Plocabulin, a novel tubulin-binding agent, inhibits angiogenesis by modulation of microtubule dynamics in endothelial cells

High concentration rat tail Collagen I was obtained from BD Biosciences (San José, CA, USA). Phorbol 12-myristate 13-acetate (PMA), Hoechst 33,342, 3-(4,5-dimethylthiazol-2-yl)-2,5-diphenyltetrazolium bromide (MTT), mouse monoclonal anti-α-tubulin (T5168), sulforhodamine B (SRB), and phalloidin-FITC (P5282) were obtained from Sigma (St Louis, MO, USA). Human recombinant Fibroblast Growth Factor-basic (FGF) was from Peprotech (Rocky Hill, NJ, USA). Alexa 594-conjugated goat anti-mouse IgG secondary antibodies (A11032) were obtained from Molecular Probes (Rockford, IL, USA). For in vitro experiments, plocabulin (PharmaMar, Madrid, Spain) was prepared as a 1 mg/ml stock solution in DMSO and stored at − 80 °C. For in vivo experiments, lyophilized vials (1.6 mg/vial) of plocabulin (PharmaMar) were used. For xenograft studies, potential antiangiogenic effect induced by the treatment was studied by Angiosense TM 680EX Fluorescent Imaging Agent (Perkin Elmer Inc., MA, USA).

Human Umbilical Vein Endothelial Cells (HUVECs) were obtained from Lonza (Basel, Switzerland) and grown in endothelial basal medium (EGM-2) supplemented with growth supplements: 2% Fetal Bovine Serum (FBS), human Epidermal Growth Factor (hEGF), Vascular Endothelial Growth Factor (VEGF), R3-Insulin-like Growth Factor-1 (R3-IGF-1), ascorbic acid, hydrocortisone human Fibroblast Growth Factor-Beta (hFGF-β), heparin, gentamicin/amphotericin-B (GA). Human microvascular endothelial cells (HMEC-1) were obtained from ATCC (Manassas, Virginia, USA) (CRL-3243) and grown in MCDB131 (without L-Glutamine) medium supplemented with hEGF (10 ng/ml), hydrocortisone (1 μg/ml), glutamine (10 mM) and 10% FBS. Only low passage cells (between passages 3 and 7) were used. For microtubule dynamics, EB3-GFP plasmid was nucleofected using Amaxa technologies with the HUVEC nucleofector kit (Lonza) according to the manufacturers’ protocols. For xenograft experiments, we used 2 human derived cell lines: NCI-H460 non-small cell lung carcinoma (HTB-177™) and MDA-MB-231 breast adenocarcinoma (CRM-HTB-26™), both from ATCC (Manassas). Before animal inoculation, cells were maintained in vitro at 37 °C with 5% CO2 in Dulbecco’s Modified Eagle’s Medium (Sigma-Aldrich) and passaged every 3 to 5 days upon reaching confluence.

Endothelial cells were treated with plocabulin at different concentrations for 6 h, 24 h and 48 h, fixed with methanol for 10 min at − 20 °C and incubated with a blocking solution (5% bovine serum albumin in PBS) for 30 min. Cells were then incubated with primary mouse anti-human α-tubulin antibody for 1 h at room temperature. After three washes with a PBS/BSA1% solution, cells were incubated with Alexa 594-conjugated goat anti-mouse IgG secondary antibody at room temperature for 1 h. Cells were then incubated with phalloidin-FITC for 1 h at 37 °C in a humid chamber. Cells were finally counterstained with addition of Hoechst 33,342 (1 μg/ml) for 5 min and mounted with Mowiol mounting medium. Pictures were taken with a Leica DM IRM fluorescence microscope equipped with a 100× oil immersion objective and a DFC 340 FX digital camera (Leica, Wetzlar, Germany).

The relative expression profile of 55 human angiogenesis-related proteins was performed using Proteome Profiler Human Angiogenesis Array kit (R&D Systems, NE, Minneapolis, U.S.A) after treatment of HUVEC endothelial cells with plocabulin for 24 h.

The MTT colorimetric assay was used for quantitative measurement of cell viability, as previously described [26]. The highest plocabulin concentration in the assay was 17 nM and then, serial dilutions 1/2.5 from this initial concentration were added to the cells.

