Abstract
Kinesin-1 is a plus-end microtubule-based motor, and defects in kinesin-based transport are linked to diseases including neurodegeneration. Kinesin can auto-inhibit via a head-tail interaction, but is believed to be active otherwise. Here we report a tail-independent inactivation of kinesin, reversible by the disease-relevant signaling protein, casein kinase 2 (CK2). The majority of initially active kinesin (native or tail-less) loses its ability to interact with microtubules in vitro, and CK2 reverses this inactivation (~ 4-fold) without altering kinesin’s single motor properties. This activation pathway does not require motor phosphorylation, and is independent of head-tail auto-inhibition. In cultured mammalian cells, reducing CK2 expression, but not its kinase activity, decreases the force required to stall lipid droplet transport, consistent with a decreased number of active kinesin motors. Our results provide the first direct evidence of a protein kinase up-regulating kinesin-based transport, and suggest a novel pathway for regulating the activity of cargo-bound kinesin.
Introduction
Kinesin-1 (conventional kinesin, KIF5) is an important microtubule-based molecular motor that enables fast transport toward the cell periphery. Kinesin-based transport is essential for cell function, and its impairment is linked to diseases including neurodegeneration. Discovered over two decades ago, the single molecule function of kinesin in vitro has been studied extensively, and is relatively well understood. How kinesin function is regulated in vivo, however, remains an important open question. Kinesin can auto-inhibit via a direct head-tail interaction (reviewed in1), but this known inhibition is released upon cargo recruitment2, and little is known about regulation that occurs without altering the motor’s presence on cargo3–5.
We report here a tail-independent inactivation for kinesin-1, reversible by Casein Kinase 2 (CK2). CK2 is a highly conserved, ubiquitous serine/threonine kinase important for cell growth and proliferation. There is a partial overlap in disease associations between kinesin and CK2 (tumorgenesis6–7, and neurodegeneration8–10), but there has heretofore been no evidence of a direct functional regulatory interaction.
Results
CK2 increases the active fraction of native kinesin in vitro
To probe for a potential effect of CK2 on kinesin function in vitro, we first incubated native kinesin (purified from bovine brains, Fig 1a–b) with recombinant CK2 holoenzyme (New England Biolabs, Fig. 1c) in a buffer optimized for CK2 kinase activity, and then diluted the reaction (>10-fold) in buffer appropriate for biophysical or biochemical measurements. Parallel reactions from which CK2 had been omitted served as controls. To control for the possibility of contamination, we verified that the recombinant CK2 employed in the current study is >90% pure by Coomassie stained SDS-PAGE gel (Fig. 1c), and observed no detectable contamination by mass spectroscopy (Supplementary Table S1).
In vitro, CK2-pretreatment significantly increased the probability of a kinesin-coated bead binding to and processing along a microtubule at 1mM ATP, in a dosage-dependent manner (Fig. 1d). To examine the effect of CK2 on kinesin in depth, we carried out extensive studies at a molar incubation ratio of ~3 CK2 per kinesin, where effects were pronounced and approached saturation (Fig. 1d). Upon completion of the CK2 treatment, kinesin samples were diluted 10–1000 times in motility buffer, and incubated with carboxylated polystyrene beads to allow for non-specific decoration of motors onto beads. Motor density on the beads was controlled by varying the input kinesin concentration, while keeping bead concentration constant. Using a conventional single beam optical trap, we positioned individual kinesin-coated beads in close vicinity of a microtubule (surface-immobilized in our flow cells) to evaluate the motor-microtubule interaction. In this confined geometry, the ability of a bead to bind to and process along the microtubule directly reflects the presence of active motors on the bead (Supplementary Discussion). Further, the more active motors present, the greater the probability of bead binding11. Remarkably, in parallel assays differing only in the presence or absence of CK2, we observed a significant enhancement in bead binding fraction associated with CK2-treatment (Fig. 1d). Control experiments using beads incubated with CK2 but without kinesin showed no specific affinity to microtubules over the dilution range tested in Fig. 1d (zero bound of 50 beads tested). In the same confined geometry, CK2 treatment did not appreciably reduce the mean time between successive binding events for a bead with a single active kinesin, suggesting that the motor’s ‘on’ rate for the microtubule was minimally affected (‘Time Lapse’, Supplementary Figure S1a). We further observed no significant effect of CK2 on the single motor travel or velocity of the active kinesin (Supplementary Figure S1a), suggesting that the increase in bead binding fraction arose from an increase in the number of active motors on the bead. To reach the single molecule range where no more than one active motor is present on the bead (~30% binding11), an additional ~8-fold dilution was required for CK2-treated kinesin samples relative to the CK2-blank case (Fig. 1d). This enhancement cannot be accounted for by the minor impact of CK2 on kinesin-bead association (Supplementary Figure S1b, and Supplementary Methods), thus suggesting an increase in the fraction of active motors on the bead.
