Analysis of Ni2+-induced arrest of Paramecium axonemes
J0RGEN LARSEN1'2 and PETER SATIR1
'Department of Anatomy and Structural Biology, Albert Einstein College of Medicine, Bronx, New York 10461, USA^Institute of Cell Biology and Anatomy, University of Copenhagen, Copenhagen 0, Denmark
Summary
Thi9 study examines the molecular basis for paral-ysis of ciliary motility by Ni2+. At concentrationsabove 0.1 mM, Ni2+ slowed and subsequently stoppedswimming of living, axenically grown Parameciumtetraurelia. However, some cilia still beat in thepresence of 0.1 mM Ni2+. When permeabilized andreactivated with 4mM ATP at pCa>7, cells resumedciliary beat and swam forward at approximately170±28/ims~1; swimming speed increased in thepresence of 10 ftM cyclic AMP. Addition of Ni2+
(pNi<5) caused rapid arrest of all ciliary beat in asingle position. This was fully reversible when EGTAwas added to raise the pNi. Axonemes were thenisolated and sliding was observed in the presence oftrypsin and ATP. When pNi was lowered to about 5,sliding was reduced dramatically. This too was
reversible with EGTA. Dynein was then extractedfrom the axonemes and used for in vitro translo-cation assays. At concentrations of Ni2+ wheremicrotubule-sliding and axonemal beat were greatlyinhibited or absent, microtubule translocation invitro by 22 S dynein was only slightly affected.However, translocation by 14 S dynein was stoppedcompletely. When pNi was raised by repeated wash-ing with solutions containing EGTA, microtubuletranslocation by 14 S dynein resumed. We concludethat Ni2+ induces a reversible paralysis by a directeffect on 14 S dynein while 22 S dynein is not aprimary target.
Paramecium is an excellent system for studying the effectsof various agents on the control of ciliary activity, sincebiochemical and mutational analysis can be combined todissect the regulatory pathways (Hinrichsen et al. 1986;Saimi and Kung, 1987; Bonini and Nelson, 1988; Satir etal. 1988). Since the original work of Gelei (1935) andTartar (1950), it has been well established that Ni2+
gradually inhibits ciliary activity in different ciliatedcells, including Paramecium, but the molecular basis ofNi2+ action has not been understood. This may proveimportant, since Ni2+ is effective in reversing the highCa block in certain axonemes to a fixed beat position(Tiands down'), in which one axonemal switch thatcontrols arm activity during beat is thought to be blocked(Satir et al. 1990). The present study has investigatedwhether Ni2+ acts directly on the Paramecium ciliaryaxoneme and, if so, whether the site of inhibition is dyneinitself. Detergent-permeabilized cells have proven valuablein addressing the first question (Naitoh and Kaneko, 1973;Bonini and Nelson, 1988; Lieberman et al. 1988). Thistreatment compromises the cell membrane so that, as wewill demonstrate, low concentrations of Ni2+ can directlyand reversibly block axonemal motility.
digestion and the complexity of structural interaction inthe axonemes limit the information obtainable about thedirect effect of Ni2+ on dynein itself in such studies. An invitro motility assay has recently been introduced by whichthe properties of isolated dynein in translocating micro-tubules can be studied directly (Paschal et al. 1987; Valeand Toyoshima, 1988; Sale and Fox, 1988). This assay is animprovement over existing motility assays, because onlypurified dynein and microtubules are involved. We haveused this assay to investigate the effect of Ni2+ onmicrotubule translocation induced by the two dyneins(22 S and 14 S) that we have isolated from Parameciumaxonemes (Larsen et al. 1991). The effect of Ni2+ on dyneinATPase activity has also been determined. We willdemonstrate that Ni2+ inhibits beat and sliding at thesame concentration, that it inhibits microtubule translo-cation by one of the dyneins (14 S), while it has a muchmore limited effect on translocation by isolated 22 Sdynein, suggesting that Ni2+ inhibits axonemal switchingby an effect on 14 S dynein.
Materials and methods
Paramecium tetraurelia were grown axenically at 27 °C in amedium adapted from Soldo et al. (1966). Cells were harvested atlate log or early stationary phase by gentle centrifugation.Paramecia were then washed three times in different buffers andallowed to equilibrate for at least 30min prior to experimentaluse.
Ni2+ was added as nickel acetate from 10 mM or 100 mM stocksolutions. When addition of Ni2+ or other components to the
33
media caused a change in pH this was readjusted by addition ofNaOH or HC1. To calculate pNi for different solutions, we haveused the Calcon computer program developed by Goldstein (1979),and modified by J. S. Tash. Stability constants are from Martelland Smith (1974).
