Floral Transition and Nitric Oxide Emission During Flower Development in

Arabidopsis thaliana is Affected in Nitrate Reductase-Deficient Plants

K. Seligman1, E. E. Saviani

1, H. C. Oliveira

1, C. A. F. Pinto-Maglio

2and I. Salgado

1,*

1 Departamento de Bioquımica, Instituto de Biologia, CP 6109, Universidade Estadual de Campinas-UNICAMP, 13083-970, Campinas,SP, Brasil2 Laboratorio de Citogenetica, Centro de Pesquisa e Desenvolvimento de Recursos Geneticos Vegetais, CP 28, Instituto Agronomico deCampinas, 13020-902, Campinas, SP, Brasil

The nitrate reductase (NR)-defective double mutant of

Arabidopsis thaliana (nia1 nia2) has previously been shown to

present a low endogenous content of NO in its leaves

compared with the wild-type plants. In the present study, we

analyzed the effect of NR mutation on floral induction and

development of A. thaliana, as NO was recently described as

one of the signals involved in the flowering process. The NO

fluorescent probes diaminofluorescein-2 diacetate (DAF-

2DA) and 1,2-diaminoanthraquinone (1,2-DAA) were used

to localize NO production in situ by fluorescence microscopy

in the floral structures of A. thaliana during floral develop-

ment. Data were validated by incubating the intact tissues

with DAF-2 and quantifying the DAF-2 triazole by fluores-

cence spectrometry. The results showed that NO is synthe-

sized in specific cells and tissues in the floral structure and its

production increases with floral development until anthesis.

In the gynoecium, NO synthesis occurs only in differentiated

stigmatic papillae of the floral bud, and, in the stamen, only

anthers that are producing pollen grains synthesize NO.

Sepals and petals do not show NO production. NR-deficient

plants emitted less NO, although they showed the same

pattern of NO emission in their floral organs. This mutant

blossomed precociously when compared with wild-type plants,

as measured by the increased caulinar/rosette leaf number

and the decrease in the number of days to bolting and

anthesis, and this phenotype seems to result from the

markedly reduced NO levels in roots and leaves during

vegetative growth. Overall, the results reveal a role for NR in

the flowering process.

Keywords: Arabidopsis thaliana — Diaminofluorescein —

Floral induction — Flower development — Nitrate

reductase (NR) — Nitric oxide (NO).

Abbreviations: cPTIO, 2-(4-carboxyphenyl)-4,4,5,5-tetra-methyl-imidazole-1-oxyl 3-oxide; 1,2-DAA, 1,2-diaminoanthraqui-none; DAPI, 40,6-diamidino-2-phenylindole; DAF-2DA,diaminofluorescein-2 diacetate; DAF-2T, diaminofluorescein-2triazole; DMSO, dimethylsulfoxide; MOPS, 3-(N-morpholino)propanesulfonic acid; NO, nitric oxide; NR, nitrate reductase.

Introduction

The free radical nitric oxide (NO) is involved in the

regulation of a broad range of physiological and patho-

physiological processes in animals, ranging from blood

pressure and immune response to neural communication

(Schmidt and Watler 1994). In the last decade, NO has also

been implicated in the modulation of several physiological

and developmental processes in plants, including flowering

(see Lamattina et al. 2003, Desikan et al. 2004, Delledonne

2005, Simpson 2005, Salgado et al. 2004, Salgado et al.

2006, Neill et al. 2007)

The switch to flowering is a major developmental

transition in the plant life cycle and has significant

consequences for reproductive success of the species

(Henderson and Dean 2004, Simpson and Dean 2002).

Exogenous and endogenous signals, such as photoperiod,

vernalization, gibberellins and the circadian clock, are

known inducers of reproductive development (Mouradov

et al. 2002, Simpson and Dean 2002). Recently, NO was

revealed as one of the signals that controls the floral

induction in Arabidopsis. He and collaborators (2004), when

investigating the effect of NO on vegetative growth, showed

that the application of the NO donor sodium nitroprusside

resulted in a delay of flowering induction. Through genetic

screening they identified Arabidopsis mutants with high or

low endogenous NO levels, which delayed flowering or

blossomed precociously, respectively, when compared with

wild-type plants. Moreover, high levels of NO suppressed

the LFY gene in the floral meristem and the CO gene for

floral promotion, which positively affected flowering and,

additionally, increased the expression of the FLC floral

repressor gene. These results suggest a role for NO in

suppressing floral transition by affecting the expression of

genes involved in the regulation of the flowering process.

