pH-Responsive Peptide-Polymer Hydrogel for Biofilm Disruption

Abstract

Biofilm formation presents a significant challenge in chronic infections, as it enables bacteria to resist conventional antibiotics and thrive in various areas of the body. The treatment is further hurdled by the acidic environment of biofilms due to anaerobic glycolysis of bacteria and the accumulation of acidic by-products. Therefore, there is a need for the development of antimicrobial materials which can selectively and preferentially eradicate biofilms in the acidic environment. Toward this aim, this study explores the use of acid-responsive double-network peptide-polymer hydrogels, encapsulated with antimicrobial peptides, to effectively target and disrupt biofilms. The hydrogel consists of two essential components: a self-assembling peptide nanofiber containing a non-natural ionic amino acid, which imparts pH responsiveness in the weakly acidic range, and a 4-arm PEG polymer that forms covalent bonds with the peptide nanofiber, enhancing the hydrogel’s mechanical strength. Upon acidification, peptide nanofibers disassembled causing an increased pore size of the hydrogel and release of encapsulated antimicrobials to the biofilm site. We expect that by leveraging the unique properties of the double network self-assembled peptide-PEG hydrogels and pH-triggered release mechanism, this innovative hydrogel approach may offer a more targeted, effective, and safer treatment option against biofilm-associated infections.

INTRODUCTION

Bacterial biofilms are a major challenge in wound healing, as they can significantly hinder recovery and complicate treatment strategies.^1–4 Biofilms are formed when bacteria adhere to surfaces and secrete extracellular polymeric substances, creating a protective environment that shields the bacteria from both the immune system and antimicrobial agents.^{5, 6} These biofilm structures are often associated with chronic infections and are difficult to treat as they are innately antibiotic-resistant.⁷ The presence of biofilm in infected wounds not only delays the healing process but also increases the risk of recurrent infections and the development of antimicrobial resistance, making it crucial to develop new, more effective prevention and treatment strategies.^{8, 9}

A promising approach to combating biofilm-associated infections is the use of antimicrobial peptide hydrogel.^10–17 These hydrogels combine the antimicrobial properties of peptides with the structural advantages of hydrogels, offering a versatile and efficient tool to treat infected wounds.¹⁸ Additionally, hydrogels mimic the extracellular matrix (ECM) found in natural tissues, providing a scaffold that supports cell growth, migration, and tissue regeneration and eventually promotes healing.^19–21 This combination of antimicrobial activity and tissue-supportive properties makes peptide-based hydrogels particularly useful in promoting wound healing and eradicating biofilms.²² In our lab, we have developed a novel self-healing injectable hydrogel by incorporating a polymer into the peptide matrix.¹² This hydrogel offers several advantages, including enhanced injectability, self-healing capabilities, and improved mechanical strength. The peptide-polymer conjugate exhibits two key mechanisms that contribute to its gel strength: first, the self-assembly of the peptides into nanostructures, which initially forms a weaker gel; and second, the covalent bonding between the peptide and the polymer, which reinforces the structure and enhances rigidity. This double network enables the hydrogel to have both flexibility for injection and stability for effective treatment. This strategy also enables the encapsulation of antimicrobial agents within the gel making them an antimicrobial hydrogel.

In spite of these advancements, combating biofilm infections still poses challenges.²³ One of the main challenges is delivering targeted antimicrobial agents to the infected site, especially if the environment is low in pH, such as those found in wound infection.^{24, 25} The acidic microenvironment commonly found at infected sites can hinder the efficacy of conventional antimicrobial treatments. Utilizing the acidic pH at the infected sites, we aim to develop a pH-responsive peptide-polymer conjugate hydrogel which can trigger controlled release of encapsulated antimicrobials at the infected sites. In our hypothesis, the pH-responsive peptide–polymer double-network hydrogels are initially formed at physiological pH (7.4), where the stable network structure facilitates efficient encapsulation of antimicrobial peptides. Upon application to biofilm-infected sites, the acidic microenvironment (lower pH) destabilizes the self-assembled peptide structure, leading to the controlled release of the encapsulated antimicrobial agent. Conversely, in non-infected regions that maintain physiological pH, the hydrogel network remains stable, thereby minimizing drug release and enhancing biocompatibility.

The peptide-polymer hydrogel often needs a high concentration of antimicrobial agents as the release of the antimicrobial is very minimal. Incorporating the pH responsivity to the gel can provide more controlled release if the hydrogel disassembles at the lower pH to deliver the antimicrobials therefore reducing the amount of antimicrobial needed for effective treatment, thereby minimizing potential cytotoxicity, and improving efficacy of the hydrogel. Although numerous hydrogel-based systems have been developed for pH-responsive application, most efforts have focused on polymer-based systems.^26–29 Although peptide-based hydrogels are often favored for their biocompatibility and tunable properties at the single amino acid level, they typically suffer from limited mechanical strength due to weak intermolecular interactions, which can restrict their ability to form stable hydrogels. Nonetheless, several pH-responsive peptide-based hydrogel systems have been successfully developed and applied across a range of biomedical applications.^30–32

In this study, we aim to generate pH-responsive double-network peptide-polymer hydrogels using multidomain peptides (MDPs) that contain non-natural ionic amino acids. The non-natural amino acid with tertiary amine side chains introduced into the peptide component will impart pH-sensitive disassembly properties, while the polymer component will provide additional structural support.^{12, 33} We hypothesize that the hydrogel will remain stable and retain its strength under neutral pH conditions but will disassemble at lower pH values due to protonation of the tertiary amine side chains. This pH-triggered disassembly will trigger an internal structural change and the controlled release of antimicrobial agents at the infection site, allowing for more efficient treatment of biofilm infections.^{34, 35} To evaluate the effectiveness of this new hydrogel, we will use Methicillin-resistant Staphylococcus aureus (MRSA) as a model organism for biofilm eradication studies. MRSA is a common and problematic pathogen that is known for its ability to form biofilms and its resistance to many conventional antibiotics.^{9, 36} By testing the hydrogel against MRSA biofilms, we can assess the hydrogel’s ability to penetrate biofilm structures, release antimicrobial agents in response to acidic pH, and ultimately reduce biofilm viability.

