Cellular Microbiology: Pathogen-Host Cell Molecular Interactions

Opsonophagocytosis of Chlamydia pneumoniae by Human Monocytes and Neutrophils

Mads Lausen, Mathilde Selmar Pedersen, Nareen Sherzad Kader Rahman, Liv Therese Holm-Nielsen, Faduma Yahya Mohamed Farah, Gunna Christiansen, Svend Birkelund
Craig R. Roy, Editor
DOI: 10.1128/IAI.00087-20

ABSTRACT

The human respiratory tract pathogen Chlamydia pneumoniae, which causes mild to severe infections, has been associated with the development of chronic inflammatory diseases. To understand the biology of C. pneumoniae infections, several studies have investigated the interaction between C. pneumoniae and professional phagocytes. However, these studies have been conducted under nonopsonizing conditions, making the role of opsonization in C. pneumoniae infections elusive. Thus, we analyzed complement and antibody opsonization of C. pneumoniae and evaluated how opsonization affects chlamydial infectivity and phagocytosis in human monocytes and neutrophils. We demonstrated that IgG antibodies and activation products of complement C3 and C4 are deposited on the surface of C. pneumoniae elementary bodies when incubated in human serum. Complement activation limits C. pneumoniae infectivity in vitro and has the potential to induce bacterial lysis by the formation of the membrane attack complex. Coculture of C. pneumoniae and freshly isolated human leukocytes showed that complement opsonization is superior to IgG opsonization for efficient opsonophagocytosis of C. pneumoniae in monocytes and neutrophils. Neutrophil-mediated phagocytosis of C. pneumoniae was crucially dependent on opsonization, while monocytes retained minor phagocytic potential under nonopsonizing conditions. Complement opsonization significantly enhanced the intracellular neutralization of C. pneumoniae in peripheral blood mononuclear cells and neutrophils and almost abrogated the infectious potential of C. pneumoniae. In conclusion, we demonstrated that complements limit C. pneumoniae infection in vitro by interfering with C. pneumoniae entry into permissive cells by direct complement-induced lysis and by tagging bacteria for efficient phagocytosis in both monocytes and neutrophils.

INTRODUCTION

Chlamydia pneumoniae is a human pathogen that frequently causes mild respiratory symptoms during infection that usually resolve spontaneously but can cause severe, long-lasting atypical pneumonia and has been associated with the development of several chronic disease conditions, including atherosclerosis and asthma (1, 2). C. pneumoniae is an obligate intracellular Gram-negative bacterium with a unique biphasic developmental cycle. The bacterium alternates between the extracellular infectious elementary body (EB) and the intracellular noninfectious reticular body (RB). During infection, C. pneumoniae EBs engage airway epithelium and induce their own uptake by secreting preformed effectors into the host cell cytoplasm by the type III secretion system (3). Intracellularly, the EBs transform into RBs and start replicating in a modified vacuole called an inclusion. After 48 to 72 h, RBs start to divide asynchronously, and EBs are formed and released from the host cell by cell lysis or by membrane extrusion (4).

Upon infection, C. pneumoniae EBs first encounter airway epithelial cells and alveolar macrophages, which respond to infection by secreting inflammatory mediators stimulating vascular endothelium activation and immune cell chemotaxis (5). Neutrophil granulocytes are the first immune cells to transmigrate from the bloodstream into the alveolar space and subsequently signal for mononuclear cell recruitment and infiltration (6, 7). Thus, neutrophils and monocytes comprise an early cellular defense against C. pneumoniae, and the interaction between extracellular chlamydial EBs and these phagocytes likely determines the course of the infection. Multiple studies have proposed that C. pneumoniae can evade intracellular killing mechanisms in both monocytes and neutrophils and thereby use these cells as cellular vectors for extrapulmonary dissemination that potentially causes chronic disease (8–10). However, these prosurvival mechanisms may be altered by the route of bacterial ingestion, since engagement with different phagocytic receptors activates different intracellular signaling and trafficking pathways.

It was previously shown that phagocytic uptake of the intracellular bacterium Mycobacterium tuberculosis through complement receptors or the mannose receptor protects the bacterium from intracellular killing by delaying phagosome maturation (11, 12). On the contrary, uptake of antibody-opsonized M. tuberculosis through Fcγ receptors causes rapid phagosome maturation and phagolysosomal fusion, leading to reduced infection outcome. Thus, opsonin-directed phagocytosis is a critical factor determining the intracellular fate of M. tuberculosis, and similar mechanisms could be involved in C. pneumoniae infections and pathogenesis, despite these pathogens being very different in nature.

The complement system consists of more than 30 soluble and membrane-bound proteins that work in a cascade-like manner to induce antimicrobial effector functions, including opsonization, chemotaxis, and cell lysis. The cascade process is initiated within minutes of engagement (13) and proceeds through three separate activation pathways: classical (recognition of antibody complexes), lectin (recognition of carbohydrate moieties), and alternative (spontaneous hydrolysis of C3) pathways (14). Following activation, complement C4 and C3 are proteolytically cleaved into small soluble C4a and C3a fragments (9 kDa each) and large C4b and C3b fragments (75 kDa and 110 kDa, respectively) that attach covalently to the activating surface and opsonize the microbe for phagocyte detection. C3b can be further cleaved into iC3b, C3dg, or C3d, which are all potent opsonins recognized by different complement receptors on phagocytic cells (14). The final stage of the complement cascade includes cleavage of complement C5 and generation of a multimeric membrane-spanning C5b-9 complex (membrane attack complex [MAC]) that induces pore formation and cell lysis (14).

During primary infection, C. pneumoniae engages the complement system in the alveolar space. Although alveolar complement concentration and activity are reduced compared to concentrations and activity seen in serum, the system is sufficiently functional to opsonize M. tuberculosis with complement C3 activation products (15, 16). Primary infection leads to C. pneumoniae-specific IgG production, but these antibodies offer limited protection against C. pneumoniae, since reinfections are frequently observed (17). Thus, during secondary infections, both antibodies and complement are present in the alveolar space, which have the potential to modulate the intracellular fate of C. pneumoniae in phagocytes and hence the outcome of the infection.

To further characterize the pathogenesis of C. pneumoniae during primary and secondary infections, we investigated how complement and antibody opsonization affect phagocytosis and intracellular survival of C. pneumoniae in human monocytes and neutrophils.

RESULTS

Complement and antibody opsonization of C. pneumoniae.To examine the role of opsonization and phagocytosis during C. pneumoniae infections, we first screened the serum samples from 10 healthy donors for IgG antibodies against C. pneumoniae and found that nine of ten donors were positive for IgG against C. pneumoniae. Serum from one seropositive individual and serum from the only identified seronegative individual were used in all further experiments. These sera are called immune serum and nonimmune serum, respectively, throughout this paper.

