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*
The Walter and Eliza Hall Institute of Medical Research, Melbourne, Victoria, Australia; and
The Edward Jenner Institute for Vaccine Research, Compton, Newbury, United Kingdom
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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-,
CD4+8-, and
CD4-8+ DC. We have tested whether these
represent stages of development or activation within one DC lineage, or
whether they represent separate DC lineages. All three DC subtypes
appear relatively mature by many criteria, but all retain a capacity to
phagocytose particulate material in vivo. Although further maturation
or activation could be induced by bacterially derived stimuli,
phagocytic capacity was retained, and no DC subtype was converted to
the other. Continuous elimination of CD4+8- DC
by Ab depletion had no effect on the levels of the other DC subtypes.
Bromodeoxyuridine labeling experiments indicated that all three DC
subtypes have a rapid turnover (half-life, 1.5–2.9 days) in the
spleen, with none being the precursor of another. The three DC subtypes
showed different kinetics of development from bone marrow precursors.
The CD8+ spleen DC, apparently the most mature,
displayed an extremely rapid turnover based on bromodeoxyuridine uptake
and the fastest generation from bone marrow precursors. In conclusion,
the three splenic DC subtypes behave as rapidly turning over products
of three independent developmental streams. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Recently, we found that the DC of mouse spleen can be separated into
three distinct populations by employing surface markers more commonly
used to segregate T cells, namely CD4 and CD8 (10). The
surface staining of splenic DC for CD4 or CD8 was shown to represent
authentic expression of these genes by the DC themselves, rather than
pick-up of Ag from T cells or autofluorescence. This was an important
issue, because the DC from other sources, such as the thymus, were
found to be especially subject to artifacts of this type
(10). The CD8, when expressed, was in the form of an
homodimer; none of the DC populations expressed CD8ß. The
three spleen DC populations were shown to have the surface phenotype
CD4+8-11b+205-,
CD4-8-11b+205-,
and
CD4-8+11b-205+,
respectively. The CD4+8-
population could be further distinguished by the expression of moderate
levels of F4/80 and by short term adherence to glass or plastic
surfaces (10). All three populations were efficient
at activating allogenic T cells into cell cycle in a conventional mixed
leukocyte culture, so all populations met one basic criterion for
classification as mature DC (10).
Previous isolation procedures from both this and other laboratories had resulted in the selective loss of one or another of these splenic DC subsets, so their biological properties had not been fully evaluated and compared. Accordingly, in this study we examine the immediate developmental origin of the CD4+8-, CD4-8-, and CD4-8+ DC subsets of mouse spleen, asking about their relative maturation state, phagocytic ability, precursor-product relationships, and developmental kinetics. The results point to three separate developmental streams leading to three rapidly turning over DC subtypes.
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Materials and Methods |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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All mice were bred under specific pathogen-free conditions at The Walter and Eliza Hall Institute animal breeding facility. They were used at 5–7 wk of age unless otherwise stated. Most experiments used male mice of the C57BL/6J Wehi strain. For studies of the kinetics of development from bone marrow precursors the irradiated recipients were C57BL/6 Ly5.1 Pep3b mice, aged 8–10 wk. The CD4-/- C57BL/6 mice with the CD4 gene eliminated by homologous recombination were provided by Dr. W. Heath (The Walter and Eliza Hall Institute) and were originally obtained from Dr. T. Mak. The anti-CD4 transgenic mice of the BM-1 strain, expressing the GK1.5 anti-CD4 gene under the human CMV promoter, have been described previously (11).
Bone marrow transfer
The recipient C57BL/6 Ly5.1 mice were gamma-irradiated with two doses of 5.5 Gy, 3 h apart. Total bone marrow cells from C57BL/6 (Ly5.2) mice were transferred i.v. (3–5 x 106 nucleated cells/recipient mouse) into groups of three irradiated recipients, which were pooled for isolation of spleen DC at the specified times after transfer. The progeny DC were detected on the basis of Ly5.2 expression in conjunction with the DC markers.
