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. 2012 Nov 9;161(1):196–209. doi: 10.1104/pp.112.206656

Deficiency in a Very-Long-Chain Fatty Acid β-Ketoacyl-Coenzyme A Synthase of Tomato Impairs Microgametogenesis and Causes Floral Organ Fusion1,[W]

Anna Smirnova 1,*, Jana Leide 1, Markus Riederer 1
PMCID: PMC3532251  PMID: 23144186

Abstract

Previously, it was shown that β-ketoacyl-coenzyme A synthase ECERIFERUM6 (CER6) is necessary for the biosynthesis of very-long-chain fatty acids with chain lengths beyond C28 in tomato (Solanum lycopersicum) fruits and C26 in Arabidopsis (Arabidopsis thaliana) leaves and the pollen coat. CER6 loss of function in Arabidopsis resulted in conditional male sterility, since pollen coat lipids are responsible for contact-mediated pollen hydration. In tomato, on the contrary, pollen hydration does not rely on pollen coat lipids. Nevertheless, mutation in SlCER6 impairs fertility and floral morphology. Here, the contribution of SlCER6 to the sexual reproduction and flower development of tomato was addressed. Cytological analysis and cross-pollination experiments revealed that the slcer6 mutant has male sterility caused by (1) hampered pollen dispersal and (2) abnormal tapetum development. SlCER6 loss of function provokes a decrease of n- and iso-alkanes with chain lengths of C27 or greater and of anteiso-alkanes with chain lengths of C28 or greater in flower cuticular waxes, but it has no impact on flower cuticle ultrastructure and cutin content. Expression analysis confirmed high transcription levels of SlCER6 in the anther and the petal, preferentially in sites subject to epidermal fusion. Hence, wax deficiency was proposed to be the primary reason for the flower fusion phenomenon in tomato. The SlCER6 substrate specificity was revisited. It might be involved in elongation of not only linear but also branched very-long-chain fatty acids, leading to production of the corresponding alkanes. SlCER6 implements a function in the sexual reproduction of tomato that is different from the one in Arabidopsis: SlCER6 is essential for the regulation of timely tapetum degradation and, consequently, microgametogenesis.

Terrestrial plants have developed the cuticle, a barrier that restrains water movements between the outer cell wall of the epidermis and the surrounding atmosphere, thus preventing uncontrolled water loss (Riederer, 2006; Samuels et al., 2008; Domínguez et al., 2011). Besides this, the cuticle protects the plant from biotic stresses and prevents ectopic interorgan fusion during flower development (Lolle et al., 1992; Lolle and Pruitt, 1999; Tanaka and Machida, 2006; Pollard et al., 2008; Javelle et al., 2011). The cuticle is a complex matrix of cutin, waxes, and polysaccharides. Cutin is a biopolymer consisting of C16 and C18 fatty acids with a varying degree of oxygenation (Pollard et al., 2008; Domínguez et al., 2011). Cuticular wax is a general term for a complex mixture of hydrophobic compounds, including homologous series of very-long-chain aliphatic lipids. The wax composition varies not only among different species but also from one organ to another and throughout the course of plant development. Environmental conditions also have an impact on the wax composition (Samuels et al., 2008). Therefore, the mechanisms by which plants control wax biosynthesis are of considerable interest.

Long-chain acyl-coenzyme A synthetases (LACS) activate the precursors of wax biosynthesis, C16 and C18 fatty acids, on the outer membrane of the plastid. The resulting long-chain acyl-CoAs undergo elongation to very-long-chain fatty acids (VLCFAs) in the endoplasmic reticulum. VLCFA biosynthesis might be a major metabolic route in the epidermis, since VLCFAs are the primary compounds for all other wax components (Kunst et al., 2006; Samuels et al., 2008). The initial condensation reaction catalyzed by β-ketoacyl-CoA synthases was shown to be rate limiting in the elongation process. β-Ketoacyl-CoA synthases form a family of proteins that are thought to have different substrate specificities (Millar and Kunst, 1997; Joubès et al., 2008). To date, three of the condensing enzymes were shown to be involved in cuticle formation in Arabidopsis (Arabidopsis thaliana) shoots: KETOACYL-COENZYME A SYNTHASE1 (KCS1; Todd et al., 1999), FIDDLEHEAD (FDH; Lolle et al., 1992; Pruitt et al., 2000; Voisin et al., 2009), and ECERIFERUM6 (CER6 [also known as CUT1 or KCS6]; Jenks et al., 1995; Millar et al., 1999; Fiebig et al., 2000; Hooker et al., 2002). Furthermore, KCS20 and KCS2/DAISY were recently described as redundant condensing enzymes implicated in VLCFA biosynthesis in shoots (Lee et al., 2009).

The importance of the contribution of cutin and cuticular waxes in cell-cell interactions remains unclear (Javelle et al., 2011). A number of cuticular mutants were isolated, but only some of them showed postgenital organ fusion, joining organ primordia after their inception at the shoot apical meristem (Lolle et al., 1998). Among Arabidopsis mutants deficient in waxes, cer10, cer13, wax1, wax2/cer3/yre, and deadhead exhibited the organ fusion phenotype (Jenks et al., 1995, 1996; Lolle et al., 1998; Yephremov et al., 1999; Zheng et al., 2005). Another contact-mediated developmental process is the pollen-stigma interaction. CER1, CER3, CER6, and CER10, which contribute to the biosynthesis and/or deposition of cuticular waxes in Arabidopsis, were shown to be required for early pollen recognition on the stigma (Preuss et al., 1993; Aarts et al., 1995; Hülskamp et al., 1995; Fiebig et al., 2000). Mutations in any of these genes, besides the waxless shoot, resulted in conditional sterility. The mutants produced viable pollen, but it failed to hydrate on the dry stigma because of altered pollen coat lipids. Under conditions of high humidity, fertility was restored. The same phenotype was observed in the lacs1 lacs4 double mutant (Jessen et al., 2011). Formation of the sporopolleninic pollen exine is another aspect of the involvement of some enzymes contributing to the biosynthesis of cuticle components in sexual plant reproduction (Ariizumi et al., 2003; Jung et al., 2006; Zhang et al., 2008; Shi et al., 2011).

