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Departments of Medicine and Physiology, Cardiovascular Research Institute, University of California, San Francisco, San Francisco, California, USA
1Correspondence: 1246 Health Sciences East, University of California, San Francisco, CA 94143-0521, USA. E-mail: verkman{at}itsa.ucsf.edu
ABSTRACT
Aquaporin (AQP) water channels are expressed in high-grade tumor cells of different tissue origins. Based on the involvement of AQPs in angiogenesis and cell migration, we tested whether AQP expression in tumor cells might enhance their migration and metastatic potential. Transfection of B16F10 and 4T1 tumor cells with AQP1 did not affect their appearance, size, growth, or substrate adherence but increased their plasma membrane osmotic water permeability by 5- to 10-fold. In vitro analysis of cell migration by transwell assay, wound healing and video microscopy showed a 2- to 3-fold accelerated migration of the AQP1-expressing tumor cells compared to control cells. In mice, AQP1 expression increased tumor cell extravasation by >1.5-fold as quantified by counting tumor cells in lung at 6 h after tail vein injection of a mixture of fluorescently tagged AQP1-expressing and control tumor cells. AQP1 expression also increased by 3-fold the number of lung metastases 14 days after tail vein injection of tumor cells, with alveolar wall infiltration seen with AQP1-expressing tumor cells. Our results provide evidence for AQP-facilitated tumor cell migration and spread, suggesting a novel function for AQP expression in high-grade tumors. AQP inhibition may thus reduce the metastatic potential of some tumors.—Hu, J., Verkman, A. S. Increased migration and metastatic potential of tumor cells expressing aquaporin water channels.
Key Words: aquaporin • water channel • tumor metastasis • cell migration • cancer
THE AQUAPORINS (AQPS) are a family of small transmembrane proteins that facilitate osmotically driven water transport, and in some cases the transport of small solutes such as glycerol. The best established role of AQPs in mammalian physiology is in transepithelial fluid transport, as occurs in the urinary concentrating mechanism and glandular fluid secretion (1
, 2)
. Several newer and in some cases unanticipated roles of AQPs have been identified, such as in the regulation of brain and corneal water balance (3
, 4)
, neural signal transduction (5)
, and glycerol metabolism in skin (6)
and fat (7)
cells. Recently, we reported impaired angiogenesis and endothelial cell migration in mice lacking AQP1 (8)
, and suggested that AQP-dependent cell migration might be a general cellular phenomenon whose mechanism involves AQP-facilitated water influx into dynamic cellular protrusions (lamellapodia) at the leading edge of migrating cells.
A potential consequence of AQP-facilitated migration, if it occurs in tumor cells, is enhanced metastatic potential produced by accelerated cell migration across microvessels and into normal tissues. Several studies have reported AQP expression in a variety of human tumors, which in some cases was correlated with tumor grade (9
10
11
12
13
14
15
16
17)
. AQP expression in tumors has also been proposed to be of diagnostic and prognostic value (18
, 19)
. The functional role of AQP expression in tumors is, however, unknown. We hypothesized that AQPs in tumor cells may facilitate their migration and thus their metastatic potential, which, if correct, would suggest AQP inhibition as a possible strategy for inhibition of tumor spread.
Here, the involvement of AQPs in tumor cell migration and spread was tested in vitro and in mice using mouse tumor cell lines without vs. with AQP1, a water-selective AQP that does not affect cell growth or other vital cell functions (8
, 20
, 21)
. In vitro studies showed increased water permeability and migration of AQP1-expressing tumor cells, yet similar cell growth, size, and substrate adherence. By differential fluorescent labeling of AQP-expressing vs. control tumor cells, we found significantly enhanced tissue extravasation of intravenously (i.v.) delivered AQP1-expressing tumor cells. A greater number of lung metastases were also found after intravenously delivered AQP1-expressing tumor cells, with evidence for alveolar wall tumor cell invasion. Subcutaneous tumor growth was not changed by AQP1 expression, though local invasion was seen with AQP1-expressing tumor cells. Our results suggest AQP expression and tumor cell water permeability as potentially important determinants of tumor spread.
