Plant Physiology and Biochemistry 135 (2019) 224–232 Contents lists available at ScienceDirect Plant Physiology and Biochemistry journal homepage: www.elsevier.com/locate/plaphy Research article Resource allocation in response to herbivory and gall formation in Linaria vulgaris T Ana Sedlarević Zorića, Filis Morinaa,b,∗, Ivo Toševskic,d, Tomislav Tostie, Jelena Jovićd, Oliver Krstićd, Sonja Veljović-Jovanovića a Institute for Multidisciplinary Research, University of Belgrade, Belgrade, Serbia Biology Centre of the Czech Academy of Sciences, Institute of Plant Molecular Biology, České Budejovice, Czech Republic c CABI, Delémont, Switzerland d Institute for Plant Protection and Environment, Department of Plant Pests, Zemun, Serbia e University of Belgrade, Faculty of Chemistry, Belgrade, Serbia b A R TICL E INFO A BSTR A CT Keywords: Carbohydrates Galls Herbivory Linaria vulgaris Rhinusa pilosa Source-sink interactions Trehalose 6-phosphate Trehalose and its precursor, trehalose 6-phosphate (T6P), are essential regulators of plant response to abiotic and biotic stress. Here we used the specific host-insect interaction between Linaria vulgaris (Plantaginaceae) and stem-galling weevil, Rhinusa pilosa (Mecinini, Curculionidae) with the aim to distinguish carbohydrate allocation patterns in response to herbivory, gall formation (G1, 24 h after oviposition), and gall development (G2, 7 days after oviposition) under controlled conditions. The hypothesis is that herbivory and galling induce distinct responses in both leaves and stems, and that shifts in carbon allocations are regulated by signaling sugars. Systemic response to herbivory was accumulation of T6P and maltose. The main feature of G1 in the stems was accumulation of trehalose, accompanied by increased T6P, turanose and glucose content, oppositely to the leaves. In G2, galls had 3-folds higher weight than controls, with further accumulation of fructose, glucose, turanose, and total water-insoluble carbohydrates (TIC), while the sucrose/hexose ratio decreased. Analysis of fast chlorophyll fluorescence kinetic (OJIP) transients in G2 showed a slight decrease in quantum yield of electron transport flux from QA to QB, and towards photosystem I acceptor side, correlated with the decreased content of photosynthetic pigments and hexoses accumulation. Redistribution of photosynthates, and accumulation of T6P were induced in response to herbivory, indicating its signaling role. The results support the hypothesis that R. pilosa can induce plant reprogramming towards the accumulation of beneficial carbohydrates in developing gall by mechanisms which include both T6P and trehalose. 1. Introduction Interactions between host plants and gall-inducing insects are remarkable examples of structural, physiological, and chemical manipulations of the plant cells resulting in abnormal plant outgrowths – galls (Tooker and De Moraes, 2008; Giron et al., 2016). Gall initiation and development are manifested by the cell hypertrophy and tissue hyperplasia, as well as by changes in the chemical composition (Giron et al., 2016; Oliveira et al., 2016; Carneiro et al., 2017). Galls are mostly initiated in the zones of active cell division (Weis et al., 1988), such as in the apical meristems (Fay et al., 1996; Barnewall, 2011; Barnewall and De Clerck-Floate, 2012). Recent studies have shown that leaf gall tissue has altered photosynthetic activity compared to the uninfected one, accompanied by ∗ accumulation of soluble sugars and changes in starch content (Oliveira et al., 2011; Castro et al., 2012; Carneiro et al., 2014; Huang et al., 2014). However, most of the studies describe leaf or root galls, while reports on biochemical modifications in stem galls in herbs are rare (St John and Shorthouse, 2000; Raman et al., 2006; Marini-Filho and Fernandes, 2012). Additionally, a number of these studies reflects the changes in the composition of nutrients (C, N), while comparative analysis of soluble sugar profile during stem gall development are scarce (e.g. Agarrwal et al. (2014)). Moreover, specific soluble sugars may act as signaling molecules which regulate gene expression thus modulating growth from the cell to the whole plant level (Rolland et al., 2006; Lastdrager et al., 2014). In this respect, the emerging regulatory roles of sucrose, glucose, trehalose and its precursor trehalose 6-phosphate (T6P) have been described recently (Lastdrager et al., 2014; Xiong et al., 2013; Figueroa and Lunn, 2016). Corresponding author. Biology Centre of the Czech Academy of Sciences, Institute of Plant Molecular Biology, České Budejovice, Czech Republic. E-mail addresses: morina@umbr.cas.cz, filis@imsi.rs (F. Morina). https://doi.org/10.1016/j.plaphy.2018.11.032 Received 7 September 2018; Received in revised form 27 November 2018; Accepted 27 November 2018 Available online 28 November 2018 0981-9428/ © 2018 The Authors. Published by Elsevier Masson SAS. This is an open