Potential conflict of interest: Nothing to report.
This work was supported by the National Institute of Alcohol Abuse and Alcoholism grants R01 AA17626 and R21 AA15683 (awarded to P.L.T.).
The liver is the major site of ethanol metabolism and thus sustains the most injury from chronic alcohol consumption. Ethanol metabolism by the hepatocyte leads to the generation of reactive metabolites and oxygen radicals that can readily adduct DNA, lipids, and proteins. More recently, it has become apparent that ethanol consumption also leads to increased post-translational modifications of the natural repertoire, including lysine hyperacetylation. Previously, we determined that alcohol consumption selectively impairs clathrin-mediated internalization in polarized hepatocytes. However, neither the step at which the block occurs nor the mechanism responsible for the defect have been identified. To identify the specific step at which clathrin-mediated internalization is impaired, we examined the distributions, levels, and assembly of selected components of the clathrin machinery in control and ethanol-treated cells. To determine whether the impairment is caused by ethanol-induced lysine acetylation, we also examined the same coat components in cells treated with trichostatin A (TSA), a deacetylase inhibitor that leads to protein hyperacetylation in the absence of ethanol. Conclusion: We determined that both ethanol and TSA impair internalization at a late stage before vesicle fission. We further determined that this defect is likely the result of decreased dynamin recruitment to the necks of clathrin-coated invaginations resulting in impaired vesicle budding. These results also raise the exciting possibility that agents that promote lysine deacetylation may be effective therapeutics for the treatment of alcoholic liver disease. (Hepatology 2012)
The liver is the major site of ethanol metabolism and thus sustains the most injury from chronic alcohol consumption. Alcohol is metabolized by alcohol dehydrogenase (ADH) and cytochrome P450 2E1 (CYP2E1). ADH-mediated metabolism results in the production of acetaldehyde, a highly reactive intermediate that can form covalent modifications on lipids, DNA, and proteins, including tubulin, actin, calmodulin, and many lysine-dependent enzymes.1-3 CYP2E1-mediated metabolism not only produces acetaldehyde, but also highly reactive oxygen and hydroxyethyl radicals and other lipid-derived reactive intermediates.1 Like acetaldehyde, all of these CYP2E1-generated by-products can form covalent modifications on various macromolecules.1 More recently, it has been shown that alcohol exposure induces protein covalent modifications that are part of the natural repertoire, including increased methylation, phosphorylation, and acetylation.4 In particular, numerous proteins have been identified that are lysine hyperacetylated upon ethanol exposure. Recently, we identified over 40 non-nuclear proteins that are hyperacetylated in livers from ethanol-fed rats and/or in WIF-B cells.5, 6 Among these are cortactin, tubulin, and actin (the latter two of which are known to be acetaldehyde adducted). Thus, one hypothesis for alcohol-induced hepatotoxicity is that the accumulated covalent modifications during chronic alcohol consumption lead to hepatic dysfunction and liver injury.
For years, it has been appreciated that chronic alcohol consumption impairs protein trafficking,7-9 and more recently, efforts have been aimed at understanding how protein adduction and acetylation may contribute to those defects. In general, two trafficking pathways are affected: secretion and receptor-mediated endocytosis. We have further shown that alcohol consumption selectively impairs clathrin-mediated internalization.10 These and our other studies were performed in polarized, hepatic WIF-B cells. Importantly, WIF-B cells efficiently metabolize ethanol using endogenous ADH and CYP2E1 and produce the many reactive intermediates and oxygen radicals described above.11, 12 We have also determined that WIF-B cells display the same alcohol-induced defects in protein trafficking (including impaired clathrin-mediated internalization) and acetylation as described in situ.
Although there are numerous proteins that participate in clathrin vesicle formation,13, 14 there are three core components: clathrin, adaptor protein 2 (AP2), and the guanosine triphosphate (GTP)ase, dynamin. In general, clathrin triskelions are recruited to and assembled at regions of the plasma membrane enriched in phosphatidylinositol 4,5-bisphosphate. AP2 is targeted to these regions and interacts directly with sorting signals on internalized proteins. Dynamin is then recruited to and assembled on the necks of coated pits and, upon coordinated GTP hydrolysis, promotes vesicle fission. The released vesicles are rapidly uncoated, allowing for coat recycling and vesicle fusion.
