Abstract

   Mild‐heat photothermal antibacterial therapy avoids heat‐induced damage
   to normal tissues but causes bacterial tolerance. The use of
   photothermal therapy in synergy with chemodynamic therapy is expected
   to address this issue. Herein, two pseudo‐conjugated polymers P^M123
   with photothermal units and P^Fc with ferrocene (Fc) units are designed
   to co‐assemble with DSPE‐mPEG[2000] into nanoparticle NP^M123/Fc.
   NP^M123/Fc under 1064 nm laser irradiation (NP^M123/Fc+NIR‐II)
   generates mild heat and additionally more toxic ∙OH from endogenous
   H[2]O[2], displaying a strong synergistic photothermal and chemodynamic
   effect. NP^M123/Fc+NIR‐II gives >90% inhibition rates against MDR
   ESKAPE pathogens in vitro. Metabolomics analysis unveils that
   NP^M123/Fc+NIR‐II induces bacterial metabolic dysregulation including
   inhibited nucleic acid synthesis, disordered energy metabolism,
   enhanced oxidative stress, and elevated DNA damage. Further,
   NP^M123/Fc+NIR‐II possesses >90% bacteriostatic rates at infected
   wounds in mice, resulting in almost full recovery of infected wounds.
   Immunodetection and transcriptomics assays disclose that the
   therapeutic effect is mainly dependent on the inhibition of
   inflammatory reactions and the promotion of wound healing. What is
   more, thioketal bonds in NP^M123/Fc are susceptible to ROS, making it
   degradable with highly favorable biosafety in vitro and in vivo.
   NP^M123/Fc+NIR‐II with a unique synergistic antibacterial strategy
   would be much less prone to select bacterial resistance and represent a
   promising antibiotics‐alternative anti‐infective measure.

   Keywords: acute wound infections, biodegradable polymer, chemodynamic
   therapy, ESKAPE pathogens, photothermal therapy
     __________________________________________________________________

   NP^M123/Fc under 1064 nm laser irradiation (NP^M123/Fc+NIR‐II)
   generates mild heat and additionally more toxic ∙OH from endogenous
   H[2]O[2], displaying a synergistic effect of PTT and CDT for
   anti‐infection. Thioketal bonds in NP^M123/Fc are susceptible to ROS,
   making it degradable with highly favorable biosafety. NP^M123/Fc+NIR‐II
   representes a biocompatible and broad‐spectrum nanoplatform for
   antibiotics‐alternative treatment of superficial infections.

   graphic file with name ADVS-11-2309624-g004.jpg

1. Introduction

   With the wide use and abuse of antibiotics to prevent and treat
   bacterial infections, antibiotic resistance is occurring increasingly.
   Particularly, ESKAPE pathogens including Enterococcus faecium (Ec.
   faecium), Staphylococcus aureus (S. aureus), Klebsiella pneumoniae (K.
   pneumoniae), Acinetobacter baumannii (A. baumannii), Pseudomonas
   aeruginosa (P. aeruginosa), and Enterobacter species (especially Eb.
   cloacae and Eb. hormaechei) are commonly characteristic of multidrug
   resistance (MDR), and they represent the leading cause of refractor
   infections in hospital settings throughout the world, leading to higher
   medical costs, prolonged hospital stays, and increased mortality.^[
   [40]^1 ^]

   Contemporary antibiotics‐alternative therapeutic strategies such as
   nanotechnology are potential options for combating MDR bacteria
   including ESKAPE pathogens.^[ [41]^2 ^] Various organic or inorganic
   nanomaterials have the ability to evade existing mechanisms of acquired
   drug resistance, taking advantages such as bare selection for
   resistance, minimal invasion, and low host toxicity.^[ [42]^3 ^] In
   addition, the unique physico–chemical properties (such as size, shape,
   and surface chemistry) of nanomaterials render the elevated targeting
   attack pathogens.^[ [43]^3 ^] Currently, nanotechnology‐based
   antibacterial agents mainly rely on four major principles of action,
   namely sonodynamic therapy (SDT), photodynamic therapy (PDT),
   chemodynamic therapy (CDT), and photothermal therapy (PTT). SDT
   utilizes senosensitizers to produce reactive oxygen species (ROS) from
   surrounding oxygen under low‐intensity ultrasound, killing bacteria
   through the synergistic action of ROS and ultrasound.^[ [44]^4 ^]
   Likewise, PDT employs photosensitizers that can transfer photon energy
   to surrounding oxygen to produce ROS.^[ [45]^5 ^]

   PTT makes use of photothermal agents to absorb energy from the
   near‐infrared region I and II (NIR‐I and NIR‐II) light for the
   production of heat.^[ [46]^6 , [47]^7 , [48]^8 ^] However, a high local
   temperature of up to 70 °C is required to kill bacteria completely if
   PTT is used alone.^[ [49]^9 ^] This will bring pain to the patients and
   even damage to surrounding tissues.^[ [50]^9 ^] On the contrary, the
   use of mild heat (commonly ≤ 48 °C in vivo) in PTT can avoid
   heat‐induced damage of normal tissues,^[ [51]^10 ^] but mild heat will
   lead to bacterial tolerance due to inducible expression of heat shock
   proteins.^[ [52]^11 ^] Combining PTT with another different strategy
   (e.g., CDT,^[ [53]^12 ^] PDT,^[ [54]^13 ^] or quaternary phosphonium^[
   [55]^14 ^]) could address the above issues and, in this case,
   CDT‐generated hydroxyl radicals (∙OH), PDT‐generated ROS or cationic
   quaternary phosphonium agent acts as a synergistic factor of
   PTT‐generated mild heat, endowing improved broad‐spectrum antibacterial
   effects.

