R. cerastii strain IEGM 1243 from the Regional Specialized Collection of Alkanotrophic Microorganisms (IEGM; WFCC-WDCM 768; UNU/CKP 73559/480868; www.iegmcol.ru accessed on 31 May 2023) was used in this study. The strain IEGM 1243 was isolated from soil, lake shore Kumnylor, Tyumen region, Russia. In preliminary studies it showed IBP-degrading (100 mg/L) activity in the presence of 0.1% n-hexadecane (data not shown). Moreover, IEGM 1243 uses n-hexadecane as a sole carbon source and is resistant to Mo⁶⁺ (5.0 mM). The experiments were conducted in 250 mL Erlenmeyer flasks containing 100 mL of “RS” (Rhodococcus Surfactant) mineral salt medium. The composition of the medium per liter included: KNO₃ 1.0 g, K₂HPO₄ 1.0 g, KH₂PO₄ 1.0 g, NaCl 1.0 g, MgSO₄ × 7H₂O 0.2 g, CaCl₂ × 2H₂O 0.02 g, FeCl₃ 0.001 g, and 0.1% (v/v) trace element solution (1.5 g/L FeCl₃ × 7H₂O, 0.1 g/L H₃BO₃, 0.01 g/L ZnSO₄ × 7H₂O, 0.05 g/L Co (NO₃)₂ × 6H₂O, 0.005 g/L CuSO₄ × 5H₂O, and 0.005 g/L MnCl₂ × 4H₂O). IBP (C₁₃H₁₇O₂Na; CAS: 31121-93-4; (RS)-2-(4-(2-methylpropyl)phenyl)propanoic acid) and DCF (C₁₄H₁₀Cl₂NNaO₂, CAS: 15307–79-6, 2-(2-(2,6-dichlorophenylamino)phenyl)acetic acid) were used as sodium salts (Sigma-Aldrich, St. Louis, MO, United States; Glentham Life Sciences, Corsham, United Kingdom) and added to the mineral salt medium, each at a final concentration of 50 mg/L. Stock solutions of IBP and DCF (50 mg/mL) were prepared individually using water and stored at 4°C before use. We opted for this high concentration of NSAIDs due to the potential application of Rhodococcus-based biocatalysts in local post-treatment technologies during drug manufacturing, where influents contain elevated pharmaceutical levels. Moreover, previous research on biodegradation and the diverse effects of pharmaceuticals on bacteria supports the use of higher dosages (Alobaidi et al., 2021; Żur et al., 2021; Mohamed et al., 2023; Suleiman et al., 2023). D-glucose was employed as an additional carbon and energy source at a concentration of 0.5 g/L. Cells of R. cerastii IEGM 1243 grown for 3 days in LB broth (Himedia Laboratories Pvt. Limited, Maharashtra, India) and washed twice with phosphate buffer (pH 7.0) were added to the medium to a concentration of 0.96 × 10⁷ cells/mL. The experiments were conducted at 160 rpm and 28°C.

The following treatments were used in the experiments: (1) mineral salt medium + R. cerastii IEGM 1243 + 0.5 g/L glucose (biotic control), (2) mineral salt medium + R. cerastii IEGM 1243 + 0.5 g/L glucose + 50 mg/L IBP (positive control 1), (3) mineral salt medium + R. cerastii IEGM 1243 + 0.5 g/L glucose +50 mg/L DCF (positive control 2), (4) mineral salt medium + R. cerastii IEGM 1243 + 0.5 g/L glucose + NSAID mixture (50 mg/L IBP + 50 mg/L DCF) (positive control 3), (5) mineral salt medium +0.5 g/L glucose + 50 mg/L NSAID (abiotic controls), (6) mineral salt medium + autoclaved R. cerastii IEGM 1243 + 0.5 g/L glucose + 50 mg/L NSAID (sorption controls). The abiotic controls were necessary to elucidate the abiotic degradation of the pharmaceuticals, while the sorption controls were essential for assessing the removal of the pharmaceuticals by sorbing them onto bacterial cells. The biotic (negative) control was essential to differentiate the effects of NSAIDs on bacterial cells from the natural variations in their life cycle.

