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Cargo binding and regulatory sites in the tail of fungal conventional kinesin

Stephan Seiler*, Jochen Kirchner*, Christian Horn*, Athina Kallipolitou*, Günther Woehlke* and ManfredSchliwa*†

*Adolf Butenandt Institute, Cell Biology, University of Munich, 80336 Munich, Germany†e-mail: schliwa@bio.med.uni-muenchen.de

Here, using a quantitative in vivo assay, we map three regions in the carboxy terminus of conventional kinesin that are involved in cargo association, folding and regulation, respectively. Using C-terminal and internal deletions, point mutations, localization studies, and an engineered ‘minimal’ kinesin, we identify five heptads of a coiled-coil domain in the kinesin tail that are necessary and sufficient for cargo association. Mutational analysis and in vitro ATPase assays highlight a conserved motif in the globular tail that is involved in regulation of the motor domain; a region preceding this motif participates in folding. Although these sites are spatially and functionally distinct, they probably cooperate during activation of the motor for cargo transport.

onventional kinesin is an ATP-dependent motor enzyme thatmoves unidirectionally and processively alongmicrotubules1,2. It was initially purified from squid neural

tissue3, bovine brain4 and sea-urchin eggs5 and is now recognized asa component of many eukaryotic cells6. The heavy chain of conven-tional kinesin is an elongated molecule with a globular motordomain (head) at the amino terminus, a stalk consisting of severalcoiled-coil segments interrupted by presumably flexible linkers,and a small C-terminal globular domain. In animal kinesins, thetwo heavy chains interact at their C-terminal portions with twolight chains7–9. Light chains are nonessential for motor activity10, butare required for in vivo function11–13.

In four species of fungi, motors have been identified that sharemany features with conventional kinesins from animal species14–17.These similarities are more extensive than those to other kinesinfamilies, indicating that these motors may be fungal counterparts ofconventional kinesin. They seem to have analogous functions,although they are not necessarily homologous at all levels. No fun-gal kinesin has so far been reported to possess light chains.

In both fungal and animal kinesins, the motor domain containsthe microtubule-binding interface18 and the nucleotide-bindingsite19, but the domains that follow, termed neck linker, neck andhinge2, are also required for efficient motor function. Thus the necklinker, a region of ~15 amino acids, undergoes conformationalchanges that are thought to drive kinesin motility20. The neck regionalso contains elements that specify the direction of movement alongmicrotubules21–23 and support processive movement24, and the flex-ible hinge that follows contributes to mechanochemical coupling ofATPase activity and microtubule movement25.

The functions of the domains that comprise the extended stalkand tail regions are less well understood. In vitro studies provideevidence of an inhibitory action of the tail on the enzymatic andmotile activities of the motor domain26–28. Inhibition requires theability of the molecule to adopt a folded conformation26–29, which inturn critically depends on the presence of two presumably flexibleregions in the stalk30. Folding is essential for in vivo function, asNeurospora kinesin containing a mutation in one of the two flexibleregions performs poorly in vivo and does not fully rescue the nullmutant phenotype30. The ability to fold is an intrinsic property ofthe kinesin heavy chain and does not depend on the presence oflight chains26–28.

Together, these findings indicate that kinesin may be in a folded,inhibited state when not bound to cargo in the cell. Is cargo bindingalone sufficient to activate the motor? Where is the cargo bindingsite, and how are cargo binding and motor activity coordinated? Toaddress these questions, we have used an in vivo model system30 that

monitors the functional consequences of manipulations that affectcargo binding and tail-mediated regulation of motor activity. Thismodel is unique in that it allows kinesin function to be studied inliving cells using a quantitative assay. It involves transformation ofkinesin complementary DNAs, modified at desired positions, intoa kinesin-deficient strain of Neurospora; growth rate is used as anindicator of functionality. In combination with localization studiesof Myc-tagged mutant kinesins and in vitro assays of folding andATPase activity, we have been able to identify a cargo-interactionsite and a motif involved in the regulatory interaction between thetail and motor domains.

ResultsLocalization of Myc-tagged kinesin. Several lines of evidence indi-cate that conventional kinesin may be predominantly involved inmembrane transport6,31,32. Its C-terminal portion has long beenthought to be responsible for cargo binding8,12,33,34, but thedomain(s) involved have not been identified. To study cargo asso-ciation in a fungal kinesin, we generated Myc-tagged versions offull-length and mutant motors and investigated their subcellularlocalization in growing hyphae after transformation into a kinesin-deficient strain of Neurospora. Previous studies using kinesin anti-bodies always resulted in a punctate staining pattern even inkinesin-null strains, implying either unspecific labelling or crossre-action with other kinesin-like proteins (S.S. and M.S., unpublishedobservations). The Myc epitope tag was fused to the kinesin N ter-minus and was recognized by a specific antibody. When probedwith an anti-Myc antibody, cells of a kinesin-null strain trans-formed with full-length Myc–kinesin exhibited a punctate labellingpattern superimposed upon weak cytoplasmic staining (Fig. 1b). Incontrast, neither wild-type nor untransformed kinesin-null cellsexhibited any labelling (Fig. 1b). Myc-tagged kinesin constructswere indistinguishable from their untagged counterparts in in vitromicrotubule-gliding assays and in the extent of their rescues of thenull phenotype after transformation, showing that the Myc tag doesnot interfere with motor function.Identification of a cargo-interaction site in the kinesin tail. Previ-ous studies have shown that the growth rate of Neurospora crassa inrace tubes can be used as a reliable reporter of the ability of mutantkinesins to suppress the null phenotype30. We combined this assaywith localization studies of kinesin deletion mutants. C-terminaldeletions ranging from only 6 (Kin(∆923–928)) to ~100 aminoacids (Kin(∆830–928)) resulted in mutants with growth rates inter-mediate between those of the wild type and of null mutants (Fig.1a), but which exhibit the same punctate staining pattern as control