LPS or monophosphoryl lipid A (MPL) treatment of mice
Groups of eight mice were injected i.v. with either LPS from Escherichia coli (L3137, Sigma-Aldrich, Castle Hill, Australia) suspended in PBS (30 µg/mouse) or MPL (12) (provided by SmithKline Beecham, Rixensort, Belgium) suspended in a buffered salt solution (20 µg/mouse). In both cases the control mice were injected with the appropriate suspension medium only. The suspension medium had been tested and verified as endotoxin free. The mice were generally studied 18 h postinjection.
Injection of fluorescent latex particles for in vivo phagocytosis
Groups of eight mice were injected with 200 µl of 2.6% solid Fluoresbrite Carboxy YG 0.5-µm latex microspheres (Polysciences, Warrington, PA). Control mice were injected with the equivalent nonfluorescent latex microspheres from the same source. One, 18, or 42 h later DC were isolated side-by-side from the spleens of pooled test mice or the pooled control mice. During analysis the DC from the control mice were labeled with the same immunofluorescent reagents as the test mice and served as the background to assess fluorescent microsphere uptake. In some experiments 2.0-µm fluorescent latex microspheres from the same source were used in a similar manner.
Isolation of DC
The procedure was performed as described recently (10, 13). Briefly, spleen fragments were digested for 20 min at room temperature with collagenase-DNase, then treated for 5 min with EDTA to disrupt T cell-DC complexes. All subsequent procedures were performed at 0–4°C in a divalent-metal free medium. Light density cells (<1.077 g/cm3 at mouse osmolarity) were selected by centrifugation in a Nycodenz (Nycomed Pharma AS, Oslo, Norway) medium. Cells not of DC lineage were depleted by incubating the cells with titrated levels of anti-CD3 (KT3), anti-Thy1 (T24/31.7), anti-B220 (RA3-6B2), anti-Gr-1 (RB6-8C5), and anti-erythrocyte (TER-119) Ab, then removing the Ab-binding cells with anti-rat Ig-coupled magnetic beads (Dynabeads, Dynal, Oslo, Norway). Note that anti-CD4, anti-F4/80, and graded levels of anti-CD11b Ab, used in earlier versions of the procedure (13), were all omitted from the depletion mix to avoid loss of the CD4+8- DC population (10). Note also that anti-B220 Ab, omitted in error from a previous description (10), was included. The splenic DC at this stage were usually about 80% pure. This preparation was then used for presorting when required or directly for immunofluorescent labeling before positive sorting or analysis.
Presorting to eliminate autofluorescent cells
The DC preparation described above contained a variable (5–30%) level of autofluorescent cells that were not DC, but could contaminate a DC sample isolated by sorting based on immunofluorescent labeling (10). In most cases these were eliminated before immunofluorescent labeling by high speed presorting, using a MoFlo instrument (Cytomation, Fort Collins, CO) as described in detail previously (10). In some cases when the level of autofluorescent cells was only about 5%, they were rejected by gating for autofluorescence in an unused fluorescence channel during the positive sorting or analyses as described previously (10).
Immunofluorescent labeling of DC preparations
The mAb, the fluorescent conjugates, and the multicolor labeling
procedures have all been specified previously (10, 13, 14). To identify and sort all DC, the pan-DC markers used were
high levels of class II MHC and/or CD11c. Anti-CD11c (N418) was used as
Cy5, FITC, or PE conjugates. Anti-class II MHC (N22 or M5/114) was used
as FITC, PE, Cy5, Texas Red, or Alexa 594 conjugates; the conjugation
levels were deliberately less than maximal to ensure that the strong
staining for class II MHC at saturation did not cause inaccurate color
compensation problems in other channels. The markers used to separate
the spleen DC subpopulations were CD8 and CD4. Anti-CD8
(YTS169.4) and anti-CD4 (GK1.5 or YTA3.1) (15, 16)
were used as FITC, PE, Cy5, Alexa 594, or biotin conjugates. The
staining for costimulator molecules employed anti-CD80 (16-10A1),
anti-CD86 (GL1), and anti-CD40 (FGK45.5), used as FITC or
biotin conjugates. The anti-Ly5.2 used for distinguishing the
progeny of donor bone-marrow cells was AL1-4A2, used as an FITC
conjugate. The second-stage stain for biotin-conjugated mAb was
PE-streptavidin (PharMingen, San Diego, CA). Propidium iodide (PI) was
included at 1 µg/ml in the final wash after immunofluorescent
staining to label dead cells.
