Changes induced by osmotic stress in the morphology ... - Springer Link

Journal of Forestry Research (2014) 25(3): 523−534 DOI 10.1007/s11676-014-0491-9

ORIGINAL PAPER

Changes induced by osmotic stress in the morphology, biochemistry, physiology, anatomy and stomatal parameters of almond species (Prunus L. spp.) grown in vitro Shakiba Rajabpoor • Soghra Kiani • Karim Sorkheh Farahnaz Tavakoli

Received: 2013-04-06;

Accepted: 2013-07-09

© Northeast Forestry University and Springer-Verlag Berlin Heidelberg 2014

Abstract: We investigated the influence of different levels of osmotic

(Browicz) A.E. Murray, Prunus scoparia Spach) and ‘Lycioides’ (Pru-

stress on growth and development in selected wild almond species (eight

nus lycioides Spach, Prunus reuteri Bossi. et Bushe) were best adapted to

Prunus spp.) grown in vitro. The study, while endorsing the efficacy of

osmotic stress. Increase in chlorophyll concentration and leaf thickness

in vitro screening of auxiliary buds of wild almond for osmotic stress

under high osmotic stress can be considered as preliminary selection

tolerance, showed species variability in its response to osmotic stress.

parameters for osmotic stress tolerance in Prunus spp. The study con-

Osmotic stress reduced growth and development of all the species. How-

firmed the efficacy of the in vitro method for screening of large number

ever, the putative tolerant Prunus spp. showed better performance than

of genotypes for osmotic stress tolerance in wild almond species.

the putative susceptible genotypes. On average there was an 80% decrease in shoot dry weight at -1.2 MPa. Reduction in shoot weight was more common in osmotic stress-susceptible species in the section labeled

Keywords: Prunus spp., proline, osmotic stress, leaf character, malondialdehyde, protoplast membrane permeability

‘Euamygdalus’. The tolerant Prunus species produced smaller changes in biochemical responses than the sensitive cultivars for malondialdehyde content, catalase activity, relative permeability of protoplast membranes, and net photosynthetic rate. The tolerant species maintained cell integrity better than drought sensitive species. Wild almond species in the section labeled ‘Spartioides’ (Prunus arabica (Olivier) Neikle, Prunus glauca

The online version is available at http://www.springerlink.com Shakiba Rajabpoor Department of Biology, Payam e Noor University, P.O.Box. 193953697, Tehran, Iran. Soghra Kiani Department of Agriculture, Payam e Noor University, P.O.Box. 193953697, Tehran, Iran. Karim Sorkheh (

)

Department of Agronomy and Plant Breeding, Faculty of Agriculture, Shahid Chamran University of Ahwaz, P.O.Box 61355/144, Iran. Tel Fax: 06113330079. E-mail: [email protected] Farahnaz Tavakoli Department of Plant Breeding and Biotechnology, Faculty of Agriculture, Shahrekord University, P.O.Box 115, Iran Corresponding editor: Hu Yanbo

Introduction Wild almond (Prunus L. spp.) is an osmotic stress-tolerant species (Rouhi et al. 2007; Sorkheh et al. 2011). Fruit trees have different morpho-biochemical, physiological and anatomical adaptations that allow them to survive osmotic stress situations (Save et al. 1995; Torrecillas et al. 1996; 1999; Sorkheh et al. 2011). Rouhi et al. (2007) found important differences between three almond species in their ecophysiological behaviour in response to osmotic stress. Adaptation to osmotic stress can vary considerably between species and even within a species (Rouhi et al. 2007; Sorkheh et al. 2011). Plants living in arid and semiarid regions use different mechanisms to survive in osmotic stress affected soils. The mechanisms involved in the control of water loss include stomatal closure, raised stomatal and cuticular resistance, changes in leaf area, orientation and anatomy (Escalona et al. 1999; Chaves et al. 2002; Lawlor 2002; Romero and Boita 2006; Vijayan et al. 2008). Liu and Zhao (2005) showed that the tolerant ramie (Boehmeria nivea L.) genotypes have higher relative water content, water potential and cell membrane stability, and accumulated more proline than the sensitive ones under 12–14 days of osmotic stress. The results of Sánchez et al. (1998) also indicated a potential role of proline in reducing the damage caused by dehydration. Lima et al. (2002) testified to increases in activity of superoxide dismutase, catalase and ascor-

