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Planta

, Volume 249, Issue 1, pp 235–249 | Cite as

Overexpression of geraniol synthase induces heat stress susceptibility in Nicotiana tabacum

  • Ashita Hamachi
  • Masahiro Nisihara
  • Shiori Saito
  • Hojun Rim
  • Hideyuki Takahashi
  • Monirul Islam
  • Takuya Uemura
  • Toshiyuki Ohnishi
  • Rika Ozawa
  • Massimo E. Maffei
  • Gen-ichiro ArimuraEmail author
Original Article
  • 414 Downloads
Part of the following topical collections:
  1. Terpenes and Isoprenoids

Abstract

Main conclusion

Transgenic tobacco plants overexpressing the monoterpene alcohol geraniol synthase exhibit hypersensitivity to thermal stress, possibly due to suppressed sugar metabolism and transcriptional regulation of genes involved in thermal stress tolerance.

Monoterpene alcohols function in plant survival strategies, but they may cause self-toxicity to plants due to their hydrophobic and highly reactive properties. To explore the role of these compounds in plant stress responses, we assessed transgenic tobacco plants overexpressing the monoterpene alcohol geraniol synthase (GES plants). Growth, morphology and photosynthetic efficiency of GES plants were not significantly different from those of control plants (wild-type and GUS-transformed plants). While GES plants’ direct defenses against herbivores or pathogens were similar to those of control plants, their indirect defense (i.e., attracting herbivore enemy Nesidiocoris tenuis) was stronger compared to that of control plants. However, GES plants were susceptible to cold stress and even more susceptible to extreme heat stress (50 °C), as shown by decreased levels of sugar metabolites, invertase activity and its products (Glc and Fru), and leaf starch granules. Moreover, GES plants showed decreased transcription levels of the WRKY33 transcription factor gene and an aquaporin gene (PIP2). The results of this study show that GES plants exhibit enhanced indirect defense ability against herbivores, but conversely, GES plants exhibit hypersensitivity to heat stress due to suppressed sugar metabolism and gene regulation for thermal stress tolerance.

Keywords

Geraniol Monoterpene Sugar metabolites Thermal stress Tobacco 

Abbreviations

CIS

Cooled injection system

CTS

Cold trap system

Fru

Fructose

FW

Fresh weight

geranyl-glc

Geranyl β-d-glucopyranoside

Geranyl-pri

Geranyl β-primeveroside

GES

Geraniol synthase

Glc

Glucose

G6P

Glucose 6-phosphate

GPP

Geranyl diphosphate

GUS

β-Glucuronidase

HSF

Heat shock factor

HSP

Heat shock protein

IPP

Isopentenyl diphosphate

Linalyl-glc

Linalyl β-d-glucopyranoside

linalyl-pri

Linalyl β-primeveroside

PDMS

Polydimethylsiloxane

PVOC

Plant volatile organic compound

qPCR

Quantitative polymerase chain reaction

ROS

Reactive oxygen species

Introduction

The majority of plant volatile organic compounds (PVOCs) consist of terpenes that play an important role in ecological interactions between plants and other organisms, including pollinators and herbivore enemies, and the surrounding environment (Arimura et al. 2016; Gershenzon and Dudareva 2007; Loreto and Schnitzler 2010). For instance, plants show induced production of volatiles in response to herbivore attack, and those PVOCs function in repelling herbivores such as ovipositing butterflies and host-seeking aphids (De Moraes et al. 2001; Kessler and Baldwin 2001; Unsicker et al. 2009) and in attracting herbivore enemies (Arimura et al. 2009). Likewise, by recruiting pollinators, PVOCs play an important role in plant reproduction (Arimura et al. 2016).

Apart from PVOCs’ infochemical functions, lipophilic volatiles are involved in plant responses to environmental stresses. Isoprene (C5 hemiterpene), one of the highly volatile terpenes, can stabilize plant cell membranes (Loreto and Schnitzler 2010), and is also able to quench tropospheric ozone to protect plants from oxidative stress (Loreto and Velikova 2001; Vickers et al. 2009; Velikova and Loreto 2005; Sharkey et al. 2008), as has been similarly shown for a suite of monoterpenes (C10) (Loreto et al. 2004; Pinto et al. 2007), sesquiterpenes (C15) (Hallquist et al. 2009) and green leaf volatiles (C6) (Hamilton et al. 2009). However, in contrast to the numerous studies on isoprenes, there have been only a few studies on the role of mono-and sesquiterpenes in such environmental stress tolerances. Monoterpene alcohols (e.g., linalool, menthol and geraniol) have been shown to exert multiple functions, e.g., apoptosis and oxidative stress induction (Ashida et al. 2002; Izumi et al. 1999), antimicrobial activity (Herman et al. 2016), mosquito repellence (Barnard and Xue 2004; Müller et al. 2009) and plant growth inhibition (Fischer et al. 2013). Indeed, land plant species have evolved mechanisms to release highly volatile compounds, including monoterpene alcohols, which may cause self-toxicity due to their hydrophobic and highly reactive activities (Yazaki et al. 2017). These compounds accumulate frequently in specific secretory structures (Iijima et al. 2004a; Maffei 2010; Yazaki et al. 2017), and product modifications, such as glycosylation and oxidation, may also contribute to the stable storage of terpenes in plant cell vacuoles to prevent self-toxicity.

