Optical Near-Field Electron Microscopy
Raphaël Marchand, Radek ?achl, Martin Kalbá?, Martin Hof, Rudolf Tromp, Mariana Amaro, Sense J. van der Molen, Thomas Juffmann
OOptical Near-Field Electron Microscopy
Rapha¨el Marchand,
1, 2
Radek ˇSachl, Martin Kalb´aˇc, Martin Hof, RudolfTromp,
4, 5
Mariana Amaro, Sense J. van der Molen, and Thomas Juffmann
1, 2 University of Vienna, Faculty of Physics, VCQ, A-1090 Vienna, Austria University of Vienna, Max Perutz Laboratories,Department of Structural and Computational Biology, A-1030 Vienna, Austria J. Heyrovsk´y Institute of Physical Chemistry of the Czech Academy of Sciences, Dolejˇskova 3, 182 23 Prague, Czech Republic IBM T.J. Watson Research Center, 1101 Kitchawan Road, Yorktown Heights, New York 10598, USA Huygens-Kamerlingh Onnes Laboratory, Leiden Institute of Physics, Leiden University, Leiden, The Netherlands
Imaging dynamical processes at interfaces and on the nanoscale is of great importance throughoutscience and technology. While light-optical imaging techniques often cannot provide the necessaryspatial resolution, electron-optical techniques damage the specimen and cause dose-induced arte-facts. Here, Optical Near-field Electron Microscopy (ONEM) is proposed, an imaging techniquethat combines non-invasive probing with light, with a high spatial resolution read-out via electronoptics. Close to the specimen, the optical near-fields are converted into a spatially varying electronflux using a planar photocathode. The electron flux is imaged using low energy electron microscopy,enabling label-free nanometric resolution without the need to scan a probe across the sample. Thespecimen is never exposed to damaging electrons.
Keywords:
Electron Microscopy, Near-field optics, LEEM.
I. INTRODUCTION
Interfaces are of utmost importance throughout sci-ence, technology, industry, biology, and medicine. Imag-ing dynamical processes happening at interfaces can yieldcrucial information on the underlying processes, rang-ing from electroplating, to corrosion, to protein dynam-ics in lipid bilayers. Despite great progress over the lastdecades, there is currently no microscopy technique thatcan image dynamics at interfaces with nanometric resolu-tion, label-free, damage-free, and over extended periods:Optical super-resolution microscopy [1–6] has shownremarkable results over the last two decades, offering aresolution in the single digit nanometer range for selectedapplications (see e.g. [7, 8] and references therein). But,besides phototoxicity, fluorescence-based methods face atrade-off between frame rate, accuracy and observationtime, due to the finite number of photons that can becollected per fluorophore. And while high labelling den-sity can affect biological function [9], low labelling densitycan lead to severe statistical artefacts due to blinking andbleaching [10]. Label-free optical techniques (such as in-terferometric scattering microscopy (iSCAT) [11, 12] orplasmonics enhanced protein characterization [13]) en-able the detection and weighing of proteins, but theirspatial resolution is diffraction limited. Resolving this is-sue by decreasing the wavelength to the X-ray regime isnot an option for dynamic single molecule studies, as theyare precluded by the trade-off between signal to noise ra-tio and damage [14].Scanning probe techniques enable atomic resolution insurface imaging [15], and sub-nanometer resolution inimaging membrane bound proteins [16], but can perturbsoft membranes in high speed imaging [17]. Also, thescanning of the probe can limit imaging frame rates andfield of view. Similar trade-offs are faced in near-field scanning optical microscopes, where additionally the fi-nite probe size often limits the spatial resolution to tensof nanometers [18].Electron optical techniques enable the determina-tion of the ensemble-averaged atomic structure of pro-teins [19], and the imaging of the proteome of a cell [20].However, these techniques require frozen samples, whichprecludes real-time imaging of dynamics. Recently, thinliquid cells have enabled the observation of proteins intheir native environment within electron microscopeswith nanometer resolution [21]. However, dose-induceddamage leads to artefacts in electrochemical studies andlimits extended dynamical studies of sensitive materials[22]. Recently, Transmission Electron Microscopy at eV-energies (eV-TEM) was developed by some of us [23], todecrease electron-induced damage while imaging. In fact,eV-TEM has been combined with Low Energy ElectronMicroscopy (LEEM) so as to image samples in electrontransmission and reflection at energies of 0 −
100 eV [23].Although first results of the combination of these tech-niques are promising, this method is yet to prove itselffor dynamical processes.Techniques that correlate light and electron mi-croscopy are promising alternatives to the techniques dis-cussed above [24]. Traditionally, light microscopy is firstused for live imaging, or for imaging with fluorescence-enabled specificity, and electron microscopy is then usedto retrieve one final high resolution snapshot of the spec-imen under study.Here, we propose a novel technique that combines non-invasive probing with light with a read-out based on elec-tron optics offering nanometric resolution. Probing andread-out are coupled via a photocathode placed in theoptical near-field of the scattered light, where resolutionis not limited by the optical wavelength. We will firstdescribe the idea in more detail, then discuss its techno- a r X i v : . [ phy s i c s . a pp - ph ] F e b Figure
1. Optical Near-field Electron Microscopy (ONEM).Visible light illuminates a specimen (e.g. protein) in a liquidcell (LC). The resulting optical near-field interference patternis converted into a spatially varying electron flux via a pho-tocathode (PC). The spatial information is retrieved usingaberration-corrected electron microscopy. Scalebar: 5 nm. logical feasibility, its theoretical resolution and contrast,and lastly explore the potential application space of Op-tical Near-field Electron Microscopy (ONEM).