A substantial part of the data presented so far were obtained from studies on Arabidopsis. This species, however, has a very thin cuticle, and its cutin composition is rather different from other plants (Bonaventure et al., 2004). Our knowledge of wax biosynthesis in other model species commonly used in cuticle research is limited. Tomato (Solanum lycopersicum) is an important horticultural crop and a valuable tool for plant research. Previously, the use of a slcer6 tomato mutant (formerly lecer6) allowed the demonstration of a direct relationship between the cuticular transpiration barrier properties and modification of the cuticular wax composition during fruit development (Leide et al., 2007). Interestingly, besides alterations in cuticular waxes, the mutation in SlCER6 impacts flower organ morphology and fertility. Reasons for the reduced fertility in the slcer6 mutant could not be the same as for atcer6 plants because tomato has a wet stigma and, consequently, a different system of pistil-pollen interaction (Heslop-Harrison, 1977). In contrast to atcer6, the slcer6 mutant demonstrates flower organ fusion and thus appears to be a perfect model for investigating cuticle integrity regulation. The study presented in this paper shows the contribution of SlCER6, previously only known to be involved in cuticle formation in vegetative organs, to the sexual reproduction and flower development in tomato.

RESULTS

Flower Phenotype of the slcer6 Mutant

The MicroTom wild-type flower contains five sepals alternating with five petals and a pistil surrounded by five anthers coupled by epidermal hairs into an anther cone. At the time of anthesis, the wild-type flowers opened while the slcer6 flowers stayed closed (Fig. 1A). In the wild type, anthers were opened by longitudinal slits that serve for anther desiccation and pollen release, whereas the slcer6 anthers remained closed (Fig. 1B). Stomium cells degenerated in slcer6 without visible disorders. However, due to fusion between the external anther walls, the mutant pollen could only move to the neighbor locule but failed to escape from the anther cone (Fig. 1C). The petals were fused with each other and partly attached to the anther epidermis (Fig. 1, D–F). Thus, the slcer6 flowers demonstrated reproducible petal-to-petal, petal-to-anther, and anther-to-anther epidermal fusions. The calyx, the stigma, and the flower buds were not subject to this process.

Figure 1.

Figure 1.

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Flower morphology of the MicroTom wild type (WT) and the slcer6 mutant. A, Opened wild-type flower and slcer6 flower with fused organs at anthesis. The arrow indicates the suture between fused petals in the mutant. B, Wild-type anthers with longitudinal stomium slits, which serve for pollen release (arrows), and slcer6 anthers closed lengthwise. C, Cross sections of anthers at anthesis. In the wild type, the anther walls bend inward. Epidermal fusion in slcer6 (arrowheads) impedes curling of the anther walls, and the pollen stays inside joint locules of adjacent anthers. Most of the pollen has been lost during specimen preparation. Ec, Epidermal cells; L, locule; S, splitted stomium. D, Cross section of a slcer6 flower bud at the stage of microspore formation. The left box marks the site of petal fusion depicted in E. The right box marks the site of petal-to-anther epidermal fusion depicted in F. AnEp, Anther epidermis; P, petal.

To further characterize the slcer6 mutant, the fused organs at different flower developmental stages were examined by scanning electron microscopy. These stages were correlated to the bud length (Table I). Epidermal cells of the wild-type anther underwent elongation at the time of microspore maturation (Fig. 2, A and B). Typical nanoridges emerged later during pollen development (Fig. 2C). In slcer6, misshapen epidermal cells were found in immature anthers (Fig. 2, D and E). The mutant epidermis at anthesis also displayed nanoridges, but its surface was covered with petal imprints (Fig. 2F). In both lines, epidermal cells of the adaxial petal surface were hardly distinguishable at the stage of microspore because of numerous anther imprints (Fig. 2, G and J). At the stage of binucleate pollen, the outlines of petal epidermal cells became apparent in the wild type, while in the slcer6 mutant, the imprints persisted (Fig. 2, H and K). At anthesis, petal epidermal cells were easily discernible in both lines. However, the petal epidermis of the mutant showed remnants of anther cells that stayed attached after mechanical disconnection of the fused organs (Fig. 2, I and L). Thus, the anther abaxial and petal adaxial surfaces dynamically interact in the course of flower development. At the time of anthesis, this connection is abolished in the MicroTom wild type but persists in the slcer6 mutant, resulting in coalescence of the anther cone and the corolla.

Table I. Stages of microsporogenesis and microgametogenesis in MicroTom flowers categorized by bud length (n = 10).

Stage Bud Length
mm
Microsporocyte meiosis 2.0–2.5
Tetrad 2.5–3.0
Early microspore 3.0–3.5
Vacuolated microspore 3.5–4.0
Late microspore 4.0–5.0
Binucleate pollen 5.0–6.0
Mature pollen ≥7.0
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Figure 2.

Figure 2.

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Scanning electron microscopy of the abaxial anther and the adaxial petal epidermis at different stages of flower development in the MicroTom wild type (WT) and the slcer6 mutant. A and D, Anther epidermal cells in the flower bud at the stage of late microspore. In slcer6, cells are misshapen (arrows). B and E, In both lines, anther epidermal cells become wider at the stage of young pollen. Misshaping of cells in the mutant is indicated (arrows). C and F, At the time of anthesis, epidermal cells have lobed margins and show cuticular ridges (arrowheads). slcer6 anther epidermal cells display petal imprints left after mechanical separation of the fused organs (arrows). G and J, The petal surface displays anther epidermal cell imprints in the flower buds at the stage of late microspore (dotted lines). Contours of petal epidermal cells are indistinguishable. H and K, The petal pattern becomes visible in the wild type at the time of young pollen formation. In slcer6, the petal surface still shows conspicuous anther imprints (dotted line). I and L, At the time of anthesis, petal epidermal cells are apparent in both lines. In slcer6, the remnants of anther cells left after mechanical disconnection are found between petal epidermal cells (arrows).

