# **Plant Physiology under Abiotic Stresses**

Edited by Yanyou Wu Printed Edition of the Special Issue Published in *Horticulturae*

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## **Plant Physiology under Abiotic Stresses**

## **Plant Physiology under Abiotic Stresses**

Editor

**Yanyou Wu**

MDPI • Basel • Beijing • Wuhan • Barcelona • Belgrade • Manchester • Tokyo • Cluj • Tianjin

*Editor* Yanyou Wu Chinese Academy of Sciences China

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This is a reprint of articles from the Special Issue published online in the open access journal *Horticulturae* (ISSN 2311-7524) (available at: https://www.mdpi.com/journal/horticulturae/ special issues/plant physiology abiotic).

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## **Contents**

Reprinted from: *Horticulturae* **2021**, *7*, 510, doi:10.3390/horticulturae7110510 ........... **119**

## **About the Editor**

#### **Yanyou Wu**

Yanyou Wu received his Ph.D. degree from Sichuan University, China, in 1994. Now he is a professor at the Research Center for Environmental Bio-Science and Technology, State Key Laboratory of Environmental Geochemistry, Institute of Geochemistry, Chinese Academy of Sciences. His research focuses on interdisciplinary of biology and geology, such as biogeochemical cycle of carbon. His recent interests concentrate on the adaptation mechanism of plants to the environment and plant electrophysiology.

## *Editorial* **Plant Physiology under Abiotic Stresses: Deepening the Connotation and Expanding the Denotation**

**Yanyou Wu**

Research Center for Environmental Bio-Science and Technology, State Key Laboratory of Environmental Geochemistry, Institute of Geochemistry, Chinese Academy of Sciences, Guiyang 550081, China<EMAIL_ADDRESS>
**Abstract:** Abiotic stress factors influence many aspects of plant physiology. The works collected in the Special Issue deepen plant physiology's connotation (such as plant electrophysiology) under abiotic stress and expand the denotation (such as environmental pollutants as abiotic stress factors). At the same time, the achievements of the selected papers published in the Special Issue also exhibit their potential application value in the production of horticultural plants.

#### **1. The Effects of Various Abiotic Stresses**

The effects of various abiotic stresses on substance metabolism, energy transformation, and the growth and development of different plants are disparate. Abiotic stress includes several adversities, such as drought, temperature, light, salinity, nutrients, and heavy metals, as well as complex stresses, such as karst environments, saline–alkali soils, and coastal wetland environments. Water is the essential substance necessary for plant survival. A water deficit adversely affects plant morphology, growth and development, and physiological and biochemical processes. Therefore, water stress is also the most common abiotic stress factor. Most of the research works collected in the Special Issue concern the effects of water stress on plant physiology, biochemistry, and growth and development [1–4]; however, these works also focus on some complex multifactorial stresses [5–7].

#### **2. Moderate Drought Stress Has a Positive Effect on Plant Secondary Metabolism**

Generally, water stress harms the primitive metabolism of plants [1–4], but appropriate water stress has a positive effect on plant secondary metabolism and adaptation. Honório et al. found that under moderate drought stress, the production of liriodenine and total alkaloids in *Annona crassiflora* Mart. increased, but photosynthesis did not decrease [3]. Al-Quraan et al. demonstrated that drought can promote γ-aminobutyric acid (GABA) accumulation, but it also inhibits the synthesis of chlorophyll a and chlorophyll b in green pea seedlings (*Pisum sativum* L.) [2]. The abovementioned works show that humans can control soil to reach a moderate humidity level to produce secondary metabolites.

#### **3. The Application of Some Exogenous Organic Compounds Can Restore Metabolic Activity Reduced Due to Drought Stress**

Drought causes damage to the cell membrane, disrupts cell function, and reduces the physiological vitality of plants. Yeast extract is rich in essential nutrients and is a natural and safe biofertilizer. Chitosan is an environmentally friendly biological polysaccharide obtained by the deacetylation of chitin, which is widely present in nature. However, Abdelaal et al. presented that the application of yeast extract (8 g/L) or chitosan (30 mM) alone or in combination can enhance the antioxidant capacity of plants and restore metabolic activity reduced due to drought stress [8]. We can expect that appropriate drought combined with the application of yeast extract and/or chitosan can increase the synthesis of secondary metabolites in plants.

**Citation:** Wu, Y. Plant Physiology under Abiotic Stresses: Deepening the Connotation and Expanding the Denotation. *Horticulturae* **2023**, *9*, 218. https://doi.org/10.3390/ horticulturae9020218

Received: 18 December 2022 Accepted: 20 January 2023 Published: 7 February 2023

**Copyright:** © 2023 by the author. 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/).

#### **4. Plant Electrophysiology Can Deepen the Understanding of Plant Physiology under Abiotic Stress**

When affected by abiotic stresses, the anatomy, function, and metabolic activities of plant cells change, and these changes are bound to be reflected in the electrical properties of plant tissues and organs. Plant electrical signals can quickly respond to water status in the environment in real-time and characterize intracellular water metabolism. The works collected in the Special Issue demonstrate plant electrophysiology's great potential in the study of plant stress biology. According to the Gibbs free energy equation and Nernst equation, the relationship model between the clamping force and the electrical signal (capacitive reactance, resistance, impedance, and inductive reactance) of plant leaves can be quantitatively reflected in real-time by the intracellular water-holding capacity, water transfer rate, water-holding time, and water use efficiency of plant leaves. *Broussonetia papyrifera* (L.) Vent. and *Morus alba* L. have different intracellular water metabolism mechanisms adapted to soil water deficits [4]. The karst-adaptable plant *Orychophragmus violaceus* (L.) O. E. Schulz adapts to karst drought environments by synergistically regulating intracellular water transport, cell elasticity, and leaf anatomy [7]. The acquisition and analysis of plant electrophysiological information have opened up a broad path for studying plant physiology under abiotic stress.

#### **5. The Environmental Pollutants Become Abiotic Stress Factors**

Climate change and environmental pollution caused by human activities have gradually become abiotic stresses, which hugely impact plant life activities. With a background of elevated atmospheric carbon dioxide, the adverse effects of low temperature on plant growth and development are much greater than the adverse effects of high temperature on plant growth and development [5]. Environmental pollutants such as sulfur dioxide and bisulfites have become abiotic stress factors that seriously affect the physiological activities of plants. The work of Li et al. shows that suitable concentrations of NaHSO3 can be used as a source of sulfur for plants to play a positive role in the physiological activities of plants, and high concentrations of NaHSO3 reduce the photosynthetic capacity of plants and adversely affect plant growth and development. The effects of bisulfites as photorespiratory inhibitors are often masked [9].

#### **6. Complexity and Diversity of Salinity Stress**

Both salt in the soil and salt spray in the atmosphere can become abiotic stress factors. The influence of salt stress on coastal plants is caused by the superposition of soil salinity and atmospheric salt spray stress, and salt composition is also compounded. The root is the earliest organ to respond to salt stress in the soil. *Tilia cordata* Mill. can only respond to salinity by adjusting the stomatal conductance of leaves and reducing photosynthetic capacity by improving water use efficiency, as its roots do not prevent salt intake. *Pyrus pyraster* L. (Burgsd.) maintains stable photosynthesis under salt stress, as its roots limit the transfer of salt ions to leaves [10]. The first organ to respond to salt spray in the atmosphere is the leaves of plants. *Callistemon citrinus* significantly decreases photosynthetic capacity under salt spray stress [11]. In fact, plants growing in soil salinization areas are not simply affected by single-salinity adversity but by salinity–alkali compound adversity. Wang et al. found that low concentrations of saline–alkali stress (NaCl:NaHCO3 = 3:1, 0~90 mM) can promote glucose metabolism and the synthesis of antioxidant enzymes, while high concentrations of saline–alkali stress (NaCl:NaHCO3 = 3:1, 120~150 mM) reduce the activities of peroxidase and catalase [6]. This is related to the fact that at an appropriate concentration, bicarbonate improves the stress tolerance of plants [12]. In horticultural plant production, saltwater dilution irrigation is an effective measure.

**Funding:** This research was funded by Support Plan Projects of Science and Technology of Guizhou Province [No. (2021)YB453].

**Data Availability Statement:** Not applicable.

**Acknowledgments:** I gratefully acknowledge all the authors that participated in this Special Issue.

**Conflicts of Interest:** The author declares no conflict of interest.

#### **References**

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## *Article* **Effect of Three Water Regimes on the Physiological and Anatomical Structure of Stem and Leaves of Different Citrus Rootstocks with Distinct Degrees of Tolerance to Drought Stress**

**Waqar Shafqat 1,2, Yasser S. A. Mazrou 3,4, Sami-ur-Rehman 5, Yasser Nehela 6,7,\*, Sufian Ikram 5,8, Sana Bibi 9, Summar A. Naqvi 5, Mansoor Hameed 9 and Muhammad Jafar Jaskani 5,\***

**Abstract:** Citrus is grown globally throughout the subtropics and semi-arid to humid tropics. Abiotic stresses such as soil water deficit negatively affect plant growth, physiology, biochemistry, and anatomy. Herein, we investigated the effect(s) of three water regimes (control, moderate drought, and severe drought) on the physiological and anatomical structure of 10 different citrus rootstocks with different degrees of tolerance to drought stress. Brazilian sour orange and Gadha dahi performed well by avoiding desiccation and maintaining plant growth, plant water status, and biochemical characters, while Rangpur Poona nucellar (*C. limonia*) and Sunki × bentake were the most sensitive rootstocks at all stress conditions. At severe water stress, the highest root length (24.33 ± 0.58), shoot length (17.00 ± 1.00), root moisture content (57.67 ± 1.53), shoot moisture content (64.59 ± 1.71), and plant water potential (−1.57 ± 0.03) was observed in tolerant genotype, Brazilian sour orange. Likewise, chlorophyll *a* (2.70 ± 0.06), chlorophyll *b* (0.87 ± 0.06) and carotenoids (0.69 ± 0.08) were higher in the same genotype. The lowest H2O2 content (77.00 ± 1.00) and highest proline content (0.51 ± 0.06) were also recorded by Brazilian sour orange. The tolerance mechanism of tolerant genotypes was elucidated by modification in anatomical structures. Stem anatomy at severe drought, 27.5% increase in epidermal cell thickness, 25.4% in vascular bundle length, 30.5% in xylem thickness, 27.7% in the phloem cell area, 8% in the pith cell area, and 43.4% in cortical thickness were also observed in tolerant genotypes. Likewise, leaf anatomy showed an increase of 27.9% in epidermal cell thickness, 11.4% in vascular bundle length, 21% in xylem thickness, and 15% in phloem cell area in tolerant genotypes compared with sensitive ones. These modifications in tolerant genotypes enabled them to maintain steady nutrient transport while reducing the risk of embolisms, increasing water-flow resistance, and constant transport of nutrients across.

**Keywords:** drought stress; citrus; oxidative stress; proline; photosynthesis; water potential; vascular bundle modifications

**Citation:** Shafqat, W.; Mazrou, Y.S.A.; Sami-ur-Rehman; Nehela, Y.; Ikram, S.; Bibi, S.; Naqvi, S.A.; Hameed, M.; Jaskani, M.J. Effect of Three Water Regimes on the Physiological and Anatomical Structure of Stem and Leaves of Different Citrus Rootstocks with Distinct Degrees of Tolerance to Drought Stress. *Horticulturae* **2021**, *7*, 554. https://doi.org/10.3390/ horticulturae7120554

Academic Editor: Yanyou Wu

Received: 21 October 2021 Accepted: 1 December 2021 Published: 6 December 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/).

#### **1. Introduction**

Citrus fruit crops in the family Rutaceae have the largest fruit industry globally [1,2]. Rootstock choice and selection for the citrus scion variety are the most valuable decisions for growers for the implication of better yield and quality with other valuable characters. Citrus rootstock controls the physiological, biochemical, morphological, and genetic characteristics of scion cultivars grafted on selected rootstocks through the rootstock scion interaction pathway [3]. Fruit juice quality and tree productivity of scion cultivars are also affected by rootstock characters. Rootstock has a significant impact on nutrition, horticultural and pathological traits of citrus cultivars, growth, vigor, stress resistance, and fruit quality of the scion [4–7]. Rootstock controls translocation of water and nutrients from the soil and distributes among different up ground plant organs, which were also disturbed negatively by the impact of stresses [8,9].

Citrus plants affected by several biotic (fungal, viral, and bacterial diseases) and abiotic (water deficit, heat, flooding, and salinity) stresses exhibit wide-ranging losses in citrus production [10,11]. Cultivars of genus *Citrus* are grown in a climate of wide range due to evergreens and perennial tree crops [12]. The capacity to tolerate the unfavorable climatic conditions of the citrus crop is high but the estimated yield loss due to environmental abiotic stresses in citrus is up to 82% [13]. Climate change conditioning causes a global rise in temperature and limited the availability of freshwater for the crop. Meanwhile, crop consumption is increasing rapidly resulting in reduced levels of groundwater ultimately limiting productivity [14]. Global warming due to climate change caused frequent extreme water deficit conditions, an important component for global agriculture [15,16]. Abiotic stresses are interlinked with each other in which soil water deficit/drought is a serious environmental factor, which frequently limits the growth and productivity of important crop species [17]. Supply in water restriction can severely limit plant cell division, plant growth, fruit development, and fruit production [18,19]. Soil water deficit for long-term events causes permanent negative changes in the plant which can be in response to previous stress. Water deficit conditions can be developed at different phenological stages which can change physiological and molecular processes of the plant [20].