access article under the CC BY-NC-ND license (http://creativecommons.org/licenses/BY-NC-ND/4.0/). Plant Physiology and Biochemistry 135 (2019) 224–232 A.S. Zorić et al. Three major hypotheses for explaining the adaptive significance of gall induction and evolution of gall morphology are nutrition, enemy, and microenvironment hypotheses (Stone and Schönrogge, 2003). The nutritional hypothesis states that the advantage of galling herbivores is having a high-quality nutrient source with less plant defensive compounds than other feeding modes (Price et al., 1987; Crespi et al., 1997). The microenvironment hypothesis states that galls shield the insect from unfavourable abiotic conditions (excess ultraviolet radiation, temperature changes and desiccation) which are the major problems for larval development (Stone and Schönrogge, 2003; Price et al., 1987; Crespi et al., 1997; Miller et al., 2009). According to the enemy hypothesis, the gall protects the gall-inducer from its predators, parasitoids and pathogens (Stone and Schönrogge, 2003). Gall-inducing insects are mostly inert during the feeding stage (Price et al., 1987); thus, they are entirely dependent on the nutrient sink formed in the host plant tissue. Rhinusa pilosa (Gyllenhal, 1838) is a highly specific stem-galling weevil associated with yellow toadflax, Linaria vulgaris (Toševski et al., 2015). Based on its narrow host range, R. pilosa was selected for investigation as a potential agent for the biological control of yellow toadflax in North America (Gassmann et al., 2014). It is overwintering in the adult stage and appears in early spring when adults start to feed on young yellow toadflax shoots and then start to mate. Oviposition takes place in the apical region of the young growing shoots where eggs are deposited inside the stem pith tissue. Gall development is visible already 48 h after oviposition, and its growth is completed after about 10 days. The complete insect development, from egg to adult, is taking place inside the gall. Finally, the adults feed on the remaining gall tissue for two weeks and create an exit hole to leave the gall (Gassmann et al., 2014). In our previous study, we used the specific interaction of R. pilosa and L. vulgaris to reveal the mechanisms of gall initiation (Sedlarević et al., 2016). In this model system, gall induction has been attributed to female ovipositional fluid (Barnewall and De ClerckFloate, 2012), and our study indicated the existence of specific signaling phenolic compounds which might be responsible for successful gall initiation. Here, the relationship between L. vulgaris and R. pilosa has been used as a model system to answer the following questions: i) are there specific changes in carbohydrate profile induced by insect feeding, gall initiation and/or subsequent gall growth; ii) how are energy demands of the insect developing in the gall secured (manipulation of source-sink interactions by the galler) iii) how is the primary metabolism (chlorophyll fluorescence, photosynthetic performance) affected by gall formation. the insects were daily transferred to new plants to ensure they have enough food supplies and optimal conditions for mating. Subsequently, the 24 copulated females were separated from the males and transferred for oviposition to 24 new plants. These plants were young and actively growing, with the average stem height of 15 cm. The act of oviposition was registered by eye, and the number of oviposition events was subsequently counted and confirmed under a stereomicroscope (Leica MS5, Mannheim, Germany). Time and day of the oviposition event for each plant were recorded. For biochemical analysis, 20 stem sections were collected from 20 different control (C) or plants affected by herbivory (H) from the same apical region (upper 1/3) of the stem, at the beginning and at the end of the experiment. Out of 20 sections, leaves and stems from each two were pooled together, to obtain ten samples of control leaves and stems (CL and CS for leaves and stems, respectively) and ten samples of plants affected by herbivory (HL and HS for leaves and stems, respectively), that is five for analysis of pigments, and five for carbohydrate analysis. Gall tissue (apical region) for investigating the initiation phase (G1) was collected 12–24 h after oviposition and it was pooled in the same manner as controls (G1L and G1S for leaves and stems, respectively). For investigating the effects of gall development (growing phase, G2), 7 days after oviposition, 12 samples from 12 individual plants were collected and gall tissue was separated in halves, one half for pigment and the other half for carbohydrate analysis. All samples were kept at −80 °C for biochemical analysis. 