524 bate peroxidase to a greater extent in the drought tolerant clone of Coffea canephora Pierre ex A. Froehner than in the sensitive one. Malondialdehyde content has a close relation with osmotic stress tolerance of plants (Liu and Zhang 1994; Jiang and Huang 2001). Barbara et al. (1999) reported that the tolerant wheat (Triticum aestivium L.) cultivar could maintain higher photosynthetic activity than a sensitive cultivar under osmotic-stress conditions. Anatomical changes induced by water deficits in higher plants are better visual indicators (Shao et al. 2008). Responses of plant tissue to water stress depend on the anatomic characteristics that regulate the transmission brought by the cells (Matsuda and Rayan 1990; Olmos et al. 2007). Anatomical alterations might occur when a plant is subject to water deficit (Makbul et al. 2011). Tissues exposed to environments with low water availability showed reduction in cell size and increase in vascular tissue and cell wall thickness (Pitman et al. 1983; Guerfel et al. 2009; Makbul et al. 2011). Modifications to cell wall architecture and alterations of xylem/phloem ratios are involved in the resistance of plants to environmental stresses (Child et al. 2003). Stomatal resistance in some trees such as almond (Prunus dulcis Mill.), peach (Prunus persica L.), olive (Olea europaea L.) and other trees increased with osmotic stress (Torrecillas et al. 1996; Giorio et al. 1999; Mahhou et al. 2005; Rouhi et al. 2006). In some plants, such as in Nerium oleander L., stomata are partly covered with outer epidermis or can even be crytoporus. The adaptations increase the distance that water vapour has to travel during transpiration. They also increase boundary layer resistance and therefore decrease transpiration. In some species, stomata lay in grooves that can be covered by scales, raphides and trichomes (Karschon 1974; Ehleringer 1980; Van Damme 1991). The anatomical features restrict water losses while simultaneously reducing daily carbon assimilation at the leaf level and decreasing the long-term net carbon gain by the whole plant (Romero and Boite 2006). Rouhi et al. (2007) and Sorkheh et al. (2009) demonstrated that non-domesticated Prunus L. species have a lower sensitivity to water stress than cultivated genotypes because of morphological and physiological characteristics, like lower leaf area, stomata density and size, and lower leaf water potential. The selection of osmotic-resistant Prunus rootstocks from nondomesticated germplasm is the focus of several almond genetic research projects (Gradziel and Kester 1998; Camposeo et al. 2010). Even so, there is little information on stomatal characteristics of Prunus L. spp. (Fanizza and Reina 1990; Guirguis et al. 1995; Rouhi et al. 2007; Sorkheh et al. 2009; Camposeo et al. 2010). The present investigation aimed to understand the influence of osmotic stress on morphological, biochemical, physiological, anatomical and stomatal characters of eight Iranian wild almond species. Different osmotic stress conditions representative of natural conditions in Iran were induced by Polyethylene Glycol 6000 (PEG 6000). The results of the study are expected to help almond breeders to select proper breeding stocks, parameters and thresholds of osmotic stress to develop osmotic-tolerant highyield genotypes in almond.

