Reactive oxygen species are involved in cell death in wheat roots against powdery mildew

Journal of Integrative Agriculture 2019, 18(9): 1961–1970 Available online at www.sciencedirect.com

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RESEARCH ARTICLE

Reactive oxygen species are involved in cell death in wheat roots against powdery mildew LI Cheng-yang1*, ZHANG Nan1*, GUAN Bin1, ZHOU Zhu-qing1, MEI Fang-zhu2 1 2

Laboratory of Cell Biology, College of Life Science and Technology, Huazhong Agricultural University, Wuhan 430070, P.R.China College of Plant Sciences & Technology, Huazhong Agricultural University, Wuhan 430070, P.R.China

Abstract Inoculation of wheat (Triticum aestivum L.) leaves with wheat powdery mildew fungus (Blumeria graminis f. sp. tritici) induces the cell death in adventitious roots. Reactive oxygen species (ROS) play a key role in respond to biotic stress in plants. To study the involvement of ROS and the degree of cell death in the wheat roots following inoculation, ROS levels and microstructure of root cells were analyzed in two wheat cultivars that are susceptible (Huamai 8) and resistant (Shenmai 8) to powdery mildew fungus. At 18 d after powdery mildew fungus inoculation, only Huamai 8 displayed the leaf lesions, while root cell death occurred in both varieties. Huamai 8 had a high level of ROS accumulation, which is associated with increased root cell degradation, while in Shenmai 8, there was little ROS accumulation correlating with slight root cell degradation. The molecular study about the expression levels of ROS scavenging genes (MnSOD and CAT) in wheat roots showed that these genes expression decreased after the leaves of wheat was inoculated. The difference between Huamai 8 and Shenmai 8 on subcellular localization of H2O2 and O2–· was corresponded with the different down-regulation of the genes encoding for superoxide dismutase and catalase in two wheat cultivars. These results suggested that ROS were involved in the process by which powdery mildew fungus induced cell death in wheat roots. Keywords: powdery mildew, wheat (Triticum aestivum L.), reactive oxygen species, ultrastructure, programmed cell death

disease (Ji et al. 2008). PM mainly infects the epidermis

1. Introduction Wheat powdery mildew, caused by powdery mildew fungus (PM) (Blumeria graminis f. sp. tritici), is a severe plant

of leaves and stems, forming a sucker apparatus to absorb nutrients from the host (Sutton et al. 2007). During the interactions between plant and fungus, a hypersensitive response (HR), the rapid death of plant cells, is often observed at the inoculated sites. The HR limits the growth of the fungus, releases signals that cause defensive responses in the surrounding cells, and induces systemic

Received 14 April, 2018 Accepted 24 August, 2018 Correspondence ZHOU Zhu-qing, E-mail: zhouzhuqing@mail. hzau.edu.cn * These authors contributed equally to this study. © 2019 CAAS. Published by Elsevier Ltd. This is an open access article under the CC BY-NC-ND license (http:// creativecommons.org/licenses/by-nc-nd/4.0/). doi: 10.1016/S2095-3119(18)62092-1

resistance throughout the plant. The cell death factors of the HR function more as a signaling system than as a direct defense mechanism (Heath 2000). Systemic programmed cell death, a special form of programmed cell death (PCD), is often observed at distant uninoculated sites during plantmicrobe interactions and it has the same characteristics as common PCD (Alvarez et al. 1998; Deng et al. 2010). It

