Determinants for Membrane Fusion Induced by ... - ACS Publications

8264

J. Phys. Chem. B 2008, 112, 8264–8274

Determinants for Membrane Fusion Induced by Cholesterol-Modified DNA Zippers Gudrun Stengel,*,† Lisa Simonsson,† Richard A. Campbell,‡ and Fredrik Ho¨o¨k*,† Department of Solid State Physics, UniVersity of Lund, 22100 Lund, Sweden, and Department of Physical Chemistry, GetingeVa¨gen 60, Box 124, 22100 Lund, Sweden ReceiVed: March 6, 2008

Intracellular membrane fusion is coordinated by membrane-anchored fusion proteins. The cytosolic domains of these proteins form a specific complex that pulls the membranes into close proximity. Although some results indicate that membrane merger can be accomplished solely on the basis of proximity, others emphasize the importance of bilayer stress exerted by transmembrane peptides. In a reductionist approach, we recently introduced a fusion machinery built from cholesterol-modified DNA zippers to mimic fusion protein function. Aiming to further optimize DNA-mediated fusion, we varied in this work length and number of DNA strands and used either one or two cholesterol groups for membrane anchoring of DNA. The results reveal that the use of two cholesterol anchors is essential to prevent cDNA strands from shuttling to the same membrane, which leads to vesicle release instead of membrane merger. A surface coverage of 6-13 DNA strands was a precondition for efficient fusion, whereas fusion was insensitive to DNA length within the tested range. Besides lipid mixing, we also demonstrate DNA-induced content mixing of large unilamellar vesicles composed of the most abundant cellular lipids phosphatidylcholine, phosphatidylethanolamine, cholesterol, and sphingomyelin. Taken together, DNA-mediated fusion emerges as a promising tool for the functionalization of artificial and biological membranes and may help to dissect the functional role of fusion proteins. Introduction Membrane fusion is essential for eukaryotic cell function and is coordinated by dedicated fusion proteins.1,2 Neuronal exocytosis requires docking of synaptic vesicles with the presynaptic plasma membrane and subsequent fusion of both membranes. Three members of the soluble N-ethyl-maleimide-sensitivefactor attachment protein (SNARE) family play a crucial role in this process: synaptobrevin (SB), located on the vesicle membrane, forms a membrane-bridging complex with syntaxin 1A (SX) and SNAP-25, the latter two both anchored in the plasma membrane.1,2 In 1998, Rothman and co-workers demonstrated that these three proteins were sufficient to catalyze lipid mixing of POPC/DOPS vesicles in vitro.3 Since then, it has become widely accepted that these proteins constitute nature’s minimal fusion machinery, albeit additional proteins are likely to be involved in vivo.2 SNARE proteins are expected to influence all stages of fusion, which in the current view are considered to be (a) membrane contact, (b) stalk formation/ hemifusion, (c) trans-monolayer contact (TMC), and (d) fusion pore formation.1,4 In the stalk intermediate, only the outer lipid leaflet has merged. The stalk can either revert to the original bilayer structure or change into a TMC when the cis monolayers expand radially, thereby bringing the trans monolayers into contact. A diaphragm is then formed, which contains only lipids of the trans monolayers. Fusion is complete when the diaphragm ruptures and a fusion pore is created. A popular strategy for the investigation of the mechanisms underlying fusion is the reconstitution of fusion-associated proteins in simple model membranes to test their ability to mediate lipid mixing.3,5 To single out the important role lipid * Corresponding authors. E-mail: [email protected] and fredrik.hook@ chalmers.se. Phone: +46-31-7726109. Fax: +46-31-7723134. † Department of Solid State Physics. ‡ Department of Physical Chemistry.

composition plays in membrane fusion,6,7 other research groups concentrate on protein-free fusion as induced by chemicals like polyethylene glycol (PEG) and bivalent ions.8–10 In some cases, alterations of the nature of the lipids alone are sufficient to induce fusion. For instance, adhesion and fusion were observed as a result of strong electrostatic interactions between oppositely charged membranes11 and between vesicles which the membrane curvature was altered by chemical cleavage of the lipid headgroup.12 Although valuable mechanistic insights were derived from such systems,8 it is important to bear in mind that methods that drive fusion by a global change in hydration state, osmotic pressure, or lipid line tension ignore the site-specific nature of fusion and expose membranes to nonphysiological conditions. As the field progresses, focus has shifted to the rational design of membrane-bound receptors that render membranes fusogenic by means of surface recognition.13,14 However, few groups have reported complete fusion of both lipid leaflets by using artificial receptors. Frequently, fusion stalled at the hemifused state, which results from the merger of the outer lipid leaflets while the inner leaflets remain separated. An example is the hemifusion observed for giant liposomes modified with monomeric thymidine and adenosine-lipid conjugates15 and for various truncated fusion proteins the transmembrane domains of which were replaced by lipid anchors.16,17 The latter result gave rise to the conclusion that the transmembrane domain of fusion proteins plays an active role in fusion and is thus indispensible. However, the community is still divided on this point, given the existing evidence for complete fusion in protein-free systems. Two examples are the membrane activiation by lipid-linked bipyridine in the presence of metalions18 or by magainin and vancomycin as membrane-anchored recognition motif.19 Recently, we introduced an artificial membrane fusion machinery consisting of DNA that hybridizes in a zipperlike fashion and is membrane-anchored via cholesterol (CH),20 see

10.1021/jp802005b CCC: $40.75  2008 American Chemical Society Published on Web 06/21/2008

Membrane Fusion Induced by CH-Modified DNA Zippers

J. Phys. Chem. B, Vol. 112, No. 28, 2008 8265

SCHEME 1: DNA-Mediated Membrane Fusiona

a (Top) Sequential hybridization of double CH-anchored DNA leading to merger of vesicle contents. (Bottom) Nucleobase sequence and nomenclature of DNA constructs.

SCHEME 2: Chemical Structure of Single (A) or Double (B) CH-Modified DNA

protein function and hence to broaden the scope of recognitionguided membrane fusion. Initially, the common motivation for synthesizing lipophilic DNA was to enhance the cellular uptake of antisense DNA and RNA by lipidation.23 As a result, numerous synthetic strategies have been developed to couple DNA to alkyl-chains, CH and derivatives, alkylated glycerols, fullerenes, and adamantane, offering a large pool of membrane anchors with a wide range of properties.24–31 Here, we focused on the design of fusogenic DNA zippers from CH-modified DNA. By using established fluorescence assays, we investigate DNA-induced lipid and content mixing of large unilamellar vesicles (LUVs) composed of either DOPC/DOPE/CH or DOPC/DOPE/SM/CH (DOPC ) 1, 2-Dioleyl-sn-glycero-3-phosphocholine, DOPE ) 1, 2-Dioleyl-sn-glycero-3-phosphoethanolamine, and SM ) sphingomyelin from bovine brain). We assess how parameters such as length of the DNA strands, anchoring strategy, and number of DNA strands affect the DNA’s ability to induce membrane fusion and discuss possible mechanistic parallels to proteininduced fusion. Materials and Methods

Schemes 1 and 2. The overall architecture of the CH-DNA aims at minimizing the distance between two DNA-bridged bilayers and is vaguely reminiscent of the SNARE core complex, where the water soluble N-terminal portions of SB, SX, and SNARE25 form a four-helix bundle that pins the bilayers together.21 The formation of this helix bundle starts at the N-terminus and proceeds toward the C-terminus in a zipperlike fashion.22 The associated coiled-coil transition is believed to make a free energy contribution to the fusion reaction. In a similar way, DNA serves to pin two bilayers together and undergoes an unstructured-helix transition upon hybridization that is energetically favorable. The goal of this work is to mimic SNARE

Material. CH and 1,2-dioleoyl-sn-glycero-3-phospho-Lserine-N-(7-nitro-2-1,3-benzoxadiazol-4-yl) diammonium salt (NBD-PS), 1,2-dioleyl-sn-glycero-3-phosphocholine (DOPC), and 1,2-dioleyl-sn-glycero-3-phosphoethanolamine (DOPE) were purchased from Avanti Lipids Inc., Birmingham. SM was obtained from Sigma Aldrich. Bodipy500/510-C5-HPC 2-(4,4difluoro-5-octyl-4-bora-3a,4a-diaza-s-indacene-3-pentanoyl)-1hexadecanoyl-sn-glycero-3-phosphocholine (Bodipy500/510), Bodipy530/550-C5-HPC 2-(4,4-difluoro-5,7-diphenyl-4-bora3a,4a-diaza-s-indacene-pentanoyl)-1-hexadecanoyl-sn-glycero3-phosphocholine (Bodipy530/550), and lissamine-rhodamine B 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine (rhodamine-DHPE) were purchased from Molecular Probes, Eugene. Oligonucleotides were purchased from Eurogentec S.A., Bel-

