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Antenna-Enhanced Fluorescence Correlation Spectroscopy Resolves Calcium-Mediated Lipid-Lipid-Interactions Stephan Block, Srdjan S. A#imovi#, Nils Odebo Länk, Mikael Käll, and Fredrik Höök ACS Nano, Just Accepted Manuscript • DOI: 10.1021/acsnano.7b07854 • Publication Date (Web): 12 Mar 2018 Downloaded from http://pubs.acs.org on March 13, 2018
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Antenna-Enhanced Fluorescence Correlation Spectroscopy Resolves CalciumMediated Lipid-Lipid-Interactions
Stephan Block1,2,*, Srdjan S. Aćimović1, Nils Odebo Länk1, Mikael Käll1,*, Fredrik Höök1,*
1
Department of Physics, Chalmers University of Technology, 412 96 Göteborg,
Sweden 2
Department of Chemistry and Biochemistry, Freie Universität Berlin, Berlin,
Germany.
*Corresponding authors: Stephan
Block,
e-mail:
[email protected];
Mikael
Käll,
e-mail:
[email protected]; Fredrik Höök, e-mail:
[email protected] Keywords: single-molecule fluorescence correlation spectroscopy, nanoplasmonics, lipid complexes, lipid mobility, hotspot
Author contributions SB performed the FCS experiments, developed and performed the FCS analysis scheme, and wrote a first draft of the text. SSA fabricated plasmonic antenna substrates. NOL performed numerical simulations. MK and FH conceived and supervised the project. All authors discussed and edited the text.
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Abstract Fluorescence correlation spectroscopy (FCS) has provided a wealth of information on the composition, structure, and dynamics of cell membranes. However, it has proved challenging to reach the spatial resolution required to resolve biophysical interactions at the nm-scale relevant to many crucial membrane processes. In this work, we form artificial cell membranes on dimeric, nanoplasmonic antennas, which shrink the FCS probe volume down to the ~20 nm length-scale. By analysing the autocorrelation functions (ACFs) associated with the fluorescence bursts from individual fluorescently tagged lipids moving through the antenna “hot spots”, we show that the confinement of the optical readout volume below the diffraction limit allows the temporal resolution of FCS to be increased by up to 3 orders of magnitude. Employing this high spatial and temporal resolution to probe diffusion dynamics of individual dye-conjugated lipids, we further show that lipid molecules diffuse either as single entities or as pairs in the presence of calcium ions. Removal of calcium ions by addition of the chelator EDTA almost completely removes the complex contribution, in agreement with previous theoretical predications on the role of calcium ions in mediating transient interactions between zwitterionic lipids. We envision that antenna-enhanced FCS with single-molecule burst analysis will enable to resolve a broad range of challenging membrane biophysics questions, such as stimuli-induced lipid clustering and membrane protein dynamics.
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Life depends on complex networks of precisely fine-tuned interactions between different types of biological molecules.1 Many of these processes are orchestrated by a 5 to 10 nm thin lipid bilayer, acting both as compartmentalizing hydrophobic barriers and as hosts for specialized membrane proteins that, for example, control selective molecular transport into and out of the cell.2 The emerging view is that many membrane functions depend on the detailed properties and local assembly of spatially and temporally dynamic nm-scale lipid domains,3,4 which, due to domain sizes being far below the diffraction limit of optical microscopy, remains challenging to characterize under physiological conditions. As an example, the salt-mediated formation of lipid-lipid complexes is well established from molecular simulations and has been also concluded from ensemble-averaged measurements of lipid mobility and lipid-mediated protein signalling,5-8 but a direct observation of single-molecule complex formation and dimension remains lacking. Recently, the combination of fluorescence correlation spectroscopy (FCS) with stimulated emission depletion (STED) microscopy was demonstrated,9,10 allowing the mobility of biomolecules to be studied within cell membranes on length scales of approximately 20 nm.11 To study lipid-lipid complexes, this length scale has to be reduced even further, which can be achieved by complementing FCS with plasmonic nanostructures, which provide highly confined fluorescence enhancement,12-14 yielding observation volumes down to the 10 nm scale.15,16 Herein, we present a means to study lipid movement within artificial cell membranes using plasmonic nanoantenna-enhanced FCS and explore with single-molecule resolution calcium-ion mediated lipid complex formation.
