Amphiphile‐Assisted Synthesis of Ruthenium Nanoparticles for Controlled Release and Enhanced Antibacterial Activity

Abstract

A novel procedure for preparing Ruthenium nanoparticles (RuNPs) based on low‐molecular‐weight amphiphilic molecules and Ru(III) complexes as antibacterial agents with controlled release properties has been developed. Two hydrophobic Ru(III) complexes, Ru‐TOA and Ru‐Benza, analogs to the NAMI‐A prodrug, are encapsulated within the core of the micelles formed through the self‐assembly of these amphiphiles. The self‐assembly of amphiphile I, which contains a double polar head, results in highly water‐stable and monodispersed RuNPs incorporating both hydrophobic Ru complexes. These RuNPs exhibit hydrodynamic sizes ranging from 26.7 to 104.2 nm for NPs derived from Ru‐TOA complex, and ≈10 nm for those derived from Ru‐Benza. Compared to Ru(III) complexes, these RuNPs offer several advantages, including protection from aqueous degradation and enhanced bacterial uptake. Moreover, post‐synthesis modification of the RuNPs with molecular staples based on polyethylene glycol chains of varying lengths enables controlled Ru release, reducing the burst effect. Interestingly, these RuNPs demonstrate excellent antibacterial activity, with minimum inhibitory concentration (MIC) values of 16 mg·L⁻¹ and minimum bactericidal concentration (MBC) values of 32 mg·L⁻¹ against a broad range of Gram‐positive bacteria, including S. aureus, Staphylococus pseudintermedius, and Enterococcus faecalis, highlighting their potential efficacy against clinically relevant bacterial strains.

A novel synthesis of Ruthenium nanoparticles (RuNPs) based on amphiphiles and Ru(III) complexes as antibacterial agents with controlled release properties has been developed. Post‐synthesis modification of the RuNPs with PEG molecular staples enables controlled Ru release, reducing the burst effect. The RuNPs exhibit excellent antibacterial activity against clinically relevant bacterial strains.

1. Introduction

The rapid rise in bacterial resistance to antibiotics is one of the major global challenges that humanity must face in the coming decades. According to recent studies, by 2050, around 10 million people could die each year from bacterial infections.^{[ 1} , ² , ^{3 ]} While subsequent studies have revised these estimates, antimicrobial resistance (AMR) remains a significant clinical and public health burden that is expected to increase over time, highlighting the urgent need for action.^{[ 4} , ^{5 ]}

In particular, several bacterial species, including Staphylococcus aureus, Escherichia coli, Klebsiella pneumonia, and Mycobacterium tuberculosis, already exhibit concerning levels of resistance, and therapeutic options for some multi‐resistant strains are limited.^{[ 6} , ^{7 ]} In this context, the search for new compounds with proven antimicrobial activity remains one of the most active areas of research.^{[ 8} , ⁹ , ^{10 ]}

Among various strategies, transition metal‐based ions are gaining significant attention in biomedical research as promising anticancer and antimicrobial agents due to their involvement in biological processes.^{[ 11} , ^{12 ]} A notable example is the Pt(II) complex cisplatin, approved worldwide in 1978, which remains the most important metallodrug for clinical use. Moreover, other platinum‐based drugs, such as carboplatin and oxaliplatin, were approved as anticancer agents in 1989 and 2003, respectively.^{[ 13 ]} Regarding antimicrobial activity, silver‐ and copper(II)‐based complexes have been extensively investigated for their potent bactericidal activity.^{[ 14} , ^{15 ]} The significance of metal‐based drugs lies in their unique electronic and stereochemical properties, which distinguish them from organic compounds. These characteristics enable them to combine with a wide variety of organic ligands in different coordination geometries. One particularly intriguing transition metal, ruthenium in the oxidation states II and III, has been extensively studied in recent years as a promising metallodrug. Among them, the Ru complexes NAMI‐A, KP1339, TLD1433, and BOLD‐100 have progressed to phase I/II in clinical trials as anticancer compounds. The Ru(III) complexes are considered inert, being necessary to activate them by reducing to Ru(II) or forming aqua complexes.^{[ 16} , ^{17 ]} In contrast, the Ru(II) complexes exhibit significant biological activity through various modes of action as interaction with nucleic acids, production of reactive oxygen species (ROS), or inhibition of enzymes. As antibacterial compounds, ruthenium(II) complexes have been demonstrated to be effective against both Gram‐positive and Gram‐negative bacteria. For instance, Ru(II) polypyridyl complexes have displayed bactericidal activity through multiple mechanisms, including DNA intercalation, interaction with DNA or RNA, inhibition of specific proteins or biofilm formation, and ROS generation.^{[ 18} , ¹⁹ , ²⁰ , ^{21 ]} However, an important drawback of these Ru(II) complexes is their high cytotoxicity in mammalian cells. On the other hand, a few Ru(III) complexes with limited antimicrobial activity have been reported as antimicrobial agents. The efficacy of the Ru(III) complexes lies in the ability to disrupt cellular metabolism through the properties of their chelating ligands and/or the activation of the Ru(III) into Ru(II).^{[ 22} , ²³ , ^{24 ]} However, significant challenges remain in this biomedical field, particularly regarding the rise in antibiotic resistance, more active compounds, and the high toxicity of developed metallodrugs, which limit their clinical applications.^{[ 14} , ^{25 ]} Nanotechnology appears to be a potential solution to tackle these issues. Nanomaterial‐based therapies offer an innovative approach to combating AMR due to their unique size, physicochemical properties, and mechanisms of action that can overcome traditional antibiotic limitations. Nanomaterials can produce direct antimicrobial activity by disrupting bacterial membranes (i.e., metallic NPs), generating ROS (i.e., metal and metal oxide NPs), or interfering with intracellular processes.^{[ 26} , ²⁷ , ²⁸ , ^{29 ]} Moreover, nanocarriers can enhance the delivery of antibacterial drugs by improving their stability, targeting, and bioavailability.^{[ 30 ]} In fact, these nanomaterials can bypass bacterial efflux pumps, target multiple cellular components, or induce physical damage to bacteria.^{[ 25} , ²⁸ , ³¹ , ^{32 ]} Advances in the engineering of nanomaterials permit tuning properties as size, shape, coating, and composition to target bacteria with different modes of action and reduced side effects. In particular, nanocarriers based on organic materials such as micelles, vesicles, and hydrogels are emerging as versatile platforms for drug encapsulation, preventing degradation, transporting drugs to the site of infection, and overcoming resistance mechanisms.^{[ 33} , ^{34 ]} With this consideration in mind, herein, we report the design and synthesis of Ru(III)‐based nanoparticles (RuNPs) that exhibit potent antibacterial activity against Gram‐positive bacteria such as Staphylococcus aureus. These RuNPs were prepared through encapsulation of hydrophobic ruthenium(III) complexes using low‐molecular‐weight (MW) amphiphiles. These Ru(III) complexes could be considered prodrugs similar to NAMI‐A, requiring activation via aquation or redox reactions. However, the use of these Ru(III) complexes as drugs is limited by their poor stability in water. To overcome this limitation, the encapsulation in organic nanomaterials offers several advantages including: i) Tuneable synthesis of RuNPs, ii) Protection of Ru(III) complexes from aqueous degradation, iii) Enhanced bacterial uptake compared to free Ru complexes, iv) Controlled Ru release, achieved by surface modification with covalently attached polyethylene glycol (PEG)‐based molecules to prevent the burst release.

