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Phagocytosis by macrophages decreases the radiance of bioluminescent Staphylococcus aureus

BMC Microbiology volume 25, Article number: 12 (2025) Cite this article

Abstract

Background

In vivo evaluations of the antimicrobial efficacy of biomaterials often use bioluminescent imaging modalities based on bioluminescent bacteria to allow follow-up in single animals. Bioluminescence production by bacteria is dependent on their metabolic activity. It is well known that several factors can influence the metabolism of bacteria, such as the use of antimicrobials and changes in bacterial growth phase. However, little is known about the influence of intracellular residence of bacteria on bioluminescence. For example, Staphylococcus aureus can survive in the peri-implant tissue and is known to survive intracellularly in macrophages.

Results

In this study, we evaluated the bioluminescent radiance of S. aureus upon phagocytosis by macrophages. We showed that S. aureus reduced its bioluminescence upon phagocytosis by macrophages compared to S. aureus in a single culture. Simultaneously, bacterial numbers as measured by colony-forming units remained constant over time. S. aureus was released extracellularly as a result of macrophage cell death. Following this release, the bacteria increased their bioluminescence again. Replenishment of fresh macrophages showed an immediate increase in bioluminescence. Moreover, the addition of fresh macrophages showed a diminished decrease in bioluminescence at 24 h of coculture, but this effect did not last.

Conclusion

Together, this study demonstrates that phagocytosis by macrophages decreases bioluminescence of S. aureus, which is an important factor to consider when using bioluminescent imaging to study the infection process in an in vivo model.

Peer Review reports

Introduction

The use of biomaterials for restoration of body functions due to oncological surgery, trauma or wear is increasing, owing to the ageing population, advances in healthcare and the increasing availability of medical treatment to the population [1]. A major complication of biomaterial implants is the occurrence of biomaterial-associated infections (BAI). These infections are notoriously hard to treat because of the physiological attributes and structure of the biofilm that reduces the susceptibility of antimicrobials [2], but also due to the continuous development of antimicrobial resistance which proposes an emerging global health threat. One of the major pathogens in BAI is Staphylococcus aureus, a major global health burden that is the cause of many different infections [3]. To prevent BAI, biomaterials are being developed that prevent bacterial adhesion, release antimicrobials, or kill bacteria upon contact with the biomaterial [4].

Validation and ultimately the translation of new materials to the clinic need extensive evaluations often including animal testing with bioluminescence imaging (BLI) as a tool to monitor infection. BLI of bacteria, genetically modified to produce bioluminescence, displaces the need for euthanasia of animals at multiple timepoints to collect material for bacterial counts. BLI can be seen as an indicator of bacterial metabolic activity, as light is produced through enzymatic processes [5]. However, current studies often directly correlate BLI with the number of surviving bacteria, therewith neglecting downregulation or upregulation of the bacterial metabolism, both in subsequent growth phases or in an attempt to escape host defense mechanisms or survive antibiotic exposure [6,7,8]. This is also illustrated by discrepancies in in vivo studies in the comparison between bioluminescence measurements and endpoint colony-forming unit (CFU) counts [9, 10]. Although this discrepancy can partly be explained by the insufficient sensitivity of BLI, the microbial environment also has a strong influence on bacterial metabolism, which hampers the relationship between CFU counts and bioluminescence.

Apart from down- or upregulation of the metabolism as a result of antimicrobials or changes in growth phase, part of the discrepancy between CFU counts and BLI may be related to bacterial survival in peri-implant tissue [11]. For instance, upon infection of tissue by S. aureus, a host immune response is initiated, for which S. aureus has several evasion mechanisms. The first line of the host defense is phagocytosis by professional phagocytes, mainly macrophages. Aside from evasion of phagocytosis, S. aureus can withstand killing by these phagocytes adopting evasion mechanisms and reside intracellularly [12]. For other members of the staphylococcus genus, survival in peri-implant tissue has been demonstrated, exhibiting colocalization with macrophages, indicating intracellular survival [11, 13].

