CD4+ as Monocytes cell surface receptors

CD4+ as Monocytes cell surface receptors

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Today in class, our teacher was discussing about the pathogenecity of the HIV virus.

He said that the binding between HIV virus and macrophages is done by the interaction between the GP120 of the virus and CD4 receptors,chemokine receptors(CCR-5 and CXCR-4) and fusin receptor present on Macrophages.

My doubt is I think CD4 receptors are present on T-helper cells only. Are they present on Macrophages too? Do i have a wrong notion about CD4 receptors? If not, then are there some other receptors responsible for the transfer of viral particles inside monocytes and dendritic cells?

Any extra information is appreciated.

Macrophages and monocytic cells can express CD4, CD8 or even co-express them. The CD4 and CD8 molecules interact with MHC complexes on neighboring cells, and as you can imagine elicit varying effect in a cell type-dependent manner.

Plainly enough, Zhen et al. demonstrated that monocytic CD4 ligation to MHC-II molecules on other cells, such as activated endothelium, induced an expression pattern conducive to macrophage differentiation (1). Gibbings et al. also demonstrated that CD8 associates with FcyR on the monocyte's surface similar to CD8/TCR associations, and stimulates the secretion of cytokines (2).

I'll try to find a model source, but CD8a on dendritic cells can associate with MHC-II molecules on T cells like HLA-DR and aid in cross-priming the T cells.

Now what CD4 does on functional, differentiated macrophages is less clear to me, and I need to review the literature on that a bit more. It could just be expression imprinted because they're monocyte-derived cells, or it could also elicit an effect like CD8/FcryR above. I do not know.

There are more cases, so these serve as selected examples.


In molecular biology, CD4 (cluster of differentiation 4) is a glycoprotein found on the surface of immune cells such as T helper cells, monocytes, macrophages, and dendritic cells. It was discovered in the late 1970s and was originally known as leu-3 and T4 (after the OKT4 monoclonal antibody that reacted with it) before being named CD4 in 1984. Ώ] In humans, the CD4 protein is encoded by the CD4 gene. ΐ] Α]

CD4+ T helper cells are white blood cells that are an essential part of the human immune system. They are often referred to as CD4 cells, T-helper cells or T4 cells. They are called helper cells because one of their main roles is to send signals to other types of immune cells, including CD8 killer cells, which then destroy the infectious particle. If CD4 cells become depleted, for example in untreated HIV infection, or following immune suppression prior to a transplant, the body is left vulnerable to a wide range of infections that it would otherwise have been able to fight.

Second-messenger regulation of receptor association with clathrin-coated pits: a novel and selective mechanism in the control of CD4 endocytosis.

CD4, a member of the immunoglobulin superfamily, is not only expressed in T4 helper lymphocytes but also in myeloid cells. Receptor-mediated endocytosis plays a crucial role in the regulation of surface expression of adhesion molecules such as CD4. In T lymphocytes p56lck, a CD4-associated tyrosine kinase, prevents CD4 internalization, but in myeloid cells p56lck is not expressed and CD4 is constitutively internalized. In this study, we have investigated the role of cyclic AMP (cAMP) in the regulation of CD4 endocytosis in the myeloid cell line HL-60. Elevations of cellular cAMP were elicited by 1) cholera toxin, 2) pertussis toxin, 3) forskolin and IBMX, 4) NaF, or 5) the physiological receptor agonist prostaglandin E1. All five interventions led to an inhibition of CD4 internalization. Increased cAMP levels did not inhibit endocytosis per se, because internalization of insulin receptors and transferrin receptors and fluid phase endocytosis were either unchanged or slightly enhanced. The mechanism of cAMP inhibition was further analyzed at the ultrastructural level. CD4 internalization, followed either by quantitative electron microscopy autoradiography or by immunogold labeling, showed a rapid and temperature-dependent association of CD4 with clathrin-coated pits in control cells. This association was markedly inhibited in cells with elevated cAMP levels. Thus these findings suggest a second-messenger regulation of CD4 internalization through an inhibition of CD4 association with clathrin-coated pits in p56lck-negative cells.

Clinical features of HIV-1 infection

The HIV-1 virus is transmitted by exchange of bodily fluids. The mode of transmission may involve the transfer of free virions or HIV-1 infected cells. Initial (acute) infection with HIV-1 results in clinical symptoms within 1 to 3 weeks in at least half of those newly infected. These symptoms are similar to influenza or mononucleosis along with a non-pruritic macular erythematous rash. 52 Shortly after acute infection, most patients undergo seroconversion. This is followed by a period of clinical latency, which may last from 3 to more than 15 years, before AIDS develops and the patient eventually dies of multiple infections and/or malignancies. Progression to AIDS is accompanied by loss of CD4 + T lymphocytes, with symptoms being noticed at blood levels less than 500 cells/L. Although the vast majority of those who are infected with HIV-1 will develop AIDS, there is mounting evidence that some people are able to live with the virus for extended periods of time without developing clinical disease. Such individuals are termed ‘long-term non-progressors’, although only time will tell if this group will also succumb to disease. 53 Factors that affect the rate of progression to AIDS (for review see Levy 54 ) include age (most HIV-1 infected infants progress relatively slowly 55 , 56 ), general health (the presence of other infections may speed progression to AIDS 57 ) and lifestyle (smoking, 58 alcohol 59 and drug use 60 may all speed progression). Differences in the infecting HIV-1 strain and the host immune response are probably also important in disease progression rates.


CD8 and CD4 are expressed by several cell types that do not express TCR, although this statement should be cautioned by the demonstration of rearranged TCR on neutrophils [ 21 ] and the presence of rearranged TCR genes in DCs [ 4 ] and NK cells [ 22 ]. Mouse and human species have significant differences in their immune systems [ 23 ]. One of these differences is in the cell types, which express CD4 and CD8. Although CD8 is found on DC and T cells in all species [ 24 25 26 ], it is only in rat and human that CD8 is found on a variety of other immune cells. Current evidence suggests that CD8α protein is expressed by NK cells in rat [ 27 ] and human [ 28 ] but not in mouse [ 29 ] (Table 1 and references therein). Similarly rat [ 42 , 44 ] and human monocytes [ 31 ] and macrophages [ 35 ] express CD8α, and the same cells in the mouse lack CD8α [ 33 ]. Intriguingly, rat macrophages and mast cells also express CD8β [ 35 , 36 ]. Analogous to the situation for CD8, CD4 in rat and human is expressed by a wider variety of innate immune cell types than in mouse (Table 1 and references therein). Interestingly, DCs also diverge among species in their expression pattern of CD4: Mouse DCs express high levels of CD4, but human and rat DCs express CD4 at low levels [ 26 ].

