www.elsevier.com/locate/jinsphys
Vitellogenic ovarian follicles of
Drosophila
exhibit a
charge-dependent distribution of endogenous soluble proteins
Russell W. Cole, Richard I. Woodruff
*Department of Biology, West Chester University, West Chester, PA 19383-8102, USA
Received 17 September 1999; accepted 19 January 2000
Abstract
In ovarian follicles ofDrosophila, soluble endogenous charged proteins are asymmetrically distributed dependent upon their ionic charge. Reversal of the normal ionic difference across the intercellular bridges which connect nurse cells to their oocyte results in a redistribution of these proteins. Twelve soluble endogenous acidic proteins were identified by 2-D gel electrophoresis as being present in both oocytes and nurse cells in samples run on four or more gels. Of these, following osmotically induced reversal of the electrical transbridge gradient the concentration of seven proteins decreased in the oocyte while nurse cell concentrations of all twelve proteins increased. Of seven basic proteins analyzed, following reversal of the electrical gradient the concentration of all seven increased in oocytes. Four of these decreased in nurse cells, while nurse cell concentrations of the remaining three basic proteins also appeared to decrease, but yielded spots too faint for measurement. Data presented here demonstrate that, as in the Saturniidae, the ionic gradient across the nurse cell-oocyte intercellular bridges of the dipteran, Drosophila, can influence the distribution of soluble endogenous charged molecules. 2000 Elsevier Science Ltd. All rights reserved.
Keywords:Ionic gradient; Cytosolic proteins; Membrane potential
1. Introduction
In insects, as throughout most of the animal kingdom, developing oocytes are most commonly supported by sibling cells called nurse cells, to which they are joined by open channels of cytoplasm called intercellular bridges (Telfer, 1975). During pre-ovulation develop-ment the oocyte nucleus becomes a dormant germinal vesicle (GV) and the oocyte acquires yolk via endo-cytosis (Telfer, 1965). Meanwhile, the nurse cell nuclei become highly endopolyploid and active, with most of the RNA produced destined to pass through the inter-cellular bridges and become sequestered in the oocyte (Bier, 1963a,b; Pollack and Telfer, 1969). Intercellular bridges, the products of incomplete cytokinesis, most often function to maintain synchrony among syncytial cells (Fawcett et al., 1959), yet in ovarian follicles this function must be circumvented as oocyte and nurse cells
* Corresponding author. Tel.:+1-610-436-2417; fax:+ 1-610-436-2183.
E-mail address:[email protected] (R.I. Woodruff).
0022-1910/00/$ - see front matter2000 Elsevier Science Ltd. All rights reserved. PII: S 0 0 2 2 - 1 9 1 0 ( 0 0 ) 0 0 0 4 4 - 5
become both morphologically and physiologically differ-ent (Fig. 1). As the two cell types differdiffer-entiate along separate lines some mechanism must exist which allows them to retain autonomy while still maintaining the open bridges needed to transport copious amounts of RNA
from their sites of synthesis in the nurse cells to the sites where these molecules will be sequestered in the oocyte. Particularly in Drosophila, the oocyte is precisely structured, eventually containing organization which influences post-fertilization development (Nusslein-Vol-hard et al., 1987). This organized “pre-programming” is achieved within a multicellular complex which must itself be highly and actively regulated. Indeed, the devel-opmental sequence of follicular activities that generate egg structure implies an exacting set of cellular controls. Morphological polarity, manifested as an anterior-pos-terior orientation (with respect to both the ovariole and the whole organism) of the oocyte-nurse cell syncytium, is apparent when the follicle is first formed. In Hyalo-phora cecropia, an electrically-based physiological polarity initiates close to the onset of vitellogenesis (Woodruff and Telfer 1973, 1980), and continues as a steady-state phenomenon for several days, until the nurse cells disintegrate about 24 h before the end of vitellogen-esis. This physiological polarity involves a metabolically driven difference in [Ca2+]
i between the oocyte and the
nurse cells (Woodruff et al., 1991; Woodruff and Telfer, 1994), which establishes an electrical gradient focused across the bridges connecting the two cell types. Recently, this transbridge ionic gradient has been shown to influence the distribution of charged endogenous cyto-solic proteins (Cole and Woodruff, 1997). This current, seemingly through action of the cytosolic proteins whose distribution it regulates, also enforces changes in the transcriptional activity of the oocyte nucleus (Woodruff et al., 1998).
