A structural role for lipids in organelle shaping.
Background and Significance
Biological membranes are formed primarily from phospholipids organized into lamellar bilayer structures. In large membranes, local regions can often be considered to approximate flat surfaces or sheets. However, biological membranes may also adopt highly curved conformations such as in tubules of the endoplasmic reticulum (ER), small vesicles, edges of ER sheets or Golgi stacks, and fusion stalks. Curvature is associated with asymmetry across the bilayer: this can be achieved by altering the types or amounts of lipids in one or both monolayers, inserting hydrophobic or amphipathic regions of proteins into the membrane, or applying external scaffolding forces (Graham and Kozlov, 2010). Curvature may also arise by other lipid-dependent mechanisms such as line tension generated at the boundaries of patches of lipids in phase states of differing degrees of disorder coexisting within a membrane (Knorr et al., 2012).
Theoretical considerations suggest that lipids in regions of high curvature may adopt non-bilayer configurations such as hexagonal HII, or if cone shaped, may "fit" bilayer curvature more or less smoothly, reducing the energetic costs of forming such regions (Basanez et al., 1996; Kozlovsky et al., 2002; Kozlovsky and Kozlov, 2002; Cherno-mordik and Kozlov, 2008). To fit a highly curved region, lipids whose effective shapes approximate cones (e.g., diacyldycerol, phosphatidylethanolamine) are better than cylindrical lipids (such as phosphatidylcholine). Conical lipids have reduced head-to-tail cross area ratios compared to cylindrical lipids. Thus they have been suggested to fit the curved regions without the energetic penalties of leaving "hydrophobic voids" in the bilayer structure (Kozlovsky and Kozlov, 2002).
Lipids whose effective shape is conical or inverted conical can be more precisely characterized by their associated negative or positive curvatures, measured by whether monolayers of these lipids bend toward (cones) or away from (inverted cones) the head groups. Conical lipids can contribute to overall bilayer bending if asymmetrically disposed across the bilayer. It has been proposed that such asymmetry would be found in regions of extreme bending such as the fusion stalk, an intermediate favored in mechanistic models of membrane fusion (Kozlovsky and Kozlov, 2002; Chernomordik and Kozlov, 2008). Theoretical considerations suggest that the critical asymmetry in fusion stalks derives from negatively curved lipids in the proximal membranes of the two interacting bilayers (Kozlovsky and Kozlov, 2002). Support for this prediction comes from experiments in which protein-free lipid bilayers are fused in vitro. Such fusion is facilitated by incorporation of lipids exhibiting negative curvature into the proximal or contacting monolayers; conversely, it is inhibited by incorporation of lipids exhibiting positive curvature (Basanez et al., 1996; Villar et ed., 2000). This suggests the possibility that similar structural effects would be found in naturally occurring membranes, perhaps modulated by fusion proteins.
A variety of experiments suggest roles for proteins in maintaining organelle curvature and thus shape (Voeltz et al., 2006; Shibata et al., 2009, 2010). Proteins have been postulated to bend membranes by providing scaffolding, exerting forces as motor proteins, or inserting hydrophobic or amphipathic regions into monolayers (Shibata et al., 2009). For example, in the ER, a single membranous network continuous with the nuclear envelope, tubules that are highly curved in cross-section coexist with relatively flat sheets. Abundant reticulons and interacting proteins (DP1/Yoplp) contain hydrophobic domains that may form hairpins that insert into tubular regions (Hu et al., 2011). They may also act as scaffolding proteins. Their depletion in cells results in conversion of tubules to sheets; conversely, overexpression results in long tubules at the expense of sheets (Voeltz et al., 2006). The vertebrate proteins CLIMP-63, which appears to serve as a luminal spacer, and p180, as a sheet stacker, may oppose the formation of tubules by stabilizing flat sheets (Shibata et al., 2009. 2010; Friedman and Voeltz, 2011).
