Uniform, Large-Area, Highly Ordered Peptoid Monolayer and BilayerFilms for Sensing ApplicationsDaniel J. Murray,†,∥ Jae Hong Kim,†,∥ Elissa M. Grzincic,† Samuel C. Kim,‡ Adam R. Abate,‡,§

and Ronald N. Zuckermann†,*†The Molecular Foundry, Lawrence Berkeley National Laboratory, 1 Cyclotron Road, Berkeley, California 94720, United States‡Department of Bioengineering and Therapeutic Sciences, University of California, San Francisco, California 94158, United States§Chan Zuckerberg Biohub, San Francisco, California 94158, United States

*S Supporting Information

ABSTRACT: The production of atomically defined, uniform,large-area 2D materials remains as a challenge in materialschemistry. Many methods to produce 2D nanomaterials sufferfrom limited lateral film dimensions, lack of film uniformity, orlimited chemical diversity. These issues have hindered theapplication of these materials to sensing applications, whichrequire large-area uniform films to achieve reliable andconsistent signals. Furthermore, the development of a 2Dmaterial system that is biocompatible and readily chemicallytunable has been a fundamental challenge. Here, we report asimple, robust method for the production of large-area,uniform, and highly tunable monolayer and bilayer films, fromsequence-defined peptoid polymers, and their application ashighly selective molecular recognition elements in sensor production. Monolayers and bilayer films were produced on thecentimeter scale using Langmuir−Blodgett methods and exhibited a high degree of uniformity and ordering as evidenced byatomic force microscopy, electron diffraction, and grazing incidence X-ray scattering. We further demonstrated the utility ofthese films in sensing applications by employing the biolayer interferometry technique to detect the specific binding of thepathogen derived proteins, shiga toxin and anthrax protective antigen, to peptoid-coated sensors.

■ INTRODUCTION

Two-dimensional (2D) nanomaterials such as graphene andmetal chalcogenides have had gained enormous interest due totheir promising structural, optical, electronic, and mechanicalproperties.1,2 Inspired by the huge opportunities of thesematerials in a variety of applications, organic analogs of these2D nanomaterials have recently been investigated by manygroups.3−5 Notably, the use of polymer building blocks toconstruct 2D materials through a self-assembly approach hasenabled high chemical tunability and biocompatibility to beachieved.6,7 One of the most attractive building blocks arepeptoid polymers, bioinspired oligomers of sequence-definedN-substituted glycines. One key feature of peptoids is theincredible diversity of side-chain residues that can beincorporated on its backbone. Additionally, precise control ofchain length and monomer sequence can be achieved throughthe solid-phase submonomer method.8 The lack of chiralityand hydrogen bond donors in the peptoid backbone enablesthe interactions between polymer chains to be programmedthrough side chain selection.9 This provides a means to designand control the self-assembly process of these polymers andresults in the ability to create supramolecular peptoidassemblies with atomically defined structural features and

high crystallinity.10−15 Adopting these unique properties,amphiphilic diblock peptoid polymers have been synthesizedwhich assemble into atomically defined 2D nanomaterials withuniform thickness (∼2.7 nm) and high area/thickness ratio(>109 nm−1) at the air−water or oil−water interface.16−19

Furthermore, molecular engineering of these nanosheets18 bychemical tailoring of the peptoid strands to integrate molecularrecognition elements creates a robust platform that is ideallysuited for biomedical applications such as biosensing and theremediation of threat agents. Nevertheless, in order tosuccessfully scale production of these materials to an industrialscale and create practical devices, there still remain significantchallenges in fabrication. The key steps requires simultaneouscontrol over several length scales. Sensing devices, for example,could require the preparation of homogeneous peptoidmonolayer or bilayer films up to the centimeter scale.Therapeutic applications would likely require production ofuniformly sized peptoid nanosheets on the micron scale.

Received: August 15, 2019Revised: September 24, 2019Published: October 11, 2019

Article

pubs.acs.org/LangmuirCite This: Langmuir 2019, 35, 13671−13680

© 2019 American Chemical Society 13671 DOI: 10.1021/acs.langmuir.9b02557Langmuir 2019, 35, 13671−13680

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This work addresses these challenges and demonstrates theutility of peptoid monolayers and bilayers films in molecularrecognition by employing the surface-based platform ofbiolayer interferometry (BLI). The technique of BLI hasattracted a great deal of interest as a real-time, label-free, high-throughput, and high sensitivity biosensing technique.20 Thismethod is based on the measurement of optical interferencepatterns generated by white light reflected off a functionalsurface and that of an internal reference surface. As analytesbind to the functional surface, a shift in the interference patternis measured and related back to specific binding interactions.In order to obtain specific and sensitive response for targets, itrequires the functional surface to be modified by theconjugation of functional chemical or biomolecular recognitionelements. Although several strategies for the conjugation offunctional biocomponents on a desired substrate have beendeveloped,21 there remain shortcomings, such as uncontrolleddisplay density and molecular orientation. This can result in aloss of binding activity and an increase of background signal.22