An increase in the active fraction of kinesins per bead is supported by two additional biophysical metrics: bead travel distance, and force production. Both experimental and theoretical studies12–13 demonstrate a positive correlation between cargo travel distance and motor number in vitro, and cargo force production conclusively measures the number of simultaneously engaged motors on a cargo both in vitro14–16 and in vivo4–5. If CK2 does increase the active fraction of kinesins without altering its single motor function, CK2-treatment should effectively increase the number of available motors per bead, and improve both bead travel and force production. Indeed, using the same motor:bead incubation ratio where there was a small contribution from a second motor for the CK2-blank control, we observed a significant enhancement in the average bead travel (>three fold, Fig. 1e) and force production (pronounced 2-motor contribution at ~9pN, Fig. 1f) associated with CK2 treatment. The unchanged one-motor peak position at ~4.5 pN in force distributions (Fig. 1f) indicates that CK2 treatment does not alter kinesin’s single motor force production.
To eliminate concerns of bead-related experimental artifacts, we carried out bead-independent microtubule affinity pull-downs17 to biochemically examine the effect of CK2 on kinesin activity. Kinesin samples (pretreated with or without CK2) were diluted 10-fold in microtubule binding buffer, and incubated with microtubules at large molar excess (>600-fold molar excess of tubulin-heterodimer to kinesin), followed by pelleting of the microtubules and associated motors. Here, AMPPNP (adenylyl imidodiphosphate, a non-hydrolysable ATP analogue) was used to increase the stability of kinesin-microtubule complexes (Supplementary Discussion). Consistent with the increased active motor fraction observed in the bead assay (Fig. 1d–f, Supplementary Figure S1a–b), CK2-pretreatment significantly increased the fraction of native kinesin co-sedimenting with microtubules (Fig. 1g). This increase is unlikely to be explained by any altered interactions between the active kinesin and the microtubule: at the physiologically relevant 1 mM ATP, single molecule biophysical measurements gave no indication of CK2 influencing either kinesin-microtubule ‘on’ rate or ‘off’ rate (as probed by time lapse between binding events, and single molecule travel distance, respectively, Supplementary Figure S1a). Combined, these data indicate that the effect of CK2 treatment is to increase the fraction of active kinesin molecules capable of binding to microtubules, rather than simply altering the association/dissociation rates of active kinesins with microtubules in the presence of AMPPNP.
Consistent with previous reports18–19, CK2 co-sediments with microtubules with high affinity under our pulldown conditions (Fig. 1g–h). We observed no significant evidence of the native kinesin altering the CK2/microtubule interaction (Fig. 1g).
The CK2 effect is independent of the kinesin tail
In principle, modulation of the known head-tail binding1 mechanism could alter the fraction of active kinesin, while leaving the single motor function intact. To conclusively test whether or not CK2 functioned by altering kinesin’s head-tail interaction, we used the functional kinesin construct (K560) lacking the tail and the light chain20–21 (Fig. 2a–b). Unless otherwise noted, a final microtubule affinity purification step22 was used to specifically select for active K560 motors (Fig. 2b). We observed no detectable contamination in the microtubule-purified K560 using Coomassie-stained gels (Fig. 2b) or via mass spectroscopy (Supplementary Table S1). The effect of CK2-pretreatment persisted for the tail-less K560, as assayed by both the bead-independent microtubule pulldowns (Fig. 2c–d), and the bead assays (specific or non-specific recruitment, Fig. 2e–f, Supplementary Figure S1c–d). Again, the single motor functions of the active K560 (Supplementary Figure. S1c) and K560/bead association (Supplementary Figure S1d) remained unaltered, indicating that the main effect of CK2 is to increase the active motor fraction for a given K560 population.
