Supported Lipid Bilayers with Phosphatidylethanolamine as the Major Component

Transcription

1 Supporting Information for: Supported Lipid Bilayers with Phosphatidylethanolamine as the Major Component Anne M. Sendecki 1, Matthew F. Poyton 1, Alexis J. Baxter 1, Tinglu Yang, 1 Paul S. Cremer 1,2 1 Department of Chemistry, 2 Department of Biochemistry and Molecular Biology, Pennsylvania State University, University Park, PA correspondence to: psc11@psu.edu S1

2 A Note on PE Distribution between Bilayer Leaflets It is expected that PE should preferentially partition to regions of high negative curvature in mixed PC and PE bilayer systems. For SLBs on planar substrates, however, there is very little curvature. Electrostatics can also drive leaflet asymmetry when charged lipids are present, 1 but both PE and PC are zwitterionic near neutral ph. As such, one would not expect any significant leaflet asymmetry for mixed PC and PE bilayers on this grounds either in planar SLB systems. The notion that the two leaflets would have roughly even lipid concentrations is supported by work from Anglin & Conboy, 2 who used mixed PC and PE bilayers to measure lipid flip-flop rates by vibrational sum frequency spectroscopy (VSFS). At equilibrium, their VSFS decay curves for mixed bilayer systems were found to go to baseline, indicating no significant leaflet asymmetry. Of course, PE should partition to the inner leaflet of curved structures, including tubules, when these structures protrude from the supported bilayer surface. S2

3 Additional Materials and Methods Materials Texas Red 1,2-dihexadecanoyl-sn-glycero-3-phosphoethanolamine, triethylammonium salt (TR- DHPE) was purchased from Life Technologies (Grand Island, NY). Poly(dimethylsiloxane) (PDMS) was obtained from Dow Corning (Sylgard, silicone elastomer-184). 1-Palmitoyl-2- oleoyl-sn-glycero-3-phosphocholine (POPC), 1-palmitoyl-2-oleoyl-sn-glycero-3- phosphoethanolamine (POPE), 1,2-dilauroyl-sn-glycero-3-phosphoethanolamine (DLPE), 1,2- dilauroyl-sn-glycero-3-phosphocholine (DLPC) and 1,2-dipalmitoyl-sn-glycero-3- phosphoethanolamine-n-(7-nitro-2-1,3-benzoxadiazol-4-yl) (NBD-DPPE) were purchased from Avanti Polar Lipids (Alabaster, AL). Vesicle Storage The small unilamellar vesicles (SUVs) were sized using dynamic light scattering and the average diameter was 140 nm with an average polydispersity of 0.1. Vesicles containing less than 25 mol% of PE were stored at 4 ºC. Vesicles containing 25 mol% or greater were unstable at 4 ºC and sediment to the bottom of container. Therefore they were stored at room temperature and used within 7 days of synthesis. Supported Lipid Bilayer (SLB) Formation SLBs were formed in PDMS wells on borosilicate glass (Fisher Scientific, Pittsburgh, PA). SUVs spontaneously rupture to form SLBs when incubated over clean glass surfaces in a matter of seconds. A reusable PDMS well was constructed to hold the lipid solution in place above the planar glass supports. Briefly, PDMS was mixed using 1 part crosslinker to 10 parts monomer, placed S3

4 under vacuum for 1.5 hrs, poured into a glass corral the thickness of a microscope slide, and then cured overnight. PDMS rings were then cut from the resulting sheet. They were stored on Scotch tape when not in use. The glass was first cleaned in a detergent solution of 1:7 7X (BP Biomedicals) and 18.2 MΩ water, rinsed copiously with 18.2 MΩ water, dried with compressed nitrogen gas, and finally annealed at 530 ºC for 5 hrs. This preparation rendered the glass surface hydrophilic. To adhere them to the glass, they were removed from the tape and pressed onto the cleaned planar glass substrates to form a shallow wells. To form an SLB, 75 μl of the vesicle solution was added and incubated for 20 min before rinsing excess vesicles away using 18.2 MΩ water. The supported bilayer formed in the well was then scratched lightly with tweezers to remove a section of lipid material, and then rinsed again. The scratch was subsequently used to provide contrast and aid in focusing when looking at the bilayer with a fluorescence microscope. Fitting Exponential Decay For the 80 mol% POPE bilayers that showed fluorescence decay over extended time periods, it was possible to fit the data to a single exponential to approximate an apparent diffusion constant. In this case, the time at which the fluorescence begun to decrease was set to t = 0, and the data was fit to: where y0 is the starting fluorescence level, A represents the final fluorescence level (usually slightly darker than before bleaching had occurred, due to photobleaching and immobile dyes trapped by surface inhomogeneities), and b was used to calculate the diffusion coefficient: / / where w is the diameter of the laser beam and γ = S4