The deposition of functionalized peptoid monolayers integrat-ing molecular recognition elements onto a BLI sensor maymitigate these issues and provide a highly tunable platform.Using this method, molecular recognition elements incorpo-rated as a “loop domain” insert in the middle of a peptoidsequence can be displayed on a hydrophilic and zwitterionic

nanosheet surface that helps prevent nonspecific binding.18,23

Furthermore, peptoids with loop domains or “loopoids” areversatile tools for the chemical conjugation and display offunctional ligands through the addition of desired aminemonomers in the loop domain. For instance, alkyne-containingloopoid nanosheets were conjugated with azide-functionalizedcarbohydrates via click chemistry and were shown to have highand specific binding affinity for target proteins owing to themultivalency originating from the high display density ofcarbohydrates on the nanosheet surface.24

Herein, we report the fabrication of large-area and uniformpeptoid nanofilms on silicon and glass substrates based onLangmuir−Blodgett techniques. A peptoid monolayer film wasprepared at the air−water interface in a Langmuir trough andthen successfully deposited on the substrate by the Langmuir−Blodgett (LB) method. Peptoid bilayer films were alsoassembled by the deposition of second monolayer onto thehydrophobic face of a monolayer nanofilm by the Langmuir−Schaefer (LS) method. The uniformity, high ordering, large-area of prepared peptoid nanofilm was demonstrated by acombination of optical microscopy, atomic-force microscopy,electron microscopy, and X-ray diffractometry. Importantly,this coating method is applicable to a wide variety of peptoidsequences, making the functionality of the resulting films easilytunable (Table 1). Using these methods, we also show that the

Table 1. Peptoids Used in This Study to Prepare Monolayer and Bilayer-Coated Surfaces

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peptoid films can be used to make peptoid nanosheets withuniform shape and size through a photolithography-basedapproach using a photoreactive peptoid. Additionally, loop-functionalized peptoid monolayers were coated onto the tip ofoptical fibers and used as a BLI biosensing platform. On thebasis of the structural properties of the resultant multivalentpeptoid nanofilms, we confirmed significant improvement ofBLI signal against multimeric toxin-related proteins, such asshiga toxin 1 subunit B (STX 1B) and anthrax protectiveantigen [(PA63)7], through a multivalent manner.

■ RESULTS AND DISCUSSIONPeptoid Monolayer Preparation and Deposition by

LB Technique. The scale-up of free-floating nanosheets inaqueous solution has been previously achieved by a vial-rocking method,15 which allows repeated lateral surfacecompressions to be applied to a key assembly intermediate, aspontaneously forming monolayer at the air−water or oil−water interface.25 Although this method provides multimilli-gram quantities of nanosheets, they are small in size (<1 mm)and have a broad distribution of shape and size. This hindersapplication of these materials for sensing applications whichrequire a uniform coating of material, often over a large-area.To address this problem, we employed the LB technique toprepare large (centimeter scale) and uniform peptoidmonolayers and subsequently bilayers at the air−waterinterface in a Langmuir trough (Figure 1).A major disadvantage of this method is the requirement of

large subphase volumes (∼100 mL or more) which cause anenormous consumption of peptoid stock if the standardmethod of peptoid adsorption from bulk solution is utilized formonolayer formation.15 In order to alleviate this problem, wedirectly spread a small volume of peptoid from a concentratedstock solution onto the air−water interface. Specifically, 10 μLof peptoid 1 stock solution (2 mM in 1:1 water/DSMO) wasapplied to the surface of 100 mL subphase (2 mM Tris buffer,pH 8.0) in a Langmuir trough. We use this amount of peptoidto provide a slight excess of molecules relative to the amountrequired to form exactly one monolayer. This accounts forsome loss to the bulk and provides a small reservoir of

molecules to allow for the monolayer equilibration process.The DMSO in the stock solution is used to prevent self-assembly in the stock solution and dissolves into the subphaseupon spreading. Upon the spreading of 1, the surface pressurerapidly increased to ∼16 mN m−1 due to immediate spreadingof peptoid strands at the interface (Supporting InformationFigure S1). After this initial spreading, the surface pressurebegan to gradually decrease, indicating that desorption ofpeptoids is faster than adsorption at this stage. After a period ofabout 2 min, the surface pressure leveled out and a gradual riseof surface pressure was observed, indicating that adsorption ofpeptoid strands to the interface eventually dominates. Thissurface pressure increase eventually stabilized to reach anequilibrium state at ∼22 mN m−1. When this equilibrium statewas reached, the film was compressed to 30 mN m−1, justbelow the collapse pressure, to form the compressed peptoidmonolayer at the interface. On our trough, this resulted in amonolayer size of 82 cm2. Details of the peptoid monolayerformation and collapse of this peptoid were studied in detail.26