In preliminary experiments we investigated the effect of Ni2+
on swimming behavior of living paramecia in different buffers: (1)in 2 mM KC1, 1 mM CaCl2, 5 ra Tris-HCl, pH 7.2; (2) in a TECKbuffer (4mM KC1, lmM CaCl2, 0.1 mM EDTA, 10 mM Tris-HCl,pH 7.2); or (3) in a buffer without calcium (5 mM Pipes, 1 mM KC1,20 mM MgCl2, pH7.2). For determination of the rate of motility,control and cation-exposed cells were placed in a small chamberand observed by dark-field or phase-contrast microscopy. Visualobservations or motion analysis were used to estimate changes inswimming behavior caused by Ni2+
Preparation and reactivation of permeabilized cellsParamecia were permeabilized with 0.01% Triton X-100 on iceaccording to the method of Naitoh and Kaneko (1972). Whenswimming and ciliary beating had ceased (25 min), the cells werewashed free of detergent. Thin sections of such permeabilized cellsindicate that the ciliary, cell and outer alveolar membranes aregreatly disrupted or entirely missing (Lieberman et al. 1988).Permeabilized cells were reactivated at room temperature in aMgATP solution (4mM MgSO4, 4mM ATP, 1.5mM EGTA, 10 mMTris-maleate, 50 mM KC1, pH 7.0). In some cases, cyclic AMP wasadded to the reactivation medium to increase the swimming speed(Bonini and Nelson, 1988). Samples of 0.4ml were withdrawn andexposed to Ni2+ and other divalent cations (Ca2+, Mg2"1", Co2+) todetermine their effect on swimming behavior and ciliary activityin reactivated cell models. After 2 mm exposure the effect wasobserved by phase-contrast and dark-field microscopy. In someexperiments EGTA was added following cation exposure, in whichcase the effect on swimming behavior and ciliary activity wasdetermined again 2 min after EGTA addition.
Observation and recording of swimming behaviorSwimming behavior was examined by dark-field microscopy andrecorded on videotape for quantitative motion analysis. Thevideotapes were analyzed using a system introduced by MotionAnalysis Systems (MAS), Inc. (Santa Rosa, CA), following theprocedure of Lieberman et al. (1988), modified after Sundberget al. (1986).
Scanning electron microscopyPreparation for SEM was performed as described by Lieberman etal. (1988) with minor modifications. Living and permeabilizedcells were quick-fixed for 30 s in 1 % OsO< in 10 mM cacodylatebuffer (pH 7.0), followed by rapid addition of 2 % glutaraldehydein 10 mM cacodylate buffer. After 10 min the OsO4-glutaralde-hyde mixture was replaced by fresh glutaraldehyde (2 %) for 1 h,and washed twice with 10 mM cacodylate buffer. Fixed cells werethen transferred to polylysine-coated coverslips and dehydratedthrough a graded series of ethanol. After dehydration the sampleswere critical-point dried and gold coated before examination in aJEOL S35 scanning electron microscope (SEM). Micrographs arerepresentative of the sample of undistorted, well-ciliated cells.
Isolation of axonemesTo eliminate the possible effect of high Ca2+ in the calcium shocktreatment, in some cases axonemes were prepared by mechanicaldeciliation of permeabilized cells as described by Hamasaki et al.(1989). Results using this preparation are essentially identical tothose obtained by a calcium shock deciliation (Adoutte et al.1980), which was used routinely. After deciliation, cilia wereisolated from cell bodies by centrifugation and purified accordingto methods of Hamasaki et al. (1989). Cilia derived from Ca2+
shock treatment were then treated with 0.8% Triton X-100 for20 min on ice to remove membranes.
Axonemal sliding experimentsThe axonemes were washed twice with 'activation buffer' (4mM
MgS04, 1.5 mM EGTA, 50 mM KCl, 10 mM Tris-maleate, pH7.0or 7.5) to remove Triton, resedimented, and resuspended inactivation buffer. Following measurement of absorbance (O.D.units) of the suspension at 350 nm, a sample corresponding to0.4 unit A350 was obtained and placed on ice for immediate use. Insome experiments, this axonemal suspension was exposed toCa2+, Mg2+, Co2+ or Ni2+ for 2 min and introduced into a 15/Jperfusion chamber (Larsen et al. 1991). The axonemes attached tothe glass were then activated by perfusion with activation buffercontaining trypsin (l/;gml-1), ATP (O.lmM—4mM) and appropri-ate cations. In other sliding experiments, the axonemal suspen-sion was first digested with trypsin. The digestion process wasmonitored turbidimetrically and halted with excess soybeantrypsin inhibitor, when the A350 decreased to 80 % of the initialvalue. The axonemes were then applied to the perfusion chamberand unadsorbed axonemes were removed by perfusing activationbuffer through the chamber. The adsorbed axonemes wereexposed to the appropriate cations for 2 min and then perfusedwith activation buffer containing ATP. In some cases, exper-iments were performed in which EGTA was added followingcation exposure. The proportion of disintegrated axonemes wasestimated by direct observation of sliding in dark-field andrecorded on videotape.
