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Assembly of silkmoth chorion proteins: In vivo patterns of disulfide bond formation
Insect Biochem. Vol. 18, No. 5, pp. 471-482, 1988 Printed in Great Britain.All rights reserved
0020-1790/88 $3.00+ 0.00 Copyright © 1988PergamonPress pie
ASSEMBLY OF SILKMOTH CHORION PROTEINS: IN VIVO PATTERNS OF DISULFIDE BOND FORMATION JEROME C. REGIER1 and JAMESR. WONG 2 Center for Agricultural Biotechnology of the Maryland Biotechnology Institute and Department of Entomology, University of Maryland, College Park, MD 20742, U.S.A. 2Dana-Farber Cancer Institute, Harvard Medical School, 44 Binney Street, Boston, MA 02115, U.S.A. (Received 30 November 1987; accepted 10 February 1988) Abstract--The developing silkmoth chorion is completely soluble in the absence of a sulfhydryl reducing agent until near the end of choriogenesis. During this time, ~<6% of all cysteine residues are present as cystine. Then, all chorion proteins rapidly (4-8 h) become completely resistant to solubilization. At the same time, cysteine is converted to cystine. Complete conversion extends beyond choriogenesis for at least 1-2 days. During choriogenesis, low-molecular-weight disulfide-bonded chorion protein multimers form that are soluble in the absence of a reducing agent. These multimers assemble inside folli~e cells within minutes after protein synthesis, are apparently secreted normally and remain intact for at least 2 h. Only a relatively minor fraction of the total chorion proteins participate in multimer formation. Even so, all classes of chorion proteins are present in multimers. Furthermore, certain linkage patterns are preferred. In particular, El and E2 crosslink only with themselves and with each other. Class B proteins crosslink abundantly with themselves and, at times, with class A proteins, but As do not abundantly crosslink with themselves. These multimers are good candidates as early intermediates in the assembly pathway of chorion. Key Word Index: silkmoth chorion, protein assembly, disulfide bond formation, cysteine content
INTRODUCrION A molecular analysis of silkmoth chorion assembly is a realistic goal of substantial interest. There are few morphogcnetic systems today that have been characterized as extensively as the chorion--at the levels of gene structure and organization, gene expression, protein synthesis, protein secondary structure, ultrastructure and evolution (see Regier and Kafatos, 1985 and Rcgier, 1988 for reviews). These studies arc largely directed at different levels of organization, and an understanding of their interrelationships is a primary goal. At present, the relationship between the structure and expression of individual macromolecules, on the one hand, and ultrastructure and evolution, on the other, is least well understood. In the case of chorion, this identifies the realm of chorion protein assembly. Assembly of the chorion may be substantially simpler to describe than that of many other complex, but biologically interesting, examples of morphogenesis. F o r example, unlike insect cuticle, the chorion consists almost entirely of protein. All of these proteins are separable without invoking procedures that break peptide bonds. In general, post-transitional modifications play a negligible role in generating additional biochemical diversity (reviewed in Rcgier and Kafatos, 1985). Furthermore, the chorion is extracellular; thus, conveniently separating the events that occur intracellularly from those that occur extracellularly. ts. IS/5--D
471
Previously, Blau and Kafatos (1979) have described two operationally defined processes that occur sequentially during assembly in the extracellular chorion--fixation and cementing. In the case of fixation, chorion proteins can be extracted into a supernatant phase with difficulty in water or sodium dodecyl sulfate (SDS) in the absence of a sulfhydryl reductant, whereas with cementing, a reductant is necessary for solubilization. Blau and Kafatos also reported that a substantial fraction (>I 70%) of chorion proteins in a choriogenic follicle can be solubilized in 1% SDS alone. However, at the time of ovulation, solubility properties change dramatically such that 80% of chorion proteins are now insoluble without a reductant. This study suggests that disulfide bond formation at ovulation leads to a more impenetrable shell. This conclusion is supported by laser Raman spectroscopy, where it was shown that free cysteine peaks clearly detectable in choriogenic follicles are absent in an ovulated follicle (Hamodrakas et al., 1982). The present report provides a quantitative basis for relating disulfide bond formation to changes in chorion solubility. By using more vigorous extraction procedures, we also demonstrate that cementing does not begin before the very end of choriogenesis and is then completed rapidly. We extend analysis of chorion assembly to the intracellular environment by identifying and characterizing a set of low-molecularweight disulfide-bonded chorion protein multimers. Upon secretion, these multimers may serve as nuclei
472
JEROMEC. REGIERand JAMESR. WONG
for additional assembly, that eventually leads to fixation and cementing. Although further studies will be required to establish their significance, these multimers provide the first biochemical handle for dissecting the pathway of chorion morphogenesis. MATERIALS AND METHODS
Animals and follicles Antheraea polyphemus pupae were stored at 5°C until needed and were allowed to initiate adult development at room temperature. Approximately 15 days later (Berger and Kafatos, 1971), choriogenic and ovulated follicles were dissected from females and placed in Grace's tissue culture medium without hemolymph (Grace, 1962). The choriogenic follicles were staged after labeling with [3H]leucine (Paul and Kafatos, 1975). Filler extraction Filler-forming proteins (called E1 and E2; see Regier, 1986) were washed off the chorion's surface in the following manner: 716 laid eggs (approx. 4 ml packed vol) were stirred vigorously in 8 ml of 6 M urea, 2*/0 SDS, 0.12 M Tris-HC1 (pH 8.45) and 0.054 M iodoacetamide at room temperature for 17 h. None of the follicles dissolved or even burst during this treatment. The wash was spun at 24,000 g for 30 rain at room temperature. The supernatant was fully reduced and the cysteine residues carboxamidomethylated with iodoacetamide. The pellet and the extracted follicles (minus oocyte) were dissolved in 6M urea, 2% SDS, 0.36M Tris-HCl (pH 8.45), 0.286 M 2-mercaptoethanol, followed by S-carboxamidomethylation. Determination of free cysteine concentrations Choriogenic follicles were cut into quarters and the oocyte removed. One quarter of each choriogenic follicle was used for developmental staging. Follicle cells were removed by forceps from the other 3 pieces after briefly (~< 1 rain) soaking the fragments in water. One chorion fragment was completely dissolved in 8 M gnanidinium hydrochloride, 0.36 M Tris-HC1 (pH 8.48), 0.04 M dithiothreitol, followed by quantitative carboxamidomethylation of cysteine residues (method A). Another piece was incubated in 8 M gnanidinium hydrochloride, 0.36M Tris-HC1 (pH 8.48), 0.027 M iodoacetamide for 1 h at room temperature (method B). The last piece was incubated in 0.36M Tris-HC1 (pH 8.48), 0.027 M iodoacetamide for I h at room temperature (method C). Then, the 3 chorion pieces (plus any associated dissolved proteins) were dialyzed against water, lyophilized, hydrolyzed in 6 M HCl and their amino acid compositions determined. The rationale of this experiment was that the sum of (reduced) cysteine plus (oxidized) cystine would be quantitated as carboxymethylcysteine by method A, but that only (reduced) cysteine would be measured as carboxymethylcysteine by methods B and C, thus permitting an indirect estimate of cystine content. Ovulated chorions were similarly analyzed. Extraction of chorion protein multimers Follicles were cultured in the presence of [3H]leucine for varying time periods (5 min-2 h), occasionally followed by a non-radioactive incubation (up to 4h). After oocyte removal, the follicle (with or without cells, depending on the stage of the follicle and on the particular experiment) was stirred vigorously with an extraction buffer for a minimum of l0 rain. Longer extraction times were without effect. The composition of extraction buffer was typically 8 M urea, 2% SDS, 0.05M Tris-HCl (pH7.5), 0.054M iodoacetamide. The inclusion of iodoacetamide in the extraction buffer was essential to prevent subsequent/n vitro disulfide bond formation. Similar results were obtained with an extraction buffer of pH 8.5. In one experiment (see Fig. 1),
[~4C]iodoacetamide was included in the extraction buffer to label multimers in vitro. Electrophoretic fraetionation began as soon as the extraction was completed.