8266 J. Phys. Chem. B, Vol. 112, No. 28, 2008 gium, see Scheme 1 for base sequences. Terbium (III) chloride, sodium citrate, dipicolinic acid (DPA), N-[tris(hydroxymethyl)methyl]-2-aminoethanesulfonic acid (TES), tris(hydroxymethyl)aminomethane hydrochloride (TRIS), and L-histidine (L-his) were from Sigma Aldrich. Vesicle Preparation. Vesicles were composed of DOPC/ DOPE/CH (50:25:25 mass ratio) or DOPC/DOPE/SM/CH (35: 30:15:20 mass ratio). Where required, vesicles were either labeled with 0.3% Bodipy500/510 and 0.3% (Bodipy530/550) or with 1.5% NBD-PS and 1.5% Rh-DHPE as FRET pairs. Chloroform was removed from the lipid mixtures in a nitrogen stream (1 h), and dry lipids were hydrated in buffer (60 min, 5 mg/mL, 10 mM Tris/HCl pH 8.0, 100 mM NaCl, 1 mM EDTA). Extrusion was performed by using the Avanti Lipid MiniExtruder equipped with 100 nm polycarbonate membranes (Whatman). Vesicles were never used for more than one day. The DNA strands ss-dc1 and ss-dc2 did not self-incorporate into the vesicles when added to the aqueous solution. Instead, DNA was added to the lipid solution in chloroform prior to drying of lipids at a lipid-to-DNA molar ratio of 5260:1. Extrusion was carried out as described above. Accordingly, DNA is presumed to be present in the inner and outer leaflet in this case. Reduction of NBD-PS-Labeled Vesicles. A 1:1 mixture of NBD-PS/Rh-DHPE-labeled vesicles (10 mg/mL) and 200 mM sodium dithionite (in 10 mM Tris/HCl pH 10.0, 100 mM NaCl) were mixed at room temperature and incubated for 5 min. Free sodium dithionite was removed by gel filtration by using Microspin S-200 HR columns from Amersham Biosciences. Reduction was carried out immediately before use of the vesicles. Lipid Mixing Assays. DNA-modified vesicles containing dyes were mixed with unlabeled DNA-modified vesicles at a ratio of 1:4; the total lipid concentration was 275 µM. Before mixing, both vesicle populations were functionalized with cDNA at a molar lipid-to-DNA ratio of 1316:1 by simply mixing the aqueous vesicle and DNA solutions. Self-incorporation of the DNA strands was allowed to take place for at least 30 min prior to use. If not stated otherwise, the reaction buffer was 10 mM Tris/HCl pH 7.4, 100 mM NaCl, 1 mM EDTA, and the temperature was 30 °C. The change in donor intensity is plotted as ID(%) ) 100 × (It - I0)/(Itotal - I0), with I0 being the donor intensity at t ) 0 before lipid mixing and Itotal the donor intensity after disruption of the vesicles in 0.8% (w/v) Triton X-100. Content Mixing Assay. The Tb/DPA content mixing and leakage assays were based on assays previously described by Wilschut et al.10 Dried lipids were hydrated in buffer A, B, or C overnight (at 4 °C), final concentration 10 mg/mL. Buffer A: 2 mM TES/L-his, 15 mM TbCl3, 150 mM sodium citrate, pH 7.4. Buffer B: 2 mM TES/L-his, 150 mM DPA, pH 7.4. Buffer C: 2 mM TES/L-his, 7.5 mM TbCl3, 75 mM sodium citrate, 75 mM DPA, pH 7.4. Extrusion was carried out as described above. Non-encapsulated material was removed by using Microspin S-200 HR columns (Amersham Biosciences) equilibrated in 3.7 mM TES/L-his, 185 mM NaCl, 2 mM EDTA, pH 7.4. Content mixing was initiated by mixing equal amounts of DNA-loaded vesicles encapsulating Tb (buffer A) and DPA (buffer B) at a final lipid concentration of 219 µM in reaction buffer (3.7 mM TES, 3.7 mM L-his, 185 mM NaCl, 2 mM EDTA, 20 mM CaCl2, pH 7.4). Content mixing was quantified by relating the fluorescence change, ∆FCM, measured upon mixing of Tb and DPA vesicles to the fluorescence intensity observed for vesicles encapsulating Tb/DPA complex, ∆Fmax: percent content mixing ) ∆FCM/0.5∆Fmax. Vesicle leakage was examined by reacting

Stengel et al. vesicles containing Tb/DPA complex with vesicles encapsulating 3.7 mM TES/L-his, 185 mM NaCl, pH 7.4. Tb/DPA complex leaking out of vesicles is quenched by the surrounding reaction buffer, which leads to an intensity decrease, ∆FL. The amount of leakage is calculated as percent leakage ) (∆Fmax - ∆FL)/ ∆Fmax. Please note that it is not straightforward to compare the total amounts measured for content and total lipid mixing because the assays are measured on different scales. Lipid mixing is scaled relative to the maximal possible donor intensity, which corresponds to the donor emission in the absence of the acceptor (or at infinitely large vesicle size). Because of the limited amount of DNA strands available for hybridization, the size of the vesicle aggregates must be small, and correspondingly, lipid mixing always remains far below 100%. Lipid mixing occurs also in complexes the lipid bilayers of which join transiently but do not fuse. In contrast, content mixing is scaled relative to the fluorescence intensity that would result from the quantitative one-to-one fusion of Tb with DPA loaded vesicles and is not sensitive to several rounds of fusion. Fluorimeter Settings. Experiments were carried out by using a QM-4/2005 spectrofluorimeter (Photon Technology International Inc., Birmingham, NJ) by using a 3 mL quartz cuvette. Bodipy500/510 donor was excited at 485 nm, and the emission was recorded at 517 nm (all slits 2 nm). The NBD donor was excited at 460 nm, whereas the emission monochromator was set to 525 nm (excitation slits 1 nm, emission slits 10 nm). The Tb/DPA complex was excited at 276 nm, and its emission was collected at 543 nm (excitation slits 2 nm, emission slits 4 nm). In all cases, a 500 nm long pass cutoff filter was used to reject scattered light (ThorLabs). Ellipsometry Measurements. Silicon dioxide surfaces were coated with a 1:1 mixture of PLL(20)-g[3.5]-PEG (2) and PLL(20)-g[3.5]-PEG (2)/PEG(3.4)-Biotin (50%) from SuSos AG, Switzerland (total concentration 10 µg/mL). A 2:1 complex of biotin-DNA (5′-CAC-TAT-ATG-TTC-GTT-CCC-biotin-3′, 0.6 µM) and neutravidin (0.3 µM) was formed prior to injection onto the PLL-PEG surface. For surface attachment, vesicle species 1 was functionalized with 3′-CH-CCC-AGG-CAGCAC-GGA-GTG ATA TAC AAG CAA-5′ and 5′-CH-CCCTCC-GTC-GTG-CCT-3′ at a DNA-to-vesicle ratio of 3:1 in addition to fusion inducing DNA zippers. Measurements were performed on a Rudolph Research thinfilm null ellipsometer (type 43603-200E) to determine the adsorbed amount of tethered and fused vesicles. The light source was a xenon arc lamp fitted with a wavelength filter of 401.5 nm. The angle of incidence was 68°. The measurements were made in situ with 5 mL of solution agitated with a magnetic stirrer bar at ∼100 rpm in a thermostatted trapezoidal cuvette. The optical properties of silicon wafers with a 30 nm native silicon dioxide layer (thermally oxidized at 920 °C for ∼1 h) were characterized numerically from the amplitude and phase changes of the light upon reflection (Ψ and ∆, respectively) in air and water according to the method of Tiberg and Landgren.32 The parameters Ψ and ∆ were acquired for the in situ experiment at a rate of 0.5 s-1 and interpreted with respect to a five-layer homogeneous, stratified interfacial model: silicon/ silicon dioxide/first adsorbed layer/second adsorbed layer/ solution. The first adsorbed layer comprised biotinylated PLLg[3.6]-PEG with neutravidin and DNA. The second adsorbed layer comprised tethered and fused vesicles. It was helpful to model the two layers separately so that variations in the coverage and conformation of the vesicles did not inflict unrealistic changes on the density of the supporting architecture. Charac-



Figure 1. DNA-mediated total lipid (A) and inner leaflet mixing (B) of DOPC/DOPE/CH vesicles. NBD/rhodamine-labeled vesicles were mixed with the 4-fold amount of unlabeled vesicles. Prior to this, vesicles had been modified with different DNA constructs at a molar vesicle-to-DNA ratio of 1:100 (lipid-to-DNA ratio 1316:1). Color coding: ds-2/3 + ds-1/4 vesicles (black), ds-mm2/3 + ds-mm1/4 (red), ss-1 + ss-2 (blue), and ss-dc1 + ss-dc-2 (orange). Vesicles carrying non-cDNA (open black circles) were monitored as controls. To test the orientation of DNA incorporation in cases where DNA was mixed with lipids in organic solvent, NBD/rhodamine-labeled ds-dc1 vesicles were incubated with 20 nM free ss-dc2 to block the DNA binding sites. Addition of unlabeled vesicles coated with ds-dc2 did not result in lipid mixing (green line, vesicle addition at arrow). Note that the final lipid mixing induced by ds-2/3 + ds-1/4 varied slightly between 35 and 40% for different vesicle batches. If it was not possible to measure all DNA constructs by using the same vesicle batch, we measured ds-2/3 + ds-1/4 as internal standard (rel lipid mixing ) 1) and scaled all other curves recorded with the same batch relative to it.

