Next Article in Journal
Biomass Allocation and Nutrients Utilization in Wheat as Affected by Phosphorus Placement and Salt Stress
Previous Article in Journal
Effects of Apatite Concentrate in Combination with Phosphate-Solubilizing Microorganisms on the Yield of Ryegrass Cultivar Izorskiy
 
 
Search for Articles:
Title / Keyword
Author / Affiliation / Email
Journal
Article Type
 
 
Advanced Search
 
Section
Special Issue
Volume
Issue
Number
Page
 
Journals
Agronomy
Volume 13
Issue 6
10.3390/agronomy13061569
agronomy-logo
Submit to this Journal Review for this Journal Propose a Special Issue
► Article Menu

Article Menu

share Share announcement Help format_quote Cite question_answer Discuss in SciProfiles thumb_up
...
Endorse textsms
...
Comment
first_page
settings
Order Article Reprints
Font Type:
Arial Georgia Verdana
Font Size:
Aa Aa Aa
Line Spacing:
Column Width:
Background:
Article

Cyto-Embryological Analysis of Wild Kentucky Bluegrass Germplasm in Gansu Province, China

by
Jinqing Zhang
and
Huiling Ma
*
College of Pratacultural Science, Gansu Agricultural University, Key Laboratory of Grassland Ecosystem, Ministry of Education, Pratacultural Engineering Laboratory of Gansu Province, Sino-U.S. Center for Grazingland Ecosystem Sustainability, Lanzhou 730070, China
*
Author to whom correspondence should be addressed.
Agronomy 2023, 13(6), 1569; https://doi.org/10.3390/agronomy13061569
Submission received: 16 May 2023 / Revised: 6 June 2023 / Accepted: 7 June 2023 / Published: 8 June 2023
(This article belongs to the Section Crop Breeding and Genetics)
Browse Figures
Versions Notes

Abstract

:
Reproduction studies, particularly embryology, represent basic information of any plant. However, the current embryological information is fragmentary for Kentucky bluegrass (Poa pratensis L.). Here, paraffin sections were used to examine the cyto-embryological characteristics, including microsporogenesis, microgametogenesis, megasporogenesis, megagametogenesis, and apomixis, of wild Kentucky bluegrass germplasm from Gannan (GN) and Longnan (LN) in Gansu Province. The study found no significant differences in pollen diameter, characteristics, viability, and stigma receptivity between the two germplasm materials. The Kentucky bluegrass consisted of three anthers, and each contained four pollen sacs that were divided into left and right halves. After meiosis, the microspore mother cells formed dyads and tetrads, which were primarily symmetrical and underwent mitosis to form three-celled pollen. Kentucky bluegrass has a one-locular ovary, two-feathery stigmas, thick nucleolar and anatropous ovules, and a typical polygonum embryo sac as its reproductive organs. The main type of apomixis observed was apospory, resulting in the coexistence of multiple embryo sacs. Polyembryonic seeds were frequently observed in Kentucky bluegrass due to apospory. Most importantly, our research found that apospory caused early embryogenesis during fertilization, which is a vital embryological feature for identifying sexual reproduction and apomixis in Kentucky bluegrass. Sexual reproduction followed strict double fertilization, while in apomixis a complete seed was only formed through pseudogamy. These embryological characteristics are documented here, and their study can aid in understanding the evolution of Kentucky bluegrass.

1. Introduction

Poa pratensis, commonly known as Kentucky bluegrass, is primarily found in the moist and cool regions of the northern temperate and frigid zones. It is a vital constituent of grassland and meadow vegetation. Owing to its well-developed rhizomes and robust resistance to trampling, it is also a crucial landscaping plant for lawns and contributes to soil and water conservation efforts [1]. Additionally, it is also utilized in stabilization of eroded soils and improving soil structure and fertility [2]. Most importantly, the remarkable grass has extreme versatility, adaptability, and evolutive capacity due to its facultative apomictic system in which both apomictic and sexual hybridization/selfing coexist [3]. Apomixis is a mode of asexual reproduction in plants where the formation of seeds occurs without the fertilization of the egg cell. This represents a significant advantage for commercial development as it ensures uniform offspring and eliminates the challenges of vegetative propagation and the requirements of sexual reproduction [4]. Kentucky bluegrass mainly exhibits aposporic apomixes in which aposporous initial cells (AICs) from the nucellus are distinguishable by their dense cytoplasm and large nucleus and form lateral embryo sacs that are situated next to the reduced embryo sac [4]. The apospory may generate seeds with multiple embryos (polyembryony) that are anatomically recognizable. Supernumerary embryo production allows plants to mitigate the effects of genetic load on fertility by producing more or higher quality offspring, which is akin to reproductive compensation in plant evolution [5]. From an adaptive perspective, apomixis is a method of multiplying gene combinations that are currently adapted, while sexuality provides a reservoir of variation that can be selected in the process of evolution [6]. Therefore, P. pratensis is considered as one of the most variable, buffered, and tolerant species.
For plants to survive in their specific habitats, they must be able to grow, develop, and reproduce under suitable conditions. The mechanism by which they reproduce in time and space is an important adaptive strategy. In polar regions, some flowering plants reproduce only vegetatively or generatively, while other species reproduce by both sexual and asexual methods [7]. The extent to which a plant relies on a mixed reproduction system is species-specific, but it can also change over time and in response to environmental stressors to maximize reproductive success [7]. For example, Kentucky bluegrass is remarkably adaptive due to a wide range of chromosomal numbers and sexual and apomictic reproductive diversity [8]. Successful anther and ovule development are required for stable plant reproduction, and abnormal development can lead to reproductive failure [9]. Embryologists are particularly interested in developmental abnormalities as their understanding can shed light on the mechanisms of ontogenetic processes. Apomictic forms of angiosperms are especially intriguing in this regard as their reproductive structures often exhibit various abnormalities [10]. Comparative analysis of the gametophytic variation rate in apomictic and sexual cereal species indicates a regular pattern of abnormalities in apomixis [10]. Based on such apomictic evidence, our main hypothesis is that anatomical analysis of sporogenesis and gametogenesis development can ontogenetically confirm embryo sac origin in Kentucky bluegrass. The objective of this study was to enhance the understanding of Kentucky bluegrass reproductive biology by describing ovule ontogeny, megasporogenesis, and embryo sac ontogeny up to the seed development stages of selected Kentucky bluegrass materials as well as to identify possible ontogenetic evidence of apomictic embryo sac development. Moreover, cyto-embryological characteristics, the peculiarities of structures and processes in the generative sphere, and the type of reproductive system are specific to each species and are more stable traits than others, such as morphological ones. Therefore, they help botanists better understand a species [11]. Reproduction studies, particularly embryology, represent basic information of any plant [12]. Due to its desirable traits and considerable importance as a turfgrass and a “model species” of apomixis research [10], it is necessary to study Kentucky bluegrass’s reproductive characteristics, which may provide a scientific basis for further exploration and utilization of the species. Thus, this study is significant for the research that aims to comprehend the development of apomixis in Kentucky bluegrass and reveal the genetic mechanism of apomixis.

