type nodules. In wild
type nodules the formation o fautophagic bodies occurs in young infected cells and
in senescing cells, but not in efficient nitrogen-fixing zone. In these cells some parts
of the cytoplasm or small freshly released symbiosomes, being in close proximity
with the vacuole, become engulfed in the vacuole lumen (Fedorova & Brown, 2007).
In some cases, such as nodules formed on the DNF1 –“defective in nitrogen fixation”
mutant, where symbiosomes do not mature, autophagic bodies become quite
numerous (Wang et al., 2010)2010, but do not specifically entrap symbiosomes.
In wild type root nodules, the quick expansion of the microsymbiont helps to
maintain the tight contact of the plasma membrane with the cell wall even after
the collapse of the vacuole. However in TIP1g:RNAi infected cells when symbiosome
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growth and maturation was blocked, the plasma membrane contact with the cell wall
was impaired. It is possible that this may cause starvation of these cells. The arrest of
symbiosome development and their consequent lysis might also be partly due to the
starvation.
In other plants, for example in A. thaliana, single and multiple knockouts of TIP
aquaporins are not lethal (Wudick et al., 2009). The severe phenotype, which we
observed in TIP1g:RNAi nodules, is most likely due to the precarious situation of
symbiotic cells, which lack a functional vacuole but at the same time are “burdened”
by thousands of symbiosomes. This phenotype suggests that infected cells do not
have a robust mechanism to maintain contact between the plasma membrane and
the cell wall and thus may be easily subjected to disruption of water transport from
the apoplast to the infected cell.
The presence of tonoplast or late endosome proteins on symbiosomes raises the
question of how they can fuse to this membrane in the absence of a HOPS tethering
complex. We showed that VPS11, which encodes the core subunit of the HOPS
complex, is temporary repressed in the first cell layer of the fixation zone. This subunit
is shared with another tethering complex, CORVET (Balderhaar & Ungermann,
2013). Therefore, it is unlikely that CORVET can replace the HOPS complex in the
symbiotic cells of the fixation zone. How the fusion of vesicles is regulated in mature
symbiosomesin the zone of fixation remains to be solved.
The non-acidic pH of the vacuole in the symbiotic cells reduces its lytic properties
and may affect endocytotic trafficking (Dettmer et al., 2006). The alteration in
endocytosis probably suppresses fusion of symbiosomeswith themselves, as well
as with other endosomes and young vacuoles. This is similar to the suppression of
fusion of bacteria-containing vacuoles with lysosomes in mammalian cells as this
also involves deacidification of phagosomes as well as lysosomes (Huynh & Grinstein,
2007; von Bargen et al., 2009). A further consequence of an increased vacuolar pH, is
that it may lead to impaired malate uptake by the vacuole, as the inward transport
is dependent on the acidic pH of the vacuolar lumen (Hurth et al., 2005; Etienne et
al., 2013). Malate is a primary carbon source for symbiosomes (White et al., 2007),
thus vacuole deacidification may boost the availability of malate for symbiosomes
by negatively affecting the transport of malate into the vacuole. The mechanism
by which the vacuolar pH is increased remains to be determined, but it seems that
a V-ATPase, regulating vacuolar pH, could be involved (Li et al., 2010; Schnitzer et al.,
2011; Tarsio et al., 2011).
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In conclusion, we report here that infected cell adjustment to accommodate
rhizobia involves the suppression and defunctionalization of its vacuole and
retargeting of some tonoplast proteins to symbiosomes. The temporary suppression
of the HOPS tethering complex is most likely part of the mechanism that facilitates
symbiosome expansion and maintenance as nitrogen fixing organelles.
Materials and Methods
Plant materials, transformation and inoculation.
Agrobacterium rhizogenes–based root transformation of M. truncatula Jemalong
A17 was performed according to Limpens et al. (2005). Roots were inoculated by S.
meliloti 2011 and with S. meliloti expressing nifH:GFP. M. truncatula plants that were
inoculated with G. intraradices were co-cultivated with Allium schoenoprasum nurse
plants in a sand/hydrobead mixture saturated with Hoagland medium according to
Ivanov et al. (2012).
Cloning
VPS11 and VPS39 open reading frames and their 2.5 kb 5′ regulatory sequence
were amplified via PCR from 7 dpi nodule cDNA and genomic DNA respectively
using Phusion highfidelity polymerase (Finnzymes). Coding sequences of VPS11 and
VPS39 were directionally cloned with SmaI –KpnI and KpnI-BamHI, respectively, into
a modified pENTR vector (pENTR2) containing a multiplecloning site. Entry clones
for VPS11 and VPS39 promoters were generated by TOPOcloning (Invitrogen). The
Gateway cloning system (Invitrogen) was used to create genetic constructs for RNA
interference, promoter-GUS and GFP-fusion. pENTR clones of VPS11 and VPS39 were
recombined into the following destination vectors using LR Clonase (Invitrogen): pE12pK7WGF2-R (containing the M. truncatula ENOD12 promoter) (Limpens etal., 2009),
pKGW-GGRR, UBQ3-pK7WGF2-R, creating N-terminal GFP-X fusions. GFP-VPS11and
GFP-VPS39 translational fusions driven by their own promoters were generated by
multiple cloning of gene and promoter sequences into pKGW-MGW.
The TIP1g open reading frame and its 2kb 5′ regulatory sequence were amplified
via PCR from 7 dpi nodule cDNA and genomic DNA respectively using Phusion
high fidelity polymerase (Finnzymes). TIP1g pENTR clones were recombined into
the following destination vectors using LR Clonase (Invitrogen): pLBpK7WGF2-R
(containing the Pisum sativum LB promoter) (Limpens et al., 2009), pKGWGGRR,
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UBQ3-pK7WGF2-R and LB-pK7WGF2-R, creating N-terminal GFP-X fusions. GFP-TIP1g
translational fusion driven its by own promoter was generated by multiple cloning of
gene and promoter sequences into pKGW-MGW. Primers are listed in Supplemental
Table 1.
GUS staining, sectioning and light microscopy
Transgenic roots and nodules were collected and washed twice in 0.1 M sodium
phosphate buffer (pH 7.2), incubated in β-glucuronidase buffer under vacuum
at room temperature for 30 minutes to allow the buffer to replace oxygen in the
tissue, incubated at 37°C for 2 hours or overnight to enable the enzymatic reaction
and embedded for sectioning following the Technovit 7100 protocol (Technovit).
Sections were mounted on microscope slides, counterstained with ruthenium red and
analyzed using a Nikon Optiphot-2 and Leica DM 5500 Flu microscopes.
The selection of transgenic roots and nodules was performed on a Leica MZFLIII
fluorescent macroscope equipped with filter cubes for the detection of GFP, YFP, RFP,
excitation/emission EGFP (Ex: 470/40 D: 495 Em: 525/50); EYFP (Ex: 510/20 D: 530
Em:560/40); DsRED (Ex: 545/30 D: 570 Em: 620/60).
3
Confocal microscopy
Confocal imaging of GFP-fused proteins was done on transgenic hand sectioned
nodules by using a Zeiss LSM 5 Pascal confocal laser-scanning microscope (Carl
Zeiss) and Zeiss Meta LSM 510 microscope. For enhancement of GFP-VPS11, GFPVPS39 and GFP-TIP1g signal driven by their own promoters we used polyclonal rabbit
anti-GFP antibody (Molecular Probes) (1:100 dilution) and secondary anti-rabbit
Alexa 488 antibody (Molecular Probes) (1:200 dilution), normal goat serum or 3%
BSA was used for blocking. Sections were counterstained with FM4-64 (30 μg/mL)
or Propidium Iodide (PI). In cases where the constructs were created using strong
promoters like ubiquitin, the localization was observed without enhancement. For
immunolocalization of VTI11 specific M. truncatula anti-VTI11 antibodies developed
in rabbit (diluted 1:100) were used (Limpens et al., 2009), and as a secondary an
anti-rabbit CY3 Ab (Molecular Probes) (diluted 1:200). To enhance the GFP signal in
this experiment anti-GFP antibodies developed in mouse (1:50) were used, followed
by secondary anti-mouse Alexa 488 (1:200). A mix of 0.5% skim milk powder with
2% BSA was used as a blocking solution. The protocol for Neutral Red staining was
adapted from Dubrovsky et al. (2006). Confocal microscopy settings with the window
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for NR excitation/emission (564/595–615nm) were used. Bacteroids were contrasted
by staining with SYTO 16 (green).
3D Reconstruction and volume/surface measurements
3D reconstruction of Z-stacks obtained by confocal microscopy of root nodules
expressing GFP-SYP22 driven by ProUBQ3 or ProLB promoters or wild type root
nodules counterstained by Sytox green (Molecular Probes) was used for quantitative
estimation of the volume and surface area of the vacuole, bacteroids, and cell.
Bacteroids and nuclei were counterstained either by PI (red fluorescence) or Sytox
green. To obtain 3D reconstructions, confocal image stacks (50 images with 0.5 μm
of z-step) were imported to Imaris 7.5 (Bitplane Inc.). After baseline subtraction, a
sub-region with a cell of interest was defined. The isosurface module was used to
reconstitute the 3D pictures. Volume and surface area measurements were performed
using the Imaris Measurement Pro module.
Sample preparation for light and electron microscopy and EM immunodetection.
The protocol for tissue processing was described previously (Limpens et al., 2009).
Semithin sections (0.6 μm) for light microscopy and thin sections (60 nm) for EM
of transgenic nodules were cut using a Leica Ultracut microtome. Nickel grids with
the sections were blocked in normal goat serum with 1% milk or 2% BSA in PBS and
incubated with the primary antibody at the dilutions given above. Goat anti-rabbit
antibody coupled with 10-nm gold (BioCell) (1:50 dilution) were used as secondary
antibody. Sections were examined using a JEOL JEM 2100 transmission electron
microscope equipped with a Gatan US4000 4K×4K camera.
Expression in Xenopus laevis Oocytes
The cDNA of TIP1g was cloned into the expression vector pGEMHE using the
restriction enzymes Bam HI and Xba I. To linearize the plasmid, pGEMHE-TIP1g was
digested with Nhe I. Complementary RNA was transcribed using 1 μg of linearised
DNA with the mMESSAGE mMachine Kit (Ambion). X. laevis oocytes were surgically
removed and defolliculated, then injected with 30 ng cRNA or water, using a microinjector (Drummond‘Nanoject II’ automatic nanolitre injector, Drummond Scientific,
Broomall, PA, USA). After injection, oocytes were incubated in ND96 for 48 h at 18°C.
To measure water permeability, oocytes were transferred to a 5-fold diluted solution
of ND96 (osmolarity 47 mOsmol kg-1). The volume change in the oocyte was derived
from images captured at 3 sec intervals for 2 min under a dissecting microscope with
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IC Capture 2.0 software (The Imaging Source, US) as AVI format video files. Image J
software (http://rsbweb.nih.gov/ij/, United States Government) was used to calculate
the change in the total area of the oocytes captured in the AVI video file. The rate of
oocyte swelling was plotted as V/V0 versus time, where V is a volume at a certain time
point and V0 is the initial volume. Water permeability coefficient values for oocytes
injected with cRNA encoding TIP1g, AQP1 or water were determined as described
by Fetter et al. (2004). The significance of results was analyzed by Tukey’s multiple
comparison test.
qRT-PCR Analysis.
Total RNA was extracted from roots and different zones of 14 dpi root nodules
using E.Z.N.A. Plant RNA Mini Kit (Omega Bio-Tek) and transcribed into cDNA using
the iScript cDNA synthesis kit (Bio-Rad). Real-time PCR was set up in a 20 μL reaction
system using iQSYBR Green Supermix (Bio-Rad). Gene-specific primers that were
designed with Primer-3-Plus software (Untergasser et al., 2007). Gene expression
profiles were normalized against the transcription level of the reference gene UBQ10.
Calibration of Lysosensor Yellow/Blue (Molecular Probes) was performed in
capillaries filled with calibration buffers, which contain Lysosensor Yellow/Blue (1:50
v/v) and 50 mM phosphate buffer adjusted to a pH scale from 3 to 8. The images
were taken with the filter cube A (DAPI). Filter cube A UV excitation filter 340-380,
dichromatic mirror 400 suppression filter LP 425.
3
Targeting of TIP1g in X. laevis oocytes
To confirm targeting of TIP1g to the plasma membrane of X. laevis oocytes the
vector pT7:YFP:TIP1 was generated. YFP-TIP1g localization was observed by confocal
microscopy. Oocytes were stained by the dye FM4-64 for plasma membrane
visualization.
NH3 flux experiment
To see whether MtTIP1g is a NH3 transporter, an NH3 flux experiment was
performed. TIP1g expressing oocytes were incubated in 10 mM N15-labeled NH4+
solution, pH 6.5, 7.5 or 8.5.
C14 malate flux
TIP1g injected oocytes were incubated in standard Ringers solution for two days.
The oocytes were then transferred to a solution containing 5 mM malic acid (C14-
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labeled), 0.15 mM CaCl2, 1 mM KCl, 5 mM MES and 230 mM mannitol, pH 7.5. After 2h
incubation, oocytes were harvested and analyzed by a scintillation counter. Data was
normalized by two blank solution readings.
Accession numbers
Sequence data can be found in the Phytozome and GenBank website and
database under the following accession numbers: VPS11, Medtr1g019500; VPS39,
Medtr5g020140; TIP1g, Medtr4g063090.
Acknowledgments
Authors thank Prof. T.W.J. Gadella (University of Amsterdam) for the help with
analysis of Imaris 3D reconstructed images. Authors are very grateful for Dr. P. Smith
(University of Sydney) for the help in editing the manuscript. We are thankful for our
colleagues: Norbert de Ruijter for his assistance with confocal imaging, Rene Geurts
for helpful discussions and Jan Hontelez for providing the line of S. meliloti expressing
nifH:GFP. A. G. received a PhD fellowship from EPS School of Biological Sciences
(Wageningen University). Authors declare that there is no conflict of interest.
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Supplemental materials
Supplemental Figure 1. Lysosensor Yellow/Blue staining to determine vacuolar pH in nodule cells. IC, infected
cells; UC, uninfected cell; (*) vacuole.
Supplemental Figure 2. The expression of ProVPS11:GUS (A) and ProVPS39:GUS (B) in transgenic roots inoculated
by G. intraradices. Neutral Red staining of the vacuole in the arbuscule (asterics) containing cell. H, hyphae.
Supplemental Figure 3. Co-localization of endosome/vacuole molecular marker VTI11 with VPS proteins in
ProVPS11:GFP-VPS11 (A) and ProVPS11:GFP-VPS39 (B) transgenic nodules. Arrow, tonoplast labelled by GFP.
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Supplemental Figure 4. The level of RNAi silencing of VPS11 (A) and VPS39 (B).
3
Supplemental Figure 5. ( A ) Phylogenetic comparison of transporters from M. truncatula and A. thaliana.
