Heat Reduces Nitric Oxide Production Required for
Auxin-Mediated Gene Expression and Fate Determination
in Tree Tobacco Guard Cell Protoplasts1[OA]
Robert A. Beard, David J. Anderson 2, Jennifer L. Bufford 3, and Gary Tallman*
Department of Biology, Willamette University, Salem, Oregon 97301
Tree tobacco (Nicotiana glauca) is an equatorial perennial with a high basal thermotolerance. Cultured tree tobacco guard cell
protoplasts (GCPs) are useful for studying the effects of heat stress on fate-determining hormonal signaling. At lower
temperatures (32°C or less), exogenous auxin (1-naphthalene acetic acid) and cytokinin (6-benzylaminopurine) cause GCPs to
expand 20- to 30-fold, regenerate cell walls, dedifferentiate, reenter the cell cycle, and divide. At higher temperatures (34°C or
greater), GCPs expand only 5- to 6-fold; they do not regenerate walls, dedifferentiate, reenter the cell cycle, or divide. Heat (38°C)
suppresses activation of the BA auxin-responsive transgene promoter in tree tobacco GCPs, suggesting that inhibition of cell
expansion and cell cycle reentry at high temperatures is due to suppressed auxin signaling. Nitric oxide (NO) has been
implicated in auxin signaling in other plant systems. Here, we show that heat inhibits NO accumulation by GCPs and that
G
L-N -monomethyl arginine, an inhibitor of NO production in animals and plants, mimics the effects of heat by limiting cell
expansion and preventing cell wall regeneration; inhibiting cell cycle reentry, dedifferentiation, and cell division; and
suppressing activation of the BA auxin-responsive promoter. We also show that heat and L-NG-monomethyl arginine reduce
the mitotic indices of primary root meristems and inhibit lateral root elongation similarly. These data link reduced NO levels to
suppressed auxin signaling in heat-stressed cells and seedlings of thermotolerant plants and suggest that even plants that have
evolved to withstand sustained high temperatures may still be negatively impacted by heat stress.
The three major interrelated abiotic plant stresses
accompanying global climate change are increasing
concentrations of atmospheric CO2, heat, and drought
(Mittler, 2006). Heat stress (Mittler, 2006; Mittler and
Blumwald, 2010) is known to damage plant tissues
(Pareek et al., 1997), interfere with plant reproductive
development (Warrington, 1983; Francis and Barlow,
1988; Commuri and Jones, 2001; Matsui and Omasa,
2002; Sato et al., 2006; Barnabás et al., 2008; Snider
et al., 2011), and cause the redistribution of native
plant populations (Marchand et al., 2006; Walker et al.,
2006; Darbah et al., 2010; Offord, 2011). Significant
consequences of heat stress include reductions in crop
yields, lower biomass production (Mittler, 2006;
Qaderi et al., 2006; Sato et al., 2006; White et al., 2006;
Barnabás et al., 2008; Mittler and Blumwald, 2010), and
1
This work was supported by the M.J. Murdock Charitable Trust
and the National Science Foundation (grant no. MCB–1021093).
2
Present address: Deininger Research Group, University of Utah,
Huntsman Cancer Institute, 2000 Circle of Hope Drive, Salt Lake City,
UT 84112.
3
Present address: Department of Botany, University of Hawaii at
Manoa, 3190 Maile Way, Room 101, Honolulu, HI 96822.
* Corresponding author; e-mail [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.plantphysiol.org) is:
Gary Tallman ([email protected]).
[OA]
Open Access articles can be viewed online without a subscription.
www.plantphysiol.org/cgi/doi/10.1104/pp.112.200089
1608
alterations to allelic frequencies in plant populations
(Jump et al., 2006).
Even though a host of heat’s negative impacts
on plant growth and reproduction have been documented, little is known about the cellular and molecular mechanisms that underlie these effects. Nor do
we know how the more frequent and sustained periods of heat stress expected from climate change
might alter these mechanisms to modify plant physiological performance. Even if crop plants can be genetically modified to be more thermotolerant, we do not
know whether they will remain fecund. Neither can we
yet predict which native plants might succumb to the
effects of sustained heat.
Many plant heat stress studies have focused on plant
thermotolerance (Penfield, 2008). There are two types:
basal and acquired (Larkindale et al., 2005; Yoo et al.,
2006; Penfield, 2008; Clarke et al., 2009). Basal thermotolerance (BT) is that inherent to each plant species.
Acquired thermotolerance (AT) is that which can be
induced by exposing plants to relatively short bouts of
less severe heat (a heat shock) to activate a program
that protects the plant transiently against a subsequent
short bout of more severe heat. BT has received less
attention than AT, and the molecular basis for BT is
not well understood. In Arabidopsis (Arabidopsis
thaliana), salicylic acid (SA) and jasmonic acid derivatives can activate BT, but SA is not always required for
BT (Clarke et al., 2009). Mutating SIZ1, a unique
Arabidopsis ubiquitin-like modifier gene required to
sumoylate specific heat shock transcription factors,
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Heat Reduces Nitric Oxide to Determine Guard Cell Fate
reduces BT independent of SA signaling (Yoo et al.,
2006). HSP101 has been implicated in Arabidopsis BT
(Queitsch et al., 2000), but it appears to be more important for AT (Queitsch et al., 2000). AT has been
studied extensively and is thought to be mediated, in
large part, through activation of heat shock promoters
(Li et al., 2010), regulation of heat shock transcription
factors (Cohen-Peer et al., 2010; Scharf et al., 2012),
and/or accumulation of certain heat shock proteins
such as HSP101 (Queitsch et al., 2000).
Beyond thermotolerance, our knowledge of plant
heat stress is inadequate, perhaps because even after
heat shock many crop plants and model plants still
succumb to extended bouts of high heat. Because such
plants comprise the bulk of those examined, virtually
nothing is known about how heat might affect hormone-directed plant growth and development long
term if these plants were able to survive. Nor do we
know how heat-induced alterations to hormonal signaling for plant growth might affect day-to-day plant
physiological processes, such as stomatal opening or
closing, by modulating components of other hormonal
signaling cascades like that/those for abscisic acid
(ABA). Therefore, there is a pressing need for good
experimental models in which plants survive high heat
long enough to allow the effects of extended heat stress
on hormone function, plant growth and development,
and a wide variety of other plant physiological processes to be studied.
Tree tobacco (Nicotiana glauca) is an equatorial perennial with a high BT (Zuloaga and Morrone, 1996). In
climate change studies, little attention has been given to
plants like tree tobacco that have evolved to survive
sustained bouts of high heat. While it is often assumed
that increases in mean global temperatures will have
little effect on these hardy plants, our previous studies
with cultured tree tobacco guard cells (Roberts et al.,
1995; Taylor et al., 1998; Gushwa et al., 2003; Dong
et al., 2007) suggest that heat can disrupt growth hormone signaling in ways that drastically alter cell fate.
Cultured tree tobacco guard cell protoplasts (GCPs)
manifest the inherent, cell-autonomous BT of the parent plant by surviving for several weeks at temperatures as high as 40°C (Roberts et al., 1995). Tree
tobacco GCPs are a useful monoculture system for
studying how sustained exposure to heat regulates the
hormonal signaling mechanisms governing cell survival, cell expansion, cell wall formation, cell cycle
reentry, and cellular dedifferentiation (Tallman, 2005).
In medium containing an auxin (1-naphthalene acetic
acid [NAA]) and a cytokinin (6-benzylaminopurine
[BAP]), GCP survival increases dramatically as culture
temperatures are increased from 20°C to 32°C (Roberts
et al., 1995). After 1 week in culture, survival at 20°C is
only 20% to 30% while cell survival at temperatures
between 32°C and 40°C is 70% to 80% (Roberts et al.,
1995; Gushwa et al., 2003). At temperatures of 32°C or
less, NAA and BAP are both required for survival; at
temperatures greater than 34°C, neither is required for
survival (Gushwa et al., 2003).
In addition to increasing survival, heat has dramatic
effects on the fate of cultured tree tobacco GCPs. At a
lower culture temperature of 32°C, GCPs regenerate
cell walls (Taylor et al., 1998), expand as much as 20- to
30-fold in “footprint” area (Roberts et al., 1995), reenter
the cell cycle (Gushwa et al., 2003), dedifferentiate
(Taylor et al., 1998), and divide to form embryogenic
callus tissue from which plants can be regenerated
(Sahgal et al., 1994). At a higher culture temperature of
38°C, GCPs expand only 5- to 6-fold (Roberts et al.,
1995). They do not regenerate cell walls (Taylor et al.,
1998), reenter the cell cycle (Gushwa et al., 2003), dedifferentiate (Taylor et al., 1998), or divide (Roberts
et al., 1995). Culturing tree tobacco GCPs for 16 h or
more at 38°C suppresses activation of the auxin-responsive BA transgene promoter (Aspuria et al., 2002;
Dong et al., 2007); BA activation is not suppressed
when GCPs are cultured similarly at 32°C. We hypothesize that sustained heat interferes with the auxin
signaling required for cell expansion, cell wall regeneration, cell cycle reentry, and cellular dedifferentiation processes that are prerequisites for cell cycle
progression.
