Extracellular Oxygen Sensors Based on PtTFPP and Four-Arm Block Copolymers

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Nanosensors for extracellular oxygen-sensing.

Abstract

Three four-arm amphiphilic block copolymers with different chain lengths, consisting of a hydrophilic chain of polyethylene glycol (PEG) and hydrophobic segment of polycaprolactam (PCL), were synthesized and used to encapsulate the high-efficient and hydrophobic oxygen probe of platinum(II)-5,10,15,20-tetrakis-(2,3,4,5,6-pentafluorophenyl)-porphyrin (PtTFPP) to form polymer micelles. This approach enabled the use of PtTFPP in aqueous solution for biosensing. Experimental results demonstrated that the particle sizes of these nano-oxygen sensors between 40.0 and 203.8 nm depend on the structures of block copolymers. PtTFPP in these micelles showed an effective quantum yield under nitrogen environment, ranging from 0.06 to 0.159. The new sensors are suitable for analyzing dissolved oxygen concentrations in the range of 0.04–39.3 mg/L by using the linear Stern–Volmer equation at room temperature. In addition, it has been shown that these sensors are capable of in situ monitoring the dissolved oxygens in the culture medium of E. coli and Romas cells during the respiration process, and distinguishing the drug activity of antibiotic ampicillin from that of antimycin A. This study showed that the use of these nanostructured multi-arm block copolymer micelles can achieve efficient biological applications without specific structural modification of the hydrophobic PtTFPP probe, which is expected to have broad prospects.

1. Introduction

Oxygen is vital to life [1,2]. It plays a crucial role in the fields of the environment [3,4,5], industry [6,7], agriculture [8], biology [9,10,11], and health [12,13]. The oxygen content in the atmosphere is 21%, and is approximately 8.86 mg/L in the water. An adult man cannot survive for more than 10 min without oxygen. Besides, oxygen is also an important "food"; the body metabolizes about 200–250 grams per day [14]. Hypoxia [15] refers to a condition in which a body or a region of the body lacks sufficient oxygen supply at the tissue level, which may lead to a series of physiological and pathological consequences, and even lead to cell death [16]. Studies have shown that hypoxia changes cell behavior and is associated with extracellular matrix remodeling and increased migration and metastatic behavior [17,18]. Therefore, the ability to measure the oxygen content of tissues and living cells plays a critical role in understanding cellular activities, the pathological causes of disease, and cancer diagnosis. On the other hand, a known fact is that cellular respiration refers to the process in which an organic substance undergoes a series of oxidative decomposition in a cell to produce an inorganic substance or a small molecule organic substance, which releases energy and generates adenosine triphosphate (ATP). Therefore, the measurement of cellular oxygen consumption is of great significance for the evaluation of aerobic glycolysis rate, oxidative phosphorylation, and high-throughput drug screening [19,20].

The methods currently used to determine oxygen content are limited. The Clark electrode method is the most common method for measuring the partial pressure of oxygen. It is based on current analysis and consists of a platinum electrode wrapped with an oxygen permeable membrane. By applying a voltage, the surface of the electrode undergoes a redox reaction, causing oxygen to be reduced at the surface of the electrode, thereby obtaining an oxygen concentration. However, the Clark electrode method may lead to oxygen consumption and tissue damage. The second common method is also an electrochemical method, and is widely used for measuring oxygen concentration in fuel cells and the automotive industry. However, it is difficult to apply in the biological field. In addition, there are radioisotope methods and magnetic resonance methods, but these techniques are complicated to operate and costly. Therefore, optical sensing and imaging based on luminescence quenching is considered to be a true alternative to the above method, and is replacing the Clark electrode method in many areas [12].