Electrophoresis SDS-polyacrylamide slab gels used for fractionating chorion proteins have been described (Goldsmith et al., 1979). In Fig. 2 only, the resolving gel composition was 15°/. acrylamide, 0.4% methylenebisacrylamide and no urea was included. SDS-polyacrylamide gels run under reducing conditions included 5 mM mereaptoacetic acid in the reservoir buffer; this was essential to prevent disulfide bond formation during electrophoresis. For electrophoresis in the second dimension, a narrow strip of the first dimension SDS-polyacrylamide gel was placed horizontally on top of the second dimension gel and embedded in sample buffer (either with 0.72 M 2-mercaptoethanol or without, depending on whether reducing conditions were required) containing 1% agarose. Gels were impregnated with 2,5-diphenyloxazole, vacuum dried and exposed to X-ray film for autoradiography (Lasky and Mills, 1975). RESULTS
Chorion solubility The chorion is highly structured throughout its morphogenesis and is quite resistant to dissolution in non-denaturing, aqueous solutions (Kawasaki et al., 1971). However, chorions isolated from all but the latest stages of choriogenesis can be almost completely dissolved with protein denaturing reagents, e.g. 6 M urea. At ovulation and thereafter, dissolution requires the further addition of a sulfhydryl reducing agent. Thus, cementing (see Introduction) is rapid and occurs around the time of ovulation. To illustrate this, a developmental series of 19 choriogenic follicles plus the first three post-choriogenic or ovulated follicles were extracted with protein denaturants in the absence of a reductant. The insoluble residue of each follicle was solubilized with the addition of a reductant and fractionated on a S D S polyacrylamide gel (Fig. 3A). Two proteins, which we have identified as E l and E2, behave differently from other chorion proteins in that they can be extracted with variable efficiency from the insoluble, ovulated chorion in the absence of a reducing agent (Fig. 3A, B; Blau and Kafatos, 1979). Despite their extractability, E 1 and E2 proteins are not soluble, based on the fact that they can be pelleted with a relatively low speed spin (Fig. 3B). As for the other chorion proteins, E1 and E2 dissolution require a reducing agent; i.e. they are cemented. Previous studies have shown that E1 and E2 uniquely assemble to form the filler substructure, the bulk of which is localized as little bundles resting on the chorion's outer surface (Mazur et al., 1980). These bundles of filler can be easily washed off the surface in distilled water. Thus, the extractability of El and E2 in the absence of reducing reagent results from lack of attachment to the bulk of the chorion rather than from solubilization. Kinetics o f disulfide bond formation A n obvious conclusion from the dramatic loss of chorion solubility at ovulation is that cystine formation has occurred. Less obvious, however, is the extent of cystine formation. To examine this, we
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Fig. 1. Autoradiograms of multimers labeled in vivo for increasing time periods. A stage Xa follicle was cut into quarters and the oocyte removed. Three of the pieces were cultured in the presence of [3Hlleucin e for 5, 30 and 120 rain followed by a 2-rain nonradioactive "chase". These samples were then treated with extraction buffer and electrophoretically fractionated as described in Fig. 6A (see panels in this figure labeled Xa,7' and Xa,122'; the Xa,32' sample was identical to the other two and is not shown). The remaining follicle fragment was treated with extraction buffer containing [n4C]iodoacetamide and fractionated as above (Xa,T). In the bottom right panel (Xc,5'), a stage Xc follicle was cut in half and the oocyte removed. The follicle was cultured in the presence of [3Hlleucine for 5 min. The cells were removed from the chorion, treated with extraction buffer and fractionated as above. The dashed line links components o f a single multimeric complex. The composition of the two prominent multimeric complexes in Xc,5' are indicated. Arrowheads identify monomeric proteins that have formed intramolecular disulfide bonds.
473
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Fig. 2. Autoradiograms of multimers from staged follicles:compositional analysis. (A) A stage Xc follicle was cultured in the presence of [3H]leucine for 2~ h, further incubated with nonradioactive leucine for 2 h, cut in half to remove the oocyte, treated with extraction buffer and fractionated by diagonal gel electrophoresis as described in Fig. 6A. (B) As above, except the follicle (stage Xd) was cultured in the presence of [3H]leucine for 75 min and "chased" for 1 h. Dashed lines identify components of individual multimeric complexes. The compositions of many multimers are indicated.
474
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Fig. 3. Electropherograms of chorion proteins from staged follicles after treatment with extraction buffer. (A) Nineteen individual follicles representing all stages of choriogenesis (the most mature labeled - I and the least mature labeled - 19) and the first three ovulated follicles ( + 1 to +3, with + 1 the most recently ovulated) were extracted in 6 M urea, 2% SDS, 0.36 M Tris-HCl (pH 8.45), 0.135 M iodoacetamide with continuous, but gentle, shaking for 1 h. The insoluble membrane was isolated, dissolved by addition of reducing agent, S-carboxamidomethylated, fractionated by SDS-PAGE, stained with Coomassie Brilliant Blue and photographed. The patterns for samples - 5 to - 1 9 were very similar to that for - 4 and are not shown. The fraction of each extracted eggshell shown is 0.29 (lanes - 3 , - 4 ) , 0.051 (lanes - 2 , - 1) and 0.077 (lanes + 1 to + 3). (B) Ovulated, laid eggshells were vigorously stirred overnight with extraction buffer (see Materials and Methods). The liquid was centrifuged. Proteins in the supernatant were reduced, S-carboxamidomethylated, fractionated by SDS-PAGE, stained with Coomassie Brilliant Blue and photographed (lane 1). Proteins in the pellet (lane 2) and in eggshells (unextracted, lane 3; extracted, lane 4) were dissolved by the inclusion of a reductant and treated as above. Lanes 1 and 2 contain proteins extracted from the equivalent of 1 eggshell. Lanes 3 and 4 contain that from the equivalent of 0.040 eggshell. Classes of chorion proteins are identified on the side margins.
475
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Fig. 5. Autoradiograms o f electrophoretically fractionated proteins from extracts ofchoriogenic follicles-without and with sulfhydryl reduction. Follicles were cultured in the presence of [3H]leucine for 2 h, further incubated with nonradioactive leucine for 1 h, treated with extraction buffer (no reductant) and split in half. One half was immediately fractionated on an SDS-polyacrylamide gel (part A). The other half was reduced prior to eleetrophoresis (part B). Bands or clusters of bands that are specific to unreduced samples are identified by solid circles and stars. Individual lanes are identified by their developmental stage (Paul and Kafatos, 1975).
476
Chorion protein assembly have directly quantitated the amount of free cysteine in staged chorions before and after sulfhydryl reduction (Fig. 4). The difference between these two measurements is the cystine content. Until the last 3% of choriogenesis, i.e. up to stage Xc +, free cysteine accounts for at least 94% of the total, Subsequently, the free cysteine concentration declines, such that by the fifteenth ovulated follicle its value is only about 9% of the total. In order to quantitate free cysteine accurately, it was necessary to modify it by carboxamidomethylation prior to acid hydrolysis of the chorion. Surprisingly, almost identical results are obtained when the carboxamidornethylation reaction occurs in a simple aqueous buffer (see solid circles) and in 8 M guanidinium hydrochloride (see solid triangles). An interpretation of this finding is in the Discussion.
Identification of low-molecular-weight disulfide-bonded multimers During choriogenesis, a small fraction of total cysteine residues form disulfide bonds, as implied by Fig. 4. Some of these bonds are intermolecular and lead to the formation of protein multimers that are soluble in the presence of protein denaturants. To illustrate this, staged choriogenic follicles were Devetopmentat
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Fig. 4. Stage-specific changes in the free cysteine content of chorions. Staged follicles were cut into sections and the chorion isolated. Cysteine concentrations were measured in three different ways for chorion sections from each follicle (see Material and Methods for details). Method A, the chofion was completely dissolved with a denaturant and a reductant. Method B, the chorion was treated with a denaturant alone. Method C, the chorion was treated with an aqueous buffer alone. The samples were S-carboxamidomethylated with iodoacetamide and hydrolyzed. The carboxymethylcysteine concentration of samples treated by methods B and C are expressed as a fraction of the total carboxymethylcysteine measured by method A (see • for the B/A ratio and • for the C/A ratio). These fractions remained uniformly high, prior to stage VIII (data not shown). From stages VIII to Xd and after, the carboxymethylcysteine content measured by method A varied from 6.6-7.1 M%. Symbols are connected only if all samples were obtained from the same animal. Amino acids other than cysteine were not affected by methods A-C (data not shown). The lower ordinate identifies follicle position within the ovariole ( - l , last choriogenic follicle; + l, first postchoriogenic follicle; L, laid follicle). The upper ordinate identifies the synthetic stage of ehoriogenic follicles (Paul and Kafatos, 1975).
