fpls-08-00001 February 2, 2017 Time: 17:32 # 1
ORIGINAL RESEARCH
published: 06 February 2017
doi: 10.3389/fpls.2017.00001
Edited by:
Shabir Hussain Wani,
Michigan State University, USA
Reviewed by:
Taras P. Pasternak,
Albert Ludwigs University of Freiburg,
Germany
Rudra Deo Tripathi,
National Botanical Research Institute
(CSIR), India
*Correspondence:
Durgesh K. Tripathi
dktripathiau@gmail.com
Vijay P. Singh
vijaypratap.au@gmail.com
Devendra K. Chauhan
dkchauhanau@yahoo.com
Prashant K. Singh
pksinghau@gmail.com
Specialty section:
This article was submitted to
Crop Science and Horticulture,
a section of the journal
Frontiers in Plant Science
Received: 15 June 2016
Accepted: 03 January 2017
Published: 06 February 2017
Citation:
Tripathi DK, Mishra RK, Singh S,
Singh S, Vishwakarma K, Sharma S,
Singh VP, Singh PK, Prasad SM,
Dubey NK, Pandey AC, Sahi S and
Chauhan DK (2017) Nitric Oxide
Ameliorates Zinc Oxide Nanoparticles
Phytotoxicity in Wheat Seedlings:
Implication of the
Ascorbate–Glutathione Cycle.
Front. Plant Sci. 8:1.
doi: 10.3389/fpls.2017.00001
Nitric Oxide Ameliorates Zinc Oxide
Nanoparticles Phytotoxicity in Wheat
Seedlings: Implication of the
Ascorbate–Glutathione Cycle
Durgesh K. Tripathi
1,2
*
, Rohit K. Mishra
2
, Swati Singh
3
, Samiksha Singh
4
,
Kanchan Vishwakarma
5
, Shivesh Sharma
2,5
, Vijay P. Singh
6
*
, Prashant K. Singh
7
*
,
Sheo M. Prasad
4
, Nawal K. Dubey
1
, Avinash C. Pandey
7
, Shivendra Sahi
8
and
Devendra K. Chauhan
3
*
1
Centre of Advanced in Botany, Banaras Hindu University Varanasi, Varanasi, India,
2
Centre for Medical Diagnostic and
Research, Motilal Nehru National Institute of Technology, Allahabad, India,
3
D D Pant Interdisciplinary Research Lab,
Department of Botany, University of Allahabad, Allahabad, India,
4
Ranjan Plant Physiology and Biochemistry Laboratory,
Department of Botany, University of Allahabad, Allahabad, India,
5
Department of Biotechnology, Motilal Nehru National
Institute of Technology, Allahabad, India,
6
Government Ramanuj Pratap Singhdev Post Graduate College, Koriya, India,
7
Nanotechnology Application Centre, University of Allahabad, Allahabad, India,
8
Department of Biology, Western Kentucky
University, Bowling Green, KY, USA
The present study investigates ameliorative effects of nitric oxide (NO) against zinc oxide
nanoparticles (ZnONPs) phytotoxicity in wheat seedlings. ZnONPs exposure hampered
growth of wheat seedlings, which coincided with reduced photosynthetic efficiency
(F
v
/F
m
and qP), due to increased accumulation of zinc (Zn) in xylem and phloem
saps. However, SNP supplementation partially mitigated the ZnONPs-mediated toxicity
through the modulation of photosynthetic activity and Zn accumulation in xylem and
phloem saps. Further, the results reveal that ZnONPs treatments enhanced levels of
hydrogen peroxide and lipid peroxidation (as malondialdehyde; MDA) due to severely
inhibited activities of the following ascorbate–glutatione cycle (AsA–GSH) enzymes:
ascorbate peroxidase, glutathione reductase, monodehydroascorbate reductase and
dehydroascorbate reductase, and its associated metabolites ascorbate and glutathione.
