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. 2015 Apr 16;145(3):429–442. doi: 10.1111/imm.12460

Peptidoglycan recognition protein–peptidoglycan complexes increase monocyte/macrophage activation and enhance the inflammatory response

Mauricio C De Marzi 1,2,3,*, Marcos Todone 1,2,3,*, María B Ganem 1, Qian Wang 4, Roy A Mariuzza 4, Marisa M Fernández 1, Emilio L Malchiodi 1,✉
PMCID: PMC4479541  PMID: 25752767

Abstract

Peptidoglycan recognition proteins (PGRP) are pattern recognition receptors that can bind or hydrolyse peptidoglycan (PGN). Four human PGRP have been described: PGRP-S, PGRP-L, PGRP-Iα and PGRP-Iβ. Mammalian PGRP-S has been implicated in intracellular destruction of bacteria by polymorphonuclear cells, PGRP-Iα and PGRP-Iβ have been found in keratinocytes and epithelial cells, and PGRP-L is a serum protein that hydrolyses PGN. We have expressed recombinant human PGRP and observed that PGRP-S and PGRP-Iα exist as monomer and disulphide dimer proteins. The PGRP dimers maintain their biological functions. We detected the PGRP-S dimer in human serum and polymorphonuclear cells, from where it is secreted after degranulation; these cells being a possible source of serum PGRP-S. Recombinant PGRP do not act as bactericidal or bacteriostatic agents in the assayed conditions; however, PGRP-S and PGRP-Iα cause slight damage in the bacterial membrane. Monocytes/macrophages increase Staphylococcus aureus phagocytosis in the presence of PGRP-S, PGRP-Iα and PGRP-Iβ. All PGRP bind to monocyte/macrophage membranes and are endocytosed by them. In addition, all PGRP protect cells from PGN-induced apoptosis. PGRP increase THP-1 cell proliferation and enhance activation by PGN. PGRP-S–PGN complexes increase the membrane expression of CD14, CD80 and CD86, and enhance secretion of interleukin-8, interleukin-12 and tumour necrosis factor-α, but reduce interleukin-10, clearly inducing an inflammatory profile.

Keywords: innate immunity, monocyte, peptidoglycan, peptidoglycan recognition protein, Staphylococcus aureus

Introduction

Innate immunity provides the first line of defence against invading microorganisms employing different strategies to discriminate self from foreign antigens. One of these strategies, known as ‘microbial non-self recognition', consists of the host's ability to recognize common microbial products such as lipopolysaccharide, CpG and peptidoglycan (PGN) among others, which are named pathogen-associated molecular patterns. The PGN interact with humoral (lysozyme, soluble CD14) and cellular (macrophages, lymphocytes) components of the immune system.1 Macrophage activation, through CD14 and Toll-like receptor 2 (TLR2), induces the production of cytokines responsible for some clinical manifestations in infections.2–5 PGN recognition proteins (PGRP) are molecules that bind PGN, such as CD14 and TLR2, or hydrolyse PGN, such as lysozymes. PGRP are highly conserved from insects to mammals and can be grouped as short PGRP [PGRP-S of 19 000–20 000 molecular weight (MW)], intermediate PGRP (PGRP-I of 40 000–45 000 MW) and long PGRP (PGRP-L of 30 000–90 000 MW).6–9 In the human genome there are three genes expressing four different PGRP designated as PGRP-S, PGRP-L, PGRP-Iα and PGRP-Iβ, also named as PGLYRP-1, PGLYRP-2, PGLYRP-3 and PGLYRP-4, respectively.10

The PGRP were described as being involved in phagocytosis by Drosophila macrophages, activation of the Toll pathway and in autophagy.7,11,12 Most PGRP are soluble proteins present in intracellular vesicles.13,14 Human PGRP-S was implicated in the intracellular destruction of bacteria by polymorphonuclear (PMN) cells. Murine and human PGRP-S were first described as bacteriostatic proteins whereas bovine PGRP-S showed bactericidal activity;1 however, it was later demonstrated that PGRP-S, PGRP-Iα and PGRP-Iβ can act as bactericidal homo- or hetero-dimers if they are N-glycosylated and in the presence of divalent cations.14–16 It was found that internalized PGN is associated with PGRP-Iα in the cytoplasm of intestinal epithelial cells.17 It was recently found that PGRP-L, PGRP-Iα and PGRP-Iβ were induced in human corneal epithelial cells in response to ligands of TLR1, TLR2, TLR3, TLR5 and TLR6 (such as PGN), which were localized predominantly in the cell membrane and cytoplasm.18 Human PGRP-L is an enzyme that hydrolyses PGN and would be present in serum, liver and intraepithelial lymphocytes.8,14,19

The three-dimensional structures of different PGRP reveal a common topology with the T7 lysozyme.13,20–23 PGRP have at least two binding sites, one for PGN recognition, and another that would interact with non-identified host proteins.21,24,25 The hydrophobic nature of the PGRP-LB groove indicates that the back face would serve for subsequent signalling after PGRP molecule clustering by binding to polymeric cell wall components.20 Binding and crystallographic studies demonstrate that CPGRP-S bind to lipopolysaccharide and PGN.26,27

Peptidoglycan is an excellent target for most clinically effective antibiotics and also for recognition by the innate immune system, which has several PGN recognition proteins including CD14, TLR2, PGRP, nucleotide-binding oligomerization domain proteins 1 and 2; and PGN-lytic enzymes like lysozymes and amidases.25 The fact that microbe recognition and phagocytosis are principal aspects of innate immunity and the poor knowledge about PGRP localization and the mechanisms in which human PGRP are involved, led us to analyse the presence of PGRP-S, PGRP-Iα and PGRP-Iβ in human samples to elucidate their effect on monocyte/macrophage activity.