To verify our hypothesis whether bacterial intracellular survival could result in a decreased bioluminescence, we studied the radiance of bioluminescent S. aureus Xen29 bacteria upon macrophage phagocytosis and related this to CFU counts. To investigate this, we cultured surface-attached S. aureus in the presence of murine macrophages in order to mimic murine infection models and measured both CFU counts and bioluminescence over time. In order to relate this to phagocytosis we determined intra- and extracellular bacterial numbers and related this to intracellular bacterial numbers and macrophage viability.

Results

Optimization of culture medium

In order to optimize the coculture medium we first studied the growth of both organisms in different ratios of bacterial and cell medium, TSB and DMEM + FBS, respectively [14]. Growth of J774 murine macrophages was observed in media containing less than 30% TSB, thus 70% DMEM + FBS or more (Fig. 1). No changes in morphology were observed in conditions containing up to 30% TSB compared to 0% TSB (Fig. S1A and B). J774 cells grown in 100% TSB showed a change in cell morphology and subsequent cell death. Since the CFU number of S. aureus Xen29 was hardly (less than 1 decade) affected by the absence of TSB, 100% DMEM + FBS medium were applied in the co-culture assay. This medium condition did not influence the bioluminescence of bacteria compared to DMEM + FBS with low percentages of TSB (Fig. S1C). (Title in Suppl methods in MOESM1 refers to older title. Should be: Belonging to "Phagocytosis by macrophages decreases the radiance of bioluminescent Staphylococcus aureus")

Fig. 1
figure 1

Growth of S. aureus and macrophages as a function of percentage of TSB in DMEM-HG + FBS. The dotted horizontal line indicates the seeding concentration of J774 macrophages. Concentrations measured in duplicate after 48 h of growth

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Macrophages alter the metabolic behavior of S. Aureus over time

To reach the sensitivity limit for bioluminescence detection, 108 bacteria were seeded. After incubation for 2 h and removal of non-adherent bacteria, the number of adhering CFUs per well was 2.4 × 107. J774 macrophages (106 per well) were added (t = 0 h), reaching a bacteria: macrophage ratio of around 24:1, and bioluminescence and CFUs were measured over the course of 48 h of coculturing (Fig. 2). At 8 h of S. aureus in coculture with macrophages an increase in CFU (Fig. 2A) and bioluminescence (Fig. 2B) was shown. Remarkably, bacteria in single culture showed the same initial increase in CFU at 8 h (Fig. 2A) but showed a slight decrease in bioluminescence. This bioluminescence continued to drop 100-fold over time, while bacteria in coculture with macrophages showed a significantly larger decrease of almost 10,000-fold after 24 h of coculture. This decrease was not persistent, as bioluminescence increased at 48 h of coculture (Fig. 2B). In order to verify whether this effect was caused by nutrient depletion alone, a control experiment was executed in which we administered at the timepoint of 12 h, spent medium that was depleted from nutrients by macrophages. This control did show a small decrease (less than a factor of 10) in bioluminescence (Fig. S2A) but far less than showed in in Fig. 2B with bacteria exposed to macrophages. However, adding fresh medium at time 12 h and 24 h did not stop the considerable drop of bioluminescence (Fig. S2B and C). Simultaneously, CFU of bacteria in both coculture and single culture remained constant after 8 h, following the initial increase (Fig. 2A). At 12 and 24 h, bioluminescence measurements were performed before and after disturbing the culture by opening and closing of the well plate, showing an immediate increase in bioluminescence after disturbing the culture (Fig. 2B).

To mimic the physiological condition where macrophages are constantly replenished, fresh macrophages were added to the coculture, either at 12 (Fig. 2C) or 24 h (Fig. 2D) with measurements performed both before and directly after replenishment. The addition of macrophages at 12 h did not show an immediate effect on the bioluminescence (Fig. 2C). However, at 24 h the decrease of bioluminescence was less prominent than the condition without adding macrophages. An influx of fresh macrophages at 24 h, however, showed a prominent increase in bioluminescence (Fig. 2D), higher than caused solely by disturbing bacteria through opening of the well plate as shown by the coculture condition. Although for both conditions, the addition of macrophages at 12–24 h, bacterial metabolism was altered at 24 h, bioluminescence at 48 h was similar to when macrophages were not replenished, indicating that this effect is not abiding.

