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Published in final edited form as: ACS Infect Dis. 2023 Oct 13;9(11):2316–2324. doi: 10.1021/acsinfecdis.3c00415

Nitric Oxide-Induced Morphological Changes to Bacteria

Huan K Nguyen 1, Samantha L Picciotti 1, Magdalena M Duke 1, Christopher A Broberg 1, Mark H Schoenfisch 1,2,*
PMCID: PMC11041245  NIHMSID: NIHMS1984780  PMID: 37831756

Abstract

Antimicrobial resistance (AMR) poses a serious threat to global health, necessitating research for alternative approaches for treating infections. Nitric oxide (NO) is an endogenously produced molecule involved in multiple physiological processes, including response to pathogens. Herein, we employed microscopy and fluorescence-based techniques to investigate the effects of NO delivered from exogenous NO donors on the bacterial cell envelopes of pathogens, including resistant strains. Our goal was to assess the role of NO donor architecture (small molecules, oligosaccharides, dendrimers) on bacterial wall degradation to representative Gram-negative bacteria (Klebsiella pneumoniae, Pseudomonas aeruginosa) and Gram-positive bacteria (Staphylococcus aureus, Enterococcus faecium) upon treatment. Depending on the NO donor, bactericidal NO doses spanned 1.5 to 5.5 mM (total NO released). Transmission electron microscopy (TEM) of bacteria following NO exposure indicated extensive membrane damage to Gram-negative bacteria with warping of cellular shape and disruption of cell wall. Among the small molecule NO donors, those providing a more extended release (t1/2 = 120 min) resulted in greater damage to Gram-negative bacteria. In contrast, rapid NO release (t1/2 = 24 min) neither altered the morphology nor roughness of these bacteria. For Gram-positive bacteria, NO treatments did not result in any drastic change to cellular shape or membrane integrity, despite permeation of the cell wall as measured by depolarization assays. The use of positively charged quaternary ammonium (QA)-modified NO-releasing dendrimer proved to be the only NO donor system capable of penetrating the thick peptidoglycan layer of Gram-positive bacteria.

Graphical Abstract

graphic file with name nihms-1984780-f0001.jpg

The graphical abstract includes electron micrographs and zoomed-in graphics, and depicts the effect of nitric oxide on Gram-negative and Gram-positive bacteria. This manuscript contributes to the understanding of using nitric oxide to treat bacterial infections.

Antimicrobial resistance (AMR) is a critical issue in modern medicine. A 2022 global collaboration surveying 204 countries and territories estimated that 1.27 million deaths were attributed to AMR in 2019 (worldwide), exceeding the combined global fatal cases of HIV (524,000) and malaria (558,000) in the same year.1–4 Antibiotic resistance is a major subset of AMR. The emergence and propagation of multidrug-resistant (MDR) bacteria pose a serious threat to human, animal, and environmental health. In 2017, a report funded by the World Health Organization (WHO) identified 20 bacteria species with strong patterns of resistance acquisition.5 Among these bacteria, a group of six pathogens (Enterococcus faecium, Staphylococcus aureus, Klebsiella pneumoniae, Acinetobacter baumannii, Pseudomonas aeruginosa, and Enterobacter spp) were designated critical or high priority for research and development of new effective antibiotics. These bacteria are collectively known as the ESKAPE pathogens.

ESKAPE pathogens include both Gram-positive and Gram-negative bacteria which differ in their bacterial cell envelopes. Gram-negative bacteria possess a three-layered cell envelope, consisting of lipid bilayer-based plasma membrane, a thin (2–3 nm) layer of peptidoglycan (PG) and an outer membrane. The outer membrane of Gram-negative bacteria is comprised of a lipid-based inner leaflet and an outer leaflet of lipopolysaccharides (LPS) that provides both structural support and protection against cells and hydrophobic molecules such as proteins. The cell envelope of Gram-positive bacteria has only two inner layers (i.e., no outer membrane). However, they possess a thick PG layer (up to 100 nm) that surrounds the plasma membrane.6 As might be expected, antibiotics interact and have different activity against Gram-negative versus Gram-positive bacteria. Narrow-spectrum antibiotics often target one specific metabolic process (e.g., PG synthesis or DNA replication). In this regard, they are effective against only one class of bacteria.7 Despite the need for next generation antibiotics to address antimicrobial resistance, the approval of such pharmaceuticals has declined due to prohibitive development costs and regulatory obstacles.8 The mounting challenges of using antibiotics to treat bacterial infections necessitates research on antimicrobial agents that offer new mechanisms of action and do not foster resistance.

