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 Previous Article | Table of Contents | Next Article 
Blood, Vol. 93 No. 9 (May 1), 1999:
pp. 3064-3073
Bone Marrow Neovascularization, Plasma Cell Angiogenic Potential, and
Matrix Metalloproteinase-2 Secretion Parallel Progression of Human
Multiple Myeloma
By
Angelo Vacca,
Domenico Ribatti,
Marco Presta,
Monica Minischetti,
Monica Iurlaro,
Roberto Ria,
Adriana Albini,
Federico Bussolino, and
Franco Dammacco
From the Department of Biomedical Sciences and Human Oncology, and
the Institute of Human Anatomy, Histology and Embryology, School of
Medicine, University of Bari, Bari; the Department of Biomedical
Sciences and Biotechnology, School of Medicine, University of Brescia,
Brescia; Advanced Biotechnology Center, National Institute for Cancer
Research, Genova; and the Institute for Cancer Research and Treatment
(IRCC), School of Medicine, University of Torino, Torino, Italy.
 |
ABSTRACT |
To assess whether the progression of plasma cell tumors is
accompanied by angiogenesis and secretion of matrix-degrading enzymes, bone marrow biopsy specimens from 20 patients with monoclonal gammopathy of undetermined significance (MGUS), 18 patients with nonactive multiple myeloma (MM), and 26 patients with active MM were
evaluated for their angiogenic potential and matrix-metalloproteinase (MMP) production. A fivefold increase of the factor VIII+
microvessel area was measured by a planimetric method of point counting
in the bone marrow of patients with active MM as compared with
nonactive MM and MGUS patients (P < .01). When serum-free conditioned media (CM) of plasma cells isolated from the bone marrow of
each patient were tested in vivo for their angiogenic activity in the
chick embryo chorioallantoic membrane (CAM) assay, the incidence
of angiogenic samples was significantly higher (P < .01) in the active MM group (76%) compared with nonactive
MM (33%) and MGUS (20%) groups. Moreover, a linear correlation
(P < .01) was found between the extent of vascularization of
the bone marrow of a given patient and the angiogenic activity exerted in the CAM assay by the plasma cells isolated from the same bone marrow. In vitro, a significantly higher fraction of the plasma cell CM
samples from the active MM group stimulated human umbilical vein
endothelial cell (HUVEC) proliferation (53%, P < .01),
migration (42%, P < .05), and/or monocyte chemotaxis (38%,
P < .05) when compared with nonactive MM and MGUS groups
(ranging between 5% and 15% of the samples). Also, immunoassay of
plasma cell extracts showed significantly higher (P < .01)
levels of the angiogenic basic fibroblast growth factor (FGF)-2 in the
active MM patients than in nonactive MM and MGUS patients (153 ± 59, 23 ± 17, and 31 ± 18 pg FGF-2/100 µg of protein, respectively).
Accordingly, neutralizing anti-FGF-2 antibody caused a significant
inhibition (ranging from 54% to 68%) of the biological activity
exerted on cultured endothelial cells and in the CAM assay by plasma
cell CM samples from active MM patients. Finally, in situ hybridization of bone marrow plasma cells and gelatin-zymography of their CM showed
that active MM patients express significantly higher (P < .01) levels of MMP-2 mRNA and protein when compared with nonactive MM
and MGUS patients, whereas MMP-9 expression was similar in all groups.
Taken together, these findings indicate that the progression of plasma
cell tumors is accompanied by an increase of bone marrow neovascularization. This is paralleled by an increased angiogenic and
invasive potential of bone marrow plasma cells, which is dependent, at
least in part, by FGF-2 and MMP-2 production. Induction of angiogenesis
and secretion of MMPs by plasma cells in active disease may play a role
in their medullary and extramedullary dissemination, raising the
hypothesis that angiostatic/anti-MMP agents may be used for therapy of MM.
© 1999 by The American Society of Hematology.
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INTRODUCTION |
ANGIOGENESIS is a required step in the
progression of tumor growth, invasion, and metastasis.1
Progression also involves secretion of the extracellular
matrix-degrading enzymes such as metalloproteinase-2 (MMP-2 or
72-kD type IV collagenase) and MMP-9 (92-kD type IV
collagenase) by tumor cells.2 In human solid tumors such as
colon, breast, lung carcinomas, and melanoma, angiogenesis and
MMP-2/MMP-9 overexpression occur simultaneously during invasion and
metastasis, but are downregulated or even absent in hyperplastic or
normal tissue and in situ tumors.3-5 In contrast, little is
known regarding angiogenesis in response to hematologic tumors.