EXPERIMENTAL SECTION

Materials.

Reagents used for the synthesis of the non-natural amino acid Fmoc-Xp, including (S)-3-amino-2-(tert-butoxycarbonylamino) propionic acid, propionaldehyde, sodium cyanoborohydride, and N-(9-fluorenylmethoxycarbonyloxy)succinimide, were obtained from Fisher Scientific. Solvents and reagents for peptide synthesis and purification, such as dimethylformamide (DMF), diisopropylethylamine (DIPEA), acetonitrile (ACN), trifluoroacetic acid (TFA), and triisopropylsilane (TIS), were also sourced from Fisher Scientific. Fmoc-protected amino acids, hexafluorophosphate benzotriazole tetramethyluronium (HBTU), MBHA rink amide resin, piperidine, pyridine, Mueller Hinton Broth (MHB), and agar were purchased from Sigma-Aldrich. 4-Arm PEG-Glutaramide Acid Succinimidyl Ester (4-Arm PEG-GAS, MW 20 k) was obtained from Creative PEG Works. Transmission electron microscopy (TEM) grids and uranium acetate dihydrate were purchased from Ted Pella, Inc. Mueller Hinton Broth (MHB) and Agar was purchased from Sigma-Aldrich. Methicillin-Resistant Staphylococcus aureas (MRSA) (ATCC 33592) and Pseudomonas aeruginosa (P. aeruginosa) (ATCC 27853) were purchased from ATCC. Bacterial Live and Dead Assay kit and Live/Dead Viability/Cytotoxicity kit were purchased from Fisher Scientific.

Synthesis and Purification of Peptides.

The peptides used in this study were synthesized on the Prelude peptide synthesizer using standard solid-phase peptide synthesis (SPPS) at a 50 μmol scale, with MBHA rink amide resin as the support. Fmoc deprotection was performed by treating the resin twice with a 20% (v/v) piperidine solution in DMF to remove the Fmoc groups. For coupling, Fmoc-protected amino acids, HBTU, and DIPEA (1:1:2) were dissolved in DMF and added to the resin. This process was repeated for each amino acid coupling. After the synthesis was complete, the N-terminus was acetylated using a mixture of acetic acid and DIPEA in DMF. The peptides were cleaved from the resin using a trifluoroacetic acid (TFA)/triisopropylsilane (TIS)/H₂O (95/2.5/2.5, v/v) solution for 3 hrs. The cleavage solution was then filtered, and the resin was washed twice with neat TFA. The TFA residue was evaporated under moderate airflow, and the resulting peptide solution was precipitated with cold diethyl ether, followed by centrifugation four times. The crude peptide was dried under vacuum overnight before being purified via HPLC. Purification was carried out using a preparative reverse-phase C4 column with a linear gradient of water/acetonitrile containing 0.05% TFA. Elution was monitored at 230 and 280 nm, and the molecular mass of the purified peptide was confirmed by electrospray ionization mass spectrometry.

Hydrogel Preparation.

To prepare 100 μL of hydrogel, first, 50 μL of a 4 wt% peptide solution in water was prepared. Weighed out the required amount of PEG-GAS and dissolved it into the 50 μL of 4 wt% peptide solution. The final concentration of PEG in the mixture should be 2 wt%. Next, 50 μL of PBS 2X (Phosphate-Buffered Saline) was added to the solution to initiate the hydrogel formation process. The hydrogel was incubated at room temperature for 30 minutes to fully form before proceeding with any experimental procedures.

FT-IR Spectroscopy.

Hydrogel samples were prepared by lyophilizing 100 μL of 2 wt% hydrogel. FTIR spectra were acquired using a Thermo Nicolet 6700 FTIR spectrometer. Each spectrum was recorded as an average of 128 scans.

Circular Dichroism (CD) Spectroscopy.

Samples were prepared by diluting 2 wt% peptide or peptide-PEG gels prepared in PBS 1X, pH 7.4 to a concentration of 150 μM in 20 mM Britton-Robinson (BR) buffer at pH 7.4 and 5.5. The samples were incubated at 4 °C overnight. Data were collected from 250 nm to 190 nm at room temperature (RT) using a 1 mm cuvette, a bandwidth at 1 nm, a scan rate at 100 nm/min, and a response time of 1 sec. Each spectrum was averaged from seven scans.

Transmission Electron Microscopy.

The samples were prepared as follows: ~10 μl of the hydrogel was deposited onto a holey carbon grid. After 2 minutes, the excess solution was quickly removed using a piece of filter paper. The samples were negatively stained by placing a 10 μl of 2 wt% uranyl acetate aqueous solution on the top for 2 minutes. Excess of the uranyl acetate solution was removed using filter paper. The samples were left to dry under ambient conditions. TEM imaging of the assembled structures was performed on a Hitachi H-9500 High-resolution TEM.

Rheology Measurements.

