1. Introduction ....................................................................................................................................................................... 503
1.1. Basic anatomy and physiology of the liver .............................................................................................................. 503
1.2. Major cell types of the liver ....................................................................................................................................... 504
1.2.1. Hepatocytes..................................................................................................................................................... 504
1.2.2. Liver sinusoidal endothelial cells .................................................................................................................. 506
1.2.3. Hepatic stellate cells ....................................................................................................................................... 507
1.2.4. Kupffer cells ..................................................................................................................................................... 508
1.2.5. Cholangiocytes ............................................................................................................................................... 509
1.2.6. Hepatic progenitor cells ................................................................................................................................. 510
1.3. Hepatocyte cytoarchitecture and cell polarity ........................................................................................................ 510
1.3.1. Cell–cell interactions ...................................................................................................................................... 511
220.127.116.11. Homotypic hepatocyte interactions ................................................................................................ 511
Organotypic liver culture models: Meeting current challenges
in toxicity testing
Edward L. LeCluyse1, Rafal P. Witek2, Melvin E. Andersen1, and Mark J. Powers2
1The Institute for Chemical Safety Sciences, The Hamner Institutes for Health Sciences, Research Triangle Park,
NC, USA and 2Life Technologies, Durham, NC and Frederick, MD, USA
Prediction of chemical-induced hepatotoxicity in humans from in vitro data continues to be a significant challenge
for the pharmaceutical and chemical industries. Generally, conventional in vitro hepatic model systems (i.e. 2-D
static monocultures of primary or immortalized hepatocytes) are limited by their inability to maintain histotypic
and phenotypic characteristics over time in culture, including stable expression of clearance and bioactivation
pathways, as well as complex adaptive responses to chemical exposure. These systems are less than ideal for longer-
term toxicity evaluations and elucidation of key cellular and molecular events involved in primary and secondary
adaptation to chemical exposure, or for identification of important mediators of inflammation, proliferation and
apoptosis. Progress in implementing a more effective strategy for in vitro-in vivo extrapolation and human risk
assessment depends on significant advances in tissue culture technology and increasing their level of biological
complexity. This article describes the current and ongoing need for more relevant, organotypic in vitro surrogate
systems of human liver and recent efforts to recreate the multicellular architecture and hemodynamic properties of
the liver using novel culture platforms. As these systems become more widely used for chemical and drug toxicity
testing, there will be a corresponding need to establish standardized testing conditions, endpoint analyses and
acceptance criteria. In the future, a balanced approach between sample throughput and biological relevance
should provide better in vitro tools that are complementary with animal testing and assist in conducting more
predictive human risk assessment.
Keywords: in vitro hepatic models, hepatocytes, hepatotoxicity, organotypic culture models, microfluidic devices,
Address for Correspondence: Edward L. LeCluyse, Ph.D., The Hamner Institutes for Health Sciences, 6 Davis Drive, P.O. Box 12137.
Tel: 919-558-1322. Fax: 919-226-3150. E-mail: firstname.lastname@example.org
(Received 15 October 2011; revised 26 March 2012; accepted 30 March 2012)
Critical Reviews in Toxicology, 2012; 42(6): 501–548
© 2012 Informa Healthcare USA, Inc.
ISSN 1040-8444 print/ISSN 1547-6898 online
502 E. L. LeCluyse et al.
Critical Reviews in Toxicology
18.104.22.168. Heterotypic cell interactions ............................................................................................................ 511
1.3.2. Liver biomatrix ................................................................................................................................................ 512
1.4. Liver biomechanical properties ............................................................................................................................... 513
1.5. Liver hemodynamics ................................................................................................................................................. 514
2. The current state of cell-based hepatic culture systems ............................................................................................... 514
2.1. Past strategies for maintaining hepatic structure and function in vitro ................................................................ 514
2.1.1. Historical perspective ..................................................................................................................................... 515
2.1.2. Extracellular matrix effects ............................................................................................................................ 515
2.1.3. Adhesive and mechanical factors .................................................................................................................. 516
2.1.4. Three-Dimensional spheroid aggregate culture .......................................................................................... 516
2.1.5. Co-culture systems ......................................................................................................................................... 517
2.1.6. Perifusion culture systems ............................................................................................................................. 518
2.2. Lessons learned and where we go from here .......................................................................................................... 518
2.3. Current challenges for today’s model systems ........................................................................................................ 518
3. Considerations for the development of organotypic liver models .............................................................................. 519
3.1. Source of cellular material ........................................................................................................................................ 519
3.1.1. Primary cells.................................................................................................................................................... 519
3.1.2. Immortalized cell lines ................................................................................................................................... 520
22.214.171.124. HepG2 ................................................................................................................................................ 520
126.96.36.199. Fa2N-4 ................................................................................................................................................ 520
188.8.131.52. HepaRG ............................................................................................................................................. 521
3.1.3. Stem cells ......................................................................................................................................................... 521
3.2. Maintenance of histotypic and phenotypic characteristics ................................................................................... 522
3.3. Zonal architecture and microenvironments ........................................................................................................... 523
3.4. Controlled flow dynamics ......................................................................................................................................... 523
3.5. Defined cellularity ..................................................................................................................................................... 524
3.6. Accessibility................................................................................................................................................................ 524
3.7. Throughput and cost-effectiveness .......................................................................................................................... 525
4. Working towards standardized methods for evaluation and validation of advanced culture models ..................... 525
4.1. Assessing cell and tissue integrity ............................................................................................................................ 526
4.2. Standardizing culture conditions ............................................................................................................................. 527
4.3. Assessing metabolic capacity of in vitro systems .................................................................................................... 527
4.4. Normalization of in vitro data across culture platforms ......................................................................................... 528
5. Advanced organotypic culture technologies ................................................................................................................. 529
5.1. Microfluidic perfusion array ..................................................................................................................................... 529
5.2. Bioengineered micro-patterned liver platform ....................................................................................................... 530
5.3. Biochip dynamic flow system ................................................................................................................................... 531
5.4. 3-D liver tissue culture scaffold ................................................................................................................................ 532
5.5. 3-D scaffolds with dynamic flow .............................................................................................................................. 532
6. Applications in drug and chemical testing .................................................................................................................... 533
6.1. Long-term study of low-dose exposures to drugs and chemicals ......................................................................... 533
6.2. Elucidation of intercellular effects on the initiation or propagation of chemical toxicity ................................... 533
6.3. ‘Gold-standard’ to compare the biological relevance of HTS assays..................................................................... 534
6.4. Continuity between studies within a single project or study ................................................................................. 534
6.5. Mimicking dynamic exposure profiles ..................................................................................................................... 534
6.6. Metabolite identification and profiling .................................................................................................................... 534
6.7. Toxicity testing and computational modeling for human risk assessment........................................................... 535
7. Conclusions and future directions ................................................................................................................................. 536
Acknowledgements ................................................................................................................................................................ 537
Declaration of interest ............................................................................................................................................................ 537
References ............................................................................................................................................................................... 537
Appendix: Abbreviations ....................................................................................................................................................... 548
Organotypic liver culture models 503
© 2012 Informa Healthcare USA, Inc.
There are increasing pressures for regulatory, economic
and practical reasons to find more effective and efficient
ways to understand and predict human response to drug
and chemical exposure. In vitro testing strategies have been
applied successfully to predict the in vivo pharmacokine-
tics and clearance of compounds for years, including
the potential of compounds to be involved in significant
adverse interactions through the induction or inhibition of
liver enzymes (Lin, 2006; Hewitt et al., 2007a; Obach, 2009;
Obach et al., 2008). Cell-based approaches and endpoint
assays to study hepatoxicity of drugs and other chemicals
in vitro have also been used and described extensively
(Castell et al., 2006; Gebhardt et al., 2003; Gomez-Lechon
et al., 2008; Guguen-Guillouzo et al., 2010; Guillouzo and
Guguen-Guillouzo, 2008). Nonetheless, there remains a
need for more relevant and sophisticated in vitro models
systems with which to probe and identify pathways that
are perturbed following acute and chronic exposure
to chemicals and to help explain species differences
in compound biotransformation and bioactivation. In
this regard, the mode of action (MOA) for many types
of chemical- or drug-induced hepatotoxic responses
often includes multiple organs and cell types involving
perturbation of pathways over prolonged exposure periods
(DeLeve et al., 1997; Kmiec, 2001; Sunman et al., 2004). For
example, chemical-induced changes in nuclear receptor
activation and the corresponding changes in target gene
expression patterns can eventually lead to overwhelming
an organism’s adaptive responses over many days or even
weeks of exposure at low, but physiologically relevant,
exposure levels (Moreau et al., 2007; Pascussi et al.,
2005). Immune-mediated responses that are associated
with reactive metabolites or that occur upon exposure
to endotoxins require interactions between hepatocytes,
endothelial cells and Kupffer cells (Sunman et al., 2004;
DeLeve et al., 1997). Clearly, there is a need to develop
more physiologically-relevant, long-term culture model
systems for assessing toxicity, conducting in vitro-in vivo
extrapolation (IVIVE) and supporting development of
physiologically-based pharmacokinetic (PBPK) models of
chemical disposition and toxicity.
The purpose of this review article is to explore the
historical evolution of hepatic culture models and the
reasons why there continues to be a need for more
advanced in vitro systems with which to study chemical-
induced hepatotoxicity. In the following sections, we (1)
review the basic anatomy and physiology of the liver,
especially those attributes or features which represent
the biological basis for the different modes of action
of hepatotoxins, (2) describe the reasons why current
standard model systems are not able to address certain
facets of chemical-induced hepatotoxicity, (3) provide a
list of the basic components or requirements that ideally
should be incorporated into the development and vali-
dation of advanced in vitro model systems, (4) describe
some examples of emerging cell culture technologies
and how they combine elements of tissue architecture,
cellular composition and hemodynamic flow with tradi-
tional and novel platforms, and (5) discuss applications
of these advanced culture systems in drug and chemical
1.1 Basic anatomy and physiology of the liver
The liver is a versatile organ which plays an impor-
tant role in a variety of critical functions, including the
detoxification of the systemic and portal blood to the
production and secretion of blood and bile components
(Rodés et al., 2007). The liver is also involved in protein,
steroid, and fat metabolism as well as vitamin, iron, and
sugar storage. The classical structural unit of the liver is
the hepatic lobule (Figure 1) (Bioulac-Sage et al., 2007).
When viewed in cross section, the lobule has the shape of
a polygon, usually a hexagon. At the corners of the polyg-
onal lobule are the portal triads consisting of the hepatic
artery, bile duct, and portal vein. The central structure
of the lobule, traversing its long axis, is the central vein.
Plates of parenchymal cells or hepatocytes radiate from
the central vein to the perimeter of the lobule to define
the basic functional unit of the liver, known as the acinus,
Figure 1. Representation of histotypic liver microstructure. (A)
Diagram of the basic hepatic lobule and acinus substructure
showing the relative direction of blood flow from portal triads
towards the central veins (red arrows). (B) Diagram illustrating the
three-dimensional architecture of the liver between a portal triad
and the central vein. The networks of bile canaliculi (yellow-green)
run parallel and counter to the blood flow through the sinusoids.
504 E. L. LeCluyse et al.
Critical Reviews in Toxicology
which also serves as a microcosm of the major hepatic
microenvironments, containing the essential cellular
and physiological features that define the unique archi-
tecture of the liver tissue. Hepatic plates or cords are
generally one hepatocyte thick and are separated from
one another by the hepatic sinusoids (the “capillaries”
of the liver) which are lined by sinusoidal endothelium
(Bioulac-Sage et al., 2007; Khan et al., 2007).
The liver acinus is demarcated into three discrete
zones: zone 1 is the periportal region; zone 2 is the
midlobular region; and zone 3 is the pericentral
region (Figure 2) (Rappaport, 1977; Ito and McCuskey,
2007). Blood enters the liver from the portal veins and
hepatic arteries at the portal triads, flows through the
sinusoidal microvasculature surrounded by the plates
of parenchymal cells, and exits from the central vein.
Due to the particular configuration of cells along the
microvasculature and the directionality of flow through
the lobular units, various chemical gradients and
microenvironments are present (Smith and Wills, 1981;
Ugele et al., 1991; Gebhardt, 1992). Cell maturation,
matrix chemistry, solute concentrations, endogenous
substrate utilization, oxygen tension, gene expression
and xenobiotic clearance mechanisms vary across the
acinus (Figure 2A) (Probst and Jungermann, 1983; Wolfe
and Jungermann, 1985; Wojcik et al., 1988; Reid et al.,
1992; Lindros, 1997; Turner et al., 2011; Wang et al., 2011).
An example of the differences in the zonal expres-
sion of specific genes in human liver is shown in
Figure 2B using antibodies against cytochrome P450
3A4 (CYP3A4). Similar to many cytochrome P450 (CYP)
enzymes, the highest levels of CYP3A4 expression are in
zone 3 (pericentral) and extend to the mid-lobule region
(in this particular case). The positional difference in
expression is partially responsible for the zonal pattern
of toxicity exhibited in vivo upon exposure to many bio-
activated compounds, such as acetaminophen, carbon
tetrachloride, bromobenzene and chloroform (Black,
1984; Tomasi et al., 1985; Anundi et al., 1993; Moon et al.,
2010). Midlobular (zone 2) necrosis is observed in rodents
exposed to natural and synthetic compounds, such as
cocaine, phytol and germander (Roth et al., 1992; Mackie
et al., 2009; Loeper et al., 1994). In the case of other hepa-
totoxins (e.g. allyl alcohol, phosphorus), zone 1 specific
toxicity may be observed, as a result of the unique oxygen,
metabolic and cellular microenvironments located near
the portal triad (Badr et al., 1986; Przybocki et al., 1992).
1.2 Major cell types of the liver
The liver is comprised of cells that are broadly divided
into two categories: parenchymal cells and nonparen-
chymal cells (NPC). The parenchymal fraction consists
of hepatocytes, which represent nearly 80% of liver
volume and 60% of the total cell population in the liver
(Kmiec, 2001; Bioulac-Sage et al., 2007). The nonparen-
chymal fraction encompasses the remaining liver cells,
representing approximately 6.5% of liver volume (the
remaining volume consisting of the vascular and ductu-
lar networks) and 40% of the total number of liver cells.
Major liver NPC include bile duct epithelial cells (or chol-
angiocytes), liver sinusoidal endothelial cells (LSEC),
hepatic stellate cells (HSC), Kupffer cells (KC) and pit
cells (intrahepatic lymphocytes or nature killer cells).
While traditionally relegated to the status of the “other”
cell types of the liver when discussing hepatocytes, NPC
are important contributors to various roles that support
and regulate hepatic growth and function (Kmiec, 2001).
These functions include production of growth factors and
other mediators of cellular function, including transport
and metabolism. NPC can serve as the primary targets of
certain hepatotoxins, or can mediate the physiological
or pathological response to other cells (Ramadori et al.,
2008; Parola and Pinzani, 2009; Ishibashi et al., 2009).
The parenchymal cells or hepatocytes are highly differ-
entiated epithelial cells that comprise the cell plates of
the liver lobule (Figure 3). They perform a majority of the
physiological functions commonly associated with the
liver, including xenobiotic biotransformation and elimi-
nation (Rodés et al., 2007). Hepatocytes are involved in
protein, steroid, and fat metabolism as well as vitamin,
iron, and sugar storage and display marked morphologic,
Figure 2. Structural and functional zonation of the liver. (A) Discrete
zones of the liver between the portal vein (PV) and central vein (CV)
illustrating the differences in cell size, phenotype and gradients in
oxygen tension and metabolism. (B) Immunostaining of human
liver tissue with antibodies again CYP3A4 (brown stain) showing
the differential expression of CYP enzymes across the zones of the
liver microstructure. The greatest expression of CYP enzymes is
predominantly in pericentral hepatocytes (zone 3) with a distinct
boundary or gradient at the mid-lobular region (zone 2).
Organotypic liver culture models 505
© 2012 Informa Healthcare USA, Inc.
biochemical and functional heterogeneity based on their
zonal location (Traber et al., 1988; Gebhardt, 1992; Ugele
et al., 1991; Jungermann and Kietzmann, 1996; Lindros
et al., 1997; Turner et al., 2011). Under healthy non-
adaptive conditions, parenchymal cell size increases
from Zone 1 to Zone 3, accompanied by distinctive zonal
variations in morphological features of the cells, such
as mitochondria, endoplasmic reticulum, lipid vesicles
and glycogen granules (Figure 2A) (Michaels et al., 1984;
Uchiyama and Asari, 1984; Ferri et al., 2005).
Much of the functional diversity of hepatocytes is
also revealed in their cytological features. Hepatocytes
are cuboidal in shape and possess one or more nuclei
with prominent nucleoli (Figure 3). The fraction of
hepatocytes that are polyploid (4N and 8N), which
results from mitotic division of the nucleus without
accompanying cytokinesis, increases across the liver
lobule from Zone 1 to Zone 3 (Gupta, 2000; Celton-
Morizur and Desdouets, 2010). Generally, hepatocytes
possess abundant mitochondria with Golgi complexes
localized mainly adjacent to the bile canaliculi. The
cytoplasm is rich in both rough endoplasmic reticulum
(RER), which is indicative of the hepatocyte’s secretory
nature, and smooth endoplasmic reticulum (SER),
with many of the enzymes involved in phase 1 and
2 biotransformation of drugs and other xenobiotics.
Lysosomes are scattered throughout the cytoplasm and
play a central role in the degradation of extracellular and
intracellular macromolecules including organelles and
proteins (autophagy) that results from environmental
stress, such as nutrient or serum deprivation (Singh,
2010; Rautou et al., 2010). Hepatocytes are also highly
polarized cells with distinct sinusoidal and canalicular
plasma membrane domains that are separated by
junctional complexes (Figures 3 & 4). These membrane
domains exhibit ultrastructural, compositional, and
functional differences (Simons and Fuller, 1985; Meijer,
1987) and are essential for the hepatocyte’s role in the
uptake, metabolism, and biliary elimination of both
endogenous and exogenous substrates (Klaassen and
Watkins, 1984; Meijer et al., 1990; van Montfoort et al.,
In the intact liver, hepatocytes exhibit efficient trans-
port of a wide variety of endogenous and exogenous
substances from blood into bile (Klaassen and Watkins,
1984; Meijer et al., 1990). Physiologically, biliary trans-
port is concerned primarily with the production and
secretion of bile components which are necessary for fat
absorption in the gut (Rodés et al., 2007) but is also an
important step in the detoxication of both endogenous
and exogenous compounds (Klaassen and Watkins,
1984). The production of bile requires the coordinated
participation of transport mechanisms selectively local-
ized to the sinusoidal and canalicular membranes of the
hepatocytes (Hubbard et al., 1985; Simons and Fuller,
1985; Klaassen and Aleksunes, 2010). Perturbation of
these transport mechanisms by drugs and other xenobi-
otics is one cause of intrahepatic cholestasis that can lead
to accumulation of substrates to toxic levels in both the
liver and plasma.
The functional and structural specialization of the
hepatocyte is related to selective activation and the
sustained expression of a distinct set of gene programs
encoding specific categories of proteins (De Simone and
Cortese, 1992; De Simone and Cortese, 1991). The expres-
sion of hepatocyte-specific genes is primarily regulated
at the transcriptional level and depends on signals from
both inside and outside the cell (De Simone and Cortese,
Figure 3. Histological and architectural structure of the liver parenchyma and endothelium. (A) Transmission electron micrograph of whole
liver showing histotypic configuration and cytoarchitecture of hepatocytes (HC), including bile canaliculi (BC) and nucleoli (arrowhead).
Sinusoids contain red blood cells (RBC) and resident macrophages (Kupffer cells, KC), and are lined with sinusoidal endothelial cells (LSEC).
(B) Diagram illustrating the diverse morphological features of the mature hepatocyte including bile canaliculi, junctional complexes, and
various subcellular organelles. Hepatocytes exhibit cellular polarity of subcellular organelles, cytoskeletal elements, and biochemical
composition of membrane domains. BLD, basolateral domain; AD, apical domain; RER, rough endoplasmic reticulum; SER, smooth
endoplasmic reticulum; Mito, mitochondria; Gly, glycogen granules; Lys, lysosomes; Sp Disse, space of Disse; Fen, fenestrations; ECM,
extracellular matrix; GJIC, gap junction intercellular communication; Desm, desmosome; AJ, adherence junction; TJ, tight junction; BC, bile
canaliculi; LSEC, liver sinusoidal endothelial cell.
506 E. L. LeCluyse et al.
Critical Reviews in Toxicology
1991; Derman et al., 1981; Xanthopoulos and Mirkovitch,
1993). Extracellular soluble (e.g. growth factors, cyto-
kines, other hormones) and insoluble (e.g. extracel-
lular matrix composition) signals play a major role in
determining which combination of genes is expressed
and, thus, the resulting phenotype (DeLeve et al., 2004;
Bissell and Choun, 1988; Bissell et al., 1990a; Bucher
et al., 1990; Martinez-Hernandez and Amenta, 1995;
Nagaki et al., 1995; Rana et al., 1994; Sidhu et al., 1994;
Sidhu and Omiecinski, 1995).