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reported that inoculation of Arabidopsis leaves with avirulent Pseudomonas syringae induced HR at the inoculated sites and led to a low frequency of systemic cell death in distant tissues (Alvarez et al. 1998). Reactive oxygen species (ROS), including superoxide (O2–·), hydrogen peroxide (H2O2), and the hydroxyl radical (·OH), are produced during the interaction of metabolism intermediates with oxygen (Rychter 2006). Accumulated ROS can damage functional proteins, DNA, and plasma membranes and they can reduce plant growth and survival. As a result, an effective antioxidant system, which comprises a series of antioxidant enzymes like catalase (CAT) and superoxide dismutase (SOD) and antioxidant substrates such as ascorbate, has evolved for clearing away the excess ROS (Noctor and Foyer 1998; Corpas et al. 2001). Normally, plants maintain a balance between ROS generation and elimination to maintain ROS concentrations at low levels. However, this balance can be disrupted under biotic or abiotic stress and ROS may accumulate rapidly to induce defense reactions (Prasad et al. 1994; Tsugane et al. 1999). During the plantmicrobe interactions, abundant ROS often occurred at the sites undergoing HR (Shetty et al. 2008). Relationships between ROS and PCD during interactions between plants and pathogens are often documented. After inoculation of Arabidopsis and soybean leaves with avirulent P. syringae, ROS accumulated, leading to the occurrence of HR and expression of defense genes (Bowling et al. 1994; Levine et al. 1996). Most studies have focused on the function of ROS in HR, while the role of ROS in systemic PCD has received far less study. Here we report systemic studies of the dynamic changes of ROS during the systemic PCD in wheat roots caused by PM. Our previous study showed that PM could induce systemic PCD and cell death in wheat adventitious roots (Deng et al. 2010). Waterlogging treatments resulted in the accumulation of ROS in the cortical cells, suggesting that cell death was accompanied by ROS production (Xu et al. 2013). Many other reports suggest that ROS might be involved in plant-pathogen responses (Alvarez et al. 1998; Corpas et al. 2001; Rychter 2006). We studied the roles played by ROS in PM induced root cell death by comparing the differences between two wheat cultivars that are susceptible and resistant to PM (Blumeria graminis f. sp. tritici). Wheat susceptible to PM is Huamai 8, and wheat with resistance is Shenmai 8 (Chen et al. 2002). We found that PM infections in wheat induced the degradation of root cortical cells in both varieties, but Huamai 8 had more serious injuries. We tested the hypothesis that cell death is accompanied by ROS production by detecting the production of H2O2 and O2–· at microscopic and ultra-cellular levels. We also studied the expression levels of genes controlling ROS homeostasis in wheat roots. Our results

suggested that ROS was involved in the process that PM used to induce cell death in wheat roots.

2. Materials and methods 2.1. Plant growth conditions, treatments, and sampling Seeds of wheat varieties Huamai 8 and Shenmai 8 which were harvested in field, were surface-sterilized with 0.25% (w/v) sodium hypochlorite for 3 min, rinsed thoroughly with deionized water and placed on wet filter paper in a glass Petri dish to germinate at 20°C in darkness. When the radicle was about 5 mm long (1 d after germination), germinating seeds were transferred to plastic pots (8 cm high, 9 cm diameter) containing soil. The soil was watered daily with 1/2 Hoagland’s nutrient solution to maintain field capacity (60%, w/w). The nutrient solution contained: 2 mmol L–1 Ca(NO3)2, 2.5 mmol L–1 KNO3, 0.5 mmol L–1 NH4NO3, 0.5 mmol L–1 KH2PO4, 1 mmol L–1 MgSO4, 2.5 μmol L–1 KI, 50.2 μmol L–1 H3BO3, 50 μmol L–1 MnSO4, 15 μmol L–1 ZnSO4, 0.52 μmol L–1 Na2MoO4, 0.05 μmol L–1 CuSO4, 0.053 μmol L–1 CoCl2, and 25 μmol L–1 EDTA FeNa. Plants were grown in a growth chamber that had been sterilized by potassium permanganate fumigation. Growth chamber settings were 20°C, 70% relative humidity, and a 14-h light/10 h dark photoperiod (Deng et al. 2010). When the 4th leaf appeared (about 18 d after germination), treatments were initiated. For the powdery mildew experiment, 20 plants of each of the two varieties were put into another growth chamber containing many wheat seedlings that were severely infected by B. graminis f. sp. tritici to let powdery mildew occur naturally. Twenty control plants of each of the two varieties were grown in a clean chamber under the same conditions, as the treatments. At 36 d after germination, the roots were washed out. The newborn adventitious roots, more than 8 cm in length, were used for this study. Root segments of about 2–3 mm in length, 6 cm from the root tip, were used for preparation of frozen sections, vibration sections semithin sections, and ultrathin sections. Similar root segments 4–8 cm from the root tip, were used for DNA and RNA extraction.

