© 2018. Published by The Company of Biologists Ltd.

Microtubule Dynamics Regulation Reconstituted in Budding Yeast

Lysates

Zane J. Bergman1, Jonathan Wong1, David G. Drubin, and Georjana Barnes

Department of Molecular and Cell Biology, University of California, Berkeley, CA 94720

Corresponding author’s email: gbarnes@berkeley.edu

1. These authors contributed equally to this work.

Keywords: microtubule, reconstitution, dynamic instability, kinesin

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JCS Advance Online Article. Posted on 5 September 2018

SUMMARY STATEMENT

We developed an in vitro assay for measuring the growth and dynamics of single

microtubules in total budding yeast cellular protein complexity.

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ABSTRACT

Microtubules (MTs) are important for cellular structure, transport of cargoes, and

segregation of chromosomes and organelles during mitosis. The stochastic growth and

shrinkage of MTs, known as dynamic instability, is necessary for these

functions. Previous studies to determine how individual MT-associated proteins (MAPs)

affect MT dynamics have been performed either through in vivo studies, which provide

limited opportunity for observation of individual MTs or manipulation of conditions, or in

vitro studies, which either focus on purified proteins, and therefore lack cellular

complexity, or on cell extracts made from genetically intractable organisms. In order to

investigate the ensemble activities of all MAPs on MT dynamics using lysates made

from a genetically tractable organism, we developed a cell-free assay for budding yeast

lysates using TIRF microscopy. Lysates were prepared from GFP-tubulin-expressing

yeast strains and MT polymerization from pre-assembled MT seeds adhered to a

coverslip was observed in real time. Through use of cell division cycle (cdc) and MT

depolymerase mutants, we found that MT polymerization and dynamic instability are

dependent upon the cell cycle state and the activities of specific MAPs.

INTRODUCTION

Microtubules (MTs) are polar cytoskeletal tracks that have many crucial functions in

cells that include trafficking of cargoes, maintenance of cell shape, and partitioning of

genetic material to daughter cells during mitosis and meiosis (Hirokawa and Tanaka,

2015). These diverse functions are achieved through the plasticity of MT structural,

biochemical, and dynamic properties that can vary between cell types, between cell

cycle stages, or even between MTs at a given time within a single cell. Mechanisms

that control MT dynamics include the stochastic growing and shrinking of ends intrinsic

to the polymer, known as dynamic instability, the combined forces of motor proteins

acting on the microtubules, and the activities of microtubule-associated proteins (MAPs)

that act as polymerases, depolymerases, stabilizers, and destabilizers (Bowne-

Anderson et al., 2015).

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The study of MT dynamics and the effects of their MAPs have historically come from in

vivo or in vitro studies. In vivo studies on dynamics are limited to either bulk analysis

through such approaches as fluorescence recovery after photobleaching (FRAP)

(Salmon et al., 1984) and speckle analysis (Grego et al., 2001; Waterman-Storer et al.,

1998) or analysis of single MTs that can be resolved by light microscopy, usually at the

cell periphery (Shaw et al., 1997). Dynamics at MT plus-ends at the cell periphery in

interphase cells or astral MTs in dividing cells can be measured, but this excludes the

plus-ends of most interphase and all kinetochore and interpolar MTs. The other source

of dynamics studies is from reconstituted in vitro systems that have produced stunning

insights into such processes as spindle assembly (Sawin and Mitchison, 1991) and plus

end regulation (Li et al., 2012; Moriwaki and Goshima, 2016). Another productive

avenue toward studies of MT dynamics is genetics, which has been particularly valuable

for discovery of key microtubule dynamics regulators (Pasqualone and Huffaker, 1994;

Wang and Huffaker, 1997), which tend to be of low abundance and therefore difficult to

identify by biochemical means. While in vivo, in vitro, and genetic approaches have

been extremely productive, being able to combine two or more of these approaches,

such as genetics and in vitro reconstitution, holds great promise to achieve a level of

analysis greater than could be achieved with any single approach alone. Development

of a budding yeast lysate system for MT dynamics studies would allow genetics to be

combined with the full complexity of total cellular protein in an open system reflecting

the activities of the full panoply of yeast proteins expressed in yeast by comparing

activities of extracts prepared from mutants of MAP and cell cycle control genes.

We developed an in vitro assay that reconstitutes MT dynamics within the high

complexity of the total soluble protein content of a cell. This approach overcomes some

of the inherent limitations of previous studies and can elucidate and examine the

emergent properties of multiple MAPs acting on MTs. Our assay utilizes cleared lysate

prepared from yeast strains and observes the polymerization and plus-end dynamics of

MTs in vitro. Budding yeast is an ideal organism for MT dynamics in vitro reconstitution

because its MT network is: relatively simple but well studied, there is a fairly complete

components list, and mutants of every key structural and regulatory protein are readily

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available (Moore et al., 2009; Winey and Bloom, 2012). Budding yeast includes

homologs of many metazoan proteins and results are relevant to these organisms. Our

system utilizes pre-formed MT seeds, allowing specific analysis of plus end MT

dynamics in readily resolved single MTs in the absence of rate-limiting nucleation

reactions.

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RESULTS

Creation of Lysates and Imaging Chambers

Our goal was to create a protein system that enabled us to visualize single MT

dynamics within the complexity of the cellular milieu. To that end, we combined aspects

of classical in vivo genetics with the expediency, control, and accessibility of an in vitro

assay. We achieved this goal by preparing cleared lysates from budding yeast strains

that natively expressed GFP-tubulin (Straight et al., 1997). We wanted to examine MT

dynamics in lysates from actively growing cells, so strains were grown to late log phase

before being harvested by centrifugation. In order to create a concentrated product,

during harvest, cells were washed with water to remove remaining medium. Any

remaining liquid on top of the cell pellet was aspirated. The concentrated cell pellet was

apportioned and flash-frozen with liquid nitrogen and then crushed using a cryogenic

impact mill, avoiding addition of any buffer or other liquid. The resulting lysate was

highly concentrated (81.5 ± 8.5 mg mL-1 protein concentration).

Preparation of microscope slides, cover glasses, and cellular lysates are discussed in

detail in Materials and Methods. Briefly, a passivated coverslip was affixed to a

microscope slide with strips of double-sided tape to create a flow chamber (Bieling et

al., 2010). Rhodamine-labelled, GMPCPP-stabilized porcine MT seeds were adhered

to the coverslip using a biotin-streptavidin system. Whole-cell lysates from strains

natively expressing GFP-tubulin were buffered with 10X PEM, cleared of insoluble

material by ultracentrifugation, and supplemented with ATP and GTP. This mixture was

then flowed into the chamber. The microtubules were imaged by total internal reflection

fluorescence (TIRF) microscopy at 28°C in an environmental chamber. The samples

were imaged using 561 nm and 488 nm laser illumination every 5 seconds for 10

minutes. In lysates capable of polymerizing MTs, we found that by the time we could

find an appropriate area to image, the GFP-tubulin had already begun to assemble off

the rhodamine-labeled seeds. The activity of the lysates remained stable for up to 45

minutes of observation.

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Initial experiments using lysate of wild-type strains growing asynchronously did not

regularly exhibit any growing MTs off of the rhodamine-labeled seeds. This was a

puzzling result, as MTs are present and dynamic in all cells, but not one without

precedent. Previous work with actin showed that in vitro assembly of filaments was

highly dependent on the cell cycle stage of the harvested cells (Miao et al.,

2013). Microtubule dynamics across phyla are also known to fluctuate throughout the

cell cycle to accommodate the variety of distinct functions they serve at each stage of

the cell cycle (Rusan et al., 2001). To test this possibility, we sought to arrest the

cultures at different points of the cell cycle before harvesting. Strains with temperature-

sensitive alleles of cell-division cycle genes were used to arrest cells at different cell

cycle stages. When utilizing these strains, incubation at the restrictive temperature for 3

hours yielded >95% of cells arrested at either G1 (cdc28-4), S phase (cdc7-1),

metaphase (cdc23-1), or in late anaphase (cdc15-2) (Goranov et al., 2009). Upon

conducting our assay with these lysates, we observed MTs growing from the seeds for

all cases except when the cells had been arrested in G1 (Fig. 1A). From here, we

analyzed MT dynamics by generating kymographs and measuring rates of growth and

shrinking and the overall dynamicity (dimers sec-1) for individual MTs.

Lysate Optimization for Reproducibility

To obtain interpretable results using this assay it was essential to have a high degree of

reproducibility, especially when comparing dynamics properties across a large number

of genetic backgrounds. It quickly became apparent that a number of factors could

affect the dynamics of samples made independently from the same strain. These

factors included added nucleotide, protein concentration, and lysing conditions during

milling.