L-Arginine and nitrite are the major substrates for NO

synthesis in various biological systems. In animals,

the synthesis of NO during the oxidation of L-arginine to

L-citrulline is well characterized and is accomplished by

a family of nitric oxide synthase (NOS) enzymes (Stuehr

1997). NOS-like activity (Barroso et al. 1999, Ribeiro et al.

*Corresponding author: E-mail, ionesm@unicamp.br; Fax, þ55-19-3521-6129.

Plant Cell Physiol. 49(7): 1112–1121 (2008)doi:10.1093/pcp/pcn089, available online at www.pcp.oxfordjournals.org� The Author 2008. Published by Oxford University Press on behalf of Japanese Society of Plant Physiologists.All rights reserved. For permissions, please email: journals.permissions@oxfordjournals.org

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1999) and a putative NOS-like enzyme have been identified

in plants (Guo et al. 2003), although the function of this

putative NOS-like enzyme in NO synthesis is controversial

(Zemotjtel et al. 2006). In addition to L-arginine, nitrite is an

important source of NO in plants and this activity was

originally described as a side reaction of the nitrate

reductase (NR) enzyme (Dean and Harper, 1986), the

major activity of which is to reduce nitrate to nitrite during

nitrogen assimilation. The nitrite-reducing activity of NR

was later demonstrated by other groups (Yamasaki and

Sakihama 2000, Neill et al. 2002, Rockel et al. 2002).

In most of these reports, the nitrite-reducing activity of NR

was substantiated by the inability of NR-deficient plants to

produce NO. However, it was recently demonstrated that

foliar extracts of Arabidopsis thaliana NR double mutant

plants, nia1 nia2, present the same capacity to produce NO

as wild-type plants when nitrite is exogenously supplied

(Modolo et al. 2005). These mutant plants have

lower endogenous contents of nitrite (Modolo et al. 2005)

and L-arginine (Modolo et al. 2006) in their leaves, which

may account for their decreased capability for NO

synthesis. These results suggested that the NR enzyme, in

addition to its nitrite-reducing activity, also has an

important role in generating substrates for NO synthesis,

and an imbalance in its endogenous content may affect

NO-mediated processes. Indeed, the lower endogenous

NO content of NR-deficient A. thaliana plants leaves

them susceptible to pathogen attack (Modolo et al. 2006),

as plant defense response to pathogens is known to be

mediated by NO (Delledonne et al. 1998, Modolo et al.

2002). In accordance with these findings, pathogen signals

were shown to induce expression of NR genes in potato

tubers (Yamamoto et al. 2003), and silencing of NR

genes significantly decreased elicitin-induced NO produc-

tion in Nicotiana benthamiana (Yamamoto-Katou et al.

2006).

Direct measurement of NO is a difficult task because of

its low concentrations and the extremely short-life of this

radical in biological systems. A series of diaminofluo-

resceins, indicator probes that are very sensitive (detection

limits of 5 nM) and non-cytotoxic, were originally devel-

oped for monitoring NO production in neuronal cells

(Kojima et al. 1998). These fluorescent probes have been

used widely for NO monitoring in plant tissues (Pedroso

et al. 2000, Pagnussat et al. 2002, Neill et al. 2002, Corpas

et al. 2004, Prado et al. 2004, Modolo et al. 2006, Zhao et al.

2007). Frequently, tissues are treated with the esterified

form of the probe, diaminofluorescein-2 diacetate (DAF-

2DA), which diffuses into cells where esterases hydrolyze

the diacetate residues, thereby trapping DAF-2 within the

intracellular space. In the presence of oxygen, NO or the

NO-derived species nitrosate DAF-2 to produce the highly

fluorescent DAF-2 triazole (DAF-2T).

