Induced Simultaneous Formation of G

DOI: 10.1002/chem.201504389

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& Electroformation | Very Important Paper|

Point-to-Plane Nonhomogeneous Electric-Field-Induced Simultaneous Formation of Giant Unilamellar Vesicles (GUVs) and Lipid Tubes Chuntao Zhu, Ying Zhang, Yinan Wang, Qingchuan Li, Wei Mu,* and Xiaojun Han*[a] Abstract: It is well-known that homogeneous electric fields can be used to generate giant unilamellar vesicles (GUVs). Herein we report an interesting phenomenon of formation of GUVs and lipid tubes simultaneously using a nonhomogeneous electric field generated by point-toplane electrodes. The underlying mechanism was analyzed using finite element analysis. The two forces play main roles, that is, the pulling force (F) to drag GUVs into lipid tubes induced by fluid flow, and the critical force (Fc) to prevent GUVs from deforming into lipid tubes induced by electric fields. In the center area underneath the needle electrode, the GUVs were found because F is less than Fc in that region, whereas in the edge area the lipid tubes were obtained because F is larger than Fc. The diffusion coefficient of lipid in the tubes was found to be 4.45 mm2 s¢1 using a fluorescence recovery after photobleaching (FRAP) technique. The method demonstrated here is superior to conventional GUV or lipid tube fabrication methods, and has great potential in cell mimic or hollow material fabrication using GUVs and tubes as templates.

Molecular self-assembly is considered a key technology in the design and fabrication of diverse nano- and microstructure materials. There are two general ways for molecules to assemble into ordered structures, that is, self-assembly through direct interactions among building blocks, or indirectly induced by an external field, such as magnetic-, electric-, and flow fields. An electric field was reported to trigger 1,3,6,8-tetrakis(1-butyl-1H1,2,3-triazol-4-yl)pyrene (TP) molecules to form a flower structure on top of a trimesic acid (TMA) matrix.[1] An electric field was also used to induce amphiphilic molecules to assemble into vesicles and tubes.

[a] C. Zhu, Y. Zhang, Y. Wang, Q. Li, Dr. W. Mu, Prof. Dr. X. Han State Key Laboratory of Urban Water Resource and Environment School of Chemical Engineering and Technology Harbin Institute of Technology 92 West Da-Zhi Street, Harbin 150001 (P. R. China) E-mail: [email protected] [email protected] Supporting information for this article is available on the WWW under http://dx.doi.org/10.1002/chem.201504389. Chem. Eur. J. 2016, 22, 2906 – 2909

Lipid vesicles and tubes, the self-assembled lipid aggregates, are very common in biological systems. The morphology of lipid aggregates is crucial in determining their biological function.[2] As two important forms of artificial bio-membrane, they have attracted a lot of attention. For several decades, lipid vesicles have been used as model systems to study cellular behavior.[3] In particular, giant unilamellar vesicles (GUVs) with similar sizes to live cells have been widely used in the studies of lipid rafts,[4] gene delivery,[5] encapsulation of biologically active compounds,[6] and protocells.[7] Lipid tubes were ideal models for the study of the transport of molecules between cells.[8] Recently, many methods have been developed to generate GUVs and lipid tubes, which can be roughly categorized into two classes: self-assembly of lipid molecules and field-induced self-assembly of lipid molecules. For GUVs, they were generated by amphiphiles through a self-assembling process in water,[9] and also formed by electroformation,[10] microfluidic jetting method,[6a,d] etc. In particular, electroformation is more popular due to its high yields, high unilamellarity and good reproducibility.[10b, 11] Meanwhile, a variety of methodologies have been developed to generate the lipid tubes, including self-assembly of lipid molecules[2a, 12] and deformation of lipid aggregates undergoing a certain field.[8a,b, 13] Molecular self-assembly is a simple and powerful technique for lipid-tube formation,[14] but it is limited to particular lipids, such as1,2-bis(10,12-tricosadiynoyl)-sn-glycero-3-phosphocholine (DC8,9PC). Molecular self-assembly induced by an external field, including hydrodynamic flow,[13b,c] electric fields,[15] etc., has been used to generate lipid tubes as well. So far, there have been no reports on the formation of GUVs and lipid tubes simultaneously. Herein we developed a novel point-to-plane electrode microsystem (Figure 1 a and b) for generating GUVs and lipid tubes simultaneously on the same substrate. The vesicle area was located in the center regions underneath the needle electrode with a diameter of 600 mm, whereas the lipid tubes were formed at the edge regions. Finite element analysis was performed to understand this phenomenon. The lipid vesicles ((4.28 œ 1.32) Õ 104 cm¢2) and tubes ((1.38 œ 0.39) Õ 104 cm¢2) can be generated with high yields, which have great potential in cell study or hollow nanomaterial synthesis. This method may extend to other amphiphilic molecules. Figure 1 c presents a fluorescence image of 1,2-dioleoyl-snglycero-3-phosphocholine (DOPC) vesicles/tubes containing 0.5 % Texas red-labeled 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt (TR-DHPE) formed on

