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Blood, 1 July 2005, Vol. 106, No. 1, pp. 86-94. Prepublished online as a Blood First Edition Paper on March 15, 2005; DOI 10.1182/blood-2004-09-3417.
HEMATOPOIESIS VCAM-1 expression in adult hematopoietic and nonhematopoietic cells is controlled by tissue-inductive signals and reflects their developmental originFrom the Division of Hematology, University of Washington, Seattle, WA; and the Program in Molecular Immunology, Institute of Molecular Medicine and Genetics, Medical College of Georgia, Augusta.
Although expression of vascular cell adhesion molecule 1 (VCAM-1) in endothelial cells and its functional implications have been previously appreciated, VCAM-1 expression in other than endothelial cells, especially hematopoietic cells, has been recently recognized and has not been explored in detail. Using normal mice and mice with a conditional ablation of VCAM-1 through a Tie2-driven cre transgene, we have studied the biodistribution and the pattern of VCAM-1 expression in circulating versus tissue-residing cells before and after their enforced mobilization. In the normal mouse, both at basal hematopoiesis or following mobilization, VCAM-1 expression is confined to myeloid cells residing in hematopoietic tissues, whereas free cells in circulation or in body cavities are devoid of VCAM-1 messenger RNA (mRNA) and protein. However, following culture, proliferating myeloid cells, but not lymphoid cells, express VCAM-1. In the VCAM-1–ablated mouse, there is an increase in circulating progenitors as a consequence of their ongoing release from bone marrow, a process enhanced by splenectomy. We postulate that the main mechanism leading to their release is the ablation of VCAM-1 by fibroblastic and by endothelial cells. Ablation of VCAM-1 in fibroblasts by Tie2-driven cre is a novel finding and likely denotes their developmental ancestry by Tie2-expressing (mesenchymal?) progenitor cells during development.
The vascular cell adhesion molecule 1 (VCAM-1), an immunoglobulin (Ig)–like transmembrane adhesion molecule, is highly conserved in evolution and participates in a variety of cellular functions in health and disease. Human VCAM-1 has 2 isoforms, the predominant 7 Ig-domain isoform and a minor, alternatively spliced isoform with 6 Ig domains, whereas in mice the second isoform consists of the first 3 domains attached to the cell membrane through a glycosylphosphatidylinositol (GPI) anchor.1,2 VCAM-1 is minimally expressed on most resting vascular endothelial cells and is inducible in many tissue vascular beds following injury or stress. Because of this activation, VCAM-1 has been implicated in the pathophysiology of certain autoimmune diseases, atherosclerosis, and allograft rejection.3-8 VCAM-1 is constitutively expressed in bone marrow (BM) stromal/endothelial cells and certain classes of hematopoietic cells (B cells, follicular dendritic cells, and macrophages9-15). Its major ligand is the integrin very late antigen 4 (VLA-4;  4 1) with binding sites located in the first and fourth Ig domains, whereas other ligands bind with less affinity and include 47, 91, and D2.16-18 Deletion of the VCAM-1 gene produced a phenotype similar to deletion of the 4 gene, with embryonic lethality due to cardiac and placental developmental abnormalities, although few VCAM-1–/– mice survived.19,20 Several investigators then adopted approaches to overcome the embryonic lethality observed in VCAM-1–null mice. Cybulsky et al generated mice with a homozygous deletion of domain 4 (VCAM-1D4D),21 resulting in expression of the VCAM-1 isoform containing one binding site. Embryonic survival was significantly reduced and was strain dependent, but surviving mice showed a reduction of VCAM-1 protein to less than 10% of wild type, allowing informative studies in the adult animals.22-24 In other studies,25 Tie2-cre transgenic mice were bred with floxed VCAM-1 mice to allow VCAM-1 deletion in hematopoietic cells and endothelium through cre-loxP–mediated recombination. These viable mice