Storage protein changes during zygotic embryogenesis in interior spruce

Tree Physiology 8,7 l-8 1 0 1991 Heron Publishing-Victoria,

Canada

Storage protein changesduring zygotic embryogenesis in interior spruce BARRY S. FLINN,’ DANE R. ROBERTS, DAVID T. WEBB* and BEN C. S. SUTTON Forest Biotechnology Centre, British Columbia Research Corporation, 3650 Wesbrook Mall, Vancouver, B.C., Canada V6S 2L2

ReceivedJune 12, 1989 Summary The major storage proteins isolated from protein bodies of embryo tissues of interior spruce Picea glauca (Moench) VosslPicea engelmanii Parry had apparent molecular weights of 41, 35, 33, 24 and 22 kD. Minor proteins of 30 and 27.5 kD were also observed. Based on their solubility characteristics, the 41 kD protein was identified as a water and buffer-soluble albumin, and the 35, 33,24 and 22 kD proteins were characterized as buffer-insoluble, high salt-soluble globulins. Two-dimensional electrophoresis revealed each protein was composed of several isoelectric variants. Developmentally specific accumulation of storage proteins was observed during embryogenesis. The 41 kD protein only accumulated during the later stages of cotyledon maturation, whereas the other storage proteins began to accumulate during the early stages of embryo development. All storage proteins showed major accumulations during cotyledon maturation.

Introduction Recently, Gifford (1988) identified the major seed storage proteins of several Pinus species. Storage proteins of several Pinus species have been characterized with respect to their solubility characteristics (Katsuta 1959, 1961), and quantitative changes in storage lipids and proteins have been reported for some Pinus species (Johnson et al. 1987). However, few qualitative analyses have been carried out to determine changes in storage proteins during conifer embryo development. The purpose of this study was to identify and characterize storage proteins in interior spruce and to describe their temporal pattern of accumulation during zygotic embryo development. Materials

and methods

Plant material Interior spruce from the interior of British Columbia represents a mixture of two closely related species (Picea glauca (Moench) Voss and Picea engelmanii Parry), ’ Author to whom correspondence should be addressed. 2 Present address: Forest Biology Division, The Institute of Paper Science and Technology, 575 14th Street NW, Atlanta, GA 30318, USA.

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FLINN,

ROBERTS,

WEBB

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SU’lTON

which hybridize with one another (Owens and Molder 1984). Seed cones from source tree EK 10, which was open pollinated and possessed a Picea engelmanii maternal background, were collected on July 13, July 27 and August 24,1987. Cones and mature seed were stored at 4 “C. Embryos were excised from the seed, classified according to their developmental stage based on morphological criteria established by Buchholz and Stiemert (1945), and stored at -80 “C until required. Stage 2 embryos were torpedo shaped, possessed a visible apical dome, but lacked cotyledon primordia. Stage 3 embryos possessed cotyledons that had not developed above the apical dome, whereas Stage 4 embryos had cotyledons that completely covered the apical dome. Stage 4 embryos from the three collection dates had cotyledons of varying degrees of development, i.e., Stage 4-3 embryos (collected on August 24) were more developed than Stage 4-2 embryos (collected on July 27), which were more developed than Stage 4-l embryos (collected on July 13). Embryos from desiccated seed were classified as mature. Plant tissues Seeds were dissected and embryos and megagametophytes removed. Mature embryos were subdivided into cotyledons, shoot apical meristem including 1 mm of hypocotyl, and the remaining hypocotyl. To study the effects of germination on embryo proteins, seeds were surface sterilized in 10% Chlorox containing 0.1% Tween 80 for 15 min, washed three times with sterile distilled water and imbibed overnight at 4 “C in the dark. Imbibed seeds were placed on a water-saturated Kimpak inside a Magenta GA7 vessel and germinated at 27 “C in a 16-h photoperiod with a light intensity of 30 l.trnol m-* s-’ . Seeds were sampled at three-day intervals, characterized by the degree of radicle emergence, and germinants of similar development were removed and stored at -80 “C until analyzed for protein. Protein body isolation and analysis Protein bodies were isolated from mature embryos as described previously (Roberts et al. 1989). The protein body pellet from 80-100 embryos was resuspended in 80-100 ~1 of extraction buffer. To separate buffer-soluble and buffer-insoluble proteins (Gifford 1988), the protein body pellet was extracted with 35 ~1 of 0.05 M sodium phosphate buffer, pH 7.5, centrifuged for 10 min at 16 000 g and the supematant removed. The pellet was re-extracted and the supematant fractions pooled to give the buffer-soluble fraction. This was mixed with an equal volume of sodium dodecylsulfate (SDS) solubilizing buffer (562.5 ~10.5 M Tris-HCl (pH 6.8), 225 pl2-mercaptoethanol, 225 ~1 glycerol, 90 mg SDS, 0.5 mg bromophenol blue) and treated as described below. The remaining pellet was extracted in 100 l.tl of a l/l mixture of sodium phosphate buffer/SDS solubilizing buffer, centrifuged, and the supematant designated the buffer-insoluble fraction. To identify the solubility characteristics of the storage proteins by Osborne’s (1924) criteria, the protein body pellet derived from 50 mature embryos was extracted with 25 ~1 of either distilled water, 0.05 M sodium phosphate buffer, buffer containing 0.2 M NaCI, or buffer containing