The catalase activity of R. cerastii IEGM 1243 was determined spectrophotometrically following the protocol described by Gogoleva et al. (2012). Bacterial cells cultured with glucose with and without NSAIDs were centrifuged at 3,000 rpm for 5 min, washed with phosphate buffer (pH 7.0), and resuspended in the same buffer to obtain an optical density (OD₄₉₂) of 0.2. Next, a 0.00125 M hydrogen peroxide (H₂O₂) solution (1 mL) was added to 200 μL of the cell suspension, and the mixture was incubated for 10 min at room temperature. To stop the decomposition of H₂O₂ by catalase, 100 μL of 2 M HCl solution was added. Subsequently, a 0.025 M potassium iodide solution (1 mL) was added to the resulting mixture, gently mixed, and centrifuged at 3,000 rpm for 15 min. Distilled water was used in control samples instead of cell suspensions. Finally, the absorbance of the supernatant was measured at 492 nm using a Lambda EZ201 spectrophotometer (Perkin-Elmer, Waltham, MA, United States).

The catalase activity was calculated using the equation:

where A_cat represents the catalase activity in μM/min × OD; OD_v is the optical density of the supernatant in the experimental sample (in arbitrary units of optical density); OD_c is the optical density of the supernatant in the control sample (in arbitrary units of optical density), T is the incubation time of bacteria in the presence of H₂O₂ (10 min); and OD_b is the optical density of the bacterial suspension (OD₄₉₂ 0.2). The catalase activity was expressed as a percentage of the initial level.

Cell respiration activity was evaluated using a 10-channel Micro-Oxymax^® respirometer (Columbus Instruments, Columbus, OH, United States) (Ivshina et al., 2019). The experiments were conducted in 300 mL Micro-Oxymax glass flasks containing 100 mL of mineral salt medium with constant agitation (160 rpm, 28 ± 2°C). The rate (μL/h) and accumulation (μL) of oxygen consumption were measured. Automated recording of respiratory activity was performed every 30 min over a period of 7 days.

The removal rates of IBP and DCF were determined using high-performance liquid chromatography (HPLC) on an LC Prominence chromatograph (Shimadzu Corporation, Kyoto, Japan) equipped with a reverse-phase C18 column, 25 cm × 4.6 mm, 5 μm (Supelco Inc., Bellefonte, PA, United States) and a diode array detector. Mobile phase consisted of a phosphate buffer solution (pH 5.0) and acetonitrile, mixed in a 60:40 ratio. Elution was performed isocratically at a flow rate of 0.5 mL/min, and detection was carried out at a wavelength of 254 nm. The injection volume was 20 μL, and the column thermostat temperature was set at 40°C. Under these conditions, the retention times for IBP and DCF were determined to be 9.28 ± 0.03 and 18.70 ± 0.02 min, respectively. Prior to analysis, samples were prepared by centrifuging for 5 min at 10,000 rpm. The resulting supernatants were then filtered using a 0.20 μm nylon membrane filter (Nantong FilterBio Membrane Co., Ltd., Nantong, China). The concentrations of DCF and IBP were calculated by comparing the areas under the peaks with those of the standard solutions.

For cell analysis and quantification, ImageJ software (US National Institutes of Health, Bethesda, Maryland, United States) was employed. A detailed protocol for image processing can be found here (https://www.allevi3d.com/livedead-assay-quantification-fiji/ accessed on 31 May 2023).

Statistical analyses were performed to determine the differences in the experimental data using one-way ANOVA followed by two-sided Dunnett’s t-test. The software used for conducting the statistical tests was SPSS 23.0 (IBM, Armonk, New York, United States). The significance level for the results was set at p < 0.05. Each experiment was performed in triplicate to ensure accuracy and reproducibility of the results. For presenting the data from repeated experiments, the values were expressed as mean ± standard deviation.

