|Cancer||Cell death||Cell cycle||Cytoskeleton||Exo/endocytosis||Differentiation||Division||Organelles||Signalling||Stem cells||Trafficking|
Hepatic stellate cell (vitamin A-storing cell) and its relative – past, present and future
Haruki Senoo*1, Kiwamu Yoshikawa*, Mayako Morii†, Mitsutaka Miura*, Katsuyuki Imai* and Yoshihiro Mezaki*
*Departments of Cell Biology and Morphology, Akita University Graduate School of Medicine, 111 Hondo, Akita City, Akita 0108543, Japan, and †Department of Pediatric Surgery, Akita University Graduate School of Medicine, 111 Hondo, Akita City, Akita 0108543, Japan
HSCs (hepatic stellate cells) (also called vitamin A-storing cells, lipocytes, interstitial cells, fat-storing cells or Ito cells) exist in the space between parenchymal cells and liver sinusoidal endothelial cells of the hepatic lobule and store 50–80% of vitamin A in the whole body as retinyl palmitate in lipid droplets in the cytoplasm. In physiological conditions, these cells play pivotal roles in the regulation of vitamin A homoeostasis. In pathological conditions, such as hepatic fibrosis or liver cirrhosis, HSCs lose vitamin A and synthesize a large amount of extracellular matrix components including collagen, proteoglycan, glycosaminoglycan and adhesive glycoproteins. Morphology of these cells also changes from the star-shaped SCs (stellate cells) to that of fibroblasts or myofibroblasts. The hepatic SCs are now considered to be targets of therapy of hepatic fibrosis or liver cirrhosis. HSCs are activated by adhering to the parenchymal cells and lose stored vitamin A during hepatic regeneration. Vitamin A-storing cells exist in extrahepatic organs such as the pancreas, lungs, kidneys and intestines. Vitamin A-storing cells in the liver and extrahepatic organs form a cellular system. The research of the vitamin A-storing cells has developed and expanded vigorously. The past, present and future of the research of the vitamin A-storing cells (SCs) will be summarized and discussed in this review.
Key words: hepatic stellate cell, retinoid-binding protein, vitamin A, vitamin A-storing cell
Abbreviations: αSMA, α smooth muscle actin, 9RA, 9-cis retinoic acid, ADRP, adipose differentiation-related protein, ARAT, acyl-CoA/retinol acyltransferase, Asc 2-P, l-ascorbic acid 2-phosphate, ATRA, all-trans retinoic acid, CRBP, cellular retinol-binding protein, CTGF, connective tissue growth factor, ECM, extracellular matrix, EGF, epidermal growth factor, EMT, epithelial to mesenchymal transition, ER, endoplasmic reticulum, FGFs, fibroblast growth factors, FoxO, Forkhead box gene, group O, FXR, farnesoid X receptor, GFAP, glial fibrillary acidic protein, HSCs, hepatic stellate cells, KCs, Kupffer cells, LRAT, lecithin/retinol acyltransferase, LSCs, lung stellate cells, LSECs, liver sinusoidal endothelial cells, MEF2, myocyte enhancer factor 2, MMPs, matrix metalloproteinases, MVB, multivesicular body, N-CAM, neural cell adhesion molecule, NPCs, non-parenchymal cells, PCs, parenchymal cells, PHx, partial hepatectomy, PI3K, phosphatidylinositol 3-kinase, PPARs, peroxisome proliferator-activated receptors, PSCs, pancreatic SCs, PXR, Pregnane X receptor, RARs, retinoic acid receptors, RBP, retinol-binding protein, SCs, stellate cells, STAP, stellate cell activation-associated protein, STRA6, stimulated by retinoic acid 6, TIMP, tissue inhibitor of metalloproteinases, VFSC, vocal fold SCs
1To whom correspondence should be addressed (email firstname.lastname@example.org).
Part of a series marking the 70th birthday of the Cell Biology International Editor-in-Chief Denys Wheatley
The research of HSCs (hepatic stellate cells) has re-started when the cells were re-discovered by Wake (1971) (reviewed by Wake, 1980). Establishment of cell isolation and culture methods of HSCs and standardization of the nomenclature (Hepatic stellate cell nomenclature, 1996), namely, HSCs, have promoted the development of HSC research (Friedman, 2008b; Senoo et al., 2010). Excellent and new reviews on HSCs have been already published (Wake, 1980, 1998; Blomhoff and Wake, 1991; Senoo et al., 1997, 2007; Li and Friedman, 2001; Sato et al., 2003; Senoo, 2004, 2007b; Friedman, 2008b; Smedsrød et al., 2009), but we would like to write this review emphasizing pioneer works, basic research and important but not well-known studies. Not only physiological roles but also roles of HSCs in pathogenesis of diseases are intriguing.
The hepatic lobule consists of liver PCs (parenchymal cells) and NPCs (non-parenchymal cells) associated with the liver sinusoids: LSECs (liver sinusoidal endothelial cells), KCs (Kupffer cells), pit cells, dendritic cells and HSCs (Wake, 1971, 1980; Bloom and Fawcett, 1994; Figure 1). LSECs (Wisse, 1970, 1972) express lymphocyte co-stimulatory molecules (Kojima et al., 2001) and form the greater part of the extremely thin lining of the sinusoids, which are larger than ordinary capillaries and more irregular in shape. KCs are tissue macrophages and components of the diffuse mononuclear phagocyte system. They are usually situated on the endothelium with cellular processes extending between the underlying LSECs. The greater part of their irregular cell surface is exposed to the blood in the lumen of the sinusoid. Pit cells are natural killer cells. Dendritic cells locate in the portal triad in human liver (Prickett et al., 1988) and in periportal and central areas in rat liver (Steiniger et al., 1984) that capture and process antigens, migrate to lymphoid organs and secrete cytokines to initiate immune responses (Banchereau and Steinman, 1998). The HSCs that lie in the space between LSECs and PCs are considered to be derived from mesenchymal origin. Both LSECs and HSCs are derived from mesenchymal tissue, namely, septum transversum. KCs are from the monocyte–macrophage system.
Mesenchymal cells that store vitamin A in their cytoplasm have been found in extrahepatic organs (kidneys, intestines, lungs, pancreas, etc.) and characterized [reviewed by Wake (1980); Nagy et al., 1997; Apte et al., 1998; Bachem et al., 1998; Matano et al., 1999]. The purpose of this review is to summarize the past and present of the HSC and extrahepatic vitamin A-storing cell research and try to describe the future of the research.
Discovery and re-discovery of HSCs
The SCs (stellate cells) (‘Sternzellen’) in the liver (HSCs) were discovered by Kupffer (1876) [reviewed by Wake (1980)], using the gold chloride method (Wake et al., 1986). As a neuroanatomist, Kupffer tried to find nervous components in the liver of rat, mouse, rabbit, ox, pig, dog and human by using the gold chloride method at that time. Accidentally, he found black-stained star-shaped cells in the liver and called the cells SCs (‘Sternzellen’). The gold chloride reaction is specific; only HSCs are clearly stained black in the background of red-stained other liver cells such as liver PCs. Kupffer described correctly that HSCs located in the space between PCs and LSECs (Figure 1). Rothe, one of his students, applied the gold chloride method to livers in other mammals and found similar cells in these livers (Rothe, 1882). However, 22 years after the discovery, a misconception by Kupffer (1898, 1899) generated new misconception and led to a great confusion in the liver histology [Wake, 1974, 1978, 1982, 2009; reviewed by Wake (1980)]. Kupffer (1898,1899) ‘corrected’ his description in 1876 of the SCs as a misconception in the 12th Congress of Anatomists held in Kiel in 1898. He compared the liver specimen stained by gold chloride method with the rabbit liver specimen injected with India ink and concluded that ‘Sternzelle’ was a special endothelial cell that phagocytosed foreign materials. Although the research on phagocytic liver cells has progressed from Kupffer's ‘correction’, the research contained errors; ‘phagocytic liver cells’ included both of phagocytic KCs and HSCs. From Kupffer's ‘correction’ of 1898, the SCs (‘Sternzellen’, HSCs) of the liver disappeared from the history of the research of the liver. The true nature of these cells remained enigmatic for over several decades until they were re-discovered by Wake (1971, 1980). Wake has revealed that the HSCs are quite different from the phagocytic Kupffer cells being located in the sinusoid forming by LSEC. HSCs locate in the space between the PCs and LSECs. HSCs are identical to the perisinusoidal cells such as fat-storing cells (Ito, 1951) and interstitial cells (Suzuki, 1958).
The history of the study of HSCs was thoroughly described and discussed by the review of Wake (1980). Wake divided the history of the study of HSCs into three periods. During the first period (1876–1897), HSCs were correctly recognized with regard to cell shape, distribution and localization, namely, in the perisinusoidal space. During the second period (1898–1970), due to Kupffer's ‘correction’ in 1898, HSCs disappeared from the liver research. In Kupffer's ‘correction’, he changed his earlier opinion and concluded that HSCs are the special endothelial cells of the sinusoids. During the third period (1970–present) HSCs have been recognized correctly. After the re-discovery of HSCs by Wake (1971), HSCs have re-appeared and gained proper position in the liver research. During the misconception period, namely, second period, many researchers ‘found’ HSCs and named the cells differently. Various names of HSCs became barriers for the researchers to enter the HSC study and, as will be discussed later, the standardization of the nomenclature that was done in 1996 (Hepatic stellate cell nomenclature, 1996). Now, many researchers use the common name of HSCs. The standardization promoted entry of new researchers to this research field and promoted scientific communications among the researchers.
HSCs originally designated as Sternzellen (Kupffer, 1876) have been given various names as listed by the review of Wake (1980). These names are SCs, granular cells, pericytes, fat-storing cells, metalophil cells, interstitial cells, lipophagic cells, perisinusoidal cells, transmittal cells, mesenchymal cells, lipocytes, adventitious connective tissue cells, Disse-space cells, sinusoidal mesenchymal cells, Ito cells, lipid-storing cells, lipocytes perisinusoideus, vitamin A-storing cells, extravascular cells and vitamin A-uptake cells [see references in the review by Wake (1980)].
The terminology of HSCs was discussed in FASEB (Federation of American Society of Experimental Biology), and HSC was judged as the most appropriate name because this name had no personal names, indicating morphology and location of the cell (Norum, 1984).
In 1996, 98 investigators in the field of liver non-parenchymal cell biology and hepatic fibrosis sent a letter to the editor of Hepatology proposing that the term ‘hepatic stellate cells’ be used in referring to the cell type formerly known as either the Ito cell, the lipocyte, the fat-storing cell, the perisinusoidal cell, the pericyte or the parasinusoidal cell (Hepatic stellate cell nomenclature, 1996). The investigators hoped that the adoption of this proposal will eliminate a source of confusion in this rapidly growing field. Their reasons for the choice of the term ‘hepatic stellate cells’ included the following: (i) It is historically correct, (ii) It is accurate and (iii) The term ‘stellate cell’ is already consistently used in the retinoid field to refer to these cells.
After this correspondence of ‘Hepatic stellate cell nomenclature’ appeared in Hepatology, using the term ‘hepatic stellate cell’ has been increased. From this epoch-making standardization of the nomenclature, the number of studies of HSCs has increased.
