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Brief Communication |
Department of Biological Sciences, University of Toronto Scarborough, 1265 Military Trail, Scarborough, Ontario M1C 1A4 Canada
Received for publication April 12, 2007. Accepted for publication October 25, 2007.
ABSTRACT
We developed an improved cytological protocol for producing high quality, light microscope images of plant meiotic chromosomes. Because the technique works on species with small genomes and thick microsporocyte cell walls, it should be useful for studying the wild relatives of Arabidopsis and other eudicots with small genomes. Combining this improved fixation protocol with our new analysis of associated substages in floral buds, we can unambiguously assign individual meiotic cells to particular substages of prophase I in Arabidopsis thaliana, even for difficult distinctions such as that between late zygotene or early diplotene. In this report we provide the first estimate of the individual duration of the zygotene and pachytene substages (4.8 h and 10.0 h, respectively) in A. thaliana. We also have examined the diffuse substage of prophase I and report that during this post-pachytene substage, nuclei retain the association of homologous nucleolus organizer regions and homologous centromeres, despite the generally diffuse chromatin and generally unpaired chromosome regions. Additionally, we have observed that centromeric regions of the chromosomes of diffuse-stage nuclei are highly condensed, more so than those of any other substage of prophase I.
Key Words: Arabidopsis thaliana (Brassicaceae) • diffuse stage • duration pachytene • duration zygotene • floral bud substage-association staging • improved cytology • meiosis
During meiosis, homologous chromosomes come together, undergo reciprocal genetic exchange, and form chiasmata. The chiasmata function as ties that hold homologous chromosomes together so they can correctly co-orient to opposite poles for the first meiotic division. The reciprocal genetic exchanges are accomplished through the enzymatic events of the double strand break (DSB) pathway (Villeneuve and Hillers, 2001
). The coordination of the steps in the DSB pathway with other chromosomal events, such as alignment of homologous chromosome regions (Storlazzi et al., 2003
), homologous chromosome recognition (vs. homeologous [Wilson et al., 2005
] or even non homologous associations [Higgins et al., 2005
]), and chromosome cohesion (Bolcun-Filas et al., 2007
) and condensation, requires careful orchestration of the action and interaction of a variety of gene products and chromatin regions.
Arabidopsis thaliana is one of the important genetic models for the study of plant meiosis. Genetic analyses of Arabidopsis meiotic mutants are extensive (e.g., Bai et al., 1999
; Couteau et al., 1999
; Azumi et al., 2002
; Cai et al., 2003
; Caryl et al., 2003
; Mercier et al., 2003
; Schwarzacher, 2003
). Light microscopic cytology (Ross et al., 1996
) in conjunction with fluorescence in situ hybridization (Lysak et al., 2001
) and immunocytochemistry (Armstrong et al., 2002
) has been used to analyze the wild type and mutants. Additionally species hybrids and induced allopolyploids (Comai et al., 2000
) can readily be created between A. thaliana and its wild relatives, presenting the opportunity to study the role of individual meiotic genes in "enforcing" chromosome homology and constraining reciprocal genetic exchange.
We report here an improved light microscopic cytological technique for studying meiosis in Arabidopsis and unambiguously identifying all substages of prophase I. Our protocol allows us to readily distinguish hard to identify stages such as zygotene, early diplotene, and the diffuse stage. We also have used our technique to confirm the estimate by Armstrong et al. (2003)
of the relative length of leptotene and to determine for the first time the duration of the zygotene and pachytene substages. We also report new features of the diffuse stage of prophase I.
MATERIALS AND METHODS
Plant material
Seeds of A. thaliana cv. Landsberg erecta (Ler) were obtained from the Arabidopsis Biological Resource Center (http://www.arabidopsis.org/abrc). Plants were grown under long-day conditions (16 h light/8 h dark); the day/night temperatures were 22°/19°C. Relative humidity was constant at approximately 55%. Primary inflorescences were harvested and fixed within 5 d after the opening of the first flower.
Preparation of unfixed anthers for meiotic stage analysis
Buds of desired size were placed in 10 mM sodium citrate (pH 4.0) in the cavity of a depression slide; sepals were teased apart with a scalpel, releasing the anthers. Individual anthers were given two parallel cuts and then transferred to 20 µL of 4'6-diamidino-2-phenyl indole (DAPI) in Vectashield antifade mounting medium (Vector Laboratories, Burlingame, California USA) placed at the center of a clean glass slide. A glass coverslip was carefully applied, and its edges were sealed with nail polish. The coverslip was then gently tapped to extrude meiocytes from the cut anthers.