Different cleavage fragments of complement C3 and C4 are known to bind and opsonize bacteria for phagocytic recognition. C3 deposition on C. pneumoniae has only been investigated by flow cytometry using an antibody recognizing various C3 cleavage forms (18). Thus, the exact C3 opsonins that bind the C. pneumoniae surface remain unknown. In addition, complement C4 deposition has never been investigated on C. pneumoniae. We therefore analyzed the deposition of these complement components on purified C. pneumoniae EBs using immunoelectron microscopy (IEM) and Western blotting.

Figure 1A demonstrates that complement C3 is abundantly deposited on the chlamydial surface when incubated in nonimmune serum (NHS−). Very limited C3 deposition was observed in heat-inactivated serum (HIHS−) (Fig. 1B), suggesting that C3 deposition was due to activation of the complement system and not due to unspecific antibody binding. This observation was confirmed by quantification of gold particles, showing a median gold binding ratio of 21.8 in NHS− (12.1 to 20.9) versus 1.9 in HIHS− (1.2 to 2.6) (Fig. 1C).

FIG 1
FIG 1

Complement opsonization of C. pneumoniae in nonimmune serum. Purified C. pneumoniae EBs were incubated in nonimmune serum (NHS−) (A) or heat-inactivated nonimmune serum (HIHS−) (B) and analyzed by immunoelectron microscopy (IEM) using anti-C3c antibodies and anti-rabbit antibody with 10-nm colloidal gold as primary and secondary antibodies, respectively. (C) Gold deposition on each bacterium was quantified relative to the background gold level for each condition (NHS− and HIHS−). (D) Purified C. pneumoniae EBs were subjected to Western blot analysis and labeled against C3c. C. pneumoniae EBs were incubated in NHS− (E) or HIHS− (F) and subjected to IEM using anti-C4c antibody as the primary antibody. (G) Complement C4 deposition was quantified as described for panel C. (H) Western blot analysis of complement C4 deposition on C. pneumoniae incubated in NHS− or HIHS−. Western blot analysis was repeated three times, and representative blots are shown. IEM images were captured from at least 8 random fields for each sample in one experiment. For each bacterium, a bacterium-to-background gold deposition ratio was calculated and plotted, with each dot representing one chlamydial organism. The median ratio is presented as black lines. Bars, 200 nm.

These observations were confirmed by Western blot analysis demonstrating solid deposition of C3 in NHS− but not in HIHS− (Fig. 1D). Several high-molecular-weight bands (>110 kDa) are seen on the Western blot, indicating cleavage of the alpha chain of native C3 and covalent attachment of C3 fragments to chlamydial surface structures (Fig. 1D). The 40-kDa fragment shown in Fig. 1D corresponds to the α′2 chain of C3, demonstrating deposition of the potent opsonin iC3b.

Interestingly, we also demonstrated deposition of C4 on the chlamydial surface when incubated in NHS− (Fig. 1E) but not in HIHS− (Fig. 1F). Compared to C3, C4 was more sporadically deposited on the chlamydial surface, indicated by a lower bacterium-to-background ratio for C4 (Fig. 1E and G). The observations were confirmed by Western blot analysis, which demonstrated deposition of the C4b opsonin. C4b deposition is indicated by the emergence of high-molecular-weight bands, suggesting cleavage and exposure of the thioester group in the alpha chain of C4 along with the presence of the C4 beta (70 kDa) and gamma (35 kDa) chains (Fig. 1H). These observations suggest involvement of the lectin-mediated pathway, since the nonimmune serum tested negative for both anti-C. pneumoniae IgG and IgM, making classical-mediated C4 activation unlikely.

To evaluate opsonization during secondary infection, we incubated C. pneumoniae EBs in immune serum and investigated both antibody and complement opsonization. Immunofluorescence microscopy demonstrated that immune serum reacted with individual chlamydial organisms located in perinuclear inclusions in infected HeLa cells (Fig. 2A). The IgG antibodies reacted with the surfaces of RBs, indicated by the ring-shaped structures surrounding a DNA core (Fig. 2A). As expected, nonimmune serum did not react with the chlamydial inclusions (Fig. 2B). These observations were made on fixed and permeabilized cells and may not accurately reflect native surface labeling.

FIG 2
FIG 2

Antibody and complement opsonization of C. pneumoniae in immune serum. C. pneumoniae (5 × 105 IFU/well) was grown in in HeLa cells for 48 h, and chlamydial inclusions were immunofluorescently stained using 2-fold serial dilutions of immune serum (A) or nonimmune serum (B) as the primary antibody. Purified C. pneumoniae EBs incubated in heat-inactivated immune serum (HIHS+) (C) or heat-inactivated nonimmune serum (HIHS−) (D) were immunolabeled with anti-human IgG with 10-nm colloidal gold and subjected to immunoelectron microscopy (IEM). (E) From IEM images, chlamydial gold deposition, for each bacterium, was quantified relative to the background gold level. Purified C. pneumoniae EBs were incubated in immune serum (NHS+) (F) or heat-inactivated immune serum (HIHS+) (G) and analyzed by IEM using anti-C3c antibodies and anti-rabbit antibodies with 10-nm colloidal gold as primary and secondary antibodies, respectively. (H) Gold deposition in IEM images was quantified as described for panel E. (I) Purified C. pneumoniae EBs were subjected to Western blot analysis and labeled against C3c. (J to M) C. pneumoniae was treated as described for panels F to I except the bacteria were labeled with an anti-C4c antibody. For immunofluorescence microcopy, each serum was tested at four different dilutions, and images were captured from five random fields for each dilution. The experiment was repeated twice. In IEM, a bacterium-to-background gold deposition ratio was calculated for each bacterium and plotted, with each dot representing one chlamydial organism. The median ratio is presented as black lines. IEM images were captured from at least 8 random fields for each sample in one experiment. Western blot analysis was repeated three times, and representative blots are shown. Bars, 10 μm (A and B) and 200 nm (C, D, F, G, J, and K).

Thus, to investigate native antibody labeling and to determine if antibodies in immune serum also bind C. pneumoniae EBs, we incubated purified C. pneumoniae EBs with immune serum and evaluated IgG binding to the surface by IEM. Gold-conjugated anti-human IgG antibodies bound to the surface of chlamydial EBs when incubated in immune serum (Fig. 2C), while only a few gold particles were associated with chlamydial EBs when incubated in nonimmune serum (Fig. 2D). Quantitative analysis demonstrated a median gold binding ratio of 27.1 (23.6 to 39.1) in HIHS+ versus 1.1 (0.6 to 2.5) in HIHS−, showing that the limited bacterial gold deposition observed in nonimmune serum equals the background gold deposition (Fig. 2E). Thus, human C. pneumoniae IgG antibodies present in serum interact with epitopes located both on fixed RBs (Fig. 2A) and on native EBs (Fig. 2C and E).

Next, we wanted to investigate if anti-C. pneumoniae IgG affects complement activation and deposition on the chlamydial surface. Complement C3 was heavily deposited when C. pneumoniae was incubated in NHS+ but not in HIHS+ (Fig. 2F to H). Similar levels of C3 deposition between immune serum and nonimmune serum were observed (median ratios, 21.8 for NHS− versus 23.8 for NHS+), suggesting a negligible role of IgG in complement C3 deposition (Fig. 1A and C and 2F and H).

