2. 2009; Cechin et al., 2015) and salt stress (Egbichi et al., 2014; Sheokand
et al., 2010). Among these enzymes is ascorbate peroxidase (APX, EC
1.11.1.11), which plays a vital role in defense against oxidative stress.
APX utilizes ascorbate (AsA) as its specific electron donor to re-
duce hydrogen peroxide (H2O2) to H2O, with the concomitant gener-
ation of monodehydroascorbate/dehydroascorbate (MDHA/DHA)
(Dalton et al., 1986; Asada, 1994; Iturbe et al., 2001). In analogy to
animals, plants have NOS enzymatic activity, which catalyzes the
conversion of L-arginine to L-citrulline, with a simultaneous release
of NO (Wendehenne et al., 2001).
Although the identity of NOS in plants has not been resolved (Bates
et al., 1995; Barroso et al., 1999; Corpas et al., 2001; Crawford and Guo,
2005; Zamojtel et al., 2006), the L-arginine-dependent NO production
provides a convenient tool to investigate a possible similar NO produc-
tion pathway in plants. In fact, several studies have used compounds
such as Nω-Nitro-L-Arginine methyl ester (L-NAME) (Corpas et al.,
2009; Leach et al., 2010) and NG-monomethyl-L-arginine acetate
(LNMMA) (Zhang et al., 2003). These analogues of L-arginine, which
function as competitive inhibitors of animal NOS-mediated NO synthe-
sis on plants, result in decreased NO content.
There are several studies on the individual role of GB and NO in me-
diating plant tolerance against various abiotic stresses. However, to our
knowledge, no investigations have been done on the effects of endoge-
nous NO on GB content and its effect thereof on the APX enzymatic ac-
tivity. In view of this fact, we investigated the effect of inhibition of NOS
activity with L-NAME on GB accumulation and the resulting effect on
APX activity in maize leaves and roots. We also investigated the effect
of L-NAME on cell viability, lipid peroxidation and H2O2 level in maize
seedlings when supplemented with an NO donor (DETA/NO).
2. Materials and methods
2.1. Plant growth
Maize (Zea mays L. cv Silverking) seeds (donated by Capstone Seeds
Pty Ltd) were surface-sterilized in 0.35% sodium hypochlorite for
10 min and then rinsed four times with sterile distilled water. The seeds
were imbibed in sterile distilled water for 20 min and sown in 2 L of
pre-soaked (distilled water) filtered silica sand (98% SiO2, Rolfes® Silica,
Brits), in 20 cm diameter plastic pots. The sand was kept moist by
watering with distilled water during germination. Germinated seedlings
(one plant per pot) were grown on a regulated condition of 25/19 °C
day/night temperature cycle under a 16/8 h light/dark cycle, at a photo-
synthetic photon flux density of 300 μmol photons.m−2
.s−1
during the
day phase. Plants were supplied with nutrient solution [1 mM K2SO4,
2 mM MgSO4, 5 mM CaCl2, 5 mM KNO3, 10 mM NH4NO3, 1 mM K2HPO4
buffer at pH 7.2, 5 μM H3BO3, 5 μM MnSO4, 1 μM ZnSO4, 1 μM CuSO4,
2 μM Na2MoO4, 1 μM CoSO4, 100 μM Fe-NaEDTA, and 10 mM 4-(2-
hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) at pH 7.2] at
the V1 stage (when the collar of the first true leaf is visible). Plants at
the V1 stage which were of similar height were selected for all
experiments.
2.2. Treatment of plants
One week after the plants had reached the V1 stage, control plants
were supplied with nutrient solution every third day. For treatments,
the nutrient solution was supplemented with the following final con-
centrations: 2 mM L-NAME, a combination 2 mM L-NAME with
200 μM DETA/NO, and finally a combination of 2 mM L-NAME with
200 μM Diethylenetriamine (DETA). DETA lacks the NO moiety and
serves as a control for NO treatments. Treatments or nutrient solution
(200 ml per pot) were applied to each plant directly to the sand at the
base of the stem of the plant in the pot every three days. After 10 days
of treatment, plants were carefully removed from the sand, then used
immediately for cell viability or snap-frozen (in liquid nitrogen) for
use in all other experiments.
2.3. Determination of protein concentration
Protein concentrations for all assays were measured in the extracts
as instructed for the RC DC™ Protein Assay Kit II (Bio-Rad Laboratories).
