Nucleic acid nanostructure sensor device

Abstract

Nanostructure sensor devices are provided that are capable of measuring mechanical properties associated with semifluid membranes. The nanoscale structures comprise a probing tip that can apply a force to the membrane to induce a deformation. The determination of the amount of deformation of the semifluid membrane can be converted into a measurement of a mechanical property associated with the semifluid membrane. The membranes may be synthetic or comprised within biological systems such as vesicles or cells. Methods for using the nanostructure sensors are also provided.

Description

[0001] NUCLEIC ACID NANOSTRUCTURE SENSOR DEVICE

[0002] FIELD

[0003] The present invention relates to novel nucleic acid nanostructures and their uses. In particular, it relates to nucleic acid nanodevices that can act as mechanical sensors of membrane dynamics for lipid or synthetic bilayer type membranes.

[0004] BACKGROUND

[0005] DNA nanostructures have shown potential to advance nanotechnology and the life sciences. Compared to other materials, DNA nanostructures have a highly controllable architecture which is based on predictable folding using base-pairing rules (Rothemund P. W. Nature 440, 297-302 (2006), Seeman, N. C.; Sleiman, H. F. Nat. Rev. Mater. (2017), 3, 17068; Hong, F. et al. Chem. Rev. 2017, 117, 12584-12640; Praetorius, F. et al. Nature (2017), 552, 84-87; Sacca, B.; Niemeyer, C. M. Angew. Chem. Int. Ed. 2012, 51 , 58-66). By exploiting these properties, functional DNA nanostructures are increasingly designed to benefit areas outside DNA nanotechnology. Examples include DNA scaffolds which precisely position proteins and other biomolecular components for research applications in biophysics and molecular biology. Furthermore, predictable changes in DNA nanostructures have been exploited as smart biosensing devices which measure pH inside cells (Bhatia, D.; et al. Nat. Commun. (2011), 2, 339) or in cellular DNA nanocages for delivery of bioactive cargo (Walsh, A. S.; et al. ACS Nano (201 1), 5, 5427-5432).

[0006] Physical materials, including cells, may be defined by their mechanical properties such as elastic resistance of substances that make up their surface. Atomic Force Microscopy (AFM) has emerged as a powerful technique for quantifying cell membrane elasticity through direct mechanical probing at the nanoscale. The method employs a sharp cantilever tip that applies controlled forces to the membrane surface while measuring the resulting deformation with subnanometer precision. The force-indentation curves generated during this process provide direct measurements of membrane mechanical properties. Young's modulus is calculated from the slope of the linear region of the force-displacement curve using contact mechanics models such as Hertz, Sneddon, or Johnson-Kendall-Roberts (JKR) theory, depending on the tip geometry and adhesion characteristics. Modern AFM systems can detect forces in the piconewton range with spatial resolution of 1-10 nanometers, enabling measurement of local variations in membrane elasticity across individual cells. The technique can quantify Young's modulus values typically ranging from 0.1 to 100 kPa for biological membranes, while also providing information about membrane tension, adhesion forces, and viscoelastic properties through dynamic measurements. AFM has been successfully applied to study membrane elasticity changes during cellular processes such as differentiation, drug treatment, and disease progression. However, limitations include potential cellular damage from tip contact, influence of cytoskeletal structures on measurements, and the need for careful model selection for data interpretation. The technique also requires specialized sample preparation and environmental control for live cell measurements and is limited to cell surface measurements, meaning that intracellular membrane measurements are out of scope.

[0007] There is a need to further develop further improved and optimized methods and compositions for quantifying lipid bilayer-type membrane elasticity through direct mechanical probing at the nanoscale. The present invention addresses the deficiencies in the art. These and other uses, features and advantages of the invention should be apparent to those skilled in the art from the teachings provided herein.

[0008] SUMMARY

[0009] According to the present disclosure, the inventors have provided a molecular machine (MM) in the form of a sensor nanodevice that enables measurement of lipid bilayer membrane dynamics. The MM can be used in in a massively parallel fashion on synthetic membranes, on vesicles, liposomes, lipid nanoparticles, as well as inside living cells.

[0010] In a first aspect the invention provides for a nucleic acid nanostructure comprising: a base module, wherein the base module is configured to adjoin a semifluid membrane that defines a surface; and a beam module, wherein the beam module is connected to the base module via one or more flexible hinge sequences such that the beam module is able to translate through a range of motion between a first and a second conformation relative to the base module; wherein translation of the beam module from the first to the second conformation causes the nucleic acid nanostructure to apply a force to the surface of the semifluid membrane.

[0011] A second aspect provides method for measuring a mechanical property associated with a semifluid membrane, wherein the semifluid membrane defines substantially planar surface, the method comprising the steps of: attaching a nanoscale structure to the semifluid membrane surface, wherein the nanoscale structure comprises a probe tip, and wherein the nanoscale structure is configured to transition through a range of motion from a first conformation to a second conformation upon application of an actuation trigger, and wherein in the second conformation the probe tip applies a load force to the surface of the membrane sufficient to induce deformation of the membrane; and determining the amount of deformation of the membrane, converting the determination of the amount of deformation into a measurement of a mechanical property associated with the semifluid membrane.

[0012] Further aspects of the invention provide for compositions comprising one or more nanostructures as defined herein, a sensor nanodevice comprising a nanostructure as defined herein, and a membrane elasticity sensor nanodevice as defined herein.

[0013] In other aspects of the invention, uses of (or methods for using) a nanostructure, a composition, a sensor nanodevice or a membrane elasticity sensor nanodevice as defined herein are provided for a method for diagnosis or prognosis of a pathological condition within a human or animal subject.

[0014] In yet further aspects of the invention, uses of (or methods for using) a nanostructure, a composition, a sensor nanodevice or a membrane elasticity sensor nanodevice as defined herein are provided for a method for assessing the effect of a xenobiotic compound or substance on a cell, cell culture or membrane containing cellular extract.

[0015] Within the scope of this application it is expressly intended that the various aspects, embodiments, examples and alternatives set out in the preceding paragraphs, in the claims and / or in the following description and drawings, and in particular the individual features thereof, may be taken independently or in any combination. That is, all embodiments and / or features of any embodiment can be combined in any way and / or combination, unless such features are incompatible.

[0016] BRIEF DESCRIPTION OF THE DRAWINGS

[0017] One or more embodiments of the invention will now be described, by way of example only, with reference to the accompanying drawings, in which:

[0018] Figure 1. Molecular Machine (MM) sensor nanodevices measures lipid bilayer nanomechanics in a massively parallel fashion and inside cells, a, The base units of the MM binds with cholesterol anchors (yellow) to bilayer membranes. Hybridization of opening strands actuates sensor cantilever to push its tip downward and locally indent the membrane. The elastic membrane deformation is measured via a fluorescent FRET readout, b, sensor indentation allows parallel quantification of local membrane elasticities, whereas the opening tilt angle is larger for softer (top) membranes than for stiffer membranes (bottom), c, In cells, the sensor nanodevice measures the elasticity of the plasma membrane (dark grey) as well as internal membranes (lighter grey islands), d, 2D connectivity map of the MM sensor nanodevice designed with CaDNAno. 5' and 3' termini of the DNA strands are represented by a square and triangle, respectively. The duplexes are numbered on the left side.

[0019] Figure 2. 3D model of the sensor nanodevice and its dimensions. The 3D model was generated with Blender and is shown in views from the a) front b) side c) top and bottom. Sensor nanodevice consists of a base, a beam, and a distal tip. The cylinders represent DNA duplexes, which are interconnected by single strands (not shown). Smaller cylinders at the bottom indicate cholesterol anchors for membrane binding of the sensor nanodevice to a semifluid membrane bilayer. The dark oval and the light oval dots annotate the location for the incorporated FRET fluorophores such as Cy5 and Cy3 fluorophores, respectively.

[0020] Figure 3. 2D duplex map of sensor nanodevice MM, its tip and base with numbered component duplexes in top view. Each circle represents a double helix within the MM. The numbering of duplexes is the same as in Figure 1d. a) Top view of the beam of sensor. The dark dark oval on duplex 23 indicates the Cy3 dye. b) Horizontally cut section of the distal tip. c) Top view of the base. The dark and light circles indicate the locations of the hairpin and singlestranded actuator sequences, respectively, and mid-grey the position of the single-stranded hinge sequences, respectively. Actuator and hinge sequences connect the beam and the base (not shown). Actuator sequences in positions 4, 5,14, 15, 24, and 25 are shorter in length than those of position 0, 1 , 18, 19, 20, and 21 . The former hybridize with short opening strands 7-12 while the latter bind to long opening strands 1-6 (see Table 5).

[0021] Figure 4. Scheme on the reversible opening and closing of sensor nanodevice MM. a) The addition of 12 oligonucleotide opening strands (orange) opens MM while the addition of 12 closing oligonucleotides (brown) closes the MM to yield side product duplexes, b) Detailed mechanism of the DNA-triggered opening and closure of MM exemplified by a single actuator hairpin located between beam (top) and base (bottom). The single-stranded opening strand binds to the dumbbell-like hairpin of complementary sequence to unzip the hairpin and form an energetically favourable duplex which pushes the beam and base apart to open MM. To revert the opening of MM to the closed state, a closing strand binds to a single-stranded toehold section of the MM- bound opening strand to unzip it from the hairpin sequence. This reforms the hairpin and reverts MM to the closed conformation. A new duplex between complementary opening and closing strands is formed as a side product.

[0022] Figure 5. Triggered reversible opening of the sensor nanodevice MM is tracked by fluorescence resonance energy transfer (FRET), a) Scheme of MM carrying FRET donor dye Cy3 (light grey) and acceptor dye Cy5 (dark grey) at the base and beam, respectively as distance markers. MM opening and closing by the addition of opening and closing strands alters the distance between Cy3 and Cy5 and causes FRET-induced changes in fluorescence emission. In the closed conformation, both fluorophores are within proximity, and excitation of Cy3 leads to FRET and Cy5 emission. In the open state, the two fluorophores are apart and FRET between Cy3 and Cy5 is weakened as the dye distance exceeds the Forster distance Ro. b) Plot displaying the distance dependency of the FRET efficiency for Cy3 and Cy5. An increase in the distance between the two fluorophores leads to a decrease in FRET efficiency. The Forster distance Ro is defined as the distance between two fluorophores to allow 50% FRET efficiency and is 5 nm for the FRET pair Cy3 Cy5.

[0023] Figure 6. Gel electrophoretic analysis on the formation and purification of sensor nanodevice MM. 1 % agarose gel in 0.5x TAE, 10 mM MgCh at 65 V for 1 h separating PhiX174 scaffold, unpurified folded MM and PEG-purified MM. The upward mobility shift for the assembled MM band relative to the scaffold band indicates successful assembly; a higher-migrating band represent a dimeric MM. The fast migrating, intense band of small staple strands, which had been added in 6x excess for MM assembly, were removed by precipitation with PEG, as confirmed by the disappearance of the gel band for staple strands.

[0024] Figure 7. Transmission electron microscopy images of negatively closed sensor nanodevice MM in the closed state. Scale bar at bottom right, 20 nm.

[0025] Figure 8. Transmission electron microscopic (TEM) analysis of negatively stained sensor nanodevice MM in the closed conformation in front view, a) Seventy TEM micrographs used for the alignment, b) Averaged image after alignment and c) standard deviation between the aligned images. Scalebar, 20 nm.

[0026] Figure 9. TEM analysis of negatively stained sensor nanodevice MM in the closed conformation in top view, a) Twenty TEM micrographs used for the alignment, b) Mean image after alignment and c) standard deviation between the aligned images. Scalebar, 20 nm.

[0027] Figure 10. Dimensions of sensor nanodevice MM obtained by analysis of TEM images, a) Mean TEM images after MM alignment from the front and top view with marked points for dimension measurements, b) Plot displaying the measured dimensions for MM. Lowercase numbers indicate the two points used for distance measurements, n = 70. c) Model of MM in front and top view with the mean dimensions obtained by TEM analysis.

[0028] Figure 11. oxDNA simulations of sensor nanodevice MM. a) Closed and b) opened MM shown in front (left) and side view (right) with oxDNA-derived dimensions, the angle between the beam and base are annotated in dashed lines. The shading in the MM structures represent the RMSF noise as indicated by the scale.

[0029] Figure 12. Thermal fluctuation analysis of sensor nanodevice MM. Cando analysis of MM in a) front, b) side, and c) bottom view of MM indicating high stability of the MM core with a minimal root mean square fluctuation (RMSF) of 0.1 nm. The bottom of the base and tip exhibit higher flexibility due to single-stranded anchoring strands and the relatively low number of bundled duplexed, respectively.

[0030] Figure 13. Actuated opening and closing of sensor nanodevice MM is tracked with fluorescence readout, a, Reversible actuation of MM by opening and closing strands changes the MM’s tilt angle and the associated fluorescence signals of Cy3 and Cy5 distance markers, b, MM opening and closing changes MM’s gel electrophoretic migration, c, oxDNA simulations of MM’s open and closed state (top) and image-averaged negative stain TEM images of MM in both states (bottom), scalebar 20 nm. d, Analysis of MM tilt angles from TEM images (n = 70 and n = 51) for closed and opened state, respectively), e, Cy3 and Cy5 kinetic emission traces tracking MM closing and subsequent opening, f, Maximum change of Cy3 and Cy5 emission upon MM opening and closing from kinetic traces (as in e); mean ± s.d from n = 4. One-way ANOVA with Tukey’s multiple comparisons test at ****P value < 0.0001 . g, Gel electrophoretic analysis of MM opening at various temperatures and reversible MM opening and closing, g, 2% Agarose gel run in 0.5 x TBE, 10 mM MgCh for 2.5 h at 65 V confirms the opening of MM at temperatures from 25 °C to 45 °C, as indicated by the gel band shift occurring for closed MM when mixed with opening strands (OS), h, 2.5% Agarose gel run for 3 h at 65 V on the reversible opening and closing of MM in 1 x PBS as indicated by repeated upward and downward gel shifts. Some MM aggregation is promoted at high agarose gel concentration. A line parallel to the wells aids as a visual guide to highlight the gel shift. A 1 kb DNA ladder at the left is a size reference.

[0031] Figure 14. TEM images of negatively stained sensor nanodevice MM in the open state. Scale bar 50 nm.

[0032] Figure 15. TEM analysis of negatively stained sensor nanodevice MM in the opened conformation showing the side view of MM. a) thirty-six TEM micrographs used for the alignment, b) Mean image after alignment and c) standard deviation between the aligned images. Scalebar, 20 nm.

[0033] Figure 16. Analysis of MM opening angle from TEM images, a) Front view of the 3D model of MM in closed and opened confirmations, indicating the corresponding opening angles (dotted lines) as obtained by TEM analysis, b) Scatter plot of the opening angles between base and beam obtained from TEM images of MM in closed and opened state.

[0034] Figure 17. Fluorescent emission spectra of Cy3 / Cy5-labelled sensor nanodevice MM with FRET dependent signatures for the open and closed state. Spectra of closed MM (dark grey) and opened MM (light grey) using Cy3 excitation at 540 nm show emission peaks at 574 nm (Cy3) and 664 nm (Cy5). Increased emission at 664 nm is a signature for the closed MM state while increased emission at 574 nm is characteristic for the open MM state. Figure 18. MM position and chemical structure of the cholesterol-tag used for membrane anchoring of MM. a) Part of the 2D DNA connectivity map of MM’s base unit. The arrows indicate the 3' and square the 5' terminus of staple and anchoring strands, b) Chemical diagram of cholesterol with hydrophobic and hydrophilic parts, c) Lipid anchoring strand comprising cholesterol attached via a flexible tetra(ethylene glycol) (TEG) linker to the DNA sequence.

[0035] Figure 19. DLS analysis on the size distribution of DOPC and POPC single unilamellar vesicles extruded at 200 nm.

[0036] Figure 20. Microscopic and gel electrophoretic analysis of sensor nanodevice MM binding to unilamellar vesicles, a) 1 % Agarose gel (0.5 x TAE 10 mM MgCI2) of MM without and with cholesterol membrane anchors; MM+chol (2.5 pM) was incubated with small unilamellar vesicles (SUVs, average diameter 200 nm) made with 50 to 300 pM DOPC or POPC phospholipid final concentration. The gel band upshift of MM+chol is caused by the interaction of cholesterol tags with the gel matrix while the upshift of MM+chol incubated with SUVs is caused by successful MM binding to SUVs which are too large to migrate into the gel. b) Plot showing MM binding extent in dependence of SUVs concentration measured as lipid concentration. The data summarise the binding gel binding analysis as in panel a (n = 3). Fitting the graph with a Langmuir isotherm yielded Kd values for MM binding of 22.0 + / - 2.2 pM for DOPC and 24.0 + / - 3.0 pM for POPC vesicles. A 1 kb DNA ladder was included as a reference.