2. Materials and Methods

2.1. Experimental Materials

The wild germplasm materials of Kentucky bluegrass from Gannan (GN) and Longnan (LN) in Gansu Province of China were collected. Previous studies have reported that the two materials exhibited significant differences in apomictic rates [3]. Before sowing, the land was leveled and plowed (with a depth greater than 20 cm) and then dried for 10 days. After that, we cracked the soil blocks, removed all debris, applied sufficient base fertilizer (mainly organic fertilizer), and then irrigated thoroughly to prepare for sowing. On 25 April 2019, GN and LN were planted in the lawn training base of Gansu Agricultural University in the northwestern suburbs of Lanzhou (103°34′ east longitude, 36°5′ north latitude) using single grain sowing. Three plots were planted in total, and each plot had five plants of each material. The highest temperature at the time of sowing was 32 °C, and the lowest temperature was 14 °C. The terrain of the test location was inclined from northwest to southeast, with an altitude of 1517.3 m. It was in a mid-temperate climate zone, with obvious inland climate characteristics, four distinct seasons, sufficient sunlight, and a dry climate that is prone to drought. The annual precipitation was 350 mm, the annual evaporation was 1664 mm, and the annual sunshine hours were 2474.4 h. The annual average temperature was 10.3 °C. The frost-free period was 171 days. The soil of the base was mainly yellow soil with a deep soil layer containing 2.83% organic matter, 0.09% total nitrogen, 49.88 mg·kg−1 available nitrogen, and 173.19 mg·kg−1 available potassium with pH = 7.69. Combined with soil preparation before sowing, 750 kg·hm−2 of phosphorus fertilizer and 375 kg·hm−2 of nitrogen and phosphorus compound fertilizer were applied. Field management included thinning seedlings, intertillage weeding, and timely irrigation. In April 2020, from the booting stage of Kentucky bluegrass, the florets were collected every 2 days for a total of 20 times.

2.2. Observation of Pollen Morphology and Size

The method described involved collecting inflorescences of Kentucky bluegrass at full-bloom stage and preparing fresh pollen on a glass slide by dropping a drop of water. One inflorescence was randomly collected from each individual plant for measurement, and there were 15 inflorescences (15 biological replicates) for each material. The morphology of the pollen was then observed under a Moticam Pro motic 205A microscope (Olympus Corporation, Tokyo, Japan), with three randomly selected fields of view observed. The diameters of 10 pollen cells were measured in each field of view, and the horizontal and longitudinal diameters of pollen were recorded separately to calculate the “pollen index” (pollen index = horizontal diameter/longitudinal diameter). The shape of the pollen was determined based on the pollen index, with a pollen index of 0.91–1.00 indicating a round shape and a pollen index less than 0.90 indicating an oval shape. This method provided a quantitative assessment of pollen shape, which can provide important information on the reproductive biology of Kentucky bluegrass.

2.3. Detection of Pollen Viability

Sampling method was the same as above. The method used for determining pollen viability involved staining the pollen with iodine–potassium iodide (KI–I2) solution and observing it under a microscope. One inflorescence was randomly collected from each individual plant for measurement, and there were 15 inflorescences (15 biological replicates) of each material. The KI–I2 solution was dropped on the pollen, and after mixing, it was covered with a cover glass and incubated at 35 °C for 15 min. The pollen was then observed under the microscope, and the number of active and inactive pollen were counted. The active pollen was round and dark brown in color, while the inactive pollen was pale yellow or colorless. The pollen viability percentage was calculated as the number of active pollen divided by the total number of pollen observed multiplied by 100%. Five fields of view were selected, and the number of pollen in each field of view was about 70 to 100. The method used was based on previous studies by Snyman et al. [13] and Xu et al. [14].

2.4. Stigma Receptivity

The benzidine–hydrogen peroxide method was used to detect the receptivity of stigmas at different flowering stages. Florets at different growth stages were randomly selected and mixed with a solution of 1% benzidine + 3% Hydrogen peroxide + water (v:v:v = 4:11:22) and added to a concave glass slide. The appearance of blue and the formation of bubbles indicated that the stigma had receptivity, while the absence of blue or bubbles indicated that the stigma did not have receptivity. A total of 60 stigmas (two stigmas were randomly collected from each individual plant) were observed for each material with three replicates of 20 stigmas each.

2.5. Anther Dehiscence and Pinnate Stigma Observation

Single florets before and after anthesis were dissected and observed under a stereomicroscope (Version: Discovery. V12, Carl Zeiss, Jena, Germany) to observe the structure of the floret. Two florets were randomly collected from each individual plant and a total of 60. Additionally, scattered anthers and intact ovaries were photographed.

2.6. Embryological Analysis

For the embryological analysis, flowers at different developmental stages from the two germplasm materials were fixed in a mixture of formalin, glacial acetic acid, and 70% ethanol in a ratio of 1:1:18 parts (FAA Fixative Solution). The samples were embedded in paraffin, cut into 9 to 15 μm sections using a Leica RM2265 electric rotary automatic microtome (Beijing Yetech Technology Co., Ltd., Beijing, China), and treated according to the classical paraffin methods [15]. Two flowers were randomly collected from each individual plant and a total of 60. The sections were stained with Ehrlich’s hematoxylin (1 min) and eosin (2 min), and the double staining was carried out to obtain permanent microscope slides. The slides were observed under a Revolve RVL-100-G inverted microscope, and the stage of development of the generative sphere was described.

2.7. Seed Germination Experiment

The seeds were disinfected. The seeds were soaked in tap water for 12 h, sterilized with 70% ethanol for 2 min, and soaked in 20% sodium hypochlorite for 15 min. Finally, the seeds were rinsed 5 to 6 times with sterile water before use.
Ten plastic culture dishes and 1000 seeds from each material were taken for the germination test at 20 °C. Two layers of filter paper were laid and 4 mL sterile water was added in each culture dish. The culture conditions were exposed to light for 14 h and dark for 10 h. Water was added by weighing every day. After 25 days, the test was completed, and the germination-related indicators of each material were recorded and counted.
Germination potential (%) = number of germinated seeds on the 13th day/1000 × 100%
Germination rate (%) = number of germinated seeds on the 25th day (N25)/1000 × 100%
One-embryo seedling rate (%) = number of one-embryo seedling/N25 × 100%
Two-embryo seedling rate (%) = number of two-embryo seedling/N25 × 100%
Three-embryo seedling rate (%) = number of three-embryo seedling/N25 × 100%
polyembryony seedling rate (%) = two-embryo seedling rate + three-embryo seedling rate

2.8. Data Statistics

The data were analyzed using the independent samples t-test in SPSS version 20.0 (IBM Corp., Armonk, NY, USA) at p ≤ 0.05 level for Windows, and the results were presented as mean ± standard error.

3. Results

3.1. Detection of Pollen Viability and Stigma Receptivity

Table 1 presents the pollen traits of the two germplasm materials. The results showed that GN had larger maximum, minimum, and average pollen diameters compared to LN. The horizontal and longitudinal diameters of the two germplasm materials were measured, and the pollen indexes were calculated (Table 1). It was observed that 83.33% of the pollen in GN had an index of approximately 1, indicating round pollen, and 76.67% of the pollen in LN was round. KI–I2 staining revealed that 89.13% of the pollen grains in GN were dyed dark brown, indicating viability, and 85.26% of the pollen in LN were viable (Figure 1). Furthermore, the results of KI–I2 staining also showed that pollen grains in GN were more yellow, while those in LN were darker (Figure 1). The stigma receptivity test found that 80.00% of stigmas in GN were receptive, and 82.00% of those in LN were receptive (Table 1). Although these indicators have numerical differences, the variance analysis showed that indicators of the two germplasm materials for pollen diameter, shape, vitality, and stigma availability show no significance (p > 0.05).

3.2. Development of Male Reproductive Organs in Kentucky Bluegrass

3.2.1. Anther Development

There are three anthers in a floret of wild Kentucky bluegrass. The anthers are attached to the top of the stamen and consist of four pollen sacs, which are divided into left and right halves (Figure 2A,B). The anther structure in the early developmental stage is relatively simple, consisting of two parts, i.e., the outer epidermis and the inner meristem (Figure 2D). As the anther continue to develop, a group of cells with larger volume, denser cytoplasm, and an obvious nuclei are found in the four corners. The division of these cells accelerates, and the horizontal section of the anther gradually changes from a nearly round shape to a quadrangular shape (Figure 2C). Furthermore, these cells shortly thereafter differentiate into longitudinally arranged sporozoites on the inside of the epidermal cells in the four corners (Figure 2E). The sporozoites form two layers of cells, the inner and outer, after one cycle of division. The outer layer is the primary parietal cells, and the inner layer is the primary sporogenous cells. The primary parietal cells undergo one parallel division and multiple vertical divisions, resulting in three layers of cells arranged in concentric circles as anther wall are layered. From the outside to the inside, the layers are the epidermis and tapetum as well as a starch-rich endothecium.