2.4. Determination of glycine betaine content
Estimation of endogenous glycine betaine content in Zea mays was
carried out by modifying a method previously described by Sairam
et al. (2002). Plant root and leaf tissue (250 mg) were ground to a fine
powder in liquid nitrogen. The tissue was incubated in tubes containing
20 ml of de-ionized water for 24 h at 25 °C. The samples were filtered
and mixed with 2 N H2SO4. An aliquot (0.25 ml) was transferred into
a test tube and cooled in ice water for 1 h. Cold potassium iodide-
iodine reagent (0.1 ml) was added, vortexed, and then centrifuged at
1000 × g for 30 min at 4 °C. The sample was incubated for 24 h at
4 °C. The formed periodite crystals were dissolved in 14 ml of 1,2-di-
chloroethane and shaken at room temperature for 48 h. The absorbance
was then read at a wavelength of 365 nm using a FLUOstar Omega UV-
visible spectrophotometer (BMG LabTech GmbH, Ortenberg, Germany).
2.5. Estimation of H2O2 content
In order to determine if the inhibition of NOS activity affects ROS ac-
cumulation, we measured H2O2 content in the maize treatments. The
H2O2 content was determined in the maize root and leaf extracts by
modifying a method previously described by Velikova et al. (2000).
Zea mays tissue (100 mg) was ground to a fine powder in liquid nitrogen
and homogenized in 400 μl of cold 6% (w/v) TCA. The extracts were cen-
trifuged at 12,000 × g for 30 min at 4 °C and 50 μl of the supernatant was
used to initiate the reaction in a mixture (total volume of 200 μl) con-
taining 5 mM K2HPO4, pH 5.0 and 0.5 M KI. The reaction was incubated
at 25 °C for 20 min and absorbance readings were recorded at 390 nm.
H2O2 content was calculated using a standard curve based on the absor-
bance (A390 nm) of H2O2 standards.
2.6. Measurement of lipid peroxidation
Lipid peroxidation was determined in Zea mays root and leaf tissue
by measuring malondialdehyde (MDA) formation, using the thiobarbi-
turic acid (TBA) method as previously described by Buege and Aust
(1978). Plant tissue (100 mg) was ground into a fine powder in liquid
nitrogen and homogenized in 400 μl of cold 5% (w/v) trichloroacetic
acid (TCA). The homogenate was centrifuged at 12,000 x g for 30 min
at 4 °C. Aliquots (100 μl) of the supernatant were mixed with 400 μl of
0.5% TBA (prepared in 20% TCA). The mixture was incubated at 95 °C
for 30 min and the reaction was stopped by placing the mixture on ice
for 5 2min. The mixture was further centrifuged at 12,000 x g for
5 min at 4 °C. The absorbance of the supernatant was measured at 532
and 600 nm. After subtracting the non-specific absorbance (A600 nm),
the MDA concentration was determined by its extinction coefficient of
155 mM−1
cm−1
and expressed as nmol g−1
of fresh weight.
2.7. Evaluation of cell viability in Zea mays roots and leaves
In order to establish if application of NO (200 mμ DETA/NO) could
maintain Zea mays cell viability after the inhibition of NOS, evaluation
of root and leaf cell viability was carried out. This cell viability assay
was estimated by modifying a method previously described by
Sanevas et al. (2007). The tissues (100 mg per treatment) were harvest-
ed and stained with 0.25% (w/v) Evans Blue for 15 min at room temper-
ature. The roots and leaves were then washed for 30 min in distilled
water, followed by extraction of the Evans Blue stain (taken up by
219S. Ullah et al. / South African Journal of Botany 105 (2016) 218–225
3. dead cells) from the tissues using 1% (w/v) sodium dodecyl sulfate
(SDS) after incubation for 1 h at 55 °C. Absorbance of the extract was
measured with a FLUOstar Omega UV-visible spectrophotometer
(BMG LabTech GmbH, Ortenberg, Germany) at 600 nm to determine
the level of Evans Blue taken up by the tissues.
2.8. Determination of APX enzymatic activity
Plant APX activities were measured in Zea mays root and leaf ex-
tracts by modifying a method previously described by Asada (1984).
