Drosophila melanogaster Scramblases modulate synaptic transmission
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《联合生物学》
1 Laboratory of Protein Dynamics and Signaling, National Cancer Institute Frederick, Frederick, MD 21702
2 Program in Gene Function and Expression, University of Massachusetts Medical School, Worcester, MA 01605
3 Centro de Estudios Científicos, Valdivia 511-0246, Chile
4 EM Facility/Image Analysis Laboratory, Science Applications International Corporation Frederick, Frederick, MD 21702
5 Universidad Austral de Chile, Valdivia 509-9200, Chile
Scramblases are a family of single-pass plasma membrane proteins, identified by their purported ability to scramble phospholipids across the two layers of plasma membrane isolated from platelets and red blood cells. However, their true in vivo role has yet to be elucidated. We report the generation and isolation of null mutants of two Scramblases identified in Drosophila melanogaster. We demonstrate that flies lacking either or both of these Scramblases are not compromised in vivo in processes requiring scrambling of phospholipids. Instead, we show that D. melanogaster lacking both Scramblases have more vesicles and display enhanced recruitment from a reserve pool of vesicles and increased neurotransmitter secretion at the larval neuromuscular synapses. These defects are corrected by the introduction of a genomic copy of the Scramb 1 gene. The lack of phenotypes related to failure of scrambling and the neurophysiological analysis lead us to propose that Scramblases play a modulatory role in the process of neurotransmission.
M.B. Edwards, R. Jorquera, and H. Silva contributed equally to this paper.
Abbreviations used in this paper: dsRNA, double-stranded RNA; ECP, exo/endo pool of SVs; EMS, ethyl methane sulfonate; GMR, glass multimer reporter; NMJ, neuromuscular junction; PS, phosphatidylserine; PTP, posttetanic potentiation; RP, reserve pool; S2 cells, Schneider cells; SV, synaptic vesicle; UAS, upstream activating sequence.
Introduction
Scramblases are a family of four proteins in humans and mice. They are conserved across species, and at least one member of this family is encoded in organisms ranging from yeast to human beings (Sims and Wiedmer, 2001). Human Scramblase 1 (PLSCR1) was initially identified as a protein thought to be involved in facilitation of transbilayer movement of phospholipids across the plasma membrane (Basse et al., 1996; Comfurius et al., 1996; Zhou et al., 1997). Many cell membranes harbor a Ca2+-dependent mechanism that can facilitate transbilayer movement of phospholipids between the two leaflets, leading to loss of membrane lipid asymmetry termed scrambling. Scrambling is observed in platelets, erythrocytes, and other cells after elevated intracellular Ca2+ and cell injury by complement and also in cells undergoing phagocytosis and apoptosis (Zwaal et al., 2005). Attempts to purify and clone proteins active in scrambling led to the identification of human PLSCR1. Scramblases are type 2 single-pass transmembrane proteins (Zhou et al., 1997). The four members identified in mice and humans have been named PLSCR1 through -4. All except PLSCR2 contain a proline-rich NH2-terminal domain with PXXP and PPXX motifs that can potentially bind to SH3 and WD40 domain–containing proteins. An intracellular EF hand, a Ca2+ binding motif, precedes the transmembrane segment. Mice lacking two of these Scramblases, 1 and 3, were recently generated. Cells derived from PLSCR1-null mice showed alterations in granulocyte production in response to growth factors and antiviral response to interferon (Zhou et al., 2002). PLSCR3-null mice and double mutants of PLSCR1 and -3 nulls showed adiposity, dyslipidemia, and insulin resistance (Wiedmer et al., 2004). Neither of the knockout mice shows defects in events associated with lipid scrambling. This suggests either that the other two members of the PLSCR family maintain Scramblase activity or that the presumed activity of this protein in scrambling is incorrect. Hence, the actual in vivo biological function of members of the Scramblase family has yet to be elucidated.
The Drosophila melanogaster genome encodes two ubiquitously expressed proteins with strong sequence homology to mice and human Scramblases. Using a reverse genetic approach, we have generated flies lacking each of the two Scramblases individually and flies lacking both Scramblases. Our studies using these mutants indicate that Scramblases do not play a deterministic role in scrambling of phospholipids that accompanies developmentally regulated apoptosis or in immune response involving phagocytosis of bacterially infected cells. Further analyses indicate that Scramblases play a modulatory role in the process of neurotransmission at the larval neuromuscular junction (NMJ).
Results
D. melanogaster Scramblases
The deduced sequences of the mammalian PLSCR family of proteins reveal them to be type 2 plasma membrane proteins with the transmembrane domain at the COOH terminus and a proline- and cysteine-rich acidic domain. The NH2-terminal segments contain PXXP and PPXY sequences that are potential SH3 and WW domain binding sites, respectively. The COOH-terminal ends of the proteins are conserved and contain a putative Ca2+ binding motif proximal to the transmembrane domain. There is considerable sequence variation at the NH2-terminal segment of Scramblases. The PLSCR2 member of the family lacks the NH2-terminal proline- and cysteine-rich sequence. D. melanogaster genome analysis identifies two genes with significant homology to mammalian Scramblases (Fig. 1 A; Drysdale et al., 2005). These are CG32056 and CG1893. Protein encoded by CG32056 Scramb 1 shows 37% identity with mouse Scramb 1 (PLSCR1) protein, whereas that of CG1893 or Scramb 2 protein shows 41% identity with Scramb 2 (PLSCR2). The Scramb 1 and 2 proteins of D. melanogaster are 56% identical and 69% similar to each other (Fig. 1 A). Although Scramb 1 includes the proline-rich region NH2-terminal domains with PXXP and PPXX motifs that bind SH3 and WD40 region, Scramb 2 protein lacks this stretch of amino acids and in this regard is closer to mammalian Scramb 2 protein. A third gene, CG9804, has been labeled as Scramblase in the annotated genome. However, this protein shows only 25% identity with either mammalian or the D. melanogaster Scramblases and seems to have branched out earlier during evolution. Moreover, unlike the other two D. melanogaster proteins, this protein has restricted expression, as we fail to see ubiquitous expression by Western analysis (unpublished data). CG10427 stated as a Scramblase in the Flybase was annotated as an uncertain gene and has been eliminated in release 3 of the genome annotation (this aberration resulted from annotation errors of CG32056 gene region).
To profile the tissue and developmental expression of Scramblases in D. melanogaster, we have generated polyclonal antibodies against both Scramb 1 and 2 proteins and in addition have obtained monoclonal antibodies against Scramb 1 protein. We have also succeeded in expressing a GFP-tagged version of Scramb 2 using the upstream activating sequence (UAS)–Gal4 system (Brand and Perrimon, 1993). Western blot analysis of wild-type D. melanogaster demonstrates that the two proteins are expressed ubiquitously and throughout development (Fig. 1 B). Lower levels are observed by Western analysis in early embryos before cellularization. The low abundance in early embryos is attributable to the fact that these are predominantly plasma membrane proteins (see below and Fig. S1, available at http://www.jcb.org/cgi/content/full/jcb.200506159/DC1). The D. melanogaster embryo begins as a single cell and a syncitium before undergoing cellularization, and the plasma membrane surface grows 30-fold during cellularization of the syncytial embryo. Immunofluorescence localization and analysis of transgenic flies (GFP–Scramb 2 fusion) localize the proteins predominantly to the plasma membrane (not depicted). We have colocalized Scramb 1 protein with HRP antigen at the plasma membrane in the larval neurons and the Scramb 2 protein with Notch that localizes to the plasma membrane at the apical margins of the cells of the larval eye-antennal anlagen (Fig. S1). Although Scramb 2 is homogeneously localized at the plasma membrane, Scramb 1 shows a nonhomogenous distribution along the plasma membrane, with areas of intense staining interspersed with areas of homogenously strong staining. The localization of the two proteins suggests a role for them in events involving the plasma membrane. Because the in vivo role of Scramblases in lipid scrambling and other cellular functions remains unresolved, we decided to generate mutant flies lacking the family of Scramblases and then carry out functional analyses.
Generation and isolation of Scramblase mutants
We have used a Western blot–based reverse genetics approach to obtain mutants for both Scramblase genes in D. melanogaster (Dolph et al., 1993; Acharya et al., 1998). The lack of predictable phenotypes defining the mutant complicates many genetic screens. Therefore, we used a previously documented Western blot–based screening strategy that monitors loss of protein expression (loss of Scramblase expression in our case) on immunoblots (Dolph et al., 1993; Acharya et al., 1998). In this protocol, male flies are randomly mutagenized, by feeding them the chemical mutagen, ethyl methane sulfonate (EMS). Mutagenized males are crossed to female balancer flies. Balanced progeny are crossed to flies with a large chromosomal deficiency that uncovers the genomic interval including the Scramb 1 gene. Single heads from flies trans-heterozygous for the deficiency and a mutagenized third chromosome are screened for the loss of Scramb 1 antigen by Western blots (Fig. 2 A). From a screen of 1,950 EMS-mutagenized third chromosomes, 1,900 lines that were viable in trans to Df(3L)AC1, rnroe-1pp were obtained. Western analysis of these lines identified a mutant line expressing no Scramb 1 protein (Fig. 2 A, bottom). The Scramb 1 gene was isolated from this mutant and subjected to sequence analysis. The scramb1-null mutant had a T-to-A transition in the first intron of the coding region. This leads to a splicing defect (confirmed by RT-PCR across intron 1) and truncation of the protein after 38 amino acids (+ 3 amino acids of the intron). Thus, the scramb1 mutant is either a null allele or a severely hypomorphic mutant.
Likewise, in the Scramb 2 screen, 2,100 lines, viable in trans over Df(3L)M21(kni[ri-1]p) were established and screened by Western analysis (Fig. 2 B, top). One of the lines showed no antigen reactivity against the Scramb 2 antibody (Fig. 2 B, bottom). Sequencing of the Scramb 2 gene from this line showed a T-to-A transition at the first exon–intron junction. In this instance, the splicing defect (confirmed by RT-PCR) leads to termination of the gene after 57 amino acids (an additional eight amino acids from read-through in the intron). Hence, the scramb2 mutant is either a null or severely hypomorphic mutant. Both mutants were backcrossed three times to w1118 (control) to outcross all background mutations.
The two mutants are null (or severely hypomorphic) for the respective normal Scramblase proteins. Double mutants of scramb1 and -2 were generated by meiotic recombination of the individual mutants. The double mutants do not express both proteins as confirmed by Western analysis and also by immunofluorescence at the larval NMJ (Fig. 2 and Fig. S2, available at http://www.jcb.org/cgi/content/full/jcb.200506159/DC1).Because the in vivo role of Scramblases in lipid scrambling has not been unequivocally established, we analyzed the mutants in events requiring scrambling of phospholipids, including the exposure of phosphatidylserine (PS) on cell surface in cells undergoing apoptosis and phagocytosis.
Scramblase mutants show no defects in developmentally regulated and ectopically induced apoptotic events in vivo
scramb1, scramb2, and the double mutants are homozy- gous viable. Programmed cell death is an important feature of D. melanogaster development and tissue homeostasis. During development, excess cells and tissues that are no longer useful are removed by apoptosis, and mutants that fail to undergo such normal apoptosis do not survive past embryogenesis (White et al., 1994). Because Scramblase mutants are viable and fertile and show no developmental defects, apoptosis during development is clearly not compromised in these mutant flies. As in mammals, apoptotic cell death in D. melanogaster is also characterized by PS exposure and clearance of cells by phagocytosis (van den Eijnde et al., 1998). In D. melanogaster, reaper, hid, and grim genes are necessary for induction of apoptosis. Ectopic overexpression of any of these genes in the fly eye using the eye-specific driver GMR (glass multimer reporter) causes normal photoreceptors to undergo apoptotic cell death, resulting in a severe eye ablation phenotype (Nordstrom et al., 1996). To determine whether Scramblase mutants can modulate this phenotype, we examined eyes from flies overexpressing reaper in each of the three mutant backgrounds. The eye sizes were not significantly different in the three mutant backgrounds compared with control flies (Fig. 3). This indicates that the dynamics of reaper-mediated apoptotic cell death is not altered in Scramblase mutant backgrounds. Because the eye sizes were not different between the control w1118 and mutant flies, PS-mediated cell death and phagocytic clearance of apoptotic cells were clearly unaffected in the mutant backgrounds. Thus, Scramblases are not required for either programmed or ectopically triggered apoptotic cell death.
