Correction

BIOPHYSICS AND COMPUTATIONAL BIOLOGYCorrection for “Voltage-dependent motion of the catalytic regionof voltage-sensing phosphatase monitored by a fluorescent aminoacid,” by Souhei Sakata, Yuka Jinno, Akira Kawanabe, andYasushi Okamura, which appeared in issue 27, July 5, 2016, ofProc Natl Acad Sci USA (113:7521–7526; first published June 21,2016; 10.1073/pnas.1604218113).The authors note that on page 7522, left column, first paragraph,

line 9, “faster” should instead appear as “slower.”

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Voltage-dependent motion of the catalytic region ofvoltage-sensing phosphatase monitored by afluorescent amino acidSouhei Sakataa,b,1,2, Yuka Jinnoa, Akira Kawanabea, and Yasushi Okamuraa,2

aLaboratory of Integrative Physiology, Graduate School of Medicine, Osaka University, Suita, Osaka 565-0871, Japan; and bInstitute for Academic Initiatives,Osaka University, Suita, Osaka 565-0871, Japan

Edited by Gail Mandel, Howard Hughes Medical Institute: Oregon Health & Science University, Portland, OR, and approved May 12, 2016 (received for reviewMarch 16, 2016)

The cytoplasmic region of voltage-sensing phosphatase (VSP) derivesthe voltage dependence of its catalytic activity from coupling to avoltage sensor homologous to that of voltage-gated ion channels. Toassess the conformational changes in the cytoplasmic region uponactivation of the voltage sensor, we genetically incorporated afluorescent unnatural amino acid, 3-(6-acetylnaphthalen-2-ylamino)-2-aminopropanoic acid (Anap), into the catalytic region of Ciona intesti-nalis VSP (Ci-VSP). Measurements of Anap fluorescence under voltageclamp in Xenopus oocytes revealed that the catalytic region assumesdistinct conformations dependent on the degree of voltage-sensoractivation. FRET analysis showed that the catalytic region remainssituated beneath the plasma membrane, irrespective of the voltagelevel. Moreover, Anap fluorescence from a membrane-facing loop inthe C2 domain showed a pattern reflecting substrate turnover. Theseresults indicate that the voltage sensor regulates Ci-VSP catalytic activ-ity by causing conformational changes in the entire catalytic region,without changing their distance from the plasma membrane.

VSP | unnatural amino acid | phosphatase

Voltage-sensor domains confer the voltage dependency topore domains of voltage-gated ion channels and voltage-

sensing phosphatase (VSP). In some proteins, the voltage-sensordomain also provides an ion permeation pathway (1–3). TheVSP is a voltage-dependent phosphoinositide phosphatase (4–8),which is composed of two regions: a voltage-sensor domain ho-mologous to that of voltage-gated ion channels and a cytoplasmiccatalytic region (4). The catalytic region shows a high degree ofsimilarity to the phosphatase tensin homolog deleted on chromo-some 10 (PTEN), which is known as a tumor suppressor. PTENand the catalytic region of VSP are both composed of a phos-phatase domain, which contains the phosphatase active center, anda C2 domain, which is reportedly responsible for membrane bind-ing (9–12). In addition, a linker region called the phosphoinosi-tide binding motif (PBM) is situated between the voltage sensorand the catalytic region of VSP (13–16).Molecular mechanisms of coupling between the voltage-sensor

domain and the catalytic region in VSP still remain unclear. Inthe recently reported crystal structures of the catalytic region ofVSP, the PBM interacts with a loop structure in the phosphatasedomain (17, 18), named the “gating loop.” This structure sug-gests two distinct conformations (18) and raises the possibilitythat movement of the voltage sensor induces conformationalchanges in the gating loop by altering the conformation of thePBM. Our previous study using a mutant with a voltage sensortrapped into an intermediate state has suggested that enzymaticactivity is graded, dependent on the state of the voltage-sensordomain (19). A recent study with rapid detection of change ofcatalytic products (PI(3,4)P2, PI(4,5)P2, and PI4P) as readout ofphosphatase activity provided evidence that distinct states of thevoltage sensor are coupled to multiple states of the catalyticregion probably with distinct preference for phosphoinositidespecies (PIP3 versus PIP2s) (20). However, little information is

available for the motion of the catalytic region of VSP driven bythe operation of the voltage-sensor domain. To address theseissues, we used a method of genetic incorporation of an environ-ment-sensitive fluorescent amino acid analog, 3-(6-acetylnaphthalen-2-ylamino)-2-aminopropanoic acid (Anap), for site-specific labelingof the catalytic region of Ciona intestinalis VSP (Ci-VSP) (21–24). Because Anap is comparable in size to other amino acids, itsincorporation causes a minimal perturbation of the protein’sstructure. Results show that the voltage sensor regulates Ci-VSPcatalytic activity by inducing conformational changes in both thephosphatase and C2 domains, not accompanied by a significantchange of a distance from the plasma membrane. We also reportthat fluorescence change of Anap is consistent with the presenceof multiple voltage-dependent states and with protein conforma-tion, which is sensitive to substrate availability in the active center.

