Isolation of the microtubule-vesicle motor kinesin from rat liver: Selective inhibition by cholestatic bile acids

GASTROENTEROLOGY 1995;108:824-833

Isolation of the Microtubule-Vesicle Motor Kinesin From Rat Liver: Selective Inhibition by Cholestatic Bile Acids DAVID L. MARKS, NICHOLAS F. LARUSSO, and MARK A. McNIVEN Center for Basic Researchin DigestiveDiseases, MayoClinic, Foundation,and Graduate School, Rochester, Minnesota

Background~Aims: Vesicular transport is supported by

microtubule-based, force-transducing adenosine triphosphatases (ATPases), such as kinesin, a ubiquitous motor enzyme that has been well studied in neuronal tissues. Although vesicular transport is important for hepatocellular secretory and clearance activities, the role of kinesin in liver function is poorly understood. Furthermore, the effects of bile acids on kinesin are unknown. M e t h o d s : Kinesin was purified from rat liver cytosol by conventional chromatography and microtubule affinity binding and was characterized by immunoblotting with domain-specific kinesin antibodies and amino acid sequencing of tryptic fragments. Kinesin activity was measured with and without bile acids using an in vitro motility assay and ATPase assays. Results: Immunoblot analysis and partial amino acid sequencing of purified kinesin showed that the sequence at the heavy chain of hepatic kinesin is nearly identical to that of brain kinesin. Purified kinesin transported microtubules in vitro with a velocity of - 0 . 5 I~m/s; this activity was significantly inhibited by 0 . 5 - 1 mmol/L taurochenodeoxycholate but not by tauroursodeoxycholate. At a dose of i mmol/L, chenodeoxycholate conjugates, but not ursodeoxycholate or cholate conjugates, directly inhibited the ATPase activities of kinesin and another microtubule motor, cytoplasmic dynein. Conclusions: Cholestatic concentrations of chenodeoxycholate conjugates directly inhibit the activity of microtubule motors, suggesting a possible mechanism for impairment of vesicular transport in cholestasis. 'epatocytes use vesicular mechanisms to perform .many critical hepatic functions, including plasma protein secretion, clearance of circulating substances, and biliary secretion. ~'2 This vesicular trafficking process is highly dependent on microtubules as has been shown by numerous studies in which plasma protein secretion, 3'4 delivery of endocytosed proteins to lysosomes, 5'6 transcytosis of immunoglobulin A, 7'8 and biliary excretion of proteins and lipids 9'1° were inhibited by microtubule poisons. In nonhepatic cells and tissues, microtubulebased transport has been shown to be driven by the motor enzymes kinesin and cytoplasmic dynein, which hy-

H

drolyze adenosine triphosphate (ATP) to generate a motive force and support the movement of vesicles along microtubules) 1'12 Kinesin has been found in all eukaryotic cells and is described as an orthograde motor believed to transport secretory vesicles away from the centrosome toward the cell surface. It is known to consist of two heavy chains ( ~ 1 2 0 kilodaltons), which contain highly conserved enzymatic domains for microtubule binding and ATP hydrolysis, and two associated light chains ( ~ 6 0 kilodaltons) at the tail of the molecule, which may mediate motor-vesicle binding) ~'13 Thus, it is predicted that the microtubule-based cytoskeleton and associated motor enzymes play important roles in hepatic vesicular transport. However, there is little information about microtubule motors in the hepatocyte. Kinesin has been previously reported in liver. 14'15 W e recently showed using immunochemical methods that kinesin is present in hepatocytes, where it may play a role in the transport of Golgi and transcytotic vesicles.16 However, kinesin from rat liver has never been purified or characterized. Indeed, it is not known if hepatic kinesin is identical to conventional kinesin, 1:-19 one of the many kinesinlike proteins identified by molecular techniques 11'19'2° or a novel gene product. Further, it has never been shown that liver kinesin is able to translocate vesicles or other particles. Thus, our first aims were to purify and characterize liver kinesin. Microtubule-dependent vesicular transport in the liver is reported to be altered during cholestasis, 21'22 possibly as a result of elevated levels of bile acids in plasma. Indeed, the effects of bile acids on vesicular transport in the liver are significant and have been well documented. It is known that elevated bile acid levels can inhibit hepatic uptake of circulating proteins, 23'24 stimulate the Abbreviations used in this paper: EGTA,ethyleneglycol-bis(~-aminoethyl ether)-N,N,N',N'-tetraacetic acid; GCDC, glycochenodeoxycholate; GUDC,glycoursodeoxycholate;HPLC, high-performanceliquid chromatography; MES, 2-(N-morpholino)ethanesulfonicacid; SOS-PAGE,sodium dodecylsulfate-polyacrylamidegel electrophoresis; TODC,taurochenodeoxycholate;TUDC,tauroursodeoxycholate. © 1995 by the American GastroenterologicalAssociation 0016-5085/95/S3.00