2.2. Anatomy of the adventitious roots Samples were collected 18 days after inoculation (DAI) from the treatments and the controls. Root segments were excised using a sharp razor blade. Samples were prepared as described by Wang et al. (2008). The root segments were prefixed in 2.5% (v/v) glutaraldehyde/0.1 mol L–1 phosphate buffer (pH 7.2) for 2 h, then rinsed three times for 15 min each with phosphate buffer (pH 7.2). They were post-fixed

LI Cheng-yang et al. Journal of Integrative Agriculture 2019, 18(9): 1961–1970

in 1% OsO4 in the same buffer for 2 h, followed by three 15-min rinses with phosphate buffer (pH 7.2). Afterwards, the samples were dehydrated through an acetone series (30, 50, 70, 90, 95, and 100%) (v/v in dd H2O) at room temperature, there was a 20-min exposure to each concentration. Then the samples were infiltrated in a graded scale of 3:1, 1:1, and 1:3 (v/v) acetone/SPI-PON 812 resin and, as the last step, in 100% (v/v) SPI-PON 812 resin (SPI Supplies, West Chester, PA, USA), for 12 h per step. Samples were embedded in SPI-PON 812 resin which was polymerized at 60°C for 24 h. Semithin sections (0.5–0.8 µm thick) were cut on an LKB 2088 Ultracut Ultramicrotome (Bromma, Sweden) and the sections were viewed using a light microscope (Nikon Eclipse 80i, Japan). For microstructural analysis, samples were prepared as described by Xu et al. (2013). Segments of roots were embedded in 5% (w/v) agarose in warm water; and cross sections, 40 µm thick, were cut with a vibrating blade microtome (Leica VT1000 S, Germany).

2.3. Detection of ROS accumulation Samples at 0, 6, 10, 12, 15, and 18 DAI were embedded using Tissue-Tek OCT 4583 compound (Sakura Finetek, USA). Transverse sections (25 µm) were made on a Leica CM1850 (Germany) freezing microtome. Unstained frozen sections were observed for auto-fluorescence in distilled H2O using a fluorescence microscope (Nikon Eclipse 80i, Japan). To analyze ROS production, we used a ROS (including H 2O 2 and O 2–·) sensitive fluorescent probe 2,7-dichlorodihydrofluorescein (H 2 DCF-DA, Sigma). This non-polar compound is converted to a membraneimpermeable polar derivative H2DCF-DA by esterases, following entry into the cell. H2DCF-DA is non-fluorescent, but is rapidly oxidized to the highly fluorescent H2DCF by intracellular ROS. Detection of ROS was performed according to the methodologies outlined in Bouranis et al. (2003). Frozen sections were incubated in H2DCF-DA (dissolved in DMSO first and then dilute with distilled H2O to 5 µmol L–1; DMSO final concentration less than 0.5%) for 1 h at 25°C first and then rinsed three times with distilled H2O. Sections were observed and photographed using a fluorescence microscope (Nikon Eclipse 80i, Japan). The excitation and emission wavelengths were 450–490 nm (blue) and 520 nm (green) respectively.

2.4. Cytochemical localization of H2O2 and O2–· Localization of H2O2 was performed according to the method described in Bestwick et al. (1997). Samples at 12, 15, and 18 DAI were incubated in freshly prepared 5 mmol L–1

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CeCl3 in 50 mmol L–1 3-(N-morpholino) propanesulphonic acid (MOPS) at pH 7.2 for 1 h at room temperature. Then the samples were fixed in sodium cacodylate buffer (pH 7.2) consisting of 1.25% (v/v) paraformaldehyde and 1.25% (v/v) glutaraldehyde for 2 h and then were washed thoroughly with sodium cacodylate buffer. Afterwards, the samples were fixed in 1% OsO4 and rinsed in buffer solution. Samples were dehydrated and embedded as described earlier. Ultrathin sections (80 nm) were prepared using an LKB 2088 Ultracut ultramicrotome (Bromma, Sweden). Sections were observed and photographed using a transmission electron microscope (Hitachi H-7650, Japan) at an accelerating voltage of 80.0 kV. Accumulation of O 2–· at the subcellular level was determined using the 3,3-diaminobenzidine (DAB)/MnCl2 method (Steinbeck et al. 1993; Romero-Puertas et al. 2004). Samples at 12, 15, and 18 DAI were incubated in freshly prepared 0.1 mol L–1 HEPES buffer (pH 7.2) containing 2.5 mmol L–1 DAB, 0.5 mmol L–1 MnCl2, and 1 mmol L–1 Na-azide for 30 min at room temperature. Samples were dehydrated and embedded as described earlier. Ultrathin sections (80 nm) were prepared using an LKB 2088 Ultracut ultramicrotome (Bromma, Sweden). Sections were observed and photographed using a transmission electron microscope (Hitachi H-7650, Japan) at an accelerating voltage of 80.0 kV.