Materials and methods

PGRP expression and purification

DNA encoding human PGRP-S, the C-terminal domain of PGRP-Iα (PGRP-IαC), which includes residues 162–341, and the N-terminal domain of human PGRP-Iβ (PGRP-IβN), which includes residues 51–208, were cloned in the pT7-7 vector resistant to ampicillin.13,24 Recombinant PGRP were expressed as inclusion bodies in Escherichia coli BL21 (DE3) cells. After purification, inclusion bodies were refolded and purified as previously described.28–30 All rPGRP were treated with polymyxin and verified to be lipopolysaccharide-free by the Limulus amoebocyte lysate assay (< 0·03 endotoxin U/ml; Pyrotell Associates of Cape Cod, Falmouth, MA).

PGN

Cultures from Staphylococcus aureus, Enterococcus faecalis, E. coli and Brucella abortus glycerol stocks were performed in luria broth (LB) medium and grown with shaking at 37° to reach an absorbance of 0·8 at 600 nm. Cells were centrifuged, washed and stored at −20° until use. PGN was obtained from the bacterial cell wall as described elsewhere.31

Human samples and cells cultures

Human blood and serum samples were obtained from healthy donors after receiving their written consent. Human peripheral blood cells (PBMC, 90% pure) were separated from heparinized blood by centrifugation through Ficoll–Hypaque. PMN cells (98% pure) were also isolated from blood by centrifugation through Ficoll–Hypaque followed by dextran sedimentation. All cell manipulations were performed under sterile conditions at 4°, so minimizing PMN cell priming and stimulation. Human PMN cells (106 cells per assay) were stimulated for degranulation with 10 mmol/l fMLP in 37° warm PBS for 15 min. Alternatively, PMN cells were lysed in 0·2 m Tris–HCl pH 7·0, with 0·2 m NaCl, 4 mm EDTA, 10% glycerol, 1% Nonidet P-40, and a mixture of protease inhibitors. Thereafter, cells and cell debris, respectively, were separated from supernatants by centrifugation.

The human monocytic leukaemia cell line THP-1 was obtained from the American Type Culture Collection (Manassas, VA) and cultured in complete medium [RPMI-1640 medium supplemented with 10% fetal bovine serum (FBS), 2 mm glutamine, 1 mm pyruvate, 100 U/ml penicillin and 100 μg/ml streptomycin]. Before each assay, cells (3 × 105 to 4 × 105/ml) were treated for 72 hr with 1,25-dihydroxyvitamin D3 (0·05 μm) or left untreated. Cells were counted after trypan blue staining in a Neubauer chamber.

Bacteria and PGN binding assay

Fresh cultured S. aureus, E. faecalis, E. coli and B. abortus were resuspended in 1 ml of PBS with 10 μg of PGRP or lysozyme. After overnight incubation at 4°, the bacterial suspensions were centrifuged, washed and resuspended in PBS. Bacteria incubated in the absence of PGRP were used as control. Insoluble PGN binding assay was performed as described by Yoshida et al.,31 incubating 0·1 mg of PGN in PBS with either 10 μg of PGRP, lysozymes or with human samples (e.g. serum, PMN cell lysates, supernatant of induced cells), overnight at 4°. Supernatant and pellets were subject to SDS–PAGE and immunoblotting analysis.

Flow cytometry and fluorescence microscopy

Binding of PGRP to PBMC and THP-1 cells was analysed by FACS/flow cytometry. Cells (2 × 106) were resuspended in PBS with 10% human serum overnight at 4°. After that, cells were incubated for 30 min with PGRP-S, PGRP-IαC, PGRP-IβN or BSA as negative control (10 μg). The presence of PGRP in the cell surface was analysed with rabbit anti-PGRP serum and FITC-conjugated anti-rabbit antibodies. Alternatively, cells were incubated for 1 hr with 10 μg of PGRP-S-FITC, PGRP-IαC-FITC or PGRP-IβN-FITC at 4° and 37°. To determine the mechanism involved in PGRP uptake, wortmannin was added at a final concentration of 100 nm and 50 μm, respectively, 15 min before PGRP-FITC. A control with DMSO was performed. Twenty thousand events per sample were collected on a Partec PAS III flow cytometer and analysed with Winmdi software (The Scripps Institute, Flow Cytometry Core Facility, La Jolla, CA). Image capture was performed using a fluorescence microscope (Olympus BX-51, Tokyo, Japan).

To determine the effect of PGN and PGRP on cell activation, THP-1 cells treated with PGN or with PGN–PGRP complexes were incubated with the following monoclonal antibodies: anti-human CD14-phycoerythrin-Cy7, CD11-phycoerythrin-Cy5, CD86-FITC and CD80-FITC. For double fluorescence experiments, the flow cytometer was appropriately compensated with FloMAX software (Informer Technologies Inc, Dominica).

Cell proliferation assays, cytokine determinations and nuclear factor-κB expression

Cell proliferation was assessed in PBMC or THP-1 cells (2 × 105/well, in 1 ml of complete medium) cultured in the presence of PGN (10 μg/ml), PGRP (10 μg/ml), PGN–PGRP or BSA (control) for 24–72 hr. Biological activity as indicator of cellular proliferation was determined with the tetrazolium assay (MTT). Cytokines interleukin-8 (IL-8), IL-12, IL-10 and tumour necrosis factor-α (TNF-α) were measured by ELISA from supernatant (SN) (R & D Systems, Oxon, UK). Each experiment was repeated at least three times.

