Skip to main content
PMC home page
ACS Omega logoLink to ACS Omega
. 2018 Feb 8;3(2):1635–1644. doi: 10.1021/acsomega.7b01641

Design and Evaluation of Short Self-Assembling Depsipeptides as Bioactive and Biodegradable Hydrogels

Kevin M Eckes 1, Kiheon Baek 1, Laura J Suggs 1,*
PMCID: PMC6044717  PMID: 30023812

Abstract

graphic file with name ao-2017-01641f_0008.jpg

Described herein is the design of a cell-adherent and degradable hydrogel. Our goal was to create a self-assembling, backbone ester-containing analogue of the cell adhesion motif, arginine–glycine–aspartic acid (RGD). Two depsipeptides containing Fmoc (N-(fluorenyl)-9-methoxycarbonyl), Fmoc-FR-Glc-D, and Fmoc-F-Glc-RGD (where “Glc” is glycolic acid) were designed based on the results of integrin-binding affinity and cell interaction analyses. Two candidate molecules were synthesized, and their gelation characteristics, degradation profiles, and ability to promote cell attachment were analyzed. We found that ester substitution within the RGD sequence significantly decreases the integrin-binding affinity and subsequent cell attachment, but when the ester moiety flanks the bioactive sequence, the molecule can maintain its integrin-binding function while permitting nonenzymatic hydrolytic degradation. A self-assembled Fmoc-F-Glc-RGD hydrogel showed steady, linear degradation over 60 days, and when mixed with Fmoc-diphenylalanine (Fmoc-FF) for improved mechanical stiffness, the depsipeptide gel exhibited improved cell attachment and viability. Though the currently designed depsipeptide has several inherent limitations, our results indicate the potential of depsipeptides as the basis for biologically functional and degradable self-assembling hydrogel materials.

Introduction

Peptide-based self-assembling materials show great promise as injectable scaffold materials for tissue regeneration applications, but little work has been done to understand their long-term fate in vivo. Such materials may not be easily degraded by the body. Indeed, the level of matrix-degrading protease expression varies with the tissue and cell type; thus, frequently remodeled tissues (e.g., skin) likely have much greater protease activity than tissues with low turnover (e.g., nerve).1,2

Recent work by our group and others suggests that it may be possible to engineer ester-mediated degradability into amphiphilic peptides without disrupting their self-assembly capability.3−5 It remains unclear, however, whether backbone ester-modified bioactive peptides retain other biologic activities, such as cell binding. To investigate this question and make a biologically interactive and degradable hydrogel, we developed several self-assembling, ester-containing depsipeptides based on a sequence with relevance to biomedical applications: arginine–glycine–aspartic acid (RGD). This sequence is found in several extracellular matrix (ECM) proteins, is known to bind to a number of cell surface integrin variants to mediate cell adhesion to the ECM,6 and acts as a handle for cellular interrogation of substrate mechanical properties that are known to affect cell behavior, migration, and differentiation.7−10

RGD and RGD-containing peptides have been incorporated into many synthetic hydrogel systems to encourage cell interaction with otherwise inert materials. For this reason and the fact that the RGD sequence is one of the shortest known biomolecular recognition sequences, we chose to develop RGD-mimicking depsipeptides as the basis for self-assembling degradable materials with the potential for peptide-like bioactivity. A secondary goal was to use the system to study the role that backbone hydrogen bonding plays in the binding of RGD peptides to an integrin protein. While ester-modified peptides have been used previously to elucidate the role of backbone hydrogen bonding in protein folding,11−14 β-amyloid formation,15,16 and other peptide–protein interactions,17,18 to our knowledge there have been no studies investigating the importance of hydrogen bonding in mediating biomechanically important peptide–protein interactions such as integrin–RGD binding.

Results and Discussion

Binding Affinity Analysis of Peptides and Depsipeptides

Our initial study was designed to test whether or not an ester substitution within the RGD sequence allowed the ligand to retain its affinity for cellular integrins. Fluorescence polarization (FP) spectroscopy was used to assess the binding affinity of the depsipeptide R-Glc-D to integrin proteins in comparison with positive (RGD) and negative arginine–glycine–glutamic acid (RGE) tripeptide controls. Molecular structures of R-Glc-D, RGD, and RGE are shown in Figure 1 to highlight the similarities between the three molecules.

Figure 1.

Figure 1

Open in a new tab

Molecular structures of RGD derivatives.

Human recombinant integrin α5β1 was found to induce measurable FP of 10 nM fluorescein Isothiocyanate (FITC)-conjugated GRGDSP peptide in Tris-buffered saline at concentrations above 200 nM (Figure S1), and the dissociation constant (Kd) of this interaction was calculated as 207 nM from a curve fit using a one-site binding assumption (GraphPad Prism 6). Competitive binding experiments, in which aliquots of a solution of 10 nM FITC-GRGDSP and 300 nM integrin were mixed with varying concentrations (10–9 to 10–2 M) of RGD, RGE, or R-Glc-D, revealed that R-Glc-D has some capacity to compete with FITC-GRGDSP/integrin binding, as evidenced by the decreasing FP with increasing R-Glc-D concentration (Figure 2).

Figure 2.

Figure 2

Open in a new tab

Half inhibition (i.e., IC50) of the fluorescence of FITC-GRGDSP occurs at a ∼40-fold lower concentration for RGD than R-Glc-D, indicating greater affinity of RGD for integrin than R-Glc-D (A). Comparison of FP inhibition by R-Glc-D and RGE peptides (negative control). The average IC50 value for R-Glc-D is 1115 ± 211 μM (mean ± standard deviation). RGE was tested once as a negative control, and the calculated IC50 value was 2840 μM (B).

RGD inhibited FP with an IC50 of 26.8 μM (Table 1), a level about 42 times greater than that of R-Glc-D, whereas RGE displayed slightly less FP inhibition relative to R-Glc-D (Figure 2). Inhibition constant (Ki) values were calculated assuming Kd = 207 nM, using the FP-specific method developed by Nikolovska-Coleska, et al.19 Glc-RGD was not included in the competitive binding experiments as it has four residues rather than three, and rigorous characterization of how ester bond substitutions flanking a bioactive peptide sequence affect the binding activity is outside the scope of the current study.

Table 1. IC50 and Ki of RGD Derivatives.