2.2. Analysis of fast chlorophyll fluorescence kinetic transients Direct fast imaging of chlorophyll fluorescence was performed using a macroscopic fluorescence imaging system (FluorCam, closed version, Photon Systems Instruments, Brno, Czech Republic), equipped with a newly developed camera, software version and a measuring protocol that allowed direct measurement of fast fluorescence induction (OJIP), as described in more detail in a publication of this new system (Küpper et al., 2018). For these experiments, we used 12 control plants and 12 plants with developed galls (growing phase, G2). Leaves were placed in a measuring chamber, ventilated by a stream of water-saturated air at 24 °C, and gently pressed to the glass window by a moist cotton pad fixed with a nylon mesh as described in detail by Küpper et al. (2007). Stems and galls were measured without the chamber, after cutting the plant at the base and wrapping the lower part of the stems in a wet cotton pad. The obtained OJIP curves of dark-adapted samples were normalised by their Fp (equivalent to Fm as saturating irradiance was used) values after background subtraction. Excitation light was supplied by two light panels with blue actinic light (LEDs with 460 nm peak) and supersaturating flashes of 4000 μmol m−2 s−1, while another two panels supplied measuring flashes with optimized control electronics for fast kinetics and synchronization with the ultrafast camera. Measurements were conducted using a custom-made protocol according to Küpper et al. (2018). Images were taken through a band-pass filter (684 nm) every 250 μs in the fast beginning of the kinetics, and towards the slower parts the frame rate was gradually reduced to save memory space in the camera. Analysis was done usingthe FluorCam software and a description of the used OJIP-test parameters used in the study, according to Stirbet and Govindjee (2011), is given in Table 1. 2. Materials and methods 2.1. Plant material and experimental conditions Experiments were performed over three years, in the period 2016–2018. Linaria vulgaris plants with young actively growing stems were selected to ensure optimal conditions for gall initiation and formation. The plants were potted individually and separated into three groups: control plants (C), plants for herbivory experiments (H), and plants for oviposition trials (G1 for 24 h, and G2 7 days after oviposition). Each potted plant was caged with a capped, ventilated plastic cylinder (10 × 35 cm), and the soil surface was covered with coarse sand to reduce excessive moisture inside the cylinder and to make the weevils more visible, as previously described in Gassmann et al. (2014). All plants were kept at 21–24 °C with a 16 h/8 h day/night photoperiod, the maximal light intensity at noon was 200 μmol m−2 s−1 in a stepwise cycle, and humidity was kept at 60% to allow normal gall development. After hibernation, newly emerged adults (females and males) were placed in pairs on 24 plants for feeding and mating. These plants had visible signs of injury due to insect feeding already after one day, and they were used for analysis of herbivory effects. During one week, 2.3. Extraction and analyses of photosynthetic pigments Frozen leaf and stem samples were extracted in 80% acetone (1:10 w/v) and centrifuged at 10 000 g at room temperature, and then the pellets were re-extracted. Supernatants were pooled together, diluted in 80% acetone, and the absorbance was measured at 663 nm, 645 nm and 470 nm. Concentrations of chlorophylls a and b (Chl a and Chl b) and carotenoids (Car) were determined spectrophotometrically (Shimadzu UV-2501 PC, Kyoto, Japan) following the equations proposed by 225 Plant Physiology and Biochemistry 135 (2019) 224–232 A.S. Zorić et al. Table 1 Calculations and definitions of JIP parameters according to Stirbet and Govindjee (2011). φPo = JoTR/JABS=1- Fo/Fm φET2o= JoET2/JABS =1- Fj/Fm φRE1o= JoRE1/JABS=1- Fi/Fm ψET2o = JoET2/ JoTR =1- Vj ψRE1o= JoRE1/ JoTR =1- Vi δRE1o= JoRE1/ JoET2 =(1- Vi)/(1- Vj) Maximum quantum yield of primary PSII photochemistry Quantum yield of the electron transport flux from QA to QB Quantum yield of the electron transport flux until the PSI electron acceptors Efficiency/probability with which a PSII trapped electron is transferred from QA to QB Efficiency/probability with which a PSII trapped electron is transferred until PSI acceptors Efficiency/probability with which an electron from QB is transferred until PSI acceptors Lichtenthaler and Wellburn (1983) and the values expressed in μg pigment per g of fresh weight (FW). carbohydrate content was obtained by comparing the absorbance of the sample at 625 nm against a standard curve of glucose (0.1–1 mg ml−1). 