Electron microscopyFor determination of the effect of Ni2+ on negatively stainedpreparations of digested axonemes two samples were withdrawnfrom the above mentioned cuvette. Ni2+ was added to one sample,whereas the other served as control. Digested axonemes wereapplied to a sheet of Formvar-coated carbon-stabilized, coppergrids. Sliding was initiated by diluting the sample with an equalvolume of activation buffer containing ATP. The axonemes wereincubated for 5 min in these solutions at room temperature, afterwhich excess fluid was removed and 1 % aqueous uranyl acetatepipetted onto the grids for negative staining. The grids weredrained, allowed to dry in air under cover, and observed in a JEOL100CX electron microscope. Sliding images were identified bycriteria discussed by Sale and Satir (1977) and Larsen et al.(1991).
Preparation of dynein from parameciumThis procedure essentially follows Larsen et al. (1991). Axonemesfrom 4-liter stationary-phase cultures were isolated as above, andresuspended in axoneme buffer (30 mM Hepes, 5mM MgS04>0.5 mM EDTA, 20 mM KCl, lmM dithiothreitol (DTT),SOKTUmT1 aprotinin, lO/zgrnP1 leupeptin, pH7.6) with 0.6 MKCl to extract the dyneins. Samples of 0.2 ml containing high-saltextracted protein were layered on top of linear 12 ml 5 % to 30 %sucrose gradients prepared in axoneme buffer without leupeptinand aprotinin, and with DTT at O.lmM. The gradient wascentrifuged for 15 h at 35 000 revs min"1 (4°C) in a SW41 Ti rotor.Fractions were collected from top to bottom of the tube. Eachfraction was assayed for protein and ATPase activity, and thosecontaining the 14 S and 22 S dynein were pooled separately. Thepurified dynein solutions were frozen for later use in in vitromotility assays or for measurement of ATPase activity.
Dynein in vitro motility assayMotility assays were carried out essentially as described by Valeand Toyoshima (1988) and Larsen et al. (1991). 14 S or 22 S dyneinsamples were adjusted to approximately O.lmgml"1 protein inaxoneme buffer and a 20 ;d sample was applied to the assaychamber in two successive portions for 2-min incubation periodseach. The unadsorbed dynein was removed by perfusing 20/dtranslocation buffer (50 mM K+-acetate, 10 mM Tris-acetate,pH7.5, lmM EGTA, 3rnM MgS04) through the chamber.Subsequently, translocation buffer containing taxol-stabilizedbovine brain microtubules and 1 mM ATP was perfused throughthe chamber. This assay system has the advantage that thechamber that has dynein bound to the glass can be perfusedseveral times to examine movement under different conditions.To test the effects of Ni2+ on microtubule motility, Ni2+ wasadded to the translocation medium in which the motility assay
34 J. Larsen and P. Satir
was performed. Translocation of microtubules was examined atroom temperature using video-enhanced dark-field microscopy.Reversibility of inhibition was tested by perfusing additionaltranslocation buffer without nickel through the chamber.
ATPase and protein assaysATPase activities of the 22 S and 14 S dyneins were tested in thepresence of Ca2+, Mg2+, Co2+, Ni2+ and the dynein inhibitorvanadate. In these experiments the dyneins were always washedfree from sucrose using a 30000Mr cutoff centricon andresuspended in activation or translocation buffer before additionof the metal ions.
Dynein ATPase activity (in the absence of microtubules) wasanalyzed by the orthophosphate determination method followingHayashi and Takahashi (1979), modified after Murphy and Riley(1962). Protein concentration was determined using the BioRadBradford reagent (BioRad, Rockville Centre, NY) using bovineserum albumin as a standard.
