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Antisense inhibition of macrophage inflammatory protein 1-α blocks bone destruction in a model of myeloma bone disease
Sun Jin Choi,1 Yasuo Oba,1 Yair Gazitt,1 Melissa Alsina,2 Jose Cruz,3 Judith Anderson,4 and G. David Roodman1,4
1Department of Medicine/Hematology, University of Texas Health Science Center, San Antonio, Texas, USA2H. Lee Moffitt Cancer Center, Tampa, Florida, USA3Texas Tech University Health Science Center, Lubbock, Texas, USA4Audie Murphy Veterans Administration Medical Center, San Antonio, Texas, USA
Address correspondence to: G. David Roodman, Research (151), E-1152 BST, 200 Lothrop Street, Pittsburgh, Pennsylvania 15213, USA.Phone: (412) 648-9712; Fax: (412) 648-9723; E-mail: roodmangd@msx.upmc.edu.
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1Department of Medicine/Hematology, University of Texas Health Science Center, San Antonio, Texas, USA2H. Lee Moffitt Cancer Center, Tampa, Florida, USA3Texas Tech University Health Science Center, Lubbock, Texas, USA4Audie Murphy Veterans Administration Medical Center, San Antonio, Texas, USA
Address correspondence to: G. David Roodman, Research (151), E-1152 BST, 200 Lothrop Street, Pittsburgh, Pennsylvania 15213, USA.Phone: (412) 648-9712; Fax: (412) 648-9723; E-mail: roodmangd@msx.upmc.edu.
Find articles by Oba, Y. in: JCI | PubMed | Google Scholar
1Department of Medicine/Hematology, University of Texas Health Science Center, San Antonio, Texas, USA2H. Lee Moffitt Cancer Center, Tampa, Florida, USA3Texas Tech University Health Science Center, Lubbock, Texas, USA4Audie Murphy Veterans Administration Medical Center, San Antonio, Texas, USA
Address correspondence to: G. David Roodman, Research (151), E-1152 BST, 200 Lothrop Street, Pittsburgh, Pennsylvania 15213, USA.Phone: (412) 648-9712; Fax: (412) 648-9723; E-mail: roodmangd@msx.upmc.edu.
Find articles by Gazitt, Y. in: JCI | PubMed | Google Scholar
1Department of Medicine/Hematology, University of Texas Health Science Center, San Antonio, Texas, USA2H. Lee Moffitt Cancer Center, Tampa, Florida, USA3Texas Tech University Health Science Center, Lubbock, Texas, USA4Audie Murphy Veterans Administration Medical Center, San Antonio, Texas, USA
Address correspondence to: G. David Roodman, Research (151), E-1152 BST, 200 Lothrop Street, Pittsburgh, Pennsylvania 15213, USA.Phone: (412) 648-9712; Fax: (412) 648-9723; E-mail: roodmangd@msx.upmc.edu.
Find articles by Alsina, M. in: JCI | PubMed | Google Scholar
1Department of Medicine/Hematology, University of Texas Health Science Center, San Antonio, Texas, USA2H. Lee Moffitt Cancer Center, Tampa, Florida, USA3Texas Tech University Health Science Center, Lubbock, Texas, USA4Audie Murphy Veterans Administration Medical Center, San Antonio, Texas, USA
Address correspondence to: G. David Roodman, Research (151), E-1152 BST, 200 Lothrop Street, Pittsburgh, Pennsylvania 15213, USA.Phone: (412) 648-9712; Fax: (412) 648-9723; E-mail: roodmangd@msx.upmc.edu.
Find articles by Cruz, J. in: JCI | PubMed | Google Scholar
1Department of Medicine/Hematology, University of Texas Health Science Center, San Antonio, Texas, USA2H. Lee Moffitt Cancer Center, Tampa, Florida, USA3Texas Tech University Health Science Center, Lubbock, Texas, USA4Audie Murphy Veterans Administration Medical Center, San Antonio, Texas, USA
Address correspondence to: G. David Roodman, Research (151), E-1152 BST, 200 Lothrop Street, Pittsburgh, Pennsylvania 15213, USA.Phone: (412) 648-9712; Fax: (412) 648-9723; E-mail: roodmangd@msx.upmc.edu.