3.2.2. Microspore and Microgametophyte Development

Microspores develop simultaneously with the differentiation and growth of the anther wall. The primary sporogenous cells divide mitotically to produce secondary sporogenous cells, which are densely arranged, polygonal in shape, and have dense cytoplasm (Figure 2C). The secondary sporogenous cells further divide to generate microspore mother cells, which initially appear as densely arranged cells and eventually develop into single free cells with large nuclei, dense cytoplasm, and distinct nucleoli. The microspore mother cell then enters the meiosis stage, and after the first meiosis, a dyad is formed (Figure 2F). Initially, only the dyad exists (Figure 2G), but after further development, a tetrad is formed following the second meiosis. At this stage, dyads and tetrads coexist (Figure 2H). After further development, symmetrical tetrads are completely formed in Kentucky bluegrass (Figure 2I), signifying the completion of microspore development.
The tetrads persist for a short period before the microspores separate from each other, forming irregularly shaped free microspores. Each diploid microspore mother cell undergoes meiosis, resulting in the formation of a single, round haploid microspore with a large central vacuole, which is known as mononuclear pollen (Figure 2J). The mononuclear pollen has a thickened cell wall and cytoplasm with a nucleus located in the center of the cell (Figure 2K). Subsequently, the mononuclear pollen undergoes mitosis to form pollen with two nuclei (Figure 2L,M) that divides asymmetrically to produce a larger germ cell and a smaller vegetative cell. Shortly after the formation of germ cells, they detach from the pollen wall, become spherical, and dissociate in the cytoplasm of vegetative cells (Figure 2N). Before anther dehiscence, the germ cells undergo a mitosis to form a sperm cell, resulting in the formation of three-cell type pollen. The mature pollen grains have an outer and inner wall. At this stage, the anthers are also mature (Figure 2O,P), and after further development, the anthers divide and release pollen grains (Figure 2Q,R).
The process of anther crack releasing pollen grains occurs gradually (Figure 3). Figure 3A show a mature and intact anther with mature pollen grains, and one side of the anther gradually dehisces (Figure 3B,C). Finally, both sides of the anther begin to dehisce, and the release of mature pollen grains is gradually completed (Figure 3D).

3.3. Development of Female Reproductive Organs in Kentucky Bluegrass

This part of the results is described in detail in our previous study [16]; therefore, the discussion is brief, and the development process is shown in Figures S1 and S2 of the Supplementary Materials.

3.3.1. Structure of Kentucky Bluegrass Ovary

Kentucky bluegrass has a one-locular ovary (Figure S1A) with two feathery stigmas (Figure 4), forming two layers of integuments (Figure S1B–D). The morphological differences of different ovary individuals are obvious, and Figure 4 shows this difference and the diversity of feather stigma. Additionally, it has a well-developed nucleolus and multi-layered cells, belonging to the thick nucleolar ovule, and it is a retrograde ovule.

3.3.2. Megaspore and Megagametophyte Development

The development of megaspores in Kentucky bluegrass follows a typical polygonum-type embryo sac pattern, starting with the differentiation of sporogenous cells and leading to the formation of functional megaspores (Figure S2). Sporogenous cells differentiate first (Figure S2A) and followed by the formation of megaspore mother cells (MMC) as the sporogenous cells enlarge (Figure S2B). The MMC then undergoes meiosis to form dyads and tetrads (Figure S2C). While three megaspores in the tetrad disintegrate, one megaspore continues to develop into a functional megaspore (Figure S2D,E). The development of the megagametophyte occurs mainly through mitosis. The functional megaspore further develops into a mononuclear embryo sac (Figure S2F), which undergoes consecutive mitosis to form a two-nucleated embryo sac (Figure S2G), a four-nucleated embryo sac (Figure S2H), and an eight-nucleated embryo sac (Figure S2I). This “7 cells and 8 nuclei” structure comprises two synergid cells, three antipodal cells, one central cell with two nuclei, and one egg cell.

3.4. Apomictic Embryo Sac Development Process

The development of the apomictic embryo sac in Kentucky bluegrass has been previously reported [3] and is further illustrated in this study with additional images (Figure 5). Initially, multiple nucleolar cells in the ovary become specialized and significantly larger than the surrounding cells (Figure 5A). These specialized cells then further develop and expand to form an aposporous initial cell (AIC), which is large in size and has a distinct nucleus. The AICs can coexist in the same ovary with the MMCs, and the two can be arranged laterally (Figure 5B) or vertically (Figure 5C). When the sexual megaspores are in the dyad stage, the nucleolar cells are specialized (Figure 5D), and there can also be a coexistence of AICs and dyads (Figure 5E). Moreover, multiple specialized nucleolar cells can co-develop to form multiple AICs. At this stage, the MMCs may disintegrate to form nutrients that are absorbed by the AICs. In Figure 5F, the traces left by the disintegration of the MMCs can be clearly observed, and in some ovaries, the MMCs develop normally and coexist with the two AICs (Figure 5G). The development of the AICs can be asynchronous with the development of the megasporocytes. After the sexual embryo sac forms a tetrad, the AICs can develop and form, and the two can coexist (Figure 5H). In Figure 5I–K, the AICs undergo one mitosis to form a longitudinal arrangement and are clearly differentiated from the dinucleate embryo sac developed by sexual reproduction. In addition, the existence of apomictic characteristics results in Kentucky bluegrass forming a trinucleate embryo sac (Figure 5L). The trinucleate embryo sac can originate from three sources: a single-nucleate embryo sac produced by sexual reproduction and two from AICs; the coexistence of a binucleate embryo sac produced by sexual reproduction and one from AIC; or the coexistence of three AICs, with the degeneration of the sexual embryo sac.

3.5. Fertilization Process of Kentucky Bluegrass

Kentucky bluegrass has two feathery stigmas (Figure 4); hence, the pollen is easily retained on it. Furthermore, it is mainly cross-pollinated. After the anthers dehisce, the mature pollen grains fall on the stigma of another flower. Within a short time, pollen germinates on the stigma and grows pollen tubes (Figure 6A). The pollen tube continues to grow and enters the embryo sac from a degenerated synergid at the micropylar end. At the same time, the egg cells and polar nuclei are clearly visible (Figure 6B). Subsequently, the egg cell and sperm nucleus approach and fuse, and the synergetic cells in the embryo sac gradually degenerate (Figure 6C). In some embryo sacs, the two polar nuclei will fuse to form a single secondary nucleus (Figure 6D), which will then fuse with the sperm nucleus during double fertilization (Figure 6E–G). The polar nucleus is located close to the sperm nucleus, and they gradually fuse to complete the fertilization process. In normal sexual reproduction, the sperm nucleus enters the polar nucleus earlier, and the fusion process is shorter (Figure 6H). However, the process of apomixis is characterized with the phenomenon of early embryogenesis (Figure 6I,G), i.e., before the polar nucleus is not fertilized. The egg cell or synergetic cell spontaneously form a proembryo (parthenogenesis), but polar nucleus needs to combine with sperm nucleus to produce endosperm (pseudofertilization). During the development of the embryo sac in Kentucky bluegrass, two proembryos may also form by either the simultaneous division of synergids and egg cells or by the division of integuments to form adventitious embryos (Figure 6K,L). Additionally, both sexual and apomictic reproduction can occur within the same ovary (Figure 6M). In sexual reproduction, the zygote undergoes secondary lateral divisions to form apical and basal cells (Figure 6N). The basal cells then undergo secondary oblique divisions, while the apical cells undergo secondary longitudinal divisions, eventually forming a “T”-shaped proembryo (Figure 6O,P). As the proembryo continues to develop, the four cells divide again, forming an eight-cell proembryo (Figure 6Q). After further division of the octad, the proembryo enlarges and takes on a pear shape (Figure 6R). As development continues, the embryo, hypocotyl, radicle, radicle sheath, and ectoderm are formed, respectively. The polar nucleus is fertilized to form the primary endosperm, and the primary endosperm continues to divide to gradually form endosperm cells (Figure 6S).

