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Vacuolar-type H⁺-ATPases at the plasma membrane regulate pH and cell migration in microvascular endothelial cells

Departments of ¹Physiology and ²Internal Medicine, Texas Tech University Health Sciences Center, Lubbock; ³Department of Medical Physiology, Texas A & M University System Health Science Center, Temple; and ⁴Department of Animal Science, Texas A & M University, College Station, Texas; ⁵Department of Ornamental Horticulture, ARO Volcani Center, Bet-Dagan, Israel; and ⁶Children's Memorial Research Center, Northwestern University Feinberg School of Medicine and Robert H. Lurie Comprehensive Cancer Center, Chicago, Illinois

Submitted 14 February 2006 ; accepted in final form 24 March 2006

	ABSTRACT

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Microvascular endothelial cells involved in angiogenesis are exposed to an acidic environment that is not conducive for growth and survival. These cells must exhibit a dynamic intracellular (cytosolic) pH (pH_cyt) regulatory mechanism to cope with acidosis, in addition to the ubiquitous Na⁺/H⁺ exchanger and HCO₃^–-based H⁺-transporting systems. We hypothesize that the presence of plasmalemmal vacuolar-type proton ATPases (pmV-ATPases) allows microvascular endothelial cells to better cope with this acidic environment and that pmV-ATPases are required for cell migration. This study indicates that microvascular endothelial cells, which are more migratory than macrovascular endothelial cells, express pmV-ATPases. Spectral imaging microscopy indicates a more alkaline pH_cyt at the leading than at the lagging edge of microvascular endothelial cells. Treatment of microvascular endothelial cells with V-ATPase inhibitors decreases the proton fluxes via pmV-ATPases and cell migration. These data suggest that pmV-ATPases are essential for pH_cyt regulation and cell migration in microvascular endothelial cells.

fluorescence spectroscopy; carboxyseminaphthorhodafluor-1; bafilomycin; sodium/hydrogen exchanger; buffering capacity; proton fluxes; migration; macrovascular endothelial cells

Changes in intracellular (cytosolic) pH (pH_cyt) are important in signal transduction mechanisms, which regulate many physiological processes, including cell growth, secretion, contraction, and invasion/migration (18, 44, 47). These processes are important in angiogenesis and vascular remodeling (13). The regulation of pH_cyt in most eukaryotic cells, including endothelial cells, is mediated by the Na⁺/H⁺ exchanger and HCO₃^–-dependent H⁺-transporting mechanisms (11, 17, 22, 68). In some specialized and highly invasive cells (metastatic cells, macrophages, neutrophils, and osteoclasts), plasma membrane vacuolar (V) H⁺-ATPases (pmV-ATPases) are also used to regulate pH_cyt (27, 35, 50, 58). These ATPases are distinguished from other proton pumps by their pharmacological inhibition (8, 33). The V-ATPases are inhibited by bafilomycin A₁, concanamycin, salicylihalamide, and 7-chloro-4-dinitrobenz-2-oxa-1,3-diazole, which have no effect on the P- or F-type ATPases (6, 9, 52, 64, 65).

Microvascular endothelial cells, similar to tumor cells, are exposed to hypoxic and acidic environments (31, 60), which are not favorable for growth and cell survival. We have shown that pmV-ATPase expression in highly invasive metastatic tumor cells provides a dynamic pH_cyt regulatory mechanism for these cells (27, 51). The similarity between metastatic cells and angiogenic microvascular endothelial cells with regard to invasion of adjacent tissue by the invading cell led us to hypothesize that 1) microvascular, but not macrovascular, endothelial cells express pmV-ATPase as a dynamic pH_cyt regulatory mechanism, which allows them to cope with acidic environments, 2) microvascular endothelial cells employ this pump's activity for cell migration, and 3) the presence of pmV-ATPases at the leading edge in microvascular endothelial cells allows them to maintain a more alkaline pH_cyt at the leading than at the lagging edge, thus creating a pH_cyt gradient favorable for cell migration.

	MATERIALS AND METHODS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
Media, buffers, and chemicals. Dulbecco's modified Eagle's medium (DMEM) was supplemented with 10% or 20% fetal bovine serum (FBS), 2 mM L-glutamine, 5 mM D-glucose, 20 U/ml heparin, 1 mM sodium pyruvate, 100 U/ml penicillin, 100 µg/ml streptomycin, and 0.2 µg/ml amphotericin B (GIBCO, Grand Island, NY). Joklik's essential medium (essentially Ca²⁺-free) was supplemented with 60 mM taurine, 20 mM creatine, and 5 mM HEPES. Cell perfusion buffer (CPB) consisted of (in mM) 110 NaCl, 1 MgSO₄, 5.4 KCl, 1.5 CaCl₂, 0.44 KH₂PO₄, 0.35 NaH₂PO₄, 5 glucose, 2 L-glutamine, and 25 HEPES at the indicated pH. Na⁺-free CPB consisted of all CPB ingredients, except those containing Na⁺ (110 mM N-methylglucamine was substituted for NaCl). CPB solutions containing HCO₃^– were continuously bubbled with 5% CO₂ at 37°C. The concentration of HCO₃^– in the buffer was determined as described elsewhere (17). High-K⁺ buffer contained (in mM) 146 KCl, 20 NaCl, 5 glucose, 2 glutamine, 10 HEPES, 10 MES, and 10 bicine. The rationale for using these organic buffers was to allow for precise buffering across a wide pH range (5.5–8.0) (27).

