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Article

Photosystem II Responses at the Whole-Potato-Leaf Level After Colorado Potato Beetle Feeding

by
Ilektra Sperdouli
1,
Stefanos S. Andreadis
1,
Julietta Moustaka
2,
Eleni I. Koutsogeorgiou
1,
Emmanuel Panteris
3 and
Michael Moustakas
3,*
1
Institute of Plant Breeding and Genetic Resources, Hellenic Agricultural Organization-Dimitra, 57001 Thessaloniki, Greece
2
Department of Food Science, Aarhus University, 8000 Aarhus, Denmark
3
Department of Botany, School of Biology, Aristotle University of Thessaloniki, 54124 Thessaloniki, Greece
*
Author to whom correspondence should be addressed.
Plants 2026, 15(8), 1159; https://doi.org/10.3390/plants15081159
Submission received: 13 March 2026 / Revised: 31 March 2026 / Accepted: 6 April 2026 / Published: 9 April 2026
(This article belongs to the Section Crop Physiology and Crop Production)
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Abstract

The damage caused by herbivores is generally measured as the amount of leaf tissue consumed, without accounting for the fate of the leftover tissue. As a result, the plant defense mechanisms that promote resistance to herbivore feeding by photosynthetically acclimating the rest of the plant to the feeding spot leaf area have not been well exploited. Plant-insect interactions are now becoming better defined with the development of visualization methods that permit spatial whole-leaf assessment of photosynthetic efficiency after herbivore attack. The purpose of our study was to evaluate the spatial heterogeneity of photosystem II (PSII) function at the whole-leaf level before and after herbivory by the Colorado potato beetles. Twenty minutes after Colorado potato beetle (Leptinotarsa decemlineata) feeding, the maximum efficiency of PSII photochemistry (Fv/Fm) decreased significantly, suggesting photoinhibition due to reduced efficiency of the oxygen-evolving complex (OEC). The decreased quantum yield of PSII photochemistry (ΦPSII) after feeding, at the neighboring area of the feeding spot and at the rest of the leaf area, was attributed to the reduced efficiency of the open PSII reaction centers (Fv′/Fm′), since there was no change in the fraction of open PSII reaction centers (qp). Nevertheless, plant defense elicitation was activated by the photoprotective mechanism of non-photochemical quenching (NPQ) that reduced the singlet oxygen (1O2) formation in potato plants in the neighboring area of the feeding spot and at the rest of the leaf area. In addition, the increased production of hydrogen peroxide (H2O2) triggered by this increase suggests that it acted as a signaling molecule in the biotic stress defense response.

1. Introduction

Herbivorous insects pose a major agricultural challenge, reducing annual crop yields and resulting in considerable losses, currently estimated at approximately 38% [1]. Traditional approaches to estimating productivity losses due to herbivory in agricultural systems often overlook the effects on photosynthesis in the remaining undamaged leaf tissue [2,3,4]. High-resolution analysis of photosynthetic function is therefore essential for elucidating how intact areas are physiologically influenced [3]. Chlorophyll fluorescence analysis has emerged as a basic methodological tool for probing the functional integrity of the photosynthetic apparatus and for quantifying plant tolerance to diverse biotic and abiotic stressors [4,5,6,7,8,9]. Since stress conditions perturb the utilization efficiency of absorbed light energy, chlorophyll fluorescence provides a sensitive, noninvasive, cost-effective, and highly accurate diagnostic tool for stress-induced impairments in photosynthetic efficiency [3,6,7,10,11,12,13,14]. However, the spatial heterogeneity of photosynthetic activity across the leaf lamina limits the usefulness of standard point-based chlorophyll fluorescence measurements [8,15,16,17,18,19]. The development of chlorophyll fluorescence imaging apparatus has substantially overcome this limitation by enabling spatially resolved quantification of photosynthetic heterogeneity at the whole-leaf scale [15,17,19,20].
Insect herbivory, like other biotic stresses, is known to alter photosynthetic activity, most commonly reducing it, although compensatory enhancements can also occur [3,7,12,13,21,22]. Because the light-dependent reactions generate the reducing power and chemical energy necessary for synthesizing a broad suite of defense-related metabolites, they constitute a critical element of the plant’s integrated response to herbivore attack [23]. Photosystem II (PSII), a multiunit thylakoid membrane complex responsible for photochemical charge separation, water oxidation, and the generation of atmospheric oxygen, is indispensable for sustaining aerobic life on Earth [5,24,25]. PSII is also widely recognized as one of the most stress-sensitive components of the photosynthetic apparatus, exhibiting pronounced susceptibility to both biotic and abiotic perturbations [15,26,27].
Insect herbivory triggers plant responses involving photosynthesis, reactive oxygen species (ROS), and hormonal signaling, which are interconnected through a complex crosstalk [28,29]. Primarily, herbivory affects photosynthesis and initiates defense mechanisms, such as ROS production and the release of defense hormones, including jasmonic acid (JA) and salicylic acid (SA) [3,28]. ROS, such as singlet oxygen (1O2), hydrogen peroxide (H2O2), and superoxide anion radical (O2•−), are continuously produced in the light reactions of photosynthesis at basal levels, and are kept under optimal growth conditions by the antioxidative enzymatic and non-enzymatic systems in homeostasis [11,29,30,31,32]. However, under environmental stress conditions, the equilibrium between ROS generation and removal is disrupted [32,33,34,35,36].
ROS, acting as key signaling molecules, combine hormonal and photosynthetic pathways to coordinate a defense response [28]. ROS generation in photosynthesis plays a fundamental role as retrograde signaling molecules, activating the plant’s defense responses to environmental stressors, contributing to the “oxidation-reduction” balance, regulating a variety of physiological functions, and activating a plethora of acclimation responses [29,30,37,38,39,40,41]. Plants not only respond to insect feeding damage but also respond to insect egg positioning at the earliest stage of insect attack, allowing the plant to prepare its defense even before the damaging feeding stages of the insect life cycle have begun [42,43].
Potato (Solanum tuberosum L.) is one of the most important world food crops, ranking third after rice and wheat [44,45], and is significant because of its starch-rich tubers, which are a principal source of the daily diet for several people [46]. Potatoes are vulnerable to many types of biotic stress, with defoliating insects posing a major threat that can diminish both yield and tuber quality [12]. Among these insect pests, the Colorado potato beetle (Leptinotarsa decemlineata Say, Coleoptera: Chrysomelidae) is particularly notable for the extensive damage it causes and its remarkable resistance to insecticides [45]. The Colorado potato beetle is currently considered to be the principal insect defoliator of potatoes. Both adults and larvae consume whole leaves without differentiating between leaf tissues. Climate change may significantly affect the behavior and distribution of Colorado potato beetles, allowing them to spread into regions where they were not previously found, thereby increasing their harmfulness [47,48]. Yet, due to global climate change, increased drought stress affects plant health, resulting in fewer resistant plants against pest attacks [4].
Here, we studied the effects of short-time-restricted feeding by the most damaging phytophagous pest of potato on light energy use efficiency and ROS generation. We aimed to assess whether the photosynthetic mechanism could respond to Colorado potato beetle feeding and prevent further damage to the remaining leaf area. We investigated whether the leaf parts not damaged by herbivory (the neighboring area to the feeding spot and the rest of the leaf area) were negatively affected after herbivore feeding, or whether the photoprotective mechanism triggered defense elicitation to prevent further damage.

2. Results

2.1. Spatial Heterogeneity of PSII Function Before and After Feeding

The potato leaflets show photosynthetic heterogeneity both before and after herbivore feeding. In the representative color-coded images of the chlorophyll fluorescence parameters, an obvious spatial heterogeneity between the left and right leaf sides was recognized, with higher values of the maximum efficiency of PSII photochemistry (Fv/Fm), and a higher amount of absorbed light energy directed to photochemistry (ΦPSII), occurring in the left leaf side (Figure 1). The higher values of the quantum yield of regulated non-photochemical energy loss in PSII (ΦNPQ) occurred at the right side, while the higher values of the quantum yield of non-regulated energy loss in PSII (ΦNO) occurred at the left leaf side (Figure 1). This spatial heterogeneity between the left and right leaf sides was also retained after herbivore feeding. Twenty min of herbivore feeding resulted in decreased whole leaf Fv/Fm and ΦPSII values and increased whole leaf ΦNPQ and ΦNO values (Figure 1). The white arrows in Figure 1 show the feeding spot area, wherein the two areas of interest (AOIs) and their associated values for the maximum efficiency of PSII photochemistry (Fv/Fm), the effective quantum yield of PSII photochemistry (ΦPSII), and the quantum yield of regulated non-photochemical energy loss in PSII (ΦNPQ) were all equal to 0.00. However, the associated values for the quantum yield of non-regulated energy loss in PSII (ΦNO) were 1.00 (Figure 1).
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Figure 1. Representative images of the maximum efficiency of PSII photochemistry (Fv/Fm), the effective quantum yield of PSII photochemistry (ΦPSII), the quantum yield of regulated non-photochemical energy loss in PSII (ΦNPQ), and the quantum yield of non-regulated energy loss in PSII (ΦNO) before and 20 min after Colorado potato beetle feeding. The white arrows show the feeding spot area in potato leaflets. The areas of interest (AOIs) measured at the leaf surface are shown as circles, while for each parameter, the overall mean value from n = 3–4 leaves (±SD) is given in white. White arrows point to feeding areas. Two areas of interest (AOIs), shown as circles with their associated measurements labeled in red, were added at the feeding spot. An asterisk denotes statistical significance difference between before and after feeding. The color code on the bottom of the images shows pixel values ranging from 0.0 to 1.0.
Figure 1. Representative images of the maximum efficiency of PSII photochemistry (Fv/Fm), the effective quantum yield of PSII photochemistry (ΦPSII), the quantum yield of regulated non-photochemical energy loss in PSII (ΦNPQ), and the quantum yield of non-regulated energy loss in PSII (ΦNO) before and 20 min after Colorado potato beetle feeding. The white arrows show the feeding spot area in potato leaflets. The areas of interest (AOIs) measured at the leaf surface are shown as circles, while for each parameter, the overall mean value from n = 3–4 leaves (±SD) is given in white. White arrows point to feeding areas. Two areas of interest (AOIs), shown as circles with their associated measurements labeled in red, were added at the feeding spot. An asterisk denotes statistical significance difference between before and after feeding. The color code on the bottom of the images shows pixel values ranging from 0.0 to 1.0.
Plants 15 01159 g001