Journal of Forestry Research (2014) 25(3): 523−534

Material and methods Wild almond species We studied eight wild almond species (Prunus L. spp.) including three osmotic-tolerant species (Prunus arabica (Olivier) Neikle, Prunus glauca (Browicz) A.E. Murray, Prunus scoparia Spach) in the section labeled ‘Spartioides’, two moderately tolerant species (Prunus lycioides Spach, Prunus reuteri Bossi. et Bushe) in the section labeled ‘Lycioides’, and three osmotic stresssusceptible species (Prunus communis (L.) Archang, Prunus eleagnifolia (Spach) Fritsch, Prunus orientalis Mill. (Syn. Prunus argenta Lam.)) in the section labeled ‘Euamygdalus’. These eight species were categorized into the above three sections based on their responses to osmotic stress in vitro cultures as reported by Sorkheh et al. (2010). Morphological studies The morphological characters of 8-9 month-old nursery-raised saplings were investigated at the Faculty of Agriculture of Shahrekord University during 2010−2012. Single saplings were planted in earthen pots containing mixed soil (white sphagnum (45%), peat (40%) and perlite (15%)) (Rouhi et al. 2007). Saplings were grown in a greenhouse at 27±5°C, and 65±10% relative humidity (RH) under normal daylight and well-watered conditions from May 2010 until the end of January 2011. Pots were then arranged in a greenhouse-based gutter system described by Ranjbarfardooei et al. (2000) and Ranjbarfardooei et al. (2002). The plants were continuously irrigated using a circulating system consisting of a water pump, gutter and a reservoir containing a standard Hoagland nutrient solution. Osmotic stress treatments according to Rouhi et al. (2007) consisted of a control treatment (osmotic potential of the nutrient solution (Ψs) = -0.1 MPa), and five osmotic stress levels (Mild osmotic stress (Ψs = -0.8, -1.0 MPa), Moderate osmotic stress (Ψs =-1.1, -1.2 MPa) and Severe osmotic stress (Ψs = -1.3 MPa). Osmotic stress levels were induced by adding non-penetrating polymers of PEG 6000 (Chazen et al. 1995) to the nutrient solution following the methods of Ranjbarfardooei et al. (2000) and Rouhi et al. (2007). The concentration of PEG 6000 in the nutrient solution was determined following Burlyn and Merrill (1973). Osmotic stress treatments started on April 5, 2012, by daily and linearly adding PEG 6000 until final osmotic stress levels were gained for all treatments after two weeks, following the method of Ashraf and Leary (1996). This meant that the osmotic potential of the nutrient solution decreased at a rate of -0.057, -0.071, 0.079, -0.086 and -0.093 MPa⋅day−1 during the two weeks to achieve a final osmotic potential of -0.8, -1.0, -1.1, -1.2 and -1.3 MPa, respectively. Once the osmotic stress levels reached the levels desired for unification in the assessment, the plants were pruned 10 cm above ground level. The osmotic stress levels were kept constant by regularly checking the electric conductivity (EC)

Journal of Forestry Research (2014) 25(3): 523−534 (HI 9933, Hanna Instruments Inc., Woonsocket, RI, USA) of the nutrient solution and adding distilled water when needed until the original EC value was reached again. Mean day and night temperature in the greenhouse were 32°C and 20°C, and humidity was 65% and 85%, respectively. Daily maximum natural photosynthetic active radiation (PAR) intensity (PAR Quantum Sensor SKP 215, Skye Instruments Ltd., Powys, UK) at plant level varied between 870 and 250 µmol⋅m−2⋅s−1. Treatments were arranged as a factorial set-up with complete random design and with three replications. In total 8×18 almond plants were used. Data on the number of primary branches sprouted, height of the longest shoot, number leaves in the longest shoot, inter-nodal length, single leaf size and weight, were recorded as reported earlier (Sorkheh et al. 2007; Sorkheh et al. 2009). Leaf dimensions (length, width) of the 6th through 11th leaf counting from the top were measured with a ruler at the onset of the experiment. Leaf area for each plant was determined by a planimeter (Li-3000, LI-COR, Lincoln, NE, USA). Biochemical studies Biochemical assays were carried out on the 5th leaf from the top of each twig on the 30th day after pruning to unify the rate of growth, which is most important for biochemical assessment in wild almond species. Chlorophyll concentration of the leaf was determined following the method of Arnon (1949) with slight modification according to Sorkheh et al. (2012). Total soluble sugar, protein, proline and phenol concentrations were determined by the methods of Morris (1948), Lowry et al. (1951), Bray and Thrope (1954) and Bates et al. (1973), respectively, with a slight modification for wild almond species according to Sorkheh et al. (2012). In vivo activity of the enzyme nitrate reductase was measured by the method of Hageman and Hucklesby (1972). Mineral concentrations (Na+ and K+) in the leaf were estimated after digesting with the tri-acid method. In this method leaves from the 9th to 11th positions from the top of the twig according to Vijayan et al. (2008) were dried at 80°C for 48 h in a hot air oven, as in most of the plants the 5th leaves were not sufficient for mineral analysis due to their use in biochemical assays. The dried leaves were then powdered with a grinder. Leaf powder (1 g) was later digested with 10 mL of tri-acid digestion mixture (nitric acid, perchloric acid and sulphuric acid in 10: 4: 1 ratio) by boiling until the solution became colourless. After cooling, the digestion mixture was diluted with distilled water, filtered through Whattman filter paper No.1 and the volume made up to 100 mL. Na+ and K+ were determined by flame ionisation with a flame photometer (Systronics, Naroda Industrial Estate, Gujarat, India). Physiological measurements Physiological measurements were made on the 5th leaf from the top of plants growing in either osmotic-stressed or well-watered conditions. All analyses were done according to Zhang (1989) and with a modification updated for wild almond species according to Liu et al. (2005) and Sorkheh et al. (2011, 2012).

