Apr 8, 1982 - colchicine, and podophyllotoxin) did not have any significant effect on the tubulin- mediated or Ca2'-enhanced aggregation and fusion of ...
THEJOURNAL
OF
BIOLOGICAL CHEMISTRY
Vol. 257, No. 24, Issue of December 25, pp. 15137-15144,1982 Printed in U.S.A.
Aggregation and Calcium-induced Fusion of Phosphatidylcholine Vesicle-Tubulin Complexes* (Received for publication, April 8, 1982)
Nirbhay KumarSB, RobertBlumenthalf, Maryanna Henkartll,John N. Weinsteinl, and RichardD. Klausnerf * * From the *Laboratory of Cell Biology, National Heart, Lung, and Blood Institute, and the (IlmmunologyBranch, Division of Cancer Biology and Diagnosis, National Cancer Institute, and the !Laboratory of Theoretical BioZogy, Division of Cancer Biology and Diagnosis, National Cancer Institute, National Institutes of Health, Bethesda, Maryland 20205
Insertion of tubulin into the bilayer of dipalmitoyl tubulin becomes an integral component of the membrane. Soifer and Czosnek ( 5 ) have reported that tubulin is synthephosphatidylcholine vesicles atthephasetransition results in the formation of stable vesicle-tubulin com- sized on rough microsomes, incorporatedinto microsomal plexes (Klausner, R. D., Kumar, N., Weinstein, J. N., membranes, and translocated to the plasma membrane as a Blumenthal, R., and Flavin,M. (1981) J.Biol. Chem 256, component of vesicles. 5879-5885). These complexes aggregated when mainWe have recently shown (6, 7 ) that purified tubulin can be tained below phase transition for 10-20 min. Addition incorporated into dipalmitoyl phosphatidylcholine vesicles at of millimolar concentrations of Ca2+,Mn2+, Zn2+, and the lipid phase transition temperature, without any requireCo2+,butnot M&+, causedthevesicle-tubulincomment for detergent or sonication, and resultsin the formation plexes to fuse into larger structures as shown by (a) of stable complexes. Similar results had been reported by electron microscopy,( b )increased trapped volume, andCaron and Berlin (8) using dimyristoyl phosphatidylcholine (c) changes in resonance energy transfer betweentwo vesicles. The insertion process is accompanied by structural fluorescentlipidprobesincorporatedintothesame vesicle. There was noloss of internal aqueous contents perturbations of both the lipid bilayer and the tubulin (6, 7 ) . from thevesicle-tubulincomplexesduring Ca2'-in- In this paperwe examine the propertiesof the vesicle-tubulin complexes. These complexes aggregated even in the absence duced fusion. Anti-tubulin drugs had no effect on the aggregation or fusion, and vesicle-bound tubulin did of divalent cations; inthe presenceof millimolar CaZ+or Mn2+ closed structures not associate with microtubules when tubulinwas as- but notMg2+,they fused to form much larger sembled in vitro. Trypsin-treated vesicle-tubulin com- without releaseof the water-soluble internal marker,carboxyplexes were incapable of supporting Ca2'-induced fu- fluorescein. We have also compared free and vesicle-bound tubulin for their ability bind to colchicine and MAPs' orto be sion. This system provides a model for Ca2'-induced tyrosinolated and detyrosinolatedby specific enzymes. andprotein-mediatednonleakyfusionofuncharged lipid bilayers. MATERIALS AND METHODS
Although tubulin, thebasic structural unitof microtubules, is generally considered to be a cytoplasmic protein, there are data whichsuggest that it is alsoassociated with various organelles and membranes(1-3). In additionto their primary role in mitosis, microtubules have also been implicated in a number of membrane-linked processes such as cell motility, secretion, transport, redistribution of membrane proteins, and maintenance of cell shapeand size. So far,there is little knowledge regardingthe specific role of membrane-associated tubulin. It has been suggested that membrane-bound tubulin could be involved in interactions between membranous vesicles and cytoskeletal elements or between vesicles and the plasma membrane(4). Of equal interest is the question of how
Preparation of Tubulin-Microtubule protein was purified from freshly obtained bovine brains by three cycles of assembly and disassembly without glycerol, according to the procedure of Asnes and Wilson (9). Tubulin2 was further purified by phosphocellulose chro(6) and stored in buffer containing matography as described elsewhere 100mM K'-2-(N-morpho1ino)ethanesulfonic acid, 1 m M MgSO.,, 1mM EGTA, 2 mM dithiothreitol, and 0.1 mM GTP, pH6.8, at -70"C. MAPS which remain bound in the column were subsequently eluted with 1.0 M KC1in the above buffer, concentrated using an Amicon PM-30 filter, and storedat -70 "C. Protein was determined according to the method ofLowry et al. (10) using bovine serum albumin as standard. Preparation of Vesicles-Small unilamellar vesicles of DPPC labeled with either [14C]DPPCor [3H]DPPC but without carboxyfluoresceinwereprepared as described elsewhere (6). DPPC vesicles labeled with N-NBD-PE and N-Rh-PE (Avanti Polar Lipids, Inc., Birmingham, AL) at aratio of 98:l:l (see below) hadthe same elution profile as unlabeled DPPC vesicles on a Sepharose 4B gel filtration column. Vesicles were kept at room temperature (25 "C) and used within 24 h. Lipid vesicle-tubulin complexes were prepared by incubating tubulin with DPPC vesicles at their liquid crystalline phase
* The costs of publication of this article were defrayed in part by the payment of page charges. This article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact. at the XI1 International §A part of this workwaspresented Congress of Biochemistry, Perth (August 1982), and travel funds were ' The abbreviations used are: MAPs, microtubule-associated proobtained from the International UnionofBiochemistry. Present teins; DPPC, dipalmitoyl phosphatidylcholine; T c , phase transition; address,Building8,Room320, National Institute of Allergy and EGTA, ethylene glycol bis(P-aminoethyl ether)-N,N,N'N'-tetraacetic InfectiousDiseases,National Institutes of Health, Bethesda, MD acid; N-NBD-PE, N-(7-nitro-2,1,3-benzoxadiazol-4-yl)phosphatidyl20205. ethanolamine;N-Rh-PE, N-(lissamine RhodamineB sulfonyl) phos**Present address, Laboratory of Biochemistry and Metabolism, phatidylethanolamine; Hepes, 4-(2-hydroxyethyl)-l-piperazineethBuilding 10, Room 9N116, National Institute of Arthritis, Diabetes, anesulfonic acid. and Digestive and Kidney Diseases, National Institutes of Health, This refers to tubulin purified by phosphocellulose chromatograBethesda, MD 20205. phy. 3 X tubulin refers to three assembly cycles purified tubulin.