Nitric oxide is an endogenously produced radical that plays an integral role in the innate immune response to foreign pathogens by eliciting both oxidative and nitrosative stress that ultimately damage the bacterial cell envelope and metabolism/viability.9,10 In physiological milieu, NO undergoes oxidation to produce reactive nitrogen species such as dinitrogen oxide that reacts with surface proteins on the bacterial outer membrane. As a result of thiol and amine nitrosation, these proteins lose vital signaling or transport functions.11–13 Additionally, NO reacts with superoxide (O2−) to create peroxynitrite (ONOO−) which disrupts the integrity of the phospholipid membranes via peroxidation.14 After traversing the cell envelope, NO may cause further damage via DNA deamination, inhibition of DNA repair processes and/or production of other genotoxic agents.15,16 As the antibacterial activity from NO is the result of multiple mechanisms of action, antimicrobial resistance is projected to be less likely than that for conventional antibiotics. To date, serial passage assays with sub-microbiocidal levels of NO donors have shown no resistance to NO for representative Gram-negative and Gram-positive bacteria, and Candida auris.17

Nitric oxide’s ability to destroy bacteria without fostering resistance makes the development of NO-based therapeutics an attractive goal in the era of antimicrobial resistance. However, clinical application of NO has been limited to inhalation of NO gas to treat pulmonary hypertension, acute respiratory distress syndrome, and ischemia/reperfusion injury.18–20 The short half-life of NO gas (a few seconds) and the requirement of specialized equipment (e.g., gas cylinders) hinder the use of NO for antibacterial applications.21 As an alternative, chemical NO donors that spontaneously release NO have become desirable systems for generating localized NO to study NO-related effects to pathogens. Such NO donors have been successfully synthesized using a variety of molecular architectures, including nanoparticles, dendrimers, coatings, and biopolymers.22–26 Further chemical modifications of the NO donors have been used to make these systems even more active against bacteria (e.g., use of antibacterial quaternary ammonium groups to compliment the bactericidal effect of NO).27,28

While prior studies have demonstrated the bactericidal effects of NO-releasing donors, study of the effects of NO on the cell envelope of different bacterial classes as a function of NO donor is scarce. Deupree et al. studied the surface morphology of adsorbed P. aeruginosa onto an NO-releasing polymer coating.29 Of note, these studies were limited to Gram-negative bacteria and carried out in the context of bacterial adhesion to abiotic surfaces. Given the structural uniqueness between Gram-negative and Gram-positive bacteria, we hypothesized that NO may impact their cell envelope integrity differently. In this study, we systematically evaluated the effects of NO on the cell envelopes of representative ESKAPE pathogens as a function of NO donor (i.e., structure, payloads, and release kinetics). Both antimicrobial assays and microscopies were employed to investigate the effects of NO treatment on the cell envelopes of Gram-negative and Gram-positive bacteria. We also evaluated multi-drug resistant (MDR) bacteria to elucidate the role of AMR on observed NO-mediated effects.

Results and Discussion

Nitric oxide exhibits antibacterial activity against both Gram-positive and Gram-negative bacteria, allowing high potential for NO-based therapeutics in the era of increasing AMR infections. While the effects of NO on Gram-negative bacteria have been reported previously to be membrane disrupting,29 the work described herein was focused on investigating whether NO donors of varying release kinetics altered changes to cell envelope as a function bacteria type (Gram-negative versus Gram-positive). We hypothesized that NO may act differently on bacterial envelopes depending on the bacteria class, NO release, and NO donor identity (i.e., structure). Three small molecules and one oligosaccharide NO donors were selected as model NO donor systems due to their high NO payloads (1.5– 10 μmol/mg) and ability to eradicate both Gram-negative and Gram-positive bacteria. The NO-release payloads and half-lives of materials utilized in this study are provided in Supporting Information (Table S1).