MATERIALS AND METHODS
Mice
Experiments were done on weight and sex-matched wild-type mice in C57BL/6 and BALB/C genetic backgrounds (Jackson Lab, Bar Harbor, ME, USA). Protocols were approved by UCSF Committee on Animal Research.
Cell lines
B16F10 melanoma cells (American Type Culture Collection CRL-6457, ATCC, Rockville, MD, USA) were cultured at 37°C in a humidified atmosphere containing 5% CO2 with Dulbecco’s modified Eagle medium (DMEM) supplemented with 4 mM L-glutamine, 100 U ml–1 penicillin, 0.1 mg ml–1 streptomycin, and 10% FBS. 4T1 mammary gland tumor cells (American Type Culture Collection CRL-2539) were cultured with RPMI 1640 supplemented with 2 mM L-glutamine, 10 mM HEPES, 1 mM sodium pyruvate, 100 U ml–1 penicillin, 0.l mg ml–1 streptomycin, and 10% FBS.
Cell transfection
Cells were plated to 
80% confluence in 12-well plates 24 h before transfection. 2 µg of recombinant plasmid pcDNA3.1-AQP1 or pcDNA3.1-yellow fluorescent protein and 4 µg of Lipofectamin 2000 (Invitrogen, Carlsbad, CA, USA) were added into 100 µl Opti-MEM medium for 5 min according to manufacturer’s instructions. After 48 h the cells were trypsinized and plated on 10 cm diameter dishes, and hygromycin B (500 µg/ml; Roche, Nutley, NJ, USA) was added for selection. At 12–18 days, hygromycin B-resistant cell clones were isolated and transferred to separate culture dishes for expansion and analysis.
Fluorescent labeling
Cells were incubated for 30 min at 37°C with Opti-MEM containing the fluorescent cell tracker dyes 5-chloromethylfluorescein diacetate (CMFDA, green fluorescent) or a rhodol-based fluorophore (CMRA, red fluorescent) at 2 µM (Molecular Probes, Eugene, OR, USA). After washing, cells were incubated for an additional 30 min with dye-free medium, washed, and trypsinized.
Tumor cell growth, size, proliferation, and substrate adherence
Cell growth curves were generated after seeding cells in 24-well plates at a density of 5 x 103 cells/ml and counting triplicate wells every 24 h for 7 days. Cell diameter was determined by photographing cells at high magnification after trypsinizing and suspending in medium. Cell proliferation was measured by 3H-thymidine incorporation. Three days after seeding cells at 105/ml, the culture medium was replaced by serum free medium for 24 h, then pulsed with 1 µCi/ml [3H]thymidine (Amersham, Arlington Heights, IL, USA) for 6 h at 37°C. Cells were washed in PBS, solubilized, and DNA was precipitated with 10% ice-cold trichloroacetic acid for determination of [3H]thymidine radioactivity by scintillation counting, and DNA content by bisbenzimide (Hoechst 33258; Sigma, St. Louis, MO, USA) fluorescence. Results of the proliferation assays are presented as the mean and SD of triplicate cultures. For measurements of cell substrate adherence, flat-bottom 96-well plates were coated overnight at 4°C with 10 µg/ml fibronectin, 10 µg/ml collagen 1, 20 µg/ml laminin-1, or 1% BSA (as control). Wells were washed with PBS and blocked with 1% BSA for 1 h at 37°C. To each well was added 0.1 ml of cell suspension in Opti-MEM (2x105 cells/ml), plates were incubated for 60 min at 37°C in 5% CO2, and nonadherent cells were gently washed away using Opti-MEM. Adherent cells were fixed with 1% glutaraldehyde, stained with 0.1% crystal violet, washed with PBS, and quantified by absorbance at 595 nm in a plate reader.
Transwell migration and invasion assays
Assays were performed using a modified Boyden chamber (Corning Costar, Rochester, NY, USA) containing a gelatin-coated polycarbonate membrane filter (6.5 mm diameter, 8 µm pore size) (8)
. The upper surface of the filter was coated with 20 µl Matrigel (0.3 mg ml–1; BD Biosciences, Bedford, MA, USA). The upper chamber contained cells in culture medium (1.5x105/ml) with 1% FBS, and the lower chamber contained culture medium with 10% FBS (chemoattractant) or 1% FBS (control). Cells were incubated for 6 h at 37°C in 5% CO2. Nonmigrated cells were scraped from the upper surface of the membrane with a cotton swab, and migrated cells remaining on the bottom surface were counted after staining with Coomassie blue. Cell counting done by fluorescence microscopy in experiments where fluorescently labeled cells were used.