Bafilomycin A₁ was obtained from Wako Chemicals (Richmond, VA), and SCH-28080 was a generous gift of Dr. A. Barnett (Schering, Bloomfield, NJ). The fluorescent dyes were obtained from Molecular Probes (Eugene, OR). All other chemicals were obtained from Sigma Chemical (St. Louis, MO), unless otherwise stated.

Isolation of micro- and macrovascular endothelial cells. Microvascular (cardiac) and macrovascular (aortic) endothelial cells were isolated from normal BioBreeding (BB) rats (Biomedical Research Models, Worcester, MA) with use of techniques previously described, with some modifications (63). Briefly, for isolation of microvascular endothelial cells, heart ventricles were minced with fine surgical scissors and digested with Liberase Blendzyme 2 [Roche, Indianapolis, IN; 0.2 mg/ml in HEPES-buffered DMEM (H-DMEM)] at 37°C for 60 min using a shaker for continuous agitation of the digestion solution. For isolation of macrovascular endothelial cells, aortic segments (dissected free of any attached connective tissue) were cut longitudinally and subjected to the digestion procedure used for the minced ventricles. Undigested tissue fragments were removed by filtration through 100-µm nylon mesh cell strainers (BD Biosciences, Bedford, MA). The resulting cell suspension was pelleted by centrifugation, rinsed twice, and resuspended in H-DMEM. Biotinylated mouse anti-rat CD31 (PECAM) antibody (clone 3A12, Research Diagnostics, Flanders, NJ) was added to a concentration of 5 µg/ml, and the suspension was incubated for 60 min at room temperature on a rotator. The cells were again pelleted by centrifugation, rinsed, and resuspended in 1 ml of H-DMEM. Streptavidin-coated magnetic beads (10 µl; Dynabeads M-280, Invitrogen, Carlsbad, CA) were added, and the suspension was incubated for 45 min at room temperature on the rotator. For selection of endothelial cells, the Eppendorf tube containing the cell suspension was placed in a magnetic stand (magnetic particle concentrator, Invitrogen), and magnetic beads attached to endothelial cells were pulled to the side of the tube (adjacent to the magnet). Nonendothelial cells that remained in suspension were removed by aspiration. Beads were resuspended in H-DMEM, and the procedure was repeated twice to ensure removal of all contaminating nonendothelial cells. Endothelial cells were resuspended in growth medium and plated on gelatin-coated tissue culture dishes for cell growth expansion. Micro- and macrovascular endothelial cells from three to four rats were pooled into one 60-mm gelatin-coated (1.5% gelatin in PBS) petri dish and cultured at 37°C under 10% CO₂ in DMEM with 20% FBS. Endothelial cell identity was confirmed as described elsewhere (63). Micro- and macrovascular endothelial cells were passaged by trypsinization and subsequently grown at extracellular pH (pH_ex) 7.4 in DMEM supplemented with HCO₃^– and 10% FBS.

Measurement of pH_cyt in cell populations. pH_cyt was determined by the fluorescence of carboxyseminaphthorhodafluor-1 (5-[and 6]carboxy-SNARF-1) as described previously (27). Briefly, two coverslips containing cells at confluency were loaded with 7.5 µM SNARF-1 in its acetoxymethyl ester (AM) form and incubated at 37°C in 5% CO₂ for 45 min and then in buffer for 30 min to ensure complete ester hydrolysis/leakage of uncleaved dye. The coverslips were placed in a holder-perfusion device, and the fluorescence of SNARF-1 was monitored with a spectrofluorometer (model SLM-8100/DMX) equipped for sample perfusion at 37°C. SNARF-1 was excited at 534 nm, with emission at 584 and 644 nm. The ratio of fluorescence at 644 nm to fluorescence at 584 nm was used to monitor pH changes. Fluorescence data were converted to ASCII format for subsequent data analysis in SigmaPlot (version 8.0, Jandel Scientific, San Rafael, CA).