   CDT is applied to convert hydrogen peroxide (H[2]O[2]) into more toxic
   ∙OH via a Fenton or Fenton‐like reaction.^[ [56]^15 ^] In this context,
   Ferrocene (Fc) has attracted the interest in developing nanomaterials
   to convert endogenous abundant H[2]O[2] at infection sites to
   participate in Fenton‐like reaction, where H[2]O[2] oxidizes Fc into
   ferrocenium (Fce) and meanwhile itself is catalyzed into ∙OH.^[ [57]^16
   , [58]^17 , [59]^18 ^]

   Herein, we developed two biodegradable polymers namely P^M123 and P^Fc,
   which were assembled with commercial DSPE‐mPEG[2000] to generate the
   nanoparticle NP^M123/Fc (Scheme [60]1 ). P^M123 contained the NIR‐II
   polymer photothermal agent M123 plus the ROS‐sensitive thioketal bonds,
   while P^Fc harbored Fc as well as the thioketal bonds. NP^M123/Fc under
   1064 nm laser irradiation (NP^M123/Fc+NIR‐II) exhibited M123‐mediated
   photothermal activity and Fc‐dependent chemodynamic effect, which
   worked in a synergistic manner. NP^M123/Fc+NIR‐II was highly efficient
   in inhibiting MDR ESKAPE pathogens in vitro and in treating infected
   wounds in a mouse model. NP^M123/Fc was degradable upon ROS, and it
   would be highly degradable, rapidly cleared, and hypotoxic in the body.
   NP^M123/Fc+NIR‐II with a unique two‐way synergistic antibacterial
   strategy would be much less prone to select bacterial resistance and
   represented a promising measure for antibiotics‐alternative therapy.

Scheme 1.

   Scheme 1
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   Schematic illustration of development of NP^M123/Fc for treatment of
   acute wound infections. a) The polymer P^M123 was synthesized from the
   four monomers namely M1, M2, M3, and M4, among which M1, M2, M3
   constituted the photothermal agent M123 while M4 contained the
   ROS‐sensitive thioketal bond. b) The polymer P^Fc was synthesized from
   another four monomers namely M5 (Fc), M6 (containing thioketal bond),
   M7, and M8 (mPEG). c) P^M123, P^Fc, and DSPE‐mPEG[2000] were
   self‐assembled together into nanoparticle NP^M123/Fc. d) NP^M123/Fc
   under 1064 nm laser irradiation (NP^M123/Fc+NIR‐II) generates mild heat
   and additionally more toxic ∙OH from endogenous H[2]O[2], displaying a
   synergistic effect of PTT and CDT for anti‐infection. The thioketal
   bonds in NP^M123/Fc are susceptible to ROS, making it degradable with
   highly favorable biosafety in vitro and in vivo. NP^M123/Fc+NIR‐II
   represented a biocompatible and broad‐spectrum nanoplatform for
   antibiotics‐alternative treatment of superficial infections.

2. Results and Discussion

2.1. Preparation and Characterization of Polymers and Nanoparticles

   The present study involved three different polymers (i.e., P^M123,
   P^Fc, and DSPE‐mPEG[2000]) and three distinct nanoparticles (i.e.,
   NP^M123, NP^Fc, and NP^M123/Fc) as shown in Scheme [62]1 and Scheme
   [63]S1 (Supporting Information). P^M123 (Scheme [64]1a) was synthesized
   from the four monomers namely M1, M2, M3, and M4. M1 was the linker
   responsible for linking M2, M3 and M4 to form P^M123. M2 mainly endowed
   P^M123 with photothermal properties. M3 caused the UV absorption
   wavelength of P^M123 to be redshifted and thus made P^M123 excitable at
   1064 nm. The M123 moiety as a whole could be recognized as a polymer
   photothermal agent. M4 contained the ROS‐sensitive thioketal bond,
   conferring the degradability of P^M123. P^Fc (Scheme [65]1b) was
   synthesized from another four monomers namely M5, M6, M7, and M8. M5
   was Fc. M6 was the linker responsible for linking M5, M7, and M8 to
   form P^Fc. M7 contained the ROS‐sensitive thioketal bond, conferring
   the degradability to P^Fc. Since M5, M6, and M7 were all hydrophobic,
   M8 (mPEG) was added to render the amphipathicity of P^Fc. The
   successful synthesis of P^M123 (Figure [66]S1, Supporting Information)
   or P^Fc (Figure [67]S2, Supporting Information) was validated by ^1H
   NMR (nuclear magnetic resonance) spectra. For P^M123, the peaks at 0.5
   to 2 ppm could be attributed to the alkane chain, the peaks at 7 to
   9.5 ppm could be attributed to the hydrogen of thiophene, and the peaks
   at 2.5 to 4.5 ppm could be attributed to the methylene group of
   ROS‐sensitive thioketal bond (Figure [68]S1, Supporting Information).
   For P^Fc, the peaks at 7 to 8 ppm belonged to Fc and the peak at
   1.5 ppm belonged to the methyl of thioketal bond (Figure [69]S2,
   Supporting Information), indicating that P^Fc contained Fc and
   ROS‐sensitive moieties. Hydrophobic P^M123 or amphipathic P^Fc was
   subsequently self‐assembled with amphipathic DSPE‐mPEG[2000],
   generating NP^M123 or NP^Fc, respectively (Scheme [70]S1, Supporting
   Information). P^M123, P^Fc, and DSPE‐mPEG[2000] were self‐assembled
   together into NP^M123/Fc (Scheme [71]1c).

   As determined by transmission electron microscopy (TEM) and scanning
   electron microscopy (SEM), NP^M123, NP^Fc, and NP^M123/Fc displayed
   uniform spherical morphology with average diameters at about 106, 108,
   and 120 nm, respectively (Figure [72] 1a; Figures [73]S3 and [74]S4,
   Supporting Information). Energy dispersive X‐ray spectral scanning
   (EDXS) showed that there were the key elements C, N, O, Fe and S in
   NP^M123/Fc (Figure [75]1b), and the presence of Fe confirmed the
   successful doping of Fc into NP^M123/Fc.

Figure 1.

