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Originally published online as doi:10.1189/jlb.0206086 on September 14, 2006

Published online before print September 14, 2006
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(Journal of Leukocyte Biology. 2007;81:893-903.)
© 2007 by Society for Leukocyte Biology

Pivotal Advance: Vasoactive intestinal peptide inhibits up-regulation of human monocyte TLR2 and TLR4 by LPS and differentiation of monocytes to macrophages

N. Foster, S. R. Lea, P. M. Preshaw and J. J. Taylor1

Oral Microbiology and Host Responses Group, Oral Biology, School of Dental Sciences, University of Newcastle upon Tyne, Newcastle upon Tyne, UK

1 Correspondence: Oral Microbiology and Host Responses Group, Oral Biology, School of Dental Sciences, University of Newcastle upon Tyne, Framlington Place, Newcastle upon Tyne NE2 4BW, UK. E-mail: j.j.taylor{at}ncl.ac.uk

ABSTRACT

Vasoactive intestinal peptide (VIP) is an immunoregulatory peptide, which inhibits LPS-induced cytokine secretion in myeloid cells and has beneficial effects in animal models of inflammatory diseases. We show for the first time that VIP decreases LPS-induced up-regulation of TLR2 and TLR4 by human monocytic THP1 cells and peripheral blood monocytes (PBM). VIP inhibited up-regulation of TLR4 expression in THP1 cells in response to LPS from Escherichia coli or the periodontal pathogen Porphyromonas gingivalis within 6 h poststimulation but had less of an effect on TLR2. After 24 h, P. gingivalis LPS-stimulated monocytic THP1 cells to differentiate into macrophages, which predominantly expressed TLR2, and E. coli LPS-stimulated THP1 differentiation to predominantly TLR4-expressing macrophages. VIP decreased monocyte differentiation to macrophages induced by LPS from either species and also reduced overall TLR2 and TLR4 expression in these cells. VIP had a similar effect on human PBM. The transcription factor PU.1 regulates TLR expression and has a central role in myeloid cell differentiation. VIP inhibited the nuclear translocation of PU.1 in LPS-stimulated THP-1 monocytes. VIP also inhibited the expression of the M-CSF receptor, which is regulated by PU.1. In summary, VIP inhibited LPS-induced differentiation of monocytes with a concomitant reduction in TLR2 and TLR4 expression. Although there was differential induction of TLR expression by LPS from P. gingivalis and E. coli, VIP inhibited the action of both of these LPS types on monocytes. The mechanism of action of VIP on monocyte differentiation may be via inhibition of the transcription factor PU.1.

Key Words: VIP • myeloid cells • Toll-like receptors • lipopolysaccharide

INTRODUCTION

Many studies have now reported on the immunomodulatory effect of vasoactive intestinal peptide (VIP) and its potential as a future therapeutic for a number of inflammatory diseases [1 ]. TLR2 and -4 are important receptors of pathogen-associated molecular patterns, and in humans, TLR2 [2 ] or TLR4 [3 ] may be activated by LPS; these receptors therefore play a pivotal and central role in proinflammatory immune responses. However, which TLR is activated depends on the LPS type, as gram-negative enteropathogens such as Salmonella minnesota or Escherichia coli predominantly activate TLR4 in human monocytes [4 ], whereas LPS from the gram-negative oral pathogen Porphyromonas gingivalis may stimulate TLR2 or TLR4 [5 ]. There have been no reports about the direct effect of VIP on TLR expression in human myeloid immune cells. However, Gutiérrez-Cañas et al. [6 ] have shown that VIP down-regulates TLR4 and TLR4-dependent cytokines in human synovial cells removed from patients with rheumatoid arthritis.

Recently, Gomariz et al. [7 ] have reported that VIP inhibits trinitrobenzene sulfonic acid (TNBS)-induced colitis in mice and that inhibition coincides with down-regulation of TLR2 and -4 on lymphocytes, macrophages, and dendritic cells. VIP decreases the mRNA of a number of inflammatory cytokines involved in TNBS-induced colitis in mice, and this coincides with a return to homeostasis in inflamed intestine [8 ]. Furthermore, Gomariz et al. [7 ] speculate that down-regulation of TLR expression may occur via inhibition of NF-{kappa}B, as previous studies had shown that VIP inhibits I{kappa}B degradation and prevents nuclear translocation of NF-{kappa}B in THP1 cells following stimulation with LPS from the gram-negative enteropathogen E. coli [9 ]. However, although a NF-{kappa}B-binding site has been detected on the promoter sequence of the murine TLR2 gene [10 ], a NF-{kappa}B site has not been found on the promoter sequence of the murine TLR4 gene, and in human cells, neither TLR2 nor TLR4 is regulated by NF-{kappa}B (reviewed in ref. [11 ]). Moreover, regulation of human TLR2 and TLR4 mainly occurs via the Ets family transcription factor PU.1 [12 ] and TLR4 via PU.1 and IFN-response factor [13 ]. The effect of VIP on TLR expression in human cells cannot therefore be extrapolated from mouse models.