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Results and Discussion The antenna substrates, fabricated by hole-mask colloidal lithography on microscope cover-glass slides,17 consist of widely spaced (>> 1 µm) pairs of silver nanodisks (average gap distance 14.4 ± 5 nm, diameter ~80 nm, thickness ~15 nm; Figure 1a). All dimers on a substrate have the same orientation. Antenna material and morphology were chosen to yield a strong plasmonic response at both the FCS excitation wavelength (561 nm) and the fluorescence emission peak (~591 nm) of the lipid tag. This design process was guided by numerical simulations and confirmed by dark-field spectroscopy of individual antennas (Figure 1b – 1d and Supporting Information Note S1), showing a clear LSPR peak split between longitudinal and transverse excitation, that is, when the incident polarization is parallel or perpendicular to the dimer axis, respectively. The peak split is a signature of plasmonic near-field coupling between the individual nanodisks composing a dimer, which is what generates the electromagnetic hotspot in the gap region. Experimental scattering spectra (Figure 1d) were found to be in good agreement with simulations based on finite-difference time-domain (FDTD) calculations (Supporting Information Figure S1). Nanoantennas were located using laser backscattering images in buffer (Figure 1c, middle). Supported lipid bilayers (SLBs) were formed on the nanoantenna substrates using the vesicle rupture approach in presence of calcium ions (5 mM),18 which is known to mediate lipid-lipid interactions even for zwitterionic lipids.8 Confocal fluorescence imaging (Figure 1c, right) of the sample at different emission intervals (hyperspectral imaging) was used to generate spatially resolved fluorescence spectra of the SLB, peaking at 591 nm, in agreement with the known emission spectrum of lissamine-rhodamine,19 but with a peak count rate that typically increased in areas containing nanoantennas (Figure 1e). 4 ACS Paragon Plus Environment
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Figure 1: (a) Scheme of the dimeric nanoantenna (grey) coated with a supported lipid bilayer (SLB; pink) containing rhodamine-conjugated lipids (red dots). (b) The design process was guided by simulations yielding spatial distributions of the enhancement (b, top), indicating the presence of bursts in FCS intensity traces (b, bottom) in presence of nanoantennas (blue) in addition to the fluctuations of the bare SLB (red). (c) Polarized dark-field spectroscopy of bare nanoantennas in air (c, left), compared with backscattering microscopy (c, middle) and confocal imaging (c, right; 585 - 594 nm emission interval; fluorescence intensity divided by the respective intensity of a bare bilayer) of the same nanoantennas after SLB formation. A grid structure (large vertical stripes) is used to keep the same field of view. (d) Dark-field spectra typically peak close to the 591 nm emission peak of the dye. (e) Spectral imaging often shows an increased fluorescence intensity close to nanoantennas (light and dark blue circles) with respect to areas lacking nanoantennas (bare SLB; red circles), confirming sufficient spectral overlap of dye and nanoantenna. Error bars indicate standard deviations. Spectra d and e originate from the nanoantenna indicated by the green circle in panel c.