525

Relative permeability (REC) of protoplast membranes REC was measured by using a conductimeter (HI 9933, Hanna Instruments Inc., Woonsocket, RI, USA) according to Liu et al. (2005) with some modifications based on Sorkheh et al. (2011). Leaf samples were cut into equal sized pieces (0.3 g for each treatment) and placed in 25 mm × 150 mm culture vessels containing 15 mL of distilled water, and shaken on an orbital shaker (100 rpm) for 24 h at room temperature. The first conductance of the bathing solution was measured using a conductivity meter (ECa). The tubes were then autoclaved at 115°C for 10 min and final readings were taken following autoclaving and incubated for 24 h at room temperature (ECb).

REC = ECa/ECb Malondialdehyde (MDA) content 1 g of fresh leaves (W) was cut into small pieces, put into a mortar with an amount of liquid nitrogen and ground into slurry. The slurry was transferred into a centrifuge tube and made up to 10 mL (V1), and centrifuged at 4000 rpm for 15 min. 1.5 mL clean solution (V2) was then collected into a cuvette from the centrifuge tube, 2.5 mL of 0.5% thiobarbituric acid (V3) was added, the cuvette boiled for 15 min and cooled quickly in cold water. Finally, the solution was centrifuged at 1800 rpm for 10 min, and optical density (OD) of the clean solution was tested under 532 and 600 nm.

MDA (nmol g

−1

) = [(OD 532 − OD600 ) × (V2 + V3 ) × V1/V2 ]/(0.155 × W )

where 0.155 is the extinction coefficient in ml nmol-1. Catalase activity One gram of fresh leaf (W) was cut into small pieces, put into a mortar containing an amount of phosphoric acid buffer (pH = 7), and ground into a slurry. The slurry was transferred into a centrifuge tube and made up to 10 mL (V1), centrifuged at 4000 rpm for 15 min and the clean solution was stored. 2 mL extraction solution (V2) was collected into flask 1, and 2 mL boiled extraction solution into flask 2 (as a control), 5 mL of 1% H2O2 was added to each flask, and the flasks were kept in water at 30°C for 10 min (T), then 5 ml of 1% H2SO4 was added to each flask to stop enzymic reaction. The left H2O2 was titrated with 0.1 mol L1 KMnO4 to keep the pink color for 30 s. H2O2 decomposed (mg, A) = (X2 − X1) ×C ×1.7 where X1 is the volume of 0.1 mol L-1 KMnO4 used for flask 1, X2 is for flask 2, while 1.7 is the equivalent of H2O2 (mg) for 1 mL of 0.1 mol L-1 KMnO4. Catalase activity (mg, H2O2 g-1 min-1) = (A ×V1/ V2)/ (W ×T). Net photosynthetic rate (Pn) Net photosynthetic rate (Pn) was measured by using half a leaf