II. CONCEPT
The proposed method is sketched in Figure 1: Visiblelight is focused onto a sample (e.g. a protein), whichis close to an ultra-thin vacuum-liquid interface (e.g. ina liquid cell (LC) [25, 26]). The sample scatters lightelastically, leading to nanometric features in the near-field. This spatial light distribution is then converted,still in the near-field, into a spatially varying electronflux via the photoelectric effect within a thin layer of low-work-function photocathode (PC) material [27, 28]. Theemitted photoelectrons are then imaged using aberration-corrected low-energy electron optics [29, 30]. The elec-trons therefore provide a non-destructive read-out of thenanometric optical near-fields.The proposed technique would be intrinsically damagefree: First, low-work-function photocathodes can be effi-ciently excited with green light [27], and thus at photon energies significantly below the absorption band of mostproteins. The inset in Figure 1 shows the simulated near-field intensity distribution caused by a 50 kDa protein inwater and at 5 nm distance from the photocathode. Near-field simulations predict a signal to background ratio ofabout 1 . × photoelectrons per spa-tially resolved area to achieve a signal to noise ratio of1. Assuming a photoelectron conversion efficiency of 3 %(A conversion efficiency of ≈
15 % was measured for a25 −
30 nm thick photocathode [27] with excitation at2 . . × photons. Fora frame rate of 1 kHz and 5 nm resolution, the requiredillumination intensity is about 2 . µ W µ m − , i. e. wellbelow intensities reported in e.g. interferometric scatter-ing microscopy (iSCAT) using light at 405 nm [11]. Thisis not surprising: Both iSCAT and ONEM are shot-noiselimited. While iSCAT detects scattered light more ef-ficiently (no photocathode), this is by far compensatedby the narrower point spread function in ONEM, whichincreases contrast and resolution.Second, the photocathode material will have a workfunction much lower than the work function of its liquidcell support (1 . CsSb [27],1 . . hν = 2 . λ = 532 nm, where h is the Planck’s constant, ν thefrequency of the excitation light, and λ its wavelengthin vacuum), such that the energy of the created pho-toelectrons will be insufficient to overcome the internalpotential barrier. The sample is therefore not exposed toelectrons. III. FEASIBILITY
The feasibility of ONEM relies on three technologiesthat have recently been developed independently.First, electrons emitted from the photocathode haveto be imaged with nanometer resolution. Interestingly,similar requirements are to be met in LEEM [29, 30] andeV-TEM [23]. Typical LEEM systems have a lateralresolution of about 5 nm. In the last decade however,aberration-corrected Low Energy Electron Microscopy(AC-LEEM) has been introduced, with an optimal res-olution down to 1 . µ m [29, 30]. Moreover, using novel algorithms, manyimages can now be stitched together smoothly. Thus, theeffective field of view can be dramatically extended, with-out loss of resolution [33]. Furthermore, we note that aresolution smaller than 3 nm has been predicted for PhotoElectron Emission Microscopy (PEEM) on such state-of-the-art systems [29]. Figure 2 shows how ONEM canbe implemented within an existing LEEM setup. The Figure
2. Schematic ONEM setup. A specimen within a liq-uid cell is illuminated (solid green line) and optically inspected(dashed green line) with a Continuous Wave (CW) laser. Pho-toelectrons (solid red line) generated in the photocathode atthe backside of the liquid cell are imaged in an aberration-corrected LEEM. For this, the electrons travel through a setof lenses and are reflected by an aberration-correcting mir-ror. The electron gun at the top is off in ONEM, but it canbe turned on to perform LEEM experiments, e. g. on thephotocathode material (dashed red line).