[0037] Figure 21 Sensor nanodevice MM indents lipid bilayers and reports on their localised elastic deformation by fluorescence lifetime imaging microscopy (FLIM). a, MM indentation of stiff POCP and soft DOPC membranes results in different tilt angles, different Cy5 emission levels, and Cy3 fluorophore lifetimes T, as indicated, b, % Kinetic Cy5 emission traces for MM bound to DOPC and POPC vesicles upon addition of OS as indicated by a grey line, in 50 mM HEPES 500 mM NaCI. c, Magnitude of Cy5 emission at the endpoint of kinetic traces of MM opening, as in b; mean ± SD from n = 6; unpaired T-test at **P < 0.05. d, Fast-FLIM microscopic image of DOPC GUVs decorated with closed MM. scalebar 50 pm e, FastFLIM images of MM decorated GUVs composed of DOPC, POPC, and phase-separated 40 / 40 / 20 DOPC / DPPC / Cholesterol. Green and magenta arrows indicate Ld and Lo phases, respectively. The FLIM image pixels are coloured for T (Cy3). Scale bar 20 pm. Microscopic FastFLIM images reports the average photon arrival time while FLIM lifetime values reported as numbers are derived from lifetime histograms, f, Bar chart on the relative change in Cy3 lifetime following OS addition showing mean ± SD of n = 10; one-way ANOVA with Tukey’s multiple comparison test at ****P <0.0001 .

[0038] Figure 22. Purification of sensor nanodevice MM by size exclusion chromatography (SEC) and gel electrophoresis analysis of SEC fractions, a) SEC elution profile for MM and component staple strands using absorption at 260 nm. The x-axis provides the names of the SEC fractions, b) 1 % Agarose gel run for 1 h at 65 V showing the SEC fractions from panel a, and as a reference, scaffold strand PhiX174 and PEG-purified MM. Fractions B9 and B10 contain purified MM without staple strands.

[0039] Figure 23. Gel electrophoresis analysis on the stability and reversible opening of MM in 1 x PBS. a) 1 % Agarose gel run (65 V, 1 h) of MM incubated in 1x PBS for up to 60 min. b) 2.5% Agarose gel (65 V, 3 h) on the reversible opening and closing of MM in 1 x PBS. Reversible opening and closing for two cycles are indicated by upward and downward shifts. The visible aggregation of the MM in the wells is due to a high agarose concentration.

[0040] Figure 24. Non-linear Gaussian fit of the DLS analysis on the size distribution of giant unilamellar vesicles (GUV) composed of DOPC (left peak) and POPC (right peak).

[0041] Figure 25. Kinetic trace of Cy5 fluorescence emission on sensor nanodevice MM opening in 1x PBS. Normalised fluorescent Cy5 trace showing the opening of MM in 1x PBS buffer by adding to closed MM opening strands in 10x excess at 5 min.

[0042] Figure 26. Scheme for measuring membrane elasticities by FRET and FLIM and estimation on the number of sensor nanodevice MM bound to GUVs. a) POPC vesicles with stiff membrane results in a low degree of indentation and MM opening, hence proximity of the fluorophore pair Cy3 / Cy5, high FRET efficiency, and low lifetime (T) which can be measured by FLIM. By contrast, DOPC vesicles with soft membrane allows for more membrane indentation, a higher degree of MM opening, greater separation of the Cy3 / Cy5 pair, low FRET efficiency, and a high lifetime. The different lifetimes T of the cells are being measured and then represented in a colourmap with red / yellow indicating high lifetimes and green / blue low lifetimes, b) FastFLIM microscopic image of DOPC GUVs decorated with closed MM showing a field of view of 590 x 590 pm2. To the GUVs, 9*1 O10MM (30 pL, 5 nM, diluted to 120 pL) were added. Assuming complete binding and a z-section of approximately 300 nm height yields 6*104MM per field of view. The foreground pixels account for 8% of the image leading to a final concentration of 8* 105MM on the membranes in the field of view. This calculation actually underestimates the amount of bound MM, as out-of-focus MM are also binding to the membranes and MM distribution is not homogenous.

[0043] Figure 27. Average Cy3 lifetime and lifetime decay curves of sensor nanodevice MM bound to GUVs. Average Cy3 lifetimes ± SD (top panel) of a) Cy3 / Cy5-MM bound to DOPC GUVs, b) POPC GUVs, and c) biphasic GUVs with disordered DOPC and ordered DPPC CL domains, and with opening strands (+OS), and after sequential addition of closing strands (+CS). The bottom panels show the lifetime decay curves with lifetime components TXand T2. The average lifetime is derived from Ai * T±and A2 * T2where A1 and A2 are weighting factors in the lifetime distribution. n = 10 GUVs analysed per condition, across three independent experiments. One-way ANOVA, Tukey’s multiple comparison test, < 0.0001 ; **P = 0.0055; *p = 0.0211 ; ns = not significant. *

[0044] Figure 28. Cy3 lifetime changes of GUV-bound sensor nanodevice MM upon actuation reveal differences in membrane elasticities. Plot on the average Cy3 lifetime changes for MM actuation from Figure 27. Datapoints are from n = 10 GUVs per condition, across three independent experiments, and show the average and SD.

[0045] Figure 29. FLIM analysis Cy3-labeled sensor nanodevice MM bound to DOPC GUVs a) Fast- FLIM image of a DOPC GUV incubated with Cy3-MM. The pixels are color-coded according to lifetime (scale 0-2 ns) and have a corresponding point in the phasor plot. Scale bar, 10 pm. b) Phasor plot of Cy3-MM, overlay of n = 10 fields of view, c) The average lifetime of Cy3-MM from phasor analysis is 3.4 ns ± 0.1 , n= 10.

[0046] Figure 30. Phasor analysis of FLIM read-out of Cy3 / Cy5-sensor nanodevice MM bound to vesicles, a) Schematic diagrams of the phasor plot showing that the phasor dot position on the universal circle line indicates the fluorescence decay lifetime. Furthermore, the position of the phasor dot on the universal circle line indicates fewer lifetime components and no FRET while a position towards the center indicates more lifetime components and FRET. A component point represents in the phasor plot represents an image pixel in the FLIM image, b) Cy3 lifetime of MM in GUVs composed of DOPC, POPC and DOPC / DPPC CL. The data is shown for MM, the addition of opening strands (+OS), and closing strands (+CS). Panels show graphical 2D phasor FLIM distributions of n = 10 GUVs (overlaid). The bottom panels for DOPC and POPC show the merge phasor plots for MM, + OS, + CS. The bottom panels in biphasic illustrate phasor points of DOPC and DPPC CL phases merged.

[0047] Figure 31. Microscopic analysis of sensor nanodevice MM preferentially binding to liquid disordered rich region, a) Schematic for biphasic vesicles. DOPC and Liss Rhodamine PE form the liquid-disordered phase, DPPC and cholesterol form the liquid-ordered phase, b) Confocal microscopy images of Cy5-tagged MM bound to GUVs composed of DOPC / RhPE / DPPC / CL GUV (1 % Rh), imaged under the Cy5 and Rhodamine channels, respectively, c) Bar chart shows mean membrane fluorescence intensity ± SD of DOPC and DPPC CL phases. N > 10 GUVs analysed across three independent experiments. Unpaired t-test, ****P <0.0001.

[0048] Figure 32. Rapid endocytosis of sensor nanodevice MM in live HeLa cells confirmed by costaining with lysosomes and late endosomes, a) Fluorescence confocal microscopy images of Cy3-tagged MM in live HeLa cells stained with green Lysotracker MM. For the MM / Lysotracker image, the MM is displayed in red, lysotracker is shown in blue. Purple indicates an overlay of MM and lysotracker. Scale bar, 10 pm. b) Average Pearson correlation coefficient (PCC) ± SD of MM to Lysotracker. Figure 33. Dynamin-inhibitor Dynasore increases the dwell time of sensor nanodevice MM at the plasma membrane. Fluorescence confocal microscopy images of Cy3-tagged MM without the addition of Dynasore and after 30 min incubation with Dynasore (50 pM). Scale bar, 10 pm.

[0049] Figure 34. FLIM analysis of the sensor nanodevice MM in live HeLa cells establishes bilayer regions of varying stiffness, a, Schematic of a HeLa cell with bound MM to the stiffer outer plasma membrane (magenta) and softer internal membrane (green) as found in FLIM analysis, b, Intensity and Fast-FLIM images of MM in live HeLa cells and upon addition of opening strands (+OS). Plasma membrane (PM) and internal (INT) membrane regions are labelled. The pixels are coloured according to the lifetime (scale 0 - 2ns) and have a corresponding point in the phasor plot in Supplementary Fig. 34. Scale bar, 10 pm. c, The line plot displays the mean lifetimes ± SD calculated by phasor analysis in the internal membrane ROI (INT)and the plasma membrane region of interest (PM). One-way ANOVA, Tukey’s multiple comparison test, ****P <0.0001 ; n = 10 fields of view GUVs per condition, across three independent experiments. FLIM lifetime analysis of MM-Cy3 bound to cells, d) Fast-FLIM image of live HeLa cells incubated with MM carrying only Cy3 but no Cy5. The pixels are coloured according to lifetime (scale 2-4 ns) and have a corresponding point in the phasor plot. Scale bar, 10 pm. e) Phasor plot of MM-Cy3, overlay of n = 15 fields of view, f) Lifetime of MM-Cy3 from phasor analysis.

[0050] Figure 35. Fast-FLIM images, corresponding phasor plots, and lifetime decays of sensor nanodevice MM bound to live HeLa cells reveal distinct lifetimes from the plasma membrane (PM) and internal vesicles (INT). a) Scheme for measuring membrane elasticities of different membrane regions via Fast-FLIM. b) Fast-FLIM images from Figure 34 (scale 0-2) and corresponding phasor plots for MM + opening strands obtained from drawing ROIs on the PM and in INT of cells; overlay of n = 10 fields of view for each condition. Scale bar, 10 pm. b) Lifetime decay curves (average of n = 10 from three independent experiments) of closed MM and following the addition of OS in PM and INT regions, c) Schematic diagram of phasor plot demonstrating how the phasor point moves with FRET and the addition of OS.

[0051] Figure 36. FRET trajectories reveal a higher FRET efficiency and number of FRET donors for sensor nanodevice MM bound at the plasma membrane (PM) than internal (INT) cell membranes, a) FRET trajectories drawn from the MM-Cy3 data to the quenched and FRET- active MM data and the MM + opening strands data in the PM and INT. A circle is placed on each data to determine the number of donors in FRET and the FRET efficiency. Autofluorescence is considered, b) Scatter plot shows results from the FRET trajectory indicating the FRET efficiency and the number of donors with FRET (%). Data shows the overlay of n = 10 fields of view for each condition from three independent experiments. Figure 37. FLIM lifetime images of sensor nanodevice MM-bound to HeLa cells, a) FLIM lifetime images of MM-Cy3 with Cy3 only as a control to determine the maximum unquenched lifetime, b) FLIM lifetime images of closed Cy3 Cy5 plus addition of opening strands. Scalebar, 20pm.

[0052] Figure 38. Sensor nanodevice MM deformation in lipid bilayers is confirmed by FliptR. a) Schematic diagram of FliptR lifetime shifting from green to red as order in the membrane increases, b) FLIM-fit images of FliptR within GUV membranes (Ctrl) and upon the addition of MM, opening strands (OS), and closing strands (CS). The images are colour-coded for 12. For reasons of clarity, the microscopy images of biphasic GUV are arranged to show the DOPC domains on the left side of the GUV, and DPPC / CL on the right side. Scale bar, 20 pm. c) Line plot showing the response of FliptR T2 to above conditions for MM bound to GUVs. The values represent the average ± SD for n > 10 GUVs per condition, across three independent experiments. Separate plots with the calculated statistical significance for each lifetime change upon addition are shown in Supplementary Figure 43. d) Bar chart on the calculated relative changes in T2 (%) following addition of OS. One-way ANOVA, Tukey’s multiple comparison test, ****P <0.0001 ; n > 10.

[0053] Figure 39. FliptR fluorescence lifetime in GUVs for control experiments using sensor nanodevice MM. a) Bar chart of mean T2 ± SD upon the addition of MM without any cholesterol tags (MM (-CL)) (n = 10) to GUVs composed of DOPC. One-way ANOVA, Tukey’s multiple comparison test, *****p < 0.0001 ; ns = not significant, b) Bar chart showing mean T2 ± SD upon the addition of CS to MM in a closed conformation (MM (closed)) to GUVs composed of DOPC and POPC. Ordinary one-way ANOVA, Tukey’s multiple comparison test, ns = not significant.

[0054] Figure 40. Response of FliptR fluorescence lifetime upon addition of sensor nanodevice MM to GUVs of different membrane composition and elasticities, a) Bar charts show mean T2 ± SD in GUVs composed of DOPC (n = 13) and upon addition of MM (n = 15), OS (n = 13), CS (n = 12). b) Bar charts show mean T2 ± SD in GUVs composed of POPC (n = 13) and upon addition of MM (n = 18), OS (n = 15), CS (n = 12). c) Bar charts show mean T2 ± SD in GUVs composed of Ld and d) Lo. (n = 10) and upon addition of MM (n = 18), OS (n = 13), CS (n = 11). a-c) One-way ANOVA, Tukey’s multiple comparison test, *****p < 0.0001 ; ns = not significant.

[0055] Figure 41. FliptR-based calibration, a) Model employed to estimate the FlipTR calibration factor t;, showing the “spherical cap”-like deformation induced by the sensor nanodevice MM. b) Plot showing how depends on the estimated membrane coverage. The black arrow indicates the estimated coverage for our system. The corresponding FlipTR calibration value lies between the ones reported by c) Schematic illustration of the footprint of the sensor nanodevice MM distal tip which touches the membrane surface. The dark discs indicate the DNA duplexes of the distal tip. The lighter grey disc describes the size and radius of the tip model used for the calibration calculations.

[0056] DETAILED DESCRIPTION

[0057] Prior to setting forth the invention, a number of definitions are provided that will assist in the understanding of the invention.

[0058] Unless otherwise indicated, the practice of the present invention employs conventional techniques of chemistry, molecular biology, microbiology, recombinant DNA technology, and chemical methods, which are within the capabilities of a person of ordinary skill in the art. Such techniques are also explained in the literature, for example, M.R. Green, J. Sambrook, 2012, Molecular Cloning: A Laboratory Manual, Fourth Edition, Books 1-3, Cold Spring Harbor Laboratory Press, Cold Spring Harbor, NY; Ausubel, F. M. et al. (Current Protocols in Molecular Biology, John Wiley & Sons, Online ISSN:1934-3647); B. Roe, J. Crabtree, and A. Kahn, 1996, DNA Isolation and Sequencing: Essential Techniques, John Wiley & Sons; J. M. Polak and James O'D. McGee, 1990, In Situ Hybridisation: Principles and Practice, Oxford University Press; M. J. Gait (Editor), 1984, Oligonucleotide Synthesis: A Practical Approach, IRL Press; and D. M. J. Lilley and J. E. Dahlberg, 1992, Methods of Enzymology: DNA Structure Part A: Synthesis and Physical Analysis of DNA Methods in Enzymology, Academic Press; Synthetic Biology, Part A, Methods in Enzymology, Edited by Chris Voigt, Volume 497, pages 2-662 (2011); Synthetic Biology, Part B, Computer Aided Design and DNA Assembly, Methods in Enzymology, Edited by Christopher Voigt, Volume 498, Pages 2-500 (2011). Each of these general texts is herein incorporated by reference.

[0059] As used herein, the term ‘comprising’ means any of the recited elements are necessarily included and other elements may optionally be included as well. ‘Consisting essentially of’ means any recited elements are necessarily included, elements that would materially affect the basic and novel characteristics of the listed elements are excluded, and other elements may optionally be included. ‘Consisting of’ means that all elements other than those listed are excluded. Embodiments defined by each of these terms are within the scope of this invention.