In the current study, we highlighted geraniol, a representative monoterpene alcohol, as our research target. We selected this monoterpene because it is the predominant volatile component of several plant taxa, including rose, citronella and herb oils. The geraniol synthase (GES) gene has been used frequently for metabolic engineering of terpenes as model cases, such as in tomato, Arabidopsis, and grapevine (Vitis vinifera) (Davidovich-Rikanati et al. 2007; Fischer et al. 2013). However, interestingly, the ecological and physiological functions (e.g., defense ability to biotic and abiotic stresses) have never been assessed using these transgenic lines; only primary analyses regarding growth phenotypes and metabolic modifications have been reported. As described above, since this monoterpene alcohol may exert both detrimental and beneficial effects on plants (Izumi et al. 1999; Müller et al. 2009), we questioned whether and how transgenic plants that constitutively overproduce geraniol show an array of phenotypes. Here, we assessed transgenic plants in terms of (1) plant growth/reproduction; (2) defense responses to biotic stresses, as an array of PVOCs are ubiquitously functional for plant defense and communication; and (3) environmental stress tolerances, as some terpenes may play a role in adaption to abiotic stresses, as described above. For one of the model cases, we transformed an Ocimum basilicum geraniol synthase (ObGES) gene into tobacco plants, which constitute the most promising model plant species for terpene engineering (Lücker et al. 2004a, b; Muroi et al. 2011). The scope of this study will provide new information to advance our understanding of the detailed features of plant terpene engineering.

Materials and methods

Production of transgenic tobacco plants

We constructed a binary vector containing the ObGES gene expressed under the control of the cauliflower mosaic virus (CaMV) 35S promoter and Arabidopsis HSP 18.2 terminator (Nagaya et al. 2010). The full-length open reading frame of ObGES (AY362553) cDNA was amplified with the primers ObGESU-XbaI and ObGESL-XhoI using plasmid pET-ObGES (Iijima et al. 2004b) as the template and cloned into pCR4-TOPO (Invitrogen, Carlsbad, CA, USA). After confirmation that the desired sequence had been cloned, the cloned DNA was digested with Xba I and Xho I, and ligated into the pSKAN221-HT vector digested with the same enzymes. pSKAN221-HT was derived from pSKAN35SGUS (Nakatsuka et al. 2012) in which the nopaline synthase (NOS) terminator was replaced by the heat shock protein (HSP) 18.2 terminator. The resulting plasmid, pSKAN35SGES, was introduced into Agrobacterium tumefaciens strain EHA101 by electroporation. ObGES-overexpressing tobacco (Nicotiana tabacum cv. SR1) was produced by Agrobacterium-mediated transformation as described previously (Mishiba et al. 2010). Briefly, leaf sections were prepared from 1-month-old plants grown aseptically in an incubator and immersed in an Agrobacterium suspension for a short period. The sections were then co-cultivated for 2 days, after which they were transferred to selection medium containing 200 mg L−1 kanamycin. The cultures were subcultured every 2 weeks, after which the regenerated shoots were transferred to rooting medium containing 100 mg L−1 kanamycin. The culture process was performed at 25 °C under a 16-h photoperiod at a light intensity of approximately 30 µE m−2 s−1. After rooting and acclimatization, the regenerated plants were grown in a controlled greenhouse to set seeds. Twelve lines of transgenic T1 seeds were tested for germination on Murashige and Skoog medium supplemented with 200 mg L−1 kanamycin at 25 °C under a 16-h photoperiod at a light intensity of ca. 30 µE m−2 s−1. T2 seeds harvested from each T1 individual plant that showed ca. 3:1 segregation ratio were tested for kanamycin resistance again. Finally, T2 and T3 homozygous plant lines were used for further analyses. A homozygous tobacco T2 line transformed with the binary plasmid pSMABR-35SpGUS (in which the reporter GFP gene of pSMABR35SsGFP was replaced by the β-glucuronidase [GUS] gene), expressing bialaphos resistance and GUS genes, was used as a control. All of the tobacco plants were grown in plastic pots in a climate-controlled room at 24 ± 1 °C (16 h photoperiod at a light intensity of 80 µE m−2 s−1) for 6 weeks and then used for experiments. We used an individual plant for preparing a single set of samples for all analyses (Figs. 1, 2).
Fig. 1

GES products. a Headspace plant volatiles. Representative gas chromatography–mass spectrometry profile of headspace volatiles of the GUS control and GES4 plants are presented. Quantitative data represent the means of GES products (linalool, nerol, β-citronellol, and geraniol) from wild-type (WT), GUS and GES (4, 7, 10) plants with standard errors (n = 3). 1: linalool, 2: nonanal; 3: dodecane; 4: decanal; 5: β-citronellol; 6: nerol; 7: geraniol. b Endogenous accumulation of monoterpene glycosides in leaves. Representative liquid chromatography–mass spectrometry profile of linalyl diglycoside (linalyl β-primeveroside [linalyl-pri]), geranyl monoglucoside (geranyl β-d-glucopyranoside [geranyl-glc]) in leaves of the GUS or GES4 plants are presented. No geranyl β-primeveroside (geranyl-pri) or linalyl β-d-glucopyranoside (linalyl-glc) was detected in any of the samples assessed. b Quantitative data represent the means of linalyl-pri and geranyl-glc from wild-type (WT), GUS and GES (4, 7, 10) plants with standard errors (n = 3). Means indicated by different small letters are significantly different, based on a one-way ANOVA with post hoc Tukey’s HSD (P < 0.05). IS internal standard, FW fresh weight, NS not significant

Fig. 2

Phenotypes of GES plants for direct and indirect defenses towards pests. Wild-type (WT), GUS control or GES (4, 7, 10) plants were exposed to a herbivore (Spodoptera litura and Besimia tabaci) or fungous Botrytis cinerea. S. litura larval weight gained (a), the probability of survival of B. tabaci (b), and B. cinerea-induced lesions (c) were determined after 3 days and 4 days, respectively. Quantitative data represent the means ± standard errors (n = 23–28 for herbivores and n = 12–16 for fungus). Means were not significantly different (NS; P > 0.05), based on a one-way ANOVA. (d) Olfactory response of Nesidiocoris tenuis adult females when offered WT plants vs GUS plants or GES plants, and when offered WT + geraniol in triethyl citrate (TEC) vs WT + TEC. The figures in parentheses represent the numbers of predators that did not choose either odor source (“no choice” subjects). A replicated G test was conducted to evaluate the significance of attraction in each experiment (*, P < 0.05; NS P > 0.05)