Plant responses to water stress are mediated by changes in root growth pattern and stomatal closure at moderate stress resulting in reduced CO2 intake, impairing photosynthesis, and loss in production [21,22]. Extreme water stress conditions alter the physiological and morphological processes of the plant. The molecular and biochemical machinery is also affected under stress conditions. Reactive oxygen species (ROS), i.e., hydrogen peroxide, superoxide, singlet oxygen, and hydroxyl radical are excessively produced and accumulate in plant cells at water deficit conditions. Photoinhibition or photooxidation caused by ROS accumulation leads to uncontrolled photosynthetic components oxidation [23,24]. Water deficit/drought damages photosynthetic apparatus and photosynthetic pigments (chlorophyll *a*, chlorophyll *b*, and carotenoids) which also reduced the photosynthetic rate [25,26]. Photosynthetic pigments are linked to stress tolerance in plants [27,28]. Resistant cultivars had a higher amount of chlorophyll contents and carotenoids when exposed to water deficit conditions [29]. Water deficit conditions decrease contents of total chlorophyll *a* and *b* contents [30]. Water deficit conditions damage thylakoid lamella and reduce active oxygen species' chlorophyll contents [31]. The chlorophyll content is linked inversely with the severity and duration of water deficit conditions [32]. After oxidative stress, accumulation of proline acts as an adaptive stress response. The intracellular levels of proline can increase by >100-fold during stress [33]. Proline accumulation acts as a compatible osmolyte to buffer cytosolic pH and to balance cell redox status. Proline can also function as a molecular chaperone stabilizing the structure of proteins and as a ROS scavenger [24,34].

Plant anatomical structure is also disturbed when the plant is subjected to water deficit which affects the regulation of water through the vascular bundles. The effect of drought stress firstly occurs on the cell structure [35,36]. Increased hydraulic resistance and decreased growth are directly associated with xylem structure [37]. Anatomical changes in citrus rootstocks under abiotic stress influence their ability to survive. In drought

conditions, the xylem vessel becomes emboli and dysfunctional. Therefore, plants with narrow and higher number of vessels are considered to be more drought tolerant [38]. The plant cells that face an environment with a shortage of water have generally shown an increase in vessel tissue, thick cell wall, reduction in cell size, and most severe condition cell wall and the cell membrane becomes ruptured [39,40]. The anatomical changes may occur to protect the plants under stress conditions. Multiple changes occur in response to water stress such as alteration of xylem phloem ratio, wall structure, lumen size, and lumen area that resist environmental stress on the plant [41]. The sensitivity of rootstock against drought stress is directly related to vessel dimension [42,43]. The vessel density of root and stem decreases with tree height as vessel diameter increases. The rootstock growth ability is dramatically affected by several xylem traits, xylem phloem ratio, vessel size, and vessel density [43,44]. Thus, traits play an important role in maintaining hydraulic conductance of root and stem and leaves [37,45]. Vessel number and size is the key factor to maintaining hydraulic conductance [46].

Strategies for drought tolerance are highly relevant in the case of rootstock selection and multiplication for ensuring continuous productivity. Rootstocks with a higher root growth ratio, better water use efficiency, higher root hydraulic conductivity, and lower osmatic conductance can withstand drought conditions while maintaining higher growth levels and mass accumulation [47,48]. After a severe drought, the recovery of tolerant rootstocks is much better than other rootstocks. Therefore, screening of drought-tolerant rootstocks is of utmost importance. For drought stress prevention, the plant increases water uptake efficiency either by increasing root density or deepening roots [26].

Climate changes shift the weather conditions by a high degree of unpredictability; water shortage in the soil and plant by a continuous increase in daily average temperature every year are inevitable, which intimidate overall globe agriculture industry stability by the negative effect on plant health and production consumer demand. Survival of the citrus industry against the water deficit needs to evaluate a tolerant/resistant citrus rootstock against the climate change scenario. The objective of the current study was to evaluate 10 citrus rootstocks against drought, based on visual changes, water potential, morphological and biochemical characters. After initial screening, the two most sensitive and tolerant genotypes were used to study the anatomical differences in leaf and stem to elucidate the drought tolerance mechanism in tolerant genotypes.

#### **2. Materials and Methods**

#### *2.1. Plant Materials, Experimental Site, and Growth Conditions*

Six months old citrus rootstocks potted plants were taken as experimental material. Ten genetically diverse citrus rootstocks were examined (details are given in Table 1). A potted plant experiment was executed at the Institute of Horticultural Science, University of Agriculture Faisalabad, Pakistan. Seeds of selected rootstock were taken from citrus rootstocks progeny block, HIS, UAF, Pakistan.

**Table 1.** Characteristics of citrus rootstocks used in this study.

**Table 1.** *Cont.*

The seeds were sown in transparent grow bags (height 12- and width 6--) and placed in a growth chamber (Model: BST/PGC-175; Bionics Scientific Technologies (p) ltd., Delhi, India) at 32 ± 2 ◦C, relative humidity oscillating between 80 and 95% and 12–14 h of light. Potted media contained sand, silt, and clay (1:1:1). These plants were placed in the chamber for six months before the treatment application and during growth, pots were regularly irrigated with tap water (75% field capacity) and fertilized weekly with nutrient solution (1.0 g L−1 Ca(NO3)2, 0.4 g L−1 KNO3, 0.6 g L−1 MgSO4 and 0.4 g L−1 NH4H2PO4 (MAP)).

#### *2.2. Water Regimes and Treatments*

Citrus rootstocks were subjected to water deficit treatment applications after six months of growth in the controlled growth condition. Plants were exposed to three different groups: controlled condition, moderate water deficit (moderate drought), and severe water deficit (severe drought). Each treatment group consists of 10 citrus rootstocks with three replications. Control plants (sufficient water and optimal temperature ~32 ◦C) were irrigated once every two days (75% field capacity). Water deficit treatments were moderate (50% field capacity) and severe (25% field capacity) deficit conditions. Water was applied with 2 days interval for control and treated plants. The controlled and water deficit exposed genotypes were kept at 32 ± 2 ◦C for day and night temperature in a controlled growth room. Thereby, three experimental groups were established as control, moderate water deficit, and severe water deficit conditions.

#### *2.3. Morphological and Biomass Measurements*

Leaves were visually observed for leaf necrosis or chlorosis on rootstock seedlings after 15 days of stress applications. Plant height (cm) and root length (cm) were measured with help of a scale after uprooting the plants at the end of the experiment (15 DAS). Leaf water potential was determined to collect the healthy leaf samples at dawn (Shafqat et al., 2021) by using a pressure chamber (Model 3000, Soil moisture Equipment, Santa Barbara, CA, USA). Root and shoot fresh biomass were weighed on electric balance (TS-200 Digital Electronic Scale), oven-dried (70 ◦C) for 48 h, and weighed again separately [48]. Shoot and root moisture content were calculated using Equation (1):

$$\text{Moisture content} \left( \% \right) = \frac{\text{Fresh weight} - \text{dry weight}}{\text{Fresh weight}} \times 100 \tag{1}$$

#### *2.4. Biochemical and Stress-Associated Biomarker Measurements* 2.4.1. Leaf Photosynthetic Pigments Contents

Leaf sample of 0.5 g was cut into small pieces and homogenized by adding 80% acetone (v/v). The extract was transferred to a 15 mL tube [48]. Test tubes were placed in the dark to avoid light for 24 h and filtered through filter paper. The absorbance was determined at 663 nm for chlorophyll a, 647 nm for chlorophyll b, and 470 nm for carotenoid. The Chl *a*, Chl *b*, and carotenoids contents were determined using Equations (2)–(4), respectively.

$$\text{Chl } a \text{ (mg/g fresh weight)} = \frac{(12.7 \text{OD663} - 2.69 \text{ OD645}) \times \text{V}}{1000 \times \text{W}} \tag{2}$$

$$\text{Chl } b \text{ (mg/g fresh weight)} = \frac{(22.9 \text{OD645} - 4.68 \text{ OD6663}) \times \text{V}}{1000 \times \text{W}} \tag{3}$$

$$\text{Carotenoids } (\text{mg/g fresh weight}) = \frac{\text{OD } 480 + 0.114 \text{ (OD } 663) - 0.638 \text{ (OD } 645)}{\text{Em} \times 100} \quad (4)$$

where V = volume of the sample, W = weight of fresh tissue, Em = 2500.

#### 2.4.2. Determination of Proline

Proline contents were assessed by following the acid ninhydrin method [49]. Fresh 0.5 g leaf material was extracted using 10 mL of 3% sulfosalicylic acid (Panreac, Barcelona, Spain) for 30 min. Centrifugation at 2000 *g* for 20 min at 4 ◦C was done. The 2 mL of filtered aqueous extract was mixed with acid ninhydrin reagent (2 mL), and glacial acetic acid (2 mL), and heated at 100 ◦C for 1 h. The reaction mixture after cooling was segregated against toluene (4 mL) and the absorbance of the organic phase was recorded at 520 nm using a spectrophotometer (IRMECO UV/VIS Model U2020). The resulting values were related with a standard curve plotted using known amounts of proline (Sigma, St Louis, MO, USA).

#### 2.4.3. Determination of Hydrogen Peroxide (H2O2)

The leaf tissues of 0.1 g were ground in 1 mL of trichloroacetic acid solution (0.1%) within an ice bath [50]. After preparation in Eppendorf, the samples were centrifuged at 9719× *g* for 15 min. The supernatant of 500 μL was transferred into a new tube having a mixture of 1 M KI (1000 μL) and 10 mM potassium phosphate buffer (500 μL). Absorbance was read at 390 nm in a UV-1900 spectrophotometer (Eppendorf BioSpectrometer® basic). H2O2 content was calculated as μmol g−1 DW by comparing the absorbance values against the standard curve of commercial H2O2.

#### *2.5. Stem and Leaf Anatomical Evaluation*

#### 2.5.1. Plant Material and Experiment

Two highly tolerant (Brazilian sour orange and Gadha dahi) and highly sensitive (Rangpur Poona nucellar and Sunki × bentake) genotypes from the first screening study were selected to screen based on the leaf and stem anatomical study. These selected rootstocks were exposed to the same treatments for 15 days then leaf and stem samples were collected for anatomical studies.

#### 2.5.2. Preservation, Sectioning, Staining, and Mounting

Stem and leaf samples (2 cm) were preserved in a formalin acetic alcohol (FAA) solution containing 5% formalin, 10% acetic acid, 50% ethanol, and 35% distilled water. Thereafter, the preserved material was subsequently transferred to an acetic alcohol solution (acetic acid 25% v/v, ethanol 75%) for the long-term preservation of samples. A free-hand sectioning technique was used to prepare permanent slides of stem and leaf transverse sections cut with the razor blade, and some fine sections were carefully picked up on wash glass for staining and dehydration through a series of washings with ethanol (30%, 50%, and then 70% for 15 min each). For staining, the lignified tissues (xylem vessels and sclerenchyma) were transferred to safranin (1.0 g dissolved in 100 mL, 70% alcohol) for 20 min, dehydrated in 90% alcohol for 5 min, and stained with fast green (1.0 g dissolved in 90% ethanol) for one minute. Finally, the tissues were washed three times with absolute alcohol and then transferred to xylene for cleaning the contrast. The sections were mounted in Canada balsam by putting a drop of resin on a slide and placing the sections on the slides and photographed with a digital camera attached to a compound microscope.

#### 2.5.3. Anatomical Traits

The stem and leaf cross-sectional area (mm2) was measured under a compound light microscope (Olympus MX63, Japan) by recording the maximum length and width of the root sections. Cells present in plants epidermis were measured with the help of ruler

or cm scales, and the length of the epidermal cell was multiplied by 41.5 to obtain a value in micrometers. Vascular bundle length was calculated by measuring xylem and phloem which were multiplied by 41.5 to obtain the value in μm. Xylem thickness was calculated by the width of the xylem region. Metaxylem area, phloem cell area, pith cell area, and the cortical area were calculated in μm2 by measuring the length and width of each trait. Cortical thickness was obtained by measuring the total width of the cortical region in micrometers. By using Equation (5), the areas of the different cells and tissues were calculated (which was modified from the area of the circle).

$$\text{Area} = \frac{\text{Maximum length} \times \text{Maximum width}}{28} \times 22 \tag{5}$$

#### *2.6. Statistical Analysis*

Throughout this study, all experiments were laid out using a full factorial split-plot design arranged in randomized complete blocks using rootstocks as main plots and water regimes in the subplots. All experiments were carried out using at least three biological replicates for each treatment. The analysis of variance (ANOVA) was used to test the significant differences among different drought levels (*p*Drought), rootstocks (*p*Rootstocks), and their interaction (*p*Drought × rootstocks). Tukey's honestly significant difference (HSD) test was used for post-hoc analysis (*p*Drought × rootstocks < 0.05). Moreover, the data matrix of all dependent variables was used to perform the principal component analysis (PCA). Finally, means of all dependent variables were used for two-way hierarchical cluster analysis (HCA). Similarities and variations between treatments were presented as a heat map.

#### **3. Results**

#### *3.1. Drought Negatively Affects the Roots and Shoots Length*

Both root and shoot length were significantly affected by varying drought levels (*p*Drought < 0.0001) and rootstocks (*p*Rootstock < 0.0001) (Table 2). Briefly, the root length of different citrus rootstocks was significantly reduced when plants were stressed with different levels of drought. Although control Brazilian sour orange had the highest root length (27.67 ± 0.58 cm; *p*Drought × Rootstock < 0.0001), shortest root lengths were recorded by Sunki × bentake when stressed with moderate (10.33 ± 0.58 cm) or severe drought (9.33 ± 0.58 cm) and Rangpur Poona nucellar rootstock under severe drought conditions (10.33 ± 0.58 cm) (Table 2). On the other hand, while there were no significant differences in shoot length of all tested rootstock, except Gabbuchini, under normal growth conditions, shoot lengths were significantly reduced under various drought levels. Rangpur Poona nucellar rootstock had the shortest shoots (5.33 ± 0.58 cm), followed by Sunki × bentake (6.00 ± 1.00 cm) when stressed under severe drought (*p*Drought × Rootstock < 0.0001) (Table 2).