For adhesion assays, HUVEC cells (75,000 cells/well) were cultured in 96-well plates covered with fibronectin (2.5 μg/ml) and collagen 0.05 μg/ml at 4 °C for 10 min. Adhesion was then allowed at 37 °C during 30 min in the absence or presence of plocabulin. After 3 washes with PBS, remaining cells were fixed with glutaraldehyde (0.1%) and stained with sulforhodamine B. As positive and negative controls, we have used MnCl₂ (0.5 mM), latrunculin A (3 μM) and cytocalasin D (2 μM), respectively. For migration and invasion assays, 6.5 mm-diameter transwell chambers (Sigma-Aldrich, St. Louis, MO, USA) with polycarbonate membrane (pore size 8.0 μm) were used. HUVEC cells (1.5 × 105) were seeded on the upper compartment of the transwell membrane in 100 μl of serum-free culture medium containing or not plocabulin. The lower compartment (well) was filled with 600 μl of serum-free culture medium containing or not a chemoattractant (FBS 2%). For the invasion assays, the upper compartment of the transwell was previously coated with 12 μg of matrigel basement membrane matrix to create a physical barrier between the two compartments of the chamber. After 24 h of incubation, culture medium was removed from the inside of the chamber, and the non-migrated cells on the upper side of the membrane were wiped off using a cotton swab. Migrated cells on the lower side of the membrane were fixed with a glutaraldehyde 1% solution, washed and stained with sulforhodamine B following standard techniques.

The tube formation or destabilization assays are based on the ability of endothelial cells to form three-dimensional capillary-like tubular structures when cultured on a basement membrane matrix. HUVEC cells (2.5 × 104 cells) were serum-starved overnight, resuspended in 100 μl of medium supplemented with FBS 2% per well and plated on 96-well plates previously coated with 50 μl of Matrigel (BD Biosciences). For tube formation assays, fresh culture medium containing FBS 2% alone or FBS 2% plus plocabulin was added after 1 or 6 h of incubation at 37 °C. For tube destabilization assays, fresh culture medium containing FBS 2% alone or FBS 2% plus plocabulin was added when capillary tube structures were well established. In both cases, after 6 h and 24 h post-treatment, cells were stained with calcein-AM to measure cell viability and analyze tube formation. Cultures were observed and photographed by phase-contrast and fluorescence microscopy.

Endothelial cell spheroids were generated overnight by culturing endothelial cells in complete medium containing 20% methylcellulose in non-adherent 96-wells plates. Harvested spheroids were then embedded into 2 mg/ml collagen gels overlaid with complete medium supplemented with 40 ng/ml FGF, 50 ng/ml PMA and with the indicated drug concentration. Angiogenic activity was quantified by measuring the cumulative length of the sprouts that had grown out of each spheroid, their mean number and length, 24 h after embedding using ImageJ software. For each condition, at least 20 spheroids were analyzed. Comparisons between different samples were analyzed by Student’s t test. Differences were considered significant at *P < 0.05, **P < 0.01 and ***P < 0.001.

All animal protocols were reviewed and approved according to regional Institutional Animal Care and Use Committees. Design, randomization and monitoring of experiments (including body weights and tumor measurements) were performed using NewLab Software v2.25.06.00 (NewLab Oncology, Vandoeuvre-Lès Nancy, France). Female athymic Nude-Foxn-1 nu/nu mice (Envigo, RMS Spain S.L.) between 4 to 6 weeks of age were s.c. xenografted with NCI-H460 or MDA-MB-231 cancer cells into their flank with ca. 5 × 106 cells or 7.5 × 10,6 respectively. In the NCI-H460 experiment, when tumors reached ca. 150 mm³, mice were intravenously administered in three consecutive weekly doses (0.08, 8 and 16 mg/kg/day) whereas the control animals received an equal volume of vehicle with the same schedule. Caliper measurements of the tumor diameters were made three times a week and tumor volumes were calculated according to the following formula: (axb2) /2, where a and b were the longest and shortest diameters respectively. Animals were humanely sacrificed when their tumors reached 2500 mm³ or if significant toxicity (e.g. severe body weight reduction) was observed. Differences in tumor volumes between treated and control group were evaluated using the Mann–Whitney U-test. Statistical analyses were performed by Graph Pad Prism® v5.03 (Graph Pad Software Inc. La Jolla, CA, USA). In addition, three randomly-selected H460 tumor-bearing animals were dedicated to study blood vessel density and to characterize vascular changes related with plocabulin administration by an in vivo imaging system, IVIS^® Spectrum (Perkin Elmer Inc., Waltham, MA, USA). When tumors reached ca. 300 mm³, animals (n = 3/group) were treated with a single dose of either placebo, or plocabulin (at 2 or 16 mg/kg) and then, they were administered via tail vein injection with AngioSense 680 EX, which is a near-infrared labeled fluorescent macromolecule that remains localized in the vasculature for extended periods of time and enables imaging of blood vessels and angiogenesis. Fluorescent signal acquisition was performed at 24 h after the treatment and AngioSense dosing. In the MDA-MB-231 experiment, six tumor bearing mice, ca. 500 mm³ were randomly selected 24 h after being administered with plocabulin at 16 mg/kg or placebo. The animals were sacrified by CO₂ asphyxiation whithin their home cages, being CO₂ flow set to displace 20% of the cage volume per minute. Death was confirmed by physical examination. Tumors were then dissected free and processed for paraffin embedding and sectioning, serial sections were cut and stained with hematoxylin/ eosin for histology evaluation.

