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From the Department of Pathology,* Faculty of Medicine, and the Faculty of Pharmacy, University of Manitoba, Winnipeg, Manitoba, Canada; the Department of General Surgery, Nanshan Hospital, Shenzhen, China; the Institute of Environmental Medicine, and the Department of Pathology and Laboratory Medicine,¶ University of Pennsylvania, Philadelphia Pennsylvania; and the Department of Pharmacology,|| University of South Alabama, Mobile, Alabama
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55% of the cells cleared from eNOS-deficient or L-NAME pretreated mice. eNOS knockout and L-NAME-treated mice had twofold to fivefold more metastases than wild-type mice, measured by the number of surface nodules or by histomorphometry. We conclude that tumor cell arrest in the pulmonary microcirculation induces eNOS-dependent NO release by the endothelium adjacent to the arrested tumor cells and that NO is one factor that causes tumor cell apoptosis, clearance from the lung, and inhibition of metastasis.
Evidence from in vitro and in vivo studies has shown that reactive oxygen and nitrogen species can be cytotoxic to neoplastic cells6-9 and reduced their adhesion to postcapillary venules.10 In vivo, we have recently demonstrated that the arrest of intravascular B16F1 melanoma cells in the liver induces the rapid local release of nitric oxide (NO) that causes apoptosis of the melanoma cells and inhibits their subsequent development into hepatic metastases.5 Because pulmonary endothelial cells generate NO in response to shear stress,11 we have postulated that there is a comparable cytotoxic mechanism in the lung.
Here we provide data showing that the arrest of B16F1 melanoma cells in the pulmonary circulation of wild-type C57B1/6 mice (WT mice) induces the local release of NO that is endothelial nitric oxide synthase (eNOS)-dependent. In turn NO causes the melanoma cells to undergo apoptosis and inhibits their development into pulmonary metastatic tumors. The existence of comparable mechanisms in the lung and the liver suggests that NO may be a natural defense against the formation of metastatic tumors in organs with complex microvascular networks.
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Fluoresbrite carboxylate YG microsphere-labeled B16F1 melanoma cells were injected into the tail veins of C57Bl/6 mice. The lungs were removed between 0 and 48 hours after cell injection to measure NO production, tumor cell apoptosis, and the clearance of tumor cells from the lung. To study metastasis, unlabeled B16F1 cells were injected and the lungs were collected 7 or 14 days later to count surface metastatic nodules or to quantify metastasis by histomorphometry. All analyses were performed blindly.
Animals and Materials
WT female C57Bl/6 mice, weighing 20 to 22 g, were purchased from Charles River Breeding Laboratories (Montreal, QC, Canada or Kingston, NY) and housed according to University of Manitoba guidelines. eNOS knockout C57Bl/6 mice (eNOS KO mice), age 5 to 6 weeks, were obtained from Jackson Laboratory (Bar Harbor, ME). The murine B16F1 melanoma cell line was obtained from the American Type Culture Collection (Rockville, MD). Fluorescent polystyrene microspheres (15 µm in diameter) were purchased from Bangs Laboratories, Inc. (Fisher, IN). Fluoresbrite carboxylate YG microspheres (0.05 µm) were purchased from Polysciences, Inc. (Warrington, PA) and MitoTracker Red from Molecular Probes (Eugene, OR). 
-MEM, Opti-MEM reduced serum medium, penicillin-streptomycin, and trypsin-ethylenediaminetetraacetic acid were purchased from Life Technologies (Burlington, ON, Canada). Avertin (2,2,2-tribromoethanol), diethyldithiocarbamate (DETC), and NG-nitro-L-arginine methyl ester (L-NAME) were purchased from Sigma-Aldrich Canada Ltd. (Oakville, ON, Canada). 4,5-Diaminofluorescein diacetate (DAF-2 DA) was obtained from Calbiochem (La Jolla, CA). The ApopTag Peroxidase in Situ Apoptosis Detection Kit S7100 was from Intergen Company (Purchase, NY).