   Figure 1
   [76]Open in a new tab

   Characterization of NP^M123/Fc. a) TEM images of NP^M123/Fc. Scale bar
   is 100 nm. b) STEM and EDXS images of NP^M123/Fc. Scale bar is 200 nm.
   c) UV–vis–NIR absorption spectra of NP^M123, NP^Fc, and NP^M123/Fc. d)
   Time–concentration–temperature curves of NP^M123/Fc under 1064 nm laser
   irradiation (1.0 W cm^−2, 5 min). e) Time‐dependent heating and cooling
   curve of 100 µg mL^−1 NP^M123/Fc under 1064 nm laser irradiation
   (1.0 W cm^−2, 11 min). The 𝜂 value was calculated, through linear
   regression, from this heating and cooling curve. f) Time–temperature
   curve of four cycles of heating and cooling of NP^M123/Fc under 1064 nm
   laser irradiation (1.0 W cm^−2, 5 min). g) Absorption curves of MB as a
   selective trapping agent to detect ∙OH in the reaction mixture of
   NP^M123/Fc+H[2]O[2] or NP^M123+H[2]O[2] after different incubation
   times. h) Trends of UV absorption at 410 nm as indicated by the above
   MB‐based detection results. DLS detection of i) diameter sizes, j)
   PDIs, and k) average Zeta potentials of NP^M123/Fc in the presence and
   absence of H[2]O[2]. l) Schematic illustration of H[2]O[2]‐mediated
   degradability, M123‐dependent NIR‐II photothermal activity, and
   Fc‐dependent chemodynamic activity of NP^M123/Fc in vitro.

   UV–vis–NIR absorption spectra denoted that each of NP^M123 and
   NP^M123/Fc had two major absorption peaks at 740 and 1064 nm,
   indicating that these nanoparticles had absorption extended in NIR‐II
   (Figure [77]1c). In order to investigate the photothermal properties,
   NP^M123 or NP^M123/Fc at 0 to 200 µg mL^−1 in the aqueous solution was
   irradiated with a 1064 nm laser at 1 W cm^−2 for 5 min, and the
   time‐temperature curves were recorded (Figure [78]1d; Figure [79]S5,
   Supporting Information). The heating process became faster with the
   increasing of NP^M123/Fc concentrations, and 100 µg mL^−1 NP^M123/Fc
   (used for the following in vitro and in vivo antibacterial assays) gave
   a temperature increase to 50 °C under laser irradiation for 3 min
   (Figure [80]1d; Figure [81]S5, Supporting Information). NP^M123 showed
   similar results (Figure [82]S6, Supporting Information). The
   photothermal conversion efficiency (𝜂) of NP^M123 and NP^M123/Fc were
   calculated as 38.16% and 37.8% respectively (Figure [83]1e; Figure
   [84]S7, Supporting Information), which were comparable to the majority
   of reported polymer‐based NIR‐II photothermal nanomaterials.^[ [85]^19
   , [86]^20 ^] After four cycles of heating (irradiation) and cooling,
   100 µg mL^−1 NP^M123/Fc gave the nearly same temperature elevation to
   about 60 °C in each cycle and the almost constant cooling rates at
   about 3.2 °C min^−1 for all cycles (Figure [87]1f). NP^M123 showed
   similar results (Figure [88]S8, Supporting Information). Therefore,
   NP^M123/Fc containing the photothermal agent M123 had an efficient and
   stable NIR‐II photothermal effect in vitro upon 1064 nm laser
   irradiation.

   Given the participation of Fc and H[2]O[2] in a Fenton‐like reaction,^[
   [89]^16 , [90]^17 , [91]^18 ^] 100 µg mL^−1 NP^M123 or NP^M123/Fc was
   mixed with 1 mm H[2]O[2] and 2 mm 1,3‐diphenylisobenzofuran (DPBF),
   followed by incubation for 0–3.5 min, to test consumption of H[2]O[2]
   (Figures [92]S9 and [93]S10, Supporting Information). DPBF with an
   absorption peak at 410 nm was used herein as a selective trapping agent
   of ROS including H[2]O[2] and ∙OH.^[ [94]^21 ^] With the prolonging of
   incubation times, the absorption peaks as detected by SpectraMax
   Microplate Reader were decreased for NP^M123/Fc, NP^M123 and NP^Fc
   (Figure [95]S9, Supporting Information), but the absorbance of
   NP^M123/Fc or NP^Fc at 410 nm decreased faster than that of NP^M123
   (Figure [96]S10, Supporting Information); therefore, both NP^M123/Fc
   and NP^Fc (but not NP^M123) would produce other ROS from H[2]O[2], and
   Fc played a role in ROS production in NP^M123/Fc. To test the
   production of ∙OH from H[2]O[2], the ∙OH‐specific trapping agent
   methylene blue (MB) was used to determine its absorption peak at
   664 nm; with the prolonging of incubation times, NP^M123/Fc or NP^Fc
   gave rapidly decreased absorption peaks, but NP^M123 gave almost
   unchanged absorption peaks (Figure [97]1g; Figure [98]S11, Supporting
   Information). Moreover, the absorbance of NP^M123/Fc or NP^Fc at 664 nm
   decreased faster than that of NP^M123 (Figure [99]1h; Figure [100]S11,
   Supporting Information). These results indicated that NP^M123/Fc or
   NP^Fc (but not NP^M123) produced ∙OH from H[2]O[2], which was dependent
   on Fc. The Fc release kinetics of NP^M123/Fc in vitro was measured by
   ICP‐MS‐based detection of dissociated Fc; as shown in Figure [101]S12
   (Supporting Information), there was a gradual increase in Fc release
   from 0 to 48 h whether there was the presence of 10 mm H[2]O[2] or not,
   but the detected Fc release rate 76.48% in the presence of 10 mm
   H[2]O[2] at 48 h was much higher the counterpart value (23.31%) in the
   presence of PBS. Therefore, Fc could be effectively released from
   NP^M123/Fc in the presence of H[2]O[2]. To test the oxidation of Fc,
   1 mg mL^−1 P^Fc was incubated with 1 mm H[2]O[2] at 37 °C for 24 h, and
   then X‐ray photoelectron spectroscopy (XPS) revealed that the addition
   of H[2]O[2] made the change of iron valence state from +2 into +3
   (Figure [102]S13, Supporting Information). Therefore, NP^M123/Fc had
   the chemodynamic effect in vitro to produce ∙OH from H[2]O[2] and
   meanwhile to oxidize Fc into Fce.

