1. Introduction
Nontuberculous mycobacteria (NTMs) are common in nature and exist in water sources, soil, and vegetation as environmental organisms. Currently, more than 200 species are recognized as NTMs, and new species are identified every year [
1,
2]. Although relatively few species of NTMs cause disease in human, the incidence of human NTM infection appears to have increased significantly in recent decades due to causes such as an aging population, immunosuppression, and broad-spectrum antibiotic use [
1,
2,
3]. In addition, most NTMs that cause infections are challenging to treat and worsen the patient’s prognosis because they are resistant to the first-line drug for tuberculosis [
4]. According to their growth rate, NTMs that grow on the media within or after 7 days are defined as rapidly growing mycobacteria (RGM) and slowly growing mycobacteria (SGM), respectively [
5]. Of these,
Mycobacterium mucogenicum (
Mmuc) belongs to the RGM group [
6].
Mmuc has rarely been reported to infect humans, and cases are most commonly seen in immunocompromised patients [
6]. Their isolation may be considered clinically important because RGMs are recovered from various environmental sources, including water, and they survive by forming biofilms and by interacting with protozoa [
6,
7]. After
Mmuc was first reported as a
Mycobacterium chelonae-like organism (MCLO) in a peritonitis outbreak in 1982,
Mmuc strains have been detected in a wide range of infection sites, such as the central nervous system, skin, and lungs [
8,
9,
10]. In particular, the most clinically significant infections are posttraumatic wound infections and catheter-related sepsis [
6,
9]. In particular,
Mmuc has been identified as a water contaminant in hospitals. Possible causes of these opportunistic infections include a poorly disinfected organ indwelling catheter and contamination of the water used for surgeries, such as dialysis, and the water used for laboratory testing [
8]. Furthermore,
Mmuc was shown to be more resistant to chlorine than other bacteria, including
M. fortuitum and
Escherichia coli, and therefore, it has a higher probability of contamination [
3]. Similar to other pathogenic NTMs,
Mmuc can physically display various colony morphologies, including rough (R) and variable smooth (S) types [
11,
12]. This colony-based distinction is dependent on the presence or absence of surface-associated glycopeptidolipids (GPLs) [
13].
Mmuc is classified as the S-type if GPLs are present and as the R-type if GPLs are not [
11,
13]. The presence or absence of GPLs is related to meaningful pathological aspects such as biofilm formation or sliding motility, interaction with host cells, intracellular trafficking in macrophages, and virulence, which affects the clinical severity of the infection [
11]. In general, the R-type of
Mycobacterium spp. causes a more severe inflammatory response in macrophages or mice than the S-type [
13,
14]. The morphological classification of
Mmuc and patient case reports have been studied [
12], but no studies have assessed immunological differences based on morphological analyses.
In the present study, the immune responses to the standard ATCC49650 strain (Mmuc-ATCC) and the R-type (Mmuc-R; KMRC 00136-76003) and S-type (Mmuc-S; KMRC 00136-76002) clinical strains were first compared. Here, we examined immune responses based on the fact that virulent mycobacteria can survive and replicate within macrophages as intracellular bacteria. Additionally, we examined the steps of apoptotic cell death because it is regarded as an innate intracellular response designed to limit the multiplication of intracellular pathogens. In addition, we attempted to compare and analyze the cytokine patterns of each strain according to the activation of macrophages. Finally, extracellular flux analysis was performed to assess the modulation of the energy metabolism of BMDMs infected with Mmuc-ATCC, Mmuc-S, and Mmuc-R.
