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Job Sharing in the Endomembrane System: Vacuolar
Acidification Requires the Combined Activity of
V-ATPase and V-PPase
Anne Kriegel,a Zaida Andrés,a Anna Medzihradszky,b Falco Krüger,a Stefan Scholl,a Simon Delang,a
M. Görkem Patir-Nebioglu,a Gezahegn Gute,c Haibing Yang,d Angus S. Murphy,c Wendy Ann Peer,c,e Anne Pfeiffer,b
Melanie Krebs,a Jan U. Lohmann,a and Karin Schumachera,1
a Department
of Plant Developmental Biology, Centre for Organismal Studies, Heidelberg University, 69120 Heidelberg, Germany
of Stem Cell Biology, Centre for Organismal Studies, Heidelberg University, 69120 Heidelberg, Germany
c Plant Science and Landscape Architecture, University of Maryland, College Park, Maryland 20742
d Horticulture and Landscape Architecture, Purdue University, West Lafayette, Indiana 47907
e Environmental Science and Technology, University of Maryland, College Park, Maryland 20742
b Department
ORCID IDs: 0000-0002-5418-6248 (H.Y.); 0000-0001-5649-7413 (A.S.M.); 0000-0003-0046-7324 (W.A.P.); 0000-0001-6484-8105 (K.S.)
The presence of a large central vacuole is one of the hallmarks of a prototypical plant cell, and the multiple functions of this
compartment require massive fluxes of molecules across its limiting membrane, the tonoplast. Transport is assumed to be
energized by the membrane potential and the proton gradient established by the combined activity of two proton pumps, the
vacuolar H+-pyrophosphatase (V-PPase) and the vacuolar H+-ATPase (V-ATPase). Exactly how labor is divided between these
two enzymes has remained elusive. Here, we provide evidence using gain- and loss-of-function approaches that lack of the
V-ATPase cannot be compensated for by increased V-PPase activity. Moreover, we show that increased V-ATPase activity
during cold acclimation requires the presence of the V-PPase. Most importantly, we demonstrate that a mutant lacking both
of these proton pumps is conditionally viable and retains significant vacuolar acidification, pointing to a so far undetected
contribution of the trans-Golgi network/early endosome-localized V-ATPase to vacuolar pH.
INTRODUCTION
The evolutionary success of higher plants is in large part due to
their unique cell architecture. Their often large cell volumes are
filled with a central vacuole containing mostly water and solutes
that allow plants to maximize collection of solar energy and mineral
nutrients by increasing the surface of their photosynthesizing and
nutrient-absorbing organs at minimal cost. Besides being lowcost space fillers, vacuoles are the main store for solutes and serve
as a hydrostatic skeleton that provides the driving force for cell
growth and reversible volume changes. They allow plants to adapt
to the fluctuating availability of essential nutrients, to detoxify the
cytosol when challenged by harmful molecules, and serve as
lysosome-like organelles in which endocytic and autophagic
cargo are digested (Marty, 1999; Martinoia et al., 2012). Although
vacuoles are generally multifunctional, a single function, like
protein storage, tends to prevail in a particular cell type or developmental stage (Zheng and Staehelin, 2011), and coexistence
of vacuoles with different functions in a single cell has been
demonstrated only in very few cases (Epimashko et al., 2004;
Frigerio et al., 2008). All vacuolar functions require massive fluxes
of molecules across the tonoplast, which are assumed to be
1 Address
correspondence to [email protected].
The author responsible for distribution of materials integral to the findings
presented in this article in accordance with the policy described in the
Instructions for Authors (www.plantcell.org) is: Karin Schumacher (karin.
[email protected]).
www.plantcell.org/cgi/doi/10.1105/tpc.15.00733
energized by the proton gradient and membrane potential created
by the combined activity of two proton pumps, the vacuolar
H+-ATPase (V-ATPase) and the vacuolar H+-pyrophosphatase
(V-PPase). Both proton pumps are highly abundant tonoplast
proteins, underlining that the amount of energy invested into vacuolar transport is substantial (Gaxiola et al., 2007; Schumacher,
2014). Pyrophosphate (PPi) is a by-product of many biosynthetic
processes, and the V-PPase could thus be the predominant vacuolar
proton pump in young, growing cells. However, the V-PPase has
also been discussed as a backup system for the V-ATPase under
ATP-limiting conditions like anoxia or cold stress, and it is generally
assumed that the combined action of V-ATPase and V-PPase enables plants to maintain transport into the vacuole even under
stressful conditions (Maeshima, 2000).
V-ATPases are multisubunit proton pumps found in all eukaryotes that consist of the peripheral V1 complex responsible for ATP
hydrolysis and the membrane-integral Vo complex responsible for
proton translocation. Localization of the V-ATPase is determined
by the isoforms of the membrane-integral Vo subunit VHA-a with
VHA-a1 mediating targeting to the trans-Golgi network/early
endosome (TGN/EE) and VHA-a2 and VHA-a3 conferring tonoplast localization (Dettmer et al., 2006). Arabidopsis thaliana
mutants that lack both tonoplast-localized isoforms are viable but
show a strong reduction in growth and nutrient storage capacity
(Krebs et al., 2010). The fact that the vha-a2 vha-a3 mutant still
maintains a 10-fold proton gradient across the tonoplast (vacuolar
pH 6.4 versus cytosolic pH 7.4) argues that the V-PPase, a homodimer of a single polypeptide chain, plays a more important
role in vacuolar acidification than the V-ATPase. Arabidopsis
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plants carrying a T-DNA insertion in the only gene encoding
a K+-stimulated Arabidopsis vacuolar PPase 1AVP1 or Arabidopsis vacuolar H+-PPase VHP1 were reported to show severe
developmental phenotypes caused by defects in auxin transport
(Li et al., 2005). However, additional independent alleles of AVP1/
VHP1, uncovered by the analysis of fugu5 mutants that failed
to support heterotrophic growth after germination, did not show
auxin-related phenotypes. Importantly, vacuolar pH in the fugu5
mutants was only mildly affected, and compensation by increased
V-ATPase activity was ruled out (Ferjani et al., 2011). Moreover, the
mild postgermination growth defect of fugu5 seedlings resulting in
a slightly different cotyledon shape could be rescued by expression of a soluble yeast pyrophosphatase (Ferjani et al., 2011),
highlighting a so far undiscovered role of the V-PPase in the removal of PPi required to avoid accumulation of inhibitory concentrations of PPi (Ferjani et al., 2012). These findings are also
highly relevant as overexpression of AVP1 in Arabidopsis, as well
as in a number of crop plants, results in improved drought and salt
tolerance (Gaxiola et al., 2001; Pasapula et al., 2011; Gamboa
et al., 2013; Schilling et al., 2014). AVP1 overexpression has also
been reported to result in increased cell division at the onset of
organ formation and increased auxin transport, which appeared to
be a consequence of increased DpHPM (visualized as cytosolic
alkalinization) resulting from altered distribution and abundance of
the plasma membrane (PM) H+-ATPase and the PIN-FORMED1
auxin efflux facilitator (Li et al., 2005). Although AVP1 clearly is an
abundant tonoplast protein (Segami et al., 2014), it has been
reported to be localized at the PM in sieve element companion
cells and upon overexpression also in other cell types (Langhans
et al., 2001; Paez-Valencia et al., 2011; Pizzio et al., 2015). The
mechanistic base of the beneficial traits achieved by overexpression of AVP1 is thus unclear, and it remains to be determined if and to what extent increased vacuolar solute
accumulation due to increased proton pumping by the V-PPase is
involved. By combining loss- and gain-of-function approaches, we
have addressed how V-ATPase and V-PPase share the job of
vacuolar acidification. Here, we show that lack of the V-ATPase
cannot be compensated for by increased V-PPase activity but also
that increased V-ATPase activity during cold acclimation requires
the presence of the V-PPase. Most importantly, we show that
a mutant lacking both tonoplast V-ATPase and V-PPase is viable and
retains significant vacuolar acidification, revealing the presence of
a so far unnoticed contribution of the TGN/EE-localized V-ATPase.