In contrast to this, the addition of SNP together with ZnONPs maintained the cellular
functioning of the AsA–GSH cycle properly, hence lesser damage was noticed in
comparison to ZnONPs treatments alone. The protective effect of SNP against ZnONPs
toxicity on fresh weight (growth) can be reversed by 2-(4carboxy-2-phenyl)-4,4,5,5-
tetramethyl- imidazoline-1-oxyl-3-oxide, a NO scavenger, and thus suggesting that NO
released from SNP ameliorates ZnONPs toxicity. Overall, the results of the present study
have shown the role of NO in the reducing of ZnONPs toxicity through the regulation of
accumulation of Zn as well as the functioning of the AsA–GSH cycle.
Keywords: amelioration, ascorbate–glutathione cycle, DNA damage, ZnONPs, nanotoxicity, nitric oxide
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Tripathi et al. NO Ameliorates ZnONPs Phytotoxicity
INTRODUCTION
In recent years, nanotechnology has emerged as a scientific trend
that could have benefits in many fields (González-Melendi et al.,
2008; Nair et al., 2010; Gogos et al., 2012; Aziz et al., 2015,
2016; Prasad et al., 2016; Tripathi et al., 2017a,c,d). Compared
to bulk particles, nano forms show unique physicochemical
properties, which are being used as chemical delivery agents
in targeting molecules to a specific cellular organelle in plants
(Monica and Cremonini, 2009; Nair et al., 2010; Subbaiah et al.,
2016; Wang et al., 2016; Tripathi et al., 2017a,b). Several studies
have demonstrated both the beneficial and harmful impacts
of nanoparticles on plants, which are due to type and size
of nanoparticles—especially their specific surface area and the
plant species (Moore, 2006; Dietz and Herth, 2011; Miller et al.,
2012; Tripathi et al., 2015, 2016; Shweta et al., 2016; Singh
et al., 2016). In the ecosystem, plants are not only significant
components but also a latent pathway for nanoparticle transport
and bioaccumulation into food chains (Zhu et al., 2008; Wang
et al., 2012; Zafar et al., 2016).
Among the variety of metal nanoparticles that are frequently
used for commercial purposes, ZnONPs are the most prominent.
They have been exploited in manufacturing paints, glass, plastics,
lubricants, ceramics, pigments, cement, rubber, foods, and
batteries (Monica and Cremonini, 2009). Therefore, extensive
usage of ZnONPs in various products enhances the probability
of their discharge into the environment, which may have
serious consequences on plant productivity. Negative impacts of
ZnONPs have been tested in various plants like ryegrass, rape,
lettuce, radish, corn, cucumber (Lin and Xing, 2007), garden cress
and broad bean (Manzo et al., 2011), zucchini (Stampoulis et al.,
2009), and wheat (Du et al., 2011). Recent studies also exposed the
effect of ZnONPs toxicity on various plant species such as Cicer
arietinum, Brassica nigra, Arabidopsis thaliana, Pisum sativum,
Zea mays, and green alga Picochlorum sp. (Burman et al., 2013;
Hazeem et al., 2016; Mukherjee et al., 2016; Subbaiah et al., 2016;
Wang et al., 2016; Zafar et al., 2016). Thus, the results of previous
studies show that ZnONPs have significant negative effects on
plant productivity, which demands for developing strategies in
order to mitigate ZnONPs toxicity in plants.
Nitric oxide (NO), a gaseous free radical, influences several
physiological and biochemical responses in plants under stressful
and non-stressful conditions. It has been found that NO
effectively alleviates toxic impacts of various stresses like UV-B,
salt, heavy metal, heat, and light in plants (Song et al., 2006;
Shi et al., 2007; Zhang et al., 2009; Xu et al., 2010; Del Río,
2015; Serrano et al., 2015; Singh et al., 2015). Chen et al.