We next transferred the peptoid monolayer onto the surfaceof hydrophilic or hydrophobic substrates. For a hydrophilicsubstrate, monolayers of 1 were transferred on the surface ofplasma cleaned Si/SiO2. The transfer of 1 monolayer wasaccomplished by the immersion of the substrate into thesubphase before spreading peptoids on the interface andwithdrawing the substrate at a rate of 1 mm/min through theinterface after peptoid monolayer formation. As the monolayerwas transferred off the air/water interface, the surface pressurewas held constant at 30 mN m−1, and a decrease in the troughsurface area was observed. This decrease in area was consistentwith the area of the substrate, giving a transfer ratio close to 1which indicates that peptoid monolayer was uniformly coatedon the surface of substrate. Transfer of the monolayer at theequilibrium spreading pressure of 22 mN m−1 resulted in lowertransfer ratios and thus a less uniform coating. The Si/SiO2substrate allows observation of the deposition of 1 monolayerby reflectance optical microscopy (Figure 1b). We found that 1monolayer was extensively covered on the substrate withoutobservable defects. Moreover, the contrast homogeneitysuggested that 1 monolayer had a uniform thickness. On the

Figure 1. (a) Schematic illustration of fabrication of peptoid monolayer film by Langmuir−Blodgett transfer method. (b) Reflectance optical imageof B28 monolayer on Si/SiO2. The coated portion appears a lighter blue on the left, and the uncoated portion is darker blue on the right. (c and d)AFM images of peptoid 1 and loop-inserted (loopoid 1) monolayer film. (e) SAED pattern of peptoid 1 monolayer film.

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basis of the height measurement by atomic force microscopy(AFM), the thickness was approximately 1.2 ± 0.2 nm, whichis consistent with the expected height of the peptoid strandsthat are laying with the backbone parallel to the substrate(Figure 1c).We also confirmed that peptoid monolayer could be coated

on to hydrophobic substrate using a 2-phenylethyl-function-alized Si/SiO2 substrate. This substrate was lowered downthrough the compressed monolayer at 1 mm/min and thenremoved after the subphase had been aspirated clean. Opticalimaging showed that monolayers transferred in this fashion areuniformly coated without observable defects just as those seenfor transfer to hydrophilic Si/SiO2 (Supporting InformationFigure S2). Static contact angle measurements of thismonolayer showed a water contact angle of 34°, in contrastto 86° for the bare phenethyl modified Si/SiO2. These contactangles were observed to be stable, indicating the films were notdelaminating from the substrate. This shows that monolayerscan be coated with either the hydrophilic side or thehydrophobic side exposed. The ability to transfer themonolayer with the hydrophilic side exposed will enable thecreation of surfaces coated with surface-functionalized peptoidmonolayers for aqueous analyte binding applications.We further demonstrated that this method of monolayer

formation and transfer was applicable to other peptoidsequences. An AFM image of a monolayer from a loopinserted peptoid, loopoid 1, which was transferred to aphenethyl modified Si/SiO2 surface, is shown in Figure 1d.The measured thickness of the monolayer was 2.5 ± 0.3 nm.This thickness is indicative of a monolayer that has the loopdomains protruding from the surface.18 Transfer to thehydrophobic surface results in the loop domains being exposedon the substrate surface. The ability to the coat surfaces in thisway leads to the possibility of fabricating surfaces with a large-area uniform coating of monolayers bearing molecularrecognition elements.To investigate the internal ordering to the monolayer, we

performed selected area electron diffraction (SAED). Themonolayers were transferred to plasma treated ultrathin carbonTEM grids, by pulling them up through the monolayer asdescribed above. Diffraction was performed at 77 K in order toreduce beam damage. The patterns show an isotropic peak at

1.4 Å−1 which corresponds to a 4.5 Å spacing (Figure 1e).Diffraction at 4.5 Å in peptoid nanosheets has previously beenshown to result from the spacing between aligned peptoidchains.13

Peptoid Bilayer Formation by Langmuir−SchaeferTechnique. Since the LB deposition of the hydrophilic face ofa peptoid monolayer on hydrophilic Si/SiO2 substrate resultsin the exposure of a uniform layer of hydrophobic 2-phenylethyl groups, we sought to use the Langmuir−Schaefer(LS) method for transferring a second peptoid monolayer inan LB trough to fabricate the peptoid bilayer structure (Figure2a) that has previously been reported in peptoid nanosheets.13

