Sexual Plant Reproduction

, Volume 22, Issue 2, pp 73–85

Floral biology of Ziziphus mauritiana (Rhamnaceae)

Authors

    • A. Katz Department of Dryland Biotechnologies, The Jacob Blaustein Institutes for Desert Research (BIDR)Ben-Gurion University of the Negev (BGU)
  • Bert Schneider
    • A. Katz Department of Dryland Biotechnologies, The Jacob Blaustein Institutes for Desert Research (BIDR)Ben-Gurion University of the Negev (BGU)
Original Article

DOI: 10.1007/s00497-009-0093-4

Cite this article as:
Tel-Zur, N. & Schneider, B. Sex Plant Reprod (2009) 22: 73. doi:10.1007/s00497-009-0093-4

Abstract

Floral development of the synchronous dichogamous species Ziziphus mauritiana, as followed by light and scanning electron microscopy (SEM), was divided into 11 stages using a series of landmark events. Main cellular events happen synchronously in the female and the male structures, such as meiosis in micro- and macrosporocyte cells, tetrad microspore formation and appearance of the functional megaspore cell, and onset of embryo sac differentiation coinciding with mitosis in the microspores. The last stage was characterized by anthesis and continued development of the flower, beginning with anther dehiscence (male phase) and proceeding to the female phase, which was characterized by style elongation. Flowers exhibit synchronous protandrous dichogamy; anthesis takes place in the morning (group A, e.g., clone Q-29) and afternoon (group B, e.g., clone B5/4). Stigma receptivity started after the male phase and occurred synchronously and complementarily with pollen dispersal in the two clones. Pollen viability and production were similar in the two clones, but the pollen diameter of Q-29 was significantly larger than that of B5/4. This study provides the basis for understanding the biological mechanisms regulating floral development, thus expanding the prospects for Z. mauritiana breeding programs and for further molecular and genetic studies of this species.

Keywords

Floral organogenesisProtandrousRhamnaceaeSynchronous dichogamy

Introduction

The genus Ziziphus (Rhamnaceae) comprises about 170 species native to the tropics and subtropics (Islam and Simmons 2006). Ziziphus mauritiana (Lamk.), known as ber, desert apple or Indian plum, is an evergreen thorny shrub or a small tree up to 15 m height, with many drooping branches. Under severe environmental conditions, it is a compact shrub.

Z. mauritiana is an economically important fruit crop, cultivated on marginal lands on a commercial scale, especially in India, where it is endemic (Arndt et al. 2001). Z. mauritiana is tolerant to extreme environmental conditions, including drought, high temperatures and saline water (Chrovatia et al. 1993; Mizrahi and Nerd 1996; Clifford et al. 1998). Cultivars introduced to Israel from India and planted in an experimental plot in the Southern Dead Sea Basin produced high yields (up to 80 kg/tree per year), despite the extreme temperatures prevailing there and saline water irrigation (Mizrahi and Nerd 1996). Thus, Z. mauritiana has a suitable genetic makeup for selection and breeding as a sustainable crop for the arid and semiarid regions of the world, which is an increasingly crucial challenge in the face of global climate change and ever-diminishing freshwater supplies (Pareek 2001).

Ziziphus has hypanthium-type flowers (a cup or tube bearing the floral parts above the base of the ovary of the flower) with five membranous hood-like petals. The ovary has two chambers, each with a single ovule, and is broadly attached at the base. Each fruit can bear two viable embryos. The five stamens are epipetalous, each being surrounded by a petal. The pistil is central, terminating in two stigmatic lobes (Galil and Zeroni 1967). Z. mauritiana flower buds emerge from an axillary position in a cyme inflorescence. The flowers open only for 1 day, with flowers at the top of the plant being the first to open. Each cyme has 12–14 buds (Vashishtha and Pareek 1979). Fruits are drupes, similar to small apples, with a crisp white flesh. Fruits may be oval or round, 2.5–6.3 cm long (Pareek 1997), depending on the cultivar. They are sweet and rich in antioxidant compounds and vitamins (Jawanda and Bal 1978; Machuweti et al. 2005).