For determination of nuclear factor-κB (NF-κB) expression level, cells were cultured for 90 min. The nuclear protein extractions were obtained with 10 mm HEPES buffer in the presence of protease inhibitors (aprotinin, pepstatin, leupeptin, PMSF, EDTA, EGTA and dithiothreitol) and Nonidet P-40, and quantified using the Bradford assay. All extracts were subjected to immunoblotting analysis for NF-κB determination employing a mouse anti-NF-κB (1 : 2000) (Santa Cruz Biotechnology, Dallas, TX) and a goat anti-mouse IgG secondary antibody conjugated to peroxidase (1 : 5000) and an enhanced chemiluminescence reagent. Images were acquired using a Kodak image station 2000R (Eastman Kodak, Rochester, NY). Histone H8 was used as control of protein expression. Image J (National Institutes of Health, Bethesda, MD) was employed for relative protein expression quantification.

Antibacterial assays

Bactericidal and bacteriostatic activity of human PGRP was assayed on logarithmically growing bacteria (S. aureus or B. abortus) at 108/ml during at least 3 hr as previously described,32 with BSA as a control protein or PGRP (10 μg/ml of PGRP-S, PGRP-Iα or PGRP-Iβ; ∽3 × 108 molecules of PGRP/bacterial cell). The standard medium was RPMI-1640 with FBS. Control without PGRP was carried out similarly. The number of live bacteria was determined by colony counts on LB agar plates at 1-hr intervals.

Infection studies

Staphylococcus aureus was pre-incubated for 1 hr at 37° in RPMI medium alone as control or in the presence of 10 μg/ml PGRP-S, PGRP-Iα or PGRP-Iβ previously treated with polymyxin. THP-1 cells (106/ml) were cultured in RPMI-1640 with 10% FBS, pyruvate and glutamine during 24 hr at 37°. Subsequently, THP-1 cells were co-cultured for 1 hr with S. aureus previously treated or not, at a multiplicity of infection of 100 (bacteria/monocytes), in a final volume of 0·2 ml of RPMI. After that, THP-1 were washed and resuspended in RPMI with gentamicin to eliminate residual bacteria present in the medium. After different end-points (12, 24, 48 and 72 hr) infected cells were washed with PBS to remove antibiotics and lysed with 2% Triton ×100. Lysates were diluted with PBS and plated on LB agar. Colony-forming unit (CFU) determination of intracellular bacteria was determined by the counting plate technique.

Phagocytosis and endocytosis assays

THP-1 cells (106/well) were cultivated in the presence of 108 CFU/ml of dead S. aureus-FITC or 20 μg/ml of PGN-FITC previously incubated with PGRP-S, PGRP-Iα, PGRP-Iβ or alone as control. All assays were performed in RPMI medium with pyruvate and glutamine in the presence of FBS. After 1–3 hr at 37°, cells were washed and resuspended in PBS-trypan blue to determine phagocytosis of FITC-bacteria or endocytosis of PGN-FITC by flow cytometry. We analysed the percentage of positive cells presented in the M1 region of cells with PGRP-S. aureus-FITC/cells with S. aureus-FITC.

Apoptosis assays

For apoptosis determination, THP-1 cells were cultured (2 × 105/ml) in the presence of PGN (10 μg/ml), PGRP (10 μg/ml), PGN–PGRP or BSA (control) for 24–72 hr. Analysis of early apoptosis was performed by flow cytometry using the Phycoerythrin Annexin V Apoptosis Detection Kit I (BD Pharmingen, San Diego, CA) following the manufacturer′s instructions. Each experiment was repeated at least three times. In addition, the LIVE/DEAD Kit (Invitrogen, Carlsbad CA) was employed for membrane integrity and metabolic activity determination.

Statistical analysis

Results were tested statistically using a one-way analysis of variance with Dunnett's multiple comparison test or the Student's t-test (Graph-Pad Prism; GraphPad Software, San Diego, CA). Results were determined to be statistically significant when a P value of < 0·05 was obtained.

Results

Recombinant PGRP are biologically active proteins

We cloned and expressed human PGRP-S, PGRP-IαC and PGRP-IβN as inclusion bodies. The refolding and purification process yielded ∽0·5 mg/l of culture. To determine the biological activity of the recombinant proteins we used a precipitation assay with insoluble S. aureus PGN. The precipitated PGRP were analysed by immunoblotting showing that both PGRP-S and PGRP-IαC exist in two functional forms, monomeric and dimeric, both capable of binding to PGN, whereas PGRP-IβN was detected only as a monomer (Fig.1a). In addition, we found that PGRP-S and PGRP-IαC also bind insoluble PGN purified from E. faecalis, E. coli and B. abortus in both monomeric and dimeric forms (see Supplementary material, Fig. S1). To confirm the capacity of recombinant PGRP to bind to natural PGN on bacteria, PGRP (10 μg/ml) were incubated with different species. All PGRP were able to strongly bind to Gram-positive S. aureus and E. faecalis, and to a lesser extent, to Gram-negative bacteria E. coli and B. abortus, as observed by immunoblotting using specific anti-PGRP antibodies (see Supplementary material, Fig. S1). This reactivity pattern correlates well with the external PGN exposure to these bacteria species. PGRP dimers are reduced to monomers when treated with dithiothreitol after detaching from insoluble PGN or bacteria (see Supplementary material, Fig. S1), demonstrating that dimerization is due to an intermolecular disulphide bridge.

Figure 1.