Despite the high sensitivity of the diaminofluoresceins

for NO, its use for monitoring NO in plants has received

criticism (Stohr and Stremlau 2006) due to the observation

that diaminofluoresceins also react with dehydroascorbic

and ascorbic acid to generate new compounds that have

fluorescence emission profiles similar to DAF-2T (Zhang

et al. 2002). Considering that dehydroascorbic and ascorbic

acid are common antioxidants, these cross-reactions may

result in confounding interpretations when the diacetate

form of the probe is used to localize in situ NO production.

However, Ye and collaborators (2004) have shown that by

incubating sample tissues directly with DAF-2 these side

effects may be avoided because NO, in contrast to the

interfering molecules, is a gas and can diffuse out of the

tissue and react with DAF-2.

In the present study, we compared the sites of NO

production during flower development and the floral

induction of wild-type A. thaliana with that of the nia1 nia2

mutant plants. We used DAF-2DA and 1,2-DAA to localize

in situ NO production in the floral buds of A. thaliana,

and the results were validated by quantifying NO emitted

from different parts of the plant using the free form of the

diaminofluorescein probe. We discuss the importance of

NR-dependent NO synthesis in the flowering process.

Results

NO production in Arabidopsis thaliana floral structures

localized with DAF-2DA

NO production in wild-type A. thaliana floral buds

in stage 12 of development was analyzed by fluorescence

microscopy of individual floral structures—gynoecium,

stamens, sepals and petals—previously treated with the

NO fluorescent probe DAF-2DA. Stage 12 of floral

development represents the last stage that precedes bud

opening, and corresponds to that in which stigmatic

papillae in the gynoecium are fully developed and stamens

begin to produce pollen grains (Smyth et al. 1990). As

shown in Fig. 1A, in gynoecium previously treated with

DAF-2DA, only the stigmatic papillae showed high levels

of NO emission, indicated by the green fluorescence of

DAF-2T. At this stage of development, NO production

appears to be restricted to certain regions of the stigmatic

cell (Fig. 1B). In the stamens, NO production was detected

in the gametes (Fig. 1C, D) that, in this phase

of development, have already undergone meiosis and the

resulting microspore cells are being differentiated to pollen

grains. When gynoecium was previously treated with the

NO scavenger, 2-(4-carboxyphenyl)-4,4,5,5-tetramethyl-

imidazole-1-oxyl 3-oxide (cPTIO), DAF-2T fluorescence

in stigmatic papillae was blocked (Fig. 1E), indicating

that it corresponded to NO emission. Green fluores-

cence emission in the anthers was also blocked by

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cPTIO (Fig. 1F). Sepals and petals exhibited no green

fluorescence that would be indicative of NO production but

only red fluorescence that corresponds to the chlorophyll

autofluorescence at the wavelength analyzed (Fig. 1G, H).

No fluorescent signals were detected in the controls where

the floral structures were treated with water instead of

DAF-2DA (not shown).

Floral buds of nia1 nia2A. thaliana mutant plants at

stage 12 of development were also imaged for NO after

treatment with DAF-2DA. As seen in the floral structures

of the wild-type plant, in this mutant, DAF-2T fluorescence

was also restricted to the stigmatic papillae of the gynoe-

cium, and in the stamens, DAF-2T fluorescence was only

detected in anthers producing pollen grains (see below).

NO production during flower development localized with

DAF-2DA

Fig. 2 shows NO fluorescent images detected in the

gynoecium and anthers of the nia1 nia2 mutant at different

stages of floral development. At stage 8, which represents

25 µm

A

25 µm

B

50 µm

C

25 µm

D

0.2 mm

E

25 µm

F

0.2 mm

G

0.2 mm

H

Fig. 1 In situ NO production in wild-type A. thaliana floral structures at stage 12 of development, detected with DAF-2DA. Floralstructures were treated with DAF-2DA and NO was detected by the green fluorescence of DAF-2T. (A, B) Gynoecium and (C, D) stamentreated with DAF-2DA. (E) Gynoecium and (F) stamen treated with cPTIO and DAF-2DA. (G) Petal and (H) sepal treated with DAF-2DA,showing the red autofluorescence of chlorophyll.

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the beginning of the late phase of flower development and,

after which most of the cellular differentiation takes

place (Smyth et al. 1990), NO emission in the gynoecium

was restricted to the site where the stigmatic papillae will

develop (Fig. 2A). Significant detection of NO emission

takes place only at stage 11 of floral development, in the

newly differentiated stigmatic papillae that appear at the

top of the stiletto (Fig. 2B). At stage 12, fully developed

stigmatic papillae continue producing NO (Fig. 2C).