2. Results and Discussion

2.1. Synthesis of Ruthenium Nanoparticles

The first step was the synthesis of non‐water‐soluble Ru(III) complexes. The hydrophobic Ru(III) complexes were prepared from hydrogen trans‐Bis(dimethyl sulfoxide) tetrachlororuthenate(III) as Ru precursor, which is used in the synthesis of the hydrophilic NAMI‐A complex.^{[ 16 ]} The incorporation of a hydrophobic counterion in the Ru(III) complex precursor could lead to the formation of the desired Ru payload. Therefore, the addition of tetraoctylammonium (TOA) bromide to a solution of the Ru precursor in a solvent mixture of H₂O/EtOH (1:4) resulted in the formation of a non‐water‐soluble Ru complex (referred to as Ru‐TOA). This new Ru complex was extracted with dichloromethane to remove the unreacted Ru precursor and ammonium salts. This simple and efficient methodology produced the exchange of the (DMSO)₂H counterion for a hydrophobic counterion with an excellent yield. For comparison, another hydrophobic Ru complex was prepared using a common counterion found in some antibiotics, such as benzathine. Therefore, the exchange of the (DMSO)₂H cation for protonated benzathine led to the desired Ru complex (referred to as Ru‐Benza) with a moderate yield.

The two new Ru complexes, Ru‐TOA and Ru‐Benza, were characterized by NMR, Ultraviolet–visible (UV–vis), Fourier transform infrared (FTIR), and Powder X‐ray diffraction (PXRD) spectroscopies (Figures S1–S3, Supporting Information). In spite of the paramagnetic behavior of the Ru(III) nucleus, the ¹H‐NMR spectra showed the signals of the counterions in the typical range from 8 to 1 ppm, and also a dramatic upfield shift and broadening of the DMSO coordinated to the Ru at −12 ppm for both Ru complexes (Figures S4 and S5, Supporting Information). FTIR spectra exhibited also the presence of the counterions, TOA and Benza, in the Ru complexes. In the spectrum of Ru‐TOA (Figure S1A, Supporting Information), the peaks in the range 3000–2840 and 1465 cm⁻¹ correspond to C‐H stretching and bending, respectively, which could be attributed to the counterion TOA. On the other hand, the counterion Benza evidences the peaks in the range 2000–1650 cm⁻¹ assigned to mono‐substituted phenyl absorptions (Figure S1B, Supporting Information). The peaks in the range 1070–1030 cm⁻¹, in both spectra are attributed to the axial sulfoxide bound to Ru. UV–vis spectra exhibited the typical absorption band of the Ru(III) complexes at 430 nm, in combination with the absorption bands of the corresponding counterions (Figure S2, Supporting Information).^{[ 16 ]} Finally, PXRD of Ru‐TOA and Ru‐Benza show the presence of the main peaks that match the spectrum of the Ru precursor, as can be observed in Figure S3 (Supporting Information). Interestingly, in both XRD spectra, these peaks exhibited a shift related to the exchange of the counterion, which influences the lattice parameters of the Ru complexes.

After the synthesis of the hydrophobic Ru complexes, the Ruthenium nanoparticles were prepared through an encapsulation procedure led by the self‐assemblies of low‐molecular‐weight amphiphilic molecules (Figure 1 ). Two different amphiphilic molecules based on oleic acid, as a hydrophobic tail, and tetraethylene glycol (TEG), as a hydrophilic polar head, were successfully synthesized following a protocol reported by some of us.^{[ 35 ]} These amphiphiles exhibit a high capacity to self‐assemble into micelles with low critical micelle concentration (CMC) values, and also, to encapsulate hydrophobic payloads, such as inorganic NPs and drugs.

Figure 1.

Structures of the amphiphilic molecules I and II, and schematic representation of the RuNPs synthetic procedure.

Thus, the sonication of a hydrophobic Ru complex (Ru‐TOA or Ru‐Benza) in dichloromethane (DCM) with a solution of the amphiphilic molecule (I or II) in water led to the encapsulation of the hydrophobic Ru complex in micelles by forming water‐stable Ru nanoparticles (RuNPs), as it is described below in the characterization. Finally, the solution was left at 4 °C overnight, and then centrifuged at 4000 rpm for 10 min to remove larger aggregates and precipitated hydrophobic Ru complex, resulting in a clear orange solution (Figure S6, Supporting Information). As commented in the introduction section, one of the main drawbacks of the Ru(III) complexes is their poor stability in aqueous solution, which is solved by the encapsulation forming NPs that protect them from degradation. Moreover, the size and stability of these RuNPs could be tuned depending on several factors, including i) sonication time, ii) the nature of the Ru complexes and amphiphilic molecules, and iii) the amount of added Ru complex.

Therefore, the first experiments to optimize the preparation of the RuNPs were conducted by varying sonication times using the hydrophobic Ru‐TOA and amphiphilic molecule I. Briefly, 1 mL of an aqueous solution of amphiphilic molecule I (1 mg mL⁻¹) was added to 0.5 mL of a solution of the Ru complex Ru‐TOA (0.5 mg) in DCM. The mixture was sonicated using a probe sonicator for a specific time. The resulting suspension was purified as commented above, and the obtained clear orange suspension of Ru nanoparticles prepared from Ru‐TOA, named Ru1NPs, was characterized by different techniques.