Structure of HSCs
Light microscopic observation of HSCs
Gold chloride method
After Kupffer (1876) and his pupil, Rothe (1882), used the gold chloride method for the detection of HSCs, no one used this method for the research of HSCs until Wake used the method and re-discovered HSCs (Wake, 1971, 1980). He improved the original method of Kupffer (Wake et al., 1986; Senoo and Wake, 1988) because the original method of Kupffer was unstable (Wake et al., 1986). By this improvement, the gold chloride method has become stable and applicable to research of HSCs even in arctic animals in Greenland and Svalbard archipelago (Higashi and Senoo, 2003; Figure 2). By this method, HSCs are stained black in the red background of the other liver cells (Figure 2). The details of the gold chloride method and histochemistry of the gold chloride reaction were described previously (Wake et al., 1986).
Fluorescence microscopy for detection of vitamin A autofluorescence
Fluorescence microscopy is a useful tool for specific demonstration of autofluorescence of vitamin A emanating from the lipid droplets of HSCs (Wake, 1980; Senoo and Wake, 1988; Higashi and Senoo, 2003; Figure 3).
Immunofluorescence and immunohistochemistry
Immunofluorescence and immunohistochemistry using specific markers of HSCs [such as desmin, αSMA (α smooth muscle actin), GFAP (glial fibrillary acidic protein), αB crystallin, nestin] are useful methods for the demonstration of HSCs (Yokoi et al., 1984; Gard et al., 1985; Tsutsumi et al., 1987; Takase et al., 1988; Aimed et al., 1991; Buniatian et al., 1996; Neubauer et al., 1996; Niki et al., 1996, 1999; Dack et al., 1997; Messing, 1999).
SMA is recognized as a marker of activated HSCs (Enzan et al., 1994). HSCs also express genes similar to that of neural cell types, e.g. N-CAM (neural cell adhesion molecule) (Knittel et al., 1996), synaptophysin (Cassiman et al., 1999), neurotrophins and neurotrophin receptors (Trim et al., 2000; Cassiman et al., 2001), or microtubule-associated protein 2 (Sato et al., 2001b). These molecules are also used as markers of HSCs.
Other light microcopic methods
Silver impregnation method (Wake, 1980), lipid staining and haematoxylin and eosin staining (Higashi and Senoo, 2003; Irie et al., 2004) and autoradiography (Hirosawa and Yamada, 1973; Hirosawa, 1977; Hirosawa et al., 1988; Matsuura et al., 1993) are useful for the detection of HSCs. HSCs are reactive to the Golgi staining, which is usually used for neurofilament staining (Wake, 1980). Recently, the laser-induced fluorescence spectroscopy has been applied to investigate intestinal and liver tissue of normal and vitamin A-fed rats (Akhmeteli et al., 2008). A special procedure based on intensity spectral functions fitting was developed for the recognition of vitamin A in different tissues. However, the usefulness of this method is not yet fully confirmed.
Ultrastructure of HSCs
Vitamin A lipid droplets are conspicuous features of the cytoplasm of HSCs. Another conspicuous feature of HSCs is their well-developed rough ER (endoplasmic reticulum) with membrane-bound ribosomes. ER lumen or ER cisternal space is moderately expanded indicating active biosynthesis of secretory polypeptides or proteins. Development of smooth ER is poor. A small amount of glycogen particles exist in the cytoplasm. Lysosome exists in HSCs, and HSCs have phagocyic activity (Mousavi et al., 2005). Early and late endosomes and MVB (multivesicular body) were demonstrated in HSCs; a function of other than action as phagolysosomes has been attributed to MVB (Wake, 1974, 1980). RBP (retinol-binding protein) has been demonstrated in endosomes and MVB by immunoelectron microscopy (Senoo et al., 1990). MVB was proposed as an essential cell organelle for the development of vitamin A-containing lipid droplets in HSCs.
Actin filaments and microtubules are distributed in the periphery and in the core of the cellular processes of HSCs, respectively (Sato et al., 1998). These filaments and microtubules have been proposed to play roles during extension and retraction of the cellular processes (Sato and Senoo, 1998; Imai and Senoo, 1998, 2000). HSCs tend to project a cilium into the perisinusoidal space; the functional significance of the cilium is not yet clearly shown.
Intermediate filaments, desmin and GFAP are markers of HSCs; however, the specific functions of these filaments are not yet thoroughly clarified.
The nucleus of HSCs is oval or elongated in electron microscopy. The nucleus is frequently indented by lipid droplets in the cytoplasm. One or more nucleoli can be seen in a HSC.
Three-dimensional structure of HSCs
HSCs distribute regularly within hepatic lobules. The cell consists of a spindle-shaped or angular cell body and long and branching cytoplasmic processes, which encompass the endothelial tubes of sinusoids (Wake, 1995, 1998). Some processes penetrate the hepatic cell plates (structures formed by PCs. When PCs adhere to each other, they become like a plate.) to reach the neighbouring sinusoids to taper off to several subendothelial processes. Accordingly, a single HSC wraps two or three, sometimes, four sinusoids with long processes. The total length of sinusoids surrounded by a single HSC is 60–140 μm in the rat liver.
The subendothelial processes of HSCs are flat and have three cell surfaces, inner, outer and lateral. The inner one is smooth and adheres to the adluminal (basal) surface of the LSECs. Between the two cells, namely, HSCs and LSECs, the basement membrane components, such as type IV collagen, nidogen and laminin are intercalated. The outer surface, facing the perisinusoidal space (space of Disse), is decorated with short microvillous protrusions. The lateral edges of the subendothelial processes are characteristically studded with numerous spike-like microprojections whose tips make contact with the microvillous facets of the PCs. Thus, HSCs adhere to LSECs through basement membrane components and, on the other hand, make spotty contacts with PCs.
Recently, HSCs have been demonstrated at molecular and morphological levels to adhere to each other by adherens junctions (Higashi et al., 2004).
Relation with other cells in the liver
Relation with LSECs
The hepatic sinusoidal wall is covered by a layer of LSECs, which separates the lumen from the underlying space of Disse (Wisse, 1970, 1972). The hepatic sinusoids are unique among microvascular beds in that the endothelial lining lacks an organized basement membrane and possesses fenestrae without diaphragms (Braet et al., 2001).
Interactions between HSCs and LSECs play an important role in liver homoeostasis (Wake, 1995, 1998). The close association between these two cell types is similar to that observed with pericytes and endothelial cells in other capillary beds. A very close juxtaposition of LSECs and HSCs indicates a supporting function of the latter (Wisse, 1970, 1972).
Conventionally, the perisinusoidal space has been understood as the space between the sinusoids and the PCs, in which the HSCs are present. There is no virtual space between the LSECs and the HSCs; the latter encircle intimately the endothelial tubes in normal mammalian livers. Thus, LSECs and HSCs form an inseparable entity. According to a revised model, the perisinusoidal space is bordered by the LSEC–HSC sheath on one side and PCs on the other (Wake, 1995, 1998). Within this limited space are contained nerve fibres and various ECM (extracellular matrix) components such as collagens (type I, III, V and VI), glycoproteins and proteoglycans. Interactions of HSCs and LSECs have been reviewed recently by Senoo (2007a).
Relation with PCs
In co-culture of HSCs and PCs, an abundant ECM deposits over PCs and between PCs and HSCs (Loreal et al., 1993). This matrix contains laminin, fibronectin and procollagens III and IV. Thus, HSC and PC interaction induces and promotes deposition of ECM. In this co-culture system, functional gap junctions between PCs and HSCs have been demonstrated (Rojkind et al., 1995). PCs in the co-culture system are polarized and maintain a cuboidal shape and have a tendency to form cords.
PCs express nerve growth factor during liver injury and may regulate the number of activated HSCs via induction of apoptosis (Oakley et al., 2003). Reversely, HSCs can regulate proliferation of PCs (Uyama et al., 2002). HSCs can stimulate proliferation of PCs through the HGF (hepatocyte growth factor), extracellular heparan sulfate and heparan sulfate proteoglycan. Thus, the relation between HSCs and PCs is mutually dynamic.
Relation with KCs
Direct contact between HSCs and KCs is rare in normal anatomical situation. The interaction of HSCs with KCs is important in pathological conditions, such as hepatic fibrosis and inflammation.
HSCs exposed to KC culture medium exhibit cellular and nuclear enlargement associated with up to a 3-fold increase in collagen and total protein synthesis (Friedman and Arthur, 1989). The KC-conditioned medium, obtained 48 h after CCl4 administration to rats, stimulates proliferation of cultured HSCs (Armendariz-Borunda, 1989).
Lipopolysaccharide stimulation of HSC/KC coculture, but not HSC monoculture, induces HSC death (Fischer et al., 2002). Activated KC can effectively kill HSC by a caspase 9- and receptor-interacting protein-dependent mechanism. The interaction of HSCs with KCs is complicated and intriguing.
Relation with nerve cells
HSCs are well innervated. During sympathetic nerve action, the neurotransmitter noradrenaline and the cotransmitter ATP cause increases in prostaglandin F2 and prostaglandin D2 release from HSCs (Athari et al., 1994).
Sympathetic hepatic nerves reach the liver through nerve bundles that accompany the large vessels in the liver hilus. They follow the vascular ramification until the portal fields. From there, they penetrate to different extents into the acinus, where they end with varicosities in the space of Disse close to PCs (Reilly et al., 1978) and HSCs [Bioulac-Sage et al., 1990; reviewed by Püschel (2004)].
Emerging evidence suggests that the sympathetic nervous system may play a role in the development of cirrhosis (Oben et al., 2003). Using cultured HSCs, the sympathetic neurotransmitters, norepinephrine and neuropeptide Y, which markedly stimulate the proliferation of activated, myofibroblastic HSCs, have been shown (Oben et al., 2003). Norepinephrine, but not neuropeptide Y, also induces collagen gene expression. Physiologically relevant concentrations of sympathetic neurotransmitters directly modulate the phenotype of HSCs. This suggests that targeted interruption of sympathetic nervous system signalling in HSCs may be useful in constraining the fibrogenic response to liver injury (Oben et al., 2003).
The effect of nerves on perisinusoidal cells has not yet been elucidated. However, the location, shape, morphology and origin of perisinusoidal cells would suggest that they play a role in the haemodynamic regulation of sinusoidal blood flow. It has recently been shown how important PC–NPC communication is in the action of nerves on glucose release by PCs; the cell-to-cell communications may also apply to nerves and sinusoidal cells for the haemodynamic regulation of sinusoidal blood flow (Bioulac-Sage et al., 1990).
The SC unit, ‘stellon’
A single HSC sends branching cytoplasmic processes to two, three and sometimes more neighbouring sinusoids. Conversly, a single LSEC receives cytoplasmic processes from one to three HSCs. On the othe hand, subendothelial processes of a single HSC appear to make contact through their spines with 20–40 PCs. The LSEC–HSC–PC (cell to cell to cell) complex may create a cellular unit in the liver tissue (Wake, 2006). He calls this unit ‘the stellate cell unit’, or in brief, ‘the stellon’. The stellon is not an independent unit, but a loosely overlapping unit with neighbouring units.
The intimate association between HSCs and the neighbouring cell types through the stellon may facilitate intercellular transport and paracrine stimulation by soluble mediators. When HSCs contract or relax, LSECs are contracted or relaxed. This wavy motion of HSCs is conducted to increase the number of LSECs and PCs in a spreading manner. The stellon may be a transducer coupled between the bloodstream and the hepatic parenchyma, which regulates various functions in the liver (Wake, 2006; Senoo, 2007a).
Function of HSCs
Vitamin A handling
Vitamin A (Figure 4) is known to regulate diverse cellular activities such as cell proliferation, differentiation, morphogenesis and tumorigenesis (Blomhoff, 1994; Chawla et al., 2001; Balmer and Blomhoff, 2002; Blomhoff and Blomhoff, 2006). In physiological conditions, HSCs store 50–80% of the total vitamin A in the whole body as retinyl palmitate in lipid droplets in the cytoplasm and regulate both transport and storage of vitamin A (Blomhoff and Blomhoff, 2006).