Angiogenesis correlates with plasma cell growth (S-phase fraction) in
patients with monoclonal gammopathy of undetermined significance (MGUS)
and multiple myeloma (MM)6 and is associated with acute
lymphoblastic leukemia7 and high-grade B-cell
non-Hodgkin's lymphomas.8 MMP-2 is activated by and MMP-9
is secreted by plasma cells in MM, potentially contributing to tumor
invasion,9 whereas MMP-9 is secreted by some high-grade lymphomas, correlating with systemic spread and shorter
survival.10
In this study, the extent of bone marrow angiogenesis was investigated
in patients with MGUS, nonactive and active MM. The in vivo and in
vitro angiogenic potential and the MMP-2/MMP-9 expression of bone
marrow plasma cells isolated from the same patients were also assessed.
The results showed increased bone marrow neovascularization during
active MM paralleled by increased angiogenic potential and MMP-2
expression by the plasma cells isolated from active MM patients when
compared with MGUS and nonactive MM patients.
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MATERIALS AND METHODS |
Patients and control subjects.
A total of 65 patients who fulfilled the South West Oncology Group
(SWOG) diagnostic criteria for MM and MGUS11 were studied (Table 1). Myeloma patients were defined as
active or nonactive, according to clinical features and M-component
level.12 Active patients were those: (1) at diagnosis, with
symptomatic disease and an increase in M-component level in the 3 months before analysis; (2) at relapse; (3) with unresponsive and
rapidly progressive disease (leukemic progression), characterized by
severe bone pain, hypercalcemia, and pancytopenia. Several patients at
relapse or with leukemic progression displayed extramedullary
localizations. Nonactive patients were those in: (1) posttreatment
complete/objective response; (2) the off-treatment plateau phase.
Control subjects were 12 patients with anemia due to iron or vitamin
B12 deficiencies.
The study was approved by the local ethics committee and all patients
gave their informed consent.
Measurement of bone marrow angiogenesis.
Blood vessels were detected in 6-µm sections of 4%
paraformaldehyde-fixed paraffin-embedded biopsy specimens by staining
endothelial cells with the antifactor VIII murine monoclonal antibody
(MoAb) M616 (IgG1; Dako, Glostrup, Denmark) and a three-layer
biotin-avidin-peroxidase system described previously.13
Megakaryocytes, very few in number, also stained by factor VIII, but
were easily distinguishable by their morphology. Angiogenesis was
measured as microvessel area without knowledge of the clinical
diagnosis. Briefly, four to six 250× fields covering each of two
sections per biopsy were examined with a superimposed 484-point square
reticulum (12.5 × 10 2 mm2) for the
presence of microvessels (capillaries and small venules). These were
identified as endothelial cells, either single or clustered in nests or
tubes, and clearly separated from one another, and either without or
with a lumen (not exceeding 10 µm). A planimetric point count
method14 with slight modifications for the computed image
analysis (Leica Quantimet 500, Wetzlar, Germany) was applied to measure
the microvessel area within the cellular area (reticulum area minus
dense connective tissue, fat, bone lamellae, necrosis, and hemorrhage
areas).13 Values are expressed as mean ± 1 standard deviation (SD) per patient, subgroups, and groups of patients.
Cell cultures and preparation of conditioned medium (CM).
Aspirates close to the biopsy site were subjected to Ficoll-Hypaque
density gradient centrifugation and plasma cell enrichment. T cells
were removed by twofold E-rosetting, monocytes and fibroblasts by
adhesion determined by culturing in plastic flasks for 90 minutes in
RPMI 1640 containing 10% fetal calf serum (FCS) at 37°C in 5%
CO2 humidified atmosphere. Enriched plasma cells were then obtained by: (1) incubating residual cells with magnetic beads (Dynal,
Oslo, Norway) coated with antibody to the plasma cell marker CD38
(Becton Dickinson, Mountain View, CA) for 30 minutes at +4°C; (2)
magnetic subtraction; (3) bead detachment by culturing cells for 12 hours in RPMI-1640 supplemented with 10% FCS at 37°C in 5%
CO2.13 Enriched plasma cells contained <2%
of T cells and monocytes, as assessed by flow cytometry and with the
anti-CD3 and anti-CD68 antibodies, respectively (FACScan; Becton
Dickinson). They consisted of greater than 95% tumor plasma cells and
their clonally related cells,15 or plasma cells in control
subjects, as assessed by morphology in May-Grünwald-Giemsa and
flow cytometry with the anti-CD38 antibody, or by immunocytochemical
staining with anti- or anti- antibody (Dako) according to the
light chain of the M-component. Cells of each patient and control
subject were cultured (1 × 107 per 25-cm2
flask) in RPMI-1640 medium (6 mL/flask) supplemented with 1% glutamine
for 24 hours, and their viability assessed by trypan blue exclusion was
greater than 90%. The CM was collected, sequentially centrifuged at
1,200 and 12,000 rpm for 10 minutes, respectively, filtered through
sterilized 0.22-µm pore-size filters (Costar, Cambridge, MA), and
stored at  80°C.