exhibited mild leukocytosis with elevated immature B cells in the peripheral blood (PB), with reduced immature B cells in the BM, consistent with the role of VCAM-1 in the retention of immature lymphocytes in the marrow. In a similar approach, mice carrying the Mx-cre transgene bred with mice homozygous for the floxed VCAM-1 gene, subsequently deleted neonatally by interferon-mediated cre recombination, led to a reduction of immature B cells in the BM, and an increase of these cells in the circulation.26 Additional in vitro studies have shown that the VLA-4/VCAM-1 interaction is involved in the adhesion of human B cells to follicular dendritic cells of germinal centers in vitro and a link has been reported between VCAM-1 function and the rescue of germinal center B cells or thymocytes from apoptosis.13,14,27 VCAM-1 may also function in maturation and costimulation of T cells.28-30 In contrast to results from chimeric 4–/– mice showing postnatal dependency of B and T cells on 4 integrins,31 B and T lymphocytes can develop in the absence of VCAM-1.25,26 Later studies reported that neonatally induced inactivation of VCAM-1 impaired responses to Toxoplasma encephalitis.32 Herein, we explored the pattern of VCAM-1 expression among hematopoietic and nonhematopoietic cells from BM and from selective tissues, before and after the use of mobilizing agents. Several novel insights about VCAM-1 regulation in these cell populations are revealed by studying both control and VCAM-1–ablated animals and these are described in the present work.
Mice
VCAM-1 mice used in our experiments were created through interbreeding of VCAM-1 floxed (VCAM-1f/f) mice with Tie2cre transgenic mice.25 When VCAM-1f/f males are bred with Tie2cre+ females, all progeny, regardless of the presence of the cre transgene, have a constitutional deletion of 1 VCAM-1 allele (VCAM-1 Mobilization treatments Granulocyte–colony-stimulating factor (G-CSF; Neupogen Filgrastim; Amgen, Thousand Oaks, CA) treatments were given via an osmotic pump (Alzet Model 2001; Durect, Cupertino, CA) for 5 days at 100µg/kg per day (3 experiments) or by intraperitoneal injection twice daily at 200 µg/kg for 4 days (1 experiment). All mice were tested on the fifth day. Flt-3 ligand alone (FL; 50 ng/kg per day; kind gift from S. Lyman, Immunex, Seattle, WA) or the combination of G-CSF (100 µg/kg per day) and FL was administered with osmotic pump for 7 days. Antibodies Anti–VCAM-1 (MK/2) antibodies, unconjugated for immunohistochemistry (IHC) and phycoerythrin (PE)–conjugated for FACS, were from Southern Biotech (Birmingham, AL). Fluorochrome-conjugated antibodies to B220, CD3, TER119, Mac-1, Gr-1, and CD117 (c-kit), and fluorescein isothiocyanate (FITC)–conjugated secondary antibodies and streptavidin allophycocyanin (APC) were from BD-Pharmingen (San Diego, CA). Antibodies used for endothelial cells were biotinylated CD105 (endoglin), MECA32 (BD-Pharmingen), biotinylated isolectin B4 (Vector Labs, Burlingame, CA), labeled acetylated low-density-lipoprotein (diI-Ac-LDL; Biomedical Technologies, Stoughton, MA), rabbit anti–human von Willebrand factor (VWF; Abcam, Cambridge, MA), and PE-conjugated anti–mouse CD202 (Tie2; eBioscience, San Diego, CA). Anti–rabbit Alexa 594, streptavidin 488 (Invitrogen/Molecular Probes, Carlsbad, CA), and streptavidin PE (Biomeda, Foster City, CA) were used as secondary reagents. FACS analysis Antibody-stained cells were analyzed on a FACSCalibur (BD Immunocytometry Systems, San Jose, CA) using the CELLQuest program. Immunohistochemistry Tissues for immunohistochemistry were fixed in 4% paraformaldehyde, sucrose treated, embedded in TissueTek (Sakura Finetechnical, Tokyo, Japan), and frozen in acetone/dry ice. Femurs/tibiae were similarly fixed, decalcified in 10% EDTA (ethylenediaminetetraacetic acid; 7 to 10 days at 4°C), sucrose treated, then embedded and frozen. Air-dried frozen sections were washed, blocked for endogenous peroxidase activity and nonspecific staining, stained with appropriately diluted antibodies, and processed following the