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Figure 2. The fluorescence images after applying AC electric field with amplitude of 2.5 V and frequency of 10 Hz with the electrode distance of 340 mm for a) 0, b) 60, c) 135, and d) 170 min, respectively. The scale bar is 100 mm. e–h) Schematic illustration of the growing process corresponding to each top image.

Figure 1. Schematic setup for lipid GUV and tube fabrication and fluorescence images of lipid GUVs and tubes. a) Schematic illustration of the setup containing the ITO electrode (bottom) and a polydimethylsiloxane (PDMS) coverslip (top) separated by a rectangular polytetrafluoroethylene frame. b) Schematic side view of the setup in (a). c) The overview fluorescence image of the DOPC (doped with 0.5 %TR-DHPE) GUVs and lipid tubes under an AC field of 2.5 V and 10 Hz with the electrode distance of 340 mm. d) Zoomed-in view of the GUV region. e) Zoomed-in view of the lipid tube region. f) FRAP experiment of the lipid tubes. Fluorescent images of DOPC nanotubes taken at i) 10 s and ii) 10 min after photobleaching, respectively. The scale bars are 500 mm (c), 50 mm (d, f) and 100 mm (e).

the indium tin oxide (ITO) electrode surface upon applying an alternating current (AC) field of 2.5 V, 10 Hz for 180 min. It is clearly seen that the GUVs were located in the center area vertically underneath the tungsten needle, whereas the lipid tubes were generated outside the GUV area and grew radially towards the edges. In a zoomed-in image of GUV area (Figure 1 d), GUVs were sphere shape with the size ranging from 16 to 56 mm in diameter (obtained from 350 random measurements), while in a zoomed-in image of the tube area, the lipid tubes were generated with a length ranging from 100 mm to 1 mm and diameter of approximately 9 œ 3.43 mm (obtained from 170 random measurements). The lipid tubes, like elongated balloons, have a similar shape to those in previous reports.[13a, 16] The GUVs and lipid tubes could survive up to 21 days at room temperature. GUVs and lipid tubes kept their spherical and tubule shapes respectively. The lipid tubes are more stable than previous results.[17] Furthermore, a fluorescence recovery after photobleaching (FRAP) technique was used to measure the diffusion coefficient (D) of TR-DHPE within the lipid tubes (Figure 1 f). The images were taken after 10 s (Figure 1 f i) and 10 min (Figure 1 f ii) after photobleaching. The lipid tubes became dark after photobleaching in the center area, whereas their fluorescence intensity became homogeneous after 10 min. The D value was calculated to be 4.45 mm2 s¢1 with a mobile fraction of 0.87, indicating the highly mobile feature of lipids in nanotubes. The diffusion conChem. Eur. J. 2016, 22, 2906 – 2909