March 1 9 9 5

biliary excretion of lysosomal contents and endocytic markers, 25-28 and alter the distribution and morphology of transcytotic vesicles and lysosomes in the canalicular region of the hepatocyte. 25'29 Currently, the mechanisms by which bile acids alter microtubule-based vesicular transport in the hepatocyte are undefined. Bile acids have been reported to directly inhibit the activities of some adenosine triphosphatases (ATPases) 3° 32 and, therefore, may also affect microtubule-based motor ATPases. Thus, using purified liver kinesin and in vitro assays, we tested the hypothesis that bile acids directly inhibit kinesin activity. Our results show that hepatic kinesin is similar to neuronal kinesin in its polypeptide composition, amino acid sequence, and biological activities, including its ability to move microtubules in vitro. Furthermore, liver kinesin activity in vitro is significantly inhibited at a 1mmol/L dose by chenodeoxycholate conjugates but not ursodeoxycholate or cholate conjugates. These results show that cholestatic concentrations of chenodeoxycholate conjugates can directly inhibit the activity of a microtubule motor and suggest that one possible mechanism for alterations of vesicular transport in cholestasis is the impairment of motor enzyme function by elevated bile acid levels.

M a t e r i a l s and M e t h o d s Materials Taxol was a gift from the National Cancer Institute (Bristol-Myers Squibb, Princeton, NJ). Tubulin was purified from bovine brain by successive polymerization and depolymerization, followed by chromatography on P l l phosphocellulose (Whatman Inc., Clifton, NJ) as previously described. 33'34 Purified tubulin (5-8 mg/mL) was polymerized into Taxol-stabilized microtubules by incubation for 30 minutes at 37°C in 0.1 mol/L piperazine-N-N'-bis-(2-ethanesulfonic acid) (pH 6.9; Calbiochem, La Jolla, CA) with 0.1 mmol/L guanosine 5'-triphosphate (Sigma Chemical Co., St. Louis, MO), 1 mmol/L MgSO4, and 30 btmol/L Taxol. Axonemes (demembran-ated flagella) were prepared from Chlamydomonas sp. as described previously. 35 Cytoplasmic dynein was prepared from rat liver as described previously. 15 Bile acids, apyrase, and all other reagents were from Sigma Chemical Co.

INHIBITION OF LIVER KINESIN BY BILE ACIDS

825

MgC12, 0.1 mmol/L ethylenediaminetetraacetic acid, and 1 mmol/L dithiothreitol (ph 7.5) plus protease inhibitors (0.5 mmoI/L phenylmethylsulfonyl fluoride, 10 btg/mL leupeptin, 10 btg/mL tosyl arginine methyl ester, 1 btg/mL pepstatin, and 0.6 mg/mL benzamidine). The following steps were performed at 4°C unless otherwise noted. Livers (80-100 g) were homogenized in 3 - 4 volumes of homogenization buffer using a Potter-Elverhjem homogenizer (two strokes at 2000 rpm). The homogenate was centrifuged for 30 minutes at 18,000g. The supernatant was then centrifuged for 60 minutes at 110,000g to yield a high-speed supernatant. The high-speed supernatant was loaded onto a P l l phosphocellulose column, which was then washed with homogenization buffer and eluted with 700 mmol/L KC1 in a homogenization buffer. The protein peak eluted from the phosphocellulose column was detected by UV absorption, pooled, and dialyzed against homogenization buffer overnight. The dialyzed phosphocellulose peak was then loaded onto a Mono Q FPLC column (Pharmacia, Piscataway NJ) and eluted with a gradient of 0.1 - 1 mol/L KC1 in homogenization buffer. Fractions enriched in kinesin were identified using sodium dodecyl sulfate-polyacrylamide gel electrophoresis (SDS-PAGE) or immunoblotting. Peak kinesin fractions were eluted at ~300 mmol/L KCI, pooled, and then dialyzed against 0. t mol/L 2-(N-morpholino)ethanesulfonic acid (MES), 0.2 mol/L KC1, 0.5 mmol/L MgSO4, 1 mmol/L ethylene glycol-bis(~-aminoethyl ether)-N,N,N',N'-tetraacetic acid (EGTA), and 2 mmol/L dithiothreitol (pH 6.9) (MES buffer) with 30% glycerol. To further purify kinesin by microtubule affinity binding, dialyzed kinesin-containing fractions were incubated for 15 minutes at 37°C with Taxol-stabilized microtubules (final concentration, 0.5-1.0 mg/mL), tripolyphosphate (final concentration, 3 mmol/L), and Taxol (final concentration, 20 btmol/ L). Samples were then centrifuged at 180,000g for 90 minutes at 30°C. To wash off contaminating proteins, pellets were resuspended in MES buffer with 3 mmol/L tripolyphosphate and 20 btmol/L Taxol by repeated passages through a 22-gauge needle; incubated for 10 minutes at 37°C; and centrifuged a t 40,000g for 30 minutes. To release kinesin from microtubules, the washed pellets were resuspended as described above in MES buffer with 10 mmol/L ATP, 10 mmol/L MgSO4, and 20 btmol/L Taxol; incubated for 15 minutes at 37°C; and centrifuged as described above. The final supernatant (purified kinesin) was adjusted to 1 mol/L glycerol; aliquots were frozen in liquid nitrogen and stored at -80°C.