Dynamic light scattering (DLS) measurements revealed that the hydrodynamic (HD) size of the Ru1NPs decreased progressively with the sonication time, and exhibited polydispersity index (PDI) values in the range 0.19–0.33 (Figure S7A, Supporting Information). In addition, turbidity was observed in mixtures with sonication times shorter than 30 min, indicating the presence of non‐stable material. To determine the optimal sonication time, an intermediate sonication time of 10 min was analyzed by TEM. After the purification process, both the precipitate and the supernatant were examined. TEM images of the precipitate revealed the formation of unstable micro‐needles (Figure S7B, Supporting Information), likely originating from the hydrophobic Ru complex, which could not be fully incorporated by the amphiphilic molecules. This incomplete surface passivation can induce anisotropic surface tensions, potentially driving anisotropic growth and subsequent collapse into fine powder upon centrifugation.^{[ 36} , ³⁷ , ^{38 ]} In contrast, TEM analysis of the supernatant showed small aggregates formed by accumulations of Ru1NPs (Figure S7C, Supporting Information). On the contrary, sonication for 30 min leads to the formation of monodispersed Ru1NPs with a diameter size of 27±1.9 nm and a PDI value of 0.19. TEM images of these Ru1NPs showed uniform and monodisperse NPs (Figure 2A). Moreover, the EDX spectrum revealed the presence of Ru in the nanoparticles (Figure 2B). To confirm that the Ru1NPs were formed from the hydrophobic Ru complexes, micro X‐ray fluorescence (µXRF) analysis was carried out. The µXRF spectrum of the synthesized Ru1NPs exhibited the characteristic signals of the Ru‐TOA complex, including peaks for Ru, the equatorial chlorides, and the DMSO axial ligands (Figure 2C). Moreover, the UV–vis spectroscopy displayed an absorbance pattern similar to that of the starting Ru‐TOA (Figure S8, Supporting Information).

Figure 2.

A) Representative TEM image and corresponding size distribution histogram (n = 100 particles) of Ru1NPs with 30 min sonication. The scale bar corresponds to 50 nm, B) EDX spectrum, and C) µXRF spectrum of Ru1NPs.

Next, we performed studies on the addition of a Ru complex to evaluate the effect on the morphology and stability of RuNPs. Following the hydrophobic Ru‐TOA complexes, we performed the synthesis of Ru1NPs with two types of amphiphilic molecules, I and II. These amphiphiles present diverse hydrophobic‐hydrophilic balances with interesting self‐assembly capacity.^{[ 35 ]} In both cases, the sizes of the Ru1NPs increase linearly with the addition of the Ru complex. The size of the Ru1NPs ranges from 26.7 to 104.2 nm, and from 103.5 to 137.8 nm for the amphiphilic molecules I and II, respectively (Figure 3 ; Figure S9, Supporting Information). Interestingly, the amphiphile I is able to encapsulate three times more Ru complexes than the amphiphile II. The addition of higher amounts of Ru complex in both amphiphiles leads to the formation of RuNPs aggregates, which precipitate over time. As commented above, the nature of the amphiphiles leads to different sizes of RuNPs. So, when comparing the same amount of Ru complex added, the amphiphile I, which presents a double hydrophilic polar head, results in smaller NPs in the presence of Ru‐TOA complex than the amphiphile II, which exhibits a double lipid tail. However, TEM analysis of both Ru1NPs showed no defined morphologies, revealing nanoflower‐like structures for the Ru1NPs with amphiphile I (Figure 3B,C,E,F).

Figure 3.

A) HD size vs [Ru‐TOA complex] using amphiphilic molecule I (n = 3 independent experiments), representative HRTEM images and corresponding size distribution histograms (n = 50 particles) of Ru1NPs synthesized using amphiphilic molecule I at a Ru‐TOA complex concentration of B) 0.5 mg mL⁻¹, scale bars correspond to 100 nm for low‐magnification, and 20 nm for the inset, and C) 6.4 mg mL⁻¹, scale bars correspond to 200 nm for the low‐magnification, and 20 nm for the inset. D) HD size vs [Ru‐TOA complex] using amphiphilic molecule II (n = 3 independent experiments), representative HRTEM images and corresponding size distribution histograms (n = 50 particles) of Ru1NPs synthesized using amphiphilic molecule II at a Ru‐TOA complex concentration of E) 0.7 mg mL⁻¹, scale bar of 250 nm, and F) 10.4 mg mL⁻¹, scale bar of 100 nm.

These studies were also performed with the hydrophobic complex Ru‐Benza. The addition of this complex resulted in the formation of smaller RuNPs compared to those obtained with the Ru‐TOA. The Ru2NPs were also characterized using µXRF spectrum, showing the characteristic signals of the Ru‐Benza complex (Figure S10, Supporting Information). In the case of the amphiphile I, the Ru2NPs exhibited HD sizes between 8 and 10 nm with a low amount of encapsulated Ru (max. 9 mg mL⁻¹). For the amphiphile II, the encapsulation of the Ru‐Benza complex was also low, resulting in the formation of larger NPs with HD sizes up to ≈1000 nm (Figure 4 ; Figure S11, Supporting Information). TEM analysis confirms the formation of Ru2NPs. In the case of the amphiphile I, the TEM images of the Ru2NPs showed a size of 8 nm with a non‐defined morphology at the lowest addition of Ru (Figure 4B). In contrast, at the highest addition of Ru, the NPs exhibited a size of 30 nm with an oval morphology (Figure 4C). The TEM images of the Ru2NPs prepared from the amphiphile II showed larger NPs sizes across the entire range of Ru addition, with aggregated structures observed at the highest addition of Ru‐Benza complex concentration (Figure 4E,F). In fact, these aggregates, on the order of micrometers, appear to be formed from a combination of individual 50 nm RuNPs.

Figure 4.

A) size vs [Ru‐Benza complex] using amphiphilic molecule I (n = 3 independent experiments), representative HRTEM images and corresponding size distribution histograms (n = 50 particles) of Ru2NPs synthesized using amphiphilic molecule I at a Ru‐Benza complex concentration of B) 0.5 mg mL⁻¹, scale bar corresponds to 10 nm, and C) 8.0 mg mL⁻¹, scale bars correspond to 50 nm. D) HD size vs [Ru‐Benza complex] using amphiphilic molecule II (n = 3 independent experiments), representative HRTEM images and corresponding size distribution histograms (n = 50 particles) of Ru2NPs synthesized using amphiphilic molecule II at a Ru‐Benza complex concentration of E) 8.0 mg mL⁻¹, scale bar corresponds to 100 nm, and F) 74.7 mg mL⁻¹, scale bars correspond to 1 µm for the low‐magnification and 200 nm for the inset.