As expected, complement C4 was deposited on C. pneumoniae incubated in NHS+ but not in HIHS+ (Fig. 2J to L). More complement C4 deposition was observed in immune serum, in the presence of IgG, than in nonimmune serum (median ratios, 3.5 for NHS− versus 27.5 for NHS+), suggesting increased classical complement activation.

Increased complement C4 deposition in immune serum compared to that in nonimmune serum was also indicated by Western blot analysis demonstrating more intense C4 bands when bacteria were incubated in immune serum (Fig. 2M versus Fig. 1H). Several high-molecular-weight bands were observed, suggesting cleavage of the alpha chain of C4 and covalent attachment of C4b to the bacterial surface. In addition, the cleaved alpha chain (a′ chain) was observed to be around 80 kDa, suggesting noncovalent attachment of C4b (Fig. 2M).

In summary, C. pneumoniae is opsonized by iC3b and C4b in the absence of anti-C. pneumoniae IgG. C. pneumoniae IgG is able to bind both EBs and RBs, leading to increased C4b deposition on purified EBs.

Complement-mediated bacterial lysis.We wanted to investigate if the initial binding of complement and antibodies alone could interfere with reproductive infection of C. pneumoniae, since antibody binding and complement activation precede immune cell infiltration.

Electron microscopy revealed that most chlamydial organisms were intact after 30 min of incubation in serum, demonstrating no signs of bacterial lysis (Fig. 3A). However, a few bacteria showed signs of complement-mediated lysis, indicated by disruption of normal bacterial morphology together with pore-forming structures (Fig. 3B, arrowheads). Thus, we aimed to investigate if the complement cascade proceeds through the terminal complement pathway leading to MAC-induced bacterial lysis in these cases. Chlamydial organisms were incubated in nonimmune and immune serum and immunogold labeled using a monoclonal antibody, recognizing a neoepitope of the C5b-9 complex. As shown in Fig. 3C, C5b-9 is asymmetrically deposited on disintegrated chlamydial organisms and is located in close proximity to 10-nm pore-like structures, indicating MAC formation and MAC-induced lysis. Bacteria incubated in heat-inactivated serum showed no signs of bacterial lysis (Fig. 3D). MAC formation was observed in both immune and nonimmune sera, but more gold was observed in NHS+ samples (median ratio, 34.5 [12.0 to 72.4]) than in NHS− samples (median ratio, 12.1 [6.0 to 36.2]), suggesting that serum IgG promotes the formation of C5b-9 complexes. Interestingly, most gold-labeled bacteria appeared healthy, with normal morphological features and with no signs of bacterial lysis. These observations suggest that nonlytic C5b-9 complexes are formed on the chlamydial surface. Chlamydial organisms with altered morphology and signs of bacterial lysis appeared generally larger than unaffected chlamydiae, possibly representing intermediate or reticular bodies inevitably located within the EB fraction during the EB purification process. These observations imply that lytic C5b-9 formation occurs primarily on intermediate or reticular bodies but leaving EBs unaffected. Thus, some C. pneumoniae organisms are sensitive to complement-mediated lysis in both immune and nonimmune sera.

FIG 3
FIG 3

Formation of membrane attack complex (MAC) on C. pneumoniae. Purified C. pneumoniae EBs were incubated in immune and nonimmune sera and processed for IEM. (A) C. pneumoniae incubated in immune serum (NHS+) and negatively stained with PTA. (B) C. pneumoniae incubated in immune serum (NHS+) and negatively stained with PTA. (C) C. pneumoniae incubated in nonimmune serum (NHS−) and immunogold labeled against C5b-9. Area with disintegrated bacterial morphology was enlarged (hatched boxes) demonstrating 10-nm pore-like structures (yellow arrowheads) (B and C) in close proximity to gold particles (C). (D) C. pneumoniae incubated in heat-inactivated immune serum (HIHS+) and immunogold labeled against C5b-9. Anti-mouse antibody with 10-nm colloidal gold was used as a secondary antibody. IEM images were used to quantify gold deposition on C. pneumoniae incubated in nonimmune serum (E) and immune serum (F). For each chlamydial organism, the gold particle deposition was quantified by calculating a bacterium-to-background ratio and the ratios were used to create scatterplots. Each dot represents the ratio from one chlamydial organism, and the black lines show the median ratios. Bars, 200 nm. HIHS−, heat-inactivated nonimmune serum.

Our observations show that complement fragments and antibodies were deposited on almost all C. pneumoniae EBs, while only a few bacteria were directly killed by complement-mediated lysis. We therefore asked if opsonin deposition on apparently intact Chlamydia EBs had any effect on chlamydial infectivity. To answer this question, we tested the ability of immune and nonimmune sera to neutralize chlamydial infection in HeLa cells by enumerating inclusion-forming units (IFU) in each experimental condition. Figure 4 shows that the complements had a significant impact on chlamydial infectivity. Only 0.38% ± 0.2% (mean ± standard deviation [SD]) and 1.19% ± 0.72% of the initial inocula were recovered after incubation in immune (NHS+) and nonimmune (NHS−) sera, respectively. These observations indicate that the combination of the complement and antibodies more efficiently reduces chlamydial infectivity than the complement alone, but the difference was not statistically significant. When sera were heat inactivated, 19.1% ± 6.7% and 44.7% ± 5.8% (Fig. 4) of inocula were recovered, demonstrating that the complement significantly inhibited C. pneumoniae infectivity. The difference between heat-inactivated immune (HIHS+) and nonimmune (HIHS−) sera further suggests that antibodies alone can reduce C. pneumoniae infectivity. To confirm this observation, we supplemented heat-inactivated nonimmune serum (HIHS−) with the IgG fraction from a seropositive donor and evaluated chlamydial infectivity. As shown in Fig. 4, supplementing nonimmune serum with anti-C. pneumoniae IgG (HIHS−+IgG) significantly reduced chlamydial infectivity compared to that by nonimmune serum alone, confirming that anti-C. pneumoniae IgG reduces infectivity of C. pneumoniae. These data demonstrate that both antibody and, especially, complement opsonization have a profound impact on C. pneumoniae infectivity.

FIG 4
FIG 4

Serum neutralization of C. pneumoniae EBs. C. pneumoniae inocula were incubated in immune serum (NHS+), nonimmune serum (NHS−), heat-inactivated immune serum (HIHS+), heat-inactivated nonimmune serum (HIHS−), or HIHS− supplemented with 1.5 mg/ml IgG from immune serum (HIHS−+IgG) for 30 min and used to infect HeLa cells. HeLa cells were inoculated with 10.8 × 104 inclusion-forming units (IFU) of NHS-opsonized C. pneumoniae and 1.2 × 104 IFU of HIHS-opsonized C. pneumoniae. IFU were quantified by immunofluorescence staining of chlamydial inclusions. Images were captured from seven random fields in each sample using a 16× lens objective. Data were obtained from duplicate samples from three independent experiments. All data are presented as means ± SDs. Differences between means were analyzed by Welch’s ANOVA with Games-Howell multiple-comparison test. *, P < 0.05.

