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HEMATOPOIESIS
From the University of Washington, Seattle, WA.
The hierarchy of cytoadhesion molecules involved in
hematopoietic/stem progenitor cell mobilization has not yet been
delineated. Previous studies have suggested an important role for
The physiologic egress of mature leukocytes from
bone marrow to peripheral blood, as well as the escape of a small
number of stem/progenitor cells from the normal bone marrow environment to the circulation, are poorly understood phenomena. The movement of
cells from the extravascular spaces of bone marrow to circulation may
require a coordinated sequence of reversible adhesion and migration
steps. The repertoire of adhesion molecules expressed by
stem/progenitor cells or by stromal cells in bone marrow is crucial in
this process. Alterations in the adhesion and/or migration of
progenitor cells triggered by diverse stimuli would likely result in
their dislodgment or redistribution between bone marrow and peripheral
blood. It has been shown over the last 3 decades that several empiric
in vivo treatments1 can evoke the release of
stem/progenitor cells from bone marrow into the circulation, although
the mechanisms involved were not uncovered. Because integrins appear to
be important regulators of adhesion and migration events in many
cellular systems,2 it has been speculated that this family
of molecules, which has a broad expression pattern in hemopoietic cells, also plays an important role in the mobilization of
stem/progenitor cells. Several in vitro3-5 as well as in
vivo6-11 observations support this contention.
Specifically, it was shown that in vivo use of function-blocking
antibodies against the Because mobilization is a complex, multistep process, it is likely that
several classes of cytoadhesion molecules are participating in this
process in a sequential and/or overlapping manner, as is the case with
mature leukocytes in well-studied models of
inflammation.13 In these latter studies the interplay of
selectins with Unlike mature leukocytes migrating through activated
endothelial surfaces, stem/progenitor cells migrate through normal
endothelia with a reverse directionality of movement (from tissues to
circulation rather than the opposite), and the hierarchy of molecules
involved in these movements has not been delineated. Despite these
differences, many of the same molecules, especially of the integrin
family (because of their wide distribution in hemopoietic cells) may work cooperatively in this process as well. To test the contribution of
Mice
Monkeys
Monoclonal antibodies Directly conjugated antimouse CD49d/ 4 (clone PS/2) was
purchased from Southern Biotechnology (Birmingham, AL). An unconjugated form of this antibody as well as another anti-4 antibody (R1/2) both
low in endotoxin and free of azide were kindly provided by Roy Lobb
(Biogen, Cambridge, MA). Other fluorescence-conjugated antibodies
against mouse cell surface epitopes, as well as low-endotoxin, azide-free forms used for injection were all purchased from Pharmingen (San Diego, CA). These included the following: CD11a/LFA-1 (clone M17/4), CD11b/Mac-1 (clone M1/70), CD18/ 2 (clones C71/16, M18/2, and
GAME-46), CD49e/5 (clone 5H10-27), and CD117/Kit (clone 2B8). An
additional antimouse CD18 (2E6) was purified from hybridoma cells
purchased from American Type Culture Collection (Manassas, VA)
and was provided by R. Winn (Seattle, WA). For depletion of lineage-committed cells, a cocktail of antibodies was used: GR-1, Mac-1
(CD11b), L3T4 (CD5), B220 (CD45R), and Ter-119, all rat antimouse
immunoglobulin G (IgG) purchased from Pharmingen. Magnetic beads
labeled with goat antirat IgG from Miltenyi Biotec (Auburn, CA) were
used to collect lineage depleted (Lin ) cells according to
the manufacturer's protocols.
Monoclonal antibody 60.3 (IgG2a)18 recognizes a functional
epitope on the common Fluorescence-activated cell sorter analysis and sorting Confirmation of genotype in CD18 knockout and CD18 hypomorphic animals was performed on peripheral blood white blood cells (WBCs) stained with anti-CD18 fluorescein isothiocynate (FITC) antibody and analyzed using a FACSCalibur (Becton Dickinson Immunocytometry Systems, San Jose, CA). Cell sorting of Lin samples was performed
on a FACSVantage (Becton Dickinson).
Genotyping Progeny of CD18+/ mice were genotyped at 3 weeks
of age by polymerase chain reaction amplification of tail DNA samples.