In situ calibration of SNARF-1. In situ calibration curves were generated as described previously (27). Briefly, cells attached to coverslips were perfused with high-K⁺ buffers at pH_ex 5.5–8.0 (at 0.2-pH unit intervals). The buffers contained 2 µM valinomycin and 6.8 µM nigericin to collapse the pH gradient. The ratio of SNARF-1 fluorescence at 644 nm to SNARF-1 fluorescence at 584 nm at each pH_ex was fitted to the following equation

(1)

where R_obs is the fluorescence ratio at any given pH, R_min is the fluorescence ratio when the dye is fully protonated, R_max is the fluorescence ratio when the dye is fully unprotonated, and pK_a is the apparent acid dissociation constant. Equation 1 is solved iteratively using nonlinear least squares analysis and yields pK_a, R_min, and R_max values for SNARF-1 in these cells. From these in situ calibration curves, the following parameters were obtained for SNARF-1 in microvascular endothelial cells (n = 33): pK_a = 7.76 (SD 0.076), R_min = 0.55 (SD 0.004), and R_max = 2.49 (SD 0.23). The in situ calibration parameters for SNARF-1 in macrovascular endothelial cells (n = 33) were as follows: pK_a = 7.69 (SD 0.089), R_min = 0.435 (SD 0.007), and R_max = 2.94 (SD 0.28). These values were significantly different between cell types (P < 0.05). pH_cyt values were obtained for each experiment by using Eq. 1 and their corresponding in situ calibration parameters with SigmaPlot.

Measurement of pH_cyt in discrete cellular regions with use of spectral imaging microscopy. Spectral imaging microscopy allows measurements of ions in discrete subcellular regions of single cells with high temporal, spectral, and spatial resolution (29, 49). The spectral imaging microscope is based on a Spectra-Pro-150 spectrograph directly coupled to the side port of an inverted microscope (model IX70, Olympus). The spectrograph has 300 groves per nanometer of grating, is blazed at 500 nm (Acton Research, Acton, MA), and is equipped with a high-dynamic-range frame transfer back-illuminated charge-coupled device (CCD) camera (model Spec10B, Princeton Instruments, Trenton, NJ) controlled by an ST133 controller (Princeton Instruments). The CCD camera has a 1,340 x 512 pixel imaging array (pixel = 9 x 9 µm). The entrance of the slit spectrograph was set at 0.2 mm throughout the experiments, except for the zero-order spectra, where the slit was set at 2.0 mm. The spectrograph and the CCD camera settings were computer controlled using commercially available software (Winspec/32 version 2.5.10.1 [EC] , Roper Scientific, Trenton, NJ). The CCD camera temperature was maintained at –100°C for all the experiments. The full spectral output of the cell can be obtained within as little as 2 ms and with 0.4-nm spectral resolution. The spatial information was obtained by alignment of a single cell along the length of the entrance slit, so that spectra were acquired from unique subcellular locations (i.e., leading edge to lagging edge). Data were collected from 15 discrete regions of interest of the cell and binned to obtain a higher signal-to-noise ratio. The optical filters were as follows: 510-nm-narrow band-pass filter and 550-nm-long band-pass dichroic filter (Omega Optical, Brattleboro, VT).

Immunocytochemistry. Macro- and microvascular endothelial cells were fixed with 4% paraformaldehyde for 15 min, washed with 25 mM glycine, and then permeabilized with 0.1% Triton X-100. The cells were sequentially incubated with primary antibody specific for the E subunit of V-H⁺-ATPase (46). Cells were washed extensively and then labeled with Alexa Fluor 568 secondary (anti-mouse IgG) antibody and Alexa Fluor 488-phalloidin, which binds to F-actin and helps delineate the cell edge (51). The cells were mounted in VectaMount solution (Vector Laboratories, Burlingame, CA) and maintained at 4°C overnight. The cells were observed with a confocal laser scanning microscope (model LSM 510 META, Zeiss) with a x63 objective (Plan-APOCHROMAT, 1.4 NA, oil differential interference contrast). Simultaneously acquired images of Alexa Fluor 488-phalloidin (actin cytoskeleton, green) and Alexa Fluor 568 (V-ATPase, red) fluorescence were collected, and each section was analyzed on a pixel-by-pixel basis utilizing Physiology software (version 3.0, Zeiss) to assess colocalization of actin and V-ATPase.

Cell migration/invasion assay. Microvascular, but not macrovascular, endothelial cells are involved in new blood vessel formation, which requires these cells to invade and migrate through extracellular matrix (ECM) proteins (20, 30, 51). To determine whether microvascular endothelial cells are more migratory and invasive than macrovascular endothelial cells, cells grown at confluence in T-25 flasks in DMEM were loaded with 5 µM calcein-AM for 30 min. The cells were then trypsinized, washed, and counted. To evaluate the degree of cell invasion through various ECMs in vitro, HTS FluoroBlok (Becton Dickinson, Bridgeport, NJ) inserts were briefly soaked in Matrigel, seeded at densities of 5 x 10⁴ cells/well, and incubated at 37°C in 5% CO₂ for 24 h. HTS FluoroBlok inserts contain a 3-µm polyethylene terephthalate membrane impregnated with dyes that absorb visible light at 490–700 nm. To evaluate cell migration, we used this approach, except the filters were not coated with Matrigel. This allows us to study the ability of the cell to deform to allow it to migrate through the filter pores. The inserts were subsequently visualized, and images of the bottom and top of the insert were obtained with a x20 objective (UPlan Fl 0.5 Ph1, Olympus) and a confocal microscope (model 1024 MRC, Bio-Rad, Hercules, CA). Calcein was excited with the 488-nm line of a 50-mW krypton-argon laser, and emission was collected using the VHS filter (Bio-Rad) blocks, which contain a 515-nm emission filter. Experiments were done in triplicate, and five images were obtained per HTS FluoroBlok. The images were subsequently analyzed, and the cells were visually counted in defined areas. Percent invasion/migration was corrected for proliferation and calculated as follows

