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Lipid Nanotubule Fabrication by Microfluidic Tweezing Jonathan West, Andreas Manz, and Petra S. Dittrich* ISAS - Institute for Analytical Sciences, Bunsen-Kirchhoff-Str. 11, D-44139 Dortmund, Germany ReceiVed February 14, 2008. ReVised Manuscript ReceiVed March 11, 2008 There is currently great interest in the development of lipid enclosed systems with complex geometrical arrangements that mimic cellular compartments. With biochemical functionalization, these soft matter devices can be used to probe deeper into life’s transport dominated biochemical operations. In this paper, we present a novel tool for machining lipid nanotubules by microfluidic tweezing. A bilayer poly(dimethylsiloxane) (PDMS) device was designed with a lipid reservoir that was loaded by capillary action for lipid film deposition. The lipid reservoir is vertically separated from an upper flow for controlled material wetting and the formation of giant tubule bodies. Three fluidic paths are interfaced for introduction of the giant tubules into the high velocity center of a parabolic flow profile for exposure to hydrodynamic shear stresses. At local velocities approximating 2 mm s-1, a 300-500 nm diameter jet of lipid material was tweezed from the giant tubule body and elongated with the flow. The high velocity flow provides uniform drag for the rapid and continuous fabrication of lipid nanotubules with tremendous axial ratios. Below a critical velocity, a remarkable shape transformation occurred and the projected lipid tubule grew until a constant 3.6 µm diameter tubule was attained. These lipid tubules could be wired for the construction of advanced lifelike bioreactor systems.
Introduction Spontaneously forming lecithin liquid crystals were first discovered by Bangham in 1965.1 These spherical crystals consist of a bimolecular lipid leaflet encapsulating an aqueous core and have become known as liposomes or vesicles. These have made an extraordinary impact in diverse applications, including cosmetic, food, and drug preparations. Beyond these industrial applications, vesicles provide scientists with a biomimetic tool to probe life’s biochemical processes. Here, the combination of a confined microscopic environment enclosed by a functional fluid membrane provide a model for collision probable and fluctuation dominated reactions occurring within cells and organelles.2 The exquisite morphologies of cellular compartments enable transport controlled reaction sequences to be undertaken for a huge variety of molecular processing operations. To further mimic cellular architectures, tubular geometries are also required. The self-assembly of lipid amphiphiles into nanotubes with lengths of tens to hundreds of micrometers is well documented.3–5 Dimensional control is largely derived from lipid choice, molecular engineering, and solvent makeup.6 Photopolymerization and other postassembly techniques are commonly used to derive templates for the fabrication of rugged and functionalized hollow nanocylinders.7 For life science investigations, lipids must be retained in a fluidic state for accurate membrane and membrane protein * To whom correspondence and requests for materials should be addressed. E-mail:
[email protected]. (1) Bangham, A. D.; Standish, M. M.; Watkins, J. C. J. Mol. Biol. 1965, 13, 238–252. (2) Chiu, D. T.; Wilson, C. F.; Ryttse´n, F.; Stro¨mberg, A.; Farre, C.; Karlsson, A.; Nordholm, S.; Gaggar, A.; Modi, B. P.; Moscho, A.; Garza-Lo´pez, R. A.; Orwar, O.; Zare, R. N. Science 1999, 283, 1892–1895. (3) Nakashima, N.; Asakuma, S.; Kim, J.-M.; Kunitake, T. Chem. Lett. 1984, 1709–1712. (4) Yager, P.; Schoen, P. E. Mol. Cryst. Liq. Cryst. 1984, 106, 371–381. (5) Yamada, K.; Ihara, H.; Ide, T.; Fukumoto, T.; Hirayama, C. Chem. Lett. 1984, 1713–1716. (6) Shimizu, T.; Masuda, M.; Minamikawa, H. Chem. ReV. 2005, 105, 1401– 1443. (7) Schnur, J. M. Science 1993, 262, 1669–1676.