FCS measurements were recorded after SLB formation in areas containing and lacking nanoantennas on the same sample (Figure 2) using a fluorescence pass band covering 570-630 nm, which overlaps the LSPR and the dye’s emission peak. Intensity traces recorded from areas lacking nanoantennas showed fluctuations around an average count rate of 9 kHz independent of the polarization angle φ (measured with respect to the dimer long axis) of the incident laser polarization
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(Figure 2a). The corresponding autocorrelation functions (ACFs) G (τ ) were well described using a 2D model:5
G (τ ) = 1 +
1 0 1 , ⋅ g (τ ) with g 0 (τ ) = N 1 + τ /τ D
(1)
as expected for the 2D lipid diffusion within SLBs, yielding a diffusion time τ D = 5.8 ± 1.1 ms and an average number of dye-conjugated lipids in the confocal volume of N = 7.5 ± 0.7 (Supporting Information Tables S1 and S2). This corresponds to a
molecular brightness, Q , of a single lissamine-rhodamine dye of Q ≈ 1.2 kHz, being far below the saturation threshold (~50 kHz, Supporting Information Figure S2). We calibrated the effective radius w0 of the confocal volume using particles of known diffusion coefficient D and then used the relation
τD =
w02 4D
(2)
to convert τ D into a lipid diffusion coefficient of 2.3 ± 0.4 µm2/s,20 in good agreement with previous FCS studies of lipid diffusion in SLBs.20,21 Similar ACFs were observed in confocal probe volumes containing nanoantennas if the laser polarization was transversal to the antenna dimer axis ( φ = 90°; Figure 2b-c, left), indicating a lack of pronounced enhancement effects for this probe geometry. However, for longitudinal excitation ( φ = 0°), nanoantennas typically showed occasional intense fluorescence bursts superimposed on the SLB background rate of ~9 kHz (Figure 2b-c). We attribute these bursts to dyes passing through the hotspot region of an antenna, 6 ACS Paragon Plus Environment
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causing a significant rise in the dye’s net brightness due to LSPR-mediated enhancement of the fluorescence excitation, emission and/or collection efficiency. In some cases, the bursts could transiently increase the total count rate by up to two orders of magnitude. The magnitude of the bursts showed pronounced variations between different nanoantennas on the same substrate (representative examples of large and small bursts are shown in Figure 2b and 2c, respectively). This heterogeneity is likely due to variations in nanoantenna properties caused by the fabrication process, in particular variations in gap width.
Figure 2: FCS intensity traces (left and middle column) and corresponding ACFs (right column; g (τ ) = [G (τ ) − 1]/[G (0) − 1] ) of bare SLBs (a) and SLB-coated nanoantennas (b, c), recorded for transverse (left) and longitudinal excitation (middle), respectively. Insets show the corresponding intensity traces at higher magnification. Bare SLBs (a), showing no dependence of the polarization angle φ , and nanoantennas using transverse excitation (left in b and c, φ = 90°) are well described by a 1-component model (Equation (1); red solid line). However, additional bursts are observed using longitudinal excitation (middle in b and c, φ = 0°), yielding ACFs requiring a 2-component model, Equation (3), for proper description (black solid line).
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Curve-fitting of ACFs obtained from intensity traces containing pronounced burst using the single-component model, Equation (1), usually failed indicating the presence of a second component due to dye’s passing through the nanoantenna hotspot. All such ACFs were therefore fitted using a two-component model (see Supporting Information Note S2)
G (τ ) = 1 +
∗ 0 2 τD + g ( ) E g ∗ (τ ) , τ 2 τD N ⋅ 1 + Eτ D∗ / τ D
(
1
)
(3)
which contains in addition to the confocal contribution g 0 (τ ) a hotspot component
g ∗ (τ ) =
1 , 1 + τ / τ D∗
(4)
characterized by a diffusion time τ D∗ and an effective brightness enhancement factor E defined by the ratio between the dye’s average brightness in the hotspot, Q ∗ , and the non-enhanced brightness Q : E = Q ∗ / Q . When fitting Equation (3), the first component ( τ D ~5 ms) was fixed (Figure 2, red solid lines) to values determined from FCS measurements in areas lacking nanoantennas on the same sample (Figure 2a). Using these reference constraints, fitting led to diffusion times of the second component, τ D∗ , that were several orders of magnitude below τ D (Supporting Information Figure S3). Assuming that the diffusion constants characterizing the two components are the same,9 this clearly indicates hotspot volumes significantly below the diffraction limit. To quantify this, we estimated the lateral hotspot radius rhs from the measured τ D∗ values using rhs = w0 τ D∗ / τ D and plotted the so obtained rhs values 8 ACS Paragon Plus Environment
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against the corresponding enhancement factors E (Supporting Information Figure S4). For longitudinal excitation ( φ = 0°), we find a significant increase in E when the rhs values go down to approximately 20 nm, being clearly in the range of the actual
