Formin-like 1 (FMNL1) is a member of the formin family of actin nucleators, and is one of the few formins for which in vitro activities have been well characterized. However, the functional roles of this mammalian formin remain ill-defined. In particular, it is unclear how the unique in vitro biochemical properties of FMNL1 relate to its regulation of cellular processes. Here, we demonstrate that FMNL1 depletion caused a dramatic increase in cellular F-actin content, which resulted in Golgi complex fragmentation. Moreover, increased F-actin and maintenance of Golgi structure were distinctly regulated by the gamma isoform of FMNL1, which required binding to actin. Importantly, in addition to Golgi fragmentation, increased F-actin content in the absence of FMNL1 also led to cation-independent mannose 6-phosphate receptor dispersal, lysosomal enlargement and missorting of cathepsin D. Taken together, our data support a model in which FMNL1 regulates cellular F-actin levels required to maintain structural integrity of the Golgi complex and lysosomes.
The actin cytoskeleton has been implicated in numerous cellular processes, including cell migration, morphogenesis, membrane trafficking, cell polarity and division (Pollard and Cooper, 2009). The ability of F-actin to regulate these events is dependent on actin nucleators that help overcome the energetic barriers impairing actin polymerization (Chhabra and Higgs, 2007). Formin proteins have emerged as important regulators of F-actin assembly and have been implicated in membrane remodeling, transcriptional activation and tumor cell proliferation and invasiveness (Carreira et al., 2006; Hannemann et al., 2008; Lizarraga et al., 2009; Tominaga et al., 2000).
Formins contain a characteristic formin homology 2 (FH2) domain, which dimerizes into a ‘donut-shape’ in order to regulate actin polymerization (Xu et al., 2004). Studies of both yeast and human formins indicate that FH2 dimers bind to barbed ends of actin filaments and move processively as filaments grow (Harris et al., 2004; Otomo et al., 2005; Xu et al., 2004). Importantly, in the yeast formin Bni1p, two highly conserved residues (I1431 and K1601) within the FH2 domain are required for actin association and proper FH2 conformation (Otomo et al., 2005; Xu et al., 2004). Additionally, the FH2 domain antagonizes capping proteins by preventing barbed-end capping, and thus alters filament elongation and/or depolymerization rates (Higgs, 2005).
Among the better-characterized formins are the yeast formins Cdc12 and Bni1 and the mammalian formins Dia1 and Dia2 (also known as protein diaphanous homolog 1 and 3, DIAPH1 and DIAPH3), and FRL1, which is a member of the formin-related protein in leukocytes (FRL) subgroup; also known as formin-like (FMNL) proteins. Interestingly, studies of the FH2 domains from each of these formins indicate that they are functionally diverse, showing variable effects on barbed-end elongation rates and actin filament dynamics. Some formins are potent actin nucleators (Dia1 and Dia2) whereas others are weak (Bni1 and FRL1) (reviewed in Higgs, 2005). Also, although Dia1 does not slow filament barbed-end elongation at all, Cdc12 completely inhibits it; whereas, FRL1, Dia2 and Bni1 slow barbed-end elongation by approximately 80, 70 and 25%, respectively (reviewed by Goode and Eck, 2007; Higgs, 2005). Moreover, the FH2 domains of FRL1 and Dia2 have actin bundling activity. Importantly, two formins, FRL1 and INF2 can sever actin filaments in vitro (Chhabra and Higgs, 2006; Harris et al., 2004). In addition to the numerous ways formins affect F-actin directly, other factors add complexity to formin function, such as distinct regulatory signals, differential interaction with a spectrum of formin-binding proteins and levels of free G-actin. The unique features of these various formin family members probably explain the diversity of actin-based structures found in formin-regulated processes (Chhabra and Higgs, 2007), which is why it is fundamentally important to understand the individual contributions of each of these formins to cell biology.