Complement-mediated phagocytosis of C. pneumoniae.As demonstrated above, neither complement nor antibody opsonization completely abrogates chlamydial infectivity. We therefore aimed to investigate how phagocytes participate in the clearance of opsonized C. pneumoniae. To quantitatively evaluate the role of opsonization in phagocytic uptake during primary and secondary infections, we analyzed the uptake of opsonized and nonopsonized C. pneumoniae EBs in monocytes and neutrophils by flow cytometry. Intracellular staining of cells for flow cytometry is troublesome as it requires cell permeabilization, which causes considerable changes to the physical characteristics of the cells. To avoid permeabilization, C. pneumoniae organisms were labeled with fluorescein isothiocyanate (FITC), and the uptake of FITC-labeled bacteria and FITC labeling specificity were evaluated by confocal microscopy.

Figure S1 in the supplemental material shows that FITC-labeled organisms appear small (around 0.5 μm) and round, in accordance with the morphology of chlamydial EBs. To confirm the intracellular location of these FITC signals, monocytes were stained for the cytoplasmic protein S100A8 to visualize the cell cytoplasm (Fig. S1B). As demonstrated in Fig. S1C, all FITC-positive organisms were located inside the cell cytoplasm.

FITC labeling is a rather unspecific fluorescent labeling technique, since the isothiocyanate group reacts with primary amine groups. Thus, to evaluate whether the FITC signal in Fig. S1 originated from chlamydial organisms and not contaminants such as cell debris from the chlamydial isolation procedure, we stained FITC-labeled chlamydiae with a monoclonal antibody against chlamydial lipopolysaccharide (LPS). As demonstrated in Fig. S1D and F, all FITC-positive structures reacted with the anti-Chlamydia LPS antibody, confirming that all FITC-positive structures were chlamydial organisms. Thus, FITC labeling of C. pneumoniae is efficient and specific, and the FITC-labeled bacteria locate inside the cells, enabling the bacteria to be used in a flow cytometric assay.

To analyze the effect of opsonization on phagocytosis of C. pneumoniae, isolated leukocytes were incubated with FITC-labeled C. pneumoniae for 30 min under different opsonizing conditions and subjected to flow cytometry analysis. To quantify the uptake, monocyte and neutrophil cell populations were gated as demonstrated in Fig. S2. Both cell types were gated based on forward- and side-scatter characteristics, and monocytes were additionally gated based on CD14 surface expression.

We first analyzed the role of complement-mediated opsonophagocytosis to experimentally mimic the infection conditions during primary infection. Thus, phagocytic uptake was quantified in medium containing either normal nonimmune serum (NHS−) or heat-inactivated nonimmune serum (HIHS−). As depicted in Fig. 5A and B, phagocytosis of C. pneumoniae in both monocytes and neutrophils was much more efficient in NHS− than in HIHS−, suggesting an important role for complement opsonization in phagocytosis of C. pneumoniae. Interestingly, neutrophil-mediated phagocytosis was almost absent under nonopsonizing conditions (Fig. 5B) (HIHS−), suggesting that opsonization is paramount for neutrophil phagocytosis of C. pneumoniae, while monocytes exploit opsonin-independent phagocytic pathways.

FIG 5
FIG 5

Opsonophagocytosis of C. pneumoniae in monocytes and neutrophils. The percentages of C. pneumoniae-positive monocytes (A) and neutrophils (B) incubated with C. pneumoniae at an MOI of 10 in nonimmune serum were quantified by flow cytometry. The combined roles of complement and anti-C. pneumoniae IgG in monocyte (C) and neutrophil (D) phagocytosis were tested using immune serum. Heat-inactivated nonimmune serum and heat-inactivated nonimmune serum supplemented with the IgG fraction from immune serum were used as the controls to determine the role of IgG in phagocytosis in monocytes (E) and neutrophils (F). The percentages of C. pneumoniae-positive monocytes (G) and neutrophils (H) incubated with C. pneumoniae at an MOI of 1 in immune serum and nonimmune serum. Data were obtained from duplicate samples from three independent experiments except for those in panels E and F, which were obtained from duplicate samples from two independent experiments. A minimum of 8,000 gated events were obtained from each sample. All data are presented as means ± SDs. One-way ANOVA with Tukey’s post hoc test, Welch’s ANOVA with Games-Howell multiple-comparison test, or Welch’s t test was used to compare column means. *, P < 0.05; NHS+, immune serum; NHS−, nonimmune serum; HIHS+, heat-inactivated immune serum; HIHS−, heat-inactivated nonimmune serum; HIHS−+IgG, heat-inactivated nonimmune serum supplemented with IgG fraction from immune serum.

Phagocytosis in the presence of antichlamydial antibodies.We demonstrated that monocyte- and, especially, neutrophil-mediated phagocytosis of C. pneumoniae was critically dependent on complement opsonization. Complement activation and activity in the alveolar space are reduced compared to those in serum and are primarily dependent on classical complement activation (15, 16). Thus, we sought to investigate whether anti-C. pneumoniae antibodies could mediate efficient phagocytosis or potentiate complement-mediated phagocytosis, as both mechanisms may be important to understand the biology of C. pneumoniae reinfections.

By comparing the heat-inactivated immune serum (Fig. 5C and D) (HIHS+) with heat-inactivated nonimmune serum (Fig. 5A and B) (HIHS−), it is evident that antibody opsonization increased phagocytic uptake of C. pneumoniae in both monocytes (3-fold increase) and, especially, neutrophils (17-fold increase); however, this effect was significantly lower than for complement alone. To confirm these observations, we purified the IgG fraction from immune serum and supplemented HIHS− with purified IgG. As demonstrated in Fig. 5E and F, supplementing nonimmune serum with the IgG fraction from immune serum increased the phagocytic uptake in both monocytes and neutrophils, confirming that the observed differences are due to anti-C. pneumoniae antibodies. We could not assess whether the presence of IgG potentiated complement-mediated phagocytosis, since both monocyte and neutrophil cell populations were saturated with bacteria when complement competent serum was used (Fig. 5A to D) (NHS±). We therefore reanalyzed these samples using C. pneumoniae at a multiplicity of infection (MOI) of 1 and showed that monocytes and neutrophils more efficiently ingest C. pneumoniae when both complement and IgG antibodies are present (Fig. 5G and H).

Intracellular survival of ingested bacteria.C. pneumoniae is ingested through both complement- and antibody-dependent mechanisms. Evidence from other intracellular bacteria shows that the intracellular fate of ingested bacteria is highly dependent on the route of uptake, and it is therefore important to study the intracellular fate of C. pneumoniae under the opsonizing conditions we have demonstrated here.

To evaluate the intracellular fate of ingested bacteria, peripheral blood mononuclear cells (PBMCs) and neutrophils containing opsonized and nonopsonized bacteria were lysed by ultrasonication, and liberated bacteria were used to infect HeLa cell monolayers.

