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. 2025 Oct 24;5(6):900–906. doi: 10.1021/acspolymersau.5c00095

Bactericidal Activities of Copolymers Incorporating Formylphenyl Motif for Targeting Surface Proteins

Esteban Bautista †, Melody Sun †, Michael Colwin ‡,§, Albert R La Spada ‡,§, Seunghyun Sim †,∥,⊥,*
PMCID: PMC12874151  PMID: 41657378

Abstract

We developed a set of intrinsic antimicrobial copolymers equipped with a dual-binding mechanism for the bacterial cell surface, targeting both the anionic bacterial membrane through electrostatic interactions and surface proteins through reversible imine formation. These copolymers, containing formylphenyl and quaternary ammonium functional groups, were systematically evaluated for their biological activity as the chemical composition and architecture were varied. These bactericidal polymers exhibit potent antimicrobial activity in NB media against both Staphylococcus epidermidis and Escherichia coli, while also demonstrating excellent in vitro biocompatibility against mammalian and red blood cells. This work expands the chemical repertoire of intrinsic antimicrobial polymers that coalesce with bacterial matter.

Keywords: antimicrobial polymers, AMR, copolymers, formylphenyl motif, quaternary ammonium

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Introduction

The overconsumption and poor stewardship of antibiotics have led to the emergence of a class of bacteria with antimicrobial resistance (AMR) properties. , With bacterial AMR, infections may have no effective treatment options available, making previously manageable minor infections or wounds potentially fatal conditions. Bacterial AMR is expected to become the next great epidemic, and it is likely that by 2050, 8 million annual deaths will be attributed to AMR. It is imperative that we expand the scope of the antimicrobial agents. Many studies have reported the development of synthetic macromolecules with intrinsic antimicrobial activity. − While small-molecule antibiotics target specific bacterial processes, nonspecific mechanisms of antimicrobial polymers (e.g., membrane disruption) can exhibit broad-spectrum activity. Inspired by chemical structures of host-defense peptides, , this is typically achieved by combining cationic residues that can adhere to anionic bacterial surfaces with hydrophobic moieties responsible for disrupting the bacterial lipid membrane. − The effect of cationic and hydrophobic groups, as well as their relative ratio, sequence, and topology, has been investigated to enrich the repertoire of antimicrobial polymers. − In addition, a number of studies have investigated the potential of incorporating active targeting mechanisms, such as sugar-lectin binding or hydrogen bonding. − Here, we ask: Can we combine an additional targeting moiety for surface proteins in the design of antimicrobial polymers? Surface proteins in pathogenic bacteria are responsible for host–pathogen interactions that result in bacterial virulence, − executing a variety of functions, including nutrient acquisition for iron extraction, adherence, invasion of host tissue, and evasion of immune responses. ,

In this article, we report the design, synthesis, and evaluation of a set of antimicrobial polymers that incorporate aldehydes to form dynamic covalent bonds with bacterial surface proteins (Figure A). In our previous work, we examined a set of intrinsic bactericidal polymers that feature an orthogonal dual-binding mechanism, enabling the formation of electrostatic interactions and reversible boronate esters with glycans on the bacterial surface through the incorporation of quaternary ammonium and phenylboronic acid residues, respectively. Building on our previous work, we envisioned a new set of antimicrobial agents that target the negatively charged bacterial membrane and surface proteins. Quaternary ammonium residues served as anion scavengers on the bacterial surface due to their permanent cationic charge, a very successful strategy in designing antimicrobial compounds. , Aromatic aldehydes have been shown to react with available primary amine groups in proteins, forming an imine. , While imine formation is a well-known example of dynamic covalent chemistry and has been used for bioconjugation, to the best of our knowledge, it has not been explored for designing intrinsically antimicrobial polymers. We chose benzaldehyde as an electrophile to target the nucleophilic primary amines in surface proteins as well as the general hydrophobic motif.

