Linking membrane microdomains to the cytoskeleton: Regulation of the lateral mobility of reggie-1/flotillin-2 by interaction with actin
Matthias F. Langhorst
*, Gonzalo P. Solis, Sylvia Hannbeck, Helmut Plattner, Claudia A.O. Stuermer
Department of Biology, University of Konstanz, Universitaetsstrasse 10, D-78457 Konstanz, Germany
Abstract The reggies/flotillins are oligomeric scaffolding pro- teins for membrane microdomains. We show here that reggie- 1/flotillin-2 microdomains are organized along cortical F-actin in several cell types. Interaction with F-actin is mediated by the SPFH domain as shown by in vivo co-localization and in vitro binding experiments. Reggie-1/flotillin-2 microdomains form independent of actin, but disruption or stabilization of the actin cytoskeleton modulate the lateral mobility of reggie-1/flo- tillin-2 as shown by FRAP. Furthermore, reggie/flotillin micro- domains can efficiently be immobilized by actin polymerisation, while exchange of reggie-1/flotillin-2 molecules between micro- domains is enhanced by actin disruption as shown by tracking of individual microdomains using TIRF microscopy.
Keywords: Reggie/flotillin; Actin; Membrane cytoskeleton;
Membrane microdomain
1. Introduction
It is generally accepted that the plasma membrane of eukaryotic cells is organized into microdomains conveying specificity to membrane-associated processes. Although the existence of membrane microdomains based solely on phase separation of lipids (‘‘lipid rafts’’) is presently under intense scrutiny [1], the existence of membrane microdomains orga- nized and scaffolded by oligomeric proteins like caveolins is indisputable [2]. Reggie-1 and -2 were discovered as proteins upregulated in retinal ganglion cells during axon regeneration after optic nerve injury [3]. They were independently described as proteins abundant in the ‘‘floating’’ detergent-resistant membrane fraction and therefore named flotillin-2 and -1, respectively [4]. The reggies/flotillins oligomerize at cellular membranes [5,6] to form clusters of approximately 50–
100 nm [7,8]. These clusters form the scaffold of specialized membrane microdomains implicated in insulin signalling [9], lymphocyte activation [10] and endocytosis of GPI-anchored proteins [11] (reviewed in [12]). It is generally assumed that plasma membrane microdomains are anchored to the actin cytoskeleton but experimental evidence is scarce. Only for cav-
eolae an indirect interaction of caveolin-1 with actin via filamin has been shown [13], for most other microdomains the anchor- ing mechanism is unclear. Here, we provide evidence for an interaction of reggie-1/flotillin-2 with F-actin, which is medi- ated by its SPFH domain and regulates its lateral mobility at the plasma membrane and thus positioning and local stabiliza- tion of reggie/flotillin microdomains.
2. Material and methods
2.1. Antibodies, plasmids and reagents, cells and transfection
Anti-actin polyclonal antibody (pAB) was from Santa Cruz (Santa Cruz, USA), anti-GST pAB from Amersham Pharmacia (Piscataway, USA), anti-reggie-1/flotillin-2 mAB from Transduction Labs (Heidel- berg, Germany), phalloidin-Alexa-568 from Molecular Probes (Lei- den, Netherlands), secondary antibodies coupled to HRP from Jackson ImmunoResearch (Soham, UK). Cytochalasin D and Jasplak- inolide were from Calbiochem (Bad Soden, Germany).a-Actin was from Sigma (Munich, Germany),b-actin from Cytoskeleton (Denver, USA). Enzymes for molecular biology were from New England Bio- labs (Beverly, USA) or Fermentas (St. Leon-Rot, Germany). Deletion mutants of reggie-1 were obtained by PCR using rat full-length reggie- 1 cloned in pEGFP-N1 (Clontech, Heidelberg, Germany) (R1FL- EGFP) as template[5] and primers introducing restriction sites for EcoRI and BamHI. Fragments were then subcloned into pEGFP- N1. N2a and HeLa cells were cultured and transfected as described previously[14].