Scramblase mutants can mount an effective response to infection with microorganisms
Immune defense mechanisms in D. melanogaster, like in other metazoan organisms, include physical barriers to infection, innate, and humoral responses (Lavine and Strand, 2002; Hoffmann, 2003; Brennan and Anderson, 2004; Leclerc and Reichhart, 2004). The immune hemocyte cells, for example, mediate cellular responses such as phagocytosis, encapsulation, and melanization, as well as produce humoral effector proteins. Phagocytic cells recognize a variety of signals on cells primed for phagocytosis (such as apoptotic and bacterially infected cells), including cell surface exposure of PS. We decided to evaluate whether the absence of Scramblases compromised any of these functions, resulting in an observable phenotype. We therefore infected wild-type and mutant cells with a mixture of gram positive and negative microorganisms as described in Materials and methods (Rodriguez et al., 1996; Bernal and Kimbrell, 2000). Immune competence is measured as viability of flies over time, as shown in Table I. The resistance to an inoculum of mixed microorganisms was comparable between the control w1118 and mutant backgrounds. Thus, the immune competence of Scramblase mutants is similar to wild-type flies and not compromised.
We conclude that Scramblases do not play a critical role in in vivo events that involve scrambling of phospholipids, such as exposure of PS to the outer leaflet and immunoreactive mechanisms for phagocytic clearance of bacterially infected cells. To demonstrate that PS exposure, a process requiring scrambling of phospholipids, was not compromised in backgrounds where Scramblase protein levels are perturbed, we conducted a series of experiments using D. melanogaster Schneider (S2) cells in which we either overexpressed the two Scramblases or depleted them using RNAi.
Increased expression or knock down of Scramblases in S2 cells does not affect PS exposure on cell surface
Exposure of cell surface PS during apoptosis has been used as a measure of the ability of cells to scramble phospholipids (Schlegel et al., 2000; Tepper et al., 2000; Wolfs et al., 2005). An early event in apoptosis is exposure of PS on the cell surface, which can be analyzed by binding of fluorescently labeled annexin V that has a high affinity for cell surface PS. The impermeant nucleic acid dye propidium iodide is used in conjunction to exclude cells whose plasma membrane integrity is compromised (necrotic or damaged). To visualize the process of scrambling (i.e., exposure of PS in cells with altered Scramblase expression), we used D. melanogaster S2 cells to knock down or overexpress Scramblase and follow PS exposure. Scramb 1 and 2 proteins were knocked down separately or together by RNAi treatment in S2 cells by transfecting with double-stranded RNA (dsRNA) as described in Materials and methods. Western blot analysis of dsRNA-treated cell lysates (+) shows significant depletion of Scramb 1 and 2 compared with untreated lysates (Fig. 4, A and B). S2 cells after dsRNA treatment were incubated in a Ca2+-rich medium containing fluorescently labeled annexin V and propidium iodide. PS exposure was measured as percentage of cells that stained positive for annexin V but negative for propidium iodide (Fig. 4 D, % scrambling). As seen in Fig. 4 D (–Act D), there is no difference between control S2 cells and Scramb 1, Scramb 2, and the double (Scramb 1, Scramb 2) knockdown cells. We then tested whether scrambling would be compromised in Scramblase knockdown cells undergoing apoptosis. It has previously been shown that D. melanogaster S2 cells undergo apoptosis and expose PS when treated with Actinomycin D (Zimmermann et al., 2002). Using similar conditions, apoptosis was induced in S2 cells by Actinomycin D treatment, and PS exposure was measured as before. There were no apparent differences in the ability of these cells to expose their cell surface PS (Fig. 4 D, +Act D) as compared with non-RNAi–treated cells undergoing apoptosis.
We then stably overexpressed Scramb 1 and 2 proteins under the control of the metallothionein promoter (Bunch et al., 1988). Fig. 4 C depicts Western blots showing dramatic inducible expression of both Scramblases. PS exposure was measured in these cells overexpressing Scramblases. There was no significant difference in the number of cells that were annexin V positive and propidium iodide negative between the control and overexpressors (Fig. 4 E, % scrambling). The baseline apoptosis was itself slightly higher, probably because of the addition of 0.5 mM copper sulfate to induce protein expression (Fig. 4 E, –Act D). Next, we induced apoptosis in S2 cells overexpressing Scramb 1 or 2 protein by Actinomycin D treatment and evaluated PS exposure by annexin V binding assays. There was no difference between the vector-transfected control and the overexpressing stable cells (Fig. 4 E, +Act D). Thus, neither knock down or overexpression of Scramblases resulted in altered exposure of PS in resting cells as well as in cells undergoing apoptosis.
The results described above lead us to believe that Scramblases do not play a determining role in scrambling of phospholipids in D. melanogaster. We then sought to evaluate the actual in vivo functions of this family of proteins. The Scramblase double-mutant flies appear visibly more active in a vial compared with either of the single mutants or the control w1118 flies. Hyperactivity has been associated with neurological alterations in D. melanogaster (Wang et al., 1997; Ghezzi et al., 2004). Because the double mutants were hyperactive and because Scramblases are ubiquitously expressed in the plasma membrane, the D. melanogaster NMJ was analyzed in our mutant for defects in synaptic structure and function. D. melanogaster larval NMJ is extremely active with intracellular and intercellular signaling involving a plethora of signal-transduction cascades active across plasma membranes at the junction (Keshishian et al., 1996; Rodesch and Broadie, 2000; Keshishian, 2002; Kidokoro et al., 2004; Kuromi and Kidokoro, 2005). We rationalized that the structure and function would be sensitive to loss of Scramblase proteins if they had an important role in events involving the plasma membrane.
Scramblase mutants show altered synaptic function and increased release of neurotransmitters
We verified the expression of Scramb 1 and 2 at the larval NMJs by immunofluorescence colocalization with synaptotagmin and HRP (Fig. 5). We also performed coimmunolocalization of Scramb 2 with disc large, cysteine string protein, syntaxin, and Fasciclin II (Fig. S4, available at http://www.jcb.org/cgi/content/full/jcb.200506159/DC1). Scramb 1 and 2 proteins are expressed in all segmental nerves and presynaptic and postsynaptic membranes of the larval NMJs (Figs. 5, S3, and S4). We then undertook ultrastructural examination and analysis of the active zones (Babcock et al., 2004; Rohrbough et al., 2004). The analysis revealed a significant increase in the number of synaptic vesicles (SVs) and more vesicles docked at the active one (Fig. 6 and Table II). The single mutants, on the other hand, did not show significant changes in the ultrastructure of NMJs, except that there was more than one docked vesicle at the active zone of most of the type 1b boutons examined (unpublished data).
We reasoned that an increased SV content in double-mutant synapse shown in EM studies should result in an increased loading of FM1-43 at nerve endings after massive vesicle mobilization. FM1-43 binds to membranes and remains trapped in SVs that undergo endocytosis. We analyzed the extent of FM1-43 loading in vesicles cycling through an exo/endo pool of SVs (ECP). Loading of ECP with FM1-43 was achieved by exposing boutons (5 min) to an external solution high in K+ in the presence of dye (Fig. 7 A; Delgado et al., 2000). Fluorescence brightness was analyzed 5 min after extensive perfusion with dye-free standard solution. Data analysis revealed that fluorescence brightness in double-mutant boutons was significantly above control (Fig. 7 A). After ECP loading, we subjected synapses to high-frequency stimulation of the nerve at 10 Hz in a standard external solution containing FM1-43, to also load vesicles cycling through a reserve pool (RP) of SVs. As shown in Fig. 7 B, this maneuver increases dye load, and fluorescence now distributes over the whole volume of synaptic boutons. Under these conditions also, fluorescence brightness in the double mutant was significantly above control (Fig. 7 B). Finally, to evaluate the extent of dye loading in SVs cycling through RP, synapses were exposed for a second round to a high K+ solution void of dye (3 min) to induce massive release of SVs in ECP. The fluorescence that remains after this second exposure to high K+ labels vesicles that are cycling through the RP (Delgado et al., 2000). Data analysis revealed that fluorescence brightness was significantly larger in double-mutant boutons (Fig. 7 C). Thus, our FM1-43–loading studies are consistent with EM data indicating an excess content of SVs in double-mutant presynaptic terminals. Importantly, as documented in Fig. 7, in all conditions, the extent of FM1-43 loading in the double mutant was rescued to control values by the introduction of a copy of the Scramb 1 gene.
We then proceeded to investigate ECP exocytosis rate by monitoring the decay of FM1-4 fluorescence brightness after dye loading of vesicles through ECP during low-frequency stimulation of the nerve at 0.5 Hz. Under this condition, transmitter release is maintained essentially by mobilization of ECP (Kuromi and Kidokoro, 2000). We found that the time course of decline of fluorescence brightness was similar in double-mutant and control boutons, indicating similar properties of ECP exocytosis (unpublished data). We also investigated the recruitment of SVs from an RP. The RP contains a majority of SVs and has been implicated in use-dependent increase in transmitter release and is mobilized during high-frequency activity at the synapse (Delgado et al., 2000; Kuromi and Kidokoro, 2000). 5 min after dye loading of SVs cycling through the RP, RP recruitment rates were assessed by monitoring the decline in fluorescence brightness during high-frequency stimulation at 10 Hz. We found that the rate of RP recruitment was markedly enhanced in double-mutant boutons, as indicated by a significantly faster rate of fluorescence brightness decline (2.4 ± 0.2 min [n = 10] vs. 8.5 ± 0.3 min [n = 10]; Fig. 7 D). This result prompted us to investigate whether RP recruitment at rest, in the absence of high-frequency stimulation of the nerve, was also enhanced in the double mutant. Therefore, we evaluated the extent of fluorescence brightness decline at rest, 5 min after SVs cycling through RP were loaded with dye. As shown in Fig. 7 E (top) at rest, the decline in fluorescence brightness was markedly enhanced in double-mutant boutons (fivefold). Moreover, the extent of fluorescence decline induced by a 5-min stimulation of the nerve at 0.5 Hz in the double mutant was also significantly enhanced relative to control (Fig. 7 E, bottom). These results reveal that RP recruitment in double-mutant synapses is abnormally enhanced. Fig. 7(D and E) also documents that RP recruitment in double-mutant larvae was rescued to control levels by the introduction of a copy of the Scramb 1 gene.
We next evaluated the properties of synaptic transmission by recording nerve-evoked postsynaptic currents from larval NMJs (Acharya et al., 1998; Delgado et al., 2000; Kidokoro et al., 2004). Single-mutant synapses displayed transmitter release and plasticity properties that did not differ significantly from control (unpublished data). In contrast, synaptic transmission in the double mutant differed in several aspects from the control. We first observed that the frequency of spontaneous postsynaptic currents was significantly enhanced in the double mutant (Table III). We also found that release probability, quantal content, and nerve-evoked postsynaptic currents were significantly larger in the double mutant (Fig. 8, A and C; and Table III). On the other hand, the amplitude of current and the charge transferred by the release of a single quantum were, within experimental error, the same in control and double mutant (Table III). Fig. 8 A shows plots of nerve-evoked postsynaptic currents as a function of external Ca2+ concentration in double logarithmic fashion in control, double mutant, and P{Scramb 1}; double mutant. Data analysis revealed a similar Ca2+ cooperativity coefficient for nerve-evoked responses. However, in double-mutant synapses, the curve was shifted toward lower Ca2+ concentrations. We also recorded postsynaptic currents evoked by exposing synapses to a hyperosmotic 0.5 M sucrose solution (Acharya et al., 1998). As documented in Fig. 8 B, neurotransmitter release evoked by application of a 20-s hyperosmotic shock in the vicinity of the synapse was dramatically increased in the double mutant. Importantly, as documented in Fig. 8 and Table III, the synaptic defects in the double mutant were rescued by the introduction of a copy of the Scramb 1 gene.