Human umbilical vein endothelial cells (HUVEC) prepared as
described16 were grown in Petri dishes coated with 1%
gelatin (Sigma Chemical Co, St Louis, MO) in M199 medium supplemented with 20% FCS, 0.02% bovine brain extract, and 0.01% porcine heparin (both from Sigma Chemical Co). A Kaposi's sarcoma spindle cell line17 was cultured in Dulbecco's modified Eagle's medium
(DMEM) supplemented with 10% FCS and 1% glutamine. Cells
were harvested in trypsin/EDTA solution (0.05/0.02% in
phosphate-buffered saline [PBS]), washed twice with PBS, and cultured
(1 × 107 per 25-cm2 flask) in DMEM for 24 hours. Cell viability and collection of the CM were performed as
described above.
HUVEC proliferation assay.
A total of 2.5 × 103 HUVEC (second passage) was
plated in 96-well plates precoated with 1% gelatin. After 24 hours,
the medium was removed and replaced on days 0, 2, and 4 in
quadruplicate with complete fresh medium (positive control) or with
starvation medium (containing only 2.5% FCS) supplemented 1:1
(vol:vol) with RPMI-1640 medium (negative control) or with the plasma
cell CM. The cell number was estimated18 on day 6 by the
crystal violet colorimetric method of Kueng et al19 with
reading at 595 nm in the Microplate Reader Model 3550 (Bio-Rad
Laboratories): the cell number was derived from a calibration curve set
up with a known number of cells. Values were expressed as mean per sample.
HUVEC chemotaxis assay.
By using the Boyden chamber technique,20 200 µL of the
plasma cell CM, of Kaposi CM as the positive control,17 or
of 0.1% BSA in DMEM as the negative control (to evaluate random
migration) were placed in triplicate in the lower compartment of the
chamber (Costar), and 1.2 × 105 cells20
in 400 µL of DMEM 0.1% BSA were seeded in the upper compartment.
Compartments were separated by a 12-µm pore-size polycarbonate filter
(polyvinylpyrrolidone-free; Costar) precoated with 0.1% gelatin. After
6 hours of incubation at 37°C, cells on the upper side of the
filter were removed by scraping, whereas those that had migrated to the
lower side were fixed in absolute ethanol, stained with toluidine blue
(Merck, Darmstadt, Germany), and counted in 10 oil 400× immersion
fields. Values were expressed as mean per sample.
Monocyte chemotaxis assay.
Monocytes were isolated from heparinized peripheral blood of healthy
donors by centrifugation on Ficoll-Hypaque gradients. After
NH4Cl lysis of residual erythrocytes, they were further purified by centrifugation on Percoll gradients (density = 1062) and resuspended in RPMI-1640 medium containing 0.1%
BSA to a final concentration of 4 × 106 cells/mL.
Chemotaxis was performed in a 48-microwell chamber consisting of two
compartments (Costar) separated by a 5-µm pore-size polycarbonate
filter (polyvinylpyrrolidone-free; Costar). The lower compartment was
filled in sextuplicate with 27 µL of: (1) the plasma cell CM; (2) the
RPMI-1640 medium/0.1% BSA as the negative control; (3) this medium
supplemented with 10 nmol/L of the strong chemoattractant formylpeptide
as the positive control.21 The upper compartment was filled
with 2 × 105 cells in 50 µL. After 2 hours of
incubation at 37°C in 5% CO2 humidified atmosphere,
the cells on the lower surface of the filter were fixed, stained, and
counted in the same way as in the HUVEC chemotaxis assay.
Chick embryo chorioallantoic membrane (CAM)-gelatin sponge assay.
CAM of fertilized White Leghorn chicken eggs was used.22 On
day 3 of incubation, a square window was opened in the shell and 2 to 3 mL of albumin was removed to detach the developing CAM from the shell.
On day 8, 3 µL of plasma cell CM was loaded onto 1 mm3
gelatin sponges (Gelfoam; Upjohn Co, Kalamazoo, MI) that were then
implanted on top of the CAM. Sponges loaded with RPMI-1640 medium alone
or containing basic fibroblast growth factor (FGF-2) (200 ng/mL) were
used as the negative and positive controls, respectively. The sponge
trapped the sample, allowing a slow release of the products contained
in the medium. CAM were examined daily until day 12, when the
angiogenic response peaked,22 and photographed in ovo with
a Zeiss SR stereomicroscope and the MC63 camera system (Zeiss,
Oberkochen, Germany). To better highlight vessels, CAM were injected
into the large allantoic vein with an India ink solution, fixed in
Serra's fluid, dehydrated in graded ethanols, and made transparent
with methylbenzoate.23
Angiogenesis was measured in the sponges by a planimetric
method22 similar to that applied for bone marrow biopsy
specimens. Briefly, on day 12, the embryos and their membranes were
fixed in ovo in Bouin's fluid. Sponges and the underlying and
immediately adjacent CAM portions were removed, embedded in paraffin,
sectioned at 3 µm along a plane parallel to the CAM surface, and
stained with 0.5% toluidine blue. Four to six 250× fields
covering almost the whole of every third section within 30 serial
sections of at least two sponges per patient were analyzed, and the
microvessel area was calculated inside the reticulum test reference
area as the mean ± 1 SD per patient, subgroups, and groups of patients.