Characterization of Rat Liver Kinesin Purification of Rat Liver Kinesin Kinesin was purified from rat liver using a modification of a method used to purify kinesin from bovine adrenal medulla. 36 Sprague-Dawley rats (150-250 g) were killed by decapitation in accordance with protocols approved by the Mayo Animal Care Committee. Rat livers were rapidly removed and then rinsed in ice-cold homogenization buffer containing 50 mmol/L imidazole, 100 mmol/L KCI, 0.5 mmol/L

The polypeptide composition of purified liver kinesin was determined by SDS-PAGE using 8.5% acrylamide gels according to Laemmli. 3v To determine partial peptide sequences of purified kinesin heavy chains, heavy chains were excised from SDS-PAGE gels and digested in situ with trypsin as previously described. 38 Tryptic fragments were then separated by high-performance liquid chromatography (HPLC) on an ABI 130A Separation System (Applied Biosystems, Foster

826

MARKS ET AL.

City, CA) using an Applied Biosystems Aquapore OD-300 C18 column (2.1 X 220 mm), with a linear gradient of 5 % 40% acetonitrile in 0.1% trifluoroacetic acid and water run over 60 minutes. The purified peptides were applied directly to Porton peptide supports and sequenced using a Porton 2090E gas phase sequencer (Beckman Instruments, Fullerton, CA). The peptide sequences were compared with published sequences of kinesin heavy chain using the DNA* program (DNAStar Inc., Madison, WI). To further characterize kinesin, we prepared domain-specific antibodies to kinesin heavy chain. Four different peptides based on conserved regions of enzymatic domain of the kinesin heavy chain (see Results for sequences) were synthesized by the Mayo Clinic Peptide Core, conjugated to keyhole limpet hemocyanin, suspended in Freund's adjuvant, and injected into New Zealand white rabbits. Sera were collected and screened for the ability to recognize kinesin on immunoblots. Partial characterization of one of these polyclonal antisera, MMR44, was recently described by us in a separate publication) 6 Purified kinesin was immunoblotted against these antibodies using established methods) 9

GASTROENTEROLOGY Vol. 108, No. 3

kDa 205 - -

116 - -

~ - Kinesin heavy chain

96--

Kinesin - 7 light chains

66--

"~- Tubulin 45--

Figure 1, SDS-PAGE showing the purification of kinesin from rat liver.

Kinesin Motility and ATPase Assays The ability of purified kinesin to move microtubules along a substrate was tested using an in vitro microtubule motility assay similar to previously described techniques. 36'4° Purified kinesin in solution was first added to a glass slide and allowed to adhere for ~ 5 minutes. Axonemes, which are demembranated flagella made up of microtubule bundles, were then added. Lines of vacuum grease were placed on the slide on two opposite sides around the kinesin, and a coverslip was placed over the slide. Axoneme motility was initiated by the perfusion of motility buffer (30 mmol/L imidazole, 2 mmol/ L EGTA, 4 mmol/L MgC12, and 4 mmol/L ATP, p H 7.5) under the coverslip. In some experiments, after perfusion with normal motility buffer, motility buffer plus various bile acids were perfused. Kinesin-driven movements of axonemes along glass slides were observed using a Hamamatsu video camera (Hamamatsu Photonics, Hamamatsu-City, Japan) coupled to a Zeiss Axiovert microscope (Zeiss, Batavia, IL) and recorded on videotape using a Panasonic super VHS recorder (Panasonic Communications Seacaucus, NJ). Images of moving axonemes were played back, and their position at discreet time points was marked on transparencies overlayed on a video screen. Distances traveled across the screen were measured and calibrated using video images of a stage micrometer viewed at the same magnification. The velocities of axoneme movement were expressed in micromoles per second. Kinesin ATPase activity was quantified by incubating purified kinesin with 0.75 mmol/L ATP for 30 minutes in the presence or absence of 0.8 mg/mL microtubules in 15 mmol/ L imidazole and 5 mmol/L MgCI2 (pH 7.0) and measuring inorganic phosphate released using a previously described colorimetric method. 4~ Cytoplasmic dynein and apyrase ATPase activities were measured in the absence of microtubules as