3. Discussion
Mmuc strains are isolated from water and capable of causing opportunistic infections and can contaminate chemical devices and equipment, which may provide a mechanism for bacteria to enter the bloodstream through wounds. Immunosuppressed patients are at risk of significant morbidity and mortality from exposure to these healthcare environmental pathogens [
8]. This study demonstrates for the first time the differences in the innate immune responses, signaling pathways, virulence, and metabolic changes of the S- and R-type clinical strains and the ATCC standard strain of
Mmuc. The
Mmuc-R grew as wrinkled and rough colonies on Middlebrook 7H10 agar, whereas
Mmuc-ATCC and
Mmuc-S grew into smooth, dome-shaped colonies. In a similar manner, electron microscopy confirmed that the surface of
Mmuc-R was more wrinkled and rough, while the surfaces of the strains
Mmuc-ATCC and
Mmuc-S were less wrinkled. Previous studies on the colony morphotypes of NTMs, including
M. abscessus, have shown that rough morphotypes lacking GPLs induce a greater inflammatory response and more virulence than smooth morphotypes [
5,
13].
Our results clearly show that the intracellular survival of
Mmuc-R within macrophages was higher than that of both S-type strains. According to our data, although
Mmuc-R had a better chance of surviving in BMDMs than S-type
Mmuc, whether the bacterial counts of both types of
Mmuc increased until 5 days after infection could not be determined. In the case of
Mmuc-R, the number of initially infected bacteria was maintained for 5 days, and the number of S-type strain bacteria decreased severely. Actually, infection of mouse BMDMs with pathogenic
Mycobacterium spp. does not always increase bacterial numbers. In particular, in the case of
M. abscessus, depending on the experimental condition or strain type, the number may decrease slightly upon macrophage infection [
21,
22]. Perhaps this difference is because even if the bacteria were sufficiently multiplied within the cell, they were released to the outside due to cell death and then removed during washing. In addition, although the bacteria are continuously multiplied, the bacterial numbers may nearly appear to be maintained because they are removed by various macrophage suppression mechanisms. Further research should be focused on elucidating the underlying mechanisms in more detail.
Another predictive marker of intracellular pathogen virulence is the confirmation of cytokine secretory capacity. Highly pathogenic bacteria generally induce the secretion of inflammatory cytokines, but they are not always correlated. All strains of
Mmuc upregulated the proinflammatory cytokines TNF-α, IL-6, and IL-12p40 upon infection, and their levels increased in an MOI-dependent manner. In particular,
Mmuc-R induced relatively high inflammatory cytokine secretion and was the only investigated strain to induce IL10 production, which is an anti-inflammatory cytokine. Almost all NTMs, including
M. abscessus and
M. massiliense, can be divided into rough and smooth types depending on the presence or absence of GPLs, and most R-type strains are highly pathogenic and have high cytokine secretory capacity [
23]. In our results, cytokine secretion was higher in the
Mmuc-R, and the pattern was the same as that for the general NTM strain.
Apoptosis is a programmed cell death mechanism controlled by host balance with a cell signal generated by the cell itself, and necrosis is an unprogrammed cell death caused by external stimuli such as infection. In addition, many reports suggest that necrosis is preferred over apoptosis within macrophages infected by virulent pathogens [
16]. In this study,
Mmuc-R promoted late cell death and a necrotic phenotype of macrophages, suggesting that
Mmuc-R is more pathogenic during infection and may sufficiently proceed to host cell lysis and tissue injury. Upon comparing the immune responses of
Mmuc strains,
Mmuc-R was more virulent than S-type
Mmuc in all aspects of intracellular viability; the production of cytokines is important for the immune response, induction of cell death, and signaling pathways, similar to the immune response of other RGMs. Furthermore, our study showed that the clinical strains were more virulent than
Mmuc-ATCC.
Several intracellular bacteria have been reported to shift the bioenergetic metabolism of macrophages via a specific approach that benefits the pathogen [
20]. The mechanism by which NTM rewires macrophage energy metabolism to facilitate survival is poorly characterized. Here, we used extracellular flux analysis to explore the modulation of the energy metabolism of BMDMs infected with
Mmuc-ATCC,
Mmuc-S, and
Mmuc-R.
Mmuc infection induced a metabolic shift of infected BMDMs to a higher energetic level, and this shift was observed for all the
Mmuc strains tested (
Figure 7 and
Figure 8).