Tubulin cytoskeleton in untreated HUVECs cells is characterized by cytoplasmic microtubules radiating from a central point (microtubule organizing center) to the cell periphery (Fig. 1, red). Treatment of HUVEC cells with different concentrations of plocabulin resulted in depolymerization of the microtubule network (representative images are shown in Fig. 1a). At low concentrations (0.01 nM), these effects were observed after 48 h of incubation with the drug while, at higher levels (0.1–1 nM), they were observed as soon as after 6 h of treatment (Additional file 1: Figure S1A). Similar results were obtained with immortalized human HMEC-1 cells (Additional file 1: Figure S1B). Of note, in these cells, the effects of plocabulin 0.01 nM were observed even after only 6 h of drug treatment. We have also evaluated the effects on the morphology and the actin cytoskeleton of plocabulin on HUVEC cultures (Fig. 1b). Differently from untreated cells, plocabulin-treated cells showed a rounded morphology with well-developed actin filament structure similar to the “dense peripheral bands” observed in mature endothelial cells [28, 29]. Moreover, plocabulin also induced endothelial cell retraction and rounding and plasma membrane blebbing (Fig. 1b). All these effects were dependent on concentration and time of incubation: the higher the concentration, the faster they appeared. Again, these effects were observed at similar levels after treatment of HMEC-1 cells with the drug (Additional file 1: Figure S1). We have finally analyzed the expression profiles of 55 angiogenesis-related proteins in supernatants of HUVEC cultured cells in the absence or presence of plocabulin 0.05 nM for 24 h using a membrane-based sandwich immunoassay. The analyses showed that none of these proteins were altered in HUVEC cells after treatment (data not shown).

To test the effect of plocabulin on microtubule dynamics in living endothelial cells, we ectopically expressed EB3-GFP using Amaxa nucleofector technology in HUVEC cells. Twenty-four hours after transfection, endothelial cells were treated with different concentrations of plocabulin for one hour. Treatment with DMSO was used as a non-treated condition. Microtubule plus-end dynamics was then live-recorded using spinning-disk confocal microscopy. As shown in Fig. 2a, with the maximum intensity projection of EB3-GFP signal during 2 min acquisition, microtubules were detected in the presence of plocabulin concentrations ranging from 0.01 nM to 0.1 nM, whereas addition of 1 nM induced an almost complete suppression of GFP-labeled microtubule plus-end signals. Plocabulin-treated cells showing remaining microtubules were then analyzed for microtubule dynamics. Kymographs were drawn for at least 100 MTs in each condition and subsequently analyzed (Fig. 2b and c and Additional file 2: Table S1). Plocabulin treatment significantly reduced the velocity and the covered distance of growth events in a concentration-dependent manner. Addition of this drug also caused a strong increase in catastrophe frequency.

HUVEC cells were left to adhere to an extracellular matrix composed of fibronectin and type I collagen for 10 min and were then exposed for a short period (30 min) to plocabulin ranging from 1 pM to 10 nM. No detachment of cells from the extracellular matrix was observed (data not shown). We then evaluated the in vitro effect of plocabulin on endothelial cell migration and invasion using transwell chambers and HUVEC primary cells. As shown in Fig. 3a, HUVEC cells showed very low basal levels of migration in serum-free medium, but migration was greatly increased in the presence of FBS 2% added to the lower chamber as chemo-attractant. HUVEC cells were then cultured in the presence of chemo-attractant and treated with different concentrations (0.01, 0.1, 1 and 10 nM) of plocabulin. As shown in Fig. 3a, the drug inhibited the migration of endothelial cells in a concentration-dependent manner. Concentrations higher than 0.1 nM resulted in a nearly complete abrogation of cell migration. For invasion experiments, the porous membrane of transwell chambers was pre-coated with a layer of Matrigel mimicking the physiological basement membrane. The results obtained were similar to those described for the migration assays (Fig. 3b). Plocabulin completely inhibited cell invasion at concentrations higher than 0.1 nM. To discard that the inhibition of cell migration and invasion exerted by plocabulin were due to a direct cytotoxic activity, cell survival of HUVEC cells was analyzed in concentration-response curves using a standard MTT method. As shown in Fig. 3c, at the effective concentrations used in the migration and invasion assays, plocabulin was not cytotoxic against HUVEC cells in a 24-h assay. With plocabulin, cells retained nearly 100% viability at 1 nM (concentration that corresponded to a complete inhibition of cell migration and invasion). Thus, the inhibitory concentrations of plocabulin on HUVEC migration and invasion were not coincident with those for inhibition of cell proliferation, indicating that the aforementioned effects were not likely mediated through unspecific cytotoxicity of the drug.