2.2. Impact of Feeding on the Efficiency of the Oxygen-Evolving Complex and the Maximum Efficiency of Photosystem II Photochemistry

The efficiency of the oxygen-evolving complex (OEC) as evaluated in the two different zones (the neighboring area at the feeding spot and the rest of the leaf area) by the ratio Fv/Fo, was significantly lower 20 min after feeding compared to before (Figure 2a). No difference between the two zones, before or after feeding, was observed. Similar results were observed for the maximum efficiency of PSII photochemistry (Fv/Fm) in the two areas before and after feeding, with lower Fv/Fm values detected after feeding (Figure 2b).
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Figure 2. Changes in the efficiency of the oxygen-evolving complex (Fv/Fo) (a) and the maximum efficiency of PSII photochemistry (Fv/Fm) (b) in the two different zones (the neighboring area at the feeding spot and the rest of the leaf area) before and 20 min after Colorado potato beetle feeding. Bars display standard deviations (SD). Significant differences are shown by different lower-case letters (p < 0.05).
Figure 2. Changes in the efficiency of the oxygen-evolving complex (Fv/Fo) (a) and the maximum efficiency of PSII photochemistry (Fv/Fm) (b) in the two different zones (the neighboring area at the feeding spot and the rest of the leaf area) before and 20 min after Colorado potato beetle feeding. Bars display standard deviations (SD). Significant differences are shown by different lower-case letters (p < 0.05).
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2.3. Impact of Feeding on the Light Energy Use Efficiency

The absorbed light energy is allocated either to photochemistry (ΦPSII), dissipated as heat (ΦNPQ), or lost in a nonregulatory way (ΦNO), with all of them being equal to 1 [49].
The quantum yield for photochemistry (ΦPSII) for the neighboring area at the feeding spot and for the rest of the leaf was significantly lower 20 min after feeding compared to before feeding (Figure 3a). In contrast to ΦPSII, the regulated non-photochemical energy loss in PSII (ΦNPQ) was significantly higher 20 min after feeding than before, in both areas (Figure 3b). Also, no difference was observed between the neighboring area at the feeding spot and the rest of the leaf in ΦNPQ values, both before and after feeding (Figure 3a).
Plants 15 01159 g003
Figure 3. Changes in ΦPSII (a), ΦNPQ (b), and ΦNO (c) in the two different zones (the neighboring area at the feeding spot and the rest of the leaf area) before and 20 min after Colorado potato beetle feeding. Bars display standard deviations (SD). Significant differences are shown by different lower-case letters (p < 0.05).
Figure 3. Changes in ΦPSII (a), ΦNPQ (b), and ΦNO (c) in the two different zones (the neighboring area at the feeding spot and the rest of the leaf area) before and 20 min after Colorado potato beetle feeding. Bars display standard deviations (SD). Significant differences are shown by different lower-case letters (p < 0.05).
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The yield of non-regulated energy loss in PSII (ΦNO) was significantly lower 20 min after feeding compared to before feeding, at the neighboring area to the feeding spot and at the rest of the leaf (Figure 3c). This decrease contrasted with the observed increase in ΦNO values after feeding compared to before, evaluated across the whole leaf area (Figure 1). This was due to the high ΦNO values (1.00) detected at the feeding spot, where the two areas of interest (AOIs) shown as circles in Figure 1 were added at the feeding spot.

2.4. Impact of Feeding on the Fraction of Open PSII Reaction Centers, Their Efficiency, the Electron Transport Rate, and the Photoprotective Mechanism

The fraction of open PSII reaction centers (RCs) (qp), representing the redox state of quinone A (QA), after 20 min of feeding, remained the same as before feeding (Figure 4a). No differences were observed between the neighboring area at the feeding spot and the rest of the leaf in qp values, both before and after feeding (Figure 4a). A reduced efficiency of the open PSII reaction centers (RCs) (Fv′/Fm′) at the neighboring area of feeding and at the rest of the leaf was observed 20 min after feeding, compared with before feeding (Figure 4b). No differences were observed between the neighboring area at the feeding spot and the rest of the leaf in the efficiency of the open PSII RCs (Fv′/Fm′), both before and after feeding (Figure 4b).
Plants 15 01159 g004
Figure 4. Changes in the portion of open PSII reaction centers (RCs) (qp) that reveal the redox state of quinone A (QA) (a), the efficiency of the open PSII RCs (Fv′/Fm′) (b), the photoprotective mechanism of non-photochemical quenching (NPQ) (c), and the electron transport rate (ETR) (d) in the two different zones (the neighboring area at the feeding spot and the rest of the leaf area) before and 20 min after Colorado potato beetle feeding on potato leaves. The bars display standard deviations (SD). Significant differences are shown by different lower-case letters (p < 0.05).
Figure 4. Changes in the portion of open PSII reaction centers (RCs) (qp) that reveal the redox state of quinone A (QA) (a), the efficiency of the open PSII RCs (Fv′/Fm′) (b), the photoprotective mechanism of non-photochemical quenching (NPQ) (c), and the electron transport rate (ETR) (d) in the two different zones (the neighboring area at the feeding spot and the rest of the leaf area) before and 20 min after Colorado potato beetle feeding on potato leaves. The bars display standard deviations (SD). Significant differences are shown by different lower-case letters (p < 0.05).
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The photoprotective mechanism of non-photochemical quenching (NPQ) increased significantly 20 min after feeding, compared with before, in both areas (Figure 4c). However, no difference was observed between the neighboring area at the feeding spot and the rest of the leaf in NPQ values, both before and after feeding (Figure 4c). The electron transport rate (ETR) reduced significantly at the neighboring area of the feeding spot and at the rest of the leaf 20 min after the feeding compared to before feeding (Figure 4d).

2.5. Correlation of the Maximum Efficiency of PSII with the Efficiency of the Oxygen-Evolving Complex

The maximum efficiency of photosystem II photochemistry (Fv/Fm) and the efficiency of the oxygen-evolving complex (Fv/Fo) were positive and significantly correlated as shown in a regression analysis for both before feeding (R2 = 0.9966, p < 0.01) (Figure 5a), and after 20 min of Colorado potato beetle feeding on potato leaves (R2 = 0.9946, p < 0.01) (Figure 5b).

2.6. Hydrogen Peroxide Imaging Before and After Herbivore Feeding

Hydrogen peroxide production in potato leaves was localized mainly in the leaf veins, as evidenced by green fluorescence (Figure 6). After 20 min of Colorado potato beetle feeding, H2O2 generation was localized at the feeding spot (Figure 6b,c) and at the leaf veins (Figure 6b), where it was slightly increased compared to control leaves (Figure 6a). H2O2 was not detected at the neighboring of the feeding spot or at the rest of the leaf area (Figure 6b,c).