P. gingivalis is commonly isolated from the inflamed periodontal pockets of patients suffering from periodontal disease and is implicated in the pathogenesis of this chronic inflammatory disorder [14 ]. A study by Linden et al. [15 ] has shown that VIP can be detected in gingival crevicular fluid in periodontal disease. This latter study is one of the few published to show that VIP is produced in response to inflammation in human tissues. Recently, we have shown that production of TNF-{alpha} by monocytic THP1 cells stimulated with LPS from P. gingivalis or E. coli is inhibited by coculture of the cells with VIP and that this inhibition coincides with inhibited nuclear translocation of NF-{kappa}B and c-jun [16 ]. However, the possibility that VIP may inhibit LPS-induced inflammatory pathways at the initial point of detection (the TLR) has not been studied in human cells. Furthermore, the comparison between the ability of different LPS types to stimulate TLR and the effect that VIP may have on these TLRs will be of critical importance in developing immunomodulatory approaches to different inflammatory disorders in man.

The aim of the work we report here was to investigate the effect of VIP on TLR2 and TLR4 expression and differentiation in human monocyes exposed to LPS from E. coli and P. gingivalis. Our findings indicate that although these two LPS variants have differential effects on TLR expression, VIP can universally inhibit LPS-induced differentiation of monocytes into macrophages and concomitant TLR up-regulation. The immunological effects of VIP are coincident with an inhibition of the myeloid transcription factor PU.1, which suggests a mechanism of action.

MATERIALS AND METHODS

Reagents
Unless otherwise stated, all laboratory reagents were purchased from Sigma (Poole, UK), and all antibodies were purchased from Serotec (Oxford, UK).

THP1 cell culture
Human promonocytic THP1 cells were purchased from the European Collection of Cell Cultures (Salisbury, Wilts, UK). Cells were cultured in RPMI media, supplemented with FCS (10% v/v), L-glutamine (2 mM), penicillin (100 U/ml), streptomycin (100 µg/ml), and amphotericin-B (2.5 µg/ml), and maintained at 37°C and 5% CO2. Prior to use, 5 x 106 THP1 cells per well were differentiated into monocytes in six-well tissue-culture plates (Greiner Bio-one Ltd., Stonehouse, UK) using 0.1 µM 1,25 dihydroxyvitamin D3 (Vit D3; Merck Biosciences, Nottingham, UK) for 24 h, as reported previously [17 ]. After 24 h, the ability of cells to adhere to the plastic culture dish (indicative of differentiation to monocytes) was assessed by microscopy. CD14 expression was also assessed periodically by FACS analysis (see below), as increased CD14 expression is also indicative of differentiation of promonocytic THP1 cells to monocytes [17 ]. During routine culture, cell viability was assessed by trypan blue exclusion and was always found to be >90%.

Isolation and culture of human PBMC and peripheral blood monocytes (PBM)
As Gomariz et al. [7 ] have shown that different murine leukocytes express different levels of TLR when mice are exposed to TNBS, we decided to use purified human buffy coats to examine the effect of VIP on LPS-induced TLR up-regulation in monocytes and lymphocytes. Venous blood (35 ml) was collected by venepuncture from healthy human volunteers into vacutainers containing EDTA. Blood volumes were diluted 2x in PBS prior to underlay with Ficol hypaque (density 1.077 g/ml). The samples were then centrifuged at 300 g for 20 min in a 3K10 centrifuge (Sigma). Buffy coats were removed and mixed with 2x PBS and centrifuged at 300 g for 15 min. To remove any erythrocyte contamination, cell pellets were then resuspended in erythrocyte lysis buffer (150 mM NH4Cl, 10 mM KHCO3, 0.1 mM EDTA, pH 7.3) for 3 min before quenching in 10 ml PBS and centrifugation at 300 g for 10 min. Platelet contamination was removed further by resuspending the cells in 50 ml platelet buffer (PBS containing 2 mM EDTA) and centrifugation at 300 g for 10 min.

After removal of the supernatant and resuspension of the cell pellet in platelet buffer, the process was repeated. PBMC within the buffy coat were counted prior to incubation in six-well plates (as stated above) at cell densities of 1 x 106 cells/ml. Cells were rested for 2 h in RPMI medium prior to stimulation with LPS or VIP. In experiments in which only purified monocytes were used, buffy coat cells were incubated in plastic culture dishes, as stated above, for 4 h. After 4 h, nonadherent cells (lymphocytes) were removed, and adherent cells were washed gently 2x with media prior to stimulation (as stated above).

Stimulation of THP1, PBMC, or PBM with LPS and VIP
All immunological parameters measured throughout this study were performed following stimulation of cells with LPS or LPS/VIP. Relevant controls were also measured in parallel. Ultrapure LPS from E. coli 0111:B4 (Invivogen, Calne, Wilts, UK) and LPS from P. gingivalis W5O (a gift from Dr. Minnie Rangarajan, Queen Mary’s School of Medicine and Dentistry, London, UK) were used to stimulate 1 x 106 cells/ml in six-well plates for 6 h or 24 h. In previous VIP titration experiments, we measured TNF-{alpha} in cell supernatants by ELISA and determined that 100 ng/ml LPS stimulated a significant increase in TNF-{alpha} production by THP1 cells, and such increase was optimally inhibited by a VIP concentration of 10–8 M [16 ]. These LPS/VIP concentrations were used in all subsequent experiments, and VIP was added simultaneously with LPS for 6–48 h. Cells treated with LPS/VIP were compared with unstimulated cells cultured for the same time period, and in some cases, cells stimulated with PMA (1 µg/ml) were used as positive controls.

Following experimental treatments, cells were stained with the fluorescent exclusion dye propidium iodide (PI; 20 µg/ml), and cell survival was analyzed by FACS (see below). PI uptake following treatment was compared with uptake in cells cultured for the same time period without LPS or VIP stimulation (unstimulated control) and cells that had been incubated in methanol for 10 min (positive control). In all cases, there were negligible differences between the numbers of dead cells following experimental treatment and unstimulated controls, and viability always remained above 80% following treatment. There was also no difference in the survival rates of cells cultured in LPS compared with those cultured in LPS and VIP.