  IC50 (μM) Ki (μM)
R-G-E 2840 1157
R-Glc-D 1115 ± 211 453.9 ± 85.8
R-G-D 26.8 ± 7.5 10.8 ± 3.0
Open in a new tab

Adhesion and Morphology of Cells on Surface-Modified Glass

Based on the binding affinity analysis above, we proceeded to evaluate cell adhesion and spreading behavior on glass surfaces covalently functionalized with peptide or depsipeptide moieties. Our evaluation included RGD and RGE as well as R-Glc-D, in which the ester bond is included within the bioactive sequence, and Glc-RGD, in which the ester bond is adjacent to the RGD ligand. Assessment of cell adhesion and spreading in two dimensions was chosen to reduce the variability encountered with the use of 3D gels. Successful coupling of Fmoc-amino acids to amino-polyethylene glycol (PEG) glass was confirmed by checking for fluorescence of the Fmoc group using a fluorimeter with a solid state sample holder (Figure S2).20 NIH 3T3 fibroblasts were cultured on these substrates for 4 h, fixed, stained with Alexa Fluor 488-phalloidin and DAPI, and imaged. Fluorescence microscopy images shown in Figure 3 demonstrate significantly increased cell number and adhesion on RGD and Glc-RGD-presenting surfaces relative to other groups (Table 2); R-Glc-D surfaces were not statistically different from PEG-only and RGE surfaces in terms of cell attachment.

Figure 3.

Figure 3

Open in a new tab

NIH-3T3 fibroblast spreading on peptide-modified glass surfaces; blue represents DAPI and green represents Alexa Fluor 488-phalloidin. Scale bar = 50 μm.

Table 2. Results of Cell Number and Shape Analysis.

  cell number (10×) 40× area (μm2) perimeter (μm) circularity (n.d.)
PEG-glass 6 ± 4.1 864 ± 396 115 ± 26 0.80 ± 0.07
R-G-E 20 ± 5.3 554 ± 125.3 96 ± 17 0.77 ± 0.12
R-Glc-D 11 ± 6.4 1369 ± 1082 151 ± 52 0.69 ± 0.11
Glc-R-G-D 33 ± 7.1 1888 ± 1241 200 ± 49 0.53 ± 0.10
R-G-D 33 ± 8.0 2445 ± 984 297 ± 116 0.39 ± 0.14
Open in a new tab

Cell spreading was assessed by measuring the area and perimeter of attached cells in 40× images. Only cells on RGD and Glc-RGD surfaces were significantly greater for both metrics than cells on other surfaces. Cell circularity, a shape parameter to describe roundness, was also calculated for each measured cell. Cells on RGD and Glc-RGD surfaces tended to be less circular because of the spreading and extension of pseudopodia and filopodia, presumably because of the strong affinity of their surface integrin proteins for the peptide surface. Finally, upon examination of the micrographs, it is clear that actin filaments within cells on RGD and Glc-RGD surfaces are bundled into stress fibers, whereas on other surfaces, actin staining is diffused and delocalized. Stress fibers are characteristic of cells generating traction forces on a surface,21 and as they are not observed in cells on PEG, RGE, and R-Glc-D surfaces, we infer that cells are not likely to interact strongly or specifically with these molecules. In other words, these nonadhesive surfaces seem not to induce intracellular signaling pathways initiated by integrin binding and/or focal adhesion complex formation that ultimately result in cytoskeletal organization and stress fiber formation. From these results, it appears that hydrogen bonding by backbone amide groups within the RGD sequence may be as critical as the side-chain identity and sequence in mediating specific and functional binding with cell surface integrin proteins. Thus, while ester incorporation may be useful for introducing hydrolytic degradability into self-assembling peptide systems, the bioactive sequences must exclude rather than incorporate esters to retain full functionality.

Gelation of Self-Assembling Peptides and Depsipeptides

Fmoc-RGD has previously been reported to promote molecular self-assembly into hydrogels, which are able to support cell attachment and spreading in culture.22 In our initial efforts to synthesize a self-assembling RGD analogue, we found that generally, ester substitution of Fmoc-containing peptides resulted in much slower gelation kinetics. For example, we synthesized Fmoc-R-Glc-D and demonstrated that at a concentration of ∼15 mg/mL, this molecule self-assembles over the course of 2–3 days to form gels by pH switch both in phosphate-buffered saline (PBS) and deionized (DI) water with 50 mM NaCl added (see Figure S3). However, the slow gelation time limited the utility of Fmoc-R-Glc-D for functional tests of cell adhesion and spreading. In part, because of the work of Gazit and co-workers demonstrating the self-assembly of Fmoc-FRGD23 and also our own previous work with longer, charged depsipeptides,5 we hypothesized that adding a phenylalanine residue to Fmoc-R-Glc-D would dramatically decrease the time of gelation for our Fmoc-depsipeptides.

On the basis of previous gelation, integrin interaction, and cell attachment studies, Fmoc-F-Glc-RGD and Fmoc-FR-Glc-D were therefore selected as the self-assembling depsipeptide candidates for investigating cell attachment and degradation. These depsipeptides, as well as Fmoc-FRGD (positive control) and Fmoc-FRGE (negative control), were synthesized using a combination of standard solid phase peptide synthesis techniques and methods that our lab previously developed.24 All of the synthesized molecules, whose structures are given in Figure 4, were capable of forming a hydrogel via self-assembly as shown by vial inversion.

Figure 4.

Figure 4

Open in a new tab

Molecular structures of self-assembling peptides and depsipeptides. Fmoc-FRGD, Fmoc-FRGE, Fmoc-FR-Glc-D, and Fmoc-F-Glc-RGD are all able to form self-supporting hydrogels under solvent-exchange or pH switch conditions.

Excluding Fmoc-F-Glc-RGD, 5 mg/mL solutions of all molecules were capable of self-assembly leading to gelation both by pH switch and solvent exchange. In the case of Fmoc-F-Glc-RGD, a hydrogel was formed by solvent exchange when the concentration was above 10 mg/mL, but gelation was not observed with the pH switch method. Comparing Fmoc-FRGD hydrogels formed by different gelation methods, the storage modulus of the solvent exchange hydrogels was higher than that of the pH-switched hydrogels (Figure S4). Because local pH changes during acid addition result in spatial variations in gel density, the pH switch method generally results in a less homogeneous hydrogel than the solvent exchange method,25 and this could give rise to different rheological characteristics. Furthermore, the resulting hydrogel formed by solvent exchange (pH ≈ 4.8) had a lower pH than the hydrogel formed by pH switch (pH ≈ 5.5). The lower final pH may increase the ratio of molecules with a protonated C-terminal −OH group, thus increasing the average hydrophobicity of the molecules, which may in turn lead to a greater propensity for intermolecular association and a resulting increase in storage modulus. In addition, Raeburn et al. showed that the mechanical properties of self-assembled hydrogels can be tuned and influenced by the identity and the final volume fraction of the organic solvent.26 They demonstrated that the structure of the hydrogel network and the kinetics of gelation were affected by gelation conditions. Thus, the properties of self-assembled hydrogels depend both on the molecular structure and the gelation method.