The extracts which were supplemented with ascorbate to a final con-
centration of 2 mM and were added to the assay buffer containing
50 mM K2HPO4, pH 7.0, 0.1 mM EDTA and 50 mM ascorbate. The reac-
tion was initiated by adding 1.2 mM H2O2 in a final reaction volume of
200 μl and APX activity was calculated based on the change in absor-
bance at 290 nm using the extinction coefficient of 2.8 mM−1
cm−1
.
For the determination of the response of Zea mays APX isoforms the
treatments, electrophoretic APX separation was carried out as previous-
ly described by Mittler and Zilinskas (1993) and non-denaturing poly-
acrylamide gel electrophoresis was performed at 4 °C in 12%
polyacrylamide mini gels. Prior to loading extracts containing 50 μg of
protein into the wells, gels were equilibrated with running buffer con-
taining 2 mM ascorbate for 30 min at 4 °C. After the electrophoresis,
gels were incubated in 50 mM potassium phosphate buffer (pH 7.0)
containing 2 mM ascorbate for 20 min and then transferred to solutions
containing 50 mM potassium phosphate buffer (pH 7.8), 4 mM ascor-
bate and 2 mM H2O2 for 20min. The gels were washed in the buffer
for 1 min and submerged in a solution of 50 mM potassium phosphate
buffer (pH 7.8) containing 28 mM N,N,N′, N′-tetra methyl ethylene di-
amine and 2.5 mM nitroblue tetrazolium for 10–20 min with gentle ag-
itation in the presence of light. The gel images were captured and ana-
lyzed by densitometry using AlphaEase FC imaging software (Alpha
Innotech Corporation).
2.9. Statistical analysis
One-way analysis of variance (ANOVA) test was used to evaluate
statistical validity of the results and means (from three independent ex-
periments) were compared according to the Tukey–Kramer test at 5%
level of significance, using GraphPad Prism 5.03 software.
3. Results
3.1. Glycine betaine content in Zea mays root and leaves
Application of L-NAME caused a decrease in GB content in both root
and leaves of Zea Mays. A decrease of approximately 19% and 17% in GB
content of roots and leaves, respectively, was recorded when compared
to the GB content of roots and leaves from untreated (control) Zea Mays
(Fig. 1A and B). The level of GB content in response to a combination of
L-NAME and 200 μM DETA were very similar to those with treatment of
only L-NAME. Exogenous application of the nitric oxide donor (DETA/
NO) reversed the negative effect of the inhibition of NOS by L-NAME
as the GB content was partially restored (Fig. 1A and B).
3.2. H2O2 content, lipid peroxidation, and cell viability in Zea mays roots
and leaves
Inhibition of NOS by L-NAME in Zea mays resulted in a significant in-
crease of H2O2 content by approximately 68% and approximately 134%
in roots and leaves, respectively, when compared to the untreated con-
trol (Fig. 2A and B). There was no significant difference in the level of
H2O2 content between L-NAME and L-NAME combined with 200 μM
DETA in Zea mays roots and leaves. However, supplementing the L-
NAME treatment with 200 μM DETA/NO resulted to a slight increase
of H2O2 only by approximately 15% in Zea mays roots when compared
with the untreated control. Furthermore, there was no significant differ-
ence in Zea mays leaf H2O2 content between the untreated and a com-
bined treatment of L-NAME with 200 μM DETA/NO.
Inhibition of NOS resulted in oxidative damage to membrane lipids,
as shown by the amount of malondialdehyde content in the L-NAME
treatment of Zea mays roots. Treatment with 2 mM L-NAME showed a
95% increase in lipid peroxidation when compared to the untreated con-
trol (Fig.2C). A similar increase in malondialdehyde content was ob-
served in roots treated with 2 mM L-NAME in combination with
200 μM DETA. Exogenous application of 200 μM DETA/NO combined
with 2 mM L-NAME resulted in complete removal of the negative effect
of NOS inhibition on lipid peroxidation, as shown in Fig. 2C. A similar re-
sponse (albeit with different degrees of change), as observed for roots,
occurred in MDA content of Zea mays leaves in all treatments (Fig. 2D).
Treatment with L-NAME caused a drastic loss of root cell viability. This
is shown by the sharp increase of approximately 287% in Evans Blue
Fig. 1. Glycine betaine content in (A) roots and (B) leaves of Zea mays. GB was measured after 10 days of treatment with either nutrient solution only (Untreated), 2 mM L-NAME, 2 mM L-
NAME + 200 μM DETA or 2 mM L-NAME + 200 μM DETA/NO. Data shown are the mean (±SE) of three independent experiments. Values sharing a common letter are not significantly
different at p ˂ 0.05.