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15138
Ca2+-inducedFusion of Vesicle-TubulinComplexes
transition temperature, i.e. 40 “C for 2 min. We have shown earlier that tubulin is inserted into the bilayer of these vesicles at thephase transition to form stable complexes (6). The trapped volume of vesicles (volume of the totalinternal aqueous compartment) was determined by preparing and incubating the vesicles at 40 “C in the presence of 20 mM carboxyfluorescein. The vesicles and vesicle-tubulin complexes werepassed over a Sephadex G-25 (PD-10, Pharmacia) column, and the total internal volume of the vesicle per mol ofphospholipid was determined by spectrofluorimetric measurement of carboxyfluorescein. The Ca*+-induced “fused” vesicles were too large to pass through the G-25 column. They were centrifuged at 100,ooO X g in an Airfuge (Beckman Instruments). The pellet was washed seven times with buffer containing 150 mM NaCl, 10 mM Hepes, and 1 mM EGTA, pH 7, dissolved with Triton X-100, and carboxyfluorescein and lipid were measured. The volume of the aqueous compartment of vesicles relative to total volume was determined by measuring the ratio between fluorescence before and after separatingthe vesicles from the medium containing 20 mM carboxyfluorescein. The phospholipid concentration (mol/ liter) was measured after the separation. The trapped volume (liters/ mol) was then determined by dividing the fluorescence ratio by the phospholipid concentration. Centrifugation Assay for Aggregation/Fusion of Vesicles or Vesicle-Tubulin Complexes-This assay made use of the change in sedimentation constant as a result of aggregation and/or fusion of vesicles. Routinely, vesicles alone or vesicle-tubulin complexes were incubated for 15-30 min below the phase transition temperature of DPPC (usually at 28-30 “C) in tubes (7 X 50 mm; cellulose acetate butyrate). The assay buffer was 100 mM K+-2-(N-morpholino)ethanesulfonic acid (pH 6.8) containing 0.5 mM MgS04 and 0.75 mM EGTA. At the end of incubation, tubes were centrifuged (using adaptors no. 408) a t 40,000 X g for 20 min a t 25 “C (SorvallRotor SS34). Supernatants were removed for counting. The bottoms of the tubes were cut off and dropped into scintillation vials for counting. The recovery of total radioactivity (pellet andsupernatant) was >98%. Results are expressed as the ratio of radioactivity in the pellet/ total radioactivity. The actual amountpelleted varied between 20 and 65% (at a tubulin to vesicle molar ratio of 101) with several vesicle and/or tubulin preparations. In general, this variability was seen only with the pelleting of vesicle-tubulin complexes in the absence of added Ca2+.However, any single preparation gave extremely reproducible results. ResonanceEnergyTransfer between Lipid Probes-Mixing of lipids was assayed according to the procedure developed by Pagano and his co-workers (11)involving resonance energy transfer between two fluorophores incorporated into the vesicle bilayer. The energy donor NBD and the energy acceptor rhodamine are coupled to the free amino group of phosphatidylethanolamine to form N-NBD-PE and N-Rh-PE (11). The two lipid probes were obtained from Avanti Polar Lipids. As shown by Struck et al. (11)when both fluorescent lipids are in lipid vesicles at appropriate surface densities (ratio of fluorescent lipid to total lipid), efficient energy transfer is observed. Steady state emission and excitation spectra were obtained by using a Perkin-Elmer MPF 44b spectrofluorometer, the excitation band slit was at 4 nm, and the emission slit at 10 nm. Following each set of measurements, vesicles were disrupted with Triton X-100 (0.1% final concentration). This treatment eliminated energy transfer (see below) and allowed the determination of the concentrations of NNBD-PE and N-Rh-PEfrom their emission intensities by usingdirect excitation. All spectra were obtained at room temperature (22 “C). Spectra of tubulin-vesicle preparations containing both N-NBDPE and N-Rh-PE (DPPC:N-NBD-PE:N-Rh-PE = 981:1), excited at 450 nm, show emission maxima at 530 nm and 585 nm (see Fig. 5 under “Results”). Essentially all of the fluorescence at 530 nm comes from N-NBD-PE, whereas the fluorescence at 585 nm arises from fluorescence energy transfer between the donor and acceptor pair. The efficiency of energy transfer (quenching of the energy donor) in such samples is defined by the relationship (12)
E
=
1 - FIFO
(1)
where F is the fluorescence at 530 nm in the presence of N-Rh-PE and Fo is the fluorescence at 530 nm in the absence of N-Rh-PE. Fig. 5 shows that in the presence of detergent, therhodamine fluorescence at 585 nm is completely abolished when excited at 450 nm, indicating complete mixing of the fluorophores with excess DPPC lipid. We define the NBD fluorescence at 530 nm in the presence of detergent as Fo. The energy transfer efficiency for the DPPC:N-NBD-PE:N-
Rh-PE = 98:l:l vesicles (86%) was about equal to that reported by Struck et al.(11)for phosphatidylserine vesicles containing the same mol per cent N-NBD-PE and N-Rh-PE. Electron Microscopy-For electron microscopy, DPPC vesicles and vesicle-tubulin complexes with or without Ca2+were negatively stained within an hour of the time that thevesicle-tubulin complexes were prepared. Uranyl acetate and phosphotungstic acid in several protocols were tested for studying the vesicles and vesicle-tubulin complexes in the electron microscope. Procedures included mixing the vesicles with stains before application to the grids or application of the vesicles to thegrids followed by blotting and/or various washes before application of the stain. (The problems and advantages of these techniques are considered in more detail under “Discussion”.) After several stains and protocols had been tested, the following procedure was adopted. All manipulations were done a t room temperature which is below Tc for DPPC. A drop (about 8 p l ) of each sample was pipetted onto carbon and Formvar-coated grids. After 1 min or less, the grids were blotted but not allowed to dry. A drop of 2%phosphotungstic acid, pH 6.5, wasapplied, and after 1 min or less the grids were blotted, allowed to dry, and examined using a Phillips 400 microscope at 60 kV. RESULTS
Aggregation of Vesicle-Tubulin Complexes Fig. 1 shows the typical turbidity changes (indicated by changes in 90” light scattering at 470 nm) of DPPC vesicles upon addition of various concentrations of tubulin at 40 “C. Without tubulin, there was no change in the light scattered by vesicles, whereas increasing amounts of tubulin resultedin an increase in the scattering, probably as a result of aggregation of the small unilamellar DPPC vesicles. There was very close correspondence in the time course for the turbidity increase and the release of encapsulated carboxyfluorescein reported previously (6). The change in turbidity takes place only when tubulin is incubated withvesicles around 37-40 “C (which is also the transition temperature), A likely interpretation of the observed increase in the turbidity is that tubulin promoted the aggregation of vesicles under conditions in which it is inserted into thelipid bilayer (6).A variety of other soluble proteins such as hemoglobin, immunoglobulin, albumin, and ovalbumin do not produce an effect similar to that of tubulin on these vesicles. Formation of larger structures was also indicated by the relative increase in the sedimentation of vesicles as shown in Table I and Fig. 2. The pelleting of the vesicle-tubulin complexes allowed us to study further the effect of divalent cations. Fig. 2 shows the effect of 2.5 mM CaC12 on the pelleting of DPPC vesicles. Under these conditions, Ca2+ did not increase the pelleting of DPPC vesicles
-
-
-!