Degradation of Self-Assembling Depsipeptides

Degradation profiles of 5 mg/mL Fmoc-FRGD (as a control), 5 mg/mL Fmoc-FR-Glc-D, and 10 mg/mL Fmoc-F-Glc-RGD hydrogels (all made by solvent exchange) were constructed by periodically analyzing gel samples using high-performance liquid chromatography (HPLC) and calculating the fractional chromatogram peak areas of the gelator molecule and its respective Fmoc-containing degradation product (Figure 5).

Figure 5.

Figure 5

Open in a new tab

Degradation profile of the self-assembling hydrogel of 5 mg/mL Fmoc-FRGD (A), 5 mg/mL Fmoc-FR-Glc-D (B), and 10 mg/mL Fmoc-F-Glc-RGD (C). Unlike Fmoc-FRGD, depsipeptides were gradually degraded over 60 days. All gels were made by solvent exchange.

In contrast with Fmoc-FRGD, which maintained its molecular structure, Fmoc-F-Glc-RGD and Fmoc-FR-Glc-D showed significant degradation over 2 months; a new peak in the chromatogram, corresponding to either Fmoc-FR or Fmoc-F, the ester hydrolysis products of Fmoc-FR-Glc-D and Fmoc-F-Glc-RGD, respectively, grew in fractional areas over time while the starting product peak decreased. Interestingly, both depsipeptides degraded relatively linearly during the early phase (approximately 30 days for Fmoc-FR-Glc-D and 50 days for Fmoc-F-Glc-RGD) according to the R2 value, which was 0.97 for Fmoc-FR-Glc-D and 0.93 for Fmoc-F-Glc-RGD, by the least squares method. At the termination of the degradation period, it was noted that the gels had not entirely collapsed. It is unknown whether the remaining gel structures are composed exclusively of the starting depsipeptides or the gels retain the degradation products Fmoc-FR and Fmoc-F in the fibrous nanostructure;27 however, no precipitates were observed visually at the end of the degradation period. If indeed Fmoc-FR and Fmoc-F are stabilized within the existing fibrous nanostructure rather than precipitated, it is possible that they contribute to the overall gel stability observed.

Cell Spreading and Viability on Self-Assembled Hydrogels

To assess the basic feasibility of using degradable self-assembling Fmoc-depsipeptides as cell-supporting matrices for tissue engineering, we performed two-dimensional (2D) cell culture experiments over the self-assembled solvent-exchanged hydrogels, and cell morphology and viability were measured (Figures 6 and 7A). For these experiments, we did not include Fmoc-FR-Glc-D, as earlier FP and cell spreading experiments described above suggested that the inclusion of the ester bond within the bioactive RGD sequence reduces its integrin-binding affinity to levels below than needed for proper biomechanical function. Thus, we chose to evaluate cell spreading and viability only on the degradable Fmoc-F-Glc-RGD gels, with the Fmoc-FRGD gels serving as the positive control.

Figure 6.

Figure 6

Open in a new tab

Bright-field and fluorescence microscopy images of fibroblasts on Fmoc-FRGD (A,D), Fmoc-F-Glc-RGD (B,E), and a mixture (10 mg/mL of Fmoc-FF and Fmoc-F-Glc-RGD) (C,F) of self-assembling hydrogels. Whereas most cells on Fmoc-F-Glc-RGD were round, extended cells were observed on Fmoc-FRGD and the hydrogel made from a mixture of Fmoc-FF and Fmoc-F-Glc-RGD. Scale bars = 200 μm (A–C) and 50 μm (D–F).

Figure 7.

Figure 7

Open in a new tab

Viability of fibroblasts on self-assembling hydrogel samples (A) and storage and loss modulus of cell-free gel samples (B). Statistical analysis was performed for several groups (** for p < 0.01 and *** for p < 0.001).

Rheological characterization of Fmoc-F-Glc-RGD and Fmoc-FRGD gels shows that at its minimal gelation concentration of 10 mg/mL, Fmoc-F-Glc-RGD is nearly 3 times softer than Fmoc-FRGD at half of that concentration (see Figures 7B and S4). To avoid confounding the effects of matrix compliance and integrin-binding affinity in assessing cell spreading,7 we opted to use Fmoc-FF as a co-gelator to provide mechanical support to the Fmoc-F-Glc-RGD gels. We chose Fmoc-FF because of its extensive characterization in the literature, relatively high storage modulus when gelled by solvent exchange (10–20 kPa), and previous use as a co-gelator for cell culture studies.28−31 We also hypothesized that Fmoc-FF molecules would interact favorably with the Fmoc- and phenylalanine side chain groups in Fmoc-F-Glc-RGD, thus enhancing the overall stability of the nanostructures formed.

Fibroblasts cultured on the Fmoc-FF hydrogel exhibited very little viable cell attachment (∼25%). In contrast, on the Fmoc-FRGD hydrogel, the viable cell attachment was near 100% of control with extended morphology. Cells on the Fmoc-F-Glc-RGD hydrogel, however, exhibited a rounded shape and an intermediate level of cell attachment at approximately 50% of control. As previously mentioned, we were concerned that the softness of the Fmoc-F-Glc-RGD hydrogel relative to the Fmoc-FRGD hydrogel (Figure 7B) could have an effect on the cell attachment, independent of the integrin binding.32 To resolve this question, Fmoc-FF, which is able to make a much stiffer hydrogel, was mixed with Fmoc-F-Glc-RGD.30 The stiffness of the hydrogel mixture depended on the relative amounts of the constituents and was reduced sharply as the concentration of Fmoc-F-Glc-RGD was increased (Figures 7B and S4). However, the Fmoc-FF/Fmoc-F-Glc-RGD mixture hydrogels were still more rigid than the single component Fmoc-F-Glc-RGD hydrogel. The mixture hydrogels also had an improved viable cell attachment and increased cell spreading above a concentration of 0.5 mg/mL of Fmoc-F-Glc-RGD compared to either the single component Fmoc-F-Glc-RGD hydrogel or Fmoc-FF hydrogel (Figures 6 and 7A). Fibroblasts on the Fmoc-FRGD hydrogel were viable after 24 h in agreement with other reports,23 but the numbers of viable fibroblasts on either the Fmoc-FF or the degradable hydrogels at longer time points were minimal. It is unclear whether this loss of viability over time for the degradable hydrogels is a result of loss of integrin-binding capacity (as the RGD ligand is cleaved thus resulting in cell detachment), a result of the effects of the degradation products on cell viability, or a combination of both factors.33