#### *3.2. Drought Stress Disrupts the Water Relations of Citrus Rootstocks*

As strong wilt phenotype was visually observed on citrus seedlings after 15 days post applications (DPA; Figure 1), the water relations (water potential, root moisture content, and shoot moisture content) were assessed (Table 2). For instance, water potential was reduced significantly by water-deficient (*p*Drought < 0.0001) and rootstocks (*p*Rootstock = 0.0018). Kirrumakki nucellar rootstock had the highest water potential (−0.11 ± 0.66 Mpa), whereas, Keen sour orange, Rough lemon, Brazilian sour orange, Sunki × bentake, X639, Kirrumakki nucellar, and Rangpur Poona nucellar rootstocks had the lowest water potential when severely stressed with water deficiency, without significant differences between them (*p*Drought × Rootstock = 0.0250; Table 2). Likewise, the moisture content of both root and shoot was significantly reduced by drought levels (*p*Drought < 0.0001) and rootstocks (*p*Rootstock < 0.0001) (Table 2). Normal irrigated Brazilian sour orange had the highest moisture content of root (75.67 ± 0.59%) and shoot (84.75 ± 0.65%), whereas Rangpur Poona nucellar rootstock had the lowest moisture content of root (36.33 ± 0.58%) and shoot (40.69 ± 0.65%) under severe drought conditions (Table 2).

**Table 2.** Effect of different water regimes on the root length (cm), shoot length (cm), and water relations of 10 differentcitrus rootstocks with distinct degrees of tolerance to drought stress.

Data presented are means ± standard deviation (mean ± SD) of three replicates. Different letters indicate statistically significant differences among treatments, while "ns" signifies no significant differences between them according to Tukey's honestly significant difference test (*p* < 0.05).

**Figure 1.** Effect of different water regimes on the leaf phenotype of 10 different citrus rootstocks with distinct degrees of tolerance to drought stress.

#### *3.3. Water Deficiency Interrupts the Photosynthetic Pigments of Citrus Rootstocks*

Drought stress considerably lessened chlorophyll *a,* chlorophyll *b,* and carotenoids content of different rootstocks (*p*Drought < 0.0001; Table 3). Normal irrigated Brazilian sour orange had the highest chlorophyll *a* content (3.74 ± 0.12 mg g−1 FW), while Rangpur Poona nucellar rootstock had the lowest chlorophyll *a* content (1.57 ± 0.04% mg g−1 FW; *p*Drought × Rootstock < 0.0001). Furthermore, there was no significant difference in chlorophyll *b* content among all tested rootstocks, except Gabbuchini and Gada dahi, under normal irrigation conditions. However, chlorophyll *b* content was significantly decreased under water deficiency conditions (Table 3). Likewise, there were no significant differences in carotenoids content of all studied citrus rootstocks, except Gabbuchini, under normal irrigation conditions. Nevertheless, carotenoids content was significantly reduced when citrus rootstocks were stressed with different drought levels. It is worth mentioning that Rangpur Poona nucellar rootstock had the lowest carotenoids content (0.14 ± 0.03 mg g−1 FW), followed by Sunki × bentake (0.23 ± 0.02 mg g−1 FW) under severe drought conditions (Table 3).

**Table 3.** Effect of different water regimes on the photosynthetic pigments, H2O2, and proline content of 10 different citrus rootstocks with distinct degrees of tolerance to drought stress.

Data presented are means ± standard deviation (mean ± SD) of three replicates. Different letters indicate statistically significant differences among treatments, while "ns" signifies no significant differences between them according to Tukey's honestly significant difference test (*p* < 0.05).

#### *3.4. Drought Stress Induced the Accumulation of Stress-Associated Biomarkers in Citrus Rootstocks*

Two major stress-associated biomarkers, including H2O2 and endogenous proline content, were assessed (Table 3). Generally, both drought levels (*p*Drought < 0.0001) and rootstocks (*p*Rootstock = 0.0261) induced the H2O2 accumulation. Plants under drought

stress conditions (moderate or severe) had significantly higher H2O2 levels, compared with non-stressed rootstocks. It is worth mentioning that all tested rootstocks had their highest H2O2 levels under severe drought conditions with no noticeable difference between them (Table 3). Similarly, endogenous proline content was significantly affected by waterdeficient (*p*Drought < 0.0001) and citrus rootstock (*p*Rootstock < 0.0001), and its levels were boosted with raising the severity of drought stress. Brazilian sour orange had the highest proline levels (0.51 ± 0.06 μmol g−1 FW) under severe drought conditions. Nevertheless, there was no significant difference in proline content of different citrus rootstocks under regular irrigation conditions (Table 3).

#### *3.5. Principal Component Analysis (PCA) and Two-Way Hierarchical Cluster Analysis (HCA) Revealed the Differences among Water Treatments and Citrus Rootstocks*

To better understand the water deficiency and citrus rootstocks interactions, principal component analysis (PCA) and two-way hierarchical cluster analysis (HCA) were carried out (Figure 2). PCA-associated scatterplot revealed a clear separation among water deficit treatment (control, moderate drought, and severe drought), as well as all studied rootstocks with respect to PC1 (approximately 81.28%) and PC2 (about 11.94%) (Figure 2A). Moreover, the PCA-associated loading plot showed that while root length, root moisture content, shoot moisture content, shoot length, carotenoids, chlorophyll a, chlorophyll b, and water potential were positively correlated with non-stressed control plants, H2O2, and proline content were positively associated with water deficit treatments (Figure 2B).

In harmony with PCA findings, the HCA and its associated heatmap revealed the differences among water-deficient treatments (Figure 2C). For example, HCA-associated dendrogram among rootstocks revealed that all studied rootstocks were clustered separately into three distinct clusters. Cluster (a) included all non-stressed rootstocks, Cluster (b) included all moderate drought-stressed rootstocks except Rangpur Poona nucellar and Sunki × bentake which were clustered with severely stressed rootstocks within Cluster (c). Additionally, the HCA-associated dendrogram among investigated variables revealed that they were clustered into two separate clusters. Cluster 'I' included root length, root moisture content, shoot moisture content, shoot length, carotenoids, chlorophyll a, chlorophyll b, and water potential which all were higher in regularly irrigated non-stressed control rootstocks. On the other hand, Cluster 'II' included only H2O2 and proline contents which were higher in citrus rootstocks grown under severe deficient water stress (Figure 2C).

#### *3.6. Water Deficiency Alters the Anatomical Structure of Citrus Rootstocks*

To better understand the mechanism of drought resistance in citrus, the effect of different drought levels on the anatomical structure of stem and leaves of two highly tolerant (Brazilian sour orange and Gadha dahi) and two highly sensitive genotypes (Rangpur Poona nucellar and Sunki × bentake) from the first screening study was investigated. As we mentioned above, under drought stress conditions, the moisture content of both roots and shoots was reduced, resulting in yellow, curled, wilted leaves, and some other adverse wilt-associated symptoms (Figure 1). Briefly, under severe drought stress, the length of epidermal cells and vascular bundles was reduced, as well as the thickness of the xylem and cortical was thinner.

#### 3.6.1. Effect of Drought Stress on Stem Anatomy of Citrus Rootstocks

Microscopic observation of stem cross-section showed that the highly tolerant genotypes (Brazilian sour orange and Gadha dahi) had some features such as water-filled cells, small cell gaps, tight and round cells under normal water conditions, as well as under drought stress conditions (Figure 3).

**Figure 2.** Principal component analysis (PCA) and two-way hierarchical cluster analysis (HCA) of individual morphological and physiological parameters were assessed in 10citrus rootstocks with distinct degrees of tolerance to drought stress under three water regimes. (**A**) PCA-associated scatterplots, (**B**) PCA-associated loading plots, and (**C**) two-way HCA. Variations in the dependent variables among studied treatments are visualized as a heat map. Rows correspond to dependent variables, whereas columns correspond to different treatments. Low numerical values are lightyellow-colored, while high numerical values are colored dark red (see the scale at the right bottom corner of the heat map).

However, both drought levels and rootstocks (*p*Drought < 0.0001 and *p*Rootstock = 0.0002, respectively) significantly affected the length of the epidermal cell and vascular bundle (Figure 4A,B, respectively), xylem thickness (Figure 4C), area of metaxylem cell (Figure 4D), phloem cell (Figure 4E), pith cell (Figure 4F), pith thickness (Figure 4G), and cortical cell (Figure 4H), and cortical thickness (Figure 4I). Under all water conditions, the anatomical structures of the highly tolerant genotypes Brazilian sour orange and Gadha dahi performed better than two highly sensitive genotypes Rangpur Poona nucellar and Sunki × bentake. For instance, under regular irrigation, both Brazilian sour orange and Gadha dahi rootstocks had the highest epidermal cell length (*p*Drought × Rootstock = 0.0255), vascular bundle length (*p*Drought × Rootstock < 0.0001), xylem thickness (*p*Drought × Rootstock = 0.488), metaxylem cell area (*p*Drought × Rootstock = 0.0140), phloem cell area (*p*Drought × Rootstock = 0.0346), pith cell area (*p*Drought × Rootstock = 0.0316), pith thickness area (*p*Drought × Rootstock = 0.0112), cortical cell area (*p*Drought × Rootstock = 0.0180), and cortical thickness (*p*Drought × Rootstock = 0.0319) with no significant differences between them. Although, both moderate and severe drought

stresses significantly reduced most, if not all, of these anatomical attributes, both highly tolerant genotypes did not show noticeable changes under drought stress. Nevertheless, the mesophyll cells of drought-sensitive Rangpur Poona nucellar and Sunki × bentake genotypes were slightly deformed and had shorter epidermal cell and vascular bundle length, as well as narrower metaxylem, phloem, pith, and cortical area (Figure 4).

**Figure 3.** Stem transverse section of two highly tolerant (Brazilian sour orange and Gadha dahi) and two highly sensitive genotypes (Rangpur Poona nucellar and Sunki × bentake). PT: pith; CR: cortex; XL: xylem; PL: phloem.

3.6.2. PCA and Two-Way HCA Divulged the Variations in Stem Anatomy of Different Citrus Rootstocks

Briefly, the PCA-associated scatter plot showed a clear separation among water deficit treatment (control, moderate drought, and severe drought), as well as all studied rootstocks with respect to PC1 (approximately 87.29%) and PC2 (about 8.41%) (Figure 5A). It is worth mentioning that the drought-tolerant rootstocks Brazilian sour orange and Gadha dahi were grouped close to each other and separately from the two sensitive genotypes Rangpur Poona nucellar and Sunki × bentake under normal irrigation and moderate drought, but not severe drought conditions. Furthermore, the PCA-associated loading plot showed that all studied stem anatomical features were positively associated with normal water application (Figure 5B). In agreement with PCA results, the HCA and its associated heatmap uncovered the differences among different rootstocks under water-deficient treatments (Figure 5C). For example, HCA-associated dendrogram among rootstocks revealed that all studied rootstocks separated into two distinct clusters. Cluster (a) included all non-stressed rootstocks and the highly tolerant (Brazilian sour orange and Gadha dahi), grown under moderate drought, whereas cluster (b) included all severely stressed rootstocks and the two highly sensitive genotypes (Rangpur Poona nucellar and Sunki × bentake) that were moderately stressed with drought (Figure 5C).

**Figure 4.** Effect of different water regimes on the stem anatomical features of two highly tolerant (Brazilian sour orange and Gadha dahi) and two highly sensitive genotypes (Rangpur Poona nucellar and Sunki × bentake). (**A**) Epidermal cell length (μm), (**B**) Vascular bundle length (μm), (**C**) Xylem thickness (μm), (**D**) Metaxylem cell area (μm2), (**E**) Phloem cell area (μm2), (**F**) Pith cell area (μm2), (**G**) Pith thickness area (μm2 × 103), (**H**) Cortical cell area (μm2), and (**I**) Cortical thickness (μm). Data presented are means ± standard deviation (mean ± SD) of three biological replicates. Different letters indicate statistically significant differences among treatments, while "ns" signifies no significant differences between them according to Tukey's honestly significant difference test (*p* < 0.05).

**Figure 5.** Principal component analysis (PCA) and two-way hierarchical cluster analysis (HCA) of individual stem anatomical features of two highly tolerant (Brazilian sour orange and Gadha dahi) and two highly sensitive genotypes (Rangpur Poona nucellar and Sunki × bentake) under three water regimes. (**A**) PCA-associated scatter plots, (**B**) PCAassociated loading plots, and (**C**) two-way HCA. Variations in the dependent variables among studied treatments are visualized as a heat map. Rows correspond to dependent variables, whereas columns correspond to different treatments. Low numerical values are light-yellow-colored, while high numerical values are colored dark red (see the scale at the right bottom corner of the heat map).

3.6.3. Effect of Drought Stress on Leaf Tissue Structure of Citrus Rootstocks

Microscopic observation of citrus leaves cross-section showed that it had an asymmetric heterogeneous structure that was characterized by two unequal palisade parenchyma (Figure 6).

**Figure 6.** Leaf transverse section of two highly tolerant (Brazilian sour orange and Gadha dahi) and two highly sensitive genotypes (Rangpur Poona nucellar and Sunki × bentake). PT: pith; CR: cortex; XL: xylem; PL: phloem; PP: palisade parenchyma.

Like stem anatomy, citrus leaf anatomical attributes were significantly altered by both drought levels and rootstocks. However, the anatomical changes in leaf tissue structure were less significant in highly tolerant rootstocks (Brazilian sour orange and Gadha dahi) than sensitive genotypes (Rangpur Poona nucellar and Sunki × bentake). Interestingly, differences in all studied anatomical features of citrus leaves under different water regimes were cultivar-dependent. These features included epidermal cell length (*p*Drought × Rootstock = 0.0315; Figure 7A), vascular bundle length (*p*Drought × Rootstock = 0.0009; Figure 7B), xylem thickness (*p*Drought × Rootstock = 0.0251; Figure 7C), metaxylem cell area (*p*Drought × Rootstock = 0.0349; Figure 7D), phloem cell area (*p*Drought × Rootstock = 0.0197; Figure 7E), pith cell area (*p*Drought × Rootstock = 0.0163; Figure 7F), pith thickness area (*p*Drought × Rootstock = 0.0335; Figure 7G), cortical cell area (*p*Drought × Rootstock = 0.0250; Figure 7H), and cortical thickness (*p*Drought × Rootstock = 0.0357; Figure 7I). Under all tested water regimes, highly tolerant rootstocks Brazilian sour orange and Gadha dahi had thicker epidermal and vascular bundle, as well as wider pith and cortical areas compared with sensitive genotypes (Rangpur Poona nucellar and Sunki × bentake). Additionally, severe drought stress significantly reduced the thickness of all leaf tissues, particularly in sensitive genotypes (Figure 7).