At stage 13, when the bud opens and petals are visible,

NO emission is decreased at the stigmatic papillae (Fig. 2D).

In stamens of nia1 nia2 mutants at stage 8 of

development, when they have already begun their

differentiation but are not yet producing pollen grains,

there is no sign of NO emission (Fig. 2E). At this stage,

stamens present only red fluorescence in the wave-

length analyzed, which corresponds to the chlorophyll

autofluorescence. At stage 11 of stamen development

(Fig. 2F) NO production is discretely seen in the micro-

spores. At stages 12 and 13, NO production is clearly

seen in the microspores and forming pollen grains

(Fig. 2G, H).

Floral structures of wild-type plants at different

stages of floral development were also analyzed for NO

imaging after treatment with DAF-2DA. The results

were similar to those observed in nia1 nia2 mutants,

A

0.1 mm

B

50 µm

F

0.15 mm

C

50 µm

G

0.1 mm

D

50 µm

H

E

0.15 mm0.2 mm

Fig. 2 In situ NO production in nia1 nia2A. thaliana gynoecium and stamen during flower development, detected with DAF-2DA. (A–D)Gynoecium and (E–H) stamen at stages 8 (A, E), 11 (B, F), 12 (C, G) and 13 (D, H) of floral development were treated with DAF-2DA andimaged for NO production.

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with maximal NO emission at stages 11 and 12 in the

gynoecium and pollen grains (as showed in Fig. 1),

indicating that the sites of NO emission during flower

development are not influenced by NR double mutation.

NO production in floral structures localized with 1,2-DAA

NO emission during floral development of A. thaliana

was checked by using the NO fluorescent probe 1,2-

diaminoanthraquinone (1,2-DAA). 1,2-DAA is considered

50 µm

A

50 µm

B

20 µm

C

25 µm

D

50 µm

E

0.1 mm

G

0.1 mm

H

10 µm 10 µm

I J K L

25 µm

F

V gg

gg V

Fig. 3 In situ NO production in wild-type A. thaliana stigmatic papillae and microspores detected with 1,2-DAA. Floral structures weretreated with 1,2-DAA, and NO was detected by the red fluorescence of the 1,2-DAA triazole. (A, C, E) Gynoecium and (B, D, F) stamen atstages 11 (A, B), 12 (C, D) and 13 (E, F) of floral development treated with 1,2-DAA. (G, H) Gynoecium and stamen, respectively, at stage12 of development treated with cPTIO and 1,2-DAA (control). (I, K) Images of pollen grains treated with 1,2-DAA and (J, L) thecorresponding images of blue fluorescent nuclear staining with DAPI. v, vegetative nucleus; g, generative nucleus.

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to be more specific than DAF-2DA for in situ NO

visualization, as it reacts selectively with NO (Heiduschka

and Thanos 1998, Dacres and Narayanaswamy 2005). The

pattern of NO emission did not differ between both

genotypes, and fluorescent images obtained for the wild-

type floral structures are shown in Fig. 3. At stage 11 of

development the gynoecium showed intense red fluores-

cence emission, indicative of NO, at the stigmatic papillae

(Fig. 3A), and in the stamens it is possible to see that the

newly formed microspores are not yet producing NO

(Fig. 3B). At stage 12, stigmatic papillae continue pro-

ducing NO (Fig. 3C) and in the anthers NO is emitted

by the mature microspores (Fig. 3D). At stage 13, NO

emission in stigmatic papillae is not seen (Fig. 3E), and

in the stamen (Fig. 3F) it is possible to see tetrads with

almost no NO production, while intense NO synthesis

is seen in mature microspores. NO emission was not

detected when gynoecium and anthers, at stage 12 of

development, were treated with cPTIO and 1,2-DAA

(Fig. 3G and H, respectively), indicating that the red

fluorescence emission detected in the stigmatic papillae and

in the microspores can be attributed to NO. In order to

follow NO production during gametogenesis, stamens were

treated with 1,2-DAA and 40,6-diamidino-2-phenylindole

(DAPI), a specific fluorescent marker of the nucleus. As

shown in Fig. 3I–L, NO production is maximal in the

uninucleate microspore and almost absent in the tri-

nucleated pollen grains, where the two more brightly

stained small generative nuclei and the larger and more

diffusely stained vegetative nucleus are seen by the blue

fluorescence of DAPI.