Sterility of the slcer6 Mutant

Seeds were present in all MicroTom wild-type fruits, while in the slcer6 mutant, seeds were found only in 27% of fruits. The quantity of seeds per fruit in the mutant was reduced to 5% ± 1% (mean ± se) of the wild type (Fig. 3A). In order to verify whether female fertility and the pollen-stigma interaction are affected by the mutation, the slcer6 plants were pollinated with the wild-type pollen. The slcer6 stigma fully supported germination of the wild-type pollen (Fig. 3B). The number of seeds in fruits obtained by the cross pollination was the same as in the wild type (Fig. 3A). To verify whether pollen release could be impaired in the mutant, slcer6 plants were hand pollinated with the slcer6 pollen. The pollen germinated on the stigma, but pollen tubes were not as abundant as in the cross-pollination experiments (Fig. 3B). The number of seeds per fruit increased almost 10 times, but it was still two times lower compared with the MicroTom wild type (Fig. 3A).

Figure 3.

Figure 3.

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Seed production and in vivo pollen germination in the MicroTom wild type (WT) and the slcer6 mutant. A, Seed production in the wild type (n = 47), the slcer6 mutant (n = 129), and slcer6 plants hand pollinated (HP) with wild-type pollen (n = 28) or slcer6 pollen (n = 51). Means ± se are shown. Significant differences are marked with different letters (P < 0.05, one-way ANOVA followed by Tukey’s HSD test). B, In vivo pollen germination. The slcer6 flowers were hand pollinated with wild-type (n = 29) or slcer6 (n = 31) pollen. Pistils were divided in three classes: pistils with numerous pollen tubes (more than 10; black bars), few pollen tubes (gray bars), and no pollen tubes (white bars).

Pollen Development and Exine Formation in the slcer6 Mutant

Microscopic analysis showed that a large number of the slcer6 pollen grains are misshapen and highly variable in size (Fig. 4, A and B). Some mutant pollen grains were stained neither with 4′,6-diamidino-2-phenylindole (DAPI) nor with fluorescein diacetate. Such cells were represented by pollen walls without nuclei or cytoplasm (Fig. 4, C and D). On average, the degraded pollen made up 68% ± 1% (mean ± se). Microsporogenesis, beginning with meiosis of the microsporocyte and completed with microspore formation, occurred without visible disorders. Microgametogenesis, which includes microspore maturation and mitosis with subsequent binuclear pollen grain formation, was heavily impaired in the mutant. Already at the stage of the early microspore, nuclei were degraded in some cells (Fig. 4, E and F). Late microspores had a highly vacuolated cytoplasm and no nuclei (Fig. 4, G and H).

Figure 4.

Figure 4.

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Microgametogenesis in the MicroTom wild type (WT) and the slcer6 mutant. A and B, Wild-type pollen with vegetative and generative nuclei and misshapen slcer6 pollen grains stained with DAPI. C and D, Semisections of wild-type pollen with electron-dense cytoplasm and the degraded slcer6 pollen. Pollen walls without cytoplasm are marked (arrows). E and F, Wild-type and slcer6 early microspores stained with DAPI. Microspores without nuclei are indicated in the mutant (arrows). G and H, Semisections of wild-type and slcer6 late microspores. Wild-type cells have structured nuclei and cytoplasm with vacuoles and mitochondria. Mutant microspores are highly vacuolated and have no nuclei.

In order to verify whether the malfunction in slcer6 pollen development is caused by impairments in sporopolleninic exine formation, scanning and transmission electron microscopy of the pollen wall were performed. Neither the slcer6 pollen exine nor that of the microspore were different from the MicroTom wild type. Pollen resistance to acetolysis was also unchanged (Supplemental Fig. S1).

Water Content, Water Loss, and Cuticle Permeability Changes during slcer6 Flower Bud Development

Since an alteration of the flower cuticle may provoke enhanced transpiration and subsequent untimely pollen dehydration, the water status of the slcer6 flower buds was addressed. The water content did not differ in the wild-type and slcer6 young flower buds at the stage of microsporogenesis (Fig. 5A). Further on, the water content in slcer6 decreased to 87% of the MicroTom wild type (bud length, 6–8 mm). Water loss in the buds with young pollen (bud length, 6.5 mm) was 1.34-fold higher in the mutant compared with the wild type (Fig. 5B). The cuticle of the slcer6 young buds at the stage of microsporogenesis (bud length, 1–4 mm) was more permeable for chlorophylls and carotenoids as compared with the MicroTom wild type: the amount of diffused pigments was 1.3-fold higher after 150 min of extraction (Fig. 5C). This difference increased 3-fold in flower buds with young pollen (bud length, 6–7 mm). The total pigment content was not altered (Supplemental Fig. S2).

Figure 5.

Figure 5.

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Analysis of the water content, transpirational water loss, and pigment leaching in MicroTom wild-type (WT) and slcer6 flower buds. A, Water content changes in the course of flower development (means ± se, n = 20–30). Significant differences in the water content are marked with asterisks (P < 0.05, one-way ANOVA followed by Tukey’s HSD test). B, Transpirational water loss of flower buds with binucleate pollen (bud length, 6.5 mm; means ± sd, n = 5; P < 0.05, Mann-Whitney U test). DW, Dry weight. C and D, Chlorophyll and carotenoid leaching for the flower buds at early stages of microsporogenesis (bud length, 1–4 mm) and for flower buds with young pollen (bud length, 6 mm). Means ± sd are shown; n = 5. Each time point is significantly different between the wild type and the slcer6 mutant for both pigments (P < 0.05, Mann-Whitney U test). FW, Fresh weight.

Anther Tapetum Development in the slcer6 Mutant

Since male sterility is usually associated with abnormal tapetum development, transmission electron microscopy was performed to characterize the slcer6 tapetum ultrastructure. In the MicroTom wild type at the early microspore stage, the cytoplasm of tapetal cells contained mitochondria, plastids, extensive endoplasmic reticulum, and vacuoles with inclusions. The cells had an irregular shape, and their walls were lined with electron-dense orbicules (Fig. 6A), tapetum-produced globular structures consisting of sporopollenin. In the slcer6 mutant, tapetum cells almost devoid of cytoplasm were detected already at the stage of early microspore (Fig. 6B). The outer tapetum facing the epidermis was generally represented by such empty cells. At the stage of late microspore, degenerating tapetal cells contained multiple spheroid lipid bodies in the wild type (Fig. 6C). In the slcer6 mutant, the cytoplasm of the inner tapetum cells contained irregularly shaped prominent bodies, while the outer tapetum cells were mostly empty (Fig. 6, D–F). At the time of anthesis, tapetal cells were devoid of cytoplasm in both lines (Fig. 6, G and H). Thus, a substantial part of the outer tapetum underwent untimely degradation in slcer6, while the inner tapetum demonstrated altered morphology.