According to our data, R. cerastii IEGM 1243 exhibited moderate biodegradative activity towards the tested pharmaceutical compounds (Figure 1). When considering individual NSAIDs, the residual content of IBP after a 7 days period was 68.4%, while that of DCF was 75.9%. Interestingly, when using the combination of NSAIDs, the degradation of IBP was higher, with a residual content of 40.4% after 7 days. However, the degradation of DCF in this case was negligible, with a residual content of 89.7%. No losses of substances were observed in abiotic and sorption controls, indicating the biocatalytic nature of the degradation process.

In our previous studies, we confirmed that R. cerastii strain IEGM 1278 was capable of completely cometabolizing 100 mg/L IBP within 6 days in the presence of n-hexadecane (Ivshina et al., 2021b). Other research showed that IBP could be efficiently degraded at high concentrations (200–1,000 mg/L) by both Gram-negative strains (Sphingomonas sp. Ibu-2, Variovorax sp. Ibu-1) and Gram-positive strains [Bacillus thuringiensis B1(2015b), Citrobacter freundii PYI-2, C. portucalensis YPI-2, Nocardia sp. NRRL 5646, Patulibacter sp. I11] (Jan-Roblero and Cruz-Maya, 2023). On the other hand, DCF is more recalcitrant to biodegradation. Previously, we isolated R. ruber strain IEGM 346, which was able to degrade only 50% of 50 mg/L DCF within 60 days (Ivshina et al., 2019). In other reports, only Gram-negative Klebsiella sp. KSC exhibited the capability to degrade high concentrations of DCF (70 mg/L) within three days, while other strains such as Pseudomonas moorei KB4, Labrys portucalensis F11, Raoultella sp. DD4 degraded much lower concentrations (ranging from 0.5 to 7 mg/L) over up to 28 days (Wojcieszyńska et al., 2023). The results obtained in our study indicate that further optimization of biodegradation conditions for IEGM 1243 is necessary, as our findings do not stand out in this regard.

However, it is important to note that the biodegradation pattern was not the primary focus of our research. We previously outlined the putative biodegradation pathways of DCF and IBP in Ivshina et al. (2019, 2021b). Additionally, Guzik and Wojcieszyńska’s (2019) thorough review covered the microbiological degradation pathways of NSAIDs. Instead, our study primarily concentrated on the physiological and morphological responses of rhodococci when exposed to NSAIDs, which are presented below.

Concurrently with the biodegradation experiments, we conducted quantitative assessments of total cell count and cell viability percentage (Figure 1; Supplementary Figures S1, S2). The initial cell count was determined to be 0.96 cells/mL × 10⁷. Throughout the duration of the experiment, an increase in biomass was consistently observed across all experimental conditions, with the most significant growth recorded in both the control and in the presence of IBP. The highest cell count was attained when IBP was present, peaking at 3.32 cells/mL × 10⁷ after 4 days of the experiment. In parallel, the viability of rhodococci in the control sample exhibited an upward trend until day 4, with viability levels rising from 59.5 to 94.9%, followed by a decline to 54.7% on day 7. In the presence of IBP, the maximum cell viability was reached on day 5 (80.2%), gradually decreasing to 65.9% on day 7. Notably, the overall cell count in the presence of DCF and the NSAID mixture on day 7 of the experiment amounted to 2.1 and 1.59 cells/mL × 10⁷, respectively. Interestingly, these conditions exhibited the highest proportion of viable cells at the end of the experiment.

Analysis of cellular respiration in the presence of NSAIDs revealed the following patterns (Figure 2). Both in the biotic control and in the presence of IBP, the lag phase of IEGM 1243 cells was less than 24 h (Figure 2A). The maximum rate of oxygen consumption in the control was reached after 1.5 days of the experiment, amounting to 4753.7 μL/h. In contrast, in the presence of IBP, this parameter was recorded after 2.5 days of the experiment, with a significantly higher value of 13125.4 μL/h.

The transition of rhodococci to the stationary growth phase (Figure 2B) correlated with a slowdown in the biodegradation of NSAIDs (Figure 1). DCF and the mixture exerted a suppressive effect on the metabolic activity of rhodococci for a period of 3–4 days. The maximum rate of oxygen consumption in the presence of DCF was 12580.9 μL/h after 4.5 days of the experiment. Under the combination of IBP and DCF, the peak oxygen consumption was observed on day 6, measuring 12936.4 μL/h.