Fig. 2
figure 2

Monitoring the effect of macrophages on S. aureus Xen29 growth and bioluminescence. (A) CFU measurements of S. aureus Xen29 in the absence or presence of macrophages. Bacterial numbers were obtained after lysing macrophages (co-culture) and disrupting bacterial clusters. (B) Bioluminescent radiance of S. aureus Xen29 in the absence or presence of macrophages. Bioluminescence of media and non-luminescent macrophages were just at or below the read-out noise of the system. (C, D) Bioluminescent radiance of S. aureus Xen29 in the presence of macrophages, with replenishment of macrophages at 12 h (C) or 24 h(D), as indicated by the colored arrows. At timepoints 12 and 24 h measurements are shown before and after the addition of macrophages (and the associated opening of the well plate). Data are depicted as the mean resided extracellular of 3 independent triplicate experiments with error bars representing SD. Statistical analysis using repeated measures two-way ANOVA (A) or mixed-effect analysis (B, C, D) with Tukey post-hoc test. * p < 0.05 for statistical significance between 2 consecutive timepoints within a condition, # p < 0.05 for statistical significance between conditions at a single timepoint (timepoint 8 h and 12 h before (B) or timepoint 24 h after (D)

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Bacterial bioluminescence is related to intracellular residence in macrophages

To visualize bacterial uptake by macrophages we applied DAPI staining of nucleic acids of both macrophages and bacteria, in combination with Phalloidin (to stain macrophage actin) and immunolabeling of S. aureus bacteria (Fig. 3). Based on DAPI staining we only observed extracellular bacterial presence after 12 h (See Fig. S3), indicating cellular uptake of bacteria during the first 8 h. To quantify the rate of bacterial phagocytosis by macrophages, we calculated the percentage of intracellular S. aureus bacteria [15] (Fig. 3A, C). At 8 h, the percentage of intracellular bacteria was nearly 100%, with some macrophages undergoing cell division (Fig. S4A, B). Already at 12 h, this percentage decreased to around 60% (Fig. 3A, C), while some macrophages appeared fused to foreign body giant cells (Fig. S4C). At 24 h, all bacteria resided extracellularly. Remarkably, at 24 h no actin was visible anymore, indicating that macrophages lost their apparent cell boundaries (Fig. 3C), indicative of macrophage cell death. This was corroborated by an assessment of the viability of macrophages (Fig. 3B, S5), which showed a similar trend as observed in the percentage of intracellular bacteria. No viable macrophages were observed at 24 and 48 h, indicating that upon macrophage cell death, intracellular bacteria are released into the extracellular environment. At 24 h, some bacteria remained in the vicinity of the dead cell, while at 48 h they left the dead cells (Fig. S2D). As the bacterial CFUs remained constant over time, this demonstrates that bacteria survive intracellularly within macrophages until macrophage cell death. Macrophage cell death occurred at 12 h already, with the complete death of macrophages at 24 h (Fig. 3B). Meanwhile, bacterial bioluminescence only increased at 48 h of coculture (Fig. 2B). Therefore, we conclude that macrophage cell death precedes induction of bacterial bioluminescence.

Fig. 3
figure 3

S. aureus phagocytosis by macrophages. (A) Intracellular bacterial counts of S. aureus residing intracellularly in J774 macrophages. Intracellular counts were determined by staining of bacteria and the actin cytoskeleton of macrophages. (B) Percentage of viable macrophages as measured by LIVE/DEAD staining in the absence and presence (coculture) of bacteria. Data (A and B) are depicted as the mean of 3 independent triplicate experiments with error bars representing SD. Statistical analysis using repeated measures one-way (A) or two-way (B) ANOVA with Tukey post-hoc test, * p < 0.05. (C) Confocal laser scanning microscopy of adherent bacteria cocultured with macrophages at different timepoints. DNA is stained blue with DAPI, S. aureus is immunolabeled against protein A in green and the actin cytoskeleton of macrophages to visualize cell contour is stained red using phalloidin. Images are a maximum projection of a z-stack. The scale bar denotes 50 μm