Phylogenetic analyses (bootstrap values of 500 replicates) were conducted using MEGA version 5. (B) ClastalW
alignment of amino acid sequence of M. truncatula aquaporins with another studied water transporters.
Transmembrane domains are shown with a line below the alignment; triangles indicate the NPA selectivity filter.
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Supplemental Figure 6.Expression of M. truncatula TIP genes in roots and different zones of 14 day old nodules
(14-1, meristem and infection zone, 14-2, fixation zone). Expression level was determined by qRT-PCR and
normalized against transcription level of reference gene M. truncatula UBQ10.
Supplemental Figure 7. YFP-TIP1g localizes on plasma membrane of injected oocytes. Confocal image of
expressing YFP-TIP1g oocytes. Plasma membrane is counterstained withFM4-64 (A). Signal overlapping in
region of interest 1 (ROI1) (B). Water permeability coefficient values for oocytes injected with cRNA encoding
TIP1g and YFP-TIP1g(C).
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Supplemental Figure 8. Water-channelactivityof TIP1g. (A) The relative volume of oocytes injected with cRNA
encoding TIP1g, human AQP1 or water following exposure to hypotonic media. The rate of oocytes swelling is
plotted as V/V0 versus time, where V is a volume at the certain time point and V0 is the initial volume. (B) Water
permeability coefficient values for oocytes injected with cRNA encoding TIP1g, AQP1or water. The difference was
significant, that show that TIP1g is a functional water transporter.
A B 3
Supplemental Figure 9. TIP1g does not transport ammonia and malate. (A) NH3 flux experiment. 15N content of
TIP1g injected oocytes were same with water control in all pH tested. This result suggests that TIP1g is not a NH3
transporter. (B) Malate flux experiment. TIP1g injected oocytes, in comparison with water control, show no
significant differences. TIP1g does not transport malate.
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Supplemental Figure 10. The induction of the rhizobial nifH gene, detectable due to GFP fluorescence, permits
discrimination between Fix+ and Fix- nodules on ProLB:RNAi-TIP1g transgenic roots. (A-C) Fix+ transgenic nodules
(TrFix+) and Fix+ non-transgenic (Ntr) nodules. (D-F) Fix- transgenic nodules (TrFix-) and Fix+ non-transgenic (Ntr)
nodules. (A) And (D) transgenic roots were detected by DsRed emission. (C) And (F) Rhizobial nifH expression
was detected by GFP emission. (B) And (E) merged image obtained by using YFP chanel. Bars: 2mm.
Supplemental Figure 11. The levelofTIP1g gene silencing in Fix+ and Fix- nodules harvested from ProLB:RNAi-TIP1g
transgenic roots in comparison with an empty vector control. The difference between the control and Fixnodules is significant (P<0.05).
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Supplemental Table 1. Primers used for cloning and qRT-PCR analysis
Gene specific primers for cloning
VPS11-F TCCCCGGGATGTATCAATGGCGGAAGTT
VPS11-R GGGGTACCTCAGAAG-CCACTGCTAGATGAT
VPS39-F CGGGATCCATGGTGCACAGTGCGTACG
VPS39-R GGGGTACCTCATCGCTTCCTCAACTGA
ProVPS11- F CACCTATTCAAATTGAAAAAACACAGAATATT
ProVPS11- R CGCCGCCGCGAT
ProVPS39- F CACCAAGGATTCAAACCCCGATCA
ProVPS39- R TTTGGTTACGAAATA-TTGAAGTTGA
TIP1g-F CACCATGCCGATTTCTAGAATTGCA
TIP1g-R TTAATAATCCGTGACAGGTAACTG
ProTIP1g- F CACCGCTTGTCTTGATTTCATGGATTG
ProTIP1g- R TGTTTTATATTTTTTCTTTTCTCAAAGA
3
qRT-PCR primers
VPS11- F TCAAGCAACGCAACTTCCTG
VPS11-R TCAGGCACAAAGCTGATTGC
VPS39-F AATCTACTCGCCGGAAACAG
VPS39-R TGACACAACAGGCTTCTTCG
TIP1a-F ATTTACGCTGCAAGGGACAC
TIP1a- R CACATGCAGGGTTGATTGAC
TIP1b-F CGTTGACCCAAAGAAGGGTA
TIP1b-R AAGTCCAGCAATTCCACCAC
TIP1c-F GTCACCTTTGGATTGGCTGT
TIP1c-R TCAGCTGCTGTGGCATAAAC
TIP1d-F TTTTGTGTTCGCTGGAGTTG
TIP1d-R ACAGCTGGGTTCAAATGTCC
TIP1e-F CCCAAAGAAAGGAG-CACTTG
TIP1e-R CCAACCCAATAAACCCAATG
TIP1f-F GGTTCCATAG-TGGCATGCTT
TIP1f-R AGCTGCTGTGGCATACACTG
TIP1g-F AACACCAG-CAGGGTTGGTAG
TIP1g-R CAAGCAACTGAGCAATCCAA
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ARP2/3-Mediated actin nucleation
associated with symbiosome
membrane is essential for the
development of symbiosomes in
infected cells of Medicago truncatula
root nodules
Aleksandr Gavrin1, Veerle Jansen1, Sergey Ivanov2, Ton Bisseling1,3 and Elena Fedorova1
1
Laboratory of Molecular Biology, Graduate School Experimental Plant Sciences, Wageningen
University, 6708 PB, The Netherlands; 2Boyce Thompson Institute for Plant Research, Tower Road,
Ithaca, NY 14853, USA; 3College of Science, King Saud University, Riyadh 11451, Saudi Arabia
Published in Molecular Plant-Microbe Interactions 2015
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Abstract
The nitrogen-fixing rhizobia in the symbiotic infected cells of root nodules are
kept in membrane compartments derived from the host cell plasma membrane,
forming what are known as symbiosomes. These are maintained as individual units,
with mature symbiosomes having a specific radial position in the host cell cytoplasm.
The mechanisms that adapt the host cell architecture to accommodate intracellular
bacteria are not clear. The intracellular organization of any cell depends heavily on
the actin cytoskeleton. Dynamic rearrangement of the actin cytoskeleton is crucial
for cytoplasm organization and intracellular trafficking of vesicles and organelles.
A key component of the actin cytoskeleton rearrangement is ARP2/3 complex,
which nucleates new actin filaments and forms branched actin networks. To clarify
the role of ARP2/3 complex in the development of infected cells and symbiosomes
we analyzed the pattern of actin microfilaments and the functional role of ARP3
in Medicago truncatula root nodules. In infected cells, ARP3 protein and actin were
spatially associated with maturing symbiosomes. Partial ARP3 silencing causes
defects in symbiosome development; in particular, ARP3 silencing disrupts the final
differentiation steps in functional maturation into nitrogen-fixing units.
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Introduction
Soil bacteria, collectively known as rhizobia, establish symbiotic relationships with
the leguminous plants. Within root nodules, the bacteria are hosted in specialized
cells in the compartments called symbiosomes. The membrane of symbiosomes is
derived from the host cell plasma membrane. The symbiosis between legume and
rhizobia represents a striking example of a prolonged intracellular lifestyle of bacteria
in plant cells inside specialized membrane compartments. However, the mechanisms
by which the architecture of symbiotic cells is modified to accommodate intracellular
bacteria are not clear.
The intracellular architecture of plant cells relies heavily on the cytoskeleton and
particularly on the actin cytoskeleton. Actin microfilament rearrangement is linked to
the positioning of organelles and influences cell shape as well as providing a roadway
for the transport of membrane vesicles (van der Honing et al., 2007; Pollard & Cooper,
2009; Yanagisawa et al., 2013). The change of actin patterns is largely dependent on
actin filament assembly and growth, which is in turn a function of actin nucleation
and polymerization (Avisar et al., 2009; Sinclair et al., 2009). Actin monomers
(globular G-actin) are polymerized into filaments with a characteristic architecture
and polarity. The rate-limiting step for polymerization is the creation of trimeric actin
seeds, a process known as nucleation (Pollard, 2007). Two complexes regulate this
process: proteins of the formin group and the actin–related proteins 2/3 (ARP2/3)
complex. The formation of straight filaments de novo depends on formin (Deeks et al.,
2002; Dominguez, 2009; Aspenström, 2010). The ARP2/3 complex creates nucleation
points on the sides of existing actin filaments, leading to branched networks (Pollard,
2007). The ARP2/3 complex consists of seven subunits: two actin-related proteins,
ARP2 and ARP3, stabilized in an inactive state by five other subunits (ARPC1, ARPC2,
ARPC3, ARPC4, and ARPC5). The complex has numerous nucleation promoting
factors (NPFs), which have been intensively studied in animals and yeasts (Pollard
& Cooper, 2009; Campellone & Welch, 2010; Rotty et al., 2013). For enteropathogenic
bacteria in animal cells, a common strategy for the establishment of an intracellular
lifestyle is the manipulation of host cell actin via the ARP2/3 actin nucleating complex
(Welch & Way, 2013).
In plants the homologues of ARP3 detected by heterologous antibodies in tobacco
and maize were localized along cortical actin filaments and to multivesicular bodies
(Van Gestel et al., 2003). The activation of ARP2/3 in most plant species is dependent
on the WAVE (WASP family verprolin homologous)/SCARs (suppressor of cAMP
receptors) family of NPFs (Yanagisawa et al., 2013).
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The role of actin and ARP2/3 in rhizobium-legume symbiosis has been mainly
studied at the early stages of the interaction. This showed for example that Nod factors,
signal molecules from rhizobia, cause the deformation and the fragmentation of a
fine mesh of actin in root hairs (de Ruijter et al., 1999; Blancaflor et al., 2006; Timmers,
2008). The importance of the ARP2/3 pathway for symbiosis was shown recently
(Yokota et al., 2009; Miyahara et al., 2010; Hossain et al., 2012). For example, Medicago
truncatula ortholog of NAP1, a component of the SCAR/WAVE complex, is operational
for the infection threads growth on the early stages of symbiosis (Miyahara et al.,
2010). Exposure of root hairs of Phaseolus vulgaris to Rhizobium etli Nod factors resulted
in a rapid increase in the number of F-actin newly available plus ends, which probably
represent sites of polymerization and in the re-localization of F-actin plus ends to
infection thread initiation sites (Zepeda et al., 2014).
However, the mechanisms of actin reformation in the intracellular stage of
symbiosis - when the rhizobia become nitrogen-fixing organelles - were not examined,
despite the documented changes in microfilament organization within the infected
cells of root nodules (Whitehead et al., 1998; Davidson & Newcomb, 2001; Fedorova et
al., 2007).
The aim of this work was to study the role of the actin nucleating factor ARP3 in
the intracellular accommodation and development of symbiosomes in M. truncatula
nodules. We found that during the transition from an immature stage of development
to the stage of nitrogen fixation, the actin pattern in infected cell undergoes prominent
reorganization. The actin microfilaments in the cytoplasm of the host cell form the
network around the mature symbiosomes. Silencing of ARP3 negatively affects the
symbiosome maturation.
Results
The actin configuration changes during intracellular accommodation of rhizobia
To determine the role of actin in the host cell development, we first visualized
filamentous actin (F-actin) in root nodules using phalloidin with a fluorescent
tag. Medicago nodules (14 and 21 dpi) were fixed and hand-sectioned along the
longitudinal axis. These sections were incubated with phalloidin (see Materials and
Methods) and subsequently analyzed by confocal microscopy. The nodule meristem
displays a typical root meristem cell actin configuration: in interphase cells, there is
a fine F-actin network in the cortex and a radial array of actin cables tethering the
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nucleus (Figure 1A). In differentiated uninfected cells, prominent thick actin cables
were located in cytoplasmic strands across the vacuoles, forming a network around
the nucleus (Figure 1B). These transvacuolar strands interconnect the perinuclear and
cortical cytoplasm (Ketelaar et al., 2010). This F-actin organization is very similar to
that of other differentiated plant cells with an isodiametric type of growth. In new
infected cells in which rhizobia colonization is just starting, the actin microfilaments
were associated with infection threads but not with freshly released or dividing
bacteria (Figure 1C, D). However at the transition from colonization to functional
maturation, actin microfilament pattern in infected cells changes markedly (Figure
1D). In latest stages of symbiosis in the cells containing growing symbiosomes, we
observed the formation of a net of fine actin microfilaments around symbiosomes
(Figure 1E, F). The final step of functional symbiosome differentiation coincides with
a radial alignment of symbiosomes in the host cell cytoplasm. The mechanisms that
may be controlling this change are discussed below.
ARP3 is localized in the cytoplasm surrounding the symbiosomes
To determine the role of the actin pattern change that causes the formation
of an actin net around the symbiosomes, we first needed the information about
the underlying mechanism that might affect the actin network. To study whether
the ARP2/3 pathway is involved in the formation of the fine F-actin surrounding
symbiosomes, we focused on the ARP3 subunit. Medicago has a single ARP3 gene
(Medtr8g089630.1). Its expression was studied by qRT-PCR in roots and nodules 7, 14,
and 21 days post inoculation (dpi).
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Figure 1. Distribution of actin (A-F) visualized by phalloidin staining (green fluorescence) and localization of
ARP2/3 (G-I) in infected cells of Medicago root nodules (green fluorescence). Bacteroids and host cell nuclei
counterstained by propidium iodide (red fluorescence). A, zonation of the root nodule, the dividing cell from the
meristem is given as insertion; B, actin network over the vacuole, the arrows indicate the vacuole; C, Infection
thread encompassed by actin filaments; D, a host cell filled with growing symbiosomesof stage 2 (Vasse et al.,
1990) highlighted by rectangle. Note that the microfilaments in the host cytoplasm are not yet organized into a
net surrounding the symbiosomes, the symbiosomes do not have a radial pattern yet; E and F, an infected cell
housing symbiosomesin stage 3/4, note the radial pattern of symbiosome sand the similar pattern in actin
microfilament distribution; F, an image displays only the green channel to show the actin pattern; G, localization
of ARP3 in infected cells of GFP-ARP3 expressing nodules (non-counterstained DIC image). The insertion is
displaying the labelling of plasma membrane (PM) which is visible in the cells where part of the cytoplasm is not
in the focal plane (arrowheads); H, ARP3 immunolocalization using anti-ARP3 antibody, the signal for ARP3 is
present in the cytoplasm of mature infected cells around the SBs and in uninfected cells cytoplasm. Note the
radial pattern of mature symbiosomes in infected cell; I, an image displays only the red channel to show the
symbiosomes arrangement. M, meristem; zII, infection zone; zIII, zone of nitrogen fixation; V, vacuole; IC,
infected cell; UnC, uninfected cell; IT, infection thread; F, freshly released bacteria; an asterisk (*) indicates
dividing cells. Bars: 20µm.
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Figure 2. Expression level of ARP3 and Western blot analysis of nodule extracts. A, ARP3 expression in root
nodules at different age, fold expression relative to uninoculated roots. Error bars represent standard deviation.
