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,¶
,¶
*
Institute for Immunology, University of Muenchen, and
GSF National Research Center for Environment and Health, Institute of Molecular Immunology, Muenchen, Germany;
Clinical Cooperation Group, Aerosols in Medicine, Institute for Inhalationbiology, GSF National Research Center for Environment and Health and Asklepios-Fachkliniken Muenchen-Gauting, Gauting, Germany;
Institute of Cancer Research and Molecular Biology, Medisinsk Teknisk Senter, Trondheim, Norway; and
¶ Department for Microbiology and Immunology, University of Leicester, Leicester, United Kingdom
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The cells of the monocyte lineage derive from myelomonocytic stem cells
in bone marrow. They mature to monocytes and, as such, they go into
blood followed by migration into tissue. In tissue these cells, now
termed macrophages, differentiate into phenotypically and functionally
distinct cell types like alveolar macrophages, osteoclasts, or
microglia cells. While the heterogeneity of these tissue macrophages is
well established, monocyte heterogeneity has been clearly demonstrated
only recently. Using mAbs and flow cytometry we could distinguish the
classical CD14++ cells and the
CD14+CD16+ monocytes
(2). In healthy donors these cells account for
50 cells/µl and 10% of all monocytes. The
CD14+CD16+ monocytes will
increase dramatically in patients with severe infection
(3, 4, 5), and excessive exercise will also increase the
cells by factor 4 due to mobilization from the marginal pool
(6). The
CD14+CD16+ monocytes appear
to be more mature, they express higher levels of HLA-DR Ags as compared
with classical monocytes (7), and they show expression of
a distinct pattern of cytokines. Specifically, the
CD14+CD16+ cells have been
shown to efficiently produce the proinflammatory cytokine TNF, while
they produce no or little of the anti-inflammatory cytokine IL-10
(8). Based on these findings we have termed these cells
proinflammatory monocytes.
Cytokine expression by human monocytes previously has been studied after isolation of the cells by density gradient separation, Ab staining, and cell sorting. Because these procedures may alter the function of these cells we have developed in the current study a system that involves stimulation of whole peripheral blood (WPB) followed by staining and analysis of cytokine expression. This system allows for a more meaningful analysis of the regulatory mechanisms involved in cytokine gene expression. Using this method we can demonstrate a higher level of TNF protein expression of the CD14+CD16+ monocytes as compared with the classical CD14++ cells. Also, depletion of these cells from PBMC populations will reduce the amount of Pam3Cys induced TNF in the supernatant by >60%, although the CD14+CD16+ monocytes account for only 10% of the monocytes. This suggests that in vitro, and even more so in vivo, during bacterial infection the minor population of CD14+CD16+ monocytes may be a major source of inflammatory cytokines like TNF.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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WPB anticoagulated with heparin at 10 U/ml was obtained from apparently healthy donors who were recruited from institute personnel and from students. Six hundred-microliter samples were incubated for 6 h at 37°C in 15-ml polypropylene tubes (catalog no. 188 261; Greiner, Frickenhausen, Germany) in the presence of brefeldin A (BFA) at 10 µg/ml final (catalog no. B-7651; Sigma, Muenchen, Germany). Samples were left untreated or were stimulated with LPS from Salmonella minnesota (catalog no. L-6261; Sigma) at 0.1–1000 ng/ml or with Pam3Cys (catalog no. L2000; ECM Microcollections, Tuebingen, Germany) at 1–100 µg/ml. For control of LPS contamination cultures were set up with and without polymyxin B (PMB; Pfizer, Karlsruhe, Germany) at 10 µg/ml. Also, using the Endosafe Limulus Gel Clot assay (catalog no. AE 010; Charles River Breeding Laboratories, Sulzfeld, Germany) (sensitivity: 0.06 EU/ml) for monitoring of our tissue culture reagents, including BFA and heparin, we found no detectable LPS.
Cell surface staining
The stimulated samples were treated with ammonium chloride
buffer (0.83% w/v) for 3 min until erythrocytes were lysed. After
washing the leukocytes with staining buffer (PBS/2% FCS), cells were
resuspended in 100 µl of staining buffer and Abs to CD14 (My-4-FITC
at 18 µg/ml final concentration; catalog no. 6603511; Beckman
Coulter, Krefeld, Germany) and HLA-DR (Immu357-PC-5 at 0.06 µg/ml
final concentration; catalog no. 2659; Beckman Coulter) were added
followed by incubation on ice for 20 min. After washing the samples
with staining buffer the cells were fixed with paraformaldehyde at 4%
in PBS for 20 min at room temperature followed by two wash steps. The
CD16 Ag was detected using Ab Leu11c-PE at 6
µg/ml final concentration (catalog no. 347617; BD Biosciences,
Heidelberg, Germany).
