*Review* **Deciphering the Role of Ion Channels in Early Defense Signaling against Herbivorous Insects**

**Akanksha Gandhi 1, Rupesh Kariyat <sup>1</sup> , Amaravadhi Harikishore 2, Marzieh Ayati <sup>3</sup> , Anirban Bhunia <sup>4</sup> and Nirakar Sahoo 1,\***


**Abstract:** Plants and insect herbivores are in a relentless battle to outwit each other. Plants have evolved various strategies to detect herbivores and mount an effective defense system against them. These defenses include physical and structural barriers such as spines, trichomes, cuticle, or chemical compounds, including secondary metabolites such as phenolics and terpenes. Plants perceive herbivory by both mechanical and chemical means. Mechanical sensing can occur through the perception of insect biting, piercing, or chewing, while chemical signaling occurs through the perception of various herbivore-derived compounds such as oral secretions (OS) or regurgitant, insect excreta (frass), or oviposition fluids. Interestingly, ion channels or transporters are the first responders for the perception of these mechanical and chemical cues. These transmembrane pore proteins can play an important role in plant defense through the induction of early signaling components such as plasma transmembrane potential (Vm) fluctuation, intracellular calcium (Ca2+), and reactive oxygen species (ROS) generation, followed by defense gene expression, and, ultimately, plant defense responses. In recent years, studies on early plant defense signaling in response to herbivory have been gaining momentum with the application of genetically encoded GFP-based sensors for realtime monitoring of early signaling events and genetic tools to manipulate ion channels involved in plant-herbivore interactions. In this review, we provide an update on recent developments and advances on early signaling events in plant-herbivore interactions, with an emphasis on the role of ion channels in early plant defense signaling.

**Keywords:** reactive oxygen species; herbivory; membrane potential; ion channel

### **1. Introduction**

Plants regularly encounter a wide range of abiotic and biotic stresses in nature. Abiotic stress includes drought, salinity, extreme temperatures, radiation, floods, and heavy metals, whereas biotic stressors include insect, animal herbivores, and microbial pathogens. Plant and insect-herbivore interactions are among the most significant species interactions found in nature [1,2], and it is estimated that, annually, herbivory causes a 20% loss in the total productivity of agricultural crops [3]. However, plants are not totally defenseless against herbivory and are able to perceive and respond to this onslaught. They can perceive the insect attack through both mechanical and chemical cues. Mechanical signals are elicited through the damage caused by herbivores by piercing, chewing, or biting of plant tissues, and chemical signals are relayed via herbivore-associated elicitors (HAEs) such as oral secretions (OS) or regurgitant, insect excreta (frass), or oviposition fluids, to name a few [4,5]. Plants not only actively respond to herbivory, but also initiate a series of biochemical

**Citation:** Gandhi, A.; Kariyat, R.; Harikishore, A.; Ayati, M.; Bhunia, A.; Sahoo, N. Deciphering the Role of Ion Channels in Early Defense Signaling against Herbivorous Insects. *Cells* **2021**, *10*, 2219. https://doi.org/ 10.3390/cells10092219

Academic Editor: Suleyman Allakhverdiev

Received: 1 July 2021 Accepted: 20 August 2021 Published: 27 August 2021

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**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

responses following the perception of herbivory. These biochemical cascades are initiated through ion channels that control the changes in the plasma membrane potential (Vm), generation of reactive oxygen species (ROS), cytosolic calcium fluxes, and ultimately induce plant defense genes to mount a multi-layered defense response that can act at both local and systemic levels [4,6–10]. In recent years, there have been several reviews on plantherbivore interactions [4,5,7,11–26]. Here we complement these existing reviews with current research and recent discoveries on plant-herbivore interactions, focusing on early plant defense signaling, with a particular emphasis on ion channels involved in early plant defense signaling.

### **2. Long-Distance Communication in Plant Defense**

During herbivory, the damaged areas of the plant need to inform the rest of the plant to keep them ready for the imminent herbivory threat. Therefore, plants need to alert their unaffected parts by sending long-distance signals from the site of damage to various parts of the plant to appraise the threat. Plants respond to diverse stimuli by communicating amongst cells from distinct tissues or organs, a process called systemic signaling [27]. Studies have revealed the existence of complex regulatory mechanisms that allow the plant to activate resistance in systemic tissues, commonly referred to as systemic acquired resistance (SAR) [28]. SAR is characterized by a more potent and faster response to future encounters with microbes, insects, or abiotic stress.

Considerable progress has been made in understanding this intricate relationship between plants and herbivores with a plethora of field and lab studies. These include studies that have dissected pairwise interactions between a specific herbivore and its host; interactions at species, genus, and community levels with multiple hosts and herbivores; and studies examining plant defense signaling networks through molecular genetics genomics, to name a few [29,30]. However, our knowledge of how plants perceive these cues and how that leads to specific and tightly regulated defense responses is still in its infancy. It has been proposed that following the insect attack, the foremost event is the recognition of the cue and its perception by specific membrane receptors and the transduction of these signals into the plant cell. These cues are termed as "early defense signaling molecules" such as the depolarization of plasma membrane along with the generation of secondary messengers such as cytosolic Ca2+ [31], reactive oxygen species (ROS), and reactive nitrogen species (RNS) [32–35] that contribute to plant defense signal transduction events.

Long-distance communication in plants has been linked with ion channels or membrane transporters. These are transmembrane pore proteins involved in the movement of ions across the cell membrane. In recent years, with electrophysiological tools, the research on ion channels in plants has been gaining momentum. Studies have reported that ion channels facilitate long-distance communication via Vm, Ca2+, and ROS (Figure 1). Ion channels have been shown to mediate systemic signaling by modulating the influx of ions into different plant tissues [36]. They sense signals from the functional cells at the site of herbivory to activate other cells, which in turn relay this signal to induce defense responses. For example, a recent study [37] identified glutamate receptor-like channels (GLRs) in *Arabidopsis thaliana* that are related to mammalian ionotropic glutamate receptors, play a role in Ca2+ signaling during herbivory, nutrient transport, root gravitropism, and plant defense [38,39]. However, in mammals, these channels are involved in neurotransmission, and their openings are stimulated by glutamate binding to the postsynaptic neuron, resulting in Ca2+ and other cations influx. The signal is transmitted because of voltage changes caused by ion flux [40]. Remarkably, these GLRs are also responsible for long-distance Ca2+ transmission in plants in response to herbivory or mechanical injury, efficiently communicating herbivore attacks to surrounding cells.

**Figure 1.** Initiation of early defense signaling mechanisms in response to insect herbivore attack. Schematic diagram showing herbivore *M. sexta* feeding induced signaling events, which include the perception of HAEs such as OS, frass, and oviposition by specialized receptors on the outer plasma membrane, which trigger modulation of Vm via H+-ATPase and Ca2+ ion influx into the cell via Ca2+ channels, GLR3.3/3.6 and/or CNGC19. The increase in cytosolic Ca2+ may trigger the further release of vacuolar Ca2+ via the TPC1 channel. The subsequent release of Ca2+ may activate nicotinamide adenine dinucleotide phosphate (NADPH oxidase) and respiratory burst oxidase homologues (RBOHDs), leading to ROS generation, and induction of plant defense responses. Illustration by Annette Diaz.

There has been considerable research on identifying the factors that are involved in long-distance signaling. Plants can appraise their unaffected parts by extensive network of intracellular regulators, Vm, Ca2+, and ROS [18,41]. The transmission rate of all these waves ranges from ~100 to >1000 μm/sec [41,42]. The process starts with the propagation of long-distance electrical signals as a result of variation in membrane potential due to potassium (K+) and Ca2+ flux. Variation in Vm is critical for plant wounding responses [43]. Finally, Ca2+ and ROS, versatile secondary messenger, were generated that plants use to sense and transform environmental stimuli into an adaptive intracellular response [44]. Insect feeding and OS can lead to changes to the cytosolic Ca2+ concentration, and these spatiotemporal variations have been shown to yield Ca2+ signatures [45–48]. On the other hand, ROS are extremely reactive and hazardous chemicals formed from oxygen. Among them are O2, H2O2, and OH−. ROS which has been demonstrated to act as a self-propagating long-distance and fast wound signal [49]. Throughout this review, we will focus on the role of ion channels, Vm, Ca2+, and ROS in plant response to herbivory and provide an overview of what is currently known about the role of ion channels in plant-herbivore interactions.

### **3. Membrane Potential (Vm)**

The Vm is an electrical potential of the cell membrane that is maintained via the balance of ion fluxes across the plasma membrane. Vm indicates whether a cell is excited or not. It is responsible for generating action potentials in tissues, muscles, and nerves in animals and plays a crucial role in diverse biological functions such as biological sensing, hearing, cell cycle, proliferation, contractility, and circadian rhythm, to name a few [50]. Unlike animals, plants use Vm to regulate plant cellular functions such as maintaining turgor pressure, osmotic balance, and stomatal closure. There is no net flux of ions through the membrane when in equilibrium, called the resting membrane potential. Changes in the resting membrane potential will occur due to an unbalanced movement of ions, thus leading to Vm being more positive (depolarization) or more negative (hyperpolarization). In general, plants maintain a negative resting membrane potential in the order of −110 to −150 mV [51,52]. It has been reported that the signal transduction mechanism of plants to respond to minor changes in Vm leads to plant defense responses. The way

plants sense insect cues and initiate defense responses has been a point of interest for many years. One hypothesis that has evolved by studying cellular responses following herbivory suggests that the first event following herbivory generates the fluctuation in Vm [53]. Maffei et al. [43] has also demonstrated that both mechanical wounding and OS of cotton leafworm (*Spodoptera littoralis*) alter Vm in lima bean (*Phaseolus lunatus* L.) at increasing distances of 5, 30, and 60 mm from the bite zone. Vm depolarization was observed within the first 15 min of feeding by *S. littoralis* in the palisade cells. The effect of *S. littoralis* regurgitant and its components were also tested on Vm in *P. lunatus* leaf and the results showed that Vm alterations were independent of regurgitate concentration. In addition, they also examined changes in Vm in response to the application of various H2O2 concentrations to mechanically damaged and herbivore-wounded *P. lunatus* leaves. H2O2 treatment induced a robust Vm that was significantly greater in herbivory-wounded plants than in mechanically injured leaves [54].

Bricchi et al. [55] studied Vm alterations in wild-type and plasmodesmata mutated *A. thaliana pdko3* lines; plasmodesmata are channels within the plant cell that allow chemicals to pass through, establishing a pathway for cell-to-cell communication. A strong Vm depolarization occurred in wild-type *A. thaliana* plants within 7 to 8 min after herbivory, but the *pdko3* mutant did not exhibit Vm depolarization in response to herbivory or application of OS from *S. littoralis*. However, Ca2+ elevation was observed in both wild types as well as in *pdko3* mutant. This observation ruled out the possibility of Ca2+ channels being involved in Vm depolarization. To dissect the dependence of Vm depolarization on potassium (K+) channels, the K+ channel activity was measured using fluorescent indicator FluxORTM. A significant increase in K+ channel activity was observed in wild-type plants, whereas a complete loss of K+ channel activity was observed in *pdko3* plants. This finding also suggests that K<sup>+</sup> channels are involved in Vm depolarization and supports the hypothesis that plant cells respond to OS by a Vm-mediated signal transduction pathway.

The fluctuation in Vm has been known to be induced by the binding of specific components from herbivore OS with the receptors present at the plasma membrane [56]. These components can alter ion channel activities, causing an imbalance in ion movement, which influences the membrane potential of the plasma membrane [43]. A study by Mohanta et al. [57] showed that Kew tree (*Ginkgo biloba*), a living fossil plant, responds to *S. littoralis* herbivory by inducing Vm depolarization, which was evident up to 6 h. Another study using *A. thaliana* also showed that the extent of Vm depolarization was the same for *S. littoralis*, green peach aphid (*Myzus persicae)*, and the plant pathogenic bacteria *Pseudomonas syringe*, but the timing of the occurrence of Vm depolarization was different for each of these biotrophs. Moreover, the magnitude of early defense response depends upon the amount of tissue damage by the biotroph. Vm depolarization was rapid upon the attack of chewing herbivore, *S. littoralis* (30 min to 2 h), as it caused substantial tissue loss, since it consumed large amounts of leaf tissue. On the other hand, less damage was observed by a phloem feeder, *M. persicae* (4 to 6 h), that delayed the plant defense response since phloem feeders with sucking mouthparts feed on vascular tissues without visible tissue damage as observed with chewing herbivores [58]. It is apparent that Ca2+ and ROS generation are directly tied to Vm when herbivores interact with plants, and Vm is essential for plant defense responses.