Analysis of TLR surface expression on THP1, PBMC, and PBM
FACS analyses were used to measure TLR2, TLR4, and CD14 expression on the surface of cells following stimulation with P. gingivalis or E. coli LPS for 6 h or 24 h (THP1 cells) or for 6–48 h (PBMC and PBM) in the presence or absence of VIP. Results were compared with results obtained from parallel experiments, whereby surface expression of these markers was measured on cells cultured in media only for the same time periods (unstimulated cells). Appropriate isotype controls were also used (for details of antibodies, Table 1).


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  		Table 1. Antibodies Used in FACS Analyses

 
Briefly, LPS/VIP-stimulated and unstimulated control monocytic THP1 cells or human PBMC (1x106 cells per group) were centrifuged at 300 g for 10 min on a 3K10 centrifuge (Sigma). The cells were then washed three times by resuspension in PBS containing BSA (1% w/v) and EDTA (2 mM; FACS buffer) and centrifuged as previously stated. Cells were then incubated in FACS buffer containing human serum (10% v/v; Sigma) for 15 min at room temperature on a sample mixer (Dynal, Wirall, Cheshire, UK) to block FcR activity. After centrifugation, the cells were washed three times in ice-cold FACS buffer before incubating in CD14.PE for 45 min in darkness and on ice on a Rotatest end-to-end shaker (Luckham Ltd., Burgess Hill, West Sussex, UK). After incubation, the cells were washed 3x in ice-cold FACS buffer and incubated for 45 min with TLR2.FITC or TLR4.FITC, as stated previously. After incubation, the cells were washed 3x in ice-cold FACS buffer and resuspended in a final volume of 300 µl FACS buffer prior to analysis. In separate experiments, FACS analyses were performed to measure the M-CSF receptor (M-CSFR) on the surface of monocytic THP1 cells, which had been cocultured with LPS or LPS/VIP for up to 48 h. These experiments were conducted in exactly the same way as FACS analyses mentioned previously, replacing TLR or CD14 antibodies with the M-CSFR antibody at concentrations specified in Table 1 .

All analyses were performed on a FACScan analyzer (Becton Dickinson, San Jose, CA, USA). Cells (1x104) were analyzed in each treatment, and all treatments were compared with relevant isotype controls (Table 1 ). Samples were acquired using CellQuest pro software (Becton Dickinson) and analyzed using WinMDI 2.8 software.

Examination of nuclear translocation of PU.1 in human THP1 cells by confocal laser-scanning microscopy (CLSM)
Previous CLSM studies have shown that nuclear translocation of PU.1 can be observed 60 min after stimulation of murine macrophages (RAW cells) with LPS [18 ]. We therefore set out to determine, by CLSM analysis, whether PU.1 was translocated to the nucleus of monocytic THP1 cells following treatment with LPS for 60–90 min or inhibited from doing so by VIP. Following experimental treatments, adherent THP1 cells were washed 3x in PBS, fixed for 1 h in ice-cold (–20°C) methanol, and then permeabilised in 0.5% (v/v) Triton X-100 for 10 min at room temperature. After washing 3x in PBS, the cells were incubated for 60 min with mouse antihuman PU.1 (1 µg/ml; Autogen Bioclear, Calne, Wilts, UK). After 60 min, the cells were washed 3x in PBS containing 0.05% (v/v) Tween 20 (PBS-Tween). Vacant binding sites were blocked for 60 min using PBS-Tween and BSA (1% w/v). After washing, the cells were incubated with rabbit antimouse IgG.FITC (1/ 400) for 45 min prior to washing in PBS-Tween (3x) and incubating with 4'-6' diamidino-2 phenylindole (10 µg/ml; w/v). The cells were washed 3x and permanently mounted in fluorescent-mounting media (Dako, Carpinteria, CA, USA) prior to analysis with a TCS SP2 UV CLSM (Leica, Heidelburg, UK). All data were analyzed using Leica software, and verification of results was achieved using an independent, expert observer.

Morphological changes in THP1 cells exposed to LPS or LPS and VIP
Following culture of cells on coverslips in Vit D3 for 24 h, the cells were cultured with LPS, with or without VIP, as stated previously. After removal of coverslips, the cells were fixed and mounted prior to examination using the transmitted light facility of the TCS SP2 microscope.

Measurement of M-CSF by THP1 cells
The concentration of M-CSF in supernatants removed from THP1 cells, which had been cultured with LPS or LPS and VIP for 6 h and 24 h, was measured by ELISA using commercially available DuoSet kits in accordance with the manufacturer’s recommendations (R&D Systems, Abingdon, Oxford, UK). These results were compared with M-CSF concentration in supernatants obtained from unstimulated cells or cells stimulated with PMA for 6 h and 24 h.

Statistical analysis
An ANOVA test with a one-way classification was used to calculate significant differences (P<0.05) between test and control samples in ELISA analyses. Minitab software was used for all statistical analyses.

RESULTS

VIP inhibits LPS-induced TLR2 and TLR4 expression in THP-1 monocytes
Incubation of monocytic (Vit D3-treated) THP1 cells with E. coli LPS for 6 h increased the percentage of cells, which expressed TLR4 by 15–18% above TLR4 expression on the surface of unstimulated (control) cells (Fig. 1A ), and P. gingivalis LPS stimulated up-regulation of TLR4 on the THP1 cell surface by 6–10% (Fig. 1C) .