477
cultured in the presence of [3H]leucine and then extracted in the absence of a reducing agent. An aliquot of each solubilized extract was subsequently reduced. Reduced and unreduced aliquots were fractionated in parallel on SDS-polyacrylamide gels (Fig. 5). A brief description of follicle staging is necessary for understanding this and subsequent figures: Cboriogenesis has been divided into 10 stages, I-X, with each stage corresponding to the accumulation of 10% of the final dry wt. Stages II-IX are approximately equal in duration (3 h), whereas I has been subdivided into five 3-h periods (Ia to Ie) and X into four (Xa to Xd). Prominent differences between the two sets of samples are observed. In particular, up to stage IX, unreduced samples contain two broad bands from apparent M, 29,000 to 35,000 (see stars) that are absent from reduced samples. These bands are embedded within a much broader and highly complex array of additional faint bands that are also restricted to the unreduced samples. Together, these bands account for approx. 5-10% of total leucine incorporation at these stages. After stage IX, these bands become much fainter and a new subset of higher apparent M r bands appear specifically in the unreduced samples (see solid circles). Compositional analysis of the two most prominent clusters of multimers demonstrates that they consist largely of A- and B-size proteins (see below). A and B proteins have been estimated to have M, ranges of 8000o13,000 and 14,000-22,000. Thus, the prominent multimers are most likely to be dimers and/or trimers, based on Mr comparisons of multimers and their monomers. Accurate Mr measurements are not possible because multimers are branched structures rather than linear structures and because Mr measurements of small proteins by SDS-polyacrylamide gel electrophoresis (PAGE) is less accurate. To increase the resolution of disulfide bonded multimers, we have utilized a 2-D electrophoretic separation technique, hereafter, called diagonal gel electrophoresis (Wang and Richards, 1974; Sommer and Traut, 1975; Hynes and Destre, 1977). In the first dimension multimers are fractionated on a SDSpolyacrylamide gel under conditions where they remain intact. Then, a gel strip from the first dimension is embedded horizontally on an identical gel that is run in a second dimension under reducing conditions (Fig. 6A). Multimers in the first dimension migrate more rapidly in the second dimension due to their reduced size, whereas, proteins that were already fully reduced in the first dimension migrate identically in both dimensions. Therefore, as a group fully reduced monomeric proteins lie on a straight line diagonal. Monomeric proteins that contain intramolecular disulfide bonds may migrate more rapidly in the first dimension than in the second because of their initially more compact structure (see arrowheads in upper left panel of Fig. l; Hynes and Destree, 1977). As controls, we have shown that all proteins lie on the straight line diagonal if disulfide bonds are reduced prior to electrophoresis in the first dimension (Fig. 6B) or if they are maintained intact in both dimensions (Fig. 6C). We have also shown that briefly boiling the non-reduced extract prior to electrophoresis is without effect on the final pattern (unpublished observation). Thus, disulfide-bonded
478
JEROMEC. REGmRand JAMESR. WONG
multimers are stable during electrophoresis. In general, prechoriogenic follicles synthesize proteins of higher Mr, and fewer multimers are detectable (Fig. 6D).
Timing and localization of multimer formation To determine how rapidly multimers form, we cultured pieces of a stage Xa follicle in the presence of [3H]leucine for different times, followed by extraction and diagonal gel electrophoresis (upper panels in Fig. 1). In parallel, an equivalent follicle piece was labeled in vitro with [t4C]iodoacetamide to identify the total population of multimers (lower left panel in Fig. 1). Surprisingly, multimer formation is detectable as early as 7 min after protein synthesis. After that, multimers do not significantly increase in concentration relative to total monomers. This is clearest when examining autoradiograms in which monomer spots are much less intense than those shown. (Such autoradiograms are not shown because the multimer pattern is too faint for good reproduction.) Furthermore, multimers do not increase in size with increasing age. That is, multimer sizes (consistent with being largely dimers) are unchanged after 122rain and, in fact, are very similar to the total multimer population (see lower left panel in Fig. 1). Thus, once formed, multimers appear to maintain a constant size throughout choriogenesis, i.e. up to 2 days (Paul and Kafatos, 1975). The composition of multimers also remains constant for at least 122min, although we cannot exclude the possibility of exchange between similarly sized multimeric subunits. Multimers containing E1 and E2 proteins are assembled more rapidly than those containing A, B and C proteins. For example, at stage Xc, multimers that are ~<5 min old consist almost exclusively of E1 and E2 (lower right panel in Fig. 1). Subsequently, A-, B- and C-containing multimers form (see Fig. 2A). The rapid formation of multimers relative to protein transport to the chorion (t~/2=20min, Blau and Kafatos, 1978) strongly suggests that multimer formation is an exclusively intracellular process. To demonstrate that multimers form intracellularly, a choriogenic follicle was cultured in the presence of [3H]ieucine for 2 h, rapidly separated into its cellular and chorion compartments, extracted and fractionated by diagonal gel electrophoresis (Fig. 7). In a similar experiment, in which the follicle was labeled for only a few minutes, multimers were found exclusively in the cellular fraction (unpublished observation). An important inference from the cellular location of multimer formation is that individual multimers must consist of proteins synthesized at similar times. Of course, once the multimer is transported to the chorion, subunit exchange via shuffling of disulfide bonds is theoretically possible. However, the observed stability of multimer composition makes this less likely. Another important inference from Fig. 7 is that multimers and monomers appear to be transported to the chorion with similar kinetics, based on the similar ratios of multimers to monomers in the two fractions. Additional timepoints are clearly needed to confirm this.
Multimer composition All proteins in a single multimeric complex will, after reduction, fall along a vertical line below the straight line diagonal (see dashed line in lower right panel of Fig. 1). If a multimer contains components of identical sizes, e.g. as in a homodimer, then only one spot below the straight line diagonal will be present. In general, the multimeric patterns are quite complex to interpret because of the large number of chorion proteins. The complexity is substantially reduced, although still high, when analyzing only newly synthesized proteins. Nevertheless, it is clear that all classes of proteins participate in multimer formation (Figs 1, 2, 6 and 7). Two particularly straightforward patterns are presented in Fig. 2, although even here substantial interpretation is required. At stage Xa (part A), AI, A4 and B groups of proteins form dimers with themselves and with each other in all combinations. The high intensity of A1-B, A4-B and B-B dimers relative to A1-A1, A1-A4 and A4-A4 dimers implies distinct preferences for disulfide bond formation, perhaps based on nearest neighbor relationships. The conclusion that B-B dimers are more abundant than A - A dimers appears to be true at all stages of choriogenesis (Figs 1, 6 and 7; unpublished observations). The relative abundance of A-B heterodimers is more variable. Multimers that are larger than dimers are clearly evident in Fig. 2A (see bracket at bottom to identify this region for A- and B-containing multimers). Nevertheless, these represent a relatively small fraction of total multimers, as is more evident when the autoradiographic exposure is not as intense. A lighter exposure also reveals that only 5-10% of the newly synthesized proteins are present in multimers, consistent with our previous interpretation of Fig. 5. At stage Xd (Fig. 2B), E1 forms a prominent homodimer and, presumably, a faint homotrimer (see also lower fight panel in Fig. 1B). El also forms a prominent heterodimer with E2. Larger muitimers are evident. DISCUSSION
Cementing of chorion proteins The importance of disulfide bond formation for stabilizing the chorion has been recognized previously (Kawasaki et al., 1971), and this process has been called cementing (Blau and Kafatos, 1979). We have extended these studies by demonstrating that cementing is confined to a relatively short period (approx. 8 h) around the time of ovulation (Fig. 3A). Previous studies by Blau and Kafatos (1979) were inconclusive as to whether or not cementing occurred prior to ovulation. For example, up to 30% of proteins in a mid-choriogenic follicle failed to be extracted in the absence of a reductant. We have shown that these proteins are not cemented, i.e. held together in an insoluble form by disulfide bonds. Their solubility in the absence of a reductant was revealed by using stronger protein denaturants (urea or guanidinium hydrochloride instead of SDS) in an extraction buffer of known pH and, possibly, by shaking more vigorously during the extraction. We have also shown that
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Fig. 6. Identification of disulfide-bonded multimers after diagonal gel electrophoresis and demonstration that they are not experimental artifacts. A follicle (stage Id) was cultured in the presence of [3H]leucine for 2 h, cut in half and the oocyte removed. One half was treated with extraction buffer (no reductant) and fractionated by SDS-PAGE under oxidizing conditions. The gel strip containing the fractionated proteins was embedded horizontally on top of a second, identical SDS-polyacrylamide gel and run under reducing conditions (part A). The other half was completely dissolved with extraction buffer plus reductant and fractionated as in part A (see part B). Another chorion (stage Xc, no cells present) was treated with extraction buffer containing [~4C]iodoacetamide and fractionated as in part A, except that the proteins were not reduced prior to or during the second dimension of electrophoresis (part C). A prechoriogenic follicle was treated as in part A (see part D).
479
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Fig. 7. Autoradiograms of newly synthesized multimers present within cellular and within chorion compartments. A stage IX follicle was cut in half and cultured in the presence of [3H]leucine for 2 h. Cells were separated from the chorion with forceps, after briefly (~< 1 min) soaking the half follicles in water; this water treatment greatly facilitates complete separation of cells and chorion, Each was treated with extraction buffer and fractionated by diagonal gel electrophoresis as described in Fig. 6A. A, cells only; B, chorion only.