Quantification of NO emission from floral buds and

vegetative tissues using DAF-2

DAF-2 was used to quantify NO emitted from the

floral structures by adapting the procedure described by Ye

and collaborators (2004). Intact structures were incubated

with DAF-2. During the incubation period, NO emitted

from the tissue can be released into the solution and trapped

by the probe (Ye et al. 2004). Scanning emission fluores-

cence of supernatants from wild-type floral buds showed

peak emission at 515 nm, upon excitation at 495 nm,

characteristic of DAF-2T emission fluorescence (Fig. 4A).

Supernatant solutions from floral buds incubated with

DAF-2DA, instead of DAF-2, showed no fluorescence

emission in this wavelength range (Fig. 4A) confirming that

the esterified form of the probe does not detect NO released

from the tissues. Fig. 4B shows DAF-2T emission fluore-

scence at 515 nm from supernatant solutions of floral

buds of both genotypes at different stages of flower

development, when previously incubated with DAF-2. As

can be seen, much larger amounts of NO are emitted from

wild-type floral buds at stage 11 (15.07� 0.65 nmol g�1 h�1)

than at stages 8 (6.55� 0.61 nmol g�1 h�1) and 13

(5.18� 0.53 nmol g�1 h�1) of floral development. In nia1

nia2 floral buds, estimated NO emission at stages 8,

11 and 13 reached 4.21� 0.54, 11.04� 0.9 and

2.75� 0.41 nmol g�1 h�1, respectively. These results corro-

borated those obtained through in situ NO localization

using DAF-2DA (Fig. 2) and 1,2-DAA (Fig. 3), showing

maximal synthesis of NO just before flower opening.

Additionally, it was possible to show that floral buds of

wild-type plants emitted more NO than nia1 nia2 mutant

plants; at stage 11 of floral development, when NO is

maximally emitted, production of the radical was reduced

by 26.74% in the NR-deficient plant. These results showed

that differences in NO emission by floral buds that could

not be detected between the two genotypes by in situ NO

A

0

4800

50

100

150

200

250

500 520 540 560 580

5

10

15

20

WT

nia1 nia2

Stage 8 Stage 11 Stage 13 Root

B

NO

(n

mo

l.g−1

.h−1

)F

luo

resc

ene

inte

nsi

ty

Floral bud

Wavelength (nm)

DAF-2 DA

DAF-2

Leaf

Fig. 4 Quantification of NO emitted from wild-type and nia1 nia2floral buds, leaves and roots of A. thaliana. Tissues were incubatedwith DAF-2 or DAF-2DA, as described in Materials and Methods,and the emission fluorescence of solutions was scanned for DAF-2T detection, upon excitation at 495 nm. (A) Spectra emissionfluorescence of solutions from wild-type A. thaliana floral budsat stage 11 of development, incubated with DAF-2 or DAF-2DA.(B) Emission fluorescence of solutions recorded at 515 nm, fromwild-type and nia1 nia2 floral buds (at the indicated stages ofdevelopment), roots and leaves, after incubation with DAF-2.

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localization could be detected with the DAF-2 spectro-

fluorimetric assay.

NO emissions from vegetative tissues were estimated

under the same experimental conditions. NO emission from

wild-type leaves was 3.51� 0.53 nmol g�1 h�1 while that of

nia1 nia2 leaves was 0.77� 0.11 nmol g�1 h�1 (Fig. 4B),

showing that NO emission was markedly reduced in the

NR-deficient mutant leaves. These results corroborate

previous observations of the very low endogenous levels

of NO in nia1 nia2 leaves of A. thaliana (Modolo et al. 2005,

Modolo et al. 2006). Fig. 4B also shows that roots of wild-

type plants also emitted much higher levels of NO

(17.08� 1.52 nmol g�1 h�1) than the corresponding tissue

in the NR-deficient plants (8.7� 0.46 nmol g�1 h�1).

