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The Walter and Eliza Hall Institute of Medical Research, Melbourne, Victoria, Australia
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Abstract |
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 Top Abstract IntroductionMaterials and MethodsResultsDiscussionReferences |
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+
DEC-205high CD11blow DC. The CD4 on the surface
of the CD4+ splenic DC subpopulation was produced by the DC
themselves, and CD4 RNA transcripts were present. Likewise, the CD8
on the surface of the splenic CD8+ DC was shown to be a
product of the DC themselves, in agreement with earlier evidence. All
three spleen DC types would be considered as mature, based on
expression of CD80, CD86, and CD40 as well as on T cell stimulating
function. Mouse thymuses appeared to contain two DC types; both were
DEC-205highCD11blow, but they differed in the
level of CD8 expression. However, as well as this authenticated
marker expression, immunofluorescent staining was also found to reflect
a series of artifacts, due to the autofluorescence of contaminating
cells and due to pickup of CD4 and CD8ß. By constructing mice
chimeric for the hemopoietic lineages using mixtures of wild-type bone
marrow with CD4null or CD8null bone marrow,
a marked pickup by thymic DC of Ags derived from thymocytes was
demonstrated. |
Introduction |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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The expression of certain cell-surface markers has already pointed to
substantial DC heterogeneity. We reported that a subpopulation of mouse
spleen DC display on their surface CD8 as an homodimer and, in
accordance with this, express mRNA for CD8 but not for CD8ß
(11). This splenic CD8+ DC
population is DEC-205+
CD11b-, in contrast to a second splenic DC
population that is CD8-
DEC-205- CD11b+
(12). Lymph node DC appear still more heterogeneous and
include an additional population that is
CD8low DEC 205+
(12, 13, 14). By contrast, our analysis of mouse thymus DC
revealed what appeared to be a single group of
CD8+ DEC-205+ DC, albeit
with a wide range of CD8 surface staining (12, 15).
Murine CD8+ DC appear to represent a
lymphoid-related DC lineage separate from the conventional
myeloid-related CD8- DC (10).
CD8+ thymic or splenic DC can be produced in
recipient mice on transfer of an early thymic lymphoid precursor
population with little, if any, capacity to form myeloid cells
(16, 17). The murine lymphoid-related DC lineage differs
from myeloid-related DC both in the cytokine requirements for
development and in transcription factor control (18, 19).
However, the expression of CD8 by the lymphoid-related DC does not
appear to be essential for either their development or their function
(18, 20), so it may be a relic of their particular
developmental history. No CD8 expression has been detected on human DC
(21).
Initially we detected only low levels of CD4 on murine DC
(11), as did Crowley et al. (22). In what at
that time appeared as a striking contrast, a proportion of human and
rat DC stained strongly for surface CD4 (21, 23, 24).
However, Salomon et al. (13) have since reported CD4
staining of a subgroup of mouse lymph node DC, and Pulendran et al.
(25) have noted the occurrence of CD4 staining on a
subgroup of Flt3 ligand-treated mouse spleen DC. We now find that a
major population of mouse spleen DC expresses relatively high levels of
CD4, a population previously eliminated from our preparations by the
immunodepletion steps used to ensure high DC purity. However, we find
that not all apparent CD4 or CD8 staining represents authentic
expression by the DC because both autofluorescence and passive pickup
of these molecules onto the DC surface confuse the staining results.
Accordingly, we have conducted a critical analysis of the CD4, the
CD8, and the CD8ß staining of mouse thymus and spleen DC. Using
these markers and appropriate controls, we identify three distinct
populations of DC in mouse spleen and two populations of DC in mouse
thymus.
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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. Most experiments used female mice of the C57BL/6J Wehi strain. For constructing bone marrow chimeras, the recipients were C57BL/6 Ly-5.1 Pep3b mice. The bone marrow donors included the above strains, CD8-/- C57BL/6 mice (originally obtained from Dr. T. Mak), and CD4-/- C57BL/6 mice (provided by Dr. W. Heath; originally obtained from Dr. T. Mak). For the allogeneic T cell stimulation experiments, the CD4 T cells were derived from CBA CaH Wehi mice.
Bone marrow chimeric mice
The C57BL/6 Ly-5.1 mice were gamma-irradiated (two doses of 550
rad, 3 h apart) and then were injected i.v. with 3 x
106 bone marrow cells from wild-type Ly-5.1 mice
together with 3 x 106 Ly-5.2 bone marrow
cells from CD4-/- mice,
CD8-/- mice, or wild-type C57BL/6 mice. For
each experimental group reconstituted with a mix of Ly-5.1 wild-type
and Ly-5.2 CD4null or CD8null
bone marrow, a parallel control group reconstituted with a mix of
Ly-5.1 wild-type and Ly-5.2 wild-type bone marrow was set up. Five to
seven weeks after bone marrow transfer, the chimeric mice from each
group were pooled, and the DC were isolated from the spleens and
thymuses for analysis.