gap sizes, 14.4 ± 5 nm, of the dimers. We find this agreement remarkable, since Equation (3) is a strong simplification of the complex spatial field distribution around the antennas. Similar reductions in the readout volume were also achieved using STED-FCS,11 which offers, in contrast to antenna-enhanced FCS, the advantage that the readout volume is not limited to certain measurement points. It should be noted, however, that the dyes are subject to orders of magnitude higher optical exposures in STEDFCS than in antenna- enhanced FCS, which is for example manifested in the total intensities used in typical experiments (~10 µW for antenna-based FCS versus ~100 mW for STED-FCS in Ref. 11) or the fact that in antenna-enhanced FCS the fluorescence enhancement is strictly localized to the hotspot, such that dyes are exposed to a strong excitation only if they pass through the hotspot and thus contribute to a burst. Hence, antenna-enhanced FCS does not outperform STEDFCS with respect to the size of the readout volume, but subjects the dyes to lower optical exposures than those typically used in STED-FCS experiments, thereby providing an interesting alternative for performing FCS beyond the diffraction limit. For transversal excitation ( φ = 90°), however, most of the ACFs did not show any hotspot component (Supporting Information Figure S5). This is expected since hotspot formation between the dimers requires longitudinal excitation.15 Hence, the second ACF contribution observed at fast lag times cannot be caused by triplet dynamics within the confocal volume, which is further supported by the fact that the excitation intensity was chosen far below the value need to observe triplet dynamics 9 ACS Paragon Plus Environment
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(Supporting Information Figure S6). Furthermore, by comparing ACFs from FCS intensity traces, which were simultaneously recorded using emission intervals of 570 – 630 nm (containing the LSPR peak of most dimers) and 630 – 690 nm (being red-shifted to the LSPR peaks), it can also be shown that the second ACF component is indeed created by the nanoplasmonic hotspot but not by triplet dynamics (see Supporting Information Note S3 for detailed discussions). The fast hotspot component in the ACF allows for an interesting application of sub-diffraction FCS based on burst analysis.22-24 To extract D values using conventional FCS, the recorded intensity trace has to be at least 2 orders of magnitude longer than τ D to cover 99% of the ACF (see Equation (1)). For a single component decay with τ D around 5 ms, the intensity trace has therefore to span at least 500 ms. In contrast, as the hotspot component offers τ D∗ values on the order of few 10 µs, antenna-enhanced FCS should in principle allow for a dramatic improvement of the temporal resolution. To test this hypothesis, we checked if the entire hotspot component of the ACF could be retrieved by considering only those 10 ms intervals of an intensity trace that contained at least a single burst. Indeed, this kind of simple post-measurement data analysis, detailed in Supporting Information Note S4, resulted in the same hotspot component of the ACF as obtained by processing a complete 10 s intensity trace (Figure 3). Further, since the single bursts included in this analysis are separated by several seconds, it is obvious that they originate from independent dye-conjugated lipids, that is, the probability that bursts observed in adjacent intervals were created by the same dye molecule is practically zero. This then automatically implies that the diffusion constant of a single dyelabeled lipid molecule passing through the hotspot can be extracted. This is in contrast to conventional confocal FCS, which is indeed based on single molecule 10 ACS Paragon Plus Environment
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fluorescence data but nevertheless requires, in a typical experiment, a large number of single dye molecules to enter and exit the confocal readout volume to yield a complete autocorrelation function. For example, based on a confocal diffusion time of 5 ms, it can be calculated that more than 1000 dye-labeled lipids contributed to the intensity trace shown in Figure 2a, which leads to an ACF (Figure 2a, right) that shows far more noise at short lag times than the antenna-enhanced ACFs (Figure 3c) derived from single dye-labeled lipids passing through the hotspot. This demonstrates a clear increase in measurement resolution by using antennaenhanced in comparison to confocal FCS. Based on the notion that each burst event represents a single molecule passing through a hotspot, we constructed τ D∗ histograms for each nanoantenna offering sufficient enhancement (Figure 4). The histograms showed broad τ D∗ distributions in presence of calcium (Figure 4a,b), often composed of two, and sometimes three, equally spaced peaks reminiscent of a Gaussian distribution, that is, a first peak at ∗ ∗ the most probable value τ D,1 , a second peak at approximately 2 ⋅τ D,1 etc (Supporting
Information Figure S7). In contrast, τ D∗
distributions obtained from control
measurements or simulations (Supporting Information Figures S8 and S9) exhibit only a single peak, strongly suggesting that the observation of multiple peaks is not an artefact caused by the burst analysis.
