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inhibiting bacteria
Accepted Manuscript Differential oxidative and antioxidative response of duckweed Lemna minor toward plant growth promoting/inhibiting bacteria Hidehiro Ishizawa, Masashi Kuroda, Masaaki Morikawa, Michihiko Ike PII:
S0981-9428(17)30257-7
DOI:
10.1016/j.plaphy.2017.08.006
Reference:
PLAPHY 4960
To appear in:
Plant Physiology and Biochemistry
Received Date: 7 June 2017 Revised Date:
7 August 2017
Accepted Date: 8 August 2017
Please cite this article as: H. Ishizawa, M. Kuroda, M. Morikawa, M. Ike, Differential oxidative and antioxidative response of duckweed Lemna minor toward plant growth promoting/inhibiting bacteria, Plant Physiology et Biochemistry (2017), doi: 10.1016/j.plaphy.2017.08.006. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
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Differential oxidative and antioxidative response of duckweed Lemna minor
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toward plant growth promoting/inhibiting bacteria
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Hidehiro Ishizawa1, Masashi Kuroda1, Masaaki Morikawa2, and Michihiko Ike1*
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Engineering, Osaka University, 2-1 Yamadaoka, Suita, Osaka 565-0871, Japan
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Division of Sustainable Energy and Environmental Engineering, Graduate School of
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Division of Biosphere Science, Graduate School of Environmental Science, Hokkaido
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University, N10-W5, Kita-ku, Sapporo 060-0810, Japan
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Corresponding author
Phone: +81-(0)6-6879-7674; fax:+81-(0)6-6879-7675
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Email:
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Hidehiro Ishizawa: [email protected]
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Masashi Kuroda: [email protected]
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Masaaki Morikawa: [email protected]
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Michihiko Ike: [email protected]
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Highlights 1
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・Growth inhibiting bacteria induced higher oxidative stress levels of duckweed than growth-promoting bacteria.
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for the effects on duckweed oxidative stress level.
・Growth promoting bacteria also induced equal or higher oxidative stress relative to aseptic control.
・Bacteria can be a significant cause of oxidative stress similar to the other
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・Two plant growth-promoting and two growth-inhibiting bacterial strains were tested
environmental factors.
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Abstract
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Bacteria colonizing the plant rhizosphere are believed to positively or negatively
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affect the host plant productivity. This feature has inspired researchers to engineer such 2
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interactions to enhance crop production. However, it remains to be elucidated whether
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rhizobacteria influences plant oxidative stress vis-a-vis other environmental stressors,
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and whether such influence is associated with their growth promoting/inhibiting ability.
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In this study, two plant growth-promoting bacteria (PGPB) and two plant
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growth-inhibiting bacteria (PGIB) were separately inoculated into axenic duckweed
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(Lemna minor) culture under laboratory conditions for 4 and 8 days in order to
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investigate their effects on plant oxidative stress and antioxidant activities. As
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previously characterized, the inoculation of PGPB and PGIB strains accelerated and
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reduced the growth of L. minor, respectively. After 4 and 8 days of cultivation,
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compared to the PGPB strains, the PGIB strains induced larger amounts of O2•−, H2O2,
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and malondialdehyde (MDA) in duckweed, although all bacterial strains consistently
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increased O2•− content by two times more than that in the aseptic control plants.
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Activities of five antioxidant enzymes were also elevated by the inoculation of PGIB,
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confirming the severe oxidative stress condition in plants. These results suggest that the
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surface attached bacteria affect differently on host oxidative stress and its response,
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which degree correlates negatively to their effects on plant growth.
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Keywords: Duckweed, Plant growth-promoting bacteria, plant growth-inhibiting
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bacteria, Oxidative stress, Reactive oxygen species, Plant-microbe interactions
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Introduction
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Plants experience a variety of environmental factors, and not all are ideal for plant
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growth (Boyer, 1982). Most of the factors, such as low temperature, salinity, ultraviolet
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radiation, and pathogen inhibit the growth of plants, commonly by inducing oxidative 3
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stress through the accumulation of reactive oxygen species (ROS). Although ROS are
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byproducts of normal cellular activities in the mitochondria, chloroplast, and
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peroxisome, they are capable of causing damage to plant cellular lipids, proteins, and
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DNA through radical reactions when present in excess. On the other hand, plants have
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developed well-tuned antioxidant systems to defend themselves from ROS, and these
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have been extensively reviewed (Mittler, 2002; You and Chan, 2015). However, it is
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also reported that the generation of ROS often exceeds the plants’ antioxidant capacity
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and causes significant loss of the plant biomass and yield even under normal
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environmental conditions (Apel and Hirt, 2004). Up-regulation of antioxidants itself
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may inhibit growth through cross-talk between developmental and stress-response
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networks (Cabello et al., 2014). Therefore, maintaining ROS and antioxidants at low
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levels is essential to achieve enhanced productivity.
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Recently, it became known that bacterial communities present in the plant
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rhizosphere are one of the factors impacting plant productivity akin to the
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environmental factors (Anderson and Habiger et al., 2012). Extensive research in this
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field has demonstrated the general occurrence of plant growth-promoting bacteria
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(PGPB) and plant growth-inhibiting bacteria (PGIB) or deleterious rhizobacteria (DRB)
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poses beneficial and deleterious effects on the host growth (Probenza et al., 1996;
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Suslow and Schroth, 1981). Our original research using duckweed Lemna minor as a
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model plant showed that both promotive and inhibitory effects by the co-existing
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bacterial community can be caused by complex and interactive influences of PGPB and
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PGIB existing in the root and frond zone of duckweed (Ishizawa et al., 2017). Therefore,
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to establish new approaches to improve crop productivity utilizing bacteria, it is critical
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that interactions between plants and bacteria, especially for PGPB and PGIB, are well
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understood. It is likely that oxidative stress plays a role in determining the beneficial and
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harmful effects of rhizobacteria on the plant, considering that oxidative stress caused by
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environmental factors also affects plant growth (Apel and Hirt, 2004). Therefore,
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understanding plant–bacterial interactions from the viewpoint of oxidative stress in
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plants may offer clues to elucidate the mechanisms leading to promotive/inhibitory
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effects on plant growth involving rhizobacteria. Such knowledge is helpful in
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developing techniques/strategies to properly regulate the rhizobacterial community to
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improve plant growth. However, studies on the relationship between plant oxidative
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stress and coexisting bacteria have been scarce. The aim of this study is to examine how
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PGPB and PGIB affect oxidative stress levels of the host plant and its response. Toward
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this, sterile L. minor was co-cultivated with previously isolated PGPB and PGIB strains
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under laboratory conditions, and the changes in plant ROS and other stress associated
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indicators in duckweed were monitored.
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Materials and Methods
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Plant and bacterial strains
Duckweed (Lemna minor, RDSC clone 5512) plants collected from a small pond
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in the botanical garden of Hokkaido University (Sapporo, Japan) were used in the
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experiments. The plants were sterilized by washing with 0.5% sodium hypochlorite for
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7 min and followed by washing twice with sterilized water. The sterilized plants were
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successively cultured in flasks containing modified Hoagland medium (36.1 mg/L
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KNO3, 293 mg L-1 K2SO4, 3.87 mg L-1 NaH2PO4, 103 mg L-1 MgSO4·7H2O, 147 mg L-1
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CaCl2·H2O, 3.33 mg L-1 FeSO4·7H2O, 0.95 mg L-1 H3BO3, 0.39 mg L-1 MnCl2·4H2O, 5
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0.03 mg L-1 CuSO4·5H2O, 0.08 mg L-1 ZnSO4·7H2O, 0.254 mg L-1 H2MoO4·4H2O, and
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5 mg L-1 EDTA·2Na; pH 7.0) and incubated at 28°C, with a light intensity of 80 µmol
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m-2s-1 and a photoperiod of 16 h/8 h day/night cycle. Four bacterial strains (Aquitalea magnusonii H3, Acinetobacter septicus M3,
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Asticcacaulis exentricus M6, Pseudomonas otitidis M12) isolated from the same
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duckweed strain were used for the study. The strains were previously characterized in
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terms of their effects on duckweed growth when co-cultivated with sterile L. minor:
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strains H3 and M12 were promotive (PGPB), while M3 and M6 were inhibitory (PGIB)
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(Ishizawa et al., 2017). These PGPB or PGIB strains were cultivated by inoculating a
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loop of bacterial colony into 20 mL of liquid LB medium and shaking the tube at 120
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rpm at 28°C until the culture reached the late exponential phase. Cells were then
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harvested by centrifugation (10,000 × g, 4°C, 10 min), washed twice with sterilized
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Hoagland medium before using them in our experiments.
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Experimental design
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Sterile L. minor plants were co-cultivated with each one of the four bacterial
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strains under similar light and nutrient conditions as mentioned above. Attachment of
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bacterial strains to L. minor was enabled by growing it along with a suspension of
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bacterial cells (optical density at 600 nm = 0.1) added to the sterilized Hoagland
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medium and maintained for 24 h prior to the experiment. Then, 10 fronds of L. minor
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with the attached bacteria were transferred to 60 mL of fresh bacteria-free medium and
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co-cultivated. During co-cultivation, the medium was replenished at 48 h intervals.