3. Discussion

Plant-insect interactions are now becoming better defined with the development of visualization methods that permit spatial whole-leaf assessment of the distribution of absorbed light energy before and after herbivore attack [3,13,15]. Under many biotic or abiotic stress factors, the absorbed light energy in photosynthesis exceeds the capacity of de-excitation processes, leading to photoinhibition and primarily damaging PSII via the formation of ROS [29,30,33,34,35,36,50]. Twenty minutes after Colorado potato beetle feeding, the maximum efficiency of PSII photochemistry (Fv/Fm) decreased significantly, suggesting possible photoinhibition. Photoinhibition appears either at the acceptor side through the excited triplet state of chlorophyll (3Chl*), resulting in singlet oxygen (1O2) formation, or at the donor side through inactivation of the OEC [51,52,53,54,55,56,57,58]. The OEC catalyzes water oxidation, located on the electron donor side of PSII, and contains the so-called Mn4CaO5 cluster [59,60,61]. Consequently, photoinhibition can emerge via a dual mechanism, with PSII as the primary target [53,55,62]. The observed photoinhibition in potato leaves 20 min after Colorado potato beetle feeding was the result of an impairment of the OEC function, as estimated by the decreased Fv/Fo ratio [63,64,65,66,67,68,69]. It is well known that reduced OEC efficiency also implies a lower Fv/Fm ratio [70,71]. In agreement with this, a significant positive correlation between the maximum efficiency of PSII photochemistry (Fv/Fm) and the function of OEC was revealed by regression analysis, both before (Figure 5a) and after feeding by the Colorado potato beetle (Figure 5b). Photoinhibition, which is associated with reduced OEC efficiency [72,73,74,75], is often associated with crop production [76].
However, since advanced analysis beyond simple Fv/Fm is required for accurate physiological assessment [77], we examined the light-use efficiency of potato plants before and after 20 min of Colorado potato beetle feeding. The observed photoinhibition in potatoes after the feeding, as observed by the reductions in Fv/Fm, was coupled with decreases in the effective quantum yield of PSII photochemistry (ΦPSII) (Figure 1). The decreased photochemical efficiency (ΦPSII), at the neighboring area to the feeding spot and at the rest of the leaf area after Colorado potato beetle feeding (Figure 3a), was overcompensated by the amplified regulated non-photochemical energy loss in PSII (ΦNPQ) (Figure 3b), which developed a drop in non-regulated loss (ΦNO) (Figure 3c). This decreased ΦNO is correlated with reduced quantity of singlet oxygen (1O2) generation [11,78,79,80]. 1O2 is created through the interaction of molecular O2 with the excited 3Chl* and is considered highly reactive and damaging [30,81,82,83,84]. Thus, the reduced 1O2 generation at the neighboring area of the feeding spot and at the rest of the leaf area, after 20 min of Colorado potato beetle feeding, confirms that the observed photoinhibition was associated with a “donor-side photoinhibition” through a reduced OEC efficiency. Otherwise, an increased 1O2 production illuminates the so-called ‘‘acceptor-side photoinhibition’’ of PSII [85,86,87,88,89]. However, a 20 min feeding by the pinworm Tuta absoluta on potato leaves did not have any effect on the OEC or Fv/Fm at the whole-leaf level [12]. Inhibition of photosynthesis due to insect herbivory was frequently reported [12,21,90,91,92], and in this context, ROS play a crucial role [12,28].
Plants can respond to disturbances in their homeostasis, triggered by biotic stress factors, by displaying adaptive responses that often result in compensatory responses [4,25,32]. Understanding these molecular mechanisms that initiate adaptive responses in plants is significant for enhancing crop productivity [4]. Thus, plants have developed several photoprotective mechanisms to counteract ROS production and hence escape PSII damage [29,32,93,94]. The dissipation of excess absorbed light energy as heat by the mechanism of non-photochemical quenching (NPQ) [31,81,95,96] protects PSII from the formation of ROS, which are considered detrimental to plant cells [31,81,97,98,99]. Twenty min after the feeding, the increased NPQ mechanism at the neighboring area to the feeding spot and at the rest of the leaf area (Figure 4c) not only resulted in decreasing the ETR (Figure 4d), but also reduced the 1O2 production (Figure 3c).
The decreased ΦPSII and ETR, at the neighboring area of the feeding spot and at the rest of the leaf area, are attributed to the decreased efficiency of the open PSII reaction centers (Fv′/Fm′) (Figure 4b), since there was no change in the number of open PSII reaction centers (qp) (Figure 4a). In contrast to our results, 20 min feeding by the leafminer Tuta absoluta on potato leaves did not have any effect on Fv′/Fm′ but decreased the number of open PSII RCs [12]. However, this difference may result from their different feeding styles: T. absoluta is a miner, causing internal damage to the leaf, while the Colorado beetle is a chewing herbivore. This is a paradigm that different feeding styles produce differential plant responses.
NPQ can be considered sufficient if the fraction of open PSII RCs can be maintained at a level similar to that in control conditions under any disturbance of plant homeostasis [100,101]. In our study, in agreement with this, the increased NPQ in the neighboring area to the feeding spot and in the rest of the leaf area (Figure 4c) appeared to be sufficient to keep the same portion of PSII RCs open as in controls (Figure 4a). In contrast, after 20 min of feeding by the leafmine Tuta absoluta on potato, the increased NPQ could not keep open the same number of PSII RCs as before feeding, suggesting that the NPQ mechanism was not efficient enough [12]. Thus, it can be concluded that potato PSII response mechanism to insect herbivores depends on the insect species.
The enhancement of NPQ after Colorado potato beetle feeding also resulted in decreased 1O2 generation (Figure 3c). It is now well documented that any disturbance of ROS production at the light reactions of photosynthesis triggers the plant’s protective defense response to environmental perturbations, and it contributes to restoring the “oxidation-reduction” balance [29,37,102,103,104,105,106,107]. The most stable ROS is hydrogen peroxide (H2O2), which can act as a long-distance signaling molecule mediating plant responses to alterations in homeostasis [108,109,110]. H2O2 is produced during photosynthesis in the electron transport chain by the electron leakage in PSI that reduces oxygen (O2) to superoxide anion (O2•−), which is rapidly converted to hydrogen peroxide (H2O2) by the superoxide dismutase (SOD) [29,30,111,112,113]. H2O2 can travel through leaf veins faster than from cell to cell, and thus it spreads through leaf veins to act as a long-distance molecule triggering the plant defense response during biotic or abiotic stress in plants [12,29,103,114,115].
1O2 and H2O2 are the main ROS that initiate various signaling networks when the light reactions of photosynthesis are dysfunctional [116]. 1O2 is very reactive and can induce but not transduce signaling [117,118]. On the contrary, foliar-produced H2O2 is less reactive but is a mobile molecule that can diffuse throughout the leaf veins to act as a long-distance messenger [114,116,119]. It seems that the H2O2 produced at the feeding zone located in the vicinity of the leaf’s midrib (Figure 6b) elicits defense responses to herbivore feeding, by its diffusion through the leaf veins [13,103,120,121]. H2O2 is the most stable ROS that can mediate plant responses to stress [84]. Since 1O2 is produced by energy transfer and H2O2 by electron transport, yet they are produced simultaneously, it appears likely that their signaling pathways can occasionally interfere or antagonize each other [13,29,37,115]. A coordinated ROS generation has been considered the major plant defense response mechanism to herbivores [122].
Plants continuously experience insect herbivory, and insect feeding causes injury and impairs crop production [123,124]. Insect herbivory induces various signals from injured tissues that are perceived in untouched tissues, which subsequently boost their defense [124,125]. Such signals include the phytohormone JA, which is synthesized via the oxygen lipid pathway [126] and regulates plant defense against diverse insect herbivores [123,125,127]. JA biosynthesis proceeds in the vascular bundles [125,128], possibly via hydrogen peroxide signaling. The JA is considered to function as a long-distance signaling molecule traveling through the phloem [125] similar to hydrogen peroxide signaling [13,103,120,121]. The need of plants to protect themselves against herbivores has generated an array of photoprotective mechanisms that permit them to compensate for herbivory [3,129]. The level of plant photoprotective mechanisms also depends on the insect species under examination. Cues derived from different insects differentially influence flavonoid production [130], underlining the importance of the insect species involved in affecting plant-insect interactions.

4. Materials and Methods

4.1. Plant Material and Growth Conditions

Potato plants (Solanum tuberosum L. cv Spunta) were grown in 5 L plastic pots as described before [4]. The growth conditions were 23 ± 2°/19 ± 2 °C (day/night) temperature, 70 ± 5/80 ± 5% day/night relative humidity, and a 14-h photoperiod with photosynthetic photon flux density (PPFD) of 630 ± 10 μmol quanta m−2 s−1.

4.2. Leptinotarsa Decemlineata

Adults of Colorado potato beetle (L. decemlineata) (approximately 200 at a 1:1 sex ratio) were collected from a potato field cultivation established at the premises of the Institute of Plant Breeding and Genetic Resources, ELGO-Dimitra (Thermi, Greece). The collected Colorado potato beetle population was maintained at 26 ± 1 °C under a light–dark photoperiod of 16:8 h and 60–65% relative humidity on non-transgenic potato seedlings (S. tuberosum L. cv Spunta) in controlled growth chambers. The adults used in the bioassays were starved for 24 h prior to each experiment.

4.3. Experimental Design

The 4th terminal leaf of each potato plant was enclosed in the measurement chamber of a fluorometer, and photosynthetic efficiency was measured. Following the first measurement, one randomly selected Colorado potato beetle adult was added and allowed to feed for 20 min without removing the leaflet from the fluorometer’s measurement chamber. After the measurement, the insect was removed, and a new measurement was conducted on the same leaflet immediately after the feeding. Three to four leaves from different potato plants were measured before and after 20 min of Colorado potato beetle feeding.