RESULTS
Lack of Tonoplast V-ATPase Activity Cannot Be
Compensated for by Increased V-PPase Activity
To investigate if a lack of tonoplast V-ATPase activity can be
compensated for by increased V-PPase activity, the vha-a2 vha-3
double mutant was crossed with AVP1-1, a transgenic line expressing AVP1 under the control of the double 35S promoter
(Gaxiola et al., 2001). In the segregating F2 generation, vha-a2 vha-a3
AVP1-1 plants were identified by genotyping and showed a small
increase in size compared with vha-a2 vha-a3 (Supplemental Figure
1A). Although it was reported previously that AVP1 protein levels are
increased in AVP1-1 plants (Gaxiola et al., 2001), we could not detect
ubiquitous AVP1 overexpression based on qPCR, RNA in situ hybridization, immunocytochemistry, and immunoblot analysis in the
vha-a2 vha-a3 mutant background as well as in the progeny of the
original transgenic AVP1-1 line (Supplemental Figures 1B to 1K).
Lack of AVP1 overexpression is most likely due to transgene silencing, as indicated by the complete absence or patchy kanamycin
resistance in AVP1-1 and AVP1-2 in subsequent generations
(Supplemental Figures 1M and 1N). We thus generated transgenic
lines expressing AVP1 under the control of the UBQ10 promoter,
which leads to constitutive and robust overexpression (Grefen et al.,
2010; Behera et al., 2015) and resulted in 2- to 3-fold higher V-PPase
activity (Figures 1A and 1B). Notably, constitutive overexpression of
AVP1 did not correlate with an increase in rosette size and fresh
weight (Figures 1C and 1D) or with altered auxin transport and
content as reported previously (Li et al., 2005; Supplemental Tables 1
and 2).
Importantly, overexpression of AVP1 in the vha-a2 vha-a3 mutant
background did not cause an increase in rosette size (Figures 2A to
2C) and did not affect leaf cell sap pH (Figure 2D) or root vacuolar pH
(Figure 2E). Taken together, these results show that constitutive
overexpression of the tonoplast V-PPase AVP1 does not necessarily
cause an increase in biomass in wild-type plants and that enhanced
V-PPase activity cannot compensate for a lack of tonoplast
V-ATPase activity under standard growth conditions.
A Second T-DNA Insertion in GNOM Is Responsible for the
avp1-1 Phenotype
The apparent discrepancy between the severe phenotype of
avp1-1 seedlings and the mild cotyledon shape phenotype of
vhp1-1 and fugu5 seedlings led us to test the hypothesis that the
avp1-1 allele is linked to a second-site mutation (Ferjani et al.,
2012). Therefore, whole-genome sequencing of the avp1-1 line
(GABI-Kat 005D04) was performed and identified a second T-DNA
insertion in the 39 end of At1g13980, the gene encoding the ARFGEF GNOM that is genetically linked to the insertion in AVP1
(At1g15690; Figure 3A; Supplemental Figure 2). GNOM is required
for polar auxin transport (Geldner et al., 2003) and weak alleles
cause phenotypes that are highly similar to the one observed in
avp1-1 (Geldner et al., 2004). To test if the additional T-DNA insertion in GNOM is indeed responsible for the avp1-1 phenotype,
we performed allelism tests among the avp1/vhp1/fugu alleles
and the gnom allele emb30-1 (Mayer et al., 1993). The gnom
phenotype was observed in 25% of seedlings in the F1 progeny
from a cross between the avp1-1/+ and emb30/+ lines, whereas
crosses of homozygous vhp1-1 and fugu5-1 plants to avp1-1/+
resulted in a 1:1 ratio of the wild type and fugu5-1/vhp1-1-phenotype seedlings (Figures 3B and 3C). Further comparison among
the AVP1/VHP1 loss-of-function alleles showed that auxin
transport and free indole-3-acetic acid (IAA) levels in the fugu5
alleles, unlike avp1-1, were similar to those in the wild type
(Supplemental Tables 3 and 4). Based on the combined results, we
concluded that the avp1-1 phenotype is caused by a linked
second site mutation in GNOM, whereas the fugu5/vhp1 phenotype reflects the lack of AVP1/VHP1 activity. Importantly, although V-PPase activity is not detectable in both vhp1-1 and
fugu5-1, we observed only a marginal increase of cell sap pH in
Job Sharing between V-ATPase and V-PPase
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Figure 1. Increased Biomass Does Not Require Constitutive AVP1 Overexpression.
(A) Increased AVP1 protein amounts in UBQ:AVP1 lines. Microsomal membrane extracts of 4-week-old wild-type, 35S:AVP1, and UBQ:AVP1 plants were
separated by SDS-PAGE and subsequently immunoblotted with anti-V-PPase antibody. Equal protein loading is indicated by anti-VHA-C detection.
(B) Elevated K+-stimulated PPase activity in UBQ:AVP1 but not in 35S:AVP1 lines. Plants were grown for 4 weeks under LD conditions. Wild type activity was
set to 100%. Graph shows result of one representative experiment of three biological replicates. Error bars indicate SD of n = 3 technical replicates.
(C) and (D) Phenotypes (C) and rosette fresh weight (D) of 4-week-old 35S:AVP1 plants compared with UBQ:AVP1 lines grown under LD conditions. Error
bars indicate SE of n = 8 to 11 plants. FW, fresh weight. Bar = 3.5 cm.
both mutants (Supplemental Figure 3B). However, under our
growth conditions, the rosette size of vhp1-1 mutants was reduced, whereas fugu5-1 was indistinguishable from the wild type
(Supplemental Figure 3A). Based on these findings, we used both
mutants, fugu5-1 and vhp1-1, in our further studies.