Wound healing assay
Cells were cultured as confluent monolayers, synchronized in 1% FBS for 24 h, and wounded by removing a 300–500 µm-wide strip of cells across the well with a standard 200 µl pipette tip. Wounded monolayers were washed twice to remove nonadherent cells. Wound healing was quantified using Image J software as the mean percentage of the remaining cell-free area compared with the area of the initial wound. In some studies wound healing was followed using time-lapse phase-contrast microscopy (181 frames over 15 min or 3 h). For these studies, cells were seeded in 35 mm Petri dishes with 14 mm diameter and glass-bottomed microwell (MatTek, Ashland, MA, USA) and kept at 37°C in a humidified atmosphere with 5% CO2. Lamellipodia "width" was quantified as lamellipodial area divided by lamellipodial length at the wound edge.
Histology
Cells, tumors, and lungs were fixed in 4% paraformaldehyde and embedded in paraffin or OCT for paraffin and frozen sections, respectively. Paraffin-embedded sections were stained with hematoxylin and eosin. Immunofluorescence localization of AQP1 was done using a purified rabbit polyclonal antibody (Chemicon, El Segundo, CA, USA) by standard procedures. AQP1 immunohistochemistry was done using ImmPRESSTM anti-rabbit immunoglobulin (Vector Labs, Portland, ME, USA) and diaminobenzidine as substrate.
Osmotic water permeability measurements
Cells grown on cover glasses were stained with calcein by incubation for 30 min with 10 µM calcein acetoxymethyl ester (AM) (Molecular Probes) and mounted in a perfusion chamber designed for rapid solution exchange. Solutions were exchanged from 300 to 600 mosM PBS (sucrose added to increase osmolality) and the rate of change of calcein fluorescence was monitored to determine water permeability, as described (22)
.
Tumor cell extravasation in mice
Control and AQP1-expressing tumor cells were labeled in vitro with CMFDA or CMRA as described above. Cells labeled with the red and green fluorescent dyes were mixed at a ratio of 1:1, and 2.5 x 106 cells suspended in PBS were introduced by tail vein injection. The ratio of green-to-red fluorescent cells in the injected suspension was measured by counting in a fluorescence microscope. Lungs were harvested 10 min or 6 h after injection. The trachea was cannulated with polyethylene PE-50 tubing and the pulmonary artery with PE-90 tubing. After transecting the left atrium, lungs were perfused in situ with PBS followed by 4% paraformaldehyde (in PBS) at constant pressure (25–35 cm H2O); 0.5 ml of 4% paraformaldehyde was also infused into the airspaces through the tracheal cannula. Lung tissue was sectioned at 5 µm in a cryostat and fluorescent cells in 5 random fields of each slice were counted.
Tumor growth and metastasis in mice
106 4T1 cells were injected i.v. by tail vein in BALB/c mice. Mice were sacrificed after 14 days, and lungs were harvested for hemotoxylin/eosin staining and AQP1 immunohistochemistry. The number of tumor colonies in lung was counted, and colony size and the alveolar wall thickness in a 50 µm peritumoral region were measured using Spot software. In some experiments, 2 x 105 4T1 or 106 B16F10 cells were injected s.c. between the shoulder blades. Tumor length (L) width (W) were measured with a caliper for estimation of tumor volume as 0.52xLxW2 every 3 days for 18 days.
RESULTS
Characterization of tumor cell lines
Two tumor cell lines were selected for the studies here (B16F10 and 4T1 cells) from more than six mouse tumor cell lines evaluated for their low water permeability, stable AQP1 expression after transfection, and metastatic potential in mice. Control and AQP1-transfected B16F10 and 4T1 cell lines were characterized. Phase-contrast micrographs in Fig. 1
A indicate a similar appearance of control and AQP1-expressing cells in each line. Figure 1B
shows plasma membrane AQP1 protein expression in the transfected cells by immunofluorescence. Osmotic water permeability was measured to verify functional plasma membrane AQP1 expression using a calcein swelling assay (22)
, in which the cytoplasm was stained with calcein prior to inducing osmotic volume changes by superfusion with solutions of different osmolalities. Cell swelling produces an immediate increase in calcein fluorescence upon dilution of cytoplasmic proteins that quench calcein fluorescence. Figure 1C
shows relatively slow osmotic equilibration in control cells with reciprocal exponential time constant 
–1 0.1 s–1. Water permeability in the AQP1-transfected cells was substantially increased, consistent with the immunofluorescence data in Fig. 1B
.