manufacturer's instructions (ABComplex/HRP; Dako, Carpinteria, CA). Culture–colony-forming unit assay Culture–colony-forming unit (CFU-C) assays were performed using a methylcellulose mixture (Methocult GF; Stem Cell Technologies, Vancouver, BC, Canada), as described previously.33 Endothelial and fibroblast cell cultures Endothelial cells from bone were derived by treating cleaned and flushed femurs with 0.125% Trypsin/0.2% EDTA for 15 minutes at 37°C; cells were discarded, bones minced, then incubated with 0.1% collagenase (1-3 hours at 37°C). After washing and removing bone fragments, cells were plated in collagen I–coated wells (BD BioCoat; BD Biosciences, Bedford, MA) using endothelial-cell medium as previously described,34 adding 250 ng/mL endothelial mitogen (Biomedical Technologies), 10 ng/mL recombinant murine vascular endothelial cell growth factor (rm-VEGF) and 7.5 ng/mL basic fibroblast growth factor (both from Peprotech, Rocky Hill, NJ). Nonadherent cells were removed after 24 hours and fresh medium was added. Adherent cells were dissociated with either 1:1 cell dissociation solution/phosphate-buffered saline (PBS) (Sigma Aldrich, St Louis, MO), or 1:10 TrypLE Express/PBS (Invitrogen). Bone fibroblasts were derived by plating bone fragments or cells from above in Dulbecco modified Eagle medium (DMEM) with 20% fetal bovine serum (FBS), cultured for 4 to 7 weeks, and dissociated with TrypLE Select. Endothelial cells from heart or lung were isolated by mincing, then washed extensively to remove blood, collagenase treated and plated in 2% gelatin-coated wells/dishes with endothelial cell medium. Endothelial cells or fibroblasts were used for evaluation by fluorescence-activated cell sorting (FACS) or for DNA and RNA studies. Bone marrow, peripheral blood, and splenic lymphocyte cell cultures Femurs from control and VCAM-1–ablated mice were flushed, and recovered cells were washed, incubated for 2 hours in Iscoves modified Dulbecco medium (IMDM) plus 10% FBS to remove adherent cells, counted and plated in IMDM with 10% FBS, 100 ng/mL murine stem-cell factor (Peprotech), 10 µL/mL of murine interleukin 3 (IL-3; kind gift from K. Kaushansky, San Diego, CA), 100 ng/mL G-CSF, 50 ng/mL granulocyte macrophage–colony-stimulating factor (GM-CSF; Peprotech), and 3 U/mL erythropoietin (Epoetin; Amgen, Thousand Oaks, CA). Mononuclear cells from blood of mobilized control or ablated animals were isolated by density gradient separation (Histopaque 1083; Sigma Chemical), washed, and cultured in the same media as BM cells. Spleens from control and VCAM-1–ablated mice were teased, triturated, washed, and plated in 10% FBS, 100 ng/mL murine IL-2, 50 ng/mL murine IL-7 (both from Peprotech), and 10–4 M 2-mercaptoethanol (Sigma Chemical), or in the same cytokine cocktail as the PB cells. Samples were collected for analysis at day 0 and after 3 to 7 days in culture. DNA/RNA studies
Genomic DNA and total RNA were isolated from mouse tails or cell cultures using DNA or RNA isolation kits (Gentra Systems, Minneapolis, MN) according to the manufacturer's instructions. Isolated RNA was reverse-transcribed using oligo-dT (Invitrogen) or random nonamer (Sigma Chemical) primer and SuperScript II enzyme (Invitrogen). The following primers were used in polymerase chain reactions (PCRs): for genomic DNA to distinguish between wild type, floxed, and Enzyme-linked immunosorbent assay (ELISA) Levels of stromal cell–derived factor 1 (SDF-1) and soluble VCAM-1 (sVCAM-1) were assayed in the cytokine analysis core facility of Fred Hutchinson Cancer Research Center, as previously described.37 SDF-1 and sVCAM-1 kits were purchased from R&D Systems (Minneapolis, MN). Cell-cycle analysis Bone marrow cells, permeabilized for 15 minutes in PBS with 10 mM EDTA and 0.1% Triton X-100, were mixed with propidium iodide at 2 µg/mL and analyzed by FACS using CELLQuest and ModFit LT (Verity, Topsham, ME) software. Statistical tests Tests of significance were performed using the Student t test (Microsoft Excel 2000 software; Microsoft, Redmond, WA).