[0060] The term ‘nucleic acid’ refers to a sequence of nucleotides that may be single or double-stranded, with the 3' and 5' ends of each nucleotide connected by phosphodiester bonds. These polynucleotides may consist of either deoxyribonucleotide or ribonucleotide bases. Nucleic acids include DNA and RNA, which are typically produced synthetically but can also be extracted from natural sources. Additionally, nucleic acids may involve modified forms of DNA or RNA, such as those that have been methylated or chemically altered, for example, 5’-capping with 7- methylguanosine or its analogues, 3’-processing like cleavage and polyadenylation, splicing, or labelling with fluorophores or other substances. Nucleic acids may also include synthetic nucleic acids (XNA) or nucleic acid analogues, such as hexitol nucleic acid (HNA), cyclohexene nucleic acid (CeNA), threose nucleic acid (TNA), glycerol nucleic acid (GNA), locked nucleic acid (LNA) and peptide nucleic acid (PNA). Hence, where the terms ‘DNA’ and ‘RNA’ are used herein it should be understood that these terms are not limited to only include naturally occurring nucleotides. Sizes of nucleic acids, also referred to herein as ‘polynucleotides’ are typically expressed as the number of base pairs (bp) for double stranded polynucleotides, or in the case of single stranded polynucleotides as the number of nucleotides (nt). One thousand bp or nt equal a kilobase (kb). Polynucleotides of less than around 100 nucleotides in length are typically called ‘oligonucleotides’.

[0061] Polynucleotides or oligonucleotides may be functionalized by, for example, using a modified phosphoramidite in the strand synthesis reaction. Enzymic modification using a terminal transferase can also be used to incorporate an oligonucleotide, which incorporates a modification such as an anchor, to the 3’ of a single stranded nucleic acid (e.g. ssDNA). The polynucleotides or oligonucleotides of the disclosure may be modified to have phosphorothioate (PS) linkage in the backbone to increase resistance to nuclease degradation, or to have 2'-O-Methyl (2'-OMe) and 2'-0-Methoxyethyl (2'-MOE) modifications on the 2' position of the ribose sugar in RNA to improve stability against ribonucleases (enzymes that degrade RNA).

[0062] As used herein, the terms ‘3" (‘3 prime’) and ‘5" (‘5 prime’) take their usual meanings in the art, i.e. to distinguish the ends of polynucleotides. A polynucleotide has a 5' and a 3' end and polynucleotide sequences are conventionally written in a 5' to 3' direction. The term ‘complements of a polynucleotide molecule’ denotes a polynucleotide molecule having a complementary base sequence and reverse orientation as compared to a reference sequence.

[0063] The term ‘duplex’ is used herein refers to double-stranded nucleic acid hybridised molecules, such as DNA (dsDNA), meaning that the nucleotides of two complimentary DNA sequences have bonded together and then coiled to form a double helix (assuming A-, B- orZ-form), or also singlestranded RNA (ssRNA) that has annealed to a complimentary DNA sequence to generate an RNA-DNA hybrid (RDH) duplex.

[0064] According to the present invention, homology to the nucleic acid sequences described herein is not limited simply to 100%, 99%, 98%, 97%, 95% or even 90% sequence identity. Many nucleic acid sequences can demonstrate biochemical equivalence to each other despite having apparently low sequence identity. In the present invention homologous nucleic acid sequences are considered to be those that will hybridise to each other under conditions of low stringency (Sambrook J. et al, Molecular Cloning: a Laboratory Manual, Cold Spring Harbor Press, Cold Spring Harbor, NY). However, it may be desired in some cases to distinguish between two sequences which can hybridise to each other but contain some mismatches - an “inexact match”, “imperfect match”, or “inexact complementarity” - and two sequences which can hybridise to each other with no mismatches - an “exact match”, “perfect match”, or “exact complementarity”. Further, possible degrees of mismatch are considered.

[0065] As used herein, the term ‘nanostructure’ refers to a geometrically predefined or ‘predesigned’ two- or three-dimensional molecular structure typically comprised from a biopolymer, suitably a naturally or non-naturally occurring nucleic acid or a polypeptide, which structure has at least one dimension or an aspect of its geometry that is within the nanoscale (i.e. 10-9metres). Nanoscale structures suitably have dimensions or geometry of less than around 100 nm, typically less than around 50 nm, and most suitably around 20 nm. Nanoscale structures suitably possess dimensions or geometry greater than around 0.1 nm, typically greater than around 1 nm, and optionally greater than around 2 nm. A nanostructure according to an embodiment of the present disclosure is able to associate with, or bind to, a synthetic or cellular or sub-cellular membrane or to a microsomal or exosomal structure within a tissue sample obtained from a subject. In an embodiment of the invention, the nanostructure associates with a membrane via insertion of a least one associated hydrophobic anchor moiety into the membrane bilayer. According to this embodiment of the invention a majority of the nanostructure is localised and optionally anchored to a surface of the membrane but does not penetrate or puncture the membrane other than through the intercalation of membrane anchors into the lipid or synthetic polymer bilayer.

[0066] Assembly of nucleic acid-based nanostructures may occur spontaneously in solution, such as by heating and cooling a mixture of DNA strands of preselected sequences, or may require presence of additional co-factors including, but not limited to, nucleic acid scaffolds, nucleic acid aptamers, nucleic acid staples, co-enzymes, and molecular chaperones. Where desired nanostructures result from one or more predesigned spontaneously self-folding nucleic acid molecules, such as DNA or RNA, this is typically referred to as nucleic acid ‘origami’. Rational design and folding of DNA to create two dimensional or three-dimensional nanoscale structures and shapes is known in the art (e.g. Rothemund (2006) Nature 440, 297-302). An embodiment of nucleic acid rational design may be DNA origami, wherein single-stranded DNA molecule is self-assembled into specific shapes by the complementary binding of multiple short "staple" strands. In embodiments of the present disclosure, scaffold strands may be based around well-known ssDNA sources such as the bacteriophage genomes of phiX174; lambda; M13mp18 and variants thereof; M13mp19; f1 ; fd; and P1 , for instance. Alternative synthetic sources of ssDNA are also available and will be known to the skilled person.

[0067] The nucleic acid sequences that form the nucleic acid nanostructures will typically be manufactured synthetically, although they may also be obtained by conventional recombinant nucleic acid techniques. DNA constructs comprising the required sequences may be comprised within vectors grown within a microbial host organism (such as E. coli). This would allow for large quantities of DNA or RNA to be prepared within a bioreactor and then harvested using conventional techniques. The vectors may be isolated, purified to remove extraneous material, with the desired DNA sequences excised by restriction endonucleases and isolated, such as by using chromatographic or electrophoretic separation.

[0068] The term ‘modular’ as used herein refers to the use of one or more units, or modules, to design or construct a whole or part of a larger complex nanostructure. In the context of the present invention it refers to the use of individual component modules, sub-units or building blocks to construct a nanostructure, suitably a nanostructure configured to interact with and bind to a semifluid or lipid bilayer membrane. The modules may be each the same or the modules may be different. To form the nanostructure, the individual modules are constructed so as to include assembly interface regions that facilitate assembly into a larger nanostructure complex via complementary base pairing at the assembly interfaces or via linkage using hinge strands of nucleic acids.

[0069] As used herein the term ‘hydrophobic’ refers to a molecule having apolar character including organic molecules and polymers. Examples are saturated or unsaturated hydrocarbons. The molecule may have amphipathic properties. As used herein, the term ‘amphipathic’ describes a molecule with both hydrophobic and hydrophilic regions.

[0070] As used herein, the term ‘hydrophobically-modified’ relates to the modification (joining, bonding or otherwise linking) of a polynucleotide strand with one or more hydrophobic moieties. A ‘hydrophobic moiety’ as defined herein is a hydrophobic organic molecule and may be synonymous with the term ‘lipophilic’ indicating the molecule has an affinity for lipids and particularly the lipid core of a membrane bilayer. The hydrophobic moiety may be any moiety comprising non-polar or low polarity aliphatic, aliphatic-aromatic or aromatic chains. Suitably, the hydrophobic moieties utilised in the present invention encompass molecules such as long chain carbocyclic molecules, polymers, block co-polymers, and lipids. The term ‘lipids’ as defined herein relates to fatty acids and their derivatives (including tri-, di-, monoglycerides, and phospholipids), as well as sterol-containing metabolites such as cholesterol. The hydrophobic moieties comprised within the embodiments of the present invention are capable of forming non-covalent attractive interactions with amphipathic semifluid membranes or phospholipid bilayers, such as the lipid- based membranes of cells and act as membrane anchors for the nanostructure. According to certain embodiments of the present invention suitable hydrophobic moieties, such as lipid molecules, possessing membrane anchoring properties may include sterols (including cholesterol, derivatives of cholesterol, phytosterol, ergosterol and bile acid), alkylated phenols (including methylated phenols and tocopherols), flavones (including flavanone containing compounds such as 6-hydroxyflavone), saturated and unsaturated fatty acids (including derivatives such as lauric, oleic, linoleic and palmitic acids), and synthetic lipid molecules (including dodecyl-beta-D-glucoside). The anchors for the polymer membrane may be the same as for lipid bilayers or they may be different. The specific hydrophobic moiety anchor may be selected based on the binding performance of the membrane chosen. In embodiments of the invention the disclosed nanostructures may comprise one or more hydrophobic or lipophilic anchors that act to attach or connect or anchor the nucleic acid nanostructure (i.e. the sensor nanodevice) to a generally hydrophobic membrane such as a semifluid or lipid bilayer of a cell, organelle or vesicle. The lipid anchors are attached to the nanostructure, typically the base module, or comprised within any other associated stabilising or functional modules that form part of overall the nanostructure. Suitably attachment is via oligonucleotides that carry the lipid anchor, suitably cholesterol, at the 5' or 3' terminus. In embodiments of the invention the base module comprises a plurality of anchors. Polynucleotides or oligonucleotides may be functionalized using a modified phosphoramidite in the strand synthesis reaction, which is easily compatible for the addition of reactive groups, such as cholesterol and lipids, or attachment groups including thiol and biotin. Enzymic modification using a terminal transferase can also be used to incorporate an oligonucleotide, which incorporates a modification such as an anchor, to the 3’ of a single stranded nucleic acid (e.g. ssDNA). These lipid modified anchor strands may hybridize via ‘adaptor’ oligonucleotides to corresponding sections of the nucleic acid sequence forming the scaffold section of the nanostructure. Alternatively, the lipid anchors are assembled with the nanostructure using lipid-modified oligonucleotides that contribute as either the scaffold or staple strands. A combination of approaches to anchoring using two or more membrane anchors may also be adopted wherein anchors are incorporated into one or all of a scaffold strand, a staple strand and an adaptor oligonucleotide. Cholesterol has been found to be a particularly suitable lipid for use as an anchor in the present invention (see Figure 1 (a)). The use of other lipids as anchors is contemplated, although it may be expected that there is a particular preference for a particular lipid, and a given number of membrane anchors, for a given membrane chemistry.

[0071] The term ‘membrane’ in the context of this application refers to a thin layer or barrier that separates two environments and selectively prevents the movement of molecular substances in solution, such as ions, small molecules, peptides, sugars, proteins and nucleic acids, between them. The membrane may of biological origin (i.e. from a cell or subcellular compartment) or comprise biological materials, synthetic materials, or a combination of these. A membrane may comprise a plurality of molecules that are charged and hydrophilic at one end, and hydrophobic and non-polar at the other end - also described as being amphipathic. In embodiments of the invention, the plurality of molecules of a membrane are all substantially amphipathic. Biological material suitable for a membrane composition may typically include lipids, specifically phospholipids. A membrane may be comprised of lipids, typically phospholipids and cholesterol. Examples of phospholipids include phosphatidylcholine (PC), phosphatidylethanolamine (PE), phosphatidylserine (PS), phosphatidylinositol (PI) or phosphatidylglycerol (PG). Examples of PC may be 1 ,2-Dimyristoyl-sn-glycero-3-phosphocholine (DMPC), 1 ,2-dioleoyl-sn-glycero-3- phosphocholine (DOPC), 1-Palmitoyl-2-oleoyl-sn-glycero-3-phosphocholine (POPC), or 1 ,2- diphytanoyl-sn-glycero-3-phosphocholine (DPhPC). An example of phosphatidylglycerol may be 1 ,2-Dimyristoyl-sn-glycero-3-phosphoglycerol (DMPG). An example of phosphatidylinositol may be phosphatidylinositol 4,5-bisphosphate (PIP2). Biological membrane material may also typically comprise cholesterol. A membrane may additionally comprise proteins, protein complexes and glycol proteins suitably embedded within or anchored to a surface of a membrane.

[0072] The membrane may also, or alternatively, comprise synthetic materials such as a synthetic polymer. In embodiments, the synthetic polymers may be synthetic lipid analogues. The synthetic polymer membrane may include a block copolymer, a di-block copolymer, a tri-block copolymer, a terpolymer, an alternating copolymer, or a combination thereof. In some embodiments, the polymer membrane is made of tri-block polymers comprising poly 2-methyl-2-oxazoline (PMOXA) or polydimethylsiloxane (PDMS). In embodiments, the tri-block polymer comprising PMOXA and PDMS can be organised into alternating layers of hydrophilic PMOXA and hydrophobic PDMS, for example, PMOXA-PDMS-PMOXA. Hydrophilic PMOXA layers can interact with water, while the hydrophobic PDMS layers provide stability between the two PMOXA layers. In some embodiments, each block of polymers may be longer than a single unit. For example, membranes used can include PMOXA6-PDMS35-PMOXA6; PMOXA6-PDMS65-PMOXA6 or PMOXA11- PDMS65-PMOXA11 . In a different embodiment, the polymer membrane is made of di-block polymers comprising poly(1 ,2-butadiene)-b-polyethylene oxide (PBD-PEO), with hydrophobic PBD and hydrophilic PEO components. The membrane may be a hybrid membrane comprising both synthetic polymers and lipids. In some embodiments, a hybrid membrane may comprise 1 :1 PBD-PEO and DPhPC. In some embodiments, a hybrid giant unilamellar vesicle comprising both lipid and polymer components may be generated. There may be more than one type of lipid and polymer respectively. A polymer membrane may comprise synthetic lipid analogues. In an embodiment, the polymer membrane is made of poly (ethylene oxide)-block-poly (butylene oxide) (PEO-b-PBO) block copolymers. PEO-b-PBO is an amphipathic molecule with hydrophilic PEO and hydrophobic PBO.

[0073] A membrane may be in a form of a variety of macromolecular structures, such as micelles, inverted micelles, monolayers, bilayers, polymersomes, or other lamellar structures. Lamellar structures may include a giant unilamellar vesicle (GUV) and a large unilamellar vesicle. In a micelle, the polar head groups are positioned on the outer surface, while the hydrocarbon chains are oriented towards the interior. Typically, the terminal methyl groups of the fatty acyl chains are located at the centre of the hydrophobic core, which can take the form of a sphere, ellipsoid, or cylinder. A lipid bilayer is characterised by orientation of the polar head groups facing the exterior surfaces in an aqueous environment, and the hydrophobic hydrocarbon chains in the interior of the bilayer. Other membranes may be comprised within lipid or lipidoid nanoparticles (LNPs).

[0074] Lipid bilayers tend to close in on themselves, preventing the exposure of hydrophobic aliphatic chains to water, which leads to the formation of a compartment enclosed by lipids. A lamellar structure refers to a layered arrangement of lipid, copolymer or synthetic lipid analogues. In an embodiment, a lamellar structure may include a unilamellar structure with a single layer of lipid, copolymer of synthetic lipid analogues. In an embodiment, a unilamellar structure may include vesicle or liposome referring to a structure made up of lipid bilayers enclosing an aqueous compartment. In an embodiment, a lamellar structure may be a multilamellar structure comprising multiple layers of lipid, copolymer or synthetic lipid analogues. In an embodiment, a vesicle-like structure made from amphipathic block copolymers are referred to polymersomes. Polymersomes typically comprise a bilayer membrane that encloses an aqueous core.

[0075] In embodiments of the disclosure, the lipid bilayer defines a planar structure having a first exterior facing surface and a second interior facing surface. In such arrangements the lipid bilayer may serve as a partition between aqueous compartments within a cell, or between the cell and its surroundings, or within a composition, or within a device that comprises a membrane. The term ‘surface’ should be understood to indicate the phase transition boundary between the lipid bilayer and the external, typically aqueous, environment.

[0076] In one embodiment of the present disclosure the nanostructures described herein interact with and adhere to a first exterior facing surface. In a further embodiment of the present disclosure the nanostructures described herein interact with and adhere to a second interior facing surface. In alternative embodiments of the present disclosure the nanostructures described herein interact with both first and second surfaces substantially simultaneously.

[0077] The term ‘fluidity’ in the context of a membrane is defined as the degree in which individual molecules of the lipid membrane are free to rotate and move in lateral directions. The fluidity of a lipid membrane can be determined by several factors including composition of the membrane such as the ratio of saturated to unsaturated fatty acids. High proportions of phospholipids with unsaturated fatty acids typically increases the fluidity of the membrane by preventing tight packing of the fatty acids using its kinked tailed. Cholesterol may impact the fluidity of the membrane by restricting the movement of phospholipid fatty acid chains. The lipid membrane comprising both cholesterol and phospholipids may exhibit a ‘semi-fluidity’, as the mosaic structure of the lipid components maintains structural organisation while at least some lipid molecules, typically phospholipids, can move laterally.