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Polymer structure and the effect of composition on the minimum inhibitory concentration (MIC) against Gram-positive and Gram-negative bacteria. (A) The copolymer structure in this study comprises N-(3-formylphenyl)­acrylamide (FPAA) and (3-acrylamidopropyl)-trimethylammonium (APTAC). Primary lysines on surface proteins are targeted by FPAA (m) residues to form dynamic covalent bonds. Quaternary ammonium groups in APTAC (n) function as anionic scavengers on the bacterial surface through electrostatic interactions. (B–E) Copolymers and homopolymers of APTAC were tested against model Gram-positive and Gram-negative strains, Staphylococcus epidermidis (ATCC 12228) and Escherichia coli (DH10B), respectively, in NB media. MIC values for (B) S. epidermidis and (C) E. coli are plotted as a function of n + m. MIC values for (D) S. epidermidis and (E) E. coli are plotted as a function of the percentage of FPAA (% FPAA). The MIC was defined as the lowest concentration that completely inhibited bacterial growth. Experiments were conducted in a biological quadruplicate (N = 4). Error bars correspond to the standard error of the mean (SEM).

Results and Discussion

The Compositional Effect of Formylphenyl Group on Antibacterial Activity

To uncover the structure–property relationship of copolymers bearing N-(3-formylphenyl)­acrylamide (FPAA) and (3-acrylamidopropyl)-trimethylammonium (APTAC) residues, linear polymers of different architectures, homopolymers, block copolymers, and gradient copolymers, were synthesized and characterized (Figures A and S1–S20, Table S1). By comparing these polymers, we aimed to isolate the effect of individual side chains by varying chemical compositions and to understand the impact of polymer architecture. A polymer library consisting of APTAC and FPAA was synthesized via reversible addition–fragmentation chain transfer (RAFT) polymerization, using 2-(dodecylthiocarbonothioylthio)-2-methylpropionic acid (DDMAT) as the chain transfer agent. FPAA monomer was synthesized using an established protocol (Figure S4). Number-average molecular weight (M n) was determined via 1H NMR by end-group analysis, while weight-average molecular weight (M w) was determined by diffusion-ordered spectroscopy (DOSY) NMR. , The molecular weight distribution of Polymer F, due to its limited water solubility, was characterized via matrix-assisted laser desorption ionization time-of-flight mass spectrometry (MALDI-TOF MS). The block architecture was verified by comparing the size of the first block and the final product by either DOSY or MALDI-TOF MS (Table S1). Kinetic analysis based on 1H NMR indicated preferential incorporation of FPAA over APTAC, resulting in a gradient architecture of copolymers (Figure S3). The reactivity of the phenylformyl moiety on FPAA toward amine residues in aqueous solution was confirmed through small-molecule model studies using 1H NMR (Figure S21). The complete list of polymers in the library is presented in Table S1.

Polymers of varying chemical compositions were evaluated for their antimicrobial activity via minimum inhibitory concentration (MIC) studies against a model Gram-positive bacterium, S. epidermidis (ATCC 12228), and a Gram-negative bacterium, E. coli (DH10B), in NB media. S. epidermidis is a commensal organism in human skin and can become an opportunistic pathogen when the host’s immune system or physical barrier has been compromised, which makes it a good target for antimicrobial polymers. Similarly, E. coli is a major pathogen associated with urinary tract infections. We used well-characterized strains of these representative bacterial species for our study. Antibacterial activities were examined as a function of the total number of repeating units (n + m) and the percentage of FPAA (% FPAA) based on NMR analysis (Figure ). The total number of repeating units (n + m), which also corresponds to polymer molecular weights, did not exhibit a clear trend in the observed MICs for both S. epidermidis and E. coli (Figure B,C). Across both Gram stains, as the percentage of FPAA increased, higher MIC values were observed, indicating that increasing APTAC residues have a positive effect on antimicrobial activity (Figure D,E and Table S2). MICs of the polymers without FPAA (0% FPAA, A1–A4 in Tables S1 and S2) were approximately 2–4 μg/mL for S. epidermidis and 4–8 μg/mL for E. coli. These results suggest that incorporating formylphenyl groups into antimicrobial polymers reduces the antimicrobial activity. This observation is also in agreement with our previous work, which involved antimicrobial polymers comprising quaternary ammonium and phenylboronic acid residues.