Figure 6.

Figure 6.

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Transmission electron microscopy of anther tapetum cells during microgametogenesis in the MicroTom wild type and the slcer6 mutant. A, Wild-type tapetum at the stage of early microspore. The tapetum cell is irregularly shaped, and electron-dense cytoplasm contains mitochondria (M), vacuoles (V), and other organelles. The tapetal wall (TW) facing the locule is lined with electron-dense orbicules (Or). B, slcer6 tapetum at the stage of early microspore. The tapetal cell is almost devoid of cytoplasm. C, Wild-type tapetum at the vacuolate microspore stage. Cytoplasm becomes more disintegrated and contains spheroid lipid drops (arrowheads). D, slcer6 inner tapetum cell with irregularly shaped prominent bodies (asterisks) at the vacuolate microspore stage. E, Outer tapetum at the vacuolate microspore stage in slcer6. The tapetum cell lined with orbicules along the wall has almost no cytoplasm. F, Cross section of a slcer6 anther with microspores (MS). The outer tapetum (OtT) is located at the side of contacting anthers. The inner tapetum (InT) is located closer to the connective tissue. G, Wild-type tapetum at anthesis and mature pollen grain (PG) with highly structured electron-dense cytoplasm. The tapetal wall facing the locule is appressed to that of the middle layer. H, slcer6 tapetum and degenerated pollen grain (DP) at anthesis.

Composition of Cuticular Waxes, Cutin, and Cuticle Ultrastructure in the slcer6 Flower

Analysis of anther cuticular waxes revealed six compound classes in both lines: (1) n-alkanes, (2) iso-alkanes, (3) anteiso-alkanes, (4) alkanoic acids, (5) sterols, and (6) triterpenoids. In the MicroTom wild type, the most prominent compounds were n- and iso-alkanes, mainly with chain lengths of C29 and C31, while in the mutant, alkanoic acids represented the major compound class (Fig. 7A). The strongest effect of the slcer6 mutation on wax constituents was found for the n-alkanes. Their total content was diminished to 19% of the wild type (Table II). The mutation affected n-alkanes with chain lengths of C25 and C27 or greater. Branched alkanes in slcer6 reached only around one-third of their amount in the wild type. The content of iso-alkanes with chain lengths of C25, C27, and C29 or greater and anteiso-alkanes with chain lengths of C28 and C30 or greater was lowered in comparison with the wild type. Deposition of n-C32 and C33 in all groups of alkanes was exclusively seen in the wild type. The content of other compound classes was comparable in both lines.

Figure 7.

Figure 7.

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Cuticular wax and cutin composition of MicroTom wild-type (WT) and slcer6 flowers at anthesis. A, Composition of the anther cuticular waxes. B, Cutin composition of the anther and the corolla (freeze-dried material). Means ± sd are shown; n = 4 to 5. Significant differences are marked with asterisks (P < 0.05, Mann-Whitney U test). DW, Dry weight.

Table II. Total cuticular wax quantities and relative wax composition of MicroTom wild-type and slcer6 anthers at anthesis (n = 5).

Values shown are means ± sd.

Compound Class MicroTom Wild Type MicroTom slcer6 Ratio
µg g−1 dry wt
n-Alkanes
 Even chained 142 ± 11 71 ± 5 0.50a
 Odd chained 565 ± 39 61 ± 1 0.11a
 Total 707 ± 46 (33%) 132 ± 5 (13%) 0.19a
iso-Alkanes
 Even chained 90 ± 5 42 ± 1 0.47a
 Odd chained 628 ± 30 209 ± 6 0.33a
 Total 718 ± 34 (33%) 251 ± 6 (25%) 0.35a
anteiso-Alkanes
 Even chained 108 ± 5 29 ± 2 0.27a
 Odd chained 17 ± 1 12 ± 2 0.71a
 Total 125 ± 5 (6%) 41 ± 3 (4%) 0.33a
Alkanoic acids
 Total 445 ± 55 (21%) 445 ± 23 (44%) 1.00
Triterpenoids and sterols
 Total 160 ± 11 (7%) 148 ± 15 (15%) 0.93a
Wax coverage 2,155 ± 92 (100%) 1,017 ± 49 (100%) 0.47a
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a

P < 0.05, Mann-Whitney U test.

In order to exclude possible effects of the altered morphology of the slcer6 anthers on the surface area, the same experiment was performed for anthers and corollas harvested from closed green flowers before anthesis. The data were comparable to those presented above, but the impact of the slcer6 mutation was detected for n- and iso-alkanes with chain lengths of C27 or greater and for anteiso-alkanes with chain lengths of C26 or greater (Supplemental Fig. S3A). Analysis of the wild-type corolla waxes revealed that the contribution of branched alkanes in total wax coverage in petals is more prominent than in the anther and makes up 56%, although the wax pattern was similar for both organs (Supplemental Fig. S3B).

Analysis of the monomeric composition of cutin was performed for anthers and corollas harvested at anthesis. In the first experimental setup, enzymatically isolated cuticles were analyzed in order to determine the relative proportions of cutin monomers (Supplemental Fig. S4). In the second, freeze-dried material was used for quantitative estimation of the cutin content (Fig. 7B). The major monomer in both lines was (9)10,16-dihydroxyhexadecanoic acid; its portion made up 25% to 30% of the total cutin content in freeze-dried material and more than 90% in the isolated cuticles. The relative proportions of different cutin monomers for the MicroTom wild type and the slcer6 mutant were essentially comparable in both types of samples.

Figure 8.

Figure 8.