The difference between the expression of 7dpi nodules versus 21 dpi nodules is significant (P<0.05); B, distribution
of normalized RNA-seq reads (%) for Medicago ARP3 (probe Mt0009_10239) in different nodule zones: FI,
meristematic zone; FIId, distal infection zone; FIIp, proximal infection zone; IZ, interzone II/III; ZIII, fixation zone
(obtained by using the Symbimix database (INRA, https://iant.toulouse.inra.fr/symbimics/). C, Western blot
analysis of extract fromGFP-ARP3 (line 1) and wild type nodules (line 2) probed with anti-ARP3 antibody shows
the bands of expected size: (#) band of 73kD – GFP-tagged ARP3, (*) band of 46 kD - endogenous ARP3; D, the
level of silencing estimated by qPCR analysis of ARP3 expression using a template of RNA extracted from empty
vector control and RNAi-ARP3 nodules. The difference between the control and RNAi-ARP3 nodules is significant
(P<0.01). Error bars represent standard deviation.
4
The expression level of ARP3 is higher in mature 21 dpi nodules in comparison
with primordia/young 7dpi nodules. This points to a role in later stages of symbiosis
(Figure 2A). To gain an insight to the expression of ARP3 in developmental zones of
the nodule we have used the database of Symbimix (INRA, https://iant.toulouse.inra.
fr/symbimics/) (Figure 2B). Symbimix is based on laser-capture microdissection (LCM)
which was coupled to RNA sequencing and gives the transcriptome of both bacterial
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and plant partners in 5 developmental zones of the nodules of M. truncatula (Roux et
al., 2014). The analysis of ARP3 expression (Figure 2B) shows that mature infected cells
have the highest level of ARP3 expression versus the meristem and the distal infection
zone which contains freshly infected cells (Figure 2B). However, in the data obtained
by Limpens et al. (2013) by similar method, the expression of ARP3 was found to be
highest in zone of infection.
To study the subcellular localization of ARP3 in infectedcells, we constructed
N-terminal fusion of ARP3 with GFP under leghemoglobin (LB) promoter, which is
active only in infected cells. GFP-ARP3-expressing nodules were analyzed by confocal
microscopy (Figure 1G). The analysis of subcellular localization of GFP-tagged ARP3 in
mature infected cells of the fixation zone has shown that ARP3 localized in the host
cell cytoplasm; the fluorescence signal of GFP was surrounding the symbiosomes and
delineating the plasma membrane (Figure 1G). Due to the activity of LB promoter, the
GFP labelling was observed in infected cell, but was absent in uninfected neighboring
cells. To verify the labelling pattern, in a separate experiment, we used a specific
anti-ARP3 antibody developed for Arabidopsis thaliana (Kotchoni et al., 2009). ARP3
proteins of A. thaliana and M. truncatula share over 87% identity at the level of amino
acid sequences. The anti-ARP3 antibodies were tested on Medicago nodule extracts
by Western blot analysis. The band of expected size has been detected, that shows
that anti-ARP3 specific antibody may be used in the analysis of Medicago nodules
(Figure 2C). Immunolabeling with specific anti-ARP3 antibody has shown a pattern
similar to that observed for GFP-tagged ARP3 (Figure 1H, I).
To study the localization of ARP3 in relation to actin in high resolution, we used
electron microscopy immunogold labeling in mature symbiotic cells of nodules
expressing GFP-ARP3. For immunolabeling of actin we used anti-actin antibody
developed in mouse followed by a secondary anti-mouse antibody coupled to
10 nm colloidal gold particles. For immunolocalization of ARP3 we used in separate
experiments either anti-GFP antibody developed in rabbit or specific anti-ARP3
antibody, which is also developed in rabbit. The secondary anti-rabbit antibody
coupled with 15nm gold particles were used to reveal the signal (Figure 3). The signals
for actin and for ARP3 were present in the cytoplasm around the symbiosomes
in stage 3 according to the developmental stage classification of Vasse et al. (1990)
(Figure 3A, B). These symbiosomes already lost the ability to divide and grow rapidly.
Actin and ARP3 were also associated with small vacuoles (Figure 3C, D). The actin
network around symbiosomes was structurally similar to the dynamic “actin scaffold”
observed around the membrane of the phagosomes by Liebl & Griffiths (2009).
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Figure 3. Immunogold localization of ARP3 and actin in infected cells. A, B double labeling, A, the localization of
actin (10 nm gold particles) is indicated by the arrowhead and ARP3 (15 nm gold particles) is indicated by the
arrow in infected cells of GFP-ARP3 expressing nodules. The signal is present in the cytoplasm surrounding the
symbiosome membrane; B, labelled for actin (10nm gold particles) microfilaments around the symbiosome
membrane; C, small vacuole labeled by anti-actin antibody; D, small vacuole labeled by anti-GFP antibody in
GFP-ARP3 expressing nodule tissue. SB, symbiosome; Vac, vacuole.
4
The actin network around symbiosomes is not formed in dnf1 mutant nodules
To clarify the link between the developmental stage of the symbiosomes and
the ARP3 pattern, we used Medicago mutant dnf1 (Wang et al., 2010)ZM. DNF1 gene
encodes a symbiosis-specific subunit of the signal-peptidase complex. The mutation
of this gene blocks the development of the symbiosomes so they are not able to reach
maturity (Van de Velde et al., 2010; Wang et al., 2010). The nodules of dnf1 mutant are
unable to fix nitrogen and their infected cells are filled with small symbiosomes at
developmental stages 1 and 2 (Figure 4A, B). Immature symbiosomes may persist in
this stage up to 10 consecutive cell layers till the senescence starts.
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Figure 4. Phenotype of dnf1 mutant nodules (A-D) and analysis of actin pattern in nodules with partial silencing
of ARP3 (E, F). A and B, light microscopy of the dnf1 nodules; C, phalloidin staining of actin in the dnf1 nodules; D,
the localization of ARP3 in the dnf1 nodules by using anti-ARP3 specific antibody, plasma membrane (PM) is
labelled in infected cell (arrow) and in uninfected cell (arrowhead); E and F, phalloidin staining of actin in RNAiARP3 nodules. The pattern of actin in the infected cell is represented by dots or very short microfilaments,
neighboring uninfected cells have actin microfilaments similar to the control nodules presented on Figure1. IC,
infected cells; UnC, uninfected cells; IT, infection thread. Bars: A = 50µm; B-F = 20µm.
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We performed the actin labeling (Figure 4C) and the immunolocalization of ARP3
using the specific anti-ARP3 antibody on nodule sections of dnf1 mutant (Figure 4D).
The actin pattern in the uninfected cells was similar to what we previously observed
(Figure 1A-C). Actin microfilaments were crossing the vacuoles and delineating the
plasma membrane. However, infected cells of dnf1 nodules do not form the subtle
actin net enveloping the symbiosomes as was observed in control nodules (Figure 1F).
The signal for ARP3 outlined the plasma membrane (Figure 4D) but did not surround
the symbiosomes in the same way as was observed in control nodules (Figure 1G, H).
The pattern of ARP3 in uninfected cells was, however, similar to that in uninfected cells
of control nodules. We conclude that ARP3 association with symbiosomes depends
on the developmental stage; it starts to be assembled in transition between stages 2-3
and forms dense actin network around mature nitrogen-fixing symbiosomes.
The results of immunolocalization of ARP3 in dnf1 mutant nodules suggest that
the ARP2/3 complex may be involved in the formation of the F-actin array around
membranes of growing and mature symbiosomes. Therefore, we tested the effect of
ARP3 knockdown on the actin configuration and determined its role in symbiosome
maturation.
The F-actin network is essential for symbiosome development
To study the role of the ARP2/3 complex in F-actin organization around
symbiosomes, we made a RNA interference (RNAi) construct driven by the LB
promoter. LB promoter becomes active in distal cell layers of the infection zone
(Limpens et al., 2009), however it is heavily up-regulated in mature infected cells of
nitrogen fixation zone. The silencing under LB promoter should reduce the level of
ARP3 only in the infected cells, and should not affect the uninfected cells. The level of
silencing estimated by qPCR using as a template RNA extracted from pooled nodules
from 25 plants is shown in Figure 2D.
To find out whether the actin configuration is affected due to partial silencing
of ARP3, we used phalloidin staining in fresh nodules collected from RNAi-ARP3
transgenic roots; five nodules were analyzed (at 14 dpi). The configuration of actin
filaments in infected cells of RNAi-ARP3 nodules was markedly different comparing
with control infected cells: actin was present as dots or very short cables (Figure 4E, F)
and the actin net encompassing the symbiosomes was absent. Actin microfilament
pattern in uninfected cells was not affected (Figure 4E, F). The intensity of the labeling
was similar to control nodules (Figure 1B, D). We concluded that the partial silencing
of ARP3 affects the actin configuration in infected cells.
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Figure 5. Light (A, B, D, and E) and electron microscopy (C, F, and G) analysis of RNAi-ARP3 and empty vector
control nodules. A-C, empty vector control nodules; D-G, RNAi-ARP3 nodules. Silencing did not affect ARP3
expression in the zone of infection; no defects in bacteria release were detected in RNAi-ARP3 nodules in
comparison with control (A, D). In the zone of fixation, mature symbiosomes were radially aligned in control
nodules (B, C). In RNAi-ARP3 nodules, small undeveloped symbiosomes are freely distributed in the host
cytoplasm (E, F, and G). The partial detachment of plasma membrane from cell wall is indicated by arrowheads
(E,G). IC, infected cell; UnC, uninfected cell; It, infection thread; V, vacuole; SB, symbiosome; S, senescent
symbiosomes, PM, plasma membrane, CW, cell wall. Bars: A, D = 50µm; B,E = 25µm; C, F,G = 2µm.
To investigate the effect of silencing of ARP3 on symbiosome development, we
analyzed RNAi-ARP3nodules by light and electron microscopy (Figure 5). A total of
25 nodules from 2 consecutive experiments were analyzed at 15 dpi. No defects in the
bacteria release from infection threads or host cell structural deviations were detected
in the meristem or zone of infection (Figure 5D). However, around 50% of the nodules
displayed an aberrant phenotype that was observed in distal cell layers, where the
maturation of symbiosomes takes place (Figure 5E, F). In the nodules with an aberrant
phenotype, symbiosomes remain markedly smaller throughout the infected zone.
They were randomly distributed in the host cytoplasm (Figure 5E, F). This contrasts
with the control nodules, in which the symbiosomes reached maturity and were
radially aligned (Figure 5B, C). Vacuole fragmentation also was observed in some of
the infected cells in RNAi-ARP3nodules. In addition to the effect on symbiosomes
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differentiation, we saw in 40% of the nodules that the plasma membrane was
locally detached from the cell wall in infected cells (Figure 5D, E, G). The effect on
symbiosomes in these cells may therefore be indirectly caused by impaired symplast/
apoplast transport. However the defects in symbiosome development were also
observed in the nodule cells without visible plasma membrane detachments.
The reduced expression of ARP3 was not lethal for the infected cells as the plant
cells maintained symbiosomes in several (approx. 8 to 15) cell layers. Ultrastructural
analysis of RNAi-ARP3 nodules (Figure 5G) showed that the host cytoplasm of infected
cells contained ribosomes, endoplasmic reticulum, and mitochondria. However, the
symbiosomes remained small and immature and some of them were prematurely
lysed.
Discussion
Actin rearrangement in infected cells of root nodules
F-actin rearrangement plays a crucial role in mediating the relationships of plant
cells in their contacts with biotic agents and pathogens (Takemoto & Hardham, 2004;
Day et al., 2011; Henty-Ridilla et al., 2013). Cárdenas et al. (1998) and de Ruijter et al.
(1999) have shown that Nod factors, the main signalling molecules of symbiosis, affect
actin microfilaments pattern in root hairs during the establishment of symbiosis.
Information about the role of actin in the later stages of symbiosis is scarce; however
some new insights into the molecular mechanism of actin cytoskeletal configuration
in infected nodule cells appear. For instance, Marchetti et al. (2013) have shown that
Medicago microsymbiont Sinorhizobium meliloti co-cultured with HeLa cells induced a
change of actin pattern. This modification was caused by queuosine, a hypermodifed
nucleoside, which was produced by bacteria. Although changes of actin pattern have
been documented in infected leguminous cells (Whitehead & Day, 1997; Davidson &
Newcomb, 2001; Fedorova et al., 2007; Timmers, 2008), but the molecular mechanisms
of actin microfilament rearrangement in cells containing rhizobia are still unknown.
To address this issue, we investigated the role of actin and ARP2/3-dependent
actin nucleation during the intracellular accommodation of rhizobia in root nodule.
We have shown that the configuration of actin cytoskeleton changes markedly
during the course of symbiosome maturation. Immature dividing symbiosomes in
young infected cells of wild type nodules and in the nodules of dnf1 mutant do not
have a net of microfilaments enveloping them. During the phase of symbiosomes
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expansion, actin network around symbiosome membranes may be highly dinamic
for supporting vesicle transport and promoting the growth. Actin is known to
organize the plant cytoplasm (Yanagisawa et al., 2013) and may be essential for the
organization of narrow zones of cytoplasm around the symbiosome membranesfor
efficient transport to the membrane.
An actin “scaffold” is gradually assembled around growing symbiosomes, and
become quite prominent around mature nitrogen-fixing units. The analysis of the
localization and the functional role of ARP3 in root nodules has shown that ARP3 is
operational in the assembly of actin around symbiosomes and is indispensable forthe
maturation to convert to a nitrogen-fixing units.
How may the ARP2/3 complex contribute to symbiosome maturation?
In A. thaliana, ARP2/3 is strongly associated with cell membranes of the microsomal
fraction from several organelles including ER, tonoplast, plasma membrane and
early endosomes (Kotchoni et al., 2009; Zhang et al., 2013). It is well established that
symbiosome membrane growth relies on host cell membranes provided in the form
of post-Golgi vesicles and ER (Kereszt et al., 2011; Ivanov etal., 2012). The integration of
ARP2/3 into branched actin filament networks may facilitate the physical anchoring
of a polymerizing actin filaments on an organelle surface (Yanagisawa et al., 2013).
Similar mechanisms may be employed during the construction of symbiosome
membranes. It is possible that the phenotype we observed in RNAi-ARP3 nodules
may be the consequence of non-optimal association of ARP3 with the membranes
of ER and post-Golgi vesicles causing the defects in their trafficking toward the
symbiosomes.
In the last phase of symbiosome development, after they reach maturity and start
to fix atmospheric nitrogen, the actin is arranged around symbiosome membranes
to form a network of microfilaments. The formation of this structure depends on the
developmental stage: it is a property of maturing and fully mature symbiosomes. Its
assembly therefore may be linked to symbiosome membrane properties, which are
developmental stage-dependent.