Inhibitory and bactericidal assays

Before analyzing the structure-altering effects of NO on bacteria, the killing doses of the NO donors were determined using MBC assays. Except for spermine, control (non-NO-releasing) scaffolds (i.e., NO donor precursor structures) did not elicit bacterial killing even at doses up to 16 mg/mL, whereas the MBCs of the corresponding NO donors spanned 0.5–2 mg/mL, corresponding to NO doses of 1.5–12 mM (Table 1). The MBCs of antibiotic-susceptible and -resistant strains of bacteria were nearly identical, indicating that antibiotic resistance does not influence the manner in which NO kills bacteria or its mechanisms of action.

Table 1.

Minimum bactericidal concentrations for NO donors and their precursors against bacteria. For the NO donors, the NO payload in mM is provided in parentheses.

Strain MBC (mg/mL)
SPER SPER/NO CD-EDA7 CD-EDA7/NO
Klebsiella pneumoniae MGH 78578 8–16 2 (11) >16 1 (1.5)
Klebsiella pneumoniae KPPR1-ΔwcaJ 8 1–2 (5.5–11) 16 2 (3.0)
Pseudomonas aeruginosa PAO1 8–16 1 (5.5) >16 1 (1.5)
Enterococcus faecium JL282 >16 1 (5.5) >16 8–16 (12–24)
Staphylococcus aureus LAC >16 1–2 (5.5–11) >16 1–2 (1.5–3)
Staphylococcus aureus SF8300 >16 0.5 (2.8) >16 1 (1.5)
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Impact of nitric oxide on bacteria class and cell envelope type

After the bactericidal doses of NO were determined via microplate bactericidal assays, transmission electron microscopy (TEM) and atomic force microscopy (AFM) were used to investigate the impact of NO donors on the integrity of the cell envelopes of the different bacteria species. Gram-negative bacteria possess both a plasma membrane and an outer membrane separated by a thin layer of peptidoglycan (PG). In contrast, Gram-positive bacteria possess only one membrane (plasma) but with a much thicker PG outer layer that maintains structural integrity and osmotic pressure. These cell envelope differences are readily observed in TEM micrographs of untreated bacteria (Figure 1). Gram-negative Klebsiella pneumoniae and Pseudomonas aeruginosa show oblong cross-sections with a thin cell envelope and apparent double membrane. The outer membranes of both species contain lipopolysaccharides (LPS), consisting of a lipid A anchoring to the membrane, an oligosaccharide core, and a polymeric side chain.31 In addition to LPS, capsule is present in K. pneumoniae, which refers to an assembly of repeating sugar subunits known as capsular polysaccharides (CPS).31 This additional capsule layer (Figure 1A), giving the bacterial cell envelope of K. pneumoniae a fuzzy appearance and slightly larger apparent thickness versus P. aeruginosa. Compared to Gram-negative bacteria, both Gram-positive species (S. aureus and E. faecium) displayed more circular-like cross-sections and thicker cell envelopes that are attributed to the PG layer.

FIGURE 1.

FIGURE 1.

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TEM micrographs of Klebsiella pneumoniae MGH78578 (A), Pseudomonas aeruginosa PAO1 (B), Staphylococcus aureus LAC (C), and Enterococcus faecium JL282 (D). Bacteria were untreated (A1-D1) or treated with 2xMBC of SPER/NO (A2-D2) and CD-EDA7/NO (A3-D3). Bacterial death (>99.9%) was confirmed via agar plating for all samples.

These structural differences altered the impact of NO. Treatment of Gram-negative bacteria with SPER/NO and CD-EDA7/NO led to extensive membrane disruption and warping of cell shape (Figure 1). Such effects were not observed for non-NO releasing materials (data not shown), proving that the disruption resulted from NO and is a key factor in killing of Gram-negative bacteria. The membrane disruption was observed in both K. pneumoniae and P. aeruginosa, despite the former having an extra protective capsule layer, indicating that membrane disruption is common for Gram-negative bacteria. These results align with a prior report that described NO’s ability to disrupt the bacteria’s envelope by phospholipid oxidation.29 In contrast, disruption of the cell envelope was not observed in Gram-positive bacteria, even though bacterial cultures proved to be nonviable via agar plating. When treated with bactericidal levels of NO, both S. aureus and E. faecium maintained circular cell shape and apparent integrity (Figure 1) despite death.