2.5. Gene expression of MnSOD and CAT RNA from root sections at 0, 6, 10, 12, 15, and 18 DAI was extracted according to the instructions of the Total RNA Isolation Kit (Yeasen, Shanghai). The concentration of RNA was measured by NANODROP 2000 Ultraviolet and Visible Spectrophotometer (Thermo Scientific, USA). Electrophoresis was carried out on a 0.8% agarose gel to check out the integrity of the RNA and 0.8 µg RNA was used to synthesize the first-strand cDNA by reverse transcription. The quality of cDNA was analyzed via PCR (Tang et al. 2007). Finally, the gene expression of MnSOD and CAT were detected in a qPCR System (Applied Biosystems, USA). SYBR Green Real-time PCR Master Mix (Yeasen, Shanghai) was used to perform the reaction. The relative expression level of each gene was quantified with the comparative threshold cycle method, using wheat β-actin as the internal reference. PCR reactions for each of the three biological replicates were performed in duplicate. The primer sequences are shown in Appendix A.

2.6. Statistical analysis Statistical analysis was performed using the GraphPad Prism (7.00) Software. Measurement of fluorescence

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intensity was accomplished with Image J (1.37c). Results were compared using the multiple t-test and data were presented as the mean±standard deviation. Significant differences were stated where P<0.05.

3. Results 3.1. Huamai 8 and Shenmai 8 sensitivity to powdery mildew infection The leaves of Huamai 8 were covered by powdery mildew at 18 DAI (Fig. 1-A), a result different from the control (Fig. 1-B). No PM lesions were observed on leaves of Shenmai 8 (Fig. 1-E), a result similar to the control (Fig. 1-F). The adventitious roots of both varieties were reduced at 18 DAI (Fig. 1-C and G) compared to the control (Fig. 1-D and H). However, fresh weight of the Huamai 8 roots were reduced more (0.43 g) than those of Shenmai 8 (Appendix B). Inoculation of wheat leaves with wheat PM can induce cell death in wheat adventitious roots, but no

fungal structures have been observed (Deng et al. 2010).  Due to the different responses of Huamai 8 and Shenmai 8 to PM infection, we studied the root anatomy of both varieties to evaluate differences. At 18 DAI, substantial cell shape deformation and degradation of the root cortex were detected in Huamai 8 (Fig. 1-J and K, arrows). However, only the cortical cell deformation of the roots was observed in Shenmai 8 (Fig. 1-M and O), perhaps due to the resistance of this variety to PM. The root structure of the controls remained intact (Fig. 1-J, L, N, and P).

3.2. Dynamic changes of ROS accumulation in Huamai 8 and Shenmai 8 ROS can play important roles in the PCD process of plants. The cell death in roots as a result of leaf PM infection prompted verification that the ROS production had accumulated in these cells. ROS formation was indicated by green fluorescence. In Huamai 8, central cells showed weak fluorescence, but there was almost no fluorescence in the root cortical

A

B

C

D

E

F

G

H

I

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L

M

N

O

P

Fig. 1 Effect of powdery mildew fungus (PM) infection on leaves and roots of Huamai 8 and Shenmai 8. A, leaves of Huamai 8, 18 days after inoculation (DAI). B, D, F, H, controls of Huamai 8 and Shenmai 8, respectively. C, adventitious roots of Huamai 8, 18 DAI. E, leaves of Shenmai 8, 18 DAI. G, adventitious roots of Shenmai 8, 18 DAI. Vibration sections (I, J, M, and N) and semithin sections (K, L, O, and P) show the microstructural changes in the root cells of both varietiies. I and K, root cells of Huamai 8 at 18 DAI. M and O, root cells of Shenmai 8 at 18 DAI. J, L, N, and P, controls of Huamai 8 and Shenmai 8, respectively. Arrows indicate the shape deformation and degradation of cortical cells. Bar=1 mm (A–H); bar=0.1 mm (I–P).