The CK2 effect does not require motor phosphorylation
Since CK2 is a kinase, we hypothesized that motor phosphorylation was responsible for CK2’s effect on kinesin activity. Kinesin is an in vitro substrate for CK223–24, and our CK2 treatment of kinesin resulted in phosphate incorporation into both the native (Fig. 3a) and the tail-less motors (Fig. 3b–e. Focusing on tail-independent activation, via mass spectroscopy we identified one significantly phosphorylated residue in K560, a serine at 520 (Fig. 3d, Supplementary Table S2, and Supplementary Methods). This serine is located in a sequence (YELLSDELN) that is a reasonable match to the CK2 phosphorylation consensus, in which acidic residues at the +1, +2 and +3 residues are important25. Using site-directed mutagenesis, we substituted serine 520 with alanine, and generated a functional mutant that was minimally phosphorylated by CK2 (Fig. 3e), suggesting serine 520 residue is indeed the main CK2 phosphorylation site in K560. Nonetheless, CK2-pretreatment still significantly enhanced the binding of this mutant kinesin to microtubules (Fig. 3f), indicating that phosphorylation of serine 520 is not the mechanism of CK2-mediated kinesin activation.
To test the possibility that additional, less prominent phosphorylation sites might be important for the observed CK2 effect, we introduced the CK2 specific inhibitor, Tetrabromocinnamic Acid26 (TBCA), to our in vitro CK2 treatment. TBCA severely limited the catalytic activity of CK2 (both motor phosphorylation and CK2 autophosphorylation, Fig. 3c), but did not impact CK2’s effects on K560 activity (Fig. 3f). The activation effect of CK2 on kinesin also persisted when the kinase activity of CK2 was limited by reducing the kinase/motor incubation temperature to 0oC (Fig. 2d, 3b), again indicating that motor phosphorylation is not required for CK2 to activate K560. A non-catalytic mechanism of CK2 action is consistent with the observation that the CK2 effect is dosage-dependent and approaches saturation at an incubation ratio of ~3 kinase per motor (Fig. 1d).
Co-sedimentation of CK2 with microtubules persisted after incubation with the mutant motor S520A, and was independent of CK2’s kinase activity (‘S520A’, Fig. 3f).
The CK2 effect is specific
Independent of motor phosphorylation, ice incubation of kinesin with CK2 introduced a ~4-fold increase in the fraction of microtubule-bound motors, for both the truncated and the native kinesins (‘Pre-Treatment’, Fig. 4a–b). Consistent with bead assay findings (Fig. 1b), the activation effect of CK2 was dosage dependent, with the active fraction of the native kinesin reaching ~80% of the overall motor population at 3:1 kinase:motor incubation ratio (‘3CK2’, Fig. 4a–b). Control experiments using an equal molar amount of bovine serum albumin (BSA) in place of CK2 demonstrated negligible effect for both the native and the truncated kinesins (‘BSA’, Fig. 4a–b). Two more independent experiments support the specificity of CK2 effect for K560. First, we verified that a different serine/threonine kinase (glycogen synthase kinase 3β, GSK3β) had no significant impact on K560 activity (3:1 GSK3β:K560, Fig. 4b). Second, we identified differential effects of the two catalytic subunits of CK2, CK2α vs. CK2α′ (Fig. 4b). Despite sharing 75% sequence identity27, the two subunits exhibited distinct effects on kinesin activity. At the same 3:1 kinase:motor incubation ratio, CK2α significantly increased the fraction of kinesin in microtubule pellets, but CK2α′ did not (Fig. 4b). We observed no evidence of contaminants in either subunits via Coomassie stained SDS-PAGE gels (Fig. 4c). This functional specialization of CK2α vs. CK2α′ again demonstrates the specificity of CK2 effect identified in the current study.
CK2 re-activates inactive kinesin
One trivial explanation for the observed CK2 effect would be that the kinase somehow prevented an irreversible inactivation of purified kinesin. As a direct test, we evaluated the effect of CK2 on already inactive kinesin (‘Re-Activation’, Fig. 4). In contrast to treating kinesin with/without CK2 in parallel prior to parallel microtubule pulldowns (‘Pre-Treatment’, Fig. 4), in ‘Re-Activation’ we specifically assayed for the active fraction of kinesin before and after incubation with CK2. Here, the microtubule pulldowns were staggered in time. To control for the possibility that CK2 functioned by reversing motor precipitation or adsorption onto plasticware, we transferred kinesin samples to new plasticware prior to CK2 incubation (so that any adsorbed motors would be left behind), and then carried out the microtubule pulldowns. Starting with samples that contained a majority of inactive motors, subsequent incubation with CK2 reversed the inactivation and resulted in a population with the majority of motors active, for both native and truncated wild-type kinesins (‘Re-Activation’, Fig. 4a–b), and the phospho-mutant S520A (Fig. 4d). These data demonstrate that tail-independent kinesin inactivation is reversible, and that the main effect of CK2 is to reverse this inactivation without requiring motor phosphorylation by the kinase. Using quantum dot (qDot) assays, we verified that CK2 could significantly increase kinesin activity after motor-qDot recruitment (Fig. 5), supporting the notion that the tail-independent activation of kinesin by CK2 may in fact be feasible for cargo-bound motors.