Atomic force microscopy (AFM) was used to quantify the surface roughness of individual bacteria by raster scanning a sharp probe across the surface of the bacterial cell. The calculated root-mean-square (RMS) roughness was then used to correlate relative roughness to cell envelope damage. In this regard, greater RMS values indicate enhanced cell envelope damage. As shown in Figure 2, the RMS roughness values of control K. pneumoniae, P. aeruginosa, S. aureus, and E. faecium were 8.2 ± 1.4, 8.4 ± 0.9, 10.6 ± 0.5 and 8.3 ± 1.3 nm, respectively. In accordance with TEM results, the RMS roughness values of Gram-negative bacteria increased for both K. pneumoniae and P. aeruginosa upon treatment with NO. In contrast, the RMS roughness values of Gram-positive S. aureus LAC and E. faecium JL282 showed no significant difference between control and bacteria treated with NO. Of note, qualitative differences in Gram-negative bacteria were observed between different NO donor architectures (i.e., small molecule versus cyclodextrin). Specifically, TEM micrographs indicated that treatment of K. pneumoniae and P. aeruginosa with CD-EDA7/NO imparted membrane disruption that was discernible but less dramatic than that for spermine/NO. Nevertheless, no significant difference between spermine/NO and CD-EDA7/NO treatment was observed for RMS roughness of Gram-negative bacteria (Figure 2).

FIGURE 2.

FIGURE 2.

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Root-mean-squared surface roughness of Klebsiella pneumoniae MGH78578 (A), Pseudomonas aeruginosa PAO1 (B), Staphylococcus aureus LAC (C), Enterococcus faecium JL282 (D), treated with non-NO-releasing (hollow) and NO-releasing (filled) spermine/NO (red) and CD-EDA7/NO (green). Areas of 500–800 nm2 were imaged at 0.5 Hz and height traces were first-ordered flattened.

As the outer membrane of Gram-negative bacteria and the PG layer of Gram-positive bacteria each play a protective role to the plasma membrane, a comparison of these two cell wall components may account for the significant disruption of the cell envelope for Gram-negative but not Gram-positive bacteria. While both LPS (residing in the outer membrane) and PG contain polymeric chains of sugar residues, the composition of these polymers differs significantly. The structure of LPS is dominated by long oligosaccharide chains that protect Gram-negative bacteria by presenting antigens that mask the bacteria and help evade innate immune response.31,32 Peptidoglycan consists of alternating layers of N-acetylglucosamine and N-acetylmuramic acid that are cross-linked by a tetrapeptide, forming a mechanically strong lattice.33 The different layers are also anchored by teichoic acid residues that increase PG rigidity.6 Additionally, the outer membrane of Gram-negative bacteria is significantly thinner (ca. 2.5 nm in P. aeruginosa) than that of PG in Gram-positive bacteria (30–100 nm), conferring the latter substantially greater rigidity under extracellular stress or damage.34

Impact of nitric oxide-release kinetics

It was hypothesized that the NO-release kinetics may lead to different effects on the bacterial cell envelope. A couple of NO donors were thus added to the study, including propylamine propylamine NONOate (PAPA/NO) and dipropylenetriamine NONOate (DPTA/NO) – fast and slow NO-releasing NO donors, respectively (t1/2 = 0.4 ± 0.02 h and 5.2 ± 0.3 h). For reference, spermine/NO (SPER/NO) is an intermediate NO-releasing molecule with a NO-release half-life of 2.0 ± 0.4 h. To isolate the effects of only NO-release kinetics, bacteria were treated with concentrations of NO donor that resulted in an NO payload of 40 mM. Micrographs from TEM experiments showed that treatment of Gram-negative K. pneumoniae led to significant warping and/or disruption of bacterial cell envelopes (Figure 3). Of note, the NO-release kinetics did not influence changes to the membrane integrity. Results from AFM experiments corroborated this finding, as we observed an increase in RMS roughness regardless of the NO donor, indicating comparable deterioration of bacterial membrane (Figure 4). Indeed, the only distinction in envelope damage was again observed between Gram-negative and Gram-positive bacteria, as each of the NO donors appeared to produce no significant changes to morphology or membrane roughness upon treating S. aureus.