LEEM sample holder will be modified to allow for opticalrear-illumination of the sample. The low work-functionphotocathode faces the LEEM objective lens, which ex-tracts the emitted photoelectrons in a strong electrostaticfield of 100 kV cm − . The electrons are accelerated to thecolumn potential of 15 keV, after which they follow thered solid path in Figure 2, from the photocathode, viaan electron mirror (for aberration correction) and elec-tron lenses, to the camera. The use of miniature objec-tives [34] within the light illumination path would allowfor a correlative read-out using light optics (dashed greenline), which could provide additional information (e.g.molecular specificity via fluorescent labelling).Second, in order to reach sub-optical-wavelength res-olution the distance of the object to the photocathodelayer needs to be much shorter than the optical wave-length. Ultra-thin, highly efficient photocathodes [27, 28]have just been developed, and have been shown to besmooth on the nanometer scale [35]. Most importantly,these photocathodes are efficient at green light excita-tion. Since these photocathodes are not to be exposed toair, they have to be prepared in situ , e.g. via evaporation,sputter deposition, or pulsed laser deposition [36] from asolid photocathode target [37]. Alternatively, alkali met-als or other photocathode materials could be stabilized in between graphene layers [27, 38], potentially allowingfor an ex situ preparation of the required photocathode.Third, the study of specimens in their natural environ-ment seems incompatible with the vacuum requirementsfor electron optics and the operation of photocathodes.Recently however, this has become possible by the use ofultrathin interfaces between vacuum and air or liquid en-vironments [39, 40]. The latter have been developed forliquid cell TEM [25], and modalities for electrochemistryexperiments, photoactivation of specimens, and in situ specimen mixing are now commercially available. Thistoolbox can be directly applied for ONEM. Note thatONEM liquid cells do not need to be ultra-thin, as itis light (and not electrons) that passes through the liq-uid, potentially allowing for more elaborate and robustsample manipulation.The advance of the three techniques described abovemakes ONEM technologically feasible, allowing fordamage-free measurements on dynamic processes in a liq-uid. Next, we discuss the contrast and resolution thatONEM could deliver. IV. CONTRAST AND RESOLUTION
In the following section we will simulate the expectedcontrast and resolution of ONEM. All simulations werecalculated using the MNPBEM toolbox [31]. We as-sume that the sample is excited with an x -polarizedplane wave travelling in the z -direction (reference field): ~ E ref ( ~ r , t ) = E ref e i ( kz − ωt ) ~ u x , where k = 2 π n m λ − ,with λ = 532 nm the wavelength of the reference fieldin vacuum, ω its angular frequency, n m = 1 .
33 the re-fractive index of water, and ~ u x a unitary vector in thex-direction. The particle of interest (e.g. protein, goldnanoparticle, copper cluster,. . . ) is assumed to be spher-ical, with its center at the origin of the coordinate sys-tem (Figure 3a). The intensity distribution I ( ~ r ) re-sults from interfering the scattered field ~ E scat ( ~ r , t ) withthe reference field ~ E ref ( ~ r , t ): I ( ~ r ) ∝ || ~ E tot ( ~ r , t ) || = || ~ E ref ( ~ r , t ) + ~ E scat ( ~ r , t ) || . Figure 3b, 3c, and 3d showthe intensity distribution obtained for a protein in wa-ter (radius R = 2 . n p = 1 . z = 5 nm, z = 50 nm, and z = 5 µ m,respectively.On the z-axis ( y = x = 0), we observe the followingbehaviour: In the near-field region ( R < z (cid:28) λ ), (Figure3a, 3b, and 3c), the scattered field is out of phase withthe reference field, resulting in a total on-axis intensity, I (0 , , z ), smaller than the intensity of the reference fieldalone I ∝ E ref . In the far field ( λ (cid:28) z , Figure 3d), thescattered field is in phase with the reference field, leadingto I (0 , , z ) > I .These results can be understood considering the modelof an ideal radiating dipole, excited by the reference field.In this case, the total dipole moment of the particle isgiven by ~ p ( t ) = (cid:15) m α ~ E ref ( ~ , t ), where (cid:15) m is the permit- Figure
3. a) 3-dimensional drawing of a spherical nanoparti-cle and the interference pattern I ( x, y ) obtained on a screen(photocathode) in the near-field ( z = 5 nm). b-d) simu-lated interference pattern I ( x, y ) for a spherical protein ofradius R = 2 . n p = 1 .
44, at: b) z = 5 nm (scale bar: 10 nm, ∆ I/I = 8 × − ), c) z = 50 nm(scale bar: 100 nm, ∆ I/I = 6 × − ), d) z = 5 µ m (scalebar: 1 µ m, ∆ I/I = 8 × − ). e) simulated Michelson con-trast C = ( I max − I min )( I max + I min ) − of the interferencepattern I ( x, y ) as a function of the distance of the screen,for gold (Au) particles, copper (cu) particles, proteins (all R = 2 . R = 22 . I ( x, y ) for a particle ofradius R = 2 . z to the par-ticle (the FWHM is found to be independent of the particlematerial). tivity of the surrounding medium, and α is the complexpolarizability of the particle, given by α = 3 V (cid:15) − (cid:15) m (cid:15) +2 (cid:15) m ,where V is the volume of the particle, and (cid:15) the com-plex permittivity of the particle (see e. g. ch. 5.2in [41]). For a protein in water, α , (cid:15) , and (cid:15) m arereal, and the dipole moment is in phase with the ex-citation field, which is consistent with the model of aLorentz oscillator driven at frequencies far below the (ma-terial dependant) resonance frequency (ch. 3.5 in [42]).The on-axis ( x = 0, y = 0) scattered field is given by ~ E scat (0 , , z, t ) = p ( t ) e i ( kz − ωt ) π(cid:15) m ( k z + ikz − z ) ~ u x (ch. 9.2. Figure
4. Michelson contrast of the simulated interfer-ence pattern I ( x, y ) for a nanoparticle, as a function of thenanoparticle diameter, for z = 25 nm. in [43]). We find that the results of the simulations agreewith the analytical expression to better than 2 % (seeSuppl. Figure S1). For z (cid:28) /k , the − /z term domi-nates, and the scattered field is anti-parallel to the refer-ence field. For 1 /k (cid:28) z , the 1 /z term dominates, and thescattered field is parallel to the reference field, in agree-ment with Figure 3b, 3c, and 3d. For λ = 532 nm and n m = 1 .