We wondered if lipids also play a role in organelle shaping as well as fusion. Such a role has been dismissed on the grounds that there is no evidence for strong lipid asymmetry in organelle membranes (Shibata et al., 2009). However, in the absence of data against localized structural lipid asymmetry in endomembranes, an experimental test of the role of conical lipids in organelle shaping seemed to us constructive. We therefore took indirect approaches to alter the amounts of one conical lipid--diacylglycerol (DAG)--in organelle membranes to assess the potential requirement for lipids exhibiting negative curvature (Doman et al., 2012). Although the function of DAG in signaling pathways is well established, especially at the plasma membrane, its potential contribution to endomem-brane structure (or membrane fusion) mostly derives from in vitro experiments on nuclear envelope assembly (Barona et al., 2005; Larijani and Poccia, 2012).
We summarize here experiments in which sea urchin oocytes and eggs are microinjected with enzymes that lower the concentration of DAG, such as cliacylglycerol kinase (DGK), followed by incorporation of exogenous 1.3 DAG to mitigate the effects of the enzyme. An analog of the signaling molecule 1,2 DAG, 1,3 DAG, while retaining conical shape, lacks the ability to bind to the CI domain of 1,2 DAG signaling target proteins (Sanchez-Pinera and Mi-col, 1999). Lowering DAG, by microinjection of enzymes that either lower its precursor phosphatidylinositol (4,5) bisphosphate ([PIP.sub.2]) or directly convert it to phosphatidic acid, delays or inhibits the membrane fusion leading to formation of the nuclear envelope, and transforms the ER into extended sheet structures at the expense of the more highly curved tubules. This suggests that membrane proteins are insufficient to maintain organelle shape or support fusion (Doman et al., 2012). The transformation of ER is prevented by delivery of exogenous 1.3 DAG or phosphatidylethanolamine, an unrelated lipid exhibiting negative curvature and not expected to be recognized by the injected DGK. Thus we emphasize a structural rather than a signaling role for DAG and propose approaches toward more detailed investigations of the roles of conical lipids in maintaining organelle shape and facilitating membrane fusion in vivo.
Materials and Methods
Specimens of the sea urchin species Lytechinus pictus were obtained from Marinus Scientific. Long Beach, California. or South Coast Bio-Marine, Terminal Island, California. Collection and manipulations of gametes and fertilization techniques have been previously described (Poccia and Green, 1986). Gamete quality was routinely monitored by fertilization and embryo culture to the blastula stages.
To soften fertilization envelopes prior to injection of fertilized eggs, eggs were incubated for I min in 0.22-mm Millipore-filtered seawater (MFSW) adjusted with freshly prepared 3-amino-1,2,4-triazole (ATA, Sigma-Aldrich) to 3 mmol [l.sup.-1] prior to fertilization. Two minutes thereafter, eggs were washed once in 3 mmol [l.sup.-1] ATA in MFSW and once in seawater lacking ATA.
Microinjection needles were prepared from 50-[micro]1 Drummond microcapillary tubes coated with Sigmacote (Sigma-Aldrich). Tubes were pulled with a Model P-87 Flaming/Brown micropipette puller (Sutter Instruments) set at 720 units Heat, 75 units Pull, 80 units Velocity, and 240 units Time. Needle tips were broken and, for aqueous injections, the needles were back-loaded with about 0.5-1.0 [micro]l of mercury (Terasaki and Jaffe, 1993).
The back of the injection needle was then firmly attached to one end of a piece of Intramedic tubing (Clay Adams) that was attached to the metal syringe tip of a Gilmont syringe, and the tubing, syringe tip, and syringe were entirely filled with Fluorinert (Sigma-Aldrich). The injection needle was then mounted onto a Narishige MN-151 joystick, and the Gilmont syringe was slowly turned to push the mercury nearly to the tip of the needle. Needles were then loaded with injection solutions (Terasaki and Jaffe, 1993).