In vivo force depends on CK2 level but not kinase activity
Might these in vitro effects be relevant in vivo? To test this, we examined the effect of a decrease in cellular CK2α levels on cargo force production, using a lipid droplet transport system in cultured mammalian cells (Fig. 6). CK2α is present on lipid droplets28, kinesin-1 is the predominant plus-end motor moving such droplets4, and the overall force driving in vivo cargos is an effective readout for the average number of engaged motors4–5. Using siRNA transfections to specifically reduce CK2α expression in Cos-1 cells down to 12% of wild-type levels (Fig. 6a–d, scrambled siRNA control introduced no significant effect on CK2α level), we measured a four-fold decrease in the percentage of droplets capable of escaping a fixed optical trap (‘siRNA1’, Fig. 6e). Using a second siRNA to target a different region of CK2α (Fig. 6c–d), we observed similar reduction in cargo force production (‘siRNA2’, Fig. 6e). Consistent with our finding that motor phosphorylation was not required for CK2-mediated stimulation in vitro, pharmacological inhibition of CK2 kinase activity(Fig. 6f) had no effect on droplet force production (‘TBCA’, Fig. 6e). Significant variations in droplet size could affect force measurements, but neither knockdown of CK2α expression nor inhibition of CK2 activity had any significant effect on lipid droplet size (Fig. 6g).
Tail-independent inactivation occurs for monomeric kinesin
What is the mechanism responsible for the tail-independent kinesin inactivation? One possibility is that kinesin might form oligomers and thereby become inactive. To address this, we utilized size-exclusion chromatography to investigate potential changes in kinesin sizing profile before and after inactivation (Supplementary Methods). The K560 peak location did not change (Fig 7a), but the associated motor activity was significantly reduced (~5-fold, assayed by bead binding fraction, Fig 7b). This result suggests that protein aggregation is not important for the reversible kinesin inactivation observed in the current study. A second possibility is that the ‘inactive’ kinesin state might occur when the motor binds to ADP, given that the equilibrium binding of kinesin to microtubules has been reported to be weak for ADP-bound motors29–30 (Kd ~10–20 μM). Using dialysis (80 mM Pipes pH 7.0, 5 mM ETDA, Supplementary Methods) to deplete potential bound nucleotides in Ni-purified K560, we again observed significant loss of motor activity for the dialyzed K560 after ice incubation (‘Dialyzed’, ‘Before’ vs. ‘After’, Fig. 7c). While the completeness of dialysis-mediated nucleotide removal was difficult to assess, the fraction of K560 capable of binding to microtubules remained the same as for the un-dialyzed samples (‘Undialyzed, After’, Fig. 7c) and for the dialyzed samples where ADP was introduced back in (‘Dialyzed+ADP, After’, Fig. 7c) that underwent identical ice incubation. Thus, kinesin-inactivation is unlikely to simply reflect ADP binding. A third possibility is that there might be an inhibitory head-head interaction, which could block the microtubule binding of the kinesin dimers. To test this, we generated a truncated kinesin (K339) that exists as a functional monomer31. For this K339 monomer, motor inactivation still persisted and was again rescued by CK2 (Fig 7d). Therefore the tail-independent kinesin inactivation identified in the current study cannot require head-head interaction. Based on these findings, we hypothesize that the tail-independent kinesin inactivation may involve a conformational change within individual kinesin heads, at least somewhat independent of their nucleotide-bound states.