Initially, only excess GTP was added to the lysate to maintain the GTPase activity

necessary for MT polymerization (Carlier and Pantaloni, 1981). However, since many

MT-associated proteins (MAPs) and the MT motors dynein and kinesin, particularly

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kinesin-8 and kinesin-14 family members, have direct effects on MT dynamics and

require ATP for their activity (Gupta et al., 2006; Maddox et al., 2003; Sproul et al.,

2005; Varga et al., 2006), we investigated the need for exogenous ATP. We observed

MT dynamics of three independent preparations of lysates from S phase-arrested cells

in the presence or absence of additional 0.9 mM ATP (Fig. 1B). In one of three lysates,

without ATP added, MTs underwent constant growth without catastrophe whereas the

other two had cycles of catastrophe and rescue. After exogenous ATP was added, all

three preparations induced cycles of polymerization and depolymerization. Variable

levels of endogenous ATP in the lysate clearly change the dynamics of MTs. Therefore,

excess ATP and GTP were added for all experiments.

We hypothesized that the efficiency of milling could affect the protein concentration in

lysates, which in turn might affect the dynamics of MTs assembled in the assay.

Variability in milling can arise at several steps. The SPEX 6875 Cryogenic Impact Mill

used for these studies allows for 3 different sized vials and corresponding impactors

and has several settings for the duration and intensity of milling. Even though we had

standardized the duration and intensity of our milling protocol (Materials and Methods),

different sized impactors had been used for different sized cell harvests based upon the

manufacturer’s instructions. To test the effect of these variables on reproducibility, we

created several independent lysates from the same cdc7-1 parent strain using different

culture volumes. These harvests were then milled in the recommended vial and

impactor set for the corresponding harvested mass according to the SPEX manual. The

amount of variability in MT activity between these samples was surprising. Samples

ranged from having no detectable activity to assembling MTs that only grew and

paused, with still other samples that had MTs that underwent growth, pausing, and

shrinking (Fig. 1C). By quantifying protein levels by Bradford assay and Western blotting

(Fig. 1D), we found that milling cells from a 2 L culture with the small milling set was

less efficient in recovering α-tubulin than using the medium milling set on a 4 L culture.

A correlation was discovered between protein concentration and these distinct

phenotypes (Fig. 1E).

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We wanted to determine if nuclear protein levels might also be sensitive to changes in

culture and milling volumes and therefore contribute to variations in the protein

concentration of lysates. In budding yeast, the nuclear membrane stays intact

throughout the cell cycle, creating separate compartments for microtubules with distinct

subpopulations of MAPs in the cytoplasm versus the nucleus. To assess levels of

nuclear proteins present in the cleared lysate, we compared protein extracts from cdc7-

1 samples created using the two different sized vials and impactors and two different

culture volumes. We then assayed for the presence of the cohesin complex subunit

Mcd1 (Guacci et al., 1997), which localizes inside the nucleus, and α-tubulin, present in

both compartments, in the lysates (Fig. 1D). We compared the amount of Mcd1 in the

cleared lysate versus the total lysate. We found that when using the small impactor and

vial set, the cleared lysate had only 23.1% of the Mcd1 found in the total lysate,

whereas cleared lysate prepared using the medium impactor and vial set had 78.3% of

the Mcd1 found in total lysate. When we increased the volume of cells grown from 2 L to

4 L and used the medium impactor and vial, essentially all of the Mcd1 was found in the

cleared lysate. We then examined MT growth in these lysates (Fig. 1E). We did not

observe any MTs growing from seeds in the 2 L sample prepared in the small vial. The

increase in culture volume and resulting inclusion of more nuclear proteins and tubulin

in the lysate significantly changed the behavior of MTs in our assay. MTs in lysate from

a 2 L culture milled with a medium impactor exhibited growth only 42.6% of observed

time, whereas MTs in lysate from a 4 L culture grew 76.6% of the time. These results

indicated that using a larger and heavier impactor and milling a greater mass of cells

were necessary to efficiently recover nuclear proteins in the cleared lysates. This result

may be attributable to increased efficiency in lysing cells and/or organelles.

To ensure consistent protein composition and concentration levels across all

preparations, we standardized our harvest at several steps. First, 4 L cultures were

grown to a standard density. Strains harvested without arrest were grown to late log-

phase (OD 600 ≈ 0.7). If the strains were to be arrested by temperature shift, they were

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first cultured to an OD 600 ≈ 0.35 and then shifted to the restrictive temperature for 3

hours. Next, when grinding cells using the cryogenic impact mill for lysate preparation,

exactly 4 g of cells was used for each preparation as this was the minimum mass of

material harvested from 4 L of culture under our conditions. And third, milling was only

done in the medium vial and impactor set. This procedure controlled for any effects on

lysis efficiency that occur due to spatial constraints of the impactor and sample within

the sample vials.

Cell-cycle Dependence of MT Activity

As mentioned above, MT growth and dynamics were only observed when otherwise

wild-type cultures were arrested at different stages of the cell cycle (Fig. 1A,

Supplemental Movies 1-3). Asynchronous and G1-arrested lysates did not consistently

show any appreciable GFP-tubulin signal growing off of the rhodamine-labeled

seeds. Table 1 lists the rates of growth and shrinkage, the frequency of catastrophe

and rescue, and the overall dynamicity of the S phase-, metaphase-, and anaphase-

arrested lysates. In addition to the above measurements, we pooled the times of

growth, shrinking, and pausing for each population of MTs to create growth profiles for

the respective cell cycle stages (Fig. 1F). These data support previous findings that all

MT dynamics parameters are dependent upon the cell cycle.

The lack of assembled MTs in asynchronous and G1-arrested lysates led us to

hypothesize that an inhibitor of polymerization might be present in these lysates but not

in lysates from later arrest points in the cell cycle. We further postulated that a mixture

of G1 and S phase lysates might have characteristics intermediate between the original

lysates if key factors are titratable, or might exhibit the characteristics of G1 lysate if the

inhibitor acts in a dominant manner. We tested this by preparing two lysates arrested in

different cell cycle stages, mixing them at different ratios, and incubating for 5 minutes

before adding nucleotides and flowing them onto slides. S phase and G1 lysates were

mixed at 9:1, 3:1, 1:1, and 1:3 ratios, respectively, run through our assay, and analyzed

by kymographs. The 9:1 lysate mixture declined in time spent growing compared to S

phase lysate, down to 63.7% from 74.5%, and an increase in shrinking time and

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pausing time (5.6% to 6.7% and 19.9% to 29.6%, respectively) (Fig. 2A). Interestingly,

the growth rate increased from 0.35 ± 0.11 to 0.42 ± 0.16 μm min-1 (Fig. 2B). When the

amount of S phase lysate was decreased to a 3:1 ratio, the growth rate was only slightly

faster than S phase (0.40 ± 0.16 μm min-1) and the time spent paused was doubled to

47.4%. An equal mixture of lysate slowed the growth rate slightly to 0.30 ± 0.15 μm

min-1 and increased the amount of time the MTs were shrinking (5.6% to 10.3%) or

pausing (19.9% to 47.6%) Importantly, none of the shrinkage rates in these experiments

was statistically different from each other (Fig. 2C). When the ratio was reversed to 1

part S phase and 3 parts G1 lysate, there were no measurable MTs grown from the

seeds, similar to G1 lysate. These data show that G1-arrested lysate has a dominant

effect on MT dynamics when constituting 75% of a mixture with S phase-arrested

lysate, but that the effect becomes titratable when constituting 50% or less of the

mixture.

Association of MAPs with Dynamic MTs

We next determined whether our assay is amenable for studies on the association of

MAPs with MTs. We used an RFP-tagged clone of the yeast homolog of the EB1 tip-

tracking protein, BIM1, to follow its localization with MTs as they assembled in S phase

lysate (Fig. 3A, Supplemental Movie 4). Bim1-TagRFP-T was found along the entire

length of the MTs but concentrated at both growing and shrinking plus-ends, just as has

been described for in vivo (Wolyniak et al., 2006) and in vitro experiments (Zimniak et

al., 2009). We also investigated the association and translocation of kinesin motors

along these MTs by utilizing a strain that expressed KIP3-TagRFP-T and GFP-TUB1.

Observation of lysates, from cells arrested in metaphase, showed Kip3-TagRFP-T

bound to a MT and moved toward the plus end (Fig. S1 and Supplemental Movie 5).