The rheological properties of the hydrogels were characterized using oscillatory rheology with an Anton Paar MCR 702 Multidrive rheometer equipped with a 10 mm stainless steel parallel plate geometry, at a temperature of 25 °C. A 200 μL sample of the hydrogel was placed onto the lower rheometer plate, and the parallel plate gap was adjusted to 1 mm. A dynamic time sweep was first performed for 15 minutes with a frequency of 6 rad/s and a strain of 0.2%. This was followed by a dynamic frequency sweep, with the frequency ranging from 1 to 100 rad/s, while maintaining a strain of 0.2%. To evaluate the hydrogel’s recovery, it was subjected to a disruption step with a 1000% strain for 30 seconds at 6 rad/s, after which a dynamic time sweep was repeated for 15 minutes at 6 rad/s and 0.2% strain to assess the storage modulus recovery. Finally, a dynamic strain sweep was conducted on each hydrogel sample to determine the yield strain, with a strain range from 0.01% to 100% and a frequency of 6 rad/s. For samples exhibiting very low storage and loss moduli, only the 15-minute dynamic time sweep (frequency: 6 rad/s, strain: 0.2%) was performed.

Scanning Electron Microscopy (SEM).

Hydrogels were incubated in their respective pH buffers for 2 hrs, after which the buffer was removed. To preserve structural integrity, the samples were initially frozen at −80 °C and subsequently lyophilized overnight. The dried hydrogels were mounted onto carbon tape and sputter-coated with a 5 nm-thick layer of gold. Both surface and cross-sectional morphologies were analyzed using a field emission scanning electron microscope (FE-SEM) operated at an accelerating voltage of 3.0 kV and a working distance of 12.0 mm.

Release Assay.

Samples for the release assay were prepared by encapsulating 100 μM Rhodamine-labelled K₈ in the hydrogel, following the same preparation method described above. A 50 μL hydrogel sample was placed in a 96-well plate, and an equal volume of PBS 1X buffer at pH 7.4 and pH 5.5 was added on top. The release of Rhodamine-labelled K₈ was monitored by measuring UV absorbance at 560 nm at various time intervals. The measurements were conducted in triplicate for each condition.

Live and Dead Staining Assay for Bacterial Cells.

400 μL of MRSA suspension (10⁸ CFU/mL) was incubated in a confocal dish for 48 hrs at 37°C to allow biofilm formation. After incubation, the suspension was removed, and the dish was washed with PBS 1X (pH 7.4). Next, 100 μL of Mueller-Hinton broth (MHB) at pH 7.4 or 5.5 was added to the dish, followed by 50 μL of hydrogel containing 100 μM K₈, which was spread evenly across the dish. The confocal dishes were then incubated at 37°C for 2 hrs. Afterward, the MHB was removed, and the dishes were washed with PBS 1X (pH 7.4). 200 μL of a live and dead dye mix (60 μM Propidium Iodide (PI) + 4 μM SYTO 9) in PBS 1X (pH 7.4) was added, and the dish was incubated at 37°C for 30 minutes. The dye mix was then removed, and the dishes were washed with PBS 1X (pH 7.4). To fix the bacteria, 100 μL of glycerol was added. Images were captured using an epifluorescence microscope and processed with ImageJ software.

Biofilm Disruption Plating Assay for Bacterial Cells.

Method 1:

96 well plate was seeded with 200 μL of bacterial broth culture (MRSA or P. aeruginosa) at the concentration of 1 × 10⁸ CFU/mL and incubated with MHB at specific pH at 37°C for 48 hrs to facilitate biofilm formation. After incubation, the supernatant media containing planktonic cells were removed, and wells were washed thrice with 1X PBS to further remove any non-adherent cells. 50 μL of hydrogel was added, and after 30 minutes of gelation, layered with 50 μL of MES or TRIS buffer to maintain a microenvironment pH of 5.5, and 7.4, respectively. The wells were incubated for 2 hrs at 37°C, following which the treatment was removed. Finally, the cells were scraped off the wells, serially diluted in MHB at specific pH and plated. The agar plates were incubated at 37°C for 18 hrs, before colony forming units (CFU) were counted to calculate the bacterial killing efficiency percentage.

Method 2:

MRSA was cultured in Mueller Hinton Broth (MHB) until it reached the log phase growth. The final concentration of the culture was adjusted to 1 × 10⁸ CFU/ml using fresh MHB. Then, in a 96 well plate, 120 μL of culture were added to the designated wells and the peg lid was placed on to the plate to enable biofilm formation on the pegs. The plate was incubated for 48 hrs at 37°C without any shaking to allow the biofilm to develop properly. The peg lid containing MRSA biofilm was removed from initial culture plate after incubation, carefully rinsed with sterile PBS to make sure it is free from all planktonic cells. The peg lid was transferred in a new treatment well plate containing 60 μL of hydrogels layered with 50 μL of MES or TRIS buffer to maintain a microenvironment pH of 5.5, and 7.4, respectively. After 2 hrs of incubation in the hydrogel, the peg lid was taken out and rinsed gently with PBS to remove any residual hydrogel. To recover the biofilm, the peg lid was then placed into fresh 96-well plate with recovery medium at two different pH conditions, followed by sonication for 15 minutes. The suspension was collected and plated for counting and calculating of the bacterial killing efficiency. Bacterial killing efficiency was calculated using the following formulae:

$$\mathit{B}\mathit{a}\mathit{c}\mathit{t}\mathit{e}\mathit{r}\mathit{i}\mathit{a}\mathit{l}\mathit{k}\mathit{i}\mathit{l}\mathit{l}\mathit{i}\mathit{n}\mathit{g}\mathit{e}\mathit{f}\mathit{f}\mathit{i}\mathit{c}\mathit{i}\mathit{e}\mathit{n}\mathit{c}\mathit{y}(\mathrm{\%})=\frac{\frac{\mathit{C}\mathit{F}\mathit{U}}{\mathit{m}\mathit{L}}\mathit{o}\mathit{f}\mathit{c}\mathit{o}\mathit{n}\mathit{t}\mathit{r}\mathit{o}\mathit{l}-\frac{\mathit{C}\mathit{F}\mathit{U}}{\mathit{m}\mathit{L}}\mathit{o}\mathit{f}\mathit{t}\mathit{e}\mathit{s}\mathit{t}}{\frac{\mathit{C}\mathit{F}\mathit{U}}{\mathit{m}\mathit{L}}\mathit{o}\mathit{f}\mathit{c}\mathit{o}\mathit{n}\mathit{t}\mathit{r}\mathit{o}\mathit{l}}\times \mathbf{100}$$ (1)

Scanning Electron Microscopy (SEM) of Biofilm.

Bacterial solution (10⁸ CFU/mL) was added in a 24-well plate with 12 mm round cover glass fitted on the bottom of each well. After 48 hrs of incubation, bacterial solution was withdrawn from the plates and washed with PBS to remove the non-adherent bacteria, followed by the addition of 200 μL of fresh media (maintained at pH 5.5 and 7.4) to each well. For the test set up, 50 μL of hydrogel encapsulating 100μM of the antimicrobial peptide was mixed with 200μL of MHB and added to the wells. The plates were incubated at 37 °C for 2 hrs and the suspension was withdrawn. The cover glasses were washed with 1X PBS twice. The bacteria were incubated with 4% glutaraldehyde solution and left overnight for fixation. The cover glasses were further washed with 1X PBS, followed by dehydration using a series of ethanol solutions with different volume contents (35, 50, 75, 90, 95 and 100%) for 5 minutes each. The sample was placed on a carbon tape, which was further coated with a 5 nm-thick gold layer. The morphology of the bacteria with and without peptide hydrogel treatments were observed using a field emission scanning electron microscope (FE SEM) operated at an accelerating voltage of 3.0 kV and a working distance of 12.0 mm.

Mammalian Cell Culture.

Human dermal fibroblasts (HDF; ATCC, PCS-201-012) were routinely cultured in complete growth medium (COM) consisting of Dulbecco’s Modified Eagle Medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 100 IU/mL penicillin, 100 μg/mL streptomycin, and 2 mM L-glutamine. To prepare the hydrogel, 50 μL of sterile peptide precursor solution was added to a 96-well plate and allowed to polymerize. The hydrogels were balanced with 1× DPBS solution and then incubated with COM overnight at 37°C. Subsequently, HDF cells were seeded onto the hydrogel surface at a density of 1 × 10⁴ cells per well. The cell-seeded hydrogels were maintained in complete medium for 24 hrs. at 37°C under humidified conditions (95% air, 5% CO₂).

Live & Dead Staining and HDF Cell Viability Assay and Statistical Analysis.

The viability of HDF cells cultured on hydrogels was evaluated using the Live/Dead Viability/Cytotoxicity Kit (Invitrogen). Following the removal of the culture medium, the samples were rinsed twice with 1× DPBS and incubated with a staining solution containing 2 μM Calcein AM (CAM) and 4 μM Ethidium Homodimer-1 (EthD-1) in DMEM for 30 minutes at 37°C in the dark. After incubation, the samples were rinsed twice with 1× DPBS and immediately visualized under an inverted fluorescence microscope (Eclipse TiS, Nikon). Six randomly selected fields of view from 3 different samples were imaged, and the acquired images were analyzed using ImageJ software. The number of live and dead cells in each image was quantified, and cell viability was calculated as the ratio between the number of viable cells and number of total cells. Statistical analyses were performed using Origin software, with data expressed as mean ± standard deviation. Comparisons between two groups were conducted using a two-tailed Student’s T-test, with p < 0.05 considered statistically significant.

RESULTS AND DISCUSSION

The peptide sequences used in this study, listed in Table 1, were designed based on our previous work on short multi-domain peptides (MDPs) capable of self-assembling into peptide-polymer conjugate hydrogels in the presence of 4 arm PEG-GAS.¹² All peptides were synthesized using standard Fmoc solid-phase peptide synthesis (Figure S1). Successful synthesis was confirmed by electrospray ionization mass spectrometry (Figure S2), and purity was verified via high-performance liquid chromatography (HPLC), with chromatograms displaying a single major peak for each peptide (Figure S2). The peptide sequences feature an alternating hydrophilic-hydrophobic QW(QL)₃ domain (Q: glutamine, L: leucine, W: tryptophan; 3 denotes the number of QL repeating units) that promotes the formation of sandwich-like nanofiber structures, with leucines and tryptophan embedded within the hydrophobic core of the assembly. Tryptophan was included as a UV-Vis reporter to enable accurate concentration determination. Additionally, lysines were incorporated at both termini of the sequence to enhance the solubility of the peptide assemblies, and facilitate crosslinking with polymers to form extended networks, resulting in robust gels.

Table 1.

Peptide Sequences used in this study.