terization of the two adsorbed layers was carried out sequentially. The thickness, d, and refractive index, n, of each layer were fitted numerically to the measured changes in Ψ and ∆,32 and then, the adsorbed amount, Γ, was calculated by using the de Feiters expression, Γ ) (n - nS)d/(dn/dc), where nS is the refractive index of the solution, and dn/dc the refractive index increment of the adsorbed layer. The value dn/dc ) 0.192 cm3/g was used for layer 1 as a weighted average of the values of PLL-g[3.6]-PEG (0.139 cm3/ g)33 and neutravidin (0.212 cm3/g),34 calculated proportionally from the experimentally determined adsorbed amounts. Reasonable variations in dn/dc for layer 1 had a negligible effect on the adsorbed amount of vesicles in layer 2. The value dn/dc ) 0.148 cm3/g was used for layer 2, taken from a literature value of DOPC35 as the major constituent of the lipid mixture. The relative adsorbed amounts of vesicles from the four experiments, and hence our entire interpretation of the data, is not affected by the error in this chosen value, because Γ is inversely proportional to dn/dc. Results Effect of Anchoring Strategy on Fusion. The objective of this work was to design a DNA zipper with improved fusion properties compared to those of the previously presented DNA construct.20 In the previous fusion study, we employed a DNA zipper consisting of a 12 base pair duplex with 15 overhanging bases as depicted in Scheme 1 (cf. double strand (ds)-2/3 and ds-1/4). The blunt end of the duplex was modified with two CH groups (one per DNA strand) that insert into lipid bilayers and act as DNA anchors. The choice of the double CH anchor was motivated by the observation that the dissociation of double CH-DNA from membranes is several orders of magnitude slower than that of single CH-DNA.36 On the basis of the complementarity of single strand (ss)-1 + ss-2 (and ss-3 + ss4) and the resulting net gain in free energy, the double-stranded regions are expected to melt as hybridization between vesicles carrying ds-2/3 and ds-1/4 occurs (see Scheme 1 for the time course of melting). If formation of the blunt-ended 12 and 27 base pair duplexes disrupts the association of the two CH groups, the membrane attachment of DNA is expected to be destabilized. As recently also addressed by Chan et al,59 further concern is

that melting of the double-stranded region is initially energy expensive and therefore constitutes an obstacle to membrane fusion. We were interested in finding out whether (a) single CH-anchored DNA mediates fusion (modification of different vesicles with ss-1 and ss-2, respectively) despite less stable membrane attachment, (b) a double CH-labeled single strand is a more efficient fusogen (ss-dc1 + ss-dc2) than ds-2/3 + ds-1/4 because the need for duplex melting is omitted, and (c) the introduction of three nucleobase mismatches adjacent to CH hampers membrane fusion based on associated changes in membrane spacing and linker flexibility (ds-mm2/mm3 + dsmm1/mm4). I. Lipid Mixing of DOPC/DOPE/CH Vesicles. First, we monitored lipid mixing induced by different DNA constructs by using 100 nm LUVs consisting of DOPC/DOPE/CH or DOPC/DOPE/SM/CH. Previously, these lipid compositions were shown to be most potent in both DNA- and PEG-mediated fusion.20,37 For DOPC/DOPE/CH vesicles, lipid mixing was probed by using the conventional NBD-PS/rhodamine-DHPE dequenching FRET assay, which, in conjunction with sodium dithionite treatment, enabled us to monitor total and inner leaflet mixing separately.38,39 However, labeling of DOPC/DOPE/SM/ CH vesicles with these dyes abolished lipid mixing completely. The inhibitory effect of headgroup labeled lipids on fusion, such as NBD and rhodamine, has been reported elsewhere and agrees with our observation.39 Instead, we probed lipid mixing of DOPC/DOPE/SM/CH vesicles by using low concentrations (0.3%) of Bodipy500/510 and Bodipy530/550 dyes covalently linked to the lipid’s fatty acid chain.39 All DNA constructs described here, except for ss-dc1 + ssdc2 (see Materials and Methods), were found to self-incorporate into lipid bilayers simply by mixing the aqueous vesicle and DNA solutions. In our standard fusion assays, dye-labeled and unlabeled vesicles were functionalized with cDNA at a molar vesicle-to-DNA ratio of 1:100 (lipid-to-DNA ratio 1316:1) after vesicle extrusion. Lipid mixing was initiated by addition of a 4-fold excess of unlabeled vesicles that had been preloaded with cDNA. For DOPC/DOPE/CH vesicles, ∼35% total lipid mixing was observed after 30 min for all DNA pairs exhibiting double CH anchors (ds-2/3 + ds-1/4 (black), ss-dc1 + ss-dc2 (orange), and

8268 J. Phys. Chem. B, Vol. 112, No. 28, 2008

Stengel et al.

TABLE 1: Rate Constants for DNA-Induced Lipid and Content Mixing of DOPC/DOPE/CH Vesicles k1 (10-3 s-1)

A1 ds-2/3 + ds-1/4 ds-mm2/3 + ds-mm1/4 ds-dc1+ ds-dc2 ss-1 + ss-2

0.482 ( 0.001 0.525 ( 0.001 0.480 ( 0.005

ds-2/3 + ds-1/4 ds-mm2/3 + ds-mm1/4 ds-dc1 + ds-dc2 ss-1 + ss-2

0.256 ( 0.003 0.237 ( 0.002 0.281 ( 0.009

ds-2/3 + ds-1/4 ds-mm2/3 + ds-mm1/4 ss-1 + ss-2

8.3 ( 0.2 6.2 ( 0.1 5.4 ( 0.1

Total Lipid Mixing 1.01 ( 0.01 1.04 ( 0.003 2.96 ( 0.02 Inner Leaflet Mixing 1.61 ( 0.02 1.04 ( 0.02 2.18 ( 0.09 Content Mixing 1.7 ( 0.05 1.2 ( 0.04 0.4 ( 0.04

A2

k2 (10-3 s-1)

0.527 ( 0.001 0.375 ( 0.002 0.384 ( 0.005 0.585 ( 0.005

12.3 ( 0.08 9.73 ( 0.09 11.7 ( 0.1 14.6 ( 0.1

0.251 ( 0.003 0.147 ( 0.003 0.22 ( 0.01 0.274 ( 0.001

15.1 ( 0.4 11.0 ( 0.4 11.4 ( 0.6 16.3 ( 0.3

8.4 ( 0.2 4.8 ( 0.2 3.1 ( 0.1

11.0 ( 0.5 7.9 ( 0.4 5.6 ( 0.3

TABLE 2: Rate Constants for DNA-Induced Lipid and Content Mixing of DOPC/DOPE/SM/CH Vesicles k1 (10-3 s-1)

A1

a

ds-2/3 + ds-1/4 ds-mm2/3 + ds-mm1/4 ss-1 + ss-2a ss-3 + ss-4a

15.11 ( 0.05 8.63 ( 0.04

ds-2/3 + ds-1/4 ds-mm2/3 + ds-mm1/4

5.0 ( 0.06 3.6 ( 0.02

Lipid Mixing 0.43 ( 0.004 0.49 ( 0.006

Content Mixing 1.0 ( 0.02 0.5 ( 0.02

A2

k2 (10-3 s-1)

4.84 ( 0.03 1.7 ( 0.03 7.88 ( 0.01 5.58 ( 0.01

13.6 ( 0.3 10.5 ( 0.5 11.0 ( 0.2 11.3 ( 0.3

5.0 ( 0.07 1.8 ( 0.03

11.8 ( 0.4 14.7 ( 0.7

Data were corrected for drift at t > 5 min before fitting with a single-exponential function.