Upregulation of V-ATPase during Cold Acclimation Depends
on the Presence and Activity of V-PPase
Under standard growth conditions and based on steady state pH,
the V-PPase does not seem to make a major contribution to vacuolar acidification (Ferjani et al., 2011), and we thus next investigated
its role during cold acclimation, a process previously reported to
require increased tonoplast proton-pumping activity (Schulze et al.,
2012). In the wild type, cold acclimation for 4 d at 4°C led to a slight
reduction of leaf cell sap pH from 5.8 to 5.7 (Figure 4A), consistent
with increased amounts and activities of both V-ATPase and
V-PPase (Figures 4B to 4E). In both fugu5-1 and vhp1-1, vacuolar
pH was slightly higher than in the wild type under control conditions
but the cold-induced drop of cell sap pH was reduced. Conversely,
overexpression of AVP1 induced an even stronger pH decrease
(Figure 4A). Surprisingly, the cold-induced increase in V-ATPase
amount and activity was reduced in the V-PPase loss-of-function
mutants and enhanced in the overexpression line (Figure 4D),
indicating that upregulation of the V-ATPase during cold acclimation depends on the presence and activity of the V-PPase.
A Triple Mutant Lacking Both Tonoplast Proton Pumps
Is Viable
A triple mutant lacking V-ATPase and V-PPase would allow us to
address two important questions. First, if the presence of proton
pumps at the tonoplast is essential and, second, if the V-PPase
does indeed not contribute to vacuolar acidification under nonstress conditions, in which case we would expect no vacuolar pH
difference between vha-a2 vha-a3 and the triple mutant. In contrast, the pH gradient at the tonoplast would be even more reduced
in the triple mutant compared with vha-a2 vha-a3 if the lack of
proton pumping in vhp1-1/fugu5-1 single mutants is masked by
a compensatory mechanism. We thus crossed the tonoplast
V-ATPase-deficient vha-a2 vha-a3 mutant with plants homozygous for vhp1-1 or fugu5-1. In the segregating F2 progeny, we
were able to identify vhp1-1 vha-a2 vha-a3 and fugu5-1 vha-a2
vha-a3 seedlings; however, the vegetative growth of both triple
mutants was strongly impaired and seedlings only survived when
germinated and grown in sterile culture for a few days before
transfer to soil (Figure 5A). Flowers of the triple mutant had smaller
petals and shorter stamen filaments as well as nondehiscent
anthers, leading to strongly reduced fertility (Figure 5B). Siliques of
fugu5-1 vha-a2/+ vha-a3 plants were slightly shorter and contained
;10% aborted ovules, indicating that gametophyte development
was impaired. This became more evident as vhp1-1 vha-a2 vha-a3
mutants did not produce any seeds, whereas fugu5-1 vha-a2 vha-a3
plants produced few but viable seeds (Figure 5C). We next measured
cell sap pH using leaf material of 3-week-old plants grown on soil
under standard long-day (LD) growth and found it to be strongly
increased to pH 7.1 in the fugu5-1 vha-a2 vha-a3 triple mutant,
whereas cell sap pH for the fugu5-1 mutant was slightly increased
(pH5.8) and the vha-a2 vha-a3 mutant showed the expected increase to pH 6.4 (Figure 5D). Cell sap pH can be used as a robust
approximation of vacuolar pH, as the vacuolar lumen normally
occupies ;90% of the cell volume. To determine if this assumption is
also true for the triple mutant, we used BCECF staining to show that
although in the fugu5-1 vha-a2 vha-a3 mutant the cell size of both
epidermis and mesophyll cells was reduced, a large central vacuole
was clearly present (Figures 5E to 5J). Based on these findings, the
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Figure 2. Overexpression of AVP1 Does Not Complement the Tonoplast V-ATPase Double Mutant vha-a2 vha-a3.
(A) Arabidopsis wild-type and vha-a2 vha-a3 mutant plants expressing AVP1 under the UBQ promoter have higher AVP1 protein level. Microsomal
membrane proteins of 3-week-old plants were extracted, separated by SDS-PAGE, and subsequently immunoblotted with anti-V-PPase antibody. Equal
protein loading is indicated by TIP1;1 detection. A quantification of AVP1 protein levels is shown below and each bar corresponds with the band in the blot
immediately above it. Error bars indicate SE of n = 3 technical replicates.
(B) UBQ:AVP1 cannot restore wild type growth in vha-a2 vha-a3. Plants were grown for 3 weeks under LD conditions at 22°C. Bar = 1.75 cm.
(C) AVP1 overexpression lines show increased V-PPase activity. K+-stimulated PPase activity was determined using microsomal membranes extracted
from 3-week-old plants. Wild type activity was set to 100%. Error bars represent SE of n = 3 biological replicates.
(D) Overexpression of AVP1 has no effect on cell sap pH. Plants for cell sap pH measurements were grown for 3 weeks under LD conditions at 22°C. Error
bars show SD of n = 3 biological replicates.
(E) Vacuolar pH in root epidermal cells is not changed upon AVP1 overexpression. Vacuolar pH was measured in roots of 6-d-old plants. Error bars represent
SD of n = 3 biological replicates.
V-PPase AVP1/VHP1 acts as a proton pump and is responsible for
the remaining vacuolar acidification observed in leaves of the vha-a2
vha-a3 mutant. However, these results also raise the important
question of if and how cell expansion and vacuole formation can
occur in the absence of the two proton pumps.
Lack of V-ATPase and V-PPase Does Not Abolish Cell
Expansion and Vacuole Acidification in Hypocotyl and
Root Cells
Hypocotyl length of etiolated seedlings is solely based on cellular
expansion (Gendreau et al., 1997), and we found it to be reduced
by 50% in the fugu5-1 vha-a2 vha-a3 mutant (Supplemental Figure
4A). Similarly, root length was also reduced by ;50% due to
reduced cell expansion as the size of the meristematic zone was
found to be not significantly different between wild-type and
fugu5-1 vha-a2 vha-a3 seedlings (Supplemental Figures 4B and
4C). To visualize vacuolar morphology and to measure vacuolar
pH in the three developmental zones, roots were stained with
BCECF and analyzed by confocal laser scanning microscopy
(CLSM) (Viotti et al., 2013). Meristematic root cells of wild-type
roots contained a complex tubular vacuolar network surrounding
the nucleus (Figure 6A, bottom). In the root elongation zone, cells
start to rapidly expand accompanied by the inflation of vacuoles
Job Sharing between V-ATPase and V-PPase
Figure 3. T-DNA Insertion in GNOM Causes Phenotypic Defects in avp1-1.
(A) Position of the two T-DNA Insertions on chromosome 1 in the GABI-Kat
005D04 line. RB, right border; LB, left border. Bars = 500 bp.
(B) Comparison of the phenotypes of 5-d-old wild-type, fugu5-1, avp1-1,
and gnomR5 seedlings. Bar = 1 cm.
(C) Allelism test between V-PPase and gnom mutants. V-PPase mutants
(vhp1-1 and fugu5-1) were crossed with avp1-1/+ or with gnom/+ (emb301/+) mutants. Phenotypes of 6-d-old F1 seedlings (n = 80 to 190) were
counted. Numbers given in the table are percentages (%).