VCAM-1 expression in hematopoietic cells
VCAM-1 expression in hematopoietic cells was assessed by flow cytometry using well-characterized antibodies. In control mice of young or old age ( In contrast to high VCAM-1 positivity in the hematopoietic tissues of normal mice, circulating cells were virtually devoid of VCAM-1 expression and as such, they were no different from VCAM-1–ablated mice (Figure 2B).
VCAM-1 expression in nonhematopoietic cells VCAM-1 expression in nonhematopoietic cells was initially assayed by immunohistochemical analyses of tissues. In control mice, endothelial cells in the BM, spleen, and lymphoid tissues (lymphatic and blood vessels) were clearly positive for VCAM-1, as were some stromal cells of the thymus (Figure 3). There were no VCAM-1+ endothelial cells in tissues from ablated mice, although rare, scattered positive cells could be seen. We next initiated endothelial cell cultures from bone, lung, or heart, and tested them after in vitro expansion. Expression of endothelial cell markers (Tie2, VWF, diI-Ac-LDL, and isolectin B4 uptake38) was seen in all cultures by immunofluorescence labeling, and the cells formed tubular networks in Matrigel (Becton Dickinson, Bedford, MA; see also Supplemental Figure S1B, available at the Blood website [see the Supplemental Materials link at the top of the online article]). VCAM-1 mRNA was detected in endothelial cell cultures from bone, lung, and heart (Figure 4 and data not shown) from control animals, and at much lower levels in endothelial cell cultures from ablated animals, in contrast to the virtual absence of VCAM-1 mRNA from hematopoietic cells residing in bone marrow from the same animals (Figure 4).
In addition to endothelial cell cultures, the level of VCAM-1 ablation was tested in stromal cell cultures (positive for DDR-2 and procollagen 1 1; Figure 4 and data not shown) from crushed bones. Unexpectedly, we found a significant level of VCAM-1 gene excision in fibroblast cultures from ablated animals (Figure 4), similar to that seen in some endothelial cell cultures. Parallel cultures of early passage fibroblasts from control animals showed VCAM-1 expression by virtually all CD45– cells (Supplemental Figure S2B). Also, Tie2 mRNA was detected in cultured fibroblasts (Supplemental Figure S2A), but at lower levels than in cultured endothelial cells. Basal hematopoiesis in VCAM-1–ablated animals in the presence or absence of the spleen Bone marrow cellularity and progenitor content were tested in VCAM-1–ablated young animals (9-14 weeks old) as well as in older animals (> 40 weeks old) to uncover age-related changes. As seen in Table 1, BM cellularity of young adult VCAM-1–ablated animals was lower than controls. This difference, likely attributable partially to their smaller (about 10%) weight, was maintained when older animals from both groups were compared. There was a moderate decrease in BM progenitor content in young ablated animals, but in older animals of either genotype, no significant differences were seen, although progenitors were significantly increased in both, consistent with previously published data.39 The proportion of lineage-affiliated cells, Gr-1+, Mac1+, and TER119+, in BM was not significantly different between the 2 groups (Supplemental Table S1). However, there were lower levels of T cells (P < .05) and B220+ cells.