Figure 1

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Peptidoglycan recognition proteins (PGRP) binding to insoluble Staphylococcus aureus peptidoglycan (PGN). (a) Recombinant PGRP-S, PGRP-Iβ and PGRP-Iα (10 μg/ml) were incubated overnight at 4° with 100 μg of insoluble PGN in PBS. Sample pellets were removed, washed and subjected to SDS–PAGE and immunoblotting analysis using a mix of specific anti-PGRP-S, PGRP-Iα and PGRP-Iβ antibodies. (b) Human serum sample and PBS were precipitated with PGN and analysed by immunoblotting. A 45 000-MW band was observed with anti-PGRP-S antiserum. Recombinant PGRP-S was also analysed as control. (c) Supernatant and pellet of cultured polymorphonuclear cells (PMN) were analysed by immunoblotting. Lane 1, supernatant of PMN without fMLP induction; lanes 2, 3, supernatant of PMN induced with fMLP at 15 and 45 min; lane 4, supernatant of induced PMN precipitated with PGN; lane 5, pellet of uninduced PMN; lanes 6, 7, pellet of induced PMN at 15 and 45 min; lane 8, supernatant of uninduced PMN precipitated with PGN; and lane 9, recombinant PGRP-S. MWM: molecular weight markers.

PGRP-S is present in human sera and PMN cells

To analyse whether native PGRP also exist as a dimer in fluids, human sera were precipitated with insoluble PGN and analysed by immunoblotting. A 45 000-MW band was detected with rabbit anti-PGRP-S-specific antibodies (Fig.1b). No PGRP-S monomer was observed in human sera; however, the treatment with dithiothreitol produced the corresponding monomer. Similar results were obtained when sera were immunoprecipitated with anti-PGRP-S-specific antibodies and protein-A–agarose (results not shown). We estimated PGRP-S concentration in serum to be ∽5 μg/ml. No PGRP-Iα or PGRP-Iβ could be detected in human sera.

When purified human PMN cells were lysed and analysed by immunoblotting employing specific anti-PGRP-S sera, a 45 000-MW band could be detected (Fig.1c). As the presence of PGRP-S in exocytic granules had been previously reported,33 we induced PMN cell degranulation with fMLP. By immunoblotting we found that the degranulated cells did not conserve PGRP-S; however, it was found in culture SN as a 45 000 MW band corresponding to a PGRP-S dimer. Precipitation with PGN of SN from degranulated PMN cells yielded a similar band at 45 000 MW (Fig.1c). We were not able to detect any PGRP-Iα or PGRP-Iβ in human PMN or SN.

PGRP are recognized and internalized by monocytes

With the aim of determining whether PGRP would be recognized by receptors on immune system cells, we analysed the binding of PGRP to PBMC by flow cytometry using specific rabbit antibodies for each PGRP and anti-rabbit FITC-conjugate as a second antibody. We observed that all assayed PGRP bound to PBMC, with PGRP-S and PGRP-Iα having the highest number of positive cells and showing an increase of 12–17-fold with respect to control cells not treated with PGRP but incubated with specific antibodies (Fig.2a–c). When we analysed the number of positive cells gated by forward/size cell characteristics, we observed that all PGRP preferentially bound to monocytes rather than to lymphocytes (data not shown). To verify monocyte–PGRP interaction, we conducted the experiment with the human monocytic leukaemia cell line THP-1 with similar results. PGRP-S, PGRP-Iα and PGRP-Iβ bound to THP-1 cells in a comparable mode, between two and four times with respect to control cells not treated with PGRP but incubated with specific antibodies (Fig.2d–f). In addition, we used FITC-labelled PGRP to circumvent any interaction of the IgG from the anti-PGRP serum and the THP-1 cells and avoid false-positive results. Hence, 1,25-dihydroxyvitamin D3-treated THP-1 cells were incubated with PGRP-FITC at 4° and 37° to inhibit and allow phagocytosis, respectively. At 4° we observed an increase in the number of positive cells in the M1 region for all PGRP with an increase of twofold to threefold (100–200%) with respect to control cells not treated with PGRP-FITC, indicating that the binding to the monocyte membrane is specific (Fig.2g). To test whether the PGRP are internalized after binding to the cell membrane we analysed the results of the assay performed at 37° by flow cytometry in the presence of trypan blue, for quenching of any extracellular fluorescence. We found a twofold to threefold increase in the FITC fluorescence signal of cells treated with PGRP-FITC with respect to non-treated controls (100–200%), suggesting that internalization was effectively occurring after binding (Fig.2h). Immunoblotting assays of THP-1 cells incubated with PGRP using rabbit anti-PGRP as a first antibody confirmed that PGRP bound to THP-1 monocytes (Fig.2i). We previously demonstrated that THP-1 cells lack endogenous PGRP (results not shown). PGRP-S-FITC internalization by THP-1 cells was also observed by fluorescence microscopy (Fig.2j). The presence of phosphoinositide 3-kinase inhibitor wortmannin in culture did not reduce the incorporation of PGRP by THP-1 (result not shown). These results suggest that macropinocytosis would not play a major role in PGRP uptake by monocytes. To study if PGRP would act as a link between PGN recognition and intracellular signalling, we analysed NF-κB expression in THP-1 cells. Figure2(k) shows that PGRP increase the level of NF-κB expression with respect to controls and, similarly, the PGRP-PGN complex increases the level of NF-κB expression with respect to control cells treated with PGN.

Figure 2.