In further studies, we investigated plasticity properties of double-mutant synapses. Analysis of paired-pulse facilitation revealed that although the extent of facilitation of neurotransmitter release in the double mutant was diminished relative to the control, this difference was not significantly different from the control (Table III). We next evaluated tetanic facilitation and posttetanic potentiation (PTP) by monitoring nerve-evoked currents using a stimulation protocol designed to monitor tetanic facilitation and PTP. Postsynaptic responses were evoked first by stimulating the nerve at 0.5 Hz for 20 s. This was followed by a period of tetanic stimulation at 10 Hz for 50 s. After the tetanic stimulation, the regimen was switched back to 0.5 Hz to monitor PTP. We found that the extent of facilitation of transmitter release elicited by high-frequency stimulation of the nerve did not differ significantly from the control (Table III). On the other hand, we observed that PTP in the double mutant was abnormally prolonged. As shown in Fig. 8 C, in control synapse, nerve-evoked transmitter release decayed to values before stimulation (Fig. 8 C, bottom, discontinuous line) in 50 s after the cessation of high-frequency stimulation. In contrast, in the double mutant, transmitter release remained significantly potentiated after 100 s, indicating altered short-term synaptic memory. Introduction of a genomic copy of the Scramb 1 gene in the double mutant rescued PTP to normal. Also expression of a UAS–Scramb 2 gene driven by elav-GAL4 and neuronal promoter (elav-GAL4; UAS–Scramb 2; double mutant) rescued the content of ECP, RP, mobilization of RP pool of vesicles, postsynaptic current amplitude, quantal content, tetanic facilitation, and posttetanic facilitation to control levels (Fig. S5, available at http://www.jcb.org/cgi/content/full/jcb.200506159/DC1). In summary, our electrophysiological inquiries indicate that several aspects of synaptic function, including spontaneous, nerve-evoked release and PTP, are altered in the double mutant.
Despite changes in synaptic structure and physiology, overall synaptic function is not compromised in that the viability of the animal is not grossly affected in the double mutants. Thus, we believe that Scramblases have modulatory function in the secretory process of NMJs.
Discussion
Lipids are asymmetrically distributed between the two leaflets of the plasma membrane. Although the outer leaflet is enriched in choline phospholipids such as phosphatidylcholine and sphingomyelin, the inner leaflet contains anionic phospholipids such as PS and phosphatidylethanolamine. Although these lipids have a tendency to equilibrate along their concentration gradient, specific proteins that catalyze uni- or bidirectional transport of lipids from one leaflet to another maintain the asymmetry. One of the important players in this process is aminophospholipid translocase. Although its molecular identity has not been definitively identified, a P-type ATPase (aminophospholipid translocase) is thought to mediate this function (Wolfs et al., 2005). However, certain physiological and pathological conditions actively cause collapse of this asymmetry, resulting in scrambling of phospholipids. The most notable situation is in cells undergoing apoptosis wherein cell surface exposure of PS is one of the earliest detectable biochemical events. In fact, the annexin V binding assays for apoptotic cells is based on this exposure of PS. PS exposure has also been proposed as one of the signals that initiate the phagocytic process in cells targeted for phagocytosis. Lipid scrambling is thought to depend on one or more membrane proteins with lipid Scramblase activity. PLSCR1 was thus purified and identified as a putative Scramblase based on in vitro assays of transfer of lipids in bilayer. Subsequently, three other members of this family were discovered in humans and mice.
Mice lacking PLSCR1 and -3 have been generated. Phenotypic analysis of the mutants indicated that PLSCR1 mice showed a defective response of hematopoietic precursors to granulocyte colony–stimulating and stem cell factors. PLSCR3 mice show insulin resistance, glucose intolerance, and dyslipidemia. Both knockout mice show no defects in lipid scrambling, suggesting that the presumed activity of this protein is not correct or that other members of the PLSCR family maintain the Scramblase activity. D. melanogaster has two definitively identified members of this family, and we have generated mutants in both proteins. Analyses of null (or severely hypomorphic) Scramblase mutants demonstrate that these proteins do not play a critical role in scrambling of phospholipids; instead, they have a modulatory role in the secretory process.
We have demonstrated that D. melanogaster Scramblases show an overlapping and ubiquitous expression throughout development, and they localize to the plasma membrane in all the tissues examined. The ubiquitous distribution suggests their participation in general housekeeping function involving plasma membrane. We used a Western blot–based genetic screen to obtain null mutants for each of the two Scramblases. We also recombined the two null mutants to generate double mutants of Scramblases. The viability and lack of any developmental defects in the single and double mutants and our studies with reaper expression in mutant animal backgrounds indicate that they do not play a deterministic role in scrambling of phospholipids. This notion is further supported by our data in S2 cells, where neither overexpression nor RNAi-mediated knock down of these proteins has any effect on the ability of resting or apoptotic cells to expose PS.
Our studies indicate that loss of both Scramblases results in alteration in both the organization and function of the larval NMJ. The synaptic phenotype of the double mutant includes an increase in the number of vesicles at the synapse (including docked vesicles), a fivefold increase in RP recruitment at rest, a twofold increase in release probability, and an abnormal PTP. The fact that the recruitment of vesicles from RP at rest in the absence of nerve stimulation is dramatically enhanced in the double mutants might explain in part the abnormally elevated number of docked vesicles seen in EM analysis. The increased cycling of vesicles through the RP, as indicated by the almost twofold increase in uptake of FM1-43 (Fig. 7 C), probably associates with the twofold increase in release probability and evoked release seen in the double-mutant synapse (Fig. 8, A and C; and Table III). Although increased resting Ca2+ levels can contribute to increased probability of release, we do not believe they are contributing to the double-mutant phenotype. For example, although RP recruitment at rest in the double mutant is enhanced fivefold relative to w1118, the probability of release is only twofold above normal. Because the cooperative factor of Ca2+ for transmitter release is approximately Ca4, an increase in Ca2+ at rest in the double mutant would be expected to have a much larger effect on release probability than on RP recruitment. On the other hand, an enhanced recruitment of SVs from RP could contribute to the increased number of SVs available for release observed in the double-mutant presynaptic terminals. An abnormal handling of Ca2+ might underlie the enhanced probability of release found in double-mutant synapses. Indeed, our results indicate a shift to left on the Ca2+ dependence or release, without a change in Ca2+ cooperativity, suggesting that Ca2+ handling might be somehow abnormal in double-mutant synapses. However, paired-pulse facilitation is only slightly reduced in the double mutant synapse, arguing against a grossly abnormal handling of Ca2+. Also, the rates of ECP exocytosis are known to depend significantly on Ca2+ (Kuromi and Kidokoro, 2003). However, contrary to what one would expect if Ca2+ levels were abnormally elevated, we observed that the rates of ECP exocytosis were the same in the control and double-mutant boutons. Thus, our observations would in principle rule out the possibility that grossly elevated Ca2+ levels underlie the abnormal synaptic phenotype observed in double-mutant synapses. In D. melanogaster, RP recruitment and transmitter release are fostered by elevations in cAMP levels at synaptic boutons, and RP recruitment is antagonized by PKA inhibitors (Kuromi and Kidokoro, 2000). Importantly, in a dunce mutant, with abnormally elevated cAMP levels, RP size is dramatically reduced, probably because of the excess translocation of SVs from RP (Kuromi and Kidokoro, 2000). We found that in spite of enhanced RP recruitment, RP size remains vigorous in the double mutant, as documented in Fig. 7 C. Thus, unlike dunce, enhanced RP recruitment in the double mutant is not accompanied by a reduced RP size. We cannot rule out the possibility that cAMP signaling is altered in double mutant synapses. If so, such alteration should allow for a significant enhancement in RP recruitment without a reduction in RP size. In short, our data seem to rule out in principle a grossly abnormal handling of Ca2+ or a severely altered operation of the cAMP cascade in double-mutant synapses. It is important to note that the effects of cAMP on transmitter release are not limited to fostering RP recruitment. There is evidence that cAMP-gated channels at the presynaptic terminal also play an important role, independent of PKA, in enhancing transmitter release in D. melanogaster (Cheung et al., 2006). In D. melanogaster, RP dynamics has also been associated to the actin cytoskeleton and to the actin binding myosin ATPase complex (Kidokoro et al., 2004; Verstreken et al., 2005). Further work should allow us to better understand the role of Scramblases in regulating RP dynamics and transmitter release in D. melanogaster. The fact that D. melanogaster Scramblases exhibit putative protein binding domains means it is possible that they are able to interact with other proteins and signaling pathways to participate in regulating transmitter secretion and SV trafficking at the presynaptic terminal. We provide the first evidence that Scramblases can play an important role in regulating RP recruitment, transmitter release, and short-term synaptic memory in D. melanogaster.
We believe that Scramblases have a general role in modulating regulated secretory process. In fact, earlier work done in mammalian Scramblases seems to support this notion, although this conclusion was not drawn in those studies. In one of the earlier studies, Scramblases were found to interact with thrombospondin 1, a secreted protein (Aho and Uitto, 1998). Interferon , a secreted protein is a known up-regulator of Scramblase (Der et al., 1998; Zhou et al., 2000). Human Scramblase has been demonstrated to interact with human salivary leukocyte protease inhibitor, a secreted protein (Tseng and Tseng, 2000). Activation of Mast cells results in degranulation and secretion of a host of vasoactive amines, proteases, and proinflammatory cytokines. Cultured mast cells RBL-2H3 activated with FcRI resulted in up-regulation and phosphorylation of Scramblase protein (Pastorelli et al., 2001). Potentiation of the antiviral action of interferon by Scramblase has also been reported (Dong et al., 2004). PLSCR1 interacts with b-secretase (b-site amyloid precursor protein–cleaving enzyme), which is involved in secretion of the amyloid -peptide (Kametaka et al., 2003). PLSCR1 mutants have no defect in scrambling but have deficient granulopoiesis, and PLSCR3 mutant mice display adiposity and dyslipidemia. Finally, B lymphocytes from a patient with Scott syndrome (whose red blood cells show defective scrambling), despite being deficient in Scramblase activity, have normal levels of PLSCR1, and the nucleotide sequence of the corresponding mRNA is identical to controls (Zhou et al., 1998). All the aforementioned results and our data from analyzing scramb1, scramb2, and the double mutants reveal a role for Scramblases in modulating regulated exocytosis and not in the scrambling of phospholipids.
Materials and methods
Identification and cloning of D. melanogaster Scramblases
A BLAST (basic local alignment search tool) search of the D. melanogaster genome database using the mammalian Scramb 2 sequence revealed the existence of two definitive members of Scramblase family of genes. The two Scramblases are Scramblase 67D (Scramb 1) and 63B (Scramb 2). D. melanogaster genome annotation project has listed these genes as CG1893 and CG32056. CG32056 (Scramb 1) is localized to the 67D region on the left arm of third chromosome, and CG1893 gene (Scramb 2) is localized to the 63B region on the left arm of third chromosome. A BLAST search against the cDNA database revealed the existence of several cDNA clones from the D. melanogaster collection maintained at the time by the Berkeley Drosophila Genome Project and Research Genetics. All clones whose 5' or 3' ends showed homology to the mouse Scramblases were obtained from Research Genetics and sequenced. Full-length cDNA clones corresponding to GM13876 for Scramb 1 and GH10494 for Scramb 2 genes were then used for all subsequent manipulations. The Scramblase cDNAs with appropriate cloning restriction sites were amplified, cloned, and sequenced. For transgenic overexpression in D. melanogaster, the appropriate cDNAs were cloned into pUAST vector, and for S2 cell overexpression they were cloned into pRMHa-3 vector (a gift from L. Goldstein, University of California, San Diego, La Jolla, CA).