4.4. Chlorophyll Fluorescence Analysis

Chlorophyll fluorescence measurements were performed as described in detail previously [131], using an Imaging-PAM Fluorometer M-Series MINI-Version (Heinz Walz GmbH, Effeltrich, Germany). The potato leaflets were dark-adapted for 20 min before each measurement. The minimum (Fo) and maximum (Fm) chlorophyll a fluorescence ιin the dark was recorded, and the variable (Fv) chlorophyll a fluorescence was calculated from Fv = Fm − Fo. The maximum chlorophyll a fluorescence in the light (Fm’) was acquired with saturating pulses (SPs) every 20 s for 5 min after application of the actinic light (AL) of 636 μmol photons m−2 s−1, which was selected to match the growing light intensity. The minimum chlorophyll a fluorescence in the light (Fo’) was computed as Fo’ = Fo/(Fv/Fm + Fo/Fm’) [132]. Steady-state photosynthesis (Fs) was measured after 5 min of illumination time with the AL of 636 μmol photons m−2 s−1. The remaining chlorophyll fluorescence parameters were estimated using Win V2.41a (Heinz Walz GmbH, Effeltrich, Germany) and are described in detail in Table S1.

4.5. Imaging of Hydrogen Peroxide

Hydrogen peroxide (H2O2) was detected before and after 20 min of Colorado potato beetle feeding on potato leaves. Potato leaflets were incubated in the dark for 30 min with 25 μM 2′,7′-dichlorofluorescein diacetate (DCF-DA, Sigma Aldrich, Chemie GmbH, Schnelldorf, Germany) as described earlier [115,133]. H2O2-specific fluorescence was observed afterwards with a Zeiss AxioImager Z2 epi-fluorescence microscope equipped with an AxioCam MRc5 digital camera (Carl Zeiss MicroImaging GmbH, Göttingen, Germany) [133].

4.6. Statistics

Statistical analysis was performed with R software (version 4.3.1, R Core Team, 2023). A two-way repeated measures ANOVA was conducted for each parameter, followed by post hoc pairwise comparisons using Fisher’s Protected Least Significant Difference (LSD) test. Values were considered significantly different at p < 0.05. A linear regression analysis was also performed.

5. Conclusions

Colorado potato beetle feeding decreased the maximum efficiency of PSII photochemistry (Fv/Fm), suggesting photoinhibition due to reduced efficiency of the oxygen-evolving complex (OEC). The observed reduced amount of absorbed light energy that was directed after feeding to photochemistry (ΦPSII) was attributed to the reduced efficiency of the open PSII reaction centers (Fv′/Fm′), since there was no alteration in the fraction of open PSII reaction centers (qp). The ability of potato plants to keep the same fraction of open PSII reaction centers as before feeding suggests an efficient photoprotective mechanism of non-photochemical quenching (NPQ), which was activated to reduce the singlet oxygen (1O2) formation in potato plants in the neighboring area to the feeding spot and at the rest of the leaf area. Concomitantly, NPQ triggered a slight increase in hydrogen peroxide (H2O2) production, which can act as a signaling molecule by diffusing through leaf veins to elicit defense responses. A coordinated signaling pathway involving 1O2 and H2O2 appears to operate as a biotic stress defense response mechanism in potatoes.
Understanding the molecular mechanisms that initiate adaptive responses in potatoes following Colorado potato beetle feeding is essential for developing new breeding approaches to enhance biotic stress resilience and crop productivity. Our findings highlight the importance of considering both the whole-leaf spatial direct effects of herbivory and the indirect effects on the neighboring area around the feeding spot and the rest of the leaf area.

Supplementary Materials

The following supporting information can be downloaded at https://www.mdpi.com/article/10.3390/plants15081159/s1. Table S1: Definitions of the chlorophyll fluorescence parameters used in the experiments.

Author Contributions

Conceptualization, M.M.; methodology, I.S., S.S.A., J.M., E.I.K., E.P., and M.M.; validation, J.M. and M.M.; formal analysis, I.S., S.S.A., J.M., E.I.K., E.P., and M.M.; investigation, I.S., S.S.A., J.M., E.I.K., E.P., and M.M.; resources, I.S., S.S.A., E.P., and M.M.; data curation, I.S., S.S.A., J.M., E.I.K., E.P., and M.M.; writing—original draft preparation, M.M.; writing—review and editing, I.S., S.S.A., J.M., E.I.K., E.P., and M.M.; visualization, I.S., J.M., E.P., and M.M.; supervision, M.M.; project administration, I.S., S.S.A., E.P., and M.M. All authors have read and agreed to the published version of the manuscript.

Funding

This research received no external funding.

Data Availability Statement

All data supporting this study are available within the paper and within its Supplementary Data published online.

Conflicts of Interest

The authors declare no conflicts of interest.

Abbreviations

The following abbreviations are used in this manuscript:
1O2Singlet oxygen
3Chl*Excited triplet state of chlorophyll
ALActinic light
AOIsAreas of interest
DCF-DA2′,7′-dichlorofluorescein diacetate
ETRElectron transport rate
FmMaximum chlorophyll a fluorescence in the dark-adapted leaf
Fm’Maximum chlorophyll a fluorescence in the light-adapted leaf
FoMinimum chlorophyll a fluorescence in the dark-adapted leaf
Fo’Minimum chlorophyll a fluorescence in the light-adapted leaf
FsSteady-state photosynthesis
FvVariable chlorophyll a fluorescence in the dark-adapted leaf (Fv = Fm − Fo)
Fv’/Fm’Efficiency of the open PSII reaction centers
Fv/FmMaximum efficiency of PSII photochemistry
Fv/FoEfficiency of the oxygen-evolving complex on the donor side of PSII
H2O2Hydrogen peroxide
JAJasmonic acid
NPQNon-photochemical quenching (dissipation of excitation energy as heat)
O2•−Superoxide anion radical
OECOxygen-evolving complex
PPFDPhotosynthetic photon flux density
PSIPhotosystem I
PSIIPhotosystem II
QAQuinone A 
qpPhotochemical quenching (fraction of open PSII reaction centers, representing also the redox state of quinone A)
RCsReaction centers 
ROSReactive oxygen species
SASalicylic acid
SDStandard deviation
SODSuperoxide dismutase
SPsSaturating pulses
ΦNOQuantum yield of non-regulated energy loss in PSII
ΦNPQQuantum yield of regulated non-photochemical energy loss in PSII 
ΦPSIIEffective quantum yield of PSII photochemistry