Results
The effect ofNi2+ on ciliary activity in living parameciaIn TECK buffer Paramecium tetraurelia swam forward ina gently curving spiral path with a swimming velocity ofabout 360 ± 26/an s"1 (mean of 3 experiments; swimmingvelocity determined by motion analysis of at least 30randomly chosen cells in each experiment). These forwardswimming cells showed characteristic metachronal wavepatterns with a wavelength of approximately 11 fan andan effective stroke toward the posterior end of the cell(Fig. 1). Ni2+ immobilized cell motility in a dose-depen-dent manner. In TECK buffer, concentrations above0.1 mM Ni2+ slowed and finally stopped swimming within5 min as a result of uncoordinated ciliary activity. Suchimmobilized cells showed some sporadic ciliary motion,but had lost the coordinated beating characteristic ofmetachronal waves. After prolonged exposure to Ni2+
concentrations above 0.1 mM an increasing deciliation wasobserved, but some cilia still beat, and the cells exhibitedrocking movements. This residual ciliary activity was,however, limited and the cilia appeared stiff, in that theirbeat was restricted to a limited arc relative to the bodysurface.
Ca2+ has been reported to abolish the immobilizing
effect of Ni2+ in Paramecium (Kuznicki, 1963; Andrivon,1972) and it could be that the residual beating we observedwas due to the presence of 1 mM CaCl2 in TECK buffer.Therefore the effect of Ni2+ on cell motility was tested in abuffer without added Ca2+. This buffer did not affect theresults.
Effect of Ni2+ and other divalent cations onpermeabilized cells
In an appropriate reactivation medium, permeabilizedparamecia were immobile in the absence of externallysupplied ATP. After reactivation with Mg2+ and ATP,ciliary activity resumed and most cell models swam. Thefew models that did not swim were reactivatable, sincetheir cilia were beating rapidly, but they appeared to beattached firmly to the glass of the chamber. The averageswimming velocity of the free-swimming cell models wasapproximately 170±28Jums~1 (mean of 3 experiments)and the models swam continuously forward with themetachronal wave pattern characteristic of living, for-ward-swimming" cells (Fig. 2A). Addition of cyclic AMP tothe reactivation medium caused an increase in theswimming speed of the free-moving permeabilized cells,and an increase in the number of free-moving cells in thepopulation. When the cells were reactivated in thepresence of 10 ,UM cyclic AMP the swimming speedincreased to about 200 jim s"1 and virtually all of the cellsin the population swam in straight or curved paths(Fig. 3). Ni2+ at low free-ion concentration (30,UM) causedrapid arrest of all ciliary activity, resulting in a completeimmobilization of the permeabilized cells (Fig. 3). Otherdivalent cations such as Co2+ did not cause a comparableinhibition. When permeabilized cells were reactivated andexposed to approximately 30 /JM free Ni2+ (pNi 4.6), therewas no deciliation. The metachronal wave was no longerobserved in SEM and cilia were captured primarily in asingle-stroke position (Fig. 2B). For most of their length,the cilia curved uniformly to point toward the posteriorend of the cell in a position corresponding to the end of theeffective stroke ('hands down', in the terminology of Wais-Steider and Satir, 1979).
The permeabilized cells resumed ciliary activity whenEGTA was added, raising the pNi to 6.4 or higher (Fig. 3;Table 1). Although beat was reactivated in the whole
Fig. 1. SEM of quick-fixed living Paramecium. Metachronal waves are seen (A=ll/an). Anterior end of the cell to the left; effectivestroke direction to the right. Bar, 10;<m.
Effect of Ni2+ on axonemes 35
Fig. 2. SEM of quick-fixed permeabilized Paramecium in reactivation medium. (A) Triton-permeabilized reactivated control cellwith normal metachronal waves. (B) After 26 /.IM free Ni2+ is added, virtually all cilia point toward the posterior of the cell.Metachronal waves are absent. Bar, 10 fun.
population, both the number of swimming cells and theiraverage swimming velocity fell substantially (Fig. 3).Even after all treatment, however, some cells resumedswimming at rates greater than 200(ms"1, suggestingthat for these cells, Ni2+ inhibition was completelyreversible (Fig. 4).
The effect of Ni2+ on axonemal slidingThe axonemes obtained from isolated cilia of Para-
mecium were not uniform in length and short fragmentswere commonly found. Without trypsin treatment, in ATPParamecium axonemes did not normally resume beatingand only slid to a limited extent. Trypsin treatment causedan increase in the proportion of axonemes that disinte-grated by sliding upon ATP addition. Ni2+ had a markedeffect on the ATP-induced disintegration of trypsin-treated axonemes. When pNi was lowered to about 5, thenumber of axonemes sliding was greatly reduced or sliding
36 J. Larsen and P. Satir
B
pNi% Swimming cells
Swimming speed (/ans~')
00
<500
>729
150
Fig. 3. Swimming tracks of permeabilized Paramecium generated by motion analysis. Swimming paths represent distancestravelled in 1 s (11 frames). For clarity, only frames 1, 4, 7 and 10 are displayed. (A) Cells in reactivation medium containing 10 (JMcyclic AMP; (B) after addition of 2mM Ni 2 + to A; (C) After addition of 2mM EGTA to B.