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After 4 and 8 days of cultivation, the plants were harvested and subjected to the
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analyses of ROS and other stress associated indicators.
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Plant growth evaluation During the growth period, the number of L. minor fronds was periodically
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counted and recorded for evaluating the effect of PGPB/PGIB strains on plant growth.
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Relative growth rate (RGR, d-1) was calculated as (ln FNt – ln 10)/t, where FNt is the
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frond number of L. minor on day t (4 or 8). In addition, fresh weight (FW) and dry
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weight (DW, 80°C for 24 h) of plants were measured at the end of 8-d growth period.
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Estimation of the amount of bacteria attached on duckweed
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The amount of bacterial cells that attached to plants was estimated as the number
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of colony forming units (CFU) per gram fresh weight of the plants. At the end of each 4
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and 8 day growth periods, 20 mg of plants were washed twice with 20 mL of sterile
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Hoagland medium and homogenized in 5 mg L-1 tripolyphosphate (TPP) using a
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BioMasher II (Nippi, Tokyo, Japan). The homogenates were spread onto solid 1:10
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diluted LB medium containing 1.5% agar. Agar plates were incubated at 28°C for 3
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days and the number of bacterial colonies were counted.
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Determination of chlorophyll content
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Total plant chlorophyll content was determined spectrophotometrically (UV-1850,
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Shimazu, Kyoto, Japan). Pigment extraction was performed by soaking 30 mg of plants
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in 3 mL of methanol for 90 min in the dark. Chlorophyll content per milligram fresh
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weight was calculated using absorbance at 650 nm (A650) and 665 nm (A665) and
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applying the formula (Grimme and Boardman, 1972).
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Chl a + b = 4.0 × A665 + 25.5 × A650 7
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Estimation of ROS Plant superoxide anion (O2•−) level was estimated by measuring the reduction of
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nitroblue tetrazolium (NBT) according to the method suggested by Doke (1983). Thirty
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milligrams of fresh plant samples were immersed in 1.5 ml of a mixture containing 10
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mM sodium phosphate buffer (pH 7.8), 0.05% NBT and 10 mM NaN3 in a test tube,
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followed by incubation at 23°C for 60 min. Following this, 1.0 mL of the reaction
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mixture was incubated at 85°C for 15 min. This reaction was stopped by cooling the
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solution with ice and absorbance at 580 nm (A580) was measured to assay the reduction
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of NBT. The level of O2•− was expressed as the increase in A580 per hour per gram fresh
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weight.
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Hydrogen peroxide (H2O2) content was measured according to Velikova et al.
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(2000). Thirty milligrams of plant material were homogenized in 1.5 mL test tube with
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1.0 mL of ice-cold 0.1% trichloroacetic acid (TCA) using a high-power homogenizer
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(ASG50. AS ONE, Aichi, Japan) at 6,000 rpm for 30 s. Homogenates were then
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centrifuged (10,000 × g, 4°C, 15 min), and 0.5 mL of supernatant was mixed with 0.5
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mL of 10 mM potassium phosphate buffer (pH 7.0) and 1.0 mL of 1 M KI. The mixture
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was incubated in the dark at 23°C for 60 min, and the absorbance at 390 nm was
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measured. The content of H2O2 was calculated using a standard curve plotted with
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known concentrations of H2O2.
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Determination of malondialdehyde (MDA) content
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The level of lipid peroxidation was estimated indirectly by measuring the content
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of MDA, a by-product of lipid peroxidation (Heath and Packer, 1968). Thirty 8
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milligrams of plant material were homogenized in 1.5 mL test tube containing 0.5%
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2-thiobarbituric acid in 1 mL of 20% TCA using a high-power homogenizer at 6,000
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rpm for 30 s. Homogenates were then transferred to centrifuge tubes and incubated at
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95°C for 15 min. The reaction was stopped by placing the tubes on ice, and followed by
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centrifugation (10,000 × g, 2°C, 10 min). The absorbance of the supernatant was
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measured at 532 nm. Absorbance at 600 nm was also measured to account for
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non-specific turbidity or background. The content of MDA was calculated using an
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extinction coefficient of 155 mM-1 cm-1.
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Antioxidant enzyme assays
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Catalase (CAT, EC 1.11.1.6) activity was determined by measuring the change in
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absorbance at 240 nm which indicates decomposition of H2O2 (e = 40 mM-1cm-1)
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according to the method of Aebi (1984). The reaction mixture consisted of 900 µL of 50
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mM potassium phosphate buffer (pH 7.0), 50 µL of 300 mM H2O2, and 50 µL of
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enzyme extract. The reaction was started by adding H2O2.
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Ascorbate peroxidase (APX, EC 1.11.1.11) activity was assayed by measuring the
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change in absorbance at 290 nm that accompanied the consumption of ascorbic acid (e
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=2.8 mM-1cm-1) as described in Nakano and Asada (1981). The reaction mixture
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contained 800 µL of 50 mM potassium phosphate buffer (pH 7.0), 0.1 mM EDTA, 0.5
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mM L-ascorbic acid, 50 µL of 100 mM H2O2, and 150 µL of enzyme extract.
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For guaiacol peroxidase (GPX, EC 1.11.1.7) activity determination, increase in
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absorbance at 470 nm indicating the formation of tetraguaiacol (e = 26.6 mM-1cm-1) was
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measured as previously described (Horvat et al., 2007). The reaction mixture consisted
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of 900 µL potassium phosphate buffer (pH = 6.5), 15 mM guaiacol, 50 µL of 300 mM 9
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H2O2, and 50 µL of enzyme extract. Glutathione reductase (GR, EC 1.8.1.7) activity was determined using the method
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described by Smith et al. (1988) by measuring the increase in absorbance at 412 nm that
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corresponds to the reduction of 5,5’-dithiobis-2-nitrobenzoic acid (DTNB) to
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2-nitro-5-thiobenzoic acid (TNB, e =13.6 mM-1cm-1). The reaction mixture contained
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500 µL of 200 mM potassium phosphate buffer (pH 7.5), 1 mM EDTA, 250 µL of 3
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mM DTNB in 10 mM potassium phosphate buffer (pH 7.5), 100 µL of ultra-pure water,
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50 µL of 2 mM NADPH, 50 µL of 20 mM glutathione disulfide (GSSG), and 50 µL of
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enzyme extract. The reaction was initiated by adding GSSG to the mixture.
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Superoxide dismutase (SOD, EC 1.15.1.1) was assayed based on the inhibition of
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NBT’s photochemical reduction by the decomposition of O2•− (Beauchamp and
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Fridovich, 1971). The reaction mixture consisted of 3 mL of 50 mM sodium phosphate
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buffer (pH 7.8), 13 mM L-methionine, 75 µM NBT, 0.1 mM EDTA, 300 µL of 24 µM
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riboflavin, and 0 – 200 µL of enzyme extract. The total volume of the mixture was
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adjusted to 3.5 mL by adding 50 mM sodium phosphate buffer (pH 7.8). The mixture
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was illuminated at a light intensity of 70 µmol m-2s-1 for 10 min, and the absorbance at
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560 nm was measured immediately to evaluate the photochemical reduction of NBT.
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One unit of SOD activity was defined as the amount of enzyme required to cause a 50%
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inhibition of the NBT photochemical reduction.
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The activity of all the above antioxidant enzymes was expressed as unit mg-1
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protein. For protein extraction, 50 mg of fresh plants in 1.5 mL test tube were
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homogenized with 1 ml of ice-cold 50 mM potassium phosphate buffer (pH 7.0)
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containing 1% (w/v) polyvinylpolypyrrolidone. The homogenate was then centrifuged
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(20,000 × g, 2°C, 20 min) and the supernatant was used for the enzyme assay. Soluble 10
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protein content of the supernatant was determined using the method of Bradford (1976)
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with bovine serum albumin as a standard.
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Statistical analysis
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All assays were performed with 3 biological replicates. Data was analyzed by
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one-way analysis of variance (ANOVA), and where it was significant, Duncan’s
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multiple-range test was performed to separate the means. Significance at p < 0.05 was
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applied to all analyses. Statistical analyses were performed in R software v3.2.3
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(http://www.r-project.org).
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Results
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Effects of bacterial inoculation on the growth and chlorophyll content of duckweed
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In this study, two PGPB strains (H3, M12) and two PGIB strains (M3, M6) were
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individually co-cultivated with sterile L. minor. The amounts of bacteria colonized on
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the plants are shown in Figure 1. Strain H3 had the highest CFU both after 4 and 8 days
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of cultivations. Although there was no carbon source to support bacterial growth in the
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medium, the CFU counts per plant fresh weight were maintained throughout the
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cultivation period, suggesting that these strains have stably attached to their host and
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established. Furthermore, as to strain M3, a notably higher number of CFU was
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observed after 8 days than at 4 days.