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stant is in the range of that obtained from giant liposomes from 1.1 to 6.3 mm2 s¢1.[18] The growth process of lipid tubes was tracked by a microscope in real-time at an amplitude of 2.5 V and a frequency of 10 Hz. Figure 2 a–d shows the sample images in the lipid tube formation area after applying the field for 0, 60, 135, and 170 min, respectively. Before applying the field, the bilayer stacked on the glass surface gave the homogenous fluorescence intensity, as shown in Figure 2 a. Just after applying the sinusoidal AC field, the bilayer stacks started to swell, and formed GUVs after 60 min, as shown in Figure 2 b. Subsequently, the mature GUVs were pulled into lipid tubes (Figure 2 c) after 135 min, and the lipid tubes extended continuously between 135 and 170 min. Finally, a typical fluorescence image of elongated balloon lipid tubes was observed as shown in Figure 2 d. The corresponding schematic illustrations are shown in Figure 2 e–h. Apart from the deformed vesicles, the lipid sources of the tubes are partially from the bilayer stacks on the electrode surface, because the tubes changed their membrane areas to respond to the strain.[19] To understand the formation process of lipid tubes, 2D finite element analysis of the electric and fluid coupling field generated by the point-to-plane electrode were performed using COMSOL software at amplitude of 2.5 V and frequency of 10 Hz. The fluid flow of the solution was simulated. Figure 3 a shows a snapshot of the simulation with the largest magnitude during each cycle, that is, j sinwt j = 1. Due to the electrode setup, the fluid close to the bottom electrode surface flows towards the edge of the bottom electrode and then swirls vertically back to the needle electrode. The video of the fluid flow is shown in the Supporting Information (Movie S1). The lipid tubes grew along with the fluid flow direction, which suggests to us that the fluid pulling force caused the lipid tube formation. In current experimental conditions, the lipid tubes were only generated when the pulling force (F) from fluid flow overcame the critical force (Fc) from the interactions among lipids and electric fields to keep its spherical shape. The pulling force acting on the GUVs (F) is related to the lateral velocity (Vx) by Equation (1) below: F ¼ 6phrV x

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ð5Þ

Further bringing Equation (5) into Equation (2), Fc is given by Equation (6) below: Fc ¼ 2p

Figure 3. a) A snapshot of fluid flow induced by point-to-plane electrode system at an amplitude of 2.5 V and a frequency of 10 Hz at t = 0.025 s. Vx components (b) and electric field strength (c) against lateral distance away from the center point above the plane electrode surface with 1, 5, 10, 25, 50, 80, and 100 mm cross-section height, respectively. d) The pulling force acting on the GUVs (F, squares) obtained using Equation (1) and the critical force (Fc, circles) obtained using Equation (6).

where r is the radius of GUV, and is about 25 mm. The value of h is 0.7208 Õ 10¢3 Pa s at 35 8C. The critical force (Fc) is given in Equation (2) below:[13b] Fc ¼ 2p

pffiffiffiffiffiffiffiffiffiffiffi 2kc sh

ð2Þ

where sh is the effective surface tension of the GUVs, and the value of bending rigidity (kc) under 35 8C is 7.58 Õ 10¢20 J.[20] The relationship between the effective surface tension sh and the pressure difference DP across the GUV surface[21] is shown in Equation (3): DP ¼ 2sh =r

ð3Þ

where DP is obtained from Equation (4) below. The derivation process is shown in the Supporting Information. DP ¼ P¢0 ¼ 0:6627 e0 ew E 2

ð4Þ

where e0 is 8.854 Õ 10¢12 F/m, and ew is 74.85. Bringing Equation (4) into Equation (3), Equation (5) is obtained. Chem. Eur. J. 2016, 22, 2906 – 2909

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pffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffiffi 0:6626kc e0 ew E 2 r