3.6. Seed Structure of Wild Kentucky Bluegrass

Observing Figure 7, it can be seen that the intact seeds of Kentucky bluegrass have palea shell, seed coat and pericarp, aleurone layer, embryo, and endosperm (Figure 7A–D). After further development, the embryo differentiates into cotyledons (single sheets), embryos, coleoptiles, hypocotyls, and radicles (Figure 7E). Due to the existence of apomixis in Kentucky bluegrass, a single seed can form double embryos and triple embryos in addition to single embryos. The single embryo cannot determine its reproductive mode, and at least one of the double embryos is derived from apomixis. Similarly, at least two of the three embryos are produced by apomixis. In double embryos, the development of the two embryos can be synchronized (Figure 7G–I,K), but there are also cases where the two embryos develop asynchronously (Figure 7F,L). Among the three embryos, the three embryos can develop at the same time (Figure 7J), and there is also a case where one embryo degenerates and decomposes (Figure 7M). The variation of di- and tri-embryos is complex, but in any case, their existence proves that Kentucky bluegrass can successfully form seeds in apomixis.

3.7. Seed Germination of Wild Kentucky Bluegrass

The seed germination is shown in Table 2. There was no polyembryony seedling, including two-embryo seedling and three-embryo seedling in GN, while there were 7.72% two-embryo seedling and 0.13% three-embryo seedling in LN. There was a significant difference in the comparison of the two germplasm materials with the same indicator. This phenomenon was consistent with the result of seed dissection. There were polyembryonic seeds in P. pratensis, which can form polyembryony seedlings. The polyembryony seedlings had different shapes, for example, there were obvious differences in the size of the two seedlings in two-embryo seedling (Figure 8). Moreover, the types were more complex in the three-embryo seedling.

4. Discussion

4.1. Pollen Viability and Stigma Receptivity in Wild Kentucky Bluegrass

Measuring pollen viability is essential for plant breeders. The commonly used pollen viability assay, the pollen germination assay, requires optimization, is time-consuming, and yields inconsistent results [17]. Consequently, chemical staining is a popular method in agriculture to assess pollen viability [17]. In this study, we employ the KI–I2 staining method to detect pollen viability. This method is straightforward, practical, and generates clear images, making it a feasible approach to assess pollen viability (Figure 1). Pollen is a small, living organism with a distinct structure and function. It has a simple structure and is easy to manipulate in vitro. Pollen is closely linked to plant genetics and reproduction [18,19]. Successful plant reproduction depends not only on pollen viability but also on stigma receptivity, which significantly impacts pollination and fertilization rates [20]. Therefore, effective plant pollination requires vigorous pollen, receptive stigma, and pollinators. Pollen must pass through the pollinator to reach the receptive stigma when it is vigorous to complete pollination.
One of the characteristics of apomixis is the absence of double fertilization, which requires viable pollen, receptive stigmas, and pollinators. There is air-pollinated flower in Kentucky bluegrass, and the two germplasm materials in this study were planted in the same environment, such as climatic factors, site conditions, and management levels, hence, their fertilization may be related to pollen vigor and stigma receptivity. In this study, the pollen viability and stigma receptivity of two wild germplasm materials with different apomictic rates were tested. The results showed that the diameter of pollen in GN was slightly larger than that in LN, and most of the pollen was round (Table 1 and Figure 1). In addition, the pollen viability of both germplasm materials was greater than 85%, and the stigma receptivity was greater than 80%, and there was no difference between the two germplasm materials (Figure 1), indicating that the reason why the apomictic rates of the two wild Kentucky bluegrass were significantly different is not related to the pollen viability or the stigma receptivity. Interestingly, related scholars have reported that pollen fertility of apomictic species is lower than that of sexual species, suggesting that low pollen fertility may be a useful indicator of apomixes [21]. This is different from the results of this study, which may be due to Kentucky bluegrass being previously established as pseudogamous, i.e., it must be pollinated to produce viable endosperm and seed. However, the species being presented in the referred literature are possibly obligate apomicts and/or not pseudogamous.

4.2. Microspore, Microgametophyte, Megaspore, and Megagametophyte Development in Wild Kentucky Bluegrass

Apomixis is a special mode of reproduction that combines with hybrid production and is considered the holy grail of agriculture due to its ability to fix heterosis of F1 hybrids in succeeding generations [22]. Therefore, conducting embryological research on species with apomictic characteristics is crucial. Breeding practices have shown that understanding the law of plant reproduction and development is essential for carrying out cross-breeding work [23]. In this study, we utilized the paraffin section method to observe the development process of microspore, microgametophyte, megaspore, and megagametophyte in the wild Kentucky bluegrass of Gansu. Our objective was to investigate whether the different apomictic rates were associated with abnormal sporogenesis and gametophyte development from the perspective of reproductive biology. These findings will provide a theoretical basis for resource development and utilization as well as research on apomixis.

4.2.1. The Developmental Process of Sexual Reproduction

The findings of this study revealed that the meiotic cytokinesis of the pollen mother cell of wild Kentucky bluegrass followed a continuous type, the tetrad was of the left-right symmetrical type, and the mature pollen grain was of the 3-cell type (Figure 2). It was also observed that the ovary had one compartment, anatropous ovules, double integuments, and a typical polygonum-shaped embryo sac with “7 cells and 8 nuclei” (Figure S2), which is consistent with the studies of Su [24] and Tian et al. [25]. Most plant microspores only undergo a single mitosis to form two cells, i.e., two-celled pollen, and the second division occurs in the pollen tube. However, some plant germ cells undergo an additional mitosis in the pollen to form two sperm, resulting in three-celled pollen when dispersed [26]. Approximately 75% of studied plants have two-celled pollen, and 25% have three-celled pollen [27]. Friedman and Williams [28] posited that the female gametophyte with 7 cells and 8 nuclei is an evolutionary trait of angiosperms, and the mature embryo sac of Kentucky bluegrass is a polygonum embryo sac with 7 cells and 8 nuclei (Figure S2), indicating its evolutionary status. Therefore, three-celled pollen could be a more advanced characteristic.