Table 1. Reversal ofNi2+ inhibition of beat ofpermeabilized cells by EGTA
Conditions beforeEGTA addition
pNi Beat
4.63.73.7
Final
pNi
7.7*6.4*9.6t
conditions
Beat}
*+2mM EGTA.t+4raM EGTA.
} + + + , Comparable to untreated permeabilized cells; + + , slightinhibition; - , no beat.
Table 2. Effects of different cations on ATP-inducedsliding of axonemes
Cation pCa' Shdingt
ControlNiCoMgCa
>74.74.33.23 7
Axonemes in buffer (pH7 or 7.6) treated with trypsin before orsimultaneously with addition of ATP.
* Free ion concentration with 2 mM of cation added.t Direct observation of sliding in darkfleld on videotape: + + + ,
marked increase in number sliding versus control; + + , comparable tocontrol; +, decrease versus control; - , little or no sliding.
Fig. 4. Distribution of swimming speeds for free-movingpermeabilized paramecia reactivated in medium containing10 tat cyclic AMP. (A) Control. (B) Cells stopped by addition of2 mM Ni2+ and restarted by addition of 2 mM EGTA (protocolas in Fig. 3).
was abolished. In contrast, marked sliding occurred at pCa4. Sliding also occurred when Co2+ was substituted forNi2+ (Table 2). The Ni2+ inhibition of sliding could bereversed by subsequent addition of EGTA.
Electron microscopy of negatively stained preparationsof disintegrated axonemes showed that both the dyneinarms and the spoke groups were easily identified aftertrypsin treatment. Sliding configurations of ciliary axon-
emes were readily identified (Larsen et al. 1991) (Fig. 5A).When pNi was lowered to about 5, sliding configurationscould no longer be found, but axonemes sometimes openedto show periodic interdoublet linkages (Fig. 5B).
The effect oftranslocation
on dynein-mediated microtubule
When dyneins were extracted from Paramecium axonemeawith 0.6 M KC1 and further purified by sucrose densitygradient centrifugation, two peaks of ATPase activity,corresponding to 22 S dynein and 14 S dynein, wereobtained. In in vitro motility assays, the translocationvelocity depends on the buffer used. Under standardconditions with the translocation buffer (see Materials andmethods) the isolated 22 S dynein induced attachment andtranslocated MAP-free, taxol-stabilized, calf brain micro-tubules with an average velocity of 2.71(±0.92)/zms~1 at22 °C in the presence of 1 mM ATP (Table 3) (Larsen et al.1991). Addition of Ni2+, to pNi 4.6, sufficient to inhibitbeat and sliding, affected translocation velocity signifi-cantly, but did not stop translocation (Table 3). In somecases, translocation by 22 S dynein could still be observedat free Ni2+ concentrations at least 10 times higher thanthose necessary to inhibit beat completely in permeabil-ized cells. In contrast, under the same standard conditionsat pNi>7, 14 S dynein translocated microtubules atvelocities of 0.79(±0.33)/ans"1 (Table 3) (Larsen et al.1991). However, translocation was inhibited at pNi 5.6 and
Effect ofNi2+ on axonemes 37
Table 4. Effects of different divalent cations on 14 S and22 S dynein Mg2*-ATPase Activity
B
Fig. 5. Negative-stain images of axonemes treated withtrypsin and ATP. (A) Control. Sliding configurations arereadily found. Note normal (+) direction displacement ofdoublet N+l. Dynein arms are indicated by arrow. Bar, 0.1/on.(B) With Ni2+ at pNi< 5, sliding configurations are absent.Axonemes are largely intact. Inset shows that axonemessometimes unroll. Spoke groups repeat is indicated. Doubletdisplacement is not seen. Bar, 0.5 fira.
Table 3. Measurements of dynein-mediated microtubuletranslocation
pNi
>7 (control)6.85,64 *
Velocity
22 S
2.71±0.92
1.47+0.21
of translocation
14 S
0.79±0.330.62±0.160.51±0.09
Addition
Control10/(M vanadate2mM Ca2+
3 HIM Ca2+
2mM Mg24-3mM Nig2*2mM Ni2+
3m M Ni 2 +
Mean of 3 experiments; s.D. less
Relative(.9,
22 S
1009
96$4
m1019382
than 10 %.
activity>)
14 S
1008
__
102S38878
The effect on Ni2+ on 22 S and 14 S dynein-induced microtubuletranslocation. Translocation of microtubules was examined at roomtemperature using video-enhanced dark-field microscopy. Measurementswere made on at least 30 microtubules. Values are given in (ons"1±s.D.