Staining for TLRs
For staining of TLR2 the Ab TL2.1 (IgG 2a) (9) was used as Alexa 488 conjugate along with an Alexa 488 isotype control both at 10 µg/ml final concentration.
Intracellular staining
Paraformaldehyde-fixed samples were permeabilized with Perm/Wash solution (catalog no. 2097 KZ; BD Biosciences) for 5 min at room temperature. For detection of intracellular TNF (icTNF) we added PE-conjugated anti-TNF Ab (MP9–20A4, catalog no. RHTNFA04; Caltag Laboratories via Medac, Hamburg, Germany) or as isotype control PE-conjugated rat IgG1 (catalog no. R104; Caltag Laboratories) both at 10 µg/ml for 20 min on ice. Samples were then washed twice and were resuspended in PBS/0.5% paraformaldehyde to be analyzed within 24 h.
Specificity control
The specificity of TNF staining was determined by incubating the anti-TNF-PE conjugate for 10 min at room temperature with a 9-fold molar excess of recombinant human TNF (kindly provided by BASF-Knoll, Ludwigshafen, Germany). This mixture of Ab and cytokine was the added to the permeabilized cells. During our studies we noted that the intensity of icTNF staining in monocytes shows variability between individuals and over time. Therefore, data were interpreted always within a given experimental series.
Flow cytometry analysis
Analysis was done on a FACScan flow cytometer (BD Biosciences)
and linearity of the instrument was shown by using Immuno-Brite
standard beads (catalog no. PN 6603473; Beckman Coulter). Cells were
analyzed by setting scatter gates around monocytes and a portion of the
adjacent lymphocytes (see www.monocytes.de for further details). At
least 500
CD14+DR++CD16+
monocytes were analyzed per sample and fluorescence intensities were
determined by subtracting the median fluorescence intensity of the
isotype control from the median fluorescence intensity of the specific
Ab. The resultant channels of specific median fluorescence intensity
were used as a semiquantitative measure of receptor expression. For
icTNF staining (see below) the specific (or 
= 
) median
fluorescence intensity was calculated by subtracting the signal for
anti-TNF staining in the presence of excess rTNF from the specific
staining.
Quantitative PCR
Quantitative PCR was performed using the LightCycler system (Roche Diagnostics, Mannheim, Germany) according to the manufacturer’s instructions by using primer pairs as noted below. In brief, mRNA was isolated and reverse-transcribed as for conventional RT-PCR. A total of 3 µl of cDNA were used for amplification in the SYBR green format using the LightCycler FastStart DNA Master SYBR Green I kit from Roche Diagnostics (catalog no. 2 239 264). For quantitative PCR, the LightCycler system offers the advantage of speed and real-time measurement of fluorescent signal during amplification. The SYBR green dye binds specifically to the minor groove of dsDNA. Fluorescence intensity is measured after each amplification cycle. During PCR, a doubling of template molecules occurs in each cycle only during the log-linear phase. Although the LightCycler displays signals from every cycle, the instrument uses fluorescent signals only generated during this informative log-linear phase to calculate the relative amount of template DNA.
The following primer pairs were used: TLR2 5' primer,
5'-GCCAAAGTCTTGATTGATTGG-3', and 3' primer,
5'-TTGAAGTTCTCCAGCTCCTG-3' (product length: 394 bp); TLR4 5'
primer, 5'-GAAATGGAGGCACCCCTTC-3', and 3' primer,
5'-TGGATACGTTTCCTTATAAG-3' (product length: 510 bp). As an internal
control the housekeeping gene 
-enolase was amplified.