4.2.2. The Developmental Process of Apomixes

In genus Poa (Poaceae), many species have apomictic characteristics, such as Poa alpina L., Poa attenuata Trin., Poa pratensis L., and Poa sterilis M. Bieb [29]. Microscopic observation is one of the most effective and direct methods to identify apomixis in plants, and it can provide the most convincing evidence for identifying apomixis. Apomixis can fix any genotype across generations and is a new direction in plant breeding [30]. The research content of apomixis in Kentucky bluegrass also involves many aspects, especially in the aspect of cell embryology, which has made remarkable progress. In the late 1970s, Yong et al. [31] outlined the technique of cleaning pistils and thick sectioning to observe apomixis in Kentucky bluegrass. Marshall and Brown [32] subsequently identified that the apomictic mode in Kentucky bluegrass is primarily apospory. In addition, a small number of parthenogenetic embryos, synergistic embryos, and antipodal embryos were identified [25]. The sporogenesis process in Kentucky bluegrass is caused by the expansion of one or more somatic cells in the ovule to restore the meristematic capacity, leading to the production of an unreduced mature embryo sac after several mitosis [33,34]. One, two, or three aposporous initial cells may occasionally be seen in Kentucky bluegrass [35]. During this process, sexual embryos and apomictic embryos can coexist in the same ovule [36,37] and develop into mature embryo sacs simultaneously, eventually producing seeds with di- or tri-embryos. As early as 1993, Kondrasho [38] had found that in the same ovule of Kentucky bluegrass the sexual embryo sac and apomictic embryo sac coexisted. Galla et al. [39] showed the polysporous embryo sac in Buxus megistophylla is a distinguishing feature of apomixis, but there is only one embryo sac at higher flowering stages. These reports support our study finding that, during the embryonic development of Kentucky bluegrass, one or more nucellus cells located in the nucellus below the epidermis and adjacent to the MMC increased in size, with obvious nuclei, and resumed the meristem ability to form an AIC (Figure 5). The AIC gradually develops into a mature apomictic embryo sac through mitosis, forming an apomictic embryo, which can coexist with the sexual embryo sac (Figure 5). Interestingly, our result found that apospory will form early embryogenesis during fertilization (Figure 6), which will become an important embryological feature for identifying sexual reproduction and apomixis in Kentucky bluegrass.
In recent years, the research on apomixis in Kentucky bluegrass has become more and more detailed. For instance, Tian et al. [25] have observed five types of embryo sacs in the reproductive process of Kentucky bluegrass ‘Barun’, and the proportion of sexual embryo sacs was 21.44%, 49.67% of somatic apomictic embryo sacs, 10.72% of parthenogenetic embryo sacs, 4.82% of synergistic embryos, and 7.66% of antipodal embryo sacs, respectively. Subsequently, Liu et al. [40] have used the wild Kentucky bluegrass and the commercial variety ‘Midnight 2’ in eight areas of Gansu Province, China as the test germplasm materials and calculated the apomictic rate. The apomictic rate ranges from 11.20–71.04%. The results of this study are similar to those of the present study. The wild germplasm materials of Kentucky bluegrass in Gansu Province of China are mainly apospory with two or three coexisting embryo sacs (Figure 5).

4.3. Pollination, Fertilization, and Seed Development of Wild Kentucky Bluegrass

Based on the observation of the pollination and fertilization process in Kentucky bluegrass, this study found that early embryogenesis in Kentucky bluegrass was influenced by the existence of apomixis. During the early stage of early embryogenesis, the development of sexual embryos was relatively conservative [41]. It was also observed that Kentucky bluegrass has a bifid feathery stigma, which exhibited a good ability to receive pollen (Figure 4). Furthermore, the sexual reproduction of Kentucky bluegrass strictly followed the double fertilization process (Figure 6). In apomixis, the maternal cells do not undergo meiosis and do not require fertilization in later stages. However, to obtain a normal endosperm of 3n, fertilization of the polar nucleus is required (Figure 6). Therefore, the apomictic embryo sac in Kentucky bluegrass can only form complete seeds after pseudofertilization.
Among the angiosperms, many plants have multi-embryo seedlings, such as Kentucky bluegrass [42], asparagus (Asparagus officinalis) [43], Suriname surinamensis (Carapa surinamensis) [44], Phoebe zhennan [45], mango (Mangifera indica) [46], and so on. Most of the methods to study plant polyembryony are judged by seed germination to form polyembryonic seedlings, such as in Figure 8. There are few reports on judging polyembryosis by dissection and observation of the number and shape of embryos. The emergence rate of polyembryos may be influenced by external environmental factors, and all embryos in polyembryonic seeds may not be able to germinate. Therefore, direct observation using the anatomical method provides more accurate results. In some species, such as Citrus reticulata [47] and Handroanthus chrysanthus [48], the direct dissection method has been used to observe polyembryos. However, in some plants with hard seed coats and small seed sizes, direct embryo counts through seed dissection may not be feasible [49]. Therefore, in this study, the polyembryonic phenomenon was observed using paraffin section method, and it was found that two embryos and three embryos were common in Kentucky bluegrass (Figure 7). Furthermore, asynchronous development of embryos was also observed. Among three embryos, all embryos can develop at the same time, and there is also a situation in which one embryo degenerates and decomposes. It shows that the changes of di-embryo and tri-embryo are complex and diverse, but in any case, their existence proves that Kentucky bluegrass can successfully form seeds by apomictic mode.

Supplementary Materials

The following supporting information can be downloaded at: https://www.mdpi.com/article/10.3390/agronomy13061569/s1. Figure S1. The ovary structure of wild germplasm materials of Kentucky bluegrass. STI: stigma; STY: stylus; AN: anther; OW: ovary wall; OV: ovule; PL: placenta; LO: lodicules; BR: bract; PG: pollen grain; II: inter integument; OI: outer integument; MI: micropyle; NU: nucellus; AC: archesporial cell; MMC: megaspore mother cell; DY: dyad; TE: tetrad; ME: megaspore; MS: mononuclear embryo sac; TS: two-nucleus embryo sac; FS: four-nucleus embryo sac; ES: eight-nucleus embryo sac; SY: synergid; EC: egg cell; PN: polar nucleus; ANC: antipodal cells. Figure S2. The development of embryo sac in wild germplasm materials of Kentucky bluegrass. AC: archesportial cell; MMC: megaspore mother cell; DY: dyad; TE: tetrad; ME: megaspore; MO: mononuclear embryo sac; TS: two-nucleus embryo sac; FS: two-nucleus embryo sac; NC: nucellar cell; AIC: aposporous initial cell; SY: synergid; EC: egg cell; PN: polar nucleus; ANC: antipodal cells.

Author Contributions

H.M. conceived the original research plans, supervised the experiments, provided funding supporting, and agreed to serve as the author responsible for contact and communication. J.Z. performed the experiments, analyzed the data, and wrote the article. All authors have read and agreed to the published version of the manuscript.

Funding

This research was supported by the National Natural Science Foundation of China (NSFC) (Funder: Huiling Ma; No. 31760699) and the “Innovation Star” Project for Outstanding Postgraduates in Gansu Province (Funder: Jinqing Zhang; No. 2021CXZX-347).

Data Availability Statement

All relevant files are included in this article and its supplementary files.

Acknowledgments

We thank Yan Liu for helping with experimental guidance. We would like to thank Joseph Elliot at the University of Kansas for his assistance with English language and grammatical editing of the manuscript.

Conflicts of Interest

The authors declare no conflict of interest.

