Ubiquitin-Proteasome Pathway
in the Pathogenesis of Liver Disease
Samuel W. French, Fawzia Bardag-Gorce
32
32.1
Introduction
The discovery of the ubiquitin-proteasome pathway as a regulated system of protein digestion within all cells [32] has led to an appreciation of the impor- tance of protein turnover control. This control is a mechanism for regulation of cellular processes and quality control of intracellular proteins. Many liver cell functions are regulated by this mechanism of protein degradation. These include cell cycle check points and activation of transcription factors such as nuclear factor- κB (NF-κB) (Chapter 29), and hy- poxia inducible factor-1 α (HIF-1α) (Chapter 26) [28, 67, 71]. The loss of proteasomes or the inhibi- tion of the ubiquitin-proteasome pathway can lead to hepatocellular injury including proliferation and apoptosis [74, 107], and hepatic inclusions of ag- gregated cytokeratins [24]. Liver cell gene expres- sions, dependent on transcription factor activation by the proteasome, could impede the inflammatory response of the liver and the response to hypoxic in- jury. The importance of the ubiquitin-proteasome pathway to the homeostatic mechanisms involved in liver injury is the focus of this review.
32.2
Ubiquitin-Proteasome Pathway
The enzymes that catalyze ubiquitin activation, conjugation and ligation are depicted as E1, E2, and E3, respectively in Fig. 32.1. Ubiquitin is indicated as a shaded circle. Note that the polyubiquitinated protein indicated by Cn+1 docks at the 19S protea- some. Alpha ( α) units of the 20S proteasome par- ticle are shown in gray, while the beta ( β) subunits are shown in white. Both types of subunits make up the 20S catalytic core of the proteasome. The arrows shown at the 19S proteasome indicate that the ubiq- uitinated protein is deubiquitinated by proteasomal deubiquitinase before the protein unfolds and enters
the chamber of the proteasome to undergo digestion.
Ubiquitin (shown as Ub) exists as a pool of free mol- ecules (upper left of Fig. 32.1), which can be attached either singly or in polyubiquitin chains to the sub- strate protein (bottom center). The polyubiquitinat- ed protein substrate is then degraded by the 26S pro- teasome to smaller peptides, which can then either be degraded to amino acids by other proteinases or be presented on the surface of the cell membrane in the immune cells. The beta subunits in the middle of the 20S proteasome include chymotrypsin-like and trypsin-like catalytic enzymes, which digest the proteins that enter the chamber. The entrance of the proteins into the digestion chamber is gated by alpha subunits. In this way the digestion of proteins is selectively regulated. Proteins that are digested by the 26S proteasomes must first be targeted to pro- teasomes by at least a 4 polyubiquitin chain where the C-terminal glycine is covalently bound to lysine residues in the protein destined to be digested by the proteasome.
32.3
Ubiquitination of Proteins Destined for Digestion by the Proteasome
Initially, in the conjugation phase of ubiquitin to the protein destined for proteolysis by the proteasome, ubiquitin is first activated through the ATP-depend- ent formation of thiol ester with a cysteine residue of the ubiquitination-activating enzyme (E
1) (Fig. 32.1).
The carboxyl group of the last amino acid of ubiqui- tin, a glycine-glycine dipeptide, is first activated by adenylation. A thiol group in the activating enzyme (E
1), which initiates the ubiquitination enzyme cas- cade, attaches the ubiquitin-carboxyl-adenosine 5’- monophosphate to form a E
1-ubiquitin thiol ester.
Ubiquitin is then transferred to a cysteine residue on a ubiquitin-conjugating enzyme (E
2) (Fig. 32.1).
Ubiquitin is then transferred to a lysine residue of
the protein substrate in a reaction catalyzed by a
ubiquitin-protein ligase (E
3) (Fig. 32.1) [38, 40]. All
known E3s utilize one of two catalytic domains, HECT domains or a RING finger [81].
