RESEARCH ARTICLE

The Ca2+-activated Cl− channel Ano1 controls microvilli lengthand membrane surface area in the oocyteRaphael Courjaret, Rawad Hodeify, Satanay Hubrack, Awab Ibrahim*, Maya Dib, Sahar Daas‡ andKhaled Machaca§

ABSTRACTCa2+-activated Cl− channels (CaCCs) play important physiologicalfunctions in epithelia and other tissues. In frog oocytes the CaCCAno1 regulates resting membrane potential and the block topolyspermy. Here, we show that Ano1 expression increases theoocyte surface, revealing a novel function for Ano1 in regulating cellmorphology. Confocal imaging shows that Ano1 increases microvillilength, which requires ERM-protein-dependent linkage to thecytoskeleton. A dominant-negative form of the ERM protein moesinprecludes the Ano1-dependent increase in membrane area.Furthermore, both full-length and the truncated dominant-negativeforms of moesin co-localize with Ano1 to the microvilli, and the twoproteins co-immunoprecipitate. The Ano1–moesin interaction limitsAno1 lateral membrane mobility and contributes to microvilliscaffolding, therefore stabilizing larger membrane structures.Collectively, these results reveal a newly identified role for Ano1 inshaping the plasma membrane during oogenesis, with broadimplications for the regulation of microvilli in epithelia.

KEY WORDS: Ano1, Microvilli, ERM, Xenopus oocyte, Moesin

INTRODUCTIONOogenesis encompasses the growth phase of vertebrate oocytesduring which time they accumulate macromolecular complexesessential for supporting early embryogenesis (Machaca, 2009). Thisis followed by a maturation phase before oocytes acquire thecompetency to be fertilized. Oocyte maturation encompasses thereductionist meiotic division in addition to cytoplasmic remodelingthat prepares the mature egg for fertilization and the egg-to-embryotransition. In the frog oocyte, maturation is initiated in response tosteroid hormones that induce a commitment to progressing throughmeiosis, until the process is arrested again at metaphase of meiosis IIup to fertilization (Machaca, 2009; Nader et al., 2013). An importantcomponent of oocyte maturation is the remodeling of the Ca2+

signaling machinery to support a specialized Ca2+ transient atfertilization (Machaca, 2009; Nader et al., 2013). The Ca2+ signal atfertilization possesses specialized spatial and temporal dynamicsthat encode cellular processes, such as the block to polyspermy andthe completion of meiosis, in a sequential fashion to mediate the

egg-to-embryo transition. Ca2+ is in fact the universal signal for eggactivation at fertilization in all sexually reproducing speciesinvestigated to date, although the fertilization-specific Ca2+

transient takes on different spatio-temporal dynamics in a species-specific fashion (Stricker, 1999).

Frogs have evolved two complimentary, but mechanisticallydistinct, Ca2+-dependent mechanisms to efficiently blockpolyspermy given their large egg size. The fast block topolyspermy occurs almost immediately after fertilization, andresults from the stimulation of Ca2+-activated Cl− channels(CaCCs) leading to membrane depolarization, which precludesfurther sperm–egg fusion (Cross and Elinson, 1980; Jaffe et al.,1983). On a slower time scale Ca2+ induces the fusion of corticalgranules, which release enzymes that modify the egg’s extracellularmatrix and mediate the more sustained slow-block to polyspermy(Grey et al., 1974; Wolf, 1974).

CaCCs were first described in the Xenopus oocyte as regulators ofthe resting membrane potential, in addition to their role in the blockto polyspermy (Duran and Hartzell, 2011; Machaca et al., 2001).CaCCs represent the primary current in the oocyte and theirbiophysical properties have been well characterized (Hartzell et al.,2005; Kuruma and Hartzell, 2000). CaCCs are broadly distributedamong different species from invertebrates to mammals, and acrossdiverse tissues in vertebrates (Kidd and Thorn, 2000; Robertson andMartin, 1996). Functionally CaCCs regulate multiple physiologicalprocesses, including epithelial secretion, vision, olfaction, vasculartone and neuronal excitability (Hartzell et al., 2005; Machaca et al.,2001). Our understanding of CaCCs was hampered for many yearsby controversy regarding their molecular identity; however, in 2008,the Xenopus oocyte CaCC was identified as TMEM16A usingan expression cloning approach (Schroeder et al., 2008), andknockdown of endogenous Xenopus TMEM16A was shown toreduce native Ca2+-activated Cl− current (Yang et al., 2008). Giventhat TMEM16A is an anion channel with eight transmembranedomains, Yang et al. renamed the protein anoctamin 1, or Ano1 (foranion+octa) (Yang et al., 2008). Mammalian genomes encode 10anoctamin homologs. Ano1 is widely expressed in various epitheliawhere it is important for Cl− secretion, as illustrated by secretiondefects in Ano1 knockout mice (Ousingsawat et al., 2009). Inaddition to its role as a secretory channel in epithelia, Ano1 has alsobeen shown to play important roles in gut smooth muscle, airwaysmooth muscle, nociception, the biogenesis of the primary cilium,and interestingly, Ano1 expression has been shown to correlate withcancer, providing a good biomarker for some cancers (Duran andHartzell, 2011; Ruppersburg and Hartzell, 2014).

Here, we describe a newly identified function of Ano1 that isindependent from its channel activity. We show that Ano1 regulatesthe length and diameter of microvilli in Xenopus oocytes through itsinteraction with ERM proteins and the actin cytoskeleton. Thenumber of microvilli in the oocyte has been estimated to beReceived 18 February 2016; Accepted 2 May 2016

Department of Physiology and Biophysics, Weill Cornell Medical College in Qatar,Education City – Qatar Foundation, Luqta Street, PO Box 24144, Doha 24144,Qatar.*Present address: University of South Alabama, Department of Pediatrics, 1700Center Street, Mobile, AL 36604, USA. ‡Present address: Qatar CardiovascularResearch Center, Qatar Science and Technology Park, Doha 26999, Qatar.