1.2.2 Liver sinusoidal endothelial cells
LSEC line the walls of hepatic sinusoids (Figure 5A–C)
and are thin, elongated cells, like most vascular
endothelial cells that possess a relatively large
number of pinocytotic vesicles, suggesting significant
endocytotic activity (DeLeve, 2007b; DeLeve, 2007a;
Perri and Shah, 2005). The intercellular adhesions
between endothelial cells of the liver sinusoids are
much less prominent than typical vascular endothelial
cells and their plasma membrane is characterized by
small pores, or fenestrations, 50–200 nm in diameter
that allow free diffusion of many substances, but not
particles of the size of chylomicrons and whole cells,
between the blood and the hepatocyte basolateral
surface (Figure 5B) (Braet and Wisse, 2002; DeLeve,
2007b; DeLeve, 2007a; Cogger et al., 2010). The greater
intercellular permeability and surface fenestrae along
with the lack of a prominent basement membrane
between the LSEC and parenchyma all contribute to
enhance hepatocyte exposure to soluble components
in the circulating blood (DeLeve, 2007b; DeLeve, 2007a;
Perri and Shah, 2005) and improve passive transport
of many endogenous and xenobiotic substrates (Braet
and Wisse, 2002). The increased access to blood permits
greater oxygenation of hepatocytes and more efficient
clearance of drugs and other xenobiotics.
LSEC are part of the reticuloendothelial system (RES)
and play three important roles in maintaining overall
hepatic homeostasis. First, they act as a “selective sieve”
for substances passing from the blood to hepatocytes and
vice versa. Second, they serve as a “scavenger system” ,
clearing the blood of macromolecular waste products that
originate from turnover processes in various tissues. LSEC
exhibit significant endocytic capacity for colloids and for
many ligands, including glycoproteins, components of
the extracellular matrix (hyaluronate, collagen, fibronec-
tin), immune complexes, transferrin and ceruloplasmin
(DeLeve, 2007b; DeLeve, 2007a). Third, LSEC play a role in
hepatic immunity to foreign pathogens and immune tol-
erance to neo-antigens formed during the metabolism of
xenobiotics (Perri and Shah, 2005; DeLeve, 2007b; DeLeve,
2007a). LSEC also function as antigen-presenting cells
(APC) in the context of both MHC-I and MHC-II restriction
with the resulting development of antigen-specific T-cell
tolerance (DeLeve, 2007a; DeLeve, 2007b). They are active
in the secretion of cytokines, eicosanoids (i.e. prostanoids
and leukotrienes), endothelin-1 (ET-1), nitric oxide, and
some extracellular matrix (ECM) components (DeLeve et
al., 2004; Deleve et al., 2008).
LSEC also play a significant role in the clearance and
bioactivation of drugs and other xenobiotics, and are a
target for some types of chemical-induced hepatotoxicities
(Deaciuc et al., 2001; Xie et al., 2010; DeLeve, 2007a;
DeLeve, 2007b; Ito et al., 2003). The LSEC-specific phase
1 enzymes have been less well characterized compared
to their epithelial counterparts, but it is clear that they do
contribute to the metabolism, clearance and bioactivation
of endogenous and exogenous substrates (DeLeve,
2007b; DeLeve, 2007a). For example, the cytotoxicity
Figure 4. Cellular structures involved in cell-cell and cell-matrix interactions. (A) Ultrastructural composition of junctional complexes between
adjacent hepatocytes. Intercellular adhesions between the basolateral (sinusoidal) and apical (canalicular) domains of adjoining hepatocytes
are composed of a series of three distinct types of junctions: the tight junction (TJ), the adherens junction (AJ) and the desmosomal belt (D).
(B) Diagram illustrating the adhesion molecules and associated proteins and pathways that mediate hepatocyte interactions with each other
(claudins/occludins, cadherins, connexins) and the extracellular matrix (integrins). (Modified from D. Dostal, Ph.D., Div. of Mol. Cardiology,
Texas A&M Health Science Center.)
Organotypic liver culture models 507
© 2012 Informa Healthcare USA, Inc.
of acetaminophen is observed in LSEC in the absence
of hepatocytes, suggesting that they are fully capable of
generating the reactive metabolite of acetaminophen and
mimic its cytotoxic effects (Ito et al., 2003; Xie et al., 2010).
In addition, LSEC are able to activate aflatoxin B1 to a
mutagenic metabolite through an Aroclor 1254-inducible
pathway (Schlemper et al., 1991; Jennings et al., 1992).
LSEC also exhibit high levels of many phase 2 conjugating
enzyme activities (Utesch and Oesch, 1992). Although
the overall metabolic capacity of LSEC is less than that
of hepatocytes (~1/10th), their overall role in hepatic
clearance of compounds and hepatoxic events has been
generally overlooked and underappreciated (Schrenk
et al., 1991; Sacerdoti et al., 2003; Wu et al., 2008).
1.2.3 Hepatic stellate cells
HSC, also called perisinusoidal cells, Ito cells or fat-stor-
ing cells, reside in the space of Disse – the perisinusoidal
space between the basolateral surface of hepatocytes
and the anti-luminal side of sinusoidal endothelial
cells (Asahina et al., 2009). Under normal physiological
conditions in the adult liver, HSC are morphologically
characterized by extensive dendrite-like extensions that
wrap around the sinusoids, essentially “embracing” the
endothelial cells (Figure 5D–F) (Friedman, 2008). This
close contact between HSC and their neighboring cell
types facilitates intercellular communication by means
of soluble mediators and cytokines. HSC store vitamin
A, control turnover and production of ECM, and are
involved in regulation of sinusoid contractility. HSC can
be identified by the expression of desmin, a typical inter-
mediate filament protein within contractile cells. Mature
HSC produce both network and fibrillar collagens (large
amounts of type I collagen and lower levels of type III,
IV and V collagen), large amounts of elastin and both
heparan sulfate proteoglycans (HS-PG) and chondroitin
Figure 5. Major nonparenchymal cell types of the liver. Top row: CD-31 staining of liver sinusoidal endothelial cells (LSEC) lining vascular walls of
whole liver (A), scanning electron micrograph of the endothelial lining of the liver sinusoids showing extensive patches of fenestrae (arrows) (B),
and primary LSEC showing typical morphology in vitro (C). Middle row: HSC (GFAP) in normal liver (D), myofibroblastic HSC (µSMA) in fibrotic
liver (E), and isolated qHSC showing storage of vitamin A as bright “floating” vesicles within the cell body. Upon activation or injury the HSC
undergo extensive morphological and biochemical changes, which include the synthesis, secretion and restructuring of ECM molecules. Bottom
row: Kupffer cells (KC) showing their dynamic morphology (G), their identification with CD68 showing extended projections on the cell bodies
used for contact with other cells (H), and a magnified view showing KC loaded with vesicles containing cytokines and other secretory factors (I).
508 E. L. LeCluyse et al.
Critical Reviews in Toxicology
sulfate proteoglycans (CS-PG)(Wang et al., 2010b; Parola
and Pinzani, 2009).
HSC also produce important cytokines and growth
factors for intercellular communication in normal and
injured liver. These include hepatocyte growth factor
(HGF), transforming growth factor-α (TGF-α) and
epidermal growth factor (EGF), three potent growth
factors for hepatocyte proliferation during liver regeneration
(Friedman, 2008; Asahina et al., 2009). TGF-α and EGF also
stimulate mitosis in stellate cells themselves, creating an
autocrine loop for cellular activation. Insulin-like growth
factor (IGF-I and II) and platelet-derived growth factor
(PDGF), among the most potent HSC mitogens, are also
secreted by stellate cells (Ramadori et al., 2008; Asahina
et al., 2009). Collectively, these factors allow HSC to
influence their own gene expression and phenotype as
well as that of other cells of the liver.
Following liver injury, HSC become activated to a
myofibroblastic (MF) phenotype characterized by a loss
of vitamin A and expression of α-smooth muscle actin
(α-SMA) (Friedman, 2008). In this activated state, MF-HSC
produce growth factors and cytokines, such as transform-
ing growth factor β (TGF-β), which play a key role in the
regulation of hepatocyte growth and the development
of inflammatory fibrotic response of the liver (Ramadori
et al., 2008; Parola and Pinzani, 2009). Connective tissue
growth factor (CTGF) is also expressed by HSC and pro-
motes fibrogenesis. HSC participate significantly in the
inflammatory response of the liver through secretion of
cytokines, such as macrophage colony-stimulating factor
(M-CSF), which regulates macrophage accumulation and
growth, interleukins-8 and -6 (IL-8, IL-6), monocyte che-
motactic peptide (MCP)-1, CCL21, RANTES, CCR5, and
the anti-inflammatory IL-10. Activated HSC express toll-
like receptors (TLRs) allowing them to recognize bacterial
endotoxin lipopolysaccharide (LPS) and function as APC.
HSC also amplify the inflammatory response by inducing
infiltration of leukocytes.
HSC are involved in the onset and progression of cir-
rhosis, which is typically associated with highly activated
cells leading to a fibrotic response, a progressive increase
in deposition of ECM proteins and scar tissue formation
throughout the liver. Mice defective in the lhx2 gene, which
regulates the fibrogenic process, have early and inappro-
priate activation of stellate cells and “spontaneous” cirrho-
sis (Wandzioch et al., 2004). A major contributing factor
includes the production of the potent vasoconstrictor
ET-1. ET-1 has a prominent contractile effect on HSC and
MF-HSC, which may contribute to portal hypertension in
the cirrhotic liver. Activated HSC also produce elevated
levels of extracellular matrix proteins (e.g. collagen types I,
III, IV, V) and various basal adhesion molecules (fibronec-
tin, and laminin α1 and γ1 chains) that contribute to scar
tissue formation throughout the liver (Friedman, 2006).
1.2.4 Kupffer cells
KC have mesenchymal origins and are the resident mac-
rophages in the liver with a pronounced endocytic and
phagocytic capacity (Figure 5G–I) (Jaeschke, 2007). They
are localized within the sinusoidal microvasculature on
the luminal side of endothelial cells; however, they have
long cytoplasmic extensions that facilitate direct cell-to-
cell contact with hepatocytes. KC are in constant contact
with gut-derived particulate materials, such as tissue
and cellular debris, and soluble bacterial products and
endotoxins (Kolios et al., 2006). These particulates and
other macromolecular complexes are rapidly and effi-
ciently extracted from the blood by KC and subsequently
processed through the endosomal and lysosomal path-
ways (Roberts et al., 2007). KC and their products are
also involved in modulating the turnover of hepatocytes
and other cell types by apoptosis (Hoebe et al., 2001).
The morphology and biocapacity of KC is highly hetero-
geneous; in the periportal area, KC are larger and more
active in phagocytosis, whereas centrilobular Kupffer
cells are more active in the production of cytokines and
inflammatory responses (Roberts et al., 2007).
KC are part of the RES and represent the largest popu-
lation of resident macrophages in the body. They play a
very important role in immune surveillance of the host
and are involved in modulating systemic responses to
severe infections and controlling concomitant immune
responses via antigen presentation and suppression of
the activation and proliferation of T-cells (Kolios et al.,
2006). In their primary scavenger role, KC endocytose
foreign particles and bacterial endotoxins, which causes
their activation and subsequent production of a number
of modulators of cell signaling pathways, such as oxygen-
derived free radicals, nitric oxide, eicosanoids, peptide
leukotrienes, prostaglandins, and various cytokines,
including TNF-α, TGF-α, IL-1, IL-6 and others (Kolios
et al., 2006). Activation of KCs is elicited also during
chemical-induced liver injury and they have been found
to play a stimulatory role in liver regeneration, can reverse
liver fibrosis, and are critical for the progression of alco-
holic injury. In addition to their phagocytic capacities,
KC process significant quantities of gut-derived antigens
and blockage of KC results in an exaggerated response to
these antigens. They also interact in complex ways with
bactericidal neutrophils that immigrate rapidly to the
liver in response to infection.
KC are capable of modulating the metabolic activity of
hepatocytes via production of cytokines (e.g. IL-1, IL-6,
TNFµ) that induce the expression of acute phase proteins
while causing the down-regulation of genes involved in
the metabolism and clearance of xenobiotics (Hoebe
et al., 2001). Proinflammatory cytokines produced by KC
can cause a potent and complete suppression of cyto-
chrome P450, Uridine 5′-diphospho (UDP)-glucuronosyl
transferase systems and uptake and efflux transporter
expression (Sunman et al., 2004; Wu et al., 2006; Higuchi
et al., 2007; Morgan, 2009). In this context, KC are an
important component in the development of hepatocyte
culture systems intended to mimic liver injury caused by
bioactivation of xenobiotics and their resultant inflam-
Organotypic liver culture models 509
© 2012 Informa Healthcare USA, Inc.
Cholangiocytes, also called intrahepatic bile duct cells, are
biliary epithelial cells that line the bile ducts (Figure 6). They
account for approximately 5% of the liver cell population
and are distinct from undifferentiated hepatoblasts that
also give rise to mature hepatocytes. Morphologically, they
make-up the cuboidal epithelium in the small interlo-
bular bile ducts, but become progressively columnar and
mucus-secreting in larger bile ducts approaching the porta
hepatis and the extrahepatic ducts. The cholangiocyte
population is heterogeneous with respect to morphology,
secretion and expression patterns, and its response to
hormones, peptides, growth factors, cytokines, bile acids,
injury or toxins (Marzioni et al., 2002; Bogert and LaRusso,
2007; Glaser et al., 2006).
Functionally, cholangiocytes play an important role
in regulating localized liver immune responses through
secretion of cytokines and other mediators that influence
invading inflammatory cells. Cholangiocytes can also
interact with immune cells directly through expression
of adhesion molecules on the cell surface (Fava et al.,
2005; Adams and Afford, 2002; Glaser et al., 2009). They
are actively involved in the absorption and secretion of
water, organic anions, organic cations, lipids, electro-
lytes, and in the regulation of ductal bile secretion (Tietz
and LaRusso, 2006). Several hormones and locally acting
mediators are known to contribute to this cholangiocyte
fluid/electrolyte secretion and these include secretin,
acetylcholine, ATP, and bombesin.
In the liver, cholangiocytes contribute to bile secretion
via the release of bicarbonate in both the canaliculi and
the bile ducts that generates bile-salt independent flow
(Tietz and LaRusso, 2006; Xia et al., 2006). Bicarbonate
is secreted from cholangiocytes through mechanisms
which involve chloride efflux through activation of Cl−
channels, and further bicarbonate secretion via anion
exchange protein 2/solute carrier family 4 member 2
and secretin are two relevant hormones that act similarly
on their target cells (hepatocytes and cholangiocytes,
respectively). These hormones interact with specific G
protein-coupled receptors, increasing intracellular lev-
els of cAMP and activation of cAMP-dependent Cl− and
cholangiocytes appear to have cAMP-responsive intra-
cellular vesicles in which AE2/SLC4A2 co-localizes with
cell-specific Cl- channels (cystic fibrosis transmembrane
conductance regulator (CFTR) in cholangiocytes and
an undetermined protein in hepatocytes) and aquapo-
rins (AQP8 in hepatocytes and AQP1 in cholangiocytes)
(Banales et al., 2006). cAMP-induced coordinated traf-
ficking of these vesicles to canalicular or cholangiocyte
luminal membranes and subsequent exocytosis results
in increased osmotic forces and passive movement of
water with net bicarbonate-rich hydrocholeresis.
Cholangiocytes are also involved in the reabsorption
of biliary constituents like glucose and glutathione (Celli
et al., 1998; Bogert and LaRusso, 2007; Strazzabosco and
Fabris, 2008). For example, glutathione is catabolized
by the ectoenzyme γ-glutamyltranspeptidase (GGT)
expressed by the apical domain of cholangiocytes (also
expressed on the apical membranes of hepatocytes).
− exchange. Glucagon
− secretory mechanisms. Both hepatocytes and
Figure 6. Representative cell types of the liver and their corresponding autocrine and paracrine signals that are secreted in both health and disease.
510 E. L. LeCluyse et al.
Critical Reviews in Toxicology
The subsequent uptake of glutamate and cysteinyl-glycine
is crucial to avoid liver depletion of GSH. The importance
of cholangiocytes in liver function and disease has been
elucidated through the development of animal and cell-
based models that have enabled elucidation of their role
in the progression of liver disease (Glaser et al., 2009;
Strazzabosco and Fabris, 2008). Their importance is
further underscored by the number of diseases for which
cholangiocytes are the primary target, including primary
biliary cirrhosis (PBC), primary sclerosing cholangitis,
AIDS cholangiopathy, disappearing bile duct syndromes,
Alagille’s syndrome, cystic fibrosis, and biliary atresia.
1.2.6 Hepatic progenitor cells
HPC are bi-potential stem cells residing in human and
animal livers that are able to differentiate towards the
hepatocytic and the cholangiocytic lineages (Figure 6)
(Gaudio et al., 2009; Turner et al., 2011). The HPC reside
in a compartment contained within the canals of Hering.
These canals represent the smallest and most peripheral
branches of the biliary tree connecting the bile cana-
licular system with the interlobular ducts (Gaudio et al.,
In normal adult liver, HPC are small, quiescent cells
with elongated or vesicular nuclei, small nucleoli and
scant cytoplasm. Under normal circumstances they have
a relatively low proliferation rate and represent a reserve
compartment that is activated only when the mature
epithelial cells of the liver are continuously damaged or
inhibited in their replication, or in cases of severe cell
loss (Zhang et al., 2008). Under these conditions, resi-
dent HPC are activated and expand from the periportal
to the pericentral zone giving rise to mature hepatocytes
and/or cholangiocytes (Vig et al., 2006; Santoni-Rugiu et
al., 2005; Libbrecht et al., 2001). In rat liver, the HPC are
activated and induced to proliferate by various hepato-
carcinogens and other noxious stimuli whereupon their
nuclei acquire an oval shape, thus the name ‘oval cell’ in
the early literature (Grisham and Hartroft, 1961; Ogawa
et al., 1974).
The HPC niche is defined as the cellular and extra-
cellular microenvironment which supports the stem
cell populations and contributes to sustain self-renewal
and is composed of numerous cells, such as LSEC, HC,
cholangiocytes, KC, pit cells and other inflammatory
cells (Moore and Lemischka, 2006). All of these cells in
combination with numerous hormones and growth fac-
tors interact and cross-talk with progenitor cells influ-
encing their proliferative and differentiative processes.
The unique microenvironment and interaction with the
specific cell types is thought to be a key mechanism in
regulating the maintenance of self-renewal and matu-
ration capacities by stem cells. Nevertheless, a number
of different types of signaling and adhesion molecules
within the niche influence stem cell quiescence, self-
renewal and cell fate decisions. In fact, this niche envi-
ronment has been associated with regulating key stem
cell functions, such as maintaining stem cell quiescence
and providing proliferation- or maturation-inducing
signals when numerous progenitor cells are required to
generate mature cell lineages.
HPC activation and proliferation occurs under a num-
ber of extenuating circumstances by chemical, physi-
cal and mechanical means, and has been described in
various acute and chronic liver diseases (Bird et al., 2008;
Katoonizadeh et al., 2006). Regardless of the cause, activa-
tion of the HPC does not normally occur unless a signifi-
cant loss of mature cell mass has occurred. A threshold of a
50% loss of mature hepatocytes, together with a significant
decrease in proliferation of the remaining mature hepa-
tocytes, is required for an extensive HPC activation event
(Bird et al., 2008; Katoonizadeh et al., 2006).
HPC and their niche represent a potential target for
chemical- and drug-induced toxicity that can effect
liver regeneration and disrupt the molecular pathways
involved in cellular maturation leading to liver disease
and carcinogenesis (Katoonizadeh et al., 2006). The
inhibition of mature hepatocyte replication in long-term
chronic liver disease and chemical exposure is associated
with HPC activation. In several chronic liver pathologies,
the extent of HPC activation and proliferation is corre-
lated with the extent of fibrosis (Libbrecht et al., 2000).
The exact role of the HPC compartment in the causes or
adaptations of the liver to chemical- or drug-induced
injury is mostly unclear at this point. However, there is
sufficient evidence that the HPC represent an important
component of liver responses to chemical exposure and
need to be included in future strategies of toxicity testing.