Preparation of injection solutions
Wesson oil saturated with [DiIC.sub.18] from Sigma-Aldrich was used to label the ER of both unfertilized and fertilized eggs (Terasaki and Jaffe, 1993). DGK and synaptojanin 1 (Synl) were diluted in LB (10 mmol [l.sup.-1] HEPES, pH 8.0, 250 mmol [l.sup.-1] NaC1, 25 mmol [l.sup.-1] EGTA, 5 mmol [l.sup.-1] Mg[C1.sub.2], 110 mmol [l.sup.-1] glycine, 250 mmol [l.sup.-1] glycerol, 1 mmol [l.sup.-1] dithiothreitol) prior to injection ([diacylglycerol kinase [DGK, Sigma-Aldrich]; recombinant phosphoinositide 5-phosphatase synaptojanin 1 [Syn 1, prepared in the Cell Biophysics Laboratory at Cancer Research UK1). Injection concentrations given are pipette concentrations that typically become diluted about 20-100-fold in the cytosol.
Kiehart injection chambers were constructed using double-sided clear adhesive tape as spacers (Kiehart, 1982; Terasaki and Jaffe, 1993). Chambers were mounted on a Zeiss Standard upright microscope. The joystick micromanipulator was attached to a stable metal base. Eggs were loaded into chambers by capillary action. Chambers were then sealed with a thin layer of mineral oil to prevent evaporation. Needles were introduced through the oil into the chamber. After alignment of the microneedles with the eggs, the microscope stage was moved horizontally to gently push eggs into the injection needle (Terasaki and Jaffe, 1993).
Eggs were pierced manually, and the needle was removed by retracting the microscope stage. Injections of enzymes into fertilized eggs were done at least 20 minutes after the oil injection (which generally was done 10 min after fertilization) in order to allow the [DiIC1.sub.18] time to spread throughout the ER. Unfertilized eggs were injected with [DiIC1.sub.18] and injected with enzymes 30 min thereafter.
Preparation of small unilamellar vesicles
Small unilamellar vesicles (SUVs) were prepared from [beta]-BODIPY 500/510 [C.sub.12]-HPC (Molecular Probes D-3793), sn-1, 3 DAG (1.3 DAG, Avanti Polar Lipids 110581), L-[alpha]-phosphatidylcholine (PC. Avanti Polar Lipids 840051), and L-[alpha]-phosphatidylethanolamine (PE, Avanti Polar Lipids 213506). As a fluorescent marker, 200 [micro]l of 5 mmol [l.sup.-1] PC in chloroform was mixed with 1 [micro]l of 5 mmol [l.sup.-1] [beta]-BODIPY 500/510 [C.sub.12]-HPC. in chloroform in a glass vial. To make 20 mole% DAG-containing SUVs, 16 ml of this solution was then mixed with 12.9 [micro]1 of 1.6 mmol [l.sup.-1] sn-1,3 DAG in a mixture of equal volumes of toluene and methanol. This solution was dried by gently blowing argon into the vial, after which the vial was placed into a vacuum desiccator for 30 min. Thereafter, 100 [micro]l of LB was added and the suspension sonicated for three 10-s intervals, waiting 10 s between each sonication to allow the suspension to cool. PE-containing SUVs were prepared as above, substituting for 1,3 DAG the equivalent amount of PE (20 mole%). Control SUVs contained 100 mole% PC.
Sonicated SUV suspension (10-[micro]ml) was then added to a 100-p.1 suspension of unfertilized eggs. The fluorescent marker was incorporated throughout the eggs within 30 min. Eggs were subsequently washed in MFSW and then inserted into the injection chambers and injected as described.
A Zeiss LSM 5 PASCAL or a Nikon Eclipse Ti confocal microscope with 40X objective was used with the rhoda-mine filter set to monitor and image the ER. Images were imported into Volocity 6.0 (PerkinElmer Improvision) and image contrast was adjusted.