Nitric oxide (NO) is an important cell signaling gas
in animals (Cáceres et al., 2011) and plants (Hayat
et al., 2009). In alfalfa (Medicago sativa) mesophyll
protoplasts, NO is required for, and promotes, auxinmediated S-phase cyclin-dependent kinase (CdK)
accumulation and activation, cell cycle reentry, and
embryogenic cell formation (Ötvös et al., 2005). NO
also modulates the auxin-mediated expression of
S-phase CdK genes in developing lateral roots of tomato (Solanum lycopersicum; Correa-Aragunde et al.,
2006). In this study, we examined whether heat might
interfere with auxin signaling in cultured tree tobacco
GCPs by blocking NO production and/or accumulation required for auxin-responsive gene expression
and auxin-mediated cell wall formation, cell cycle reentry, and dedifferentiation. We show that heat reduces NO accumulation by tree tobacco GCPs and that
a known inhibitor of plant NO production, L-NGmonomethyl arginine (L-NMMA; Mur et al., 2005),
mimics nearly all of the effects of sustained heat (culture at 38°C) on these cells. We also show that in tree
tobacco seedlings, heat and L-NMMA reduce the mitotic indices of root apical meristems and inhibit lateral
root elongation to similar extents.
RESULTS
Heat Reduces NO Accumulation by GCPs
We used the NO-sensitive fluorescent dye diaminofluorescein-2 diacetate (DAF-2DA; Kojima et al.,
1998) to determine whether heat might inhibit NO
accumulation by cultured tree tobacco GCPs; NO results in the conversion of DAF-2DA to the fluorescent
derivative, triazolofluorescein (DAF-2T). After 16 h at
32°C in medium containing NAA and BAP, 35.3% 6
2.9% of GCPs exhibited DAF-2T fluorescence, a value
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Beard et al.
not significantly different from the 39.3% 6 3.1% of cells
that exhibited DAF-2T fluorescence in medium containing the NO generator S-nitroso-N-acetyl-DL-penicillamine
(SNAP; Figs. 1, A and D, and 2). The majority of fluorescence was localized to chloroplasts, but about 15%
of total cells showed cytoplasmic fluorescence, most
of it concentrated around nuclei (perinuclear; Figs.
1A and 2). Only 1.1% 6 2.9% of cells showed both
chloroplastic and perinuclear fluorescence (Fig. 2).
L-NMMA and 2-phenyl-4,4,5,5,-tetramethylimidazoline-1-oxyl-3-oxide (PTIO) each reduced significantly
the percentages of cells fluorescing to 7.4% 6 2.1% and
7.3% 6 1.7%, respectively (Figs. 1, B and C, and 2). In
medium with L-NMMA or PTIO, DAF-2T fluorescence
was restricted to chloroplasts (Fig. 2).
After 16 h at 38°C in medium containing NAA and
BAP, 10.0% 6 3.6% of GCPs exhibited DAF-2T fluorescence (Fig. 1E), significantly lower than the 35.3% 6
2.9% of cells that exhibited DAF-2T fluorescence at 32°
C (Figs. 1A and 2) or the 38.6% 6 9.2% that showed
DAF-2T fluorescence at 38°C in medium containing
SNAP (Figs. 1H and 2). The mean percentage of cells
that exhibited DAF-2T fluorescence after 16 h at 38°C
was not significantly different from those of L-NMMA
and PTIO controls at 38°C (Figs. 1, F and G, and 2),
and fluorescence was confined to chloroplasts in heat-,
L-NMMA-, or PTIO-treated cells (Fig. 2). Only SNAP
controls showed perinuclear fluorescence at 38°C
(Figs. 1H and 2).
L-NMMA
Prevents GCP Cell Wall Regeneration
GCPs cultured for 1 week at 32°C in medium with
NAA and BAP regenerated cell walls (Fig. 3A); those
cultured at 32°C in medium with NAA, BAP, and
L-NMMA did not (Fig. 3B).
Figure 1. Effect of heat on NO production by cultured tree tobacco
GCPs. GCPs were cultured for 16 h at 32˚C or at 38˚C and then stained
with the NO-detecting fluorescent dye DAF-2DA (5 mM). The presence
of NO results in the production of a fluorescent product, DAF-2T
(green). Red indicates chlorophyll autofluorescence. A, At 32˚C, GCPs
produced NO. B and C, In parallel cultures, NO production at 32˚C
was reduced by L-NMMA and by the NO scavenger PTIO. D and H,
The NO generator SNAP produced dye fluorescence in GCPs at 32˚C
and at 38˚C. E, Culture at 38˚C reduced NO to levels similar to those
observed at 32˚C in medium with L-NMMA (B) or PTIO (C). F and G,
Neither L-NMMA nor PTIO further reduced NO levels in cells cultured
at 38˚C. All magnifications are 2003.
Figure 2. Effects of heat on NO accumulation in cultured tree tobacco
GCPs. Percentages are shown for protoplasts fluorescing at various
cellular locations after treatment with the NO-reporting dye DAF-2DA
following temperature pretreatment for 16 h at 32˚C or 38˚C in medium alone (control) or in medium containing L-NMMA, PTIO, or
SNAP. Values are means 6 SE. n = 4 except for SNAP at 32˚C, where
n = 3. “Both” indicates cells with both chloroplast and perinuclear
fluorescence. A, Significantly different from the corresponding mean at
32˚C; B, significantly different from the corresponding control mean at
the same temperature (ANOVA; Fisher PLSD; P # 0.05)
Effects of L-NMMA on Hormone-Mediated Cell
Expansion, Cell Morphology, and Survival
After 1 week at 32°C in medium with NAA, BAP,
and L-NMMA, mean cell footprint area (Roberts et al.,
1995; hereafter, “size”) was 16.6 6 0.9 3 1022 mm (Fig.
4C). This value was approximately six times that of
freshly isolated GCPs (Fig. 4A) but was significantly
lower than the 34.9 6 2.7 3 1022 mm mean area of cells
cultured at 32°C in medium with NAA and BAP alone
(Fig. 4B; ANOVA; partial least square difference
[PLSD]; P # 0.0001). It was not, however, significantly
different from those of cells cultured at 38°C in medium with NAA and BAP (Fig. 4D; 18.5 6 1.2 3 1022
mm; ANOVA; Fisher PLSD; P = 0.40) or at 38°C in
medium with NAA, BAP, and L-NMMA (13.1 6 0.5 3
1022 mm; ANOVA; Fisher PLSD; P = 0.11).
The morphologies of GCPs cultured for 1 week at
32°C in medium with NAA, BAP, and L-NMMA were
very similar to those of GCPs cultured for 1 week at
38°C in medium with NAA and BAP (Fig. 4, C and D).
After 1 week of culture at 32°C in medium with
NAA, BAP, and 1 mM L-NMMA, mean cell survival
was 47.2% 6 1.3%, significantly lower than the
66.4% 6 0.4% observed for the same treatment at 38°C
(ANOVA; Fisher PLSD; P # 0.004). Survival at 32°C in
medium with 1 mM L-NMMA and BAP, but lacking
NAA, was 32.6% 6 3.7%, significantly lower than with
NAA, BAP, and L-NMMA at 32°C (ANOVA; Fisher
PLSD; P # 0.02). Survival at 32°C in medium with
1 mM L-NMMA and NAA, but lacking BAP, was
40.3% 6 6.6%; survival at 32°C in medium with 1 mM
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Heat Reduces Nitric Oxide to Determine Guard Cell Fate
Figure 3. Effects of L-NMMA on cell wall regeneration by cultured tree
tobacco GCPs. After 1 week in culture at 32˚C in medium without
1 mM L-NMMA (A) or with 1 mM L-NMMA (B), cells were stained with
0.05% Calcofluor White and examined microscopically at 4003 under blue epifluorescence illumination.
but neither NAA nor BAP, was 38.3% 6 2.9%.
Neither survival percentage was statistically different
from that of 32°C cultures containing NAA, BAP, and
L-NMMA (ANOVA; Fisher PLSD; P = 0.21 and 0.11,
respectively).
L-NMMA,
L-NMMA
Blocks Hormone-Mediated Cell Cycle Reentry
In tree tobacco GCP cultures, S-phase typically begins
within 72 to 96 h after cultures are established (Gushwa
et al., 2003). In the experiments reported here, after
5-bromo-deoxyuridine (BrdU) pulse labeling at 32°C for
24 h beginning 72 h after cultures were established, an
average of 27.9% 6 8.6% (mean 6 SE; n = 3) nuclei from
cells cultured in medium with NAA and BAP exhibited
fluorescein isothiocyanate-anti-BrdU fluorescence (Fig.
5, A and B). In parallel experiments with cells from the
same isolates, no labeling was detected in cells cultured
at 38°C (Fig. 5C) or at 32°C in medium containing 1 mM
L-NMMA (Fig. 5, D and E).
Suppresses Activation of the BA
Auxin-Responsive Promoter at the Normally
Permissive Temperature of 32°C
transformants cultured at 32°C (Fig. 6D; Table I) or at
38°C (Fig. 6, D and F; Table I). L-NMMA (1 mM) significantly reduced the mean percentage of BA-mgfp5ER transformants that accumulated mGFP5-ER at 32°C
to 5.9% 6 2.4% (Fig. 6B; Table I), a mean percentage
similar to that of cells cultured at 32°C in medium
without NAA (Table I) or at 38°C (Fig. 6C; Table I).