INTERIOR

SPRUCE

STORAGE

PROTEINS

73

1 M NaCl. After extraction, the samples were centrifuged as described above, the supematant removed and the pellet re-extracted. The two supematant fractions per extraction were pooled, mixed with an equal volume of SDS solubilizing buffer and treated as described below. Electrophoresis

For analysis of proteins by sodium dodecylsulfate-polyacrylamide gel electrophoresis (SDS-PAGE), embryonic or megagametophytic tissues were homogenized with 30 to 40 l.tl SDS solubilizing buffer per mg tissue. For extractions under non-reducing conditions, 2-mercaptoethanol was omitted from the SDS solubilizing buffer. Samples were boiled for 6 min, then centrifuged for 10 min at 16 000 g and stored at -80 “C. Protein was determined by a modification of the method described by Ghosh et al. (1988), in which protein dot blots were incubated in 1 ml of 1% SDS for 1.5 h and measured spectrophotometrically at 595 run. Bovine serum albumin was used as a standard. For two-dimensional electrophoresis, embryos were homogenized in modified 2D-MH extraction buffer (9.6 ml of the modified extraction buffer contained 100 pl Pharmalyte 3-10, 175 mg NaCl, 3.7 mg NaaDTA, 3.8 mg EGTA, 200 p.1 Triton X-100, 8.8 mg ascorbic acid, 154 mg dithiothreitol, 100 ltg leupeptin, 100 Itg a2-macroglobulin, 0.2 M SDS and distilled water) (Mayer et al. 1987) at a ratio of 30 ~1 buffer per mg tissue in microfuge tubes. The homogenate was incubated with 2 mg protamine sulfate ml-’ extraction buffer for 10 min at room temperature and then centrifuged for 10 min at 16 000 g. The supematant, excluding the floating lipid layer, was removed and solid urea was added to a final concentration of 9.5 M. Extracts were stored at -80 “C until analyzed. Just before sample application, additional Pharmalyte (5-8 or 3-10) was added at a ratio of 4 ~1 for each 96 ~1 of sample. For one-dimensional SDS-PAGE analysis, the Laemmli buffer system (Laemmli 1970) was used. For two-dimensional analysis (IEF followed by SDS-PAGE), samples were run through a 10 cm tube gel (2 mm diameter). The gel solution contained 48.6 g urea, 28.8 ml H20, 11.8 ml acrylamide-bis-acrylamide (31% T, 4.3% C), 20.3 ml 10% Triton X-100,4.5 ml Pharmalyte 5-8 and 0.5 ml Pharmalyte 3-10 and was polymerized with ammonium persulfate and TEMED. The sample was applied to the cathodic end and covered with overlay buffer (2 ml 10% Triton X-100, 450 ul Pharmalyte 5-8, 50 ~1 Pharmalyte 3-10,7.5 ml H20). The cathode buffer was 0.1 N NaOH and the anode buffer was 0.06% phosphoric acid. Gels were run at 400 V constant voltage to a total of 7100 Vh. After electrophoresis, tubes were incubated on ice for 20 min, gels were then extruded, incubated in SDS solubilizing buffer for 20 min, and applied to a 4% stacking gel. This was overlaid with 1% agarose in 12.5 mM Tris-HCl, pH 6.8. After electrophoresis, gels were fixed and stained with Coomassie R-250 or silver stained (Wray et al. 1981). Microscopy