1.3 Hepatocyte cytoarchitecture and cell polarity
Unlike other epithelia, which typically exhibit apical
(luminal) and basolateral (blood-facing) domains on
opposing surfaces of an epithelial sheet, hepatocytes
possess two basolateral domains that interface with the
sinusoidal microvasculature on opposite sides of the sin-
gle cell layers or plates (Figure 3) (Wolkoff and Novikoff,
2007). This configuration establishes a relatively unique
cytoarchitecture among epithelial tissues in that the api-
cal domain (i.e. bile canaliculus) lies midway between the
lateral domains of opposing epithelial cells, and there-
fore is wholly contained within the hepatic plates. The
canalicular domains, which have well-formed microvilli,
tight, intermediate and gap junctions, and desmosomes
clearly delineating their boundaries, begin as minute
intercellular channels which arise between adjacent cells
(Figure 4) (Khan et al., 2007). Most canaliculi form a belt-
like structure around the periphery of each hepatocyte
and interconnect with canaliculi from adjacent cells to
form an elaborate, anastomosing network of small tubu-
lar compartments (~0.5–1.0 µm diameter) throughout
the cell plates of the liver lobule. The networks of cana-
liculi within a cell plate terminate at the portal triad and
interconnect with bile ductules via the canals of Hering,
eventually draining into the common bile duct and the
gall bladder (Rodés et al., 2007). Both the canalicular
and sinusoidal surfaces have distinct cytochemical,
Organotypic liver culture models 511
© 2012 Informa Healthcare USA, Inc.
immunological and biochemical characteristics that are
crucial for maintaining normal hepatic function (Roman
and Hubbard, 1983; Hubbard et al., 1985; Maurice et al.,
1985; Stevenson et al., 1986; Chapman and Eddy, 1989).
1.3.1 Cell–cell interactions
184.108.40.206 Homotypic hepatocyte interactions
Adhesion of epithelial cells to one another is an important
process for the differentiation of multicellular organisms.
Disorders in intercellular communications and contact
are believed to play an important role in carcinogenesis
and the loss of normal growth control mechanisms
(Leibold and Schwarz, 1993; Krutovskikh et al., 1995).
Perturbations in the normal cell–cell interactions,
whether environmentally or intrinsically induced, can
affect a cell’s ability to respond normally to toxic insult
and, therefore, affect the normal disposition of drugs and
other xenobiotics (Kinch et al., 1995; Volberg et al., 1992).
In the intact liver, there are extensive lateral contacts
between hepatocytes. These include traditional epi-
thelial junctions such as tight junctions (which define
the barrier between apical and basolateral domains),
gap junctions (which facilitate direct intercellular com-
munication), intermediate (adherens) junctions and
desmosomes (which provide structural support and
integrity to eptithelial sheets) (Figures 3 and 4) (Hughes
and Stamatoglou, 1987). At the molecular level, the abil-
ity of cells to associate in a cell-specific manner involves
membrane-bound cell adhesion molecules. Cadherins
and connexins are the primary mediators of cell–cell
contacts and intercellular communication in epithelial
cells, respectively, and are considered to be essential
for maintaining tissue homeostasis and growth control
(Figure 4B) (Fladmark et al., 1997; Geiger and Ayalon,
1992). In addition to their role in maintaining the polarity
of hepatocytes and integrity of the epithelium, connexin-
mediated gap junction formation and membrane-
associated E-cadherin expression and distribution play
a critical role in the maintenance of normal cytochrome
P450 gene expression in hepatocytes and its regulation
by xenobiotic receptors (Hamilton et al., 2001).
The overall phenotype of hepatocytes and their
responsiveness to xenobiotic exposure is determined,
in part, by members of the Wnt signaling pathway and
the ρ-family of small GTPases (Nelson and Nusse, 2004;
Bustelo et al., 2007; Klaus and Birchmeier, 2008; Popoff
and Geny, 2009). Clustering of E-cadherin receptors
during cell contact and the establishment of cadherin-
mediated cell junctions depends simultaneously on
endogenous small GTPases (rhoA, rac1) and members
of the Wnt pathway, such as β-catenin (Braga and Yap,
2005; Popoff and Geny, 2009; MacDonald et al., 2009).
The Wnt pathway is necessary for maintaining nor-
mal regulation of liver regeneration and proliferation
of hepatocytes. Wnt signaling is regulated inside the
cell mainly by unbound levels of β-catenin, which is a
membrane-associated transcription factor that upregu-
lates genes involved in cell cycle control as well as cell
proliferation and motility (Baum and Georgiou, 2011). In
normal liver, β-catenin is one of the key proteins associ-
ated with E-cadherin-based junctional complexes (Braga
and Yap, 2005). Under circumstances where cell–cell
contacts are lost or perturbed (e.g., tissue damage, cer-
tain cell culture conditions), β-catenin is released from
the junctional complexes at the cell periphery and begins
to accumulate inside affected cells. When intracellular
concentrations of the unbound protein increase, nuclear
translocation and transcriptional activation of specific
target genes occurs, which in turn results in a switch in
phenotype from a quiescent to a proliferative state (Baum
and Georgiou, 2011).
Further evidence as to its role in tissue homeostasis
comes from studies that implicate somatic mutations in
the β-catenin gene, its cellular redistribution, and nuclear
accumulation in tumor formation and progression by
constitutively stimulating cell proliferation (Clevers,
2006). The activity of the small GTPases also play a role
in regulating the dynamics of the cytoskeleton as well as
interactions between cell junctions, cell-surface recep-
tors, and adhesion-dependent signaling pathways (e.g.
catenins, integrins) (Baum and Georgiou, 2011). Overall,
hepatocytes, like other epithelial cells, are dependent on
homotypic cell–cell contacts and the regulation of asso-
ciated signaling pathways for the maintenance of normal
structure and function.
220.127.116.11 Heterotypic cell interactions
Direct and indirect interactions and communications
between the different cell types of the liver play a much
greater role than originally appreciated in the main-
tenance of normal liver function and in xenobiotic-
induced hepatotoxicity both in vivo and in vitro (DeLeve
et al., 2004; McCuskey et al., 2005; Sunman et al., 2004).
Liver cells can affect one another through secretion of a
variety of paracrine factors (Figure 6). Paracrine signaling
is the primary form of regulation between parenchymal
cells and their partner NPC and represents the clas-
sic epithelial-mesenchymal relationship described by
embryologists and developmental biologists (Golosow
and Grobstein, 1962; Wang et al., 2010b). Coordinate
maturation of the parenchymal and NPC partners occurs
in association with lineage-dependent gradients of para-
crine signals (Wang et al., 2010b).
Although major emphasis has been placed on the role
of matrix chemistry and configuration in hepatocyte cul-
ture and differentiation (see section “Liver biomatrix”),
more than a dozen soluble signals have been identified
that change qualitatively and quantitatively with differ-
entiation (Wang et al., 2010b; Turner et al., 2011). Matrix
molecules such as proteoglycans (PG), and especially
heparin sulfate proteoglycan (HS-PG) and heparin pro-
teoglycan (HP-PG), have many growth factor-binding
sites determining growth factor storage, release, con-
formation, stability, affinities for specific receptors,
and other aspects of the signal transduction processes.
Therefore, soluble paracrine signals work synergistically
512 E. L. LeCluyse et al.
Critical Reviews in Toxicology
with the matrix components to dictate specific cell gene
expression patterns and the resulting phenotype (Reid
et al., 1992; Turner et al., 2011).
During drug- or chemical-induced liver injury,
injurious stimuli and stress signals cause activation
of LSEC, KC, polymorphonuclear leukocytes (PMN),
and platelets (PLT) and a release of various aggressive
mediators, enhancing the intrahepatic accumulation
of inflammatory cells (Vollmar and Menger, 2009; Shaw
et al., 2010). When activated during liver injury or
disease, KC produce oxygen-derived free radicals, nitric
oxide, eicosanoids, peptide leukotrienes, prostaglandins,
and various cytokines, including tumor necrosis factor
α (TNF-α), TGF-α, IL-1, IL-6 and others. KC and their
products are involved in modulating cell death by
inducing apoptosis in hepatocytes and other cell types
(Kolios et al., 2006; Roberts et al., 2007).
PMN, PLT, and LSEC can interact and bind to each
other, leading to further adhesion and accentuation of
mediator release. Upregulation of adhesion molecules
allows the firmly attached PMN to migrate towards
chemotactic signals (IL-8, cytokine-induced neutrophil
chemoattractant-1 [CINC-1], macrophage inflamma-
tory protein-2 [MIP-2], and KC), being released by intact
stress-exposed hepatocytes (De Pablo et al., 2010).
Tissue-infiltrating PMN exert direct hepatotoxicity by
reactive oxygen species (ROS) and hydrolytic enzyme
release. Hepatocytes undergo necrosis, apoptosis, or
mixed aponecrosis, depending on the severity of insult
and their intracellular ATP stores. Apoptotic and necrotic
hepatocytes further attract PMN by either surface expo-
sure of phosphatidylserine (PS) or leakage of high-mobil-
ity group box-1 (HMGB-1) (Klune and Tsung, 2010).
The interdependency of parenchymal cells and
their NPC companions places a severe constraint on
zone-specific patterns of gene expression and, thus, on
the inherent response to hepatotoxins in vivo. During
development, the NPC mature coordinately with the
epithelium, maturation associated with changes in the
paracrine signaling. The recent identification of specific
paracrine signals (both matrix and soluble) that control
the fate of liver stem and mature cells has been critical
to generating uniform cultures of liver parenchymal
cells maintained at a precise maturational stage (Wang
et al., 2010b; Turner et al., 2011). These advances are an
important consideration for the development of more
advanced in vitro model systems for studying chemical-
induced hepatotoxicity, toxicity pathway perturbations,
and networks of interactions among various cell types.
HPC activity is affected in a large part by both direct
and indirect heterotypic interactions (Gaudio et al.,
2009). The cellular environment surrounding the hepatic
stem cell niche is composed of numerous distinct cell
types, including HSC, LSEC, cholangiocytes, KC, pit cells
and other inflammatory cells, that provide signals to the
HPC influencing their proliferation and differentiation
through the provision of numerous signals within the
niche (Alison et al., 2009). The cellular microenvironment
causes changes in the surrounding matrix and endocrine
signal profiles, which in turn affect Wnt-mediated path-
ways involved in the HPC response (Yang et al., 2008; Hu
et al., 2007; Apte et al., 2008).
Inflammatory cells are responsible for producing a
range of cytokines and chemokines that may influence
the HPC response to liver injury (Alison et al., 2009).
For instance, T-cells express a TNF-like weak inducer of
apoptosis (TWEAK) which can stimulate HPC prolifera-
tion by engaging specific death receptor pathways. Other
inflammatory signals (e.g., IFN-γ and TNF-α) may stimu-
late hepatic progenitor cells to proliferate (Francis et al.,
2008; Jakubowski et al., 2005). Moreover, a resistance to
the growth-inhibitory effects of TGF-β may allow HPC
to proliferate under conditions which would otherwise
inhibit hepatocyte proliferation (Nguyen et al., 2007).
1.3.2 Liver biomatrix
Extracellular matrix directs and maintains both architec-
ture and phenotypic gene expression of liver cells. As with
all epithelia, cells are anchored to an insoluble matrix
that enables physical attachment to a substratum. In the
liver, this matrix is found in the Space of Disse between
the hepatocytes and the LSEC. Immunochemical analy-
sis of the Space of Disse from rat liver has shown that
the basal surface of hepatocytes in vivo are in intimate
contact with several extracellular matrix proteins such as
collagen (types I–IV), laminins, fibronectin, and HS-PG
(Martinez-Hernandez, 1984; Martinez-Hernandez and
Amenta, 1993; Bissell et al., 1990a; Bissell et al., 1987).
Actual matrix composition is typically a function
of zonal position, with discernible gradients from the
periportal region (zone 1) to pericentral region (zone 3)
(Figure 7) (Wang et al., 2010b; Turner et al., 2011). The
portal triads are dominated by fibrillar collagens (types I
and III), laminins (weak levels), vimentin, hyaluronans,
and less sulfated forms of CS-PG and HS-PG transition-
ing in gradient fashion through the Space of Disse to a
matrix chemistry around the central vein comprised of
type IV and VI collagens (with weak expression of type
III), syndecans 1 and 4, highly sulfated PG, especially
heparin PG, and no hyaluronans or laminins (Reid et al.,
1992). The fully differentiated hepatocyte lineages are
associated with network collagens (e.g. type IV and VI)
and forms of HS-PG with increasing sulfation ending in
HP-PG in zone 3. In addition, elastin is found generally
throughout the acinus, as is collagen type I8, a form of
HS-PG, both closely associated with the blood vessels.
These gradients in matrix chemistry are paralleled by
those of soluble signals, most being bound to various
matrix components, particularly the glycosaminoglycans
(GAGs) that are part of the PGs (McClelland et al., 2008;
Capila and Linhardt, 2002; Taipale and Keski-Oja, 1997).
The chemistry of the matrix works synergistically with the
soluble signals to dictate specific biological responses
from the cells. Indeed, the soluble factors are biphasic,
yielding mitogenic effects when complexed with the less
sulfated PG and causing growth arrest and differentiation
Organotypic liver culture models 513
© 2012 Informa Healthcare USA, Inc.
when complexed with the highly sulfated ones. These
effects are mediated by classic signal transduction path-
ways complemented by the mechanical effects of the
matrix (Wang et al., 2010b; Lozoya et al., 2011).
Although typical structures of epithelial basement
membranes are not uniformly observed along the
sinusoids from portal triads to central veins, collagen type
IV and some bound, small fibrils can be found forming
net-like, porous three-dimensional (3-D) lattices, serving
as scaffolding for the heaptocytes. Collagen type I bundles
can be viewed as the principal structure of the scaffolds
to which other collagen types, glycoproteins, and PG are
attached. In the space of Disse, small bundles of collagen
type I and fibers of collagen types III and VI can be identified
as well as some collagen type V, which is more abundant
near portal triads and central veins. Laminin, entactin/
nidogen, perlecan and collagen type IV are found in the
portal triad, whereas only perlecan and some collagen
type IV are found in the space of Disse. Fibronectins are
ubiquitous and prevalent throughout the scaffolds and are
especially abundant in the space of Disse, where they form
either fine filaments or granular deposits.
In addition to the direct role of the biomatrix in differ-
entiation, hepatic phenotype in vitro can also be affected
by biomechanical, adhesive, and structural aspects of a
biomatrix. The cytoskeletal structure and overall architec-
ture is integral to cellular phenotype (Ingber, 1993; Huang
and Ingber, 1999), and is largely regulated by the matrix
biology and chemistry (Mooney et al., 1992; Moghe et al.,
1996). Hepatocyte and hepatic plate-like architecture has
been reproduced in diverse culture systems that provide
cell attachment sites on two opposing sides of the cells
to allow for the appropriate localization of cell adhesion
molecules and cytoskeletal components (e.g. collagen
sandwich and EHS Matrix overlay) (Dunn et al., 1991;
Ezzell et al., 1993; LeCluyse et al., 1996a; Hamilton et
al., 2001). Modulation of adhesivity (Powers et al., 1997)
or compliance (Coger et al., 1997) of the biomatrix also
enhances functionality by enabling cells to spontaneously
aggregate and polarize with appropriate architecture.
Clearly, the influence of a biomatrix on hepatocytes is
more complex than chemical composition alone.
1.4 Liver biomechanical properties
The behavior of liver tissue and component cells is greatly
influenced by the biophysical and biomechanical prop-
erties of the extracellular environment. These phenom-
ena play a critical role in all aspects of tissue maturation,
from development to differentiation, and play an active
role throughout the life of an organism (Engler et al.,
2006; Griffith and Swartz, 2006; Swartz and Fleury, 2007).
In the liver, significant differences in the biomechanical
properties of extracellular matrix are observed. At the
hepatocyte/endothelial interface in the space of Disse,
matrix is soft and porous, while in the vicinity of cholan-
giocytes and stellate cells more rigid, cross-linked prop-
erties are present (Hayes et al., 2007; Reid, 1990; Turner et
al., 2011) (Figure 7). These properties are also manifest in
the behavior of hepatic stem cells, which differentiate as
hepatoblasts on more compliant materials, while matur-
ing towards cholangiocytes in more rigid microenviron-
ments (Turner et al., 2007; Turner et al., 2008; Turner
et al., 2011; Lozoya et al., 2011).
Transduction of biomechanical forces from matrices to
cells is primarily mediated through cell-surface recep-
tors (e.g. integrins) and the cytoskeleton (Figure 4).
A distinguishing feature of epithelial cells, including
hepatocytes, is the formation of “adhesion belts” around
the circumference of these cells at the lateral interface
(Stamatoglou and Hughes, 1994). These structures are
composed of actin filaments anchored to adherens junc-
tions and serve to provide a coordinated mechanical
interaction between epithelial cells. In addition to pro-
viding structural integrity for hepatic tissues, the cyto-
skeleton itself affects intracellular signaling cascades,
for example through the activation of the Wnt signaling
Figure 7. Diagram representing the zonal differences in lineage biology of hepatocytes, as well as the corresponding zonal differences in
extracellular matrix chemistry. The stem cell compartment is located in the portal triad region associated with the Canals of Hering (see section
“Hepatic progenitor cells”). These pluripotent stem cells can be stimulated to differentiate into either bile duct epithelial cells (cholangiocytes)
or parenchymal cells (hepatocytes). The cellular and biochemical composition of the different zones of the liver between the portal triad
and central vein is partially determined by the chemical make-up of the Space of Disse as well as other factors in the microenvironment,
such as other cell types, oxygen, nutrients, and endogenous/endogenous substrates. PV, portal vein; HA, hepatic artery; LAM, laminin; FN,
fibronectin; CS, chondroitin sulfate; HS, heparan/heparin sulfate.
514 E. L. LeCluyse et al.
Critical Reviews in Toxicology
and rho- and rac-associated proteins, which affect cellu-
lar phenotype (Klaus and Birchmeier, 2008; MacDonald
et al., 2009; Bustelo et al., 2007; Turner et al., 2011).
Overall, these studies highlight the importance of the
biomechanical properties of the extracellular matrix and
their effect on intra- and inter-cellular signal transduction
pathways in the context of tissue architecture and gene
regulation. The biochemical signaling effects imparted
by the chemical composition, geometrical configuration
and plasticity of the surrounding extracellular matrix is
a fundamental determinant of its overall influence on
tissue compliance and liver function during chemical
exposure and subsequent adaptive responses.
1.5 Liver hemodynamics
The liver is a highly-vascularized organ, receiving
~25–30% of the total blood volume at any given time.
Approximately 100 mL of blood passes through 100 g
of liver every minute, or a total of ~1.5 L per 1.5 kg adult
liver tissue (Bradley et al., 1945). The fundamental unit
of interest from a hemodynamic perspective on tissue
phenotype is the sinusoid, in which most molecular
delivery and transport occurs. Sinusoids commonly have
diameters ranging from 7 (periportal) to 15 µm (pericen-
tral) (Li, 2010; Vollmar and Menger, 2009). The resulting
shear stresses present in the liver are estimated to be on
the order of ~0.1–0.5 dyne/cm2 (Lalor and Adams, 1999).
These values are at the low end of shear stresses found in
other capillary systems of the body, which are typically
on the order of 15 dyne/cm2 (Koutsiaris et al., 2007). Liver
sinusoidal shear increases dramatically under condi-
tions such as reperfusion and partial hepatectomy, and
may play a role in initiating the liver regeneration cas-
cade (Schoen et al., 2001).
Physiological shear appears to play an important role
in facilitating phenotypic behaviors of vascularized tis-
sues under healthy and diseased conditions (Griffith and
Swartz, 2006; Hastings et al., 2007). However, the direct
shear stresses experienced by hepatocytes is difficult to
gauge as the effects of flow are mitigated by the separation of
hepatocytes from sinusoidal blood by LSEC and the space of
Disse. Convective introduction and removal of blood-borne
molecules, and the resulting gradients that are established in
the process, are more likely to be of direct relevance to hepato-
cyte phenotype and function – particularly when attempting
to reestablish such environments in vitro. However, LSEC
phenotype is affected in a significant way by the presence or
absence of direct flow and/or shear forces (Hastings et al.,
2007; Hastings et al., 2009; DeLeve et al., 2004).
The liver is continuously perfused with blood and all
associated nutrients, wastes, xenobiotics, etc. The result
is the formation of a dynamic environment within the
acinus where distinct gradients are created (Figures 1
and 2). Positional gradients within the sinusoid (i.e. from
zone 1 to zone 3) occur as molecules are metabolized
or synthesized along the sinusoids, or due to preferen-
tial consumption and/or transport in specific zones.
These gradients are generally a function of biochemical
properties and residence times within the liver alone
and do not typically rely on the interplay of other organ
systems. Oxygen tension provides an example of a posi-
tional gradient within the liver. Sinusoidal oxygen gra-
dients are fairly well established and range from ~60–70
mmHg (periportal) to 25–35 mmHg (perivenous) (Allen
and Bhatia, 2003). Temporal gradients, on the other hand
are typically the result of systemic synthesis/clearance,
and are a function of the rates of appearance/disap-
pearance and residence times in multiple tissues and
organs (e.g. absorption in the intestine, metabolism in
the liver, excretion in the kidney). Both types of gradients
are relevant for depicting hepatic physiological micro-
environments. The bioavailability, exposure levels and,
ultimately, the toxicity profile of most compounds are
influenced by the dynamic temporal and positional gra-
dients (e.g. clearance) resulting from the inherent ADME
properties of various tissues. In vivo these parameters are
dependent on hepatic and extrahepatic factors (ADME
profiles, heart rate, etc.).