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qRT-PCR analysis of SlCER6 transcriptional level in different organs of the MicroTom wild type (WT) and its slcer6 mutant. Flower organs (sepal, petal, pistil, anther, and pollen), fruits (whole immature green, peel of mature green, and mature red fruits), young leaf, stem, and root were sampled in independent biological replicates (n = 3 for the wild type, n = 2 for slcer6). Petal and pollen are shown only for the wild type. Means ± sd are shown.

To verify whether the adhesion response in the slcer6 mutant is associated with defects of the cuticular membrane, the cuticle ultrastructure was inspected (Supplemental Fig. S5). The cuticle was observed as a thin continuous and regular layer with a thickness around 60 nm in the wild type anther with early microspores. At the time of anthesis, the cuticle membrane thickened to 270 ± 41 nm. However, the slcer6 anther cuticle membrane showed the same structure throughout development, even in the fusion suture zone. The cuticle thickness in the mutant was not altered and made up 270 ± 26 nm at anthesis.

Expression of the SlCER6 Gene

Quantitative reverse transcription (qRT)-PCR analysis was carried out to examine the expression profile of SlCER6 among flower organs, pollen, immature green fruits, the peel of mature green and red fruits, young leaves, stems, and roots (Fig. 8). SlCER6 transcripts were detected in all MicroTom wild-type samples except the pollen. The highest level of SlCER6 transcripts was found in the anther. High SlCER6 expression was evident in young leaves and mature and immature green fruits, where it made up 76% and 57% to 59% of the expression in the anther, respectively. In the mature red fruit, the transcript amount decreased to 32%. The SlCER6 expression pattern for the wild-type fruits was similar to the one obtained earlier (Mintz-Oron et al., 2008). SlCER6 was expressed at intermediate levels in sepals and petals of 39% to 42% compared with the anther. The root and the pistil expressed SlCER6 only weakly. In the slcer6 mutant, SlCER6 expression was negligible in all samples and did not exceed 0.01%; only in the anther, it reached 6% of the expression level in the wild-type anther.

To characterize SlCER6 spatial expression in the MicroTom flower, transgenic plants containing a GUS reporter construct fused to the upstream genomic promoter sequence of SlCER6 were analyzed. GUS expression was found in anthers, sepals, and petals of flowers at anthesis (Fig. 9A). Positive blue GUS staining bordered the anther stomial slit coinciding with the sites of anther-to-anther epidermal fusion in the slcer6 mutant (Fig. 9B). The most intensive GUS staining was observed at the distal part of the anther where two apical pores are located (Fig. 9C). Examination of cross sections revealed that GUS is expressed in the epidermal cells of anther, sepal, petal, and in the ovary (Fig. 9, D–H). Positive blue staining was also detected in the endothecium, a subepidermal cell layer specifically formed in the upper one-third of the tomato anther, but neither in the developing pollen nor in tapetal cells. For further analysis, GUS expression was examined in flower buds at early developmental stages. GUS staining became visible in sepals and anthers already in 1-mm flower buds with newly differentiated organs (Fig. 9I). Blue positive staining highlighted the whole sepal surface before meiosis of microsporocytes (Fig. 9J). In contrast to the sepals, GUS staining became initially apparent at petal edges (Fig. 9, K–M), which are subject to the organ fusion in the slcer6 mutant.

Figure 9.

Figure 9.

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Temporal and spatial expression pattern of SlCER6 detected by a GUS reporter gene assay. A, Inflorescence at anthesis with reporter gene expression in sepals, petals, and the upper part of the anther (blue). B, Adaxial side of the anther at anthesis. The staining borders longitudinal stomium slits, sites of anther desiccation and pollen distribution (arrows). C, Androecium and gynoecium at the stage of young microspore. GUS is expressed in the region of the future apical pores, longitudinally between anthers (arrow), and in the ovary. D, Fragments of an anther cross section showing GUS staining in the epidermal and endothecial cells with characteristic ribs. En, Endothecium; Ep, epidermis; L, locule; T, tapetum. E, Cross section of the anther apex above the pores showing GUS expression in the epidermis. F and G, Cross sections with staining of the petal (F) and the sepal (G) epidermis in flower buds with microspores. H, Cross section of an ovary with GUS staining in the ovary wall in a flower at anthesis. EW, External ovary wall; IW, internal ovary wall. I, Cross section of a flower bud with newly formed flower organs. GUS is expressed in the sepal epidermis and the anther. An, Anther; S, sepal. J, Flower bud with microsporocytes. Sepals are stained. K to M, Development of GUS expression in petals. The expression emerges initially in petal margins.

DISCUSSION

Contribution of Cuticular Waxes to the Prevention of Flower Organ Fusion in Tomato

The slcer6 tomato exhibits prominent epidermal fusion of flower organs. Intimate interaction between epidermal cells is well reflected in changes of the petal and anther surface architectures in the course of flower development. SlCER6 transcripts are detected in the petal and the anther fusion sites in tomato, similar to the CER6 transcripts in Arabidopsis (Hooker et al., 2002). Additionally, SlCER6 is expressed in organs not subject to the fusion process (e.g. sepals). This might be explained by early and asynchronous formation of the sepal primordia in flower development. As a result, each sepal stays apart from the other organs (Sekhar and Sawhney, 1984). SlCER6 transcripts were also detected in the walls of the multilocular ovary, which is formed as a result of a routine carpel fusion process. Similarly, expression of a FDH ortholog was found in the Antirrhinum majus ovary, which has the same ontogeny as tomato (Efremova et al., 2004). The data on the SlCER6 expression pattern in vegetative organs of tomato are similar to the results obtained for Arabidopsis by qRT-PCR and the analysis of GUS expression under the control of the CER6 promoter (Hooker et al., 2002; Joubès et al., 2008).