DC suspension and isolation media
These were all isoosmotic with mouse serum and identical with those described previously (11).
DC isolation procedure
The method was similar to our previous procedures (11, 12) with modifications to avoid loss of any CD4-bearing DC.
Spleens (normally eight) were cut into small fragments and then
digested with frequent mixing for 25 min at room temperature (22°C)
in 10 ml modified RPMI 1640-FCS medium containing collagenase (1 mg/ml;
type II; Worthington Biochemical, Freehold, NJ; verified as free of
trypsin-like protease activity) and DNase I (Boehringer Mannheim,
Mannheim, Germany). To disrupt DC-T cell complexes, EDTA (1 ml, 0.1 M
(pH 7.2)) was added, and mixing continued for 5 min. Undigested fibrous
material was removed by filtration through a stainless steel sieve. All
subsequent steps were at 0–4°C using a divalent metal-free balanced
salt solution containing EDTA (EDTA-BSS). Cells were recovered from the
digest by centrifugation, the pellet was resuspended in a 1.077
g/cm3 isoosmotic Nycodenz medium and centrifuged
at 1700 x g for 15 min, and then the low-density
fraction was collected. The low-density cells (3–5% of the total)
were diluted in EDTA-BSS, recovered by centrifugation, and then
incubated for 30 min with the following mAb: anti-CD3 (KT3-1.1);
anti-Thy 1 (T24/31.7, a pan-Thy 1); anti-Gr1 (RB68C5); and
anti-erythrocyte (TER-119). All mAb were titrated and used at
near-saturating concentrations, except anti-Thy 1, which was used
at 25% of this level to avoid removing any DC that had absorbed low
levels of Thy 1. Free mAb was removed by centrifuging the DC through a
cushion of EDTA-FCS, and then the cells coated with mAb were removed
using anti-rat Ig coupled magnetic beads (Dynabeads, Dynal, Oslo,
Norway). The beads and cells in a 5:1 ratio, respectively, were mixed
as a concentrated slurry by continuous slow rotation for 20 min. The
slurry was then diluted with EDTA-BSS, and the beads and bound cells
were removed with a Dynal magnet using several removal steps. The
splenic DC (around 85% pure) were suspended in EDTA-BSS for
presorting, staining, and analysis or sorting. For the isolation of
thymic DC a similar procedure was used in this study, starting with
thymuses and doubling the initial quantities to give a DC preparation
only 20–30% pure. However, because thymic DC were finally shown to
lack the CD4+ F4/80+
population, it was possible to produce an equivalent population of DC
80% pure by using the following mAb in the depletion mixture:
anti-CD3 (KT3-1.1); anti-Thy 1 (T24/31.7, a pan-Thy 1);
anti-CD4 (GK1.5); anti-CD8ß (53-5.8); anti-CD25 (PC/61);
anti-CD11b (Mac-1; M1/70.15); anti-F4/80 (F4/80);
anti-Gr1 (RB68C5); and anti-erythrocyte (TER-119). Anti-Thy 1,
anti-CD4, anti-CD8ß, anti-CD11b, and anti-F4/80 were
all used at 25% of the usual near-saturation level to avoid depleting
cells that either absorbed or expressed low levels of these markers on
the surface.
Immunofluorescent labeling of DC preparations
The mAb, the fluorescent conjugates, and the labeling procedure
have all been given in detail elsewhere (11, 12). The
hybridomas were all grown and the mAb were purified and conjugated in
this laboratory. The mAb used as pan-DC markers for segregating and
sorting DC from non-DC contaminants were: anti-CD11c (N418), used
as a FITC or Cy5 conjugate, and/or anti-class II MHC (N22), used as
a Cy5 or Texas Red conjugate. Anti-class II MHC, which normally stains
DC very strongly, was deliberately conjugated at less than maximal
levels to keep fluorescence at saturation staining to only medium-high
levels with which accurate compensation for fluorescence in other
channels could be maintained. CD11c staining alone was used where DC
were to be used for stimulating CD4 T cells to avoid blocking class II
MHC. The mAb normally used to divide the DC into subpopulations were:
anti-CD8 (YTS169.4), used as a FITC or an Alexa 594 conjugate;
anti-CD4 (GK1.5), used as a PE or an Alexa 594 conjugate;
anti-DEC-205 (NLDC145), used as a FITC or a biotin conjugate; and
anti-CD11b (M1/70), used as a FITC, biotin, or Cy5 conjugate. Other
mAb used were as follows: anti-CD8ß (53-5.8), used as a FITC or
biotin conjugate; anti-CD80 (16-10A1), used as a FITC or biotin
conjugate; anti-CD86 (GL1), used as a FITC or biotin conjugate;
anti-CD40 (FGK45.5), used as a FITC or biotin conjugate;
anti-CD24 (M1/69), used as a FITC or biotin conjugate;
anti-CD49d (PS/2), used as a FITC or biotin conjugate;
anti-CD49f (EA-1; obtained from B. Imhof, Department of Pathology,
Université de Génève, Switzerland), used as a FITC or
biotin conjugate; and anti-F4/80 (F4/80), used as a FITC or biotin
conjugate. The second-stage stain for the 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. DC were labeled in EDTA-BSS-FCS medium and before analysis were passed through a 26-gauge needle or repetitively passed through a pipette tip to minimize doublet formation.