Depletion of CD16+ monocytes
For depletion of CD14+CD16+ monocytes from PBMC were isolated by Ficoll-Hypaque (Pharmacia, Freiburg, Germany) density gradient separation. A total of 10 x 106 cells each were resuspended in 100 µl of PBS containing 5 µl of CD16 microbeads (catalog no. 130-045-701; Miltenyi Biotec, Bergisch-Gladbach, Germany) or for control CD8 microbeads (catalog no. 130-045-201; Miltenyi Biotec). After incubation for 30 min at 4°C cells were washed and resuspended in 0.5 ml PBS and this was loaded onto a RS column (catalog no. 130-042-201; Miltenyi Biotec) that was positioned in a MiniMACS magnet (catalog no. 130-042-102; Miltenyi Biotec). Nonadherent cells were recovered and total number of CD14-positive monocytes and percentage of CD14+CD16+ monocytes were determined by FACS. Cells were then incubated for 5 h at 37°C without or with stimulation by either Pam3Cys (2 µg/ml) or LPS (2 ng/ml). Supernatant TNF was determined by ELISA (PeliKine, catalog no. M1923; Hiss Diagnostics, Freiburg, Germany) and was expressed in picograms per 106 monocytes. Average depletion of CD14+CD16+ monocytes with this procedure was 94%.
Culture of isolated CD14++ monocytes for demonstration of the stability of phenotype
For culture of classical monocytes, PBMC were isolated by Ficoll-Hypaque density gradient separation and were depleted of CD16-positive cells using CD16 microbead Abs and MACS as given in the preceding paragraph. Depleted cells were then enriched for classical monocytes using CD14 microbead Ab (catalog no. 130-050-201; Miltenyi Biotec). Purified cells were then cultured in the presence of BFA (10 µg/ml) for 6 h without and with LPS at 100 ng/ml. The monocytes were then stained for CD14-PE (catalog no. 6603262; Beckman Coulter) and HLA-DR-PE-Cyan5 conjugate (catalog no. 2659; Beckman Coulter) and analyzed by FACS.
Statistics
For statistical analysis Student’s t test was used.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Monocyte subpopulations can be readily defined as
CD14+CD16+ and
CD14++CD16 cells using CD14 and CD16 Abs in
two-color immunofluorescence. However, when WPB cells are stimulated
with LPS for several hours, then the CD16 cell surface molecule is
down-regulated such that the
CD14+CD16+ monocytes cannot
be clearly identified anymore (data not shown). Under the same
conditions cells coexpressing low levels of CD14 plus high levels of
HLA-DR retained their staining pattern. To demonstrate that
theseCD14+DR++
cells are identical to the
CD14+CD16+ monocytes we
performed three-color immunofluorescence analyses. In the two-color
plot for CD14 and DR (Fig. 1
, middle panel) a population that is DR-only (DR-positive
lymphocytes), a population coexpressing high levels of CD14 and low
levels of DR (CD14++DR+
classical monocytes), and a population with the
CD14+DR++ phenotype are
visible. When the CD14+DR++
cells are gated and analyzed for CD16, the majority of cells were found
CD16 positive (Fig. 1, upper panel).
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, lower
panel) (average 2.1 ± 0.5%). Thus gating on the
CD14+DR++ population can be
used as an alternative approach for the analysis of the
CD14+CD16+ monocytes. To
further substantiate this point we have isolated
CD14++DR+ monocytes and
have cultured them in the presence of BFA without and with LPS for
6 h. The data in Fig. 2 show that
there is no relevant generation of
CD14+DR++ cells from
CD14++DR+ monocytes under
these conditions.
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We then stimulated whole blood samples with LPS at 100 ng/ml for
6 h in the presence of BFA and stained the cells for CD14, DR, and
TNF. A typical example, shown in Fig. 3, gives the TNF expression for the two monocyte populations as depicted
in Fig. 3, middle panel. The single parameter histograms
(Fig. 3, upper and lower panels) give the signal
obtained with the anti-TNF-Ab (Fig. 3, upper and lower
panels, right curve) as compared with the negative control,
which is the anti-TNF staining in the presence of a 9-fold molar
excess of rTNF (Fig. 3, upper and lower panels, left curve).
It demonstrates a strong expression of TNF by both monocyte
populations, but the
CD14+DR++ monocytes exhibit
a clearly higher level of expression. In five donors, the
average median fluorescence intensity was 37 ± 18.8 in the
CD14++ monocytes while the
CD14+DR++ monocytes
expressed TNF with a median fluorescence intensity of 125.4 ±
55.1 channels (p < 0.05); i.e., the signal for
TNF protein was 3-fold higher in these cells.