480
Chorion protein assembly all proteins in the ovulated chorion are cemented, even E 1 and E2 that remain extractable due to their unusual surface localization (Fig. 3B). Based on these findings, future studies directed at understanding the mechanism of cementing should focus on changes associated with ovulation. In particular, at the very end of choriogenesis, follicle cell contacts with the surface of the oocyte are loosened. Then, at ovulation, the follicle cells slough off, leaving the chorion exposed to hemolymph (King and Aggarwal, 1965). It would be reasonable to test whether this new environment favors cystein¢ oxidation, possibly through the introduction of a catalyst, e.g. metal ion or enzyme (see Margaritis, 1985 and references therein), or by a shift in pH.
Disulfide bond formation --general features The cementing of the chorion at ovulation correlates with the onset of a major decline in (reduced) cysteine (Fig. 4), Interestingly, the decline in cysteine that leads to complete cementing in the first postchorionating follicle is quite small in absolute terms, i.e. 9% of total. We estimate that it takes 1-2 additional days before cysteine has fallen to < 10% of its original value (Paul and Kafatos, 1975). In the course of quantitating cysteine residues in staged chorions, it was discovered that iodoacctamide was as effective in modifying free cysteine when dissolved in a non-denaturing buffer as in a denaturing buffer (Fig. 4). This is surprising because the uitrastructure of the chorion is unaffected by a nondenaturing buffer, whereas, in a denaturing buffer, choriogenic follicles are completely dissolved. This demonstrates that virtually all cysteine residues, which are largely confined to the arm portions of chorion sequences (see Regier and Kafatos, 1985), are freely accessible to the aqueous environment, even in an ovulated chorion.
The significance of multimer formation The formation of small amounts (5-10% of total) of generally low-molecular-weight, disulfide-bonded protein multimers throughout most of choriogenesis has been documented (Fig. 5). Multimer formation during choriogenesis and cementing around the time of ovulation appear to represent quite distinct phenomena, despite the fact that both depend on formation of disulfide bonds. We base this on the observation that multimers increase neither in size nor in concentration (relative to monomers) as choriogenesis proceeds. That is, we find no evidence that multimers eventually become insolubilized any more rapidly than monomers, e.g. see lower left panel in Fig. 1. Furthermore, insolubilization at ovulation appears to affect all chorion proteins equally, whereas multimer formation shows selectivity (see below). Thus, the biological significance of multimers may be quite different than that of cementing. Several lines of evidence are consistent with the hypothesis that multimers represent normal intermediates in the pathway of chorion protein assembly. For one, multimers form in vivo; i.e. they are not artifacts of extraction or of electrophoresis, as great care was taken to stabilize simultaneously both free cystein¢ (by carboxamidomethylation) and disulfide bonds (by exposure to air, by avoiding strongly
481
alkaline pH, by avoiding exposure to metal ions). Secondly, multimer formation occurs intracellularly, and probably not at all in the chorion (Figs 1 and 7; unpublished observations). This observation implies specificity of interaction because chorion protein concentrations are undoubtedly much higher in the chorion than intracellularly. There is precedent for the hypothesis that disulfide bonds form intracellularly. Tartakoff et al. (1979) showed that immunoglobulin M subunits, which are secreted, form disulfide bonds with each other while still in the rough endoplasmic reticulum. This covalent association appears to be preceded by noncovalent association. The secretory kinetics of chorion proteins are also consistent with disulfide bond formation occurring in the rough endoplasmic reticulum or in the Golgi (Blau and Kafatos, 1978). Thirdly, multimers appear to be secreted into the extracellular space at a rate similar to that for monomers (Fig. 7). That is, multimers do not represent abortive intermediates that are not secreted or else are secreted abnormally. Fourthly, we have directly demonstrated the specificity of multimer formation. Most convincingly, we have shown that E1 and E2 proteins form multimers only with themselves, and that E multimers form more rapidly than do A and B protein multimers, even though E1 and E2 have the lowest concentrations of cysteine (lower fight panel in Fig. 1; Fig. 2A; Regier et al., 1980). The complexity of A- and B-size components makes their definitive identification more difficult. Nevertheless, the most reasonable interpretation of our data is that As and Bs form dimers with each other and with themselves. However, their ratios are not uniform. In particular, B-B dimers are always abundant relative to A - A dimers (Figs 1, 2, 6 and 7). Fifthly, and more theoretically, for a disulfide bond to be formed, the protein(s) must have the proper conformation and be in an environment with a favorable redox potential (see Creighton, 1984 for a discussion of this point). Taken together, the evidence suggests that multimers represent intermediates in chorion protein assembly. Protein assembly is generally thought to be directed via weak bonds, e.g. hydrogen bonds. This appears to be the case for chorion as well, based on the ability of 8 M urea alone to dissolve rapidly a choriogenic follicle. With this in mind, it is reasonable to propose that disulfide-bonded multimers form parts of larger intracellular complexes held together by weak bonds. Upon secretion into the extracellular chorion, these complexes could serve as nuclei for additional noncovalently-mediated assembly that leads to fixation within several hours (Blau and Kafatos, 1979). Thus, the finding that a relatively small fraction of chorion proteins form disulfide bonds is not an argument against their role as assembly intermediates, and, in fact, may provide insight into the actual mechanism. In particular, it provides a means of limiting the size of the secreted intermediate, thus, avoiding the problem of passaging the narrow-pored sieve layer that is attached to the ¢xtracellular surface of the secretory microvilli (Mazur et al., 1980; unpublished observations). In fact, previous studies have established that E proteins don't passage this sieve layer. This directly correlates with their more rapid assembly (Figs 1 and 2), with
482
JEROMEC. REGIER and JAMESR. WONG
their interactions only with themselves and not with As, Bs or Cs (see also Mazur et al., 1980), and with the unusually large size of E2 (Regier, 1986). Future studies will have to be directed at identifying the putative noncovalently-linked multimers. This will be greatly facilitated by our finding that assembly is initiated within the cell. Thus, the initial protein interactions within the cell can be separated from the latter interactions in the chorion. It should be interesting, for example, to perform diagonal gel electrophoresis of chorion multimers crosslinked in vitro with a cleavable crosslinking reagent. This might directly reveal whether the disulfide bonded multimers that form in vivo are part of a larger, noncovalently-bonded complex. Finally, it is apparent that disulfide bonds form intramolecularly as well as intermolecularly (Figs 1 and 7). These could represent intermediate stages in protein folding, either as an unassembled m o n o m e r or as a multimer joined by noncovalent bonds. Acknowledgements--We gratefully acknowledge Drs V. Veletza for participation in the early phases of the project and F. C. Kafatos for providing laboratory space and helpful comments on the manuscript. This research was supported by a grant from the National Institutes of Health to JCR and from the National Science Foundation to F. C. Kafatos. REFERENCES
Bcrger E. and Kafatos F. C. (1971) Quantitative studies of prococoonase synthesis and accumulation during development. Devl Biol. 25, 377-397. Blau H. M. and Kafatos F. C. 0978) Secretory kinetics in the follicular ceils of silkmoths during eggshell formation. J. Cell Biol. 78, 131-151. Blau H. M. and Kafatos F. C. (1979) Morphogenesis of the silkmoth chorion--pattems of distribution and insolubilization of the structural proteins. Devl Biol. 72, 211-225. Creighton T. E. (1984) Disulfide bond formation in proteins. In Methods in Enzymology, Vol. 107 (Edited by Wold F. and Moldave K.), pp. 305-329. Academic Press, New York. Goldsmith M. R., Rattner E. C., Koehler M. D., Balikov S. R. and Bock S. (1979) Two-dimensional electrophoresis of small-molecular-weight proteins. Analyt. Biochem. 99, 33-40. Grace T. D. C. (1962) Establishment of four strains of cells from insect tissues grown in vitro. Nature, Lond. 195, 788-789. Hamodrakas S. J., Asher S. A., Mazur G. D., Regier J. C. and Kafatos F. C. (1982) Laser Raman studies of protein
conformation in the silkmoth chorion. Biochim. Biophys. Acta 703, 216-222. Hynes R. O. and Destree A. (1977) Extensive disulfide bonding at the mammalian cell surface. Proc. natn Acad. Sci. U.S.A. 74, 2855-2859. Kawasaki H., Sato H. and Suzuki M. (1971) Structural proteins in the silkworm egg-shells. Insect Biochem. 1, 130-148. King R. C. and Aggarwal S. K. (1965) Oogenesis in Hyalophora cecropia. Growth 29, 17-83. Lasky R. A. and Mills A. D. (1975) Quantitative film detection of 3H and 14C in polyacrylamide gels by fluorography. Eur. J. Biochem. 56, 335-341. Margaritis L. (1985) The egg-shell of Drosophila melanogaster III. Covalent crosslinking of the chorion proteins involves endogenous hydrogen peroxide. Tissue Cell 17, 553-559. Mazur G. D., Regier J. C. and Kafatos F. C. (1980) The silkmoth chorion: morphogenesis of surface structures and its relation to synthesis of specific proteins. Devl Biol. 76, 305-321. Paul M. and Kafatos F. C. (1975) Specific protein synthesis in cellular differentiation. II. The program of protein synthetic changes during chorion formation by silkmoth follicles, and its implementation in organ culture. Devl Biol. 42, 141-159. Regier J. C. (1986) Evolution and higher-order structure of architectural proteins in silkmoth chorion. EMBO Jl 5, 1981-1989. Regier J. C. (1988) Evolution in steps--The role of regulatory alterations in the diversification of the moth chorion morphogenetic pathway. In Self-Assembling Architecture. Forty-sixth Symposium of the Society for Developmental Biology (Edited by Varner J.), Liss, New York. In press. Regier J. C. and Kafatos F. C. (1985) Molecular aspects of chorion formation. In Comprehensive Insect Physiology, Biochemistry and Pharmacology, Vol. 1 (Edited by Kerkut G. A. and Gilbert L. I.), pp. 113-151. Pergamon Press, Oxford. Regier J. C., Mazur G. D. and Kafatos F. C. (1980) The silkmoth chorion: Morphological and biochemical characterization of four surface regions. Devl Biol. 76, 286-304. Sommer A. and Traut R. R. (1975) Identification by diagonal gel electrophoresis of nine neighboring protein pairs in the Escherichia coil 30S ribosome crosslinked with methyl-4-mercaptobutyrimidate. J. molec. Biol. 97, 471-481. Tartakoff A., VassaUi P. and I~traz M. (1979) Plasma cell immunogiobulin M molecules--their biosynthesis, assembly, and intracellular transport. J. Cell Biol. 83, 284--299. Wang K. and Richards F. M. (1974) An approach to nearest neighbor analysis of membrane proteins. J. biol. Chem. 249, 8005-8018.