The ubiquitin-proteasome system of regulated protein digestion is highly selective where signal- dependent proteolysis directs the time and location of protein turnover in the cell [38, 40]. The selection of proteins to be degraded by the proteasome oc- curs at the point of conjugation, though the E
2s and E
3s have different substrate specificities. Each E
3, in conjunction with its cognate E
2, binds substrates that share a particular ubiquitination signal, usual- ly a small primary sequence motif [53, 62, 80]. Rec- ognition of the primary signal targets the substrate to be labeled with a second signal, i.e. the polyu- biquitin chain formation. Only the polyubiquitin chain signal is recognized by the proteasome. The phenomenon of polyubiquitinated secondary signal formation provides a large number of substrates for the proteasome, which are predestined for selective digestion.
The recognition of the secondary signal by the proteasome is true only for the proteasome and differs in this way from other ATP-dependent pro- teases in the cell [100]. The ubiquitin molecules are linked covalently to one another through K48-G76 isopeptide bonds. The first ubiquitin binds a lysine in the substrate. This supports residues in the sub- strate to generate an isopeptide bond [16]. The af- finity of binding of the polyubiquitin chain to the proteasome increases 100-fold as the chain length increases from two to four ubiquitins, but increases
only tenfold when eight more ubiquitin molecules are added [100]. Therefore, the length of the polyu- biquitin chain determines the rate of substrate deg- radation by the proteasome, after the chain binds to the 19S proteasome, possibly because of the nature of the deubiquitinating enzyme chain reaction. If the chain is too short before degradation is initiat- ed, the substrate will be deubiquitinated, but not de- graded. A longer ubiquitin chain would allow more time for unfolding of the substrate and opening the gate to allow entrance of the substrate into the cata- lytic chamber of the 20S proteasome for digestion.
The ubiquitin-proteasome system is regulated at the level of ubiquitination or at the level of the proteasome activity. The targeting motif can be the N-terminal residue, a sequence, a domain, or a post- translational change such as phosphorylation that results from externally generated signal transduc- tion. Phosphorylation of either the substrate or the ligase can activate binding of the ligase. Binding can be enhanced by ubiquitin-like proteins (UBL) by in- creasing the affinity of the ligases to the E
2compo- nent of the conjugating apparatus, especially in the case of phosphorylated substrates. UBL proteins in- teract with the polyubiquitinin domain (UBA). The UBL domain probably acts to facilitate the interac- tion with UB-containing proteins, targeting them to the proteasome [17]. Linking elements on proteins such as UBA and UBL may increase the efficiency of the proteasome process.
ATP
Deubiquitination
Fig. 32.1. Scheme of the ubiquitin-proteasome pathway
32.4
Ubiquitin-Proteasome Mechanics
The processing of short-lived regulatory and struc- tural proteins within the cell for degradation by the ubiquitin-proteasome system is extraordinarily complex and has recently been reviewed elsewhere in detail [32]. The process involves the folding of intracellular proteins by heat shock proteins and chaperones [35, 37]; polyubiquitination by three enzymes in sequence (ubiquitin-activating enzyme, E1; ubiquitin-conjugating enzyme, E2s; and ubiqui- tin-protein ligase, E
3s); assembly of the 20S protea- some containing the catalytic subunits; assembly of the 19S proteasome including the base and lid;
assembly of the 26S subunit; ATP, and deubiquiti- nating enzyme. There are thousands of proteasomes in hepatocytes that are mobile in the cytosol and localized to the nuclear pore, endoplasmic reticu- lum, and the nucleus [33]. In pathologic conditions, where proteins accumulate to form an aggresome, the proteasomes localize in the aggresomes [42].
When isolated from the liver they are found as 20S, 20S with one 19S, and 20S with two 19S attachments (26S) [28]. The 20S proteasome does not require ubiquitination of the proteins or ATP to digest pro- teins. This includes oxidized proteins [91, 97]. Thus, the digestion of proteins by the 26S proteasome is regulated by the polyubiquitinated protein and ATP-dependent mechanisms.
Other proteins involved in delivering the ubiq- uitinated proteins to the proteasome for digestion include CHIP, Rad 23, AG-1, and P62 (ZIP). These proteins dimerize and bind to polyubiquitin chains and the proteasome to facilitate proteolysis through a hypothetical UBA/UBL [18, 30, 35–37, 83, 102]. The protein substrates for proteasome digestion are nu- merous and the list is still growing [24, 101]. These include p53, Cdc 25, Cdc 18, IKB α, β-catenin, CD4, HIF- α, EGF-R, PDGF-R, Mdm
2, and MAT α2, which are involved in liver diseases. Each has a different ubiquitin ligase.