Figure 2

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Peptidoglycan recognition proteins (PGRP) binding to peripheral blood mononuclear cells (PBMC) and monocytes/macrophages. PBMC and THP-1 cells, previously incubated with serum to block Fc receptors, were cultured in the presence of PGRPs or BSA as negative control (10 μg) and then analysed by flow cytometry or immunoblotting. Binding of PGRP to PBMC (a–c) and THP-1 cells (d–f) determined by FACScan. The presence of PGRP was analysed with rabbit anti-PGRP sera and anti-rabbit-FITC conjugate antibodies. Flow cytometry results were analysed as the % of positive cell population presented in the M1 region of cells treated with PGRP/untreated cells. Binding of PGRP-S-FITC, PGRP-Iα-FITC and PGRP-Iβ-FITC to THP-1 cells at 4° (g) and 37° (h) determined by flow cytometry. Insets: Representative cytometry histogram. **P < 0·01, ***P < 0·001. (i) Immunoblotting of THP-1 cells incubated with PGRP at 4°. (j) Image capture of cells incubated with PGRP-S-FITC in the presence of DiD and DAPI staining. (k) Determination of nuclear factor-κB (NF-κB) expression by immunoblotting. Lane 1, PGN-PGRP-S; lane 2, PGN; lane 3, PGRP-S; lane 4, untreated cells. Figures show a representative three or four experiments.

Recombinant PGRP have no bactericidal or bacteriostatic activity

To determine whether human PGRP have bacteriostatic or bactericidal activity, S. aureus, E. faecalis, E. coli and B. abortus (108 CFU/ml) were cultivated in RPMI with FBS during 1–3 hr in the presence of PGRP (10 μg/ml). Control without PGRP was performed similarly. Bacterial CFU/ml were analysed by the counting plate method at a 1-hr interval, observing similar results in the presence or absence of PGRP (Fig.3a). No CFU differences were observed with culture of 24–48 hr incubation with PGRP (result not shown). We next analysed whether the PGRP would cause membrane damage to bacteria. Using the Sytox-Green assay labelling method, which is a membrane-non-permeable dye that binds to DNA, we observed that PGRP-S only produced low levels of membrane damage after 1 hr of exposure, and that PGRP-Iα began to injure the membrane after 2 hr of incubation (data not shown). Nevertheless, a strong increase in Sytox-Green incorporation, determined as the number of positive cells present in the M1 region for assay/number of positive cells in the M1 region for control was detected after 3 hr of culture for both PGRP-S and PGRP-Iα compared with controls (Fig.3b). No membrane damage was observed with PGRP-Iβ in all the conditions assayed.

Figure 3.

Figure 3

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Peptidoglycan recognition proteins (PGRP) effect on Staphylococcus aureus and incorporation by monocytes/macrophages. (a) S. aureus (108/ml) were cultured in the presence or absence of PGRP (10 μg/ml) and colony-forming units (CFU)/ml were determined for bactericidal and/or bacteriostatic activity. (b) Incorporation of membrane-non-permeable marker Sytox-green by S. aureus cultivated after 3 hr in the presence or absence of PGRP and analysed by flow cytometry. We analysed the increase of Sytox-green incorporation as the % of bacteria treated with PGRP present in M1 region/% of bacteria not treated with PGRP present in the M1 region. Inset: cytometry histogram. (c) Phagocytosis assay of S. aureus-FITC by THP-1 in the presence or absence of PGRP. We analysed by flow cytometry the % of positive cell population presented in the M1 region of cells culture with PGRP-S. aureus-FITC/cells cultured with S. aureus-FITC (control). Insets: cytometry histogram. (d) THP-1 infection studies of S. aureus pre-incubated with PGRP-S for 48 hr. Figures show a representative of three experiments. **P < 0·01, ***P < 0·001.

PGRP enhance phagocytic activity and bacteria elimination

With the aim of studying whether PGRP modify bacterial phagocytosis by monocytes, 106 THP-1 cells were cultivated with 108 CFU of killed FITC-labelled S. aureus previously treated or not with PGRP (10 μg/ml). Flow cytometry analysis showed that the ability of monocytes to phagocytose S. aureus in the presence of any of the PGRP analysed is increased by ∽130% (Fig.3c). To determine the effect of PGRP facilitating phagocytosis by THP-1 on infecting bacteria, we used a model of monocyte infection by live S. aureus, which were pre-incubated 1 hr at 37° in the presence of 10 μg/ml PGRP-S or in medium alone as control. Then, THP-1 cells were co-cultured for 1 hr with PGRP-S-treated S. aureus at a multiplicity of infection of 100 (bacteria/monocytes). Infected THP-1 were washed and suspended in RPMI with gentamicin to eliminate residual bacteria present in the medium or bound to the membrane. After different incubation times, infected cells were washed with PBS to remove antibiotic and lysed with 2% Triton-X-100 in PBS. Lysates were diluted with PBS and plated on LB agar. CFU determination of viable intracellular bacteria was determined by the counting plate technique. Figure3(d) shows that at 12 hr the number of live bacteria incorporated in the presence of PGRP-S is 10 times higher than in the absence of PGRP-S, which is in agreement with the incorporation of dead bacteria in the previous assay. At 24 hr the THP-1 cells could only reduce the number of live bacteria in a small proportion, irrespective of the presence or absence of PGRP-S. However, at 48 hr the THP-1 cells that incorporated bacteria with PGRP-S were able to kill as much as 10 times more bacteria than cells that incorporated bacteria without PGRP-S.