Statistically significantly fewer IFU were recovered when bacteria were ingested by PBMCs in the presence of immune serum (NHS+, 45 ± 19 IFU/μl) and nonimmune serum (NHS−, 70 ± 16 IFU/μl) than when bacteria were ingested in media supplemented with heat-inactivated immune serum (HIHS+, 315 ± 49 IFU/μl) and nonimmune serum (HIHS−, 452 ± 25 IFU/μl) (Fig. 6A). Similar results were obtained for neutrophils, with 99 ± 4 IFU/μl and 141 ± 32 IFU/μl recovered IFU in immune and nonimmune sera, respectively (Fig. 6B). Serum heat inactivation caused a statistically significant increase in IFU recovery from both immune serum (Fig. 6B) (HIHS+, 452 ± 139 IFU/μl) and nonimmune serum (Fig. 6B) (HIHS−, 1,151 ± 259 IFU/μl). Although more bacteria were present intracellularly in cells incubated in NHS than in those in HIHS (Fig. 5A to D), more reproductive bacterial organisms were recovered from cells in HIHS. These data suggest that complements not only facilitate rapid and efficient uptake of C. pneumoniae but also mediate effective intracellular neutralization. We also observed statistically significantly fewer IFU recovered from bacteria ingested in the presence of heat-inactivated immune serum (HIHS+) than in the presence of heat-inactivated nonimmune serum (HIHS−) in both cell types (Fig. 6A and B), suggesting that antibodies also play a role in bactericidal activity against C. pneumoniae.

FIG 6
FIG 6

Intracellular neutralization of opsonized C. pneumoniae in PBMCs and neutrophils. C. pneumoniae (MOI = 10) was incubated with PBMCs (A) and neutrophils (B) under different culture conditions for 30 min, and cell lysates were prepared by ultrasonication. Lysates were used to infect HeLa cell monolayers, and inclusion-forming units (IFU) were enumerated after 48 h by immunofluorescence microscopy. Intracellular localization of C. pneumoniae in monocytes after 30 min was investigated by immunofluorescence staining and confocal microscopy of C. pneumoniae (C) and LAMP1 (D). (E) Overlay image. (F) Monocyte incubated with mock control. Confocal microscopy was repeated twice for all four (NHS+, NHS−, HIHS+, and HIHS−) conditions, and images were captured from five random fields in each sample. Bars, 5 μm. Quantitative data were obtained from duplicate samples from three independent experiments. Data are presented as means ± SDs. Differences between means were analyzed by Welch’s ANOVA with Games-Howell multiple-comparison test. *, P < 0.05; NHS+, immune serum, NHS−, nonimmune serum; HIHS+, heat-inactivated immune serum; HIHS−, heat-inactivated nonimmune serum.

Studies performed on M. tuberculosis suggest that bacterial uptake by complement receptors delays phagosomal maturation and phagolysosomal fusion, thereby facilitating intracellular bacterial survival. Our results suggest the opposite: that complement-mediated uptake leads to rapid intracellular neutralization of C. pneumoniae. We therefore tested whether C. pneumoniae was targeted to lysosomal compartments in monocytes after complement-mediated ingestion. Figure 6C to E shows that C. pneumoniae localizes within LAMP1-positive vesicular structures in monocytes after a 30-min incubation in medium supplemented with nonimmune serum (NHS−). Staining of mock-infected cells demonstrates that the FITC-signal in LAMP1-positive structures originates from chlamydial organisms (Fig. 6F). Thus, complement-mediated uptake of C. pneumoniae in monocytes directs ingested bacteria to lysosomal compartments.

DISCUSSION

In the present study, we demonstrated that C. pneumoniae EBs are opsonized with both IgG and the complement opsonins iC3b and C4b when incubated in human serum. We showed that complement activation limits C. pneumoniae infection in multiple ways: by interfering with C. pneumoniae entry into permissive cells, by direct MAC-induced lysis, and by tagging bacteria for efficient phagocytosis by both monocytes and neutrophils.

The activation and deposition of complement C3 and the terminal complement complex on C. pneumoniae demonstrated in this study were previously shown by Cortes and colleagues (18). Interestingly, using electron microscopy, we observed C5b-9 deposition without pore formation and bacterial lysis. C5b-9 complexes associated with pore-like structures and altered bacterial morphology were primarily observed on reticular and intermediate bodies. Thus, our observations suggest that extracellular EBs may be resistant to MAC-induced lysis. It is likely that the cysteine-rich proteins in the highly cross-linked chlamydial outer membrane of EBs provide a physical barrier that hinders components of the terminal complement complex to interact with the outer membrane lipid bilayer. Alternatively, MAC formation could be inhibited by recruitment of host-derived complement regulators to the EB surface. Thus, it was previously demonstrated that Chlamydia trachomatis binds the MAC inhibitor vitronectin; however, it was not investigated if vitronectin modulates complement deposition on the chlamydial surface (19).

Cortes et al. (18) concluded that properdin and alternative complement activation were indispensable for C3b deposition and complement-mediated neutralization. Interestingly, we found that complement C4 is deposited on the surface of C. pneumoniae. Deposition of complement C4 in both immune and nonimmune sera suggests that also the lectin-mediated activation pathway is involved in C. pneumoniae-induced complement activation. This observation is supported by previous studies demonstrating that different lectins can bind the surface of C. pneumoniae (20, 21). Moreover, quantification of gold labeling and semiquantitative Western blot analysis suggested that more C4 was deposited in immune serum than in nonimmune serum, indicating activation of the classical pathway. This is supported by the finding that IgG1 is the predominant IgG subclass raised against C. pneumoniae and, to a lesser extent, IgG3 and both IgG1 and IgG3 are efficient complement activators (22). The IgG subclasses raised against C. pneumoniae may also explain why we observed an IgG-dependent inhibition of C. pneumoniae infectivity in HeLa cells (Fig. 4). Studies on C. trachomatis showed that HeLa cells express FcγRIII and that Fc-mediated endocytosis through this receptor promotes reproductive infection of C. trachomatis (23, 24). It is likely that C. trachomatis is more prone to FcγRIII-mediated uptake than C. pneumoniae, since IgG3 is the predominant IgG subclass raised against C. trachomatis (22). Thus, the IgG subclass distribution seems important for the infectivity and the reproductive outcome of Chlamydia spp. in vitro.

A possible explanation for the discrepancies between our results and the findings by Cortes et al. (18) could be that classical- and/or lectin-mediated activation pathways are abrogated at the C4 level, leading to pathway termination before C3 cleavage. C4 binding protein (C4bp), a fluid-phase negative complement regulator, can terminate complement activation at the C4 level by inducing C4b cleavage and C2a dissociation from the classical C3 convertase (25). Several bacterial pathogens are able to recruit and bind C4bp, leading to complement resistance, but this has not yet been demonstrated for chlamydial organisms (26, 27).