The hypothesis of the involvement of cuticular waxes in postgenital organ fusion was proposed some time ago (Lolle and Pruitt, 1999; Tanaka and Machida, 2006). However, it was found that the abnormal adhesion response in Arabidopsis is not necessarily accompanied by defective wax accumulation but rather is characterized by impairments in cutin formation and/or the inability to form the continuous cuticle proper. Irregular depositions in the cuticle were observed in wax2 (Chen et al., 2003), hothead/ace (Kurdyukov et al., 2006b), bodyguard (Kurdyukov et al., 2006a), lacerate (Voisin et al., 2009), cutinase-expressing plants (Sieber et al., 2000), and atwbc11 defective in the wax transporter (Luo et al., 2007). The wax-deficient fdh mutant showed an unaltered cuticle structure in the fusion zones but, in contrast to slcer6, accumulated abnormally high levels of cutin and wax (Voisin et al., 2009). It is a peculiarity of slcer6 that it has a reduced wax content but unchanged structure and quantity of cutin. Thus, neither perturbation in the cuticle structure nor altered cutin, but a malfunction of wax biosynthesis, is primarily responsible for the flower fusion phenomenon in slcer6.

Putative Role of the SlCER6 Enzyme in Cuticular Wax Production in Tomato

Research on wax biosynthetic pathways focuses mainly on the biosynthesis of linear aliphatics, since they are characteristic components of plant cuticular waxes (Samuels et al., 2008). However, large amounts of iso- and anteiso-alkanes were found in waxes of Solanaceae members, specifically in leaves of tobacco (Nicotiana tabacum) and eggplant (Solanum melongena; Grice et al., 2008; Haliński et al., 2009). The content of branched alkanes is low in leaves and mature fruits of tomato, as in the majority of model plants used in cuticle research so far (Vogg et al., 2004; Leide et al., 2007). It is a new finding that these compounds are abundant in tomato flower waxes.

The slcer6 mutation mostly affects n-alkanes in flowers, leaves, and fruits. However, the slcer6 mutation has a high impact on the content of branched alkanes in flowers and immature green fruits (Leide et al., 2007). Since waxes of leaves and mature fruits contain branched alkanes only in trace amounts (Vogg et al., 2004; Leide et al., 2007), the effect of the slcer6 mutation is masked in these organs. One can suggest that SlCER6 is involved in the production of both linear and branched alkanes with chain lengths of C27 or greater. Importantly, most of these alkanes are still present in the mutant, although in reduced quantities, indicating that redundant enzymes or other β-ketoacyl-CoA synthases, whose substrate specificities overlap with SlCER6, might contribute to the biosynthesis of these compounds.

The current vision of the biosynthesis of branched aliphatics states that iso- and anteiso-long-chain fatty acids are synthesized by the successive elongation of CoA thioesters of short branched-chain acids. These fatty acids are derived from branched-chain amino acids (Val, Leu, and Ile). Thus, iso-, anteiso-, and n-alkanes have different CoA precursors, while the subsequent elongation reactions are similar (Kolattukudy, 1970; Grice et al., 2008). Recent research on Arabidopsis showed that overexpression of CER1, involved in the reduction reaction of VLCFAs to alkanes (Bernard et al., 2012), sharply increases the production of n- and iso-alkanes in leaves, indicating that CER1 is able to produce both linear and branched hydrocarbons (Bourdenx et al., 2011). This presumed ability to work with different substrates might be shared between the SlCER6 and CER1 enzymes.

Thus, the SlCER6 enzyme specificity seems to be broader than was suggested before. The data testify that SlCER6 elongates fatty acids leading to n- and iso-alkanes with total chain lengths of C27 or greater and to anteiso-alkanes with total chain lengths of C28 or greater. Probably, the resulting product of the condensation reaction depends not only on the specificity of the enzyme but also on the accessibility of linear or branched-chain acids, which in turn depends on the CoA precursors.

The Impact of the SlCER6 Gene on Sexual Reproduction in Tomato

Hand-pollination experiments revealed that reduced seed production in the slcer6 mutant is caused exclusively by male sterility. One could assume that the SlCER6 gene product is involved in the pollen exine formation because most male-sterile mutants are impaired in sporopollenin biosynthesis or transport (Piffanelli et al., 1998; Ariizumi and Toriyama, 2011). Enzymes participating in both wax biosynthesis and pollen exine formation are known for Arabidopsis and rice (Oryza sativa), such as FACELESS POLLEN1 (Ariizumi et al., 2003), WAX DEFICIENT ANTHER1 (Jung et al., 2006), and DEFECTIVE POLLEN WALL (Shi et al., 2011). However, this suggestion is not confirmed, since the slcer6 exine does not differ from the wild type. The absence of SlCER6 transcripts in the developing pollen indicates that SlCER6-derived aliphatics are not produced by the male gametophyte itself, as in Arabidopsis (Hooker et al., 2002).

Successful production of viable pollen depends on the programmed cell death of the tapetum. Both premature and delayed death of tapetal cells result in malfunction of the nutrient supply followed by death of the microgametophyte (Parish and Li, 2010; Wilson et al., 2011). We found that impairment in the tapetal cell degeneration process contributes to male sterility in the slcer6 tomato. In contrast to slcer6, the atcer6 mutant has viable pollen and its tapetum does not display any abnormalities (Preuss et al., 1993). However, the content of long aliphatics is reduced in the atcer6 pollen coat, causing conditional sterility (Preuss et al., 1993; Fiebig et al., 2000). The finding of CER6 transcripts in the tapetum by in situ hybridization led to the conclusion that CER6 is necessary for the biosynthesis of pollen coat lipids by tapetal cells (Hooker et al., 2002). Surprisingly, SlCER6 expression is not detected in the tapetum of the tomato GUS reporter line. Probably, this reflects different contributions of the condensing enzymes to the formation of the lipidic pollen coat in these two species. Indeed, the pollen coat is feebly developed in tomato pollen, while the Arabidopsis pollen grain has a thick pollen coat with lipidic droplets (Preuss et al., 1993). Zheng et al. (2005) found that the cer10 mutant, deficient in an enoyl-CoA reductase required for VLCFA biosynthesis, has unviable pollen similar to slcer6. One can speculate that aliphatics produced by the fatty acid elongase complex are involved in the regulation of tapetum disintegration. A few years ago, evidence for the involvement of VLCFAs in the regulation of cell death appeared. It was shown that epidermal misexpression of the FATTY ACID ELONGATION1 gene in Arabidopsis leads to the death of trichomes via a lipidic pathway. An assumption was made that the death program is triggered by the presence of a critical threshold of certain fatty acids or other lipids (Reina-Pinto et al., 2009).