Flow cytometric analysis, sorting, and presorting of DC
The fluorescent-labeled DC preparations were analyzed on a FACStarPlus instrument (Becton Dickinson, San Jose, CA). Color compensation was initially set using Calibrite beads (Becton Dickinson) and then was checked using appropriate stained cell controls. Up to four fluorescent channels were used for the immunofluorescence staining (FL1 for fluorescein, FL2 for PE, FL3 for Cy5, and FL4 for Texas Red or Alexa 594), with the FL5 channel set to exclude PI-positive dead cells and autofluorescent cells. Care was taken in setting this gating to ensure that any cells staining very brightly in FL3 and spilling over into FL5 were not gated out as dead cells.
In many cases the DC preparations were presorted rather than just gated to remove autofluorescent cells with the presorting being performed before immunofluorescent staining and analysis. Presorting was conducted at high speed on a MoFlo instrument (Cytomation, Fort Collins, CO) or on the FACStarPlus instrument using the FL1 and FL2 channels. In some cases the thymic DC preparations were labeled with anti-CD11b before presorting, and cells expressing high levels of CD11b were excluded along with the autofluorescent cells. This same channel of fluorescence could then be used for anti-class II MHC on subsequent staining because only cells showing high fluorescence were included as DC. Such presorting of thymic DC preparations was not required once it was established that immunomagnetic bead depletion employing anti-CD4, anti-F4/80, and anti-CD11b could be performed without eliminating any significant thymic DC population.
Short-term culture of DC
To wash passively bound proteins from the DC surface, the DC-enriched preparation, after presorting to remove autofluorescent cells but before labeling and sorting, was incubated for 16 h at 37°C in modified RPMI 1640-FCS medium, in a 10% CO2-in-air atmosphere, and at a concentration of 5 x 105 cells/ml. The DC were then harvested, and dead cells were removed by selecting light-density viable DC using the Nycodenz centrifugation procedure described above. Recoveries of viable DC were around 50%.
DC-allogeneic T cell mixed leukocyte cultures
The mixed leukocyte culture system for determining the stimulatory capacity of DC was as described previously (5, 6). Briefly, DC were sorted from the C57BL/6 mice as CD11c+ rather than MHC Class II+ cells. Various numbers of DC were cultured with 20,000 CBA mouse lymph node CD4 T cells and purified as described previously (5) in 200 µl medium in V-well plates. At various times the cultures were pulsed for 6 h with [3H]TdR, and then thymidine incorporation into cellular DNA was measured by liquid scintillation counting. The data shown are for day 3 of culture, the response peak, but similar relative stimulatory activity was observed at all time points.
Assay for DC adhesion properties
DC sorted from spleen DC preparations on the basis of class II
MHC, CD4, and CD8 expression were suspended in a HEPES-buffered
mouse osmolarity RPMI 1640 medium containing 10% FCS. The suspensions
were placed in shallow chambers consisting of coverslips fastened to
glass slides by double-sided tape along two opposite edges. After
filling the chambers with the DC suspension, the other two edges of the
coverslip were sealed with nail polish. After 1 h of incubation at
37°C, the DC in the chambers were viewed under phase contrast
microscopy using a x40 objective, and adherent cells were counted.