The alternative activation pathway seems important for C. pneumoniae-induced complement activation in vitro, but the significance of this pathway during in vivo lung infections is still elusive. In the lungs, complement activation proceeds primarily through the classical pathway, and alternative complement activation fails to activate and deposit C3 on both mycobacteria and streptococci in bronchoalveolar lavage fluid (15, 28). It remains uncertain whether C. pneumoniae is opsonized in the alveolar space of the lungs, but the classical complement activation is functional, and during secondary infection, complement-activating IgG is present in the alveolar space (15).

We show that the complement efficiently facilitates phagocytosis of C. pneumoniae in both monocytes and neutrophils and that most opsonized bacteria are unable to cause reproductive infection when liberated from phagocytes. Complement receptor 3 (CR3, CD11b/CD18) recognizes different C3 cleavage fragments, including iC3b identified on C. pneumoniae EBs here. CR3 is, together with Fcγ receptors, the most important phagocytic receptor and is widely expressed on both monocytes and neutrophils. It is therefore likely that complement-mediated phagocytosis takes place via CR3, leading to intracellular destruction. This is different from CR-mediated uptake of other intracellular bacteria. CR-mediated uptake of M. tuberculosis into human monocytes and macrophages allows safe entrance by delaying phagosomal maturation and phagolysosomal fusion (11). We observed that C. pneumoniae is rapidly targeted to LAMP1-positive intracellular compartments showing efficient phagosomal trafficking, facilitating intracellular neutralization of the chlamydial reproductive potential. Thus, safe entrance into host cells through CR-mediated phagocytosis is not a conserved mechanism for all intracellular bacteria. This is supported by the observation that the facultative intracellular bacterium Francisella tularensis fails to escape phagosomal trafficking in murine bone marrow macrophages when opsonized with either complement or IgG (29). Knockout of CD11b, a subunit of the heterodimeric CR3, leads to phagosomal escape with cytosolic localization of F. tularensis, supporting that CR3 plays opposing roles during experimental infections with different intracellular bacteria (29).

Direct interaction between opsonized bacteria and phagocytic opsonin receptors provides one explanation for the increased phagocytic uptake and intracellular neutralization observed here. Phagocytic priming by the complement anaphylatoxins C3a and C5a may provide another likely explanation. In an in vivo Chlamydia psittaci lung infection model, C5aR−/− mice displayed slightly worse clinical scores than wild types (WT) during early-stage infection (30). The same authors later demonstrated that C3a and its receptor C3aR are critically involved in antichlamydial immunity in the same C. psittaci lung infection model (31). Both receptors are expressed on monocytes and neutrophils and can potentiate phagocytic functions by both inducing an increased surface expression of complement receptors and inducing a hostile intracellular environment (32, 33). C5a induces increased surface expression of CD11b and increased phagocytosis of Escherichia coli in neutrophils (34) and can also potentiate intracellular killing of E. coli by increasing the oxidative burst in a whole-blood assay (35).

In addition, priming neutrophils with C5a leads to translocation of CR1-containing secretory vesicles to the cell surface (36, 37). CR1 recognizes the activation products C3b and C4b together with several other complement-related proteins (38). We demonstrate, for the first time, that C4b is deposited on the surface of C. pneumoniae; hence, C5a-induced translocation of CR1 to the plasma membrane may induce increased bacterial phagocytosis. However, the role of C4b in opsonophagocytosis remains elusive.

We demonstrated that C. pneumoniae ingested in the presence of complement-competent serum had impaired ability to cause reproductive infection in HeLa cells compared to that of bacteria ingested under nonopsonizing conditions. Intracellular survival and replication of C. pneumoniae within phagocytes have previously been evaluated in several in vitro studies; however, these have been conducted under various experimental conditions. Both C. pneumoniae and C. trachomatis are able to replicate in human neutrophils when ingested under nonopsonizing conditions (10, 39). Rajeeve et al. showed that C. trachomatis uses the chlamydial protease-like activating factor (CPAF) to inhibit neutrophil activation and cell death, thereby creating a replicative environment (39). Since CPAF is highly conserved among chlamydiae, C. pneumoniae may use a similar CPAF-dependent mechanism to survive and replicate in neutrophils.

Similarly, coincubating monocytes and C. pneumoniae under nonopsonizing conditions leads to recovery of chlamydial progeny up until 48 h postinfection (9, 40), suggesting that opsonin-independent uptake also facilitates C. pneumoniae survival in monocytes. This idea was further supported by the observation that differentiated macrophages support intracellular replication of C. pneumoniae. Differentiated macrophages, but not peripheral blood monocytes, express the mannose receptor (MR), which facilitates intracellular survival of M. tuberculosis, and the MR has also been linked to phagocytic uptake of chlamydial organisms (12, 41, 42). Others used commercially available human serum from AB blood-type donors in the infection medium combined with centrifugation. These studies were unable to recover chlamydial progeny after 6 and 48 h but detected 16S rRNA, suggesting metabolic but not replicative activity (8, 43). However, neither complement functional activity nor serostatus was evaluated for these sera, making it difficult to draw any conclusions on the role of opsonization. We demonstrated that both complement and IgG opsonization potentiates phagocytosis and intracellular killing of C. pneumoniae, suggesting that the uptake mechanism is important for the intracellular fate of C. pneumoniae, previously demonstrated for other chlamydial organisms (44).

Here, we demonstrated that complement opsonization and, to a lesser extent, IgG opsonization inhibit C. pneumoniae progeny infection and facilitate C. pneumoniae uptake in human monocytes and neutrophils. Ingested bacteria are rapidly trafficked to destructive intracellular compartments and eliminated, showing that opsonization and phagocytosis are efficient means of controlling extracellular C. pneumoniae organisms. Thus, opsonization is a critical factor to include in future in vitro studies exploring the interaction between C. pneumoniae and human phagocytes.

MATERIALS AND METHODS

Antibodies and reagents.The following primary antibodies were used in this study: phycoerythrin (PE) anti-human CD14 (clone M5E2; BioLegend, CA, USA), monoclonal antibody (MAb) 15.2.3 against chlamydial LPS (45), polyclonal antibody (PAb)198 against C. pneumoniae outer membrane (46), anti-MRP8 (S100A8) (Abcam, UK), anti-LAMP1 (Sino Biological, China), polyclonal rabbit anti-human C3c (Agilent Technologies, Glostrup, Denmark), polyclonal rabbit anti-human C4c (Agilent), and monoclonal mouse anti-human C5b-9 (clone aE11; Agilent).

The following secondary antibodies were used in this study. Goat anti-human IgG and goat anti-rabbit and goat anti-mouse antibodies conjugated to 10-nm colloidal gold (British BioCell, Cardiff, UK) were used for immunoelectron microscopy. Horseradish peroxidase-conjugated AffiniPure goat anti-human Fcγ-specific IgG was used for enzyme-linked immunosorbent assay (ELISA). Anti-rabbit IgG conjugated to alkaline phosphatase was used for Western blotting (Sigma-Aldrich, MO, USA). FITC-conjugated goat anti-human IgG, Fcγ fragment specific (Jackson ImmunoResearch, Cambridge, UK), Alexa Fluor (AF)555 goat anti-rabbit, AF555 goat anti-mouse, and AF647 goat anti-rabbit (Invitrogen of Thermo Fisher Scientific, MA, USA) antibodies were used for immunofluorescence staining.

