BRCA1 regulates microtubule dynamics and taxane-induced apoptotic cell signaling

Abstract

The taxanes are effective microtubule-stabilizing chemotherapy drugs used in the treatment of various solid tumors. However, the emergence of drug resistance hampers their clinical efficacy. The molecular basis of clinical taxane resistance remains poorly understood. Breast cancer 1, early onset gene, BRCA1, is a tumor-suppressor gene, whose expression has been correlated with taxane sensitivity in many solid tumors including non-small cell lung cancer. However, the molecular mechanism underlying the relationship between BRCA1 (B1) expression and taxane activity remains unclear. To this end, we created a stable B1 knockdown A549 cell line (B1-KD) to investigate B1’s role in microtubule biology and response to taxane treatment. We show that B1-KD rendered A549 cells resistant to paclitaxel (PTX), phenocopying clinical studies showing that low B1 expression correlated with taxane resistance. As previously reported, we show that loss of B1 enhanced centrosomal γ-tubulin localization and microtubule nucleation. Interestingly, we found that the B1-KD cells exhibited increased microtubule dynamics as compared with parental A549 cells, as assessed by live-cell confocal microscopy using enhanced green fluorescent protein-tagged α-tubulin or EB1 protein. In addition, we showed that loss of B1 impairs the ability of PTX to induce microtubule polymerization using immunofluorescence microscopy and a cell-based tubulin polymerization assay. Furthermore, B1-KD cells exhibited significantly lower intracellular binding of a fluorescently labeled PTX to microtubules. Recent studies have shown that PTX-stabilized microtubules serves as a scaffold for pro-caspase-8 binding and induction of apoptosis downstream of induced-proximity activation of caspase-8. Here we show that loss of B1 reduces the association of pro-caspase-8 with microtubules and subsequently leads to impaired PTX-induced activation of apoptosis. Taken together, our data show that B1 regulates indirectly endogenous microtubule dynamics and stability while its loss leads to microtubules that are more dynamic and less susceptible to PTX-induced stabilization conferring taxane resistance.

INTRODUCTION

The microtubule is arguably one of the most important targets for cancer chemotherapy owing to the clinical success of microtubule-targeting drugs such as the taxanes and vinca alkaloids.¹ In solid tumors, the taxanes are among the most effective chemotherapeutic agents routinely used in the treatment of a wide range of tumors including breast cancer,^2–7 non-small cell lung cancer^8,9 and prostate cancer.^10–13 Despite the efficacy of the taxanes, many tumors display either inherent or acquired resistance to them, hampering their clinical activity. The molecular basis underpinning the taxanes’ clinical resistance has not been fully elucidated.

Interestingly, in clinical studies, loss of function and/or low expression of the breast cancer 1, early onset gene (BRCA1) have been associated with resistance to taxane and sensitivity to DNA-damaging-based chemotherapies (for example, cisplatin, anthracycline) in a variety of tumor types.^14–18 BRCA1 is a tumor-suppressor gene with well-defined nuclear functions that include DNA damage response and repair, as well as cell cycle checkpoint activation.¹⁹ The central role of BRCA1 in DNA repair can readily explain why its loss leads to enhanced cisplatin sensitivity.²⁰ However, the molecular basis of taxane resistance conferred by loss of BRCA1, or the taxane sensitization observed on restoration of wild-type BRCA1²¹ is not well understood.

Recently, cytoplasmic functions of BRCA1 have been identified including focal adhesion and centrosome regulation.^22,23 BRCA1 was shown to inhibit centrosomal microtubule nucleation in a cell cycle-dependent manner by ubiquitinating gamma(γ)-tubulin and preventing its centrosomal localization.^24,25 Beyond these data, little is known about any part BRCA1 has in the regulation of microtubule biology and, especially, the microtubule response to taxane treatment.

Taxanes bind beta (β)-tubulin and stabilize microtubules leading to mitotic arrest and cell death. Therefore, the taxanes have been classically thought to exert their antitumor activity by inhibiting mitosis. However, recent evidence suggests that microtubule-stabilizing drugs have alternative mechanisms of action that lead to cell death, primarily through signaling pathways occurring during interphase.^26,27 One such pathway is the activation of caspase-8 following taxane-induced microtubule stabilization. Mielgo et al.²⁸ showed that paclitaxel (PTX)-stabilized microtubules served as a scaffold for the binding of the death-effector domain of pro-caspase-8. Thus, the microtubule polymers provided a platform for sufficient spatial proximity of pro-caspase-8 molecules, enabling activation of the downstream caspase-8 proteolytic cascade, via induced-proximity activation, which is the main mechanism of caspase-8 activation. Although this report provided a mechanistic explanation for interphase-induced cell death following PTX treatment, the potential role BRCA1 has in modulating this pathway affecting taxane sensitivity and resistance remains to be elucidated.

To better understand the mechanism of BRCA1-mediated taxane resistance, we set out to investigate the effects of BRCA1 on microtubule biology and its potential relationship with taxane activity. To this end, we have generated an isogenic pair of cell lines with and without BRCA1 protein expression. We have found that BRCA1 knockdown (B1-KD) in A549 lung cancer cells confers resistance to PTX and sensitivity to cisplatin treatment, recapitulating the reported clinical phenotypes. We show that BRCA1 regulates microtubule dynamics and stability and that by modulating the microtubule cytoskeletal network, BRCA1 controls how microtubules respond to PTX treatment and, consequently, alter drug-induced microtubule-based activation of caspase-8.