ResultsVoltage-Dependent Fluorescence Change of Anap Introduced into theCytoplasmic Region of Ci-VSP. Earlier crystallographic studies ofthe catalytic region of Ci-VSP suggest operation of the gatingloop in the phosphatase domain regulates the opening andclosing of the active center, and this movement is controlled bythe voltage sensor (18). To determine whether voltage-sensor

Significance

Voltage-sensing phosphatase (VSP) dephosphorylates phosphoi-nositides in a voltage-dependent manner. The molecular mech-anisms by which the voltage-sensor domain of VSP activates thecatalytic activity of the cytoplasmic region still remain unknown.Using a method of incorporation of a fluorescent unnaturalamino acid, 3-(6-acetylnaphthalen-2-ylamino)-2-aminopropanoicacid (Anap), in the catalytic region, we revealed that some loopsin the catalytic region move on membrane depolarization andthat the catalytic region is located beneath the plasmamembraneirrespective of the membrane potential. Furthermore, fluores-cence change of Anap in the C2 domain showedmultiple voltage-dependent activated states and protein conformation, which issensitive to substrate availability in the active center. Thesefindings provide novel insights into the mechanisms of voltage-dependent catalytic activity of VSP.

Author contributions: S.S. and Y.O. designed research; S.S., Y.J., and A.K. performed re-search; S.S. analyzed data; and S.S. and Y.O. wrote the paper.

The authors declare no conflict of interest.

This article is a PNAS Direct Submission.

Freely available online through the PNAS open access option.1Present address: Department of Physiology, Division of Life Sciences, Faculty of Medicine,Osaka Medical College, Takatsuki, Osaka 569-8686, Japan.

2To whom correspondence may be addressed: Email: sakatas@osaka-med.ac.jp oryokamura@phys2.med.osaka-u.ac.jp.

This article contains supporting information online at www.pnas.org/lookup/suppl/doi:10.1073/pnas.1604218113/-/DCSupplemental.

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movement induces gating loop motion, we incorporated Anap atfour sites within the gating loop (D400, F401, F407, and Q408)of Ci-VSP containing an enzyme-inactive mutation (C363S) andexpressed the protein in Xenopus oocytes. We used two band-pass emission filters, 420–460 nm and 460–510 nm, to detectAnap fluorescence (Fig. S1). We found that the fluorescenceintensity changed in a voltage-dependent manner in all four sites(Fig. 1 and Fig. S2). Kinetics of the fluorescence change ofAnap-incorporated F401 were faster than that of the chargemovement of the voltage sensor of WT Ci-VSP in lower voltagesthan 50 mV but exhibit similar kinetics to those of the chargemovement at higher voltages (Fig. 1B). On the other hand, nofluorescence changes were detected when another mutation(D129R) was introduced that immobilized the voltage sensor(25) (Fig. S3). Furthermore, we introduced a mutation into thePBM motif (R253A/R254A) that reportedly causes an uncou-pling between the voltage-sensor movement and the catalyticactivity (13, 14). No voltage-dependent fluorescence changeswere detected when Anap was incorporated at F401 with theR253A/R254A mutations of the PBM motif (Fig. S3).We also examined whether the C2 domain, which is not directly

connected to the voltage-sensor domain, moves upon voltage-sensor activation. Anap was incorporated into the 515 loop (fromS513 to R520) and Cα2 loop in the C2 domain (Fig. 2 and Fig. S4).Voltage-dependent fluorescence changes were observed at six ofthe eight labeled sites within the 515 loop, but not at S513 or L518(Fig. 2A), whereas fluorescence changes were observed at all of thelabeled sites in the Cα2 loop (Fig. 2B). When Anap was incorpo-rated in the background of a voltage-sensor immobile mutation(D129R) or PBM mutation, the voltage-dependent changes influorescent signal from both S515Anap and K555Anap, whichotherwise showed the largest fluorescence change in the 515 andCα2 loops, respectively, were lost (Fig. S3). We found the fluo-rescence changed in both same and opposite directions betweenthe 420–460 nm and the 460–510 nm filters. The direction of fluo-rescence change may include the information of how the confor-mation changed. However, we analyzed the only magnitude of thefluorescence without paying much attention to the directionof change in this study because detailed mechanisms of changeof Anap fluorescence are not known so far.We next determined how fast the phosphatase and C2 domains

move upon membrane depolarization (Fig. 3 and Fig. S5). Therising phases of the normalized fluorescence changes at F401Anap,S515Anap, and K555Anap, which showed the largest fluorescence

changes in each loop, were fitted by single exponential equation.Estimated delays in the fluorescences through both the 420–460 nmand 460–510 nm emission filters upon the depolarizing step to160 mV were from 2 ms to 4 ms, on average, in all three constructs,and did not differ significantly (Fig. 3B).