Mechanisms of activating intracellular signaling of CD4 and CD8

At different temporal stages of TCR signaling, CD4 and CD8 may function inside or outside lipid rafts to affect intracellular signaling through lck or LAT. In one of the first steps of TCR activation, CD45, located outside lipid rafts, dephosphorylates lck. CD4 and CD8 immunoprecipitate with CD45 [ 45 , 46 ] and may be involved in early lipid raft-independent dephosphorylation of lck. Dephosphorylated lck then moves into lipid rafts to phosphorylate two sites on CD3ζ. Dynamic palmitoylation of lck and/or CD4 and CD8 may regulate the transition of lck into lipid rafts. Subsequently, ZAP-70 can be recruited by phosphorylated CD3ζ. ZAP-70 can then phosphorylate LAT, which is potentially associated with CD4 and CD8 [ 12 ]. This lipid raft-associated LAT can center a scaffolding network of intracellular signaling complexes, indirectly recruiting SLP-76 to TCR complexes. SLP-76 recruits non-catalytic region of tyrosine kinase (NCK) and allows activation of recruited Itk, contributing to the phosphorylation and activation of PLCγ [ 47 ]. Subsequent binding of PLCγ to LAT along with Grb2 together will strongly influence calcium flux, cytoskeletal reorganization, proliferation, and gene transcription [ 48 ].

In addition to activating intracellular signaling through lck or LAT, CD8 may affect TCR-mediated T cell activation via its extracellular binding to MHC class I. TCR binds MHC-peptide with a slow association rate (10 3 –10 5 /Ms −1 ), typical of induced fit binding [ 9 ], whereas CD8 has a faster association rate typical of rigid body binding (1.4×10 5 /Ms −1 ) [ 49 ]. CD8, statistically, should contact MHC class I on an apposing cell before TCR, as experimental data confirm [ 50 ]. The binding kinetics of CD4 for MHC class I is less clear but resembles that of CD8 in general [ 9 ], suggesting similar models may be applicable to it as well. Interestingly, two-dimensional crystallography of MHC class I on lipid rafts suggests MHC class I may adopt a supine orientation on the plasma membrane with the CD8 binding site exposed and the TCR binding site partially obscured [ 51 ]. Thus, association rates and spatial considerations suggest CD8 may establish first interactions with MHC class I and heighten the probability of TCR scanning for and bending to productively engage MHC class I at a later time-point.


In this study, the MRM MS and SEM measurements are used to assess two human blood cell preparations in search of optimal cell reference materials for quantitative flow cytometry that are more stable and easier to maintain than fresh whole blood. Because of the lyophilization process, the CD4 density value on CD4+ lymphocytes from Cyto-Trol cells is lower than the value from cryopreserved PBMC, most likely explained by the truncation of the CD4 receptor proteins and damaged and/or broken microvilli where CD4 receptors reside. On the other hand, steric hindrance of antibody binding and the association of CD4 receptors with other biomolecules likely contribute significantly to the close to 50% lower CD4 receptor density value for cryopreserved PBMC determined from flow cytometry than the value obtained from MRM MS.

The consistent CD4 expression on T cells in normal donor PBMC, serving as the biological control enhances the reliability of clinical diagnostics and immunotherapies. This CD4 receptor protein also plays a crucial role in the progression of HIV-1 viral infection in that the gp120 viral protein binds to the CD4 receptor on T cells, leading to the viral entry and cell disintegration [15]. Though numerous efforts have been put in the development of vaccines against the infection, there is limited success largely because of the complexity of the viral infection process and limited robust measurement techniques supporting the understanding of the underlying mechanisms of trial vaccines. The three powerful techniques used in this study, flow cytometry, MRM MS and scanning electron microscopy, allowed us to better understand the changes caused by the lyophilization process on CD4+ lymphocytes. These techniques would enable the measurements of CD4 receptor density and the number of anti-CD4 monoclonal antibodies, e.g., Ibalizumab [16] and Bispecific Ibalizumab [17], bound to cells bearing CD4 receptors. These measurements would greatly help to shed light on the underlying mechanisms of two trial vaccines for the treatment and prevention of HIV-1. Furthermore, down modulated CD4 cell surface expression and subcellular localization [18], and depletion of the surface CD4 protein [19] have been reported in the literature in some cases of HIV infection. It would certainly be more challenging to apply these techniques to measure internalized CD4 proteins in different cell compartments.


Specific Coimmunoprecipitation of CD4 and CCR5 from Cell Lines.

It was previously found that the soluble D1D2 fragment, but not the entire extracellular portion of CD4, interferes with the chemokine MIP-1α for binding to CCR5, indicating possible interactions between CD4 and CCR5 (11). Although the differences between the two-domain and the four-domain fragments of CD4 could be attributed to a better exposure of conformational epitopes in the two-domain CD4 fragment, an alternative possibility is that the CD4D1D2–CCR5 interaction does not reflect the properties of the wild-type (membrane-associated) CD4 binding to CCR5. Therefore, we used an immunoprecipitation assay, optimized for high efficiency and reproducibility, to directly detect the interactions between native membrane-associated CD4 and CCR5.

To demonstrate coimmunoprecipitation of cell surface-associated CD4 and CCR5, we used 3T3 cell lines transfected to express CD4 (3T3.CD4), CD4 and CXCR4 (3T3.CD4.CXCR4) or CD4 and CCR5 (3T3.CD4.CCR5). The CD4 concentrations at the surfaces of these cells were very similar: their ratios were 1.26:1.2:1, respectively, as measured by using flow cytometry (data not shown see also Fig. 1A, lane II). The surface concentrations of CCR5 and CXCR4 were also very similar (flow cytometry data not shown). Because CD4 is easier to detect by Western blotting and biotinylation than CCR5, in most cases we used anti-CCR5 antibodies to coimmunoprecipitate CD4. We found that the N terminus-specific mAb 5C7 (29) was the most efficient antibody in immunoprecipitating CCR5 compared with a battery of other anti-CCR5 antibodies (data not shown). This mAb coimmunoprecipitated surface-associated CD4 in 3T3 cell lines coexpressing CD4 and CCR5 (Fig. 1A, lane IV). The band was CD4 because it aligns with the CD4 band obtained by direct immunoprecipitation using an anti-CD4 mAb (OKT4) (Fig. 1A, lane II), has the expected molecular weight (55 kDa), and reacts specifically in the Western blot assay (Fig. 1A, lane V). CD4 was highly and specifically enriched in the coimmunoprecipitates as was observed by comparison of cell-surface biotinylated lysates that were not subjected to immunoprecipitation (Fig. 1A, lane I) with the coimmunoprecipitates (Fig. 1A, lane IV). In these experiments, CCR5 was not detected because of the low efficiency of biotinylation however, it was observed by using Western blotting (Fig. 1A, lane VI). Similar results were obtained for a number of cell lines, including PM1 and L1.2 transfectants, that coexpress CCR5 and CD4 (data not shown).