   To test sensitivity to H[2]O[2] in vitro, NP^M123, NP^Fc, and
   NP^M123/Fc were incubated with 1 mm H[2]O[2] for 3.5 min. TEM and SEM
   disclosed nearly complete disintegration of nanoparticles (Figures
   [103]S14 and [104]S15, Supporting Information). As determined by
   dynamic light scattering (DLS), the average diameter sizes
   (Figure [105]1i) and the polydispersity indexes (PDIs, Figure [106]1j)
   of NP^M123/Fc were changed from 123.07 to 2531.33 nm (giving a huge
   fold increase of 20.57) and from 0.16 to 0.93 (approaching nearly 1^[
   [107]^22 ^]), respectively, and moreover the average Zeta potentials of
   NP^M123/Fc were changed from −31.08 mV to + 7.22 mV (Figure [108]1k).
   In addition, NP^M123 and NP^Fc gave similar DLS‐detected results
   (Figure [109]S16, Supporting Information). Therefore, the ROS‐sensitive
   thioketal bonds in NP^M123, NP^Fc, and NP^M123/Fc were degradable upon
   H[2]O[2] in vitro and then these nanoparticles were disintegrated in
   the solutions; as the degradation of NP^Fc or NP^M123/Fc, Fc was
   released in the solution and then oxidized by H[2]O[2] into positively
   charged Fce, thereby causing the change in overall Zeta potentials from
   negative charge into positive. In addition, the degradation of NP^M123
   or NP^M123/Fc was along with the release of photothermal agent M123.

   Taken together, as determined by comprehensive in vitro
   characterizations, NP^M123/Fc was degradable upon H[2]O[2] and,
   moreover, it exhibited the photothermal activity to produce mild heat
   from NIR‐II laser energy and also the Fc‐dependent chemodynamic
   activity to produce ∙OH from H[2]O[2] (Figure [110]1l).

2.2. Antibacterial Efficacy Against ESKAPE Pathogens In Vitro

   The in vitro antibacterial efficacies of NP^M123, NP^Fc, and NP^M123/Fc
   were measured by using plate counting method with clinical MDR ESKAPE
   isolates as model pathogens, and a total of eight treatment groups
   namely FF1 (Mock), FF2 (NIR‐II), FF3 (NP^M123), FF4 (NP^M123+NIR‐II),
   FF5 (NP^Fc), FF6 (NP^Fc+NIR‐II), and FF7 (NP^M123/Fc), and FF8
   (NP^M123/Fc+NIR‐II) was set (Figure [111] 2 ). 1 mm H[2]O[2] was added
   for all in vitro antibacterial experiments, which was needed to degrade
   NP^M123, NP^Fc, and NP^M123/Fc to release M123 or/and Fc for subsequent
   bactericidal actions. No significant inhibition effects on all six
   ESKAPE bacteria (Figure [112]2a–f) were found for the five scheduled
   control groups FF1, FF2, FF3, FF5, and FF7, confirming that each
   nanoparticle along with 1 mm exogenous H[2]O[2] had very limited
   bactericidal activity in the absence of laser irradiation. FF4 (PTT
   alone) had a higher bactericidal activity relative to FF6 (CDT alone);
   notably, when PTT and CDT worked together (FF8), the highest
   antibacterial efficacy against both Gram‐positive (Figure [113]2a,b)
   and Gram‐negative (Figure [114]2c–f) ESKAPE bacteria were detected with
   inhibition rates at 97.79%, 91.42%, 92.82%, 97.6%, 94.54%, and 93.53%
   for Ec. faecium, S. aureus, K. pneumoniae, A. baumannii, P. aeruginosa,
   and Eb. hormaechei, respectively.

Figure 2.

   Figure 2
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   Antibacterial efficacy against ESKAPE pathogens in vitro. FF1: Mock,
   FF2: NIR‐II, FF3: NP^M123, FF4: NP^M123+NIR‐II, FF5: NP^Fc, FF6:
   NP^Fc+NIR‐II, FF7: NP^M123/Fc, FF8: NP^M123/Fc+NIR‐II. NIR‐II
   represents 1064 nm laser irradiation (1.0 W cm^−2, 3.5 min). a–f)
   Representative photos of plate count agars for ESKAPE pathogens post
   treatment, and corresponding statistical analysis of bacterial counts.
   Data are presented as mean ± SD (n = 6). ns: p ≥ 0.05, *: p < 0.05, **:
   p < 0.01, ***: p < 0.001. g) Representative CLSM images (n = 3) of
   live/dead bacteria post treatment. Red and green fluorescence signals
   correspond to dead and live bacterial cells, respectively. Scale bar is
   100 µm. 1 mm H[2]O[2] are added for all experiments.

   P. aeruginosa and S. aureus were then selected as the representatives
   of Gram‐negative and Gram‐positive ESKAPE pathogens, respectively, to
   investigate whether FF8 had the PTT/CDT synergistic effect to combat
   bacteria. First, the synergistic factor Q values (see Supporting
   Information for computational formula and evaluation criterion) were
   calculated as 1.24 and 1.23 for P. aeruginosa and S. aureus,
   respectively. These two Q values were larger than 1.15 which was
   commonly used as a cutoff indicator of “strong synergy.”^[ [116]^23 ^]
   Second, staining with live/dead double fluorescent dyes
   SYTO‐9/propidium iodide (PI) and imaging with confocal laser scanning
   microscopy (CLSM) were carried out and the results indicated that, for
   either P. aeruginosa or S. aureus (Figure [117]2g), a few red
   fluorescences (dead) and abundant green fluorescence (live) were
   observed for bacterial cells treated with FF6, while abundant red
   fluorescence (dead) and a few green fluorescence (live) were observed
   for bacterial cells treated with FF4; by contrast, nearly all bacterial
   cells treated with FF8 gave red fluorescence (dead), further confirming
   the synergy.

   Taken together, NP^M123/Fc employed 1064 nm laser‐triggered
   photothermal effect and Fc‐dependent chemodynamic effect in a
   synergistic manner, achieving the excellent bactericidal efficacy
   against MDR ESKAPE pathogens in vitro.