FRET Between Anap and Dipicrylamine Embedded in the PlasmaMembrane Does Not Detect a Large Change in Distance Betweenthe Catalytic Region of Ci-VSP and the Membrane. The results so farshow that voltage-sensor movement regulates the conformationof the catalytic region. However, because Ci-VSP substrates aremembrane phosphoinositides, it is also possible that the voltagesensor regulates phosphatase activity, changing the distance be-tween the catalytic region and the plasma membrane, therebymodulating the availability of substrate close to the active site ofthe enzyme. To test this possibility, we measured FRET betweenthe Ci-VSP catalytic region and the plasma membrane. The cata-lytic region was labeled with Anap at S513, as the site where Anapfluorescence was unchanged by membrane depolarization (Fig.2A), and the plasma membrane was stained with dipicrylamine(DPA) as FRET acceptor (Figs. S1C and S6A) (26–28). Wemeasured Anap fluorescence to evaluate the FRET efficiency be-cause DPA is not a fluorescent molecule. It is known that DPAtranslocates between the outer and inner leaflets of the plasmamembrane in a voltage-dependent manner (26, 27) (Fig. S6 B andC). To detect voltage-dependent movement of the catalytic regionusing FRET, membrane potential was held at 0 mV, a level atwhich the voltage sensor of Ci-VSP is in a nearly resting state andthe majority of DPA is on the inner leaflet (Fig. 4A). When themembrane potential is hyperpolarized from 0 mV to −120 mV, theintensity of the Anap fluorescence should increase because ofthe decrease in FRET efficiency due to the transition of DPAtoward the outer leaflet. When the membrane is depolarized from0 mV to 160 mV, the Anap fluorescence should decrease due tothe increase in FRET efficiency as the catalytic region is pulledtoward the plasma membrane (Fig. 4A, Left). On the other hand,Anap fluorescence should not be changed if the catalytic regiondoes not move toward the membrane, even when the membrane isdepolarized to 160 mV (Fig. 4A, Right). We measured Anap fluo-rescence in a wider bandwidth for emission (the 420–520 nmemission filter) to increase the fluorescence intensity detectedby the photomultiplier tube (PMT) (Fig. S1A). An increase inAnap fluorescence was observed in the presence of DPA whenthe membrane was hyperpolarized to −120 mV in oocytesexpressing Ci-VSP (S513Anap/C363S) (Fig. 4B, Upper). Next,we changed the membrane potential from 0 mV to 160 mV andfound that the fluorescence slightly decreased (Fig. 4B, Lower).To examine whether this decrease resulted from movement of

the catalytic region, we also tested a Ci-VSP mutant (D129R/S513Anap/C363S) in which the voltage sensor was immobilized

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Fig. 1. Voltage-dependent changes of Anap fluorescence. (A) Representa-tive changes in fluorescence from Anap incorporated at F401. Red and blackare the fluorescences detected using 420–460 nm and 460–510 nm band-passemission filters, respectively. Voltage steps were applied from −60 mV to160 mV in 20-mV increments. (Inset) Structure of Anap. (B) Time constant ofthe fluorescence change of Anap incorporated into F401 and the chargemovement of WT Ci-VSP. The moved charges were estimated by the in-tegration of on-sensing currents. The fluorescence elicited by the step pulsesand the integrated currents were fit by the single exponential equation. Thedata of sensing charge measured from the cut-open oocyte was derivedfrom a previous paper (the same dataset used for figure 1 C and D of ref. 4)(gray circle). Data are shown as the mean ± SD [n = 5, 5, and 5 for the 420–460 nm, the 460–510 nm filter, and on-sensing charge with TEVC (triangle),respectively].

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Fig. 2. Voltage-dependent changes in fluorescence from Anap incorporatedinto the 515 and Cα2 loops in the C2 domain.Changes in the fluorescence ofAnap in the 515 loop (A) and the Cα2 loop (B) evoked by 160-mV test pulses,respectively.

7522 | www.pnas.org/cgi/doi/10.1073/pnas.1604218113 Sakata et al.