Under the same conditions, L-NMMA did not reduce
the mean percentage of 35S-mgfp5-ER transformants
that accumulated mGFP5-ER at 32°C (Table I). With or
without L-NMMA, culturing BA-mgfp5-ER transformants at 38°C restricted mGFP5-ER accumulation to less
than 1% of cells, but the same high percentage (approximately 50%) of 35S-mgfp5-ER transformants accumulated mGFP5-ER at 38°C as at 32°C.
Our previous studies indicated that 12 to 16 h of
culture at 38°C were required before NAA-mediated
BA activation was suppressed in all GCPs examined
(Dong et al., 2007). While L-NMMA would be expected
to act almost immediately, it could be argued that in
heat experiments, long-lived transcripts produced
early in the 38°C culture period could be responsible
for the mGFP5-ER accumulation in 35S controls, masking
inhibitory effects of heat on generalized transcription
and translation processes. To address this possibility, as
additional controls we preincubated GCPs for 16 h at
38°C, transformed them with the BA- or 35S-mgfp5-ER
construct, and then administered hormonal treatments.
Similar high percentages of 35S-mgfp5-ER transformants accumulated mGFP5-ER regardless of whether
cells were transformed initially (Fig. 6D; Table I) or
after 16 h of culture at 38°C (Fig. 6F; Table I). In a
single control experiment, pretransformation and
then posttransformation incubation at 32°C did not
inhibit auxin-induced mGFP5-ER accumulation; 42%
of GCPs accumulated mGFP5-ER in the 21-h posttransformation incubation in medium with NAA. Only
0.3% of cells transformed with BA-mgfp5-ER accumulated mGFP5-ER when a 16-h pretransformation incubation at 32°C was followed by an additional 21-h
posttransformation incubation at 38°C in medium with
NAA (Fig. 6E).
L-NMMA
Acts Stereospecifically to Inhibit Cell Division
Animal NO synthase enzymes catalyze NO formation
from L-Arg (Besson-Bard et al., 2008) but not from
G
D-Arg; L-NMMA inhibits these enzymes but D-N monomethyl arginine (D-NMMA) does not (Mur et al.,
2005). D-NMMA (1 mM) did not prevent GCPs from dividing at 32°C in medium with NAA and BAP (Fig. 4E).
L-NMMA
At 32°C in medium with NAA and BAP, 48.8% 6
4.2% of GCPs transformed with the transgene reporter
BA-mgfp5-ER accumulated mGFP5-ER (Fig. 6A; Table
I), a mean percentage similar to that of 35S-mgfp5-ER
L-NMMA-Mediated
Inhibition of Cell Division Is Partially
Reversed by 1 mM L-Arg at 32°C, But Arg Does Not Reverse
Heat-Mediated Inhibition of Cell Division
We examined whether L-Arg could reverse the inhibition of cell division by L-NNMA at 32°C. After
1 week of culture at 32°C in medium with NAA and
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Figure 4. Effects of culture temperature and L-NMMA, D-NMMA, or L-NMMA and L-Arg in combination on the expansion and
division of tree tobacco GCPs after 1 week in culture. GCPs were cultured in medium containing both an auxin (NAA) and a
cytokinin (BAP). A, Freshly isolated GCPs. B, GCPs dividing at 32˚C. C and D, Limited expansion and similar morphologies of
GCPs cultured at 32˚C in medium with 1 mM L-NMMA (C) and those cultured at 38˚C (D). E, Unlike L-NMMA, D-NMMA does
not prevent cell division at 32˚C. F, At 32˚C, 5 mM L-Arg rescues some GCPs from L-NMMA-mediated inhibition of cell division.
Magnification is 2003 in B and 4003 in all other panels. Bar = 40 mm for A and 25 mm for B to F.
BAP, approximately 40% of living GCP were dividing
(Fig. 7); 1 mM L-NMMA reduced cell division to less
than 0.5% (Fig. 7). In medium lacking L-NMMA, concentrations of L-Arg in the range of 0.25 to 5 mM reduced
division to approximately 32% to 36%; 20 mM L-Arg
reduced division to 1% to 2% (Fig. 7).
At a lower concentration (0.25 mM), L-Arg did not
reverse L-NMMA-mediated inhibition of cell division,
but 1 mM L-Arg prevented L-NMMA from inhibiting
cell division in approximately 50% of cells based on the
corresponding control without L-NMMA (Figs. 4F and
7). With or without L-NMMA, L-Arg concentrations of
1 mM or greater reduced cell size substantially and in
inverse proportion to L-Arg concentration (data not
shown), which probably prevented GCPs from reentering the cell cycle.
When cells cultured at 38°C were exposed to the
same concentrations of L-Arg, none of the concentrations tested, up to and including 20 mM, reversed the
inhibition of cell division by heat (data not shown).
Sustained Heat Arrests the Growth of Tree
Tobacco Seedlings
When 10- to 14-d-old seedlings grown at 26°C were
transferred to 38°C, their growth was arrested but
seedlings survived (Fig. 8A). Heat significantly reduced
petiole length, leaf area, and primary root length (Table
II). Heat also prevented lateral root elongation (Fig. 8B)
and root hair formation (data not shown), although it
did not prevent the development of lateral root primordia (Table II). Growth at 38°C reduced root mitotic
index significantly over those of 2- and 4-week-old
seedlings maintained at 26°C as controls (Table II).
Seedlings transferred to 32°C grew similarly to those
maintained at 26°C but appeared to produce fewer
lateral roots and somewhat larger shoots (Fig. 8).
L-NMMA
Reduces Shoot Growth and Prevents Lateral
Root Development
Transferring 10- to 14-d-old seedlings grown at 26°C
to 32°C and treating them with 1 mM L-NMMA reduced petiole length and leaf expansion significantly
over those of plants grown similarly at 32°C without
L-NMMA (Table II) and those of 26°C and 32°C sham
delivery controls (Table II). L-NMMA prevented the
formation of lateral root primordia and lateral root
elongation (Table II; Fig. 8B). Interestingly, while the
treatment medium in which roots were growing contained 1 mM L-NMMA, the only significant reduction
in number of lateral root primordia occurred when
L-NMMA was applied to the shoot meristem in an agar
block. L-NMMA did not prevent root hair formation
(data not shown), and beyond its effects on lateral
roots, no other aspects of plant growth, including
primary root growth, were reduced as drastically by
L-NMMA as by heat (Table II). At 32°C, L-NMMA reduced root mitotic index significantly over that of 2and 4-week-old controls maintained at 26°C (Table II).
DISCUSSION
Table III contains a comparative summary of the
effects of heat and L-NMMA on cultured GCP and
seedlings referred to in this study and our previous
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Heat Reduces Nitric Oxide to Determine Guard Cell Fate
Figure 5. Effects of heat or L-NMMA on cell cycle reentry in cultured tree tobacco GCPs. The G1-to-S cell cycle transition
typically begins 72 h after cultures are established. A, Nuclei from cells cultured for 96 h at 32˚C stained with Hoechst 33342.
B, The same nuclei (see arrows) shown in A but stained with a fluorescein isothiocyanate-conjugated rabbit anti-BrdU antibody
after 24 h of pulse labeling with BrdU in the 24-h period 72 to 96 h after cultures were established. C, Parallel BrdU-treated
culture similar to B but at 38˚C. D, Hoechst staining of nuclei in parallel culture at 32˚C in medium containing 1 mM L-NMMA.
E, The same as D (see arrows) except stained with anti-BrdU antibody. F, Hoechst in cells at 38˚C in medium with L-NMMA.
Results are typical of those from three separate experiments. Magnification is 2003 for A to C and 4003 for D to F.
studies. Taken together, these data point to NO as a
central determinant of auxin-mediated cell fate in tree
tobacco GCPs cultured at lower temperatures and indicate that sustained heat can alter cell fate by reducing NO levels (Figs. 1 and 2). At low to moderate
culture temperatures (32°C or less), reducing GCP NO
production/accumulation with L-NMMA mimics the
inhibitory effects of heat (38°C) on hormone-directed
cell expansion (Fig. 4, C and D), cell wall regeneration
(Fig. 3B), dedifferentiation (Fig. 4C), cell cycle reentry
(Fig. 5, D and E), and cell division (Fig. 4, B–D). At 32°
C in medium lacking NAA, BAP, or both, adding
L-NMMA to the medium increased cell survival 10- to
110-fold over survival reported previously for GCPs
cultured without NAA and/or BAP (Table III;
Gushwa et al., 2003). Without NAA and/or BAP,
survival at 32°C in medium with L-NMMA was approximately one-half to two-thirds of the hormone-
Figure 6. Effects of heat or 1 mM L-NMMA on activation of the auxin-responsive BA transgene promoter from pea (Pisum
sativum) in cultured tree tobacco GCPs. GCPs were transformed with either BA-mgfp5-ER or with 35S-mgfp5-ER in which the
constitutive 35S cauliflower mosaic virus promoter drives the synthesis of mGFP5-ER. Green indicates mGFP5-ER fluorescence,
and red indicates chlorophyll autofluorescence. GCPs were cultured in medium with an auxin (NAA) and a cytokinin (BAP). A,
mGFP5-ER accumulation after 21 h at 32˚C. B, Parallel 32˚C culture to A but in medium with 1 mM L-NMMA. C, Parallel culture
to A but at 38˚C. D, Field representative of parallel 35S constitutive control cultures at 32˚C or 38˚C in medium with or without
L-NMMA. E, Preincubation at 32˚C for 16 h before transformation with BA-mgfp5-ER followed by transformation and then
incubation at 38˚C for an additional 21 h. F, Control similar to E, but preincubated at 38˚C for 16 h followed by transformation
with 35S-mgfp5-ER and then incubation for another 21 h at 38˚C. All magnifications are 4003.