2.5. Statistical analysis 2.4. Extraction and analysis of carbohydrates The significant effects of herbivory, gall initiation and development on carbohydrates, chlorophyll fluorescence parameters and photosynthetic pigments, were tested with the Mann-Whitney U test. Tukey's post hoc test was used to test for significant differences in the contents of individual carbohydrates, starch and TIC among different sample groups. Mann–Whitney U test and Tukey's post hoc test were conducted with IBM SPSS statistics software (Version 20.0, SPSS Inc., Chicago, IL, USA). Significance threshold value was set at 0.05. For soluble sugar analysis, frozen tissues were homogenized in liquid nitrogen and extracted in ultrapure water (1/6, w/v) for 30 min in an ultrasonic bath at 25 °C. Following centrifugation at 10 000 g for 10 min at 4 °C, the residues were re-extracted under the same conditions and the supernatants were pooled before analysis similarly as described in Vidović et al. (2015). Standard solutions of glucose, fructose and sucrose were prepared in Ultrapure TKA water at a concentration of 1000 ng ml−1, whereas standard solutions of the other analyzed sugars were prepared at a concentration of 100 ng ml−1. Trehalose 6-phosphate dipotassium salt was purchased from Sigma and prepared at the concentration of 100 ng ml−1. The quality control mixture used for monitoring instrument performance was prepared by diluting standards to concentrations in the range 0.9–100 ng ml−1 (depending on the concentration in the samples). Chromatographic separations were performed using a DIONEX ICS 3000 DP liquid chromatography system (Dionex, Sunnyvale, CA, USA) equipped with a quaternary gradient pump (Dionex, Sunnyvale, CA, USA). The carbohydrates were separated on a CarboPac®PA100 pellicular anion-exchange column (4–250 mm) (Dionex, Sunnyvale, CA, USA) at 30 °C. The mobile phase consisted of the following linear gradients (flow rate, 0.7 ml min−1): 0–5 min,15% A, 85% C; 5.0–5.1 min,15% A, 2% B, 83% C; 5.1–12.0 min, 15% A, 2% B, 83% C; 12.1 min, 15% A, 4% B, 81% C; 12.1–20.0 min 15%A, 4% B, 81% C; 20.0–20.1 min 20% A, 20% B 60% C; 20.1–30.0 min 20% A, 20% B 60% C; where A was 600 mM sodium hydroxide, B was 500 mM sodium acetate and C was ultrapure water. Before the analyses, the system was preconditioned with 15% A and 85% C for 15 min. Each sample (25 ml) was injected with an ICS AS-DV 50 Autosampler (Dionex, Sunnyvale, CA, USA). The electrochemical detector consisted of gold as the working and Ag/AgCl as the reference electrode. For the total water-insoluble carbohydrates (TIC) and starch determination, the pellets from soluble sugar extraction were washed in 80% ethanol at 80 °C, centrifuged, rewashed four times under the same conditions and dried at 40 °C. For starch analysis, the pellet was extracted in water and perchloric acid (1:1 v/v) for 20 min on ice, centrifuged, and re-extracted under the same conditions (modified protocol according to Hansen and Møller (1975)). The combined supernatants were used for starch determination by anthrone method (Sadasivam and Manickam (1992)): 50 μl of the sample was mixed with 50 μl water and 400 μl of anthrone (40 mg of anthrone in 20 ml of H2SO4), and the mix was shaken and heated for 8 min at 100 °C. The reaction was stopped by immersion in ice. The starch content was obtained by comparing the absorbance of the sample at 625 nm using glucose as a standard. For TIC analysis, the samples were extracted in 2.5 N HCl at 90 °C for three hours, according to Hedge and Hofreiter (1962) and Sadasivam and Manickam (1992). Following centrifugation, acid neutralization was performed using sodium carbonate. Total water-insoluble carbohydrate (TIC) content was determined by anthrone method (Sadasivam and Manickam (1992)):10 μl of samples were mixed with 90 μl water and 400 μl of anthrone (40 mg of anthrone in 20 ml of H2SO4), and the mix was shaken and heated for 8 min at 100 °C. The reaction was stopped by immersion in ice for 5 min. The 3. Results 3.1. Gall induction In this experiment, 100 L. vulgaris plants were submitted for gall induction using 24 mated R. pilosa females. During the three-week period, 87 galls were successfully initiated. The galled plants were subsequently collected for analysis following the time scale of oviposition as described in M&M. Average fresh weights of upper 1/3 of control stems (relative to oviposition localization) were 0.12 ± 0.01 g. As expected, after 24 h (G1) there was no significant increase in the stem weight, while 7 days after oviposition (G2) gall tissue increased to 0.35 ± 0.06gFW. 3.2. Soluble sugar analysis The content of total soluble carbohydrates in control stems (CS) was 4.4 ± 0.4 mg gFW−1, in the stems after herbivory (HS) 4.5 ± 0.7 mg gFW−1, while it gradually increased from 5.5 ± 0.6 mg gFW−1 in the gall initiation phase (GS1) to 7.0 ± 0.8 mg gFW−1 in the seven days-old galls (GS2). In control leaves (CL) the soluble carbohydrates content was about 6.0 ± 1.1 mg gFW−1, in the leaves of plants after herbivory (HL) it was 5.2 ± 0.2 mg gFW−1, while both in GL1 and GL2 leaves the content was decreased (3.9 ± 0.1 mg gFW−1 and 4.7 ± 0.1 mg gFW−1, respectively). The main carbohydrates detected were monosaccharides: arabinose, fructose, galactose, glucose, mannose, rhamnose, and xylose; disaccharides: maltose, sucrose, trehalose and its intermediary trehalose-6-phosphate, and