Late maturation

stage embryos were fixed, embedded, sectioned and stained as

FLINN,

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described previously (Roberts et al. 1989). Results

Identification

and characterization

of storage proteins

Both embryo and megagametophyte tissues contained prominent proteins with apparent molecular weights of 41, 33,24 and 22 kD (Figures 1A and 1B). A protein band of 35 kD, which was prominent in the megagametophyte protein profile (Figure lB), was also detectable in the whole embryo protein profile (cf. Figures 1A and 1B). Two major protein bands of 17 and 16 kD were observed in embryo tissues, but not in megagametophyte tissue. No major protein differences were observed among the various embryonic tissues. To determine if any of the proteins were storage proteins, embryo protein profiles were examined during germination (Figure 2). By Day 3 of germination, at which time radicle emergence had occurred, the 41,35,33,24 and 22 kD proteins were almost undetectable. However, the 17 and 16 kD proteins were still visible by Day 9 of germination (Figure 2). To test the assumption that the 41, 35, 33, 24 and 22 kD proteins were storage proteins, proteins were extracted from protein bodies isolated from embryonic tissue and analyzed electrophoretically. The isolated protein body profile was composed of several prominent bands and revealed proteins of 41, 35, 33,24 and 22 kD (Figure 3). Minor protein bands of 30 and 27.5 kD were also observed in the protein body fraction and these proteins were prominent in megagametophyte tissue (cf. Figures 1B and 3).

Figure 1. SDS-PAGE of total proteins stained with Coomassie blue from (A) embryonic tissues and (B) megagametophyte tissue. (A) Lane 1, cotyledon tissue; Lane 2, shoot apical meristem and 1 mm of hypocotyl; Lane 3, remaining hypocotyl tissue. (B) Lane 1, megagametophyte tissue. MW, molecular weight standards. Ten pg protein was applied to each lane. Arrows denote the 41, 35,33,24,22, 17 and 16 kD proteins.

INTERIOR SPRUCE STORAGE PROTEINS

i

204

5

MW

75

kD

97.4 66.2 42.7

31.0

21.5 14.4

Figure 2. SDS-PAGE of Coomassie-stained total embryo proteins during germination. Lane 1, mature embryo; Lane 2,3 days after sowing; Lane 3,6 days after sowing; Lane 4,9 days after sowing; Lane 5, 12 days after sowing. MW, molecular weight standards. Lane 1 was loaded with 15 pg protein, all other lanes were loaded with 10 pg protein. The 41,35,33,24 and 22 kD proteins (solid arrows) as well as the 17 and 16 kD proteins (open arrows) are indicated.

E PB MW kD

Figure 3. SDS-PAGE of proteins stained with Coomassie blue from mature embryos (E) and isolated protein bodies (PB). MW, molecular weight standards. Fifteen pg protein was applied to each lane. Arrows denote the 41,35,33, 30,27.5,24 and 22 kD proteins.

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To determine possible structural associations between individual proteins, extracts from isolated protein bodies were examined by SDS-PAGE under both reducing and non-reducing conditions (Figure 4). The characteristic patterns of storage proteins observed under reduced conditions were not seen under non-reduced conditions. In non-reduced conditions, the 33,24 and 22 kD proteins were absent, but proteins with apparent molecular weights in the range of 55-57 kD were evident, suggesting that disulfide linkages exist between the 33 kD protein and the 24 and 22 kD proteins (Figure 4). When isolated protein bodies were extracted with buffer, only the 41 kD protein was extracted (Figures 5 and 6). The remaining proteins were extracted with SDS-containing buffer (Figure 5) or with buffer containing 1 M NaCl (Figure 6). Microscopic analysis of protein bodies from nearly mature embryos revealed two distinct protein-staining regions (Figure 7), indicating the heterogeneous nature of the proteins within these organelles.