Find articles by Anderson, J. in: JCI | PubMed | Google Scholar
1Department of Medicine/Hematology, University of Texas Health Science Center, San Antonio, Texas, USA2H. Lee Moffitt Cancer Center, Tampa, Florida, USA3Texas Tech University Health Science Center, Lubbock, Texas, USA4Audie Murphy Veterans Administration Medical Center, San Antonio, Texas, USA
Address correspondence to: G. David Roodman, Research (151), E-1152 BST, 200 Lothrop Street, Pittsburgh, Pennsylvania 15213, USA.Phone: (412) 648-9712; Fax: (412) 648-9723; E-mail: roodmangd@msx.upmc.edu.
Find articles by Roodman, G. in: JCI | PubMed | Google Scholar
Published December 15, 2001 - More info
J Clin Invest. 2001;108(12):1833–1841. https://doi.org/10.1172/JCI13116.
© 2001 The American Society for Clinical Investigation
Received: April 24, 2001; Accepted: October 15, 2001
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Methods
Construction of ARH cell lines stably transfected with an antisense construct to human MIP-1α or empty vector. The MIP-1α antisense clone was constructed in the pcDNA3 vector. The first exon of the human MIP-1α cDNA (159 bp) (6) was generated by standard PCR techniques using MIP-1α–specific primers, 5′- CAG AAG GAC ACG GGC AGC AG-3′ (sense) and 5′-GTG ATG CAG AGA ACT GGT TG-3′ (antisense) (6). The PCR product was cloned into the TA cloning vector (Invitrogen Corp., Carlsbad, California, USA), followed by DNA sequence analysis. The first exon of the MIP-1α cDNA was then digested with the EcoRI restriction enzyme and inserted in the reverse orientation into the vector that was linearized with the EcoRI restriction enzyme (Figure 1). The orientation of the MIP-1α construct was determined using PCR techniques with the T7 and MIP-1α sense and antisense primers, followed by DNA sequencing. ARH cells (105) in exponential growth phase were plated in RPMI-1640 media containing 10% FBS (Life Technologies Inc., Rockville, Maryland, USA) into six-well plates for 24 hours and then were transfected with the MIP-1α antisense construct or the empty vector using Lipofectamine Plus (Life Technologies Inc.) in serum-free media, according to the manufacturer’s protocol. Twenty-four hours after transfection, the media was removed, and fresh media containing 10% FBS and 500 μg/ml of G418 was added. After 2 weeks of culture with G418, ARH cells were diluted into 96-well plates to isolate single clones, the clones expanded, and expression levels of MIP-1α were determined using a human MIP-1α (hMIP-1α) immunoassay kit (R&D Systems Inc., Minneapolis, Minnesota, USA). Transfection of ARH cells with pcDNA3–β–GAL reporter constructs and a β-galactosidase assay were used for determination of the transfection efficiency. The empty vector control ARH cell line was produced as described above, except that the cells were stably transfected with the empty pcDNA3 vector. We established two permanent cell lines from both antisense (AS) and empty vector (EV) transfected cells. Growth patterns for the parental ARH cells (WT-ARH) and the antisense MIP-1α (AS-ARH) or empty vector control cells (EV-ARH) were determined by culturing the cells (104 cells/well) for 7 days in RPMI-1640 media containing 10% FBS (Life Technologies Inc.) in the presence or absence of the ST2 murine marrow stromal cell line (104 cells/dish) on dentin slices in 48-well plates. The ST2 cells cultured on dentin were used as a model of the bone microenvironment. The number of cells at each time point was then counted. MIP-1α levels in the conditioned media at the end of 3 days of culture were determined in triplicate cultures using a commercial ELISA assay (R&D Systems Inc.). There was no significant difference in the growth characteristics of either the WT-ARH, AS-ARH, or EV-ARH cell lines. The AS-ARH cell line that produced the lowest levels of MIP-1α in the conditioned media was then used for in vivo studies.
Figure 1Construction of MIP-1α antisense. The first exon of MIP-1α cDNA was generated by standard PCR techniques as described in Methods. The identity of the MIP-1α cDNA was confirmed by sequence analysis, and it was subcloned into the pcDNA3 vector. Clones that had the reverse orientation for the MIP-1α cDNA were screened by PCR, and the orientation was confirmed by DNA sequence analysis.