0020-1790/88 $3.00+ 0.00 Copyright © 1988PergamonPress pie
ASSEMBLY OF SILKMOTH CHORION PROTEINS: IN VIVO PATTERNS OF DISULFIDE BOND FORMATION JEROME C. REGIER1 and JAMESR. WONG 2 Center for Agricultural Biotechnology of the Maryland Biotechnology Institute and Department of Entomology, University of Maryland, College Park, MD 20742, U.S.A. 2Dana-Farber Cancer Institute, Harvard Medical School, 44 Binney Street, Boston, MA 02115, U.S.A. (Received 30 November 1987; accepted 10 February 1988) Abstract--The developing silkmoth chorion is completely soluble in the absence of a sulfhydryl reducing agent until near the end of choriogenesis. During this time, ~<6% of all cysteine residues are present as cystine. Then, all chorion proteins rapidly (4-8 h) become completely resistant to solubilization. At the same time, cysteine is converted to cystine. Complete conversion extends beyond choriogenesis for at least 1-2 days. During choriogenesis, low-molecular-weight disulfide-bonded chorion protein multimers form that are soluble in the absence of a reducing agent. These multimers assemble inside folli~e cells within minutes after protein synthesis, are apparently secreted normally and remain intact for at least 2 h. Only a relatively minor fraction of the total chorion proteins participate in multimer formation. Even so, all classes of chorion proteins are present in multimers. Furthermore, certain linkage patterns are preferred. In particular, El and E2 crosslink only with themselves and with each other. Class B proteins crosslink abundantly with themselves and, at times, with class A proteins, but As do not abundantly crosslink with themselves. These multimers are good candidates as early intermediates in the assembly pathway of chorion. Key Word Index: silkmoth chorion, protein assembly, disulfide bond formation, cysteine content
INTRODUCrION A molecular analysis of silkmoth chorion assembly is a realistic goal of substantial interest. There are few morphogcnetic systems today that have been characterized as extensively as the chorion--at the levels of gene structure and organization, gene expression, protein synthesis, protein secondary structure, ultrastructure and evolution (see Regier and Kafatos, 1985 and Rcgier, 1988 for reviews). These studies arc largely directed at different levels of organization, and an understanding of their interrelationships is a primary goal. At present, the relationship between the structure and expression of individual macromolecules, on the one hand, and ultrastructure and evolution, on the other, is least well understood. In the case of chorion, this identifies the realm of chorion protein assembly. Assembly of the chorion may be substantially simpler to describe than that of many other complex, but biologically interesting, examples of morphogenesis. F o r example, unlike insect cuticle, the chorion consists almost entirely of protein. All of these proteins are separable without invoking procedures that break peptide bonds. In general, post-transitional modifications play a negligible role in generating additional biochemical diversity (reviewed in Rcgier and Kafatos, 1985). Furthermore, the chorion is extracellular; thus, conveniently separating the events that occur intracellularly from those that occur extracellularly. ts. IS/5--D
471
Previously, Blau and Kafatos (1979) have described two operationally defined processes that occur sequentially during assembly in the extracellular chorion--fixation and cementing. In the case of fixation, chorion proteins can be extracted into a supernatant phase with difficulty in water or sodium dodecyl sulfate (SDS) in the absence of a sulfhydryl reductant, whereas with cementing, a reductant is necessary for solubilization. Blau and Kafatos also reported that a substantial fraction (>I 70%) of chorion proteins in a choriogenic follicle can be solubilized in 1% SDS alone. However, at the time of ovulation, solubility properties change dramatically such that 80% of chorion proteins are now insoluble without a reductant. This study suggests that disulfide bond formation at ovulation leads to a more impenetrable shell. This conclusion is supported by laser Raman spectroscopy, where it was shown that free cysteine peaks clearly detectable in choriogenic follicles are absent in an ovulated follicle (Hamodrakas et al., 1982). The present report provides a quantitative basis for relating disulfide bond formation to changes in chorion solubility. By using more vigorous extraction procedures, we also demonstrate that cementing does not begin before the very end of choriogenesis and is then completed rapidly. We extend analysis of chorion assembly to the intracellular environment by identifying and characterizing a set of low-molecularweight disulfide-bonded chorion protein multimers. Upon secretion, these multimers may serve as nuclei
472
JEROMEC. REGIERand JAMESR. WONG
for additional assembly, that eventually leads to fixation and cementing. Although further studies will be required to establish their significance, these multimers provide the first biochemical handle for dissecting the pathway of chorion morphogenesis. MATERIALS AND METHODS
Animals and follicles Antheraea polyphemus pupae were stored at 5°C until needed and were allowed to initiate adult development at room temperature. Approximately 15 days later (Berger and Kafatos, 1971), choriogenic and ovulated follicles were dissected from females and placed in Grace's tissue culture medium without hemolymph (Grace, 1962). The choriogenic follicles were staged after labeling with [3H]leucine (Paul and Kafatos, 1975). Filler extraction Filler-forming proteins (called E1 and E2; see Regier, 1986) were washed off the chorion's surface in the following manner: 716 laid eggs (approx. 4 ml packed vol) were stirred vigorously in 8 ml of 6 M urea, 2*/0 SDS, 0.12 M Tris-HC1 (pH 8.45) and 0.054 M iodoacetamide at room temperature for 17 h. None of the follicles dissolved or even burst during this treatment. The wash was spun at 24,000 g for 30 rain at room temperature. The supernatant was fully reduced and the cysteine residues carboxamidomethylated with iodoacetamide. The pellet and the extracted follicles (minus oocyte) were dissolved in 6M urea, 2% SDS, 0.36M Tris-HCl (pH 8.45), 0.286 M 2-mercaptoethanol, followed by S-carboxamidomethylation. Determination of free cysteine concentrations Choriogenic follicles were cut into quarters and the oocyte removed. One quarter of each choriogenic follicle was used for developmental staging. Follicle cells were removed by forceps from the other 3 pieces after briefly (~< 1 rain) soaking the fragments in water. One chorion fragment was completely dissolved in 8 M gnanidinium hydrochloride, 0.36 M Tris-HC1 (pH 8.48), 0.04 M dithiothreitol, followed by quantitative carboxamidomethylation of cysteine residues (method A). Another piece was incubated in 8 M gnanidinium hydrochloride, 0.36M Tris-HC1 (pH 8.48), 0.027 M iodoacetamide for 1 h at room temperature (method B). The last piece was incubated in 0.36M Tris-HC1 (pH 8.48), 0.027 M iodoacetamide for I h at room temperature (method C). Then, the 3 chorion pieces (plus any associated dissolved proteins) were dialyzed against water, lyophilized, hydrolyzed in 6 M HCl and their amino acid compositions determined. The rationale of this experiment was that the sum of (reduced) cysteine plus (oxidized) cystine would be quantitated as carboxymethylcysteine by method A, but that only (reduced) cysteine would be measured as carboxymethylcysteine by methods B and C, thus permitting an indirect estimate of cystine content. Ovulated chorions were similarly analyzed. Extraction of chorion protein multimers Follicles were cultured in the presence of [3H]leucine for varying time periods (5 min-2 h), occasionally followed by a non-radioactive incubation (up to 4h). After oocyte removal, the follicle (with or without cells, depending on the stage of the follicle and on the particular experiment) was stirred vigorously with an extraction buffer for a minimum of l0 rain. Longer extraction times were without effect. The composition of extraction buffer was typically 8 M urea, 2% SDS, 0.05M Tris-HCl (pH7.5), 0.054M iodoacetamide. The inclusion of iodoacetamide in the extraction buffer was essential to prevent subsequent/n vitro disulfide bond formation. Similar results were obtained with an extraction buffer of pH 8.5. In one experiment (see Fig. 1),
[~4C]iodoacetamide was included in the extraction buffer to label multimers in vitro. Electrophoretic fraetionation began as soon as the extraction was completed.