Heat treatment

For heat stress treatment, the potted plants were incubated at 45 °C or 50 °C under the light condition (80 µE m−2 s−1). The experimental period was 14 h for heat susceptibility assays (Fig. 3 and Supplemental Fig. S3). For all of the physiological analyses, heat treatment of plants was performed for 1 h (and 2 h for metabolite analysis; Fig. 4 and Supplemental Fig. S5) before severe leaf damage was detected (see Fig. 3), to exclude indirect effects, for instance, due to cellular damage.
Fig. 3

Monoterpene alcohols affect plant heat susceptibility. Wild-type (WT), GUS control, GES and OS2 plants were incubated at 50 °C. Three replicates were carried out, and representative images are shown

Fig. 4

Heat maps and hierarchical clustering of metabolites extracted from leaves of wild-type (WT) and GES4 plants before and after heat treatment at 50 °C for 1 h. Heat maps of 73 metabolites at each time point were compared. Normalized data of metabolite concentrations are represented by the Z scale. The absolute metabolite concentrations are reported in Supplemental Table S1. Three biological replications of data were clustered and analyzed. ACC 1-aminocyclopropane-1-carboxylate

Biotic stress

Spodoptera litura (Fabricius) (Lepidoptera, ) was transferred to our laboratory in 2014 from a culture reared at the Sumika Technoservice Co. Ltd. (Takarazuka, Japan). The insects were reared on artificial diet (Insecta LF, Nihon Nosan Kogyo Ltd., Tokyo, Japan) in a climate-controlled room at 24 ± 1 °C. Third-instar S. litura larvae that weighed from 1.8 to 2.3 mg were reared on the shoots of potted plants in the above room conditions for 3 days. When a larva was dead or lost during assays, we excluded those samples, and final replicate analyses were conducted with 12–15 independent samples.

A young adult female of Bemisia tabaci (Gennadius) (Hemiptera: Aleyrodidae) B-type, within 2 h after its emergence, was placed on a leaf of a tobacco plant and covered with a polyethylene cylinder cage with mesh windows (height: 4 cm, diameter: 4.5 cm). The insect was observed every day, and its presence was recorded until it was dead. Fourteen replications were made using potted, independent plants of the same line.

Botrytis cinerea (strain IuRy-1) was grown on potato sucrose agar under black light at 25 °C for 1 week to promote the formation of conidia. For inoculation, 10 µL of conidial spore suspension (1 × 107 conidia mL−1, in 2.5% glucose) was spotted onto the left and right sides of the upper surface of a leaf of the potted plants. The inoculated plants were covered with a plastic sheet to enable the maintenance of humidity. The plants were placed in a growth chamber at 24 ± 1 °C for 4 days. The leaf lesion area was analyzed using ImageJ.

Y-tube olfactometer

The generalist predator Nesidiocoris tenuis (Reuter) (Hemiptera: Miridae) was reared, and Y-tube olfactometer assays were performed according to the method described in Rim et al. (2015). N. tenuis was introduced into the olfactometer with a Y-shaped wire (3.5 cm inner diameter, 13 cm long for each branch tube and 13 cm long for the main tube). Briefly, a healthy WT plant and a GES4, GES7 or GES10 plant, or a WT plant and a GUS plant, each in a pot, were placed in one volatile source bottle each, and used for the bioassays to compare the attractivity of their volatiles for the predators. Otherwise, 10 mg of geraniol in 1 g of triethyl citrate (TEC) in a 4 ml glass vial and a potted WT plant, or a control glass vial containing 1 g of TEC and a potted WT plant, were placed in the respective source bottles. N. tenuis adult females were naïve to tobacco plants and GES products. Predators that did not make a choice within 5 min (“no choice” subjects) were excluded from the statistical analysis. The orientation of the odor source containers relative to the olfactometer arms was changed after every five bioassays. Assays using 20 predators were carried out as a single replicate in a day. Four replicates using four independent plants were carried out on different days (so 80 predators in all were assessed). The experiments were performed in a climate-controlled room (24 ± 1 °C).

Measurement of photosynthetic electron flow

Plants grown for 6 weeks (see above) were dark-adapted for 20 min before chlorophyll fluorescence measurements. Measurements were made at 24 ± 1 °C on the upper surface of the fully mature leaves (leaf positions #7) using a photosynthesis yield analyzer (MINI- PAM, Walz, Effeltrich, Germany).

Determination of leaf chlorophyll and carotenoid contents

Chlorophylls a and b, and carotenoids were extracted from leaves (100 mg) with 80% acetone and centrifuged (12,000 g, 15 min, 4 °C). The supernatant was used for measurement according to the method in Wellburn (1994).

PVOC analysis

PVOCs emitted from the potted tobacco plants were collected using the volatile collection system of GERSTEL-Twister [polydimethylsiloxane (PDMS) coated stir bar, film thickness 0.5 mm, 10 mm length, GERSTEL GmbH & Co. KG, Mülheim an der Ruhr, Germany]. PVOCs from potted plants were collected on Twister in a closed glass container (2 L) in a laboratory room (24 ± 1 °C, light intensity 80 µE m−2 s−1) for 1 h. n-Tridecane (0.1 μg) was added to the glass container as internal standard. The adsorbed volatiles were analyzed using a 6890/5972 gas chromatography/mass spectrometry (GC–MS) system (Agilent Technologies, Santa Clara, CA, USA) with an HP-5MS capillary column (Agilent Technologies), equipped with a thermo desorption system (TDS), a cooled injection system (CIS), and a cold trap system (CTS) (GERSTEL GmbH & Co. KG). Headspace volatiles collected on a Twister were released from the PDMS by heating in the TDS at 280 °C for 4 min, within a He flow. The desorbed compounds were collected in the CIS at − 130 °C, and then the collected compounds were released from the CIS by heating (230 °C). The desorbed compounds were collected again in the CTS at − 50 °C, and then flash heating of the CTS (200 °C) provided sharp injection of the compounds into the capillary column of the gas chromatograph to which the CTS was connected. GC-oven temperature was programmed to rise from 40 °C (9 min hold) to 280 °C at 10 °C min−1. All of the PVOCs were identified and quantified by comparing their mass spectra and retention times with those of authentic compounds. Replicate analyses were conducted with four to five independent leaf samples.