* Translocation almost completely abolished.
was almost completely abolished at pNi 4.6, i.e. the sameconcentration of Ni2+ that blocked ciliary beat in MgATP-reactivated permeabilized cells and sliding of isolatedtrypsin-treated axonemes. When pNi was raised byrepeated perfusion with translocation buffer (containingEGTA), translocation by 14 S dynein resumed at controlvelocities.
Effect of divalent cations on dynein ATPase activityThe Mg2+-ATPase activity of purified 22 S or 14 S dyneinwas measured before and after addition of Ni2+. Intranslocation buffer, at pNi values below 5, dynein activitydecreased to about 80 % of the control value for both 22 Sand 14 S dynein (Table 4). Comparable addition of Ca2+
had no effect on ATPase activity. Addition of 10,UMvanadate nearly abolished activity, as anticipated. Thusat pNi values where motility, sliding and microtubuletranslocation by 14 S dynein were greatly inhibited,ATPase activity of dynein was much less affected.
Discussion
Ni2+ has long been known to inhibit ciliary activity ofciliates (Gelei, 1935; Kuznicki, 1963) and other cells,although the precise mechanism involved was unknown.In living paramecia, Ni2+ inhibits ciliary activity in agradual and diffuse way, probably acting first at the cellmembrane, by competing with other divalent cations atmembrane channel sites. The use of detergent-permeabil-ized, MgATP-reactivated cells has permitted experimentalaccess to the axonemal effects of Ni2+. In contrast to itseffects on living cells, Ni2+ rapidly arrests ciliary activityin reactivated Triton-extracted cells (Naitoh and Kaneko,1973; Andrivon, 1974). Hence Ni2+ appears to act directlyon the axoneme. In SEM, almost all nonbeating cilia ofNi2+-immobilized permeabilized paramecia point pos-teriorly ('hands down'). Naitoh and Kaneko (1973) showedthat Ni2+-immobilized cilia could be moved into a 'handsup' position by addition of Ca2+ (above 10~ 4 M). Thisbehavior would be consistent with the switching ofmammalian sperm cells by Ca2+ and Ni2+ (Lindemannand Goltz, 1988) and of 'hands down' mussel gill cilia(Wais-Steider and Satir, 1979; Reed and Satir, 1986). Incilia, Ni2+ affects a switch point of axonemal activity sothat cilia are blocked in a position corresponding to thefinal stage of an effective stroke, while Ca acts to blockmotility at the end of a recovery stroke (Satir et al. 1990).
ATP-induced sliding of trypsin-treated axonemes isinhibited by the same concentration of Ni2+ that inhibitsthe motility of demembranated cells under the same
38 J. Larsen and P. Satir
conditions. When pNi is lowered to about 5,* sliding isreduced dramatically or abolished. In contrast, slidingoccurs at pCa 4 or in comparable Co2+ concentrations.Therefore, Ni2+ specifically affects either dynein mecha-nochemistry and/or some trypsin-insensitive structuralcomponents of the axoneme. Lindemann et al. (1980)showed that low concentrations of Ni2+ did not completelyinhibit microtubule sliding in reactivated bull spermflagella, but did inhibit beat. This suggests that theconcentration thresholds for Ni2+ inhibition of sliding andbeat may not coincide exactly, although in our exper-iments they are in the same range.
To investigate whether Ni2+ affects dynein directly, weisolated 22 S and 14 S dynein from Paramecium axonemes.22 S dynein is a three-headed bouquet with three heavychains that are u.v. photocleavable in the presence ofvanadate, while 14 S dynein is a single-headed species thatmay be heterogeneous (Larsen et al. 1991). Two u.v.photocleavable heavy chains are found in the 14 S regionin our preparations. Travis and Nelson (1988) reportedthat a shoulder of the 22 S peak could be identified as 19 Sdynein, but this is not apparent in our preparations wherethe fraction size is larger. Their 12 S peak probablycorresponds to our 14 S dynein (see Larsen et al. 1991, forfurther details).