220 S. Ullah et al. / South African Journal of Botany 105 (2016) 218–225
4. uptake compared to untreated plants (Fig. 2E). A combination treatment
with 2 mM L-NAME and 200 μM DETA did not reverse the suppressive ef-
fect of L-NAME on Zea mays root cell viability. This is indicated in Fig. 2E by
an approximately 296% in Evans Blue uptake when compared to untreat-
ed plants. On the other hand, the combination treatment of 2 mM L-
NAME and 200 μM DETA/NO resulted in partial reversal of the reduction
in Zea Mays root cell viability as shown in Fig. 2E. A similar trend (albeit
with different degrees of change) as observed in Zea mays roots for all
these treatments was also observed for Zea mays leaves (Fig 2F).
3.3. Effect of inhibition of NOS activity on APX enzymatic activity
In view of the fact that there was a marked decrease in the level of
H2O2 in plants treated with a combination of 2 mM L-NAME and
200 μM DETA/NO, we analyzed the effect of L-NAME on total APX enzy-
matic activity in Zea mays roots and leaves. As shown in Fig 3A and B,
there was marked decrease of total APX activity in response to L-
NAME. Application of L-NAME decreased total APX activity by approxi-
mately by 71% and approximately 30% in roots and leaves, respectively,
when compared to the untreated controls.
There was no significant difference in the level of total APX activ-
ity in response to L-NAME when combined with 200 μM DETA in
comparison to treatment with L-NAME alone (Fig. 3A and B). On
the other hand, treatment with 2 mM L-NAME combined with
200 μM DETA/NO resulted in partial negation of the suppressive ef-
fect of L-NAME on total APX enzymatic activity in Zea mays roots,
whereas complete reversal in this treatment was observed for leaves
(Fig 3A and B).
Fig. 2. H2O2 content in (A) roots and (B) leaves, together with malondialdehyde (MDA) content in (C) roots and (D) leaves and cell viability in (E) roots and (F) leaves of Zea mays. Assays
were measured after 10 days of treatment with either nutrient solution only (Untreated), 2 mM L-NAME, 2 mM L-NAME + 200 μM DETA or 2 mM L-NAME + 200 μM DETA/NO. Data
shown are the mean (±SE) of three independent experiments. Bars sharing a common letter are not significantly different at p ˂ 0.05.
221S. Ullah et al. / South African Journal of Botany 105 (2016) 218–225
5. Further analysis on the response of individual APX isoforms to NO in
NOS-inhibited Zea mays roots was carried out using in-gel APX enzy-
matic assay coupled with pixel intensity analyses. On visualization,
bands of APX1, APX2, and APX3 isoforms indicated that the untreated
Zea mays had the greatest activity as evidenced by their high band in-
tensity (Fig. 3C). Further observation suggests that APX activity bands
from the combination treatment with L-NAME and 200 μM DETA/NO
had the second highest intensity, with the band intensity from
ZmAPX3 being the highest among the three isoforms in this treatment.
However, Zea mays root APX isoforms from treatment with either 2 mM
L-NAME or 2 mM L-NAME in combination with 200 μM DETA had the
least APX activity (Fig. 3C). This is visually evident from their low
band intensities when compared to the untreated control. Further ob-
servation suggests that APX activity bands from 2 mM L-NAME and
2 mM L-NAME in combination with 200 μM DETA treatments have
slightly different levels of activity. Similar observation as those made
for roots were made for leaves across all the experiments (Fig 3D).