|
Figure 1D
summarizes proliferation and growth studies. AQP1 expression did not significantly effect the proliferation or growth of B16F10 or 4T1 cells as assayed by 3H-thymidine incorporation and cell counting.
Increased in vitro migration of AQP1-expressing tumor cells
In vitro analysis of tumor cell migration was done using transwell migration and wound healing assays. In the transwell migration assay, cells in medium containing 1% serum were added to the upper surface of a Boyden chamber and allowed to adhere. Migration was measured over 6 h by contacting the chamber (containing a porous membrane with 8 µm diameter pores) with medium containing 1% (control) or 10% serum. Cells were stained with Coomassie blue, the number of adherent cells was counted, then nonmigrated cells were scraped off the upper surface of the porous filter to reveal the migrated cells (Fig. 2
A, top). Figure 2A
(bottom) shows significantly greater migration of AQP1-transfected cells. Migration was relatively low in the absence of a chemoattractant stimulus (1% serum). Cell labeling with the fluorescent cell tracker dyes CMFDA (green fluorescent) or CMRA (red fluorescent), to be used in in vivo mouse studies below, did not significantly affect migration, nor did cell transfection with a yellow fluorescent protein (YFP) -encoding plasmid ("mock"). Figure 2B
shows that cell adherence to filters coated with collagen, fibronectin, or laminin was not different in control vs. AQP1-tranfected cells. Figure 2C
shows that AQP1 expression increased tumor cell migration in an "invasion" assay, in which cells migrated though serial barriers consisting of a Matrigel layer and the porous filter.
|
A complementary would healing assay of in vitro cell migration was done in which an 300 µm-wide linear strip of cells was scraped from confluent monolayers using a pipette tip. Wound closure was quantified from serial micrographs as shown in Fig. 3
A. In agreement with the transwell assay, the wound healing assay showed significantly accelerated wound closure in the AQP1-transfected vs. control tumor cells at 15 h after scratch (Fig. 3B
). Figure 3C
shows polarized expression of AQP1 at the leading edge of AQP1-expressing migrating cells at the wound edge, which was seen in 40% of AQP1-expressing B16F10 and 4T1 cells. Cell protrusions accompanying migration were quite different in the two cell types, with 4T1 cells showing multiple bleb-like lamellipodia (not shown), as seen with CHO cells (8)
, whereas B16F10 cells showed wide lamella (Fig. 3D
), as seen in glial cells (23)
. Video imaging of migrating B16F10 and 4T1 cells indicated more frequent bleb/lamella formation after AQP1 transfection (not shown), as we reported previously for AQP-expressing CHO, glial and proximal tubule cell types (8
, 21
, 23)
. AQP1-expressing B16F10 cells showed consistently greater lamellipodial area per leading edge surface (lamellipodial width) than control cells (Fig. 3D
).
|
Increased extravasation and metastasis of AQP1-expressing tumor cells in vivo
Tumor cells were labeled with fixable fluorescent dyes in order to identify them after i.v. injection in mice. Cells were stained with the green fluorescent dye CMFDA or the red fluorescent dye CMRA, each of which is cell permeable and, within cells, becomes entrapped by covalent reaction with cytoplasmic proteins. Initial studies were done to establish labeling conditions to give bright, stably labeled cells without effect on migration and to establish a ratio procedure to measure at the same time the migration of control and AQP1-expressing cells. As mentioned above (Fig. 2A
), the fluorescence labeling did not affect cell migration in vitro. The labeled cells remained fluorescent, with easily distinguishable red vs. green color for >24 h after labeling.