The FRL/FMNL subgroup of formins share similar domain organization to diaphanous proteins (Katoh, 2003) and consist of FMNL1 (FRL1), FMNL2 (FRL3) and FMNL3 (FRL2; here referred to as FMNL1-3 for consistency with recent literature and databases). Although their specific cellular functions are not well understood, emerging studies so far suggest that FMNL3 stimulates filopodia formation (Harris et al., 2010), and FMNL2 regulates cancer invasiveness (Kitzing et al., 2010). FMNL1 is overexpressed in T-cell malignancies, and is a proposed tumor antigen (Favaro et al., 2006; Schuster et al., 2007). Previous work from our group showed that FMNL1 accumulated near the microtubule-organizing center (MTOC) in T cells and regulated MTOC polarization toward the immunological synapse (Gomez et al., 2007). More recently, a study has described a novel FMNL1 isoform, FMNL1γ, in addition to the previously described FMNL1α and FMNL1β isoforms (Yayoshi-Yamamoto et al., 2000). These alternatively spliced isoforms of FMNL1 differ in their C-termini. Interestingly, the FMNL1γ isoform was reported to localize to the plasma membrane (PM), to regulate membrane blebbing and to be constitutively active (Han et al., 2009). FMNL1γ was also found to regulate podosome dynamics and adhesion in macrophages (Mersich et al., 2010). Thus, FMNL1 functions to regulate actin at various cellular locations, and has the potential to associate with membranous structures.
Here, we report a previously uncharacterized role for FMNL1-mediated regulation of actin dynamics in maintaining the structural integrity of the Golgi complex and lysosomes. Furthermore, this study yields biological relevance for the well-understood in vitro features of actin regulation by the FH2 domain of FMNL1 and provides insight into the crucial relationship between actin dynamics and structural maintenance of cellular organelles.
FMNL1 is expressed in epithelial cells
FMNL1 expression has been observed in T cells, B cells, malignant B cells from patients with chronic lymphocytic leukemia (CLL) and macrophages (Gomez et al., 2007; Han et al., 2009; Mersich et al., 2010; Schuster et al., 2007). However, northern blot analysis has shown FMNL1 expression in non-hematopoietic malignant cells (Krackhardt et al., 2002). We therefore investigated FMNL1 expression in Jurkat T cells as a positive control and other epithelial-derived cancer cell lines by immunoblotting. We found that FMNL1 is expressed in Jurkat T cells as expected, but also in HeLa cells and some of the pancreatic cancer cell lines analyzed (Fig. 1A). We also compared the expression of FMNL1 with that of the diaphanous related formins (Drfs), FMNL2, FMNL3, Dia1 and Dia2. We found that all the cell lines expressed these formins to varying degrees (Fig. 1B–D). Interestingly, we observed that among the Drfs analyzed, FMNL1 expression is predominant in HeLa cells. We have also observed FMNL1 expression in NK cells, as well as several breast and colorectal cancer cell lines (data not shown). We conclude that FMNL1 is expressed in a variety of epithelial cancer cells in addition to hematopoietic cells.
FMNL1 depletion increases cellular F-actin
The high expression of FMNL1 in HeLa cells, prompted us to use this cell line in further studies. We next examined the effect of FMNL1 depletion on F-actin using short hairpin RNA (shRNA) vectors (shFMNL1; Fig. 2A), coexpressing GFP, which allowed us to identify transfected cells (Gomez et al., 2005). The shFMNL1 decreased FMNL1 expression by 95% and 74%, in HeLa and Jurkat cells, respectively, and resulted in diminished FMNL1 immunostaining, thus confirming the antibody specificity (supplementary material Fig. S1). Depletion of FMNL1 modestly affected the expression of FMNL2 and FMNL3 (by 20 and 25%, respectively), whereas it had no effect on Dia1 or Dia2 expression (supplementary material Fig. S2A–C).
Strikingly, knockdown of FMNL1 induced the formation of massive actin stress fibers in HeLa cells (Fig. 2B). Similarly, suppression of FMNL1 in Jurkat T cells resulted in a substantial increase in cortical F-actin compared with control transfected cells (Fig. 2B). In order to obtain a quantitative measure of this effect, we used FITC-conjugated phalloidin to measure the F-actin content by flow cytometry in Jurkat T cells following suppression of FMNL1. Indeed, suppression of FMNL1 caused a significant increase in F-actin content (Fig. 2C,D). The observed increased in F-actin upon FMNL1 depletion did not correspond to increased levels of actin in cells (supplementary material Fig. S3). Thus, FMNL1 depletion results in increased F-actin content, suggesting that one role for FMNL1 is to negatively regulate cellular F-actin.