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cells at 6 DAI (Fig.  2-A). In contrast, fluorescence was

results (Fig. 2-V). The negative control showed almost no

present in some outer cortical cells at 10 DAI, (Fig. 2-B). At

fluorescence (Fig. 2-F and L).

12 DAI, intensive fluorescence was present on most cortical cells and stele (Fig. 2-C) with the maximum fluorescence intensity in both tissues (Fig. 2-M). At 15 DAI, fluorescence was generally weaker than that at 12 DAI but many

3.3. Genes regulating the homeostasis of ROS are differently regulated in wheat roots of Huamai 8 and Shenmai 8 during PM infections

cortical cells still showed intense fluorescence (Fig. 2-D). At 18 DAI, fluorescence remained in outer cortical cells

Due to ROS are toxic motabolic products, there are genes

(Fig. 2-E). The fluorescence intensity of ROS was detected

encoding ROS scavening enzymes in plant cells that can

in the sections and we found that the strongest ROS signal

control ROS levels. ROS fluorescence results on both wheat

was at 12 DAI. There were significantly different results

varieties indicated that ROS accumulate differently in roots

(P<0.01) of ROS fluorescence signal between infected root

during PM infections. We therefore studied the expression

and the control at 12 and 15 DAI (Fig. 2-U). The results

levels of genes controlling ROS homeostasis. The expression levels of gene SOD, encoding for the

indicate a dynamic change of ROS dependent fluorescence signal during the test period. Compared with Huamai 8, the

H2O2-producing enzymes, which scavenge superoxide

fluorescence in the roots of Shenmai 8 remained weaker

anions, displayed a similar pattern of change in both

during the entire period of infection (Fig. 2-G, H, I, J, and

varieties. It was repressed from 0 to 18 DAI (Fig. 3-A and

K) and were similar to the control (Fig. 2-Q, R, S, and

B). The transcripts of SOD in both varieties were strongly

T). The fluorescence intensity of ROS showed the same

inhibited during the entire infection period except at 12 DAI

B

G

H

M

N

Fluorescent intensity

U

Huamai 8

O

30

C

D

E

I

J

K

P

Infected roots Control

20 10 0

6 12 15 18 Days after inoculation (DAI)

Q

Shenmai 8

V

30

F

L

R

Fluorescent intensity

A

S

T

** *

Infected roots Control

20 10 0

6 12 15 18 Days after inoculation (DAI)

Fig. 2 Root sections treated with H2DCF-DA for the detection of reactive oxygen species (ROS). Higher ROS levels results in higher green fluorescence intensity. A, B, C, G, and H, ROS detection in Huamai 8 at 6, 10, 12, 15, and 18 days after inoculation (DAI). Arrows indicate the enrichment of ROS. Asterisks indicate the degradation of cortical cells. M–P, controls (uninoculated roots) of Huamai 8 at 6, 12, 15, and 18 DAI. D, E, F, J, and K, ROS detection in Shenmai 8 at 6, 10, 12, 15, and 18 DAI. Arrows indicate the enrichment of ROS. Asterisks indicate the shape deformation of cortical cells. I and L, negative controls, not treated with H2DCF-DA. Q–T, controls (uninoculated roots) of Shenmai 8 at 6, 12, 15, and 18 DAI. U and V, the fluorescent intensity of ROS in the root sections of Huamai 8 and Shenmai 8, respectively. Data are the mean±SD, n=3. Results are significantly different between inoculated and uninoculated (control) roots. 0.01
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Huamai 8

**

**

0.0

* **

0.5 0.0

MnSOD

1.5 1.0

***

0.5 ***

0.0

***

*** ***

0 6 10 12 15 18 Days after inoculation (DAI)

***

***

0 6 10 12 15 18 Days after inoculation (DAI)

D

CAT

1.5 1.0

***

0 6 10 12 15 18 Days after inoculation (DAI)

C Relative expression level

***

***

Relative expression level

1.0 0.5

B

MnSOD

1.5

Relative expression level

Relative expression level

A

Control

CAT

1.5 1.0

**

** **

**

**

0.5 0.0

0

6 10 12 15 18 Days after inoculation (DAI)