(29). We found that the decrease in fluorescence persisted in thevoltage-sensor immobile form (Fig. 4C). Plots of the normalizedfluorescence changes in S513Anap/C363S and D129R/S513Anap/C363S against the membrane voltage showed no significant dif-ference (Fig. 4D). The voltage dependencies of the FRET effi-ciency (Fig. 4D) were more like that of DPA movement than thecharge–voltage (Q–V) relationship of Ci-VSP (Fig. S6C), sug-gesting no voltage-dependent change in the distance between thecatalytic region and the plasma membrane. The Q–V relationshipfor DPA was not completely saturated around 0 mV (Fig. S6C),consistent with an earlier study (27). This finding indicates thatDPA is not situated entirely within the inner leaflet at 0 mV, buttranslocates to the inner leaflet at 160 mV, which would explainwhy we observed a decrease of the fluorescence at 160 mV in thevoltage-sensor immobile mutant. Thus, FRET experiments revealedno voltage-dependent change in the distance between the cata-lytic region and the plasma membrane.

Distinct Fluorescence–Voltage Relationships Between the Enzyme-Active and -Inactive Forms of Ci-VSP. We next compared changesin Anap fluorescence in enzymatically active and inactive forms(C363S) of Ci-VSP and systematically analyzed their voltage de-pendency. One site in the gating loop of the phosphatase domain(F401) and three sites in the C2 domain (S515, R520, and K555)were analyzed. The kinetics of the fluorescence changes atF401Anap did not differ significantly between the active and in-active forms of the enzyme (Fig. 5A and Fig. S7). The fluores-cence–voltage (F–V) relationship for fluorescence measuredusing the 420–460 nm filter was identical to that measured usingthe 460–510 nm filter both in the active and inactive forms (Fig.5B). With the active form, the F–V relationships for the 420–460 nmand 460–510 nm signals were shifted toward higher voltages com-pared with the inactive form (Fig. 5B).The F–V relationship at S515Anap (in the 515 loop of the C2

domain) showed a slight rightward shift in the 460–510 nmfluorescence from the active enzyme compared with the inactiveform (Fig. 5 C and D). For the active enzyme, the 420–460 nmsignal was too small to analyze its voltage dependence (Fig. S7).The F–V relationship for R520Anap, which is also located in the515 loop, had two components (Fig. S7I). The fluorescence inthe bandwidth of 460–510 nm from both the inactive and active

forms of the enzyme decreased at around 50 mV and then in-creased at higher membrane voltages.The F–V relationship for the 460–510 nm signal fromK555Anap

(Cα2 loop of the C2 domain) had one component in the inactiveenzyme, but two components in the active form (Fig. 5 E and F).The F–V relationship for the active enzyme was shifted slightlytoward higher voltages compared with the inactive form (Fig. 5F),as was the case with the 420–460 nm signals from F401Anap andS515Anap. The F–V relationships of K553Anap and K558Anap,other sites in Cα2 loop, were also examined (Fig. S8). BothK553Anap and K558Anap exhibited two components of fluores-cence change in the active enzyme, but only one in the inactivemutant as in the case of K555Anap (Fig. S8). We verified that Ci-VSP mutants into which Anap was incorporated in the above sitesall retained the voltage-dependent catalytic activity using theKir3.2 channel (Fig. S9).

The Voltage-Dependent Movement of the Cα2 Loop Is Sensitive toSubstrate Availability. Our analyses revealed that the F–V rela-tionships were shifted between the enzyme-active and -inactiveforms in all sites we examined, including F401, S515, and K555. TheF–V relationship for F401Anap, S515Anap showed only one com-ponent in both the enzyme-inactive and -active forms, whereas theF–V relationship for the 460–510 nm fluorescence from K555Anapin the Cα2 loop showed one and two components in the enzyme-inactive and -active forms of Ci-VSP, respectively (Fig. 5F). In thepulse protocol with a 1.2-s pulse interval, phosphoinositide sub-strates were depleted after activation of phosphatase of the Anap-incorporated enzyme-active form of Ci-VSP and there was littleavailable substrate in the plasma membrane due to the incompleterecovery of phosphoinositides (Fig. S9 D and E) mediated by en-dogenous kinases (30, 31). Therefore, one possible explanation forthe difference in the F–V relationships between the active and in-active forms is that the observed fluorescence was affected by thelevel of substrates in the plasma membrane.To determine how the level of available substrate affects Anap

fluorescence, we measured the signal using longer pulse intervals

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Fig. 4. FRET between Anap and DPA embedded in the plasma membrane.(A) Strategy for examining the voltage-dependent movement of the cata-lytic region toward the plasma membrane. Text provides details. (B and C)Changes in the fluorescence of Anap incorporated at S513 within active (B)and immobilized (C) forms of the voltage sensor in the presence of DPA.(Upper and Lower) Anap fluorescence evoked by the test pulses to −120 mVand +160 mV, respectively. (Insets) Anap fluorescence in the absence ofDPA at −120 mV. Vertical and horizontal bars in the Insets indicate 0.25%and 0.2 s, respectively. (D) Voltage dependency of Anap fluorescence inthe presence of DPA. Data are shown as the mean ± SD (n = 7 for bothS513Anap/C363S and D129R/S513Anap/C363S).

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Fig. 3. Estimation of the delay to onset of the fluorescence changes elicitedby depolarization. (A) Magnified views of the changes in fluorescence fromF401Anap/C363S. (Left and Right) Fluorescence changes detected using 420–460 nm and 460–510 nm emission filters, respectively. (Insets) Overviews of thefluorescence data. Black lines above the fluorescence data are actual mem-brane potentials, and the median time of the voltage change from −60 mV to160 mV was defined as the time of depolarization, which is indicated by anarrow. The fluorescence changes were fitted by the single exponential equa-tion (the blue curve). The dotted line indicates the initial level, which wasdefined as the average of the fluorescence intensity from 0 ms to 30 ms afterthe beginning of the protocol, during which the voltage was clamped at−60 mV. The intersection of the exponential curve (blue) and the dotted linewas defined as the onset of the fluorescence change. Trace was shown fromthe middle of the first 30 ms. (B) Quantitative comparison of the delay to onsetof the fluorescence change after depolarization. Red and black bars indicatethe delay estimated from the data obtained using 420–460 nm and 460–510 nmfilters, respectively. Data are shown as the mean ± SD (n = 4, 4, and 6 forF401Anap/C363S, S515Anap/C363S, and K555Anap/C363S, respectively).

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(60 s). We reasoned that with this pulse protocol with a longinterval the level of phosphoinositides recovers until the nextepisode of the test pulse, because the Kir current could be re-stored within around 50 s to its original amplitude by clampingthe membrane voltage at −60 mV, a voltage at which Ci-VSPis silent (Fig. S9 D and E). The F–V relationships for bothF401Anap and S515Anap appeared to be unaffected by thedifferent pulse intervals (Fig. S10 A and B). By contrast, the F–Vrelationship for K555Anap showed one component when fluo-rescence was measured with a 60-s pulse interval, but two com-ponents when fluorescence was measured with a 1.2-s interval(Fig. 6A). In addition to the change in the F–V relationship, wefound that the kinetics of the fluorescence change also differedbetween the 1.2-s and 60-s intervals in active Ci-VSP. Thefluorescence from K555Anap measured at 60-s intervals showeda transient decrease just after the large increase evoked by thedepolarizing test pulse to 160 mV (Fig. S10C, Bottom). Thistransient decrease was not observed when the fluorescence wasmeasured at 1.2-s intervals (Fig. 5E).To confirm whether the transient decrease in fluorescence was

dependent on the level of substrate in the plasma membrane, wemeasured the fluorescence change in the absence and presenceof a depolarizing prepulse in the same oocytes (Fig. 6B and Fig.S10D). We found the fluorescence decrease was not seen in theprotocol with a prepulse (Fig. 6B) but was seen in the absence ofa prepulse. Moreover, the hump could be restored by clampingthe membrane voltage for 1 min at −60 mV, a voltage at whichCi-VSP is in a resting state (Fig. 6B). The prepulse-dependentchange in fluorescence kinetics was not observed at F401Anap orS515Anap (Fig. S10E). These results show that substrate avail-ability did not affect the F–V relationship or the kinetics of thefluorescence change at F401Anap or S515Anap, but did affect

fluorescence from K555Anap, which suggests that movement ofonly the Cα2 loop is affected by substrate availability.Crystal structures showed that the Cα2 loop is indirectly bound

to the substrate by several hydrogen bonds mediated by Tyr522(18) (Fig. 7A). To confirm that these hydrogen bonds are re-sponsible for the Cα2 loop movement associated with the substratemetabolism, Tyr522 was mutated to alanine and the fluorescencechange of Y522A/K555Anap was measured. The hump was notfound by the depolarization test pulse to 160 mV in the absence ofprepulse with a 60-s interval in the Y522A mutant (Fig. 6B), andthe F–V relationship of Y522A/K555Anap measured with a 60-sinterval did not have two components and was similar to thatmeasured with a 1.2-s interval (Fig. 7B). Because the Y522A mu-tant was reported to retain the phosphatase activity to a levelsimilar to that of WT protein but with the positive shift of voltagedependence (18), substrates should be depleted from the bindingpocket on the 60-s membrane depolarization. These findings sug-gest that interactions between the Cα2 loop and the substrate viaTyr522 are involved in the substrate-dependent change of thefluorescence of K555Anap. The F–V relationships of three siteslabeled in the Cα2 loop, K553, K555, and K558, were composedof one or two components dependent on the catalytic activity,whereas those in F401 (gating loop) and S515 (515 loop) were onecomponent irrespective of the catalytic activity. These findingssuggest that the catalytic activity of Ci-VSP is accompanied bymovement of the Cα2 loop within the C2 domain.

DiscussionTo detect the voltage-dependent motion of the cytoplasmic cata-lytic region of Ci-VSP, we incorporated a fluorescent unnaturalamino acid, Anap, into the protein using a genetic method with aXenopus oocyte expression system. Our approach brought thefollowing findings. First, three loops in the catalytic region move ina voltage-dependent manner (Figs. 1 and 2) with similar timing(Fig. 3). Second, changes in the voltage dependency of the fluo-rescence introduced into some sites revealed two components(Fig. 5F and Figs. S7I and S8 D and G), suggesting distinct con-formational changes in the catalytic region depending on the de-gree of voltage-sensor activation. Third, the FRET experimentprovided no evidence of a change in the distance between thecatalytic region and plasma membrane upon voltage-sensor acti-vation (Fig. 4). Fourth, Anap on a single site, K555, of the Cα2loop showed fluorescence change sensitive to the availability ofsubstrates in the enzyme active center (Fig. 6), reflecting confor-mational changes associated with catalysis and/or substrate turn-over (Fig. 7C).

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Fig. 6. Effect of phosphoinositide availability on themotion of the cytoplasmicregion of Ci-VSP probed by Anap at K555. (A) F–V relationships measured using1.2-s test pulse intervals (circles) or 60-s test pulse intervals (triangles). Data areshown as the mean ± SD (n = 4 and 9 for the fluorescence measured using 1.2-sand 60-s test pulse intervals, respectively). (B) Changes in the kinetics with al-tered phosphoinositides level induced by Ci-VSP activities in the same oocyte(Left, K555Anap; Right, Y522A/K555Anap). (Top) Timing of the depolarization.Arrows indicate decay in fluorescence that depends on a depolarizing prepulse.(Left Bottom) Fluorescence change in the absence of the prepulse, recorded1 min after the measurement in the presence of the prepulse, which is shown inthe second panel from the Bottom. Data are taken from the same oocytes.

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Fig. 5. F–V relationship of the enzyme-active and -inactive forms of Ci-VSP.(A, C, and E) Representative fluorescence changes of Anap incorporated intoF401 (A), S515 (C), and K555 (E) in the enzyme-active or -inactive forms. (B, D,and F) F–V relationship for F401Anap (B), S515Anap (D), and K555Anap (F). Redand black are the fluorescences detected using 420–460 nm and 460–510 nmband-pass emission filters, respectively. Open and filled circles indicate datafrom the inactive and active enzymes, respectively.

7524 | www.pnas.org/cgi/doi/10.1073/pnas.1604218113 Sakata et al.

A Fluorescent Unnatural Amino Acid Suggests Distinct Voltage-DependentStates of the Catalytic Region of Ci-VSP. An earlier X-ray crystallo-graphic study suggested a model in which the operation of thegating loop is associated with voltage-sensor movement (18). Wenow provide direct evidence that upon the voltage-sensor activation,the gating loop moves (Fig. 1 and Fig. S2) along with the 515 andCα2 loops in the C2 domain (Fig. 2). Interestingly, the fluorescenceof K553Anap and K555Anap decreased when the membrane wasdepolarized to around 50 mV, whereas those increased at highervoltages, and the fluorescence of K558Anap increased at around50 mV and decreased at higher voltages (Fig. 5F and Fig. S8 D andG). In addition, these two components of the F–V relationships werealso observed in both the active and inactive forms of the enzymewith incorporated R520Anap (Fig. S7I). These observations indicatethat the catalytic region does not have only two states correspondingto the resting and activated state of the voltage sensor but couldhave an intermediate state, which depends on the degree of acti-vation of the voltage sensor. We have previously conducted an ex-periment of trapping the intermediate state of the voltage sensor(19) to show that the catalytic activity is graded. Experimentsof monitoring the levels of phosphoinositides using GFP-fusedpleckstrin homology domains suggests that the substrate specificitycould be altered in a voltage-dependent manner (18, 32, 33). Arecent study of several voltage-sensor mutants showing altered statetransitions among three distinct states of voltage sensor com-bined with rapid detection of phosphoinositide change, as thereadout of enzyme activity suggested that a distinct stateof voltage sensor induces a distinct enzyme state with biasedpreference toward phosphoinositide species (PI(3,4,5)P3 versusPI(4,5)P2 and PI(3,4)P2) (20). It will be interesting in the future toaddress whether bidirectional changes of Anap fluorescencedependent on membrane voltage in our study reflect multiplestates of the catalytic region with distinct enzyme properties.