2.3. Bacterial Morphological and Biochemical View of Antibacterial Mechanism
In Vitro

   The underlying mechanism of antibacterial action in vitro was explored
   with respective to bacterial morphological and biochemical changes
   using MDR S. aureus and P. aeruginosa as representatives by setting the
   abovementioned eight treatment groups (Figure [118] 3 ). First, SEM
   (Figure [119]3a) and TEM (Figure [120]S17, Supporting Information) was
   employed to observe the integrity of bacterial morphology and cell
   membrane after treatment: i) NP^M123/Fc+NIR‐II resulted in very severe
   morphological wrinkling and cell membrane disruption; ii)
   NP^M123+NIR‐II induced relatively lower levels of wrinkling and
   disruption; and iii) only slight wrinkling and concavity on bacterial
   cell surfaces was observed for the those treated with NIR‐II and no
   changes could be observed for those treated with the remaining 5
   treatment groups. The damage to bacterial cell membrane by
   nanoparticles would lead to the leakage of bacterial cytoplasmic
   components.^[ [121]^23 ^] Second, the contents of DNA, proteins, K^+,
   and Na^+ in the supernatants of P. aeruginosa (Figure [122]3b–e) and S.
   aureus (Figure [123]3f–i) suspensions after treatment. The results
   showed that the largest amounts of DNA, proteins, K^+, and Na^+ were
   detected for those treated with NP^M123/Fc+NIR‐II, denoting that
   NP^M123/Fc+NIR‐II resulted in the most severe bacterial damage. Third,
   CLSM (Figure [124]3j) was utilized to characterize DNA damage by
   staining bacterial DNA with 4′,6‐diamidino‐2‐phenylindole (DAPI) dye
   with blue fluorescence: i) only a little fluorescence intensity was
   observed for those treated with NP^M123/Fc+NIR‐II, indicating the
   occurrence of most severe DNA damage; and ii) bright fluorescence could
   be observed for all the other treatment groups, among which the
   fluorescence intensity of those treated with NP^M123+NIR‐II was much
   lower than the remaining six treatment groups. Taken together,
   NP^M123/Fc+NIR‐II exhibited the best efficacy in damaging bacterial
   cell membranes (causing leakage of cytoplasmic components) and
   meanwhile inducing bacterial DNA damage.

Figure 3.

   Figure 3
   [125]Open in a new tab

   Bacterial morphological and biochemical view of antibacterial mechanism
   in vitro. FF1: Mock, FF2: NIR‐II, FF3: NP^M123, FF4: NP^M123+NIR‐II,
   FF5: NP^Fc, FF6: NP^Fc+NIR‐II, FF7: NP^M123/Fc, FF8: NP^M123/Fc+NIR‐II.
   NIR‐II represents 1064 nm laser irradiation (1.0 W cm^−2, 3.5 min). a)
   SEM images of P. aeruginosa and S. aureus post treatment. Scale bar is
   500 nm. Leakage of b) DNA, c) protein, d) K^+, and e) Na^+ by P.
   aeruginosa post treatment. Leakage of f) DNA, g) protein, h) K^+, and
   i) Na^+ by S. aureus post treatment. j) CLSM images of DAPI‐stained
   bacterial DNAs post treatment. Scale bar is 1 µm. Data are presented as
   mean ± SD (n = 6). ns: p ≥ 0.05, *: p < 0.05, **: p < 0.01, ***:
   p < 0.001. 1 mm H[2]O[2] are added for all experiments.

2.4. Bacterial Metabolomic View of Antibacterial Mechanism In Vitro

   To characterize bacterial metabolomic response after treatment in
   vitro, a metabolomics analysis was performed for P. aeruginosa treated
   with NP^M123/Fc+NIR‐II in relative to Mock (Figure [126] 4 ). First,
   the principal component analysis (PCA) indicated the datasets of these
   two groups were lasted at two different clusters, indicating the
   existence of obvious parallelism within each group but high‐level
   differences between the two groups (Figure [127]4a). Second, a total of
   116 differentially regulated metabolites (DRMs) were identified for
   NP^M123/Fc+NIR‐II compared Mock (Figure [128]S18, Supporting
   Information), and they could be divided into 62 up‐regulated
   metabolites and 54 down‐regulated ones (Figure [129]4b,c). Third, as
   shown by Kyoto encyclopedia of genes and genomes (KEGG) enrichment
   analysis, there were the four enriched up‐regulated metabolic pathways
   (i.e., biosynthesis of secondary metabolites, arginine and proline
   metabolism, aminobenzoate degradation, and nicotinate and nicotinamide
   metabolism), and also the nine enriched down‐regulated metabolic
   pathways especially including pyrimidine metabolism, β‐alanine
   metabolism, alanine, aspartate and glutamate metabolism, glutathione
   metabolism, and purine metabolism (Figure [130]4d). Fourth, the major
   DRMs and related pathways were mapped, aiming to interpret bacterial
   metabolic dysregulation (Figure [131]4e): i) purine metabolism‐related
   GTP, dGMP, and deoxyguanosine were down‐regulated, thereby obstructing
   DNA synthesis and causing DNA damage^[ [132]^24 ^]; ii) pyrimidine
   metabolism CMP and cytosine were down‐regulated, hence hampering RNA
   synthesis^[ [133]^25 ^]; iii) riboflavin metabolism‐related riboflavin
   were up‐regulated, denoting over‐production of ROS^[ [134]^26 ^]; iv)
   glutathione metabolism‐related putrescine and NADP^+ were up‐regulated,
   consequently leading to enhancement of oxidative stress; and v) carbon
   metabolism‐related succinate and O‐acetyl‐L‐serine were down‐regulated,
   therefore resulting in disturbance of energy metabolism.^[ [135]^27 ,
   [136]^28 ^] In addition, as revealed by Receiver Operating
   Characteristic (ROC) curve analysis (Figure [137]S19, Supporting
   Information), putrescine gave an AUC (area under the ROC curve) value
   of 0.990 (> the cutoff 0.9^[ [138]^29 ^]); therefore, the up‐regulation
   of putrescine (as a biomarker) denoted the pathological state in P.
   aeruginosa after treatment. Taken together, NP^M123/Fc+NIR‐II induced
   inhibited nucleic acid synthesis, disordered energy metabolism,
   enhanced oxidative stress, and elevated DNA damage in bacteria.

Figure 4.