Kymograph analysis of Kip3-TagRFP-T molecules showed that these motors would

move to the end of the MT and accumulate there as previously reported (Gupta et al.,

2006). We calculated the rate of Kip3-TagRFP-T movement to be 2.8 μm min-1, similar

to reported rates (Varga et al., 2009). These observations indicate that endogenous

MAPs localize in this assay in a similar way to what has been reported in live cells.

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Next, we determined whether our assay allows analysis of dynamic MAP exchange

when different lysates are flowed sequentially over assembled MTs. For these

experiments, two different lysates were thawed and cleared simultaneously before the

addition of nucleotide. The first lysate was loaded into the chamber and the reaction

was observed for 10 minutes so the MTs could assemble and become dynamic. Next,

a new field of MTs was found and imaged for 2 minutes. Then nucleotide was added to

the second lysate and this mixture was flowed into the chamber, replacing the first

lysate, all while the slide was mounted on the microscope. Imaging of the same field

resumed immediately, allowing us to follow effects on one population of MTs as the

lysate was changed. In control experiments of S phase lysate followed by fresh S

phase lysate, we observed MTs that continued to have dynamic MT activity (Fig.

S2). However, MTs assembled in an S phase lysate showed arrested growth and

began to shrink back to the seeds when G1 lysate was flowed into the chamber (Fig.

3B). The opposite was seen for the reciprocal shift experiment in which MT seeds in G1

lysate began to assemble MTs after the addition of S phase lysate (Fig. 3C).

These effects could be explained if the exchange of lysates were sufficient to dissociate

and/or exchange MAPs on the MTs. We tested this possibility by starting our assay

with a BIM1-TagRFP-T lysate from S phase-arrested cells and then flowing onto the

slide a similarly arrested lysate from cells that express unlabeled Bim1. In this

sequence of events, the Bim1-TagRFP-T signal was lost from the MTs after flow-

through of the second lysate (Fig. 3D).

Role of Motor Depolymerases

Having determined that MT polymerization and dynamics are dependent on the cell

cycle stage of cells from which lysates were prepared, we next investigated the possible

cause of the absence of MT assembly in our assay in asynchronous and G1-arrested

lysates. We tested the possibility that MT depolymerases prevent MT polymerization in

the lysates. Two MT depolymerases have been identified in budding yeast, Kip3

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(kinesin-8) (Gupta et al., 2006; Varga et al., 2009) and Kar3 (kinesin-14) (Endow et al.,

1994). We examined MT dynamics in the absence of these proteins in asynchronous

and cell cycle-arrested lysates.

Kip3 is a highly processive plus-end directed kinesin that destabilizes the plus end of

the MT (Gupta et al., 2006). Absence of Kip3 in cells leads to longer cytoplasmic MTs

(cMTs) as well as longer spindles that do not properly breakdown until after telophase

(DeZwaan et al., 1997; Huyett et al., 1998; Woodruff et al., 2010). We deleted KIP3 in

our strains to determine if it is responsible for preventing MT assembly in lysates

prepared from asynchronous and G1-arrested cells, and to determine how it affects MT

dynamics in lysates arrested in other cell cycle stages. Surprisingly, the asynchronous

kip3Δ lysates showed little MT assembly (Fig. 4A). The MTs that were observed had

muted dynamics (0.35 ± 0.18 μm min-1 growth rate, 0.55 ± 0.28 μm min-1 shrinkage rate,

and Table 2). In keeping with our previous results, the G1-arrested kip3Δ lysates

lacked any MT activity. However, in S phase-arrested kip3Δ lysates, the shrinkage rate

decreased 25% (from 0.84 ± 0.29 to 0.63 ± 0.24 μm min-1) without any major change in

the assembly profile. This result is in contrast to both the metaphase-arrested and

anaphase-arrested kip3Δ lysates, wherein the shrinkage rates increased (from 0.82 ±

0.23 to 1.31 ± 0.39 μm min-1 and 0.45 ± 0.21 to 0.59 ± 0.18 μm min-1,

respectively). Another interesting finding was that the growth rate increased in kip3Δ

when compared to WT for S phase- (from 0.35 ± 0.11 to 0.49 ± 0.15 μm min-1),

metaphase- (from 0.44 ± 0.16 to 0.85 ± 0.19 μm min-1), and anaphase- (from 0.30 ±

0.16 to 0.44 ± 0.17 μm min-1) arrested lysates. Moreover, in S phase kip3Δ lysates,

MTs spent less time growing (70.0%) and more time shrinking or pausing (7.2% and

22.8%, respectively) than in wild-type (Fig. 4B). This result was in contrast to what we

observed for metaphase- and anaphase-arrested kip3Δ lysates, wherein MTs spent the

majority of their time growing (Figs. 4C and 4D). The catastrophe frequencies mirrored

this change in growth profile for all 3 phases (Table 2). Based on these results, we

conclude that in our assay, Kip3 contributes to MT destabilization, but it alone does not

prevent MT growth in lysates from G1-arrested cells. Additionally, Kip3’s effects on MT

dynamics are cell cycle stage dependent.

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The other budding yeast protein suggested to have MT depolymerase activity is

Kar3. This kinesin-14 moves predominantly towards MT minus-ends (Molodtsov et al.,

2016) and is involved in kinetochore capture (Tanaka et al., 2005) and spindle formation

and stability (Hoyt et al., 1993; Saunders et al., 1997). Early work with Kar3 suggested

that it has MT depolymerase activity (Chu et al., 2005; Endow et al., 1994; Sproul et al.,

2005), but recent work from Mieck and colleagues has questioned this possibility (Mieck

et al., 2015). Though KAR3 is not necessary for cell viability, kar3Δ strains exhibit

mitotic delays (Meluh and Rose, 1990). To avoid any complications from cell cycle

defects in KAR3 mutants, we used the auxin-induced degron (AID) system to tightly

control the depletion of Kar3 activity from yeast (Fallis et al., 2009; Morawska and

Ulrich, 2013). Our Western blots showed that the vast majority of Kar3-9myc-AID

(referred to as Kar3-AID) protein is depleted upon treatment with indole acetic acid

(IAA) for 15 min (Fig. S3). We paired our KAR3-AID allele with cdc mutants and added

IAA during the last 30 minutes of a 3-hour temperature shift. Interestingly,

asynchronous KAR3-AID lysates displayed robust MT growth and dynamics (0.47 ±

0.17 μm min-1 growth rate, 0.90 ± 0.29 μm min-1 shrinkage rate, and Table 2). However,

G1-arrested lysates depleted of Kar3 did not assemble MTs. In S phase-arrested Kar3-

depleted lysates, the growth (0.66 ± 0.23 μm min-1) and shrinkage (1.05 ± 0.47 μm min-

1) rates increased 188.5% and 125%, respectively, when compared to wild-type, and

the time spent pausing increased (27.4%) at the expense of shrink time (3.0%) (Fig.

4B). For lysates of metaphase-arrested cells, we found that depletion of Kar3 led to an

increase in growth rate similar to the rate of kip3Δ lysates (0.84 ± 0.27 μm min-1) and a

slight increase in shrinkage rate compared to WT (1.08 ± 0.33 μm min-1). Though the

growth profile of this lysate appeared very similar to that of WT at the same arrested

stage, the overall dynamics were elevated (Table 2). Dramatic differences were seen in

anaphase-arrested lysates depleted of Kar3. Both the growth rate and shrinkage rate

increase significantly over WT and kip3Δ rates (0.68 ± 0.20 and 1.13 ± 0.29 μm min-1,

respectively) (Fig. 4D). MTs, in lysate of the Kar3-depleted anaphase-arrested cells,

spent the majority of their time (87.0%) growing and only seemed to change this

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behavior once they intersected or overlapped with another MT. This phenomenon

became more prevalent over time. These data point to Kar3 having different roles

throughout the cell cycle, with the most pronounced contribution to MT dynamics

observed in anaphase.