Ziziphus flowers exhibit synchronous protandrous dichogamy, i.e., flowers of an individual plant mature in synchrony and anther dehiscence precedes stigma receptivity with little or no overlap between the sexual stages (Renner 2001). Such temporal separation of sexual functions has been referred to as a “temporal dioecism” (Cruden 1988). Genotypes of Ziziphus species are divided into two groups according to the timing of anthesis—morning or afternoon—designated groups A and B, respectively. In Z. mauritiana, anther dehiscence begins shortly after anthesis and terminates within 2 h (Desai et al. 1986) to 4 h (Josan et al. 1980). Flowering phases of the two types overlap, thus, utilizing complementary sexual morphs in orchard design are vital for successful cross-pollination and optimum yields. Z. mauritiana is reported to be self-incompatible (Godara 1980). The flowering period is prolonged; in Israel, the flowering season begins early July and continues till the end of October. Late pruning may delay the flowering season (unpublished data). Flowers are visited by different species of insects, including wasps, flies, butterflies, and bees (Mishra et al. 2004).

Galil and Zeroni (1967) found that two processes take place rapidly in the pistil of Z. spina-christi following anthesis: elongation of the style and stigma development. Stigmas are considered receptive when they can support germination of compatible pollen grains. Based on the observations of a sticky, shiny secretion from the stigma and on hand pollination tests, Z. mauritiana is reported to be receptive on the day of anthesis (Vashishtha and Pareek 1979; Desai et al. 1986). Yet, detailed studies of style elongation and stigma development and the relationship of these two processes to stigma receptivity in Ziziphus are lacking.

Similarly, almost no information is available on the genetic and molecular factors controlling synchronous dichogamy. A detailed morphological description of flower development would therefore provide a vital foundation for future studies that would ultimately give a better understanding of the mechanisms that regulate synchronous dichogamy. To this end, we undertook a systematic anatomical analysis of flower development based on histological sections and scanning electron microscopy (SEM) techniques. We followed the course of events through the period of flower initiation, development, and differentiation and divided the developmental process into 11 specific stages. In parallel, we studied pollen production, diameter, and viability and followed the temporal sequence of stigma receptivity after anthesis.

Materials and methods

Study site and plant material

This study was conducted in an experimental plot at the Sede-Boqer Campus of Ben-Gurion University of the Negev, located in the Negev Highlands, Israel (30°52″N, 34°46″E, 430 m above sea level). The Negev Highlands are characterized by cold and mostly sunny winters with mean daily maximum/minimum temperatures of 14.9/3.8°C, and by hot, dry summers with mean daily maximum/minimum temperatures of 32/17°C. Average annual rainfall is 80 mm with considerable deviation from year to year. Two Z. mauritiana clones, Q-29 (morning anthesis, group A) and B5/4 (afternoon anthesis, group B), previously selected in trial plantings in Israel, were used in the present work (voucher specimens are kept at the herbarium of the Hebrew University of Jerusalem, Israel). These clones were grafted onto Z. mauritiana seedlings and planted among other Ziziphus species in the experimental plot in native loess soil. Chemical fertilization (Poly-Feed DRIP 23:7:23 + 2MgO with micronutrients, Haifa Chemicals Ltd.) was supplied through a drip system. The plants were irrigated weekly with 56 l during the hot season and 16 l during the cold wet season (November–March). The grafted trees were 1.5–2.5 years old at the time of the study.

Flower development

Clone Q-29 was chosen among several genotypes planted at the experimental plot in Sede Boqer, all with similar flower morphology, due to its profuse flower production and prolonged flowering season. Shoot tips, floral buds, and flowers from Q-29 were sampled for histological and SEM analysis at various stages of development. For histological analysis, tissues were fixed with FAA (formalin:acetic acid:alcohol, 1:1:3), and then placed on ice in a vacuum desiccator to facilitate the penetration of the fixative into the plant tissue. Hard plant tissues, characteristic of desert plants, hindered embedding and resulted in collapsed tissues, thus, we experimented with several embedding and cutting methods including cryostat sectioning using the Leica CM 3050 S with Tissue-Tek® O.C.T. The optimum method was fixation, dehydration with an ethanol series, and embedding in Paraplast Plus® (McCormick™ Scientific) according to Jackson (1991). Blocks were sectioned at 8–10 μm (Leica RM2235 rotation microtome) and stained with Safranin-Fast Green (Rusin 1999). Several microtome blade models made by various manufacturers were tried for sectioning and the best results were obtained with Patho Cutter-II 350 80 mm microtome blades (ERMA Inc., Tokyo, Japan). The sections were studied and photographed with an Axioimagera1 LED (Zeiss) microscope and an Axiocam HRC camera (Zeiss).