3.6.4. PCA and Two-Way HCA Revealed the Differences in Leaf Tissue Structure of Different Citrus Rootstocks

In brief, the PCA-associated scatter plot showed a clear separation among water deficit treatment (control, moderate drought, and severe drought), as well as all studied rootstocks with respect to PC1 (approximately 93.24%) and PC2 (about 3.95%) (Figure 8A). It is worth mentioning that the drought-tolerant rootstocks Brazilian sour orange and Gadha dahi were grouped close to each other and separately from the two sensitive genotypes Rangpur Poona nucellar and Sunki × bentake under all investigated water regimes. Moreover, the PCA-associated loading plot showed that all studied anatomical features of citrus leaves were positively associated with normal water application (Figure 8B). Like PCA, the HCA and its associated heatmap revealed the differences among different rootstocks under water-deficient treatments (Figure 8C). For example, HCA-associated dendrogram among rootstocks revealed that all studied rootstocks separated into two distinct clusters. Cluster (a) included all regularly irrigated genotypes and the two highly tolerant rootstocks (Brazilian sour orange and Gadha dahi) that were grown under moderate drought. On the other hand, cluster (b) included all severely stressed genotypes and the two sensitive

rootstocks (Rangpur Poona nucellar and Sunki × bentake) that were moderately stressed with drought (Figure 8C).

**Figure 7.** Effect of different water regimes on the leaf anatomical features of different two highly tolerant (Brazilian sour orange and Gadha dahi) and two highly sensitive genotypes (Rangpur Poona nucellar and Sunki × bentake). (**A**) Epidermal cell length (μm), (**B**) Vascular bundle length (μm), (**C**) Xylem thickness (μm), (**D**) Metaxylem cell area (μm2), (**E**) Phloem cell area (μm2), (**F**) Pith cell area (μm2), (**G**) Pith thickness area (μm2 × 103), (**H**) Cortical cell area (μm2), and (**I**) Cortical thickness (μm). Data presented are means ± standard deviation (mean ± SD) of three biological replicates. Different letters indicate statistically significant differences among treatments, while "ns" signifies no significant differences between them according to Tukey's honestly significant difference test (*p* < 0.05).

**Figure 8.** Principal component analysis (PCA) and two-way hierarchical cluster analysis (HCA) of individual leaf anatomical features of two highly tolerant (Brazilian sour orange and Gadha dahi) and two highly sensitive genotypes (Rangpur Poona nucellar and Sunki × bentake) under three water regimes. (**A**) PCA-associated scatter plots, (**B**) PCA-associated loading plots, and (**C**) two-way HCA. Variations in the dependent variables among studied treatments are visualized as a heat map. Rows correspond to dependent variables, whereas columns correspond to different treatments. Low numerical values are light-yellow-colored, while high numerical values are colored dark red (see the scale at the right bottom corner of the heat map).

#### **4. Discussion**

Water deficit conditions are a major environmental factor, which frequently limits the growth and productivity of important crop species [17]. Restriction of water supply can severely limit plant growth, development, and production [18,19]. Choice of rootstock is among the most important decisions a grower makes, and implications for yield and quality are enormous. Rootstock in citrus trees influences the morphological, biochemical, physiological, and genetic characteristics of grafted scion cultivars through the rootstock scion interaction pathway [51]. Citrus rootstocks with better drought tolerance ability can greatly reduce production losses [52]. In this study, plant material consisted of 10 genetically diverse citrus rootstocks belonging to different citrus categories, i.e., oranges, pummelo, lemon, lime, their hybrids, and originating from diverse localities. The leaf shape and size of these rootstocks also varied. These rootstocks are reported to have different tolerance towards some biotic and abiotic stresses. The drought tolerance of these rootstocks was studied in this investigation. Leaf water potential in plants is directly related to water availability [53]. Leaf water potential indicates the whole plant water status, and maintenance of high leaf water potential is found to be associated with dehydration avoidance mechanisms. Our results demonstrated the decrease in leaf water potential as drought conditions become severe compared with control. The maintenance of water potential in leaves is a direct indicator of a plant dehydration avoidance mechanism, as genotypes with better water potential at stress conditions are regarded as drought tolerant [23,54,55]. In our studies, Brazilian sour orange showed drought tolerance by performing best at severe drought and Rangpur Poona nucellar at moderate drought. For insensitive genotypes, the decrease in leaf water potential indicated the mechanical injury of leaf chloroplasts caused by stress conditions which result in reduced transpiration rate and oxidative stress [16].

Citrus rootstocks with high chlorophyll *a* and *b*, and carotenoid contents against the stresses, especially the water stress, are regarded as tolerant rootstocks [56]. The normal functioning of photosynthetic machinery is affected by drought stress, the degradation, and photo-oxidation of chlorophyll caused by transpirational imbalance at water stress hamper the plant's ability to harvest light reducing total photosynthetic output [21,57]. Results showed that photosynthetic pigments chlorophyll *a*, chlorophyll *b,* and carotenoids reduced significantly at elevated stress conditions and overall genotype Brazilian sour orange and Gada dahi had the highest chlorophyll contents at drought conditions highlighting their ability to tolerate drought stress. Plants with dark green leaves (chlorophyll) under drought stress are considered tolerant. Visual assessments indicated the Brazilian sour orange as a tolerant rootstock without changes in leaf green color and leaf necrosis; while, Savage citrange emerged as the most sensitive, with maximum plant death and leaf shedding during stress treatments. The compromised photosynthetic machinery reduces carbohydrate transport and as a result plant growth is also reduced. The ability of plants to maintain growth under limited water supply reveals their tolerance ability [43,58]. Results showed that among citrus rootstocks, Brazilian sour orange and Gada dahi at water stress conditions maintained steady root and shoot growth. The root and shoot moisture content of these rootstocks were also high at stress conditions showing their tolerant nature. While, Rangpur Poona nucellar had the lowest shoot and root growth and moisture content emerged as the most sensitive.

Metabolic imbalances triggered by drought stress cause oxidative stress and as a result ROS are produced and accumulated [59]. The increased oxidation greatly reduces metabolic activities and the normal functioning of cell organelles.  Proline is an osmoprotectant that is trigged as a result of ROS production in the cell, its production and accumulation work as ROS scavenging, redox balance, and reduce cell damage which normalizes the functionality of plant cells [60]. In drought-tolerant genotypes, the ROS production is reduced, and proline concentration is increased with increasing severity of stress. The results show that Brazilian sour orange has minimal ROS production and the highest proline concentration at severe drought stress.

In citrus under drought stress, leaves are observed to be shorter with thick epidermal cells which facilitates reducing the transpirational rate and oxidative stress [61,62]. Modifications in vascular anatomy are important for plant acclimation potential. Vascular bundles present in mid rid of leaves serve as a source to distribute nutrients and water. At

stress conditions with reduced leaf size, the reduction in the vascular bundle is an indicator of plants' abilities to modify their anatomy under stress. In the vascular bundle, the xylem acts as a source of water transport. The plants with greater xylem vessel diameter are unable to survive harsh environmental conditions [63–65]. At the onset of drought, stress transpiration, water uptake from roots, and stem hydraulic capacitance begin to decline. That reduces growth, the vascular bundles in the stem are observed to be reduced along with pith cell area and cortical thickness [66,67]. In this study, the results showed that with increasing drought intensity the anatomy of both sensitive and tolerant genotypes was modified, interestingly the two tolerant genotypes Brazilian sour orange and Gada dahi had greater values for all the leaf and stem anatomy parameters at severe drought stress than sensitive genotypes. This could be because of the continuous adaptability of tolerant genotypes which enabled them to maintain growth and function as the amount of water became limited, in response to sensitive genotypes in which the response could have been triggered at very later stages which abruptly affect their growth. These modifications in tolerant genotypes enabled them to maintain steady nutrient transport while reducing the risk of embolisms, increasing water-flow resistance, and constant transport of nutrients across [64].

#### **5. Conclusions**

Drought stress adversely affected plant water status, photosynthetic machinery, biochemical balance, and anatomical structure of all the citrus rootstocks studied. The intensifying drought reduced leaf water potential, and compromised the photosynthetic apparatus by damaging photosynthetic pigments (chlorophyll "*a*", "*b*", and carotenoid) apparent from lighter green color. Oxidative stress caused by ROS production which triggered production of proline. Alteration in anatomical structures of leaf and stem were observed. Citrus rootstocks Brazilian sour orange and Gada dahi performed best under drought stress, mitigated damage at molecular biochemical and anatomical levels, while rootstocks Sunki × bentake and Rangpur Poona nucellar were the most sensitive rootstocks.

**Author Contributions:** Conceptualization, M.J.J., S.A.N., M.H. and Y.N., methodology, W.S., M.J.J., Y.S.A.M. and M.H.; software, W.S., S.A.N., Y.N. and S.I.; formal analysis, Y.S.A.M., W.S., S.-u.-R. and Y.N.; resources, M.J.J., M.H., S.A.N. and Y.S.A.M.; data curation, S.-u.-R., S.B., S.I. and W.S.; writing—original draft preparation, W.S., S.I., S.-u.-R. and Y.N.; writing—review and editing, M.J.J., Y.S.A.M., S.A.N., Y.N. and M.H.; supervision, M.J.J. All authors have read and agreed to the published version of the manuscript.

**Funding:** We would like to acknowledge the Higher Education Commission, Islamabad and Endowment Fund Secretariat, University of Agriculture, Faisalabad, Pakistan for providing funding. The authors extend their appreciation to the Deanship of Scientific Research at King Khalid University for funding this work through the Program of Research Groups under grant number (RGP 2/165/42).

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

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** The data that supports the findings of this study are contained within the article and available from the corresponding author upon reasonable request.

**Acknowledgments:** Y.N. would like to extend his appreciation to the Graduate Student and Research Affairs Sector of Tanta University, Egypt. We also thank all staff members of our laboratories for their helpful discussions and comments. The authors extend their appreciation to the Deanship of Scientific Research at King Khalid University for funding this work through the Program of Research Groups under grant number (RGP 2/165/42).

**Conflicts of Interest:** The authors declare that there is no conflict of interest, and they have no known competing financial interests or personal relationships that could have appeared to influence the work reported in this paper.

#### **References**

## *Article* **Physiological and Biochemical Characterization of the GABA Shunt Pathway in Pea (***Pisum sativum* **L.) Seedlings under Drought Stress**

**Nisreen A. AL-Quraan 1,\*, Zakaria I. Al-Ajlouni 2 and Nima F. Qawasma 1**

**Abstract:** The physiological and biochemical role of the γ-aminobutyric acid (GABA) shunt pathway in green pea seedlings (*Pisum sativum* L.) was studied in response to soil water holding capacity levels: 80%, 60%, 40%, 20%, and 10% grown under continuous light at 25 ◦C for 7 days and 14 days, separately. Characterization of seeds germination pattern, seedlings growth (plant height, fresh and dry weight, and chlorophyll contents), GABA shunt metabolite (GABA, glutamate, and alanine) levels, total protein and carbohydrate levels, and oxidative damage (MDA level) were examined. Data showed a significant effect of drought stress on seed germination, plant growth, GABA shunt metabolites level, total protein and carbohydrate contents, and MDA level. A significant decline in seed germination percentage was recorded at a 20% drought level, which indicated that 20% of soil water holding capacity is the threshold value of water availability for normal germination after 14 days. Seedling fresh weight, dry weight, and plant height were significantly reduced with a positive correlation as water availability was decreased. There was a significant decrease with a positive correlation in Chl *a* and Chl *b* contents in response to 7 days and 14 days of drought. GABA shunt metabolites were significantly increased with a negative correlation as water availability decreased. Pea seedlings showed a significant increase in protein content as drought stress was increased. Total carbohydrate levels increased significantly when the amount of water availability decreased. MDA content increased slightly but significantly after 7 days and sharply after 14 days under all water stress levels. The maximum increase in MDA content was observed at 20% and 10% water levels. Overall, the significant increases in GABA, protein and carbohydrate contents were to cope with the physiological impact of drought stress on *Pisum sativum* L. seedlings by maintaining cellular osmotic adjustment, protecting plants from oxidative stress, balancing carbon and nitrogen (C:N) metabolism, and maintaining cell metabolic homeostasis and cell turgor. The results presented in this study indicated that severe (less than 40% water content of the holding capacity) and long-term drought stress should be avoided during the germination stage to ensure proper seedling growth and metabolism in *Pisum sativum* L.

**Keywords:** drought stress; GABA: gamma aminobutyric acid; metabolism; pea; *Pisum sativum* L.; seedling growth; water deficit

#### **1. Introduction**

Pea (*Pisum sativum* L.) was used by Mendel to lay the foundation of modern genetics [1]. It is one of the major food legumes that can grow in different regions and rich in proteins, vitamins, minerals, carbohydrates, and seed oil [2]. Pea is predominantly a self-pollinated crop with limited variation in the number of flowers per node [3]. Most garden pea germplasm/varieties lines have either one or two flowers per node [4]. The

**Citation:** AL-Quraan, N.A.; Al-Ajlouni, Z.I.; Qawasma, N.F. Physiological and Biochemical Characterization of the GABA Shunt Pathway in Pea (*Pisum sativum* L.) Seedlings under Drought Stress. *Horticulturae* **2021**, *7*, 125. https:// doi.org/10.3390/horticulturae7060125

Academic Editor: Yanyou Wu

Received: 24 April 2021 Accepted: 25 May 2021 Published: 27 May 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/).

agro-ecological importance of pea is connected with its capability to form symbiotic nitrogen fixation with rhizobial bacteria that promotes and enhances seed germination, seedling vigor and emergence, root and shoot growth, plant stand and biomass, and seed weight [5].