Mmuc accelerated OXPHOS to enter a metabolic energetic state and consequently increased the basal respiration rate, maximum respiration, and spare respiratory capacity of the macrophages (
Figure 7C,D,G). However, we also showed that
Mmuc infection upregulated proton leakage and that the ATP production levels did not significantly differ between the control and infection groups (
Figure 7E,F). We also showed that
Mmuc-R infection significantly increased the nonmitochondrial oxygen consumption rate, unlike
Mmuc-ATCC and
Mmuc-S (
Figure 7B). The nonmitochondrial oxygen consumption rate increases in the presence of reactive oxygen species (ROS) and reactive nitrogen species (RNS), and mitochondria are damaged due to the deleterious effects of these reactive intermediates [
24,
25]. Given these results,
Mmuc-R more negatively affects the bioenergetic health of macrophages than
Mmuc-ATCC or
Mmuc-S. Upregulation of the aerobic glycolysis response is considered a hallmark of proinflammatory signals in both myeloid and lymphoid cells [
25]. We showed that infection with all
Mmuc strains increased glycolysis in infected macrophages, as evidenced by the induced glycolysis and glycolysis capacity rate (
Figure 8B,C). In particular,
Mmuc-R infection significantly increased both glycolysis and the glycolytic capacity rate, unlike the S-type
Mmuc strains. However, we also showed that
Mmuc infection reduced the glycolytic reserve rate and the glycolytic reserve as percentage parameters (
Figure 8D,F). The glycolytic reserve rate is an important bioenergy source that is activated in response to increases in both glycolysis and the glycolysis capacity rate. Therefore, the reduction in the glycolytic reserve may limit macrophage energy supply when BMDMs need ATP, especially after
Mmuc-R infection.
4. Materials and Methods
4.1. Bacterial Culture
The Mmuc reference strain (ATCC49650) was purchased from American Type Culture Collection (ATCC, Manassas, VA, USA). The two clinical strains (KMRC 00136-76002 and KMRC 00136-76003) were obtained from The Korean Mycobacteria Resource Center (KMRC) of the Korean Institute of Tuberculosis (Osong, Korea). Mmuc strains were grown in Middlebrook 7H9 medium (Difco Laboratories, Detroit, MI, USA) supplemented with 10% oleic albumin dextrose catalase (OADC; BD Biosciences, San Diego, CA, USA) and 0.5% glycerol at 37 °C for 7 days. The number of colony-forming units (CFU)/mL was determined on 7H10 agar supplemented with OADC at 37 °C.
4.2. Generation of Bone Marrow-Derived Macrophages
Mouse bone marrow-derived macrophages (BMDMs) were isolated from the femurs and tibias of 7-week-old female C57BL/6 mice (DBL, Chungcheongbuk-do, Korea) [
26]. The cells were washed with Dulbecco’s modified Eagle’s medium (DMEM; Biowest, France; L0103-500) and then centrifuged at 440×
g for 3 min. The pellet was resuspended in 40 mL of DMEM containing glutamine, 1% antibiotic–antimycotics (Biowest, France; L0010-100), 10% fetal bovine serum (FBS; Welgene Co.; Daegu, Korea, S001-01), and 10 ng/mL recombinant mouse M-CSF (JW CreaGene). BMDMs were differentiated in complete medium for a total of 6 days at 37 °C in the presence of 5% CO
2. On day 3, the media were added, and the culture was maintained for an additional 3 days. After 6 days, nonadherent cells were removed, and the differentiated macrophages were detached by adding trypsin-0.25% EDTA (Biowest, France; L0931-100) for 5 min in an incubator. The detached cells were centrifuged at 440×
g for 3 min, resuspended, and seeded in complete DMEM.