functionality. As a self-assembly alternative, mechanical approaches can be used to manipulate lipid assemblies. Umbilical nanotubes tethered to maternal vesicles can be formed by the application of a point load. This can be achieved by the relative translation of an adhered microparticle8,9 or micropipet10,11 away from a vesicle. Initial efforts focused on the use of these point load mechanisms to elucidate the fundamental biophysics of lipid bilayer dynamics.12 The micropipet can also provide additional lipid material and be used for directly writing complex conjugated vesicle-nanotube networks on a surface. These can be photopolymerized to serve as templates for metal and molecular structuring.13 More recently, these remarkable soft matter nanoand microfluidic systems were retained in their native fluidic state to represent geometrically sophisticated lifelike cellular models.14,15 Biochemically functionalized membranes and interiors were used to investigate biochemistry at the nanoscale,16,17 and molecular transport was made possible using Marangoni18 or electrophoretic flows.19 For a greater appreciation of these biomimetic networks, readers are directed to the outstanding review by Karlsson et al.20 (8) Roopa, T.; Kumar, N.; Bhattacharya, S.; Shivashankar, G. V. Biophys. J. 2004, 87, 974–979. (9) Heinrich, V.; Bozˇicˇ, B.; Svetina, S.; Zˇeksˇ, B. Biophys. J. 1999, 76, 2056– 2071. (10) Waugh, R. E. Biophys. J. 1982, 38, 29–37. (11) Frusawa, H.; Fukagawa, A.; Ikeda, Y.; Araki, J.; Ito, K.; John, G.; Shimizu, T. Angew. Chem., Int. Ed. 2003, 42, 72–74. (12) Evans, E.; Yeung, A. Chem. Phys. Lipids 1994, 73, 39–56. (13) Evans, E.; Bowman, H.; Leung, A.; Needham, D.; Tirrell, D. Science 1996, 273, 933–935. (14) Karlsson, A.; Karlsson, R.; Karlsson, M.; Cans, A. S.; Stro¨mberg, A.; Ryttse´n, F.; Orwar, O. Nature 2001, 409, 150–152. (15) Hurtig, J.; Gustafsson, B.; Tokarz, M.; Orwar, O. Anal. Chem. 2006, 78, 5281–5288. (16) Davidson, M.; Karlsson, M.; Sinclair, J.; Sott, K.; Orwar, O. J. Am. Chem. Soc. 2003, 125, 374–378. (17) Bauer, B.; Davidson, M.; Orwar, O. Langmuir 2006, 22, 9329–9332. (18) Karlsson, R.; Karlsson, A.; Orwar, O. J. Am. Chem. Soc. 2003, 125, 8442–8443. (19) Tokarz, M.; Åkerman, B.; Olofsson, J.; Joanny, J.-F.; Dommersnes, P.; Orwar, O. Proc. Natl. Acad. Sci. U.S.A. 2005, 102, 9127–9132. (20) Karlsson, M.; Davidson, M.; Karlsson, R.; Karlsson, A.; Bergenholtz, J.; Konkoli, Z.; Jesorka, A.; Lobovkina, T.; Hurtig, J.; Voinova, M.; Orwar, O. Annu. ReV. Phys. Chem. 2004, 55, 613–649.
10.1021/la8004823 CCC: $40.75 2008 American Chemical Society Published on Web 05/27/2008
High Axial Ratio Lipid Tubules
Figure 1. (a) Illustration of the microfluidic device assembly. The device comprises a lipid reservoir, a basal microfluidic circuit, and an upper linear microfluidic channel. (b) Microscopy image of the aligned PDMS layers, with the upper flow path indicated with a black arrow and the basal converging flow paths indicated with white arrows.
A rival strategy considers fluid flow for wetting lipid films and the directed growth of multiple lipid tubules. This parallel fabrication method can be used to provide the unit elements that could be interconnected for the elaboration of more complex fluidic systems. A simple method involves the mechanically driven growth of giant membrane lobes, some tens of micrometers in diameter, across surfaces by wetting fronts.21 Alternatively, hydration gradients between two glass plates can be used for the slow growth of tubules.22 Other methods use microfluidics to apply hydrodynamic forces for the rapid formation of high axial ratio tubules with heterogeneous diameters.23 In a similar study, flow rate control was used to improve the modulation of tubule dimensions.24 For greater geometric control, the lipid film can be interfaced with the flow using a 100 nm thick silicon nitride membrane with an array of micropores for extruding tubules of prodigious lengths (several centimeters) with diameters matching those of the micropores.25 In this contribution, we present a new microfluidic format for the formation of giant lipid tubule bodies and introduction of these into a high velocity flow region for lipid nanotubule fabrication by hydrodynamic tweezing. This soft matter machining capability can be used to fabricate membrane enclosed aquaduct systems with scales appropriate for cellular processes.