FMNL1 localizes to the Golgi complex
We next wanted to study the functional consequence of FMNL1-mediated regulation of F-actin in cells. To get a better idea of possible biological systems relying on FMNL1, we investigated the localization patterns of FMNL1 by immunofluorescence in HeLa cells. Consistent with previous studies in T cells (Gomez et al., 2007; Han et al., 2009), FMNL1 accumulated at a punctate structure next to the nucleus and also throughout the cytoplasm (Fig. 3A,B). Although there was no obvious overlap of FMNL1 with F-actin stress fibers or radiating microtubules, we observed that FMNL1 accumulated with punctate perinuclear F-actin structures surrounding the MTOC (Fig. 3A,B). This localization of FMNL1 resembled the characteristic morphology of the Golgi complex. Indeed, FMNL1 strongly colocalized with the trans-Golgi network marker Golgin97 (G97) and the cis-Golgi network marker GM130 (Fig. 4A,B). Colocalization with Golgi markers was also observed in Jurkat T cells (Fig. 4C,D). Additionally, we found that FMNL1 colocalized with endosomal and lysosomal markers (EEA1 and LAMP1, respectively) but only in the perinuclear region (supplementary material Fig. S4A,B), but for the most part we have not identified localization to a specific subcellular structure for the remaining FMNL1 that is dispersed throughout the cytoplasm.
The striking localization of FMNL1 to the Golgi complex prompted us to investigate this relationship. First, we analyzed the effect of brefeldin A, a toxin that causes Golgi dispersal, on FMNL1 localization. Upon brefeldin A treatment, FMNL1 was dispersed throughout the cytoplasm, resembling the expected affect of brefeldin A on GM130 (Fig. 4B). This observation provided further evidence that FMNL1 accumulates at the Golgi and behaves like Golgi-resident proteins upon Golgi dispersal. Second, we determined the effect of F-actin stabilization on the Golgi complex. HeLa cells transfected with YFP–actin were treated with DMSO or the actin stabilizer jasplakinolide (supplementary material Fig. S5). As expected, jasplakinolide altered the normal organization of actin stress fibers as well as the morphology of the Golgi, consistent with a previous report (Lazaro-Dieguez et al., 2006). FMNL1 remained localized to the Golgi, as can be observed by colocalization of FMNL1 with G97. Taken together, F-actin stabilization leads to altered Golgi morphology, suggesting a possible role of FMNL1 regulation of F-actin in maintaining the structure of the Golgi.
FMNL1 regulates Golgi structure
In order to determine the possible effects of FMNL1 suppression on Golgi morphology we imaged HeLa cells following FMNL1 depletion. Compared with control shRNA-transfected (shControl) cells, we observed, using the G97 marker (Fig. 5A), that shFMNL1-transfected cells displayed a dramatic fragmentation of the trans-Golgi network (TGN). This effect was also observed in FMNL1-depleted Jurkat T cells (supplementary material Fig. S6A). We scored the Golgi phenotype by categorizing it as compact or fragmented, according to its appearance. The Golgi shown in Fig. 5A exemplify the criteria used for classification. Quantification of these results showed that 31% of GFP-positive cells expressing control shRNA had a fragmented Golgi structure (Fig. 5B). However, 76% of shFMNL1-transfected cells (GFP-positive) had a fragmented Golgi phenotype. This quantification approach produced very similar results to those obtained using ImageJ quantification of Golgi fragments in 100 cells (supplementary material Fig. S6B,C). The analysis showed a significant increase in the number of Golgi fragments in FMNL1-depleted cells (supplementary material Fig. S6B). When we analyzed the distribution of Golgi fragments in shControl cells, we observed that the majority of cells (66%) contained fewer than 40 fragments per cell, whereas only 35% of shFMNL1-transfected cells fell into this category (supplementary material Fig. S6C). From these results, we concluded that 34% of shControl and 65% of shFMNL1 cells had a fragmented Golgi phenotype. Thus, we decided to utilize the visual quantification approach (Fig. 5B) for all future experiments in which the Golgi phenotype is used as a functional readout.