The change of symbiosome membrane identity coincides with the gradual change
in actin arrangement in the cytoplasm surrounding symbiosomes
In symbiotic infected cells, the identity of freshly released symbiosomes is similar
to plasma membrane, but over the course of maturation they accept late endosome/
tonoplast proteins into their membranes (Limpens et al., 2009). Such relocation
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may be the consequence of host cell vacuole de-functionalization that causes the
retargeting of tonoplast-residing proteins toward the symbiosomes (Gavrin et al.,
2014; Chapter 3). This change in membrane identity therefore coincides with the
gradual change in actin arrangement around symbiosomes.
The dense actin network around the mature symbiosomes may contribute to
maintenance of them as individual units, for example by limiting fusion with the
lytic vacuoles as well as homotypic fusion of symbiosomes. The pattern of actin
rearrangement around symbiosomes structurally resembles the actin “scaffold”
formed around endosomes and phagosomes in animal cells, which prevents fusion
with lysosomes (Kumar & Valdivia, 2008; Liebl & Griffiths, 2009).
In some RNAi-ARP3nodules, we observed vacuole fragmentation similar to that
observed in yeast cells with mutations of actin and proteins involved in the regulation
of nucleation by the ARP2/3 pathway (Eitzen et al., 2002). Vacuole formation defect
in the case of actin mutations may be explained by an abnormality of the endosome
positioning, maturation and migration towards vacuoles (Taunton et al., 2000; Morel
et al., 2009; Gopaldass et al., 2012).
Recently Deeks et al. (2012) and Hawkins et al. (2014) described the plant networked
(NET) superfamily of actin-binding proteins. Members of this family couple the actin
cytoskeleton with specific membranes: tonoplast, nuclear membrane, or plasma
membrane. It seems worthwhile to test whether the NET protein specific for late
endosomes/tonoplasts is retargeted to the SBs at the transition from infection to
fixation.
The ARP2/3 complex in plants is essential for some cell types such as trichomes
(Zhang et al., 2005) and may be dispensable in others (Deeks & Hussey, 2005). For
example, NAP1, a component of the SCAR/WAVE complex, that regulates actin
polymerization, is operational for appropriate root-hair and infection-thread growth
during early stages of infection, but dispensable for the nodulation (Miyahara et
al., 2010). ACTIN-RELATED PROTEIN COMPONENT1 (ARPC1), which constitutes a
subunit of the ARP2/3, is also essential for rhizobial infection but not for arbuscular
mycorrhiza symbiosis in Lotus japonicas (Hossain et al., 2012).
We believe that infected cells of root nodules have specific requirements for
ARP2/3 due to the extremely high demand for membrane traffic and the role that
the actin network plays in reformation of cytoplasm architecture and symbiosome
maturation. We have shown that actin configuration in infected nodule cells changes
markedly during symbiosome development. When symbiosomes mature, they
become surrounded by actin network. The ARP2/3 complex is operational in the
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rearrangement of actin around symbiosomes. We hypothesize that the retargeting
of late endosome/tonoplast proteins to the symbiosome membranes allows the
recruitment of actin to the symbiosomes and that this recruitment is necessary for
their functional maturation.
Material and Methods
Plant growth conditions, transformation and inoculation
Plants of Medicago truncatula Jemalong A17 were grown in the growth chamber
with 16h of light, temperature 21°C. Plants were inoculated with Sinorhizobium
meliloti 2011, and harvested 15/21 days post inoculation (dpi). Agrobacterium rhizogenes
MSU440 based root transformation of M. truncatula roots was performed according
to Limpens et al. (2004).
Nodulation assay was performed from two independent experiments. The batch
of plants used for transformation contains around 50 plants. The nodules from the
transgenic roots of these plants were collected for analysis at 15 and 21 dpi. The
nodules from at least 10 plants were used for immunolocalization experiments. For
analysis of nodule phenotype in the experiments with the silencing of ARP3 were used
15 nodules from transgenic roots of different plants.
Cloning
cDNA for cloning was prepared according to the iScript™ cDNA Synthesis Kit
protocol (Bio-Rad Laboratories, Inc.) with 1 µg RNA template for cloning and 500
ng RNA template for qPCR. The desired sequence was amplified by using Phusion™
High-Fidelity DNA Polymerase (Finnzymes, Finland). Cloning primers: Arp3-cl-F
5’CACCATGGCTCCGGGTATACAAAA3’; Arp3-cl-R 5’TTAATACATCCCCTTAAAAACAGGA3’.
PCR products were purified using a High Pure PCR Products Purification Kit (Roche
Diagnostics, Germany).
Entry clone of ARP3 gene was generated by TOPO cloning (Invitrogen). The
Gateway® technology (Invitrogen, USA) was used to create the desired GFP fusion and
RNAi destination vectors. pENTR clone of ARP3 was recombined into the following
destination vectors using LR Clonase (Invitrogen): pK7WGF2-R (containing LB
promoter) creating N-terminal GFP-X fusions and pK7GWIWG2 for RNAi silencing
driven by LB promoter (Limpens et al., 2009).
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ARP2/3-mediated actin nucleation at symbiosomes is essential for their development
qPCR Analysis
Total RNA was extracted from roots and different zones of 14 dpi root nodules
using E.Z.N.A. Plant RNA Mini Kit (Omega Bio-Tek) and transcribed into cDNA using
the iScript cDNA synthesis kit (Bio-Rad). qPCR was set up in a 20-μL reaction system
using iQ SYBR Green Supermix (Bio-Rad). As an internal control for qPCR analysis
was used the expression of Ubiquitin. The analysis was performed in three or six
replicates. The normalization of results and the creation of the graphs displaying the
results were performed automatically with the software of Bio-Rad CFX Manager
3.0 protocol. Gene-specific primers were designed with Primer-3-Plus software
(Untergasser et al., 2007). qPCR primers: Arp3-q-F 5’AGCAATGGA-GAGGCGTTAAA3’;
Arp3-q-R 5’CGCAGACTGAATG-CACTTGT3’.
Sample preparation for transmission electron microscopy
Tissue preparation was performed as described earlier (Fedorova et al., 2007).
For immunogold analysis with electron microscopy, nodules were fixed in 1%
paraformaldehyde in a 50 mM phosphate buffer, embedded in LR white resin, and
polymerized with UV light at –20°C. Thin sections (60 nm) were cut using a Leica
Reichert Ultracut S microtome; nickel grids with the sections were blocked in 3% BSA
in PBS, and then incubated overnight at 4°C with the primary antibody.
Primary antibody: anti-actin developed in mouse (Molecular Probes), dilution
1:20; anti-Arp3 developed in rabbit (generously donated by Prof. Szymanski) dilution
1:50, anti-GFP developed in rabbit (Molecular Probes), dilution 1:100. The signal was
revealed by incubation with secondary antibody: gold-labeled (15 nm) goat antirabbit (BioCell), dilution 1:40; or gold-labeled (10 nm) goat anti-mouse (BioCell),
dilution 1:40; incubation proceeded for 1 hour at room temperature. Sections were
contrasted with 2% aqueous uranyl acetate and lead citrate and examined using a
JEOL JEM 2100 transmission electron microscope.
4
Confocal laser–scanning microscopy
Transgenic roots and nodules were selected using a fluorescent stereomacroscope
(Leica MZ FLIII, Leica). As a selection marker for the transformed roots and nodules
was used red fluorescent protein DsRED1 (Limpens et al.,2004). Harvested nodules
were manually sectioned in phosphate buffer with 3% sucrose and counterstained by
propidium iodide 0.01%. Samples were washed with phosphate buffer with sucrose
and transferred to microscope slides. For immunolabelling nodules were fixed in 1%
paraformaldehyde in a 50 mM phosphate buffer, washed with the same buffer (4°C)
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Chapter 4
and manually sectioned. Sections were transferred to the blocking solution (3% BSA
in PBS) and then incubated overnight at 4°C with the primary antibody. Primary
antibody: anti-GFP developed in rabbit (Molecular Probes), dilution 1:250, anti-Arp3
developed in rabbit (generously donated by Prof. Szymanski) dilution 1:50. Sections
were counterstained by propidium iodide 0.01%. Confocal imaging was done on a
Zeiss LSM 5 Pascal confocal laser–scanning microscope (Carl Zeiss, GmbH).
Actin labeling with a phalloidin probe
Nodule sections were incubated (for 2h at room temperature and in darkness)
in phosphate buffer pH 7.2 with 200 mM MBS (3-maleimidobenzoic acid
N-hydroxysuccinimide ester, Sigma), 0.05%Triton X-100, and Alexa FluorR 488
phalloidin (Molecular Probes), dilution 1:100. Sections were counterstained and
examined as described earlier.
Western blot analysis
The proteins were extracted from root nodules in 0.025M Tris-HCl buffer containing
1mM EDTA, 1mM DTT and protease inhibitors cocktail (Roche). The probes loaded to
the gel: 15 mg/well for extracts from GFP-tagged ARP3 and 45mg/well for extracts
from wild type nodules to detect the endogenous ARP3. The GFP-tagged isoform was
more abundant that the endogenous protein due to the over expression caused by a
strong LB promoter used in N-terminal fusion of ARP3 with GFP. The proteins were
separated by 12% SDS-PAGE and blotted to nitrocellulose (Bio-Rad). The membrane
was incubated in 3% BSA as a blocking agent followed by primary anti-ARP3 specific
antibody, 1:50 dilution; followed by secondary antibody, Anti-Rabbit IgG peroxidase
antibody produced in goat (Sigma), 1:5000 dilution. The immunosignal was revealed
by incubation with Immuno-Star Western Chemiluminescent Kit (Bio Rad, 170-5070).
The blots were photographed by Molecular Imager Chemi Doc XRS+ using Image Lab
Software, in Chemi mode or EpiWhite (for the Prestained Protein Ladder).
Accession number of Medicago ARP3 nucleotide sequence is Medtr8g089630.1.
Acknowledgments
The authors greatly appreciated a generous gift of anti-ARP3 antibody from Prof.
Szymanski. The authors would like to thank Dr. T. Ketelaar and Dr. H. Franssen for
reading and commenting the manuscript and Dr. R. Geurts for fruitful discussions.
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ARP2/3-mediated actin nucleation at symbiosomes is essential for their development
A.G. received a PhD fellowship from EPS School of Biological Sciences (Wageningen
University). E.F. and T.B. are supported by the European Research Council (ERC-2011AdG294790). English spellcheck: http://www.tessera-trans.com.
4
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The establishment of symbiosis:
the role of Synaptotagmin1
Aleksandr Gavrin1, Olga Kulikova1, Ton Bisseling1,2, Elena E. Fedorova1
Laboratory
of
Molecular
Biology,
Department
of
Plant
Sciences,
Graduate
School
Experimental
Plant
Sciences,
Wageningen
University,
6708
PB,
The
Netherlands; 2College of Science, King Saud University, Riyadh 11451, Saudi Arabia
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Abstract
During formation of infected cells new membrane protrusions of plasma
membrane as infection threads, cell wall-free unwalled droplets and symbiosomes
are formed. These membrane compartments separate the bacteria from the
host cytoplasm. Symbiosomes rapidly increase in number and volume due to the
expanding bacteria and may represent putative sites of increased membrane stretch
or tension. We hypothesized that membrane tension may create a vector for vesicle
fusion to the interface membrane. To test this hypothesis, we studied MtSyt2 and
MtSyt3, two Medicago truncatula homologs of Arabidopsis thaliana synaptotagmins
from the group of synaptotagmin 1 (SYT1). MtSyt2 and MtSyt3 are expressed in
nodule primordia and apical parts of root nodules. The proteins were localized in the
plasma membrane regions of meristematic and elongating cells, showing that they
are involved in the houskeeping membrane fusion. However, in the infected cells
MtSyt2 and MtSyt3 were very abundant over the plasma membrane protrusions infection threads and unwalled droplets. The double silencing of MtSyt2 and MtSyt3
shows that they are operational during the growth of infected cells and intracellular
rhizobia accommodation. We assume that MtSyt2 and MtSyt3 may be co-opted by the
symbiosis into the process of interface membrane formation.
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The establishment of symbiosis: the role of Synaptotagmin 1
Introduction
The legume-rhizobium symbiosis is one of the rare examples in the plant kingdom
where bacteria are accommodated inside host cells. Specialized infected cells of the
root nodule, a de novo symbiotic organ, contain bacteria that are able to fix atmospheric
nitrogen. The host cell undergoes morphological changes as a response to enter of
rhizobia. Bacteria penetrate developing nodule cells through cell wall-bound tubular
structures called infection threads. The release of bacteria into the plant cell takes
place from unwalled infection droplets, local cell wall-free regions of the infection
thread. Released bacteria become surrounded by a host cell-derived membrane and
form so-called symbiosomes (Gibson et al., 2008; Kondorosi et al., 2013). Due to the
rhizobia expanding in the host cell, the surface of the interface membrane, separating
the symbionts, increases and may reach up to eight times the area of the plasma
membrane (Roth & Stacey, 1989).
It is well known that the plasma membrane is inelastic; its capacity to stretch is only
around 1-3%. Therefore, exocytosis of new membrane material is involved in changes
in the membrane surface and in repairing damaged membrane loci (Apodaca, 2002;
Grefen et al., 2011). According to the hypothesis of Morris and Homann (2001), in
animal cells the modification of membrane surface area depends on the membrane
tension. A local increase of tension leads to an increase of membrane surface area due
to the fusion of the available endomembrane reserves with that specific region of
the membrane by Ca2+-dependent targeted exocytosis. Membrane vesicles including
lysosomes are exocytosed to seal the wound and to ease the membrane tension
(Reddy et al., 2001; McNeil & Kirchhausen, 2005; Idone et al., 2008; Gauthier et al.,
2009).
In plants, the typical examples of targeted exocytosis are tip growth of root hairs
and pollen tubes (Qin & Dong, 2015) and the formation of new membranes of the
cell plate during cell division (El Kasmi et al., 2013). The study on tip growth showed
the establishment of a tip-focused Ca2+ gradient, as well as actin microfilament
reorganization and redirection of small GTPases of the Rab, Rop and ARF families
involved in membrane fusion (Ketelaar et al., 2003; Šamaj et al., 2006). Local higher
concentration of cytoplasmic calcium is required for the targeted vesicular fusion.
This highlights the crucial role of calcium sensors in targeted exocytosis.
In animals, the role of calcium sensors from the synaptotagmin I and VII group in
membrane fusion is well established (Südhof & Rothman, 2009; Wang et al., 2014).