Cumulative oxygen uptake analysis demonstrated that cells exhibited higher metabolic activity in the presence of NSAIDs (Figure 2B). For instance, on day 9, the total amount of oxygen consumed in the control was 611911.4 μL, whereas in the presence of IBP, DCF, and their mixture, these values were 956805.2 μL, 1,043,623 μL, and 1,070,803 μL, respectively. The increased amount of oxygen consumed in the presence of DCF and the NSAID mixture after 7 days of the experiment correlates with data on higher cell viability under these conditions (Figure 1; Supplementary Figure S2).

Catalase, as a vital antioxidant enzyme, plays a crucial role in the reduction of hydrogen peroxide (H₂O₂) to oxygen (O₂) and water (H₂O). This enzyme holds significant importance in bacterial stress response and the degradation of xenobiotics (Wang et al., 2000, 2020; Kaushal et al., 2018). Rhodococcus spp. are well-known for their high abundance of catalases (Yuan et al., 2021), which makes them robust and adaptable microorganisms. In the context of xenobiotic degradation, including the biodegradation of NSAIDs, the high levels of catalases in rhodococci provide significant benefits. The presence of NSAIDs can lead to the production of harmful reactive oxygen species in the bacteria. However, thanks to their abundant catalases, Rhodococcus spp. may efficiently neutralize these ROS, ensuring their metabolic activity and successful degradation of the xenobiotics.

The initial catalase activity of R. cerastii IEGM 1243 was 0.66 relative units corresponding to 100%. Within the first day, a decrease in catalase activity was observed in all tested variants. However, the most significant (p < 0.05) changes in catalase activity of rhodococci were observed in the presence of DCF and the NSAID mixture (Figure 3). The decrease in catalase activity under these conditions during the first 3 days can be attributed to the cells being in the lag phase of growth (refer to Figure 2A). Interestingly, on the second day, there was an increase in catalase activity in the presence of NSAIDs, while in the control group, the activity continued to decrease. The opposite pattern was observed on the third day. The increase in catalase activity in the samples with NSAIDs on day 4 can be explained by the bacteria entering the exponential growth phase, active degradation of NSAIDs (as seen in Figure 1), and possible accumulation of metabolites. The decrease in catalase activity in all samples on day 7 could be associated with the transition to the stationary growth phase, slowing down of cellular metabolism, and the formation of multicellular aggregates (Supplementary Figure S2) (Wood and Sørensen, 2001). Moreover, it is hypothesized that cell aggregation enhances intercellular synergy, leading to a reduced need for cells to secrete the necessary enzymes (D’Souza et al., 2023). By forming multicellular aggregates, bacterial cells can establish communication networks that facilitate the exchange of genetic material and metabolic by-products. This can lead to a more efficient distribution of tasks among the cells, allowing some individuals to focus on enzyme production while others specialize in different functions. Consequently, this division of labor within the aggregated community can reduce the overall demand for enzyme secretion by individual cells.

Studies on cell morphology and morphogenesis are of great importance for understanding cell growth, reproduction, and adaptation to the environment (Young, 2006; Levin and Angert, 2015; Ojkic et al., 2019; Shi et al., 2021; Govers et al., 2023). In this study, we analyzed the morphometric parameters of live cells of R. cerastii IEGM 1243 exposed to NSAIDs, and the results are presented in Figures 4–7, Table 1, and Supplementary Figures S3–S5. Initially, in their native state, the R. cerastii IEGM 1243 cells exhibited an elongated rod-shaped morphology (Figures 4A,B) with distinct peaks and valleys (Figure 4D), as well as a rough cell surface.