In the same manner as the synthesis of Ru1NPs, the addition of more Ru (9 and 4.5 mg mL⁻¹ for the amphiphiles I and II, respectively) leads to the precipitation of the NPs. Therefore, the size of the RuNPs could be determined following Equation (1).

$$Size\left(nm\right)=Fx{C}_{Ru}\left(m{g}_{Ru}/m{L}_{solution}\right)+b$$ (1)

Equation (1) Size dependence of RuNPs on the concentration of Ru complex.

Where F is the growing factor (reported in nm·mL·mg⁻¹) that determines the variation of size with respect to the added mass of Ru, and b is a constant which depends on the nature of the amphiphilic molecules.

For instance, to demonstrate that the synthesis of RuNPs reported here follows a linear growth and that it is possible to prepare a known size of RuNPs, the addition of 2 mg mL⁻¹ of Ru‐TOA complex with the amphiphilic molecule I was carried out. The Ru1NPs exhibited a size of 32.9 nm, very similar to the predicted size of 32.8 nm.

2.2. Stability of RuNPs

The stability of the RuNPs formed using amphiphilic molecule I in solution was studied by placing them in cuvettes in different media and analyzing them by DLS and UV–vis spectroscopy at 0 h and 5 days. In water, both RuNPs remained unaltered, exhibiting HD sizes and UV–vis spectra similar to the initial particles (data not shown). However, in phosphate‐buffered saline (PBS), both RuNPs exhibited a significant decrease in size from 150 to ≈40 nm and from 15 to 8 nm for Ru1NPs and Ru2NPs, respectively (Figure 5A,B). Moreover, the UV–vis spectra showed a decrease in the main absorption bands for the respective Ru complexes after 5 days of exposure in PBS (Figure 5C,D). For instance, the absorption bands of the Ru1NPs, peaking at 300 and 600 nm, and the band at 350 nm, along with a small shoulder ≈500 nm for the Ru2NPs, all decreased. This indicates that the Ru complex was released from within the micelles into the surrounding buffered solution, leading to the precipitation of the Ru in the cuvette.

Figure 5.

HD size vs time graph in PBS for A) Ru1NPs and B) Ru2NPs (n = 3 independent experiments) formed using amphiphile I. UV–vis absorption spectra in PBS at t = 0 h (black), and t = 5 days (blue), for Ru1NPs C) and Ru2NPs D) formed using amphiphile I.

To demonstrate the loss of Ru from within the micelles, Inductively Coupled Plasma‐Mass Spectrometry (ICP‐MS) analysis was conducted. ICP‐MS confirmed a Ru release of ≈50% from the Ru1NPs within 60 min. Interestingly, the Ru release from the nanoparticles takes place in the presence of salts (Figure 6A). The cations of these salts could be exchanged for the hydrophobic counterion of the Ru (TOA or Benza), thus triggering the Ru release from the systems (Figure S12, Supporting Information).^{[ 39} , ^{40 ]}

Figure 6.

In vitro release study of Ru1NPs using amphiphile I: A) Plot of Ru release in PBS (black) and H₂O (red), B) Comparative Ru release of the Ru1NPs (black) and stapled Ru1NPs with molecular staples of PEG₁₅₀₀ with n = 34 (red), PEG₆₀₀ with n = 14 (blue), TEG with n = 4 (pink), and DEG with n = 2 (green). All data represent the mean of 3 independent experiments (n = 3).

Therefore, further investigations for a more controlled Ru release were carried out. The possibility to control the entrance of cations from the bulk to the interior of the nanosystems could lead to slower exchanges with the consequent effect on the Ru release profile. In this sense, molecules designed with similar functions to staples were synthesized to minimize the cation exchange. These molecules are based on PEG with n repetitions ranging from 2 to 34, ending with reactive groups to bind the amino groups displayed from the RuNPs. Thus, depending on the nature of these molecular staples, the release profile could be tuned. First, the formation of the staples on the surface of the Ru1NPs formed using amphiphilic molecule I (chosen as the RuNPs model) was confirmed by variations on the HD size and ζ‐potential. As expected, the size of the Ru1NPs increases significantly with the molecular weight of the staples. In fact, the PEG‐derived staples with the highest molecular weight, PEG₁₅₀₀ with n = 34, exhibited the highest increase in size, reaching 138.4 nm. Additionally, the polydispersity index (PDI) values of the stapled Ru1NPs were similar to those of the starting Ru1NPs, with a PDI value of 0.4. However, the shortest staple, DEG with n = 2, resulted in Ru1NPs with a larger size of 113.4 nm, and a PDI value of 0.6. The presence of a short length in the staple leads to interlinking NPs rather than interactions with the same NP. Moreover, the success of the stapling could be also followed by the ζ‐potential. All the stapled Ru1NPs resulted in less positive ζ‐potential than the starting Ru1NPs. Most of the staples led to negative ζ−potential NPs except to the staple with n = 34. The stapling effect on the Ru1NPs in the ζ‐potential values could depend on mainly two effects, the steric hindrance and the number of staples per NPs. Low MW PEG staples exhibited negative ζ‐potential values, likely due to the presence of a high number of PEG molecules. In contrast, high MW PEG staples could not be incorporated in similar numbers as the low MW staples, resulting in positive ζ‐potential value (Table 1 ).

Table 1.

DLS analysis of Ru1NPs using amphiphile I with and without stapling.

	HD Size (nm)	PdI	ζ Potential [mV]
Ru1NPs	39.8 ± 2.8	0.39	18.20
Ru1NPs‐PEG1500	138.4 ± 18	0.41	2.56
Ru1NPs‐PEG600	94.2 ± 3	0.40	−1.05
Ru1NPs‐TEG	81.1 ± 1.4	0.60	−7.93
Ru1NPs‐DEG	113.4	0.34	−1.62

The Ru release analysis of the stapled Ru1NPs was carried out using ICP‐MS. As observed in Figure 6B, the highest MW PEG molecular staples (PEG₁₅₀₀ with n = 34) led to a slow release profile, preventing the initial burst release. In contrast, as the MW of the staples decreases, the Ru releases became similar to those of the initial Ru1NPs. Regarding the greatest Ru release, it was observed that the highest MW staples exhibited the maximal Ru release, reaching 87% at 24 h. The other stapled Ru1NPs, with n = 4 (TEG) and 14 (PEG₆₀₀), showed similar release percentages, 53% and 61%, respectively. These differences in release profiles are due to the generation or modification of channels in the nanosystem caused by the stapling of the PEG molecules that connect the micelle core to the external environment and accelerate drug diffusion, supporting the concept that PEG content can tune not just the total extent but also the timing and kinetics of release.^{[ 41} , ⁴² , ⁴³ , ^{44 ]} The lowest MW molecular staple (DEG) could not be compared with the other staples as it led to the formation of small RuNPs aggregates (Table 1).