The ovarian follicles ofDrosophilaresemble those of Hyalophora, but possess more nurse cells (n=15) and reside in a more conventional blood ion environment. Reports from different labs on the existence of a trans-bridge gradient inDrosophilahave varied. Bohrmann et al. (1986) and Sun and Wyman (1987, 1993), found no significant difference, while Woodruff et al. (1988), Woodruff (1989), Verachtert et al. (1989), Verachtert and De Loof (1989) and Singleton and Woodruff (1994) all found significant differences between the steady-state potentials of nurse cells and the oocyte to which they are attached. An experimentally supported answer explaining the differing results has now been put forth, and centers around the composition of the media in which the measurements were performed (Singleton and Woodruff, 1994). Experimental evidence revealed that the steady-state membrane potential (Em) of nurse cells
was more affected by osmolarity than Emof oocytes. The
osmolarity of adult female hemolymph was measured to be 250 mOsmol, at which osmolarity nurse cells were shown to be more electronegative than the oocyte to which they were attached. At increasingly higher osmol-arity the difference between cell types first decreased to 0 and then reversed.
Microinjection of fluorescently labeled lysozyme, in
either the positive or the negative form, has provided evidence that a charge-dependent asymmetric distri-bution of proteins can occur inDrosophila(Woodruff et al., 1988), but lysozyme is an exogenous protein in this system, and soluble endogenous proteins might be regu-lated by other means. Thus in the present study we have utilized 2-D gel electrophoresis to analyze the distri-bution of charged endogenous cytosolic proteins from the ovarian follicles of Drosophila. Soluble proteins could be susceptible to iontophoretic effects, while pro-teins bound to cytoskeletal elements, membranes and other cytoplasmic structures would not be. Bound pro-teins are present in such perfusion that, if not removed, they obscure the influence of the electrical gradient upon the soluble proteins. As in a previous study (Cole and Woodruff, 1997), a necessary step was to separate sol-uble proteins from those which were bound. To achieve this, we harvested soluble proteins from nurse cell or from oocyte extracts by centrifugation and ultrafiltration. We furthermore took advantage of the effect on the transbridge electrical gradient wrought by changes in osmolarity (Singleton and Woodruff, 1994). This pro-vided a non-invasive non-pharmacological means to reverse the direction of the gradient. If the transbridge gradient actually does influence the distribution of charged soluble molecules, the distributions of both acidic and basic proteins should be affected in opposite manners. Relative to controls incubated in a 255 mOs-mol. medium, in follicles incubated at 400 mOsmOs-mol. the concentrations of soluble acidic proteins should diminish in the oocytes, and increase in the nurse cells. Similarly, the relative concentrations of soluble basic proteins should decrease in the nurse cells and increase in the oocytes of follicles incubated at high osmolarity.
The experiments reported here show that the distri-butions of most of the soluble proteins responded to changes in the osmolarity of the incubation medium exactly as if they were responding to the transbridge electrical gradient.
2. Materials and methods
2.1. Animals
2.2. Drosophila physiological salt solution (PSS)
The physiological salt solution used during dissections and incubations was designed to have the same major ion composition and osmolarity as theDrosophila hemo-lymph which normally bathes developing follicles (Singleton and Woodruff, 1994). This PSS was made up of: 100 mM Na-glutamate, 25 mM KCl, 15 mM MgCl2,
5 mM CaSO4, 2 mM sodium phosphate buffer (pH 6.9).
Sucrose was added as needed to bring the osmolarity to 250–255 mOsmol (PSS255). For reversal of the trans-bridge ionic gradient, sucrose was added to bring the osmolarity to 400 mOsmol (PSS400). Osmolarity of sol-utions was checked using commercial osmometers; either an Osmette A (Precision Systems Inc, Natick, MA) or a Vapro 5520 (Wescor, Logan, UT).
2.3. Follicle collection and preparation
Vitellogenic follicles were dissected from female Dro-sophila in PSS255 and transferred to either fresh PSS255, or to PSS400. For this report we selected only stage 10A or 10B follicles. Following one hour incu-bation at room temperature, control or high osmolarity treated follicles were transferred to a homogenization medium consisting of 20% w/v sucrose, 2.5% w/v poly-vinylpolypyrrolidone, 10 mM CaCl2, 5 mM HEPES and
5 mM AEBSF protease inhibitor (Calbiochem, La Jolla, CA). A fresh 26-gauge hypodermic needle was attached to the needle holder of a Narishige MN-151 Emerson-type micromanipulator (Narishige Instruments, Japan), and adjusted so that the sharp edge of the tip bevel was positioned as a guillotine above the follicles. The micromanipulator was used first to lower the sharp edge onto the oocyte-nurse cell junction until the extreme tip touched the glass bottom of the working chamber. Then the tip was drawn back in a slicing motion, severing the follicle so that the nurse cell cap was precisely separated from the oocyte. Following separation neither the nurse cell cap nor the oocyte showed overt signs of leakage, suggesting that a seal was achieved as the opposing sides of the intercellular bridges were pressed together as the cut was made. Nor did Lucifer Yellow CH iontophoret-ically microinjected into oocyte or nurse cell cap reveal any sign of leakage. Since no detectable leakage was observed from either fraction, several follicles were sequentially “decapitated”, the oocytes being concen-trated in one area of the chamber and the nurse cell caps in another. Oocytes (OOC) or nurse cell caps (NCC) were drawn into a microtransfer pipette and transported to a drop of homogenization medium, volume of which was adjusted to obtain a ratio of 20µl for every 80 OOC or 80 NCC, and thence to a chilled microcentrifuge tube. The cells were pressed with a chilled glass pestle to express their cytoplasm with only minimal rupture of epithelial cells, of yolk spheres in the case of oocytes,
and of nuclei in the case of nurse cells. The samples were centrifuged at 12,000 g for 20 min at 4°C to remove intact epithelial cells, yolk spheres, nuclei, and other cellular debris. When resuspended pellets from centrifuged ooplasm were examined microscopically using SSEE optics (Ellis, 1978), yolk spheres and col-umnar follicle epithelial cells were present in abundance, while resuspended nurse plasm pellets contained many squamoid epithelial cells but few if any yolk spheres. Nor had nurse cell nuclei been lysed, but instead remained intact within the cytoplasm of nurse cells, plasma membranes of which had been ruptured (Fig. 2). No yolk spheres, epithelial cells, nuclei nor other debris were found in the supernatant. Samples were pooled until each pool contained the protein extracted from 240 follicles in each 60µl. In some cases, to more stringently insure the sample contained only the soluble proteins, the supernatant was centrifuged through a 300 kDa cut-off filter (Millipore, Bedford, MA)(6,000 g, 20–40 min, 4°C). However, because the amounts of sample were so small, and to include a small number of bound proteins which served as references as described below, this was not routinely done.