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Table 1 summarizes the growth indices of L. minor. In the experiment, RGR of
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control plants (0.354 for 8-day cultivation) was at the same level as reported by Ziegler
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et al. (2014) which used the optimized culture condition for duckweeds. Similar to the
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previous study (Ishizawa et al., 2017), PGPB strain H3 and M12 accelerated L. minor 11
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growth, while PGIB strains M3 and M6 significantly delayed growth compared to
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aseptic control, in terms of the frond number on day 4 and 8. One notable difference is
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that the growth-promoting effect of strain M12 was smaller than reported previously.
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We believe this might be due to the higher growth rate of control plants owing to the
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difference in nutrient conditions.
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Plant chlorophyll content did not show any significant differences among the
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plants co-cultivated with the four bacterial strains (Figure 2), indicating the little effects
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of PGPB and PGIB on the pigment synthesis and/or destruction of L. minor. It should
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be noted that plant had lower levels of chlorophyll content after day 8 than day 4 for all
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the tests, suggesting some change in duckweed morphology or nutritional status have
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occurred in the later part of cultivation period, possibly due to the high plant density
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and/or formation of ethylene (Farber and Kandeler, 1990).
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Plant ROS levels
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We observed significant changes in the superoxide anion (O2•−) and hydrogen
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peroxide (H2O2) levels of L. minor, depending on the bacterial strains used for
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co-cultivation. These free radicals serve as the major forms of ROS in plants. As shown
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in Figure 3a, all bacterial strains significantly elevated O2•− content of the plants
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compared to the aseptic control. We have confirmed that simple introduction of bacterial
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cells (105~106 cfu) to the test reagent of O2•− did not affect the result of the assay (data
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not shown). Among the bacterial strains, PGIB strain M6, which had the strongest
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growth-inhibitory effects in this experiment, caused the largest increase in the O2•−
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content of L. minor. Another PGIB strain M3 also tended to induce more than PGPB
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strains, though the trend is not statistically significant.
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As for H2O2 content, significantly higher value compared to the control were
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recorded for the plants cultivated with PGIB strain M6 at days 4 and 8 as well as strain
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M3 at day 4 (Figure 3b). Although not significant, plants co-cultivated with PGPB
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strains H3 and M12 also showed higher H2O2 content than control plants, especially in
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the later part of growth period (day 8).
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Lipid peroxidation
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The level of duckweed lipid peroxidation was determined by estimating the
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amount of MDA accumulation in plant tissues (Figure 4) as an indicator of cellular
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damage caused by ROS. Though the difference could not be confirmed due to large
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variation in results for both on day 4 and 8, PGPB strains increased MDA content in the
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range of 2.6 - 16.3%, while PGIB strains increased MDA in the range of 11.4 - 39.7%
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compared to the control plants. Thus, the general trend of the changes in MDA content
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during the cultivation period was similar to that of O2•− and H2O2 content, showing that
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the degree of lipid peroxidation is more or less correlated to ROS production.
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Specifically, it is worth noting that the significant increase of MDA content observed for
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the plants co-cultivated with M6 on day 4 coincided with the highest accumulation of
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O2•− (Figure 3a).
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Antioxidant enzyme activities
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The effects of bacterial strains on the activities of five important antioxidative
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enzymes of the plants were assayed. Although we could not exclude bacterial-originated
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enzymes in these assays, their effects on the results are considered to be limited because
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the amount of bacterial protein extracted in the assays would be extremely smaller than 13
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plant proteins. Similar to the ROS and lipid peroxidation, the higher values were
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recorded for the plants cultivated with PGIB strains M3 and M6 in all enzyme studies
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except CAT (Figure 5). Since plants are known to up-regulate antioxidant enzymes
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responding to adverse environments or ROS itself (Babu et al., 2003), the result would
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reflect the plant activation of antioxidant enzymes by perceiving the higher ROS content.
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In addition, we observed that activities of each enzyme were quite different between the
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plants collected after 4 and 8 days. Because the values were expressed based on the
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amount of total extracted protein, it suggests different protein allocation in plants
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depending on the growth stage.
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Discussion
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Despite a number of publications on growth promoting and growth inhibiting
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effects of plant rhizobacteria, their effects, particularly on plant oxidative stress
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response, are poorly understood. This study highlights the effects of bacterial strains on
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plant oxidative stress and antioxidant activities using duckweed L. minor, which is a
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useful model plant for investigating plant-microbe interactions (Appenroth et al., 2016).
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As previously characterized (Ishizawa et al., 2017), the bacterial strains used in
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this study either promoted growth or inhibited the growth of duckweed (Table 1), while
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the influence on chlorophyll content was limited (Figure 2). Interestingly, both PGPB
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and PGIB caused significantly different oxidative stress levels in L. minor, as seen by
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measuring the ROS content, MDA content, and antioxidant enzymatic activities (Figure
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3-5). In our experiments, the numbers of living bacterial cells colonized on the plants,
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which might be a kind of bacterial dose to the plant, were estimated as CFU counts. The
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CFU counts showed a large variation up to about ten times between treatments (Figure
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1). However, their effect on the plant oxidative stress levels could not be correlated to
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the CFU counts. Therefore, it is likely that the observed changes in plant oxidative
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stress levels depend on the nature and type of bacterial strains introduced rather than the
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amount of bacterial cells attached.
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Among four bacterial strains used in this study, two PGPB strains, Aquitalea
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magnusonii H3 and Pseudomonas otitidis M12, showed relatively moderate influence
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on plant oxidative stress. However, these strains led to very high levels of O2•− content
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in inoculated plants relative to control plants, as well as PGIB strains (Figure 3a). It is
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likely that bacteria generally stimulated the plant O2•− generation, though the degree was
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different among bacterial strains. Plants possess the ability to generate O2•− as an
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immune reaction when they recognize microorganisms through perception of
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microbe-associated molecular patterns (MAMPs) such as presence of chitin or flagellins
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(Smith et al., 2015). Other studies have shown that plants generate O2•− from the activity
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of NADPH oxidase during the establishment of plant–bacterial symbiosis as signal
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molecules (Nanda et al., 2010). This suggests that the observed phenomena were a
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result of spontaneous generation of O2•− by plants in response to bacterial colonization,
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which did not occur in the aseptic control plants.
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On the other hand, there are some recent studies of PGPB associated with
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terrestrial plants that act to alleviate plant oxidative stress under harsh stress conditions
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through the regulation of antioxidant activities (Habib et al., 2016; Islam et al., 2014). In
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our study, we used favorable growth conditions for L. minor. As a result, the role of
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PGPB strains in alleviating plant oxidative stress compared to the aseptic control was
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not detected, suggesting the presence of growth-promoting mechanisms of PGPB other
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than through stress alleviation. However, in a few studies, other duckweed PGPB strains 15
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were reported to enhance the growth of duckweed even in the presence of toxic phenol
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(Yamaga et al., 2010) and chromium (Tang et al., 2015). Considering all these studies,
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the possibility that duckweed PGPB strains play a role in alleviating plant stress when
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plants are exposed to stressful environments cannot be ruled out. Therefore, to gain
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more insights on beneficial plant–microbe interactions additional studies are required.
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Compared to the PGPB strains, two PGIB strains, Acinetobacter ursingii M3
367
and Asticcacaulis excentricus M6, caused considerably higher oxidative stress levels in
368
L. minor (Figure 3-5). Although there are several studies about the mechanism of plant
369
growth promotion by PGPB such as nutrient supply and modulation of plant hormone
370
(Glick, 2012), very little is known about how PGIB negatively affect plant growth and
371
productivity. Thus far, hydrogen cyanide production (Astrom, 1991) and ammonia
372
production (Weise et al., 2013) were proposed as possible mechanisms of existing PGIB
373
strains. In this study, we show that PGIB strains induce higher levels of oxidative stress
374
in the host plant than by the other bacterial strains, which explain at least some of the
375
growth-inhibitory mechanisms employed by PGIB. The knowledge that PGIB induce a
376
considerably high levels of plant oxidative stress may provide clues toward establishing
377
strategies for crops to defend PGIB. For example, engineering plants for elevated
378
abiotic stress tolerance or introducing outcompeting rhizobacterial strains with less
379
influence on plant stress might be effective for managing the adverse effects of PGIB.
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Duckweeds have been used in plant stress research and for chemical toxicity tests
381
for a long time (Ziegler et al., 2016), and their responses toward environmental stress
382
factors have been extensively studied. Thus far, a variety of heavy metals (Parlak et al.,
383
2012; Tkalec et al., 2008; Duman et al., 2010), salinity (Cheng, 2011), ammonium
384
(Wang et al., 2016), and chemical compounds (Obermeier et al., 2015) have been shown 16
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to induce a dose-dependent response to oxidative stress, while the degree and type of
386
activated antioxidants seemed to vary and depend highly on the culture conditions
387
and/or duckweed strains used. In this study, similar responses of duckweed to the
388
above-mentioned abiotic stressors are being reported for the first time for
389
plant-associated bacteria, particularly in suppressing the growth of the host plant.