Flow cytometric analysis and sorting of DC
As described previously (10, 13), most analyses and some sorting were performed on a FACStarPlus instrument (Becton Dickinson, San Jose, CA), using up to four fluorescent channels for the immunofluorescent staining (FL4 for FITC, FL2 for PE, FL3 for Cy5, and FL1 for Texas Red or Alexa 594), with the FL5 channel set to exclude PI-positive dead cells and autofluorescent cells. Care was taken during gating that any cells brightly fluorescent in FL3 and spilling over into FL5 were not gated out as dead cells. As well as the class II MHC and CD11c markers, the forward and side scatter gates were set to select for the light scatter characteristics of DC. For presorting and for sorting in up to three colors when large DC numbers were needed, the MoFlo instrument (Cytomation) was used.
Bromodeoxyuridine (BrdU) labeling and analysis
The procedure was similar to that developed for the labeling
kinetics of T cells (17). Groups of eight mice were
initially injected i.p. with 1 mg of BrdU (Sigma, St. Louis, MO) in
saline to ensure immediate availability of the precursor, then
continuously given BrdU (0.8 mg/ml) in sterile drinking water, which
was changed daily. The DC were isolated and presorted to remove
autofluorescent cells as described above. They were then surface
stained as described above with either anti-CD11c or anti-class
II MHC (Cy5 conjugate), anti-CD8 (PE conjugate), and
anti-CD4 (Alexa 594 conjugate); PI staining was omitted. The DC
were then stained for BrdU uptake as follows. The surface-stained DC
were washed, resuspended in cold 0.15 M NaCl, and fixed by dropwise
addition of cold 95% ethanol. They were incubated on ice for 30 min,
washed with PBS, then incubated for 30 min at room temperature with PBS
containing 1% paraformaldehyde and 0.01% Tween 20. They were
pelleted, then incubated for 10 min at room temperature with 50 U of
DNase I (Roche, Mannheim, Germany) in 0.15 M NaCl containing 4.2 mM
MgCl2, then washed. They were then incubated for
30 min at room temperature with FITC-conjugated anti-BrdU mAb
(Becton Dickinson), washed, and resuspended in PBS. The stained DC were
then analyzed on the FACStarPlus flow cytometer
(Becton Dickinson) for the number of FITC-positive (BrdU positive)
cells in each gated DC subset. The FITC fluorescence distribution of
BrdU-labeled cells showed only 3% overlap with the background in the
FITC channel when analyzing DC subsets from non-BrdU-treated
controls.