I .-c C
-
I
1
I
1
15
30
45
60
TIME Is)
FIG. 1. Effect of tubulin on light scatteringof DPPC vesicles. Freshly prepared DPPC vesicles(4.5 lipid) containing carboxyfluorescein were incubated with various amounts of t u b u l i in the cuvettes. The numbers in the figure indicate molar ratio of tubulin to vesicles (assuming 4500 lipid per vesicle). Temperature of the cuvette holder was maintained by recirculating water at 42 “C. Ninety degree light scattered at 470 run was determined using an Aminco-Bowman spectrofluorometer with 2-mm slits and a two-channel Y-t recorder. Tubulin was added a t time zero (belowphase transition temperature). Experiments were performed in the presence of 1m~ EGTA. Tubulin alone did not show any light scattering changes.
Cu2’-induced Fusion of Vesicle-Tubulin Complexes
15139
was also a substantial population of free small vesicles. Within
TABLEI
the clumps, individual small vesicles of the original size were Effect of varying tubulin/vesicle ratioon the pelleting of vesicles Vesicle-tubulin complexesof varying tubulin to vesicle ratios were clearly distinguishable (Fig. 4C). Fig. 4C was selected from an prepared in buffer containing 0.75 mM EGTA. They were incubated at 30 “C for 15 min before centrifugation (40,000 X g for 30 min) to determine pelleting of vesicles, as described under “Materials and Methods.” Tubulin/vesicle
[“CIDPPC (pellet/total)
mol/mol 0
0.09
3 6 15 30
1.0
0.17 0.25 0.36 0.48
-
1
10.971
10.691
1
2
3
4
I
1 5
FIG. 2. Effect of Caz+ on pelleting of vesicles and vesicletubulin complexes. Lipidconcentrationwas 2.7 mM. 1, vesicles alone; 2, vesicles + 2.5 m~ CaC12;3, vesicles + tubulin (6 PM) without passage through the phase transition;4, vesicle-tubulin complexes;5, vesicle-tubulin complexes + 2.5 mM CaClz. All the incubations and centrifugation were at 30 “C. Numbers in parentheses represent the results obtained when incubation and centrifugation were done at 0 “C.
alone nor was there any significant effect on the pelleting when tubulin and vesicles were mixed belowTc. When tubulin was inserted into the lipid bilayer at the endothermic Tc approximately 40 to 65% of the vesicles could be pelleted in the absence of Ca2+.Addition of Ca2+to the vesicle-tubulin complexes, however, caused nearly complete pelleting of the vesicles (Fig. 2). The effect of Ca2+was dependent upon the final concentrations added to vesicle-tubulin complexes. As shown in Fig. 3, maximum pelleting occurred above 2.5 mM concentration. Since these assays were performed in buffer containing 0.75 mM EGTA, the threshold concentration of free Caz+for maximum pelleting was above 1.0 mM. The inset of Fig. 3 also shows that Mg2’ was without any effect, whereas Mn2+,Zn2+,and Co2+enhanced the pelleting of the vesicletubulin complexes. Effectiveness of various cations testedwas Mn2+2 Ca2+2 Zn2+> Co2+> Mg2+. Evidence for Ca2+-induced Fusion Electron Microscopy-Fig. 4 is a series of electron micrographs preparedby the technique described under “Materials and Methods.” It illustrates the effect of Ca2+ on vesicletubulin complexes. The population of DPPC vesicles eluted from the Sepharose 4B column contained primarily single spherical vesicles 18-30 nm in diameter.Among these vesicles, there was a small number of larger vesicles (up to 10 nm in diameter) each of which appeared to contain several of the small vesicles (Fig. 4A). DPPC vesicles incubated with 5 mM Ca2+ were indistinguishable from thosewithout Ca2’ (Fig. 4B).DPPC vesicles into which tubulin had been incorporated by passage through Tc showed marked clumping, but there
area including some of the largest aggregates found. Tubulincontaining vesicles incubated in the presence of 5 mM Ca2+ completely gathered into clumps with no free vesicles apparent. Within these clumps in addition to some distinguishable small vesicles there were also vesicles of a variety of larger sizes as well as much larger structures apparentlycomposed of sheets of membrane folded into topologically complex forms (Fig. 40). Fig. 4 0 was selected from an area showing the highest proportion of distinguishable small vesicles in addition to thelarger membrane forms. The most obvious interpretation is that the larger vesicles and complex membrane structures are formed by fusion of the small vesicles. The differences between vesicles and vesicle-tubulin complexes with or without ea2+were obvious and reproducible. Trapped Volume-Small unilamellar vesicles as well as vesicle-tubulin complexes at a 30:l ratio (tubulin to vesicle) were formed in 150 mM NaC1-10 m~ Hepes buffer, pH 7.0, containing 20 m~ carboxyfluorescein and 1mM EGTA. Fusion was induced by adding Ca2+to a final concentration of 10 mM to the vesicle-tubulin