Specific coimmunoprecipitation of cell surface-associated CD4 and CCR5 from 3T3 cells coexpressing these two molecules. (A) Equal numbers of 3T3 cells expressing CD4 and CCR5 (+) or CD4 only (−) were biotinylated, processed as described in Materials and Methods, and either used as a whole-cell lysate (0.25% of total, gel I) or immunoprecipitated with an anti-CD4 mAb (OKT4) (gel II), anti-CXCR4 mAb (4G10) (gel III), or anti-CCR5 mAb 5C7 (gels IV–VI). The biotinylated proteins were detected by using streptavidin–horseradish peroxidase (gels I-IV). CD4 and CCR5 were detected in an aliquot of the same samples as in gel IV by using Western blotting with an anti-CD4 polyclonal antibody (T4-4) (gel V) or an anti-CCR5 polyclonal antibody [CKR5(C20)] (gel VI). (M denotes molecular markers, and the numbers are in kDa). (B) Coimmunoprecipitation of CD4 by the anti-CCR5 mAbs m180 (lane 1) and m181 (lane 2). The coimmunoprecipitated CD4 and the immunoprecipitated CCR5 were detected by sing Western blotting as in A. (C) Coimmunoprecipitation of CCR5 from 3T3.CD4.CCR5 cells with anti-CD4 antibodies. 3T3.CD4.CCR5 cells (lanes 1, 2, and 4) or 3T3.CD4 cells (lane 3) were used for immunoprecipitation by OKT4 (lanes 2 and 3) or by a control antibody (CG10) (lane 4). Lane 1 shows for comparison immunoprecipitation with the anti-CCR5 mAb 5C7. CD4 and CCR5 were detected by using Western blotting as in A. (D) Cell lysates were immunoprecipitated by the anti-CCR5 mAb 5C7, the immunoprecipitation product was analyzed by using a silver stain kit (lanes 1 and 2) and compared with proteins detected by streptavidin–horseradish peroxidase in biotinylated lysates (lanes 3 and 4). M denotes molecular weight marker, and + and − denote 3T3.CD4.CCR5 or 3T3.CD4 cells, respectively. ∗, bands caused by CD4 and CCR5. The two bands above and below CD4 are caused by the 5C7 mAb heavy and light chain, respectively. Lane 2 represents lane 1 at higher sensitivity, where CCR5 is clearly seen. (E) CD4–CCR5 coimmunoprecipitation is not significantly affected by cholesterol depletion. 3T3.CD4.CCR5 cells were treated with 10 mM methyl-β-cyclodextrin for 1 hr at 37°C (which caused significant cytotoxicity) and used for immunoprecipitation by the anti-CCR5 antibody 5C7 (lane 2), and compared with untreated cells (lane 1) and 3T3.CD4 cells (lane 3). CD4 and CCR5 were detected by using Western blotting as in A. Bottom shows Western blotting of CD4 from whole-cell lysates.

The specificity of the CD4 coimmunoprecipitation was demonstrated through several control experiments. The anti-CCR5 mAb 5C7 did not coimmunoprecipitate CD4 from 3T3.CD4 cells that do not express CCR5 (Fig. 1A, lanes IV and V). In another experiment, an antibody (4G10) to a related chemokine receptor (CXCR4) was used. This antibody is able to efficiently immunoprecipitate CXCR4 and coimmunoprecipitate CD4 and CXCR4 in cells expressing these two molecules (Fig. 3A) but did not coimmunoprecipitate CD4 in cells expressing CD4 and CCR5 (Fig. 1A, lane III). In addition, the amount of coimmunoprecipitated CD4 was proportional to the amount of immunoprecipitated CCR5, demonstrated by using two anti-CCR5 mAbs with different immunoprecipitating activities (Fig. 1B), further suggesting a specific CD4–CCR5 interaction. We coimmunoprecipitated CCR5 with an anti-CD4 antibody (OKT4) (Fig. 1C, lane 2), but not with a control antibody (CG10), which does not bind to CD4 (lane 4), in cells coexpressing these two molecules (lane 2), but not in CCR5 negative cells (lane 3).

To further evaluate the specificity of the CD4–CCR5 interaction and to address the question of whether other proteins interact with CCR5 and potentially influence the CD4–CCR5 association, we used silver staining of proteins immunoprecipitated by the anti-CCR5 mAb 5C7 in parallel with biotinylation (some proteins, particularly chemokine receptors, are poorly labeled by biotin and at low concentrations may not be detected). Apart from the bands corresponding to CCR5 and CD4, the only other major bands are those representing the heavy and light chains of the precipitating mAb 5C7 and bands that are apparently not specific to CCR5, because they were immunoprecipitated in CCR5-negative cells (Fig. 1D). These data not only imply that the interaction between CD4 and CCR5 is not mediated by another molecule but also indicate that the coimmunoprecipitation of CD4 and CCR5 is unlikely to be due to compartmentalization of these two molecules within defined membrane microdomains, which would lead to coimmunoprecipitation of a greater number of proteins. Another experiment supporting this notion shows that cell lysis with two different detergents (Brij97 and NP40) did not disrupt the CD4–CCR5 interaction (data not shown). Furthermore, depletion of cholesterol from the cell membrane with methyl-β-cyclodextrin, a procedure that was shown to disrupt microdomain structure (see, e.g., ref. 30), did not block the coimmunoprecipitation of CD4 and CCR5 (Fig. 1E).

Coimmunoprecipitation of CD4 and CCR5 in Primary CD4 + T Cells, Macrophages, and Monocytes.

Results similar to those described above for cell lines were also obtained with primary human cells susceptible to HIV-1 entry. Our initial attempts to coimmunoprecipitate CD4 and CCR5 from the surface of primary T lymphocytes resulted in very weak bands that were at the limit of assay sensitivity, because of the low level expression of CCR5 at the surface of these cells and the relatively small percentage of cells expressing it as evaluated by flow cytometry (data not shown). By using two alternative procedures for activating the CD4 + T cells, as described in Materials and Methods, expression of CCR5 was significantly increased to high (+) and very high (++) levels, corresponding on average to ≈2–4 × 10 3 and 3–5 × 10 4 molecules, respectively, as estimated by using quantitative flow cytometry. In these cells, CD4 levels were similar, and the amount of CD4 coimmunoprecipitated with the anti-CCR5 mAb 5C7 correlated with their cell fusion efficiency (Fig. 2A). The amount of coimmunoprecipitated CD4 in human macrophages and monocytes also correlated with the efficiency of their fusion with cells expressing the HIV-1 JRFL Env (Fig. 2B) but not with the surface concentration of CD4. Indeed, the monocytes expressed similar or higher levels of CD4, but the amount of coimmunoprecipitated CD4 was undetectable or barely detectable in our assay (Fig. 2B). The larger amount of coimmunoprecipitated CD4 in macrophages is likely related to the higher levels of CCR5 compared with monocytes—on average ≈5–10 × 10 3 vs. <2 × 10 3 molecules per cell as estimated by using quantitative flow cytometry. However, the CCR5 levels in macrophages were lower compared with the ++ CD4 + T cells, and we were not able to detect the immunoprecipitated CCR5 by Western blotting (data not shown).