RT-PCR for CD4 mRNA expression in DC
Total cytoplasmic RNA was prepared from purified DC populations using Qiagen RNeasy Mini Kit (Qiagen, Clifton Hill, Australia). The first-strand cDNA synthesis from extracted RNA was performed in a final volume of 50 µl using an AMV Reverse Transcription System (Promega, Madison, WI). The PCR was performed in a final volume of 30 µl containing 5 µl cDNA (equal to 1000 cells), MgCl2 (1.5 mM), thermo-reaction buffer (1x), dNTP mixture (0.2 mM each), 1 µg of each oligonucleotide primer, and 1 U of Taq polymerase (Promega). Each set of 25–35 cycles consisted of 1 min at 94°C (for the first cycle this step was 4 min), 1 min at 60°C, and then 1 min at 72°C (for the final cycle this step was 11 min). A DNA thermal cycler was used (Perkin-Elmer/Cetus, Norwalk, CT). A sample (15 µl) of each reaction was electrophoresed through a 2% agarose gel. PCR for ß-actin was performed (using the same PCR conditions) as controls for the amount of cDNA. The relative intensity of each CD4 PCR product band was then normalized on the basis of the ratio of the intensity of the CD4 band compared with the corresponding ß-actin band. The sequence and the position of the oligonucleotides used as primers for the PCR were as follows: CD4 forward primer 828-846, 5'-GAG AGT CAG CGG AGT TCT C-3'; CD4 reverse primer 1156-1135, 5'-CTC ACA GGT CAA AGT ATT GTT G-3; ß-actin forward primer 25-45, 5'-GTG GGC CGC TCT AGG CAC CAA-3'; ß-actin reverse primer 541-564, 5'-CTC TTT GAT GTC ACG CAC GAT TTC-3'.
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Results |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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In earlier studies we had seen only marginal staining of mouse
spleen DC for CD4 (11), even when anti-CD4 was omitted
from the immunomagnetic bead depletion procedure used to remove
nondendritic lymphoid and myeloid cells from the preparation. However,
when both CD4 and F4/80 were omitted during this depletion step, the
overall yield of DC in the enriched preparation doubled (to a mean of
2.5 x 106 DC per spleen), and over half of
the DC obtained stained for CD4 (Fig. 1
).
In all respects other than the appearance of this additional
CD4+ population, the properties of the
CD4-CD8+ and the
CD4-CD8- DC were
identical with those we have reported previously. The proportion of
splenic DC that were CD4+ was higher than with
lymph node DC of which only around 15% were CD4+
(data not shown). The CD4-bearing DC were among the
CD8- DC rather than the putative
lymphoid-related CD8+ DC, as shown in Fig. 1.
This figure also provides an estimate of the cutoff point where the
anti-CD4 and anti-F4/80 immunomagnetic bead procedure
eliminated the high-staining CD4+ DC in our
earlier studies.
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, with no new subpopulations
apparent. However, longer sorting or analysis was then required to
reject the high level of contaminants. Autofluorescent cells in DC-enriched preparations
One cost of omitting immunomagnetic bead depletion, or reducing
the number of mAb used for depletion, was a reduced purity before
sorting and, as a consequence, a reduced resolution and an increasing
difficulty in eliminating all contaminants. This was particularly the
case for autofluorescent cells, which sometimes were at high levels in
the DC preparations. Fig. 1 represents a favorable case in which most
of the 8% autofluorescent cells could be gated out along with
PI-positive dead cells during analysis. However, in many cases as
illustrated in Fig. 2, which shows the
autofluorescence of an unstained DC preparation, autofluorescent cells
represented up to 30% of the DC-enriched sample. The autofluorescence
overlapped both the class II MHC and the CD11c fluorescent staining
used to define DC, as well as the CD4 and CD8 fluorescent staining used
to separate DC subpopulations. Autofluorescence was brightest in the
most useful PE and FITC channels. It could not be completely gated out
using other channels without concurrent loss of DC because of
limitations in the accuracy and linearity of the color-compensation
systems. The autofluorescence was a property of intact, viable cells
that could not be gated out on the basis of high PI staining.
Accordingly, the most effective (albeit time-consuming) approach was to
presort to eliminate autofluorescent cells before immunofluorescent
staining.
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), and the surface
phenotype of both was examined (data not shown). All class II
MHChigh, CD11chigh,
DEC-205+, and CD40+ cells
that could be classed as mature DC were within the nonautofluorescent
fraction. The autofluorescent cells were positive but low for class II
MHC and were almost negative for the DC markers CD11c, DEC-205, and
CD40. They expressed both CD80 and CD86. They were F4/80 strong
positive and most, but not all, were strongly positive for CD11b.