FIGURE 3.

FIGURE 3.

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TEM micrographs of Klebsiella pneumoniae MGH78578 (A-C), and Staphylococcus aureus LAC (D-F) treated with PAPA/NO (A, D), SPER/NO (B, E) and DPTA/NO (C, F). Bacterial death (>99.9%) was confirmed via agar plating for all samples.

FIGURE 4.

FIGURE 4.

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Root-mean-squared surface roughness of Klebsiella pneumoniae MGH78578 (A), and Staphylococcus aureus LAC (B) treated with PAPA/NO (blue), SPER/NO (red) and DPTA/NO (green) or untreated (black). Areas of 500–800 nm2 were imaged at 0.5 Hz and height traces were first-ordered flattened.

Role of antibiotic susceptibility

We hypothesized that because NO acts on bacteria via multiple mechanisms, its effects on bacterial cell envelopes would be similar regardless of antibiotic susceptibility. To investigate this hypothesis, we exposed NO-releasing donors to two different strains of K. pneumoniae (MGH 78578 and KPPR1-ΔwcaJ). Compared to K. pneumoniae strain MGH 78578, KPPR1-ΔwcaJ is resistant to both rifampicin and streptomycin. Similar experiments were carried out with LAC (methicillin-susceptible) and SF8300 (methicillin-resistant) S. aureus strains. Comparing the TEM images of MGH 78578 and KPPR1-ΔwcaJ before and after NO donor treatment indicated that the use of SPER/NO (Figure 1, Figure S2) led to similarly significant disruption of the cell membrane (e.g., warping and shearing of cell membranes) while the effect of CD-EDA7/NO treatment on membrane disruption was far less (Figure 1, Figure S2). Similar results were observed for S. aureus LAC and SF8300 strains, which showed that the bacterial cell envelopes were not affected by NO (Figure S3). These findings agree with other literature that indicates NO kills bacteria by multiple pathways10 and therefore does not differentiate between antibiotic resistant and susceptible bacteria. In the context of increasing antibiotic resistance worldwide, this result suggests that NO-releasing prodrugs may be a promising alternative strategy for treating bacterial infections.

Staphylococcus aureus membrane depolarization under treatment by nitric oxide

TEM and AFM images of bacteria treated with NO donors revealed that regardless of the NO-release kinetics, the cell envelope of Gram-positive bacteria was not visibly affected. This finding contradicts some previous work, including a study by Park et al. who reported via TEM that treatment of S. aureus with a NO-releasing polyethylenimine-based tri-block copolymer (F68-BPEI-NONOates) resulted in disruption of cell envelope accompanied by discharge of cytoplasmic material.35 However, Park et al. did not comment on the status of S. aureus when treated with non-NO-releasing F68-BPEI. Given that the concentration-dependent kill-curve of F68-BPEI-NONOates closely resembles that of its non-NO-releasing analog, it is likely that the membrane damage and toxicity were caused by the polymeric scaffold and not NO.