ds-mm2/mm3 + ds-mm1/mm4 (red) in Figure 1A). DNAspecificity of lipid mixing was demonstrated by the absence of lipid mixing for vesicles carrying non-cDNA or for vesicles the DNA binding sites of which were blocked by incubation with free cDNA. The time courses of lipid mixing were well described by double-exponential functions of the general form y ) A1(1 - e-k1t) + A2(1 - e-k2t) with slow and fast rate constants of k1 ≈ 0.001 s-1 and k2 ≈ 0.011 s-1 (see Tables 1 and 2). For DNA that exhibited three consecutive, terminal C-C base mismatches (ds-mm2/mm3 + ds-mm1/mm4), k2 was slightly lower (0.0097 s-1) than for matched DNA (ds-2/3 + ds-1/4), whereas k1 was preserved. For ss-dc1 + ss-dc2, k1 was instead larger (0.003 s-1), whereas k2 remained unchanged. Interestingly, the time course of lipid mixing induced by single CH single strands (ss-1 + ss-2) was described by a singleexponential function with k2 ≈ 0.012 s-1. Here, lipid mixing stalled after 5 min at a significantly lower final level (∼21%) than that observed for double CH-anchored DNA (∼35%). It is also interesting to note that the kinetics of ss-dc1 + ss-dc2 clearly resembles the kinetics of ds-2/3 + ds-1/4 guided lipid mixing, although no duplex melting is required for complete hybridization in the former case. Thus, melting of the short double-stranded region can be excluded as the rate-limiting step for lipid mixing induced by ds-2/3 + ds-1/4, and the biphasic nature of the time course is implicated with the fusion mechanism characteristic for double CH-DNA rather than with duplex melting. For DOPC/DOPE/CH vesicles, inner leaflet mixing was monitored by using the same assay employing labeled vesicles the donor dye of which, NBD, was selectively bleached by dithionite treatment. Upon this treatment, vesicles typically lost ∼60% of their NBD fluorescence intensity. The inner leaflet mixing rates for the individual types of DNA were identical with those recorded for total lipid mixing (Figure 1B). This finding distinguishes DNA-mediated fusion from SNARE-driven

fusion in that inner leaflet mixing was found to be slower than outer leaflet mixing in the latter process.40 In all cases, inner leaflet mixing was in the range of ∼50% of the total lipid mixing yet slightly lower for the mismatched and single CH-labeled DNA constructs. This observation suggests the following two scenarios: (i) 50% of the vesicles undergo complete fusion immediately, whereby inner and outer leaflet mixing is perfectly synchronized, and (ii) intermediates were formed, in which the inner and outer leaflets exchanged lipids, however, without permanent opening of a fusion pore. A hypothetical structure that could account for inner leaflet mixing without pore opening is a highly disordered transmonolayer contact intermediate (see further below). II. Content Mixing and Leakage of DOPC/DOPE/CH Vesicles. To obtain a more detailed picture of the events associated with DNA-mediated fusion, we probed mixing of the aqueous vesicle contents and leakage by employing a conventional Tb/DPA assay.10 In brief, a nonfluorescent Tb/ citrate complex was encapsulated in one and DPA in the other vesicle population. Fusion, marked by mixing of vesicle contents, produces a strongly fluorescent Tb/DPA complex, the emergence of which was recorded as a function of time. The reaction buffer contained quenching agents to eliminate the signal arising from leakage out of the vesicles. Leakage was examined independently by coencapsulating the Tb/DPA complex and following the decrease in intensity as unlabeled vesicles with cDNA were added. For DOPC/DOPE/CH vesicles, content mixing was observed for all tested DNA constructs. The rate constants and extents of content mixing decreased in the order ds-2/3 + ds-1/4 (17%, Figure 2A) > ds-mm2/3 + ds-mm1/4 (11%, Figure 2B) > ss-1 + ss-2 (5%, Figure 2C). For double CH-anchored DNA, the kinetics of inner leaflet and content mixing were in good agreement, implying that inner leaflet mixing truly represents fusion pore opening. However, the significant divergence



Figure 2. Content mixing (black) and leakage (red) for DOPC/DOPE/CH vesicles as induced by different DNA constructs. Vesicles, loaded with Tb and DPA, were functionalized with cDNA strands at a molar vesicle-to-DNA ratio of 1:100 and mixed in a 1-to-1 ratio. Formation of the fluorescent Tb/DPA complex as a consequence of content mixing was recorded as a function of time. DNA-induced leakage was tested by reacting vesicles loaded with Tb/DPA complex and unlabeled vesicles. DNA modifications: ds-2/3 + ds-1/4 (A), ds-mm2/3 + ds-mm1/4 (B), and ss-1 + ss-2 (C). Vesicles modified with non-cDNA were monitored for content mixing (open black circles) and leakage (open red circles).

between the rate constants of content and inner leaflet mixing in the case of ss-1 + ss-2 suggests the existence of a fusion intermediate, such as a flickering fusion pore, that allowed for inner leaflet mixing but only for little and slow exchange of contents. This is an important observation in light of the abundance of studies that interpret inner leaflet mixing assays in terms of complete fusion. Membrane fusion is strictly defined as nonleaky merger of lipid bilayers, and hence, the model system for native fusion should ideally be leakage free. However, leakage is a common observation for vesicle-vesicle fusion assays in vitro, and the origins are debatable. In the DNA-mediated case (Figure 2A-C), leakage occurs with a significant lag time relative to content mixing. The lower the final level of content mixing monitored for a DNA construct, the earlier leakage sets in until it finally exceeds content mixing after about 60 min. Unfortunately, the leakage assay cannot identify which mode of vesicle interaction is the leaky one. As we know from the comparison of total and inner leaflet mixing above, nonproductive vesicle contacts exist that lead to outer but not to inner leaflet mixing. Leakage can equally well originate from those linked vesicles or be a consequence of fusion pore opening itself. Possible factors that facilitate leakage are (i) formation of nonbilayer phases because of large amounts of CH and DOPE, (ii) refolding of the fusion products into multilamellar structures, and (iii) liquid exchange with the ambient medium to adjust the inner volume of the fusion products according to the shape of the resulting vesicles. An important question is whether DNA-mediated fusion traverses through structural intermediates that are relevant for biological fusion. For the process induced by ds-2/3 + ds-1/4, the absence of a lag phase between inner and outer leaflet mixing and content mixing and the agreement of the corresponding rate constant point toward a nonsequential, hence largely concerted, fusion mechanism. Weinreb and Lentz recently published evidence that experimental data of this kind can very well be compatible with a sequential fusion mechanism as assumed in the framework of the stalk theory.41 The authors stress that a fusion intermediate state does not correspond to a fixed structure but describes an ensemble of rapidly interconverting structures with different functional properties such as leakage and content and lipid mixing. III. Lipid/Content Mixing and Leakage of DOCP/DOPE/ SM/CH Vesicles. Experiments involving DOPC/DOPE/SM/CH vesicles reproduce the trends reported above for DOPC/DOPE/ CH. In agreement with the results above, lipid mixing is most efficient for ds-2/3 + ds-1/4, yielding ∼18% lipid mixing after 60 min (Figure 3A). In the same time interval, ds-mm2/mm3

+ ds-mm1/mm4 induced only ∼7% lipid mixing. Both DNA substrates display double-exponential lipid mixing time courses, with k1 ≈ 0.004-0.005 s-1 and k2 ≈ 0.011-0.014 s-1. Single CH-DNA-induced lipid mixing is characterized by a rapid, single-exponential increase (k2 ≈ 0.011 s-1) that reaches a maximum after only 5 min. The longer, 27-mer, single strand is more efficient hereby with its amplitude even exceeding matched double CH-DNA during the initial phase. Content mixing was only observed for double CH-anchored DNA, yielding 10% with ds-2/3 + ds-1/4 (Figure 3B) and 5% with ds-mm2/mm3 + ds-mm1/mm4 (Figure 3C); the single strands produced a negligible amount of content mixing. Remarkably, the latter two DNA constructs induced significant leakage despite low extents of content mixing. In summary, the data presented above demonstrate the importance of double CH-anchoring for DNA-mediated fusion. Studies with two different model membranes, DOPC/DOPE/ CH and DOPC/DOPE/SM/CH, confirmed that single CH-DNA facilitates short-lived lipid mixing but is nearly ineffective in content mixing. For the latter lipid mixture, the associated leakage increases relative to the amount of content mixing, suggesting vesicle rupture or leaky DNA reorganization as a domineering process. Previously, SM was shown to reduce vesicle leakage in PEG-induced fusion while maintaining the fusogenity of the membrane.42 In the present work, SM diminished the total amount of leakage; however, leakage relative to the total amount of content mixing remained almost identical. The elongation of the linker region of double CHanchored DNA by only three base mismatches, which increases the membrane spacing by at most 1 nm, decreases the amount of content mixing by ∼33% for DOPC/DOPE/CH vesicles and even by ∼50% for DOPC/DOPE/SM/CH vesicles. Thus, the nature of the linker connecting the transmembrane and cytosolic segment and the membrane proximity play a crucial role in DNA-mediated fusion. Effect of DNA Length. It has been speculated that secretory membrane fusion is fueled by a coiled-coil transition of the four helix bundle in the core SNARE complex.2 We addressed the question whether the fusogenity of the DNA zippers will depend on the magnitude of free energy made available from DNA hybridization. To this end, we varied the length of the DNA strands and examined the resulting lipid mixing for DOPC/ DOPE/SM/CH vesicles. The complementary strands ss-3 + ss4, ss-1 + ss-2, and ss-5 + ss-6 were 12, 27, and 42 bases long, with a GC content of ∼50%. To maintain the possibility for double CH anchoring by partial duplex formation, we preserved the base sequence closest to the membrane.


Stengel et al.

Figure 3. Fusion of DOPC/DOPE/SM/CH vesicles as a function of DNA length and CH anchoring. (A) Lipid mixing. Bodipy500/Bodipy530labeled vesicles were mixed with a 4-fold amount of unlabeled vesicles after they had been functionalized with different DNA constructs at a molar vesicle-to-DNA ratio of 1:100. Color coding: ds-2/3 + ds-1/4 (black), ds-3/6 + ds-4/5 (green circles), ds-mm2/3 + ds-mm1/4 (red), ss-1 + ss-2 (blue), ss-3 + ss-4 (cyan), ss-5 + ss-6 (magenta circles), and vesicles with non-cDNA (black circles). (B-D) Content mixing (black) and leakage (red) of DOPC/DOPE/SM/CH vesicles under conditions identical to those described in Figure 2. DNA modifications: ds-2/3 + ds-1/4 (B), dsmm2/3 + ds-mm1/4 (C), and ss-1 + ss-2 (D).

Figure 3A shows lipid mixing of DOPC/DOPE/SM/CH vesicles induced by DNA constructs of different length. 12mer single strands induced less lipid mixing than 27-mer single strands; however, lipid mixing induced by the 42-mer exactly matched that of the 27-mer. Similarly, lipid mixing triggered by the partial duplexes ds-2/3 + ds-1/4 and ds-3/6 + ds-4/5 is identical. Because we know that single CH-DNA does not induce content mixing, we will limit our discussion to the double CH-tagged DNA constructs. Above, we hypothesized that melting of the duplex regions of ds-2/3 and ds-1/4 in order to form the blunt-ended duplexes ds-1/2 and ds-3/4 is energetically favorable. This results from the inspection of the Gibbs free energy, ∆G, of the reaction educts and products: two 12 base pair duplexes (2 × 16.3 kcal/mol, if we exclude additional stabilizing effects due to the overhanging bases) are rearranged into a 12 (16.3 kcal/mol) and a 27 (37.3 kcal/mol) base pair duplex upon DNA zipper formation, yielding a net free energy gain, ∆(∆G) of 53.6 - 32.6 ) 21 kcal/mol. The same calculation yields a free energy gain of 45.9 kcal/mol for the structural rearrangement of ds-3/6 + ds-4/5. As the kinetic traces agree perfectly for the two DNA pairs, one can conclude that the energy released upon DNA hybridization cannot be transmitted to the membrane in the given geometry. The strain imposed onto the membrane when the end of the DNA that is proximal to the membrane hybridizes seems to be more relevant for membrane fusion. Thus, the efficiency of membrane fusion cannot be improved by increasing the length of the 27-mer DNA strand in the partial duplex. From these experiments, we cannot rule out that alteration of the base sequence nearest to the membrane would change the outcome of fusion. However, because of the need for double CH anchoring, this cannot be

tested conveniently without altering the energy for duplex melting in our system. Effect of DNA Surface Coverage. Another relevant question is whether a minimum number of DNA strands is required to induce fusion. So far, we only presented DNA-induced fusion data obtained with a lipid-to-DNA ratio of 1316:1, which averages ∼100 DNA strands per vesicle when quantitative binding is assumed. By employing FRET-based affinity titrations, we derived a KD of ∼20 nM with a maximal coverage of ∼450 DNA strands for binding of double CH-anchored DNA to 100 nm LUVs (data not shown). At such a low DNA coverage, the presence of DNA is not likely to induce phase changes of the lamellar bilayer structure. The fact that vesicle contents are retained in the presence of DNA gives evidence for the integrity of the membrane. By using ds-2/3 + ds-1/4 and DOPC/DOPE/SM/CH vesicles, the molar DNA-to-vesicle ratio was varied between 100:1 and 3:1 (Figure 4). The time courses of lipid mixing were nearly identical down to DNA coverages as low as 13 DNA strands per vesicle. and DNA-to-vesicle ratios larger than 100:1 did not result in faster or more efficient lipid mixing (not shown). At coverages below 13:1, lipid-mixing rates and efficiencies decreased significantly. The weak dependence of lipid mixing on the number of incorporated DNA strands suggests that only a few DNA tethers are needed to perturb the membrane sufficiently to induce lipid mixing. According to the literature, activation barriers for membrane fusion range between 25 and 40 kcal/mol. To overcome this barrier, fusion is commonly a concerted effort of protein assemblies in nature. The number of v-SNAREs in synaptic vesicles is widely quoted as 30-100 per vesicle,43 and


Figure 4. Effect of DNA surface coverage on lipid mixing. Bodipy500/ Bodipy530-labeled and unlabeled DOPC/DOPE/SM/CH vesicles were modified with ds-2/3 and ds-1/4 at different molar vesicle-to-DNA ratios. Vesicles without (magenta) or with non-cDNA (molar vesicleto-DNA ratio 1:100) were monitored as controls. DNA-free vesicles were less stable than vesicles covered with non-cDNA, presumably because of electrostatic repulsion between the latter. We did not run negative controls with non-cDNA at different surface concentrations but only captured the extreme cases without or with 100 DNA strands. The estimated numbers of DNA strands per vesicle are indicated in the plot.

fusion pore opening catalyzed by the influenza hemaggluttinin virus requires 6-8 virus trimers.44 In our system, about 12 DNA strands seem to be required for significant fusion. Assembly and Stability of Intervesicle Contacts. We used ellipsometry to investigate the stability of DNA-bridged DOPC/ DOPE/SM/CH vesicles, by immobilizing one of the vesicle populations on a solid support. In particular, we examined the stability of the intervesicle contact mediated by ds-2/3 + ds1/4 at 18, 30, and 37 °C and compared it with vesicle stability mediated by ss-1 + ss-2 at 30 °C. Ellipsometry is a surface-sensitive optical method that allows for the quantification of surface-bound mass based on refractive index changes. Bulk vesicles were transferred to the surface by using a well characterized PEG-grafted polylysine (PLL-PEG)based binding matrix.45 In the first step, biotin-doped PLL-PEG was electrostatically adsorbed to a silicon dioxide surface. PLLPEG coated surfaces have been demonstrated to be inert to vesicle binding.45 Subsequently, a biotin-DNA/neutravidin complex was immobilized and used to capture bulk vesicles carrying a low copy number of complementary binding-DNA (see Materials and Methods) to avoid tether-induced vesicle deformations (∼3 per vesicle). Figure 5A displays the time trace for surface binding of vesicle species V1 (∼3 binding DNA, ∼100 ds-2/3). After 2 h, the V1 coverage averaged 962 ng/cm2 ( 10%. Assuming a spatial requirement of 55 Å2 per lipid (which takes into account the condensing effect of CH in the presence of SM),46 this amount corresponds to a surface coverage of 51%, which is in good agreement with the coverage expected from the random sequential adsorption of spheres. Thus, the surface is densely packed with vesicles. In a second step, vesicle species V2 was injected. V2 was modified with ∼100 ds-1/4, which bound specifically to V1-anchored ds-2/3. At 18 °C, V2 remained stably bound to V1 for more than 12 h. At 30 °C, 10% of the total lipid mass was lost after 12 h, and the lipid release was even more pronounced at 37 °C, yielding 19% lost lipid mass after only 3 h. Because the surface attachment of V1 was absolutely stable over the entire temperature range, the lost lipid mass can be attributed to release of V2. The stochiometry at maximal V2 coverage was ∼1.2 (V2:V1) at all temperatures. The rate of V2 release increased dramatically when V1 and V2

J. Phys. Chem. B, Vol. 112, No. 28, 2008 8271 were linked by hybridization of ss-1 and ss-2 at 30 °C. In this case, the release was so rapid that V2 barely accumulated on the surface and was completely released after 15 min. The presented ellipsometry data show that coupling of vesicles by DNA in a zipperlike geometry is accompanied by vesicle release and the magnitude of the release depends strongly on the temperature. This kind of destabilization has not been reported for vesicle networks sustained by membrane-anchored DNA duplexes that are oriented parallel to the surface normal47,48 (cDNA strands both attached via the 3′ end). A plausible explanation is that the zipperlike geometry puts strain on the CH anchors, for instance by inducing a tilt of CH, which is relieved by shuttling of the CH anchors to the opposite membrane. Formation of a transient hemifusion state would inevitably relocate the CH anchors to the same membrane, even without exposing the CH anchors to water. As expected, elevated temperatures provide the intrinsic energy to promote DNA shuttling and thus vesicle release. Double CH anchoring of the DNA is clearly an advantage because it counteracts dissociation of the DNA-bridged vesicle complex or makes fusion faster than dissociation. Thus, complete melting of double CHanchored DNA such as that implied in Scheme 1 may not take place. Possibly, a fast equilibrium between different DNA configurations exists that holds the CH anchors in proximity. The fact that single CH-DNA tethered vesicles are released within 15 min may explain the distinct shapes of lipid mixing curves for the same construct (Figures 1A and 3A). There, lipid mixing is initially rapid but levels off after only 5 min, followed by a plateau phase. The plateau phase may correspond to a situation where the majority of vesicles have dissociated. To determine whether vesicle release is a process that competes with membrane fusion, we carried out lipid-mixing assays at 18, 30, and 37 °C by employing ds-2/3 + ds-1/4 and DOPC/DOPE/SM/CH vesicles. No lipid mixing was observed at 18 °C (Figure 5B). The rate of lipid mixing at 37 °C was larger than the rate observed at 30 °C (Figure 5B), but the efficiencies were identical. Thus, thermal energy simultaneously facilitates both vesicle release due to CH shuttling and lipid mixing. Discussion The presented membrane fusion concept is a reductionist approach toward mimicking protein-mediated membrane fusion. Unlike fusion proteins, DNA can easily be modified to test certain parameters that may effect fusion, such as DNA length, DNA complementarity and base sequence, and the nature of the anchoring group. Even though it is tempting to infer similarities from a comparison of the overall architecture of the SNARE complex and the DNA zippers, the mechanism underlying DNA-mediated fusion had not been investigated in detail. In the following discussion, we will identify possible driving forces and discuss parallels to native membrane fusion. Kinetics of Fusion. To address whether lipid mixing is instantaneous upon vesicle docking, we first approximate the vesicle collision rate. The rate, kdiff, of diffusion-controlled collision in three dimensions is given by kdiff ) 4πNA(RA + RB)(DA + DB). Using NA ) Avogadro’s number, RA ) RB ) 50 nm as vesicle radius, and DA ) DB ) 3 × 10-8 cm2 s-1 as vesicle diffusion constant49 gives a kdiff of ∼5 × 109 M-1 s-1. Our lipid-mixing assays were carried out by using an excess of unlabeled, CU, over labeled vesicles, CL, with CL ) 0.6 nM and CU ) 2.4 nM. Under these pseudo-first-order conditions, vesicle docking occurs initially with an effective rate keff ) kdiffCU ) 12 s-1. Naturally, not every collision leads to vesicle


Stengel et al.

Figure 5. (A) DNA-mediated interactions of DOPC/DOPE/SM/CH vesicles probed by ellipsometry. To prevent vesicle binding or spreading, silicon dioxide surfaces were made inert with PLL-PEG. The PLL-PEG layer was doped with biotin to allow the immobilization of DNA-neutravidin complex. The plot shows the injection and subsequent surface binding of vesicles (V1) modified with ∼100 strands of ds-2/3 and a small amount of unrelated DNA that was complementary to neutravidin-coupled DNA. In the next step, vesicles modified with ∼100 strands of ds-1/4 were added to examine the interaction between vesicles carrying fusion-inducing DNA. Vesicle interactions were investigated at 18 (red), 30 (black), and 37 °C (blue). The green curve corresponds to vesicles modified with complementary single CH-DNA (V1 + ss-1 and V2 + ss-2) at 30 °C. (B) Lipid mixing of DOPC/DOPE/SM/CH vesicles induced by ds-2/3 + ds-1/4 at different temperatures.

binding. Boxer and co-workers recently reported a docking probability of P ) 0.06 for vesicles that were diffusing laterally in two dimensions (Dves ≈ 0.2 × 10-8 cm2 s-1) and were modified with 50 copies of complementary 24 base single strands.47 A conservative estimate then yields a docking probability of about 0.01 in three dimensions. If we apply P ) 0.01 to our system, the effective rate of docking decreases to keff ) 0.12 s-1. Hence, vesicle docking is at least one order of magnitude faster than the fastest rate constant observed for lipid mixing (0.01 s-1). Consequently, lipid mixing is unlikely to occur instantaneously upon vesicle docking, which is consistent with the observation that the lipid-mixing rates did not accelerate under conditions that promote the rate of vesicle docking, such as increasing DNA length or DNA coverage. Boxer and coworkers have also shown that the probability for vesicle docking scales strongly with the length of the DNA strands, ∝ (lDNA)4.47 Although we monitored a difference in the final extent of lipid mixing for 12-mer and 27-mer DNA, there was no change in rate when the DNA length was extended beyond 27 bases. Furthermore, we did not observe rate changes when varying the number of DNA strands between 100 and 25 for DOPC/ DOPE/SM/CH vesicles, whereas the docking rate changed by a factor of 6 when 50 instead of 25 DNA strands were employed in the 2-dimensional docking assay mentioned above.47 Taken together, these observations suggest that lipid mixing itself must be the rate-limiting step in the fusion pathway. In addition to thermal energy constraints, the fusion reaction may be slowed down by the requirement for several DNA tethers at the vesicle-vesicle interface. Although diffusion of double CHDNA is quite fast (DCH-DNA ≈ 0.2 × 10-8 cm2 s-1), DNA could have difficulties entering the vesicle contact zone because of repulsive hydration forces. In our study, lipid and content mixing of DOPC/DOPE/CH vesicles induced by double CH-DNA was a biphasic process with a slow, k ≈ 0.001 s-1, and a fast, k ≈ 0.01 s-1, component. Overall, these rates are in line with rates previously reported for in vitro vesicle fusion. In PEG-mediated fusion, lipid-mixing rates range between 0.07 and 0.005 s-1 depending on the lipid composition and reaction conditions, whereas content mixing tends to be slower with a typical value of 0.002 s-1.9 In the first report of SNARE-mediated vesicle fusion, one round of fusion (measured as lipid mixing) was accomplished after 20-50 min, which is comparable to what we observed.3 However, in later publications, the fusion rates were scattered

between ∼10-5 and 0.06 s-1,50 which makes it difficult to make conclusive statements. Interestingly, when Lentz and co-workers reconstituted SNAREs for the first time at a low SNARE coverage, which was similar to our DNA coverage, SNAREs catalyzed neither docking nor fusion.51 In the presence of 6% PEG, SNARE-modified DOPC/DOPE/SM/CH/DOPS vesicles displayed lipid-mixing rate constants of ∼0.003 s-1 and contentmixing rate constants of 0.004 s-1, with a low extent of content mixing of 5% after 10 min. By using DOPC/DOPE/SM/CH vesicles, DNA-induced lipid mixing proceeded with rate constants of k1 ≈ 0.004 s-1 and k2 ≈ 0.0136 s-1, whereas content mixing took place with k1 ≈ 0.0001 s-1 and k2 ≈ 0.0118 s-1. The extent of content mixing was 7% after 10 min. Hence, at similar receptor coverage and lipid composition, DNAmediated fusion is slightly more effective than SNARE-mediated fusion in the presence of PEG. Membrane Distance and DNA Anchoring. Because of the repulsive hydration forces and surface roughness, the natural separation distance of biological membranes is ∼5-10 nm. Experiments using the surface force apparatus57 and theoretical calculations58 have demonstrated the presence of strong attractive forces at separation distances below 2 nm, especially when the membrane is laterally stressed at the same time. In fact, the resulting attraction is strong enough to merge bilayers. DNA, lipid-anchored parallel to the vesicle’s surface normal, was previously used to connect vesicles.48,52 The corresponding separation distance was about 8 nm.53 Fusion has not been observed in this geometry, which perfectly agrees with our own results obtained for parallel anchoring of double CH-DNA. The fact that fusion can be accomplished by simply reversing the direction of the cDNA strands to form a DNA zipper strongly suggests a decrease in membrane separation distance as one driving force for fusion. This is further supported by the observation that a DNA construct the linker of which, connecting CH and DNA, was extended by the introduction of only three base mismatches fuses membranes more slowly and less efficiently. In addition to reducing membrane distance by switching from the parallel to the zipperlike geometry, we have likely added drag and/or tilt to the CH groups. The ellipsometry experiments give evidence for the existence of such pulling force in that the dissociation rate of DNA-bridged vesicle complexes is faster than the dissociation rate of DNA from the membrane under stress-free (unhybridized) conditions. This is true for both single and double CH-DNA; however, vesicle complexes

Membrane Fusion Induced by CH-Modified DNA Zippers tethered by the former are very short-lived. Our data support that lipid mixing can occur during shuttling of the DNA from trans into cis position, but the membranes do not interact long enough to open a fusion pore. DNA release requires that the repulsion acting between two membranes is larger than the energy needed for DNA detachment. Thus, the hydrophobicity of the DNA anchor is a crucial design criterion for an artificial fusion machinery. Phillips et al. measured the rate of spontaneous CH transfer between DOPC bilayers under equilibrium conditions and found a strong correlation with the bilayer packing density.46 Addition of SM to PC/CH membranes and the associated membrane condensation reduced the CH transfer rates up to two orders of magnitude. At 37 °C, CH was released at ∼0.00034 s-1 from a DOPC membrane, and the van der Waals attractive energy between cholesterol and neighboring phospholipids was estimated to 5.6 kcal/mol on the basis of the theory from Salem.54 Although CH has fewer hydrocarbons than phospholipids, the van der Waals interactions between CH and phospholipids are comparable to the interaction of phospholipids with each other. Confirming that hydrophobic interactions scale with the length of the hydrocarbon chain and with the surface area of the membraneembedded molecule (which is large for CH’s fused ring system),55 nearly identical limiting areas, ∼40 Å, were reported for CH and phospholipids in compression experiments.46 Because CH is a fairly simple membrane anchor that does not span both lipid leaflets, DNA-mediated fusion is likely to be primarily proximity driven. This observation seems to collide with previous work, in which the transmembrane domains of SNARE proteins were identified to play a crucial role. Rothman and co-workers replaced the transmembrane domains of SB and SX with isoprenes or PE to examine the importance of the SNARE transmembrane domains. The PE-linked proteins did not catalyze lipid mixing,56 whereas the isoprene-linked ones did. The isoprene group was theoretically long enough to span both lipid leaflets; however, no proof was provided that it actually did. Besides the depth of membrane penetration, the cross section of the hydrophobic group might be very important, which would explain why double CH-anchored DNA is more effective in fusion than single CH-DNA. Even though we managed to accomplish membrane fusion by using membrane-anchored DNA, one should be aware of the crucial role membrane composition plays in this process. Simple PC bilayers, which were not activated by the addition of CH and PE, did not undergo DNA-mediated fusion. Among the miscellaneous effects CH and PE have on membrane properties, the tendency to form inverted bilayer phases, the reduction of bilayer rupture tension, and the negative intrinsic curvature are probably the most prominent ones. The link between fusion and lipid properties is treated in several excellent reviews,57,58 and we foresee that the DNA-based approach presented here will help further elucidate the important role played by lipids. Conclusions In this work, we demonstrated that short DNA oligonucleotides, membrane-attached via CH in an orientation that mimics the overall zipperlike architecture of fusion-inducing proteins, induce fusion of 100 nm LUVs composed of DOPC/DOPE/ CH and DOPC/DOPE/SM/CH. Membrane fusion was demonstrated by established lipid-mixing and content-mixing assays employing small amounts of DNA (ca. 100 DNA strands per vesicle). Anchoring of DNA with two CH groups was found to be essential for fusion, whereas single CH-anchored DNA

J. Phys. Chem. B, Vol. 112, No. 28, 2008 8273 triggered primarily mixing of the outer lipid leaflet and leakage, owing to the rapid dissociation of the DNA-bridged vesicle complex in the latter case. The relaxation of the linker region proximal to CH by nucleobase mismatches caused a significant reduction in content mixing, thus emphasizing the importance of close bilayer apposition and linker stiffness in DNA-mediated fusion. Although the use of longer DNA strands increases the probability for vesicle docking, membrane fusion was not furthered when using 42/12-mers duplex instead of 27/12-mers. This observation, together with fusion rate constants falling below the rate constants expected for vesicle docking, suggests that the DNA and lipid rearrangements taking place at the vesicle-vesicle contact zone are limiting for fusion. We anticipate that DNA, being a convenient modular building block that allows the control of vesicle interactions, will help shed light on the mechanism of biological membrane fusion in the future. Acknowledgment. We thank Pauline Vandoolaeghe and Tommy Nylander for support for the ellipsometry measurements and Peter Jo¨nsson for valuable comments. Funding was provided by the Marie Curie Grant MEIF-CT-2006-039909 and the Swedish Research Council (Grant 2005-3140). We thank LayerLab AB, Gothenburg, Sweden, for help with the design of some of the cholesterol-modified oligonucleotides. References and Notes (1) Jahn, R.; Grubmuller, H. Membrane fusion. Curr. Opin. Cell Biol. 2002, 14 (4), 488–495. (2) Sollner, T. H. Intracellular and viral membrane fusion: A uniting mechanism. Curr. Opin. Cell Biol. 2004, 16 (4), 429–435. (3) Weber, T.; Zemelman, B. V.; McNew, J. A.; Westermann, B.; Gmachl, M.; Parlati, F.; Sollner, T. H.; Rothman, J. E. SNAREpins: Minimal machinery for membrane fusion. Cell 1998, 92 (6), 759–772. (4) Siegel, D. P. The modified stalk mechanism of lamellar/inverted phase transitions and its implications for membrane fusion. Biophys. J. 1999, 76 (1), 291–313. (5) Xu, Y. B.; Zhang, F.; Su, Z. L.; McNew, J. A.; Shin, Y. K. Hemifusion in SNARE-mediated membrane fusion. Nat. Struct. Mol. Biol. 2005, 12 (5), 417–422. (6) Chernomordik, L. V.; Kozlov, M. M. Protein-lipid interplay in fusion and fission of biological membranes. Annu. ReV. Biochem. 2003, 72, 175–207. (7) Salaun, C.; James, D. J.; Chamberlain, L. H. Lipid rafts and the regulation of exocytosis. Traffic 2004, 5 (4), 255–264. (8) Lee, J.; Lentz, B. R. Secretory and viral fusion may share mechanistic events with fusion between curved lipid bilayers. Proc. Natl. Acad. Sci. U.S.A. 1998, 95 (16), 9274–9279. (9) Lentz, B. R. PEG as a tool to gain insight into membrane fusion. Eur. Biophys. J. Biophys. Lett. 2007, 36 (4-5), 315–326. (10) Wilschut, J.; Duzgunes, N.; Fraley, R.; Papahadjopoulos, D. Studies on the Mechanism of Membrane-FusionsKinetics of Calcium-Ion Induced Fusion of Phosphatidylserine Vesicles Followed by a New Assay for Mixing of Aqueous Vesicle Contents. Biochemistry 1980, 19 (26), 6011–6021. (11) Pantazatos, D. P.; MacDonald, R. C. Directly observed membrane fusion between oppositely charged phospholipid bilayers. J. Membr. Biol. 1999, 170 (1), 27–38. (12) Kunishima, M.; Tokaji, M.; Matsuoka, K.; Nishida, J.; Kanamori, M.; Hioki, K.; Tani, S. Spontaneous membrane fusion induced by chemical formation of ceramides in a lipid bilayer. J. Am. Chem. Soc. 2006, 128 (45), 14452–14453. (13) Sommerdijk, N.; Hoeks, T. H. L.; Synak, M.; Feiters, M. C.; Nolte, R. J. M.; Zwanenburg, B. Stereodependent fusion and fission of vesicles: Calcium binding of synthetic gemini phospholipids containing two phosphate groups. J. Am. Chem. Soc. 1997, 119 (19), 4338–4344. (14) Vandenburg, Y. R.; Smith, B. D.; Perez-Payan, M. N.; Davis, A. P. Non-leaky vesicle fusion and enhanced cell transfection using a cationic facial amphiphile. J. Am. Chem. Soc. 2000, 122 (13), 3252–3253. (15) Pincet, F.; Lebeau, L.; Cribier, S. Short-range specific forces are able to induce hemifusion. Eur. Biophys. J. Biophys. Lett. 2001, 30 (2), 91–97. (16) Kemble, G. W.; Danieli, T.; White, J. M. Lipid-Anchored Influenza Hemagglutinin Promotes Hemifusion, Not Complete Fusion. Cell 1994, 76 (2), 383–391.

8274 J. Phys. Chem. B, Vol. 112, No. 28, 2008 (17) Salzwedel, K.; Johnston, P. B.; Roberts, S. J.; Dubay, J. W.; Hunter, E. Expression and Characterization of Glycophospholipid-Anchored HumanImmunodeficiency-Virus Type-1 Envelope Glycoproteins. J. Virol. 1993, 67 (9), 5279–5288. (18) Richard, A.; Marchi-Artzner, V.; Lalloz, M. N.; Brienne, M. J.; Artzner, F.; Gulik-Krzywicki, T.; Guedeau-Boudeville, M. A.; Lehn, J. M. Fusogenic supramolecular vesicle systems induced by metal ion binding to amphiphilic ligands. Proc. Natl. Acad. Sci. U.S.A. 2004, 101 (43), 15279– 15284. (19) Gong, Y.; Luo, Y. M.; Bong, D. Membrane activation: Selective vesicle fusion via small molecule recognition. J. Am. Chem. Soc. 2006, 128 (45), 14430–14431. (20) Stengel, G.; Zahn, R.; Hook, F. DNA-induced programmable fusion of phospholipid vesicles. J. Am. Chem. Soc. 2007, 129 (31), 9584–9585. (21) Sutton, R. B.; Fasshauer, D.; Jahn, R.; Brunger, A. T. Crystal structure of a SNARE complex involved in synaptic exocytosis at 2.4 angstrom resolution. Nature 1998, 395 (6700), 347–353. (22) Sorensen, J. B.; Wiederhold, K.; Muller, E. M.; Milosevic, I.; Nagy, G.; de Groot, B. L.; Grubmuller, H.; Fasshauer, D. Sequential N- to C-terminal SNARE complex assembly drives priming and fusion of secretory vesicles. EMBO J. 2006, 25 (5), 955–966. (23) Shea, R. G.; Marsters, J. C.; Bischofberger, N. Synthesis, Hybridization Properties and Antiviral Activity of Lipid-Oligodeoxynucleotide Conjugates. Nucleic Acids Res. 1990, 18 (13), 3777–3783. (24) Boutorin, A. S.; Guskova, L. V.; Ivanova, E. M.; Kobetz, N. D.; Zarytova, V. F.; Ryte, A. S.; Yurchenko, L. V.; Vlassov, V. V. Synthesis of Alkylating Oligonucleotide Derivatives Containing Cholesterol or Phenazinium Residues at their 3′-Terminus and their Interaction with DNA within Mammalian-Cells. FEBS Lett. 1989, 254 (1-2), 129–132. (25) Gosse, C.; Boutorine, A.; Aujard, I.; Chami, M.; Kononov, A.; Cogne-Laage, E.; Allemand, J. F.; Li, J.; Jullien, L. Micelles of lipidoligonucleotide conjugates: Implications for membrane anchoring and base pairing. J. Phys. Chem. B 2004, 108 (20), 6485–6497. (26) Kurz, A.; Bunge, A.; Windeck, A. K.; Rost, M.; Flasche, W.; Arbuzova, A.; Strohbach, D.; Mueller, S.; Liebscher, J.; Huster, D.; Herrmann, A. Lipid-anchored oligonucleotides for stable double-helix formation in distinct membrane domains. Angew. Chem., Int. Ed. 2006, 45 (27), 4440–4444. (27) Letsinger, R. L.; Chaturvedi, S. K.; Farooqui, F.; Salunkhe, M. Use of Hydrophobic Substituents in Controlling Self-Assembly of Oligonucleotides. J. Am. Chem. Soc. 1993, 115 (16), 7535–7536. (28) Mackellar, C.; Graham, D.; Will, D. W.; Burgess, S.; Brown, T. Synthesis and Physical-Properties of Anti-Hiv Antisense Oligonucleotides Bearing Terminal Lipophilic Groups. Nucleic Acids Res. 1992, 20 (13), 3411–3417. (29) Scheidt, H. A.; Flasche, W.; Cismas, C.; Rost, M.; Herrmann, A.; Liebscher, J.; Huster, D. Design and application of lipophilic nucleosides as building blocks to obtain highly functional biological surfaces. J. Phys. Chem. B 2004, 108 (41), 16279–16287. (30) Tomkins, J. M.; Barnes, K. J.; Blacker, A. J.; Watkins, W. J.; Abell, C. Lipophilic modification of oligonucleotides. Tetrahedron Lett. 1997, 38 (4), 691–694. (31) Zhang, G. R.; Farooqui, F.; Kinstler, O.; Letsinger, R. L. Informational liposomes: Complexes derived from cholesteryl-conjugated oligonucleotides and liposomes. Tetrahedron Lett. 1996, 37 (35), 6243–6246. (32) Tiberg, F.; Landgren, M. Characterization of Thin Nonionic Surfactant Films at the Silica Water Interface by Means of Ellipsometry. Langmuir 1993, 9 (4), 927–932. (33) Baumeler, S. Sugar Adsorption on PLL-g-PEG-coated Nb2O5 Surfaces; ETH: Zu¨rich, 2004. (34) Xu, F.; Zhen, G. L.; Yu, F.; Kuennemann, E.; Textor, M.; Knoll, W. Combined affinity and catalytic biosensor: In situ enzymatic activity monitoring of surface-bound enzymes. J. Am. Chem. Soc. 2005, 127 (38), 13084–13085. (35) Grant, L. M.; Tiberg, F. Normal and lateral forces between lipid covered solids in solution: Correlation with layer packing and structure. Biophys. J. 2002, 82 (3), 1373–1385. (36) Pfeiffer, I.; Hook, F. Bivalent cholesterol-based coupling of oligonucletides to lipid membrane assemblies. J. Am. Chem. Soc. 2004, 126 (33), 10224–10225. (37) Haque, M. E.; Lentz, B. R. Roles of curvature and hydrophobic interstice energy in fusion: Studies of lipid perturbant effects. Biochemistry 2004, 43 (12), 3507–3517.

Stengel et al. (38) Duzgunes, N. Fluorescence assays for liposome fusion. Liposomes, Part B 2003, 372, 260–274. (39) Malinin, V. S.; Haque, M. E.; Lentz, B. R. The rate of lipid transfer during fusion depends on the structure of fluorescent lipid probes: A new chain-labeled lipid transfer probe pair. Biochemistry 2001, 40 (28), 8292– 8299. (40) Lu, X. B.; Zhang, F.; McNew, J. A.; Shin, Y. K. Membrane fusion induced by neuronal SNAREs transits through hemifusion. J. Biol. Chem. 2005, 280 (34), 30538–30541. (41) Weinreb, G.; Lentz, B. R. Analysis of membrane fusion as a twostate sequential process: Evaluation of the stalk model. Biophys. J. 2007, 92 (11), 4012–4029. (42) Haque, M. E.; McIntosh, T. J.; Lentz, B. R. Influence of lipid composition on physical properties and PEG-mediated fusion of curved and uncurved model membrane vesicles: “Nature’s own” fusogenic lipid bilayer. Biochemistry 2001, 40 (14), 4340–4348. (43) Brunger, A. T. Structure and function of SNARE and SNAREinteracting proteins. Q. ReV. Biophys. 2005, 38 (1), 1–47. (44) Bentz, J.; Mittal, A. Architecture of the influenza hemagglutinin membrane fusion site. Biochim. Biophys. Acta. 2003, 1614 (1), 24–35. (45) Stadler, B.; Falconnet, D.; Pfeiffer, I.; Hook, F.; Voros, J. Micropatterning of DNA-tagged vesicles. Langmuir 2004, 20 (26), 11348– 11354. (46) Lundkatz, S.; Laboda, H. M.; McLean, L. R.; Phillips, M. C. Influence of Molecular Packing and Phospholipid Type on Rates of Cholesterol Exchange. Biochemistry 1988, 27 (9), 3416–3423. (47) Chan, Y. H.; L, P.; Boxer, S. G. Kinetics of DNA-mediated docking reactions between vesicles tethered to supported lipid bilayers. Proc. Natl. Acad. Sci. U.S.A. 2007, 104 (48), 18913–18918. (48) Yoshina-Ishii, C.; Chan, Y. H. M.; Johnson, J. M.; Kung, L. A.; Lenz, P.; Boxer, S. G. Diffusive dynamics of vesicles tethered to a fluid supported bilayer by single-particle tracking. Langmuir 2006, 22 (13), 5682– 5689. (49) Schuette, C. G.; Hatsuzawa, K.; Margittai, M.; Stein, A.; Riedel, D.; Kuster, P.; Konig, M.; Seidel, C.; Jahn, R. Determinants of liposome fusion mediated by synaptic SNARE proteins. Proc. Natl. Acad. Sci. U.S.A. 2004, 101 (9), 2858–2863. (50) Liu, T. T.; Tucker, W. C.; Bhalla, A.; Chapman, E. R.; Weisshaar, J. C. SNARE-driven, 25-ms vesicle fusion in vitro. Biophys. J. 2005, 89 (4), 2458–2472. (51) Dennison, S. M.; Bowen, M. E.; Brunger, A. T.; Lentz, B. R. Neuronal SNAREs do not trigger fusion between synthetic membranes but do promote PEG-mediated membrane fusion. Biophys. J. 2006, 90 (5), 1661–1675. (52) Svedhem, S.; Pfeiffer, I.; Larsson, C.; Wingren, C.; Borrebaeck, C.; Hook, F. Patterns of DNA-labeled and scFv-antibody-carrying lipid vesicles directed by material-specific immobilization of DNA and supported lipid bilayer formation on an Au/SiO2 template. ChemBioChem 2003, 4 (4), 339–343. (53) Ajo-Franklin, C. M.; Yoshina-Ishii, C.; Boxer, S. G. Probing the structure of supported membranes and tethered oligonucleotides by fluorescence interference contrast microscopy. Langmuir 2005, 21 (11), 4976–4983. (54) Salem, L. Attractive Forces between Macromolecular Chains of Biological Importance. Nature 1962, 193 (4814), 476–477. (55) Israelachvili, J. N. Intermolecular and Surface Forces; Academic Press: London, 1992. (56) McNew, J. A.; Weber, T.; Parlati, F.; Johnston, R. J.; Melia, T. J.; Sollner, T. H.; Rothman, J. E. Close is not enough: SNARE-dependent membrane fusion requires an active mechanism that transduces force to membrane anchors. J. Cell Biol. 2000, 150 (1), 105–117. (57) Chernomordik, L. Non-bilayer lipids and biological fusion intermediates. Chem. Phys. Lipids 1996, 81 (2), 203–213. (58) Tamm, L. K.; Crane, J.; Kiessling, V. Membrane fusion: A structural perspective on the interplay of lipids and proteins. Curr. Opin. Struct. Biol. 2003, 13 (4), 453–466. (59) Chan, Y.-H. M.; Lengerich, B.; Boxer, S. G. Lipid-anchored DNA mediates vesicle fusion as observed by lipid and content mixing. Biointerphases 2008, 3 (2), FA17–FA21.

JP802005B

Determinants for Membrane Fusion Induced by ... - ACS Publications

Recommend Documents