526 (main leaf vein left), trichloroacetic acid 5% was applied on the petiole to stop transport of photosynthetic products and the petiole was wrapped with tinfoil to support the leaf in an erect position. We then oven-dried and weighed a certain area of leaf (leaf discs from the cut half-leaf, leaf area (A), leaf weight (W1). Four hours (T) later, the left half-leaf was cut off, oven-dried and weighed to the same size of leaf (W2). Pn (mg CO2 dm-2 h-1) = (W2 − W1) ×1.5/A ×T where 1.5 is the index for converting assimilated dry matter into CO2 according to Liu et al. (2005). Leaf anatomical studies Leaf samples were fixed in formalin-acetic acid-alcohol (1:1:18 ratio) for 16 h. Samples were dehydrated by passing through alcohol grades of 30, 50, 70, 80, 90 and 95% for 1 h each and finally transferred to absolute alcohol for 16 h. Dehydrated samples were transferred to alcohol-chloroform mixtures (3:1; 1:1; 1:3) and finally to pure chloroform. Thin slices of paraffin (56°C melting point) were added to the chloroform at 10 min intervals until full saturation was gained. Later, the leaf-chloroformparaffin mixture was kept on a hot plate at 37°C for 48 h and shifted to 45°C for 16 h. The samples were then transferred to 60°C bath, the molten paraffin was poured out and fresh paraffin slices were added. This was repeated 3-4 times at 30 min intervals, until no smell of chloroform remained in the leaf sample. After adding fresh paraffin, the molten paraffin along with the leaf samples were transferred to paper boats. Proper orientation of the material was carried out with a hot needle. After proper solidification, sections were prepared from the samples with a rotary microtome and stained with Delafield’s Haematoxylin, counter stained with 1.00% methylene blue and mounted with euparol. Thickness of epidermis, palisade and spongy layers were recorded under a microscope (OLYMPUS BX51, Japan). Stomatal studies Imprints of epidermal cells and stomata of both sides of the leaf were taken from the 8th leaf from the top by using clear nail polish (Deborah 77600, Italy) at the end of the second stress week. A thin layer of polish was applied with a small brush onto the leaf surface so that it was distributed on a rectangular area of about 1 cm2. A transparent film with a replica print of the epidermis and stomata was formed after evaporation of the solvent. Shrinkage of the replicas might have occurred during drying, so that the surface area of the reproduced structure might have been smaller than the original on the leaf. This phenomenon was avoided by using adhesive tape to fix the replica on a microscope slide (Weyers and Meidner 1990; Jones 1992; Elagoz et al. 2006). In this study, a fully expanded leaf (upper and lower sides) of each plant from each treatment was used for measurement of stomatal density. Replicas were looked at under a light microscope (Olympus GX 31, Japan) with magnification of 40 × 10. To make stomatal counting easier, the image from the micro-

Journal of Forestry Research (2014) 25(3): 523−534 scope was transferred onto a TV screen by a video camera (JVC TK - 860 E). Stomata were directly counted on the TV screen and converted to stomatal density (the number of stomata for each mm2) by means of a calibration plate (Graticules LTD, England, 200 × 0.01= 2 mm). The minimum number of microscopic fields needed for determination of stomatal density was determined according to the method of Zaid and Hughes (1995). Based on this method, for each treatment, the stomata of 96 fields were counted (3 replications in each treatment × 16 areas and imprint × 2 counts for each area). Data analysis Analysis of variance (ANOVA) was carried out to assess differences between Prunus species, osmotic stress levels, and their interactions on measured characters. Data were statistically analyzed using a factorial design using SAS program (SAS Institute 2000). The mean separations were assessed with a Duncan multiple range test at the 1% statistical level of significance.