Interestingly, depletion of the closely related FMNL2 and FMNL3 did not lead to similar alterations in Golgi structure (Fig. 5B; supplementary material Fig. S7A,B), suggesting that FMNL1 functions independently of FMNL2 and FMNL3 in this context. We also used additional Golgi markers to confirm the Golgi fragmentation phenotype in FMNL1-depleted cells. The structural integrity of the cis-Golgi network was also compromised in shFMNL1-transfected cells, as observed by immunofluorescence using anti-GM130 (supplementary material Fig. S7C). An additional trans-Golgi marker, TGN46, also confirmed the effect of FMNL1 depletion (supplementary material Fig. S7D). Taken together, these results demonstrate that FMNL1-dependent processes maintain the structural integrity of both the trans and cis compartments of the Golgi complex.
Depletion of FMNL1 and the resultant Golgi fragmentation had no affect on the secretory trafficking pathway because we did not observed defects in the vesicular stomatitis virus glycoprotein transport assay (not shown). However, FMNL1 depletion altered the distribution of other membranous organelles involved in intracellular trafficking. Interestingly, in FMNL1-suppressed cells, lysosomes appeared swollen and bundled together in the perinuclear area (supplementary material Fig. S8A). Moreover, FMNL1 depletion caused a significant enlargement of lysosomes containing the lysosomal enzyme cathepsin D, increased intracellular levels of cathepsin D and missorting of cathepsin D (supplementary material Fig. S8B–D). Transport of cathepsin D is mediated by the cation-independent mannose 6-phosphate receptor (CI-MPR), which carries newly synthesized lysosomal enzymes from the Golgi to early endosomes, and then cycles back to the TGN to retrieve additional cargo (Pfeffer, 2009). At steady state, CI-MPR has been reported to localize to a juxtanuclear area, colocalizing with either TGN markers or EEA1-positive vesicles (supplementary material Fig. S9A,C). Suppression of FMNL1 caused a dramatic redistribution of endosomes and CI-MPR (supplementary material Fig. S9A). CI-MPR showed decreased localization with TGN46 staining in FMNL1-depleted cells, but maintained its localization to early endosomes (supplementary material Fig. S9B,D). This suggests that, in addition to Golgi structure, FMNL1 also regulates lysosomal trafficking pathways and lysosome structure.
Regulation of Golgi structure by FMNL1 is isoform specific and requires actin binding
To date, three isoforms of FMNL1 have been identified: FMNL1α, β and γ (Han et al., 2009). Using RT-PCR, all isoforms were found to be expressed in HeLa and Jurkat cells (supplementary material Fig. S10). Thus, we analyzed the morphology of the Golgi in HeLa cells transfected with suppression–rescue vectors, which express shRNA against FMNL1, and simultaneously re-express resistant HA–YFP–FMNL1α, β or γ (supplementary material Fig. S11A,B). The sequences of the FMNL1 isoforms used in this study have been previously described (Han et al., 2009). Analysis of YFP-positive cells, showed that cells re-expressing FMNL1α or β had Golgi fragmentation, whereas FMNL1γ accumulated near the Golgi and rescued the Golgi phenotype (Fig. 6A,B). In order to determine the contribution of actin binding to the role of FMNL1 in regulating Golgi morphology we generated an actin-binding mutant lacking the C-terminus of FMNL1 (ΔFH2). Significantly, FMNL1ΔFH2 failed to rescue the fragmented Golgi phenotype (Fig. 6A; supplementary material Fig. S11B), suggesting that the ability of FMNL1 to bind F-actin is crucial for its role in maintaining Golgi architecture.
Interaction of the formin Bni1p with barbed ends has been mapped to two highly conserved residues, Ile1431 and Lys1601, in the knob and lasso/post regions of the FH2 domain (Otomo et al., 2005; Xu et al., 2004). Further studies have confirmed the relevance of those sites (Harris et al., 2010; Harris et al., 2006; Otomo et al., 2005; Shimada et al., 2004). Mutations corresponding to this conserved Ile in FMNL1 lead to faster disassociation from barbed ends, decreasing the ability of FMNL1 to compete with capping protein (CP) (Harris et al., 2006). Thus, we investigated the biological relevance of the corresponding sites in FMNL1γ for the structural maintenance of the Golgi by generating FMNL1γ barbed-end binding mutants, I720A and K871D. Analysis of YFP-positive cells indicated that re-expression of either FMNL1γI720A or FMNL1γK871D failed to rescue the fragmented Golgi phenotype but did accumulate with the fragmented Golgi (Fig. 6B,C). Thus, the ability of FMNL1γ to regulate actin through barbed-end association is necessary for the structural integrity of the Golgi complex.