5 Attenuated Total Reflection Infrared Spectroscopy Corroborating data using attenuated total reflection infrared spectroscopy (ATR-IR) was taken to confirm that the SLBs contained similar amounts of POPE and POPC to the corresponding vesicles form which they were formed. Briefly, palmitoyl chain perdeuterated POPE (d31-palmitoyl) was mixed with POPC and used to make vesicles, as described in the main text. The vesicles were then injected into a home-built multi-bounce ATR flow cell. This formed SLBs on the native oxide surface of a clean silicon wafer. IR spectra were taken before and after rinsing away the excess bulk vesicles, and the relative signal from the CD2 and CH2 stretch regions was compared against their expected ratio to determine if the supported bilayer which was formed had the intended composition. Preparation of Silicon Attenuated Total Reflection (ATR) Device Double-sided polished silicon (001) wafers (10 20 Ω cm resistivity, 500 μm thick, University Wafers) were cut to 1 cm 4.71 cm. Wafers were placed in an angled holder in order to bevel the short ends to 45. This was accomplished by using 600 grit silicon carbide sandpaper, followed by 3 μm alumina lapping film (3M) on an electric turntable with a constant flow of water. The wafers were cleaned by rinsing with ethanol, followed by 18 MΩ water. To render the surface more hydrophilic, wafers were exposed to a 3:1 mixture of piranha etchant for 60 minutes. The wafers were then placed in a custom-made flow cell machined from polycarbonate; screws held the ATR crystal and flow cell together. The flow cell was placed inside a Thermo Nicolet 470 FT-IR setup and secured to a XY-translational stage; this allows for the signal to be optimized prior to spectra S5

6 being obtained. The temperature was controlled and monitored using TEC plates and a thermocouple. Collection of ATR-FTIR Spectra The flow cell was placed in the center of the Nicolet sample cavity and then sealed so that dry nitrogen could purge the system. Samples were introduced into the cell by use of a syringe and polytetrafluoroethylene tubing. Buffer (10 mm PBS, 300 mm NaCl) was first added to the cell in order to record a background spectra. All samples were normalized to the buffer spectra. Vesicles (1 ml of 1 mg/ml) containing mixtures of POPC and palmitoyl chain perdeuterated POPE (1- palmitoyl-d31-2-oleoyl-sn-glycero-3-phosphoethanolamine, Avanti Polar Lipids) were added to the cell. Spectra was recorded after incubating for 1 hour. This is referred to as the SLB & Bulk Vesicles spectrum in Table S1. Such spectra were used to confirm that the unilamellar vesicles that were made, had the nominal lipid composition that was expected. This could be done because the contribution to the signal from the surface was small compared to the vesicles in the bulk. Next, rinsing was performed using 3 ml of buffer. After this, the bilayer was incubated for an hour and another spectrum was recorded for just the SLB in the absence of bulk vesicles, to measure the composition of the SLB alone. Data was exported from the collection software, OMNIC, and analyzed on OriginPro. Percent signal was obtained by the following equation: CD 2 %= Area under CD 2 Area under CD 2 + Area under CH 2 x 100% where "CD2 % " wass the percent signal due to CD2 stretches, "Area under CD2" was the total integrated peak area for the CD2 asymmetric and symmetric stretches, and "Area under CH2" was the total integrated peak area for the CH2 asymmetric and symmetric stretches. S6