(2015) have reported that NO reduces ZnONPs toxicity in rice
seedlings by regulating oxidative damage and antioxidant defense
Abbreviations: APX, ascorbate peroxidase; AsA, reduced ascorbate; AsA+DHA,
total ascorbate; DHA, dehydroascorbate; DHAR, dehydroascorbate reductase;
F
v
/F
m
, maximum photochemical efficiency of PS II; GR, glutathione reductase;
GSH, reduced glutathione; GSH+GSSG, total glutathione; GSSG, oxidized
glutathione; H
2
O
2
, hydrogen peroxide; MDA, malondialdehyde; MDHAR,
monodehydroascorbate reductase; NPQ, non-photochemical quenching; qP,
photochemical quenching; ROS, reactive oxygen species; ZnONPs, zinc oxide
nanoparticles.
systems. Sufficient literature is available regarding NO-mediated
mitigation of metal stress (Saxena and Shekhawat, 2013; Singh
et al., 2015; Zhao et al., 2016). However, little is still known about
NO-mediated alleviation of nanoparticle toxicity in plants.
In the present study, we have investigated how sodium
nitroprusside (SNP; a donor of NO) regulates toxicity of ZnONPs
in wheat seedlings by examining: (i) Zn accumulation and
physiological and biochemical responses of wheat seedlings
under ZnONPs stress, (ii) estimation of enzymes and metabolites
of the ascorbate–glutatione cycle (AsA–GSH) cycle under
ZnONPs and NO treated seedlings and (iii) mechanisms related
to NO-mediated alleviation of ZnONPs stress.
MATERIALS AND METHODS
Synthesis and Characterization of Zinc
Oxide Nanoparticles (ZnONPs)
Synthesis of ZnONPs was performed by using the procedure of
Sharma et al. (2011) with slight modification. An appropriate
amount (21.9 g) of zinc acetate was taken in 100 ml of
methanol and dissolved while continuously stirring for 2 h
at room temperature. Simultaneously, 140 mM KOH solution
was prepared in 100 ml of methanol with refluxing through
a water condenser with constant stirring for 2 h at 50
◦
C.
Thereafter, both solutions were mixed with constant stirring for
2 h. This mixing was done while refluxing through the water
condenser at 50
◦
C. The final solution was allowed to cool at
room temperature and agitated overnight. This solution was
centrifuged and pellets were washed several times with absolute
ethanol and water in order to remove impurities. The obtained
product was placed in a vacuum oven for 24 h at 50
◦
C to
get powder of ZnONPs. The characterization of ZnONPs was
performed by transmission electron microscopy (TEM), UV-Vis
spectrophotometer, Raman and X-ray diffraction (XRD) analysis,
thermal analysis, and XRD. XRD was performed on a Rigaku
D/max-2200 PC diffractometer operated at 40 kV/40 mA, using
CuKα1 radiation with wavelength of 1.54 Å in the wide angle
region from 20 to 80
◦
on 2θ scale.
Plant Material and Growth Conditions
Wheat (Triticum aestivum L.) seeds were purchased from a
certified supplier of local market in the Allahabad district
of India. Seeds were surface sterilized in 10% (v/v) sodium
hypochlorite solution for 5 min, then washed thoroughly
and soaked for 2–4 h in distilled water. After sterilization
and soaking, uniform-sized seeds were sown in plastic
trays containing sterilized sand. Thereafter, trays were
kept in the dark for seed germination at 25 ± 2
◦
C. When
germination reached maximum of percentage, seedlings
were grown in a growth chamber under photo synthetically
active radiation (PAR) of 250 µmol photons m
−2
s
−1
and
60% relative humidity with 16:8 h light–dark regime at
25 ± 2
◦
C for 8 days. During the growth period, seedlings
were sprayed with water whenever required. Uniformed-sized
seedlings were used to analyze the impact of NO on various
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Tripathi et al. NO Ameliorates ZnONPs Phytotoxicity
physiological and biochemical parameters under ZnONPs
toxicity.
Nitric Oxide and ZnONPs Treatments
Uniform-sized seedlings were gently uprooted from sand and
their roots washed in tap water. Thereafter, seedlings were
acclimatized in half-strength Hoagland’s nutrient solution for
7 days. After this, ZnONPs and NO treatments were given.
SNP was used as a donor of NO. Selection of SNP dose
(100 µM) was based on earlier studies (Singh et al., 2013, 2015).