Increased F-actin in FMNL1-depleted cells is decreased by FMNL1γ expression
We next investigated whether the morphological perturbations that we observed in the structure of the Golgi complex are related to the increased F-actin in FMNL1-depleted cells. To test this, we re-expressed FMNL1 isoforms and mutants in cells depleted of FMNL1 and examined F-actin in these cells (Fig. 7). HeLa cells re-expressing YFP-tagged FMNL1α or FMNL1β had increased F-actin, similar to shFMNL1 cells (Fig. 7A,B). However, re-expression of FMNL1γ was sufficient to decrease F-actin when compared with non-transfected cells (Fig. 7C). Some cells expressing higher levels of FMNL1γ showed striking parallel increases in F-actin (not shown), but these cells represented a minority of the transfection pattern obtained with this isoform. Deletion of the FH2 domain of FMNL1 (FMNL1ΔFH2) resulted in high levels of F-actin in cells and the most variable pattern of F-actin across transfected cells (Fig. 7D). This mutant is highly stable and there is a diverse range of expression that might explain these differences. Indeed, we were unable to find non-transfected cells to compare phenotypes. Importantly, abolition of actin binding by FMNL1γ (using FMNL1γK871D) failed to decrease F-actin (Fig. 7E). Thus, we concluded that FMNL1γ re-expression was able to reduce F-actin in FMNL1-depleted cells by its ability to interact with actin barbed ends.
F-actin increases at the Golgi in FMNL1-depleted cells
The observed increase in F-actin in FMNL1-depleted cells (Fig. 2), is probably responsible for the observed fragmentation of the Golgi complex (Fig. 5) as there is evidence that pharmacological stabilization of F-actin and increased F-actin in cells causes Golgi vesiculation (Lazaro-Dieguez et al., 2006; von Blume et al., 2009) (supplementary material Fig. S5). Thus, we investigated whether the consequences of increasing cellular F-actin were distally affecting the structural integrity of the Golgi, or if we could observe F-actin alterations near the Golgi. To assess this, we transfected HeLa cells with shControl–YFP or shFMNL1–YFP and observed F-actin at the Golgi by immunofluorescence (Fig. 8A,B). We obtained z-stack images of a magnified area around the Golgi. Control cells had low levels of F-actin, mostly short filaments, which is consistent with previous reports suggesting that F-actin at the Golgi is a pool of short and highly dynamic actin filaments (Musch et al., 2001). By contrast, we observed increased size and number of actin filaments traversing the Golgi in shFMNL1 knockdown cells, suggesting that F-actin is increased at the Golgi upon FMNL1 depletion (Fig. 8B). Although re-expression of FMNL1α and FMNL1β did not diminish these increased levels of F-actin seen at the Golgi (Fig. 8C,D), FMNL1γ efficiently decreased F-actin at the Golgi, but not a FMNL1γ mutant deficient in actin barbed-end binding (Fig. 8D,E). These data suggest that FMNL1γ regulates the actin cytoskeleton at the Golgi, and maintains a pool of short actin filaments required for structural integrity of the Golgi.
In this study, we have identified a novel role for FMNL1 in the regulation of F-actin and the morphology of the Golgi complex. Specifically, we showed that FMNL1 localized to the Golgi complex, where it maintained the structure of the Golgi through its ability to interact with actin barbed ends. Loss of FMNL1 led to increased actin content, which seemed to underlie defects in Golgi structural maintenance. Taken together, our data support a model in which the ability of FMNL1 to regulate F-actin is crucial for structural maintenance of membranous organelles, including the Golgi complex, endosomes and lysosomes (supplementary material Fig. S12).
We propose that the observed Golgi fragmentation results from the dramatic increase in F-actin observed in FMNL1-depleted cells. This is supported by the phenotype obtained by re-expressing FMNL1γ, which decreased F-actin at the Golgi and rescued the Golgi fragmentation phenotype. Moreover, we have observed a significant time-dependent decrease in the percentage of shFMNL1-treated cells exhibiting Golgi fragmentation when also treated with latrunculin B (not shown).