Preparation of fixed anthers and chromosome spreads
The primary inflorescences were harvested when the first, or first and second, flower buds opened, by placing the inflorescences into a small vial containing tap water. When the harvest was completed, the water was exchanged for freshly prepared 4% paraformaldehyde (PFA) (pH 8.2). Buds were gently agitated in PFA for 15 min. The PFA was then exchanged with a mixture of methanol : glacial acetic acid (3 : 1), and the buds were again gently agitated for 15 min. The 3 : 1 fixative was replaced with fresh fixative twice after a 15-min agitation. At this point, the inflorescences appeared white. The inflorescences were then stored in the acidified methanol at 4°C until needed. Storage time did not exceed 1 mo.
Primary inflorescences were transferred from 3 : 1 acidified methanol into the cavity of a depression slide containing tap water (pH 7.0). The inflorescences were repeatedly rinsed with tap water until the pH of the water in the depression slide cavity was 
7 (pH test strips were used). The larger yellow (pollen containing) buds were discarded, and the remaining buds were rinsed a few more times with tap water. Buds in the appropriate size range (0.3–0.6 mm) were cut away from the inflorescences. The tap water was exchanged with 10 mM sodium citrate buffer (pH 5.3) and then subsequently exchanged with 200 µL of cell wall digestion enzyme mixture (0.5% pectinase, 0.5% hemicellulase, 0.5% cellulose, 0.5% cytohelicase, 0.5% polyvinylpyrrolidone MW 40 000 [PVP 40] dissolved in citrate buffer [pH 5.3]). The depression slide with the buds then was placed in a humid chamber, and the chamber was placed in an oven at 37°C for 3 h. After the bud digestion, the enzyme mixture was exchanged with ice cold bud-stabilizing buffer (1000 µL 10 mM sodium citrate [pH 7.0], 10 µL 500 mM ethylene glycol bis(2-aminoethyl ether)-N,N,N'N'-tetraacetic acid [EGTA] and 1/8 protease inhibitor tablet [Roche Diagnostic, Indianapolis, Indiana USA]), and the depression slide was placed in a covered petri dish sitting on ice. Using a fine wire loop and a dissecting scope, we selected and transferred a bud to another depression slide containing freshly prepared 4% PFA (pH 8.2) for 60 s. The bud was then immediately transferred into 5 µL of 45% acetic acid situated at the center of a clean glass slide. The bud was gently prodded (usually less than 1 min) to facilitate its dispersal in the acid, and then the nuclear suspension was immediately dried with a hair dryer (on low setting) positioned 30 cm from the specimen. DAPI (10 µg/mL) in Vectashield antifade mounting medium was then applied to the spread area, a coverslip was applied, and the edges of the coverslip were sealed with nail polish. The slides were examined by fluorescence microscopy using a Zeiss (Toronto Ontario, Canada) Axiophot epifluorescence microscope with filter set 2 for DAPI (G365/ST395/LP420) and a plan achromat 100x oil immersion lens. Images were captured with a cooled charge coupled device camera (Empix Imaging, Mississauga, Ontario, Canada) and processed with Northern Eclipse software (Empix Imaging).
Collection of floral bud substage-association data
The minimum number of chromosome spreads per bud (per slide) included in the substage-association analysis that would be representative of the large and smaller anther meiocytes within that bud was determined to be 25 using the statistical sampling formula: n = Nx/[(N + 1)E2 + x]; where n = sample size (number of chromosome spreads obtained per bud), N = population size (total number of meiocytes per bud), E = error (20%), and x = confidence level (95%) ([Danesoft [London, UK] statistical software). The population size (N) of the meiocytes per bud was calculated to be 720 (six anthers per bud x four locules per anther x 30 meiocytes per locule). Slides were not included in the analysis if all the chromosome spreads appeared to come from a single anther.