complexes in a medium containing 20 mM carboxyfluorescein. The trapped volume was measured after separation of the vesicles from the medium. As shown in Table 11, vesicle-tubulin complexes had a 2-fold increase in trapped volume over the original vesicles, and Ca2+-induced fusion produced a 10-fold increase in trapped volume. This result indicatesthat at least some of the large structures seen in Fig. 4 0 are closed vesicles. Mixing of Lipid Probes-In Fig. 5 we show spectra of a tubulin-vesicle preparation containing both N-NBD-PE (donor) and N-Rh-PE (acceptor) (DPPC:N-NBD-PE:N-Rh-PE = 98:1:1), excited at 450 nm. Emission maxima at 530 nm and 585 nm areobserved. Essentially all of the fluorescence at 530 nm comes from N-NBD-PE, whereas the fluorescence at 585 nm arises from fluorescence energy transfer between the donor and acceptor. The tubulin-vesicle complexes were mixed with tubulin-vesicle complexes (without probes)at a (vesicle/vesicle) ratio of 1:20, and fusion was initiated by addition of Ca“. the As shown inFig. 5 there was amarkedreductionin emission peak at 585 nm and a concomitant increase in fluorescent yield from N-NBD-PE at 530 nm upon addition of Ca2+.These changes indicate a reduction inthe efficiency of energy transfer between N-NBD-PE and N-Rh-PE. This is consistent with fusion between the DPPC/N-NBD-PE/NRh-PE-tubulin complexes and pure DPPC-tubulincomplexes, followed by lateral diffusion of the fluorescent lipids in the 1.0
c
COCI,
0.51
I
1
I
2
I
3
I
4
5
CaCIZ CONCENTRATION LmMl
FIG. 3. Effect of varying CaClz concentration on vpiicle-tubulin complexes. Vesicle-tubulin complexes were incubated for 30 min at 30 “C in the presence of various concentrations of CaC12.Data in the inset shows the pelleting of vesicles in the presence of 2.5 mM concentration of other cations. Assay buffer contained 0.75 mM EGTA.
15140
Ca2+-inducedFusion of Vesicle-Tubulin Complexes
FIG. 4. Electron micrographsof negatively stained vesicles.165,000 x: A, DPPC vesicles; B, DPPC vesicles in the presence of 5 mM CaC12; C, DPPC vesicle-tubulin complexes containing 10 tubulin molecules per vesicle in the presence of 5 m~ EGTA; D, DPPC vesicletubulin complexes containing 10 tubulin molecules per vesicle in the presence of 5 mM CaCL. The bar indicates 200 nm.
TABLE I1 Trapped volume of vesicles and vesicle-tubulin complexes with and without Ca'+ The vesicles and vesicle-tubulin complexes were prepared in 20 mM carboxyfluorescein, 150 mM NaCI, 10 mM Hepes, 1 mM EGTA, pH 7. Trapped volume was measured after separation of vesicles or vesicletubulin complexes from the incubation medium. Tubulin to vesicle ratio was 301. Trapped volume liter/mol
Vesicles Vesicle-tubulin complexes Vesicle-tubulin complexes + 10 mM ca2+
0.36 0.85 4.33
plane of the newly formed membrane. Struck et al. (11)have shown that the energy transfer efficiency of vesicles containing 1% N-NBD-PEincreases linearly as a function of mol % (surface density) N-Rh-PE up to about 1 mol % N-Rh-PE. Therefore, a 2-fold decrease in energy transfer efficiency rep-
resents a 2-fold decrease in surface density, consistent with fusion of one probe-containingvesicle with one purevesicle. In Fig. 6B we have plotted the energy transfer efficiency calculated according toEquation 1 as a function of Ca2+ concentration. At about 1 mM Ca2+ there isa 2-fold decrease in transfer efficiency, indicating that on the average each fluorescent vesicle has fused with an unlabeled vesicle. The Ca2+ dependence shownin Fig. 6 is quite similar to the Ca2+ dependence of pelleting shown in Fig. 1. In accordance with the divalent cation specificity of pelleting we also see that Mg2+had littleeffect on energy transfer, whereas the effect of Mn2+ was similar to that of Ca". When the probe-containing vesicles were incubated with Ca2+ in the absence of pure vesicles there was no change in energy transfer, indicating that theeffect was not due to,for instance, a change in the radius of curvature of the vesicles. However, when we incubated probe-containing vesicle-tubulin complexes with DPPC vesicles without tubulin we did see Ca2+-dependentchangesinenergytransfer (Fig. 6). This
Ca”-induced
WAVE
LENGTH
Fusion of Vesicle-Tubulin
(nm)
FIG. 5. Effect of Ca’+-induced fusion on energy transfer. The complexes were formed at a tubulin to vesicle ratio of 3O:l. The labeled complexes (DPPC/N-NBD-PE/N-Rh-PE (98:l:l) and tubulin), were mixed with the unlabeled DPPC vesicle-tubulin complexes at a vesicle:vesicle ratio of 1:20, and CaClz was added to induce fusion. The emission spectra were obtained by exciting the samples at 450 nm immediately after mixing, as described under “Materials and Methods.” The numbers indicate final Ca2+ concentrations.