The studies described here were aimed at identifying the specific step at which ethanol exposure impairs clathrin-mediated internalization and thus the potential mechanism(s) responsible for that impairment. We examined the protein expression, distributions, and assembly of the three core components of the clathrin machinery. Because both actin and cortactin are hyperacetylated upon alcohol exposure and participate in vesicle fusion, we also examined their distributions.5 The distribution and assembly of clathrin-coated vesicles was compared to that of asialoglycoprotein receptor (ASGP-R), whose clathrin-mediated internalization is impaired by ethanol exposure.15, 16 To determine whether ethanol-induced protein acetylation could explain the internalization defect, we also examined cells treated with trichostatin A (TSA), a pan-deacetylase inhibitor.
5′NT, 5′ nucleotidase; ADH, alcohol dehydrogenase; AP2, adaptor protein 2; ASGP-R, asialoglycoprotein receptor; CCD, charge-coupled device; CHC, clathrin heavy chain; CYP2E1, cytochrome P450 2E1; GTP, guanosine triphosphate; HDAC6, histone deacetylase-6; HEPES, N-2-hydroxylethylpiperazine-N′-ethanesulfonic acid; IgA, immunoglobulin A; mAbs, monoclonal antibodies; PBS, phosphate-buffered saline; PFA, paraformaldehyde; pIgA-R, polymeric IgA receptor; ROI, regions of interest; RT, room temperature; TEM, transmission electron microscopy; TIRF, total internal reflection fluorescence; TSA, trichostatin A.
Materials and Methods
Reagents and Antibodies.
F12 (Coon's) medium, TSA, and horseradish-peroxidase–conjugated secondary antibodies were from Sigma-Aldrich (St. Louis, MO). Fetal bovine serum was from Gemini Bio-Products (Woodland, CA), and N-2-hydroxylethylpiperazine-N′-ethanesulfonic acid (HEPES) was from HyClone (Logan, Utah). Cy3, Alexa Fluor 488– and Alexa Fluor 568–conjugated secondary antibodies were purchased from Invitrogen (Carlsbad, CA), and the Texas Red–conjugated secondaries were from Jackson ImmunoResearch (West Grove, PA). The clathrin heavy-chain (CHC) (X22) and AP2 (AP6) monoclonal antibodies (mAbs) were from Novus Biologicals (Littleton, CO), and the dynamin-2 mAbs were from BD Biosciences (San Jose, CA). The monoclonal actin and cortactin antibodies were from Abcam (Cambridge, MA) and Millipore (Billerica, MA), respectively. The polyclonal ASGP-R antibodies and the monoclonal polymeric immunoglobulin A (IgA)-receptor (pIgA-R), CE9, and 5′nucleotidase (5′NT) antibodies were provided by Dr. A. Hubbard (Johns Hopkins University School of Medicine, Baltimore, MD).
WIF-B Cell Culture.
Cells were grown as previously described.17 On day 7, cells were treated with 50 mM of ethanol buffered with 10 mM of HEPES (pH 7.0) at 37°C for 72 hours, as previously described.12
Recombinant adenovirus encoding pIgA-R was provided by Dr. A. Hubbard. The dynamin wild-type and K44A dominant negative recombinant adenoviruses were provided by Drs. S. Schmid and H. Damke (Scripps, La Jolla, CA). After 48 hours of ethanol exposure, cells were infected for 1 hour at 37°C, as previously described.18 Cells were washed with complete medium and incubated for an additional 18-20 hours in the continued absence or presence of ethanol to allow protein expression. Then, 50 nM of TSA was added during the last 30 minutes of virus expression.
Western Blotting and Immunoprecipitations.