Both 22 S dynein and 14 S dynein support translocationof purified bovine brain microtubules in an in vitromotility assay using a standard translocation buffer, but22 S dynein translocates microtubules at a rate aboutthree times as fast as 14 S dynein. Addition of Ni2+ atconcentrations sufficient to inhibit beat and slidingabolishes translocation by 14 S dynein in in vitro assaysbut only partially affects translocation by 22 S dynein.Translocation by 22 S dynein is observed at Ni2+ concen-trations at least ten times higher than those required forcomplete beat inhibition. In contrast, Vale and Toyoshima(1988) demonstrated that addition of vanadate completelyinhibited 22 S dynein-induced microtubule translocationin Tetrahymena, indicating that Ni2+ does not inhibitciliary motility in the same way as vanadate, byabolishing general dynein ATPase activity. In fact ATPaseactivities of purified 22 S and 14 S dyneins are only weaklyinhibited by Ni2+, although the extent of inhibition seemsdependent on the exact assay conditions. Using somewhatdifferent conditions, Travis and Nelson (1988) showed that14 S dynein ATPase activity was inhibited by Ni2+ to amuch greater extent than 22 S dynein. Therefore, inhi-bition of beat, sliding and probably ATPase activity byNi2+ correspond to the specific inhibition of 14 S dynein, asdemonstrated by inhibition of microtubule translocationin in vitro assays. This is supported by the observation thatthe effect of Ni2+ on translocation, as well as on beat andsliding, is reversible when EGTA is added to raise the pNi.Unfortunately, the axonemal localization of 14 S dynein,which now seems critical to an understanding of switchingactivity in the axoneme, and of the Ni effects, isuncertain. Many investigators believe that, on the basis ofrebinding studies, 14 S dynein is a component of the innerdynein arm (Warner et al. 1985). However, dyneins oftenrebind promiscuously to several structures in the axoneme(Satir et al. 1981). Another possibility is that 14 S dynein isa component of the spoke head. There is some suggestionthat antibodies to 14 S Tetrahymena dynein are localizedat either the spoke head or the outer midwall of thedoublets. It is unlikely that dynein at the outer midwallcould be responsible for switching arm activity, but theinner arm-spoke relationships are probably fundamental
to this process. It would be useful to have the appropriateaxonemal mutants in Paramecium to be able to pursuethis question further. It is also interesting that 22 Sdynein, which is found in the outer arm, is evidently notcritically inhibited by Ni2+ at low concentrations.
This work was supported in part by grants from the USPHS toP.S. and from the Danish Natural Science Research Council toJ.L. We are grateful to J. Avolio, K. Barkalow and T. Hamasakifor their help with various aspects of the study and to M. AnnHolland for secretarial assistance. Assistance and support for themotion analysis work reported was kindly provided by Dr J. L.Spudich and his laboratory.
References
ADOUTTB, A., RAMANATHAN, R, LEWIS, R. M., DUTE, R. R., LING, K Y ,KUNG, C. AND NELBON, D. L. (1980). Biochemical studies of theexcitable membrane of Paramecium tetraurelia, HI. Proteins of ciliaand ciliary membranes. J. Cell Bwl. 84, 717-738.
ANDRIVON, C. (1972). The stopping of ciliary movements by nickel saltsin Paramecium caudatum: the antagonism of K+ and Ca++ ions. Actaprotozool 11, 373-386.
ANDETVON, C. (1974). Inhibition of ciliary movements by Ni2+ ions intnton-extracted models of Paramecium caudatum. Archs Int Physwl.Biochem. 82, 843-852.
BONINI, N. M. AND NELSON, D L. (1988). Differential regulation ofParamecium ciliary motility by cAMP and cGMP. J. Cell Bwl. 106,1615-1623.
GELEI, J. (1935). Ni-Infusion in Dienste der Forschung und desUntemchtes. Biol. Zbl. 66, 57-74.
GOLDSTEIN, D. A. (1979) Calculation of the concentrations of the cationsand cation-ligand complexes in solutions containing multiple divalentcations and ligands Biophys. J. 28, 235-242
HAMASAKI, T , MUBTAUCH, T. J., SATIR, B. H. AND SATDI, P. (1989). Invitro phosphorylation of Paramecium axonemes and penneabilizedcells. Cell Motd. Cytoskel. 12, 1-11.
HAYASHI, M. AND TAKAHASHI, M. (1979). Ciliary adenosinetriphosphatase from a slow swimming mutant of Parameciumcaudatum J biol Chem 254, 11561-11665.
HINRICHSEN, R D , BURGESS-CASSLER, A., SOLTVEDT, B. C, HENNESSEY,T. AND KUNG, C. (1986). Restoration by calmodulin of a Ca2+-dependent K+ current missing in a mutant of Paramecium. Science232, 503-506.
KUZNICKI, L (1963). Reversible immobilization of Parameciumcaudatum evoked by nickel ions. Acta protozool. 1, 310—312.
LAKSEN, J., BAKALOW, K., HAMASAKI, T. AND SATIR, P. (1991). Structuraland functional characterization of Paramecium dynein: initial studies.J. Protozool. 38, 55-61.
LIEBERMAN, S. J., HAMASAKI, T. AND SATIR, P. (1988). Ultrastructureand motion analysis of penneabilized Paramecium capable of motilityand regulation of motility. Cell Motd Cytoskel. 9, 73-84.