Figure 1
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  		Figure 1. FACS analysis of LPS-induced TLR expression by monocytic THP1 cells and the effect of VIP (6 h postculture). (A) TLR4 expression on the surface of THP1 cells incubated with E. coli 0111:B4 LPS (100 ng/ml) or LPS and VIP (10–8 M). (B) Measurement of TLR2 expression on THP1 cells incubated with E. coli 0111:B4 LPS (100 ng/ml) or LPS and VIP (10–8M). (C) TLR4 expression on the surface of THP1 cells incubated with P. gingivalis W50 LPS (100 ng/ml) or LPS and VIP (10–8 M). (D) Measurement of TLR2 expression on THP1 cells incubated with P. gingivalis W50 LPS (100 ng/ml) or LPS and VIP (10–8 M). Histograms are representative of results obtained on more than five separate occasions.

 
However, TLR2 up-regulation was not as pronounced when THP1 cells were stimulated with LPS for 6 h, and E. coli LPS only induced a 2–3% increase (Fig. 1B) , and P. gingivalis LPS induced increases of 3–6% (Fig. 1D) . The effect of VIP on TLR inhibition was, therefore, most prominent when measuring TLR4, following stimulation of THP1 cells with P. gingivalis LPS (Fig. 1C) and in particular, E. coli LPS (Fig. 1A) . In both cases, VIP reduced surface expression of TLR4 to levels similar to unstimulated (control) cells. Quantitative analyses of TLR expression on the surface of THP1 populations showed that after 6 h, VIP significantly inhibited P. gingivalis LPS-induced TLR4 (P<0.05) in comparison with the numbers of cells expressing TLR4 following culture with only P. gingivalis LPS, but VIP had a much greater effect on the number of THP1 cells expressing TLR4 following culture with E. coli LPS (P<0.05; see Fig. 4A ).


Figure 4
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  		Figure 4. Quantitative analyses of the effect of LPS or LPS and VIP on the number of TLR4- and TLR2-expressing cells in the THP1 cell population after 6 h or 24 h. CD14/TLR4 and CD14/TLR2 expression on the surface of THP1 cells incubated with E. coli 0111:B4 LPS (100 ng/ml; Ec), P. gingivalis W50 LPS (100 ng/ml; Pg), or LPS and VIP (10–8 M). (A) VIP significantly decreases the number of THP1 cells expressing P. gingivalis LPS-induced TLR4 and E. coli LPS-induced TLR4 after 6 h. (B) VIP does not significantly decrease the number of THP1 cells expressing P. gingivalis LPS-induced TLR2 after 6 h, but VIP does significantly reduce the number of THP1 cells expressing TLR2 following incubation with E. coli LPS. (C) VIP did not significantly decrease the number of THP1 cells expressing P. gingivalis LPS-induced TLR4 after 24 h, but the effect of VIP on E. coli LPS-induced TLR4 after 24 h was highly significant. (D) VIP decreased the number of THP1 cells expressing TLR2 when cocultured with P. gingivalis LPS for 24 h but did not significantly decrease the number of THP1 cells expressing TLR2 when cocultured with E. coli LPS for 24 h. Data obtained are means of experiments in which TLR was measured on 104 cells on five separate occasions. *, P < 0.05; **, P < 0.005. NS, Not significant.

 
VIP inhibits the LPS-induced differentiation of THP-1 monocytes into macrophages
When THP1 cells were stimulated with E. coli LPS for 24 h, a second cell population was generated in culture. FACS analyses showed that this second population was TLR4-dominant, representing >30% of the total CD14 population (Fig. 2B ), and a much lower number of cells in this second population expressed TLR2 (10–15% of total CD14 cells; Fig. 2E ). However, coculture of THP1 cells with E. coli LPS and VIP for 24 h dramatically reduced the expression of TLR4 (Fig. 2C) and TLR2 (Fig. 2F) to levels similar to those expressed by THP1 cells cultured for 24 h in media only (unstimulated controls; Fig. 2A and 2D ). Conversely, stimulation of THP1 cells with P. gingivalis LPS for 24 h induced a second cell population, which was TLR2-dominant, representing >25% of the total CD14 population (Fig. 3E ) compared with approximately half of the cell population (10–13%) expressing TLR4 (Fig. 3B) . However, when THP1 cells were cultured with P. gingivalis LPS and VIP, TLR4 expression was only reduced moderately to 8–11% (Fig. 3C) , whereas TLR2 expression was reduced by more than half (to 10–13%; Fig. 3F ). In both cases, VIP decreased TLR expression to levels similar to those measured on the surface of unstimulated controls (Fig. 3A and 3D) . Quantitative analyses of TLR expression on the surface of THP1 populations showed that after 24 h, VIP did not significantly inhibit P. gingivalis LPS-induced TLR4 populations in comparison with the numbers of cells expressing TLR4 following culture with only P. gingivalis LPS, but VIP significantly inhibited (P<0.005) the number of THP1 cells expressing TLR4 following culture with E. coli LPS (Fig. 4C ). Conversely, VIP significantly inhibited (P<0.005) the number of THP1 cells expressing TLR2 following stimulation of cells with P. gingivalis LPS for 24 h but did not significantly inhibit the number of THP1 cells expressing TLR2 following incubation of cells with E. coli LPS for 24 h (Fig. 4D) . These differences were a result of the fact that E. coli LPS induced THP1 populations, which were much more TLR4-dominant than TLR2-dominant, and P. gingivalis LPS had the opposite effect, stimulating TLR2-dominant THP1 cells after 24 h (Fig. 4C and 4D) . Therefore, when monocytic THP1 cells were stimulated with E. coli or P. gingivalis LPS for 24 h, a second adherent cell population was generated, which was TLR4- or TLR2-dominant, depending on the LPS type used to stimulate the cells.