Assay of adhesion properties and mRNA expression of human β1 integrins in WT-ARH, EV-ARH, or AS-ARH cells in vitro. The adhesion assay was performed according to the methods of Kim et al. (7) with minor modifications. ST2 cells (106) were plated in six-well plates in αMEM (Life Technologies Inc.) containing 10% FBS. After 24 hours, WT-, EV-, and AS-ARH cells (106) were added to the ST2 cells and the cells cocultured in RPMI-1640 media containing 10% FBS for 3 days. The culture plates were then washed extensively with 3 ml of serum-free RPMI-1640 five times. The remaining cells were fixed with acetone, stained with hematoxylin, and counterstained with eosin. Plasma cells (EV-ARH, AS-ARH, or WT-ARH) that were stained blue and attached to the ST2 cells were scored in ten random microscopic fields (×400).
Human β1 integrin mRNA expression levels in WT-, EV-, or AS-ARH cells were determined by RT-PCR analysis. PCR primers and PCR conditions for the human α3, α4, α5, α6, α7, and β1 integrins are shown in Table 1. Human-specific GAPDH was used as an internal control using human-specific GAPDH primers (208 bp), 5′-CTC TGA CTT CAA CAG CGA CA-3′ (sense) and 5′-TCT CTC TCT TCC TCT TGT GC-3′ (antisense). All PCR conditions were in the linear phase of the reaction. In selected experiments blocking Ab (20 mg/ml) to the α5 of the α5β1 integrin (P1D6; Life Technologies Inc.) was added to the cocultures.
Western blot analysis of the α5 integrin in WT-, EV-, and AS-ARH cells. The protein levels of human α5 integrin were tested by Western blot analysis. WT-, EV-, and AS-ARH cells (105) were suspended in 200 μl of SDS gel loading buffer and subjected to PAGE analysis. Gels were transferred to nitrocellulose membranes and the membranes blotted with an anti-CD49e (α5 integrin) mAb (1:200) (P1D6; Life Technologies Inc.), followed by anti-mouse IgG conjugated to peroxidase (Sigma Chemical Co., St. Louis, Missouri, USA) and visualized by chemiluminescence on x-ray films. Anti–β-actin mAb (Sigma Chemical Co.) was used as an internal control on the same membranes after stripping the anti-CD49e Ab with 1% SDS.
Development of myeloma bone disease and tumor burden in SCID mice transplanted with AS-ARH or EV-ARH cells in vivo. Three million WT-ARH cells, AS-ARH cells, or EV-ARH cells were injected intravenously into SCID mice that had received 3.75 Gy of total body irradiation 24 hours before transplantation. We have shown previously that this protocol is sufficient to induce myeloma bone disease in these mice within 28–35 days after transplantation (1). The animals were then observed for development of myeloma bone disease, hypercalcemia, and survival as described previously (1). Once the animals developed paraplegia, they were euthanized, and the vertebrae and long bones were subjected to histomorphometric analysis for OCL and osteoblast surfaces, area eroded, OCLs per unit area, and tumor volume, as described previously (8). At the end of the experiment, MIP-1α levels were measured in marrow plasma, and human IgG (hIgG) levels were measured in serum of animals by commercially available ELISA assays (MIP-1α; R&D Systems Inc.) (hIgG; AlerCHEK Inc., Portland, Maine, USA). In addition, marrow plasma samples from the animals were tested for their capacity to simulate OCL formation in human marrow cultures, as described below. In selected experiments, mice were transplanted with AS-ARH or EV-ARH cells as described above and sacrificed at 3, 6, 9, or 15 days after transplantation and evaluated for the presence of tumor cells by PCR using human-specific GAPDH primers (see above) and by histomorphometry. GAPDH primers that detected a consensus sequence in both murine and human GAPDH were used as an internal control for total GAPDH expression (sense 5′-ACC ACA GTC CAT GCC ATC AC-3′; antisense 5′-TCC ACC ACC CTG TTG CTG TA-3′).