Cell Culture and Fluorescent Labeling
B16F1 cells were cultured in -MEM supplemented with 10% fetal bovine serum and 1% penicillin/streptomycin solution, in a T25 culture flask (Corning Glass, Corning, NY) to 95% confluence. The culture medium was then replaced by Opti-MEM serum-reduced medium and fluoresbrite carboxylate microspheres were added to the flask in a ratio of 100 µl of microspheres per 5 ml of Opti-MEM medium (microsphere stock, 2.5% solids-latex). The flask was gently mixed and put back in the incubator for 2 hours to label the cells with occasional shaking of the flask. The cells were rinsed three times with Opti-MEMI medium to wash off the unincorporated beads and left in -MEM culture medium overnight to reduce cell aggregation on detaching. The labeled cells were detached on the following day with 0.5x trypsin-ethylenediaminetetraacetic acid at 37°C for 5 minutes. The cell suspension was centrifuged at 170 x g for 3 minutes. The cell pellet was resuspended in pyrogen-free saline and kept on ice before injection. Cells with a viability >95%, measured by trypan blue exclusion, were used in the study.
B16F1 Melanoma Cell Injection
Mice were anesthetized with avertin (30 mg/ml; 0.2 to 0.3 ml/mouse, IP). A fluorescent-labeled cell suspension of 5 x 105 B16F1 melanoma cells in a volume of 150 µl of saline was injected into the tail vein throughout 2 to 3 minutes, using a 30G1/2 needle and 1-ml syringe. The lung was collected under anesthesia at specific times (0 to 48 hours) after injection. For 0-hour samples, the lung tissue was removed immediately at the end of the injection (within 30 seconds to 1 minute). To determine whether the shear stress of tumor cells may also play a role in the induction of NO by lung microvascular endothelial cells, we injected 5 x 105 15-µm fluorescent-labeled polystyrene microspheres in a volume of 150 µl of saline intravenously. The lung tissue was collected 20 minutes afterward for electron paramagnetic resonance (EPR) measurement.
L-NAME Administration
Based on our previous study,5 the competitive NO synthase inhibitor L-NAME was administered to inhibit NO production and to test its effects on cell clearance, apoptosis, and metastasis formation. For cell clearance and apoptosis analysis, 5 mg/kg of L-NAME was injected intraperitoneally 1 hour before cell injection and 2.5 mg/kg of L-NAME was co-injected with the cells. For the metastasis assays, 2.5 mg/kg of L-NAME was co-injected with the cells and 5 mg/kg of L-NAME was injected intraperitoneally 20 hours after the cell injection.
EPR Measurement of NO Production
EPR spectroscopy was used to quantitatively measure the production of NO. The NO trapping agents DETC (400 mg/kg in saline, IP) and FeSO4/sodium citrate (40 mg/kg and 200 mg/kg, respectively, mixed in water, SC) were given to each mouse 30 minutes before obtaining the lung sample. The mice were anesthetized 10 minutes before sacrifice. The thorax was opened and the lungs were removed (between 30 seconds to 1 minute) and placed onto a precooled Petri dish on ice. The lungs were quickly sliced into smaller pieces, placed into a precooled 1-ml syringe, and transferred into a Suprasil synthetic quartz tube (2.4-mm inner diameter; Heraeus Amersil, Duluth, GA) by pushing the tissue gently through the syringe to fill the tube up to 3 to 4 cm in height. The tube was then immediately placed into liquid nitrogen until NO was measured by EPR spectroscopy.12
EPR spectroscopy, used for the measurement of trapped NO-Fe2+-(DETC)2 complex, was performed at 125 K (temperature controller model BVT-3000; Bruker Spectrospin Ltd., Karlsruhe, Germany). The spectra were measured with a Bruker model EMX EPR X-band spectrometer system operating at 9.25 GHz with 100 kHz modulation. The instrument settings were as follows: 1) microwave power, 5 mW; 2) modulation amplitude, 5 G and signal level, 1 x 103; and 3) scan range, 500 G. The concentration of the NO-Fe2+-(DETC)2 complex in each sample was assumed to be proportional to the signal amplitude (peak-to-trough) of the triplet-hyperfine structure (hyperfine splitting of 13 G) observed at g = 2.04 (see below). Data were expressed as relative EPR signal intensities (arbitrary units) after subtracting the Cu2+-(DETC)2 complex signal observed in all samples. As described in our previous work,5
we performed a double-integration calibration to calibrate the conversion from peak-to-peak EPR amplitudes to nmol ON-Fe2+-(DETC)2/gram of wet tissue concentrations (see the caption to Figure 1
). The only difference is that we used the Bruker EPR spectrometer for our present work, whereas the Varian E-12 spectrometer was used previously.5
We have performed EPR measurements on the same samples at 125 K on both the Varian and Bruker EPR systems to relate the peak height measurements from one instrument to the other. AWIN-EPR software (Bruker Spectrospin Ltd.) was used for nearly all EPR data manipulation on a Compaq Deskpro P500 computer (Compaq Computer Corp., Houston, TX).