(Figure 6A, middle). Finally, in the differentiation zone, cells reach
their final size and are almost completely filled by the large central
vacuole (Figure 6A, top). No strong differences in vacuole morphology between the wild type and vha-a2 vha-a3 or fugu5-1
mutants were observed (Figures 6B and 6C). Only in the elongation
and transition zone of fugu5-1 vha-a2 vha-a3 roots was vacuole
morphology severely altered (Figure 6D, middle and top). Here,
vacuoles appeared as multiple spheres of different sizes distributed within the cell, whereas vacuoles in fully elongated cells of
the triple mutant have regained a completely normal shape. For pH
measurements, CLSM images of stained roots were analyzed and
the obtained emission ratios were converted to pH values using an
in situ calibration curve (Krebs et al., 2010). In general, all genotypes showed an increased vacuolar pH in the elongation zone,
but surprisingly the vacuolar pH in the different regions of fugu5-1
vha-a2 vha-a3 roots was not significantly increased when compared with vha-a2 vha-a3 (Figures 6E to 6G).
Vacuolar Acidification Is Abolished by the V-ATPase
Inhibitor Concanamycin A
The fact that the triple mutant is able to maintain a 10-fold proton
gradient across the tonoplast of root cells points to the contribution of an additional source of proton import into the vacuole. In
recent years, it has become clear that P-type H+-ATPases can be
found at the tonoplast in some cell types (Verweij et al., 2008;
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Appelhagen et al., 2015), and we thus assessed the effect of
ortho-vanadate, an inhibitor of P-type ATPases on media acidification (Supplemental Figure 5A), and vacuolar pH in roots using
BCECF. Although a slight increase of vacuolar pH was detected in
the roots of wild-type, vha-a2 vha-a3, and fugu5-1 plants, vacuolar
pH in the triple mutant fugu5-1 vha-a2 vha-a3 was not affected
by ortho-vanadate (Figure 7A). Moreover, although transcripts of
AHA10, the only tonoplast-localized P-type H+-ATPase of Arabidopsis identified to date (Appelhagen et al., 2015), are detectable
in roots, transcript levels are not increased in any of the V-ATPase
and/or V-PPase mutants (Supplemental Figure 5B). In contrast, the
V-ATPase inhibitor concanamycin A (ConcA) abolished vacuolar
acidification in all genotypes tested, including vha-a2 vha-a3 and
fugu5-1 vha-a2 vha-a3 (Figure 7B), leading to the accumulation of
autophagic bodies without affecting BCECF accumulation or
vacuolar morphology (Figure 7C). Importantly, TGN/EE-localized
V-ATPase complexes containing VHA-a1 are the only remaining
targets of ConcA in vha-a2 vha-a3 and fugu5-1 vha-a2 vha-a3. We
cannot rule out that a minor proportion of VHA-a1 is relocated to
the tonoplast in vha-a2 vha-a3; however, we could not detect
VHA-a1-GFP at the tonoplast using highly sensitive hybrid detection CLSM (Supplemental Figures 5C to 5F). We thus conclude
that the V-ATPase in the TGN/EE contributes to vacuolar acidification either by controlling trafficking of a yet unknown proton
pump or by vesicular transport of protons.
DISCUSSION
How Does AVP1 Overexpression Enhance Crop Yield?
Although constitutive overexpression of AVP1 was reported for
the two transgenic lines AVP1-1 and AVP1-2 (Gaxiola et al., 2001),
we could not detect increased AVP1 RNA or protein levels in
leaves and several other tissues. Given that a vector containing
several copies of the 35S promoter was used in the original
construct (Yoo et al., 2005) and that multiple copies of the T-DNA
have been detected in the transgenic lines (Gaxiola et al., 2001), it
is not unexpected that expression of the transgene was lost over
time presumably due to 35S-siRNA-mediated transgene silencing
(Daxinger et al., 2008; Mlotshwa et al., 2010). Silencing of
transgenes expressed under the control of the 35S promoter is
common in Arabidopsis; however, it is widely used in crop plants,
as it has been shown to cause constitutive and stable overexpression of many transgenes, including AVP1. 35S-driven
overexpression of AVP1 is a successful strategy to improve crop
performance. Increased biomass, higher stress tolerance, and
enhanced P solubilization are highly desirable traits that have been
achieved in many crop plants including rice (Oryza sativa), barley
(Hordeum vulgare), wheat (Triticum aestivum), cotton (Gossypium
hirsutum), sugarcane (Saccharum officinarum) (Yang et al., 2007;
Pasapula et al., 2011; Kumar et al., 2014; Li et al., 2014), and
tomato (Solanum lycopersicum), for which stable expression of
the transgene up to the T8 generation has been reported (Park
et al., 2005; Yang et al., 2014).
Despite the fact that constitutive overexpression of AVP1 in
Arabidopsis was lost due to transgene silencing in the original
lines, the progeny still display the significant increase in rosette
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Figure 4. Increase of V-ATPase Activity after Cold Acclimation Depends on V-PPase.
(A) Lack of V-PPase activity prevents full acidification upon cold acclimation. Plants for cell sap pH measurements were grown for 6 weeks under short-day
conditions at 22°C. Error bars represent SD of n = 3 biological replicates. Asterisk indicates significant difference at P < 0.05 (Student’s t test).
(B) V-PPase activity affects the induction of V-ATPase activity upon cold acclimation. KNO3-inhibited ATPase activity was determined with microsomal
membranes extracted from 6-week-old, short day-grown plants that were cold-acclimated for 4 d at 4°C. Error bars show SD of n = 3 biological replicates.
Asterisk indicates significant difference at P < 0.05 (Student’s t test).
(C) V-PPase activity of the wild type and UBQ:AVP1 increases after cold exposure. K+-stimulated PPase activity was determined from microsomal
membranes extracted from 6-week-old, short day-grown plants that were cold-acclimated for 4 d at 4°C. Error bars represent SD of n = 3 biological replicates.
Asterisk indicates significant difference at P < 0.05 (Student’s t test).
(D) and (E) Protein levels of V-ATPase and V-PPase increase upon cold exposure in the wild type and UBQ:AVP1. Microsomal membrane proteins of
6-week-old plants were extracted, separated by SDS-PAGE, and subsequently immunoblotted with VHA-C antibody (D) and anti-V-PPase antibody (E).
TIP1;1 detection was used as loading control. Protein levels were measured using ImageJ and normalized to TIP1;1. Protein levels at 22°C were set to 100%.