§Author for correspondence (Khm2002@qatar-med.cornell.edu)

K.M., 0000-0001-6215-2411

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≈40×106, and they expand the surface of the oocyte three- tofourfold (Kado et al., 1981). The structure of a microvillus has beenextensively described in the intestinal brush border (Brown andMcKnight, 2010; Crawley et al., 2014). Microvilli are shapedaround parallel bundles of actin fibers that stabilize the thin, highlycurved microvillus membrane against significant physical forcesincluding surface tension, bending stiffness and membrane–cytoskeleton adhesion energy (Crawley et al., 2014; Nambiaret al., 2010). Single actin filaments are insufficient to overcomethese forces, generating the need for actin bundles that are cross-linked and stabilized by a family of actin-binding proteins, includingvillin, fimbrin and espin (Crawley et al., 2014). In turn, actinbundles are physically linked to integral plasma membrane proteinsand membrane lipids through two primary classes of proteins, theunconventional myosins and ERM proteins. The ERM (ezrin,radixin andmoesin) family of proteins share 70-80% identity amongthe different members, and are highly conserved throughout theanimal kingdom (Arpin et al., 2011). The ERM proteins bindmembrane lipids such as PIP2 and also connect the cytoskeleton toa wide variety of transmembrane proteins such as ions channels(e.g. CFTR) (Guerra et al., 2005), G-protein coupled receptors (e.g.κ-opioid receptor and β2-adrenergic receptor) (Heydorn et al., 2004;Li et al., 2002), as well as adhesion proteins (e.g. ICAM1, CD44)(Bretscher et al., 2002). The ERM proteins share a commonstructure; the N-terminal (Nter) domain [termed FERM (4.1 proteinand ERM) or N-ERMAD (ERM associated domain)] is composedof three lobes that mediate ERM binding to the plasma membrane.The C-terminal, or C-ERMAD, domain contains the binding site foractin (see Fig. 3A). At rest the protein is folded and the C-ERMADand FERMdomains are in contact, resulting in auto-inhibition of theERM crosslinking function. ERM proteins need to unfold to

unmask the functional sites allowing binding to their specificpartners, a process promoted by phosphorylation of a threonineresidue in the C-ERMAD domain (Fehon et al., 2010). Theinteraction of the ERM protein with its target at the plasmamembrane involves either direct binding to the cytoplasmic portionof membrane proteins, or requires linker proteins such as EBP50(ERM-binding phosphoprotein 50) (also known as NHERF; Na-H+

exchanger regulatory factor) or E3KARP (NHE type 3 kinaseregulatory protein) that contain a PDZ domain (Fehon et al., 2010).

Data presented herein reveal a newly discovered function of Ano1that is independent of channel activity in modulating the structure ofmicrovilli. Ano1 supports microvilli elongation through itsinteraction with ERM in the oocyte. This results in an expansionof membrane surface area as a function of Ano1 expression. Hence,Ano1 plays a crucial role in regulating membrane surface area in theoocyte by physically linking to the cytoskeleton through ERMproteins. Oocyte maturation is associated with the loss of microvilliresulting from endocytosis, a process that is crucial for thebiogenesis of the first polarized epithelium leading to blastocoelformation.

RESULTSAno1 overexpression increases membrane capacitanceThe effect of expression of Xenopus laevis Ano1 (10 ng RNA) onCaCC currents in stage VI oocytes was evaluated using a voltagejump from the resting membrane potential of −30 mV to +40 mV(Fig. 1A). There was a significant increase in the resting currentrecorded at +40 mV in cells expressing Ano1 (Fig. 1B), that wasassociated with a decrease in membrane resistance (Fig. 1C) and asmall but significant depolarization (Fig. 1D). At the restingmembrane potential [i.e. −35.8±0.8 mV (mean±s.e.m.), n=42] the

Fig. 1. Overexpression of Ano1 in Xenopus oocytes increasesmembrane capacitance. Various physiological parameters were recorded on uninjected cells(naïve) and cells expressing Ano1–mCherry (Ano1). (A) Chloride currents recorded at rest (left traces) and during store depletion induced by ionomycin (Ion;10 µM, right traces) following a voltage jump from −30 mV to +40 mV. (B) Amplitude of the resting chloride currents (ICl1) recorded at +40 mV. (C) Membraneresistance (Rm). (D) Membrane potential (Em). (E) Bar chart summarizing the amplitude of the current (ICl1) following mobilization of store Ca2+ with ionomycin innaïve and Ano1-expressing cells. (F) Increase in the membrane capacitance of oocyte expressing Ano1. (G) Overexpression of the Ca2+ channel Orai1 does notincrease cell capacitance (Cm). Statistics were performed using Student’s t-test; *P<0.05; ***P<0.001; ns, not significant. The number of cells tested (n) isindicated over the bars or in the plotting area when identical. Data in B–G represented as means±s.e.m.

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driving force for chloride ions is minimal, with the reversal potentialfor Cl− (ECl) in our recording conditions being between −25 mVand −30 mV (Barish, 1983; Costa et al., 1989). Furthermore, thecurrent through CaCC in the Xenopus oocyte displays a strongoutward rectification at depolarizing potential and low Ca2+

concentration. Taken together, this explains the moderate effect ofAno1 expression on the membrane potential.To increase intracellular Ca2+ and stimulate Ano1, we mobilized

intracellular Ca2+ stores using ionomycin, and recorded Cl− currentsat the peak of the Ca2+ release phase (Courjaret andMachaca, 2014).As illustrated in Fig. 1A and E, the current recorded at +40 mV wasgreatly increased in response to intracellular Ca2+ release in Ano1-expressing cells. This indicates that the over-expressed Ano1increases the Ca2+-activated chloride currents in the oocyte withsimilar properties to the endogenous CaCC, as previously reportedin axolotl oocytes (Schroeder et al., 2008). Strikingly, however,these expected changes in membrane conductance propertiesfollowing Ano1 expression were associated with a significantincrease in the membrane capacitance (from 230.6±4.4 nF to311.8±9.2 nF; Fig. 1F). This increase in membrane surface area isspecific to Ano1, as overexpression of another ion channel, Orai1,does not affect membrane capacitance (Fig. 1G).