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Discussion

Since bioluminescent imaging enables the assessment of infection at subsequent timepoints in the same animal, the use of BLI is expanding with the current societal debate on reducing animal testing. Still, the use of murine models to study in vivo infection proliferation has been discussed because of the often applied extremely high bacterial inoculum needed that does not reflect the number of pathogens that in clinical cases will lead to BAI [16]. This high number of bacteria (107 – 109 CFU’s per inoculum) is needed to be able to infer a significant chronic BAI in mice and rats and also to surpass the noise level of bioluminescence imaging instrumentation [17]. Based on the same considerations the MOI in our assay was taken relatively high as well (1:24). Nevertheless, many studies overlook the implications of bioluminescence in living organisms like bacteria as bioluminescence is not a stationary process which can be directly translated to numbers of organisms. Rather, bioluminescence is heavily influenced by the metabolic activity of organisms [5]. For instance, direct evidence of the tight link between the metabolism and bioluminescence is the observation of immediate increases of bioluminescence after lifting the lid for adding fresh macrophages at t = 12 and 24 h. The S. aureus Xen29 has been genetically modified to obtain the Photorhabdus luminescence Lux ABCD operon, which constitutes bioluminescence in the presence of oxygen, ATP and NADPH. In particular, oxygen and an aldehyde are necessary to oxidize reduced flavin, FMNH2, to obtain flavin, FMN, that acts as a substrate for producing light in the presence of luciferase (LuxAB). LuxCDE is necessary for the production of aldehydes [18]. So, by lifting the lid in the bioluminescence assay, oxygen supply is increased which boosts the bioluminescence quite immediately. Also, the slight decrease in the bioluminescence from a steady number of culturable bacteria in the present study, points to an earlier and often found effect in both Gram-negative and Gram-positive strains. The main origin of this decline of luciferase activity, also in S. aureus, was shown to be connected to a depletion of FMNH2 which directly impacts the availability of FMN as a substrate for bioluminescence [19].

In the present study, we show that changes in the bioluminescence of bacteria also take place as a result of phagocytosis by macrophages, which induces a 10,000-fold decrease in bioluminescence while bacteria remained culturable. This reduction of metabolism was not lasting and luminescence increased again after longer coculture times, which coincided with macrophage cell death and release of intracellular bacteria, confirming our hypothesis that macrophage phagocytosis silences bacterial luminescence, rather than bacterial nutrient deprivation in the coculture, as was suggested earlier [20].