RESULTS

Loss of BRCA1 in A549 cells mimics clinical taxane resistance

To better understand the role of BRCA1 in taxane sensitivity and resistance, we have generated an isogenic pair of A549 non-small cell lung cancer cell lines consisting of the parental cell line with functional wild-type BRCA1 and a derivative cell line with stable BRCA1 knockdown (B1-KD) following lentiviral infection with empty and BRCA1-specific short hairpin RNAs (shRNAs) and subsequent selection of stable clones using puromycin (Figure 1). Several clones were isolated and expanded for the empty and BRCA1-specific shRNAs. The BRCA1-specific shRNA clone with the highest BRCA1 knockdown by reverse transcriptase–PCR and western blot analyses, namely B1-KD, was selected for further characterization, and the empty shRNA clone, henceforth named A549, with no change in BRCA1 mRNA and protein levels was used as the control (Supplementary Figure S1a). Initial characterization of B1-KD cells revealed a 70% decrease in BRCA1 mRNA expression as assessed by quantitative PCR and a concomitant decrease in BRCA1 protein expression (Figures 1a and b). To ensure that the loss of BRCA1 protein was associated with loss of function, we assessed the extent of γ-tubulin ubiquitination and localization as well as centrosome amplification, as they represent hallmarks of BRCA1 cytoplasmic function.^24,25 Recent reports identified that the BRCA1/BARD1 complex serves as the E3-ubiquitin ligase for γ-tubulin and that ubiquitination of γ-tubulin results in the displacement of the latter from the centrosome. Thus, BRCA1 loss of function is associated with centrosome amplification via accumulation of γ-tubulin at the centrosome. Indeed, our results revealed significantly higher amounts of γ-tubulin localized at the centrosome in B1-KD cells, as compared with the A549 cells (Figure 1c). In agreement with the established role of BRCA1 in γ-tubulin ubiquitination, our results showed that immunoprecipitation of γ-tubulin from B1-KD cells was associated with a loss of γ-tubulin mono-ubiquitination in contrast to the results obtained using the A549 cell line (Supplementary Figure S1b, arrow points to mono-ubiquitinated form of γ-tubulin). In addition, immunofluorescence staining with pericentrin (used as a marker of the centrosome) revealed that B1-KD cells had an increased number of cells with amplified centrosomes (that is, >2 centrosomes per cell), as compared with the A549 cells (P<0.01), in agreement with published reports (Figure 1d). Next, to determine whether the loss of BRCA1 had an impact on the cells’ sensitivity to PTX and cisplatin, we performed a 30-h cytotoxicity assay, which revealed that B1-KD cells were threefold more resistant to PTX (PTX IC₅₀ = 30 and 10 nM for B1-KD and A549, respectively) and fourfold more sensitive to cisplatin (CDDP IC₅₀ = 0.5 μM and 2 μM for B1-KD and A549, respectively, as compared with the A549 cells; Figure 1e). The doubling times for the A549 and B1-KD cells were 25 and 31 h, respectively. A second B1-KD clone with similar BRCA1 protein knockdown also showed PTX resistance in the 30-h cytotoxicity assay (Supplementary Figure S2). To confirm BRCA1 knockdown renders cells PTX-resistant was not limited to A549 cells, transient B1-KD with BRCA1-specific shRNA in HOP62 cells rendered them PTX-resistant compared with empty controls, although to a lesser extent than in the A549 model (Supplementary Figure S3) because of the <30% transfection efficiency observed. Similar results were obtained in a 72-h cytotoxicity assay using sulforhodamine B (data not shown). In a soft agar colony formation assay, the B1-KD cells grew into well-formed, larger size colonies in the presence of PTX while the A549 cells did not (Supplementary Figure S4). Taken together, these data suggest that the B1-KD cells we have generated exhibited loss of function of BRCA1 and phenocopy clinical taxane resistance and platinum sensitivity.

Figure 1.

Characterization of B1-KD in A549 cells. (a) Relative BRCA1 mRNA expression levels in A549 and B1-KD cells determined by real-time quantitative PCR (qPCR; P<0.01). Total RNA was extracted from A549 and B1-KD cells and subjected to real-time qPCR using primers specific for BRCA1 or actin. (a) relative quantification of BRCA1 and actin mRNA levels was calculated using a comparative Ct method. BRCA1 data were normalized to actin. Results are displayed as percentage of control (A549 cells). (b) Immunoblot of BRCA1 protein expression in A549 and B1-KD cells. Actin is used as a loading control. In all, 50 μg of total cell lysates from A549 and B1-KD cells were resolved by sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) and immunostained using an antibody against BRCA1 (C-20) and actin as a loading control. (c) Localization of γ-tubulin in A549 and B1-KD cells was assessed by γ-tubulin immunofluorescence. One representative cell is shown per condition, centrosomes are labeled by γ-tubulin staining. Scale bar = 1 μm. Bar graph shows extent of γ-tubulin staining as measured by diameter of each centrosome for 30 cells per condition. P<0.001. (d) Left panel: A549 and B1-KD cells were immunostained for pericentrin to visualize the centrosomes and counterstained with 4,6-diamidino-2-phenylindole (DAPI). Representative confocal microscopy images are shown. Arrows, indicate amplified centrosomes. White scale bar = 10 μm. Right panel: percentage of cells with more than two centrosomes was determined in A549 and B1-KD cells (P<0.01). (e) Drug sensitivity assay in A549 (black line) and B1-KD cells (red line) treated with a range of different PTX or cisplatin concentrations for 30 h. Cell viability was measured using the CellTiter-Glo assay and plotted as % survival.