Coimmunoprecipitation of CD4 and CCR5 in primary cells. (A) Human CD4 T cells expressing high (+) and very high (++) amounts of CCR5 were used for immunoprecipitation with the anti-CCR5 mAb 5C7 (Center) or fusion with HeLa cells expressing the HIV-1 JRFL Env (Left and Right). The coimmunoprecipitated CD4 (Center Top) and immunoprecipitated CCR5 (Center Middle) were detected by using Western blotting as in Fig. 1A. The CD4 Western blotting of whole-cell lysates is shown (Bottom) as a measure of the level of CD4. The average number of syncytia for ++ cells was 92 ± 10.5, for + cells was 29 ± 7, and for control HeLa cells was 6 ± 3. The average diameter of syncytia from the ++ cells was about 4-fold larger than that for the + cells. (B) Human macrophages (Left, lane 1) and monocytes (Left, lane 2) were used for CCR5 immunoprecipitation. CD4 coimmunoprecipitation was detected as in A. The CD4 Western blotting of whole-cell lysates is shown (Bottom) as a measure of the level of CD4. (Right) Fusion of these cells with HeLa cells expressing the HIV-1 JRFL Env as quantitated by the β-galactosidase assay. The control represents HeLa cells that do not express HIV-1 Env.

CD4 Interaction with CCR5 Is Stronger than That with CXCR4 and Is Not Increased in the Presence of gp120.

We previously found that CD4 could associate weakly with CXCR4 even in the absence of gp120 (10). However, the results varied in a cell line- and assay condition-dependent manner. To evaluate the strength of the CD4–CCR5 association relative to the CD4–CXCR4 interaction, 3T3 cell lines expressing CD4 and either CCR5 or CXCR4 at approximately the same surface concentrations were used. In the 3T3.CD4.CXCR4 cells, CD4 associated weakly with CXCR4, but the association was dramatically increased by addition of X4 HIV-1 Env gp120 (IIIB) (Fig. 3A). In contrast, the amount of CD4 coimmunoprecipitated by the anti-CCR5 mAb 5C7 in the 3T3.CD4.CCR5 cells was high even in the absence of gp120, and the addition of X4R5 (89.6) or R5 (JRFL) HIV-1 Env gp120 did not significantly increase the CD4–CCR5 coimmunoprecipitation (Fig. 3 B and C). The quantity of CD4 coimmunoprecipitated by anti-CCR5 mAbs in the absence of gp120 was about the same as the quantity of CD4 coimmunoprecipitated by anti-CXCR4 mAbs in the presence of gp120. These results indicate that the CD4–CXCR4 association is weaker than the CD4–CCR5 interaction and that the preformed complexes between CD4 and CCR5 are close to a saturation level, where the addition of gp120 cannot further increase their complex formation.

The effect of gp120 on the CCR5–CD4 and CXCR4–CD4 association. (A) 3T3.CD4.CXCR4 cells were incubated with the anti-CXCR4 mAb 4G10 in the absence (−) or presence (+) (5 μg/ml) of HIV-1 IIIB gp120. The CD4 and CXCR4 were detected by using Western blotting with either the polyclonal anti-CD4 Ab T4–4 or 4G10. (B) 3T3.CD4.CCR5 cells were incubated with the anti-CCR5 mAb 5C7 in the absence or presence (5 μg/ml) of HIV-1 89.6 gp120. The CD4 and CCR5 were detected by using Western blotting as described in Fig. 1A. (C) The same as in B but gp120 from the R5 HIV-1 JRFL was used instead of the dual tropic 89.6.

The First Two Domains of CD4 and the Second Extracellular Loop of CCR5 Are Probably Involved in the Formation of the CD4–CCR5 Complex.

To identify possible regions of CD4 that are responsible for the interaction with CCR5, we used a cell line (A2.01.T4.T8) expressing a hybrid CD4–CD8 molecule containing the first two domains of CD4, which was previously shown to support HIV-1 Env-mediated fusion (31) although at a lower rate than the wild-type CD4 (16). The A2.01.T4.T8 cells were induced to express CCR5 with a recombinant vaccinia virus encoding the CCR5 gene. These cells, coexpressing CCR5 and the CD4-CD8 hybrid molecule, fused with cells expressing the R5 HIV-1 Env Bal, although at somewhat lower efficiency compared with cells expressing wild-type CD4 (data not shown). These results are analogous to our previously reported observations of fusion between the A2.01.T4.T8 cells and cells expressing the X4 HIV-1 Env IIIB (16). The CD4–CD8 molecules were coimmunoprecipitated by an anti-CCR5 mAb from the A2.01.T4.T8 cells expressing CCR5 but not from those infected with control wild-type vaccinia virus (Fig. 4A). The surface levels of CD4–CD8 molecules in the cells infected with the CCR5 and wild-type vaccinia viruses were not significantly different as quantified by flow cytometry (data not shown) and Western blotting (Fig. 4B). To establish that the CD8 portion of the CD4–CD8 hybrid molecule was not involved in the interaction with CCR5, we used HeLa cells expressing either CD4 or CD8. CCR5 was again expressed in these cells by recombinant vaccinia virus. In those cells only CD4 but not CD8 was coimmunoprecipitated with an anti-CCR5 mAb (5C7), demonstrating that the CD8 portion of the hybrid CD4–CD8 molecule was unlikely to be involved in the interaction with CCR5 (data not shown). Together, these results suggest that the first two domains of CD4 associate with CCR5.