OCL formation assays. Nonadherent human bone marrow mononuclear cells were obtained from normal donors as described previously (9) and tested for their capacity to form OCL-like multinucleated cells (MNCs) in long-term marrow cultures. These studies were approved by the Institutional Review Board of the University of Texas Health Science Center at San Antonio. The cultures were treated with either 200 pg/ml of recombinant hMIP-1α (R&D Systems Inc.) or 10% vol/vol marrow plasma from animals transplanted with WT-ARH cells, EV-ARH cells, or AS-ARH cells. Marrow plasma was obtained by flushing the femurs with 0.5 ml αMEM-10% FBS containing 100 U/ml preservative-free heparin, pelleting the cells by centrifugation for 10 minutes at 1,500 g, and collecting the supernatants. In selected experiments, the cultures were also treated with a neutralizing Ab to human MIP-1α (5 ng/ml) (R&D Systems Inc.), which is sufficient to neutralize up to 500 pg/ml of MIP-1α. At the end of the 3 weeks of culture, the number of MNCs that cross-reacted with the 23c6 mAb was determined. The 23c6 mAb identifies OCL-like cells that express calcitonin receptors and resorb bone (3).
Fluorescence-activated cell sorter analysis of ARH cells for expression of CD29 (β1 integrin), CD49e (α5 integrin), and CD18 (β2 integrin). Analyses were performed as outlined previously (10). Briefly, tubes containing 106 AS-ARH or EV-ARH cells were prepared and 20 μl of Ab were added to 100 μl of cell suspension (107 cells/ml). The cells were incubated for 30 minutes at 4°C. After washing to remove unbound Ab, the percentage of cells expressing the different integrins was determined by flow cytometry (FACScalibur; Becton Dickinson Immunocytometry Systems, Mountain View, California, USA). Twenty thousand events were collected. Analysis of results was done using the Cell-Quest Program (Becton Dickinson Immunocytometry Systems), and the percentage of positive cells obtained was corrected for background staining by subtracting the value obtained from the isotype Ab control. Anti-β1 integrin (4B4, CD29-FITC), anti-α5 (CD49e-FITC), and anti-β2 integrin (CD18-FITC) were from Immunotech (Immunotech, Marseilles, France). Ab’s and isotype-matched controls were from the same company.
Statistical analysis. In vitro and in vivo results are reported as the mean plus or minus SEM for five replicate samples and were compared by the Student t test. Results were considered significantly different for P values less than 0.05.
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Results
As shown in Figure 2a, there was no significant difference in the growth characteristics of WT-ARH, EV-ARH, or AS-ARH cells. Furthermore, coculture of the WT-ARH, AS-ARH, or EV-ARH cells with the ST2 cells, a murine marrow stromal cell line, on dentin slices demonstrated similar growth characteristics (Figure 2b). However, MIP-1α levels in media conditioned for 3 days by the cells were significantly different (Figure 2c). AS-ARH cells produced approximately 30–50 pg/ml of MIP-1α in the 3-day conditioned media, in contrast to 1,000–1,200 pg/ml of MIP-1α for the WT-ARH cell line or EV-ARH cells.
Figure 2Growth characteristics of WT-, EV-, and AS-ARH cells in vitro. (a) WT-, EV-, and AS-ARH cells (105) were cultured in six-well plates containing RPMI-1640 media containing 10% FBS. At days 3 and 5 of the culture, the cells were sampled, stained with trypan blue, and counted. (b) ST2 cells (104) in αMEM containing 10% FBS were plated onto dentin slices in 48-well plates. After 24 hours, WT-, EV-, and AS-ARH cells (104) in RPMI-1640 media containing 10% FBS were added to the culture. At days 4 and 7 of the culture, viable cells were scored as described above. Growth rates of WT-, EV-, and AS-ARH cells were not significantly different in the presence or absence of ST2 cells cocultured on dentin slices. (c) Conditioned media from WT-, EV-, or AS-ARH cells (105/ml) cultured in RPMI-1640 media containing 10% FBS were harvested at day 3, and the expression levels of MIP-1α were measured with a MIP-1α ELISA kit according to the manufacturer’s protocol. Similar results were seen in three independent experiments (*P < 0.0001).