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Table I. Like heat, L-NMMA inhibits auxin-mediated accumulation of thermostable mGFP5-ER in reporter-transformed tree tobacco GCPs
Protoplasts were transformed with BA-mgfp5-ER, in which mgfp5 expression is regulated by the auxin-responsive BA promoter from pea, or with
35S-mgfp5-ER, in which mgfp5 expression is regulated by the constitutive 35S cauliflower mosaic virus promoter. Values are means 6 SE from three
separate experiments with cells from parallel cultures from the same protoplast isolates. *Significantly different from 32˚C +NAA, +BAP control;
**significantly different from the corresponding 35S (ANOVA; Fisher PLSD; P # 0.05). ND, Not determined.
Treatment
35S-mgfp5-ER
BA-mgfp5-ER
%
24
24
24
24
24
16
h
h
h
h
h
h
at
at
at
at
at
at
32˚C
32˚C
32˚C
38˚C
38˚C
38˚C
+ NAA + BAP (control)
+ NAA + BAP + 1 mM L-NMMA
2 NAA + BAP
+ NAA + BAP
+ NAA + BAP + 1 mM L-NMMA
+ NAA + BAP before transformation, then 21 h at 38˚C + NAA + BAP
independent survival percentage observed at 38°C in
medium with or without L-NMMA (Table III; Gushwa
et al., 2003; this study). The results of preincubation
control experiments eliminate the possibility that heatmediated suppression of generalized transcription
and/or translation mechanisms is/are responsible for
the lack of auxin-inducible mGFP5-ER accumulation
after heat or L-NMMA treatment. Thus, both heat and
L-NMMA suppress auxin-responsive transgene expression in reporter-transformed GCPs (Fig. 6; Table I;
Dong et al., 2007), suggesting that many of the inhibitory phenomena observed under both treatments are
likely due to failed NO-dependent auxin signaling.
Plants can produce NO nonenzymatically and
through multiple enzymatic pathways (Besson-Bard
et al., 2008). The intracellular origins of, and biosynthetic pathways for, the NO observed in our 32°C
culture experiments are uncertain. The effects of
L-NMMA were stereospecific (Fig. 4E), and a substantial
fraction of L-NMMA-treated cells cultured at 32°C
could be rescued by 1 or 5 mM L-Arg in competition
with 1 mM L-NMMA (Fig. 7), suggesting that NO accumulation was Arg dependent. Recently, the first
plant kingdom Arg-dependent NO synthase gene was
isolated from Ostreococcus species (Foresi et al., 2010),
and it has been suggested that chloroplasts of higher
plants play a central role in NO metabolism (Gas et al.,
2009). Consistent with those studies, in our 32°C experiments, DAF-2T localized primarily in or around
chloroplasts (Fig. 2). However, even in control experiments with the NO generator SNAP, the maximum
percentage of cells in which dye was observed was
approximately 35% to 40%, and perinuclear cytoplasmic dye localization was less common (Fig. 2), raising
the question of whether the dye reliably reports
physiological NO levels and localization.
The advantages and limitations of diaminofluorescein dyes for evaluating NO production have been reviewed (Hilderbrand et al., 2005; Planchet and Kaiser,
2006a). Dyes are the only tractable method for detecting NO in single cells or very small populations of cells
such as those used here, for analyzing distributions
of NO-producing cells in cellular subpopulations, and
for localizing potential sites of NO production or
accumulation within cells (Hilderbrand et al., 2005;
50.6 6 6.4
50.4 6 7.8
ND
49.1 6 6.9
48.0 6 6.1
57.4 6 9.7
48.8
5.9
6.3
0.8
0.2
6 4.2
6 2.4*,**
6 0.4*
6 0.4*,**
6 0.2*,**
ND
Planchet and Kaiser, 2006a). We chose DAF-2DA because other methods for measuring NO require much
larger amounts of tissue than were available here, are
often destructive, and/or rely on specialized detectors
that are generally unavailable and/or prohibitively
expensive (Planchet and Kaiser, 2006a).
It seems unlikely that failure to observe DAF-2T in
60% to 65% of SNAP-treated cells was due to undersaturation with SNAP. At a culture pH of 6.1 (Tallman,
2005) and a SNAP concentration of 2.5 mM, and assuming a half-life of approximately 5 h (Ignarro et al.,
1981) and an estimated conversion efficiency of SNAP
to NO of 0.5 (Feelisch, 1991), NO concentrations 16 h
after SNAP addition to cultures would be approximately 125 to 150 nM, well above the 3 to 10 nM
Figure 7. Effects of increasing concentrations of L-Arg on the division
of cultured tree tobacco GCPs treated with 1 mM L-NMMA. GCPs were
cultured in medium containing an auxin (NAA) and a cytokinin (BAP)
for 1 week at 32˚C without (black bars) or with (white bars) 1 mM LNMMA and the L-Arg concentrations indicated. Values are means 6 SE
from three separate experiments; in each experiment, 300 to 380 cells
were scored microscopically at 2003. A, Significantly different from
no-L-NMMA, no-Arg control; B, significantly different from the corresponding Arg control lacking L-NMMA (ANOVA; Fisher PLSD; P #
0.05).
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Heat Reduces Nitric Oxide to Determine Guard Cell Fate
reactive oxygen chemistry of some cellular and/or
subcellular compartments (Planchet and Kaiser, 2006a),
ascorbic acid and dehydroascorbic acid (Planchet and
Kaiser, 2006a), PTIO (Vitecek et al., 2008), and unidentified compounds (Planchet and Kaiser, 2006b)
can yield false-positive readings with at least some
diaminofluorescein dyes. Thus, full controls are required when diaminofluorescein dyes are used to
evaluate cellular NO levels. The fact that the volume of
the cytoplasm of a guard cell is largely occupied by a
vacuole with acidic content may explain, at least in
part, why little fluorescence was detectable in the cytoplasm. The layer of cytosol beneath the plasma
membrane of the guard cell is also quite thin, so that
cytoplasmic fluorescence is more easily observed near
nuclei where the endoplasmic reticulum is aggregated.
In contrast, the alkaline stroma of a microscopically
illuminated chloroplast might be an ideal environment
for producing DAF-2T fluorescence. The observation
that even SNAP-treated cells exhibited little cytosolic/
perinuclear DAF-2T fluorescence further suggests that
caution may be warranted in interpreting subcellular
localization results. It should be noted, however, that
upon SNAP treatment at 38°C, approximately the
same percentage of chloroplasts showed DAF-2T fluorescence as did chloroplasts of cells cultured at 32°C,
suggesting that any heat-induced alterations to the
environment of this compartment did not affect the
ability of the dye to report NO.
Even in experiments with L-NMMA and PTIO, 7% to
9% of GCPs exhibited low-level background DAF-2T
fluorescence in chloroplasts. While such fluorescence
could be attributed to sources that are insensitive to
L-NMMA, PTIO results suggest that the dye is promiscuous, that PTIO is not always effective as a NO
scavenger, and/or that conditions in certain chloroplasts unrelated to NO result in the chemical conversion of DAF-2DA to DAF-2T. From the standpoint of
protein saturation, the 1 mM concentration of L-NMMA
used in these experiments is typical of literature values
and is high enough to fully suppress NO production in
tobacco (Nicotiana tabacum) plants when measured
with a laser photoacoustic detector (Mur et al., 2005).
Furthermore, the observation that fluorescence persisted in chloroplasts of a small but similar fraction of
Figure 8. Effects of heat (38˚C) or L-NMMA on tree tobacco shoot and
root development. A, Plants grown for 14 d at 21˚C before growth for
another 2 weeks at 21˚C, at 32˚C, at 32˚C with roots and shoots exposed to 1 mM L-NMMA, or at 38˚C. B, Effects of heat and L-NMMA on
primary root growth and lateral root elongation.
detection limit of the fluorescein-based dyes (Hilderbrand
et al., 2005).
The maximum DAF-2DA concentration that could
be loaded in our cell culture medium without creating
obscuring background fluorescence was 5 mM. While
this limitation may have prevented us from detecting
NO in cells with very low NO levels or caused us to
miss NO that was more diffusely distributed throughout the cytoplasm, DAF-2DA concentrations in the
range of 1 to 10 mM are typical in such experiments
(Hilderbrand et al., 2005), and because the dye is freely
cell permeable, over the 30 min period of dye loading
even a concentration of 5 mM should oversaturate the
picomolar-to-nanomolar NO-producing capacity of the
cell (Planchet and Kaiser, 2006a).
DAF-2T fluorescence can be quenched drastically at
lower pH levels (Hilderbrand et al., 2005), and the
Table II. Heat arrests the growth of tree tobacco seedlings, L-NMMA reduces petiole length and leaf expansion, and both prevent lateral root
elongation and reduce the mitotic index of the primary root
Values are means 6 SE. *Significantly different from 32˚C; **significantly different from 38˚C; ***significantly different from 32˚C + 1 mM L-NMMA
(ANOVA; Fisher PLSD; P # 0.05).