Although there are numerous optical oxygen probes on the market, including some newly synthesized oxygen probes, most of them are hydrophobic [21,22]. Although hydrophobic probes have high quantum yield, they are difficult to be used in biology. Some hydrophobic probes can be modified directly into hydrophilic oxygen probes; however, the quantum yields of such probes are quite low. For example, the quantum yield of a hydrophilic oxygen probe made of metalloporphyrin (PtCPK, PtCP, PdCPK, PtTCPP) [23,24] is as low as 0.001–0.0095. Therefore, Vinogradov et al. developed a series of dendritic oxygen probes to improve the quantum yields of oxygen probes and give them some additional characteristics [25,26]. These oxygen probes have hydrophobic shells with platinum–porphyrin as the core. For these dendrimers, phosphorescent metalloporphyrins were encapsulated inside hydrophobic dendrimers, which form disclosure shells by enveloping the chromophores to control oxygen diffusion to the excited triplet states. Polyethylene glycol (PEG)ylation or the carboxylation of dendritic molecules can not only ensure the high water solubility of sensors, but also prevent direct interactions between probes and biomacromolecules. Therefore, the quantum yields of these porphyrin-containing dendritic molecules were raised to between 0.017 and 0.073 [26]. Since 2012, in order to enable the application of hydrophobic probes in biological research, we have started to use micelles formed by amorphous block copolymers to encapsulate highly efficient hydrophobic oxygen probes. At present, we mainly use amphiphilic polymers, including grafted polymers [27,28] and simple block copolymers [29]. The quantum yields of platinum(II)-5,10,15,20-tetrakis-(2,3,4,5,6-penta- fluorophenyl)-porphyrin (PtTFPP) in these micelles depend on whether there is energy transfer in the micelles [28] and the structures of the polymers [29], which are approximately between 0.107 and 0.231. Other research groups have reported the use of conjugated polymers or metalloporphyrins nanoparticles to achieve a high quantum yield of 0.18 for intracellular oxygen sensing [30,31,32].

Considering the abundance of polymer architectures, herein we used multi-arm block copolymers as carriers for achieving high quantum yields of PtTFPP in water. In recent years, multi-arm block polymers have gradually attracted considerable interest. It has been reported that micelles composed of multi-armed polymers may exhibit various morphologies and assemblies than linear polymers [33,34]. Also, multi-arm star polymers have higher terminal groups’ concentrations than conventional linear polymers [35], which enables further modification or the functionalization of the multi-arm polymers. Herein, we used a multi-arm poly(ethylene glycol) as the initiator to initialize the polymerization of ε-caprolactone to obtain multi-arm block copolymers (Scheme 1). The micelles formed from these polymers were used to incorporate the hydrophobic and widely-used oxygen probe of PtTFPP to enable the application of PtTFPP in biological conditions (Figure 1). For demonstrating the bioapplication capability of these nanostructured sensors, the sensors were applied to in situ monitoring of cell respiration with and without drug inhibition through a commercially available plate reader.

2. Materials and Methods

2.1. Materials and Reagents

The hydrophobic oxygen probe PtTFPP used in the experiment was ordered from Frontier Scientific (Logan, UT, USA). The chemical reagents and solvents used in these experiments were of analytical grade. 4-(2-Hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES) was purchased from Sigma-Aldrich (St. Louis, MO, USA). 4-Arm poly(ethylene glycol) (M_n = 10,000, M_w/M_n = 1.06) was ordered from Creative PEGWorks (Chapel Hill, NC, USA). ε-Caprolactone (CL) was ordered from Sigma-Aldrich together with the antibacterial drug Ampicillin and Antimycin A. Stannous octoate was commercially available from J & K Scientific Ltd, Shanghai. Dialysis membranes (regenerated cellulose, Mw cut off 10000) were purchased from Pierce (Rockford, IL).

Double-distilled water was utilized to prepare the HEPES buffer and Lysogeny Broth (LB) medium. To accurately control gas proportion, a gas manipulator (measuring error: ±1%) which was purchased from Alicat Scientific (Tucson, AZ, USA) was used. All experimental sensing measurements were performed under normal atmospheric pressure (760 mmHg, or 101 kPa) at room temperature (25 °C) or 37 °C conditions.

The Lysogeny Broth medium used for the cultivation of E. coli is made by dissolving 10 g of tryptone, 5 g of yeast extract, and 10 g of sodium chloride in 1 L of double-distilled water and adjusting the pH to 7.0 with sodium hydroxide. The medium could be used for culturing bacteria after high-temperature and high-pressure sterilization. Ramos cells (human B lymphocyte cells) were cultivated in a 37 °C thermostatic cultivation box containing 5% carbon dioxide by using culture medium RPMI 1640 containing 10% fetal bovine serum (FBS) and 1% penicillin. HEPES buffer was prepared by dissolving 2.383 g HEPES in 1 L of double-distilled water and adjusting the pH to 7.4 with NaOH.