$u Y lo 0 dz 1----D------
-- -___
o--------o-
i
NO ADDED VESICLES ------a
FUSION
I 0
I
,
L
,
L
2
4
6
8
10
C&b
CONCENTRATION
WITH VESICI .ES
I 12
14
(rnbl)
FIG. 6. Effect of Ca2+ on fusion of DPPC-tuhulin complexes. A, DPPC-tubulin complexes were formed by bringing DPPC vesicles containing 120 rnM carboxyfluorescein to Tc in the presence of tubulin at a vesicle to tubulin ratio of 1:30. The complexes were then immediately rechromatographed on a Sephadex G-25 (PD-10) column to remove carboxyfluorescein released at the phase transition. The ratio of fluorescence of the rechromatographed complexes before and after treatment with 0.1% Triton X-100 was 0.23. Leakage was assessed by measuring fluorescence after mixing with Ca*+ and subtracting the background fluorescence of the complexes (no Ca*‘). B, energy transfer was assessed as described in the legend to Fig. 5. Transfer efficiency was calculated according to Equation 1 (see under “Materials and Methods”). The numbers were not corrected for the effect of Triton X-100 on NBD fluorescence. DPPC-tubulin complexes containing the
probes were mixed with unlabeled DPPC-tubulin DPPC vesicles efficiency in
alone
(0)
in the
presence
complexes (0) and
of CaC12. Energy
transfer
the presence of Mn” (A) or Mg2+ (V) is also shown. Energy transfer in the presence of Ca” but without any acceptor vesicles is indicated by n . result tween
indicates that vesicle-tubulin
Cax+-induced complexes
and
fusion can protein-free
also
occur vesicles,
15141
Complexes
be mixed immediately after formation with the unlabeled vesicles to produce efficient Ca’+-dependent energy transfer changes. When they were allowed to stand for a while in the absence of Ca2+ they presumably aggregated. When those were mixed with unlabeled vesicle-tubulin complexes there was no marked change in energy transfer efficiency upon addition of Ca” (not shown). The aggregated labeled complexes had presumably fused with each other and not with the unlabeled complexes. Other controls (not shown) which did not produce Ca*+dependent energy transfer changes include mixing vesicles with Ca” without tubulin and mixing vesicles with tubulin (without phase transition) and Ca”. Ca “-induced Fusion of Vesicle-Tubulin Complexes was Nonleaky-An important question about vesicle-vesicle fusion is whether the fusion process is accompanied by leakage of vesicle contents. This is the case with CaZf-induced fusion of phosphatidylserine vesicles (19). In order to assess leakage we formed vesicle-tubulin complexes by bringing DPPC vesicles containing 120 mM carboxyfluorescein to Tc in the presence of tubulin at a tubulin-vesicle ratio of 30~1. We then immediately rechromatographed the complexes on a Sephadex G-25 (PD-10) column to remove dye released during complex formation. All this was done in a buffer containing 1 mM EDTA. The ratio between the fluorescence of the rechromatographed complexes before and after disrupting them with Triton X-100 was 0.23. This is consistent with 30 mM carboxyfluorescein remaining in the vesicles (see Fig. 3 in reference 6). This indicates that about 70% carboxyfluorescein was released from the vesicles during complex formation. Fig. 6A shows that addition of Ca2+ at concentrations which induced efficient fusion (see Fig. 6B) did not cause leakage of carboxyfluorescein from those complexes. The average leakage, probably due to dilution of the complexes into the Ca’+ buffer, was 4.6 + 3.3% (dashed line in Fig. 6A). The leakage at the different Ca*’ concentrations was not significantly different from that background level. Further Characterization of Aggregation and Ca’+-induced Fusion-In the next set of experiments we tested whether the pelleting of lipid is also accompanied by a proportional amount of tubulin in the pellets. Results of one such experiment in which we used ‘H-DPPC vesicles and tyrosinolated [“‘Cltubulin are shown in Table III. There is no difference between tyrosinolated and detyrosinolated tubulins in their interaction with DPPC vesicles (see below). Tyrosinolated [‘4C]tubulin was prepared according to the method described elsewhere (13). This modification of tubulin involves addition of a tyrosine residue at the COOH terminus of the (Y subunit, catalyzed by tubulin tyrosine ligase in the presence of ATP. No significant differences in the in vitro microtubule assembly properties so far have been observed between maximally tyrosinolated and detyrosinolated tubulins (14). Tubulin alone under these conditions did not pellet, with or TABLE III Co-pelleting of[3H]DPPC and tyrosinolated [%]tubulin Tubulin to vesicle ratio was 9:l. Concentration of Mn2+ or Ca“ was 2.5 mM and that of EGTA was 5 mM. Tyrosinolated [‘4C]tubulin was prepared as described elsewhere (13). Radioactivity Condition
beas
was suggested by the pelleting experiments (see below). Mixing of vesicle-tubulin complexes at 44 “C produced a similar Ca*+ effect on energy transfer as that seen at room temperature (below transition). The probe-containing vesicle-tubulin complexes needed to
pellet/total Vesicles Vesicles + Ca*+ Vesicles + Ca2+ + EGTA Vesicle-tubulin + Ca’ Vesicle-tubulin + Mn2+
0.17 0.19 0.13 0.96 0.99
0.8 0.93
Ca2+-inducedFusion of Vesicle-Tubulin Complexes
15142
without Ca2+.There was no pelleting of the tyrosinolated ["C] tubulin if the tubulin was added to vesicles below Tc. However, tubulin did pellet with the lipid whenit was inserted into vesicles. When the vast majority of the lipid was pelleted in the presence of divalent cation, a comparable amount of tubulin accompanied the lipid. Results in Table IV show that actin, another cytoskeletal protein which interacts with DPPC vesicles, also mediated the Ca2+-induced sedimentation of vesicles. On the other hand, serum apolipoprotein Al, which also inserts into these vesicles, did not behave like tubulin and actin. Agents which affect polymerization of tubulin (polylysine, colchicine, and podophyllotoxin) did not have any significant effect onthe tubulin- mediated or Ca2'-enhanced aggregation and fusion of vesicles.MAPs reproducibly caused 10-15% inhibition of pelleting of vesicles (Table V). Bovine serum albumin and NaCl had no effect onthe vesicle sedimentation with or without added Ca2+. Brief treatment with trypsin abolished the Ca2+-inducedsedimentation of tubulin-containing vesicles (Table VI). This result suggests a role for the exposed portion of the tubulin molecule (i.e.that is not buried in the bilayer) in mediating the Ca2+effect. To testwhether the Ca2+effect reflected enhanced proteinprotein interaction or whether inserted tubulin could interact with pure lipid, we made vesicle-tubulin complexes with unlabeled lipid and examined the pelleting of 14C-labeled(protein-free DPPC) vesicles. Data in Table VI1 show that Ca2+induced pelleting of vesicles can take place between tubulincontaining and protein-free vesicles. However, it is necessary that tubulin be inserted in at least some participating vesicles. Finally we have examined whether vesicle-tubulin complexes interact with microtubules during polymerization of tubulin. Results shown in Table VI11 suggest that neither esicles nor tubulin-containing vesicles bindtightly to microtubb'os.Assembled microtubules were separated from the bulk of unassembled contents by centrifugation through a 50% sucrose cushion. In these experiments more than 60% of the tubulin was recovered in the microtubule pellet, and 10 PM colchicine resulted in 90% inhibition of this polymerization. Radioactive lipid in the pellets using either vesicles or vesicletubulin complexes was the same with or without colchicine and represented only about 6% of the total lipid. Further Characterization of Tubulin-Vesicle InteractionAssociation of tubulin with DPPC vesicles was studied as described earlier (6) by measuring the release of encapsulated dye (carboxyfluorescein) into the medium. The amounts of protein required to cause 50% (of total) release of the dye are shown in Table IX. Three assembly-cycle purified tubulin (3 X tubulin) which has approximately 80% tubulin and 20% MAPs was 9 to 10 times less effective than purified tubulin, and MAPs (up to3 mg/ml) were totally ineffective ininducing
TABLEV Further characterization of tubulin-mediated a n d Ca'+-induced aggregation/fusion of DPPC vesicles Tubulin to vesicle ratio was 101. Vesicle-tubulin complexes were incubated with various substances tested for 20 min before centrifugation to determine pelleting of vesicles. CaZi was added to a concentration of 2.5 mM. -CaC12 +CaCl, pellet/total
Vesicles Vesicle-tubulin Vesicle-tubulin + tubulin (0.4 to 20 p ~ ) Vesicle-tubulin + MAPs (0.75 mg/ml) 0.78 Vesicle-tubulin + Poly-L-lysine (10.87 mg/ml) Vesicle-tubulin + colchicine (100 p ~ ) Vesicle-tubulin + podophyllotoxin (100
Condition
Vesicles Vesicles + actin" Vesicle-actin Vesicle-actin + Ca'+ Vesicles + apolipoprotein Al" Vesicle-apolipoprotein AI Vesicle-apolipoprotein A1 + Ca2' "Added together at 30 "C (below Tc).