Synaptotagmins have two calcium-binding C2 domains (C2A and C2B) with different
5
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affinities to calcium and the N-terminal transmembrane domain. Synaptotagmins are
able to bind to the integral membrane proteins (t-SNAREs), as well as to the vesicular
v-SNAREs (vesicle-associated membrane protein, VAMP) in a process that is regulated
by Ca2+ (Adolfsen et al., 2004; Maximov et al., 2009; Falkowski et al., 2011). The response
of the cells to mechanical stimulation involves Ca2+ spiking, phospholipase signaling,
rapid remodeling of the actin skeleton and quick retargeting of exocytosis to the
place where the membrane is overstretched or damaged (Jaffe et al., 2002; Telewski,
2006). The SNARE complex lacks specific Ca2+-binding sites, so the association of
SNAREs with synaptotagmin creates a vector for local membrane fusion. In this case,
synaptotagmins provide Ca2+ sensitivity to direct the rapid response that is necessary
for the fusion of the membrane vesicle with the region of the overstreached or
damaged membrane (Draeger et al., 2011).
In recent years synaptotagmins have been also found in plants (Craxton, 2004;
Nakagawa et al., 2007; Schapire et al., 2008; Yamazaki et al., 2008). The structure of
plant synaptotagmin 1 differs from that of conventional mammalian orthologs.
Plant synaptotagmin 1 lacks an N-terminal extension in the extracellular region, but
there is an SMP (plant synaptotagmin-like, mitochondria proteins) domain just next
to the C-terminal side of the trans-membrane domain (Yamazaki et al., 2010)2010.
In Arabidopsis roots synaptotagmin 1 (SYT1) is especially expressed in the quiescent
center, columella, epidermis and vasculature and in the tip of growing root hairs. It is
localized on the plasma membrane and cell plates of dividing cells. Synaptotagmin
is involved in membrane repair in case of osmotic misbalance and cold stress and
interacts with viral movement proteins (Schapire et al., 2008; Yamazaki et al., 2010).
Microarray data also show a large overlap between responses to cold, drought and
osmotic stresses and the mechanical wounding (Walley et al., 2007).
The development of infected nodule cells requires formation of membrane
interface between the symbionts. The plasma membrane of the host cells becomes
asymmetric and the new membrane protrusions (infection threads, cell wall-free
unwalled droplets and symbiosomes) are formed. These membrane compartments,
separating the bacteria from the host cytoplasm, rapidly increase in number and
volume and become putative sites of increased membrane stretch or tension. To
respond to the membrane tension, the cell has to redirects part of the exocytotic
traffick to the site of membrane overstretch/damage. The fusion of post-Golgi vesicles
as well as ER cisterns to infection threads, unwalled droplets and symbiosomes is well
documented and shows that this redirection is a real biological event (Roth & Stacey,
1989). How is the fusion with these target membranes coordinated? It is known that
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The establishment of symbiosis: the role of Synaptotagmin 1
the plant cells respond to contacts with the microbes by redirection of vesicular
traffic, reorganization of the cytoskeleton and changes in the endomembrane
structure of the infected cell. This response has a common pattern with mechanical
stimulation (Hardham et al., 2008). It is possible that the membrane tension may
create a vector for vesicle fusion to the interface membrane, which is overstreched
by expanding bacteria. To test this hypothesis, we studied MtSyt2 and MtSyt3, two
M. truncatula homologs of Arabidopsis thaliana synaptotagmins from the SYT1 group
(Schapire et al., 2008)2008. We showed that MtSyt2 and MtSyt3 are expressed in
nodule primordia and the apical part of root nodules. The proteins are present in the
plasma membrane regions of meristematic and elongating cells and can be involved
in housekeeping membrane fusion insuring the plasma membrane expansion. In
the infected nodule cells MtSyt2 and MtSyt3 are present on plasma membrane,
infection threads and unwalled droplets. The double silencing of MtSyt2 and MtSyt3
shows that they are operational during the growth of infected cells and intracellular
rhizobia accommodation. We hypothesise that MtSyt2 and MtSyt3 may be co-opted
by rhizobium symbiosis into the process of interface membrane formation.
Results
The selection of Medicago truncatula orthologs of Synaptotagmin 1
Five homologs of Synaptotagmin 1 genes were retrieved from M. truncatula
databases by BLAST analysis using Arabidopsis thaliana synaptotagmin 1 (SYT1)
sequences. Phylogenetic analysis shows that the MtSyt1 and MtSyt2 genes and A.
thaliana AtSYT1 and AtSYT2 belong to the same group; further sub-grouping is
not clearly defined. MtSyt3 is an orthologue of AtSYT3, they form a separate group
(Yamazaki et al., 2010)2010. MtSyt4 and MtSyt5 are grouped with AtSYT4 and
AtSYT5, however the expression of MtSyt4 and MtSyt5 in the nodules is very low. The
phylogenetic tree is presented in Supplementary Figure 1.
For this study we selected MtSyt1, MtSyt2 andMtSyt3 genes. The expression of
MtSyt1, MtSyt2 and MtSyt3 were analyzed by qRT-PCR (Supplementary Figure 2) in
roots and nodules at 14 days post inoculation (dpi). The expression of MtSyt1 and
MtSyt2 was lower in nodules than in roots, whereas expression of MtSyt3 was higher
in nodules. To specify the expression pattern of these homologs in developmental
zones of the nodule, we used database Symbimix (INRA, https://iant.toulouse.inra.
fr/symbimics/) (Supplementary Figure 3A). Symbimix is based on laser-capture
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microdissection which was coupled to RNA sequencing and gives the transcriptome
of both bacterial and plant cells in 5 developmental zones of M. truncatula nodules
(Roux et al., 2014)2014. MtSyt1, MtSyt2 and MtSyt3 show the same expression pattern
as we obtained by qRT-PCR analysis (Supplementary Figure 3B). MtSyt1 and MtSyt2
expression in nodules was lower than in roots, and the expression of MtSyt3 was
higher in nodules than in roots. MtSyt2 and MtSyt3 are mainly expressed in the apical
zones of the nodule: in distal and proximal parts of the zone of infection and in the
interzone, in contrast to the expression of MtSyt1, with the highest expression in the
fixation zone.
To study the role of synaptotagmin 1 during the symbiosis establishment in the
nodule, we have selected MtSyt2 and MtSyt3. They are expressed in the infection zone,
where membrane expansion by exocytosis is most prominent. MtSyt1 in contrast was
mainly expressed in the fixation zone of the nodule where the membrane expansion
has already stopped.
Expression analysis of MtSyt2 and MtSyt3
For the analysis, we created constructs containing the region 2.5 kb upstream of the
translational start of MtSyt2 and MtSyt3 fused to β-glucuronidase (GUS). Transgenic
roots were obtained using Agrobacterium rhizogenes-mediated transformation.
Transgenic roots and 14 dpi nodules were used for analysis of MtSyt2 and MtSyt3
expression (Figure 1A-G). pMtSyt2:GUS and pMtSyt3:GUS are strongly expressed in
young primordia (Figure 1A). In mature nodules, pMtSyt2:GUS is expressed in the
meristem and distal cell layers in the zone of infection (Figure 1B). Expression of
pMtSyt3:GUS occurs in the infection zone and distal layers of the fixation zone (Figure
1C).
In the roots MtSyt2 and MtSyt3 were expressed in all cells and most strongly in
the meristem. MtSyt2 was expressed in meristem and the epidermis of the elongation
zone, MtSyt3 in epidermal cells of the elongation zone including the root hairs. (Figure
1D, F). To see whether the MtSyt2 and MtSyt3 expression pattern may be affected by
tension, we performed a bending assay. Bended root was harvested after 48h. Bending
shifted the GUS staining to the site of bending (Figure 1E, G).
Specify whether the Nod-factor signaling pathway may have some concomitant
link with synaptotagmin 1, we analyzed the expression pattern of MtSyt2 and MtSyt3
in roots of the mutants of the Nod factor signaling pathway (dmi1, dmi2 and dmi3).
Previously, it was reported that root hairs of dmi2 and dmi3, the mutants with defects
in symbiotic responses to rhizobial Nod factors, are hypersensitive to mechanical
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The establishment of symbiosis: the role of Synaptotagmin 1
stimulation (Esseling et al., 2004). The GUS expression pattern in roots of the
uninoculated mutant plants does not show any difference from the wild type and
occured in meristem and elongation zone of the roots (Supplementary Figure 4). The
bending of the roots of dmi1, dmi2 or dmi3 mutants showed changes in the promoter
GUS expression similar to what was observed in the control roots (Supplementary
Figure 5). We have concluded that the roots of the mutants do not show defects in the
expression of MtSyt2 and MtSyt3.
Figure 1. Promoter-GUS expression analysis of MtSyt2 and MtSyt3 in nodules and roots. A, the pattern of MtSyt2
and MtSyt3 promoter-GUS expression in nodule primordia. B, expression of MtSyt2 in the nodule. C, expression
of MtSyt3 in the nodule. Expression pattern of MtSyt2 in the root before (D) and after bending (E). Expression
pattern of MtSyt3 in the root before (F) and after bending (G). Bars: A, B, C: 100 μm.
5
Cellular localization of MtSyt2 and MtSyt3
To investigate the localization of MtSyt2 and MtSyt3 in nodules and roots,
constructs expressing GFP translational fusions of these genes under the control
of their native 5’regulatory sequences were created. For confocal imaging, 14-21 dpi
root nodules were hand sectioned and fixed in 1% paraformaldehyde in a phosphate
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buffer as previously described (Gavrin et al., 2014). The GFP signal was enhanced by
anti-GFP antibody coupled with ALEXA 488. Confocal microscopy of the localization of
MtSyt2-GFP and MtSyt3-GFP revealed local accumulation of GFP fluorescence in the
meristems of roots and nodules in plasma-membrane regions; the strongest signal
was found in the cell plates of dividing cells (Figure 2). MtSyt2 labelling was present in
the cells directly adjacent to the meristem and was associated with infection threads.
MtSyt3 signal was found in all cell layers of the infection zone and the first cell layers of
the fixation zone. It was stronger than the signal of MtSyt2. In young infected cells, the
MtSyt3-GFP signal delineated the infection threads and the sites of bacteria release.
Some fluorescence also was present in the host cell cytoplasm. The symbiosomes
were not specifically outlined by the GFP signal. MtSyt2 and MtSyt3 were thus
strictly localized on cells undergoing expansion and new membrane formations: in
meristematic cells, in dividing cells where the signal was observed over the forming
cell plate, as well as in the zone of infection in the nodules, but not on symbiosomes.
Figure 2. Confocal microscopy showing the cellular localization of GFP-tagged MtSyt2 (A-D) and MtSyt3 (E-H). (A,
B) In root meristems MtSyt2-GFP labels the plasma membrane region, note intense GFP labelling over cell plates
(*) in dividing cells. (C) In the nodule MtSyt2 is localized in the meristem and distal part of the infection zone. (D)
The signal is associated with the infection thread tip. (E, F) In the nodule the MtSyt3-GFP signal is present in the
meristem, proximal and distal parts of infection zone, in the young infected cells (G). GFP signal delineates
infection threads and unwalled droplets (H). ZI, zone of infection; YIC, young infected cells; IT, infection threads;
(*) cell plates; R, release of bacteria. Bacteria and host cell nuclei were stained by propidium iodide. Bars: A:
20μm; B: 10μm; C: 40μm; D: 20μm; E: 40μm; F, G: 20μm; H: 5μm.
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Functional analysis by RNA interference (RNAi)
To knock down the transcription levels of MtSyt2 and MtSyt3 genes in the zone of
infection, we made RNAi silencing constructs under the control of the Enod12 promoter
(Limpens et al., 2005). Constructs were initially created that individually targeted the
expression of MtSyt2 and MtSyt3. The composite plants with Enod12:RNAi-MtSyt2 and
Enod12:RNAi-MtSyt3 expressing roots were inoculated with S. meliloti. The root nodules
from the transgenic roots were analyzed at 14 dpi by light microscopy. Microscopic
sections of these nodules do not display distinct irregularities in nodule structure
and were similar to control nodules (results not shown). To better understand the
roles of MtSyt2 and MtSyt3 in symbiosis, we created an RNAi construct to silence
MtSyt2 and MtSyt3 simultaneously. The level of silencing for both genes is shown in
Supplementary Figure 5. Analysis of the transgenic roots showed a markedly lower
number of nodules per transgenic root: 2.81±1.65 versus 5.32±1.91 in the control.
The transgenic nodules had a distinct phenotype showing anatomical aberrations
in nodule structure. The transgenic nodules were smaller than the control nodules.
The most prominent anomalies were the distortion in the number of the cell layers
in nodule developmental zones and the delay in rhizobia release and symbiosome
maturation. The RNAi-affected nodules display a short meristem, extended zone of
infection, diminished or aborted zone of fixation (Figure 4; Table 1). The symbiosomes
were not able to differentiate further than stage 2 according to the classification
of Vasse et al. (1990). The double silencing phenotype was observed in 65% of the
nodules (n=20).
Table 1. The number of cell layers in different nodule zones in control and Syt2/Syt3 RNAi nodules
Meristem
Infection zone and
interzone II/III
Zone of nitrogen
fixation
Control nodules
3.5±0.23
5.75±0.44
21.3±2.74
RNAi nodules
0.8±0.20
10.75± 1.74
2.08±1.3
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Figure 3. Light microscopy of the nodules elicited on the transgenic roots where both MtSyt2 and MtSyt3 were
silenced by using double RNAi construct. (A, B) MtSytB/MtSytC double silencing nodules. (B) The magnification
of (A) showing the extended zone of infection (arrows). (C, D) Control nodule. (B) Magnification of infection zone.
Bars: A, C: 50 μm; B, D: 25 μm.
Discussion
We hypothesized that targeted exocytosis which is dependent on the proteins
of the group of synaptotagmin 1, may support the formation of rapidly growing
membranes during the establishment of legume-rhizobium endosymbiosis. The plant
cell responses to pathogenic and symbiotic microbes exhibit striking similarities
to responses to mechanical stimuli (Hardham et al., 2008; Chehab et al., 2009). To
address this hypothesis we performed promoter-GUS analyses of homologues of
Arabidopsis SYT1, MtSyt2 and MtSyt3, in Medicago root nodules. Both are strongly
expressed in nodule primordia and apical parts of mature root nodules where cell
expansion occurs. MtSyt2 and MtSyt3 are present in expanding and dividing cells and
are associated with plasma membrane, cell plates, infection threads and unwalled
droplets. This pattern of synaptotagmin distribution makes it difficult to determine
whether MtSyt’s directly affect “symbiotic” membrane formation or this is an indirect
effect of disturbed cell division and elongation. However, the MtSyt2 and MtSyt3 that
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The establishment of symbiosis: the role of Synaptotagmin 1
are localized on membranes delineating infection threads and unwalled droplets
may provide the vector for targeting exocytosis to the interface membrane. It is worth
to mention that the localization of Nicotiana benthamiana Syt1 envelops the haustoria
of Phytophthora infestans by the pattern similar to the pattern on infection threads (Lu
et al., 2011).
Double silencing of MtSyt2 and MtSyt3 shows that they are operational during the
accommodation of rhizobia in the root nodule and infected cell growth. Observed
phenotype, the widening of the infection zone and the delay in symbiosome release/
maturation, may be explained by the defects in the tip growth of infection threads and
concomitant low level of bacteria present in the infected cells. It is also may reflect the
compromised growth of plasma membrane of infected cells.