2.3. Analysis of the Ru Release Profiles

The Ru release of the Ru1NPs can be divided into two profiles following the Korsmeyer‐Peppas model.^{[ 45} , ^{46 ]} The non‐stapled Ru1NPs and the low MW stapled Ru1NPs, with n = 4 (TEG) and 14 (PEG₆₀₀), exhibited release index values ranging from 0.14, for the non‐stapled Ru1NPs, to 0.24 and 0.28 for the stapled Ru1NPs with n = 4 and 14, respectively. These values correspond to a Fickian diffusion model (release index ≤ 0.45), which allows us to calculate the released rate constants for each Ru release profile, being 27.3 for the non‐stapled Ru1NPs, 9.50 and 6.12 for the stapled Ru1NPs with n = 4 and 14, respectively (Figure S13, Supporting Information).

On the other hand, the stapled Ru1NPs with n = 34 (PEG₁₅₀₀) follow a Super Case II transport model (release index > 0.89) as this system exhibited a release index value of 0.9 (Figure S13, Supporting Information). This system showed a released rate constant of 0.116, indicating a slow Ru release profile, which is quite similar to the zero‐order release kinetics model, ideal for the drug delivery systems.

2.4. Microbiological Studies

Three bacterial species have been selected for this work: The Gram‐positive Staphylococcus aureus and two Gram‐negative species, Escherichia coli and Pseudomonas aeruginosa. These species are commonly employed in antimicrobial resistance studies due to their clinical relevance. The bacterial strains used in this work were type strains from the American Type Culture Collection (ATCC), specifically S. aureus ATCC 25923, E. coli ATCC 25922, and P. aeruginosa 27853.

Preliminary susceptibility tests were performed to provide valuable initial insights into the antimicrobial activity of our RuNPs formed using amphiphile I without staples. Therefore, disks were loaded with 100 micrograms of Ru1NPs and Ru2NPs, based on the Ru complex content, and they were placed on a plate inoculated with the corresponding microorganism. Interestingly, the formation of growth inhibition halos for S. aureus by Ru2NPs suggested a promising susceptibility to these NPs, whereas the absence of halos for E. coli and P. aeruginosa indicated limited or no activity against Gram‐negative bacteria (Figure 7A–C). The average diameter of the halo for S. aureus was 11 mm for Ru2NPs. Similarly, Ru1NPs showed no inhibition halos for any of the tested bacteria (Figure 7D–F), suggesting lower antimicrobial efficacy compared to Ru2NPs.

Figure 7.

Antimicrobial susceptibility test using Ru2‐NPs for S. aureus A), E. coli B), and P. aeruginosa (in this panel, all the disks are loaded with Ru2‐NPs). C) Antimicrobial susceptibility test using Ru1NPs for S. aureus D), E. coli E), and P. aeruginosa F) (in this panel, the clear disks correspond to blank disks without NPs).

To quantify more precisely this potential activity of our RuNPs toward Gram‐positive bacteria, and determine the minimum concentration required for bacterial growth inhibition, MIC and MBC measurements were performed. The results of MIC determination indicated good antimicrobial activity of both RuNPs against Staphylococcus aureus, with MIC values of 64 and 32 mg L⁻¹ for Ru1NPs and Ru2NPs, respectively (Table 2 ; Figure S14, Supporting Information). For the MBC measurements, MBC values were determined to be 64–128 and 64 mg L⁻¹ for Ru1NPs and Ru2NPs against S. aureus, respectively. Due to the promising MIC and MBC values observed for S. aureus, other Gram‐positive bacteria were tested, namely, Staphylococus pseudintermedius LMG 22219 and Enterococcus faecalis ATCC 29212. These bacteria are clinically relevant, causing infections both in animals and humans.^{[ 47} , ^{48 ]} The Ru2NPs exhibited MIC values between 16 and 32 mg L⁻¹ and MBC values of 32–64 mg L⁻¹ against these Gram‐positive bacteria, indicating potential efficacy against a broader range of Gram‐positive bacteria (Table 2).

Table 2.

Results of Minimal Inhibitory Concentration (MIC) and Minimal Bactericidal Concentration (MBC) against RuNPs for Gram‐negative and Gram‐positive bacteria. Concentrations are based on Ru complex content.

Compound	Bacteria	MIC [mg L−1]	MBC [mg L−1]
Ru1NPs	Staphylococcus aureus ATCC 25923	64	n.d.
	Staphylococcus pseudintermedius LMG 22219	16‐32	n.d
	Enterococcus faecalis ATCC 29212	32‐64	n.d.
	Escherichia coli ATCC 25922	64/128	128
	Pseudomonas aeruginosa ATCC 27853	128	512
Ru2NPs	Staphylococcus aureus ATCC 25923	32	64
	Staphylococcus pseudintermedius LMG 22219	16	32
	Enterococcus faecalis ATCC 29212	16	32
	Escherichia coli ATCC 25922	256	512
	Pseudomonas aeruginosa ATCC 27853	512	>512

As observed in the initial susceptibility test with disks for Gram‐negative bacteria, the RuNPs exhibited limited antibacterial activity. The MIC values for Ru1NPs ranged from 64 to 128 mg L⁻¹, while Ru2NPs showed ever higher MIC values from 256 to 512 mg L⁻¹ (Table 2). High MBC values for Gram‐negative bacteria were similarly elevated, ranging from 512 mg L⁻¹ to even higher, suggesting that RuNPs were probably less effective for Gram‐negative bacteria. Notably, Ru1NPs showed a lower MIC value of 128 mg L⁻¹ against E. coli, while Pseudomonas exhibited significant resistance to both Ru compounds, particularly to Ru2NPs (Figure S14, Supporting Information). These results agree with the use of cetrimide, another quaternary ammonium salt, as an inhibitor of some culture media for isolating resistant Pseudomonas aeruginosa while inhibiting other sensitive microorganisms.^{[ 49 ]}