Immunoprecipitations were performed as previously described.20 In general, cells were lysed in 1% nonyl phenoxypolyethoxyethanol, 150 mM of NaCl, 50 mM of Tris, and 1 mM of ethylene diamine tetraacetic acid (pH 7.5) on ice for 30 minutes and cleared by centrifugation at 120,000×g for 30 minutes at 4°C. Antidynamin antibodies (0.5-1 μg) were added and recovered with Protein G agarose (Thermo Fisher Scientific Inc., Waltham, MA). The precipitated fractions were resuspended in Laemmli sample buffer and boiled for 3 minutes. Samples were immunoblotted with antibodies specific to AP2 (1:1,000), CHC (1:2,000), cortactin (1:2,500), actin (1:2,500), or dynamin (1:2,500). Immunoreactivity was detected using enhanced chemiluminescence (PerkinElmer, Crofton, MD).
Cells were fixed on ice with 4% paraformaldehyde/phosphate-buffered saline (PFA/PBS) for 1 minute and permeabilized with ice-cold methanol for 10 minutes. Cells were processed for indirect immunofluorescence, as previously described,21 using antibodies against ASGP-R (1:1,000), pIgA-R (1:200), AP2 (1:100), or CHC (1:1,000). Fluorophore-conjugated secondary antibodies were used at 5 μg/mL. To label cortactin (1:100), cells were permeabilized with PEM (100 mM of PIPES, 1 mM of ethylene glycol tetraacetic acid, 1 mM of MgCl2; pH 6.8), containing 0.1% saponin and 8% sucrose for 2 minutes and fixed at room temperature (RT) with 4% PFA/PBS for 30 minutes. To visualize membrane-associated dynamin (1:100), cells were permeabilized with 0.1% Triton X-100/ PEM/sucrose for 2 minutes at RT and fixed in methanol for 5 minutes at −20°C. Epifluorescence was visualized using an Olympus BX60 Microscope (Opelco, Inc., Dulles, VA). Images were collected using a Coolsnap HQ2 digital camera (Photometrics, Tucson, AZ) and IPLabs image analysis software (BioVision Technologies, Inc., Chester Springs, PA). Confocal microscopy was performed on a Zeiss Axiovert 200 inverted microscope with a 510 meta laser scanning confocal module (Carl Zeiss GmbH, Oberkochen, Germany).
To quantitate relative AP2 membrane staining, random fields were visualized by epifluorescence and digitized. From micrographs, membrane fluorescence was traced using the segmented line tool, and intracellular staining regions of interest (ROI) were measured using the ImageJ Measure ROI tool. The averaged background pixel intensity was subtracted from both the averaged membrane and intracellular intensities, and the ratio of basolateral versus intracellular fluorescence intensity was determined.
K+ Depletion/Repletion Assays.
The K+ depletion/repletion assays were performed as previously described.22 For ASGP-R antibody trafficking studies, K+-depleted cells were surface labeled with anti-ASGP-R antibodies (1:25) for 20 minutes on ice. Cells were reincubated in prewarmed medium supplemented with 10 mM of KCl, and the ASGP-R antibody-antigen complexes were allowed to traffic for desired times at 37°C. Cells were fixed, permeabilized, and the trafficked ASGP-R antibodies were labeled with secondary antibodies.
Total Internal Reflectance Fluorescence and Transmission Electron Microscopy.
Cells were stained as described above and mounted in Tris-buffered saline (pH 10.5) containing 5% glycerol and 4 mg/mL of phenylenediamine. Fluorophores were excited with a 2.5-W Kr/Ar laser (Spectra Physics, Irvine, CA) and visualized using an Olympus 1X 71 inverted microscope and total internal reflection fluorescence (TIRF) illuminator (Olympus, Center Valley, PA). Images were collected using a Photometrics Evolve EM-CCD (charge-coupled device) camera (Photometrics, Tuscon, AZ) and Metamorph software (Molecular Devices, Sunnyvale, CA). Puncta were counted using the FociPicker three-dimensional ImageJ plugin. Fully covered 10,000 px2 ROIs were selected from random images. In general, five images/experiment were acquired and two to five fields/image were counted. For transmission electron microscopy (TEM), cells were fixed and processed using standard Epon embedding techniques. Ultrathin sections were cut and stained with uranyl acetate, followed by lead citrate. Grids were viewed on a Hitachi 7600 transmission electron microscope (Hitachi, Tokyo, Japan), and images were captured with an AMT CCD camera (Advanced Microscopy Techniques, Woburn, MA).