References

  1. Junaid, M.D.; Gokce, A.F. Global agricultural losses and their causes. Bull. Biol. Allied Sci. Res. 2024, 9, 66. [Google Scholar] [CrossRef]
  2. Mauch-Mani, B.; Baccelli, I.; Luna, E.; Flors, V. Defense priming: An adaptive part of induced resistance. Annu. Rev. Plant Biol. 2017, 68, 485–512. [Google Scholar] [CrossRef] [PubMed]
  3. Moustaka, J.; Meyling, N.V.; Hauser, T.P. Induction of a compensatory photosynthetic response mechanism in tomato leaves upon short time feeding by the chewing insect Spodoptera exigua. Insects 2021, 12, 562. [Google Scholar] [CrossRef] [PubMed]
  4. Moustaka, J.; Sperdouli, I.; Andreadis, S.S.; Stoikou, N.; Giannousi, K.; Dendrinou-Samara, C.; Moustakas, M. The compensatory response of photosystem II photochemistry to short-term insect herbivory is suppressed under water deficit. Insects 2025, 16, 984. [Google Scholar] [CrossRef]
  5. Moustakas, M.; Sperdouli, I.; Moustaka, J. Early drought stress warning in plants: Color pictures of photosystem II photochemistry. Climate 2022, 10, 179. [Google Scholar] [CrossRef]
  6. Thomson, V.P.; Cunningham, S.A.; Ball, M.C.; Nicotra, A.B. Compensation for herbivory by Cucumis sativus through increased photosynthetic capacity and efficiency. Oecologia 2003, 134, 167–175. [Google Scholar] [CrossRef]
  7. Retuerto, R.; Fernández-Lema, B.; Obeso, J.R. Changes in photochemical efficiency in response to herbivory and experimental defoliation in the dioecious tree Ilex aquifolium. Int. J. Plant Sci. 2006, 167, 279–289. [Google Scholar] [CrossRef]
  8. Lenk, S.; Chaerle, L.; Pfündel, E.E.; Langsdorf, G.; Hagenbeek, D.; Lichtenthaler, H.K.; Van Der Straeten, D.; Buschmann, C. Multispectral fluorescence and reflectance imaging at the leaf level and its possible applications. J. Exp. Bot. 2007, 58, 807–814. [Google Scholar] [CrossRef]
  9. Saglam, A.; Chaerle, L.; Van Der Straeten, D.; Valcke, R. Promising monitoring techniques for plant science: Thermal and chlorophyll fluorescence imaging. In Photosynthesis, Productivity, and Environmental Stress, 1st ed.; Ahmad, P., Ahanger, M.A., Alyemeni, M.N., Alam, P., Eds.; John Wiley & Sons Ltd.: Hoboken, NJ, USA, 2020; pp. 241–266. [Google Scholar]
  10. Moustaka, J.; Sperdouli, I.; Moustakas, M. Light energy use efficiency in photosystem ΙΙ of tomato is related to leaf age and light intensity. Crops 2024, 4, 623–635. [Google Scholar] [CrossRef]
  11. Moustaka, J.; Sperdouli, I.; Panteris, E.; Adamakis, I.D.S.; Moustakas, M. Aspirin foliar spray-induced changes in light energy use efficiency, chloroplast ultrastructure, and ROS generation in tomato. Int. J. Mol. Sci. 2025, 26, 1368. [Google Scholar] [CrossRef]
  12. Sperdouli, I.; Andreadis, S.; Moustaka, J.; Panteris, E.; Tsaballa, A.; Moustakas, M. Changes in light energy utilization in photosystem II and reactive oxygen species generation in potato leaves by the pinworm Tuta absoluta. Molecules 2021, 26, 2984. [Google Scholar] [CrossRef] [PubMed]
  13. Sperdouli, I.; Andreadis, S.S.; Adamakis, I.S.; Moustaka, J.; Koutsogeorgiou, E.I.; Moustakas, M. Reactive oxygen species initiate defence responses of potato photosystem II to sap-sucking insect feeding. Insects 2022, 13, 409. [Google Scholar] [CrossRef] [PubMed]
  14. Garab, G. Revisiting the QA model of chlorophyll-a fluorescence induction: New perspectives to monitor the photochemical activity and structural dynamics of photosystem II. Photosynth. Res. 2025, 163, 54. [Google Scholar] [CrossRef] [PubMed]
  15. Moustaka, J.; Moustakas, M. Early-stage detection of biotic and abiotic stress on plants by chlorophyll fluorescence imaging analysis. Biosensors 2023, 13, 796. [Google Scholar] [CrossRef]
  16. Rolfe, S.A.; Scholes, J.D. Chlorophyll fluorescence imaging of plant-pathogen interactions. Protoplasma 2010, 247, 163–175. [Google Scholar] [CrossRef]
  17. Gorbe, E.; Calatayud, A. Applications of chlorophyll fluorescence imaging technique in horticultural research: A review. Sci. Hortic. 2012, 138, 24–35. [Google Scholar] [CrossRef]
  18. Guidi, L.; Calatayud, A. Non-invasive tools to estimate stress-induced changes in photosynthetic performance in plants inhabiting Mediterranean areas. Environ. Exp. Bot. 2014, 103, 42–52. [Google Scholar] [CrossRef]
  19. Moustakas, M.; Calatayud, A.; Guidi, L. Chlorophyll fluorescence imaging analysis in biotic and abiotic stress. Front. Plant Sci. 2021, 12, 658500. [Google Scholar] [CrossRef]
  20. Pérez-Bueno, M.L.; Pineda, M.; Barón, M. Phenotyping plant responses to biotic stress by chlorophyll fluorescence imaging. Front. Plant Sci. 2019, 10, 1135. [Google Scholar] [CrossRef]
  21. Delaney, K.J.; Higley, L.G. An insect countermeasure impacts plant physiology: Midrib vein cutting, defoliation and leaf photosynthesis. Plant Cell Environ. 2006, 29, 1245–1258. [Google Scholar] [CrossRef]
  22. Delaney, K.J. Injured and uninjured leaf photosynthetic responses after mechanical injury on Nerium oleander leaves, and Danaus plexippus herbivory on Asclepias curassavica leaves. Plant Ecol. 2008, 199, 187–200. [Google Scholar] [CrossRef]
  23. Lu, Y.; Yao, J. Chloroplasts at the crossroad of photosynthesis, pathogen infection and plant defence. Int. J. Mol. Sci. 2018, 19, 3900. [Google Scholar] [CrossRef] [PubMed]
  24. Barber, J. Photosynthetic energy conversion: Natural and artificial. Chem. Soc. Rev. 2009, 38, 185–196. [Google Scholar] [CrossRef] [PubMed]
  25. Moustakas, M.; Moustaka, J.; Sperdouli, I. Hormesis in photosystem II: A mechanistic approach. Curr. Opin. Toxicol. 2022, 29, 57–64. [Google Scholar] [CrossRef]
  26. Moustaka, J.; Ouzounidou, G.; Sperdouli, I.; Moustakas, M. Photosystem II is more sensitive than photosystem I to Al3+ induced phytotoxicity. Materials 2018, 11, 1772. [Google Scholar] [CrossRef]
  27. Zlobin, I.E.; Ivanov, Y.V.; Kartashov, A.V.; Sarvin, B.A.; Stavrianidi, A.N.; Kreslavski, V.D.; Kuznetsov, V.V. Impact of weak water deficit on growth, photosynthetic primary processes and storage processes in pine and spruce seedlings. Photosynth. Res. 2019, 139, 307–323. [Google Scholar] [CrossRef]
  28. Kerchev, P.I.; Fenton, B.; Foyer, C.H.; Hancock, R.D. Plant responses to insect herbivory: Interactions between photosynthesis, reactive oxygen species and hormonal signalling pathways. Plant Cell Environ. 2012, 35, 441–453. [Google Scholar] [CrossRef]
  29. Moustaka, J.; Moustakas, M. ROS generation in the light reactions of photosynthesis triggers acclimation signaling to environmental stress. Photochem 2025, 5, 28. [Google Scholar] [CrossRef]
  30. Moustakas, M. Plant photochemistry, reactive oxygen species, and photoprotection. Photochem 2022, 2, 5–8. [Google Scholar] [CrossRef]
  31. Müller, P.; Li, X.P.; Niyogi, K.K. Non-photochemical quenching. A response to excess light energy. Plant Physiol. 2001, 125, 1558–1566. [Google Scholar] [CrossRef]
  32. Moustakas, M. Molecular mechanisms of plant abiotic stress tolerance. Int. J. Mol. Sci. 2025, 26, 2731. [Google Scholar] [CrossRef] [PubMed]
  33. Gill, S.S.; Tuteja, N. Reactive oxygen species and antioxidant machinery in abiotic stress tolerance in crop plants. Plant Physiol. Biochem. 2010, 48, 909–930. [Google Scholar] [CrossRef] [PubMed]
  34. Hasanuzzaman, M.; Bhuyan, M.H.M.B.; Parvin, K.; Bhuiyan, T.F.; Anee, T.I.; Nahar, K.; Hossen, M.S.; Zulfiqar, F.; Alam, M.M.; Fujita, M. Regulation of ROS metabolism in plants under environmental stress: A review of recent experimental evidence. Int. J. Mol. Sci. 2020, 21, 8695. [Google Scholar] [CrossRef] [PubMed]
  35. Mittler, R. Oxidative stress, antioxidants and stress tolerance. Trends Plant Sci. 2002, 7, 405–410. [Google Scholar] [CrossRef]
  36. Apel, K.; Hirt, H. Reactive oxygen species: Metabolism, oxidative stress, and signal transduction. Annu. Rev. Plant Biol. 2004, 55, 373–399. [Google Scholar] [CrossRef]
  37. Mittler, R. ROS are good. Trends Plant Sci. 2017, 22, 11–19. [Google Scholar] [CrossRef]
  38. Mittler, R.; Vanderauwera, S.; Gollery, M.; van Breusegem, F. Reactive oxygen gene network of plants. Trends Plant Sci. 2004, 9, 490–498. [Google Scholar] [CrossRef]
  39. Inupakutika, M.A.; Sengupta, S.; Devireddy, A.R.; Azad, R.K.; Mittler, R. The evolution of reactive oxygen species metabolism. J. Exp. Bot. 2016, 67, 5933–5943. [Google Scholar] [CrossRef]
  40. Lee, K.P.; Kim, C. Photosynthetic ROS and retrograde signaling pathways. New Phytol. 2024, 244, 1183–1198. [Google Scholar] [CrossRef]
  41. Foyer, C.H.; Noctor, G. Redox regulation in photosynthetic organisms: Signaling, acclimation, and practical implications. Antioxid. Redox Signal. 2009, 11, 861–905. [Google Scholar] [CrossRef]
  42. Hilker, M.; Meiners, T. Early herbivore alert: Insect eggs induce plant defense. J. Chem. Ecol. 2006, 32, 1379–1397. [Google Scholar] [CrossRef] [PubMed]
  43. Bruce, T.J. Interplay between insects and plants: Dynamic and complex interactions that have coevolved over millions of years but act in milliseconds. J. Exp. Bot. 2015, 66, 455–465. [Google Scholar] [CrossRef] [PubMed]
  44. Ye, M.; Peng, Z.; Tang, D.; Yang, Z.; Li, D.; Xu, Y.; Zhang, C.; Huang, S. Generation of self-compatible diploid potato by knockout of S-RNase. Nat. Plants 2018, 4, 651–654. [Google Scholar] [CrossRef] [PubMed]
  45. Del Mar Martínez-Prada, M.; Curtin, S.J.; Gutiérrez-González, J.J. Potato improvement through genetic engineering. GM Crops Food 2021, 12, 479–496. [Google Scholar] [CrossRef]
  46. Jiao, Q.; Xin, L.; Bai, H.; Zong, W.; Jia, T.; Hu, X. Heterologous expression of α-AtCAH1 improves resistance to drought stress and increases tuber yield in potato (Solanum tuberosum L.). Plant Biol. 2026, 28, 91–101. [Google Scholar] [CrossRef]
  47. Wang, C.; Hawthorne, D.; Qin, Y.; Pan, X.; Li, Z.; Zhu, S. Impact of climate and host availability on future distribution of Colorado potato beetle. Sci. Rep. 2017, 7, 4489. [Google Scholar] [CrossRef]
  48. Liao, J.; Liu, J.; Li, C. Effects of repeated short-term heat exposure on life history traits of Colorado potato beetle. Insects 2022, 13, 455. [Google Scholar] [CrossRef]
  49. Kramer, D.M.; Johnson, G.; Kiirats, O.; Edwards, G.E. New fluorescence parameters for the determination of QA redox state and excitation energy fluxes. Photosynth. Res. 2004, 79, 209–218. [Google Scholar] [CrossRef]