Figure 4. SDS-PAGE of isolated protein bodies analyzed under reduced (Lane 1) and non-reduced (Lane 2) conditions. The 41, 35, 33, 30, 27.5, 24 and 22 kD proteins are indicated (solid arrows). The 55-57 kD protein band observed under non-reduced conditions is indicated (open arrow). Five pg protein was applied to each lane.

MWl

2

42.7

Figure 5. SDS-PAGE of buffer-soluble isolated protein bodies. A 10 pl sample

(Lane 1) and buffer-insoluble (Lane 2) proteins extracted from was applied to each lane. MW, molecular weight standards.


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Figure 6. Coomassie-stained SDS-PAGE of water soluble (Lane l), buffer soluble (Lane 2), low salt soluble (Lane 3) and high salt soluble (Lane 4) proteins extracted from isolated protein bodies. A 10 ~1 sample was applied to each lane. MW, molecular weight standards.

Two-dimensional electrophoresis of storage proteins Protein body samples were subjected to two-dimensional electrophoresis using both narrow (5-8 ampholytes) and broad (3-10 ampholytes) pH ranges (Figure 8). Many of the single protein bands observed with one-dimensional electrophoresis contained several isoelectric variants. The 41 kD protein consisted of several polypeptides, which were found toward the basic end of the 5-8 gel. The 35 and 33 kD proteins consisted of several isoelectric variants, with the 33 kD protein consisting of several variants that extended from the acidic to the basic end of the 5-8 gel. The 30 and 27.5 kD proteins were located toward the acidic end of the 5-8 gel, whereas the 24 and 22 kD proteins contained several variants, some of which were very basic and more readily observed in the 3-10 gel (Figure SB).

Storage protein accumulation during embryo development To follow the temporal appearance of storage proteins, embryos ranging in development from Stage 2 to maturity were analyzed by one- and two-dimensional electrophoresis. The results are summarized in Table 1. Major accumulations of storage proteins, especially the 41 kD protein (Figure 9) occurred after Stage 4-1, when cotyledons were already well developed. The 24 and 22 kD proteins were detectable by Stage 3, the first stage at which cotyledons were visible (Figure 9). Although the 35,33,30 and 27.5 kD proteins did not show major accumulations until Stage 4, they were detectable in Stage 2 embryos (Figure 9).

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FLINN, ROBERTS, WEBB AND SUTTON

Figure 7. Light micrograph of a longitudinal cotyledon section from a late maturation stage embryo, stained by the periodic acid-Schiff’s technique and aniline blue black. Protein bodies (PB) containing light and dark staining zones are visible. N = nucleus, x 1350.

Figure 8. Silver-stained, two-dimensional electrophoretogram of protein body extracts examined using (A) pH 5-8 ampholytes and (B) pH 3-10 ampholytes. The acidic (+) and the basic (-) ends of the gel are indicated. The storage proteins are enclosed in boxes to allow easy visualization. Twenty ug of protein was applied to each lane. MW, molecular weight standards.