Summary and Conclusions

Our results demonstrate the potential of depsipeptides as the basis for self-assembling hydrogel materials with biological function and controlled hydrolytic degradation. We found that at least in the case of the RGD sequence, ester substitution within the bioactive sequence reduces the affinity for integrin in a manner that evades cell attachment and spreading. However, the depsipeptide with the ester flanking the bioactive RGD sequence, Fmoc-F-Glc-RGD, was able to form a hydrogel with relatively short kinetics and degrade approximately linearly over 60 days, in contrast with its nondegradable analogue. By mixing this engineered depsipeptide hydrogelator with the nonRGD containing Fmoc-FF, we were able to increase the stiffness of the resulting gel, which supported greater viable cell attachment and spreading than either the Fmoc-FF or Fmoc-F-Glc-RGD gels alone. The major limitations of the current design are the potential for RGD cleavage and loss of cell-binding capacity prior to the hydrogel disassembly as well as the unknown cytotoxicity of the degradation products, in particular, Fmoc-F. One factor confounding the decoupling of these effects is that Fmoc-F is not readily soluble in water at physiological pH and temperature and is not known to be able to directly form gels that might be used directly for cytotoxicity analysis. Our group is therefore investigating alternatives to the Fmoc-F terminus to drive the self-assembly.

Experimental Section

Solution-Phase Synthesis of Depsipeptide Units (Fmoc-R-Glc and Fmoc-F-Glc)

Depsipeptides were synthesized using a combinatorial solution- and solid-phase synthesis approach previously developed by our lab.24 Through this approach, the Fmoc-protected depsipeptide units of a specific amino- and α-hydroxy-acid combination can be incorporated into peptides synthesized on a solid resin support using well-established coupling chemistries and automated peptide synthesizing instruments. For the present study, we synthesized side-chain-protected Fmoc-arginine(pentamethyldihydrobenzylfuran-5-sulfonyl)-glycolic acid (abbreviated as Fmoc-R(Pbf)-Glc) and Fmoc-phenylalanine-glycolic acid (Fmoc-F-Glc) depsipeptide units in solution. Glycolic acid (Acros Organics) was first carboxyl-protected. Glycolic acid and 1.5 equiv benzyl chloride (Sigma-Aldrich) were dissolved in a minimal volume of ethyl acetate, and then, 1.5 equiv triethylamine (TEA, Acros Organics) was added. The mixture was refluxed at 85 °C overnight, filtered to remove TEA-chloride salts, and then distilled to remove excess benzyl chloride. Excess benzyl chloride was removed from the mixture by vacuum distillation to yield pure benzyl glycolate (Glc-Bn). Glc-Bn was then dissolved in dichloromethane (DCM) with 1.2 equiv Fmoc-Arg(Pbf)-OH (Novabiochem) and 0.01 equiv dimethylaminopyridine, and the mixture was chilled on ice. In the case of a reaction with Glc-Bn and Fmoc-Phe-OH (Novabiochem), a minimal volume of dimethylformamide (DMF) was used for dissolving Fmoc-Phe-OH. To the chilled mixture, was added 1.2 equiv dicyclohexylcarbodiimide. The reaction was allowed to warm to room temperature overnight, after which the mixture was filtered to remove dicyclohexylurea crystals and concentrated in vacuo. The carboxyl-protected Fmoc-depsipeptide product (either Fmoc-R(Pbf)-Glc-Bn or Fmoc-F-Glc-Bn) was then purified by flash chromatography (av yield ≈ 85–90%) and concentrated in vacuo. Pure benzylated Fmoc-depsipeptides were benzyl-deprotected by catalytic hydrogenolysis, following the general method developed by Bajwa.34 Fmoc-R(Pbf)-Glc-Bn or Fmoc-F-Glc-Bn was dissolved in a minimal volume of absolute ethanol with DCM added to aid the dissolution. To this mixture, 10 equiv 1,4-cyclohexadiene (Acros Organics) was added, followed by a mass of palladium on carbon (Pd/C, 10% Pd, Acros Organics) equivalent to the mass of the reactant. This mixture was stirred at room temperature for at least 3 h, and the reaction was monitored by thin layer chromatography (TLC). The mixture was then filtered in vacuo through a Celite pad and triturated with hot ethanol and hot DCM, which was then reconcentrated in vacuo. The crude product was then dissolved in <10 mL of dimethylsulfoxide (DMSO) and purified by reversed-phase (RP) flash chromatography using a RediSep Gold C18 column on a CombiFlash instrument (Teledyne Isco) with a 45 min linear gradient of 0–95% acetonitrile in water. The pure product was then lyophilized from a concentrated solution in acetonitrile and stored as a powder at −20 °C for future use.

Solid-Phase Coupling and Purification of Fmoc-Peptides and Fmoc-Depsipeptides

All peptides and depsipeptides were coupled on solid phase using diisopropylcarbodiimide (DIC)/OxymaPure (ethyl 2-cyano-2-(hydroxyimino)acetate) amide coupling chemistry. Depending on the desired sequence, either Fmoc-Asp(OtBu)-Wang resin or Fmoc-Glu(OtBu)-Wang resin was swelled in DCM in a fritted, capped syringe for 20 min. The resin was rinsed with DMF, and then Fmoc groups were removed with 20% piperidine in DMF (5 mL). After addition to the resin, the capped syringe was placed in a BioWave scientific microwave oven (Ted Pella, Inc.) operating at ∼100 W and subjected to microwave energy in 30 s increments for a total of 2 min. At the end of each 30 s increment, the mixture was vortexed. This process was repeated twice more with fresh piperidine solution to ensure complete Fmoc removal. The resin was then washed (vortexed 4 × 15 mL DMF followed by 4 × 15 mL DCM) and a coupling solution containing the next Fmoc-amino acid was added to the resin. Coupling solutions consisted of 3 equiv of the Fmoc-amino acid and 3 equiv of OxymaPure in less than 10 mL of DMF, to which 3 equiv of DIC was added. The mixture was allowed to pre-activate for at least 10 min before adding to resin (for most amino acids, a yellow color develops upon DIC addition). After microwaving (5 min at ∼100 W with mixing every 30 s), the resin was washed (vortexed 4 × 15 mL DMF followed by 4 × 15 mL DCM). The subsequent coupling steps followed the same Fmoc-deprotection, washing, and coupling methods. For coupling the solution-synthesized Fmoc-depsipeptide units, a similar protocol was used, with a slight difference. For the coupling solution, only 2 equiv of Fmoc-R(Pbf)-Glc or Fmoc-F-Glc was used in an effort to conserve the material.