Fly stocks and husbandry
D. melanogaster stocks were cultured on standard maize meal agar and maintained at 25°C unless otherwise mentioned. The Df(3L)AC1 rnroe-1pp/TM3, Sb Df(3L)M21, kni[ri-1]p/In(3L)Rt33[L]f19, w–; P{w + mC = Actin}-Gal4 stocks were obtained from Bloomington Stock Center. Cyo, 2x GMR Rpr/Sco; MKRS/TM6B flies were obtained from K. White (Massachusetts General Hospital, Boston, MA).
Genetic screens and isolation of Scramblase mutants
w1118 males were aged for 3–4 d, starved overnight in empty vials, and subjected to chemical mutagenesis by feeding them on a diet of 25 μM EMS in 3% sucrose as previously described (Dolph et al., 1993; Acharya et al., 1998). These flies were then crossed en masse to virgin females with balanced third chromosomes. Single F1 males carrying a mutagenized chromosome over a third chromosome balancer were then crossed to virgin Df(3L)AC1, rnroe-1pp/TM3, Sb, flies in individual vials. The deficiency uncovers the 67D region. From each of these vials, a single progeny that carried the mutagenized chromosome over the deficiency was subjected to Western blot analysis for the loss of respective Scramblase antigen (Dolph et al., 1993; Acharya et al., 1998). Vials with lethal mutations in the interval uncovered by the deficiency were set aside to be used in transgenic rescue experiments, if necessary. Single fly heads were homogenized in SDS-PAGE buffer and loaded on gels. The separated proteins were transferred to nitrocellulose and incubated with affinity-purified rabbit polyclonal antibodies raised against the Scramb 1 protein. A single null mutant was isolated in the screen. To outcross all incidental and irrelevant mutations, the null mutant was backcrossed three times to w1118 flies and selected by Western analysis. A similar approach was used to isolate the scramb2 mutant. However, Df(3L)M21, kni[ri-1]p/In(3L)Rt33[L]f19, a deficiency that uncovers the 63B region, was used instead. Again, the null mutant was backcrossed three times to w1118 controls to outcross all other mutations and selected by Western analysis.
Transgenic flies
The Scramb 2 cDNA was amplified in frame with open reading frame of GFP gene and ligated to generate GFP–Scramb 2 fusion. The gene was then cloned into pUAST vector and injected into embryos to generate transgenic flies expressing GFP–Scramb 2 fusion. pUAST–Scramb 2–transgenic flies were generated. These lines were subsequently crossed to actin-Gal4 lines to drive its expression ubiquitously. A similar approach was attempted for Scramb 1 gene, but no protein was detected in these flies either by Western analysis or by immunofluorescence. We presumed that the protein was misfolded and degraded. An 11.8-Kb Sal1–Kpn1 fragment of genomic DNA spanning the Scramb 1 gene was cloned into pUAST, and transgenic lines were obtained. One of these mapped to the second chromosome and was used in the transgenic rescue experiments. The transgenic expression of Scramb 1 protein was confirmed by Western analysis in the double-mutant background.
Antibodies
Rabbit polyclonal and monoclonal antibodies were raised against bacterially expressed, affinity-purified MBP–Scramb 1 protein in which the Scramb 1 protein was further cleaved off from MBP by factor Xa cleavage and purified by electro elution from polyacrylamide gels. Additionally, a COOH-terminal peptide antibody was raised against Scramb 1–derived peptide CFFEKAGNQETDRPGML and affinity purified. Rabbit polyclonal antibodies were raised against Scramb 2 protein expressed using a pET3a expression system and purified by SDS-PAGE and electroelution. For Western analysis, affinity-purified anti-peptide Scramb 1 antibody was used at 1:500 dilution and anti–Scramb 2 antibody was used at 1:2,000–1:4,000. IPP antibodies were a gift from C. Zuker (University of San Diego, San Diego, CA). Anti–Fas II (1:500), anti-synaptotagmin (1:200), anti-cysteine string protein (1:25), anti-synataxin (1:200), and anti-DLG (1:200) monoclonal antibodies were obtained from Iowa Hybridoma Bank. Affinity-purified rabbit anti-synaptotagmin (1:1,000) antibody was a gift from M. Ramaswami and S. Sanyal (University of Arizona, Phoenix, AZ). Anti-HRP (1:500) and secondary antibodies (1:500) were obtained from Jackson ImmunoResearch Laboratories. Anti-CDase (1:3,000) antibodies have been described previously (Acharya et al., 2003, 2004; Rohrbough et al., 2004). The Notch antibody was a gift from M. Fortini (National Cancer Institute Frederick, Frederick, MD).
Immunofluorescence
Embryos and imaginal discs were collected and prepared for immunofluorescent staining as described previously (Patel, 1994). Tissues were incubated overnight with the antibodies (1:100–1:500 for anti–Scramb 1 monoclonal and anti–Scramb 2 rabbit polyclonal) at 4°C. These were costained with Alexa 568 or 488–conjugated goat anti–rabbit or goat anti–mouse IgG for 2 h at room temperature, washed, and mounted on Vectashield. For GFP visualization, the wing or eye imaginal discs were isolated from wandering third instar larvae, fixed in 4% formaldehyde, washed, and stained with propidium iodide for nuclear staining. For confocal microscopy, laser-scanning confocal microscopes (models 410 and 510; Carl Zeiss MicroImaging, Inc.) were used in most experiments, using a 63x oil objective, and pictures were obtained at resolutions ranging from 512 x 512 to 2048 x 2048. The pictures were pseudocolored in Photoshop (Adobe) for use in figures.
Electron microscopic examination
For electron microscopic examinations, wandering third instar larvae grown at 25°C were dissected and fixed using 2% glutaraldehyde and 4% formaldehyde in sodium cacodylate buffer. The tissues were postfixed in 1% osmium tetroxide and dehydrated in graded ethanol and propylene oxide. Specimens were embedded in epoxy resin (Embed-812) and thin sectioned. Transverse sections (90 nm) were stained with uranyl acetate and lead citrate and examined.
Genetic crosses
Mutant Scramblase flies (individual and double mutants) were crossed into GMR-Rpr background to evaluate the effects of these mutations in reaper-induced apoptosis. UAS–Scramb 2 GFP flies were crossed to actin-Gal4 to drive expression ubiquitously.
Bacterial infections
Bacterial infection experiments were done as described previously (Rodriguez et al., 1996; Bernal and Kimbrell, 2000). Escherichia coli, Enterobacter cloacae B12, Staphylococcus epidermidis, and Micrococcus reoseus were gifts from D. Kimbrell and R. Schoenfeld (University of California, Davis, Davis, CA). They were individually grown and pelleted. They were subsequently resuspended in LB collectively and pelleted again. The microorganism mixture was then injected into the thorax of individual flies using tungsten needles. A group of 10 individual flies were maintained in each vial, and the vials were replaced with a fresh vial every day. A total of 100 flies were infected for individual samples, and the experiment was repeated three times.
Scrambling index
Scramb 1 and 2 were cloned into pRMHA3 vector, and stable lines were established. The proteins were induced with 0.5 mM copper sulfate, and cells were harvested after 48 h of induction for Western analysis. A 900-bp fragment of Scramb 1 and a 600-bp fragment of Scramb 2 were PCR amplified with E. coli T7 polymerase site as described previously (Clemens et al., 2000; Worby et al., 2001) with minor modifications. RNA was synthesized using the T7 RNA polymerase kit (QIAGEN). The dsRNA were annealed by heating to 75°C for 20 min and slow cooling to room temperature. 1 μg of dsRNA was transfected to S2 cells in a 3-cm dish by Ca2+ phosphate transfection and media changed after 24 h. dsRNA-treated cells were evaluated for protein levels by Western analysis at 24, 48, and 72 h after transfection. For scrambling (apoptotic) index, stable cells induced for protein production for 48 h or cells treated with dsRNA for 48 h were treated with Actinomycin D. They were then incubated in annexin V binding buffer (10 mM Hepes, pH 7.4, 150 mM NaCl, and 2.5 mM CaCl2) with annexin V–Alexa 488 propidium iodide. Stained cells were visualized using a confocal microscope (model 510). Approximately 300 cells were counted in three different fields for each experimental sample. Cells that were annexin V positive and propidium iodide negative were then used to calculate the percentage of cells that exposed PS on the surface (scrambling index).
Electrophysiology
Postsynaptic responses evoked by stimulation of the nerve were recorded from segments A2 or A3 of ventral longitudinal muscle 6 or 7 in third instar larvae using a two-electrode voltage clamp as reported previously in a standard working solution containing 128 mM NaCl, 2 mM KCl, 4 mM MgCl2, 0.2 mM CaCl2, 5 mM Hepes, and 36 mM sucrose, pH 7.3. Osmotic release of neurotransmitter was achieved by application of a hyperosmotic standard solution containing 500 mM sucrose and 50 μM CaCl2 (Acharya et al., 1998; Delgado et al., 2000). The currents were recorded at –80 mV holding potential, and current amplitudes and integrals were analyzed using pClamp software (Axon Instruments, Inc.). Nerves were cut close to the ventral ganglia and sucked into the stimulating pipette. Evoked currents were elicited by stimulation of the nerve at the frequencies indicated using a programmable stimulator (Master-8; MPI). Data acquisition and analysis were done using pClamp software. The quantal content of nerve-evoked responses was estimated by dividing the current integral of individual nerve-evoked currents by the integral of the current evoked by the release of an individual quantum, as described previously (Acharya et al., 1998; Delgado et al., 2000).
Loading and unloading of FM1-43
Loading of ECP with FM1-43.
Dye loading of vesicles cycling through ECP was achieved by exposing synaptic boutons for 5 min to high K+ external containing 10 μM FM1-43 (Invitrogen) as described by Kuromi and Kidokoro (2000).
Loading of RP with FM1-43.
After dye loading of SVs cycling through ECP, nerve was stimulated at 10 Hz for 5 min in standard working solution containing FM1-43. FM1-43 was then removed by extensive rinsing with dye-free standard solution. Then, synaptic boutons were exposed for a second time to high K+ for 3 min to release SVs cycling through ECP, after which the external solution was extensively rinsed with standard working solution. The remaining FM1-43 fluorescence corresponds to SVs cycling through the RP (Delgado et al., 2000).
FM1-43 unloading.
In all cases, FM1-43 unloading was monitored in a standard working solution at rest, in the absence of nerve stimulation or by stimulating the nerve at low (0.5 Hz) or high frequency (10 Hz) using a programmable stimulator.
Imaging
Synaptic boutons were imaged using an epi-fluorescence microscope (BX50WI; Olympus) with a 60x objective. Samples were excited for 200–300 ms via a controlled shutter system (Uniblitz; Vincent Associates). Images were captured with a 12-bit cooled charge-coupled device camera (SensiCam; PCO Imaging). Images were acquired and analyzed using Workbench 2.2 software (Axon Imaging). Results are from at least three synaptic boutons from a minimum of six different larvae per experimental condition. Fluorescence analysis was performed after correcting for bleaching and background.
Online supplemental material
Acknowledgments
We thank Drs. Charles Zuker, Kristin White, Vivian Budnik, Mark Fortini, Mani Ramaswami, Subhabrata Sanyal, and Deborah Kimbrell for flies reagents and flies; Drs. Shyam Sharan, Mark Fortini, and Ira Daar for critical reading of the manuscript; and Jason de la Cruz for technical assistance.
This work was supported by funding from the intramural division of the National Cancer Institute. Work in the laboratory of P. Labarca is supported by FONDECYT and the Howard Hughes Medical Institute. Centro de Estudios Científicos is Millennium Institute funded in part by grants from Fundacion Andes and the Tinker Foundation. P. Labarca is a Howard Hughes Medical Institute international scholar. R.A. Jorquera was the recipient of a doctoral fellowship from Mejoramiento de la Calidad y la Equidad de la Educacion Superior.
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2 Program in Gene Function and Expression, University of Massachusetts Medical School, Worcester, MA 01605
3 Centro de Estudios Científicos, Valdivia 511-0246, Chile
4 EM Facility/Image Analysis Laboratory, Science Applications International Corporation Frederick, Frederick, MD 21702
5 Universidad Austral de Chile, Valdivia 509-9200, Chile
Scramblases are a family of single-pass plasma membrane proteins, identified by their purported ability to scramble phospholipids across the two layers of plasma membrane isolated from platelets and red blood cells. However, their true in vivo role has yet to be elucidated. We report the generation and isolation of null mutants of two Scramblases identified in Drosophila melanogaster. We demonstrate that flies lacking either or both of these Scramblases are not compromised in vivo in processes requiring scrambling of phospholipids. Instead, we show that D. melanogaster lacking both Scramblases have more vesicles and display enhanced recruitment from a reserve pool of vesicles and increased neurotransmitter secretion at the larval neuromuscular synapses. These defects are corrected by the introduction of a genomic copy of the Scramb 1 gene. The lack of phenotypes related to failure of scrambling and the neurophysiological analysis lead us to propose that Scramblases play a modulatory role in the process of neurotransmission.
M.B. Edwards, R. Jorquera, and H. Silva contributed equally to this paper.
Abbreviations used in this paper: dsRNA, double-stranded RNA; ECP, exo/endo pool of SVs; EMS, ethyl methane sulfonate; GMR, glass multimer reporter; NMJ, neuromuscular junction; PS, phosphatidylserine; PTP, posttetanic potentiation; RP, reserve pool; S2 cells, Schneider cells; SV, synaptic vesicle; UAS, upstream activating sequence.
Introduction
Scramblases are a family of four proteins in humans and mice. They are conserved across species, and at least one member of this family is encoded in organisms ranging from yeast to human beings (Sims and Wiedmer, 2001). Human Scramblase 1 (PLSCR1) was initially identified as a protein thought to be involved in facilitation of transbilayer movement of phospholipids across the plasma membrane (Basse et al., 1996; Comfurius et al., 1996; Zhou et al., 1997). Many cell membranes harbor a Ca2+-dependent mechanism that can facilitate transbilayer movement of phospholipids between the two leaflets, leading to loss of membrane lipid asymmetry termed scrambling. Scrambling is observed in platelets, erythrocytes, and other cells after elevated intracellular Ca2+ and cell injury by complement and also in cells undergoing phagocytosis and apoptosis (Zwaal et al., 2005). Attempts to purify and clone proteins active in scrambling led to the identification of human PLSCR1. Scramblases are type 2 single-pass transmembrane proteins (Zhou et al., 1997). The four members identified in mice and humans have been named PLSCR1 through -4. All except PLSCR2 contain a proline-rich NH2-terminal domain with PXXP and PPXX motifs that can potentially bind to SH3 and WD40 domain–containing proteins. An intracellular EF hand, a Ca2+ binding motif, precedes the transmembrane segment. Mice lacking two of these Scramblases, 1 and 3, were recently generated. Cells derived from PLSCR1-null mice showed alterations in granulocyte production in response to growth factors and antiviral response to interferon (Zhou et al., 2002). PLSCR3-null mice and double mutants of PLSCR1 and -3 nulls showed adiposity, dyslipidemia, and insulin resistance (Wiedmer et al., 2004). Neither of the knockout mice shows defects in events associated with lipid scrambling. This suggests either that the other two members of the PLSCR family maintain Scramblase activity or that the presumed activity of this protein in scrambling is incorrect. Hence, the actual in vivo biological function of members of the Scramblase family has yet to be elucidated.
The Drosophila melanogaster genome encodes two ubiquitously expressed proteins with strong sequence homology to mice and human Scramblases. Using a reverse genetic approach, we have generated flies lacking each of the two Scramblases individually and flies lacking both Scramblases. Our studies using these mutants indicate that Scramblases do not play a deterministic role in scrambling of phospholipids that accompanies developmentally regulated apoptosis or in immune response involving phagocytosis of bacterially infected cells. Further analyses indicate that Scramblases play a modulatory role in the process of neurotransmission at the larval neuromuscular junction (NMJ).
Results
D. melanogaster Scramblases
The deduced sequences of the mammalian PLSCR family of proteins reveal them to be type 2 plasma membrane proteins with the transmembrane domain at the COOH terminus and a proline- and cysteine-rich acidic domain. The NH2-terminal segments contain PXXP and PPXY sequences that are potential SH3 and WW domain binding sites, respectively. The COOH-terminal ends of the proteins are conserved and contain a putative Ca2+ binding motif proximal to the transmembrane domain. There is considerable sequence variation at the NH2-terminal segment of Scramblases. The PLSCR2 member of the family lacks the NH2-terminal proline- and cysteine-rich sequence. D. melanogaster genome analysis identifies two genes with significant homology to mammalian Scramblases (Fig. 1 A; Drysdale et al., 2005). These are CG32056 and CG1893. Protein encoded by CG32056 Scramb 1 shows 37% identity with mouse Scramb 1 (PLSCR1) protein, whereas that of CG1893 or Scramb 2 protein shows 41% identity with Scramb 2 (PLSCR2). The Scramb 1 and 2 proteins of D. melanogaster are 56% identical and 69% similar to each other (Fig. 1 A). Although Scramb 1 includes the proline-rich region NH2-terminal domains with PXXP and PPXX motifs that bind SH3 and WD40 region, Scramb 2 protein lacks this stretch of amino acids and in this regard is closer to mammalian Scramb 2 protein. A third gene, CG9804, has been labeled as Scramblase in the annotated genome. However, this protein shows only 25% identity with either mammalian or the D. melanogaster Scramblases and seems to have branched out earlier during evolution. Moreover, unlike the other two D. melanogaster proteins, this protein has restricted expression, as we fail to see ubiquitous expression by Western analysis (unpublished data). CG10427 stated as a Scramblase in the Flybase was annotated as an uncertain gene and has been eliminated in release 3 of the genome annotation (this aberration resulted from annotation errors of CG32056 gene region).
To profile the tissue and developmental expression of Scramblases in D. melanogaster, we have generated polyclonal antibodies against both Scramb 1 and 2 proteins and in addition have obtained monoclonal antibodies against Scramb 1 protein. We have also succeeded in expressing a GFP-tagged version of Scramb 2 using the upstream activating sequence (UAS)–Gal4 system (Brand and Perrimon, 1993). Western blot analysis of wild-type D. melanogaster demonstrates that the two proteins are expressed ubiquitously and throughout development (Fig. 1 B). Lower levels are observed by Western analysis in early embryos before cellularization. The low abundance in early embryos is attributable to the fact that these are predominantly plasma membrane proteins (see below and Fig. S1, available at http://www.jcb.org/cgi/content/full/jcb.200506159/DC1). The D. melanogaster embryo begins as a single cell and a syncitium before undergoing cellularization, and the plasma membrane surface grows 30-fold during cellularization of the syncytial embryo. Immunofluorescence localization and analysis of transgenic flies (GFP–Scramb 2 fusion) localize the proteins predominantly to the plasma membrane (not depicted). We have colocalized Scramb 1 protein with HRP antigen at the plasma membrane in the larval neurons and the Scramb 2 protein with Notch that localizes to the plasma membrane at the apical margins of the cells of the larval eye-antennal anlagen (Fig. S1). Although Scramb 2 is homogeneously localized at the plasma membrane, Scramb 1 shows a nonhomogenous distribution along the plasma membrane, with areas of intense staining interspersed with areas of homogenously strong staining. The localization of the two proteins suggests a role for them in events involving the plasma membrane. Because the in vivo role of Scramblases in lipid scrambling and other cellular functions remains unresolved, we decided to generate mutant flies lacking the family of Scramblases and then carry out functional analyses.
Generation and isolation of Scramblase mutants
We have used a Western blot–based reverse genetics approach to obtain mutants for both Scramblase genes in D. melanogaster (Dolph et al., 1993; Acharya et al., 1998). The lack of predictable phenotypes defining the mutant complicates many genetic screens. Therefore, we used a previously documented Western blot–based screening strategy that monitors loss of protein expression (loss of Scramblase expression in our case) on immunoblots (Dolph et al., 1993; Acharya et al., 1998). In this protocol, male flies are randomly mutagenized, by feeding them the chemical mutagen, ethyl methane sulfonate (EMS). Mutagenized males are crossed to female balancer flies. Balanced progeny are crossed to flies with a large chromosomal deficiency that uncovers the genomic interval including the Scramb 1 gene. Single heads from flies trans-heterozygous for the deficiency and a mutagenized third chromosome are screened for the loss of Scramb 1 antigen by Western blots (Fig. 2 A). From a screen of 1,950 EMS-mutagenized third chromosomes, 1,900 lines that were viable in trans to Df(3L)AC1, rnroe-1pp were obtained. Western analysis of these lines identified a mutant line expressing no Scramb 1 protein (Fig. 2 A, bottom). The Scramb 1 gene was isolated from this mutant and subjected to sequence analysis. The scramb1-null mutant had a T-to-A transition in the first intron of the coding region. This leads to a splicing defect (confirmed by RT-PCR across intron 1) and truncation of the protein after 38 amino acids (+ 3 amino acids of the intron). Thus, the scramb1 mutant is either a null allele or a severely hypomorphic mutant.
Likewise, in the Scramb 2 screen, 2,100 lines, viable in trans over Df(3L)M21(kni[ri-1]p) were established and screened by Western analysis (Fig. 2 B, top). One of the lines showed no antigen reactivity against the Scramb 2 antibody (Fig. 2 B, bottom). Sequencing of the Scramb 2 gene from this line showed a T-to-A transition at the first exon–intron junction. In this instance, the splicing defect (confirmed by RT-PCR) leads to termination of the gene after 57 amino acids (an additional eight amino acids from read-through in the intron). Hence, the scramb2 mutant is either a null or severely hypomorphic mutant. Both mutants were backcrossed three times to w1118 (control) to outcross all background mutations.
The two mutants are null (or severely hypomorphic) for the respective normal Scramblase proteins. Double mutants of scramb1 and -2 were generated by meiotic recombination of the individual mutants. The double mutants do not express both proteins as confirmed by Western analysis and also by immunofluorescence at the larval NMJ (Fig. 2 and Fig. S2, available at http://www.jcb.org/cgi/content/full/jcb.200506159/DC1).Because the in vivo role of Scramblases in lipid scrambling has not been unequivocally established, we analyzed the mutants in events requiring scrambling of phospholipids, including the exposure of phosphatidylserine (PS) on cell surface in cells undergoing apoptosis and phagocytosis.
Scramblase mutants show no defects in developmentally regulated and ectopically induced apoptotic events in vivo
scramb1, scramb2, and the double mutants are homozy- gous viable. Programmed cell death is an important feature of D. melanogaster development and tissue homeostasis. During development, excess cells and tissues that are no longer useful are removed by apoptosis, and mutants that fail to undergo such normal apoptosis do not survive past embryogenesis (White et al., 1994). Because Scramblase mutants are viable and fertile and show no developmental defects, apoptosis during development is clearly not compromised in these mutant flies. As in mammals, apoptotic cell death in D. melanogaster is also characterized by PS exposure and clearance of cells by phagocytosis (van den Eijnde et al., 1998). In D. melanogaster, reaper, hid, and grim genes are necessary for induction of apoptosis. Ectopic overexpression of any of these genes in the fly eye using the eye-specific driver GMR (glass multimer reporter) causes normal photoreceptors to undergo apoptotic cell death, resulting in a severe eye ablation phenotype (Nordstrom et al., 1996). To determine whether Scramblase mutants can modulate this phenotype, we examined eyes from flies overexpressing reaper in each of the three mutant backgrounds. The eye sizes were not significantly different in the three mutant backgrounds compared with control flies (Fig. 3). This indicates that the dynamics of reaper-mediated apoptotic cell death is not altered in Scramblase mutant backgrounds. Because the eye sizes were not different between the control w1118 and mutant flies, PS-mediated cell death and phagocytic clearance of apoptotic cells were clearly unaffected in the mutant backgrounds. Thus, Scramblases are not required for either programmed or ectopically triggered apoptotic cell death.