32.5
Experimental Alcoholic Liver Disease and Proteasomal Catalytic Activity
Donohue [22] has recently reviewed the literature summarizing the evidence implicating the ubiq- uitin-proteasome system in ethanol-induced liver disease. He first introduced the concept [21] but, to date, it has largely been ignored in recent reviews of the mechanisms of alcohol-induced liver injury
[69]. The concept of liver injury through this mech- anism applies to both alcoholic and non-alcoholic liver disease in man [98, 99], as well as experimental ethanol-induced liver disease in rats and mice [4, 6, 21, 23, 28].
32.6
Functional Consequences of Ethanol Inhibition
of the Proteasome in the Liver
Donohue et al. [21] first showed that the activities of three proteolytic enzymes located in the 20S pro- teasome, i.e., chymotrypsin-like, trypsin-like, and peptidyl-glutamyl-peptide hydrolase, were inhibited in rats fed ethanol intragastrically for 2 months but not in rats fed ethanol ad lib. The inhibition of these proteasome enzyme activities correlated with an in- crease in liver weight, compared to pair-fed controls.
The liver total proteins also increased significantly compared to pair-fed controls, suggesting that the increase in liver weight and protein content was a result of the inhibition of protein proteolysis asso- ciated with ethanol intake. The relative amounts of proteasome subunits did not change with ethanol, suggesting that the inhibition of catalytic activity was not due to reduced enzyme protein levels. The proteasome inhibition was shown in the standard activities, i.e. in the SDS-stimulated 20S proteasome activities, and in the ATP-stimulated 26S protea- some activities, indicating that the defect involved the 26S proteasome as well as the 20S proteasome.
The inhibition of proteasome activity corre- lated with an increase in malondialdehyde in liver homogenates, suggesting that malondialdehyde ad- ducts resulting from increased lipid peroxidation caused by alcohol could be involved in proteasome inhibition. None of these changes was noted in the rats fed ethanol ad lib, suggesting that a dose-re- sponse relationship is responsible for proteasome inhibition [21]. The studies of Fataccioli et al. [23]
corroborated those of Donohue et al. [21]. In addi-
tion, when CYP2E1 inhibitors were fed with ethanol,
inhibition of the proteasome chymotrypsin-like ac-
tivity was significantly reduced, suggesting that free
radical damage may be the mechanism of inhibition
of the proteasome caused by ethanol feeding. Li-
pid peroxidation correlated inversely with chymot-
rypsin-like activity. Again, ad lib feeding of ethanol
did not cause these associations with proteasome
inhibition, nor was there an increase in formation of
malondialdehyde, protein thiols, or protein carbon-
yls. Also, ad lib feeding of ethanol did not increase
the total liver protein levels.
Rouach et al. [88] found that rats fed ethanol intragastrically had increased levels of malondial- dehyde, protein thiols, and protein carbonyls in the liver. The increase in oxidized proteins correlated positively with the severity of the liver pathology.
Thus, the inhibition of proteasome activity and the increase in protein oxidation may be related because the proteasome eliminates oxidized proteins [91].
32.7
Other Functional Consequences of Proteasome Inhibition
Ethanol stabilizes proteins normally degraded by the proteasome. Hypothetically, this would include CYP2E1, NF- κB, and HIF-1α. In the case of CYP2E1, it has been shown that ethanol withdrawal leads to an overnight reduction in the ethanol-induced CYP2E1 and that this rapid loss in CYP2E1 in the liver can be prevented by injecting a potent proteasome inhibi- tor, PS-341 [7]. This change in CYP2E1 levels cor- related with a return of the proteasome, inhibited by ethanol, to normal levels of activity 24 h after ethanol withdrawal. This is compelling evidence that CYP2E1 was stabilized by ethanol inhibition of the proteasome. Proteasome inhibitor treatment of HepG2 cells in vitro increases the level of CYP2E1 threefold by preventing degradation by the protea- some (stabilizes CYP2E1) [79]. In the case of NF- κB stabilization, it was shown that ethanol fed intragas- trically for 2 months was associated with the lack of NF- κB activation [34], which could be the result of inhibition of proteasome digestion of I κBα that pre- vents NF- κB activation. NF-κB would be expected to be activated by oxidative stress caused by ethanol in this model. In the case of HIF-1 α, the nuclear activa- tion of this transcription factor is increased at high blood alcohol levels and expression of genes regu- lated by HIF-1 α is also increased, including eryth- ropoietin, iNOS, and VEGF gene expression [7, 70], suggesting that the normal elimination of HIF-1 α by the proteasome is inhibited by ethanol feeding.