This seeks to provide a large-area and uniform peptoid bilayerfilm that is not achievable using the current vial-rockingmethod.15 Before the deposition of the second peptoidmonolayer, the first peptoid monolayer film was dried undervacuum at 50 °C for 2 h to remove any remaining aqueoussolution between the monolayer and substrate to ensureadhesion. Subsequently, the LS deposition method was appliedto deposit a second peptoid monolayer. A transfer ratio couldnot be calculated since the subphase was aspirated clean beforeremoval of the substrate. Reflectance optical microscopeimages of the prepared film showed that it was homogeneouslydeposited across the substrate (Figure 2b). The edges of thepeptoid bilayer film were delaminated and folded back over onthe substrate, indicating that the resulting film has themechanical strength necessary to remain intact once it hasbeen removed from the support substrate (Figure 2c). Thethickness of the bilayers was shown to be 3.0 ± 0.2 nm byAFM (Figure 2d). This is consistent with previously reportedbilayer nanosheets. Moreover, the bilayer has a surfaceroughness similar to the underlying SiO2. This points to auniform packing of peptoids within the film.Staining the bilayer film with Nile red, known to stain

nanosheets in solution, and imaging by fluorescencemicroscopy reveals a direct correlation of emission withreflectance optical microscopy images (Supporting Informa-tion Figure S3). This gives further evidence for bilayerformation, as Nile red has been shown to selectivelyincorporate into the hydrophobic core of bilayer nanosheets.13

Staining of the monolayer films results in no fluorescenceemission.

Figure 2. (a) Schematic illustration of peptoid bilayer film by Langmuir−Schaeffer transfer method. (b, c) Reflectance optical microscopic imagesof peptoid 1 bilayer film on Si/SiO2. (d) AFM images and height profile of peptoid 1 bilayer film. (e) SAED pattern of peptoid 1 bilayer film.

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We further investigated the internal ordering of monolayersand bilayers of peptoid 1 using grazing-incidence wide-angle X-ray scattering (GIWAXS) and selected area electron diffraction(SAED) by transmission electron microscopy (TEM). In theGIWAXS patterns (Supporting Information Figure S4), bothmonolayer and bilayer on Si(100) have an in-plane peak at1.37 Å−1 corresponding to a 4.6 Å spacing which is not seen inthe out-of-plane direction. The higher intensity of this peak forthe bilayer is consistent with having an additional layer present.This spacing is indicative of the chain to chain spacing ofparallel polymers that was previously reported in floatingpeptoid monolayers.25 By comparing this data to that fromdiffraction from peptoid monolayers at the air−water inter-face,25 we can conclude that the chain alignment is preservedduring the LB transfer process. The absence of this peak in the

out-of-plane direction indicates alignment of the peptoidchains in the plane of the substrate, which is also consistentwith AFM height measurements discussed previously. Addi-tionally, a peak corresponding to the interlayer spacing ofmonolayers is seen at 0.41 A−1 with the second and thirdharmonic seen at 0.83 A−1 and 1.26 A−1 in the out-of-planedirection for the bilayer (Supporting Information Figure S4).This set of peaks is absent in the pattern for the monolayer.Analysis of the bilayer films by selected area electron

diffraction (SAED) was also performed. As for the monolayers,we performed the diffraction experiments at 77 K to minimizebeam damage to the films. Diffraction patterns of bilayers of 1(Figure 2e) show an isotropic peak at 1.4 Å−1 corresponding toa 4.5 Å spacing as seen in the monolayer films. This isconsistent with the above GIWAXS data and shows that the

Figure 3. SAED patterns of thermally annealed peptoid 1 monolayer (a) and bilayer (b).

Figure 4. (a) Schematic illustration of photopatterning of peptoid bilayer film by UV cross-linking method. The use of photoreactive peptoid 2allows bilayers to be deposited and then cross-linked with the desired pattern. (b, c, and d) Fluorescent microscopic images of peptoid nanosheetswith uniform shape such as squares, rectangles, and circles.