S. aureus is known to survive intracellularly and although most studies have been performed using non-immune cells [12], earlier studies have shown S. aureus to reside and even replicate intracellularly in macrophages [21,22,23]. Here, we also observed intracellular survival and replication in the first hours after phagocytosis, coinciding with increasing bioluminescence. At later time points, we did not observe growth, but apparently, macrophages were not lethal for S. aureus bacteria in this assay. It is well known that S. aureus bacteria are notorious survivors in a harsh environments [24]. In particular, S. aureus is able to counteract NO production by phagocytes and can evade pattern recognition by macrophages in biofilms. Also, biomaterial-associated infection animal models show that even after 28 days high amounts of culturable bioluminescent S. aureus Xen29 bacteria were found in the tissue of mice in contact with a non-degradable biomaterial, whereas bioluminescence remained low [10]. In our assay, after 24 h, bioluminescence dropped significantly, while bacterial numbers remained similar. Previously, studies of bacteria residing within macrophages have shown a reduced ATP production with a factor of 1.5 [21]. This limited reduction in ATP cannot explain the 10,000-fold reduction in bioluminescence, which is corroborated by simulations that show that ATP only has a small impact on bioluminescence [18]. Rather than on ATP, changes in bioluminescence depend on NADPH production as the Photorhabdus luminescens luxACBDE operon of S. aureus Xen29 requires NADPH for the production of aldehydes as the substrate for light production [25]. The production of reactive oxygen species (ROS) by phagocytic cells is an important killing mechanism in the fight against intracellular pathogens, which can be neutralized by bacterial enzymes such as glutathione reductase, thioredoxin reductase and antioxidant systems that require NADPH for their reductive activity [26]. ROS production is induced by a low pH that bacteria encounter within the phagosome, upon which bacteria increase their enzyme activity [27]. It has already been shown that in multiple bacterial species a low pH decreases the bioluminescence production by bacteria [28]. Moreover, a reduction in NADPH has been shown to impair S. aureus intracellular survival in macrophages [29], as this allows ROS to induce damage to the bacteria. As a result of this high demand for NADPH, we suggest that by reducing bioluminescence production bacteria increase their pool of NADPH that is available to combat ROS killing, as is consistent with the delayed increase in bioluminescence upon macrophage death. The decline in NADPH production available for bioluminescence is suggested to be a follow up of an initial increase of the NADPH redox pool, and initial increase in bioluminescence, needed for bacterial respiration. This was suggested by Daghighi et al. to explain an often found increase in bioluminescence upon exposure to sub-inhibitory concentrations of antibiotics [7, 17, 30]. Peyrussen et al. also showed that the production of ROS by macrophages leads to deeper dormancy of S. aureus [31], resulting in prolonged lag times which also explains the delayed increase in bioluminescence following macrophage cell death [31]. Simultaneously, we found that bacteria in single culture also showed a slight drop in bioluminescence, but less prominent and we did not measure an initial increase as we did in the presence of macrophages. Within the first hours, bacteria in both coculture and a single culture replicated upon which they reach the stationary phase, which causes an increase in bioluminescence. However, bacteria in a single culture follow the replication rate and reach stationary phase and subsequent peak in bioluminescence in 4 to 6 h of subculture, already exhibiting a small decline in bioluminescence at 8 h. Concurrently, bacteria phagocytosed by macrophages have delayed exponential growth and therefore the peak of bioluminescence can be detected around 8 h of coculture [32].

We show that bacteria also regain their bioluminescence after macrophage cell death. Different studies have reported different lengths of intracellular survival of S. aureus within macrophages, ranging from 24 h [21, 23] to 5 days [22]. Here, we report a bacterial intracellular state between 12 and 24 h, although at 12 h already a population of macrophages died and released bacteria. These differences in intracellular survival might depend on bacterial strains that differ in their expression of certain virulence factors, for instance, phenol-soluble modulins that are necessary for phagolysosomal escape, avoiding killing and enabling cytoplasmic replication of bacteria [33]. It has been shown previously [23] that macrophage cell death precedes the emergence of S. aureus bacteria. Cells underwent apoptosis, characterized by activation of caspase-3 and membrane blebbing. Here, we have obtained evidence of macrophage cell death in two ways. First, by staining of cells with a nucleic acid stain that only reaches the target upon cell membrane disruption but which not always correlate to cell death. Second, we showed a loss of actin after 24 h, indicating cells have lost their viability as actin is maintained through a dynamic process [34]. After 12 h, some macrophages showed reduced membrane integrity based upon the nucleic acid stain, but did not completely lose their actin cytoskeleton, indicating that bacteria require only small membrane disruptions to escape phagocytosis, as illustrated by the increased numbers of extracellular bacteria after 12 h of coculture. Our results do not show clear signs of apoptosis, as indicated by the lack of membrane blebbing. Rather, our results point in the direction of types of cell death that are characterized by loss of membrane integrity, such as necrosis. As the dense actin cytoskeleton offers support and movement to the cell membrane, the observed loss of this cytoskeleton can be indicative of disrupted membrane integrity, which is something that can be induced by the secretion of bacterial toxins [35, 36].