BRCA1 modulates microtubule dynamics

As the microtubule is the cellular target of PTX, we hypothesized that loss of BRCA1 protein renders cells more PTX-resistant because of alterations of the microtubule cytoskeleton. The role of BRCA1 on centrosome function^24,25 prompted us to investigate centrosomal microtubule nucleation in A549 and B1-KD cells. In each cell line, microtubules were completely depolymerized using nocodazole, and allowed to regrow after nocodazole was washed out from the cells. Our results revealed that the B1-KD cells displayed an increased microtubule regrowth from the centrosome following nocodazole washout as compared with the A549 cells. Quantitative analysis of the average length of individual microtubules revealed a significant increase in centrosomal microtubule regrowth in the B1-KD cells compared with A549 cells (Figure 2a, P <0.0001). Similar results were seen in another B1-KD clone as well in the HOP62 cell line (Supplementary Figure S5). The steady-state length of individual microtubules is determined by their intrinsic dynamic instability reflected by the balance between the rates of tubulin dimer addition and removal from microtubule plus-ends.^1,29 To examine the behavior of microtubule plus-end dynamics as a function of BRCA1 expression we assessed the velocity of the microtubule plus-end-associated protein EB1 in A549 and B1-KD cells, using live-cell confocal microscopy. EB1 tracking is utilized as a marker of microtubule dynamics and it binds to the plus-end of growing microtubules while it dissociates from microtubules during phases of pause or shortening.³⁰ Therefore, EB1 tracking is considered a highly sensitive and reliable surrogate marker of microtubule dynamics. We transfected A549 and B1-KD cells with a plasmid encoding enhanced green fluorescent protein-tagged EB1 and assessed microtubule growth dynamics using live-cell spinning disk confocal microscopy. Our data involving analyses of approximately 30 representative cells per condition revealed that the EB1 comet velocity was significantly increased in B1-KD cells at a rate of 20 μm/min as compared with 10 μm/min observed in the A549 line (Figure 2b). Although these results indicated enhanced microtubule growth rates in B1-KD cells, the other properties of microtubule dynamics such as shortening rates or time spent at pause could not be determined because EB1 only associates with the growing microtubule.³¹ To assess if BRCA1 regulated other parameters of microtubule dynamicity, we monitored microtubule dynamics in both cell lines, by tracking the tips of enhanced green fluorescent protein (EGFP)-labeled microtubules using live-cell confocal microscopy. Our results showed that loss of BRCA1 led to an increase in both the rates of microtubule polymerization (14 vs 18 μm/min), corroborating the EB1 comet velocity data, as well as depolymerization (15 vs 22 μm/min) in A549 and B1-KD cells, respectively. In addition, microtubules from the B1-KD cells spent less total time at pause (24 vs 46% for B1-KD and A549 cells, respectively) and displayed enhanced overall microtubule dynamicity (16 μm/min) as compared with microtubules from A549 cells (9 μm/min; Figure 2c and Table 1). Taken together, these data suggest that BRCA1 has a role in microtubule biology by dampening centrosomal microtubule nucleation and overall microtubule dynamics.

Figure 2.

BRCA1 regulates microtubule dynamics. (a) Microtubule regrowth following nocodazole-induced depolymerization and wash-out, was assessed in A549 and B1-KD cells by immunostaining with antibodies against α-tubulin and γ-tubulin to mark the centrosome (arrows mark the center of the centrosome). The length of each microtubule originating from the centrosome was measured and graphically displayed (right panel, ****P<0.001). White scale bar = 10 μm. (b) Time-lapse confocal microscopy of EB1-EGFP puncta was performed for 1-min recording 1 frame per second following transient transfection of A549 and B1-KD cells. The tracks of EB1-EGFP puncta movement are shown with the following designations: Red = start of track; green = path of puncta during time lapse; blue = end of track. EB1-EGFP comet velocity was determined for tracks that remained in focus for the duration of the recording and had a discernible end point (blue). Left: representative single cell images from each condition are shown. Right: graphic display of average comet velocity calculated as the ratio of total distance of travel (μm) divided by the duration of recording time (1 min; ****P<0.001). Average values represent measurements in approximately 30 individual cells per condition. White scale bar = 10 μm. (c) Microtubule dynamics in A549 or B1-KD cell lines were measured with live-cell imaging following transfection of EGFP-tubulin. Shown are representative examples of frames from consecutive time points with the arrow highlighting the tip of a microtubule. Microtubule dynamics were measured by tracking tips of microtubules using EGFP-tubulin and live-cell spinning disk confocal microscopy. Quantitation of distinct microtubule dynamics parameters is shown in Table 1. All parameters measured were significantly different between A549 and B1-KD cell lines (P<0.001) except the catastrophe frequency (P<0.05; italicized). Scale same as in panel b.

Table 1.

Analysis of microtubules dynamics in the presence and absence of BRCA1 in A549 cells

Parameter	A549	B1-KD	% Changea	P-value
Mean rates, μm/min
Polymerization	13.99±0.57	18.30±0.74	30.81	<0.001
Depolymerization	14.52±0.55	21.67±0.79	32.99	<0.001
% Time spent
Polymerization	25.64±1.48	39.26±1.41	34.69	<0.001
Depolymerization	28.44±1.35	36.76±1.50	29.25	<0.001
Pause	45.92±2.38	23.98±1.36	−47.78	<0.001
% Transition frequency
Rescue	28.64±0.69	25.93±0.57	−9.46	<0.001
Catastrophe	29.66±0.78	27.41±0.59	−7.59	<0.05
Dynamicity, μm/min	9.15±0.27	15.79±0.52	72.57	<0.001

^aThe percentage difference relative to the corresponding parameter in A549 cells.

Loss of BRCA1 impairs PTX-induced microtubule stabilization

To examine whether the effects of B1-KD on microtubule dynamics affected the ability of PTX to engage its target and induce microtubule stabilization, we examined the extent of drug-induced microtubule stabilization in A549 and B1-KD cells. We used microtubule bundling (Figure 3a, arrows) as the readout for cellular taxane activity in an immunofluorescence assay, as microtubule bundle formation is the result of drug-mediated microtubule polymer stabilization and as such it represents a hallmark of taxane cellular activity. Using this assay, we observed that the ability of PTX to induce microtubule bundling in the B1-KD cells was markedly compromised as compared with the abundant microtubule bundles present in PTX-treated A549 cells (Figure 3a and Supplementary Figure S6). To quantitate the effect of PTX treatment on microtubule polymer mass, we performed a cell-based tubulin polymerization assay, in which cells were exposed to PTX and microtubule polymers (P) were separated from soluble tubulin dimers (S) by centrifugation. As shown in Figure 3b, PTX treatment of A549 cells resulted in a shift of total tubulin recovered in the polymer fraction, from 10% in control to 60% following PTX treatment. In contrast, PTX treatment of B1-KD cells resulted in a modest increase in microtubule polymer mass, from 8% in untreated cells to 36% in drug-treated cells. Taken together, these results indicate that loss of BRCA1 renders microtubules more dynamic and less responsive to PTX treatment suggesting that the drug–tubulin interaction might be compromised under these conditions.