Nitrate reductase activity

NR activity was evaluated in floral buds, leaves and

roots by monitoring NO3� reduction to NO2

�. While leaves

and roots of wild-type plants had an NR activity of

646.57� 17.47 and 579.30� 9.30 pmol NO2�min�1mg�1,

respectively, in floral buds, NR activity, if any, was very

low. NR activity in floral buds at stage 8, when detected,

was in the range of 11.82� 5.46 pmol NO2� min�1mg�1,

and at stage 11 and in opened flowers (stage 13 and later)

it was not detected. As expected, NR activity was not

measurable in any of the extracts of nia1 nia2 double

mutants (floral buds, leaves or roots), confirming the

original description by Wilkinson and Crawford (1993) of

the extremely low NR activity of the NR double-deficient

A. thaliana plants.

Effect of NR mutation on floral induction

As shown in Fig. 5, in plants grown under our

experimental conditions (12 h light/12 h dark), bolting in

the wild-type cultivar occurred 37.09� 0.78 d after seeding,

while in nia1 nia2 mutant plants bolting occurred earlier,

at 31.69� 0.31 d after seeding. Similarly, anthesis, the

opening of the floral bud, occurred earlier in nia1

nia2 mutants (35.56� 0.31 d after seeding) compared with

wild-type plants (40.87� 0.99 d after seeding). Thus, on

average, the number of days to bolting and anthesis

was anticipated by 5.4 and 5.3, respectively, in the nia1

nia2 mutant, which represents an important difference

in the timing of floral induction, for a species that has

a life cycle of about 6 weeks under the used growth

conditions. Additionally, on average, wild-type and

nia1 nia2 presented 10.26� 0.28 and 8.56� 0.32 rosette

leaves and 1.74� 0.18 and 1.97� 0.12 cauline leaves,

respectively, at the day of anthesis (Fig. 5). Thus, the

ratio of cauline/rosette leaf number at the day of anthesis

increased from 0.17 in wild-type to 0.23 in nia1 nia2

plants. The overall results showed that NR double

mutation in A. thaliana resulted in an earlier flowering

phenotype when measured either by the number of days to

bolting and anthesis or by the leaf number at the time of

anthesis.

Discussion

In the present study, by using two fluorescent probes,

we localized the sites of NO production in floral structures

of wild-type A. thaliana and showed that NO is emitted by

specific cell types of the gynoecium and stamens, while

sepals and petals do not show any significant NO emission.

Stigmatic papillae and mature microspores are prone to NO

emission, indicating that only structures directly involved

with the reproductive system synthesize this signaling

radical. Moreover, by following NO production during

the late stages of flower development we could show that

maximal NO production coincides with the morphological

differentiation of the reproductive organ, i.e. the stigmatic

papillae differentiation and microspore maturation. The

observation that NO is produced in specific structures in the

plant reproductive system and at a developmental stage that

precedes pollination suggests a role for NO in plant

reproduction. Our data corroborate previous observations

in Senecio and Arabidopsis stigmas, where a decrease in NO

emission was observed at advanced stages of flower

development while developing pollen grains presented an

intense NO emission when followed with DAF-2DA

(McInnis et al. 2006).

By comparing NO production in floral structures of

wild-type plants with that of the NR double-deficient

mutant plants we observed that the sites and stages where

NO is maximally produced in floral structures of A. thaliana

are not affected by NR mutation. In order to quantify and

check the specificity of the in situ NO localization when

treating floral structures of A. thaliana with DAF-2DA,

WT nia1 nia20 30

35

40

45

Lea

f n

um

ber

Days

5

10

15Cauline leafRosette leafBolting time (days)Anthesis time (days)

Fig. 5 Floral induction parameters of wild-type and nia1 nia2A. thaliana plants. The parameters analyzed were the number ofdays to bolting (when the plant had a bolt of 1 cm), the numberof days to anthesis (bud opening) and the number of rosetteand cauline leaves (counted at the day of anthesis), as indicated.Data represent the mean of three independent experiments, withn¼ 35–40. The error bars represent �SE.