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films consist of aligned peptoid chains lying in the plane of thesubstrate they were transferred to.Monolayer and Bilayer Annealing. To date, all of the

reported diffraction data on films of peptoid 1 has shown anisotropic peak at 1.4 Å−1 which has been shown to result fromthe alignment of the peptoid backbones. Since this peak isisotropic, we know that the domain size of the aligned peptoidchains is much smaller than the spot size in the SAEDexperiments. In turn, this shows that the monolayer and bilayerfilms have little to no long-range ordering. We nextinvestigated if the long-range ordering of the films could beimproved using thermal annealing. Because the chain orderingis known to occur during the monolayer formation step,25 weused an annealing process in which the monolayer wasannealed at 50 °C for 3 h while being compressed to 30 mN/mat the air water interface. After the 3 h annealing, the

monolayer was transferred off the interface as previouslydescribed. SAED over an area of about 0.75 μm2 at 77 K of thisannealed monolayer shows a set of sharp spots at 1.4 Å−1 incontrast to the diffraction ring seen in the unannealedmonolayer (Figure 3a). This spot disappears after prolongedexposure to electron doses of ∼10 e−/Å2. The dramaticincrease of long-range order in the monolayer, as indicated bythe diffraction spots, led us to prepare bilayers using thismethod. For this, we used the LS method with two annealedmonolayers. The SAED patterns for the annealed bilayers showa set of spots at 1.4 Å−1 as well; however, there is now anadditional set at 1.4 Å−1 that is oriented in a different direction(Figure 3b). This second set is weaker in intensity indicatingthat most of the peptoid alignment is in the direction of themore intense spot. This demonstrates the importance of

Figure 5. Biosensing platform based on loop-functionalized peptoid monolayer-coated optical fiber by interferometry. (a) Scheme for the surfacemodification of an optical fiber by the LB technique to deposit a large and uniform peptoid monolayer. (b) Schematic illustration of the peptoidmonolayer-deposited optical fiber for the detection of protein binding by interferometry. (c) Binding (solid) and fitting (dashed) curve of loopoid2 monolayer against 500 nM STX 1B, loopoid 3 monolayer against 500 nM (PA63)7 (red and pink), and negative control sample (e.g., peptoid 1and lactose loopoid monolayers) against 500 nM (PA63)7 and Stx 1B (black). (d) Plot of maximum intensity of wavelength shift against theaverage of shortest distance between loops. The shortest distance between loops were adapted from the literature.20 (e) Demonstration of theregeneration and reusability of loopoid 2 and loopoid 3 coated sensor tips. Sensor tips were immersed into 10 mM HCl solution for 10 s at roomtemperature and then neutralized by using PBS buffer containing 0.1% tween 20 for 10 s.

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thermal annealing at the monolayer stage to create long-rangeorder in the subsequent films.Photopatterning of Peptoid Nanosheets for Size and

Shape Uniformity. By taking advantage of a methodpreviously described by our group27 for the production ofcovalently cross-linked nanosheets, we show that bilayer filmscan be photopatterned to control their lateral dimensions.Patterning of these films should enable the production ofbilayer peptoid structures of defined size and shape. Thegeneral method for photopatterned film production is shownin Figure 4a. First, bilayer films of peptoid 2 were deposited onSi/SiO2. Optical microscopy (Supporting Information FigureS5) showed that these films behave similar to those of peptoid1. Some delamination was observed along the edges of thesubstrate, but the bulk of the substrate was covered with acontinuous film. AFM imaging (Supporting Information FigureS5) showed that this film is slightly thicker than that of peptoid1. This is consistent with previous reports of nanosheets madein solution.Following bilayer film production, the films were irradiated

through a photomask with 254 nm light to photo-cross-link inthe hydrophobic core. The cross-linking renders the filminsoluble in organic solvents, allowing the unirradiated portionof the film to be removed by washing with ethanol. Finally, thedeveloped films were stained with Nile red and imaged withfluorescence microscopy (Figure 3b−d). We demonstrated thistechnique to produce three different shapes (squares,rectangles, circles), but the size and shape are purely functionsof the photomask and can be tailored to the desiredapplication.Loop-Functionalized Peptoid Monolayer as Bio-Layer

Interferometric Sensing Platform. The use of ourdeveloped method resulted in the large-area and uniformityof peptoid monolayer and bilayer; therefore, it can be useful forthe precise surface display of molecular recognition elements(e.g., peptides, peptoids, drugs, carbohydrates, etc.). Weapplied our method for the deposition of functionalizedloopoid monolayers to the tip of glass optical fibers to detectanalytes using BLI technique as shown in Figure 5a. For thedeposition of the loopoid monolayer, we first cleaned thesurface of optical fiber by immersion into 0.01 M NaOH for 1h, and then aromatic groups were attached to the surface bythe reaction with 2-phenylethylsilane vapor overnight. Usingthe resultant hydrophobic surface of the optical fiber, weintroduced the loopoid monolayer by Langmuir−Schaeferdeposition as described above. To demonstrate that themonolayer deposition occurred with the proper solutiondisplay of ligands, we first introduced a biotin-conjugatedpeptoid 3 monolayer and then observed the binding of AlexaFluor 647 conjugated streptavidin by fluorescent microscopy.We confirmed the binding activity of biotin groups on thesurface by the resultant increase of Alexa Fluor 647’sfluorescence within biotin-B36 monolayer (Supporting In-formation Figure S7). For the generation of interferometricsignal against interested targets as shown in Figure 5b, weemployed globotriose-conjugated loopoid 2 and peptide 6mer(TYWWLD)-inserted loopoid 3 which are well-knownfunctional ligands as binders for toxin-related proteins, suchas shiga toxin subunit 1B (Stx) and anthrax protective antigen(PA63)7, respectively.