Results Morphological characters Mean length and width of fully expanded leaves of the eight investigated species are shown in Fig. 1. Leaf size differed significantly (p 0.01) (Fig. 5 and 7). Stomatal density differed significantly between the control and osmotic-stressed Prunus communis and Prunus lycioides. Under well-watered condition, stomatal density was highest for Prunus scoparia and lowest for P. lycioides. While under osmotic stresses, P. lycioides and P. communis had the lowest and the highest stomatal densities, respectively. P. scoparia lost its leaves in all drought stress treatments.

5 0

C arote noide s (mg g FW-1)

0.50 0.40

c

**

**

**

**

0.30

**

**

**

**

0.20

*

0.10

a

is ien t al P. o r

un i s

i fo l i a gn P. e le

ri eu te

omm P. c

P. r

i de s ycio

a

r ia

P. l

co pa P. s

la u c P. g

P. a

r a bi

ca

0.00

[ Spartioides section] [Lycioides se ction] [Euamygdalus se ction]

[Prunus L . Fig. 4: Nitrogen reductase activity (a), sodium: potassium ratio (b) and carotenoid (c) content of fully expanded leaves for the eight investigated wild almond species induced by osmotic stress at different drought levels. Mean values are given with one standard error of the mean. **significant at 0.01 level. The values shown in the figure were means of studied traits and

Fig. 6: Stomatal density of fully expanded leaves for the eight investigated wild almond species induced by drought stress at different osmotic levels. Mean values are given with one standard error of the mean. **significant at 0.01 level. The values shown in the figure were means of studied traits and denote statistically differences at the 1% level between species for a certain osmotic stress level.

denote statistically differences at the 1% level between species for a certain osmotic stress level.

Fig. 5: Anatomical features of control and severe osmotic stress wild almond species of Prunus scoparia leaf. a (Control) and b (severe osmotic stress: 1.3 Mpa)- Cross sections of leaf. le: lower epiderma, ue: upper epiderma, pp: palisade parenchyma, sp: spongy parenchyma. Bare indicate 100 μm

Fig. 7: Anatomical features of fully expanded leaves stomatal imprint for Prunus arabica (a), Prunus glauce (b), Prunus scoparia (c), Prunus lycioides (d), Prunus reuteri (e), Prunus communis (f), Prunus eleagnifolia (g) and Prunus orientalis (h) at control level. Bare indicate 100 μm.


531

Table 3: Total leaf thickness, palisade and spongy layers in fresh leaf samples of in vitro selected almond species in control and stressed treatments. Species

Tolerant to drought

P. communis P. orientalis P. eleagnifolia P. lycioides P. reuteri P. arabica P. glauca P. scoparia

Sensitive Sensitive Sensitive Moderate Moderate Tolerant Tolerant Tolerant

Mild drought -0.8 Mpa -1.0 Mpa 132bb† 138dd 136ac 147de 132ab 139bd 125aa 130ba 130ac 134bb 149bf 149bf 134ad 136bc 144be 142ag



-0.8 Mpa 42.7ad 50.2ae 40.1ac 35.6aa 38.4ab 52.9cf 52.0af 47.6de

-1.0 Mpa 50.2bc 56.9ee 45.8cb 38.5ba 39.4ba 51.1bd 51.6bd 47.6dc



-0.8 Mpa 54.2ae 57.8bg 50.6ad 40.2ab 35.5aa 56.7af 48.4ac 50.2bd

-1.0 Mpa 55.6be 56.0af 52.8bd 45.3bb 40.8ba 60.9cg 51.6bc 45.3ab

Leaf thickness (μm) Moderate drought -1.1 Mpa -1.2 Mpa 137da 133ca 152ee 144cc 143cd 145ec 138ca 145ec 137ca 142db 148ab 148ad 149cc 155df 147cb 150de Palisade layer (μm) -1.1 Mpa -1.2 Mpa 50.2bc 47.1dc 54.2de 52.9ce 50.3dc 45.2cb 40.2ca 52.3ee 40.2ca 39.6ba 52.9cd 49.8ad 55.6cf 61.3ef 41.3ab 45.3cb Spongy layer (μm) -1.1 Mpa -1.2 Mpa 56.4dc 57.3ee 57.3bd 57.8be 57.1dd 56.4cd 50.8cb 54.3db 45.2da 42.3ca 59.9be 64.4df 57.3dd 56.0cd 57.3dd 52.4cc