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Table 2 summarizes the oxidative and antioxidative responses induced by PGIB
391
strains, in comparison to Panda (2006) who investigated the response of L. minor fronds
392
to copper (Cu2+) exposure. Copper is one of the most typical and well-studied stress
393
factors for duckweed, which induces severe oxidative stress in the plant at higher
394
concentrations (Panda et al., 2008; Babu et al., 2003; Razinger et al., 2007). As shown
395
in Table 2, increase in O2•− content and GPX caused by PGIB were much higher than
396
that caused by copper, although the responses to H2O2 content was comparable to
397
moderate concentrations of Cu2+. Upregulation of antioxidant enzymes induced by
398
PGIB were also comparable to that by copper, except for GPX. Even though we
399
observed a considerable increase in MDA content for plants co-cultivated with PGIB,
400
the level of lipid peroxidation did not seem to be as high as that caused by Cu2+.
401
Considering these information, we propose that the effects on the plant oxidative stress
402
by PGIB strains are generally equivalent to the moderate dose of environmental
403
contaminants such as copper, which should be certainly a focus of future research.
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In summary, this is the first study to highlight the different effects of PGPB and
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PGIB strains on the oxidative stress and antioxidant enzyme activities of L. minor. Two
406
PGIB strains were found to increase plant oxidative stress and antioxidant enzyme
407
activities similar to abiotic stress factors reported previously. On the other hand, PGPB
408
strains were less stressful, though they also caused certain levels of oxidative stress on 17
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the plant when compared to the control plant. Although the detailed mechanisms are
410
still unknown, oxidative stress and its response may play important roles in growth
411
inhibition caused by PGIB. In addition, a series of oxidative stress assays performed in
412
this study can serve as useful tools to evaluate beneficial/deleterious effects of bacteria
413
on duckweed productivity and possibly extend to studies on other plant species.
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Lemna aoukikusa. Environ Sci Technol 44:6470–6474.
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39. You J, Chan Z (2015) ROS regulation during abiotic stress responses in crop plants. Front Plant Sci 6:1092.
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40. Ziegler P, Adelmann K, Zimmer S, Schmidt C, Appenroth KJ (2014) Relative in
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vitro growth rates of duckweeds (Lemnaceae) - The most rapidly growing higher
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plants. Plant Biol 17:1–9.
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41. Ziegler P, Sree KS, Appenroth KJ (2016) Duckweeds for water remediation and toxicity testing. Toxicol Environ Chem 2248:1–28.
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Acknowledgement
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This study was supported by the Advanced Low Carbon Technology Research and
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Development Program (ALCA) of the Japan Science and Technology Agency (JST).
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We would like to thank Editage (www.editage.jp) for English language editing.
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Abbreviations
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ANOVA, analysis of variance; APX, ascorbate peroxidase; CAT, catalase; CFU, colony
22
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forming
5,5’-dithiobis-2-nitrobenzoic
acid;
DRB,
528
rhizobacteria; DW, dry wight; FW, fresh weight; GPX, guaiacol peroxidase; GR,
529
glutathione reductase; GSSG, glutathione disulfide; MAMP, microbe associated
530
molecular pattern; MDA, malondialdehyde; NBT, nitroblue tetrazolium; PGIB, plant
531
growth-inhibiting bacteria; PGPB; plant growth-promoting bacteria, RGR, relative
532
growth rate; ROS, reactive oxygen species; SD; standard deviation; SOD, superoxide
533
dismutase; TCA, trichloroacetic acid.
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DTNB,
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unit;
deleterious
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ACCEPTED MANUSCRIPT Table 1 Effects of bacterial strains on the growth of L. minor, where the initial frond number = 10. Mean ± SD are shown. Frond number 4d
RGR 8d
4d
FW (mg)
DW (mg)
8d
8d
8d
45.8 ± 2.5 a
190.0 ± 4.9 a
0.380 ± 0.014 a
0.368 ± 0.003 a
236.0 ± 1.6 a
12.62 ± 0.20 a
M12
42.3 ± 2.2 b
177.8 ± 9.4 b
0.360 ± 0.013 b
0.360 ± 0.007 b
220.7 ± 11.9 b
12.26 ± 0.32 ab
M3
40.7 ± 1.2 c
158.7 ± 4.8 d
0.351 ± 0.008 c
0.345 ± 0.004 d
198.3 ± 2.9 c
11.33 ± 0.21 c
M6
39.3 ± 1.4 d
152.3 ± 5.9 d
0.342 ± 0.009 d
0.340 ± 0.005 d
194.3 ± 6.2 c
10.98 ± 0.33 c
Control
41.7 ± 1.7 bc
169.5 ± 3.9 c
0.357 ± 0.010 bc
0.354 ± 0.003 c
215.3 ± 3.3 b
11.94 ± 0.22 b
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Significant differences between treatments and control are designated by different letters (ANOVA, p < 0.05).
Table 2 Change of oxidative stress associated indicators induced by PGIB strains in comparison with copper (Panda 2008). Mean percent change from control (non-treated) plants are shown. Exposure term
PGIB (M3)
4d
PGIB (M6)
H 2O 2
MDA
CAT
APX
GPX
GR
SOD
Reference
98.3
15.9
14.8
8.3
15.0
33.4
10.3
-6.7
This study
8d
119.2
5.7
11.4
27.8
52.8
52.8
33.6
28.4
This study
4d
167.8
23.3
39.7
6.5
15.9
29.7
5.3
10.8
This study
8d
126.8
44.3
18.3
22.3
53.3
58.5
45.5
22.8
This study
0.0
15.3
64.5
-
-
9.8
32.4
7.7
Panda (2008)
13.0
24.7
89.4
-
-
0.0
50.9
6.3
Panda (2008)
31.3
40.0
95.8
-
-
-9.8
82.4 -13.0
Panda (2008)
2+
2d
2+
2d
20 µM Cu 50 µM Cu
2+
2d
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100 µM Cu
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Stress factor
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a
8.E+05 a
6.E+05 b
4.E+05 c c
b
2.E+05
b
b
0.E+00 H3
M3
M6
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CFU (g-1-FW)
1.E+06
M12
Figure 1 Amount of bacterial strains that colonized on plant bodies after 4 days (grey bars) and 8 days (black bars)
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cultivations. Error bars show the standard deviations (n = 3). Significant differences between treatments are
1.2
a
a
0.8 0.6
a
a
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Chl a+b (mg g-1-FW)
a
1.0
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designated by different letters (ANOVA, p < 0.05).
a
a a
a a
M6
M12
Control
0.4 0.2 0.0
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H3
M3
Figure 2 Chlorophyll content of L. minor cultivated with and without inoculation of bacterial strain after 4 days (grey bars) and 8 days (black bars) of growth. Error bars show the standard deviations (n = 3). Values share the
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same letter indicate no significant difference (ANOVA, p<0.05).
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5 4
b 0.20
a
a
b b b
a
ab
b
b
3 c c
2 1
0.15
b a b
0.05 0.00
0 H3
M3
M6
M12
H3
Control
b b
b
0.10
b
a
b
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H2O2 (mg g-1-FW)
O2•− (∆A580 h-1 g-1-FW)
a
M3
M6
M12
Control
Figure 3 Effect of bacterial strains on the plant superoxide anion level (a), hydrogen peroxide content (b) after 4
SC
days (grey bars) and 8 days (black bars) of cultivations. Error bars show the standard deviations (n = 3). Significant
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differences between treatments are designated by different letters (ANOVA, p < 0.05).
30
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20
ab
ab
a
a
ab a
a
b a
15 10
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MDA (µg g-1-FW)
25
5 0
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H3
M3
M6
M12
Control
Figure 4 Effect of bacterial strains on the plant malondialdehyde (MDA) content after 4 days (grey bars) and 8 days (black bars) of growth. Error bars show the standard deviations (n = 3). Significant differences between treatments are designated by different letters (ANOVA, p < 0.05).
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a a
a
0.15
a
0.10
a
a
a a
a a
0.05 0.00 M12
0.40
a
b b
b
0.30 a
a
0.20
b
b
0.10
ab
b
0.10 0.08 0.06
a
ab
ab
0.04
M3
M6
M12
ab
a
Control
ab
b a bc
c
0.02 0.00
M3
M6
a
a
M12
120 100
ab
c a
a a
40
H3
a
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20 0
Control
M3
H3
M3
M6
M12
Control
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80 60
b
GR (U mg-1-protein)
a
a
a
a
0.2
H3
d
H3
SOD (U mg-1-protein)
b
0.4
Control
0.50
b
b
0.6
SC
M6
0.00
e
0.8
M AN U
GPX (U mg-1-protein)
M3
a
a
0.0 H3
c
1.0
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b
0.20
APX (U mg-1-protein)
CAT (U mg-1-protein)
a
M6
M12
bc
a
Control
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Figure 5 Effect of bacterial strains on the plant enzymatic antioxidant activities after 4 days (grey bars) and 8 days (black bars) of cultivations. The activities of catalase (a), ascorbate peroxidase (b), guaiacol peroxidase (c), glutathione reductase (d), and superoxide dismutase (e) are shown. Error bars show the standard deviations (n = 3). Significant differences between treatments were designated by different letters (ANOVA, p < 0.05).