Figure 2
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  		Figure 2. FACS analysis of E. coli 0111:B4 LPS-induced monocytic THP1 cell differentiation and TLR expression and the effect of VIP (24 h postculture). CD14/ TLR4 and CD14/TLR 2 expression on the surface of THP1 cells incubated with E. coli 0111:B4 LPS (100 ng/ml) or LPS and VIP (10–8 M). Arrows denote expansion of a differentiated population following E. coli LPS exposure. Coculture of THP1 cells with E. coli LPS and VIP inhibits the generation of this differentiated population. Increased TLR expression is predominantly a result of the differentiation of a CD14high/TLR4high population. Dot plots are representative of results obtained on more than five separate occasions.

 

Figure 3
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  		Figure 3. FACS analysis of P. gingivalis W50 LPS-induced monocytic THP1 cell differentiation and TLR expression and the effect of VIP (24 h postculture). CD14/ TLR4 and CD14/TLR 2 expression on the surface of THP1 cells incubated with P. gingivalis W50 LPS (100 ng/ml) or LPS and VIP (10–8 M). Arrows denote a differentiated population following P. gingivalis LPS exposure. Coculture of THP1 cells with P. gingivalis LPS and VIP inhibits the generation of this differentiated population. Increased TLR expression is predominantly a result of the differentiation of a CD14high/TLR2high population. Dot plots are representative of results obtained on more than five separate occasions.

 
To determine further whether this second cell population expressed macrophage characteristics, we compared our test samples with THP1 cells exposed to PMA (a promoter of THP1 cell differentiation to macrophages). After stimulating monocytic THP1 cells with PMA for 24 h, a second population was generated, which was TLR4high (Fig. 5A ), and morphological examination showed that a high proportion of these adherent cells had differentiated into macrophages (Fig. 5B) . We then compared the morphological appearance of these cells with that of unstimulated THP1 cells and THP1 cultured with LPS or LPS + VIP. E. coli LPS (Fig. 5C) and P. gingivalis LPS (Fig. 5D) induced macrophage morphology, and this was inhibited when the cells were cocultured with E. coli LPS + VIP (Fig. 5E) or P. gingivalis LPS + VIP (Fig. 5F) . In the presence of LPS + VIP, THP1 cells were more rounded and had a morphological appearance similar to unstimulated THP1 monocytes (Fig. 5G) .


Figure 5
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  		Figure 5. VIP inhibits morphological changes associated with LPS-induced differentiation of monocytic THP1 cells to macrophages (24 h postculture). (A) FACS analysis showing the generation of a CD14high/TLR4high cell population (arrowed) in culture following treatment of THP1 cells with PMA. (B) PMA induced THP1 cells to differentiate into macrophages (elongated macrophages arrowed). (C) E. coli 0111:B4 LPS (100 ng/ml) induced macrophage differentiation (elongated macrophages arrowed) in populations of Vit D3-treated THP1 cells. (D) P. gingivalis W50 LPS (100 ng/ml) induced Vit D3-treated THP1 cell populations to differentiate into macrophages (elongated macrophages arrowed). (E) Macrophage differentiation is prevented when Vit D3-cultured THP1 cells are cocultured with E. coli LPS and VIP (10–8M). (F) Macrophage differentiation is prevented when Vit D3-cultured THP1 cells are cocultured with P. gingivalis LPS and VIP (10–8M). (G) Rounded (control) THP1 cells treated only with Vit D3. All images are representative of results obtained on more than four separate occasions. Original scale bar = 10 µm

 
LPS-stimulated nuclear translocation of PU.1 is inhibited by VIP
PU.1 is known to be an important transcription factor in human myeloid cell TLR expression and differentiation. Therefore, we visualized PU.1 localization via confocal microscopy. Our results clearly showed that E. coli (Fig. 6 A-C ) and P. gingivalis LPS (Fig. 6G 6H 6I) stimulated nuclear translocation of PU.1 in THP1 cells after only 90 min and had a similar effect to PMA (Fig. 6N) . However, following coculture with VIP, E. coli LPS (Fig. 6D 6E 6F) and P. gingivalis LPS (Fig. 6J 6K 6L) failed to induce similar nuclear translocation of PU.1, and in these experiments, a perinuclear localization of PU.1 was observed, which was similar to that seen in unstimulated cells (Fig. 6M) .