Materials and Methods Concept. The method involves the introduction of reservoirtethered giant lipid tubules into a high velocity flow path where localized shear forces act to tweeze lipid nanotubules. The device is illustrated in Figure 1a and incorporates a basal lipid reservoir surrounded by a microfluidic circuit. A linear microchannel in the upper layer is positioned over the lipid reservoir. This enables controlled giant tubule production, without a microporous interface, by vertical separation from the flow. As shown in Figure 1b, the upper channel is also aligned to the basal channel to produce an (21) Suzuki, K.; Masuhara, H. Langmuir 2005, 21, 537–544. (22) Tan, T.-C.; Shen, A. Q.; Li, Y.; Elson, E.; Ma, L. Lab Chip 2008, 8, 339–345. (23) Brazhnik, K. P.; Vreeland, W. N.; Hutchison, B.; Kishore, R.; Wells, J.; Helmerson, K.; Locascio, L. E. Langmuir 2005, 21, 10814–10817. (24) Lin, Y.-C.; Huang, K.-S.; Chiang, J.-T.; Yang, C.-H.; Lai, T.-H. Sens. Actuators, B 2006, 117, 464–471. (25) Dittrich, P. S.; Heule, M.; Renaud, P.; Manz, A. Lab Chip 2006, 6, 488– 493.
Langmuir, Vol. 24, No. 13, 2008 6755 arrangement akin to a hydrodynamic focusing system where a central flow is flanked by parallel flows. However, in the described device, the three flows instead become interfaced as two basal paths and one upper path. The multilayer system is used to transport giant lipid tubule bodies into the center of a Poiseuille flow. Here, the local high velocities impart shear stresses for tweezing lipid tubules. Fabrication. The bilayer poly(dimethylsiloxane) (PDMS, Sylgard, Dow Corning) device was prepared by replica molding each layer from a SU-8 master. The upper and basal microchannels were both 500 µm wide with a depth of 55 µm. The SU-8 master was fabricated by standard methods involving spin-deposition, photolithography, and development. The PDMS prepolymer was mixed with the curing agent at a ratio of 10:1 (w/w) and then thoroughly degassed using a vacuum desiccator. To enable optical access, the basal layer was prepared as a thin (∼150 µm) PDMS cookie, produced by aliquoting a ∼200 µL volume of uncured PDMS onto the SU-8 master followed by placing a glass coverslip above and pressing firmly to spread the material as a thin film. The coverslip also provides a convenient handle for subsequent assembly steps. The PDMS cookies were cured on a hotplate at 70 °C for 5 min. The upper PDMS channel was molded on the SU-8 master as a robust 5 mm thick layer. Assembly and Operation. Phospholipids are common to life. In this study, the phospholipid 1,2-dilauryl-sn-glycero-3-phosphocholine (DLPC, Avanti Polar Lipids) was used. DLPC was dissolved in pure (spectroscopy grade) chloroform at concentrations of 50, 100, and 200 mM. The lipid reservoir was deposited by suspending a 500 nL droplet of chloroform-solved DLPC above the end of the channel. Upon contact with the PDMS, capillary and gravity forces act to propel the mixture along the length of the open microchannel. The chloroform quickly evaporated, leaving a dried film of lipid material. Reservoir films were characterized by white light interferometry (NewView 5000 microscope, Zygo). Fluidic interconnection was achieved by punching 1 mm diameter through holes in the thick upper PDMS layer. To remove particulate debris, an ethanol, distilled water, and N2 stream rinse sequence was used. An air plasma (1 mbar) treatment for 5 min was used to activate the surface of both PDMS layers with bonding achieved by visual alignment and pressing the layers together for a few seconds. Plasma treatment of the lipid film did not interfere with giant tubule formation. For simplicity of instrumentation, a single digitally controlled syringe pump (PHD 2000, Harvard Apparatus) was interfaced via Tygon tubing to the microfluidic device using Luer ports (Scandinavian Micro Biodevices ApS). These were attached using the polyvinyl siloxane based elastomer Panasil (Kettenbach) and aligned to the through holes using a pin. Fluid introduction from a common inlet port enters both the upper and basal channels. The fluid follows the upper linear channel, while in the basal channel it first diverges before converging in front of the lipid reservoir where it is reunited with the upper microchannel, with the now singular flow exiting from a common outlet port. The microfluidic device was positioned on an in-house machined stage for video analysis using an inverted microscope (Olympus IX 71), with image acquisition using a charge-coupled device (CCD) camera (EMCCD DV-887, Andor). The aqueous flows were seeded with 0.005% (w/v) 2 µm diameter polystyrene particles for local flow velocity estimation by image pair and integration time analysis.