Involvement of the first two domains of CD4 and the second extracellular loop of CCR5 in the CD4–CCR5 association. (A) Coimmunoprecipitation of CD4–CD8 hybrid molecules containing the first two domains of CD4 by an anti-CCR5 mAb. The A2.01.T4.T8 cells expressing the hybrid CD4–CD8 molecule were infected with a recombinant vaccinia virus (vvCCR5–1107) (lane 1), encoding the gene for CCR5, and a control wild-type (WR) vaccinia virus (lane 2). The anti-CCR5 mAb 5C7 was used to immunoprecipitate CCR5 and T4–4 for detection of CD4 by Western blotting. (B) The amount of CD4–CD8 molecules in A2.01.T4.T8 cells infected with the CCR5 (lane 1) or WR (lane 2) vaccinia virus was not significantly different as demonstrated by Western blot with an anti-CD4 Ab (T4–4). (C) Coimmunoprecipitation of CCR5 by an anti-CD4 mAb (OKT4) is inhibited in the presence of another anti-CD4 mAb (CG7). For comparison, lane 1 shows CCR5 immunoprecipitated by 5C7. Lanes 2, 3, and 4 represent the amount of CCR5 coimmunoprecipitated by a mixture of OKT4 (ascites fluid 3.5 μl/ml) and increasing concentrations of the anti-CD4 mAb CG7 (0, 5, and 10 μg/ml, respectively). (D) CD4 coimmunoprecipitation by 5C7 is inhibited in the presence of another anti-CCR5 mAb (2D7) (29) directed to the second extracellular loop. Equal amounts (3 μg/ml) of 5C7, which does not affect HIV entry, were mixed with increasing amounts (0, 3, and 6 μg/ml) (lanes 1, 2, and 3, respectively) of the HIV-1-blocking mAb 2D7 and used for immunoprecipitation. The sample obtained from 9 × 10 6 3T3.CD4.CCR5 cells was divided into two portions, and the smaller one (1/6 of total) was used for Western blot of CD4 by T4–4, and the rest were used for Western blot of CCR5 by the CKR5(C20). (E) The CCR5-terminus-specific mAb 5C7 (lane 2) immunoprecipitates CCR5 much more efficiently than the CCR5 ecl-2-specific mAb 2D7 (lane 1). Equal amounts (4 μg/ml) of these two mAbs were used for immunoprecipitation of CCR5 in 3T3.CD4.CCR5 cells. The molecular markers are shown on the right side (lane M). (F and G) Differential inhibition by two anti-CCR5 mAbs, m182 and m183 (which do not immunoprecipitate CCR5 as measured by our assay) of the CCR5 coimmunoprecipitation by the anti-CD4 mAb OKT4. Lanes 1, 2, and 3 represent 1, 2, and 4 μg/ml of m182 and m183, respectively. CCR5 was detected by Western blotting with CKR5(C20).

We also confirmed the previous observations that a soluble fragment of CD4 consisting of the first two domains (sCD4D1D2) competes with macrophage inflammatory protein (MIP)1-α for CCR5 (11). Interestingly, we found that the HIV-1 infection inhibiting mAb CG7 at 60 nM almost completely blocked the ability of sCD4D1D2 to displace the chemokine MIP-1α (Table 1). The same antibody significantly inhibited the coimmunoprecipitation of CCR5 by the anti-CD4 mAb OKT4 (Fig. 4C). Unlike CG7, a second anti-CD4 mAb (CG1) and a control mAb (CG10) were not effective in preventing the displacement of MIP1-α by sCD4D1D2 (Table 1). Although we do not know whether sCD4D1D2 represents a good model for native CD4 with respect to CCR5 binding, these results do suggest that specific regions of CD4, potentially those overlapping the epitope of CG7, probably located within the first CD4 domain (32), are involved in the association with CCR5.

CG7 displacement inhibition of MIP-1α

To localize regions of CCR5 interacting with CD4, we used mixtures of mAbs recognizing the N terminus and the first (ecl-1) and second (ecl-2) extracellular loops of CCR5. Increasing the concentration of a mAb against the ecl-2 of CCR5 (2D7) in a mixture with the 5C7 reduced the total amount of coimmunoprecipitated CD4 (Fig. 4D), whereas the amount of immunoprecipitated CCR5 was slightly increased (Fig. 4D), probably resulting from the additional, yet weak immunoprecipitation of CCR5 with 2D7 (Fig. 4E). Another HIV-1-blocking mAb specific for the CCR5 ecl-2 (m182 R&D Systems) showed similar although weaker inhibitory effects (Fig. 4F) in contrast to the non-HIV-1-blocking anti-CCR5 mAb m183, which did not interfere with the CD4–CCR5 coimmunoprecipitation (Fig. 4G). Other mAbs to the N terminus and mAbs to ecl-1 of CCR5 did not decrease the amount of coimmunoprecipitated CD4 when used in combination with 5C7 (data not shown). These results suggest that the ecl-2 of CCR5 is involved in the interaction with CD4. However, even at the highest concentration of mAb (2D7) used (50 μg/ml), the inhibition of the CD4–CCR5 coimmunoprecipitation was not complete, indicating that regions additional to the second extracellular loop of CCR5 are probably involved in the interaction with CD4.

Membrane-Associated CD4 Colocalizes with CCR5.

We further examined whether the CD4 and CCR5 molecules associate by using confocal laser scanning microscopic analysis of fluorescently labeled molecules. Colocalization of the two molecules was observed as demonstrated by the yellow (red-green colocalization) staining, suggesting formation of large multimolecular complexes between these two molecules (Fig. 5). A correlation map was prepared by using software developed in this laboratory. The regions of true overlap of green and red staining were selected from the noncolocalized staining and the background fluorescence and represented as white dots. A relatively high degree of colocalization is evident from this analysis (Fig. 5 Lower). Less colocalization was observed between CD4 and CXCR4 (Fig. 5 Left) than with CCR5. It was previously shown that addition of X4 HIV-1 Env gp120 increases the colocalization of CD4 and CXCR4 (13). The extent of this increase appears to be comparable with the colocalization between CD4 and CCR5 in the absence of gp120 in a manner reminiscent of the immunoprecipitation data shown in Fig. 3. No significant colocalization was observed between CD45 and CCR5 or HLA class I and CCR5, suggesting specificity in the interaction between CD4 and CCR5 (data not shown). These results suggest that CCR5 (and to a lesser extent CXCR4) not only associates with native membrane-associated CD4 but that the two molecules form large multimolecular complexes possibly because of their dimeric structures (ref. 33, and data not shown).

Colocalization of CD4 and CXCR4 (Left) or CCR5 (Right). A CXCR4 (or CCR5)-myc tag-expressing HeLa cell, double stained for CD4 (green) and CXCR4 (or CCR5) (red) (Upper). Colocalization of the two molecules is demonstrated by the yellow (red-green colocalization) staining, suggesting their clusterization. Correlation maps of these images (Bottom) show the regions of true overlap of green and red staining selected from the noncolocalized staining and the background fluorescence, represented as white dots.

Inhibition of the CD4–CCR5 Interaction Correlates with the Inhibition of HIV-1 Env-Mediated Fusion.