Figure 6
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  		Figure 6. VIP inhibits nuclear translocation of the transcriptional regulator PU.1 in LPS-stimulated monocytic THP1 cells, which were stimulated with E. coli LPS (100 ng/ml; A–C), E. coli LPS + VIP (10–8M; D–F), P. gingivalis LPS (100 ng/ml; G–I), and P. gingivalis LPS + VIP (10–8M; J–L) for 90 min. Images in the left column show localization of PU.1, images in center column show transmitted light views, and images in the right column are overlaid images. Nuclear translocation of PU.1 is evident in THP1 cells stimulated with E. coli LPS (A–C) but is inhibited when cells are cocultured with E. coli LPS + VIP (D–F). Nuclear translocation of PU.1 is evident in THP1 cells stimulated with P. gingivalis LPS (G–I) but is inhibited when cells are cocultured with P. gingivalis LPS + VIP (J–L). Unstimulated monocytic THP1 cells (M) demonstrated a perinuclear localization of PU.1 (overlay image of anti-PU.1 FITC and transmitted light image). Cells stimulated with PMA (1 µg/ml) for 90 min (positive control cells) demonstrated PU.1 nuclear translocation (overlay image of anti-PU.1 FITC and transmitted light image; N). All results obtained are representative of results obtained on more than three separate occasions. Original scale bar = 10 µm. Closed arrows denote cell membrane; open arrows denote nuclear membrane.

 
We further analyzed the inhibitory effect of VIP on LPS-induced nuclear translocation of PU.1 by measuring the expression of M-CSFR (a PU.1 gene target) on the surface of E. coli LPS or LPS + VIP-treated THP1 monocytes. The results we obtained showed that LPS had no effect on M-CSFR expression within the first 24 h of culture (data not shown). However, after 48 h, LPS induced an increase in M-CSFR expression by FSChigh cells in the THP1 population (Fig. 7B ) compared with unstimulated THP1 cell cultures in which fewer FSChigh/M-CSFRhigh cells were differentiated (Fig. 7A) . However, when THP1 cells were cocultured with LPS + VIP, there was a significant (P<0.05) decrease in the number of FSChigh/M-CSFRhigh cells (Fig. 7C and 7D) . The FSChigh/M-CSFRhigh THP1 cells induced by LPS were also CD14high compared with unstimulated controls, and this CD14 signal was also reduced in the presence of VIP (data not shown). Therefore, our results show that the greatest M-CSFR expression occurs in the LPS-induced CD14high/FSChigh (macrophage) population of THP1 cells, and differentiation of these cells is inhibited by the presence of VIP. However, LPS did not increase the production of M-CSF protein by monocytic THP1 cells, and this was also not affected by coculture with VIP but was increased significantly (P<0.05) by PMA following culture for 6–24 h (Fig. 7E) .


Figure 7
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  		Figure 7. FACS analysis of M-CSFR expression on the surface of THP1 monocytes cultured for 48 h with E. coli LPS and the effect of VIP. (A) M-CSFR expression on the surface of unstimulated forward-scatter (FSC)high (R1) and FSClow (R2) monocytic THP1 cell populations. (B) Increase in M-CSFRhigh/FSChigh population (R1) following incubation of monocytic THP1 cells with LPS. (C) Decrease in M-CSFRhigh/FSChigh (R1) population when monocytic THP1 cells are cocultured with LPS and VIP. (D) Quantitative analysis of changes in the number of M-CSFRhigh/FSChigh THP1 cells (R1) following incubation with LPS or coculture with LPS and VIP. Histograms are means of three replicate experiments performed on five separate occasions. Increase in CD14 expression was also measured in a M-CSFRhigh/FSChigh (macrophage) population following incubation with LPS, and this population was also decreased following coculture of THP1 cells with LPS and VIP (data not shown). (E) ELISA analysis of mean (±SD) M-CSF protein in monocytic THP1 cell supernatants following treatment with E. coli LPS, E. coli LPS and VIP, PMA, or unstimulated controls. Only PMA induced a significant (P<0.05) increase in M-CSF production above the levels measured in unstimulated cell supernatants. Each data point is a mean of M-CSF concentration measured in triplicate on four separate occasions. *, P < 0.05.

 
Effects of VIP on PBM
Although our data clearly showed that VIP inhibited up-regulation of TLR in LPS-stimulated THP1 cells, this transformed cell line may be inherently different to human monocytes isolated from healthy volunteers. Therefore, we repeated FACS analyses to study TLR expression by PBM isolated from healthy human volunteers. In general, PBM were more refractory to LPS culture than THP1 cells prestimulated with Vit D3, and we measured no change in TLR4 expression in PBM cultured with E. coli LPS for up to 24 h (data not shown). However, after 48 h in culture, E. coli LPS increased TLR4 expression above TLR4 expression measured on the surface of unstimulated control cells, and this was decreased by the addition of VIP into the PBM/LPS cultures (Fig. 8A ). As with THP1 cells, TLR4 expression was greatest in FSChigh cells and was increased following incubation of PBM with E. coli LPS (Fig. 8C) compared with unstimulated control cells (Fig. 8B) . However, the addition of VIP to PBM, cocultured with LPS, significantly (P<0.05) decreased the number of cells in the TLR4high/FSChigh population (Fig. 8D and 8E) . There was also a parallel increase in CD14 expression in the TLR4high/FSChigh population following culture of cells with LPS, and the CD14 signal was also decreased following coculture of PBM with LPS and VIP (data not shown).


Figure 8
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  		Figure 8. FACS analysis of LPS-induced TLR4 and CD14 expression on the surface of human PBM. (A) FACS histogram showing increase in TLR4 expression on the surface of human PBM cultured for 48 h with E. coli LPS and inhibition of LPS-induced TLR4 expression by coculture with VIP. (B) The TLR4high/FSChigh population (R1) in unstimulated PBM cells, compared with the R2 population, which is smaller (FSClow) and expresses less TLR4. (C) Increased TLR4high/FSChigh population (R1) following incubation of PBM with E. coli LPS for 48 h. (D) The TLR4high/FSChigh population (R1) is decreased when PBM are cocultured with LPS and VIP. (E) Analysis of changes in the number of TLR4high/FSChigh THP1 cells (R1) following incubation with LPS or coculture with LPS and VIP. Histograms are means of three replicate experiments performed on five separate occasions. *, P < 0.05. An increase in CD14 expression was also measured in the TLR4high/FSChigh (macrophage) population following incubation with LPS, and this population was also decreased following coculture of THP1 cells with LPS and VIP (data not shown).