Discussion The major storage proteins in embryo and megagametophyte tissues of interior spruce had apparent molecular weights of 41,35,33,24 and 22 kD. Some of these proteins were similar to those described in Pinus by Gifford (1988) who identified two groups of proteins of 31-34.5 kD and 21.5-22.5 kD as storage proteins, as well as a 43 kD protein, which disappeared during germination and was suggested to play a storage role. We also identified minor proteins around 30 and 27.5 kD in protein


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Table 1. The presence (+) or absence (-) of various storage proteins during embryo development. Storage protein

Embryo developmental stage

WI

2

3

3-4

4-l

4-2

4-3

Mature

41 35 33 30 27.5 24 22

+ + + + -

+ + + + + +

+ + + + + +

+ + + + + +

+ + + + + + +

+ + + + + + +

+ + + + + + +

body preparations that disappeared during germination. The 30 and 27.5 kD proteins were more prominent in megagametophyte than in embryo protein profiles. The 41 kD protein could be classified as a buffer-soluble albumin, and the other major proteins as globulins. The major proteins in mature embryos of Pinus densifora and Pinus thunbergiana are albumins (Katsuta 1959, 1961). Based on the criteria described by Gifford (1988), the buffer-soluble proteins could be considered as matrix proteins, and the buffer-insoluble, high salt-soluble proteins as crystalloid proteins. Microscopic examination of the protein bodies revealed two protein-staining zones similar to the matrix and crystalloid regions described by Lott (1980). Structural relationships between proteins of the crystalloid fraction, similar to those observed with interior spruce under non-reduced conditions, appear common among different species (Tully and Beevers 1976, Gifford 1988). Two dimensional electrophoresis of storage proteins revealed that they were heterogeneous in isoelectric point, which is characteristic of storage proteins from many species (Spencer and Higgins 1982).

kD

MW

97.4 *~~. ” 66.2 -

1

2

,3”

4...

5

,cJ ,,;a ‘.

Figure 9. SDS-PAGE of total proteins stained with Coomassie blue from embryos at different developmental stages. Lane I, Stage 2; Lane 2, Stage 3; Lane 3, Stage 3-4; Lane 4, Stage 4-1; Lane 5, Stage 4-2; Lane 6, Stage 4-3; Lane 7, Mature embryos. Arrows denote the storage proteins. Twelve pg protein was applied to each lane. MW, molecular weight standards.

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FLINN, ROBERTS, WEBB AND SUTTON

Numerous studies with angiosperms have shown that storage proteins accumulate in protein bodies during cotyledon development (Harris and Boulter 1976, Adler and Muntz 1983, Greenwood and Chrispeels 1985). We detected low amounts of the 35, 33,30 and 27.5 kD proteins as early as Stage 2, although major accumulations of all proteins did not occur until the late stages of development. The accumulation of storage proteins during the early stages of embryo development has been observed in some angiosperm species (Dure and Galau 198 1, Rahman et al. 1982, Spencer et al. 1980). Dure (1985) reported that storage protein messenger RNAs begin accumulation during early embryogenesis. In contrast, the major matrix protein (41 kD) in interior spruce embryos was not detected in pre-cotyledonary embryos, but only accumulated during the later stages of embryo maturation. Although most storage proteins in conifer seeds are located in megagametophyte tissue (Bewley and Black 1985), this study was initiated to identify embryonic storage proteins and to follow their accumulation as markers for comparing development in somatic and sexually produced embryos. Such comparisons have been useful in evaluating somatic embryogenesis in alfalfa (Redenbaugh et al. 1986, Stuart et al. 1988). Acknowledgments The authors thank Dr. Gyula Kiss and Giselle Phillips of the British Columbia Ministry of Forests for the supply of interior spruce material, and Gerry O’Neil for photo preparations. This work was partially funded by the Canadian Forestry Service and the British Columbia Ministry of Forests through the Forest Resource Development Agreement. References Adler, K. and K. Muntz. 1983. Origin and development of protein bodies in cotyledons of Viciufuba. Planta 157:401-410. Bewley, J.D. and M. Black. 1985. Seeds: physiology of development and germination. Plenum Press, New York, 367 p. Buchholz, J.T. and M.L. Stiemert. 1945. Development of seed and embryos in Pinus ponderosa, with special reference to seed size. Trans. IIl. Acad. Sci. 38:27-50. Dure, L. 1985. Embryogenesis and gene expression during seed formation. Oxford Surveys Plant Mol. Cell Biol. 2: 179-197. Dure, L.S. and G.A. Galau. 1981. Developmental biochemistry of cottonseed embryogenesis and germination. XIII. Regulation of biosynthesis of principal storage proteins. Plant Physiol. 68:187194. Ghosh, S., S. Gepstein, J. Heikkila and E.B. Dumbroff. 1988. Use of a scanning densitometer or an ELISAplate reader for measurement of nanogram amounts of protein in crude extracts from biological tissues. Anal. B&hem. 169:227-233. Gifford, D.J. 1988. An electrophoretic analysis of the seed proteins from Pinus mnticolu and eight other species of pine. Can. J. Bot. 66:1808-1812. Greenwood, J.S. and M.J. Chrispeels. 1985. Immunocytochemical localization of phaseolin and phytohemagglutinin in the endoplasmic reticulum and Golgi complex of developing bean cotyledons. Planta 164:295-302. Harris, N. and D. Boulter. 1976. Protein body formation in cotyledons of developing cowpea Vignu unguiculutu seeds. Ann. Bot. 40:739-744. Johnson, M.A., J.A. Carlson, J.H. Conkey and T.L. Noland. 1987. Biochemical changes associated with zygotic pine embryo development. J. Exp. Bot. 38:518-524.