Presence of the wild-type allele was detected by the primers
5'-CTGGACTGTTCTTCCTGGGATC-3' (forward) and 5'-GTACTTGGTGCATTCCTGGGAC-3'
(reverse), whereas presence of the targeted allele was detected by the
primers 5'-CTGGACTGTTCTTCCTGGGATC-3' (forward) and
5'-CTCGATGCGATGTTTCGGTTGGTG-3' (reverse). Amplifications were performed
on a Stratagene Robocycler (La Jolla, CA) for 35 cycles with an
oligonucleotide annealing temperature of 54°C. Of the 123 animals
tested in this manner, 16% were CD18/, a frequency
consistent with previous reports.16 Genotypes of the
knockout mice were routinely confirmed by fluorescence-activated cell
sorter (FACS) analysis of peripheral blood samples with an anti-CD18
antibody as described above.
Clonogenic progenitor cell assays Blood from mice was collected in preservative-free heparin. The WBC count was measured with a Coulter (Hialeah, FL) particle counter using a Zap-oglobin II-lysed sample, which was then corrected for heparin volume. The remainder was washed, lysed with hemolytic buffer, washed, and cultured in methylcellulose (MC) as previously described.20 Cells from the femoral bone marrow and spleen were similarly washed and cultured, without lysis. Blood from primates was collected in ethylenediaminetetraacetic acid for complete blood counts or preservative-free heparin for cultures.11 Cultures were incubated at 37°C in 5% CO2 and saturating humidity for 7 days (murine cultures) or 14 days (primate cultures), and colonies were counted based on morphologic criteria under a dissecting microscope. Colonies were classified as burst-forming units-erythroid (BFU-E), colony-forming units-granulocyte macrophage (CFU-GM), or CFU-granulocyte, erythroid, macrophage, megakaryocyte (CFU-GEMM) or were totaled and reported as CFU-culture (CFU-C).Assays for chemokines and cytokines Chemokine and cytokine levels in primate plasma were analyzed by the Cytokine Analysis Laboratory, Fred Hutchinson Cancer Center, Seattle, WA.Statistics Statistical analysis was performed using Student t test as found in Microsoft Excel software, using 2-tailed analysis assuming unequal sample variance.
Treatment of mice with anti-integrin antibodies Groups of at least 5 mice received intravenous treatments with various purified (endotoxin-free) anti-integrin antibodies using the same dose and schedule, ie, 3 injections over 3 days (1 injection daily of 2 mg/kg of body weight, Figure 1). The animals were bled the following day for determining total leukocyte count and for inoculation into clonogenic MC cultures for quantitation of circulating progenitors. Doses of antibodies used were derived from our previous experience with anti-VLA48,9,20 or from data by other investigators.10 Although a single injection of anti-4
(clone PS/2) induced a significant mobilization, single injections of
each of 2 anti-CD18 antibodies (GAME-46 and 2E6) were ineffective
(Figure 2 and data not shown).
Also, a single injection of anti-CD11a (LFA-1) was previously reported
to be ineffective.21 Results with groups of mice injected
3 times with anti-CD11a, anti-CD18 (GAME-46), anti-5, or anti-4
are presented in Figure 1. Although
differences before and after treatment seen with all tested antibodies
were statistically significant (P < .002), anti-4
treatment, as seen in Figure 1, was superior to the others in
mobilization (P < .0001). A different anti-4 antibody
(clone R1/2) in an independent experiment gave results similar to PS/2:
pretreatment, 101.7 ± 58.8 CFU-C/mL; posttreatment (2 mice), 375 and
1709 CFU-C/mL. Most CFU-C mobilized by PS/2 were of GM-type (92%),
similar to the pattern seen in baseline circulating CFU-C (95% are
CFU-GM). However, in these mice, BFU-E increased from 5.3 ± 1.8/mL
of blood at baseline to 35 ± 8.2/mL, and CFU-GEMM increased from
2.5 ± 1.0/mL to 19.4 ± 6.2/mL, N = 15-17. The increase of BFU-E
in mice after anti-4 is consistent with preferential BFU-E increase
in primates reported earlier.11 During anti-CD11a
treatment, there was also a notable increase in circulating BFU-E (from
2.0 ± 0.8 to 52.2 ± 7.7 BFU-E/mL, N = 5 mice). To test whether
BFU-E or other progenitors were indeed expressing LFA-1, we sorted
Lin/kit+/LFA-1+ or
kit+/LFA-1 cells and inoculated these in
clonogenic MC media. The data (Table 1) show that although the
frequency of progenitors is higher (2-fold) in the
Lin/kit++/LFA-1 fraction, the
total number of progenitors present in the
kit++/LFA-1+ fraction is higher because more
kit++ cells were LFA-1+ than
LFA-1. Of interest, all BFU-E were present in the
kit++/LFA-1+ population. In addition to the
anti-CD18 antibody used in Figure 1 (GAME-46), a 3-day treatment with
another anti-CD18 (M18/2) yielded a smaller increase in CFU-C from
102 ± 15/mL at baseline to 158 ± 60/mL of blood. This antibody
was reported to inhibit metastasis22 when used in vivo.