2.2. Instruments

The structures and molecular weights of the four-arm block copolymers were determined by 400M ¹H Nuclear Magnetic Resonance (¹H-NMR, Ascend™ 400, Bruke) and gel permeation chromatography (GPC) (Water 1515, Waters), respectively. Dynamic light scattering (DLS) (Nano ZS, Malvern) was used to measure the diameters of the micelles. Before measuring the micelle diameters, the micelles were dedusted by a 0.45-μm nylon microfilter membrane. The absorbance and phosphorescence intensity was determined by UV/Vis spectrometer (Lambda 25, PerkinElmer) and spectrofluorophotometer (FluoroMax-4, Horiba), respectively. The FLS980 spectrometer (Edinburgh Instruments) was used to determine the lifetime of micelles. The oxygen consumption of E. coli and Ramos cells was measured by a Cell imaging multi-mode microplate reader (Cytation 3, Cytation™).

2.3. Synthesis

2.3.1. Synthesis of P1

First, 1.0 g of dry 4-arm PEG-OH was placed into a clean shrink tube together with the magnetic stirrers, and was vacuumed for 10 min. Then, 2.591 g ε-caprolactone monomer was mixed with a drop of stannous octoate (Sn(Oct)₂) as a catalyst in a small flask, and injected into the shrink tube with the protection of nitrogen gas. The air in the pipe was removed through a Schlenk Line. Under the condition of nitrogen in a 110 °C oil bath, the polymerization was kept under stirring for 24 hours. Then, the reaction was quickly ventilated and cooled in an ice bath to halt. The crude solid products were re-dissolved with 1–2 mL dichloromethane (DCM) and precipitated two to three times with ice ethyl ether. White powder solid products were obtained. This procedure was repeated twice for getting a pure white polymer P1 of 2.645 g with a yield of 73.6%. ¹H NMR (400 MHz, Chloroform-d, δ(ppm)): 4.22 (t, 8H, -COOCH₂CH₂O-), 4.05 (t, 404H, -COOCH₂CH₂-), 3.64 (s, 900H, -CH₂O-), 2.30 (t, 402H, -C=OCH₂-), 1.69–1.59 (m, 808H, -CH₂CH₂O- and -COCH₂CH₂-), 1.38 (m, 403H, -CH₂CH₂CH₂-). n = 51; M_n(NMR) = 33000; M_n(GPC) = 13610, M_w(GPC) = 17800, M_w/M_n = 1.31.

2.3.2. Synthesis of P2

P2 was synthesized following the same procedure of P1, except 1.0 g of four-arm PEG-OH was used to initialize 1.296 g of ε-caprolactone. Yield is 45.6%. ¹H NMR (400 MHz, Chloroform-d, δ (ppm)): 4.22 (t, 8H, -COOCH₂CH₂O-), 4.05 (t, 153H, -COOCH₂CH₂-), 3.64 (s, 900H, -CH₂O-), 2.30 (t, 153H, -C=OCH₂-), 1.69–1.59 (m, 311H, -CH₂CH₂O- and -COCH₂CH₂-), 1.38 (m, 155H, -CH₂CH₂CH₂-). n = 19; M_n(NMR) = 18700; M_n(GPC) = 11940, M_w(GPC) = 12680, M_w/M_n = 1.27.

2.3.3. Synthesis of P3

P3 was synthesized following the same procedure of P1, except 1.0 g of four-arm PEG-OH was used to initialize 0.648 g of ε-caprolactone. Yield is 44.0%. ¹H NMR (400 MHz, Chloroform-d, δ (ppm)): 4.22 (t, 8H, -COOCH₂CH₂O-), 4.05 (t, 59H, -COOCH₂CH₂-), 3.64 (s, 900H, -CH₂O-), 2.30 (t, 64H, -C=OCH₂-), 1.69–1.59 (m, 126H, -CH₂CH₂O- and -COCH₂CH₂-), 1.38 (m, 58H, -CH₂CH₂CH₂-). n = 8; M_n(NMR) = 13600; M_n(GPC) = 8970, M_{w (GPC)} = 9960, M_w/M_n = 1.11.