Scramblase mutants can mount an effective response to infection with microorganisms
Immune defense mechanisms in D. melanogaster, like in other metazoan organisms, include physical barriers to infection, innate, and humoral responses (Lavine and Strand, 2002; Hoffmann, 2003; Brennan and Anderson, 2004; Leclerc and Reichhart, 2004). The immune hemocyte cells, for example, mediate cellular responses such as phagocytosis, encapsulation, and melanization, as well as produce humoral effector proteins. Phagocytic cells recognize a variety of signals on cells primed for phagocytosis (such as apoptotic and bacterially infected cells), including cell surface exposure of PS. We decided to evaluate whether the absence of Scramblases compromised any of these functions, resulting in an observable phenotype. We therefore infected wild-type and mutant cells with a mixture of gram positive and negative microorganisms as described in Materials and methods (Rodriguez et al., 1996; Bernal and Kimbrell, 2000). Immune competence is measured as viability of flies over time, as shown in Table I. The resistance to an inoculum of mixed microorganisms was comparable between the control w1118 and mutant backgrounds. Thus, the immune competence of Scramblase mutants is similar to wild-type flies and not compromised.
We conclude that Scramblases do not play a critical role in in vivo events that involve scrambling of phospholipids, such as exposure of PS to the outer leaflet and immunoreactive mechanisms for phagocytic clearance of bacterially infected cells. To demonstrate that PS exposure, a process requiring scrambling of phospholipids, was not compromised in backgrounds where Scramblase protein levels are perturbed, we conducted a series of experiments using D. melanogaster Schneider (S2) cells in which we either overexpressed the two Scramblases or depleted them using RNAi.
Increased expression or knock down of Scramblases in S2 cells does not affect PS exposure on cell surface
Exposure of cell surface PS during apoptosis has been used as a measure of the ability of cells to scramble phospholipids (Schlegel et al., 2000; Tepper et al., 2000; Wolfs et al., 2005). An early event in apoptosis is exposure of PS on the cell surface, which can be analyzed by binding of fluorescently labeled annexin V that has a high affinity for cell surface PS. The impermeant nucleic acid dye propidium iodide is used in conjunction to exclude cells whose plasma membrane integrity is compromised (necrotic or damaged). To visualize the process of scrambling (i.e., exposure of PS in cells with altered Scramblase expression), we used D. melanogaster S2 cells to knock down or overexpress Scramblase and follow PS exposure. Scramb 1 and 2 proteins were knocked down separately or together by RNAi treatment in S2 cells by transfecting with double-stranded RNA (dsRNA) as described in Materials and methods. Western blot analysis of dsRNA-treated cell lysates (+) shows significant depletion of Scramb 1 and 2 compared with untreated lysates (Fig. 4, A and B). S2 cells after dsRNA treatment were incubated in a Ca2+-rich medium containing fluorescently labeled annexin V and propidium iodide. PS exposure was measured as percentage of cells that stained positive for annexin V but negative for propidium iodide (Fig. 4 D, % scrambling). As seen in Fig. 4 D (–Act D), there is no difference between control S2 cells and Scramb 1, Scramb 2, and the double (Scramb 1, Scramb 2) knockdown cells. We then tested whether scrambling would be compromised in Scramblase knockdown cells undergoing apoptosis. It has previously been shown that D. melanogaster S2 cells undergo apoptosis and expose PS when treated with Actinomycin D (Zimmermann et al., 2002). Using similar conditions, apoptosis was induced in S2 cells by Actinomycin D treatment, and PS exposure was measured as before. There were no apparent differences in the ability of these cells to expose their cell surface PS (Fig. 4 D, +Act D) as compared with non-RNAi–treated cells undergoing apoptosis.
We then stably overexpressed Scramb 1 and 2 proteins under the control of the metallothionein promoter (Bunch et al., 1988). Fig. 4 C depicts Western blots showing dramatic inducible expression of both Scramblases. PS exposure was measured in these cells overexpressing Scramblases. There was no significant difference in the number of cells that were annexin V positive and propidium iodide negative between the control and overexpressors (Fig. 4 E, % scrambling). The baseline apoptosis was itself slightly higher, probably because of the addition of 0.5 mM copper sulfate to induce protein expression (Fig. 4 E, –Act D). Next, we induced apoptosis in S2 cells overexpressing Scramb 1 or 2 protein by Actinomycin D treatment and evaluated PS exposure by annexin V binding assays. There was no difference between the vector-transfected control and the overexpressing stable cells (Fig. 4 E, +Act D). Thus, neither knock down or overexpression of Scramblases resulted in altered exposure of PS in resting cells as well as in cells undergoing apoptosis.
The results described above lead us to believe that Scramblases do not play a determining role in scrambling of phospholipids in D. melanogaster. We then sought to evaluate the actual in vivo functions of this family of proteins. The Scramblase double-mutant flies appear visibly more active in a vial compared with either of the single mutants or the control w1118 flies. Hyperactivity has been associated with neurological alterations in D. melanogaster (Wang et al., 1997; Ghezzi et al., 2004). Because the double mutants were hyperactive and because Scramblases are ubiquitously expressed in the plasma membrane, the D. melanogaster NMJ was analyzed in our mutant for defects in synaptic structure and function. D. melanogaster larval NMJ is extremely active with intracellular and intercellular signaling involving a plethora of signal-transduction cascades active across plasma membranes at the junction (Keshishian et al., 1996; Rodesch and Broadie, 2000; Keshishian, 2002; Kidokoro et al., 2004; Kuromi and Kidokoro, 2005). We rationalized that the structure and function would be sensitive to loss of Scramblase proteins if they had an important role in events involving the plasma membrane.
Scramblase mutants show altered synaptic function and increased release of neurotransmitters
We verified the expression of Scramb 1 and 2 at the larval NMJs by immunofluorescence colocalization with synaptotagmin and HRP (Fig. 5). We also performed coimmunolocalization of Scramb 2 with disc large, cysteine string protein, syntaxin, and Fasciclin II (Fig. S4, available at http://www.jcb.org/cgi/content/full/jcb.200506159/DC1). Scramb 1 and 2 proteins are expressed in all segmental nerves and presynaptic and postsynaptic membranes of the larval NMJs (Figs. 5, S3, and S4). We then undertook ultrastructural examination and analysis of the active zones (Babcock et al., 2004; Rohrbough et al., 2004). The analysis revealed a significant increase in the number of synaptic vesicles (SVs) and more vesicles docked at the active one (Fig. 6 and Table II). The single mutants, on the other hand, did not show significant changes in the ultrastructure of NMJs, except that there was more than one docked vesicle at the active zone of most of the type 1b boutons examined (unpublished data).
We reasoned that an increased SV content in double-mutant synapse shown in EM studies should result in an increased loading of FM1-43 at nerve endings after massive vesicle mobilization. FM1-43 binds to membranes and remains trapped in SVs that undergo endocytosis. We analyzed the extent of FM1-43 loading in vesicles cycling through an exo/endo pool of SVs (ECP). Loading of ECP with FM1-43 was achieved by exposing boutons (5 min) to an external solution high in K+ in the presence of dye (Fig. 7 A; Delgado et al., 2000). Fluorescence brightness was analyzed 5 min after extensive perfusion with dye-free standard solution. Data analysis revealed that fluorescence brightness in double-mutant boutons was significantly above control (Fig. 7 A). After ECP loading, we subjected synapses to high-frequency stimulation of the nerve at 10 Hz in a standard external solution containing FM1-43, to also load vesicles cycling through a reserve pool (RP) of SVs. As shown in Fig. 7 B, this maneuver increases dye load, and fluorescence now distributes over the whole volume of synaptic boutons. Under these conditions also, fluorescence brightness in the double mutant was significantly above control (Fig. 7 B). Finally, to evaluate the extent of dye loading in SVs cycling through RP, synapses were exposed for a second round to a high K+ solution void of dye (3 min) to induce massive release of SVs in ECP. The fluorescence that remains after this second exposure to high K+ labels vesicles that are cycling through the RP (Delgado et al., 2000). Data analysis revealed that fluorescence brightness was significantly larger in double-mutant boutons (Fig. 7 C). Thus, our FM1-43–loading studies are consistent with EM data indicating an excess content of SVs in double-mutant presynaptic terminals. Importantly, as documented in Fig. 7, in all conditions, the extent of FM1-43 loading in the double mutant was rescued to control values by the introduction of a copy of the Scramb 1 gene.
We then proceeded to investigate ECP exocytosis rate by monitoring the decay of FM1-4 fluorescence brightness after dye loading of vesicles through ECP during low-frequency stimulation of the nerve at 0.5 Hz. Under this condition, transmitter release is maintained essentially by mobilization of ECP (Kuromi and Kidokoro, 2000). We found that the time course of decline of fluorescence brightness was similar in double-mutant and control boutons, indicating similar properties of ECP exocytosis (unpublished data). We also investigated the recruitment of SVs from an RP. The RP contains a majority of SVs and has been implicated in use-dependent increase in transmitter release and is mobilized during high-frequency activity at the synapse (Delgado et al., 2000; Kuromi and Kidokoro, 2000). 5 min after dye loading of SVs cycling through the RP, RP recruitment rates were assessed by monitoring the decline in fluorescence brightness during high-frequency stimulation at 10 Hz. We found that the rate of RP recruitment was markedly enhanced in double-mutant boutons, as indicated by a significantly faster rate of fluorescence brightness decline (2.4 ± 0.2 min [n = 10] vs. 8.5 ± 0.3 min [n = 10]; Fig. 7 D). This result prompted us to investigate whether RP recruitment at rest, in the absence of high-frequency stimulation of the nerve, was also enhanced in the double mutant. Therefore, we evaluated the extent of fluorescence brightness decline at rest, 5 min after SVs cycling through RP were loaded with dye. As shown in Fig. 7 E (top) at rest, the decline in fluorescence brightness was markedly enhanced in double-mutant boutons (fivefold). Moreover, the extent of fluorescence decline induced by a 5-min stimulation of the nerve at 0.5 Hz in the double mutant was also significantly enhanced relative to control (Fig. 7 E, bottom). These results reveal that RP recruitment in double-mutant synapses is abnormally enhanced. Fig. 7(D and E) also documents that RP recruitment in double-mutant larvae was rescued to control levels by the introduction of a copy of the Scramb 1 gene.
We next evaluated the properties of synaptic transmission by recording nerve-evoked postsynaptic currents from larval NMJs (Acharya et al., 1998; Delgado et al., 2000; Kidokoro et al., 2004). Single-mutant synapses displayed transmitter release and plasticity properties that did not differ significantly from control (unpublished data). In contrast, synaptic transmission in the double mutant differed in several aspects from the control. We first observed that the frequency of spontaneous postsynaptic currents was significantly enhanced in the double mutant (Table III). We also found that release probability, quantal content, and nerve-evoked postsynaptic currents were significantly larger in the double mutant (Fig. 8, A and C; and Table III). On the other hand, the amplitude of current and the charge transferred by the release of a single quantum were, within experimental error, the same in control and double mutant (Table III). Fig. 8 A shows plots of nerve-evoked postsynaptic currents as a function of external Ca2+ concentration in double logarithmic fashion in control, double mutant, and P{Scramb 1}; double mutant. Data analysis revealed a similar Ca2+ cooperativity coefficient for nerve-evoked responses. However, in double-mutant synapses, the curve was shifted toward lower Ca2+ concentrations. We also recorded postsynaptic currents evoked by exposing synapses to a hyperosmotic 0.5 M sucrose solution (Acharya et al., 1998). As documented in Fig. 8 B, neurotransmitter release evoked by application of a 20-s hyperosmotic shock in the vicinity of the synapse was dramatically increased in the double mutant. Importantly, as documented in Fig. 8 and Table III, the synaptic defects in the double mutant were rescued by the introduction of a copy of the Scramb 1 gene.
In further studies, we investigated plasticity properties of double-mutant synapses. Analysis of paired-pulse facilitation revealed that although the extent of facilitation of neurotransmitter release in the double mutant was diminished relative to the control, this difference was not significantly different from the control (Table III). We next evaluated tetanic facilitation and posttetanic potentiation (PTP) by monitoring nerve-evoked currents using a stimulation protocol designed to monitor tetanic facilitation and PTP. Postsynaptic responses were evoked first by stimulating the nerve at 0.5 Hz for 20 s. This was followed by a period of tetanic stimulation at 10 Hz for 50 s. After the tetanic stimulation, the regimen was switched back to 0.5 Hz to monitor PTP. We found that the extent of facilitation of transmitter release elicited by high-frequency stimulation of the nerve did not differ significantly from the control (Table III). On the other hand, we observed that PTP in the double mutant was abnormally prolonged. As shown in Fig. 8 C, in control synapse, nerve-evoked transmitter release decayed to values before stimulation (Fig. 8 C, bottom, discontinuous line) in 50 s after the cessation of high-frequency stimulation. In contrast, in the double mutant, transmitter release remained significantly potentiated after 100 s, indicating altered short-term synaptic memory. Introduction of a genomic copy of the Scramb 1 gene in the double mutant rescued PTP to normal. Also expression of a UAS–Scramb 2 gene driven by elav-GAL4 and neuronal promoter (elav-GAL4; UAS–Scramb 2; double mutant) rescued the content of ECP, RP, mobilization of RP pool of vesicles, postsynaptic current amplitude, quantal content, tetanic facilitation, and posttetanic facilitation to control levels (Fig. S5, available at http://www.jcb.org/cgi/content/full/jcb.200506159/DC1). In summary, our electrophysiological inquiries indicate that several aspects of synaptic function, including spontaneous, nerve-evoked release and PTP, are altered in the double mutant.
Despite changes in synaptic structure and physiology, overall synaptic function is not compromised in that the viability of the animal is not grossly affected in the double mutants. Thus, we believe that Scramblases have modulatory function in the secretory process of NMJs.
Discussion
Lipids are asymmetrically distributed between the two leaflets of the plasma membrane. Although the outer leaflet is enriched in choline phospholipids such as phosphatidylcholine and sphingomyelin, the inner leaflet contains anionic phospholipids such as PS and phosphatidylethanolamine. Although these lipids have a tendency to equilibrate along their concentration gradient, specific proteins that catalyze uni- or bidirectional transport of lipids from one leaflet to another maintain the asymmetry. One of the important players in this process is aminophospholipid translocase. Although its molecular identity has not been definitively identified, a P-type ATPase (aminophospholipid translocase) is thought to mediate this function (Wolfs et al., 2005). However, certain physiological and pathological conditions actively cause collapse of this asymmetry, resulting in scrambling of phospholipids. The most notable situation is in cells undergoing apoptosis wherein cell surface exposure of PS is one of the earliest detectable biochemical events. In fact, the annexin V binding assays for apoptotic cells is based on this exposure of PS. PS exposure has also been proposed as one of the signals that initiate the phagocytic process in cells targeted for phagocytosis. Lipid scrambling is thought to depend on one or more membrane proteins with lipid Scramblase activity. PLSCR1 was thus purified and identified as a putative Scramblase based on in vitro assays of transfer of lipids in bilayer. Subsequently, three other members of this family were discovered in humans and mice.
Mice lacking PLSCR1 and -3 have been generated. Phenotypic analysis of the mutants indicated that PLSCR1 mice showed a defective response of hematopoietic precursors to granulocyte colony–stimulating and stem cell factors. PLSCR3 mice show insulin resistance, glucose intolerance, and dyslipidemia. Both knockout mice show no defects in lipid scrambling, suggesting that the presumed activity of this protein is not correct or that other members of the PLSCR family maintain the Scramblase activity. D. melanogaster has two definitively identified members of this family, and we have generated mutants in both proteins. Analyses of null (or severely hypomorphic) Scramblase mutants demonstrate that these proteins do not play a critical role in scrambling of phospholipids; instead, they have a modulatory role in the secretory process.
We have demonstrated that D. melanogaster Scramblases show an overlapping and ubiquitous expression throughout development, and they localize to the plasma membrane in all the tissues examined. The ubiquitous distribution suggests their participation in general housekeeping function involving plasma membrane. We used a Western blot–based genetic screen to obtain null mutants for each of the two Scramblases. We also recombined the two null mutants to generate double mutants of Scramblases. The viability and lack of any developmental defects in the single and double mutants and our studies with reaper expression in mutant animal backgrounds indicate that they do not play a deterministic role in scrambling of phospholipids. This notion is further supported by our data in S2 cells, where neither overexpression nor RNAi-mediated knock down of these proteins has any effect on the ability of resting or apoptotic cells to expose PS.
Our studies indicate that loss of both Scramblases results in alteration in both the organization and function of the larval NMJ. The synaptic phenotype of the double mutant includes an increase in the number of vesicles at the synapse (including docked vesicles), a fivefold increase in RP recruitment at rest, a twofold increase in release probability, and an abnormal PTP. The fact that the recruitment of vesicles from RP at rest in the absence of nerve stimulation is dramatically enhanced in the double mutants might explain in part the abnormally elevated number of docked vesicles seen in EM analysis. The increased cycling of vesicles through the RP, as indicated by the almost twofold increase in uptake of FM1-43 (Fig. 7 C), probably associates with the twofold increase in release probability and evoked release seen in the double-mutant synapse (Fig. 8, A and C; and Table III). Although increased resting Ca2+ levels can contribute to increased probability of release, we do not believe they are contributing to the double-mutant phenotype. For example, although RP recruitment at rest in the double mutant is enhanced fivefold relative to w1118, the probability of release is only twofold above normal. Because the cooperative factor of Ca2+ for transmitter release is approximately Ca4, an increase in Ca2+ at rest in the double mutant would be expected to have a much larger effect on release probability than on RP recruitment. On the other hand, an enhanced recruitment of SVs from RP could contribute to the increased number of SVs available for release observed in the double-mutant presynaptic terminals. An abnormal handling of Ca2+ might underlie the enhanced probability of release found in double-mutant synapses. Indeed, our results indicate a shift to left on the Ca2+ dependence or release, without a change in Ca2+ cooperativity, suggesting that Ca2+ handling might be somehow abnormal in double-mutant synapses. However, paired-pulse facilitation is only slightly reduced in the double mutant synapse, arguing against a grossly abnormal handling of Ca2+. Also, the rates of ECP exocytosis are known to depend significantly on Ca2+ (Kuromi and Kidokoro, 2003). However, contrary to what one would expect if Ca2+ levels were abnormally elevated, we observed that the rates of ECP exocytosis were the same in the control and double-mutant boutons. Thus, our observations would in principle rule out the possibility that grossly elevated Ca2+ levels underlie the abnormal synaptic phenotype observed in double-mutant synapses. In D. melanogaster, RP recruitment and transmitter release are fostered by elevations in cAMP levels at synaptic boutons, and RP recruitment is antagonized by PKA inhibitors (Kuromi and Kidokoro, 2000). Importantly, in a dunce mutant, with abnormally elevated cAMP levels, RP size is dramatically reduced, probably because of the excess translocation of SVs from RP (Kuromi and Kidokoro, 2000). We found that in spite of enhanced RP recruitment, RP size remains vigorous in the double mutant, as documented in Fig. 7 C. Thus, unlike dunce, enhanced RP recruitment in the double mutant is not accompanied by a reduced RP size. We cannot rule out the possibility that cAMP signaling is altered in double mutant synapses. If so, such alteration should allow for a significant enhancement in RP recruitment without a reduction in RP size. In short, our data seem to rule out in principle a grossly abnormal handling of Ca2+ or a severely altered operation of the cAMP cascade in double-mutant synapses. It is important to note that the effects of cAMP on transmitter release are not limited to fostering RP recruitment. There is evidence that cAMP-gated channels at the presynaptic terminal also play an important role, independent of PKA, in enhancing transmitter release in D. melanogaster (Cheung et al., 2006). In D. melanogaster, RP dynamics has also been associated to the actin cytoskeleton and to the actin binding myosin ATPase complex (Kidokoro et al., 2004; Verstreken et al., 2005). Further work should allow us to better understand the role of Scramblases in regulating RP dynamics and transmitter release in D. melanogaster. The fact that D. melanogaster Scramblases exhibit putative protein binding domains means it is possible that they are able to interact with other proteins and signaling pathways to participate in regulating transmitter secretion and SV trafficking at the presynaptic terminal. We provide the first evidence that Scramblases can play an important role in regulating RP recruitment, transmitter release, and short-term synaptic memory in D. melanogaster.
We believe that Scramblases have a general role in modulating regulated secretory process. In fact, earlier work done in mammalian Scramblases seems to support this notion, although this conclusion was not drawn in those studies. In one of the earlier studies, Scramblases were found to interact with thrombospondin 1, a secreted protein (Aho and Uitto, 1998). Interferon , a secreted protein is a known up-regulator of Scramblase (Der et al., 1998; Zhou et al., 2000). Human Scramblase has been demonstrated to interact with human salivary leukocyte protease inhibitor, a secreted protein (Tseng and Tseng, 2000). Activation of Mast cells results in degranulation and secretion of a host of vasoactive amines, proteases, and proinflammatory cytokines. Cultured mast cells RBL-2H3 activated with FcRI resulted in up-regulation and phosphorylation of Scramblase protein (Pastorelli et al., 2001). Potentiation of the antiviral action of interferon by Scramblase has also been reported (Dong et al., 2004). PLSCR1 interacts with b-secretase (b-site amyloid precursor protein–cleaving enzyme), which is involved in secretion of the amyloid -peptide (Kametaka et al., 2003). PLSCR1 mutants have no defect in scrambling but have deficient granulopoiesis, and PLSCR3 mutant mice display adiposity and dyslipidemia. Finally, B lymphocytes from a patient with Scott syndrome (whose red blood cells show defective scrambling), despite being deficient in Scramblase activity, have normal levels of PLSCR1, and the nucleotide sequence of the corresponding mRNA is identical to controls (Zhou et al., 1998). All the aforementioned results and our data from analyzing scramb1, scramb2, and the double mutants reveal a role for Scramblases in modulating regulated exocytosis and not in the scrambling of phospholipids.
Materials and methods
Identification and cloning of D. melanogaster Scramblases
A BLAST (basic local alignment search tool) search of the D. melanogaster genome database using the mammalian Scramb 2 sequence revealed the existence of two definitive members of Scramblase family of genes. The two Scramblases are Scramblase 67D (Scramb 1) and 63B (Scramb 2). D. melanogaster genome annotation project has listed these genes as CG1893 and CG32056. CG32056 (Scramb 1) is localized to the 67D region on the left arm of third chromosome, and CG1893 gene (Scramb 2) is localized to the 63B region on the left arm of third chromosome. A BLAST search against the cDNA database revealed the existence of several cDNA clones from the D. melanogaster collection maintained at the time by the Berkeley Drosophila Genome Project and Research Genetics. All clones whose 5' or 3' ends showed homology to the mouse Scramblases were obtained from Research Genetics and sequenced. Full-length cDNA clones corresponding to GM13876 for Scramb 1 and GH10494 for Scramb 2 genes were then used for all subsequent manipulations. The Scramblase cDNAs with appropriate cloning restriction sites were amplified, cloned, and sequenced. For transgenic overexpression in D. melanogaster, the appropriate cDNAs were cloned into pUAST vector, and for S2 cell overexpression they were cloned into pRMHa-3 vector (a gift from L. Goldstein, University of California, San Diego, La Jolla, CA).
