Comparison of Oligo (ethylene glycol) alkanethiols versus n

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Comparison of Oligo(ethylene glycol)alkanethiols versus n-Alkanethiols: Self-Assembly, Insertion, and Functionalization Mitchell J. Shuster,†,‡ Amit Vaish,†,§ Megan L. Gilbert,|| Michelle Martinez-Rivera,|| Roya M. Nezarati,|| Paul S. Weiss,*,†,‡,||,^,# and Anne M. Andrews*,†,^,r,O †

California NanoSystems Institute, ^Department of Chemistry & Biochemistry, #Department of Materials Science & Engineering, Department of Psychiatry, and OSemel Institute for Neuroscience & Human Behavior, University of California, Los Angeles, Los Angeles, California 90095, United States ‡ Department of Physics, §Department of Bioengineering, and Department of Chemistry, The Pennsylvania State University, University Park, Pennsylvania 16802, United States )

r

bS Supporting Information ABSTRACT: We describe the self-assembly and chemical functionalization of oligo(ethylene glycol)alkanethiol (OEG) molecules. Insertion of OEGs into n-alkanethiolate monolayer matrices depends considerably on terminal functionality, unlike insertion of n-alkanethiols. Thus, inserted fractions of OEGs cannot be inferred from related systems, yet tuning, to some extent, is possible by controlling insertion parameters. Furthermore, while the in situ reactivities of dilute inserted carboxyor amine-terminated OEGs versus n-alkanethiols protruding from the surrounding matrix are similar in amide bond formation reactions, complete monolayers of OEGs react to a greater extent compared to n-alkanethiols with similar terminal functionalities. We interpret these differences in terms of the reduced crystalline packing of terminal ethylene glycol groups of OEGs.

1. INTRODUCTION Self-assembled monolayers (SAMs) have found widespread application to impart tailored properties to noble metal surfaces because of their simplicity, ease of use, and patterning capabilities.116 In particular, n-alkanethiolate SAMs have been extensively studied because they are highly ordered, stable, and chemically versatile; a wide variety of terminal functional groups can be presented at the exposed interface, thereby determining the chemical, physical, and biological properties of these films.3,1726 Similar to n-alkanethiols, oligo(ethylene glycol)alkanethiols (OEGs) self-assemble on Au substrates. However, unlike n-alkanethiol SAMs, self-assembled monolayers of OEGs are characterized by resistance to biofouling.2733 Therefore, OEG monolayers are often used to produce surfaces targeted toward biological applications, especially for sensors and biomedical devices.11,13,14,16,3438 Previous studies have focused on OEG SAM structure, formation, and the mechanism of resistance to protein adsorption.7,30,31,33,3943 However, how the chemical reactivities of OEG molecules on surfaces compare with the reactivities of widely studied n-alkanethiols and their terminally functionalized derivatives has received relatively little attention. Molecular insertion into SAMs is used to place individual molecules, pairs of molecules, or larger clusters into controlled molecular environments.12,20,25,35,4451 The insertion technique enables control, albeit stochastic, at the molecular level by exploiting r 2011 American Chemical Society

defects in preformed monolayers to allow dilute intercalation of new molecules into a host SAM. The extent of insertion of alkanethiols can be controlled by modifying the type and density of defects in the initial monolayer. Deposition conditions and monolayer processing are used to control initial SAM quality. Exposure time and concentration of the molecule(s) to be inserted are also varied to control insertion.49 Molecules can be inserted across entire substrates or patterned using hierarchical strategies for lithography and insertion.22,26,49,52,53 Previously, we have used insertion-directed self-assembly to create biofunctionalized capture surfaces.35,36,52,54 While singlecomponent monolayer and codeposition preparations have been investigated as methods for biofunctionalized surface preparation,5,7,5558 there has been comparatively little study on the properties of preparations utilizing insertion-directed self-assembly for the creation of biocapture surfaces. Insertion is advantageous as a method for functional substrate fabrication because it offers control over the final surface composition and molecular distributions on substrates. In contrast, codeposition can suffer from the effects of phase separation, interactions of buried molecular functionalities, molecular geometries, for example, n-alkanethiol molecules outcompeting cage molecules because Received: August 3, 2011 Revised: October 30, 2011 Published: November 03, 2011 24778

dx.doi.org/10.1021/jp207396m | J. Phys. Chem. C 2011, 115, 24778–24787

The Journal of Physical Chemistry C

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Figure 1. Self-assembled monolayers. Molecular self-assembly and chemical reactivity of surface functional groups were compared under different densities and with respect to different base SAM chemistries. (A) Single-component (high density) OEG vs n-alkanethiolate carboxy- or aminefunctionalized monolayers. (B) Molecular insertion (low density; dilute) of OEG vs n-alkanethiolate carboxy- or amine-terminated thiols into preformed n-alkanethiol SAMs.

of their higher packing density, and surface dipole moments, for example, carboranethiol molecules with favorably interacting dipole moments outcompeting carboranethiol molecules with repulsive dipole interactions on the surface. All of these can result in molecular ratios on surfaces that do not reflect those in solution and surface distributions of molecules that are not random, such as linear chains or phase-separated clusters.6,7,5964 Better understanding of the properties of OEG molecules, both in single-component and mixed monolayer systems, will facilitate their utilization in surface preparations for a wide range of functional applications. Directly imaging OEG supramolecular assemblies with molecular resolution via scanning probe techniques is challenging, owing to the wide tunneling barrier imparted by the length of OEG molecules and the polarity and disorder of the OEG moieties.6569 As an alternative, we utilized infrared spectroscopy to investigate and to compare the properties of n-alkanethiolate and OEG SAMs.28,56,7072 Infrared spectroscopy enables ensemble measurements of the properties of single-component OEG SAMs or mixed monolayers composed of both OEG and n-alkanethiolate molecules. Moreover, it enables the measurement and quantification of the presence of specific chemical bonds, which is useful for monitoring chemical functionalization and for determining protonation states. Here, we compare the chemical availability and reactivity of OEG versus n-alkanethiol molecules with carboxyl or amine termination with respect to the assembly environment. We form single-component monolayers of mercaptohexadecanoic acid (MHDA), 23-(9-mercaptononyl)-3,6,9,12,15,18,21-heptaoxatricosanoic acid (HEG), 1-(9-mercaptononyl)-3,6,9,12,15,18hexaoxaundecan-11-amine (AEG), or 11-amino-1-undecanethiol (AAT) (Figure 1A). Otherwise, these molecules are inserted into n-alkanethiolate monolayers (Figure 1B). The chemistry of the terminal functional groups is not altered by their surface density. However, the availability of functional groups for chemical reaction depends on the local chemical environment. For insertion studies, the matrix molecules are selected to be shorter than molecules for insertion, enabling inserted molecules to protrude from the surrounding SAM matrix. This configuration results in greater conformational degrees of freedom and less steric hindrance than for single-component monolayers composed entirely of the functional molecules, where all molecules