32.8
The Mechanism of Ethanol-Induced Inhibition of Proteasome Activity in the Liver
Ethanol feeding does not inhibit proteasome cata- lytic activity directly [10]. The inhibition does not occur after 2 weeks of ethanol feeding. Inhibition of the proteasome was not found until 1 month of feed- ing rats ethanol. This suggested that an oxidized product of CYP2E1 generated by ethanol CYP2E1
would be a likely mechanism of inhibition. Indeed, transgenic mice, in which the gene for CYP2E1 had been knocked out, failed to develop proteasome activity inhibition even after 1 month of ethanol feeding [4]. One candidate product is 4-hydrox- ynonenal (4-HNE), an end product of lipid peroxi- dation, which results from the hydroxyethyl radical generated by CYP2E1 during ethanol oxidation [2].
Indeed, 4-HNE is increased in the liver of rats fed ethanol intragastrically [26]. 4-HNE progressively increased in the liver during 1 month of ethanol feeding, peaking when the proteasome catalytic ac- tivity had become inhibited [10]. 4-HNE adducts are formed in the liver in alcoholic liver disease. Their presence correlated with the morphologic severity of liver disease [89].
The possibility of adduct forming between a pro- teasome subunit and 4-HNE accounting for the loss of proteolytic activity was investigated. When the 20S proteasome was isolated from the livers of rats fed ethanol and controls, no loss of catalytic activity was found. Also, no 4-HNE adduct was found when the 20S proteasome was studied by two-dimension- al (2D)-electrophoresis and Western blot. However, when the 26S proteasome was isolated, and studied by 2D-electrophoresis and Western blot, there was loss of proteasome activity, and a 4-HNE adduct to a 44-kDa subunit was found in the alcohol-fed rats.
The 2D membrane, stripped of the 4-HNE antibody and stained with an antibody to Rpt4, an ATPase subunit of the regulatory complex 19S part of the 26S proteasome, showed that 4-HNE was adducted to this subunit in the ethanol-fed rat liver. Thus, proteasomal catalytic activity could be lost because the subunit that controls the opening of the catalytic channel was altered by 4-HNE adduction [10].
Another possible mechanism that could explain
the loss of proteasome catalytic activity after ethanol
feeding is phosphorylation of a regulatory subunit of
the proteasome. Previous in vitro studies had shown
that ethanol stimulated phosphorylation of proteins
in liver cells through a protein kinase C (PKC) route
[49]. Okadaic acid, which inhibits phosphatase ac-
tivity and increases phosphorylated proteins in the
liver, caused profound inhibition of proteasome
activity in vivo [8]. Proteasomes isolated from the
liver of rats fed ethanol and controls were studied
for subunit phosphorylation using an antibody to
phosphothreonine. Using 2D-electrophoresis and
Western blotting, a hyperphosphorylated subunit,
alpha 7, was found in the proteasomes isolated from
the liver. Inhibition of proteolytic activity of the
20S proteasome could result from phosphorylation
of this subunit because the α subunits are also in-
volved in opening the mouth of the catalytic cham-
ber of the 26S proteasome.
32.9
Some Consequences of Inhibition of the Proteasome: Apoptosis
Apoptosis in the liver is increased in alcoholic liver disease [27]. Apoptosis is also increased in the liver from viral hepatitis and drug hepatitis. Inhibition of the proteasome is one mechanism involved in apoptosis. When rats were injected with the potent proteasome inhibitor PS-341 (IP 0.5 mg/kg), apop- tosis increased compared to controls [28] (Fig. 32.2).