Upon completion of coupling, the peptide (or depsipeptide) was cleaved from the resin and side chains were simultaneously deprotected by adding 5 mL of a solution of 95:2.5:2.5 trifluoroacetic acid/water/triisopropylsilane (TFA/H2O/TIPS) and mixing for 1.5–2 h. The mixture was then collected in a clean round bottom flask, and subsequent resin washes (1 × 5 mL TFA/H2O/TIPS and 5 × 10 mL DCM) were added to the flask before concentrating in vacuo on a rotary evaporator. Excess TFA and H2O were removed by repeatedly adding acetone or DCM and reconcentrating. The resulting oil was dissolved in <5 mL DMF and the product was precipitated with cold diethyl ether to form a white product. The mixture was split into separate centrifuge tubes, centrifuged and triturated and vortexed with cold ether, and then recombined into a single tube, followed by three subsequent rounds of centrifugation and trituration/vortexing to remove excess TFA and scavenged side-chain protecting groups. After drying the product under a gentle stream of N2, the product was redissolved in DMF or DMSO and purified by RP-HPLC (0% H2O initially for 10–15 min to remove excess DMF/DMSO, followed by a 30–40 min linear gradient of 0–90% acetonitrile in water). Pure product fractions were collected and lyophilized at a low temperature (−100 °C collector) for 2 days.

FP Assays

RGD, R-Glc-D, and RGE peptides were synthesized and purified as their Fmoc-terminated versions, as described above. After purification by HPLC and lyophilization, Fmoc-RGD, Fmoc-R-Glc-D, or Fmoc-RGE was dissolved in a minimal volume of DMF in a round-bottom flask, and 200 μL of piperidine was added to the mixture. The mixture was stirred, and within 2 min, a white precipitate (peptide) formed. Cold diethyl ether was added to the flask, and the suspension was divided among four 50 mL centrifuge tubes. The precipitate was centrifuged and then resuspended and combined in one tube. The aggregated precipitate was then washed and centrifuged four times using fresh, cold diethyl ether each time. After decanting the ether supernatant from the final wash, nitrogen gas was gently blown over the pellet to remove the excess diethyl ether. The pellet was dissolved in a minimal volume of DI water and filtered to remove colloidal dibenzofulvene particles. Removal by filtration was confirmed using TLC. The clear filtrate was then lyophilized, and the peptide identity and purity was confirmed using TLC.

For FP assays, inhibitor peptides (RGD—positive control and R-Glc-D and RGE—negative control) were diluted in series from 3 × 10–9 to 3 × 10–2 M in Tris with 10 mM MgCl2, 10 mM CaCl2, 1 mM MnCl2, and 100 mM NaCl at pH 7.4. Human recombinant integrin α5β1 (R&D Systems) was diluted to 900 nM in the same Tris buffer. FITC-GRGDSP (Anaspec, Inc.) was also diluted in the same Tris buffer to 30 nM. To each test well in a black-bottom 384-well plate, 10 μL of the chosen dilution of RGD, R-Glc-D, or RGE was added to 20 μL of a 1:1 integrin/FITC-GRGDSP solution. For the positive control well, 10 μL of buffer was added in the place of inhibitor solution. For the negative control well, 10 μL of FITC-GRGDSP alone was added to 20 μL of Tris buffer. Plates were covered and incubated at room temperature at least 30 min before use. The FP of the wells was read using a BioTek Synergy H4 instrument operating at default FITC excitation (λ = 485 nm) and emission (λ = 528 nm) wavelengths. Fluorescence anisotropy data from different experiments were normalized by calculating the percentage of each well’s value relative to the highest anisotropy value observed in any given experiment.

Covalent Functionalization of Glass Coverslips with Amine-Terminated PEG

Glass surfaces were functionalized with RGD peptides or R-Glc-D and Glc-RGD depsipeptides to assess cell adhesion and spreading in a system that facilitates imaging and quantitative assessment. Functionalization was accomplished using the methods described by Todd et al.35 Circular glass coverslips (18 mm diameter, thickness “2”) were cleaned by sonicating for 10 min in acetone, followed by rinsing with DI water. Coverslips were then submerged in 3 M NaOH for 5 min, rinsed with DI water, and submerged in a piranha solution (30 v/v % concentrated sulfuric acid and 70 v/v % hydrogen peroxide). After 1 h, coverslips were removed and rinsed in a large excess of DI water and air-dried completely. An epoxide-terminated silane, (3-glycidyloxipropyl)trimethoxysilane (GOPTS, Sigma-Aldrich), was pipetted onto the surface of half the coverslips, and the remaining coverslips were placed on top of the GOPTS to reduce air exposure. The sandwiched coverslips were then placed in an oven at 37 °C for at least 2 h, after which the coverslips were washed with dry acetone and dried under N2. Immediately following, homobifunctional diamino-PEG (H2N–PEG–NH2) in dry powder form was placed directly on the silane-functionalized surface of half of the coverslips, which were then placed in an oven at ∼75 °C. The PEG powder melted into a semiviscous liquid, and while still in the oven, the remaining coverslips were placed on top of the PEG-covered coverslips with the silane side facing down. The coverslip–PEG assemblies were incubated at 75 °C for 48 h to provide sufficient time for complete coupling of the PEG-terminal amine groups to the epoxide groups on the silanated glass. After incubation, coverslips were disassembled while still hot (to prevent PEG recrystallization/hardening) and were rinsed with a large excess of DI water. Coverslips were then marked to indicate the non-PEGylated side.