Drought is a major abiotic stress that affects plant growth and yield [6]. Plant species and genotype adaptation and level of tolerance vary according to the type of drought stress (meteorological, agricultural, hydrological, or socioeconomic drought) [7]. Drought limits the productivity of many crops, especially during the seedling stage [8]. Water scarcity negatively affects vegetative growth by direct influence on its metabolic sink strength and fruit composition [9]. It causes reduced quality and quantity of the crop yield, growth rate, leaf expansion, stem elongation [8,10], stomatal conductance [11], and grain filling) [12]. Increasing the level of CO2 in plants mitigates the impact of drought stress in many legume species [13].

Physiological, biochemical, and morphological processes in plants can occur as a result of drought stress causing changes in the expression of genes that lead to alteration in protein production [8]. In addition, respiration, photosynthesis, enzyme activities, and mineral nutrition, Redox (oxidation/reduction) homeostasis, and chloroplast metabolism are influenced by drought [12,14]. Analysis of proteomics data in plant leaves showed that many drought-responsive proteins that are involved in osmotic regulation, cell structure modulation, ROS scavenging, drought signal transduction, as well as carbohydrate metabolism, were upregulated under drought stress [15].

Drought stress reduces the quality and yield of many crops. Extended periods of water deficit results in a reduction in plant growth, photosynthesis efficiency and initiates a series of actions to maintain plant survival. These actions include regulation of stomatal conductance, osmotic adjustment, cell turgor maintenance, and protection of cellular membranes, enzymes, and macromolecules from oxidative damage [16]. During drought stress, plants close their stomata which leads to plasma membrane damage; consequently, the internal CO2 concentration will be decreased, and excessive generation of reactive oxygen species will be increased, leading to a reduction in the photosynthetic rate and plant growth [17,18]. Water deficiency in maize caused a reduction in seedling survival rate, acceleration of the post-pollination embryo abortion rate, and ultimately, yield loss by postponing silking through the increase in the anthesis to the silking interval that leads to a reduction in ear and kernel number per plant [19–22]. Furthermore, water scarcity decreased photosynthetic efficiency, chlorophyll content, and CO2 exchange in maize seedlings [21,23].

Drought stress decreased total barley (*Hordeum vulgare* L.) grain yield through a drop in tillers, spikes, and grains number per individual plant [24]. In soybean, drought stress decreased branch seed and total crop yields [25]. Cell elongation in higher plants can be inhibited through water movement interruption to the surrounding elongating cells from the xylem under severe water deficiency [26].

The development of candidate drought-tolerant associated genes is based on precise screening for germplasm and breeding materials in limited water environments using bioinformatics [27]. Various studies on some plant genes subjected to drought stress described the biochemical pathways that are involved in drought acclimation. Proteins and metabolites that were involved in protective mechanisms against drought conditions were identified. These mechanisms include detoxification enzymes, redox status regulation, signaling pathways, protein folding and degradation, photosynthesis stability, and primary metabolism [18]. In addition, drought and low water content reduction lead to an increase in sugar and amino acid concentrations in plants [28].

GABA is a four-carbon non-protein amino acid that increases in plant tissues under stress [29]. The GABA shunt has a functional role in biotic and abiotic stress in plants through the improvement in the antioxidant activity to restrict ROS species production, plant cell signaling, and metabolism under stress [29,30]. The GABA shunt pathway is composed of three enzymatic reactions. GABA is largely produced through glutamate decarboxylation that is catalyzed by glutamate decarboxylase enzyme, then GABA is catabolized

inside the mitochondrial matrix to succinic semialdehyde (SSA) by GABA transaminase enzyme. Then, succinate is produced from SSA oxidization by the mitochondrial succinic semialdehyde dehydrogenase enzyme or reduced to gamma-hydroxybutyrate (GHB) [30].

GABA is made from glutamate by glutamate decarboxylase that is present in symbiotic rhizobia bacteroids [31]. Furthermore, GABA was shown to be the second most plentiful amino acid in detached pea (*Pisum sativum* L.) nodules through the nuclear magnetic resonance analysis, which, in turn, reflects the vital role of GABA in amino acid cycling during bacteroid metabolism [31]. Data showed that large amounts of GABA is accumulated in root nodules, while GABA that is newly made appeared to be limited to pea nodules that are metabolically active [32]. Bound forms of GABA accumulated in nodules of many legume species, reaching 20% of the total nitrogen content [32].

Drought stress could increase the activation of GABA transporters (ProTs and AAP3) that regulate the entry of GABA across the cell membrane. The entrance of GABA decreased drought leaf wilting and improved membrane stability via the reduction in oxidative damage in plants [33]. Endogenous GABA enhanced drought tolerance through the inhibition of lipid peroxidation and photosynthesis [33]. In addition, GABA stabilized the intracellular pH during drought stress and provided a source of nitrogen and carbon during the Krebs cycle and carbon–nitrogen metabolism in the maturation of green peas [34]. Drought stress directly influences metabolic sink strength during vegetative growth, which triggers an imbalance in redox homeostasis that affects overall plant growth and development.

In this study, the effects of drought stress on the physiological and biochemical characterization of the GABA shunt pathway in green pea (*Pisum sativum* L.) were investigated. Characterization of seeds germination pattern, seedling growth (plant height, fresh weight, dry weight, and chlorophyll content), oxidative damage (Malondialdehyde level), GABA shunt metabolite levels (GABA, Glutamate and Alanine), total proteins, and total carbohydrate levels were determined.

#### **2. Materials and Methods**

#### *2.1. Plant Materials and Growth Conditions*

The green pea seeds (*Pisum sativum* L.) that were used in this study were harvested in 2018 and obtained from local pea growers in Irbid/Jordan. All experiments were performed in the laboratory using a plant growth medium. The growth medium was a mixture of peat moss and perlite growth soil with a ratio of (2:1) wt/wt, later referred to as "soil" in a pot (7 g). Seeds were surface disinfected with 6% (*v*/*v*) sodium hypochlorite for 10 min and then washed five times with sterile distilled water. Surface sterilized seeds were planted in soil. Tap water was used to irrigate the seeds at a full water holding capacity level (80–100%) for two weeks until the appearance of seedlings) [35]. The two-week-old seedlings were then subjected to drought treatments according to soil water holding capacity levels: 80%, 60%, 40%, 20%, and 10% separately for 7 days and another set for 14 days. Plants were grown at 25 ◦C under continuous cool white fluorescent lamp illumination (40 μmol m−2 s −1). For assays and parameters determinations, three replicates of 6 pea (*Pisum sativum* L.) seeds for each replicate were used.

#### *2.2. Seed Sensitivity to Drought Stress Assay*

Three replicates of 15 pea (*Pisum sativum* L.) surface-sterilized seeds were placed on three filter papers as artificial growth surface in sealed Petri dishes and irrigated with tap water in drought treatments according to full water holding capacity levels: 80%, 60%, 40%, 20%, and 10% separately for 14 days and allowed to grow at 25 ◦C. Radicle emergence from the seeds was recorded daily for 14 days. Drought treatment effect on seed germination was calculated.

#### *2.3. Seedling Physiological Growth Parameters Assay*

After each drought treatment for the 7 and 14 days separately, plant height (cm), fresh weight (g), and dry weight (g) were determined. Shoot fresh weight (g) was determined by directly weighing the collected seedlings' shoot tissues. Shoot dry weight (g) was determined following weighing the seedlings' shoot tissues after oven drying at 70 ◦C for 48 h.

#### *2.4. Chlorophyll Content Determination*

After each drought treatment for the 7 and 14 days separately, fresh shoots tissues were harvested. Chlorophyll pigments (Chl *a* and Chl *b*) extraction and determination were performed according to Jeffrey et al. protocol [36] with the following adjustments: 300 mg fresh leaves mixed with one ml of ice-cold 90% acetone were ground in an Eppendorf tube using a micro centrifuge tube pestle to prepare the chlorophyll extract. The extracted liquid was placed in a new Eppendorf tube and centrifuged at 11,000× *g* for 5 min. The supernatant was used for the determination of both chlorophyll pigments. The resulted supernatant absorbance was measured spectrophotometrically at 647 nm, 664 nm, and 750 nm wavelengths. Equation and extension coefficients [36] were used to calculate the concentration of Chl *a* and Chl *b* (μg/mL). The average of three replicates for each treatment was calculated.

#### *2.5. Metabolites Extraction*

After each drought treatment for the 7 and 14 days separately, fresh shoot tissues were harvested. GABA shunt metabolites were extracted and determined according to Zhang and Bown [37] with the following adjustments: 500 mg of fresh shoot tissues were ground in an Eppendorf tube using a micro centrifuge tube pestle until a fine powder was obtained, and then placed in 1.5 mL Eppendorf tubes and vortexed with 400 μL methanol for 10 min. Methanol from samples was removed by overnight evaporation at room temperature. To each tube, 500 μL of 70 mM lanthanum chloride was added and vortexed for 15 min. Then tubes were centrifuged at 12,400× *g* for 5 min. One hundred and sixty microliters of 1 M KOH was added to the collected supernatants and vortexed for 10 min. Tubes were centrifuged at 12,400× *g* for 5 min. The resulted supernatant (metabolites extract) was used for the determination of GABA shunt metabolites (GABA, alanine, and glutamate). The average of three replicates was used for each drought treatment.

#### *2.6. GABA (γ-Aminobutyric Acid) Level Determination*

The protocol of Zhang and Bown [37] was used for GABA determination with the following adjustments: 50 μL of metabolites extract, 14 μL of 4 mM NADP+, 19 μL of 0.5 M potassium pyrophosphate, pH (8.6), 10 μL of (2 u/μL) GABASE enzyme (GABASE enzyme (Sigma-Aldrich Corp., St. Louis, MO, USA) was suspended in 0.1 M potassium pyrophosphate, pH (7.2), containing 12.5% Glycerol and 5 mM β-mercaptoethanol), and 10 μL of α-ketoglutarate were mixed in an Eppendorf tube to prepare the reaction mixture. The change in absorbance at 340 nm after the addition of α-ketoglutarate was measured after 90 min incubation at 25 ◦C using a microplate reader. The GABA level (nmol/mg FW) was determined using the NADPH standard curve. For each drought treatment, the average of three replicates was used.

#### *2.7. Alanine Level Determination*

The Bergmeyer protocol [38] was used for alanine determination with the following adjustments: 180 μL of 0.05 M Na-carbonate buffer, pH (10), 7 μL of 30 mM β-NAD+, 10 μL of sample extract, and 1 μL of 0.3 u/μL alanine dehydrogenase (Sigma-Aldrich, St. Louis, MO, USA) enzyme suspension were mixed in an Eppendorf tube to prepare the reaction mixture. The change in absorbance at 340 nm after the addition of alanine dehydrogenase was measured after 60 min incubation at 25 ◦C using a microplate reader. The alanine level (nmol/mg FW) was determined using the NADH standard curve. For each drought treatment, the average of three replicates was used.

#### *2.8. Glutamate Level Determination*

The Bergmeyer protocol [38] was used for glutamate determination with the following adjustments: 180 μL of 0.1 M Tris-HCl pH (8.3), 8 μL of 7.5 mM β-NAD+, 10 μL of sample extract, and 1 μL of 0.8 u/μL glutamate dehydrogenase enzyme suspension (Sigma-Aldrich, St. Louis, MO, USA) were mixed in an Eppendorf tube to prepare the deamination reaction mixture. The change in absorbance at 340 nm after the addition of glutamate dehydrogenase was measured after 60 min incubation at 25 ◦C using a microplate reader. The glutamate level (nmol/mg FW) was determined using the NADH standard curve. For each drought treatment, the average of three replicates was used.

#### *2.9. Total Protein Content Determination*

After each drought treatment for the 7 and 14 days separately, fresh shoot tissues were harvested. Total protein content was determined using a SMART BCA Protein Assay Kit (Intron Biotechnology, Gangnam-gu, Seoul, Korea) according to the manufacturer's instructions. Five hundred milligrams of fresh shoots tissues mixed with 100 μL of distilled water were ground in an Eppendorf tube using a micro centrifuge tube pestle. Fifty microliters of the extracted liquid was obtained and mixed well with 1 mL of the kit working solution. All tubes were incubated at 37 ◦C for 30 min and then kept at room temperature for 5 min. The absorbance of the samples was measured at 562 nm using a microplate reader. The BSA standard curve was used to determine the total protein concentration (μg/mL). For each drought treatment, the average of three replicates was used.

#### *2.10. Total Carbohydrates Content Determination*

After each drought treatment for the 7 and 14 days separately, fresh shoot tissues were harvested. Total carbohydrates content was determined using the Total Carbohydrate Quantification Assay Kit (catalog number: ab155891, Abcam, Cambridge, MA, USA) according to the manufacturer's instructions. Fifty milligrams of fresh shoot tissues mixed with 200 μL ice-cold kit assay buffer was ground in an Eppendorf tube using a micro centrifuge tube pestle. Tubes were then centrifuged at 12,400× *g* for 5 min. The resulted supernatant was collected. In a 96-well microplate, 30 μL of collected supernatant from each sample and the kit standard solution wells were mixed with 150 μL of concentrated H2SO4. The microplate was mixed well for 5 min on a shaker and then incubated at 90 ◦C for 15 min. The microplate was cooled at room temperature for 15 min. Thirty microliters of the kit developer solution was added to all treated samples and kit standard solution wells, mixed on a shaker at room temperature for 15 min, and measured OD at 490 nm. A glucose standard curve was used to determine the total carbohydrate concentration (μg/mg FW). For each drought treatment, the average of three replicates was used.