It had been generally accepted that vitamin A was stored in KCs (reticulo-endothelial cells) belonging to the monocyte–macrophage lineage in the liver (Popper, 1941; Popper et al., 1943), until Wake (1964) demonstrated that HSCs are the main storage site of vitamin A.
More than 95% of vitamin A in the HSC is present in the form of retinyl esters packed together in cytoplasmic lipid droplets. Retinyl esters account for 30–50% of the content of lipid droplets.
The normal reserve of retinyl esters in HSCs represents an adequate supply of vitamin A for most individuals for several weeks or months (Blomhoff et al., 1990). This extensive storage of retinyl esters in HSCs, and the cells' ability to control mobilization of retinol, ensures a steady blood plasma retinol concentration of about 1–2 μM in spite of normal fluctuation in daily intake of vitamin A.
Dietary vitamin A intake, but not triglyceride intake, markedly influences the number and size of HSC lipid droplets (Moriwaki et al., 1988).
Various vitamin A-binding proteins expressed in the inside and outside of cells
Metabolic events are critical to maintaining physiological concentrations of retinoic acid. In the cytoplasm, retinol, retinaldehyde and retinoic acid are associated with retinoid-binding proteins, most of which are approximately 15-kDa proteins belonging to the fatty acid-binding protein/cellular retinol-binding protein family. The ligand binding properties and molecular features of these proteins have been well characterized. Recent experiments have highlighted the importance of the cellular retinoid-binding proteins in controlling the concentration of free retinoids and in directing protein-bound retinoids to key enzymes responsible for their metabolism. For example, CRBP (cellular retinol-binding protein) has been implicated in retinol uptake, retinol esterification, mobilization of retinyl esters and the initial oxidation of retinol to retinaldehyde. The ligands bound to other retinoid-binding proteins have also been shown to be available for enzymatic transformation. The new knowledge of the various ways these cytoplasmic proteins buffer the concentration of ligand, control their distribution and determine their metabolism by specific enzymes is contributing to an improved understanding of the physiological control of retinoid action (Ross, 1993; Noy, 2000).
Multiple types of retinoid-binding proteins that associate with different chemical and isomeric forms of retinoids have been identified in both intracellular and extracellular compartments. Hence, retinoids are found in vivo either associated with cellular membranes or bound to a specific retinoid-binding protein. The parent vitamin A molecule, all-trans-retinol, circulates in blood bound to serum RBP (also called RBP4) (Kanai et al., 1968; Muto et al., 1982). Inside cells, all-trans-retinol and its oxidation product, all-trans-retinal, are associated with one of the three isoforms of cellular retinol-binding proteins (CRBP-I, CRBP-II and CRBP-III (Vogel et al., 2001; Zizola et al., 2008), also called RBP1, RBP2 and RBP3, respectively). All-trans-retinoic acid is found intracellularly bound to one of the two cellular retinoic acid-binding protein isoforms (CRABP-I and CRABP-II). The visual pigment 11-cis-retinal and its precursor, 11-cis-retinol, associate in several cell types in the eye with CRALBP (cellular retinal-binding protein) (Saari et al., 1982). Another ocular protein that can bind retinoids, the IRBP (interphotoreceptor retinoid binding protein), is present in the extracellular space separating the pigment epithelium and the photoreceptor cells. iLBP (intracellular lipid-binding protein) and ERABP (epididymal retinoic acid-binding protein) (Newcomer and Ong, 1990; Ong et al., 2000) are also reported. Retinoid-binding proteins are believed to share a common role, in that they act to solubilize and stabilize their hydrophobic and labile ligands in aqueous spaces. However, a growing body of information indicates that, in addition to this general role, specific retinoid-binding proteins have distinct functions in regulation of the transport, metabolism and action of the particular retinoids with which they associate.
Cellular uptake of vitamin A
The concentration of vitamin A in the bloodstream is regulated within the physiological range in the body (1–2 μM). Uptake mechanism of vitamin A is still controversial. By receptor-mediated endocytosis, the cells take up retinol from the blood, where it circulates as a complex of retinol and a specific binding protein called RBP (Blomhoff, 1994) (Figure 5). Once inside the cell, free retinol has several fates, one of which is reformation of the complex with RBP, and returns to the bloodstream (Blomhoff et al., 1991, 1992a, 1992b; Senoo et al., 2000).
When chylomicron retinyl ester was injected intravenously into normal rats, the radioactivity was cleared from blood with t1/2 approximately equal to 10 min (Blomhoff et al., 1982). Retinyl ester was taken up initially by the liver, which, after 30 min, contained 80–90% of the radioactivity injected. Initially, most of the radioactivity was in PCs, but after 30 min, it disappeared from these cells and reappeared in NPCs. Newly administered retinol is initially taken up by the PCs and subsequently (within 1–2 h) transferred to NPCs. When liver cells were prepared between 5 and 18 h after intraduodenal administration of [3H]retinol, the radioactive retinol was recovered mainly in the HSC. Other liver cells (i.e. PCs, LSECs and KCs) contained only small amounts of [3H]retinol. Newly administered [3H]retinol, which is initially located in the PCs, is transferred to the HSCs and stored there (Blomhoff et al., 1988a). Chylomicron retinyl ester is not transported directly from PCs to HSCs without first undergoing hydrolysis to reinol (Blaner et al., 1987). Retinol delivered to HSCs from RBP is preferentially esterified, and the retinol and RBP complex is handled differently to free retinol by the HSCs (Trøen et al., 1994). Immunoelectron microscopic studies suggest that RBP mediates the paracrine transfer of retinol from PCs to the HSCs and that HSCs bind and internalize RBP by receptor-mediated endocytosis (Senoo et al., 1990, 1993; Malaba et al., 1995). The hypothesis that the cellular uptake of retinol involves the specific interaction of a plasma membrane receptor with RBP at the extracellular surface followed by ligand transfer to cytoplasmic CRBP has been investigated (Sundaram et al., 1998). Optimal retinol uptake is achieved with holo-RBP, the membrane receptor and apo-CRBP. The RBP receptor, through specific interactions with the binding proteins, participates (either directly or via associated proteins) in the mechanism which mediates the transfer of retinol from extracellular RBP to intracellular CRBP. Receptors for RBP have been reported previously (Båvik et al., 1991, 1992, 1993; Smeland et al., 1995), and a new receptor, STRA6 (stimulated by retinoic acid 6), has been reported (Kawaguchi et al., 2007; Pasutto et al., 2007; Blaner, 2007). In bovine retinal pigment epithelium cells, STRA6, a multitransmembrane domain protein, as a specific membrane receptor for RBP, has been identified. STRA6 binds to RBP with high affinity and has robust retinol uptake activity from the retinol–RBP complex. It is widely expressed in embryonic development and in adult organ systems. The RBP receptor represents a major physiological mediator of cellular vitamin A uptake (Kawaguchi et al., 2007). STRA6 mutations define a pleiotropic malformation syndrome representing the first human phenotype associated with mutations in a gene from the ‘STRA’ group (Pasutto et al., 2007). Two unrelated consanguineous families with malformation syndromes sharing anophthalmia and distinct eyebrows as common signs, but differing for alveolar capillary dysplasia or complex congenital heart defect in one and diaphragmatic hernia in the other family, have been reported. How retinol is taken up by cells has been an area of active research, since the first report of a cell-surface receptor for RBP in the mid-1970s. Two recent reports, by Kawaguchi et al. (2007) and Pasutto et al. (2007), provide compelling data regarding the molecular identity and properties of a RBP receptor and open new research avenues for a better understanding of retinoid biology (Blaner, 2007).
Cellular storage of vitamin A
Vitamin A is stored as retinyl esters in the lipid droplets in the cytoplasm of HSCs. PCs play a minor role in liver storages of retinol, and HSCs store more than 90% of liver retinoid (Blomhoff et al., 1988a, 1988b, 1988c). Retinol esterification is performed by enzymes such as ARAT (acyl-CoA:retinol acyltransferase) (Ross, 1982) and LRAT (Ong, 1994). It is now well documented that LRAT is the physiologically important enzyme involved in the esterification of retinol in the liver (Blaner et al., 1990). HSCs are found to contain the highest level of LRAT-specific activity among liver cells. LRAT-positive cells have been found immunohistochemically to be confined in the space of Disse, corresponding with the location of desmin-positive HSC in rodent and human livers (Nagatsuma et al., 2009). LRAT in HSCs is mainly distributed within the rough ER and MVBs. Retinol esterification activity of LI90 (Murakami et al., 1995), a human HSC-like cell line, is demonstrated to be similar to that of primary cultures of rat HSC and higher than that of a human hepatoma cell line (Matsuura et al., 1999).
HSCs had a significantly higher esterification capacity than myofibroblasts. Both LRAT and ARAT participate in retinol esterification in HSCs: LRAT's activity correlates with the vitamin A status, while ARAT depends upon the availability of fatty acyl-CoA and the overall lipid metabolism in HSCs (Fortuna et al., 2001). The fatty acid composition of retinyl-esters suggested a preferential inclusion of exogenous fatty acids into retinyl esters. Hepatic LRAT activity is negligible during vitamin A deficiency, and all-trans-retinoic acid rapidly induces the activity of liver LRAT in retinoid-deficient rats. Retinol and all-trans-retinoic acid strongly induce LRAT activity.
Mouse Lrat gene expression has been disrupted by targeted recombination and a homozygous Lrat knockout (Lrat−/−) mouse is generated (Batten et al., 2004). The Lrat−/− mouse develops normally and has trace levels of all-trans-retinyl esters in the liver, lungs, eyes and blood, whereas the circulating all-trans-retinol is reduced only slightly. The level of CRBPIII has been up-regulated in adipose tissue of Lrat−/− mice (O'Byrne et al., 2005). Large vitamin A lipid droplets that are normally seen within the HSCs were totally absent in Lrat−/− mice. Lrat−/− mice absorb dietary retinol primarily as free retinol in chylomicrons; however, retinyl esters are also present within the chylomicron fraction obtained from Lrat−/− mice. The fatty acyl composition of these chylomicron retinyl esters suggests that they are synthesized via an acyl-CoA-dependent process suggesting the existence of a physiologically significant ARAT (O'Byrne et al., 2005). Retinol bound to CRBPs may be esterified with long-chain fatty acids by the enzyme, LRAT, or may be oxidized to retinoic acid metabolites used in the mechanism of action of vitamin A. HSCs transfected with the vector for sense CRBP-I esterified more retinol than cells transfected with the expression vector for antisense CRBP-I, while retinol esterification in control cells was intermediate (Nilsson et al., 1997). CRBP-I influences retinol esterification in HSCs. Retinyl ester storage is altered in HSCs transfected with CRBP-I. HSCs are the site of the liver β-carotene storage and release, which can be used for retinoic acid production as well as for maintenance of the homoeostasis of circulating carotenoids in periods of low dietary uptake (Martucci et al., 2004).