2.4. Electrophoresis
2.4.1. Sample preparation
The procedures used for two dimensional gel electro-phoresis were essentially the same as those described in an earlier study of the soluble endogenous proteins of the luna moth, Actias luna(Cole and Woodruff, 1997). Because of their importance to the results, they are repeated here. Either oocyte or nurse cell cap extracts
were mixed 1:1 with acidic or basic overlay buffers (Basic—0.2% Bio-Lyte 3/10 (Bio-Rad, Laboratories, Richmond, CA), 1.8% ampholine 8/10.5 (Sigma), 9.5 M urea, 5.0% ß-mercaptoethanol, 0.4% Nonidet P-40 (Sigma); (Acidic—0.6% Bio-Lyte 3/5, 0.7% Bio-Lyte 3/10, 9.5 M urea, 5.0% ß-mercaptoethanol, 0.4% Non-idet P-40). Buffered samples were then incubated at room temperature for 10 min.
2.4.2. First dimension: isoelectric focusing (IEF) Different acrylamide monomer solutions were pre-pared for acidic and basic proteins. For basic gels, 600 µl of 9/11 ampholine and 50 µl of 3/10 Bio-Lyte were added to a monomer solution consisting of 4.3% acryla-mide and 0.2% piperazine diacrylaacryla-mide (PDA), 9.5 M urea, 3% Nonidet P-40. For acidic gels, 450 µl of 3/5 Bio-Lyte and 50 µl of 3/10 Bio-Lyte were added to the monomer solution. Twenty microliters of TEMED (Sigma) and 60 µl 10% ammonium persulfide (APS) were added and the gels allowed to polymerize. Ten microliters of an overlay solution consisting of 1:1 mix-ture of the appropriate overlay buffer and distilled water plus enough Bromophenol Blue for visibility were added to the sample well and 30 µl of prepared sample was then introduced between the gel and the overlay solution. (Thus each gel contained the soluble proteins from 60 follicles.) Acidic protein gels were run at 5°C on a Mini PROTEAN II 2-D electrophoresis unit (Bio-Rad) at 500 V for 10 min, then 750 V for 3 h 20 min. Due to the inherent instability of gradients for high pH isoelectric points (O’Farrell et al., 1977), basic gels were run at 500 V for 10 min, then 800 V for only 1 h 30 min. Some gels from each run were extruded onto parafilm and cut into eight pieces of equal length; each piece was eluted overnight in a separate container with 0.50 ml of 50 mM KCl, and the elutate measured with a pH meter. After IEF, the pH gradient of acidic gels went from pH 3.85±0.06 (S.E.M.) to 5.87±0.12 (S.E.M.), while that of basic gels ranged from 8.0±0.1 (S.E.M.) in the first seg-ment to 9.6±0.1 (S.E.M.) in the seventh segseg-ment. Seg-ment eight, in contact with the lower tank buffer, aver-aged 11.1±0.02 (S.E.M.).
For acidic proteins, the cathode chamber was filled with 20 mM NaOH and the anode chamber with 10 mM H3PO4. For basic proteins, the anode chamber was filled
with 250 mM HEPES (Sigma) solution and the cathode chamber with 1.0 N NaOH.