turanose; trisaccharide raffinose, and sugar alcohols: galactitol, mannitol and sorbitol (Fig. 1). The content of carbohydrates in both CS and CL at the beginning and at the end of the experiment was not significantly changed (data not shown). In the stems, insect feeding (HS) induced a significant increase in the content of trehalose 6-phosphate (T6P), and maltose, accompanied by an elevated content of galactose (p < 0.01) compared to CS, while the content of galactitol decreased (Fig. 1). In initiating gall (G1S), the most striking change in the soluble carbohydrates composition was about 14-fold increased accumulation of trehalose, accompanied by significant transient accumulation of mannitol, raffinose and rhamnose compared to CS, HS and GS2 stems. In the same time, sorbitol content decreased, while sucrose level was slightly higher compared to CS (Fig. 1). The accumulation of glucose and turanose was correlated with gall development, reaching the 226 Plant Physiology and Biochemistry 135 (2019) 224–232 A.S. Zorić et al. Fig. 1. The content of soluble carbohydrates in L. vulgaris control stems (CS), stems after insect feeding (HS), in initiating gall (GS1, less than 24 h) and growing gall (GS2, 7 days after oviposition). Values represent the means ± SE (n ≥ 8). Different letters denote statistically significant differences between the treatments according to Tukey's post-hoc test. In the leaves of plants with initiating galls (GL1), a transient increase of maltose, mannitol, rhamnose and xylose was observed, while contents of fructose, galactose, galactitol, raffinose, sorbitol, trehalose and turanose declined. Sucrose content remained at the similar level as in HL. In both GL1 and GL2, the content of glucose slightly decreased, while sucrose levels slightly increased. Further, in the leaves of plants with developing gall (GL2) the levels of galactitol, fructose and turanose increased compared to GS1, while sorbitol and xylose remained at the similar level. highest values in GS2. Moreover, fructose content in GS2 galls increased significantly compared to both CS and GS1, while xylose remained at the higher level than in CS. Galactitol and galactose contents decreased in developing gall, as well as sucrose. In both GS1 and GS2, the level of T6P was significantly higher compared to CS (p < 0.01), although lower than in HS. In comparison to GS1, in GS2 the contents of trehalose, sucrose, mannitol and rhamnose decreased significantly. The content of arabinose significantly decreased only in the developing gall, while mannose was not significantly affected by herbivory or gall development. In the leaves of plants which were exposed to herbivory (HL), the level of T6P was the highest, similarly as in HS (p < 0.05 compared to CL). Further, an increase in arabinose, maltose, mannose, mannitol and rhamnose levels was observed along with a decrease in fructose, galactitol and raffinose compared to CL (Fig. 2), while sucrose content slightly increased. Contrary to the stem galls, the content of T6P was significantly lower in the leaves of plants with initiating and developing galls (GL1 and GL2) than in CL (p < 0.05 and p < 0.01, respectively). 3.3. The content of starch and total water-insoluble carbohydrates Starch content in control stems (CS) contributed about 80% to total water-insoluble sugars (TIC), while in galled stems (GS2) and in the stems which were exposed to herbivory this ratio decreased to about 45–50% (Fig. 3). Moreover, starch content significantly decreased after herbivory (HS) compared to CS, while in the galls no difference in starch content was detected. On the other hand, the content of TIC 227 Plant Physiology and Biochemistry 135 (2019) 224–232 A.S. Zorić et al. Fig. 2. The content of soluble carbohydrates in L. vulgaris control leaves (CL), leaves of plants after insect feeding (HL), and leaves of plants with initiating gall (GL1, less than 24 h) and growing gall (GL2, 7 days after oviposition). Values represent the means ± SE (n ≥ 8). Different letters denote statistically significant differences between the treatments according to Tukey's post-hoc test. 3.4. Analysis of OJIP chlorophyll fluorescence kinetic transients gradually increased with gall development by 1.5- and 1.7-fold compared to control stems (p < 0.05 for the significant difference between CS and GS2). In the leaves, there was no significant change in starch or TIC content due to feeding or gall development, and starch contributed about 45–50% to TIC (Fig. 3). The fast chlorophyll fluorescence is commonly used as a sensitive tool for detection of changes in photosynthetic electron flow and in the efficiency of photochemistry. The OJIP measurements showed no differences in the leaves regarding the gall development compared to Fig. 3. Content of starch (white bars) and total insoluble carbohydrates (TIC, grey bars) in L. vulgaris stems (S) and leaves (L) of control plants (CS, CL), plants