Rat liver samples were electrophoresed on 8.5% gels and stained with Coomassie blue. Fractions loaded were liver cytosol, kinesincontaining peak fractions visualized by phosphocellulose and Mono Q chromatography. Precipitation of kinesin from the Mono Q peak fractions with exogenous microtubules yielded a pellet significantly enriched in kinesin. Purified kinesin was released into the supernatant from the microtubule pellet with ATP. Protein loads were 200, 25, 25, 25, and 10 ~g (from left to right). Numbers indicate the position of molecular weight standards.

described earlier except that the ATPase buffer had p H 8.0 for dynein; for apyrase, pH was 6.5 and MgCI2 was replaced by 2 mmol/L CaCl2 . In some experiments, various bile acids were added to the incubation solutions just before the start of the incubation. For all experiments, appropriate blanks were carried through all steps of the assay, so that absorbances due to microtubules, bile acids, and proteins alone could be subtracted.

Kinesin Microtubule Pelleting Assay To test the effects of bile acids on the ability of kinesin to bind microtubules in vitro, purified kinesin (1.25 ~g) was incubated with Taxol-stabilized microtubules (3.25 mg/mL) in the presence or absence of bile acids for 15 minutes at 37°C in MES buffer with 3 mmol/L tripolyphosphate and 20 ~mol/ L Taxol (final volume, 200 ~L). Samples were then centrifuged at 70,000g in a Beckman TLA-100 rotor. Pellets were then probed for kinesin by immunoblotting as previously described. 16

Results Purification and Characterization of Rat Liver Kinesin M a n y different m e t h o d s have been used to isolate k i n e s i n from a variety of different organisms and tissues.

March 1995

A

14

INHIBITION OF LIVER KINESIN BY BILE ACIDS

I

32

- -- RFRPLNE -- --

ILl

71

KFQGEDTVVIA . . . .

827

B

II1

KDVLEGYNGTIFAYGQTSSGKTHTMEGK-MMR43

204 I V

221

I K L S G K L Y L V D L A G S E K V S K T G A E G A V L D E A K N I N K S L S A L G N V I S A L A EGS - - RIS H S l F,,, - - 01

K

MMR48

MMR4$

V 273 I XXXm TYVPYRDSKMTRILQDS LGGNCRT17-HY L E 'A~

430

VI

D-I

43 45 48 50 I Liver kinesin

I

43 45 48 50I

Brain homogenate

-- KQLDD . . . .

MMR$O 915

VII

I

RGHSAQIAKP

X-Y~ IRPGQ-

Figure 2. Biochemical characterization of purified liver kinesin. (A) Tryptic fragments of purified liver kinesin heavy chain and amino acid sequences of peptides used to prepare kinesin antibodies; comparison with human kinesin heavy chain. Amino acid sequences shown are portions of a sequence of human kinesin heavy chain reported previously. 18 Dotted lines indicate portions of the sequence not shown (distances are not to scale). Numbers above the sequences indicate the position of the first residue in each region of interest. Lines above the sequences (labeled I-VII) indicate tryptic fragments homologous to overlined regions of human kinesin. X, amino acid residues in tryptic fragments that could not be unambiguously identified. Lines below the sequences indicate homologous regions of peptide sequences used to prepare polyclonal antibodies against kinesin heavy chain. The reference numbers of the antibodies produced are indicated in bold letters below these lines. Amino acid symbols interrupting lines showing tryptic fragments and peptides indicate a lack of identity with the human sequence at those residues. (B) Purified liver kinesin is recognized by antibodies against four different regions of kinesin heavy chain. Purified liver kinesin and crude rat brain homogenate samples were run on SDS-PAGE gels, transferred to polyvinylidene difluoride, and probed with antibodies (MMR 43, 45, 48, and 50) produced against synthetic peptides based on four enzymatic regions of kinesin heavy chain (see A). Note that each antibody recognizes a single band in liver but three bands in brain.

We adapted the purification scheme of Urrutia and Kachar (1991), 36 which has proven useful for the isolation of kinesin in relatively high yields from small quantities of tissue, for the purification of kinesin from rat liver. After successive phosphocellulose and Mono Q chromatography of rat liver cytosol, the kinesin heavy chain was visible on SDS-PAGE gels by Coomassie blue staining (Figure 1). Precipitation of kinesin from the Mono Q peak fractions with exogenous microtubules yielded a pellet that was significantly enriched in kinesin, with few contaminating proteins (Figure 1). Final extraction of kinesin from microtubule pellets with ATP yielded kinesin preparations with purities ranging from 80% to 90% as assessed by densitometry of SDS-PAGE gels (Figure 1). The purified kinesin fractions contained varying amounts (10%-15 % of total protein) of residual exogenous tubulin, but no other major contaminating proteins were visible on Coomassie blue-stained gels (Figure 1). The yield of purified kinesin was ~ 5 0 ng/mg of total liver protein (data not shown). We previously determined using immunoblotting that kinesin is present in liver at a concentration of ~0.3 btg kinesin/mg total protein. 16 Thus, our recovery of kinesin from liver was ~17%. Liver kinesin consisted of a heavy chain with a molecular weight of ~ 120 kilodaltons and light chains with molecular weights in the range of 60-80 kilodaltons, similar to size ranges reported for kinesin heavy and light chains