We next analyzed MT dynamics in lysates made from kip3Δ KAR3-AID strains. The

kip3 kar3 phenotype has not been reported previously due to a synthetic lethal

genetic interaction (Cottingham and Hoyt, 1997), though a kip3ts kar3Δ analysis did not

show any morphological defects in MTs (DeZwaan et al., 1997). Surprisingly, lysates

from the asynchronous culture did not support MT assembly (Fig. 4A). However,

lysates prepared from G1-arrested kip3Δ KAR3-AID lysate supported MT assembly

from the seeds with moderate growth and shrinkage rates (0.48 ± 0.16 and 0.79 ± 0.32

μm min-1, respectively), and the MTs only grew about 55% of the time. In lysates

prepared from the double mutant arrested in S phase, the MTs grew at a rate over twice

that of WT (0.76 ± 0.21 μm min-1). S phase lysate lacking Kar3 and Kip3 spent <2% of

the time shrinking (Fig. 4B). The dynamicity of MTs in the S phase-arrested double

mutants was the highest of any S phase lysate we observed (18.9 dimers sec-1). For

anaphase lysates lacking these two motors, the growth rate doubled and the shrinkage

rate increased 2.5-fold (0.74 ± 0.17 and 1.03 ± 0.34 μm min-1, respectively) as

compared to anaphase-arrested WT lysate, but the frequencies of catastrophe and

rescue remained similar. Despite this, MTs in the double mutant lysate spent 71.3% of

the time growing, 12.1% shrinking, and 16.6% pausing, which was quite different from

WT lysates in anaphase, but not that different from the single kip3Δ mutant (Fig.

4D). Based on the above observations, we conclude that these two kinesin-family

proteins contribute to MT destabilization, but in different capacities. When both are

missing, MTs grow more often than compared to what we observed for WT lysates and

for single mutant lysates, and at faster rates. Unexpectedly, in the absence of these

motors, MTs also depolymerize at a faster rate in anaphase lysates. Thus, their MT-

associated activities are more complex than previously appreciated.

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Two Types of Kar3 Heterodimers Behave Distinctly During the Cell Cycle

The Kar3 kinesin-family protein is unique in that it has two distinct binding partners that

create two different heterodimers with distinct cellular localizations and functions

(Manning et al., 1999; Page et al., 1994; Shanks et al., 2001). Cik1 has a nuclear

localization sequence that sequesters the Kar3/Cik1 heterodimer in the nucleus during

vegetative growth, where it acts on nuclear MTs (nMTs) (Manning et al.,

1999). Conversely, in mating cells, Cik1 is transcribed from an alternate start site,

which omits the NLS, sending the heterodimer to the cytoplasmic face of the SPB

(Benanti et al., 2009) where it forms the cMT array necessary for karyogamy (Hepperla

et al., 2014). Neither of these functions requires depolymerase activity. Kar3’s other

binding partner is Vik1. While both binding partners resemble kinesin-like proteins, Vik1

lacks the NLS found on Cik1 and its motor homology domain is truncated. Function of

the Kar3/Vik1 heterodimer is not well understood, but its localization seems to be limited

to the cytoplasmic face of the SPB and along cMTs (Manning et al., 1999). Using our

assay, we attempted to dissect the different activities of these two heterodimers by

creating single deletion mutants of CIK1 or VIK1 in our strains and prepared lysates

from each strain. In lysates from asynchronously grown cik1Δ cells, there was no MT

growth, similar to other asynchronous lysates. However, in lysates prepared from

asynchronously grown vik1Δ cells, MTs grew as well as they did in lysates from

asynchronously grown KAR3-AID cells (Fig. 5A). These data indicate that in the context

of a cellular lysate, Kar3/Vik1 but not Kar3/Cik1, possesses depolymerase

activity. Moreover, comparing the MT growth profiles for lysates prepared from

asynchronously grown KAR3-AID and vik1Δ cells, MTs were found to spend less time

growing (69.0% and 41.2%, respectively) and more time pausing (21.2% and 43.7%,

respectively) and shrinking (9.8% and 15.0%, respectively) when Vik1 was absent (Fig.

5B). Thus, both Kar3/Cik1 and Kar3/Vik1 heterodimers contribute to MT dynamics

regulation.

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DISCUSSION

A MT Dynamics Assay Using Budding Yeast Cleared Lysates

We created an assay that allows dynamics of individual MTs to be analyzed in the

context of a budding yeast cleared lysate made from total cellular proteins. Studies of

MT dynamics often involve in vivo imaging of MTs or in vitro analysis of activities of

purified proteins. Studies in Xenopus extracts have also made major contributions

(Belmont et al., 1990). Each approach has advantages and disadvantages. In vivo

imaging reveals the biological behavior of MTs, but often suffers from the inability to

distinguish individual MTs. Studies of purified proteins reveal the functional capacity of

individual proteins or small collections of proteins, but typically do not account for the full

complexity of proteins and regulators. Studies in Xenopus extracts allow the impact of

the full complexity of the cytoplasm to be explored, but it can be challenging to identify

the roles of individual proteins due to incomplete knockdown of activity using RNAi or

antibodies. An extract system from budding yeast has great promise because activities

can be explored in the total cellular protein complexity, and the functions of individual

proteins can be tested by making extracts from mutants. Moreover, mutants can be

exploited to explore several different cell cycle stages, and proteins can be fluorescently

tagged and expressed at endogenous levels for visualization in the extract system.

Our lysate preparation procedure excludes any non-native material and minimizes the

amount of added liquid. Frozen cells are pelleted, then crushed using a cryogenic

impact mill, creating a lysate that is expected to contain the full complement of soluble

cellular proteins. Minimizing protein dilution by adding only concentrated buffer and

protease inhibitors was emphasized to better mimic the native conditions within a cell.

The specific components added were intended to maintain conditions competent for MT

dynamics despite the potential disruption of vacuoles, mitochondria, peroxisomes, and

other organelles that might potentially release proteases, hydrolyze nucleotides, or

lower the pH of the lysate. The importance of maintaining the high protein

concentration on reproducibility might be due to an activity being near threshold levels

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at lower protein concentrations so minor differences resulted in substantial variation in

activity, or might be due to other differences related to lysis efficiency.

Our assay was developed to specifically enable analysis of plus end dynamics and not

nucleation. Ultracentrifugation removes membranes and large cellular debris from the

lysates so spindle pole bodies (SPBs), the yeast MT organizing centers, are not likely to

be present. The assay bypasses the nucleation steps of MT assembly because stable

MT seeds are affixed onto a coverslip. Cleared lysate was supplemented with excess

ATP, to keep ATPases including motor proteins and protein kinases active, and GTP,

for tubulin assembly. When the lysates were added to the seeds, microtubules could

grow and exhibit dynamics without the complication of a nucleation step. In budding

yeast, the minus ends of MTs are anchored into the SPB and are likely not dynamic

(Bergman et al., 2012; Byers et al., 1978; Maddox et al., 2000). In our assay, we mainly

observed MT growth only on one end of the seeds. Only rarely did tubulin assemble off

of both ends of a seed and the dynamics of the two ends were easily distinguishable by

the lower dynamicity of the minus end. Only the plus end was measured in these

cases.

In order to preserve physiological conditions, we monitored MT dynamics using a GFP-

tagged copy of TUB1, one of the two α-tubulin genes in yeast. Use of endogenously

tagged tubulin has the advantage of preserving the physiologically relevant mix of

tubulin isotypes and any relevant post-translational modifications. Recent work

supports the “tubulin-code” hypothesis: that the differences in MT and MAP behavior

between cell types can be a direct result of differences in the levels of tubulin isotypes

incorporated into MTs and the actions of acetylases, methylases, and detyrosinases

(Garnham and Roll-Mecak, 2012; Sirajuddin et al., 2014; Vemu et al., 2017). In S.

cerevisiae, levels of the α-tubulins, Tub1 and Tub3, affect the dynamics of MTs they

compose (Bode et al., 2003). Another advantage of using a homogeneous extract

system is that recent work has shown that MAPs from one species may interact

differently with tubulins from different species (Howes et al., 2018; Kollman et al., 2015;

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Podolski et al., 2014). For these reasons, our system is likely to have advantages in

preservation of physiological MT dynamic behavior and regulation.

Cell Cycle Dependence of MT Dynamics

A few key conclusions can be drawn from these data. First and foremost is that MT

dynamics in our reconstituted system approach known values collected from in vivo

data within 1-2 fold (Kosco et al., 2001). Table 1 reports the observed rates of growth

and shrinkage, the frequencies of rescue and catastrophe, and the overall dynamicity of

MTs during different phases of the cell cycle. When combined with the growth profiles

shown in Fig. 1F, these values approximate those reported in live cells. However, MT

dynamics in our assay are significantly less than what has been reported in purified

protein reconstitution systems that can recapitulate all phases of polymerization

dynamics (Moriwaki and Goshima, 2016). Additionally, upon comparison of catastrophe

events in our study versus those observed in previous in vitro systems, kymographs of

MTs from our assay show a slower rate of shrinking during catastrophe and MTs

typically undergo a rescue before reaching the GMPCPP-stabilized seed (Figs. 1, 3,

and 4). What has commonly been reported previously is a catastrophe that results in

disassembly completely back to the seed almost instantaneously. Our system more

closely resembles what is observed in vivo and in fully reconstituted systems than what

has previously been shown in vitro.