CK2 effect is mediated by weak K560/kinase interaction
Thus far, our data indicate that the neither the kinesin tail (Fig. 2–5) nor phosphorylation (Fig. 3, 4d) is required for CK2-mediated kinesin activation. Using immunoprecipitation assays (Supplementary Methods) against kineisn’s C-term His tag (Fig. 8a) or heavy chain (Fig. 8b), we observed complex formation/co-immunoprecipitation between CK2 and K560, independent of CK2’s kinase activity. Intriguingly, when tested separately, of the two catalytic subunits, only CK2α′ co-immunoprecipitated with K560 (Fig. 8b). Nonetheless, CK2α--but not CK2α′-- demonstrated similar potency for kinesin activation as the CK2 holoenzyme (immunoblot for one of two trials shown in Fig. 8c, quantified in Fig. 4b). We conclude from these data that the functional interaction between CK2 and the tail-less kinesin is specific but weak. Immunofluorescence staining of untreated wild-type Cos-1 cells demonstrated some co-localization between CK2α and kinesin heavy chain (Fig 8d–e, Supplementary Methods), and provides support for the likelihood of a weak interaction in vivo. We note that there appeared to be cell to cell variation in the amount of co-localization of CK2 and kinesin, as judged either by eye or the cross-correlation analysis. Thus, we suspect that the CK2-kinesin interaction may be regulated, and may be an interesting topic for future studies.
Under our microtubule pulldown conditions, CK2 exhibited strong interaction with microtubules (Fig. 1h). In principal, CK2 might function simply by acting as a microtubule associated protein that recruited kinesin to the microtubule, and an initial kinesin-CK2 interaction might not be relevant. To test this possibility, we prepared identical CK2/K560 mixtures (3:1 kinase:motor), and evaluated the importance of a 1.5hr incubation of CK2 with kinesin prior to introducing microtubules (Fig. 8f). Here, the same amount of CK2, K560, and microtubules were used, and microtubule pulldown assay conditions were identical (that is, the mixture of CK2 and K560 was in the presence of the microtubules for the same amount of time). We observed a significant increase in K560 activity only for samples that had undergone extended incubation with the kinase (Fig. 8f) before being exposed to microtubules. However, CK2 binding to microtubules was not affected by the motor/kinase incubation, and there was no increase in the amount of CK2 on the MTs correlating with the ~3.8-fold increase in kinesin activity (Fig. 8f). These data indicate that an initial functional interaction between CK2 and K560 is necessary, and a model where CK2 first binds to MTs, and then recruits kinesin, is inconsistent with our data. However, we cannot rule out other contributions of the microtubule to this process (see below).
Although CK2 and kinesin co-immunoprecipitate in vitro, the presence of kinesin did not enhance the fraction of CK2 binding to microtubules (Fig. 1g). We hypothesize that CK2 may have a stronger binding to the microtubule than to the K560. This possibility might provide a mechanism for dissociating the CK2/kinesin complex after functional interaction.
Activation effect of CK2 is enhanced for ADP-bound kinesins
The hypothesized CK2-mediated alteration in kinesin conformation (within individual motor heads), coupled with the weak interaction between kinesin and CK2 (Fig. 8g), suggests that an additional change occurs to drive the motor activation. Although kinesin’s inactivation itself may not depend on ADP presence, part of the CK2-mediated activation process could involve a change in kinesin’s nucleotide-bound state. To test this, we returned to the dialyzed K560 motors that approximates the motor’s nucleotide-free state (‘Dialyzed’, quantified in Fig. 8g, immunoblot for one dialysis trial shown in Fig. 7c), and tested whether CK2’s ability to activate kinesin was affected when we introduced ADP back into the dialyzed sample (‘Dialyzed+ADP’, quantified in Fig. 8g, immunoblot for one dialysis trial shown in Fig. 7c). Intriguingly, CK2 was able to activate kinesin more effectively when ADP was present (Fig. 7c, 8g). Additionally, when ADP was present in the dialyzed sample, the effect of CK2 approached that for the un-dialyzed samples (54± 4% vs. 67 ±10% respectively, mean ±SEM, n=3 trials each, immunoblots for one dialysis trial shown in Fig. 7c), and we observed no significant impact of additional ADP on CK2 effect in un-dialyzed kinesin samples (data not shown). The exact sensitivity of the CK2 effect to ADP may in fact be larger, depending on the completeness of nucleotide depletion in dialysis. Nonetheless, the observed difference is statistically significant (p = 0.02, t-test, Fig. 8g), tested for three independent dialysis trials (immunoblot for one of three trials shown in Fig. 7c), and is consistent with the hypothesis that ADP binding/release may contribute to the activation process. Additional possibilities include conformation changes in CK2 as a result of motor binding, or competition between microtubules and K560 for CK2 binding. Complete mechanistic details remain to be investigated in future studies.