Severe drought -1.3 Mpa 130aa 140bb 140db 143dc 147ed 151ce 157eg 156ef -1.3 Mpa 42.2cb 51.1be 40.3ba 47.9dd 40.1ca 52.9cf 58.2dg 44.0bc -1.3 Mpa 55.6bd 58.2ce 52.1bc 50.1cb 40.8ba 60.4cf 63.1eg 52.9cc

†Each mean value is calculated from three replications. The first letter denote statistical differences at the 5% level between the different osmotic stress levels for a certain species while the second letter denote statistical differences at the 5% level between species for a certain osmotic stress level.

Discussion Leaf morphological traits related to osmotic tolerance include density of fine hairs on the leaf surface, thickness of the cuticle, darker lamina color and more erect leaf architecture (Li 1993; He et al. 1995; Adam et al. 2002). The present study showed that wild almond (Prunus L. spp.) species responded differently to osmotic stress levels. The morphological changes observed in this study agreed with the findings of Sorkheh et al. (2010). However, in contrast to the results of Sorkheh et al. (2010) on biochemical changes, in the present study we found drastic changes in leaf biochemical constituents. Accumulation of soluble sugars is a common phenomenon for plants growing at high salinity (Liu and Zhao 2005). Sucrose and other simple sugars are effective in stabilizing proteins and in the adjustment of cellular osmotic potential (Garg et al. 2002; Taji et al. 2002). The decline in a sugar concentration observed in this study at a greater drought stress level (-1.2 MPa) might have been caused by excessive use of carbohydrate substrates in energy production to counteract the adverse effect of osmotic stress (Shalhevet et al. 1974). Abiotic stress, for example, osmotic stress and salinity, is known to influence protein synthesis through inhibition of amino acid incorporation (Garg et al. 1997) and proteolysis (Uprety and Sarin 1976). Our results showed that, at higher osmotic stress

levels, leaf protein concentration declined significantly in osmotic-susceptible species but rose in osmotic stress-tolerant species, particularly in wild almond species of the ‘Spartioides’ section. The increased leaf chlorophyll concentration under high osmotic stress levels in osmotic stress-tolerant species suggests the possibility of using the chlorophyll concentration as a preliminary selection criterion in Prunus spp. for osmotic stress tolerance (Sorkheh et al. 2011; Sorkheh et al. 2012). The increase in proline concentrations observed at all osmotic stress levels can be explained, as proline is one of the most prominent osmolytes that accumulate during osmotic adjustment under abiotic stress. Proline is also involved in stabilizing sub-cellular structures, scavenging free radicals, and buffering the cellular redox potential under stress conditions. It can also act as a protein compatible hydrotrope (Srinivas and Balasubramanian 1995) to alleviate cytoplasmic acidosis, and to maintain proper NADP+/NADPH ratios (Hare and Cress 1997). Rapid break-down of proline on relief of stress could also provide enough reducing agents to support mitochondrial oxidative phosphorylation and generation of ATP for recovery from stress and repair of stress-induced damage (Hare and Cress 1997; Hare et al. 1998). The increase in proline concentration was greater in osmotic-tolerant species such as Prunus species of the ‘Spartioides’ section. This species-