(2)

We also evaluated cell migration using the wounded monolayer model in micro- and macrovascular endothelial cells (48, 51), which allows us to study the ability of cells to migrate to close the wound. The cells were grown on 12-mm coverslips to confluency and subsequently wounded with a micromanipulator to induce a 300-µm gap (51). The cells were allowed to close the wound for up to 24 h in the absence or presence of bafilomycin to inhibit V-ATPase. At selected times, the cells were fixed, permeabilized, and incubated with FITC-phalloidin. Images of wounded monolayers were then obtained with a x20 objective and a Bio-Rad confocal microscope (excitation at 488, emission at 515 nm). Migration was assessed as wound distance at selected times from three randomly selected areas.

Assay of cell-doubling times. Because differences in cell migration/invasion may be due to differences in cell growth between micro- and macrovascular endothelial cells, we evaluated cell growth. Cells were plated onto 24-well plates (Falcon, Becton Dickinson) at an initial density of 2 x 10⁴ cells/well in triplicates. After 12 h, the cells were fixed with 1% glutaraldehyde to obtain values at time 0. Thereafter, the cells were fixed at 24-h intervals for up to 120 h. At the end of the experiment, the cells were stained with 0.1% crystal violet for 20 min, destained with running water for 5 min, and air-dried. The absorbance at 590 nm is linearly related to the number of cells; thus cell number can be estimated to obtain the kinetics of cell growth (16, 51). The data were fitted to the following sigmoid (3-parameter) equation to obtain the cell-doubling times using SigmaPlot software

(3)

From these experiments, we determined that the doubling time in microvascular endothelial cells (n = 4) was 27.44 h (SD 2.24), whereas the doubling time in macrovascular endothelial cells (n = 4) was 40.45 h (SD 6.99). Thus doubling times were significantly faster in microvascular than in macrovascular endothelial cells (P < 0.05).

Statistical analysis. Data were analyzed by nonparametric and parametric tests and ANOVA (SigmaStat 2.03, Jandel Scientific, Richmond, CA) as appropriate. Statistical significance was assigned at P < 0.05.

	RESULTS

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Migration/invasion is greater in microvascular than in macrovascular endothelial cells. The degree of migration/invasion of cells through an artificial basement membrane matrix is significantly higher in microvascular than in macrovascular endothelial cells (Fig. 1, A and B). These experiments allowed us to study the ability of the cells to degrade ECM proteins and undergo dynamic morphological changes that would enable them to traverse the filter's pores. We also performed wounding monolayer experiments to quantify the extent of cell migration after a wound. A wounded monolayer labeled with FITC-phalloidin at time 0 and 18 h after wound closure is shown in Fig. 1, C and D. From such experiments, we determined the extent of wound closure (i.e., migration) as a function of wound distance. We observed significantly more complete wound closure at 24 h in microvascular than in macrovascular endothelial cells (n = 3; Fig. 1E). Treatment of wounded monolayers with 20 nM bafilomycin A₁ resulted in a significant inhibition of wound closure in microvascular, but not macrovascular, endothelial cells (Fig. 1, E and F). These data indicate that V-ATPases are important in the migration of microvascular endothelial cells.

View larger version (22K): [in this window] [in a new window]   Fig. 1. A: invasion of micro- and macrovascular endothelial cells. Cells (5 x 104) were plated onto extracellular matrix protein-coated polycarbonate 3-µm-pore filters (Osmonics, Minnetonka, MN) in a modified Boyden chamber and incubated for 24 h at 37°C. Invading cells were removed with 2 mM EDTA-PBS, fixed, stained, and visually counted. Values are means ± SE (n = 4). *P < 0.05 vs. microvascular. B: migration of micro- and macrovascular endothelial cells. Cells were handled as described in A, except filters were not coated. Cell migration was determined as described in A. Values are means ± SE (n = 4). *P < 0.05 vs. microvascular. C and D: wounded monolayer model study of migration in micro- and macrovascular endothelial cells. Cells were grown on coverslips; at confluency, a micromanipulator was used to produce a 300-µm wound in cell monolayer. Cells were fixed, permeabilized, and stained with FITC-phalloidin at time 0 (C) and after 18 h (D). Confocal microscopy images were obtained with a x20 objective, and migration was analyzed on 3 different fields per coverslip. E and F: quantification of migration of wounded monolayers of micro- and macrovascular endothelial cells without and with bafilomycin (Baf). Data were derived from experiments similar to those described in C and D. Values are means ± SE (n = 5). *P < 0.05 vs. Baf.