References

  1. Gillespie, L.J.; Soreng, R.J. Phylogenetic analysis of the bluegrass genus Poa based on DNA restriction site date. Syst. Bot. 2005, 30, 84–105. [Google Scholar] [CrossRef]
  2. Wieners, R.R.; Fei, S.-z.; Johnson, R.C. Characterization of a USDA Kentucky Bluegrass (Poa pratensis L.) Core Collection for Reproductive Mode and DNA Content by Flow Cytometry. Genet. Resour. Crop Evol. 2006, 53, 1531–1541. [Google Scholar] [CrossRef]
  3. Zhang, J.Q.; Ma, H.L. The Female Gametophyte Characteristics and Gene Expression Analysis Involved in Apomixis of Wild Germplasm Materials of Kentucky Bluegrass in Gansu Province of China. J. Plant Growth Regul. 2022, 42, 2283–2304. [Google Scholar] [CrossRef]
  4. Souza Perez, M.; Speroni, G. New apomictic pathway in Myrtaceae inferred from Psidium cattleyanum female gametophyte ontogeny. Flora 2017, 234, 34–40. [Google Scholar] [CrossRef]
  5. Porcher, E.; Lande, R. Reproductive compensation in the evolution of plant mating systems. New Phytol. 2005, 166, 673–684. [Google Scholar] [CrossRef] [Green Version]
  6. Mazzucato, A.; Falcinelli, M.; Veronesi, F. Evolution and adaptedness in a facultatively apomictic grass, Poa pratensis L. Euphytica 1996, 92, 13–19. [Google Scholar] [CrossRef]
  7. Giełwanowska, I.; Kellmann−Sopyła, W. Generative reproduction of Antarctic grasses, the native species Deschampsia antarctica Desv. and the alien species Poa annua L. Pol. Polar Res. 2015, 36, 261–279. [Google Scholar] [CrossRef] [Green Version]
  8. Ghanbari, M.A.; Salehi, H.; Jowkar, A. Genetic Diversity Assessment of Iranian Kentucky Bluegrass Accessions: II. Nuclear DNA Content and Its Association with Morphological and Geographical Features. Mol. Biotechnol. 2023, 65, 84–96. [Google Scholar] [CrossRef]
  9. Yang, Y.; Guo, X.; Wang, K.-l.; Liu, Q.-H.; Liu, Q.-C. Anther and ovule development in Clematis terniflora var. mandshurica (Ranunculaceae). Flora 2019, 253, 67–75. [Google Scholar] [CrossRef]
  10. Yudakova, O.I. Abnormalities of Female Gametophyte Development in Apomictic Bluegrass Forms. Russ. J. Dev. Biol. 2009, 39, 150–156. [Google Scholar] [CrossRef]
  11. Semerdjieva, I.; Sidjimova, B.; Jankova, E.; Kostova, M.; Jeliazkov, V. Study on Galanthus species in the Bulgarian flora. Heliyon 2019, 5, e03021. [Google Scholar] [CrossRef] [Green Version]
  12. Zhang, Q.; Hao, Q.; Guo, X.; Liu, Q.; Sun, Y.; Liu, Q.; Wang, K. Anther and ovule development in Camellia japonica (Naidong) in relation to winter dormancy: Climatic evolution considerations. Flora 2017, 233, 127–139. [Google Scholar] [CrossRef]
  13. Snyman, S.J.; Komape, D.M.; Khanyi, H.; van den Berg, J.; Cilliers, D.; Lloyd Evans, D.; Barnard, S.; Siebert, S.J. Assessing the Likelihood of Gene Flow from Sugarcane (Saccharum Hybrids) to Wild Relatives in South Africa. Front. Bioeng. Biotechnol. 2018, 6, 72. [Google Scholar] [CrossRef]
  14. Xu, D.; Mondol, P.C.; Uzair, M.; Tucker, M.R.; Zhang, D. Agrobacterium-Mediated Genetic Transformation, Transgenic Production, and Its Application for the Study of Male Reproductive Development in Rice. J. Vis. Exp. JVE 2020, 164, e61665. [Google Scholar] [CrossRef]
  15. Yang, X.; Wei, J.; Xia, J.; Fang, Q.; Zhang, B. Histochemical characteristics and differentiation of the belowground buds of Medicago archiducis-nicolai during overwintering. Pratacult. Sci. 2022, 39, 300–308. [Google Scholar] [CrossRef]
  16. Zhang, J.; Ma, H.; Liu, Y. Analysis on characteristics of female gametophyte and functional identification of genes related to inflorescences development of Kentucky bluegrass. Protoplasma 2021, 259, 1061–1079. [Google Scholar] [CrossRef]
  17. Ross, P.; Slovin, J.; Chen, C. A simplifed method for differential staining of aborted and non-aborted pollen grains. Int. J. Plant Biol. 2010, 1, 13. [Google Scholar] [CrossRef] [Green Version]
  18. Faure, J.E.; Aldon, D.; Rougier, M.; Dumas, C. Emerging data on pollen tube growth and fertilization in flowering plants. Protoplasma 1996, 193, 132–143. [Google Scholar] [CrossRef]
  19. Shukla, A.K.; Vijayaraghavan, M.R.; Chaudhry, B. Biology of Pollen; APH Publishing: Delhi, India, 1998. [Google Scholar]
  20. Quan, H.G.; Yu, K.L.; Hong, G.; Jin, Y.H.; Quan, X.L. Studies on the development of male gametophyte and pollen viability of datura stramonium. Agric. Sci. J. Yanbian Univ. 2021, 43, 25–30. [Google Scholar] [CrossRef]
  21. Palumbo, F.; Draga, S.; Vannozzi, A.; Lucchin, M.; Barcaccia, G. Trends in Apomixis Research: The 10 Most Cited Research Articles Published in the Pregenomic and Genomic Eras. Front. Plant Sci. 2022, 13, 878074. [Google Scholar] [CrossRef]
  22. Chahal, L.S.; Conner, J.A.; Ozias-Akins, P. Phylogenetically Distant BABY BOOM Genes from Setaria italica Induce Parthenogenesis in Rice. Front. Plant Sci. 2022, 13, 863908. [Google Scholar] [CrossRef] [PubMed]
  23. Xiong, H.Y.; Liu, Z.X. Mega- and Microsporogenesis and Development of Female and Male Gametophytes in Michelia maudiae Dunn. Bull. Bot. Res. 2018, 38, 212–217. [Google Scholar]
  24. Su, Q.D. Study on Rudimentary Panicle Development and Embryology in ‘Nassua’ Kentucky Bluegrass (Poa pratensis L.). Master’s Thesis, Huazhong Agricultural University, Wuhan, China, 2008. [Google Scholar]
  25. Tian, C.X.; Ma, H.L.; Zhang, Y.M. Embryo Types and Characteristics of Apomixis in Poa pratensis L. Sci. Agric. Sin. 2013, 46, 2633–2642. [Google Scholar] [CrossRef]
  26. Dumas, C. Developmental Biology of Flowering Plants, by Valayamghat Raghavan, Springer-Verlag, Berlin. ISBN 0-387-98781-9); DM 159.00. Plant Sci. 2000, 157, 267. [Google Scholar] [CrossRef]
  27. Liu, M.Q.; Wang, X.Q.; Luo, X.Z.; Dai, L.P.; Chen, S.Q. Microsporo genesis and development of male gametophyte of rabdosia rubescens (Hemsl.) Hara. Seed 2020, 39, 43–47. [Google Scholar] [CrossRef]
  28. Friedman, W.; Williams, J. Modularity of the angiosperm female gametophyte and its bearing on the early evolution of endosperm in flowering plants. Evol. Int. J. Org. Evol. 2003, 57, 216–230. [Google Scholar] [CrossRef]
  29. Brožová, V.; Koutecký, P.; Doležal, J. Plant apomixis is rare in Himalayan high-alpine flora. Sci. Rep. 2019, 9, 14386. [Google Scholar] [CrossRef] [Green Version]
  30. Solantzeva, M.V. Apomixis and hemigamy as one of its forms. Proc. Indian Natl. Sci. Acad. 1978, 44, 78–80. [Google Scholar]
  31. Young, B.A.; Sherwood, R.; Bashaw, E. Cleared-pistil and thick-sectioning techniques for detecting aposporous apomixis in grasses. Can. J. Bot.-Rev. Can. Bot. 1979, 57, 1668–1672. [Google Scholar] [CrossRef]
  32. Marshall, R.D.; Brown, A. The evolution of apomixis. Heredity 1981, 47, 1–15. [Google Scholar] [CrossRef] [Green Version]
  33. Savidan, Y. Apomixis: Genetics and Breeding; Wiley Publisher: Hoboken, NJ, USA, 2010; Volume 18, pp. 13–86. [Google Scholar]
  34. Miles, J. Apomixis for Cultivar Development in Tropical Forage Grasses. Crop Sci. 2007, 47, S238. [Google Scholar] [CrossRef]
  35. Albertini, E.; Barcaccia, G.; Porceddu, A.; Sorbolini, S.; Falcinelli, M. The mode of reproduction is detected by path1 and sex1 SCAR markers in a wide range of facultative apomictic Kentucky biuegrass varieties. Mol. Breed. 2001, 7, 293–300. [Google Scholar] [CrossRef]