The treatments include: control (no added ZnONPs and SNP),
SNP (100 µM), 100 µM ZnONPs, 100 µM ZnONPs+100 µM
SNP, 200 µM ZnONPs and 200 µM ZnONPs+100 µM
SNP. In the case of ZnONPs+SNP treatments, seedlings
were pretreated with SNP prepared in a nutrient solution
for 24 h under PAR of 250 µmol photons m
−2
s
−1
. They
were then given ZnONPs treatments. Just after ZnONPs and
SNP treatments, seedlings were further grown in a growth
chamber for 7 days under 250 µmol photons m
−2
s
−1
of
PAR and 60% relative humidity with 16:8 h light–dark regime
at 25 ± 2
◦
C. During this growth period, the medium of
various treatments was changed twice and aerated daily to avoid
root anoxia. After 7 days of ZnONPs and SNP treatments,
seedlings were harvested and various parameters were analyzed
immediately. To test whether NO released from SNP is involved
in ZnONPs toxicity alleviation, 2-(4-carboxy-2-phenyl)-4, 4, 5,
5-tetramethylimidazoline-1-oxyl-3-oxide (c-PTIO, 100 µM), a
scavenger of NO, was used.
Estimation of Growth, Photosynthetic
Pigments, Chlorophyll Fluorescence, and
NO Content
Growth was measured in terms of fresh weight. Seedlings
were selected randomly from control and treated samples and
then their fresh weight was determined. For the estimation
of photosynthetic pigments (total chlorophyll, chlorophyll a +
chlorophyll b), the method of Lichtenthaler (1987) was adopted.
For the assessment of photosynthetic performance, chlorophyll
a fluorescence measurements were taken in the dark adapted
leaves of control and treated seedlings using hand held leaf
fluorometer (FluorPen FP 100, Photos System Instrument, Czech
Republic). The estimation of NO was performed according to
the method of Zhou et al. (2005) as described in Singh et al.
(2015).
Estimation of Zn Content in Seedlings
For the determination of Zn content, dried samples (50 mg)
from control and treated seedlings were digested in mixed
acid (HNO
3
:HClO
4
; 85:15, v/v) until transparent solution was
obtained. The volume of the digested sample was maintained up
to 30 ml with double distilled water. The content of Zn in digested
samples was determined by an atomic absorption spectrometer
(iCE 3000 Series, model-3500 AAS, Thermo scientific, UK) fitted
with a specific lamp of particular metal using the appropriate drift
blank.
Collection of Phloem and Xylem Saps,
and Estimation of Zn Content
The collection of phloem and xylem saps from wheat seedlings
was performed according to the method of Hazama et al. (2015).
Treated and untreated seedlings of 27 days cut on surfaces of
the stems near the petioles of mature leaves with a razor blade
and exuded drops (excluding the first drop) collected as phloem
sap using micropipettes. Samples of phloem sap were stored in
eppendorf previously washed with 0.1 M HNO
3
for 2 days and
then with double distilled water three times to eliminate metals.
The samples were stored at −20
◦
C until analysis.
After the collection of phloem sap, stems were cut at 2 cm
above the interface of the shoot and root, and xylem sap exudates
were collected for 30 min using micropipettes. The measured pH
of the xylem sap was 6.0–6.4. Samples of xylem sap were also
stored at −20
◦
C until analysis.
For the estimation of Zn in phloem and xylem saps above, the
described procedure (for Zn) was followed.
Estimation of Hydrogen Peroxide (H
2
O
2
)
and Lipid Peroxidation
For the estimation of H
2
O
2
, fresh samples (50 mg) from control
and treated seedlings were crushed in 0.1% (w/v) trichloroacetic
acid (Velikova et al., 2000). Absorbance of the reaction
mixture was recorded at 390 nm. The H
2
O
2
concentration was
calculated by using a standard curve prepared with H
2
O
2
. Lipid
peroxidation as MDA content was estimated according to the
method of Heath and Packer (1968). The values of non-specific
absorption recorded at 600 nm were subtracted from the values
recorded at 532 nm. The content of MDA was determined by
using an extinction coefficient of 155 mM
−1
cm
−1
.