Based on evidence that NO treatment disrupts the cytoplasmic membrane of Gram-negative bacteria, we postulated that NO diffuses through the PG layer of Gram-positive bacteria and induces killing by permeabilizing the cytoplasmic membrane. Even upon NO-mediated killing, the PG layer provides rigidity and maintains the shape of the cell envelope. To investigate this claim, we studied the depolarization of S. aureus cytoplasmic membrane using 3,3′-dipropylthiadicarbocyanine iodide (DiSC3-5). In this fluorescent assay, DiSC3-5 was added to a black 96-well plate containing S. aureus treated with an NO donor. The ensuing fluorescence intensity at 672 nm was measured every 30 s for 20 min. During this observation period, a slight overall decrease in fluorescence intensity was observed across samples due to DiSC3-5 adsorption onto the surface of the 96-well plates (Figure 5A). In uncompromised (i.e., viable) bacteria, DiSC3-5 is readily transported inside the cytoplasmic membrane, which leads to rapid fluorescence quenching (Figure 5A, blue). Chloramphenicol, a bacteriostatic antibiotic, was also added to untreated bacteria to prevent growth and distortion of the fluorescent readings. Treatment of S. aureus by both SPER/NO and CD-EDA7/NO led to slower quenching of fluorescence intensity. To provide a clear comparison between the samples, steady-state measurements were plotted (Figure 5B). Staphylococcus aureus samples treated with SPER/NO and CD-EDA7/NO had 4–5 times greater fluorescence levels than chloramphenicol-treated bacteria, indicating that the ionic balance across the cytoplasmic membrane had been disrupted (i.e., the bacteria are no longer able to retain DiSC3-5). In separate control experiment, Staphylococcus aureus was treated with rifampicin, an antibiotic that deactivates RNA polymerase and causes bacterial death by inhibiting RNA synthesis. As shown in Figure 5B, the fluorescence of bacteria with DiSC3-5 remained low as might be expected because rifampicin’s mechanism of action does not compromise the cytoplasmic membrane. The results of these experiments demonstrate that NO-mediated killing in Gram-positive bacteria is achieved via depolarization of cytoplasmic membrane because of NO’s diffusion through the PG layer. Diffusion through the approx. 0.1-micron thick PG layer supports earlier findings that somewhat larger NO doses are generally required to eradicate Gram-positive versus Gram-negative bacteria.

FIGURE 5.

FIGURE 5.

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(A) Changes of DiSC3-5 fluorescence intensity in Staphylococcus aureus vs. time. Graph depicts a representative plot of one independent experiment. Error bars represent standard deviation of four technical replicates. (B) Fluorescence intensity of DiSC3-5 at t = 25 min (20 min after addition of DiSC3-5). Error bars represent standard deviation for n ≥ 3 independent experiments. * p < 0.05. **** p < 0.0001.

Effects of nitric oxide-releasing quaternary ammonium-modified dendrimers

Given that CD-EDA7/NO and the small molecule NO donors did not appear to inflict major structural damage to Gram-positive bacteria, we sought to investigate whether other NO donor systems might do so despite the thick PG layer. Worley et al. previously reported the unprecedented antibacterial activity of quaternary ammonium (QA)-modified dendrimers.27 These QA-modified dendrimers were originally envisioned as “dual-action” antimicrobial agents as they combined the bactericidal action of NO with the ability of alkyl chains to target hydrophobic bacterial envelopes. The alkyl chain length of the QA modification proved to be an important factor in determining the antibacterial activity of the dendrimers with longer alkyl chains lowering the MBC for Gram-positive bacteria. Based on the results by Worley et al., K. pneumoniae and S. aureus were treated with a C10-QA modified dendrimer (dendrimer-QA) in a manner analogous to the studies reported above. Of note, the MBCs of the NO-releasing dendrimer-QA (Table S2) were comparable to those reported by Worley et al.27 As shown in Figure 6, TEM experiments revealed membrane morphology distinct from that of bacteria treated by the NO-releasing cyclodextrin and small molecule NO donors. Compared to these NO donors, the dendrimer-QA/NO system showed less significant membrane disruption to Gram-negative bacteria. However, visible delamination of membrane from the cytoplasm was observed (Figure 6B, C). The disruption of Gram-positive bacteria was characterized by puncturing of the thick PG layer at multiple points. Such effects were not seen with any of the other NO donors. As Gram-positive bacteria lack double-layered membranes, the ability of dendrimer-QA to disrupt the PG layer substantiates its lower bactericidal concentration.

FIGURE 6.

FIGURE 6.

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TEM micrographs of Klebsiella pneumoniae MGH78578 (A-C), and Staphylococcus aureus LAC (D-F). Bacteria were untreated (A, D) or treated with 2x MBC of dendrimers-QA (B, E) or dendrimers-QA/NO (C, F). Bacterial death (>99.9%) was confirmed via agar plating for all samples.

Conclusions

Nitric oxide is a potent antibacterial agent with activity against both Gram-negative and Gram-positive bacteria. While NO inflicts membrane disruptions to both Gram-negative and Gram-positive bacteria, the extent of disruption is largely independent of NO donor architecture but dependent on Gram designation. In Gram-negative bacteria that lack a thick PG layer, NO donors effectively rupture both the outer and cytoplasmic membranes, causing puncture and loss of cell shape. In Gram-positive bacteria, the disruption of cell envelope by NO is more nuanced: the thick PG layer maintains cell rigidity despite cell membrane depolarization. These results suggest that the PG layer is a diffusion barrier to killing by NO, which aligns with past findings where exogenous NO is found to be more effective in Gram-negative bacteria.