Fig. 3 Expression levels of reactive oxygen species (ROS) homeostasis controlling genes in wheat roots. A and B, expression levels of superoxide dismutase (MnSOD) in both wheat varieties. C and D, expression levels of catalase (CAT) in both wheat varieties. Transcript levels relative to 0 day after inoculation (DAI) for each time point were normalized to the levels of actin. Data are the mean±SD, n=3. Results are significantly different between inoculated and uninoculated (control) roots. 0.01
in Huamai 8. Results were significantly different (P<0.01) between inoculated and uninoculated roots, indicating that ROS production was strongly inhibited in both varieties except at 12 DAI in Huamai 8.  The expression levels of CAT in Huamai 8 gradually decreased after treatment except at 18 DAI, which showed a sharp CAT increase (Fig. 3-C).

localization of H2O2 and O2-· in Huamai 8 and Shenmai 8.

Subcellular localization of H 2O 2 in root cells The

formation of H2O2 was indicated by CeCl3 precipitation. In Huamai 8, intensive precipitates were found on the cell

walls, comparing with the control at 12 DAI (Fig. 4-A and B). At 15 DAI, H2O2 accumulation remained evident in the

The expression levels of CAT were slight, but significant

inner surface of plasma membranes and tonoplasts of root

(P<0.01), repressed during the whole period of PM treatment

cell (Fig. 4-C). However, fewer precipitates were detected

in Shenmai 8 (Fig. 3-D). However, the significant difference

on the plasma membranes of the control (Fig. 4-D). In

of Shenmai 8 (P<0.01) was less than that of Huamai 8

Shenmai 8, H2O2 precipitates on plasma membranes and

(P<0.001) between inoculated and uninoculated roots

cell walls, similar to the controls, were almost and weak

at 12 and 15 DAI (Fig. 3-C and D). Owing to its strong

detected at 12 and 15 DAI (Fig. 4-G, H, I, and J), which

ROS-scavenging capacity, the down-regulation of CAT and

was in consistent with ROS fluorescence results. No H2O2

reduction of catalase might promote ROS accumulation in

precipitates were detected in the root cells of negative

the root cells.

controls (Fig. 4-E, F, K, and L). And observing almost no cellular materials, markedly different to the control at 18 DAI

3.4. Subcellular localization of H2O2 and O2–· in Huamai 8 and Shenmai 8

of root cell occurred after PM infected the plant.

Results of ROS fluorescence and the expression levels of

formation of O2-· was indicated by the precipitates. In

genes controlling ROS homeostasis on both wheat varieties

in Huamai 8 (Fig. 4-E and F), means that the degradation Subcellular localization of O 2–· in root cells The

Huamai 8, O2-· precipitates in root cells were strongly

indicated that ROS accumulate differently in roots especially

detected on plasma membrane at 12 and 15 DAI (Fig. 5-A

at 12 and 15 DAI. We therefore tested the subcellular

and C). Similarly, O2-· precipitates were only clearly

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Huamai 8

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Shenmai 8

A

B

G

H

C

D

I

J

E

F

K

L

Fig. 4 H2O2 ultracytochemical localization in Huamai 8 (A–F) and Shenmai 8 (G–L). A, C and E, root cell ultrastructure at 12, 15 and 18 days after inoculation (DAI). B, D and F, controls, uninoculated root at corresponding time. G, I and K, root cell ultrastructure at 12, 15 and 18 DAI. H, J and L, controls, uninoculated root at corresponding time. Arrows indicate H2O2 precipitates. E, F, K, and L, negative control, not treated with CeCl3. Bar=2 µm (B, D, E, F, H, J, and L); bar=1 µm (A, C, G, I, and K). Huamai 8

Shenmai 8

A

B

G

H

C

D

I

J

E

F

K

L

-

Fig. 5 O2 · ultracytochemical localization in Huamai 8 (A–F) and Shenmai 8 (G–L). A and C, root cell ultrastructure at 12, 15, and 18 days after inoculation (DAI). B, D and F, control, uninoculated root at corresponding time. G, – I and K, root cell ultrastructure at 12, 15 and 18 DAI. Arrows indicate O 2 · precipitates. E, F, K, and L, negative control, not treated with either DAB or MnCl 2. Bar=2 µm (C, E, F, G, K, and L); bar=1 µm (A, H, I, and J); bar= 0.5 µm (B and D).