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Figure 3: Panel a and b show a representative FCS intensity trace and the corresponding ACF (using the entire intensity trace), respectively. Complementary, ACFs were calculated (c) from 10 ms intervals of the intensity trace a that contain single bursts exceeding a userdefined threshold (a, red dashed line). Due to a fast hotspot component ( ≈ 18 µs) and high enhancement, already 10 ms intervals contain sufficient data to retrieve the entire hotspot component (numbers mark the bursts used to calculate the indicated ACFs). The τ D∗ distribution (d) shows a pronounced peak at 19 µs, close to average τ D∗ value in panel b.
We thus pooled the τ D∗ distributions obtained from different nanoantennas, which ∗ ∗ usually differ in their τ D,1 value, into a single histogram by normalizing τ D∗ to τ D,1 . The ∗ ∗ so-obtained τ D∗ / τ D,1 histogram (Figure 4c) clearly shows peaks at τ D∗ / τ D,1 = 1 and
∗ ∗ τ D∗ / τ D,1 = 2 (and an indication for a peak at τ D∗ / τ D,1 = 3 ), demonstrating that the
double peak structure is statistically robust and shared among individual ∗ nanoantennas. As τ D∗ is inversely proportional to D , Equation (2), a peak at 2 ⋅τ D,1
formally corresponds to a decrease in D by a factor of 2 with respect to the first peak. This suggests that the first peak is due to a subpopulation of lipid-conjugated 12 ACS Paragon Plus Environment
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dyes diffusing with the "confocal" D of approximately 2.5 µm2/s, while the second peak corresponds to a subpopulation diffusing with only 1.25 µm2/s. This is a strong indication for lipid oligomer formation within the SLB. A quantitative correlation between D and the area of a lipid oligomer immersed in an SLB was recently ∗ derived,5, 25 suggesting that the peak at 2 ⋅τ D,1 corresponds to complexes containing
two lipids (in the present context corresponding to one dye-conjugated lipid in complex with one non-labelled lipid; Supporting Information Figure S10).
Figure 4: (a, b) τ D∗ distributions of individual nanoantennas are composed of multiple peaks in presence of calcium ions (5 mM), but become single-peaked after addition of 10 mM of the calcium chelator EDTA (d, e; showing the same nanoantennas as in a, b), indicative for the observation of single lipids (leftmost peak) and dimeric lipid-lipid complexes diffusing through the hotspot. The same behavior is seen in histograms (c, f) pooling the data of multiple ∗ nanoantennas (after normalizing the τ D∗ axis by the diffusive time of the first peak, τ D,1 ).
Solid lines indicate Gaussian distributions.