A modest but significant increase was seen in spleen cellularity of ablated animals, which persisted with age, when the total cellularity increased in both groups (Table 1). There were also more splenic progenitors in ablated animals, both young and old, but no significant differences in the proportion of B cells, T cells, myeloid, or TER119+ erythroid cells in the spleen. Significant and consistent differences in circulating cells were found between ablated and control groups. Circulating white blood cells (WBCs) were significantly higher in the ablated group (Table 1), with a higher proportion of B220+ cells and a relative decrease in Mac-1+ cells (Supplemental Table S1). More pronounced and sustained differences were seen in the number of circulating hematopoietic progenitors (CFU-Cs). It is notable that the progenitor differences in peripheral blood between VCAM-1–ablated animals and controls were maintained with age (Table 1).
Since cellularity and progenitor content in the spleen were higher in VCAM-1–ablated mice, we suspected that the spleen continuously sequesters progenitor cells from circulation, so we studied splenectomized animals. Both WBCs and circulating progenitors increased (Figure 5) in splenectomized mice (5.3-fold for CFU-GMs, 22.4-fold for BFU-Es, and 6.2-fold for CFU-GEMMs). In the BM of splenectomized animals, there was an increased CFU-C content per femur (149 376 ± 15 131/femur in controls compared with 112 065 ± 17 447 in the ablated group), but no significant differences in the proportion of different lineages within this population. It is speculated that the increase in progenitor content in the marrow after splenectomy is attributable at least in part to some rehoming of circulating progenitor cells.40 Mobilization of hematopoietic progenitors in VCAM-1–deficient adult mice
To test the ability of VCAM-1
We then examined the response to G-CSF (G-CSF-5d; Figure 6). WBCs increased similarly in VCAM-1f/f and VCAM-1
Since there was considerably less CFU-C accumulation in the spleens of VCAM-1
To further pursue the issue of decreased splenic response in ablated mice, we used more intense mobilization with combinations of G-CSF and FL.42-44 Following a 7-day mobilization with G-CSF plus FL, a strong response was elicited by both control and VCAM-1–ablated animals well above levels achieved with either cytokine alone (Figure 6). However, the total progenitor content in BM, spleen, and blood was again higher in control mice (7.64 ± 2.1 x 106 CFU-Cs in f/f vs 4.43 ± 1.35 x 106 in
There was very little VCAM-1 expression in circulating cells after mobilization in both control and deficient animals (Figure 2B). To explore whether VCAM-1 expression could be induced in mobilized cells, the latter were cultured in a cytokine cocktail. After 3 days in culture and at a time when there was no appreciable increase in cell number, there were significant increases in VCAM-1 mRNA (Figure 7) and protein expression (from < 1% to 10.1%). There was an additional increase (to 28.6%) in VCAM-1+ cells at day 6 following cell proliferation. To test whether all proliferating cells express or activate VCAM-1 expression, we also cultured splenocytes under IL-7 and IL-2 stimulation. Despite significant proliferation, the latter failed to show VCAM-1 expression (Figure 7). Further, since VCAM-1 can be up-regulated in nonproliferating endothelial cells by inflammatory stimuli, we also tested induction of VCAM-1 expression in mature hematopoietic cells from control mice following an inflammatory stimulus (thioglycolate-induced peritonitis). Cells accumulated in high numbers in the inflamed peritoneum, but no expression of VCAM-1 was seen at either 4 or 16 hours after thioglycolate injection (data not shown). SDF-1 and sVCAM-1 levels after mobilization
Because of the previously emphasized role of SDF-1 in mobilization, we explored changes in SDF-1 levels in BM and PB after mobilization. Significant decreases in BM SDF-1 levels at the peak of mobilization were seen in wild-type, VCAM-1f/f, and VCAM-1
VCAM-1 expression in hematopoietic cells is tissue associated Although VCAM-1+ cells of hematopoietic origin have been previously noted, no systematic studies of VCAM-1 expression patterns among hematopoietic cells at baseline and following stress have been reported. VCAM-1 expression in normal BM is present among cells of all lineages, both immature and mature. The overall expression of VCAM-1 in almost half of BM cells is consistent with VCAM-1 expression in a subset of progenitor cells or kit+ cells, and likely a subset of primitive stem cells, although further studies are needed to document this notion. Thus, VCAM-1 expression in at least a common myeloid progenitor (CMP), may question a prior conclusion26 about the expression and functional influence of VCAM-1 only in late cells, beyond the stage of common progenitors.