Figure 3.

B1-KD impairs PTX-induced microtubule stabilization. (a) A549 and B1-KD cells were treated with vehicle (dimethylsulfoxide (DMSO)) or 1 μM PTX for 1 h, fixed and immunostained with α-tubulin and imaged using confocal microscopy. Images were acquired under identical settings with a 100X/1.4NA objective. Representative cells from each condition are displayed. Arrows point to PTX-induced microtubule bundles. White scale bar = 10 μm. (b) Cell-based tubulin polymerization assay in A549 and B1-KD cells. Upper panel: cells were treated with vehicle (DMSO) or 100 nM PTX for 1 h and subjected to cell-based tubulin polymerization assay. Briefly, following lysis and centrifugation the pellet fraction (P), containing polymerized microtubules, was separated from the supernatant fraction (S), containing soluble tubulin dimers and equal volumes of each were loaded on adjacent wells on sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) and immunoblotted for α-tubulin. Ladder band represents 50 kDa. Lower panel: graphic display of % tubulin in the pellet fraction, which was calculated as the densitometric ratio of the pellet fraction (P) divided by the total tubulin content (P + S) for each condition. Data show the average of three biological repeats. (c) Live-cell-based PTX-binding affinity assay: A549 and B1-KD cells were incubated with 250 nM TubulinTracker for 16 h at 37 °C. After the incubation, TubulinTracker was washed out three times and replaced with phenol red-free RPMI. Cells were imaged using live-cell confocal microscopy using identical conditions and images were acquired with a 40X/0.75NA objective. Drug binding to tubulin was assessed on a single cell by dividing the total cellular fluorescence intensity by the total area of the cell determined by phase-contrast imaging. One hundred cells per condition were analyzed. ***P<0.001. (d) Microtubule co-sedimentation assay: A549 cells were lysed, precleared (HSS) and subjected to a microtubule co-sedimentation assay following addition of exogenous purified bovine brain microtubule protein. A cycle of tubulin polymerization was performed at 37 °C for 30 min in the presence of 20 μM PTX and was followed by centrifugation to separate the microtubule polymer fraction (WP) from the soluble tubulin dimer fraction (WS). A cycle of depolymerization was performed by subjecting the WP fraction to cold- and calcium-induced depolymerization. Following centrifugation the cold-insoluble microtubule pellet (CP) was separated from the cold supernatant (CS). Equal volumes from each fraction were loaded onto adjacent wells of an SDS–PAGE gel and immunoblotted for α-tubulin and BRCA1. Ladder band in tubulin Immunoblot represents 50 kDa. CS, cold supernatant; CP, cold pellet; HSP, high-speed pellet; HSS, high-speed supernatant; WS, warm supernatant.

To further investigate why loss of BRCA1 makes microtubules less responsive to PTX treatment, we examined the extent of PTX cellular binding affinity to microtubules in A549 or B1-KD cells. To do so, we treated cells with a fluorescently tagged PTX for 16 h, and measured the average fluorescence intensity in the cell’s cytoplasm in both cell lines. We observed that B1-KD cells had a ~50% decrease in average fluorescence intensity compared with the A549 cells when normalized for cell area (Figure 3c). These data corroborate and extend our earlier observations and demonstrate that loss of BRCA1 diminishes the binding of PTX to cellular microtubules resulting in impaired drug-induced microtubule stabilization.

Microtubules serve as binding sites for many microtubule-associated proteins (MAPs), which regulate endogenous microtubule dynamics and control changes in microtubule behavior in response to stress. Our results indicate that BRCA1 directly or indirectly affects microtubule biology and response to microtubule-stabilizing agents. Accordingly, BRCA1 may be a MAP that promotes microtubule stability, similar to the von Hippel-Lindau tumor suppressor protein,³² and whose loss could render microtubules hypostable. To test the hypothesis that BRCA1 is a MAP, we performed a microtubule co-sedimentation assay, which is a more sensitive and robust alternative to co-immunoprecipitations, for the identification of cellular proteins that have affinity for the microtubule polymer. This assay utilizes successive cycles of tubulin polymerization and depolymerization based on the physical and chemical properties of tubulin, whereas the addition of exogenous purified tubulin provides a ‘microtubule polymer core seed’ for MAP binding. Here, briefly pre-cleared total cell lysate (high-speed supernatant) from A549 cells was supplemented with exogenous purified microtubule protein and subjected to a cycle of polymerization in the presence of PTX and guanosine-5′-triphosphate (GTP) at 37 °C. Microtubule polymers and any endogenous proteins with affinity for them were sedimented in the pellet fraction (warm pellet (WP)) following high-speed centrifugation, whereas soluble tubulin dimers and all other cellular proteins remained in the supernatant (warm supernatant). To ensure that the resulting WP fraction contained functional microtubules as opposed to nonspecific protein aggregate, we subjected microtubule polymers from the WP fraction to a cycle of cold-, calcium-induced depolymerization followed by high-speed centrifugation in order to separate the soluble tubulin (cold supernatant) from any remaining cold-stable microtubules (cold pellet). Surprisingly, our results showed that no BRCA1 was recovered in the WP fraction under conditions that resulted in near 100% tubulin polymerization and as such no BRCA1 was recovered in the cold supernatant and cold pellet fraction, which are derivatives of WP. Instead, all of BRCA1 protein was recovered in the supernatant fraction (warm supernatant) suggesting that BRCA1 is not directly associated with microtubules but that it exerts its effects on microtubules indirectly (Figure 3d).