However, when tested in vitro,23 it did not inhibit
adhesion of LFA-1+ cells to intercellular adhesion molecule
(ICAM)-1, thus questioning its mode of action in vivo. In all anti-CD18
treatments, no increase in BFU-E or CFU-GEMM was seen. Collectively,
the data from all 3 anti-CD18 treatments would support the concept that
if there is an increase in CFU-C, it is likely modest and is not
accompanied by a concurrent increase in WBCs, questioning their in vivo
effectiveness.
Combined anti- 4 administered in combination with the
other antibodies had an additive or synergistic effect on mobilization, mice (at least 5 each) were treated with the same schedule and doses as
above but either with or without the anti-4 (ie, 3 injections, once
daily) and bled the day after the third injection. In all treatments
with coadministered anti-4, augmentations in mobilization were seen
(Figure 1). Interestingly, the combined anti-4 and anti-CD11a
treatment was highly effective in increasing further BFU-E mobilization
from 52.2 ± 7.7 BFU-E/mL with anti-CD11a alone to 292.2 ± 43
BFU-E/mL after the combined treatment (also see Table 2).
Treatment of monkeys with anti-CD18, either alone or in combination
with anti- 4
(clone HP1/2) antibodies of known efficacies. Either 1, 2, or 3 injections of anti-CD18 alone were given. Observations before, during,
and several days after treatment were made. Treatment of 2 monkeys with
either 1 (Figure 2, lower panel) or 2 (Figure
3, upper panel) injections of anti-CD18
antibody did not elicit significant mobilization during the first 3 to
4 days despite a prominent increase in WBC count, confirming our
earlier observations.11 Assays of blood continued for
several days after treatment revealed a significant elevation in
mobilized CFU-C between days 5 and 8 posttreatment in the animal given
2 injections. This increase was not as evident after only 1 injection
(data not shown). At the time the CFU-C peak was noted, the WBC count
had returned to normal and circulating antibody was no longer
detectable in these animals, as indicated by specific ELISA performed
in R. Winn's laboratory.26 (These 2 monkeys had plasma
levels of circulating 60.3 antibody of 0.96 µg/mL and 3.1 µg/mL 24 hours after a single injection.) One of these 2 monkeys was given a
second injection of 60.3 at 2 mg/kg (Figure 3, upper panel), which then
resulted in a plasma level of 5.0 µg/mL 24 hours later. (The data
from another monkey given 3 injections of a-CD18 antibody at 2 mg/kg
[data not shown] did not differ significantly from this animal.) When
anti-CD18 was combined with anti-VLA4 in another animal (one daily
injection at 2 mg/kg each for 2 days), a dramatic synergy was seen with a delayed mobilization peak (Figure 3, lower panel). Because the anti-4 antibody used in this animal was the humanized version with a
long half-life of over 10 days8,27 (unpublished
data, 1993), its influence on mobilization was prolonged
(Figure 3, middle panel). Whatever the mechanism for the delayed peak
observed with anti-CD18, it worked synergistically with the continued
inhibition of VLA4 function to augment mobilization.
Cytokine/chemokine levels in monkeys following anti-CD18 treatment To test whether the delayed peak observed in monkeys after 2 injections of anti-CD18 was due to the potential release of cytokines or chemokines in circulation, we performed ELISAs to test for the presence of interleukin (IL)-8, macrophage inflammatory protein (MIP)-1, monocyte chemoattractant protein (MCP)-1, IL-1, and stromal cell-derived factor (SDF)1 chemokines that have been
previously implicated in mobilizationor for cytokines like
granulocyte colony-stimulating factor (G-CSF), kit ligand, or
macrophage colony-stimulating factor (M-CSF). The results are shown in
Table 2. No consistent or significant increases were noted from this study to allow firm conclusions.
Treatment of CD18-hypomorphic or CD18-deficient mice Because antibody treatments in vivo are difficult to interpret and may differ on occasion from results in gene-targeted mice,28 we tested whether synergy in mobilization between1 and 2 integrins could be seen in CD18-hypomorphic or in
CD18/ mice. CD18 expression in the peripheral blood
cells of the CD18-hypomorphic or CD18/ mice was tested
by cell surface staining with anti-CD18 antibody (C71/16). As expected
from previous studies, it was approximately 15% in hypomorphic
mice,16 whereas no expression of CD18 was seen in white
cells of CD18-deficient mice17 (Figure
4). The bone marrow cellularity of the
latter showed a mild increase with a predominance of myeloid elements
(data not shown). Baseline WBC counts were higher, similar to
previously reported values in CD18 null mice17 (6 mice:
WBC/µL 33 831 ± 3677) compared with CD18 hypomorphic mice
(16 animals: WBC 11 480 ± 522). Also, circulating CFU-C were higher
in CD18/ mice (6 mice: 399 ± 24 vs 227 ± 18.9/mL
of blood in hypomorphic mice). However, in our antibiotic-treated
colony of CD18 null mice, both the WBCs and the CFU-C were lower
(5 mice: WBC = 17 050 ± 1261 and
CFU-C = 154 ± 47/mL of blood). Treatment of these mice
with anti-4 antibody elicited a significant mobilization that was
several-fold higher (Figure 5) than that
observed in control mice. Treatment of 14 B6.129+/+ mice
with anti-4 alone elicits an increase of an average of 5-fold above
baseline (range 3- to 12-fold). In 5 anti-4-treated CD18
hypomorphic mice, the increase was about 7-fold (201 ± 34 to 1387 ± 107 CFU-C/mL of blood), not significantly
different from controls. However, in 5 similarly treated
CD18/ mice, CFU-C increased 49-fold (from 154 ± 47
to 7527 ± 2411 CFU-C/mL [Figure 5]; WBC: from 17 050 ± 1261 to
74 694 ± 11 003). This difference was unlike any increase in other
strains of mice that we have treated thus far (25 mice of 6 different
strains gave an average of 5.4-fold increase with a range of 3.2- to
12-fold). To test whether the synergy in mobilization was delayed or
extended in CD18/ mice treated with anti-4 (as
observed in primates), a single injection of anti-4 was given with
assessment of circulating CFU-C daily. Anti-4 alone given to +/+
littermates showed a 12- to 24-hour mobilization peak (Figure 2). By
contrast, mobilization in CD18/ mice after a single
anti-4 injection was maintained at peak levels over the first 3 days
postinjection (2 animals, data not shown). Collectively, the data in
gene-targeted mice and those with antibody blockade are consistent with
the cooperativity between 1 and 2 integrins and suggest that both
1 and 2 integrins are likely responsible for progenitor anchoring
within the bone marrow in vivo and that abrogation of their function
leads to enhanced migration out of the bone marrow.
In the present study, we have documented that, in addition to
Although a synergistic effect in mobilization was seen with inhibition
of both Are stem/progenitor cells the targets of the combined anti- Studies in monkeys were particularly instructive. The synergistic
effect of Because of the difficulties in pinpointing the in vivo effects of
antibodies,28 we sought to complement our data using mice deficient in CD18 integrins. If abrogation of CD18 function is important for the synergy with anti- Several examples on the cooperativity between
We kindly thank Arthur L. Beaudet for providing the CD18 knockout mice and Margaret Oppenheimer for her expert secretarial assistance.
Submitted August 7, 2000; accepted October 24, 2000.
Supported by National Institutes of Health Grants AI32177, DK46557, and RR00166.
The publication costs of this article were defrayed in part by page charge payment. Therefore, and solely to indicate this fact, this article is hereby marked "advertisement" in accordance with 18 U.S.C. section 1734.
Reprints: Thalia Papayannopoulou, Division of Hematology, University of Washington, Box 357710, Seattle, WA 98195-7710; e-mail: thalp{at}u.washington.edu.
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Abstract
Introduction