#### *2.11. Oxidative Damage Assay*

After each drought treatment for the 7 and 14 days separately, fresh shoot tissues were harvested. Lipid peroxidation as malondialdehyde (MDA) equivalent was determined by estimating the TBA reactive substances (TBARS) as described by Heath and Packer [39] with some adjustments: 20 mg of fresh shoots tissues was mixed with 0.25 mL of 0.5% (*w/v*) thiobarbituric acid in 20% (*w/v*) trichloroacetic acid and 0.25 mL of 175 mM NaCl in 50 mM Tris-HCl (pH 8) and ground in an Eppendorf tube using a micro centrifuge tube pestle. Tubes were placed in a water bath at 90 ◦C for 25 min. Tubes were then centrifuged at 12,400× *g* for 20 min and the supernatant was collected. A microplate reader was used to measure the absorbance of the supernatant at 532 nm. A standard curve of MDA was used to calculate the MDA level (nmol/mg FW). For each drought treatment, the average of three replicates was used.

#### *2.12. Experimental Design and Data Analysis*

A completely randomized design (CRD) was used for all experiments in this study. All treatments and assays were replicated three times. For all assays parameters, mean

and standard deviation (SD) values were determined. Results were expressed as mean. Data analysis was performed by one-way analysis of variance (ANOVA) using the Least Significant Difference (LSD) multiple comparison tests on the means using the SPSS version 16.0 software. For all data analyses, *p*-value ≤ 0.05–0.01 was considered significant, and *p*-value < 0.01 was considered highly significant.

#### **3. Results and Discussion**

*3.1. Seed Sensitivity to Drought Stress*

Germination percentage of *Pisum sativum* L. was affected significantly (*p* < 0.05) by drought treatments to all saturation levels (80%, 60%, 40%, 20%, and 10%) in pea seeds for 14 days. In general, the germination percentage significantly decreased with a positive correlation as water content decreased (Table 1). The highest seed germination percentage was observed at the 80% saturation level (*p* < 0.05). A significant decline was recorded at 40% water saturation level. After 14 days, the germination percentage was 100, 80, and 30 at 60, 40, and 20% soil water saturation levels, respectively. No germination was observed at 10% drought treatment point (Table 1). These results indicated that the 40% water saturation level is the threshold value for normal pea seed germination after 14 days (*p* = 0.0001, *r* = 0.929).

**Table 1.** Germination percentage of pea (*Pisum sativum* L.) in response to soil water holding capacity levels: 80%, 60%, 40%, 20%, and 10% grown under continuous light at 25 ◦C. Means followed by different letters are significantly different (*p* ≤ 0.05) by LSD; *r* = correlation coefficient.

\* means α < 0.05.

Seed germination is a critical stage for plant survival. Drought stress greatly affects seed germination, but the response intensity and harmful effects of stress depend on the plant species [40]. A decrease in the germination of *Eremosparton songoricum* was observed under drought stress [40]. The results of the current study indicated that as water saturation decreased, a significant drop in the germination percentage in pea (*Pisum sativum* L.) was observed. Similar to our finding, Liu et al.'s [41] study revealed that seed germination of two maize cultivars (Liansheng15 cultivar and Zhengdan 958 cultivar) was reduced under drought stress. Increased water deficit minimized absorption of water by seeds, therefore preventing their germination [42]. However, it has been shown that failure of the emergence of the root was due to a reduction in water level between the soil and the seeds [43]. In addition, a reduced water gradient affected enzymatic reactions, which caused a delay in seed germination [44].

Muscolo et al. [45] found significant differences between two lentil cultivars, *Ustica* and *Pantelleria,* collected from Sicilian islands subjected to drought stress with a remarkable decrease and delay in seed germination. Inhibition of seed germination was directly related to energy production through respiration, enzyme and hormonal activity, reserve mobilization, and dilution of the protoplasm to increase metabolism for successful embryonic growth [45]. Water availability is a limiting factor for non-dormant seed germination. It also affected the speed, uniformity, and percentage of seed emergence [46]. Dornbos et al. [47] showed that the germination percentage was decreased during the seed filling stage in soybean (*Glycine max* L.c.v Merr.) under severe drought stress. Inhibition of starch catabolism under drought and salt stress decreased seed germination [48,49]. AL-Quraan et al. [50] showed a significant decrease in seed germination of five wheat cultivars (*Triticum aes-* *tivum* L.) under salt and osmotic stress. Furthermore, AL-Quraan et al. [51] reported that cold, heat, salt, and osmotic stresses significantly reduced seed germination in two lentil (*Lens culinaris* Medik) cultivars. The results of this study showed that reduction in water content had a severe impact on seed germination and early seedling growth in pea (*Pisum sativum* L.) that might be caused by a reduction in water absorption by seeds during metabolic processes and enzymatic activity in germinating pea seeds.

#### *3.2. Seedling Physiological Growth in Response to Drought Stress*

Seedling fresh weight, dry weight, and seedling height of *Pisum sativum* L. were measured in response to drought treatments according to soil water holding capacity levels of 80%, 60%, 40%, 20%, and 10% for 7 and 14 days, separately. Seedling fresh weight and seedlings height were significantly reduced with a positive correlation as water availability decreased (Table 2). The seedling fresh weight (*p* = 0.001, 0.05, *r* = 0.957, 0.979) and seedling height (*p* = 0.001, 0.001, *r* = 0.867, 0.992) were reduced significantly after 7 and 14 days, respectively. Dry weight reduction was not significant under all drought treatment after 7 (*p* = 0.096) and 14 days (*p* = 0.228), respectively.

**Table 2.** Seedling height (cm), fresh weight, and dry weight (gm), and water content (%) of pea (*Pisum sativum* L.) in response to soil water holding capacity levels: 80%, 60%, 40%, 20%, and 10% grown under continuous light at 25 ◦C for 7 days and 14 days, separately. Means followed by different letters are significantly different (*p* ≤ 0.05) by LSD; *r* = correlation coefficient.

\* means α ≤ 0.05.

In agreement with our study, Embiale et al. [52] reported that unavailability of water resulted in a significant decline in pea's basal diameter increment, area, width, and expansion length of leaf, number of branches and leaves, and total seedlings length. Plant cells save water by avoiding active growth. Stomatal closure, diminished leaf water potential, turgor loss, reduction in cell enlargement and growth [53], and inhibition of shoot and root growth [54] are common plant responses due to water stress. In general, reduction in plant biomass is positively linked with prolonged water deficiency and inhibition of cell expansion per cell growth due to low turgor pressure [55]. Reduction in plant dry weight under water stress could be due to imbalanced stomatal conductance that leads to a reduction in carbon assimilation per unit leaf area and low biomass production [56,57]. Drought stress was associated with reduced cellular division and elongation during germination, causing a reduction in root length [58].

Khorasaninejad et al. [59] reported a reduction in shoot fresh and dry weight, root dry weight, internodes, and internodes length of peppermint (*Mentha piperita* L.) under drought stress. A decrease in root and shoot fresh and dry weights and shoot length was observed in sensitive bean genotypes under water stress [60]. Zhang et al. [61] reported that the length and width of maize leaves on seedlings were shorter under water stress due to a reduction in chlorophyll content. A reduction in water content decreased plant height, stem diameter, stem and root biomass, and total biomass of *Populus nigra* (poli) [62]. In agreement with our study, Riad et al. [63] reported that decreasing water content resulted in a reduction in plant growth parameters (root length, root, and leaves fresh and dry

weights, and plant length), biomass accumulation, and vegetative growth of green peas as compared with 100% water availability.

#### *3.3. The Effect of Drought on Chlorophyll Content*

Table 3 shows that the chlorophyll content were significantly decreased after 7 and 14 days as water content decreased. *Pisum sativum* L. seedlings possessed the highest total Chl *a* and Chl *b* contents at 80% water saturation level (*p* = 0.020, 0.003) with positive correlation (*r* = 0.938, 0.986) after 7 and 14 days, respectively. There was a significant decrease with a positive correlation in Chl *a* and Chl *b* contents in response to 7 days and 14 days of drought treatments.

**Table 3.** Chlorophyll content (Chl *a*, Chl *b*, total chlorophyll) (μg/mL) of pea (*Pisum sativum* L.) seedlings in response to soil water holding capacity levels: 80%, 60%, 40%, 20%, and 10% grown under continuous light at 25 ◦C for 7 days and 14 days, separately. Means followed by different letters are significantly different (*p* ≤ 0.05) by LSD; *r* = correlation coefficient.

\* means α < 0.05.

Chlorophyll is a major chloroplast component and has a positive relationship with photosynthetic rate. Our results agreed with Nyachiro et al.'s [64] study, which found a significant reduction in Chl *a* and Chl *b* caused by drought stress in six wheat (*Triticum aestivum* L.) cultivars. Shinde and Thakur [65] reported that drought stress significantly reduced Chl *a* and Chl *b* contents in three varieties of chickpeas during vegetative growth or anthesis. Our data indicated that decreased water availability caused a significant decrease in Chl *a* and Chl *b* contents in *Pisum sativum* L. after 7 and 14 days. This reduction in total chlorophyll content leads to a reduction in photosynthesis and plant growth which was correlated with a significant reduction in seedling growth under all water stress levels. Depending on the duration and severity of drought stress, plant genotypes, and environmental conditions, a reduction in Chl *a* and *b* contents has been reported in other plant species [66,67]. Previous studies on *Triticum aestivum* and *Zea mays* [68], *Gossypium hirsutum* [69], and *Tectona grandis* [70] found that the content of photosynthetic pigments reduced as water availability decreased. It has been reported that metabolic imbalance occurred as a result of a decrease in chlorophyll synthesis and an increase in chlorophyll degradation under drought stress [71–73].

Nitrogen is a component of the chlorophyll structure in plant tissues. Under stress conditions, a decline in chlorophyll content was associated with changes in nitrogen metabolism as a result of proline synthesis to maintain osmotic adjustment [74]. Low nitrogen levels reduced photosynthetic rates due to a reduction in chlorophyll synthesis [75]. Severe water stress decreased stomatal conductance, transpiration, and photosynthetic rate [56,76]. As a drought response mechanism, chloroplasts reduced light absorption by decreasing the chlorophyll content [77]. Studies revealed that the inhibition of photosynthesis efficiency as a result of an imbalance between light capture and usage under water shortage enhances oxidative stress. Furthermore, under water stress, stomatal closure and variation in photosynthetic metabolism lead to a reduction in CO2 availability and directly impacted the photosynthetic rate [78]. Under severe drought conditions, chlorophyll content in wheat seedlings was reduced as a result of the increased activity of chlorophillase and peroxidase enzymes [79]. In response to drought stress, chlorophyll content was

reduced in the vegetative and flowering stages of the *Avena* species [80]. Alberte et al. [81] reported a reduction in Chl *a*/*b* contents occurred in lamellar content in response to water deficit. Patro et al. [82] and Ghotbi-Ravandi et al. [83] reported that stomatal closure due to water deficit is a major factor inhibiting photosynthesis in *Arabidopsis thaliana* and barley (*Hordeum vulgare* L.) seedlings, respectively. Moreover, drought stress significantly decreased the chlorophyll content in *Pisum sativum* L. seedlings [84] and three wheat varieties (*Triticum aestivum* L.) [85]. The results of our study indicated that a water imbalance due to drought stress caused a reduction in chlorophyll content that might be attributed to the photosynthetic apparatus activity disturbance.

#### *3.4. GABA Shunt Activation in Response to Drought Stress*

The GABA shunt metabolites increased significantly as a result of drought stress caused by different water saturation levels (Table 4). There were significant differences in GABA, alanine, and glutamate content at 80%, 60%, 40%, 20%, and 10% soil water content between the 7- and 14-day treatments. In general, there was a significant increase in GABA, alanine, and glutamate (*p* = 0.001, 0.0001, 0.002, respectively) with a negative correlation (*r* = −0.987, −0.968, −0.978, respectively) with water availability after the 7-day treatment. A significant increase was also found in GABA, alanine, and glutamate (*p* = 0.0001, 0.003, 0.870, respectively) with a negative correlation (*r* = −0.906, −0.989, −0.939, respectively) after the 14-day treatment. The increase in GABA content was about 4 and 5 times higher at the 20% and 10% water levels, respectively, after the 14-day treatment when compared with GABA content at the same water levels after the 7 day treatment. The increase in GABA levels was accompanied by a slight but significant (*p* ≤ 0.01–0.05) increase in the levels of alanine and glutamate after 7- and 14-day treatments under all water saturation levels.

**Table 4.** Levels of GABA shunt metabolites (GABA, alanine, and glutamate) of pea (*Pisum sativum* L.) seedlings in response to soil water holding capacity levels: 80%, 60%, 40%, 20%, and 10% grown under continuous light at 25 ◦C for 7 days and 14 days, separately. Metabolite levels were calculated as nmol/mgFW. GABA (γ-Aminobutyric acid), Ala (alanine), Glu (glutamate). Means followed by different letters are significantly different (*p* ≤ 0.05) by LSD; *r* = correlation coefficient.

\* means α < 0.05.

The GABA shunt has been associated with physiological responses, such as cytosolic pH regulation [86], nitrogen metabolism, and carbon fluxes into the Krebs cycle [87], protection against ROS production [88], and osmoregulation and signaling [89]. The GABA shunt is also involved in carbon and nitrogen metabolism, maintenance of cell membrane integrity [90], and minimizing the negative effects of abiotic stresses on plant metabolism [91].