For SEM analysis, the plant material was fixed overnight with 3.7% formaldehyde, 50% ethanol, and 5% acetic acid and then dehydrated with an increasing ethanol gradient (up to 100%). The fixed tissues were critical-point-dried with liquid CO2, mounted on aluminum stubs with double-sided tape, coated with 11 nm of gold by using an automated sputter coater (Polaron E6700, high-vacuum evaporator), and then examined with a SEM (JEOL JSM-5610LV).

Stigma receptivity

Flowers were collected at regular periods following anthesis. Whole flowers from the two cultivars were dissected under a stereomicroscope (SV6 FL, Zeiss) and stigma-style tissues were removed into 20 mM phosphate buffer pH 4.5, containing 0.1 M guaiacol and 0.1 M H2O2, until an orange-brown color was observed (approximately 1–2 min). At least ten stigmas from each clone were tested at each developmental stage. The tissues were studied and photographed with a stereomicroscope (SV6 FL, Zeiss) and an Axiocam HRC camera (Zeiss).

Pollen performance

To determine the number of pollen grains per anther, ten flowers from each clone were harvested before stamen dehiscence, and the five anthers from each flower (50 anthers) were gently removed and vortexed in a tube containing 300 μl of 0.5 M sucrose. The pollen suspension was injected into a hemocytometer and the total number of grains within the 25-square counting area was determined. The number of pollen grains per anther was calculated from the following formula according to the hemacytometer procedure (Sigma, Product information Z359629): (mean number of pollen grains per square × 104 × 0.3)/50.

The viability of the pollen grains of the two clones was assayed using fluorescein diacetate (FDA) at a final concentration of 2 μg/ml (Heslop-Harrison and Heslop-Harrison 1970). Pollen grains from undehisced anthers were collected at the beginning of anthesis, brought to the lab and dispersed on a microscope slide in a drop of FDA stain. The slide was incubated for 4–5 min at room temperature and examined with an Axioimagera1 LED (Zeiss) fluorescent microscope, with a blue excitation (450–490 nm) filter set. Brightly fluorescing grains were scored as viable, while weakly fluorescing grains or those not exhibiting fluorescence were scored as non-viable. At least 500 grains were examined per flower. In addition, the equatorial diameter of 100 viable pollen grains was measured using the AxioVision AC Rel. 4.5 program (Zeiss).

Data analysis

Differences between the two clones in pollen performance—viability, diameter, and grains per anther—were analyzed using Student’s t test with confidence limits set at α ≤ 0.05.

Results

Flower development

We examined and characterized the morphological events in flower buds of Z. mauritiana, clone Q-29, starting from the appearance of floral meristems until 24 h after anthesis. We divided the developmental process into 11 stages based on SEM and optical microscope observations of medial longitudinal sections (Table 1) as described below.
Table 1

Morphological indications and respective lengths of floral buds in Ziziphus mauritiana, clone Q-29, at various stages of development

Stage

Morphological indications and developmental activity

Bud length (mm)

Figure numbers

1

Transition to reproductive stage and meristem initiation

<0.12

1a

2

Initiation of sepal primordium

≈0.150

1b–e

3

Initiation of petal and stamen primordia

≈0.460

1f

4

Initiation of carpel primordium

≈0.490

1a, b

5

Stamen differentiation into anther and filaments

≈0.60

2c, d

6

Ovule initiation

≈0.780

2e, f

7a

Meiosis: microsporocyte at early-prophase I

≈0.90

3a, b

7b

Microsporocyte at mid-prophase I. Macrosporocyte initiation.