Methods

Materials

β-cyclodextrin (CD), bis(3-aminopropyl)amine (DPTA), N-(2-hydroxyethyl)ethylenediamine (HEDA), N-propyl-1,3-propanediamine (PAPA), ethylenediamine (EDA), spermine (SPER), epoxy medium, dodecenylsuccinic anhydride, methyl nadic anhydride, 2,4,6-tris(dimethylaminomethyl)phenol were purchased from Sigma-Aldrich (St. Louis, MO). Sodium methoxide (5.4 M in methanol), triphenylphosphine, and common laboratory salts and solvents were purchased from Thermo Fisher Scientific (Fair Lawn, NJ). Tryptic soy broth (TSB), cation-adjusted Mueller Hinton broth (CAMHB), and tryptic soy agar (TSA) were obtained from Becton, Dickinson, and Company (Franklin Lakes, NJ). Unless otherwise specified, all chemicals were used as received without further purification. Nitrogen (N2), oxygen (O2), NO calibration (25.87 ppm balance N2), and pure NO (99.5%) gas cylinders were purchased from Airgas National Welders (Raleigh, NC). Distilled water was purified to a resistivity of 18.2 MΩ-cm and a total organic content of ≤6 ppb using a Millipore Milli-Q UV Gradient A10 system (Bedford, MA).

Planktonic Bacterial Inhibition and Eradication Assays

The minimum inhibitory concentrations (MIC) of the NO-releasing donors and controls (i.e., non-NO-releasing forms) were determined using the broth microdilution method.30 Overnight cultures were created from stock agar plates made fresh for each experiment. Overnight solutions were diluted in 5 mL of TSB, grown to 108 CFU/mL, and then diluted to 5×105 CFU/mL in CAMHB. These bacterial solutions were subsequently distributed to 96-well plates. The NO-releasing donors were prepared in CAMHB, pH adjusted to 7, and transferred to the 96-well plates at volumes to result in serially dilution with the bacteria solutions. The 96-well plates were then incubated at 37°C for 24 h. The MIC of the NO-releasing donor was determined as the lowest concentration leading to no visual growth. The minimum bactericidal concentrations (MBCs) were determined by plating (10 μL) of concentrations greater than or equal to the MIC on TSA, incubating overnight at 37°C and counting to determine the concentration resulting in 3-log reduction of bacterial viability.

Transmission Electron Microscopy

Gram-negative bacteria were grown to OD ~0.7 and Gram-positive bacteria were grown to OD ~1.0 in TSB to generate a bacterial pellet for sample preparation. To ensure killing of high bacterial density, 5-mL bacterial suspensions in CAMHB were treated with 1.5–2x MBC NO donors or equal concentrations of controls for 24 h at 37 °C under aerobic conditions. Subsequently, 10-μL aliquots were plated to determine viability after treatment.

Sample preparation: Bacterial pellets were washed three times with PBS, fixed in 2.5 % w/v glutaraldehyde in PBS at 4 °C overnight, then washed three times with 0.1 M cacodylate buffer to remove glutaraldehyde. The pellets were post-fixed in 2% aqueous osmium tetroxide solution for 2.5 h, then washed three times with 0.1 M cacodylate buffer. The pellets were serially dehydrated with 50, 70, 90%, and twice with 100% ethanol. For each dehydration, the pellets were suspended in the ethanol solution in a centrifuge tube and gently agitated on a tube rotator for 10 min at room temperature. The pellets were then recollected via centrifugation and the supernatant was replaced with the next ethanol solution. After serial dehydration, a transition step was conducted where the pellets were incubated for 15 min at room temperature with 1:1 ethanol/propylene oxide and twice with 100% propylene oxide.