The two-color cell labeling strategy for simultaneous detection of control and AQP1-expressing cells was tested using the in vitro transwell assay in which a mixture of CMRA-labeled control 4T1 cells (red) and CMFDA-labeled AQP1-expressing 4T1 cells (green) was added to the upper chamber. Figure 4
A (left panel) shows the easily recognizable red and green adherent cells at 6 h after cell addition. More green than red cells remained after scraping the upper surface of the filter (middle panel). From counting many cells on multiple filters, the ratio of AQP1-expressing vs. control cells was significantly increased after scraping cells (Fig. 4A
, right), in agreement with the data above showing increased migration of AQP1-expressing cells.
|
The same approach was used in mice to study tumor cell extravasation, with CMRA labeling of control and CMFDA labeling of AQP1-expressing cells, and with the labeling reversed (CMFDA labeling of control and CMRA labeling of AQP1-expressing cells). Figure 4B
shows a low magnification fluorescence micrograph of the cell suspension used for tail vein injection, with approximately equal numbers of red and green fluorescent cells. The two lower photographs show labeled tumor cells in lung at 6 h after tail vein injection, with more AQP1-expressing than control tumor cells for both labeling schemes. The data are summarized in Fig. 4C
as cell count ratios and total number of cells per low power microscope field for control and AQP1-expressing 4T1 and B16F10 cells. Migration of AQP1-expressing cells was greater than that of control cells for both labeling schemes (orange bars). In control studies, lungs harvested at 10 min (instead of 6 h) showed relatively few fluorescent cells (purple bars), most representing adherent cells to lung endothelium that have not yet migrated. The cell count ratio (purple bars) in lung at 10 min was not significantly different from that in the cell suspension used for tail vein injection (open bars). When a mixture of red- and green-labeled control cells was used for tail vein injections (right panels) the cell count ratio at 6 h was similar to that at 10 min and in the original cell suspension, as expected. These data provide evidence for increased lung extravasation of AQP1-expressing tumor cells after tail vein injection.
Tumor cell metastasis and growth were also studied in vivo. In one set of experiments lung metastasis was evaluated 15 days after tail vein injection of control or AQP1-expressing 4T1 cells. Figure 5
A (left) shows a greater number of well-demarcated tumor metastases in lungs of mice injected with AQP1-expressing tumor cells. Also, in most mice receiving the AQP1-expressing cells there was evidence for tumor infiltration of alveolar walls (middle). Immunocytochemistry in Fig. 5A
(right) shows AQP1 expression in alveoli (microvascular endothelia), as expected, with AQP1 expression in the 4T1-AQP1 cells but not in control 4T1 cells. The data summary in Fig. 5B
shows that AQP1-expressing tumor cells produced a remarkably greater number of lung metastases, but without altered tumor colony size. Alveolar wall thickness within 50 µm of metastases was significantly greater for the AQP1-expressing tumor cells due to tumor cell invasion.
|
In another set of experiments mice were implanted subcutaneously with control or AQP1-expressing 4T1 or B16F10 cells to study tumor growth and local invasion. Figure 5C
shows that tumor growth, as assessed by tumor volume at different times after implantation, was not affected by AQP1 expression. However, finger-like projections into subcutaneous adipose tissue were often seen with AQP1-expressing but not control 4T1 cells (Fig. 5D
). Both AQP1-expressing and control B16F10 tumors were well encapsulated, without evidence of local invasion.
DISCUSSION
As mentioned above, the expression of various AQPs has been found in high grade tumors of different tissue origins. Tumor cell AQP expression has been correlated with metastatic potential, and in some instances AQPs are expressed in tumor cells whose tissue of origin does not normally express AQPs (10
, 12
, 14)
. The possible involvement of tumor cell AQPs in water movement between tumor parenchyma and microvessels has been proposed, as well as in tumor cell volume regulation (24
, 25)
. However, the substantially slower rates of tumor mass hydration and cell volume regulation compared to cell osmotic water equilibration, which generally occurs in seconds, make it unlikely that the water permeability of tumor cells can be rate-limiting for their growth or volume homeostasis. The data here support a novel role for AQPs in tumor cells in their migration, which occurs in metastasis during tumor cell invasion across tissue barriers such as blood vessels. One consequence of AQP involvement in tumor cell migration is the possibility of AQP inhibition to limit tumor spread, though testing of this possibility will require development of suitable AQP-selective inhibitors.