Densitometry analyses (Fig. 4) were used to compare APX enzymatic
activity for each isoform. Pixel intensities obtained indicate that root
ZmAPX1 was slightly decreased by approximately 8% in response to a
combination of 2 mM L-NAME and 200 μM DETA/NO when compared
to root ZmAPX1 enzymatic activity of untreated roots (Fig. 4A). On the
contrary, the enzymatic activity of root ZmAPX1 was decreased by ap-
proximately 18% in response to a combination of 2 mM L-NAME and
200 μM DETA when compared to ZmAPX1 enzymatic activity of un-
treated roots. The enzymatic activity of root ZmAPX2 was significantly
decreased by approximately 14% and approximately 16% in response
to 2 mM L-NAME and a combination of 2 mM L-NAME with 200 μM
DETA, respectively (Fig. 4B). There was no significant difference in the
enzymatic activity of ZmAPX2 between the untreated control and Zea
mays treated with a combination of 2 mM L-NAME and 200 μM DETA/
NO. The enzymatic activity of ZmAPX3 was decreased by approximately
40% in response to 2 mM L-NAME when compared to the activity of
ZmAPX3 in the untreated sample (Fig. 4C). The activity of ZmAPX3 in re-
sponse to a combination of 2 mM L-NAME and 200 μM DETA/NO was
slightly decreased by approximately 20% when compared to the activity
of ZmAPX3 in the untreated sample (Fig. 4C). A marked decrease of
ZmAPX3 activity of approximately 30% in response to a combination
of 2 mM L-NAME and 200 μM DETA was observed when compared to
the activity of ZmAPX3 in the untreated sample (Fig. 4C).
Densitometric analysis of ZmAPX isoforms in leaves showed that
ZmAPX1 and ZmAPX2 had no statistically significant responses to any
of the treatments (Fig 4D and E). The response of leaf ZmAPX3 followed
a similar trend as observed in root ZmAPX3 (Fig 4F and C). Thus, for the
leaves, the activity of ZmAPX3 in response to a combination of 2 mM L-
NAME and 200 μM DETA/NO decreased by approximately 20%, whereas
it was decreased by approximately 45%, when compared to the activity
of ZmAPX3 in untreated leaves (Fig. 4F). Leaf ZmAPX3 activity was
Fig. 3. Effect of inhibition of NOS on APX activity in roots and leaves of Zea mays. Total APX activity was determined in (A) roots and (B) leaves whereas the activities of individual isoforms
of APX were evaluated in (C) roots and (D) leaves. The three isoforms are referred to as ZmAPX1, ZmAPX2, and ZmAPX3 on the basis of their migration on the native PAGE gel. APX activity
was measured after 10 days of treatment with either nutrient solution only (Untreated), 2 mM L-NAME, 2 mM L-NAME + 200 μM DETA or 2 mM L-NAME + 200 μM DETA/NO. Data shown
are the mean (±SE) of three independent experiments. Bars sharing a common letter are not significantly different at p ˂ 0.05.
222 S. Ullah et al. / South African Journal of Botany 105 (2016) 218–225
6. decreased by approximately 35% in response to a combination of 2 mM
L-NAME and 200 μM DETA when compared to the activity of leaf
ZmAPX3 in the untreated sample (Fig. 4F).
4. Discussion
In plants, NOS is an important enzyme that is responsible for the pro-
duction of NO from arginine, using O2 and NADPH as co-substrates (Del
Rio et al., 2004). However, to date, no homologues of the animal NOS
protein exist in higher plants. Furthermore, Foresi et al. (2010) discov-
ered the first and only NOS in green algae Ostreococcus tauri. Neverthe-
less, studies on NOS in higher plants involved the use of classical
chemical inhibitors of eukaryotic NOS (Corpas et al., 2009). Exogenous
application of NOS inhibitors leads to decreased NO content in higher
plants, which suggests that plants possess proteins with NOS-like activ-
ity (Cueto et al., 1996; Corpas et al., 2009). These unidentified NOS-like
proteins play diverse roles in plants and regulate many important cellu-
lar processes. These processes include the development of functional
soybean nodules (Leach et al., 2010), regulating superoxide dismutase
activity (Hao et al., 2008), regulating cysteine protease activity (Leach
et al., 2010), and modulating free amino acid levels (Boldizsár et al.,
2013). On the other hand, one of the most important compatible solutes
in plants, GB, is synthesized by the oxidation of choline (Weigel et al.,
1986; Weretilnyk et al., 1989). It accumulates in the cytosol, chloro-
plasts, and plastids of halo-tolerant plants (Rhodes and Hanson, 1993;
Allard et al., 1998).