Effect of Polymer Architecture on Self-Assembly and Antibacterial Activity

Having established the effect of copolymer composition on antibacterial activity, we sought to understand the effect of the polymer architecture on antibacterial activity. We selected copolymers with comparable molecular weights and different polymer architectures: gradient copolymer (G) and diblock copolymer (AF) (Figure A) for detailed comparison. These polymers exhibited identical MIC values of 8 and 16 μg/mL against S. epidermidis and E. coli in NB, respectively (Figure B). We note that they exhibited an increased MIC in caMHB media (Table S3). Despite their architectural difference, their identical MIC in NB media motivated us to investigate whether there are any nanoscale differences between these two polymers in this chemical environment. Transmission electron microscopy (TEM) of polymers AF and G showed that G assembles into spherical nanoparticles, whereas AF forms anisotropic nanoscale aggregates (Figure C,D). Dynamic light scattering (DLS) measurements revealed the average hydrodynamic radii (R h) for self-assembled antimicrobial polymers AF and G at their respective MIC in the range of hundreds of nanometers (Figures E and S22). G showed a z-average of 224.7 nm with a narrow distribution and a polydispersity index (PDI) of 0.107, whereas AF showed a z-average of 557.6 nm with a PDI of 0.235. Incorporation of FPAA, regardless of the gradient or block copolymer architecture, reduces antimicrobial activity compared to APTAC homopolymers (Table S2), which possess only a single binding mechanism. To assess the colloidal stability of the nanoparticles, ζ-potential measurements were conducted at a concentration of 16 μg/mL. Polymers G and AF both showed positive average ζ-potential values of approximately +32 and +63 mV, respectively, due to the cationic quaternary ammonium residues (Figure F). Considering that ζ-potential measurements for APTAC homopolymer are around +11 mV, we speculate that incorporation of hydrophobic FPAA increases the colloidal stability of the nanostructures and attenuates antibacterial activity compared to APTAC homopolymer. Interestingly, while these two polymers did not show any difference in MIC, G seemed to be slightly more potent than AF in terms of the minimum bactericidal concentrations (MBC) (Figure B), indicating the general mode of action of these copolymers is bactericidal agents (i.e., causing cell death).

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Effect of copolymer architecture on self-assembly and antimicrobial properties. (A) Molecular weights of the selected copolymers with similar compositions. The degrees of polymerization (n and m) and M n were determined via 1H NMR end-group analysis. M w was determined via a DOSY NMR. (B) Comparison of minimum inhibitory concentration (MIC) and minimum bactericidal concentration (MBC) among the selected polymers with similar compositions and different polymer architectures in NB. Antimicrobial polymers were tested against S. epidermidis (ATCC 12228) and E. coli (DH10B). (C) Representative transmission electron microscopy (TEM) image of polymer G. (D) Representative TEM image of polymer AF. (E) Dynamic light scattering (DLS) of antimicrobial polymers in NB media. (F) ζ-Potential measurements of G and AF in H2O. TEM and measurements for DLS and ζ-potential were performed at 16 μg/mL. Scale bars: 0.5 μm.

Mechanical Damage to Bacterial Cells by Copolymers Incorporating Formylphenyl Groups