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Contributions Hidehiro Ishizawa designed and performed the experiments, interpreted the results, and drafted the manuscript. Masashi Kuroda designed the experiments, interpreted the
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results, drafted and revised the manuscript. Masaaki Morikawa interpreted the results, critically revised the manuscript, and supervised the project. Michihiko Ike designed the experiments, interpreted the results, drafted and revised the manuscript, and supervised
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the project.
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S0981-9428(17)30257-7
DOI:
10.1016/j.plaphy.2017.08.006
Reference:
PLAPHY 4960
To appear in:
Plant Physiology and Biochemistry
Received Date: 7 June 2017 Revised Date:
7 August 2017
Accepted Date: 8 August 2017
Please cite this article as: H. Ishizawa, M. Kuroda, M. Morikawa, M. Ike, Differential oxidative and antioxidative response of duckweed Lemna minor toward plant growth promoting/inhibiting bacteria, Plant Physiology et Biochemistry (2017), doi: 10.1016/j.plaphy.2017.08.006. This is a PDF file of an unedited manuscript that has been accepted for publication. As a service to our customers we are providing this early version of the manuscript. The manuscript will undergo copyediting, typesetting, and review of the resulting proof before it is published in its final form. Please note that during the production process errors may be discovered which could affect the content, and all legal disclaimers that apply to the journal pertain.
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Differential oxidative and antioxidative response of duckweed Lemna minor
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toward plant growth promoting/inhibiting bacteria
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Hidehiro Ishizawa1, Masashi Kuroda1, Masaaki Morikawa2, and Michihiko Ike1*
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Engineering, Osaka University, 2-1 Yamadaoka, Suita, Osaka 565-0871, Japan
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Division of Sustainable Energy and Environmental Engineering, Graduate School of
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Division of Biosphere Science, Graduate School of Environmental Science, Hokkaido
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University, N10-W5, Kita-ku, Sapporo 060-0810, Japan
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Corresponding author
Phone: +81-(0)6-6879-7674; fax:+81-(0)6-6879-7675
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Email:
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Hidehiro Ishizawa: [email protected]
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Masashi Kuroda: [email protected]
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Masaaki Morikawa: [email protected]
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Michihiko Ike: [email protected]
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Highlights 1
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・Growth inhibiting bacteria induced higher oxidative stress levels of duckweed than growth-promoting bacteria.
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for the effects on duckweed oxidative stress level.
・Growth promoting bacteria also induced equal or higher oxidative stress relative to aseptic control.
・Bacteria can be a significant cause of oxidative stress similar to the other
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・Two plant growth-promoting and two growth-inhibiting bacterial strains were tested
environmental factors.
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Abstract
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Bacteria colonizing the plant rhizosphere are believed to positively or negatively
48
affect the host plant productivity. This feature has inspired researchers to engineer such 2
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interactions to enhance crop production. However, it remains to be elucidated whether
50
rhizobacteria influences plant oxidative stress vis-a-vis other environmental stressors,
51
and whether such influence is associated with their growth promoting/inhibiting ability.
52
In this study, two plant growth-promoting bacteria (PGPB) and two plant
53
growth-inhibiting bacteria (PGIB) were separately inoculated into axenic duckweed
54
(Lemna minor) culture under laboratory conditions for 4 and 8 days in order to
55
investigate their effects on plant oxidative stress and antioxidant activities. As
56
previously characterized, the inoculation of PGPB and PGIB strains accelerated and
57
reduced the growth of L. minor, respectively. After 4 and 8 days of cultivation,
58
compared to the PGPB strains, the PGIB strains induced larger amounts of O2•−, H2O2,
59
and malondialdehyde (MDA) in duckweed, although all bacterial strains consistently
60
increased O2•− content by two times more than that in the aseptic control plants.
61
Activities of five antioxidant enzymes were also elevated by the inoculation of PGIB,
62
confirming the severe oxidative stress condition in plants. These results suggest that the
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surface attached bacteria affect differently on host oxidative stress and its response,
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which degree correlates negatively to their effects on plant growth.
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Keywords: Duckweed, Plant growth-promoting bacteria, plant growth-inhibiting
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bacteria, Oxidative stress, Reactive oxygen species, Plant-microbe interactions
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Introduction
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Plants experience a variety of environmental factors, and not all are ideal for plant
71
growth (Boyer, 1982). Most of the factors, such as low temperature, salinity, ultraviolet
72
radiation, and pathogen inhibit the growth of plants, commonly by inducing oxidative 3
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stress through the accumulation of reactive oxygen species (ROS). Although ROS are
74
byproducts of normal cellular activities in the mitochondria, chloroplast, and
75
peroxisome, they are capable of causing damage to plant cellular lipids, proteins, and
76
DNA through radical reactions when present in excess. On the other hand, plants have
77
developed well-tuned antioxidant systems to defend themselves from ROS, and these
78
have been extensively reviewed (Mittler, 2002; You and Chan, 2015). However, it is
79
also reported that the generation of ROS often exceeds the plants’ antioxidant capacity
80
and causes significant loss of the plant biomass and yield even under normal
81
environmental conditions (Apel and Hirt, 2004). Up-regulation of antioxidants itself
82
may inhibit growth through cross-talk between developmental and stress-response
83
networks (Cabello et al., 2014). Therefore, maintaining ROS and antioxidants at low
84
levels is essential to achieve enhanced productivity.
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Recently, it became known that bacterial communities present in the plant
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rhizosphere are one of the factors impacting plant productivity akin to the
87
environmental factors (Anderson and Habiger et al., 2012). Extensive research in this
88
field has demonstrated the general occurrence of plant growth-promoting bacteria
89
(PGPB) and plant growth-inhibiting bacteria (PGIB) or deleterious rhizobacteria (DRB)
90
poses beneficial and deleterious effects on the host growth (Probenza et al., 1996;
91
Suslow and Schroth, 1981). Our original research using duckweed Lemna minor as a
92
model plant showed that both promotive and inhibitory effects by the co-existing
93
bacterial community can be caused by complex and interactive influences of PGPB and
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PGIB existing in the root and frond zone of duckweed (Ishizawa et al., 2017). Therefore,
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to establish new approaches to improve crop productivity utilizing bacteria, it is critical
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that interactions between plants and bacteria, especially for PGPB and PGIB, are well
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understood. It is likely that oxidative stress plays a role in determining the beneficial and
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harmful effects of rhizobacteria on the plant, considering that oxidative stress caused by
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environmental factors also affects plant growth (Apel and Hirt, 2004). Therefore,
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understanding plant–bacterial interactions from the viewpoint of oxidative stress in
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plants may offer clues to elucidate the mechanisms leading to promotive/inhibitory
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effects on plant growth involving rhizobacteria. Such knowledge is helpful in
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developing techniques/strategies to properly regulate the rhizobacterial community to
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improve plant growth. However, studies on the relationship between plant oxidative
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stress and coexisting bacteria have been scarce. The aim of this study is to examine how
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PGPB and PGIB affect oxidative stress levels of the host plant and its response. Toward
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this, sterile L. minor was co-cultivated with previously isolated PGPB and PGIB strains
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under laboratory conditions, and the changes in plant ROS and other stress associated
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indicators in duckweed were monitored.
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Materials and Methods
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Plant and bacterial strains
Duckweed (Lemna minor, RDSC clone 5512) plants collected from a small pond
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in the botanical garden of Hokkaido University (Sapporo, Japan) were used in the
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experiments. The plants were sterilized by washing with 0.5% sodium hypochlorite for
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7 min and followed by washing twice with sterilized water. The sterilized plants were
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successively cultured in flasks containing modified Hoagland medium (36.1 mg/L
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KNO3, 293 mg L-1 K2SO4, 3.87 mg L-1 NaH2PO4, 103 mg L-1 MgSO4·7H2O, 147 mg L-1
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CaCl2·H2O, 3.33 mg L-1 FeSO4·7H2O, 0.95 mg L-1 H3BO3, 0.39 mg L-1 MnCl2·4H2O, 5
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0.03 mg L-1 CuSO4·5H2O, 0.08 mg L-1 ZnSO4·7H2O, 0.254 mg L-1 H2MoO4·4H2O, and
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5 mg L-1 EDTA·2Na; pH 7.0) and incubated at 28°C, with a light intensity of 80 µmol
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m-2s-1 and a photoperiod of 16 h/8 h day/night cycle. Four bacterial strains (Aquitalea magnusonii H3, Acinetobacter septicus M3,
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Asticcacaulis exentricus M6, Pseudomonas otitidis M12) isolated from the same
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duckweed strain were used for the study. The strains were previously characterized in
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terms of their effects on duckweed growth when co-cultivated with sterile L. minor:
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strains H3 and M12 were promotive (PGPB), while M3 and M6 were inhibitory (PGIB)
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(Ishizawa et al., 2017). These PGPB or PGIB strains were cultivated by inoculating a
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loop of bacterial colony into 20 mL of liquid LB medium and shaking the tube at 120
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rpm at 28°C until the culture reached the late exponential phase. Cells were then
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harvested by centrifugation (10,000 × g, 4°C, 10 min), washed twice with sterilized
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Hoagland medium before using them in our experiments.