Electrophoresis SDS-polyacrylamide slab gels used for fractionating chorion proteins have been described (Goldsmith et al., 1979). In Fig. 2 only, the resolving gel composition was 15°/. acrylamide, 0.4% methylenebisacrylamide and no urea was included. SDS-polyacrylamide gels run under reducing conditions included 5 mM mereaptoacetic acid in the reservoir buffer; this was essential to prevent disulfide bond formation during electrophoresis. For electrophoresis in the second dimension, a narrow strip of the first dimension SDS-polyacrylamide gel was placed horizontally on top of the second dimension gel and embedded in sample buffer (either with 0.72 M 2-mercaptoethanol or without, depending on whether reducing conditions were required) containing 1% agarose. Gels were impregnated with 2,5-diphenyloxazole, vacuum dried and exposed to X-ray film for autoradiography (Lasky and Mills, 1975). RESULTS
Chorion solubility The chorion is highly structured throughout its morphogenesis and is quite resistant to dissolution in non-denaturing, aqueous solutions (Kawasaki et al., 1971). However, chorions isolated from all but the latest stages of choriogenesis can be almost completely dissolved with protein denaturing reagents, e.g. 6 M urea. At ovulation and thereafter, dissolution requires the further addition of a sulfhydryl reducing agent. Thus, cementing (see Introduction) is rapid and occurs around the time of ovulation. To illustrate this, a developmental series of 19 choriogenic follicles plus the first three post-choriogenic or ovulated follicles were extracted with protein denaturants in the absence of a reductant. The insoluble residue of each follicle was solubilized with the addition of a reductant and fractionated on a S D S polyacrylamide gel (Fig. 3A). Two proteins, which we have identified as E l and E2, behave differently from other chorion proteins in that they can be extracted with variable efficiency from the insoluble, ovulated chorion in the absence of a reducing agent (Fig. 3A, B; Blau and Kafatos, 1979). Despite their extractability, E 1 and E2 proteins are not soluble, based on the fact that they can be pelleted with a relatively low speed spin (Fig. 3B). As for the other chorion proteins, E1 and E2 dissolution require a reducing agent; i.e. they are cemented. Previous studies have shown that E1 and E2 uniquely assemble to form the filler substructure, the bulk of which is localized as little bundles resting on the chorion's outer surface (Mazur et al., 1980). These bundles of filler can be easily washed off the surface in distilled water. Thus, the extractability of El and E2 in the absence of reducing reagent results from lack of attachment to the bulk of the chorion rather than from solubilization. Kinetics o f disulfide bond formation A n obvious conclusion from the dramatic loss of chorion solubility at ovulation is that cystine formation has occurred. Less obvious, however, is the extent of cystine formation. To examine this, we
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Fig. 1. Autoradiograms of multimers labeled in vivo for increasing time periods. A stage Xa follicle was cut into quarters and the oocyte removed. Three of the pieces were cultured in the presence of [3Hlleucin e for 5, 30 and 120 rain followed by a 2-rain nonradioactive "chase". These samples were then treated with extraction buffer and electrophoretically fractionated as described in Fig. 6A (see panels in this figure labeled Xa,7' and Xa,122'; the Xa,32' sample was identical to the other two and is not shown). The remaining follicle fragment was treated with extraction buffer containing [n4C]iodoacetamide and fractionated as above (Xa,T). In the bottom right panel (Xc,5'), a stage Xc follicle was cut in half and the oocyte removed. The follicle was cultured in the presence of [3Hlleucine for 5 min. The cells were removed from the chorion, treated with extraction buffer and fractionated as above. The dashed line links components o f a single multimeric complex. The composition of the two prominent multimeric complexes in Xc,5' are indicated. Arrowheads identify monomeric proteins that have formed intramolecular disulfide bonds.
473
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Fig. 2. Autoradiograms of multimers from staged follicles:compositional analysis. (A) A stage Xc follicle was cultured in the presence of [3H]leucine for 2~ h, further incubated with nonradioactive leucine for 2 h, cut in half to remove the oocyte, treated with extraction buffer and fractionated by diagonal gel electrophoresis as described in Fig. 6A. (B) As above, except the follicle (stage Xd) was cultured in the presence of [3H]leucine for 75 min and "chased" for 1 h. Dashed lines identify components of individual multimeric complexes. The compositions of many multimers are indicated.
474
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Fig. 3. Electropherograms of chorion proteins from staged follicles after treatment with extraction buffer. (A) Nineteen individual follicles representing all stages of choriogenesis (the most mature labeled - I and the least mature labeled - 19) and the first three ovulated follicles ( + 1 to +3, with + 1 the most recently ovulated) were extracted in 6 M urea, 2% SDS, 0.36 M Tris-HCl (pH 8.45), 0.135 M iodoacetamide with continuous, but gentle, shaking for 1 h. The insoluble membrane was isolated, dissolved by addition of reducing agent, S-carboxamidomethylated, fractionated by SDS-PAGE, stained with Coomassie Brilliant Blue and photographed. The patterns for samples - 5 to - 1 9 were very similar to that for - 4 and are not shown. The fraction of each extracted eggshell shown is 0.29 (lanes - 3 , - 4 ) , 0.051 (lanes - 2 , - 1) and 0.077 (lanes + 1 to + 3). (B) Ovulated, laid eggshells were vigorously stirred overnight with extraction buffer (see Materials and Methods). The liquid was centrifuged. Proteins in the supernatant were reduced, S-carboxamidomethylated, fractionated by SDS-PAGE, stained with Coomassie Brilliant Blue and photographed (lane 1). Proteins in the pellet (lane 2) and in eggshells (unextracted, lane 3; extracted, lane 4) were dissolved by the inclusion of a reductant and treated as above. Lanes 1 and 2 contain proteins extracted from the equivalent of 1 eggshell. Lanes 3 and 4 contain that from the equivalent of 0.040 eggshell. Classes of chorion proteins are identified on the side margins.
475
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Fig. 5. Autoradiograms o f electrophoretically fractionated proteins from extracts ofchoriogenic follicles-without and with sulfhydryl reduction. Follicles were cultured in the presence of [3H]leucine for 2 h, further incubated with nonradioactive leucine for 1 h, treated with extraction buffer (no reductant) and split in half. One half was immediately fractionated on an SDS-polyacrylamide gel (part A). The other half was reduced prior to eleetrophoresis (part B). Bands or clusters of bands that are specific to unreduced samples are identified by solid circles and stars. Individual lanes are identified by their developmental stage (Paul and Kafatos, 1975).
476
Chorion protein assembly have directly quantitated the amount of free cysteine in staged chorions before and after sulfhydryl reduction (Fig. 4). The difference between these two measurements is the cystine content. Until the last 3% of choriogenesis, i.e. up to stage Xc +, free cysteine accounts for at least 94% of the total, Subsequently, the free cysteine concentration declines, such that by the fifteenth ovulated follicle its value is only about 9% of the total. In order to quantitate free cysteine accurately, it was necessary to modify it by carboxamidomethylation prior to acid hydrolysis of the chorion. Surprisingly, almost identical results are obtained when the carboxamidornethylation reaction occurs in a simple aqueous buffer (see solid circles) and in 8 M guanidinium hydrochloride (see solid triangles). An interpretation of this finding is in the Discussion.