3,3′,5,5′-Tetramethylbenzidine (TMB-ONE) and 5-bromo-4-chloro-3-indolyl phosphate (BCIP)-Nitro Blue Tetrazolium (NBT) were purchased from Kementec (Taastrup, Denmark) and used for ELISA and Western blotting, respectively.

Cell lines and culture.Baby hamster kidney (BHK) and HeLa cell lines were purchased from the American Type Culture Collection (ATCC, Manassas, VA) and cultured in complete medium consisting of RPMI 1640 supplemented with 5% fetal calf serum (FCS) and 0.01 mg/ml gentamicin and maintained at 37°C and 5% CO2. Cell lines were tested to determine they were free of mycoplasma by Hoechst (33258) staining.

Bacterial organism and propagation.C. pneumoniae (CDC/CWL-029/VR1310) was purchased from ATCC and cultivated in BHK cells in complete medium with 2 μg/ml cycloheximide and 0.1% glucose. The infected cells were cultured at 37°C and 5% CO2 for 72 h. C. pneumoniae EBs were purified by density gradient centrifugation as previously described (46). Mock inoculum was prepared by processing uninfected BHK cells in parallel with infected cells.

Serum samples.Serum samples were obtained from 10 healthy volunteers at Aalborg University, Denmark. Blood was drawn by venipuncture and collected in S-Monovette serum tubes (Sarstedt, Nümbrecht, Germany), and serum was isolated immediately according to the manufacturer’s instructions. Isolated serum was immediately stored on ice before freezing at −80°C for later use. Serum heat inactivation was performed for 30 min at 56°C. Serum containing anti-C. pneumoniae IgG is denoted immune serum, and serum without anti-C. pneumoniae IgG is denoted nonimmune serum. Throughout this paper, the following abbreviations were used to describe the different sera: NHS+, immune serum; NHS−, nonimmune serum, HIHS+, heat-inactivated immune serum; and HIHS−, heat-inactivated nonimmune serum.

All protocols were approved by the regional ethics committee of Region Nordjylland (N-20150073), and all experiments were carried out according to the Declaration of Helsinki. All participants gave written informed consent to participate in the study.

ELISA.C. pneumoniae-specific IgG was measured using Chlamydia pneumoniae-IgG ELISA Medac plus (Medac, Wedel, Germany) with C. pneumoniae outer membrane complex as the antigen. Absorbances were measured at 450 nm on a Sunrise microplate reader (Tecan, Männedorf, Switzerland).

Immunofluorescence validation of serum reactivity.HeLa cell monolayers, seeded on coverslips, were infected with 5 × 105 inclusion-forming units (IFU) C. pneumoniae by centrifugation at 1,000 × g for 30 min at 37°C followed by a 30-min incubation at 37°C and 5% CO2. Medium containing extracellular bacteria was removed, and cells were incubated in complete medium with 2 μg/ml cycloheximide and 0.1% glucose for 48 h and subsequently fixed in 3.7% formaldehyde for 20 min at 4°C. Cells were permeabilized in ice-cold methanol for 10 min, blocked in 0.5% bovine serum albumin (BSA) for 15 min at 37°C, and incubated with serial dilutions of immune and nonimmune sera for 30 min at 37°C. Cells were washed three times in phosphate-buffered saline (PBS) and incubated with FITC-conjugated anti-human IgG secondary antibody diluted 1:200 in 0.1% BSA in PBS for 30 min at 37°C. Cells were washed three times in PBS and counterstained with 2 μM To-Pro-3 iodide (Invitrogen of Thermo Fisher Scientific) for 20 min at room temperature. Cells were imaged using a Leica TCS SP5 confocal laser scanning microscope with HCX PL Apo 63×/1.40 and CX PL Apo 100×/1.47 lens objectives (Leica Microsystems, Wetzlar, Germany).

Immunogold labeling and transmission electron microscopy.Purified C. pneumoniae EBs were incubated in 50% NHS or HIHS for 30 min at 37°C and washed three times in PBS, with centrifugation at 15,000 × g for 15 min at 4°C between each wash. Processing and immunogold labeling of bacteria for transmission electron microscopy (TEM) were performed as previously described (47). Briefly, serum-coated (NHS or HIHS) C. pneumoniae EBs were added to carbon-coated 400-mesh glow-discharged nickel grids and washed three times in PBS before blocking in 1% ovalbumin (Sigma-Aldrich) in PBS (pH 6.5). To determine the binding of human antibodies to the chlamydial surface, grids were incubated for 30 min at 37°C with goat anti-human IgG conjugated to 10-nm colloidal gold diluted 1:25 in 0.5% ovalbumin in PBS. To determine complement binding, samples were incubated with primary antibodies (anti-C3c, 1:200; anti-C4c, 1:200; anti-C5b-9, 1:20) diluted in 0.5% ovalbumin in PBS for 30 min at 37°C. Grids were washed in three drops of PBS and subsequently incubated for 30 min at 37°C with secondary antibodies (goat anti-rabbit or goat anti-mouse antibody conjugated to 10-nm colloidal gold) diluted 1:25 in 0.5% ovalbumin in PBS. The grids were washed in three drops PBS, incubated in three drops of 0.5% cold fish gelatin in PBS, and negatively stained with 0.5% phosphotungstic acid (PTA). Grids were blotted dry on filter paper and investigated using a Jeol 1010 transmission electron microscope (Jeol, Tokyo, Japan). Gold deposition was quantified as previously described (47). Briefly, a minimum of seven random fields were imaged for each grid containing at least 12 chlamydial organisms in total. For each chlamydial organism, gold particles on the bacteria and in the background were counted, and the density of deposition was calculated for both (gold per area). From these numbers, a bacterium-to-background ratio was calculated to quantitatively describe bacterial gold deposition relative to background deposition. From these ratios, the median and interquartile ranges (IQRs) were calculated for each sample.

Western blotting.SDS-PAGE and Western blotting were performed essentially according to Lausen et al. (47). Briefly, purified C. pneumoniae EBs were incubated in 50% serum for 30 min at 37°C, and unbound complement was removed by washing three times in PBS, with centrifugation at 15,000 × g for 15 min between each wash. The chlamydial pellet was lysed in SDS sample buffer plus 5% β-mercaptoethanol, and proteins were separated on an 8% polyacrylamide gel. Proteins were blotted onto a nitrocellulose membrane (GE Healthcare Life Sciences, Marlborough, MA) in Tris-glycine buffer (25 mM Tris, 192 mM glycine) plus 20% methanol for 2 h at 100 V using a TE22 Mighty Small transfer tank (Hoefer, Inc., Holliston, MA).