Radioactivity in the pellet W total
22 30
51 88 25 16
15
0.18 0.61 0.63
0.17 0.89 0.92
0.56 0.59 0.61 0.64
0.87 0.89
PM)
Vesicle-tubulin + bovine serum albumin 0.88 (1 mg/ml) Vesicle-tubulin + NaCl (1.25 M)
0.57 0.89
0.6
TABLE VI Trypsin treatment of vesicle-tubulin complexes abolishes Ca2+enhanced aggregation/fusion of vesicles Tubu1in:vesicle ratio was 1O:l. Trypsin concentration was 3 pg/ml. After 10 min at 30 "C,trypsin was inhibited by the addition of soybean trypsin inhibitor (9 pg/ml). Trypsin-treated or freshly prepared vesicle-tubulin complexes were further incubated in the presence of 5 mM CaCI2 for 30 min at 30 "C and processed for the determination of pelleting of vesicles. Soybean trypsin inhibitor alone had no effect. Condition
Radioactivity pellet/total 0.06
Vesicles Vesicle-tubulin Vesicle-tubulin 0.98 + Ca2' Trypsin-treated (vesicle-tubulin) Trypsin-treated (vesicle-tubulin) + Ca'+
.
TABLEIV Ca'+-induced aggregation/fusion of vesic1es:protein specificity Actin to vesicle ratio was 5 and that of apolipoprotein AI tovesicle was 3. Vesicle-protein complexes were prepared as for tubulin at the phase transitiontemperature (38-40 "C). Ca2' was added to a concentration of 2.5 mM.
Radioactivity (["Cllipid)
Condition
0.58 0.58 0.60
TABLEVI1 Aggregatwn/fusion can also take placebetween vesicle-tubulin and protein-free vesicles Tubulin to vesicle ratio was 1 0 1 and CaC12 concentration, 2.5 mM. In the sets in which radioactive and nonradioactive vesicles were combined, equal lipid concentrations for both were used. Incubations were done at 30 "C. Radioactivity in pellet
Condition +CaCla
-CaC12
76 of total
Vesicles" Vesicles" + vesicles Vesicle"-tubulin Vesicle-tubulin + vesicles" a
16 15 72
58
11 15
92 90
I4C-labeledvesicles.
dye -*elease.In fact, addition of MAPs to tubulin during phase transition release resulted in the partial inhibition of release (not shown). There was no difference in the ability of maximally tyrosinolated and detyrosinolated tubulins to cause release of the dye from DPPC vesicles. Tyrosinolated and detyrosinolated tubulins were prepared by incubating tubulin with tubulin tyrosine ligase and carboxypeptidase A as described (13). There was no significant effect of GTP and various anti-tubulin drugs such as colchicine and podophyllotoxin in the phase transition release assay. On the other hand, heat inactivation of the tubulin (90 min at 45 "C) dramatically destroyed phase transition release activity. Since there was no difference between tyrosinolated and detyrosinolated tubulins in their interaction with vesicles, we also
Ca2+-inducedFusion of Vesicle-TubulinComplexes TABLEVI11 Vesicle-tubulincomplexes do not associate with microtubules 3 x tubulin (1.5 mg/ml) was assembled for 45 min at 30 "C in the presence or absence of 10 pM colchicine in a total volume Of 300 pl. (c) and (d) contained 25% tubulinasvesicle-tubulincomplex (10 tubulinspervesicle).Polymerizedmicrotubules were collected by centrifugationof the assembled solution througha 50%sucrose cushion (Beckman rotor 65 at 50,000 rpm for 2 h, 25-30 "c). Pellets were redissolved in 0.1 N NaOH and analyzed for total protein and radioactivity. In (c) to (f) total "C counts(['4C]DPPCvesicles)added during polymerization were7300. Pellet Assembly condition
Protein
"C-lipid % total
(a) 3 X tubulin
(b) (a) + 10 ~ L colchicine M (c) 3 X tubulin + vesicle-tubulincomplexes (10%v/v) (d) (c) + 10 PM colchicine (e) (a) + vesicles (lo%,v/v) ( f ) Vesicles alone
61 8 60
9 62
6.6 6.5 4.9 4.0
TABLEIX Concentration to cause ~ ( W Oof dye release (pg/ml) Release refers to phase transition release as described in reference 6 using 30 ~ L DPPC M lipid and 100 mM entrapped carboxyfluorescein. Values are mean f S.D. of 3 to 4 experiments. Tubulin-drug complexes were prepared by incubating tubulinwith 2-fold molar excess of drugs at 30 "C for 60 min. Free (unbound) drug was removed by gel chromatography on a Sephadex G-50 column. 11 f 5 Tubulin" 105 f 20 3 X tubulin No release MAPS Tubulin-colchicine complex 12 f 4 Tubulin-podophyllotoxin complex 10 Similar concentration range was observed for maximally tyrosinolated (45%tyrosine) and detyrosinolated(0-2% tyrosine) tubulins. 'I
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as described (17,18). Aliquots 3 x tubulin and heat treatment of tubulinor vesicle-tubulincomplexes (approximately 10 tubu1in:vesicle) were incubated at 30 "C for 60 minwith varying amounts of ["2P]MAPz and 0.5 mM GTP. Tubeswere centrifugedin a n Airfuge (20 p.s.i.,30 min a t 25 "C) and supernatants and pellets were analyzed for radioactivity and protein. The amount of tubulin inpelletedvesicle-tubulin complexes was estimated on thebasis of results described in Table 111. Under these conditions, microtubules had bound approximately 1 mol of MAP2 per 9 mol of tubulin, a value similar to the one reportedelsewhere (14, 19). DISCUSSION