The livers were examined for apoptosis 4 h post in- jection using TUNEL staining. Positive-stained nuclei were increased, as were cells that stained positive for activated caspase-3 (hepatocytes and sinusoidal cells). Alanine aminotransferase (ALT) was increased. Proteasome chymotrypsin-like ac- tivity was decreased in liver homogenates (~ 85%
decrease). The way in which proteasome inhibition leads to apoptosis is complex. Many pro- and anti- apoptosis proteins are regulated by the proteasome [28]. These include Bax, Bcl-2, p53, mdm 2, IKB α, caspase 3, caspase 8 c-1AP1, XiAP, NF- κB precursor, and Bid [74]. Pro-apoptosis proteins are p53, Bax, and Bid. Anti-apoptosis proteins are c-IAPI, XIAP,
Bd-2, and NF- κB. Thus, the ubiquitin-proteasome system is an essential player in the regulation of proteins involved in programmed cell death in the liver and other organs.
32.10
Role of the Ubiquitin-Proteasome Pathway in Liver Cell Aggresome (Mallory Body) Formation
Mallory bodies are formed by damaged liver cells in a great variety of different chronic liver diseases [41]. Thus, they are a sign of more severe liver injury [3, 90]. In alcoholic liver disease, ubiquitin conju- gates (polyubiquitinated proteins) are increased in the blood [99] and ubiquitin in the liver correlates with the presence of Mallory bodies in non-alco- holic steatohepatitis [3]. Thus, there is a general in- crease in polyubiquitinated proteins in an environ- ment in which Mallory bodies form. The association of Mallory bodies with ubiquitinated cytokeratins in liver cells has been known since 1988 [64, 75].
Only recently has the role of the ubiquitin-protea- some pathway in the pathogenesis of Mallory body
Fig. 32.2a –f. Comparison of ethanol feeding and proteasome inhibition effects on liver cells.
a–d Hematoxylin and eosin-stained sections from the liver of rats fed ethanol for 1 month (a), or fed ethanol for 1 month and given PS- 341 (b) or rats given PS-341 only (c, d). Note that both ethanol and PS-341 caused apoptosis (a, c). e, f Hepatocytes stained by TUNEL from the liver of a rat fed ethanol for 1 month (e) or a rat fed ethanol for 1 month and given PS- 341 (f). PS-341 treatment caused apoptosis (f) mainly when combined with ethanol (e)
formation been appreciated. The new concept is based on the aggresome phenomenon, which biolo- gists have described in a variety of tissues where protein aggregates accumulate around the centro- some [42, 104, 106]. The idea was first formulated as a consequence of events in the degradation pathway of liver cell cytokeratins. It started with a conforma- tional change in the cytokeratin proteins caused by phosphorylation, where α helices decreased and β sheets increased, causing the cytokeratins to stick to each other [109]. According to the concept of ag- gresome formation, β-sheet conformational change is an important mechanism for protein aggregation [54]. Conformational changes in Mallory bodies have been documented by infrared spectroscopy [44]. Hyperphosphorylation of cytokeratin 8 by hepatocytes from mouse livers forming Mallory bodies was documented by
32P incorporation into cytokeratins of hepatocytes in tissue culture using 2D-electrophoresis [15]. Mallory bodies containing hyperphosphorylated proteins have been found in a model of Mallory body formation in hepatocytes.
Phosphothreonine antibody was used in Western blots of Mallory body protein [72]. Hyperphosphor- ylation of cytokeratins in Mallory-body-forming liv- ers has been characterized further by finding phos- phorylation of multiple sites on amino acid residues in the cytokeratin proteins [94]. The turnover of cy- tokeratins is signaled by phosphorylation, followed
by ubiquitination. Proteolysis of the cytokeratins by the proteasome then occurs [56].
To test the hypothesis that hyperphosphorylation induces Mallory body formation, drug-primed mice were given an acute dose of okadaic acid and the liv- ers were monitored for the appearance of Mallory bodies. Fifteen minutes after okadaic acid injection, phosphorylated protein aggregates formed in liver cells. The aggregates stained positive with cytokera- tin antibody [109]. True Mallory bodies formed by day 2 and progressed with daily injections of oka- daic acid. The Mallory bodies stained positive for phosphothreonine in tissue slices and by Western blot in the Mallory body band. Human liver cell Mallory bodies also stained positive with the phos- phothreonine antibody. Similar cytokeratin aggre- gates formed in hepatocytes in tissue culture when exposed to okadaic acid [13, 92].