are of the same length. This configuration is typically applied where subsequent reactions are used to prepare capture surfaces based on the availability of the functionality of the inserted molecules protruding from the surrounding monolayer matrix.10,35,36,49,54,73,74 For OEG and n-alkanethiolate monolayers, the differences in crystalline packing at the solution/monolayer interface significantly impact reactivity. The properties of OEG versus n-alkanethiol molecules are contrasted in three ways. First, we compare protonation/deprotonation and dimer configurations between carboxy-terminated OEG and carboxy-terminated n-alkanethiol molecules inserted at low density into n-alkanethiol SAMs.72,75 Next, we explore differences in the inserted fractions of OEGs into n-alkanethiolate SAMs based on differences in the terminal functional groups of the inserted molecules. Finally, we examine variations in chemical reactivity for carboxy-terminated versus amine-terminated molecules. Functional group reactivity is studied in both singlecomponent monolayers and for insertion at low density into n-alkanethiolate monolayers. Throughout this study, we employ methyl-terminated n-alkanethiolate monolayer matrices for investigating insertion into SAMs. Although OEG SAMs are typically used for biofunctionalized surfaces because of their antibiofouling properties,7,33,34,37,38 studies of insertion into OEG SAMs are complicated by a number of factors. Of relevance to this study, infrared spectroscopy is impeded by the use of OEG SAMs because of spectral interference between peaks from the backbone of the OEG SAM molecules and peaks of interest arising from the inserted molecules. We used n-alkanethiol SAMs to reduce the complexity of infrared spectra to facilitate understanding the observed properties. The use of n-alkanethiol SAMs for insertiondirected studies therefore represents a compromise between investigating systems relevant to biologically directed applications and systems for which metrology and analyses are practicable.

2. MATERIALS AND METHODS 2.1. Substrate Preparation. Substrates were prepared by 1 Å/s electron-beam deposition of 100 Å thick Cr followed by 1000 Å thick Au on silicon wafers (Silicon Quest, Santa Clara, CA) using a Kurt J. Lesker (Pittsburgh, PA) evaporator system. Immediately prior to SAM formation, substrates were flame annealed using a hydrogen flame. 24779

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Figure 2. Thiols used for insertion and/or monolayer formation. The red carbon chain indicates that all hydrogen atoms attached to the chain are substituted with deuterium atoms.

2.2. Chemicals. 1-(9-Mercaptononyl)-3,6,9,12,15,18-hexaoxaundecan-11-biotin (BEG) and AEG were purchased from ProChimia (Sopot, Poland), while HEG was purchased from Toronto Research Chemicals (Toronto, ON). Mercaptoundecanoic acid (MUDA), MHDA, 1-octanethiol (C8), N-ethyl-N-(dimethylaminopropyl)carbodiimide (EDC), diisopropylcarbodiimide (DIC), N-hydroxysuccinimide (NHS), 5-hydroxytryptamine hydrochloride (5-HT), glacial acetic acid, potassium hydroxide (KOH), triethylamine, and dimethylformamide (DMF) were obtained from Sigma-Aldrich (St. Louis, MO). Commercial grade ethanol was purchased from Pharmaco-AAPER (Brookfield, CT). Perdeuterated 1-dodecanethiol (D12) and AAT were obtained from Asemblon (Redmond, WA). 9-Fluorenylmethoxycarbonyl-5-hydroxytrytophan (FMOC-5-HTP) was purchased from Anaspec (San Jose, CA). All chemicals were used as received. Schematics of the thiol molecules are shown in Figure 2. 2.3. Monolayer Formation. Initial single-component SAMs of C8, D12, MHDA, HEG, AEG, and BEG were formed by immersing Au substrates in 1 mM ethanolic solutions of each thiol for 1 h. Over this time frame, these SAMs are expected to achieve 90100% of their maximum packing density and achieve a well-ordered state.76 Solutions of carboxy-terminated and amineterminated thiols included 5% glacial acetic acid or triethylamine, respectively, to improve monolayer quality.75 For dilute, mixed component SAMs, molecular insertion was achieved by incubating Au substrates with preformed monolayers in 1 mM ethanolic solutions of the inserted thiol for 1 h in the absence of acetic acid or triethylamine. After SAM formation and where applicable, substrates were thoroughly rinsed with ethanol and dried under a stream of N2 gas. 2.4. Acid/Base Rinsing. Exposure of carboxy-terminated thiols to alternating acidic and basic solutions was performed to investigate correlations between infrared spectral peaks and carboxy group protonation/deprotonation states. Acidic rinses consisted of 5% glacial acetic acid in 18.2 MΩ deionized water

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(v/v), while basic rinses were in 100 mM KOH(aq). Samples were submerged in either solution for 15 s and then carefully dried with a stream of N2. 2.5. Chemical Functionalization. Reactivity of carboxy and amine terminal functional groups was assessed by monitoring amide bond formation. For carboxy-terminated thiols, the terminal group was reacted with the primary amine of 5-HT, as described previously.35 Briefly, samples were immersed in an aqueous solution containing 15 mM NHS and 25 mM EDC for 1 h, rinsed with deionized water, and then transferred to a PBS pH 9.5 solution of 25 mM 5-HT. This sequence is shown in Scheme 1 (Figure 3). Monolayers containing or composed of amineterminated thiols were functionalized with FMOC-5-HTP to form amide bonds by incubation in solutions of 25 mM NHS, 25 mM EDC, 22 mM DIC, and 20 mM FMOC-5-HTP in DMF for 4 h.36 Upon removal, samples were rinsed with ethanol, dried under a stream of N2, and then incubated in 1:5 piperidine/DMF (v/v) for 15 min for FMOC deprotection. Samples were rinsed with ethanol and dried again under a stream of N2. This sequence is designated as Scheme 2 (Figure 3). 2.6. Infrared Reflection Adsorption Spectroscopy. Infrared (IR) spectra were obtained using a Thermo Nicolet 6700 infrared spectrometer (Thermo Electron Corp., Waltham, MA) equipped with a liquid nitrogen-cooled HgCdTe detector. Infrared reflectance adsorption spectroscopy (IRRAS) was performed using a Seagull variable-angle reflection accessory (Harrick Scientific Inc., Ossining, NY) at an angle of 84 to the surface normal and with p-polarized light. The spectrometer and accessories were purged with dry, CO2-free air from a FTIR purge gas generator (Parker-Balston, Cleveland, OH). Spectra are averages of 1024 scans at 2 cm1 resolution and 1.27 cm/s mirror speed. Spectra were normalized against D12 monolayers to avoid spectral interference with the methylene stretches of the alkyl backbones of the SAMs under study. Spectra intended for direct comparison were collected on the same day to minimize signal intensity variations due to day-to-day IR source variations.