Bar charts represent quantification of one representative immunoblot.
biomass that was originally reported. In contrast, the lines overexpressing AVP1 established in this study only showed a marginal
increase in biomass, suggesting that constitutive overexpression
is neither necessary nor sufficient for improved growth and
possibly also stress tolerance. One possible scenario to explain
this is that overexpression is maintained in particular cell types or
tissues of AVP1-1 and AVP1-2 plants. It was shown recently that
guard cell-specific overexpression of the plasma membrane H+ATPase AHA2 causes enhanced stomatal aperture and increased
photosynthetic activity, ultimately resulting in larger leaves (Wang
et al., 2014). Similarly, based on the fact that AVP1 has been
proposed to function as a PPi synthase at the PM of sieve element
companion cell (Pizzio et al., 2015), restricted overexpression of
AVP1 in the phloem could be causative. The positive effects of
AVP1 overexpression were initially assumed to be solely due to the
increased capacity for cation transport into the vacuole, but it is
now clear that other roles of AVP1, e.g., in PPi homeostasis, need
to be considered. Under stress conditions such as salinity or
phosphate starvation, endogenous AVP1 expression is induced
along with an increase in rhizosphere acidification in the case of
phosphate starvation (Yang et al., 2007). In support of the acid
growth hypothesis, for the AVP1-1 line, auxin transport was reported to display increased acidification of the apoplast caused
by the higher presence and activity of P-type H+-ATPase at the
PM (Li et al., 2005). Importantly, no correlation between increased
AVP1 expression and altered auxin transport and content was
detected for the transgenic lines expressing AVP1 under control of
the UBQ10 promoter. This is in line with the fact that, based on
DR5:GUS expression and its response to exogenous IAA, the
fugu5-1 mutant has been shown to have a normal auxin distribution and response (Ferjani et al., 2011). In addition, we have
shown here that auxin transport and auxin content are indistinguishable from that of the wild type in all fugu5 alleles and
that the severe developmental phenotype of avp1-1 is due to the
presence of a second T-DNA insertion close to the 39 end of the
gene encoding the ARF-GEF GNOM that is required for PIN cycling. Taken together, these results show that AVP1 is not required
for auxin transport and that auxin might thus not be directly involved in and responsible for the enlarged root systems observed
in Arabidopsis and different crop species. Clearly, further insight
into the mechanisms underlying the beneficial traits is urgently
required and could lead to better-defined strategies for AVP1
expression in crop improvement.
Does AVP1 Act as a Proton Pump?
Based on the facts that high concentrations of PPi, a by-product of
many biosynthetic reactions, are present and that the V-PPase is
Job Sharing between V-ATPase and V-PPase
7 of 14
Figure 5. Vacuolar Proton Pump Triple Mutant Is Viable and Has a Neutral Cell Sap pH.
(A) fugu5-1 vha-a2 vha-a3 growth is even more inhibited than vha-a2 vha-a3. Wild-type and proton pump mutant plants were grown for 26 d under LD
conditions at 22°C. Bar = 1.75 cm.
(B) Proton pump triple mutants exhibit defects in flower development. Bright-field micrographs of dissected flowers are shown. Stamen filaments reached the
pistil in the wild type and the segregating proton pump triple mutant flower, whereas they are shortened in the fugu5-1 vha-a2 vha-a3 flower. Bar = 1 mm.
(C) Silique size of fugu5-1 vha-a2 vha-a3 is strongly reduced. Excised siliques of the wild type, fugu5-1 vha-a2/+ vha-a3, and fugu5-1 vha-a2 vha-a3 are
shown. Bar = 3 mm.
(D) Cell sap pH of fugu5-1 vha-a2 vha-a3 is nearly neutral. Plants were grown for 3 weeks under LD conditions at 22°C. Error bars show SD of n = 3 biological
replicates.
(E) to (J) Vacuoles of epidermal cells ([E] to [G]) and mesophyll cells ([H] to [J]) of rosette leaves of 5-week-old wild-type and vacuolar proton pump mutants
stained with BCECF (green). Autofluorescence of chloroplasts is shown in magenta. Bars = 25 µm.
more abundant than the V-ATPase in young tissues, the V-PPase
has been considered to be the main tonoplast proton pump in such
tissues (Maeshima, 2000). This notion was supported by our
previous work in which we found that the tonoplast V-ATPase-
deficient mutant vha-a2 vha-a3 maintained a 10-fold proton
gradient across the tonoplast (vacuolar pH 6.4 versus cytosolic pH
7.4; Krebs et al., 2010). In contrast, the role of the V-PPase as
a major proton pump was challenged by the finding that loss of
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Figure 6. Root Vacuole Morphology in Tonoplast Proton Pump Mutants.
(A) to (D) The shape of vacuoles was monitored in the root differentiation zone, elongation zone, and meristematic zone of 6-d-old wild-type (A), vha-a2 vhaa3 (B), fugu5-1 (C), and fugu5-1 vha-a2 vha-a3 (D) seedlings. Roots were stained with BCECF (green) and FM4-64 (red). Bars = 20 µm.
(E) to (G) Vacuolar pH measurement in three different root zones. pH values were determined in the root differentiation zone (E), elongation zone (F), and
meristematic zone (G) of 6-d-old wild-type and vacuolar proton pump mutant seedlings. Error bars represent SD of n = 3 biological replicates.
V-PPase activity in the fugu5-1 and vhp1-1 mutants only had
a mild effect on vacuolar pH. Whereas the vacuolar pH in seedlings
of fugu5-1 and vhp1-1 was shown to be 0.25 pH units more alkaline than in the wild type (Ferjani et al., 2011), we showed here
that the cell sap pH of mature rosette leaves of the V-PPase
mutants is only marginally changed (<0.1 pH units). The contribution of AVP1 to vacuole acidification, at least under standard
growth conditions, thus seemed questionable. However, it is
important to keep in mind that due to the flexible ATP/H+-coupling
rate of the V-ATPase, the lack of V-PPase activity might be masked
by increased V-ATPase H+-pumping activity. Although it has been
shown that in the fugu5 mutants the ATP hydrolysis rate was not
increased, proton-pumping activity was not determined. Moreover, we demonstrate here that overexpression of AVP1 does not
ameliorate the vha-a2 vha-a3 mutant phenotype, which also argues against an important role of the V-PPase in vacuolar acidification. Interestingly, the fact that in Arabidopsis lack of tonoplast
V-ATPase activity cannot be compensated for by increased
V-PPase activity stands in contrast to the fact that expression of
AVP1 rescues vph1, a yeast mutant lacking the vacuolar isoform of
the V-ATPase (Pérez-Castiñeira et al., 2011; Coonrod et al., 2013).
This is most easily reconciled by assuming that the activities of
V-ATPase and V-PPase depend on each other and our results
concerning cold acclimation support such a scenario. The massive increase in the concentration of sugars and organic acids in
the vacuole, one of the most prominent metabolic changes that
reaches a maximum after 4 to 5 d of cold acclimation, goes along
with an increase in V-ATPase amount and activity, whereas the
V-PPase remained largely unaffected (Schulze et al., 2012).
Surprisingly, we could show here that cold-induced upregulation of the V-ATPase depends on the presence of the V-PPase.