28,29 We also used plain peptoid strandsand lactose-conjugated loopoid as negative controls. Upon Stxbinding to the monolayer of loopoid 2, the interferometricsignal increased and decreased in accordance with association

and dissociation phase (Figure 5c). In addition, there was nobinding to loopless control monolayers from peptoid 1 andlactose-conjugated monolayers from loopoid 3. These resultsindicate that globotriose moiety of loopoid 2 monolayers wereadequately exposed on the surface of optical fiber, whilemaintaining their binding activity and preventing nonspecificbinding. We also observed that selective response against(PA63)7 only appeared in case of loopoid 3. Therefore, large-area and uniform loopoid monolayer is suitable for the surfacemodification of optical fiber and the label-free real-timemeasurement of target binding by the BLI technique.On the basis of the BLI response curves, we studied binding

parameters (e.g., association/dissociation rate and bindingconstant) of monolayers of loopoid 2 and loopoid 3 againstStx 1B and (PA63)7. Using a two-to-one heterogeneous ligandmodel, the binding constants (KD) of the loopoid 2 andloopoid 3 monolayers were determined to be, approximately,15.5 and 31.1 nM, respectively. By contrast, the KD ofunivalent globotriose and half-maximal inhibitory constant(IC50) of TYWWLD for Stx 1B and (PA63)7 are known to beconsiderably weaker at 2 mM and 150 μM, respectively.30,31

The enhanced binding affinity from the millimolar ormicromolar range to the nanomolar range suggests that thesurface displayed ligands on the loopoid monolayer interactwith their target proteins in a multivalent manner.31,32 Inaddition, Battigelli et al. recently reported the enhancement ofbinding affinity to sugar-functionalized nanosheets throughmultivalent interactions with lectins and that the binding wasdependent upon the ligand display density.24 Thus, weexamined the relationship between the binding affinity andligand display density using a series of monolayers prepared bydilution of loopoid 2 with peptoid 1. We found a dramaticincrease of signal against Stx 1B binding when the globotrioseunits are less than 1.6 nm apart (on average) from each other.This threshold distance is in good accordance with the shortestdistance between binding sites on Stx 1B (∼1.4 nm as shownin Supporting Information Figure S8), showing that theaccumulation of globotriose on peptoid nanosheets inducedcooperative binding behavior of multiple carbohydrates. Thus,the multivalency from the molecular structure of the loopoidmonolayer can improve the sensitivity for multivalent targets.We further studied the reusability of BLI sensor tips made

from loopoid 2 and loopoid 3 via stripping of bound proteinsby the immersion of sensor tips into 10 mM HCl solution for10 s at room temperature. The binding efficiency was wellmaintained for three cycles at each different concentration ofproteins (Figure 5e), hence indicating good capacity forreusability of loopoid monolayers for BLI-based biosensingplatform. We also tested the stability of these loopoidmonolayer coated tips by the measurement of the bindingactivity using stored loopoid monolayer in 10 mM Tris buffer(pH 8.0) for 9 days at 4 °C. We obtained a similar bindingsignal and KD (∼20 nM) with the stored sample as those of thefresh sample (Supporting Information Figure S9). Takentogether, large-area and uniform loopoid monolayer depositioncan be versatile tools for the surface modification of the BLIsensor format to detect bioanalytes with high sensitivity andreliability.

■ CONCLUSIONHerein, we demonstrated the fabrication of large-area peptoidmonolayer and bilayer films with long-range order usingLangmuir techniques. These methods have been shown to

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result in films that are highly uniform as shown by opticalmicroscopy and AFM. The internal ordering of the films wasinvestigated by SAED and GIWAXS and revealed that thetransferred films consist of aligned peptoid strands that lie inthe plane of the substrate. This structure is locally similar tothat reported previously for the large-scale preparation of free-floating nanosheets in solution. Films can be fabricated from awide variety of peptoids that display surface features atcontrollable densities that can be used as molecular recognitionelements for sensing. As such, we have shown that by coatingsensors with monolayers of loopoids for use in BLI, we canfabricate highly sensitive, selective, and robust platforms for thedetection of select targets. The wide applicability of the filmfabrication methods to peptoids of very diverse chemicalfunctionalities results in the ability to tune the sensor chemistryto the desired analyte.