2 the current state of cell-based hepatic
Tissue and cell culture provide an in vitro environment
for the maintenance, manipulation, and assessment of
cells under controlled conditions. For the purposes of
studying toxicological response to xenobiotic exposure,
an in vitro environment that mimics the inherent prop-
erties and natural relationship of tissues and cells as
they exist in vivo will generally provide a more accurate
portrayal of primary and secondary events depending
on the cellular and molecular complexity of the system.
In this section, the development of hepatocyte cell cul-
ture is reviewed from a historical perspective, noting the
advantages and disadvantages of conventional in vitro
hepatic culture models evaluated from the context of
basic biological principles and the resulting performance
limitations of the cells in culture. We also discuss recent
advances in tissue and cell culture technologies and how
they are being applied to address more complex cellular
and molecular events.
2.1 Past strategies for maintaining hepatic structure
and function in vitro
Cultured primary and immortalized hepatocytes have
been used for decades to address a wide variety of phar-
macological and toxicological research topics (Gebhardt
et al., 2003; Hewitt et al., 2007b; Hewitt et al., 2007a;
Guillouzo and Guguen-Guillouzo, 2008; Gomez-Lechon
et al., 2010). One shortcoming of conventional 2-D mono-
cultures of hepatocytes utilized traditionally for com-
pound testing is the partial or complete loss of viability
and phenotype over time in culture. When reflecting on
the various factors that dictate the expression of normal
hepatic phenotype in vivo, it is easy to understand that
much of the conditional loss of structure and function
in vitro is due to the loss of physiological context under
Organotypic liver culture models 515
© 2012 Informa Healthcare USA, Inc.
conventional culture conditions. In many respects, the
loss of normal cell structure and function in vitro is in
reality an adaptation to the preparation and cultivation
process that causes a shift in the gene program expressed
in the cells as requisite contextual signals are lost.
Hepatotoxicity in vivo is often dependent on specific
anatomical, morphological and phenotypic properties
of the individual cell types that comprise the liver
microenvironments in vivo. The three-dimensional
relationships of the unique cell types within the
microenvironments of the liver (e.g. periportal versus
pericentral), the regional hemodynamic flow patterns,
and other physiological factors, such as oxygen tension
and cytokine profiles, all play important roles in
determining the toxicokinetics and toxicity of particular
compounds. Current cell-based models that are
routinely utilized to perform toxicity testing in vitro are
generally simple culture platforms (typically standard
microtiter plate formats) employed under static,
nonphysiologic conditions. Due to their simplicity,
these static, monoculture model systems often
represent suboptimal models for drug and chemical
safety testing that are not able to mimic or predict
more complex MOA. One of the biggest challenges to
the development of more organotypic in vitro models
of the liver is the integration of the architectural and
cellular complexities of the liver, while incorporating
the important elements of the localized hemodynamics
of the regional microenvironments.
2.1.1 Historical perspective
Historically, several major complications have con-
founded the use of cultured hepatocytes for conduct-
ing long-term metabolism and toxicity testing. First,
there is variable attachment and rapid deterioration
of histotypic architecture, cellular polarity and func-
tionality of hepatocytes maintained on plastic culture
dishes (LeCluyse et al., 1996a). A second problem is the
lack of other relevant cell types (i.e. NPC) required for
mimicking normal functions and toxic mechanisms.
A third challenge involves supplying cultures with
adequate nutrients to carry out the wide array of cel-
lular functions performed by hepatocytes in vivo (e.g.
phase 1 and 2 biotransformation reactions, synthesis
of bile acids and serum proteins). A number of these
issues have been addressed to some extent for certain
applications, while others continue to be biologically
and technically challenging (Guillouzo and Guguen-
Guillouzo, 2008; Gebhardt et al., 2003).
Over the past several years several modifications to
conventional culture conditions have improved hepatic
function and longevity of primary rat hepatocytes
(LeCluyse, 2001; LeCluyse et al., 1996a). Co-culture
with fibroblasts or rat liver biliary epithelial cells
(Guguen-Guillouzo et al., 1983; Donato et al., 1990;
Kuri-Harcuch and Mendoza-Figueroa, 1989) and the
use of complex extracellular matrix substrata (liver bio-
matrix, EHS extracts) (Rojkind et al., 1980; Bissell et al.,
1987) prolong the functional lifespan of hepatocytes.
In addition, several different approaches have been
employed in an effort to preserve hepatocyte function
by manipulating the extracellular matrix geometry or
configuration. Overlaying the hepatocyte cultures with
an additional layer of extracellular matrix (sandwich
configuration) results in striking improvements in
hepatocyte morphology and liver-specific gene expres-
sion (Dunn et al., 1991; Sidhu et al., 1994; Musat et al.,
1993; LeCluyse et al., 1994).
2.1.2 Extracellular matrix effects
An important influence on the maintenance of normal
hepatic structure and function in vitro are the cellular
interactions with the surrounding extracellular matrix
(Bucher et al., 1990; Ichihara, 1991; Martinez-Hernandez
and Amenta, 1995). The nature of the extracellular matrix
interactions with cultured hepatocytes determines both
cell shape and cytoarchitecture which, in turn, are related
to the expression of transcription factors and gene pro-
grams (Ingber, 1993; Nagaki et al., 1995; Rana et al., 1994).
The chemical composition and biophysical characteris-
tics of extracellular matrices used in hepatocyte cultures
profoundly affect both liver-specific gene expression and
cellular response to extracellular soluble signals.
In vivo, extracellular matrix consists of a mosaic of
lipids, proteins and carbohydrates in a complex, hetero-
geneous and dynamic environment (see section “Liver
biomatrix”). The simplest approach to cultivating mature
hepatocytes in vitro has been on individual components
of extracellular matrix, or combinations of these vari-
ous forms. Use of films of individual extracellular matrix
proteins (collagens type I, III or IV, fibronectin, laminin)
as substrata does not greatly improve preservation of
differentiated functions (Bissell et al., 1987; Sawada
et al., 1987; Ben-Ze’ev et al., 1988). However, addition of
individual components of liver biomatrix, such as PG or
related GAGs, to hormone- and nutrient-enriched media
increases the levels of mRNA for albumin and some
other liver-specific proteins, while lowering abnormally
high mRNA levels for cytoskeletal proteins, such as actin,
suggesting that the presentation of hepatic matrix com-
ponents in vitro affects cellular function. Type I collagen
in hydrated gel form rather than as a dried film enhances
the stabilization of liver-specific mRNA (Zaret et al., 1988)
and delays, but does not prevent, loss of differentiation
(Michalopoulos and Pitot, 1975; Sirica et al., 1979; Ben-
Ze’ev et al., 1988). However, it is unclear whether this
effect is due to the specific presence of type I collagen or
to the mechanical properties of the hydrated gel.
Impressive results have also been obtained with
complex mixtures of extracellular matrix components,
notably with liver-derived “biomatrix” (Enat et al., 1984;
Wang et al., 2011) or with Matrigel, a biomatrix preparation
derived from the Engelbreth-Holm-Swarm (EHS)
sarcoma (Orkin et al., 1977). Used as a substratum for cell
attachment, cells do not flatten but retain a rounded shape
and aggregate in clusters or columns that thicken with
516 E. L. LeCluyse et al.
Critical Reviews in Toxicology
time (Bissell et al., 1987). Many liver-specific functions
appear to be wholly or partly preserved although some are
still lost progressively. For instance, albumin secretion is
maintained at 40–100% of in vivo levels (partly depending
on the medium formulation employed), reflecting partial
preservation of normal mRNA levels (Bissell et al., 1987;
Schuetz et al., 1988). The abnormally high expression of
actin and tubulin genes and appearance of α-fetoprotein
frequently observed in simple culture conditions are also
absent or suppressed on Matrigel (Schuetz et al., 1988;
Ben-Ze’ev et al., 1988; Lindblad et al., 1991).
Biomatrix is a complex, partially purified extract of
extracellular matrix material prepared from whole rat
liver and contains types I–IV collagen, fibronectin and
extracellular matrix glycoproteins, including a number
of important PG and growth factors (Rojkind et al., 1980;
Wang et al., 2011). When compared with gelled collagen,
liver biomatrix enhances hepatocyte attachment and
survival for longer periods (3 weeks or more) in culture.
Albumin gene expression also has been reported to be
significantly higher in cultures maintained on biomatrix
compared to those on collagen. Hepatocytes cultured
on liver biomatrix attached preferentially to areas of the
substratum which stained intensely for glycoproteins
(PAS-positive) (Reid et al., 1980). In addition, these
PG-rich areas of the substratum supported long-term
survival of hepatocyte cultures. The levels of liver-specific
functions of mature liver cells maintained on biomatrix
scaffolds for weeks proved to be the same or similar to
those of freshly isolated, adult hepatocytes (Wang et al.,
2011). Hepatocytes on type I collagen deteriorated rap-
idly within 2 weeks, whereas those on biomatrix scaf-
folds remained stable morphologically and functionally
for more than 8 weeks. Overall, the presence of tissue-
specific matrix components similar to those found in
the space of Disse may be a key factor in the success of
biomatrix as a substratum for hepatocyte cultures.
2.1.3 Adhesive and mechanical factors
Physical properties of a matrix can play an equally impor-
tant role in vitro. Notable among these are the impact of
cell-surface adhesion and matrix compliance on cellu-
lar architecture and phenotype. These properties affect
hepatic function through control of hepatocyte spread-
ing and permissiveness towards formation of intercellu-
lar adhesion and junctional complexes. The mechanical
and adhesive properties of matrices influence function-
ality of cultured hepatocytes on rigid surfaces regardless
of molecular composition. Hepatocytes on matrices that
are both rigid and highly adhesive tend to exhibit spread
morphologies with extensive stress fibers (LeCluyse et
al., 1996a; Hamilton et al., 2001). Cells attach and spread,
and do not generally exhibit epithelial polarity. Rodent
cells typically lose function and viability rapidly over
hours-to-days. These results appear to be due solely to
the physical properties of the matrix, as similar behav-
iors can be observed on matrices traditionally associ-
ated with “physiological” composition, such as Matrigel,
when their physical properties are modified to increase
cell spreading (Powers et al., 1997; Coger et al., 1997).
Hepatocytes on compliant or lower adhesive matrices
exhibit more extensive intercellular interactions, histo-
typic cytoskeletal organization and physiological pheno-
type (Mooney et al., 1992; Powers et al., 1997). Functional
properties typically associated with culture on a specific
matrix type may also be attributed to the physical prop-
erties of that matrix. Both the composition of a matrix
(collagen, laminin, fibronectin, HSPG, etc.) and physical
and mechanical properties of that matrix (surface ligand
density, compliance, etc.) are important.
Matrix geometry can also play a significant role in cel-
lular function. Adult rat hepatocytes cultured in a ‘sand-
wich’ configuration, i.e. between two layers of gelled
collagen type I, reconstructing the opposing sinusoidal-
facing domains of hepatic plates in vivo, remain viable
for prolonged periods of time and maintain normal levels
of secretion of several liver-specific proteins and organic
compounds (Dunn et al., 1989; Dunn et al., 1991). In
addition, hepatocytes maintained in a sandwich configu-
ration exhibit a more normal distribution of microtubules
and actin filaments, and they respond to prototypical
cytochrome P450 enzyme inducers in a more physiologi-
cal fashion (Dunn et al., 1991; Ezzell et al., 1993; LeCluyse
et al., 1999; Hamilton et al., 2001).
2.1.4 Three-dimensional spheroid aggregate culture
Primary hepatocytes when cultured on non-adhesive
surfaces under appropriate conditions will form small
sphere-shaped aggregates or “spheroids” over several
days (Landry et al., 1985; Li et al., 1992; Dilworth et al.,
2000). These structures deposit extracellular matrix
material on the outer surface essentially encapsulating
the spheroids. Immunolabelling techniques have shown
that the entire spheroidal structure is delineated by a
discrete zone of extracellular matrix material containing
laminin, fibronectin, and collagen (Landry et al., 1985;
Tong et al., 1990). In addition, cell survival and many
differentiated functions are maintained for prolonged
periods of time in spheroid culture (Brophy et al., 2009;
Sakai et al., 2010a; Sakai et al., 2010b).
Many of the beneficial effects of spheroidal aggregate cul-
ture are attributed to the retention of a three-dimensional
cytoarchitecture, the presence of key extracellular matrix
components, and the establishment of important cell–cell
contacts (Landry et al., 1985; Takabatake et al., 1991; Yuasa
et al., 1993; Luebke-Wheeler et al., 2009; Sakai et al., 2010a;
Sakai et al., 2010b). Spheroids also appear to facilitate the
segregation of cells in a histotypic manner. Mixed cultures
of hepatocytes and endothelial cells have been shown to
self-sort in a histotypic fashion, with the endothelial cells
forming a thin layer at the tissue-fluid interface, i.e. the
periphery of the structure, and bile canaliculi forming at
the center cores (Powers and Griffith, 1998).
dexamethasone, glucagon, insulin, and EGF enhances
integrity of the spheroids and maintains production
Organotypic liver culture models 517
© 2012 Informa Healthcare USA, Inc.
of albumin, glucokinase, and transferrin for up to 60
days in suspension culture (Tong et al., 1990; Yuasa
et al., 1993). Some liver-specific functions, such as
albumin production and tyrosine aminotransferase
(TAT) induction by glucagon and dexamethasone,
are maintained at high levels for up to two months
compared to hepatocyte cultures maintained on a
simple collagen substratum where a complete loss of
TAT activity and albumin production is observed by 1
and 2 weeks, respectively (Landry et al., 1985; Koide
et al., 1989; Tong et al., 1992; Yagi et al., 1995; Koide et
al., 1990). Induction of CYP enzymes by prototypical
inducers (e.g. 3-methycholanthrene, dexamethasone,
phenobarbital [PB]) is retained in spheroid cultures and
remains relatively constant for up to 3 weeks. Spheroid
cultures also retain their ability to respond to peroxisome
proliferators even after 12 days in culture. For example,
treatment with the peroxisome proliferator nafenopin
caused a 4.5-fold increase in cytoplasmic volume
fraction of peroxisomes with a concomitant induction of
peroxisomal bifunctional enzyme and CYP4A, enzyme
markers associated with peroxisome proliferation
(Roberts and Soames, 1993). Similarly, thyroid hormone
(T3) induced the rate of transcription of 5′-deiodinase
mRNA in spheroid cultures, mimicking its response in
vivo (Menjo et al., 1993).
Recent developments in spheroid generation have
focused on the use of scaffolds to facilitate the aggrega-
tion of hepatocytes while controlling the resulting size
of the structures (Dvir-Ginzberg et al., 2003; Napolitano
et al., 2007). Because of oxygen diffusion limitations and
extensive metabolism by hepatocytes, it is important to
limit the dimensions of hepatocyte spheroids to less than
~300 microns in diameter to avoid necrosis of the central
regions (Yarmush et al., 1992; Powers et al., 2002a; Powers
et al., 2002b; Glicklis et al., 2004). Specific scaffolds have
been designed to overcome challenges inherent with
suspension culture (i.e. uncontrolled or inconsistent
diameter), and typically shorten the time required for
spheroid formation by facilitating intercellular contact.
Although spheroid aggregate culture techniques have
been utilized for many years, they have found application
mainly in bioreactor or artificial liver devices (Tostões et
al., 2011; McIntosh et al., 2009). There are few publica-
tions that report employment of this technique for tox-
icity testing; however, spheroids have been used in the
generation of culture systems that appear to be promising
for toxicology applications (Powers et al., 2002b; Powers
et al., 2002a). Overall, further evaluation of spheroids as
a tool to study chemical effects over prolonged exposure
periods appears worthwhile (Meng, 2010).
2.1.5 Co-culture systems
Adult rat hepatocytes when cultured in combination
with rat liver epithelial cells (RLEC) have higher levels
of albumin secretion and survive for longer periods
of time compared to monocultures of hepatocytes on
collagen-coated plates (Guguen-Guillouzo et al., 1983;
Clément et al., 1984; Lescoat et al., 1985). Co-cultures of
hepatocytes and RLEC exhibit a more robust response
of acute phase response genes to treatment with inflam-
matory cytokines than hepatocytes alone (Peters et al.,
2010). Notably, restoration of hepatic function in co-
cultured hepatocytes is concomitant with the deposition
of an elaborate, highly organized network of extracellular
matrix material between the two cell types (Guguen-
Guillouzo et al., 1983). These fibrils has been determined
to contain most of the elements of the extracellular matrix
components found in vivo, such as collagen types I, III,
and IV as well as fibronectins and laminins (Clément et
al., 1988c; Clément et al., 1988b; Clément et al., 1988a).
Co-culturing also appears to enhance gap junctional
intercellular communication as indicated by increased
dye-coupled communication (Rojkind et al., 1995).
Co-cultures of rat hepatocytes cope better with oxidative
stress than hepatocyte cultures alone as indicated by the
significantly lower levels of glutathione peroxidase and
reductase expressed after exposure to different oxygen
levels (Mertens et al., 1993). Moreover, levels of glutathi-
one are also more stable in co-cultures of hepatocytes
and epithelial cells (Mertens et al., 1991).
Most of the facilitative effects of the co-culture system
can be provided by a variety of primary and transformed
epithelial and mesenchymal cell lines (Morin and
Normand, 1986; Kuri-Harcuch and Mendoza-Figueroa,
1989; Donato et al., 1990; Donato et al., 1991; Khetani
and Bhatia, 2008). The particular combination of cell
adhesion molecules (i.e. cadherins, integrins) involved
in the cell–cell and cell–matrix contacts and soluble fac-
tors (autocrine and paracrine) expressed in co-cultures
of hepatic cells may regulate the re-establishment of
integral signal transduction pathways in concert with
cytoskeletal redistribution, which in turn lead to the
expression of liver-specific transcription factors and
sustained phenotypic structure and function (DeLeve et
al., 2004; Rosales et al., 1995; Khetani and Bhatia, 2008;
Mammoto and Ingber, 2010; Wang et al., 2009).
In addition to the supportive role of other cell types
in maintaining normal hepatic structure and function
in vitro, co-cultures have also been utilized to study the
direct and indirect signaling pathways involved in certain
drug- and cytokine-induced effects on hepatocyte func-
tion (Wandzioch et al., 2004; Sunman et al., 2004; Tukov
et al., 2006). For example, co-cultures of primary hepato-
cytes and Kupffer cells were employed to elucidate and
reproduce the in vivo effects of interleukin-2 (IL-2) on
CYP3A4 clearance of drugs (Sunman et al., 2004). Parallel
cultures of hepatocytes alone did not produce the same
effects, suggesting that Kupffer cell activation and pro-
duction of secondary mediators (e.g. IL-6, TNFα) were
necessary to elicit the response.
For the most part, co-culture systems satisfy nearly
all of the biophysical requirements for optimal mainte-
nance of hepatocytes in vitro (i.e. cell shape, cell–cell and
cell–matrix contacts). However, the loss of key hepatic
functions (e.g. cytochrome P450 isoforms) serves to
518 E. L. LeCluyse et al.
Critical Reviews in Toxicology
underscore the role of other hemodynamic, microen-
vironmental and/or soluble factors in determining the
overall hepatocyte phenotype in vivo (Guillouzo et al.,
1990; Rogiers et al., 1990; LeCluyse et al., 1996a).
2.1.6 Perifusion culture systems
Maintenance of primary liver cells under dynamic flow
conditions dates back over three decades (Gebhardt and
Mecke, 1979a). Perifusion culture allowing the continuous
perfusion of the cell monolayer with culture medium avoids
many disadvantages of conventional static culture systems
and generally improves viability, lifespan, and metabolic
performance of cultured primary hepatocytes (Gebhardt
and Mecke, 1979a; Gebhardt et al., 1996). Comparable
findings have been observed with considerably different
perifusion systems (De Bartolo and Bader, 2001; Dich and
Grunnet, 1992; Gebhardt et al., 1996; Takeshita et al., 1998).
Perifused hepatocytes regain normal hormonal sensitivity
under conditions where the same cells maintained under
conventional static culture conditions did not respond to
hormones at all (Gebhardt and Mecke, 1979b). They also
showed an enhanced sensitivity toward mitogens and a
more physiological response to the growth-promoting
effects of several carcinogens (Gebhardt and Fischer, 1995;
Klein and Gebhardt, 1997). Cytochrome P450 levels and
EROD activity were stabilized or could be easily induced
in the perifused hepatocytes, suggesting that this culture
method has great promise for long-term metabolism
studies (Gebhardt and Mecke, 1979b; Gebhardt et al.,
2.2 Lessons learned and where we go from here
The means to cultivate well-differentiated hepatocytes
in vitro for prolonged periods of time (>2 weeks) with
full metabolic capacity has been a goal for some time.