To analyze how plocabulin affects the formation of new capillaries, HUVEC cells were grown on Matrigel in the presence or absence of the compound. Untreated, control HUVEC cells rapidly formed a network of angio-tube like structures that were visualized by fluorescence microscopy after staining the cells with calcein-AM (Fig. 4). Plocabulin interfered with the correct formation of the HUVEC capillary network at concentrations as low as 0.1 nM (Fig. 4a). At higher concentrations, the intercellular connections were mostly absent with drastic effects on the formation of the capillary network. Having established that plocabulin interferes with the formation of capillary networks from HUVEC cells when cultured on Matrigel, we wondered if the compound was also affecting an already established capillary network. At the concentration of 0.1 nM some effects on the capillary network could be observed (Fig. 4b). Treatment with plocabulin 1 nM or higher concentrations resulted in a significant disruption of the formed capillary-like network, with the majority of cells forming small clumps with a few number of intercellular connections. A complete dissapearance of the network was observed at concentrations higher than of 1 nM; in this case, cells appeared retracted and distinct cords were no longer observed. Altogether, these results indicate that plocabulin interfere with the formation and stability of the capillary networks formed by endothelial cells when cultured on a matrigel basement membrane matrix.

We finally performed a 3D sprouting assay in the presence of increasing concentrations of plocabulin. Twenty-four hours after embedding in collagen gel, non-treated spheroids of endothelial cells had the ability to produce and to extend capillary-like sprouts (Fig. 5). Addition of plocabulin affected the global angiogenic activity of endothelial cells in a concentration-dependent manner, as quantified by the mean cumulative sprout length per spheroid (Fig. 5a). This effect was the consequence of a reduced number as well as the reduced length of sprouts, indicating that plocabulin impeded both the formation of primary sprouts and their extension or stabilization (Fig. 5a). Treatment of pre-established angiogenic sprouts with the same drug concentrations lead to their collapse (data not shown). To determine if these effects were mediated by the antimitotic properties of the compound, we added thymidine in the medium to block cell division. Control spheroids treated with thymindine still actively produced long sprouts (Fig. 5b). Interestingly, treatment with plocabulin severely affected sprouting activities compared to the non-treated spheroids, precluding a mitosis effect of the compounds at the concentrations used.

We then performed xenograft studies to test whether the in vitro antiangiogenic activity of PM060184 was also involved in its in vivo antitumor activity. NCI-H460 lung tumor cells were xenografted into the right flank of athymic nu/nu mice. Once the tumors reached ca. 150 mm³, the mice were randonmly assigned into groups of 10 mice each and either vehicle or plocabulin (0.08, 8 and 16 mg/kg/day) was intravenously administered for 3 consecutive weeks. At the drug doses used in the experiment, no significant toxicity or body weight loss was observed in the treated animals (data not shown). As shown in Fig. 6a, the highest doses of plocabulin presented antitumor activity with a statistically significant inhibition (p ≤ 0.0001 vs placebo) of tumor growth as well as, a rapid, extensive and irreversible hemorrhagic necrosis 4 days after the first dose (Fig. 6b). Complete tumor regressions were also observed after the administration of plocabulin at 16 mg/kg:Out of 10 mice, 1, 2, 3, 7 and 7 animals experienced complete tumor remissions on days 5, 7, 10, 14 and 21, respectively (Fig. 6c). We then studied if the observed effects on tumors were in part due to a reduction in functional vascular volume induced by the drug. For this purpose, randomly selected mice bearing H-460 xenografts were treated with a single dose of placebo, 2 or 16 mg/kg of plocabulin (n = 3/group). As shown in Fig. 6d, a strong and dose-dependent decreases in their vasculature as assessed by the percentage of reductionin the intratumoral fluorescence of plocabulin-treated animals (ca. 65 and 45% compared to placebo for 2 and 16 mg/kg, respectively) after the administration of Angiosense™ 680. Finally, the antiangiogenic activity of plocabulin was further evaluated in nude mice bearing MDA-MB-231 breast cancer xenografts (n = 6). Again, in this tumor xenograft, the administration of a single dose of plocabulin (16 mg/kg) induced a very strong reduction in the number of vessels after 24 h of treatment (Additional file 3: Figure S2). Altogether, these results confirmed the antiangiogenic activity of plocabulin.