In contrast to VCAM-1 positivity of myeloid cells in BM and spleen, it is of interest that PB cells do not display VCAM-1 (Figure 2B). This is true not only for cells during steady-state hematopoiesis, but also after cytokine-induced mobilization, when significant increases in the numbers of progenitor and mature cells are seen in periphery. To explore the reasons for the absence of VCAM-1 expression by PB cells, we considered the following possibilities: whether VCAM-1– cells were preferentially mobilized, both at steady state and after G-CSF; whether VCAM-1 positivity was masked or VCAM-1 expression was silenced once the cells were in the PB environment; or whether another common characteristic of cells in the PB was responsible for the VCAM-1– phenotype. If preferential mobilization of VCAM-1– cells occurred, one would expect enrichment for VCAM-1+ cells in the BM of controls at the peak of mobilization. Instead, a decrease in VCAM-1 expression was found in BM at the peak of mobilization (Figure 2) in G-CSF and a further decrease in G-CSF plus FL (from 49.6% ± 2.8% to 14.8% ± 0.4%). The possibility of a masking epitope was a viable one; therefore, we examined transcription of VCAM-1 in PB cells. We found that PB cells, from animals both mobilized and at steady state, were not only negative for VCAM-1 protein, but had no detectable levels of VCAM-1 mRNA (Figure 7). Next, we explored whether the known differences in cell cycling between PB and BM46,47 might be responsible for differences in VCAM-1 expression. Indeed, after cytokine-induced proliferation in vitro, mobilized cells displayed a significant expression of VCAM-1 mRNA and protein. However, in vitro culture could have generated new progeny with a VCAM-1–expressing phenotype from VCAM-1– cells, but this was unlikely since VCAM-1 levels were increased prior to increases in cell numbers, suggesting activation of VCAM-1 before cell division. Although these data cannot entirely dissociate activation alone versus cells in cycle under a strong cytokine influence, the fact that proliferating lymphoid cells in vitro do not express VCAM-1 suggests some specificity for myeloid cells, or the activation of molecular pathways leading to VCAM-1 expression only in myeloid cells. Whatever the mechanism, differences in VCAM-1 expression between PB and BM or spleen suggest that VCAM-1 expression is actively maintained by tissue-derived signals, but is down-regulated or silenced in the periphery. This view is consistent with absence of VCAM-1 expression in free cells (macrophages) in peritoneal fluid in thioglycolate-induced aseptic peritonitis. Role of VCAM-1 in hematopoietic progenitor retention at basal hematopoiesis and in mobilization
Studies of basal hematopoiesis in VCAM-1–ablated mice and their age-matched controls reveal certain notable differences. Among them, the most conspicuous is a modest increase in circulating WBCs, but a significant and sustained increase in circulating hematopoietic progenitors. Although an increased progenitor content is seen in the spleen, its enhancement following splenectomy suggests release from the BM with subsequent sequestration in the spleen. Given the extravascular location of hematopoietic cells in BM, how is the enhanced release of progenitor cells explained? Is it secondary to VCAM-1 absence from hematopoietic cells themselves or to VCAM-1 ablation from endothelial cells? We have already argued that a preferential release of VCAM-1– hematopoietic cells present in BM is an unlikely explanation for this phenomenon. Are VCAM-1– endothelial cells responsible? We believe that absence of VCAM-1 not only from endothelial cells, but especially from BM stromal cells, tested for the first time in the present studies, is likely responsible for this process, suggesting a key role for these cells in the retention of (