We expect that our in vitro results can be translated to in vivo bacterial infections and that the engulfment of bacteria by macrophages can lead to a drastic reduction in bioluminescence. Still, relating in vitro to in vivo results is notoriously difficult. In an in vivo situation, macrophages are continuously replenished from the bloodstream and new macrophages can phagocytose released bacteria. Therefore, we attempted to mimic this physiological condition by adding new macrophages to the coculture after different lengths of coculture. This resulted in an immediate increase in bioluminescence, potentially reflecting a transient NADPH overexpression as we observed with the first exposure of bacteria to macrophages, but later coculture times did not show any differences compared to when macrophages were not replenished.

Conclusion

In summary, we evaluated the relationship between bioluminescence measurements and intracellular survival of bacteria within macrophages. As a main conclusion, we show that while CFUs of bacteria do not fluctuate over time, bioluminescence significantly reduces as an apparent result of the loss of bacterial metabolic activity upon phagocytosis or, as we suggest by an active defense mechanism to reduce the NADPH reductase pool. They only slowly return to their regular bioluminescence upon macrophage cell death and subsequent bacterial release. These results show that caution is warranted when using bioluminescence as a tool for in vivo studies on bacterial infections for replacement of CFU counts, not only when studying biofilms but also for infection of surrounding soft tissue.

Methods

Culturing and harvesting of bacteria

Bacterial cultures of bioluminescent S. aureus Xen29 (PerkinElmer, Waltham, USA) were grown on trypticase soy agar (TSA, Oxoid, Basingstoke, UK / BD, Franklin lakes, USA) supplemented with 200 µg/mL kanamycin (KAN, Sigma-Aldrich, St. Louis, USA) at 37 °C for 24 h under aerobic conditions. A preculture was prepared by inoculation of a single bioluminescent colony (confirmed with IVIS) in 10 mL of trypticase soy broth (TSB, Oxoid, Basingstoke, UK) containing 200 µg/mL KAN and incubated at 37 °C, 150 rpm for 24 h under aerobic conditions. A main culture was prepared by inoculation of 200 mL of TSB with 10 mL of preculture and cultured at 37 °C, 150 rpm for 16 h. Bacterial cultures were harvested from the main culture by centrifugation at 6500 g for 5 min at 10 °C. The pellet was resuspended in sterile phosphate buffered saline (PBS; 5 mM K2HPO4, 5 mM KH2PO4, 150 mM NaCl, pH 7.0) and washed twice with the abovementioned centrifugation. Bacteria were counted in a Bürker-Türk counting chamber and diluted in sterile PBS to a concentration of 108 bacteria/mL.

Culturing and harvesting of macrophages

J774A.1 (ATCC TIB-67) murine macrophages were cultured in Dulbecco’s Modified Eagle Medium containing 4.5 g/L glucose, pyruvate and glutaMAX (DMEM, Gibco Thermofisher, Waltham, USA) supplemented with 10% Fetal Bovine Serum (FBS, Gibco Thermofisher, Waltham, USA), referred to as DMEM + FBS. Cells were cultured in polystyrene flasks (Greiner Bio-One, Kremsmünster, Austria) at 37 °C and 5% CO2. Cells were passaged upon reaching a confluency of 70–80% by means of scraping. Prior to experiments, cells were harvested by centrifugation at 175 g for 5 min. Cells were counted with a Bürker-Türk hemocytometer using Trypan Blue (Gibco Thermofisher, Waltham, USA) and diluted to 106 cells/mL.

Development of modified culture medium

To enable the growth of both S. aureus and J774A.1 cells simultaneously, an optimal medium had to be formulated. Bacterial medium (TSB) and cell medium (DMEM + FBS) were combined in different ratios and the growth rates of both bacteria and cells were determined. For S. aureus growth, precultures and main cultures were made in different percentages of TSB in DMEM + FBS media (0, 2, 4, 6, 8, 10, 20, 30 and 100%) from a single colony on a TSA plate. Precultures and main cultures were cultured as described above. The main culture was then 10-fold serially diluted in PBS and three drops of 10 µL were plated on TSA plated, incubated for 16 h at 37 °C under aerobic conditions after which colonies were counted.