Many studies reported that various stresses caused an increase in endogenous GABA accumulation in plants [92]. The current study showed a significant increase in GABA content as a result of a reduction in water availability to ease the metabolic damage in pea seedlings during vegetative growth. Studies showed that GABA effectively decreased leaf wilting and improved membrane stability induced by drought stress in perennial ryegrass (*Lolium perenne*) [93], black pepper [94], and white clover *(Trifolium repens*) [33], which confirmed the beneficial effect of GABA in protecting plants from oxidative stress. Insufficient availability of oxygen resulted in significant suppression of melon seedling

growth. However, seedling growth was significantly improved when exogenous GABA was applied [95]. Several amino acid syntheses are regulated by the TCA cycle [96]. Glutamate, as a precursor amino acid of GABA production, is produced from the αketoglutaric acid (an intermediate of the TCA cycle) transamination [90]. Glutamate metabolic pathway activation via the TCA cycle was confirmed by GABA production and defense of creeping bentgrass against drought stress [97]. An increase in endogenous GABA levels due to an increase in glutamate content occurred in white clover in response to drought stress [33]. Increased content of glutamic acid and GABA were found in six winter wheat genotypes due to drought stress [98]. In addition, alanine was increased at the beginning of drought stress due to an increase in glutamate levels [99]. In this study, the increase in GABA content as water availability decreased mitigated the damaging effects of drought stress on pea (*Pisum sativum* L.) seedlings by supplying enough carbon/nitrogen source to the TCA cycle and amino acid synthesis.

#### *3.5. The Effect of Drought on Seedling Proteins Level*

In general, the total protein level in *Pisum sativum* L. seedlings increased with increasing water deficit. A significant difference was found in the total protein level of pea seedlings between the 7-day (*p* = 0.0001, *r* = −0.984) and 14-day (*p* = 0.0001, *r* = −0.982) treatments (Figure 1). The total protein level was 2 to 3 times higher after 7-day treatments when compared with the 14-day treatments under all water saturation levels. The significant increase in protein content after 7 days indicated that *Pisum sativum* L. seedlings could accumulate protein to lower the osmotic potential to maintain cellular structures and metabolic stability under drought stress. However, the steady-state level of protein content after 14 days indicated the ability of pea seedlings to tolerate prolonged water deficit by keeping stable protein metabolism to adjust cellular osmolarity and provide protection from further dehydration damage.

**Figure 1.** Total proteins level of pea (*Pisum sativum* L.) seedlings in response to soil water holding capacity levels: 80%, 60%, 40%, 20%, and 10% grown under continuous light at 25 ◦C for 7 days and 14 days, separately. Protein levels were calculated as μg/mL. Means (columns) with different letter scripts are significantly different (*p* ≤ 0.05) by LSD.

Under environmental stress factors, plants stimulated specific changes in protein synthesis [100]. In agreement with our study, Guttieri et al. [101] reported increased protein concentration under water deficit due to higher rates of nitrogen accumulation in hard wheat (*Triticum aestivum* L.) grains. Ozturk and Aydin [102] also reported an increase in protein and gluten content in winter wheat cultivars grain in response to drought stress when compared with the fully irrigated treatment. During a combination of heat and drought stress, glutamine, ornithine, tyrosine, valine, and tryptophan were accumulated in *Arabidopsis thaliana* and purslane plants to maintain cellular osmotic adjustment and keep leaf turgor in response to such stress combination [103,104]. Drought stress induced the accumulation of protein by enhancing the protein biosynthesis in wheat and barley [105,106]. However, a reserve of available substrates for protein synthesis could be associated with amino acid accumulation to facilitate quick retrieval of osmotic adjustment and plant metabolism in response to water deficiency [107]. In a study that used three Australian bread wheat (*Triticum aestivum* L.) cultivars, Ford et al. [108] suggested that the cultivar RAC875 had the highest capacity to withstand drought stress by increasing the cellular protein synthesis. Although there was an increase in total protein levels which were involved in ROS scavenging, they observed a decrease in proteins involved in the Calvin cycle and photosynthesis. In contrast to our results, Goł ˛ebiowska et al. [109] observed a decrease in the number of differential proteins in leaves of winter barley subjected to drought stress for three weeks. Mohammadkhani and Heidar [110] reported an initial increase followed by a decrease in protein concentration in maize varieties (*Zea mays* L.) subjected to drought stress. The initial increase in total soluble proteins might be due to the expression of new proteins involved in stress adaptation. However, the decrease in protein concentration might be due to a severe reduction in photosynthesis. Under water stress, proline amino acid accumulated in plant cells prevents cellular oxidation through scavenge ROS [111] and regulates plant osmotic pressure for efficient water absorption [112]. Free proline level was reported to be increased under water deficit in wheat [113,114]. The amount of proline was increased significantly in ginger (*Zingiber officinale)* in response to a reduction in water availability and prolonged duration of drought stress [112].

#### *3.6. The Effect of Drought on Seedling Carbohydrates Content*

In the current study, data showed that the total carbohydrate level in *Pisum sativum* L. seedlings increased as water availability decreased (Figure 2)., the total carbohydrate level increased significantly (*p* = 0.0001) with a negative correlation (*r* = −0.970) in response to drought stress for 7 days. Similar results were obtained under drought stress for 14 days (*p* = 0.0001, *r* = −0.0980).

**Figure 2.** Total carbohydrates level of pea (*Pisum sativum* L.) seedlings in response to soil water holding capacity levels: 80%, 60%, 40%, 20%, and 10% grown under continuous light at 25 ◦C for 7 days and 14 days, separately. Total carbohydrate level was calculated as μg/mg FW. Means (columns) with different letter scripts are significantly different (*p* ≤ 0.05) by LSD.

Water stress induces carbohydrate accumulation and osmolytes synthesis to maintain the water potential of plants [115,116]. Soluble sugar levels in plants also increased in response to drought stress. Plants respond to water shortage by balancing their potential osmotic proportion with the external environment by increasing their soluble sugars at the cellular level, reducing activities in roots, reducing the metabolism of carbohydrates as a result of severe pressure, and reducing the transfer of sugars in rinsing vessels [117]. In agreement with our study, an increase in total soluble sugars in durum (*Triticum durum* L.) wheat [118] and oligosaccharides in two sesame cultivars [119] were reported. In contrast, Akinci and Losel [120] reported a decrease in total sugar in cucumber cultivars under water stress. However, a major reserve of carbohydrates was detected in the leaves of cucumber seedlings. Under drought stress, sugar accumulation might be due to the fact that sucrose content increased because the enzyme activities involved in sucrose breakdown were diminished [121]. Furthermore, sugars protect the cells during drought by two physiological mechanisms. The first mechanism involves the hydroxyl group of sugars, which substitute the water to maintain hydrophilic interactions and hydrogen bonding in membranes and proteins during dehydration to prevent protein denaturation [122]. The second mechanism involves sugar vitrification in dehydrated cells through the formation of a biological glass in the cytoplasm to reduce water permeability [123]. Mohammadkhani and Heidari [110] reported an increase in soluble sugar concentrations as a result of starch degradation in two maize (*Zea mays* L.) cultivars. Soluble sugars accumulated only in roots of eucalyptus trees under drought stress [124]. Regier et al. [62] reported an increase in soluble sugar concentrations in the roots of two divergent clones of *Populus nigra* to maintain osmotic adjustment under water deficit. Soluble sugars were also reported to be increased and participated in plant metabolic regulation and stress signaling in response to osmotic stress [125]. Additionally, increased soluble sugars may act as osmoprotectants to stabilize the membranes and sustain cell turgor in response to water deficit [126]. Li and Li [127] showed an increase in the glucose, fructose, and sucrose content in micropropagated apple plants *(Malus domestica* Borkh) in response to water stress. It is concluded

from these previous studies that fluctuations of sugar concentrations in different plants under drought stress might be due to variations in CO2 assimilation, partitioning of carbon source-sink, activity of enzymes involved in sucrose–starch partitioning, and inhibition of Calvin cycle enzymes.

#### *3.7. Oxidative Damage in Response to Drought Stress*

The accumulation of MDA, which is a byproduct of membrane lipids' oxidative damage, can be used as a marker for oxidative stress [128]. In this study, the MDA levels in the pea seedlings were determined to evaluate the lipid peroxidation in response to drought stress. Figure 3 shows that the malondialdehyde (MDA) level of pea (*Pisum sativum* L.) seedlings was slightly but significantly increased after 7 days (*p* = 0.0001, *r* = −0.967). However, the MDA level was increased sharply and significantly under all water stress levels reaching the highest level (140 nmol/mgFW at 10% water saturation level) (*p* = 0.0001, *r* = −0.975) after 14 days. Our results showed that the pea seedlings suffered minor damage under mild (60% water saturation level) drought stress, but severe lipid peroxidation and oxidative damage of the cell membrane occurred with increased (40%, 20%, and 10% water saturation levels) water deficit (Figure 3).

**Figure 3.** Malondialdehyde (MDA) level of pea (*Pisum sativum* L.) seedlings in response to soil water holding capacity levels: 80%, 60%, 40%, 20%, and 10% grown under continuous light at 25 ◦C for 7 days and 14 days, separately. The MDA level was calculated as nmol/mg FW. Means (columns) with different letter scripts are significantly different (*p* ≤ 0.05) by LSD.

Many abiotic stresses lead to the accumulation of MDA in plant tissues due to lipid peroxidation of cellular membranes and overproduction of ROS [39]. Severe water stress caused agitation and instability in the metabolic processes in the mitochondria and chloroplasts, leading to high ROS production [129]. ROS resulting from oxidative stress are toxic and highly reactive molecules that can damage cellular macromolecules and subcellular structures [130,131]. The levels of MDA and H2O2 are indicators of the free radical reactions occurring in plant-stressed tissues [132]. Similar to our findings, Pandey et al. [133] reported that the MDA level was increased in *Avena* species leaves under drought stress, increasing membrane leakage. In addition, Zlatev et al. [134] reported an increase in membrane damage and ROS production in three bean (*Phaseolus vulgaris* L.) cultivars as a result of water deficit. Similarly, Tatar and Gevrek [135] reported an increase in the

level of MDA in wheat as the duration of drought stress increased. Leaves of growing carrots (*Daucus carota* L.) accumulated high levels of MDA content under water deficit stress [91]. Morabito and Guerrier [136] reported that oxidative stress occurred 12 h after the application of drought stress. However, after a longer period of drought stress, an antioxidants defense system was initiated. Abid et al. [137] showed that a higher MDA concentration in wheat (*Triticum aestivum* L.) was associated with higher H2O2 content and greater rate of O2 •− generation. Hernandez et al. [138] reported that salinity stress in plant tissues could lead to cell membrane rupture due to the accumulation of ROS and lipid peroxidation.

#### *3.8. Correlation between GABA Level and All Physiological and Metabolic Parameters*

Under all water treatment levels used in this study, the GABA level was negatively correlated with seed germination, seedlings height, fresh and dry weight, and chlorophyll content (Table 5). On the other hand, GABA level was positively correlated with protein and carbohydrate contents and MDA level under all water treatments. The elevated level of GABA metabolites (GABA, alanine, and glutamate), total proteins, and total carbohydrates content might be involved in cellular osmotic adjustment, protecting plants from oxidative stress, balancing of carbon and nitrogen (C:N) metabolism, and maintaining cell metabolic homeostasis and cell turgor under water stress [9,22].

**Table 5.** Correlation analysis between the GABA level and all physiological and metabolic parameters that were measured in this study under drought stress treatments after 7 and14 days, separately of pea (*Pisum sativum* L.) seedlings. G% = germination percentage; *r* = correlation coefficient.

#### **4. Conclusions**

Our study showed that water deficit had a suppressive effect on *Pisum sativum* L. growth and metabolism. Drought stress significantly inhibited seed germination, decreased fresh and dry weight and plant height due to a reduction in water absorption by seeds during metabolic processes and enzymatic activities in germinating pea seeds and early seedling growth. In addition, water deficit caused a significant decrease in chlorophyll pigments that might be attributed to disturbances in the activity of the photosynthetic apparatus. Pea seedlings suffered minor cellular damage under mild drought stress. However, the oxidative damage and ROS production were more severe with increased duration and levels of water deficit. The increase in GABA content as water availability decreased mitigated the damaging effects of drought stress on pea (*Pisum sativum* L.) seedlings by supplying enough carbon/nitrogen sources to the TCA cycle and amino acid synthesis. The significant increase in protein content might lower the osmotic potential to maintain cellular structures and metabolic stability under drought stress. Pea seedlings tolerate prolonged water deficit by keeping stable protein metabolism to adjust cellular osmolarity and to provide protection from further dehydration damage. Water deficit induced carbohydrate accumulation to maintain water potential and osmotic adjustment via the activation of osmolytes synthesis. Overall, the significant increases in GABA, protein and carbohydrate contents were to cope with the physiological impact of drought stress on

*Pisum sativum* L. seedlings by maintaining cellular osmotic adjustment, protecting plants from oxidative stress, balancing carbon and nitrogen (C:N) metabolism and maintaining cell metabolic homeostasis and cell turgor. According to data presented in the current study, a sufficient water supply is vital for normal growth and metabolism in pea seedlings. Severe (less than 40% water content of the holding capacity) and long-term drought stress should be avoided during the germination stage to ensure proper seedling growth.

**Author Contributions:** N.A.A.-Q.: Study design, statistical analysis, result interpretation, writing and revision of the manuscript. Z.I.A.-A.: Methodology assistance, data analysis, and result discussion. N.F.Q.: Performing the experimental work, data collection and writing the first draft of the manuscript. All authors have read and agreed to the published version of the manuscript.

**Funding:** This research was financially supported by the Deanship of Research, Jordan University of Science and Technology, Jordan, grant number [118/2019].