2.2. Microscopy
For immunofluorescence, cells were grown on poly-LL-lysine coated coverslips (VWR, Darmstadt, Germany), fixed with 4% paraformalde- hyde, permeabilized by brief immersion in 0.1% Triton-X100 (Sigma) and incubated with phalloidin as indicated. For TIRF microscopy the TIRF slider system and aa-Plan Fluar 100·/1.45 objective from Carl Zeiss were used in combination with an Axiovert 200M and an Axiocam MRm (all Carl Zeiss). Images were analysed and processed using Axiovision 4.6 (Carl Zeiss) and ImageJ[15]. For tracking of indi- vidual microdomains, the feature point tracking algorithm[16]inte- grated in an ImageJ plugin was used[17]. FRAP experiments were carried out on a LSM 510 META using a 63·/1.4 Plan-Apochromat objective as described previously[10].
2.3. Electron microscopy
N2a cells were treated with 1lM Cytochalasin D as indicated and processed for immuno-gold electron microscopy as described previ- ously[7,18].
2.4. In vitro actin binding assay
DR1NT, the isolated SPFH domain of reggie-1/flotillin-2, was amplified by PCR using pR1FL-EGFP as template and subcloned into pQE31 (Qiagen) and pGEX (Amersham) for N-terminally tagging with a His6- and a GST-tag, respectively. The fusion protein was ex- pressed inEscherichia coliBL21(DE)pLysS and purified according to
*Corresponding author. Present address: Carl Zeiss MicroImaging GmbH, Koenigsallee 9-21, D-37081 Goettingen, Germany.
E-mail address:Matthiaslanghorst@email.de (M.F. Langhorst).
Konstanzer Online-Publikations-System (KOPS) URN: http://nbn-resolving.de/urn:nbn:de:bsz:352-opus-119206
the instructions provided by the vector’s manufacturers. Actin binding was assayed by co-precipitation with F-actin. Briefly,a- or b-actin stock solution in G-actin buffer (200lM CaCl2, 200lM ATP, 100lM DTT, 5 mM Tris, pH 8) were mixed with polymerisation buf- fer (100 mM KCl, 2 mM MgCl2, 1 mM EGTA, 1 mM ATP, 5 mM
Tris, pH 7.5) and incubated for 1 h at room temperature. The indicated amount of protein was added and incubated for 1 h at room tempera- ture. The mixture was centrifuged for 1 h at 100 000·gand 4C in a Beckmann Optima MaxE ultracentrifuge using a TLA 55 rotor. The supernatant was mixed with 5·sample buffer, the pellets were washed
Fig. 1. Association of reggie-1/flotillin-2 with F-actin is mediated by the SPFH domain. (A) Schematic representation of the reggie-1/flotillin-2 deletion mutants. (B, C) Confocal views of the basal plasma membrane of N2a cells: Plasma membrane microdomains demarcated by R1FL-EGFP closely associated with cortical F-actin (B), deletion of the C-terminal flotillin-domain did not abolish the association with F-actin (C), but deletion mutants lacking the SPFH domain (R1WTSH or R1MCT) still localized to the plasma membrane and did not form actin-associated clusters (D + E, all bars 2lm).
once in buffer, recentrifuged and solubilized in 2·sample buffer. Sam- ples were analysed by SDS-gel electrophoresis, Coomassie blue or Western blotting. Actin polymerisation was assayed using the Actin Polymerisation Biochem Kit (Cytoskeleton) and a Spectrofluor Plus 96-well plate reader (Tecan, Zurich, Switzerland) according to the manufacturer’s protocols.
3. Results and discussion
3.1. Reggie-1/flotillin-2 associates with filamentous actin in cells Reggie-1/flotillin-2 fused to EGFP (R1FL-EGFP) clearly co- localized with phalloidin-stained F-actin at the plasma mem- brane of N2a (Fig. 1B), HeLa, HepG2 and 3T3 cells (data not shown). Reggie-1/flotillin-2 microdomains at the plasma membrane were organized along actin filaments of the mem- brane cytoskeleton (Fig. 1B). Identical results were obtained with HA-tagged reggie-1/flotillin-2, and no co-localization of reggie-1/flotillin-2 with microtubuli or vimentin-type interme- diate filaments could be observed (data not shown). Endoge- nous reggie-1/flotillin-2 also co-localized with F-actin (Supplementary Fig. 1A), but to a small extent only. This dif-
ference is most probably caused by the incompatible fixation procedures for F-actin recovery and reggie-1/flotillin-2 stain- ing. Overexpressed reggie-1/flotillin-2, however, did no longer associate into microdomains, but localized along F-actin fibres (Supplementary Fig. 1B), corroborating the inherent ability of reggie-1/flotillin-2 to interact with F-actin. We tried to exclude fixation artefacts by live cell imaging of N2a cells expressing actin-mRFP and R1FL-EGFP, but the diffuse background caused by G-actin-mRFP excluded clear co-localization analy- sis (Supplementary Fig. 1C). Co-localization with F-actin was previously also shown for other members of the SPFH family, i.e. stomatin-1 [19] or podocin [20].
Using different deletion constructs we next identified the reg- gie-1/flotillin-2 domains involved in actin interaction. R1NT- EGFP comprising the SPFH domain (Fig. 1A) localized to the plasma membrane. Microdomain/cluster formation was, however, less pronounced as this construct lacks the flotillin domain known to mediate oligomerization [6]. R1NT-EGFP localized along F-actin fibres (Fig. 1C), demonstrating that the flotillin domain is dispensable for association with F-actin.
The membrane anchor region alone (aa 1–30) also localized to
Fig. 2. In vitro interaction of reggie-1/flotillin-2 with F-actin. (A) Schematic representation ofDR1NT representing the domain necessary for actin interaction. (B) Co-precipitation of 10lM GST-DR1NT with 5lM F-actin was carried out as detailed in Section 2. GST-DR1NT specifically precipitated with F-actin but not with G-actin or the respective buffer controls. A representative blot of five independent experiments is shown. (C) Polymerisation of pyrene-actin was monitored by fluorescence as detailed in Section2. Addition ofDR1NT had no significant effects on actin polymerisation. Addition of BSA or elution buffer was without effect (data not shown).
the plasma membrane (although it accumulated at the Golgi complex to some extent), but its distribution at the plasma membrane was diffuse and showed no correlation with the underlying membrane cytoskeleton (Fig. 1D). Similarly, R1MCT, a construct lacking the SPFH domain (Fig. 1A) showed diffuse localization at the plasma membrane without association with F-actin (Fig. 1E). Thus, the SPFH domain is apparently necessary for the association of reggie-1/flotil- lin-2 with F-actin at the plasma membrane.
3.2. In vitro binding of the SPFH domain of reggie-1/flotillin-2 to F-actin
To investigate whether the co-localization of R1FL-EGFP with F-actin observed in vivo is the result of a direct interac- tion of reggie-1/flotillin-2 with F-actin, we purified the minimal actin binding domain identified above (DR1NT, Fig. 2A) as a His- or GST-tagged protein recombinantly expressed in E. coli and tested its ability to bind to actin in vitro by co-precipita- tion with F-actin. Both the His-tagged and the GST-tagged
DR1NT, but not BSA as negative control (data not shown),co-precipitated specifically with filamentous
a- and b-actinbut not with G-actin or the respective buffer controls (Fig. 2B). Thus, the SPFH domain of reggie-1/flotillin-2 is capable of binding to F-actin. However,
DR1NT had no dis-cernible effect on the kinetics of in vitro actin polymerisation assayed by pyrene-actin fluorescence (Fig. 2C), suggesting that binding of reggie-1/flotillin-2 to F-actin mediates anchoring only. Most interestingly, actin-binding might be a general function of the SPFH-domain to which so far no general func- tion was assigned.
3.3. Reggie-1/flotillin-2 microdomains are formed by oligomerization rather than by binding to actin
To test whether reggie/flotillin microdomains are formed by interaction of reggie-1/flotillin-2 with the cortical actin cyto- skeleton, we treated N2a cells with Cytochalasin D to destroy
the actin cytoskeleton, which we verified by phalloidin-staining and fluorescence microscopy (data not shown). We then ana- lysed the localization of reggie-1/flotillin-2 in Cytochalasin D-treated vs. non-treated control cells by immuno-gold label- ling and electron microscopy. In both control and CytD-trea- ted cells, numerous reggie/flotillin clusters were observed (Fig. 3A) and no significant difference in number, size or fre- quency of reggie/flotillin microdomains (Fig. 3B) was ob- served, suggesting that reggie/flotillin microdomains are most probably formed by oligomerization of the reggies/flotillins rather than by clustering via binding to the cortical actin cyto- skeleton.
3.4. Interaction with F-actin regulates the lateral mobility of reggie-1/flotillin-2
Next, we investigated the influence of the actin cytoskeleton on the lateral mobility of reggie-1/flotillin-2 by fluorescence recovery after photobleaching (FRAP) in N2a cells. A mobile fraction of 0.83 ± 0.02 (n = 25, ±S.E.M.) and a half time of fluorescence recovery of only t
1/2= 16.2 ± 1.6 s indicated high lateral mobility of R1FL-EGFP in N2a cells (Fig. 4A) as pre- viously reported for PC12 and Jurkat cells [10]. Disruption of the actin cytoskeleton by Cytochalasin D treatment signifi- cantly decreased the half time of fluorescence recovery to t
1/2= 11.1 ± 1.5 s (n = 25), while having no significant influ- ence on the mobile fraction (M
F= 0.80 ± 0.02) (Fig. 4A and B). This suggests that a breakdown of the actin cytoskeleton increases the lateral mobility of reggie-1/flotillin-2. Incubation with Jasplakinolide, an actin stabilizing compound [21], signif- icantly increased the half time of fluorescence recovery to t
1/2= 20.2 ± 1.5 s (n = 30) (Fig. 4A and B). Thus actin stabil- ization reduced the lateral mobility of reggie-1/flotillin-2.
The reggies/flotillins are associated with the detergent-resis- tant membrane fraction. Therefore, we tested the role of cho- lesterol on the mobility of the reggies/flotillins. Cholesterol depletion using methyl-b-cyclodextrin led to a significant
Fig. 3. F-actin is not necessary for reggie-1/flotillin-2 cluster formation HeLa cells were treated with 1lM Cytochalasin D and processed for electron microscopy and immuno-gold labelling as detailed in Section2. (A) Representative images of reggie-1/flotillin-2 microdomains in Cytochalasin D- treated and untreated cells, 5 nm gold grains indicate reggie-1/flotillin-2 labelling (bars 0.1lm). (B) Quantitative comparison of immuno-gold labelling, showing no significant effect of Cytochalasin D-treatment on reggie/flotillin microdomains.
reduction of reggie-1/flotillin-2 mobility at the plasma mem- brane (Fig. 4B) as shown before for a variety of other mem- brane proteins [22], thus confirming the sensitivity and validity of our FRAP approach. Disruption of microtubules by nocodazole was without effect on reggie-1/flotillin-2 mobil- ity (Fig. 4B).
To test whether the observed effects of actin on reggie-1/flo- tillin-2-mobility were caused by molecular interactions between reggie-1/flotillin-2 and F-actin or by an indirect, general effect of an actin-anchored picket fence on membrane protein mobil- ity, we performed similar experiments with the deletion mutant R1MCT. R1MCT has the same membrane anchor as the wild- type protein, but lacks most of the SPFH domain which is necessary for actin interaction (Fig. 1A). FRAP experiments in untreated control cells showed a generally faster lateral diffusion of R1MCT compared with the full length protein (t
1/2= 6.9 ± 0.7 s, (n = 26)) (Fig. 4C and D). Jasplakinolide- treatment had no effect on the lateral mobility of R1MCT (t
1/2= 6.7 ± 0.7 s, (n = 30)) (Fig. 4C and D). Thus, R1MCT mobility is unaffected by F-actin stabilization, confirming that the lateral diffusion of full-length reggie-1/flotillin-2 is regu- lated by interaction of its SPFH domain with F-actin.
3.5. Reggie-1/flotillin-2 microdomains can be trapped by F-actin stabilization
The changes in reggie-1/flotillin-2 mobility observed in FRAP experiments might be caused by changes in mobility
of reggie-1/flotillin-2 microdomains at the plasma membrane or by modulation of exchange of individual molecules between microdomains. We therefore measured the mobility of reggie/
flotillin microdomains directly by total internal reflection fluo- rescence (TIRF) microscopy. We used HeLa cells in these experiments, because reggie-1/flotillin-2 microdomains are less densely spaced in these cells (Fig. 5A), so that individual microdomains can be unambiguously followed. The general lateral mobility of reggie-1/flotillin-2 and its dependence on ac- tin as observed by FRAP experiments in HeLa cells is similar to N2a cells (data not shown).
Reggie-1/flotillin-2 microdomains at the plasma membrane of HeLa cells exhibited a biphasic diffusional behaviour.
Phases of confined diffusion (Fig. 5B, pictures 1,2 and 4,5) were interrupted by phases of long range diffusion (Fig. 5B, pictures 3 and 6). This behaviour is reminiscent of the hop-dif- fusion in lateral confinement zones observed in single molecule studies [23].
Neither stabilization of the actin cytoskeleton by Jasplakino-
lide nor disruption by Cytochalasin D changed the steady state
number or appearance of reggie-1/flotillin-2 microdomains as
observed by TIRF microscopy, confirming our results by elec-
tron microscopy (Fig. 3). Disruption of F-actin by Cytochala-
sin D had no effect on the lateral mobility of reggie-1/flotillin-2
microdomain mobility. Diffusional behaviour was indistin-
guishable from untreated control cells (Fig. 5C), suggesting
that the increased mobility observed in FRAP experiments is
Fig. 4. F-actin regulates reggie-1/flotillin-2 mobility at the plasma membrane. (A) N2a cells were transfected with R1FL-EGFP and treated with 1lM Cytochalasin D or Jasplakinolide 24 h after transfection. Averaged FRAP tracings for 25–30 cells from three independent experiments are shown. (B) Summary of the effects of F-actin disruption (1lM Cytochalasin D,n= 25), F-actin stabilization (1lM Jasplakinolide,n= 30), cholesterol extraction (12 mM methyl-b-cyclodextrin (MbCD),n= 25) and microtubuli disruption (1lM nocodazole,n= 25) on the average half time of fluorescence recovery. F-Actin stabilization and cholesterol extraction decreased, while F-actin disruption increased R1FL-EGFP mobility, disruption of microtubules was without effect. (C and D) Cells were transfected with R1MCT and treated with 1lM Jasplakinolide, averaged tracings and average half times of fluorescence recovery are shown (n= 30, three independent experiments). Jasplakinolide-treatment was without effect on R1MCT mobility. *indicates significant differences (Student’st-test,P< 0.05), error bars indicate S.E.M.caused by increased exchange of individual molecules between clusters. This is further corroborated by the observation, that in Cytochalasin D-treated cells many tracks of individual microdomains were lost, because their fluorescent signal be- came diffuse, suggesting lateral dispersion of the molecules.
Thus, interaction with F-actin might regulate the exchange of reggie-1/flotillin-2 molecules and potentially proteins bound to reggie-1/flotillin-2 between individual microdomains.
Stabilization of the actin cytoskeleton by Jasplakinolide- treatment significantly decreased the lateral mobility of reggie-1/flotillin-2 microdomains. The mean displacement in Jasplakinolide-treated cells was significantly lower compared with untreated control cells (Fig. 5C). Many microdomains were almost completely immobilized and phases of long-range diffusion were very rare (Fig. 5D). Thus, by local stabilization of the cortical actin cytoskeleton, cells can efficiently immobi- lize reggie-1/flotillin-2 microdomains. We have previously described a dramatic immobilization of reggie/flotillin micro- domains in T lymphocytes, where many reggie/flotillin micro- domains are pre-assembled in a polarized macrodomain, which is stabilized by the actin cytoskeleton [10].
In summary, our results unravel the anchoring of reggie-1/
flotillin-2 microdomains to the cortical actin cytoskeleton by binding of its SPFH domain to F-actin. This mechanism may apply also to other membrane microdomains scaffolded by SPFH-proteins like stomatins, podocin, erlins or prohibitin.
This provides a mechanism ensuring spatial specificity in path- ways depending on these different microdomains, as cells can restrict and steer lateral diffusion of these microdomains sim- ply by regulation of actin dynamics.
Acknowledgement:This work was supported by Grants from the Deut- sche Forschungsgemeinschaft DFG (SFB-TR11), the Ministerium Forschung, Wissenschaft und Kunst Baden-Wu¨rttemberg (TSE pro- gram) and the Fonds der Chemischen Industrie. We gratefully acknowledge the help of Friederike Jaeger in reggie/flotillin co-locali- zation experiments.
Appendix A. Supplementary data
Supplementary data associated with this article can be found, in the online version, at doi:10.1016/j.febs- let.2007.08.074.
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