32.11
Role of Heat Shock Proteins in Mallory Body Formation
Heat shock proteins act as chaperones for altered cytoplasmic proteins that have been targeted for digestion by the proteasome after polyubiquitina- tion (Fig. 32.3) [105]. Heat shock treatment of drug-
Fig. 32.3a, b. Theoretical model of the role of heat shock pro- teins (hsp) in normal protein degradation and MB formation. (a) Normal cytokeratin degradation. (b) Cytokeratin aggregation and formation of MBs in the presence of stress and/or UBB+1. Ub
ubiquitin, hsp 70 heat shock protein 70, hsp 90 heat shock protein 90, MB Mallory body, UBB+1 mutant ubiquitin, cytokeratin. (Re- printed from [86], with permission from Elsevier)
primed mice induces Mallory bodies after upregula- tion of heat shock protein expression [108]. When human or mice livers containing Mallory bodies were stained with antibodies to heat shock proteins 70 and 90, they stained positive, indicating that both heat shock proteins were pathogenetically linked to Mallory body formation [86, 95]. The expression of heat shock protein 70 and 90 as determined by West- ern blot correlated with the number of Mallory bod- ies expressed [9].
32.12 Role of p62
in Mallory Body Formation
P62 (Z1P) links polyubiquitinated proteins to the proteasome. P62 is a major component of hyaline bodies in hepatocellular carcinomas [93], Mallory bodies [96], as well as Alzheimer’s disease neurofi- brillary tangles and synucleipathies [59, 60] are in- volved in p62-ubiquitin binding in aggresome for- mation. ZIP/p62 is a scaffolding protein that binds to polyubiquitin at its UBA domain [30, 82]. Human Mallory bodies stain strongly with p62 antibody (Fig. 32.4).
32.13
Role of Microtubules in Mallory Body Formation
Aggresomes require microtubules to be present, in order for them to form. Colchicine and nocoda-
zole prevent aggresome formation [42, 104, 106].
Miniaggregates are transported on microtubules towards the microtubule organizing center at the centrosome, driven by the minus-end motor protein dynein [29, 43]. Using a new tissue culture model of Mallory body formation, microtubule inhibitors totally blocked Mallory body formation [87]. Only small cytokeratin-ubiquitin positive aggregates formed at the cell periphery, whereas the control liv- er cultures formed numerous Mallory bodies in the absence of the microtubular inhibitors (Fig. 32.5).
Tubulin and Tau isoforms located in microtubules localized in the Mallory body [51, 84].
32.14
Role of Transglutaminase in Mallory Body Formation
Transglutaminase was found in the Mallory bodies and in Western blots and was induced when Mallory bodies were induced [9]. Northern blots showed that transglutaminase gene expression was upregulated when Mallory bodies were formed at the same time that cytokeratin 8 was overexpressed. Increased transglutaminase activity and cross-linking of cy- tokeratins were reported earlier [19, 110, 111]. Cross- linking of cytokeratin by transglutaminase in hepa- tocytes may account for the insolubility of Mallory bodies in the SDS cocktail used in Western blots [78].
Fig. 32.4a–c. Human liver cells with numerous Mallory bodies due to alcohol abuse. The liver section was double stained with antibodies to cytokeratin 18 (a) and to p62 (b) and viewed with
a confocal microscope. a, and b images were merged in c. Note the merged image shows the Mallory bodies, indicating colo- calization of p62 and cytokeratin 18 (×2,500)
32.15
Role of Proteasome Inhibition in the Formation of Mallory Bodies
Proteasome inhibitors induced aggresome forma- tion in vitro, presumably because the inhibition of proteolysis causes an accumulation of proteins in the cell [42, 106]. This means that inhibition of the proteasome is central to Mallory body formation [109]. In fact, injection of a potent proteasome in- hibitor in drug-primed mice induces Mallory body formation after 2 days [24]. The proteasome inhibi- tor PS-341 is selective in its action on the protea- some and is an effective anticancer drug because proteasome inhibition causes apoptosis of cancer cells [1]. PS-341 also induces aggresome formation in hepatocytes in vitro after going through a stage of collapse of cytokeratin from the plasma membrane followed by retraction of the cytokeratin around the nucleus [85] at 5 h. Later (24 h), aggresomes are formed in place of the contracted cytokeratin. When Mallory bodies form in mouse liver, the cytokeratin disappears in the cytoplasm (ghost cell formation).