Results and Discussion Phospholipids can be interfaced with hydrodynamic flows by the deposition of a bulk lipid film within a microchannel. This can be achieved by straightforward spotting followed by solvent evaporation to leave a dried lipid film.23,24 For greater geometrical control, the lipid reservoir can be deposited using vesicles positioned by wetting fronts22 or deposited within a micromachined cavity, interfaced with a microchannel via a microporous membrane.25 In this research, we have developed a novel lipid film deposition method that enables positional and film thickness control. The interfacial tensionx between the hydrophobic chloroform and PDMS materials can be exploited for the capillary
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driven loading of a bulk lipid film within an open channel. Simple positioning of a droplet of chloroform-solved lipid material at the microchannel origin produces a capillary driven and gravity assisted flow. A flow front in a 55 µm deep channel is documented in Figure 2a. ?>The video is provided in the Supporting Information (SI1). Transport velocities with a 50 mM DLPC solution were typically 700 µm s-1. With the use of higher concentrations (100 and 200 mM) with higher viscosities, the flow is significantly retarded with evaporation acting to limit transport lengths to only 2-4 mm. Solvent flow is therefore a dynamic interplay between capillary, gravity, and evaporation effects. Desirable transport lengths are thus achieved with low surface area to volume channel structures and low lipid concentrations to limit the effects of evaporation. The chloroform rapidly evaporated leaving a dried multilayer lipid film that hugged the channel walls. The dried lipid film is shown in figure 2b. The film thickness increased with increased lipid loading concentrations. White light interferometry of a 200 mM lipid deposit with a thickness of approximately 36 µm is shown in the Supporting Information (SI2). Loading with a 50 mM DLPC solution enables lipid deposition along the complete length of the reservoir, and it was therefore preferred for tubule tweezing experiments. At this loading concentration, the lipid film was typically 12 µm deep, containing an estimated 6000 molecular layers for the provision of large quantities of material for giant tubule body formation. Assembly of the basal PDMS layer with the upper microchannel provides a recessed lipid material reservoir. Fluid actuation through the device was used for wetting and swelling the lipid material in the reservoir. The lipid reservoir was vertically separated and protected from the major upper flow to enable the formation of near-uniform giant tubule bodies with diameters of 9-12 µm. Viscous drag acts to couple the wetted reservoir with the upper flow and transport the giant tubules out from the reservoir. Giant tubules aligned with the flow are shown in Figure 3. At the PDMS plateau between the reservoir and the converged microfluidic paths, frictional forces at the no-slip boundary act
Figure 2. Lipid reservoir deposition: (a) video microscopy frame of a capillary flow of chloroform-solved DLPC along the length of the reservoir channel and (b) the resultant dried lipid film.
Figure 3. Video microscopy frame of three well-developed giant tubule bodies with the central body deflected by the converging flows.
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Figure 4. (a) Hydrodynamic tweezing a nanoscopic jet of lipid material from the tip of a giant tubule body at 0.4 s and (b) tubule diameter growth to e1 µm at 2.1 s.