[0078] As used herein, the term ‘polymer’ comprises repeating structural units or monomers that are chemically bonded typically in a chain structure. The term ‘copolymer’ refers to a polymer comprising two or more different monomers that are bonded in the same polymer chain. Copolymers can be classified into various types, including a ‘random copolymer’ wherein the monomers are distributed randomly along the chain, an ‘alternating copolymer’ wherein the different monomers alternate in a regular pattern, and a ‘block copolymer’ wherein two or more chemically distinct polymer segments, or ‘blocks’ are covalently bonded. The term ‘di-block polymer’ is a type of block copolymer that consists of two distinct polymer segments, or blocks.

[0079] The term ‘terpolymer’ refers to a polymer having three units that are different from each other.

[0080] As used herein, the term ‘membrane vesicle’ refers to a substantially spherical membrane structure that encloses a fluid-filled interior compartment.

[0081] As used herein a ‘sensor’ device, as used herein, refers to a detector that responds to changes physical parameters of a specific target by producing a measurable and / or quantifiable output signal. In the context of the present disclosure, a sensor device comprises a nanoscale structure fabricated from nucleic acid, such as by using DNA origami techniques, wherein nucleic acid strands are precisely folded and assembled into predetermined three-dimensional configurations. The sensor operates by undergoing conformational changes, or other detectable molecular events upon contact with actuator molecules, thereby generating a corresponding output that can be measured through various detection modalities including, but not limited to, fluorescence, electrical conductivity, plasmon resonance, magnetic resonance, detection of mechanical displacement, or quantum / optical properties. The measurable output provides a direct or indirect indication of the properties of one or more target membranes, enabling quantitative or qualitative analysis of chemical, biological samples or environments.

[0082] The term 'sensor nanodevice' as used herein refers to a molecular machine comprised of nucleic acid nanostructure modules that cooperate to provide a sensor function. In the specific context of cellular membrane elasticity determination, a sensor nanodevice of the present disclosure may operate as a mechanical probe capable of detecting and quantifying the physical properties of synthetic membranes, vesicles, LNPs, cellular membranes and / or sub-cellular organellar structures through direct interaction. The nucleic acid nanostructure modules are engineered to possess defined mechanical properties, including predetermined stiffness, flexibility, and forceresponse characteristics, enabling the sensor nanodevice to apply a mechanical stress to a target membrane surface while simultaneously enabling measurement of the resulting deformation or resistance. The cooperative function of the modules comprised within the sensor nanodevice can allow for single or multiple time point measurements or even real-time monitoring of membrane properties through measurable changes in the sensor nanodevice's conformation. The sensor nanodevice may incorporate multiple functional domains, including membrane-targeting sequences or moieties for specific cellular or organellar localization, force-generating elements that apply mechanical pressure upon actuation, and reporter functionalities that transduce mechanical deformation into detectable signals such as fluorescence changes, conformational switches, or binding state alterations. Through this integrated molecular architecture, the sensor nanodevice provides quantitative assessment of a range of membrane parameters, such as elasticity including Young's modulus, membrane tension, and viscoelastic properties, thereby enabling precise characterization of cellular mechanical properties at the nanoscale level. In embodiments of the present disclosure, the quantitative assessment of membrane property parameters, such as elasticity including Young's modulus, membrane tension, and viscoelastic properties, is achieved through the sensor nanodevices acting as nanomolecular probes. This affords the ability to apply precisely controlled mechanical forces and measure the corresponding membrane responses at the molecular level. Young's modulus determination, for instance, comprises actuation of the sensor nanodevice to initiate the application of a defined normal force perpendicular to a membrane surface while measuring the resulting elastic deformation of that surface. The modulus calculated as the ratio of applied stress to the measured strain within the linear elastic region of the membrane's mechanical response. The sensor nanodevice allows quantification of membrane tension by detecting the resistance to lateral deformation when tangential forces are applied across the membrane plane, providing measurements of the membrane's in-plane stress state and its capacity to resist stretching or compression. Viscoelastic property assessment encompasses both the elastic (recoverable) and viscous (time-dependent) components of membrane behaviour, wherein the sensor nanodevice can applies oscillatory or step-function mechanical stimuli allowing a temporal response characteristics, including stress relaxation, creep behaviour, and frequency-dependent modulus variations to be measured. The nucleic acid architecture of the sensor nanodevice enables real-time monitoring of these parameters through measurable conformational changes that correlate with specific mechanical responses. Advanced measurement capabilities include determination of membrane bending rigidity through detection of curvature-induced conformational changes, assessment of lateral heterogeneity in mechanical properties across different membrane domains, and characterization of temperature-dependent viscoelastic transitions that reflect changes in lipid phase behaviour and membrane organization.

[0083] In specific embodiments of the present disclosure a sensor nanodevice is used to elucidate information on lipid-bilayer type, as well as mechanical properties such as membrane indentation dynamics and elasticity. The sensor nanodevice of one embodiment is configured to comprise a movable module that may be actuated to bear upon and apply a force, in the form of an applied load, to a surface of an adjacent membrane. The angular rotation, or ‘tilt angle’ or ‘tilting motion’ of the movable module may be converted into a detectable signal, such as via fluorescence. In one embodiment, to measure an angular rotation (ortilt angle) the distance-reporting fluorophores Cy3 and Cy5 are positioned at a sensor nanodevice's static base module and movable beam, respectively, so that fluorescence resonance energy transfer (FRET) can be used to track changes in fluorophore distance occurring during sensor nanodevice actuation. FRET operates on the principle of non-radiative energy transfer between a donor fluorophore (Cy3) and an acceptor fluorophore (Cy5), where the efficiency of energy transfer is inversely proportional to the sixth power of the distance between the fluorophores, making it exquisitely sensitive to nanometer-scale distance changes. When the sensor nanodevice is in its relaxed state, the Cy3 and Cy5 fluorophores are positioned at a baseline distance that produces a characteristic FRET efficiency and corresponding fluorescence intensity ratio. Upon actuation, membrane contact is initiated and there is a subsequent tilting of the nanodevice beam module to bear upon the membrane surface. In response to this application of force there is an elastic resistance exerted by the membrane. At equilibrium the spatial separation between the fluorophores changes proportionally to the degree of angular deflection of the beam module, resulting in measurable alterations in FRET efficiency. Small conformational changes in the nanostructure, on the order of 1-10 nanometers, translate into significant changes in FRET signal due to the distance dependence, enabling detection of subtle mechanical deformations that would otherwise be below the resolution limit of conventional optical techniques, or even of other approaches such as atomic force microscopy (AFM). The ratiometric nature of FRET measurements provides inherent correction for variations in fluorophore concentration, excitation intensity, and photobleaching effects, while the real-time monitoring capability allows for dynamic tracking of membrane elasticity changes, such as during cellular processes or in response to cellular stimuli. The positioning of fluorophores at strategic locations within the sensor nanodevice architecture ensures that the FRET signal specifically reports on the mechanical parameter of interest (tilt angle) while minimizing interference from other conformational changes, thereby providing a direct and quantitative readout of membrane indentation depth and the corresponding elastic response forces.

[0084] Alternative FRET arrangements suitable for implementation in the sensor nanodevice, according to embodiments of the present disclosure, may include various fluorophore pairs that provide distance-dependent energy transfer capabilities comparable with conventional Cy3 / Cy5 systems of the type described herein. Alexa Fluor dye pairs, such as Alexa Fluor 488 / 555 or Alexa Fluor 555 / 647, offer superior photostability and reduced photobleaching while maintaining comparable FRET efficiency and spectral characteristics suitable for real-time monitoring of nanostructure conformational changes. Quantum dot-based FRET systems utilizing semiconductor nanocrystals as donors paired with organic acceptor fluorophores may provide enhanced brightness, size-tunable emission properties, and extended operational lifetimes, enabling prolonged measurement periods without signal degradation. Lanthanide-based FRET arrangements incorporating terbium or europium complexes as donor species offer the advantage of long-lived luminescence that permits time-gated detection, thereby eliminating background fluorescence and autofluorescence interference from biological samples. Additionally, ATTO dye series and DyLight fluorophore pairs may provide effective alternatives with high extinction coefficients and optimized spectral overlap integral values that enhance FRET sensitivity to distance changes. The selection of specific FRET pairs may be tailored to particular end-use sensor requirements, including desired spectral range, photostability demands, detection sensitivity, and compatibility with biological environments, while maintaining the fundamental distance-reporting capability essential for quantifying conformational change of the sensor nanodevice during membrane elasticity measurements. In other embodiments of the present disclosure small conformational changes in the sensor nanodevice nanostructure may be detected and quantified using one or more alternative signal generating technologies. In an embodiment, single-molecule force spectroscopy (SMFS) using optical tweezers or magnetic tweezers represents a highly sensitive approach where the sensor nanodevice may be tethered between a membrane surface and a solid-state substrate (such as a trapped microsphere or chip). In this embodiment, direct measurement of force-extension relationships with piconewton resolution and sub-nanometer displacement sensitivity are facilitated. This technique favours detection of conformational changes in the sensor nanodevice through fluctuations in mechanical force due to elastic resistance of the membrane.

[0085] In a further embodiment of the disclosure plasmonic coupling-based detection utilizes gold nanoparticles or plasmonic nanostructures integrated into the nucleic acid sensor nanodevice, where mechanical deformation alters the plasmonic coupling between nanoparticles, resulting in measurable shifts in surface plasmon resonance frequencies or scattering intensities. The nearfield coupling is highly distance-dependent, providing excellent sensitivity to nanometer-scale structural changes thereby providing a direct and quantitative readout of membrane indentation depth and the corresponding elastic response forces. Plasmonic coupling offers particular advantages for nucleic acid nanostructures due to its compatibility with aqueous biological environments, high sensitivity, and potential for multiplexed detection.

[0086] In other embodiments of the present disclosure electrochemical sensing of mechanical deformations of the sensor nanodevice may be detected and quantified through the use of redoxactive labels. This approach comprises incorporating one or more redox-active moieties (such as methylene blue or ferrocene) at specific positions within the nanostructure of the sensor nanodevice, where conformational changes alter the electron transfer efficiency to an adjacently placed electrode surface. The resulting changes in electrical current provide quantitative readout of structural deformation of the sensor nanodevice with high temporal resolution.

[0087] In a further embodiment of the disclosure quantum dot-based photonic rulers employ semiconductor quantum dots with size-dependent emission properties positioned within the nanostructure of the sensor nanodevice. In this embodiment mechanical stress induces changes in quantum confinement or inter-dot coupling, leading to measurable shifts in photoluminescence spectra or quantum yield. The signal output may be used to providing a direct and quantitative readout of membrane indentation depth and the corresponding elastic response forces.

[0088] In yet a further embodiment of the disclosure mechanical resonance detection may be utilized. In this approach the sensor nanodevice itself may function as a nanomechanical resonator, such that conformational changes in the sensor nanodevice alter the resonant frequency, which can be detected through integrated piezoresistive elements or optical interferometry with femtometerscale displacement sensitivity. The sensor nanodevices of embodiments of the present disclosure are provided with an actuatable hinge region that can be activated via a binding interaction with an actuation trigger. The hinge region is located between the base module and the beam module and acts as a mechanism for facilitating the transition of the nanostructure from at least a first conformation (termed “closed”) to at least a second conformation (termed “open”) whereby the beam bears upon and deforms an adjacent semifluid membrane. It will be appreciated that depending upon the precise configuration of the sensor nanodevice, there may be one or more intermediate transitional conformations that exist between the first and second conformations. In a specific embodiment of the invention, the trigger mechanism is comprised of a region of at least one single stranded nucleic acid, more typically a plurality, within a linking structure between the adjacent base and beam modules. This region of single stranded nucleic acid lacks the rigidity of a double helix and contributes to the properties of flexure required by the hinge. This region, is nonetheless readily accessible to the surrounding solution such that actuation triggers in solution may bind to the flexible hinge sequence(s), thereby providing the trigger, and lead to a change in its structural conformation. The flexible hinge region may comprise a plurality of sequences that vary from single stranded to incorporating small hairpin loops. Precise configuration of the hinge region will control the range of movement of the beam module relative to the base module. The change in structural conformation leads to initiation of rotation about a fulcrum point on the leading edge of the base module causing the beam module as a whole to rotate in an arc about the fulcrum point from first to second conformation. For example, in one embodiment of the invention the actuation trigger comprises a single stranded nucleic acid present in solution which is capable of hybridising fully or partially to a sequence comprised within the hinge region. The binding of the one or more actuation trigger nucleic acid molecules to the hinge sequence results in hybridisation and formation of the conventional double helical structure through Watson-Crick base pairing. This increases steric repulsive forces between the beam and base nanostructure modules which exceed the tension of the hinge sequences and the surface tension of the adjacent semifluid membrane which bears upon a distal tip of the beam, thereby causing the sensor nanodevice to transition to a more open second configuration.

[0089] In an alternative embodiment, the hinge region may comprise one or more linkers to a binding moiety. The binding moiety may be linked covalently to a single stranded or hairpin loop containing nucleic acid of the hinge region, such as via a nitriloacetate (NTA) conjugation, which has high affinity to a His-tagged binding moiety via complexation of Ni2+or Co3+. Upon binding of the actuation trigger to the binding moiety steric effects in the hinge region can result in initiation of an actuation event causing the sensor nanodevice transition from a first to a second conformation.

[0090] In embodiments of the present invention, the base module comprises a plurality of bundled DNA duplexes. In embodiments of the disclosure, the base module incorporates at least 10 DNA duplexes, with alternative configurations utilizing at least 15, at least 20, at least 25, and optimally around 30 or more DNA duplexes. This multi-duplex architecture enables precise control over the nanostructure's mechanical properties, surface area, and binding capacity while maintaining structural stability through intermolecular interactions between the bundled DNA components. The base module is generally cuboid in shape but may assume any regular polygon or cylindrical shape that facilitates the relative movement of the associated beam module. The dimensional parameters of the base module may be optimized to achieve desired performance characteristics, including stability and adhesion via the membrane anchors, relative to the semifluid membrane surface. The vertical height dimension, which determines the nanostructure's projection outwardly from the membrane interface, is configured to be at least 5 nm in specific embodiments, with alternative embodiments with vertical heights of at least 7 nm, at least 10 nm, at least 12 nm, or at least 15 nm. The horizontal width dimension of the base module, measured along an axis parallel to the plane of the membrane surface, ranges from at least 3 nm in minimal configurations to at least 4 nm, at least 6 nm, at least 8 nm, at least 10 nm, at least 12 nm, at least 14 nm, or at least 16 nm in alternative embodiments. This width parameter influences the base module's contact area with the membrane and may contribute towards its lateral stability. The horizontal length dimension extends from at least 5 nm in compact designs to at least 8 nm, at least 10 nm, at least 15 nm, at least 20 nm, or at least 25 nm in extended configurations. The length parameter, in conjunction with the width, determines the overall footprint of the base module relative to the membrane surface. An exemplary configuration of a base module according to the present disclosure is shown in Figures 2 and 3(c).