Short term culture of isolated DC
DC preparations were stained with the following combinations to
avoid background and autofluorescence problems postculture, which were
minimal in the Cy5 channel: 1) anti-CD11c-PE, anti-CD8-Cy5, and
anti-CD4-FITC; or 2) anti-CD11c-PE, anti-CD8-FITC, and
anti-CD4-Cy5. A small sample of each was analyzed immediately using
FACStarPlus (Becton Dickinson) and compared with
unstained controls. The bulk of each group was then separately sorted,
using the MoFlo instrument (Cytomation), collecting from group 1
CD11c+ CD4-
CD8+ DC and CD11c+
CD4- CD8- DC, and from
group 2 CD11c+ CD4+
CD8- DC. Cells from each of the sorted groups
were then incubated at 37°C for 16 h at
106 cells/ml in modified RPMI 1640 medium
(13) containing either GM-CSF (100 U/ml) or GM-CSF plus
CpG-DNA (oligonucleotide containing a CpG motif (18),
phosphorothioate modified, synthesized by GeneWorks (Adelaide,
Australia; 0.5 µM)) or GM-CSF plus a stimulatory anti-CD40 mAb
(FGK45.5; 100 µg/ml). After incubation the DC were collected, and
each sample was divided into two parts. One part was restained with the
same reagent combinations used initially, and the other was not
restained. All samples were then analyzed on the
FACStarPlus, gating on viable (PI excluding) DC
(CD11c+) and using the Cy5 channel to analyze CD4
or CD8 expression.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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In a previous study (10) we distinguished three DC
subtypes in mouse spleen, namely
CD4+8-
DEC-205- CD11b+,
CD4-8-
DEC-205- CD11b+, and
CD4-8+
DEC-205+ CD11b-. We showed
that the CD4 and CD8 on the surface of these DC were genuine DC
products and had not merely been picked up from associated T cells
(10). The existence of these three DC subtypes is
confirmed in the analysis of control spleen DC preparations, shown in
Fig. 1
. In a separate study (data not
shown) we have confirmed by immunofluorescent staining of spleen
sections that the majority of these DC are located in the marginal
zones of the spleen, with only the
CD4-8+DC being
concentrated in the T cell areas. An important issue was whether these
DC subpopulations derive from separate developmental lineages or
whether they are just maturation or activation stages of the one DC
lineage. We had provided some evidence, but no formal proof, that the
DC bearing CD8 have a different developmental origin from those that
lack CD8 (19, 20, 21). The relationship of the CD4-bearing DC
to the other DC subpopulations had not been examined. Accordingly, we
first examined their relative maturation status. All three DC
subpopulations could be classed as mature, because they were able to
efficiently activate allogenic T cells into cell cycle
(10) and because, as shown in Fig. 2, they all expressed similar levels of
CD40, CD80 (B7.1), and CD86 (B7.2) when freshly isolated from normal
spleen. However the CD4-8+
DC could be considered more mature by some criteria, because some cells
of this subpopulation expressed higher surface levels of class II MHC
(Fig. 2), and they were larger and showed more pronounced dendritic
extensions on microscopic examination. Thus, it remained a possibility
that some of the DC subsets simply reflected differences in the
relative state of terminal DC activation or maturation. Accordingly, we
tested the effect of various bacterially derived adjuvants that induce
DC activation and maturation to determine whether one subpopulation
could then be transformed into another. Bacterial LPS is known to
induce DC maturation; it also causes a shift of DC from the marginal
zones into the T cell areas of spleen, followed by DC death
(22). Injection of LPS produced 18 h later a more
mature or activated state in all three splenic DC subpopulations, as
judged by increases in surface class II MHC, CD40, CD80, and
particularly CD86 (Fig. 2). There was a drop in the level of CD4 on the
CD4+ cells recovered to the point where overlap
with the CD4- DC was substantial together with a
small drop in the CD8 level per CD8+cell
recovered. Both the proportion and absolute number of
CD4+8- and
CD4-8+ DC were reduced
(Table I). However, the marked drop in DC
recovery at this point made it more likely that most of the shift was
due to the selective death of these subpopulations, rather than to
complete down-regulation of CD4 or CD8 and transformation into a more
mature CD4-8- DC form.
Clearer results were obtained with injection of MPL, a less toxic
derivative of LPS and used as an adjuvant (12). This also
produced further activation of all DC subpopulations (Fig. 2), but
without any significant drop in overall DC recovery (Table I). In this
case, although there was a decrease in the level of CD4 and CD8 per
cell, all three subpopulations defined by CD4 and CD8 expression
remained present, and there was little evidence for transformation of
one into another (Fig. 1).