33, the transition occurs at z = 1 /k ≈
64 nm (seeSuppl. Figure S2).The simulated Michelson contrast C = ( I max − I min ) / ( I max + I min ) as a function of the distance to thescreen (i.e. the photocathode) is shown in Figure 3e fordifferent particle materials. The refractive indices of theprotein and the virus were taken from [11] and [44], re-spectively. For gold (Au) and copper (Cu), the complexrefractive indices were interpolated at 532 nm from datain [45] and [46], respectively. In the near-field the con-trast is found to drop as C ∝ /z , which will enableexcellent suppression of signal from scatterers that arenot bound to the interface. Figure 3f shows the resolu-tion of ONEM as a function of z , evaluated by the FullWidth at Half Maximum (FWHM) of the central featurein the interference pattern along the x -direction. Thecalculation was again performed assuming a R = 2 . ∂F W HM/∂z = 0 . R dependenceof the Michelson contrast. For metallic nanoparticles thecontrast is seen to saturate for particle radii >
20 nm.
V. POTENTIAL APPLICATION SPACE
By enabling dynamic studies of interfaces on thenanometer scale, ONEM has applications ranging frommaterials science to membrane biology.One of them is the study of plasmonic fields. It ischallenging to characterize light-matter interactions anddevices based on optical near-fields, on the nanometerscale. Laser triggered electron microscopy has provento be a versatile tool for mapping optical near-fields onnanometer spatial and femtosecond temporal scales [47–49]. ONEM could enable this in liquid environments, andat much lower optical excitation powers, allowing for aprecise characterization, and consequent further develop-ment, of plasmonic (bio)sensors [13, 50, 51] .ONEM could also be applied in electrochemistry,studying for example the nucleation and growth ofnanoscale copper clusters in a liquid environment. Suchexperiments were the first application of in situ liquidcell TEM [39], starting a new area of research that hasgrown rapidly over the last decade [25], both in materialscience and in biology. However, a serious and unresolvedissue with such experiments is that the high-energy elec-tron beam passes through the electrochemical cell abovethe working electrode, creating a large number of free,solvated electrons. These can strongly affect the electro-chemistry [22, 52], and may even lead to the formationof hydrogen bubbles [53]. ONEM could be used to ob-serve the nucleation and growth during electrodepositionwith nanometer resolution, and without electron-dose in-duced artefacts. Successful implementation of this pro-totype liquid cell experiment will open the door to in-vestigating many electrochemical problems that are ofsignificant industrial interest and that are currently outof reach for high-resolution, real-time studies (e.g. corro-sion, mass transport in batteries, swelling, liquid crystalswitching...).Finally, ONEM could also be used to image proteinsinteracting on, in, or with biomembranes under condi-tions mimicking a native membrane environment. It hasbeen shown that lipid bilayers can be formed on sur-faces with the use of cushions or tethers to allow for theembedding of proteins in the membrane, in particulartransmembrane proteins, while maintaining the mobilityand functionality of the inserted proteins [54–56]. If thisis done on the vacuum-liquid interface, ONEM could beused for continuous high resolution imaging. First ex-amples could include the visualization of supramolecu-lar protein assemblies formed on lipid bilayers. In gen-eral, oligomerization of membrane proteins into multi-component units is often critical for their function, ordysfunction. For instance, a change in the aggregationbehavior of a pro-apoptotic protein Bax, subsequent for-mation of Bax complexes with a broad distribution ofoligomerization numbers, and finally the opening of func-tional pores in the mitochondrial membrane representkey steps in programmed cell death [57]. Similarly, supra-molecular in-membrane assemblies consisting of 7 − /z , unwanted protein signal from the solution is ef-ficiently suppressed. It can therefore be used for imag-ing membrane related processes even in the cases wherethe equilibrium between bound and unbound protein isshifted in favour of the unbound protein. VI. DISCUSSION AND OUTLOOK