   Figure 4
   [139]Open in a new tab

   Metabolomics analysis of P. aeruginosa after treatment. a) PCA plot of
   clustered datasets between the two treatment groups NP^M123/Fc+NIR‐II
   (FF8) and Mock (FF1). NIR‐II represents 1064 nm laser irradiation
   (1.0 W cm^−2, 3.5 min). b) Volcano plot of DRMs identified for
   NP^M123/Fc+NIR‐II compared to Mock; Up: Up‐regulated metabolites. Down:
   Down‐regulated metabolites. c) Heatmaps of major DRMs. d) KEGG pathway
   enrichment analysis of DRMs. e) Diagrams of major DRMs and related
   pathways. Red represents up‐regulation and blue stands for
   down‐regulation. Experiments were performed with five independent
   biological replicates. 1 mm H[2]O[2] are added for all experiments.

2.5. Therapeutic Efficacy Against Infected Wounds in Mice

   Endogenous H[2]O[2] at infection sites would trigger the degradation of
   nanoparticles around infected bacteria, which would be along with the
   abovementioned synergistic antibacterial action under 1064 nm laser
   irradiation. Given this, in vivo therapeutic efficacy against infected
   wounds in mice was evaluated with an orderly program^[ [140]^23 ^] of
   wound fabrication, bacterial inoculation, infection establishment, and
   anti‐infective therapy (Figure [141] 5a). The infections involved two
   clinical MDR isolates: S. aureus USA300‐FPR3757^[ [142]^30 ^] and P.
   aeruginosa F291007.^[ [143]^31 ^] USA300‐FPR3757 belonged to the S.
   aureus USA300 clone, which had a highly virulent phenotype and
   represented a major source of community‐acquired S. aureus infections
   in the USA, Canada, and Europe.^[ [144]^30 ^] F291007 belonged to the
   high‐risk P. aeruginosa ST235 clone, which displayed a hypervirulent
   phenotype and caused worldwide nosocomial infections with poor clinical
   outcomes.^[ [145]^32 ^] Herein, a total of 5 therapeutic groups namely
   Mock, NIR‐II, NP^M123, NP^M123+NIR‐II, NP^M123/Fc, NP^M123/Fc+NIR‐II
   were employed. First, NP^M123/Fc+NIR‐II and NP^M123+NIR‐II gave almost
   the same tendency of temperature increase to about 48 °C within 3.5 min
   at P. aeruginosa‐infected wounds upon 1064 laser irradiation (Figure
   [146]S20, Supporting Information). These local mild‐heat temperatures
   would readily cause thermal injury to bacterial cells but not
   surrounding tissues.^[ [147]^10 ^] Second, the wounds were photographed
   and their areas were counted for mice infected by P. aeruginosa
   (Figure [148]5b,c) or S. aureus (Figure [149]S21a,b, Supporting
   Information) at sequential time points post‐therapy; compared to all
   the other therapeutic groups, NP^M123/Fc+NIR‐II gave the most obvious
   decrease in wound areas and the smoothest scabs on day 10 post‐therapy.
   As for body weights, there was no statistically significant difference
   between all five groups (Figure [150]5d; Figure [151]S21c, Supporting
   Information). Third, the bacterial loads at wounds on day 2
   post‐therapy were counted for P. aeruginosa‐ (Figure [152]5e,f) and S.
   aureus‐infected mice (Figure [153]S21d,e, Supporting Information);
   NP^M123/Fc+NIR‐II gave the largest inhibition rates (i.e., 94.88% and
   90.98% for P. aeruginosa and S. aureus, respectively) in relative to
   all the other four groups. Fourth, as determined by hematoxylin and
   eosin (H&E) staining of wound tissues on day 10 post‐therapy for P.
   aeruginosa‐ (Figure [154]5g) and S. aureus‐infected mice (Figure
   [155]S21f, Supporting Information), NP^M123/Fc+NIR‐II exhibited the
   best histopathologic outcome—abundant fibroblasts, regenerated blood
   vessels, and new hair follicles in granulation tissues; by contrast,
   there were abundant neutrophils, macrophages, and lymphocytes at wound
   tissues for all the other four groups, disclosing the persistence of
   infection‐induced inflammation at wounds. Taken together,
   NP^M123/Fc+NIR‐II gave the best in vivo antibacterial and wound healing
   efficacy.

Figure 5.

   Figure 5
   [156]Open in a new tab

   Therapeutic efficacy against P. aeruginosa‐infected wounds. FF1: Mock,
   FF2: NIR‐II, FF3: NP^M123, FF4: NP^M123+NIR‐II, FF7: NP^M123/Fc, FF8:
   NP^M123/Fc+NIR‐II. NIR‐II represents 1064 nm laser irradiation
   (1.0 W cm^−2, 3.5 min). a) Schematic illustration of the orderly
   program of infected wound establishment and anti‐infective therapy, as
   well as the time points for different tests post therapy. b)
   Representative photos of infected wounds post therapy. Statistical
   analysis of c) wound areas and d) body weights post therapy. Data are
   presented as mean ± SD (n = 5). e) Representative photos of plate count
   agars for infected wounds on day 10 post‐therapy, and f) corresponding
   statistical analysis of bacterial load counts. Data are presented as
   mean ± SD (n = 6). g) Representative photos of H&E staining of infected
   wounds (n = 3) on day 10 post‐therapy. Scale bar is 100 µm. Black
   arrows denote inflammatory cells such as neutrophils, macrophages, and
   lymphocytes. Green arrows represent hair follicles. Yellow arrows show
   fibroblasts. Red arrows stand for new blood vessels. ns: p ≥ 0.05, *:
   p < 0.05, **: p < 0.01, ***: p < 0.001.