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. The phagocytic capacity of splenic DC subpopulations
A well-accepted model of DC maturation is that DC lose much of
their capacity for phagocytosis of particulate and antigenic material
when they migrate to lymphoid organs and develop the capacity to
stimulate T cells. As a further test of the maturation state of the
three splenic DC populations, fluoresceinated latex beads of two sizes
as well as fluoresceinated protein and polysaccharide Ags were injected
i.v. into mice. The entry of label into the three splenic DC
populations 2, 18, and 24 h later was ascertained. As illustrated
in Fig. 3, a surprising high proportion
of all three splenic DC populations showed a capacity to take up the
0.5-µm latex beads, at levels ranging from one to many beads per DC,
within 2 h of injection. The
CD4-8+ DC population
showed the lowest proportion of phagocytic cells, but even here about
30% of the DC contained beads, and the number of beads per positive
cell was similar to that in other DC. It was verified by fluorescence
microscopy that the fluorescent beads were within the DC rather than
simply adherent to the cell surface. Similar results were obtained with
the larger 2-µm beads and with a range of protein and polysaccharide
Ags (data not shown). In the case of the nondigestible fluorescent
latex beads, the label persisted 18 h later (Fig. 3). In the case
of protein Ags, the label was markedly reduced at 18 h and was no
longer detectable at 48 h (data not shown).
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Because it was surprising that these relatively mature DC already
within the spleen should still be capable of phagocytosis, LPS was
administered to the mice to induce further DC maturation. This was
expected to cause a reduction in DC phagocytic capacity. However, as
illustrated in Fig. 3
, even the splenic DC of LPS-treated mice showed a
remarkable capacity to ingest the fluorescent latex beads; the
stimulant caused little change in the proportion showing bead uptake or
the number of beads per cell. The same was found for fluorescent
protein uptake (data not shown). In additional experiments the effect
of MPL administration was similar to that of LPS. There were no marked
differences among the three DC subpopulations in the effect of
bacterially derived adjuvants on their capacity to ingest latex
beads.
Overall, all three splenic DC subtypes showed a capacity to ingest particulate material as well as antigenic proteins, and this ability persisted despite bacterially derived signals that promote further DC activation or maturation. There was no evidence from these experiments that the three DC subtypes represented sequential stages of DC maturation.
Further maturation of splenic DC in culture
A more direct test of possible precursor-product relationships
among the three DC subpopulations would be first to separate them by
fluorescence-activated cell sorting, then to transfer each population
into an environment allowing further development. Our attempts to
perform such experiments by reintroducing splenic DC subpopulations
into the environment of a lymphoid organ have been unsuccessful to date
due to poor reseeding back into the spleen following i.v. injection and
to pick-up of surface Ags from thymocytes following direct injection
into the thymus. However, it is possible to obtain further maturation
of the isolated DC in short term culture before DC death intervenes;
the presence of GM-CSF in the medium improves DC life span and
recovery. Under these conditions the DC show changes akin to those seen
in vivo on LPS or MPL injection, namely increased expression of class
II MHC and of costimulator molecules; one difference is that CD80,
rather than CD86, shows the most marked increase on culture
(13). The effect of such terminal maturation during short
term (overnight) culture on the CD4 or CD8 expression of the separated
DC subtypes, either incubated with GM-CSF alone or with two effective
activators of DC (the bacterial DNA motif CpG-DNA or stimulatory
anti-CD40 mAb) is shown in Fig. 4.
Under these conditions DC recovery ranged from 60–80%; incubation
longer than 1 day resulted in marked DC death, particularly within the
CD4-8+ subpopulation.
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, IL-4, Flt-3 ligand,
and combinations of these) was included in the culture medium along
with GM-CSF. Overall, activating or maturing the splenic DC
subpopulations in culture did not involve any measurable
transformation between the
CD4+8- and the
CD4-8+ phenotypes or
between the CD4-8- and
CD4-8+ phenotypes. The
relative stability of CD8 expression on culture is in line with our
previous results (13). Although there was a marked
reduction in CD4 expression by the
CD4+8- DC in culture, most
cells retained some CD4 expression, and no distinct subgroup of
CD4-8- DC was
generated. Effect of elimination of CD4-bearing DC
Our original expectation was that the new population of splenic
CD4+8- DC, being adherent
and expressing several monocyte/macrophage markers, would be the least
mature form and would give rise eventually to
CD4-8- DC or even to
CD4-8+ DC. Although the
forgoing studies did not entirely support this concept, the partial
drop in CD4 surface expression on incubation or induced maturation left
the possibility that a complete loss of CD4 might nevertheless occur
normally in vivo. If
CD4+8- DC were the
obligatory precursors of other populations, elimination of this subset
should prevent the development of the downstream DC forms. It was
possible to test this using mice made transgenic for a depleting
anti-mouse CD4 mAb, GK1.5 (11). This anti-CD4
mouse has been found to be selectively depleted of all peripheral
CD4+ T cells (11). Therefore, the DC
levels in the spleen of these continuously depleted mice was determined
and compared with that in mice lacking CD4 because of elimination of
the CD4 gene by homologous recombination. During analysis the DC were
stained with an mAb, YTA3.1, that recognizes a different CD4 epitope to
GK1.5 and is not blocked by GK1.5 (15, 16). The results
are shown in Fig. 5 and Table II.