 
The CD14high/FSChigh phenotype is indicative of differentiation to macrophages, and the finding that far fewer macrophages were generated in PBM cultures stimulated with LPS (compared with THP1 cells stimulated with Vit D3 prior to LPS) was also observed by microscopy. However, LPS had no effect on TLR4 expression on the surface of non-CD14-expressing PBM in the buffy coat (lymphocytes) at any of the time-points measured (data not shown).

DISCUSSION

Although there is a wealth of reported data showing the inhibitory effect of VIP on inflammatory immune responses in murine myeloid cells or whole animal (murine) models exposed to E. coli or Salmonella LPS (reviewed in refs. [19 , 20 ]), there are surprisingly few studies of this kind using human cells. VIP has been shown to inhibit IL-8 production from THP1 cells stimulated with LPS from E. coli 0111:B4 [21 ], and recently, we have shown that VIP also inhibits TNF-{alpha} production by THP1 cells cultured with LPS from E. coli 0111:B4 or P. gingivalis W50 [16 ]. However, for the first time, our current study clearly shows that VIP inhibits TLR2 up-regulation following incubation of THP1 cells with P. gingivalis W50 LPS and TLR4 up-regulation following incubation of human THP1 cells or PBMC with E. coli 0111:B4 LPS.

Within 6 h, E. coli and P. gingivalis LPS up-regulated TLR4 expression on the surface of THP1 cells, but when these cells were cocultured with LPS and VIP, TLR4 expression was decreased to levels similar to those measured on the surface of unstimulated (control) cells. The effect on TLR2, however, was less apparent. P. gingivalis LPS up-regulated TLR2 expression on the surface of THP1 cells more than E. coli LPS after 6 h, but the increase was not as great as that measured for TLR4 following exposure to E. coli LPS. VIP, therefore, reduced the expression of TLR2, proportionately much less than was the case for TLR4. This suggests that monocytes may respond to LPS (whether TLR2- or TLR4-dominant), first by up-regulating TLR4, possibly explained by our previous work, which has shown (at least in mice) that functional TLR4 is needed for TLR2 expression [22 ].

The most dramatic effect of LPS on TLR2 and TLR4 expression was seen after 24 h. Muthukuru et al. [23 ] have shown that TLR4 and TLR2 are up-regulated on the surface of human monocytes incubated with E. coli LPS or P. gingivalis LPS for 24 h, and studies using THP1 monocytes have shown that up-regulation of TLR2 in response to P. gingivalis LPS is dose-dependent over the 24-h incubation period [24 ]. However, both of these studies show that following a washing and settling period, TLR hyporesponsiveness occurs on rechallenge with LPS. Our studies, using THP1 cells pretreated with Vit D3, show that both LPS types stimulated the generation of a second adherent cell population, which was clearly discernible by FACS analysis and had macrophage morphology when compared, by microscopy, to THP1 cells cultured with Vit D3 followed by PMA, a technique commonly used to convert promonocytic THP1 cells to macrophages [25 ]. In a previous study, P. gingivalis and E. coli LPS have also been reported to induce differentiation of human monocytes to macrophages [26 ].

However, in our study, we show that in the presence of different LPS types, monocytic THP1 cells differentiate into macrophages, which express the correct TLR receptor for the LPS type present, such that P. gingivalis LPS induced predominantly TLR2-expressing macrophages, and THP1 cells exposed to E. coli LPS predominantly gave rise to TLR4-expressing macrophages. However, we did not study the effect of LPS rechallenge on TLR expression in these newly differentiated macrophage populations. Coculture of monocytic THP1 cells with LPS and VIP inhibited TLR surface expression, and our results suggest that probably the greatest single effect of VIP on inhibition of TLR expression is a result of the ability of VIP to inhibit differentiation of monocytes to macrophages following Vit D3/LPS stimulation. We also studied the effect of VIP on Vit D3 differentiation of promonocytic THP1 cells to monocytic THP1 cells. Our results showed that VIP (10–8 M) had no effect on Vit D3-induced promonocyte differentiation.

A number of naturally occurring TLR inhibitors have been discovered, which inhibit the effect of TLRs at various subcellular levels (reviewed in ref. [27 ]). Mechanisms that inhibit TLR at the level of the cell membrane include the production of soluble TLR2 (sTLR2) [28 ] and sTLR4 [29 ], which inhibit LPS/TLR engagement. In human embryonic kidney 293 cells, TLR4 surface expression is controlled by increased ubiquitination by the protein Triad3A [30 ]. Gomariz et al. [7 ] have suggested that a possible mechanism for inhibition of TLR2 and TLR4 by VIP in TNBS-induced colitis of mice may involve inhibition of NF-{kappa}B, and previous studies have shown that VIP inhibits degradation of I{kappa}B and therefore prevents nuclear translocation of NF-{kappa}B [31 ]. Although we have shown that VIP inhibits nuclear translocation of NF-{kappa}B and c-Jun in THP1 cells, cultured with LPS from P. gingivalis or E. coli [16 ], there is an absence of NF-{kappa}B response sites on the murine TLR4 and human TLR4 promotor [11 ].