Treatment
Petiole Length
(n = 81)
mm
26˚C, initial, at 10 to 14 d
3.5 6 0.2*,**,***
26˚C, final, at 4 weeks
6.3 6 0.3*,**
32˚C at 4 weeks
10.1 6 0.5
38˚C at 4 weeks
1.6 6 0.1*
32˚C + 1 mM L-NMMA at 4 weeks 6.6 6 0.3*,**
32˚C + medium only at 4 weeks
11.1 6 0.6**,***
Leaf Area
(n = 81)
1˚ Root Length
(n = 27)
mm 2
60.5 6
316.5 6
285.7 6
49.0 6
116.1 6
263.8 6
4.0*,***
28.3**,***
31.2
4.4*
6.5*,**
22.8**,***
No. of Lateral Root
Primordia (n = 27)
Root Mitotic Index
(n = 9)
mm
33.8
65.1
53.2
25.1
47.7
57.9
6
6
6
6
6
6
2.4*,**,***
2.5*,**,***
2.9
1.3*
3.0**
2.7**,***
7.0
7.07
12.4
11.7
7.7
11.3
Plant Physiol. Vol. 159, 2012
6
6
6
6
6
6
0.7*,**
0.7*,**
1.3
1.2
0.8*,**
1.9
0.51
0.51
0.42
0.34
0.36
0.42
6
6
6
6
6
6
0.03**,***
0.04**,***
0.04
0.05
0.03
0.09
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Beard et al.
Table III. At 32˚C, many (but not all) of the effects of L-NMMA on the development of cultured tree tobacco GCPs and seedlings are similar to those
of heat (38˚C)
ND, Not determined.
Characteristic/Treatment
GCPs
Cell wall regeneration?
Cell expansion, 1 week
Cell survival, 1 week with
NAA and BAP
Cell survival, 1 week
No NAA
No BAP
No NAA or BAP
Cell cycle reentry?
Cell division?
Cell division with D-NMMA?
Cell division with L-Arg?
BA auxin-responsive promoter
activated?
DAF-2DA fluorescence
Shoots
Shoot development: leaf expansion,
petiole elongation
Roots
Primary root grows?
Lateral root primordia develop?
Lateral roots develop?
Root hairs develop?
Mitotic index, primary root
a
Taylor et al. (1998).
b
This study.
c
32˚C
32˚C + 1 mM L-NMMA
38˚C
Yes (Fig. 3A)
12.5-foldb; up to 29-foldc
69.3% 6 5.5%d
No (Fig. 3B)
5.9-foldb
47.2% 6 1.3%b
Noa
6.6-foldb
71.2% 6 5.5%d
0.3% 6 0.3%d
3.9% 6 3.7%d
2.3% 6 2.3%d
Yes (Fig. 5B)
Yes (Fig. 4B)
Yes (Fig. 4E)
Yes (Fig. 7)
Yes (Fig. 6A; Table I)
32.6% 6 3.7%b
40.3% 6 6.6%b
38.3% 6 2.9%b
No (Fig. 5E)
No (Fig. 4C)
ND
Yes (Fig. 7)
No (Fig. 6B; Table I)
77.3% 6 1.2%d
75.3% 6 2.5%d
69.7% 6 10.1%d
No (Fig. 5C)
No (Fig. 4D)
ND
Nob
No (Fig. 6C; Table I)
Yes (Figs. 1A and 2)
No (Figs. 1B and 2)
No (Figs. 1E and 2)
Control (Table II)
Reduced (Table II)
No; arrested (Table II)
Yes (Table II)
Yes (Table II)
Yes (Fig. 9B)
Yes
Same as 26˚C control (Table II)
Yes; reduced (Table II)
No (Table II)
No (Fig. 9B)
Yes
Reduced versus 26˚C
control (Table II)
No; arrested (Table II)
Yes (Table II)
No (Fig. 9B)
No
Reduced versus 26˚C
control (Table II)
Roberts et al. (1995).
cells despite treatment with L-NMMA and PTIO suggests that this residual fluorescence is unrelated to
Arg-dependent NO production and that, indeed, it
may not be related to NO production at all. Results
with PTIO and L-NMMA at 38°C indicate that the residual fluorescence observed in chloroplasts of cells at
high temperature is probably caused by processes
similar to those that cause the residual fluorescence
observed in chloroplasts at 32°C and not by unique
interactions of the dye with PTIO or L-NMMA at high
temperature.
Despite limitations of the dye, there are plausible
physiological explanations for the observed dye results. NO is a component of the guard cell ABA
(García-Mata and Lamattina, 2002; Hancock et al.,
2011), extracellular Ca2+ (Wang et al., 2012), and SA
(Hao et al., 2010) signaling pathways that cause stomata to close in darkness or under various abiotic
stress conditions. ABA and Ca2+ induce NO production by first activating the production of hydrogen
peroxide (H2O2; Bright et al., 2006; Wang et al., 2012).
In Arabidopsis, production of the NO used in both
ABA and SA signaling is thought to be catalyzed primarily by nitrate reductase (Bright et al., 2006; Ribeiro
et al., 2009; Hao et al., 2010). Interestingly, NO generated by nitrate reductase appears to be required for
ABA-induced stomatal closure in turgid leaves but not
d
Gushwa et al. (2003).
for stomatal closure in leaves that are rapidly wilting
(Ribeiro et al., 2009).
Presumably NO levels must be carefully controlled to
regulate normal stomatal function. Thus, GCPs may
have powerful metabolic, scavenging, and/or sequestration mechanisms (Song et al., 2011) that, in the first
16 h of culture, render levels of endogenous and exogenously supplied NO undetectable with DAF-2DA
in a high percentage of cells. In our studies, L-NMMA
would be expected to inhibit Arg-dependent NO production, not NO production catalyzed by nitrate reductase, suggesting that the sources of NO for guard
cell responses to ABA, Ca2+, and SA differ from the
sources of NO required for auxin- and cytokininmediated cell cycle reentry. Adjusted for background
fluorescence, the 28% of cells that produced NO by an
L-NMMA-sensitive mechanism after 16 h of culture at
32°C is very similar to the 27.9% 6 8.6% of GCP that
entered S-phase in the period 72 to 96 h after the initiation of cultures. This observation is consistent with the
hypothesis that these fluorescing cells represent the
earliest subpopulation of GCP that generates NO by an
L-NMMA-sensitive, Arg-dependent mechanism in preparation for cell cycle reentry (Ötvös et al., 2005; CorreaAragunde et al., 2006). We do not yet know whether
Arg-dependent NO production/accumulation might be
auxin and/or cytokinin dependent in these cells.
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Heat Reduces Nitric Oxide to Determine Guard Cell Fate
If it were possible to use NO from any source to
accomplish ABA, Ca2+, or SA signaling in guard cells,
an interesting prediction of our results would be that
reduction of guard cell NO levels at high temperatures
might reduce guard cell sensitivity to these compounds and ions and thereby limit the capacity of tree
tobacco stomata to close under stressful conditions.
However, it appears that NO-independent pathways
for stomatal closure may dominate under conditions of
extreme stress (Ribeiro et al., 2009).
Studies of the role of NO in auxin signaling (Ötvös
et al., 2005; Correa-Aragunde et al., 2006; Terrile et al.,
2012), the results reported here, and the results of our
previous studies (Table III) suggest a working model
for how heat and L-NMMA might regulate the fate of
cultured GCPs by reducing NO levels (for summary,
see Fig. 9). Regardless of hormone or temperature
conditions, in the absence of Arg-dependent NO production/accumulation, GCPs survive in high percentages (Table III) but do not make the G1-to-S cell cycle
transition (Fig. 4, B–D); they become quiescent instead
(Fig. 4, C and D). It is envisioned (Fig. 9) that at lower
temperatures (32°C or less), NO produced through a
heat/L-NMMA-sensitive, Arg-dependent mechanism
acts as a bifunctional signaling gas that can, at the G1to-S cell cycle checkpoint, either direct cultured GCPs
into S-phase or, depending on the conditions, into a
NO-mediated cell death pathway (below). High temperature (34°C or greater ) or L-NMMA treatment at
lower temperature (32°C or less) blocks Arg-dependent
NO production, leaving cells in, or directing cells into, a
quiescent state (Fig. 9). At lower temperatures, NO
produced from Arg is used to nitrosylate critical residues in the TIR1 auxin receptor (Terrile et al., 2012).
Binding of auxin to the nitrosylated receptor then activates the transcription of S-phase CdK genes (CorreaAragunde et al., 2006), the CdK proteins themselves
(Ötvös et al., 2005), and, at the same time, transcription
of the gene(s) for 1-aminocyclopropane-1-carboxylic
acid synthase (ACS; Arteca and Arteca, 2008), the ratelimiting enzyme in ethylene biosynthesis (Arteca and
Arteca, 2008). We hypothesize that the resulting ethylene production blocks the NO-directed cell death
pathway. Consistent with published studies, auxin is
required to activate ACS transcription (Arteca and
Arteca, 2008), while cytokinin is required to stabilize
the ACS protein (Hansen et al., 2009), perhaps
explaining why tree tobacco GCPs die at lower temperatures in medium lacking auxin and/or cytokinin
(Gushwa et al., 2003). Failure to produce ethylene may
be a signal to cells initiating cell cycle reentry that the
hormonal conditions needed to complete cell cycle
progression are inadequate; both auxin and cytokinin
are known to be required for the production of certain
phase-specific CdKs required for cell cycle progression
beyond S-phase (Francis, 2001).