Our analysis is admittedly complicated in two ways by the fact that we have created a

system without nucleo-cytoplasmic boundaries and spatial cues. Firstly, we are

assaying nuclear and cytoplasmic activities together. Since budding yeast undergo a

closed mitosis, the composition of MTs in the nucleus can be different than those found

in the cytoplasm. Our method of lysis pools all available tubulin and MAPs, possibly

creating unnatural mixtures of proteins and protein modifications. The pools of MAPs

acting on the nMTs and cMTs normally remain separate throughout the cell cycle.

Secondly, the activity of some MAPs and the overall behavior of MTs has been shown

to be spatially regulated in the cytoplasm to facilitate specific cellular functions (Estrem

et al., 2017; Fukuda et al., 2014). Our assay lacks spatial cues other than proximity to

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other MTs. We can, nevertheless, speculate on the possible biological underpinnings of

our results with the caveat that, in the future, it might be important to develop an

approach that separates the cytoplasmic and nuclear pools of proteins. During S

phase, cMTs are rapidly growing so they can probe the cytoplasmic space in order to be

captured by Myo2 and transported toward the bud neck (Hwang et al., 2003; Yeh et al.,

2000; Yin et al., 2000). This may be why we observed primarily growing MTs.

Meanwhile, kMTs growing from the SPB probe the nuclear space to get captured by

kinetochores (Huang and Huffaker, 2006; Tanaka et al., 2005), which happens earlier in

the cell cycle than in cells with an open mitosis, which may also place a premium on MT

growth. In metaphase cells the kMTs must fluctuate between growing and shrinking

states to align the sister chromatids at the metaphase plate (Kosco et al., 2001;

Sprague et al., 2003), and this may explain the increased dynamics observed in our

system. Finally, during late anaphase, kMTs are short and stable after they have been

disassembled during chromosome separation. This may reflect the prolonged pause

time observed in lysates representing this stage. As for the ipMTs, in arrested cdc15-2

mutants, they are mostly stable as they have slid across each other to push the poles to

the daughter cells, though some muted plus-end dynamics continue through anaphase

(Fridman et al., 2009; Higuchi and Uhlmann, 2005; Rizk et al., 2014). Accordingly, the

growth profile for anaphase-arrested lysates were paused for >80% of the observed

time, possibly reflecting a blended state of the dynamics observed in vivo of kMTs and

ipMTs.

Despite the similarities between MTs observed in live cells and those in our assay

during later parts of the cell cycle, it was curious that MTs did not assemble in lysates

made from asynchronous and G1-arrested cells (with the exception of kip3Δ KAR3-

AID). Clearly, MTs exist and are dynamic in the cytoplasm during G1. There is

evidence that in budding yeast the nuclear MTs during this stage of the cell cycle are

firmly attached to kinetochores and are not dynamic (Dorn et al., 2005; Jin et al., 2000;

O’Toole et al., 1999). This MT behavior could be modulated through MAPs that are

only present in either the cytoplasm or the nucleus. Recent work suggests that the

nuclear-localized fraction of Stu2, the yeast XMAP215 homolog, is in part responsible

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for maintaining the short length and muted dynamics of the nMTs during mating (van

der Vaart et al., 2017). Results from our lysate mixing experiments (Fig. 2) suggest

that there is a titratable factor that can inhibit the growth and formation of MTs. The

sequential flow-through experiments also point to a factor(s) in G1 lysates that can stop

the growth of existing MTs that will then depolymerize all the way back to the seeds. In

the future, it will be important to use genetics and our extract system to identify this

factor or factors.

In total, these studies reveal pronounced changes in MT dynamics with different cell

cycle stages, the molecular and mechanistic basis for which can now be determined

using our budding yeast lysate system.

Emergent Properties of Two Kinesin Depolymerases

One approach to investigate the basis for the absence of MT assembly in lysates from

asynchronous and G1-arrested cells was to genetically remove the two known MT

depolymerases. Budding yeast has a kinesin-8, Kip3, and a kinesin-14, Kar3, which

have both been shown to destabilize MTs (Endow et al., 1994; Gupta et al., 2006). In

our assay, deleting the KIP3 locus resulted in only modest MT dynamics in

asynchronous lysates and no detectable MT growth in G1 lysates (Fig. 4). However, in

asynchronous lysates, knocking-down Kar3 protein with a KAR3-AID allele showed

robust MT assembly and dynamics. Interestingly, lysate from asynchronous kip3Δ

KAR3-AID cells lacks the ability to assemble MTs, unlike both single mutant

asynchronous lysates. The intriguing interplay of Kip3 and Kar3 activities became even

more apparent when the cells were arrested in S phase. When Kar3 is the only

depolymerase present, MTs shrunk for a greater portion of time. Without Kar3, MTs

paused longer and shrunk faster. Lysate generated from S phase-arrested cells lacking

both depolymerases spent more time growing and had an increased disassembly rate.

These observations are consistent with the possibility that Kip3 increases the

disassembly rate in S phase-arrested lysates. However, there are further layers of

complexity that need to be investigated because the microtubule disassembly rates of

kip3Δ lysates in other cell cycle phases increase (Table 2), as previously reported in

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cells (Fukuda et al., 2014; Gupta et al., 2006; Su et al., 2013). Upon examination of the

MT growth rates in these lysates, it is evident that they increase in either single mutant,

and even more so in the double mutant. An increase in growth rate in the absence of

depolymerases suggests that Kar3 and Kip3 do not only promote MT disassembly, but

they also modulate assembly rates.

This new technique may also begin to address questions that have been difficult to

answer using in vivo approaches alone. Kar3 has long been thought to exhibit MT

depolymerase activity. Direct evidence of this activity has thus far eluded the field,

perhaps due to the presence of two different Kar3 heterodimers (Kar3/Cik1 and

Kar3/Vik1) in separate cellular localizations and due to cell-cycle-dependent contexts.

In our assay, the presence of MT assembly in vik1Δ but not cik1Δ asynchronous cell

lysates indicates that the Kar3/Vik1 heterodimer has MT depolymerase activity at some

point in the cell cycle, likely during G1. It is now important to investigate the

contributions of the activity of these two heterodimers at different points in the cell cycle

in order to determine the nature of this phenotype.

Our assay has successfully reconstituted MT dynamics in cell lysate from a genetically

tractable organism. We have shown that MT behavior is greatly influenced by the cell

cycle stage, with roles for depolymerases. While the mixing of nuclear and cytoplasmic

contents is a complication in the system, this system can be further exploited to dissect

nuclear and cytoplasmic regulatory mechanisms, both of which are likely reflected in our

observations. This is particularly valuable for nuclear MT regulation since this MT

population is very difficult to visualize in live cells. Another consideration is that in

budding yeast, each SPB emanates 3-5 cytoplasmic MTs in G1 (Byers and Goetsch,

1975) or 2 during mitosis (O’Toole et al., 1999), and about 20 nuclear MTs, and this

major MT population cannot be readily observed in vivo. Since we are observing single

MTs in the complexity of soluble cellular lysates, the emergent properties of protein

populations on MTs can be studied. We have shown that MAPs can be visualized and

tracked on MTs in this system. During our work, we also observed MT bundling in

parallel and anti-parallel orientations, and MTs zippering together after meeting at their

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plus ends (not shown), suggesting that there is even more potential to elucidate

biologically important MT-based activities using the complex protein environment of

yeast lysate. As there are methods to disrupt every yeast gene, it is now possible to

assay, in an open extract system, the MT-related activities of essentially all yeast

proteins, and to adapt the assay for the study of additional cellular processes.

MATERIALS AND METHODS

Yeast Strains, Culturing and Harvesting

Yeast strains used in this study can be found in Table 4. Fluorescent and degradation

tags were integrated by homologous recombination as previously described. Strains

were grown in standard rich medium (YPD) at either 30°C or at 25°C if they contained a

temperature-sensitive allele. Strains containing the KAR3-9myc-AID allele were treated

with 250 μM 3-indole acetic acid (Sigma) in DMSO and buffered with 50 mM potassium

phosphate buffer at pH 6.2 for the 30 minutes just prior to harvesting.