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Indeed, after removal of calcium ions the τ D∗ distributions narrowed down and were dominated by the first peak (Figure 4d-f and Supporting Information Figure S11). This indicates a decrease of the lipid-lipid-complex fraction induced by calcium removal and thus the involvement of calcium in the formation of lipid-lipid complexes. This type of detailed information is essentially inaccessible to conventional confocal FCS since several different dye molecules will enter and leave the confocal volume during the long measurement time, leading to an ACF that is averaged over the different species. Nevertheless, as the relative occurrence of single lipids and lipid-lipid complexes was observed to be modified by calcium removal, even time-averaged ACF should show a shift to faster diffusion time τ D and higher diffusion constants D after EDTA addition. Using confocal FCS, we indeed measured an increase in D from (2.47 ± 0.13) µm2/s to (3.06 ± 0.05) µm2/s by addition of 10 mM EDTA ( n = 3 independent experiments, p < 0.01; Supporting Information Figure S12), thus confirming the trend observed in the antenna-based results.
Conclusion We have demonstrated that the combination of plasmonic nanoantenna-enhanced FCS and burst analysis allows the diffusion of individual dye-conjugated lipids to be studied with single lipid resolution. We showed that the entire hotspot ACF component can be extracted from fluorescence bursts, which allowed us to obtain the diffusion constants of single lipids entering and leaving the hotspot and to resolve lipid-lipid complexes in presence of calcium ions. We envision that nanoantennaenhanced FCS will allow challenging membrane biophysics questions to be addressed, such as the influence of mono- and bivalent salts on lipid complex
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formation in bilayers, being only one particular example of ion-specific effects (Hofmeister phenomena) often observed in nature.26
Materials and Methods Fabrication of dimeric nanoantennas. Plasmonic antenna substrates were fabricated by a variant of the standard hole-mask colloidal lithography (HCL) method in which material deposition is performed at two opposite off-normal angles to create dimer pairs.17 Clean glass slides (150 µm thickness) were spin-coated with PMMAA4 (MicroChem) and covered with a dilute layer of polystyrene beads (PS; 80 nm average diameter) with an average spacing large enough to allow for optical single particle analysis. The attachment of the PS beads is facilitated through interaction with a positively charged PPDA polymer layer (Sigma-Aldrich) pre-formed on top of the PMMA. The PS bead concentration was 0.0002% and we used a settling time of ca. 40 s before rinsing the substrates with deionized water. A 10 nm thin Cr layer was then deposited on the samples, after which the beads were removed through tape stripping. The hole-mask thus formed is transferred to the resist by oxygen plasma etching with a duration sufficient to obtain a pronounced undercut profile. Nanodisk dimers were formed by first depositing a 1 nm Ti adhesion layer at ±10° deposition angle and then depositing 15 nm of Ag at angles alternating between 10° and -10° after every 5 nm of deposited material, keeping the dimer thickness low enough to enable lipid bilayer formation across the antenna structures. Lift-off was performed in a pre-warmed remover (mr REM 400, Micro Resist technology) at 55°C and followed by an isopropanol rinsing step. The substrates were dried under a stream of nitrogen and immediately sealed in a nitrogen atmosphere inside opaque containers. The substrates were stored for a maximum of two weeks prior to any use. The final 15 ACS Paragon Plus Environment
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averaged edge-to-edge gap distance between the dimer constituents was 14.4 ± 5 nm, as determined by imaging in a scanning electron microscope. The variation mainly results from the slight polydispersity in PS bead diameter. Vesicle preparation – POPC (1-palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine), DSPE-PEG(2k) (1,2-distearoyl-sn-glycero-3-phosphoethanolamine-N-[biotinyl (poly (ethylene glycol))-2000] and rhodamine-DOPE (1,2-dioleoylsn-glycero-3-phosphoethanolamine-N-(lissamine rhodamine B sulfonyl)) were obtained from Avanti Polar Lipids Inc. (Alabaster, AL). Tris(hydroxymethyl)-aminomethane hydrochloride (TRISHCl), sodium chloride and