NO is a well-known mediator of cell death in plants.
For example, in rice (Oryza sativa) leaves, NO is required to nitrosylate proteins that license H2O2induced leaf cell death (Lin et al., 2012). In Arabidopsis
suspension cultures, NO triggers programmed cell
death as a response to pathogenic stress, apparently
through a mechanism independent of a H2O2-induced
mitogen-activated protein kinase cascade (Clarke et al.,
2000). NO also activates programmed cell death in
Zinnia cultures used to study xylogenesis (Neill, 2005).
Our previous results (Roberts et al., 1995) suggest
that at 32°C, cell survival depends largely on whether
ethylene is present in appropriate amounts as cells
approach the S-phase cell cycle checkpoint (Fig. 9). Our
data (Table III) also show that survival is highest at
lower temperatures when hormonal conditions favor
both NO and ethylene production, suggesting that
each modulates the other in a delicate balance that
positively regulates survival at such temperatures.
Inducing the transcription of ACS with an auxin
(NAA; Arteca and Arteca, 2008) and stabilizing the
ACS protein with a cytokinin (BAP; Hansen et al.,
2009) may create an ethylene signaling cross talk with
NO and/or a parallel condition for NO scavenging
(Song et al., 2011) that results in high cell survival
(approximately 70%), auxin-responsive gene expression,
S-phase reentry (Ötvös et al., 2005; Correa-Aragunde
et al., 2006), and cell dedifferentiation so that cells that
successfully traverse S-phase go on to divide (Ötvös
et al., 2005). If threshold NO levels for cell cycle reentry
were lower than those required to trigger cell death
(Bai et al., 2012), ethylene would only need to reduce
NO levels or limit NO activity enough to prevent cell
death. There is some evidence that ethylene produces conditions that cause guard cell NO to be
scavenged. In concert, exogenous auxin and cytokinin have been shown to activate guard cell ethylene
production that can block ABA-induced stomatal
closure (Tanaka et al., 2006). Treating guard cells
Figure 9. Proposed model for NO-dependent, auxin-mediated cell
fate determination in cultured tree tobacco GCPs. At 32˚C (blue), NO
is available to nitrosylate critical residues in the TIR1 auxin receptor.
Auxin binding to nitrosylated TIR1 results in the transcription of genes
for S-phase CdKs and ACS. Stabilized by cytokinin, ACS catalyzes
ethylene biosynthesis that prevents NO-mediated cell death. At 38˚C,
reduced NO production prevents cell death, auxin-mediated cell cycle
reentry, and auxin-dependent ethylene production. The effects of heat
can be mimicked at 32˚C by treating cells with 1 mM L-NMMA.
Plant Physiol. Vol. 159, 2012
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Beard et al.
with ethephon, which degrades to form ethylene, or
with 1-aminocyclopropane-1-carboxylic acid partially
inhibits dark-induced stomatal closure while it reduces
NO levels (Song et al., 2011). Thus, GCPs cultured at
lower temperatures might be expected to default into a
NO-mediated cell death pathway under any condition
that would perturb the delicate NO/ethylene balance
by inhibiting ethylene synthesis or action when auxin
and/or cytokinin are omitted from culture medium
(Table III), when ethylene production is inhibited
with 1-aminoethoxyvinylglycine (AVG), an inhibitor
of ACS (Yu and Yang, 1979), or when ethylene production or signaling is antagonized with exogenous
ABA (Cheng et al., 2009). Consistent with this hypothesis, each of these treatments is lethal to GCPs
cultured at 32°C (Roberts et al., 1995; Gushwa et al.,
2003). At 32°C, regardless of the combination of NAA,
BAP, and/or ABA used in culture medium, reducing
NO accumulation with L-NMMA probably vastly reduces the number of GCPs that enter a cell death
pathway (Table III). It is also possible that by reducing
NO levels, heat and L-NMMA simply block the progression of cells toward a developmental stage at which
NAA, BAP, and ethylene are strictly required for GCPs
to survive.
Our data are consistent with another hypothesis that
when GCPs are under heat stress (38°C), high levels of
ethylene reduce NO levels (Song et al., 2011) within 12
to 16 h and that, thereafter, the loss of NO results in a
gradual reciprocal reduction in levels of ethylene (Fig.
9). Heat reduces NO accumulation to levels similar to
those observed when cells are cultured at lower temperatures in medium with 1 mM L-NMMA (Figs. 1 and
2), and survival at 38°C is not affected by L-NMMA
(this study). If NO is required for auxin signaling, an
expected result of heat treatment would be failed auxin
signaling for the transcription of genes for ACS (Arteca
and Arteca, 2008). In some systems, heat inhibits ACS
transcription (Suzuki et al., 2005) and ethylene biosynthesis (Yu et al., 1980), and at 38°C, GCP survival is
unaffected by whether NAA or BAP is present
(Gushwa et al., 2003), by AVG (Roberts et al., 1995), or
by the ethylene antagonist ABA (Roberts et al., 1995),
while both AVG and ABA are lethal at 32°C (Roberts
et al., 1995). If heat were to reduce the production or
accumulation of NO and then ethylene in turn, the
demand for each to modulate the effects of the other
for optimum survival would be eliminated. Interestingly, 1 or 5 mM L-Arg reversed the inhibition of cell
division by L-NMMA in a substantial fraction of cells
at 32°C (Fig. 7), but concentrations of L-Arg as high as
20 mM did not reverse the inhibition of cell division by
heat. Among the possible explanations is that both NO
and ethylene production must be restored before cells
can reenter the cell cycle at high temperature. Confirmation of these working hypotheses awaits the development of reliable, sensitive techniques for making
specific and simultaneous quantitative measurements
of changes in NO and ethylene levels over physiologically relevant time courses.
Plant cells contain three genomes, nuclear, mitochondrial, and plastidial, each of which has been
evolutionarily conserved to a different extent. Each
genome must be replicated as a part of organelle division in preparation for cell division, each must be
transcribed to fulfill its cellular function, and each has
programming potential to disrupt cellular homeostasis
and cause cell death. In cultured plant cells, NO is
required for DNA replication and auxin-mediated cell
cycle reentry (Ötvös et al., 2005; Correa-Aragunde
et al., 2006), but it can also mediate programmed cell
death (Clarke et al., 2000). It seems plausible that endosymbiotic cellular evolution has resulted in gaseous
cross-signaling among plant organelles that, under
normal conditions, prevents cell death and synchronizes DNA replication in, and division of, three different organelles that share the same cytoplasm.
Rapidly diffusible, volatile gases like NO and ethylene
that can leave their sites of origin and cross membranes readily would be ideally suited to this role.
Some NO-regulated signaling proteins such as NOA1
(formerly NOS1; Guo et al., 2003; Moreau et al., 2008),
a plastidial GTPase thought to be required for plastid
ribosome function (Gas et al., 2009), have already been
identified.
Failed auxin signaling at high temperatures could
explain why GCPs expand only 5- to 6-fold at 38°C
compared with the 20- to 30-fold expansion observed
at 32°C (Fig. 4, B and D). There is a limit to how far
GCPs can expand before they must regenerate cell
walls to maintain osmotic and water potentials. It is
plausible that without the NO-modulated auxin signaling for gene expression required for cell wall formation (Correa-Aragunde et al., 2008; Fig. 3), cell
expansion would be limited to that achievable with
proteins such as ABP1 that can facilitate some aspects
of auxin-mediated growth that are gene independent
(Jones et al., 1998).
The extent to which results with cultured GCPs and
seedlings can be compared remains to be investigated,
but intriguing similarities exist at some levels. For example, heat clearly reduces the number of cells entering mitosis in GCP cultures and in primary root apical
meristems. The effects of heat and L-NMMA on seedling development that occurred in darkness, such as
lateral root formation (Fig. 8; Table II), were more
similar to their effects on GCPs (also cultured in
darkness) than to those such as petiole elongation and
leaf expansion that occurred in light. NO production in
the green alga Ostreococcus tauri is light level and
growth phase dependent (Foresi et al., 2010), but the
effects of light on NO metabolism, scavenging, and
sequestration in cultured tree tobacco GCPs and tree
tobacco plants have not been investigated. The formation of lateral root primordia and lateral roots in
tomato (Correa-Aragunde et al., 2004, 2006) and the
formation of root hairs (Lombardo et al., 2006) in lettuce (Lactuca sativa) and Arabidopsis have been linked
previously to NO and NO-dependent auxin signaling.
In the case of lateral root primordia and lateral root
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Heat Reduces Nitric Oxide to Determine Guard Cell Fate
development in tomato, it is not clear whether the
source of NO is Arg and, if so, whether Arg-dependent
NO production is strictly required (Correa-Aragunde
et al., 2006). Our results with tree tobacco would suggest
that in this particular tobacco species Arg-dependent
NO production might be needed for auxin-mediated
formation of lateral root primordia at moderate temperatures. Although heat did not reduce the number of
lateral root primordia, NO is required for very early
stages of primordia formation, not later stages (CorreaAragunde et al., 2006). Thus, the apparent conflict in
the L-NMMA and heat results could be explained by
the rapid action of L-NMMA versus the slower effects
of heat on root development. The observation that
both heat and L-NMMA inhibited lateral root formation to similar extents in tree tobacco indicates that
Arg-dependent NO production is probably required
for lateral root formation in this species.