These results showed that our Ru2NPs, synthesized from the Ru‐Benza complex and, to a lesser extent, Ru1NPs, from the Ru‐TOA complex, exhibited good antimicrobial activity against S. aureus and other Gram‐positive bacteria, with no cytotoxicity effects in human cells (Figure S15, Supporting Information). In contrast, both RuNPs displayed lower antibacterial activity against Gram‐negative bacteria. Notably, the value of MIC obtained in our work, particularly that for the Ru2NPs, 32 mg L⁻¹, is comparable to, or even lower than, MIC values reported for other Ru(II) complexes in the literature (Table 3 ), which range from 2 to 100 mg L⁻¹. It is important to remark that Ru(II) complexes with lower MIC values exhibit high cytotoxicity in mammalian cells (IC₅₀ = 5–100 µm, such as [Ru(bb₇)(dppz)]²⁺ and [Ru(bb₇)(Me₂phen)]²⁺) or lack cytotoxicity data altogether (such as [Ru(2,9‐Me₂phen)₂dppz]²⁺ and [Ru(phen)₂(dppz)]²⁺). In contrast, Ru(III) complexes exhibited significantly higher MIC values against S. aureus, such as 400 mg L^{−1[ 22 ]} or 600 mg L^{−1[ 50 ]} (Table 3). Analogously, MBC values in the literature vary from 4 to 170 mg L⁻¹, while the MBC values for Ru2NPs were 32 mg L⁻¹. In this context, our results suggest that encapsulating the Ru complex may enhance its internalization within bacterial cells compared to the Ru(III) complex alone. To evaluate the contribution of individual compounds, control experiments were carried out with the starting salt (RuCl₃), RuCl₄(DMSO)₂‐(DMSOH₂)⁺ and amphiphile I. The MIC values for these compounds were ≥512 mg L⁻¹ for all three tested bacteria, with the exception of the amphiphile I, which exhibited moderate activity against S. aureus (MIC = 128 mg L⁻¹) (Table S1, Supporting Information). This selectivity is likely due to the amino groups on the surface of the micelles, which facilitate interaction with the Gram‐positive bacterial wall. These results support the hypothesis that encapsulation enhances the internalization and antimicrobial efficacy of the Ru(III) complexes, particularly in the RuNP formulations. However, additional studies are underway to further investigate the mechanism of bacterial growth inhibition, particularly whether the metal is internalized by the bacteria and how RuNPs influence cellular redox status.

Table 3.

Comparison of MIC and MBC values for S. aureus with different Ru complexes reported in the literature.

Ru(II) complex	MIC [mg L−1]	MBC [mg L−1]	Refs.
[Ru(bb7)(dppz)]2+	2	4	[51]
[Ru(phen)2(dppz)]2+	4	8‐16	[51]
[Ru(bb7)(Me2phen)]2+	8	64	[51]
[Ru(phen)2dpq]2+	64	128	[52]
[Ru(bpy)2dpqC]2+	32	128	[52]
[Ru(2,9‐Me2phen)2dppz]2+	8	32	[52]
[[22]2(tpphz)]4+	107	182	[53]
[{Ru(5‐Mephen)2}2(tpphz)]4+	165	281	[53]
[{Ru(2,9‐Dimephen)2}2(tpphz)]4+	86	171	[53]
[{Ru(3,4,7,8‐Tetramephen)2}2(tpphz)]4+	73	146	[53]
[{Ru(4,7‐diphenylphen)2}2(tpphz)]4+	44	88	[53]
[Ru(Phen)2(Guanide)(Cl)]Cl2·H2O	400	n.d.	[22]
mer‐[Ru(2‐bimc)3] ⋅ H2O	600	n.d.	[50]
Ru1NPs	64	n.d.	This work
Ru2NPs	32	64	This work

Table 3 shows a comparative analysis of MIC and MBC values for S. aureus between our RuNPs and previously reported Ru complexes. Ruthenium complexes are well known for their antimicrobial activity against Gram‐positive bacteria.^{[ 51 ]} Both mononuclear Ru complexes with dipyrophenazine (dppz)^{[ 52 ]} or dinuclear/oligonuclear compounds with flexible linkers between metal centres^{[ 54 ]} have demonstrated good antibacterial activity against S. aureus. The mechanism does not appear to involve membrane lysis but rather the intercalation of the metal into DNA or its binding to mRNA,^{[ 55 ]} in addition to disrupting the bacterial membrane structure before internalization, which ultimately results in DNA damage.^{[ 53 ]} On its side, Ru compounds complexed with lipolytic ancillary ligands or with the dppz ligand displayed better activity against both Gram‐positive and Gram‐negative strains, due to their effect on membrane structure and DNA binding.^{[ 55} , ^{56 ]}

3. Conclusion

To summarize, we have demonstrated the amphiphile‐assisted synthesis of Ruthenium Nanoparticles, Ru1NPs and Ru2NPs, with potential biomedical applications. These RuNPs are based on hydrophobic Ru(III) complexes, Ru‐TOA and Ru‐Benza, analogs of the NAMI‐A prodrug, with hydrophobic counterions such as tetraocylammonium or benzathine. Oleic acid‐derived amphiphiles I and II self‐assemble around the hydrophobic Ru complexes, forming monodispersed RuNPs that protect them from aqueous degradation. Both RuNPs can be prepared with varying sizes and morphologies by adjusting sonication time, amphiphilic molecule, and the amount and type of the hydrophobic Ru complex. Their stability and Ru release are controlled through PEG‐based molecular staples on the RuNP surface. The stapling process influences Ru release profiles, which depend on the nature of the PEG molecular staples. The PEG1500‐stapled RuNPs exhibit the slowest Ru release, similar to zero‐order kinetics, ideal for drug delivery applications. Additionally, both RuNPs showed good antimicrobial activity against Gram‐positive bacteria, especially Ru2NPs. However, they are not efficient against Gram‐negative bacteria, likely due to the dual‐membrane structure of Gram‐negative cell walls, which may prevent NP penetration. The encapsulation of the Ru complexes as NPs likely enhances cellular uptake in Gram‐positive strains, contributing to their antimicrobial effect. Further experiments are underway to elucidate the precise mechanism of action in Gram‐positive bacteria and the controlled release behavior.

4. Experimental Section

Ru Complexes Synthesis—Synthesis of Ru‐TOA Complex

To a solution of hydrogen trans‐Bis(dimethyl sulfoxide) tetrachlororuthenate (III) (0.36 mmol) in 5 mL of a solvent mixture of H₂O/EtOH (1:4) was added TOABr (0.36 mmol). The mixture was stirred for 30 min, and then 10 mL of H₂O was added. The product was extracted with dicloromethane (DCM) (3 × 20 mL), and the combined organic phases were dried over anhydrous Na₂SO₄. The solution was filtered, and the solvent was evaporated under reduced pressure. The product was obtained as dark orange oil (0.73g, 85% yield). ¹H NMR (300 MHz, DMSO‐d₆): δ (ppm) 3.16 (brs, 8H), 1.58 (brs, 8H), 1.42‐1.12 (m, 40H), 0.94‐0.74 (m, 12H). In CDCl₃, an additional signal at δ (ppm) ‐9.22 (brs, 6H) was observed; this signal was absent in DMSO‐d₆ due to exchange with the deuterated solvent. Elemental analysis calcd for C₃₆H₈₀Cl₄NO₂RuS₂·2.5 Et₂O: C: 52.86, H: 9.85, N: 1.79, found: C: 52.47, H: 10.04, N: 1.34.