[0091] According to embodiments of the present disclosure, the beam module comprises an elongate member that extends as a cantilever outwardly along an axis that is substantially parallel to the plane of the membrane surface when the sensor nanodevice is in the first (closed) configuration. The beam comprises a plurality of bundled DNA duplexes. The beam module may incorporate at least 20 DNA duplexes, with alternative configurations utilizing at least 30, at least 40, at least 50, at least 60, at least 70, at least 80, and optionally around 90 or more DNA duplexes. Selection of the size and configuration of the beam may depend upon the desired length and, thus, the degree of rotation about the fulcrum point represented by leading edge of the base module. It follows that a shorter length of the beam will result in a great range of movement, whereas a longer length of beam will result in a smaller range of movement. Hence, selection of beam module length may be tailored to accommodate the sensor output modality that is selected (e.g. choice of FRET pairing fluorophores, quantum dots, or other detection approaches). The beam module may be L-shaped, including a downwardly extending distal tip portion that may provide a probe functionality required when interfacing with the adjacent semifluid lipid bilayer membrane. The beam module is generally elongated cuboid in shape but may assume any regular polygon or cylindrical shape that facilitates the movement relative to the base portion and the membrane. The vertical height dimension, which determines the nanostructure's projection outwardly from the membrane surface (e.g. the membrane interphase), is configured to be great than that of the base module which sits below the beam module relative to the surface of the membrane. Typically, the vertical height of the beam module is at least 10 nm in specific embodiments, with alternative embodiments with vertical heights of at least 20 nm, at least 25 nm, at least 30 nm, at least 35 nm, at least 40 nm, at least 45 nm, or at least 50 nm. The horizontal width dimension of the beam module, measured along an axis parallel to the plane of the membrane surface, ranges from at least 3 nm in minimal configurations to at least 4 nm, at least 6 nm, at least 8 nm, at least 10 nm, at least 12 nm, at least 14 nm, or at least 16 nm in alternative embodiments. This width parameter influences the beam module's contact area with the underlying base module and also the surface of the membrane at the distal tip portion. The horizontal length dimension should be greater than the underlying base portion so as to create a cantilever. In embodiments the beam portion extends in length from at least 10 nm in compact designs to at least 15 nm, at least 20 nm, at least 25 nm, at least 30 nm, at least 35 nm, at least 40 nm, at least 45 nm, at least 50 nm, at least 55 nm, or at least 60 nm in extended configurations. The length parameter, in conjunction with the width, contributes to the overall footprint of the downwardly extending distal tip portion which in turn will establish a surface area available for contact with the surface of the membrane. In embodiments of the invention the surface area of the distal tip that is available to bear upon and deform the membrane surface may be configured to be at least around 9 nm2, around 12 nm2, around 14 nm2, around 16 nm2, around 18 nm2, around 20 nm2, around 24 nm2, around 28 nm2, around 36 nm2, around 42 nm2, around 56 nm2, around 64 nm2, around 72 nm2, around 84 nm2, around 92 nm2, or around 128 nm2. The selection of probe tip area may determine the mechanics of the sensor nanodevice configuration as well as the type of informational output that may be obtained from the sensor as a whole. An exemplary configuration of a beam module according to the present disclosure is shown in Figures 2 and 3(c).

[0092] As will be apparent, the ability to select different sizes and configuration of the sensor nanodevices of the present disclosure allows for nanostructural optimisation of the sensor to a range of different types of semifluid or lipid bilayer-type membranes. By way of example, highly fluid, labile bilayer membranes having high elasticity and low stiffness may benefit from configurations of the sensor nanodevice that exhibit a lower range of movement, whereas the opposite may be true for stiffer less compliant membranes.

[0093] The specific combination of DNA duplex quantity and dimensional parameters creates a tunable platform wherein the structural and functional properties can be precisely adjusted for particular applications. The bundled DNA duplex architecture provides inherent biocompatibility, programmable sequence-specific interactions, and the ability to incorporate functional modifications through standard nucleic acid chemistry techniques. The dimensional flexibility allows optimization for different membrane types, target molecule sizes, and operational requirements while maintaining the fundamental structural integrity provided by the DNA duplex framework. In specific embodiments of the present disclosure the sensor nanodevice is configured such that the DNA duplexes comprised within the beam and the base portion are substantially aligned such that they project outwardly, in a vertical orientation, from the membrane surface. In alternative embodiments the DNA duplexes of beam and / or base may be aligned so as to be oriented in parallel with the plane of the membrane.

[0094] According to the present disclosure, the sensor nanodevices described in specific embodiments may be used to obtain detailed quantitative data relating to elastic membrane deformation. In accordance with Examples 2 and 3 (described in more detail below) a fluorescence decay value (T value) from FRET following actuation of the sensor nanodevice when anchored to different semifluid lipid bilayer membranes was converted to membrane indentation depths values. These indentation values correspond to inferred bending rigidities for the membranes tested. The bending rigidities are closely related to the Youngs’ moduli which can be calculated. It will be appreciated that, as described previously, alternative signalling modalities may be used instead of FRET such that the invention is not limited.

[0095] The sensor nanodevice of the present disclosure can be used in a diverse range of applications that vary from research tools for assessing membrane dynamics of compositions, through to diagnostic compositions. The following represent non-limiting indications of utility for the sensor nanodevices disclosed herein. It will be apparent to the skilled reader that the presently disclosed devices may have additional very wide-ranging uses that are not disclosed explicitly herein.

[0096] Cardiovascular Diagnostics - early detection of atherosclerosis is made by measuring endothelial membrane stiffness changes that precede visible plaque formation. Sensor nanodevices may be incorporated into point-of-care tests that utilise liquid biopsies that comprise circulating endothelial cells or membrane vesicles as biomarkers, potentially identifying cardiovascular risk in subjects.

[0097] Cancer Detection and Monitoring - malignant transformation fundamentally alters membrane composition, elasticity and mechanics within cancerous or pre-cancerous cells. Compositions comprising the presently disclosed sensor nanodevices allow for detection of circulating tumour cells based on their distinct membrane properties, or by analysis of membrane stiffness patterns in solid or liquid biopsies to identify cancer types, progression and to monitor treatment response (e.g. as a companion diagnostic). This is particularly valuable for many early-stage or hard to treat cancers where current biomarkers may be insufficient. By way of example, it is known that the physical microenvironment contributes to the aggressiveness of recurrent aggressive glioblastoma (GBM). Recurrent GBM cells are reported to express extracellular matrix proteins such as collagen, MMP2 and MMP9 which contribute to increased stiffness in brain tissue and a higher Young's modulus (Acharekar et al. Matrix Biol. (2023) 115:107-127). Neurological Applications - Neurodegeneration involves significant membrane pathology. Hence, sensor nanodevices as described may be incorporated into point-of-care tests for neurodegenerative diseases, such as Alzheimer's, Parkinson's, multiple sclerosis and amyotrophic lateral sclerosis (ALS), that analyse membrane mechanics in cerebrospinal fluid- derived vesicles or even accessible peripheral cells that reflect central nervous system changes.

[0098] Infectious Disease Diagnostics - Pathogens alter host cell membrane properties in characteristic ways. Compositions based around the presently disclosed sensor nanodevices can enable rapid pathogen identification and antimicrobial susceptibility testing by measuring how treatments affect host cell membrane integrity, potentially providing results in hours rather than days. This may be of particular benefit in cases where direct detection of pathogen titre may be close to or below available detection levels. By way of example, envelope viral fusion with host cell plasma membrane can proceed to lipid mixing but not complete fusion, whereas internalized viruses undergo complete fusion with endosomes. This may result in detectable alteration of cell membrane properties during the viral infection process - e.g. for enveloped viruses such as coronaviruses (e.g. SARS-CoV-2), HIV, RSV, influenza etc. It is reported that HIV-1 increases the number of intercellular structures to infect new cells, including tunnelling nanotubes and filopodia, indicating that membrane mechanics can play a role in viral transmission (Bracg et al., Front. Immunol. (2018) vol. 9:260).

[0099] Drug Development and Personalized Medicine - For pharmaceutical applications, compositions comprising the sensor nanodevices may be used in assays to screen drug candidates for membrane-disrupting or -altering effects, to optimize drug delivery systems, or to develop personalized treatment protocols based on individual membrane characteristics. This approach may be of particular value for drugs targeting membrane proteins, cell surface receptors or lipid metabolism. In this embodiment of the present disclosure, uses of a nanostructure, a composition, a sensor nanodevice or a membrane elasticity sensor nanodevice as defined herein are provided in a method for assessing the effect of a xenobiotic compound or substance on a cell, cell culture or membrane containing cellular extract. Xenobiotic compounds may include a pharmaceutical agent, such as a pharmaceutical compound (e.g. a small molecule), a biological or other drug formulation, as well as toxins, toxoids, and / or micronutrients. In specific embodiments, the xenobiotic compound or substance is a therapeutic compound or a candidate therapeutic compound or substance, and the use is in a method for drug discovery or drug development.

[0100] Aging and Metabolic Research The sensor nanodevices disclosed herein can be used to assess and even guantify membrane aging signatures, thereby serving as a biomarker for biological age versus chronological age, or to help diagnose metabolic disorders through characteristic membrane lipid composition or other biomechanical changes. It will be appreciated that the massively parallel screening capability is a significant benefit for the applications described herein because it allows us analysis thousands of cells, vesicles or membrane samples simultaneously. This makes high-throughput and even population-scale studies feasible enabling the statistical power needed for robust diagnostics development. Samples of tissues, cells or lipid containing vesicles may obtained from solid or liquid biopsies obtained from a human or animal subject. Typically, a liquid biopsy comprises a sample of a bodily fluid selected from one of the group consisting of: blood; urine; saliva; semen; tears; sweat; lymphatic fluid; bile; cerebrospinal fluid; ascites; pleural effusion; stool; and a mucus secretion. In particular embodiments of the disclosure a liquid biopsy comprises whole blood, serum and / or plasma.

[0101] In embodiments of the present disclosure the sensor nanodevices may be comprised is a composition that combines the nanostructure in an aqueous buffer solution. Standard base buffer may include any suitable electrophoresis, PCR or nucleic acid carrier buffer, including but not limited to: tris-acetate-EDTA (TAE) buffer; TRIS-HCI buffer; Tris-EDTA (TE) buffer; and / or HEPES-based buffers. Suitably the buffers are modified by addition the cofactor magnesium chloride (MgCh), suitably at a working concentration in 1X buffer of at least 5 nM, 10 nM, 25 nM, 50 nM, 75nM, 100 mM, optionally at least 150 nM. For use with cells or biologically derived tissues the sensor nanodevices may be comprised within a solution of phosphate buffered saline (1X PBS). In a specific embodiment of the invention, the composition comprises DNA sensor nanodevices in solution at a concentration of around at least 5 nM, 10 nM, 20 nM, 30 nM, 40 nM, 50 nM, 75 nM, 100 nM, 150 nM, 200 nM, 300 nM, 500 nM, or 1000 nM,

[0102] The invention is further illustrated by the following non-limiting examples.

[0103] EXAMPLES

[0104] Example 1 - Construction and characterisation of a nucleic acid sensor nanodevice

[0105] A multifunctional molecular machine (MM) that operates as a sensor nanodevice is provided. The sensor nanodevice indents bilayer-type membranes allowing measurement of their local elasticity in a massively parallel fashion and inside cells (Fig. 1). To design the device, a bottom-up DNA nanotechnology approach was selected which yields nanometer dimensions, precise nanomechanical movement, controllable forces34-38, and defined interactions with lipid bilayer membranes3940, as successfully demonstrated with other nanodevices that puncture41-44, shape45“50, or fuse bilayers51- 53, and serve as artificial cytoskeletons454654.

[0106] The presently exemplified DNA nanodevice sensor comprises as a first component a box-like shaped base module (Fig. 1 a) which carries a plurality of lipid anchors for membrane location. The base measures 20 x 12 x 15 nm and is composed of 30 interconnected DNA duplexes. The sensor nanodevice design is based on caDNAno DNA origami software55(Figs. 1 (d) and 2) where a scaffold strand (Table 1) and complementary, shorter staple strands (Table 2) anneal into a DNA nanostructure3435.

[0107] For active membrane indentation, the sensor nanodevice comprises as a second component, an L- shaped cantilever beam module with a vertical distal tip (Fig. 1 a) made of interconnected DNA duplexes (Fig. 2). The cantilever’s proximal end is 15 nm high and its horizontal beam is 40 nm wide (Fig. 1 a, Fig. 2). The beam is connected to the sensor nanodevice base via hinge sequences comprised of single-stranded DNA (Figs. 2 and 3, and Table 3) to allow for a controlled seesawlike rotational cantilever movement (Fig. 1 a). The sensor nanodevice is kept in a closed, nonindenting state (Fig. 1 a) by 12 actuator single-stranded and structurally condensed sequences located between the beam and the base module (Figs. 3 and 4, Table 4).

[0108] To actuate the sensor nanodevice into the open indenting state, the added opening actuator trigger strands hybridise to the condensed actuator hinge strands and form extended DNA duplexes (Fig. 1 a, Fig. 4, Table 5) which displaces the beam upwardly and about a fulcrum point located at one side of the base (referred to as the leading side). This serves to move the distal cantilever tip downward with a designed maximum tilt angle of around 30°. The opening is designed to be reversible by adding closing strands (Table 6) which remove opening strands by toehold-mediated competitive hybridization.

[0109] As sensor nanodevice actuation pushes the cantilever tip downwardly such that it bears on the membrane, it was surmised that the indentation degree depends on the membrane elasticity and associated energetic cost of deformation resulting in a bigger tilt angle for a softer membrane and a smaller angle for stiffer membrane (Fig. 1 b). This allows probing local elasticity in a massively parallel fashion (Fig. 1 b) and inside cells (Fig. 1 c). The sensor nanodevices design provides sufficient energy for membrane indentation due to configuration of actuator strands and tip dimensions, using as rational design basis the Hertz’ biophysical model on indenting elastic membranes (see Example 2). Furthermore, tip compression during indentation should be small as the tip’s duplexes are parallel to the indentation direction56.

[0110] To report on membrane indentation and elasticity, the sensor nanodevice was designed so as to convert the tilt angle into a fluorescence signal (Fig. 1 a). To measure the tilt angle, the distancereporting fluorophores Cy3 and Cy5 are positioned at sensor nanodevice’s base and beam, respectively, so that fluorescence resonance energy transfer (FRET) can track changes in fluorophore distance occurring during sensor nanodevice actuation (Fig. 1 a, Fig. 2a, Figs. 5). In this design, weak membrane indentation of a stiff membrane would keep Cy3 and Cy5 proximate, resulting in a strong FRET response and an enhanced Cy5 signal (Fig. 2a) while a soft membrane would lead to weaker FRET response and a stronger Cy3 signal57 according to a quantitative model (see Example 3). The design was validated by fabricating a sensor nanodevice via annealing a scaffold strand and staple strands (Tables 1 and 2). Self-assembly was successful as indicated by a single band in agarose gel electrophoresis (Fig. 6). The sensor nanodevice was subsequently purified from nonincorporated staple strands via PEG-mediated precipitation and structurally analyzed by transmission electron microscopy (TEM) after negative staining. The TEM-derived dimensions of 33 x 9 x 28 nm (Figs. 7-10, Table 14) compare well to the caDNAno design55(40 x 12 x 30 nm) as well as the dimensions predicted by oxDNA coarse-grain simulations58(40 x 8 x 31 nm) (Fig. 11 a). In Cando simulations59, the sensor nanodevice is stable with few thermal fluctuations (Fig. 12).

[0111] The design was also validated by actuating the sensor nanodevice to reversibly open and close. The experiments were conducted initially without any membranes or anchoring. Successful actuation was confirmed with gel electrophoresis as the gel band of sensor nanodevice shifted after adding opening or closing strands (Fig. 13h). The shifts were maintained at temperatures from 25°C to 45°C (Fig. 13g) and across multiple open-closing cycles (Fig. 13h). Furthermore, the sensor nanodevice ’s tilt angle - as quantified by TEM - shifted for the closed sensor nanodevice from 4.3 ± 3.6 ° (n = 70) to the designed value of 30.0 ± 10.4 ° (n = 50) for the open structure (Fig. 13c, d, Figs. 14-16). The open tilt angle matches well the value of around 29° obtained by oxDNA simulations (Fig. 11 b); the tilt angle of the closed state is in lower agreement as the condensed actuator strands are not well replicated in the simulations (Fig. 11 a). The dynamics of triggered closing and opening was established using sensor nanodevice carrying Cy3 and Cy5 distance markers at base and beam, respectively (Fig. 13a). Mixing a closed-state sensor nanodevice with opening strands led to the transition from high-FRET with strong Cy5 emission to low-FRET with weaker Cy5 and stronger Cy3 signal (Fig. 13a,e,f). The fluorescence changes were reversed after the addition of closing strands (Fig. 13e,f, Fig. 17).

[0112] Sensor nanodevice actuation and indentation was tested to determine if it was possible to sense a difference between bilayer-type membranes of different elasticities.

[0113] Sensor nanodevices were equipped with a plurality of membrane anchors (Fig. 18, Tables 7 and 8). The sensor nanodevices were added to vesicles composed of stiffer POPC and softer DOPC membranes of 200 nm diameter (Fig. 21 a, Fig. 19). The two membranes’ elastic Young’s moduli of ~12.36 MP and 9.72 MPa, respectively, differ as a DOPC’s fatty acid chain includes an extra double bond (Table 9).

[0114] Sensor nanodevices bound equally well to both POPC and DOPC vesicles with Kd values of 24.0 ± 3.0 pM and 22.0 ± 2.2 pM, respectively, as established with gel electrophoresis (Fig. 20). To test whether sensor nanodevice indentation distinguishes membrane elasticity (Fig. 21 a), sensor nanodevices carrying Cy3 / Cy5 distance markers were bound to vesicles and then actuated by adding opening strands. This led to a Cy5 emission decrease (Fig. 21 b) indicating the successful opening of the membrane-bound sensor nanodevice . Strikingly, the fluorescence drop at 20.4 ± 1.0 % for POPC was smaller than for DOPC at 25.8 ± 4.0 % (Fig. 21 b,c) in agreement with a smaller indentation degree for stiffer POPC membranes. Surprisingly, the 24 % difference in the fluorophore signals matches the 24 % difference in Young’s moduli of the two membranes.