Quantification of endogenous aroma glycosides in transgenic tobacco plants

Endogenous amounts of geranyl β-d-glucopyranoside (geranyl-glc), linalyl β-d-glucopyranoside (linalyl-glc), geranyl β-primeveroside (geranyl-pri) and linalyl β-primeveroside (linalyl-pri) in tobacco plants were analyzed according to (Ohgami et al. 2015) with minor protocol modifications. Tobacco leaves were finely chopped, crushed in an auto miller crusher (TK-AM5, TAITEC, Saitama, Japan), suspended in 80% methanol and centrifuged (12,000 g, 20 min, 4 °C). Methanol in the supernatant was removed in vacuo and the aqueous phase was separated with n-hexene. The aqueous layer was concentrated in vacuo, dissolved in distilled water and purified with Oasis HLB solid phase extraction sorbent (Waters, MA, USA). The glycosidic fractions were concentrated in vacuo and dissolved in distilled water prior to liquid chromatography/mass spectrometry (LC–MS) analysis. LC–MS analysis was performed using the LCMS-2020 system (Shimadzu, Kyoto, Japan) equipped with an InertSustain AQ-C18 reversed phase column (2.1 mm i.d. × 150 mm, 5 μm, GL Science, Tokyo, Japan), and the following electrospray operating conditions were used: dry gas 1.5 l/min, capillary voltage 1.5 kV, dry gas temperature 250 °C. The endogenous aroma glycosides were quantified with the sulfide indole motility (SIM) negative mode with gradient elution with aqueous formic acid (0.1%, v/v) as solvent A and acetonitrile as solvent B at a flow rate of 0.2 ml/min at 40 °C. The gradient condition started with isocratic conditions of 9% solvent B for 30 min and then increased up to 24% solvent B for 5 min and kept at 24% for 35 min. The m/z 361 ion for geranyl-glc (tR 57.4 min) and linalyl-glc (tR 54.2 min), and the m/z 471 ion for geranyl-pri (tR 53.0 min) and linalyl-pri (tR 51.0 min) were analyzed.

Metabolite analysis

Metabolites were extracted using the method of Takahashi et al. (2014). Twenty milligrams of freeze-dried leaves were ground to a fine powder using a Micro Smash M100 (Tomy, Tokyo, Japan) and homogenized at 4 °C for 10 min with ice-cold 50% (v/v) methanol containing 10 μM 1,4-piperazineethanesulfonate and methionine sulfone as internal standards for quantification. After centrifugation at 20,000 × g for 5 min, the supernatant was filtered through a 3-kDa-cutoff filter (Millipore, Billerica, MA, USA) by centrifuging at 16,000 × g for 60 min, and the filtrates were used for metabolite analysis. Organic acids, amino acids, phosphates, and other metabolites were separated with a ZORBAX Eclipse plus-C18 column (250 mm × 4.6 mm; Agilent Technologies, Santa Clara, CA, USA) and sugars were separated with a Hypercarb column (4.6 × 100 mm; Thermo Scientific, Waltham, MA, USA) according to the method of Takahashi et al. (2017). Metabolite detection and quantification was performed using a qTOF-MS quadrupole time-of-flight mass spectrometry instrument (Agilent Technologies). Accuracy was verified with reference standard compounds, and the absolute concentrations were calculated by comparing the peak areas with known concentrations of reference compounds using Agilent MassHunter Quantitative Analysis Software (Rev. B.05.02).

For hierarchical clustering analysis, three biological replications of metabolome data were normalized by a Z score transformation and clustered using Ward’s method with the squared Euclidean distance as a dissimilarity measure using the Statistical Package for the Social Sciences (SPSS v.10.0). Heatmaps were created using Microsoft Excel 2011 software.

Determination of invertase activity

Total proteins were extracted from leaves (about 80 mg) using extraction buffer (50 mM Tris–HCl (pH 8.0), 150 mM NaCl, 0.5% TritonX-100). Invertase activity of the extracts was assayed at 37 °C in an assay buffer (pH 4.9), using an Invertase Activity Colorimetric Assay Kit (BioVision, Inc., Milpitas, CA, USA) following the manufacturer’s protocol. The absorbance was measured at 570 nm. Protein concentration of the extracts was determined according to the method of Bradford (1976).

Light and transmission electron microscopy

Leaf segments were sampled from tobacco WT and GES4 plants before and after heat treatment at 50 °C for 1 h and fixed in 2.5% glutaraldehyde in 0.1 M sodium cacodylate buffer (pH 7.4) for 2 h at room temperature under vacuum, and overnight at 4 °C. Samples were then washed twice in sodium cacodylate buffer (10 min each wash) and postfixed for 1 h at RT in 1% osmium tetroxide in the same buffer. Leaf segments were dehydrated in a graded ethanol series (30, 50, 70, 90, and 100%, 15 min for each wash), rinsed twice in acetone 100% and embedded in Epon/Araldite resin (8 g of Araldite 502, 12.5 g of Epon 812, 24 g of dodecenylsuccinic anhydride, eight drops of the Epoxy embedding medium accelerator DMP30).