calcium chloride were obtained from Sigma Aldrich (Steinheim, Germany). If not otherwise stated, all solutions were prepared or diluted using a TRIS-HCl buffer consisting of 100 mM Tris-HCl, 50 mM NaCl, 5 mM CaCl2 that was adjusted to pH = 7.4 using HCl. Small unilamellar vesicles (SUV) were prepared by the extrusion method as described earlier.27 In brief, lipid films were formed in round bottom flasks under flowing nitrogen and dried in vacuum, hydrated by adding 1 mL of the Tris-HCl buffer, followed extruding the mixture through polycarbonate membranes (400 nm pore size; Avanti Polar Lipids Inc., Alabaster, AL). Vesicles were based on POPC, supplemented with 0.25 mol% of PEG-DSPE- and either 0.6 mol% (for confocal imaging) or 2x10-4 mol% (for FCS) of lissamine-rhodamine-DOPE-conjugated lipids. SLB formation, confocal imaging, and FCS – All confocal imaging and FCS measurements were conducted on nanoantenna surfaces (fabricated as described above), supplemented with a home-made 10 µL PDMS well,28 using a LSM780 confocal microscope (ZEISS, Germany) equipped with a 561 nm solid state laser and a C-Apochromat 40x/1.2 water immersion FCS objective (ZEISS, Germany). All measurements were performed using a pinhole size equivalent to 1 Airy unit and laser excitation at 561 nm. SLBs were formed by injecting vesicles (lipid 16 ACS Paragon Plus Environment
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concentration 0.3 mg/mL), followed by rinsing (10 times) with Tris-HCl buffer. SLBs were based on POPC supplemented by 0.25 mol% of DSPE-PEG(2k) and either 0.6 mol% (for confocal imaging; Fig. 1d) or 2×10-4 mol% (for FCS) of lissaminerhodamine-DOPE-conjugated lipids. PEG-conjugated lipids form polymer cushions above and below the SLB, reducing the influence of substrate on lipid diffusion.29 The spatial distribution of nanoantennas on the surface was recorded using elastically backscattered light, i.e., by configuring the LSM780 in reflective mode imaging using a 20% reflective (instead of a dichroic) mirror in the optical path and a detector range of 555 – 565 nm, allowing to collect the light elastically backscattered by the nanoantennas (Fig. 1c, middle). This allowed areas containing and lacking nanoantennas to be reliably identified. After choosing a suitable area, FCS was then performed by replacing the reflective with a dichroic mirror (MBS 488/561/633; ZEISS, Germany), directing the fluorescence intensity via a user-definable monochromator to 2 GaAsP detectors, allowing intensity fluctuations to be recorded simultaneously in 2 readout channels: one ranging between 570 and 630 nm (therefore containing the LSPR and the dye’s emission peak) and one ranging between 630 and 690 nm. Raw FCS data as well as the results of two in-built hardware autocorrelators (one for each FCS read-out channel) were stored for further data analysis as described in detail in Supporting Information Notes S2 – S5. Calibration of the confocal volume was done directly after the FCS measurements by recording FCS ACFs of TetraSpeck beads (0.2 µm diameter; ThermoFisher Scientific) in buffer. Attempts to directly characterize the SLB quality within the gap region (e.g., using AFM imaging) failed so far. This is attributed to the nanoscopic size (average gap distance of ~15 nm) and the notable aspect ratio of the gap region, making a characterization of the membrane within the gap very challenging. We therefore have 17 ACS Paragon Plus Environment
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no direct information about the state of the membrane within the gap, but the indirect evidence of proper membrane formation based on its capability to provide a fluid phase is fairly convincing. In particular, antenna-based FCS measurements typically lack signs of bleaching, i.e., thousands of single intensity bursts can be recorded over extended periods. Hence, with only 2×10-4 mol% of the lipids being fluorescently labelled, continuous lipid exchange of lipids must originate from a continuous fluid bilayer with a dimension of at least 0.2 µm2, which is multiple orders of magnitude larger than the observed hotspot size. This observation would hardly be possible for defective membranes that usually show impaired fluidity such as immobile lipid fractions.29 In this respect, formation of defective membranes in the gap region will most likely not result in a sustained observation of fluorescence bursts, since lipid diffusion will be impaired and bleached