Ru Complexes Synthesis—Synthesis of Ru‐Benza Complex

To a solution of hydrogen trans‐Bis(dimethyl sulfoxide) tetrachlororuthenate (III) (0.36 mmol) in 4 mL of EtOH was added benzathine (0.18 mmol). Then, 0.5 mL of HCl solution (1 m) was added. The reaction mixture was stirred for 30 min, and then 10 mL of H₂O was added. The product was collected via vacuum filtration as an orange solid (0.18 g, 47% yield). ¹H NMR (300 MHz, DMSO‐d₆, ppm): δ 8.85 (brs, 4H), 7.51 (s, 10H), 4.32 (brs, 4H). In CDCl₃, an additional signal at δ (ppm) −9.19 (brs, 6H) was observed; this signal was absent in DMSO‐d₆ due to exchange with the deuterated solvent. Elemental analysis calcd for C₂₄H₄₆Cl₈N₂O₄Ru₂S₄·2.5 H₂O: C: 26.55, H: 4.73, N: 2.58 found: C: 26.48, H: 4.57, N: 2.54.

Synthesis of RuNPs

The RuNPs were synthesized by sonication with a probe in a glass vial. Briefly, a 5 mL solution of the corresponding amphiphilic molecule (1 mg mL⁻¹) with a solution of the hydrophobic Ru complex in DCM with the desired amount of Ru complex was sonicated. After the sonication, the RuNPs were left in the fridge for 12 h, and were centrifuged at 1500 rpm for 10 min to get rid of the suspended microcrystals.

Synthesis of Molecular Staples

• Molecular staple 1: To a solution of HO‐PEG1500‐OH (100 mg, 0.067 mmol) in dry tetrahydrofuran (THF) (10 mL) in a 50 mL round‐bottom flask under argon atmosphere was added sodium hydride (4.8 mg, 0.2 mmol) (Scheme S1, Supporting Information). The mixture was stirred for 15 min, and succinic anhydride was added (13.4 mg, 0.134 mmol). After stirring overnight, the reaction was quenched, adding 20 mL of a saturated solution of NaHCO₃, and then was extracted with DCM (3 × 15 mL). The organic layers were dried over anhydrous Na₂SO₄ and filtered. The solvent was evaporated under reduced pressure to obtain 102 mg (0.06 mmol, 90% yield). ¹H‐NMR (300 MHz, MeOD‐d₄): δ (ppm): 4.22‐4.19 (m, 4H), 3.71‐3.55 (PEG), 2.62‐2.57 (m, 4H), 2.49‐2.45 (m, 4H).

• Dimesyl‐PEGn derivatives 2, 3, and 4.^{[ 57} , ⁵⁸ , ^{59 ]}: Molecular Staples 11, 12, and 13. To a solution of HO‐PEGn‐OH (2 mmol) and 500 mmol% of trimethylamine (TEA) in DCM (10 mL) in a 50 mL round‐bottom flask under argon atmosphere was added 250 mmol% of methanesulfonyl chloride was added dropwise at 0 ⁰C (Scheme S2, Supporting Information). After stirring for 15 min, the mixture was heated up to room temperature and stirred overnight. Then, the reaction was quenched by adding 20 mL of a saturated solution of NaHCO₃ and was extracted with DCM (3 × 15 mL). The organic layers were dried over anhydrous Na₂SO₄ and filtered. The solvent was evaporated under reduced pressure to obtain the desired product.

• Diazide‐PEGn derivatives 5, 6, and 7.^{[ 60} , ⁶¹ , ^{62 ]}: To a solution of 2, 3, or 4 (1 mmol) in EtOH (10 mL) was added 250 mmol% of NaN₃. The reaction mixture was refluxed overnight. The solvent was removed under reduced pressure, and the crude was dissolved in DCM (30 mL). The mixture was washed with H₂O (1 × 20 mL) and brine (1 × 20 mL). The organic layer was dried under anhydrous Na₂SO₄ and filtered. The solvent was evaporated under reduced pressure to obtain the desired product.

• Diamine‐PEGn derivatives 8, 9, and 10.^{[ 63} , ^{64 ]}: To a solution of 5, 6, or 7 (1 mmol) in THF (5 mL) was added 250 mmol% of triphenylphosphine. The reaction mixture was stirred for 30 min, and then H₂O (10 mL) was added to the mixture. After stirring overnight, the organic solvent was removed under reduced pressure, and the aqueous solution was acidified to pH 1. Then, the aqueous solution was washed with DCM until the PPh₃ and PPh₃O were completely removed. Next, the aqueous solution was basified with KOH to pH 14, and the product was extracted with DCM (3 × 15 mL). The organic layer was dried over anhydrous Na₂SO₄, filtered, and the solvent was removed under reduced pressure to obtain the desired product.

• Dicarboxilic acid‐PEGn derivatives 11, 12, and 13: To a solution of 8, 9, or 10 (1 mmol) in DCM (5 mL) was added 200 mmol% of succinic anhydride, and the reaction mixture was stirred overnight. The product was filtered or decanted to obtain the desired product. Molecular staple 11: ¹H‐NMR (300 MHz, CDCl₃): δ (ppm): 2.57–2.44 (m, 4H), 1.81–1.62 (m, 12H). Molecular staple 12: ¹H‐NMR (300 MHz, MeOD‐d₄): δ (ppm): 3.66–3.60 (m, 8H), 3.54 (t, J = 5.5 Hz, 4H), 3.36 (t, J = 5.5 Hz, 4H), 2.61–2.53 (m, 4H), 2.50–2.45 (m, 4H). Molecular staple 13: ¹H‐NMR (300 MHz, MeOD‐d₄): δ (ppm): 3.68–3.59 (PEG), 3.54 (t, J = 5.6 Hz, 4H), 3.36 (t, J = 5.6 Hz, 4H), 2.61–2.55 (m, 4H), 2.50‐2.45 (m, 4H).