Currently, however, there is an even greater sense of
urgency to develop improved in vitro model systems to
better understand the temporal relationship between
physiologic exposure to a compound or its metabolites
and the ensuing sequence of subcellular events that even-
tually lead to adverse responses in humans. In addition,
a multicellular system that reproduces or mimics subse-
quent adaptive and immune responses, as well as cellular
regenerative processes (i.e. stem cell responses) would be
of great value as well. Significant progress has been made
towards understanding the key interactions and synergis-
tic roles between the various cell types, important aspects
of flow dynamics and zonal microenvironments, and the
role of biochemical composition and configuration of the
extracellular matrix on hepatocyte biology and chemical-
induced toxicity. Understanding these parameters is a pre-
requisite to creating hepatic culture models that mimic the
types of chemical-induced toxicities observed in vivo and
that are capable of reproducing the complex perturbations
of key cellular pathways along with the subsequent adap-
tive responses over time. Consequently, the focus of recent
efforts has been the development of organotypic cell cul-
ture platforms that allow the maintenance of hepatocytes
and other cell types under more physiologic conditions
whereby the native architecture and phenotype is restored
and maintained for weeks if not months.
2.3 Current challenges for today’s model systems
Many drugs and other xenobiotics present in the portal
and systemic blood are taken up by hepatocytes by spe-
cific transport proteins where they can be metabolized
by CYP and other enzymes involved in phase 1 and 2
biotransformation reactions prior to elimination from
the cell or tissue by efflux transporters. Much remains
to be learned about these detoxication systems, includ-
ing the basis for species differences in their activity and
specificity, pathways that control their expression and
regulation, their endogenous and exogenous substrates,
and the rate-controlling step(s) in hepatic uptake,
metabolism and excretion of hepatotoxins. More sophis-
ticated in vitro systems that retain these important facets
of liver biology are needed to evaluate hepatic uptake
and metabolism, cytochrome P450 induction, chemical
interactions affecting hepatic metabolism, hepatotoxic-
ity, and cholestasis (Gebhardt et al., 2003; Houck et al.,
2009; Judson et al., 2010). The development of more
physiologic, organotypic hepatic culture systems that
maintain liver structure and function over longer periods
of time will also permit further understanding of com-
plex mechanisms of hepatotoxicity, the identification of
key stress pathways and development of more predictive
computational models (Judson et al., 2010; Clewell et al.,
2008; Houck and Kavlock, 2008). In fact, activation of the
molecular pathways involved in determining whether
chemical-induced perturbations translate into an adap-
tive or toxic response often occurs over a period of many
days or weeks. Therefore, culture systems are needed
that maintain phenotypic function for weeks or even
months. Likewise, some toxic responses require direct or
indirect interactions between the multiple cell types to
mimic an in vivo-like response (e.g. acetaminophen- and
LPS-induced toxicity) (DeLeve et al., 1997; Kmiec, 2001;
Roberts et al., 2007).
The response of individual liver cells to chemical
insult in vivo depends on the microanatomy and
the local microenvironments within the organ. The
complex relationship between cell types and how they
adapt to chemical exposure is dynamic (involving both
concentration and rate changes over time) and cannot
easily be mimicked by conventional static monolayer
culture systems. For instance, monocultures of
hepatocytes maintained under static culture conditions
lack important direct and indirect communications
with other relevant cell types, namely LSEC, HSC and
KC, which are involved in mediating many mechanisms
of chemical-induced toxicity. Using simple cultures of
hepatocytes alone does not account for the concerted
response between cells that occurs in vivo, nor does it
reflect the differential sensitivity to known hepatotoxins
that occurs in a zone-specific fashion (Anundi et al.,
1993; DeLeve et al., 1997; Edling et al., 2009). These
Organotypic liver culture models 519
© 2012 Informa Healthcare USA, Inc.
systems also lack the replenishment of crucial nutrients
and other cofactors (e.g. GSH, PAPS and UDP-GA) and
the removal of waste products from the break-down of
both endogenous and exogenous substrates, a deficiency
that can make the exposed cells more vulnerable to self-
generated and artificial levels of a multitude of stress-
inducing substances. Notably, hepatocytes maintained
under simple static culture conditions often exhibit
greater sensitivity to hepatotoxins than those maintained
under more physiologic conditions (Richert et al., 2002).
Mimicking the intricate anatomy of the liver involv-
ing the hepatobiliary elimination of compounds and the
nature of the interface between the canalicular networks,
the canals of Hering and the bile duct system is a daunt-
ing bioengineering challenge. An in vitro system that can
recreate the complex three-dimensional architecture of
the liver and multicellular relationship of the sinusoidal
and bile duct systems does not exist at the present time.
Moreover, compounds may cause cholestasis through
either impairment of bile secretion by the hepatocyte or
bile duct injury involving the ductules or the interlobular
ducts (Chazouilleres and Housset, 2007). The complex
mechanisms of chemical-induced canalicular and chol-
angioidestructive cholestasis are nearly impossible to
reproduce within the context of our current cell culture
devices (Cullen and Ruebner, 1919; Chazouilleres and
Liver function and disease is also tied intimately
with the endocrine system and endocrine disorders are
often associated with liver abnormalities (Silverman et
al., 1989; DeSantis and Blei, 2007). Non-alcoholic fatty
liver disease and diabetes mellitus are two of the most
prevalent diseases in the US today. Both endocrine-
based diseases of the liver can cause major changes in
liver function, cytokine levels and response to chemi-
cal exposure which cannot currently be reproduced
in vitro unless cells are procured directly from those
tissues (Angulo and Lindor, 2002; DeSantis and Blei,
2007). Moreover, synthetic oral androgen and estrogen
compounds can be associated with cholestasis and the
development of benign and malignant tumors of the liver
(Steinbrecher et al., 1981; Ishak and Zimmerman, 1987;
Nakao et al., 2000). The complex systems biology involv-
ing multiple organ systems and cellular pathways in vivo
makes it difficult, if not impossible, to reproduce or study
their role in xenobiotic-induced toxicity in vitro.
Despite these challenges and limitations for recreat-
ing all of the key structural and functional components
of the liver inside the laboratory, major efforts to create
improved model systems of the hepatic micro-architec-
ture and to reproduce some of the key cellular interactions
involved in many xenobiotic-induced hepatotoxicities
are currently part of several academic and industry pro-
grams. For developing in vitro systems intended to model
the basic liver sinusoidal architecture and related cell
interactions, lessons learned from past experiences indi-
cate that there are at least four key areas that need to be
considered for recreating and maintaining liver-specific
structure and function in vitro: (1) extracellular matrix
composition and geometry, (2) cell–cell interactions
(both homo- and heterotypic), (3) dynamic flow and (4)
medium formulation, including various endocrine fac-
tors. In addition, there are a growing number of studies
suggesting the importance of histotypic architecture and
tissue organization in the restoration of phenotypic gene
expression and responsiveness to chemical exposure
(DeLeve et al., 2004; Hamilton et al., 2001; Hastings et al.,
2009; Peters et al., 2010; Wang et al., 2010a).
3 Considerations for the development of
organotypic liver models
3.1 Source of cellular material
3.1.1 Primary cells
Freshly isolated primary hepatocytes are the preferred
cell model for recapitulating the functional responses of
the liver, especially for in vitro studies to predict in vivo
drug metabolism and clearance (Lin, 2006; Hewitt et al.,
2007b; Hewitt et al., 2007c; Gomez-Lechon et al., 2008;
Obach, 2009). Primary cells when cultured under proper
conditions express all the major metabolizing enzymes
and transporter proteins in their native configuration
(LeCluyse et al., 1996a; Hamilton et al., 2001). Primary
hepatocytes are typically isolated from intact liver tissue
by collagenase digestion, and purified through a series
of low-speed, density-gradient centrifugations steps
(LeCluyse et al., 1996b; LeCluyse et al., 2005). Using this
method, it is possible to obtain primary hepatocytes from
animal and human liver tissues that retain high levels of
transport and metabolic functions. Although primary
hepatocytes initially do contain metabolic enzymes at
their physiological levels immediately after isolation,
most liver-specific gene expression and CYP-related
functions decrease during the initial stages of cultivation.
This loss of gene expression and concomitant decrease
in function over time can be mitigated by culturing the
cells with an appropriate medium formulation, on com-
plex extracellular matrices and/or with other cell types as
discussed earlier in this document.
While primary hepatocytes offer significant functional
benefits, their routine use in cell culture systems presents
several challenges. The most significant is the difficulty
in obtaining primary human cells and the tissues from
which they are isolated. As a result, commercial ven-
dors have become an important source for obtaining
primary cells, particularly those of human origin. Recent
advances in the cryopreservation of human hepatocytes
has enhanced the convenience and capabilities associ-
ated with the use of primary cells, removing limitations
requiring culture of hepatocytes within hours of isolation
and enabling repeat experimentation with cells from
an individual donor. When properly prepared, cryo-
preserved hepatocytes typically exhibit similar viability
and function after thawing compared to freshly isolated
hepatocytes (Li et al., 1999; McGinnity et al., 2004;
Richert et al., 2006; Li, 2010). However, locating a reliable
520 E. L. LeCluyse et al.
Critical Reviews in Toxicology
commercial source of fresh or cryopreserved hepato-
cytes from other key toxicology species, particularly dog,
monkey and mouse, can be problematic.
3.1.2 Immortalized cell lines
A cell line is a permanently established, transformed
clonal lineage, where the daughter cells will proliferate
indefinitely when given proper medium and growth
substratum conditions. In contrast to primary cell
cultures, cell lines are not restricted to a limited number
of cell divisions due to mutations in one or more growth
control pathways (Mees et al., 2009; Shay and Wright,
2005), and therefore have become immortalized. Liver cell
lines in conventional systems are a very popular in vitro
model to study liver function and general mechanisms
of toxicity. However, they are typically unsuitable for
drug metabolism and toxicity prediction, due to the fact
that cell lines do not contain all the metabolic enzyme
families, and the enzymes that are present are not at their
physiological levels. Disadvantages are the dependence of
gene expression on passage number, genomic instability,
leading to dedifferentiated cells whose phenotype no
longer resembles that of the cell in vivo.
While cell lines have a number of undesirable proper-
ties, an important benefit in the use of human cell lines
is that they provide a readily available source of cells that
can generate data relevant to humans. Moreover, they
are easy to handle and replace the use of animals. Several
hepatic cell lines including HepG2, C3A (a sub-clone
of the hepatoma-derived HepG2 cell line), HepaRG
(Vermeir et al., 2005; Castell et al., 2006; Kanebratt and
Andersson, 2008b; Kanebratt and Andersson, 2008a)
and the Fa2N-4 cell line (Mills et al., 2004; Ripp et al.,
2006; Youdim et al., 2007; Hariparsad et al., 2008) have
been assessed as candidates to replace primary human
hepatocytes in CYP induction and metabolism stud-
ies (Kanebratt and Andersson, 2008b; Kanebratt and
Andersson, 2008a; Jennen et al., 2010; Wilkening et al.,
2003). These hepatic cell lines can be a reasonable alter-
native to primary cells for use in dynamic flow organo-
typic devices, especially for the initial proof of concept
studies, because they are readily available, support long-
term culture, and maintain some hepatocyte functions in
vitro. The caveat is that important metabolic and recep-
tor pathways are likely to be deficient in these cell lines
compared to primary liver cells.
The most commonly used and best characterized
human liver cell line is the HepG2 cell line. HepG2
cells are derived from a liver tissue with a well differ-
entiated hepatocellular carcinoma. They are adherent,
epithelial-like cells when grown as monolayers and in
small aggregates, have a model chromosome number of
55, and are non-tumorigenic. HepG2 cells secrete typi-
cal hepatic plasma proteins, such as albumin, transfer-
rin, fibrinogen, α-2-macroglobulin, and plasminogen,
and carry out biotransformation of many, but not all,
xenobiotic compounds. They are capable of bioactivating
mutagens and carcinogens, and carry no p53 mutations
enabling them to activate DNA damage response, induce
growth arrest, and initiate apoptosis (Hsu et al., 1993b;
Knasmuller et al., 1998; Wilkening et al., 2003). Because
HepG2 cells are easy to maintain compared with primary
human hepatocytes, they are frequently employed in
various toxicogenomics studies and, despite their insen-
sitivity to TNF-α, they are the most frequently used cell
type for examining the effects of cytokine regulation on
hepatic acute phase protein synthesis (Burczynski and
Penning, 2000; Harries et al., 2001; Hong et al., 2003;
Jennen et al., 2010).
Comparisons between HepG2 and primary hepato-
cytes at the transcriptome level show substantial dif-
ferences in basal gene expression (Harris et al., 2004;
Liguori et al., 2008; Olsavsky et al., 2007). For instance,
HepG2 show higher expression of genes involved in cell
cycle regulation, DNA, RNA, and nucleotide metabo-
lism, transcription, transport, and signal transduction,
and lower transcription levels are associated with cell
death, lipid metabolism, and xenobiotic metabolism. By
contrast, basal gene expression levels of phase 1 and 2
biotransformation enzymes (CYP1A1, CYP1A2, CYP2C9,
CYP2E1, and CYP3A4) are substantially lower in HepG2
when compared to primary hepatocytes. The inherent
lack of bioactivation potential leads to an underestima-
tion of metabolic-dependent toxicity for particular com-
pounds, such as aflatoxin B1, making HepG2 cells a less
predictive in vitro model system (Olsavsky et al., 2007;
Westerink and Schoonen, 2007; Wilkening et al., 2003).
Several alternative variants or subclones of HepG2
were introduced to address some of the shortcomings
with the original cell line (e.g. HepG2/C3A). For example,
HepG2 can be transfected with constructs which express
increased levels of phase 1 enzymes (such as CYP1A1,
CYP1A2, CYP2E1 and CYP3A4) or glutathione-S-trans-
ferases (Vermeir et al., 2005; Knasmuller et al., 1998).
Nevertheless, HepG2 and the various subclones that
have been identified (including C3A) provide a biologi-
cal model that enables some rudimentary approxima-
tion of hepatic function that offers some value in certain
applications, but these cells typically fall well short of
recapitulating many important aspects of primary hepa-
tocyte function and phenotype.
Fa2N-4 cells are derived from primary human hepatocytes
immortalized by transfection with the SV40 large
T-antigen (Mills et al., 2004; Vermeir et al., 2005; Ripp et
al., 2006). The Fa2N-4 cell line maintains morphological
characteristics of primary human hepatocytes and is non-
tumorigenic. Initial studies have shown that various P450
enzymes (CYP3A4, CYP1A2, and CYP2C9) are inducible
in Fa2N-4 cells after exposure to several prototypical
inducers (Mills et al., 2004). In addition, expression of
PXR and AhR, which are important transcription factors
Organotypic liver culture models 521
© 2012 Informa Healthcare USA, Inc.
involved in the regulation of drug-metabolizing enzymes
and transporters, have been detected in Fa2N-4 cells
and were shown to be at the same levels as primary
human hepatocytes (Mills et al., 2004). In this regard,
Fa2N-4 cells provide an acceptable model system to
predict the potential of compounds to be CYP1A and
CYP3A inducers (Ripp et al., 2006). However, both CAR
and several hepatic uptake transporters including the
OATPs are deficient in Fa2N-4 cells relative to primary
hepatocytes (Hariparsad et al., 2008), and ultimately, as
with HepG2, they are not a very representative system
for recapitulating hepatocyte function and response to
HepaRG is a cell line derived from a hepatocellular car-
cinoma (Guillouzo et al., 2007). When confluent, mono-
layers of HepaRG cells consist of two distinct cell types.
One type is flattened and morphologically resembles
cholangiocyte-like cells that retain a clear cytoplasm. The
second cell type shares similar morphological character-
istics with primary human hepatocytes. Both cell types are
equally represented within the cell population at conflu-
ency. In order to obtain liver-like function, HepaRG cells
must be differentiated into hepatocyte-like morphology
by treatment with dimethyl sulfoxide (DMSO) (Guillouzo
et al., 2007). Under this treatment, HepaRG cells become
quiescent and exhibit more hepatocyte-like functions.
For example, differentiated HepaRG cells express various
cytochrome P450 enzymes, such as CYP1A2, CYP2B6,
CYP2C9, CYP2E1, and CYP3A4, at substantially higher
levels than other cell lines described in previous sec-
tions. They also exhibit functional phase 2 conjugation
pathways and contain many membrane transporters
normally found in primary hepatocytes (Guillouzo et
al., 2007; Kanebratt and Andersson, 2008a; Turpeinen et
al., 2009). In addition, HepaRG cells express functional
receptor pathways involved in xenobiotic metabolism
and clearance, including CAR, PXR, and AhR. This
improved suite of ligand-activated nuclear receptors
results in more hepatic-like or in vivo-like induction of
CYP1A1, CYP1A2, CYP2B6, CYP2C8, CYP2C9, CYP2C19
and CYP3A4 in HepaRG compared to most other hepatic
cell lines (Kanebratt and Andersson, 2008b; Kanebratt
and Andersson, 2008a; Turpeinen et al., 2009).
Altogether, HepaRG cells exhibit an adult hepatocyte-
like phenotype, more so than any other hepatic cell line
currently available. However, HepaRG cells are not with-
out limitations or caveats. The presence of high concen-
trations of DMSO (1%) is essential for cell differentiation
and optimal expression of metabolic enzymes and, in its
absence, CYP activities decrease markedly. High DMSO
exposure artificially supports high CYP gene expression
resulting in the activation of receptor pathways involved
in the regulation of phase 1 and 2 biotransformation
enzymes (e.g. CAR and PXR) (LeCluyse, 2001; LeCluyse
et al., 2000). Under these conditions, CYP3A4 expres-
sion in HepaRG cells is unable to respond to prototypical
inducers, such as PB and RIF. Therefore, cell culture con-
ditions must be adequately modified to accurately model
a normal hepatocyte response. Nevertheless, HepaRG
represents the most promising surrogate to primary
human hepatocytes and has served as a valuable tool for
conducting some preclinical development studies that
require extended treatment periods and consistent per-
formance (Kanebratt and Andersson, 2008b; Kanebratt
and Andersson, 2008a).
3.2 Stem cells
Because of the challenges associated with the procure-
ment of primary hepatocytes and the limitations in
functionality of cell lines, the use of stem-cell derived
hepatocytes is a seemingly attractive alternative. Stem
cells are capable of extensive self-renewal through cell
division, while maintaining the capacity to differentiate
into tissue- or organ-specific cells. For the purposes of
this review, we classify the starting cell populations as
pluripotent stem cells and adult stem cells. A thorough
assessment of differentiation strategies and outcomes
for these cells is provided by Snykers and colleagues
(Snykers et al., 2009).
Pluripotent stem cells (PSC) include embryonic stem
cells (ESC) and induced pluripotent stem cells (iPSC).
Both ESC and iPSC are capable of nearly limitless self-
renewal and retain the ability to differentiate into each
of the three germ layers (endoderm, mesoderm, and
ectoderm). In theory these cells can differentiate to any
cell type. Protocols have been fairly well established for
the differentiation of PSC towards hepatocytes (Shiraki et
al., 2008; Tsutsui et al., 2006; Ogawa et al., 2005; Sullivan
et al., 2010; Agarwal et al., 2008). These commonly focus
on initiating the endodermal differentiation process
through application of activin A, followed by the inclu-
sion of fibroblast growth factors and Wnt3a to facilitate
differentiation towards hepatic lineages. Subsequent
culture in medium containing traditional hepatocyte
culture supplements (e.g. insulin, dexamethasone, hepa-
tocyte growth factor) in addition to oncostatin M facili-
tate functional maturation into hepatocyte-like cells over
a 1–2 week period.
Adult stem cells can both self-renew and differenti-
ate to form some or all of the cell types in specific tis-
sues or organs and are generally considered to assist in
repair and regeneration of the tissue or organ in which
they reside. From this perspective, oval cells, or hepatic
progenitor cells, are significant for their putative role in
repopulating liver epithelium (Gaudio et al., 2009) (see
also section “Hepatic progenitor cells”). These resident
stem cells of the liver are bi-potent, and have the ability to
differentiate down biliary or hepatic lineages (Figure 6)
(Thorgeirsson, 1996; Vessey and de la Hall, 2001; Forbes
et al., 2002). Hepatic differentiation is facilitated by treat-
ment with “cocktails” that commonly include HGF, EGF,
oncostatin M and fibroblast growth factor (FGF) (Snykers
et al., 2009), and by increased cellular confluency (Strick-
Marchand and Weiss, 2002; Yamasaki et al., 2006). There
522 E. L. LeCluyse et al.
Critical Reviews in Toxicology
are several examples of extrahepatic stem cells being
able to differentiate into hepatocyte-like cells, such as the
multipotent adult progenitor cells (MAPC) or mesenchy-
mal stem cells (MSC) from bone marrow (Ong et al., 2006;
Lee et al., 2004; Schwartz et al., 2002; Snykers et al., 2006)
and adipose tissue (Banas et al., 2007). Differentiation
protocols for these cells are similar to those for oval cells.