Estimation of Activities of APX, GR,
MDHAR, and DHAR: the
Ascorbate–Glutathione Cycle Enzymes
Fresh samples (1.0 g) from control and treated seedlings
were homogenized in 10 ml of chilled 50 mM potassium
phosphate buffer (pH 7.0) containing 1 mM EDTA and 1% (w/v)
polyvinylpyrrolidone in mortar and pestle under cool conditions.
In the case of APX and DHAR activities, 1 mM ascorbic acid
and 2 mM 2-mercaptoethanol were added into the above buffer,
respectively. The homogenate was centrifuged at 20,000 g for
10 min at 4
◦
C and supernatant was used as an enzyme. All
enzymatic measurements were carried out at 25
◦
C by using a
Shimadzu, UV-VIS Spectrophotometer (UV-1700 Pharma Spec)
(Gangwar et al., 2011).
APX (EC 1.11.1.11) activity was determined according to the
method of Nakano and Asada (1981). The decrease in absorbance
was measured at 290 nm. The enzyme activity was calculated by
using an extinction coefficient of 2.8 mM
−1
cm
−1
. One unit (U)
of enzyme activity is defined as 1 nmol ascorbate oxidized min
−1
.
Glutathione reductase (EC 1.6.4.2) activity was assayed
according to the method of Schaedle and Bassham (1977). The
decrease in absorbance was read at 340 nm, and GR activity was
calculated using an extinction coefficient of 6.2 mM
−1
cm
−1
. One
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Tripathi et al. NO Ameliorates ZnONPs Phytotoxicity
unit (U) of enzyme activity is defined as 1 nmol NADPH oxidized
min
−1
.
Monodehydroascorbate reductase (EC 1.6.5.4) activity was
estimated according to the method of Hossain et al. (1984). The
enzyme activity was calculated using an extinction coefficient of
6.2 mM
−1
cm
−1
. One unit (U) of enzyme activity is defined as
nmol NADPH oxidized min
−1
.
Dehydroascorbate reductase (EC 1.8.5.1) activity was assayed
by the method of Nakano and Asada (1981). An increase
in absorbance was read at 265 nm, and DHAR activity was
calculated using an extinction coefficient of 7.0 mM
−1
cm
−1
. One
unit (U) of enzyme activity is defined as 1 nmol DHA reduced
min
−1
.
Estimation of Metabolites of the
Ascorbate–Glutathione Cycle: Ascorbate
and Glutathione
Total ascorbate, AsA, and DHA were determined by the method
of Gossett et al. (1994). This assay is based on the reduction
of Fe
3+
into Fe
2+
with ascorbic acid in acid solution followed
by formation of red chelate between Fe
2+
and 2, 2
0
- bipyridyl.
Control and treated samples were homogenized in 10 ml of
5% (w/v) m-phosphoric acid using a mortar and pestle in
cool conditions. The homogenate was centrifuged at 22,000 g
for 15 min. AsA+DHA was determined in a reaction mixture
consisting of 0.2 ml of supernatant, 0.5 ml of a 150 mM potassium
phosphate buffer (pH 7.4) containing 5 mM EDTA and 0.1 ml of
10 mM DTT to reduce DHA to AsA. After 10 min of incubation
at room temperature, 0.1 ml of 0.5% (w/v) N-ethylmaleimide
was mixed in samples and stirred for 5 min to remove excess
DTT. AsA was assayed in a similar manner, except that DTT
was substituted by 200 µl deionized H
2
O. Color developed in
the reaction mixture through the addition of 0.4 ml of 10%
(w/v) TCA, 0.4 ml of 44% (v/v) o-phosphoric acid, 0.4 ml of
4% (w/v) 2, 2
0
-bipyridyl in 70% (v/v) ethanol and 0.2 ml of
3% (w/v) FeCl
3
. The reaction mixture was incubated at 40
◦
C
for 1 h and quantified spectrophotometrically at 525 nm. DHA
was determined by subtracting AsA from AsA+DHA. Ascorbate
content was found out by using a standard curve prepared with
L-ascorbic acid.