For light microscopy, thick sections of resin-embedded leaf tissues (0.5 µm thick) were cut with a glass knife using an ultramicrotome, stained with 1% toluidine blue, and examined using a Nikon Eclipse Ci light microscope equipped with a digital camera CCD 5.1 Mpixels. For transmission electron microscopy, thin sections (70 nm thick) were cut with a diamond knife (Diatome), stained with uranyl acetate and lead citrate, and examined using a Philips CM10 Transmission Electron Microscope.

Measurement of reactive oxygen species (ROS)

Leaf discs (0.785 cm2) prepared from the fresh leaves of the potted plants were incubated overnight in distilled water in a 96-well titer plate (one disc per well) (Schwessinger et al. 2011; Smith and Heese 2014). After water was replaced with fresh distilled water (25 µL per well), the plate was incubated at room temperature (24 °C) or at 50 °C under the light condition (80 µE m−2 s−1) for 15 min. ROS production was determined using a luminol-based assay system in 50 µL of 10 mM MES buffer (pH 6.0) containing 0.2 µM luminol (Wako Pure Chemical Industrials, Ltd., Osaka, Japan) and 20 µg L−1 horseradish peroxidase (Sigma-Aldrich, St. Louis, MO, USA). Luminescence was immediately measured using a 1420 Luminescence Counter ARVO Light (PerkinElmer, Boston, MA, USA), and the signal integration time was 1 s.

RNA extraction, cDNA synthesis and quantitative polymerase chain reaction (qPCR)

Approximately 100 mg of leaf tissues were homogenized in liquid nitrogen, and the total RNA was isolated and purified using Sepasol®-RNA I Super G (Nacalai Tesque, Kyoto, Japan) following the manufacturer’s protocol. Reverse transcription-qPCR was performed according to the method described in Uemura et al. (2018). Primers used are listed in Supplemental Table S1. Relative transcript abundances were determined after normalization of the raw signals with the housekeeping transcript abundance of an actin gene (GQ281246). Replicate analyses were conducted with 6–9 independent samples. We did not use samples or data when sufficient amounts or quality of RNA (> 83 ng µL−1; A260/A280 = 1.9–2.1) were not isolated from leaves or when abnormal quantification cycle (Cq) values for the actin gene were obtained.

Statistical analyses

A replicated G test was conducted to evaluate the data in Fig. 2d. Data sets from Fig. 4 were analyzed using Student’s t test. For all other data sets, we performed one-way ANOVA with post hoc Tukey’s HSD using online programs (http://astatsa.com/OneWay_Anova_with_TukeyHSD/ and http://vassarstats.net/anova1u.html).

Results

Generation of transgenic tobacco plants constitutively producing geraniol and other monoterpene alcohols

We prepared gene constructs consisting of ObGES inserted downstream of the constitutive CaMV 35S promoter, and generated a set of independent transgenic tobacco lines (GES plants) (Supplemental Fig. S1). Three representative examples exhibiting substantial geraniol emission are shown in Fig. 1a: the emission levels of GES plants ranged between 6.6 nmol g−1 fresh weight (FW) h−1 (GES10) and 10.0 nmol g−1 FW h−1 (GES4 and GES7), whereas control plants (WT and GUS-transformed plants) showed no emission of this monoterpene (Fig. 1a). Nerol [cis-isomer of geraniol], β-citronellol and linalool were also detected in the headspace of all GES plants only at lower levels (~ 2% of the geraniol emission level). Linalool, the other enzymatic product of GES (Fischer et al. 2013), was similarly emitted from all plants.

Leaves of GES plants also exhibited the accumulation of linalyl diglycoside (linalyl-pri) and geranyl monoglycoside (geranyl-glc); however, these compounds were completely absent in control plants (Fig. 1b).

Phenotypic characterization of GES plants

GES plants and control plants showed similar growth, development and vegetative stage, resulting in similar numbers of flowers and harvested seeds (Supplemental Figs. S2a and S2b). These observations were in agreement with the photosynthetic activity, total chlorophyll and carotenoid contents in mature leaves, which were not different between GES and control plants (Supplemental Figs. S2c, S2d, and S2e).

We next assessed phenotypic responses of GES plants towards biotic stresses. To first evaluate the direct defense traits of GES plants against biotic stresses, we focused on the generalist herbivores S. litura and silverleaf whitefly (B. tabaci), and the necrotic pathogen B. cinerea, which are frequently used to study plant defense responses (Mitsunami et al. 2014; Shpakovski et al. 2017; Li et al. 2017). As shown in Fig. 2a–c, GES plants did not show visually detectable differences in either the performance of larvae of the herbivores or necrotic lesions made by the pathogen, in comparison to control plants.

We then assessed olfactory responses of herbivore enemies because monoterpenes often play an important role in indirect defenses by attracting such enemies (Arimura et al. 2016). For these assays, we focused on N. tenuis, a carnivorous predator of various herbivore pests such as S. litura (Rim et al. 2015), thrips (Itou et al. 2013) and spider mites (Urbaneja et al. 2003). N. tenuis adults did not discriminate between PVOCs from GUS plants and WT plants, but preferred PVOCs from GES plants over those from WT plants (Fig. 2d). These olfactory responses were confirmed by assays using the vapor of synthetic geraniol (releasing 27 nmol h−1) + WT plant vs WT plant, which resulted in significant attraction of N. tenuis towards the geraniol compound (Fig. 2d).

Next, we assessed the ability of GES plants to withstand environmental stresses. The potted GES plants were more susceptible than control plants to heat treatment at 50 °C but not at 45 °C (Fig. 3 and Supplemental Fig. S3), indicating that the GES plants suffered from the extremely high temperature (> 50 °C). Moreover, GES plants grown aseptically on 1/2 Murashige and Skoog medium supplemented with 3% sucrose at 25 °C for 2 weeks were susceptible to cold treatment at 1 °C for 1 week in comparison to control plants following the same treatment (Supplemental Fig. S4).