detected in root cells at 12 DAI in Shemai 8 (Fig. 5-G). Little O2-· precipitates were detected on plasma membrane in root cell of Shenmai 8 at 15 DAI, which may be due to the high reactivity and short half-life of O2-· in cells. While in uninoculated wheat roots of both varieties, almost no O2-· precipitates were observed in root cells (Fig. 5-B, D, H, and J). No O2-· precipitates were detected in root cells of negative controls (Fig. 5-E, F, K, and L).

4. Discussion 4.1. PM in the leaves of wheat induced the cell death of wheat roots PM lesions in the leaves of Huamai 8 but not in Shenmai 8. This is probably related to their different levels of pathogen resistance capacities (Fig. 1-A–H).  Their genetic

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background both from Yangmai 158. Huamai 8 was successfully selected by Huazhong Agricultural University in 1995, which is resistant to waterlogging, cold, and lodging. Shenmai 8 is obtained through traditional breeding in 2004 (Zhang et al. 2004). Previous results suggested that, even without pathogen spread, the infection could induce systemic PCD in adventitious roots (Deng et al. 2010). In this study, cell shape deformation and degradation of the root cortical cells occurred at 18 DAI in Huamai 8, and to a lesser extent, in Shenmai 8 (Fig. 1-I–P). Although the two varieties differ greatly in their pathogen resistance, they both displayed root cell death when the leaves were infected by PM, indicating that similar signaling events might be involved in both varieties. This cell death appears to be necrotic cell death, different from developmentally controlled cell death, one form of HR-related non-autolytic PCD seen under biotic stress (Doorn 2011; Doorn et al. 2011; Olveracarrillo et al. 2015). In general, resistant cultivars have strong HR-PCD in leaves than that of susceptible cultivars because the HRPCD reaction of the cells caused by biotic stress in leaves can cause the cells around the infection site to rapidly die, thereby limiting the spread of the pathogen (Lam 2004; Angel et al. 2018). However, the case of the root in this study is opposite. This cell death more likely to be a systemic PCD due to diseased plants. It seems to be caused by changes in the homeostasis of the plants after the leaves have been diseased (Zhou et al. 2008). This PCD reaction is different to the usual HR-PCD in the time of occurrence and synchronization. It also differs from HR-PCD in the cause and manifestation (Rogers 2010). It seems to be the chain reaction caused by the disease of the plant and it is indeed induced under biotic stress.

4.2. ROS was involved in the cell death of wheat roots After inoculation of Arabidopsis leaves with avirulent P. syringae, a low frequency of systemic cell death in distant tissues occurred and that ROS were involved in the process (Alvarez et al. 1998). In this study, we also found that the ROS fluorescence intensity was the highest at 12 DAI in Huamai 8 and there were dynamic overall changes of ROS fluorescence (Fig. 2-U). However, slight ROS fluorescence was observed in Shenmai 8, which has resistance to powdery mildew, during the entire test period (Fig. 2-V). ROS fluorescence only revealed ROS dynamic changes at the cellular or tissue levels. Because of sensitivity limitations, very low ROS levels in the cells might not be detected. To clarify the involvement of ROS during PM-induced ROS production and its significance in cell death, we analyzed the subcellular localization of H2O2 and O2-· production. In plant cells, the antioxidant system composition of SOD and CAT is important in maintaining ROS homeostasis. First, O2-· is

quickly catalyzed to H2O2 by SOD, then the accumulated H2O2 is eliminated by CAT (Willekens et al. 1997). Plants normally maintain a balance between ROS generation and elimination so ROS concentrations in cells are at a low level. However, accumulation of ROS such as O2-· and H2O2 increases quickly under stress and the plant then faces the threat of oxidative damage. Therefore, it is crucial to maintain a balance of antioxidant enzymes in cells for suppressing toxic ROS levels and allowing plant survival in adverse conditions (Apel and Hirt 2004). These results indicated that ROS was involved in the occurrence of cell death after inoculation of leaves with PM.