View larger version (71K): [in this window] [in a new window]   Fig. 2. Immunocytochemical localization of vacuolar (V)-ATPase in microvascular (A–C) and macrovascular (D–F) endothelial cells. Cells were grown on glass coverslips; at confluency, cells were wounded as described in Fig 1C. After 12–16 h, cells were fixed and permeabilized. A and D: cytoskeleton labeled with Alexa Fluor 488-phalloidin. B and E: subunit of V-ATPase (red). C and F: sequential images of Alexa Fluor 488-phalloidin (green) and Alexa Fluor 568 (red) fluorescence obtained with a laser scanning confocal microscope (Ar/HeNe lasers at 30 and 1 mW, respectively, with excitation at 488 and 543 nm and emission at 515 and 585 nm) using a x60 objective. Merge images of red (V-ATPase) and green (actin) were assessed on a pixel-by-pixel basis. Lamellipodia are clearly defined at wound site (long arrows). Plasma membrane V-ATPase (pmV-ATPase) is conspicuous at the leading edge of microvascular endothelial cells (long white arrows). V-ATPase is inconspicuous at the lagging edge of the same cell (short arrows, B and C); cells behind the wound do not show apparent V-ATPase at their plasma membranes (short arrows).

Acid-loading experiments allow identification of Na⁺- and HCO₃^–-independent pH_cyt recovery. To study the differences in the mechanisms of pH_cyt regulation between micro- and macrovascular endothelial cells, we selected conditions in which the two main pH_cyt regulatory mechanisms should be dormant. Thus acid-loading experiments utilizing the NH₄Cl prepulse technique (47) were used to evaluate the characteristics of pH_cyt recovery in the absence of Na⁺ and HCO₃^–. Our expectations were that this experimental strategy could reveal a novel Na⁺- and HCO₃^–-independent pH_cyt regulatory mechanism. Cells loaded with SNARF-1 were perfused with CPB until steady-state pH_cyt was achieved. Addition of 25 mM NH₄Cl caused a rapid intracellular alkalinization (Fig. 3A), whereas acute removal of NH₄Cl reversed the condition and caused rapid acidification in micro- and macrovascular endothelial cells. In the absence of Na⁺ and HCO₃^–, macrovascular endothelial cells (n = 11) did not recover from acidification [dpH/dt = 0.001 (SD 0.005), proton flux (J) = 0.01 mM H⁺/min (SD 0.1)], but microvascular endothelial cells did recover from this acid load (Fig. 3). The H⁺ buffering capacity (_i) was significantly higher in microvascular than in macrovascular endothelial cells (n = 11): _i = 36.7 (SD 1.08) vs. 30.9 (SD 1.37) mM (P < 0.05). To determine whether pH_cyt recovery in microvascular endothelial cells was mediated by V-H⁺-ATPases, we examined pH_cyt recovery from acid loads in an Na⁺- and HCO₃^–-free buffer in the presence of bafilomycin to inhibit V-H⁺-ATPase and found a significant decrease in J_H⁺ (Fig. 3B). P-type H⁺-ATPase inhibitors such as SCH-28080 had no effect on J(Fig. 3B). To determine whether Na⁺/H⁺ exchange and HCO₃^–-based H⁺ transport contributed to pH_cyt regulation in microvascular endothelial cells, we performed acid-loading experiments in the presence of Na⁺ and HCO₃^– and found J values similar to those observed in medium with Na⁺ and without HCO₃^– (cf. Fig. 3B). In the presence of Na⁺ and HCO₃^–, macrovascular endothelial cells also recovered from an acid load [J= 1.44 (SD 0.34) mM H⁺/min (n = 5)]. These J values are similar to those observed in microvascular endothelial cells. Collectively, these data indicate ubiquitous Na⁺- and HCO₃^–-dependent pH_cyt regulatory mechanisms in micro- and macrovascular endothelial cells. Importantly, microvascular, but not macrovascular, endothelial cells exhibited an additional Na⁺- and HCO₃^–-independent pH_cyt regulatory system that improved their ability to cope with acid loads (cf. Fig. 3, A and B). To further demonstrate that neither Na⁺/H⁺ exchanger nor HCO₃^–-based H⁺-transporting mechanisms were involved in the pH_cyt recoveries, we performed experiments in the absence of Na⁺ and HCO₃^– with 5-(N,N-hexamethylene)-amiloride and DIDS, blockers of Na⁺/H⁺ exchanger and anion transport, respectively (Fig. 3B). Neither 5-(N,N-hexamethylene)-amiloride (not shown) nor DIDS significantly altered the kinetics of pH_cyt recovery in an Na⁺- and HCO₃^–-free buffer. These data indicate the presence of an Na⁺- and HCO₃^–-independent pH_cyt regulatory mechanism in microvascular endothelial cells that allows them to recover from acid loads; this mechanism is absent in macrovascular endothelial cells.