7 Microfluidic Device Fabrication The microfluidic devices used for the nickel titration were made from PDMS and glass (cleaning procedure described above). Briefly, photolithography was used to pattern photoresist on a glass slide, which was then etched using HF and cleaned to form a master. PDMS was prepared as described above and poured over the master before being placed in the oven at 55 ºC overnight. Inlets and outlets for each channel were punched through the PDMS block using a polished needle. The PDMS was attached to a cleaned glass slide following the cleaning of both in an oxygen plasma cleaner (PDC-32G, Harrick, Pleasantville, NY) for 30 seconds. The block and slide were immediately pressed together and held in place for 1 minute, then heated for 1 minute at 100 ºC. Making Supported Lipid Monolayers For the experiments shown in Figure S2, supported lipid monolayers were used. In a microfluidic device made as described above, acetonitrile was first used to rinse the channel. Then, 3- cyanopropyldimethylchlorosilane (Acros Organics, Geel, Belgium) was directly introduced into the microfluidic channel and was incubated for 20 min. Following incubation, the channel was successively washed with acetonitrile, ethanol, and water. The device was allowed to air dry, and then vesicles made from 99.5 mol% POPC and 0.5 mol% TR-DHPE were injected. Underwater Tapping Mode Atomic Force Microscopy (UWTM-AFM) AFM images were taken with a Nanoscope IIIa Multimode SPM (Digital Instruments, Santa Barbara, CA) equipped with a type J scanner. To perform an experiment, a clean glass coverslip was mounted onto a stainless steel disk. A 1 mg/ml vesicle solution was introduced onto the glass substrate to form a bilayer. After an incubation time of 20 min., the vesicle solution was replaced S7

8 with 18.2 MΩ water or PBS buffer. Next, the steel disk with the supported bilayer covered by a dome of water was carefully placed on the J scanner. By forming a meniscus between the cantilever and the steel disk, the bilayer was kept well hydrated for at least 4-5 hours. All AFM images were obtained with 100 nm long oxide-sharpened triangular probes (silicon nitride, spring constant: 0.58 N/m) in fluid tapping mode at a scan rate of 1.25 Hz. The drive frequency was 7-9 khz and the drive amplitude was ~ 1.5 V. The only treatment applied to the images was flattening. Labeled Protein Incubation Experiments For the experiments shown in Figure S4, SLBs were first prepared in wells as described above and imaged by fluorescence microscopy. Next, bovine serum albumin (BSA) conjugated with Alexa Fluor 488 (Life Technologies, Grand Island, NY) dissolved in buffer at 5 µg/ml was introduced above the bilayer, and incubated for 15 min. Extra protein was washed away using 18.2 MΩ water, and the bilayer was imaged again under excitation wavelengths first for TR-DHPE and then for Alexa Fluor 488. S8

9 Supporting Figures Figure S1. The data from Figure 2 of the main text fit to a double exponential. A) Bilayers were formed with 0 mol% POPE, 99.5 mol% POPC and 0.5 mol% TR-DHPE. B) Bilayers were formed with 70 mol% POPE, 29.5 mol% POPC, and 0.5 mol% TR-DHPE. The solid lines are double exponential fits to the data: 1 1 where, for an SLB with two diffusing populations, A and C are the mobile fractions and b and d were used to calculate the corresponding diffusion coefficients: / / where w was the diameter of the laser beam and γ = As expected, the double exponential gave a better fit to the data than the single exponential shown in Figure 2. However, the meaning of these results was not exactly clear. For 0 mol% POPE SLBs formed at 23 C, the mobile fractions were 0.70 and 0.27, and the diffusion coefficients were 3.4 µm 2 /s and 0.4 µm 2 /s, respectively. For 70 mol% POPE, the mobile fractions and diffusion coefficients were substantially different (mobile fractions of 0.63 and 0.30, with diffusion coefficients of 2.5 µm 2 /s and 0.2 µm 2 /s, respectively). At 37 C, the mobile fractions for 0 mol% POPE were 0.68 and 0.29, with respective diffusion coefficients of 9.0 µm 2 /s and 1.0 µm 2 /s. For 70 mol% POPE, the mobile fractions were 0.71 and 0.20, with respective diffusion coefficients of 7.0 µm 2 /s and 0.9 µm 2 /s. For 70 mol% POPE bilayers at 23 C, it might be tempting to assign the two diffusing populations to the liquid-ordered and liquid-disordered phases, but this argument cannot not apply at 37 C, or for 0 mol% POPE bilayers. Moreover, it is highly unlikely that a 99.5 mol% POPC bilayer undergoes phase segregation. Indeed, no domains were observed with fluorescence S9