Ano1 expression changes the structure of the microvilliindependent of channel functionThe theoretical surface area of a smooth oocyte of 1.2∼1.3 mm indiameter would result in a cell capacitance of 45–55 nF (with aconversion factor of 1 µF/cm2; Hille, 1992). This value is close tothe measured capacitance of Xenopus eggs where the microvillihave been lost during oocyte maturation. When the stage VI oocytematures, a significant portion of the membrane area in microvilli isinternalized, leading to a loss of microvilli and flattening of the cellmembrane (Kado et al., 1981). Whereas the apparent diameter of thecell does not change during oocyte maturation, the cell capacitancedrops from 224.0±4.5 nF (n=51) to 74.3±2.8 nF (n=67) (see alsoKado et al., 1981; Machaca and Haun, 2000; Peres and Bernardini,1985). This indicates that the microvilli surface accounts for themajority of the cell capacitance.We imaged oocyte microvilli and evaluated their dimensions

under different conditions. We first measured the size and shape ofmicrovilli on the animal pole of naïve cells using wheat germagglutinin to stain the plasma membrane of live oocytes (El-Jouniet al., 2008). Cells were cooled to 4°C during the staining process tolimit vesicular trafficking (Fig. 2A). This was then compared withmembrane structure in cells expressing mCherry-tagged Ano1 byimaging the red fluorescence of mCherry. The first strikingobservation was the presence of very long microvilli and, onsome cells, of large vesicular-shaped structures consisting of broadmembrane extensions of shorter length, similar to the length ofmicrovilli in control cells (Fig. 2A, arrowheads). These structuresare likely fused microvilli. Both features were absent in naïve cells(Fig. 2A). We first quantified the maximum microvillus length innaïve cells, Ano1-expressing cells, and cells expressing the Ca2+

channel Orai1–GFP. The length of the 10 longer microvilli wasmeasured in a single orthogonal section for each cell and averaged.There was a significant increase in the maximum microvillus lengthwhen Ano1–mCherry was expressed but not with Orai1–GFP(Fig. 2B). Additionally, the maximum microvillus diameter wasalso increased in Ano1-expressing, but not Orai1-expressing, cells(Fig. 2C). To obtain an estimate of the average increase in the lengthof the microvilli resulting from Ano1 expression, we analyzedconfocal z-stacks by plotting the intensity of the fluorescent signal

across orthogonal sections from the membrane intracellularboundary to the tip of the microvilli (Fig. 2D–F). The width ofthe curves at half-maximum intensity provides a good estimate forthe average microvillus length, which was significantly increased inAno1-expressing cells (Fig. 2F). In addition, we used a meshreconstruction algorithm of the oocyte surface (Fig. 2G) that allowsthe quantification of the actual membrane surface. In naïve cells theestimated actual membrane surface was 2880±260 µm2 (n=17) for a400 µm2 section, indicating that the convolutions increase themembrane surface by ∼7.2× at the animal pole of naïve cells. Thisvalue is larger than the theoretical value of an ∼fivefold increase forthe whole cell but this probably results from the uneven distributionof the microvilli and their higher density at the animal pole, and thatthe measure of the surface is highly sensitive to the meshparameters. In cells expressing Ano1 the increase factor reached∼10.3× (Fig. 2H). The mesh analysis suggests that the increasedlength of microvilli in Ano1-expressing cells is associated with adecrease in their density. To obtain an estimate of microvilli densityin control versus Ano1-expressing cells, we used the averagemicrovilli length and maximal diameter measured as discussedabove to calculate microvilli density for a theoretical oocyte of1.25 mm in diameter (surface area of 4.9 mm2, equivalent to 49 nFat 1 µF/cm2). The measured surface area of a control oocyte is∼23 mm2 (230 nF), yielding a total microvilli surface area of18.1 mm2. In control cells the average microvillus length is 1.74 μmwith a maximum diameter of 0.42 μm. If we model the microvillusas a cone, this results in an individual microvillus surface area of1.29 µm2, and a total number of ∼14×106 (low estimate) with adensity of 2.9×106 villi/mm2. Ano1-expressing cells have a surfacearea of 31.2 mm2 (312 nF) with an average microvillus length of2.87 µm and a maximum diameter of 0.91 µm. This gives amicrovillus surface area of 4.8 µm2 and a total of 5.5×106 microvilliwith a density of 1.1×106 villi/mm2. This represents a two- tothreefold reduction in microvilli density following Ano1expression. These calculations are consistent with the imagingdata (Fig. 2A), the mesh analysis (Fig. 2G) and the dome-likestructures, which could result from several microvilli fusingtogether (Fig. 2A, arrowheads).

To assess whether Ano1 Cl− channel function is required for itsrole in modulating microvilli length, we blocked channel functionusing the Ano1 inhibitor MONNA (Oh et al., 2013) during theexpression of Ano1. MONNA (1 µM) effectively blocked theresting CaCC current in both naïve oocyte (Fig. 3A), and in oocytesexpressing Ano1 (Fig. 3B). However, despite the fact that MONNAblocked the CaCC current it had no effect on the increasedcapacitance observed following Ano1 expression (Fig. 3C). This isalso visible in the confocal images recorded on cells overexpressingAno1 where no change in the microvilli density or structure isvisible (Fig. 3D). Collectively, these results support the conclusionthat Ano1 increases the membrane capacitance by increasing thelength and diameter of microvilli while decreasing their overalldensity and that this effect does not require the activation of thechannel.