View larger version (14K): [in this window] [in a new window]   Fig. 3. A: microvascular, but not macrovascular, endothelial cells exhibit a Na+- and HCO3–-independent cytosolic pH (pHcyt) regulatory system. Cells were grown on glass coverslips to confluency, intracellularly loaded with carboxyseminaphthorhodafluor-1 (SNARF-1)-AM, and transferred to a spectrofluorometer for pHcyt measurements. Cells were superfused with cell perfusion buffer (CPB) until steady-state pHcyt was reached. At 1st arrow, superfusate was exchanged with 25 mM NH4Cl. At 2nd arrow, superfusate was exchanged for Na+- and HCO3–-free CPB. Data are representative of 34 and 11 experiments on micro- and macrovascular endothelial cells, respectively. B: effect of inhibitors of primary and secondary H+ transport systems on proton flux (J) in microvascular endothelial cells. Cells were handled as described in A. At the time indicated by 2nd arrow in A, superfusate was exchanged with Na+- and HCO3–-free CPB containing 50 nM bafilomycin (n = 6), 100 µM DIDS (n = 3), or 5 µM SCH-28080 (n = 5). Recovery of pHcyt (J) after acid loading were determined during the first 3 min from experiments similar to those described in A. J was obtained by multiplying dpH/dt in the first 5 min of recovery by apparent intrinsic buffering capacity (i) as described elsewhere (17). Values are means ± SE. *P < 0.05 vs. Na+- and HCO3–-free medium.

View larger version (9K): [in this window] [in a new window]   Fig. 4. A: effect of acute Na+ removal in micro- and macrovascular endothelial cells. Cells were handled as described in Fig. 3A, except at the arrow, perfusate was changed to Na+-free CPB. Data are representative of 11 and 6 experiments for micro- and macrovascular endothelial cells, respectively. Microvascular, but not macrovascular, endothelial cells recover from Na+-induced acidification. B: effect of inhibitors on J in microvascular endothelial cells. Recovery of pHcyt after removal of Na+ (J) was estimated during the first 3 min from experiments described in A. Values are means ± SE of 3 experiments for 100 µM DIDS and 5 experiments for 50 nM bafilomycin. *P < 0.05 compared with Na+-free medium.

View larger version (36K): [in this window] [in a new window]   Fig. 5. Spectral imaging microscopy study of pHcyt in leading and lagging edges of the cell. Microvascular endothelial cells were grown on glass coverslips to confluency and loaded with SNARF-1. A: a single cell was aligned onto the entrance slit (2.0 mm) of the spectrograph, allowing identification of its leading and lagging edges. Inset: selected cell, where leading domain is the wounded side and lagging edge is the rear of the cell (B). B and C: slit width decreased to 0.5 mm (B) and then to 0.2 mm (C) to allow for increased spatial resolution and increased signal-to-noise ratio. D: emission filters were removed, and fluorescence spectra were collected and deconvoluted from individual regions of interest (ROI). E: emission spectra were collected and recorded from 2 distinct ROI in the cell (Lead and Lag). F: spectral pHcyt changes of SNARF-1 [arbitrary fluorescence units (auf)] from a single ROI. G: recovery from an acid load in Na+- and HCO3–-free CPB. Spectra similar to those in F were collected from 12 different ROI at 50-ms sampling rates. At 1st arrow, cells were superfused with CPB-25 mM NH4Cl; at 2nd arrow, superfusate was exchanged for Na+- and HCO3–-free CPB.

View larger version (23K): [in this window] [in a new window]   Fig. 6. In situ calibration of 12 individual ROI from single cells. Microvascular endothelial cells were handled as described in Fig. 5. A: at the end of the experiment, in situ titrations were performed. Full SNARF-1 spectra were collected at 12 different regions of the cell from leading to lagging edge, and ratios were plotted. pHex, extracellular pH. Data from only 6 ROI were fit to Eq. 1, which was solved iteratively using nonlinear least squares analysis, and apparent acid dissociation constant (pKa, B) and fluorescence ratios when dye is fully protonated and unprotonated (Rmin and Rmax, respectively, C) were determined and plotted as a function of distance from leading to lagging edge.

	DISCUSSION

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

Because migration and invasion through the ECM are important elements in angiogenesis, we evaluated whether microvascular endothelial cells could penetrate an artificial basement membrane more readily than macrovascular endothelial cells. Our data indicated that microvascular endothelial cells exhibiting pmV-ATPase were more migratory/invasive than macrovascular endothelial cells, which do not exhibit pmV-ATPase. Because vascular remodeling after a lesion involves migration of endothelial cells from the noninjured area to repair the lesion, we employed a wounded monolayer experiment. In this experiment, scraping off a 300-µm region in a confluent monolayer of endothelial cells results in cell migration toward the damaged region to repair the wound (48). This orderly movement of cells adjacent to the lesion occurs immediately and reveals an easily distinguishable leading edge. When healing was allowed to continue, the wound was closed in 24 h. Importantly, bafilomycin treatment significantly decreased wound closure in microvascular endothelial cells. These data indicate that a bafilomycin-sensitive component is involved in migration of microvascular endothelial cells.