2.4.3. Second dimension: separation by molecular weight
Second dimension separation was in SDS mini slab gels (8.5×7 mm, 1 mm thick) containing 12.0% acrylam-ide and 0.26% PDA dissolved in a degassed stock sol-ution of 0.6 M Tris(hydroxymethyl)aminomethane (Aldrich Chemical Company, Inc., Milwaukee, WS), 0.27 M Trizma hydrochloride (Sigma), and 1% lauryl
sulfate (Sigma). To polymerize the gel solutions, 50 µl of TEMED and 250 µl of 10% ammonium persulfate solution were added. Some second dimension separ-ations were done using pre-cast 12% acrylamide “ready-gels” (Bio-Rad).
An IEF gel was placed on the top of each slab gel, and a 1µl aliquont of SDS-PAGE low molecular weight protein standards (Bio-Rad) diluted with SDS running buffer was placed in the left hand sample well. Running buffer contained 25 mM Tris(hydroxymethyl) aminomethane, 192 mM glycine (Sigma), 0.1% lauryl sulfate), 5 µl glycerol, and Bromophenol Blue. Second dimension gels were run at 100 V for 10 min then 150 V for 1 h.
2.5. Fixing and staining
Slab gels were fixed for 20 min with 50% methanol for 12 h. Gels were than rehydrated and stained with a highly sensitive silver stain (Morrissey, 1981) reported to be 200 times more sensitive than Coomassie Blue (Deely, 1989). Stained gels were dried between two sheets of transparent cellulose (Promega, Madison, WI). Simultaneously run and stained gels of a control and of a treated sample were dried side-by-side to aid in the computer analysis of protein content.
2.6. Criteria for acceptability of protein spots for analysis
For analysis we chose proteins which gave evidence of being soluble by being detectable in ultrafiltrated sam-ple, and for acidic proteins, appearing in at least control oocyte sample and nurse cell sample from follicles incu-bated in PSS400, and in four or more gels of each type. Basic proteins were chosen if they met the ultrafiltrate criterion, and were detectable in control nurse cells and oocytes from follicles incubated in PSS400.
2.7. Computer analysis and calculations for determination of protein amounts
Dried gels were scanned with either a Kodak digital sciences DC40 camera for electrophoresis gels (Kokak, Rochester, NY) or an Epson ES-1000c flat-bed scanner (Epson Accessories, Torrance, CA) to digitize the image. No differences were found between data derived from a single gel by either device. The image was stored using a Power Macintosh computer (Apple Computer, Inc., Cupertino, CA) and retrieved using IP Lab Gel LC Ver-sion 1.1.2f (Signal Analytics Corporation, Vienna, VA). Using the IP Lab Gel LC analysis program, images of protein spots were evaluated to determine the density of each protein relative to a reference standard (Cole and Woodruff, 1997) and below.
considerable difference from fly to fly in the size of developmentally equivalent follicles. Fortunately, as in our earlier study on soluble proteins of Luna moth fol-licles, in centrifuged samples there was a small popu-lation of proteins that showed little or no change that could be correlated with changes in the transbridge ionic gradient resulting from experimental treatment, and which were removed by ultrafiltration through a 300 kDa filter. We assumed these to be proteins which in situ were bound to cytoplasmic organelles remaining in the 12,000gsupernatant, and which were solublized by urea during first dimension electrophoresis. The density of these spots thus reflected only the amount of sample loaded. Absorbancy of a bound protein spot in either control or experimental gel could thus be used as a “stan-dard” against which the absorbancy of truly soluble pro-teins could be compared. For acidic propro-teins, the average of several bound proteins (shown as “B” in Fig. 3) pro-vided a final normalization factor for each gel, and the absorbancy of each soluble protein was corrected by that factor. Thus if the average density of the bound proteins in an experimental gel was found to be 1.5 times greater than the average density of the same bound proteins in a control gel, the absorbancy of each soluble protein spot in the experimental gel was reduced by the appropriate amount. In basic gels there were far fewer proteins, and only one could be identified as a suitable reference pro-tein. Thus for basic proteins, concentration is simply reported as a percentage of the density of that reference protein. By the strategies outlined above, we determined the relative absorbancy of proteins from control follicles or from follicles incubated at high osmolarity.
3. Results
3.1. Acidic proteins
As shown in Fig. 3, by centrifugation and ultrafiltr-ation we were able to identify four distinct proteins meeting the criteria for bound reference proteins (designated as “B”). In addition we identified 12 accept-able proteins (designated by numbers 1–12), each of which met the criteria above for analysis as a cytosolic protein. Of these, in oocyte samples from 400 mOsmol incubations, there was an overall 2±1% decrease from control levels. Seven of the proteins decreased in con-centration, three increased slightly, and two increased markedly relative to their concentrations in controls (Fig. 4). Among the seven proteins which did decrease, the average change was 3.7±1%, the largest change being 9% and the smallest being 1%. By pairedt-test compar-ing the 255 mOsmol vs 400 mOsmol concentration of each protein on four or more gel pairs, P=0.28. For the seven which did decrease, P=0.01. Fig. 5 shows an enlarged portion of a gel on which was run sample from
control oocytes, and a similar section of a gel on which was run oocyte sample from follicles incubated in PSS400. One of the bound acidic proteins (marked B) can be seen, as well as several soluble acidic proteins. For convenience, the proteins of interest have been marked by arrowheads. Note that these proteins do not in all cases correspond to those proteins analyzed in Fig. 3 and Fig. 4, since not all of the proteins so clearly seen in this pair of gels were sufficiently resolved on four or more gel pairs to allow them to be used in statistical analysis. Fig. 6 shows the absorbancy profiles from a PSS255 gel and a PSS 400 gel for the bound reference protein and a cytosolic protein, spots of which appear along the cursor line marked in Fig. 5.