Peptide Synthesis on Glass Substrates

For solid-phase synthesis of peptides on PEGylated glass substrates, coverslips were immersed in preactivated coupling solutions and subjected to microwave energy. Coupling solutions were similar to those prepared for normal resin-bound peptide synthesis, with 0.2–0.8 mmol Fmoc-amino acid or glycolic acid and 2 equiv of OxymaPure dissolved in ∼6 mL of DMF, followed by addition of 2 equiv of DIC and 10 min of preactivation prior to coupling. For each amino acid coupling step, 2 mL of coupling solution was added to a glass Petri dish containing several coverslips with the PEGylated side facing up, and the coverslips were microwaved for 3 min at 250 W (very little heating was observed). This process was repeated two more times with a fresh coupling solution each time. Coverslips were then washed thoroughly with DMF. To remove Fmoc groups, ∼2 mL 20% piperidine in DMF was added to the Petri dish and the dish was microwaved for 2 min. This process was repeated two more times with fresh piperidine solution, and then, coverslips were washed thoroughly with DMF. Subsequent Fmoc-amino acid or Fmoc-depsipeptide couplings were performed as described above until the full peptide was generated. In the case of coupling glycolic acid for Glc-RGD-functionalized PEG-glass, glycolic acid was used directly as a reactant with the standard coupling reagents, and the reaction time was monitored using the Kaiser color test for free amines.36

Solid-state fluorescence spectroscopy was used to confirm the coupling of peptides to the PEG-glass surfaces through the fluorescence of the Fmoc group at an excitation wavelength of 270 nm and emission spectrum from 290 to 360 nm. Finally, to cleave peptide side-chain protecting groups but leave the peptide C-terminus attached to the PEG-glass, a 95:2.5:2.5 TFA/H2O/TIPS mixture was applied to the coverslips and allowed to incubate at room temperature for 2 h, followed by thorough washing with DMF, DCM, and water. Coverslips were sterilized by soaking in 70% ethanol in DI water prior to seeding with cells.

Cell Culture and Spreading Assessment on Functionalized Glass

NIH 3T3 fibroblasts were cultured from the frozen stock in standard tissue culture flasks and passaged at least twice before experimental use. For adhesion and spreading on glass substrates, cells were trypsinized, centrifuged, and resuspended in a serum-free medium. Peptide-functionalized glass substrates were placed in separate wells of 12-well plates and washed 2× with PBS. Cell suspensions were added at a seeding density of ∼6500 cells/cm2 and incubated for 4–5 h. Nonadherent cells were washed away with PBS, and adherent cells were fixed with 4% formalin in PBS for 10 min. After fixation, coverslips were washed with PBS and cells were permeabilized with a 0.1% Triton X-100 solution. After washing 3× with PBS, cell actin filaments were stained with Alexa Fluor 488 phalloidin (Life Technologies) according to the manufacturer’s protocol and incubated for 30 min. The solution was removed, and a DAPI nuclear stain (5 μg/mL) was applied and allowed to incubate for 10 min. After washing 3× with PBS, coverslips were removed from wells, mounted on glass microscope slides, and imaged on a microscope with fluorescence excitation using DAPI and FITC filter cubes. Images were analyzed using ImageJ. The cell number in 15 separate 10× images was assessed by automated counting of DAPI-stained nuclei through the use of a custom thresholding and particle counting macro. Cell area, perimeter, and circularity were measured using the ImageJ trace and a measure tool on 10 representative cells from images at 40× magnification. The pixel-to-distance relationship used to calculate the actual distance and area metrics in micrometer was established by capturing a photo of a hemocytometer grid at 40× using the same microscope and attached camera used for cell imaging. Statistical analysis was performed using GraphPad Prism 6. A one-way analysis of variance test was performed, followed by Tukey’s test for multiple comparisons. P-values corrected for multiple comparisons are reported.

Gelation of Self-Assembling Peptides and Depsipeptides

Two methods, pH switch and solvent exchange, were employed to generate the hydrogels. For the pH switch method, a solution of 1 equiv 0.5 M NaOH with 1 equiv gelator molecule solution was prepared and mixed until it became transparent, after which 1 equiv 0.1 M HCl was added to the solution and gently mixed. The solution was left undisturbed until gelation was observed. In the case of the solvent exchange method, a 50× solution (i.e., 50 times the desired final concentration) of the gelator molecule in the DMSO solution was prepared. A 49-to-1 volume ratio of DI water to the gelator solution was added and gently mixed to achieve the final desired concentration, and the solutions were left undisturbed until gelation was observed. When making the mixed “co-gelator” hydrogels by solvent exchange, 100× solutions of each gelator molecule in DMSO were prepared individually and then premixed before adding DI water in the proper volume ratio. In this manner, the gels with varying co-gelator ratios could be made, but all had the same final DMSO content.

Degradation Profile Assessment

The degradation characteristics of the Fmoc-F-Glc-RGD, Fmoc-FR-Glc-D, and Fmoc-FRGD hydrogels (made by solvent exchange with DI water, at a concentration of 5 mg/mL except Fmoc-F-Glc-RGD at 10 mg/mL) were assessed by the following method. The gels were formed in tight-sealing microcentrifuge tubes and placed in a 37 °C incubator, and at specified time points, 50 μL of each sample gel was dissolved in 450 μL methanol and analyzed by LC–MS. Fractional peak area percentages were calculated from peak integrations and plotted as a function of time. Evaporation of liquid was not visually evident, nor was any obvious bacterial contamination.

Gel Rheology

Storage and loss modulus of the Fmoc-FF, Fmoc-FRGD, Fmoc-F-Glc-RGD, and mixtures of Fmoc-FF and Fmoc-FRGD or Fmoc-F-Glc-RGD hydrogels (made by solvent exchange) were measured by an Anton-Paar MCR101 rheometer with a parallel plate geometry (top plate diameter of 8 mm). Briefly, self-assembled hydrogels were formed in polydimethylsiloxane molds (3 mm deep, 8 mm diameter). Oscillatory shear stress rheometry was performed (1% strain, 0.5–100 Hz), and the storage and loss modulus of the hydrogels at 1.99 Hz were used for analysis.

2D Cell Morphology Analysis and Viability Assay

Two-dimensional cell spreading and viability experiments were performed by seeding 3T3 fibroblasts on the surfaces of hydrogels of Fmoc-FF, Fmoc-FRGD, Fmoc-F-Glc-RGD, and mixtures of Fmoc-FF and Fmoc-FRGD or Fmoc-F-Glc-RGD. DMSO/gelator solutions were sterile-filtered to prevent bacterial contamination. Before cell seeding, 100 μL of each hydrogel (made by solvent exchange using sterile DI water) was formed in a 48-well microplate and equilibrated with serum-free Dulbecco’s modified Eagle’s medium-containing antibiotics. This equilibration was performed twice to remove most of the DMSO and equilibrate the pH of the gels. Aliquots of 2 × 104 fibroblasts in serum-free media were transferred to each hydrogel-containing well, and the complete medium was added after 2 h of incubation. The cell morphology was analyzed using bright-field and fluorescence imaging (Alexa Fluor 488 phalloidin and DAPI), and an MTS [3-(4,5-dimethylthiazol-2-yl)-5-(3-carboxymethoxyphenyl)-2-(4-sulfophenyl)-2H-tetrazolium, inner salt, CellTiter 96, Promega] assay was performed to measure the viable cell attachment after 6 h.