10 microscopy down to the diffraction limit of light (Figure S6). Moreover, fluorescence microscopy did not reveal any heterogeneity on the order of 27% in the bilayer that could account for two diffusing populations. Another possibility for two populations would be that the upper and lower leaflets in a bilayer have different diffusion coefficients. In an SLB, it is possible that any TR-DHPE in the lower leaflet, closest to the underlying glass surface, would have its diffusion hampered by interactions with the glass. To try to address this issue, data was taken with supported lipid monolayers in place of the bilayers. Figure S2 shows FRAP data taken on a 99.5 mol% POPC and 0.5 mol% TR-DHPE monolayer formed on a PDMS support that has been rendered hydrophobic by silanization as described above. The resulting FRAP curve is clearly represented better by the double exponential fit than a single exponential fit, just as the bilayers were. Therefore, it is not straight forward to assume that two populations abstracted from a double exponential fit, represent the two individual leaflets in a supported bilayer. Figure S2. FRAP data of a 99.5 mol% POPC and 0.5 mol% TR-DHPE monolayer formed on a hydrophobic support. The red line is a fit to a single exponential, and the green is a fit to a double exponential. The single exponential fit has been employed in the main text as an empirical approximation that fits reasonably well to bilayers data. Adding a second exponential mathematically improves the fit to the data. However, it would also be possible to 3, 4, or even n exponentials. In that case, each population and diffusion coefficient might be assumed to represent the different local microenvironments that fluorophores experience, influenced by slight heterogeneities in the solid support. Such an explanation seems quite plausible, but the finite signal-to-noise ratio of the FRAP experiments only allows a noticeable improvement to the fit for the addition of a second exponential. As such, we judged it best to simply fit the data with a single S10

11 exponential, since most of the standard explanations for just two distinct populations seem implausible. As such, the mobile fractions presented in Figure 3 and Figure 6 can be regarded as a lower bound to the true mobile fractions of the SLBs. Indeed, the data tend to diverge on the high side from the single exponential fits at longer time periods. Also, the diffusion coefficients should reflect an average of the diffusion of individual TR-DHPE molecules over a range of environments. Figures S3 and S4 explore the quality of SLBs formed with POPE lipids via AFM and fluorescence microscopy experiments. No holes were found under any of the conditions observed, but there was evidence for increasing amounts of three-dimensional structures as the mol% of POPE was increased. S11

12 Figure S3. UWTM-AFM images of SLBs (left) and their corresponding line profiles (right). A) 0 mol% POPE shows a flat surface, free of defects and three dimensional structures. Threedimensional structures appear at B) 10 mol% POPE, and increase in number with C) 25 mol% POPE and D) 50 mol% POPE. E) 70 mol% POPE shows the highest surface roughness, with corresponding with lipid domains. S12

13 Figure S4. SLBs incubated with Alexa Fluor 488 BSA confirmed the absence of holes (40x magnification). Indeed, following incubation with the labeled protein, the SLB fluorescence was unchanged over the solid support in regions where the bilayer was present. The Alexa Fluor 488 BSA did, however, denature on the bare glass in the diagonal scratches where the bilayer was absent. These results demonstrate that there were very few, if any holes within the SLB large enough to allow BSA to come down and denature on the glass support. Figures S5 and S6 contain fluorescence images of the POPE SLBs, and can be used to qualitatively asses the bilayers. Figure S5 specifically investigates the phase separation, and shows that the dark spots, presumed to be liquid ordered domains, are not holes in the bilayer. Figure S6 shows fluorescence images of POPE SLBs under 10x and 100x magnification. Figure S7 shows an example ATR-IR spectrum, and Table S1 summarizes the ATR-IR analysis. S13