Secretion of vitamin A from cells
The mechanism of release of vitamin A from HSCs to plasma is also still controversial. HSCs isolated from liver contained RBP mRNA (Andersen et al., 1992). By Western blot analysis, cultivated HSCs were found to secrete RBP into the medium. Cultivated HSCs loaded in vitro with [3H]retinyl ester mobilized radioactive retinol as a complex with RBP. Furthermore, exogenous RBP added to the medium of cultured HSCs increased the secretion of retinol to the medium. These data suggest that HSCs in vivo mobilize retinol directly to the blood and that a transfer to PCs for secretion as holo-RBP is not required. The data suggest that the same mechanism for retinol mobilization may exist in hepatic and extrahepatic SCs. Vitamin A-storing SCs in liver, lungs and probably also in other organs may synthesize their own RBP (or alternatively use exogenous RBP) and mobilize holo-RBP directly to the blood (Andersen et al., 1992). The opposite data have been reported (Sauvant et al., 2001). Primary cultures of HSCs have been demonstrated to express immunoreactivity against RBP when cultured in a medium containing RBP, but were unable to synthesize RBP transcripts and proteins. Pulse and chase-type experiments demonstrated that radioactive retinol was released in culture medium without binding proteins. Inhibition of protein secretion by brefeldin A did not modify quantitatively retinol release. This data ruled out the direct involvement of RBP in retinol mobilization from HSCs. Moreover, HSCs co--cultured with primary isolated PCs displayed an increase of retinol transfer from HSCs to PCs, when they established direct physical contacts, compared with co-cultures without contact. Based on these latter data, a mechanism of retinol mobilization from HSCs via the PCs using retinol transfer through cellular membranes is proposed (Sauvant et al., 2001). Results from knockout mice indicate that RBP is involved (Quadro et al., 1999). The mechanism responsible for retinol mobilization from HSCs has not yet been elucidated in detail.
Lipid droplets and HSC
Lipids present in rat HSC lipid droplets include retinyl ester, triglyceride, cholesteryl ester, cholesterol, phospholipids and free fatty acids. Retinyl ester and triglyceride are present at similar concentrations, and together, these two classes of lipid account for approximately three quarters of the total lipid in HSC lipid droplets (Blaner et al., 2009). The function of ADRP (adipose differentiation-related protein) is known to be the uptake of long-chain fatty acids and formation of lipid droplets in lipid-accumulating cells. ADRP overexpression induced the formation of lipid droplets in activated HSCs (Fukushima et al., 2005).
Two types of vitamin A–lipid droplets have been described in HSCs. Type I lipid droplets are membrane-bound (surrounded with unit membrane, namely, phospholipid bilayer), whereas type II lipid droplets are not membrane-bound (not surrounded with unit membrane) (Wake, 1974, 1975, 1980; Yamamoto and Ogawa, 1983; Yamada et al., 1987; Geerts et al., 1990; Blomhoff and Wake, 1991; Friedman, 2008a). The relation between the two lipid droplets has not yet been elucidated. What kinds of molecules are involved in the formation of them is poorly understood. According to Wake (1974,1975,1980), type I lipid droplets are derived from MVBs, which belong to the endosome–lysosome system, and type II lipid droplets are derived from type I lipid droplets after losing their surrounding unit membrane. Matsuura et al. (1993) reported that MVBs are important organelles for retinoid storage as lipid droplets. On the other hand, Yamamoto and Ogawa (1983) reported that type I lipid droplets are the autophagolysosome of type II lipid droplets. As mentioned above, LRAT, an enzyme responsible for all retinyl ester synthesis within the liver, is required for HSC lipid droplet formation, since LRAT-deficient mice completely lack HSC lipid droplets. When HSCs become activated in response to hepatic injury, the lipid droplets and their retinoid contents are rapidly lost. HSC lipid droplets are suggested to be specialized organelles for hepatic vitamin A storage and that loss of HSC lipid droplets may contribute to the development of hepatic disease (Blaner et al., 2009).
Nuclear receptors and HSC
As mentioned above, HSCs store vitamin A as a form of retinyl ester within the lipid droplets of their cytoplasm. This characteristic feature of HSC led to the idea that HSC functions are regulated by two types of transcriptional factors, RARs (retinoic acid receptors) and PPARs (peroxisome proliferator-activated receptors), both belonging to the nuclear receptor superfamily, because RARs mediate most physiological functions of vitamin A and PPARγ is known to be a master gene for adipose cell differentiation.
Quiescent HSCs store not only vitamin A, but also lipids such as triglycerides. Upon activation, both vitamin A and trigycerides are depleted from HSC. PPARγ protein is present in quiescent HSCs, and it decreases drastically during HSC activation (Everett et al., 2000; Galli et al., 2000).
Vitamin A (retinol) is hydrolysed to retinal by retinol dehydrogenases and further hydrolysed to all-trans-retinoic acid, an active form of retinoids, by retinal dehydrogenases (Napoli, 1999). Retinoic acid binds to the RARα, β and γ within the nucleus to enhance their transcriptional activity.
Presence of mRNA for RARα, β and γ in HSC was reported (Weiner et al., 1992; Ohata et al., 1997; Hellemans et al., 2004). Protein presence was also evidenced by EMSA (electrophoretic mobility shift assay) (Ohata et al., 1997; Hellemans et al., 2004). Recently, we have reported that RAR expression is suppressed in quiescent HSCs at the post-transcriptional level, and only activated HSCs are responsive to retinoids (Mezaki et al., 2007, 2009). We documented that RARα proteins show a punctate or speckled cytosolic distribution during rat HSC activation in vitro (Mezaki et al., 2009). In those studies, we used a rabbit polyclonal antibody (sc-551) raised against a peptide targeting the C-terminus of human RARα (Figures 6A–6C). For confirmation, we used a mouse monoclonal antibody (Rα10) raised against a synthetic peptide corresponding to residues 444–462 of mouse RARα, a sequence which is identical to that in the rat (441–459). Almost the same distribution of RARα proteins was observed with this monoclonal antibody (Figures 6D–6F), verifying the speckled cytosolic distribution of RARα in activated HSCs. The biological meaning of this phenomenon is now under investigation in our laboratory.
The inhibitory effect of estrogen on activation of rat HSCs is reported (Shimizu et al., 1999, 2003). This effect is mediated by oestrogen receptor β subtype because oestrogen receptor α is negative in both quiescent and activated rat HSCs (Zhou et al., 2001).
The presence of GR (glucocorticoid receptor) in HSC is demonstrated (Raddatz et al., 1996).
VDR (vitamin D receptor) content is very low in liver compared with those in small intestines and colon (Sandgren et al., 1991). However, HSC, as well as other NPCs such as LSECs and KCs, harbour considerable amounts of VDR mRNA and protein (Gascon-Barre et al., 2003).
Bile acids work not only as a detergent, which facilitates dietary fat absorption, but also as ligands for FXR (farnesoid X receptor) (Chiang, 2002). FXR up-regulates BSEP (bile salt export pump) in bile canaliculi and down-regulates CYP7A1 (cholesterol 7α-hydroxylase), the rate-limiting enzyme for bile acid synthesis, thus reducing a concentration of toxic bile acids within the liver (Chiang, 2002). Besides this bile acid homoeostatic function of FXR in PCs, there exists FXR in HSCs, playing an antifibrotic role in porcine serum-administrated, bile duct-ligated and CCl4-induced hepatic fibrosis (Fiorucci et al., 2004, 2005).
PXR (Pregnane X receptor), or SXR (steroid X receptor) in human, has been categorized in an orphan nuclear receptor, and its ligands include pregnane steroids, bile acids and drug ligands such as rifampicin (Wright, 2006). Recently, vitamin K is also shown to be a PXR ligand (Tabb et al., 2003; Igarashi et al., 2007). In rodents, pregnenolone-16α-carbonitrile, a PXR activator, inhibits liver fibrogenesis in spite of the absence of PXR expression in HSC (Marek et al., 2005). However, a recent report tells that PXR is actually expressed in quiescent rodent HSCs, but it suddenly decreases upon activation (Wright, 2006), indicating that the antifibrogenic effect of pregnenolone-16α-carbonitrile might be mediated by PXR. In human, PXR is expressed constitutively throughout HSC activation, and rifampicin, a species-specific PXR activator, inhibits HSC activation as well (Haughton et al., 2006). Functions of the nuclear receptors in HSCs will be clarified in more detail in the near future.
Production and degradation of ECM components by HSCs
In pathological conditions such as liver cirrhosis, HSCs lose vitamin A, proliferate vigorously and synthesize and secrete a large amount of ECM components such as collagen, proteoglycan, glycosaminoglycan and glycoprotein. The structure of the cells also changes from star-shaped HSCs to that of fibroblast-like cells or myofibroblasts (Majno, 1979) with well-developed rough-surfaced ER and Golgi apparatus (Senoo and Wake, 1985; Blomhoff and Wake, 1991; Senoo et al., 1997; Sato et al., 2003). In order to elucidate cell type(s) responsible for collagen metabolism among NPCs in the liver, collagen production by HSCs, KCs and LSECs was analysed (Senoo et al., 1984). HSCs were found to produce collagen on day 8 in primary culture, although collagen production was not induced at an earlier stage of culture (day 2). Capability of collagen production by cells was retained in the secondary culture, suggesting that the HSC is a candidate cell responsible for collagen production. KCs and LSECs produced little collagen either on day 2 or day 8 in primary culture under the conditions employed.
Types of collagen produced by HSCs in secondary culture were analysed by fluorography after SDS/polyacrylamide slab gel electrophoresis under non-reducing and reducing conditions (Figure 7; Senoo et al., 1984). Type I collagen is the major component synthesized (Figure 7, lane a). Minor components include type III collagen, which remained at the γ-region under non-reducing conditions (Figure 7, lane a), but migrated to the α1-region after reduction (Figure 7, lane b), and type IV collagen, which remained slightly below the origin under non-reducing conditions (Figure 7, lane a), but migrated to region slightly lower than the β-region after reduction (Figure 7, lane b). All these bands were susceptible to purified bacterial collagenase (Figure 7, lanes c and d). Quantitation of these collagen bands by densitometry indicated that the percentage of type I, type III and type IV collagens was 88.2%, 10.4% and 1.4%, which is consistent with an observation on collagen types in human liver cirrhosis.
Thus, HSCs have been shown to be able to produce ECM components.
Degradation of ECM by HSCs
MMPs (matrix metalloproteinases) and TIMP (tissue inhibitor of metalloproteinases) have been reported to be synthesized by HSCs (Iredale et al., 1998; Friedman, 2000; Benyon and Arthur, 2001; Pinzani and Marra, 2001; Arthur, 2002; Friedman and Arthur, 2002; Poynard et al., 2002). Recent reports indicate that a differential expression of MMP activity, hence the remodelling of ECM components, is dependent on the substratum used for the culture of HSCs (Li et al., 1999; Wang et al., 2003). HSCs can take up collagen fibrils by enodocytosis (Mousavi et al., 2005) and phagocytosis (Higashi et al., 2005a) for degradation.
Reversible regulation of morphology, proliferation and function of the HSCs by three-dimensional structure of ECM
Tissues are not composed solely of cells. A substantial part of their volume is intercellular space that is largely filled by an intricate network of macromolecules constituting ECM. This matrix comprises a variety of polysaccharides and proteins that are secreted locally and assembled into an organized meshwork (Alberts et al., 2008). ECM was considered to serve mainly as a relatively inactive scaffolding to stabilize the physical structure of tissues until recently. But now, it is clear that ECM plays a far more active complex role in regulating the behaviour of the cells that contact, influencing their morphology, development, migration, proliferation and functions (Hata and Senoo, 1992; Senoo and Hata, 1993, 1994a, 1994b; Senoo et al., 1996).