A corollary to the decrease of VCAM-1 in normal BM after G-CSF mobilization is the increase in sVCAM-1 in circulation.48 If VCAM-1 cleavage was critical for G-CSF mobilization, one would have expected that VCAM-1–ablated mice would respond less to such a treatment. Contrary to expectations, our mice responded as well as, or, at suboptimal stimulation, better than controls (Figure 6) by displaying enhanced levels of circulating progenitors. Thus, the proteolytic modulation of VCAM-1 in the bone marrow is not the driving force in G-CSF mobilization, a view consistent with results obtained in protease-deficient mice.49 Furthermore, our data suggest that absence of VCAM-1 may facilitate exit from the bone marrow in response to G-CSF–induced signals. This conclusion is consistent with a sustained hematopoietic progenitor egress from the bone marrow in these mice at baseline, similar to one seen in mice with conditional deletion of In contrast to BM showing decreased VCAM-1 levels, there was an increase in VCAM-1 expression by myeloid cells residing in the spleens of normal animals following G-CSF (Supplemental Figure S3). Therefore, proteolytic conditions for VCAM-1 cleavage were only present in BM, in agreement with prior data.48-50 However, since progenitor cells were also sequestered in the spleens of VCAM-1–ablated mice either before or after G-CSF, VCAM-1 in the spleen may not be responsible for their sequestration or entry into the spleen, in contrast to a previous speculation.48 Nevertheless, since mobilized normal mice accumulated more progenitor cells in their spleens compared with mobilized VCAM-1–ablated mice, one has to postulate that in the absence of VCAM-1, either there is a ceiling in splenic sequestration, or that the proliferative expansion of progenitors within the spleen is limited in the ablated animal. Consistent with this notion is the increased proportion of kit+ cells and increased cycling in the spleens of control animals. In mobilized mice, levels of soluble VCAM-1 in BM were reduced after mobilization, in contrast to PB. Presence of measurable levels of soluble VCAM-1 in VCAM-1–deficient animals (Table 2) provided confirmatory evidence that VCAM-1 is not completely ablated in the vascular beds of all tissues in the deficient mice, as was independently concluded by our RNA studies (Figure 7). Furthermore, SDF-1 levels in BM were reduced in all mice at the time of peak mobilization (Table 2). Whether these observed SDF-1 changes represent the main cause or just another facilitating pathway in mobilization is difficult to state for our or other murine models. In summary, we have uncovered new information about the biology of VCAM-1 expression in hematopoietic cells by demonstrating that it is constitutively expressed in myeloid cells confined to tissues. Although an association with their proliferative status was seen in vitro and in vivo, the functional significance of VCAM-1 expressed in hematopoietic cells has yet to be explored. Ablation of VCAM-1, in at least a subset of BM fibroblasts, suggests their developmental origin from a Tie2-expressing (mesenchymal?) progenitor, and further studies to pursue this issue are in progress. Increased egress of BM hematopoietic progenitors in the VCAM-1–ablated mouse validates the role of the VCAM-1/VLA-4 pathway in intramarrow retention of hematopoietic progenitors. Further, our postmobilization data provide further clarification about the role of VCAM-1 in G-CSF–induced mobilization.
We are grateful to Devra Batdorf for her assistance with mouse care, husbandry, and mouse splenectomy.
Submitted September 3, 2004; accepted March 9, 2005.
Prepublished online as Blood First Edition Paper, March 15, 2005; DOI 10.1182/blood-2004-09-3417.
Supported by National Institutes of Health grants DK46 557, HL58 734 (T.P.), and AI047 379 (P.A.K.).
T.U. and L.M.S. contributed equally to this work.
The online version of the article contains a data supplement.
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 357 710, Seattle, WA 98195-7710; e-mail: thalp{at}u.washington.edu.
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Abstract
Introduction
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