≈1.4

3c, d

7c

Microsporocyte at late-prophase I. Meiosis in ovules

≈1.6

4a–c

7d

Microsporocyte at metaphase I

≈1.7

4d

8

Tetrad formation of microspores and functional megaspore cell and three degenerating megaspores

≈1.7

4e, f

9a

Microspores development. Free young microspore with small vacuoles

≈1.77

5a

9b

Ovule showing large megaspore cell, and inner and outer integuments visible

≈1.9

5b

9c

Microspores with a large vacuole and a visible nucleus, the ovule with both inner and outer integument fully differentiated, micropyle visible, embryo sac formed

≈2.1

5c–f

10

Before anthesis: nearly mature pollen and enlarged ovule

≈3.0

6a–e

11

Anthesis and post-anthesis: 3 and 24 h after anthesis

 

7a–e

Stage 1: floral initiation

Flower initiation became apparent at the shoot apex with an increase in meristem size. The floral apex was visible in a longitudinal section of shoot tips (Fig. 1a).
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Fig. 1

Stages 1–3 of floral initiation and flower development in Ziziphus mauritiana, as seen in medial longitudinal sections (ac, e, f) and SEM (d). Stage 1: transition to reproductive stage. a Floral meristem initiation and bud scale. Stage 2: intitation of sepal primordium. b Floral broadening. c Flower primordium consisting of sepal primordial, covered by trichomes. d Five-sepal primordia develop in a spiral arrangement on the floral apex. e Sepal elongation. Stage 3: intitation of sepal and stamen primordial. f Petal and stamen primordia, a central concavity becomes visible (arrow). Scale bars 20 μm in a, d; 50 μm in b, c, e; and 100 μm in f. bs bud scale, fm floral meristem, pe petal, s sepal, st stamen

Stage 2: initiation of sepal primordium

The floral meristem became broadened and undulated on the upper surface, these protuberances will become the sepal primordia (Fig. 1b). The floral meristem developed into a circular structure with sepal primordia (Fig. 1c). The five-sepal primordia arose in a spiral arrangement at the border of the floral primordium (Fig. 1d). Stage 2 continued with the elongation of sepals; the floral apex is nearly flat with a slight central depression (Fig. 1e).

Stage 3: initiation of petal and stamen primordia

Differentiation of petal and stamen primordia and a central concavity (arrow) was clearly distinguishable (Fig. 1f).

Stage 4: initiation of carpel primordium

Stamen and petal whorls arose alternately to the sepal whorl and the central concavity (arrow) began to deepen (Fig. 2a). A single carpel primordium occupied most of the center of the flower bud, the petals and stamens also increased in size (Fig. 2b).
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Fig. 2

Stages 4–6 of flower development in Z. mauritiana, as seen in SEM (a, b, f) and medial longitudinal sections (ce). Stage 4: carpel primordia initiation. a Lateral view of the petal and stamen primordia development within a deep concavity (arrow). b Lateral view of carpel development. Stage 5: stamen differentiation into filaments and anthers. c Stamen differentiation and carpel elongation. d Detail of the sporogeneus cells. Stage 6: ovule initiation. e Further carpel development, ovule initiation and anther development. f Detail of top view showing the development of the anther and stigma. The five sepals, two petals and one stamen were removed. Scale bars 10 μm in d; 50 μm in a; 100 μm in b, c; and 200 μm in e, f. a anther, c carpel, op ovule primordium, pe petal, ppl primary parietal layer, s sepal, sgc sporogeneus cell, sm stigma, st stamen

Stage 5: stamen differentiation into anther and filament

Stamens became constricted at the base, indicating the onset of differentiation between the anther and the filament. In addition, elongation of the carpel was observed (Fig. 2c). At this stage, sporogenous cells were visible (Fig. 2d).

Stage 6: ovule initiation

Ovule primordium initiation was clearly visible and the anthers increased in size, becoming lobe shaped (Fig. 2e), followed by additional carpel elongation and development of bi-lobed anthers (Fig. 2f).