The phenotype of the nodule has some similarities with the mutant dnf1 (Wang
et al., 2010)2010 where the development of the symbiosomes was restricted to stage
1-2 according to the classification of (Vasse et al., 1990). Dnf1 encodes a symbiosisspecific subunit of the signal-peptidase complex (Van de Velde et al., 2010; Wang et al.,
2010). Being an early component of the protein secretory pathway in the endoplasmic
reticulum, it cleaves signal peptides of polypeptides that are exocytosed. Similarities
between the phenotype of double RNAi silencing of MtSyt2 and MtSyt3 and dnf1 may
suggest a link between the vesicular fusion to the plasma membrane and symbiosisspecific secretory pathway dependent on DNF1.
Recent studies showed that mechanical stimuli may be a potential signal
monitored by plants during endosymbiosis establishment (Hardham et al., 2008).
For example, root hairs of dmi2 and dmi3 mutants, are hypersensitive to mechanical
stimulation when treated with Nod factors (Esseling et al., 2004). The promoter GUS
expression of MtSyt2 and MtSytC3 in uninoculated roots of dmi2 and dmi3 showed no
difference from wild type roots. However, these studies were performed in the absence
of Nod factors. Such Nod factor studies remain to be done to resolve a potential role of
synaptotagmin in these mutants.
In conclusion, the membrane tension created by the growing bacterial colony
in an infection thread may be perceived by the plant cell as a signal activating the
transport and fusion of the host cell membrane vesicles to the sites of the contact.
The localization of MtSyt2 and MtSyt3 at the site of bacterial release, around infection
threads and in the plasma membrane of growing cells supports the hypothesis that
this calcium sensor may be involved in the creation of a vector for targeted exocytosis
in young infected cells.
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Materials and Methods
Plant Materials, Transformation, and Inoculation
The Medicago truncatula accession Jemalong A17 was grown in perlite saturated
with Färhaeus medium without nitrate in a growth chamber at 21°C and 16/8-hour
light/darkness cycle. These plants were inoculated with Sinorhizobium meliloti strain
Sm2011 (OD600 0.1, 2 mL per plant). Root nodules were collected for analysis 14
days post inoculation (dpi). Agrobacterium rhizogenes MSU440-mediated hairy root
transformation was performed according to Limpens et al. (2005).
Blast analysis
The synaptotagmin homologs of Medicago truncatula named MtSyt1, MtSyt2 and
MtSyt3 were retrieved by BLAST analysis based on homology with Arabidopsis SYT1
(Schapire et al., 2008)2008.
Cloning
Gateway technology (Invitrogen) was used to produce all genetic constructs.
The primers used in cloning procedures are shown in Supplementary Table 1. DNA
fragments of putative MtSyt2 and MtSyt3 promoters were generated by PCR on M.
truncatula genomic DNA isolated from root nodules as a template and using Phusion
High-Fidelity DNA Polymerase (Finnzymes) and specific primers. PCR fragments
were introduced in pENTR-vector (Invitrogen) and subsequently recombined into the
destination vectors for promoter GUS fusion.
MtSyt2 and MtSyt3 genes were amplified on cDNA made from root nodule
RNA. The pMtSyt2:MtSyt2-GFP and pMtSyt3:MtSyt3-GFP translational fusions were
generated by multiple cloning.
To create single RNAi constructs, 600 bp long DNA fragments were amplified
and introduced into entry clones which were subsequently recombined into Gateway
pK7GWIWG2(II) binary vector, where expression of RNAi cassette is driven by the
ENOD12 promoter (Limpens et al., 2004). The double RNAi construct was created by
fusing MtSyt2 and MtSyt3 PCR-generated sequences using protocol adopted from
In-Fusion® HD Cloning,www.clontech.com. In a first round of PCR the short overlaps
(15 bp) were introduced to PCR products by using specific primers. A mixture of two
obtained PCR fragments diluted in 1:500 was used as a template in second PCR to
create a single DNA fragment. The primers are listed in Supplementary Table 1.
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Quantitative PCR Analysis
Total RNA was extracted from roots and 14 dpi root nodules using the E.Z.N.A.
Plant RNA Mini Kit (Omega Bio-Tek) and transcribed into cDNA using the iScript cDNA
synthesis kit (Bio-Rad). Real-time PCR was set up in a 20mL reaction system using iQ
SYBR Green Supermix (Bio-Rad). Gene-specific primers were designed with Primer3-Plus software (Untergasser et al., 2007). Gene expression profiles were normalized
against the transcription level of the reference gene UBQ10. The primers are listed
in Supplementary Table 1. Data were compared with M. truncatula Gene Expression
Atlas (http://mtgea.noble.org/v2/) and Symbimics database (https://iant.toulouse.
inra.fr/symbimics/).
GUS Staining
Transgenic roots and nodules were collected and washed twice in 0.1M sodium
phosphate buffer (pH7.2) incubated in GUS buffer under vacuum at room temperature
for 30 min to allow the buffer to replace oxygen in the tissue, incubated at 37°C for 2
hours. Hand-cut sections of processed nodules were analyzed using Leica DM 5500
Flu microscopes.
Confocal Laser-Scanning Microscopy
GFP-fused proteins were visualized on transgenic roots and hand-sectioned
nodules. Imaging was done on a Zeiss LSM 5 Pascal confocal laser-scanning
microscope (Carl Zeiss). Polyclonal rabbit anti-GFP antibodies (Molecular Probes) at
a dilution of 1:100 and secondary anti-rabbit Alexa 488 antibodies (Molecular Probes)
at a dilution of 1:200 were used for signal enhancement. Goat serum or 3% (vol/wt)
BSA was used as the blocking agent. Sections were counterstained by FM4-64 (30 μg/
mL) or propidium iodide (0.001%).
Sample Preparation for Light Microscopy
Tissue preparation was performed as described previously (Limpens et al., 2009).
Semi-thin (0.6 μm) sections were cut using a Leica Ultracut microtome and examined
using a Leica FL light microscope.
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Acknowledgements
A.G. received a PhD fellowship from EPS School of Biological Sciences (Wageningen
University). E.F. and T.B. are supported by the European Research Council (ERC-2011AdG294790).
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Supplementary materials
Supplementary Figure 1. The phylogenetic tree of M. truncatula synaptotagmin orthologs MtSyt1, MtSyt2 and
MtSyt3 grouped with A. thaliana AtSYTs.
Supplemetary Figure 2. Expression level of MtSyt1, MtSyt2, MtSyt3 in roots and 14 dpi nodules. Error bars
represent standard deviation. The difference between the expression of SytC in roots and nodules is significant
(P<0.05).
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Supplementary Figure 3. A, expression level of MtSyt1, MtSyt2, MtSyt3 in roots and nodules. B, distribution of
normalized RNA-seq reads (%) for MtSyt1, MtSyt2, MtSyt3 in nodule developmental zones: FI, meristematic zone;
FIId, distal infection zone; FIIp, proximal infection zone; IZ, interzone II/III; ZIII, fixation zone. The data obtained
by using the Symbimix database (INRA, https://iant.toulouse.inra.fr/symbimics/).
Supplementary Figure 4. The analysis t of GUS-promoter expression for MtSyt2 and MtSyt3 in the roots of DMI 1,
2 and 3 mutant plants.
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Supplementary Figure 5. The level of silencing for both MtSyt2 and MtSyt3 estimated by q-PCR analysis
Supplementary Table 1. List of primers
Cloning primers
Syt1-ORF-F
ATGGGTTTCTTCAGTACAATTTTTGG
Syt1-ORF-R
TTATGCAGTTCTCCACTGCAAC
Syt2-ORF-F
ATGAGTATTTTAAGTACTATAGCTAGTTTTTTAGG
Syt2-ORF-R
TTATGGGGTTCTCCACTGAA
Syt3-ORF-F
ATGGGGTTCTTTGAAAGTTTCTT
Syt3-ORF-R
TTAAACCACCTTCCATTTTATCTCA
Syt1-Pr-A-F
AGAATCCCTTATTGTACTTTTTGC
Syt1-Pr-A-R
TTGGAATGGATCTAAATGATTTC
Syt2-Pr-B-F
CGACGTTCCTTCCTCTTTGG
Syt2-Pr-B-R
TTCAACTTCAATATCAATGTCTATGG
Syt3-Pr-C-F
AGCAACTAAAAATGGCAAGAAAA
Syt3-Pr-C-R
TTTTGGTGAACAGAGCAATGA
qPCR primers
Syt1 -F
AGGAGGCCCGTTGGAATTTT
Syt1 -R
TGTGATGCTTCCCCTCAACA
Syt2 -F
TGGATCCATCACAGGCCATG
Syt2 -R
CGCAAACGGATTTGTGTGGT
Syt3 -F
TTCCGGCGAAGAAAACCACT
Syt3 -R
ATGCCCCAGGGATTCCTTTG
5
Primers for double silencing
Syt2 -F
TGGATCCATCACAGGCCATG
Syt2 -R
CGCAAACGGATTTGTGTGGT
Syt3 -F
TTCCGGCGAAGAAAACCACT
Syt3 -R
ATGCCCCAGGGATTCCTTTG
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Chapter 6
General Discussion:
The adaptation of the
endomembrane system of host cell
to intracellular bacteria
Aleksandr Gavrin
Laboratory of Molecular Biology, Graduate School
Wageningen University, Droevendaalsesteeg 1, 6708PB
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Experimental Plant Science,
Wageningen, The Netherlands
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Chapter 6
In my thesis I have focused on the molecular mechanisms that control the
adaptation of infected cells to accommodate the microsymbiont. This was studied in
determinate as well as indeterminate nodules. During nodule development the host
cell architecture dramatically changes. However, how this is coordinated and what
mechanisms are involved is not yet completely clear. I will first summarize my major
observations that are related to subsequent steps of the adaptation of host cells to
accommodate intracellular rhizobia.
• Rhizobia enter nodule cells by cell wall bound infection threads. At these threads,
locally cell wall free regions are created which are named unwalled droplets.
This local modification of the host cell wall is accompanied by a change in pectin
turnover (Chapter 2). In this process a symbiosis specific exocytotic process,
involving VAMP721d is operational. At the unwalled droplets, rhizobia surrounded
by a host membrane are released into the cytoplasm of the host. These are the
symbiosomes that subsequently divide. Release and division of symbiosomes also
requires VAMP721d.
• The vacuole volume of the infected cells markedly decreases, creating space for the
intracellular rhizobia, at the transition of infection to fixation zone. Expression of
HOPS genes is switched off in the same cell layer. The vacuole defunctionalization
changes the traffic toward tonoplast and part of the vacuole-residing proteins
appears on symbiosome membranes. Symbiosome membranes accept for
example endosomal/vacuolar markers Rab7, and the vacuolar aquaporin TIP1g.
When the vacuolar aquaporin TIP1g is re-targeted to the symbiosomesthe water
flow is redirected from the vacuole towards symbiosomes, which facilitates their
growth. At the final stage when symbiosis is terminated, the proteins from the
HOPS complex, VPS11 and VPS39, are being recruited to symbiosome membranes.
This may allow the lytical machinery of the vacuole/autolysosomes to process the
remnants of symbiosomes and the collapsing host cell (Chapter 3).
• Growth of infected cells results in a volume several times larger than that of
uninfected cells. By default, growth of any plant cell requires turgor pressure,
which depends on a functional vacuole. How is growth achieved in infected cells
where the vacuole is markedly diminished and non-acidic? The strategy, which is
employed in symbiosis, seems the substitution of osmotic pressure of the vacuole
by a mechanical pressure of expanding symbiosomes (Chapter 3).
• When symbiosomes expand an actin network is formed around them. This is
associated with the accumulation of ARP3, member of an actin nucleating complex.
This actin network may be involved in vesiclae transport towards symbiosomes
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General Discussion
and promoting their growth. The dense actin network becomes most prominent
in mature infected cells, where a net of microfilaments practically envelop each
symbiosome. This may ensure their maintenance as individula units (Chapter 4).
• During the development of infected cell the infection threads, unwalled droplets,
and symbiosomes rapidly increase in volume and may be the putative sites of
increased membrane tension. Two Medicago truncatula homologs of calcium
sensor Synaptotagmin 1 of A. thaliana, MtSyt2 and MtSyt3, are expressed in nodule
primordia and apical parts of root nodules. The proteins are located in the plasma
membrane regions of meristematic and elongating cells, showing that they are
involved in the housekeeping membrane fusion. In the infected cells MtSyt2
and especially MtSyt3 is very abundant over the plasma membrane protrusion
- infection threads and unwalled droplets. The double silencing of MtSyt2 and
MtSyt3 shows that they are operational during the growth of infected cells and
intracellular rhizobia accommodation. The localization of MtSyt2 and MtSyt3 at
the site of bacterial release, around infection threads and in the plasma membrane
of growing cells supports the hypothesis that this calcium sensor may be involved
in the creation of a vector for targeted exocytosis in young infected cells (Chapter 5).
• In the next part of Chapter 6, I will discuss my studies on endomembrane system
of infected cell in the framework of root nodule development and endomembrane
biology in general.
The role of Nod factor signalling in control of rhizobia release from infection threads
Nodule organogenesis is initiated by reprogramming of differentiated root
cells and this is induced by rhizobial Nod factors (Xiao et al., 2014). These Nodfactors probably also have a role in the nodule, since components of the Nod-factor
signalling pathway are expressed in the nodules in the infection zone (Bersoult et al.,
2005; Capoen et al., 2005; Limpens et al., 2005; Riely et al., 2006). Mutation or knock
down of expression of Nod-factors signaling pathway genes, using nodule-specific
promotershas no effect on nodule organogenesis, but it affects the release of rhizobia
from infection threads. For instance, knockdown of SymRK (DMI2) causes a block
of release of rhizobia into host cells, while nodule formation and infection thread
growth still occur (Capoen et al., 2005; Limpens et al., 2005). Specific silencing of the
Nod factor receptor NFP in the infection zone blocks rhizobial release in nodule cells
(Moling et al., 2014). The Nod factor signalling components IPD3 and CCaMK (DMI3)
also are essential for release of rhizobia (Morzhina et al., 2000; Voroshilova et al.,
2009; Ovchinnikova et al., 2011).
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Formation of cell wall-free regions at infection threads and expansion of infected
cells
Rhizobia enter host cells by infection threads and at these threads locally
unwalled regions are formed, which involves cell wall modification. These are
the sites where rhizobia are released. It was hypothesized that the VAMP721d/e
exocytotic pathway, identified in M. truncatula (Ivanov et al., 2012), has a role in this
local modification of the cell wall. To clarify the role of VAMP721d/e we used Glycine
max, which forms determinate nodules. The silencing of GmVAMP721d blocked the
release of rhizobia. Instead of symbiosomes, the infected cells contained big clusters
of bacteria embedded in a matrix of methyl-esterified and de-methyl-esterified
pectin. Surprisingly, GmVAMP721d-positive vesicles were not transporting methylesterified pectin to symbiosoems, but most likely deliver enzymes involved in pectin
turnover, as GmVAMP721d vesicles partly co-localize with pectate lyase GmNPL1. This
is an ortholog of the nodule pectate lyase (LjNPL) from Lotus japonicus (Xie et al., 2012).