Synthesis of Molecular Staples—Molecular Staples Conjugation on RuNPs

To a solution of EDC·HCl (0.06 µmol) in 250 µL MES buffer (10 mm, pH 5.5), a solution of NHS (0.195 µmol) in MES (250 µL) and a solution of the molecular staple (0.11 µmol) in MES (500 µL) were added. The mixture was stirred with an orbital shaker at 190 rcf for 30 min. The mixture was then added to a suspension of RuNPs (1.39 mg in Ru) in MES (1 mL), and stirred with an orbital shaker at 190 rcf for 1.5 h. After this time, the suspension was purified using centrifuge filters, washing with MES buffer (3 × 2 mL). The RuNPs were resuspended in H₂O (2 mL).

Synthesis of Molecular Staples—In vitro Release Studies of RuNPs

The Ru release studies were carried out in a dialysis membrane with a molecular weight cut‐off of 50 kDa. The corresponding solution of RuNPs (2 mL) in water transferred into a dialysis bag was dialyzed under magnetic stirring against 2 L of H₂O‐MilliQ or PBS buffer at room temperature. After a defined time lapse, an aliquot of 200 µL was taken from the dialysis bag, and the Ru concentration was determined using ICP‐MS. The measurements were performed in triplicate.

Microbiological Studies

Bacteria were replicated on plates of tryptone soy agar (TSA) and incubated at 37 °C for 24 h. For liquid cultures, a loop of a bacterial colony was cultivated in 3 mL of tryptone soy broth (TSB) for 24 h at 37 °C and 150 rpm. Before starting the experiments, the optical density at 600 nm of the cultures was determined and adjusted to 1.0 with sterile TSB.

Microbiological Studies—Preliminary Observation of Susceptibility Toward RuNPs by Disk Test

To assess whether the bacterial growth would be inhibited by the developed RuNPs, a preliminary study by disk susceptibility test was conducted according to CLSI (2015). Sterile paper blank disks (Oxoid, ThermoFischer, UK) were loaded with 100 µg of the different RuNPs, based on the Ru complex content, dissolved in 20 µL and allowed to dry at room temperature (in triplicate each). Independent cultures of the three bacteria were grown overnight in 3 mL of TSB (tryptone soy broth) at 37 °C and 200 rpm. The optical density at 600 nm was adjusted to 1 with sterile TSB. Plates of Müeller‐Hinton agar (MHA) were inoculated with the cultures three times using sterile swabs and allowed to dry in a biosecurity cabin for 10 min. The disks were applied on the surface of the inoculated agar with the help of sterile tweezers. Plates were sealed and incubated at 37 °C for 24 h. After incubation, plates were inspected to see the formation of growth inhibition halos around the disks, and the diameter of the halos was recorded.

Microbiological Studies—Determination of MIC and MBC

The determination of the MIC was done by the microdilution technique in 96‐well microtitre plates in Müeller‐Hinton broth (MHB).^{[ 65 ]} Serial dilutions of the compounds were prepared in MHB following a 2‐base logarithmic gradient of concentrations from 512 to 0.5 mg L⁻¹. A column of the plate was reserved for MHB without RuNPs as a control. Once prepared, the wells were inoculated with 5 µL of the bacterial cultures (optical density at 600 nm adjusted to 1). A row of the plate was kept without inoculation as a control of proper sterility. Three replicates were done for every bacterium. The plates were sealed around with tape and incubated at 37 °C for 24 h. After incubation, the wells were visually inspected for turbidity. The MIC was considered the concentration of the first well for which turbidity was not observed.

For the determination of MBC, 100 µL of the content of wells was streaked on TSA plates and the plates were incubated at 37 °C for 24 h. After incubation, the plates were inspected for the appearance of colonies. The MBC was considered the minimal concentration of the well whose corresponding plate gave no colonies.

Cytotoxicity Assays

In short, the HFF‐1 cells were plated at a density of 1 × 10⁴ cells per well in a 96‐well plate at 37 °C in 5% CO₂ atmosphere (200 µL per well, number of repetitions = 3). After 24 h of culture, the medium in the wells was replaced with fresh medium containing RuNPs at varying concentrations, ranging from 8 to 256 µg L⁻¹. After 24 h, the supernatant of each well was replaced by 100 µL of fresh medium with 3‐[4,5‐dimethylthiazol‐2‐yl]‐2,5‐diphenyl tetrazolium bromide (MTT) (1 mg mL⁻¹). After 2 h of incubation at 37 °C and 5% CO₂, the medium was removed, the formazan crystals were solubilized with 100 µL of DMSO, and the solution was mixed to dissolve the reacted dye. The absorbance of each well was read on a microplate reader (Dynatech MR7000 instruments) at 570 nm. The relative cell viability (RCV) and its associated standard deviation, calculated with respect to control wells containing cell culture medium without RuNPs, were determined using the following equations:

$$RCV\left(\%\right)=\frac{Ab{s}_{sample}-Ab{s}_{Pos.Control}}{Ab{s}_{Neg.Control}-Ab{s}_{Pos.Control}}\times 100$$ (2)

$${\sigma }_{RCV}\left(\%\right)=RV{C}_{sample}\times \sqrt{{\left(\frac{{\sigma }_{sample}}{Ab{s}_{Sample}}\right)}^2+{\left(\frac{{\sigma }_{Neg.Control}}{Ab{s}_{Neg.Control}}\right)}^2}$$ (3)

Statistical Analysis

All experiments were performed in triplicate (n = 3) unless otherwise stated. Data were expressed as mean ± standard deviation (SD). Statistical analysis was performed using Jamovi software (version 2.6.44) with the Mann–Whitney U statistical test. Differences were considered statistically significant at p < 0.05.

Conflict of Interest

The authors declare no conflict of interest.

Author Contributions

R.G.F. and M.P.L. conceived the idea and designed the experiments. R.G.F. synthesized and characterized the RuNPs. M.E.B. characterized the Ru complexes. E.P. carried out the bacterial tests. M.L.G.M. performed the cellular viability. M.P.L. supervised the work and coordinated the contributions. The manuscript was written through the contributions of all authors. R.G.F., M.E.B., E.P., I.F., M.L.G.M., and M.P.L. have given approval to the final version of the manuscript.

Supporting information

Supporting Information

Gimeno‐Ferrero R., Estruch‐Blasco M., Pajuelo E., Fernández I., García‐Martín M. L., and Pernia Leal M., “Amphiphile‐Assisted Synthesis of Ruthenium Nanoparticles for Controlled Release and Enhanced Antibacterial Activity.” Small Methods 9, no. 12 (2025): e02207. 10.1002/smtd.202502207

Data Availability Statement

The data that support the findings of this study are available in the supplementary material of this article.