When both bacteria were treated with polymer G at MIC, spherical structures were observed under an optical microscope, regardless of the original shape of the bacteria (spherical S. epidermidis vs rod-shaped E. coli) (Figure A–F). While S. epidermidis and self-assembled polymer G alone are both spherical, the resulting spherical microstructures showed a broader distribution of sizes compared to cells alone (Figure A,B). The rod-shaped morphology of E. coli completely disappeared upon treating polymer G (Figure D,E). On the other hand, cells treated with polymer AF formed anisotropic, fibril-like structures on a micrometer scale across both Gram-positive and Gram-negative cells, regardless of their original shape. Both bacteria completely lost their original structure and dimensions when treated with polymer AF (Figure A,F). These structures far exceed the sizes of the polymer nanoparticles (Figure B–D). This is in contrast to the homopolymer of APTAC from our previous study, which did not coalesce with bacterial matter. Taken together, copolymer nanoparticles incorporating formylphenyl groups interact with bacterial surfaces, and the subsequent assembly of synthetic polymers and bacteria is the physical cause of its bactericidal mode of action. Notably, the resulting structure, following bactericidal activity (Figure B,F), generally mirrors the self-assembled nanostructures of each polymer alone (Figure C,D).

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Physical lysis of bacteria upon treatment with copolymers containing formylphenyl groups. (A–F) Optical micrographs were taken after treating S. epidermidis and E. coli with polymer solutions at their respective MICs in NB for 16 h at 37 °C and 250 rpm. (A) S. epidermidis without polymer treatment. (B) S. epidermidis incubated with G (8 μg/mL). (C) S. epidermidis incubated with AF (8 μg/mL). (D) E. coli without polymer treatment. (E) E. coli incubated with G (16 μg/mL). (H) E. coli incubated with AF (16 μg/mL). Scale bars = 10 μm. (G–L) Scanning electron microscope (SEM) micrographs taken at 0.5 MIC to observe the intermediate structures and deduce the mechanism of action of the copolymers. (G) S. epidermidis without polymer treatment. (H) S. epidermidis incubated with polymer G (4 μg/mL). (I) S. epidermidis incubated with polymer AF (4 μg/mL). (J) E. coli without polymer treatment. (K) E. coli incubated with polymer G (8 μg/mL). (L) E. coli incubated with polymer AF (8 μg/mL). Scale bars are all 1 μm, except for (L), which is 5 μm.

To gain additional structural insight into the observed structures in our optical microscopy studies, scanning electron microscopy (SEM) was employed (Figure G–L). SEM micrographs were taken at half the MIC to observe intermediate structures, as the typical cellular structure becomes invisible when bacteria are treated with polymers at their respective MIC. For the cells treated with polymer G, we observed the deformation of cellular structures and pore formation on the cell surface (Figure H,K), indicating that the mechanical damage caused by the antibacterial polymer results in physical lysis of the bacteria. Cellular deformation and anisotropic aggregates (Figure I,L), including ribbon-like microstructures in E. coli (Figure L), were observed in the cells treated with polymer AF. Altogether, the amphiphilic nature of the copolymers and their tendency to self-assemble drive mechanical disintegration of the bacterial structure and subsequent coalescence with cellular debris.

Biocompatibility of Bactericidal Copolymers Incorporating Formylphenyl Groups

Antibacterial polymers were evaluated for their cytocompatibility in model mammalian cells via cytotoxicity studies against human embryonic kidney (HEK 293). Polymers AF and G are generally not toxic to HEK 293 at MICs or MBCs (Figure A,B). In particular, polymer AF demonstrated cell viability above 90% across all experimental concentration ranges against HEK 293, indicating excellent cytocompatibility (Figure A). Gradient polymer G was found to be moderately cytotoxic at 512 μg/mL, a 32-fold difference from its observed bacterial MIC, with an average viability of 55% in HEK 293 cells (Figure B). It is noteworthy that even with the presence of anionic charges and surface proteins on the mammalian cell surface, the copolymers AF and G do not exert similar effects, namely, lysis and coalescence, as they do for bacterial cells.

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Biocompatibility of antibacterial copolymers. (A, B) Cytotoxicity study with HEK cells treated with (A) AF and (B) G polymer. Optical micrographs of RBCs (C) without polymers and upon exposure to polymers (D) AF and (E) G. Micrographs were taken at their respective C H for each polymer. Scale bars = 40 μm. (F) Summary table of biocompatibility results based on cytotoxic, hemolytic, and hemagglutination activities of antibacterial polymers. Cytotoxicity (IC50) is the minimum concentration of antibacterial polymers needed for 50% cell viability of HEK 293 cells. Hemolysis (IC10) is the minimum concentration of antibacterial polymers needed for the 10% lysis of RBCs. Hemagglutination (C H) is determined by the minimum concentration of antibacterial polymers needed for 10% aggregation of RBCs.