Rod-shaped bacteria have the ability to readily adapt their shape in response to changes in the surrounding environment (Chang and Huang, 2014; Ojkic et al., 2019). In our study, we observed that the control cells underwent shape alterations on day 4, with a reduction in length and an increase in width (Figure 5 and Table 1). However, according to height measurements, no significant differences in cell sizes were observed between the NSAID-treated cells and the control (Figures 6, 7 and Table 1). Notably, when exposed to DCF and the NSAID mixture, cells produced a significant amount of extracellular polymeric substances, which hindered the assessment of cell profile and height (Figure 7).

On the seventh day, significant differences in morphometric parameters of cells in the presence of NSAIDs were observed, particularly a decrease in surface area, volume, and an increase in the surface area-to-volume ratio (SA/V) (Table 1). SA/V is an important physical parameter that regulates the influx and efflux of substances and metabolites (Ojkic et al., 2022). The cell size, shape, and SA/V are interconnected, and individual cells can change their size and shape to achieve an optimal SA/V (Harris and Theriot, 2018; Shi et al., 2021). Previously, we observed alterations in the SA/V ratio in rhodococci exposed to pharmaceuticals (Ivshina et al., 2019; Tyumina et al., 2019; Ivshina et al., 2021b, 2022b). For example, when exposed to toxic substrates such as DCF or naproxen, cells tend to reduce their SA/V to minimize the exposed cell surface for contact with the stressor (Ivshina et al., 2019, 2022b). Studies by other authors have shown that in the presence of ampicillin, Escherichia coli cells maintain an ellipsoidal shape and a low SA/V to reduce their metabolic activity, conserve energy, and prevent division (Uzoechi and Abu-Lail, 2019). The reduction in metabolic activity, in turn, leads to cell persistence and resistance (Ojkic et al., 2022). On the other hand, in the presence of less toxic compounds or under carbon source limitation, SA/V ratio may increase for more efficient contact between cells and a substrate (Ivshina et al., 2019, 2021b). In the short term, a cell is capable of regulating its shape by adjusting turgor pressure. However, over longer time scales, changes in cell size are predominantly determined by alterations in cell width (Oldewurtel et al., 2021). In our case, on day 7, the NSAID-treated cells reduced their width (p < 0.05), which ultimately resulted in smaller cell volume and larger SA/V. Possible reasons for the change in SA/V ratio include the availability of precursors for peptidoglycan biosynthesis and the presence of inhibitors of fatty acid biosynthesis (Harris and Theriot, 2018). Additionally, cell size is influenced by the levels of transport proteins, metabolic activity, and the accumulation of cell division protein (FtsZ) (Ojkic et al., 2019; Bertaux et al., 2020).

The effect of IBP on the ultrastructure of R. cerastii IEGM 1243 cells was investigated using transmission electron microscopy (TEM) analysis. TEM provides high-resolution imaging of cellular structures, allowing for detailed observations of cellular ultrastructure. Comparative analysis of ultrathin sections revealed that the morphological traits of rhodococci were similar in cultures developed under both control and IBP-containing conditions (Figure 8; Supplementary Figures S6–S8). As evident from the Figure 8E, strain IEGM 1243 exhibits an apical mode of cell elongation, resulting in the increased accumulation of new cell wall material at the cell poles (Donovan and Bramkamp, 2014). The cells exhibited characteristic features, including the presence of an outer capsular layer, which was visually discernible as protrusions. Furthermore, they displayed a stratified cell wall with an outer layer that appeared electron-dense, indicating the existence of a well-defined cell envelope. Additionally, the cytoplasmic membrane remained intact, indicating the overall structural integrity of the cells.

In both the control and IBP-treated cells, TEM revealed the presence of two distinct types of structures: electron-transparent bodies, likely lipid inclusions, and electron-dense bodies, believed to be polyphosphates. Intracellular lipid inclusions important energy storage compounds that fulfill carbon requirements and maintain redox homeostasis, enabling bacteria to survive for extended periods (Alvarez et al., 1996, 2000; Mallick et al., 2021). Furthermore, they play a crucial role in the pathogenesis of mycobacteria (Mallick et al., 2021). Rhodococcus spp. commonly utilize polyhydroxyalkanoates, triacylglycerols, and glycogen as intracellular carbon reserves (Hernández et al., 2008; Cappelletti et al., 2020).