For lysate preparation, strains were grown overnight in starter cultures and then diluted

into two parallel cultures of 2 L of YPD and grown until either OD600 ≈ 0.7 or OD600 ≈

0.35 if cultures were to be shifted for arrest. To arrest cells, cultures were shifted to

37°C for 3 hours before being harvested. Strains that contained an auxin-inducible

degron had a final concentration of 50 mM potassium phosphate buffer pH 6.2 and 250

μM indole acetic acid in DMSO added for the last 30 minutes of culturing. Cells were

then harvested by serial centrifugation at 6,000 RPM in a Sorvall RC5B with a SLA-

3000 rotor for 10 minutes at 4°C. Cells were then resuspended in ddH2O, transferred to

a 50 mL conical tube, and pelleted in a ThermoFisher CR3i table-top centrifuge for 3

minutes at 3,000 RPM. This wash and pelleting was repeated. After the second wash,

all standing moisture was removed from the cell pellet by aspiration. Cells were then

flash frozen in liquid nitrogen and stored at -80°C.

Lysates were prepared from frozen cell pellets by cryogenic impact milling in a SPEX

6875 Freezer mill High Capacity Cryogenic Grinder with a liquid nitrogen reservoir. 4 g

of frozen cells were weighed and placed into a medium-sized SPEX vial that had been

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pre-chilled in liquid nitrogen. Milling consisted of a 3 minute pre-chill followed by 10

cycles of 3 minutes of grinding at 30 impacts per second (15 cps) and 1 minute of

rest. The sample vial remained submerged in liquid nitrogen throughout the process.

Powdered lysate was collected in a 50 mL conical tube and stored at -80°C. Lysate

preparations were stable for >1 year.

Generating Rhodamine-labeled Tubulin Seed Mix

Previously isolated porcine tubulin was cycled to ensure the absence of non-functional

tubulin. This was mixed with both biotin-conjugated and rhodamine-labeled porcine

tubulin (Cytoskeleton Inc.) resuspended in PEM (80 mM PIPES pH 6.9, 1 mM EGTA, 1

mM MgCl2). Final concentrations of tubulin were 5 mg mL-1 unlabeled, 1 mg mL-1 biotin-

labeled, and 1 mg mL-1 rhodamine-labeled. GMPCPP (Jena Biosciences) was added to

a final concentration of 1 mM. Aliquots were flash frozen in liquid nitrogen and stored at

-80°C.

Cleaning of Glass Slides and Passivation of Coverslips

Prior to use, microscope slides (Corning Inc.) were washed in acetone and then 100%

ethanol for 15 minutes each. Slides were left to air dry before being stored in an airtight

container. 1.5 thickness coverglass (Corning Inc.) was first cleaned by submerging in

isopropanol and subjected to 20 minutes of sonication. Coverslips were then washed

twice with ddH2O and then in 70% ethanol for 1 minute each. The coverslips were then

blow dried with nitrogen gas and placed into a ceramic coverslip holder. This step was

followed by 10 minutes of illumination in a plasma cleaner chamber (Harrick Plasma

PDC-32G). A 0.1 mg mL-1 solution of PLL-g-PEG:PEG(3.4)-biotin (50%:50%) (SuSoS

AG) in 10 mM HEPES was prepared and 50 μL drops were placed onto

parafilm. Coverslips were placed on top of the drops and covered in a humidity

chamber for 1 hour. Passivated cover glass was then washed for 2 minutes in PBS and

rinsed in ddH2O for 1 minute. The coverslips were again air dried with nitrogen and

stored in an airtight container at 4°C. Passivated coverslips could be used up to 2

weeks later.

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Assembly of Flow Chamber

Double-sided tape was cut into thirds lengthwise and placed parallel on the center of a

cleaned microscope slide to create to equally sized channels. A passivated cover glass

was placed on top of the tape and a cotton-tipped applicator was used to gently press

on the cover slip at the lines of contact with the tape to ensure a continuous bond and

prevent leaks. A single channel was first washed with 30 μL PEM and wicked through

with Whatman paper. Neutravidin (Invitrogen) was diluted 1:400 in PEM and 30 μL was

pulled through the channel. The slide was then kept in a humidified chamber for 10

minutes. The channel was then washed twice with 30 μL of Pluronic™ F-127

(Invitrogen) diluted to 0.1% in PEM. This assembly was again incubated under humidity

for 10 minutes. During this incubation, porcine tubulin was assembled into seeds by

incubating Tubulin Seed Mix (above) at 37°C for 10 minutes. A 10X oxygen scavenging

(OS) solution was freshly prepared by mixing 10 μL of PEM with 5 μL of 40X glucose

oxidase + catalase (8 mg mL-1 glucose oxidase, 1.4 mg mL-1 catalase in PEM) and 5 μL

of 40X Glucose + 2-mercaptoethanol (180 mg mL-1 glucose, 20% 2-mercaptoethanol in

PEM). MT Seed mix was made by addition of 0.5 μL of assembled seeds to 36.5 μL of

PEM, 5 μL of 5 mg mL-1 casein, 5 μL of 10X OS, and 2.5 μL of 2% methylcellulose. 20

μL of MT Seed mix was then flowed through the channel. Excess MT Seed mix was

placed on the ends of the channel to prevent drying of the channel and the slide was

incubated at 37°C for 5 minutes. The channel was then washed with 50 μL of Warm

PEM (1X OS solution diluted with PEM at 37°C), leaving it ready for addition of lysate.

Preparation of Whole Cell Lysates

To prepare lysate for use in the assay, 0.22 g of powdered lysate was weighed out into

a 1.5 mL tube pre-chilled in liquid nitrogen. 25 μL of cold 10X PEM (800 mM PIPES pH

6.9, 10 mM MgCl2, 10 mM EGTA) and 0.5 μL of Protease Inhibitor Cocktail IV

(Calbiochem) were added to the lysate and spun down briefly. Lysate was thawed on

ice for 10 minutes before loading into the pre-chilled polycarbonate ultracentrifuge

tube. Lysate was then cleared of insoluble material by spinning at 346,000 x g for 25

minutes at 4°C. After the spin, 20 μL of cleared lysate was aliquoted into a fresh 1.5 mL

tube pre-chilled on ice. Cleared lysates were stable for >1 hr. 1 μL each of 20 mM ATP

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and 20 mM GTP (both in PEM) were added to cleared lysate before flowing through

prepared chamber.

Immunoblotting To compare relative protein concentrations in lysates, equal amounts of thawed and

diluted lysate powder were run on a 10% polyacrylamide gel. Preparation of cleared

lysate is described above. Total lysate was prepared by adding 220 µL of cold 2X PEM

and 0.5 µL of Protease Inhibitor Cocktail IV to 0.22 g of lysate powder. The mixture was

solubilized with an equal volume of 2X SDS sample buffer, boiled for 1 minute, and

briefly centrifuged before the supernatant was loaded onto the gel. Immunoblotting was

performed with 1:10,000 rabbit anti-Mcd1 (gift from Dr. Vincent Guacci, University of

California, Berkeley) and 1:1000 rat anti-α-tubulin (Santa Cruz Biotechnology, Cat.# 5C-

53030). Band intensities were measured with Image Studio Lite (LI-COR).

For tracking the amount of Kar3-9myc-AID protein in cells, cultures were grown to mid-

log phase in rich media and treated with 250 µM indole acetic acid in DMSO and

buffered with 50 mM potassium phosphate buffer at pH 6.2. Cells were then harvested

over a time course by centrifugation in a ThermoFisher CR3i table-top centrifuge for 3

minutes at 3,000 RPM. Cell pellets were resuspended in 1 mL 20% trichloro acetic acid

(TCA, Sigma) and transferred to a 2 mL screw-top tube. Cells were again pelleted in a

microfuge at top speed for 2 min. The supernatant was removed and the cell pellet was

resuspended in 200 µL 20% TCA. Approximately 200 µL of 425-600 µm acid-washed

glass beads (Sigma) was added to the tube which was then agitated on a vortexer for

10 min at 4°C. In order to collect lysate, a hole in the bottom of the tube was made with

a 25G needle and the entire tube was placed in a 5 mL round-bottom tube. This

apparatus was then centrifuged at 2500RPM for 3 min in the table-top centrifuge.

Beads were washed with 200 µL 5% TCA and collected in the same 5 mL tube, twice.

The pellet in the 5 mL tube was resuspended in the standing liquid and transferred to a

1.5 mL tube. This was spun at 5000 RPM for 10 min in a microcentrifuge. The

supernatant was discarded and the pellet was resuspended in 2X SDS sample buffer. 1

M Tris base was used to neutralize any remaining TCA. Equivalent OD amounts were

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then separated electrophoretically on a 10% polyacrylamide gel and transferred to

nitrocellulose. 9E10 mouse anti-myc primary antibody (prepared from hybridoma

supernatant) was used at 1:750 dilution for detecting KAR3-9myc-AID protein.