Quantification of plasma cell FGF-2 and vascular endothelial growth
factor (VEGF-A).
A total of 2 to 5 × 106 cells of each patient washed
twice in PBS were sonicated in 1 mL of ice-cold PBS with three
15-second 60-W bursts (Labsonic 2000 U, B-Braun, Germany), and
clarified by centrifugation at 12,000 rpm for 10 minutes at
4°C.24 For each sample, 100-µg proteins measured with
the Bradford method (Bio-Rad Laboratories, Richmond, CA) were tested in
duplicate for FGF-2 and VEGF-A concentrations by using the sandwich
enzyme-linked immunosorbent assay (ELISA) (Quantikine Human FGF-2,
Quantikine Human VEGF-A; R & D Systems, Minneapolis, MN) according to
the manufacturer's instructions.
MMP-2 and MMP-9 sodium dodecyl sulfate-polyacrylamide gel
electrophoresis (SDS-PAGE) zymography.
Gelatin-zymography of plasma cell CM was performed to visualize the
gelatinolytic activity of the secreted MMP-2 and MMP-9.25 A
total of 10 µg of proteins from each CM were applied in duplicate to
7.5% SDS-PAGE gels copolymerized with type A gelatin from porcine skin
(Sigma Chemical Co) at a final concentration of 0.6 mg/mL. After
electrophoresis, gels were washed in 2.5% Triton 1× for 1 hour
to remove SDS, incubated for 18 hours at 37°C, and stained in 0.1%
Coomassie brilliant blue. The gelatinolytic regions were observed as
white bands against a blue background. The levels of MMP activity were
assessed by scoring the intensity of the bands by a computerized image
analysis (APPLE, Cupertino, CA).
In situ hybridization of MMP-2 and MMP-9 mRNA.
This was performed as described previously.26
Cytocentrifuged plasma cells (1 to 2 × 105 per slide)
were fixed in 4% paraformaldehyde, washed in PBS, made permeable with
10 µg/mL of proteinase K (Sigma Chemical Co) in CaCl2 (2 mmol/L)-tris(hydroxymethyl)aminomethane (TRIS) (20 mmol/L) for 5 minutes at 37°C, acetylated with 0.25% acetate in 0.1 mol/L
triethanolamine, washed in 2X standard saline citrate (SSC), dehydrated in graded ethanols and air-dried. Cells
were hybridized overnight at 50°C with 5 µg/mL of two
5'-biotinylated oligonucleotides (Genenco Life Sciences,
Florence, Italy), the first of 42 bases complementary to the ninth exon
sequence 446-459 of the MMP-2 mRNA, the second of 48 bases
complementary to the ninth exon sequence 445-460 of the MMP-9
mRNA.27 Fifty percent deionized formamide, 600 mmol/L NaCl,
80 mmol/L TRIS, 4 mmol/L EDTA, 10 mmol/L dithiothreitol,
1X Denhardt's solution, and 100 µg/mL salmon sperm DNA were used as
hybridization medium. After washing in 2X to 0.01X SSC and in PBS,
cytospins were incubated overnight at 4°C with
streptavidin-alkaline phosphatase conjugate (Promega Co, Madison, WI).
After alkaline phosphatase activity was revealed by Western blue
stabilized substrate (Promega Co), cytospins were mounted in buffered
glycerin and evaluated by two observers with a double-headed Leitz
Dialux 20 photomicroscope (Leitz, Wetzlar, Germany). Plasma cells were
scored positive for the hybridization signal relative to the background
signal of RNAse-treated control cytospins hybridized with the same oligonucleotides.
| |
RESULTS |
Bone marrow microvessel area.
Biopsy sections stained with antifactor VIII antibody were examined
planimetrically to determine their microvessel area as normalized to
the total cellular area. Table 2 shows that
this area was much larger in patients with active MM than in those with
nonactive MM, MGUS, or control subjects (virtually identical). Figure 1 shows that microvessels in active
MM bone marrow were thin, winding, often without visible lumina, and
that single, spread endothelial cells and small endothelial sprouts
without lumen were evident, as opposed to straight vessels and no
sprouts in nonactive MM, MGUS, and control subject bone marrow.
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| Fig 1.
Staining with factor VIII of bone marrow from patients
with (A) MM at relapse, (B) MM at plateau, (C) MGUS, and (D) a control
subject (patient with pernicious anemia). Note in (A) numerous
microvessels, whereas in (B), a microvessel and some rare endothelial
cell clusters and in (C), (D) the lack of vessels in the presence of
strongly stained megakaryocytes. Bar, (A) to (C) 40 µm; (D) 55 µm.
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Angiogenic potential of bone marrow plasma cells.
The intense neovascularization in the bone marrow of active MM patients
was followed by assessment of the angiogenic potential of their bone
marrow plasma cells harvested in close proximity to the biopsy site.
The CM of plasma cell cultures was evaluated in vivo to determine its
ability to induce an angiogenic response in the chick embryo
CAM-gelatin sponge assay.22 As shown in Table 3, plasma cell CM of 20/26 (76%)
active MM patients induced a pronounced angiogenic response. This
presented as a dense growth with numerous capillaries converging like
spokes toward the sponge (Fig 2A and B). In
contrast, only 33% and 20% of CM from nonactive MM and MGUS patients,
respectively, induced the response and none from control subjects. Each
CM was assayed on two to three eggs and gave similar results.
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| Fig 2.
Angiogenic activity of plasma cell CM: chick embryo
CAM-sponge assay. (A) The positive control (RPMI-1640+FGF-2) and (B)
the CM of an active MM patient (progression) were loaded onto gelatin
sponges implanted on top of the CAM on day 8. Macroscopic appearance of
the CAM on day 12: note the presence of numerous blood vessels with a
"spoked wheel" pattern around both sponges, highlighted by India
ink injection. (C) Histologic section of the sponge sub (B), showing a
collagenous matrix pierced by winding blood vessels (arrowheads)
containing circulating cells and surrounded by a dense mononuclear cell
infiltrate. (D) Histologic section of a sponge loaded with negative
control medium (RPMI-1640) and devoid of vessels among the trabeculae.
Bar, 3 mm in (A) and (B); 90 µm in (C) and (D).
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Planimetric quantification of the average microvessel area also showed
the more intense neovascularization of the CAM loaded with CM of active
MM patients (Table 3). As observed in their bone marrow, the angiogenic
response elicited by their plasma cell CM consisted of thin, winding,
and branching microvessels, either without or with a lumen containing
circulating cells (Fig 2C). Interestingly, this angiogenic activity was
always significantly correlated with the corresponding bone marrow
microvessel area (Fig 3).
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| Fig 3.
Relationship between bone marrow neovascularization and
bone marrow plasma cell angiogenic activity. For each patient, bone
marrow microvessel area was measured and plotted against the
microvessel area measured in the CAM-sponge assay after loading of the
CM of his/her plasma cells isolated close to the biopsy site. Each
symbol (bold square, active MM; triangle, nonactive MM; circle, MGUS)
corresponds to one patient. Significance of the regression analysis was
calculated by the Pearson (r) test.
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Furthermore, the plasma cell CM of active MM patients more strongly
induced in vitro cell functions correlated with angiogenesis. CM of
14/26 (53%) active MM patients stimulated enhanced HUVEC proliferation
(Fig 4A), as compared with CM
of 2/18 (11%) nonactive MM and 3/20 (15%) MGUS patients (P < .01,  2 Cochrane test), and a higher percentage
(P < .05) of CM from active MM patients induced
enhanced chemotaxis in both HUVEC (Fig 4B) and human monocytes (Fig
4C). Plasma cell CM of control subjects overlapped the negative
controls in all of the assays.
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| Fig 4.
Effect of the CM of bone marrow plasma cells on
(A) HUVEC proliferation, (B) HUVEC chemotaxis, and (C) human monocyte
chemotaxis. In (A), low-density cultures of HUVEC (2.5 × 103 cells per well) were exposed on days 0, 2, and 4 with
complete medium (positive control), starvation medium (negative
control), and negative control medium supplemented 1:1 (vol/vol) with
plasma cell CM. HUVEC were counted on day 6. Each dot represents the
mean of four determinations for each CM or control. In (B), Kaposi cell
CM (positive control), DMEM.1% BSA (negative control), and plasma
cell CM were added to the lower compartment, and 1.2 × 105 HUVEC were placed in the upper compartment. Cells that
migrated to the lower surface of a gelatin-coated filter separating the
compartments were counted after 6 hours. Each dot is the mean of three
determinations for each CM or control. In (C), a formylpeptide solution
(positive control), RPMI-1640.1% BSA (negative control), and plasma
cell CM were added to the lower compartment, and 2 × 105
human monocytes were placed in the upper compartment. Cells that
migrated to the lower surface of a polycarbonate filter were counted
after 2 hours. Each dot is the mean of six determinations for each CM
or control. In all assays, reproducibility was  ±10% of the mean
value of each CM. For all assays, the cutoff corresponds to the mean
plus 3 SD of the negative control medium. The number of CM tested for
each group of patients is given in brackets. The mean ± 1 SD is given
for each group.
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Seven samples from active MM patients induced all three responses
compared with one from nonactive MM and none from MGUS patients. In
addition, a small percentage of CM that induced neovascularization in
the CAM assay failed to induce chemotaxis in HUVEC and/or human monocytes.
Plasma cell FGF-2 and FGF-2 antagonism.
FGF-2 is a potent angiogenic factor28 and stimulator of
monocyte chemotaxis.29 Evaluation of its levels in plasma
cell lysates by immunoassay showed that they were significantly higher (P < .01, Student's t-test) in the plasma cells of
active MM patients (153 ± 59 pg FGF-2/100 µg protein) compared
with nonactive MM and MGUS patients (23 ± 17 and 31 ± 18 pg
FGF-2/100 µg protein, respectively).
An assessment was therefore made of the effect of a neutralizing
polyclonal anti-FGF-2 antibody (kindly provided by Dr D.B. Rifkin, New
York University, New York, NY) on the CM samples from seven active MM
patients that induced both endothelial cell and monocyte functions in
vitro and angiogenesis in vivo in the CAM. The antibody (at 400 µg/mL) partly inhibited CM-induced HUVEC proliferation (from 22 ± 5 × 103 to 12 ± 7 × 103
cells/dish, 46%), HUVEC chemotaxis (from 146 ± 33 to 65 ± 18 cells/filter, 56%), monocyte chemotaxis (from 1,473 ± 321 to 486 ± 175 cells/filter, 68%), and CAM
neovascularization (from 2.72 ± 0.59 × 102
to 1.27 ± 0.38 × 102 mm2 of
microvessel area, 54%). Its inhibition of CM-induced CAM neovascularization is illustrated in Fig 5
(see page 3067).
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| Fig 5.
Macroscopic appearance of a CAM (day 12) implanted
simultaneously at day 8 with a sponge loaded with the plasma cell CM of
an active (progression) MM patient alone (*) and with a second sponge
loaded with the same CM added with an anti-FGF-2 antibody (**). Note
the angiogenesis toward the one-asterisk sponge (some neovessels are
arrowheaded), and its inhibition by the anti-FGF-2 antibody. Bar, 2 mm.
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FGF-2 may thus play a role in plasma cell-mediated angiogenesis. When
all patients were considered, however, there was no significant
correlation between their individual plasma cell FGF-2 levels and bone
marrow neovascularization, suggesting that other angiogenic factors may
be involved. The angiogenic activity of VEGF-A can be excluded, as its
levels were low in plasma cells from active MM patients (12 ± 9 pg
VEGF-A/ 100 µg proteins), nonactive MM patients (19 ± 10 pg), and
MGUS patients (21 ± 11 pg).
MMP-2 and MMP-9 production by bone marrow plasma cells.
The invasive potential of bone marrow plasma cells during MM
progression was assessed by SDS-PAGE gelatin-zymography of plasma cell
CM. Bone marrow plasma cells from patients secrete activated (62-kD
form) MMP-2 and lower levels of activated (88-kD form) MMP-9, whereas
those from control subjects secrete only marginal levels of activated
MMP-2 (Fig 6A). Accordingly, bone marrow
plasma cells from patients expressed MMP-2 and MMP-9 mRNA, whereas in plasma cells from control subjects, the MMP-9 mRNA appeared to be
absent and the MMP-2 mRNA to be either poorly expressed or absent, as
evaluated by in situ hybridization (Fig 7).
Quantification of secreted MMP-2 and MMP-9 was performed by
computerized image analysis of the gelatinolytic bands, which showed
that MMP-2 was present in plasma cell CM of 22/26 (84%) active MM
patients at 22.5 ± 7.4 × 103 optical density
(OD), compared with 5/18 (27%) nonactive MM patients at 12.4 ± 3.6 × 103 OD, 5/20 (25%) MGUS patients at 13.4 ± 5.4 × 103 OD, and 2/12 (16%) control subjects at 4.0 ± 1.7 × 103 OD (P < .01, 2 Cochrane test). MMP-9 was cosecreted in 6 (23%)
active MM, 3 (16%) nonactive MM, and 2 (10%) MGUS patients. Its
levels, usually lower than MMP-2, overlapped between active MM (10.5 ± 2.7 × 103 OD), nonactive MM (10.0 ± 2.8 × 103 OD), and MGUS (9.0 ± 1.4 × 103 OD). The MMP-2 and MMP-9 quantification in
representative patients and a control subject is shown in Fig 6B.
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| Fig 6.
MMP-2 and MMP-9 secretion by bone marrow plasma cells.
The patient with nonactive MM was in the plateau phase; the patient
with active MM was at relapse; the control was a patient affected with
anemia due to iron deficiency. (A) SDS-PAGE gelatin zymography of
plasma cell CM samples. Note the white bands against a dark background
with an apparent molecular weight of 62 kD and 88 kD, corresponding to
the gelatinolytic regions of activated (cleaved) MMP-2 and MMP-9,
respectively. Measurement of the intensity of the bands, as evaluated
by computerized image analysis, is shown in (B). The assay was
performed in duplicate for each CM. Reproducibility was ±20% of
the mean intensity value. The lowest (third) band in the lane of the
active MM patient represents the 54-kD cleaved form of
MMP-2,2 which is sometimes present in the CM of both
nonactive and active MM patients, but is not correlated with disease
activity.
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| Fig 7.
Enriched bone marrow plasma cells isolated from patients
with (A) active (relapsing) MM, (D) nonactive (plateau) MM, and (G)
MGUS stained with May-Grünwald-Giemsa. Expression of (B) MMP-2
and (C) MMP-9 mRNA by the plasma cells of the active MM patient; (E)
MMP-2 and (F) MMP-9 mRNA by those of the nonactive MM patient; (H)
MMP-2 mRNA by those of the MGUS patient; (I) MMP-2 mRNA by a control
subject (a patient with pernicious anemia). Note the weaker MMP-2
expression in the nonactive MM patient, a weak expression in the MGUS
patient, and no expression in the control. Bar, 22 µm in (A), (D),
(G); 16 µm in (B), (C), (E), (F), (H), and (I).
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| |
DISCUSSION |
This study showed a significant increase of bone marrow angiogenesis
(evaluated as microvessel area) in patients with active MM compared
with nonactive MM and MGUS patients. Their bone marrow plasma cells
also displayed a stronger angiogenic potential, as assessed by the
ability of their CM to stimulate angiogenesis in the chick embryo CAM,
cell proliferation of HUVEC, and HUVEC, and human monocyte chemotaxis.
They also produced higher levels of angiogenic FGF-2 and of MMP-2.
These properties of active MM plasma cells were associated with MM
activity rather than with the tumor cell mass or burden, as very
similar numbers of plasma cells were assayed for each patient of each group.
As the progression from in situ to invasive and metastatic solid tumors
is accompanied and enhanced by the switch from the prevascular to the
vascular phase,30 our findings suggest that active MM may
represent the "vascular phase" of plasma cell tumors, and
nonactive MM and MGUS the "prevascular phase." Bone marrow angiogenesis may therefore be involved in progression from MGUS or
nonactive MM to active MM.
The angiogenic activity exerted by the CM of plasma cells isolated from
a given patient correlated significantly with the corresponding bone
marrow microvessel area in all patients. Plasma cells may thus be a
major source of angiogenic stimuli in the bone marrow microenvironment,
although macrophages, T cells, and mast cells may also play a role.
Plasma cell CM, in fact, stimulates endothelial cell proliferation and
motility, functions required for vascular sprouting31 and
monocyte chemotaxis, with the consequent recruitment of activated cells
able to secrete a variety of angiogenic factors.32
It must be pointed out, however, that significant variability in the
ability of plasma cell CM to exert a biological response on endothelium
and/or monocytes was observed in the same experimental group. For
instance, 76% of the plasma cell CM samples obtained from the 26 active MM patients induced an angiogenic response in the CAM assay,
while 53%, 42%, and 38% induced HUVEC proliferation, HUVEC
chemotaxis, or human monocyte chemotaxis. Thus, several CM samples were
unable to exert a response in all of the assays, a characteristic
shared by 7 of the 26 active MM samples tested. This may reflect
differences in the sensitivity of our biological assays and/or the
presence of different angiogenesis factors in the plasma cell CM of
different patients.
Although none of the parameters tested may represent an independent
prognostic factor, our data clearly indicate a significant increase of
bone marrow plasma cell angiogenic potential in the cohort of active MM
patients when compared with nonactive MM patients and MGUS patients.
This is reflected by the increased neovascularization of the bone
marrow in MM progression. Interestingly (data not shown), the plasma
cell labeling index (LI), a progression marker12 studied in
45 patients, was found to be highly correlated with their bone marrow
microvessel area (Pearson's [r] = .83, P < .001). Indeed, an LI (%) of 3.43 ± 1.86 was associated with an area of 0.42 ± 0.21 mm2 × 102 in 19 patients with active MM, whereas LI (%)/area (mm2 × 102) of 0.57 ± 0.38/0.09 ± 0.03, and 0.36 ± 0.25/0.11 ± 0.02 matched in 12 nonactive MM and 14 MGUS
patients, respectively. Conceivably, other progression markers, such as
interleukin-6 (IL-6) and IL-1 ,33 which also act as
angiogenic factors,30-32 might be correlated with the bone
marrow microvessel area.
Plasma cells isolated from the bone marrow of active MM patients
produce higher levels of FGF-2, suggesting that this angiogenic factor
plays a role in bone marrow neovascularization during MM progression.
Interestingly, we have found that neutralizing anti-FGF-2 antibody
causes a partial, but significant, decrease in the angiogenic activity
exerted by the plasma cell CM in both the chick embryo CAM in vivo
assay and in vitro assays. No significant correlation was observed,
however, between FGF-2 levels in plasma cell extracts and bone marrow
microvessel area, or between FGF-2 levels and the angiogenic potential
exerted by plasma cell CM samples in vitro and/or in vivo. Taken
together, these data suggest that other factors secreted by plasma
cells able to induce angiogenesis directly and/or indirectly (for
instance, via monocyte recruitment and activation), namely
VEGF-A,34 tumor necrosis factor-
(TNF-),35 macrophage-colony stimulating
factor,36 IL-1,37 and transforming growth
factor- (TGF-),38 may act together with FGF-2.
Further studies are required to characterize these factors. Preliminary observations have shown that production of VEGF-A by plasma cells does
not increase significantly during MM progression (see above), although
its ability to act in synergy with FGF-239,40 suggests that
even if its levels remain fairly constant, increased FGF-2 production
may result in a potent stimulation of bone marrow angiogenesis in
active MM patients. The support of the human bone marrow
microenvironment for the growth of both myeloma plasma cells and new
vessels into the tumor mass has recently been shown.41
As observed for tumor cells from invasive and metastatic solid
tumors,2 the bone marrow plasma cells of active MM patients express and secrete high levels of MMP-2. In some patients, sizable levels of MMP-9 were coexpressed and cosecreted. Both enzymes were
present in the plasma cell CM in their cleaved, activated form,42 suggesting that they are rapidly cleaved after
secretion, possibly by membrane-type (MT)-MMP,43 with which
plasma cells are likely equipped. Yet, additional activators such as
plasmin, kallicrein, trypsinlike serine proteinase, cathepsin G, and
-chymotrypsin,2 possibly carried by fetal calf serum,
are unlikely to be involved because the CM contained no serum. Previous
results showed a prominent production of activated MMP-9 in
MM.9 Differences in the sensitivity of the detecting system
may account for this discrepancy. Nevertheless, given the ability of
both activated MMP-2 and MMP-9 to degrade types IV, V, VII, and X
collagens, as well as fibronectin,2 the data suggest that
plasma cells of active MM patients can invade interstitial stroma and
the subendothelial basement membrane. Additional proof of the invasive
capability will be the positive balance between one or both MMPs and
their antagonists, or tissue inhibitors of MMPs. This is under ongoing study.
The increased bone marrow neovascularization, increased angiogenic and
proteolytic potential of plasma cells, together with their poor
adhesiveness to fibronectin (due to the lower expression of VLA-4
integrin),44 and their enhanced ability to adhere to the
vascular endothelium and thereby extravasate (due to the high expression levels of LFA-1 and CD44)13 may explain the
frequent occurrence of extramedullary localizations in active MM, eg,
peripheral blood, skin, liver, lung, and kidney,45 as in
five of our patients. Thus, combination therapy with
angiostatic/protease inhibitor molecules plus cytolytic drugs
previously shown to be effective in animal solid tumor
models46 could perhaps be considered for clinical
application in active MM.
| |
FOOTNOTES |
Submitted March 12, 1998; accepted December 29, 1998.
Supported in part by the Associazione Italiana per la Ricerca sul
Cancro-A.I.R.C., Milan (Project Diagnosis and Prognosis in Clinical
Oncology to F.D. and Special Project Angiogenesis to M.P.), and
Ministry of Education-M.U.R.S.T., (Grants 40% 1997 to
A.V. and M.P.), Rome, Italy. M.M. is the recipient of a fellowship from
the Fondazione Italiana per la Ricerca sul Cancro (F.I.R.C.), Milan, Italy.
The publication costs of this
article were defrayed in part by
page charge payment. This article
must therefore be hereby marked
"advertisement"
in accordance with 18 U.S.C. section
1734 solely to indicate this fact.
Address reprint requests to Franco Dammacco, MD, Department of
Biomedical Sciences and Human Oncology, Section of Internal Medicine
and Clinical Oncology, Policlinico, Piazza G. Cesare, 11, I-70124 Bari,
Italy; e-mail: dimoclin{at}cimedoc.uniba.it.
| |
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TOP
ABSTRACT
INTRODUCTION