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NO generation was monitored in situ by labeling the pulmonary endothelium with DAF-2 DA that is de-esterified intracellularly to DAF-2. NO and its higher oxides, such as NOx or nitrous anhydride (N2O3), provide the third nitrogen to form a triazo ring from the two amino groups of the nonfluorescent DAF-2 and convert it to diaminotriazolofluorescein (DAF-2T) that is fluorescent and can be monitored at 490 nm excitation and 530 nm emission.13
We used an established intact organ microscopy technique to image microvascular endothelial cells in situ in isolated, ventilated, blood-free mouse lungs in real time using an epifluorescence microscope.14 Briefly, female WT mice, weighing 20 to 22 g were anesthetized with intraperitoneal sodium pentobarbital (50 mg/kg). A tracheostomy was performed, and artificial ventilation with 95% air and 5% CO2 (BOC Group, Inc., Murray Hill, NJ) was started through a cannula. The abdomen was opened and the animal was exsanguinated by transection of the major abdominal vessels. A cannula was inserted into the main pulmonary artery via a puncture in the right ventricle, and another was inserted into the left atrium. The lung was cleared of blood by gravity perfusion via the pulmonary artery with an artificial medium (Krebs-Ringer bicarbonate buffer KRB: NaCl, 118.45; KCl, 4.74; MgSO4/7H2O, 1.17; CaCl2/2H2O, 1.27; KH2PO4, 1.18; and NaHCO3, 24.87, in mmol/L, with 5% dextran and 10 mmol/L glucose at pH 7.4). The flow-through perfusate left the lung via the left atrial cannula. Once the lung became visibly cleared of blood, the heart-lung preparation was dissected en bloc and was placed onto a 48 x 60 x 0.16-mm coverglass window in a specially designed Plexiglas chamber with ports for the tracheal, pulmonary, and left atrial cannulas. The cardiovascular ports were connected to a peristaltic pump that recirculated 30 ml of perfusate at a constant flow rate of 2 ml/min through the pulmonary vascular bed. The chamber was placed on the stage of an epifluorescence microscope fitted with a x60 objective (Nikon Diaphot TMD) and equipped with an optical filter changer (Lambda 10-2; Sutter Instrument Co., Novato, CA). A local anesthetic (0.05 mg of xylazine, Sigma) was injected subepicardially into the posterior wall of the right atrium to abolish lung movement artifact because of contraction of remaining cardiac muscle. Excitation of the lung surface was accomplished with a mercury lamp fiber optic light source, a fluorescein isothiocyanate filter set for DAF-2T (HQ41001 with 480/40 excitation filter, 505 LP dichroic mirror, and 535/40 emission filter; Chroma Technology Corp., Brattleboro, VT). The integrity of the preparation was continuously monitored by online measurements of intratracheal and pulmonary artery perfusion pressures. Endothelial cells in the subpleural vasculature were positively identified by labeling with DiI-acetylated low-density lipoprotein added to the perfusate (a tetramethylrhodamine isothiocyanate filter set for DiI-acetylated low-density lipoprotein, Chroma Technology Corp.). We used a Nikon Diaphot TMD epifluorescence microscope, a Hamamatsu ORCA-100 digital camera (Hamamatsu Corp., Bridgewater, NJ), and MetaMorph imaging software (Universal Imaging Corp., Downingtown, PA) for imaging. After an equilibration period of 45 minutes with the isolated lung to allow uptake of DAF-2 DA (5 µmol/L), intravascular dye was removed by perfusion with dye-free medium for 5 minutes to reduce background fluorescence. Mitotracker-Red labeled B16F1 cells (2 x 106) were injected through a pulmonary artery port by peristaltic pump at 2 ml/min. Images of DAF-2T-stained vascular endothelial cells were taken from the same area every 3 seconds for up to 20 minutes during which ventilation was stopped. As a control, after the equilibration period, images were taken during continuous perfusion.