The Core Clathrin Machinery Redistributes in Ethanol-Treated WIF-B Cells.
We previously determined that ethanol exposure led to the dramatic redistribution of ASGP-R from intracellular endosomes to the basolateral membrane in WIF-B cells.15 Closer examination using confocal microscopy revealed that the membrane-associated ASGP-R in ethanol-treated cells was present in discrete puncta (Fig. 1A). Because these puncta resembled clathrin-coated pits visualized at the light level, we examined the distributions of core clathrin-coat proteins. In control cells, CHC localized primarily to an intracellular compartment (Fig. 1A). As observed for ASGP-R, CHC redistributed to the basolateral membrane in discrete puncta in ethanol-treated cells. Although AP2 primarily localized to the basolateral membrane in control cells (Fig. 1A), there was a 19% ± 4% increase (P ≤ 0.02) of the adaptor at the membrane in ethanol-treated cells with a reciprocal decrease in diffuse cytosolic staining. Similarly, increased basolateral cortactin-positive puncta were observed in ethanol-treated cells (Fig. 1A).
Because of its large, soluble pool, we permeabilized cells with Triton X-100 before fixation to detect membrane-associated dynamin. In control cells, dynamin was detected at the basolateral membrane (Fig. 1A). However, virtually no dynamin was observed at the basolateral surface in ethanol-treated cells. Coimmunoprecipitations confirmed these results. In control cells, both CHC and cortactin coimmunoprecipitated with dynamin, indicating interactions among these proteins (Fig. 1C). In contrast, the coprecipitated levels of CHC and cortactin were decreased after ethanol exposure, reflecting decreased interactions. To further confirm that decreased interactions were not the result of decreased expression levels, we immunoblotted cell lysates for coat components. No changes in levels of dynamin, CHC, AP2, cortactin, or actin were observed (Fig. 1D), ruling out this possibility. Together, these results suggest that the clathrin-coated structures are late-stage invaginations unable to bud from the membrane because of impaired dynamin recruitment.
To test whether these altered distributions required ethanol metabolism, we treated cells with the ADH inhibitor, 4-methyl pyrazole. 4-methyl pyrazole prevented CHC and dynamin redistribution, indicating that the defect was likely mediated by acetaldehyde (Supporting Fig. 1).
Protein Hyperacetylation May Explain the Ethanol-Induced Internalization Defect.
Previously, we determined that ASGP-R internalization is impaired by treatment with TSA, a pan-deacetylase inhibitor.15 To determine whether TSA also induces the redistribution of ASGP-R and the clathrin machinery, we immunostained control and cells treated for 30 minutes with 50 nM of TSA at 37°C, conditions that hyperacetylate proteins to the same extent as ethanol.15 As for ethanol-treated cells, TSA addition led to the redistribution of ASGP-R, CHC, AP2 (38% ± 17% increase) and cortactin to the basolateral membrane in discrete puncta (Fig. 1B). Also, as for ethanol-treated cells, virtually no membrane-associated dynamin was observed in TSA-treated cells (Fig. 1B). This suggests that not only are these structures late-stage intermediates, but also that hyperacetylation may explain the internalization defect.
The Clathrin-Coated Structures Are Continuous With the Basolateral Membrane and Recruit Receptors.
If the structures are late-stage intermediates, the prediction is that they are continuous with the plasma membrane. To test this prediction, we used TIRF microscopy to visualize the bottommost 100 nm of the cell, the approximate diameter of a clathrin-coated pit. In control cells, few discrete ASGP-R-positive puncta were observed at the cell surface (Fig. 2A). Additional profiles were also detected, albeit smaller and dimmer, likely representing budding vesicles or receptors not clustered into pits. In ethanol-treated cells, increased ASGP-R-positive puncta were observed that were more uniform in size (Fig. 2A). The basolateral membrane also had decreased diffuse staining, suggesting fewer unclustered receptors.