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we measured DAF-2T fluorescence after incubating floral

structures with the free form of the probe. When DAF-2

is directly used to quantify NO emission, interfering

molecules such as dehydroascorbic and ascorbic acids can

be avoided (Ye et al. 2004). The results obtained validated

the experiments of in situ NO localization using DAF-2DA

and 1,2-DAA showing that total NO emission by floral

structures decreases with floral senescence. Additionally,

NR-deficient plants were shown to produce less NO in their

floral structures when compared with wild-type plants. The

reduction of approximately 26% in NO emission by nia1

nia2 floral buds could not be attributed to the decreased

activity of NR in these tissues because, even in wild-type

floral buds, NR activity was almost unmeasurable. Thus,

NO synthesis in floral structures seems to depend on

substrate remobilization from vegetative tissues. Possibly,

the role of NR in NO synthesis in floral structures would

be to provide the substrates resulting from the process of

nitrogen assimilation in vegetative tissues. Previous works

have highlighted the importance of NR in providing the

substrates for NO synthesis (see Salgado et al. 2006).

Although the limiting factor for NO production appears to

depend at first glance on substrates (nitrite and L-arginine),

the main mechanism by which NO is produced during

flower development still needs to be elucidated. As

mitochondria have been shown to be an important

organelle in the mechanism of NO synthesis in plants

(Planchet et al. 2005, Modolo et al. 2005), fluctuations

in NO synthesis observed during flower development could

be related to mitochondrial activity, but this remains to be

determined.

We also checked the effect of NR mutation on floral

timing and showed that nia1 nia2 mutant plants had a

decreased number of days to bolting and anthesis and an

increased ratio of caulinar/rosette leaf number compared

with the wild-type plants. As NO was recently identified

as a repressor of floral induction (He et al. 2004) we

additionally checked the effect of NR deficiency on the

levels of NO in the plant during vegetative growth. The

NR-deficient plants were previously shown to present a

lower endogenous content of NO in their leaves when NO

was estimated by gas spectrometry (Magalhaes et al. 2000),

electron paramagnetic resonance (Modolo et al. 2005) and

in situ localization with DAF-2DA (Modolo et al. 2006).

Here, the lower ability of nia1 nia2 leaves to emit NO was

confirmed, and extended to roots, when NO emission was

checked with the DAF-2 probe. The effect of the NR double

mutation in reducing NO emission was clearly seen in these

vegetative tissues that have high levels of NR activity in the

wild-type genotype. These results suggest that the earlier

flowering phenotype of nia1 nia2 mutant plants could result

from the reduced NO content in their leaves and roots

during vegetative growth.

Overall, the present results suggest that NO production

during flower development and floral induction is affected

in nia1 nia2 mutant plants, revealing a role for NR in

flowering control and plant reproduction.

Materials and Methods

Plant material and culture conditions

The assays were conducted with wild-type plants ofA. thaliana L. ecotype Columbia-0 and with nia1 nia2 double-mutant plants of the structural genes of NR, developed byWilkinson and Crawford (1993). The seeds were sterilized withsodium hypochloride (0.3%) for 10min, and the plants cultivatedin pots with vermiculita : perlita (1 : 1). Wild-type plants wereirrigated with half-strength Murashige and Skoog (1962) liquidmedium without the addition of glucose and glycine. The nia1 nia2plants were cultured in the medium described by Wilkinson andCrawford (1991) with reduced amounts of nitrate and ammonia asthe main nitrogen sources. The plants were cultivated undercontrolled conditions in growth chambers at a relative humidity of100%, 248C and a photoperiod of 12 h light and 12 h dark. Floralbuttons at stages 8, 11, 12 and 13 of development (Smyth et al.1990) were used in the experiments.

Nitrate reductase activity

Floral buds, leaves and roots were used to determine the NRactivity, as described by Su et al. (1996), with some modifications.Arabidopsis thaliana tissues (150mg) were ground with a mortarand pestle in liquid N2 and then homogenized in 1ml of cooledextraction buffer, that contained 50mM 3-(N-morpholino) propa-nesulfonic acid (MOPS), pH 7.4, 1mM EDTA, 5mM FAD and7mM cysteine. The mixture was centrifuged at 10,000� g for10min at 48C. An equal volume of assay buffer, composed of50mM MOPS, pH 7.4, 10mM KNO3, 1mM EDTA and 1mMNADH, was added to the resulting supernatant and the mixtureincubated in the dark at room temperature. After 60min, thereaction was stopped by the addition of the Griess reagent, andnitrite production was quantified at 540 nm (Green et al. 1982).The NR activity assay was done in the dark to avoid direct nitrateconversion into ammonium (Dembinski et al. 1996).