We then transplanted SCID mice, which had been previously sublethally irradiated, with 3 × 106 AS-ARH, EV-ARH, or WT-ARH cells to determine their effects on development of myeloma bone disease and tumor burden in these animals. SCID mice transplanted with AS-ARH cells had a median survival that was longer compared with the mice injected with WT-ARH or EV-ARH cells (AS-ARH, 28 ± 4 days vs. WT-ARH, 20 ± 2 days, or EV-ARH, 23 ± 3 days). These results did not reach statistical significance (P = 0.06) because of the large variability in survival of these animals in three independent experiments. However, in all experiments, AS-ARH mice lived longer than EV-ARH or WT-ARH mice. Animals receiving WT- or EV-ARH cells all developed paraplegia before death due to vertebral involvement by myeloma. In contrast, animals receiving AS-ARH cells never developed paraplegia, but died of a wasting disease most likely reflecting extramedullary involvement by their myeloma. As shown in Figure 3a, MIP-1α levels were reduced to almost undetectable levels (< 10 pg/ml) in marrow plasma from the vertebrae or femurs of animals transplanted with AS-ARH cells. In contrast, the levels of human MIP-1α, although variable, were markedly elevated in marrow plasma from animals transplanted with either the WT-ARH or the EV-ARH cells. Furthermore, as shown in Figure 3b, the levels of human IgG, which are an indicator of tumor burden, were significantly decreased in the marrow plasma of animals implanted with AS-ARH cells (0.1–1.0 μg/ml) compared with EV-ARH cells or WT-ARH cells (80–120 μg/ml).
Figure 3Survival of SCID mice implanted with WT-, ET-, or AS-ARH cells and expression of MIP-1α in vivo. WT-, EV-, and AS-ARH cells were infused intravenously into SCID mice (n = 10 per group) as described in Methods and were sacrificed when they became paraplegic. Femurs and vertebrae were then removed and bone marrow plasma obtained by flushing the bones with 1 ml of serum-free αMEM. Expression levels of hMIP-1α (a) and human IgG (b) were measured with ELISA kits. hMIP-1α expression in mice implanted with AS-ARH cells was reduced to almost undetectable levels. Human IgG levels, which are indicators of tumor burden, were significantly reduced in AS-ARH mice compared with WT- or EV-ARH mice, but were still detectable (0.1–1 μg/ml). Similar results were seen in three independent experiments (*P < 0.0001).
We then determined if the marrow plasma from SCID mice implanted with the various ARH cell lines could stimulate OCL formation in human marrow cultures. As shown in Figure 4, marrow plasma from animals bearing the WT-ARH cells or animals implanted with EV-ARH cells stimulated OCL formation in human marrow cultures. The OCL stimulatory activity present in marrow plasma from these animals was totally inhibited by a neutralizing Ab to MIP-1α (Figure 4). In contrast, marrow plasma from SCID mice implanted with AS-ARH cells did not stimulate OCL-like cell formation in human marrow cultures.
Figure 4Effect of bone marrow plasma from SCID mice implanted with WT-, ET-, or AS-ARH cells on human OCL-like formation. Bone marrow plasma (10% vol/vol) from SCID mice infused with WT-, ET-, or AS-ARH cells that were obtained at the time they developed paraplegia, was added to human bone marrow cultures in the absence of exogenously added osteoclastogenic factors. rhMIP-1α (200 pg/ml) stimulated OCL-like MNC formation, which was blocked by the MIP-1α–neutralizing Ab (5 ng/ml). Bone marrow plasma from WT- or EV-ARH mice stimulated OCL-like MNC formation that was blocked by the MIP-1α neutralizing Ab. In contrast, bone marrow plasma from AS-ARH mice did not stimulate OCL-like MNC formation. Similar results were seen in two independent experiments (*P < 0.05; **P < 0.01).