Quantitation of the puncta confirmed these observations (Table 1). Ethanol exposure led to a 1.3-fold increase in membrane-associated ASGP-R puncta. A similar increase was observed with TSA (1.2-fold), although this was not statistically significant. To confirm that the effect was shared by other receptors internalized via clathrin-coated vesicles, we examined the transcytosing receptor, pIgA-R. As for ASGP-R, the number of discrete pIgA-R-positive puncta was increased in alcohol-exposed cells (Fig. 2A) by 1.4-fold (Table 1).
Table 1. ASGP-R, pIgA-R, and CHC Accumulate at the Plasma Membrane in Ethanol and TSA-Treated Cells
Cell-surface puncta positive for the indicated proteins were imaged using TIRF microscopy. To quantitate the profiles, ROI with areas of 10,000 px2 were selected and counted using the FociPicker 3D ImageJ plugin with the following settings: tolerance = 10, minimum pixels in focus = 1, and scale of image in X, Y, and Z planes = 10 pixels/μm. In general, two to five fields were selected per image, with five images acquired per experiment. Values for ASGP-R are expressed as averages ± standard error of the mean determined from three (EtOH) or four (TSA) independent experiments. Values for pIgA-R, CHC (EtOH), and AP2 are expressed as averages determined from two independent experiments. Values for CHC (TSA) are expressed as averages from four independent experiments. Values for 5′NT and CE9 were determined from n = 1.
The total number of CHC-positive puncta was also increased in ethanol-treated cells (Fig. 2B; Table 1). These profiles were also larger and tended to form aggregates. Interestingly, the number and size of AP2-positive puncta did not change in treated cells (Fig. 2B; Table 1). At present, we cannot reconcile these disparate observations, but they may represent differences in their assembly kinetics into coats. For comparison, we examined two non-clathrin-associated proteins: 5′NT (glycosylphosphatidylinositol-anchored transcytosing protein) and CE9 (basolateral resident). In control cells, both 5′NT and CE9 were detected in abundant, small puncta with a diffuse background (Fig. 2C). The distributions (Fig. 2C) or amounts of CE9 or 5′NT-positive puncta (Table 1) did not change in ethanol-treated cells, confirming the selectivity of the defect.
To determine whether receptors were properly recruited to clathrin-coated pits in treated cells, we examined the degree of colocalization between ASGP-R and CHC. In both ethanol- and TSA-treated cells, ASGP-R and CHC partially colocalized (r = 0.38) (Fig. 2D). Together, these results indicate that the puncta are continuous with the plasma membrane, and that internalization is impaired after receptor recruitment and pit assembly.
Impaired ASGP-R Internalization Correlates With Impaired Dynamin Membrane Binding.
To examine the kinetics of ASGP-R vesicle recruitment and internalization, we synchronized endocytosis by the sequential disassembly/reassembly of clathrin lattices by K+ depletion/repletion, as previously described.22 After clathrin lattices were disrupted by K+ depletion for 30 minutes, live cells were surface labeled with antibodies specific to external ASGP-R epitopes to detect only the membrane-associated receptors. Cells were then incubated with K+-containing medium for up to 15 minutes to allow rapid coated pit assembly and internalization. After K+ depletion and surface labeling (0 minutes), ASGP-R in control and treated cells was detected only at the basolateral membrane (Fig. 3A,B). ASGP-R membrane distribution was more uniform than at steady state, consistent with clathrin lattice disassembly preventing receptor clustering. In control cells after 5 minutes of repletion, ASGP-R accumulated at or near the plasma membrane in large, discrete puncta (Fig. 3A,B). After 15 minutes, there was a significant decrease in surface labeling with a reciprocal increase in intracellular labeling, indicating successful, rapid ASGP-R internalization. In contrast, after 5 minutes of repletion in treated cells, there were fewer large puncta, indicating delayed receptor clustering and/or clathrin vesicle assembly (Fig. 3A,B). This delay persisted for 15 minutes with fewer ASGP-R-positive puncta and little intracellular staining, consistent with a block in late internalization.