Identification of low-molecular-weight disulfide-bonded multimers During choriogenesis, a small fraction of total cysteine residues form disulfide bonds, as implied by Fig. 4. Some of these bonds are intermolecular and lead to the formation of protein multimers that are soluble in the presence of protein denaturants. To illustrate this, staged choriogenic follicles were Devetopmentat
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Fig. 4. Stage-specific changes in the free cysteine content of chorions. Staged follicles were cut into sections and the chorion isolated. Cysteine concentrations were measured in three different ways for chorion sections from each follicle (see Material and Methods for details). Method A, the chofion was completely dissolved with a denaturant and a reductant. Method B, the chorion was treated with a denaturant alone. Method C, the chorion was treated with an aqueous buffer alone. The samples were S-carboxamidomethylated with iodoacetamide and hydrolyzed. The carboxymethylcysteine concentration of samples treated by methods B and C are expressed as a fraction of the total carboxymethylcysteine measured by method A (see • for the B/A ratio and • for the C/A ratio). These fractions remained uniformly high, prior to stage VIII (data not shown). From stages VIII to Xd and after, the carboxymethylcysteine content measured by method A varied from 6.6-7.1 M%. Symbols are connected only if all samples were obtained from the same animal. Amino acids other than cysteine were not affected by methods A-C (data not shown). The lower ordinate identifies follicle position within the ovariole ( - l , last choriogenic follicle; + l, first postchoriogenic follicle; L, laid follicle). The upper ordinate identifies the synthetic stage of ehoriogenic follicles (Paul and Kafatos, 1975).
477
cultured in the presence of [3H]leucine and then extracted in the absence of a reducing agent. An aliquot of each solubilized extract was subsequently reduced. Reduced and unreduced aliquots were fractionated in parallel on SDS-polyacrylamide gels (Fig. 5). A brief description of follicle staging is necessary for understanding this and subsequent figures: Cboriogenesis has been divided into 10 stages, I-X, with each stage corresponding to the accumulation of 10% of the final dry wt. Stages II-IX are approximately equal in duration (3 h), whereas I has been subdivided into five 3-h periods (Ia to Ie) and X into four (Xa to Xd). Prominent differences between the two sets of samples are observed. In particular, up to stage IX, unreduced samples contain two broad bands from apparent M, 29,000 to 35,000 (see stars) that are absent from reduced samples. These bands are embedded within a much broader and highly complex array of additional faint bands that are also restricted to the unreduced samples. Together, these bands account for approx. 5-10% of total leucine incorporation at these stages. After stage IX, these bands become much fainter and a new subset of higher apparent M r bands appear specifically in the unreduced samples (see solid circles). Compositional analysis of the two most prominent clusters of multimers demonstrates that they consist largely of A- and B-size proteins (see below). A and B proteins have been estimated to have M, ranges of 8000o13,000 and 14,000-22,000. Thus, the prominent multimers are most likely to be dimers and/or trimers, based on Mr comparisons of multimers and their monomers. Accurate Mr measurements are not possible because multimers are branched structures rather than linear structures and because Mr measurements of small proteins by SDS-polyacrylamide gel electrophoresis (PAGE) is less accurate. To increase the resolution of disulfide bonded multimers, we have utilized a 2-D electrophoretic separation technique, hereafter, called diagonal gel electrophoresis (Wang and Richards, 1974; Sommer and Traut, 1975; Hynes and Destre, 1977). In the first dimension multimers are fractionated on a SDSpolyacrylamide gel under conditions where they remain intact. Then, a gel strip from the first dimension is embedded horizontally on an identical gel that is run in a second dimension under reducing conditions (Fig. 6A). Multimers in the first dimension migrate more rapidly in the second dimension due to their reduced size, whereas, proteins that were already fully reduced in the first dimension migrate identically in both dimensions. Therefore, as a group fully reduced monomeric proteins lie on a straight line diagonal. Monomeric proteins that contain intramolecular disulfide bonds may migrate more rapidly in the first dimension than in the second because of their initially more compact structure (see arrowheads in upper left panel of Fig. l; Hynes and Destree, 1977). As controls, we have shown that all proteins lie on the straight line diagonal if disulfide bonds are reduced prior to electrophoresis in the first dimension (Fig. 6B) or if they are maintained intact in both dimensions (Fig. 6C). We have also shown that briefly boiling the non-reduced extract prior to electrophoresis is without effect on the final pattern (unpublished observation). Thus, disulfide-bonded
478
JEROMEC. REGmRand JAMESR. WONG
multimers are stable during electrophoresis. In general, prechoriogenic follicles synthesize proteins of higher Mr, and fewer multimers are detectable (Fig. 6D).
Timing and localization of multimer formation To determine how rapidly multimers form, we cultured pieces of a stage Xa follicle in the presence of [3H]leucine for different times, followed by extraction and diagonal gel electrophoresis (upper panels in Fig. 1). In parallel, an equivalent follicle piece was labeled in vitro with [t4C]iodoacetamide to identify the total population of multimers (lower left panel in Fig. 1). Surprisingly, multimer formation is detectable as early as 7 min after protein synthesis. After that, multimers do not significantly increase in concentration relative to total monomers. This is clearest when examining autoradiograms in which monomer spots are much less intense than those shown. (Such autoradiograms are not shown because the multimer pattern is too faint for good reproduction.) Furthermore, multimers do not increase in size with increasing age. That is, multimer sizes (consistent with being largely dimers) are unchanged after 122rain and, in fact, are very similar to the total multimer population (see lower left panel in Fig. 1). Thus, once formed, multimers appear to maintain a constant size throughout choriogenesis, i.e. up to 2 days (Paul and Kafatos, 1975). The composition of multimers also remains constant for at least 122min, although we cannot exclude the possibility of exchange between similarly sized multimeric subunits. Multimers containing E1 and E2 proteins are assembled more rapidly than those containing A, B and C proteins. For example, at stage Xc, multimers that are ~<5 min old consist almost exclusively of E1 and E2 (lower right panel in Fig. 1). Subsequently, A-, B- and C-containing multimers form (see Fig. 2A). The rapid formation of multimers relative to protein transport to the chorion (t~/2=20min, Blau and Kafatos, 1978) strongly suggests that multimer formation is an exclusively intracellular process. To demonstrate that multimers form intracellularly, a choriogenic follicle was cultured in the presence of [3H]ieucine for 2 h, rapidly separated into its cellular and chorion compartments, extracted and fractionated by diagonal gel electrophoresis (Fig. 7). In a similar experiment, in which the follicle was labeled for only a few minutes, multimers were found exclusively in the cellular fraction (unpublished observation). An important inference from the cellular location of multimer formation is that individual multimers must consist of proteins synthesized at similar times. Of course, once the multimer is transported to the chorion, subunit exchange via shuffling of disulfide bonds is theoretically possible. However, the observed stability of multimer composition makes this less likely. Another important inference from Fig. 7 is that multimers and monomers appear to be transported to the chorion with similar kinetics, based on the similar ratios of multimers to monomers in the two fractions. Additional timepoints are clearly needed to confirm this.
Multimer composition All proteins in a single multimeric complex will, after reduction, fall along a vertical line below the straight line diagonal (see dashed line in lower right panel of Fig. 1). If a multimer contains components of identical sizes, e.g. as in a homodimer, then only one spot below the straight line diagonal will be present. In general, the multimeric patterns are quite complex to interpret because of the large number of chorion proteins. The complexity is substantially reduced, although still high, when analyzing only newly synthesized proteins. Nevertheless, it is clear that all classes of proteins participate in multimer formation (Figs 1, 2, 6 and 7). Two particularly straightforward patterns are presented in Fig. 2, although even here substantial interpretation is required. At stage Xa (part A), AI, A4 and B groups of proteins form dimers with themselves and with each other in all combinations. The high intensity of A1-B, A4-B and B-B dimers relative to A1-A1, A1-A4 and A4-A4 dimers implies distinct preferences for disulfide bond formation, perhaps based on nearest neighbor relationships. The conclusion that B-B dimers are more abundant than A - A dimers appears to be true at all stages of choriogenesis (Figs 1, 6 and 7; unpublished observations). The relative abundance of A-B heterodimers is more variable. Multimers that are larger than dimers are clearly evident in Fig. 2A (see bracket at bottom to identify this region for A- and B-containing multimers). Nevertheless, these represent a relatively small fraction of total multimers, as is more evident when the autoradiographic exposure is not as intense. A lighter exposure also reveals that only 5-10% of the newly synthesized proteins are present in multimers, consistent with our previous interpretation of Fig. 5. At stage Xd (Fig. 2B), E1 forms a prominent homodimer and, presumably, a faint homotrimer (see also lower fight panel in Fig. 1B). El also forms a prominent heterodimer with E2. Larger muitimers are evident. DISCUSSION
Cementing of chorion proteins The importance of disulfide bond formation for stabilizing the chorion has been recognized previously (Kawasaki et al., 1971), and this process has been called cementing (Blau and Kafatos, 1979). We have extended these studies by demonstrating that cementing is confined to a relatively short period (approx. 8 h) around the time of ovulation (Fig. 3A). Previous studies by Blau and Kafatos (1979) were inconclusive as to whether or not cementing occurred prior to ovulation. For example, up to 30% of proteins in a mid-choriogenic follicle failed to be extracted in the absence of a reductant. We have shown that these proteins are not cemented, i.e. held together in an insoluble form by disulfide bonds. Their solubility in the absence of a reductant was revealed by using stronger protein denaturants (urea or guanidinium hydrochloride instead of SDS) in an extraction buffer of known pH and, possibly, by shaking more vigorously during the extraction. We have also shown that
-SH v
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(A)
(B)
(C)
(D)
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Fig. 6. Identification of disulfide-bonded multimers after diagonal gel electrophoresis and demonstration that they are not experimental artifacts. A follicle (stage Id) was cultured in the presence of [3H]leucine for 2 h, cut in half and the oocyte removed. One half was treated with extraction buffer (no reductant) and fractionated by SDS-PAGE under oxidizing conditions. The gel strip containing the fractionated proteins was embedded horizontally on top of a second, identical SDS-polyacrylamide gel and run under reducing conditions (part A). The other half was completely dissolved with extraction buffer plus reductant and fractionated as in part A (see part B). Another chorion (stage Xc, no cells present) was treated with extraction buffer containing [~4C]iodoacetamide and fractionated as in part A, except that the proteins were not reduced prior to or during the second dimension of electrophoresis (part C). A prechoriogenic follicle was treated as in part A (see part D).