C

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cells when probed with anti-Myc antibody (Fig. 1b). In contrast,Kin(∆740–928), which lacks the globular tail and the coiled-coildomain preceding it (coil 3), failed to rescue the null phenotype atall. In this mutant, the punctate staining pattern was absent and wasreplaced by uniform cytoplasmic staining (Fig. 1b). In accordancewith several localization studies using other cell types, in which thecargo of kinesin appeared as puncta or small particles35–37, we pro-pose that the punctate structures observed here represent the cargoof kinesin, which includes small vesicles destined for secretion at thehyphal tip15,38. Thus, the region between residues 740 and 830 seemsto contain a crucial site for cargo binding.

To locate the cargo-binding element more accurately within thiscoiled-coil region, we generated stepwise deletions of two heptadseach, ensuring that the remaining heptads were joined such that acontinuous coiled coil was formed. Deletion of the two heptadscomprising residues 787–800 resulted in a mutant kinesin that wasunable to rescue the null phenotype (Fig. 1c), indicating that thisregion may be essential for cargo binding.

To investigate the specificity of the cargo-interaction site, wegenerated a series of point mutations in the region between residues780 and 830. We targeted charged amino acids for mutations andavoided substituting residues at positions a or d within heptads(Fig. 2a). In single mutations we reversed the charge; in double andtriple mutations we replaced lysine or arginine with alanine andglutamate with glutamine. The propensity to form a coiled-coileven in a stringent 14-amino-acid window39 was unaltered in all sin-gle and double point mutants and reduced only in triple mutants.

Point mutations in the region between residues 785 and 814resulted in a severely reduced growth rate (15–55% that of wild-type cells; Fig. 2b), whereas point mutations in regions N-terminalor C-terminal to this had hardly any effect. Thus, the triple muta-tions Kin(KKE812–814AAQ) and Kin(ERK819–821QAA) arelocated in identical positions within adjoining heptads and yetresulted in vastly different growth rates (Fig. 2b). Immunofluores-cence microscopy of Myc-tagged point mutants showed that punc-tate staining was reduced, but not abolished (data not shown).

In general, deletions or point mutantions resulted in a loss-of-function phenotype. To confirm the crucial function of the kinesin783–820 region in cargo binding, we produced a ‘minimal’ kinesinconstruct composed of the functional motor domain25 (head/neck/hinge), the beginning of coil 1 (to ensure dimerization) and theputative cargo-binding element consisting of residues 783-820 (Fig.2c). Transformation of cells with this severely truncated version ofkinesin resulted in a gain-of-function phenotype with a hyphalgrowth rate of 2.2 cm day–1 (Fig. 2d). This construct also restored the

Figure 1 Rescue of the kinesin-null phenotype and immunolocalization of Myc-tagged kinesin constructs. a, Overview of the domain organization of Neurospora kinesin (top) and rescue by C-terminal-deletion constructs. WT represents both wild-type Neurospora and a kinesin-deletion strain transformed with full-length kinesin; ∆Kin represents a kinesin-deletion mutant. Values are means from at least three experiments; s.d. ≤10% of the mean. b, Immunofluorescence microscopy of Myc-tagged constructs using an anti-Myc antibody. ‘Wild type’ represents kinesin-null mutants transformed with Myc-tagged full-length kinesin. Labelling specificity is shown by the lack of staining in ∆Kin-transformed cells (bottom panel) and in untransformed kinesin-null mutants (data not shown). Scale bar represents 10 µm. c, Complementation of kinesin-null mutants by constructs containing double-heptad deletions in coil 3. Values are means from at least three experiments; s.d. ≤10% of the mean.

Wild type

Kin(∆830–928)

Kin(∆740–928)

∆Kin

a b

c

Head Stalk TailCoil 1 Coil 2 Coil 3 Growth

(cm day–1)

6.8

3.6

3.5

1.2

1.4

WT

Kin(∆923–928)

Kin(∆830–928)

Kin(∆740–928)

Kin(∆757–770)

Kin(∆771–786)

Kin(∆787–800)

Kin(∆801–814)

Kin(∆815–828)

∆Kin

3.8

4.1

1.3

4.6

5.2

Figure 2 Point mutations in the cargo-binding region, and ‘minimal’ kinesin.