Expression of the ERM protein, moesin, modulates oocytesurface areaPerez-Cornejo et al. comprehensively defined the Ano1interactome, and identified the ERM protein family as robustAno1-binding partners, thus providing a molecular link between theplasma membrane and the cytoskeleton (Perez-Cornejo et al.,2012). They also found that the interaction between Ano1 andmoesin influences the presence of both proteins at the plasma

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membrane. As discussed in the Introduction the microvillusmembrane associates with the actin cytoskeleton, allowing it toovercome the physical forces against its bending to fit the structureof the microvillus. The observation that Ano1 expression increasesmicrovilli length suggests that Ano1 links the plasma membrane tothe cytoskeleton through ERMs, thus supporting the increasedmicrovillus length and area. To test whether ERM proteins providethe molecular link between Ano1 and the cytoskeleton within themicrovilli, we expressed different human moesin constructs andtested the effect on membrane surface area. We chose moesin as theERM protein as it is known to be expressed in Xenopus oocytes(Thorn et al., 1999). Human and Xenopusmoesin share 88% overall

identity, reaching 97% in the functional FERM domain and 100% inthe F-actin binding site (Fig. 4A; Fig. S1). Cells were injected withcRNAs coding for three different proteins; full-length moesin (FL-moesin), a truncated form containing the FERM domain (the first381 residues; Nter-moesin), and a protein encoding the F-actin-binding domain (382 to C-terminal end; Cter-moesin) (Fig. 4A)(Amieva et al., 1999). Oocytes tolerated the expression of FL-moesin well; however, prolonged overexpression of Nter-moesinlead to depigmentation of the animal pole and cell death at longertime points (Fig. 4B). Cter-moesin was poorly expressed (Fig. 6).Microvilli were visualized using wheat germ agglutinin (WGA)staining and orthogonal reconstruction of confocal z-stacks. There

Fig. 2. Microvilli structure followingoverexpression of Ano1 in Xenopus oocytes.(A) Confocal imaging was performed across theplasma membrane of naïve cells stained with wheatgerm agglutinin (WGA) or expressing Ano1–mCherry (Ano1). Two confocal planes areillustrated; one at the very top of the microvilli (Tip)and one close to the base (Base). The lower panel(Ortho) presents orthogonal projections across theconfocal z-stack. There is a pronounced increase inmicrovilli length, as well as the appearance of dome-like structures (white arrowheads) presumablyresulting from microvilli fusion. (B) The maximumlength of 10 microvilli was averaged on each cell. Athird data set obtained from cells expressing theOrai1–GFP protein was used as an additionalcontrol. Ano1 expression specifically extended themaximum length of the microvilli. (C) The maximumdiameter of the microvilli identified at the ‘base’plane was measured in the same cells as in B andreveals an increase in Ano1-expressing cells.(D) Average microvillus length was evaluated usingthe fluorescence intensity across a z-stack, the baseof the plasma membrane is used as a verticalreference. (E) The intensity measured in D is plottedas a function of the z-axis illustrating microvilliextension. (F) The thickness of microvilli was thenevaluated as the width of the curve in E at half-maximal intensity (dotted line). (G) The oocytesurface at the animal polewas reconstructed using amesh technique (see Materials and Methods).(H) Histogram summarizing the increase in surfaceinduced by the membrane convolutions. Statisticswere performed using either Student’s t-test orANOVA; **P<0.01; ***P<0.001; ns, not significant.Data in B,C,F,H represented as means±s.e.m.

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was no obvious effect of FL-moesin and Cter-moesin expression onthe plasma membrane (Fig. 4B). For cells expressing Nter-moesin,membrane structure depended on the extent of the discoloration ofthe animal pole of the oocyte. In non-discolored cells, somemicrovilli were longer, although patches of the membrane lackedmicrovilli (Fig. S2). When the cells were discolored the membranebecame more chaotic, showing large round-shaped structuresstained by WGA and no visible microvilli (Fig. S2). Therefore,

for the rest of the experiments with Nter-moesin, analysis waslimited to cells with no discoloration on the animal pole to avoidnon-specific toxicity effects. The cell capacitance was significantlyincreased following expression of FL-moesin (Fig. 4C). In contrast,expression of the N-terminal truncation resulted in a reduction ofmembrane capacitance (Fig. 4C). The increase in membrane areafollowing FL-moesin expression is likely a result of its crosslinkingfunction between the cytoskeleton and plasma membrane. The

Fig. 3. The increase in membrane area does notrequire Ano1 Cl− channel function. Naïve cellsand cells expressing Ano1 where incubated withthe Ano1 inhibitor MONNA (1 µM) for the durationof the experiment. (A,B) The resting chloridecurrent (ICl1) was significantly reduced by MONNAin both naïve (A) and Ano1-expressingcells (B). (C) MONNA incubation had no effect onthe membrane capacitance (Cm) of naïve cells nordid it influence the increase in Cm recordedfollowing Ano1 expression. (D) Confocal imaging ofthe Ano1–mCherry protein across the plasmamembrane reveals no change in the microvillistructure in cells incubated in MONNA. Statisticswere performed using either Student’s t-test (A,B)or ANOVA (C); **P<0.01; ***P<0.001; ns, notsignificant. Data in A,B (right panels) and Crepresented as means±s.e.m.

Fig. 4. Expression of full-length and truncatedmoesin in Xenopus oocytes. (A) Cartoonrepresenting the structure of the ERM protein moesin.The FERM domain contains the binding site to plasmamembrane proteins whereas the C-terminal end(C-ERMAD) binds actin. The arrow marked ‘381’indicates the position of the separation between theNter- and Cter-moesin fragments used in this study.(B) Photographs of oocytes expressing the differentmoesin proteins (top panels). Prolonged expression ofNter-moesin induces a discoloration of the animal pole.The lower panels are orthogonal sections of the sameooctyes obtained from confocal z-stacks ofrepresentative cells stained with WGA.(C) Capacitance (Cm) of oocytes injected with thedifferent moesin constructs. The number of cells used(n) is indicated in parentheses above the bars.Statistics were performed using ANOVA; ***P<0.001;ns, not significant. (D) Dose dependence of themembrane capacitance (Cm) upon injection of differentamounts of cRNA coding for Nter-moesin. Data in C,Drepresented as means±s.e.m.