  50. Vass, I.; Cser, K.; Cheregi, O. Molecular mechanisms of light stress of photosynthesis. Ann. N. Y. Acad. Sci. 2007, 1113, 114–122. [Google Scholar] [CrossRef]
  51. Kato, M.C.; Hikosaka, K.; Hirotsu, N.; Makino, A.; Hirose, T. The excess light energy that is neither utilized in photosynthesis nor dissipated by photoprotective mechanisms determines the rate of photoinactivation in photosystem II. Plant Cell Physiol. 2003, 44, 318–325. [Google Scholar] [CrossRef] [PubMed]
  52. Hakala, M.; Tuominen, I.; Keränen, M.; Tyystjärvi, T.; Tyystjärvi, E. Evidence for the role of the oxygen-evolving manganese complex in photoinhibition of Photosystem II. Biochim. Biophys. Acta 2005, 1706, 68–80. [Google Scholar] [CrossRef] [PubMed]
  53. Ohnishi, N.; Allakhverdiev, S.I.; Takahashi, S.; Higashi, S.; Watanabe, M.; Nishiyama, Y.; Murata, N. Two-step mechanism of photodamage to photosystem II: Step 1 occurs at the oxygen-evolving complex and step 2 occurs at the photochemical reaction center. Biochemistry 2005, 44, 8494–8499. [Google Scholar] [CrossRef] [PubMed]
  54. Tyystjärvi, E. Photoinhibition of Photosystem II and photodamage of the oxygen evolving manganese cluster. Coord. Chem. Rev. 2008, 252, 361–376. [Google Scholar] [CrossRef]
  55. Oguchi, R.; Terashima, I.; Chow, W.S. The involvement of dual mechanisms of photoinactivation of photosystem II in Capsicum annuum L. plants. Plant Cell Physiol. 2009, 50, 1815–1825. [Google Scholar] [CrossRef]
  56. Vass, I. Role of charge recombination processes in photodamage and photoprotection of the photosystem II complex. Physiol. Plant. 2011, 142, 6–16. [Google Scholar] [CrossRef]
  57. Campbell, D.A.; Tyystjärvi, E. Parameterization of photosystem II photoinactivation and repair. BBA-Bioenergentics 2012, 1817, 258–265. [Google Scholar] [CrossRef]
  58. Zavafer, A.; Koinuma, W.; Chow, W.S.; Cheah, M.H.; Mino, H. Mechanism of photodamage of the oxygen evolving Mn cluster of photosystem II by excessive light energy. Sci. Rep. 2017, 7, 7604. [Google Scholar] [CrossRef]
  59. Vinyard, D.J.; Ananyev, G.M.; Dismukes, G.C. Photosystem II: The reaction center of oxygenic photosynthesis. Annu. Rev. Biochem. 2013, 82, 577–606. [Google Scholar] [CrossRef]
  60. Najafpour, M.M.; Ghobadi, M.Z.; Haghighi, B.; Eaton-Rye, J.J.; Tomo, T.; Shen, J.R.; Allakhverdiev, S.I. Nano-sized manganesecalcium cluster in photosystem II. Biochemistry 2014, 79, 324–336. [Google Scholar]
  61. Watanabe, S.; Noguchi, T. Intermediate formation via proton release during the photoassembly of the water-oxidizing Mn4CaO5 cluster in photosystem II. J. Phys. Chem. B 2024, 128, 8145–8161. [Google Scholar] [CrossRef] [PubMed]
  62. Oguchi, R.; Terashima, I.; Kou, J.; Chow, W.S. Operation of dual mechanisms that both lead to photoinactivation of photosystem II in leaves by visible light. Physiol. Plant. 2011, 142, 47–55. [Google Scholar] [CrossRef] [PubMed]
  63. Govindachary, S.; Bukhov, N.G.; Joly, D.; Carpentier, R. Photosystem II inhibition by moderate light under low temperature in intact leaves of chilling-sensitive and -tolerant plants. Physiol. Plant. 2004, 121, 322–333. [Google Scholar] [CrossRef] [PubMed]
  64. Pellegrini, E.; Carucci, M.G.; Campanella, A.; Lorenzini, G.; Nali, C. Ozone stress in Melissa officinalis plants assessed by photosynthetic function. Environ. Exp. Bot. 2011, 73, 94–101. [Google Scholar] [CrossRef]
  65. Siddiqui, H.; Ahmed, K.B.M.; Hayat, S. Comparative effect of 28-homobrassinolide and 24-epibrassinolide on the performance of different components influencing the photosynthetic machinery in Brassica juncea L. Plant Physiol. Biochem. 2018, 129, 198–212. [Google Scholar] [CrossRef]
  66. Mosadegh, H.; Trivellini, A.; Lucchesini, M.; Ferrante, A.; Maggini, R.; Vernieri, P.; Mensuali Sodi, A. UV-B physiological changes under conditions of distress and eustress in sweet basil. Plants 2019, 8, 396. [Google Scholar] [CrossRef]
  67. Gohari, G.; Farhadi, H.; Panahirad, S.; Zareei, E.; Labib, P.; Jafari, H.; Mahdavinia, G.; Hassanpouraghdam, M.B.; Ioannou, A.; Kulak, M.; et al. Mitigation of salinity impact in spearmint plants through the application of engineered chitosan-melatonin nanoparticles. Int. J. Biol. Macromol. 2023, 224, 893–907. [Google Scholar] [CrossRef]
  68. Kalisz, A.; Kornaś, A.; Skoczowski, A.; Oliwa, J.; Jurkow, R.; Gil, J.; Sękara, A.; Sałata, A.; Caruso, G. Leaf chlorophyll fluorescence and reflectance of oakleaf lettuce exposed to metal and metal(oid) oxide nanoparticles. BMC Plant Biol. 2023, 23, 329. [Google Scholar] [CrossRef]
  69. Zia, A.; Farrag, E.S.; Mahmoud, S.Y. Dieback of royal poinciana (Delonix regia) trees induced by Alternaria tenuissima and its impact on photochemical efficiency of photosystem II. Physiol. Mol. Plant Pathol. 2024, 133, 102357. [Google Scholar] [CrossRef]
  70. Tóth, S.Z.; Nagy, V.; Puthur, J.T.; Kovács, L.; Garab, G. The physiological role of ascorbate as photosystem II electron donor: Protection against photoinactivation in heat-stressed leaves. Plant Physiol. 2011, 156, 382–392. [Google Scholar] [CrossRef]
  71. Széles, E.; Kuntam, S.; Vidal-Meireles, A.; Nagy, V.; Nagy, K.; Ábrahám, Á.; Kovács, L.; Tóth, S.Z. Single-cell microfluidics in combination with chlorophyll a fluorescence measurements to assess the lifetime of the Chlamydomonas PSBO protein. Photosynthetica 2023, 61, 417–424. [Google Scholar] [CrossRef]
  72. Callahan, F.E.; Becker, D.W.; Cheniae, G.M. Studies on the photo-inactivation of the water-oxidizing enzyme. II. Characterization of weak light photoinhibition of PSII and its light-induced recovery. Plant Physiol. 1986, 82, 261–269. [Google Scholar] [CrossRef] [PubMed]
  73. Chen, G.X.; Kazimir, J.; Cheniae, G.M. Photoinhibition of hydroxylamine-extracted photosystem II membranes: Studies of the mechanism. Biochemistry 1992, 31, 11072–11083. [Google Scholar] [CrossRef] [PubMed]
  74. Anderson, J.M.; Park, Y.I.; Chow, W.S. Unifying model for the photoinactivation of photosystem II in vivo: A hypothesis. Photosynth. Res. 1998, 56, 1–13. [Google Scholar] [CrossRef]
  75. Sarvikas, P.; Hakala, M.; Pätsikkä, E.; Tyystjärvi, T.; Tyystjärvi, E. Action spectrum of photoinhibition in leaves of wild type and npq1-2 and npq4-1 mutants of Arabidopsis thaliana. Plant Cell Physiol. 2006, 47, 391–400. [Google Scholar] [CrossRef]
  76. Hamdani, S.; Khan, N.; Perveen, S.; Qu, M.; Jiang, J.; Govindjee; Zhu, X.G. Changes in the photosynthesis properties and photoprotection capacity in rice (Oryza sativa) grown under red, blue, or white light. Photosynth. Res. 2019, 139, 107–121. [Google Scholar] [CrossRef]
  77. Garab, G.; Magyar, M.; Sipka, G.; Lambrev, P.H. New foundations for the physical mechanism of variable chlorophyll a fluorescence. Quantum efficiency versus the light-adapted state of photosystem II. J. Exp. Bot. 2023, 74, 5458–5471. [Google Scholar] [CrossRef]
  78. Klughammer, C.; Schreiber, U. Complementary PS II quantum yields calculated from simple fluorescence parameters measured by PAM fluorometry and the Saturation Pulse method. PAM Appl. Notes 2008, 1, 27–35. [Google Scholar]
  79. Kasajima, I.; Ebana, K.; Yamamoto, T.; Takahara, K.; Yano, M.; Kawai-Yamada, M.; Uchimiya, H. Molecular distinction in genetic regulation of nonphotochemical quenching in rice. Proc. Natl. Acad. Sci. USA 2011, 108, 13835–13840. [Google Scholar] [CrossRef]
  80. Gawroński, P.; Witoń, D.; Vashutina, K.; Bederska, M.; Betliński, B.; Rusaczonek, A.; Karpiński, S. Mitogen-activated protein kinase 4 is a salicylic acid-independent regulator of growth but not of photosynthesis in Arabidopsis. Mol. Plant 2014, 7, 1151–1166. [Google Scholar] [CrossRef]
  81. Demmig-Adams, B.; Adams, W.W., III. Photoprotection and other responses of plants to high light stress. Annu. Rev. Plant Physiol. Plant Mol. Biol. 1992, 43, 599–626. [Google Scholar] [CrossRef]
  82. Krieger-Liszkay, A. Singlet oxygen production in photosynthesis. J. Exp. Bot. 2005, 56, 337–346. [Google Scholar] [CrossRef] [PubMed]
  83. Ogilby, P.R. Singlet oxygen: There is indeed something new under the sun. Chem. Soc. Rev. 2010, 39, 3181–3209. [Google Scholar] [CrossRef] [PubMed]
  84. Telfer, A. Singlet oxygen production by PSII under light stress: Mechanism, detection and the protective role of β-carotene. Plant Cell Physiol. 2014, 55, 1216–1223. [Google Scholar] [CrossRef]
  85. Barber, J.; Andersson, B. Too much of a good thing: Light can be bad for photosynthesis. Trends Biochem. Sci. 1992, 17, 61–66. [Google Scholar] [CrossRef]
  86. Aro, E.M.; Virgin, I.; Andersson, B. Photoinhibition of Photosystem II. Inactivation, protein damage and turnover. Biochim. Biophys. Acta 1993, 1143, 113–134. [Google Scholar] [CrossRef]
  87. Yamamoto, Y. Quality control of photosystem II. Plant Cell Physiol. 2001, 42, 121–128. [Google Scholar] [CrossRef]
  88. Yamamoto, Y.; Aminaka, R.; Yoshioka, M.; Khatoon, M.; Komayama, K.; Takenaka, D.; Yamashita, A.; Nijo, N.; Inagawa, K.; Morita, N.; et al. Quality control of photosystem II: Impact of light and heat stresses. Photosynth. Res. 2008, 98, 589–608. [Google Scholar] [CrossRef]
  89. Chan, T.; Shimizu, Y.; Pospíšil, P.; Nijo, N.; Fujiwara, A.; Taninaka, Y.; Ishikawa, T.; Hori, H.; Nanba, D.; Imai, A.; et al. Quality control of photosystem II: Lipid peroxidation accelerates photoinhibition under excessive illumination. PLoS ONE 2012, 7, e52100. [Google Scholar] [CrossRef]
  90. Velikova, V.; Salerno, G.; Frati, F.; Peri, E.; Conti, E.; Colazza, S.; Loreto, F. Influence of feeding and oviposition by phytophagous pentatomids on photosynthesis of herbaceous plants. J. Chem. Ecol. 2010, 36, 629–641. [Google Scholar] [CrossRef]
  91. Schröder, R.; Forstreuter, M.; Hilker, M. A plant notices insect egg deposition and changes its rate of photosynthesis. Plant Physiol. 2005, 138, 470–477. [Google Scholar] [CrossRef][Green Version]
  92. Nabity, P.D.; Zavala, J.A.; DeLucia, E.H. Herbivore induction of jasmonic acid and chemical defences reduce photosynthesis in Nicotiana attenuata. J. Exp. Bot. 2013, 64, 685–694. [Google Scholar] [CrossRef] [PubMed]
  93. Pinnola, A.; Bassi, R. Molecular mechanisms involved in plant photoprotection. Biochem. Soc. Trans. 2018, 46, 467–482. [Google Scholar] [CrossRef] [PubMed]
  94. Khan, I.; Sohail; Zaman, S.; Li, G.; Fu, M. Adaptive responses of plants to light stress: Mechanisms of photoprotection and acclimation. A review. Front. Plant Sci. 2025, 16, 1550125. [Google Scholar] [CrossRef] [PubMed]
  95. Kumari, A.; Njuguna, J.N.; Zheng, X.; Kromdijk, J.; Sacks, E.J.; Glowacka, K. Genetic basis of non-photochemical quenching and photosystem II efficiency responses to chilling in the biomass crop Miscanthus. GCB Bioenergy 2025, 17, e70015. [Google Scholar] [CrossRef]
  96. Zuo, G. Non-photochemical quenching (NPQ) in photoprotection: Insights into NPQ levels required to avoid photoinactivation and photoinhibition. New Phytol. 2025, 246, 1967–1974. [Google Scholar] [CrossRef]
  97. Schreiber, U.; Klughammer, C. Non-photochemical fluorescence quenching and quantum yields in PSI and PSII: Analysis of heat-induced limitations using Maxi-Imaging PAM and Dual-PAM-100. PAM Appl. Notes 2008, 1, 15–18. [Google Scholar]
  98. Ruban, A.V. Nonphotochemical chlorophyll fluorescence quenching: Mechanism and effectiveness in protecting plants from photodamage. Plant Physiol. 2016, 170, 1903–1916. [Google Scholar] [CrossRef]
  99. Ruban, A.V. Light harvesting control in plants. FEBS Lett. 2018, 592, 3030–3039. [Google Scholar] [CrossRef]
  100. Lambrev, P.H.; Miloslavina, Y.; Jahns, P.; Holzwarth, A.R. On the relationship between non-photochemical quenching and photoprotection of photosystem II. Biochim. Biophys. Acta 2012, 1817, 760–769. [Google Scholar] [CrossRef]
  101. Moustakas, M.; Panteris, E.; Moustaka, J.; Aydın, T.; Bayçu, G.; Sperdouli, I. Modulation of photosystem II function in celery via foliar-applied salicylic acid during gradual water deficit stress. Int. J. Mol. Sci. 2024, 25, 6721. [Google Scholar] [CrossRef] [PubMed]
  102. Asada, K. Production and scavenging of reactive oxygen species in chloroplasts and their functions. Plant Physiol. 2006, 141, 391–396. [Google Scholar] [CrossRef] [PubMed]
  103. Mittler, R.; Vanderauwera, S.; Suzuki, N.; Miller, G.; Tognetti, V.B.; Vandepoele, K.; Gollery, M.; Shulaev, V.; Van Breusegem, F. ROS signaling: The new wave? Trends Plant Sci. 2011, 16, 300–309. [Google Scholar] [CrossRef] [PubMed]
  104. Foyer, C.H.; Noctor, G. Redox signaling in plants. Antioxid. Redox Signal. 2013, 18, 2087–2090. [Google Scholar] [CrossRef]
  105. Zandalinas, S.I.; Mittler, R. ROS-induced ROS release in plant andanimal cells. Free Radic. Biol. Med. 2018, 22, 21–27. [Google Scholar] [CrossRef]
  106. Janků, M.; Luhová, L.; Petřivalský, M. On the origin and fate of reactive oxygen species in plant cell compartments. Antioxidants 2019, 8, 105. [Google Scholar] [CrossRef]
  107. Fedoreyeva, L.I. ROS as signaling molecules to initiate the process of plant acclimatization to abiotic stress. Int. J. Mol. Sci. 2024, 25, 11820. [Google Scholar] [CrossRef]
  108. Mittler, R.; Berkowitz, G. Hydrogen peroxide, a messenger with too many roles? Redox Rep. 2001, 6, 69–72. [Google Scholar] [CrossRef]
  109. Li, H.; Jiang, X.; Lv, X.; Ahammed, G.J.; Guo, Z.; Qi, Z.; Yu, J.; Zhou, Y. Tomato GLR3.3 and GLR3.5 mediate cold acclimation induced chilling tolerance by regulating apoplastic H2O2 production and redox homeostasis. Plant Cell Environ. 2019, 42, 3326–3339. [Google Scholar] [CrossRef]
  110. Foyer, C.H. How plant cells sense the outside world through hydrogen peroxide. Nature 2020, 578, 518–519. [Google Scholar] [CrossRef]
  111. Ivanov, B.N.; Borisova-Mubarakshina, M.M.; Kozuleva, M.A. Formation mechanisms of superoxide radical and hydrogen peroxide in chloroplasts, and factors determining the signalling by hydrogen peroxide. Funct. Plant Biol. 2018, 45, 102–110. [Google Scholar] [CrossRef] [PubMed]
  112. Smirnoff, N.; Arnaud, D. Hydrogen peroxide metabolism and functions in plants. New Phytol. 2019, 221, 1197–1214. [Google Scholar] [CrossRef] [PubMed]
  113. Breeze, E.; Mullineaux, P.M. The Passage of H2O2 from chloroplasts to their associated nucleus during retrograde signalling: Reflections on the role of the nuclear envelope. Plants 2022, 11, 552. [Google Scholar] [CrossRef] [PubMed]
  114. Wilson, K.E.; Ivanov, A.G.; Öquist, G.; Grodzinski, B.; Sarhan, F.; Huner, N.P.A. Energy balance, organellar redox status, and acclimation to environmental stress. Can. J. Bot. 2006, 84, 1355–1370. [Google Scholar] [CrossRef]
  115. Adamakis, I.D.S.; Sperdouli, I.; Eleftheriou, E.P.; Moustakas, M. Hydrogen peroxide production by the spot-like mode action of bisphenol A. Front. Plant Sci. 2020, 11, 1196. [Google Scholar] [CrossRef]
  116. Mullineaux, P.M.; Exposito-Rodriguez, M.; Laissue, P.P.; Smirnoff, N. ROS-dependent signalling pathways in plants and algae exposed to high light: Comparisons with other eukaryotes. Free Radic. Biol. Med. 2018, 122, 52–64. [Google Scholar] [CrossRef]
  117. Karim, M.F.; Johnson, G.N. Acclimation of photosynthesis to changes in the environment results in decreases of oxidative stress in Arabidopsis thaliana. Front. Plant Sci. 2021, 12, 683986. [Google Scholar] [CrossRef]
  118. Foyer, C.H.; Hanke, G. ROS production and signalling in chloroplasts: Cornerstones and evolving concepts. Plant J. 2022, 111, 642–661. [Google Scholar] [CrossRef]
  119. Gorecka, M.; Alvarez-Fernandez, R.; Slattery, K.; McAusland, L.; Davey, P.A.; Karpinski, S.; Lawson, T.; Mullineaux, P.M. Abscisic acid signalling determines susceptibility of bundle sheath cells to photoinhibition in high light-exposed Arabidopsis leaves. Philos. Trans. R. Soc. Lond. B Biol. Sci. 2014, 369, 20130234. [Google Scholar] [CrossRef]
  120. Baxter, A.; Mittler, R.; Suzuki, N. ROS as key players in plant stress signaling. J. Exp. Bot. 2014, 65, 1229–1240. [Google Scholar] [CrossRef]
  121. Dietz, K.J.; Pfannschmidt, T. Novel regulators in photosynthetic redox control of plant metabolism and gene expression. Plant Physiol. 2011, 155, 1477−1485. [Google Scholar] [CrossRef]
  122. Ali, M.; Cheng, Z.; Ahmad, H.; Hayat, S. Reactive oxygen species (ROS) as defenses against a broad range of plant fungal infections and case study on ROS employed by crops against Verticillium dahliae wilts. J. Plant Interact. 2018, 13, 353–363. [Google Scholar] [CrossRef]
  123. Erb, M.; Reymond, P. Molecular interactions between plants and insect herbivores. Annu. Rev. Plant Biol. 2019, 70, 527–557. [Google Scholar] [CrossRef] [PubMed]
  124. Pang, S.; Zhai, J.; Song, J.; Rong, D.; Hong, Y.; Qiu, Y.; Ma, J.; Qi, T.; Huang, H.; Song, S. bHLH19 and bHLH20 repress jasmonate-mediated plant defense against insect herbivores in Arabidopsis. Plant J. 2024, 120, 2623–2638. [Google Scholar] [CrossRef] [PubMed]
  125. Fürstenberg-Hägg, J.; Zagrobelny, M.; Bak, S. Plant Defense against Insect Herbivores. Int. J. Mol. Sci. 2013, 14, 10242–10297. [Google Scholar] [CrossRef] [PubMed]
  126. Wasternack, C.; Feussner, I. The oxylipin pathways: Biochemistry and function. Annu. Rev. Plant Biol. 2018, 69, 363–386. [Google Scholar] [CrossRef]
  127. Zhu, L.; Yang, H.; Li, P.; Dong, L.; Zhao, S.; Lv, H.; Crickmore, N.; Zhou, X.; Zhang, Y.; Guo, Z. Plant strategies against herbivorous insects. J. Integr. Plant Biol. 2026, in press. [Google Scholar] [CrossRef]
  128. Hause, B.; Stenzel, I.; Miersch, O.; Maucher, H.; Kramell, R.; Ziegler, J.; Wasternack, C. Tissue-specific oxylipin signature of tomato flowers: Allene oxide cyclase is highly expressed in distinct flower organs and vascular bundles. Plant J. 2000, 24, 113–126. [Google Scholar] [CrossRef]
  129. Dorey, T.; Deluigi, J.; Maccagni, A.; Rasmann, S.; Glauser, G.; Willi, Y. Variation in herbivore defense strategies among plant species differing in elevational distribution and the role of temperature in defense. New Phytol. 2026, 249, 3091–3103. [Google Scholar] [CrossRef]
  130. Shinde, S.; Kundu, P.; Shrestha, K.; Kaler, E.; Verma, K.; Block, A.K.; Louis, J. Flavonoids at the crossroads of plant defense: A multifunctional shield against insect pests. J. Exp. Bot. 2026, erag096. [Google Scholar] [CrossRef]
  131. Moustaka, J.; Panteris, E.; Adamakis, I.D.S.; Tanou, G.; Giannakoula, A.; Eleftheriou, E.P.; Moustakas, M. High anthocyanin accumulation in poinsettia leaves is accompanied by thylakoid membrane unstacking, acting as a photoprotective mechanism, to prevent ROS formation. Environ. Exp. Bot. 2018, 154, 44–55. [Google Scholar] [CrossRef]
  132. Oxborough, K.; Baker, N.R. Resolving chlorophyll a fluorescence images of photosynthetic efficiency into photochemical and non-photochemical components—Calculation of qP and Fv’/Fm’ without measuring Fo’. Photosynth. Res. 1997, 54, 135–142. [Google Scholar] [CrossRef]
  133. Moustaka, J.; Tanou, G.; Adamakis, I.-D.; Eleftheriou, E.P.; Moustakas, M. Leaf age-dependent photoprotective and antioxidative response mechanisms to paraquat-induced oxidative stress in Arabidopsis thaliana. Int. J. Mol. Sci. 2015, 16, 13989–14006. [Google Scholar] [CrossRef]
Plants 15 01159 g005
Figure 5. The relationship of the maximum efficiency of PSII photochemistry (Fv/Fm) with the efficiency of the oxygen-evolving complex (Fv/Fo) before (a) and after 20 min of Colorado potato beetle feeding on potato leaves (b). The blue dots represent the corresponding measurements of the variables, while the red line is the regression line that shows the relationship between the two variables.
Figure 5. The relationship of the maximum efficiency of PSII photochemistry (Fv/Fm) with the efficiency of the oxygen-evolving complex (Fv/Fo) before (a) and after 20 min of Colorado potato beetle feeding on potato leaves (b). The blue dots represent the corresponding measurements of the variables, while the red line is the regression line that shows the relationship between the two variables.
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Plants 15 01159 g006
Figure 6. H2O2 production in potato leaves before (a) and after 20 min of Colorado potato beetle feeding (b,c). The light green color denotes H2O2 generation. The magnification in (c) is the feeding spot area.
Figure 6. H2O2 production in potato leaves before (a) and after 20 min of Colorado potato beetle feeding (b,c). The light green color denotes H2O2 generation. The magnification in (c) is the feeding spot area.
Plants 15 01159 g006
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MDPI and ACS Style

Sperdouli, I.; Andreadis, S.S.; Moustaka, J.; Koutsogeorgiou, E.I.; Panteris, E.; Moustakas, M. Photosystem II Responses at the Whole-Potato-Leaf Level After Colorado Potato Beetle Feeding. Plants 2026, 15, 1159. https://doi.org/10.3390/plants15081159

AMA Style

Sperdouli I, Andreadis SS, Moustaka J, Koutsogeorgiou EI, Panteris E, Moustakas M. Photosystem II Responses at the Whole-Potato-Leaf Level After Colorado Potato Beetle Feeding. Plants. 2026; 15(8):1159. https://doi.org/10.3390/plants15081159

Chicago/Turabian Style

Sperdouli, Ilektra, Stefanos S. Andreadis, Julietta Moustaka, Eleni I. Koutsogeorgiou, Emmanuel Panteris, and Michael Moustakas. 2026. "Photosystem II Responses at the Whole-Potato-Leaf Level After Colorado Potato Beetle Feeding" Plants 15, no. 8: 1159. https://doi.org/10.3390/plants15081159

APA Style

Sperdouli, I., Andreadis, S. S., Moustaka, J., Koutsogeorgiou, E. I., Panteris, E., & Moustakas, M. (2026). Photosystem II Responses at the Whole-Potato-Leaf Level After Colorado Potato Beetle Feeding. Plants, 15(8), 1159. https://doi.org/10.3390/plants15081159

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