3. RESULTS AND DISCUSSION 3.1. AcidBase Chemistry of Carboxy-Terminated Thiols in Self-Assembled Monolayers. The carboxy-terminated thiols

MHDA and HEG were selected for study based on their frequent use in the fabrication of biofunctionalized surfaces.35,36,52,56,58,67,77 These and other carboxy-terminated thiols are often employed because of the wide variety of chemistries available for tethering molecules of interest to carboxy groups. The presence of prominent and well-resolved diagnostic infrared spectral peaks in monolayer systems makes carboxy-terminated molecules excellent for IRRAS analysis.28,35,56,72,78 Carbonyl stretching bands arising from carboxylic acid groups, which are often dimerized, are typically observed between 1700 and 1740 cm1 for molecules in solid or liquid phases.79 In dilute solutions or in the vapor phase, the carbonyl stretching band is shifted to 1760 cm1.79 Carboxy-terminated molecules in SAMs display a range of conformations. Monomer, acyclic dimer, and cyclic dimer forms of the carboxyl group (Figure 4) can coexist and appear as doublet or triplet peaks in IR spectra; specific peak assignments are made based on relative peak positions.72,78 While acyclic dimers are generally uncommon, the conformational constraints created by SAMs make single hydrogen bonds between adjacent carboxyl groups more likely. It has been proposed that as many as 50% of neighboring carboxyl groups in carboxyl-terminated 24780



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Figure 3. Reaction schemes for carboxy and amine terminal functional groups.

SAMs take on this configuration.80 The appearance of carboxyl IR absorption peaks depends on a number of factors in monolayer systems, including SAM preparation conditions and surface coverage. For example, MHDA inserted into a D12 matrix, as shown in Figure 5A(i), appears as a doublet peak at 1740 and 1720 cm1, indicative of carboxyl groups in monomer and acyclic dimer configurations, respectively (see Table 1).72 In contrast, HEG inserted into a D12 matrix, as shown in Figure 5B(i), appears as a weaker single peak at 1740 cm1. While this peak is attributable to the carboxyl carbonyl, a more detailed assignment of this peak cannot be made without further investigation. Additionally, a strong absorption peak at 1630 cm1 is observed in this spectrum, which does not appear in the corresponding MHDA spectrum (vide infra). These peak positions are not dependent on the use of D12 as a background SAM. We chose to use D12 here for consistency with later experiments where the use of D12 is necessary to avoid spectral overlap of the background SAM methylene peaks with those from the inserted molecules in the 28002950 cm1 region of the spectrum. Others and we have observed the 1630 cm1 peak in lowcoverage HEG self-assembled monolayers.35,56 However, a definitive peak identification has not been reported. Here, by sequentially rinsing with acidic (glacial acetic acid) and basic (KOH) solutions, we identify the 1630 cm1 peak as being associated with the deprotonated form of the HEG carboxy termini. In Figure 5B, we demonstrate the reversible acid/base characteristics of the carboxyl and carboxylate moieties, respectively, by monitoring the shifts between the 16101630 cm1 and 17401760 cm1 peaks, where we assign the 1760 cm1 peak in the HEG system to monomer carboxyl groups and the 1740 cm1 peak to acyclic dimers (see Table 1). Similar peak shifts occur with the MHDA system, as seen in Figure 5A (with the exception of the initial acid rinse, which has no effect on peak structure because MHDA deposits from ethanol in the protonated carboxyl form). Additional confirmation of a shift between carboxylate and carboxyl forms is provided by the appearance and disappearance, respectively, of peaks in the 14101470 cm1 range. We attribute these peaks to symmetric carboxylate modes (Table 1).72 We conclude that while isolated carboxyl groups behave similarly in OEG and n-alkanethiolate inserted systems, these two molecular species do not necessarily insert in identical forms. Another difference between the inserted MHDA and HEG systems is the relative prevalence of monomer and dimer forms of the carboxyl groups, which has previously only been examined for single-component monolayers.80,81 The relative intensities of the ∼1740 cm1 doublets show that a higher fraction of the inserted HEG molecules appear in the monomer form, indicated

Figure 4. Carboxyl monomer and dimer configurations. Carboxyl groups can adopt (A) monomer, (B) acyclic dimer, and (C) cyclic dimer configurations in SAMs.72,78,80

by the more pronounced 1760 cm1 peak of the doublet (Figure 5B(ii),(iv)). By contrast, a higher fraction of the MHDA molecules are in the acyclic dimer configuration, as indicated by the higher intensity 1720 cm1 peak (Figure 5A(ii),(iv)). We attribute this observation to the greater conformational degrees of freedom for the HEG molecules30,39,40 compared to MHDA. The additional freedom leads to a larger number of possible conformations than direct dimerization, making the dimer form less likely. We also consider how differences in insertion properties (such as insertion into proximate defect sites) might play a role in governing the quantity and distribution of inserted molecules. Insertion in small bundles and as single molecules is expected for the preparation conditions used here, as both domain boundaries and void spaces in the host matrix are expected for short SAM formation times.20,70,8284 It is possible that the longer HEG molecules interfere sterically with further insertion in proximate defect sites, resulting in larger spacings between HEG molecules and, consequently, fewer HEG dimers as compared to MHDA. It is also possible that the greater conformational degrees of freedom of the HEG molecules impact their insertion properties. However, the spectra of HEG and MHDA monolayers, which show the same relative monomer/dimer fractions as the inserted systems (see the Supporting Information), do not provide evidence to support this explanation. 3.2. Variation of Inserted Fractions of Oligo(ethylene glycol)alkanethiols Based on Terminal Group Functionality. While qualitative differences in the properties of inserted OEG and n-alkanethiol molecules are evident, quantitative differences are also apparent between different types of inserted OEG thiols. Figure 6 shows IRRAS peaks at 2870 and 2920 cm1, corresponding to methylene symmetric and asymmetric stretches, respectively, for both HEG and AEG molecules inserted into D12 SAMs. A deuterated thiol was selected for the host matrix to avoid overlap of methylene IR peaks from the SAM with those arising from the 24781



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Figure 5. Carboxyl/carboxylate cycling in carboxy-terminated n-alkanethiol vs oligo(ethylene glycol)alkanethiol inserted systems. (A) Mercaptohexadecanoic acid inserted from ethanol into a D12 SAM (i) shows characteristic carboxyl monomer (1740 cm1) and acyclic dimer (1720 cm1) peaks. (ii) Rinsing with an acidic solution does not alter the spectrum. (iii) Rinsing with basic solution results in a change to the carboxylate form and a corresponding shift in the spectral peaks. (iv) Rinsing with acidic solution recovers the original spectrum. (B) Carboxy-terminated OEG inserted from ethanol into a D12 SAM (i) shows spectral peaks indicative of a mixture of carboxyl and carboxylate forms with a bias toward the latter. (ii) Rinsing with an acidic solution shifts the molecules into the carboxyl form, with carboxyl monomer (1760 cm1) and acyclic dimer (1740 cm1) peaks apparent. (iii) Rinsing with basic solution shifts the inserted HEG molecules to the unprotonated carboxylate form. (iv) Rinsing again with an acidic solution demonstrates that the transition is reversible, with recovery of the spectrum indicative of the carboxyl form.