Stage 7: meiosis

To facilitate the description of morphological events of meiosis, we further divided stage 7 into four sub-stages, designated a–d. Floral buds at stage 7a produced anthers divided by clearly formed locules and an elongated carpel (Fig. 3a). Anther cells comprised sporogenous tissue with microsporocyte cells at early-prophase I, which were visibly different in shape from tapetal and anther wall cells (Fig. 3b). Stage 7b was defined by further ovule and anther development showing microsporocytes at mid-prophase I (Fig. 3c) and a large macrosporocyte cell in the young ovule (Fig. 3d) became discernable. In stage 7c, the anther tissues included microsporocytes at late-prophase I, illustrated by two red points clearly visible in the nucleolus (Fig. 4a). These two points stained strongly with Safranin-Fast Green and were probably two nucleolus organizer regions (NORs), indicating that NORs were active until the late stage of the prophase. The ovule was bent outward due to additional cell division upon initiation of the integument, displaying basal placentation (Fig. 4b). At this stage, the macrosporocyte cell was visible and entered meiosis (Fig. 4c). In stage 7d, floral bud development continued and microsporocytes synchronously entered metaphase I when the chromosomes moved to their equatorial position, and the nucleolus disappeared (Fig. 4d).
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Fig. 3

Flower development in Z. mauritiana, as seen in medial longitudinal sections. Stage 7a: meiosis I. a Anther showing differentiated locules. Carpel elongated. b Detail of anther cells showing microsporocytes at early-prophase I. Stage 7b: meiosis. c Detail of the anther cells, microsporocytes at middle prophase I. d Detail of the ovule cells: macrosporocyte initiation (arrow). Scale bars 10 μm in bd and 200 μm in a. a anther, c carpel, end endothecium, epi epidermal cell, mc microsporocyte, ml middle layers, msc macrosporocyte cell, ne nucellar epidermis, pc parietal cell, pe petal, s sepal, tap tapetum

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Fig. 4

Stages 7 (continued) and 8 of flower development in Z. mauritiana, as seen in medial longitudinal sections. Stage 7c: meiosis. a Detail of microsporocytes at late-prophase I, showing two NORs in the nucleolus (arrow). b Detail of the ovule showing the initiation of the inner integument. c Detail of the ovule showing the macrosporocyte cell (arrow). Stage 7d: meiosis. d Microsporocytes at metaphase I. Stage 8: tetrad formation. e Detail of the anther cells showing microspore tetrad enveloped by callose wall (arrow). f Detail of the ovule showing the functional megaspore cell and three degenerated megaspores (arrows). Scale bars 10 μm in a, c, d, f; 50 μm in b; and 20 μm in e. fmc functional megaspore cell, ii inner integument, mc microsporocyte, msc macrosporocyte cell, ne nucellar epidermis, oi outer integument, pc parietal cell, pla placenta, tap tapetum, tet tetrad

Stage 8: tetrad formation

Anthers and ovules continued to develop. At the end of meiosis, each microsporocyte formed a tetrad of four haploid microspores isolated by a massive callose wall around the tetrad and between each monad (Fig. 4e), thereby completing the microsporogenesis phase. At the end of the meiotic division, when the macrosporogenesis phase was complete, a large functional megaspore and three degenerating megaspores were evident (Fig. 4f).

Stage 9: development of microspores and ovule, and embryo sac formation

We further divided stage 9 into three sub-stages. Stage 9a was characterized by continued enlargement of the sepals, petals, and stamens. At this stage, the microspores were released into the anther locules by dissolution of the callose wall. The irregularly shaped microspores became triangular in shape (Fig. 5a). Stage 9b was characterized by additional floral bud enlargement, making the ovule with its inner and outer integuments and a single large megaspore cell clearly visible (Fig. 5b). Stage 9c was characterized by a further bud differentiation (Fig. 5c), involving the development of a nucleus and a large vacuole in the microspore cells (Fig. 5d). At this stage, the ovule showed the inner and outer integuments, both fully differentiated, a micropyle, and a funiculus (Fig. 5e). Figure 5f illustrates the formed Polygonum-type embryo sac, with egg apparatus, antipodals, and polar nuclei.
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Fig. 5