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). While at all time
points the CD14+DR++
monocytes exhibited a higher immunofluorescence signal for TNF, this
difference was only significant for the 4- and 6-h time points.
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, these cells
did, however, show a first response at the 1 ng/ml dose similar to the
classical CD14++DR+ cells.
At this dose the cells produced TNF with an average median of 17
channels, expression increased to 261 channels at 100 ng/ml and reached
365 channels at the highest dose of 1000 ng/ml. Again, TNF
expression was clearly higher in the
CD14+DR++ monocytes as
compared with the classical monocytes at all doses (Fig. 5). Hence,
when comparing the two monocyte populations, the
CD14+DR++ monocytes respond
at the same low dose of LPS and the response was even higher despite
the low-level expression of the CD14 LPS receptor.
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We next asked whether the higher expression of TNF in the
CD14+DR++ monocytes is also
seen after stimulation with other microbial products like Pam3Cys. To
exclude the possibility that a LPS contamination of the Pam3Cys
preparation causes monocyte stimulation we have used PMB for LPS
neutralization. PMB, in fact, is very active in that it completely
abrogates the LPS-induced TNF production (Fig. 6, lower panel). When looking
at TNF production by the two monocyte subsets we observed that Pam3Cys
almost exclusively stimulated the
CD14+DR++ proinflammatory
monocytes. While the classical
CD14++DR+ monocytes
responded minimally with an average median immunofluorescence of
28 ± 12 channels, the proinflammatory monocytes exhibited an
average median immunofluorescence intensity of 304 ± 166 channels
(Fig. 6, upper panel). Admixture of PMB to the Pam3Cys did
not appreciably affect the activity, thereby excluding a significant
LPS contamination. Hence, the proinflammatory monocytes show a 10-fold
signal for TNF protein after stimulation with Pam3Cys. In dose response
analysis Pam3Cys efficiently stimulated TNF expression down to a dose
of 1 µg/ml. Even at this low dose the
CD14+DR++ cells gave a
median specific immunofluorescence signal for TNF at 109 ± 68
channels compared with 13 ± 6 channels for the classical
CD14++DR+ monocytes.
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Staining for TLR4 using Ab HTA-125 (10) in
three-color immunofluorescence in our hands revealed no signal both on
the CD14++ monocytes and the
CD14+CD16+ monocytes (data
not shown). By contrast, TLR2 was clearly detectable and gave a
2-fold-higher specific median fluorescence intensity level in the
CD14+CD16+ monocytes as
compared with the classical CD14++ cells
(Fig. 7). In five donors the average
signal for TLR2 in
CD14+CD16+ monocytes was
58.7 ± 9.1 channels as compared with 33.4 ± 9.8 channels
for the classical CD14++ monocytes
(p < 0.05), reflecting a 2-fold level of
specific median immunofluorescence intensity. A similar pattern was
seen when the monocyte subsets were identified as
CD14+DR++ and
CD14++DR+ cells (data not
shown).
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the
CD14+CD16+ monocytes
exhibited slightly lower levels of mRNA for TLR2 and almost identical
levels for TLR4. In four donors the average mRNA level was
calculated to be 1.7-fold lower for TLR2 (p <
0.05) and 1.2-fold lower for TLR4 (not significant). Thus
CD14+CD16+ proinflammatory
monocytes do not express higher amounts of TLR mRNA; rather, they tend
to show slightly lower expression as compared with the classical
monocytes.
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Because the
CD14+CD16+DR++
monocytes are much more efficient producers of TNF as detected by
intracellular staining we asked whether these cells also contributed a
major portion to supernatant TNF secreted by stimulated PBMC. For this
we reacted the PBMC with a microbead-conjugated CD16 Ab and depleted
these cells using MACS. Upon LPS stimulation this led to a reduction of
supernatant TNF by 28% (p < 0.05) as compared
with control depletion with a CD8 Ab (Fig. 9). When stimulating the same cell
preparations with Pam3Cys the CD16 depletion resulted in a much
stronger reduction of secreted TNF, i.e., by 64%
(p < 0.005). These data show that the minor
population of CD14+CD16+
monocytes, which form 10% of all monocytes in blood, account for
the major proportion of the TNF protein that is secreted after
stimulation via TLR2.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Analysis of monocytes directly in WPB can avoid these problems. We therefore have developed a system that employs stimulation of WPB followed by identification of monocytes with Abs to cell surface markers and by detection of cytokines with intracellular staining. The direct addition of stimuli and inhibitors to the freshly drawn blood then allows for the study of cytokine expression unbiased by any manipulation.