Optical Near-field Electron Microscopy (ONEM) rep-resents a fundamentally new microscopy technique thatexploits light and electron optics in an unprecedentedway, and according to their respective strengths: lightfor non-invasive probing, electrons for high-spatial-resolution read-out. Light and electrons are coupled us-ing an ultra-thin photocathode, and superresolution isenabled by the near-field components of the scatteredlight. ONEM is therefore ideally suited for the damage-and label-free study of physical, chemical, or biologicalprocesses happening at interfaces. A resolution below5 nm, and dynamic imaging over extended periods, seemfeasible. Various experimental difficulties will have tobe overcome to realize such specifications. A low energyelectron microscope has to be modified to include a cus-tom sample chamber, speckle-free optical excitation, aswell as in situ photocathode coating capabilities. Coat-ing procedures will have to be optimized for sensitivityand homogeneity, and potential issues such as photocath-ode ageing, poisoning or charging will have to be ad-dressed.ONEM can be operated, and extended, in many ways:Measurements can be performed in liquid, gas, or vac-uum. They can be performed label-free, characterizingthe refractive index distribution of a sample, or use labels(e.g. metal nanoparticles, fluorophores) bound to an ob-ject of interest. Correlative in situ light microscopy canprovide additional information, e.g. on the 3-dimensionalenvironment of the interface, optionally with fluorescenceenabled specificity. Polarization-dependent scatteringcross-sections (e.g. chiral molecules or chiral photonicstructures) could enable shot-noise limited measurementsin challenging environments. Pump-probe measurementscould make studies on ultra-short timescales possible.ONEM offers unique measurement opportunities onthe dynamics of various processes, by allowing for the useof liquid cells: Optical (plasmonic) properties of nano-structured materials can be probed in a liquid environ-ment, which can yield information that is essential forbio-optical sensor design. Electrochemical experimentswill yield novel insights into physical and chemical pro-cesses at interfaces, some of which play an importantrole in the energy transition. Finally, the exploration oftethered lipid bilayers will allow for the investigation ofbiological systems, offering information on protein clus-tering and dynamics within biological membranes.Given recent developments in technology and method-ology, we consider ONEM a feasible new form of mi-croscopy. Clearly, several practical challenges are stillto be overcome, but once these are addressed, ONEMwill offer unique opportunities for damage-free imagingof dynamic processes at interfaces.
Acknowledgements
We thank Peter S. Neu for help with the figures. Thisproject has received funding from the European Union’sHorizon 2020 research and innovation programme undergrant agreement No 101017902. TJ acknowledges sup-port from the ERC MicroMOUPE Grant 758752. RMacknowledges funding from the European Union’s Frame-work Programme for Research and Innovation Horizon2020 (2014-2020) under the Marie Curie Sk lodowskaGrant Agreement Nr. 847548. MA and MH acknowl-edge GA ˇCR grant 19-26854X. [1] S. W. Hell and J. Wichmann, “Breaking the diffrac-tion resolution limit by stimulated emission: stimulated-emission-depletion fluorescence microscopy,”
Optics let-ters , vol. 19, no. 11, pp. 780–782, 1994.[2] R. Heintzmann and T. Huser, “Super-resolution struc-tured illumination microscopy,”
Chemical Reviews ,vol. 117, no. 23, pp. 13890–13908, 2017. PMID: 29125755.[3] M. J. Rust, M. Bates, and X. Zhuang, “Sub-diffraction-limit imaging by stochastic optical reconstruction mi-croscopy (storm),”
Nature methods , vol. 3, no. 10,pp. 793–796, 2006.[4] E. Betzig, G. H. Patterson, R. Sougrat, O. W. Lind-wasser, S. Olenych, J. S. Bonifacino, M. W. Davidson,J. Lippincott-Schwartz, and H. F. Hess, “Imaging in-tracellular fluorescent proteins at nanometer resolution,”
Science , vol. 313, no. 5793, pp. 1642–1645, 2006.[5] T. Dertinger, R. Colyer, G. Iyer, S. Weiss, and J. Ender-lein, “Fast, background-free, 3d super-resolution opticalfluctuation imaging (sofi),”