Another parameter provoking untimely tapetum degeneration in the slcer6 mutant could be water stress in flowers, similar to that in the fruits (Vogg et al., 2004; Leide et al., 2007). It was shown that anther water deficiency causes abnormal tapetal cell development in wheat (Triticum aestivum; Lalonde et al., 1997; Koonjul et al., 2005). Loss of turgor by anther epidermal cells, reduced water content, higher pigment leaching, and water loss might be signs of water stress in the slcer6 flower. It is hypothesized that this is mainly due to changes in cuticular wax chemistry and amount. However, some contribution of stomatal transpiration cannot be excluded, since tomato sepals have stomata (Sekhar and Sawhney, 1984).

Morphological studies together with cross-pollination experiments showed that impediment in pollen release from the anther is a further reason for male sterility in the slcer6 mutant. The absence of longitudinal stomium slits in the mutant anther could also influence the pollen desiccation process, reducing pollen lifetime and viability. Tomato plants with a mutated POSITIONAL STERILE (PS) gene, responsible for blocking of the decarbonylation pathway leading to alkane, aldehyde, ketones, and secondary alcohol biosynthesis, have sterility and unopened flowers similar to slcer6 (Leide et al., 2011). However, the ps mutant produces ample seeds under artificial self-pollination. Natural pollination does not take place in ps because the stomium cells stay intact, thereby preventing the anther split (Larson and Paur, 1948). In contrast, breakage of the stomium cells in the slcer6 mutant occurs without abnormalities.

Expression of the SlCER6 gene in the endothecium poses a new question regarding the participation of aliphatic compounds in anther dehiscence and opening. Endothecium development is coordinated with pollen maturation and degeneration of the tapetum. At the stage of a young pollen grain, endothecium cells undergo secondary wall thickening by the incorporation of lignocellulose (Dawson et al., 1999). It imposes mechanical stress on the anther, which leads to desiccation and opening (Bonner and Dickinson, 1989; Wilson et al., 2011). Interestingly, SlCER6 is expressed in endothecial cells already at the early microspore stage, before the beginning of the secondary thickening. This may hint at the presence of an aliphatic domain in these cells. However, SlCER6 loss of function does not prevent endothecium thickening. Similarly, specific expression of the SHINE2 transcription factor regulating wax biosynthesis was detected in the stomium region of the Arabidopsis anther (Aharoni et al., 2004). One can speculate that the aliphatics produced by SlCER6 are necessary to prevent water loss by endothecial cells before secondary wall deposition.

This work shows that SlCER6 is involved in the biosynthesis of cuticular waxes not only in vegetative organs, fruits, and leaves but also in flowers. Biosynthesis of the pollen coat lipids in Arabidopsis is not the only function of CER6 during pollen development: SlCER6 appeared to be essential for timely tapetal cell degeneration and successful microgametogenesis in tomato. Here, we report that wax deficiency, but not disturbance in cutin accumulation or cuticle formation, is the primary reason for the adhesive response between epidermal cells in the tomato flower. Based on chemical analysis, the substrate specificity of the SlCER6-condensing enzyme may be reevaluated. It might be involved in the elongation of linear and branched VLCFAs with chain lengths of C26 or greater, leading to the production of linear and branched alkanes, respectively. These findings broaden our current knowledge of wax biosynthetic pathways and show new aspects of the involvement of wax-producing enzymes in sexual plant reproduction.

MATERIALS AND METHODS

Plant Material and Growth Conditions

Plants of tomato (Solanum lycopersicum ‘MicroTom’) wild type and its mutant slcer6 deficient in a β-ketoacyl-CoA synthase were investigated. Isolation of the mutant referred to as lecer6 and generation of the MicroTom plants with upstream CER6 fragments fused to a GUS reporter were described previously (Vogg et al., 2004; Mintz-Oron et al., 2008). The plants were grown in a controlled-climate chamber at 75% to 80% relative humidity with a 14-h photoperiod at 450 µmol m−2 s−1 and a day/night temperature regime of 22°C/18°C. Plants were watered daily and fertilized with Hakaphos Blau nutrient solution once per week.

Scanning and Transmission Electron Microscopy

For scanning electron microscopy, plant material was fixed in 3% (v/v) glutaraldehyde, 0.1 m sodium phosphate buffer, pH 6.8, at 4°C for 4 h, washed in the same buffer, and postfixed in 2% (w/v) osmium tetroxide, 50 mm cacodylate buffer, pH 6.8, at 4°C overnight. Samples were then dehydrated in a graded acetone series and critical point dried with liquid CO2. Before examination with the scanning electron microscope (Zeiss DSM 962), samples were sputtered with gold palladium at 25 mA for 300 s (Bal-Tec SCD 005).

For transmission electron microscopy, plant material was fixed in Karnovsky solution at 4°C overnight and postfixed for 1.5 h as for scanning electron microscopy. The samples were then dehydrated in a graded ethanol series, treated with propylene oxide, and embedded in Spurr’s resin. Semisections were done for light microscopy. Ultrathin sections were cut, mounted on copper grids, and contrasted with 2% (w/v) uranyl acetate and lead citrate. Specimens were examined with a Leo 912 AB microscope (Carl Zeiss).

Fluorescence Microscopy

For the detection of pollen development stage and the pollen viability test, 1 µg mL−1 DAPI (Sigma) and 2 µg mL−1 fluorescein diacetate (Sigma) were used, respectively. The samples were analyzed with Leica DMR (Leica Microsystems) and Axio Observer Z1 microscopes supplied with AxioCam Mrm or Mrc digital cameras and AxioVision 4.8 software (Zeiss).