532 specific difference in proline accumulation under stress has been reported for several other crops (Ahmad et al. 1981; Petrusa and Winicov 1997; Sorkheh et al. 2012). The increased activity of NRase, observed in this study, agrees with Levitt (1980). Higher NRase activity in response to abiotic stress such as salinity was also reported in barley, tomato and many other crops (Arad and Richmond 1976; Tal 1997). Our results showed increased accumulation of K+ in tolerant species. In contrast, Agastian and Vivekananda (1997) showed decline in K+ concentration in some mulberry varieties planted on saline soils. Under salt stress, plants maintain high concentrations of K+ and low concentrations of Na+ in the cytosol, a high cytosolic K+ /Na+ ratio is regarded as a key requirement for growth on high salinity soils (Weinberg et al. 1982). This is achieved by regulating the expression and activity of K+ and Na+ transporters along with H+ pumps that generate the driving force for transport (Zhu et al. 1993). Malondialdehyde is the last product in membrane liposome peroxidation, its content is a measure of the peroxidation of the membrane bound liposome: the higher the malondialdehyde content, the greater the membrane damage (Liu and Zhang 1994; Jiang and Huang 2001; Liu et al. 2005). Catalase activity is associated with elimination of active oxygen caused by stress factors such as osmotic stress. Catalase, along with superoxide dismutase, protects plant cells by scavenging superoxide radicals, H2O2, and other superoxide compounds, and prevents or reduces production of free hydroxyl radicals (Liu and Zhang 1994; Sorkheh et al. 2011). Permeability of the protoplast membrane assists in governing electrolyte leakage from plant cells, so less increase in permeability under stress conditions implies greater integrity of plant cell membranes (Liu and Zhang 1994; Sorkheh et al. 2011). Osmotic-tolerant Prunus species produced smaller changes in biochemical responses than did osmotic-sensitive cultivars for malondialdehyde content, catalase activity, relative permeability of protoplast membranes, and net photosynthetic rate. These results imply that osmotic-tolerant species maintain cell integrity better than osmotic-sensitive species when almond plants are subjected to osmotic stress. Net photosynthetic rate and carbon deposition culminate all these biological responses and chemical processes (Liu et al. 2005). Increased in stomatal density might reduce the moisture retention capacity of harvested wild almond species leaves if stomata remain open on harvesting (Vijayan et al. 1997b). The increased stomatal density at higher osmotic stress levels agrees with the reports for barley (Belkhodja et al. 1999) and bread wheat (Bhagwat and Bhatia 1993). The increase in leaf thickness under high osmotic stress conditions was mainly due to a rise in the spongy layers, while the length of palisade cells remained unaffected. Wignarajah et al. (1975) reported similar results for beans. The highest stomatal density (380 mm-2) was reported by Ranjbarfardooei et al. (2002) for Pistacia khinjuk stocks, which is higher than the highest stomatal density (257 mm-2) recorded in our study. Zamani et al. (2002) reported stomatal density in plum (600 mm-2) to be three times higher than that in almond, peach and apricot, the latter three species had the same stomatal density of 200 mm-2. Also, Ferdinand et al. (2000) reported a range of stomatal density between 233-305 mm-2 for Prunus serotina

Journal of Forestry Research (2014) 25(3): 523−534 Ehrh. But stomatal density, which might influence stomatal resistance, did not differ consistently by osmotic stresses levels in our experiment. Thus, differences in stomatal resistance between all three species including almond, peach and apricot at control and all osmotic stress levels can be mainly attributed to differences in stomatal closure. In arid and semi-arid regions where osmotic stress is the major growth limiting factor, increasing stomatal resistance and early closing of stomata could be advantageous (Sean et al. 1998). In conclusion, Prunus species are moderately tolerant to osmotic stress, but significant differences exist in the osmotic tolerance of wild almond species. Our results confirmed the findings of Sorkheh et al. (2010) that in vitro culture of seed is an easy and efficient means to screen large numbers of Prunus L. spp. for osmotic tolerance. Osmotic tolerant species used in this study can now be used for breeding osmotic stress tolerant rootstocks in almond. Acknowledgement The authors would like to thank the ANRR Center of Shahrekord for material assistance. We are also thankful to Ms. Kh. Chenaneh-Hanoni for helping in arranging the facilities and for their kind help in undertaking this study. Thanks are also due to Dr. Dario Kremer, University of Zagreb and Prof. Craig Ledbetter, USDA, for their helpful comments on an earlier draft of the manuscript and improvement of English grammar.

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