Discussion
Taken together, our data uncovers a tail-independent inactivation for kinesin-1, reversible by CK2. Although other members of the kinesin family may have modes of regulation in addition to the tail-mediated auto-inhibition32, this is the first evidence to our knowledge of a second regulatory mode for kinesin-1. When tested with freshly purified K560 that was specifically selected for motor activity (via microtubule affinity purification step22, Fig. 2b), the time scale of this tail-independent inactivation and re-activation (~4 hr and 1.5 hr, respectively, Supplementary Discussion) was significantly faster than the turn-over rate of the native kinesin in vivo (half-life of 18–20 hrs33). Inactivation/activation could occur more rapidly; our temporal resolution is limited by assay conditions including motor-kinase incubation.
The finding that kinase activity of CK2 was not required for kinesin activation was unexpected. However, there are precedents: kinase-independent regulation has been reported for CK2 affecting microtubules19 and other kinases/targets34–36. It remains possible that phosphorylation by CK2 might serve as an intermediate state for additional kinases37 to enable further spatial and/or temporal regulation of the kinesin motor.
The tail-independent activation of kinesin identified here suggests a novel pathway for cell signalling to reversibly regulate kinesin activity while the motors remain cargo-associated. A recent report demonstrates that the number of engaged motors is actively regulated during axonal transport, even though the overall motor presence on cargo remains constant3. Other regulatory pathways could also take advantage of this activation pathway, either by blocking access of CK2 to the kinesin (to promote inactivation) or alternatively, by directly interacting with kinesin and mimicking the CK2 effect.
Since kinesin activation is tuned by CK2 levels (Fig. 1d, 4a–b), changes in cellular CK2 concentration likely lead to corresponding changes in kinesin activity. This is consistent with live cell force measurements (Fig. 6), and suggests the possibility of a link between CK2 enrichment38 and the preferential recruitment of kinesin to microtubules in the initial segment of axons39. Our study also suggests an intriguing role for altered CK2 signaling7–8,10 in disease progression: by influencing transport processes crucial for cell function. The extent to which this occurs for both neurodegenerative diseases (where CK2 is often decreased and transport likely impaired) and cancers (where CK2 is increased and transport is likely unregulated to support metastasis) remains an exciting area for future research.
Methods
Proteins
Native kinesin and tubulin were purified from bovine brain15,40. Functional tail-less K56020–21, monomer K33931, and phospho-mutant S520A were constructed using plasmid pET17b_k56020 (Addgene) as template. Protein was bacterially expressed and Ni-NTA purified21. To specifically select for motor activity, unless otherwise noted Ni-eluted K560 was further purified by microtubule affinity purification22 prior to flash freezing in 10% sucrose and storage at −80°C.
Human recombinant CK2 holoenzyme (CK2αα′, un-tagged), CK2α & GSK3β (both C-term His-tagged), and CK2α′ (N-term His-tagged) were purchased from New England Biolabs, Invitrogen, and Millipore, respectively. The same activation effects for kinesin were observed for three different lots of CK2 holoenzyme tested (NEB, Lot 0130802, 0140810, 14/021309).
Immunoblot
Primary antibodies used were SUK4 and tubulin-α (Developmental Studies Hybridoma Bank, University of Iowa), AKIN01 (Cytoskeleton), CK2α (Cell Signaling), and CK2α′ (Novus Biologicals). For immunoblots, primary antibody concentrations used were: 1:500 for SUK4; 1:300–500 for tubulin-α; and 1:1000 for AKIN01, CK2α, and CK2α′. Immunoblots were quantified using Odyssey (Li-Cor Biosciences).
In vitro CK2 pre-treatment
75 nM purified kinesin (native or truncated) was incubated with recombinant CK2 (holoenzyme or subunit as indicated, up to 250nM, New England Biolabs) in 20μL reaction buffer (20mM Tris-HCl pH7.5, 50mM KCl, 10mM MgCl2, 0.5mM EGTA, plus 500μMATP) at 30°C for 40 min (optimal for CK2 kinase activity), or 0°C for 1.5 hr. Reactions from which CK2 had been omitted served as controls. To block CK2 kinase activity, 100μM CK2-specific inhibitor TBCA26 (EMD Chemicals, Inc.) was introduced to above assays, with ethanol as a control. For functional characterizations (bead and qDot assay, microtubule pulldown), samples were placed on ice after allotted incubation time without further quenching of CK2 kinase activity.
To assay for phosphate incorporation, motor/kinase reaction (0.5μg kinesins per 250U CK2, 30°C for 40 min as suggested by manufacture) were supplemented with 1μCi of [γ-32P]-ATP (Perkin Elmer), quenched with reducing SDS sample buffer, separated by SDS-PAGE, and visualized by PhosphorImager (Amersham).