Proteasomes are localized at the edge of the Mallory body and the proteasomes in the cytoplasm are di- minished [5].
32.16
Role of the Frameshift Mutant
of Ubiquitin in Mallory Body Formation
Recently, a frameshift mutant of ubiquitin, UBB
+1, was shown to be present in Mallory bodies in hu-
man and mouse livers [24, 68]. The UBB
+1mutant was first reported to be present in neuronal cell ag- gresomes in Alzheimer’s disease [103]. This mutant is a result of molecular misreading of the ubiquitin B gene during transcription (a dinucleotide dele- tion), which results in a mutant ubiquitin that has lost the binding site in the C prime end of ubiquitin.
It therefore cannot function to bind misfolded pro- teins destined for proteolysis by the ubiquitin-pro- teasome pathway. This is not a functional ubiquitin.
Worse, UBB
+1is polyubiquitinated by wild-type ubiquitin, but the resultant polyubiquitin chains are refractory to disassembly by deubiquitinating enzymes (Fig. 32.6). This potentially inhibits the degradation of a polyubiquitinated substrate by pu- rified 26S proteasomes [61]. In vitro studies using primary liver cultures showed that UBB
+1transfec- tion induced cytokeratin-ubiquitin aggresomes in hepatocytes within 4 h of incubation [11]. In vitro studies used a ubiquitin-proteasome pathway mix- ture to form a reconstituted ubiquitin-proteasome, which includes a cytokeratin immunoprecipitate.
This mixture contained Mallory bodies, added ubiquitin, a ubiquitination system with the enzymes E
1, E
2, and E
3, purified proteasomes, and an enzyme mixture for generating ATP. The mixture was in- cubated for 4 and 24 h [9]. The Mallory body band (insoluble protein at the top of the gel located in the loading gel) increased when the mixture was added.
Likewise, the ubiquitinated protein fraction was in- creased. Adding UBB
+1to the mixture increased the Mallory body band further [11]. Adding PS-341 also increased the Mallory body band. The combination of UBB
+1and PS-341 did not increase the Mallory body band further, suggesting that the two treat- ments were acting through the same mechanism.
Fig. 32.5a–c. Primary culture of mouse liver cells forming Mal- lory bodies. The liver cells were double stained for cytokeratin 18 (green, a) and for ubiquitin (red, b) and viewed with a confocal microscope. c a merged picture from a and b. Note the merged
image shows the Mallory bodies in yellow, indicating colocaliza- tion of ubiquitin and cytokeratin 18. Confocal fluorescent micro- scopy (×2,500)
Ubiquitination of UBB
+1occurred. The proteasomes localized within the Mallory body, indicating that, as occurs in aggresomes in vivo, the proteasomes were bound to the aggresome. This might explain how UBB
+1interferes with the ubiquitin-proteasome pathway of proteolysis.
The question arises: does UBB
+1expression lead to cell death? In the case of neurons, UBB
+1expres- sion does lead to cell death [20]. Aggresome forma- tion under many differing circumstances may cause death of the cell [12, 63].
32.17
Role of Cytokeratin 8 and 18 Overexpression in Mallory Body Formation
During primary tissue culture of liver cells from Mallory-body-forming livers, there was an increase in synthesis of cytokeratin 8 and cytokeratin 18 as measured by Western blot [15]. Cytokeratin 8 gene expression was also markedly upregulated in Mal- lory-body-forming livers [72]. The increase of cy- tokeratins in hepatocytes may overload the ubiqui- tin-proteasome pathway of cytokeratin proteolysis, which then may cause cytokeratin accumulation to form the Mallory body. Several studies, where tissue culture cells were transfected with plasmids con- taining cDNA coded for cytokeratin 8 or cytokera-
tin 18 to cause overexpression, led to the formation of cytokeratin aggresomes. But these aggresomes failed to form the characteristic filaments seen in Mallory bodies [14, 39, 73].