Table 1. Summary of Observed FT-IRRAS Carboxyl(ate) Absorption Peaks maximum [cm1] band

MHDA

HEG

CdO str, monomer

1740

1760

CdO str, acyclic dimer

1720

1740

COO asym str

1570

16101630

COO sym str

14201460

14101470

inserted molecules. The methylene peaks provide a measure of relative insertion of molecules with equal chain lengths. Based on the differences in peak areas, the inserted fraction of AEG molecules is approximately 60% of the fraction of inserted HEG under similar insertion conditions (Figure 6). Because HEG and AEG have identical lengths and structures (with the exception of the terminal groups), we conclude that differences in relative insertion arise from differences between the terminal groups. Differences in relative insertion based on tail group functionality are distinct from what is observed for n-alkanethiol insertion. The same spectral region for two n-alkanethiol molecules, MUDA and AAT, with equal chain lengths but different terminal groups inserted into D12 SAMs, is also shown in Figure 6. While we cannot quantitatively compare spectra from inserted OEGs and n-alkanethiols directly because of their differing chain lengths, we can conclude that, based on the similar peak areas of the methylene peaks for the two different n-alkanethiol molecules, relative inserted fractions are not affected by differences in terminal functional groups for this class of molecules.

Figure 6. Oligo(ethylene glycol)alkanethiol inserted fractions vary compared to consistent inserted fractions of n-alkanethiols. Substantial differences in insertion fraction exist between OEG molecules with identical chain lengths inserted under identical conditions based on differences in the terminal functional groups. The area under the methylene stretch peaks is greatest for BEG, indicating that this molecule inserts to the greatest degree, while HEG inserts at 70% and AEG at 50% of the BEG level. In contrast, n-alkanethiols with different terminal functional groups but similar chain lengths (MUDA and AAT) do not show significant variations in inserted fractions. All molecules were inserted into identically prepared D12 monolayers.

Because OEG insertion into n-alkanethiolate SAMs is affected by the type of terminal functional group, we considered chemical functionalization prior to insertion as a factor that could also influence insertion properties. To test this, insertion of a biotinylated OEG (BEG) thiol was compared to HEG and AEG insertion 24782



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Figure 7. Comparative reactivities of carboxy-terminated thiols in inserted vs single-component monolayers show reduced reactivity for the n-alkanethiol single-component monolayer. (A) Molecules of HEG or MHDA inserted into D12 monolayers show no appreciable differences in chemical reactivity. (B) Monolayers of HEG show qualitatively similar chemical behavior with the inserted systems. In contrast, single-component MHDA monolayers show substantially reduced reactivity in the second step of the functionalization scheme.

(shown schematically in Figure 1B). For BEG, amine-terminated OEG molecules are functionalized with biotin prior to insertion. We observed greater BEG insertion compared to the other OEGs (30% higher than HEG and 90% higher than AEG based on comparisons of the combined 2870 and 2920 cm1 peak areas, as shown in Figure 6). This is striking, given that amine-terminated OEG without biotin functionalization, that is, AEG, displays the lowest level of insertion among the three OEGs investigated. We note that differences in the inserted fractions for all of these molecules are found under identical insertion conditions. None of these systems reach equilibrium,3,20,26,83 and it is therefore possible to use kinetic control by adjusting the insertion conditions (e.g., time, concentration) to select the amount of insertion (see the Supporting Information, Figure S3).49,70,82 Further study will be required to elucidate the underlying factors that determine the rates and extent of functional thiol insertion. Nonetheless, several observations can be made based on the current findings. First, because different OEG terminal functional groups significantly impact the extent of insertion under otherwise identical conditions, experiments investigating OEG insertion with respect to terminal groups cannot be directly compared without first accounting for differences in the inserted fractions, independent of other effects attributable to differences in chain length or incubation time. This is in contrast to n-alkanethiol systems, where expectations of equal insertion quantities for equal chain lengths appear valid, at least based on the data for MUDA and AAT. Second, the parameters for a targeted surface composition need to be based on the participating molecules and cannot be extrapolated from conditions used for other OEG thiol systems. This is particularly important for biological applications, where specific and varied terminal group functionality is required.7,32,35,36,56,85,86 Third, terminal functionality needs to be taken into account when determining protocols for the creation of biofunctionalized surfaces, since choices of molecular functionality and chemistries carried out prior to insertion are anticipated to alter final surface compositions significantly. As a result, comparison of in situ and preinsertion chemistries should be made to

determine which represents the optimal strategy for a given application. 3.3. Chemical Reactivity Comparisons of Carboxy-Terminated n-Alkanethiol vs Oligo(ethylene glycol)alkanethiol. Amide coupling at terminal SAM carboxyl groups via reactive NHS-ester intermediates was selected for study based on widespread use. We used HEG and MHDA as prototypical systems because previous work has shown that both molecules are amenable to chemical reaction when assembled on surfaces.28,35,56,78,8789 As above, these molecules were inserted into preformed D12 matrices, producing dilute coverage of carboxyl groups on either n-alkanethiol or OEG tethers. Functionalization was carried out according to Scheme 1 (Figure 3). Spectra of these two different inserted monolayers prior to functionalization with 5-HT are qualitatively distinct in the 15001850 cm1 wavenumber region (Figure 7A(i)). Despite differences in the spectral peaks for the HEG and MHDA samples, the peaks diagnostic for the NHS reaction are qualitatively similar for both types of samples after the formation of SAM-tethered NHS-esters, as shown in Figure 7A(ii). Peaks attributed to NHS typically appear at 1745 cm1 (the asymmetric imide carbonyl stretch), 1790 cm1 (the symmetric imide carbonyl stretch), and 1820 cm1 (the NHS-ester carbonyl stretch). The 1820 cm1 peak is diagnostic for the covalent attachment of NHS and is comparable for both insertion systems, indicating similar reactivity (assuming similar insertion quantity). This is noteworthy because it indicates that conformational differences between inserted HEG and MHDA molecules do not appear to affect the availability of these molecules for further reaction. Because the NHS-ester is particularly reactive with primary amines, we incubated HEG- or MHDA-inserted SAMs in solutions of 5-HT (serotonin), a monoamine neurotransmitter used previously to create biofunctionalized capture surfaces.35,36 Figure 7A(iii) shows the spectra of the resulting surfaces, with the peak at 1663 cm1 arising from the carbonyl stretch of the resulting amide bond with 5-HT (amide I band, the diagnostic peak for the reaction). Smaller peaks between 1500 and 1550 cm1 (amide II band) are associated with secondary 24783