Effects of altering DAG levels on endoplasmic reticulum morphology Figure 1 shows the pattern of ER in an unfertilized sea urchin egg revealed after redistribution of the fluorescent dye DiIC18 from a microinjected oil droplet saturated with the dye. Since the dye is highly water insoluble, it labels only adjacent membranes that contact the oil (Terasaki and Jaffe, 1991). Since the only membrane system that is continuous throughout the cytoplasm is the ER, the dye, which is non-disruptive to development of the embryo, serves as a specific marker of ER. Note the delicate pattern of interconnected tubules that predominates. Sheets are seen as broad, highly fluorescent areas continuous at the edges with the tubules.
Since they are relatively dormant metabolically (Ernst, 1997), unfertilized eggs are ideal for studying alterations of the ER upon manipulation of lipid content. Moreover, results are not complicated by progression of the cell cycle, because eggs are arrested in a GO state until fertilized. Morphology is stable in the unfertilized eggs even though the ER is poised to undergo widespread disassembly and reformation into a continuous network at about 5 min postfertilization (Terasaki and Jaffe, 1991).
To alter the amount of DAG in unfertilized or fertilized egg membranes, we microinjected enzymes that would lower DAG either by converting it to phosphatidic acid or by depleting the major precursor of DAG,[ PIP.sub.2] (4,5), by converting the latter to PI(4)P (Fig. 2). These enzymes are DAG kinase (DGK) and the [PIP.sub.2], 5-phosphatase Syn 1, respectively. Morphological transitions were similar for either enzyme. At the lowest concentrations used there were no changes in ER morphology, but as enzyme concentration increased, the ER was converted to increasing amounts of sheets at the expense of the more highly curved tubules. A representative unfertilized egg treated with Synl is shown in Figure 3. Such sheets often form in large, gently curved stacks that appeared to "collapse" into denser and flatter regions (Fig. 4 and Domart et al., 2012). Since similar results occurred with DGK (see the following), the alteration is probably due to a decrease in DAG rather than to an increase in phosphatidic acid or P1(4)P.
To test if the morphological effects were due to DAG depletion, we pre-incubated eggs with 1,3 DAG carried in small unilamellar vesicles (SUVs), which are fusigenic. This isomer of the signaling molecule 1,2 DAG displays similar physical properties (conical shape, negative curvature) but does not bind the typical signaling targets of 1,2 DAG (Sanchez-Pinera and Micol. 1999). Under these conditions, fluorescent marker lipids rapidly dispersed throughout the egg. Figure 5 shows the effects of preloading the eggs using SUVs of 20 mole% 1,3 DAG: 80 mole% phophatidylcholine prior to the Syn 1 treatment. These eggs showed no alteration of ER morphology (compare Fig. 5 with Fig. 3).
These experiments were repeated to evaluate the ability of DGK to alter ER morphology in unfertilized eggs, to test for the dose-dependence of DGK, and to evaluate the ability of SUVs containing PE (which is estimated at half the degree of negative curvature as DAG; see table 1 in Churchward et al., 2008) to prevent the transition from tubule to sheet. We first established the range of DGK pipette concentrations that altered ER morphology. At a concentration greater than 50 [micro]g/ml. DGK is effective in altering the tubule-to-sheet ratio by at least 90 min post-injection (Fig. 6). The degree of alteration was also dose-dependent. Figure 7 shows a comparison of two eggs injected with either 100 or 250 [micro]g/m1 DGK. The higher dose decreased the average time to achieve an equivalent stage of transition by about 30-40 min as observed in four injected eggs.
Table 1 compares the ability of pre-treatment of eggs with SUVs of various compositions to prevent the tubule-to-sheet transition induced by DGK. SUVs containing 1.3 DAG were able to completely prevent the transition at < 500 [micro]g/ml DGK and delayed sheet formation at 500 pg/ml. Treatment with SUVs containing 20 mole% PE delayed or prevented the transition induced by 250 [micro]g/ml, whereas 100% PC SUVs or no treatment were unable to prevent the transition even at 100 [micro]g/ml DGK.