We also performed hemocompatibility studies including hemolysis and hemagglutination assays against red blood cells (RBCs). These polymers are generally not hemolytic across all concentrations tested (Figure S23). Hemagglutination assays revealed that both polymers AF and G aggregated RBCs at concentrations below their MIC range (Figure C–F). Since the homopolymer of APTAC did not exhibit this behavior, formylphenyl groups should be responsible for this hemagglutination phenomenon. We speculate that surface-exposed proteins in RBCs interact with formylphenyl handles on these copolymers, driving the aggregation. Interestingly, although polymers G and AF aggregate RBCs, they do not cause hemolytic activity. The toxic effect of polymers AF and G against bacteria over mammalian cells can be attributed to the fact that bacteria have a greater negative surface potential than mammalian cells. , Permanently charged quaternary ammonium may interact more strongly with the bacterial cell surface than with mammalian cells.

Conclusions

In this work, we examined the utility of FPAA as a hydrophobic and surface protein binding motif in antimicrobial polymers. Our work systematically investigated on cationic copolymers, examining both composition and architecture. From the MIC analysis of a polymer library with varying compositions, we found that increasing the portion of FPAA in copolymers leads to decreased antimicrobial activity. At the expense of a slight decrease in the antibacterial activity of cationic polymers, incorporating FPAA into polymer chains increases the tendency for polymers to self-assemble and results in micrometer-scale aggregates coalescing with bacterial matter. Our work also establishes that these copolymers are bactericidal by physically lysing cells. The preclinical screening of these novel polymeric antibiotics revealed that they possess excellent biocompatibility and exhibit the tendency to aggregate RBCs without lysing them, hinting at their potential application as bioadhesives for infected wounds.

Methods

DOSY (Diffusion-Ordered Spectroscopy) NMR

Weight-averaged molecular weight (M w) was calculated using an established protocol. Briefly, polymer stock solutions were prepared in D2O at a 1 mg/mL concentration to obtain diffusion coefficients. Once obtained, M w was calculated as follows: log­(D) + log­(η) = log­(c) + νlog­(M). D = diffusion coefficient, η = viscosity of D2O, c = first calibration coefficient for PEG in D2O, and ν = second calibration coefficient for PEG in D2O.

MALDI-TOF MS (Matrix-Assisted Laser Desorption/Ionization Time-of-Flight Mass Spectrometry)

MALDI-TOF/TOF MS analysis was performed using an established protocol. Briefly, polymer stock solutions were prepared in dimethyl sulfoxide (DMSO) (1 mg/mL) and then serially diluted with a-cyano-4-hydroxycinnamic acid. Spectra were collected using the reflected positive ion mode, and peaks were analyzed by using Data Explorer software.

Dynamic Light Scattering (DLS)

Polymer stock solutions were prepared in NB media, and aliquots of the stock solutions were diluted to the desired concentrations. Polymer solutions were then placed on a heating block (37 °C and 250 rpm) overnight. DLS measurements were taken the next day at room temperature with a 90° scattering detector angle for the size measurements.

Minimum Inhibitory Concentration (MIC)

Antibacterial polymers were evaluated for their antimicrobial activity using established protocols. Briefly, stock solutions of the antimicrobial polymers were created in NB media and then diluted in 96-well plates with 2-fold dilutions from 512 to 1 μg/mL. Bacteria were grown (37 °C, 250 rpm) to a 1 × 108 CFU/mL to match the McFarland 0.5 standard. Bacterial suspensions were diluted 100-fold, and then 50 μL of the bacterial suspensions were added to the 96-well plate, creating a final inoculum size of 5 × 105 CFU/mL. Bacterial growth was determined after 16 h of incubation at 37 °C by optical density (OD600) using a BioTek Synergy H1 Hybrid Multi-Mode microplate reader. The MIC was defined as the lowest concentration that completely inhibited bacterial growth (no increase in the OD). Each experiment was conducted in biological quadruplicate per combination of bacterial strain, polymer, and concentration.