■ METHODSMaterials: Peptoid Synthesis, Protein Expression. All peptoid

strands were synthesized and purified according to previouslyreported procedures.18 β-Alanine tert-butyl ester was free-basedfrom the HCl salt, using 1 M aq NaOH, and extracted with DCMbefore preparation of 1 M solutions in DMF. All other amines werepurchased from commercial suppliers and used without furtherpurification. Peptoids were prepared using a Rink amide resin (200mg) from Protein Technologies with a loading of 0.64 mmol/g.Standard peptoid coupling proceeded in two steps, initialbromoacetylation with bromoacetic acid (0.8 M) and N,N-diisopropylcarbodiimide (DIC, 0.8 M) in N,N-dimethylformamide(DMF) for 2.5 min. Subsequent displacement with amine (1 M) inDMF lasted for 5 min. Chemical structures for all loopoids in thisstudy are summarized in Table 1.Shiga Toxin 1 Expression. The gene for shiga toxin 1, subunit B

(STX 1B), was cloned into a pMAL-c5x vector (New EnglandBiosciences, Ipswich, MA) with a 6X histidine tag, flexible (GGS)4linker, and tobacco etch virus (TEV) protease cleavage site directlyupstream of the gene (no maltose binding protein tag from the vectorwas included). The plasmid was constructed via Gibson Assembly(Gibson Assembly Master Mix, New England Biosciences, Ipswich,MA) of the linearized vector and the inset as a gene block with 20base pair overlaps (custom gBlocks Gene Fragments, Integrated DNATechnologies, Coralville, IA). The plasmid was transformed intochemically competent BL21 (DE3) E. coli (New England Biosciences,Ipswich, MA). For expression, 500 mL batches of Luria−Bertani (LB)medium were inoculated to 0.05 OD600 and allowed to grow to 0.6−0.8 OD600 at 37 °C with shaking at 250 rpm. The temperature wasthen lowered to 30 °C, and overexpression was induced with 1 mMisopropyl β-D-1-thiogalactopyranoside (IPTG) for 3 h. Cells wereharvested by centrifugation and stored as pellets at −80 °C untilprotein purification. Cells were redispersed in phosphate bufferedsaline pH 7.4 (PBS; 0.01 M phosphate buffer, 0.0027 M potassiumchloride and 0.137 M sodium chloride) with 10 mM imidazole andlysed with an EmulsiFlex C3 homogenizer (Avestin Inc., Ottawa, ON,Canada). Insoluble cell debris was removed by centrifugation, and thesupernatant was applied to a 1 mL HisTrap Excel (GE Healthcare LifeSciences, Marlborough, MA) Ni-NTA column on an AKTA FPLCsystem (GE Healthcare Life Sciences, Marlborough, MA) equilibratedwith PBS pH 7.4 with 10 mM imidazole. The column was washedwith 40 mM imidazole, and bound proteins were eluted with PBS pH7.4 with 500 mM imidazole. The protein was then dialyzed overnightinto PBS pH 7.4 at 4 °C; the histidine tag was cleaved with TEVprotease overnight at 4 °C, and the protein was then purified as apentamer via gel filtration with a Superdex 200 Increase 10/300 GLcolumn (GE Healthcare Life Sciences, Marlborough, MA). Sampleswere characterized by SDS-PAGE and Western blot (6X-His tagantibody-HRP conjugate; Invitrogen); concentrations were measuredby UV−vis spectroscopy, and samples were stored in PBS pH 7.4 at 4°C (Supporting Information Figure S10).

Preparation of Peptoid Monolayer by Langmuir−BlodgettTechnique. The Langmuir trough (Mini-trough, KSV Nima,Finland) was filled with a subphase of 2 mM tris buffer (pH 8),equipped a paper Wilhelmy plate, and the surface was aspirated. Aplasma-cleaned Si wafer with 300 nm of SiO2 was next submerged inthe subphase. Using a syringe, 10 μL of 2 mM peptoid solution (1:1water/DSMO) was spread on the interface. After spreading, 30 minwas allowed for the monolayer to come to equilibrium. Once this wasreached, the film was compressed at a rate of 10 cm2/min until apressure of 30 mN/m was obtained. After reaching this pressure, theSi/SiO2 wafer was pulled through the interface at a rate of 1 mm/min.During this process, the surface pressure was maintained at 30 mN/m,and the area of the trough decreased in direct correlation with thearea of the wafer being pulled though the interface.