Name		N-	Sequence	C-
141		CH3CO	KQWQLQLQLK	CONH2
141Xp		CH3CO	KQWQLQXpQLK	CONH2
K8		CH3CO	KKKKKKKKQFQFQFQFQFQFKKKKKKKK	CONH2

Our previous study demonstrates the hydrogel formation of 141 when mixed with 4 arm PEG-GAS.¹² Hydrogel is formed through two types of interactions. First the physical interaction through self-assembly of peptide 141 leads to the formation of weak hydrogel. Second, the covalent bond formation upon reacting to the amine groups in 141 with the NHS group in the PEG connects the self-assembled nanofibers to make it a rigid robust hydrogel. To impart pH-responsive properties to the hydrogel, we incorporated a non-natural amino acid, X_p, developed by our group, by substituting it for a leucine residue in the peptide sequence (Figure S3). The Fmoc-X_p amino acid was synthesized following the method previously described in our literature and introduced into the peptide sequence via standard solid-phase peptide synthesis (SPPS).^{33, 37} Based on our previous studies, when X_p is integrated into the hydrophobic core of a self-assembled structure, the surrounding hydrophobic microenvironment plays a crucial role in stabilizing the assembly at neutral pH.^{33, 37} The unique properties of X_p allow for precise tuning of its pK_a, bringing it into the weakly acidic range. As the pH drops below the pK_a, the tertiary amine group in X_p becomes protonated. This protonation induces electrostatic repulsion, disrupting the structure and promoting disassembly of the hydrogel. Building on this mechanism, we hypothesize that introducing X_p into the peptide component of the hydrogel will allow for pH-responsive behavior. The hydrogel will remain stable at neutral pH, but as the pH decreases, the protonation of X_p will trigger disassembly, enabling the hydrogel to dynamically respond to local changes in pH (Scheme 1).

Scheme 1.

Schematic representation of the self-assembly of 141X_p into nanofibers, which undergo pH-dependent disassembly into monomers as the tertiary amine side chain (highlighted in red) becomes protonated. Upon the addition of 4-arm PEG-GAS to 141X_p, the self-assembling property is preserved, while the assembled structures are cross-linked by 4-Arm PEG-GAS. The resulting polymer-peptide hydrogel (141X_p-PEG) retains pH-dependent disassembly, guided by the peptide component (141X_p) of the hydrogel.

To test our hypothesis on the new system, we performed circular dichroism (CD) spectroscopy on 141X_p at pH 5.5 and 7.4. pH 5.5 was selected for this study because bacterial infection sites typically exhibit lower pH levels as a result of anaerobic fermentation under hypoxic conditions. Additionally, the host immune response contributes to local acidification through lactic acid production during phagocytosis. Thus, these two pH values were chosen to model infected versus uninfected environments relevant to biofilm-associated infections. The CD spectra revealed β-sheet formation, indicated by the characteristic negative peak at 215 nm, along with some random coil content, as evidenced by a peak around 195 nm, as shown in Figure 1a. At pH 5.5, complete disassembly of the nanostructures was observed. In contrast, for 141 alone, β-sheet formation was detected at both pH 7.4 and 5.5, as shown in Figure S4, suggesting that the pH-responsive behavior is influenced by the introduction of X_p into the self-assembling peptide sequence. This was further validated by transmission electron microscopy (TEM). At pH 7.4, long nanofibers were observed for 141X_p (Figure 1b), as the tertiary amines in X_p remain unchanged at this pH, allowing for stable assembly. However, at pH 5.5, complete disassembly occurred (Figure 1c) due to the protonation of the tertiary amines, which triggers disruption of the structure. For 141, long nanofibers were observed at both pH 7.4 and 5.5 (Figure S5a–b), indicating that the disassembly mechanism is specific to the presence of X_p and its pH-sensitive properties. These findings support the hypothesis that the introduction of X_p in the peptide sequence imparts pH-responsive behavior to the self-assembled structures.

Figure 1.

(a) CD spectra of 141X_p at pH 7.4, showing β-sheet content along with some random coil structure. At pH 5.5, only random coil content is observed. (b) TEM of 141X_p at pH 7.4, showing long nanofibers, indicating successful self-assembly at neutral pH. (c) TEM of 141X_p at pH 5.5, showing the absence of nanofibers, indicating pH-triggered disassembly. (d) CD spectra of 141X_p-PEG at pH 7.4, showing β-sheet formation and the absence of random coil content, suggesting a more ordered structure upon the introduction of PEG. At pH 5.5, disassembly into monomers is still observed, indicating that the pH-responsive behavior is retained in the presence of PEG. (e) TEM of 141X_p-PEG at pH 7.4, showing assembled structures in the form of nanofibers, and (f) TEM of 141X_p-PEG at pH 5.5, showing the absence of nanofibers, although some precipitates are observed, likely from the PEG component (TEM scale bar: 100 nm).

In our previous study, we found that although 141 at 2 wt % forms a weak hydrogel, incorporating PEG to form 141-PEG results in a much stronger, more rigid hydrogel due to peptide self-assembly.¹² We also observed that a 2:1 (w/w) ratio of peptide to polymer produces the strongest hydrogel.¹² The increase in hydrogel strength was attributed to the formation of covalent bonds between the amines in the peptide and the NHS groups in the PEG. This dual network, consisting of both self-assembled nanofibers and covalent bonds, significantly enhances the strength of the hydrogel. Following these results, we applied the same strategy to 141X_p, which forms a weak hydrogel at 2 wt %. By incorporating PEG into 141X_p to create the 141X_p-PEG gel, we observed a significant improvement in hydrogel strength. The combination of self-assembly and covalent bonding in the 141X_p-PEG system led to a stronger hydrogel, consistent with what we observed with 141-PEG. The formation of the conjugate was confirmed using FT-IR spectroscopy (Figure S6). The characteristic NHS carbonyl (C=O) stretch peak at 1738 cm⁻¹, present in PEG, disappeared after the reaction of the NHS group on PEG with the free amine side chains of 141X_p, indicating successful conjugation to form 141X_p-PEG. Additionally, a more defined and sharper peak corresponding to the amide I band appeared at 1622 cm⁻¹, along with a distinct peak at 1527 cm⁻¹ corresponding to the amide II band, both indicating an increase in amide bond content.