Fly stocks and husbandry
D. melanogaster stocks were cultured on standard maize meal agar and maintained at 25°C unless otherwise mentioned. The Df(3L)AC1 rnroe-1pp/TM3, Sb Df(3L)M21, kni[ri-1]p/In(3L)Rt33[L]f19, w–; P{w + mC = Actin}-Gal4 stocks were obtained from Bloomington Stock Center. Cyo, 2x GMR Rpr/Sco; MKRS/TM6B flies were obtained from K. White (Massachusetts General Hospital, Boston, MA).
Genetic screens and isolation of Scramblase mutants
w1118 males were aged for 3–4 d, starved overnight in empty vials, and subjected to chemical mutagenesis by feeding them on a diet of 25 μM EMS in 3% sucrose as previously described (Dolph et al., 1993; Acharya et al., 1998). These flies were then crossed en masse to virgin females with balanced third chromosomes. Single F1 males carrying a mutagenized chromosome over a third chromosome balancer were then crossed to virgin Df(3L)AC1, rnroe-1pp/TM3, Sb, flies in individual vials. The deficiency uncovers the 67D region. From each of these vials, a single progeny that carried the mutagenized chromosome over the deficiency was subjected to Western blot analysis for the loss of respective Scramblase antigen (Dolph et al., 1993; Acharya et al., 1998). Vials with lethal mutations in the interval uncovered by the deficiency were set aside to be used in transgenic rescue experiments, if necessary. Single fly heads were homogenized in SDS-PAGE buffer and loaded on gels. The separated proteins were transferred to nitrocellulose and incubated with affinity-purified rabbit polyclonal antibodies raised against the Scramb 1 protein. A single null mutant was isolated in the screen. To outcross all incidental and irrelevant mutations, the null mutant was backcrossed three times to w1118 flies and selected by Western analysis. A similar approach was used to isolate the scramb2 mutant. However, Df(3L)M21, kni[ri-1]p/In(3L)Rt33[L]f19, a deficiency that uncovers the 63B region, was used instead. Again, the null mutant was backcrossed three times to w1118 controls to outcross all other mutations and selected by Western analysis.
Transgenic flies
The Scramb 2 cDNA was amplified in frame with open reading frame of GFP gene and ligated to generate GFP–Scramb 2 fusion. The gene was then cloned into pUAST vector and injected into embryos to generate transgenic flies expressing GFP–Scramb 2 fusion. pUAST–Scramb 2–transgenic flies were generated. These lines were subsequently crossed to actin-Gal4 lines to drive its expression ubiquitously. A similar approach was attempted for Scramb 1 gene, but no protein was detected in these flies either by Western analysis or by immunofluorescence. We presumed that the protein was misfolded and degraded. An 11.8-Kb Sal1–Kpn1 fragment of genomic DNA spanning the Scramb 1 gene was cloned into pUAST, and transgenic lines were obtained. One of these mapped to the second chromosome and was used in the transgenic rescue experiments. The transgenic expression of Scramb 1 protein was confirmed by Western analysis in the double-mutant background.
Antibodies
Rabbit polyclonal and monoclonal antibodies were raised against bacterially expressed, affinity-purified MBP–Scramb 1 protein in which the Scramb 1 protein was further cleaved off from MBP by factor Xa cleavage and purified by electro elution from polyacrylamide gels. Additionally, a COOH-terminal peptide antibody was raised against Scramb 1–derived peptide CFFEKAGNQETDRPGML and affinity purified. Rabbit polyclonal antibodies were raised against Scramb 2 protein expressed using a pET3a expression system and purified by SDS-PAGE and electroelution. For Western analysis, affinity-purified anti-peptide Scramb 1 antibody was used at 1:500 dilution and anti–Scramb 2 antibody was used at 1:2,000–1:4,000. IPP antibodies were a gift from C. Zuker (University of San Diego, San Diego, CA). Anti–Fas II (1:500), anti-synaptotagmin (1:200), anti-cysteine string protein (1:25), anti-synataxin (1:200), and anti-DLG (1:200) monoclonal antibodies were obtained from Iowa Hybridoma Bank. Affinity-purified rabbit anti-synaptotagmin (1:1,000) antibody was a gift from M. Ramaswami and S. Sanyal (University of Arizona, Phoenix, AZ). Anti-HRP (1:500) and secondary antibodies (1:500) were obtained from Jackson ImmunoResearch Laboratories. Anti-CDase (1:3,000) antibodies have been described previously (Acharya et al., 2003, 2004; Rohrbough et al., 2004). The Notch antibody was a gift from M. Fortini (National Cancer Institute Frederick, Frederick, MD).
Immunofluorescence
Embryos and imaginal discs were collected and prepared for immunofluorescent staining as described previously (Patel, 1994). Tissues were incubated overnight with the antibodies (1:100–1:500 for anti–Scramb 1 monoclonal and anti–Scramb 2 rabbit polyclonal) at 4°C. These were costained with Alexa 568 or 488–conjugated goat anti–rabbit or goat anti–mouse IgG for 2 h at room temperature, washed, and mounted on Vectashield. For GFP visualization, the wing or eye imaginal discs were isolated from wandering third instar larvae, fixed in 4% formaldehyde, washed, and stained with propidium iodide for nuclear staining. For confocal microscopy, laser-scanning confocal microscopes (models 410 and 510; Carl Zeiss MicroImaging, Inc.) were used in most experiments, using a 63x oil objective, and pictures were obtained at resolutions ranging from 512 x 512 to 2048 x 2048. The pictures were pseudocolored in Photoshop (Adobe) for use in figures.
Electron microscopic examination
For electron microscopic examinations, wandering third instar larvae grown at 25°C were dissected and fixed using 2% glutaraldehyde and 4% formaldehyde in sodium cacodylate buffer. The tissues were postfixed in 1% osmium tetroxide and dehydrated in graded ethanol and propylene oxide. Specimens were embedded in epoxy resin (Embed-812) and thin sectioned. Transverse sections (90 nm) were stained with uranyl acetate and lead citrate and examined.
Genetic crosses
Mutant Scramblase flies (individual and double mutants) were crossed into GMR-Rpr background to evaluate the effects of these mutations in reaper-induced apoptosis. UAS–Scramb 2 GFP flies were crossed to actin-Gal4 to drive expression ubiquitously.
Bacterial infections
Bacterial infection experiments were done as described previously (Rodriguez et al., 1996; Bernal and Kimbrell, 2000). Escherichia coli, Enterobacter cloacae B12, Staphylococcus epidermidis, and Micrococcus reoseus were gifts from D. Kimbrell and R. Schoenfeld (University of California, Davis, Davis, CA). They were individually grown and pelleted. They were subsequently resuspended in LB collectively and pelleted again. The microorganism mixture was then injected into the thorax of individual flies using tungsten needles. A group of 10 individual flies were maintained in each vial, and the vials were replaced with a fresh vial every day. A total of 100 flies were infected for individual samples, and the experiment was repeated three times.
Scrambling index
Scramb 1 and 2 were cloned into pRMHA3 vector, and stable lines were established. The proteins were induced with 0.5 mM copper sulfate, and cells were harvested after 48 h of induction for Western analysis. A 900-bp fragment of Scramb 1 and a 600-bp fragment of Scramb 2 were PCR amplified with E. coli T7 polymerase site as described previously (Clemens et al., 2000; Worby et al., 2001) with minor modifications. RNA was synthesized using the T7 RNA polymerase kit (QIAGEN). The dsRNA were annealed by heating to 75°C for 20 min and slow cooling to room temperature. 1 μg of dsRNA was transfected to S2 cells in a 3-cm dish by Ca2+ phosphate transfection and media changed after 24 h. dsRNA-treated cells were evaluated for protein levels by Western analysis at 24, 48, and 72 h after transfection. For scrambling (apoptotic) index, stable cells induced for protein production for 48 h or cells treated with dsRNA for 48 h were treated with Actinomycin D. They were then incubated in annexin V binding buffer (10 mM Hepes, pH 7.4, 150 mM NaCl, and 2.5 mM CaCl2) with annexin V–Alexa 488 propidium iodide. Stained cells were visualized using a confocal microscope (model 510). Approximately 300 cells were counted in three different fields for each experimental sample. Cells that were annexin V positive and propidium iodide negative were then used to calculate the percentage of cells that exposed PS on the surface (scrambling index).
Electrophysiology
Postsynaptic responses evoked by stimulation of the nerve were recorded from segments A2 or A3 of ventral longitudinal muscle 6 or 7 in third instar larvae using a two-electrode voltage clamp as reported previously in a standard working solution containing 128 mM NaCl, 2 mM KCl, 4 mM MgCl2, 0.2 mM CaCl2, 5 mM Hepes, and 36 mM sucrose, pH 7.3. Osmotic release of neurotransmitter was achieved by application of a hyperosmotic standard solution containing 500 mM sucrose and 50 μM CaCl2 (Acharya et al., 1998; Delgado et al., 2000). The currents were recorded at –80 mV holding potential, and current amplitudes and integrals were analyzed using pClamp software (Axon Instruments, Inc.). Nerves were cut close to the ventral ganglia and sucked into the stimulating pipette. Evoked currents were elicited by stimulation of the nerve at the frequencies indicated using a programmable stimulator (Master-8; MPI). Data acquisition and analysis were done using pClamp software. The quantal content of nerve-evoked responses was estimated by dividing the current integral of individual nerve-evoked currents by the integral of the current evoked by the release of an individual quantum, as described previously (Acharya et al., 1998; Delgado et al., 2000).
Loading and unloading of FM1-43
Loading of ECP with FM1-43.
Dye loading of vesicles cycling through ECP was achieved by exposing synaptic boutons for 5 min to high K+ external containing 10 μM FM1-43 (Invitrogen) as described by Kuromi and Kidokoro (2000).
Loading of RP with FM1-43.
After dye loading of SVs cycling through ECP, nerve was stimulated at 10 Hz for 5 min in standard working solution containing FM1-43. FM1-43 was then removed by extensive rinsing with dye-free standard solution. Then, synaptic boutons were exposed for a second time to high K+ for 3 min to release SVs cycling through ECP, after which the external solution was extensively rinsed with standard working solution. The remaining FM1-43 fluorescence corresponds to SVs cycling through the RP (Delgado et al., 2000).
FM1-43 unloading.
In all cases, FM1-43 unloading was monitored in a standard working solution at rest, in the absence of nerve stimulation or by stimulating the nerve at low (0.5 Hz) or high frequency (10 Hz) using a programmable stimulator.
Imaging
Synaptic boutons were imaged using an epi-fluorescence microscope (BX50WI; Olympus) with a 60x objective. Samples were excited for 200–300 ms via a controlled shutter system (Uniblitz; Vincent Associates). Images were captured with a 12-bit cooled charge-coupled device camera (SensiCam; PCO Imaging). Images were acquired and analyzed using Workbench 2.2 software (Axon Imaging). Results are from at least three synaptic boutons from a minimum of six different larvae per experimental condition. Fluorescence analysis was performed after correcting for bleaching and background.
Online supplemental material
Acknowledgments
We thank Drs. Charles Zuker, Kristin White, Vivian Budnik, Mark Fortini, Mani Ramaswami, Subhabrata Sanyal, and Deborah Kimbrell for flies reagents and flies; Drs. Shyam Sharan, Mark Fortini, and Ira Daar for critical reading of the manuscript; and Jason de la Cruz for technical assistance.
This work was supported by funding from the intramural division of the National Cancer Institute. Work in the laboratory of P. Labarca is supported by FONDECYT and the Howard Hughes Medical Institute. Centro de Estudios Científicos is Millennium Institute funded in part by grants from Fundacion Andes and the Tinker Foundation. P. Labarca is a Howard Hughes Medical Institute international scholar. R.A. Jorquera was the recipient of a doctoral fellowship from Mejoramiento de la Calidad y la Equidad de la Educacion Superior.
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