a, Sequence alignment of two fungal kinesins from Neurospora (Nkin) and Syncephalastrum (Synkin) and two animal kinesin heavy chains from the mouse (MmKHC) and Drosophila (DmKHC) in the presumptive cargo-binding region (residues 781–830 in Neurospora kinesin). * denotes residues conserved across all species shown. Point mutations were introduced in the positions marked in red and transformed into the null mutant. The region marked in grey was used for the construction of minimal kinesin. b, Growth rates of strains transformed with full-length kinesin carrying the indicated mutations in the presumptive cargo-binding region. Growth rate of the wild type was designated as 100% and that of the null mutant as 0% (see ref. 30). c, Upper panel, schematic representation of the minimal kinesin construct, consisting of residues 1–481 (head/neck/hinge and the beginning of coil 1) fused directly to residues 783–820 (marked in grey in a). Lower panel, rescue of the null phenotype by minimal kinesin, measured as in b. d, Western blotting,using an anti-Myc antibody, of hyphal cytoplasmic extracts from strains transformed with minimal (MMK) and full-length kinesin (Nkin), both carrying the Myc tag. e, Immunolocalization of minimal kinesin within a hypha. Scale bar represents 10 µm.

a

b

c

d e

Point mutation

Wild type 6.8 100

Kin(R781E) 6.8 100

Kin(KK785–786AA) 3.6 ~40

Kin(ER791–792QA) 2.9 ~25

Kin(E795K) 3.9 ~45

Kin(R802E) 4.6 ~55

Kin(E806K) 3.8 ~40

Kin(KKE812–814AAQ) 2.3 ~15

Kin(ERK819–821QAA) 6.3 ~90

Kin(ER827–828QA) 6.1 ~90

Growth(cm day–1)

Null-phenotyperescue (%)

Head Neck Coil 1783–820

481

Growth(cm day–1)

Null-phenotyperescue (%)

2.2 ~15

116 –96 –

66 –

45 –

MMK Nkin

NkinSynkin

* * ** * * * ** * * ** * * * * * **781794824849

Mr (K)

MmKHCDmKHC

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punctate staining pattern (Fig. 2e), showing that the region betweenresidues 783 and 820 is both necessary and sufficient to produce astaining pattern consistent with cargo association.An essential regulatory site in the tail. The presence of a cargo-binding site is insufficient for full complementation of the null phe-notype, as shown by deletions more C-terminal to the cargo-bind-ing region (Kin(∆830–928); Fig. 1a). These domains may thereforeinclude sequence elements with further important, but separate,functions. Several C-terminal mutations resulted in an unusualstaining pattern when probed with anti-Myc antibody. In additionto the characteristic punctate labelling superimposed upon weakcytoplasmic staining, a ~40-fold increase in staining intensity wasobserved in the apical 10–20 µm of tip hyphae (Fig. 3a).

This hyphal accumulation was observed in all deletion mutantslacking >30 residues from the extreme C terminus. The transitionfrom a wild-type staining pattern to localized staining at the tipoccured in deletions close to a highly conserved motif in the C ter-minus (886RIAKPLR892 in Neurospora kinesin). Thus, Kin(∆907–928) exhibits uniform staining, whereas Kin(∆885–928) exhibitsintense tip staining (Fig. 3a). As the deletion mutant that lacks the

cargo-association domain (Kin(∆740–928)) also causes tip stain-ing, this may represent accumulation of unloaded, cargo-freekinesin.

To confirm the importance of the conserved RIAKPLR motif intip localization, we generated single (Kin(K889E)) and double(Kin(R886E, K889E)) point mutations in this motif. These mutantswere indistinguishable from the deletion mutant Kin(∆885–928).They exhibited similarly reduced growth rates (4.1 and 3.6 cm day–1,respectively, compared to 3.8 cm day–1 for Kin(∆885–928)) andshowed intense tip staining (Fig. 3a).

To investigate the function of the RIAKPLR motif, we measuredthe microtubule-stimulated ATPase activity of truncated kinesinKin(1–433) in the presence of wild-type kinesin tail peptide or of amutant containing two point mutations at charged residues withinthe RIAKPLR motif, giving the sequence EIAEPLR. The wild-typetail caused a substantial reduction in the ATPase activity of the headconstruct (mean activity 41% of that of wild-type kinesin; Table 1),which is consistent with previous findings27. The mutant EIAEPLRmotif produced only a small reduction in ATPase turnover (meanactivity 83% of that of wild-type; Table 1), despite the fact that, inall experiments, the concentration of mutant tail peptide was higherthan or equal to the concentration of wild-type tail. We also meas-ured the ATPase rate of the head construct in the presence of 15 µMpolymerized tubulin over a range of tail-peptide concentrations(40–200 nM). Although saturating concentrations of tail peptidecould not be reached, results showed that the mutant tail constructhas a consistently lower affinity for the motor domain (data notshown).Consequences of impaired folding for in vivo function. Tip stain-ing was observed in a mutant (Kin(∆551–634)) that lacks the ‘kink’,a flexible domain linking the two principal coiled coils of the stalk.When assayed in sucrose gradients (Fig. 3b), the kink-deletionmutant was unable to attain a compact conformation and did notalter its salt-dependent sedimentation behaviour. Although it pos-sesses fully functional N and C termini, including the cargo-bindingregion and the RIAKPLR motif, it rescued the null phenotype onlypartially (growth rate 4.2 cm day–1) and resulted in apical mislocali-zation (Fig. 3c).