Minimum Bactericidal Concentration (MBC)

Immediately after the MIC assay in NB media, polymer-cell samples at the MIC, as well as at subsequent higher concentrations up to eight times the MIC, were removed from the 96-well plate. Each sample was diluted 100,000-fold, and then 100 μL was streaked onto an LB agar plate. After overnight incubation at 37 °C, the number of cell-forming units (CFU) on the plate was counted and quantified as CFU/mL. The MBC was defined as the minimum concentration of polymer at or above the MIC that kills 99.9% of bacteria on the agar plate (no colony formation). Each experiment was performed in triplicate per combination of the bacterial strain, polymer, and concentration.

ζ-Potential

Polymer stock solutions were prepared in doubly distilled H2O, and aliquots of the stock solutions were diluted to the desired concentrations, the highest MIC observed across all bacterial cell lines. Samples were then placed on a heating block overnight (37 °C at 250 rpm), and ζ-potential measurements were performed at room temperature in triplicate.

TEM

Before sample deposition, TEM grids were treated for glow discharge to hydrophilize the support film for ∼90 s. Next, 5 μL of the sample was deposited onto the TEM grid and was allowed to sit for 10 min. The grid was washed with 200 μL of doubly distilled H2O and then blotted dry with filter paper. Once dried, 5 μL of 2% uranyl acetate was deposited onto the grid for 15 s and then blotted dry with filter paper. TEM imaging was performed using a JEOL JEM-2100F TEM equipped with a Schottky-type field emission electron source at an accelerating voltage of 200 kV. Images were taken using a Gatan Oneview camera.

SEM

Interactions between antibacterial polymers and bacteria were studied using an established protocol. SEM micrographs of bacterial samples were taken at their respective 0.5 MIC after completion of the MIC assay (see the Minimum Inhibitory Concentration (MIC) section for details). Briefly, MIC assays were conducted in biological quadruplicates per combination of bacterial strain, polymer, and concentration for 16 h at 37 °C in NB media. Bacterial suspensions had a final inoculum size of 5 × 105 CFU/mL. Samples were concentrated at 5000 rpm for 10 min. The medium was discarded, and the cell pellet was resuspended in 4.0% glutaraldehyde solution (in 1× PBS) for 1 h. Next, a glass cover slide was precoated with poly-d-lysine (50 μg/mL in H2O) for 15 min, and the excess was removed with doubly distilled H2O. After the sample was fixed for 1 h, 30 μL was deposited onto the glass slide for 15 min and then washed with doubly distilled H2O. The sample/glass slide was treated with ethanol solutions (30, 50, 70, 90, and 100% in doubly distilled H2O) for 10 min each. The samples were air-dried and sputter-coated with iridium. SEM was performed on a Tescan GAIA3 SEM-FIB instrument equipped with a field emission gun at an operating voltage of 4 kV and an in-beam SE detector.

Optical Microscopy

A 2% low-melt agarose solution was prepared in 1 L of water. The solution was microwaved until it was completely dissolved. On a glass slide, a Grace Bio-Laboratories Press-To-Seal silicone isolator (25 mm × 25 mm) was placed with ∼1 mL of the molten agarose deposited. A second glass slide was quickly placed on top, sandwiching the agarose pad between the two glass slides. After 30 min, the second glass slide and isolator were carefully removed without tearing the agarose pad. Next, 2 μL of the bacterial sample was added, followed by a glass coverslip. Samples were then imaged on an RVL-100-M model ECHO Revolve microscope at 60× magnification with a phase-contrast objective. Optical micrographs of bacterial samples were taken at their respective MIC values after completion of the MIC assay. Briefly, MIC assays were conducted in biological quadruplicates per combination of bacterial strain, polymer, and concentration for 16 h at 37 °C in NB media. Bacterial suspensions had a final inoculum size of 5 × 105 CFU/mL.