After the transition step, an infiltration step was conducted. The pellets were incubated for 1 h without shaking at room temperature in a 50/50 propylene oxide/epoxy resin solution. The epoxy resin solution is EPON 812 substitute (MilliporeSigma, St. Louis, MO) comprising a mixture of 46.5% epoxy medium/28.5% dodecenylsuccinic anhydride/25% methyl nadic anhydride, supplemented with 2% v/v of 2,4,6-tris(dimethylaminomethyl)phenol. All components were mixed thoroughly, and the mixture was sonicated to remove air bubbles. After the infiltration step, each bacteria pellet was deposited at the tip of a BEEM capsule size 00 (TedPella, Redding, CA) using either forceps or a micropipette, embedded in 100% epoxy resin solution, and incubated at 70 °C for 24 h. The epoxy samples were trimmed, sectioned into 60-nm thick slices using ultramicrotome and imaged using Talos F200X transmission electron microscope.

Atomic Force Microscopy

Bacterial overnight cultures were diluted in fresh TSB (30 mL), grown to 108 CFU/mL, and treated with control or NO-releasing test agents in CAMHB for 24 h at 37° C. After treatment, the cultures were centrifuged (3,000 × g for 10 min, 25° C). The pellet was resuspended in 20 mL of PBS at pH 7.4. Freshly cleaved mica was incubated first with 250 μL of 0.1 mg/mL PLL for 1 h, then with a 250-μL aliquot of the bacterial suspension in PBS for 1 h. Excess liquid was carefully removed with a micropipette after each incubation period. Immediately before AFM imaging, the mica specimen was gently rinsed with 250 μL of ultrapure water from a micropipette and dried with a gentle jet of nitrogen gas.

Tapping-mode AFM was conducted using an Asylum Research MFP-3D AFM system (Santa Barbara, CA) and AC240TS silicon beam cantilevers (Olympus, Center Valley, PA). Survey scans (40 μm × 40 μm) were conducted before surface roughness images (500 × 500 nm for Gram-positive bacteria, 800 × 800 nm for Gram-negative bacteria) were acquired at 0.5 Hz scan rate and a resolution of 512 × 512 pixels. Images were first-order flattened before root-mean-square roughness measurements were determined using software provided by Asylum Research.

Membrane depolarization assay

Overnight cultures of Staphylococcus aureus were diluted in fresh TSB, grown to 108 CFU/mL, aliquoted into microcentrifuge tubes and treated with NO-releasing materials in CAMHB for 24 h at 37° C. An inhibition of bacterial growth with 5 μg/ml chloramphenicol was found to result in stable fluorescence levels and thus used as a control condition. After treatment, the cultures were centrifuged (21,300 × g for 10 min, 25 °C) and the pellets were resuspended in 1 mL of 37 °C LB supplemented with 0.5 mg/ml bovine serum albumin. The suspension (135 μL) was transferred to the microtiter plate and the background fluorescence was measured for 5 min. After a baseline was obtained, DiSC3-5 (100 μM in DMSO) was added to each well to a final concentration of 1 μM DiSC3-5 and 1% v/v DMSO. The fluorescence quenching was measured for 20 min until a stable signal intensity was achieved.

Supplementary Material

Supporting Information

Acknowledgements

This work was performed in part at the Chapel Hill Analytical and Nanofabrication Laboratory, CHANL, a member of the North Carolina Research Triangle Nanotechnology Network, RTNN, which is supported by the National Science Foundation, Grant ECCS-2025064, as part of the National Nanotechnology Coordinated Infrastructure, NNCI

Support Statement

This work was funded by the National Institutes of Health (DK132778 and DE032060).

Footnotes

Supporting Information

The Supporting Information is available free of charge on the ACS Publications website.

Additional materials and methods, reaction schemes, nitric oxide-release payloads, transmission electron micrographs, root-mean-squared roughness of K. pneumoniae and S. aureus, and minimum bactericidal concentrations.

Conflicts of Interest

The corresponding author declares competing financial interest. M. H. Schoenfisch is a co-founder, is a member of the board of directors, and maintains a financial interest in KnowBIO, LLC and Vast Therapeutics. Vast Therapeutics is commercializing macromolecular nitric oxide storage and release scaffolds for the treatment of respiratory infections.

References

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Supplementary Materials

Supporting Information
NIHMS1984780-supplement-Supporting_Information.pdf (604.8KB, pdf)

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