LINDEMAN, C. B. AND GOLTZ, J. (1988). Calcium regulation of flagellarcurvature and Bwimming pattern in Triton-X-100-extracted rat spermCell Motil. Cytoskel. 10, 420^431.
LINDEMANN, C. B., FENTIB, I. AND RIKMBNSFOEL, R. (1980). A selectiveeffect of Ni2+ on wave initiation in bull sperm flagella. J. Cell Biol.87, 420-426.
MARTELL, A E. AND SMITH, R. M (1974). Critical Stability Constants,vol. 1. Plenum Press, NY.
MURPHY, J. AND RILEY, J. P. (1962). A modified single solution methodfor the determination of phosphate in natural waters. Analytica Chim.Acta 27, 31-36.
NAITOH, Y. AND KANEKO, H. (1972) Triton-extracted models ofParamecium. Modification of ciliary movement by calcium ions.Science 176, 523-524.
NAITOH, Y. AND KANEKO, H. (1973). Control of ciliary activities byadenosine tnphosphate and divalent cations in Triton-extractedmodels of Paramecium caudatum. J. exp. Bwl. 58, 657-676.
PASCHAL, B. M., KING, S. M., MOSS, A. G., COLLINS, C. A., VALLEE, R. B.AND WITMAN, G. B. (1987). Isolated flagella outer arm dyneintranslocates brain microtubules m vitro. Nature 330, 672-674.
REED, W. AND SATIR, P. (1986). Spreading ciliary arrest in mussel gillepithelium: characterization by quick fixation. J. cell. Physiol. 126,191-205.
SAIMI, Y. AND KUNG, C. (1987). Behavioral genetics of Paramecium. A.Rev. Genet. 21, 47-65
SALE, W. S. AND FOX, L. A. (1988). The isolated /3-heavy chain subunit
Effect of Ni2+ on axonemes 39
of dynein translocates microtubules in vitro. J. Cell Biol. 107,1793-1797.
SALS, W. S. AND SATIE, P. (1977). Direction of active sliding ofmicrotubules in Tetrahymena cilia. Proc. natn. Acad. Sci. U.S A. 74,2045-2050.
SATIR, P., GOLTZ, J., ISSAC, N. AND LINDEMANN, C. B. (1990). Switchingarrest in cilia and the calcium response of rat sperm: a comparison.Proc. VI Int. Cong. Comp. Physiology Siena, Italy (in press)
SATDI, P., HAMASAKI, T. AND LDJBKRMAN, S. J. (1988). Signaltransduction in microtubule-based motility: Switching mechanisms inthe control of axonemal activity in Paramecium. In CytoplasmicOrganization and Cell Motility (ed. Satir, P. et aU, pp. 219-234 AlanLias, N.Y.
SATIB, P., WAIS-STEIDBH, J., LBBDUSKA, S., NASR, A AND AVOLIO, J.(1981). The mechanochemical cycle of the dynein arm. Cell MotU. 1,303-327
SOLDO, A. T., GODOY, G. A. AND VAN WAGTENDONK, W. J. (1966). Growthof particle-bearing and particle-free Paramecium aurelia in aienicculture J. Protozool. 13<3), 492-497.
SUNDBKHO, S. A., MAQSUDUL, A. AND SPUDICH, J L. (1986). Excitationsignal processing times in Halobacterium halobium phototaxis.Bwphys. J. 50, 895-900.
TABTAH, V (1950) Methods for study and cultivation of protozoa. InStudies Honoring Trevor Kincaid (ed. Hatch, M. E.), pp. 104-105.University of Washington Press, Seattle, pp. 104-105.
TRAVIS, S. M. AND NELSON, D. L. (1988). Regulation of axonemal Mg5"1"-ATPase from Paramecium cilia- effect of Ca2+ and cyclic nucleotides.Biochim. biophys. Acta 966, 84-93.
VALE, R. D. AND TOYOSHIMA, Y. Y. (1988) Rotation and translocation ofmicrotubules in vitro induced by dyneins from Tetrahymena cilia. Cell52 459-469.
WAIS-STBIDER, J. AND SATIR, P. (1979). Effect of vanadate on gill cilia:switching mechanism in ciliary beat. J. supramolec. Struct. 11,339-347.
WARNER, F. D., PERREAULT, J. G. AND MCILVAJN, J. H. (1985). Rebindingof Tetrahymena 13S and 21S dynein ATPases to extracted doubletmicrotubules. The inner and outher row dynein arms. J. Cell Sci. 77,263-287
(Received 23 February 1990 - Accepted, in revised form,11 February 1991)