Loss of BRCA1 reduces PTX-induced caspase-8 activation

Caspase-8 activation has been recently implicated in the mechanism of PTX-induced apoptosis, via the association of the death-effector domain of pro-caspase-8 with stable microtubules.²⁸ This report together with our results showing that BRCA1 loss impairs PTX-induced microtubule stabilization and sensitivity to drug treatment prompted us to examine the involvement of the caspase-8 death-effector domain in the mechanism of BRCA1-mediated PTX resistance. Immunostaining of A549 and B1-KD cells with the death-effector domain of caspase-8 revealed that caspase-8 colocalized with PTX-induced microtubule bundles in the A549 cells, but not in the B1-KD cells (Figure 4A). As caspase-8 accumulation on PTX-stabilized microtubules has been reported to lead to zymogen cleavage and activation of caspase-8, we next investigated the ability of PTX to induce caspase-8 activation in the B1-KD line. PTX treatment of A549 cells resulted in caspase-8 activation as indicated by the presence of the active cleaved form of caspase-8 (Figure 4B). In contrast, no activation of caspase-8 was observed in B1-KD cells following PTX treatment. To confirm that caspase-8 activation is responsible for PTX-induced cell death, we pre-treated cells with a caspase-8 inhibitor before PTX treatment and assessed the ability of the drug to induce cell death by performing flow cytometry analysis for markers of cell death, namely annexin-V and propidium iodide. Our results showed that caspase-8 inhibitor decreased PTX-induced cell death (Figure 4C, P <0.01, and Supplementary Figure S7). Taken together, these data suggest that caspase-8 cleavage, and subsequent cell death, is activated on short-term PTX treatment in the A549 cell line and that loss of BRCA1 prevents this activation possibly due to the loss of PTX-stabilized microtubule, which serves as a platform for caspase-8 accumulation and induced-proximity activation.

Figure 4.

B1-KD impairs PTX-induced caspase-8 microtubule localization and activation. (A) A549 and B1-KD cells were treated with dimethylsulphoxide (DMSO) vehicle (a–f) or 100 nM PTX (a′–f′) for 18 h, fixed and processed for double immunofluorescence staining using antibodies specific for the N-terminal domain of caspase-8 (DED domain) or α-tubulin. 4,6-Diamidino-2-phenylindole (DAPI) is used to counterstain the nucleus. Images were acquired under identical settings with a 100X/1.4NA objective of a ZEISS LSM 700 confocal microscope. White scale bar = 10 μm. Insets depict a higher magnification of the indicated areas. (B) A549 and B1-KD cells were treated with 100 nM PTX for 18 h, lysed and 50 μg of total cell protein was run on sodium dodecyl sulfate–polyacrylamide gel electrophoresis (SDS–PAGE) and immunoblotted with antibodies against total and cleaved caspase-8. Actin is used as a loading control. (C) A549 cells were treated with 10 nM PTX for 36 h in the presence or absence of 30 μM caspase-8 inhibitor (C8i), which was added 2 h before onset of PTX treatment. Cells were stained for propidium iodide (PI) and annexin-V and analyzed for apoptosis via flow cytometry. Cell viability (PI and annexin-V negative) was rescued with C8i pre-treatment. **P<0.01.

DISCUSSION

PTX is the prototype of microtubule-stabilizing drugs and, since the discovery of its unique mechanism of action, researchers worldwide have been focusing on the discovery of small molecules with a similar mechanism of action. Ever since the introduction of PTX in clinical oncology, an equally intensive research effort has been undertaken to understand the molecular basis of clinical activity and resistance. To date, despite the publication of >20 000 research reports on PTX, the mechanisms underlying clinical taxane resistance remain poorly understood. Acquired drug resistance is the number one cause of cancer-related deaths in metastatic patients. Therefore, a better understanding of the pathways responsible for drug resistance is imperative, in order to design improved therapeutic strategies.

BRCA1 has recently emerged as a clinical biomarker predictive of taxane activity. Several studies have reported that low BRCA1 expression levels and/or BRCA1 loss of function is associated with a lower patient response to taxane-based chemotherapy. BRCA1 is a tumor-suppressor protein mutated in 5–10% of breast and ovarian cancer patients, whereas low expression levels of BRCA1 have been reported in over 20–30% of patients with a variety of solid tumors.^19,33–40 The BRCA1 protein is involved in several cellular pathways including DNA repair, checkpoint activation and protein ubiquitination.¹⁹ Interestingly, low BRCA1 levels or mutation-driven loss of function have been correlated with chemosensitivity to platinum and chemoresistance to taxane chemotherapy in numerous clinical and preclinical studies.^14–17 As platinum and taxane chemotherapies are often used in combination for the treatment of different solid tumors and in particular lung cancer, current prospective clinical trials are utilizing BRCA1 expression to assign patients to platinum or taxane chemotherapy in an effort to personalize chemotherapy and enhance clinical response (ClinicalTrials.gov ID#’s NCT01424709, NCT00478699, NCT01276769). The sensitization of tumors with BRCA1 loss of function to platinum agents has been attributed primarily to the central role of BRCA1 in DNA repair pathways such as homologous recombination and non-homologous end-joining.²⁰ DNA-damaging drugs such as cisplatin are known to work by cross-linking DNA strands, therefore, their activity is enhanced when DNA repair is compromised. In contrast, little data exist to explain the relationship behind the resistance of tumors with BRCA1 loss of function to taxanes, partly due to the lack of understanding of the role that BRCA1 has in microtubule biology. Here, we provide evidence that will help shed some light on the role of BRCA1 in taxane resistance. We show that BRCA1 modulates microtubules such that loss of BRCA1 leads to increased microtubule dynamicity. We demonstrate that the increase in microtubule dynamicity renders microtubules less sensitive to PTX-induced stabilization and in turn prevents the stabilized microtubule-based activation of caspase-8 and subsequent activation of cell death (Figure 5).

Figure 5.