As shown in Figure 5, animals infused with EV-ARH cells developed lytic bone lesions and increased OCL formation (upper right panel; shown by the red tartrate-resistant acid phosphatase [TRAP] stain). In contrast, animals infused with AS-ARH cells did not demonstrate increased OCL formation or bone resorption (lower right panel). In addition, tumor burden in the bones of animals treated with AS-ARH cells (upper left panel) was markedly decreased compared with animals infused with empty vector-transduced cells (lower left panel). AS-ARH cells could be detected histologically in the bone marrow sections from animals transfected with the antisense construct to MIP-1α, but they were rare. Histomorphometric analysis of the vertebral bodies from these animals demonstrated that OCL numbers per square millimeter of bone and per millimeter of bone surface area were significantly reduced in animals receiving AS-ARH cells compared with EV-ARH cells (Figure 6). The percentage of tumor per total bone area was also significantly decreased in animals treated with AS-ARH cells compared with animals treated with EV-ARH cells. There was no increase in extramedullary tumor as determined by metastases to liver and spleen (data not shown).
Figure 5Histology of bone sections from SCID mice implanted with EV- or AS-ARH cells. As shown in the panels stained with hematoxylin and eosin (H&E), mice implanted with AS-ARH cells had significantly reduced tumor burden compared with mice implanted with EV-ARH cells. OCL number was markedly reduced in SCID mice implanted with AS-ARH cells compared with the mice implanted with EV-ARH cells.
Figure 6Histomorphometric analysis of bone sections from SCID mice. Mice treated with AS-ARH cells had markedly reduced tumor burden compared with mice receiving EV-ARH cells. Furthermore, the number of OCLs per square millimeter of surface, the number of OCLs per square millimeter of bone area, and the percentage of active OCL surface were significantly reduced in AS-ARH mice compared with the EV-ARH mice (*P < 0.05).
To determine if the decreased tumor burden and bone destruction in SCID mice implanted with AS-ARH cells compared with EV-ARH cells was due to decreased homing and engraftment of the AS-ARH cells, mice were transplanted with the cells and sacrificed at 3-day intervals after transplantation. As shown in Figure 7, mRNA expression levels of human-specific GAPDH in mice transplanted with AS-ARH cells were decreased by day 3 compared with animals infused with EV-ARH cells, but were detectable. The level of human GAPDH decreased further by days 9 and 15.
Figure 7Expression levels of human-specific GAPDH mRNA from SCID mice implanted with EV- or AS-ARH cells at 3 to 15 days after cell infusion. Mice were infused with EV- or AS-ARH as described in Methods and sacrificed at 3-day intervals. Marrow samples were harvested and tested for human-specific GAPDH (hsGAPDH) mRNA expression by serial dilution RT-PCR analysis. The amount of hsGAPDH present in the bone marrow of SCID mice at days 3, 9, and 15 was significantly decreased compared with that of EV-ARH. The hsGAPDH was detectable up to 15 days in bone marrow of SCID mice infused with AS-ARH cells. In contrast, hsGAPDH levels in mice transplanted with EV-ARH cells were at least two logs higher compared with that in mice infused with AS-ARH cells. GAPDH primers that could detect both murine and human GAPDH mRNA were used as an internal control (tGAPDH).
Since MIP-1α enhances expression of adhesion molecules on cells (11, 12), we then assessed if the decreased tumor burden in AS-ARH cell–transplanted animals resulted from decreased adherence of AS-ARH cells to marrow stromal cells. As shown in Figure 8a, adherence of AS-ARH cells to ST2 marrow cells was decreased compared with that of EV-ARH cells. We then measured β1 integrin mRNA levels in EV-ARH and AS-ARH cells. The VLA-4 (α4β1) and VLA-5 (α5β1) integrins have been shown to mediate adherence of myeloma cells to marrow stromal cells (13, 14). As shown in Figure 8b, α4 mRNA expression levels were similar in AS-ARH cells and EV-ARH cells regardless of treatment with a neutralizing Ab to MIP-1α or rhMIP-1α. In contrast, the expression of α5 and β1 mRNA was decreased and was increased by addition of MIP-1α in AS-ARH cells. Furthermore, treatment of WT-ARH and EV-ARH cells with the anti–MIP-1α Ab decreased α5β1 expression. To confirm that α5 expression was decreased at the protein level, Western blot analysis was performed. As shown in Figure 9a, α5 expression levels were markedly decreased in AS-ARH cells compared with WT- or EV-ARH cells.