We then assessed whether the susceptibility of GES plants to severe heat stresses was caused either by the geraniol overexpression or by other factors: e.g., lack of terpene precursors such as geranyl diphosphate (GPP) and/or isopentenyl diphosphate (IPP) due to competition with other tobacco terpene-derived primary metabolites, including carotenoids and phytohormones such as gibberellin and abscisic acid. To assess this, we examined the susceptibility of transgenic tobacco plants (OS2) producing the monotepene β-ocimene (Muroi et al. 2011). This line appeared to release 3 nmol g−1 FW h−1 of β-ocimene, as similarly shown in the geraniol levels of GES lines. The OS2 plants, however, exhibited heat tolerance to 50 °C similarly to control plants for up to 14 h. A similar result was also obtained by cold treatment at 1 °C, at which temperature no growth difference was observed between OS2 and control plants.

Metabolic changes in GES plants

Along with examining the physiological responses of GES plants to heat stress, we carried out a targeted metabolome analysis in leaflets of GES4 and GUS control plants before or after heat treatment at 50 °C (Fig. 4, Supplemental Fig. S5, and Supplemental Table S2).

We identified 73 metabolites, including amino acids, organic acids, phosphates, sugars, and other low-molecular weight compounds listed in Supplemental Table S1. A comparison between GES4 and GUS plants either before or after heat treatment showed an array of differently accumulated metabolites. Before heat treatment, the levels of Cys and carnosine were significantly higher in the leaves of GES4 plants than in those of GUS plants. After 1 h of heat treatment, numerous metabolites, including caffeic acid, sucrose, glucose 6-phosphate (G6P), Orn, His, adenosine, and guanosine, were highly accumulated in GES4 leaves. In contrast, glucose (Glc), fructose (Fru), and γ-aminobutyric acid (GABA) were accumulated more highly in GUS leaves. After 2 h of heat treatment, other metabolites, including pantothenate and some tricarboxylic acid cycle intermediates (citrate and isocitrate), increased in GES4 plants, while inositol increased in GUS leaves.

Since heat treatment resulted in modulation of the ratio of sucrose (Suc)/Glc/Fru levels between GES plans and GUS plants after only 1 h of heat treatment, we determined the enzyme activity levels of soluble invertase, an enzyme involved in the hydrolysis of Suc into Glc and Fru in the cytosol. As expected, the enzyme activity was slightly lower in the leaves of GES plants in comparison to those of control plants at 24 °C, whereas 1 h of heat treatment significantly decreased the enzyme activity in GES plants with respect to control plants (Fig. 5).
Fig. 5

Invertase activity in leaves of wild-type (WT), GUS and GES (4, 7, 10) plants before and after heat treatment at 50 °C for 1 h. Data are shown as the mean ± standard error (n = 6). Means indicated by different small letters are significantly different, based on a one-way ANOVA with post hoc Tukey’s HSD (P < 0.05)

Furthermore, to observe the cytological changes during heat treatment, light and transmission electron microscopic observations were conducted. Light microscopy analysis showed, in general, a large number of starch granules inside the chloroplasts and lipid globules both inside and outside the chloroplasts in WT leaf cells at room temperature (Fig. 6). After 1 h of heat treatment at 50 °C, the size of lipid globules increased, whereas that of starch granules decreased. GES4 leaf mesophyll cells were always characterized by a lower number and smaller size of starch granules in the chloroplasts and by the presence of myelin-like bodies (Fig. 7). After heat treatment, GES4 leaves showed large lipid globules both inside and outside the chloroplasts, which also showed fewer thylakoids than control plants (Fig. 7).
Fig. 6

Light microscopy thick sects. (0.5 µm) of tobacco leaf cells. ac Wild-type (WT) plants incubated at room temperature (24 °C) (WT 24 °C; a and b metric bar 37.5 μm; c metric bar 25 μm) showed chloroplasts (single arrows) filled with starch granules (double arrow). df WT plants exposed to heat treatment at 50 °C for 1 h (WT 50 °C; d and e metric bar 37.5 μm; f metric bar 25 μm) show evident signs of heat stress with reduced chloroplast (single arrows) and starch granule size (double arrow). giGES4 plants incubated at room temperature (24 °C) (GES 24 °C; g and h metric bar 37.5 μm; i metric bar 25 μm) show the absence of starch granules (double arrow) in the chloroplasts (single arrow). (jl) GES4 plants exposed to heat stress at 50 °C for 1 h (GES 50 °C; j and k metric bar 37.5 μm; l metric bar 25 μm) show a consistent reduction of chloroplast size (single arrow). Microscopic analysis of GUS plants showed similar cellular conditions as those of WT plants at both 24 °C and 50 °C

Fig. 7

Transmission electron microscopy of thin sections (70 nm) of tobacco leaf cells. ac Wild-type (WT) tobacco plants incubated at room temperature (24 °C) (a metric bar 3 μm; b, metric bar 1.75 μm; c, metric bar 1.2 μm) show large starch granules (single arrows) inside the chloroplasts along with large plastoglobules (double arrows). df WT plants exposed to heat treatment at 50 °C for 1 h (d metric bar 2 μm; e metric bar 2.9 μm; f metric bar 0.8 μm) show the reduction of starch granules (single arrows) and the appearance of osmiophilic deposits (asterisks) outside the chloroplasts. gjGES4 plants incubated at room temperature (24 °C) (g metric bar 4.4 μm; h metric bar 1.44 μm; i metric bar 3 μm; j metric bar 2 μm) show chloroplasts with a reduced number of starch granules (single arrows) and the presence of dark osmiophilic cytosolic structures resembling myelin bodies (double asterisks). knGES4 plants exposed to heat treatment at 50 °C for 1 h (k metric bar 2 μm; l metric bar 2.5 μm; m metric bar 1.2 μm; n metric bar 1.8 μm) show evident signs of chloroplast degeneration (kl single arrows) and the production of large plastoglobules (m and n, double arrows). Microscopy analysis of GUS control plants showed similar cellular conditions to those of WT plants at both 24 °C and 50 °C