Giemsa staining of cytospin preparations indicated that most
autofluorescent cells had a morphology resembling macrophages. In
contrast to DC, the autofluorescent cells showed a strong and permanent
(>3-day) adherence to the plastic surface of culture vessels. We
conclude that the majority of the autofluorescent cells were
macrophages, although a proportion may have been immature DC. In the case of thymic DC preparations, omission of CD4 and F4/80 immunomagnetic bead depletion caused still greater contamination of the DC preparations. As well as the autofluorescent cells, CD11bhigh (Mac-1high) nonfluorescent macrophages and thymic lymphoid precursor cells caused problems in subsequent analysis and sorting. This was overcome in some experiments by staining the cells with anti-CD11b and eliminating the CD11bhigh cells, the smaller-sized lymphoid cells, and the autofluorescent cells in the presort. Possible CD11blow DC were left in the preparation. The fluorochrome and the channel originally used for the CD11b stain could be used again for staining class II MHC on subsequent analysis or sorting because only class II MHChigh cells were then selected as DC. These maneuvers, although used in this study, finally proved unnecessary because, as shown later, thymic DC lacked the CD4+, F4/80+, and CD11b+ DC subsets found in spleen. Therefore, a full immunomagnetic bead depletion was a safe procedure for thymic DC.
Apparent CD8int cells in DC preparations
One recurring puzzle had been the appearance in DC preparations of
cells selected as class II MHC+ or
CD11c+ DC and apparent staining at intermediate
levels for CD8, rather than showing the clear-cut negative or strong
positive CD8 staining shown in Fig. 1. This usually was seen when
autofluorescent cells appeared in the background, as shown in Fig. 3, making it difficult to assess the
significance of the apparent CD8 staining. As shown in Fig. 3, presorting the autofluorescent cells from the splenic DC preparations
removed most of the cells that produced the peak of apparent
CD8int DC, although some low-level staining with
the anti-CD8 mAb sometimes persisted, as in Fig. 3. In the case
of thymic DC, the effect was less pronounced, but again some of the
apparent lower staining with anti-CD8 mAb could be attributed to
the autofluorescent cells (Fig. 3). Some DC, especially those in the
thymus, express the low-affinity Fc
receptor CD16/CD32. Blocking
this receptor during staining caused some slight further reduction in
the low-level staining with anti-CD8, but most of this staining
remained.
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int splenic DC, they were sorted directly
from an enriched splenic DC preparation and were tested for their
capacity to stimulate allogeneic CD4 T cells in a mixed leukocyte
reaction. They were compared with DC that were gated to remove
autofluorescent cells and were sorted as CD11c+
DC but were not separated further. The autofluorescent and apparent
CD11c+ CD8int cells
showed a relatively poor ability to stimulate allogeneic T cell
proliferation compared with the nonautofluorescent DC (Fig. 4). A low level of cross-contamination
with authentic DC or the presence of some immature DC could have
explained the residual stimulatory activity. Accordingly, most of the
apparent CD8int DC in splenic DC preparations
were considered to be non-DC artifacts due mainly to autofluorescence.
Presorting to eliminate autofluorescent cells was then used in most
subsequent experiments.
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The major group of CD4+ splenic DC we had
previously lost from our preparations was assessed for its ability to
stimulate proliferation in a mixed leukocyte reaction to verify that
these cells could be classified as mature DC. As shown in Fig. 4, the
DC-bearing CD4 were as effective at stimulating allogeneic CD4 T cells
as the total populations lacking CD4 when CD11c+
splenic DC were sorted simply on the basis of CD4 expression.
The splenic DC-bearing CD4 were all CD8-,
with the putative lymphoid-related CD8+ DC
lacking significant CD4 staining (Fig. 1). Therefore, splenic DC could
be subdivided into the following three populations:
CD4+8-,
CD4-8-, and
CD4-8+. One distinguishing
feature of the splenic
CD4+8- DC was their
adherence properties (Table I). After
1 h at 37°C, the majority of the
CD4+8- DC adhered and
spread out on a glass or plastic surface, much like macrophages did but
with the maintenance of a convoluted outer membrane with dendritic
extensions. Despite these adherence properties, the
CD4+8- DC showed little
capacity to phagocytose zymosan particles (data not shown). Although
some CD4-8- and
CD4-8+ DC also adhered,
the proportion was always much lower (Table I). Thus, isolation of
splenic DC based on an initial adhesion to glass or plastic, as in an
earlier DC isolation protocol (26), likely would have
selected for the CD4+8- DC
(those above the broken line in Fig. 1).
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The surface antigenic phenotypes of the three spleen DC
subpopulations were compared using a panel of mAb in three- and
four-color immunofluorescent staining (Figs. 5 and 6).
The splenic DC, including the CD8+ DC, were
all CD8ß-. The
CD8+ß- DC were
CD4- and CD11b-
DEC-205+, in accordance with our earlier results
(12). The
CD4-8- and the
CD4+8- DC were
indistinguishable by most markers, both differing from the
CD8+ DC in being CD11b+
and DEC-205- or
DEC-205low. They also both differed from the
CD8+ DC in expressing lower levels of the
heat-stable Ag CD24 and the 6 integrin CD49f.