[0115] Following successful bilayer indentation, it was assessed whether elasticity differences can also be visually mapped to membranes. Therefore, Cy3 / Cy5-tagged purified sensor nanodevices (Figs. 20 and 21) were bound to micron-sized giant unilamellar vesicles (GUVs) composed of either POPC or DOPC. sensor nanodevice -decorated vesicles were then imaged with fluorescence lifetime imaging microscopy (FLIM) to obtain for each image pixel the average fluorescence decay lifetime T of Cy3 (Fig. 21 a, Fig. 26). This key parameter was expected to reveal the sensor nanodevices Cy3-Cy5 distance, the underpinning sensor nanodevice tilt angle and membrane indentation, and ultimately membrane elasticity (Fig. 21 a, Fig. 26, Example 2). As an additional benefit, FLIM does not suffer from interference by laser power fluctuations or varying localized fluorophore concentrations60.

[0116] To ease visualizing membranes with different elasticities, FLIM images were colour-coded so that each image pixel reflects the key lifetime T for the sensor nanodevice at a given membrane position (Fig. 26). For example, in the FLIM image for membrane- bound and closed sensor nanodevice s T was uniform at 2.3 ± 0.1 ns for POPC and DOPC vesicles as indicated by blue / green hues (Fig. 21 d-f, Figs. 27 and 28). The image in Fig. 21 d shows membrane vesicles with approximately 1 million sensor nanodevices. This highlights the ability of the molecular machine to probe membrane elasticity in a massively parallel fashion.

[0117] Upon sensor nanodevice actuation with opening strands, T strikingly increased to 3.3 ± 0.1 ns for DOPC yet to solely 3.1 ± 0.1 ns for POPC (Fig. 27 and 28), representing significant changes by 50% and 34%, respectively, as well as higher deformability of softer DOPC than stiffer POPC membranes (Fig. 21 e,f, Figs. 27 and 28). The changes to higher lifetimes were reverted upon adding closing strands (Figs. 27 and 28). As other experimental support that lifetime changes represent sensor nanodevice cantilever tilting, control sensor nanodevice -Cy3 lacking Cy5 was used to mimic a maximally tilt-opened sensor nanodevice and found the corresponding T value above all previous values at 3.4 ± 0.1 ns (Fig. 29).

[0118] To obtain further quantitative insight into elastic membrane deformation by sensor nanodevice, the T values from DOPC and POPC vesicles were converted (see Example 3) to membrane indentation depths of -1 .0 ± 0.2 nm and -0.8 ± 0.1 nm, corresponding to inferred bending rigidities K! of 1.1 ± 0.2 x 10-20 J and 1.4 ± 0.2 x 10-20 J for both bilayers, respectively. The bending rigidities are closely related to the Youngs’ moduli (see Example 3) and are slightly lower compared to values obtained with other techniques, possibly due to some flexibility in the anchoring of sensor nanodevice to membranes (Fig. 18). A higher indentation degree for DOPC than POPC was also found in sensor nanodevice’s Cy3 fluorophore decay traces and FLIM phasor plots and indicated less FRET and hence more opening of the sensor nanodevice at the softer membrane (Fig. 30). The differential indentation depths were independently confirmed by determining the indentation’s effect on bilayer organisation using membrane tension dye FliptR (see Example 4).

[0119] Mapping of different membrane elasticities within single vesicles that mimic biological cells was next pursued. As a synthetic model, GUVs were used with softer DOPC-rich liquid-disordered (Ld) domains as well as stiffer DPPC / cholesterol liquid-ordered (Lo) domains (Fig. 31 a). Closed- state sensor nanodevices bound to both membrane regions yielding a uniform T value of 2.3 ns ± 0.1 (Fig. 21 e). A slight preferential sensor nanodevice binding to Ld regions was confirmed by specific staining of Ld (Fig. 31 b,c) and was in line with expectations61. When actuated, sensor nanodevices strikingly differentiated softer Ld and stiffer Lo regions with T increases to 3.2 ± 0.1 and 2.8 ± 0.2 ns, respectively, in single vesicles (Fig. 21 d-f, Fig. 27c). As side note, the relative T changes at 38% and 18% are lower than for the previous monolipid vesicles (50% and 34%) since cholesterol increases membrane stiffness in Lo DPPC / cholesterol regions and has the same effect after partial partitioning into Ld DOPC regions62, as supported by analysing membrane tension with FliptR (see Example 4).

[0120] As final test of the sensor nanodevice’s utility for measuring bilayer nanomechanics, live eucaryotic cells were probed to determine any elasticity differences between the plasma membrane and internal organelle membranes, after sensor nanodevice uptake by cells. When HeLa cells were incubated, DNA-made sensor nanodevices were rapidly internalized into late endosomal and lysosomes (Fig. 32) as also found for other DNA nanostructures63-66. Addition of endocytosis inhibitor Dynasore67-70slowed down sensor nanodevice internalization to allow for a 40 min-long FLIM analysis (Fig. 33). In FLIM images of cells with closed sensor nanodevice , signals were found for plasma membrane (PM) as well as membranes of internal vesicles (INT) (Fig. 34a, b) with a uniform lifetime of 0.8 ± 0.1 ns across the membrane regions; the T value is lower than for synthetic vesicles likely due to the different physicochemical membrane environments71 72. Actuation of cell-bound sensor nanodevice with opening strands uncovered lifetime increases of 1.0 ± 0.1 ns for PM and 1.3 ± 0.1 ns for INT regions (Fig. 34a-c) which is below 1 .5 ± 0.1 ns from control sensor nanodevice -Cy3 (Fig. 34). The lower lifetime of 1 .0 ± 0.1 ns at PM reveals that the plasma membrane is less elastic than the softer internal membranes at 1 .3 ± 0.1 ns73. Different elasticities at cellular membrane regions were also apparent in the FLIM decay curves and FLIM phasor plots showing more FRET efficiency and lower sensor nanodevice tilt angles at PM than INT membrane regions (Figs. 35-37). The phasor changes were reversible upon the addition of closing strands (Figs. 36-37) providing proof that internalised sensor nanodevices are not digested by cellular nucleases. In the present example bottom-up DNA nanotechnology has been used investigate the biologically and disease-relevant nanomechanics of lipid bilayer membranes. To be functional, the new 40 nm- sized cantilevers have been integrated into a molecular sensor nanodevice which binds to, actuates, senses, and reports on associated membrane elasticity. Compared to serial top-down analysis, the molecular devices’ small size and self-contained multifunctional operation allow deploying millions of sensor nanodevice copies for massively parallel analysis as well as for examining internal cellular membranes; simultaneous analysis of external and internal cell membranes is novel. The observed higher stiffness of the plasma membrane is likely related to its high content of cholesterol and saturated glycerophospholipids, while the more elastic internal membrane may reflect lower cholesterol levels and more unsaturated lipids74-76which alter membrane lipid order77 78. It is also possible that the stiffer PM membrane is interlinked to the underlying cytoskeleton which responds to external deforming force7980.

[0121] The sensor nanodevices described in the present disclosure indent membranes solely by around 1 nm which yields a more fine-grained picture on bilayer elasticity than othertop-down techniques whose greater indentation depth affects both membranes and the underlying protein cell cortex layer. The current sensor nanodevices report exceptionally well on relative elastic Young’s moduli; determining absolute membrane elasticities requires calibration with synthetic membranes as typical for other techniques. The present sensor nanodevices complement existing DNA nanodevices for measuring biophysical cell parameters including DNA duplexbased pulling force sensors82-86and are expected to allow new insight into biologically and biomedically relevant topics including blebbing cells, cell division, stem cell differentiation, and embryogenesis, and with adaption also to probe polymeric soft interphases.

[0122] METHODS

[0123] Materials. DNA strands, 3' labelled DNA-TEG-Chol and 5' labelled DNA-Cy3 oligonucleotides as well as Cy3 and Cy5 labelled DNA oligonucleotides were purchased from Integrated DNA Technologies (UK) or Eurogentec (BE) with HPLC or PAGE purification. Phix174 scaffold was purchased from New England Biolabs (UK). Lipids 1-palmitoyl-2-oleoyl-glycero-3-phosphocholine (POPC), 1 ,2-dioleoyl-sn- glycero-3-phosphocholine (DOPC), 1 ,2-dipalmitoyl-sn-glycero-3- phosphocholine (DPPC) and cholesterol (plant-derived) were purchased dissolved in chloroform from Avanti Polar Lipids (UK). All other reagents were purchased from Sigma-Merck (UK) unless stated otherwise.

[0124] Sensor nanodevice fabrication. The DNA sequences and 2D connectivity map of the DNA sensor nanodevice are provided in Table 2- 6 and Figure 1 (d). To self-assemble sensor nanodevice at a concentration of 50 nM, Phix174 virion DNA scaffold (10 pL, 500 nM, Table 1 [SEQ ID NO:1]) was mixed with 6 x excess of 112 staple strands (32.5 pL, 100 pM; Table 2; [SEQ ID NOs: 2-113]), 5 x excess of cholesterol anchoring strands (6 pL, 100 pM; Table 7; [SEQ ID NOs: 150-169]), 1 x TAE 160 mM MgCh (10 pL) and topped up with deionised H2O to a total volume of 100 pL. For fluorescence experiments, staple strands main025 [SEQ ID NO: 26] and main078 [SEQ ID NO: 79] were replaced with Cy3 and Cy5-labelled versions at 3 x fold excess (15 pL, 10 pM). 12 opening strands (Table 5; [SEQ ID NOs: 126-137]) and 12 closing strands (Table 6; [SEQ ID NOs: 138-149]) were added in 5x excess (3.6 pL, 100 pM). The DNA origami was folded in a Thermocycler (Biorad, US) by heating up to 80° C for 10 min following 10 min at 65°C. Next, the mixture was cooled down for one degree every hour to 25°C, and then cooled one degree every 10 min to 15°C. Volumes were adopted accordingly for assembly at the 20 nM scale.

[0125] Sensor nanodevice purification. For purification by PEG precipitation, folded sensor nanodevice (100 pL, 20-50 nM) was mixed with 200 pL of buffer 1 (15% PEG 800, 1x TAE, 0.5 mM NaCI), vortexed for 3 s and centrifuged for 30 min at 10,000 r.c.f. at 25 °C. The orientation of the tube placed in the centrifuge was labelled to indicate the position of the forming pellet. The supernatant was removed after centrifugation and 100 pL of buffer 2 (1x TAE, 16 mM MgCh) was added, vortexed for one min and incubated at 37°C and 1000 rpm in a heating block (Biorad, US) for 20 h.

[0126] Purification by size exclusion chromatography (SEC) was conducted for microscopy experiments. Assembled sensor nanodevice was purified from excess staple strands using SEC via an KTA purifier (Cytiva, US) with a Superdex 75 increase packed 10 / 300 GL column (GE Healthcare, US). The sample was eluted with degassed 16 mM MgCI2 buffer at a flow rate of 0.5 mL I min and by monitoring the absorbance at 260 and 280 nm. The eluent was collected on a 96-well plate and fractions containing sensor nanodevice were analysed via agarose gel electrophoresis.

[0127] Membrane vesicle formation. To form large unilamellar vesicles (LUV), a solution of POPC or DOPC lipids in chloroform (10 mM, 100 pL) was added to a 2 mL screw-top glass vial. The solvent was removed using argon gas, while rotating to yield a thin lipid film. The lipid was resuspended in LUV buffer (50 mM HEPES 500 mM NaCI, or 1 x TAE 16mM MgCh; 1 mL), the solution was vortexed for 30 s and then sonicated for 1 min at RT. The LUV vesicle solution was extruded 25 times through an Avanti Mini- Extruder (Avanti Polar Lipids, US) with 200 nm filter paper (Avanti Polar Lipids, US). Dynamic light scattering (DLS) was used to confirm the vesicle diameter via a Zetasizer Nano S (Malvern Panalytical, UK).

[0128] To form giant unilamellar vesicles (GUV), a solution of POPC or DOPC lipids (36 pL, 10 mg / mL) in chloroform were placed within a rubber O-ring on an indium-tin-oxide (ITO) (Nanion Technologies, DE) coated glass slide and allowed to dry to form a lipid film. The ITO slide was inserted in a vesicle prep device (Nanion Technologies, DE). Sucrose (250 pL, 400 mM) was added to the dried lipid and another ITO glass slide was placed on top to close the chamber. The GUVs were formed via electroformation by using the voltage program with 10 Hz, 4V amplitude, 1 min rise, 100 min main time and 10 min fall time. The temperature was set at 70°C forthe mixed- phased GUVs and 27 °C for DOPC and POPC GUVs.

[0129] Characterization of sensor nanodevices with agarose gel electrophoresis. A 1 % agarose gel in 0.5 x TBE 10 mM MgCh with ethidium bromide (1 :30000, 5 pL in 150 mL) was used to determine the assembly of sensor nanodevice. 5-10 pL of DNA (20 nM) were mixed with gel loading dye (purple 6x, no SDS; New England Biolabs, UK) and loaded onto the gel. The gel was run for 1-2 hours (depending on the degree of the desired separation) at 65 V and 4°C. To distinguish between the opened and closed conformation of the DNA sensor nanodevice, a 2.5% agarose gel was run for 2.5 hours at 65 V and 4 °C. The bands were analysed using an Azure Biosystems C Series Imaging Systems (Azure Biosystems, US).

[0130] Agarose gel electrophoresis was also used to confirm the binding of sensor nanodevice to LUV membrane vesicles. Purified sensor nanodevice (7 pL, 20 nM) were incubated with cholesterol strands in 5x access (4.2 pL, 10 pM) for 15 min and mixed with LUVs (POPC or DOPC) (7 pL, 20-200 pM). The DNA-sensor nanodevice LUV mixture was incubated for 1 hr at RT. The mixture was analysed using 1% agarose gel with Ethidium bromide (1 :30000, 5 pL in 150 mL) in TBE (0.5 x TBE 10 mM MgCh). The gel was run at 65 V for 90 min at 4°C. The gel bands were visualised using an Azure Biosystems C Series Imaging Systems (Azure Biosystems, US).

[0131] Transmission electron microscopy. Purified sensor nanodevice (6 pL, 3-5 nM) were placed on a glow-discharge treated carbon-coated Cu300 mesh grid and negatively stained with 2 % uranyl formate. The images were obtained with a JEM 2100 electron microscope and Orius SC200 CCD camera at 200Kv (JEOL, JP). TEM images were analyzed using a custom MATLAB code. Briefly, fiducial markers (Fig. 10a) were manually selected by the user, and a rigid-body transformation was then applied to align the images corresponding to the sensor nanodevice. Opening angle was estimated from the dot-product between vectors

[0132] Spectrometric FRET analysis of sensor nanodevice. The fluorescence emission was measured with a Varian Eclipse fluorescence spectrophotometer (Agilent, UK). The excitation slit was set at 5 nm, the emission slit at 10 nm, the scan rate at 600 nm I min and excitation at 540 nm and 600 PMT voltage. 100 pL of the DNA sensor nanodevice was added to a quartz cuvette with a 10 mm path length (Hellma, DE). For kinetics, the excitation wavelength was set at 550 nm, the emissions were scanned between 570 and 670 nm. Opening or closing strands in 5x excess were added after 5 min. The data was normalised onto the average intensity before the addition of the strands and percentual change to the initial intensities recorded.

[0133] GUV experiments with sensor nanodevice, sensor nanodevices (15 pL, 50 nM) were added to a solution of GUVs (15 pL) in 1x PBS buffer (150 pL) in a p-Dish 35 mm, high (Ibidi, US) fitted with a 6.96 mm diameter insert. For FliptR FLIM experiments, the sensor nanodevice (15 pL, 50 nM) was added to a solution of GUVs in 1x PBS supplemented with FliptR (2 mM) underthe same conditions, sensor nanodevice and GUVs were left for 15 min at RT and subjected to microscopy and lifetime imaging. For the sensor nanodevice actuating experiments, Fluorescent lifetime imaging microscopy (FLIM) measurements were carried out only after 10 min incubation with opening strands (OS) (0.9 pL, 8.3 pM) or closing strands (CS) (0.9 pL, 8.3 pM).