Oxidative stress associated with heat susceptibility

To evaluate whether the heat susceptibility of GES plants depends on the ability of GES products to act as potential cellular agents for apoptosis and oxidative stress induction (Ashida et al. 2002), we measured the endogenous level of ROS. In both GES and control plants, a similar increase of ROS was detected in leaves after heat treatment (Supplemental Fig. S6), indicating that GES products do not strongly affect the oxidative stress level of leaves exposed to heat stress.

Gene regulation associated with heat susceptibility

Next, we determined the transcript accumulation levels in GES and control plants under normal and heat stress conditions. HSPs and their regulators [heat shock factors (HSFs)] are ubiquitous proteins that play key roles in protecting against heat stress-induced denaturation of other proteins (Feder and Hofmann 1999). Transcripts of two representative heat shock-related genes, HSP101 and HSF24-like, were similarly accumulated in the leaves of GES and control plants (Fig. 8). In contrast, the transcript level of WRKY33, the gene encoding one of the WRKY transcription factors involved in plant heat responses (He et al. 2016), was reduced in all of the GES plants before heat treatment when compared to control plants, whereas after the heat treatment only GES4 plants showed a significantly lower level of WRKY33 transcript compared to controls (Fig. 8).
Fig. 8

Transcript levels of genes for heat shock protein (Hsp101), heat shock factor (Hsf24-like), aquaporins (PIP1 and PIP2), and WRKY transcription factor (WRKY33) in leaves of wild-type (WT), GUS and GES (4, 7, 10) plants before and after heat treatment at 50 °C for 1 h. Data are shown as the mean ± standard errors (n = 6–9). Means indicated by different small letters are significantly different, based on a one-way ANOVA with post hoc Tukey’s HSD (P < 0.05). NS not significant

We also examined aquaporins, based on the possibilities of altered membrane permeability to water in GES plant leaves. The aquaporin-coding gene PIP2 showed a lower transcript level at room temperature in GES plants when compared to control plants. Heat treatment at 50 °C resulted in decreased PIP2 transcript levels in all plants. PIP1, another aquaporin gene, however, showed no difference of transcript levels between control and GES plants, irrespective of temperature conditions.

Discussion

Terpenes frequently confer on plants direct and indirect defenses against herbivores and pathogens, and moreover, a certain suite of terpenes may also play key roles in plant growth, development and responses to the abiotic environment (Arimura et al. 2016). In some transgenic plants heterologously producing terpene(s), however, biosynthesis and accumulation of terpenes do not affect the viability or growth, in comparison to those of WT plants. This is the case, for example, in Arabidopsis transgenic plants overexpressing GES. However, in some other plants, for example, grapevine (Vitis vinifera), terpenes do affect the viability or growth (Fischer et al. 2013). Our findings here showed that tobacco GES plants showed no phenotypic changes of viability or growth as compared to WT plants when they were incubated under the normal growth conditions at 24 °C. When grown under these conditions, GES plants exhibited a superior ability to produce indirect defense against herbivores, which enabled the plants to attract the predator N. tenuis (Fig. 2d). To the best of our knowledge, geraniol has not previously been reported as an airborne cue that attracts herbivorous predators.

In contrast, compared with the control plants, the GES plants suffered notably from heat susceptibility. Our comparative analyses involving tobacco plants heterologously producing another monoterpene hydrocarbon (β-ocimene) confirmed our conclusion that GES products and their glycoside conjugates, rather than changes in terpene metabolism due to overproduction of monoterpenes, are potential causes of the heat susceptibility of the GES plants (Fig. 3). Moreover, although we observed that β-ocimene is not effective at altering the thermotolerance ability of tobacco, this hydrocarbon monoterpene plays rather important roles in some plants’ thermotolerance ability, as β-ocimene can function as an effective antioxidant in tropical forests (Jardine et al. 2017). Presumably, because the desert plant tobacco is inherently resistant to thermal stress, overproduction of β-ocimene does not confer greater thermotolerance ability to tobacco.

GES plants produce large amounts of free and sugar-bound monoterpenes. Both free and glycosylated monoterpenes are considered to be mostly responsible for plants’ increased susceptibility to heat stress. Our findings do not exclude the possibility that their glycosylated products are effective in causing the heat susceptibility and that the consumption of uridine diphosphate (UDP) sugar might cause the susceptibility and possibly a disordered sucrose/glucose level. This hypothesis would be reflected by the fact that transgenic tobacco plants heterologously producing β-ocimene did not show the heat susceptibility. β-ocimene cannot be conjugated with sugars due to the lack of hydroxyl groups.

Moreover, with regards to the correlation between terpene metabolic engineering and changes in osmotic cell status, our metabolome and ultrastructural analyses indicate abnormal sugar metabolism in GES plants. First, we found a possible correlation between the heat susceptibility and invertase activity (Fig. 5). The role of this enzyme in plant stress responses has been well-clarified using transgenic plants (Albacete et al. 2014; Essmann et al. 2008; Liu et al. 2016; Sun et al. 2014). Reduced invertase activity implies impaired sugar metabolism with alterations in endogenous Suc, Glc and Fru levels, as observed in GES plants exposed to heat stress. We hypothesize that the sugar and energy metabolism, as a consequence of Glc and Fru phosphorylation, could be suppressed in the GES plants due to suppressed invertase activity under heat stress, presumably leading to a heat susceptibility trait. Regarding this issue, intriguingly, our metabolome data even showed a higher G6P level in GES plants than in WT plants at 1 h after heat treatment (Fig. 4 and Supplemental Fig. S5). Although the regulation of sugar metabolism is highly complex in leaves, all of these metabolites should eventually contribute to starch accumulation. This accords with our ultrastructural observations confirming that GES plants produce a lower than normal amount of primary starch, as shown by the significant reduction of starch granules in their mesophyll chloroplasts (Figs. 6 and 7).