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decrease in membrane capacitance correlated with the levels ofexpression of Nter-moesin in a dose-dependent fashion (Fig. 4D).To better control the expression levels of the different moesin

truncations, we further normalized for the cRNA copy numberinjected per oocyte, based on the size of the different constructs(5 ng full length, 4 ng Nter-moesin and 3 ng Cter-moesin). Theresults were comparable with those obtained on injection of 10 ng ofcRNA; a slight increase in the capacitance for the expression of theFL-moesin and a reduction for the Nter-moesin (Fig. S3A). Therewas no significant change in the membrane potential or in themembrane resistance (Fig. S3B,C). Importantly, expression of thedifferent moesin constructs did not significantly alter the levels ofthe CaCC current at rest or after mobilization of store Ca2+ storeswith ionomycin (Fig. S3D,E). This indicates that expression of FL-moesin, Nter-moesin or Cter-moesin does not significantly alter themembrane residence of Ano1, and that the observed decrease inmembrane capacitance in Nter-moesin-expressing cells results froma disruption of the molecular link between the cytoskeleton and theplasma membrane. The absence of any phenotype with Cter-moesinexpression likely results from the poor expression of this constructas shown below.

Moesin interacts with Ano1We next co-expressed Ano1–mCherry with FL-moesin and itsfragments to evaluate their co-localization and influence on Ano1expression and function. We did not observe any synergistic effectof co-expression of Ano1 with FL-moesin or Cter-moesin, with bothresulting in the same level of enhancement of membranecapacitance as observed with expression of Ano1 alone (Fig. 5B).In contrast, co-expression of Nter-moesin with Ano1 reverses theenhancement of membrane surface area observed following Ano1expression (Fig. 5A,B). We interpret this result to indicate that Nter-moesin saturates the new Ano1 ERM binding sites generatedfollowing Ano1 overexpression, and as such inhibits microvilli

elongation and the associated increase in membrane surface area.However, another potential explanation is that moesin is somewhatinvolved in the trafficking of Ano1 to the plasma membrane andwith the co-expression of the dominant-negative Nter-moesin,trafficking is inhibited leading to a block of the increase insurface area. However, this is not the case, as revealed bymeasurements of the amplitude (Fig. 5C) and density (Fig. 5D) ofthe CaCC current (measured at the peak of the ionomycin-inducedCa2+ release event). Co-expression of Nter-moesin did not reduce theamplitude (Fig. 5C) or density (Fig. 5D) of the current, showing thatoverexpressed Ano1 reaches the plasma membrane to the sameextent whether expressed alone or co-expressed with Nter-moesin.

We further investigated the interaction between Ano1 and moesinin vivo using co-localization and immunoprecipitation. Thedifferent moesin constructs used above were tagged with GFP andco-expressed with Ano1–mCherry to evaluate their respectivesubcellular localization. When Nter-moesin–GFP was expressed thediscoloration of the cell and reduction of the microvilli could beobserved only after substantially longer periods of expression (3 to5 days as opposed to 24 to 48 h in the case of untagged Nter-moesin), although the fluorescence was clearly observed earlier. Wetherefore used cells with clear expression of Nter-moesin but withoutany discoloration for our analysis. In the case of the Cter-moesin–GFP, the fluorescence of the GFP could be detected on some cellsbut was very faint or absent in most of them (Fig. 6A). The co-localization of the Ano1 and moesin was then quantified usingorthogonal projections from z-stacks (Fig. 6A). Two values wereused as a reference; for maximum co-localization we measured thevalues obtained from an experiment where Ano1–mCherry-expressing cells were stained with WGA. For minimum co-localization (exclusion) we used the Pearson’s coefficientobtained from two proteins that are not located in the samecellular compartment, the Ca2+ sensor STIM1 (located in theendoplasmic reticulum) and the plasma membrane Ca2+ channel

Fig. 5. Co-expression of moesin and Ano1. Oocytes wereinjected with Ano1–mCherry alone or with either FL-moesin,Nter-moesin or Cter-moesin. (A) Confocal images across themicrovilli and orthogonal reconstruction of z-stacks of cellsexpressing Ano1–mCherry with or without Nter-moesin.(B) Capacitance (Cm) of oocytes expressing Ano1–mCherryand the three different forms of moesin, only the N-terminalfragment influences the capacitance. (C) Maximum Cl−

current (ICl1) recorded on cells expressing Ano1–mCherry andmoesin constructs following store Ca2+ mobilization withionomycin. (D) Cl− current densities obtained on cellsexpressing Ano1 and the moesin constructs. Statistics wereperformed using ANOVA; *P<0.05; **P<0.01; ***P<0.001.Data in B–D represented as means±s.e.m.

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Orai1 (Fig. 6B, S/O) (Courjaret and Machaca, 2014). There was ahigh degree of co-localization between Ano1 and either FL-moesinor Nter-moesin (Fig. 6B). For the Cter-moesin the co-localizationanalysis was limited only to selected cells that expressed asignificant GFP signal and showed reduced co-localization(Fig. 6B); however, this could be caused by poor expression ofCter-moesin in the oocyte. Nonetheless, these results indicate thatmoesin is located at the plasma membrane and is bound through its

N-terminal domain to its target. Next, to confirm that one of thetargets of moesin in the plasma membrane is Ano1, we tested theability of both proteins to co-immunoprecipitate. The experimentswere performed on cells expressing the GFP-tagged moesinconstructs. As illustrated in Fig. 6C, immunoprecipitation of FL-moesin–GFP or Nter-moesin–GFP pulls down Ano1, revealing asingle immunoreactive band close to the theoretical value of113 kDa for Ano1. These results show that Ano1 interacts withmoesin in vivo through its N-terminal domain leading to co-localization of both proteins to microvilli.

To further support the finding that moesin interacts with Ano1,we next performed FRAP experiments in order to measure Ano1diffusion in the plane of the membrane (seeMaterials andMethods).We used cells expressing Ano1–mCherry with or without untaggedmoesin proteins and bleached a selected membrane area whilemonitoring the rate of fluorescence recovery over time (Fig. 7A,B).Proteins responsible for the recovery of fluorescence afterphotobleaching can originate from the cytoplasm (fast diffusion)and from the plasma membrane (slower) (Goehring et al., 2010). Inthis case, the bleaching was performed using a line scan over the

Fig. 6. Co-localization and co-immunoprecipitation of Ano1 and moesin.(A) Orthogonal sections through z-stacks of confocal images from oocytesco-expressing Ano1–mCherry with GFP-tagged FL-moesin (wild type), Nter-moesin or Cter-moesin as indicated above the section and in the cartoonrepresentation of the constructs. The overlay image (merge) shows that bothFL-moesin and Nter-moesin co-localize with Ano1, whereas the Cter-moesinexpression levels are low and do not co-localize. (B) The co-localization levelwas assessed using Pearson’s coefficient on orthogonal sections. The level ofco-localization for STIM1 (ER) and Orai1 (plasma membrane) (S/O) was usedas a reference ‘minimal’ value, and the co-localization of WGA with Ano1–mCherry as a ‘maximal’ value (WGA, see text for details). FL-moesin and Nter-moesin, but not Cter-moesin, show strong co-localization with Ano1. Statisticswere performed using ANOVA; ***P<0.001. Data represented as means±s.e.m. (C) GFP-tagged FL-moesin and Nter-moesin co-immunoprecipitate withendogenous Ano1. Lysates from oocytes expressing the different moesinconstructs as indicated were western blotted and probed with either anti-Ano1or anti-GFP antibodies before (Input) and after (IP Anti-GFP) pull-down usingan anti-GFP antibody.

Fig. 7. Fluorescence recovery after photobleaching. FRAP experimentswere performed using a line scan over the z-axis to obtain fast orthogonalplanes (see Materials and Methods). (A) A defined area (dotted rectangle) wasbleached using increased laser power over a zoomed area, and the recoverymonitored for several minutes in cells expressing Ano1–mCherry alone or incombination with Nter-moesin. (B) Timeline of recovery from bleaching whenthe different Nter-moesin, Cter-moesin and FL-moesin constructs are co-expressed. (C) Bar chart summarizing the half-time of recovery of thefluorescence when Ano1–mCherry is co-expressed with various untaggedmoesin proteins. Statistics are performed using ANOVA; *P<0.05; **P<0.01;***P<0.001; ns, not significant. Data in C represented as means±s.e.m.

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z-axis and consequently, the intracellular pool of proteins under themembrane is bleached as well. The existence of a substantiveintracellular Ano1 pool ready to fuse with the plasma membrane isalso unlikely as, even under conditions of Ano1 overexpression, nodetectable vesicular Ano1-positive fluorescence is visible (Fig. 6A).Consequently, most of the fluorescence recovery afterphotobleaching is likely to occur from lateral diffusion in theplane of the plasma membrane. There was a significant increase inthe rate of recovery in cells expressing the Nter-moesin but no majorchange for cell expressing the full-length or the C-terminal fragment(Fig. 7B,C). This indicates that the linkage of Ano1 to thecytoskeleton through ERM reduces its mobility in the membraneand is disrupted when the Nter-moesin competes for the Ano1binding site.To confirm the role of ERM–Ano1 interactions in mediating the

increase in membrane surface area following Ano1 expression, wesought to express different fragments of Ano1 that would act asbinding sites for ERM. Given that the Ano1 binding site for ERMproteins is not known, we expressed the large cytoplasmicN-terminal fragment of Ano1 (amino acids 1–322) and a longerfragment that ends just before the presumed third transmembranedomain of Ano1 (Fig. S4). Expression of the 1–322 fragment ofAno1 effectively reduced membrane capacitance (Fig. S4C),without affecting CaCC current (Fig. S4B). This supports the roleof ERM–Ano1 interactions in modulating membrane surface area.

DISCUSSIONTo increase the exchange capacity of a cell while limiting volumevariations (i.e. increase the surface/volume ratio) epithelial cells,oocytes, lymphocytes, hepatocytes and placental cells grow thinextensions termed microvilli. They extend the membrane surfaceto facilitate exchanges (absorption and secretion), and dependingon the cell type, can also serve mechanical functions, includingphysical support and mechanotransduction. Densely packedmicrovilli form a brush border extending the surface area ofepithelial cells such as the ones lining the small intestine and theproximal tubule of the kidney by ∼tenfold (Crawley et al., 2014;Helander and Fändriks, 2014). Xenopus oocytes are covered withmicrovilli that vary in shape and density during the growth anddevelopment of the gamete. This is particularly obvious followingoocyte maturation in preparation for fertilization. During thatprocess the number of microvilli is drastically and rapidly reduced(2000/s), inducing a large drop in the membrane capacitance in afew hours, without any change in the apparent diameter of theoocyte (Kado et al., 1981; Machaca and Haun, 2000). This rapidinternalization of membranes forming the microvilli generates anintracellular vesicular pool that is essential for the rapidcellularization during early embryogenesis (Müller and Hausen,1995). Furthermore, this membrane internalization forms the initialstep in the biogenesis of the first polarized epithelium in the embryothat supports vectorial transport of Na+ and water into the blastocoel(Muller, 2001). This is because endocytosis during oocytematuration is associated with a selective internalization ofmembrane proteins resulting in two populations of membranes;the egg plasma membrane, which represents the future apicalmembrane of the epithelium, and the internalized vesicular pool,which represents the future basolateral portion, with distinct proteincontent. For example, Orai1 and PMCA are internalized duringoocyte maturation but not Ano1 (El-Jouni et al., 2008; Yu et al.,2009; Yu et al., 2010).Here we show that Ano1 regulates microvilli length and, thereby,

membrane surface area, independently of its channel function. Ano1

expression increases the oocyte surface area andmicrovilli length anddiameter (Figs 1 and 2). The Ano1-dependent effect on microvilli ismediated through ERM proteins, as co-expression of the dominant-negative N-terminal domain of moesin with Ano1 abolishes theAno1-dependent increase in microvilli length and surface area(Fig. 5). Furthermore, full-length moesin and the N-terminal domainco-localize and co-immunoprecipitate with Ano1 (Fig. 6). Inaddition, co-expression of Ano1 with Nter-moesin increases Ano1lateral mobility in the plane of the membrane, supporting theconclusion that moesin links Ano1 to the cytoskeleton, and as suchanchors it and limits its lateral diffusion (Fig. 7). Collectively, ourresults support a model in which Ano1 traffics to the microvillus,where it binds ERMproteins and cross-links to the cytoskeleton. Thisstabilizes the microvillus membrane, revealing an important role ofAno1 in microvilli biogenesis in the oocyte.

Not surprisingly, expression of Nter-moesin alone decreasesmembrane surface area (Fig. 4). This likely partly results frominhibition of Ano1–ERM interaction, but also from the inhibition ofother interactions between ERM and plasmamembrane proteins andlipids. This is highlighted by the discoloration observed in theoocyte at high levels or after prolonged expression of Nter-moesin(Fig. 4). Interestingly, however, co-expression of Ano1 with full-length moesin does not show a synergistic effect over Ano1expression alone. This could be caused by a physical limit on thelength of the microvilli, independent of the amount of availablescaffolding proteins. The effect of Ano1 on microvilli can be readilydissociated from its channel function. Blocking the activation ofAno1 using MONNA does not affect the ability of overexpressedAno1 to increase microvilli length and membrane capacitance.Conversely, when the Ano1–ERM link is perturbed, either byexpressing the dominant-negative Nter-moesin mutant (Fig. 5) or theAno1 N-terminal cytosolic fragment (Fig. S4), the global chloridecurrent is not affected, yet the increased capacitance resulting fromAno1 expression is reversed. This shows that Ano1 regulatesmicrovilli length through a physical/scaffolding interaction and notthrough its Cl− property of conducting channel ions.

The role of plasma membrane channels in shaping themorphology of microvilli is largely unknown, with the exceptionof channelrhodopsin-2, which has been shown to enlarge microvilli(Zimmermann et al., 2008). Expression of other plasma membranechannels, such as CFTR or the sodium channel ENaC, increases thecell capacitance, but this process is attributed to the increased fusionof transport vesicles at the plasma membrane (Awayda, 2000;Weber et al., 1999). Several integral membrane proteins areinvolved in shaping the microvillus structure through interactionswith ERM proteins such as CD43, CD44, ICAM-2 (Bretscher et al.,2002; Yamane et al., 2011; Yonemura et al., 1999) or CD9 in theoocyte (Runge et al., 2007). The microvillus is a spatially definedcompartment with some proteins, including ions channels,localizing to microvilli (or even in microvillus sub-regions),through binding to intermediate scaffolding proteins such asERMs (Lange, 2011). The binding of membrane proteins to thecytoskeleton would therefore have the dual role of stabilizing themicrovilli by reinforcing the scaffolding link between the plasmamembrane and the actin backbone, and of precisely distributingchannels to specific functional locations.

What is the function of the anchoring of Ano1 to thecytoskeleton? The distribution of Ano1 is not homogenous inthe oocyte and it is known that CaCCs localize preferentially to theanimal pole of the oocyte at the expense of the vegetal pole(Machaca and Hartzell, 1998). This might in fact explain theincreased microvilli density at the animal compared with the vegetal

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pole of the oocyte (Bluemink et al., 1983). We recently showed thatCa2+ influx through store-operated Ca2+ entry (SOCE) specificallyactivates distally located Ano1 through Ca2+ transfer within theER lumen and release through IP3 receptors (Courjaret andMachaca, 2014). We refer to this as ‘mid-range Ca2+ signaling’ asit spatially bridges elementary and global Ca2+ signaling. SOCE‘clusters’ do not co-localize with Ano1. Consequently, the clustersare unlikely to make it to the microvilli and constitute a specificcompartment in terms of Cl− and Ca2+ signaling. The anchoring ofAno1 to the cytoskeleton would therefore allow a very preciselocalization of the channel in the vicinity or away from specific Ca2+

release sites.Another important parameter is the function of the microvilli. In

the oocyte the microvilli contact the macrovilli arising from thefollicle cells and protruding through the vitteline envelope (Browneand Werner, 1984; Dumont and Brummett, 1978). At the contactpoint, gap junctions allow the passage of numerous molecules,particularly IP3 and Ca2+ produced by the stimulation of PLC infollicular cells, a complex coupling that varies depending on thesubtypes of follicular cells involved and/or gap junctions (Arellanoet al., 2012; Sandberg et al., 1992; Supplisson et al., 1991). Precisepositioning of Ano1 within the microvilli by anchoring it to thecytoskeleton is likely to enhance the efficiency of a signaltransferred from the follicular cells to the oocyte through gapjunctions, avoiding cytoplasmic buffers.In summary, we have newly identified a function of Ano1 that

regulates microvilli length, diameter and density within the oocytethrough linkage to ERM proteins and the actin cytoskeleton. This inturn modulates the surface area of the cell. Given the widedistribution of Ano1 in different epithelia, which are oftenenriched in microvilli, and the conservation of the role of ERMsand the actin cytoskeleton, these findings are likely to translate tomost epithelia.

MATERIALS AND METHODSExpression in Xenopus oocytesStage VI oocytes were harvested from wild-type and lab-bred Xenopuslaevis (Xenopus Express, France and Nasco, U.S.A) using previouslydescribed procedures (Machaca and Haun, 2000). Oocytes were kept in 0.5×L15 (Sigma) and used no earlier than 24 h after surgery. Animals werehandled according to Weill Cornell Medical College IACUC approvedprocedures.

Molecular biologyHuman moesin (Accession number: NP_002435, a gift from Criss Hartzell,Emory University, Atlanta, GA) was sub-cloned from pcDNA3.1 throughXba1 and HindIII restriction sites into the pSGEM vector using thefollowing set of primers: 5′-ACTGCTCTAGAATGCCCAAAACGATC-3′;5′-TCGATAAGCTTTTACATAGACTCAAATTCG-3′. Human moesinN-terminus (1–381) was sub-cloned from pcDNA3.1 through Xba1 andHindIII restriction sites into the pSGEM vector using the followingset of primers: 5′-ACTGCTCTAGAATGCCCAAAACGATC-3′; 5′-TCG-ATAAGCTTTTAACGCTTCCGTTCCTGC-3′. Human moesin C-terminus(382–577) was sub-cloned from pcDNA3.1 through Xba1 and HindIIIrestriction sites into a psGEM vector using the following primers:5′-ACTGCTCTAGAATGGCCCAGAGCGAGG-3′; 5′-TCGATAAGCT-TTTACATAGACTCAAATTCG-3′. To create moesin–GFP fusionconstructs, human moesin, moesin (1–381) and moesin (382–577) weresub-cloned from pcDNA3.1 using SacII and XbaI in-frame into pSGEM-GFP. The primers used were: for pSGEM-Moesin-GFP, 5′-ACTGCCCG-CGGATGCCCAAAACGATC-3′, 5′-TCGATTCTAGACATAGACTCA-AATTCGTCAATGCGCTGC-3′; for pSGEM-Moesin (1-381)-GFP,5′-ACTGCCCGCGGATGCCCAAAACGATC-3′, 5′-TCGATTCTAGAA-CGCTTCCGTTCCTGCTCAAGTTC-3′; for pSGEM-Moesin (382-577)-

GFP, 5′-ACTGCCCGCGGATGGCCCAGAGCG-3′, 5′-TCGATTCTAG-ACATAGACTCAAATTCGTCAATGCGCTGC-3′

ElectrophysiologyThe ionic currents were recorded using standard two-electrode voltage-clamp recording technique. Recording electrodes were filled with 3 M KCland coupled to a Geneclamp 500B controlled with pClamp 10.5 (AxonInstruments). CaCC currents were recorded using depolarizing pulses to+40 mV and a previously described ‘triple jump’ protocol (Courjaret andMachaca, 2014; Machaca and Hartzell, 1999). Membrane capacitance wasmonitored using the built-in routine from pClamp to measure membraneparameters. Cells were continuously superfused with saline during voltage-clamp and imaging experiments using a peristaltic pump. The standardextracellular saline contained (in mM) 96 NaCl, 2.5 KCl, 1.8 CaCl2,2 MgCl2, 10 HEPES, pH 7.4.

ImagingConfocal imaging of live oocytes was performed using a LSM710 (Zeiss,Germany) fitted with a Plan Apo 63×/1.4 oil immersion objective or a TCSSP5 (Leica, Germany) fitted with a 63×/1.4-0.6 oil immersion objective.Confocal z-stacks were taken in 0.5 µm sections using a 1 Airy unit pinholeaperture. The images were recorded using Zen black 2012 (Zeiss) or LASAF 2.4.1 (Leica) software. Micrographs of oocytes were taken using a ZeissLumar V12 Stereomicroscope controlled using Zen blue 2012 (Zeiss).Images were analyzed with ImageJ 1.46q (Schneider et al., 2012). Cellsurface calculation was performed using the ‘iso-surface’ function of theBone J plugin (Doube et al., 2010). A 400 µm2 image planewas selected anda mesh created to fit the 3D reconstruction of the surface. A resamplingfactor of 2 was used and a detection threshold for surface change of 20pixels. The results are expressed as fold increase of the theoretical surfacearea (measured surface/400).

Wheat germ agglutinin stainingFor WGA staining, the cells were cooled down to 4°C for 15 min thenstained for 30 min at 4°C in Ringer solution containing 5 µg ml−1 of Alexa-Fluor-488−WGA conjugate (Cat no. W11261, Thermo Fischer Scientific).The cells were then washed in Ringer solution and kept on ice prior toimaging to minimize internalization of the membrane marker.

Fluorescence recovery after photobleaching (FRAP)For FRAP experiments, orthogonal images across the membrane wereobtained directly using x-z scans: a line scan is performed at different zpositions and the z -positioning is achieved using a ‘Super-Z’ galvanometricstage (Leica) that allows for fast monitoring of the fluorescence across theplasma membrane. To bleach a defined region of the membrane, the zoomfactor of the scanner was increased two times and the laser power (561 nm)set at 100% for three scans. Scanning speed was set a 400 Hz and imagefrequency to 0.1 Hz for both bleaching and recording.

Immunoprecipitation and western blotOocytes were lysed in lysis buffer containing Triton X-100 (10 mM Hepes,150 mMNaCl, 1 mM EGTA, 0.1 mMMgCl2, 0.5% Triton X-100, pH 7.4).The immunoprecipitation was performed by magnetic associated cellsorting (MACS) using microbeads according to the manufacturer’sinstructions (Miltenyi Biotec, Germany). The lysates were incubated withmicro beads conjugated to an anti-GFP monoclonal antibody (µMACS GFPIsolation Kit 130-091-125; Miltenyi Biotec) for 30 min on ice. The lysate–beads complexes were then sorted in the magnetic field of a µMACS(Miltenyi Biotec). After the washing steps, the proteins were eluted with thedenaturing elution buffer provided in the kit. Western blotting wasperformed as previously described (Courjaret et al., 2013) and repeatedthree times on different samples. The anti-Ano1 antibody was used at adilution of 1:1000 (SAB2102460, QC13356 Sigma).

StatisticsValues are given as means±s.e.m. Statistical analysis was performed whenrequired using either Student’s paired and unpaired two-tailed t-test or

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ANOVA and corrected for multiple comparison. P-values are indicated asfollows: *P<0.05; **P<0.01; ***P<0.001 and ns, not significant. Statisticswere obtained using Prism 6.05 (GraphPad Software, La Jolla, USA).

AcknowledgementsWe are grateful to Criss Hartzell for sharing the human moesin clones.

Competing interestsThe authors declare no competing or financial interests.

Author contributionsR.C. designed and performed experiments, analyzed data andwrote the paper. R.H.performed and analyzed experiments. S.H., A.I., M.D. and S.D. performedexperiments. K.M. designed experiments, analyzed data and wrote the paper.

FundingThis work was funded by the Qatar national research fund (QNRF) [grant number:NPRP 5-475-3-127]. The statements made herein are solely the responsibility of theauthors. Additional support for the authors comes from the Biomedical ResearchProgram (BMRP) at Weill Cornell Medical College in Qatar, a program funded byQatar Foundation.

Supplementary informationSupplementary information available online athttp://jcs.biologists.org/lookup/doi/10.1242/jcs.188367.supplemental

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RESEARCH ARTICLE Journal of Cell Science (2016) 129, 2548-2558 doi:10.1242/jcs.188367

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