It has been demonstrated that the anti-CCR5 mAb 2D7 inhibits entry of R5 and R5X4 HIV-1 into U87MG-CD4 cells expressing transfected CCR5 (24). To investigate the possibility of a relationship between inhibition of the CCR5–CD4 interaction and HIV-1 Env-mediated fusion, we used a recombinant vaccinia virus-based reporter gene assay for quantitation of cell–cell fusion (19). The mAb 2D7 inhibited fusion between 3T3.CD4.CCR5 cells and cells expressing R5 (Bal and JRFL) and R5X4 (89.6) HIV Envs. The inhibition was concentration-dependent, and cell–cell fusion was significantly decreased at concentrations in the range of 0.5–50 μg/ml, a concentration range similar to that observed for inhibition of the CD4–CCR5 interaction (data not shown). For several other anti-CCR5 mAbs, there was correlation (r = 0.98, P = 0.001) between inhibition of fusion and inhibition of the CD4–CCR5 interaction (Table 2). These results indicate that the CD4–CCR5 interaction may play a role in membrane fusion mediated by the HIV-1 Env. However, as was similar to the inhibition of the CD4–CCR5 interaction, even at the highest mAb concentration used (50 μg/ml), the inhibition of fusion was not complete. The lack of complete inhibition of fusion suggests that multiple interactions, possibly both CD4–CCR5 and gp120–CCR5, and multiple interaction sites involving other extracellular regions of CCR5, are involved in the initial steps of HIV-1 entry.

Correlation between inhibition of HIV-1 (Bal)-mediated fusion and CD4–CCR5 interactions by anti-CCR5 mAbs

Materials and methods

Cell-culture reagents

Cells were cultured in RPMI 1640 culture medium supplemented with 10% fetal calf serum (FCS Hyclone Laboratories, Logan, UT), 2 mM l -glutamine, 50 μg/mL gentamicin, 1 mM sodium pyruvate, and 1% nonessential amino acids (complete medium). DCs were stimulated with 1 μg/mL lipopolysaccharide (LPS Escherichia coli 0111:B4 Sigma Chemical, Saint Louis, MO) CD154-transfected J558L cells at a ratio of 4:1 and 5 μM cytosine phosphate guanosine (CpG) oligonucleotide 2216 type A or CpG oligonucleotide 2006 type B (InvivoGen, San Diego, CA). CD4 + T cells were stimulated with anti–human CD3 mAb (clone TR66). Crystalline 1,25(OH)2D3 was a gift of Milan Uskokovic (BioXell, Nutley, NJ), dexamethasone was purchased from Sigma Chemical, and recombinant human (rh) IL-10 from PharMingen (San Diego, CA).

Cell purification

Peripheral-blood mononuclear cells (PBMCs) were isolated from buffy coats by Ficoll gradient (Pharmacia Biotec AB, Uppsala, Sweden). Peripheral-blood myeloid DCs (M-DCs) and plasmacytoid DCs (P-DCs) were magnetically sorted with blood dendritic-cell antigen-1 (BDCA-1) and BDCA-4 cell-isolation kits (Miltenyi Biotec, Bergish Gladblach, Germany), respectively, as described, 27 to a purity of 95% to 98% in both cases. DC subsets were analyzed either immediately or after culture in complete medium in the presence of 10 ng/mL recombinant human granulocyte-macrophage colony-stimulating factor (rhGM-CSF) or 20 ng/mL IL-3 (Pharmingen), respectively. To generate immature monocyte-derived DCs, monocytes, obtained from PBMCs by positive selection with CD14 beads (Miltenyi Biotec), were grown for 6 to 7 days in complete medium plus 10 ng/mL rhGM-CSF and 10 ng/mL IL-4 (PharMingen). CD4 + T cells were purified from PBMCs by negative selection with CD4 T-cell-isolation kit (Miltenyi Biotec), and CD4 + CD25 + T cells were subsequently positively selected with CD25 beads (Miltenyi Biotec).

Flow cytometric analysis

Flow cytometric analysis was performed as previously described, 7 in the presence of 100 μg/mL mouse IgG, using the mAbs anti-CD1c (BDCA-1) fluoroscein isothiocyanate (FITC) or phycoerythrin (PE), anti–BDCA-2 FITC or PE (Miltenyi Biotec), anti-CD1a FITC/PE, and anti-CD83 FITC/PE, all from Pharmingen, or with mAbs specific for ILT1 (clone 135.1), ILT2 (clone GHI75), ILT3 (clone ZM3.1), ILT4 (clone 42D.1), and ILT5 (clone 7H5). Cells were analyzed with an LSR flow cytometer (Becton Dickinson, Mountain View, CA) using CellQuest software (Becton Dickinson).

T-cell activation

Peripheral-blood DC subsets were cultured for 5 days in complete medium and for an additional 7 days in the presence of CD4 + T cells with or without 20 μg/mL anti-ILT3 antibody. Intracellular cytokine production by CD4 T-cell lines was analyzed as described. 28 CD4 + T cells were cultured in 96-well flat-bottom plates with graded amounts of immature monocyte-derived DCs, which had been cultured for the last 48 hours with or without 10 nM 1,25(OH)2D3. The coculture of DCs and T cells was carried out in the presence or absence of anti-ILT3 mAb. After 5 days of culture, interferon-γ (IFN-γ) production was measured by 2-site enzyme-linked immunosorbent assay (ELISA). Alternatively, CD4 + T cells were cultured in the presence of immature monocyte-derived DCs (ratio, 1:10) with or without anti-ILT3 and 7 to 10 days after primary stimulation, were restimulated under the same conditions. After 2 rounds of stimulation, CD4 + T cells were cultured in 96-well round-bottom plates precoated by overnight incubation with anti–human CD3 mAb. After 72 hours, cytokine production was quantified by 2-site ELISA. Paired mAbs specific for IFN-γ, IL-4, and IL-10 (Pharmingen), were used as described. 7 The detection limit for all cytokines was 15 pg/mL.

Suppression assay

The read-out system was composed of CD4 + CD25 – cells from donor A PBMCs labeled with Vybrant CFDA (carboxyfluorescein diacetate) succinimidyl ester (SE) cell tracer kit (Molecular Probes, Leiden, the Netherlands) and cultured in the presence of 1:10 LPS-matured monocyte-derived DCs from donor B. To evaluate the induction of regulatory T cells, graded amounts of CD4 + T cells from donor C were generated by 3 rounds of restimulation with allogeneic immature monocyte-derived DCs from donor D, which were cultured for the last 48 hours with or without 1,25(OH)2D3. The coculture was carried out with or without anti-ILT3 mAb (20 μg/mL). The suppression assay was performed in the presence of 1 μg/mL anti–human CD3 mAb. After 72 to 96 hours of culture, cells were recovered and analyzed with an LSR flow cytometer (Becton Dickinson) using CellQuest software.