Model for the role of BRCA1 in taxane resistance. Here, we show that BRCA1 regulates microtubules such that the presence of BRCA1 dampens microtubule dynamics. Less dynamic microtubules encourages the formation of PTX-stabilized microtubule bundles, which serve as a platform for pro-caspase-8 binding. The accumulation of multiple pro-caspase-8 molecules, via the induced-proximity activation model, leads to zymogen cleavage and transactivation of adjacent caspase-8 subunits. Once activated, caspase-8 cleaves downstream substrates that lead to cell death. The absence (or low expression levels) of BRCA1, leads to more dynamic microtubules by negating BRCA1-mediated suppression of dynamics. More dynamic microtubules are in turn, poorer substrates for PTX binding. Lower PTX binding results in fewer stabilized microtubules (bundles), which prevents the formation of the microtubule-based platforms for pro-caspase-8 accumulation and activation. Ultimately, the loss of BRCA1 prevents the formation of PTX-stabilized microtubule bundles rendering the mechanism for caspase-8 activation and downstream signaling ineffective. Such a mechanism can potentially explain the clinically observed lower response rate of cancer patients with low BRCA1 expression to taxane-based chemotherapy.

A major question arising from our data is how BRCA1 regulates microtubule biology. Proteins that regulate microtubules in cells are the so-called MAPs that bind microtubules directly in order to promote microtubule stabilization (for example, MAP4)⁴¹ or destabilization (for example, stathmin).⁴² In addition to these classically defined MAPs, along with others, we have identified numerous other proteins that associate with the microtubule cytoskeleton for the purposes of intracellular trafficking (for example, p53, AR, HIF),^43–46 protein–protein interactions (for example, von Hippel-Lindau tumor suppressor protein)^32,47 and signaling (for example, GSK3β, APC).^48,49 Therefore, the first question that we asked was whether BRCA1 associates with microtubules in order to suppress microtubule dynamics. However, our results argue against BRCA1 being a MAP as we clearly show that BRCA1 does not co-sediment with microtubule polymers (Figure 3d) suggesting that BRCA1 does not bind tubulin directly. These results have been corroborated by lack of colocalization between the two proteins as assessed by BRCA1 and tubulin immunofluorescence staining as well as lack of co-immunoprecipitation using antibodies specific for each respective protein (data not shown). However, complete lack of interaction between the two proteins in the cell’s cytoplasm cannot be entirely ruled out as our assays might not have detected a transient or weak interaction between the two proteins. Taken together, these data suggest that BRCA1 is indirectly modulating cellular microtubule dynamics. Although BRCA1 has been shown to associate with some MAPs (for example, Nlp, TPX2, NuMA, XRHAMM),^50,51 these are mitotic spindle proteins, which do not have a regulatory role in the modulation of interphase microtubule dynamics. BRCA1 is associated with several protein complexes that perform multi-functional processes in the cell, primarily in the nucleus. For BRCA1 to modulate microtubule stability, we reasoned that a cytoplasmic pathway functionally links the two proteins. An intriguing possibility for such a cytoplasmic pathway is that of the E3 ubiquitin ligase activity of BRCA1. BRCA1 in association with BARD1 forms a functional E3 ubiquitin ligase and this complex has recently been identified to regulate breast cancer cell spreading and motility by interacting with focal adhesion proteins.²² Interestingly, one of the targets of the BRCA1/BARD1 E3 ubiquitin ligase is γ-tubulin,⁵² whose ubiquitination impairs γ-tubulin’s centrosomal localization and ability to nucleate microtubules.^24,25 Indeed, we show here that B1-KD cells have impaired γ-tubulin ubiquitination (Supplementary Figure S1b), and consequently enhanced γ-tubulin centrosomal localization (Figure 1c) and microtubule nucleation (Figure 2a). Interestingly, a recent report showed that a decrease in neuronal γ-tubulin levels dampened microtubule dynamics, functionally implicating the centrosomal nucleation that occurs at the minus-end of microtubules with regulation of their plus-ends dynamicity.⁵³ In support of a role of γ-tubulin in the modulation of microtubule dynamics, another report showed that in Drosophilia S2 cells, γ-tubulin localizes to interphase microtubules lattice regulating dynamics by limiting microtubule catastrophe frequencies.⁵⁴ Our data also support of role for γ-tubulin in the modulation of endogenous microtubule dynamics whereby loss of BRCA1 impairs γ-tubulin ubiquitination leading to enhanced γ-tubulin centrosomal localization, which in turn results in increased microtubule dynamicity.

Another possibility as to how the BRCA1/BARD1 complex may regulate microtubule dynamics is by controlling the ubiquitination status of the tubulin α/β heterodimer. Recent data have revealed that α-tubulin exists in a ubiquitinated state, however, the E3 ubiquitin ligase responsible for this α-tubulin modification or the functional significance of tubulin ubiquitination, remain unknown.⁵⁵ Intriguingly, our preliminary results using α-tubulin immunoprecipitation from A549 and B1-KD whole-cell lysates show a strikingly decreased amount of ubiquitinated α-tubulin from the B1-KD cells compared with the A549 cells (highlighted boxes, Supplementary Figure S8a). Although we showed that BRCA1 is not a MAP (Figure 3d), it stands to reason that, because of the high homology between γ-tubulin and α/β tubulin, the α/β heterodimer may also be a substrate for the BRCA1/BARD1 E3 ubiquitin ligase complex. Transfection of 293 cells with a ubiquitinligase defective mutant BRCA1 B1 (I26A) plasmid showed a trend toward increased PTX resistance, however, additional experiments are required in order to draw a firm conclusion (Supplementary Figure S8b). Our preliminary data in Supplementary Figure S8a support the notion that BRCA1 controls the ubiquitination status of the tubulin α/β heterodimer, whether directly or indirectly. Future work in our laboratory will focus on the role of α/β tubulin ubiquitination in relation to regulation of microtubule dynamics and downstream microtubule signaling pathways.

Importantly, we show here that loss of BRCA1 cells confers taxane resistance, likely due to the fact that dynamic microtubules are a relatively poor substrate for PTX binding (Figure 3c). In agreement with our data, earlier reports have shown that enhanced microtubule dynamics are associated with taxane resistance in A549 cells, following in vitro selection for PTX resistance.⁵⁶ Similarly, earlier studies have demonstrated that the homeostasis of microtubule dynamics and the equilibrium between microtubule polymers and the pool of soluble tubulin dimers, changes following selection with a microtubule-stabilizing or -destabilizing drug, resulting in cells with hypo-or hyper-stable microtubules, respectively.⁵⁷ Together these studies provide further evidence for the notion that highly dynamic microtubules serve as a poor substrate for taxane binding and are in agreement with our results showing lower TubulinTracker labeling of the microtubule cytoskeleton in B1-KD cells (Figure 3c). Interestingly, our results also show that loss of BRCA1 renders microtubules hypostable, more sensitive to the effects of the microtubule-depolymerizing drug nocodazole (Supplementary Figure S9) and more resistant to the microtubule-stabilizing drug PTX (Figures 3a and b).