Stage 9 in flower development in Z. mauritiana, as seen in medial longitudinal sections. Stage 9a: microspores development. a Detail of the anther locule containing free young microspores with small vacuoles: microspores begin to acquire a trihedral shape. Stage 9b: ovule development. b Detail of the ovule showing the megasporocyte cell (arrow) and development of inner and outer integuments. Stage 9c: further microspore development and embryo sac formation. c Further floral development. d Detail of the anther locule containing free microspores with a visible nucleus and a large vacuole. e Detail of the ovule showing both inner and outer integuments, micropyle, and funiculus. f Detail of the Polygonum-type embryo sac, showing the egg apparatus, the antipodals and the polar nuclei. Antipodas, synergids and one polar cell were observed in adjacent sections of the same ovule (not shown). Scale bars 10 μm in a, d, f; 20 μm in b; 50 μm in e; and 500 μm in c. a anther, at antipodals, c carpel, e+s egg apparatus, f funiculus, ii inner integument, m micropyle, mcp microspore, mgs megasporocyte cell, ml middle layers, ne nucellar epidermis, o ovule, oi outer integument, pc parietal cell, pe petal, pn polar nuclei, s sepal, tap tapetum

Stage 10: before anthesis

By this stage, all the floral bud tissues were almost mature (Fig. 6a, c). Mature tricolporate pollen cells were formed and some became bicellular (Fig. 6b). The style and stigma developed further, showing young stigma papilla cells whose shape is clearly different than the style cells (Fig. 5d). The anther locule and the nearly mature pollen grains were visible in a section of the anthers (Fig. 6e).
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Fig. 6

Stage 10 of flower development in Z. mauritiana, floral bud before anthesis as seen in medial longitudinal sections (a, b) and SEM (ce). a Further development of the microspores and ovule and stigma elongation. b Detail of the nearly mature pollen grains showing some bicellular grains (arrow). c Longitudinal section of a nearly mature floral bud showing the petals and stigma. d Detail of the stylar apex. Note the differences in cell shape between the style and stigma. e Detail of an anther section, showing microspores and filament. Scale bars 20 μm in b; 100 μm in c, d; and 500 μm in a. a anther, fi filament, o ovule, pe petals, s sepal, sl style, sm stigma

Stage 11: anthesis and post-anthesis

Anthesis started with a slit at the top of the bud between the sepals and a gradual opening of the flower. Three hours after anthesis, at the male phase (Fig. 7a), the dehiscing anthers (Fig. 7b) were still releasing pollen grains (Fig. 7c). Three hours after anthesis the style was composed of small cells (Fig. 7d); 24 h after anthesis it achieved maximum elongation via cell expansion (Fig. 7e).
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Fig. 7

Stage 11 of flower development in Z. mauritiana, post-anthesis as seen in SEM (ac) and medial longitudinal sections (d, e). a Top view of a flower 3 h after anthesis, at the male phase. Four petals were removed. Petals were erect and stamens were curved toward the stigma. b Detail of opened anthers and filaments. c Detail of the mature pollen grains. d Longitudinal section of the style 3 h after anthesis. e Longitudinal section of the style 24 h after anthesis. Scale bars 10 μm in c; 100 μm in b, d, e; and 1 mm in a. a anther, fi filament, o ovule, pe petals, s sepal, sl style, sm stigma

Stigma receptivity

Peroxidase activity increases as pistils mature and reaches a peak when the stigma is most receptive to pollen. Anthesis in clone Q-29 takes place in the morning, around 9:30 a.m. under local conditions. At this time, when each stamen was covered by a petal, dehiscence was observed (Fig. 8a), the style was short, and no staining of the undeveloped stigma was observed (Fig. 8b). At noon, when the stamens were reflexed and most of the pollen grains had dispersed (Fig. 8c), the style was still short, and there was no visible stain on the immature stigma (Fig. 8d). Five hours following anthesis, when the stamens were erect and the style was fully elongated (Fig. 8e), peroxidase activity was clearly visible as spots on the apical part of the bifurcated stigma (Fig. 8f). Twenty-four hours after anthesis, the petals and stamens curved between the sepals (Fig. 8g), and the stigma was stained with large spots, indicating high peroxidase activity (Fig. 8h).
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Fig. 8

Localization of peroxidase activity in stigmas in clone Q-29. a, c, e, and g Flower at anthesis (9:30 a.m.), and 2.5, 5, and 24 h after anthesis, respectively. b, d, f, and h Stigma-style tissues were removed from the whole flower and stained with 0.1 M guaiacol and 0.1 M H2O2 in 20 mM phosphate buffer pH 4.5 to visualize peroxidase activity, at anthesis and 2.5, 5, and 24 h after anthesis, respectively. Scale bars 1 mm in a, c, e, g and 0.2 mm in b, d, f, h