Because a faster rate of wound closure could be explained by distinct doubling times between micro- and macrovascular endothelial cells, we evaluated this parameter. Our data indicated that doubling time in microvascular endothelial cells is 27 h, whereas doubling time in macrovascular endothelial cells is 40 h. Wound closure in 24 h in microvascular endothelial cells and doubling times >24 h in micro- and macrovascular endothelial cells support our contention that the different rates of wound closure are not due to differences in cell growth between micro- and macrovascular endothelial cells but, rather, to a distinct difference in migratory rate between these cell types.

The data reported by Bai et al. (3) suggest that wound closure is faster in microvascular than in macrovascular endothelial cells. Bai et al. estimated that human microvascular endothelial cells migrate at 15 µm/h, whereas human umbilical vein endothelial cells (HUVEC) migrate at 10 µm/h under nonstimulating conditions. There are, however, controversies regarding rates of cell migration. Specifically, Morales et al. (34) reported 27% wound closure within 24 h in wounded monolayers of HUVEC. In this case, the wound size was 2.5 mm. However, faster migration rates in a wounded monolayer of HUVEC and human coronary aortic endothelial cells, where wound closure is 40% and 30%, respectively, within 6 h have also been reported (1). The wound size in these other studies was 150–200 µm. Thus it appears that a major reason for the different rate of migration is wound size, because larger wounds are associated with slower migration rates, possibly because of the release of chemoattractants from cells that work in a paracrine fashion stimulating cell migration. It is possible that distinct differences in rates of wound closure between micro- and macrovascular endothelial cells may be due to distinct sensitivity of microvascular endothelial cells to chemokines (59). Microvascular endothelial cells from lung and kidney produce more chemokines, such as fractalkine, interleukin-1, tumor necrosis factor-, and interferon-, than macrovascular endothelial cells (HUVEC) (5). Human dermal microvascular endothelial cells are more sensitive to cytokines, such as oncostatin and IL-6, basic fibroblast growth factor, and IL-1, than HUVEC (59). Significant cell division as a mechanism of wound healing is not likely to occur in cells before 15–24 h (1, 25), because endothelial cells exhibit slower doubling times. Thus our study extends previous observations by indicating that expression of pmV-ATPase at the leading edge in microvascular endothelial cells is a mechanism that explains the faster rates of migration in microvascular than in macrovascular endothelial cells.

Our observations that bafilomycin decreases the rate of cell migration are in agreement with a recent study in which higher concentrations of bafilomycin (100–500 nM) than those used in our study suppressed cell motility in NIH 3T3 A31 mouse fibroblasts (57). The authors hypothesized that the effect of bafilomycin on cell motility was due to alterations of pH gradients in endocytic structures, which are known to exhibit V-ATPase. Recently, endosome fusion to the plasma membrane has been suggested as an important mechanism for wound healing in fibroblasts (45). Furthermore, overexpression of the 16-kDa subunit of V-ATPase in 10T1/2 fibroblasts has been shown to enhance invasion and the secretion of matrix metalloproteinase-2, an enzyme needed for protein degradation during invasion (23). Although the subcellular location of the overexpressed 16-kDa subunit was not evaluated in that study, these data indicate that overexpression of V-ATPase may be important for invasion. Our immunocytochemical data show that V-ATPase colocalizes with actin filaments at the cell's cortex and at the leading edge. This is consistent with previous studies that have indicated that V-ATPase colocalizes with actin at the cell's cortex in the ruffled border of activated osteoclasts (21, 24) and in the apical region of the middle gut epithelium of Manduca (62). In human breast cancer cells, we recently showed that pmV-ATPase expression is important for migration/invasion of highly metastatic human breast cancer cells (51). Therefore, our study complements these observations to indicate that pmV-ATPase is important for migration in microvascular endothelial cells.

Regulation of pH_cyt in most cells is accomplished by the relative contribution of Na⁺/H⁺ exchanger and HCO₃^–-based H⁺-transporting mechanisms (18, 44, 47). Microvascular endothelial cells are not the exception, because they exhibited Na⁺/H⁺ exchanger and HCO₃^–-based H⁺-transporting mechanisms. In addition to these important pH_cyt regulatory mechanisms, pmV-ATPases are also used to regulate pH_cyt in microvascular, but not macrovascular, endothelial cells. Furthermore, use of wounded monolayer experiments to reveal the leading edge of migrating cells indicate that pmV-ATPase is present at the leading edge. As a result, cells exhibit a more alkaline pH_cyt at the leading than at the lagging edge. Differences in pH_cyt regulation at the leading and lagging edges are predicted by flux ratio equations, because the passive H⁺ influx is 45 and 56 times the passive efflux at the leading edge and lagging edges, respectively (if it is assumed that membrane potential is –90 mV and pH_cyt values in Fig. 5G are used for leading and lagging edges at pH_ex 7.4). This suggests that H⁺ influx is larger at the lagging edge, consistent with a more dynamic pH_cyt regulatory system at the leading edge. Further support for a dynamic mechanism to maintain such pH_cyt differences in leading and lagging edges is based on the fact that although H⁺ permeability is extremely high (10^–3 cm/s), the actual J across the plasma membrane is very low because of the low free H⁺ concentration in the cytosol and in the extracellular environment (if we assume pH_ex 7.4). Under these conditions, the passive H⁺ influx is 0.02 pH unit/h, yet the observed difference in pH_cyt between the leading and the lagging edge is 0.2 pH unit within the time frame of our experiments (i.e., 5–20 ms). Thus it is unlikely that such differences in pH_cyt values from the leading to the lagging edge are due to simple H⁺ diffusion. We interpret these data to suggest that pmV-ATPase at the leading edge is a dominant pH_cyt regulatory system that allows these pH_cyt gradients to exist in microvascular endothelial cells.