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Overall, these results suggest that CD4 expression is not essential for the development of spleen DC and that the CD4+8- DC subtype is not a precursor of the splenic CD4-8- or CD4-8+ DC found in normal spleen.
Turnover rates of the splenic DC populations
The rate of generation and the life span of the splenic DC
populations had not been established, although there was evidence from
a published study that CD8+ DC have a much longer
life span than CD8- DC (23). We
determined the rate of turnover of the three splenic DC subpopulations
by following the kinetics of labeling of DC with the DNA precursor BrdU
under conditions of continuous administration to mice. At various times
after initiating BrdU administration, splenic DC were isolated, and the
DC preparation was stained for class II MHC (to label all DC) and for
CD4 and CD8 (to segregate the DC subtypes) and, after
permeabilization, was stained with fluorescent anti-BrdU. The
majority of BrdU-labeled DC could be readily distinguished from
unlabeled DC, but because of a small overlap with the background
fluorescence, a fully labeled DC population would score 97%, rather
than 100%, BrdU positive. The labeling kinetics of the individual
spleen DC subpopulations is summarized in Fig. 6.
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represents the entry of nondividing progeny cells into the
splenic DC pool. It was notable that none of the spleen DC subpopulations showed any lag in BrdU labeling. A lag would be expected if one subpopulation was the product of another, as labeled DC progressively entered a single pathway consisting of sequential DC subtypes. Instead, all three subpopulations showed continuous and surprisingly rapid labeling, suggesting there were three independent streams of development, at least as far back as the dividing precursors.
In contrast to expectations from published data (23) the
CD4-8+ DC showed the
fastest turnover (half-life, 1.5 days) with near-linear uniform
labeling kinetics, so that almost all the
CD4-8+ DC had been
replenished by 3 days. The
CD4-8- and
CD4+8- DC had half-lives
of 2.0 and 2.9 days, respectively, but with a nonlinear labeling curve,
indicating that some of the DC of these populations had a longer
residence in the spleen than the majority. The data in Fig. 6 are from
mice 5–7 wk of age. Older mice (20 wk) showed a 15–20% slower rate
of DC turnover but with similar relative rates of labeling for the
subpopulations. Even for older mice, this indicated a surprisingly
rapid turnover rate for all spleen DC subtypes.
Effects of LPS and MPL on the rate of turnover of splenic DC
Because LPS and MPL promoted DC maturation or activation and
initiated DC movement into the T cell areas of spleen
(22), we checked the effects of their administration to
mice on the rate of spleen DC subpopulation turnover. A single critical
time measurement (1 day) was chosen to compare with the turnover data
in Fig. 6; the results are given in Fig. 7. LPS enhanced the labeling at 1 day of
all DC subpopulations, but especially of the
CD4-8+ subset. In view of
the DC loss following LPS administration (Table I), this result could
have been due as much to selective elimination and death of the oldest
DC as to enhanced DC production or entry into the spleen. MPL had
little effect on the labeling of
CD4-8- and
CD4+8- DC, but, like LPS,
caused a marked increase in the labeling of
CD4-8+ DC. Because MPL did
not cause measurable DC loss at this time point, not even in the
CD4-8+ subpopulation
(Table I), it appeared to promote genuine turnover, enhancing both
entry and loss of CD4-8+
DC from the mature splenic DC compartment.