14 Figure S5. (Top) Fluorescence image of a 70 mol% POPE SLB imaged with a 100x objective. The green box indicates the area of the SLB analyzed in the bottom image. (Bottom) Histogram of fluorescence showing three distinct regions. The blue line in the histogram indicates the fluorescence level of the scratch region, where no bilayer material is present. The large peak in the histogram reflects the intensity of the nearby bilayer, where the liquid-ordered domains are colored orange (top and bottom) and the surrounding liquid-disordered phase is grey. From the relative difference in fluorescence between the blue and orange regions, it is clear that the orange spots in the top image are liquid-ordered domains, and not holes in the bilayer. Figure S6. Images of POPE/POPC SLBs under two different magnifications (top, 10x; bottom, 100x). The dark lines are from scratches made with tweezers to remove lipid bilayer material and create contrast. S14

15 Figure S7. An ATR-IR spectrum showing an SLB with 90 mol% PEd31 and 10 mol% PC. The absorbance spectrum is referenced to a buffer spectrum, resulting in the negative water peaks on the left side of the spectrum. The two positive peaks at 2920 cm -1 and 2850 cm -1 are the CH2 asymmetric and symmetric stretches, respectively, and the two peaks at 2200 cm -1 and 2100 cm -1 are the CD2 asymmetric and symmetric stretches, respectively. The peak areas were calculated by integrating under the curves, and the asymmetric and symmetric are added together to get total CH2 and CD2 areas. The percent CD2 is then calculated, and compared to the percent CD2 expected based upon the initial vesicle composition. The theoretical % of IR signal reported in Table S1 is calculated by taking into account the all CD2 and CH2 stretches in the lipids, including head groups and tails. For example, a 70 mol% PEd31 and 30 mol% PC SLB will have 14 CD2 moieties out of a total of 32 CH2 + CD2 moieties in the lipid, multiplied by its mole fraction, resulting in a theoretical 31% of the IR signal for the CH2 and CD2 peaks % 31% S15

16 70 mol % of PE d31 Theoretical % of IR Signal IR Signal for SLB & Bulk Vesicles IR Signal for SLB 23 C 31 % 30% ± 2.5% 30% ± 3.4% 37 C 31 % 31% ± 1.8% 31% ± 3.7% 80 mol % of PE d31 Theoretical % of IR Signal IR Signal for SLB & Bulk Vesicles IR Signal for SLB 23 C 35 % 26% ± 0.8% 30% ± 0.8% 90 mol % of PE d31 Theoretical % of IR Signal IR Signal for SLB & Bulk Vesicles IR Signal for SLB 23 C 39 % 38% ± 0.9 % 32% ± 2.2 % 37 C 39 % 40% ± 3.7 % 41% ± 1.7 % Table S1. Summary of ATR-IR results for three high mole percentage POPE bilayers. The theoretical % IR signal was calculated as described above. The IR signal for SLB and bulk vesicles was calculated from spectra where an SLB has formed by spontaneous rupture of the vesicles, but the bulk vesicles were not yet rinsed away, so signal arises from both, but is dominated by the bulk vesicles. This indicates whether the vesicle-making process resulted in the expected lipid composition. The last column, IR signal for SLB, has values calculated from spectra taken after the excess bulk vesicles were washed away. This is the measure of whether the SLBs formed had the nominal concentrations of PEd31. These experiments were conducted on separate bilayers at two different temperatures. S16

17 Making 100 mol% PE SLBs Three different lipid-tethered dyes, TR-DHPE, NBD-DPPE, and 1-palmitoyl-2-{12-[(7- nitro-2-1,3-benzoxadiazol-4-yl)amino]dodecanoyl}-sn-glycero-3-phosphoethanolamine (16:0-12:0 NBD PE) at 1 mol%, were used to make vesicles that also contained 99% POPC. This included both head-labeled and tail-labeled dyes. The SLBs formed with these fluorophores resulted in little or no fluorescent material on the planar solid support even after 20 minutes of vesicles incubation. Moreover, vesicles made with tail-labeled NBD chromophores noticeably lost their yellow color upon vesicle extrusion, even when the extruder was heated above the Tm to ensure the vesicles were in the fluid phase. As such, it does not appear that any of these dyes can reside in otherwise pure POPE membranes. Figures S8 shows fluorescence images comparing SLBs formed at 23 C and 37 C for various amounts of POPE. SLBs formed at higher temperatures tended to have less visual defects and brighter levels of fluorescence. Figure S9 shows that the annealing out of detects in high mol% POPE SLBs can occur at 37 C. In fact, after 20 min, there were less bright spots on the SLBs and they appeared to be more uniform. S17