There are several studies highlighting the individual roles of NO and
GB under various abiotic stress conditions (Strid et al., 1994; Quan et al.,
2004; Zhang et al., 2006; Zhang et al., 2009; Ali and Ashraaf, 2011; Nazia
et al., 2014). However, the interaction between GB and NO is not well
understood. Gupta et al. (2011) hypothesized that NO derived from
NOS-like proteins might regulate osmolyte accumulation and
Fig. 4. Pixel intensities signifying the level of enzymatic activity of maize root and leaf APX isoforms derived from analysis of the intensity of the activity bands. Activities of (A) ZmAPX1,
(B) ZmAPX2, (C) ZmAPX3 in roots together with (D) ZmAPX1, (E) ZmAPX2, and (F) ZmAPX3 in leaves in response to treatment with 2 mM L-NAME, 2 mM L-NAME + 200 μM DETA or
2 mM L-NAME + 200 μM DETA/NO. Error bars represent the means (±SE; n = 3) of three densitometric values. Values sharing a common letter are not significantly different at p ˂ 0.05.
223S. Ullah et al. / South African Journal of Botany 105 (2016) 218–225
7. contribute to ROS-NO regulation mechanisms. In order to elucidate on
this mechanism, Zhang et al. (2013) inhibited NOS activity with 25 μM
L-NAME and measured NO content and GB content. Results from that
study clearly showed that NO and GB content did not change in re-
sponse to the L-NAME treatment when compared to the control. How-
ever, we deduced from that work and our previous work (Leach et al.,
2010) that the inhibition of NOS activity in plants is dependent on the
concentration of the inhibitor (i.e. more than 25 μM L-NAME is required
to inhibit NOS activity in plants). We also considered the fact that Zhang
et al. (2015) observed a positive correlation between endogenous NO
and GB content in maize leaves under osmotic stress. Therefore, in the
present study, we have shown that inhibition of NOS activity affects
GB content in maize. This is evident from the fact that application of a
NOS inhibitor in the form of 2 mM L-NAME significantly decreased GB
content in maize roots and leaves. This observation is true due to the
fact that not only was the NO released from DETA/NO able to restore
the GB content to levels similar to those of the roots and leaves of un-
treated controls but also in the fact that such improvement was not ob-
served with the “L-NAME + DETA” combined treatment. The decrease
in GB content is rather interesting since it is known that GB accumulates
in a variety of plant species in response to various stresses (Ashraaf and
Foolad, 2007). Our data are in support of studies in maize by Zhang et al.
(2014, 2015), which showed a correlation between GB and NO content.
Whether the alteration of GB content in response to changes in NO con-
tent is a consequence of transcriptional regulation of betaine aldehyde
dehydrogenase (BADH) genes or post-transcriptional regulation (for
example by nitrosylation) of the BADH protein remains to be deter-
mined and forms part of the investigations that we will undertake in
the near future. Ruan et al. (2004) have shown an NO-induced compat-
ible solute accumulation in plant tissue under salt stress, which impli-
cates NO regulation of compatible solute biosynthesis under these
conditions and creates a link between salinity-induced NO accumula-
tion and compatible solute-mediated protection of plants against salin-
ity stress.
In view of the fact that exogenous application of NO to Zea mays
plants led to reduced levels of H2O2 and improved the cell viability in
NOS-inhibited Zea mays suggests that NO mobilized the enzymatic anti-
oxidant defense system. For efficient scavenging of ROS in the cell, APX
(which has a low Km for hydrogen peroxide) catalyzes the conversion of
H2O2 into water. In this reaction, APX utilizes one molecule of ascorbate
as its electron donor (Mehlhorn et al., 1996). The influence of the poorly
resolved isoforms (ZmAPX1 and ZmAPX2) on the accuracy of the densi-
tometry is noted. However, potential inaccuracy in the densitometry (as
a result of poor resolution of these two isoforms) should be of little sig-
nificance since their activity is magnitudes less than the activity of the
fast migrating isoform (ZmAPX3). Small (even if significant) variations
in the activities of APX1 and APX2 as a result of inaccurate densitometry
would make little difference to the total activity of APX in maize. This is
because the bulk of the total APX activity is derived from APX3 and this
isoform is the one with the most pronounced changes in response to the
various treatments. It is worth noting that the statistically significant
but very small interference of DETA on only the activity of the APX3 iso-
form in our experiments does not influence any of the other measured
biochemical, physiological, or cellular parameters, which confirms the
validity of the use of DETA as a valid physiological control for DETA/
NO. Such slight influence has also been reported by Boldizsár et al.