The results obtained using Kin(∆551–634) indicate that thekink may be essential for the ability of the motor to adopt a com-pact conformation in vitro. However, in addition to the kink, fold-ing requires a region of the C terminus between residues 859 and885. Although the deletion mutant Kin(∆885–928) was able toadopt a compact conformation in vitro, Kin(∆859–928) exhibiteda higher sedimentation coefficient (Fig. 3d) and may thereforeremain extended even at low salt concentrations. The precisemanner in which this region participates in the formation of a sta-bly folded state is unclear, but it may interact with the head/neck/

Figure 3 Mutants exhibiting kinesin accumulation at hyphal tips. a, Localization of the indicated kinesin mutants in distal regions of hyphae. Brightness and contrast were optimized for tip staining; staining along the lengths of hyphae therefore appears much reduced. Scale bar represents 10 µm. b, Sucrose-gradient sedimentation of wild-type kinesin and the kink-deletion mutant at high (1 M KCl; HS) and low (buffer only; LS) salt concentrations. Sedimentation coefficients of the peak fractions from the wild type are 8.8 S (LS) and 6.3 S (HS); that of the kink-deletion mutant is 6.0 S at both salt concentrations. The positions of the marker proteins catalase (11.3 S), aldolase (7.4 S) and BSA (4.2 S) are also shown. c, Tip localization of the kink deletion mutant. Scale bar represents 10 µm. d, Sucrose-gradient sedimentation of the indicated deletion mutants at high and low salt concentrations as in a. Both mutants exhibit tip localization in vivo (see a, Kin(∆885–928).

Wild type, LS

a

bc

d

Wild type Kin(∆907–928)

Kin(∆885–928) Kin(K889E)

Kin(∆551–634)

Wild type, HS

11.3 S

7.4 S 4.2 S

11.3 S

7.4 S 4.2 S

Sta

inin

g in

tens

ity(a

rbitr

ary

units

)S

tain

ing

inte

nsity

(arb

itrar

y un

its)

10 15 20 25 3015% sucrose Fraction no. 5% sucrose

Kin(∆551–634), LSKin(∆551–634), HS

Kin(∆885–928), LSKin(∆885–928), HSKin(∆859–928), LSKin(∆859–928), HS

15% sucrose Fraction no. 5% sucrose

Figure 4 Summary of the functional features of kinesin tail domains and the effects of the mutations studied here. The schematic drawing (top) shows an overview of the C-terminus of Neurospora kinesin. Positions of the C-terminal-deletion mutants (long arrows) and of point mutations or internal deletions (short arrows) are indicated. The second row of arrows indicate the functional consequences of the mutations.

Stalk

(not toscale)

Tail coiled coil Globular tail

Kink(folding)

Neckinteraction

RIAKPLR(Regulation)

MinimalkinesinCargo binding 928

Kin(∆551–634)

Kin(∆740–928)

Null-pheno-type rescuePunctatestainingTip staining

Partial Partial Partial Partial Partial Partial Partial

Folding

+ + + + + +

+ + + + +

+ ++– – –

– – –

–

–

Reduced

+ n.a.

Pointmutation

Pointmutation

Kin(∆830–928)

Kin(∆859–928)

Kin(∆885–928)

Kin(∆907–928)

Kin(∆923–928)

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hinge region when the motor is inactive26,30.

DiscussionThe Neurospora model system has allowed a functional analysis ofthe kinesin C terminus in an in vivo context. The data presentedhere are summarized in Fig. 4, which lists the relative positions ofthe various mutants and their effects on kinesin localization andfunction. A region essential for cargo binding lies in a conservedcoiled-coil segment (red). In addition to the kink (yellow), a smallportion of the tail (blue) participates in maintenance of the foldedconformation; a presumably regulatory motif (purple) is immedi-ately C-terminal to this. As discussed below, the activities associatedwith these regions can be functionally and, to some extent, experi-mentally separated from one another. These findings contribute toa refined model for kinesin-tail inhibition26–28 and regulation ofkinesin activity in vivo.

Unexpectedly, the cargo-binding region is located in anextended coiled coil adjacent to the globular tail domain. Cargoassociation is unlikely to involve melting of the coiled coil, as thecargo-binding region consists of a classical leucine zipper motifthat is predicted to form a stable coiled coil39. Binding seems to bemediated by ionic interactions, as point mutations at charged res-idues in this region strongly affect the in vivo function of themotor. In animal kinesins, salt-sensitive interactions with pre-sumptive cargo(es) have been observed in some cases34,40,41,although other reports indicate that cargo association may bemore stable42,43. These differences may be explained in part by dif-ferences in the assay system, or the presence or absence of lightchains. In Neurospora, kinesin seems to be involved in apicaltransport of submicroscopic vesicles that form the‘Spitzenkörper’. These vesicles, which are ultimately destined forsecretion44, must be continuously supplied to the tip to sustainapical growth. We propose that the observed punctate stainingpattern (Fig. 1b) represents these vesicles, although we cannotrule out the association of kinesin with other cellular components.Note that the potential binding site identified here is highly con-served among animal (~90% identity) and fungal (~85% identity)conventional kinesins, and well conserved between the twogroups (57% identity). Whether this also implies conservation ofthe mechanisms of cargo binding across species remains to bedetermined. Thus, in animal kinesins, light chains may have afunction in cargo binding, as antibodies against, or the deletion of,light chains interfere with cargo association and cause heavy-chain mislocalization11-13,45. Interestingly, the site for light-chain-binding has been mapped to a region preceding the cargo-interac-tion site identified here9. The corresponding region in fungalkinesins shares no similarity with that of animal kinesins, but ishighly conserved within this group46. This highlights a significantdiversion between fungal and metazoan kinesins.