CD spectra of 141X_p-PEG were used to assess changes in secondary structure formation upon the introduction of PEG (Figure 1d). At pH 7.4, β-sheet formation was observed, as indicated by the negative peak at 215 nm. Compared to 141X_p at pH 7.4, 141X_p-PEG exhibited a higher β-sheet content, with the characteristic random coil peak for 141X_p completely disappearing in 141X_p-PEG at pH 7.4. This suggests that the addition of PEG promotes the formation of a more ordered structure, enhancing self-assembly and leading to stronger hydrogel formation. At pH 5.5, similar to 141X_p, 141X_p-PEG also underwent disassembly, indicating that the hydrogel’s assembly mechanism is primarily driven by the peptide primary sequence, even after the introduction of PEG. Along with the random coil structure, a very small amount of β-sheet content was observed at pH 5.5, suggesting that disassembly in 141X_p-PEG can still be influenced by the covalent bonds connecting the peptides, which may partially stabilize the structure even at a lower pH. In contrast, 141-PEG exhibits β-sheet structures at both pH 7.4 and 5.5, as shown in Figure S7, indicating that the assembly mechanism of 141-PEG is not pH-dependent, highlighting the unique pH-responsive behavior of 141X_p-PEG. TEM analysis of 141X_p-PEG at pH 7.4 confirms the formation of β-sheet secondary structures, with the presence of long nanofibers, indicative of successful assembly (Figure 1e). However, at pH 5.5, disassembly was observed, as evidenced by the absence of nanofibers (Figure 1f). Instead, TEM images show the presence of precipitates at this lower pH, which may result from the aggregation of disassembled conjugates. This suggests that, under acidic conditions, 141X_p-PEG undergoes a transition from ordered structures to less organized aggregates. In contrast, 141-PEG exhibited nanofibers at both pH 7.4 and 5.5 (Figure S5c–d), further confirming that the self-assembly of 141-PEG is pH-independent.

To evaluate the mechanical properties of the hydrogels, oscillatory rheological measurements were conducted. As shown in Figure 2a, a dynamic time sweep was performed for 900 seconds at an angular frequency of 6 rad/s and a strain of 0.2% for both 141-PEG and 141X_p-PEG. Both hydrogels exhibited a higher storage modulus (G’) compared to the loss modulus (G”), indicating the formation of elastic hydrogels. Furthermore, 141X_p-PEG displayed similar high storage (5508 Pa) and loss modulus (786 Pa) values compared to 141-PEG (storage modulus: 6012 Pa, loss modulus: 965 Pa). At the 900th second, a 1000% strain was applied to completely disrupt the gel, after which the strain was reduced back to 0.2% for an additional 900 seconds to measure hydrogel recovery. Both 141-PEG (~80%) and 141X_p-PEG (~85%) showed excellent recovery, suggesting high injectability of these materials. Dynamic oscillatory frequency sweep analysis of 141X_p-PEG (Figure 2b) revealed weak frequency dependence across a range of 0.1 to 100 rad/s. Dynamic strain sweep experiments from 0.01% to 100% demonstrated shear-yielding behavior at high strain, with a crossover of storage and loss moduli occurring around 19%, consistent with the behavior observed in previously studied 141-PEG gels (Figure 2c).¹²

Figure 2.

Oscillatory Rheological Measurements. (a) Dynamic time sweep of 141X_p-PEG and 141-PEG over 900 s (frequency: 6 rad/s, strain: 0.2%). At 900 s, a 1000% strain was applied to completely disrupt the hydrogel, followed by continuous recovery of the hydrogel strength. (b) Dynamic frequency sweep measurement of 141X_p-PEG (strain 0.2%), and c) strain sweep of 141-PEG (frequency 6 rad/s). (solid line: storage modulus, dash line: loss modulus).

The pH-dependent hydrogelation process was first evaluated using the tube inversion method. Peptide or peptide-PEG gels were prepared in 1X PBS at pH 7.4 and pH 5.5, and the hydrogel strength was assessed by tilting the tube. As shown in Figure S8, 141X_p at pH 7.4 forms a viscous solution, but at pH 5.5, it completely transitions into a solution, demonstrating the pH-dependent assembly-disassembly process. However, when PEG is introduced to form 141X_p-PEG at pH 7.4, there is a significant increase in hydrogel strength. Based on our previous study on the assembly mechanism of 141-PEG gels, we hypothesize that the increased strength of 141X_p-PEG follows the same mechanism: the PEG interconnects the self-assembled structure of 141X_p, enhancing rigidity.¹² At pH 5.5, although covalent bonding still exists, the unfolding of 141X_p and subsequent disassembly prevents 141X_p-PEG from forming a strong gel. In contrast, for 141, no significant difference in gel strength was observed between the two pH conditions (Figure S8e and f). When PEG is added to form 141-PEG, strong hydrogel formation is observed at both pH 5.5 and pH 7.4, with the gel at pH 7.4 exhibiting greater strength (Figure S8g and h).