For quantification, each endothelial cell was outlined and its fluorescence intensity was measured throughout time. For each lung, mean fluorescence intensities of four to seven endothelial cells were averaged. Fluorescence intensity of three to four lungs for each condition was averaged. Data are expressed as relative fluorescence intensities (arbitrary units). All results are expressed as mean ± SE for each condition.
Assessment of B16F1 Cell Apoptosis in Vivo
Lung tissues for in situ DNA end-labeling were collected at 0, 4, 8, 24, and 48 hours after tail vein injection of fluorescent-labeled B16F1 cells. No trapping agent was given to these animals. For each time point five animals were used. Frozen sections (12 µm) were obtained from the upper and median lobes of the right lung and upper lobe of the left lung. In situ DNA end labeling using a digoxigenin-peroxidase detection system (ApopTag S7100 kit, Intergen) was performed on these sections that contained injected fluorescent-labeled melanoma cells. To achieve optimal double labeling, the procedures were performed according to the manufacture’s instructions with the following modifications, as previously described.5 1) Proteinase K digestion (20 µg/ml) was added before postfixing the sections with ethanol/acetic acid (2:1, v/v), to facilitate the unmasking of fragmented DNA in apoptotic tumor cells. 2) Incubation for in situ DNA end labeling reaction was performed overnight in a humidified chamber at 37°C, with subsequent 1 hour anti-digoxigenin peroxidase conjugate incubation at room temperature in a humidified chamber and 12 minutes peroxidase substrate color development. 3) The counter stain specimen and mount specimen steps were omitted to prevent the quenching of fluorescence in the microspheres by organic solvents. 4) The slides were mounted with Gel/Mount (Biomeda Corp, Foster City, CA). To generate positive control samples for apoptosis, labeled B16F1 cells were exposed to 2 µm of NO donor S-nitroso-N-acetyl-penicillamine (SNAP) for 24 hours. Detached and collected cell pellets were resuspended in 1% paraformaldehyde at 1 x 106/150 µl of saline. Lung tissues were collected immediately after injection and fixed in 1% paraformaldehyde.
The total number of both single fluorescent- and double-labeled cells (fluorescent/ApopTag DNA end-labeling) was scored using a fluorescence microscope with dual illumination of the fluorescence and incandescence (x200 magnification). More than 100 total cells from both left and right lobes were counted for most samples. The percentage of in situ DNA end-labeled melanoma cells was calculated by the formula: in situ end-labeled tumor cells (%) = 100 x [number of double-labeled cells]/[total number of cells (double labeled + single labeled)].
Analysis of B16F1 Cell Clearance and Fragmentation
Frozen sections (12 µm) were obtained from the upper and median lobes of the right lung and the upper lobe of the left lung, collected at 0, 4, 8, 24, and 48 hours as described above. Fluorescent-labeled B16F1 cell fragments and the intact fluorescent-labeled B16F1 melanoma cells in 100 microscopic fields (x200 magnification) were counted from each lobe to determine the number of B16F1 cells in the lung at each time point. The ratio of cell fragments to intact cells at 8 hours and 48 hours was calculated as follows: cell fragments (%) = (cell fragments/cell fragments + intact cells) x 100. Data were expressed as the mean of five values.
Metastasis Studies
Unlabeled B16F1 cells (5 x 105/150 µl saline) were injected into the tail vein of the mice and allowed to grow in vivo for 7 or 14 days. All animal surgical procedures were approved by the Bannatyne Campus Protocol Management and Review Committee at the University of Manitoba. All animal care was given according to the guidelines of the Central Animal Care Services at the University of Manitoba. The mice were sacrificed under anesthesia and the lungs were fixed in 10% neutral buffered formalin. All metastatic nodules identifiable on the surface of all lung lobes were counted using a dissecting microscope on the day 7 after tumor cell injection. Fourteen days after injection, histomorphometric analysis was performed on histological sections from all five lobes using a Merz Graticule to quantify the percentage of tissue area occupied by the metastases.
Statistical Analysis
Prism software was used for statistical analysis. For EPR, tumor cell apoptosis and clearance, an unpaired t-test was used. The Mann-Whitney U-test was used for metastasis studies. Comparisons of DAF-2T fluorescence intensity were made using analysis of variance with Bonferroni’s test using SigmaPlot 2000 (SPSS Inc., Chicago, IL).