Proceedings of the NationalAcademy of Sciences , vol. 106, no. 52, pp. 22287–22292,2009.[6] N. Gustafsson, S. Culley, G. Ashdown, D. M. Owen,P. M. Pereira, and R. Henriques, “Fast live-cell conven-tional fluorophore nanoscopy with ImageJ through super-resolution radial fluctuations,”
Nature Communications ,vol. 7, pp. 1–9, aug 2016.[7] Y. M. Sigal, R. Zhou, and X. Zhuang, “Visualizing anddiscovering cellular structures with super-resolution mi-croscopy,”
Science , vol. 361, no. 6405, pp. 880–887, 2018.[8] S. J. Sahl, S. W. Hell, and S. Jakobs, “Fluorescencenanoscopy in cell biology,” sep 2017.[9] E. A. Specht, E. Braselmann, and A. E. Palmer, “A crit-ical and comparative review of fluorescent tools for live-cell imaging,”
Annual Review of Physiology , vol. 79, no. 1,pp. 93–117, 2017. PMID: 27860833.[10] F. Baumgart, A. M. Arnold, B. K. Rossboth,M. Brameshuber, and G. J. Sch¨utz, “What we talk aboutwhen we talk about nanoclusters,”
Methods and applica-tions in fluorescence , vol. 7, no. 1, p. 013001, 2018.[11] M. Piliarik and V. Sandoghdar, “Direct optical sensing ofsingle unlabelled proteins and super-resolution imagingof their binding sites,”
Nature communications , vol. 5,no. 1, pp. 1–8, 2014.[12] G. Young, N. Hundt, D. Cole, A. Fineberg, J. Andrecka,A. Tyler, A. Olerinyova, A. Ansari, E. G. Marklund,M. P. Collier, S. A. Chandler, O. Tkachenko, J. Allen,M. Crispin, N. Billington, Y. Takagi, J. R. Sellers, C. Eichmann, P. Selenko, L. Frey, R. Riek, M. R. Galpin,W. B. Struwe, J. L. P. Benesch, and P. Kukura, “Quanti-tative mass imaging of single biological macromolecules,”
Science , vol. 360, no. 6387, pp. 423–427, 2018.[13] R. Gordon, “Biosensing with nanoaperture optical tweez-ers,”
Optics & Laser Technology , vol. 109, pp. 328–335,2019.[14] R. Henderson, “The potential and limitations of neu-trons, electrons and x-rays for atomic resolution mi-croscopy of unstained biological molecules,”
Quarterlyreviews of biophysics , vol. 28, no. 2, pp. 171–193, 1995.[15] G. Binnig, H. Rohrer, C. Gerber, and E. Weibel, “7 × Phys.Rev. Lett. , vol. 50, pp. 120–123, Jan 1983.[16] A. Engel and H. E. Gaub, “Structure and mechanicsof membrane proteins,”
Annu. Rev. Biochem. , vol. 77,pp. 127–148, 2008.[17] T. Ando, “High-speed atomic force microscopy and itsfuture prospects,”
Biophysical reviews , vol. 10, no. 2,pp. 285–292, 2018.[18] Y.-C. Yong, Y.-Z. Wang, and J.-J. Zhong, “Nano-spectroscopic imaging of proteins with near-field scan-ning optical microscopy (nsom),”
Current Opinion inBiotechnology , vol. 54, pp. 106 – 113, 2018. AnalyticalBiotechnology.[19] E. Nogales, “The development of cryo-em into a main-stream structural biology technique,”
Nature methods ,vol. 13, no. 1, pp. 24–27, 2016.[20] J. Mahamid, S. Pfeffer, M. Schaffer, E. Villa, R. Danev,L. K. Cuellar, F. F¨orster, A. A. Hyman, J. M. Plitzko,and W. Baumeister, “Visualizing the molecular sociol-ogy at the hela cell nuclear periphery,”
Science , vol. 351,no. 6276, pp. 969–972, 2016.[21] U. M. Mirsaidov, H. Zheng, Y. Casana, and P. Matsu-daira, “Imaging protein structure in water at 2.7 nm reso-lution by transmission electron microscopy,”
Biophysicaljournal , vol. 102, no. 4, pp. L15–L17, 2012.[22] T. J. Woehl, T. Moser, J. E. Evans, and F. M. Ross,“Electron-beam-driven chemical processes during liquidphase transmission electron microscopy,” sep 2020.[23] D. Geelen, J. Jobst, E. Krasovskii, S. Van Der Molen,and R. Tromp, “Nonuniversal transverse electron meanfree path through few-layer graphene,”
Physical reviewletters , vol. 123, no. 8, p. 086802, 2019.[24] A. Walter, P. Paul-Gilloteaux, B. Plochberger, L. Sefc,P. Verkade, J. G. Mannheim, P. Slezak, A. Unterhu-ber, M. Marchetti-Deschmann, M. Ogris, K. B¨uhler,
D. Fixler, S. H. Geyer, W. J. Weninger, M. Gl¨osmann,S. Handschuh, and T. Wanek, “Correlated multimodalimaging in life sciences: Expanding the biomedical hori-zon,”
Frontiers in Physics , vol. 8, p. 47, 2020.[25] N. De Jonge and F. M. Ross, “Electron microscopyof specimens in liquid,”
Nature nanotechnology , vol. 6,no. 11, pp. 695–704, 2011.[26] M. Textor and N. de Jonge, “Strategies for prepar-ing graphene liquid cells for transmission electron mi-croscopy,”
Nano letters , vol. 18, no. 6, pp. 3313–3321,2018.[27] H. Yamaguchi, F. Liu, J. DeFazio, M. Gaowei, C. W.Narvaez Villarrubia, J. Xie, J. Sinsheimer, D. Strom,V. Pavlenko, K. L. Jensen, et al. , “Free-standing bial-kali photocathodes using atomically thin substrates,”