Studies that elucidate the mechanism involved in the increased F-actin in the absence of FMNL1 will be of great interest for the field. By western blot analysis, we did not observe an effect of FMNL1 on β-actin levels in the cell, suggesting that the increase in F-actin is not related to activation of the serum response factor known to be regulated by Dia1 (Copeland and Treisman, 2002). It is possible that upon removal of FMNL1 from actin barbed ends, these are free to interact with another formin efficient at F-actin polymerization. Another possibility is that the increase in F-actin levels might be a result of the reported in vitro actin-severing activity of FMNL1 (Harris and Higgs, 2006). Notably, when the severing proteins cofilin-1 and ADF are depleted from non-muscle mammalian cells, actin stress fibers increased in length and thickness (Hotulainen et al., 2005). Moreover, von Blume et al., showed that ADF/cofilin depletion altered the structure and functions of the Golgi (von Blume et al., 2009). Consistent with this, ADF/cofilin and FMNL1 depletion lead to similar defect in cathepsin D sorting. Interestingly, neither ADF/cofilin nor FMNL1 appear to regulate Golgi function to secrete membrane-associated cargoes as FMNL1 depletion did not prevent VSV-G trafficking to the plasma membrane (not shown).
As mentioned above, FMNL1 depletion led to dramatic fragmentation of the Golgi complex. Recently, the use of drugs that affect the actin cytoskeleton demonstrated the requirement of an intact and dynamic actin cytoskeleton to maintain Golgi structure and function (Lazaro-Dieguez et al., 2007; Lazaro-Dieguez et al., 2006). Also, many actin regulators and effectors that orchestrate actin remodeling at the Golgi have been reported (reviewed by Campellone et al., 2008; Egea et al., 2006; Kondylis et al., 2007). We now add FMNL1 to this list of Golgi-resident actin regulators. Our group and others have reported FMNL1 localization around the MTOC using antibodies raised against two different epitopes, an accumulation that we now know corresponds to the Golgi (Gomez et al., 2007; Han et al., 2009). In addition to FMNL1, other regulators of actin nucleation have been implicated in Golgi structure maintenance, including WHAMM, WASH and the Abi/Scar/Wave complex (Campellone et al., 2008; Gomez and Billadeau, 2009; Kondylis et al., 2007). Contrary to the ability of FMNL1 to nucleate linear actin, these factors promote nucleation of actin branches through the Arp2/3 complex. The Arp2/3 complex localization is restricted to the cis-Golgi network and to vesicles that traffic from and into the Golgi complex (Gomez and Billadeau, 2009; Matas et al., 2004). In this case, cells need to coordinate spatial and temporal regulation of the Arp2/3 complex with its activators for actin regulation (Matas et al., 2004).
Our study provides further evidence that FMNL1 isoforms regulate diverse cellular functions, and suggest potential differences in their relative abilities to regulate actin filaments. One important aspect of our findings is that Golgi structural integrity appears to be regulated exclusively by the FMNL1γ isoform. Strikingly, only re-expression of the FMNL1γ isoform appeared to rescue the dramatic increased in F-actin observed in FMNL1-depleted cells. Certainly, these results raise a number of questions regarding the role of the three FMNL1 isoforms in cells and the specificity rather than redundancy of their functions. Further studies in vivo and in vitro are required to elucidate any possible differential activities of these isoforms in elongation, severing or capping of actin, and the effects of their C-termini, specifically of the unique sequence of the FMNL1γ isoform, in such functions.
Our results support a model in which a balance between actin polymerization and depolymerization or capping is required to maintain Golgi architecture. It has been proposed that actin can be nucleated at the Golgi in the form of short, Golgi-associated microfilaments, and a role for those have been proposed in vesicle generation (Percival et al., 2004). Instead of highly stable actin filaments, a pool of highly dynamic actin filaments with free barbed ends has been proposed to exist at the Golgi (Musch et al., 2001). In this context, it is possible that FMNL1 is at the Golgi to support the formation or maintenance of such a pool of short and highly dynamic actin filaments. How FMNL1 balances its potential polymerizing, capping, bundling and severing activities at the Golgi remains to be investigated. The effects of FMNL1 on actin dynamics might be regulated differently in diverse cellular structures. In macrophages, FMNL1 depletion, decreases actin at podosomes (Mersich et al., 2010). Additionally, FMNL1 does not regulate filopodia formation (Harris et al., 2010). Thus, further analysis of the effects of FMNL1 in various cellular structures must be carried out to better understand these differences and what triggers them.