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To determine whether NO is released in the lung after B16F1 cell arrest, we injected 5 x 105 B16F1 cells, in a volume of 150 µl of saline, into the lateral tail vein of the WT C57Bl/6 mice. We injected the NO spin trap DETC/FeSO4 30 minutes before harvesting the tissue to detect NO generation by EPR spectroscopy. As shown in Figure 1
, NO generation was induced as soon as tissue could be harvested after cell injection, increased sevenfold within 20 minutes after cell injection compared to the saline control at 0 minutes, and approached basal levels by 4 hours. Rapid and significant production of NO was also detected in mice injected with 5 x 105 fluorescent microspheres with similar diameter to that of B16F1 cells (16 ± 4 µm). In these animals, the peak NO level was significantly lower than that induced by B16F1 cell injection (66% of peak NO level of B16F1 cells, P < 0.01). The injection of saline alone was not associated with significantly increased NO production. Pretreatment of animals with the NO synthase inhibitor L-NAME inhibited NO production at 20 minutes by 64%. NO induction was completely absent in mice defective in the eNOS gene. NO was not detected in B16F1 cells (data not shown).
To identify the cellular origin of NO, 2 x 106 fluorescent-labeled B16F1 cells were introduced into the perfusate of isolated intact WT mouse lungs in which the endothelium had been preloaded with DAF-2 DA. A low level of fluorescence, indicative of NO production, was seen after loading in all groups. Enhanced fluorescence was seen within 2 minutes of attachment to the pulmonary endothelium of WT mice. This fluorescence increased throughout the next 20 minutes at the site of arrest and also in adjacent endothelial cells. Fluorescence did not change from basal levels after arrest of the tumor cells in eNOS-deficient mice (Figure 2
, Table 1
).
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To determine whether there was a correlation between NO production and apoptosis, lung tissues were harvested at various times after injecting 5 x 105 fluoresbrite microsphere-labeled cells. Cryostat sections of the lung tissue were stained for apoptotic cells with the ApopTag kit. Tumor cells were identified by their fluorescence (Figure 3A)
and scored for apoptosis (Figure 3B)
. Tumor cells became fragmented by 48 hours after injection (Figure 3C)
. Fragmentation was greater in WT mice than in eNOS KO mice or L-NAME-treated mice (Table 2)
. As shown in Figure 4A
, apoptosis of tumor cells increased with time after injection in WT mice. Enhanced apoptosis could be seen as soon as 4 hours after cell injection, reached a peak value threefold greater than baseline at 8 hours, and remained detectable throughout a 48-hour period. Apoptosis of tumor cells did not increase above baseline values in mice defective in the eNOS gene or in mice pretreated with L-NAME.
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47% of tumor cells had been cleared from the lungs of WT mice; 83% had been cleared by 48 hours. Clearance was less in mice defective in the eNOS gene or in mice pretreated with L-NAME (Figure 4B)
. Metastasis
To evaluate the effect of NO induction on metastasis, we injected 5 x 105 unlabeled B16F1 melanoma cells intravenously into WT mice, eNOS KO mice, or WT mice treated with L-NAME. Lung metastases were measured by counting the number of nodules on the visceral pleural surfaces at 7 days or by histomorphometry at 14 days. Seven days after tumor cell injection eNOS KO mice had 2.3-fold more pleural surface metastases than WT mice (P < 0.01) and WT mice pretreated with L-NAME had 2.4-fold more metastases (P < 0.05). Fourteen days after B16F1 cell injection there was a twofold greater tumor burden in the lungs of L-NAME-treated WT mice (P < 0.003) and 4.9-fold greater tumor burden in the lungs of eNOS KO mice (P < 0.001), compared to the lungs of untreated WT mice. The data are shown in Table 3
.
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As in the liver, the arrest of tumor cells in the lung microvasculature induced an immediate burst of NO. However, unlike the liver, where increased NO production was detected for less than 5 minutes, the expression of NO in the lung increased throughout a period of 20 minutes and then decreased subsequently throughout a period of 4 hours. Use of the DAF-2 staining method allowed us to further determine that, in the lung, NO originated from microvascular endothelial cells. Involvement of eNOS was confirmed by the complete absence of a response in eNOS KO mice and by the inhibition of the NO burst in L-NAME-treated mice.