Ad-vanced Materials Interfaces , vol. 5, no. 13, p. 1800249,2018.[28] H. Yuan, S. Chang, I. Bargatin, N. C. Wang, D. C. Riley,H. Wang, J. W. Schwede, J. Provine, E. Pop, Z.-X. Shen, et al. , “Engineering ultra-low work function of graphene,”
Nano letters , vol. 15, no. 10, pp. 6475–6480, 2015.[29] R. Tromp, J. Hannon, A. Ellis, W. Wan, A. Berghaus,and O. Schaff, “A new aberration-corrected, energy-filtered leem/peem instrument. i. principles and design,”
Ultramicroscopy , vol. 110, no. 7, pp. 852–861, 2010.[30] R. Tromp, J. Hannon, W. Wan, A. Berghaus, andO. Schaff, “A new aberration-corrected, energy-filteredleem/peem instrument ii. operation and results,”
Ultra-microscopy , vol. 127, pp. 25–39, 2013.[31] U. Hohenester and A. Tr¨ugler, “Mnpbem–a matlab tool-box for the simulation of plasmonic nanoparticles,”
Com-puter Physics Communications , vol. 183, no. 2, pp. 370–381, 2012.[32] N. Barrett, E. Conrad, K. Winkler, and B. Kr¨omker,“Dark field photoelectron emission microscopy of micronscale few layer graphene,”
Review of Scientific Instru-ments , vol. 83, no. 8, p. 083706, 2012.[33] T. A. de Jong, D. N. Kok, A. J. van der Torren, H. Schop-mans, R. M. Tromp, S. J. van der Molen, and J. Jobst,“Quantitative analysis of spectroscopic low energy elec-tron microscopy data: High-dynamic range imaging,drift correction and cluster analysis,”
Ultramicroscopy ,vol. 213, p. 112913, 2020.[34] L. Yang, J. Wang, G. Tian, J. Yuan, Q. Liu, andL. Fu, “Five-lens, easy-to-implement miniature objec-tive for a fluorescence confocal microendoscope,”
Opt.Express , vol. 24, pp. 473–484, Jan 2016.[35] J. Feng, S. Karkare, J. Nasiatka, S. Schubert, J. Smed-ley, and H. Padmore, “Near atomically smooth alkali an-timonide photocathode thin films,”
Journal of AppliedPhysics , vol. 121, no. 4, p. 044904, 2017.[36] D. Dijkkamp, T. Venkatesan, X. D. Wu, S. A. Shaheen,N. Jisrawi, Y. H. Min-Lee, W. L. McLean, and M. Croft,“Preparation of y-ba-cu oxide superconductor thin filmsusing pulsed laser evaporation from high tc bulk mate-rial,”
Applied Physics Letters , vol. 51, no. 8, pp. 619–621,1987.[37] M. Gaowei, Z. Ding, S. Schubert, H. B. Bhandari, J. Sin-sheimer, J. Kuehn, V. V. Nagarkar, M. S. J. Marshall,J. Walsh, E. M. Muller, K. Attenkofer, H. J. Frisch,H. Padmore, and J. Smedley, “Synthesis and x-ray char-acterization of sputtered bi-alkali antimonide photocath-odes,”
APL Materials , vol. 5, no. 11, p. 116104, 2017.[38] M. Kalb´aˇc, L. Kavan, M. Zukalov´a, and L. Dunsch, “Two positions of potassium in chemically doped c60 peapods:An in situ spectroelectrochemical study,”
The Journal ofPhysical Chemistry B , vol. 108, no. 20, pp. 6275–6280,2004. PMID: 18950111.[39] M. Williamson, R. Tromp, P. Vereecken, R. Hull, andF. Ross, “Dynamic microscopy of nanoscale clustergrowth at the solid–liquid interface,”
Nature materials ,vol. 2, no. 8, pp. 532–536, 2003.[40] Y. Han, K. X. Nguyen, Y. Ogawa, J. Park, and D. A.Muller, “Atomically thin graphene windows that enablehigh contrast electron microscopy without a specimenvacuum chamber,”
Nano letters , vol. 16, no. 12, pp. 7427–7432, 2016.[41] C. F. Bohren and D. R. Huffman,
Absorption and scat-tering of light by small particles . John Wiley & Sons,1998.[42] E. Hecht,
Optics . Pearson Education, 2017.[43] J. D. Jackson,
Classical electrodynamics . John Wiley &Sons, 1998.[44] H. Ewers, V. Jacobsen, E. Klotzsch, A. E. Smith, A. He-lenius, and V. Sandoghdar, “Label-free optical detectionand tracking of single virions bound to their receptorsin supported membrane bilayers,”
Nano letters , vol. 7,no. 8, pp. 2263–2266, 2007.[45] P. B. Johnson and R.-W. Christy, “Optical constantsof the noble metals,”
Physical review B , vol. 6, no. 12,p. 4370, 1972.[46] E. D. Palik,
Handbook of optical constants of solids II ,vol. 3. Academic press, 1998.[47] B. Barwick, D. J. Flannigan, and A. H. Zewail, “Photon-induced near-field electron microscopy,”
Nature , vol. 462,no. 7275, pp. 902–906, 2009.[48] L. Piazza, T. Lummen, E. Quinonez, Y. Murooka,B. Reed, B. Barwick, and F. Carbone, “Simultaneousobservation of the quantization and the interference pat-tern of a plasmonic near-field,”