Table 1 Effect of preincubation of eggs with small unilamellar vesicles on tubule-to-sheet transitions [DGK] SUV Number Sheets Delayed Remained [mu]g/ml Composition Eggs Formed Sheets Tubular Injected Formed 100 20% 1,3 DAG 4 0 0 4 250 20% 1,3 DAG 5 0 0 5 500 20% 1,3 DAG 6 0 4 2 250 20% PE 4 0 3 1 100 100% PC 4 4 0 0 100 none 5 5 0 0
We conclude that enzymes that lower DAG content of membranes in vivo alter ER toward a less curved morphology, an effect that can be prevented by exogenous incorporation of phospholipids exhibiting negative curvature but lacking 1,2 DAG signaling capacity. Thus the morphology of the ER is not dependent solely on proteins.
Effects of altering DAG levels on membrane fusion and nuclear envelope formation
Since membrane curvature has also been postulated to play a role in membrane fusion because of the requirement for extreme bending in the fusion stalk, we tested the effects of microinjection of DGK and Synl on fusion of the nuclear envelopes of karyomeres. Karyomeres are individual chromosomes surrounded by nuclear envelopes that form in many but not all early embryos (Wilson, 1898). Fusion of the outer nuclear membranes followed by the inner has been well documented for the sea urchin by electron microscopy (Longo, 1972). Figure 8 shows the formation of karyomeres, each of which is surrounded by a fluorescent rim continuous with the ER at anaphase in sea urchin embyros. The cytoplasmic region around the karyomeres is characterized by a concentration of ER sheets. As the cell cycle progresses, karyomeres continually fuse into larger structures until two single nuclei with single complete nuclear envelopes are formed at late telophase in each daughter cell. This process is completed by about 85 min post-fertilization for the first embryonic cell cycle and by about 144 min for the second cycle in L. pictus at 22 [degrees]C (Fig. 9).
Enzymes were titrated over a 10,000-fold concentration range. An example is shown in Figure 10 in which karyomere fusion is greatly delayed in eggs injected with 100 [micro]g/m1 DGK. Under these conditions the extent of karyomere fusion between 97 and 134 min is equivalent to control embryos at about 70-80 min. However, by 134 min, control embryos exhibited almost complete karyomere resolution of the second cell cycle. Similar effects were observed with Syn 1 (Doman et al., 2012). The extent of retardation of karyomere resolution was dose-dependent with either enzyme. Therefore, lowering DAG content inhibits karyomere membrane fusion. Moreover, concomitant dose-dependent effects on ER morphology similar to those seen in unfertilized eggs were also apparent in the embryos, with the formation of extensive sheet regions at the expense of tubules (Fig. 11).
Although there is considerable evidence that proteins can alter bending of biological membranes and are required for maintenance of organelle morphology, such evidence does not rule out a role for membrane lipids. Manipulation of protein levels in cells through mutation or transfection is widely used to investigate requirements for proteins. Alteration of lipid levels to investigate their potential roles in membrane structure requires techniques that are rapid, direct, and specific in vivo.
The experiments outlined in preceding sections using the sea urchin egg and early embryo as a model system suggest a requirement for diacylglycerol in maintaining ER morphology or facilitating membrane fusion. We interpret these phenomena as having a common basis in the ability of DAG to contribute to the high degrees of membrane curvature required for maintenance of ER tubules or fusion stalks. That these effects appear rapidly without concomitant attempts to alter resident proteins strongly suggests that proteins, although they may be required, are not sufficient for either phenomenon.