isolated from other sources. 13'42'43 Recovery of kinesin light chains in our preparations of purified liver kinesin was greater than those reported previously for kinesin purified from bovine adrenal medulla by a similar method, 36 possibly as a result of our extensive use of protease inhibitors. To confirm that we had isolated kinesin, we digested gel-purified rat liver kinesin heavy chain with trypsin, separated the tryptic peptides by HPLC, and performed amino acid sequencing on the peptides. Seven peptides that showed high homology to regions of published amino acid sequences of human kinesin heavy chain were identified (Figure 2A). The fragments were also highly homologous to the same regions of mouse kinesin heavy chain (data not shown). To our knowledge, no published sequences are available for rat kinesin heavy chain for comparison with our data. We identified peptides that corresponded to regions of the head (peptides I-V), stalk (peptide VI), and tail (peptide VII) of the kinesin heavy chain (Figure 2A), indicating that we had isolated the intact polypeptide. To further characterize purified kinesin, it was immunoblotted with polyclonal antibodies prepared against synthetic peptides (peptide sequences are shown in Figure 2A) representing consensus sequences of four different regions of the kinesin heavychain motor domain. Purified liver kinesin was recognized as a single band by all four of our kinesin peptide

828

MARKS ET AL.

A

GASTROENTEROLOGY Vol. 108, No. 3

B 0.6-

0.5

"3--,

0.4 ......... ........

e a#::~ { ,

~0.3

i:~,: { { {'g{

¢~ 0.2 > 0.1 ~a

1.......... N

I

I

No bile acids

I

0.3 0.5 1.0 I TUDC

I 0.3

0.5

1.01

TCDC

in,vii Bile Acid

Figure 3. Kinesin-mediated axoneme motility in vitro. As described in Materials and Methods, purified liver kinesin was allowed to adhere to glass cover slips. Axonemes and ATP were then added, and motility was observed by video microscopy. (A) Time lapse video micrograph showing kinesin-driven motility of axonemes. The positions of several moving axonemes are shown at 8-second intervals. One axoneme is indicated with an arrow to emphasize its movement over time (bar = 20 #m). (B) Kinesin-driven motility in vitro is inhibited by TCDC but not by TUDC. Bile acids were added to cover slips on which immobilized kinesin was transporting axonemes. Axoneme movements were recorded on videotape. Axoneme velocities were then measured and calibrated using a stage micrometer. Values are means _+ SE. *Values significantly different (P < 0.05) from controls in two-tailed t tests.

antibodies, whereas three kinesin heavy-chain isoforms (mol wt, 1 1 6 - 1 3 0 kilodaltons) were recognized in rat brain preparations (Figure 2B). These results provide evidence that the protein we purified from rat liver is indeed a conventional kinesin rather than a kinesinlike protein. Furthermore, they validate the use of our antibodies as sensitive and specific tools for the study of kinesin in hepatocellular vesicular trafficking. To confirm that purified liver kinesin was biologically active, we first assessed the ability of purified kinesin to support movement in vitro using an axoneme (demembranated flagella) gliding assay, a method frequently used to test the viability of molecular motor enzymes. As shown by time-lapse video microscopy in Figure 3A, purified kinesin indeed was able to move axonemes in vitro. The mean velocity of axoneme movement was ~ 0 . 5 btm/s, similar to velocities reported for kinesins from other sources. 36'40'44 W e also assessed the biological activity of purified kinesin using ATPase assays. In the absence of microtubules, kinesin ATPase activity was low ( 2 0 - 4 0 btmol" min 1. mg-i); however, in the presence of 0.8 mg/mL microtubules, the ATPase activity of different preparations of purified kinesin ranged from 300 to 700 btmol" min -1" mg -1 (data not shown). This 1 5 20-fold stimulation ofkinesin ATPase activity by microtubules is consistent with previous reports of the micro-

tubule dependence of kinesin ATPase activity. 36'42'45 These results show for the first time that liver kinesin is biologically active and support the prediction that liver kinesin mediates vesicle movement in vivo.