In nurse cell samples from follicles incubated in PSS400, all 12 of the analyzed proteins increased in con-centration. The overall average change for these 12 pro-teins was a 6.5±0.8% increase above control levels. By paired t-test, P=0.0001. The largest change was a 10% increase and the smallest change was 2%. Fig. 4 shows the changes in all 12 acidic proteins analyzed.
3.2. Basic proteins
Generally, the number and concentration of cytosolic proteins is small compared to the great number and abundance of proteins which are bound in some way within the cytoplasm and its organelles. Of cytosolic proteins, the vast majority are acidic or neutral, thus it was not surprising that there were few basic proteins which met our criteria of being present in both cell types and in four or more gels of one type. We were able to identify one bound protein which served as a reference, and seven cytosolic proteins (numbered 1+–7+) with pI’s of pH 8 or greater. Of these, three were in such low concentration in nurse cells that we could not meaning-fully analyze their concentrations as they decreased in follicles incubated in PSS400. Thus we provide data for seven proteins in oocytes incubated in PSS255 or in PSS400, but only for four of these in nurse cells. Because there was only one formerly bound basic pro-tein to serve as a reference, concentrations are given sim-ply as a percentage of that reference protein (Fig. 7).
Offsetting the small number of basic proteins accept-able for analysis was the clarity of their response. In oocytes incubated in PSS400, all seven basic cytosolic proteins increased their concentration. The average increase was 290±20% (2.9 times) above the levels found in control oocytes from PSS255. The maximum change was an increase to 385% of controls, while the minimum change was an increase to 222% of control concentration. By paired t-test, P=0.008.
con-Fig. 3. Examples of second dimension gels silver stained to show soluble endogenous proteins. (A) Oocyte acidic proteins. (B) Nurse cell acidic proteins. (C) Oocyte basic proteins. (D) Nurse cell basic proteins. Numbered spots are proteins which met the criteria described in the text for analysis of soluble proteins. The designation “+” after the identification numbers for basic proteins indicates only that they are positively charged, not their charge density. Spots marked “B” are proteins which were removed from samples passed through a 300 kDa filter, and thus are assumed to have been bound in situ. Solubilized during sample preparation, they varied only dependent upon the total amount of protein loaded in the first dimension, and thus serve as reference points. (Note: the nurse cell basic protein gel shown as “D” had greater separation in the second dimension than did its oocyte counterpart shown as “C”. In “D” protein no. 7+, which would otherwise not have shown in this composite figure, appears in the box at lower left. For A and B the pH gradient runs in each gel from pH 6 at left to pH 4 at right. For C and D the pH gradient runs in each gel from pH 8 at left to pH 9.6 at right.
trol concentrations (P=0.003). The maximum change was to only 6% of control values, while the minimum change was to 30% of control values.
4. Discussion
The data reported here reveal that: (1) oocytes and nurse cells of Drosophilaovarian follicles share several
endogenous proteins which are charged and soluble; (2) the concentrations of these proteins in each cell type changed on average in a manner consistent with their distributions being influenced by an electrical gradient in the intercellular bridges and (3) this transbridge elec-trical gradient operates as a “leaky” selective gate.
Fig. 4. Changes in the relative absorbancy (concentration) of 12 endogenous soluble acidic proteins caused in 1 h by the osmotically induced reversal of the transbridge electrical gradient between nurse cells and their attached oocyte.
concluded that “...only a small fraction of acidic and basic proteins may be subject to intrafollicular electro-phoresis, but the bulk of the proteins clearly does not migrate by way of electrophoresis”. Their methods, how-ever, included treatments which would have solubilized all proteins within the various cells of the follicle, obscuring in their number and concentrations those pro-teins which were naturally dissolved in the cytosol. Indeed, the small fraction of the proteins which they noted as perhaps being subject to the electrical gradient may well have constituted nearly all of the soluble charged proteins. It was for this reason that we took great care in obtaining only the cytosolic fraction before employing any treatments containing chemicals such as urea which would solubilize previously bound proteins.