For macrophages, J774A.1 cells were collected from a culture as described above. After centrifugation, cells were resuspended in cell medium or TSB, counted and diluted to 6 × 105 cells/mL. Then, 1 mL of cell suspension was added to a T25 flask containing 3 mL of combined culture media ((DMEM + FBS) media percentages the same final percentages as for bacteria. Cells were incubated for 48 h and morphology, cell spreading and adhesion were checked at 24 and 48 h. After 48 h, cells were collected from the flask using scraping and viable cells were counted using Trypan Blue staining.

Coculture assay

First, 1 mL containing 108 bacteria was added to each well of a 12-well plate in PBS containing 2% TSB to maintain bioluminescence. After 2 h of incubation at 37 °C, 5% CO2 the bacterial suspension was removed from the wells and washed once with PBS to remove non-adherent bacteria. Subsequently, 106 macrophages were added in 1 mL of DMEM + FBS to the adherent bacteria and the coculture was incubated for 48 h at 37 °C, 5% CO2. In order to replenish macrophages, 106 fresh macrophages in 1 mL of media were added to the well at 12–24 h. Bioluminescence and CFU measurements were performed at 0, 8, 12, 24 and 48 h. As a control, 1mL of media without macrophages was added. When macrophages were administered to the culture, bioluminescence measurements were performed before and after the addition of macrophages and the associated opening of the well plate, within 5 min after the addition of macrophages. Controls included macrophages, media and bacteria in DMEM + FBS. For the control of bacteria with spent macrophage media, macrophages were cultured in a 12-well plate. After 12 h, media was collected from macrophages and filtered through a 0,22 μm filter. Media was removed from bacteria and the filtered spent media was added to the bacteria.

Bioluminescence imaging (BLI)

Bioluminescence imaging was performed using the IVIS Lumina (PerkinElmer, Waltham, USA). First, grey-scale photographs were taken and subsequently a bioluminescence image was obtained using a 10 × 10 cm field of view, 1/f aperture, binning of 4 and exposure time of 3 min. Read-out noise was at a level of around photons/sec/cm2/sr. Manual regions of interest were drawn on the edges of the well and a background region of interest was drawn on an empty well using the Living Image 4.8.0 software (PerkinElmer, Waltham, USA).

Colony-forming units

Bacteria and macrophages were scraped from the well surface using the back of a sterile pipet tip. Although a limited amount of biomass remained visible after microscopical inspection, the amount was small and comparable under all conditions. The culture suspension was collected in an Eppendorf tube and sonicated using a sonication bath for 5 min to lyse macrophages and disrupt bacterial clusters. Samples were then serially diluted using a 10-fold dilution in PBS and 10 µL drops of each dilution were plated in triplicate on TSA plates. Plates were incubated for 16 h at 37 °C under aerobic conditions, after which colonies were counted.

Macrophage viability

Bacteria and macrophages were cocultured as described in the coculture assay. At timepoints 8, 12, 24 and 48 h the medium was gently removed from the wells. The coculture was washed twice with PBS to remove any traces of culture medium. Cells in one well were killed with 70% EtOH and used as a control for the staining of dead cells. Macrophages without bacteria were used as a control for cell viability without bacteria. After washing, 500 µL of a PBS solution (137 mM NaCl, 1.47 mM KH2PO4, 8.10 mM Na2HPO4·2H2O, 2.68 mM KCl, pH 7.4) was added to each well, containing 0.5 µM Calcein AM and 4 µM Ethidium Homodimer-1 (EthD-1) (LIVE/DEAD Viability/Cytotoxicity kit, Thermofisher, Waltham, USA) to determine the viability of the macrophages and incubated for 20 min at 37 °C, 5% CO2 in the dark. EthD-1 is a nucleic acid stain that can only enter cells with a damaged membrane. Fluorescence imaging was performed using a Leica DM4000B fluorescence microscope with a 40x water immersion objective. Triplicate wells were imaged for each condition, averaging 3 images per well. The percentage of culturable macrophages was taken as a percentage of the total of live and dead macrophages combined. Analysis and statistics were performed on triplicate experiments.