ð6Þ

The pulling force (F) and critical force (Fc) relate to lateral fluid velocity (Vx) and electric field strength (E), respectively. Therefore Vx and E were simulated. The simulated velocity components on the x axis (Vx, parallel to the plane electrode surface) at a cross-section height of 1, 5, 10, 25, 50, 80, and 100 mm are shown in Figure 3 b. It is noted that Vx increases with decreasing solution height from 1.39 mm s¢1 at 100 mm to 5.67 mm s¢1 at 1 mm. It means that Vx component become more and more predominant approaching the bottom electrode. Figure 3 c shows the electric field strength at a cross section of 1, 5, 10, 25, 50, 80, and 100 mm above the plane electrode surface. E increases with increasing cross-section height and mainly centralizes in the center area below the needle electrode. It increases from the edge to the center regions of the plane electrode and reaches the maximum at the center of the electrodes, which indicates that E becomes more and more pronounced towards the center areas. According to the reality, we chose the cross plan with 25 mm above the bottom electrode to calculate the pulling force (F) and critical force (Fc) using Equations (1) and (6), respectively, as shown in Figure 3 d. There are two intersections in Figure 3 d and the corresponding positions are 130 and 885 mm away from the center point, respectively. It is clear to see that pulling force gradually increases while the critical force gradually decreases between 0 and 130 mm, where F is always smaller than Fc. Consequently in this region, the GUVs retain their spherical shape, which is consistent with the experimental results in Figure 1 c. In the region between 130 to 885 mm, F is bigger than Fc, which causes GUVs to be pulled into lipid tubes. In the region outside 885 mm, although F is smaller than Fc in the calculation, lipid tubes were observed. The reason may be that the lipid tubes at the border near 885 mm extended into that area. The GUV area decreased as the distance between electrodes became smaller. We also investigated the influence of voltage on lipid tube growth. Once the lipid tubes formed, the further growth speed was investigated as a function of amplitudes of electric fields, as shown in Figure 4 a. The rate of formation of lipid tubes at each given amplitude was measured out at least three times. The higher amplitude causes bigger growth speed. The speeds were 0.133, 0.275, 0.527, 0.629, and 0.734 mm s¢1 at 1.0, 2.0, 2.5, 3.0, and 4.0 V, respectively. This result is consistent with the simulated data, as shown in Figure 4 b. At 25 mm above the surface of the electrode, the Vx increases against the field amplitude. The consistency is further evidence that the lateral fluid flow dragged the vesicle into lipid tubes. In conclusion, we have demonstrated that a point-to-plane electrode system can be used to simultaneously induce lipid molecular self-assemblies into GUVs and lipid tubes in one go. The mechanism was analyzed in detail using simulations. The

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[4]

[5] [6] Figure 4. (a) Experimental data of relative growth length at different voltage: 1.0 V (&), 2.0 V (*), 2.5 V (~), 3.0 V (*), and 4.0 V (*). (b) Curve of Vx components of fluid flow field at a 25 mm cross section above the electrode surface at amplitudes of 1.0 (&), 2.0 (*), 2.5 (~), 3.0 (*), and 4.0 V (*), with a frequency of 10 Hz.

critical force (Fc) generated by electric field prevents GUVs from deforming into lipid tubes, whereas the pulling force (F) generated from lateral fluid flow drags GUVs to form lipid tubes. In the area where Fc is less than F, the lipid tubes were formed, otherwise the GUVs were formed. A FRAP study showed that the fluidity of internal lipid is as good as that in other model cell membrane systems, which suggests that the lipid GUVs and tubes obtained using our method have great potential as cell mimics.

[7]

[8]

[9] [10]

Acknowledgements This work was supported by the National Natural Science Foundation of China (Grant No. 21273059, 21003032, 21528501, 21511130060), State Key Laboratory of Urban Water Resource and Environment (Harbin Institute of Technology) (Grant No. 2014DX09), Harbin Science and Technology Research Council (Grant No. 2014RFXXJ063), and the Fundamental Research Funds for the Central Universities (Grant No. HIT. KISTP. 201407).

[11] [12]

[13]

[14]

Keywords: electroformation · finite element analysis · fluorescence recovery after photobleaching · giant unilamellar vesicles · lipid tube

[15] [16] [17]

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Received: November 1, 2015 Published online on January 28, 2016

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