Fig. 5. Sections of a pair of 2-D gels illustrating the change in oocyte concentration of cytosolic acidic proteins caused by reversal of the transbridge electrical gradient. The gel at left shows proteins (arrowheads) from oocytes incubated at control osmolarity of 255 mOsmol. The gel at the right shows these same proteins harvested from oocytes incubated for 1 h in medium of 400 mOsmol. Protein “B” is a normally bound protein, the concentration of which serves as a normalization factor. After incubation in medium of 400 mOsmol, while density (concentration) of protein “B” is nearly unchanged, each of the eight acidic soluble proteins shown here decreased in concen-tration, presumably due to back-diffusion into nurse cells. The cursors connecting “B” and a cytosolic protein mark the lines of the absorbancy profiles shown in Fig. 6.
By centrifugation and ultrafiltration we were able to limit our investigations to those proteins which, being soluble in the cytoplasm, would be available to be influ-enced by an electrophoretic field confined to the inter-cellular bridges. Of the acidic proteins which appeared to be soluble, all 12 increased in nurse cells following reversal of the transbridge electrical gradient. In oocytes, seven behaved in a charge-dependent manner, decreas-ing in their concentrations. Four others, each of which increased strikingly in nurse cells, also increased in oocytes, but only slightly. Another protein (designated as no. 6, Fig. 4) increased markedly in concentration in both cell types after incubation at high osmolarity with the strongest increase in the nurse cells. Two possible causes, both speculative at this time, present themselves. Firstly, all five may in fact be proteins which were very loosely bound in situ, with binding constants decreased by incubations at high osmolarity. Alternatively, these five may represent a group of proteins for which rates of synthesis are strongly affected by the osmolarity of the medium (Mizuno et al., 1984; Prince and Villarejo, 1990; Hengge-Aronis et al., 1993; Muffler et al., 1996). Osmolarity of the incubation medium has been shown to cause changes in the rates of uptake of certain precur-sor amino acids, and precurprecur-sor availability would pre-sumably have an effect on rates of protein synthesis. Methionine uptake in particular has been shown to be higher at increased osmolarity (Bohrmann, 1991). Osmolarity also has an effect on the uptake of K+, the principal ion setting the value of Em(Miyazaki and
Hagi-wara, 1976). Changes in [K+]i would, in turn, cause
pro-Fig. 6. Absorbancy profiles of gel spots representing a bound refer-ence protein and near-by cytosolic acidic protein no. 12. Area under each peak is determined by the amount of protein in the spot. The plots are from digital data of pixel intensity taken along the cursor lines shown in Fig. 5. Upper plot shows the profiles of the two proteins extracted from follicles incubated in PS255, while the lower profiles are from follicles incubated 1 h in PSS400.
Fig. 7. Changes in the relative absorbancy (concentration) of seven endogenous soluble basic proteins caused by the osmotically induced reversal of the transbridge electrical gradient between nurse cells and their attached oocyte.
Following incubation in PSS400, the concentrations of all seven basic proteins assayed increased in oocytes (P=0.008). Basic proteins 4, 5 and 6 were barely visible in nurse cell samples from control follicles, and after incubation in PSS400 their concentrations had dimin-ished to unmeasurable levels. Thus we could not obtain reliable densitometric measurements for them, and they have not been included in averages for basic proteins from nurse cells.
Concerning the operation of a transbridge electrical gradient, there are some misconceptions which have entered the literature. Firstly, the gradient has been seen by some as a mechanism by which molecules and cyto-plasmic particles might be moved generally throughout the germ line cells of follicles. Yet there is absolutely no evidence that the reported electrical gradients in Dro-sophila or any other organism exist from one end of an ovarian follicle to the other. Instead, in every reported case the entire measured gradient was focused across intercellular bridges, specifically those between oocyte and nurse cells in the case of polytrophic ovaries in insects (Woodruff and Telfer 1973, 1980; Verachtert, 1988; Woodruff et al., 1988; Verachtert et al., 1989; Woodruff, 1989; Singleton and Woodruff, 1994) and marine polychaete worms (Emanuelsson and Anehus, 1985). In the telotrophic ovaries of insects, the electrical gradient is focused between groups of co-bridged nurse cells and the core of the tropharium (Telfer et al., 1981; Woodruff and Anderson, 1984; Munz and Dittman, 1987). Thus the gradient could have no part in the gen-eral movement of organelles, granules and other micro-scopically visible particles, nor of molecules, from dis-tant regions of nurse cells to locals in oocytes more than a few micrometers from an intercellular bridge.
It has been suggested that negatively charged enzymes such as those used in glycolysis would be eliminated from the nurse cell cytoplasm (and positively charged enzymes eliminated from the ooplasm) (Sun and Wyman, 1993). The data presented in this study show that nurse cells are not swept clear of acidic soluble pro-teins. This is because the gradient can serve only as a “leaky” gate mechanism, influencing the eventual con-centrations of charged molecules. Without an interven-ing membrane or membrane-like structure, at any tem-perature above absolute zero an electrical gradient, no matter how narrowly focused, could only oppose dif-fusion to a limited extent. The establised asymmetry must be in the form of a Gibbs–Donnan equilibrium, mathematically described by the Nernst equation. Thus a transbridge electrical gradient could serve to influence the concentration, but could not bring about the absolute absence of a charged molecule.