Although changes in microtubule homeostasis are established cellular responses to drug selection with microtubule inhibitors, here we show that a biological perturbation, namely loss of BRCA1, rather than drug-selection pressure, increases microtubule dynamicity resulting in taxane resistance. We have recently shown that knockdown of protein farnesyltransferase, which unlike BRCA1 binds microtubules, also resulted in enhanced microtubule dynamicity and taxane resistance.^58,59 Clinically, we have also found a correlation between lower expression of farnesyltransferase and lack of clinical response to taxane-based chemotherapy,⁶⁰ similar to the existing clinical studies with BRCA1. Although, the exact pathway that links BRCA1 with microtubule dynamics remains to be fully elucidated, we provide here an explanation that links loss of BRCA1 with taxane insensitivity via the lack of microtubule-dependent activation of caspase-8 by the induced-proximity model (Figure 5).

Taken together, these data continue to define the microtubule as a platform for signal transduction. Long has it been appreciated the microtubule’s roles in basic cell biology including the motor transport of vesicles and organelles and mitosis. A role for microtubule biology as being central to signal transduction pathways is emerging.^45,59,61 Here, we show that PTX-induced cell death signaling through caspase-8 is directly controlled by how microtubules respond to the drug, and that the behavior of these microtubules is regulated by BRCA1.

Identifying the precise mechanism by which low BRCA1 levels lead to taxane resistance could allow the development of therapeutic agents that could re-sensitize tumors with low BRCA1 expression levels to taxane agents, or potentially identify a combination therapy that would be synergistic for taxane response in both taxane-sensitive and taxane-resistant patients.

MATERIALS AND METHODS

Reagents

A549 and HEK 293T cells were obtained from ATCC (Manassas, VA, USA). The EB1-EGFP construct was a gift from the Kreitzer lab at Weill Cornell Medical College, New York, NY, USA. The EGFP-α-tubulin construct a kind gift from the Jordan lab at the University of California, Santa Barbara, CA, USA. Primary antibodies for BRCA1 (C-20, Santa Cruz Biotechnology, Santa Cruz, CA, USA), tubulin (YL1/2, Novus Biologicals, LLC, Littleton, CO, USA), actin (A2066, Sigma-Aldrich, St Louis, MO, USA), pericentrin (ab4448, Abcam, Cambridge, MA, USA), total caspase-8 (1C12, Cell Signaling Technology, Danvers, MA, USA) and cleaved caspase-8 (18C8, Cell Signaling Technology) were obtained. Alexa Fluor secondary antibodies (Invitrogen, Carlsbad, CA, USA) for immunofluorescence and western blots were obtained. Puromycin, PTX and nocodazole were obtained from Sigma-Aldrich. Fugene 6 Transfection Reagent was obtained from Roche Diagnostics (Basel, Switzerland). The caspase-8 inhibitor II (Cat#218759) was obtained from Calbiochem (Gibbstown, NJ, USA). TubulinTracker Green was obtained from Invitrogen. Protease inhibitor cocktail (REF 11836170001) was obtained from Roche Diagnostics. Propidium iodide and annexin-V were obtained from BD Biosciences (San Jose, CA, USA).

Cell culture and transfections

A549 cells were cultured in RPMI supplemented with 10% fetal bovine serum and 1% pencillin/streptomycin. BRCA1-specific shRNAs were obtained from Open Biosystems (Huntsville, AL, USA). Lentiviral particles were generated and harvested from HEK293T cells using standard protocols. A549 cells were infected with BRCA1-specific lentiviral shRNA particles for 24 h, and colonies were selected with 2 μg/ml puromycin. The EB1-EGFP and EGFP-tubulin constructs were transfected using the Fugene 6 reagent according to the manufacturer’s protocol.

Real-time PCR

mRNA was isolated from the A549 and B1-KD cells using the RNeasy Mini Kit (Qiagen, Hilden, Germany). In all, 1 μg of RNA was used to generate complementary DNA using the Protoscript First Stand cDNA Synthesis Kit (New England Biolabs, Ipswich, MA, USA). A 1:10 dilution of the complementary DNA was used as template in an iQ SYBR Green Supermix (Bio-Rad, Hercules, CA, USA) reaction with BRCA1-specific primers.

• BRCA1 forward primer = 5′-AGCCAGCCACAGGTACAGAG-3′

• BRCA1 reverse primer = 5′-AGTAGCCAGGACAGTAGAAGGAC-3′

A relative quantification of BRCA1 mRNA levels, using β-actin as the internal control, was calculated using a comparative Ct method.

Western blot

Following drug treatments, cells are collected and lysed in lysis buffer (50 mM Tris pH 7.5, 100 mM NaCl, 2 mM EDTA, 1% NP-40) supplemented with 7 × Protease Inhibitor cocktail (Roche Diagnostics). Following separation via gel electrophoresis, proteins were transferred onto polyvinylidene difluoride and blocked in 5% milk, 1% Tween-20 and 0.1% fetal bovine serum. Target proteins were detected using secondary antibodies compatible with the Odyssey Imaging System from LICOR Biosciences (Lincoln, NE, USA). Densitometric analysis was performed where appropriate with ImageJ (National Institutes of Health, Bethesda, MD, USA).

Immunofluorescence

Cells were plated on #1.5 coverslips (Electron Microscopy Services, Hatfield, PA, USA). Following drug treatment, cells were pre-extracted with 0.1% Triton X-100 for 30 s and fixed with PHEMO buffer as described previously.⁶² Cells were stained with primary antibodies at room temperature for 1 h and secondary antibodies at room temperature for 30 min. Coverslips were then mounted onto slides with MOWIOL. Images were acquired with a Zeiss LSM 700 (Carl Zeiss Inc., Thornwood, NY, USA).

