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The Pex16p Homolog SSE1 and Storage Organelle Formation in Arabidopsis Seeds

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Science  09 Apr 1999:
Vol. 284, Issue 5412, pp. 328-330
DOI: 10.1126/science.284.5412.328

Abstract

Mature Arabidopsis seeds are enriched in storage proteins and lipids, but lack starch. In the shrunken seed 1(sse1) mutant, however, starch is favored over proteins and lipids as the major storage compound. SSE1 has 26 percent identity with Pex16p in Yarrowia lipolytica and complementspex16 mutants defective in the formation of peroxisomes and the transportation of plasma membrane– and cell wall–associated proteins. In Arabidopsis maturing seeds, SSE1 is required for protein and oil body biogenesis, both of which are endoplasmic reticulum–dependent. Starch accumulation in sse1 suggests that starch formation is a default storage deposition pathway.

To support young seedling growth flowering plants deposit storage compounds (principally composed of carbohydrates, proteins, and lipids) in their seeds. Cereal plants deposit relatively more carbohydrates, whereas legume seeds and oilseeds contain relatively more proteins and lipids, respectively (1). Understanding the molecular mechanisms underlying the cellular differentiation programs involved in storage deposition would allow better manipulation of seed quality and the nutritional value of crop seeds.

In Arabidopsis, proteins and lipids are the major reserves in mature seeds (2) (Fig. 1, A and B). We have isolated ashrunken seed 1 (sse1) mutant that alters this seed storage profile, accumulating starch over proteins and lipids (Fig. 1, C and D). The cotyledon and the hypocotyl cells of sse1contain no recognizable protein bodies and few oil bodies. Starch granules, membrane stacks, vesicles, and vacuoles, all of which are absent in wild-type cells, are present in sse1 cells, and the oil bodies in sse1 contain higher electron density substances than the wild type. The sse1 seeds shrink upon desiccation (a likely consequence of insufficient deposition of storage molecules), whereas the wild-type seeds are desiccation tolerant (Fig. 1, E and F).

Figure 1

Abnormal storage deposition and the shrunken phenotype of sse1 seeds. Transmission electron micrograph (21) of a representative cell from the wild-type cotyledon (A) or hypocotyl (B) and thesse1 cotyledon (C) or hypocotyl (D). Wild-type cells are filled with numerous oil bodies (OB) and a few large protein bodies (PB). sse1 cells contain few oil bodies and additional structures such as starch granules (St), vacuoles (Vc), stacks of membranes (M), and vesicles (Vs). Wild-type C24 (E) and sse1 (F) seeds. Bar, 3.1 μM (A to D).

The sse1 mutant was identified in a transferred DNA (T-DNA) transgenic line [T line (3)] that exhibited the shrunken seed phenotype. Among the T2 seeds on the T1 plant (the primary transgenic plant), 90% of the seeds were shrunken and 10% were normally rounded. The shrunken seeds were not viable, and plants grown from the round seeds produced ∼90% shrunken seeds. This pattern of inheritance continued for generations after self-pollination (4). However, after backcrossing the T2 line to wild-type plants, sse1 behaved as a typical single-recessive Mendelian gene (5) and, when the segregation patterns of F3 families were observed, sse1 cosegregated with the T-DNA (6). The SSE1 gene and its cDNA were cloned (7), and the sequences obtained were used to design three primers for determining the genotypes of shrunken and round seeds by single-seed polymerase chain reaction (PCR) (8) (Fig. 2A). Sixty-six percent (n = 6) of round F2 seeds were heterozygous and 33% were homozygous for the wild-type allele, whereas 100% (n = 5) of shrunken F2 seeds were homozygous for the T-DNA insertion (Fig. 2B). T4 seeds were also analyzed. All round seeds (n = 13) were heterozygous and all shrunken seeds (n = 21) were homozygous for the T-DNA insertion (Fig. 2C). Thus, sse1 is recessive and in the self-pollinated T line, the mutant allele is transmitted at a higher frequency than the wild-type allele (9).

Figure 2

Genotype determination by single-seed PCR. (A) PCR diagram. Primers A and B amplify a 0.9-kb fragment from the wild-type SSE1 allele. Primers C and B amplify a 1.6-kb fragment from the T-DNA interruptedsse1 allele. (B) Single-seed PCR of round (R) and shrunken (S) seeds in a F2 population derived from a backcross between a T2 and a wild-type plant. (C) Representative single-seed PCR results of round (n =13) and shrunken (n = 21) T4 seeds. A control reaction from a wild-type (WT) seed is also shown.