Protector function of PGRP

When we used the LIVE/DEAD Cell Vitality Assay Kit to determine membrane integrity and THP-1 cell viability we observed that PGRP decreased by ∽60% the incorporation of Sytox Green by THP-1 cells compared with non-treated cells (Fig.4a–d). Similarly, PGRP were able to protect membrane damage produced by PGN (Fig.4e–h) decreasing Sytox Green incorporation by ∽40%. Moreover, PGRP or PGN–PGRP increased by 10–46% the metabolic activity of THP-1 cells with respect to controls determined as resazurin incorporation (Fig.4). To verify these results we used the Annexin/7AAD Kit to determine PGRP influence in apoptotic phenomena. We observed that PGRP were able to protect cells from apoptosis induced by PGN with a reduction of ∽40% compared with non-treated cells (Fig.4m–p). In addition, PGRP not only diminished the apoptotic process produced by PGN but they also diminished the apoptotic processes in cells cultured without PGN by 42–58% (Fig.4i–l).

Figure 4.

Figure 4

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Cell membrane damage and apoptosis. Monocytes were incubated with peptidoglycan (PGN) or PGN–peptidoglycan recognition protein (PGRP) complex, stained with Sytox-Green/resazurin and finally analysed by flow cytometry. Live/dead analysis of THP-1 cells cultures without (a) or with (b–d) PGRP. Likewise, cultures of THP-1 cells with PGN (e) or with PGN pre-incubated with PGRP (f–h). Apoptosis analysis of THP-1 cell cultures without (i) or with PGRP (j–l). Apoptosis was assessed in THP-1 cell cultures with PGN (m) or with PGN pre-incubated with PGRP (n–p). All cultures were treated with Annexin V/7AAD and then analysed by flow cytometry.

PGRP increase PGN-induced monocyte activation and proinflammatory cytokine secretion

To determine the effect of PGRP on monocyte proliferation, THP-1 cells not treated with 1,25-dihydroxyvitamin D3, were incubated for 24–72 hr with 20 μg/ml PGN previously treated or not with 10 μg/ml PGRP or BSA as a control. PGN produced a 50% inhibition in THP-1 proliferation, whereas the PGN pre-treatment with all the PGRP analysed, significantly reduced the inhibition by PGN (Fig.5a). To analyse the effect of PGRP in cell activation, THP-1 monocytes were incubated with PGN treated or not with PGRP during 24–72 hr and membrane expression of CD11, CD80, CD86 and CD14 was analysed by flow cytometry. PGRP increased CD11 expression by 50–150% (Fig.5b); however, the complex PGN–PGRP did not (result not shown). An increased expression of CD80 (among 100–200%) and CD86 (∽50%) for PGRP-S, PGRP-Iα and PGRP-Iβ at 48 hr was observed compared with control cells (Fig.5c,e). Similarly, the PGN–PGRP complex increased the expression of CD80 (> 50%) and CD86 (40–100%) produced by cells cultivated with PGN only (Fig.5d,f). When CD14 was determined, we observed that PGRP increased by ∽100% the expression of this molecule (Fig.5g) whereas PGN–PGRP complexes increased by ∽50% CD14 expression compared with controls of PGN-treated cells (Fig.5h).

Figure 5.

Figure 5

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Effect of peptidoglycan recognition proteins (PGRP) over THP-1 cells. To determine the effect of PGRP over THP-1 proliferation, cells were incubated with PGN–PGRP complexes (a) during 48 hr and then biological activity was determined with the tetrazolium assay (MTT). For cell activation analysis, THP-1 cells were treated with PGN, PGRP or with PGN–PGRP and then incubated with anti-human CD80, CD86, CD11 and CD14 antibodies. Flow cytometry results were analysed as the % of positive cell population presented in the M1 region of cells treated with PGRP or PGN-PGRP/control cells (untreated or treated with PGN, respectively). (b) CD11 expression on THP-1 cell membrane. Insets: cytometry histogram. CD80 (c) and CD86 (e) expression on THP-1 cell membrane treated with PGRP. Insets: cytometry histogram. Expression of CD80 (d) and CD86 (f) produced by PGN-PGRP. Insets: cytometry histogram. (g) Analysis of CD14 expression produced by PGRP. Insets: cytometry histogram. (h) Expression of CD14 produced by PGN–PGRP. Insets: cytometry histogram. Figures show a representative of three or four experiments. *P < 0·05, **P < 0·01, ***P < 0·001.

Culture SN from PGN-treated THP-1 cells in the presence and absence of PGRP was analysed to determine cytokine secretion. We found that PGN increased the production of IL-8, IL-12, TNF-α and IL-10 by THP-1 compared with control cells, but PGRP by themselves did not (Fig.6a–d). Surprisingly, an enhanced response of IL-8 (100–150%), IL-12 (50–150%) and TNF-α (50–100%) was detected when the three PGRP were added to PGN. All differences have statistical significance except for TNF-α when PGRP-Iα and PGRP-Iβ were used; however, these proteins increased TNF-α level consistently in all assay repetitions. We also observed that PGN–PGRP-S complexes increased IL-1 and IL-6 levels (data not shown). Conversely, the addition of the three PGRP analysed produced a significant 30–45% reduction of IL-10 compared with PGN alone (Fig.6d).

Figure 6.

Figure 6

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Cytokine secretion. To determine the effect of peptidoglycan recognition proteins (PGRP) over cytokine production, SN of monocytes/macrophages cultured with peptidoglycan (PGN), PGRP and PGN–PGRP complexes during 48 hr were tested by ELISA for interleukin-8 (IL-8) (a), IL-12 (b), tumour necrosis factor-α (TNF-α) (c) and IL-10 (d) production. Figures show a representative of three or four experiments. *P < 0·05, **P < 0·01, ***P < 0·001.