Cytotoxicity assay

Cells were plated in 96-well plates and treated with drugs for 30 h. Cell viability was measured using the CellTiter-Glo Luminescent Cell Viability Assay from Promega (Madison, WI, USA) according to the manufacturer’s protocol. Percentage survival was calculated as the treatment divided by control for each concentration of drug.

Microtubule regrowth assay

Cells on coverslips are treated with 10 μM nocodazole at 4 °C for 1 h to completely depolymerize microtubules. Nocodazole was washed out, replaced with warmed media followed with incubation at 37 °C for 10 min. Cells then fixed, stained and imaged as described above. Microtubule length was measured as the length of a single microtubule originating from the centrosome to the most distal tip (MetaMorph Software, Universal Imaging, Downingtown, PA, USA).

Live-cell imaging with EB1-EGFP

EB1-EGFP transfected A549 and B1-KD cells were imaged with Hamamatsu Orca R2 camera (Middlesex, NJ, USA) mounted onto a CSU-X1 (Yokogawa Electric Corporation, Inc., Tokyo, Japan) spinning disk head on a Zeiss SD Observer I equipped with a 100 × Plan-Apochromat (NA = 1.46) oil objective (Carl Zeiss Inc.). Time lapses were recorded and measured with Metamorph. EB-1 comet velocity was measured as the distance each EB1 puncta traveled over time lapse divided by the total time of the time lapse. Only those puncta tracks with a discernible beginning and end were measured.

Microtubule dynamics assay

EGFP-α-tubulin transfected A549 and B1-KD cells were imaged with the same microscope setup mentioned above and imaging experimental conditions were recorded as previously described.⁶³ Briefly, the tips of growing and shrinking microtubules were tracked in time lapse movies recorded every 5 s for a total of 5 min. Polymerization and depolymerization mean rates were calculated as total distance of growth or shrinkage, respectively, over time. A catastrophe is defined as a transition into microtubule shortening whereas a rescue is a transition from shortening to growth or pause. To calculate catastrophe frequency per unit time, the number of catastrophes was divided by the total time in growth. Conversely, the rescue frequency was calculated by dividing the number of rescues by the total time spent. Microtubule dynamicity is defined as the total length grown and shortened during the life (measured in minutes) of an individual microtubule.

TubulinTracker assay

For TubulinTracker experiments, A549 and B1-KD cells were treated with 250 nM TubulinTracker Green for 16 h at 37 °C. Following this incubation, the TubulinTracker was washed three times with phosphate-buffered saline and replaced with phenol-red-free RPMI. Images were acquired with the same spinning disk microscope with a 40 × Plan-Neofluar (NA = 0.75) objective (Carl Zeiss Inc.). To determine the area of the cell, an image with transmitted light was acquired. The average fluorescence intensity of the TubulinTracker was measured following a statistical background correction (MetaMorph) by using the transmitted light image to define the area of the cells. The fluorescence intensity of the TubulinTracker was normalized by the total area of the cell. For each condition ~150 cells were measured.

Tubulin polymerization assay

Following drug treatment, cells were washed warm phosphate-buffered saline once. Cell were scraped and lysed in low salt buffer (20 mM Tris–HCl pH 6.8, 1 mM MgCl₂, 2 mM EGTA, 0.5% NP-40, 1X protease inhibitor cocktail). Samples spun at max speed in a tabletop centrifuge for 30 min at room temperature. The supernatant (S) was separated from the pellet (P). The pellet was resuspended in 1 × Laemmli buffer and sonicated. Equal volumes of supernatant and pellet samples were loaded onto a gel for western blot. % Pellet is calculated as the densitometric value of the pellet band divided by the total densitometric value of the pellet and supernatant bands. Three biological repeats were performed for this assay.

Microtubule co-sedimentation assay

A549 cell lysates were subjected to the microtubule co-sedimentation assay as previously described.⁴⁵ Briefly, A549 cells were scraped in PIPES-EGTA-MgCl₂ buffer (80 mM PIPES, 1 mM EGTA, 1 mM MgCl₂ pH 6.8) supplemented with 1X cocktail protease inhibitors. Scraped cells were spun at ~16 000 × g for 15 min at 4 °C and the high-speed pellet and high-speed supernatant were separated. Exogenous purified bovine tubulin (suspended in warm PIPES-EGTA-MgCl₂ buffer and 0.5 mM guanosine-5′-triphosphate (GTP)) was added to cell lysates (high-speed supernatant) and one cycle of microtubule polymerization was carried out at 37 °C with 20 μM PTX for 30 min. Microtubules were pelleted (WP) at ~100 000 × g for 35 min at room temperature and separated from the supernatant (warm supernatant). Microtubules in the WP were depolymerized by sonication and incubation with 12 mM CaCl₂ on ice for 30 min. Properly depolymerized microtubules (cold supernatant) were separated from aggregated tubulin protein and polymerized debris (cold pellet) by a cold (4 °C) centrifugation at ~30 000 × g for 15 min.

Flow cytometry

A549 cells were pre-treated with vehicle or 30 μM caspase-8 inhibitor (C8i) for 2 h in serum-supplemented media followed by addition of vehicle or 10 nM PTX for 36 h. Cells were collected, stained with annexin-V and propidium iodide for 15 min at room temperature and flowed through the LSR II system (BD Biosciences) to determine cell viability. Results were analyzed with the flow cytometry analysis software FlowJo (TreeStar, Ashland, OR, USA).

Statistics

Graphpad Prism 4 (Graphpad Software, Inc., La Jolla, CA, USA) was used to calculate Student’s t-test and analysis of variance test (where appropriate) to determine if the mean of the measurements were significantly different between the A549 and B1-KD cell lines. P-values are demonstrated as * = <0.05, ** = <0.01, *** = <0.001 and **** = <0.0001. All error bars shown in the plots are the standard error of the mean.