(2013). Our enzyme activity results unambiguously showed that Zea
mays plants supplemented with an NO donor in the presence of the
NOS inhibitor exhibited an increased APX activity as a result of reversal
of NOS inhibition by the supplemented NO, which explains the efficient
quenching of ROS and this is in accordance with Farooq et al. (2009)
who demonstrated that NO improves drought tolerance via regulating
ROS scavenging through antioxidant enzyme activity. This is implies
that the NO could mediate in accumulation of GB, which has the capac-
ity of scavenging the reactive oxygen species under unfavorable condi-
tions. We thus conclude that NO derived from NOS activity is required
for the biosynthesis of GB and influences APX activity to regulate H2O2
levels in maize.
Acknowledgments
This work was supported by Walter Sisulu University (Institutional
Research Grant Award), the University of the Western Cape, and the Na-
tional Research Foundation (South Africa).
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![2009; Cechin et al., 2015) and salt stress (Egbichi et al., 2014; Sheokand
et al., 2010). Among these enzymes is ascorbate peroxidase (APX, EC
1.11.1.11), which plays a vital role in defense against oxidative stress.
APX utilizes ascorbate (AsA) as its specific electron donor to re-
duce hydrogen peroxide (H2O2) to H2O, with the concomitant gener-
ation of monodehydroascorbate/dehydroascorbate (MDHA/DHA)
(Dalton et al., 1986; Asada, 1994; Iturbe et al., 2001). In analogy to
animals, plants have NOS enzymatic activity, which catalyzes the
conversion of L-arginine to L-citrulline, with a simultaneous release
of NO (Wendehenne et al., 2001).
Although the identity of NOS in plants has not been resolved (Bates
et al., 1995; Barroso et al., 1999; Corpas et al., 2001; Crawford and Guo,
2005; Zamojtel et al., 2006), the L-arginine-dependent NO production
provides a convenient tool to investigate a possible similar NO produc-
tion pathway in plants. In fact, several studies have used compounds
such as Nω-Nitro-L-Arginine methyl ester (L-NAME) (Corpas et al.,
2009; Leach et al., 2010) and NG-monomethyl-L-arginine acetate
(LNMMA) (Zhang et al., 2003). These analogues of L-arginine, which
function as competitive inhibitors of animal NOS-mediated NO synthe-
sis on plants, result in decreased NO content.
There are several studies on the individual role of GB and NO in me-
diating plant tolerance against various abiotic stresses. However, to our
knowledge, no investigations have been done on the effects of endoge-
nous NO on GB content and its effect thereof on the APX enzymatic ac-
tivity. In view of this fact, we investigated the effect of inhibition of NOS
activity with L-NAME on GB accumulation and the resulting effect on
APX activity in maize leaves and roots. We also investigated the effect
of L-NAME on cell viability, lipid peroxidation and H2O2 level in maize
seedlings when supplemented with an NO donor (DETA/NO).
2. Materials and methods
2.1. Plant growth
Maize (Zea mays L. cv Silverking) seeds (donated by Capstone Seeds
Pty Ltd) were surface-sterilized in 0.35% sodium hypochlorite for
10 min and then rinsed four times with sterile distilled water. The seeds
were imbibed in sterile distilled water for 20 min and sown in 2 L of
pre-soaked (distilled water) filtered silica sand (98% SiO2, Rolfes® Silica,
Brits), in 20 cm diameter plastic pots. The sand was kept moist by
watering with distilled water during germination. Germinated seedlings
(one plant per pot) were grown on a regulated condition of 25/19 °C
day/night temperature cycle under a 16/8 h light/dark cycle, at a photo-
synthetic photon flux density of 300 μmol photons.m−2
.s−1
during the
day phase. Plants were supplied with nutrient solution [1 mM K2SO4,
2 mM MgSO4, 5 mM CaCl2, 5 mM KNO3, 10 mM NH4NO3, 1 mM K2HPO4
buffer at pH 7.2, 5 μM H3BO3, 5 μM MnSO4, 1 μM ZnSO4, 1 μM CuSO4,
2 μM Na2MoO4, 1 μM CoSO4, 100 μM Fe-NaEDTA, and 10 mM 4-(2-
hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) at pH 7.2] at
the V1 stage (when the collar of the first true leaf is visible). Plants at
the V1 stage which were of similar height were selected for all
experiments.