RESULTS AND DISCUSSION
In a preliminary study, meiosis was examined in unfixed, DAPI-stained, gently squashed, individual anthers of single buds. Each bud has six anthers, four larger anthers and two smaller anthers. Each anther has four chambers. The meiocytes within a single chamber were always at the same stage; meiocytes in different chambers of the same anther were usually, but not always, at the same stage. While the four larger anthers were usually at the same meiotic stage, the smaller anthers often were at an earlier meiotic stage (Appendix S1 [see Supplemental data accompanying online version of the article]). Thus, there is often some asynchrony within an individual bud.
Substage associations in floral buds
The slight asynchrony observed in individual buds, in conjunction with an improved fixation protocol, was used to identify the substages of prophase I that were found within the same flower bud, thereby defining the substage associations in the floral buds. For identifying the floral bud substage, 59 wild-type buds were collected and analyzed. Each bud was treated as indicated in the Materials and Methods and placed on a single microscope slide. Only slides with at least 25 recognizable meiocytes were included in the bud substage analysis; in total, 3073 nuclei were examined. For each bud, we noted which meiotic substages were found together; the associated substages are summarized in Table 1.
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Thus with knowledge gained from previous fluorescence in situ hybridization (FISH) studies (Armstrong et al., 2001
; Armstrong and Jones, 2003
), our high quality and detailed images, and substage associations, we can present a complete sequence of prophase I substages in the bud as seen in Fig. 1. A loose bouquet is seen in late leptotene (Armstrong et al., 2001
) and persists during zygotene, where there is an asymmetric distribution of the chromosomes and nucleolus to one side of the nucleus. The homologous chromosomes become associated during zygotene, with the nuclei containing a mixture of paired and unpaired chromosome regions; the centromere regions of all five pairs of chromosomes are aggregated, forming one or two chromocenters (Fig. 1D, E). By late zygotene (Fig. 1F), homologous chromosomes are extensively aligned along most of their length. In early pachytene, the chromosomes are maximally paired; the centromere regions, while paired with their homolog, are no longer all aggregated in chromocenters (Fig. 1G). By late pachytene, the five paired centromere regions appear as five distinct DAPI–bright regions. The chromosomes in general are uniformly distributed within the spread, except the nucleolus organizer regions (NORs) of chromosomes 2 and 4 are usually still associated (Fig. 1H).
By early diplotene, small regions of desynapsis begin to occur in noncentromeric regions of the chromosome spreads (Fig. 1I). Using FISH probes, Armstrong and Jones (2003)
showed that centromere regions of homologous chromosomes remain associated throughout diplotene. Our improved cytological protocol and substage associations allow us to confirm this feature without the use of FISH probes. Thus, our relatively simple fixation protocol makes it is possible to see that, excepting chiasmata themselves, the centromere regions of homologous chromosomes are the last regions to desynapse.
By mid-diakinesis, the chromosomes have condensed, and five distinct, chiasmate bivalents are apparent (Fig. 1L). At metaphase I, all five bivalents are distinct, and at anaphase I, the chromosomes have segregated properly to opposite sides of the metaphase I plate (data not shown).
Duration of meiotic stages
We examined the stages of meiosis in a set of primary inflorescences of wild-type Arabidopsis. Inflorescences were selected that had only one or two open flowers so that all meiotic stages would be represented (Smyth et al., 1990
). Meiosis I stages were found in buds that ranged from 0.39 to 0.50 mm, ±0.01 mm). To avoid sampling bias, we processed all buds in this size range, with a size distribution as follows (with the number of buds given in parentheses): 0.39 mm (6), 0.40 mm (5), 0.41 mm (5), 0.42 mm (4), 0.43 mm (4), 0.44 mm (5), 0.45 mm (6), 0.46 mm (4), 0.47 mm (5), 0.48 mm (4), 0.49 mm (6), and 0.50 mm (5). For each bud used in the analysis, the number of meiocytes at each meiotic substage was recorded. In total, 3073 meiocytes from wild type were included. We used the percentage of the total meiocytes at a particular stage to estimate the relative duration of each stage (Fig. 2). From this, we estimated that leptotene would be only slightly longer in duration than zygotene, and each would be about half as long as the pachytene stage.