Other metabolic processes should also be considered as contributors to the heat susceptibility. Cys, GABA and inositol were significantly reduced in GES4 leaves compared to GUS control leaves (Supplemental Fig. S5), as were some low-molecular weight metabolites, including amino acids, which function as energy sources and signaling molecules in environmental stress tolerance responses of several plant taxa (Shulaev et al. 2008; Wind et al. 2010).

Chaperone proteins (e.g., HSPs and HSFs) as well as ROS production are frequently responsible for heat responses (Jacob et al. 2017; Pospíšil 2016) but are not likely to be involved in the heat susceptibility of GES plants (Fig. 8 and Supplemental Fig. S6). Rather, altered permeability of the cell membrane to water in plants may involve changes in transcript accumulation of aquaporins involved in integral membrane proteins mainly facilitating the transport of water between cells (Agre 2006), and our data showing the constitutively lower transcript level of PIP2 (an aquaporin gene) in GES leaves supports the possibility of impaired water transport in these plants. Likewise, WRKY33, involved in plant heat responses (He et al. 2016), was transcribed at a lower level in the GES plants than in the control plants before heat treatment. We therefore deduced that the constitutively lower transcript levels of this WRKY gene play a significant role only in the early heat susceptibility. Taken together, these results were in accord with the finding in a recent report that cell wall invertase plays a key role in heat stress tolerance through suppression of ROS-independent programmed cell death as well as enhanced Suc import and catabolism, and expression of HSP genes in tomato fruits (Liu et al. 2016). We therefore hypothesize that invertase plays a master role in the acquisition of heat tolerance through sugar metabolism and transcriptional regulation of genes involved in cellular homeostasis.

We also propose that increased monoterpene alcohol levels and thermal stress result in the impairment of plant thermal tolerance. Beyond the physiological changes in GES plants shown here, it remains to be understood how the vapor pressure of GES products might change at higher temperatures and whether this increased volatility might play a role in the temperature dependence of their effects. Moreover, although it has not been assessed here, membrane stabilization modulated by volatile terpenes may also contribute to heat tolerance.

Concluding remarks

Metabolic engineering of terpenes within plants is expected to provide agricultural and pharmacological benefits due to the multifaceted functionality of terpenes in human health and in the environment (Lange and Ahkami 2013). Nevertheless, making use of the powerful and constitutive 35S promoter for monoterpene engineering may be tricky because this promoter does not restrict the timing and place for de novo biosynthesis, and this may cause detrimental effects on the host plants. One of the solutions is to use promoters that allow the specific expression of the target gene, for instance, in glandular trichomes and resin ducts, as shown in (Ennajdaoui et al. 2010; Zhang et al. 2018). Although we still need to unravel various puzzling effects of monoterpenes, such multidirectional outcomes should be considered for the development of true monoterpene engineering.

Author contribution statement

AH, MN, MEM and GA designed the experiments; AH, MN, SS, HR, HT, MI, TU, TO, RO, MEM and GA performed experiments; MN, MEM and GA prepared the initial manuscript draft; MN, HR, HT, TO, MEM and GA produced the final manuscript version. All of the authors approved the final manuscript.

Notes

Acknowledgements

We would like to thank Dr. Eran Pichersky (University of Michigan) for kindly providing GES cDNA clones; and Dr. Ko Kato (Nara Institute of Science and Technology, Japan) for providing a plasmid containing Arabidopsis HSP terminator. This work was financially supported in part by a JSPS KAKENHI to GA (16K07407) and a MEXT Grant-in-Aid for Scientific Research on Innovative Areas to GA (18H04630 and 18H04786).

Supplementary material

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Supplementary material 1 (DOCX 22 kb)
425_2018_3054_MOESM2_ESM.pdf (2.3 mb)
Supplementary material 2 (PDF 2363 kb)
425_2018_3054_MOESM3_ESM.xlsx (20 kb)
Supplementary material 3 (XLSX 21 kb)
425_2018_3054_MOESM4_ESM.xls (81 kb)
Supplementary material 4 (XLS 82 kb)

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Copyright information

© Springer-Verlag GmbH Germany, part of Springer Nature 2018

Authors and Affiliations

  • Ashita Hamachi
    • 1
  • Masahiro Nisihara
    • 2
  • Shiori Saito
    • 1
  • Hojun Rim
    • 1
  • Hideyuki Takahashi
    • 2
  • Monirul Islam
    • 3
  • Takuya Uemura
    • 1
  • Toshiyuki Ohnishi
    • 4
    • 5
  • Rika Ozawa
    • 6
  • Massimo E. Maffei
    • 3
  • Gen-ichiro Arimura
    • 1
    Email author
  1. 1.Department of Biological Science and Technology, Faculty of Industrial Science and TechnologyTokyo University of ScienceTokyoJapan
  2. 2.Iwate Biotechnology Research CenterKitakamiJapan
  3. 3.Department of Life Sciences and Systems Biology, Plant Physiology UnitUniversity of TurinTurinItaly
  4. 4.College of Agriculture, Academic InstituteShizuoka UniversityShizuokaJapan
  5. 5.Research Institute of Green Science and TechnologyShizuoka UniversityShizuokaJapan
  6. 6.Center for Ecological Research, Kyoto UniversityOtsuJapan

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