The defect in bacterial release, which we have observed in GmVAMP721dsilenced nodules, can be partly explained by a defect in pectin turnover that leads
to excess deposition of both forms of pectin in the lumen of bacterial clusters. The
physical constraints and compromised rheological characteristics of the matrix of
bacterial clusters, which become too adhesive, may explain the observed block of
bacterial release. The importance of pectin turnover in symbiosis has previously been
demonstrated in M. truncatula (Rodríguez-Llorente et al., 2004) and in L. japonicus
(Xie et al., 2012). The observation of infection process in the roots of LjNPL mutants
shows the defect in nodule formation. Two LjNPL mutants produced uninfected
nodules and most infections arrested as infection foci in root hairs or roots. Authors
show the diminished density of the labelling of de-methyl-esterified pectin on the
site of bacteria release implying that the pectin can be degraded locally in cell wallfree regions of infection thread (unwalled droplets). According to the data obtained
on Medicago nodules by Ivanov et al. (2012) VAMP721-positive vesicles are located
near the unwalled droplets in Medicago. In soybean nodule, they are found around
unwalled regions of the infection threads (Chapter 2) and show spatial and temporal
co-localization with pectate lyase GmNPL1, the ortholog of LjNPL of L. japonicus.
The analysis of pectin localization in soybean nodules also showed that young
infected cells contain endosomes carrying the de-methyl-esterified pectin that
implies that these cells have high pectin turnover. Endocytosis of pectin has
previously been shown in Arabidopsis thaliana root meristem cells and is assumed to
be important for the maintenance of plant cell wall integrity during cell expansion
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General Discussion
and division (Baluška et al., 2005). It may have similar functions in young infected cells
of root nodule. De-methylated pectin is less flexible and more adhesive (Wolf et al.,
2009) than methyl-esterified pectin. The development of infected cells results in the
coordinated expansion of infected host cells, microsymbiont and its surrounding cell
wall-free symbiosome membrane (Ivanov et al., 2012; Dörmann et al., 2014; LeborgneCastel & Bouhidel, 2014). This requires modification of plant cell wall (Brewin,
2004; Giordano & Hirsch, 2004). Replenishing the pool of methyl-esterified pectin
and depleting the de-methyl-esterified pectin may give to cell walls more elasticity
(Peaucelle et al.; Bárány et al., 2010). Therefore, the biological role of VAMP721d in part
is explained by its action in delivering pectin modifying enzymes to the site of release
of rhizobia (Chapter 2).
Is membrane tension may create a vector for vesicle fusion to the
interfacemembranein infected cells?
A strong indication that mechanical sensing plays a role in rhizobial infection
process came from studies of Esseling et al. (2004). They showed that root hairs of
mutants of SymRK and CCaMK are upon Nod factor perception, are very sensitive
to mechanical stimuli. Due to the expansion of rhizobia ininfected cells the putative
points of membrane tension are infection threads, unwalled droplets and young
symbiosomes. To determine whether it is the case we studied MtSyt2 and MtSyt3,
two M. truncatula homologs of Arabidopsis thaliana synaptotagmins from the group of
synaptotagmin 1 (SYT1) (Schapire et al., 2008)2008.
Synaptotagmins is a family of membrane trafficking proteins that function as Ca2+
sensors in a number of SNARE-dependent plasma membrane vesicle fusion processes,
such as exocytosis and neurotransmitter release (Lynch et al., 2007). In plant cells it is
situated in the plasma membrane region (Schapire et al., 2009; Yamazaki et al., 2010).
In M. trincatula nodules, synaptotagmin 1 components named MtSyt2 and MtSyt3
occur in the meristematic region and infection zone. RNAi silencing of MtSyt2 and
MtSyt3 caused the distortion in the number of the cell layers in nodule developmental
zones and the delay in symbiosome maturation. The RNAi-affected nodules display
a short meristem, extended zone of infection, diminished or aborted zone of fixation
and delay in symbiosome growth and maturation (Chapter 5). The phenotype of the
nodule has some similarities with the mutant dnf1 (Wang et al., 2010)2010 where
the development of the symbiosomes was restricted to stage 1-2. DNF1 encodes a
symbiosis-specific subunit of the signal-peptidase complex (Van de Velde et al., 2010;
Wang et al., 2010) and is important for the targeting of proteins to the symbiosomes.
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The morphological similarity may point to a link between the vesicular fusion to the
plasma membrane and symbiosis-specific secretory pathway dependent on DNF1. It
ispossible that the membrane tension created by expanding microsymbionton the
membranes of infection threads/unwalled droplets may create a vector for vesicle
fusion to these interface membranes (Chapter 5).
A plant cell without a functional vacuole; biological curiosity or a requirement for
successful symbiosis?
The development of infected cells starts when rhizobia are released into cells of
the cell layer directly adjacent to the meristem. Only in this layer release occurs and it
does not occur anymore in older cells despite the presence of infection threads (Vasse
et al., 1990; Monahan-Giovanelli et al., 2006). The colonization of host cell by rhizobia
has a profound effect on the endomembrane system of the host cell. A major change
concerns the biggest organelle of the plant cell, the vacuole. In determinate nodules
(soybean) the proliferation of symbiosomes in young infected cells is accompanied by
the fragmentation of vacuoles and a reduction of their size. In mature infected cells
the vacuole is completely absent (Chapter 2). In indeterminate nodules (Medicago)
the fragmentation of the vacuoles occurs at the transition of infection to the fixation
zone. In these cell layers the vacuoles of infected cells lose up to 75% of their volume
and this is accompanied by tonoplast folding. The reduction of vacuole volume
permits the expansion of bacteria in the host cell cytoplasm that can completely fill
the host cell (Gavrin et al., 2014; Chapeter 3).
To answer the question what causes the impairment in vacuole features during
the intracellular bacteria accommodation we have studied components of the HOPS
complex, the key regulator of vacuole formation (Nickerson et al., 2009; Balderhaar
& Ungermann, 2013). The expression of HOPS genes is switched off at the transition
of infection to fixation zone, so the same cell layer where vacuole collapses (Chapter
3). After being infected the vacuole undergoes a deacidification that may reduce
its lytic properties and affect endocytotic trafficking (Dettmer et al., 2006). This
probably suppresses the fusion of symbiosomes with themselves as well as with other
endosomes and young vacuoles and ensures the maintenance of the symbiosomes
as individual units. This is similar to the suppression of fusion of bacteria-containing
vacuoles with lysosomes in mammalian cells, as this also involves the deacidification
of phagosomes as well as lysosomes (Huynh & Grinstein, 2007; von Bargen et al.,
2009). Hence, vacuole defunctionalization seems a general mechanism by which
microbes are hosted in eukaryotic cells.
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General Discussion
Symbiosome membrane identity depends on membrane retargeting and
developmental stage of the symbiosomes
Rhizobia and the host cell are separated by the symbiosome membrane, the
membrane interface, which facilitates the exchange of nutrients and solutes. Such
an interface is also formed around intracellular arbuscles of endomycorrhizae and
haustoria from biotrophic fungi and oomycetes. The formation of the interface
in plant-microbe interactions has become an important topic (Ivanov et al., 2012;
Leborgne-Castel & Bouhidel, 2014). The interface membrane features, similar to
other membranes, are dependent on biosynthesis and the traffic of proteins and
lipids. For temporary membrane units such as symbiosomes it is especially important
to be recognized by the host cell as a genuine host structure to avoid the default
elimination of bacteria. On the other hand the membrane should be quite dynamic
in its transport and morphological features to support maintenance of symbiosomes
through different functional and developmental stages: from the small freshly
entered dividing bacteria to the mature nitrogen-fixing bacteroids and finally as lytic
compartment during senescence. This requires constant changes in the membrane
interface features. How is this achieved in infected nodule cells?
Important regulators of membrane properties are the proteins, which are
involved in the correct fusion with vesicles and hence regulate features of membrane
compartments. These are termed “identity markers” (Behnia & Munro, 2005). Different
cell membrane compartments have specific sets of identity markers. These proteins
are involved in the process of membrane fusion: regulatory GTPases of the Rab family
and Rab-interacting integral proteins of SNARE (soluble NSF attachment protein
receptor, where NSF stands for N-ethyl-maleimide-sensitive fusion protein) families
(Lipka et al., 2007). A SNARE complex includes t-SNAREs (syntaxins) and v-SNAREs
(VAMPs). Several other proteins associated with the membrane fusion are involved in
the positioning of the complex on the membrane like complexin and synaptotagmins
(Südhof & Rothman, 2009). Recent studies on SNARE localization in plants showed
that most SNAREs are associated with specific intracellular compartments (Ueda,
2004). Exocytotic plasma membrane targeted traffic in plants is mediated by
v-SNAREs belonging to the VAMP72 (vesicle-associated membrane protein) family.
It has been shown that specific VAMP72 genes are recruited in the interaction of
Arabidopsis with biotrophic fungi and are indispensable for the symbiosis with
rhizobia and with endomycorrhizae fungi (Sanderfoot et al., 2001; Ivanov et al., 2010).
An integral characterization of the symbiosome membrane shows that that during
their lifespan, symbiosomes recruit via vesicle-mediated transporta plethora of
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proteins, which are involved in the transport of water, ions, and other vital compounds
(Brear et al., 2013; Udvardi & Poole, 2013). Further, these studies show that it has a
mixed identity (Catalano et al., 2004; Colebatch et al., 2004). This identity changes at
specific developmental stages. Young freshly released symbiosomes accept plasma
membrane-residing Syntaxin134 and plasma membrane-targeted vesicle associated
proteins of the VAMP72 group (Limpens et al., 2009; Ivanov et al., 2012). By the end of
symbiosome differentiation symbiosomes accept endosomal/vacuolar marker Rab7.
On the later stage of development they recruit tonoplast markers, vacuolar SNAREpin complex (SYP22, SYP51 and VTI11) (Limpens et al., 2009). During the symbiosis
termination and the lysis of symbiosomes (Pérez Guerra et al., 2010; Laurence Dupont
& Sarra El Msehli, 2012), the symbiosome membrane receives HOPS complex proteins,
VPS11 and VPS39, that allow the lytic machinery of the vacuole/autolysosomes to
process the remnants of the bacteria and collapsing host cell (Gavrin et al., 2014;
Chapter 3).
At the transition of infection to fixation zone tonoplast proteins are retargeted
toward the symbiosome membrane (Limpens et al., 2009; Gavrin et al., 2014; Chapter
3). This indicates a loss of specificity of vesicle fusion. This may be caused by the
repression of the HOPS complex and vacuole defunctionalization in infected cells.
However, the loss of correct targeting to tonoplast, which would be detrimental for a
normal non-infected cell, appears to have a special function in symbiosis. For example
the retargeting of aquaporin TIP1g from the tonoplast to the symbiosome membrane
in the nodules of M. truncatula is crucial for symbiosome maturation to the nitrogenfixing stage (Gavrin et al., 2014; Chapter 3). Therefore, it is very likely that modulation
of vesicle traffic towards the symbiosome membrane is carefully crafted to support
microsymbiont accommodation and maintenance.
Changing the cell architecture: temporary/spatial modifications of actin
microfilaments network in infected cell
To accommodate thousands of rhizobia in infected nodule cells the cytoskeleton
of the host cell is rearranged (Dörmann et al., 2014) . For example the pattern of actin
microfilaments in the infected cells is modified (Whitehead & Day, 1997; Davidson
& Newcomb, 2001; Fedorova et al., 2007; Timmers, 2008). Our studies on ARP3 actin
nucleating factor in infected cells showed that the configuration of actin cytoskeleton
network has a dynamics that is related to symbiosome maturation. In the cource of
the development of infected cell the actin-nucleating factor ARP3 is taking part in the
formation of a subtle net surrounding each symbiosome (Roux et al., 2014; Gavrin et
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General Discussion
al., 2015; Chapter 4). During symbiosome growth, the subtle actin network starts to be
formed. This results in a network of microfilaments that envelop mature symbiosomes.
This actin network around the symbiosomes may be highly dynamic and support
vesicle transport to support growth of symbiosomes. Actin is known to organize the
plant cytoplasm (Yanagisawa et al., 2013) and may be essential for the organization
of narrow zones of cytoplasm around the symbiosomes for efficient transport
of membrane vesicles. The dynamic cortical cytoskeleton adheres to the plasma
membrane and controls the mobilisation of material into this expanding membrane
supporting growth of infected cells. The localization and functional analyses of ARP3
has shown that it is operational in the assembly of actin around symbiosomes and
this is indispensable for their maturation and maintenance. Growth of symbiosomes
is dependent on host cell membranes provided in the form of post-Golgi vesicles and
ER (Kereszt et al., 2011; Ivanov et al., 2012). The integration of ARP2/3 into branched
actin filament networks may facilitate the physical anchoring of a polymerizing actin
filaments on an organelle surface (Yanagisawa et al., 2013). The ARP2/3 complex is
strongly associated with membranes from several organelles including ER, tonoplast,
plasma membrane and early endosome in Arabidopsis (Kotchoni et al., 2009; Zhang
et al., 2013) that may facilitate membrane traffic. It might have a similar function in
infected symbiotic cells.
During the transition from an immature stage of development to the stage
of nitrogen fixation, the actin pattern in infected cell undergoes prominent
reorganization. The actin microfilaments in the cytoplasm of the host cell form the
network around the mature symbiosomes and ARP2/3 complex is operational in
this actin rearrangement. We hypothesize that the retargeting of late endosome/
tonoplast proteins to the symbiosome membranes allows the recruitment of actin
to the symbiosomesand that this recruitment is necessary for their functional
maturation. The ARP2/3 complex is functional in the rearrangement of actin around
symbiosomes. Silencing of ARP3 negatively affects the symbiosome maturation
(Gavrin et al., 2015; Chapter 4).
So, studies on mechanisms controlling rhizobial accommodation in specialized
infected cells shows that this is a multi-step process. The adaptation of the host cell
endomembrane system for intracellular bacteria comes not as a single tune, but as
a big orchestra playing a carefully arranged symphony to ensure efficient symbiosis.
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Moling S, Pietraszewska-Bogiel A, Postma M, Fedorova E, Hink MA, Limpens E, Gadella TWJ, Bisseling T.
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6
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Yamazaki T, Takata N, Uemura M, Kawamura Y. 2010. Arabidopsis Synaptotagmin SYT1, a Type I Signal-anchor
Protein, Requires Tandem C2 Domains for Delivery to the Plasma Membrane. The Journal of Biological
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Summary
The symbiosis of legumes with rhizobia leads to the formation of root nodules.