2.4. Preparation of Micelles

Micelles containing oxygen probes were prepared according to the method described in reference [27]. Generally, 1 mg of PtTFPP and 15 mg of polymer were dissolved in 600 μL of tetrahydrofuran and slowly injected the mixed solution into 3 mL of MilliQ distilled water stirred at high speed (1400 rpm) to form a uniform and transparent micelle solution. Then, the micelles were transferred to a dialysis bag, and the dialysis was carried out at a low speed in the pure water of Mili Q for two days to remove the organic solvent in the micelles. Change the water every 12 hours during the dialysis process. After the completion of the dialysis, the 0.45-μm filter was used to remove the unformed PtTFPP and the polymer. Then, we used an ultraviolet spectrophotometer to measure the concentration of PtTFPP in micelles by comparing the absorbance of micelles with the standard curve of PtTFPP in tetrahydrofuran. The polymer concentrations for micelle stock solutions were determined by freeze-drying the micelles solutions. Three kinds of micelles—M1, M2, and M3—were prepared by using the polymers P1, P2, and P3, respectively.

2.5. Oxygen Sensing Performance

The mixed gas of nitrogen and oxygen controlled by the Alicat gas flow controller is injected into the solution to adjust the oxygen concentration in the solution. The sensing performance of all the oxygen sensors was tested at atmospheric pressure (760 mmHg or 101.3 kPa) and at room temperature (25 °C) or 37 °C. Under atmospheric pressure, the dissolved oxygen concentration in water is 8.25 mg/L at room temperature and 6.8 mg/L at 37 °C, respectively.

2.6. Critical Micelle Concentration (CMC) Determination

The determination of critical micelle concentrations in double-distilled water of P1, P2, and P3 was according to the traditional determination method using pyrene as a hydrophobic fluorescence probe [36]. Generally, the acetone solution of pyrene with a concentration of 6 × 10⁻⁵ M was prepared under dark conditions. First, 30 μL of the solution was placed in a 5-mL vial, and the acetone was removed by evaporation. Polymers with concentrations from 1.0 × 10⁻⁴ to 0.04 mg/mL (3 mL) were added to the above vials. After stirring for 4 hours and ultrasonically shaking for 15 min, the vials were placed in a dark place at room temperature for 24 hours, waiting for measurement. The final pyrene concentration was adjusted to 6.0 × 10⁻⁷ M. The emission spectra of these solutions were measured by a spectrofluorophotometer under 334-nm excitation at room temperature. The relationship between the ratios of fluorescence intensities at 373 nm and 384 nm and polymer concentrations was analyzed to calculate the CMCs of the polymers.

2.7. Determination of Quantum Yields

The phosphorescence quantum yields of micelles were determined according to the “relative method” used in the literature [37]. The quantum yield of PtTFPP in dichloromethane (0.088 [38]) was used as a reference standard sample, and was calculated by the following formula [37]:

$${\eta }_s={\eta }_r\left(\frac{A_r}{A_s}\right)\left(\frac{I_s}{I_r}\right)\left(\frac{n_s^2}{n_r^2}\right)$$

(1)

where η_r and η_s are the phosphorescence quantum yields of the reference standard sample and the measured sample, respectively, and A_r and A_s represent the absorbance of the standard and the measured sample at the excitation wavelength. I_r and I_s are the integral values of the fluorescence intensity of the standard sample and the measured sample, respectively, and n_r and n_s are the refractive indices of the corresponding solvents in the standard sample and measured sample, respectively. The refractive indices of water and methylene chloride are 1.333 and 1.424, respectively.

2.8. Response Time Measurements

First, 3 mL of PtTFPP containing micelles were placed in a quartz cuvette, and its emission intensity at 650 nm was measured every 0.5 seconds under excitation at 405 nm to detect the real-time change of phosphorescence intensity. We use a gas flow controller to control the change of dissolved oxygen concentrations in the solutions by blowing gas with different oxygen and nitrogen ratios [11,39]. The sum velocity of the oxygen and nitrogen was set to 50 cubic centimeters per minute. The measurement starts from the state of oxygen saturation (oxygen ratio is 100%. The dissolved oxygen concentration is 39.3 mg/L and 32.4 mg/L at 25 °C and 37 °C, respectively). After reaching equilibrium, the gas was immediately changed to 100% nitrogen (oxygen ratio is 0%, and dissolved oxygen concentration is 0mg/L at both 25 °C and 37 °C). These two states were constantly switched to see the real-time changes in fluorescence at 650 nm. For these measurements, t_0.95 represented the time when the fluorescence intensities reached 95% of the fluorescent maximum or minimum.