after insect feeding (HS, HL), in plants with initiating gall (GS1,GL1, less than 24 h) and in plants with developing gall (GS2, GL2, 7 days after oviposition). Values represent the means ± SE (n ≥ 8). Different letters denote statistically significant differences between the treatments according to Tukey's posthoc test. 228 Plant Physiology and Biochemistry 135 (2019) 224–232 A.S. Zorić et al. Table 3 −1 Chlorophyll a and b (μg g−1 FW ), and their ratio, and carotenoid (Car, μg gFW ) content in L. vulgaris control leaves (CL) and stems (CS) and in leaves and stems of plants with seven days old galls (GL2 and GS2). Values represent the means ± SE (n ≥ 8) *p < 005, **p < 001 and ***p < 0001 indicate the statistically significant difference between control tissues to respective gall tissues. CL GL2 CS GS2 Chl a Chl b Chl a + b Chl a/b Car 1418 ± 200 1190 ± 403 179 ± 14 62 ± 3∗∗∗ 496 ± 68 520 ± 225 47 ± 5 24 ± 2∗∗ 1914 ± 265 1518 ± 734 226 ± 18 86 ± 5∗∗∗ 2.9 2.3 4.1 2.9 254 ± 22 202 ± 16 49 ± 4 28 ± 5∗ ± ± ± ± 0.1 0.6 0.6 0.8∗ content of total carotenoids (Car) decreased in the developing gall tissue. However, Car/Chl ratio increased from 0.38 in control stems to about 0.57 in gall tissue. In the leaves of plants with developing gall (GL2), a 2.3-fold decrease of carotenoid content compared to controls was observed, while Chl a and Chl b were not changed (Table 3). Over the course of the experiment (7 days) the level of photosynthetic pigments was not altered in control stems or leaves (data not shown). 4. Discussion 4.1. Possible roles of trehalose, T6P and hexoses on gall initiation and development The L. vulgaris - R. pilosa system presented here allows comparisons of carbohydrate profiles in the time course, during gall initiation and growth, as well as distinguishing the effect of herbivory using plants without galls after R.pilosa feeding. Gall initiation provoked immediate, transient accumulation of trehalose in the stems, accompanied by about 5-fold increased content of its intermediate, T6P (Fig. 1). In insects, trehalose is the primary sugar in the haemolymph (Tang et al., 2018); however, to our knowledge, there are no reports on its content in ovipositional fluid. Analysis of R. pilosa cecidogen indicated that trehalose was the main sugar component; nevertheless, its concentration was very low (data not shown). Therefore, considering the 14-fold increase in trehalose stem content after oviposition, its accumulation is probably induced by effectors which trigger gall initiation. Such increase in trehalose content during gall initiation might act as an initial signal for cell differentiation and carbon re-allocation. Trehalose is among the crucial virulence factors for Pseudomonas aeruginosa in plants (Djonović et al., 2013), while it is also involved in early regulation of metabolic processes in beneficial plant-microbe interactions (reviewed by Figueroa & Lunn, 2016). Additionally, trehalose was crucial for initial rice infection by Magnaporthe grisea, while its subsequent degradation allowed pathogen development (Foster et al., 2003). In this study, trehalose content decreased in developing gall after seven days (Fig. 1). On the other hand, trehalose may induce plant defense responses (Morkunas and Ratajczak, 2014), which could explain the transient increase in the initiating gall, prior to expected transcriptional re-programming by yet unknown effectors from R. pilosa. It has been reported that pathogenic bacteria Xanthomonas citri which induces citrus canker, synthesize trehalose to modify host plant metabolism to their advantage, while in the same time plants perceive trehalose as a sign of pathogen attack (Piazza et al., 2015). Besides trehalose and T6P, accumulation of hexoses (glucose, xylose, rhamnose, and raffinose and turanose) was observed upon gall initiation in L. vulgaris stem (G1S), without significant changes in the starch content and with the slight accumulation of sucrose (Figs. 1 and 3). Trehalose, T6P and sucrose are essential signaling metabolites which regulate specific responses to environmental signals such as diurnal changes and biotic and abiotic stress (Rolland et al., 2006; Griffiths et al., 2016). Sucrose is the primary photosynthetic product and transport sugar in plants. Majority of sucrose produced in Fig. 4. Kinetic representation of the OJIP transients in A) control L. vulgaris leaves and leaves on plants with developing gall, and B) control stems and stem galls seven days after oviposition. Values present average ± SE; n = 12. Table 2 JIP-test parameters of leaves and stems of control Linaria vulgaris plants and leaves and galls seven days after oviposition (G2), numbers represent average ± SE; n = 12; *p < 005, **p < 001,***p < 0001. φPo ψET2o ψRE1o δRE1o φET2o φRE1o Control leaves Gall leaves Control stem Gall stem 0.74 0.57 0.19 0.34 0.42 0.14 0.73 0.56 0.18 0.32 0.41 0.13 0.77 0.60 0.17 0.28 0.47 0.13 0.76 0.58 0.14 0.24 0.44 0.10 ± ± ± ± ± ± 0.02 0.03 0.03 0.05 0.03 0.02 ± ± ± ± ± ± 0.02 0.04 0.02 0.04 0.04 0.01 ± ± ± ± ± ± 0.01 0.01 0.01 0.02 0.01 0.01 ± ± ± ± ± ± 0.04 0.01∗ 0.01∗∗∗ 0.02∗∗∗ 0.03∗ 0.01∗∗∗ Calculations of these parameters are given in Table 1, according to Stirbet and Govindjee (2011). control plants (Fig. 4A). In contrast, in the seven days old galls, increased Chl fluorescence at F300μs, Fj and Fi, indicated changes in primary photochemical reactions and electron transport (Fig. 4B). Although φPo was not significantly affected, a decrease in quantum yield of electron transport flux from QA to QB was reduced, as well as further electron transport flux to PSI acceptors indicative by decreased φET20 and φR10 (Table 2). 