Acknowledgments

K.M.E. acknowledges the National Science Foundation for support through the Graduate Research Fellowship Program. This work was also supported by the National Science Foundation under award number DMR-1609212.

Supporting Information Available

The Supporting Information is available free of charge on the ACS Publications website at DOI: 10.1021/acsomega.7b01641.

  • Fluorescence polarization and fluorescence spectroscopy experiments, Fmoc-R-Glc-D purification and gelation, rheometric data, and depsipeptide purification (PDF)

Author Contributions

K.M.E. and K.B. contributed equally to this work. All authors have given approval to the final version of this manuscript.

The authors declare no competing financial interest.

Supplementary Material

ao7b01641_si_001.pdf (982.6KB, pdf)

References

  1. Page-McCaw A.; Ewald A. J.; Werb Z. Matrix metalloproteinases and the regulation of tissue remodelling. Nat. Rev. Mol. Cell Biol. 2007, 8, 221–233. 10.1038/nrm2125. [DOI] [PMC free article] [PubMed] [Google Scholar]
  2. Sekhon B. Matrix metalloproteinases—an overview. Res. Rep. Biol. 2010, 1, 1–20. 10.2147/RRB.S12043. [DOI] [Google Scholar]
  3. Eckes K. M.; Mu X.; Ruehle M. A.; Ren P.; Suggs L. J. β Sheets not required: Combined experimental and computational studies of self-assembly and gelation of the ester-containing analogue of an Fmoc-dipeptide hydrogelator. Langmuir 2014, 30, 5287–5296. 10.1021/la500679b. [DOI] [PMC free article] [PubMed] [Google Scholar]
  4. Tian Y. F.; Hudalla G. A.; Han H.; Collier J. H. Controllably degradable β-sheet nanofibers and gels from self-assembling depsipeptides. Biomater. Sci. 2013, 1, 1037–1045. 10.1039/c3bm60161g. [DOI] [PMC free article] [PubMed] [Google Scholar]
  5. Nguyen M. M.; Eckes K. M.; Suggs L. J. Charge and sequence effects on the self-assembly and subsequent hydrogelation of Fmoc-depsipeptides. Soft Matter 2014, 10, 2693–2702. 10.1039/c4sm00009a. [DOI] [PMC free article] [PubMed] [Google Scholar]
  6. Ruoslahti E.; Pierschbacher M. D. Arg-Gly-Asp: a versatile cell recognition signal. Cell 1986, 44, 517–518. 10.1016/0092-8674(86)90259-x. [DOI] [PubMed] [Google Scholar]
  7. Pelham R. J.; Wang Y.-l. Cell locomotion and focal adhesions are regulated by substrate flexibility. Proc. Natl. Acad. Sci. U.S.A. 1997, 94, 13661–13665. 10.1073/pnas.94.25.13661. [DOI] [PMC free article] [PubMed] [Google Scholar]
  8. Maheshwari G.; Brown G.; Lauffenburger D. A.; Wells A.; Griffith L. G. Cell adhesion and motility depend on nanoscale RGD clustering. J. Cell Sci. 2000, 113, 1677–1686. [DOI] [PubMed] [Google Scholar]
  9. Engler A. J.; Sen S.; Sweeney H. L.; Discher D. E. Matrix elasticity directs stem cell lineage specification. Cell 2006, 126, 677–689. 10.1016/j.cell.2006.06.044. [DOI] [PubMed] [Google Scholar]
  10. Huebsch N.; Arany P. R.; Mao A. S.; Shvartsman D.; Ali O. A.; Bencherif S. A.; Rivera-Feliciano J.; Mooney D. J. Harnessing traction-mediated manipulation of the cell/matrix interface to control stem-cell fate. Nat. Mater. 2010, 9, 518–526. 10.1038/nmat2732. [DOI] [PMC free article] [PubMed] [Google Scholar]
  11. Beligere G. S.; Dawson P. E. Design, synthesis, and characterization of 4-ester CI2, a model for backbone hydrogen bonding in protein α-helices. J. Am. Chem. Soc. 2000, 122, 12079–12082. 10.1021/ja001648e. [DOI] [Google Scholar]
  12. Jenkins C. L.; Vasbinder M. M.; Miller S. J.; Raines R. T. Peptide bond isosteres: Ester or (E)-alkene in the backbone of the collagen triple helix. Org. Lett. 2005, 7, 2619–2622. 10.1021/ol050780m. [DOI] [PubMed] [Google Scholar]
  13. Koh J. T.; Cornish V. W.; Schultz P. G. An experimental approach to evaluating the role of backbone interactions in proteins using unnatural amino acid mutagenesis. Biochemistry 1997, 36, 11314–11322. 10.1021/bi9707685. [DOI] [PubMed] [Google Scholar]
  14. Deechongkit S.; Nguyen H.; Powers E. T.; Dawson P. E.; Gruebele M.; Kelly J. W. Context-dependent contributions of backbone hydrogen bonding to β-sheet folding energetics. Nature 2004, 430, 101–105. 10.1038/nature02611. [DOI] [PubMed] [Google Scholar]
  15. Gordon D. J.; Meredith S. C. Probing the role of backbone hydrogen bonding in β-amyloid fibrils with inhibitor peptides containing ester bonds at alternate positions. Biochemistry 2003, 42, 475–485. 10.1021/bi0259857. [DOI] [PubMed] [Google Scholar]
  16. Elgersma R. C.; Meijneke T.; Posthuma G.; Rijkers D. T. S.; Liskamp R. M. J. Self-assembly of amylin (20–29) amide-bond derivatives into helical ribbons and peptide nanotubes rather than fibrils. Chem.—Eur. J. 2006, 12, 3714–3725. 10.1002/chem.200501374. [DOI] [PubMed] [Google Scholar]
  17. Eildal J. N. N.; Hultqvist G.; Balle T.; Stuhr-Hansen N.; Padrah S.; Gianni S.; Strømgaard K.; Jemth P. Probing the role of backbone hydrogen bonds in protein–peptide interactions by amide-to-ester mutations. J. Am. Chem. Soc. 2013, 135, 12998–13007. 10.1021/ja402875h. [DOI] [PubMed] [Google Scholar]
  18. Pedersen S. W.; Pedersen S. B.; Anker L.; Hultqvist G.; Kristensen A. S.; Jemth P.; Strømgaard K. Probing backbone hydrogen bonding in PDZ/ligand interactions by protein amide-to-ester mutations. Nat. Commun. 2014, 5, 3215. 10.1038/ncomms4215. [DOI] [PubMed] [Google Scholar]