4.3. Growth of M. mucogenicum in BMDMs
BMDMs were plated at a density of 1.5 × 105 cells per 48-well plate. After 24 h, BMDMs were infected at a multiplicity of infection (MOI) of 10 for 4 h at 37 °C. After 4 h, the cells were washed with PBS to remove all extracellular bacteria and further cultivated in fresh complete medium for 5 days. At days 0, 1, 2, 3, 4, and 5, the culture supernatants were aspirated, and cells were lysed by 0.05% Triton X-100 (Sigma, St. Louis, MO, USA). Then, the lysates were plated in tenfold serial dilutions onto 7H10 agar (Becton Dickinson, Franklin Lakes, NJ, USA) to quantify the number of viable bacteria. Colonies were counted after 4 days of incubation at 37 °C. The resultant values were reported as the mean log10 CFU ± standard deviation (SD) per 1.5 × 105 cells.
To determine whether Mmuc exhibits similar infection kinetics during the first 4 h, BMDMs were infected at an MOI of 10 per 5 × 105 cells for 30, 60, 120, 180, and 240 min at 37 °C. After incubation, the cells were washed 2–3 times with PBS prewarmed serum-free media to remove extracellular bacteria. The cells were lysed with distilled water containing 0.05% Triton X-100, and the lysates were plated in tenfold serial dilutions onto 7H10 agar to quantify the number of infected bacteria. Colonies were counted after 4 days of incubation at 37 °C. The resultant values are reported as the mean log10 CFU ± standard deviation (SD) per 1.5 × 105 cells.
4.4. Scanning Electron Microscopy and Thin-Layer Chromatography (TLC)
Scanning electron microscopy was performed as described with slight modifications [
27]. Briefly, 5 mL culture aliquots were concentrated by centrifugation at 1644×
g for 5 min, and the supernatant was discarded. Then, 1 mL of 4% paraformaldehyde was added to the concentrated cells, and the samples were incubated at room temperature for 2 h. After centrifugation, the concentrated cells were washed twice with 0.05 M sodium cacodylate buffer (pH 7.4). The samples were dehydrated with a sequential ethanol series for 10 min each (50%, 70%, 95%, and 100%). These samples were mixed well with hexamethyldisilazane, a volatile solution, and then placed on the grid before drying. The samples were coated using gold sputter and imaged with a SEM3500M scanning electron microscope.
Bacterial total lipids and GPLs were purified, solubilized, and confirmed as described previously [
13]. Total lipids were extracted from
Mmuc with a chloroform/methanol mixture (2:1,
v/
v) by ultrasonication for 20 min and phase-separated by centrifugation. GPLs were purified from total lipid extracts by acetone precipitation. The purified lipids were separated by TLC (Millipore, Billerica, MA, USA) in chloroform/methanol (9:1,
v/
v) and detected by spraying with 10% H
2SO
4 and heating at 200 °C for 10 min.
4.5. Flow Cytometric Analysis
To determine the type of cell death followed by Mmuc infection, BMDMs were infected with the bacteria at an MOI of 10. After 24 h, the infected cells were centrifuged at 440× g for 3 min, and the supernatant was discarded. Next, the cells were detached by treatment with trypsin for 10 min, and then, 300 µL of complete DMEM was added to each well. Then, we performed experiments according to the manufacturer’s protocol (BD Biosciences, San Jose, CA, USA). Briefly, cells were harvested, washed with ice-cold PBS to remove extracellular bacteria, and then resuspended in 300 µL of 1× binding buffer. Next, 1 × 106 cells were stained with 3 µL of Annexin V-FITC and 3 µL of propidium iodide (PI). Then, these samples were incubated at room temperature for 15 min. After 15 min, 400 µL of binding buffer was added to each tube, and the samples were analyzed using a Cytoflex flow cytometer (Beckman Coulter, Villepinte, France). The cells were assigned to one of four states: alive, annexin V-negative, and PI-negative; early apoptotic, annexin V-positive, and PI-negative; late apoptotic, annexin V-positive, and PI-positive; or necrotic, annexin V-negative, and PI-positive.