  36. Yahara, K.; Horie, R.; Kobayashi, I.; Sasaki, A. Evolution of DNA double-strand break repair by gene conversion: Coevolution between a phage and a restriction-modification system. Genetics 2007, 176, 513–526. [Google Scholar] [CrossRef] [Green Version]
  37. Pepin, G.; Funk, C. Evaluation of Turf, Reproductive, and Disease-Response Characteristics in Crossed and Selfed Progenies of Kentucky Bluegrass1. Crop Sci. 1974, 14, 356–359. [Google Scholar] [CrossRef]
  38. Kondrashov, A. Classification of hypotheses on the advantage of amphimixis. J. Hered. 1993, 84, 372–387. [Google Scholar] [CrossRef] [Green Version]
  39. Galla, G.; Siena, L.A.; Ortiz, J.P.A.; Baumlein, H.; Barcaccia, G.; Pessino, S.C.; Bellucci, M.; Pupilli, F. A Portion of the Apomixis Locus of Paspalum Simplex is Microsyntenic with an Unstable Chromosome Segment Highly Conserved Among Poaceae. Sci. Rep. 2019, 9, 3271. [Google Scholar] [CrossRef] [Green Version]
  40. Liu, Y.; Zhang, J.Q.; Niu, K.J.; Dong, W.K.; Ma, H.L.; Li, Y.Z. Identification of apomictic characteristics of wild Kentucky bluegrass germplasm Resources in Gansu. Grassl. Turf. 2020, 40, 84–89. [Google Scholar] [CrossRef]
  41. Iudakova, O.I.; Shakina, T.N. Specific features of early embryogenesis in apomictic Poa pratensis L. Ontogenez 2007, 38, 5–11. [Google Scholar] [CrossRef]
  42. Zhang, J.Q.; Jia, X.F.; Li, F.; Li, Y.Z.; Ma, H.L. Effects of temperature and glume status on germination and polyembryonic seedling frequency in seven wild germplasm of Kentucky bluegrass native to Gansu. Grassl. Turf. 2021, 41, 70–77. [Google Scholar] [CrossRef]
  43. Takeuchi, Y.; Kosaza, M.; Ozaki, Y.; Tomiyoshi, K.; Matsuishi, T.; Okubo, H. Origin of polyembryonic seeds and production of haploids in asparagus. Acta Hortic. 2020, 1301, 57–66. [Google Scholar] [CrossRef]
  44. Ferreira, D.; Camargo, J.L.; Ferraz, I. Do polyembryonic seeds of Carapa surinamensis (Meliaceae) have advantages for seedling development? Acta Amaz. 2019, 49, 97–104. [Google Scholar] [CrossRef] [Green Version]
  45. Tan, F.; Chen, H.; Hu, H.L.; Liao, Y.H.; Hu, T.X.; Chen, Y.F.; Zhou, G.L.; Wang, X. A Study on the Correlation among Germination Rate, Polyembryony Rate, Seed Size of Phoebe Zhennan S. Lee. J. Agric. Univ. 2018, 36, 640–647. [Google Scholar] [CrossRef]
  46. Reshma, U.R.; Simi, S. Screening of Mango Landraces for Polyembryony and Confirmation of Seedling Origin using Microsatellite Markers. Agric. Sci. Dig. 2021, 42, 128–136. [Google Scholar] [CrossRef]
  47. Kishore, K.; Monika, N.; Rinchen, D.; Lepcha, B.; Pandey, B. Polyembryony and seedling emergence traits in apomictic citrus. Sci. Hortic. 2012, 138, 101–107. [Google Scholar] [CrossRef]
  48. Mendes-Rodrigues, C.; Sampaio, D.; Costa, M.; Caetano, A.; Ranal, M.; Bittencourt Junior, N.; Oliveira, P. Polyembryony increases embryo and seedling mortality but also enhances seed individual survival in Handroanthus species (Bignoniaceae). Flora-Morphol. Distrib. Funct. Ecol. Plants 2012, 207, 264–274. [Google Scholar] [CrossRef]
  49. Mendes-Rodrigues, C.; Oliveira, P.E. Polyembryony in Melastomataceae from Brazilian Cerrado: Multiple embryos in a small world. Plant Biol. 2012, 14, 845–853. [Google Scholar] [CrossRef]
Agronomy 13 01569 g001 550
Figure 1. The KI–I2 staining of the Kentucky bluegrass pollens represents the viable pollen grains with dark brown dye and nonviable pollen grains with yellow dye. For example, the red box-type pollen indicates that the pollen has vitality, while the blue box-type pollen indicates that the pollen has no vitality. (A) The wild Kentucky bluegrass collected from Gannan (GN); (B) The wild Kentucky bluegrass collected from Longnan (LN).
Figure 1. The KI–I2 staining of the Kentucky bluegrass pollens represents the viable pollen grains with dark brown dye and nonviable pollen grains with yellow dye. For example, the red box-type pollen indicates that the pollen has vitality, while the blue box-type pollen indicates that the pollen has no vitality. (A) The wild Kentucky bluegrass collected from Gannan (GN); (B) The wild Kentucky bluegrass collected from Longnan (LN).
Agronomy 13 01569 g001
Agronomy 13 01569 g002 550
Figure 2. The microsporogensis and microgametogenesis development of Kentucky bluegrass. (A) Horizontal section of floret; (B) Longitudinal section of floret; (C,D) Formation of the primary cell wall and primary sporogenic cell; (E) Secondary sporulation cells and primary wall cells form anther triple wall; (F,G) The dyad period; (H) Period of coexistence of the dyad and the tetrad; (I) The tetrad period; (J,K,M) Mononuclear pollen period; (L,N) The dinuclear pollen period; (O–S) The period of mature anthers; (T) Longitudinal section of mature anthers. The black arrows represent important structures at different developmental stages in each subfigure.
Figure 2. The microsporogensis and microgametogenesis development of Kentucky bluegrass. (A) Horizontal section of floret; (B) Longitudinal section of floret; (C,D) Formation of the primary cell wall and primary sporogenic cell; (E) Secondary sporulation cells and primary wall cells form anther triple wall; (F,G) The dyad period; (H) Period of coexistence of the dyad and the tetrad; (I) The tetrad period; (J,K,M) Mononuclear pollen period; (L,N) The dinuclear pollen period; (O–S) The period of mature anthers; (T) Longitudinal section of mature anthers. The black arrows represent important structures at different developmental stages in each subfigure.
Agronomy 13 01569 g002
Agronomy 13 01569 g003 550
Figure 3. Opening process of anther in Kentucky bluegrass. (A) Intact mature anther; (B,C) Dehiscence on one side of anther; (D) Dehiscence on both sides of anther.
Figure 3. Opening process of anther in Kentucky bluegrass. (A) Intact mature anther; (B,C) Dehiscence on one side of anther; (D) Dehiscence on both sides of anther.
Agronomy 13 01569 g003
Agronomy 13 01569 g004 550
Figure 4. External morphology of intact ovary in Kentucky bluegrass. Figure (A–C) are three different ovaries.
Figure 4. External morphology of intact ovary in Kentucky bluegrass. Figure (A–C) are three different ovaries.
Agronomy 13 01569 g004
Agronomy 13 01569 g005 550
Figure 5. Cyto-embryological observation of apomictic ovule of Kentucky bluegrass. (A) The stage of formation of the archesportial cell and nucellar cell specialization; (B) The stage of coexistence of the archesportial cell and specialized nucellar cell; (C) The period of coexistence of the megaspore mother cell and apospore initiation cell; (D,E) The period of coexistence of the apospore initiation cell and dyad; (F) The coexistence period of two apospore initiation cells; (G) The coexistence period of two apospore initiation cells and megaspore mother cell; (H) The period of coexistence of apospore-initiating cell and dyad; (I) The stage of formation of the two-nucleus embryo sac by apomixis; (J–L) The period of the multi-nucleus embryo sac. The black arrows represent important structures at different developmental stages in each subfigure.