The use of two distinct methods to detect NO production provided consistent evidence that endothelial-derived NO is released after the arrest of tumor cells in the pulmonary microvasculature. NO is a small diatomic-free radical, but it plays complex roles in diverse biological processes in organisms ranging from bacteria to mammals. The main difficulty in measuring NO production is caused by its extremely labile nature and short half-life, which is in the millisecond range. Among the indirect NO detection methods widely accepted in the literature, we chose to use spin trapping of a NO adduct in vivo and then measure the trapped NO signal ex vivo by EPR spectroscopy. The main advantages of spin trapping are that NO has a high affinity for the spin trapping iron complex and can instantaneously form a much more stable NO-Fe2+-(DETC)2 complex. This complex generates a characteristic three-line structure on the EPR spectrum that can be distinguished from other signals. This technique has been shown to be a reproducible and sensitive NO detection method. We argue that it is the sensitivity and specificity of the EPR spin trapping method that helped us to capture the transient burst of NO release in the lung after the melanoma cell injection. We further quantified the NO peak concentration to be in the range of 50 to 100 nmol/gram of wet tissue using a well-calibrated in vitro standard. In addition to the three-line NO-Fe2+-(DETC)2 spectrum, a quartet hyperfine spectrum of Cu2+-(DETC)2 was also detected, which partially overlapped with the NO-Fe2+-(DETC)2 spectrum. Our data were expressed as the relative EPR signal intensity (equivalent to the peak-to-trough amplitude of the three-line spectrum) with the Cu2+-(DETC)2 spectrum subtracted, so that they represent the pure NO-Fe2+-(DETC)2 signal intensity. To our knowledge, this is a more specific approach because most analyses measure the NO concentration on the basis of the uncorrected EPR spectrum alone. The measurements obtained by using DAF-2 DA gave direct morphological evidence for the release of NO by endothelial cells that were in direct contact with arrested tumor cells. Because there was a very low but measurable amount of NO in eNOS knockout mice by EPR spectroscopy, it is possible that minimal amounts of NO were produced by other NO synthase isoforms.
We postulate that the mechanism responsible for the rapid release of NO from the pulmonary endothelium may be similar to that in the liver, ie, at least partially triggered by local shear forces produced by cell arrest in the microvasculature. In our previous and present studies, the NO burst could be elicited by injection of inert microspheres into the portal5 and pulmonary circulation.
Our data support the hypothesis that NO is an effector of B16F1 melanoma cell apoptosis in the lung. There is a large body of evidence that generation of NO, mediated by inducible NOS, can have cytotoxic effects on neoplastic cells and inhibit their potential for growth in vivo.15-24 In addition to this mechanism, which generates NO throughout a sustained period of time,24 we have shown here that tumor cell arrest can trigger the immediate release of NO via eNOS-dependent mechanisms and subsequently trigger apoptosis in these cells. Previous experiments in the liver have inferred a role for eNOS as an effector of B16 melanoma cytotoxicity on the basis that NO release occurred immediately (<5 minutes) after tumor cell arrest in the hepatic sinusoid.5 In a recent study, Carretero and colleagues9 have used liver endothelial cells from eNOS-/-, eNOS+/+, and WT mice, treated with cytokines, to observe the release of NO and H2O2 and the cytotoxicity of these molecules to B16F1 cells. They found that H2O2 was not cytotoxic in the absence of NO, while NO-induced tumor cytotoxicity was increased by H2O2 because of the formation of potent oxidants, likely OH- and -OONO. Their work was conducted in vitro. In contrast, our work has focused on observations in vivo showing that tumor cell arrest in the lung triggers immediate NO release that is eNOS-dependent and cytotoxic. This is consistent with the observation that even a very brief exposure (5 minutes) to NO in vitro is capable of triggering apoptosis in the B16F1 cells.5 It may be relevant to our observations that, at a clinical level, expression of immunoreactive eNOS in peritumoral microvessels has been identified as a favorable prognostic indicator in premenopausal breast cancer patients.25
In addition to direct DNA damage caused by NO, its cytotoxic effects can also be mediated through generation of peroxynitrite (ONOO-) in the presence of superoxide anion (O·2-).26 As a potent oxidant, peroxynitrite can produce tissue damage through nitration of tyrosine residues in protein to form nitrotyrosine.27 We are currently conducting experiments to determine whether nitrotyrosine can be identified in plasma28 or at sites of NO release as a mediator of cytotoxicity, and as a reflection of low level NO production in eNOS knockout mice.