Coregulation of V-ATPase and V-PPase has previously been
proposed based on the observation that in the cassulacean acid
metabolism plant Kalanchoe blossfeldiana H+-pumping by the
V-ATPase could be increased by ;300% when vesicles were
preenergized by a short incubation with PPi (Fischer-Schliebs
et al., 1997). The underlying mechanism remains to be elucidated, but a direct physical interaction as suggested previously
(Fischer-Schliebs et al., 1997) seems likely to be involved. Although the membrane potential difference between the vacuolar lumen and the cytosol is generally reported to be rather low
(#30 mV), it will be important to determine how this physiological parameter is affected in mutants lacking one or both
vacuolar H+-pumps. Determination of the tonoplast membrane
potential is challenging as it can currently only be measured via
impalement with double-barreled microelectrodes (Miller et al.,
2001; Wang et al., 2015) and novel tools that allow in vivo imaging of tonoplast membrane potential based on voltagesensitive fluorescent proteins are highly desirable (Baker et al.,
2008).
Job Sharing between V-ATPase and V-PPase
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Figure 7. Inhibition of V-ATPase Abolishes Vacuolar Acidification.
(A) Inhibition of P-type ATPases has minor effect on vacuolar pH. Vacuolar pH was measured in roots of 6-d-old seedlings after treatment with (+) or without
(2) 2 mM vanadate for 18 h. Error bars represent SD of n = 3 biological replicates.
(B) ConcA-mediated inhibition of V-ATPase blocks vacuolar acidification. Vacuolar pH was measured in roots of 6-d-old plants. Seedlings of Arabidopsis
wild-type and vacuolar proton pump mutants were incubated for 20 h in 1 µM ConcA (+) or DMSO (2) prior to pH measurements. Error bars represent SD of
n = 3 biological replicates.
(C) Vacuolar morphology after ConcA treatment. Wild-type seedlings were incubated for 20 h in 1 µM ConcA or DMSO. Roots were stained with BCECF
(green) and FM4-64. Bars = 20 µm.
What Do We Learn from a Mutant Lacking Both Vacuolar
H+- Pumps?
Given that all vacuolar functions rely on transport across the tonoplast that is either energized by the proton gradient or the
membrane potential generated by the two tonoplast proton
pumps, it is surprising that the fugu5-1 vha-a2 vha-a3 triple mutant
is viable. The analysis of this important genetic resource allowed
us to draw several important conclusions. First of all, the nearly
neutral cell sap pH in leaves of the triple mutant is most easily
explained by a significant contribution of the V-PPase to vacuolar
acidification that is not revealed in the V-PPase single mutants.
Although mesophyll cells of the triple mutant are substantially
smaller, they develop a large central vacuole, and it remains to be
determined if vacuolar pH in these cells is always high or if an initial
pH gradient is established but then consumed by secondary
active transport processes. Second, our finding that cell expansion in the hypocotyl and in the root is reduced but not abolished
demonstrates that vacuolar functions can be maintained in the
absence of both pumps. In contrast to the situation in leaves,
vacuolar pH in roots was largely unaffected by the additional lack
of the V-PPase in the triple mutant. This indicates either that the
V-PPase only plays a minor role in the root or that a compensatory
mechanism for vacuolar acidification acts in roots but not in
leaves. However, we have also shown here that in the root
elongation zone, vacuole morphology is affected in the triple
mutant but not in the vha-a2 vha-a3 double mutant, indicating
a function of the V-PPase in roots, the nature of which remains to
be identified.
How Can Vacuoles Be Acidic without V-ATPase and
V- PPase?
V-ATPase and V-PPase have long been considered to be the only
tonoplast proton pumps, but it is by now clear that members of the
P3A subgroup of P-type ATPases previously assumed to be
exclusively present at the plasma membrane can also be found at
the tonoplast (Verweij et al., 2008; Faraco et al., 2014). In Arabidopsis, AHA10 is required for the vacuolar accumulation of
proanthocyanidins (Baxter et al., 2005) and has recently been
shown to be localized at the tonoplast of seed coat endothelial
cells (Appelhagen et al., 2015). In agreement with its preferential
expression in the integument layers of developing seeds, no other
phenotypes than the altered seed color have been reported for
aha10 mutants. Although we could detect expression of AHA10 in
roots, it is not upregulated in the triple mutant. Given that vacuolar
pH was not affected by vanadate treatment, compensation by
a P3A-type ATPase is unlikely to be responsible for the vacuolar
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The Plant Cell
acidification observed in the triple mutant. In contrast, inhibition of
the remaining activity of V-ATPase complexes containing VHA-a1
by ConcA in the double and triple mutant led to complete loss of
vacuolar acidification.
Relocalization of Stv1p, the otherwise strictly Golgi-localized
isoform of yeast subunit a, to the vacuole has been reported for
yeast mutants lacking Vph1p (Perzov et al., 2002). A similar relocalization of VHA-a1 to the tonoplast could explain both the
remaining vacuolar acidification and its ConcA sensitivity. Within
the detection limit of the method, we could not observe a shift in
VHA-a1 localization to the tonoplast and thus favor an alternative
hypothesis in which the TGN/EE-localized V-ATPase contributes
directly or indirectly to vacuolar pH. ConcA specifically inhibits
V-ATPase activity; as a consequence, secretory and endocytic
trafficking from the TGN/EE to the vacuole are impaired (Dettmer
et al., 2006; Viotti et al., 2010). Assuming the existence of a so far
unidentified proton pump that is delivered via the TGN/EE to the
tonoplast, the effect of ConcA could be explained by reduced tonoplast levels of this unknown protein. An alternative explanation
would be that vesicular trafficking from the TGN/EE to the vacuole
delivers a substantial amount of protons and/or organic acids to the
vacuolar lumen. The TGN/EE is the most acidic compartment of the
plant endomembrane system (Martinière et al., 2013; Shen et al.,
2013), and using a pH-sensor with suitable resolution at low pH, we
have recently shown that in Arabidopsis roots pH in the TGN/EE is
indeed lower than the vacuolar pH (Luo et al., 2015). At least three
independent, parallel trafficking pathways exist from the TGN/EE to
the vacuole (Pedrazzini et al., 2013; Rojas-Pierce, 2013), and it remains to be determined if the combined flux of protons delivered to
the vacuolar lumen would be sufficient to maintain a proton gradient
even in the absence of tonoplast-localized proton pumps.
METHODS
Plant Materials and Growth Conditions
Arabidopsis thaliana Col-0 ecotype was used in all experiments as control. The
transgenic 35S:AVP1 lines (AVP1-1 and AVP1-2) were previously established
(Gaxiola et al., 2001). The two V-PPase mutant lines fugu5-1 and vhp1-1 were
previously described (Ferjani et al., 2007, 2011). The loss-of-function mutant
avp1-1 (GK_005D04) was obtained from the GABI-KAT collection (www.
gabi-kat.de/). The GNOM mutants gnomR5 and emb30-1 were described
previously (Mayer et al., 1993). The tonoplast V-ATPase double mutant
vha-a2 vha-a3 has been previously characterized (Krebs et al., 2010).
Standard growth medium for plate experiments contained 0.53 Murashige
and Skoog (MS), 0.5% sucrose, 0.5% phyto agar, and 10 mM MES, and the
pH was set to 5.8 using KOH. Agar and MS basal salt mixture were purchased
from Duchefa. Seeds were surface sterilized with ethanol and stratified for 48 h
at 4°C. Plants were grown under LD conditions (16 h light/8 h dark).