The advantages of using the sea urchin system described here are several. (1) Fusion of nuclear envelopes to resolve multiple karyomeres into single nuclei represents a well-documented case of membrane fusion occurring in the first cell cycle. A second extensively studied membrane fusion system is the cortical granule-plasma membrane system that has been exploited in vitro by Vacquier (1975). In several studies, evidence was presented for the importance of cholesterol and other lipids that impart negative curvature in cortical granule fusion (reviewed in this issue by Abbenini et al., pp. 200-217). (2) Fusion of nuclear membrane precursor vesicles in vitro has been well documented in the sea urchin: a class of vesicles highly enriched in the precursors and enzymes necessary to generate DAG that initiates membrane fusion has been biochemically characterized (Collas and Poccia, 1996: Larijani and Poccia, 2009: Dumas et al., 2010). (3) The unfertilized egg is arrested in stage GO of the cell cycle, is relatively dormant metabolically, and lacks multiple signaling interactions occurring in the activated embryonic cells, allowing a more direct and less ambiguous readout of the effects of lipid alterations on ER morphology. (4) Dose-dependent effects are easily evaluated by control of concentrations in solutions used for microinjection into the large egg cells. (5) The ability to incorporate lipids of negative curvature into the egg by fusion with SUVs prevents the effects of lipid depletion by microinjected enzymes. (6) All altered cells in a given experiment can be monitored and compared with adjacent unaltered control cells.
The experiments discussed here have their parallels in work done in mammalian cells using an inducible dimerization device (Domart et al., 2012). The expression of fusion proteins containing FKBP12, cyclophilin, and FRB domains can be used in varying combinations with homo- or hetero-dimerizing ligands to induce acute dimerizafion of exogenous proteins (Schreiber, 1991; Fili et al., 2006). To deplete DAG, lipid-modifying enzymes were fused to FKBP12 and targeted specifically to the ER and its nuclear membrane subdomain. The results of such experiments are remarkably similar to those described here in the sea urchin. When DAG was depleted by either DGK or the phosphoinositide 5-phosphatase SKIP, nuclear envelope formation was inhibited and the ER was reorganized into multilamellar sheets. Cells could be rescued by incubation with SUVs containing either 1,2 DAG or 1,3 DAG, emphasizing again a structural rather than a signaling role for this neutral lipid in endomembranes.
Since membrane bending is predicted to depend on asymmetry of lipid disposition across the bilayer (i.e., different amounts of conical lipids in each monolayer), one interpretation of the experiments discussed here is that DAG depletion alters the ability of membranes to form highly curved structures that depend on DAG asymmetry across the bi-layer. However, since DAG represents a lipid with the greatest potential to flip-flop across the bilayer, what would keep DAG from equilibrating? One possibility would be for DAG to bind to a protein that is asymmetrically disposed across the membrane. Another possibility in naturally occurring membranes is that proteins that provide energy for bending, but do not insert into the bilayer, might at some point be limited in their bending ability by the penalty to be paid for creating hydrophobic voids in the bent regions, unless appropriate lipids could be concentrated there (Koz-lovsky and Kozlov, 2002). In such a case, a lipid like DAG would be ideal for recruitment to the extremely curved regions. DAG could re-equilibrate across the bilayer to fill the necessary gaps, thereby generating the predicted lipid asymmetry. Here. DAG asymmetry would be a consequence of bending, not its cause. Bending would depend on proteins that are necessary but not sufficient in the absence of adequate quantities of lipids with high negative curvatures, such as DAG.