With appropriate cell surface markers monocytes can be subdivided into two populations, i.e., the CD14+CD16+ and the CD14++ monocytes (2). These cells show a differential pattern of cytokine expression in that the CD14+CD16+ monocytes fail to produce significant amounts of the anti-inflammatory cytokine IL-10 while they are good producers of the proinflammatory TNF (8). This has led us to term these cells proinflammatory monocytes.
The earlier studies on cytokine expression by monocyte
subpopulations were done with cells that were first stained for cell
surface markers, then sorted, then stimulated with LPS. The current
protocol involves initial stimulation followed by staining. However,
LPS stimulation leads to the disappearance of the CD16 molecule from
the monocyte cell surface such that the
CD14+CD16+ monocytes cannot
be readily identified anymore. The
CD14+CD16+ monocytes are
also characterized by a high level of HLA-DR and expression of this
molecule is maintained with LPS stimulation. When purified
CD14++DR+ cells with 0.71%
remaining CD14+DR++ cells
were cultured for 6 h in the presence of LPS, there was only a
slight increase of the
CD14+DR++ cells to 1.59%
(n = 3). This indicates that the 10%
CD14+DR++ cells that we
analyze for TNF production in whole blood after 6 h of stimulation
with LPS are not derived from the CD14++DR+.
Hence, cytokine expression of the CD14+CD16+
monocytes can be analyzed by looking at the
CD14+DR++ cells, and this
can be compared with the
CD14++DR+ cells that are
identical with the classical CD14++
monocytes.
Previous studies with isolation of monocytes followed by stimulation have demonstrated comparable levels of TNF mRNA and secreted protein for the two monocyte populations after stimulation by LPS (8). Using the WPB stimulation and staining strategy we can demonstrate a 3-fold higher level of LPS-induced TNF protein in the CD14+DR++ monocytes as compared with the classical monocytes. The detection of higher levels of TNF protein in the current study might be explained by advantages of this system, like the absence of interfering signals due to purification, or it could be due to the higher precision of the flow cytometry method.
Time course analysis revealed a very rapid induction of TNF protein at
0.5 h post-LPS stimulation, and this is most obvious for the
CD14+DR++ monocytes. A
maximum TNF production is observed at 6 h, and when prolonging the
incubation to 12 h there is still some increase (data not shown).
However, for practical reasons we have restricted our analysis to the
6-h time point. When looking at dose response at this point in time we
were surprised to see a pronounced TNF production already at 1 ng/ml
LPS in the CD14+DR++
monocytes while the
CD14++DR+ cells still were
close to background (see Fig. 5).
In addition to LPS a dramatically higher response with respect to
TNF was seen with Pam3Cys, a product of Borrelia and of
other types of bacteria (12, 13, 14, 15, 16). Also, Zymosan derived
from yeast and mycobacterial lipoarabinomannan both induced 2-fold
higher levels of specific median immunofluorescence intensity for TNF
in the proinflammatory monocytes (data not shown). The higher
expression of TNF by the
CD14+CD16+ monocytes might
be due to a higher expression of cell surface receptors for these
different microbial products. However, the pattern recognition receptor
CD14 is lower in the
CD14+CD16+ subset
(17) (see Fig. 1). Hence, we have looked for the TLR
coreceptors (1, 18, 19, 20, 21, 22, 23). LPS responses are mediated by the
TLR4 coreceptor. The Ab HTA-125 (10) in our hands
unfortunately gave no staining on blood monocytes. Because LPS
responses are mediated by TLR4 and because the blood monocytes showed
high levels of TNF expression after stimulation with S.
minnesota LPS in our experiments we expect that with an
appropriate reagent TLR4 protein will be detectable on both monocyte
subpopulations.
By contrast, levels of TLR2, the coreceptor used by Pam3Cys (18), were readily detected and were 2-fold higher in the immunofluorescence signal in the CD14+CD16+ monocytes. When we assume that the level of TLR2 is crucial in determining the strength of cell activation, then the higher TLR2 level may, in fact, be responsible in part for the higher response of the CD14+CD16+DR++ monocytes to Pam3Cys.
Of note, the LPS preparation used in this study has not been repurified (24) and therefore it may also partially act via TLR2 due to contaminant lipoproteins.