Effects of Bile Acids on Kinesin To test if kinesin activity is altered by elevated concentrations of bile acids, we examined the effects of bile acids on kinesin-driven in vitro motility using the axoneme gliding assay described earlier. For these experiments, axonemes were added to slides coated with kinesin; motility buffer containing ATP was then perfused under the cover slip. All slides were observed by video microscopy before the infusion of bile acids to ensure that normal axoneme motility was initially present. Using this assay, we compared the effects of taurochenodeoxycholate (TCDC), a common secondary cholestatic bile acid, and the taurine conjugate of ursodeoxycholate, an uncommon choleretic bile acid used in the treatment of cholestatic liver diseases. 46 As shown in Figure 3B, tauroursodeoxycholate (TUDC) at a dose of 0 . 3 - 1 . 0 mmol/L had no significant effect on kinesin-driven motility velocity. In contrast, TCDC at doses of 0.5 and 1.0 mmol/L significantly inhibited motility velocity (Figure 3B). In fact, on slides treated with these concentrations of TCDC, very few moving axonemes were observed, suggesting that most

March 1995

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INHIBITION OF LIVER KINESIN BY BILE ACIDS 829

~

~{

P S P S P S 0.2 mM 1.0 mM 0.2 mM TUDC TUDC TCDC

P S 1.0 mM TCDC

Control 0.2 mM 1.0 mM 0.2 mM TUDC TUDC TCDC

1.0 mM TCDC

P s Control

B

8060 40

~" 20

Figure 4. Bile acids do not inhibit the binding of kinesin to microtubules in vitro. Purified kinesin (1.25 pg) was incubated with Taxolstabilized microtubules (3.25 mg/mL) with or without bile acids and centrifuged, and the pellet was probed for kinesin by immunoblotting as described in Materials and Methods. (A) An immunoblot showing the detection of kinesin in microtubule pellets (P) and supernatants (s) after incubation with or without the indicated concentrations of bile acids. (B) Densitometric quantitation of the binding of kinesin to microtubule pellets. Values are expressed as the percent of total detectable kinesin bound to microtubules [100 × P/(P + S)] in each treatment and are means of two replicates. Note that bile acids had no significant effect on the extent of binding to microtubules.

axoneme movement was completely inhibited by TCDC (data not shown). To explore the mechanism by which TCDC may inhibit kinesin-driven motility, we next determined if TCDC inhibits the ability of kinesin to bind microtubules. W e used an in vitro binding assay in which purified kinesin was incubated with exogenous, Taxol-stabilized microtubules; the microtubules were pelleted by centrifugation; and the pellet was probed by immunoblotting for kinesin. W e found that neither TCDC nor T U D C at doses of 1 mmol/L had any significant effect on the binding of kinesin to microtubules in vitro (Figure

4). We then measured the effects of bile acids on the microtubule-stimulated ATPase activity of kinesin. W e found that TCDC caused a dose-dependent inhibition of kinesin ATPase activity with ~ 6 0 % inhibition at a 1mmol/L dose of TCDC (Figure 5). In contrast, TUDC did not significantly affect kinesin ATPase activity at this concentration. Because ursodeoxycholate is protective against some of the toxic effects of chenodeoxycholate

conjugates, 4v'4s we also tested the effects of TCDC (1 mmol/L) on kinesin pretreated with TUDC (0.5 mmol/ L). T U D C did not protect kinesin against inhibition by TCDC (data not shown). In fact, ATPase activity was lower in kinesin samples treated with both TUDC and TCDC than with TCDC alone (data not shown), most likely because of the total amount of bile acids present were higher in the former samples. To further understand the mechanisms by which bile acids affect kinesin activity, we next compared the effects of several bile acids on the ATPase activities of kinesin, cytoplasmic dynein (another motor enzyme from rat liver) prepared by the method of Collins and Vallee, t5 and apyrase, a commercially available Ca 2+ ATPase. As we reported above, liver kinesin has little ATPase in the absence of microtubules. Because of this low activity we could not conduct bile acid inhibition studies on kinesin activity in the absence of microtubules. In contrast, cytoplasmic dynein has significant basal ATPase activity without microtubules and is only stimulated 2 - 4 - f o l d by microtubules. 15'49'5° Apyrase is an ATP diphosphohydrolase with no known dependence on microtubules for activity. Thus, we could measure dynein and apyrase activity with no microtubules present to determine if bile acids could affect ATPase activity even in the absence of microtubules. When six bile acids were tested at doses of 1 mmol/L for their effects on kinesin, dynein, and apyrase ATPase activities, the results were surprising. Both TCDC and glycochenodeoxycholate (GCDC) significantly inhibited (-->50%) both kinesin and dynein ATPase activities (Table 1). In addition, glycoursodeoxycholate (GUDC) also inhibited dynein ATPase activity,

120~

lod

~ ~

i

~~TUDC

60 4~

TCDC

2( 0' 0.0

012

Bileacid[raM] 0'.4

0~6

0:8

110

Figure 5. Dose-dependent inhibition of kinesin ATPase activity by TCDC. Purified rat liver kinesin (--1 pg) was incubated for 30 minutes at 37°C with 0.8 mg/mL Taxol-stabilized microtubules and 0.75 mmol/L Mg2+-ATP in the presence or absence of bile acids. Reactions were stopped with SDS, and ATPase activity was measured by colorimetric quantitation of inorganic phosphate released. Results are expressed as percent of control values (ATPase activity in the absence of bile acids). Values are means _+ SE. *Values significantly different (P < 0.05) from controls in two4ailed t tests.