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4 integrin CD49d (Fig. 6).
They were all positive and expressed similar levels of the costimulator
molecules CD40, B7.1 (CD80), and B7.2 (CD86) (Fig. 6). Thus, they could
all be classed as mature DC, in agreement with their ability to
activate T cells. The only feature found to distinguish the
CD4+ DC, apart from the expression of CD4 itself
and the adhesion properties (Table I), was a higher level of expression
of the F4/80 Ag (Fig. 6). Nevertheless, this level was considerably
lower than it was on macrophages. Some F4/80 fluorescence was obtained
with all DC populations, and even the
CD4-8- DC showed clear
F4/80 staining. However, the differences in expression were sufficient
to produce extensive depletion of the
CD4+8- DC with
anti-F4/80 and immunomagnetic beads and allowed recovery of most
CD4-8- and
CD4-8+ DC. Immunofluorescent staining of thymic DC
In contrast to splenic DC, thymic DC were not segregated into
clear subpopulations by immunofluorescent staining. As reported
previously (12, 15), a wide spread of CD8 staining was
obtained from low to the same high levels as T cells and splenic
CD8+ DC (Fig. 5). Staining for both CD8ß and
CD4 was obtained but at levels well below those characteristic of T
cells and thymocytes (Fig. 5). However, the majority of thymic DC were
CD11blow and DEC-205high,
resembling in this respect the
CD8+ß- splenic DC
(Fig. 5). Only a trail of CD8low thymic DC
showed lower staining for DEC-205 and higher staining for CD11b,
approaching but never equivalent to the splenic
CD8- populations. Staining for a series of
other markers and gating for CD8 expression (Fig. 7) showed that
CD8low, CD8int, and
CD8high thymic DC all expressed the markers
expected of mature DC and all were generally similar to the
CD8high DC of spleen; only the
CD8low DC showed some differences for a
proportion of the cells present, including some cells that were
F4/80low CD49flow, like
CD8- splenic DC, and some cells that were
class II MHChigh, CD11cint,
and CD86high, like activated DC.
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The moderate staining for CD4 and CD8ß on thymic DC (Fig. 5) led
to suspicion that these Ags may have been picked up from associated
thymocytes, as we had demonstrated previously for the moderate level of
Thy 1 on the surface of thymic DC (27). Accordingly, a
critical test for Ag pickup was applied to the CD8, CD8ß, and CD4
staining of all the DC, using CD8null mice (in
which neither CD8 nor CD8ß should be expressed on the cell
surface) and CD4null mice. In both of these gene
knockout mouse strains it could be demonstrated (using other markers)
that all DC subtypes were present despite lacking their characteristic
markers (20). Bone-marrow chimeras were constructed by
injecting irradiated mice with equal numbers of Ly-5.2,
CD8null, or CD4null and Ly-5.1
wild-type bone marrow cells (or as control chimeras, with Ly-5.2
wild-type and Ly-5.1 wild-type bone marrow cells) and then analyzing
the DC populations 5 wk later using the Ly-5 marker to separately
analyze the DC progeny. The test was to see whether the marker
expression disappeared when the gene was disrupted (in which case the
expression had been authentic) or if the staining persisted (in which
case it had been picked up from wild-type cells, presumably T lineage
cells). The DC in the chimeras derived from the wild-type bone marrow
gave staining results similar to those already demonstrated for normal,
nonirradiated mice in both the test and control chimeras. The results
for the DC derived from the CD8null or
CD4null bone marrow are compared with these
side-by-side wild-type DC in Fig. 8.
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staining largely disappeared from the
"CD8high DC" derived from
CD8null bone marrow, indicating that the
expression had been authentic. A variable but usually very slight level
of CD8 staining above background persisted, indicating a very low
level of Ag pickup. On thymic DC, most of the
CD8high staining also disappeared, again
indicating authentic CD8 expression; however, the staining did not
drop to the background, with all DC now showing a low to intermediate
staining, corresponding to the lower staining levels seen for about
half of the DC from wild-type bone marrow (Fig. 8). Thus, the CD8
staining of thymic DC appeared to be a mix of high-level authentic
expression by about half of the DC together with low-level pick-up of
Ag from associated thymocytes by all DC.
CD8ß staining was not seen above marginal levels on splenic DC but
was seen at moderate levels on thymic DC. This staining of thymic DC
persisted at the same level on the DC derived from
CD8null bone marrow, in which the absence of
the CD8 chain should prevent expression of CD8ß on the cell
surface (Fig. 8). Thus, all CD8ß staining on DC could be attributed
to Ag pickup.