18 Figure S8. 10x fluorescent images of SLBs made with varying mol% of POPE that were formed at either 23 ºC or 37 ºC. The bilayers formed at 23 ºC had more visible defects than those formed at 37 ºC. The dark lines are scratches made by tweezers removing bilayer material to create contrast. Figure S9. 90 mol% POPE SLBs at 37 C. After rinsing, the SLB platforms were placed on a heating mantle for at least 20 min to ensure the temperature had equilibrated. Over this time period, it was possible to observe the annealing of defects, such as unruptured vesicles or uneven fluorescence, improving the quality of the SLBs. S18

19 DLPE Bilayers To further explore the effects of unsaturation in lipid tails and corroborate the results seen with POPE SLBs, we conducted FRAP experiments using DLPE, which eliminated all double bonds and had phase transition temperatures comparable to POPE (the Tm of DLPC is 29 C). 3 Generally, similar results were seen with DLPE as for POPE at 23 and 37 C (Figure 6). The SLBs maintained high mobility (Figure 6A) and diffusion coefficients (Figure 6B) with up to 50 mol% PE. Like POPE, domains were clearly visible at 70 mol% DLPE and above (Figure S10). However, the mobile fractions and diffusion coefficients revealed somewhat different DLPE behavior at room temperature compared to POPE. Specifically, there was noticeably more variation in fluorescence recovery from bilayer to bilayer for the 23 C data, even when formed from the same vesicle stock solution (Figure S11). Some recovery curves fit to a single exponential and others showed double exponential behavior (two populations recovering, one relatively fast and the other one to two orders of magnitude slower). These behaviors should be the result of phase separation in which the DLPE/DLPC SLBs could form both gel phase and liquid-ordered phase domains. 4 Gel phases were effectively immobile on the timescale of FRAP measurements, 5 resulting in only about 70% total recover, as seen in Figure S11. On the other hand, liquid-ordered phases were only an order of magnitude slower than the rest of the fluid bilayer. This resulted in the two recovering populations that were observed in the black curve in Figure S11. Interestingly, the red curve seems to show just one population. As such, there appears to be less liquid-ordered phase in this case. Finally, no tubules were observed with DLPE. To improve homogeneity and reproducibility of the SLBs, DLPE bilayers were also formed at 37 C. In this case, the DLPE bilayers behaved similarly to POPE, showing increased mobile fractions (Figure 6A, red data points), less visible defects and brighter fluorescence (Figure S12). In fact, the mobile fraction for 90 mol% DLPE bilayers was 95% at 37 C. As such, fluid SLBs S19

20 with high mobile fraction for fully saturated DLPE can be made when working near physiological temperatures. Figure S10. Images of DLPE/DLPC bilayers under 10x (top row) and 100x (bottom row) magnification taken at 23 C. The dark lines are scratches made with tweezers to remove bilayer material and provide contrast. 90 mol% DLPE at 23 C 0.8 Fluorescence Recovery SLB 1 SLB Time (s) Figure S mol% DLPE heterogeneity in FRAP curves. The two curves represent SLBs made from the same stock of vesicles on two separate pieces of glass. The noticeable difference speaks to the difficulty in making fluid, reproducible SLBs of a predominately gel-phase lipid composition. Such behavior was only seen at 23 C, and resulted in the large error bars seen from 70 to 90 mol% DLPE at room temperature in Figure 6. S20

21 Figure S12. Fluorescence images of 80 mol% DLPE SLBs formed at 23 C and 37 C. SLBs formed at 37 C were noticeably brighter, even when all experimental conditions such as the vesicles used, exposure time, contrast, etc. were all kept constant. Figure S13. Comparison of two different fluorophores at room temperature and 37 ºC in a POPC bilayer. Both probes used showed a significant increase in diffusion coefficient with the increase in temperature. S21