Nor is the isoelectric point (pI) of a protein the para-mount determinant of the degree to which it would be affected by such a gradient. Rather it is the charge den-sity on the molecule which is most important. Across an
intercellular bridge with an electrical gradient of 5 mV, a protein with a pI of pH 10, but which has an ionic charge of only 2+ would reach an equilibrium in which
it would be 1.5 times greater in concentration on the electronegative side of the bridge than on the electro-positive side (Woodruff, 1989). Conversely, a protein with a pI of only pH 8, but with 12 exposed positively charged sites, would also be positively charged in cyto-plasm. Having a charge density of 12, it would eventu-ally reach an equilibrium in which its concentration on the electronegative side of the bridge would be ten times higher than on the electropositive side. This is the most likely reason that some charged proteins showed a greater response then did others to changes in the trans-bridge gradient. For instance, concentrations of acidic proteins no. 9 and no. 10 each increased in nurse cells and declined in oocytes following osmotically induced reversal of the transbridge gradient. However, the changes exhibited by protein no. 9 in both cell types were about twice those which occurred to protein no. 10, behavior which would be expected if protein no. 9 has a greater charge density.
Many cytoplasmic particles and the molecules which attach to them have been shown to be moved about the nurse cells and the oocyte via cytoskeletal transport sys-tems. However, movement within the bridges seems to be regulated by other means (Theurkauf and Hazelrigg, 1998). While we would be surprised if distribution of bound molecules was also regulated in the same manner, data reported here strongly support the hypothesis that, within germ-line cells, the transbridge electrical gradient does influence the distribution of charged endogenous cytosolic proteins inDrosophila ovarian follicles.
Acknowledgements
The authors wish to thank Dr G. Cassotti and R. W. Woodruff for helpful suggestions in the perpetration of this manuscript. This work was supported in part by a grant from the National Science Foundation, no. IBN-9903994.
References
Bier, K., 1963a. Autoradiographische untersuchungen u¨ber die leistung des follikelepithels und der na¨hrzellen bei der dotterbildung und eiwiss synthese im fliegenovar. Arch Entwicklungsmech Org (Wilhem Roux’s) 154, 552–575.
Bier, K., 1963b. Synthese, interzellular transport, und abbau von ribonukleinsaure im ovar der stubenfliegeMusca domestica. Jour-nal of Cell Biology 16, 436–440.
Bohrmann, J., 1991. In vitro culture ofDrosophilaovarian follicles: the influence of different media on development, RNA synthesis, protein synthesis, and potassium uptake. Roux’s Archives of Devel-opmental Biology 199, 315–326.
as the principal mode of protein transport in vitellogenic ovarian follicles ofDrosophila. Development 101, 279–288.
Bohrmann, J., Huebner, E., Sander, K., Gutzeit, H.O., 1986. Intracellu-lar electrical potential measurements inDrosophilafollicles. Jour-nal of Cell Science 81, 207–221.
Cole, R.W., Woodruff, R.I., 1997. Charge dependent distribution of endogenous proteins within ovarian follicles ofActias luna. Journal of Insect Physiology 43, 275–287.
Cummings, M.R., Brown, N.M., King, R.C., 1969. The cytology of the vitellogenic stages of oogenesis inDrosophila melanogaster. I. General staging characteristics. Journal of Morphology 128, 427– 442.
Deely, D., 1989. Optimized reagents for protein analysis. Strategies in Molecular Biology 2, 44–45.
Ellis, G.W., 1978. Advances in visualization of mitosis in vivo. In: Dirksen, E., Prescott, D.M., Fox, C.E. (Eds.), Cell Reproduction. Academic Press, New York, pp. 465–476.
Emanuelsson, H., Anehus, S., 1985. Development in vitro of the female germ cells of the polychaeteOphryotrocha labronica. Jour-nal of Embryology and Experimental Morphology 85, 151–162. Fawcett, D.W., Ito, S., Slautterback, D., 1959. The occurrence of
inter-cellular bridges in groups of cells exhibiting syncytial differen-tiation. Journal of Biophysical and Biochemical Cytology 5, 453–463.
Hengge-Aronis, R., Lange, R., Henneberg, N., Fischer, D., 1993. Osmotic regulation of rpoS-dependent genes inEscherichia coli. Journal of Bacteriology 175, 259–265.
Miyazaki, S., Hagiwara, S., 1976. Electrical properties of the Droso-philaegg membrane. Developmental Biology 53, 91–100. Mizuno, T., Chou, M., Inouye, M., 1984. A unique mechanism
reg-ulating gene expression: translational inhibition by a complemen-tary RNA transcript (micRNA). Proceedings of the National Acad-emy of Science USA 81, 1966–1970.
Morrissey, J.H., 1981. Silver stain for proteins in polyacrylamide gels: a modified procedure with enhanced uniform sensitivity. Analytical Biochemistry 117, 307–310.