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Figure 8. Comparative reactivities of amine-terminated thiols in inserted vs single-component monolayers show reduced reactivity for the n-alkanethiol single-component monolayer. (A) Molecules of AEG and AAT inserted into C8 monolayers show similar levels of reactivity, mirroring the behavior of inserted HEG and MHDA in Figure 7A. (B) Monolayers of AEG show significantly higher levels of functionalization than monolayers of AAT.

amines but are not always observed because of IR spectral sensitivity to orientation. As with the NHS spectra, the diagnostic peaks are virtually identical for both tethers, indicating no appreciable differences in chemical reactivity for these carboxyl groups when isolated in and protruding from n-alkanethiolate SAM matrices. In contrast to similar reactivities for different inserted carboxyl molecules, single-component HEG and MHDA monolayers behave differently from one another. As shown in Figure 7B(i), NHS-ester formation is alike for both monolayers, as with inserted molecules. However, on subsequent 5-HT functionalization (Figure 7B(ii)), differences in monolayer reactivity are apparent. The HEG SAM exhibits a spectrum that is qualitatively similar to that seen for the inserted molecule systems shown in Figure 7A(iii). Quantitative differences are attributed to the significantly higher quantity of the relevant functional groups present in monolayers of HEG as compared to HEG inserted into D12 SAMs. The MHDA SAM spectrum, in contrast, is qualitatively different from that of inserted MHDA surfaces, HEG monolayers, and monolayers with inserted HEG. The diagnostic 1663 cm1 peak for 5-HT attachment is either absent or shifted to lower energy, indicating little to no functionalization of this monolayer. Further, while reduced in intensity, peaks indicating the continued presence of NHS-functionalized molecules are present in the MHDA monolayer after reaction with 5-HT. This is surprising because NHS-MHDA ester hydrolysis is favorable; in the absence of amide bond formation, the MHDA carboxyl terminus is expected to be recovered.56 Some recovery of carboxyl groups is likely occurring, although it is difficult to quantify because of interference of the 1745 cm1 peak from the residual NHS imide with the 1740 cm1 carboxyl carbonyl absorption for MHDA. Regardless, the persistence of NHS diagnostic peaks indicates that although the NHS reaction yield is highly similar in the two monolayers, further reaction is suppressed for the MHDA monolayer.

We hypothesize that reduction in reactivity for MHDA monolayers arises from steric hindrance; the ester bond is shielded by the closely packed, well-ordered SAM. In contrast, the oligo(ethylene glycol) moieties of HEG SAMs result in looser, less crystalline molecular packing at the terminal ends of the monolayers relative to n-alkanethiolate monolayers.30,39,40 This conformational freedom could account for the observed differences in reactivity between the two types of monolayers. Molecules of MHDA inserted into D12 SAMs lack this crystalline packing, consistent with the higher reactivity of MHDA under these conditions. The results also indicate that dilute tether coverage is necessary in some cases for in situ chemistries to occur to create biofunctionalized surfaces. 3.4. Chemical Reactivity Comparison of Amine-Terminated n-Alkanethiol and Oligo(ethylene glycol)alkanethiol. We also compared the reactivities of amine-terminated n-alkanethiol (AAT) versus amine-terminated oligo(ethylene glycol)alkanethiol (AEG). To minimize differences in comparisons between the carboxy- and amine-terminated systems, amide bond formation was again used as a representative chemical reaction. Because the NHS intermediate ester is formed with a carboxyl group, and monolayers composed of or containing AAT or AEG present amine groups, a carboxyl-presenting reagent was required. For this reason, 5-hydroxytryptophan (5-HTP) was used in place of 5-HT. With the exception of the additional carboxyl group, 5-HTP is identical to 5-HT. The additional carboxyl group is used in the formation of the NHS intermediate ester in solution and, subsequently, in the formation of an amide bond with the amine terminal groups of the thiol molecules (Scheme 2, Figure 3). The use of 5-HTP also enables the synthesis of a biologically relevant test system, as 5-HTP-functionalized monolayer systems have been used previously for biological capture applications.36 To avoid undesirable byproducts, a FMOC protecting group was attached to the primary amine of 5-HTP, resulting in molecules somewhat larger than 5-HT. The NHS ester was formed in solution, thus eliminating a step from 24784


The Journal of Physical Chemistry C the surface functionalization scheme used for carboxy-terminated tethers. As with the carboxyl-based systems, inserted and single-component monolayers were used to ascertain the chemical reactivity of amine-terminated molecules. For the inserted systems, we used C8 for the initial SAM matrix to account for the shorter AAT molecule (11 carbon chain length for AAT vs 16 carbon chain length for MHDA). By using C8 for the background SAM, the amine groups of inserted AAT molecules protrude from the surrounding matrix, better emulating the corresponding system of MHDA inserted into a D12 SAM. Results for the inserted systems are shown in Figure 8A. As before, the 1663 cm1 peak is diagnostic for amide bond formation on the surfaces. Peaks between 1520 and 1550 cm1 are indicative of unreacted primary amine groups. The peak at 1720 cm1 results from carbonyls associated with the FMOC protecting groups. As with the carboxy-terminated insertion systems, reactivity, as measured by the amide bond peak intensities, appears similar for amine-terminated OEG and n-alkanethiol molecules. This is consistent with our hypothesis of similarity in chemical availability of the molecules when protruding at low density from surrounding SAM matrices.35,36 As with the HEG/MHDA system, single-component monolayers of AEG and AAT were fabricated and functionalized using the same reactions as the inserted amine molecules. As shown in Figure 8B, a lower level of functionalization is observed for amine-terminated n-alkanethiol monolayers compared to amineterminated OEG monolayers. This is consistent with the behavior observed for the single-component carboxy-terminated monolayers. In the case of the amines, however, we observe the difference in functionalization yield at the first step of the chemistry (Scheme 2, Figure 3), as opposed to differences becoming apparent at the second step of the two-step chemistry used for carboxyl functionalization (Scheme 1, Figure 3). We hypothesize that it is difficult for the larger NHS-FMOC-5-HTP complex to achieve a favorable orientation for reaction with the tightly packed amine terminal groups in AAT monolayers compared to the smaller NHS molecule reacting with the tightly packed carboxyl groups in MHDA monolayers. Because there is no steric shielding of NHS ester bond formation in solution, as opposed to that which arises from crystalline packing on surfaces (as with the two-step chemistry of the MHDA monolayer), limited amide bond formation appears to occur on AAT monolayers. This is in contrast to the MHDA system, where amide bond formation is not observed. This behavior would account for the inverse relationship between the 1663 cm1 amide carbonyl and 1550 cm1 primary amine peak intensities evident in Figure 8B. This behavior might be of considerable importance for functionalization schemes involving large molecules or proteins, especially those carried out on densely packed functional surfaces. Specifically, reaction yields on solid substrates might be improved by modifying surface compositions to improve functional group density and availability. As an aside, we note that a primary amine peak at 1550 cm1 is absent in the initial scan of the amine-terminated monolayers. This peak is apparent only after functionalization. However, the nearly identical character of scans taken before and after removal of the FMOC protecting group on the 5-HTP molecule, converting a secondary amine to a primary amine, indicates that the appearance of this peak cannot be attributed only to the presence of the amine group of the 5-HTP molecule. Therefore, some aspect of the functionalization procedure, most likely changes in orientation, is responsible for the IR spectral change due to the

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existing primary amine tail groups. While interesting, the unpredictable behavior of primary amine groups in IRRAS experiments has been observed previously90 and is not unique to these monolayer systems.