Previous studies have shown that animal conventional kinesincan adopt a folded conformation at low salt concentrations invitro29. Folding requires the kink and allows the tail domain to exertan inhibitory effect on the motor domain26–28. Here we have

extended the importance of folding to a fungal kinesin and to the invivo situation, although further requirements have to be met as cer-tain mutant motors that possess both the kink and the cargo-bind-ing region still perform poorly in vivo. These findings argue againstmodels in which cargo binding alone is sufficient to activate themotor, in accordance with observations of embryonic, membrane-associated kinesin that becomes activated in a subsequent step47.

Our observations indicate that the conserved RIAKPLR motifin the globular tail domain may have an important function thatis unrelated to cargo binding. When this motif is deleted ormutated, kinesin accumulates at the tip. The same phenotype isobserved upon the deletion of the kink, which impairs the abilityof the motor to adopt a folded conformation in vitro. In addition,the ATPase activity of the motor domain can be repressed by a tailpeptide with a wild-type RIAKPLR motif, but much less so by atail peptide with two point mutations in this motif. There are sev-eral possible explanations of the tip-staining phenotype. First, theRIAKPLR motif may be involved in kinesin degradation36, whichwould lead to accumulation of kinesin at the tip if this motif isdeleted or mutated. However, the fact that this motif has an activefunction in regulation of the ATPase activity of the motor domainseems to be inconsistent with an exclusive degradation function.Unless unfolding renders the RIAKPLR motif unable to effectdegradation, observations of the kink mutant, in which this motifis intact, are also inconsistent with this possibility. Alternatively,the RIAKPLR motif may have a function in folding that involvesits interaction with the head/neck region of the motor. The factthat the deletion mutant Kin(∆885–927), which contains theRIAKPLR motif, can still fold at low salt concentrations seems toargue against this possibility. However, the equivalent deletion inDrosophila kinesin (DKH945), which splits the conserved RIAK-PLR motif, can still fold but is more labile (that is, it unfolds atintermediate salt concentrations)26. Thus, the RIAKPLR motif ofNeurospora kinesin may contribute to stabilization of the head–tail interaction, but is not absolutely required. A final possibility,which we currently favour, is that the RIAKPLR motif may beinvolved in regulation of motor activity. The strongest support forthis idea comes from the in vitro ATPase assays, which indicatethat this motif may directly influence the motor domain. If theinteraction between the tail and the head/neck regions of themotor is mediated primarily by the the 30 amino acids that pre-cede the RIAKPLR motif, as indicated by the data presented hereas well as in previous work26, the globular C terminus that carriesthe RIAKPLR motif would come to lie close to the motor domain,where it could exert a direct influence on motor activity. In animalkinesins, the interaction of the tail with the motor domain seemsto regulate both the ATPase activity26,27 and the motilityproperties28 of the motor. The data presented here indicate that, infungal kinesins, this function may be specifically associated withthe RIAKPLR motif. This idea is also supported by results fromthe deletion and point mutations, which cause misregulation andmislocalization at the tip (Fig. 3a), but do not interfere with theability of the motor to fold (Fig. 3d) or to bind cargo (Fig. 1b).Conversely, mutations in the cargo-binding site do not result in

Table 1 ATPase rates of a truncated Neurospora kinesin head construct (Kin(1–433)) in the presence of wild-type (RIAKPLR) or mutant (EIAEPLR) tail constructs.

Experiment Kin(1–433) alone Kin(1–433) + RIAKPLR Kin(1–433) + EIAEPLRkcat (s

–1)*

* kcat (s–1) refers to the rate of ATP turnover per kinesin dimer.

Tail conc. (nM) kcat (s–1) % of Kin(1–433)

aloneTail conc. (nM) kcat (s

–1) % of Kin(1–433) alone

1 79.2 125 18.2 23 300 54.3 692 79.8 158 41.6 52 200 70.6 893 113.6 15 32.2 28 15 85.0 754 61.0 38 37.4 61 45 60.0 97

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tip staining (that is, misregulation) because impaired cargo bind-ing does not necessarily also affect the function of the RIAKPLRmotif.

A simple model that explains how the sites identified here arefunctionally linked is that the first step in activation (or suppressionof inhibition) of the folded motor is occupation of the cargo-bind-ing region. This may be sensed by the globular tail, which, possiblythrough a conformational change, releases the tail site involved infolding and activates (or derepresses) the motor domain by way ofthe RIAKPLR motif. All of these sites cooperate in, and are thereforerequired for, in vivo function. Undoubtedly, other regulatorymechanisms (such as phosphorylation, light chains, other associ-ated proteins, and receptors) are involved and will modify the basicactivation scheme proposed here. These levels of regulation are notexcluded by the model; on the contrary, they can be expected totune further the activity of kinesin by affecting the sequence ofevents outlined here at various levels. Future work will have to iden-tify the partner domains that interact with the key sites specifiedhere. h

MethodsCells.Media, growth conditions for hyphae, and transformation procedures were as described44. The linear

growth rate was assayed in ‘race tubes’ — hollow glass tubes of 1-cm diameter, half-filled with Vogel’s

minimal agar and inoculated with conidia at one end30. Mycelial growth was recorded over time; at least

three experiments were carried out for each construct30.