That L-NMMA did not inhibit root hair formation at
a moderate temperature in tree tobacco but heat did
inhibit root hair formation may indicate that the NO
needed for root hair formation is not produced by an
Arg-dependent mechanism. In Arabidopsis, root hair
elongation was significantly reduced in the nia1 nia2
nitrate reductase double mutant grown at moderate
temperatures (Lombardo et al., 2006). While root hair
elongation was also reduced in the nos1 mutant (Guo
et al., 2003; Lombardo et al., 2006), this mutant has
since been shown to modulate NO signaling rather
than to catalyze Arg-dependent NO production, as
originally thought (Moreau et al., 2008). Failed root
hair formation in heat-treated tree tobacco could be
explained by generalized NO scavenging induced by
heat-activated ethylene production, as proposed (Song
et al., 2011), and/or by a mechanism other than the
inhibition of NO production. For example, the more
extreme inhibition of primary root elongation by heat
than by L-NMMA (Fig. 8B) may mean that heat-treated
cells never elongate enough to developmentally trigger
the formation of root hairs (Lombardo et al., 2006).
In developing anthers of barley (Hordeum vulgare)
and Arabidopsis, heat stress suppresses the expression
of auxin biosynthetic genes (Sakata et al., 2010; Oshino
et al., 2011). Auxin depletion then represses DNA
proliferation in organelles and nuclei (Oshino et al.,
2011). Although the study that included Arabidopsis
(Sakata et al., 2010) was not designed to test whether
reduced growth might also be linked to altered NO
production or accumulation, consistent with the results of that study, L-NMMA, which would not necessarily be expected to alter indole-3-acetic acid
biosynthesis, was less effective than heat in reducing
tree tobacco shoot growth and leaf expansion (Table
II), and L-NMMA did not fully inhibit elongation of the
primary root (Table II).
That the effects of heat and L-NMMA differed
somewhat from those of cultured GCPs in complex
tissues and organs is not surprising. The synthetic
growth regulator NAA was added to GCP cultures
exogenously to function as an auxin because NAA
does not require a membrane influx transporter
(Yamamoto and Yamamoto, 1998), and NAA still produces embryogenesis despite its metabolism (Ribnicky
et al., 1996). Indole-3-acetic acid, the major auxin in
planta, requires an influx transporter. Furthermore,
indole-3-acetic acid acts in a fluctuating background
of signaling molecules that are being metabolized, sequestered, and transported and for which there are
tissue-specific receptors, signaling pathways, and modulators as well as redundant signaling systems. As
one example, acting through a mitogen-activated
protein kinase cascade, H2O 2 can prevent auxinresponsive gene expression in Arabidopsis and tobacco mesophyll protoplasts (Kovtun et al., 1998,
2000), although our earlier studies provided no evidence for the involvement of H2O2 in suppressed auxin
signaling during sustained heat stress in cultured GCPs
(Dong et al., 2007).
The role of NO in signaling abiotic and biotic plant
stresses (Grün et al., 2006) is an active area of investigation. Plants grow both by cell division and by expanding existing cells; in GCP cultures, heat inhibits
both. Whether heat inhibits aspects of seedling development such as lateral root elongation by suppressing
auxin-responsive gene expression, as it does in cultured GCPs, remains to be determined, but NO has
certainly been implicated in these types of auxinrelated processes. To the extent that the results presented here can be translated to seedling development,
they indicate that even perennial equatorial plants that
have evolved to possess a high BT may be susceptible
to modest increases in temperature, like those projected to accompany the next century of global climate
change (Solomon et al., 2007), and that such changes
may alter the growth and distribution of even the most
inherently thermotolerant plants.
CONCLUSION
Tree tobacco is a highly thermotolerant equatorial
perennial, and its inherent thermotolerance is retained
at the cellular level in cultured GCPs. Heat inhibits NO
accumulation by cultured tree tobacco GCPs, and NO
is required for auxin-mediated gene expression and
cell cycle reentry in these cells. Taken together with
our previous data and those of others, our results
suggest that NO is a central determinant of cell fate in
cultured GCPs both at lower temperatures and under
heat stress. At lower temperatures, NO participates in
auxin-mediated signaling for gene expression that,
when cytokinin is present, leads to stable levels of
ethylene production. It is likely that ethylene then
modulates NO activity to prevent cell death and promote hormone-directed cell cycle reentry. At high
temperatures, high ethylene levels may vastly reduce
NO levels. Subsequently, reducing levels of NO
probably leads to reduced auxin signaling and reduced ethylene production so that cells survive in
high percentages in a quiescent state. Reducing NO
Plant Physiol. Vol. 159, 2012
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Beard et al.
accumulation with L-NMMA mimics the effects of heat
on GCPs. Heat and L-NMMA also reduce the mitotic
indices of root apical meristems and inhibit lateral root
elongation to similar extents in seedlings, suggesting that
heat may inhibit at least some aspects of plant development through Arg-dependent NO production. Tree tobacco is an equatorial perennial that has evolved a high
BT, but modest temperature increases like those projected to accompany global climate change over the next
century are sufficient to alter important aspects of tree
tobacco seedling development. Thus, despite their high
BT, perennial plants that cannot escape bouts of high
heat may be susceptible to the effects of climate change.
MATERIALS AND METHODS
Plants
Tree tobacco (Nicotiana glauca) was grown from seed (Companion Plants)
and maintained as described (Tallman, 2005; Dong et al., 2007).
Protoplast Isolation and Culture
Except for GCPs used in dye experiments aimed at determining whether
heat inhibits NO accumulation (below), GCPs were isolated from adaxial leaf
epidermis as described (Tallman, 2005; Dong et al., 2007). To isolate GCPs for
dye experiments, 0.125% (w/v) Macerozyme R-10 (PhytoTechnology Laboratories) was included in the second enzyme solution (Takemiya and Shimazaki,
2010), and no Pectolyase Y-23 was used in either enzyme solution. In all
experiments, after isolation, GCPs were cultured in darkness as described
(Tallman, 2005). Cell densities, times, temperatures, and hormone and chemical
treatments for individual experiments are provided below.
Effect of Heat on NO Production/Accumulation
We used the NO-detecting dye DAF-2DA (Assay Designs/Stressgen) to
determine whether heat inhibits NO accumulation by GCPs. In three separate
experiments, isolates containing 5.4 3 105 to 12 3 105 GCPs were collected by
centrifugation and resuspended in 2.5 mL of culture medium containing NAA
and BAP. Aliquots (300 mL) were distributed among eight cryogenic vials
divided among four treatments. One vial in each set was cultured for 16 h at
32°C and the other was cultured for 16 h at 38°C.
The NO generator SNAP was dissolved in dimethyl sulfoxide and added to
each of two vials, one per temperature, to a final concentration of 2.5 mM (Lin
et al., 2012) when cultures were first initiated. The NO scavenger PTIO was
dissolved in culture medium and added to a final concentration of 1 mM to two
other vials both when cultures were first initiated and again after the 16-h
culture period (below). Higher concentrations of PTIO typically used in whole
plant experiments (e.g. 1 mM; Flores et al., 2008) were toxic to cultured cells.
L-NMMA was added to two more vials at a concentration of 1 mM (above) at
the beginning of the 16-h culture and again after 16 h at 32°C or 38°C. Two
untreated vials, one at each temperature, were cultured in parallel as controls.
After 16 h of incubation and all other chemical additions, 0.8 mL of 0.25 mM
DAF-2DA was added to each vial to a final concentration of 5 mM (Kojima
et al., 1998); higher concentrations produced high levels of background fluorescence. Cells were returned to 32°C or 38°C incubators for 30 min to load the
dye before the contents of each were transferred to individual chamber slide
wells for examination by fluorescence microscopy under blue actinic light
(Dong et al., 2007). Two hundred randomly selected cells per well per experiment were scored for fluorescence and fluorescence localization.
Cell Wall Regeneration
The fluorescent dye Calcofluor White (Fluorescent Brightener 28; Sigma) was
used to test the effect of L-NMMA on cell wall regeneration. GCPs were cultured
as described above for 1 week at 32°C in medium with NAA and BAP 6 1 mM
L-NMMA or at 32°C in medium with NAA and BAP only. After 1 week, 200 mL
of culture medium was carefully removed from the top of each well and then
replaced by overlayering the more dense culture medium with 200 mL of a 0.1%
(w/v) stock solution of the dye in 0.01 N NaOH to achieve a final dye concentration of 0.05% (v/v). After approximately 5 min at room temperature, 200 mL of
the dye solution was carefully removed and replaced with 200 mL of culture
medium, pH 6.1. This dilution procedure was repeated twice more to reduce
background fluorescence before cells were examined microscopically using actinic
UV light to excite dye fluorescence (Gushwa et al., 2003).