NO detection by fluorescence microscopy

Samples of A. thaliana floral buds were collected and thefloral structures were separated (gynoecium, stamen, sepals andpetals) with the aid of a stereomicroscope, and were immediatelytreated for the visualization of NO using DAF-2DA (Calbiochem-Novabiochem, San Diego, CA, USA). Samples of each structurewere incubated with 10 mM DAF-2DA dissolved in 0.1Mphosphate (pH 7.2) buffer, for 30min, under agitation, at258C, in darkness. After this period the solution was removedand the floral structures were washed once with distilledwater. Alternatively, gynoecium and stamen were treated with50mM 1,2-DAA (Calbiochem-Novabiochem), prepared in 100%dimethylsulfoxide (DMSO), instead of using DAF-2DA. For thenegative controls, the floral structures were submitted to the sameconditions, using deionized water instead of the fluorescent probe.To test the specificity of the fluorescent probes, the samples werealso treated with the NO scavenger, cPTIO (Calbiochem), at 1mMfor 30min before incubation with the fluorescent probe. After thetreatments, the samples were mounted on glass coverslips inVectashield (which delays the burning of the fluorescent probe)

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containing DAPI (for DNA staining) and analyzed using aOlympus BX50 fluorescent microscope, equipped with anOlympus CCD (charge-coupled device) digital chamber Q-Color,with refrigeration. For analysis of treated samples, the followingOlympus filters were used: U-MNB2 (excitement at 450 nm andemission at 570 nm) for analysis of the DAF-2DA-treated samples(Kojima et al. 1998); U-N41005 (excitement at 535–550 nm andemission 645–675 nm) for 1,2-DAA-treated samples (Halbach2003) and U-MWU for DAPI (excitement at 372 nm and emissionat 456 nm) (Lakowicz 1999). Images were acquired using thecomputer software Image-Pro Plus, version 6.0 (MediaCybernetics, Inc., Silver Spring, MD, USA) and processed usingthe software Corel Photo Paint. Images shown in the figures arerepresentative of at least five (DAF-2DA) and three (1,2-DAA)replicates per treatment.

Spectrofluorimetric assays

Quantification of NO emission was carried out by incubatingplant tissues directly with DAF-2 in order to eliminate theinterference of intracellular compounds which may react withNO (Zhang et al. 2002), and these assays were based on the methoddeveloped by Ye and collaborators (2004). Floral buds, roots andleaves were incubated for 1 h at 258C, in darkness, with 75 mMDAF-2 (or DAF-2DA for control) prepared in 0.1M phosphatebuffer (pH 7.2). Reaction mixtures from floral buds were thenfrozen and incubated for an additional period of 24 h at –808C, toincrease total NO emission, according to Ye and collaborators(2004). As this additional step did not significantly increase NOemission from floral buds (not shown), it was omitted in the assayswith roots and leaves. The same buffer was then added to thereaction mixture to dilute the probe to a final concentrationof 10 mM, and the suspension was centrifuged at 7,500� g toprecipitate the tissues. DAF-2T fluorescence of the supernatantwas analyzed in a Hitachi F-4500 spectrofluorometer (Hitachi,Tokyo, Japan). Emission fluorescence was scanned from 500 to560 nm upon excitation at 495 nm. Estimation of NO concentra-tion was calculated from a calibration curve using the NOdonor DEA-NONOate, 2-(N,N-diethylamine)-diazenolate-2-oxide(Keefer et al. 1996).

Measurement of flowering time

Floral induction parameters were analyzed by counting thedays to bolting, the days to anthesis, and rosette and caulinarleaf number at day of anthesis. The time of bolting was determinedas the day when the plant had a bolt of 1 cm. Parameters of35–40 plants were measured, in three different experiments.

Funding

Fundacao de Amparo a Pesquisa do Estado de

Sao Paulo (05/54246-4); Conselho Nacional de

Desenvolvimento Cientıfico e Tecnologico (to I.S.).

Acknowledgments

The authors thank Dr. Eneida de Paula for the use of thespectrofluorometer.

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(Received May 7, 2008; Accepted June 4, 2008)

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