Membranes were blocked for 30 min at 37°C in 3% gelatin in Tris-buffered saline (TBS). Antibodies diluted in TB plus 0.05% Tween 20 (TBST) plus 0.2% gelatin were added to the membranes and left for incubation for 1 h at 37°C. Blots were washed three times in TBST and incubated with alkaline phosphatase-conjugated secondary antibody for 1 h at 37°C. Antigen-antibody complexes were visualized by adding BCIP-NBT (Kementec) alkaline phosphatase substrate.

Serum neutralization assay.C. pneumoniae EBs were incubated in RPMI 1640 supplemented with 10% human serum for 30 min at 37°C. Opsonized bacteria (10.8 × 104 IFU for NHS samples and 1.2 × 104 IFU for HIHS samples) were used to infect HeLa cell monolayers as described above. Cells were processed for immunofluorescence microscopy as described above with minor modifications. PAb198 (1:400) and FITC-conjugated goat anti-rabbit (1:200) antibodies were used as primary and secondary antibodies, respectively, and cell nuclei were stained with 2 μg/ml 4′,6-diamidino-2-phenylindole (DAPI).

The cells were inspected using a Leica LSM550 fluorescence microscope, and images were captured from seven random fields in each sample. All samples were analyzed in duplicates and repeated in three independent experiments. The number of IFU was enumerated and expressed as a percentage of initial inoculum. Data from the three experiments are expressed as means ± standard deviations (SDs).

FITC-labeling of bacteria.C. pneumoniae inoculum was suspended in 0.1 M NaHCO3 (pH 9) and centrifuged for 15 min at 15,000 × g. Fluorescein isothiocyanate isomer 1 (FITC; Sigma, MO, USA) was added at a concentration of 0.1 mg/ml in 0.1 M NaHCO3 buffer, and bacteria were incubated for 1 h protected from light. The labeled bacteria were washed three times in PBS to remove unbound FITC, and the final bacterial pellet was suspended in sucrose phosphate buffer (2SP), aliquoted, and stored at −80°C.

Immune cell isolation and culture.Peripheral blood was obtained from two donors (one seropositve and one seronegative) and collected in S-Monovette (Sarstedt) EDTA tubes. Blood was layered on Polymorphrep density gradient medium (Axis-Shield, Dundee, UK) and centrifuged for 40 min at 300 × g at 20°C. Layers with peripheral blood mononuclear cells (PBMCs) and polymorphonuclear cells were harvested and collected in the same tube when used for flow cytometry. The two cell populations were kept separate when used for immunofluorescence microscopy. Platelets were removed by centrifugation at 120 × g for 10 min at 20°C. Cells were resuspended in complete RPMI 1640 medium.

Immunofluorescence staining and microscopy of immune cells.For immunofluorescence staining, PBMCs were seeded in 8-well Nunc Lab-Tek chamber slides at a density of 9 × 105 cells/well. Cells were left to adhere for 90 min, and nonadherent cells were removed by washing twice in PBS. Bacteria were added at a multiplicity of infection (MOI) of 10 in RPMI 1640 plus 10% human serum.

Nonphagocytized bacteria were removed by washing three times in PBS, and cells were processed for immunofluorescence staining as described above. PAb198 (1:400), anti-LAMP1 (1:400), anti-S100A8 (1:300), and MAb 15.2.3 (1:20) antibodies were used as primary antibodies and AF555-conjugated goat anti-mouse, AF555-conjugated goat anti-rabbit, and AF647-conjugated goat anti-rabbit antibodies were used as secondary antibodies at a 1:200 dilution. Cell nuclei were stained using 2 μM To-Pro-3 iodide (Invitrogen of Thermo Fisher Scientific). Cells were imaged using a Leica TCS SP5 confocal laser scanning microscope with a CX PL Apo 100×/1.47 lens objective (Leica Microsystems).

Phagocytosis assay.FITC-labeled C. pneumoniae EBs were opsonized in RPMI 1640 plus 10% human serum for 15 min at 37°C. Bacteria (1 × 107 or 1 × 106 bacteria/tube) and immune cells (1 × 106 cells/tube) were mixed in 5-ml polypropylene tubes (MOI = 10 and MOI = 1, respectively) and coincubated for 30 min at 37°C, and cells were subsequently processed for flow cytometry.

Flow cytometry.Cells were washed in cold PBS plus 0.05% sodium azide to remove nonphagocytized bacteria. Fc receptors were blocked using 20 μg/ml human IgG (Sigma) for 15 min at 4°C. Primary fluorochrome-conjugated antibodies, diluted in PBS plus 0.05% sodium azide plus 0.01% BSA, were added, and cells were stained for 30 min at room temperature. Unbound antibody was removed by washing in PBS, and cell viability was assessed using eBioscience fixable viability dye eFluor 450 according to the manufacturer’s instructions. The cells were washed twice in PBS, fixed in 1% formaldehyde for 20 min at 4°C, and analyzed on a CytoFLEX flow cytometer (Beckman Coulter, CA, USA).

IgG purification.Serum from one seropositive donor was mixed 1:1 with binding buffer (1 M glycine, 150 mM NaCl, pH 8.5), and the serum IgG fraction was isolated on an affinity chromatography column packed with GammaBind Plus protein G Sepharose (GE Healthcare). Bound IgG was eluted from the column using elution buffer containing 100 mM glycine-HCl (pH 2.7) and collected in 25 fractions in tubes containing 1 M Tris-HCl (pH 9) to restore neutral pH. The protein concentration in each fraction was determined using a Pierce bicinchoninic acid (BCA) protein assay kit (Thermo Scientific) according to the manufacturer’s instructions.

Intracellular survival assay.PBMCs and neutrophils were coincubated with bacteria as described above. Cells were washed three times in PBS to remove nonphagocytized bacteria. The cell pellet was suspended in 2SP buffer, and cells were lysed by ultrasonication to liberate ingested bacteria. The cell lysates were used to infect HeLa cells, and IFU were quantified from duplicate samples in three independent experiments as described above.

Data analysis.Flow cytometry data were analyzed using FlowLogic v.7.2.1 (Inivai Technologies, Mentone, Australia). A one-way analysis of variance (ANOVA) with Tukey’s post hoc or Welch ANOVA with Games-Howell multiple-comparison test was used to investigate differences between multiple means. Differences between means from two independent groups were investigated using a Student’s independent t test or Welch’s t test. All statistical analyses were performed in SPSS Statistics 25 (IBM, Armonk, NY).

ACKNOWLEDGMENTS

We thank Ditte Bech Laursen for technical assistance.

This study was funded by the Aase and Ejner Danielsen Foundation, the Obel Family Foundation, and the Joint Strategic Research Program on Fighting Antibiotic Resistance (Aalborg University and University of Southern Denmark).

We declare no conflict of interest.

FOOTNOTES

    • Received 14 February 2020.
    • Returned for modification 16 March 2020.
    • Accepted 7 April 2020.
    • Accepted manuscript posted online 13 April 2020.

  • Supplemental material is available online only.

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KEYWORDS

Chlamydia pneumoniae
opsonization
phagocytosis
monocytes
neutrophils
complement

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