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Experimental design
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Sterile L. minor plants were co-cultivated with each one of the four bacterial
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strains under similar light and nutrient conditions as mentioned above. Attachment of
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bacterial strains to L. minor was enabled by growing it along with a suspension of
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bacterial cells (optical density at 600 nm = 0.1) added to the sterilized Hoagland
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medium and maintained for 24 h prior to the experiment. Then, 10 fronds of L. minor
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with the attached bacteria were transferred to 60 mL of fresh bacteria-free medium and
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co-cultivated. During co-cultivation, the medium was replenished at 48 h intervals.
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After 4 and 8 days of cultivation, the plants were harvested and subjected to the
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analyses of ROS and other stress associated indicators.
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Plant growth evaluation During the growth period, the number of L. minor fronds was periodically
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counted and recorded for evaluating the effect of PGPB/PGIB strains on plant growth.
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Relative growth rate (RGR, d-1) was calculated as (ln FNt – ln 10)/t, where FNt is the
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frond number of L. minor on day t (4 or 8). In addition, fresh weight (FW) and dry
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weight (DW, 80°C for 24 h) of plants were measured at the end of 8-d growth period.
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Estimation of the amount of bacteria attached on duckweed
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The amount of bacterial cells that attached to plants was estimated as the number
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of colony forming units (CFU) per gram fresh weight of the plants. At the end of each 4
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and 8 day growth periods, 20 mg of plants were washed twice with 20 mL of sterile
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Hoagland medium and homogenized in 5 mg L-1 tripolyphosphate (TPP) using a
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BioMasher II (Nippi, Tokyo, Japan). The homogenates were spread onto solid 1:10
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diluted LB medium containing 1.5% agar. Agar plates were incubated at 28°C for 3
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days and the number of bacterial colonies were counted.
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Determination of chlorophyll content
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Total plant chlorophyll content was determined spectrophotometrically (UV-1850,
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Shimazu, Kyoto, Japan). Pigment extraction was performed by soaking 30 mg of plants
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in 3 mL of methanol for 90 min in the dark. Chlorophyll content per milligram fresh
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weight was calculated using absorbance at 650 nm (A650) and 665 nm (A665) and
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applying the formula (Grimme and Boardman, 1972).
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Chl a + b = 4.0 × A665 + 25.5 × A650 7
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Estimation of ROS Plant superoxide anion (O2•−) level was estimated by measuring the reduction of
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nitroblue tetrazolium (NBT) according to the method suggested by Doke (1983). Thirty
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milligrams of fresh plant samples were immersed in 1.5 ml of a mixture containing 10
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mM sodium phosphate buffer (pH 7.8), 0.05% NBT and 10 mM NaN3 in a test tube,
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followed by incubation at 23°C for 60 min. Following this, 1.0 mL of the reaction
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mixture was incubated at 85°C for 15 min. This reaction was stopped by cooling the
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solution with ice and absorbance at 580 nm (A580) was measured to assay the reduction
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of NBT. The level of O2•− was expressed as the increase in A580 per hour per gram fresh
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weight.
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Hydrogen peroxide (H2O2) content was measured according to Velikova et al.
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(2000). Thirty milligrams of plant material were homogenized in 1.5 mL test tube with
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1.0 mL of ice-cold 0.1% trichloroacetic acid (TCA) using a high-power homogenizer
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(ASG50. AS ONE, Aichi, Japan) at 6,000 rpm for 30 s. Homogenates were then
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centrifuged (10,000 × g, 4°C, 15 min), and 0.5 mL of supernatant was mixed with 0.5
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mL of 10 mM potassium phosphate buffer (pH 7.0) and 1.0 mL of 1 M KI. The mixture
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was incubated in the dark at 23°C for 60 min, and the absorbance at 390 nm was
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measured. The content of H2O2 was calculated using a standard curve plotted with
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known concentrations of H2O2.
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Determination of malondialdehyde (MDA) content
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The level of lipid peroxidation was estimated indirectly by measuring the content
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of MDA, a by-product of lipid peroxidation (Heath and Packer, 1968). Thirty 8
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milligrams of plant material were homogenized in 1.5 mL test tube containing 0.5%
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2-thiobarbituric acid in 1 mL of 20% TCA using a high-power homogenizer at 6,000
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rpm for 30 s. Homogenates were then transferred to centrifuge tubes and incubated at
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95°C for 15 min. The reaction was stopped by placing the tubes on ice, and followed by
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centrifugation (10,000 × g, 2°C, 10 min). The absorbance of the supernatant was
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measured at 532 nm. Absorbance at 600 nm was also measured to account for
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non-specific turbidity or background. The content of MDA was calculated using an
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extinction coefficient of 155 mM-1 cm-1.
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Antioxidant enzyme assays
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Catalase (CAT, EC 1.11.1.6) activity was determined by measuring the change in
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absorbance at 240 nm which indicates decomposition of H2O2 (e = 40 mM-1cm-1)
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according to the method of Aebi (1984). The reaction mixture consisted of 900 µL of 50
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mM potassium phosphate buffer (pH 7.0), 50 µL of 300 mM H2O2, and 50 µL of
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enzyme extract. The reaction was started by adding H2O2.
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Ascorbate peroxidase (APX, EC 1.11.1.11) activity was assayed by measuring the
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change in absorbance at 290 nm that accompanied the consumption of ascorbic acid (e
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=2.8 mM-1cm-1) as described in Nakano and Asada (1981). The reaction mixture
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contained 800 µL of 50 mM potassium phosphate buffer (pH 7.0), 0.1 mM EDTA, 0.5
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mM L-ascorbic acid, 50 µL of 100 mM H2O2, and 150 µL of enzyme extract.
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For guaiacol peroxidase (GPX, EC 1.11.1.7) activity determination, increase in
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absorbance at 470 nm indicating the formation of tetraguaiacol (e = 26.6 mM-1cm-1) was
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measured as previously described (Horvat et al., 2007). The reaction mixture consisted
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of 900 µL potassium phosphate buffer (pH = 6.5), 15 mM guaiacol, 50 µL of 300 mM 9
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H2O2, and 50 µL of enzyme extract. Glutathione reductase (GR, EC 1.8.1.7) activity was determined using the method
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described by Smith et al. (1988) by measuring the increase in absorbance at 412 nm that
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corresponds to the reduction of 5,5’-dithiobis-2-nitrobenzoic acid (DTNB) to
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2-nitro-5-thiobenzoic acid (TNB, e =13.6 mM-1cm-1). The reaction mixture contained
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500 µL of 200 mM potassium phosphate buffer (pH 7.5), 1 mM EDTA, 250 µL of 3
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mM DTNB in 10 mM potassium phosphate buffer (pH 7.5), 100 µL of ultra-pure water,
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50 µL of 2 mM NADPH, 50 µL of 20 mM glutathione disulfide (GSSG), and 50 µL of
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enzyme extract. The reaction was initiated by adding GSSG to the mixture.
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Superoxide dismutase (SOD, EC 1.15.1.1) was assayed based on the inhibition of
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NBT’s photochemical reduction by the decomposition of O2•− (Beauchamp and
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Fridovich, 1971). The reaction mixture consisted of 3 mL of 50 mM sodium phosphate
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buffer (pH 7.8), 13 mM L-methionine, 75 µM NBT, 0.1 mM EDTA, 300 µL of 24 µM
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riboflavin, and 0 – 200 µL of enzyme extract. The total volume of the mixture was
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adjusted to 3.5 mL by adding 50 mM sodium phosphate buffer (pH 7.8). The mixture
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was illuminated at a light intensity of 70 µmol m-2s-1 for 10 min, and the absorbance at
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560 nm was measured immediately to evaluate the photochemical reduction of NBT.
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One unit of SOD activity was defined as the amount of enzyme required to cause a 50%
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inhibition of the NBT photochemical reduction.
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The activity of all the above antioxidant enzymes was expressed as unit mg-1
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protein. For protein extraction, 50 mg of fresh plants in 1.5 mL test tube were
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homogenized with 1 ml of ice-cold 50 mM potassium phosphate buffer (pH 7.0)
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containing 1% (w/v) polyvinylpolypyrrolidone. The homogenate was then centrifuged
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(20,000 × g, 2°C, 20 min) and the supernatant was used for the enzyme assay. Soluble 10
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protein content of the supernatant was determined using the method of Bradford (1976)
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with bovine serum albumin as a standard.
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Statistical analysis
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All assays were performed with 3 biological replicates. Data was analyzed by
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one-way analysis of variance (ANOVA), and where it was significant, Duncan’s
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multiple-range test was performed to separate the means. Significance at p < 0.05 was
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applied to all analyses. Statistical analyses were performed in R software v3.2.3
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(http://www.r-project.org).
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Results
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Effects of bacterial inoculation on the growth and chlorophyll content of duckweed
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In this study, two PGPB strains (H3, M12) and two PGIB strains (M3, M6) were
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individually co-cultivated with sterile L. minor. The amounts of bacteria colonized on
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the plants are shown in Figure 1. Strain H3 had the highest CFU both after 4 and 8 days
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of cultivations. Although there was no carbon source to support bacterial growth in the
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medium, the CFU counts per plant fresh weight were maintained throughout the
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cultivation period, suggesting that these strains have stably attached to their host and
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established. Furthermore, as to strain M3, a notably higher number of CFU was
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observed after 8 days than at 4 days.