Real-time quantitative RT-PCR

RNA was extracted using Trizol (Invitrogen, Carlsbad, CA) according to the manufacturer's instruction, followed by a cleanup with the RNeasy Kit (Qiagen, Hilden, Germany). Reverse transcription (RT) was performed, and real-time quantitative RT–polymerase chain reaction (PCR) of total cDNA using specific primers was carried out using an ABI PRISM 7000 Sequence Detection System (Applied Biosystems, Foster City, CA) and Taqman chemistry. The primers used are commercially available from Applied Biosystems as assays on demand. Relative quantification of target cDNA was determined by arbitrarily setting the control value to 1 and changes in cDNA content of a sample were expressed as a multiple thereof. Differences in cDNA input were corrected by normalizing to β actin or GAPDH (glyceraldehyde-3-phosphate dehydrogenase) signals. To exclude amplification of genomic DNA, RNA samples were treated with DNAse (Sigma Chemical).


Skin biopsies were obtained from untreated psoriatic plaques or from psoriatic lesions of 5 patients treated topically for 30 days with 0.005% calcipotriol cream (Psorcutan Schering, Milan, Italy) twice daily, or for 20 days with 0.1% mometasone furoate cream (Elocon Schering-Plough, Milan, Italy) twice daily, and snap-frozen in liquid nitrogen. Immunohistology was performed on 5-μm-thick cryostat sections fixed in acetone and dried overnight, as described. 29 Before incubation with specific mAbs or control isotype mouse Ig's, sections were hydrated for 10 minutes with 0.05 M Tris (tris(hydroxymethyl)aminomethane)–aminomethane saline buffer (TBS), pH 7.6, and incubated for 10 minutes with a mixture of normal human AB serum and normal rabbit serum (20% in TBS), to minimize nonspecific staining. Then, after washing in TBS, the sections were incubated for 30 minutes with specific mAbs, followed by polyclonal rabbit antimouse serum diluted 1/30 in TBS (Dako-Patt, Copenhagen, Denmark) and alkaline phosphatase anti–alkaline phosphatase complex diluted 1/50 in TBS (Dako-Patt). Finally, stainings were revealed using new fucsin (75 μL of a new fucsin–sodium-nitrite solution) in 10 mL of 0.05 M TBS, pH 8.7 5 mg naphthol AS-BI (7-bromo-3-hydroxy-2-naphtho- o -anisidine) sodium salt and 1 mM levamisole (Sigma Chemical). Sections were counterstained with Mayer haematoxylin, washed in water, and dried at room temperature before mounting. Slides were observed with a Leica light microscope (DMR Leica Microsystems, Milan, Italy) equipped with an HC Plans 10×/2.2 ocular and an N plan 20×/0.40 objective lens.

Immunofluorescent stainings were performed on 5-μm-thick frozen sections from skin biopsies of calcipotriol-treated psoriatic plaques. After fixing for 15 minutes in 4% paraformaldehyde and blocking with 0.1 M glycine for 10 minutes at room temperature, slides were incubated overnight at 4°C in blocking solution with affinity-purified rabbit anti–mouse CD11c (Jackson Immunoresearch, West Grove, PA), washed 5 times with wash buffer (0.45 M NaCl, 0.24 M Na2HPO4, 0.24 M NaH2PO4, and 0.3% Triton X-100), and incubated for 60 minutes with polyclonal antirabbit FITC (Sigma Chemical). Alternatively, slides were incubated overnight at 4°C with biotinylated rat anti–mouse CD123 (Jackson Immunoresearch) followed by Rhodamine Red-X-streptavidin (Jackson Immunoresearch). Negative controls were performed by incubation with appropriate isotype-matched primary antibodies. The slides were then washed again and mounted with 90% glycerol/phosphate-buffered saline (PBS). Slides were analyzed with an MRC-1024 laser-scanning confocal microscope (Bio-Rad Laboratories, Hercules, CA). Images were acquired and processed with Laser Sharp 3.2 software (Bio-Rad).

1,25(OH)2D3 enhances ILT3 expression on DCs. Monocyte-derived human DCs were incubated for 48 hours with medium alone or containing 10 nM 1,25(OH)2D3 or 10 ng/mL IL-10, either when immature or during LPS-induced (100 ng/mL) maturation. Surface expression of the indicated ILT molecules was determined by cytofluorimetry. Histograms represented by broken lines show staining with an isotype control those with thick lines show staining with the indicated anti-ILT mAbs. Geometric mean fluorescence intensity (MFI) values are shown in the top right corner. A representative experiment out of 6 performed is shown.

1,25(OH)2D3 enhances ILT3 expression on DCs. Monocyte-derived human DCs were incubated for 48 hours with medium alone or containing 10 nM 1,25(OH)2D3 or 10 ng/mL IL-10, either when immature or during LPS-induced (100 ng/mL) maturation. Surface expression of the indicated ILT molecules was determined by cytofluorimetry. Histograms represented by broken lines show staining with an isotype control those with thick lines show staining with the indicated anti-ILT mAbs. Geometric mean fluorescence intensity (MFI) values are shown in the top right corner. A representative experiment out of 6 performed is shown.


Contribution: J.M.M. designed and performed experiments, analyzed results, made figures, and wrote the manuscript B.R.L. designed experiments, participated in discussion of the results, and reviewed the paper J.E.S.-C. designed experiments and reviewed the paper A.J.C. performed experiments and reviewed the paper V.A.Y. performed experiments L.C.N., J.M., and D.F.N. participated in discussion of the results and reviewed the paper and L.L.L. participated in discussion of the results and edited the paper.

Conflict-of-interest disclosure: The authors declare no competing financial interests.


A novel anti-TLR5 monoclonal antibody, ACT5, revealed the cell surface expression of TLR5 on immune cells. The recruitment of TLR5 to the cell surface was completely dependent on TLR-specific chaperone PRAT4A. In Ba/F3 cells, PRAT4A physically associated with TLR5. PRAT4A knock-down resulted in marked suppression of NF-κB activation in response to flagellin. In addition, in a macrophage cell line J774, PRAT4A silencing abolished cell surface expression of endogenous TLR5 and flagellin-dependent cytokine production. These results suggest that PRAT4A-dependent cell surface expression of TLR5 is required for flagellin-induced cytokine production. Previous studies report that PRAT4A associates with a general chaperone gp96 and restricts gp96 client to TLRs ( 21). Taken together with the requirement of gp96 for flagellin response ( 20), the present results demonstrate that TLR5 is another client of the chaperone complex PRAT4A–gp96.

It is well established that TLR5 activates MyD88-dependent signaling pathways to induce cytokine production and DC maturation ( 4, 12). While cell surface TLR2/4-signaling requires a ‘sorting adaptor’, TIRAP, to recruit MyD88 to the cell surface TLR ( 17), a previous report demonstrated that TLR5 does not need TIRAP for cytokine production ( 16). Here, we confirm that TLR5 is another cell surface TLR, suggesting that MyD88 pathway from cell surface does not necessarily require TIRAP. TIRAP-independent MyD88 signaling from cell surface TLR5 may be a reason why flagellin, in contrast to LPS, is poor inflammatory and less relation to severe organ failure ( 37).