The storage (G’) and loss modulus (G”) were measured using a rheometer to evaluate changes in hydrogel strength after incubation in buffer at neutral and acidic pH. Initially, G’ and G” were measured for 141X_p-PEG without incubation, and these values were compared with those of the gel incubated in equal volumes of PBS 1X at pH 7.4 and pH 5.5. For the incubated samples, the solution on top of the gel was carefully removed without disturbing the gel prior to rheological measurements. After 2 hrs of incubation, the 141X_p-PEG at pH 7.4 showed no significant reduction in strength. However, at pH 5.5, there was a noticeable decrease in hydrogel strength (Figure 3a). After 18 hrs., a slight reduction in strength was observed at pH 7.4, whereas a more significant reduction occurred at pH 5.5(Figure S9a). Figure 3b and Figure S9b show that the overall reduction in 141-PEG hydrogel strength after incubation is modest. Both pH 5.5 and pH 7.4 exhibited similar reductions in strength, with pH 5.5 showing a slightly greater decrease. These results suggest that pH has a more substantial effect on the hydrogel strength of 141X_p-PEG compared to 141-PEG.

Figure 3.

Measurement of storage and loss moduli of hydrogels over 2 hrs. (a) 141Xp-PEG incubated in 1X PBS at pH 7.4 and 5.5. (b) 141-PEG incubated in 1X PBS at pH 7.4 and pH 5.5.

The pH-responsive morphological changes of the hydrogel were investigated using scanning electron microscopy (SEM). Initially, SEM images revealed that the hydrogel surface exhibited a fibrillar structure interspersed with porous features (Figure 4a–b). Cross-sectional imaging further highlighted a layered internal architecture with well-defined pores, characteristic of its native morphology (Figure 4c–d). To evaluate the effect of acidic conditions, the hydrogel was incubated in 1X PBS at pH 5.5 for 2 hrs, followed by SEM imaging. Post-incubation surface images demonstrated notable morphological alterations, including increased pore size and a disrupted, non-uniform surface topology (Figure 4e–f). Cross-sectional views also revealed significantly enlarged internal pores, indicative of pH-dependent degradation or disassembly of the 141Xp hydrogel network under mildly acidic conditions (Figure 4g–h).

Figure 4.

Scanning electron microscopy (SEM) images of the 141Xp-PEG hydrogel prepared at pH 7.4. (a, b) Surface morphology at 100× and 500× magnifications, respectively, showing a fiber-like surface structure. (c, d) Cross-sectional views at 100× and 500× magnifications, highlighting the porous internal architecture of the hydrogel. SEM images after incubation at pH 5.5 for 2 hrs show altered morphology: (e, f) surface views at 100× and 500× magnifications reveal non-uniform surface features with enlarged pores; (g, h) cross-sectional views at 100× and 500× magnifications display significantly enlarged internal pores, indicating pH-triggered disassembly of the hydrogel network.

It has been previously observed in our reports that antimicrobial peptides (AMPs) could be incorporated into the peptide hydrogels without largely affecting its structural integrity. A cationic AMP with the sequence K₈(QF)₆K₈, features the amphiphilic structure of a conical AMP was used a model AMP as previously reported.^{12, 38} To further evaluate the pH-dependent disassembly process, we performed a release assay on rhodamine-labelled K₈ peptide encapsulated in the hydrogel. The release of rhodamine labelled K₈ into the surrounding buffer was measured by UV absorbance at 560 nm. As shown in Figure 5a, 141X_p-PEG demonstrates some initial release of rhodamine-labelled K₈, followed by a plateau in release at pH 7.4. In contrast, at pH 5.5, there is a significant increase in release over time, which could benefit from the increase of the internal pore size as observed by SEM (Figure 4h). In contrast, 141-PEG exhibits a similar release profile at both pH 7.4 and pH 5.5. There is an initial release of Rhodamine-labelled K₈, but the release eventually plateaus, with no significant difference between the two pH conditions (Figure 5b). It is noteworthy that while the storage moduli reduced for all samples upon incubation with buffers (Figure 3), significant release was observed only for 141Xp-PEG at pH 5.5. This observation seems to suggest that the interaction between K₈ and self-assembled fibers could also play an important role due to the amphiphilic nature of both K₈ and 141-Xp. Upon disassembly of the nanofiber, K₈ was released at a much faster rate. For future studies, ITC experiments will be conducted to characterize and quantify the interaction between K₈ and self-assembled 141-Xp as well as K₈ and monomeric 141-Xp under different pH conditions.

Figure 5.

(a) Release profile of 100 μM rhodamine labelled-K₈ encapsulated in 141X_p-PEG, and (b) Release profile of 100 μM rhodamine labelled-K₈ encapsulated in 141-PEG.

To further assess pH-dependent disassembly and bacterial killing, we performed a live and dead assay on MRSA biofilm. As illustrated in Scheme 2, the 141X_p-PEG hydrogel disassembles under acidic conditions due to the protonation of its tertiary amine groups. This disassembly weakens the hydrogel network, triggering the release of the encapsulated antimicrobial peptide K₈. Owing to its cationic and amphiphilic properties, K₈ can electrostatically interact with the negatively charged biofilm matrix, disrupting its structure, and promoting the release of embedded bacterial cells. Following biofilm disruption, the K₈ peptide exerts its antimicrobial activity on the liberated bacteria, effectively inhibiting their growth and further biofilm formation.

Scheme 2.

Schematic of the anti-biofilm mechanism of the anti-microbial peptide (K₈) encapsulated 141Xp-PEG hydrogels.