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used bromodeoxyuridine (BrdU) labeling at S phase, combined with average durations to the appearance of BrdU labeling in the various meiotic stages, to estimate the duration of some of the substages of prophase I in A. thaliana. They estimated that leptotene lasts an average of 6 h. They were unable to distinguish between the individual durations of zygotene and pachytene but did conclude that the combined stages of zygotene/pachytene lasted an average of 15.3 h. They also estimated the total duration of meiosis to be 24 h, of which the pooled substages of zygotene and pachytene constituted 63.7% of the total. When we combined the number of nuclei in our study in both zygotene and pachytene substages (to make them comparable to the pooled zygotene/pachytene sample of Armstrong et al.), our results agreed well with theirs for the relative durations of leptotene vs. the combined stages of zygotene/pachytene.
However, in our study we were able to distinguish zygotene from pachytene preparations, and we thus have data on the relative numbers of each of these two substages and the percentage of total meiocytes that each substage represents. By using our data and combining it with the estimate of Armstrong et al. (2003)
that meiosis lasts 24 h in A. thaliana, we provided the first estimate of the separate durations of the zygotene and pachytene stages. Zygtotene nuclei represent 20.1% of all meiocytes; if meiosis lasts 24 h, we estimated the zygotene substage stage to be 4.8 h. Similarly, because pachytene nuclei represent 41.9% of the total meiocytes, we estimated the pachytene substage to last 10 h.
Our estimates of the duration of the interval spanning from diplotene- metaphase I are similar to that of Armstrong et al. (2003)
, but we estimated that diplotene is longer than diakinesis/metaphase I combined. In contrast, Armstrong et al. (2003)
found diplotene to be shorter than diakinesis/metaphase I. This relatively small difference might be due to our classifying diffuse stage nuclei as part of diplotene.
The diffuse stage
Our substage-association analysis and improved fixation protocol have allowed us to produce high quality images of the meiotic substage known as the diffuse stage (Fig. 1K) and to document a novel feature of chromosome centromeres at this stage. The diffuse stage has been reported in passing many times in many organisms (e.g., Moens, 1968
; Klasterska and Ramel, 1980
). In typical acid-alcohol preparations, the diffuse stage appears as an ill-defined stage in which the chromosomes are not highly condensed and often have sticky and weblike connections to each other. In our preparations, a PFA fixation both proceeds and follows the acetic alcohol treatment, yielding superior retention of certain chromosomal features and producing striking, diffuse stage images. At the diffuse stage, despite an overall loss of pairing of the homologous chromosome axes, the NORs of chromosomes 2 and 4 remain associated with each other, and the centromeres of homologous chromosomes are still associated in the region of their centromeres (Fig. 1K). More striking is that, despite an overall reduction in the level of chromosome condensation, the centromere regions actually appear to be in a more condensed state than at any other prophase I stage. The paired centromeres are very thick and ovoid (Fig. 1K) and are more condensed and rounded than even the subsequent stage of diakinesis.
The diffuse stage may be a time of chromatin remodeling (Qureshi and Hasenkampf, 1995
), a part of the chromosomal transition from mildly condensed, recombinogenic structures to the more highly condensed metaphase I chromosomes ready to segregate. This stage could also be a time to resolve any residually interlocked chromosomes. Retention of the association of homologous chromosomes at the NORs and centromeres and increased condensation in the centromere regions may help ensure that important homologous chromosome associations are maintained during this period of chromosome "remodeling."
In conclusion, we have provided detailed pictorial documentation of the chromosome morphology during prophase I of meiosis in wild-type A. thaliana. Our improved cytology and substage-association technique allow an easy and unambiguous identification of all of the substages of prophase I. Our protocol has enabled us to provide the first estimates of the durations of the important meiotic substages of zygotene and pachytene and has revealed several interesting features of the diffuse stage of meiosis, raising questions about what events might occur in this relatively overlooked meiotic stage. Our improved chromosome preservation and substage-association technique for identifying all meiotic substages will be particularly useful in studies of new meiotic mutants. As well, this improved cytological procedure will be useful in assessing chromosome pairing and reciprocal genetic exchange in wild relatives, species hybrids, and synthetic and natural polyploids of Arabidopsis. Because of the wealth of widely conserved meiotic genes identified in Arabidopsis, this improved cytological procedure can be used to determine the effect of specific meiotic genes on homologous, homeologous, and nonhomologous chromosome pairing and reciprocal genetic exchange.
FOOTNOTES
1 The authors acknowledge support from The National Science and Engineering Research Council of Canada to C.A.H. and the University of Toronto School of Graduate Studies to P.E.S. 
2 Author for correspondence (e-mail: hasenkampf{at}utsc.utoronto.ca
)
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