Microscopy.For immunofluorescence microscopy, cells were grown overnight on dialysis membrane and fixed for 30

min with 2% formaldehyde and 0.5% glutaraldehyde in PEM-buffer (50 mM PIPES, 5 mM MgSO4 and

5 mM EGTA, pH 6.8). Cell walls were digested with 1 mg ml–1 novozym 234 (InterSpex, Foster City,

California) for 15 min at room temperature, in buffer containing a proteolysis-inhibitor cocktail14.

Subsequent steps were as described48. Preparations were viewed using a Zeiss Axiophot microscope

(Zeiss, Oberkochen, Germany). Staining intensities were quantified using NIH Image software.

Wild-type and mutant kinesins were isolated from hyphal extracts and tested in in vitro microtubule-

gliding assays as described14. They were found to generate microtubule gliding at rates comparable to

those of wild-type kinesin (~2.5 µm s–1 ), showing that motor activity was unaffected.

Sucrose density gradients and blotting.For analysis of motor conformation, 100 µl of cytosolic extract was fractionated on linear 5–15% sucrose

gradients in the presence or absence of 1 M KCl, in a buffer containing 10 mM PIPES, 0.5 mM EGTA and

1 mM MgCl2, pH 6.9. These two extremes were chosen for optimal demonstration of differences in

sedimentation. Concentrations lower than 1 M KCl led to peak broadening. Catalase, aldolase and BSA

were used as standards. Gradients were centrifuged for 2.5 h at 55,000 r.p.m. using a Beckman TLS55

rotor. Fractions were collected from the bottom, separated on 7.5% polyacrylamide gels, blotted onto

nitrocellulose membranes using standard procedures14, and probed with an antibody against the Myc-

epitope tag49. Blots were scanned using the EagleEye II system (Stratagene) and analysed using NIH

Image software.

Generation of kinesin constructs.DNA encoding the 11-amino-acid Myc epitope was fused to Neurospora kinesin cDNA, at the end

encoding the N terminus, by the polymerase chain reaction (PCR), and was cloned into a vector derived

from pCSN43 (Fungal Genetics Stock Center, Kansas City, Kansas) using a ClaI restriction site

immediately 3′ of the Aspergillus bimC promoter. The vector also contained a bleomycin-resistance

cassette downstream of the coding sequence. For C-terminal deletions, the clones used were as

described30. EcoRI–AatII restriction fragments of the C termini were excised and used as replacements for

the C terminus in the Myc–kinesin vector. C-terminal, two-heptad deletions, as well as the kink deletion,

were generated by PCR, using a 5′-primer containing 18 base pairs upstream and downstream of the

desired deletion. The PCR product was then used as a primer in a second PCR reaction30 and cloned into

the Myc–kinesin vector. Point mutations were generated using the same protocol, using a shorter primer

containing the base exchange. All PCR-generated fragments were sequenced (TopLab, Munich) and

found to be error-free. As shown previously, expression levels of mutant constructs varied between 10%

and 250% of those of the wild type, but were not rate-limiting30.

ATPase assays.The constructs used in ATPase assays included a C-terminal construct, comprising residues 782–928,

that was cloned into the pGEX 2T vector and expressed in E. coli as a glutathione-S-trasferase (GST)

fusion protein. A homologous construct carrying two point mutations in the RIAKPLR motif

(EIAEPLR), generated by excision from the correponding full-length mutant, was also cloned into the

pGEX 2T vector. An N-terminal construct of the motor domain, Kin(1–433), was cloned into the pT7-7

vector and expressed in E. coli using the T7 expression system. The N-terminal construct was expressed

in E. coli strain BL-21 and purified by phosphocellulose (P11)-column chromatography, followed by S-

sepharose-column chromatography. To determine microtubule-dependent ATPase activity, a coupled

enzymatic assay50 was used as described25. Assays were carried out in the presence of 0–15 µM

microtubules and a fixed concentration of tail peptide to determine the kcat, or of near-saturating

concentrations of microtubules (15 µM) to assay the dependence on tail-peptide concentration. The

assay mixture contained 20 µM Taxol and 0.5 mM ATP in ACES buffer (12.5 mM ACES/KOH pH 6.8, 2

mM Magnesium acetate and 0.5 mM EGTA). The motor domain construct Kin(1–433) was used at 11–

44 nM. Wild-type and mutant tail constructs were used immediately from bacterial extracts because

degradation occurred during further purification steps; concentrations were determined in Coomassie-

stained polyacrylamide gels by comparison to known protein standards. kcat values were determined by

fitting a hyperbolic curve to a plot of microtubule concentration against ATPase rate.

RECEIVED 13 OCTOBER 1999; REVISED 31 MARCH 2000; ACCEPTED 14 APRIL 2000; PUBLISHED 9 MAY 2000.