We followed the peroxidase activity 26.5 h after anthesis, but by this time degeneration of the papilla cells was visible (Fig. 9a), and the entire stigma and part of the style had become brown following the immersion treatment (Fig. 9b).
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Fig. 9

Localization of peroxidase activity in stigmas in clone Q-29. a Flower at 26.5 h after anthesis. b Stigma-style tissues were removed from the whole flower and stained with 0.1 M guaiacol and 0.1 M H2O2 in 20 mM phosphate buffer pH 4.5 to visualize peroxidase activity. c Similar to b, 5 h after anthesis, but the style failed to elongate. Scale bars 1 mm in a and 0.2 mm in b, c

We observed cessation of normal style elongation in 40–50% of the flowers during the flowering season. Styles that failed to elongate did not significantly change in length from anthesis, but peroxidase activity was clearly evident as spots on the apical part of the stigma 5 h after anthesis (Fig. 9c). Styles that did not elongate 24 h after anthesis retained peroxidase activity.

Peroxidase activity was also tested for clone B5/4, in which anthesis takes place in the afternoon, at 2:30 p.m. under local conditions. Peak of peroxidase activity occurred the following day, between 9:30 a.m. and 2:30 p.m., thus, stigma receptivity in this clone occurred synchronously with pollen dispersal in Q-29 (group A), facilitating pollen transfer and pollination between the flowers of the two different groups.

Pollen performance

Pollen grain production and pollen viability were similar for the two clones; differences between the two clones for mean number of grains per anther and mean percentage of viable pollen (as determined by the FDA test) were not significant (Table 2). Mean equatorial diameter of the viable pollen grains (as determined using FDA stain) of Q-29 was significantly greater than that of B5/4 (Table 2).
Table 2

Pollen performance in clones Q-29 and B5/4

Parameter

Clone

Q-29

B5/4

Pollen grains per anther

1073 ± 130a

1170 ± 279a

Pollen viability (%)

50.3 ± 3.6a

55.7 ± 6.1a

Pollen diameter (μm)

27.3 ± 0.4a

23.1 ± 0.3b

Values in this table are mean ± SE. t test, α ≤ 0.05. Values followed by different letters are statistically different

Discussion

Flower development

The morphological characteristics of flower development in Z. mauritiana, from transition and reproductive stages to the mature flower, were followed by applying histological and SEM techniques. We demonstrated parallel occurrence of the main cellular events in the female (carpel) and the male (stamens) structures, i.e., meiosis in both micro- and macrosporocyte cells occurred at the same stage, tetrad microspore formation occurred simultaneously with the appearance of the functional megaspore cell and the three degenerating megaspores and the onset of embryo sac differentiation coincided with mitosis in the microspores.

Petals of Colubrina species (Rhamnaceae) were reported to arise from a common stamen–petal primordium (Sattler 1973). Similar petal development was described for Z. mauritiana, i.e., each petal/stamen pair seems to arise by tangential splitting of an individual primordium (Medan and Hilger 1992). The link between petals and stamens is quite strong in Rhamnaceae, as both organs tend to be fused basally (Medan and Aagesen 1995), a phenomenon which was corroborated in our work by the observed simultaneous movement of both organs following anthesis.

Our staging system provides important information of the relative timing of morphological events that accompany Z. mauritiana flower development and may facilitate future investigation of the relationships between genes and morphological traits.

Stigma receptivity

Knowledge of the timing and duration of stigma receptivity is vital for developing breeding programs for crops (Stone et al. 1995). Stigma receptivity or the ability of the stigma to support pollen germination was monitored from anthesis up to tissue necrosis. The reproductive strategy of Ziziphus species is synchronous dichogamous protandry; thus, pollen is shed before the stigma is receptive. This mechanism eliminates or significantly reduces the incidence of self- or cross-pollination between individuals of the same group. We observed and described three different stigmatic stages—immature, mature, and degenerated—in the synchronous maturation of the stigma of the two groups. Peroxidase activity increased as the stigma “matured”, reaching a peak when the stigma was most receptive to pollen (Dupuis and Dumas 1990; Dafni and Motte Manues 1998; McInnis et al. 2006). In Z. mauritiana, peroxidase activity was clearly visible after the end of the male phase and peaked 5–24 h following anthesis. A process of cell degeneration started in the stigmatic tissue shortly thereafter and the tissues finally became necrotic, showing a brown color. Previous work on other species has demonstrated different relationships between the two processes. For example, in Actinidia deliciosa, cessation of stigma receptivity occurs with degeneration of the papillae and loss of cellular integrity (Gonzalez et al. 1995), while Petunia flowers remain receptive despite necrotic stigmata (Herrero and Dickinson 1979). A relationship between stigma degeneration and stigma receptivity was not revealed in this study. Further work is thus needed to determine whether cell degeneration results in a cessation of stigma receptivity in Z. mauritiana.