A potential advantage of cells that are readily obtained
from adult sources, particularly including iPSC and
adipose derived mesenchymal stem cells, is the ability
to create donor panels that represent key polymorphic
variants within a target population. Because these cells
can be easily obtained from pre-selected donors, the
possibility of creating such a panel is very likely in the
foreseeable future. Such panels are difficult to achieve
with primary cells due to obvious limitations on selection
of source material.
In spite of advantages in sourcing and expansion of
these cells, significant barriers still exist to their imple-
mentation as reliable surrogates for primary hepatocytes.
One such barrier is around the persistent fetal phenotype
that many of these cells exhibit (Snykers et al., 2009;
Guguen-Guillouzo et al., 2010), although current iPSC-
based approaches appear to minimize this effect (Hay et
al., 2008; Sullivan et al., 2010). From the perspective of
toxicity testing, it is important that the cells being used as
“hepatocytes” express phase 1 and 2 enzymatic activities,
as well as uptake and efflux transporter activity. To date,
all stem cell based approaches exhibit suboptimal phase
1 activity, with no clear information on phase 2 or trans-
porter activity (Guguen-Guillouzo et al., 2010). Reported
deficiencies in phase 1 activity in stem cell populations
may be due to heterogeneity of the “differentiated” popu-
lation, as purified populations maintain CYP3A4 activity
at levels similar to those in primary human cells (Basma
et al., 2009).
3.2 Maintenance of histotypic and phenotypic
Ideally, advanced models of the liver should possess tis-
sues or cells that retain most, if not all, of the characteris-
tic biochemical machinery and molecular pathways that
allow for a normal phenotype to be expressed in vitro.
This requirement will most likely be accomplished with
primary cells that have not been altered in any significant
way, along with a suitable composition and configura-
tion of biomatrix and medium formulation while under
dynamic flow. Any evaluation and validation of surrogate
hepatic culture systems must incorporate proper charac-
terization measures to confirm the presence and func-
tionality of these biochemical and molecular processes
prior to determining the suitability of a new culture
model for specific types of toxicity testing.
Much is known about the cellular and molecular fac-
tors that dictate and regulate the overall architecture and
phenotype of hepatocytes and other epithelial cells (see
section “Hepatocyte cytoarchitecture and cell polarity”).
Elucidation of the optimal conditions for the long-term
cultivation of rat hepatocytes in standard 2-D static cul-
ture systems has also helped define requirements for key
components in the matrix and medium for expression of
the differentiated phenotype. A number of tissue-specific
functions, for instance, can be directly regulated by the
conditions in the culture environment, including cell–cell
contact and communication (gap and tight junctions),
various signal transduction pathways, the distribution of
surface receptors and adhesion molecules (e.g. integrins,
cadherins), the organization of cytoskeletal elements
(microfilaments, microtubules, intermediate filaments)
and the localization of cellular organelles (e.g. Golgi, ER,
nucleus). Important lessons were learned from work in
2-D, spheroid and periperfusion models. First, hepatocytes
typically express a more cuboidal shape and three-dimen-
sional architecture in those systems that are more sup-
portive of normal liver structure and function (Khan et al.,
2007; Wolkoff and Novikoff, 2007). In contrast, hepatocytes
maintained under most traditional culture conditions
exhibit an abnormal flattened shape that is often associated
with loss of the adult phenotype than with either prolifera-
tive or reparative states (Bucher et al., 1990; Ichihara, 1991).
Second, culture systems supportive of the adult phenotype
facilitate the retention of cell shape and architecture by
enhancing cell–cell and cell–matrix interactions (Hamilton
et al., 2001; Oda et al., 2008).
However, one of the biggest hurdles that must be over-
come with developing in vitro systems is the loss of con-
stitutive CYP activity during the initial 24–48 h of culture
even though other components of the P450-dependent
monooxygenase system, NADPH-cytochrome P450
reductase and cytochrome b5, are relatively well main-
tained (Akrawi et al., 1993; Grant et al., 1985). Before
preservation of hepatic functions can be achieved suc-
cessfully in vitro, the cellular, biochemical and molecular
factors that are involved in their expression and regulation
in vivo must be understood. An appreciation for these in
vivo modulators of hepatic function will help distinguish
whether the phenotype exhibited in vitro is due to the
natural course of events that occurs in the absence of
exogenous or endogenous stimuli (e.g. growth hormone
regulation of rat CYP expression patterns), or whether
it is due to a failure of the culture system to sustain the
architecture and functionality of the cell(s) (e.g. cell–cell
and cell–matrix interactions) (Wang and LeCluyse, 2003;
Waxman and Holloway, 2009; Hamilton et al., 2001).
Overall, liver-specific gene expression and the
resultant phenotype depend on maintenance of
histotypic morphology; however, manifestation of the
differentiated state, in both its structural and functional
form, cannot be entirely interpreted as a function of cell
cytoarchitecture alone. Certainly, the type, density, and
biophysical state of distinct matrix components as well
as the constitution and relative proportions of individual
soluble factors, both paracrine and autocrine, combine
to form a complete picture of the local environment.
Many of the intermediate events that occur upon cell-
ligand binding (both soluble and insoluble in nature) that
Organotypic liver culture models 523
© 2012 Informa Healthcare USA, Inc.
eventually lead to regulation of gene transcription are
primarily mediated by cross-talk between the elements of
the various signal transduction pathways (MacDonald et
al., 2009; Wang et al., 2009; Niessen et al., 2011). Current
efforts in the development of organotypic hepatic model
systems also need to consider these key factors that
dictate the expression of liver-specific phenotype in vitro.
3.3 Zonal architecture and microenvironments
The anatomical architecture of the liver establishes
unique microenvironments of hemodynamics (portal
and arterial), nutrients, oxygen tension, hormones,
metabolites, matrix biology, and endogenous and exog-
enous substrates. These anatomical features lead to dif-
ferences in cell lineage, gene expression, metabolism
and transport function. As such, a simple static culture
system in a standard CO2 incubator at atmospheric oxy-
gen levels cannot truly represent the type(s) of microen-
vironments that a cell or compound encounters in vivo.
Also, the native configuration of the different cell types,
both in terms of diversity and three-dimensional nature,
is not typically represented in traditional 2-D culture
systems, which most often are monocultures of a single
cell type (usually hepatocytes) or, at best, co-cultures of
hepatocytes with KC or SEC.
From a practical and technical perspective, the zonal
architecture and microenvironments exhibited in mam-
malian liver represent difficult challenges to incorporate
simultaneously into a single device or culture platform.
Consequently, incorporating all these features into a
single system to determine a compound’s potential for
zone-specific hepatotoxicity cannot be accomplished at
this time. However, it may be possible to mimic a single
microenvironment to some degree in an individual
device or incubator by controlling oxygen tension, levels
of nutrients and other soluble components, composition
of cell types, and dynamic flow conditions. Perhaps, with
significant advances in microfluidics and microlithog-
raphy, more complete systems with integrated designs
of cellular architecture and precise control over zonal
environments on a microscale will be possible (Rhee et
al., 2005). For the foreseeable future, we anticipate that
comprehensive replicas of the whole organ in vitro will
remain a significant hurdle to overcome for engineers
and biologists alike.
3.4 Controlled flow dynamics
Recapitulating key aspects of local flow dynamics of
the liver will likely be a critical element of establishing
organotypic culture systems for at least three reasons.
First, most static culture methods do not allow for the
constant replenishment of medium and important nutri-
ents. This issue is inherently a source of concern and a
potential cause of organelle dysfunction and degradation
of cellular integrity over short periods of time given the
high rates of metabolism, oxygen consumption (respira-
tion), gene transcription and protein synthesis that occur
in normal, healthy liver. Second, most static culture
systems do not allow for the continual removal of spent
medium and toxic by-products of catabolic and xenobi-
otic metabolism, including metabolites of both endoge-
nous and exogenous substrates. Both the buildup of ROS
and a reduction in intracellular glutathione levels may
cause oxidative stress and lead to the rapid loss of cellular
function and viability over relatively short time periods
in vitro (Richert et al., 2002; Richert et al., 2006). Several
genes associated with a cellular oxidative stress response,
such as superoxide dismutase 2, glutathione reductase,
p53, and peroxiredoxin, become over-expressed after
plating of primary hepatocytes under static culture con-
ditions. Moreover, pathways leading to cellular apoptosis
are activated under standard culture conditions and are
due, at least in part, to oxidative stress-induced changes
(Ishihara et al., 2005). Third, most static culture models
only allow cells to be exposed to a constant concentra-
tion of a drug or chemical when testing for toxic proper-
ties. In vivo, cellular or tissue exposure to most chemicals
that are ingested orally or through other portals (e.g.
skin or lungs) involves dynamic processes, including
absorption, biotransformation and elimination, where
local concentrations of parent compound are changing
over time. Although many of the biochemical processes
expressed by the different cells in the liver may be present
in vitro, changes in chemical or drug concentration over
time are not well-represented under static conditions.
In addition to dynamic processes related to compound
delivery and exposure, another compelling reason for
incorporating dynamic flow is that local hemodynamic
forces will affect the biology and phenotype of endothelial
and epithelial cells (see section “Liver hemodynamics”).
Basic cell phenotype and response to xenobiotics and
other modulators are different in vascular endothelial
cells under flow versus static conditions (Hastings et al.,
2009). Expression of detoxification genes in primary cul-
tures of human hepatocytes maintained under flow reach
levels close to or higher than those measured in freshly
isolated hepatocytes (Vinci et al., 2011). Intuitively, the
more efficient removal of waste products and metabo-
lites under dynamic flow conditions would help prevent
their accumulation inside the cells that would otherwise
adversely affect cell health and integrity under static
conditions. Under dynamic flow conditions, cells should
be more tolerant of higher concentrations of direct hepa-
totoxins, assuming that they are able to metabolize and
clear compounds more effectively, and be more sensitive
to compounds that are bioactivated to reactive metabo-
lites that cause hepatotoxicity. This differential sensitivity
to cytotoxicity has been observed with acetaminophen
and troglitazone, as well as in studies conducted in vitro
comparing differences in culture models (Richert et al.,
2002; Novik et al., 2010).
In this regard, hepatic culture platforms that
incorporate microfluidics into the system should have
advantages over those that continue to rely on traditional
culture technologies or formats. Currently, microfluidic
524 E. L. LeCluyse et al.
Critical Reviews in Toxicology
culture devices are being developed that incorporate
computer-controlled valve and pump systems, which will
allow for the concentration of a compound in medium to
be continuously adjusted, thus reproducing any form of
toxicokinetic profile (i.e. AUC) for parent or metabolite
exposure observed in vivo (Balagadde et al., 2005; Wu
et al., 2010; Huang et al., 2011). Another advantage of
these microfluidic systems is that they can be coupled
with sensitive analytical instrumentation (e.g. LC-MS/
MS) to analyze changes in parent concentrations over
time as well as identify the rate of production of specific
Future generations of culture devices or platforms that
couple some type of flow control with the relevant biol-
ogy mentioned previously should better maintain cell
health and phenotype for prolonged periods and provide
a more accurate depiction of the hepatotoxic potential
of compounds. Theoretically, such systems would allow
toxicologists to control cellular and tissue concentra-
tions of drugs and xenobiotics over time to better mimic
dynamic exposure levels known to occur in vivo. These
modifications are challenging, but likely worth the effort,
if they lead to stable organotypic culture systems that dis-
play physiologically-relevant biology and better predic-
tion of xenobiotic hepatotoxicity in humans.
3.5 Defined cellularity
Chemical-induced hepatotoxicity often occurs in spe-
cific regions of the liver and is due, in part, to the natural
configuration and relationship of the different cell types
in the zonal microenvironments (DeLeve et al., 1997;
Przybocki et al., 1992; Roberts et al., 2007). Most standard
two-dimensional static culture models of the liver are
monocultures of immortalized or primary hepatocytes.
Co-culture of hepatocytes and NPC or immortalized cell
line adds to the biological complexity, but often in an
undefined or non-prescribed fashion. Likewise, bioreac-
tors often incorporate single or multiple cell types from
the liver or other tissue into a closed system that contains
some form of scaffold or undefined stromal layer as a
substratum. In the case where immortalized cell lines
or primary hepatocytes alone are incorporated into the
system, the biological complexity of the intact liver is
lacking. In the multicellular systems, often the biomass
inside the bioreactors is composed of an undefined ratio
of cell types that often doesn’t reflect that of the original
seed stock because of inherent differences in cell health
From the standpoint of sample collection for toxic-
ity testing and “-omic” analyses, the advantage of the
conventional monoculture static and bioreactor sys-
tems is that they contain a relatively pure population
of a single cell type (e.g. hepatocytes) and, therefore,
fewer complications regarding cross-contamination
with sample collection for proteomic, transcriptomic
or metabolomic analysis. These cultures also minimize
the confusion about the cellular source of a particular
biochemical response or the order in which biochemical
and molecular events occur in closed, multicellular
systems. On the other hand, they are by design simple
systems and do not represent the multicellular complex-
ity of the liver in response to compound exposure. The
advantage of the co-culture static and bioreactor sys-
tems is that they better reflect the biological complexity
and response to compound exposure in an organotypic
fashion. However, as mentioned above they often do not
have a controlled ratio or mass of the different cell types,
which can confound the accurate interpretation of com-
It would be ideal to have the means to define and
control the configuration and the proportions of the dif-
ferent cell types inside the culture system. In vivo, each
of the different cell types along the sinusoidal structure
co-exists in a prescribed architecture, geometry, quantity
and proportion relative to one another. The relationship
changes across zones of the liver lobule and under dif-
ferent disease states (Badr et al., 1986; Xanthopoulos
and Mirkovitch, 1993; Ishibashi et al., 2009; Kolios et al.,
2006). Assessing drug- or chemical-induced effects on
liver function or adaptive responses would be enhanced
by the ability to control the presence or absence of NPC
in order to determine their relative role for specific bio-
logical responses of the liver. An added benefit, if the
different cell types were accessible separately from one
another, would be the ability to collect or harvest specific
cell types for transcriptomic or proteomic analysis. Novel
approaches to micropatterning cell attachment factors
on the growth surface of culture platforms and config-
uring different cell types across permeable membranes
or on transwell inserts will help alleviate some of these
issues in the future (Hastings et al., 2009; Khetani and
Another advantage of the standard 2-D culture systems is
that they are generally accessible to perform a number of
imaging procedures, including standard phase contrast,
differential interference contrast (DIC), and fluorescence
microscopy, as well as high-content screening. The cells
are also easily accessible for harvest and preparation of
RNA, protein or subcellular fractions to perform either
‘-omic’ or biochemical analyses. However, the loss of
histotypic architecture and phenotypic gene expression
combined with the absence of other relevant cell types
and dynamic flow makes it a poor choice for certain
types of toxicity testing and identification of certain
modes of action. In contrast, most bioreactors, which
possess many beneficial features that are lacking in the
2-D model systems, are closed systems and/or difficult to
access for most standard microscopic, biochemical and
Ideally, a culture system would be designed in such a
way as to enable microscopic evaluations on intact living
cells/tissues inside the device. In addition, the systems
would be amenable to the collection of medium samples or
the harvest of cellular material for subsequent biochemical
Organotypic liver culture models 525
© 2012 Informa Healthcare USA, Inc.
and metabolic analyses during or after completing
experimental protocols. In the absence of the ability to
perform direct microscopic evaluation through a viewing
or camera port, a culture device should be amenable to the
fixation and removal of intact cells/tissue for histological
or immunostaining procedures. For some applications,
such as, metabolic stability, metabolite identification,
changes in serum proteins, and profiling changes in CYP
expression, the ability to interface the device(s) directly
with analytical tools would be a great advantage. Such
capabilities would also be more conducive to establishing
an integrated, automated robotic system for routine
bioanalysis of compounds and their metabolites. These
systems theoretically could be setup and run under
different incubation conditions to mimic the zonal
microenvironments established in the liver as well as
different disease or stressor conditions.
3.7 Throughput and cost-effectiveness
The past decade has seen the emergence of a multi-
tude of assays and facilities to support high throughput
screening (HTS) of macromolecules to assess the effi-
cacy of compounds on specific molecular and cellular
targets or their perturbation of biological pathways.
Recently, an emphasis was placed in an NRC report on
developing in vitro assays and tools that would be cost
effective and allow high-throughput assessment of a
large number of chemicals (National Research Council,
2007; Andersen and Krewski, 2010). The initial Toxcast
program was performed primarily with a large num-
ber of in vitro models that possessed both attributes
(Houck et al., 2009; Judson et al., 2010). With about
25% of compounds, regulatory decisions are based on
liver effects – hypertrophy, enzyme induction, cancer,
etc. – in rodent liver. New in vitro test systems for liver
would improve screening of hepatic responses to these
compounds and the understanding of the relationship
of hepatic responses to organism-level toxicity.
Unfortunately, the drive for greater efficiency and
cost-effectiveness of the chemical or drug screening
process has often overshadowed the need for screen-
ing tools that are more biologically relevant from a
complexity and contextual standpoint. The biological
context of the test system, whether protein, cell or tis-
sue based, may or may not directly apply to the in vivo
situation. For example, if a compound is bioactivated
by phase 1 or 2 enzymes in vivo to a reactive metabolite
before causing hepatotoxicity, then most HTS systems,
which typically are deficient in these metabolic capa-
bilities, will not be suitable for identifying these types
of toxic events. Likewise, if the MOA involves interac-
tive responses between resident macrophages (KC) and
parenchymal cells (HC), then simple monocultures
of immortalized hepatocytes will not reproduce these
effects under most HTS conditions. Obviously, in vitro
based toxicity testing needs to incorporate a balanced
approach of throughput and relevance (regardless of
the speed at which the data can be generated). In many
respects, a ‘gold-standard’ in vitro system that mim-
ics complex modes of action and cellular phenotypes
could aid in putting the results from more molecular-
and biochemical-based HTS screening tools in better
perspective regarding their physiological relevance. For
testing of hepatic responses, a range of in vitro methods
are likely to be required to achieve useful results: some
supporting higher throughput and others designed to
maintain better correspondence with in vivo biology.
4 working towards standardized methods
for evaluation and validation of advanced
One of the current challenges with determining the suit-
ability and relevance of the various types of advanced
hepatic culture systems for different types of toxicologi-
cal applications has been the lack of consensus on a vali-
dation paradigm and corresponding acceptance criteria.
Whenever adopting a new in vitro model system for
industry-wide screening or testing purposes, standard-
ized conditions and benchmarks are typically defined
and established as part of the evaluation and validation
process prior to its adoption for particular toxicological
or pharmacological applications (Lilienblum et al., 2008;
Schechtman, 2002; Kinsner-Ovaskainen et al., 2009b;
Kinsner-Ovaskainen et al., 2009a). As such, definitions
and criteria for relevant biological and toxicological
parameters and quality control measures need to be
put into place for at least four key areas: (1) overall cell
health and integrity, (2) cell type specific morphological
and architectural integrity, (3) phase 1 and 2 xenobiotic
metabolic capacity and 4) key response pathways that are
mechanistically relevant. Related to these topics, stan-
dards are also needed to qualify suitable cell culture con-
ditions, define minimum acceptable levels of basic cell
functions, such as albumin production or basal phase 1
and 2 enzyme activity, and relevant concentration ranges
for important intermediaries of chemical-induced stress,
such as ATP and GSH. In addition, these measures need
to be benchmarked against those observed or obtained
from in vivo observations or experimentation, where
available, or from freshly isolated tissues and cells.
To facilitate the evaluation of new or existing toxicity
models, programs such as the Interagency Coordinating
Committee on the Validation of Alternative Methods
(ICCVAM) and European Centre for the Validation of
Alternative Methods (ECVAM) have developed pro-
cesses and criteria for validating new test systems and
methods (Schechtman, 2002; Lilienblum et al., 2008;
Kinsner-Ovaskainen et al., 2009a). It is not the intention
of this article to reproduce or review those strategies here
or prescribe an all-inclusive list of conditions and func-
tions that can or should be assessed prior to choosing the
most appropriate in vitro system for specific toxicological
applications. However, the following section is intended
to provide guidance and recommendations regarding
minimal validation methods and acceptance criteria for
526 E. L. LeCluyse et al.
Critical Reviews in Toxicology
providing assurances that a particular hepatic culture
system is likely to generate relevant and valid results.
An initial assessment of the global phenotype of
the cells in a novel culture device could, and possibly
should, be assessed by thorough transcriptomic and
proteomic analysis to give some assurances that the
relevant cell types are performing at or near the levels of
their counterparts in vivo. Admittedly, it is impractical
to perform these types of analyses on a routine basis.
However, a subset of basic hepatic functions and
biochemical pathways should be assessed routinely and
the results compared to a ‘gold-standard’ , which generally
equates to those reported from in vivo experiments
(when available) or from freshly isolated cells or tissues.
In some cases, benchmark levels and standards for
specific endpoints or changes in gene profiles in response
to prototype hepatotoxins are published in the literature
(e.g. (LeCluyse et al., 1999; Richert et al., 2002; Binda et
al., 2003; Pearce et al., 1996; Donato et al., 1993; Dunn
et al., 1991; Guguen-Guillouzo et al., 1983). In general,
system validation efforts should focus on specific cellular
functions and biochemical pathways, particularly for
hepatocytes, that are most relevant for mimicking
responses to hepatotoxic compounds. It can often be the
case where a particular in vitro system cannot address
or mimic a specific type of toxicity due to some inherent
limitation such as a deficiency or absence of a particular
cell type, biochemical pathway or nuclear transcription
factor related to a compound’s MOA. For example, a
monoculture of hepatocytes cannot be utilized to explore
the role of endotoxins in exacerbating chemical toxicity.
Neither can an immortalized cell line, such as HepG2,
be employed to examine the bioactivation of aflatoxin
B1 or induction of liver enzymes by PB. In either case,
the system is lacking in some key component that is
necessary, such as another relevant cell type (e.g. Kupffer
cells), CYP enzymatic activity, or nuclear receptor
expression (e.g. CAR).
4.1 Assessing cell and tissue integrity
The initial quality and integrity of primary and immor-
talized liver cells should be assessed prior to and dur-
ing their use for experimental purposes. Obviously, the
condition of the cells and the quality of the subsequent
cultures prior to treatment with compounds are impor-
tant when it comes to determining their suitability and
capability to respond in a normal fashion to a hepato-
toxic compound. Also, the metabolic capacity of the cells,
including phase 1 and 2 metabolizing enzyme function,
depends on the integrity of the cells and their ability to
generate key cofactors involved in the detoxication and
elimination of xenobiotics (e.g. NADPH, UDPGA, PAPS,
GSH etc.), as well as the production of reactive metabo-
lites. As such, assessment of the viability and integrity of
the cellular components should be an important part of
evaluating and validating the stability and robustness of
any hepatic culture model system. Moreover, the valida-
tion regimen ought to be performed at regular intervals
(e.g. daily initially, then weekly if warranted) over a
relevant period of time that mimics that of a standard
Assessment of the integrity and biological fidelity of
cells within a culture device should be conducted on
multiple levels and can be performed by either invasive
or non-invasive means. The simplest method of testing
cell viability or functional integrity is to use probe sub-
strates for metabolic or respiratory pathways that require
living cells in order to observe substrate turnover. Glucose
utilization, urea synthesis, albumin secretion, mitochon-
drial function and enzyme activity are all examples of
simple endpoint measurements that can be performed
by adding commercially-available probe substrates and
collecting media samples to determine the health and
robustness of the cells without resorting to sacrificing
a device or batch of cells. More complex responses to
modulators of liver functions, such as enzyme inducers
or bacterial endotoxins, can be probed indirectly by pro-
filing changes in substrate turnover or cytokine patterns
over time in the medium. However, definitive determi-
nations of cell numbers, cell types and basic function-
ality must be based on total protein or cell content and
can only be provided by sacrificing devices to measure
those parameters or to make histological observations
directly in extracted cellular material. Endpoints such as
those listed above should be compared or benchmarked
against those obtained from freshly isolated hepatocytes,
short-term cultures or whole liver tissue from the same
donor whenever possible (LeCluyse et al., 1999; Richert
et al., 2002; Binda et al., 2003; Pearce et al., 1996; Donato
et al., 1993; Dunn et al., 1991). Once the relationship
between corresponding functional activities of the initial
cell stocks and subsequent cell culture layers has been
established then surrogate markers and endpoints can
substitute for the more invasive measures in subsequent
To assess the morphological identity and integrity of
cells within a complex device and their biological fidel-
ity with the tissue or cells of origin, invasive means must
be utilized initially until the reproducibility of the culture
conditions can be confirmed. In some cases, if the culture
device is amenable to light and fluorescence microscopy
then adequate assessment of cell morphology and integ-
rity may be performed directly. However, if the culture
device involves the maintenance of cells as three-dimen-
sional aggregates or on scaffolds then the tissue will likely
require prefixation by standard procedures (e.g. buffered
formalin) followed by routine processing for histochemi-
cal staining and histological assessment. Likewise, some
cellular features, such as microvilli, intercellular connec-
tions (e.g. junctional complexes), phagocytosis by KC,
and fenestrations on the surface of sinusoidal endothelial
cells, can only be viewed using specialized microscopic
techniques (LeCluyse et al., 1994; LeCluyse et al., 1999;
Braet and Wisse, 2002; Cogger et al., 2010). Comparisons
in cell morphology, tissue architecture, antigenic mark-
ers and cell ratios should be benchmarked against the
Organotypic liver culture models 527
© 2012 Informa Healthcare USA, Inc.
relevant cell type(s) of the liver in vivo (Table 1) (see also
section “Major cell types of the liver”).
A benefit of organotypic culture systems with a
relative longevity of several weeks, if not months, is
that the basal levels of gene expression and other facets
of cellular phenotype have time to stabilize or reach
a definable steady-state level for specific metabolic
or intrinsic functions prior to initiating experimental
protocols. These could more readily be compared to the
corresponding levels in the intact liver or freshly isolated
cells. Longer-term organotypic model systems also allow
for the restoration and stabilization of cellular and tissue
architecture, such as cell polarity, cell–matrix and cell–
cell interactions, as well as other important subcellular
elements, such as mitochondria, Golgi (Hamilton et al.,
2001; Desai et al., 2009; Niessen et al., 2011), stabilization
of rates of protein synthesis (Bayad et al., 1991; Strey et
al., 2010), and intracellular glutathione and ATP levels
(Richert et al., 2002; Tuschl et al., 2009). In the future,
data generated from studies utilizing in vitro organotypic
model systems should be judged or scrutinized in
light of a system’s ability to maintain or exhibit certain
biochemical properties at physiologic levels, not just as
the presence or absence of key functions or components,
which so often occurs today in published reports.
During the initial time period of cell integration into
complex culture devices and their adaptation to the cul-
tivation conditions (often requiring days, if not weeks, to
stabilize), there is an opportunity to assess the recovery
and suitability of cells for particular applications. In
addition, a pre-defined incubation period would allow
baseline conditions and other experimental variables
to be defined and standardized for specific toxicological
applications and assays prior to initiating studies. This
also provides the opportunity to establish baseline values
for basic hepatic functions and enzymatic activities, such
as specific phase 1 and 2 biotransformation reactions,
which will in turn enable easier and more accurate inter-
pretation of results and comparisons to in vivo outcomes.
4.2 Standardizing culture conditions
One of the major obstacles to evaluating and validating
the suitability of in vitro hepatocyte culture systems for
pharmaceutical and toxicological applications has been
the lack of standardized culture conditions and experi-
mental methods for the proper maintenance of hepato-
cytes prior to and during in vitro testing of compounds.
Unfortunately, there have been a wide array of culture
conditions employed in the laboratory (e.g. supple-
ments, media formulations, ECM, 3-D scaffolds etc.) but
few reports with comprehensive and systematic compar-
isons of pharmacologically and toxicologically relevant
endpoints. Published studies often claim a particular
combination of culture conditions maintains adult hepa-
tocytes without loss of differentiation or, alternatively,
with “full” expression of normal liver-specific functions,
when in fact only a small number of applicable endpoints
typically have been examined (e.g. albumin production,
urea synthesis, or singular CYP activities). Moreover, the
measured endpoint(s) may or may not indicate suitabil-
ity of the system for a specific type or types of toxicologi-
An important area for standardization is choice of
media formulation and supplementary additives, includ-
ing hormones, cofactors and antioxidants. Many media
formulations have been employed for the cultivation of
primary and immortalized hepatocytes over the years;
however, few comprehensive studies comparing the
effects of formulation on the maintenance of liver-spe-
cific functions as they relate to toxicity testing have been
performed. In vivo, hepatocytes and other liver cells are
continually exposed to a variety of hormones and other
soluble factors which, alone and in combination, pro-
foundly affect cell function and growth in an additive,
synergistic or antagonistic manner (Rodés et al., 2007;
Gaudio et al., 2009; Turner et al., 2011). Complex nutri-
tional and hormonal influences help govern the normal
activities and responses of hepatocytes in vivo, including
species-specific metabolic capacity (Zaphiropoulos et al.,
1990a; Zaphiropoulos et al., 1990b; Bullock et al., 1991).
Consequently, when hepatocytes and other liver cells are
placed into culture, there typically is a considerable shift
in the factors that regulate hormone-dependent genes
Determining the critical components in media respon-
sible for enhancing hepatocyte survival and function has
been the emphasis of numerous efforts to culture adult
hepatocytes long term (Guguen-Guillouzo et al., 1983;
Dich and Grunnet, 1990; Berry et al., 1991; LeCluyse
et al., 1996a). Overall, it is clear that more enriched media
formulations support basic hepatocyte functions and the
maintenance of metabolic enzymes to a greater extent
than basal medium formulations (Sidhu et al., 1994;
Yan et al., 1995; Liu et al., 1996; Zangar et al., 1995). The
formulation of the medium in conjunction with other
appropriate culture conditions, such as ECM composi-
tion, growth factor and hormone levels, affects the estab-
lishment and maintenance of histotypic cell morphology
and cytoarchitecture of cultured hepatocytes (Sidhu
et al., 1994; Arterburn et al., 1995; Hamilton et al., 2001).
Nearly all attempts at developing or identifying optimal
media formulations for the maintenance of primary and
immortalized liver cells in vitro have been performed
under static culture conditions. The question remains as
to whether or not these optimized conditions would nec-
essarily be optimal for more organotypic cultures of liver
cells, especially under dynamic flow conditions.
4.3 Assessing metabolic capacity of in vitro systems
Another area in need of standardization is the assessment of
the metabolic functions of the liver tissue or cells in relation
to in vivo values. Frequently, the stated goal of an in vitro
model development project is to maintain both phase 1
and 2 biotransformation enzyme reactions at or near those
levels exhibited in vivo. However, this assessment can be
a daunting task for most laboratories and often requires
528 E. L. LeCluyse et al.
Critical Reviews in Toxicology
extensive time and analytical commitment. First, the choice
of probe substrates is critical due to the variety of phase
1 and 2 enzymes and the marked differences in substrate
specificity among species and individual subfamilies of
CYP enzymes (Gemzik et al., 1992; Pearce et al., 1992;
Waxman and Holloway, 2009). Second, there are significant
interindividual and species differences in the expression
and regulation of the metabolizing enzymes as well as the
transporter proteins. For example, CYP1A1/2, CYP2E1 and
CYP4A enzymes have similar substrate specificities and
catalytic rates across species; however, regulation of CYP1A
and CYP4A enzymes through activation of the nuclear
receptors AhR and PPARα, respectively, can exhibit marked
species differences (LeCluyse and Rowlands, 2007). On
the other hand, CYP2B, CYP3A and CYP4A isoforms show
some substrate similarities within subfamilies across
species, while exhibiting substrate diversity in other cases,
implying that the proper choice of substrates for assessing
the presence and functionality of these enzymes is crucial
to properly assess the relevant levels in an in vitro system
(Langsch et al., 2009). Members of the CYP2C enzyme
subfamily exhibit the greatest species differences in
expression, substrate specificity, and regulation compared
to other CYP subfamilies involved in the metabolism and
bioactivation of xenobiotics (Mugford and Kedderis, 1998;
Tsao et al., 2001; Waxman and Holloway, 2009).
Another problematic issue when trying to compare
results from in vitro model systems or between labora-
tories using similar culture systems is that a number of
media constituents, including several that have been used
routinely as supplements or solvents, are inducers of CYP
enzymes (e.g. tryptophan, ethanol, DMSO, metyrapone)
(LeCluyse et al., 1996a). Likewise, hormonal supplements,
such as dexamethasone and insulin, regulate the expres-
sion and/or activity of a number of enzymes and transport
proteins involved in xenobiotic disposition (Gebhardt
et al., 2003; Woodcroft et al., 2002; Abdelmegeed et al.,
2005). Therefore, many of the media components used to
enhance the survival of cultured hepatocytes, especially
those that seemingly “maintain” total cytochrome P450
content, must also be considered in light of their capac-
ity to modulate individual P450 enzymes in an ‘artificial’
manner, especially at nonphysiologic levels.
As such, straightforward and relatively simple meth-
ods are required to evaluate and validate the metabolic
and transport capacity of a particular hepatic model
system. Minimally, a selective subset of isoform-specific
probe substrates that evaluate several of the phase 1 and
2 enzyme activities of the cell types would be a valuable
asset to instill some confidence that important basic path-
ways are restored and maintained over time prior to chal-
lenges with a test agent. A few probe substrates do exist
and have been historically utilized to probe the metabolic
capacity of in vitro cultures of both primary and immor-
talized hepatocytes (Gebhardt et al., 2003). One of the
most commonly used probe substrates is 7-ethoxycou-
marin, which is metabolized to 7-hydroxycoumarin by
multiple cytochrome P450 enzymes and then conjugated
by sulfotransferase and UDP-glucuronosyltransferase
enzymes (Edwards et al., 1984). This approach allows one
to non-invasively determine several basic phase 1 and 2
enzyme functions of the cell culture system with a single
probe substrate. However, these assays require HPLC or
LC-MS analytical methodologies.
Alternatively, one can use fluorescent- or biolumines-
cent-based probe substrates, such as 7-alkyoxyresorufins
(CYP1A and CYP2B probes) and Luciferin-IPA (CYP3A)
from commercial sources (Sakai et al., 2010a; Donato et
al., 1993; Doshi and Li, 2011). These substrates can be
added directly to media or buffers prior to exposure of
cell cultures. The rate of metabolism, as represented by
increased fluorescence or luminescence, can be deter-
mined over time in the presence or absence of a test
article or prototypical inducer of CYP enzymes (e.g. PB,
3MC). The disadvantages or limitations of these probe
substrates is generally their lack of specificity from spe-
cies-to-species, low-turnover rates when utilized to mea-
sure metabolic activity in cell cultures, and quenching or
interference by cellular or medium components (Donato
et al., 1991; Donato et al., 1993). Clearly, a robust, stan-
dardized set of probe substrates that measure relevant
levels of CYP and phase 2 metabolic capacity in dynamic
culture systems is needed. Validated probe substrates to
measure the functional activities of major human CYP
isoforms, as well as some phase 2 conjugation pathways,
in primary hepatocytes with corresponding recom-
mendations for relevant concentrations, specific activity
levels and analytical methods can be found in a number
of related publications (Li et al., 1999; Zhang et al., 2009;
Huang et al., 2007; Smith et al., 2012).
4.4 Normalization of in vitro data across culture
A significant challenge facing scientists using advanced
organotypic model systems, especially those incorporat-
ing 3-D scaffolds and/or flow chambers, is normalizing
the data to compare on equivalent terms to those gener-
ated in traditional 2-D systems or in vivo. For conventional
2-D culture systems, this comparison is done by normal-
izing rates or quantities on the basis of the total amount
of protein (i.e. per milligram protein) or total number of
cells (i.e. per million cells) (e.g. Li et al., 1999). However,
tissues or cells in bioreactors and other closed systems
are not easily accessible or they are a mixture of cell types
in undefined proportions. To compare such data from
complex culture systems with those of traditional culture
models researchers have resorted to ‘sacrificing’ a certain
number of devices or making certain assumptions about
the amount of cellular material inside devices in order to
make calculations about particular amounts of protein,
RNA/DNA or rates of reactions (Wang et al., 2010a; Novik
et al., 2010; Dash et al., 2009). Unfortunately, these prac-
tices may need to continue pending the availability of
alternative non-invasive approaches.
To properly evaluate and validate novel organotypic
liver model culture systems, especially those that
Organotypic liver culture models 529
© 2012 Informa Healthcare USA, Inc.
involve closed systems or inaccessible tissues, our
recommendation is to design initial studies to include the
sacrifice of a certain number of devices and/or samples
in order to examine their cellular, protein, DNA and RNA
content, depending on the type and the accessible nature
of the device. This validation scheme should include a
morphological and histological characterization of the
types and configurations of the different cell types within
the device as well as their histological features (e.g. bile
canaliculi of hepatocytes, fenestrations of sinusoidal
endothelial cells and processes of the stellate cells) (see
also section “Major cell types of the liver”) (Rodés et al.,
2007). Likewise, the consistency, reproducibility and
robustness of the expression levels of marker proteins
and mRNAs should be examined. For each cell type, the
amounts and ratios of specific markers for each cell type
ought to be determined and followed over time to confirm
their presence and levels of expression. Examples of
specific functional or histotypic markers that could be
utilized for this purpose are listed in Table 1.
Alternatively, the ability to assess metabolism by
examining effluent compounds from the culture systems
could be coupled with other metabolic analysis to evalu-
ate fidelity between the in vivo and in vitro pathways.
A well-designed liver bioreactor could function in a
manner similar to isolated-perfused liver preparations
(Bessems et al., 2006). Analysis of metabolites produced
in a bioreactor might also serve to benchmark expected
metabolic pathways. Evaluation of the fidelity of the bio-
reactor and new organotypic systems could be verified
by assessing metabolite profiles with prototype test com-
pounds, i.e. those whose metabolism has already been
well-documented in vivo or through other systematic
approaches (Choi et al., 2012).
5 advanced organotypic culture technologies
There has been a recent surge in the creation of organo-
typic culture devices, especially for maintenance and
growth of liver primary and immortalized cells for toxi-
cological research (Sivaraman et al., 2005; Khetani and
Bhatia, 2008; Chao et al., 2009; Domansky et al., 2010;
Baker, 2011; van Midwoud et al., 2011). Here we summa-
rize a few relevant examples of innovative or improved
in vitro hepatic culture systems intended to support the
long-term maintenance of cell viability, morphology and
functionality. Each of these systems is either commer-
cially available or destined for commercial release. These
technologies are not meant to be all-inclusive as the
level of investment and effort in developing more predic-
tive in vitro culture models has increased immensely in
the past several years (Baker, 2011; van Midwoud et al.,
2011). However, our intention is to introduce some of
the more advanced technologies that (1) have shown
improved functionality,( 2) are reasonably mature in
their development, and (3) have undergone some valida-
tion for toxicological applications. All of these systems
have incorporated one or more of the features that are
typically missing from conventional 2-D static culture
models of the liver and attempt to address the limita-
tions of most conventional in vitro 2-D model systems.
The different model systems described in this section are
compared in Table 2 relative to phenotypic and practical
5.1 Microfluidic perfusion array
The Perfusion Array Liver system (PEARL) was designed
for automated long-term culture of primary hepatocytes
and other cell types (Figure 8A). The system is built on a
Table 2. Comparison of organotypic models of the liver with respect to structural, functional and practical considerations.
Microscale 3-D livere
aPerfusion Array Liver System (PEARL) developed by CellASIC (see section “Microfluidic perfusion array”) (Lee et al., 2007).
bMicropatterned co-culture system developed by Hepregen (see section “Bioengineered micro-patterned liver platform”) (Khetani and
cBiochip dynamic flow system developed by HμREL® (see section “Biochip dynamic flow system” ) (Chao et al., 2009).
d3-D tissue co-culture platform developed by RegeneMed (see section “3-D liver tissue culture scaffold”) (Naughton et al., 1994).
eCombination fluid flow and 3-D cell culture system developed by Griffith and colleagues (see section “3-D scaffolds with dynamic flow”)
(Powers et al., 2002a,b; Sivaraman et al., 2005).
Basic liver Defined
Table 1. Functional and histotypic markers of the different cell
types that compose the liver microstructure.
Hepatocyte Albumin, CYP3A,
Bile canaliculi Khan et al.
DeLeve et al.
et al. (1993)
Hendriks et al.
(2001), Sato et al.
Roberts et al.
530 E. L. LeCluyse et al.
Critical Reviews in Toxicology
standard 96-well plate format with 32 independent flow
units per plate, each housing 30,000 cells exposed to
continuous perfusion of 100 µl per day (Lee et al., 2007).
Within each unit, the cells are loaded into a set of micro-
fluidic structures designed to mimic the liver acinus, with
16 parallel 60 × 60 × 3,000 µm cords separated from a set of
flow sinusoid channels by an artificial porous endothelial-
like partition. The microfabricated porous barrier retains
the cells in high density 3-D aggregates while maximizing
nutrient and gas transport via diffusion through 2 µm pores.
The flow is gravity driven, eliminating the need for external
connections or pumps, making the operation compatible
with existing automation equipment and assay types. The
bottom surface of the plate is a 170 µm thick glass slide,
enabling high magnification microscopy of cultured cells.