On splenic DC, CD4 staining largely disappeared from the large
population of "CD4+ DC" when the CD4 gene was
disrupted, indicating that this had been authentic CD4 expression (Fig. 8). In contrast, the moderate staining of thymic DC with anti-CD4
persisted at normal levels even when the CD4 gene was disrupted,
indicating that all the CD4 on thymic DC, just like all the CD8ß on
these cells, could be attributed to pickup from associated
thymocytes.
Tests for authentic marker expression: overnight incubation
Another test for authentic DC surface-marker expression was to
check whether the marker persisted, rather than being washed off, after
a short culture period. However, this approach had the disadvantages of
incomplete elimination of absorbed Ags, changes in authentic marker
expression due to DC maturation in culture, and the known differential
loss of the CD8+ DC due to their slightly
faster rate of death in culture (28). The basic CD4 and
CD8 staining pattern of splenic DC persisted after culture, an
argument for authentic marker expression (Fig. 9). The level of CD4 staining dropped
noticeably but did not disappear, a result compatible with
"maturation." The level of CD8 staining remained high, as we had
noted previously (12, 15), but the relative number of
CD8+ DC decreased, compatible with a higher
proportion of CD8+ DC being among the 50% of
DC that died on overnight culture. The CD4 and CD8ß staining of
thymic DC was markedly reduced by overnight culture (Fig. 9) and in
some experiments was almost completely eliminated, a result more
compatible with surface adsorption than with real synthesis and
expression. Of particular note was the CD8 staining of thymic DC
after incubation because the DC then clearly fell into two staining
populations (Fig. 9). One group displayed persistent high CD8
staining compatible with authentic expression, as was seen with
CD8+ splenic DC. Although the number of such
CD8high thymic DC appeared to be reduced, this
may have been due to differential death in culture. The other group
after incubation showed low but still above background CD8 staining.
It could not be ascertained whether these were
CD8- DC that had not lost all the CD8
acquired from thymocytes or whether their low CD8 staining was a
consequence of low but still endogenous CD8 synthesis.
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A final test for authentic marker expression was to determine
whether the DC expressed mRNA coding for the surface molecules stained.
Such tests had already been performed for CD8 and CD8ß expression
with the conclusion that thymic DC and splenic
CD8+ contained mRNA for CD8 but not for
CD8ß (11). This approach was extended for CD4
expression, using RT-PCR (Fig. 10).
Splenic CD4+8- DC clearly
contained RNA transcripts for CD4, a fact that was evident even with a
low-sensitivity PCR, although the level of mRNA appeared less than that
seen with thymocytes. This would accord with a lower CD4 surface
expression on these DC compared with that on T cells. With a
low-sensitivity PCR, no band corresponding to CD4 mRNA could be seen
with thymic DC, CD4-8-
DC, or CD4-8+ DC. However,
by increasing the sensitivity with more PCR cycles, all DC populations
showed evidence of some CD4 mRNA, although at much lower levels than
with the CD4+8- splenic DC
population.
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Discussion |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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One important variable already recognized by most workers in the field is the procedure used for DC isolation, which can select particular DC subsets or alter their phenotype. In our earlier procedures we paid the price for overzealous elimination of all macrophage-like and lymphoid cells by the loss of a major subgroup of CD4-bearing F4/80+ adherent DC. However, insufficient DC enrichment before analysis makes flow cytometry and sorting difficult due to overlap of markers with other more numerous cell types to the presence of autofluorescent cells. It is of interest that this study has also pointed to a limitation of the earlier, adhesion-dependent isolation procedure of Steinman and coworkers (26), namely a strong selection for this same CD4-bearing adherent DC population. Thus, the earlier studies on mouse spleen DC from this laboratory and from the Rockefeller University laboratory would have been addressing different DC types with little overlap.
Another important problem when nonlymphoid cells are extracted from lymphoid tissues is autofluorescence. We have now demonstrated that the mature DC themselves are not markedly autofluorescent, at least when freshly extracted from tissues. Accordingly, the problem is finding the optimal means of eliminating the autofluorescent cells. Although reserving a fluorescent channel to gate out such cells is one solution, the fluorescence overlap between analysis channels and the limitation of color compensation procedures means this is a less-than-perfect option. Prior elimination of autofluorescent cells, for example by presorting, is a more effective but time-consuming option.
A problem that is directly associated with the DC themselves is the
pickup of surface Ags from other cells. This is especially marked with
thymic DC, which exist in the thymus tightly associated with thymocytes
as "rosettes" (29, 30) and which are in an environment
replete with dying thymocytes. It appears that DC are able to pick up
onto their outer surface Thy-1 (27), CD4, and CD8ß.