2.9. Lifetime Determination

The lifetimes of the micelles M1, M2, and M3 were determined by using an FLS980 Spectrometer [40,41]. First, 3 mL of PtTFPP micelles were placed in a quartz cuvette, and the phosphorescence lifetime of the luminescent particles was measured respectively under air and nitrogen conditions. Before measuring the lifetime of the micelle under nitrogen conditions, the quartz cuvette needs to be purged for 5 min with nitrogen and sealed with a rubber stopper. The lifetimes of M1, M2, and M3 were obtained from collecting the lifetime data of 10,000 photons emitted at 650 nm under the excitation of 405 nm. Usually, the lifetime of the micelles is calculated by fitting the attenuation curve.

2.10. Micelles’ Stability

The stability of the micelles was measured by observing the absorption changes of the probe PtTFPP in micelles and size changes of micelles [39,42]. Micelles M1, M2, and M3 were stored in a refrigerator at 4°C. Then, 20 μL of micelles were taken out and diluted 100 times for the stability test.

2.11. Culture of E. coli for Extracellular Sensing

E. coli was cultured in liquid Lysogeny Broth (LB) medium according to the method previously published [43]. The absorbance of the bacterial solution at 600 nm (OD₆₀₀) is linearly related to the cell density of E. coli in the range of 0.1–1. An OD₆₀₀ of 1 corresponds to the E. coli’s cell density of 5.0 × 10⁸ cfu/mL. The bacterial solution was diluted with the LB medium to obtain different E. coli densities for the following experiments.

2.12. Culture of Mammalian Cells for Extracellular Sensing

Ramos cells (human B Lymphocyte cells) were cultured in RPMI 1640 supplemented with 10% FBS and 1% penicillin at 37 °C in a 5% CO₂ atmosphere. Cells with different densities suspended in the medium were used for cell respiration measurements.

2.13. Oxygen Respiration Experiments

First, 3 μL of oxygen sensor micelles (concentrations of P1, P2, and P3 in M1, M2, and M3 are 1.4 mg/mL, 2.1 mg/mL, and 2.7 mg/mL, respectively. The concentration of PtTFPP is 200 μg/mL (170 μM)) was mixed with different concentrations of cells or bacterial solution at a ratio of 3:200 and added to 96-well plates for cellular respiration measurements. Then, 200 μL of mixed solution was added to each well in a 96-well plate to ensure that the concentration of PtTFPP in each well was 3 μg/mL (2.55 μM). A thin layer of mineral oil (100 μL per well) was usually added to the top of the medium to prevent the oxygen exchange in the medium. It is recognized that the method of using mineral oil for oxygen respiration detection is generally applicable to biological research [44,45]. The 96-well plate was stabilized in a plate reader at 37 °C for 20 min to remove the influence of temperature. The emission of PtTFPP was detected at 650 nm using top reading under the excitation at 405 nm.

3. Results and Discussion

3.1. Synthesis and Structure Characterization of the Polymers

The polymers were prepared through ring-opening polymerization by using the four-arm PEG-OH as the initiator (Scheme 1). ¹H NMR and GPC were used to characterize the polymers’ structures and molecular weights.

Figure 1 gave the ¹H NMR spectra of the polymers. By using the integration of the peaks a, b, c from polyethylene glycol (PEG) sites to the integration of the peaks d and h from ε-caprolactone (CL) moieties, molecular weights of the block copolymers and the weight ratios of the hydrophilic segments to hydrophobic chains were obtained (

Table 1. Materials’ properties of P1, P2, and P3. GPC: gel permeation chromatography.

Product-	Molar Ratioa	Yield	GPC			1H-NMR	Mass ratiob	CMC (μg/mL)
			Mn (g/mol)	Mw (g/mol)	PDI	Mn (g/mol)
P1	1:224	72.4%	13610	17800	1.31	33300	1:2.33	3.10
P2	1:112	45.6%	11940	12680	1.27	18700	1:0.87	2.49
P3	1:56	44.0%	8970	9960	1.11	13600	1:0.36	3.31

^a Feed molar ratio of 4-arm poly(ethylene glycol) (PEG) to ε-caprolactone (CL). ^b Molar ratio of hydrophilic chain to hydrophobic chain determined by ¹H NMR.

). According to the feed ratios of PEG to CL, three polymers were successfully obtained with P1 exhibiting the highest molecular weight and P3 possessing the lowest molecular weight.