1. Howard, J. The movement of kinesin along microtubules. Annu. Rev. Physiol. 58, 703–729 (1996).

2. Vale, R. D. & Fletterick, R. The design plan of kinesin motors. Annu. Rev. Cell Dev. Biol. 13, 745–777

(1997).

3. Vale, R. D., Reese, T. S. & Sheetz, M. P. Identification of a novel force-generating protein, kinesin,

involved in microtubule-based motility. Cell 42, 39–50 (1985).

4. Brady, S. T. A novel brain ATPase with properties expected for the fast axonal transport motor.

Nature 317, 73–75 (1985).

5. Scholey, J. M., Porter, M. E., Grissom, P. M. & McIntosh, J. R. Identification of kinesin in sea urchin

eggs and evidence for its localization in the mitotic spindle. Nature 318, 483–486 (1985).

6. Hirokawa, N. Kinesin and dynein superfamily proteins and the mechanism of organelle transport.

Science 279, 519–526 (1998).

7. Amos, L. A. Kinesin from pig brain studied by electron microscopy. J. Cell Sci. 87, 105–111 (1987).

8. Hirokawa, N. et al. Submolecular domains of bovine brain kinesin identified by electron microscopy

and monoclonal antibody decoration. Cell 56, 867–878 (1989).

9. Diefenbach, R. J., Mackkay, J. P., Armati, P. J. & Cunningham, A. L. The C-terminal region of the

stalk domain of ubiquitous human kinesin heavy chain contains the binding site for kinesin light

chain. Biochemistry 37, 16663–16670 (1998).

10. Yang, J. T., Saxton, W. M., Stewart, R. J., Raff, E. C. & Goldstein, L. S. B. Evidence that the head of

kinesin is sufficient for force generation and motility in vitro. Science 249, 42–47 (1990).

11. Gindhart, J. G., Desai, C. J., Beushausen, S., Zinn, K. & Goldstein, L. S. B. Kinesin light chains are

essential for axonal transport in Drosophila. J. Cell Biol. 141, 443–454 (1998).

12. Verhey, K. J. et al. Light-chain-dependent regulation of kinesin’s interaction with microtubules. J.

Cell Biol. 143, 1053–1066 (1998).

13. Rahman, A., Kamal, A., Roberts, E. A. & Goldstein, L. S. Defective kinesin heavy chain behavior in

mouse kinesin light chain mutants. J. Cell Biol. 146, 1277–1288 (1999).

14. Steinberg, G. & Schliwa, M. The Neurospora organelle motor: a distant relative of conventional

kinesin with unconventional properties. Mol. Biol. Cell 6, 1605–1618 (1995).

15. Lehmler, C. et al. Identification of a motor protein required for filamentous growth in Ustilago

maydis. EMBO J. 16, 3464–3473 (1997).

16. Steinberg, G. A kinesin-like mechanoenzyme from the zygomycete Syncephalastrum racemosum

shows biochemical similarities with conventional kinesin from Neurospora crassa. Eur. J. Cell Biol. 73,

124–131 (1997).

17. Wu, Q. et al. A fungal kinesin required for organelle motility, hyphal growth and morphogenesis.

Mol. Biol. Cell 9, 89–101 (1998).

18. Woehlke, G., Ruby, A. K., Hart, C. L., Hom-Booher, N. & Vale, R. D. Microtubule interaction site of

the kinesin motor. Cell 90, 207–216 (1997).

19. Kuznetsov, S. A., Vaisberg, Y. A., Rothwell, S. W., Murphy, D. B. & Gelfand, V. I. Isolation of a 45

kDa fragment from the kinesin heavy chain with enhanced ATPase and microtubule binding

affinities. J. Biol. Chem. 264, 589–595 (1989).

20. Rice, S. et al. A structural change in the kinesin motor protein that drives motility. Nature 402, 778–

784 (1999).

21. Case, R. B., Pierce, D. W., Hom-Booher, N., Hart, C. L. & Vale, R. D. The directional preference of

kinesin motors is specified by an element outside of the motor catalytic domain. Cell 90, 959–966

(1997).

22. Henningsen, U. & Schliwa, M. Reversal in the direction of movement of a molecular motor. Nature

389, 93–96 (1997).

23. Endow, S. A. & Waligora, K. W. Determinants of kinesin motor polarity. Science 281, 1200–1202

(1998).

24. Romberg, L., Pierce, D. W. & Vale, R. D. Role of the kinesin neck region in processive microtubule-

based motility. J. Cell Biol. 140, 1407–1416 (1998).

25. Grummt, M. et al. Importance of a flexible hinge near the motor domain in kinesin -driven motility.

EMBO J. 17, 5536–5542 (1998).

26. Stock, M. F. et al. Formation of the compact confomer of kinesin requires a COOH-terminal heavy

chain domain and inhibits microubule-stimulated ATPase activity. J. Biol. Chem. 274, 14617–14623

(1999).

27. Coy, D. L., Hancock, W. O., Wagenbach, M. & Howard, J. Kinesin’s tail domain is an inhibitory

regulator of the motor domain. Nature Cell Biol. 1, 288–292 (1999).