TIRF Microscopy

Once lysate was flowed through the prepared chamber, the slide was loaded onto an

Olympus IX81 inverted microscope with an environmental chamber pre-warmed to

28°C. Images were acquired with a 100X PlanApo objective (NA 1.45) and an Orca

CCD camera (Hamamatsu) using Metamorph software (Molecular Devices). TIRF was

used to illuminate a single plane of the field with 488 nm and 561 nm light every 5

seconds for 10 minutes.

Sequential Lysate Flow Assays

For sequential flow experiments, double-stick tape was used to make a perpendicular

chamber across the microscope slide. A passivated rectangular coverslip was then

placed on top of the tape to make a chamber that runs the entire width of the slide with

overhangs on both sides. The whole assembly was flipped with the coverslip-side down

in order to flow solutions through chamber as described above. All flow solution

volumes were increased by 10 μL to account for the larger volume of the chamber. 30

μL of the first lysate was flowed through the chamber and imaged for 10 minutes to

observe initial behavior. This was followed by 40 μL of the second lysate while the

sample remained mounted on the microscope.

Image and Data Analysis

Image files were analyzed using Fiji (NIH). Kymographs were constructed from all MTs

whose entire length was trackable for the entire movie after registration (StackReg,

Thévenaz et al. 1998). Dynamics parameters were calculated as in Moriwaki &

Goshima 2016. To calculate values, data from independent technical and biological

repeats from one genotype were pooled unless otherwise indicated. Growth and

shrinkage rates are reported as mean ± standard deviation. Statistical significance was

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determined by using an unpaired t-test. P values are reported as: * < 0.05, ** < 0.01, ***

< 0.001, and **** < 0.0001.

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ACKNOWLEDGMENTS

The authors would like to thank Itziar Ibarlucea Benitez for discussion and methods

development. We also thank members of the Drubin lab for discussion on experiments

and analysis. Strains with temperature-sensitive alleles of CDC genes were a gift from

Wei Guo (U. Penn).

COMPETING INTERESTS

The authors declare no competing interests in the completion of this work.

FUNDING

This work was supported by the National Institutes of Health (Grant R01 GM 47842) to

G.B..

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Figures

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Figure 1. Conditions for Assembly and Dynamics of MTs in Lysate. A) Two

examples of kymographs showing GFP-Tub1 (green) assembly from rhodamine-labeled

MT seeds (magenta) and in lysates from each stage in the cell cycle. Minus ends are

on the left, plus ends on the right. Time on the y-axis, 5 sec pixel-1, totaling 10 min,

starting from the top. B) Kymographs of S phase lysate without additional ATP (left)

and with 0.9 mM ATP. C) Effect of culturing and grinding conditions on MT behavior in

S phase-arrested lysate. D) Western blot probing for levels of tubulin and Mcd1, a

nuclear protein, in S phase lysates. Total lysate (T) and lysate cleared by

ultracentrifugation (C) were analyzed. The first row of numbers indicates the percent of

Mcd1 present in the cleared lysate compared to the total lysate normalized to the 4

L/Medium Vial lane. The second row of numbers indicates the percent of GFP-Tub1

present in the cleared lysate compared to the total lysate normalized to the 4 L/Medium

Vial lane. E) MT growth profiles in S phase lysates based on culture volume and milling

vial size. F) MT growth profiles for lysates by cell cycle. Percent of time spent in

phases for the entire population of MTs is compared across stages of the cell cycle. MT

measurements were pooled from three lysates generated from independent harvests of

cells. MT counts for each condition are listed in Table 1.

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Figure 2. MT Activity in Mixed Lysates is Titratable. A) MT growth profiles for mixed

lysates. The 1:3 S:G1 lysate lacks any MT activity. B) Growth rates of MTs in mixed

lysates compared to WT lysates arrested in S phase. C) MT shrinkage rates in S

phase-arrested and mixed lysates. None are statistically different from the others, with

the exception of the 1:3 S:G1 lysate. MT counts for each condition: 9:1 = 30, 3:1 = 25,

1:1 = 30.

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Figure 3. MAPs Dynamically Associate with MTs in Lysates. A) Kymograph of

GFP-Tub1 (green) with Bim1-TagRFP-T and rhodamine-labeled seeds (magenta).

Bim1-TagRFP-T is along MTs and decorates the plus end during growth and can be

present on shrinking ends. Horizontal scale bar is 2 μm, vertical bar is 2 minutes. B)

Fields of MTs in flow-through experiments. Dynamic MTs in S phase lysate arrested

and some began to shrink back to the seeds after flow through of G1 lysate. C) In the

reciprocal experiment, there were no MTs growing from seeds after 10 minutes in G1

lysate. 20 minutes after flow-through of S phase lysate, MTs have grown and were

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dynamic. D) Kymograph of S phase-arrested lysate expressing Bim1-TagRFP-T being

replaced with S phase-arrested lysate with untagged Bim1 (white line).

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Figure 4. MT Dynamics Depend Upon the Cell Cycle and Motor Activity in Lysates.

A) Kymographs of kip3Δ, KAR3-AID, and kip3Δ KAR3-AID lysates made from cells that

were treated with auxin (KAR3-AID genotype cells) and were either asynchronous or

arrested in S phase, metaphase, or anaphase. Rhodamine-labeled seeds in magenta

and GFP-Tub1 in green. Growth rates, shrinkage rates, and growth profiles for lysates

made from cells with these genotypes in S phase (B), metaphase (C), and anaphase

(D). MT measurements were pooled from three lysates generated from independent

harvests of cells. MT counts for each condition are listed in Table 2.

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Figure 5. Kar3 Binding Partners Alter MT Polymerization Activity. A) Fields of

rhodamine-labelled seeds (magenta) and GFP-Tub1 (green) using asynchronous cik1Δ

and vik1Δ lysates. Scale bar is 2 μm. B) MT growth profiles for KAR3-AID, cik1Δ and

vik1Δ lysates from asynchronous cultures. KAR3-9myc-AID data from asynchronous

lysate is presented again for comparison. No MT polymerization observed in three

separate cik1Δ lysates. Measurements for MTs in vik1Δ lysates were pooled from two

lysates. A total of 62 MTs were counted.

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TABLES

Table 1. Parameters of MT Dynamics in WT Strain Lysates in Each Cell Cycle Phase

Asynch. G1 S phase

n = 91

Metaphase

n = 58

Anaphase

n = 84

Growth rate

(μm min-1) - - 0.35 ± 0.11 (140) 0.44 ± 0.16 (109) 0.30 ± 0.16 (18)

Shrink rate

(μm min-1) - - 0.84 ± 0.29 (38) 0.82 ± 0.23 (102) 0.45 ± 0.21 (41)

Freq. of Cat. (sec-1)x103 - - 0.8 5.3 2.0

Freq. of Rescue (sec-1)x103 - - 14.1 12.5 13.4

Dynamicity

(dimers sec-1) - - 8.2 11.6 2.1

Footnote: No MT polymerization observed in asynchronous or G1-arrested lysates from

2 independent harvests of cells. MT measurements for each of S phase, metaphase,

and anaphase were pooled from three lysates generated from independent harvests of

cells. Number of events noted in parentheses.

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Table 2. Parameters of MT Dynamics in KIP3 and KAR3 Mutant Lysates

Asynch. G1 S phase Metaphase Anaphase

kip3Δ n = 69

KAR3-AID n = 152

kip3Δ KAR3-AID

n = 37

kip3Δ n = 90

KAR3-AID

n = 180

kip3Δ KAR3-AID

n = 86

kip3Δ n = 87

KAR3-AID

n = 108

kip3Δ n = 87

KAR3-AID

n = 240

kip3Δ KAR3-AID

n = 101

Growth rate (μm min-1)

0.35 ± 0.18 (71)

0.47 ± 0.17 (220)

0.48 ± 0.16 (95)

0.49 ± 0.15 (619)

0.66 ± 0.23 (425)

0.76 ± 0.21 (165)

0.85 ± 0.19 (245)

0.84 ± 0.27 (356)

0.44 ± 0.17 (210)

0.68 ± 0.20 (438)

0.74 ± 0.17 (238)

Shrink rate (μm min-1)

0.55 ± 0.28 (50)

0.90 ± 0.29 (55)

0.79 ± 0.32 (39)

0.63 ± 0.24 (118)

1.05 ± 0.47 (94)

0.94 ± 0.42 (10)

1.31 ± 0.39 (91)

1.08 ± 0.33 (291)

0.59 ± 0.18 (28)

1.13 ± 0.29 (72)

1.03 ± 0.34 (92)

Freq. of Cat. (sec-1)x103 2.6 3.2 3.1 1.4 1.4 0.3 2.7 9.1 1.0 0.7 2.3

Freq. of Rescue (sec-1)x103 16.4 21.7 15.2 15.6 25.1 17.4 18.2 19.4 20.4 8.4 17.9

Dynamicity (dimers sec-1)

3.8 10.5 10.1 10.8 14.3 18.9 20.7 17.7 8.7 19.2 18.0

Footnote: MT measurements were pooled from three lysates generated from

independent harvests of cells. Number of events noted in parentheses.