3.2. Micelle Preparation and Characterization

As described in the experimental section (Figure 2), micelles were prepared by incorporating PtTFPP into an amphiphilic block copolymer and injecting into distilled water under high-speed stirring. The tetrahydrofuran (THF) was removed by dialysis. The particle sizes of the micelles were measured by DLS with average diameters of about 40.0–203.8 nm (Figure 3), which depends on the polymers used. The more hydrophobic segments of the polymer, the larger the size of the micelles. By observing the stability of the micelles (Figure S1), the micelles were found to be stable at 4 °C for at least 24 days with little change in the sizes or physical properties. The concentration of PtTFPP in the micelles was determined by lyophilizing the micelles, re-dissolving in tetrahydrofuran, measuring the absorption spectrum of PtTFPP, and calculating from the standard curve of PtTFPP in THF. The detailed polymer concentration and PtTFPP concentration in the micelles were provided in

Table 2. Properties of micelles M1–M3. DLS: dynamic light scattering.

Micelles-	Concentration of Polymers (mg/mL) a	Concentrations of PtTFPP (mg/mL) b	Quantum Yield (%)	Encapsulation efficiency for PtTFPP (%) c	PtTFPP loading ratio (wt%) d	Size (nm) e	PDI e
M1	1.4	0.219	15.9	65.7	15.4	203.8	0.221
M2	2.1	0.226	11.4	79.1	9.7	89.3	0.176
M3	2.7	0.195	6.0	82.9	6.7	40.0	0.213

^a Polymer concentration in micellar solutions. ^b PtTFPP in micellar solutions. ^c The weight ratio of PtTFPP encapsulated in micelles to the fed PtTFPP for micelles’ preparation. ^d The weight percentage of PtTFPP to the block copolymers. ^e Determined by DLS.

.

For amphiphilic polymers, CMC is an important concern. The CMCs of the polymers P1–P3 were measured to be from 2.5 to 3.3 μg/mL (Figure S2). These CMCs are compatible with other amphiphilic block copolymers [46,47]. These micelles containing PtTFPP have high storage stability. At 37 °C, at least 95% of the PtTFPP was still in the micelles after five days of dialysis.

3.3. Photophysical Properties

Figure 4 shows the absorbance and phosphorescence spectra of PtTFPP at a concentration of 2.2 μM in micelles, THF, and DCM, respectively. The results demonstrated that at the same PtTFPP concentration, the PtTFPP’s emission peak in micelles is significantly higher than that in tetrahydrofuran and dichloromethane, indicating that PtTFPP in micelles has a higher quantum yield than that in organic solvents THF and DCM. The quantum yield of PtTFPP in micelle M1 under nitrogen conditions was measured to be 0.159 (Table 2), which is much higher than its quantum yield in dichloromethane (0.088 [38]). This situation may be explained by PCL providing a suitable micro-environment for PtTFPP because of its polarity compatibility [38]. Therefore, the use of micelles not only enabled the use of PtTFPP in aqueous solution, but also preserved the quantum yield of PtTFPP. Owing to the different ratios of hydrophobic chains of PCL to hydrophilic segments of PEG, the quantum yields of PtTFPP in the three micelles were also affected by the polymer structures. Among them, the M1 micelles with the longest hydrophobic segment PCL can alleviate more significant interactions between PtTFPP and water, so that PtTFPP in M1 showed the highest quantum yield among the three micelles.

3.4. Oxygen Sensing Properties

Typical oxygen sensing using micelle M3 as a representative example was given in Figure 5. With the increase of oxygen concentration, the emission of PtTFPP at 650 nm decreased, showing the oxygen sensitivity of the sensors (Figure 5a). The intensity ratio (I₀/I) curves showed in Figure 5b followed the Stern–Volmer equation:

$$\frac{I_0}{I}= 1+{\mathrm{K}}_{SV}\left[O_2\right]$$

(2)

where K_SV is the Stern–Volmer quenching constant, and [O₂] is the dissolved oxygen concentration adjusted through the nitrogen and oxygen gas mixer. Linear Stern–Volmer curves were observed, and the curves were not affected much by the polymer structures (Figure 5b). The limits of detection (LOD) for oxygen were found to be 0.07, 0.09, and 0.04 mg/L, for M1, M2, and M3, respectively by using three folds of standard errors of K_SV divided by K_SV. The temperature has some influences on sensitivity. Figure 5c gave the Stern–Volmer curves of M3 at 37 °C and 25 °C, respectively. Slightly faster KSV at 37 °C than that at 25 °C was observed due to the higher interaction activity of oxygen molecules with PtTFPP at higher temperatures.