4. CONCLUSIONS AND PROSPECTS Predicting the availability and reactivity of surface functional groups for SAMs by extrapolating from similar but distinct systems is not straightforward. Different degrees of protonation were observed for carboxy groups on the termini of oligo(ethylene glycol) versus alkyl chains after insertion from ethanolic solutions under otherwise identical conditions. Protonation variability arises from differences in the local chemical environment, while differences in conformational degrees of freedom lead to differences in carboxyl monomer/dimer ratios. Insertion kinetics of OEGs were found to be dependent on terminal functionality, which was not observed with analogous n-alkanethiol molecules. As a consequence, we conclude that achieving equivalent inserted fractions of differently functionalized OEGs will require different insertion conditions. One cannot use the behavior of either equivalent n-alkanethiols or other OEG molecules with different terminal functionalities as predictors of the extent of insertion of specific OEGs. Furthermore, we demonstrate that chemical variation of the OEG terminal group prior to insertion alters the degree of insertion of OEG molecules. Significant differences were apparent between the reactivities of OEG and n-alkanethiolate monolayers with similar exposed terminal functional groups. We have shown that the availability of terminal functional groups for reaction on single-component OEG monolayers is distinctly greater than that of similarly functionalized single-component n-alkanethiolate monolayers for both carboxy- and amine-terminated SAMs. The greater degree of crystalline packing of n-alkanethiol functional groups likely leads to decreases in reactivity compared to the less tightly packed ethylene glycol moieties. Similar reactivities of inserted n-alkanethiols and OEGs indicate that the chemical behavior of the functional groups is not intrinsically different between the two systems. These findings highlight the importance of molecular environment for determining reactivity of SAMs. By establishing significant variations in the insertion and chemistries of OEG molecules compared to n-alkanethiol molecules, we have uncovered factors important for the creation of biofunctionalized surface systems and multicomponent self-assembled systems. In the future, gaining deeper understanding of the mechanisms governing these differences will enable more effective uses of these molecules and monolayers in a wide variety of applications, including biofunctional capture and sensing surfaces. ’ ASSOCIATED CONTENT

bS

Supporting Information. Detailed information for carboxyl monolayer hydrogen-bonding fractions, carboxyl reorganization over time, and control of relative OEG inserted fractions. This material is available free of charge via the Internet at http://pubs.acs.org.

’ AUTHOR INFORMATION Corresponding Author

*(P.S.W.) E-mail: [email protected]; phone: +1 (310) 267-4838; fax: +1 (310) 267-4918. (A.M.A.) E-mail: [email protected]. edu; phone: +1 (310) 794-9421; fax: +1 (310) 983-1133. 24785



’ ACKNOWLEDGMENT The authors gratefully acknowledge funding from the National Science Foundation (no. 1013042) and the Kavli Foundation. This work was supported by the Pennsylvania State University Materials Research Institute Nanofabrication Lab and the National Science Foundation Cooperative Agreement no. ECS-0335765. ’ REFERENCES (1) Porter, M. D.; Bright, T. B.; Allara, D. L.; Chidsey, C. E. D. J. Am. Chem. Soc. 1987, 109, 3559–3568. (2) Bertilsson, L.; Liedberg, B. Langmuir 1993, 9, 141–149. (3) Stranick, S. J.; Parikh, A. N.; Tao, Y. T.; Allara, D. L.; Weiss, P. S. J. Phys. Chem. 1994, 98, 7636–7646. (4) Atre, S. V.; Liedberg, B.; Allara, D. L. Langmuir 1995, 11, 3882–3893. (5) Jung, L. S.; Nelson, K. E.; Stayton, P. S.; Campbell, C. T. Langmuir 2000, 16, 9421–9432. (6) Lewis, P. A.; Smith, R. K.; Kelly, K. F.; Bumm, L. A.; Reed, S. M.; Clegg, R. S.; Gunderson, J. D.; Hutchison, J. E.; Weiss, P. S. J. Phys. Chem. B 2001, 105, 10630–10636. (7) Nelson, K. E.; Gamble, L.; Jung, L. S.; Boeckl, M. S.; Naeemi, E.; Golledge, S. L.; Sasaki, T.; Castner, D. G.; Campbell, C. T.; Stayton, P. S. Langmuir 2001, 17, 2807–2816. (8) Mark, S. S.; Sandhyarani, N.; Zhu, C.; Campagnolo, C.; Batt, C. A. Langmuir 2004, 20, 6808–6817. (9) Mullen, T. J.; Dameron, A. A.; Saavedra, H. M.; Williams, M. E.; Weiss, P. S. J. Phys. Chem. C 2007, 111, 6740–6746. (10) Mullen, T. J.; Dameron, A.; Andrews, A. M.; Weiss, P. S. Aldrichimica Acta 2007, 40, 21–31. (11) Larsson, A.; Liedberg, B. Langmuir 2007, 23, 11319–11325. (12) Weiss, P. S. Acc. Chem. Res. 2008, 41, 1772–1781. (13) Ladd, J.; Taylor, A. D.; Piliarik, M.; Homola, J.; Jiang, S. Anal. Chem. 2008, 80, 4231–4236. (14) Frasconi, M.; Mazzei, F.; Ferri, T. Anal. Bioanal. Chem. 2010, 398, 1545–1564. (15) Nishino, T.; Umezawa, Y. Anal. Sci. 2010, 26, 1023–1032. (16) Ekblad, T.; Liedberg, B. Curr. Opin. Colloid Interface Sci. 2010, 15, 499–509. (17) Haag, R.; Rampi, M. A.; Holmlin, R. E.; Whitesides, G. M. J. Am. Chem. Soc. 1999, 121, 7895–7906. (18) MacBeath, G.; Koehler, A. N.; Schreiber, S. L. J. Am. Chem. Soc. 1999, 121, 7967–7968. (19) Schaferling, M.; Riepl, M.; Pavlickova, P.; Paul, H.; Kambhampati, D.; Liedberg, B. Microchimica Acta 2003, 142, 193–203. (20) Smith, R. K.; Lewis, P. A.; Weiss, P. S. Prog. Surf. Sci. 2004, 75, 1–68. (21) Yan, L.; Huck, W. T. S.; Whitesides, G. M. J. Macromol. Sci., Polym. Rev. 2004, C44, 175–206. (22) Srinivasan, C.; Mullen, T. J.; Hohman, J. N.; Anderson, M. E.; Dameron, A. A.; Andrews, A. M.; Dickey, E. C.; Horn, M. W.; Weiss, P. S. ACS Nano 2007, 1, 191–201. (23) Zorn, S.; Martin, N.; Gerlach, A.; Schreiber, F. Phys. Chem. Chem. Phys. 2010, 12, 8986–8991. (24) Zorn, S.; Skoda, M. W. A.; Gerlach, A.; Jacobs, R. M. J.; Schreiber, F. Langmuir 2011, 27, 2237–2243. (25) Kim, M.; Hohman, J. N.; Cao, Y.; Houk, K. N.; Ma, H.; Jen, A. K.-Y.; Weiss, P. S. Science 2011, 331, 1312–1315. (26) Saavedra, H. M.; Mullen, T. J.; Zhang, P.; Dewey, D. C.; Claridge, S. A.; Weiss, P. S. Rep. Prog. Phys. 2010, 73, 36501–36540. (27) Palegrosdemange, C.; Simon, E. S.; Prime, K. L.; Whitesides, G. M. J. Am. Chem. Soc. 1991, 113, 12–20. (28) Chapman, R. G.; Ostuni, E.; Yan, L.; Whitesides, G. M. Langmuir 2000, 16, 6927–6936. (29) Schwendel, D.; Dahint, R.; Herrwerth, S.; Schloerholz, M.; Eck, W.; Grunze, M. Langmuir 2001, 17, 5717–5720.