In vitro and in vivo evidence supports AQP-facilitated tumor cell migration and extravasation across blood vessels. The migration of two tumor cell lines was significantly enhanced after transfection with AQP1, as assessed by both transwell and wound healing assays. AQP1 transfection did not alter tumor cell growth, proliferation, morphology, size, or substrate adherence but, as expected increased their water permeability. The in vitro data agree with the increased migration of AQP-transfected fibroblasts and epithelial cells, and the reduced migration of AQP1-deficient endothelial cells (8)
and proximal tubule epithelial cells (21)
and AQP4-deficient astroglial cells (23)
. The in vivo studies in mice showed a greater number of AQP1-transfected vs. control tumor cells in lung parenchyma after tail vein infusion, producing a remarkably greater number of lung metastases. The extravasation measurements were done by a ratiometric method in which mice were injected with an 1:1 mixture of control and AQP1-expressing tumor cells that were labeled with different color fluorescent dyes. This ratio counting procedure reduced data variance by controlling for intrinsic mouse-to-mouse differences, as well as variability related to the tail vein injection procedure and tissue processing.
Cell migration involves a series of cellular events including actin polymerization and depolymerization, and the generation of cell protrusions such as lamella, lamellipodia, and blebs (26)
. We proposed that AQP-facilitated water permeability in cell protrusions enhances their formation and thus the rate of cell migration (2
, 8)
. Various AQP-expressing cells, including the tumor cells studied here, show AQP polarization to lamellipodia and an increased number/size of lamellipodia (8
, 21)
. Local actin depolymerization, which creates new osmoles, might provide the osmotic driving force responsible for water influx into cell protrusions, as might transmembrane solute transport and/or altered solute osmotic coefficients in submembrane cytoplasm (26
, 27)
. Thus, it is predicted that cell membrane water permeability at its migrating surface may be an important determinant of tumor cell migration, which would depend on membrane lipid composition, the presence and membrane polarization of AQPs, and the biophysical properties of submembrane cytoplasm. Of these factors, AQP-dependent water permeability would be the most amenable to control by use of pharmacological inhibitors.
Tumor metastasis involves a series of complex host-tumor interactions (28)
. Tumor cell extravasation into surrounding tissue has been considered a rate-limiting step in metastasis (29
, 30)
that involves the active migration of tumor cells across the endothelial barrier and penetration through the underlying basement membrane. The interaction of tumor cells with the basement membrane has been thought of in terms of three steps: cell attachment, matrix dissolution, and migration (29
, 31)
. Tumor cell binding to the basement membrane surface involves tumor cell surface receptors of integrin and nonintegrin types that recognize basement membrane glycoproteins such as laminin, collagen, and fibronectin. Locomotion propels tumor cells across the basement membrane and stroma through the zone of matrix dissolution. The formation of pseudopodial protrusions at the leading edge of migrating cells is the earliest step in locomotion. We found here that AQP1-expressing tumor cells have increased metastatic potential and local invasiveness compared to control cells, but similar adherence to various basement membrane substrates and similar growth rates. The in vitro studies provided evidence for increased cell migration as the mechanism for the enhanced metastatic potential and local invasiveness of AQP1-expressing tumor cells, which was related to their increased formation of lamellapodial extensions. Thus, manipulation of the expression or function of tumor AQPs is predicted to alter their migration and metastatic potential.
In summary, we found a novel role for AQP1 in tumor cell extravasation and spread, which may provide a functional explanation for the expression of AQPs in many tumor types. Tumor cell water permeability and AQP expression may thus be important determinants of their migration and spread in vivo, suggesting AQPs as targets in tumor therapy.
ACKNOWLEDGMENTS
We thank Dr. Eugene Solenov for help in water permeability measurements, Dr. Tonghui Ma for help in cell transfections, and Liman Qian for mouse breeding. This work was supported by grants DK35124, EY13574, DK72517, EB00415, HL59198, and HL73856 from the National Institutes of Health, and a Research Development Program grant from the Cystic Fibrosis Foundation.
Received for publication February 7, 2006. Accepted for publication April 10, 2006.
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rom atomic structure to clinical medicine. J. Physiol. 542,3-16This article has been cited by other articles:
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