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Katsuta, M. 1959. Physiological studies of the ripening and germination processes of pine seeds. I. Changes of seed proteins. Bull. Tokyo Univ. Forests 55: 125-159. Katsuta, M. 1961. The synthesis of reserve protein in ripening pine seeds. J. Jap. For. Sot. 43:157-161. Laemmli, U.K. 1970. Cleavage of structural proteins during the assembly of the head of bacteriophage T4. Nature 227:680-685. Lott, J.N.A. 1980. Protein bodies. In The Biochemistry of Plants, AComprehensive Treatise: Volume 1, The Plant Cell. Eds. P.K. Stumpf and E.E. Conn. Academic Press, New York, pp 589-623. Mayer, J.E., G. Hahne, K. Palme and J. Schell. 1987. A simple and general plant tissue extraction procedure for two-dimensional gel electrophoresis. Plant Cell Rep. 6:77-8 1. Osborne, T.B. 1924. The vegetable proteins. In Monographs in Biochemistry. Eds. R.H. Plummer and F.G. Hopkins. Longmans, Green and Co., London, 154 p. Owens, J.N. and M. Molder. 1984. The reproductive cycle of interior spruce. B.C. Ministry of Forests, Victoria, B.C. Rahman, S., P.R. Shewry and B.J. Miflin. 1982. Differential protein accumulation during barley grain development. J. Exp. Bot. 33:717-728. Redenbaugh, K., B.D. Paasch, J.W. Nichol, M.E. Kossler, P.R. Viss and K.A. Walker. 1986. Somatic seeds: encapsulation of asexual plant embryos. BioTechnology 4797-801. Roberts, D.R., B.S. Flinn, D.T. Webb, F.B. Webster and B.C.S. Sutton. 1989. Characterization of immature embryos of interior spruce by SDS-PAGE and microscopy in relation to their competence for somatic embryogenesis. Plant Cell Rep. 8:2875-288. Spencer, D. and T.J.V. Higgins. 1982. Seed maturation and deposition of storage proteins. In The Molecular Biology of Plant Development, Botanical Monographs, Vol. 18. Eds. H. Smith and D. Grierson. University of California Press, Berkeley, pp 306-336. Spencer, D., T.J.V. Higgins, S.C. Button and R.A. Davey. 1980. Pulse-labeling studies on protein synthesis in developing pea seeds and evidence of a precursor form of legumin small subunit. Plant Physiol. 66:510-515. Stuart, D.A., J. Nelsen and J.W. Nichol. 1988. Expression of 7s and 11s alfalfa seed storage proteins in somatic embryos. J. Plant Physiol. 132: 134-139. Tully, R.E. and H. Beevers. 1976. Protein bodies of castor bean endospenn. Plant Physiol. 58:710-716. Wray, W., T. Boulikas, V.P. Wray and R. Hancock. 1981. Silver staining of proteins in polyacrylamide gels. Anal. Biochem. 118: 197-203.