Little or No Change in Distance Between the Ci-VSP Catalytic Regionand the Plasma Membrane upon the Voltage-Sensor Activation. Weanalyzed FRET between the Ci-VSP catalytic region and the plasma

membrane to determine whether the distance between themchanges during enzyme activation (Fig. 4). Before testing for thiseffect, the membrane was hyperpolarized to −120 mV from aholding potential of 0 mV. We noted an increase in Anap fluores-cence at −120 mV, which was caused by the displacement of DPAfrom the inner to the outer leaflet of the membrane (Fig. 4 and Fig.S6). This finding means that when the voltage sensor is in a restingstate, the catalytic region remains close enough to the membrane forFRET to occur, and that Anap and DPA can serve as a FRET pair.In addition, there was no significant difference in the voltage-dependent change in the fluorescence between the voltage-sensor active and immobile mutants (Fig. 4D), which impliesthat activation of the voltage sensor does not lead to a change inthe distance between the catalytic region and the membrane.However, we do not completely exclude the possibility that thecatalytic region moves toward the membrane upon voltage-sensoractivation within the detection limit of the FRET system. We thusconclude that our FRET experiment showed that the catalytic regionremains beneath the plasma membrane while the voltage sensor is inthe resting state and that any possible change in the distance be-tween the catalytic region and the membrane upon voltage-sensoractivation is smaller than the detection limit of our system.

The Substrate Availability Is Associated with the Movement of theCα2 Loop in the C2 Domain.We found that, in three sites in the Cα2loop, the F–V relationships of the active enzyme contained twocomponents, whereas those of the inactive enzyme had only onecomponent (Fig. 5F and Fig. S8 D and G). The influence of thesubstrate availability on K555Anap fluorescence was lost by themutation of Y522, which mediates an indirect binding betweenthe Cα2 loop and the substrate (18) (Figs. 6B and 7B). Further-more, it has been reported that the neutralization of positive charges,K553, K554, K555, and K558 affected the substrate selectivity ofthe enzyme (32). These findings suggest that the concerted move-ment of the Cα2 loop is accompanied by the conformational changeof the phosphatase domain for the catalytic activity of Ci-VSP. Wealso examined the effect of substrate availability on the fluorescencechange of other sites of the Cα2 loop, K553Anap and K558Anap.The F–V relationships were distinct between the enzyme-active and-inactive forms, like K555Anap. However, the F–V relationships ofboth the enzyme-active and -inactive K553Anap had two compo-nents irrespective of whether the interval duration was 1.2 s or 60 s(Fig. S8D), and the depolarizing prepulse did not affect the kineticsof the fluorescence (Fig. S10E). The incorporation of Anap intoK553 or K558 does not impair the catalytic activity as shown bymeasurements of the catalytic activities (Fig. S9C). One possiblereason that only K555Anap among several sites in the Cα2 loopshowed the substrate availability is that the side chain of Anap in-corporated at K555 is located at a position exposed to an environ-ment that is altered by the motion of the Cα2 loop, whereas theK553 and K558 side chains are not.Based on a crystallographic study of the catalytic region of Ci-

VSP, it was proposed that movement of the voltage sensor changesthe gating loop conformation, which in turn opens a substrate-binding pocket near the active center, enabling substrate phos-phoinositides to bind (18). We found that the fluorescent signalfrom Anap introduced into the gating loop changed in a voltage-dependent manner (Fig. 1 and Fig. S2), which supports the ideathat voltage-sensor activation leads to movement of the gatingloop. However, voltage-dependent movement was also seen in the515 and Cα2 loops. The similar onsets of fluorescence changes inthe three is consistent with the idea that the phosphatase and C2domains move as a single unit (Fig. 3). We also found thatmovement of the Cα2 loop differed between the active and inactiveenzymes (Fig. 5F and Fig. S8 D and G), and that a hump in thefluorescence change at K555Anap in the Cα2 loop was seen whenphosphoinositides are not depleted (Fig. 6B). The kinetics of thechange in K555Anap fluorescence when phosphoinositides are not

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Fig. 7. Structure of the Cα2 loop and a model of the conformational changein the catalytic region of Ci-VSP. (A) Structure around the active center andCα2 loop. The Cα2 loop is shown in cyan. IP3 is located at the active center(PDB ID code 3V0H) (18). (B) Voltage dependence of the changes in fluo-rescence from Y522A/K555Anap. Circles and triangles denote the normal-ized fluorescence measured using 1.2-s and 60-s test pulse intervals,respectively. Data are shown as the mean ± SD (n = 4 for both fluorescencedetected using 1.2-s and 60-s test pulse intervals). (C) Proposed model of theconformation change in the catalytic region. Voltage-sensor activation in-duces conformational changes in the catalytic region such that the 515 andCα2 loops as well as the gating loop all move, and the substrate is able tobind to the active center (“substrate binding form”). Once the substrate isbound, it is dephosphorylated (“dephosphorylation form”), which may beaccompanied by a conformational change in the Cα2 loop (shown in red).