To visualize the kinetics of only those receptors at the cell surface, we examined K+-repleted cells using TIRF. In control cells, ASGP-R was detected in discrete structures at the cell surface after 0 minutes of repletion (Fig. 4A). After 5 minutes, there was a noticeable decrease in receptor labeling, and the remaining puncta were dimmer and smaller. By 15 minutes, little surface ASGP-R was detected, indicating its rapid, successful internalization. Consistent with the confocal images, ethanol exposure led to increased ASGP-R-positive puncta at 0 minutes (Fig. 4A). After 5 minutes, most of the ASGP-R-positive structures remained at the surface and were brighter and larger than control. Although at 15 minutes there were decreased levels of puncta, there was significantly more ASGP-R remaining in ethanol-treated cells than in control, indicating impaired internalization. To quantitate internalization, the ASGP-R-positive puncta were counted at each time point postrepletion and were plotted as the percent of total surface-labeled puncta at time 0. In control cells, the number of ASGP-R-positive puncta steadily decreased after K+ repletion, and by 15 minutes, only 34% of the puncta remained (Fig. 4B). In contrast, the number of puncta remained relatively constant during the first 5 minutes of repletion in ethanol-treated cells, and by 15 minutes, over 60% of ASGP-R-positive puncta remained (Fig. 4B).
To directly test whether impaired dynamin membrane recruitment could explain the internalization defect, we monitored dynamin distributions after K+ depletion/repletion in control and treated cells. Consistent with the disassembly of clathrin lattices, there was little dynamin detected at the basolateral membrane after K+ depletion in control and treated cells (Fig. 5A). In control cells, both after 5 and 15 minutes of repletion, there was a significant increase in membrane dynamin staining, indicating its recruitment to the necks of coated pits. In contrast, in treated cells, much less dynamin membrane staining was observed at all time points. Together, these results suggest that dynamin is not properly recruited to coated pits, thereby preventing vesicle fission.
Internalization Is Blocked at a Late Stage of Vesicle Assembly.
To directly confirm a block in late-stage vesicle budding, we visualized clathrin-coated profiles by TEM. Images of clathrin-coated profiles were acquired as encountered and grouped into three classes. Class 1 profiles represent early-stage clathrin structures. The electron-dense clathrin coat was visible, but there was little invagination. Class 2 profiles represent intermediate structures characterized by increased invagination, but no neck constriction. Finally, class 3 profiles represent highly invaginated coated pits with constricted necks. In control cells, the profiles distributed evenly among the three categories, with 30%-36% identified as class 3 (Fig. 6A). In contrast, in both ethanol- and TSA-treated cells, 56% of profiles were class 3, with a reciprocal loss in class 1 (Fig. 6A). Examples of the late-stage coated pits in ethanol (Fig. 6B) and TSA-treated cells (Fig. 6C) are shown. They are deeply invaginated, with partially constricted necks. Unlike the collared, coated invaginations observed in cells expressing GTPase-deficient dynamin, the profiles in treated cells had no “collars” and the necks were not directly apposed, as described below.23, 24
Dynamin Membrane Recruitment Is Impaired.
If dynamin membrane recruitment is impaired in treated cells, another prediction is that dynamin overexpression would not rescue the defect. To test this hypothesis, we overexpresseed wild-type dynamin in control and ethanol-treated cells. As for the endogenous protein, wild-type dynamin was detected at the plasma membrane (albeit in lesser amounts than endogenous) in control cells (Fig. 7A). As predicted, overexpression failed to rescue the ethanol-induced internalization defect. Little to no membrane-associated wild-type dynamin was detected, and ASGP-R redistributed to the basolateral surface (Fig. 7A) in ethanol-treated cells. For comparison, we examined the distribution of dominant negative K44A dynamin. Significantly more of the mutant dynamin was detected at the cell surface than wild type (Fig. 7B). This result is consistent with the findings that this mutant can oligomerize at the necks of invaginated pits, but cannot undergo the conformational change required for vesicle fission.23
Here, we report that ethanol exposure blocks the internalization of clathrin-coated pits at a late stage of assembly by impairing dynamin recruitment to the necks of invaginated pits, thereby preventing vesicle fission. Treatment with TSA led to remarkably similar alterations in vesicle assembly and dynamin recruitment, such that we conclude that protein hyperacetylation may explain, in part, the ethanol-induced defect in clathrin-mediated internalization.
Dynamin Membrane Recruitment, but Not GTPase Activity, Is Impaired.