dyes cannot be efficiently exchanged by nonbleached ones. Further, for each experiment the field of view was imaged using fluorescence microscopy in order to rule out that the observation of large count rates is caused by the accidental characterization of an unruptured, fluorescent vesicle located above the gap region. Spectral imaging of the samples (Fig. 1c, right) used the same optical setup as FCS, but employed only a single GaAsP detector in combination 4.5 nm thin detection interval created by the monochromator. To record a spectral stack, the same field of view was imaged multiple times using a detection interval that was shifted between 569 and 659 nm in 4.5 nm steps. Note that due to the differences in sensitivity, spectral imaging and FCS cannot be performed on the same sample: for FCS a dye concentration on the order of 2x10-4 mol% was sufficient to create sufficient signal to noise ratio (SNR), while spectral imaging required dye concentrations as high as 0.6 mol% to yield a sufficient SNR over the entire spectral stack. 18 ACS Paragon Plus Environment
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Since the polarization axis was defined by the polarization of the excitation laser in the LSM780, the sample had to be rotated within the plane of the sample stage to choose between longitudinal and transversal excitation of the nanoantennas. This process was supported by grid structures (enclosed area 100 x 100 µm2) and the random nanoantenna distribution, allowing individual nanoantennas to be identified again, even after sample rotation, buffer exchange, or using different measurement approaches. Data analysis and statistics – All data analysis was done using home-made scripts written in MATLAB (MathWorks, Natick, MA) and described in detail in Supporting Information Notes S1 – S5. In total, FCS measurements from 54 individual nanoantennas were included in this study, performed on 10 independently functionalized nanoantenna substrates. For each nanoantenna, at least 6 FCS traces (each covering 10 s of measurement time) were recorded using 2 independent detectors allowing to simultaneously collect photons emitted within the 2 independent emission intervals (570 – 630 nm and 630 – 690 nm). Hence, for each nanoantenna and each readout channel, at least 6 ACF were calculated, and used to determine average value ± standard deviations of the fitting parameter of Eq. (1) – (4) presented in Supporting Information Figures S2 + S4 and Tables S1 + S2.
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Associated Content Supporting Information The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsnano.xxx. Details on the simulation of antenna-enhanced FCS (Note S1, Figure S1). FCS count rate per dye versus excitation intensity (Figure S2). Details on the analysis of 2-component autocorrelation functions and on the extracted parameters (Note S2, Figure S3 – S5, and Table S1 and S2). Details on how to distinguish hotspot contributions and triplet dynamics (Note S3, Figure S6). Details of the burst analysis (Note S4 and S5 and Figure S7 - S10). Temporal evolution of complex dissolution upon EDTA addition (Figure S11). Impact of calcium removal on confocal FCS measurements (Figure S12). (PDF)
Author Information Corresponding Authors *E-mail:
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Acknowledgements This work was funded by the Knut and Alice Wallenberg Foundation (grant “Optical nanoantennas shine light on the nanoworld”), the Swedish Research Council (via Linneaus center SUPRA), the Focus Area Nanoscale (Freie Universtät Berlin), and the German Science Foundation (BL1514/1). We acknowledge the Centre for Cellular Imaging at the University of Gothenburg and the National Microscopy Infrastructure, NMI (VR-RFI 2016-00968) for providing assistance in microscopy. 20 ACS Paragon Plus Environment
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TOC-Image
for the manuscript
Antenna-enhanced Fluorescence Correlation Spectroscopy Resolves Calciummediated Lipid-lipid-interactions
Stephan Block1,2,*, Srdjan S. Aćimović1, Nils Odebo Länk1, Mikael Käll1,*, Fredrik Höök1,*
1
Department of Physics, Chalmers University of Technology, 412 96 Göteborg,
Sweden 2
Department of Chemistry and Biochemistry, Freie Universität Berlin, Berlin,
Germany.
*Corresponding authors: Stephan
Block,
e-mail:
[email protected];
Mikael
Käll,
[email protected]; Fredrik Höök, e-mail:
[email protected] 23 ACS Paragon Plus Environment
e-mail:
