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Experimental Therapeutics |
Departments of Surgery [R. M. S., N. T. J., V. K. G., M. C. P.] and Radiation and Cellular Oncology [H. J. M., S. S., M. A. B., A. M. K., D. M. H., S. H., R. R. W.], University of Chicago, Chicago, Illinois 60637; IlEX Oncology Inc., Boston, Massachusetts 02215 [C. R., R. K.]; and Dana Farber Cancer Institute, Harvard Medical School, Boston, Massachusetts 02115 [D. W. K.]
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ABSTRACT |
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 Top ABSTRACT INTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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INTRODUCTION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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0.01% of endothelial cells
undergo cell division. By contrast, the fraction of proliferating
endothelial cells in tumors is proposed to be 50-fold higher (1
, 2)
. These differences between proliferating endothelial cells in
tumor tissue versus normal tissue can be exploited through
the use of angiogenesis inhibitors. Furthermore, a single tumor vessel
may supply as many as 104
tumor cells, thereby
amplifying the antitumor effects of antiangiogenic compounds (1
, 2)
. Importantly, tumor endothelium is derived from normal host
cells and, in contrast to tumor cells, is genetically stable, which
suggests that tumor endothelium is unlikely to develop resistance to
cytotoxic agents (3)
. Recent investigations have focused
on the development of antiangiogenic agents, particularly those that
can be administered p.o. and for prolonged periods, as alternatives to
standard cytotoxic anticancer therapies. Investigations of antiangiogenic compounds have been conducted in preclinical and clinical trials. Strategies to inhibit angiogenesis have included the use of neutralizing antibodies to angiogenic proteins, integrin molecules, and growth factor receptors. (4, 5, 6) Also, kinase inhibitors, natural products such as TNP-470, and antibiotic-derivatives such as minocycline have been used (7 , 8) . Angiostatin and endostatin, enzymatic degradation products of plasminogen and type XVIII collagen, respectively, are reported to induce tumor regression and prolong tumor dormancy in murine model systems (9 , 10) . However, in most reports, angiostatin and endostatin are tumoristatic, not tumoricidal, and tumor regrowth ensues after cessation of treatment. Tumor cures have been limited when most angiogenesis inhibitors are used as a sole method of treatment.
Radiotherapy is an important standard cancer treatment for local and regional disease. Sixty percent of all cancer patients receive radiotherapy during the course of their disease. The importance of improving radiotherapy is further highlighted by the fact that approximately 20–30% of cancer deaths are attributable to local/regional failures and 50–60% of cancer patients die with local tumor as a component of treatment failure (11) . Recent advances in radiotherapy are attributed to improved methods of physical delivery, which allow for dose escalation and combination of radiotherapy with commonly used antineoplastic agents (12 , 13) . Combining radiotherapy and cytotoxic agents has improved local control and cure rates in a variety of human cancers but at the cost of significant acute normal tissue toxicity (14, 15, 16, 17, 18, 19) . Nonetheless, these results verify that local control can translate into an increase in cures.
Mauceri et al. (20) have recently demonstrated that combined treatment with angiostatin enhances the effects of IR4 on tumor regression. The combination of endostatin and IR has resulted in similar findings (Hanna, et. al., in press). Moreover, Gorski et al. (21) reported that the use of vascular endothelial growth factor antibody and IR produces greater than additive antitumor effects when compared with either treatment alone. These findings collectively indicate that the combination of radiotherapy and angiogenic blockade enhances the therapeutic ratio of IR by targeting both tumor cells and tumor vessels. Importantly, the therapeutic gains of the combined treatment modalities are achieved without increased toxicity when compared with treatment with radiotherapy alone.
Recently, an isocoumarin derivative, 2-(8-hydroxy-6-methoxy-1-oxo-1H-2-benzopyran-3-yl) propionic acid (NM-3), was synthesized from cytogenin, a compound isolated from the culture filtrate of Streptoverticillum eurocidium (22 , 23) . Antiangiogenic effects after p.o. administration of NM-3 have been demonstrated in the mouse dorsal air sac assay system. Although modest antitumor effects were reported when NM-3 was used as the sole therapeutic agent (22) , recent investigations suggest that antiangiogenic compounds effectively enhance the antitumor effects of IR (20 , 21) . The low-toxicity profile of NM-3, its oral bioavailability, and the ease of production, suggest that NM-3 might be a clinically useful antiangiogenic agent for combination studies with IR.
Here, we report that when NM-3 is combined with radiotherapy, an increase in local tumor control is observed without an increase in normal tissue toxicity. We also show that NM-3 is selectively cytotoxic to endothelial cells but not to tumor cells. We propose a new paradigm for enhancement of radiocurability whereby standard risk/benefit analysis, in terms of calculating the therapeutic index, is not applicable because the radio-enhancing agent has little or no antitumor effect when administered alone but enhances the antitumor effect of IR without increasing normal tissue effects.
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MATERIALS AND METHODS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Preparation and Administration of NM-3.
NM-3 was obtained from T. Nakashima (Central Research Lab.,
Mercian Corp., Fujisawa, Japan) and ILEX Oncology. For in
vitro experiments, NM-3 was prepared at a stock concentration of 1
mg/ml in PBS and filter sterilized. Serial dilutions were prepared in
sterile PBS as needed.
For in vivo studies, 1.25 mg/ml NM-3 was dissolved in PBS and filter sterilized. Each animal received 400 µl (equivalent to 25 mg/kg) by i.p. injection. Control mice received an equal volume of PBS. In one study, mice bearing Seg-1 xenografts were treated with 100 mg/kg/day NM-3 (suspended in 40% polyethylene glycol-400 in PBS).
Matrigel Assay.
The human glioblastoma cell line (U373 MG) was incubated overnight with
the hypoxia-mimetic compound cobalt chloride (250 µM;
Refs. 24, 25, 26
), harvested, and mixed with ice-cold Matrigel
(Collaborative Biomedical Products) at a concentration of 2 x 106 cells/ml. Athymic nude NCRNU female
mice (Taconic Farms) were anesthetized and injected s.c. on the midline
back with 0.25 ml of Matrigel/U373 MG. Soon after injection, the
Matrigel implant solidified and persisted without apparent
deterioration throughout the 4-day assay interval. Four h
postinjection, animals (10 per group) were given injections s.c. with
NM-3 (1–100 mg/kg/day) or PBS. Four days later, animals were
sacrificed, and Matrigel implants were dissected and photographed as
whole mounts using a stereomicroscope, allowing the gross observation
of neovascularization.
Clonogenic Assay.
Clonogenic assays were conducted as described previously
(20)
. Briefly, HUVECs and HAECs were plated in EGM-2
media. LLC and Seg-1 cells were plated in their respective growth
media. Eighteen h after plating, NM-3 was added (10–10,000 ng/ml) for
4 h. Cells were then rinsed and returned to their respective
growth media. After 7–14 days, cells were stained with crystal violet.
Colonies greater than 50 cells were scored as positive and the
surviving fraction was determined. In an additional experiment, HUVECs,
LLC cells, and Seg-1 cells were plated and treated with 1,000 ng/ml of
NM-3 for 4 h. Cells were irradiated with doses of 0–900 cGy using
a General Electric Maxitron X-ray generator operating at 250 kV,
26 mA, with a 0.5 mm copper filter, at a dose rate of 118 cGy/min.
Cells were rinsed and incubated in complete media for 7–14 days. Cells
were stained with crystal violet, and the surviving fraction was
determined by counting colonies.
Endothelial Cell Migration Assay.
Migration assays were conducted as described previously
(27)
. Briefly, HUVECs were cultured in EGM-2. To assess
migration, cells were starved overnight in minimal growth medium
containing 0.1% BSA. Cells in the NM-3 treatment groups were exposed
to 100 ng/ml of NM-3 for 4 h. The IR-treated and the
NM-3/IR-treated cultures were then irradiated with 900 cGy. The cells
were harvested, suspended in minimal growth medium with 0.1% BSA
(containing 100 ng/ml NM-3 in the NM-3 treated wells), and plated at
7 x 105 cells per well on the
lower surface of a gelatinized 5.0-µM filter (Nucleopore
Corp., Pleasanton, CA) in an inverted modified Boyden chamber. After
incubation for 1–2 h at 37°C, during which time the cells adhere to
the filter, the chamber was reinverted, medium containing 20 ng/ml
vascular endothelial growth factor was added to the top well, and the
chamber was incubated for 16 h at 37°C. The chambers were then
disassembled, and the filters fixed and stained with Diff-Quick Stain
Set (Dade International, Inc., Miami, FL). Migration was scored as the
total number of cells that migrated to the top of the membrane as
counted in 10 hpfs.
Animal Studies.
LLC cells (1 x 106 cells in 100
µl of PBS) were injected into the right hind limb of C57BL/6 female
mice (Frederick Cancer Research Institute, Frederick, MD). Seg-1 cells
(3 x 106 cells in 100 µl of
serum-free medium) and SQ-20B cells (5 x 106 cells in 100 µl of PBS) were injected into
the right hind limb of athymic nude female mice (Frederick Cancer
Research Institute, Frederick, MD). LLC, Seg-1, and SQ-20B tumors were
grown to a volume of 297.3 ± 13.3
mm3
(n = 43),
564.4 ± 22.2 mm3
(n = 31), or 681.8 ± 17.2
mm3
(n = 45),
respectively, before the mice were divided into experimental groups.
Tumor volume was determined by direct measurement with calipers as
described previously (28)
.
Mice bearing LLC tumors were given injections with 25 mg/kg/day of NM-3 as a single i.p. injection 4 h prior to IR treatment. These mice received a total dose of 100 mg/kg NM-3, with treatment beginning two days prior to IR treatment. Seg-1 tumor-bearing animals were treated with 100 mg/kg NM-3 i.p. for 4 consecutive days. In Seg-1 tumor experiments, both NM-3 and IR were started on the same day (day 0), with NM-3 given 3–4 h prior to IR. Mice bearing SQ-20B tumors received 50 mg/kg/day NM-3 (in two doses) for 8 days. In an additional experiment to assess local tumor control, C57BL/6 mice bearing LLC tumors (mean initial volume, 151.1 ± 4.6 mm3 ; n = 29) received 5 days of NM-3 at 50 mg/kg/day (split into two doses). In all of the experiments, control and IR animals were given i.p. injections with an equivalent volume PBS. Tumors were irradiated using a General Electric Maxitron X-ray generator operating at 150 kV, 30 mA, using a 1 mm aluminum filter at a dose rate of 188 cGy/min. Fractionation schemes were based on tumor-doubling times and tumor volume. LLC tumors were treated with a total dose of 40 Gy (20 Gy on 2 consecutive days); Seg-1 tumors were treated with 20 Gy (four 5-Gy fractions); and SQ-20B tumors were treated with 65 Gy (5 Gy for 2 days, 15 Gy for 2 days with 3 days between, a single dose of 10 Gy and 5 Gy for 3 days). To assess local control in LLC, a total dose of 66 Gy was given (12 Gy for 3 days, then 15 Gy for 2 days). Mice were shielded with lead except for the tumor-bearing hind limb. The care and treatment of the mice were in accordance with institutional guidelines.
Tumor Histology.
Animals were euthanized and tumors excised at specified time points
throughout the experiments. Tumor samples were preserved in paraffin or
OCT Embedding Compound (VWR Scientific, Chicago, IL). Four-µm
sections were cut from OCT blocks and mounted on
poly-L-lysine coated slides. Briefly, sections were fixed
in 1% paraformaldehyde (methanol-free; Polysciences, Warrington, PA).
After blocking with goat serum and quenching endogenous peroxidase
activity, slides were incubated at 4°C overnight with rat-antimouse
CD31 monoclonal antibody (PharMingen, San Diego, CA) diluted to 0.1
µg/ml and were then incubated for 25 min at 37°C with
rabbit-antirat IgG biotinylated secondary antibody (2 µg/ml; Vector,
Burlingame, CA). Appropriate negative controls were performed with rat
IgG2a isotype control (PharMingen), CD31 monoclonal antibody alone, and
secondary antibody alone. ABC Vectastain Standard Elite was applied and
the tissues were developed with the Vector DAB kit. Each slide
was counterstained with a solution (0.03%) of light green SF
yellowish (Fisher, Hanover Park, IL). Ten high-power fields (x400)
from each tumor section were examined using a Nikon Microphot-FX
microscope equipped with a Sony digital camera. Vessels were counted
using Macintosh Image Pro-Plus imaging software. Additional tumor
tissue was placed in 10% neutral-buffered formalin, embedded in
paraffin, sectioned at 6-µm thickness, stained with H&E, and examined
by light microscopy at x400.
Statistical Methods.
Significance was determined using one-way ANOVA, Student’s
t test, and Fisher’s exact test using Jandel Scientific
software.
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RESULTS |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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, upper panel) An extensive
network of vessels was observed both within the Matrigel matrix as well
as in the stromal interface between the skin and the Matrigel. In
contrast, Matrigel plugs from mice receiving twice daily injections of
NM-3 (100 mg/kg/day) showed a dramatic reduction in neovascularization.
(Fig. 1
, lower panel) The few vessels present showed minimal
vascular branching, which were predominantly in the stromal interface.
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). The selectivity of NM-3
cytotoxicity for endothelial cells, but not tumor cells, was consistent
at all concentrations tested (100, 1000, and 10,000 ng/ml;
P < 0.05). NM-3 was also selectively
cytotoxic to HAECs in a dose-dependent fashion (data not shown). When
HUVECs were exposed to NM-3 prior to IR, additive killing was observed
at 500, 700, and 900 cGy (Fig. 2B)
. No enhancement of
IR-induced cytotoxicity was observed when LLC cells were exposed to
NM-3 and 100–900 cGy IR (Fig. 2C)
. Similar results were
observed for Seg-1 cells (data not shown). Taken together, these data
support a selective additive interaction between NM-3 and IR in
endothelial cells. Notably, DAPI staining of HUVEC cultures
revealed that NM-3-induced endothelial cell death was not mediated by
apoptosis (data not shown).
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).
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).
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. The combined treatment group demonstrated a growth
delay of 14 days compared with the control group and 3 days compared
with IR-alone group. In experiments conducted using SQ-20B tumors (initial tumor volume = 681.8 ± 17.2 mm3 ; n = 45), control tumors grew to 5.2 ± 0.94 times original volume by day 20. A similar pattern of tumor growth was observed in the NM-3-alone treatment group (4.3 ± 1.03, day 20). Animals in these two groups were subsequently sacrificed because of tumor burden. Tumors in the IR treatment group (65 Gy) doubled in size by day 6 and regressed to 50% of original volume at day 23. By day 38, IR-treated tumors regrew to twice original volume. Animals treated with NM-3 and IR doubled in size by day 6 and then regressed to 77% of original volume at day 23. Unlike the tumors treated with IR alone, tumors receiving combined therapy with NM-3 and IR failed to reach original volume by day 38 (P = 0.006; t test; data not shown).
Experiments were next conducted to confirm that growth delay
translates to local control in a multifractional experiment. C57BL/6
mice bearing LLC tumors (initial volume, 151.1 ± 4.6
mm3
; n = 29) were
treated with 50 mg/kg/day of NM-3 and 66 Gy over 5 consecutive
days. Tumors in the control group grew to 11.83 ± 1.1
times original volume by day 7. Tumors in the IR and NM-3/IR groups
grew initially, but began regressing on day 7 (IR = 1.36 ± 0.12 times original volume and NM-3/IR = 1.08 ± 0.08 times original volume; Fig. 4C
). On day 21 after initiation of treatment, no measurable
tumor was present in 5 of 12 mice in the IR-alone group. However, when
animals were treated with NM-3 plus IR, 9 of 11 appeared to be
tumor-free (P = 0.089 by Fisher’s exact
test). The day 21 time point was chosen in these studies because, after
this time, mice in both treatment groups began dying of lung
metastases.
Combination of NM-3 and IR Leads to Vessel Disruption in LLC
Tumors.
To assess the effect of NM-3/IR on tumor vessels, LLC tumors were
excised on day 5 or 11 from C57BL/6 mice that were treated with NM-3,
IR, or NM-3/IR. CD31 immunohistochemistry was performed to obtain
microvessel counts and H&E slides were prepared for morphological
analyses. At day 5, tumors in the UTC group had a mean of
22.2 ± 6.6 vessels, NM-3-alone treated group had a mean
of 19.2 ± 3.6, the group treated with IR alone had a
mean of 18.7 ± 2.7, and the combined group had a mean
of 14.2 ± 1.2. No further reduction in vascular density
was detected in any treatment group at day 11. However, the vessels of
tumors in the combined treatment group (NM-3 + IR) at day 11
appeared disrupted compared with those in the UTC group (Fig. 5)
. In addition, there were fewer
distinct small diameter vessels in tumors from the IR alone and NM-3/IR
treated groups compared with that in the control and NM-3 alone groups
(data not shown).
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DISCUSSION |
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TopABSTRACTINTRODUCTIONMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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The mechanism by which NM-3 exerts antitumor effects is not known at this time. However, our in vitro data suggest that NM-3 alters several stages of the angiogenic process including endothelial cell survival, migration, and tube formation. Studies are under way in our laboratory to elucidate the pathways underlying interaction between NM-3 and IR.
An important focus of current clinical radiotherapy is selective targeting of tumors. This approach relies on physical techniques to achieve high doses of radiation to the tumor. Examples are intensity-modulated radiotherapy and stereotactic radiotherapy. Improved tumor control depends on radiation dose escalation. However, the application of dose escalation is limited to relatively small tumors or larger tumors in which normal tissues in the field can tolerate the high radiation doses.
Another current clinical strategy to enhance radiocurability is the use of chemical modifiers. One class of chemical modifiers is proposed to directly radiosensitize hypoxic tumor cells (29) . However, the available hypoxic tumor cell sensitizers possess significant toxicity and have yet to yield positive clinical results. A third strategy of radiosensitization is the use of standard cytotoxins as chemical modifiers of the radiation response. These agents are proposed to directly sensitize tumor cells to IR. Clinical protocols combine IR treatment with cis-platinum, mitomycin C, 5-fluorouracil, hydroxyurea, or combinations of these agents. Although these drugs are likely to be additive with radiation in most clinical circumstances, they have improved cures in head and neck cancer, lung cancer, carcinoma of the cervix, and anal cancer (14, 15, 16, 17, 18, 19) . Clinical results from these combined modality trials reinforce the concept that increased local control increases overall cures. However, these combination regimens have considerable acute tissue toxicities, which lead to patient deaths, morbidity, and increased cost of cancer care when compared with treatment with radiotherapy alone.
The present results describe a new paradigm in which a nontoxic agent potentiates the effects of IR. In this context, selective antitumor effects are achieved with NM-3 and IR without adverse normal tissue effects. These findings indicate that standard risk/benefit analyses may not be relevant to the investigation of treatment with NM-3 and IR because NM-3 has no known normal tissue toxicity and little antitumor effect alone but enhances the antitumor effects of IR. The combination of NM-3 with radiotherapy improves both tumor growth delay and radiocurability. In vitro assays of angiogenesis support mechanisms of NM-3/IR interaction through increased endothelial cell killing and decreased migration. These data are supported by reports of protein fragment angiogenesis inhibitors that potentiate radiotherapy and strengthen the concept of targeting both the tumor vasculature and tumor cells with IR.
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FOOTNOTES |
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1 Supported by ILEX Oncology Inc., by Grant
DE/CA11921 from the Chicago Oral Cancer Center, and by a grant from the
Lederer Foundation. R. R. W., R. K., and D. W. K. have a financial
interest in ILEX. 
2 R. M. S. and N. T. J. contributed equally to
the article.
3 To whom requests for reprints should be
addressed, at Department of Radiation and Cellular Oncology, University
of Chicago, 5841 South Maryland Avenue, Chicago, IL 60637. Phone: (773)
702-0817; Fax: (773) 834-7233; E-mail: rrw{at}rover.uchicago.edu
4 The abbreviations used are: IR, ionizing
radiation; HUVEC, human umbilical vein endothelial cell; LLC, Lewis
lung carcinoma; HAEC, human aortic endothelial cell; hpf, high-power
field(s); UTC, untreated control.
Received 5/31/00. Accepted 10/19/00.
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