28. Friedman, D. S. & Vale, R. D. Single-molecule analysis of kinesin motility reveals regulation by the

cargo-binding tail domain. Nature Cell Biol. 1, 293–297 (1999).

29. Hackney, D. D., Levitt, J. D. & Suhan. J. Kinesin undergoes a 9S to 6S conformational transition. J.

Biol. Chem. 267, 8696–8701 (1992).

30. Kirchner, J., Seiler, S., Fuchs, S., & Schliwa, M. Functional anatomy of the kinesin molecule in vivo.

EMBO J. 18, 4404–4413 (1999).

31. Bloom, G. S. & Endow, S. A. Motor proteins 1: kinesins. Protein Profile 2, 105–171 (1995).

32. Lane, J. & Allan, V. Microtubule-based membrane movement. Biochim. Biophys. Acta 1376, 27–55

(1998).

33. Goldstein, L. S. B. With apologies to Sheherazade: tails of 1001 kinesin motors. Annu. Rev. Genet. 27,

319–351 (1993).

34. Skoufias, D. A., Cole, D. G., Wedaman, K. P. & Scholey, J. M. The carboxy-terminal domain of

kinesin heavy chain is important for membrane binding. J. Biol. Chem. 269, 1477–1485 (1994).

35. Pfister, K. K., Wagner, M. C., Stenoien, D. L., Brady, S. T. & Bloom, G. S. Monoclonal antibodies to

kinesin heavy and light chains stain vesicle-like structures, but not microtubules, in cultured cells. J.

Cell Biol. 108, 1453–1463 (1989).

36. Hirokawa, N. et al. Kinesin associates with anterogradely transported membraneous organelles in

vivo. J. Cell Biol. 114, 295–302 (1991).

© 2000 Macmillan Magazines LtdNATURE CELL BIOLOGY | VOL 2 | JUNE 2000 | www.nature.com/ncb 337

articles

37. Wright, B. D. et al. Subcellular localization and sequence of sea urchin kinesin heavy chain —

evidence for its association with membranes in the mitotic apparatus and interphase cytoplasm. J.

Cell Biol. 113, 817–833 (1991).

38. Seiler, S., Plamann, M. & Schliwa, M. Kinesin and dynein mutants provide novel insights into the

roles of vesicle traffic during cell morphogenesis in Neurospora. Curr. Biol. 9, 779–785 (1999).

39. Berger, B. et al. Predicting coiled coils by use of pairwise residue correlations. Proc. Natl Acad. Sci.

USA 92, 8259–8263 (1995).

40. Jellali, A. et al. Structural and biochemical properties of kinesin heavy chain associated with rat brain

mitochondria. Cell Motil. Cytoskeleton 28, 79–93 (1994).

41. Yu H., Toyoshima I., Steuer E. R. & Sheetz M. P. Kinesin and cytoplasmic dynein binding to brain

microsomes. J. Biol. Chem. 267, 20457–20464 (1992).

42. Schnapp, B. J., Reese, T. S. & Bechtold, R. Kinesin is bound with high affinity to squid axon organelles

that move to the plus end of microtubules. J. Cell Biol. 119, 389–399 (1992).

43. Leopold, P. L., McDowall, A. W., Pfister, K. K., Bloom, G. S. & Brady, S. T. Association of kinesin with

characterized membrane-bound organelles. Cell Motil. Cytoskeleton 23, 19–33 (1992).

44. Seiler, S., Nargang, F., Steinberg, G. & Schliwa, M. Kinesin is essential for cell morphogenesis and

polarized secretion in Neurospora crassa. EMBO J. 16, 3025–3034 (1997).

45. Stenoien, D. S. & Brady, S. T. Immunochemical analysis of kinesin light chain function. Mol. Biol. Cell

146, 1277–1288 (1997).

46. Kirchner, J., Woehlke, G. & Schliwa, M. Universal and unique features of kinesin motors: insights

from a comparison of fungal and animal kinesins. Biol. Chem. 380, 915–921

(1999).

47. Lane, J. D. & Allan, V. J. Microtubule-based endoplasmic reticulum motility in Xenopus laevis:

activation of membrane-associated kinesin during development. Mol. Biol. Cell. 10, 1909–1922

(1999).

48. Tinsley, J. H., Minke, P. F., Bruno, K. S. & Plamann, M. p150 Glued, the largest subunit of the

dynactin complex, is nonessential in Neurospora but required for nuclear distribution. Mol. Biol. Cell

7, 731–742 (1996).

49. Evan, G. I., Lewis, G. K., Ramsey, G. & Bishop, J. M. Isolation of monoclonal antibodies specific for

human cMyc-proto-oncogene product. Mol. Cell. Biol. 5, 3610–3616 (1985).

50. Huang, T. & Hackney, D. D. Drosophila kinesin minimal motor domain expressed in E. coli.

Purification and kinetic characterization. J. Biol. Chem. 269, 16493–16501 (1994).

ACKNOWLEDGEMENTS

We thank S. Fuchs for technical assistance and U. Euteneuer for critical reading of the manuscript. This

work was supported by the Deutsche Forschungsgemeinschaft, the Volkswagen Stiftung and the Fonds

der Chemischen Industrie.

Correspondence and requests for materials should be addressed to M.S.

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