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GASTROENTEROLOGY Vol. 108, No. 3

Table 1. C o m p a r a t i v e Effects o f Bile Acids on Kinesin, C y t o p l a s m i c Dynein, and A p y r a s e ATPase Activities ATPase activity (% o f controls) a

Bile acid (1 mmol/L)

Kinesin b

Dynein °

Apyrase ~

40.4 + 9.9 e 77.5 + 10.5

50.5 _+ 1.4 e 106.1 + 2.5

91.8 + 5.5

32.7 + 5.1 e 104.7 _+ 10.0

21.8 _+ 1.7 e 88.9 + 2.1

87.1 _+ 3.8

mmol/L mmol/L

75.8 + 10.6 100.1 + 6.8

60.7 + 8.0 e 104.8 + 0.8

mmol/L mmol/L

93.6 + 6.9 108.1 + 11.2

91.9 + 7.5 109.8 + 2.5

92.6 + 4.8

88.4 _+ 8.8 87.2 + 1.4

109.9 _+ 2.2 109.0 + 1.8

96.5 + 4.7

91.0 + 8,1 90.2 + 1.1

113.6 + 1.8 107.6 + 2.8

105.3 + 6.9

GCDC 1.0 mmol/L 0.2 mmol/L

TCDC 1.0 mmol/L 0.2 mmol/L

GUDC 1.0 0.2 TUDC 1.0 0.2

90.9 -- 3.5 --

Glycocholate 1.0 mmol/L 0.2 mmol/L Taurocholate 1.0 mmol/L 0.2 mmol/L

NOTE. Values are means _+ SE for three or more experiments.

aFor each protein, results were expressed as a percent of controls (the same protein tested in the absence of bile acids). ~Kinesin activity was measured in 15 mmol/L imidazole and 2 mmol/L MgCI2 (pH 7.0) with 0.8 mg/mL Taxol-polymerized microtubules. CDynein activity was measured in 15 mmol/L imidazole and 2 mmol/L MgCI2 (pH 8.0) without microtubules. dApyrase activity was measured in 15 mmol/L imidazole and 4 mmol/L CaCI2 (pH 6.5) without microtubules. "Values significantly different (P < 0.05) from controls in two-tailed t tests.

although to a lesser extent (~40%). In contrast, TUDC, glycocholate, and taurocholate had no significant effect on kinesin or dynein activity (Table 1). Selective inhibition of dynein activity by chenodeoxycholate conjugates suggest that microtubules need not be present for bile acids to inhibit the activity of microtubule motor proteins. The unrelated ATPase, apyrase was not significantly affected by any of the six bile acids tested (Table 1). This result suggests that the inhibition by bile acids of kinesin and dynein activities is not caused by a general effect on all ATPases.

Discussion In this study, we purified the microtubule-based motor kinesin from rat liver and tested the effects of bile acids on the enzymatic activities of kinesin. Purified kinesin was shown to be similar to kinesins from other tissues by several biochemical and immunological criteria and was proven to be biologically active. We found that kinesin-based motility was significantly inhibited by

TCDC but not by TUDC. The binding of kinesin to microtubules in vitro was unaffected by bile acids; however, kinesin ATPase activity was significantly inhibited by TCDC and GCDC but not by TUDC, GUDC, glycocholate, or taurocholate. Most interestingly, TCDC and GCDC also inhibited the ATPase activity of cytoplasmic dynein, another microtubule motor, but had no effect on the ATPase of the nonmotor ATPase, apyrase. Our results show that high concentrations of bile acids, such as those present in cholestasis, could directly inhibit the ATPase activities of microtubule motors and suggest a possible mechanism for the impairment of vesicular transport in cholestasis. Although kinesin has been previously reported to be present in liver, to our knowledge, this is the first report in which liver kinesin has been purified and characterized. Liver kinesin was shown to have heavy (N 120 kilodaltons) and light (~60 kilodaltons) chains typical of kinesin from other sources13'42'43; however, unlike brain kinesin, which contains heavy-chain isoforms of several different molecular weights (116-130 kilodaltons), liver kinesin appears to contain only a single heavy-chain isoform (Figure 2). The amino acid sequences of tryptic fragments of liver kinesin heavy chain were highly homologous to regions present in published sequences of human (Figure 1) and mouse kinesin heavy chain (data not shown), as expected, because several domains of the kinesin heavy chain are highly conserved between species. Rat liver kinesin was found to be biologically active in terms of its ability to move axonemes in vitro and its microtubule-stimulated ATPase activity. Both the motility velocity (0.5 ~m/s) and the microtubule-stimulated ATPase activity (300-700 btmol" min -1" mg -1) of rat liver kinesin were in the ranges reported for other kinesins. 36'40'42'44'45'51 Thus, our results show that liver kinesin is similar to kinesins from other tissues and suggest that liver kinesin can probably support vesicle motility in the hepatocyte. The concentrations of bile acids in serum in severe clinical and experimental cholestasis have been reported to be as high as 0.5-1.0 mmol/L. 52'53 Concentrations within the hepatocyte during cholestasis may be even higher; however, it is problematic to obtain estimates of the intracellular concentrations of bile acids in hepatocytes during cholestasis, because any measurements entail disruption of the liver and possible mixing of cytosol with bile. One report suggests that isolated hepatocytes can accumulate levels of bile acids as high as 1 mmol/ L. 54 Thus, our findings that only relatively high concentrations of bile acids ( ~ 0.5-1.0 mmol/L) inhibit kinesin motility or ATPase activity suggest that bile acids may