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The BrdU labeling kinetics shown in Fig. 6 monitored the relative
rates of DC subtype development from the point of entry of the label
into the dividing DC precursors of unknown location. An alternative
approach was to follow the relative rates of generation of the splenic
DC subsets from precursors in bone marrow by transfer of bone marrow
into irradiated recipients differing at the Ly5 locus. The precursors
in bone marrow would include both multipotent hemopoietic stem
cells giving long term DC reconstitution and more committed DC
precursors giving faster, but transient, reconstitution. The results of
such a study are shown in Fig. 8.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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Several possible minimal models invoking only a single DC stream seemed possible. One preferred view was that the CD4+8- DC would be the least mature type, which on loss of CD4 would transform into CD4-8- DC and possibly also into CD4-8+ DC. The arguments for this were some similarities of these DC subtypes to monocytes and macrophages, including adherence properties and expression of CD11b and F4/80 (10). In addition, the partial loss of CD4 expression in culture or on stimulation by bacterially derived adjuvants in vivo encouraged the view that a complete loss of CD4 was possible. This model of the source of CD4-8- DC found in normal spleen was refuted by both the BrdU labeling kinetics and the bone marrow transfer studies. This indicated that CD4+8- DC had the slowest turnover in the spleen and that they were the slowest, rather than the fastest, to be generated following bone marrow transfer. Finally, the continuous and near complete elimination of CD4+8- DC in anti-CD4 transgenic mice without affecting the other DC subtypes was a strong argument against the view that expression of CD4 was an early step in the development of either CD4-8- DC or CD4+8- DC.
A second view was that
CD4-8- DC would be the
least mature DC subtype and generate the others by acquisition of
either CD4 or CD8 during further maturation. We consider that some
form of immature DC must at some stage do just this, changing from
CD4-8- to
CD4+8- or
CD4-8+. However, all the
data we obtained from the BrdU labeling kinetics and from the forced
maturation experiments indicate that the majority of
CD4-8- DC in normal
spleen are not immature DC, are not the direct precursors of the other
subtypes, and cannot be induced to express either of these markers.
They behave as a separate mature DC subtype.
If anything, the results we obtained were the exact opposite of these expectations, with the one population that appeared marginally more mature, the CD4-8+ DC, being the first to be generated from bone marrow precursors and having the fastest turnover rate. However, we do not consider it likely that these were precursors of the other DC subtypes. There was no kinetic evidence from BrdU labeling that these served as precursors, and all the attempts at precursor-product analyses suggested only that they died faster, not that they produced other DC forms.
A final possibility was that CD4 and CD8 are markers of no
interpretive value, being randomly acquired on a single DC type by some
functionally irrelevant process. However, the fact that the three DC
subsets displayed differing and characteristic BrdU labeling and bone
marrow reconstitution kinetics argues strongly against this, as does
the published evidence of differences in biological function
(4, 5, 6, 7, 8, 9). Current work from this laboratory (in preparation)
indicates that each of these subsets secretes a different pattern of
cytokines, strengthening the view that they are functionally
distinct.
Thus, the evidence to date, especially the lack of any obvious precursor to product labeling pattern in the BrdU labeling kinetics, points to the existence of three separate, independent DC developmental streams. It should be explained that the BrdU labeling data and induced maturation studies only make this point for the last steps of DC maturation, downstream from the last dividing DC precursor. This evidence does not extend back to the hemopoietic precursor cells and does not bear on the controversial issue of whether these DC subtypes are originally of myeloid or lymphoid precursor origin. However, the fact that our study of the rate of DC generation from bone marrow precursors also revealed very different generation kinetics for the different DC subtypes, in line with the BrdU labeling studies, does suggest the three distinct DC developmental streams extend back far enough to consider them different hemopoietic sublineages. In our current studies we are attempting to identify the bone marrow precursors of these separate sublineages.