Galil and Zeroni (1967) described the failure of the style to elongate in Z. spina-christi, accompanied by an undeveloped stigma. In practice, those flowers perform as male flowers, increasing the availability of pollen, then abscising. Our observations were consistent with those of Galil and Zeroni (1967), i.e., the process of style elongation in both clones stopped in 40–50% of flowers. However, in Z. mauritiana, peroxidase activity was observed in the stigmas of those styles, and the flowers did not drop off. Further experiments, such as hand pollination of flowers with undeveloped stigmas are needed in Z. mauritiana to determine whether such flowers contribute only as pollen donors or whether they can be pollinated and hence bear fruits.

Pollen performance

Pollen stainability with the FDA experiment revealed low pollen viability in both clones. Low pollen viability may be associated with chromosome variations: for example, the meiotic irregularities in the tetraploid vine cactus Selenicereus megalanthus (Lichtenzveig et al. 2000) or with extreme environmental conditions, as reported for Mangifera indica L. (Issarakraisila and Considine 1994). Pollen viability in Z. mauritiana varies among the different cultivars: it may be as high as 91%, as was reported for the cultivars Chauhara, Sanur-1, and Shamber (Desai et al. 1986) or as low as 10%, as was observed for the octaploid cultivar Illaichi (Khoshoo and Singh 1963). Further research should include cytological work and plant development studies under a range of environmental conditions in order to identify the factor(s) that affect pollen viability in Z. mauritiana.

One of the factors affecting fertilization is pollen limitation, which seems to be quite a common phenomenon in many plant species. Burd (1994) showed that 62% of the 258 species of angiosperms reviewed were pollen limited at some time or place. The number of pollen grains per anther varies widely among the different plant species. An average of 993 pollen grains per anther were observed in the insect-pollinated Euphorbia boetica (Narbona et al. 2005), a number similar to that in Z. mauritiana. Since Ziziphus flowers contain a two-chambered ovary, each with a single ovule, the number of viable total pollen grains per ovule is about 1,350–1,600. Under our local conditions, a ratio of 1,350–1,600 viable pollen grains per ovule may be a limitation; however, if the undeveloped flowers in which the process of style elongation was halted are virtually male flowers, then pollen availability may not be a limiting factor at the whole plant level. Thus, further work to elucidate the contribution of flowers with undeveloped stigmas—as pollen donors or as normal flowers—as well as testing pollen germination in vivo will contribute valuable information toward answering the still open question regarding a possible pollen limitation.

Pollen grains in Z. mauritiana are tricolporate, as was previously reported in other Rhamnaceae species such as Ziziphus lotus L., Z. spina-christi, Rhamnus alaternus, and R. utilis (Lobreau-Callen 1976; Nasri-Ayachi and Nabli 1995). Similar to previous studies reporting that pollen grains ranged in diameter from 27 to 30 μm (Desai et al. 1986), our study found an average pollen diameter of 23.1 and 27.3 μm for B5/4 and Q-29, respectively. The importance of pollen size lies in the fact that it can reflect the storage capacity for particular nutrients that may affect pollen tube growth (Roulston et al. 2000). Further work is needed to elucidate whether genotypes with large pollen grains have a competitive advantage over those with smaller grains under extreme environmental conditions. Should this hypothesis prove correct, then the pollen diameter trait may be used for selecting superior cultivars in breeding programs.

Acknowledgments

The authors thank Mr. J. Mouyal and Ms. R. Jeger (Ben-Gurion University of the Negev) for their skillful technical assistance.

Copyright information

© Springer-Verlag 2009