2.6. Host‐Response View of Anti‐Infective Mechanism In Vivo

   To determine the expression of the four major proinflammatory cytokines
   including TNF‐α, IL‐6, IFN‐β, and IL‐17A,^[ [157]^33 , [158]^34 ^]
   immunohistochemistry was conducted with P. aeruginosa‐infected wound
   tissues on day 8 post‐therapy by Mock, NIR‐II, NP^M123, NP^M123+NIR‐II,
   NP^M123/Fc, and NP^M123/Fc+NIR‐II, and the results showed that the
   NP^M123/Fc+NIR‐II resulted in the lowest production of each of these
   cytokines (Figure [159] 6 ), indicating that NP^M123/Fc+NIR‐II enabled
   the best anti‐inflammatory effect. To measure the expression of the six
   major wound‐healing protein factors including α‐SMA and collagen III
   (extracellular matrix deposition), VEGF‐A and FGF‐2 (angiogenesis), and
   KGF and EGF (re‐epithelialization),^[ [160]^35 , [161]^36 ^]
   immunofluorescence staining was carried out with P. aeruginosa‐infected
   wound tissues on day 8 post‐therapy by the above six therapeutic
   groups, and the results indicated that NP^M123/Fc+NIR‐II displayed the
   highest production of each of these six wound‐healing factors
   (Figure [162] 7 ), denoting that NP^M123/Fc+NIR‐II brought the fastest
   wound‐healing process. Taken together, NP^M123/Fc+NIR‐II exhibited the
   best efficacy to inhibit inflammatory reaction and meanwhile to promote
   wound healing.

Figure 6.

   Figure 6
   [163]Open in a new tab

   Immunohistochemistry assay of major proinflammatory cytokines in mice
   on day 8 post‐therapy. FF1: Mock, FF2: NIR‐II, FF3: NP^M123, FF4:
   NP^M123+NIR‐II, FF7: NP^M123/Fc, FF8: NP^M123/Fc+NIR‐II. NIR‐II
   represents 1064 nm laser irradiation (1.0 W cm^−2, 3.5 min). (a)
   Representative immunohistochemical staining photos for TNF‐α, IL‐6,
   IFN‐β, and IL‐17A. Yellow brown represents positive detection. Scale
   bar = 100 µm. Corresponding statistical analysis for d) IL‐6, e) TNF‐α,
   f) IFN‐β, and g) IL‐17A. Data are expressed as mean ± SD (n = 3). ns:
   p ≥ 0.05, *: p < 0.05, **: p < 0.01, ***: p < 0.001.

Figure 7.

   Figure 7
   [164]Open in a new tab

   Immunofluorescence staining assay of major wound‐healing protein
   factors in mice on day 8 post‐therapy. FF1: Mock, FF2: NIR‐II, FF3:
   NP^M123, FF4: NP^M123+NIR‐II, FF7: NP^M123/Fc, FF8: NP^M123/Fc+NIR‐II.
   NIR‐II represents 1064 nm laser irradiation (1.0 W cm^−2, 3.5 min).
   Representative immunofluorescence staining photos for a) α‐SMA and
   Collagen III, b) VEGF‐A and FGF‐2, and c) KGF and EGF. Red represents
   positive detection. Scale bar = 100 µm. Corresponding statistical
   analysis for d) α‐SMA, e) Collagen III, g) VEGF‐A, f) FGF‐2, h) KGF,
   and i) EGF. Data are expressed as mean ± SD (n = 3). ns: p ≥ 0.05, *:
   p < 0.05, **: p < 0.01, ***: p < 0.001.

   To characterize the global host response after anti‐infective therapy,
   RNA‐seq‐determined transcriptomes of P. aeruginosa‐infected wound
   tissues on day 8 post‐therapy was compared between the two therapeutic
   groups NP^M123/Fc+NIR‐II and Mock (Figure [165] 8 ). First, the PCA
   plot (Figure [166]8a) showed the far distance between the clustered
   datasets of these two groups, indicating the existence of a high‐level
   of data reproducibility within each group as well as a high‐degree of
   data differentiation between these two groups. Second, a total of 236
   differentially expressed genes (DEGs) were identified for
   NP^M123/Fc+NIR‐II in relative to Mock, and they were composed of 131
   down‐regulated genes that included those encoding the above four
   proinflammatory cytokines, and additionally 105 up‐regulated genes that
   included those encoding the above six wound‐healing factors
   (Figure [167]8b). Third, as determined by gene ontology (GO) pathway
   enrichment analysis of these DEGs, there were the five enriched
   down‐regulated pathways including NF‐κB signaling, IL‐2 production,
   regulation of cytokine production involved in inflammatory response,
   negative regulation of activation‐induced cell death of T cells, and
   negative regulation of glucocorticoid receptor signaling pathway
   (Figure [168]8c), and meanwhile the seven enriched up‐regulated
   pathways including positive regulation of apoptotic process,
   angiogenesis, actin filament organization, positive regulation of cell
   differentiation, cortical cytoskeleton organization, endothelial tube
   morphogenesis, and regulation of hair cycle (Figure [169]8d). Fourth,
   KEGG enrichment analysis gave similar results—there were the seven
   enriched down‐regulated pathways including the TNF signaling pathway,
   NF‐κB signaling pathway, IL‐17 signaling pathway, JAK‐STAT signaling
   pathway, antigen processing and presentation, B cell receptor signaling
   pathway, leukocyte transendothelial migration (Figure [170]8e), and
   simultaneously the nine enriched up‐regulated pathways including
   ECM‐receptor interaction, regulation of actin cytoskeleton, adherens
   junction, cell adhesion molecules, VEGF signaling pathway, TGF‐β
   signaling pathway, Notch signaling pathway, Apelin signaling pathway,
   and Hippo signaling pathway (Figure [171]8f). Notably, all the above
   enriched down‐regulated pathways were involved in the resolution of
   excessive inflammation, while all these enriched up‐regulated pathways
   contributed to wound healing, giving a global view of the coordinated
   process of clearance of pathogenic stimuli along with restoration of
   tissue integrity and function. Fifth, the heatmap in Figure [172]S22
   (Supporting Information) presented not only the ten factors
   simultaneously revealed by the traditional immunoassay and the RNA‐seq
   assay, but also the 12 selected down‐regulated proinflammatory factors
   and the 19 selected up‐regulated wound‐healing factors additionally
   disclosed by RNA‐seq, all of which would be the major contributors to
   the highly favored therapeutic effect of NP^M123/Fc+NIR‐II. Taken
   together, the therapeutic effect with respective to host response was
   mainly composed of inhibition of inflammatory reaction and meanwhile
   promotion of wound healing.