Mallory bodies incorporate newly synthesized cytokeratin directly in tissue culture as shown by [
35S]methionine pulse labeling using Western blots and electron microscopy autoradiography of deter- gent-extracted hepatocytes in culture where only the cytoskeleton remained [45]. In Western blots, the radiolabel was found in the Mallory band as well as in normal cytokeratin 8 and cytokeratin 18 bands.
Similarly, the radiolabel was incorporated into the Mallory filaments as well as the cytokeratin fila- ments. Cytokeratin 18 cDNA was used to transfect Mallory-body-forming cells in mouse liver tissue culture using a green fluorescent protein tag. The cytokeratin localized in both the normally distrib- uted cytokeratin but was also mainly concentrated in the Mallory body. This suggested that the nascent human cytokeratin went both to the normal fila- ments as well as directly to the Mallory body after synthesis [84, 85].
Fig. 32.6. Molecular misreading and Mallory body formation. Representation of how target proteins can be tagged efficiently for protea- somal destruction (right side) and how this oc- curs less efficiently (left side) when the mutant ubiquitin UBB+1 is involved in the ubiquitina- tion. UBB+1 has lost its ability to ubiquitinate because it does not have a terminal Gly resi- due. It is refractory to deubiquitination by the deubiquitinase (isopeptidase) enzyme. In fact, the mutant ubiquitin becomes ubiquitinated itself, and inhibits the proteasome in a domi- nant-negative way. (Reprinted from [68], with permission from Elsevier)
32.18
Functional Consequence
of the Loss of Cytokeratin Organization Due to Aggresome (Mallory Body) Formation
The function of liver cell cytokeratins and the vul- nerability of liver cells exposed to toxins and viral damage have been reviewed elsewhere [52, 57, 76, 77]. Mutations in cytokeratin 8 and cytokeratin 18 were found in patients with cryptogenic cirrhosis, hepatitis C, and other liver diseases. The question is: do changes in the cytokeratin organization due to mutations or other etiologies interfere with liver function and increase vulnerability of the liver to in- jury? For instance, hyperphosphorylation of the liv- er cell cytokeratins changes their organization, but does this lead to loss of function [55]? Cytokeratin reorganizes extensively during mitoses, apoptosis and polarization of the cell [66]. In in vitro studies using transfection of cells with mutant cytokeratins, the cells were more vulnerable to injury. Cytokerat- ins have viscoelastic properties, which make cells resistant to physical stress. Lack of cytokeratins in K18/8-deficient mice hepatocytes disturbs the pro- gression of the cell cycle. These cells are more likely to develop apoptosis in response to FAS [31, 58, 65].
The cytokeratins of the liver anchor and support the bile canaliculus, forming the pericanalicular sheath supporting bile flow [47, 48]. Likewise the cytokera- tin filaments of the liver cell are involved in transhe- patic transport of vesicles [50].
The cytokeratin cytoskeleton of liver cells that form Mallory bodies is profoundly altered so that only the Mallory body stains for keratin, leaving the rest of the cell unstained, the so-called “empty cell”
or “ghost cell” [46]. Mallory body cells lose their bile canaliculi and cytokeratin in the cytoplasm. This is associated with cell enlargement and rounding. The cytokeratin network supports the nucleus, the nu- clear pores and the centrioles. The Mallory-body- containing cells in primary tissue culture were un- able to form bile canaliculi or secrete fluorescent diacetate, compared with controls [25].
32.19
Concluding Remarks
It can be concluded that the inhibition of the ubiq- uitin-proteasome pathway of protein degradation has profound effects on the structure and function of liver cells, including loss of cell shape and inter- nal organization of organelles, polarity, and the bile secretory apparatus. Vulnerability of the liver cell
to FAS-induced apoptosis is increased. Cytokeratin aggresome formation (Mallory bodies) may further reduce proteolysis by the proteasome.
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