Nile Red staining of the bacteria revealed the presence of a small amount of lipid inclusions, mainly located at the periphery of the cells, at the initial time point (Figure 9). By day 4 of the experiment, there was a tendency for the accumulation of lipid inclusions in all treatments, particularly evident in the control and in the presence of IBP. The number of inclusions reached up to 10 per cell, predominantly localized in the central part of the cell. In the case of DCF and the NSAID mixture, the quantity and size of lipids were smaller, and the inclusions were often located at the periphery of the cells. After 7 days of observation, the control cells maintained a significant amount of lipid inclusions; however, individual cells without intracellular lipids were also observed. When exposed to NSAIDs, cells primarily depleted their lipid reserves, with individual lipid granules located at the periphery of the cells (IBP and NSAID-mixture) or at the center of the cells (DCF). The red fluorescence around the cell perimeter could be associated with the lipids of the cell membranes (Presentato et al., 2018).

According to a previous study (Wältermann et al., 2005), the process of lipid-body synthesis involves the formation of small lipid droplets that remain attached to membrane-associated enzymes. Over time, these droplets aggregate and merge, leading to the formation of larger structures known as membrane-bound lipid prebodies. Eventually, these lipid prebodies are released into the cytoplasm, becoming mature entities within the cell. Our current study corroborates these findings (Supplementary Figure S9). For instance, at the beginning of the experiment, IEGM 1243 cells grown in the presence of glucose (control) exhibited small inclusions at the periphery, which likely represent small lipid droplets. As time progressed, these droplets transitioned into the cytoplasm, forming larger lipid prebodies in the early stages and eventually maturing into large lipid inclusions on days 2 and 3. The formation of lipid inclusions in the presence of IBP followed a similar scenario, while under the influence of DCF and the NSAID mixture, the maturation of lipid inclusions occurred on day 4.

When IBP was present, cells exhibited the formation of intracellular membrane-like structures (Figure 8D; Supplementary Figure S7). These structures appeared as loop-like formations and are believed to be primarily involved in the transportation of complex compounds and their subsequent degradation through the action of membrane-bound enzymes (Royes et al., 2020; Thi Mo et al., 2022). Similar structures have been observed in previous studies involving rhodococci grown in the presence of benzoate (Solyanikova et al., 2017), oleanolic acid (Luchnikova et al., 2022), as well as liquid and solid n-hexadecane (Thi Mo et al., 2022). This suggests a common mechanism or adaptive response in the formation of these membrane-like structures across different environmental conditions and compound exposures.

Polyphosphates were represented as compartments mainly located at the cell poles (Figures 8A,C). Similar observations of polyphosphates were made in cells of R. rhodochrous IEGM 1362 grown in a nutrient-rich medium and in the presence of (−)-isopulegol (Ivshina et al., 2022a), R. rhodochrous IEGM 757 in the presence of oleanolic acid (Luchnikova et al., 2022), and R. erythropolis N9T-4 in a basal medium without any additional carbon, nitrogen, sulfur, and energy sources (Yoshida et al., 2017). In the latter case, the authors referred to these compartments as oligobodies and confirmed their high phosphorus and potassium content using X-ray spectroscopy. Phosphorus-rich granules are composed of linear chains of polyphosphate and cations such as magnesium, potassium, and calcium (Keim et al., 2005). Polyphosphate inclusions serve as an internal phosphate reserve in cells and play an important role in the adaptive mechanisms of bacteria under suboptimal environmental conditions. In our case, particularly significant accumulations of phosphorus were observed in cells grown on mineral salt agar supplemented with IBP. TEM-EDX analysis showed distinct zones of increased phosphorus accumulation at the cell poles, along with significant amounts of potassium and magnesium (Figure 10; Supplementary Figures S10–S13). Furthermore, the elevated potassium content in the external environment in the presence of IBP may indicate its efflux from the cells due to the disruption of cell membrane permeability (Luchnikova et al., 2022). However, it is essential to note that further investigations are needed to fully understand the underlying mechanisms behind these observations.