Total (GSH + GSSG), reduced (GSH), and GSSG contents
were estimated using the enzyme recycling method of Brehe and
Burch (1976). This method is based on the sequential oxidation
of GSH by 5, 5-dithiobis-2-nitrobenzoic acid (DTNB) and the
reduction of GSSG in the presence of NADPH and GR (type
III from bakers’ yeast; Sigma Chemical Company). Control and
treated samples were crushed in 3 ml of 6% (w/v) m-phosphoric
acid containing 1 mM EDTA using a mortar and pestle and
centrifuged at 1000 g for 10 min. The reaction mixture (2.4 ml)
contained 0.8 ml of reagent A [110 mM Na
2
HPO
4.
7H
2
O, 40 mM
NaH
2
PO
4
. H
2
O, 15 mM EDTA, 0.3 mM DTNB and 0.04% (w/v)
BSA], 0.64 ml reagent B [1 mM EDTA, 50 mM imidazole and
0.02% (w/v) BSA], which contained an equivalent of 1.5 units GR
activity ml
−1
and 0.8 ml of a 1:20 dilution of acid extract in 5%
(w/v) Na
2
HPO
4
(pH 7.5). The dilution of acid extract was done
immediately prior to starting the reaction through the addition
of 0.16 ml of 3 mM NADPH. The change in absorbance of the
reaction mixture was measured at 412 nm for 5 min. GSSG was
estimated by first incubating 1 ml of 1:20 diluted extract with
40 µl of 2-vinylpyridine for 1 h at 25
◦
C with vigorous shaking.
The samples incubated with 2-vinylpyridine were used for the
assay of GSSG content. The amount of GSH was determined
by subtracting GSSG from GSH+GSSG using a standard curve
prepared with GSH.
Statistical Analysis
Results were statistically analyzed by an analysis of variance
(ANOVA). Duncan’s multiple range test was applied for mean
separation for significant differences among treatments at the
P < 0.05 significance level. The results presented are the
mean ± standard error of six replicates (n = 6).
RESULTS
Observation of ZnONPs
The UV-Vis spectrum of prepared ZnONPs solution is shown in
Supplementary Figure S1A. The prepared ZnONPs have a blue-
shifted absorbance peak approximately at 340 nm compared with
that of the bulk ZnO, which has an absorbance peak at 370 nm
corresponding to a 3.35 eV band gap at room temperature.
Supplementary Figure S1B shows typical Raman-scattering
spectrum of the ZnONPs performed at room temperature.
A sharp, strong, and dominant peak located at about 438 cm
−1
was observed, which was characteristic of the scattering peak of
the Raman active dominant E
2
(high) mode of wurtzite hexagonal
ZnONPs. In addition, some very weak peaks at 331 cm
−1
were
also observed, which were assigned to E
2H
–E
2L
(multi-phonon).
Some broadened and weak peaks at 580 cm
−1
also appeared in
the spectrum.
The available reflections of the present XRD phases have
been fitted with Gaussian distribution. The broadening of XRD
peaks (i.e., Scherrer’s broadening) attributes the formation of
ZnONPs. The crystallite size, d, of ZnONPs was estimated by
Debye–Scherrer’s equation:
d =
0.9λ
β Cosθ
where, d is the crystallite size, λ is the wavelength of radiation
used, θ is the Bragg angle, and β is the full width at half maxima
(FWHM) on 2θ scale. The crystallite size was estimated for the
most prominent X-ray diffraction, corresponding to a peak at
36.25
◦
, and was deduced to be 15–18 nm (Supplementary Figure
S1C).
The variations in XRD results were well-supported by
TEM measurements. Supplementary Figure S1D shows the
representative TEM image of the prepared sample. The
morphology of the sample was found to be nearly spherical in
nature having diameters ranging from 5 to 20 nm. Supplementary
Figure S1E shows the particle size distribution with the mean at
15.37 nm.
Supplementary Figure S1F shows that thermal analysis
of ZnONPs. The ZnONPs had a very small weight loss
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Tripathi et al. NO Ameliorates ZnONPs Phytotoxicity
FIGURE 1 | Impact of SNP and cPTIO interaction on alleviation of ZnONPs toxicity. Data are mean ± standard error of six replicates (n = 6). Bars followed by
different letter(s) show a significant difference at the P < 0.05 significance level according to the Duncan’s multiple ranges test.