Altogether, we have identified that FMNL1γ localizes to the Golgi complex of mammalian cells where it regulates the structural architecture of the Golgi. Moreover, our data suggest that FMNL1γ-mediated dynamic regulation of the actin cytoskeleton, through its ability to bind barbed ends of F-actin, is responsible for the structural maintenance of other cellular organelles. Through the continued characterization of the roles and regulation of key actin regulatory proteins, we can continue to develop a picture of the physiological contributions of distinct F-actin structures in the architecture of membranous organelles.
Materials and Methods
Reagents and cell culture
We generated polyclonal antibodies against FMNL1, FMNL2 and FMNL3 (Gomez et al., 2007). In addition, we used polyclonal antibodies against Dia1 and Dia2 (dilution 1:5000; Bethyl Laboratories), EEA1 (1:500 dilution; BD Biosciences), TGN46 (1:500 dilution; Sigma) and cathepsin D (dilution 1:1000 for IF and 1:3000 for WB; EMD). Monoclonal antibodies against Golgin97 (1:300 dilution; Invitrogen), α-tubulin (1:3000 dilution; Sigma), EEA1 (1:500 dilution; BD Biosciences), LAMP1 (1:500 dilution; Santa Cruz) and GM130 (1:500 dilution; BD Biosciences) were used. Phalloidin 647 (1:50 dilution), fluorescein and Rhodamine-phalloidin (1:100 for immunofluorescence and 1:1000 for flow; Invitrogen) were used to detect F-actin. The following reagents were used as indicated in the corresponding figure legend: brefeldin A (BioLabs) and jasplakinolide (Invitrogen).
HeLa, Jurkat, Su86.86, BxPc3 and HupT3 cells were cultured in Roswell Park Memorial Institute (RPMI) medium supplemented with 10% FBS, at 37°C and 5% CO2. DAN-G cells were cultured in DMEM containing 10% FBS. Panc04.03 were cultured in RPMI supplemented with 15% FBS. Whole cell lysates were prepared in 1% NP40 (Sigma) lysis buffer, containing phenylmethylsulfonyl fluoride (PMSF; 175 μg/ml; Sigma), leupeptin (10 μg/ml; Sigma), aprotinin (10 μg/ml; Sigma), and empigen (3.3%; Bio-Rad). For experiments in which we immunoblotted for β-actin, lysates were prepared in 1% NP40, 0.1% SDS (Bio-Rad) and 0.5% deoxychocolate (Sigma), supplemented as described above.
HeLa and Jurkat cells were transiently transfected by electroporation (350 V and 330 V respectively, 1 pulse of 10 milliseconds). For overexpression studies using YFP–actin (Sigma), experiments were performed 16 hours after transfection. For shRNA transfections 30 μg/3 × 106 HeLa cells and 40 μg/10 × 106 Jurkat cells were used respectively, and cells were used 68–72 hours later.
The vectors pFRT.H1p and pCMS3.H1p have been described previously (Gomez et al., 2005). The vector pCMS3.H1p coexpresses GFP under the CMV promoter to allow identification of transfected cells. Information about the oligonucleotides used for cloning, mutations and shRNA transfections can be found in supplementary material Table S1. FMNL1β and FMNL1γ were cloned from HeLa and NK cell cDNA, respectively. FMNL1α was made from FMNL1γ using the QuikChange Site-Directed Mutagenesis Kit (Stratagene). The sequences obtained correspond to the reported sequences in the NCBI databases for the FMNL1 isoforms (FMNL1α, GI:33356147; FMNL1β, GI: 212843399; FMNL1γ, GI: 237780681). Because we used an oligonucleotide that corresponds to the 3′UTR of FMNL1β, we found two discrepancies with the NCBI sequence near the C-termini (NCBI: QVTSDLSL; ours: QVTSEVSL). FMNL1ΔFH2 corresponds to amino acids 1–632. shFMNL1 have been described and targets FMNL1 at bp 1306–1325 (Gomez et al., 2007). For rescue experiments, shFMNL1 and FMNL1 isoforms were cloned in the pCMS3.H1p/HA–YFP suppression–re-expression vector previously described (Gomez and Billadeau, 2009). All FMNL1 cDNAs were mutated to be shFMNL1 resistant (CGgGAcGCgGAgAAcGAATC) using the QuikChange Site-Directed Mutagenesis Kit.