Counts of fluorescent-labeled B16 cells in lung sections, prepared at various times after intravenous injection, confirmed that metastatic cells are cleared from the lungs, as they are in the liver. Tumor cell clearance was regulated in part by NO because fewer cells were cleared from the lungs of L-NAME-treated and eNOS KO mice than from the lungs of WT mice. The mechanisms responsible for this may involve alterations in adhesion between the cancer cells and endothelium because NO has been reported to reduce the attachment of tumor cells to the endothelium.10 NO-mediated vasodilatation of pulmonary arterioles and venules29 could also diminish cell trapping because of mechanical restriction. NO cytotoxicity to tumor cells may also contribute to the tumor cell clearance because morphologically, cell clearance was associated with fragmentation of the tumor cells and this was also greater in WT mice, compared with L-NAME-treated mice and eNOS KO mice. Previous flow cytometric analysis of B16 cells indicates that necrosis and apoptosis both occur in response to short exposure of these cells to NO.5 As in our previous studies, we found that apoptosis and clearance of tumor cells from the lung was associated with fewer metastatic nodules at 7 days and a lesser tumor burden measured by histomorphometry at 14 days.
Extensive experimental and clinical data indicate that the majority of tumor cells die rapidly in the circulation, and only a few of them can survive and proliferate to form distant metastases.23,30,31 Wong and colleagues32 demonstrated that this cell death in the lung was because of apoptosis and the observations of our lab in the liver would suggest that the death of these cells by apoptosis is a frequent event.5 The importance of apoptosis in the regulation of pulmonary metastasis is evident both because inhibition of apoptosis by overexpression of bcl-2 leads to more metastasis and because metastatic cells undergo markedly less apoptosis than nonmetastatic cells after arrest in the lung.32,33 Deformation-induced mechanical trauma and immune-mediated host cell killing have also been implicated in the intravascular death of tumor cells.31,34,35 The data that we have presented here demonstrate that the local release of NO at the site of tumor cell arrest contributes to this intravascular cytotoxicity although the limited survival of tumor cells even in mice deficient in eNOS would suggest that other factors must play a role as well.
Although there is evidence that NO may be more cytotoxic to weakly metastatic tumor cells than to highly metastatic cells,8 in this study, we have used the B16F1 cell line rather than B16F10 cells because the number of metastatic nodules induced by our protocols was amenable to statistical analysis and because the B16F1 cells are susceptible to NO in vitro and in vivo.5 Preliminary experiments in our laboratories suggest that related cell lines of varying metastatic competence elicit NO release of differing magnitude. This is the subject of a current project.
In conclusion, we provide direct evidence showing that the arrest of B16F1 melanoma cells in the pulmonary circulation of C57Bl/6 mice induces the local release of eNOS-dependent NO that is cytotoxic to the cells and inhibits their development into pulmonary metastatic tumors. The existence of comparable mechanisms in the lung and the liver suggests that NO may be a natural defense against the formation of metastatic tumors in organs with complex microvascular networks.
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Supported by the Canadian Institutes of Health Research (CIHR) of Canada (grants MT-14356 to F. W. O. and D. M. N., and MT-14477 to B. H.), the National Cancer Institute (grants NCI CA-46830 and NCI CA-89188 to R. J. M.), and the Susan G. Komen Breast Cancer Foundation (BASIC99-003201 to A. A.).
Accepted for publication October 14, 2002.
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  L. Ying and L. J. Hofseth An Emerging Role for Endothelial Nitric Oxide Synthase in Chronic Inflammation and Cancer Cancer Res., February 15, 2007; 67(4): 1407 - 1410. [Abstract] [Full Text] [PDF]
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, saline injection into WT mice; 
, B16F1 cell injection into L-NAME-treated WT mice; 
), B16F1 cells injected into eNOS KO mice (n = 5 to 10 mice/point). Data are expressed as the mean ± SE. *, P < 0.05 as compared to saline control at 0 minutes; **, P < 0.01 as compared to microspheres, L-NAME, saline controls, and eNOS KO mice at 20 minutes; ***, P < 0.01 as compared to L-NAME, saline controls, and eNOS KO mice at 20 minutes. B: Representative complete EPR spectra of NO-Fe2+-(DETC)2 complex (triplet-hyperfine structure at g = 2.04) measured 20 minutes after B16F1 melanoma cell injection [Cu2+-(DETC)2 signal was subtracted].