We reported that HSCs proliferated better and synthesized more collagen on type I collagen-coated culture dishes than on polystyrene dishes (Senoo and Hata, 1994a). We also demonstrated that the HSCs formed a mesh-like structure, proliferated slowly and synthesized a small amount of collagen on Engelbrech–Holm–Swarm murine tumour ECM containing the basement membrane components (Matrigel) (Senoo and Hata, 1994a), a gel consisting largely of laminin, type IV collagen, heparan sulfate proteoglycan and nidogen (Kleinman et al., 1986).
Uptake of [3H] retinol into the cells and esterification into retinyl palmitate was enhanced when the cells were cultured on type IV collagen-coated dishes (Davis and Vucic, 1989). The Matrigel was reported to be able to maintain the differentiated phenotype, such as storage of lipids in cultured HSCs (Friedman et al., 1989b). Types of collagen synthesized by HSCs (Davis et al., 1987; Friedman et al., 1989a) and responses of the HSCs to cytokines (Davis, 1988) are also modulated by ECM. Our studies and other works clearly show that ECM can regulate morphology, proliferation and functions of HSCs.
We reported that regulation of morphology, proliferation and collagen synthesis of the HSCs by three-dimensional structure of ECM is reversible (Imai and Senoo, 1998, 2000; Kojima et al., 1998, 1999; Imai et al., 2000b). Cultured HSCs exhibit mainly three types of morphology according to the substratum used for culture (Sato and Senoo, 1998; Sato et al., 2003). The cells displayed a drastic change in cell shape accompanying the development of cytoplasmic processes by culturing in the presence of type I and type III collagen gel as a substratum (Figure 8). On the other hand, HSCs have a flattened, fibroblast-like shape with well-developed stress fibres on the polystyrene surface, as well as on the lysine- or aminoalkylsilane-coated surface, and they show neither signs of cell spreading nor elongation of cell processes and maintain a round shape in or on Matrigel. Therefore, the ECM components can differentially regulate the morphology of HSCs, as well as at least a part of in vivo function (Senoo et al., 1996).
The cellular processes of the HSCs were demonstrated to be extended and retracted according to the ECM and speculated to have important functions in transport and storage of vitamin A and transport of metalloproteinases (Sato et al., 1997, 1998, 1999, 2001b, 2003, 2004). These data also indicate that the HSCs are not static, but dynamic, in the changeable three-dimensional structure of ECM in the space between PCs and LSECs. The dynamic movement of cultured HSCs was analysed with a video-enhanced optical microscopy (Miura et al., 1997; http://www.med.akita-u.ac.jp/˜kaibo2/default.html). When cultured on polystyrene surface, the HSCs spread well, flattened with extensive stress fibres. The cell surface ruffling activity of filopodia and lamellipodia was prominent (Figure 9), reflecting weak adhesion to the substratum. All filopodia remained dynamic throughout the 4-h recording and extended and retracted repeatedly. Within 1 h after inoculating in or on type I collagen gel, the HSCs began to extend cellular processes, and the cellular processes appeared to adhere to and extend along type I collagen fibres. After repeated extension and retraction of cellular processes, HSCs displayed a number of long cellular processes with distal fine branches by 4-h culture on type I collagen gel. The cellular processes also extended in or on type III collagen gel, but not in type IV collagen-coated dishes or on Matrigel. Such different responses may be functionally important, since HSCs, as mentioned above, extend their processes in vivo along the perisinusoidal space of Disse containing mainly type I and type III collagen, whereas these cells display smooth cell surfaces in contact with the imperfect basal membrane components of LSECs (Wake, 1980). In vivo morphology of HSCs might be regulated by three-dimensional structure of ECM.
Cell surface integrin interactions with ECM are central to the migration, localization and function of several types of cells within tissues (Kornberg et al., 1991; Hynes, 1992; Jones et al., 1993; Rosales et al., 1995; Roskelley et al., 1995). Human HSCs have been reported to express integrin α1β1, α2β1, α(V)β1 and α6β4 (Carloni et al., 1996). The extension of the processes was inhibited by the addition of anti-integrin α2 antibody or oligopeptide DGEA (Sato et al., 1998, 1999), which sequence in the type I or type IV collagen molecules has been reported to be responsible for cell surface integrin binding (Staatz et al., 1991). Therefore, the process elongation for HSCs appears to depend on integrin α2β1 binding to extracellular collagen fibrils. Since no process elongation is seen in the culture on type I or type III collagen-coated surface (monometric collagen), a native fibrillar form of interstitial collagen is a prerequisite for process elongation, as reported for integrin binding to a native form of collagen fibrils but not to monometric collagen in other cell types (Grab et al., 1996; Mercier et al., 1996; Vogel et al., 1997).
Retraction of elongated processes is induced by the treatment with herbimycin A and staurosporin, a protein tyrosine kinase inhibitor and a general protein kinase inhibitor, respectively, or with wortmannin, a PI3K (phosphatidylinositol 3-kinase) inhibitor (Miura et al., 1997, Sato et al., 1998; Kojima et al., 1998). When HSCs are inoculated in the presence of these inhibitors, the induction of process elongation is also inhibited, suggesting that both the induction and sustenance of processes depend on successive signalling events including protein and/or PI phosphorylation. Protein tyrosine phosphorylation was enhanced in HSCs cultured on type I collagen gel, compared with the culture on the polystyrene surface (Kojima et al., 1998). In HSC culture, PI3K has been found to be involved in ERK (extracellular-signal regulated kinase) activation by PDGF (platelet-derived growth factor) (Marra et al., 1995). Interestingly, H-7 or hypericin, a specific inhibitor for protein kinase C, shows only a partial effect on the induction and sustenance of cytoplasmic processes in HSCs cultured on interstitial collagen gel, but they cause a significant morphological change in HSCs cultured on a polystyrene or molecular-coated surface accompanying the F-actin degradation. These results suggest that a distinct pathway involving protein tyrosine kinase, PI kinase and/or protein serine/threonine kinase is responsible for the cell spreading or the process elongation. It has been well documented that the reorganization of F-actin is triggered by integrin binding to ECM components, followed by signal transduction mechanisms including protein phosophorylation (Hynes, 1992; Yamada and Miyamoto, 1995). In HSC culture, interstitial collagen signals, triggered by cell surface integrin, appear to finally result in the reorganization of microtubules, as well as F-actin assembly (Miura et al., 1997; Sato et al., 1998).
Cytoskeleton organization in cellular processes of HSCs
The role of cytoskeleton organization in the sustenance of the process structure have been demonstrated by a video recording of the HSC culture after the addition of cytoskeleton-degrading drugs (Miura et al., 1997; http://www.med.akita-u.ac.jp/˜kaibo2/default.html). Colchicine treatment induces the retraction of the elongated processes. The effects are dose- and time-dependent, and almost all cells are changed to round shapes within a few hours in the presence of 1 μM colchicine. The cold treatment at 4°C, which is known to induce the degradation of a cold-labile form (dendritic type) of microtubules (Rosette and Karin, 1995) also causes the retraction of elongated process within 3 h. The elongated processes are also partly retracted after the treatment with cytochalasin B (Sato et al., 1998). Although the effects of cytochalasin B appear to be weaker than that of colchicine, almost all cells are changed to a round shape after overnight treatment with cytochalasin B. Dual fluorescence staining of microtubule and F-actin in HSCs revealed that the core of processes consists of microtubules, while the periphery contained F-actin (Miura et al., 1997; Sato et al., 1998).
The elongated front of the processes displays a similar filopodia and lamellipodia containing F-actin structure and microtubules to those of the growth cone of the extending neurites. These results, in addition to the effects of cytoskeleton degradation, suggest that the elongation of cellular processes results from F-actin and the microtubule assembly in the HSC culture.
Interstitial collagen gel induces the process of elongation, mimicking the in vivo cell shape. It has been reported that interstitial type I collagen gel, used as a substratum, alters the HSC function to that of the quiescent, normal in vivo phenotype, such as the slight proliferation activity and the suppressed ECM production (Senoo et al., 1996). HSCs are not able to spread out or elongate processes on or in Matrigel (Miura et al., 1997; Sato et al., 1998), but display rather normal in vivo functions (Friedman et al., 1989a; Senoo and Hata, 1994a, 1994b). Therefore, HSCs require the basement membrane components for a normal, inactivated phenotype, whereas they need interstitial collagen fibres for the process of elongation, as mentioned above. Such an interpretation is consistent with the in vivo ECM status around HSCs in liver tissue.
HSCs cultured on interstitial collagen gel also expressed MMP 1 (Sato et al., 1998), which was demonstrable in cytoplasmic processes. HSCs are the primary ECM-producing cells in NPCs in liver fibrosis, and during liver injury, HSCs undergo activation accompanying cell proliferation and enhanced fibrogenesis. Phenotypes including morphology of HSCs can be regulated by the use of ECM components.
Stimulation of proliferation of HSCs and tissue formation of the liver by a long-acting vitamin C derivative
A long-acting vitamin C derivative, Asc 2-P (l-ascorbic acid 2-phosphate), was found to stimulate cell proliferation, collagen accumulation and tissue formation (Hata and Senoo, 1989; Kurata et al., 1993). On the basis of this discovery, Asc 2-P was added to the medium in which HSCs were cultured (Senoo and Hata, 1994a). The cells in the medium supplemented with Asc 2-P stretched better than the cells in the control medium. Asc 2-P stimulated cell proliferation and collagen synthesis of the HSCs and formation of the liver tissue-like structure in co-culture of PCs and fibroblasts (Senoo et al., 1989).
Effects of retinoid to HSC function
Effects of vitamin A on HSC function both in vivo and in vitro are still controversial. Antagonistic relationship between capability of storage of vitamin A lipid droplets and ability of ECM production in HSCs has been reported (Senoo and Wake, 1985; Imai et al., 2000a). Okuno et al. (1990) reported that acyclic retinoid (polyprenoic acid) reduced the development of hepatic fibrosis induced by CCl4 in rats. In rats, it shows that vitamin A status, reflected by hepatic retinoid content (retinol and retinyl esters), modulates the development of hepatic fibrosis induced by CCl4. In rats with low hepatic retinoid levels, CCl4-induced liver fibrosis was more pronounced than in rats with sufficient hepatic retinoid levels. Retinoids modulate collagen synthesis and deposition irrespective of the degree of hepatocellular necrosis induced by CCl4. The reduction of retinoid levels in HSCs and fibroblast-like cells by an enhanced secretion of retinol from the liver into the circulation during CCl4 treatment may stimulate the transformation of these cells to fibroblasts and, in this way, contribute to fibrogenesis of the liver (Seifert et al., 1994a, 1994b). Thus, vitamin A can modulate HSC proliferation in vitro (Davis and Vucic, 1989). However, the effects of vitamin A to the HSC function are not yet thoroughly clarified.
Retinol (1 μM) and retinoic acid (1 μM) supplementation of primary rat HSC cultures inhibited the spontaneous increase in collagen synthesis associated with HSC activation. Extracellular retinoids suppress HSC-activated collagen synthesis and cell proliferation and support the interpretation that retinoids, themselves, are regulatory factors in maintenance of the HSC in its native, differentiated state (Sato et al., 1995).
In HSCs isolated from rat fibrotic livers, the amount of RXR-α mRNA is greatly reduced. However, the effectiveness of retinoids in the treatment of liver fibrosis is controversial. Increasing the expression levels of RXR-α in livers improve the response of liver fibrosis to retinoid treatment. Hepatic fibrosis of RXR-α-transfected group, tapered off remarkably (Chen et al., 2004).