Cytotoxicity Assay

Cytotoxic activities of antibacterial polymers were evaluated using established protocols. Briefly, HEK 293 cells were plated in a 96-well plate at a density of 4000 cells/well and left to settle overnight. Cells were cultured in DMEM + GlutaMAX. The following day, the cells were treated with varying concentrations of the polymer dissolved in the media, with blank media being used as a negative control. After 24 h of treatment with the polymer, 10 μL of Presto Blue was added to the cells and the cells were left to incubate for 3 h. Absorbance readings at 570 nm were taken for the plate after that time using a Varioskan LUX Multimode Microplate Reader. Cell viability was expressed as a percentage relative to the cells that were treated with the blank media.

Hemolysis

Hemolytic activity of antibacterial polymers was evaluated using established protocols. Defibrinated sheep blood was centrifuged at 4500g for 1 min with subsequent washes with 1 × PBS until the supernatant of the blood was clear, at which point the RBCs were resuspended at 6% (v/v). 100 μL of the RBCs was added to each well of a 96-well plate. 100 μL of the polymer solution was added to each well to reach the desired concentration. 1× PBS was used as a negative control for hemolysis, while 1% Triton X-100 was used as a positive hemolytic control. After the polymer was added to the RBCs, the plate was incubated at 37 °C for 2 h, after which time the plate was centrifuged at 600g for 10 min. 100 μL of the supernatant was carefully withdrawn and transferred to a new plate. Absorbance readings at 540 nm were measured using a Varioskan LUX Multimode Microplate Reader. Readings were normalized to the Triton X-100 group, which was considered to be 100% hemolysis. The percentage of hemolysis was calculated as follows: % Hemolysis = [(sample absorbance – negative control)/(positive control – negative control)] × 100%.

Hemagglutination

Hemagglutination assays were performed using established protocols. Sheep RBCs were prepared as described above. 50 μL of the 6% (v/v) RBCs were added to each test well of a 96-well U-bottom plate, followed by 50 μL of polymer at the specified concentration. 0.05 mg/mL Concanavalin A was used as a positive control, and PBS was used as a negative control. The plates were incubated at 37 °C for 1 h, after which time they were visually assessed for hemagglutination. Representative microscopy images of several conditions were also taken to demonstrate the hemagglutination.

Supplementary Material

lg5c00095_si_001.pdf (2.5MB, pdf)

Acknowledgments

This work was supported by the UC Irvine start-up fund and by the National Institutes of Health under Award Numbers R35GM150770 and R35NS122140. The content is solely the responsibility of the authors and does not necessarily represent the official views of the National Institutes of Health. E.B. acknowledges support from the National Science Foundation Graduate Research Fellowship Program (DGE-1839285). M.S. was supported by UROP fellowship from UC Irvine. The authors acknowledge the use of facilities and instrumentation at the UC Irvine Materials Research Institute (IMRI) supported in part by the National Science Foundation Materials Research Science and Engineering Center program through the UC Irvine Center for Complex and Active Materials (DMR-2011967). Dynamic light scattering and ζ-potential measurements were performed at the Laser Spectroscopy Laboratories. Mass spectroscopy and the nuclear magnetic resonance measurements were done in the NMR and MS facility, both in the Department of Chemistry, University of California, Irvine.

The Supporting Information is available free of charge at https://pubs.acs.org/doi/10.1021/acspolymersau.5c00095.

  • Materials, instrumentation, experimental and synthetic methods, 1H and DOSY NMR spectra, and hemolysis assay (PDF)

The manuscript was written through contributions of all authors, and all authors have given approval to the final version of the manuscript.

The authors declare no competing financial interest.

Published as part of ACS Polymers Au special issue “2025 Rising Stars in Polymers”.

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