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inhibit kinesin activity when they are in the high pathophysiological range but not at lower concentrations of bile acids such as are normally present in the hepatocyte. It is also probable that other factors, such as the presence of multiple bile acids, bile acid-binding proteins, and other cholephilic ions, can influence the inhibitory effects of bile acids on microtubule motor activity. That the cholestatic bile acids, TCDC and GCDC, selectively inhibit both kinesin and dynein ATPase activities suggests a common inhibitory mechanism. It is doubtful that chenodeoxycholate conjugates inhibit motor activity by interfering with binding to microtubules because (1) kinesin was able to bind normally to microtubules in vitro in the presence of inhibitory concentrations of TCDC and (2) dynein ATPase activity was inhibited by chenodeoxycholate conjugates in the absence of microtubules. The inhibition of kinesin ATPase activity by bile acids at a dose of 1 mmol/L was not significantly correlated with bile acid hydrophobicity or critical micellar concentration (P > 0.05; data not shown). These results are in contrast with other studies that related hydrophobicity of bile acids to certain toxic effects.46'5%56 It may be significant that both dynein and kinesin are Mg2+-ATPases, whereas apyrase, which was not affected by bile acids, is a Ca2+-ATPase. It has been previously reported that TCDC was more inhibitory than taurocholate at concentrations of >--1 mmol/L to Mg2+-ATPase activity present in isolated rat plasma membranes. 3°'31 Similarly, GCDC at a dose of 1 mmol/L but not glycocholate or taurocholate were reported to inhibit the activities of Mg 2+- and Na+/K+-ATPases in membranes isolated from intestinal mucosa. 32 Thus, selective inhibition by chenodeoxycholate conjugates may be characteristic of certain (e.g., Mg 2+- and Na+/K +-) but not all (e.g., Ca 2+-) ATPases. In conclusion, our results suggest the possibility that high concentrations of certain bile acids such as those present in cholestasis inhibit vesicular transport in the hepatocyte via a direct effect on the activities of microtubule motors. W e have recently shown that kinesin is localized to the Golgi apparatus and transcytotic carriers in hepatocytes. .6 Based on these observation, we predict that cholestatic bile acids interfere with both plasma protein secretion and transcytosis via direct inhibition of kinesin. Direct effects of bile acids may not be the only mechanism by which bile acids affect microtubule motors. It has been shown that kinesin activity can be modulated both by binding to calmodulin and by phosphorylation with 5'-cyclic adenosine monophosphate (cAMP)-dependent protein kinase. 5v Relatively low concentrations (--<100 ~tmol/

INHIBITION OF LIVER KINESIN BY BILE ACIDS

831

L) of bile acids are reported to stimulate increases of intracellular Ca 2+ content in hepatocytes. 26'58'59 Thus, one consequence of treatment of hepatocytes with bile acids may be activation of calmodulin by Ca 2+ and subsequent regulation of kinesin activity by calmodulin binding. An interaction of bile acids with the cAMP signaling pathway has not been shown. However, the transport of transcytotic vesicles toward the bile canaliculus, a process that is stimulated by bile acids, 25'27'6° is also stimulated by cAMP and its analogues. 61 Additional studies are required to determine if bile acids can indirectly regulate the activity of kinesin molecules involved in transport of apically directed vesicles by activation of intracellular secondmessenger cascades.

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Received July 7, 1994. Accepted October 17, 1994. Address requests for reprints to: Mark A. McNiven, Ph.D., Center for Basic Research in Digestive Diseases, Mayo Clinic, Rochester, Minnesota 55905. Fax: (507) 284-0762. Supported by a Thompson-Mayo fellowship (to D.L.M.) and National Institutes of Health grants DK44650 and AA09227-02 (to M.A.M.) and DK24031 (to N.F.L.). The authors thank Tushar Patel for helpful comments and the Mayo Clinic Peptide Core Facility, Rochester, Minnesota, for partial amino acid sequencing of liver kinesin heavy chain.