Based on the higher expression of TNF protein in the
CD14+CD16+DR++
monocytes as detected by intracellular staining we may assume that
these cells, with their proportion of 10% among all monocytes, could
account for a large part of the TNF secreted by stimulated PBMC. Our
studies, in fact, demonstrate that depletion of these cells by MACS
will reduce secreted TNF by 30%, consistent with a 3-fold-higher
icTNF level in the
CD14+DR++ cells as compared
with the CD14++DR+
monocytes. When stimulated by Pam3Cys the reduction by depletion of
CD16+ cells was more pronounced (60%), again
consistent with the 10-fold-higher levels of icTNF seen with Pam3Cys.
Hence, for Pam3Cys stimulation the minor population of
CD14+CD16+DR++
monocytes is responsible for the major part of secreted TNF.
The blood drawn from donors at rest does not cover all leukocytes in circulation because it is restricted to the central pool. A fair amount of additional cells resides in the marginal pool, which comprises cells that are loosely attached to endothelium. These cells can be mobilized, for instance, by stress or excessive exercise. We reported earlier that compared with the CD14+CD16+ monocytes in the central pool there are three times the number of these cells found in the marginal pool (6). Hence, when looking at total monocytes in circulation (marginal plus central pool), the CD14+CD16+ monocytes account for 30% of all monocytes. When bacteria or their products seed into blood they will stimulate all CD14+CD16+ monocytes, including those in the central and in the marginal pool. Hence, in vivo the importance of the proinflammatory monocytes is even higher than demonstrated herein for cell preparations derived from the central pool of blood.
Our data suggest that the CD14+CD16+ monocytes contribute significantly to LPS-stimulated cytokine production that occurs in Gram-negative infection. The contribution is more pronounced when it comes to Pam3Cys, a compound that represents lipopeptides that are expressed by both Gram-positive and Gram-negative bacteria (12, 13, 14, 15, 16). Hence, lipopeptide-induced cytokines may contribute to any type of bacterial infection. However, while in Gram-negative infection the effects of LPS may predominate, the lipopeptides may be dominant in TLR2-mediated monocyte activation in Gram-positive infection. Based on the present findings, which demonstrate the CD14+CD16+ monocytes to be the main source or Pam3Cys-stimulated TNF production, we suggest that these proinflammatory monocytes are major players in Gram-positive infection.
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Footnotes |
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2 M.S. is on leave from the Department of Clinical Immunology, Jagiellonian University Medical College, Krakow, Poland.
3 Address correspondence and reprint requests Dr. Löms Ziegler-Heitbrock, Department for Microbiology and Immunology, University of Leicester, University Road, Leicester, LE1 9HN, U.K. E-mail address: ziehei{at}gmx.de
4 Abbreviations used in this paper: Pam3Cys, S-(2,3-bis(palmitoyloxy)-(2-RS)-propyl)-N-palmitoyl-(R)-Cys-(S)-Ser-(S)-Lys4-OH,trihydrochloride; BFA, brefeldin A; PMB, polymyxin B; TLR, Toll-like receptor; icTNF, intracellular TNF; WPB, whole peripheral blood.
Received for publication August 6, 2001. Accepted for publication January 25, 2002.
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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P. Ancuta, P. Autissier, A. Wurcel, T. Zaman, D. Stone, and D. Gabuzda CD16+ Monocyte-Derived Macrophages Activate Resting T Cells for HIV Infection by Producing CCR3 and CCR4 Ligands J. Immunol., May 15, 2006; 176(10): 5760 - 5771. [Abstract] [Full Text] [PDF]
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G. Soares, A. Barral, J. M. Costa, M. Barral-Netto, and J. Van Weyenbergh CD16+ monocytes in human cutaneous leishmaniasis: increased ex vivo levels and correlation with clinical data J. Leukoc. Biol., January 1, 2006; 79(1): 36 - 39. [Abstract] [Full Text] [PDF]
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F. Suzuki, T. Nanki, T. Imai, H. Kikuchi, S. Hirohata, H. Kohsaka, and N. Miyasaka Inhibition of CX3CL1 (Fractalkine) Improves Experimental Autoimmune Myositis in SJL/J Mice J. Immunol., November 15, 2005; 175(10): 6987 - 6996. [Abstract] [Full Text] [PDF]
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