Flow cytometry, immunofluorescence and microscopic quantification
To quantify F-actin levels by flow cytometry, Jurkat T cells were transfected with shRNA vectors, and 48 hours post-transfection, cells were washed and incubated overnight in serum-free medium supplemented with 4 mM L-glutamine. Cells were washed with PBS, fixed in 4% paraformaldehyde, permeabilized in 0.15% surfact-amps (Pierce), and incubated with fluorescein-phalloidin (1:1000) for 1 hour in FACS buffer (0.5% BSA in PBS). Data were collected using a FACS Canto II Cytometer (BD Biosciences).
For immunofluorescence, HeLa cells were grown on coverslips, and Jurkat cells were placed on poly-L-lysine-coated coverslips for 5 minutes in serum free medium at 37°C. Cells on coverslips were fixed in 4% paraformaldehyde in PBS (10 minutes at room temperature), washed with PBS (1 minute), permeabilized in 0.15% surfact-amps (3 minutes at room temperature; Thermo Fisher), washed with PBS (1 minute) and incubated in blocking buffer (5% normal goat serum, 1% glycerol, 0.1% BSA, 0.1% fish skin gelatin, 0.04% sodium azide) for 30 minutes (at room tempemperature) and incubated with primary antibodies in blocking buffer overnight at 4°C. After three PBS washes (5 minutes each), cells were incubated with secondary antibodies (1:800 dilution in blocking buffer) for 1 hour. The coverslips were then washed three times with PBS, followed by a 1 minute incubation with Hoechst 33342 (1:20,000 in water) and a final rinse with water. When Phalloidin was used, it was incubated with secondary antibodies. For some experiments, cells were pretreated with brefeldin A (10 nM) or jasplakinolide (1 μM).
Images were obtained using a LSM-710 laser scanning confocal microscope and analyzed using the Zen software package (Carl Zeiss). For Golgi fragmentation analysis, a cohort of shControl-transfected cells was used to establish image parameters. The rest of the samples were analyzed with the previously defined parameters. Over 50 cells, in triplicate, for each transfected population were scored blindly for Golgi fragmentation, in at least three independent experiments. Detailed analysis of the Golgi fragments were performed using ImageJ (100 cells per sample) as described previously (Kondylis et al., 2007) excluding fragments smaller than 2 pixels. LAMP1- and cathepsin-D-containing fragments were analyzed using ImageJ (34 cells). The same threshold was applied to all images and fragments smaller than 2 pixels were excluded from the analysis. Results are representative of at least three independent experiments. Mean fluorescence intensity and overlap coefficients (OC) were determined using the Zen software (Zeiss). For OC, values range from 0 to 1, where 0 denotes no colocalization and 1 denotes total colocalization. For Z-stack images, slices were obtained at 0.3 μm intervals, and represented as maximum intensity projections using the processing tool of the Zen software (Zeiss). All statistical analyses of the results used JMP software (SAS Institute).
Cathepsin D secretion assay
HeLa cells were transfected with the specified shRNA vectors. Cells were plated in six-well plates at a density of 2.5 × 106 cells/well 24 hours after transfection. The next day, medium was removed and cells were washed three times with serum-free medium and then grown in serum-free medium overnight. The experiment was performed 72 hours after transfection. Medium was collected and cells were harvested for whole cell lysates. Medium was filtered to remove debris (0.22 μm pore size; Millipore) and equal volumes were concentrated by centrifugation using a centrifugal filter (Ultracel-10K, Amicon Ultra, 3000 rpm, 45–90 minutes, 4°C). Whole cell lysates and medium were separated in a gradient gel (4–15%, Bio-Rad) and immunoblotted for cathepsin D. Densitometry analysis was performed using the ImageJ software.
We would like to thank Sabrice Guerrier for critical reading of the manuscript. This work was supported by NIH grants R01-AI065474 and R01-CA47752 to D.D.B., the Mayo Foundation and Allergic Diseases Training Grant NIH-T32-AI07047 (T.S.G.). D.D.B. is a Leukemia and Lymphoma Society Scholar. Deposited in PMC for release after 12 months.