to impede transport. In addition, tensile forces further serve to anchor the giant tubule body to the lipid reservoir. With mean velocities of 1-1.5 mm s-1, the giant tubules were slowly introduced to the region of converged flow and became positioned within the vertical center of the Poisueille flow for nanotubule tweezing. Figure 4 shows video microscopy frames of a lipid nanotubule continuously elongated, from left to right, from a near-stationary giant tubule body. The local flow velocity is estimated to be ∼2 mm s-1. At the limits of optical microscopy resolution, the nanotubule became apparent once it attained a diameter of 300-500 nm (see Figure 4a). As the tweezing progressed, the giant tubule was transported further forward and the nanotubule grew in size to e1 µm in diameter (see figure 4b). This indicates that the giant tubule became positioned within a lower flow velocity region. This first generation prototype therefore requires refinement for diameter tuning by improved giant tubule positioning. The dimensions approximate those observed during nanotubule self-assembly by cooling lipid materials through the phase transition temperature (Tm).3,6,7 However, tubule tweezing is observed at room temperature, well above the Tm of DLPC (-1 °C). The mechanism of material organization is therefore more likely to be mechanical in nature, offering the opportunity to tune diameters by simple manipulation of the local velocity. This lipid nanotubule fabrication method differs markedly from those previously reported. Other approaches apply an axial load by adherence of a vesicle body to a surface followed by material pulling using a micropipet,13 or by the application of point loads by optical8 or magnetic tweezing9 using a microparticle handle. Alternative methods use hydrodynamic flows for vesicle transport while the tubule remains anchored to a surface.10,26 These methods involve transport of the vesicle body by the flow, with lipid material released behind as a nanotubular tether that connects to the surface. In contrast, our fully microfluidic strategy involves a stationary lipid body (giant tubule) with the nanotubule tweezed and transported with the flow. The hydrodynamic tweezer method satisfies the Evan’s criteria, with both the application of a highly localized force on a lipid body and the availability of excess material.12 The hydrodynamic drag force imparted by the converging flows on the near-stationary giant tubule body is difficult to accurately derive. For simplicity, the drag force Fd can be crudely approximated by Stoke’s drag on a sphere: Fd ) 6πηrV, where η is the viscosity (1 × 10-3 Pa · s for water at room temperature), r is the giant tubule radius (5 µm), and V is the local velocity (2 mm s-1). Under these conditions, the force is estimated to be ∼190 pN. In comparison, reported values for pulling a tubule from a bilayer are in the range of 10-40 pN.12,27,28 The larger value we obtain is likely to be accounted for by the primitive model assumptions and/or the (26) Rossier, O.; Borghi, C. N.; Puech, P. H.; Dere´nyi, I.; Buguin, A.; Nassoy, P.; Brochard-Wyart, F. Langmuir 2003, 19, 575–584. (27) Heinrich, V.; Waugh, R. E. Annu. ReV. Biomed. Eng. 1996, 24, 595–605. (28) Roux, A.; Cappello, G.; Cartaud, J.; Prost, J.; Goud, B.; Bassereau, P. Proc. Natl. Acad. Sci. U.S.A. 2002, 99, 5394–5399.
High Axial Ratio Lipid Tubules
multilamellar character of the giant tubule body (as indicted by the strong opacity, see Figure 3), where multiple interlayer shear stresses can combine. On occasion, it was possible to observe microscopic blebs originating and extending from the front of the nanotubule tether. These bleb-terminated nanotubules are best observed on video. One such structure can just be discerned in the video provided in the Supporting Information (SI3). Viscous tugging at the onset of tubulation could induce a tension (and material loss) that is sufficient to form a spherical bleb structure. Indeed, Bar-Ziv et al. have used high intensity laser trapping to produce multiple pearling instabilities that propagate along the length of a microtubule.29 The video (Supporting Information, SI3) also documents a remarkable shape transformation that occurs as the local velocity relaxes. This transformation is documented as video microscopy frames in Figure 5. A bleb-terminated nanotubule first emerged at a local velocity of ∼2 mm s-1. Following nanotubule formation, the velocity was reduced to 1.1 mm s-1 within a period of 1.3 s. In this time frame, the terminus of the giant tubule takes on a hydrodynamic profile, demonstrating reduced axial tension,26 before returning to a hemispherical shape. During this shape transformation, the tubule diameter enlarges from 300-500 nm to a constant diameter of 3.6 µm. This microscale diameter is maintained as the velocity slows to a stop. The critical velocity at which the shape transition occurs is estimated to be 1.8 mm s-1. Local velocity manipulation could be used for nano- and microscale diameter tuning. However, unlike optical tweezers,
Figure 5. Shape transformation during velocity relaxation. Initial tweezing of a lipid nanotube (with bleb) at ∼2 mm s-1 is followed by a rapid shape transformation as the velocity is reduced below 1 mm s-1. From top to bottom, frames were recorded at 0.76, 1.26, 1.64, and 7.46 s. Local velocities were determined by microparticle tracking. In the inset is an illustration (to scale) of a nanotubule projecting from the giant tubule body with an adjoining bleb structure.