The changes in the lifetimes of PtTFPP in micelles were also measured at room temperature under nitrogen and air conditions (Figure 6). Under air condition, the three micelles’ lifetimes are 7.64 to 7.93 μs for M1, M2, and M3, which were slightly affected by the micellar structures. Under nitrogen conditions, the differences of lifetimes among the three micelles are much more obvious; those are 23.65, 28.02 and 35.98 μs for M1, M2, and M3, respectively.

3.5. Response Time and Reversibility

The oxygen sensor’s response time was measured by the oxygen/nitrogen saturation method (Figure 7), where t_0.95 is the time required to generate 95% of the total phosphorescence intensity change. The oxygen sensor recovers from the oxygen saturation state to the deoxygenation state for a relatively slow time. As shown in Figure 7, according to the calculation, the response time t_0.95 of M1–M3 in the aqueous solution from the deoxidation state to the oxygen saturation state is between 20 and 30 s, and the response time from the oxygen saturation state to the deoxygenated state is between 170 and 220 s (t_0.95-r). This response time is faster than that of PtTFPP physically doped in poly(2-hydroxyethyl methacrylate) (PHEMA) matrix and polystyrene (PS) matrix, where t_0.95 from the deoxygenation state to the oxygen saturation state is 50 s [43]. It is shown that the encapsulation of PtTFPP in micelles formed from amphiphilic multi-arm block polymers is a kind of reasonable oxygen sensing material. Besides, the repeatability of these oxygen sensors from oxygen saturation state to deoxygenation state is excellent, indicating that these micellar oxygen sensors can be reused for continuous dissolved oxygen detection.

3.6. Monitoring of Oxygen Consumption of E. coli

E. coli was used as a biological model in the experiment to measure the oxygen consumption of bacterial cells. According to the literature, E. coli has a shorter generation time, which is distributed approximately 20–90 min depending on temperature and cell density [34]. This makes it easier to detect changes in the extracellular oxygen concentration of E. coli. During the growth of E. coli, oxygen is consumed by aerobic metabolism. In the experiment, a layer of mineral oil was added to the upper layer of the LB medium to reduce the gas exchange between oxygen and the outside air. In order to observe the changes in the oxygen consumption rate at different concentrations, the initial cell densities of E. coli JM109 were set to 5 × 10⁶, 2.5 × 10⁶, 1.3 × 10⁶, 6.3 × 10⁵, and 0 cfu/mL, respectively, and the concentration of PtTFPP was 3 μg/mL (2.56 μM). The experiment was carried out in a 96-well plate at 37 °C, and its phosphorescence intensity was detected at an excitation wavelength of 405 nm by a microplate reader (Figure 8). Even when the concentration of E. coli was as low as 6.3 × 10⁵ cfu/mL, a significant change in phosphorescence intensity was observed after 80 min due to the aerobic metabolism of E. coli in LB medium. As the initial E. coli density increases, the completeness of oxygen consumption becomes faster and faster.

Mineral oil seals are commonly used to measure oxygen consumption and cell acidification during cell growth. The experiment also studied the effect of oil seals on the rate of oxygen consumption. The experiment was conducted in two parallel experiments under comparable conditions, one with an oil-free test and one with an oil-sealing test (Figure S3). The results showed that the time of depletion of oxygen in the oil-sealing group is shorter than that in the oil-free group, indicating that oil sealing can effectively isolate the oxygen exchange between the medium and the outside air, which is essential for completing the experiment in a shorter amount of time.

3.7. Monitoring Oxygen Consumption of Romas Cells

This oxygen sensor can also be used to measure the oxygen consumption of mammalian cells. We utilized Romas cells as a representative of mammalian cells for measurement. In order to reduce the experimental time, we use the mineral oil sealing. As shown in Figure 9, the growth of eukaryotic cells typically takes a longer time than prokaryotic cells. The rate of oxygen consumption of cells is strongly correlated with cell density. When the cell density is 19,000/well, the phosphorescence intensity does not change much. When the cell density increased to 1,500,000/well, the oxygen consumption of the cells increased significantly within 0.6 hours, and the phosphorescence intensity varied greatly.