3.5. The content of photosynthetic pigments The contents of Chl a and Chl b significantly decreased in developing gall, GS2 (2.7 and 2-fold, respectively) compared to control stems, and a lower Chl a/b ratio was observed (Table 3). Similarly, the 229 Plant Physiology and Biochemistry 135 (2019) 224–232 A.S. Zorić et al. On the other hand, the main systemic response to herbivory was the accumulation of T6P in both HS and HL (Figs. 1 and 2), accompanied by the starch breakdown and maltose accumulation in the stems. The ratio of starch to TIC decreased upon insect feeding (Fig. 3) indicating the differential distribution of carbohydrates. Aphids feeding elicited changes in gene expression associated with carbohydrate metabolism and chloroplasts as well as cell wall modifications in Arabidopsis (Appel et al., 2014). Moreover, the expression of four trehalose phosphate synthase genes was up-regulated by Brevicoryne brassicae feeding. Our findings indicate T6P signaling function induced by R.pilosa herbivory, which supports the previous reports on the role of trehalose phosphate synthase11 in promoting Arabidopsis defense against the phloemfeeding insects (Singh et al., 2011). Interestingly, increase in galactose was also observed in L. vulgaris stems upon herbivory. Galactose has been indicated in herbivore resistance in Pseudotsuga menziesii (Zou and Cates, 1994) and negatively affected the growth rate of Dineura pullior (Riipi et al., 2005) and Epirrita autumnata (Henriksson et al., 2003). The content of both galactose and its reduction product, galactitiol, significantly decreased in the developing galls. mesophyll cells is released to the phloem and distributed to the sink tissues where it is cleaved by different invertases to yield monomer hexoses, glucose and fructose (Figueroa and Lunn, 2016; Granot et al., 2013a, 2013b). Trehalose 6-phosphate (T6P) levels are closely related to sucrose concentration in plants (Lunn et al., 2006). T6P has been recently designated as the central signaling molecule that modulates metabolic and developmental processes, and stress responses in plants including source-sink carbon partitioning (Figueroa and Lunn, 2016; Lunn et al., 2014). T6P plays a central role in regulating carbohydrate metabolism, and the role of T6P in orchestrating cell cycle activity and cell wall biosynthesis with cellular metabolism during embryo development has also been implicated (Ponnu et al., 2011). It was shown that T6P inhibits the sucrose non-fermenting 1-related protein kinase1 (SnRK1) complex in Arabidopsis (Zhang et al., 2009). SnRK1 acts as a metabolite sensor to continually adapt the metabolism to the supply and demand of energy (Wurzinger et al., 2018). Moreover, the changes in hexose levels or sucrose: hexose ratio may trigger hexose signaling pathways mediated by hexokinases, target of rapamycin (TOR) and other transcription factors (Xiong et al., 2013; Figueroa and Lunn, 2016; Yadav et al., 2014). Concerning these hypotheses, in our model system, the ratio of sucrose: hexoses decreased from 0.21 in control stems to 0.13 and 0.04 in the initiating (GS1) and developing gall (GS2), respectively. Glucose accumulation in GS1 and GS2 could have been associated with early organ growth and increased mitotic activity as observed in developing cotyledons (Rolland et al., 2006; Horacio and MartinezNoel, 2013). The interplay between glucose and phytohormones (auxins, cytokinins, abscisic acid) during plant development has also been shown (Sairanen et al., 2012; Kushwah and Laxmi, 2017). Additionally, turanose accumulation was correlated with gall development (Fig. 1). Stimulating effect of turanose on auxin accumulation has been observed (Gonzali et al., 2005), which may be of importance considering the involvement of auxin and cytokinins in cell growth and division (Oliveira et al., 2016). The decrease of trehalose and sucrose contents was correlated with the increase of sucrose metabolites, glucose and fructose in developing gall (GS2). The activity of invertase, which catalyses sucrose breakdown to glucose and fructose, was detected in the nutritive tissue of young and mature galls (Rehill and Schultz, 2003; Oliveira et al., 2010). Increased accumulation of soluble