[0134] Cell tissue experiments with sensor nanodevice. Wild-type HeLa cells were cultured in Dulbecco's Modified Eagle Medium (DMEM) (Gibco, Life Technologies, US), supplemented with 10 % fetal bovine serum (FBS) and 1 % Penicillin-Streptomycin in T-75 flasks (Sigma-Merck (UK)). The cultures were maintained in a humidified atmosphere containing 5% CO2 at 37°C and passaged every ~3 days. Cells were plated at a density of 5,000 cells per well in a p-Dish 35 mm, high (Ibidi, US) fitted with a 6.96 mm diameter insert. The following day, the cells were washed with 1x PBS and treated with Dynasore (80 mM) for 30 min to inhibit clathrin- coated endocytosis. The cells were washed three times with 1x PBS and then incubated with sensor nanodevices in 1x PBS (5 nM) for 5 min at 37 °C. The cells were washed three times again with PBS to remove any non-bound DNA and subjected to microscopy and lifetime imaging. For the sensor nanodevice actuating experiments, FLIM measurements were carried out after 10 min incubation with opening strands (OS) (8.3 pM, 0.9 pL). FLIM imaging with sensor nanodevice opening and closing was carried out within a 45 min timeframe after adding sensor nanodevice to avoid readings of low DNA signal and photon counts. Statistical analysis was conducted using Prism GraphPad (https: / / www.graphpad.com / features).

[0135] Confocal laser scanning microscopy. Fluorescence microscopy images were taken using a 60x oil objective on the Zeiss Airyscan LSM 880 or Nikon A1 . Both imaging systems are equipped with an environmental chamber to maintain cells at 37 °C and 5 % CO2.

[0136] Images were taken using laser excitation of 532 nm (Cy3) to determine any DNA- membrane interaction. All images were taken using the same exposure and gain settings. Image analysis was carried out using Imaged ((https: / / imagej.net)) and statistical analysis using Prism GraphPad.

[0137] Fluorescence lifetime imaging microscopy. Multiphoton fluorescence lifetime imaging microscopy (FLIM) was conducted on the Leica TCS SP8-FALCON (FAst Lifetime CONtrast) system (Leica Microsystems GmbH, Wetzlar, DE) using a 40x air objective. Sensitive hybrid detectors (SMD, Leica Microsystems CMS GmbH, Mannheim, DE) allowed for photon detection, while a 910 nm pulsed laser (80 MHz) was used to excite the FliptR and 760 nm to excite Cy3. FLIM images were acquired at 512 x 512 pixels. At least 10,000 photon events were recorded in each measurement. The photon events were time-binned into a histogram and a fluorescence decay curve obtained. FastFLIM images (average photon arrival times per pixel) are shown. Regions of interest (ROI) were drawn to determine the lifetime of different vesicle phases or cell compartments. For FliptR FLIM fit analysis, the SP8 Falcon fitting model ‘n-exponential reconvolution’ was used for the biexponential fitting of the fluorescence decay curve, to extract the FliptR lifetimes T1 and T2. The parameter chi-squared (x2) was used as an indicator for the quality of the curve fitting. The longest lifetime (T2) was used to report on membrane tension, while T1 only accountant for the minority of the signal due to a low photon number. Prism GraphPad was used to plot the average lifetime and conduct statistical analysis. For FRET-FLIM, phasor analysis, the data was analysed using an integrated FLIM-Phasors-analysis, enabling a global view of the fluorescence decay in each pixel of the measured image. FRET efficiency was determined using a FRET trajectory, which combines the phasor of the unquenched donor, background signal, and the experimental points in the phasor plot, to determine the number of donors undergoing FRET and the corresponding FRET efficiency. This fit-free approach does not resolve the decay in exponential components.

[0138] Example 2 - Rational design of sensor nanodevice informed by free energy calculations

[0139] The design of the DNA-based sensor nanodevice is defined by the relationship between the free energy gained by DNA hybridization E.Gopening, driving the indentation process, and the energy penalty of deforming a lipid membrane of given thickness H and bending rigidity KC. The maximum deformation (indentation) that could be imposed on the membrane before it would break and form a toroidal pore around the DNA structure was estimated. Based on Hertz’s theory describing the elastic indentation of a 2D half-space by an axisymmetric flat indenter, it was assumed that the sensor’s tip would be cylindrical, and thus described the indentation force as in Acerbi et al.96: in terms of Young’s elastic modulus (E), Poisson ratio (v), indenter radius (R), indentation depth (5) and applied force (F).

[0140] Because the stiffness of a closely packed DNA nanostructure is much stiffer than lipid bilayers (i.e. EDNA» Emembrane), the indentation force can be expressed as:

[0141] Equation (1.2)

[0142] The indentation force will be associated to a mechanical energy of the form: Equation (1.3)

[0143] As one can consider the membrane to be incompressible, it follows that vmembrane = 0.5. It is also assumed no indentation at time zero, so that 60= 0. Then, this results with the expression:

[0144] Equation (1.4)

[0145] According to thin-plate theory, the membrane’s elastic modulus can be related to its bending rigidity (xc) by means of the following identity96:

[0146] Equation (1.5)

[0147] Therefore, the indentation energy can be expressed as a function of the membrane’s bending rigidity as:

[0148] Equation (1.6)

[0149] On the other hand, for a sufficient deformation the membrane would eventually be punctured, and a hydrophilic toroidal pore will be formed, in which lipid headgroups will rearrange to face the lumen. The energy cost for such a process is:

[0150] UPR) = 2nyLR - nysR2

[0151] Equation (1.7)

[0152] Where R is the pore radius, YL is the membrane’s pore line tension and ys is the membrane’s surface tension. For lipid membranes where no external tension is applied, the squared term can be disregarded; so the energy cost for opening a pore grows linearly with R."

[0153] According to Wohlert et al." line tension can be compared to the bending rigidity KCSO that the energy for pore formation is simplified as:

[0154] Kcn2R

[0155] UP(R)

[0156] H

[0157] Equation (1.8)

[0158] Therefore, we can calculate the ratio (p between the energy cost for both processes as:

[0159] Equation (1.9)

[0160] From this relation, it then follows that indentation depth and membrane thickness would be the only parameters determining whether the bilayer will be punctured. Indeed, penetration would occur if Ui > Up (i.e. (p > 1). Hence, a critical indentation value 6Ccan be calculated as:

[0161] Equation (1.10)

[0162] Assuming a characteristic membrane thickness H~ 4 nm, the maximum allowed indentation is calculated to be 8C~ 2.6 nm. Considering a typical bending rigidity KC~ 10-19J and H ~ 4 nm, Eq. 10 can be used to relate the indentation energy with the tip radius, which for the above values is:

[0163] U,(R)~ 2.5 * IO10 / ?

[0164] Equation (1.11)

[0165] On the other hand, the hybridization free energy for a DNA duplex of length 15-30 bp is about 1 O’19J / duplex, hence the number of duplexes required to actuate the molecular machine and create an indentation 8Con lipid membranes can be expressed as: opdupiex~ 2’5 nm R

[0166] Equation (1.12)

[0167] To lower the device’s complexity, N0Pduplexshould be kept to a minimum. The lower bound for R is limited by the honeycomb lattice used in the DNA origami design, i.e. a single hexagonal arrangement of the DNA duplexes with an equivalent radius of R ~ 3 nm. Based on design considerations, however, it was decided to extend the tip’s cross-section (Fig. 2c) to increase the rigidity of the DNA nanostructure, to an equivalent radius Req.

[0168] Req= ^ab / n

[0169] Equation (1.13) where a, b correspond to the 2 dimensions (i.e. “width” and “length”, 7.2 nm and 9 nm, respectively) of the sensor’s tip; resulting in Req~ 4.54 nm, thus requiring ~ 12 opening strands. Example 3 - Converting FLIM-FRET lifetimes into membrane indentation depths

[0170] Fluorescence lifetime imaging microscopy (FLIM) does not depend on the local fluorophore concentration, and is robust against intensity-based FRET readouts, such as channel bleed through; hence FLIM-FRET is a suitable technique to quantify the degree of indentation by sensor nanodevices.

[0171] Lifetime-based FRET efficiency can be calculated as:

[0172] Equation (2. 1) where TDArepresents the donor’s (Cy3) lifetime in the presence of the acceptor (Cy5), and TDrepresents the fluorescence lifetime of the donor on its own. Importantly, one notes that Cy3 is a well-known environmentally sensitive probe100 101; its photophysical behaviour will depend on the local DNA geometry (Fig. 5). Therefore, specific Cy3 lifetime calibration can be performed when attached to different nucleic acid nanostructures.

[0173] On the other hand, the FRET efficiency between a dye with Forster radius Docan be related to the donor-acceptor separation d as:

[0174] Equation (2.2)

[0175] Therefore: d — Do

[0176] Equation (2.3)

[0177] Based on sensor’s design, the distance between the FRET pair can be used to calculate the opening angle 9 as:

[0178] Equation (2.4) where LDand LArepresent the distance from the dyes to the pivot’s point (Fig. 13g-h). Once the opening angle is known, it is possible to calculate the indentation depth 8 at the point Ptipas:

[0179] 8 = Pysin(0) + Pxcos(0) — Py— p

[0180] Equation (2.5) where Px= - 20 nm and Py= - 15 nm represents the tip position relative to the pivot point (Fig. 5, Fig. 13d) and p = - 5 nm represents any offset (e.g. due to the cholesterol-TEG linker).

[0181] Taking the total indentation as the difference between the estimated indentation depths before and after the addition of the OS:

[0182] A<5 = 60S— 60

[0183] Equation (2.6)

[0184] Then Eq. 1 .2, Eq. 1 .5 and Eq. 2.61 .5 can be combined to estimate the bending rigidity, where Fosis derived from the Worm-Like Chain model (Eq. 3.1):

[0185] =F0SH3

[0186] ^c, estimate 247? A A

[0187] Equation (2. 7)

[0188] Combining the estimated force and the inferred A<5 from FRET measurements, recalling the tip radius Req~ 4.54 nm and setting the membrane thickness H = 4.52 nm we estimated the bending rigidity Kc estimatefor the different lipid membranes (Table 10). The bending rigidities are slightly lower compared to values obtained with other techniques (Table 11), possibly due to some flexibility in the anchoring of the sensor to membranes.

[0189] Example 4 - Mechanical characterization of the sensor nanodevice

[0190] To assess the capability of the sensor nanodevice to indent lipid membranes, the force exerted by the addition of actuation triggering opening strands was quantified.

[0191] The force caused by opening of the sensor nanodevice was modelled as that of an entropic spring. According to the Worm-Like Chain model, the addition of each opening strand will contribute:

[0192] (Equation 3. 1) where Lpis the persistence length for ssDNA (1 .5 nm), R = 0.347V - 2 nm / bp is difference in the end-to-end extension upon hybridisation of the opening strands, and Lc= 0.65 N nm / bp is the contour length for the strands connecting the tip and base of the MM where N is the number of base pairs. Using these values, we estimated that the total force to be 31 ,97pN upon full actuation of the sensor nanodevice. Similar forces have been reported by other DNA origami-based nanoactuators.

[0193] Using this insight, it was explored whether the readout from the present nanodevice followed Hertz’s indentation model by recording the fluorescence readout from nanodevice bound to DOPC and POPC membranes upon the addition of six short, six long, or all opening strands. Using Eq. 2.1 , the force was estimated for a given set of opening strands and compared that to the change in Cy5 signal (Fig. 21d, Table 12).

[0194] If comparing the change in relative indentation depth AAA = 8p0PC- 8p0PCthen, for a given force:

[0195] (Equation 3.2)

[0196] On the other hand, the FRET efficiency from intensity-based measurements may be approximated as:

[0197] (Equation 3.3)

[0198] Therefore:

[0199] (Equation 3.4)

[0200] Now, considering the relative change in Cy5 acceptor intensity, then:

[0201] (Equation 3.5)

[0202] Hence:

[0203] (Equation 3.6) where d0is the FRET distance in the absence of opening strands, and that is taken as 6.5 nm from the FRET-FLIM measurements. If it is assumed that the indentation depth and FRET distance are proportional, e.g. dos~ 80Sand d0~ 60, then one can express:

[0204] (Equation 3. 7)

[0205] Finally, combining Eq. 3.7 and Eq. 3.2 yields Eq. 3.8, which is plotted in Fig. 41 b:

[0206] (Equation 3.8)

[0207] Comparison of experimental Cy5 intensity changes in Fig. 21 a-f yields a good match in terms of two observables. Increasing the number of OS increased the change of Cy5 intensity, and the change is less prominent for stiffer vesicles (POPC). Altogether, this indicates that the sensor nanodevice has the characteristics required to report on the bending rigidity of lipid membranes.

[0208] Example 5 - Analysis of FlipTR® probe response to membrane deformation by the sensor nanodevice

[0209] Actuation of sensor nanodevices is expected to deform the lipid bilayer. Deformation was measured by membrane-tension probe FliptR to explore whether the degree of deformation correlated with the opening angle and membrane indentation as quantified by FLIM analysis of sensor signalling. FliptR reports on changes in membrane tension by its fluorescence lifetime (Fig. 38).102 103The twist angle between the two FliptR dithienothiophene (DTT) groups is affected by the membrane’s deformation, so that a higher degree of lipid packing within a deformed membrane should lead to an increased lifetime of the dye. Fluorescence emission decay curves of FliptR were recorded under four conditions: (1) incubation with GUVs (Ctrl), followed by sequential addition (2) of cholesterol-tagged sensor, (3) opening strands (+ OS), and (4) closing strands (+ CS). The decay curves were fitted with double exponentials to obtain two lifetimes (n and T2) (Table 13). The FLIM-fit images and bar charts (Fig. 38) correspond to the longer lifetime (T2) with greater number of emission events and directly reports on membrane tension.

[0210] Lipid composition profoundly affected FliptR lifetime (Figs. 38, 39, Table 13) and inferred membrane tension. In the absence of DNA (Ctrl), DOPC membranes had the lowest membrane tension (2.83 ± 0.06 ns) while POPC bilayers featured higher tension (3.23 ± 0.04 ns). In the case of phase-separated GUVs, FliptR lifetimes in the DOPC-rich Ld domains and DPPC-CL rich Lodomains were 3.4 ± 0.14 ns and 4.32 ± 0.13 ns, respectively, in agreement with existing reports104with slight deviations attributed to buffer conditions. The increased lifetime in the Ld region compared to pure DOPC GUVs could be due to a small proportion of cholesterol residing in the DOPC domain, as also suggested in the FRET-FLIM dataset (Figure 38).

[0211] After experimentally proving by the FlipTR lifetime changes the different bilayer deformations caused by the sensor (Fig. 40), it was explored whether the measured changes could be, indeed, explained by the nanoscale membrane deformations in the vicinity of the DNA nanostructure’s tip. If this were to be the case, then the calibration factor describing FlipTR lifetime change response to membrane deformation would for the sensor nanodevice-mediated indentation be within the range found in literature. To this end, the the relative FlipTR lifetime x between DOPC and POPC GUVs was first defined after sensor addition as:

[0212] TDOPC „ z:= - - 1

[0213] LPOPC

[0214] Equation (4. 1) where the lifetime after sensor actuation can be described as:

[0215] Tx= TXQ + AX

[0216] Equation (4.2) with TX,o being the lifetime afterthe sensor is attached to the bilayer, but before the opening strands are added, and ATXbeing the change in lifetime coming from sensor action and membrane indentation and resulting membrane tension.

[0217] ATXmay then be related to the membrane area strain uxas:

[0218] Equation (4.3) where KA xrepresents the area stretching modulus, uxrepresents the area strain and (xrepresents a calibration constant accounting for the tension / lifetime dependence of the FliptR probe. Therefore, the measured lifetime ratio after sensor opening can be expressed as:

[0219] Equation (4.4)

[0220] This can be expanded as: Equation (4.5)

[0221] TakingDOPC O= 2.26 ns andDOPC O= 2.29 ns, x can be rewritten as:

[0222] Equation (4.6)

[0223] Given that both membranes are in the same phase and do not exhibit phase separation, it is reasonable to assume the calibration factor will be similar for both DOPC and POPC bilayers, and therefore:

[0224] Equation (4. 7)

[0225] Assuming membrane coverage and area extension moduli will be the similar in both cases (i.e. CIDOPC = apopc and KDOPC = KPOPC) allows to express the calibration factor as: _ XTPOPC,O _

[0226] A(.TPOPC,OUDOPC +TDOPC,OUPOPC)

[0227] Equation (4.8)

[0228] On the other hand, one also needs to consider which percentage of GUV membrane surface is experienced sensor nanodevice-mediated deformation. Towards this end, membrane area strain u can be estimated as the transition from a planar membrane patch of radius a to that of a spherical cap whose height is determined by the indentation depth 8 (Fig. 41). By taking the values of 8 derived previously for DOPC and POPC membranes and using a = 9.8 nm (based on the sensor tip geometry, Fig. 41) it is possible to obtain the area of the spherical cap after deformation (As) which yields AS.DOPC = 328 nm2and AS.POPC = 320 nm2respectively, which would correspond to an area strain UDOPC = 0.085 and UPOPC = 0.062, respectively. We note that these values are around the limit for bilayer breaking (~8%), although they are a limiting case, as we reckon the height of the cap base will be larger, and consequently the area strain will be reduced.