### **4. Calcium (Ca2+)**

Ca2+ is a ubiquitous signaling molecule in plants. It functions as a secondary messenger in cellular pathways that regulate plant growth and development, cell polarity, cytoskeleton organization, ion transport across membranes, stomatal regulation, root growth, fertilization, nutrient signaling, and plant immunity [59]. Consequently, each of these processes has its own "Ca2+ signature," linked with distinct fluctuations in Ca2+ concentration in the cytosol and sometimes in a particular intracellular compartment. Therefore, Ca2+ fluxes, especially oscillations between calcium stores and the cytosol, are important for cell signaling [60–62].

In plants, the cytosolic Ca2+ concentration is maintained at or below 100 nM; however, the majority of Ca2+ is stored in the apoplast, vacuole, endoplasmic reticulum (ER), and Golgi apparatus. The apoplast serves as the first Ca2+ reservoir of a cell that can store 0.33 mM free resting Ca2+ and the first area that responds to stimuli, while the vacuole serves as the largest Ca2+ pool of a cell that can store up to 0.2–5 mM free resting Ca2+ [60,63].

The Ca2+ signature plays an important role in long-distance signal transduction during herbivore attack through which HAEs such as OS, oviposition, and frass is sensed by the cell membrane, and then, a Ca2+ is rapidly propagated in the cytosol and travels throughout the plant to induce defense responses. The shaping of this "Ca2+ signature" during plantherbivore interactions is achieved through the amplification and integration of Ca2+ signals. The amplification step is mediated via specific ion channels or transporter proteins and enhances Ca2+ fluxes at sites of herbivore attack, whereas the integration step is mediated via Ca2+ sensor proteins, which allow efficient transmission of Ca2+ signals from one cell to another in a tissue or organ. Herbivory induces Ca2+ entry from the apoplast to the cytosol via plasma membrane Ca2+ channels which stimulates Ca2+ signals in the cytosol leading to the amplification of Ca2+ signals. The localized Ca2+ signals from the cytosol are distributed throughout the whole plant. In this way, amplification, and integration of Ca2+ signals constitute two important ways by which "Ca2+ signature" contributes as a signaling molecule during plant-herbivore interactions [64].

The amplification of intracellular Ca2+ signal requires selective Ca2+ sensor proteins that respond to changes in cytosolic Ca2+ levels and encipher the frequency, amplitude, and signal localization of Ca2+ signatures. It is estimated that *A. thaliana* contains around 250 Ca2+ sensor proteins [65]. These can be classified into three main categories: (1) the calcineurin B-like proteins (CBLs) [66]; (2) the calmodulin (CaM), and calmodulin-like proteins (CMLs) [67]; and (3) the Ca2+ dependent protein kinases (CPKs) and the Ca2+ and calmodulin-dependent protein kinase (CCPK) [68]. All of these sensors contain EF-hand motifs, which enable Ca2+ binding and cause conformational changes in their structure [69].

CaM functions as a sensor relay protein since it lacks an enzymatic function. The *Arabidopsis* genome has seven calmodulin genes encoding four different isoforms (CaM1/4; CaM2/3/5; CaM6; and CaM7) [70]. CaM/CaM-like proteins (CML) regulate a variety of transcription factors, protein kinases, phosphatases, metabolic enzymes, ion pumps, and ion exchangers [71]. *A. thaliana* signal responsive (AtSR1) proteins [67], also known as CaM-binding transcription activators (AtCAMTAs) [72], have been shown to participate in wound-mediated defense responses. *Atsr1* mutants of *A. thaliana* were sensitive to attack by dark winged fungus gnats (*Bradysia impatiens*), suggesting the role of CaM as an important sensor in the early stages of the insect-plant attack [73]. Along with CaM, the plant has CML that undergo secondary structural changes in response to Ca2+ binding and act as Ca2+ relays/sensors [74]. CML and CAM share a 16% amino acid sequence similarity and include two to six EF-hand motif [70]. CML42 gene expression was shown to be increased in *A. thaliana* upon *S. littoralis* OS treatment, implying a function in early defense plant signaling [75]. CPKs have been classified as sensor responders because they combine a Ca2+ binding domain and a serine/threonine kinase domain into a single protein that performs the fundamental function of converting Ca2+ signals to phosphorylation events [76,77]. *A. thaliana* contains 34 CPK family genes that play a role in plant defense responses. CPK 3 and CPK 13 both participate in signaling after Ca2+ influx upon *S. littoralis* attack through regulation of plant defensin gene (PDF1.2) by phosphorylation of the transcription factor, HsfB2a [78]. The *cpk3* and *cpk13* mutants had much lower transcript levels of the plant defensin gene PDF1.2 in comparison to wild-type plants.

### *Tools Used to Monitor Ca2+ Signaling in Plant-Herbivore Interactions*

In recent years, the research on Ca2+ signaling has gained momentum with the advance in Ca2+ imaging techniques. Therefore, it is important to discuss different plant Ca2+ imaging methods, which are widely used in the context of plant-herbivore interactions to observe and record cytosolic Ca2+ concentration in herbivore-infested plants. These

techniques include the use of Ca2+ sensing fluorescent dyes and genetically encoded Ca2+ indicators. Various fluorescent Ca2+ sensing dyes, such as Fluo-3, Calcium Orange, etc., have been used to investigate the dynamics of cytosolic Ca2+ signals in plant- herbivore interaction [33,43,55,57,58,79–81]. For example, the Ca2+ indicator Ca2+ orange was utilized to identify changes in cytosolic Ca2+ concentrations in *P. lunatus* following *S. littoralis* herbivory. The changes in Ca2+ concentration were compared in response to a single wounding (MD) event, continual mechanical damage caused by a robotic worm (MecWorm, MW), and herbivory. After 30 min, a considerable increase in Ca2+ fluorescence was observed due to herbivory in the wounding zone, which persisted for 4 h, but in MD and MW plants, just a faint fluorescence was noticed [33]. Even though these dye-based markers have been demonstrated to be quite effective, these Ca2+ sensing dyes have some limitations, including toxicity, fragility, low fluorescence signals, and they cannot be imaged in living plants without permeabilization. To overcome these limitations, researchers have initiated research on the use of genetically encoded Ca2+ indicators. The most widely used Ca2+ imaging method includes genetically encoded Ca2+ indicators, such as GCaMP, Yellow Cameleon (YC) Ca2+-sensors. The Ca2+ sensors were developed from GFP by combining them with calmodulin. These Ca2+ sensors can be expressed in the whole plant and are functional throughout the entire plant. Therefore, it can be used to monitor cytosolic Ca2+ in plants subjected to various herbivore attack conditions [37,42,82–84]. For example, Toyota et al. [37] showed that the *P. rapae* caterpillars induced cytosolic Ca2+ responses in the leaves of *A. thaliana* can be monitored with GCaMP3. This study reported that the increases in cytosolic Ca2+ concentration were associated with ion influx through plasma membrane Ca2+ channels such as GLR3.3/GLR3.6. Another example is Verrillo et al. [83], who showed that Ca2+ induction could be monitored with YC3.60, a YC-based Ca2+ sensor, following application of *S. littoralis* OS on mechanically damaged *A. thaliana* leaves. By using these tools, it is now possible to study the dynamics of Ca2+ signaling in plant-herbivore interactions at single-leaf, whole-plant, and whole-plant-insect herbivore attack conditions.

Intracellular Ca2+ level is controlled by the influx of Ca2+ ions from extracellular through apoplastic and vacuolar membranes. Therefore, plant ion channels play an important role in regulating plant development and the perception of many stimuli, including herbivory.

### **5. Plant Ion Channels**

Ion channels are macromolecular pores in the membrane that regulate the influx and efflux of ions across the membrane at a rate of 106 ions per second. Ion channels can control ion fluxes in their target compartment and, thus, modify cellular homeostasis, and are vital in osmoregulation, development, signaling, mobility, and uptake of nutrients by the root and long-distance communication [85,86]. The first plant ion channel discovered, in 1984, is a K+ channel, Stelar K+ outward rectifier (SKOR) [87]. The last two to three decades have seen a dramatic increase in the number of ion channel subfamilies and their diverse functions. A large proportion of plant ion channel families have an analogous expression in animals. Ion channels are arranged into large families and are generally classified as cation, anion, or ligand-gated channels. Cation channels include voltagegated K<sup>+</sup> channels such as the shaker family (AKT1, AKT2, AKT6, KAT1, KAT2, KAT3, GORK, and SKOR; K<sup>+</sup> transport), tandem pore, and two-pore K<sup>+</sup> channels (TPK1, TPK4; K<sup>+</sup> transport and TPC1; Ca2+ and other cation transport), are responsible for permeation of K<sup>+</sup> ion across the plasma membrane and tonoplast membrane. Anion channels include slowly activating anion channels (SLAC1, SLAH1, SLAH2, SLAH3; Cl−/NO3 − transport), aluminum-activated malate transporters (ALMT1, ALMT6, ALMT9, ALMT12; Malate, Cl− transport), chloride channels/transporters (CLCc, CLCg, CLCe; Cl− transport), and detoxification efflux carrier (DTX33, DTX35; Cl− transport). Ligand-gated channels include cyclic nucleotide-gated channel (CNGC2, CNGC4, CNGC5, CNGC14, CNGC15, CNGC18, CNGC19, CNGC20; Ca2+/Ba2+ transport) and glutamate receptor-like channels (GLR3.1,

GLR3.3, GLR3.4, GLR3.5, GLR3.6; Ca2+ and other cations transport) [88] (Figure 2). These channels are responsible for setting up membrane potential, signal transduction, water, and solute transport [89], stomatal opening and closure [90,91], pollination [92], salt tolerance [93], and plant defense [94], to name a few. However, four distinct families of Ca2+-transporting ion channels have been shown to play a role in plant-herbivore interactions, including cyclic nucleotide-gated channels (CNGC19) [95,96], glutamate receptor-like channels (GLR3.3, GLR3.6) [37,42,97], two-pore channel 1 (TPC1) [59,84,98], and annexins (ANNEXIN 1) [99,100].

**Figure 2.** Phylogeny of plant ion channels. Representation of the phylogenetic tree of plant ion channels listed in Pantoja, 2020 [88], based on the analysis of protein homologs extracted from Uniprot.org. Progressive alignment and BLOSUM30 scoring method were used for multiple sequence alignment. The distance between the aligned sequences was calculated using Jukes-Cantor method. The phylogenetic tree was created by using the distance matrix. Unweighted pair group method average (UPGMA) was used to calculate group distance in the tree. Different colors represent different families of ion channels.

### *5.1. Cyclic Nucleotide Gated Channels (CNGC)*

The cyclic nucleotide-gated channels (CNGCs) are ligand-gated Ca2+ channels, first discovered in retinal photoreceptors and olfactory neurons [101]. They play a role in signal transduction in animals and are also present in other non-neuronal tissues [102]. These ion transport proteins have also been identified in plants [74,103,104] and have been known to be involved in a variety of biological processes, ranging from plant development and stress tolerance, disease resistance [105,106], thermal tolerance [107], and salt stress [108]. These channels are typically localized at the plasma membrane and in the model plant *A. thaliana,* which consists of 20 family members [109].

CNGC channel is composed of four subunits, and each of these subunits consists of six membrane-spanning regions and a pore domain [110]. There is a cyclic-nucleotide binding (CNB) and a calmodulin-binding domain (CaMB) present at the *C*-termini of the channel (Figure 3) [111]. In contrast, the animal system has a CaMB domain at the *N*-termini [112,113]. The plant and the animal CNGC differ in their pore amino acid sequence as well as the selectivity for various cations [105,114]. The amino acids that form the CaM binding domain overlap with the polypeptide region that includes the CNBD [115]. This overlapping affects the channel activation as the binding of CaM at the C termini hinders cyclic nucleotide-binding, suggesting variability in plant and animal CNGC channel regulation [116,117]. These channels are activated by the binding of cyclic nucleotides such as cAMP (cyclic adenosine monophosphate) and cGMP (cyclic guanosine monophosphate) [118–120], and inhibited by calmodulin binding [121]. These channels also show similarity with shaker-like K+ channels [105]. Patch-clamp recordings on plant cell protoplasts membrane directly show that CNGC activation can be achieved by the application of hyperpolarizing potentials (more negative than −120 MV), which allow Ca2+ entry into the cell [111,121].

**Figure 3.** Putative structure of CNGC19 channel. (Top) Schematic cartoon representation of CNGC19 channel subunit showing six membrane-spanning regions (S1–S6) and a large pore domain (S5–S6). Functionally relevant sites in the *C*-terminus consist of a CNB, cyclic nucleotide-binding domain which can bind cAMP/cGMP, and a CaMBD, calmodulin-binding domain which can bind calmodulin. The functional channel is formed by four subunits. (Bottom) The structure of CNGC19 has not been solved to date but is likely to show similarities with the animal CNG family of channels. Therefore, the structure shown in the figure is an approximation based on homology to other channels. The predicted CNGC19 secondary 3D structure model, showing four subunits in transparent surface view, was developed from the closest homolog PDB structure, 5VA1 (human ether-a-go-go related K<sup>+</sup> channel) using PHYRE 2.0 program. The image was prepared using Chimera software [122]. Created with BioRender.com (accessed on 30 August 2021).