Intracellular bacteria counting

Bacteria and macrophages were cocultured as described above. At timepoints 8, 12, 24 and 48 h medium was gently removed, and cells were washed twice with PBS. Cells were fixed with 4% formaldehyde (VWR, Radnor, USA) for 15 min at room temperature, washed, permeabilized with 0.2% Triton X-100 (Sigma-Aldrich, St. Louis, USA) for 15 min, washed and incubated for 10 min with 100 mM glycine (Sigma-Aldrich, St. Louis, USA) followed by 3% bovine serum albumin (BSA; Sigma-Aldrich, St. Louis, USA) in PBS for 30 min at room temperature. Then, samples were stained for S. aureus with 100 µL of 8 µg/mL of anti-S. aureus-FTIC antibody (ab68950 Abcam, Cambridge, UK) diluted in a solution of 5% BSA in PBS + 0.1% (v/v) Tween-20 (PBS-T, Boom, Meppel, The Netherlands), and incubated for 1 h at room temperature in the dark. After washing 3 times for 5 min in PBS-T, 100 µL of 1 µg/mL TRITC-phalloidin (P5282, Sigma-Aldrich, St. Louis, USA) was added and incubated for 30 min at room temperature in the dark to stain actin to visualize cell boundaries of the macrophages. Samples were washed with PBS 3 times for 5 min after which counterstaining with 2 µg/mL DAPI (D9542, Sigma-Aldrich, St. Louis, UK) was performed, staining nucleic acids of both macrophages and bacteria. After washing, samples were submerged with PBS and confocal images were taken using Leica Stellaris 5 on DM6 CS with a HCX APO L 40x/0.80 UVI free water immersion lens, working distance 3.3 mm at 1024 × 1024 resolution using the Leica LasX software (Leica, Wetzlar, Germany). Three z-stack images were taken from each well, with each condition measured in three wells. Image analyses and calculations were performed on triplicate experiments using Imaris 9.7 software (Oxford Instruments, Abingdon, UK). The S. aureus staining was used to generate 1 μm spots and the actin staining was used to generate a surface representing the macrophage membranes. The intra- and extracellular bacteria were calculated by counting the number of spots inside and outside the actin surface, respectively. To obtain the percentage of intracellular bacteria, the number of intracellular bacteria was divided by the total number of bacteria.

Statistical analysis

Statistical analyses were conducted using GraphPad Prism version 8.0 (Dotmatics, Boston, USA). Statistical tests used are specified in the captions of the figures. Significance was tested compared to the control groups and defined as significant when p ≤ 0.05 or non-significant when p > 0.05.

Data availability

The datasets used and/or analysed during the current study are available from the corresponding author on request.

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Funding

This publication is part of the DARTBAC project (with project number NWA. 1292.19.354 of the research programme NWA-ORC which is (partly) financed by the Dutch Research Council (NWO).

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Authors and Affiliations

  1. Department of Orthopedics, University Medical Center Groningen, Hanzeplein 1, Groningen, The Netherlands

    Elles C. Boonstra & Paul C. Jutte

  2. Department of Biomaterials and Biomedical Technology, University Medical Center Groningen, Antonius Deusinglaan 1, Groningen, The Netherlands

    Liliana Agresti, Henny C. van der Mei & Jelmer Sjollema

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  1. Elles C. Boonstra
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Contributions

ECB contributed to conceptualization of the study, methodology, data collection, formal analysis, visualization, writing-original draft, writing-review and editing. LA contributed to conceptualization of the study, methodology, data collection, writing-review and editing. PCJ contributed to conceptualization of the study, supervision, writing-review and editing. HCvdM contributed to conceptualization of the study, supervision, writing-review and editing. JS contributed to conceptualization of the study, supervision, visualization, writing-review and editing. All authors read, reviewed and approved the final manuscript.

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Correspondence to Jelmer Sjollema.

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Boonstra, E.C., Agresti, L., van der Mei, H.C. et al. Phagocytosis by macrophages decreases the radiance of bioluminescent Staphylococcus aureus. BMC Microbiol 25, 12 (2025). https://doi.org/10.1186/s12866-024-03674-x

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Keywords

BMC Microbiology

ISSN: 1471-2180

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