We suggest that lipids, in addition to their well-known signaling roles, are critical for maintaining organelle shape and promoting fusion through effects on membrane bending. Although the effects of lipids on fusion of protein-free membranes have long been investigated, such studies of natural membranes are still too few. One reason is that it is not a simple matter to alter lipid composition in specific biological membranes. The alternate approaches taken with sea urchin and mammalian cells to rapidly alter membrane lipids in vivo each have their distinct and complementary advantages. In the case of the sea urchin, alteration of lipids and assessment of morphology is very rapid, within minutes, and can be titrated by simply controlling the concentration of microinjected enzymes. The system is naturally synchronous, timing is precise, and cell cycle events can be eliminated or studied in the same cell type by comparing unfertilized and first-cycle fertilized eggs. The time of enzyme introduction into phases of the cell cycle can be controlled with a precision of minutes post-fertilization. However, injected enzymes are not directed to specific organelles. Major advantages of the rapalogue dimerization device used in mammalian cells are the ability to target enzymes to any organelle for which a specific membrane protein can be identified and the ability thereafter to induce it acutely with rapalogue. However, the levels of modifying enzymes or timing of the cell cycle cannot be as precisely controlled as in the sea urchin. Additional disadvantages are the relatively long cell cycles (24 h vs. 1 h) and the few cells at mitotic stages in an unsynchronized population that would otherwise require harsh treatments to induce synchrony.
Each system can be further exploited. In the sea urchin, for example, one can attempt to rescue depletion of DAG by various chemically unrelated lipids of various degrees of negative curvature. Rescue could be quantified by titration to determine if the degree of curvature is related to the effectiveness of rescue or whether other considerations should be taken into account. Other membrane fusion events such as male-female pronuclear fusion or cortical granule exocytosis could be evaluated in eggs depleted of DAG. Would exocytosis at fertilization be blocked? Would fertilization or parthenogenetic activation of DAG-depleted eggs result in perturbation of the disassembly or reassembly of the ER that normally occurs soon after fertilization?
In mammalian cells, specific targeting of the dimeiization device with various lipid-modifying enzymes to different organelles could reveal whether DAG or other lipids are generally required to maintain organelle shaping or membrane fusion. Although at least some proteins involved in shaping or fusion are likely to be membrane-specific, it remains to be established how either process might depend on structural roles of specific membrane lipids in individual organelles. It is also not clear if all shaping or fusion phenomena would share common mechanisms regarding lipid participation. Distinguishing structural and signaling roles will be of paramount importance.
The relative contributions of proteins and lipids to creation or maintenance of membrane curvature in different organelles and to fission and fusion processes will need to be assessed in future experiments. These experiments should lead to a fuller understanding of how lipids of various shapes might contribute to the structure and function of membranes and the organelles they delimit.
We gratefully acknowledge Marie-Charlotte Domart for discussions and Karen Yeh for the data of Fig. 11. Support was received from an Amherst College Schupf Fellowship to A.W., a Faculty Research Award of the Axel Schupf '57 Fund for Intellectual Life to D.P., and Cancer Research UK Core Funding to the London Research Institute to B.L.
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Received 17 January 2013; accepted 7 June 2013.
* To whom correspondence should be addressed. E-mail: email@example.com
[dagger] Current addresses: A.S.W., National Institutes of Health, National Cancer Institute, 6120 Executive Boulevard, Rockville, MD 20892: B.F.. Nova Southeastern Univ.. 3301 College Ave. Fort Lauderdale, FL 33314.
Abbreviations: DAG, diacylglycerol; ER, endoplasmic reticulum: PI P2, phosphatidylinositol (4.5) bisphosphate: DGK, diacylglycerol kinase: Syn 1. synaptojanin 1; SUV, small unilamellar vesicle.
ALAN S. WANG (1), [dagger] AUPOLA KUNDU (2), BURR FONG (1), [dagger], JULIE FITZGERALD (1), BANAFSHE LARIJANI (3), AND DOMINIC POCCIA (1), (2), (*)
(1) Department of Biology and (2) Program in Neuroscience, Amherst College, Amherst, Massachusetts 01002; and (3) Cell Biophysics Laboratory, London Research Institute, Cancer Research UK, 44 Lincoln's Inn Fields, WC2A 3LY, London, United Kingdom
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|Author:||Wang, Alan S.; Kundu, Aupola; Fong, Burr; Fitzgerald, Julie; Laruani, Banafshe; Poccia, Dominic|
|Publication:||The Biological Bulletin|
|Date:||Aug 1, 2013|
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