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

**Informed Consent Statement:** Not applicable.

**Data Availability Statement:** Raw data is available on request.

**Conflicts of Interest:** No conflict of interest to declare.

#### **References**

## *Article* **Impact of Drought and Flooding on Alkaloid Production in** *Annona crassiflora* **Mart**

**Ana Beatriz Marques Honório 1, Iván De-la-Cruz-Chacón 2, Mariano Martínez-Vázquez 3, Magali Ribeiro da Silva 4, Felipe Girotto Campos 1, Bruna Cavinatti Martin 1, Gustavo Cabral da Silva 1, Carmen Sílvia Fernandes Boaro 1,\* and Gisela Ferreira 1**

**Abstract:** The Brazilian Cerrado is the second largest Brazilian biome. In recent decades, a reduction in rainfall has indicated an extension of the dry season. Among the many native species of the Cerrado of the Annonaceae family and used in folk medicine, *Annona crassiflora* Mart. has fruits of high nutritional value and its by-products are sources of bioactive compounds, such as alkaloids. The aim of the study was to investigate how water stress impacts the production of alkaloids. The study was carried out in a nursery, and the knowledge was flood, field capacity and drought. Gas exchange, chlorophyll *a* fluorescence, antioxidant enzymes, total soluble sugars, starch, reducing sugars, sucrose, total alkaloids and liriodenine were analyzed. We observed that plants subjected to drought had an increase in the production of total alkaloids and liriodenine, without a reduction in photosynthetic metabolism. Plants kept under drought and flood conditions dissipated higher peroxidase activity, while catalase was higher in flooded plants. Starch showed the highest concentration in flooding plants without differing from drought plants; the lowest trehalose concentrations were found in both drought and flooding plants. The drought stimulated the synthesis of total alkaloids and liriodenine without reducing the primary metabolism, which suggests adaptation to Cerrado conditions.

**Keywords:** Annonaceae; antioxidant enzymes; carbohydrates; liriodenine; photosynthesis

#### **1. Introduction**

The Brazilian Cerrado is the second largest Brazilian biome, considered one of the 25 global biodiversity hotspots, present in more than twelve states and occupying approximately 25% of the national territory, with native flora characterized by small and twisted trees [1].

This biome covers important aquifers and rivers [2] and is located in the central area of the country, being the origin of large hydrographic regions in Brazil and in the South American continent [3,4]. However, Lee et al. [5], Debortoli et al. [6] and Penereiro et al. [7] reported reduction of approximately 70 mm in annual precipitation in the Cerrado region between 1979 and 2006, indicating an extension of the dry season. Furthermore, although with great biological diversity, the biome is under continuous threat of extinction due to the expansion of agriculture and pastures, as occurs with other biomes [2,8].

**Citation:** Honório, A.B.M.; De-la-Cruz-Chacón, I.; Martínez-Vázquez, M.; da Silva, M.R.; Campos, F.G.; Martin, B.C.; da Silva, G.C.; Fernandes Boaro, C.S.; Ferreira, G. Impact of Drought and Flooding on Alkaloid Production in *Annona crassiflora* Mart. *Horticulturae* **2021**, *7*, 414. https://doi.org/10.3390/ horticulturae7100414

Academic Editor: Yanyou Wu

Received: 26 September 2021 Accepted: 14 October 2021 Published: 19 October 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/).

47

The Cerrado vegetation is exposed to high irradiances (1500 to 2500 μmol. m−2 s−1), high temperatures (25–40 ◦C at midday) and in the dry season, low relative humidity (10 to 20%) [9]. Although there is abundance of light, the seasonality of the rainfall regime is one of the factors that limits vegetation growth, leading to greater investment in root formation to explore deeper soil layers [3]. As a consequence, species present in the Cerrado biome tend to have smaller specific leaf area; on the other hand, they invest more in the bark in order to guarantee their survival in situations of water and temperature stress [10], and present lower growth rates and greater hydraulic conductance per leaf area unit when compared to species from other biomes [11,12].

Many native Cerrado species belong to the Annonaceae family and are widely used in folk medicine for the treatment of different diseases [13,14]. Native *Annona crassiflora* Mart., widely spread throughout the Cerrado biome, popularly known as araticum do cerrado, marolo, araticum cortiça or bruto, is among the species with the most consumed fruits in this biome, with pleasant sensory characteristics and high nutritional value, rich in phenolic and oligosaccharide compounds [15], carotenoids and vitamins [16], in addition to alkaloids found in by-products such as leaves and stem bark [17], representing a natural source of bioactive compounds due to their antioxidant properties [14] and seeds with high lipid yield [18].

Alkaloids make up the most diverse group among nitrogenous compounds. Multiple biological activities have alkaloid origins, and there are several drugs available on the market produced from natural plant alkaloids [19]. Several alkaloids are found in *Annona crassiflora* Mart. leaves, peels and stems. Gonçalves et al. [20] isolated two alkaloids, namely atherospermidine and liriodenine, from the stem; Pereira et al. [21] isolated and characterized alkaloid stephelagin from the fruit peel and Egydio et al. [22] and Ferraz et al. [23] identified dimethoxy-dihydroxy-tetrahydroprotoberberine, isolaurelin, xylopine, anonaine, anoretin and romucosine in leaves.

Liriodenine, an alkaloid found in abundance in the Annonaceae family [20,24–27], has several potent biological activities [28,29], including potential antibacterial [24], antiprotozoal [30,31], cytotoxic [32,33] and antifungal activities [34]. In particular, it has activity against more than 20 phytopathogens, including *Rhizopus stolonifer* and *Aspergillus glaucus*, fungi that impair seed germination [25].

Although there are several reports of alkaloids in Annonaceae, so far, there are no reports on how drought conditions, similar to those periodically found in the Cerrado, impact alkaloid production in the species, or how the species tolerate flooding conditions and how these conditions would reflect on the synthesis of specialized metabolites such as alkaloids. Thus, the aim of this study was to investigate how water stress impacts the production of total alkaloids and liriodenine in *Annona crassiflora* Mart.

#### **2. Materials and Methods**

#### *2.1. Plant Material*

*Annona crassiflora* Mart. seedlings were obtained in the municipality of Paraguaçu— Minas Gerais (April 2019) and transported to the nursery of the Department of Forest Science of the Faculty of Agronomic Sciences, Unesp—Botucatu (coordinates 22◦51 latitude S and 48◦26 longitude W), where they were submitted to a 6-month acclimation period. Transplantation to 5-L polyethylene pots was carried out in November 2019. To fill the pots, medium-texture Dystrophic Red Latosol was used [35,36], collected from the surface layer (0–20 cm in depth).

During the period in which the experiment was conducted, plants were submitted to humidity and temperature conditions shown in Figure 1.

**Figure 1.** Average humidity and temperature between the 13th and 31st of May 2020 in the seedling nursery of the Department of Forestry Science of the Faculty of Agronomic Sciences (FCA) (DAT: days after transplanting).

#### *2.2. Experimental Design*

The experimental design was completely randomized, with three water stress levels (Flooding (>0.01 MPa); Field Capacity (−0.01 MPa) (Control) and Drought (−1.5 MPa)), with six replicates of two plants per plot. Plants remained in treatment (13–31 May 2020) until the permanent wilting point was reached. Two days after this stress condition was reached, the entire experiment was collected.

From the water retention curve, the percentage of water needed for the soil to reach −0.01 MPa (Field capacity) (Control) and −1.5 MPa (Drought) was calculated, which corresponded to 16% and 8% of water, respectively. After placing pots under the established conditions, they were weighed daily and the evapotranspiration difference was replaced to maintain previous conditions (−0.01 MPa and −1.5 MPa). To maintain plants under flooding, pots were kept in flooded trays throughout the experiment.

The moisture values corresponding to the water retention tension obtained through the soil water retention curve are shown below (Table 1).

#### *2.3. Gas Exchanges*

Gas exchanges were monitored weekly in all treatments from 9:00 a.m. to 11:00 a.m., with the aid of a CO2 gas and infrared gas analyzer ("InfraRed Gas Analyzer—IRGA", model GSF 3000 Fl WALZ, Germany) with saturating light of 450 m−2 s−1 determined by means of a light curve. For monitoring, six replicates (1 plant per replicate) of each treatment were evaluated, taking measurements on the 2nd and 3rd fully expanded leaves.

CO2 assimilation rate (*Anet*, μmol CO2 m−2 s−1), transpiration rate (*E*, mmol water vapor m−2 s−1) and stomatal conductance (*gs*, mol m−2 s−1) were determined. Water use efficiency (WUE, μmol CO2 (mmol H2O−1)) was calculated using the relationship between assimilated CO2 and the transpiration rate (*Anet/E*). The apparent carboxylation efficiency was calculated according to the relationship between CO2 assimilation rate and leaf intercellular CO2 concentration (*Anet/Ci*, mol m−2 s−1 Pa−1).

#### *2.4. Chlorophyll a Fluorescence*

Chlorophyll *a* fluorescence was performed from 9:00 a.m. to 11:00 a.m. using a fluorometer (LED-Array/PAM-Module3055-FL) on 18 plants (six replicates of 1 plant each) and leaves were acclimated to a period of 30 min in the dark by covering them with aluminum foil; then, an actinic light pulse of 4500 μmol m−2 s−1 was applied to obtain Fm (maximum dark-adapted fluorescence) and Fm- (maximum light-adapted fluorescence). In addition to the maximum leaf light-adapted and dark-adapted fluorescence, Fo (minimum dark-adapted fluorescence) and Fo' values (minimum light-adapted fluorescence) were also obtained.

The maximum quantum yield (Fv/Fm) [37], effective quantum yield (фPSII) [38], photochemical quenching (qP) [39], non-photochemical quenching (NPQ) [40] and electron transport rate (ETR) were calculated through Fm, Fo, Fm and Fo- , considering that 84% of light is absorbed by chlorophyll, with 50% of photons activating photosystem II chlorophyll and 50% photosystem I and photosystem II energy that cannot be dissipated (*Ex*), quantum yield of unregulated non-photochemical energy loss in photosystem II (фNO) and quantum yield of regulated non-photochemical energy loss in photosystem II (фNPQ) [41].

#### *2.5. Carbohydrate Concentration*

Total soluble sugars were extracted from the leaf material obtained from a pool of samples of two plants per replicate (six replicates per treatment), according to Garcia et al. [42], with minor modifications, and starch was extracted according to Clegg [43]. The procedure to determine the concentration of total soluble sugars was performed according to Morris [44]; for starch, it was described by Yemm and Folkes [45]; for reducing sugars, it was determined by Miller [46]; and for sucrose, it was established by Passos [47], with minor modifications.

#### *2.6. Activity of Antioxidant Enzymes, Hydrogen Peroxide and Lipid Peroxidation*

The extraction of antioxidant enzymes was performed as described by Kar and Mishra [48] from leaf material obtained from a pool of samples of two plants per replicate (six replicates per treatment). The activities of superoxide dismutase, EC <IP_ADDRESS> and catalase EC <IP_ADDRESS> enzymes were determined by the method of Peixoto et al. [49]; the activity of the peroxidase EC <IP_ADDRESS> enzyme was established according to Teisseire and Guy [50]; and soluble proteins were quantified as described by Bradford [51].

The hydrogen peroxide content was determined by the method of Alexieva et al. [52] and lipid peroxidation was determined according to methodology proposed by Heauth and Packer [53], and both analyses were obtained using a pool of leaf material from two plants per replicate (six replicates per treatment).

#### *2.7. Extraction of Total Alkaloids*

Total alkaloids were extracted from the root material of 18 *A. crassiflora* plants (six replicates of 2 plants each); the material was stored in a greenhouse with forced air circulation at 30 ◦C for ten days, and subsequently ground to obtain 1 g of dry mass for each replicate. Alkaloids were extracted from roots previously dried using the acid–base method. After thorough grinding, the plant material was moistened with a saturated sodium carbonate (Na2CO3) solution and left to dry for 48 h at room temperature. Alkaloids were extracted with chloroform (CHCl3) by constant stirring for 1 h and then filtered and washed with distilled water. The CHCl3 phases were extracted into a 1 M hydrochlo-ric acid (HCl) solution before being alkalinized to pH 9.5 with a saturated solution of Na2CO3. The alkaline solution was then re-extracted with CHCl3, dried with anhydrous sodium sulfate (Na2SO4), filtered and evaporated at approximately 25 ◦C to obtain total alkaloids [25].

#### *2.8. Quantification of Total Alkaloids and Liriodenine*

To determine the total alkaloid content, the 18 samples were stored at room temperature, re-solubilized with CHCl3 and transferred to quartz cuvettes. The absorbance of each

solution was obtained by spectrophotometer at 254 nm wavelength using liriodenine as the standard for the elaboration of the standard curve (y = 0.0881x − 0.0112, R2 = 0.9949).

After obtaining the extract, liriodenine was quantified using ultra-high-performance liquid chromatograph (UHPLC—Thermo Fisher-Scientific®, Waltham, MA, USA) with a gradient pump and UV-Vis detector using C 18 reverse phase column (150 × 4, 6 mm and 5 μm in particle diameter). The mobile phase was 30:70 water (pH 3.5 with acetic acid) and cratic isomethanol, with a flow rate of 1 mL/min, keeping the column temperature at 30 ◦C. Detection was carried out in UV at 254 nm. For liriodenine quantification, calibration curves were performed by analyzing the stock solution series (y = 0.3595x − 0.0011; R2 = 0.9989 for samples with up to 10 μg of liriodenine in the extract and y = 0.3658x + 1.142; R2 = 0.9992 for samples with more than 10 μg [25].

#### *2.9. Statistical Analysis*

Data were submitted to analysis of variance (ANOVA) using the SigmaPlot software Version 12 and means were compared by the Tukey test at 5% (*p* < 0.05) [54]. To present the biochemical variables, a radar chart was used. Input variables were initially standardized as a result of the different units, using the scale command from the basic package of the R computing environment, which centers the mean at zero and changes the scale to standard deviation [55].

#### **3. Results**

*Annona crassiflora* plants showed, in general, that the ability to adapt to water restriction conditions (drought: −1.5 MPa) reflected in the increase in specialized metabolism, unlike what occurred under flooding conditions (Figures 2 and 3, Table 2). In this experiment, an increase in the production of total alkaloids without the occurrence of reductions in the photosynthetic metabolism of plants (gas exchange and chlorophyll *a* fluorescence) was observed when *A. crassiflora* plants were kept under drought stress conditions (Figures 4 and 5).