It has been demonstrated that CNGC channels are important in modulating biotic stress responses such as Ca2+ influx in plant responses mediated by insect herbivore feeding [95]. A recent study by Meena et al. [96] has shown that the *A. thaliana* CNGC19 is responsible for generating and transmitting Ca2+ signals in local and systemic leaves mediated by the herbivore *S. litura*. A loss-of-function CNGC19 mutant in which the Ca2+ signals were attenuated was found to be more susceptible to attack by *S. litura*. In addition, jasmonic acid, a key signaling molecule in plant defense, was also observed in lower amounts in the CNGC19 mutant. These results suggest that CNGCs are involved in modulating plant resistance to insect herbivores, thus playing a role in the modulation of plant-herbivore interactions.

### *5.2. Glutamate Receptor-Like Channels*

Glutamate receptor-like (GLR) is a non-selective ion channel responsible for permeating Ca2+ ions across the plasma membrane of animals and plants. Plant glutamate receptor-like (GLR) channels are ionotropic glutamate receptor homologs in mammals (iGluRs). The iGluRs have been extensively studied for their central nervous system and have been known to play a vital role in synaptic transmission, learning, and memory [123,124]. It is intriguing that GLRs also exist in plants despite the absence of the central nervous system [125]. In plants, GLRs play a crucial role in carbon and nitrogen metabolism [126], gravitropism [127], pollen tube growth [128,129], immune defense reactions [38,130–133], and wound-induced intracellular signaling [97]. *Arabidopsis* consists of 20 GLR genes; each subunit hosts a *N*-terminal domain, two extracellular ligand-binding sites (L1, L2), and transmembrane domains (S1–S4), including a pore region (P) and the *C*-terminal domain [134] (Figure 4). In mammals, iGluRs are divided into three groups according to their sequence diversity and ligand specificities [124]. These include N-methyl-D-aspartate (NMDA), α-amino-3-hydroxy-5-methyl-4-isoxazole propionic acid (AMPA), and Kainate receptors. Plant GluRs share a high degree of similarity with the NMDA receptors that range from 16 to 63% in the ligand-binding domains and the transmembrane domains [135]. These channels are not only present at the plasma membrane but can also be found in chloroplasts, mitochondria [136], and vacuolar membranes [129]. Unlike their mammalian counterparts, the plant GLRs have much broader ligand selectivity. The major difference in plant and animal iGLR is the pore region. These non-selective cation channels are activated by amino acid glutamate, which acts as a metabolite, energy source, and neurotransmitter in animals [137,138].

Electrophysiological studies have shown the involvement of GLRs in inducing a Ca2+ influx in plants that leads to the modulation of plant defense signaling to insect herbivores [139,140]. A study by Vasta et al. [140] showed that the application of GLR agonists such as glutamate induced a strong and rapid cytosolic Ca2+ increase in tobacco (*Nicotiana tabacum)* var *xanthi* while the application of lanthanum and Ca2+ chelator, BAPTA, inhibited glutamate-induced Ca2+ increase. This observation suggests that the plant GLR has a role in the modulation of Ca2+ influx that ensures plant defense responses against insect herbivores.

GLR3.3 has been implicated in the transmission of signals in the form of Ca2+ waves from wounded to unwounded sections of the plant. When *S. littoralis* larvae were allowed to feed on *A. thaliana* wild-type plants, wound-induced surface potential alterations were detected. However, wounding reduced the surface potential alterations in the four GLR mutants GLR3.1, GLR3.2, GLR3.3, and GLR 3.6. [97]. This suggests that GLR3.3 plays an important role in the modulation of plant defense signaling to insect herbivores. Recently, Toyota et al. [37] showed that GLRs are activated by wounding and upon herbivory by cabbage butterfly (*Pieris rapae*) caterpillars in *A. thaliana* leaf expressing genetically encoded Ca2+ sensor GCaMP3. The cytosolic Ca2+ elevation and subsequent defense gene expression were observed after the application of glutamate and not with other amino acids such as sorbitol. Furthermore, the Ca2+ signals were completely abolished in the GLR3.3/GLR3.6 double mutant in *A. thaliana,* suggesting that GLR3.3 and GLR3.6 are

essential for transmitting Ca2+ signals induced by wounding and herbivory. Another recent study by Shao et al. [42] demonstrated that wounding of the main root at a distance of 2 cm from the root-shoot junction increased the Ca2+ elevation and surface wave potential (SWP) in *A. thaliana* expressing calcium sensor GCaMP6. Additionally, the application of glutamate to the wound site in the root induced an increase in Ca2+ and SWP in all leaves. Interestingly, in the GLR3.3/GLR3.6 double mutant, this wound and glutamate-induced rise in root to shoot Ca2+ was attenuated. This finding suggests that GLR3.3 and GLR3.6 are involved in propagating systemic Ca2+ signaling from leaf to leaf and root to shoot. These results provide evidence for the role of plant GLRs in the modulation of Ca2+ signaling during plant defense responses against insect herbivores.

**Figure 4.** Putative structure of GLR3.3/3.6 channel. (Top) Schematic cartoon representation of GLR3.3/3.6 channel subunit showing extracellular *N*-terminus, four membrane-spanning regions (S1–S4), 2 extracellular ligand-binding sites (L1, L2), and intracellular *C*-terminus. (Bottom) The structure of GLR3.3/3.6 has not been solved to date but is likely to show similarities with the animal NMDA receptor family of channels. Therefore, the structure shown in the figure is an approximation based on homology to other channels. The predicted GLR3.3/3.6 secondary 3D structure model showing four subunits in transparent surface view was developed from closest homolog PDB structure 4TLL (*Xenopus laevis* GluN1/GluN2B NMDA receptor), using PHYRE 2.0 program. The image was prepared using PyMol software (PyMOL Molecular Graphics System, Version 2.4, Schrödinger, LLC, New York, NY, USA). Created with BioRender.com (accessed on 30 August 2021).

### *5.3. ANNEXIN1*

Annexins are the phospholipid-binding proteins and are considered novel mechanosensitive Ca2+ channels [141,142]. In animal cells, annexins are present in the cytoplasm and cellular membranes [143]. They are involved in vital cellular processes such as membrane trafficking, ion flux, mitotic signaling, and cytoskeleton rearrangement [143,144]. Eight annexin genes have been identified in *A. thaliana* by genome sequencing [145]. Plant annexins are structurally different from their animal homologs but have a conserved primary

sequence. These 32–42 kDa proteins have two major domains: a *N*-terminal head and a *C*-terminal annexin core [143] (Figure 5). The annexin core is composed of four annexin domains (I–IV), each of which is 70 amino acids in length and contains five short helices and a conserved endonexin fold (G-X-G-T-{38-40}-D/E). Ca2+ binding activity occurs in type II and III binding sites of annexin proteins [141,143]. Plant annexins have a shorter *N*-terminal region than their animal counterparts [146] and are crucial for actin binding, inhibition of callose synthase, and oxidative stress responses [147–150]. The functional diversity of annexins is due to the variable *N*-terminal region that interacts with other proteins.

**Figure 5.** Putative structure of ANNEXIN1 channel. ANNEXIN1 secondary 3D structure model showing two subunits (homodimer) in transparent surface view was developed from PDB structure 1YCN (*Arabidopsis thaliana* ANNEXIN). The presence of Ca2+ or H2O2 appears to be required for homodimerization. The image was prepared using Chimera software [122].

A recent study by Malabarba et al. [100] reported the role of ANNEXIN1 (ANN-1) in initiating systemic defense in *A. thaliana* in response to Egyptian cotton leafworm (*S. littoralis*) herbivory. The study found that annexin 1 was responsible for inducing cytosolic free Ca2+ elevation upon wounding and simulated herbivory in *A. thaliana*. ANN-1 knock-out and ANN-1 overexpressing lines were employed in this work to evaluate their role in herbivory-mediated Ca2+ signaling. The result showed that in the ANN-1 deletion line, the increase in cytosolic Ca2+ upon herbivory by *S. littoralis* was impaired, and the larvae gained more weight than those fed on wild-type plants. On the other hand, weight increase was significantly lower in larvae that fed on the ANN-1 overexpressed line compared to the wild type. Additionally, jasmonate accumulation and defense responses were diminished in ANN-1 systemic leaves, demonstrating that ANN-1 is involved in systemic cytosolic Ca2+-dependent jasmonate induction. This finding suggests that ANN-1 modulates plant defenses against herbivore damage through the Ca2+-dependent jasmonate signaling pathway and is required for systemic rather than local defense activation in plants attacked by herbivorous insects.

### *5.4. Two Pore Channel 1 (TPC1)*

Two pore channels (TPCs) are organellar cation channels that are widely expressed in animals and plants. In animals, they are localized in the endolysosomal membrane, while in plants they reside in the tonoplast of plant vacuoles [151–154]. They are members of the voltage-gated ion channel superfamily. The vacuolar TPC1 channel, also known as the slowly activating vacuole (SV) channel, has been implicated in a variety of processes in plants, including nutrient sensing, pH homeostasis, and modulation of the membrane potential. The first plant TPC1 gene was cloned in *A. thaliana* (AtTPC1), with 733 amino acids identical to the rat TPC1 sequence [152].

Plant and animal TPCs are similar in sequence to voltage-gated Ca2+ and Na+ channels and feature two shaker-like units with six transmembrane domains (S1–S6), each joined

by a cytosolic linker containing two Ca2+-binding EF-hands (EF1 and EF2). (Figure 6). Voltage and an increase in the cytosolic Ca2+ level both influence the activity of plant TPCs. Ca2+ binding to the cytosolic EF-hand domain induces conformational changes in the pair of pore-lining inner helices from the first 6-TM domains, whereas membrane potential activates the second voltage-sensing domain, which undergoes conformational changes and facilitates pore opening [155]. The SV channel transports Ca2+ in addition to Na+ and K<sup>+</sup> and has a permeability ratio of 3:1 for Ca2+ to K<sup>+</sup> [156,157]. Ca2+ release is substantially dependent on the concentration of cytosolic free Ca2+, indicating that this channel is involved in Ca2+-induced Ca2+ release [156,158]. The plant TPC1 has been implicated in insect-plant interactions. A study by Kiep et al. [98] has shown that an increase in local cytosolic Ca2+ and systemic Ca2+ response was induced in response to *S. littoralis* feeding on *A. thaliana*. By using real-time imaging in *A. thaliana* expressing the Ca2+ reporter aequorin to monitor the induction of local and systemic cytosolic Ca2+ signals, this study showed that simulated herbivory by wounding inhibited the systemic Ca2+ signal in the *tpc1* knockout mutant. These results indicated that the TPC1 channel plays a key role in the systemic [Ca2+] cyt signal induced by insect herbivory in *A. thaliana*. Another study by Vincent et al. [84] employed *A. thaliana* plants expressing the GFP-based Ca2+ indicator GCaMP3 to visualize Ca2+ accumulation in response to aphid *M. persicae* feeding. Within 95 s of the aphids settling, a robust fluorescence burst was seen, indicating cytosolic Ca2+ elvation. The rise in cytosolic Ca2+ was strongly dependent on Brassinosteroid Insensitive Associated Kinase I (BAK1), the plasma membrane Ca2+ permeable ion channels glutamate receptor-like 3.3 and 3.6 (GLR3.3 and GLR3.6), which are critical regulators of extracellular Ca2+ import into the cytoplasm of plant cells. In addition, this study also revealed that the increase in cytosolic Ca2+ induced TPC1 mediated vacuolar Ca2+ release in response to aphid feeding, suggesting that the TPC1 channel operates in conjunction with the plasma membrane Ca2+ permeable ion channels GLR3.3 and GLR3.6 in mediating cytosolic Ca2+ increase during insect herbivory [84].

**Figure 6.** Putative structure of TPC1 channel. (Top) Schematic cartoon representation of individual plant TPC1 channel subunit comprising two repeated domains showing six membrane-spanning regions (S1–S6), two pore loops (P), and joined via a cytosolic linker containing two Ca2+ binding EF-hands (EF1 and EF2). (Bottom) TPC1 secondary 3D structure model showing two subunits in transparent surface view was developed from PDB structure 5DQQ (*Arabidopsis thaliana* TPC1). The image was prepared using Chimera software [122]. Created with BioRender.com (accessed on 30 August 2021).

### *5.5. H+-ATPase*

The proton-pumping ATPases (H+-ATPases) are the primary pumps responsible for the generation of a proton gradient across cellular membranes. This electrogenic transporter uses energy from ATP hydrolysis to drive the translocation of protons against their concentration gradient from the cytosol to the external aqueous environment [159]. The H+-ATPase is located in the plasma membrane (PM) of plant cells. It has been demonstrated that the activation and suppression of the H+-ATPase activity in the plant plasma membrane modulate Vm, resulting in the alteration of PM ion channels and transporters functions [160]. The PM H+-ATPase is a single 100 kDa polypeptide and a member of the large family of phosphorylation (P)-type ATPases. It is composed of six transmembrane helices (M1–M6) and a cytoplasmic domain containing phosphorylation (P), nucleotidebinding (N), and actuator (A) domains involved in ATP hydrolysis. The PM H+-ATPase has been implicated in various physiological processes, including cell development, intracellular pH regulation, food uptake, stomatal opening, salt tolerance, and cellular expansion [161–165].

Plant PM H+-ATPase has been shown to contribute in the propagation of the intracellular defense signaling cascade by modifying Vm in response to herbivore feeding [166]. A study by Camoni et al. [167] demonstrated that *S. littoralis* oral secretions effectively inhibited *Phaseolus lunatus* PM H+-ATPase, resulting in decreased H+ extrusion from the cytosol and modification of the Vm. This observation implied that H+ extrusion by the plant H+-ATPase was involved in Vm regulation and might initiate a plant defensive response to herbivory. Another recent study by Kumari et al. [168] has revealed that *Arabidopsis* H+-ATPase 1 (AHA1) is involved in the formation of slow wave potentials (SWPs), which are required for long-distance electrical transmission during herbivoreinduced plant defense. Fusicoccin, a PM H+-ATPase activator, prolonged the SWP repolarization phase in leaves distal to wounds. The repolarization phase was significantly prolonged in reduced function *aha1* mutants, whereas the duration of SWP repolarization was reduced in the presence of a gain-of-function mutant *ost2-2D*. Additionally, *S littoralis* larvae performed better on *aha1-7* mutants than on wild-type plants. Overall, these observations suggest that the PM H+-ATPase is required for the regulation of the Vm and electrical signal propagation between different parts of a plant during insect herbivory.

### **6. Reactive Oxygen Species (ROS)**

Reactive oxygen species (ROS) are highly reactive molecules generated by redox reactions. They are part of several biological processes, such as photorespiration, oxidative phosphorylation, the electron transport chain (ETC), as well as a plant defense against pathogens and herbivores. ROS is produced in the mitochondria, chloroplast, and peroxisomes. There are several forms of ROS like superoxide anion (O2−•−), hydrogen peroxide (H2O2 •), hydroxyl radical (HO•), peroxynitrite (ONOO), and singlet oxygen (1O2) [169]. ROS is typically produced by the nicotinamide adenine dinucleotide phosphate (NADPH) oxidase complex, which catalyzes the reduction of molecular oxygen to superoxide anion, which is then converted to H2O2. In plants, respiratory burst oxidase homologs (RBOHs) were found to be the key enzymes that catalyze the formation of ROS, which is a key step in plant protection against herbivores [170–172]. The respiratory burst oxidase homolog D (RBOHD) has been found to be essential for the propagation of ROS waves [173]. The significance of RBOHs in organizing responses against chewing insect herbivores was verified in *N. attenuate* where tobacco hornworm (*Manduca sexta*) OS enhanced NaRBOHD (*N. attenuata* NADPH oxidase homolog) on damaged leaves. ROS accumulation was diminished in *M. sexta* OS treated NaRBOHD-silenced *N. attenuata* plants without affecting OS-induced gene expression of defense-related genes [174].

The production of ROS is an inevitable by-product of metabolism in many cell types. Previously, it was assumed that ROS are toxic molecules that cause cellular damage to macromolecules [175]. Still, the role of ROS in plant defense has only recently emerged. It is well established that ROS can act as early defense signaling molecules that promote

plant defense responses against a variety of pathogens and herbivores [54,176]. ROS act as secondary messengers that can penetrate up to 8.4 cm/min in *A. thaliana* [177]. Plants use ROS to alert the non-injured tissue about a plant attack by either releasing small quantities, which activates certain defense responses or prevent cell death by limiting the production of ROS [178]. ROS production has also been suggested to be involved in plant-microbe interactions as ROS can activate or repress the expression of defense-related genes [179,180]. The role of ROS in plant resistance to herbivores has been demonstrated in resistant and near-isogenic susceptible wheat after the attack of Russian wheat aphid (*Diuraphis noxia*). A strong burst of H2O2, as well as NADPH oxidase activity, was observed in resistant plants 3 h after infestation in comparison to susceptible plants. Treatments of plants with diphenyleneiodonium (DPI), an inhibitor of NADPH oxidase, suppressed the H2O2 production. Elevation in H2O2 levels (47%) was observed by treating resistant wheat plants with a mixture of glucose and glucose oxidase [181], suggesting that H2O2 plays a role in the defense response against *D. noxia* infestation.

Studies have shown that ROS serve as early defense signaling molecules in response to herbivore-induced wounding and secretions such as OS and oviposition. Imbiscuso et al. [182] investigated the effect of brake fern (*Pteris vittata*) response to herbivory by *S. littoralis*. The *P. vittata* plants responded to the attack of *S. littoralis* by activating peroxidases which produced H2O2. The concentration of H2O2 in leaves was lower in mechanically wounded young leaves than herbivory wounded leaves, suggesting that *P. vittata* can distinguish between mechanical and herbivory wounding by modulating the amount of ROS production. A study by Shinya et al. [183] demonstrated that the application of OS isolated from generalist herbivore, nightfeeding rice armyworm, (*Mythimna loreyi*)*,* caused a strong intracellular ROS generation on rice cells, and a similar effect was obtained upon application of synthetically prepared N-linolenoyl-L-Glu, the most abundant FAC present in OS of *M. loreyi,* indicating that FAC from *M. loreyi* OS promoted ROS production in rice cells.

Recently, our group Gandhi et al. [184] demonstrated that *M. sexta* oral secretions (OS) induced ROS generation in isolated tomato protoplasts. Interestingly, our study showed that the application of tomato plant-fed (PF) *M. sexta* OS enhanced ROS generation while artificial diet-fed (DF) OS could not induce ROS in tomato protoplasts, suggesting that the oral secretions of *M. sexta* play an indispensable role in inducing ROS generation in tomato protoplasts. Our study also showed that the *M. sexta* PF-OS induced ROS increase was diminished in the presence of a Ca2+ chelator, BAPTA-AM, suggesting that there is a link between Ca2+ and ROS signaling. Several lines of evidence have indicated the existence of a positive feedback mechanism between ROS and Ca2+ production. In a heterologous expression system, treatment with ionomycin, an ionophore that leads to Ca2+ influx into cells, resulting in activation of RHD2 NADPH oxidase (root hair defective 2 reduced nicotinamide adenine dinucleotide phosphate) in root tips of *A. thaliana* confirming Ca2+ triggered RHD2 NADPH oxidase activity. These observations suggest that Ca2+ acts upstream of ROS production [185].

Compelling evidence indicates that ROS production by RBOHD is dependent on the Ca2+ binding [186,187]. RBOHD carries 2 EF-hands which are known to participate in Ca2+ dependent modulation [188]. Abscisic acid (ABA) signaling in guard cells involves both Ca2+ and ROS. *A. thaliana* mutants lacking certain NADPH oxidases (AtRBOHD and AtRBOHF) do not close their stomata and produce ROS, Ca2+, and Ca2+ channel activation when they are exposed to ABA. Supplementation of H2O2 to guard cells rescues the mutant phenotype, implying that Ca2+ entry proceeds downstream of ROS generation in ABA signaling [189,190].

In *A. thaliana*, the production of H2O2 was observed in leaves 72 h after oviposition by cabbage moth (*Pieris brassicae*) and was recognized by the formation of a reddish-brown precipitate. This result indicates that oviposition can trigger a localized response that resembles the hypersensitive response induced by pathogens [191]. A recent study by Stahl et al. [192] showed that eggs of *P. brassicae* induced generation of H2O2, salicylic acid and defense gene expression in *A. thaliana*. This study also revealed phosphatidylcholines (PCs)

released from eggs is the key signaling molecule that activates gene expression and triggers various defenses in the plants.

### *Tools Used to Monitor ROS Signaling in Plant-Herbivore Interactions*

While ROS relevance in plant-herbivore interaction is gaining momentum, the detection and characterization of ROS are still a significant bottleneck in this field. The early detection and quantification of ROS can be carried out by either utilizing genetically encoded fluorescent ROS sensors such as redox-sensitive green fluorescence protein (Ro-GFP), or synthetic fluorescent probes, such as 3,3 -diaminobenzidine (DAB) and 2 ,7 dichlorofluorescein diacetate (H2DCFDA). Genetically encoded ROS sensors "Ro-GFP" can monitor the cellular redox status at a high spatiotemporal resolution [193–199]. A recent study by Hipsch et al. [200] measured the whole plant ROS generation in response to high light, cold, and drought by using a chloroplast-targeted redox-sensitive green fluorescence protein 2 (RoGFP2). This finding suggests that whole-plant redox imaging using genetically encoded ROS sensors can be applied in a wide range of abiotic and biotic stress conditions, including plant-herbivore interaction. Despite the promising findings, the application of genetically encoded ROS sensors in plant-herbivore interactions is still limited due to the laborious and time-consuming method of its application. In contrast, synthetic fluorescent probes such as DAB and H2DCFDA are easier to use and can measure ROS in real-time with high sensitivity [201]. DAB has been used in many studies on plants as a reliable biomarker for reactive oxygen species (ROS) production [202–204]. However, in recent years, H2DCFDA has gained attention for its potential to measure the ROS levels in real-time on whole plants and as well as plant protoplasts [184,205,206]. Fichman et al. [205] measured the effect of light stress, injury, and pathogen, *P. syringae* pv. tomato DC 3000 on ROS signaling in H2DCFDA dye sprayed *A. thaliana* by using whole plant-live imaging. This study suggests that the combination of live-cell imaging and the use of H2DCFDA enables real-time monitoring of ROS in plants in response to various stress and pathogen treatments. This study also utilized an RBOHD (*rbohD*) knockout, and upon treatment with different stimuli, less ROS generation was observed. In contrast, another cytosolic ascorbate peroxidase 1 (*apx*) knockout produced more local as well as systemic ROS upon wounding or light stress treatments implying that this mutant had less ROS quenching capacity.

### **7. Conclusions**

Recent years have witnessed immense progress in identifying the early defense signaling components in plant defense against herbivores, but studies on the molecular identification and characterization of these components are still a work in progress. However, with the advent of state-of-the-art imaging techniques, physiological and biochemical assays, and genomics may help us to understand the early defense signaling events by coordinating the plasma membrane potential changes, ion channels modulation, intracellular Ca2+ and ROS generation, gene expression, and, ultimately, the host plant defense response against herbivores. Transforming plants with biosensors such as GCaMP-Ca2+ and Ro-GFP-ROS sensors can help in the early identification of the plant defense responses. HAEs such as OS, frass, and oviposition could be used to develop strategies for early detection of the impending herbivory. So far, only a handful of Ca2+ permeable channels have been identified that plays a role in plant-herbivore interactions. Further studies are needed to unravel other ion channels that may be contributing to the modulation of Vm, Ca2+, and ROS, the downstream signaling cascade, and, more importantly, the role of these ion channels in triggering a rapid defense response. A deeper understanding of these early signaling events will eventually help us to minimize herbivory by developing pest management strategies based on plant-herbivore monitoring systems. Such knowledge can be instrumental in the design of plants with improved resistance against herbivores. As such, in the future, it will be important to develop effective small-molecule modulators that can inhibit or enhance the early signaling events in plant-herbivore interactions. Such

an approach would not only facilitate research on early plant signaling events but also help in developing novel strategies for the development of herbivore-resistant crops.

**Author Contributions:** N.S. conceptualized and wrote the manuscript with A.G.; N.S. designed the final figures and edited the final manuscript draft with A.G. and R.K.; A.G., A.H., M.A., A.B. and N.S. contributed to the preparation of figures. A.G., R.K. and N.S. proofread and revised the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was funded by the College of Sciences, University of Rio Grande Valley startup fund and the University of Texas System Rising STARs Award to N.S. and College of Sciences Seed grant to R.K.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Not applicable.

**Conflicts of Interest:** The authors declare no conflict of interest.

### **References**


**Abdul Hameed <sup>1</sup> , Muhammad Zaheer Ahmed <sup>1</sup> , Tabassum Hussain <sup>1</sup> , Irfan Aziz 1, Niaz Ahmad 2,3 , Bilquees Gul <sup>1</sup> and Brent L. Nielsen 4,\***


**Abstract:** Salinity is a growing problem affecting soils and agriculture in many parts of the world. The presence of salt in plant cells disrupts many basic metabolic processes, contributing to severe negative effects on plant development and growth. This review focuses on the effects of salinity on chloroplasts, including the structures and function of these organelles. Chloroplasts house various important biochemical reactions, including photosynthesis, most of which are considered essential for plant survival. Salinity can affect these reactions in a number of ways, for example, by changing the chloroplast size, number, lamellar organization, lipid and starch accumulation, and interfering with cross-membrane transportation. Research has shown that maintenance of the normal chloroplast physiology is necessary for the survival of the entire plant. Many plant species have evolved different mechanisms to withstand the harmful effects of salt-induced toxicity on their chloroplasts and its machinery. The differences depend on the plant species and growth stage and can be quite different between salt-sensitive (glycophyte) and salt-tolerant (halophyte) plants. Salt stress tolerance is a complex trait, and many aspects of salt tolerance in plants are not entirely clear yet. In this review, we discuss the different mechanisms of salt stress tolerance in plants with a special focus on chloroplast structure and its functions, including the underlying differences between glycophytes and halophytes.

**Keywords:** salinity stress; photosynthesis; chloroplast; plastid; osmolytes; osmotic adjustment

### **1. Introduction**

Soil quality in many parts of the U.S. and worldwide is susceptible to a variety of stresses, including drought, temperature, deterioration due to erosion and other factors, and increasing salinity due to evaporation and/or irrigation practices. At the same time the human population is growing and in many regions high-quality agricultural land is decreasing due to the expansion of urban areas [1].

Salinity is inhibitory to the growth and development of many plants, including most crops [2–5]. It affects all cellular processes, including disruption of cellular homeostasis, impairment of photosynthesis, mRNA processing, transcription, protein synthesis, disruption of energy metabolisms, amino acid biosynthesis as well as lipid metabolism [6–10]. In response to increasing salt, plant cells activate specific Na+ and Cl<sup>−</sup> ion transporters and adjust the accumulation of cytosolic K+ [10–12]. Plant cells must also undergo osmotic adjustment, which is accomplished in many ways, including the production of organic osmolytes such as glycine betaine, proline, some sugars, and polyamines, of which most are synthesized in the chloroplast [3,10].

**Citation:** Hameed, A.; Ahmed, M.Z.; Hussain, T.; Aziz, I.; Ahmad, N.; Gul, B.; Nielsen, B.L. Effects of Salinity Stress on Chloroplast Structure and Function. *Cells* **2021**, *10*, 2023. https:// doi.org/10.3390/cells10082023

Academic Editor: Suleyman Allakhverdiev

Received: 30 July 2021 Accepted: 5 August 2021 Published: 7 August 2021

**Publisher's Note:** MDPI stays neutral with regard to jurisdictional claims in published maps and institutional affiliations.

**Copyright:** © 2021 by the authors. Licensee MDPI, Basel, Switzerland. This article is an open access article distributed under the terms and conditions of the Creative Commons Attribution (CC BY) license (https:// creativecommons.org/licenses/by/ 4.0/).

Chloroplasts belong to a family of cellular organelles commonly found in plant and algal cells known as plastids. Green plastids—chloroplasts—are the site where atmospheric CO2 fixation occurs through a series of biochemical reactions called the Calvin–Benson cycle by utilizing the energy produced by the light reactions of photosynthesis [13]. Elevated salinity levels affect many cellular processes, including photosynthesis, the major function of chloroplasts. The presence of salt in the soil may cause both osmotic and ionic stresses [14], which may hinder photosynthesis through the diffusional (stomatal, mesophyll and boundary layer resistance to CO2) and/or non-diffusional (photochemical and biochemical) limitations of carbon fixation [6,15–20]. Salinity exposure is also known to decrease the chlorophyll content in many plants [21,22]. However, salt-resistant plants, particularly those with a C4 mechanism, may overcome the inhibitory effect of salinity on CO2 fixation more effectively [6,23].

In general, when plants are exposed to salt stress, the very first response is osmotic shock followed by induction of stomatal closure. Stomatal closure, in turn, limits photosynthetic capacity by the restriction of CO2 supply. However, research has shown that increasing the external CO2 concentration under salt stress did not lead to an increase in photosynthesis rates in many cases. This observation suggests the involvement of some non-stomatal components in photosynthesis reduction under salinity, such as overproduction of reactive oxygen species (ROS) and the depletion of K<sup>+</sup> inside plant cells due to the accumulation of Na+ [24,25]. This results in the disruption of ionic homeostasis in chloroplasts.

Besides CO2 fixation, thylakoid reactions are also affected by salinity [6,18,26]. The most commonly studied parameters in this context are the maximum quantum efficiency of the PSII reaction centers (*Fv/Fm*), quantum efficiency of PSII (ΦPSII), non-photochemical quenching (NPQ), photochemical quenching (qP) and electron transport rate (ETR), which defines the overall performance of plants under different stresses [27]. Salt-resistant plants are known to possess resilient thylakoid reactions to overcome salinity effects such as photodamage [28] and protection of the reaction centers [29]. This may include protective mechanisms such as cyclic electron flow, photorespiration in C3 plants and regulation of NPQ [18,30]. CO2 fixation and thylakoid reactions of photosynthesis take place in thylakoids and the stroma of the chloroplast, providing the essential carbon skeleton for growth, energy for driving various metabolic reactions as well as the biosynthesis of different metabolites. Salt-induced toxicity negatively affects all these processes, resulting in poor plant growth and reduction in yield. Chloroplasts are also major reactive oxygen species (ROS) production sites at the reaction centers of PSII and PSI, where charge separation occurs, and the electron transport chain (ETC) from PSII to PSI are highly sensitive to salt-induced toxicity under which ROS production is further increased [31]. Higher concentrations of ROS cause oxidative damage to membranes, lipids, nucleic acids, proteins and some photosynthetic enzymes, resulting in reduced CO2 fixation, slower plant growth and consequently low crop yields. The ROS-scavenging system includes both enzymatic and non-enzymatic antioxidants that prevent oxidative damage. Therefore, manipulation of the components of this system holds great implications for improving the photosynthetic rates under salt stress in crop plants. This has been tested by overexpression of Cu/Zn superoxide dismutase (SOD) in the chloroplasts of tobacco [32,33] and Chinese cabbage [34]. Since chloroplasts are largely under the control of nuclear gene expression for growth and metabolic activities, chloroplasts have evolved a sophisticated signaling network to coordinate with the nucleus to control gene expression and maintain the balanced expression of genes in the two compartments. Chloroplasts also act as global sensors relaying changes in their own developmental status as well as in the environmental conditions, including light intensity and stresses to the nucleus. As a result, the nucleus adjusts the expression of its genes to ensure optimal plant performance under changing environmental conditions [35]. Until recently, this chloroplast–nucleus communication has been largely viewed as bilateral, ignoring the pivotal role of chloroplasts in adjusting gene expression and metabolic processes that affect photosynthesis and ultimately crop yields.

In this review, we discuss the effect of salt stress on chloroplasts, their structures, and various biochemical reactions occurring in them. We also compare the differences in how chloroplasts of glycophytes and halophytes respond to salinity stress.

### **2. Effects of Salinity on Chloroplast Ultrastructure**

### *2.1. Changes in Chloroplast Structure in Plants*

Chloroplasts are roughly 1–2 μm thick and 5–7 μm in diameter. They are enclosed in a chloroplast envelope, which consists of a double membrane with outer and inner layers; the space in between is called the intermembrane space. A third, internal membrane, extensively folded and characterized by the presence of closed disks (or thylakoids), is known as the thylakoid membrane. In higher plants, the thylakoids are arranged in tight stacks called grana. Grana are connected by stromal lamellae extended from one granum through the stroma into a neighboring granum. The thylakoid membrane envelops a central aqueous region known as the thylakoid lumen. The space between the inner membrane and the thylakoid membrane is filled with stroma, a matrix containing dissolved enzymes, starch granules and copies of the chloroplast genome [36].

Several changes have been associated with chloroplast structure in response to environmental factors and the availability of water and minerals [37]. These include modifications in the lamellar organization, resulting in chloroplast shrinkage [37], swelling of chloroplast lamellae and an unrecognizable grana structure under highly saline conditions [38]. Some plants, such as *Atriplex* spp., may undergo lipid deposition to counter the harmful effects of salt-induced toxicity [39,40]. In some instances, starch accumulation under high salinity has also been reported, such as in chloroplasts of wheat cultivars, which was related to damage to the sucrose-phosphate synthase in the cytosol, triggering the triose-phosphate pathway towards starch synthesis [41]. Changes in the ionic composition of starch-degrading enzymes may also be linked with excessive starch deposition [42]. Under saline conditions, reactions involving starch and sucrose biosynthesis are also known to be regulated by changes in the orthophosphate concentration [43,44]. Stress-induced destruction of the chloroplast envelope and an increase in the numbers of plastoglobuli in thylakoid membranes have also been reported in cucumber leaves [45]. Accumulation of starch grains in the chloroplasts of *Thellungiella* and tobacco plants is known to play an important role as osmotica in maintaining the structural integrity of the chloroplasts [46].

### *2.2. Changes in Ultrastructure of Chloroplasts in Glycophytes and Halophytes*

Salt stress-induced alterations in the structure of chloroplasts or thylakoid membranes have been extensively examined in various salt-sensitive plants [47,48] as well as in facultative halophytes [49]. Swelling of thylakoids under salt stress (~200 mM NaCl) was reported in rice [50]. However, recent 3D analysis confirmed that rice chloroplasts became spherical under salt stress without any changes in the overall chloroplast volume [51]. Contrasting observations regarding chloroplast volume have been reported among different species. For example, chloroplasts of salt-sensitive cultivars of wheat exhibited an increase in volume possibly due to changes in the ionic composition of the stroma [42]. Spinach chloroplasts showed a decrease in volume with concomitant changes in light-scattering during electron transport [47]. Arabidopsis seedlings grown in the presence of salt also exhibited swollen chloroplasts with less developed granum structures [41]. Changes in the thylakoid ultrastructure of potato [52] and maize [53] under salinity have been previously related to perturbed ion homeostasis in chloroplasts.

In the case of halophytes, salt entry into the chloroplast stroma may be critical for grana formation and photosystem II activity, as halophytes have been reported to accumulate more chloride (Cl<sup>−</sup>) than glycophytes and use sodium (Na+) in different functions [54]. Chloroplast swelling of *Atriplex* leaf cells at 345 mM NaCl appeared to be a likely result of the osmotic effect of salinity while few changes were reported in the chloroplasts of hair cells [40]. Similarly, distinct thylakoid swelling in *Thellungiella* under saline conditions (400 mM NaCl) was attributed to the disturbance in osmotic equilibrium [54]. Other

notable changes in halophyte chloroplasts include the formation of 'slim spindle-shaped' grana with a clear stromal matrix in the halophyte *Kandelia candel* at 400 mM and increases in the plastoglobuli numbers at 600 mM NaCl with intact grana thylakoids [49]. In *Bruguiera parviflora*, no major alterations in the structural integrity or absorption characteristics of the thylakoid membranes were noted at 400 mM NaCl; however, a reduction in antenna size, as well as cytochrome (Cyt) *f* and Cyt *b*<sup>6</sup> were observed [55].

### *2.3. Changes in the Chloroplast Ultrastructure of C4 Plants*

Plants with C4 photosynthesis possess chloroplasts both in the bundle sheath cell (BSC) and the mesophyll cell (MC). Both of these chloroplast types, BSC and MC, have been reported to respond differently to salt stress. C4 plants are further divided into three subtypes, namely, NADP-malic enzyme (NADP-ME), NAD-malic enzyme (NAD-ME) and phosphoenolpyruvate carboxykinase (PCK) types, with peculiar leaf anatomical characteristics [56]. The NADP-ME type 4 species showed little damage to BSC chloroplasts compared to MC chloroplasts [57]. The BSC chloroplasts showed the development of grana when exposed to salt stress [53,58]. On the other hand, thylakoid swelling and disruption of envelopes in MC chloroplasts of both the NAD-ME and PCK types were observed under saline conditions [57]. It has also been reported that exposure to salinity enhanced granal development in BSC chloroplasts and appressed thylakoid density [57]. However, granal development in the NAD-ME and PCK type species is not as pronounced as in the NADP-ME type species. It is believed that granal development in BSC chloroplasts could compensate for the loss of PSII activity in MC chloroplasts under salt stress [53]. In glycophyte plants of the NADP-ME and NAD-ME subtypes, salt stress causes grana unstacking in MC chloroplasts but induces grana formation in BSC chloroplasts. Interestingly, in halophytes of the same subtypes, the grana are constitutively present in BSC chloroplasts and the unstacking of grana is absent in MC chloroplasts [53,58].

### *2.4. Effects of Salinity on Chloroplast Multiplication*

Although the number of chloroplasts per leaf cell varies from a few to hundreds [59], they may occupy more than half of the cytoplasm volume in mesophyll cells [60]. Salinity may increase the number of chloroplasts per cell, e.g., in spinach, sugar beet [61], wheat [62], *Arabidopsis* [63] and *Thellungiella halophila* [46]. Bose et al. [29] proposed that halophytes have a greater capacity to increase chloroplast number than glycophytes under salinity, which may help in storing Na+ and Cl<sup>−</sup> without compromising photosynthesis [63]. Increases in succulence help in cell expansion and thus enable housing more chloroplasts [29]. Halophytes can accumulate over 20-fold higher Na+ levels in chloroplasts compared to glycophytes [63–68]. In addition to compensating for reduced photosynthesis, increases in chloroplast number may also contribute to increased energy demands for osmotic adjustment and ion homeostasis under salinity [29]. Although information about the genes involved in binary fission of chloroplasts is plentiful [68–70], the detailed effects of salinity on the mechanism/regulation of chloroplast multiplication, particularly among halophytes and glycophytes, are limited.

### **3. Effects of Salinity on Transport across Chloroplast Membranes**

Most of the nuclear-encoded proteins destined for chloroplasts are synthesized as 'cytosolic preproteins' and imported by a major pathway consisting of transmembrane protein complexes or channels, '*Toc*' (translocons on outer chloroplast) and '*Tic*' (translocons on inner chloroplast). The '*Toc*' transmembrane channels import larger molecules (including nuclear-encoded proteins) while the '*Tic*' complexes are more restrictive, with transport limited to targeted proteins [71]. Located at the interface between the stroma and the cytosol, the envelope is also the site for the transport and exchange of ions and metabolites required for the integration of the plastid metabolism within the plant cell. In general, chloroplasts harbor three types of membranes and each of them is equipped with a unique set of ion channels and transporters enabling the transport of nutrients, solutes, and

metabolites in and out of it. This is achieved by coordinated regulation of a variety of transport systems located in chloroplast membranes, such as porins, solute channels, ion-specific cation/anion channels and various primary and secondary active transport systems [29].

### *3.1. Protein Transport across Chloroplast Membranes*

The chloroplast proteome comprises 3000 different proteins, including components of the photosynthetic apparatus, which are highly abundant [72]. Most chloroplast proteins are nuclear-encoded, synthesized in the cytosol, and their import is mediated by multiprotein complexes in the envelope membranes that surround each organelle. The *Toc* complex mediates client protein recognition and early stages of the import. The *Toc* apparatus is regulated by the ubiquitin-proteasome system (UPS) in a process controlled by the envelope-localized ubiquitin E3 ligase SUPPRESSOR OF PPI1 LOCUS1 (*SP1*) [73]. Salinity stress depletes the *Toc* apparatus by enhancing the expression of *SP1*, which in turn may result in the suppression of photosynthesis activity [74].

### *3.2. Ion Transport across Chloroplast Membranes*

The proper ionic (K+, Na+, Cl<sup>−</sup>) balance is essential to control chloroplast volume [73]. For example, Cl− influx from stroma to the lumen is required for thylakoid swelling, while lumen shrinkage is due to K<sup>+</sup> (or Na+) efflux from the lumen to the stroma [75]. The outer membrane is not freely permeable to ions as some porins (OEP23, OEP37) are reported to have high cation selectivity [76], although information regarding their role in plant salt tolerance is lacking. The literature reports several nucleus-encoded candidate ion channels and transporters that regulate Na+, K+ and Cl<sup>−</sup> transport through the chloroplast envelope and thylakoid membranes [75–79]. A several-fold increased Na+ and Cl<sup>−</sup> concentration in barley chloroplasts under salt stress has been reported [76]. Slabu et al. [79] reported that salt-induced damage in broad bean chloroplasts is due to the accumulation of Na<sup>+</sup> and not of Cl<sup>−</sup> or K+. In contrast, salt toxicity and inhibition of photosynthesis in soybean were found associated with the hyperaccumulation of Cl− but not that of Na+ in chloroplasts [80,81].

### *3.3. Chloroplast Trafficking of Ions in Glycophytes vs. Halophytes*

Halophytes preferentially accumulate ~20-fold higher Na+ levels than glycophytes [64,65,67]. This high ion level is known to have some effect on chloroplast functions [63,64,67], especially in the case of CAM and C4 plants [81]. The Na+ contribution in the transport of pyruvate [82,83], ascorbate [84] and phosphate [85] into chloroplasts has been reported but the effect of salt stress on transport requires further elaboration. Salt stress induces K<sup>+</sup> loss from chloroplasts in both glycophytes and halophytes. Chloroplasts isolated from halophytes revealed better tolerance to high Na+ (100 mmol L−<sup>1</sup> Na+) and low K+ (50 mmol L−<sup>1</sup> K+) in the cytosol than chloroplasts of glycophytes [86]. Likewise, halophytes accumulate more Cl<sup>−</sup> than glycophytes under low salt conditions (≤1 mmol L−<sup>1</sup> Cl−), while at higher salinities some halophytes maintain steady Cl− concentrations, and others show a slight increase within the chloroplasts [63,65]. These findings indicate that halophytes have mechanisms to regulate the Cl− concentrations; however, the candidate transporters for Cl− regulation during salt stress remain uncharacterized.

### 3.3.1. Aquaporins and Non-Selective ion Channels

Aquaporins (PIP2;1, PIP2.3, PIP2;7, PIP1;3 and PIP1;2) are reported on the chloroplast membrane [77,78]. Expression of both PIP2;1 and PIP2;7 is altered by salinity [87]. Some aquaporins also have the ability to transport ions [88], but little is known about their function/regulation.

Non-selective ion channels include mechanosensitive channels (MSL2 and MSL3) that help reduce chloroplast swelling during hypo-osmotic conditions by releasing ions from the stroma [89]. In general, the ion selectivity of MSLs varies from non-selective to Cl−, K+, Na+ or Ca2+ selective channels [75].

### 3.3.2. Na+, K+ and Cl<sup>−</sup> Transporters

Sodium ions (Na+) can be transported into chloroplasts through an inner envelope membrane-localized Na+-dependent pyruvate transporter (*BASS2*) that is abundantly found in halophyte species compared to glycophytes [82]. Introduction of a halophyte BASS2 gene into glycophyte chloroplasts resulted in improved salt tolerance [83]. The inorganic phosphate transporters (thylakoid membrane-localized PHT4;1 and inner envelope localized PHT4;4 and PHT4;5) can use Na+ or H<sup>+</sup> as a co-transporting ion [79], thereby changing the Na+ concentration inside the chloroplasts. The existence of the Na+/H+ antiporter (NhaD; hereafter NHD)-type transporters at the chloroplast membrane mediating Na<sup>+</sup> efflux from the stroma was also reported in a halophytic tree, *Populus euphratica* [82]. In Arabidopsis, salt stress did not alter the expression of *NHD1* but silencing *NHD1* resulted in high chloroplast Na<sup>+</sup> and poor growth and photosynthetic performance [67]. In contrast, analysis of *Mesembryanthemum crystallinum* (a halophyte) under salt stress showed an increase in *NHD1* expression that resulted in higher Na+ accumulation, indicating the involvement of *NHD1* in Na<sup>+</sup> import into the chloroplasts instead of Na<sup>+</sup> export [41]. Such opposite regulation of ion transport mechanisms requires further investigation for a more complete understanding of the salt tolerance mechanisms.

Two K+ efflux antiporters (*KEA1 and KEA2*) located at the membrane of Arabidopsis have been suggested to function as K+/H+ exchangers mediating K+ export out of the stroma [90]. The Arabidopsis double loss-of-function *kea1kea2* mutant showed better growth under salt stress as compared to the wild type, due to low K+ efflux in the mutant resulting in increased K<sup>+</sup> retention as well as maintenance of pH in the stroma leading to improved photosynthetic performance and growth [91]. Arabidopsis *KEA3*, located in the thylakoid membrane, has been suggested to import K+ into the lumen in exchange for H<sup>+</sup> [91,92] and support in PSII quantum efficiency and CO2 assimilation under low light [93]; however, no information is available regarding *KEA3* function during salt stress.

Electrophysiological studies have shown the existence of Cl− permeable channels in the chloroplast envelope and thylakoid membranes [75]. A bestrophin-like protein from Arabidopsis has been discovered and shown to alter PMF portioning by functioning as a voltage-dependent Cl− channel (*AtVCCN1*) on the thylakoid membrane [93]. The effects of salinity on chloroplasts are summarized in the model in Figure 1.

**Figure 1.** A model that summarizes the effects of salinity stress on chloroplasts in salt-sensitive (glycophyte) and salt-tolerant (halophyte) plants.

### **4. Effect of Salinity on Osmotic Adjustment in Chloroplasts**

*4.1. What Is Osmotic Adjustment and How Is It Achieved?*

Hyper-osmotic stress due to salinity is well-known in plants and bacteria and may cause disrupted cell metabolism, turgor loss and growth arrest. However, an adaptive mechanism for water retention exists among organisms under stressed environments whereby they increase their osmolality, a phenomenon commonly termed as 'osmotic adjustment' [94]. Increases in osmolality are achieved by either of the following three mechanisms: (1) micro-organisms, such as bacteria or yeast, accumulate a range of osmolytes or compatible solutes available from the external medium; (2) plants activate genes for de novo synthesis of organic osmolytes (so-called 'compatible solutes'), such as glycine betaine, proline, sugars, polyols, etc.; and (3) plants regulate ion flux across cellular membranes [20,95].

### *4.2. Localization, Trafficking and Functions of Organic Osmolytes in Membrane-Bound Organelles*

Among the organic osmolytes, glycine betaine (GB), sugars (mannitol, sorbitol and trehalose), polyamines and proline are the most important and are accumulated under abiotic stresses and confer tolerance to cells without interfering with the cellular machinery of the plant [96]. Of these osmolytes metabolism of proline (PRO) depends upon two important enzymes, catalyzing its synthesis from glutamate in the cytoplasm or chloroplast and two enzymes catalyzing proline catabolism back to glutamate in the mitochondria along with an alternative pathway of its synthesis via ornithine [97]. During water deficit or physiological drought under salinity PRO is known to protect the photosynthetic apparatus as well as in cytokinin-dependent photorespiration [98]. Studies on other osmolytes suggest that sugar alcohols, such as sorbitol and mannitol, and quaternary ammonium compounds, such as GB and their precursors, are highly localized in chloroplasts [99,100] and are somehow involved in protecting the photosystem (PSII) and membrane proteins against ROS under stress conditions [95,101,102]. The impairment of thylakoid membranes that results from salt stress may be alleviated by GB probably via protection and stabilization of the protein complexes as well as changes in lipid composition of the thylakoid membrane, thereby improving photosynthesis [102]. The accumulation of GB in higher concentrations in the chloroplasts of young leaves suggests that these are the main sites of its biosynthesis [98,103] from where it is translocated to other plant parts via phloem [104]. Sugar alcohols and polyols, such as mannitol, sorbitol, etc., regulate osmotic balance by sequestering Na<sup>+</sup> in the vacuole or apoplast, thus protecting membranes against drought [105] and salt stress [106]. These osmolytes also scavenge ROS, particularly hydroxyl radicals that do not require high concentrations of osmolytes as needed for osmotic adjustment [97]. This leads to the conclusion that such compounds may be more important in 'osmoprotection' rather than 'osmotic adjustment'.

### *4.3. Are Osmolytes Compatible for Osmotic Adjustment in Planta?*

The classical concept of osmotic adjustment via accumulation of organic solutes in non-halophilic organisms still prevails [107,108] though it has been challenged by many physiologists [97,109,110]. A major shift in energy balance usually causes severe losses in growth yields of crop plants at the expense of other metabolic processes, raising the serious question of whether osmolytes are compatible in a real sense. Physiologists argue that conventional water retention under saline stress is not directly related to the contribution of organic solutes for many reasons. The first reason is the concentration of organic osmolytes, which seems to be too low compared to the inorganic solutes in cells. For instance, 3–10 mM in plants contributes less than 3% [111,112], while ~120–150 mM glycine betaine (GB) in plants contributes <50%, often ranging between 10 and 30% of the total cell solutes [112]. Even if it is assumed that most of the osmolytes are contained in the cytosol and chloroplasts (collectively constituting 10–15% of the cell volume) compared to a larger vacuolar fraction (~85%), this seems low given that 500–600 mM concentrations of Na<sup>+</sup> alone exist within the vacuole [99]. Osmolyte concentrations (GB in particular) between 200 and 300 mM may

be sufficient to prevent cytoplasmic dehydration, thereby achieving osmotic adjustment. In some of the succulent halophytes (which accumulates up to 1000 mM Na<sup>+</sup> and Cl−), ~200 mM plant water GB was reported in *Suaeda fruticosa* and about 600 mM in *Haloxylon stocksii* (sensu lato *recurvum*) under extreme saline conditions, which are exceptional as in other plants, including *Halopyrum mucronatum* and *Atriplex stocksii* (sensu lato *griffithii*), GB ranged between 100 and 150 mM [113]. The second reason for not considering organic osmolytes as 'compatible' is the cost of their synthesis, which is too high. For instance, 30–109 molecules of ATP may be required for osmolyte synthesis compared to one molecule of ATP for one K<sup>+</sup> and two Cl<sup>−</sup> in bacteria [114], while plants require approximately 41 molecules of ATP for proline, 50 for glycine betaine and 52 for sucrose [115]. Thirdly, the synthesis of such organic solutes is very slow, often ranging from hours to many days while plants growing in water-stressed environments require rapid turgor recovery [102]. Moreover, salt-sensitive genotypes of many crop plants, e.g., rice, wheat, barley, etc., accumulate comparatively higher amounts of osmolytes than salt-tolerant varieties, which also creates ambiguity in the role of osmolytes in achieving osmotic adjustment [81,109,116].

### *4.4. Effects of Osmolytes on Organelles*

Although the osmotic adjustment is based on the notion that osmolytes should not interfere with other metabolic processes, some studies on exogenous application of osmolytes suggest their toxic effects on plant growth as well as cell organelles [116,117]. Application of some L-amino acids (L-proline, L-alanine, etc.) in millimolar concentrations caused growth inhibition in suspension cultures of *Nicotiana silvestris* [116]. In another instance, a disruptive effect of PRO on the ultrastructure of chloroplasts in *Arabidopsis thaliana* was linked to feedback inhibition of its synthesis due to over-reduction of the photosynthetic electron acceptor pools [117]. In the same plant, exogenously supplied PRO was thought to have increased the rates of mitochondrial electron transport, resulting in elevated levels of ROS causing subcellular damage [117]. On the contrary, endogenous PRO did not seem to have a negative impact on the ultrastructure of chloroplasts and mitochondria in transgenic tobacco, indicating that this level of PRO had no toxic effects [118]. Though the assumption of osmolyte toxicity is largely inconclusive, it seems that plants treated with exogenous application of osmotica may suffer from an 'overdose' compared to their endogenous levels. In fact, in certain cases, exogenous application (both foliar as well as through the rooting medium) of osmolytes such as GB, PRO, inositol, and mannitol have indicated stress alleviation in many plants [29,119]. Exogenous application of osmotica, such as GB, may also enhance the membrane integrity of chloroplasts and also increase PS II efficiency [97,98], suggesting an osmoprotective role. Experiments on exogenous application of osmolytes have intrigued geneticists to manipulate the biosynthetic pathway of compatible solutes to enhance salt tolerance as osmolyte accumulation is often controlled by only one gene [102].

### *4.5. Possible Role of Osmolytes in Ion Regulation*

Although the published literature has contradicting reports on the role of osmotic adjustment via osmolytes for maintaining turgor, recent patch-clamp studies suggest that osmolytes may have a significant contribution in regulating ion transporters such as K<sup>+</sup> outward rectifying channels (KORs), though this requires further investigation [120]. Thus, ion regulation via osmolytes may prove to be an important aspect in conferring salt tolerance. In plants, K<sup>+</sup> appears to be the most abundant cation in the cytosol (100–150 mM), which may account for osmotic adjustment [121], though under stressed conditions, the electrochemical gradients may lead to the loss of K+. In halophytes, Na+ and Cl<sup>−</sup> seem to play a major role in osmotic adjustment [105]. Of these, Na<sup>+</sup> may enter the cell passively and could be used as a cheap osmoticum for maintaining cell turgor. Since Na+ is toxic and may cause an imbalance in the cytosolic K+/Na+ ratio and interferes with cell metabolism, its efficient sequestration in the vacuole is thus essential. Pumping of one mole of Na+ against the electrochemical gradient requires only 3.5 mol of ATP compared with 30–50 mol

of ATP for one mole of organic osmolyte [115]. As mentioned above, some of the sugar alcohols and polyols regulate osmotic balance by sequestering Na+ in the vacuole or apoplast. It appears that osmotic adjustment is collectively achieved by maintaining a balance between ion regulation, synthesis and accumulation of organic solutes, as well as maintenance of K+ in the cytosol [122].

### **5. Effects of Salinity on Function and Protection of Photosystems**

Under saline conditions, decreases in CO2 assimilation via the Calvin cycle accompany a decrease in photochemical electron sink, which in the presence of light impacts the functioning/efficiency of photosystems [31]. In some sensitive plants such as olives, decreases in the *Fv/Fm* ratios indicate the incidence of photodamage under saline conditions [35]. Likewise, increases in salinity resulted in a gradual decrease in activities of PSI and PSII in four rice cultivars [123]. However, unchanged *Fv/Fm* hints towards sustained PSII under saline conditions [124], such as in the Mangalamahsuri variety of rice [125]. PSII-mediated electron transport increased in low salinity followed by a decrease at high salinity in the halophyte *Bruguiera parviflora* [55]. In other instances, inhibition of de novo protein synthesis, especially of the D1 protein, indicated a lack of efficient PS II repair under saline conditions [2,126,127]. A compensation mechanism of PsbO protein induction has been observed in some studies to stabilize the PSII structure under salinity [128]. Among C3 plants, salinity reportedly resulted in poor PSII function in glycophytes such as rice and Arabidopsis [41,129] but not in the halophyte *Arthrocnemum macrostachyum* [130,131]. Several tolerant species, including halophytes such as *Sarcocornia fruticosa* [132] and *Atriplex centralasiatica* [133], also employ the xanthophyll cycle for non-photochemical quenching that dissipates excess excitation energy of PSII in the form of heat as a 'first line of defense' [31,133], thus preventing the formation of potentially cytotoxic reactive ROS. The xanthophyll cycle enzyme violaxanthin de-epoxidase consumes NADPH, which if accumulated may cause the over-reduction of reaction centers, and thereby enhance ROS (especially superoxide) formation [134]. Hence, the timely induction of the xanthophyll cycle may protect plants under stressful conditions in multiple ways. Many halophytes are reported to exhibit reversible midday photoinhibition of PSII activity to limit excitation of the PSII reaction centers [130,134]. This mechanism also minimizes the possibility of ROS formation in salt-stressed plants under high light and is considered an important ecophysiological adaptation to salinity [31]. A decrease in the antennae size due to decreased chlorophyll content was also observed in *Arthrocnemum macrostachyum* to limit PSII excitation [132].

PSI is reportedly more stress-resistant than PSII and seems to impart salt tolerance by increasing cyclic electron flow to generate ATP while avoiding the accumulation of toxicreducing species [135–137]. Information about PSI in halophytes is scarce. PSI reaction center subunit IV protein (PsaE) was upregulated under salinity in wild halophytic rice *Porteresia coarctata* but not in conventional sensitive rice [137]. Similarly, salinity treatment caused an increase in PSI transcripts in *M. crystallinum* [138]. Formation of ATP via cyclic electron flow around PSI helped to prevent overaccumulation of Na<sup>+</sup> in chloroplasts of soybean [139].

Cultured plant cell lines have also been utilized to study salt-adapted tobacco cells [140,141]. Heterotrophic tobacco cells adapted to grow at 428 mM NaCl showed elevated levels of chlorophyll and lower levels of starch along with increased CO2 fixation, oxygen evolution and photorespiration, compared to unadapted cells [140]. This was coupled with higher levels of PS-I- and PS-II-associated proteins, including Rubisco. These cells were found to have acquired a significant level of salt-tolerant photosynthetic competence [140]. Further analysis showed that oxygen evolution and CO2 fixation were more resistant to inhibition by NaCl in the salt-adapted cells [141].

### **6. Effects of Salinity on CO2 Assimilation Enzymes**

Information on the effects of salinity on chloroplast CO2 assimilation enzymes is limited among halophytes. Generally, CO2 assimilation reactions are considered more sensitive to salinity than photochemical reactions of photosynthesis [31]. Several studies have reported that salinity generally inhibits many enzymes of the Calvin cycle [137,142,143].

Ribulose-1,5-bisphosphate carboxylase/oxygenase (Rubisco) is the key photosynthetic enzyme that catalyzes the fixation of atmospheric CO2 in plants during the Calvin cycle [144]. It is the most abundant protein in leaves that accounts for 30% (C4 plants) to 50% (C3 plants) of total soluble protein in leaves [145]. In C3 plants, it is localized in all chloroplasts while in C4 plants with Kranz anatomy, Rubisco is localized specifically in the bundle sheath but not mesophyll chloroplasts [65]. In single-cell C4 species, Rubisco mRNA could be targeted to the proximal or central compartment of chloroplasts [146]. The activity of Rubisco was mostly examined by direct measurement of the enzyme activity or protein levels and measurement of its carboxylase activity (Vcmax) [12]. Salinity exposure causes a decrease in Rubisco activity in most plant species regardless of C3 or C4 type [19,146–148]. In addition, the Rubisco levels also decreased under saline conditions in both halophytes and glycophytes. For example, salinity caused an inhibition (~50%) of Rubisco activity in maize, a glycophyte, and in *Atriplex spongiosa*, a halophyte [149]. In some other instances, Rubisco activity was improved both in either low [13,20] or high salinity [150]. Rubisco activity also depends on the function of a supporting enzyme, Rubisco activase, which revitalizes the active sites of Rubisco by removing inhibitory sugar phosphates [151,152]. The enhanced activity of Rubisco activase was found in rice as well as in many halophytes, such as *S. salsa* [143] and *Thellungiella salsuginea*, under saline conditions [153]. More efficient Rubisco activation was found in *T. salsuginea* compared to *Arabidopsis thaliana* [153].

Chloroplastic fructose-1,6-bisphosphatase is considered a metabolic control point of the Calvin cycle [44,154]. In vitro salt sensitivity of this enzyme was higher in saltsensitive rice (*Oryza sativa* cv. IR26) than its wild halophytic relative *Porteresia coarctata* [142]. However, the inhibitory effects of salinity could be reversed by preincubation of the enzyme with osmolytes (effectiveness order: polyol>sugars) [142], suggesting a lower level of in vivo inhibition of chloroplastic fructose 1,6-bisphosphatase under salinity in halophytes with higher amounts of osmolytes compared to glycophytes.

Phosphoenolpyruvate carboxylase (PEPC) is the key enzyme of C4 photosynthetic metabolism that catalyzes the β-carboxylation of phosphoenolpyruvate to form four-carbon acid oxaloacetate in the mesophyll cells [144,155]. It is considered more sensitive to salinity than Rubisco [149]. Furthermore, PEPC isolated from the halophyte *Atriplex spongiosa* was found more salt-sensitive in the in vitro studies than the one from the glycophyte maize [149]. Contrary to these observations, an increase in PEPC activity was reported in the halophyte *Mesembryanthemum crystallinum* [156] and in the C4 species *Bienertia sinuspersici* under salinity [157]. Increased PEPC activity helps concentrate CO2 around Rubisco and substantially reduces the incidence of photorespiration, a major cause for growth reduction and ROS formation under environmental stresses in plants.

Pyruvate orthophosphate dikinase is the rate-limiting enzyme of the C4 cycle that catalyzes a reversible reaction to regenerate the primary CO2 acceptor phosphoenolpyruvate (PEP) [158]. However, its role in C3 plants is not fully understood [159]. Pyruvate orthophosphate dikinase is found in both chloroplasts and the cytoplasm irrespective of C3 or C4 types [160]. In C4 plants, it can comprise up to 10% of the total protein pool [161]. Interestingly, both isoforms are encoded by a single nuclear gene [162]. The labeling of the pyruvate orthophosphate dikinase protein was observed both in mesophyll and bundle sheath chloroplasts of kranz type C4 plant maize, albeit with higher levels in the latter rather than the earlier-mentioned chloroplasts [163]. In single-cell C4 species, pyruvate orthophosphate dikinase mRNA could be targeted to the peripheral or distal compartment chloroplasts [146]. Information about the impacts of salinity on the abundance and activity of this enzyme is scant. Salinity caused an increase in pyruvate orthophosphate dikinase levels in both types of chloroplasts in maize [163]. These enzymes are widely studied and

are important for the biochemical reactions of photosynthesis [17,164,165]. Induction of PEP activity would also help maintain C4 functionality under salinity stress and facilitate CO2 assimilation for biomass buildup and reduce photorespiration, as mentioned above.