3.8. Cell Viability Test

For E. coli, we tested bacterial growth at different sensor concentrations. No significant cytostatic or cytotoxicity was observed with a constant cell density (5 × 10⁵ cfu/mL), while different sensor concentrations ranging from 8–120 μg/mL, corresponding to PtTFPP concentrations of 0.9 to 6.8 μM (Figure 10). It should be noted that when these micelles are used to measure the oxygen concentration, the content of PtTFPP is usually 2.5 μM, which is much lower than 6.8 μM.

We use the methyl thiazolyl tetrazolium (MTT) method to measure the biocompatibility of the sensors to Ramos cells. In this experiment, cells were assayed for cell activity after incubation with different concentrations of M2 (from 35 μg/mL to 280 μg/mL) for 24 hours in a 37 °C cell culture incubator. It was found that 90% cell viability was observed after 24 hours of incubation with M2 at a concentration of 280 μg/mL (eight times higher than that of the normal concentration), indicating the superior biocompatibility of this nano-oxygen sensor (Figure 11). This result is compared with similar probes by Fengyu Su et al. (higher than 80%) and Tingting Pan (higher than 88% with three times higher than normal concentration), suggesting that no obvious cell toxicity is observed [39,48]. It should be pointed out that these micelles did not have cell permeability during the eight-hour experimental period in which we performed cell respiration testing. Therefore, the sensor can be utilized for extracellular applications.

3.9. Bacterial Drug Resistance Test

We also did experiments on the effects of different common antibacterial drugs (ampicillin and antimycin A) on bacterial respiration to test the drug screening ability of oxygen sensors. Ampicillin is a common antibiotic that inhibits bacterial growth by inhibiting the electron transport chain (ETC) [49,50]. Antimycin A is also a common bacteriostatic drug that inhibits the growth of various cells by stimulating oxidative stress-mediated death [49,50]. We tested the effects of two drugs on the oxygen consumption ability during the respiratory process of E. coli with constant cell densities of 2 × 10⁶ cfu/mL, wherein the concentration of antimycin A was from 0.16 to 20 μg/mL, and the concentration of ampicillin was from 20 to 50 μg/mL. A total of 3 μg/mL (2.6 μM) PtTFPP in the LB medium was used for observing antibiotics’ concentration-dependent oxygen consumption velocities (Figure 12). When there is no drug in medium, the bacteria breathe normally and consume oxygen completely within 60 min. As the drug concentration increases, oxygen consumption was inhibited significantly. When the activity of the two drugs was compared, it can be found that antimycin A can inhibit oxygen respiration more significantly than ampicillin. Thus, the oxygen sensors can be used to differentiate antibiotics’ activity by using oxygen consumption as an indicator. It was noted here that after reaching the emission maximum when ampicillin was used as the antibiotics, the emission of PtTFPP decreased. Even the detailed reason for this is not fully understood; one possibility is that most likely, there are some interactions between the ampicillin with PtTFPP.

4. Conclusions

Multi-arm amphiphilic block copolymers were prepared, and their micelles were used to encapsulate PtTFPP to enable the use of PtTFPP for oxygen sensing in biological conditions. The micelles prepared by using these block copolymers were around 40 to 200 nm, depending on the molar ratios of the hydrophobic and hydrophilic segments in the block copolymers. The micelles showed high storage stability. The PtTFPP encapsulated by micelle shows reasonable high quantum yields which are from 0.06 to 0.159. The micelles’ response time is 20–30 s from deoxygenated solutions to oxygenated solutions, showing a rapid response to changes of dissolved oxygen concentrations. These new micellar biosensors were capable of in situ monitoring dissolved oxygen and oxygen consumption of E. coli and living cells. The sensors were further demonstrated to be useful for drug screening through differentiating the influence of drug doses and species (antimycin A and ampicillin as two representative examples) on cells’ metabolic activity. Since the sensor is in liquid form, it is easily used in many general commercial instruments, such as plate readers, and can be used for further biological understanding, disease diagnosis, and metabolic investigation.

In this article, we introduce a novel multi-arm amphiphilic oxygen-sensitive probe for real-time measurement of dissolved oxygen concentration in extracellular or aqueous solutions. Compared with previous studies [27], ring-opening polymerization has the advantages of simple operation, high yield, and low cost. Furthermore, unlike linear polymers, multi-arm PEG that has higher terminal group concentrations than conventional linear polymers [35] is used, which enables the further modification or functionalization of the multi-arm polymers. Besides, this sensor shows excellent biocompatibility and faster response time for both cell and bacterial respiration measurements, indicating a good prospect in the biological field.