In vitro CK2 re-activation
CK2-blank kinesins (thawed, 150nM native or truncated dimer, or 300nM monomer) were first incubated on ice for 1.5 hours. This ice incubation does not impact the active fraction of native kinesin (~20% immediately after thawing), but significantly reduces the active fraction of truncated kinesins (dimer or monomer, down to ~20%). After ice incubation, 10μL each of the kinesin samples are identically transferred to two sets of new eppendorf tubes. For one set, 10μL CK2-blank reaction buffer was added, followed immediately by a microtubule pulldown assay to determine the active motor fraction. For the second set containing identical kinesin samples, 10μL CK2 (500nM) was added, followed by 1.5 hr kinase:motor incubation on ice, then assayed for active motor fraction via microtubule pulldown.
Microtubule affinity pull-down
Kinesin (20μL of 75nM native or truncated dimer or 150nM monomer, with indicated amounts of CK2 holoenzyme or subunit, and indicated incubation conditions) were incubated with microtubule binding solution (180μL containing 5μM microtubule, 80mM Pipes pH6.9, 50mM CH3CO2K, 4mM MgSO4, 1mM DTT, 1mM EGTA, 20μM taxol), supplemented with 4mM AMPPNP (EMD), for 15 minutes at room temperature (RT). The reaction was centrifuged in a TLS55 rotor for 10 min at 170,000g at 25°C. The resulting microtubule pellet was dissolved in 30μL 1x reducing SDS sample buffer, separated by SDS-PAGE, and analyzed by immunoblot. Addition of 1mg/ml casein in the microtubule binding solution did not alter CK2-mediated kinesin activation.
In vitro optical trap
Kinesin was first incubated with CK2 (75nM native or K560, indicated CK2 amount and conditions), then titrated and incubated with 0.6pM carboxylated polystyrene beads (0.5μm, Polysciences) in 100μL motility buffer (80mM Pipes pH6.9, 50mM CH3CO2K, 4mM MgSO4, 1mM DTT, 1mM EGTA, 10μM taxol, 1mg/ml casein) for 15min at RT for non-specific recruitment. Here, bead concentration was chosen to optimize the number of beads in our field of view, and was kept constant while we varied kinesin concentration (up to ~2000-fold dilution starting from 75nM) to vary motor:bead incubation ratio, and thereby control motor presence on individual beads. In case of K560, we also used anti-his beads41 (0.4μm) to specifically recruit the motor via its C-terminal his-tag. Anti-his beads41 were prepared by incubating 20μL penta-His biotin conjugate (Qiagen) with 10μL streptavidin-coated polystyrene beads (1% solids, Spherotech) in 100μL 1xPEM80 buffer for 30min at 4°C, and washed five times in 1xPEM80 buffer containing 8mg/ml BSA or 5.55mg/ml casein before use. K560 concentration was titrated after indicated treatment (starting at 75nM) and incubated with 1μL anti-his beads in total 15μL motility buffer for 15min at RT. After incubation, motor-bead mixture were flown into flowcells with preassembled microtubules15, and all measurements were carried out at RT in motility buffer, supplemented with 1mM ATP, and an oxygen-scavenging system (250μg/ml glucose oxidase, 30μg/ml catalase, 4.5mg/ml glucose). Bead-microtubule binding was tested using an optical trap (Ktrap ~ 0.005pN/nm) to position each bead near a microtubule. A binding event is scored if the bead processes away from the trap within a wait time of 30 sec. Extending wait times to 2minutes had no effect on binding fraction measured (see Supplement Discussion). Bead force production was monitored by the bead positions in the trap (Ktrap = 0.0436 ± 0.001pN/nm). Bead motility was measured with trap turned off upon a binding event. Where appropriate, travel distributions were fitted to a single exponential decay to extract the characteristic run length and associated uncertainties. For measurements where CK2-treatment significantly increased the population exceeding our 8μm field view, algorithmic mean (and associated error) was used to determine the minimum average travel and associated error. Stall force distributions were fitted to Gaussians to extract mean one- and two-motor forces, and the associated errors. Error on binding fractions (p) was determined as , for n beads tested.