Because dynamin self-assembly at the necks of coated pits promotes its GTPase activity leading to vesicle budding,25 it is tempting to speculate that ethanol (or TSA) exposure impairs dynamin activity. However, our ultrastructural analysis indicated that this is likely not the case. Expression of dominant negative dynamin or treatment with dynamin GTPase inhibitors (e.g., dynasore) lead to the formation of highly invaginated pits with elongated, highly constricted necks with apposed membranes.23, 24 In some cases, the dynamin oligomers are readily visible, wrapping around these elongated necks. Similarly, in cells of the temperature-sensitive shibire flies, “collars” of defective dynamin oligomers were observed on the necks of invaginated pits at the restrictive temperature.26, 27 However, in ethanol- or TSA-treated cells, no such collars were detected. Although the class 3 profiles were invaginated, the necks were not elongated nor were the sides of the necks apposed, indicating that dynamin oligomers were likely not assembling there. Furthermore, overexpressed wild-type dynamin failed to rescue the ethanol-induced defect and was not detected at the plasma membrane, indicating impaired dynamin membrane recruitment.
Possible Mechanisms for Impaired Dynamin Recruitment.
Previously, we determined that impaired clathrin-mediated internalization required ethanol metabolism and was likely mediated by acetaldehyde (see Supporting Fig. 1).15, 28 Thus, one exciting possibility is that a critical clathrin-coat component(s) is prone to adduction by acetaldehyde or other reactive metabolites, thereby impairing proper dynamin recruitment. Alternatively, (additionally?), hyperacetylation of key coat components may be at fault. This hypothesis is supported by the findings that actin and cortactin are hyperacetylated upon ethanol exposure.4 Although how cortactin, actin, and dynamin function to promote vesicle release is not completely elucidated, acetylation of cortactin is known to prevent its association with actin.29 Thus, we propose that alcohol-induced hyperacetylation leads to decreased interactions between actin, cortactin, and/or dynamin, thereby inhibiting dynamin recruitment and subsequent vesicle fission. Although our coimmunoprecipitation results are fully consistent with this hypothesis, identification of the hyperacetylated lysines in both actin and cortactin (and dynamin?) is needed to test this hypothesis.
Previously, we found that ethanol exposure led to increased microtubule acetylation and stability.6 In an effort to determine the mechanism responsible for this observation, we examined the distributions and biochemical properties of histone deacetylase-6 (HDAC6), a tubulin (and cortactin) deacetylase. We found that HDAC6 binding to endogenous microtubules was impaired in ethanol-treated cells, whereas its ability to bind or deacetylate exogenous tubulin did not change, suggesting that tubulin from ethanol-treated cells was modified, thereby preventing HDAC6 binding.30 Because both impaired HDAC6 microtubule binding and tubulin hyperacetylation require ethanol metabolism and are likely mediated by acetaldehyde6, 30 and because tubulin can be acetaldehyde adducted,31, 32 we propose that tubulin-acetaldehyde adducts impede HDAC6-tubulin binding, thereby preventing deacetylation. It is possible that an analogous scenario may explain impaired clathrin-mediated internalization in ethanol-treated cells, a possibility we are currently exploring.
Potential Therapeutics for Alcoholic Liver Disease.
With the expanding list of ethanol-induced hyperacetylated proteins and a strong correlation between hyperacetylation and defects in hepatic protein trafficking, lysine deacetylation may provide a novel therapeutic target for the treatment of alcoholic liver disease. Currently, many naturally occurring and synthetic deacetylase agonists (e.g., resveratrol and SRT-501) are in clinical trials for treatment of a host of human diseases.33 Furthermore, resveratrol has been shown to attenuate fatty liver and oxidative stress in alcohol-exposed mice.34 An exciting possibility is that specific deacetylase activators or acetyltransferase inhibitors will be useful in treating alcoholic liver disease.
The authors thank Dr. Scot Kuo, Mike Delannoy, and Barbara Smith (Johns Hopkins School of Medicine Microscope Facility) for assistance with TEM and instrument training. The authors also thank Dr. Ann Hubbard (Johns Hopkins School of Medicine) for providing lab space for some of the studies and for providing the many antibodies and viruses used in these studies.