ARTICLE

(30) Herrwerth, S.; Eck, W.; Reinhardt, S.; Grunze, M. J. Am. Chem. Soc. 2003, 125, 9359–9366. (31) Li, L.; Chen, S.; Zheng, J.; Ratner, B. D.; Jiang, S. J. Phys. Chem. B 2005, 109, 2934–2941. (32) Holzl, M.; Tinazli, A.; Leitner, C.; Hahn, C. D.; Lackner, B.; Tampe, R.; Gruber, H. J. Langmuir 2007, 23, 5571–5577. (33) Schilp, S.; Rosenhahn, A.; Pettitt, M. E.; Bowen, J.; Callow, M. E.; Callow, J. A.; Grunze, M. Langmuir 2009, 25, 10077–10082. (34) Zareie, H. M.; Boyer, C.; Bulmus, V.; Nateghi, E.; Davis, T. P. ACS Nano 2008, 2, 757–765. (35) Shuster, M. J.; Vaish, A.; Szapacs, M. E.; Anderson, M. E.; Weiss, P. S.; Andrews, A. M. Adv. Mater. 2008, 20, 164–167. (36) Vaish, A.; Shuster, M. J.; Cheunkar, S.; Singh, Y. S.; Weiss, P. S.; Andrews, A. M. ACS Chem. Neurosci. 2010, 1, 495–504. (37) Zhao, C.; Burchardt, M.; Brinkhoff, T.; Beardsley, C.; Simon, M.; Wittstock, G. Langmuir 2010, 26, 8641–8647. (38) Zhang, X.; Yadavalli, V. K. Biosens. Bioelectron. 2011, 26, 3142–3147. (39) Harder, P.; Grunze, M.; Dahint, R.; Whitesides, G. M.; Laibinis, P. E. J. Phys. Chem. B 1998, 102, 426–436. (40) Zwahlen, M.; Herrwerth, S.; Eck, W.; Grunze, M.; Hahner, G. Langmuir 2003, 19, 9305–9310. (41) Malysheva, L.; Onipko, A.; Liedberg, B. J. Phys. Chem. A 2008, 112, 728–736. (42) Valiokas, R.; Malysheva, L.; Onipko, A.; Lee, H. H.; Ruzele, Z.; Svedhem, S.; Svensson, S. C. T.; Gelius, U.; Liedberg, B. J. Electron Spectrosc. Relat. Phenom. 2009, 172, 9–20. (43) Brand, I.; Nullmeier, M.; Kluner, T.; Jogireddy, R.; Christoffers, J.; Wittstock, G. Langmuir 2010, 26, 362–370. (44) Bumm, L. A.; Arnold, J. J.; Cygan, M. T.; Dunbar, T. D.; Burgin, T. P.; Jones, L.; Allara, D. L.; Tour, J. M.; Weiss, P. S. Science 1996, 271, 1705–1707. (45) Cygan, M. T.; Dunbar, T. D.; Arnold, J. J.; Bumm, L. A.; Shedlock, N. F.; Burgin, T. P.; Jones, L.; Allara, D. L.; Tour, J. M.; Weiss, P. S. J. Am. Chem. Soc. 1998, 120, 2721–2732. (46) Donhauser, Z. J.; Mantooth, B. A.; Kelly, K. F.; Bumm, L. A.; Monnell, J. D.; Stapleton, J. J.; Price, D. W.; Rawlett, A. M.; Allara, D. L.; Tour, J. M.; Weiss, P. S. Science 2001, 292, 2303–2307. (47) Pollino, J. M.; Stubbs, L. P.; Weck, M. J. Am. Chem. Soc. 2004, 126, 563–567. (48) Moore, A. M.; Dameron, A. A.; Mantooth, B. A.; Smith, R. K.; Fuchs, D. J.; Ciszek, J. W.; Maya, F.; Yao, Y.; Tour, J. M.; Weiss, P. S. J. Am. Chem. Soc. 2006, 128, 1959–1967. (49) Mullen, T. J.; Srinivasan, C.; Hohman, J. N.; Gillmor, S. D.; Shuster, M. J.; Horn, M. W.; Andrews, A. M.; Weiss, P. S. Appl. Phys. Lett. 2007, 90, 063114. (50) Lewis, P. A.; Inman, C. E.; Yao, Y. X.; Tour, J. M.; Hutchison, J. E.; Weiss, P. S. J. Am. Chem. Soc. 2004, 126, 12214–12215. (51) Moore, A. M.; Mantooth, B. A.; Donhauser, Z. J.; Maya, F.; Price, D. W.; Yao, Y.; Tour, J. M.; Weiss, P. S. Nano Lett. 2005, 5, 2292–2297. (52) Vaish, A.; Shuster, M. J.; Cheunkar, S.; Weiss, P. S.; Andrews, A. M. Small 2011, 7, 1471–1479. (53) Mullen, T. J.; Srinivasan, C.; Shuster, M. J.; Horn, M. W.; Andrews, A. M.; Weiss, P. S. J. Nanopart. Res. 2008, 10, 1231–1240. (54) Shuster, M. J.; Vaish, A.; Cao, H. H.; Guttentag, A. I.; McManigle, J. E.; Gibb, A. L.; Martinez, M. M.; Nezarati, R. M.; Hinds, J. M.; Liao, W. S.; Weiss, P. S.; Andrews, A. M. Chem. Commun. 2011, 47, 10641– 10643. (55) Haeussling, L.; Ringsdorf, H.; Schmitt, F. J.; Knoll, W. Langmuir 1991, 7, 1837–1840. (56) Lahiri, J.; Isaacs, L.; Tien, J.; Whitesides, G. M. Anal. Chem. 1999, 71, 777–790. (57) Lahiri, J.; Ostuni, E.; Whitesides, G. M. Langmuir 1999, 15, 2055–2060. (58) Segui, S. R.; Pla, M.; Minic, J.; Pajot-Augy, E.; Salesse, R.; Hou, Y.; Jaffrezic-Renault, N.; Mills, C. A.; Samitier, J.; Errachid, A. Anal. Lett. 2006, 39, 1735–1745. 24786