Figure 8.

   Figure 8
   [173]Open in a new tab

   RNA‐seq assay of P. aeruginosa‐infected wounds on day 8 post‐therapy.
   a) PCA plot of clustered datasets between the two therapeutic groups
   NP^M123/Fc+NIR‐II (FF8) and Mock (FF1). NIR‐II represents 1064 nm laser
   irradiation (1.0 W cm^−2, 3.5 min). b) Volcano plot of DGEs for
   NP^M123/Fc+NIR‐II compared to Mock. Up: Up‐regulated genes. Down:
   Down‐regulated genes. GO pathway enrichment analysis was performed for
   c) down‐regulated and d) up‐regulated genes, and KEGG pathway
   enrichment analysis was conducted for e) down‐regulated and f)
   up‐regulated genes. Experiments were performed with four independent
   biological replicates.

2.7. Biosafety at Cellular and Animal Levels

   First, the embryonic mouse normal fibroblast cell line NIH‐3T3 and the
   human normal ovarian epithelial cell line IOSE‐80 were used to test the
   potential cytotoxicity of NP^M123/Fc by using Cell Counting Kit‐8
   (CCK8), and each cell line gave a cell viability >80%^[ [174]^14 ,
   [175]^23 , [176]^31 ^] after co‐incubation with 100 µg mL^−1 NP^M123/Fc
   for 12 h (Figure [177]S23, Supporting Information). The additional
   hemolysis assay with mouse red blood cells (RBCs) revealed that
   100 µg mL^−1 NP^M123/Fc presented a hemolytic rate <5% (Figure
   [178]S24, Supporting Information).^[ [179]^14 , [180]^23 , [181]^31 ^]
   Second, mice with infection‐free wounds were treated with Mock, NIR‐II,
   NP^M123, NP^M123+NIR‐II, NP^M123/Fc, NP^M123/Fc+NIR‐II using the
   administration route and dosage same as anti‐infective therapy. H&E
   staining on day 8 post‐treatment showed no inflammatory or pathological
   lesions in heart, liver, spleen, lung, and kidney (Figure [182]S25,
   Supporting Information). After treatment with 1 mg kg^−1
   lipopolysaccharide (LPS), obvious infiltration of neutrophils,
   macrophages, and lymphocytes were observed at wounds (positive control)
   but not in all the five organs (Figure [183]S25, Supporting
   Information). Biochemical blood test at the same time point exhibited
   no abnormal changes in the two major hepatic function indexes namely
   alanine aminotransferase (ALT), aspartate aminotransferase (AST) and
   the two major renal function indexes namely blood urea nitrogen (BUN)
   and creatinine (CREA) (Figure [184]S26, Supporting Information). Third,
   for S. aureus‐ or P. aeruginosa‐infected wounds treated with the above
   six groups, and H&E staining (Figure [185]S27, Supporting Information)
   and biochemical blood test (Figure [186]S28, Supporting Information) on
   day 8 post‐therapy gave the similar biosafety results; it should be
   noted that, after wounds were infected with bacteria, there was
   significant inflammatory reaction at infected wounds, but no obvious
   pathological injury was found in the heart, liver, spleen, lung, and
   kidney whether the therapeutic measures were applied or not. Taken
   together, NP^M123/Fc had highly favorable biosafety at cellular and
   animal levels.

3. Conclusion

   The present work developed a novel polymer nanomaterial NP^M123/Fc for
   treatment of acute wound infections. We used two distinct polymers,
   namely the photothermal agent M123‐containing P^M123 and the
   Fc‐containing P^Fc both containing ROS‐sensitive moieties, to be
   assemble into NP^M123/Fc. Fc was built in a polymerized manner at the
   center locations of NP^M123/Fc, and thus Fc introduced was highly
   controllable at high consternations. In addition, Fc as a CDT agent has
   been showed higher levels of biocompatibility relative to inorganic CDT
   metals.^[ [187]^16 , [188]^17 , [189]^18 ^] As expected, M123 in
   NP^M123/Fc endowed the efficient and stable photothermal effect upon
   1064 nm laser irradiation and, Via Fenton‐like reaction, Fc in
   NP^M123/Fc enabled the chemodynamic effect to produce more toxic ∙OH
   from endogenous H[2]O[2] abundant at infection sites. These two
   distinct antibacterial contributors PTT and CDT worked in a strong
   synergistic manner. NP^M123/Fc+NIR‐II gave >90% in vitro inhibition
   rates against MDR ESKAPE pathogens. The metabolomics analysis disclosed
   NP^M123/Fc+NIR‐II in vitro induced bacterial metabolic dysregulation
   especially including inhibited nucleic acid synthesis, disordered
   energy metabolism, enhanced oxidative stress, and elevated DNA damage.
   NP^M123/Fc+NIR‐II possessed >90% bacteriostatic rates in a mouse model
   of cutaneous wounds infected by P. aeruginosa and S. aureus, resulting
   in almost full recovery of infected wounds. The systematic
   immunohistochemistry, immunofluorescence staining, and transcriptomics
   assays with respective to host response disclosed that the therapeutic
   effect was mainly dependent on inhibition of inflammatory reaction and
   meanwhile promotion of wound healing at wound sites. The ROS‐sensitive
   thioketal bonds in NP^M123/Fc made it degradable upon ROS. Endogenous
   H[2]O[2] at infection sites would trigger the degradation of NP^M123/Fc
   located around infected bacteria, and ∙OH generated by CDT would
   further promote this biodegradation. On the one hand, the rapid
   degradation of NP^M123/Fc endowed highly favorable biosafety as
   observed at both cellular and animal levels. On another hand, this
   rapid degradation would be along with the quick release of M123 (PTT)
   and Fc (CDT) for subsequent rapid bactericidal actions. NP^M123/Fc with
   a unique synergistic PTT and CDT strategy represented a biocompatible
   and broad‐spectrum nanoplatforms for antibiotics‐alternative treatment
   of acute superficial infections.

Conflict of Interest

   The authors declare no conflict of interest.

Supporting information

   Supporting Information
   [190]ADVS-11-2309624-s001.pdf^ (3.1MB, pdf)

Acknowledgements