The SSE1 cDNA (GenBank accession numberAF085354) encodes a predicted protein of 367 amino acids (Fig. 3A). Expression of SSE1 cDNA in transgenic sse1 plants (10) complements the shrunken seed phenotype (Fig. 3B). Similar to wild type, seeds are tolerant of desiccation and cells are filled with storage proteins and lipids, but lack starch. The SSE1 sequence showed similarity to Pex16p, a membrane-associated protein required for the assembly and proliferation of peroxisomes (11) and for the trafficking of plasma membrane– and cell wall–associated proteins (12), in the yeast Y. lipolytica (Fig. 3A). Pex16p is glycosylated and transiently localized in the endoplasmic reticulum (ER) (13). Despite the limited amino acid sequence similarity (26% identity), the two proteins have similar arrangements of their hydrophobic and hydrophilic regions (Fig. 3A). A predicted glycosylation site was found in SSE1 (Fig. 3A). SSE1 complements the growth of pex16 mutants on oleic acid as sole carbon source (Fig. 3C), indicating restoration of peroxisomal function (11). The restoration of limited growth of the disruption allele P16KO-8A (11) indicates that SSE1 cannot fully replace Pex16p in peroxisome formation, probably because of the functional difference (or differences) between the two proteins. In addition, SSE1 partially complemented the pex16-1 mutant for the dimorphic transition from yeast to the mycelia form (Fig. 3D). Pex16p is normally required for mycelia phase–specific cell surface protein transport.

Figure 3

SSE1 amino acid sequence analysis andSSE1 complementation of the Arabidopsis sse1 andY. lipolytica pex16 mutants. (A) Alignment of SSE1 and Pex16p. Dots indicate gaps. Identical residues are boxed. Hydrophobic (single line) and hydrophilic (double line) regions for both proteins are underlined (22). The predicted glycosylation site of SSE1 is indicated with an asterisk. Single-letter abbreviations for the amino acid residues are as follows: A, Ala; C, Cys; D, Asp; E, Glu; F, Phe; G, Gly; H, His; I, Ile; K, Lys; L, Leu; M, Met; N, Asn; P, Pro; Q, Gln; R, Arg; S, Ser; T, Thr; V, Val; W, Trp; and Y, Tyr. (B)sse1 seeds complemented by the SSE1 transgene (10). (C) SSE1 complementation ofpex16 mutants pex16-1 and P16KO-8A(11) for growth on oleic acid as sole carbon source (23). E122 is the wild-type strain. (D) SSE1 complementation of pex16-1mutant for the dimorphic transition from yeast to mycelia form. Cells were grown at 30°C in YND liquid medium (11). TheSSE1 transformant underwent dimorphic transition at a lower frequency than the wild-type strain E122.

Peroxisomes are not generally found in dry seeds (14) (Fig. 1). Protein and oil bodies are the most abundant organelles in mature Arabidopsis seeds, and the formation of both is ER-dependent (2, 15,16). By analogy to the function of Pex16p in peroxisome assembly and cell surface protein transport, SSE1 could participate in oil body formation and storage protein delivery. The vesicles and stacks of membranes in the sse1 cells (Fig. 1D) resemble the subcellular structures in the pex16-1 mutant of Y. lipolytica (11). The similarities between oil body and peroxisome biogenesis are consistent with their related functions in germinating seedlings of fat-metabolizing plants, where oil bodies are broken down by glyoxysomes (14).

SSE1 gene expression was analyzed by competitive reverse transcription–polymerase chain reaction (RT-PCR) (Fig. 4). The amount of SSE1mRNA is reflected in the target-to-competitor cDNA ratio (17). SSE1 steady-state mRNA level in the siliques increases during seed maturation to a maximum in mature 19- and 21-day-old brown siliques. The level of mRNA is also high in cotyledons of germinating seedlings and flowers, but low in expanding leaves and roots. Glyoxysomes are assembled in germinating seedlings (14); therefore, SSE1 is likely to be required in this process. The low expression in expanding leaves, where leaf peroxisomes are formed, may be due to low peroxisome abundance. Alternatively, SSE1 may not normally be involved in peroxisome and glyoxysome formation; rather, expression in germinating seedlings may be required for maintenance of the remaining oil bodies. The high expression levels in flowers suggests additional functions of SSE1, possibly the formation of oil body–like organelles in tapetum and pollen (16).

Figure 4

Competitive RT-PCR analyses ofSSE1 expression profiles. RNA was isolated from flowers before (B), on the day of (0), or 1 day after pollination (1); from siliques 3 to 21 days after pollination; from cotyledons of 2-day-old seedlings; and from expanding rosette leaves and roots. An equal amount of competitor cDNA template was included in each reaction. The SSE1 target (T)-to-competitor (C) cDNA ratios reflect the relative expression levels of the SSE1gene (17).

Efficient use of limited amounts of assimilates for seed storage deposition requires coordinated metabolic pathways and organelle assembly. In sse1 mature embryos, cotyledon and hypocotyl cells accumulate excess starch (Fig. 1). The functional similarity of SSE1 and Pex16p argues against SSE1 being a direct inhibitor of starch synthesis; rather, it implies that protein and oil body proliferation repress starch accumulation. Starch accumulation may also be a secondary effect of the lec mutations (18). Consistent with the observations inArabidopsis, simultaneous reduction in storage proteins and increase in starch content were also observed in a soybean shriveled seed mutant (19). Thus, in at least some species of flowering plants, starch accumulation maybe a default storage deposition pathway during seed development.

  • * Present address: Phylos, Lexington, MA 02138, USA.

  • Present address: Biosources Technologies, Vacaville, CA 95688, USA.

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