Discussion

We produced, refolded and purified human recombinant PGRP-S, the C-terminal domain of PGRP-Iα and the N-terminal domain of PGRP-Iβ, all proteins with PGN binding capacity. PGRP-S and PGRP-IαC were expressed as monomeric and dimeric forms of ∽20 and ∽40 kDa, respectively. Treatment of the dimers with dithiothreitol yields the monomers, suggesting that the 40 000-MW bands are disulphide dimers of PGRP. These results are in agreement with those obtained by Lu et al.15 describing that recombinant PGRP-S is secreted as a homodimer, whereas recombinant PGRP-Iα is secreted as a homodimer or as a heterodimer with PGRP-Iβ, when co-expressed. The existence of dimers is consistent with the free Cys present in the recombinant PGRP. PGRP-S have seven Cys but only Cys8 is free in the crystal structure and it is probably responsible for the dimerization of the recombinant as well as the natural PGRP-S in serum because it is exposed to the solvent in the molecule surface.21 PGRP-IαC amino acid sequence shows the presence of five Cys but only Cys300 is free.13 Conversely, PGRP-IβN is only expressed as a monomer in spite of its five Cys. It is possible that the free Cys was not on the molecule surface; however, no structure is known to date. A second affinity constant for PGRP–PGN interaction was determined, suggesting a cooperative binding effect that would be caused by PGRP aggregation or binding of multiple PGRP molecules to each PGN molecule.26,27 In this sense, PGRP dimerization could have a physiological role in PGN recognition.

Until now, PGRP-L was mentioned as the only serum PGRP, produced in the liver and secreted to blood.34 This manuscript is the first to report the presence of PGRP-S in normal human serum samples as a 45 000-MW dimer protein, and that they bind to insoluble PGN and are recognized by specific anti-PGRP-S polyclonal mouse and rabbit antibodies without cross-reaction with other PGRP or lysozymes. The difference in electrophoretic mobility of natural and recombinant PGRP-S (38 000 MW) could be attributed to glycosylation, since PGRP-S has one potential N-glycosylation and one potential O-glycosylation site, calculated with NetNGlyc 1.0 and NetOGlyc 4.0 (Technical University of Denmark, Lyngby, Denmark), respectively. Hence, a 45 000-MW band would be expected for a glycosylated dimer of PGRP-S in the native condition. Therefore, results presented here and previous findings by others15 indicate that human PGRP-S is a serum dimeric protein.

PGRP-S was first described as a protein present in PMN cells and in the bone marrow.1 The presence of PGRP-S in the bone marrow could be a result of the generation and maturation of neutrophils before reaching circulation. We also detected the presence of PGRP-S in human PMN cells when induced with fMLP and the PGRP-S was secreted into the extracellular medium by degranulation. Neutrophil degranulation is a physiological mechanism employed to attack microorganisms but it could also take place spontaneously. PGRP-S detected in neutrophil granules has the same molecular weight (45 000 MW) as that found in serum, suggesting that PGRP-S is secreted from PMN cells as a dimer and that these cells could be responsible for the PGRP-S present in the serum.1,35 We detected PGRP-S at low concentrations, in accordance with PGRP concentration in mouse serum that was described as lower than 5 μg/ml.1 We also detected PGRP-S, PGRP-Iα and PGRP-Iβ in human saliva (results not shown) probably secreted from granulocytes, salivary glands or tongue epithelial cells.15 The presence of PGRP-S in serum could be justified just by the recognition of bacterial PGN. However, the absence of a bacteriolytic effect1,15 and a reduced or absent bactericidal or bacteriostatic activity, as discussed below, suggest an unknown after binding function of PGRP, if any.

We tested whether PGRP-S–bacteria and PGRP–PGN complexes would be taken up by blood cells. We found that PGRP can bind to the membrane of PBMC; however, the analysis of these heterogeneous populations separately showed that the binding was predominantly to monocytes rather than to lymphocytes. To confirm these observations, we determined that all PGRP analysed can bind to the THP-1 cell line of monocytes/macrophages. The lower binding of PGRP to THP-1 cells compared with PBMC could be the result of a higher number of receptors in the heterogeneous population of peripheral blood monocytes rather than in the homogeneous THP-1 cell line. Using FITC-labelled PGRP we demonstrated that PGRP could bind to THP-1 cell membrane and are endocytosed by them. Wortmannin does not inhibit PGRP incorporation, suggesting that macropinocytosis is not mainly responsible for PGRP internalization. We also found that PGRP increase NF-κB activation, suggesting that a cellular receptor for PGRP would exist to sense the presence of bacteria or PGN fragments. However, several attempts to cross-link and co-precipitate the putative receptor were unsuccessful (results not shown).

Several authors have defined PGRP as bacteriostatic or bactericidal agents.1,33,35 It was reported that they need Ca2+ and/or Zn2+ for bactericidal action or that they lose this activity when they are not glycosylated. In addition, it has been reported that they are bactericidal for different Gram-positive pathogenic (Listeria monocytogenes and S. aureus) or non-pathogenic (Bacillus and Lactobacillus) bacteria; however, they do not affect normal flora.15 Recently, Bosco-Drayon et al.36 described the importance of PGRP-LE and PGRP-LB in a balanced response to bacterial infection and tolerance to normal flora. However, the reasons for and fundamentals of these actions are not known. In this regard, the amount of PGRP needed to conserve bactericidal function is controversial. Some authors employed 45–200 μg/ml of protein,1,15,32 which is far from physiological levels. PGRP bactericidal and bacteriostatic activities described by Wang et al.32 were done in TRIS buffer, which is not adequate for bacterial growth. On the contrary, we conducted our experiments in growth medium, using a PGRP-S concentration of 10 μg/ml, the maximum level of PGRP-S detected in serum. In these conditions we did not detect bacteriostatic or bactericidal activity of human PGRP. These results demonstrate that, in contrast to antibacterial peptides,37 human PGRP do not kill bacteria by permeabilizing their cytoplasmic membranes or by disrupting their cell wall. However, PGRP-S and PGRP-Iα produce slight damage in the S. aureus cell wall, as treated bacteria incorporated Sytox-green.