The CD8ß picked up is in addition to the CD8 that they make
themselves as an authentic surface protein. This "pickup" may be no
more subtle than fragments of thymocyte cell membranes attached to the
DC surface. Alternatively, there could be more specific binding of
CD8ß to class I MHC and CD4 to class II MHC. There is no evidence
that this pickup is of any functional significance.
These artifacts have previously obscured the analysis of thymic DC
populations and have led to the picture of a single DC group with a
wide distribution of CD8 expression. It now appears that there are two
more distinct groups, one CD8high and one
CD8low or even
CD8-. However, both of these are
CD8ß-, CD4-,
DEC-205+, and CD11blow and
appear very similar by a series of other surface markers. They may all
be one basic DC lineage, some of which have been exposed to factors
inducing CD8 expression and others that have not. In favor of this
view is the finding that all DC grown with high efficiency in culture
from the thymic DC/T cell early precursor population lack CD8
expression, whereas DC produced from the same precursor population on
intrathymic transfer into an irradiated recipient generate the normal
spread of CD8 expression seen in intact mice (16, 17, 18).
However, we cannot exclude the possibility that one of these DC
subtypes found in the thymus is of different origin and might migrate
into the thymus from the bloodstream.
The splenic DC seem less prone to pickup of T cell Ags, perhaps because
of a less-tight association with T cells or because many are not
located in the T cell zones of the spleen. In particular, the CD4 on
the surface of about half of the splenic DC appears to be authentic
expression of CD4 synthesized by the DC themselves. This resolves an
apparent discrepancy between murine DC and those of human or rat
origin: it is now clear that all three species have a proportion of DC
expressing CD4. As we have indicated previously (11), the
high CD8 on the surface of about one quarter of splenic DC reflects
true synthesis by these DC, an apparent point of difference with other
species. What does this expression of CD4 or CD8 imply?
We have argued previously that the lymphoid marker CD8 on splenic and
thymic DC reflects their origin from a lymphoid precursor cell similar
to the early lymphoid precursor we isolated from mouse thymus
(16, 17, 31). Although formal clonal proof of a common
lymphoid/DC precursor is still missing, it is clear that these DC
represent a distinct lineage with distinct cytokine requirements and
transcription factor control (18, 19). They also have a
very much greater potential to produce IL-12 than either of the
CD8- DC populations (Ref. 25 and H.
Hochrein, manuscript in preparation). Although CD4 is also considered
as a lymphoid marker, CD4 on DC does not necessarily reflect a lymphoid
origin. All the other markers, including F4/80 and CD11b, suggest that
these CD4+ but CD8- DC
resemble the CD4-8-
DC, which are presumed to be of myeloid origin. It should also be noted
that CD4 can also be expressed by monocytes.
The simplest explanation would be that these CD4+8- splenic DC are an earlier and less mature form of the CD4-8- myeloid DC type. By this argument, they should appear as immature by other markers and should on maturation lose CD4 to become the CD4-8- DC. However, several facts argue against this. First, although CD4 expression does drop on incubation of these cells, the DC remain clearly CD4+ with little evidence of an increase in the number of CD4-8- DC. Second, the CD4+8- DC show expression similar to that of the CD4-8- DC of the maturation markers class II MHC, CD80, and CD86, and they appear fully mature in their capacity to stimulate allogeneic T cells. Finally, in preliminary labeling experiments to determine the turnover and lifespan of these DC types in vivo, there is no evidence for a precursor-product relationship between CD4+8- DC and CD4-8- DC (A. Kamath, D. Tough, and K. Shortman, manuscript in preparation). Therefore, they may represent a third independent developmental stream. One possibility is that they, rather than the CD8+ DC, correspond to the human CD4+ plasmacytoid DC2 lineage (32). The lineage relationships between these three splenic DC types and their individual biological functions are currently under active investigation.
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Footnotes |
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2 Address correspondence and reprint requests to Prof. Ken Shortman, The Walter and Eliza Hall Institute of Medical Research, Post Office, Royal Melbourne Hospital, Victoria 3050, Australia. E-mail address:
3 Abbreviations used in this paper: DC, dendritic cell; PI, propidium iodide; int, intermediate.
Received for publication September 20, 1999. Accepted for publication January 5, 2000.
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References |
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TopAbstractIntroductionMaterials and MethodsResultsDiscussionReferences |
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+ and CD8- subclasses of dendritic cells direct the development of distinct T helper cells in vivo. J. Exp. Med. 189:587.- dendritic cells but not of lymphoid-related CD8+ dendritic cells. Immunity 9:839.[Medline]
+ or CD8- dendritic cells. Eur. J. Immunol. 27:3350.[Medline]
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50%
of the total) were removed, and the DC were stained and analyzed as in
Fig. 5