2.2. Treatment of plants
One week after the plants had reached the V1 stage, control plants
were supplied with nutrient solution every third day. For treatments,
the nutrient solution was supplemented with the following final con-
centrations: 2 mM L-NAME, a combination 2 mM L-NAME with
200 μM DETA/NO, and finally a combination of 2 mM L-NAME with
200 μM Diethylenetriamine (DETA). DETA lacks the NO moiety and
serves as a control for NO treatments. Treatments or nutrient solution
(200 ml per pot) were applied to each plant directly to the sand at the
base of the stem of the plant in the pot every three days. After 10 days
of treatment, plants were carefully removed from the sand, then used
immediately for cell viability or snap-frozen (in liquid nitrogen) for
use in all other experiments.
2.3. Determination of protein concentration
Protein concentrations for all assays were measured in the extracts
as instructed for the RC DC™ Protein Assay Kit II (Bio-Rad Laboratories).
2.4. Determination of glycine betaine content
Estimation of endogenous glycine betaine content in Zea mays was
carried out by modifying a method previously described by Sairam
et al. (2002). Plant root and leaf tissue (250 mg) were ground to a fine
powder in liquid nitrogen. The tissue was incubated in tubes containing
20 ml of de-ionized water for 24 h at 25 °C. The samples were filtered
and mixed with 2 N H2SO4. An aliquot (0.25 ml) was transferred into
a test tube and cooled in ice water for 1 h. Cold potassium iodide-
iodine reagent (0.1 ml) was added, vortexed, and then centrifuged at
1000 × g for 30 min at 4 °C. The sample was incubated for 24 h at
4 °C. The formed periodite crystals were dissolved in 14 ml of 1,2-di-
chloroethane and shaken at room temperature for 48 h. The absorbance
was then read at a wavelength of 365 nm using a FLUOstar Omega UV-
visible spectrophotometer (BMG LabTech GmbH, Ortenberg, Germany).
2.5. Estimation of H2O2 content
In order to determine if the inhibition of NOS activity affects ROS ac-
cumulation, we measured H2O2 content in the maize treatments. The
H2O2 content was determined in the maize root and leaf extracts by
modifying a method previously described by Velikova et al. (2000).
Zea mays tissue (100 mg) was ground to a fine powder in liquid nitrogen
and homogenized in 400 μl of cold 6% (w/v) TCA. The extracts were cen-
trifuged at 12,000 × g for 30 min at 4 °C and 50 μl of the supernatant was
used to initiate the reaction in a mixture (total volume of 200 μl) con-
taining 5 mM K2HPO4, pH 5.0 and 0.5 M KI. The reaction was incubated
at 25 °C for 20 min and absorbance readings were recorded at 390 nm.
H2O2 content was calculated using a standard curve based on the absor-
bance (A390 nm) of H2O2 standards.
2.6. Measurement of lipid peroxidation
Lipid peroxidation was determined in Zea mays root and leaf tissue
by measuring malondialdehyde (MDA) formation, using the thiobarbi-
turic acid (TBA) method as previously described by Buege and Aust
(1978). Plant tissue (100 mg) was ground into a fine powder in liquid
nitrogen and homogenized in 400 μl of cold 5% (w/v) trichloroacetic
acid (TCA). The homogenate was centrifuged at 12,000 x g for 30 min
at 4 °C. Aliquots (100 μl) of the supernatant were mixed with 400 μl of
0.5% TBA (prepared in 20% TCA). The mixture was incubated at 95 °C
for 30 min and the reaction was stopped by placing the mixture on ice
for 5 2min. The mixture was further centrifuged at 12,000 x g for
5 min at 4 °C. The absorbance of the supernatant was measured at 532
and 600 nm. After subtracting the non-specific absorbance (A600 nm),
the MDA concentration was determined by its extinction coefficient of
155 mM−1
cm−1
and expressed as nmol g−1
of fresh weight.
2.7. Evaluation of cell viability in Zea mays roots and leaves
In order to establish if application of NO (200 mμ DETA/NO) could
maintain Zea mays cell viability after the inhibition of NOS, evaluation
of root and leaf cell viability was carried out. This cell viability assay
was estimated by modifying a method previously described by
Sanevas et al. (2007). The tissues (100 mg per treatment) were harvest-
ed and stained with 0.25% (w/v) Evans Blue for 15 min at room temper-
ature. The roots and leaves were then washed for 30 min in distilled
water, followed by extraction of the Evans Blue stain (taken up by
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