We next studied the role of PGRP in the presence of infecting bacteria, which is a controversial issue. Different authors have suggested that PGRP could inhibit phagocytosis1 or they do not increase bacteria uptake by PMN cells.33 However, others reported that different Drosophila PGRP isoforms (PGRP-SC1, PGRP-SA and PGRP-LC) could be involved in phagocytosis.7,11 We found that all human PGRP analysed were able to increase the monocyte/macrophage phagocytic activity for dead S. aureus-FITC, or infective S. aureus. In addition, the THP-1 cells were able to kill 10 times more bacteria than cells that incorporated S. aureus without PGRP-S. In this sense, Kashyap et al.38 recently showed that human PGRP are able to kill bacteria by a combination of mechanisms including induction of reactive oxygen species, depletion of thiols and by increasing intracellular concentration of metals.

It was previously reported that PGRP-S forms a potent cytotoxic complex with heat-shock protein 70, which induces apoptotic death in various tumour lines.21,39–41 We observed that PGRP are able to protect membrane damage produced by PGN in normal cells. Similarly, PGRP protect cells from apoptosis induced by PGN or bacterial infection, and they diminish the cell apoptotic process in the absence of PGN, suggesting that PGRP could also be considered an anti-apoptotic protein.

We also analysed the effect of PGRP on the activation of THP-1 cells and cytokine secretion. We found that PGRP are able to revert the PGN inhibition of monocyte proliferation. The proliferative effect is associated with cell activation in the presence of PGRP, shown by an increased expression of CD11, CD80, CD86 and CD14 molecules on the THP-1 cell membrane. This evidence suggests a pro-inflammatory role of PGRP. The inflammatory reaction is increased in the presence of PGRP–PGN complexes compared with PGN alone, which was demonstrated by an increased expression of CD80, CD86 and CD14 molecules on the THP-1 cell membrane. Enhanced CD80 and CD86 molecules would increase co-stimulatory signal for antigen presentation. In addition, the increased expression of CD14, a molecule involved in PGN recognition, is highly significant in the context of a bacterial infection. These results are in agreement with the cytokine analysis in supernatant from PGN-treated THP-1 in the presence or absence of PGRP. Hence, PGN by itself triggers the secretion of pro-inflammatory IL-8, IL-12 and TNF-α by monocytes/macrophages, which experienced 1·5-fold to 3-fold increase in the presence of PGRP. PGN also increases the secretion of anti-inflammatory IL-10, perhaps as a homeostatic response of the cells to inflammation. However, the presence of PGRP reduces IL-10 secretion induced by PGN, strengthening the pro-inflammatory role of PGRP.

It is important to note that PGRP-Iα and PGRP-Iβ are composed of two domains with PGN recognition capacity. We worked with one of these domains. Further studies are needed to determine whether full-length proteins have a similar effect on PGN recognition by THP-1 cells.

We conclude that human PGRP-S is a natural dimer expressed in PMN cells, which is secreted by PMN degranulation, and that their presence can be detected in serum and saliva. PGRP-Iα and PGRP-Iβ are known to be natural homo or heterodimers secreted by epithelial cells and can be found in saliva but not in serum. We hypothesized that soluble PGRP-S in serum would bind to bacteria by its PGN, so facilitating uptake and killing by macrophages. Figure7 shows a scheme summarizing the main findings of this work, including the effects induced by PGN on monocytes/macrophages and the role of PGRP driving the innate response to increased inflammation. Thus, PGRP–PGN complexes would interact with an unknown putative receptor blocking the inhibitory effects of PGN over monocyte proliferation. PGN–PGRP complexes increase activation markers such as CD80/86, and innate immune receptor CD14, which will improve the immune response to attack pathogens. The complexes also increase pro-inflammatory IL-8, IL-12, TNF-α and decrease anti-inflammatory IL-10. This inflammatory reaction will in turn attract PMN cells, which would increase the local concentration of PGRP-S, producing higher bacteria uptake and clearance. The PGN–PGRP complexes would also rescue cells from PGN-induced apoptosis and would increase immune cell survival.

Figure 7.

Figure 7

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Representation of innate immune response induced by peptidoglycan (PGN) (a) and how this response is modified in the presence of peptidoglycan recognition proteins (PGRP) (b).

Acknowledgments

This work was supported by the University of Buenos Aires, the National Research Council of Argentina (CONICET), and Agencia Nacional de Promoción Científica y Técnica, grants PICT-2010-373 (to MMF), PICT-2007-0721 (to MCDM) and PICT-2008-1139 and PICT-2010-0657 (to ELM). ELM is also supported by the Fogarty International Center (TW007972) and International Centre for Genetic Engineering and Biotechnology (CRP/ARG09-02). RAM is supported by National Institutes of Health grant AI47990. MCDM is supported by Universidad Nacional de Luján (CDD-CB 230-08). The funders had no role in study design and analysis, decision to publish, or preparation of the manuscript. The authors declare no competing financial interests.

Disclosures

The authors declare that they have no conflict of interests.

Supporting Information

Figure S1. Peptidoglycan recognition proteins binding to insoluble peptidoglycan and bacteria.

imm0145-0429-sd1.tif (1.8MB, tif)

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Associated Data

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Supplementary Materials

Figure S1. Peptidoglycan recognition proteins binding to insoluble peptidoglycan and bacteria.

imm0145-0429-sd1.tif (1.8MB, tif)

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