Two observations in this study appear to contradict the conventional views on DC maturation. The first is our finding that even the mature DC resident in the spleen retain a significant capacity for uptake of particulate material (even up to 2-µm diameter) and foreign molecules; this occurred even when further maturation was induced by bacterial products. It might have been argued that the uptake of the latex beads and the Ags had occurred while a less mature form of DC was in the bloodstream, and labeled DC only later seeded into the spleen and matured further. Against this is the finding that 30–70% of splenic DC showed uptake of beads and Ags within 2 h of injection, a period very much shorter than the rate of turnover of DC within the spleen. In addition, the DC populations isolated directly from the spleen showed a similar capacity to take up latex beads in culture. Leenen et al. (23) have also reported that DC within lymphoid organs retain phagocytic activity. In the latter case only the marginal zone DC (presumed CD8-) appeared phagocytic, whereas the DC in the T cell areas (presumed CD8+) did not, based on the loss of DC following i.v. injection of clodronate-loaded liposomes. We found that the CD8+ DC also showed a phagocytic ability for latex particles, although somewhat less than that of CD8-DC. One possible explanation for this difference is a reduced access of DC in the T cell zones to the i.v. injected liposomes in the Leenen et al. study (23). Another is that CD8+ DC may have an inherently low affinity for the liposomes they used. A third is that the CD8+ DC that showed uptake of the latex particles in our study were those outside the T cell area, although the finding of similar results after LPS injection makes this less likely. There are several studies that indicate that even mature DC retain some capacity to endocytose and process Ags (24, 25). Although it is doubtless true that the capacity of DC for phagocytosis and endocytosis drops during maturation, the conventional model seems overstated, because even relatively mature DC already situated in lymphoid organs are capable of significant uptake of foreign or particulate material.
The second contradictory result was that the CD8+ DC have the shortest life span of any spleen DC subtype. This is in direct conflict with the conclusions of Leenen et al. (23), who followed the persistence of DC in transgenic mice with a conditional ablation of dividing DC precursors. They found the interdigitating DC in the T cell zones of mouse spleen (presumed to be the CD8+ DEC-205+ DC) persisted much longer than the DC found in the marginal zones (presumed to be the CD8-DEC-205- DC). This fitted with suggestions that CD8+ DC could be long term lymphoid tissue-resident DC with regulatory functions, such as presenting self-Ags and inducing self-Ag T cell tolerance (5, 6, 26, 27). In contrast we find the CD8+ DC have by far the shortest, rather than the longest, life span. We now question whether the conditional ablation system of Salomon et al. (28), as used by Leenen et al. (23), was functioning for splenic CD8+ DC as efficiently as it functioned for CD8- DC. This system requires the expression of a viral thymidine kinase in dividing cells, in this case dividing DC precursors, making them sensitive to ganciclovir, a nucleoside analogue. The survival of spleen T cell zone DC 7 days after ganciclovir administration may simply have reflected a poor expression of the enzyme in the separate precursors of the CD8+ DC lineage. Our data indicate that virtually all CD8+ DC and most other DC turn over within 3 days and so should have been eliminated by 7 days if the ablation of dividing DC precursors was effective. Regardless of the explanation of the earlier results, both our BrdU uptake approach and our studies on the rate of subset reconstitution by bone marrow precursors indicate that we must now reassess the immunological role of CD8+ DC in view of their exceptionally fast turnover.
A point of general importance revealed in this study is the surprisingly rapid turnover of all DC subtypes in mouse spleen. Most resident DC were replaced within 3–4 days. We are currently assessing whether this is true of DC in other lymphoid tissues, such as lymph nodes, and whether it is true when DC interact with activated T cells. This result has important implications for the persistence of Ag presentation following an antigenic challenge. Clearly, the stabilization of the MHC-peptide complex on the surface of DC as DC mature (29, 30, 31) does not imply prolonged presentation of the antigenic peptide if the presenting DC itself has a very short life span.
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2 Abbreviations used in this paper: DC, dendritic cells; MPL, monophosphoryl lipid A; PI, propidium iodide; BrdU, bromodeoxyuridine.
Received for publication August 1, 2000. Accepted for publication September 15, 2000.
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