ARTICLE

(59) Smith, R. K.; Reed, S. M.; Lewis, P. A.; Monnell, J. D.; Clegg, R. S.; Kelly, K. F.; Bumm, L. A.; Hutchison, J. E.; Weiss, P. S. J. Phys. Chem. B 2001, 105, 1119–1122. (60) Kim, M.; Hohman, J. N.; Morin, E. I.; Daniel, T. A.; Weiss, P. S. J. Phys. Chem. A 2009, 113, 3895–3903. (61) Hohman, J. N.; Zhang, P. P.; Morin, E. I.; Han, P.; Kim, M.; Kurland, A. R.; McClanahan, P. D.; Balema, V. P.; Weiss, P. S. ACS Nano 2009, 3, 527–536. (62) Saavedra, H. M.; Thompson, C. M.; Hohman, J. N.; Crespi, V. H.; Weiss, P. S. J. Am. Chem. Soc. 2009, 131, 2252–2259. (63) Kim, M.; Hohman, J. N.; Serino, A. C.; Weiss, P. S. J. Phys. Chem. C 2010, 114, 19744–19751. (64) Hohman, J. N.; Claridge, S. A.; Kim, M.; Weiss, P. S. Mater. Sci. Eng., R 2010, 70, 188–208. (65) Sprik, M.; Delamarche, E.; Michel, B.; Rothlisberger, U.; Klein, M. L.; Wolf, H.; Ringsdorf, H. Langmuir 1994, 10, 4116–4130. (66) Gorman, C. B.; He, Y.; Carroll, R. L. Langmuir 2001, 17, 5324–5328. (67) Li, L. Y.; Chen, S. F.; Jiang, S. Y. Langmuir 2003, 19, 2974–2982. (68) Yadavalli, V. K.; Forbes, J. G.; Wang, K. Langmuir 2006, 22, 6969–6976. (69) Bumm, L. A.; Arnold, J. J.; Dunbar, T. D.; Allara, D. L.; Weiss, P. S. J. Phys. Chem. B 1999, 103, 8122–8127. (70) Dunbar, T. D.; Cygan, M. T.; Bumm, L. A.; McCarty, G. S.; Burgin, T. P.; Reinerth, W. A.; Jones, L.; Jackiw, J. J.; Tour, J. M.; Weiss, P. S.; Allara, D. L. J. Phys. Chem. B 2000, 104, 4880–4893. (71) Nyquist, R. M.; Eberhardt, A. S.; Silks, L. A.; Li, Z.; Yang, X.; Swanson, B. I. Langmuir 2000, 16, 1793–1800. (72) Arnold, R.; Azzam, W.; Terfort, A.; Woll, C. Langmuir 2002, 18, 3980–3992. (73) Weck, M.; Jackiw, J. J.; Rossi, R. R.; Weiss, P. S.; Grubbs, R. H. J. Am. Chem. Soc. 1999, 121, 4088–4089. (74) Mrksich, M. ACS Nano 2008, 2, 7–18. (75) Wang, H.; Chen, S. F.; Li, L. Y.; Jiang, S. Y. Langmuir 2005, 21, 2633–2636. (76) Himmelhaus, M.; Eisert, F.; Buck, M.; Grunze, M. J. Phys. Chem. B 2000, 104, 576–584. (77) Su, X. L.; Li, Y. B. Biosens. Bioelectron. 2004, 19, 563–574. (78) Frey, B. L.; Corn, R. M. Anal. Chem. 1996, 68, 3187–3193. (79) Socrates, G. Infrared and Raman Characteristic Group Frequencies, 3rd ed.; John Wiley and Sons, LTD: New York, 2001. (80) Nuzzo, R. G.; Dubois, L. H.; Allara, D. L. J. Am. Chem. Soc. 1990, 112, 558–569. (81) Smith, E. L.; Alves, C. A.; Anderegg, J. W.; Porter, M. D.; Siperko, L. M. Langmuir 1992, 8, 2707–2714. (82) Fuchs, D. J.; Weiss, P. S. Nanotechnology 2007, 18, 1–7. (83) Bumm, L. A.; Arnold, J. J.; Charles, L. F.; Dunbar, T. D.; Allara, D. L.; Weiss, P. S. J. Am. Chem. Soc. 1999, 121, 8017–8021. (84) Donhauser, Z. J.; Price, D. W.; Tour, J. M.; Weiss, P. S. J. Am. Chem. Soc. 2003, 125, 11462–11463. (85) Perez-Luna, V. H.; O’Brien, M. J.; Opperman, K. A.; Hampton, P. D.; Lopez, G. P.; Klumb, L. A.; Stayton, P. S. J. Am. Chem. Soc. 1999, 121, 6469–6478. (86) Otsuka, H.; Nagasaki, Y.; Kataoka, K. Curr. Opin. Colloid Interface Sci. 2001, 6, 3–10. (87) Lahiri, J.; Isaacs, L.; Grzybowski, B.; Carbeck, J. D.; Whitesides, G. M. Langmuir 1999, 15, 7186–7198. (88) Mrksich, M.; Dike, L. E.; Tien, J.; Ingber, D. E.; Whitesides, G. M. Exp. Cell Res. 1997, 235, 305–313. (89) Park, T.; Choo, J.; Lee, M.; Kim, Y. S.; Lee, E. K.; Lee, H. S. Anal. Sci. 2004, 20, 1255–1258. (90) Hooper, A. E.; Werho, D.; Hopson, T.; Palmer, O. Surf. Interface Anal. 2001, 31, 809–814.

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Comparison of Oligo (ethylene glycol) alkanethiols versus n

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