Figure 5. Cyto-embryological observation of apomictic ovule of Kentucky bluegrass. (A) The stage of formation of the archesportial cell and nucellar cell specialization; (B) The stage of coexistence of the archesportial cell and specialized nucellar cell; (C) The period of coexistence of the megaspore mother cell and apospore initiation cell; (D,E) The period of coexistence of the apospore initiation cell and dyad; (F) The coexistence period of two apospore initiation cells; (G) The coexistence period of two apospore initiation cells and megaspore mother cell; (H) The period of coexistence of apospore-initiating cell and dyad; (I) The stage of formation of the two-nucleus embryo sac by apomixis; (J–L) The period of the multi-nucleus embryo sac. The black arrows represent important structures at different developmental stages in each subfigure.
Agronomy 13 01569 g005
Agronomy 13 01569 g006 550
Figure 6. Cyto-embryological observation of double fertilization process in Kentucky bluegrass. (A) Pollen germinates on the stigma and pollen tube grows; (B) Egg cells and polar nuclei; (C) Fertilization of egg cell and synergid degeneration; (D) Two polar nuclei were fused to one secondary nucleus; (E,F) From the same floret, double fertilization process; (G) Double fertilization process; (H) The polar nucleus predates fertilization of the egg cells; (I–L) Embryo formation predates polar-nuclear fertilization; (M) The coexistence of sexual reproduction and apomixis; (N,O) The polar nucleus is fertilized, and the zygote is dividing; (P) Four-cell protoembryo; (Q) Eight-cell protoembryo; (R) Pear-shaped embryo; (S) Endosperm cell. The black arrows represent important structures at different developmental stages in each subfigure.
Figure 6. Cyto-embryological observation of double fertilization process in Kentucky bluegrass. (A) Pollen germinates on the stigma and pollen tube grows; (B) Egg cells and polar nuclei; (C) Fertilization of egg cell and synergid degeneration; (D) Two polar nuclei were fused to one secondary nucleus; (E,F) From the same floret, double fertilization process; (G) Double fertilization process; (H) The polar nucleus predates fertilization of the egg cells; (I–L) Embryo formation predates polar-nuclear fertilization; (M) The coexistence of sexual reproduction and apomixis; (N,O) The polar nucleus is fertilized, and the zygote is dividing; (P) Four-cell protoembryo; (Q) Eight-cell protoembryo; (R) Pear-shaped embryo; (S) Endosperm cell. The black arrows represent important structures at different developmental stages in each subfigure.
Agronomy 13 01569 g006
Agronomy 13 01569 g007 550
Figure 7. Histological observation of seeds of Kentucky bluegrass. (A–D) A whole seed at different development periods of the embryo; (E) Complete structure of the embryo; (F–I) Two embryo seed; (J) Three embryo seed; (K) Synchronous development of two embryos in the same seed; (L) The absence of simultaneous development of the two embryos in the same seed; (M) Disintegration of one embryo in the three embryo seed. GL: Glume; SP: seed coat and peel; EN: endosperm; AL: aleurone layer; EM: embryo; COT: cotyledon; COL: coleoptile; PL: plumule; EMA: embryonal axis; RA: radicle.
Figure 7. Histological observation of seeds of Kentucky bluegrass. (A–D) A whole seed at different development periods of the embryo; (E) Complete structure of the embryo; (F–I) Two embryo seed; (J) Three embryo seed; (K) Synchronous development of two embryos in the same seed; (L) The absence of simultaneous development of the two embryos in the same seed; (M) Disintegration of one embryo in the three embryo seed. GL: Glume; SP: seed coat and peel; EN: endosperm; AL: aleurone layer; EM: embryo; COT: cotyledon; COL: coleoptile; PL: plumule; EMA: embryonal axis; RA: radicle.
Agronomy 13 01569 g007
Agronomy 13 01569 g008 550
Figure 8. Polyembryo seedling of wild germplasm materials of Kentucky bluegrass in Gansu Province. (A) One-embryo seedling; (B–J) Two-embryo seedling; (K,L) Three-embryo seedling. The bar for all figures was the same as L.
Figure 8. Polyembryo seedling of wild germplasm materials of Kentucky bluegrass in Gansu Province. (A) One-embryo seedling; (B–J) Two-embryo seedling; (K,L) Three-embryo seedling. The bar for all figures was the same as L.
Agronomy 13 01569 g008
Table
Table 1. Pollen traits and stigma receptivity of wild Kentucky bluegrass collected from Gannan (GN) and Longnan (LN).
Table 1. Pollen traits and stigma receptivity of wild Kentucky bluegrass collected from Gannan (GN) and Longnan (LN).
MaterialsPollen Diameter//μmPollen ShapePollen ViabilityStigma Receptivity
MaxMinMeanRoundOval
GN30.3616.2423.12 ± 1.12 a83.33% ± 3.33 a16.67% ± 3.33 a89.13% ± 1.92 a80.00% ± 4.41 a
LN28.4215.6521.77 ± 1.03 a76.67% ± 3.33 a23.33% ± 3.33 a85.38% ± 1.97 a82.00% ± 5.00 a
Note: Different letters indicate statistical significance where the same letter indicates no significant difference between different materials, according to independent samples t-test (p < 0.05).
Table
Table 2. Statistics of germination-related indicators of wild Kentucky bluegrass collected from Gannan (GN) and Longnan (LN).
Table 2. Statistics of germination-related indicators of wild Kentucky bluegrass collected from Gannan (GN) and Longnan (LN).
MaterialsGermination PotentialGermination RateOne-Embryo Seedling RateTwo-Embryo Seedling RateThree-Embryo Seedling RatePolyembryony Seedling Rate
GN51.60% ± 1.63 a83.10% ± 0.98 a100% ± 0.00 a0 ± 0.00 b0 ± 0.00 b0 ± 0.00 b
LN53.40% ± 1.67 a79.00% ± 2.24 b92.15% ± 0.99 b7.72% ± 1.00 a0.13% ± 0.11 a7.85% ± 0.99 a
Note: Different letters indicate statistical significance where the same letter indicates no significant difference between different materials, according to independent samples t-test (p < 0.05).
Disclaimer/Publisher’s Note: The statements, opinions and data contained in all publications are solely those of the individual author(s) and contributor(s) and not of MDPI and/or the editor(s). MDPI and/or the editor(s) disclaim responsibility for any injury to people or property resulting from any ideas, methods, instructions or products referred to in the content.

Share and Cite

MDPI and ACS Style

Zhang, J.; Ma, H. Cyto-Embryological Analysis of Wild Kentucky Bluegrass Germplasm in Gansu Province, China. Agronomy 2023, 13, 1569. https://doi.org/10.3390/agronomy13061569

AMA Style

Zhang J, Ma H. Cyto-Embryological Analysis of Wild Kentucky Bluegrass Germplasm in Gansu Province, China. Agronomy. 2023; 13(6):1569. https://doi.org/10.3390/agronomy13061569

Chicago/Turabian Style

Zhang, Jinqing, and Huiling Ma. 2023. "Cyto-Embryological Analysis of Wild Kentucky Bluegrass Germplasm in Gansu Province, China" Agronomy 13, no. 6: 1569. https://doi.org/10.3390/agronomy13061569

APA Style

Zhang, J., & Ma, H. (2023). Cyto-Embryological Analysis of Wild Kentucky Bluegrass Germplasm in Gansu Province, China. Agronomy, 13(6), 1569. https://doi.org/10.3390/agronomy13061569

Note that from the first issue of 2016, this journal uses article numbers instead of page numbers. See further details here.

Article Metrics

Back to TopTop