Retinoic acid suppresses α2(I) collagen expression in HSCs through the binding of RARβ and RXRα to RAREs (retinoic acid response elements) in the α2(I) collagen promoter (Wang et al., 2004b). This study determined the influence of co-activators and co-repressors to RARβ and RXRα on the regulation of the α2(I) collagen promoter. In the presence of retinoic acid, the co-activators SRC-1 (steroid receptor coactivator-1) and GRIP-1 (growth hormone receptor interacting protein-1) augment, while the corepressor N-CoR abolishes the suppressive effects of RARβ and RXRα on α2(I) collagen promoter activity.
ATRA (all-trans-retinoic acid) exerted a significant inhibitory effect on the synthesis of procollagens types I, III and IV, fibronectin and laminin of cultured HSCs, but did not influence HSC proliferation, whereas, 9RA (9-cis-retinoic acid) showed a clear but late effect on proliferation. 9RA increased procollagen I mRNA 1.9-fold, but did not affect the expression of other matrix proteins. Results showed that ATRA and 9RA exert different, often contrary, effects on activated HSCs. These observations may explain prior divergent results obtained following retinoid administration to cultured HSCs or in animals subjected to fibrogenic stimuli (Hellemans et al., 1999).
Opposite results have been reported as follows (Vollmar et al., 2002). Although there is strong implication that retinoids regulate HSC proliferation and collagen synthesis, results from in vivo studies on the relationship between vitamin A and liver fibrosis are conflicting. In animals with high dietary hepatic retinoid levels, liver fibrosis was more pronounced and was associated with an increased CCl4 toxicity resulting in high mortality (73%). It was further associated with decreased bile flow and increased PC damage. CCl4 reduced hepatic retinoid levels in high vitamin A diet-fed animals, but restored hepatic retinoid levels in animals fed with a vitamin A-deficient diet, implying major interference of vitamin A metabolism with hepatotoxic agents such as CCl4. Low vitamin A feeding did not modulate liver fibrogenesis and caused no mortality. These results show that the vitamin A status of the liver plays an important role in liver fibrogenesis. While dietary vitamin A shortage does not promote liver fibrogenesis, high levels of vitamin A have the potential to increase systemic and hepatic toxicity of CCl4. Thus, the narrow therapeutic window for nutritional vitamin A substitution must take into account that liver fibrotic patients may display enhanced susceptibility to the adverse effects of vitamin A.
Ontogeny and phylogeny of HSCs
In mammalian embryos, the liver arises as a ventral outgrowth – hepatic diverticulum – from the caudal or distal part of the foregut early in the fourth week (Elias, 1955; Du Bois, 1968; Severn, 1971, 1972; Fukuda-Taira, 1981; Zaret, 2001; Moore et al., 2008; Varet and Grompe, 2008). FGFs (fibroblast growth factors) secreted by the developing heart, interact with the bipotential cells and induce formation of the hepatic diverticulum. The diverticulum extends into the septum transversum, a mass of splanchnic mesoderm between the developing heart and midgut. The septum transversum forms the ventral mesentery in this region. The proliferating endodermal cells give rise to interlacing cords of PCs and to the epithelium lining of the intrahepatic part of the biliary apparatus. The hepatic cords anastomose around endothelium-lined spaces, the primordial of the hepatic sinusoids.
Liver mesothelial cells give rise to mesenchymal cells, which intermingle between the growing hepatoblast cords and become incorporated to the sinusoidal wall, contributing to both LSEC and HSC populations. HSCs, which likely derive from preexisting mesenchymal cells of the septum transversum, slowly differentiate, as shown in the progressive increase in the number of intracytoplasmic lipid droplets, a characteristic feature of fully mature HSCs (Collardeau-Frachon and Scoazec, 2008). The developing liver is vascularized through a complex process of vasculogenesis that leads to the differentiation of the sinusoids. Both intrinsic or extrinsic mesothelium-derived cells have the developmental potential to contribute to the establishment of liver sinusoids (Pérez-Pomares et al., 2004). At day of 10 of gestation in mice and rats, or at 5 weeks of gestation in humans, the hepatic cord grow into the mesenchymal tissue of the septum transversum, and the primitive sinusoid-like structure is simultaneously observed between the liver cell cords (Enzan et al., 1997). In the margin of the growing liver primordium, mesenchymal cells in the septum transversum are trapped in the subendothelial space. These subendothelial cells are at the early stages of organogenesis and become progenitors of the HSCs. Several reports have suggested that HSCs are derived from the neural crest, since HSCs express glial fibrillary acidic protein (GFAP; Gard et al., 1985; Buniatian et al., 1996; Neubauer et al., 1996; Niki et al., 1996) and nestin (Messing, 1999; Niki et al., 1999), and that neural crest cells can differentiate into myofibroblasts expressing αSMA, which is a marker of activated HSCs (Carlson, 1999). HSCs are reactive to the Golgi staining, which is usually used for neurofilament staining (Wake, 1980).
HSCs also display unique characteristics of gene expression similar to that of neural cell types, e.g. expression of N-CAM (Knittel et al., 1996), synaptophysin (Cassiman et al., 1999), neurotrophins and neurotrophin receptors (Cassiman et al., 2001; Trim et al., 2000), GFAP or microtubule associated protein 2 (Sato et al., 2001b). These findings also suggest the neural crest origin of HSCs. Activated HSCs, a major ECM-producing cell type in liver fibrogenesis, shows common phenotypic features to smooth muscle cells and myofibroblasts, both of which are originated from the neural crest and express vimentin, desmin and αSMA.
However, yellow fluorescent protein that is expressed in all neural crest cells and their derivatives has not been detected in the developing liver or in HSCs from embryonic day 11.5 until adulthood of mice (Cassiman et al., 2006). This finding suggests that HSCs do not descend from the neural crest and, therefore, may derive from the septum transversum mesenchyme (Kalinchenko et al., 2003; Varet and Grompe, 2008), from endoderm or from the mesothelial liver capsule.
Recently, haematopoietic origin of HSCs has been proposed (Baba et al., 2004; Suskind and Muench, 2004; Miyata et al., 2008). In mouse cirrhotic liver, the bone marrow contributes significantly to HSCs (68%) and myofibroblast (70%) populations (Russo et al., 2006). However, in response to the bile duct ligation-induced liver injury of mice, HSCs do not originate in the bone marrow, and collagen-producing fibrocytes are recruited from the bone marrow to damaged liver (Kisseleva et al., 2006). Contribution of bone marrow to HSC transdifferentiation might be different between the disease models. Additional works might be necessary for these studies.
A hypothesis that HSCs are a type of oval cell and capable of generating PCs to regenerate injured livers was evaluated (Yang et al., 2008). The data suggest that HSCs are a type of oval cell that transitions through a mesenchymal phase before differentiating into PCs during liver regeneration. This report suggests that HSCs and PCs have the same progenitor, but more researches are needed to conclude that this hypothesis is correct or not.
The phenotypic characteristics of HSCs in rat fetal livers, in which hepatic development and haematopoiesis occur in parallel, have been determined, using a strategy focused on vitamin A (Kubota et al., 2007). These results demonstrated that the vitamin A autofluorescence-positive cells are fetal HSCs with extensive proliferative activity. Furthermore, the vitamin A autofluorescence positive cells strongly express hepatocyte growth factor, stromal-derived factor-1, and Hlx (homeobox transcription factor), indicating that they play important roles for hepatic development and haematopoiesis. Researchers have done studies as mentioned above, but ontogeny of HSCs is still controversial (Geerts, 2004).
To demonstrate the origin of SCs in phylogeny, SCs in non-mammalian and mammalian vertebrates have been investigated. We examined vitamin A and vitamin A-storing cells in arrowtooth halibut (Atheresthes evermanni Jordan et Starks) (Yoshikawa et al., 2006), lamprey (Lampetra japonica) (Wake and Senoo, 1986; Wake et al., 1987; Wold et al., 2004; Higashi et al., 2005b), ascidian (Halocynthia roretzi) (Irie et al., 2004) and polar animals (Higashi and Senoo, 2003). In the arrowtooth halibut, the highest concentration of stored vitamin A is in vitamin A-storing cells in the pyloric caecum, a teleost-specific organ protruding from the intestine adjacent to the pylorus. Considerable amounts of vitamin A were also stored in the cells in the lamina propria mucosae of the intestine and HSCs in the liver. In lamprey, retinoid is stored in retinoid-storing cells in the intestines, liver (HSCs), kidneys, gills and heart. In ascidian, retinal is the essential form of retinoid for storage, and no SCs are observed (Irie et al., 2004). Thus, the distribution of SCs with retinoid-storing capacity differs between non-mammalian vertebrates and mammals, suggesting that the SCs appeared in lamprey, and vitamin A-storing site has shifted and concentrated from the whole body to the liver during vertebrate evolution.
In arctic animals such as polar bears and arctic foxes, a large amount of vitamin A is stored in the HSCs. However, other organs store only a little amount of vitamin A (Higashi and Senoo, 2003).
Xenobiotics (such as PCBs and dioxins) may reduce the threshold of vitamin A toxicity (Nilsson et al., 2000), and both vitamin A and fat-soluble xenobiotics have a tendency to accumulate in the food chain (Dewailly et al., 1989; Barrie et al., 1992; Jarman et al., 1992; Muir et al., 1992; Holden, 1998; Wiig et al., 1998; Skaare et al., 2001). Kidney total vitamin A, which may be used as a biomarker for retinoid-related toxicity or excess, in polar bear and bearded seal, was below 1% of their liver value, which is in the normal range for most animals (Senoo et al., 2004). Arctic fox and glaucous gull, however, had kidney levels of about 9% and 42% of the liver values, respectively. This increased kidney concentrations and decreased capacity for storage in HSCs of total vitamin A in arctic fox and glaucous gull are most likely signs of vitamin A toxicity that deserve attention. Nuclear deviation has been reported in PCs on sinusoidal surface in arctic animals (Sato et al., 2001a). These data are alarming and have not been observed previously in free-living animals.
Vitamin A storage in HSCs in polar animals reflects the environmental and climate changes in the arctic area.
Relatives of HSCs
Previous studies using fluorescence microscopy, TEM (transmission electron microscopy) and electron microscopic autoradiography showed that cells that stored vitamin A distribute in extrahepatic organs, namely, lungs, digestive tract, spleen, adrenal glands, testis, uterus, lymph nodes, thymus, bone marrow, adventitia of the aorta, lamina propria of the trachea, oral mucosa, urinary bladder and tonsils (Popper, 1941, 1944; Tanaka, 1955; Kusumoto and Fujita, 1977; Yamamoto et al., 1978; reviewed by Wake, 1980; Yorifuji et al., 1980; Yamada, 1982; Nagy et al., 1997; Matano et al., 1999). Morphology of these cells is similar to that of fibroblasts. These cells emanate autofluorescence of vitamin A and contain lipid droplets in the cytoplasm. These cells and HSCs might form the SC system that regulates homoeostasis of vitamin A in the whole body. Extrahepatic SCs also can synthesize and secrete ECM components. PSCs (pancreatic SCs) (Apte et al., 1998; Bachem et al., 1998), one sort of extrahepatic SCs, are now considered to be responsible for the induction of chronic pancreatitis and pancreatic fibrosis. These extrahepatic SCs are now to be targets of the treatment of inflammation and organ fibrosis.
Although HSCs are particularly important for vitamin A storage in many animals, some other cells in organs such as lungs, kidneys and intestines of higher vertebrates may accumulate retinyl esters in lipid droplets after ingestion of large amounts of vitamin A (Blomhoff and Wake, 1991; Nagy et al., 1997). Such extrahepatic storage of retinyl esters may even be an important local supply of vitamin A to organs with a huge demand, for example, retinyl ester storage in retinal pigment epithelial cells as a prerequisite for normal visual function.
In human, as mentioned above, PSCs, one sort of extrahepatic SCs, are now considered to be responsible for the induction of chronic pancreatitis and pancreatic fibrosis (Apte et al., 1998; Bachem et al., 1998). In the pancreas sections, stellate-shaped cells (SCs) are observed; these are positive for desmin and GFAP and negative for αSMA. PSCs have a periacinar and interlobular distribution. They comprise 3.99% of all pancreatic cells. The PSCs from rat pancreas grow rapidly in culture. PSCs cultured for 24 h have an angular appearance, contain lipid droplets manifesting positive vitamin A autofluorescence and stained positively for desmin, but negatively for αSMA. At 48 h, PSCs are positive for αSMA (Apte et al., 1998). During primary culture, PSCs change from a quiescent vitamin A-storing phenotype to a highly synthetic myofibroblast-like cell expressing αSMA and desmin and biosynthesize collagen types I and III, fibronectin and laminin. PSCs are now to be targets of the treatment of inflammation and organ fibrosis (Wells and Crawford, 1998; Pinzani, 1999, 2006).
Phagocytosis by PSCs has been reported (Shimizu et al., 2005). PSCs act as resident phagocytic cells, and CD36 promotes troglitazone-induced phagocytic activity via PPAR-γ transactivation. Enhancement of phagocytic activity may provide an important approach to the treatment of pancreatic diseases.
The expression of STAP (stellate cell activation-associated protein)/cytoglobin (Kristensen et al., 2000; Kawada et al., 2001; Maeda et al., 2003) was recently confirmed in all splanchnic vitamin A-storing cells including HSCs in normal conditions (Nakatani et al., 2004). In the hepatic fibrous lesion, the expression of STAP/cytoglobin has been shown to be in activated HSCs and myofibroblasts, which have synthesized ECM. Furthermore, splanchnic vitamin A-storing cells have been reported to be distributed in the kidneys. The contribution of vitamin A-storing cells to renal fibrosis by focusing on STAP/cytoglobin has been reported (Kida et al., 2007). In adult mice, STAP-positive cells (splanchnic vitamin A-storing cells) significantly increased in kidneys after unilateral ureteral obstruction suggesting that splanchnic vitamin A-storing cells contribute to renal fibrogenesis in the obstructed kidney.
Physiologically, the most important components of the lungs are the thin-walled, terminal saccular compartments, the pulmonary alveoli (Fawcett, 1986). The interstitium of the alveolar septa is the tissue between the two layers of pulmonary epithelium lining adjacent alveoli. It includes a very close-meshed network of capillaries, pericytes, septal cells (also called as interstitial fibroblasts, interstitial contractile cells, interstitial myofibroblasts or lipid-containing interstitial cells), mast cells, monocytes and occasional lymphocytes. The septal cells found in the connective tissue space of the alveolar septa are closely applied to the capillary endothelium and to the alveolar epithelium with elongated cell processes and contain lipid droplets that incorporate 3H-labelled vitamin A (Yamada, 1982). CRBP, CRABP and several isoforms of nuclear retinoic acid receptors are expressed in these cells (Chytil, 1992; Redlich et al., 1996; Ross, 2004). The septal cells that store vitamin A–lipid droplets in the cytoplasm are the LSCs (lung SCs).
The isolated rat LSCs grown in vitro retain well the overall morphology characteristic of the vitamin A-storing cells found in lung tissues (Okabe et al., 1984).
Thus, the septal cells that store vitamin A–lipid droplets in the cytoplasm and synthesize and secrete ECM components are the LSCs, one sort of extrahepatic SCs.
Smoking habit effects the storage of vitamin A in HSCs and promotes hepatic fibrosis (Fujiwara et al., 1997), but the effects on LSCs are not yet examined. The details of morphology, function and roles in pathogenesis, ontogeny and phylogeny of LSCs are still unknown and remain to be investigated.
The main sources of vitamin A in the diet are carotenoids (provitamin A) from vegetables and retinyl esters from animal tissues (Blomhoff, 1987; Blomhoff and Blomhoff, 2006). Vitamin A is absorbed mainly from the intestine (Drummond and Bell, 1935; Popper and Volk, 1944). Cellular and molecular mechanisms of the absorption of vitamin A and provitamin A by absorptive cells have been extensively investigated (Goodman et al., 1966; Lawrence et al., 1966; Hollander and Muralidhara, 1977; Ong, 1994).
Vitamin A is absorbed by absorptive epithelial cells in the intestine and transported to the lymphatic system and stored as retinyl palmitate in lipid droplets in the cytoplasm in the HSCs. However, the route of vitamin A to the lymphatics within the villi is still problematic like that of fat (Popper and Volk, 1944). When the absorption of vitamin A was observed in the rat by fluorescence microscopy, Popper and Volk (1944) found that the cells in the lamina propria mucosae of the intestine emitted autofluorescence of vitamin A and suggested that vitamin A is carried by mesenchymal cells, leucocytes or histiocytes to the lymphatics. It was also suggested that in those cells, vitamin A might be stored for a considerable time. We have demonstrated that αSMA and desmin-positive fibroblastic cells exist in the lamina proproa mucosae of the intestine of rats (Matano et al., 1999). However, the morphology and roles of these cells were not yet investigated thoroughly.
Previously, we reported that vitamin A-storing cells exist not only in the liver (HSCs) but other organs in lower vertebrates such as lamprey (L. japonica) (Wold et al., 2004; Higashi et al., 2005b) and arrowtooth halibut (A. evermanni) (Yoshikawa et al., 2006). Other morphological studies on the mesenchymal cells in lamina propria mucosae of the digestive tract were reported [Popper and Volk, 1944; reviewed by Wake (1980), Hirosawa et al. (1988)], but the systematic study was not yet performed.
CTGF (connective tissue growth factor) is a 38-kDa protein involved in several human fibrotic disorders including atherosclerosis and skin and renal fibrosis. The role of small intestinal carcinoid tumour-derived fibrotic mediators, TGF-β1 and CTGF, have been investigated in the mediation of fibrosis via activation of an ‘intestinal’ LSCs (Kidd et al., 2007). Gastrointestinal carcinoid tumours were collected and analysed for CTGF and TGF-β1. Message levels of both CTGF and TGF-β1 in small intestinal carcinoid tumours were significantly increased compared with normal mucosa and gastric (non-fibrotic) carcinoids. Small intestinal carcinoid tumour fibrosis is a CTGF/TGF-β1-mediated SC-driven fibrotic response. The delineation of the biology of fibrosis will facilitate diagnosis and enable development of agents to obviate its local and systemic complications.
Vitamin A-storing cells in the intestine of lamprey digest ECM and are involved in the atrophy of the intestine in the lamprey during the spawning migration stage (Higashi et al., 2005a). Activity of acid hydrolase has been demonstrated in the lysosome of these cells. Phagocytosis by extrahepatic SCs was also reported in the pancreas (Shimizu et al., 2005).
Extrahepatic SCs in lungs and intestines of normal rats contain lipid droplets, and these lipid droplets increase in area when high doses of vitamin A are fed to the animals. These data suggest that not only HSCs, but also extrahepatic SCs, play an important role in vitamin A storage in normal as well as vitamin A-fed animals (Nagy et al., 1997).
The maculae flavae of the human vocal folds include dense ECM and compacted cells with a stellate morphology (Fuja et al., 2005). These VFSC (vocal fold SCs) are thought to participate in the metabolism of ECM essential in maintaining vocal fold viscoelasticity required for phonation. Isolated and cultured VFSCs maintain a distinct cellular and biochemical phenotype.
The regulation of ECM constituency is critical in maintaining vocal cord biomechanical viscoelasticity required for phonation. VFSC are thought to play a central role in laryngeal ECM metabolism, aging, scarring and cancer.
Diseases and HSCs
‘Activation’ of HSCs
Although several cell types in the liver contribute to fibrosis, it is generally accepted that the activated HSC is the source of the excess ECM. In the normal liver, the HSC is in a quiescent state, where it is the primary storage site for vitamin A. Following liver injury, the HSC undergoes an activation process with changes in cell morphology and gene expression, including increased cell proliferation, expression of new receptors and synthesis of ECM of which type I collagen predominates (Rippe and Brenner, 2004). ‘Activation’ of HSCs to a myofibroblastic phenotype is a key event in liver fibrosis (Friedman, 2006).
Identification of transcription factors such as Zf9 (Ratziu et al., 1998) with activities that are modulated during HSC activation will improve our understanding of the molecular events controlling HSC activation. Thus, HSC is now well established as the key cellular element involved in the development of hepatic fibrosis, and because of this, there is considerable interest in establishing the molecular events that trigger and perpetuate HSC activation. Molecular mechanisms of the HSC activation are investigated vigorously as follows. PDGF, EGF (epidermal growth factor), TGF-α and TGF-β and bFGF (basic fibroblast growth factor) induced a dose-dependent increase in DNA synthesis of rat HSCs. TGF-β did not affect DNA synthesis of HSC; however, TGF-β markedly potentiated the stimulatory effects of both EGF and PDGF.
PDGF is a key mitogen for HSC, and the co-ordinate release of other growth factors, together with PDGF by inflammatory cells, represents a potent potential stimulus for HSC proliferation in conditions of chronic self-perpetuating liver inflammation (Pinzani et al., 1989). HSC activation at the level of gene transcription requires the co-ordinated activity of several key transcriptional regulators of the HSC genome (Mann and Smart, 2002).
Candidates of triggering or perpetuating of HSC activation areas are as follows: PI3K (Marra et al., 1995), TGF-β1-induced activation of Ras, Raf-1, MEK and MAPK p42 and p44 (Reimann et al., 1997), ibuprofen (cyclooxygenase-2 inhibitor) (Mallat et al., 1998), activation of NF-κB (Elsharkawy et al., 1999), TNF-α (Hellerbrand et al., 1998; Varela-Rey et al., 2002), TIMP-1 (Bahr et al., 1999), MyoD protein (Vincent et al., 2001), discoidin domain tyrosine kinase receptor 2 (Olaso et al., 2001), Smad 7 (Tahashi et al., 2002), p38 MAPK (Varela-Rey et al., 2002), CTGF (Paradis et al., 2002) and FGF9 (Antoine et al., 2007).
A direct interaction between HCV proteins and HSCs may contribute to HCV-induced liver fibrosis (Bataller et al., 2004).
Transcriptional regulation plays a key role in the HSC activation process. Transcription factor MEF2 (myocyte enhancer factor 2) is correlated with HSC activation. MEF2 regulates multiple aspects of HSC activation (Wang et al., 2004a). The FoxO (Forkhead box gene, group O) family of Forkhead transcription factors, plays a crucial role in the transdifferentiation and proliferation of HSCs in liver fibrosis. Hyperinsulinaemia inactivates FoxO1 in HSCs, resulting in HSC activation and may result in the fibrosis in non-alcoholic fatty liver disease (Adachi et al., 2007).
Signals for HSC proliferation are transduced through FAK (focal adhesion kinase), PI3K and Akt. Finally, expression of type I collagen is regulated by the PI3K–Akt signalling pathway (Reif et al., 2003).