(−2.5 wt %) below 300
◦
C, probably due to the removal
of physically and chemically adsorbed water on their
surfaces. The second weight loss (−15 wt%) at about
300
◦
C is assigned to the decomposition of remaining
Zn(OH)
2
.
Growth and NO and Zn Accumulation
Growth was measured in terms of fresh weight and declined
significantly (P < 0.05) following ZnONPs treatments
(Figure 1). Treatment of the wheat seedling with 100 and
200 µM ZnONPs resulted in a significant decline in fresh
weight by 19 and 28% respectively as compared to the
control. The addition of NO donor, i.e., SNP significantly
(P < 0.05) alleviated the ZnONPs-induced decline and showed
a decrease of only 9 and 17% in fresh weight respectively
over the value of the control (Figure 1). However, the
addition of c-PTIO, a scavenger of NO, reverses the NO-
mediated alleviative effect on ZnONPs toxicity (Figure 1),
suggesting a role for SNP released NO in ameliorating ZnONPs
toxicity.
Wheat seedlings grown under 100 and 200 µM ZnONPs
treatments accumulated about 391.4 ± 3.9 and 542.3 ± 5.4 µg
Zn g
−1
dry weight. On the other hand, the addition of SNP
(100 µM ZnONPs + SNP and 200 µM ZnONPs +SNP)
significantly lowered the excess enhancement of ZnONPs in
plants. Accumulation was only187.2 ± 1.8 and 289.5 ± 2.9 µg
Zn g
−1
dry weight (Table 1).
Similar to the results of growth, the exposure of 100 and
200 µM ZnONPs decreased NO content by 27 and 45%,
respectively compared to the control (Table 1). The addition of
SNP together with both doses of ZnONPs significantly raised
levels of NO in wheat seedlings compared to the even control
sample (Table 1).
Impact of NO on Photosynthetic
Pigments and Chlorophyll A
Fluorescence under ZnONPs
Phytotoxicity
Total chlorophyll content decreased by 17 and 28% exposed
to 100 and 200 µM of ZnONPs respectively over the
value of control. However, the addition of SNP significantly
alleviates the adverse impact, and reductions were only 6
and 12% under combinations of SNP+100 µM ZnONPs
and SNP+200 µM ZnONPs respectively. SNP alone showed
significant improvement in chlorophyll content (Table 2).
The results pertaining to chlorophyll fluorescence showed that
the exposure of ZnONPs (100 and 200 µM) significantly declined
F
v
/F
m
by 8 and 24% and qP by 11 and 31%compared to their
TABLE 1 | Effect of SNP on NO and Zn contents in seedlings and xylem
and phloem saps exposed to ZnONPs phytotoxicity.
Treatments NO content
(ng g
− 1
dry weight)
Zn in
seedlings
(mg kg
− 1
dry)
Zn content (ng ml
− 1
)
Phloem sap xylem sap
ZnONP (µM)
0 76.5 ± 2.6d 92.6 ± 2.6f 2011.2 ± 24.1e 242.3 ± 3.8e
100 55.8 ± 1.4e 391.4 ± 3.9b 4484.9 ± 27.2b 566.9 ± 4.6b
200 41.8 ± 1.3f 542.3 ± 5.4a 6395.6 ± 32.4a 811.7 ± 5.3a
ZnONP (µM)+SNP(µM)
0 96.7 ± 3.8b 98.6 ± 3.5e 2005 ± 19.6e 245.3 ± 3.6e
100 88.9 ± 2.7c 187.2 ± 1.8d 3077.1 ± 24.5d 346.5 ± 3.8d
200 127.8 ± 3.1a 289.5 ± 2.9c 3881.6 ± 27.3c 513.8 ± 4.2c
Data are mean ± standard error of six replicates (n = 6). The values followed
by different letters within the same column are significantly different at P < 0.05
significance level according to the Duncan’s multiple ranges test.
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