[0229] Taking K ~ 250 mN / m, it is possible to calculate the FliptR calibration factor as a function of membrane coverage, as shown in Fig. 41. One can then define GUV coverage as the ratio between the total “indentation area” and the accessible bilayer surface so that:

[0230] Equation (4.9) Given VMM = 15 pL, CMM = 50 nM, VGUV = 15 pL, CGUV ~ 2 mM (see Methods section) and considering the area per lipid to be 72.4 A2and 69.3 A2for DOPC and POPC, respectively, with an indentation area of 361 nm2per MM, it is possible to estimate under the assumption of complete binding of sensor nanodevice to GUVs a vesicle coverage of ODOPC ~ 2.5 % and apopc ~ 2.6 % respectively. These coverage values correspond to a calibration value of ~-1 .36 ns m / mN.

[0231] Therefore, it can be concluded that the observed changes in FliptR lifetime are indeed compatible with a sensor nanodevice-induced membrane deformation.

[0232] Tables

[0233] Table 1 - Sequence of phiX174 scaffold strand [SEQ ID NO:1]

[0234] Table 2 - Names and sequence of the DNA oligonucleotides used as staple strands to form sensor nanodevice. Oligonucleotide main025 [SEQ ID NO:26] carries a 3' Cy3 modification and main078 [SEQ ID NO: 79] carries an internal Cy5 modification.

[0235] Table 3 - Sequence of hinge sequences within phiX174 sequence Table 4 - Sequences of actuator sections within phiX174 sequence and their position within the 2D duplex map in Figure 5. Actuator sequences form either a hairpin or a simple singlestranded loop, as indicated. The bold letters represent the bases which are hybridising to form the hairpin’s stem.

[0236] Table 5 - Names and sequences of the opening strand oligonucleotides. Strands 01-06 are long OS and strands 07-12 are short opening strands.

[0237] Table 6. Names and sequences of the closing strand oligonucleotides

[0238] Table 7 - Names and sequences of anchoring strand oligonucleotides which bind to cholesterol-TEG-modified DNA oligonucleotides. The binding position to duplexes within the 2D map of Figure 5 is also shown. The lowercase letters in the sequence indicate the sections to which cholesterol-TEG-modified oligonucleotides bind.

[0239] Table 8 - Names and sequences of the cholesterol-TEG-modified DNA oligonucleotides which bind to the anchoring strands listed in Table 7.

[0240] Table 9. Names, structure, and phase transition temperatures of lipids used in the example.

[0241] *Phase transition temperatures values obtained from Avanti polar lipids (https: / / avantilipids.com)

[0242] Table 10. Parameters used to calculate bending rigidity kc. The indentation depths were derived from FLIM analysis. The same membrane thickness was used throughout the calculations for kcfor reasons of consistency. A total force of 32 pN was assumed, based on Eq. 3.1 . Error represents ±SD except for KC, representing SEM.

[0243] Table 11. Known bending rigidities for DOPC and POPC Table 12 Estimated indentation force and change in Cy5 intensity upon OS addition for different OS sets. Contrast refers to Cy5_POPC-Cy5_DOPC and is related to the sensitivity of the sensor nanodevice to the membrane’s elasticity.

[0244] Table 13. Lifetime components (7) extracted from FliptR analysis of the sensor nanodevice molecular machine (MM) binding to membrane vesicles. The components are extracted from double-fitted photon lifetime histograms. Table 14. Summarising the nominal dimensions from the sensor nanodevice MM design and dimensions obtained from TEM analysis (n = 128 for side view, n = 20 for top view; ± SD) as shown in Figure 10a and b.

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[0344] Although particular embodiments of the invention have been disclosed herein in detail, this has been done by way of example and for the purposes of illustration only. The aforementioned embodiments are not intended to be limiting with respect to the scope of the appended claims, which follow. The choice of nucleic acid starting material, the scaffold sequence(s), or type of fluorescent readout used is believed to be a routine matter for the person of skill in the art with knowledge of the presently described embodiments. It is contemplated by the inventors that various substitutions, alterations, and modifications may be made to the invention without departing from the spirit and scope of the invention as defined by the claims.

Claims

WHAT IS CLAIMED IS:1 . A nucleic acid nanostructure comprising: a base module, wherein the base module is configured to adjoin a semifluid membrane that defines a surface; and a beam module, wherein the beam module is connected to the base module via one or more flexible hinge sequences such that the beam module is able to translate through a range of motion between a first and a second conformation relative to the base module; wherein translation of the beam module from the first to the second conformation causes the nucleic acid nanostructure to apply a force to the surface of the semifluid membrane.

2. The nanostructure of claim 1 , wherein the one or more flexible hinge sequences are comprised of single stranded nucleic acid sequences.

3. The nanostructure of claims 1 or 2, wherein the one or more flexible hinge sequences comprise at least one hairpin loop containing sequence.

4. The nanostructure of any one of claims 1 to 3, wherein translation of the beam module from the first to the second conformation occurs in response to an actuation trigger.

5. The nanostructure of any one of claims 1 to 4, wherein the range of motion of the beam module from the first to the second conformation relative to the base module is measurable.

6. The nanostructure of any one of claims 1 to 5, wherein the base module comprises a fulcrum point, and the translation of the beam module comprises an angular rotation about the fulcrum point in response to an actuation trigger.

7. The nanostructure of claim 6, wherein the angular rotation comprising a tilting motion whereby the beam module comprises a distal tip, and the distal tip moves through an arc relative to the base module.

8. The nanostructure of any one of claims 6 or 7, wherein the range of angular rotation is measurable.

9. The nanostructure of any one of claims 1 to 8, wherein the actuation trigger is comprised of a nucleic acid sequence, optionally a single stranded nucleic acid sequence.

10. The nanostructure of any one of claims 1 to 9, wherein the actuation trigger is comprised of a single stranded nucleic acid sequence that completely or partially hybridizes with one or more of the flexible hinge sequences.11 . The nanostructure of any one of claims 1 to 10, wherein the beam module comprises a distal tip portion.

12. The nanostructure of claim 11 , wherein the distal tip portion functions as a probe.

13. The nanostructure of any one of claims 1 to 12, wherein the applied force is measurable.

14. The nanostructure of any one of claims 1 to 13, wherein the applied force is perpendicular to a plane defined by the surface of the semifluid membrane.

15. The nanostructure of any one of claims 1 to 14, wherein the applied force is sufficient to cause a deformation of the semifluid membrane.

16. The nanostructure of claim 15, wherein the deformation of the semifluid membrane is measurable.

17. The nanostructure of any one of claims 6 to 16, wherein the range of angular rotation of the beam module relative to the base module is correlated to an amount of deformation of the semifluid membrane.

18. The nanostructure of any one of claims 15 to 17, wherein the amount of deformation of the semifluid membrane is correlated to an amount of an applied force.

19. The nanostructure of any one of claims 1 to 18, wherein the amount of applied force corresponds to a mechanical property of the semifluid membrane.

20. The nanostructure of any one of claims 9 to 19, where the length of the single stranded nucleic acid sequence actuation trigger correlates to degree of deformation of semifluid membrane.21 . The nanostructure of any one of claims 19 or 20, wherein the mechanical property is selected from: elastic stiffness, including Young's modulus; membrane tension; and / or a viscoelastic property.

22. The nanostructure of claim 21 , wherein the mechanical property is a Young’s modulus of the semifluid membrane.

23. The nanostructure of any one of claims 1 to 22, wherein the applied force comprises an applied load.

24. The nanostructure of any one of claims 1 to 23, wherein the base module and the beam module comprise distance-reporting fluorophores.

25. The nanostructure of claim 24, wherein the distance-reporting fluorophores are selected from the group consisting of: Cy3 / Cy5; Alexa Fluor 488 / 555; Alexa Fluor 555 / 647; quantum dotbased FRET; lanthanide-based FRET; ATTO dye series; and DyLight fluorophore pairs.

26. The nanostructure of any one of claims 1 to 25, wherein the base module and the beam module comprise quantum dots.

27. The nanostructure of any one of claims 24 to 26, wherein the range of motion of the beam module relative to the base module is correlated to a FRET fluorescence intensity ratio.

28. The nanostructure of any one of claims 6 to 27, wherein the beam module is configured to undergo an angular rotation about the fulcrum point from the second conformation to the first conformation in response to removal of the actuation trigger.

29. The nanostructure of claim 28, wherein the removal of the actuation trigger comprises addition of one or more opening strands that are comprised of a single stranded nucleic acid sequence that completely or partially hybridize with the one or more actuation trigger sequences.

30. The nanostructure of any one of claims 4 to 29, wherein the actuation of the nanostructure may be repeated over a plurality of cycles in response to sequential addition and removal of the actuation trigger.31 . The nanostructure of any one of claims 1 to 30, wherein the nanostructure is constructed using DNA origami techniques.

32. The nanostructure of any one of claims 1 to 31 , wherein the base module is comprised of plurality of bundled DNA duplexes.

33. The nanostructure of any one of claims 1 to 32, wherein the base module comprises at least 10, at least 15, at least 20, at least 25 and around 30 or more DNA duplexes.

34. The nanostructure of any one of claims 1 to 33, wherein the base module has a vertical height relative to the surface of the semifluid membrane of at least 5 nm, at least 7nm, at least 10 nm, at least 12 nm, or at least 15 nm.

35. The nanostructure of any one of claims 1 to 34, wherein the base module has a horizontal width along an axis parallel to the surface of the semifluid membrane of at least 3 nm, at least 4nm, at least 6nm, at least 8nm, at least 10nm, at least 12nm, at least 14nm, or at least 16nm.

36. The nanostructure of any one of claims 1 to 35, wherein the base module has a horizontal length along an axis parallel to the surface of the semifluid membrane of at least 5 nm, at least 8nm, at least 10nm, at least 15nm, at least 20nm, at least 25nm.

37. The nanostructure of any one of claims 1 to 36, wherein the beam module is comprised of plurality of bundled DNA duplexes.

38. The nanostructure of any one of claims 1 to 37, wherein the beam module comprises at least 20 DNA duplexes, at least 30, at least 40, at least 50, at least 60, at least 70, at least 80, and optionally around 90 or more DNA duplexes.

39. The nanostructure of any one of claims 1 to 38, wherein the beam module has a vertical height relative to the surface of the semifluid membrane of around at least 10 nm, at least 20 nm, at least 25 nm, at least 30 nm, at least 35 nm, at least 40 nm, at least 45 nm, or at least 50 nm.

40. The nanostructure of any one of claims 1 to 39, wherein the beam module has a horizontal width along an axis parallel to the surface of the semifluid membrane of at least 3 nm, at least 4nm, at least 6nm, at least 8nm, at least 10nm, at least 12nm, at least 14nm, or at least 16nm.41 . The nanostructure of any one of claims 1 to 40, wherein the beam module has a horizontal length along an axis parallel to the surface of the semifluid membrane of at least 5 nm, at least 8nm, at least 10nm, at least 15nm, at least 20nm, at least 25nm.

42. The nanostructure of any one of claims 1 to 41 , wherein the beam module comprises a distal tip that defines a surface area configured to bear upon and deform the surface of the semifluid membrane.

43. The nanostructure of claim 42, wherein the surface area of the distal tip is configured to be at least around 9 nm2, around 12 nm2, around 14 nm2, around 16 nm2, around 18 nm2,around 20 nm2, around 24 nm2, around 28 nm2, around 36 nm2, around 42 nm2, around 56 nm2, around 64 nm2, around 72 nm2, around 84 nm2, around 92 nm2, or around 128 nm2.

44. The nanostructure of any one of claims 1 to 43, wherein the nanostructure comprises a scaffold strand comprising all or part of a bacteriophage genome sequence.

45. The nanostructure of claim 44, wherein the bacteriophage genome comprises a sequence is selected from: phiX174; lambda; M13mp18 and variants thereof; M13mp19; f1 ; fd; and P1 .

46. The nanostructure of any one of claims 1 to 45, wherein the nanostructure comprises a scaffold strand having a sequence comprised within SEQ ID NO: 1 and complimentary staple strands comprising SEQ ID NOs: 2-113.

47. The nanostructure of any one of claims 1 to 46, wherein the base module comprises at least one hydrophobic anchor that facilitates adjoining to the surface of the semi-fluid membrane, optionally wherein the base module comprises a plurality of hydrophobic anchors.

48. The nanostructure of claim 47, wherein the at least one hydrophobic anchor is attached to a membrane-facing side of the base module or a portion thereof such that the at least one hydrophobic anchor molecule is orientated to interact with and / or extend perpendicularly into the semi-fluid membrane.

49. The nanostructure of any one of claims 47 or 48, wherein the at least one hydrophobic anchor molecule is selected from the group consisting of: a lipid; and a porphyrin.

50. The nanostructure of claim 49, wherein the lipid is selected from the group consisting of: sterols; alkylated phenols; flavones; saturated and unsaturated fatty acids; and synthetic lipid molecules (including dodecyl-beta-D-glucoside).51 . The nanostructure of claim 50, wherein: the sterols are selected from the group consisting of: cholesterol; derivatives of cholesterol; phytosterol; ergosterol; and bile acid; the alkylated phenols are selected from the group consisting of: methylated phenols; dolichols and tocopherols; the flavones are selected from the group consisting of: flavanone containing compounds; and 6-hydroxyflavone; the saturated and unsaturated fatty acids are selected from the group consisting of: derivatives of lauric acid; oleic acid; linoleic acid; and palmitic acids; and / orthe synthetic lipid molecule is dodecyl-beta-D-glucoside.

52. A composition comprising a nanostructure as defined in any one of claims 1 to 51 .

53. A composition comprising a plurality of nanostructures as defined in any one of claims 1 to 51 maintained in suspension.

54. A sensor nanodevice comprising a nanostructure as defined in any one of claims 1 to 51.

55. A membrane elasticity sensor nanodevice comprising a nanostructure as defined in any one of claims 1 to 51.

56. A method for measuring a mechanical property associated with a semifluid membrane, wherein the semifluid membrane defines substantially planar surface, the method comprising the steps of: attaching a nanoscale structure to the semifluid membrane surface, wherein the nanoscale structure comprises a probe tip, and wherein the nanoscale structure is configured to transition through a range of motion from a first conformation to a second conformation upon application of an actuation trigger, and wherein in the second conformation the probe tip applies a load force to the surface of the membrane sufficient to induce deformation of the membrane; and determining the amount of deformation of the membrane, converting the determination of the amount of deformation into a measurement of a mechanical property associated with the semifluid membrane.

57. The method of claim 56, wherein the applied load force is perpendicular to a plane defined by the surface of the semifluid membrane.

58. The method of any one of claims 56 or 57, wherein the amount of deformation of the semifluid membrane is correlated to an amount of the applied load force.

59. the method of any one of claims 56 to 58, wherein the mechanical property is selected from: elastic stiffness, including Young's modulus; membrane tension; and / or a viscoelastic property.

60. The method of claim 59, wherein the mechanical property is a Young’s modulus of the semifluid membrane.61 . The method of any one of claims 56 to 60, wherein the nanoscale structure is as defined in any one of claims 1 to 51 .

62. The method of any one of claims 56 to 60, wherein the method comprises a plurality of nanoscale structures as defined in any one of claims 1 to 51.

63. The method of any one of claims 56 to 62, wherein the semifluid membrane comprises a biological cellular or sub-cellular membrane.

64. The method of any one of claims 56 to 63, wherein the semifluid membrane is contained within a cell or sub-cellular compartment.

65. The method of any one of claims 56 to 62, wherein the semifluid membrane comprises a synthetic membrane.

66. Use of a nanostructure, a composition, a sensor nanodevice or a membrane elasticity sensor nanodevice as defined in any one of claims 1 to 51 in a method for diagnosis or prognosis of a pathological condition within a human or animal subject.

67. The use of claim 66, wherein the pathological condition is selected from: cancer; cardiovascular disease; neurodegenerative disease; or infectious disease.

68. Use of a nanostructure, a composition, a sensor nanodevice or a membrane elasticity sensor nanodevice as defined in any one of claims 1 to 51 in a method for assessing the effect of a xenobiotic compound or substance on a cell, cell culture or membrane containing cellular extract.

69. The use of claim 68, wherein the xenobiotic compound or substance is a therapeutic or candidate therapeutic compound or substance, and the use is in a method for drug discovery or drug development.

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