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Figure 6. Diameters of the giant tubule body (O) and the elongated nano- and microtubule (2) during local velocity (broken line) relaxation plotted as a function of time.
this hydrodynamic method does not have sufficient temporal resolution to determine whether the shape transformation is continuous or discontinuous in nature.9 The dimensions of the depleting giant tubule body and the growth of the elongating tubule during velocity relaxation are recorded in Figure 6. This indicates that lipid material is recruited from the giant body during tubule fabrication, and not the remote lipid reservoir. The multilamellar giant tubule therefore acts as a local material source: an individual reservoir of abundant and wetted lipid material. This coupled with the high local velocities enables the rapid fabrication of nanotubules with tremendous lengths and gigantic axial ratios (>1000). This contrasts with the use of vesicle sources, where material rapidly becomes limited and even the smallest diameter tubules are limited to a few 100 µm in length. The presented microfluidic tweezing capability provides a rapid means for the parallel fabrication of high axial ratio lipid nanotubules. Such phospholipid structures are known to be stable for weeks at room temperature.23 The described lipid nanotubules can therefore serve as building blocks for the construction of geometrically complex systems that mimic cellular structures. Fluidic systems are ideally suited for patterning these structures using microfluidic routing30 to program the position and shape of the tubules. Alternatively, surface patterning in combination with biotin-avidin coupling can be used for the self-organization of lipid bodies.31,32 More elaborate networks comprising differently functionalized compartments can also be constructed by using lipids decorated with DNA motifs for encoded assembly by surface hybridization with complementary sequences.33,34 Critical to the success of these networks is the requirement to connect the different compartments. This can be achieved by triggering fusion using optical,35 electrical,36 or (bio)chemical37,38 methods. (29) Bar-Ziv, R.; Moses, E.; Nelson, P. Biophys. J. 1998, 75, 294–320. (30) Mahajan, N.; Fang, J. Langmuir 2005, 21, 3153–3157. (31) Stamou, D.; Duschl, C.; Delamarche, E.; Vogel, H. Angew. Chem., Int. Ed. 2003, 42, 5580–5583. (32) Sott, K.; Karlsson, M.; Pihl, J.; Hurtig, J.; Lobovkina, T.; Orwar, O. Langmuir 2003, 19, 3904–3910. (33) Yoshina-Ishii, C.; Miller, G. P.; Kraft, M. L.; Kool, E. T.; Boxer, S. G. J. Am. Chem. Soc. 2005, 127, 1356–1357. (34) Dusseiller, M. R.; Niederberger, B.; Sta¨dler, B.; Falconnet, D.; Textor, M.; Vo¨ro¨s, J. Lab Chip 2005, 5, 1387–1392. (35) Kulin, S.; Kishore, R.; Helmerson, K.; Locascio, L. Langmuir 2003, 19, 8206–8210. (36) Tresset, G.; Takeuchi, S. Biomed. MicrodeVices 2004, 6, 213–218.
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Conclusions and Outlook In this contribution, we describe a novel microfluidic strategy for the rapid and continuous fabrication of nanoscale lipid tubules with high axial ratios. The microfluidic device incorporates a lipid reservoir that is vertically separated from the upper flow for the controlled growth of giant lipid tubule bodies. These can be introduced to high velocity flows for exposure to axial shear stresses for tweezing lipid nanotubules. Beneath a critical velocity, a marked shape transformation occurs, with the tubules growing to microscale dimensions. Velocity manipulation may therefore hold promise for diameter tuning. This soft matter 3-D structuring capability can be used to create fluidic systems that would otherwise be difficult to realize with even the most advanced fabrication techniques that are currently available. This research forms the basis of our ambition to develop methods for the fabrication of membrane enclosed bioreactor networks. These could find application in the investigation, and ultimately the exploitation, of biochemical processes unique to life. (37) Evans, K. O.; Lentz, B. R. Biochemistry 2002, 41, 1241–1249. (38) Estes, D. J.; Lopez, S. R.; Fuller, A. O.; Mayer, M. Biophys. J. 2006, 91, 244–254.
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Acknowledgment. The authors are grateful to Helmut Lindner and Joachim Franzke for their learned insight into mechanics, Uli Marggraf for SU-8 master fabrication, and Norman Ahlman for white light interferometry studies. Financial support from the European Community (CellPROM project, Contract No. NMP4CT-2004-500039) under the 6th Framework Programme for Research and Technology Development is gratefully acknowledged. In addition, funding from the Ministerium fu¨r Innovation, Wissenschaft, Forschung and Technologie des Landes NordrheinWestfalen and from the Bundesministerium fu¨r Bildung and Forschung is also gratefully acknowledged. Supporting Information Available: SI1: Video microscopy of lipid reservoir deposition. SI2: White light interferometry depth profiling of a 300 µm wide reservoir with lipid deposited from a 200 mM DLPC in chloroform solution. The lipid film becomes depleted at longer transport distances. SI3: Video microscopy of lipid tubule shape transformation during velocity relaxation. This material is available free of charge via the Internet at http://pubs.acs.org. LA8004823