Plant material used for in situ hybridization, immunohistochemistry, immunoblotting, qRT-PCR, V-PPase activity assay, cell sap pH, leaf vacuolar
morphology, and fresh weight determination was grown in soil at 22°C under
LD conditions.
For auxin transport assays and free IAA quantification, plants were
grown as previously described (Kubeš et al., 2012). Briefly, plants were
grown on 0.253 MS media, pH 5.7, 0.5% sucrose, and 1% phyto agar for
14 h under 100 µmol m22 s21 light unless otherwise indicated.
Seedlings for root vacuolar pH measurements, vacuolar morphology,
root zone determination, and AHA10 RNA level were grown for 6 d on plates
containing standard growth medium.
Etiolated seedlings were grown on medium containing 10 mM MESKOH, pH 5.8, solidified with 1% phyto agar. Seeds were exposed to light for
4 h and then grown in the dark at 22°C for 4 d.
For root length measurements, single seeds were placed on standard
growth medium solidified with 0.8% phyto agar and then grown vertically
for 10 d.
Seedlings for external media acidification were cultivated for 5 d on
medium containing 0.53 MS, 0.5% sucrose, and 2.5 mM MES-KOH, pH
5.8, solidified with 0.6% phyto agar before they were transferred to a 96well plate containing 0.53 MS and 0.5% sucrose with the pH set to 6.4.
Construct Design and Plant Transformation
To generate transgenic plants expressing AVP1 under the control of the
UBQ10 promoter, the plant binary vector UBQ:AVP1 was generated. First,
a 2331-bp fragment of AVP1 was amplified from cDNA using AVP1-XbaI-Fw
and AVP1-SalI-Rv (Supplemental Table 3). After subcloning into pJET1.2blunt
(Thermo Scientific), the fragment was released using the introduced XbaI and
SalI restriction sites and inserted into the backbone of the binary vector pTKan
(Krebs et al., 2012) to generate pTKan-AVP1. In addition, a 661-bp fragment of
the Arabidopsis UBQ10 promoter was amplified using UBQ10-KpnI-Fw and
UBQ10-KpnI-Rv. After subcloning of UBQ10 into pJET1.2blunt, the fragment
was released via the KpnI site and inserted into pTKan-AVP1 to finally yield
UBQ:AVP1. The UBQ:AVP1 construct was introduced into the Agrobacterium
tumefaciens strain GV3101:pMP90 and selected on 5 mg/mL rifampicin,
10 mg/mL gentamycin, and 100 mg/mL spectinomycin. Arabidopsis
ecotype Col-0 and vha-a2 vha-a3 plants were used for transformation
using standard procedures. Transgenic plants were selected on MS
medium containing 50 µg/mL kanamycin. For creating the template for in situ
hybridization, the cDNA of AVP1 was amplified from the pJET1.2blunt
construct using AVP1-SalI-Fw and AVP1-XbaI-Rv primers and cloned into
pGEM-T Easy.
Genome Sequencing
For genome sequencing, avp1-1 seedlings showing strong shoot and root
abnormalities were collected and used for DNA preparation. DNA was
isolated according to the manufacturer’s instructions using the DNeasy
Plant Mini Kit (Qiagen).
The library was produced at the CellNetworks Deep Sequencing Core
Facility (Heidelberg) and sequenced at the EMBL Genomics Core Facility
(Heidelberg) as 50-bp single-end reads, multiplexed on a HiSeq2000 (Illumina).
Sequences were mapped to the TAIR10 genome using bowtie2 (version 2.0.5)
(Langmead and Salzberg, 2012) with the fast-local option, and 21.7 mio reads
mapped for the avp1 mutant line. The avp1 mutant was additionally mapped
against the GABI-KAT plasmid pAC106 (Kirik et al., 2006; Kleinboelting et al.,
2012). The mappings were converted to bam format, sorted and indexed using
samtools (version 0.1.16) (Li et al., 2009), and displayed on the Integrative
Genomics Viewer (Robinson et al., 2011; Thorvaldsdóttir et al., 2013). Reads
that showed partial mapping to the T-DNA borders were compared with gaps in
the genomic alignment to identify the insertion sites.
RNA Isolation and cDNA Synthesis
For the analysis of AVP1 and AHA10 transcript levels, RNA was isolated
according to the manufacturer’s instructions using the RNeasy Plant Mini
Kit (Qiagen). cDNA was synthesized from 2 µg of total RNA using M-MuLV
reverse transcriptase (Thermo) and an oligo(dT) primer.
Real-Time RT-PCR
Real-time PCR reactions were performed using the DNA Engine Opticon
System (DNA Engine cycler and Chromo4 detector; Bio-Rad) and Absolute
qPCR SYBR Green Mix (Thermo Scientific). The real-time PCR reaction
Job Sharing between V-ATPase and V-PPase
mixture with a final volume of 20 mL contained 0.5 µM of each forward and
reverse primer, 10 mL of SYBR Green Mix, 4 mL of cDNA, and 4 mL of
RNase-free water. The thermal cycling conditions were composed of an
initial denaturation step at 95°C for 15 min followed by 40 cycles at 95°C for
15 s, 59°C for 30 s, and 72°C for 15 s and ended with a melting curve. For the
analysis of each sample, three analytical replicas were used. Target genes
were normalized to the expression of Actin2. Primer sequences for AVP1,
AHA10, and Actin2 are listed in Supplemental Table 3.
11 of 14
analyze whether avp1-1 mutants also carry a T-DNA insertion in GNOM,
a T-DNA right border primer (GN.T-DNA) and a GNOM-specific reverse
primer (GN.REV) were used. The GNOM wild-type allele was identified
using GNOM forward and reverse primer (GN.For and GN.Rev; Supplemental
Table 3).
Amplification of VHA-a3 using VHA-a3-specific forward (S029786.for)
and reverse (S029786.rev) primer served as a control.
Pharmacological Treatments and Stains
In Situ Hybridization
In situ hybridizations were performed as previously described (Medzihradszky et al., 2014). For AVP1 probe preparations, AVP1-pGEM-T Easy
(see constructs) was used. The template for the transcription was PCR
amplified with T7 and SP6 primers from these vectors.
Immunohistochemistry on Tissue Sections
Meristems were fixed and embedded in Steedman’s wax according to
a previously described protocol (Paciorek et al., 2006) and sectioned at
8 mm by a microtome (Leica CM3050S). Sections were mounted on slides
(X-Tra; Leica) and dried overnight at room temperature. Antibody binding
was performed as described (Paciorek et al., 2006), with the anti-AVP1
antiserum (Cosmo Bio Co.; 1:10,000 in 2% BSA-PBS) covered with Parafilm
at room temperature for 4 h and an AP-conjugated secondary antibody [goat
anti-rabbit IgG (H+L)-alkaline phosphatase; Dianova 111-055-045; 1:2500 in
2% BSA-PBS] for 1 h. After washing the sections for 90 min in microtubulestabilizing buffer (50 mM PIPES-KOH, pH 6.8, 6.5 mM EDTA, and 5 mM
MgSO4) and equilibrating in the substrate buffer, staining was developed with
NBT-BCIP solution (Roche) at room temperature for 1 h.