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Table 3. Parameters of MT Dynamics in cik1Δ and vik1Δ Asynchronous Lysates

cik1Δ vik1Δ

n = 62

Growth rate

(μm min-1) -

0.43 ± 0.24

(125)

Shrink rate

(μm min-1) -

0.76 ± 0.47

(67)

Freq. of Cat. (sec-1)x103

- 3.2

Freq. of Rescue (sec-1)x103

- 16.8

Dynamicity (dimers sec-1) - 7.5

Footnote: No MT polymerization observed in three separate cik1Δ lysates.

Measurements for MTs in vik1Δ lysates were pooled from two lysates. Number of

events noted in parentheses.

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J. Cell Sci.: doi:10.1242/jcs.219386: Supplementary information

Kip3-TagRFP-T GFP-Tub1Rhodamine-seeds

GFP-Tub1 Kip3-TagRFP-T Rhodamine-seeds

2 �m

2 m

in

Figure S1. Kymograph of Kip3-TagRFP-T along GFP-Tub1 MTs. Motor proteins

(magenta) will bind the MT (green) and move along the remaining length to the plus

end.

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S S

t = 10 min t = 20 min

Rhodamine-seeds GFP-Tub1

Figure S2. Sequential Flow-through of S-phase Lysate. Fields of MTs and seeds in

control experiments for lysate flow-through. MTs grew and were dynamic in lysate

from S phase-arrested cells. More lysate was washed through the chamber and

slightly interrupted the growth of existing MTs. After a short period, growth and

dynamics resumed similar to those observed before the lysate replacement.

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Kar3-9myc-AID

0 5 15 30 45 60 75 90 min

Figure S3. Depletion of Kar3 from Cells. Western blot of protein lysate from fixed

KAR3-9myc-AID cells treated with 250 μM 3-indole acetic acid. Cells were harvested

every 15 minutes and fixed.

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Movie S1. MTs in lysate from S phase-arrested cells. GFP-Tub1 (green) and

rhodamine-labeled seeds (magenta). Corresponds to kymographs in Figure 1A.

Scale bar is 2 μm. Playback is 20fps.

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Movie S2. MTs in lysate from metaphase-arrested cells. GFP-Tub1 (green) and

rhodamine-labeled seeds (magenta). Corresponds to kymographs in Figure 1A.

Scale bar is 2 μm. Playback is 20fps.

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Movie S3. MTs in lysate from anaphase-arrested cells. GFP-Tub1 (green) and

rhodamine-labeled seeds (magenta). Corresponds to kymographs in Figure 1A.

Scale bar is 2 μm. Playback is 20fps.

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Movie S4. Association of Bim1 with MTs in lysate. A zoomed-in view of Bim1-

TagRFP-T (magenta) associated with MTs (green) along their length and accumulated

at the plus ends from lysate of cells arrested in S phase. Rhodamine-labeled seeds

are also in magenta at the minus ends. Corresponds to kymographs in Figure 3A.

Scale bar is 2 μm. Playback is 20fps.

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Movie S5. Translocation of Kip3 along MTs in Lysate. Kip3-TagRFP-T (traveling

dots) and rhodamine-labeled seeds (bright bars) on the left and GFP-tubulin MTs on the

right from lysate of cells arrested in metaphase. Kip3-TagRFP-T puncta bound along

the MT and moved towards the plus end. Corresponds to kymographs in Figure S1.

Scale bar is 2 μm. Playback is 20fps.

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Table S1. Strains Used in this Study

Strain Name Genotype

DDY3435 MATα, lys2-801, his3Δ-200, leu2-3, 112, ura3-52::GFP-TUB1::URA3

DDY5662 MATa, lys2-801, his3Δ-200, leu2-3, 112, ura3-52::GFP-TUB1::URA3, cdc28-4

DDY5663 MATa, lys2-801, his3Δ-200, leu2-3, 112, ura3-52::GFP-TUB1::URA3, cdc7-1

DDY5664 MATa, lys2-801, his3Δ-200, leu2-3, 112, ura3-52::GFP-TUB1::URA3, cdc23-1

DDY5665 MATa, lys2-801, his3Δ-200, leu2-3, 112, ura3-52::GFP-TUB1::URA3, cdc15-2

DDY5666 MATa, lys2-801, his3Δ-200, leau2-3, 112, ura3-52::GFP-TUB1::URA3, BIM1-TagRFP-T::kanMX, cdc7-1

DDY5667 MATa, lys2-801, his3Δ-200, leu2-3, 112, ura3-52::GFP-TUB1::URA3, KIP3-TagRFP-T::HIS3MX, cdc23-1

DDY5668 MATa, lys2-801, his3Δ-200, leu2-3, 112, kip3Δ::kanMX, ura3-52::GFP-TUB1::URA3

DDY5669 MATa, lys2-801, his3Δ-200, leu2-3, 112, kip3Δ::kanMX, ura3-52::GFP-TUB1::URA3, cdc28-4

DDY5670 MATa, lys2-801, his3Δ-200, leu2-3, 112, kip3Δ::kanMX, ura3-52::GFP-TUB1::URA3, cdc 7-1

DDY5671 MATa, lys2-801, his3Δ-200, leu2-3, 112, kip3Δ::kanMX, ura3-52::GFP-TUB1::URA3, cdc 23-1

DDY5672 MATa, lys2-801, his3Δ-200, leu2-3, 112, kip3Δ::kanMX, ura3-52::GFP-TUB1::URA3, cdc 15-2

DDY5673 MATa, lys2-801, his3Δ-200, leu2-3, 112, KAR3-9myc-AID::HIS3, TIR1-::LEU2, ura3-52::GFP-TUB1::URA3

DDY5674 MATα, lys2-801, his3Δ-200leu2-3, 112, KAR3-9myc-AID::HIS3, TIR1-::LEU2, ura3-52::GFP-TUB1::URA3, cdc 28-4

DDY5675 MATα, lys2-801, his3Δ-200, leu2-3, 112, KAR3-9myc-AID::HIS3, TIR1-::LEU2, ura3-52::GFP-TUB1::URA3, cdc7-1

DDY5676 MATa, lys2-801, his3Δ-200, leu2-3, 112, KAR3-9myc-AID::HIS3, TIR1-::LEU2, ura3-52::GFP-TUB1::URA3, cdc23-1

DDY5677 MATa, lys2-801, his3Δ-200, leu2-3, 112, KAR3-9myc-AID::HIS3, TIR1-::LEU2, ura3-52::GFP-TUB1::URA3, cdc 15-2

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DDY5678 MATa, lys2-801, his3Δ-200, leu2-3, 112, KAR3-9myc-AID::HIS3, TIR1-::LEU2, ura3-52::GFP-TUB1::URA3, kip3Δ::kanMX

DDY5679 MATa, lys2-801, his3Δ-200, leu2-3, 112, KAR3-9myc-AID::HIS3, TIR1-::LEU2, ura3-52::GFP-TUB1::URA3, kip3Δ::kanMX, cdc28-4

DDY5680 MATα, lys2-801, his3Δ-200, leu2-3, 112, KAR3-9myc-AID::HIS3, TIR1-::LEU2, ura3-52::GFP-TUB1::URA3, kip3Δ::kanMX, cdc7-1

DDY5681 MATa, lys2-801, his3Δ-200, leu2-3, 112, KAR3-9myc-AID::HIS3, TIR1-::LEU2, ura3-52::GFP-TUB1::URA3, kip3Δ::kanMX, cdc15-2

DDY5682 MATα, lys2-801, his3Δ-200, leu2-3, 112, ura3-52::GFP-TUB1::URA3, cik1Δ::HIS3

DDY5683 MATα, lys2-801, his3Δ-200, leu2-3, 112, ura3-52::GFP-TUB1::URA3, vik1Δ::HIS3

All strains were constructed for this study, except for DDY3435 (source: Drubin/Barnes

lab). All strains are S288C background.