The biofilm was grown on a confocal dish by incubating bacteria in MHB medium for 2 days. Afterward, the hydrogel was added along with pH adjusted MHB media. After 2 hrs. of incubation, a live and dead dye mix was used to differentiate between live and dead bacteria. Dead bacteria were stained red, while live bacteria were stained green. Confocal images of 141X_p-PEG encapsulated with 100 μM of antimicrobial peptide K₈, incubated with biofilm at different pH levels, are shown in Figure 6a. At pH 7.4, we observed similar numbers of both live and dead bacteria, likely due to the diffusion of K₈ or its presence on the surface of the biofilm. However, at pH 5.5, improved bacterial killing was observed, as the gel disassembled and released the active monomer. It is worth mentioning that the minimum inhibitory concentration (MIC) was found to be 10 μM against MRSA at both pH values. The difference in antibiofilm activities of the K₈ encapsulated hydrogels is attributed to the increased release of antimicrobials at the acidic condition. The anti-biofilm activity was also confirmed upon conducting two different plating assays using the pH responsive 141X_p-PEG hydrogel. In the first method (detailed in the experimental section), hydrogels were placed on top of the biofilm and in the second method a peg lid coated with biofilms was applied on the top of the hydrogels. In both assays, samples include hydrogels with and without K₈ while a control set was taken where the hydrogel was absent against biofilms. Interestingly, the hydrogel alone also displayed a certain amount of killing efficiency though it was much lower than that of the hydrogel encapsulating K₈. Moreover, the killing efficiency was higher for the wells maintained at a lower pH of 5.5 compared to those maintained at the physiological pH of 7.4 (Figure 6b and Figure S10).

Figure 6.

(a) Live and dead assay of 100 μM K₈ encapsulated in 141X_p-PEG hydrogel at pH 5.5 and pH 7.4 (scale bar: 75 μm). (b) Bacterial killing efficiency of 141X_p-PEG hydrogel against MRSA biofilms obtained through the plating assay (Method 1).

The anti-biofilm effect of the K₈ encapsulated 141Xp-PEG hydrogel was further confirmed by SEM. As expected, robust formation of biofilm was observed at pH 7.4 for the control without hydrogel treatment (Figure S11a), while it was slightly reduced for the control maintained at pH 5.5 likely due to the acidic environment (Figure S11b). Upon treatment with the pH-responsive hydrogel at pH 5.5, bacterial cell density was dramatically reduced along with the disappearance of the dense extracellular matrices (Figure S11d). Notably, the hydrogel exhibits some antibiofilm activity but to a much lesser degree at pH 7.4 likely due to the passive release of the antimicrobial peptide from the hydrogel, or diffusion from the hydrogel surface (Figure S11c).

To evaluate the broad-spectrum antimicrobial activity of the K₈-encapsulated hydrogel, we extended our study to include the Gram-negative bacterial model P. aeruginosa. The pH-responsive hydrogel exhibited a similar trend in antimicrobial activity against P. aeruginosa as observed with MRSA (Figure S12). Specifically, the highest killing efficiency was observed at pH 5.5, consistent with increased AMP release from the hydrogel under acidic conditions compared to pH 7.4. Importantly, the K₈-encapsulated hydrogel showed significantly greater antimicrobial activity than the hydrogel without K₈. The consistent performance of the hydrogel against both Gram-positive and Gram-negative bacteria underscores its broad-spectrum antimicrobial potential.

The cytotoxicity of the hydrogel at pH 7.4 was evaluated using HDF cells through a Live/Dead staining assay. The results indicated that most cells remained viable after 24 hrs. of incubation with the hydrogels (Figure 7a). However, a higher number of cells adhered to the tissue culture plate (TCP) compared to the hydrogel. This difference in cell attachment may be attributed to the relatively hydrophobic nature of the hydrogel surface, which could affect initial cell adhesion. Nevertheless, qualitative analysis demonstrated that the hydrogel exhibited good cytocompatibility (Figure 7b), with an average cell viability of approximately 97%.

Figure 7.

(a) Live/dead staining images of HDF cells seeded on the TCP, and 141X_p-PEG encapsulated with 100 μM K₈ for 24 hrs. Live cells are stained green with calcein AM, and dead cells are stained red with ethidium homodimer-1. (b) Cell viability of HDF cells incubated on TCP and 141Xp-PEG encapsulated with 100 μM K₈ for 24 hrs.

CONCLUSIONS

We have demonstrated the synthesis of a pH-responsive double network hydrogel based on the self-assembly of peptide-PEG conjugates. The peptide contains a newly designed non-natural ionic amino acid for tuning the transition pH of peptides in the weakly acidic range. While these peptides have been studied in solutions in the past, the current work expanded their utility for the construction of bulk hydrogel materials. Notably, the coupling of pH-responsiveness peptide self-assembly with covalent synthetic polymers allows dynamic control over assembly, structure, and rheological properties. By leveraging the unique properties of antimicrobial peptides, self-assembling nanostructures, and pH-triggered release mechanisms, this innovative hydrogel approach may offer a more targeted, effective, and safer treatment option for patients with biofilm-associated infections. We hope that this research will lead to the development of more advanced biomaterials that can support wound healing while combating bacterial infections in a more precise and efficient manner.

Supplementary Material

ASSOCIATED CONTENT

Supporting Information.

Characterizations of the chemical compositions and physical properties of PEG-peptide conjugates. This material is available free of charge via the Internet at http://pubs.acs.org.