GUS Histochemical Assay and Cryosectioning

For GUS staining, samples were placed in 90% (v/v) acetone and vacuum infiltrated on ice, transferred to 50 mm sodium phosphate buffer, pH 7.2, containing 10 mm EDTA, 0.1% (w/v) Triton X-100, and 0.5 mm K3Fe(CN)6, and vacuum infiltrated. Samples were transferred to the buffer containing 0.5 mg mL−1 5-bromo-4-chloro-3-indolyl β-d-glucuronide sodium salt (Sigma-Aldrich). After infiltration, the samples were incubated at 37°C overnight. Chlorophyll was washed out in a graded series of ethanol. Afterward, samples were fixed in formalin-acetic acid-alcohol solution at 4°C for 24 h. Prior to examination, samples were washed three times in sodium phosphate buffer. Flower buds were dissected with a Leica MZ16 stereomicroscope and a Leica DC500 digital camera. For cryosectioning, material was frozen in Tissue-Tek O.C.T. (Sakura Fintek Europe) and mounted on stubs. Ten- to 30-µm sections were cut in a Leica CM1900 cryostat (Leica Microsystems) at –20°C. Sections were transferred on glass slides, washed with sodium phosphate buffer, and analyzed.

Analysis of Pollen

To investigate pollen development, flower buds with different lengths were harvested. At least 10 buds were analyzed for each developmental stage. Pollen was collected from flowers at anthesis using a vacuum cleaner (Johnson-Brousseau and McCormick, 2004). For acetolysis, pollen grains were treated with a mixture of sulfuric acid and acetic anhydride at 100°C (Hesse et al., 2009). In vivo and in vitro pollen germination assays were performed (Mori et al., 2006; Smirnova et al., 2009).

Measurement of Water Content, Water Loss, and Leaching of Pigments

Immediately after harvesting, the flower bud length and fresh weight were determined. Afterward, samples were dried to a constant weight at 60°C.

Water loss was recorded for flower buds approximately 6.5 mm in length. For one sample, three buds from different plants were pooled. Pedicles were sealed with high-melting paraffin. The amount of water loss versus time (seven data points per sample) was measured gravimetrically. Subsequently, the dry weight was determined. Between measurements, samples were stored at 25°C on silica gel.

The pigment-leaching protocol was adapted from Lolle et al. (1998). Three flower buds per sample were weighed and immersed in 80% (v/v) ethanol. Aliquots of the supernatant were removed at different time points. To determine the total pigment amount, plant material was frozen in liquid nitrogen, homogenized, dissolved in 100% ethanol, and pelleted. The content of chlorophylls and carotenoids in the supernatant was determined by measuring absorption at 664, 649, and 470 nm (Lichtenthaler and Buschmann, 2001) on a Unicam UV/Vis Spectrophotometer UV4 (Thermo Scientific).

Analysis of Flower Cuticular Waxes and Cutin Monomers

About 10 to 30 mg of freeze-dried material was submersed in 10 mL of chloroform containing 3 mg of heptatriacontane (Fluka) as an internal standard for 1 min and filtered through a paper filter. The solvent was evaporated under a flow of nitrogen. Hydroxyl-containing compounds were transformed into the corresponding trimethylsilyl derivatives using N,O-bis-trimethylsilyl-trifluoroacetamide (Macherey-Nagel) in pyridine (Merck). The wax composition was analyzed as described previously (Leide et al., 2007).

For cutin analysis, freeze-dried material or enzymatically isolated cuticles were used. About 25 mg of freeze-dried anthers and corollas was washed with chloroform at room temperature and at 50°C for 30 min. Afterward, samples were incubated in chloroform, which was changed daily, for 1 week. Cuticles from fresh anthers and corollas were isolated according to Leide et al. (2007). Dried samples were transesterified with 1 mL of 1.25 m methanol-HCl (Fluka) at 80°C overnight. Subsequent extraction of cutin components and derivatization were performed as described by Leide et al. (2007).

Gene Expression

Three biological samples for MicroTom wild type and two for the slcer6 mutant were compared. Plant material was harvested from 2- to 4-month-old plants. RNA isolation, complementary DNA synthesis, qRT-PCR, and data analysis were performed as described in detail by Leide et al. (2012). TIP41 was chosen as a reference gene among RPS26C, EFa1, RPL2, CAC, EXP, TUB, and UBQ (Supplemental Table S1) according to the expression stability analyzed by NormFinder version 20.

Statistical Analysis

Significant differences (P < 0.05) between multiple data sets were tested by one-way ANOVA followed by Tukey’s honestly significant difference (HSD) test for unequal n. Pairwise comparisons were tested by Mann-Whitney U test. Analysis was performed with STATISTICA 20 (StatSoft).

Sequence data from this article can be found in the GenBank database under the following accession number: CER6, GQ214500.

Supplemental Data

The following materials are available in the online version of this article.

  • Supplemental Figure S1. Scanning and transmission electron microscopy of the pollen grains and microspores in the MicroTom wild type and the slcer6 mutant.

  • Supplemental Figure S2. Total content of chlorophylls and carotenoids in the MicroTom wild type and the slcer6 flower buds at the stage of microgametogenesis (6.5 mm in length).

  • Supplemental Figure S3. Cuticular wax composition of the corollas and anthers harvested before flower opening in the MicroTom wild type and slcer6 and the wild-type corolla at anthesis.

  • Supplemental Figure S4. Cutin composition of the cuticles isolated from anther and corolla of the MicroTom wild type and the slcer6 mutant.

  • Supplemental Figure S5. Transmission electron microscopy of the anther cuticle at different stages of flower development in the MicroTom wild type and the slcer6 mutant.

  • Supplemental Table S1. Primer pairs used for gene expression analysis by qRT-PCR.

Acknowledgments

We thank Asaph Aharoni for providing us with the GUS reporter plants and Avraham A. Levy for supplying seeds for the slcer6 mutant (Weizmann Institute of Science). We are indebted to Georg Krohne for the possibility to perform electron microscopy (Theodor-Boveri-Institut für Biowissenschaften). We thank Daniela Bunsen, Claudia Gehrig, and Olga Frank for technical assistance and Jutta Winkler-Steinbeck for plant care. We are grateful to Gerd Vogg and Anton Hansjakob for criticism of the manuscript and to Ulrich Hildebrandt for comments on the manuscript (Julius-von-Sachs-Institut für Biowissenschaften).

Glossary

VLCFA

very-long-chain fatty acid

DAPI

4′,6-diamidino-2-phenylindole

qRT

quantitative reverse transcription

HSD

honestly significant difference

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