ACT5 revealed highly restricted expression of TLR5 on in vivo immune cells. Cell surface TLR5 was mainly detected on neutrophils, Ly6C hi classical monocytes, splenic CD8 − CD4 + /CD8 − CD4 − cDCs, splenic pDCs and CD11b hi CD11c hi lamina propria DCs. In acute inflammatory lesion, neutrophils and monocytes are swiftly recruited from the circulation and have a role in primary defense against microbial organisms. Neutrophils, Ly6C hi and Ly6C low monocytes infiltrated into the peritoneal cavity 12h after thioglycollate injection. Cell surface expression of TLR5 was detected on neutrophils and Ly6C hi classical monocytes, but not on Ly6C low monocytes. At 72h after thioglycollate injection, almost all exudate cells were TLR5 − Ly6C low monocytes. These findings indicate that cell surface TLR5 is one of the hallmarks of innate immune cells that are recruited early in inflammation.

All TLRs except TLR3 are expressed in human neutrophils and induce cytokine release, superoxide generation and CD62L shedding ( 38). However, neutrophils from mouse BM or an inflammatory lesion did not produce cytokines in response to flagellin despite the expression of TLR5 on the cell surface. Given a role for TLR5 in phagocytosis and ROS production ( 39), cell surface TLR5 in neutrophils may principally contribute to direct pathogen clearance rather than to cytokine production.

Based on cell surface markers, mice monocytes are subcategorized into two major subsets ( 40). The Ly6C hi CCR2 hi CX3CR1 low subset is called classical monocytes, which are recruited to a site of tissue injury or infection via CCR2-signaling and differentiate into macrophages or DCs. The other Ly6C low CCR2 low CX3CR1 hi subset, nonclassical monocytes, has a remarkable patrolling activity and gives rise to resident macrophages after entering tissues. This study showed that TLR5 is expressed on Ly6C hi , but not on Ly6C low monocytes in the BM, circulation and inflammatory lesions. Flagellin is able to induce IL-6 and MCP-1 production via cell surface TLR5 from Ly6C hi monocytes, suggesting that classical monocytes are one of the main sources of flagellin-induced cytokines in acute inflammation. Considering that Ly6C hi classical monocytes contributes to the gut immune system and have a role in protection against an enteric bacterium ( 41), flagellin-induced cytokines from classical monocytes have an important role in augmenting acute inflammation at mucosa.

In addition to neutrophils and classical monocytes, very restricted expression of cell surface TLR5 was also found in DCs. First, cell surface TLR5 was detected on CD8α − cDCs, but not on CD8α + cDCs, in the spleen. CD8α − cDCs are more potent in activating CD4 + helper T cells than CD8α + cDCs, and associate with induction of Th2-type response ( 42). Considering that flagellin has been shown to have the potent adjuvant activity with a tendency to induce Th2 response ( 12, 28), CD8α – cDCs, but not CD8α + cDCs, are likely to be required for the adjuvant activity of flagellin. Unexpectedly, splenic plasmacytoid DCs (pDCs), which is specialized to secrete large amounts of type I interferon, also express cell surface TLR5. As one report suggests that flagellin can induce IFN-β from BM-macrophages ( 43), flagellin may induce or augment the production of type I interferon in pDCs.

Against oral infection with a flagellated pathogen, TLR5 has a role in mounting adaptive immune responses ( 10, 36). CD11b hi CD11c hi DCs in the lamina propria express TLR5 mRNA and have a key role in activating flagellin-dependent adaptive immune responses. Consistent with these previous reports, our results showed that only CD11b hi CD11c hi DCs expressed TLR5 on the cell surface. In contrast to these in vivo DCs, TLR5 is barely detectable on BM-DCs. Similarly, TLR5 is detectable on Ly6C hi monocytes in vivo but not on BM-macrophages. These results reveal that BM-derived DCs/macrophages are distinct from in vivo DCs/macrophages. Unlike TLR2 and TLR4/MD-2, in vitro maturation induced by GM-CSF or M-CSF does not accompany cell surface expression of TLR5, implying that a signal is missing in in vitro maturation that induces cell surface TLR5.

In summary, our results show that TLR5 is expressed on the cell surface of neutrophils, classical monocytes and specific subpopulations of DCs, in a PRAT4A-dependent manner. The present result, however, does not necessarily exclude the possibility that intracellular TLR5 is functional. Although BM-DCs did not express cell surface TLR5, BM-DCs could respond to flagellin and induce DC maturation ( 12, 35). In the case of TLR4/MD-2, intracellular TLR4/MD-2 in BM-macrophages is responsible for upregulation of costimulatory molecules and certain chemokine production ( 22). Future studies need to focus on TLR5 inside the cells.

Japanese-Korean Cooperative Programme on Basic Medical Research Ministry of Education, Culture, Sports, Science and Technology for Program of Japan Initiative for Global Research Network on Infectious Diseases Grant-in-Aid for Scientific Research (B), grant reference 21117001 Grant-in-Aid for Exploratory Research Grant-in-Aid for Scientific Research on Innovative Areas, grant reference 23790526.

Classical monocytes, but not neutrophils, are the predominant source of flagellin-induced cytokines. (A) Gating strategies to sort neutrophils and classical monocytes from BM cells by Gr-1 and Ly6G staining. Red dots mean total CD11b + BM cells. Overlaid blue and green dots on red dots mean Ly6C int Ly6G + neutrophils and Ly6C hi Ly6G - classical monocytes, respectively (left dot plots). Since neutrophils and classical monocytes matched to Gr-1 hi Ly6G + and Gr-1 hi Ly6G – population (right dot plots), respectively, these populations were sorted and used for cytokine assay. (B and C) Sorted classical monocytes (B) and neutrophils (C) from BM cells were stimulated by 100ng ml –1 flagellin and cytokine production was analyzed by ELISA. Asterisks mean no signals were detected.

Classical monocytes, but not neutrophils, are the predominant source of flagellin-induced cytokines. (A) Gating strategies to sort neutrophils and classical monocytes from BM cells by Gr-1 and Ly6G staining. Red dots mean total CD11b + BM cells. Overlaid blue and green dots on red dots mean Ly6C int Ly6G + neutrophils and Ly6C hi Ly6G - classical monocytes, respectively (left dot plots). Since neutrophils and classical monocytes matched to Gr-1 hi Ly6G + and Gr-1 hi Ly6G – population (right dot plots), respectively, these populations were sorted and used for cytokine assay. (B and C) Sorted classical monocytes (B) and neutrophils (C) from BM cells were stimulated by 100ng ml –1 flagellin and cytokine production was analyzed by ELISA. Asterisks mean no signals were detected.