Figure 8Adhesion of WT-, EV-, or AS-ARH cells to ST2 mouse marrow stromal cells and expression levels of human β1 integrin mRNAs. (a) WT-, EV-, or AS-ARH cells (106) in RPMI-1640 media containing 10% FBS were cocultured with ST2 mouse stromal cells (106) in six-well plates. After 4 days, the plates were extensively washed with 3 ml of serum-free RPMI-1640 media five times to remove nonadherent ARH cells. The cells were fixed with acetone followed by H&E staining. Blue-stained plasma cells attached onto ST2 cells were scored using an inverted microscope by counting ten random fields (×400). The number of AS-ARH cells attached onto ST2 mouse stromal cells was significantly reduced compared with the EV- or WT-ARH cells. (b) Expression levels of the mRNAs for the β1 integrin were measured as described in Methods. Expression levels for the human α3, α4, α6, α7 integrins mRNAs were not affected by MIP-1α. In contrast, mRNA expression levels for the human α5 and β1 integrins were significantly decreased in AS-ARH cells and increased two- to fourfold in AS-ARH cells treated with MIP-1α. Treatment of WT- or EV-ARH cells with an anti–MIP-1α Ab decreased mRNA expression of the human α5 and β1 integrins by approximately 50% (*P < 0.05). Similar results were seen in two independent experiments.
Figure 9Expression of human integrin α5 in WT-, EV-, or AS-ARH cells and the effects of an anti-CD49e Ab (human integrin α5) on the adherence of ARH cells to ST2 mouse marrow stromal cells. (a) Western blot analysis for expression of the α5 integrin of VLA5 in WT-, EV-, and AS-ARH cells. Western blot analysis was performed as described in Methods. A greater than 70% reduction in α5 was seen in AS-ARH cells compared with WT- or EV-ARH cells. A similar pattern of results was seen in two independent experiments. (b) Effects of an Ab to the α5 integrin of VLA5 on adherence of ARH cells to ST2 marrow stromal cells. Adhesion assays were performed as described in Methods. Ab to α5 but not to αvβ3 blocked the adherence of WT- or EV-ARH cells to ST2 cells. A similar pattern of results was seen in two independent experiments. *P < 0.05.
Results from PCR studies suggested that transfection of ARH cells with antisense to MIP-1α resulted in downregulation of (α5β1) mRNA. We then analyzed surface expression of β2 integrin (CD18), β1 integrin (CD29), and α5 integrin (CD49e) by immunofluorescence staining and flow cytometry. The results are depicted in Figure 10. As can be seen in Figure 10, a marked decrease in the fluorescence intensity (peak mean channel, or PMC) of β1 integrin was observed in AS-ARH compared with EV-ARH (from 28–31, to 14 units of fluorescence, respectively) with a concomitant decrease in the percentage of positive cells from 58% to 34%. In contrast to β1 integrins, no decrease in PMC was observed in surface staining for β2 integrins between EV-ARH and AS-ARH. Similarly, a decrease in α5 was also observed in AS-ARH cells compared with EV-ARH cells from 58% positive cells to 22% positive cells, although the overall staining for α5 was much lower compared with β1. A concomitant decrease in PMC from 9.3 to 7.1 for α5 expression by AS-ARH cells was observed, consistent with the observed decrease in the percentage of positive cells.
Figure 10Immunofluorescence staining for CD18, CD29, and CD49e. Cells were stained and analyzed as described in Methods. Staining for EV-ARH cells was compared with AS-ARH cells for a given Ab. For each Ab, cells were also stained with isotype-matched control Ab to correct for background staining. Twenty thousand cells were analyzed for each sample. (a and c) EV-ARH cells. (b and d) AS-ARH cells.
These data suggested that decreased expression of the α5β1 integrin in AS-ARH cells was responsible for their decreased adherence to marrow stromal cells. To test this possibility, WT-, EV-, or AS-ARH cells were cocultured with ST-2 cells in the presence or absence of a blocking Ab to the α5 integrin, an isotype control Ab, or anti-αvβ3. As shown in Figure 9b, treatment of the cocultures with the anti-α5 Ab significantly decreased the binding of WT- or EV-ARH cells to marrow stromal cells. Anti-αvβ3 or the isotype control Ab did not affect the adherence of WT or EV-ARH cells to ST2 cells.
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