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Table 1 summarizes the growth indices of L. minor. In the experiment, RGR of
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control plants (0.354 for 8-day cultivation) was at the same level as reported by Ziegler
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et al. (2014) which used the optimized culture condition for duckweeds. Similar to the
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previous study (Ishizawa et al., 2017), PGPB strain H3 and M12 accelerated L. minor 11
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growth, while PGIB strains M3 and M6 significantly delayed growth compared to
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aseptic control, in terms of the frond number on day 4 and 8. One notable difference is
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that the growth-promoting effect of strain M12 was smaller than reported previously.
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We believe this might be due to the higher growth rate of control plants owing to the
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difference in nutrient conditions.
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Plant chlorophyll content did not show any significant differences among the
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plants co-cultivated with the four bacterial strains (Figure 2), indicating the little effects
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of PGPB and PGIB on the pigment synthesis and/or destruction of L. minor. It should
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be noted that plant had lower levels of chlorophyll content after day 8 than day 4 for all
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the tests, suggesting some change in duckweed morphology or nutritional status have
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occurred in the later part of cultivation period, possibly due to the high plant density
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and/or formation of ethylene (Farber and Kandeler, 1990).
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Plant ROS levels
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We observed significant changes in the superoxide anion (O2•−) and hydrogen
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peroxide (H2O2) levels of L. minor, depending on the bacterial strains used for
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co-cultivation. These free radicals serve as the major forms of ROS in plants. As shown
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in Figure 3a, all bacterial strains significantly elevated O2•− content of the plants
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compared to the aseptic control. We have confirmed that simple introduction of bacterial
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cells (105~106 cfu) to the test reagent of O2•− did not affect the result of the assay (data
285
not shown). Among the bacterial strains, PGIB strain M6, which had the strongest
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growth-inhibitory effects in this experiment, caused the largest increase in the O2•−
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content of L. minor. Another PGIB strain M3 also tended to induce more than PGPB
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strains, though the trend is not statistically significant.
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As for H2O2 content, significantly higher value compared to the control were
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recorded for the plants cultivated with PGIB strain M6 at days 4 and 8 as well as strain
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M3 at day 4 (Figure 3b). Although not significant, plants co-cultivated with PGPB
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strains H3 and M12 also showed higher H2O2 content than control plants, especially in
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the later part of growth period (day 8).
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Lipid peroxidation
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The level of duckweed lipid peroxidation was determined by estimating the
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amount of MDA accumulation in plant tissues (Figure 4) as an indicator of cellular
298
damage caused by ROS. Though the difference could not be confirmed due to large
299
variation in results for both on day 4 and 8, PGPB strains increased MDA content in the
300
range of 2.6 - 16.3%, while PGIB strains increased MDA in the range of 11.4 - 39.7%
301
compared to the control plants. Thus, the general trend of the changes in MDA content
302
during the cultivation period was similar to that of O2•− and H2O2 content, showing that
303
the degree of lipid peroxidation is more or less correlated to ROS production.
304
Specifically, it is worth noting that the significant increase of MDA content observed for
305
the plants co-cultivated with M6 on day 4 coincided with the highest accumulation of
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O2•− (Figure 3a).
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Antioxidant enzyme activities
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The effects of bacterial strains on the activities of five important antioxidative
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enzymes of the plants were assayed. Although we could not exclude bacterial-originated
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enzymes in these assays, their effects on the results are considered to be limited because
312
the amount of bacterial protein extracted in the assays would be extremely smaller than 13
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plant proteins. Similar to the ROS and lipid peroxidation, the higher values were
314
recorded for the plants cultivated with PGIB strains M3 and M6 in all enzyme studies
315
except CAT (Figure 5). Since plants are known to up-regulate antioxidant enzymes
316
responding to adverse environments or ROS itself (Babu et al., 2003), the result would
317
reflect the plant activation of antioxidant enzymes by perceiving the higher ROS content.
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In addition, we observed that activities of each enzyme were quite different between the
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plants collected after 4 and 8 days. Because the values were expressed based on the
320
amount of total extracted protein, it suggests different protein allocation in plants
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depending on the growth stage.
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Discussion
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Despite a number of publications on growth promoting and growth inhibiting
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effects of plant rhizobacteria, their effects, particularly on plant oxidative stress
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response, are poorly understood. This study highlights the effects of bacterial strains on
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plant oxidative stress and antioxidant activities using duckweed L. minor, which is a
328
useful model plant for investigating plant-microbe interactions (Appenroth et al., 2016).
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As previously characterized (Ishizawa et al., 2017), the bacterial strains used in
330
this study either promoted growth or inhibited the growth of duckweed (Table 1), while
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the influence on chlorophyll content was limited (Figure 2). Interestingly, both PGPB
332
and PGIB caused significantly different oxidative stress levels in L. minor, as seen by
333
measuring the ROS content, MDA content, and antioxidant enzymatic activities (Figure
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3-5). In our experiments, the numbers of living bacterial cells colonized on the plants,
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which might be a kind of bacterial dose to the plant, were estimated as CFU counts. The
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CFU counts showed a large variation up to about ten times between treatments (Figure
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1). However, their effect on the plant oxidative stress levels could not be correlated to
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the CFU counts. Therefore, it is likely that the observed changes in plant oxidative
339
stress levels depend on the nature and type of bacterial strains introduced rather than the
340
amount of bacterial cells attached.
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Among four bacterial strains used in this study, two PGPB strains, Aquitalea
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magnusonii H3 and Pseudomonas otitidis M12, showed relatively moderate influence
343
on plant oxidative stress. However, these strains led to very high levels of O2•− content
344
in inoculated plants relative to control plants, as well as PGIB strains (Figure 3a). It is
345
likely that bacteria generally stimulated the plant O2•− generation, though the degree was
346
different among bacterial strains. Plants possess the ability to generate O2•− as an
347
immune reaction when they recognize microorganisms through perception of
348
microbe-associated molecular patterns (MAMPs) such as presence of chitin or flagellins
349
(Smith et al., 2015). Other studies have shown that plants generate O2•− from the activity
350
of NADPH oxidase during the establishment of plant–bacterial symbiosis as signal
351
molecules (Nanda et al., 2010). This suggests that the observed phenomena were a
352
result of spontaneous generation of O2•− by plants in response to bacterial colonization,
353
which did not occur in the aseptic control plants.
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On the other hand, there are some recent studies of PGPB associated with
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terrestrial plants that act to alleviate plant oxidative stress under harsh stress conditions
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through the regulation of antioxidant activities (Habib et al., 2016; Islam et al., 2014). In
357
our study, we used favorable growth conditions for L. minor. As a result, the role of
358
PGPB strains in alleviating plant oxidative stress compared to the aseptic control was
359
not detected, suggesting the presence of growth-promoting mechanisms of PGPB other
360
than through stress alleviation. However, in a few studies, other duckweed PGPB strains 15
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were reported to enhance the growth of duckweed even in the presence of toxic phenol
362
(Yamaga et al., 2010) and chromium (Tang et al., 2015). Considering all these studies,
363
the possibility that duckweed PGPB strains play a role in alleviating plant stress when
364
plants are exposed to stressful environments cannot be ruled out. Therefore, to gain
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more insights on beneficial plant–microbe interactions additional studies are required.
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Compared to the PGPB strains, two PGIB strains, Acinetobacter ursingii M3
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and Asticcacaulis excentricus M6, caused considerably higher oxidative stress levels in
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L. minor (Figure 3-5). Although there are several studies about the mechanism of plant
369
growth promotion by PGPB such as nutrient supply and modulation of plant hormone
370
(Glick, 2012), very little is known about how PGIB negatively affect plant growth and
371
productivity. Thus far, hydrogen cyanide production (Astrom, 1991) and ammonia
372
production (Weise et al., 2013) were proposed as possible mechanisms of existing PGIB
373
strains. In this study, we show that PGIB strains induce higher levels of oxidative stress
374
in the host plant than by the other bacterial strains, which explain at least some of the
375
growth-inhibitory mechanisms employed by PGIB. The knowledge that PGIB induce a
376
considerably high levels of plant oxidative stress may provide clues toward establishing
377
strategies for crops to defend PGIB. For example, engineering plants for elevated
378
abiotic stress tolerance or introducing outcompeting rhizobacterial strains with less
379
influence on plant stress might be effective for managing the adverse effects of PGIB.
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Duckweeds have been used in plant stress research and for chemical toxicity tests
381
for a long time (Ziegler et al., 2016), and their responses toward environmental stress
382
factors have been extensively studied. Thus far, a variety of heavy metals (Parlak et al.,
383
2012; Tkalec et al., 2008; Duman et al., 2010), salinity (Cheng, 2011), ammonium
384
(Wang et al., 2016), and chemical compounds (Obermeier et al., 2015) have been shown 16
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to induce a dose-dependent response to oxidative stress, while the degree and type of
386
activated antioxidants seemed to vary and depend highly on the culture conditions
387
and/or duckweed strains used. In this study, similar responses of duckweed to the
388
above-mentioned abiotic stressors are being reported for the first time for
389
plant-associated bacteria, particularly in suppressing the growth of the host plant.