4.6. Measurement of Cytokines by Enzyme-Linked Immunosorbent Assay (ELISA)
Culture media from the bacteria-infected BMDMs (1.5 × 105 cells/48-well plate) and noninfected BMDMs were collected at 24 h post-infection. Supernatants were collected after centrifugation at 848× g for 5 min. The levels of tumor necrosis factor (TNF)-α, interleukin (IL)-6, IL-12p40, and IL-10 were analyzed using commercial ELISA kits and OptEIA ELISA kits (BD Biosciences, San Diego, CA, USA) according to the manufacturers’ instructions.
4.7. Western Blot Analysis
After infection with bacteria, adherent cells were washed twice with PBS and then lysed in ice-cold PRO-PREP™ Protein Extraction Solution (Intron, Gyeonggi-do, Korea). After incubation for 10 min, the samples were gently scraped from dishes and then centrifuged at 15,928× g for 5 min. The supernatant was collected and stored at −80 °C. The protein concentrations of the lysates were determined using the Pierce BCA Protein Assay kit (Thermo Scientific, Waltham, MA, USA). The protein was mixed with 5× SDS–PAGE loading buffer (LPS solution) and denatured by heating to 100 °C for 15 min. Ten to 30 μg of protein was subjected to electrophoresis on 10~15% polyacrylamide gels containing SDS under reducing conditions. Separated proteins were electroblotted onto 0.22 µm polyvinylidene difluoride (PVDF) membranes (BD), and blots were blocked with 5% skim milk (wt/vol) for 1 h and then washed three times with Tris-buffered saline containing 0.1% Tween 20 (TBS/T). Then, the membranes were incubated overnight at 4 °C with the following antibodies: rabbit anti-p-p38 MAPK (#4631), rabbit anti-p-ERK1/2 (#9101S), rabbit anti-p-JNK (#9251, 1:1000; Cell signaling technology, Boston, MA, USA), and mouse anti-β-actin (#A1978, 1:5000; Sigma-Aldrich, Burlington, MA, USA). Antibody binding was detected using the appropriate secondary antibody coupled with HRP, as described by the manufacturer. Enhanced chemiluminescence was used to detect relevant proteins using the EZ-Western LumiFemto Kit (DG-WF100).
4.8. Seahorse Extracellular Flux Analysis
The oxygen consumption rate (OCR) and extracellular acidification rate (ECAR) were measured using a Seahorse XFp Metabolic Flux Analyzer (Seahorse Bioscience, North Billerica, MA, USA). BMDMs were seeded in Seahorse XFp cell culture plates at a density of 80,000 cells per well and cultured at 37 °C in a 5% CO
2 incubator overnight. The next day, the BMDMs were infected with
Mmuc for 24 h at an MOI of 10. On the day of analysis, glycolysis stress and mito stress tests were performed according to the manufacturers’ protocols. For the mito stress test, the medium was replaced with XF DMEM supplemented with 10 mM glucose, 1 mM pyruvate, and 2 mM glutamine (pH 7.4) followed by incubation at 37 °C in a non-CO
2 incubator for 45 min. Oligomycin, carbonyl cyanide phospho-(p)-trifluoromethoxy phenylhydrazone (FCCP), and rotenone/antimycin A were subsequently injected into the medium at final concentrations of 1 μM, 2 μM, and 0.5 μM, respectively. For the glycolysis stress test, the medium was replaced with XF DMEM supplemented with 2 mM glutamine (pH 7.4) followed by incubation at 37 °C in a non-CO
2 incubator for 45 min. Glucose, oligomycin, and 2-deoxyglucose (2-DG) were subsequently injected into the medium at final concentrations of 10 mM, 1 μM, and 50 μM, respectively. The OCR and ECAR were automatically recorded and calculated by Seahorse XFp software. At the end of the analysis, the medium was removed, and BMDMs were washed with PBS and lysed in RIPA buffer. The protein content in BMDM lysates was measured by the Bradford assay and used for normalization. Data were derived from two independent experiments. Determinants of respiratory and acidification parameters were calculated using the following equation (
Supplemental Table S1).