Muffler, A., Traulsen, D.D., Lange, R., Hengge-Aronis, R., 1996. Post-transcriptional osmotic regulation of the sigma(s) subunit of RNA polymerase in Escherichia coli. Journal of Bacteriology 178, 1607–1613.
Munz, A., Dittman, F., 1987. Voltage gradients and microtubules both involved in intercellular protein and mitochondria transport in the telotrophic ovariole ofDysdercus intermedius. Roux’s Archives of Developmental Biology 196, 391–396.
Nusslein-Volhard, C., Frohnhofer, H.G., Lehmann, R., 1987. Determi-nation of anteroposterior polarity in Drosophila. Science. 238, 1675-1681.
O’Farrell, P.Z., Goodman, H.M., O’Farrell, P.H., 1977. High resol-ution two-dimensional electrophoresis of basic as well as acidic proteins. Cell 12, 1133–1142.
Pollack, S.B., Telfer, W.H., 1969. RNA in cecropia moth ovaries: sites of synthesis, transport, and storage. Journal of Experimental Zoology 170, 1–23.
Prince, W.S., Villarejo, M.R., 1990. Osmotic control of proU transcrip-tion is mediated through direct actranscrip-tion of potassium glutamate on the transcription complex. Journal of Biological Chemistry 265, 17673–17679.
Singleton, K., Woodruff, R.I., 1994. The osmolarity of adult
Droso-phila hemolymph and its effect on oocyte-nurse cell electrical polarity. Developmental Biology 161, 154–167.
Sun, Y.A., Wyman, R.J., 1987. Lack of an oocyte to nurse cell voltage difference inDrosophila. Neuroscience 13, 1139.
Sun, Y.A., Wyman, R.J., 1993. Reevaluation of electrophoresis in the
Drosophilaegg chamber. Developmental Biology 155, 206–215. Telfer, W.H., 1965. The mechanism and control of yolk formation.
Annual Review of Entomology 10, 161–184.
Telfer, W.H., 1975. Development and physiology of the oocyte-nurse cell syncytium. Advances in Insect Physiology 11, 223–319. Telfer, W.H., Woodruff, R.I., Huebner, E., 1981. Electrical polarity
and cellular differentiation in meroistic ovaries. American Zoolo-gist 21, 675–686.
Theurkauf, W.E., Hazelrigg, T.L., 1998. In vivo analyses of cytoplas-mic transport and cytoskeletal organization duringDrosophila oog-enesis: characterization of a multi-step anterior localization path-way. Development 125, 3655–3666.
Tilney, L.G., Tilney, M.S., Guild, G., 1996. Formation of actin fila-ment bundles in the ring canals of developingDrosophilafollicles. Journal of Cell Biology 133, 61–74.
Verachtert, B. 1988. Electrical polarity in the ovarian follicles of Sar-cophaga bullata, Drosophila melanogaster, and Locusta migratoria. Doctoral Dissertation, Katholieke Universiteit Leuven, 207 p.
Verachtert, B., Amelinckx, M., De Loof, A., 1989. Potassium and chloride dependence of the membrane potential of vitellogenic fol-licles of Sarcophaga bullata (Diptera). Journal of Insect Physi-ology 35, 143–148.
Verachtert, B., De Loof, A., 1989. Intra- and extracellular electrical fields of vitellogenic poloytrophic insect follicles. Biological Bull-etin 176 (S), 91–95.
Woodruff, R.I., 1989. Charge-dependent molecular movement through intercellular bridges inDrosophilafollicles. Biological Bulletin 176 (S), 71–78.
Woodruff, R.I., Anderson, K.L., 1984. Nutritive cord connection and dye-coupling of the follicular epithelium to the growing oocytes in the telotrophic ovarioles in Oncopeltus fasciatus, the milkweed bug. Roux’s Archives of Developmental Biology 193, 158–163. Woodruff, R.I., Dittmann, F., Telfer, W.H., 1998. Ca2+current from
oocyte to nurse cells and suppression of uridine incorporation in the germinal vesicle ofHyalophora cecropia. Invertebrate Repro-duction and Development 34, 157–164.
Woodruff, R.I., Kulp, J.H., LaGaccia, E.D., 1988. Electrically mediated protein movement in Drosophila follicles. Roux’s Archives of Developmental Biology 197, 231–238.
Woodruff, R.I., Miller, A.L., Jaffe, L.F., 1991. Differences in free cal-cium concentration between oocytes and nurse cells revealed by corrected aequorin luminescence. Biological Bulletin 181, 349– 350.
Woodruff, R.I., Telfer, W.H., 1973. Polarized intercellular bridges in ovarian follicles of the cecropia moth. Journal of Cell Biology 58, 172–188.
Woodruff, R.I., Telfer, W.H., 1980. Electrophoresis of proteins in intercellular bridges. Nature (London) 286, 84–86.