In vitro quantum dot
Quantum dots (QDots) (qDot-655 Carboxyl, Invitrogen) were excited with 488nm laser (Ti:Sapphire, Coherent) and imaged via a custom Total Internal Reflection Fluorescence (TIRF) microscope (Nikon 1.49NA, 100x), recorded at 4.25fps (Photmetrics Quantem 512SC). Motility assays utilized the same motility buffer and flow chamber (with preassembled microtubules) as for bead assays, and analysis was carried out using a custom-tracking program (Gross Lab) that identifies qDot positions via two-dimensional Gaussian fitting of their brightness profile.
For CK2 pretreatment, K560 was first incubated with CK2 (75nM K560, 0 or 250nM CK2, 1hr on ice), then diluted (25-fold) to incubate with qDots at 4.3:1 qDot:motor ratio in 6μL motility buffer for 12min at RT. The reaction was then brought up to 20μL in motility buffer, supplemented with 1mM ATP and oxygen-scavenging system, followed by motility experiments.
For CK2 effect on motors already in complex with qDot, 75nM K560 was first incubated with qDots at 4.3:1 qDot:motor ratio in 6μL moltility buffer for 30 minutes at 4°C. The kinesin/qDot mixture was then divided into equal 3μL portions and incubated with 3μL each of 0 or 250nM CK2 for 1 hr at 4°C. The reaction was then diluted (35-fold) in motility buffer, and 20μL each was supplemented with 1mM ATP and oxygen-scavenging system, followed by motility experiments.
Cell culture and CK2α knockdown
Cos-1 cells were grown in DMEM (Invitrogen) supplemented with 10% fetal bovine serum (FBS) at 37°C in 5% CO2. Gene silencing was achieved by transfection with two different commercially available CK2α siRNAs, from Qiagen (with sense strand 5′-CAUUGAAGCUGAAAUGGUATT-3′) and Santa Cruz Biotechnology, Inc (a pool of 3 different siRNA duplexes with sense strands 5′-GAAGCCAUCAACAUCACAATT-3′, 5′-GAUCCACGUUUCAAUGAUATT-3′ and 5′-CCUCAGUCUUGUAAAUGUATT-3′), using the Hiperfect transfection reagent and according to manufacturer’s instructions. Non-silencing siRNA and non-transfected cells were included as controls. Final dosing concentrations of all siRNAs provided were 50nM, unless otherwise noted, and the cells were incubated with the siRNA transfection complexes for 24 hours in complete media at normal growth conditions. After 24 hr in complete media at normal growth conditions (additional 50μM TBCA in growth media for TBCA-treatment), the cells were treated with 60μg/ml of fatty acid-albumin complex to induce the lipid droplet formation. The cells were then exposed to DMEM without glucose and fatty acid for 16 hr, before immediate lipid droplet trapping experiments. For cell lysates, scrapped off cells were washed with PBS and lysed in ice cold lysis buffer (25mM TrisHCl pH7.5, 150mM NaCl, 1% NP40, 1mM EDTA, 1mM PMSF, 1mM Na3VO4, and 1x protease inhibitor cocktail).
Live cell escape force measurements
Lipid droplets in Cos-1 cells were visualized by differential interference contrast microscopy, the force driving individual lipid droplets was determined by monitoring the droplet motion inside an optical trap4–5. An escape event was scored if the droplet moved out of the optical trap within the 30 sec wait period. A no-escape event was scored if the trapped droplet could not escape the trap, but resumed motion once the trap was turned off. We empirically tuned the trap power (980nm, 24mW output) such that the majority (~80–90%) of lipid droplets in wild-type cells could escape the trap, and measured escape events for all cells under the same trap power for direct comparison. Droplet sizes were manually quantified in ImageJ. Errors in escaped fraction (f) were determined as , for n droplets tested.
Supplementary Material
Acknowledgments
We thank Thomas Whisenant, Robyn Kaake, Lei Fang, Elizabeth McReynolds, and Chi-In Kris Ngai for helpful discussions and technical help. This work is supported by NIGMS grants GM64624 to SPG, GM74830 to LH, GM76516 to LB, NS048501 to SJK, and AHA grant 825278F to JX.
Footnotes
Author Contributions
JX designed, performed, and analyzed most of the experiments and data. BJNR, PA, and ZS contributed equally to the paper: BJNR and PA performed live cell studies. BJNR and SKV performed quantum dot assays. ZS, MKM, MTH, NJ, and SJK purified proteins. SC performed mass spectroscopy analysis. JX, MKM, and MTH carried out tests for inactivation/activation mechanisms. SPG designed and analyzed experiments. JX, LB, and SPG wrote the paper.
Competing Financial Interests
The authors declare no competing financial interests.
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