The studies reported in this paper show that the insertion of tubulin into lipid bilayers of uncharged sonicated small unilamellar vesicles results inaggregation of the vesicles, measured by light scattering and increased pelleting of the vesicles. Simply mixing theproteinand vesicles was not sufficient to cause the aggregation of vesicles. In thepresence of millimolar Ca'+ or Mnz+ but not Mg2+, there was nearly complete pelleting of the vesicles. The effects of Ca'+ on the DPPC vesicle-tubulin complexes could be related to the fact that tubulin is a Ca2+-binding protein(20). Data in Table V suggest that protein-protein interaction during aggregation of vesicles does not involve polymerization of tubulin as judged by the lack of effect of polymerization inhibitors and that vesicle-tubulin complexes do notassociate with microtubules. Results in Table VIII,however, do not rule out the possibility of dissociation of weakly bound vesicles from microtubules during centrifugation. Actin, another componentof the cytoskeletal system when incubated with lipid vesicles and passed through the phase transition also promoted increased pelleting of vesicles, which was enhanced by Cap+.We have recently observed that clathrin, a component of coated vesicles can produce fusion of dioleoyl phosphatidylcholine vesicles at pH below 6.:' Apolipoproteins,ontheotherhand, while they formvesicular recombinants with DPPC by passage through the phase transition, do not promote increased pelleting. Based onpelleting and light-scattering data,it is notpossible to differentiate between aggregation and fusion of vesicles. Electron microscopy, however, shows that inserted tubulin results in the aggregation of vesicles, and only the additionof divalent cations induces membrane fusion. Although these results with electronmicroscopy were clearand reproducible, a number of authors have pointed out a variety of artifacts that can be produced by negative staining of phospholipid preparations (see for example reference 21); and itis perhaps warranted to discuss the question of the reliability of our results in more detail. The results are internally consistent and showcleardifferences betweenthe controls: vesicles alone, vesicles with Ca2+, and tubulin-vesicle complexes withwhich did not fuse. If the final fusionof vesicles were out ea2+ caused by phosphotungstic acid it would have been a phosphotungstic acid-induced fusion dependent upon the insertion of tubulin into the DPPC vesicles and requiring calcium. In our protocol the effects of the stainon aggregation and fusion of the vesicles were minimized by applying the vesicles or vesicle-tubulin complexes to grids and blotting orwashing off the excess fluid before applying the stain. This should have left primarily vesicles that were adsorbed to the substrate to be outlined by thestain. (Washing the gridsbearing the vesicles or complexes with water before application of the stain orrinsing away floating vesicles in a stream of stain gave essentially the same results as those shown in Fig. 4). On the
tested if there would be differences in the ability of these tubulins to act as substrates for the enzymes which cause these modifications. In several experiments, we did not see any differences between the abilityof free and vesicle-bound tyrosinolated tubulin to be detyrosinolated by purified carboxypeptidase (13) or in the ability of detyrosinolated tubulin to be tyrosinolated by tubulin-tyrosine ligase (15). Vesicle-bound tubulin was equally active in binding colchicine, and the kineticsof binding of [3H]colchicine to free and vesicle bound tubulinwas identical. [3H]Colchicine binding to tubulin was tested according to the proceduredescribed elsewhere (16). Aliquots of tubulin or vesicle-tubulin complexes were incubatedin 150 pl of abuffer containing 10 mM NaH2P04, 5.0 mM MgS04, 0.24 M sucrose, and 0.1 mM GTP, pH 6.9, and 1.2 pCi/ml of ['H]colchicine (final concentration 12 p ~ at) 30 "C for 15,30,60,120, and 180 min. At each time point a 100-pl aliquot was transferred onto a DE-23 column (0.5 ml in Bio-Rad econo-column 731-1110), pre-equilibrated with 15 ml of the same buffer but without sucrose and GTP. The columnswere washed immediately with1.5 ml of buffer. Tubulin was retained in the column. The column contents were quantitatively transferred intoscintillation vials. Radioactivity was measured using Aquasol (New England Nuclear). Nonspecific binding assessed by incubating as above but in the presence of 50-fold excess of nonradioactive colchicine was less than 2-3%of total binding. A similarly low value was obtained when [3H]colchicine alone (no tubulin) was identic d y incubated andprocessed. Tubulin in the form of vesicletubulin complexes bound a significant amount of [32p] MAP2 (up to approximately 1 mol of MAPz per 15 mol of R. Blumenthal, M. Henkart, and C. Steer, unpublished observatubulin). [:'*PIMAP2 was prepared by autophosphorylation of tions.