Rhizobia which are hosted inside specialized infected cells are surrounded by hostderived membranes, forming symbiosomes. Although it is known that symbiosome
formation involves proliferation of membranes and changing of host cell architecture
the mechanisms involved in these processes remain largely uncovered.
In this thesis, I studied in more detail the adaptation of the endomembrane
system of infected cells to intracellular rhizobia. I have shown that in the first cell layer
of the nitrogen-fixing zone, the vacuole of the infected cells shrinks, creating space for
the expanding symbiosomes. Here the expression of homotypic fusion and vacuole
protein sorting complex (HOPS) genes VPS11 and VPS39 are switched off, whereas
tonoplast proteins, like the vacuolar aquaporin TIP1g, are targeted to the symbiosome
membrane. These observations suggest that tonoplast-targeted traffic in infected
cells is altered. This retargeting is essential for the maturation of symbiosomes.
Accommodation of intracellular rhizobia requires also the reorganization of
the actin cytoskeleton. I have shown that during symbiosome development the
symbiosomes become surrounded by a dense actin network and in this way, the actin
configuration in infected cells is changed markedly. The actin nucleating factor ARP3
is operational in the rearrangement of actin around the symbiosome.
It is known that the plasma membrane is inelastic; its capacity to stretch is
only around 1-3%. Exocytosis of new membrane material is therefore involved in
changes in the size of the membrane surface and in repair of damaged membrane
loci . Membrane tension may create a vector for the fusion of membrane vesicles.
To test this, the localization of proteins from the group of synaptotamin calcium
sensors involved in membrane fusion, was studied. I have shown that the Medicago
synaptotamins, MtSyt2 and MtSyt3, are localised on protrusions of the host plasma
membrane created by expanding rhizobia (infection threads, cell wall-free unwalled
droplets). Hence, at these sites of contact between symbionts membrane tension may
create a vector for exocytosis.
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It is known that the host cell wall is modified during the development of infected
cells. This process is mediated by the exocytotic pathway employing vesicle-associated
membrane proteins (VAMPs) from the VAMP721 family. Previously it was shown in
Medicago nodules, that cell-wall free interface membrane formation during bacterial
release is dependent on these proteins. I have shown that the pectin modifying
enzyme pectate lyase is delivered to the site of bacterial release in soybean nodules
by VAMP721-positive vesicles.
My study uncovered new mechanisms involved in the adaptation of host cells
to intracellular rhizobia: defunctionalization of the vacuole, actin cytoskeleton
rearrangement and the retargeting of host cell proteins to the interface membrane.
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Samenvatting
De symbiose van vlinderbloemige planten met rhizobium bacteriën leidt tot
de vorming van stikstof bindende wortelknollen. In deze knollen zijn de bacteriën
gehuisvest in gespecialiseerde geïnfecteerde cellen. Ze zijn in deze cellen aanwezig
als organel-achtige structuren omringd door een membraan van de gastheer. Deze
organel-achtige structuren worden symbiosomen genoemd. Het is bekend dat de
vorming van een symbiosoom gepaard gaat met membraanproliferatie en aanpassing
van de celarchitectuur. Echter, de mechanismen die deze processen aansturen zijn
onbekend. In dit proefschrift heb ik deze mechanismen bestudeerd.
Ik heb laten zien dat in de eerste cellaag van de fixatiezone de vacuole van de
geïnfecteerde cellen plotseling in elkaar krimpt. Hiermee wordt ruimte gecreëerd voor
de groeiende symbiosomen. Dit proces is strikt gecorreleerd met het uitschakelen
van genen die coderen voor subunits van het HOPS-complex, dat essentieel is voor de
vorming van vacuolen. Verder komen eiwitten die normaal naar de vacuolemembraan
worden getransporteerd op de symbiosoommembraan terecht. Dit laatste bleek
essentieel voor een volledige ontwikkeling van symbiosomen.
Voor de huisvesting van rhizobia binnen de geïnfecteerde cellen is een
reorganisatie van het actineskelet nodig. Ik heb laten zien dat symbiosomen
tijdens hun ontwikkeling omringd worden door een dicht actinenetwerk. Bij deze
verandering van de organisatie van het actineskelet speelt ARP3 (actin nucleating
factor) een belangrijke rol.
Een belangrijke vraag bij de vorming van membraancompartimenten is hoe
vesicles met het juiste targetmembraan fuseren. Spanning in een targetmembraan
kan een mechanisme zijn. Omdat rhizobia delen en groeien, is het mogelijk dat zij
het gastheermembraan onder spanning brengen. Synaptomines spelen een rol bij de
fusie van vesicles met membranen die onder spanning staan. Ik heb laten zien dat in
Medicago de synaptotamines MtSyt2 en MtSyt3 te vinden zijn op de infectiedraden
in knollen daar waar rhizobia vrij komen uit deze draden. Spanning op deze plaatsen
in de infectiedraden zou daarom een vector kunnen vormen voor de targetting van
vesicles.
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Bij het vrijkomen van de rhizobia uit infectiedraden worden er celwandvrije
membraandomeinen gecreëerd. Eerder is aangetoond dat eiwitten van de VAMP721familie hierbij een cruciale rol spelen. Ik heb laten zien dat vesicles gemerkt met met
VAMP721 pecate lyase, een enzym dat pectine afbreekt, transporteren naar deze
celwandvrije membraandomeinen. Het is waarschijnlijk dat dit enzym een rol speelt
bij de afbraak van materiaal in de plantencelwand op de plaats waar celwandvrije
membraandomeinen worden gevormd.
Mijn studies zoals die in dit proefschrift beschreven staan, hebben nieuwe
inzichten opgeleverd in mechanismen die ten grondslag liggen aan processen
waarmee de plantencel zich aanpast aan de intracellulare infectie door rhizobium.
Dit betreft het uitschakelen van de vacuole, de reorganisatie van het actineskelet
en een veranderde targetting van eiwitten in het vacuolemembraan naar de
symbiosoom.
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Acknowledgments
Working at the Laboratory of Molecular Biology was an amazing experience,
which is impossible to overestimate. First of all, I would like to thank my promoter,
Prof. Dr. Ton Bisseling, and my co-promoter, Dr. Elena Fedorova for the opportunity
to do my PhD at MolBi. I am very grateful for all their knowledge and experience that
they have been shearing with me during these years.
Years of my PhD were quite challenging but very interesting. This was a time of
never-ending experiments, hot disputes and true friendship. I am very thankful to
all people from the Laboratory of Molecular Biology for nice working atmosphere,
support and help with experiments, critical discussions and a great time at social
events such as lab trips, Friday drinks, barbeques, dinners and etc. Without this
support my work would have been very dull and unexciting.
Very special thanks to Maria and Marie-Jose for their great help with all nonscientific issues, documents and other administrative work. You make life of
international students much and much easier.
Wageningen is a very particular place, small town where you can meet people
from all parts of the world. I would like to thank all of my friends, soulmates and
housemates who made my life in Wageningen unforgettable.
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List of publications
Aleksandr Gavrin, David Chiasson, Evgenia Ovchinnikova, Brent N. Kaiser, Ton
Bisseling, Elena E. Fedorova. (2015) VAMP72d protein influences bacterial release
from infection threads by coordinating pectin dynamics in soybean nodules. (Accepted
with revision in New Phytologist)
Clarke, V.C., Loughlin, P.C., Gavrin, A., Chen, C., Brear, E.M., Day, D.A., and Smith,
P.M.C. (2015). Proteomic analysis of the soybean symbiosome identifies new symbiotic
proteins. Molecular & Cellular Proteomics.
Gavrin, A., Jansen, V., Ivanov, S., Bisseling, T., and Fedorova, E. (2015). ARP2/3Mediated Actin Nucleation Associated with Symbiosome Membrane is Essential for
the Development of Symbiosomes in Infected Cells of Medicago Truncatula Root
Nodules. Molecular Plant-Microbe Interactions.
Gavrin, A., Kaiser, B.N., Geiger, D., Tyerman, S.D., Wen, Z., Bisseling, T., and Fedorova,
E.E. (2014). Adjustment of Host Cells for Accommodation of Symbiotic Bacteria:
Vacuole Defunctionalization, HOPS Suppression, and TIP1g Retargeting in Medicago.
The Plant Cell 26, 3809-3822.
Kuyukina, M.S., Ivshina, I.B., Gavrin, A.Y., Podorozhko, E.A., Lozinsky, V.I., Jeffree, C.E.,
and Philp, J.C. (2006).Immobilization of hydrocarbon-oxidizing bacteria in poly(vinyl
alcohol) cryogels hydrophobized using a biosurfactant. Journal of Microbiological
Methods 65, 596-603.
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Curriculum Vitae
Aleksandr Gavrin was born on 15th of May 1983 in Perm, Russia, a city near the
Ural Mountains, the natural boundary between Europe and Asia. After finishing the
school in 2000, he started his study in microbiology at Perm State University. During
this time Aleksandr performed the first research project as a BCs and MSc student
at the Laboratory of Alkanotrophic Microorganisms, Institute of ecology and genetics
of microorganisms, under the supervision of Prof. Dr. Irina B. Ivshina and Dr. Maria S.
Kuyukina. At that time, Aleksandr obtained his first experience in scientific research,
laboratory work, as well as critical thinking and scientific analysis. Soon after obtaining
the MSc degree in Microbiology, Aleksandr joined the Laboratory of Molecular
Biology at Wageningen University, The Netherlands. He started his PhD in December
2008 under the supervision of promoter Prof. Ton Bisseling and co-promoter Dr. Elena
Fedorova. Within his PhD project he studied molecular mechanism of intracellular
accommodation of bacteria during development of Rhizobium-legume symbiosis. This
work was presented on several conferences and in the form of available publications.
In April 2013 he continued investigation of symbiosis during his first post-doctoral
research in University of Sydney, Australia.
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Education Statement of the Graduate School
Experimental Plant Sciences
Issued to:
Date:
Group:
University:
Aleksandr Gavrin
2 September 2015
Laboratory of Molecular Biology
Wageningen University & Research Centre
1) Start-up phase
►
First presentation of your project
Rhizobial symbiosome formation and its relation to plant defens
►
►
►
►
Writing or rewriting a project proposal
Writing a review or book chapter
MSc courses
Laboratory use of isotopes
date
May 2009
Subtotal Start-up Phase
2) Scientific Exposure
►
►
►
►
►
►
date
EPS PhD student days
EPS PhD student day, Leiden University
EPS PhD student day, Utrecht University
EPS PhD student day, Amsterdam University
EPS theme symposia
EPS theme 1 'Developmental Biology of Plants', Leiden University
EPS theme 1 'Developmental Biology of Plants', Wageningen University
EPS theme 1 'Developmental Biology of Plants', Leiden University
NWO Lunteren days and other National Platforms
ALW meeting 'Experimental Plant Sciences', Lunteren
ALW meeting 'Experimental Plant Sciences', Lunteren
ALW meeting 'Experimental Plant Sciences', Lunteren
Feb 26, 2009
Sep 30, 2010
Nov 30, 2012
Feb 26, 2009
Jan 28, 2010
Jan 20, 2011
Apr 07-08, 2009
Apr 19-20, 2010
Apr 04-05, 2011
Seminars (series), workshops and symposia
Invited seminar Florian Frugier
Invited seminar Matteo Brilli
EPS workshop “Plant Endomembranes”
Invited seminar Peter Cook
Invited seminar Jan Henderson
Invited seminar 'Virtues and vices of plant modelling'
Invited seminar Angus Buckling
Mar 24, 2010
Apr 29, 2010
Jul 02, 2010
Oct 27, 2010
Dec 13, 2010
Oct 11, 2011
Oct 20, 2011
Plant Sciences Seminar ‘Does gender matter (in management)?'
Invited seminar Veronica Grieneisen
Invited seminar Ruth Finkelstein
Invited seminar Larry Griffing
Invited seminar Cristel Carles
‘Writing for high impact journals’ by Andrew Sugden, Editor of Science
International symposia and congresses
2rd European retreat of PhD students in Plant Sciences, Cologne, Germany
9th European Nitrogen Fixation Conference, Geneva, Switzerland
3rd European retreat of PhD students in Plant Sciences, Orsay, France
Exocytosis in Animals, Fungi and Plants, London, United Kingdom
Nov 08, 2011
Nov 17, 2011
Nov 14, 2012
Nov 16, 2012
Jan 16, 2013
Feb 08, 2013
Apr 15-17, 2010
Sep 06-10, 2010
Jul 05-08, 2011
Sep 19-21, 2011
Nov 27-Dec 01, 2011
17th Nitrogen Fixation Congress, Perth, Australia
Presentations
Plant Retreat for PhD students 2010, Cologne, Germany (Poster)
9th European Nitrogen Fixation conference, Geneve (Poster)
Course The Legume-Rhizobium Symbiosis: from molecules to farmers fields (Poster)
Plant Retreat for PhD students 2011, Orsay, France (Talk)
Exocytosis in Animals, Fungi and Plants, London, United Kingdom (Poster)
►
►
Apr 15-17, 2010
Sep 06-10, 2010
Oct 18-22, 2010
Jul 05-08, 2011
Sep 19-21, 2011
IAB interview
Meeting with a member of the International Advisory Board of EPS
Excursions
Febr 18, 2011
Subtotal Scientific Exposure
3) In-Depth Studies
►
►
►
►
►
16.9 credits*
date
EPS courses or other PhD courses
PhD training course "Bioinformatics - A User's Approach" Wageningen University, Wageningen, The Netherlands
Postgraduated Course The Legume-Rhizobium Symbiosis: from molecules to farmers fields
Journal club
Member of the literature discussion group of Mol. Biology
Individual research training
Rainer Hedrich lab, Wurzburg, Germany
Brent Kaise lab, Adelaide, Australia
Subtotal In-Depth Studies
4) Personal development
►
1.5 credits*
Aug 30- Sep 03, 2010
Oct 18-22, 2010
2009-2012
Aug, 2011
Feb-Jul, 2012
9.0 credits*
date
Skill training courses
Academic Writing II
English listening and speaking IV
Organisation of PhD students day, course or conference
Membership of Board, Committee or PhD council
2010
2010
Subtotal Personal Development
TOTAL NUMBER OF CREDIT POINTS*
Herewith the Graduate School declares that the PhD candidate has complied with the educational requirements set by the
Educational Committee of EPS which comprises of a minimum total of 30 ECTS credits
3.6 credits*
31,0
* A credit represents a normative study load of 28 hours of study.
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Cover art:
El Lissitzky: painting "Proun"
Alexander Rodchenko: font "Rodchenko"
Cover and Lay-out: Ferdinand van Nispen, Citroenvlinder-dtp.nl, Bilthoven,
the Netherlands
Printed by: GVO drukkers & vormgevers, Ede, the Netherlands
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