Human TLR4 is regulated by PU.1 and also the IFN-response factor [11 , 13 ]. Similarly, human TLR2 is also regulated by PU.1 [12 ], and so, these studies previously reported led us to investigate the effect of VIP on PU.1. Confocal microscopy showed that when THP1 cells were cultured with P. gingivalis or E. coli LPS for 90 min, PU.1 was translocated from the cell cytoplasm to the nucleus. However, in the presence of VIP, nuclear translocation of PU.1 was prevented. We, therefore, hypothesize that inhibition of LPS-induced TLR2 and TLR4 up-regulation by VIP may occur (at least in part) as a result of inhibited nuclear translocation of PU.1. However, the inhibitory effect of VIP on nuclear translocation of PU.1 may also explain the inhibitory effect of VIP on THP1 differentiation to macrophages, as apart from its role in the expression of human TLR2 and TLR4, PU.1 is critical for differentiation of human myeloid cells [32 ]. Previous confocal studies have shown that nuclear accumulation of PU.1 occurs after 60 min when murine RAW cells (macrophages) are cultured with 1 µg/ml LPS [18 ], and in accordance with these results, our data clearly showed that LPS from E. coli or P. gingivalis (or PMA) stimulated nuclear accumulation of PU.1 within 90 min, but when THP1 cells were cocultured with LPS and VIP, nuclear accumulation of PU.1 was inhibited.

Further evidence that VIP inhibited LPS-induced nuclear translocation of PU.1 was obtained by studying M-CSFR expression. The M-CSFR gene is a known target for PU.1, and murine myeloid cells isolated from PU.1 null mice lack M-CSFR, and differentiation of these cells into macrophages is prevented [33 , 34 ]. In our study, VIP inhibited LPS-induced nuclear translocation of PU.1 and subsequent LPS-induced up-regulation of M-CSFR. However, the downstream effect of LPS-induced translocation of PU.1 (M-CSFR expression) and the inhibitory effect of VIP on M-CSFR expression were only apparent 24–48 h after incubation. Neither LPS nor LPS/VIP affected production of M-CSF protein, thus indicating that differentiation of THP1 cells, under our experimental conditions, is PU.1-dependent but does not involve M-CSF protein. Furthermore, c-Jun, which has autonomous, transcriptional activity and activity as part of the activator protein 1 complex [35 ], has also been shown to synergize with PU.1 for up-regulation of M-CSFR [36 ]. We have previously reported that VIP inhibits nuclear translocation of c-Jun in THP1 monocytes stimulated with LPS from E. coli 0111:B4 or P. gingivalis W50 [16 ]. Therefore, we cannot rule out the possibility that VIP may affect THP1 differentiation by inhibiting PU.1 and c-Jun.

We tested the inhibitory effect of VIP on TLR4 up-regulation by E. coli LPS on total human PBMC populations (buffy coat cells) and isolated PBMC. We found that LPS did not induce TLR up-regulation in non-CD14-expressing buffy coat cells (lymphocytes), which is in contrast to Gomariz et al. [7 ], who reported that murine lymphocytes up-regulated TLR receptors in mice with TNBS-induced colitis. This may be explained by species differences or the difference between LPS and TNBS (although TNBS-induced colitis probably activated TLR in these mice by exposing myeloid and lymphoid cells to intestinal bacteria). Caron et al. [37 ] have shown that highly purified human T cells up-regulate TLR5, -7, and -8 in response to appropriate ligands, and so, it is conceivable that E. coli LPS may have also up-regulated TLR4 on the surface of the T cells we used. However, we consistently found that PBMC were much less responsive to LPS than were THP1 cells. We believe that this may be a result of the Vit D3 stimulus that THP1 cells received prior to stimulation with LPS, as Vit D3 stimulates the differentiation required to convert promonocytic THP1 cells to monocytic THP1 cells, and following further stimulation with LPS, a number of macrophages were observed in culture, which increased the TLR signal. Far fewer macrophages were observed in culture when PBMC were cultured with LPS, and this coincided with a reduced TLR4 signal. Nevertheless, if monocytes were cultured for up to 48 h with LPS, there was an increase in the TLR4 signal in FSChigh/CD14high cells, and this was inhibited in the presence of VIP.

In conclusion, our data show, for the first time, that VIP inhibits LPS-induced TLR2 and -4 expression on the surface of THP1 monocytes and macrophages and also inhibits LPS-induced TLR4 expression on the surface of macrophages derived from human PBMC. The mechanism for this inhibition is most likely a result of decreased nuclear translocation of PU.1 and in light of our previously published data [16 ], possibly also c-Jun. These data highlight a novel, immunomodulatory mechanism, which may be a useful, therapeutic target in chronic inflammatory disorders mediated by LPS.

ACKNOWLEDGEMENTS

This work was supported by a UK Department of Health Clinician Scientist Award (DHCS/030G121/46) to P. M. P. The authors acknowledge the help of Dr. Trevor Booth (Confocal Microscopy) and Mr. Ian Harvey (FACS).

Received February 7, 2006; revised July 11, 2006; accepted August 4, 2006.

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N. Foster, K. Andreadou, L. Jamieson, P.M. Preshaw, and J.J. Taylor
VIP Inhibits P. gingivalis LPS-induced IL-18 and IL-18BPa in Monocytes
J. Dent. Res., September 1, 2007; 86(9): 883 - 887.
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