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Ca2+-inducedFusion of Vesicle-Tubulin Complexes
other hand, when DPPC vesicles were mixed with phosphotungstic acid in suspension, we observed stacking of the vesicles as described by Melchior et al. (21). Uranyl acetate which is generally considered the negative stain of choice for studies of microtubules was not useful for this study because the divalent cation UOZ2+is known in many systemsto mimic the effects of CaZt which was the object of these experiments. In fact, with uranyl acetate staining there was extensive reorganization of the tubulin-vesicle complexes with or without calcium while the DPPCvesicles alone or with calcium resembled those stained with phosphotungstic acid. The electron microscope shows morphologic evidence of fusion but does not allow us to distinguish between the fusion of small vesicles and the somewhat less likely possibility that thesmall vesicles disintegrate andthen form large vesicles. The increased trapped volume (see Table 11) and theconservation of encapsulated carboxfluorescein upon addition of Ca2+to vesicletubulin complexes (see Fig. 6A),however, indicate that fusion is the most likely mechanism for the formation of the large structures. Additional evidence for Ca2+-inducedfusion of DPPC vesicle-tubulin complexes was provided by the lipid mixing assay developed by Pagano and his coworkers (11).Efficient energy transfer was observed between two fluorescent lipid analogues incorporated into the same vesicle containing one of each probe molecule per 100 phospholipid molecules. Upon fusion of these vesicles with a second vesicle population containing no fluorescent lipid, the efficiency of energy transfer was reduced since lateral diffusion of the probes in the plane of the newly formed larger membrane effectively lowers its surface density. In theabsence of added Ca2+the vesicle-tubulin complexes aggregated but did not fuse, and we did not see any changes in energy transfer efficiency. However, upon addition of CaZ+we observed marked changes in energy transfer. This observation provides a built-in control that aggregation of vesicles does not change the energy transfer pattern. We could not use the resonance energy transfer assay developed by Wilschut et al. (22) based on mixing of intravesicular compartments during fusion; one of the probes used in that assay bound too strongly to tubulin, interfering with this assay. The energy transfer assaystrengthens the interpretation of our data in terms of fusion. Both gel phase and fluid phase (at 44 "C) DPPC vesicles exhibit tubulin-mediated Ca2'-dependent fusion. It it interesting that tubulin, which has been inserted into vesicles, is capable of inducing fusion with pure lipid vesicles. This suggests that the tubulin is altered by its insertion such that the exposed region is more capable of Caz+-dependent perturbationof another lipid bilayer. Such an alteration is consistent with our previous findings of an overall change in the conformation of the molecule upon insertion into thelipid bilayer. In the majority of earlier studies (22) the effect of divalent cations, mainly Ca", in inducing aggregation and fusion of vesicles has been studied with vesicles made of negatively charged phosphatidylserine or of mixed phospholipids (phosphatidylserine, phosphatidylcholine, and phosphatidylethanolamine). Small unilamellar DPPC vesicles have been reported to fuse (in the absence of Ca2+)to vesicles about 70 nm
in diameter, when held below Tc (23).This spontaneous fusion does not produce the large structures seen in Fig. 4 0 . We have earlier reported that the presence of Ca2+in the medium did not affect the insertion of tubulin into DPPC vesicles at the phase transition (6). In a recent paper,Hong et al. (24) have presented evidence for the enhancement of Caz+-dependent fusion of vesicles made of phosphatidylserine and phosphatidylethanolamine (1:3) by synexin (M,= 47,000), a water-soluble protein isolated from the adrenal medulla. In another study Zimmerberg et al. (25) observed fusion of phospholipid multilamellar vesicles with a planar phospholipid bilayer membrane that contained a water-soluble Ca2+-bindingprotein (Mr = 16,000) purified from calf brain. This is, however, the first example of nonleaky fusion of phosphatidylcholine vesicles mediated by a protein and induced by Ca2+. Acknowledgments-We thank Drs. E. D. Korn and S. J. Morris for valuable suggestions and helpful discussions and Karen Marconi for skillful secretarial assistance. REFERENCES 1. Zisapel, N., Levi, M., and Gozes, I. (1980). J. Neurochem. 34,2632 2. Bhattacharyya, B., and Wolff, J. (1975) J. Biol. Chem. 250, 76397646 3. Walters, B. B., and Matus, A. I. (1975) Nature (Lond.)257,496498 4. Soifer, D., and Czosnek, H. (1980) in Microtubules and Microtubule Inhibitors (DeBrabander, M., and DeMey, J., eds) pp. 429-447, Elsevier/North-Holland Biomedical Press, Amsterdam 5. Soifer, D., and Czosnek, H. (1980). J. Neurochem. 35, 1128-1136 6. Klausner, R. D., Kumar, N., Weinstein, J. N., Blumenthal, R., and Flavin, M. (1981) J. Bwl. Chem. 256,5879-5885 7. Kumar, N., Klausner, R. D., Weinstein, J. N., Blumenthal, R., and Flavin, M. (1981) J. BioZ. Chem. 256, 5886-5889 8. Caron, J. M., and Berlin, R. D. (1979) J . Cell Biol. 81, 665-671 9. Asnes, C. F., and Wilson, L. (1979) Anal. Biochem. 98.64-73 10. Lowry, 0. H., Rosebrough, N. J., Farr, A. L., and Randall, R. J. (1951) J. Biol. Chem. 193, 265-275 11. Struck, D. K., Hoekstra, D., and Pagano, R. E. (1981) Biochemistry 20,4093-4099 12. Fung, B. K.-K.,and Stryer, L. (1978) Biochemistry, 17,5241-5248 13. Kumar, N., and Flavin, M. (1981) J. Biol. Chem. 256, 7678-7686 14. Kumar, N., and Flavin, M. (1982) Eur. J. Biochem., in press 15. Kohayashi, T., and Flavin, M. (1978) J. Cell Biol. 79,285a 16. Garland, D., and Teller, D. C. (1975) Ann. N . Y. Acad. Sci. 253, 232-238 17. Sloboda, R. D.,Rudolph, S . A,, Rosenbaum, J. L., and Greengard, P. (1975) Proc. Natl. Acad.Sci. U. S. A . 72, 177-181 18. Vallee, R. (1980) Proc. Natl. Acad. Sci. U. S. A . 77, 3206-3210 19. Kim, H., Binder, K. L., and Rosenbaum, J. L. (1979) J. Cell Biol. 80, 266-276 20. Solomon, F. (1977) Biochemistry 16, 358-363 21. Melchior, V., Hollingshead, C. J., and Cahoon, M. E. (1980) J . Cell Biol. 86, 881-884 22. Wilschut, J., Duzgunes, N., Fraley, R., and Papahadjopoulos, D. (1980) Biochemistry 19,6011-6021 23. Schullery, S. E., Schmidt, C. F., Felgner, P., Tillack, T. W., and Thompson, T. E. (1980) Biochemistry 19, 3919-3923 24. Hong, K., Duzgunes, N., and Papahadjopoulos, D. (1981) J. B i d . Chem. 256,3641-3644 25. Zimmerberg, J., Cohen, F. S., and Finkelstein. A. (1980) Science 210,906-908