Effects of L-NMMA on Cell Expansion and Survival
To test the effects of L-NMMA on cell expansion and survival under various
hormone and temperature conditions, in each well of eight-well chamber
slides 3.75 3 104 GCPs were suspended in 400 mL of culture medium containing NAA (0.81 mM) and/or BAP (0.166 mM) or neither hormone. To achieve
a final L-NMMA concentration of 1 mM (Garcês et al., 2001), 2 mL of a 0.2 M
L-NMMA stock solution in culture medium was added to the treatment wells;
controls were left untreated. Cell areas were measured from 4003 microscopic
images by digitizing cross-sectional areas of 30 cells in each of three separate
experiments (n = 90) at the widest optical sections of cells (Roberts et al., 1995)
using IPLab software (version 3.9.4 r3) calibrated with a stage micrometer.
Images were collected with an Olympus IX70 inverted microscope with an
RGB color slider (Q-Imaging; model RGB-HM-S) situated between the side
port of the microscope and a Q-Imaging camera (Retiga 2000R). Survival was
estimated after 1 week of culture in darkness at 32°C or 38°C by scoring 300
randomly chosen cells per treatment microscopically at 4003 as living or dead
in each of three separate experiments (n = 900).
Cell Cycle Analysis
BrdU pulse labeling was used to determine whether L-NMMA would
prevent hormone-directed cell cycle reentry (Gushwa et al., 2003). GCPs (5 3
4
10 per well) were cultured for 72 h in 400 mL of medium with NAA and BAP
at 32°C 6 1 mM L-NMMA or at 38°C with NAA and BAP but without LNMMA. After 72 h, 1 mL of 10 mM BrdU (Sigma) was added to each chamber
slide well, and cultures were returned to their respective temperatures for an
additional 24 h. As described previously (Gushwa et al., 2003), at the end of
the incubation period (96 h total), cells were collected, fixed for at least 24 h,
and stained first for DNA and then for BrdU incorporation using Hoechst
stain and an anti-BrdU fluorescein isothiocyanate-labeled antibody (Invitrogen), respectively. In each of three separate experiments, approximately 300
to 500 randomly selected nuclei were scored first with fluorescence microscopy under UV actinic light for Hoechst staining and then for BrdU incorporation under blue actinic light (Gushwa et al., 2003).
Effect of L-NMMA on Auxin-Regulated
Transgene Expression
To test whether L-NMMA would suppress activation of the auxinresponsive BA promoter, GCPs were first transformed as described (Dong
et al., 2007) with BA-mgfp5-ER, a reporter construct in which the thermostable
mgfp5-ER gene is regulated by the BA auxin-responsive promoter, or with 35Smgfp5-ER, a constitutive control construct. After cells were cultured for 24 h at
32°C or at 38°C in medium with NAA and BAP 6 1 mM L-NMMA, 300 healthy
GCP per well were scored microscopically under blue actinic light for mGFP5ER fluorescence in each of three replicate experiments. Additional control
experiments with heat were also conducted. Sixteen hours at 38°C are required
to fully inhibit auxin-responsive mgfp5-ER expression driven by the BA promoter (Dong et al., 2007). To ensure that (1) heat effects were not caused by
generalized suppression of transcription and/or translation and (2) no early,
long-lived mgfp5-ER transcripts accumulated during the transformation protocol that would explain 35S-regulated mGFP5-ER accumulation at 38°C, in
these additional experiments freshly isolated GCPs were preincubated for 16 h
at 32°C or at 38°C in medium with NAA and BAP before they were transformed with BA or 35S constructs. After transformation, cells were cultured at
32°C or at 38°C in medium containing NAA and BAP for an additional 21 h
before mGFP5-ER accumulation was evaluated microscopically.
Specificity of L-NMMA Action
To determine whether L-NMMA acted stereospecifically to inhibit cell division, in some experiments cells were cultured at 32°C in a medium with
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Heat Reduces Nitric Oxide to Determine Guard Cell Fate
NAA, BAP, and 1 mM D-NMMA. To determine whether L-Arg could reverse
L-NMMA-mediated inhibition of cell division, in other experiments cells were
cultured at 32°C or at 38°C in a medium with NAA and BAP 6 1 mM
L-NMMA with or without 0.25, 1, 5, or 20 mM L-Arg.
Effects of Heat or L-NMMA on Seedling Development
To grow seedlings, tree tobacco seeds were surface sterilized as described
above. Sand columns were prepared by securing 300-mm 3 300-mm mesh
nylon around the base of a 2-inch 3 2-inch Pyrex glass cylinder (GREATGLAS) with a gum band and filling cylinders to the top with fine, 30-grade
quartz silica sand (Tallman, 2005). Cylinders were sterilized by autoclaving in
Magenta GA7 boxes fitted with vented lids (PhytoTechnology Laboratories).
Two milliliters of sterile 0.3% Gelzan (Sigma) in S&T II medium (Tallman,
2005) was pipetted in a thin, discontinuous layer onto the top of the sand
column and allowed to solidify in a laminar flow hood. After the medium
solidified, 5 mL of liquid S&T II medium was added to the top of the column
and allowed to percolate through the sand. To ensure that the sand remained
moist, 20 mL of S&T II was then added to the box. Six to eight sterile tree
tobacco seeds were pipetted onto the top of the sand column before boxes
were sealed with vented lids and incubated under continuous white fluorescent light (photosynthetic photon flux density = 100 mmol m22 s21) at room
temperature (approximately 26°C) for 14 to 21 d to establish seedlings of
similar size in each of three replicate trials. The medium in each box was
replaced with fresh medium in the sterile hood every Monday, Wednesday,
and Friday. To allow for sufficient column drainage, after the depleted medium was removed by pipetting, each box was allowed to stand open in the
laminar flow hood for at least 30 min. Fresh medium (25 mL) was then added
to the column before boxes were sealed and returned to light banks.
In temperature experiments, boxes were maintained at 26°C or transferred
to 32°C or 38°C incubators under continuous white fluorescent light (photosynthetic photon flux density = 100 mmol m22 s21). The medium was replaced
in each box every Monday, Wednesday, or Friday as described above.
In some experiments with L-NMMA, only roots were treated; in others,
both roots and shoots were treated. To treat roots, the medium in each box
was replaced with liquid S&T II medium containing 1 mM L-NMMA. LNMMA was delivered to shoots in solidified agar blocks containing 0.2%
Gelzan and 1 mM L-NMMA in S&T II. Medium containing L-NMMA was filter
sterilized, Gelzan was added, and the mixture was autoclaved. Two milliliters
was allowed to solidify to a soft state in a sterile, 5.5-cm-diameter glass petri
dish in the sterile hood. Blocks (1 cm 3 1 cm) of agar were excised with a
sterile scalpel and placed atop the shoot apical meristems of each of the three
largest seedlings in each treatment column. Blocks with medium but without
L-NMMA were used as controls. Agar blocks were replaced with fresh blocks
every Monday, Wednesday, or Friday for 14 d.
Growth measurements were taken 14 d after seedlings were transferred to
higher temperatures or 14 d after the beginning of L-NMMA treatments. All leaf
measurements were made on the three largest plants in each box. Leaf area
expansion was measured using a Bio-Rad GS710 Calibrated Imaging Densitometer and Bio-Rad Quantity One software. Leaves were removed with a
scalpel and placed on the densitometer. The instrument was set to “Live focus,
white transillumination,” and an image was captured. Outlines of each leaf
were made using the volume contour tool as if they were highly distorted gel
bands. Leaf area was defined as the area within the outlined leaf and was
calculated by the software in a volume analysis report. Petiole length was
measured with digital calipers from the base of the leaf to the point where the
petiole connected to the primary shoot.
On each of the three largest plants, primary root length was measured with
digital calipers using a magnifying lens to determine the point at which each
seedling entered the sand column, continuing to the tip of the root meristem.
The number of lateral root primordia on primary roots was also counted using a
magnifying lens. Root hair density was estimated by visualizing roots using a
microscope on wet mount slides under oil immersion at 1,0003.
To evaluate the effects of heat or L-NMMA on cell division, root mitotic
indices were estimated by a slight modification of the procedure described
(Fiskesjö, 1985). From each of the three largest plants in each box, a 2-cm
section from the tip of each primary root was excised using a scalpel and fixed
immediately by complete immersion in three to four drops of ice-cold 1 N HCl
and ethanol (60:40) in a 2-mL microcentrifuge tube. The tube was incubated in
a water bath at 60°C for 5 min before the root was washed with three to four
drops of deionzed water no fewer than three times to remove excess HCl. The
root section was placed on a glass slide, excess water was blotted away with
a Kimwipe, and the section was covered with three or four drops of a 0.4%
(w/v) toluidine blue solution. The dye was set by floating the slide on a
perforated raft directly above the 60°C water for 5 min. Finally, the root was
squashed by placing a coverslip over the section and pressing, and the slide
was viewed with a microscope at 4003. Root mitotic indices were calculated
after counting the number of root tip cells in each of three fields per root tip
that were in interphase, prophase, metaphase, anaphase, and telophase and
then dividing the total number of cells in all phases of mitosis by the total
number of cells counted.
Statistics
Data were analyzed statistically using Statview 4.01 (Abacus Concepts).
ACKNOWLEDGMENTS
We thank P. Gaudin, K. Shimazaki, B. Stebbins-Boaz, M. Dong, and I.H.
Street for technical assistance and helpful discussions.
Received May 10, 2012; accepted June 20, 2012; published June 22, 2012.
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