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Table 2 summarizes the oxidative and antioxidative responses induced by PGIB
391
strains, in comparison to Panda (2006) who investigated the response of L. minor fronds
392
to copper (Cu2+) exposure. Copper is one of the most typical and well-studied stress
393
factors for duckweed, which induces severe oxidative stress in the plant at higher
394
concentrations (Panda et al., 2008; Babu et al., 2003; Razinger et al., 2007). As shown
395
in Table 2, increase in O2•− content and GPX caused by PGIB were much higher than
396
that caused by copper, although the responses to H2O2 content was comparable to
397
moderate concentrations of Cu2+. Upregulation of antioxidant enzymes induced by
398
PGIB were also comparable to that by copper, except for GPX. Even though we
399
observed a considerable increase in MDA content for plants co-cultivated with PGIB,
400
the level of lipid peroxidation did not seem to be as high as that caused by Cu2+.
401
Considering these information, we propose that the effects on the plant oxidative stress
402
by PGIB strains are generally equivalent to the moderate dose of environmental
403
contaminants such as copper, which should be certainly a focus of future research.
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In summary, this is the first study to highlight the different effects of PGPB and
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PGIB strains on the oxidative stress and antioxidant enzyme activities of L. minor. Two
406
PGIB strains were found to increase plant oxidative stress and antioxidant enzyme
407
activities similar to abiotic stress factors reported previously. On the other hand, PGPB
408
strains were less stressful, though they also caused certain levels of oxidative stress on 17
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the plant when compared to the control plant. Although the detailed mechanisms are
410
still unknown, oxidative stress and its response may play important roles in growth
411
inhibition caused by PGIB. In addition, a series of oxidative stress assays performed in
412
this study can serve as useful tools to evaluate beneficial/deleterious effects of bacteria
413
on duckweed productivity and possibly extend to studies on other plant species.
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Acknowledgement
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This study was supported by the Advanced Low Carbon Technology Research and
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Development Program (ALCA) of the Japan Science and Technology Agency (JST).
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We would like to thank Editage (www.editage.jp) for English language editing.
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Abbreviations
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ANOVA, analysis of variance; APX, ascorbate peroxidase; CAT, catalase; CFU, colony
22
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forming
5,5’-dithiobis-2-nitrobenzoic
acid;
DRB,
528
rhizobacteria; DW, dry wight; FW, fresh weight; GPX, guaiacol peroxidase; GR,
529
glutathione reductase; GSSG, glutathione disulfide; MAMP, microbe associated
530
molecular pattern; MDA, malondialdehyde; NBT, nitroblue tetrazolium; PGIB, plant
531
growth-inhibiting bacteria; PGPB; plant growth-promoting bacteria, RGR, relative
532
growth rate; ROS, reactive oxygen species; SD; standard deviation; SOD, superoxide
533
dismutase; TCA, trichloroacetic acid.
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DTNB,
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unit;
deleterious
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23
ACCEPTED MANUSCRIPT Table 1 Effects of bacterial strains on the growth of L. minor, where the initial frond number = 10. Mean ± SD are shown. Frond number 4d
RGR 8d
4d
FW (mg)
DW (mg)
8d
8d
8d
45.8 ± 2.5 a
190.0 ± 4.9 a
0.380 ± 0.014 a
0.368 ± 0.003 a
236.0 ± 1.6 a
12.62 ± 0.20 a
M12
42.3 ± 2.2 b
177.8 ± 9.4 b
0.360 ± 0.013 b
0.360 ± 0.007 b
220.7 ± 11.9 b
12.26 ± 0.32 ab
M3
40.7 ± 1.2 c
158.7 ± 4.8 d
0.351 ± 0.008 c
0.345 ± 0.004 d
198.3 ± 2.9 c
11.33 ± 0.21 c
M6
39.3 ± 1.4 d
152.3 ± 5.9 d
0.342 ± 0.009 d
0.340 ± 0.005 d
194.3 ± 6.2 c
10.98 ± 0.33 c
Control
41.7 ± 1.7 bc
169.5 ± 3.9 c
0.357 ± 0.010 bc
0.354 ± 0.003 c
215.3 ± 3.3 b
11.94 ± 0.22 b
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Significant differences between treatments and control are designated by different letters (ANOVA, p < 0.05).
Table 2 Change of oxidative stress associated indicators induced by PGIB strains in comparison with copper (Panda 2008). Mean percent change from control (non-treated) plants are shown. Exposure term
PGIB (M3)
4d
PGIB (M6)
H 2O 2
MDA
CAT
APX
GPX
GR
SOD
Reference
98.3
15.9
14.8
8.3
15.0
33.4
10.3
-6.7
This study
8d
119.2
5.7
11.4
27.8
52.8
52.8
33.6
28.4
This study
4d
167.8
23.3
39.7
6.5
15.9
29.7
5.3
10.8
This study
8d
126.8
44.3
18.3
22.3
53.3
58.5
45.5
22.8
This study
0.0
15.3
64.5
-
-
9.8
32.4
7.7
Panda (2008)
13.0
24.7
89.4
-
-
0.0
50.9
6.3
Panda (2008)
31.3
40.0
95.8
-
-
-9.8
82.4 -13.0
Panda (2008)
2+
2d
2+
2d
20 µM Cu 50 µM Cu
2+
2d
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100 µM Cu
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Stress factor
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a
8.E+05 a
6.E+05 b
4.E+05 c c
b
2.E+05
b
b
0.E+00 H3
M3
M6
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CFU (g-1-FW)
1.E+06
M12
Figure 1 Amount of bacterial strains that colonized on plant bodies after 4 days (grey bars) and 8 days (black bars)
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cultivations. Error bars show the standard deviations (n = 3). Significant differences between treatments are
1.2
a
a
0.8 0.6
a
a
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Chl a+b (mg g-1-FW)
a
1.0
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designated by different letters (ANOVA, p < 0.05).
a
a a
a a
M6
M12
Control
0.4 0.2 0.0
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H3
M3
Figure 2 Chlorophyll content of L. minor cultivated with and without inoculation of bacterial strain after 4 days (grey bars) and 8 days (black bars) of growth. Error bars show the standard deviations (n = 3). Values share the
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same letter indicate no significant difference (ANOVA, p<0.05).
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5 4
b 0.20
a
a
b b b
a
ab
b
b
3 c c
2 1
0.15
b a b
0.05 0.00
0 H3
M3
M6
M12
H3
Control
b b
b
0.10
b
a
b
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H2O2 (mg g-1-FW)
O2•− (∆A580 h-1 g-1-FW)
a
M3
M6
M12
Control
Figure 3 Effect of bacterial strains on the plant superoxide anion level (a), hydrogen peroxide content (b) after 4
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days (grey bars) and 8 days (black bars) of cultivations. Error bars show the standard deviations (n = 3). Significant
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differences between treatments are designated by different letters (ANOVA, p < 0.05).
30
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a
20
ab
ab
a
a
ab a
a
b a
15 10
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MDA (µg g-1-FW)
25
5 0
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H3
M3
M6
M12
Control
Figure 4 Effect of bacterial strains on the plant malondialdehyde (MDA) content after 4 days (grey bars) and 8 days (black bars) of growth. Error bars show the standard deviations (n = 3). Significant differences between treatments are designated by different letters (ANOVA, p < 0.05).
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a a
a
0.15
a
0.10
a
a
a a
a a
0.05 0.00 M12
0.40
a
b b
b
0.30 a
a
0.20
b
b
0.10
ab
b
0.10 0.08 0.06
a
ab
ab
0.04
M3
M6
M12
ab
a
Control
ab
b a bc
c
0.02 0.00
M3
M6
a
a
M12
120 100
ab
c a
a a
40
H3
a
EP
20 0
Control
M3
H3
M3
M6
M12
Control
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80 60
b
GR (U mg-1-protein)
a
a
a
a
0.2
H3
d
H3
SOD (U mg-1-protein)
b
0.4
Control
0.50
b
b
0.6
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M6
0.00
e
0.8
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GPX (U mg-1-protein)
M3
a
a
0.0 H3
c
1.0
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b
0.20
APX (U mg-1-protein)
CAT (U mg-1-protein)
a
M6
M12
bc
a
Control
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Figure 5 Effect of bacterial strains on the plant enzymatic antioxidant activities after 4 days (grey bars) and 8 days (black bars) of cultivations. The activities of catalase (a), ascorbate peroxidase (b), guaiacol peroxidase (c), glutathione reductase (d), and superoxide dismutase (e) are shown. Error bars show the standard deviations (n = 3). Significant differences between treatments were designated by different letters (ANOVA, p < 0.05).
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Contributions Hidehiro Ishizawa designed and performed the experiments, interpreted the results, and drafted the manuscript. Masashi Kuroda designed the experiments, interpreted the
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results, drafted and revised the manuscript. Masaaki Morikawa interpreted the results, critically revised the manuscript, and supervised the project. Michihiko Ike designed the experiments, interpreted the results, drafted and revised the manuscript, and supervised
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the project.
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