22 Figure S14. Fluorescence images (left) and linescans (right) from the nickel titration experiments done in 10 mm Tris buffer with 100 mm NaCl at ph 8.5. The fluorescence of the two SLBs containing 99.5 mol% POPC and 0.5 mol% TR-DHPE gave rise to only a slight decrease in fluorescence with the addition of 100 µm Ni 2+. This small change was due to collisional quenching between the fluorophore and the metal ion when higher concentrations of Ni 2+ were used. Notably, the two SLBs containing 50 mol% POPE, 49.5 mol% POPC, and 0.5 mol% TR-DHPE were considerably darker than the POPC SLBs, even without the addition of NiCl2 to the solution. This is due to a combination of factors, including quenching from putative Cu 2+ contamination, and the increase in liquid-ordered domains seen with higher amounts of PE in the bilayer (as seen in Figure 4), as TR-DHPE was excluded from the more tightly packed regions of the SLBs. With the addition of 100 µm NiCl2, there was more marked quenching of the fluorescence as Ni 2+ bound to the PE head groups. The following section details the subtraction of the contribution from collisional quenching and the fitting of the residual data to a Langmuir binding isotherm. S22

23 Subtraction of Collisional Quenching and Fitting to a Langmuir Isotherm At high concentrations of Ni 2+, the fluorescence quenching of 50 mol% POPE was due to two causes: 1. quenching from Ni 2+ -PE complexes and 2. collisional quenching from unbound Ni 2+ that interacted directly with the fluorophores. To isolate the quenching that was the result of Ni 2+ - PE complexes, the collisional contribution needed to be subtracted off from the total quenching. The quenching of 0 mol% POPE bilayers was solely the result of collisional quenching, which was independent of PE concentration, and so could be taken as the collisional quenching of 50 mol% POPE bilayers. We adapted the Stern-Volmer equation to include both quenching contributions: 1 where is the fluorescence intensity of the 50 mol% POPE bilayers with no added NiCl2, is the fluorescence intensity in the presence of NiCl2, is the quencher rate coefficient due to Ni 2+ -PE binding, is the fluorescence lifetime, and is the quencher rate coefficient due to collisional quenching. Therefore 1 which can be rearranged to give: 1 where represents the 50 mol% POPE quenching and the quenching from 0 mol% POPE bilayers. S23

24 Figure S15. Stern-Volmer plots representing the fluorescence quenching with increasing NiCl2 concentrations. The data for 0 mol% POPE were fit to a line, indicating collisional quenching. All error bars were smaller than the data points. We could then solve for and take the inverse and subtract it from 1 to get the quenched fraction due to Ni 2+ -PE binding. The plot of the binding quenched fraction vs. [Ni 2+ ] were fit to a Langmuir isotherm to calculate the value of KD: 1 Ni Ni S24

25 Supplementary Movie Movie S1. The video file was taken with fluorescence microscopy on an 80 mol% POPE SLB at room temperature under 100x magnification. It begins with the focal plane at the supported bilayer interface. After this, the focal plane is slowly adjusted upward in the z direction above the bilayer to observe the tubules. Using this procedure, it was possible to estimate that the maximum length the tubules extend above the bilayer was ~20 μm. At that point there was no longer any tubule fluorescence observed. No intentional photobleaching or FRAP was done for this movie. References (1) Brown, K. L.; Conboy, J. C., Electrostatic Induction of Lipid Asymmetry. J. Am. Chem. Soc. 2011, 133 (23), (2) Anglin, T. C.; Conboy, J. C., Kinetics and Thermodynamics of Flip-Flop in Binary Phospholipid Membranes Measured by Sum-Frequency Vibrational Spectroscopy. Biochemistry 2009, 48 (43), (3) Koynova, R.; Caffrey, M. Phases and phase transitions of the hydrated phosphatidylethanolamines. Chem. Phys. Lipids 1994, 69 (1), (4) Ratto, T. V.; Longo, M. L. Obstructed Diffusion in Phase-Separated Supported Lipid Bilayers: A Combined Atomic Force Microscopy and Fluorescence Recovery after Photobleaching Approach. Biophys. J. 2002, 83 (6), (5) Bennun, S. V.; Longo, M. L.; Faller, R. Molecular-Scale Structure in Fluid-Gel Patterned Bilayers: Stability of Interfaces and Transmembrane Distribution. Langmuir 2007, 23 (25), S25