The activated HSC becomes responsive to both proliferative (PDGF) and fibrogenic (TGF-β) cytokines. It is becoming clear that these cytokines activate both MAPK (mitogen-activated protein kinase) signalling, involving p38 and FAK–PI3K–Akt–p70S6K (p70 S6 kinase) signalling cascades. Together, these regulate the proliferative response, activating cell cycle progression as well as collagen gene expression. It is anticipated that by understanding the molecular mechanisms responsible for HSC proliferation and excess ECM production, new therapeutic targets will be identified for the treatment of liver fibrosis (Friedman, 1993, 2000; Bauer and Schuppan, 2001; Parsons et al., 2007).
STAP/cytoglobin expression in HSCs
Recently, proteome analysis (Kristensen et al., 2000; Kawada et al., 2001; Maeda et al., 2003; Nakatani et al., 2004) and gene expression analysis (Jiang et al., 2006; Minicis et al., 2007) have been induced to the study investigating molecular mechanisms of HSC ‘activation’.
Using two-dimensional PAGE, a novel protein named STAP was reported (Kawada et al., 2001). STAP is a cytoplasmic protein with molecular weight of 21496 and shows about 40% amino acid sequence homology with myoglobin. STAP is demonstrated to be induced in in vivo- and in vitro-activated HSCs. STAP, also called cytoglobin, was demonstrated in fibroblast-like cells in splanchnic organs, namely, vitamin A-storing cells, but neither in epithelial cells, endothelial cells, muscle cells, blood cells, macrophage nor dermal fibroblasts (Nakatani et al., 2004). Roles of STAP in HSCs are intriguing and will be clarified more in the near future.
Hepatic fibrosis and HSCs
The molecular composition of the scar tissue in cirrhosis is similar regardless of aetiology and consists of the ECM constituents, collagen types I and III (i.e. ‘fibrillar’ collagens), sulfated proteoglycans and adhesive glycoproteins. These scar constituents accumulate from a net increase in their deposition in liver and not simply collapse in existing stroma. Although the cirrhotic bands surrounding nodules are the most easily seen form of scarring, it is actually the early deposition of matrix molecules in the subendothelial space of Disse – so-called ‘capillarization’ of the sinusoid – that more directly correlates with diminished liver function.
Chronic tissue damage often results in a deregulated wound healing, characterized by an imbalance between ECM synthesis (fibrogenesis) and degradation (fibrolysis) that leads to scar formation. Excessive scarring finally results in architectural distortion and failure of organs such as liver, lungs and kidneys.
HSCs have been identified as an important cellular source of ECM in liver fibrosis. Upon injury (e.g. by toxins or chronic hepatitis), the normally ‘quiescent’ HSC become ‘activated’ and start to proliferate. The ‘activated’ HSC undergo a phenotypic transdifferentiation to contractile myofibroblasts that express αSMA and an excess of ECM molecules.
Activation of HSCs and other resident mesenchymal cells into myofibroblasts expressing αSMA and collagen I is a key event in liver fibrogenesis. By using transgenic animals, heterogeneity of gene expression in myofibroblastic cells during active fibrogenesis has been demonstrated (Magness et al., 2004). The different cell types were distinguished on the basis of their immunophenotypic pattern with a combination of marker antibodies and on the basis of ultrastructural characteristics. Myofibroblasts and HSCs appear to be the main cell types involved in the initial phase of liver fibrogenesis induced by CCl4. Both myofibroblasts and HSC divide and transform (Seifert et al., 1994b).
Substantial improvements in the treatment of chronic liver disease have accelerated interest in uncovering the mechanisms underlying hepatic fibrosis and its resolution. Activation of resident HSCs into proliferative, contractile and fibrogenic cells in liver injury remains a dominant theme driving the field.
Not only HSCs, but also other cells, are able to produce ECM components for hepatic fibrosis and cirrhosis (Friedman, 2008a). Several new areas of rapid progress in the past 5–10 years also have taken root, including identification of different fibrogenic populations apart from resident HSCs, for example, portal fibroblasts, fibrocytes and bone marrow-derived cells, as well as cells derived from epithelial mesenchymal transition. In 1976, PCs were demonstrated to synthesize collagen for the first time (Sakakibara et al., 1976). Then, Hata et al. (1978,1980), Hata and Nagai (1980), and Hata and Ninomiya (1984) analysed collagen biosynthesis by PCs thoroughly. PCs and HSCs were demonstrated to be able to biosynthesize collagen type XVIII (Musso et al., 1998). The role of PCs in hepatic fibrosis is intriguing. EMT (epithelial to mesenchymal transition) is a central mechanism for diversifying the cells found in complex tissues. This dynamic process helps organize the formation of the body plan, and while EMT is well studied in the context of embryonic development, it also plays a role in the genesis of fibroblasts during organ fibrosis in adult tissues (Kalluri and Neilson, 2003).
The role of adult PCs as contributors to the accumulation of fibroblasts in the fibrotic liver is yet undetermined. Zeisberg et al. (2007) provide evidence that the profibrotic growth factor, TGF-β1, induces adult mouse PCs to undergo phenotypic and functional changes typical of EMT. These results provide direct evidence for the functional involvement of adult PCs in the accumulation of activated fibroblasts in the fibrotic liver. Furthermore, these findings suggest that EMT is a promising therapeutic target for the attenuation of liver fibrosis.
Roles of HSCs during liver regeneration
It is well known that liver cells, including PCs and HSCs, show a remarkable growth capacity after PHx (partial hepatectomy). Following 70% PHx in rodents, liver mass is almost completely restored after 14 days. PC proliferation starts after ∼24 h in the areas surrounding portal tracts and proceeds to the pericentral areas by 36–38 h. As a result of the early PC proliferation, avascular clusters of PCs are observed from 3 days after PHx. NPCs enter DNA synthesis ∼24 h after PCs, with peak activity at 48 h or later. Not only proliferation of PCs, but also activation of sinusoidal liver cells, including HSCs, are involved in the regeneration process through cell–cell interaction and cytokine networks (Mabuchi et al., 2004; Balabaud et al., 2004). We investigated PC and HSC interaction and the HSC activation at different time points after 70% PHx in the rat (Mabuchi et al., 2004).
PC clusters were often seen in vivo at 3 days after 70% PHx. The distance between HSCs fell from 61.7±2.1 μm in controls to 36.1±1.4 μm (P<0.001), while the HSCs/PCs ratio rose [0.71±0.01 to 1.08±0.03 (P<0.001)]. In >80% of in vivo microscopic fields at 3 days after 70% PHx, clusters of HSCs were observed especially near PC clusters. At 1 and 3 days after PHx, >20% of cells in the PC-enriched fraction were HSCs, which adhered to PCs. At 3 days after PHx, in addition to desmin staining, isolated HSCs were also positive for BrdU and αSMA and formed clusters suggesting that these HSCs were activated. At 3 days after PHx, HSCs in the HSC fraction in the isolated liver cells were positive for only desmin, which indicated that adherence to PCs is required for HSC activation. Thus, these data suggest that HSCs are activated by adhering to PCs during the early phase of hepatic regeneration.
As mentioned earlier, under physiological conditions, HSCs within liver lobules store about 50–80% of the total body vitamin A in lipid droplets in their cytoplasm, and these cells show zonal heterogeneity in terms of vitamin A-storing capacity. We have examined the status of vitamin A storage in HSCs in the liver regeneration (Higashi et al., 2005a). Morphometry at the electron microscopic level, fluorescence microscopy for vitamin A autofluorescence and immunofluorescence microscopy for desmin and αSMA were performed on sections of liver from rats at various times after the animal had been subjected to 70% PHx.
Under the electron microscope, the mean area of vitamin A-storing lipid droplets per HSC gradually decreased towards 3 days after PHx and then returned to normal within 14 days after it. However, the heterogeneity of vitamin A-storing lipid droplet area per HSC within the hepatic lobule (Higashi and Senoo, 2003) disappeared after PHx and did not return to normal by 14 days thereafter, even though the liver volume had returned to normal.
These results suggest that HSCs alter their vitamin A-storing capacity during liver regeneration and that the recovery of vitamin A homoeostasis requires a much longer time than that for liver volume.
Biliary atresia and HSCs
Biliary atresia is the most common and serious neonatal hepatobiliary disorder resulting from a fibro-inflammatory cholangiopathy. The biliary atresia is corrected by hepatoportoenterostomy developed by Kasai; however, more than half the patients ultimately require liver transplantation due to liver cirrhosis. Little is known about the aetiology and pathogenesis of biliary atresia. Recently, HSCs are suggested to play important roles in both progression and recovery of fibrogenesis in biliary atresia (Ramm et al., 1998; Issa et al., 2001). In patients with biliary atresia, αSMA-positive HSCs are localized in fibrotic septa and surrounding hyperplastic bile ducts and produce mRNA for collagen (Ramm et al., 1998). The production of collagen mRNA is seemed to be mediated by TGF-β secreted by HSCs, biliary epithelial cells and PCs adjacent to the fibrotic septa. On the other hand, HSC apoptosis are also shown to be involved in the recovery from biliary fibrosis in a bile duct-ligated animal model (Issa et al., 2001). In this experimental model, TGF-β induces apoptosis of HSCs, leading to the reversal of liver fibrosis.
The lamprey can be a model system for studying human biliary atresia because the entire bile-transport apparatus is completely lost during metamorphosis (Youson et al., 1978). Fibrosis by vitamin A-storing cells around the bile duct might contribute to the onset and progress of the atresia (Yamamoto et al., 1986). We have recently discovered that the degeneration of bile ducts depends on the apoptosis of biliary epithelial cells (Morii et al., 2010). The apoptosis starts in the cystic duct. Obstruction of the cystic duct and the largest intrahepatic bile duct precedes degeneration of more smaller intrahepatic bile ducts. Subsequent cholestasis observed in lamprey indicates the similarity between human and lamprey biliary atresia. In spite of this biliary atresia and intrahepatic cholestasis, the lamprey never develops biliary cirrhosis. Recently, there are several reports stating that biliary apoptosis may play an important role in obstructive cholangiopathy (Erickson et al., 2008; Harada et al., 2008). Revealing the mechanism for the apoptosis of lamprey biliary epithelia may help reveal the aetiology and pathogenesis of human biliary atresia (Morii et al., 2010).
After the re-discovery of the HSCs, the research of SCs has developed vigorously. Especially relevant to clinical medicine, the research has advanced rapidly as described in this review. In the standard textbook of internal medicine, Harrison's Principles of Internal Medicine, 17th edition. (Mailliard and Sorrell, 2008), HSCs have been described as a key player in hepatic fibrogenesis. Clinically, SCs are now targets to the therapy of organ fibrosis, such as liver and pancreas. However, in one of the standard textbooks of cell biology, Molecular Biology of the Cell, 5th edition (Alberts et al., 2008), description on HSCs is still lacking. As we introduced in this review, HSc and vitamin A-storing cells are intriguing from many directions of basic research. The authors hope more basic researchers join the SC research.
We thank Dr Kenjiro Wake and Dr Hajime Manner (Emeritus Professors of Tokyo Medical and Dental University) who introduced Haruki Senoo into basic medical research and still continue to encourage the authors.
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Received 14 May 2010; accepted 22 July 2010
Published online 11 November 2010, doi:10.1042/CBI20100321
© The Author(s) Journal compilation © 2010 Portland Press Limited
ISSN Print: 1065-6995
ISSN Electronic: 1095-8355
Published by Portland Press Limited on behalf of the International Federation for Cell Biology (IFCB)
Figure 2 Gold chloride staining specifically demonstrating black-stained hepatic SCs of polar bears (a and b), arctic foxes (c and d) and rats (e and f)
Figure 3 Fluorescence micrographs demonstrating vitamin A autofluorescence in hepatic SCs of polar bears (a), arctic foxes (b) and rats (c and d)