4.3. ROS was cleared at high levels of MnSOD and CAT gene expression At 12 DAI, the high level of MnSOD gene expression and low expression of CAT led to accumulation of abundant H2O2 in Huamai 8 (Figs. 4-A and 5-A). Afterwards, with the increasing CAT gene expression (Fig. 3-C), H 2O2 accumulation in cells gradually decreased (Figs. 4-C and 5-C). CAT gene expression in tobacco cells undergoing HR due to microbial attack was suppressed and an ROS burst occurred (Dorey et al. 1998). Compared with Huamai 8, slight accumulation of ROS was observed in Shenmai 8 at 12 and 15 DAI (Figs. 4-G, I and 5-G, I). The high levels of CAT gene expression might be responsible for the small amount of ROS accumulation in Shenmai 8 (Fig. 3-D). In cells, O2–· has a very short half-life and can be catalyzed quickly by MnSOD. H2O2 is the most stable of ROS and it can rapidly diffuse across cell membranes to adjacent cells (Gechev et al. 2006). As a result, a high level of CAT gene expression after infection might contribute to the removal of excess H2O2 and avoidance of the oxidative damage caused by H2O2. Similar results have been reported in other studies. Our previous study showed that the two genes encoding NADPH oxidase and Mn-SOD involved in ROS production were up-regulated at the initial stage of waterlogging. Expression of the genes encoding CAT and MT, involved in ROS detoxification, was strongly repressed at this stage (Xu et al. 2013). Yamauchi et al. (2011) observed that NADPH oxidase was up-regulated, while MT was down-regulated in maize cortical cells. These genes control ROS homeostasis and their different expressions likely resulted in ROS accumulation in the maize root cortical cells, which triggered the PCD process (Yamauchi et al. 2011). Transgenic tomato plants overexpressing a bacterial catalase were more tolerant than wild-type plants to oxidative damage caused by paraquat, drought, or chilling stress (Mohamed et al. 2003). However, the detailed mechanism leading to this cell death is unclear. We are also unable to clarify the reason why ROS

LI Cheng-yang et al. Journal of Integrative Agriculture 2019, 18(9): 1961–1970

homeostasis was damaged in wheat roots after inoculation of leaves with PM. In recent studies, PmU region related genes were associated with resistance to wheat PM (Zhang et al. 2018). Meanwhile, a resistance gene PmTm4 has been identified (Xie et al. 2017). The presence of HR in plants was accompanied by an accumulation of salicylic acid (SA) and an increase in ROS levels (Kovács et al. 2016). Excessive accumulation of SA induces cell death in Arabidopsis leaves (Chaouch et al. 2010). SA plays an important role in the immune response of plants and the activation of plant defense responses, especially when the plant is infected with a pathogen (Durner et al. 1997). SA has been described as a catalase inhibitor and selective degradation of peroxisomal catalase is reported to be the major mechanism allowing ROS accumulation to trigger autophagic programmed cell death in mouse cells (Yu et al. 2006). We speculate that other long distance signals, such as SA, may suppress the expression of antioxidant enzyme genes, leading to the accumulation of ROS and the occurrence of systemic PCD. Further studies on these mechanisms may provide greater insight into these complex phenomena.

5. Conclusion Based on our results, we proposed a model for wheat roots response to biotic stress. ROS homeostasis was compromised leading to the accumulation of ROS in root cells after inoculation of wheat leaves with PM. The combination of ROS accumulation and gene expression of antioxidant enzymes indicated that the low expression level of genes encoding antioxidant enzymes in root cells lead to the excess accumulation of ROS. At the same time, cell death and degradation events began in the root cortical cells. ROS was involved in the cell death during the days after inoculation.

Acknowledgements This work was supported by the National Nature Science Foundation of China (31071347 and 31171469). We thank Prof. Long Hong of Tianjin Agricultural University, China for providing the frozen section technology; Dr. Cao Jianbo of Huazhong Agricultural University for providing the transmission electron microscope technology and Zhang Shouzhong of Nanjing Agricultural University for providing the seeds of Shenmai 8. We would like to thank LetPub (www.letpub.com) for providing linguistic assistance during the preparation of this manuscript. Appendices associated with this paper can be available on http://www.ChinaAgriSci.com/V2/En/appendix.htm

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