Preparation of Peptoid Bilayer by Langmuir SchaefferTechnique. Bilayer deposition was accomplished through theLangmuir−Schaeffer technique. First, the monolayer was dried at50 °C under a vacuum for 2 h to remove trapped water from betweenthe film and the substrate and ensure maximum adhesion of themonolayer to the substrate. A peptoid film was prepared on theLangmuir trough, as described above, with the exception of thesubstrate placement. In this case, the substrate with a precoatedmonolayer is placed horizontally above the interface. Once the film iscompressed to 30 mN m−1, the substrate was lowered until it justtouched the interface. At that point, the interface was aspirated toremove the excess film and the substrate was lifted until the subphasedetached.

Fluorescence Microscopy. For the imaging of the bilayer filmsusing fluorescence microscopy, they were first stained with Nile red.For this, the substrates with the attached bilayer were submerged in 1μm Nile red in water for 10 min. They were then removed andwashed with pure water. After being stained, the films were imagedusing 590 nm excitation.

AFM. Bilayer and monolayer films were directly imaged by AFM.Films were dried at 50 °C under vacuum for 2 h before imaging. AFMimages were obtained on an Asylum MFP-3D AMF operated intapping mode. Budget Sensors Tap150G probes were used for allimaging.

TEM. Monolayer and bilayer samples were prepared on a plasma-etched continuous carbon TEM grid to allow for TEM analysis. Themonolayer was deposited by pulling the grid through a compressedfilm as described above. The bilayer was produced by dropping thegrid with the deposited monolayer onto a fresh film on the interface.The grid was then removed from the interface by placing a piece ofpaper over the grid on the interface and removing it. The grid wasattached to the paper and could be removed once the paper dried.The grid was washed 3× with pure water before analysis to ensurethat any residual buffer was removed. Electron diffraction patternswere obtained with a Libra 200MC at the National Center forElectron Microscopy.

GIWAXS. Grazing incidence wide-angle X-ray scattering data wascollected at the Advanced Light Source beamline 7.3.3. For theseexperiments, monolayers and bilayers were deposited on Si(111)using the methods described above. X-rays of 10 keV with an energybandwidth of E/ΔE = 100 were used. A Pilatus 2 M detector fromDectris was used. The calibration of sample detector distance andbeam center was performed with a silver behenate standard. The Nikapackage for IGOR pro was used to process the data.

Details of BLI Sensing Platform. BLI experiments wereperformed on an OctetRED96 BLI system (ForteBio, CA) usingamine-coated optical sensors. Sensors were cleaned by immersion into0.01 M sodium hydroxide solution for 1 h. Then, 2-phenylethylgroups were functionalized on the surface of the glass sensors by thevaporization of 2-phenylethyltrichlorosilane for 2 h. After the sensorswere functionalized, an annealed monolayer of the desired peptoidwas deposited using the LS method. Before observation of proteinbinding, sensors were equilibrated for 20 min in PBS buffer containing0.1% Tween 20. After reaching the equilibrium state, sensors weremoved into the solutions containing protein targets to obtain theassociation curve. Thereafter, the sensors were moved into PBS buffer

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to obtain a dissociation curve. The regeneration of the sensor tips wascarried out by using HCl treatment. Briefly, the surfaces of the sensortips were transferred into 10 mM HCl solution for 10 s at roomtemperature for the removal of bound proteins. The sensors were thenneutralized for 10 s in PBS buffer containing 0.1% Tween 20.

■ ASSOCIATED CONTENT*S Supporting InformationThe Supporting Information is available free of charge on theACS Publications website at DOI: 10.1021/acs.lang-muir.9b02557.

Surface pressure vs time for peptoid spreading, transferof monolayer B28 to phenethyl modified Si/SiO2,fluorescence imaging of nile red stained B28 bilayers,GIWAXS patterns for monolayer and bilayer B28, B28-p-Cl7 bilayer films, biotin-B36 coated BLI sensors,molecular structure of STX 1B and globotriose (PDB#∼1BOS), binding curves for monolayer coated tips after9 days, characterization of expressed STX 1B (PDF)

■ AUTHOR INFORMATIONCorresponding Author*E-mail: rnzuckermann@lbl.gov.ORCIDSamuel C. Kim: 0000-0001-5237-6358Adam R. Abate: 0000-0001-9614-4831Ronald N. Zuckermann: 0000-0002-3055-8860Author Contributions∥These authors contributed equally to this work.NotesThe authors declare no competing financial interest.

■ ACKNOWLEDGMENTSThis work was supported by the DARPA Fold F(x) programand conducted at the Molecular Foundry and Advanced LightSource at Lawrence Berkeley National Laboratory, both ofwhich are supported by the Office of Science, Office of BasicEnergy Sciences, U.S. Department of Energy under ContractNo. DEAC02-05CH11231. The authors thank Mark Kline andMichael Connolly for assistance with peptoid synthesis.

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