479
(A)
(B)
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Fig. 7. Autoradiograms of newly synthesized multimers present within cellular and within chorion compartments. A stage IX follicle was cut in half and cultured in the presence of [3H]leucine for 2 h. Cells were separated from the chorion with forceps, after briefly (~< 1 min) soaking the half follicles in water; this water treatment greatly facilitates complete separation of cells and chorion, Each was treated with extraction buffer and fractionated by diagonal gel electrophoresis as described in Fig. 6A. A, cells only; B, chorion only.
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Chorion protein assembly all proteins in the ovulated chorion are cemented, even E 1 and E2 that remain extractable due to their unusual surface localization (Fig. 3B). Based on these findings, future studies directed at understanding the mechanism of cementing should focus on changes associated with ovulation. In particular, at the very end of choriogenesis, follicle cell contacts with the surface of the oocyte are loosened. Then, at ovulation, the follicle cells slough off, leaving the chorion exposed to hemolymph (King and Aggarwal, 1965). It would be reasonable to test whether this new environment favors cystein¢ oxidation, possibly through the introduction of a catalyst, e.g. metal ion or enzyme (see Margaritis, 1985 and references therein), or by a shift in pH.
Disulfide bond formation --general features The cementing of the chorion at ovulation correlates with the onset of a major decline in (reduced) cysteine (Fig. 4), Interestingly, the decline in cysteine that leads to complete cementing in the first postchorionating follicle is quite small in absolute terms, i.e. 9% of total. We estimate that it takes 1-2 additional days before cysteine has fallen to < 10% of its original value (Paul and Kafatos, 1975). In the course of quantitating cysteine residues in staged chorions, it was discovered that iodoacctamide was as effective in modifying free cysteine when dissolved in a non-denaturing buffer as in a denaturing buffer (Fig. 4). This is surprising because the uitrastructure of the chorion is unaffected by a nondenaturing buffer, whereas, in a denaturing buffer, choriogenic follicles are completely dissolved. This demonstrates that virtually all cysteine residues, which are largely confined to the arm portions of chorion sequences (see Regier and Kafatos, 1985), are freely accessible to the aqueous environment, even in an ovulated chorion.
The significance of multimer formation The formation of small amounts (5-10% of total) of generally low-molecular-weight, disulfide-bonded protein multimers throughout most of choriogenesis has been documented (Fig. 5). Multimer formation during choriogenesis and cementing around the time of ovulation appear to represent quite distinct phenomena, despite the fact that both depend on formation of disulfide bonds. We base this on the observation that multimers increase neither in size nor in concentration (relative to monomers) as choriogenesis proceeds. That is, we find no evidence that multimers eventually become insolubilized any more rapidly than monomers, e.g. see lower left panel in Fig. 1. Furthermore, insolubilization at ovulation appears to affect all chorion proteins equally, whereas multimer formation shows selectivity (see below). Thus, the biological significance of multimers may be quite different than that of cementing. Several lines of evidence are consistent with the hypothesis that multimers represent normal intermediates in the pathway of chorion protein assembly. For one, multimers form in vivo; i.e. they are not artifacts of extraction or of electrophoresis, as great care was taken to stabilize simultaneously both free cystein¢ (by carboxamidomethylation) and disulfide bonds (by exposure to air, by avoiding strongly
481
alkaline pH, by avoiding exposure to metal ions). Secondly, multimer formation occurs intracellularly, and probably not at all in the chorion (Figs 1 and 7; unpublished observations). This observation implies specificity of interaction because chorion protein concentrations are undoubtedly much higher in the chorion than intracellularly. There is precedent for the hypothesis that disulfide bonds form intracellularly. Tartakoff et al. (1979) showed that immunoglobulin M subunits, which are secreted, form disulfide bonds with each other while still in the rough endoplasmic reticulum. This covalent association appears to be preceded by noncovalent association. The secretory kinetics of chorion proteins are also consistent with disulfide bond formation occurring in the rough endoplasmic reticulum or in the Golgi (Blau and Kafatos, 1978). Thirdly, multimers appear to be secreted into the extracellular space at a rate similar to that for monomers (Fig. 7). That is, multimers do not represent abortive intermediates that are not secreted or else are secreted abnormally. Fourthly, we have directly demonstrated the specificity of multimer formation. Most convincingly, we have shown that E1 and E2 proteins form multimers only with themselves, and that E multimers form more rapidly than do A and B protein multimers, even though E1 and E2 have the lowest concentrations of cysteine (lower fight panel in Fig. 1; Fig. 2A; Regier et al., 1980). The complexity of A- and B-size components makes their definitive identification more difficult. Nevertheless, the most reasonable interpretation of our data is that As and Bs form dimers with each other and with themselves. However, their ratios are not uniform. In particular, B-B dimers are always abundant relative to A - A dimers (Figs 1, 2, 6 and 7). Fifthly, and more theoretically, for a disulfide bond to be formed, the protein(s) must have the proper conformation and be in an environment with a favorable redox potential (see Creighton, 1984 for a discussion of this point). Taken together, the evidence suggests that multimers represent intermediates in chorion protein assembly. Protein assembly is generally thought to be directed via weak bonds, e.g. hydrogen bonds. This appears to be the case for chorion as well, based on the ability of 8 M urea alone to dissolve rapidly a choriogenic follicle. With this in mind, it is reasonable to propose that disulfide-bonded multimers form parts of larger intracellular complexes held together by weak bonds. Upon secretion into the extracellular chorion, these complexes could serve as nuclei for additional noncovalently-mediated assembly that leads to fixation within several hours (Blau and Kafatos, 1979). Thus, the finding that a relatively small fraction of chorion proteins form disulfide bonds is not an argument against their role as assembly intermediates, and, in fact, may provide insight into the actual mechanism. In particular, it provides a means of limiting the size of the secreted intermediate, thus, avoiding the problem of passaging the narrow-pored sieve layer that is attached to the ¢xtracellular surface of the secretory microvilli (Mazur et al., 1980; unpublished observations). In fact, previous studies have established that E proteins don't passage this sieve layer. This directly correlates with their more rapid assembly (Figs 1 and 2), with
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their interactions only with themselves and not with As, Bs or Cs (see also Mazur et al., 1980), and with the unusually large size of E2 (Regier, 1986). Future studies will have to be directed at identifying the putative noncovalently-linked multimers. This will be greatly facilitated by our finding that assembly is initiated within the cell. Thus, the initial protein interactions within the cell can be separated from the latter interactions in the chorion. It should be interesting, for example, to perform diagonal gel electrophoresis of chorion multimers crosslinked in vitro with a cleavable crosslinking reagent. This might directly reveal whether the disulfide bonded multimers that form in vivo are part of a larger, noncovalently-bonded complex. Finally, it is apparent that disulfide bonds form intramolecularly as well as intermolecularly (Figs 1 and 7). These could represent intermediate stages in protein folding, either as an unassembled m o n o m e r or as a multimer joined by noncovalent bonds. Acknowledgements--We gratefully acknowledge Drs V. Veletza for participation in the early phases of the project and F. C. Kafatos for providing laboratory space and helpful comments on the manuscript. This research was supported by a grant from the National Institutes of Health to JCR and from the National Science Foundation to F. C. Kafatos. REFERENCES
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