  19. Nikolovska-Coleska Z.; Wang R.; Fang X.; Pan H.; Tomita Y.; Li P.; Roller P. P.; Krajewski K.; Saito N. G.; Stuckey J. A.; Wang S. Development and optimization of a binding assay for the XIAP BIR3 domain using fluorescence polarization. Anal. Biochem. 2004, 332, 261–273. 10.1016/j.ab.2004.05.055. [DOI] [PubMed] [Google Scholar]
  20. Zelzer M.; Scurr D. J.; Alexander M. R.; Ulijn R. V. Development and validation of a fluorescence method to follow the build-up of short peptide sequences on solid 2D surfaces. ACS Appl. Mater. Interfaces 2012, 4, 53–58. 10.1021/am2015266. [DOI] [PubMed] [Google Scholar]
  21. Pellegrin S.; Mellor H. Actin stress fibres. J. Cell Sci. 2007, 120, 3491–3499. 10.1242/jcs.018473. [DOI] [PubMed] [Google Scholar]
  22. Cheng G.; Castelletto V.; Jones R. R.; Connon C. J.; Hamley I. W. Hydrogelation of self-assembling RGD-based peptides. Soft Matter 2011, 7, 1326–1333. 10.1039/C0SM00408A. [DOI] [Google Scholar]
  23. Orbach R.; Adler-Abramovich L.; Zigerson S.; Mironi-Harpaz I.; Seliktar D.; Gazit E. Self-assembled Fmoc-peptides as a platform for the formation of nanostructures and hydrogels. Biomacromolecules 2009, 10, 2646–2651. 10.1021/bm900584m. [DOI] [PubMed] [Google Scholar]
  24. Nguyen M. M.; Ong N.; Suggs L. A general solid phase method for the synthesis of depsipeptides. Org. Biomol. Chem. 2013, 11, 1167–1170. 10.1039/c2ob26893k. [DOI] [PubMed] [Google Scholar]
  25. Adams D. J.; Butler M. F.; Frith W. J.; Kirkland M.; Mullen L.; Sanderson P. A new method for maintaining homogeneity during liquid–hydrogel transitions using low molecular weight hydrogelators. Soft Matter 2009, 5, 1856–1862. 10.1039/b901556f. [DOI] [Google Scholar]
  26. Raeburn J.; Mendoza-Cuenca C.; Cattoz B. N.; Little M. A.; Terry A. E.; Cardoso A. Z.; Griffiths P. C.; Adams D. J. The effect of solvent choice on the gelation and final hydrogel properties of Fmoc–diphenylalanine. Soft Matter 2015, 11, 927–935. 10.1039/c4sm02256d. [DOI] [PubMed] [Google Scholar]
  27. Sutton S.; Campbell N. L.; Cooper A. I.; Kirkland M.; Frith W. J.; Adams D. J. Controlled release from modified amino acid hydrogels governed by molecular size or network dynamics. Langmuir 2009, 25, 10285–10291. 10.1021/la9011058. [DOI] [PubMed] [Google Scholar]
  28. Raeburn J.; Pont G.; Chen L.; Cesbron Y.; Lévy R.; Adams D. J. Fmoc-diphenylalanine hydrogels: understanding the variability in reported mechanical properties. Soft Matter 2012, 8, 1168–1174. 10.1039/c1sm06929b. [DOI] [PubMed] [Google Scholar]
  29. Jayawarna V.; Ali M.; Jowitt T. A.; Miller A. F.; Saiani A.; Gough J. E.; Ulijn R. V. Nanostructured hydrogels for three-dimensional cell culture through self-assembly of fluorenylmethoxycarbonyl–dipeptides. Adv. Mater. 2006, 18, 611–614. 10.1002/adma.200501522. [DOI] [Google Scholar]
  30. Zhou M.; Smith A. M.; Das A. K.; Hodson N. W.; Collins R. F.; Ulijn R. V.; Gough J. E. Self-assembled peptide-based hydrogels as scaffolds for anchorage-dependent cells. Biomaterials 2009, 30, 2523–2530. 10.1016/j.biomaterials.2009.01.010. [DOI] [PubMed] [Google Scholar]
  31. Zhou M.; Ulijn R. V.; Gough J. E. Extracellular matrix formation in self-assembled minimalistic bioactive hydrogels based on aromatic peptide amphiphiles. J. Tissue Eng. 2014, 5, 204173141453159. 10.1177/2041731414531593. [DOI] [PMC free article] [PubMed] [Google Scholar]
  32. Yeung T.; Georges P. C.; Flanagan L. A.; Marg B.; Ortiz M.; Funaki M.; Zahir N.; Ming W.; Weaver V.; Janmey P. A. Effects of substrate stiffness on cell morphology, cytoskeletal structure, and adhesion. Cell Motil. Cytoskeleton 2005, 60, 24–34. 10.1002/cm.20041. [DOI] [PubMed] [Google Scholar]
  33. Salinas C. N.; Anseth K. S. The enhancement of chondrogenic differentiation of human mesenchymal stem cells by enzymatically regulated RGD functionalities. Biomaterials 2008, 29, 2370–2377. 10.1016/j.biomaterials.2008.01.035. [DOI] [PMC free article] [PubMed] [Google Scholar]
  34. Bajwa J. S. Chemoselective deprotection of benzyl esters in the presence of benzyl ethers, benzyloxymethyl ethers and N-benzyl groups by catalytic transfer hydrogenation. Tetrahedron Lett. 1992, 33, 2299–2302. 10.1016/s0040-4039(00)74195-5. [DOI] [Google Scholar]
  35. Todd S. J.; Scurr D. J.; Gough J. E.; Alexander M. R.; Ulijn R. V. Enzyme-activated RGD ligands on functionalized poly(ethylene glycol) monolayers: surface analysis and cellular response. Langmuir 2009, 25, 7533–7539. 10.1021/la900376h. [DOI] [PubMed] [Google Scholar]
  36. Kaiser E.; Colescott R. L.; Bossinger C. D.; Cook P. I. Color test for detection of free terminal amino groups in the solid-phase synthesis of peptides. Anal. Biochem. 1970, 34, 595–598. 10.1016/0003-2697(70)90146-6. [DOI] [PubMed] [Google Scholar]

Associated Data

This section collects any data citations, data availability statements, or supplementary materials included in this article.

Supplementary Materials

ao7b01641_si_001.pdf (982.6KB, pdf)

Articles from ACS Omega are provided here courtesy of American Chemical Society

RESOURCES