The variation in the steady-state deprotonated-to-protonated SNARF-1 ratio may be due to actual pH_cyt differences, variations in regional cytoplasmic microviscosity (28, 43, 55, 66), or even a different proportion of dye bound to cytoplasmic proteins (4). To properly interpret the differences in SNARF-1 protonated-to-deprotonated ratios, we have taken into account the behavior of the pH fluoroprobe in the cytoplasm, because it is heterogeneous in terms of composition and organization. Regional intracellular microenvironments may differ in viscosity, which in turn could result in distinct behavior of the fluoroprobes (39). Indeed, it is known that ion-sensitive fluoroprobes may display spectral differences not only between in vitro and in situ environments (7, 42), but also within the distinctive intracellular organelles (2, 12, 40, 56). Viscosity values for the leading edge (3.8 mPa·s), lagging edge (0.5 mPa·s), and soma (0.5 mPa·s) of locomoting neutrophils have been documented (66). Thus regional differences in cytoplasmic viscosity and/or the interactions between cytoplasmic proteins and the fluorescent dye may contribute to regional variations in the ratio and the in situ calibration parameters (, R_max, and R_min) used to estimate pH_cyt.

Important effects of protein on the in vitro calibration parameters have been described for several ion indicators, including pH fluoroprobes (4, 41, 54). However, it has also been suggested that SNARF-1 does not bind to bovine serum albumin but, rather, that a contaminant present in the commercially available SNARF-1 binds to bovine serum albumin (67). Furthermore, in the cytoplasm of cardiac myocytes, a major fraction of the fluoroprobes (0.5–0.9) appears to be bound to proteins (4). Other variables that could cause variations in the in situ titration parameters include partition of the dye between cytoplasm and endomembranous compartments (37, 61), the amount of dye bound to proteins, quenching agents, and inner filter phenomena (7, 53). The reasons for the distinct in situ calibration parameters in different cellular domains are not immediately apparent. However, differences in the cytoplasmic microenvironment in terms of protein composition and viscosity could cause distinct diffusion mobility of fluorescent probes and distinct spectral properties. Indeed, studies by fluorescence recovery after photobleaching have shown that the translational diffusion of intracellular 2',7'-bis(2-carboxyethyl)-5(6)-carboxyfluorescein near the membrane and in the bulk cytoplasm is 6–10 times and 4 times lower than in water, respectively (55). However, the fluid-phase cytoplasmic viscosity in the absence of collisions or binding to cytoplasmic macromolecules is similar to the viscosity of water (26, 55). There are also important differences in molecular crowding within the different subcellular compartments, suggesting considerable diffusional heterogeneity for small metabolites and, thereby, fluoroprobes within different intracellular organelles (14). In addition, viscosity can alter the spectra of ion indicators (28, 43, 55).

To compensate for differences in the cytoplasmic microenvironment in terms of viscosity and protein composition, we used specific regional calibration parameters (pK_a, R_max, and R_min) to convert subdomain fluorescence ratios to pH_cyt. These data indicate that the heterogeneities in regional pH_cyt values are associated with physiological pH_cyt differences, where the leading edge exhibits a more alkaline pH_cyt than the lagging edge.

In conclusion, our data indicate that pmV-ATPase expression in microvascular endothelial cells is relevant for pH_cyt regulation and migration. pmV-ATPases have also been found in highly invasive tumors (51). Thus pmV-ATPase has physiological significance and could provide a target for pharmacological intervention in angiogenesis and cancer.

	GRANTS

TOP ABSTRACT MATERIALS AND METHODS RESULTS DISCUSSION GRANTS REFERENCES

 
This study was supported by American Heart Association (National) Grants 9750558N and 0555070Y, American Cancer Society Grant RPG-00-035-01CNE, National Heart, Lung, and Blood Institute Grant HLB-RO1-HL-65695, and Texas Higher Education Coordinating Board Advanced Research Program Grant 010674-0012-2001 (to R. Martínez-Zaguilán), American Heart Association (National) Grant 0630207N (to S. R. Sennoune), and National Cancer Institute Grant RO1 CA-59702 (to M. J. C. Hendrix).

	FOOTNOTES

 

Address for reprint requests and other correspondence: R. Martínez-Zaguilán, Dept. of Physiology, Texas Tech Univ. Health Sciences Center, 3601 4th St., Lubbock, TX 79430-6551 (e-mail: Raul.MartinezZaguilan{at}ttuhsc.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.

^* J. D. Rojas and S. R. Sennoune contributed equally to this work.

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