carbohydrates in leaf galls compared to uninfected leaves has been reported in several plants (Castro et al., 2012; Hartley, 1998; Huang et al., 2015). Additionally, starch as carbon reserve accumulated in root galls (Griesser et al., 2015), and leaf galls (Oliveira et al., 2010), which was not observed in the stem gall in this study. Similarly as in our study, no increase in the starch content was also observed in leaf galls of Copaifera langsdorffii and Litsea acuminata (Castro et al., 2012; Huang et al., 2015). Interestingly, in plants with initiating galls, mannitol, rhamnose, sorbitol and xylose contents have similar trends in both leaves and stems (GL2 and GS2), which may be a systemic response to oviposition. Here the antioxidative function of mannitol should be emphasized (Keunen et al., 2013) considering the increased ROS production in the first stages of enhanced tissue proliferation and growth (Mittler, 2017). Furthermore, T6P, fructose and glucose have opposite (decreasing) response to gall initiation in the leaves (GL1) compared to stems (GS1), which may further support re-allocation of hexoses towards the gall tissue. Galls provide not only nutrition to the developing insect but also a shelter against enemies, e.g. through fortification of the cell wall (Oliveira and Isaias, 2010; Carneiro et al., 2015). Previously, lignin accumulation has been observed in a mature L. vulgaris gall (Barnewall, 2011) which is supported by the increased content of structural sugars, TIC (Fig. 3), and xylose in the developing gall detected in this study. In type I cell wall (Dicot species) the main carbohydrates are cellulose and xyloglucans (Carpita, 2011), while lignin, cross-linked with polysaccharides, confers mechanical strength to the cell wall. 4.2. Effects of gall development on OJIP chlorophyll fluorescence kinetic transients Analysis of chlorophyll fluorescence kinetic transients showed significant changes in photosystem II (PSII) photochemistry and electron transport (ET) efficiency in developing galls compared to control ones (Fig. 4, Table 2). This was correlated with a decrease in the amount of Chl a and Chl b as well as carotenoids in the developing galls (Table 3). Similarly, Aldea et al. (2006) showed decreased photosystem II (PSII) efficiency in the leaf galls of several tree species compared to uninfected leaves. Besides, lower maximum quantum efficiencies for PSII and lower content of pigment-protein complexes were observed in gall leaf portions than ungalled ones (Yang et al., 2007; Huang et al., 2011). Decreased contents of chlorophylls and carotenoids have been observed in galls on Psidium myrtoides (Carneiro et al., 2014), leaf galls of Machilus thunbergii (Huang et al., 2011) and on Copaifera langsdorffii (Castro et al., 2012), while net photosynthetic rate decreased in oak galls (Jiang et al., 2018). On the other hand, no effect on photosynthesis rate was observed in Aspidosperma australe following gall formation (Oliveira et al., 2011), while even increased photosynthesis rates were observed in phyllodes with clustered wasp-induced galls than in control ones in Acacia pycnantha (Dorchin et al., 2006). Inhibition of photosynthetic performance in L. vulgaris galls may be regulated by accumulated hexoses in the same tissue. It is proposed that hexokinases and fructokinases can sense the accumulated sugars, and repress the expression of photosynthetic genes such as Rubisco and Chl a/b binding proteins (Granot et al., 2013a, 2013b), which may be related to observed changes in the galls in this study. Our results on the decreased quantum yield of electron transport flux from QA to QB and further until PSI acceptors after seven days compared to control L. vulgaris stems (Table 3) indicate limited ability for photosynthesis to support intensive tissue expansion in the gall. On the other hand, in the leaves of galled plants, there was no change in photosynthetic performance compared to control ones, which was correlated with unchanged levels of starch and total insoluble carbohydrates. Considering that plant growth is retarded in relation to dramatic increase of gall biomass during development (about 3-fold compared to control stems), we suggest that the allocation of photoassimilates to the gall as a sink tissue is the main supply of energy for enhanced metabolic processes (Castro et al., 2012; Huang et al., 2015). The low photosynthesis rate in the galls should provide oxygen and consume CO2 to stabilize the tissue and protect R. pilosa from hypoxia and hypercarbia, as suggested by Oliveira et al. (2016) and Carneiro et al. (2017). The results obtained in this study show that R. pilosa can manipulate 230 Plant Physiology and Biochemistry 135 (2019) 224–232 A.S. Zorić et al. plant resource allocation pattern from the moment of oviposition to gall development by mechanisms which involve T6P and trehalose accumulation. 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