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depleted exhibited two phases: In the earlier phase, the fluores-cence increased just after the onset of depolarization, whereas inthe later phase, the fluorescence declined (Fig. 6B, arrow). It maybe that the increase in fluorescence was associated with the bindingof substrate to the active center, which involves voltage-sensoractivation and movement of the gating, 515 and Cα2 loops,whereas the decrease in fluorescence may be associated with aconformation change to catalyze dephosphorylation of phosphoi-nositides, which is a process occurring just after substrate binding(Fig. 7C). An operation of the Cα2 loop may correspond to eachstep of a catalytic cycle of the phosphatase, such as an opening ofthe active center and a removal of the phosphate from the sub-strate. Future studies of analyzing correlation between detectionof local structural change and substrate turnover will be necessaryto test this idea. Detailed analyses using the fluorescent unnaturalamino acid in Ci-VSP will also lead to understanding conservedmolecular mechanisms shared by other phosphatases, such asPTEN, which has a high similarity to the catalytic region of Ci-VSP.

MethodscDNAs. The cDNA Ci-VSP was identical to that used in our earlier work (4–6). AKir3.2d cDNA was a gift from Yoshihisa Kurachi (Osaka University, Osaka,Japan). G protein β- and γ-plasmids were provided by Toshihide Nukada(Tokyo Metropolitan Institute of Medical Science, Tokyo, Japan). pAnap, aplasmid that encodes tRNA and aminoacyl-tRNA synthetase, was kindlyprovided by Peter G. Schultz (The Scripps Research Institute, La Jolla, CA).

Electrophysiology. Electrophysiology and fluorescence measurements wereperformed with a two-electrode voltage clamp (TEVC) as we did previously (5, 6).ND 96 solution (5 mM Hepes, 96 mM NaCl, 2 mM KCl, 1.8 mM CaCl2, 1 mMMgCl2, pH 7.5) was used as the bath solution for measurements of fluorescenceand sensing currents. For measurement of the sensing current, leak currents andthe current for charging the cell capacitance were cancelled using the P/−10 or P/−5 protocol, in which a step pulse for subtraction was applied from the holdingpotential. The sensing charge was calculated by the integration of on- or off-sensing current. The holding potential was −60 mV unless otherwise noted.

Incorporation and Measurement of Anap Fluorescence. All experiments wereperformed in compliance with the Animal Research Committees of theGraduate School of Medicine, Osaka University. Xenopus oocytes were pre-pared as described previously (4–6). For Anap (FutureChem) incorporation intoCi-VSP, 20 nL of pAnap solution (10 ng/μL) was injected into the nucleus ofthe oocytes. One day later, 1 mM Anap and cRNA encoding Ci-VSP in whichthe target site was mutated to a TAG codon were mixed 1:1, and the 20 nLof the mixture was injected into the oocytes. The oocytes were then in-cubated for 2 d in the dark. Oocytes were imaged using an IX71 invertedmicroscope (Olympus) equipped with a 10× 0.3 N.A. objective lens and amercury lamp under TEVC. Fluorescence was detected using one or twoPMTs (H10722-20; Hamamatsu Photonics). The output of the PMTs wasdigitized using the AD/DA converter (Digidata1440A) shared with the TEVCsetup. Fluorescence changes were elicited by a set of test pulses of 20-mVincrements except for the prepulse experiment. The set of fluorescence datawas averaged from 4 to 16 times. There was no time interval between theset of test pulses. The fluorescence data of the experiment with pre-conditioning pulse (the protocol shown in Fig. S10D) were also averaged byrepeating from 4 to 16 times. All fluorescence data were digitally filtered at300 Hz. DPA was purchased from Tokyo Chemical Industry for use in theFRET experiments.

Note Added in Proof. After the paper was accepted, a paper characterizingvoltage dependence of phosphatase activity of VSP shared among four sub-reactions toward distinct phosphoinositides was published (34).

ACKNOWLEDGMENTS. We thank Dr. Peter G. Schultz (The Scripps ResearchInstitute) for giving us the pAnap plasmid; Dr. Yoshihisa Kurachi (OsakaUniversity) for providing the Kir3.2d plasmid; Dr. Mari Sasaki for providingoriginal data of the sensing current of wild-type Ci-VSP measured by the cut-open oocyte method; Dr. Masafumi Minoshima (Osaka University) for givingus advice for the Anap experiment; Dr. Fumihito Ono (Osaka MedicalCollege) for encouragement and critical reading of the manuscript; andDr. Yasushi Sako (RIKEN) for critical reading of the manuscript. This work wassupported by Grants-in-Aid for Scientific Research (to S.S. and Y.O.) and CoreResearch for Evolutional Science and Technology from the Japan ScienceTechnology Agency (Y.O).

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