### *Effects on Salinity on the Gas Exchange Ecophysiology of Photosynthesis*

The effects of salinity on photosynthetic synthetic gas exchange, which eventually supports CO2 assimilation at the chloroplast level, varies not only among species but also depends on the magnitude of the salinity. For instance, the net CO2 assimilation rate (*PN* or *A*) and stomatal conductance (*Gs*) in sugar beet improved under low (75 mM NaCl) salinity while high (250 mM NaCl) was inhibitory [166]. An increase in *PN* but not in transpiration (*E*) under low salinity resulted in improved water-use efficiency (WUE) in sugar beet plants [166]. Salinity stress decreased the *PN* and *Gs* in wild-type wheat plants [24]. *PN* and *Gs* increased transiently at 200 mM NaCl in comparison to controls and 400 mM NaCl in the halophyte *Sesuvium portulacastrum* [167]. Similarly, in many other halophyte species, such as *Arthrocnemum macrostachyum* (in up to 510 mM NaCl) [133] and *Atriplex portulacoides* (200 mM NaCl) [168], low to moderate salinity improved *PN*. In contrast, salinity exposure resulted in decreased *PN* and *Gs* in the halophytes *Panicum antidotale* [20] and *Aster tripolium* [169]. Hence, impacts of salinity not only vary among glycophytes but also halophyte species. In many cases, decreased *Gs* improves the WUE of plants under stress conditions as a trade-off at the expense of *PN*. For instance, in *Sarcocornia fruticosa*, increased WUE accompanied a decline in *PN* [170]. Similarly, many halophytes exhibit C4 and CAM modes of photosynthetic CO2 assimilation, which not only reduce wastage of photosynthetic energy through photorespiration but also decrease the consequent H2O2 (a common ROS) production at the peroxisome level [31,171].

### **7. Effects of Salinity on Chloroplast ROS Homeostasis**

Exposure of plants to salinity results in a reduction in CO2 assimilation rates, which in turn leads to the overreduction of PSII along with diversion of electrons to molecular oxygen, which generates reactive oxygen species (ROS), particularly singlet oxygen [31,172,173]. In photosynthesizing leaves, chloroplasts are the major site for ROS production during the daytime [174]. In C3 plants, photorespiration resulting from the oxygenase activity of Rubisco in chloroplasts is another source of ROS generation in peroxisomes [31]. Salinity-induced stimulation of electron flow to molecular oxygen has been reported in several plant species [31,175,176]. Major ROS produced in chloroplasts include singlet oxygen (1O2), superoxide radical (O2 •−), hydrogen peroxide (H2O2) and hydroxyl radical (•OH) [31,173]. Since detection of radicle-type ROS is difficult, most studies examine H2O2 (non-radicle ROS) formation following salinity exposure [31]. In addition, studies on ROS formation in isolated chloroplasts, particularly of halophytes, are very limited. Wiciarz et al. [153] reported that isolated thylakoids from a halophyte *Thellungiella salsuginea* produced higher H2O2 levels than the model glycophyte *Arabidopsis thaliana*. However, when both plant types were exposed to salt stress, even at the low level of 100 mM NaCl, Arabidopsis produced a higher H2O2 than *T. salsuginea* and at a 300 mM NaCl concentration. Similarly, a substantially higher H2O2 level was observed in chloroplasts of wild salt-tolerant tomato *Lycopersicon pennellii* compared to chloroplasts of sensitive tomato *L. esculentum* under stress-free growth conditions. However, under NaCl stress, a decrease in H2O2 level was noted for wild tomato while the levels were increased in the sensitive species [171]. This indicates that halophyte species have efficient mechanisms to control the production of ROS or detoxify them compared to glycophytes, either through the dissipation of excess excitation energy to alternative electron sinks, such as the plastid terminal oxidase [29,172–180] (PTOX) or ROS-scavenging system [31,174]. Alternative electron sinks not only provide 'safety valves' for the efficient functioning of the photosynthetic machinery but also act as an 'avoidance' tool for control of ROS formation. Tightly regulated levels of ROS are now acknowledged as 'signals' for the regulation of different plant processes, including the defense/tolerance response of plants [3,99]. For

instance, ROS modulate the function of some plasma membrane ion transporters, such as those regulating cytosolic Na+ and K+ [177,181–185]. Similarly, a ROS 'surge' in response to salinity exposure may also activate chloroplast retrograde signaling pathways [180].

In order to prevent oxidative damage due to ROS accumulation, chloroplasts possess many enzymatic and nonenzymatic antioxidants [29,138,182,185]. Key enzymatic antioxidants are superoxide dismutases (SOD), enzymes of the Foyer–Halliwell–Asada pathway (also known as the ascorbate–glutathione cycle), and glutathione peroxidase (GPX), whereas ascorbate and glutathione are common nonenzymatic antioxidants of chloroplasts (Figure 2) [138,182,186]. Antioxidants in various combinations play an important role to keep the levels of ROS in 'functionally useful' ranges for signaling various plant processes and stress responses [31]. The water–water cycle is among the key processes responsible for ROS homeostasis in chloroplasts and is essential for salinity tolerance (Figure 2) [31,187]. Ground state molecular oxygen (O2) produced during photolysis of water in chloroplasts can accept electrons from excited photosystems, particularly the thylakoid membrane-bound primary electron acceptor of PSI to form O2 •− through a reaction called the Mehler reaction [188]. The acceptor side of the electron transport chain in PSII may also contribute to electron leakage to O2 to generate O2 •−. Thylakoid membranebound copper/zinc superoxide dismutase (Cu/Zn SOD) converts O2 •− into H2O2, which is finally reduced into the water by the action of thylakoid membrane-bound ascorbate peroxidase (tAPX), thus completing the 'water–water cycle' [189]. The Foyer–Halliwell–Asada pathway (also known as the ascorbate–glutathione cycle) in chloroplasts is an extension of the water–water cycle and involves quenching of ROS in chloroplasts by consuming NADPH, which also contributes to relaxing the 'overreduction of photosystems' by providing NADP (the final electron acceptor of PSI), and thereby minimizing the chances of further ROS generation (Figure 2) [167,190]. In this pathway, the H2O2 generated from dismutation of O2 •− by SOD is neutralized into water by the action of stromal ascorbate peroxidase (APX) using ascorbate (AsA) as the electron donor. Oxidized ascorbate is recycled by monodehydroascorbate reductase (MDHAR) and/or dehydroascorbate reductase (DHAR). The latter consumes glutathione (GSH), which is finally recycled by the action of glutathione reductase (GR) that uses NADPH as an electron donor [167,190,191]. Often, an upregulation of enzymes involved in antioxidant processes is reported in chloroplasts under environmental stresses, with a higher magnitude of tolerance compared to sensitive species [31,181]. For instance, salinity exposure resulted in enhanced activities of SOD, APX and MDHAR in chloroplasts of halophytic wild tomato *Lycopersicon pennellii* compared to conventional sensitive tomato *L. esculentum* [172]. In addition, thioredoxin/peroxiredoxin (Trx/Prx) and glutathione peroxidase (GPX) also reportedly quenched salinity-induced excess H2O2 in chloroplasts [187]. Lipophilic tocopherol can protect chloroplast thylakoid membranes from oxidative damage [189]. The 1O2 produced by PSII is mainly detoxified by carotenoids and tocopherols found in the chloroplast membranes [192]. Carotenoids detoxify 1O2 not only through the xanthophyll cycle (NPQ) but also by direct quenching of 1O2 [193]. However, some C4 plants, especially those with NADP-malic enzyme (NADP-ME) subtypes, lack PSII in their bundle sheath chloroplasts and hence supposedly lack 1O2 production [194].

**Figure 2.** The Foyer–Halliwell–Asada pathway (also known as the ascorbate–glutathione cycle) and the water–water cycle are responsible to quench the superoxide radicles and hydrogen peroxide in the chloroplasts.

### **8. Summary**

Soil salinity is one of the major challenges to the sustainable development of agriculture in different parts of the world. Salinity has detrimental effects on plant growth by imposing several constraints. For instance, salt-induced toxicity impairs the normal functioning of the organelles, such as chloroplasts—the green plastids—which house several important biochemical reactions, including photosynthesis. Chloroplast dysfunction as a result of various environmental stresses, including salinity, has been reported to have detrimental effects on plants [173]. Chloroplasts, in addition to being a site of various metabolic reactions, also act as global sensors to sense and communicate the developmental, operational and environmental changes to the nucleus.

Understanding the effect of salinity on chloroplast function and the response of various metabolic reactions to salt stress is necessary for the development of salt-tolerant crops. Little attention has been paid to how salinity affects chloroplasts and the stromal metabolic reactions. Salinity-related changes in the size, number, lamellar organization, lipid and starch accumulation, and trafficking across the chloroplast membrane are dependent on the plant species and its level of salt tolerance. Chloroplast swelling or alteration in thylakoid membranes of glycophytes may be linked with the ionic component of salinity while some halophytes are affected by the osmotic effect of high salinity (Figure 1). Most halophytes either maintain chloroplast structure or enhance grana development under salinity stress. Swelling of thylakoids and disruption of chloroplast envelopes in mesophyll cells along with intact chloroplasts in bundle sheath cells is a general C4 response under salinity, irrespective of the subtype.

Halophytes and glycophytes have evolved different pathways to respond to salinity stress. For example, halophytes are much better adapted at maintaining a lower salt concentration in the cytoplasm compared to glycophytes. Likewise, chloroplasts in halophytes seem to have a better antioxidant system than those of glycophytes, and consequently more protected photosynthetic apparatuses under salt stress. Similarly, salinity-triggered starch

deposition appears to be a damage symptom in glycophytes but a survival strategy in halophytes. The salinity-induced influx of Na+ and Cl<sup>−</sup> appears beneficial for halophytes but lethal for glycophytes (Figure 1). Accumulation of Na+ or Cl<sup>−</sup> disrupts ionic homeostasis, impairs protein synthesis and interferes with the enzymatic activities of the organelle. However, recent work suggests that the negative effects of these ions on plant health are not because of toxicity per se but are the result of interference with the absorption or metabolism of other essential ions [28]. This view stems from the evidence that K<sup>+</sup> influx in chloroplasts is reduced with excessive Na+ or Cl<sup>−</sup> accumulation. K<sup>+</sup> is an essential element for the plant cell and is not only required for chloroplast development but also for pH regulation, maintenance of the electron transport chain and thylakoid restacking [28,29]. Osmolyte synthesis suggests that organic solutes may help in fine adjustment along with ion transport (vacuolar compartmentation) and accumulation of cytosolic K+ in stressed environments rather than osmotic adjustment. However, osmolytes are certainly involved in the osmoprotection of membrane transport proteins and the scavenging of ROS. Despite ion regulation and osmotic adjustment, salinity induces many changes in chloroplast functions and signaling.

Chloroplastic CO2 fixation is generally more sensitive to salinity than the thylakoid reactions. However, CO2 fixation in many halophytes is reportedly less prone to salinity compared to glycophytes. One major evolutionary adaptation that seems to operate in halophytes is the switching of CO2 concentration around Rubisco under stressful environmental conditions, including salinity. The reduced photosynthetic efficiency is considered a major salt-induced constraint inhibiting plant growth, and ultimately crop productivity. However, it is not yet clear whether the decrease in photosynthesis is the cause of growth reduction or the reduction in the growth rate causes a decrease in photosynthesis under salt stress. Nevertheless, a reduced rate of photosynthesis leads to higher production of ROS and also triggers the activity of ROS-scavenging enzymes. The higher activity of the ROS-detoxifying enzymes maintains a level of these species in a functionally useful range required for cell signaling. These enzymatic systems are naturally present in plants. Although differences in the activity of these enzymes have been reported in different genotypes, it is believed to be associated with responses such as stomatal closures, reduction in the CO2 fixation rates and an increase in photorespiration under stressful conditions [188,194]. Tight regulation of ROS alongside many chloroplastic metabolites also function as 'putative' signals for communication between chloroplasts and the nucleus (as well as other organelles) via so-called 'retrograde signaling'. Despite information on crop and model plants, our knowledge about such signaling in halophytes is still far from full comprehension. Chloroplast functions, including photosynthesis, are integrated with other basic plant metabolic mechanisms of the plant in response to stresses, including salinity, and multiple factors work together to confer tolerance against salinity [195]. These factors include ion regulation that controls uptake and transport of salt and other ions to compartments within the plant cell, synthesis of compatible solutes, antioxidative enzymes and plant hormones and changes in photosynthesis and membranes in the cell [195]. Some of these occur within the chloroplast but are not limited to that location. These mechanisms are quite complicated, and many questions remain unanswered [195,196]. Some of these questions include how the plant senses salinity to initiate the signaling process, the precise details of how salinity stress leads to stomatal closure and growth reduction and the specific targets of ion toxicity in plant cells [196]. While advances are being made, a detailed understanding of the mechanisms behind salt tolerance is not yet clear. A comprehensive understanding of these mechanisms by employing multidisciplinary approaches is necessary for their effective incorporation into salt-sensitive crops for better crop yields under stressful environments.

**Author Contributions:** All authors participated in the writing of different sections, editing, and proofreading of the manuscript; M.Z.A. (Figure 1) and A.H. (Figure 2) prepared the figures. All authors have read and agreed to the published version of the manuscript.

**Funding:** No specific funding was received for this work.

**Institutional Review Board Statement:** Not applicable.

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** As a review paper all data is available in the work referenced.

**Acknowledgments:** We are grateful to Narendra K. Singh, Auburn University (emeritus), for helpful discussions in developing this review.

**Conflicts of Interest:** The authors declare no conflict of interest.

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