Nature communications ,vol. 6, no. 1, pp. 1–7, 2015.[49] O. Kfir, H. Louren¸co-Martins, G. Storeck, M. Sivis,T. R. Harvey, T. J. Kippenberg, A. Feist, and C. Rop-ers, “Controlling free electrons with optical whispering-gallery modes,”
Nature , vol. 582, no. 7810, pp. 46–49,2020.[50] R. W. Taylor, R. J. Coulston, F. Biedermann, S. Maha-jan, J. J. Baumberg, and O. A. Scherman, “In situ sersmonitoring of photochemistry within a nanojunction re-actor,”
Nano letters , vol. 13, no. 12, pp. 5985–5990, 2013.[51] S.-H. Oh and H. Altug, “Performance metrics and en-abling technologies for nanoplasmonic biosensors,”
Na-ture communications , vol. 9, no. 1, pp. 1–5, 2018.[52] T. J. Woehl, K. L. Jungjohann, J. E. Evans, I. Arslan,W. D. Ristenpart, and N. D. Browning, “Experimentalprocedures to mitigate electron beam induced artifactsduring in situ fluid imaging of nanomaterials,”
Ultrami-croscopy , vol. 127, pp. 53–63, 2013.[53] J. M. Grogan, N. M. Schneider, F. M. Ross, and H. H.Bau, “Bubble and pattern formation in liquid induced byan electron beam,”
Nano Letters , vol. 14, pp. 359–364,jan 2014.[54] R. Mach´aˇn and M. Hof, “Recent developments in fluo-rescence correlation spectroscopy for diffusion measure-ments in planar lipid membranes,”
International journalof molecular sciences , vol. 11, no. 2, pp. 427–457, 2010.[55] F. Roder, S. Waichman, D. Paterok, R. Schubert,C. Richter, B. Liedberg, and J. Piehler, “Reconstitu- tion of membrane proteins into polymer-supported mem-branes for probing diffusion and interactions by sin-gle molecule techniques,”
Analytical chemistry , vol. 83,no. 17, pp. 6792–6799, 2011.[56] K. R. Poudel, D. J. Keller, and J. A. Brozik, “Single par-ticle tracking reveals corralling of a transmembrane pro-tein in a double-cushioned lipid bilayer assembly,”
Lang-muir , vol. 27, no. 1, pp. 320–327, 2011.[57] Y. Subburaj, K. Cosentino, M. Axmann, E. Pedrueza-Villalmanzo, E. Hermann, S. Bleicken, J. Spatz, andA. J. Garc´ıa-S´aez, “Bax monomers form dimer units inthe membrane that further self-assemble into multiple oligomeric species,”
Nature communications , vol. 6, no. 1,pp. 1–11, 2015.[58] J. P. Steringer, S. Lange, S. ˇCujov´a, R. ˇSachl, C. Poo-jari, F. Lolicato, O. Beutel, H.-M. M¨uller, S. Unger,¨U. Coskun, et al. , “Key steps in unconventional secretionof fibroblast growth factor 2 reconstituted with purifiedcomponents,”
Elife , vol. 6, p. e28985, 2017.[59] B. Antonsson, S. Montessuit, B. Sanchez, and J.-C. Martinou, “Bax is present as a high molecularweight oligomer/complex in the mitochondrial mem-brane of apoptotic cells,”
Journal of Biological Chem-istry , vol. 276, no. 15, pp. 11615–11623, 2001.
Supplemental Materials forOptical Near-Field Electron Microscopy
I. ON-AXIS SCATTERED FIELD: COMPARISON BETWEEN ANALYTICAL EXPRESSION ANDNUMERICAL SIMULATIONS
Here we demonstrate the excellent agreement obtained between simulations using the MNPBEM toolbox [1] andthe analytic expression describing the field scattered by a dipole. Taking the expression of the x-polarized referencefield from the main text, as well as the expression of the dipole moment ~ p ( t ), the polarizability α , and the on-axisscattered field ~ E scat (0 , , z, t ), leads to: ~ E scat (0 , , z, t ) = 3 V π (cid:15) − (cid:15) m (cid:15) + 2 (cid:15) m E ref e i ( kz − ωt ) ( k z + ikz − z ) ~ u x ( S1 )The amplitude of the on-axis scattered field, normalized to the amplitude of the reference field is therefore: || ~ E scat (0 , , z, t ) || E ref = 3 V π (cid:12)(cid:12)(cid:12)(cid:12)(cid:12) (cid:15) − (cid:15) m (cid:15) + 2 (cid:15) m ( k z + ikz − z ) (cid:12)(cid:12)(cid:12)(cid:12)(cid:12) ( S2 )Figure S1 shows the excellent agreement between simulations and the analytical result from Eq. S2 , in the case ofa spherical protein. Note that the permittivities are real in this case. The terms ∝ /z n are plotted to distinguish thenear-field domain ( E scat ∝ z − ) and the far-field domain ( E scat ∝ z − ). The boundary between the near-field andthe far-field domains occurs at z ≈
64 nm.
FIG. S1. Amplitude of the on-axis scattered field for a spherical protein, normalized to the reference field: comparison betweendata simulated with the MNPBEM toolbox and the analytical expression S2 . The dashed lines give the individual contributionsof the terms ∝ /z n . The following parameters were used: (cid:15) = n , (cid:15) m = n m , n = 1 .