Arabidopsis seedlings were incubated in liquid 0.53 MS medium with
0.5% sucrose, pH 5.8, containing 1 µM ConcA, 1 µM FM4-64, 10 µM
BCECF, or the equivalent amount of DMSO in control samples for the
indicated time at room temperature. Stock solutions were prepared in
DMSO. Sodium orthovanadate (Sigma-Aldrich) was applied in a concentration of 2 mM and liquid 0.53 MS without any supplement served as the
mock control.
Confocal Microscopy
Images were recorded using a Leica TCS SP5II microscope equipped with
a Leica HCX PL APO lambda blue 63.0 3 1.20 UV water immersion objective. GFP as well as BCECF was excited at 488 nm using a VIS-argon
laser, and FM4-64 was excited at 561 nm with a VIS-DPSS 561 laser diode.
Fluorescence emission of GFP and BCECF was detected between 500 and
555 nm, and the emission of FM4-64 was detected between 615 and 676
nm. The Leica Application Suite Advanced Fluorescence software was
used for image acquisition. Post processing and measurements of images
were performed using Fiji (based on ImageJ 1.50a).
SDS-PAGE and Immunoblotting
pH Measurements
Microsomal membrane proteins were analyzed by SDS-PAGE and subsequent immunoblotting. Upon gel electrophoresis, the proteins were
transferred to a nitrocellulose membrane (Whatman). The primary antibody
against the V-PPase was purchased from Cosmo Bio (for details, see
immunohistochemistry section); the primary antibody against VHA-C
(Schumacher et al., 1999) and anti-TIP1;1 were previously described (Jauh
et al., 1998). Antigen on the membrane was visualized with horseradish
peroxidase-coupled anti-rabbit IgG (Promega) and chemiluminescent
substrate (Peqlab). Immunostained bands were analyzed using a cooled
CCD camera system (Intas).
Vacuolar pH measurements were performed as previously described
(Viotti et al., 2013) with minor modifications. BCECF fluorescence was
detected using a Leica SP5II confocal laser scanning microscope
equipped with a HCX PL APO CS 20.0 3 0.70 IMM UV water immersion
objective. All images were exclusively recorded within the root differentiation zone except when root vacuolar pH was determined
additionally in the root meristematic and elongation zone (Figure 6). In
situ calibration was performed separately for each root zone. Cell sap
pH measurements were conducted as previously described (Krebs
et al., 2010).
Microsomal Membrane Preparation and Enzyme Assay
Extracellular Acidification Assay
Extraction of microsomal membranes and V-PPase enzyme activity
measurements were performed as described (Krebs et al., 2010).
Media acidification was measured according to Haruta et al. (2010) with
minor modifications. Five-day-old single seedlings were transferred to
a 96-well plate containing 200 mL of 0.53 MS and 0.5% sucrose, pH 6.4,
supplemented with 30 µg/mL fluorescein dextran (molecular weight =
10.000; Life Technologies) and 2 mM vanadate. Before and after an incubation time of 18 h, fluorescence emission was detected at 521 nm after
excitation at 494 and 435 nm using a microplate reader (Tecan Infinite
M1000). The media pH was calculated using a calibration curve ranging
from pH 3.5 to 6.5.
Auxin Transport Assays and Free IAA Determinations
Auxin transport assays and free IAA determinations were conducted as
described (Kubeš et al., 2012). Statistics were performed with SigmaStat,
using one-way ANOVA followed by Dunnett’s post-hoc analyses set to P <
0.05. Data shown are means 6 SD of two or three independent replicates, as
indicated in the figure legends.
Accession Numbers
Genotyping
Genomic DNA was extracted as described before (Edwards et al., 1991).
To check for the presence of the T-DNA in the AVP1 gene, PCR was
performed using a T-DNA left border-specific primer (LB-JL202) and an
AVP1-specific reverse primer (AVP1.Rev). To identify the AVP1 wild-type
allele, two gene-specific primers were used (AVP1.For and AVP1.Rev). To
Sequence data from this article can be found in the Arabidopsis Genome
Initiative or GenBank/EMBL databases under the following accession
numbers: AVP1/VHP1 (At1g15690), GNOM (At1g13980), VHA-a1
(At2g28520), VHA-a2 (At2g21410), VHA-a3 (At4g39080), VHA-C
(At1g12840), AHA10 (At1g17260), TIP1;1 (At2g36830), CNX (At5g61790),
and ACT2 (At3g18780).
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The Plant Cell
Supplemental Data
Supplemental Figure 1. AVP1 transgene is silenced in 35S:AVP1
lines.
Supplemental Figure 2. T-DNA insertions in AVP1 and GNOM.
Supplemental Figure 3. Phenotypes of the T-DNA insertion line
vhp1-1 and the point mutation allele fugu5-1.
Supplemental Figure 4. Cell expansion in fugu5-1 vha-a2 vha-a3 is
independent of the tonoplast V-ATPase and V-PPase.
Supplemental Figure 5. Transcript level of tonoplast P-type ATPase
AHA10.
Supplemental Table 1. Rootward and shootward auxin transport
Supplemental Table 2. Free IAA content
Supplemental Table 3. Primer list.
ACKNOWLEDGMENTS
We thank B. Schöfer and K. Piiper for excellent technical assistance and
R. Gaxiola, A. Ferjani, and G. Jürgens for providing published materials.
This work was supported by the Deutsche Forschungsgemeinschaft within
FOR1061 to K.S. and the ERC by a starting grant to J.U.L. Funding to
W.A.P. was provided by the Maryland Agricultural Research Station.
AUTHOR CONTRIBUTIONS
A.K., Z.A., A.M., J.U.L., and K.S. designed experiments. A.K., Z.A., A.M.,
F.K., S.S., S.D., and M.G.P.-N. performed experiments and analyzed data.
M.K. established UBQ:AVP1 lines. A.P. analyzed sequencing data. G.G.,
H.Y., A.S.M., and W.A.P. contributed with auxin content and transport data.
A.K. and K.S. wrote the article.
Received August 17, 2015; revised October 21, 2015; accepted November
5, 2015; published November 20, 2015.
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Job Sharing in the Endomembrane System: Vacuolar Acidification Requires the Combined
Activity of V-ATPase and V-PPase
Anne Kriegel, Zaida Andrés, Anna Medzihradszky, Falco Krüger, Stefan Scholl, Simon Delang, M.
Görkem Patir-Nebioglu, Gezahegn Gute, Haibing Yang, Angus S. Murphy, Wendy Ann Peer, Anne
Pfeiffer, Melanie Krebs, Jan U. Lohmann and Karin Schumacher
Plant Cell; originally published online November 20, 2015;
DOI 10.1105/tpc.15.00733
This information is current as of June 18, 2017
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