Hepatocytes cultured in the microfluidic array
retain liver functions for 3–4 weeks. With both primary
and cryopreserved (human and rat) hepatocytes, cell
viability, 3-D morphology, CYP metabolic activity and
induction/inhibition potential, gene expression, albumin
production, and drug metabolism are maintained
over 28 days and found superior to sandwich-cultured
hepatocytes. Under these conditions, the hepatocytes
are likely responding to both cell-cell contact created by
the 3-D configuration and to the continuous perfusion
mass transport environment. Experiments to date have
focused predominantly on hepatocyte monocultures,
although the format of the system may be amenable to
addition of NPC along the fluidic channels. Additional
experiments regarding drug metabolism, transporter
expression, model toxicity mechanisms, and co-culture
are currently underway to further characterize the
microfluidic culture system.
5.2 Bioengineered micro-patterned liver platform
Khetani and Bhatia (Khetani and Bhatia, 2008) first
reported a miniaturized, multiwall culture system for
human liver cells with optimized microscale architec-
ture that are functional for several weeks. This approach
utilizes microtechnologies adapted from the semicon-
ductor industry to both optimize and miniaturize an
in vitro model of the liver in a multiwell format, called
Figure 8. Hepatic cell culture model systems represented in “Advanced organotypic culture technologies” . (A) Perfusion array liver system
(PEARL) (Lee et al., 2007), (B) bioengineered micropatterned liver platform (Khetani and Bhatia, 2008), (C) biochip dynamic flow system
(Chao et al., 2009; Novik et al., 2010; Maguire et al., 2009), (D) 3-D liver tissue culture scaffold (Sibanda et al., 1993; Sibanda et al., 1994;
Naughton et al., 1994), and (E) 3-D scaffolds with dynamic flow (Sivaraman et al., 2005; Domansky et al., 2010).
Organotypic liver culture models 531
© 2012 Informa Healthcare USA, Inc.
HepatoPac. Specifically, primary hepatocytes are orga-
nized into colonies of prescribed, empirically-optimized
dimensions using microfabrication tools and subse-
quently surrounded by supportive stromal cells (mouse
3T3-J2 cells) (Figure 8B). While these cells are not of
liver origin, they appear to provide many of the benefits
typically associated with co-culture of parenchymal and
nonparenchymal cells. Hepatocytes in the HepatoPac
platform retain their in vivo-like morphology, express
liver genes at high levels, metabolize compounds using
active phase 1 and 2 drug-metabolizing enzymes, secrete
diverse liver-specific products, and display functional
bile canaliculi for 4–6 weeks in vitro. The system exhibits
greater longevity and stability of liver-specific functions
relative to conventional culture models (i.e. collagen gel
sandwich, matrigel overlay) (Khetani and Bhatia, 2008;
Wang et al., 2010a).
The sensitivity and utility of the micropatterned
co-culture system can be further enhanced by (1)
longer-term dosing regimens (i.e. 2–4 weeks) for drugs
that are slowly turned over and produce secondary
metabolites; (2) use of more sensitive, high-content and
mechanistic endpoints; and (3) incorporation of liver-
derived NPC (e.g. KC) into the stromal compartment
surrounding micropatterned hepatocytes in order to
sensitize hepatocytes to drug-induced toxicity as occurs
in vivo. In addition, the longevity of the cultures and the
stability of the enzymatic and transport functions of the
primary hepatocytes in the HepatoPac platform allows
complete metabolism of compounds to occur resulting
in identification of clinically-relevant liver metabolites
missed in traditional culture systems. Indeed, long-term
incubation in HepatoPac have been shown to produce
75–80% of the clinically-relevant metabolite profile, as
opposed to less than 50% in traditional model systems,
including suspension hepatocytes, S9 and microsomal
fractions (Wang et al., 2010a).
The long-term viability (>3 weeks) and maintenance
of physiologically-relevant levels of drug-metabolizing
enzyme (DME) and transporter activity exhibited by this
system offer an attractive option for investigation of drug-
induced liver injury (DILI). In a recent study, the effects
of known human hepatotoxicants and non-hepatotoxi-
cants in micropatterned human hepatocyte co-cultures
using automated multispectral fluorescence imaging
technology to monitor perturbation of intracellular indi-
cators of hepatotoxicity. The microscale architecture was
optimized to facilitate efficient high-content imaging
(HCI) of hepatocytes without compromising longev-
ity or drug-metabolizing enzyme activity. Compounds
with well-established mechanisms of toxicity exhibited
expected changes in the targeted parameters (Khetani
et al., 2010; Kemper et al., 2011).
5.3 Biochip dynamic flow system
A biochip dynamic flow system on which resides one
or more separate, but microfluidically interconnected,
compartments has been developed to contain cultures
of living cells drawn from and/or representing different
organs or tissues of a living animal (Chao et al., 2009;
Novik et al., 2010; Maguire et al., 2009). The multi-cham-
ber device is designed on the basis of a PBPK model of a
220 g rat with the corresponding dimensions of the indi-
vidual chambers (W × L in mm): lung, 2 × 2; liver, 3.5 ×
4.6; fat, 0.42 × 50.6; and other tissues, 0.4 × 109. The lung
and liver compartments both have a depth of 20 µm and
the channels for other tissues, fat and interconnecting
network have a depth of 100 µm. A simpler two chamber
chip, with built-in grooves to minimize leakage, has also
been designed and is fabricated from polystyrene to pro-
vide a standard culture surface and enhance transpar-
ency (Figure 8C).
The biochips are incorporated into a four-chip hous-
ing which can be connected to a peristaltic pump.
Microfluidic channels interconnecting the compart-
ments permit compounds contained in “blood surro-
gate” culture medium, to re-circulate through and past
the respective cell chambers, emulating the circulatory
system of a living animal. The geometry of the system is
mathematically configured to simulate pharmacokinetic
parameters under a flow condition, such as drug resi-
dence time, circulatory transit time, organ cell density,
relative tissue size, shear stress, and others. This provides
an in vitro representation of relevant aspects of the blood
flow and pharmacokinetics of the living animal. The sys-
tem could potentially be further improved by enhancing
the physiological relevance of the tissue compartments
themselves (e.g. the use of more relevant cell systems,
implementation of more appropriate tissue architecture).
The flow system has been utilized to determine
whether the clearance data obtained using the integrated
co-culture and flow platform, would better predict in vivo
clearance (Novik et al., 2010). To accomplish this goal,
the clearance of nine compounds by human hepatocytes
was assessed while cultured under four different condi-
tions: flow-based culture in the presence and absence of
nonparenchymal cells, and static culture in the presence
and absence of the nonparenchymal cells. The intrinsic
rates determined for the static system were scaled and
the extraction ratios for the flow system were calculated
and scaled to in vivo values. The R2 coefficient of 0.9 was
obtained for the co-culture system under flow, whereas
poorer correlations were obtained for the monoculture
flow and static co-culture systems (0.7), and for the static
monoculture system (0.6). Overall, this flow-based co-
culture system cleared compounds with high-, medium-,
and low-clearance values with improved resolution and
predictive value. In addition, when co-culture was cou-
pled with flow, higher metabolite production rates were
obtained than in static systems.
For the question of insuring metabolic fidelity in
repeat exposure in vitro toxicity test systems, it may be
necessary to develop a microfluidic co-culture system
that maintains metabolism, recirculation, continuous
addition of test compound and ongoing loss from the cul-
ture system. The microfluidic, body-on-a-chip design has
532 E. L. LeCluyse et al.
Critical Reviews in Toxicology
the potential for creating custom in vitro toxicity evalua-
tions for multiple cells plated onto different parts of the
microfluidic plate (Maguire et al., 2009). The system was
designed based on PBPK model structures developed by
Shuler and colleagues (Esch et al., 2011). However, these
systems require more development, especially to scale
from a laboratory research device to a low- to medium
throughput screening tool.
5.4 3-D Liver tissue culture scaffold
Another platform for 3-D hepatocyte mono- and co-
culture with NPC is the Liver3 system, which consists of
co-cultures of liver cells from either animal or human
sources grown on a 3D scaffold under static or flow condi-
tions (Sibanda et al., 1993; Sibanda et al., 1994; Naughton
et al., 1994) (Figure 8D). The co-cultures are created by
procuring a donor liver from animal or human sources.
The cells of the native liver are isolated and separated into
hepatocyte and NPC populations. Hepatocytes attach to
and reside within a scaffold populated and modified by
the NPC and work in concert with these cells through
cell–cell and cell–matrix interactions to form a functional
tissue. The tissues are grown in multiwell plates or in cir-
culating long-term systems for metabolism and toxicity
assessment of new drug candidates or environmental
The concept for the Liver3 technology design is based on
biological principles of tissue and cell biology as outlined
previously (see “Past strategies for maintaining hepatic
structure and function in vitro”), where cells are cultured
on 3-D interconnecting porous structures and acquire a
more multidimensional, cuboidal configuration. Under
these conditions, cells are induced to express relevant
ECM proteins, migrate and co-locate with other cell types
to form native cell-cell interactions, and eventually form
a 3-D tissue structure over the course of several weeks.
The resultant tissue structure maintains differentiated
function of the different cell types, express liver-specific
proteins (e.g. albumin) and respond to inducers of CYP
enzymes (e.g. TCDD) and mediators of inflammatory
responses for up to 2 months in culture.
Overall, this culture platform provides enhanced biolog-
ical relevance and cellular function and longevity through
two key mechanisms: co-culture with nonparenchymal
cells and formation of 3-D cellular structures. A fundamen-
tal benefit of the system, the direct use of primary tissue
isolates, also presents a challenge in that the variability
present in proportions and numbers of cells from prepara-
tion to the next may present unintended variability in the
composition and functionality of the system. In addition,
the throughput of the current static and flow systems is typ-
ically medium to low. However, these issues may be minor
for many applications relative to the functional benefits
and biological relevance offered by the system.
5.5 3-D scaffolds with dynamic flow
A culture system that combines dynamic flow and
3-D cellular organization in a scalable platform that
attempts to mimic the dynamic environment of the aci-
nus was designed by Griffith and colleagues (Powers
et al., 2002a; Powers et al., 2002b; Sivaraman et al.,
2005; Hwa et al., 2007; Domansky et al., 2010; Dash
et al., 2009) (Figure 8E). The core of the platform is a
silicon or polycarbonate scaffold that contains circular
pores that are ~300 µm wide and ~300 µm deep (Powers
et al., 2002a). Primary hepatocytes are pre-formed into
3-D spheroids and seeded into these channels (Powers
et al., 2002a; Sivaraman et al., 2005; Hwa et al., 2007).
NPC can be included in these structures and will self-
sort to the outer margins of the spheroids (Powers and
Griffith, 1998) and of the 3-D structures within the scaf-
fold (Hwa et al., 2007). Cells in this system retain the
capacity to sort in a physiologically-relevant manner
with endothelial cells or other NPC localized at the tis-
sue-fluid interface. Scalability is achieved by increas-
ing the number of through-channels within the system,
and a higher-throughput version can be implemented
in multiwell plates (Sivaraman et al., 2005; Domansky
et al., 2010).
The system focuses on three key aspects of hepatic
physiology in enhancing hepatic function.
1) Dynamic flow: Perfusion rates and shear stresses
have been applied to authentically model those
experienced within the sinusoid.
2) Histotypic cellular structures: Cells are organized into
masses of cells that attempt to mimic the architec-
ture observed in sinusoids. The system is not able to
reliably or reproducibly create single-cell-thickness
structures as seen in hepatic plates, but those struc-
tures that are present appear to achieve appropriate
polarity in the context of fluid flow (Powers et al.,
3) Histotypic cellular organization: NPC can be read-
ily added to the system, and LSEC, KC and HSC all
appear to organize appropriately within this system.
A potential drawback is that it can be difficult to
control the precise ratios of these cells that eventu-
ally develop within the system over time, but this
requires further exploration.
Functional results with this system showed that gene
expression data of a panel of key hepatic markers, includ-
ing key transcription factors, CYP enzymes, transporters,
and nuclear receptors, mimic in vivo expression levels
more closely than collagen sandwich cultures. Likewise,
assessment of testosterone metabolism showed that
CYP activity is in better alignment with in vivo and
freshly isolated cells (Sivaraman et al., 2005). Further
assessment of metabolic clearance showed good cor-
relation with human in vivo clearance data. A panel of 9
compounds with disparate in vivo clearance profiles was
assessed in the system and a strong in vitro-in vivo cor-
relation was observed for eight of the nine compounds,
indicating good predictivity of hepatic clearance (Dash
et al., 2009).
Organotypic liver culture models 533
© 2012 Informa Healthcare USA, Inc.
6 applications in drug and chemical testing
Significant effort has been focused on developing a
strategy for predicting human pharmacokinetics and
toxicokinetics using a combination of in vivo animal and
in vitro human models. Although advances have been
made in both our biological understanding of chemical
toxicity and bioengineering principles, there continues
to be a need to fill major gaps in our scientific and tech-
nical understanding of complex mechanisms of hepato-
toxicity and our ability to extrapolate what is observed in
an artificial culture system to what may or may not occur
in vivo. Hepatic culture systems that are more organo-
typic in design and that incorporate the multicellular
and hemodynamic features of the tissue in vivo broaden
the scope of the basic and mechanistic studies that can
be conducted during chemical and drug testing. The
development of more physiologic, organotypic hepato-
cyte culture systems should also permit advances in our
understanding of how other cell types may be the cause
or downstream victim of chemical-induced perturba-
tions of key cellular pathways. For applications requiring
longer experimental periods, cells stably adapted to a
defined in vitro environment, or where cell architecture
and polarity are likely to be important, more sophisti-
cated culture systems will likely be far more relevant and
responsive for confirming or identifying chemical mode
6.1 Long-term study of low-dose exposures to drugs
Much of what we understand today about the toxicity
of environmental chemicals comes from subchronic
exposure of animals to relatively high doses. Chronic
exposure to low-doses of chemicals and drugs are likely
to cause completely different cellular and molecular
responses than those elicited under therapeutically
relevant doses in the patient population. In addition,
the primary, secondary and tertiary effects on gene
expression, stress pathways and adaptive responses will
require a model system that can allow for prolonged
culture periods while maintaining normal hepatic
function and adaptive responses. For certain types of
hepatotoxicity we now realize that the toxicity exhibited
under high-dose conditions does not easily extrapolate
back to those biochemical and molecular events that
will be involved in long-term, low-dose exposures that
occur under most conditions to humans (Slikker et al.,
2004a; Slikker et al., 2004b). As such, one of the more
beneficial uses of these more stable, long-term culture
systems will be the ability to study the time course and
kinetics of the initial onset of pathway perturbation
after exposure to compounds at physiological levels that
we know or assume to occur in vivo. In addition, the
secondary pathways and adaptive events that occur upon
activation of these initial pathways can be followed under
appropriate conditions. In some cases, having a second
cell type (e.g. KC), or appropriate microenvironmental
conditions, can allow for further exploration into the
causes and solutions to chemical-induced toxicity.
6.2 Elucidation of intercellular effects on the initiation
or propagation of chemical toxicity
The interactions between hepatocytes and other cell
types have significant consequences in the initiation
and progression of hepatotoxicity in vivo. For example,
the toxicity exhibited by acetaminophen (APAP) in the
liver has two phases, one beginning with its effects on the
LSEC and the second phase involving the classical necro-
sis of hepatocytes in zone 3 surrounding the central veins
(DeLeve et al., 1997; McCuskey et al., 2005). APAP hepa-
totoxicity predominantly occurs due to the bioactivation
of the parent compound by CYP enzymes to a highly
reactive quinoneimine (N-acetyl-p-benzoquinoneimine,
NAPQI), which depletes cellular GSH levels and begins
to attack nucleophilic targets under conditions of stress
(Mitchell et al., 1974; Corcoran et al., 1980). In the pres-
ence of cytochrome P450 inducers (such as PB), the
conversion of APAP to NAPQI and the consequential for-
mation of protein adducts leading to hepatotoxicity are
accelerated (Zhang et al., 2002; Kostrubsky et al., 2005).
Under normal circumstances, the combined metabolic
clearance capacity of healthy HC and LSEC can toler-
ate significant exposure to APAP (Mitchell et al., 1974).
For this reason, APAP represents an excellent candidate
compound to validate the robustness and metabolic
capacity of surrogate liver models that contain cell types
other than hepatocytes.
Many hepatotoxic responses are caused or exacer-
bated by corresponding immune system activation and
released paracrine factors, which cannot be mimicked
in simple monocultures of hepatocytes. For example,
KC activation contributes to a number of adverse effects
produced by hepatotoxic compounds (Jaeschke et al.,
2002; Jaeschke, 2007). They are also activated by many
exogenous and endogenous agents, such as cytokines,
endotoxins, and xenobiotics, including a number of
drugs (Wandzioch et al., 2004; Sunman et al., 2004;
Tukov et al., 2006). Activated KC contribute to hepato-
toxicity by producing free radicals (including superoxide
and nitric oxide) and cytokines, including TNF-α, IL-1,
and IL-6. TNF-α and, to a lesser extent IL-1, are major
mediators of cytotoxicity, and IL-6 is the major regulator
of the acute phase response (Streetz et al., 2001a; Streetz
et al., 2001b; Laskin et al., 2001; Dhainaut et al., 2001).
Activated KC also release chemokines, which attract and
activate neutrophils and lymphocytes that can potenti-
ate hepatotoxicity (Jaeschke et al., 2002). Even at subtoxic
doses macrophage activators can dramatically affect liver
function. For example, macrophage activation leads to a
robust down-regulation of xenobiotic-handling path-
ways in the liver, including many CYP and transporter
proteins (Morgan, 2001; Morgan, 2009; Renton, 2001).
Organotypic liver systems would allow the interac-
tion and adaptive responses between hepatocytes and
immune cells (e.g. KC and pit cells) under controlled
534 E. L. LeCluyse et al.
Critical Reviews in Toxicology
conditions. In addition, the role of infectious disease
and changes in cytokine levels can be examined more
systematically in organotypic model systems. Using toxi-
cogenomic approaches in conjunction with organotypic
co-culture systems, important relationships between
genes and biological pathways involving complex mecha-
nisms could be better defined. In addition, these systems
could provide new information about potential MOA’s of
prototypical and new hepatotoxicants and help create a
dataset of gene signatures that could be used to moni-
tor and identify potential hepatotoxic agents (McMillian
et al., 2004).
6.3 ‘Gold-standard’ to compare the biological
relevance of HTS assays
One of the more important roles that the advanced
culture models of human liver may serve is as a ‘gold-
standard’ for validating and confirming the relevance
of higher-throughput models. Admittedly, most of the
organotypic models described in this article will not be
easily scalable or adaptable to HTS. However, they poten-
tially represent an ideal human-surrogate with which
to compare and contrast data generated from simple
protein- or cell-based systems to provide some context
or confidence that the results are relevant to the in vivo
6.4 Continuity between studies within a single project
It is often burdensome when trying to repeat studies
utilizing the same cells from a particular donor for
repeat dosing or exposure to related compounds over
prolonged periods. As such, an added benefit of having
access to a long-term culture model that stably maintains
a consistent phenotype and genotype over prolonged
periods is that multiple studies or multiple repeat doses
can be performed with a single batch of tissues or cells.
This benefit would greatly increase the confidence and
reproducibility of study results within a particular project
as well as between compounds within a single series.
6.5 Mimicking dynamic exposure profiles
With advanced culture systems that allow control over
dynamic flow parameters, mimicking physiologically-
relevant exposure levels of a compound over time as well
as different physiologic and disease conditions becomes
theoretically possible. One of the shortcomings of tra-
ditional static culture models is the inability to repro-
duce the dynamic exposure levels that are experienced
by tissues and cells in vivo. If properly configured and
designed, studies could be conducted to mimic known in
vivo exposure levels of a compound over time (i.e. AUC)
that would better reflect the time and kinetic events that
lead to the perturbation of specific pathways of toxicity.
Although not currently achieved, future dynamic flow
culture systems that maintain overall metabolic capac-
ity of the tissues at or near in vivo levels would have the
added advantage of producing and recirculating poten-
tially active metabolites. In addition, these approaches
would allow more accurate descriptions of the onset of
events and subsequent adaptations to realistic exposure
6.6 Metabolite identification and profiling
Species differences in the expression and induction of
individual or multiple biotransformation and elimina-
tion pathways can lead to the production of different
metabolite profiles in humans compared to animal
models. The USFDA considers that the quantitative and
qualitative differences in metabolite profiles are impor-
tant when comparing exposure and safety of a drug in
a nonclinical species relative to humans during risk
assessment. When the metabolic profile of a parent drug
is similar qualitatively and quantitatively across species,
it is generally assumed that potential clinical risks of the
parent drug and its metabolites have been adequately
characterized during standard nonclinical safety
evaluations. However, because metabolic profiles and
metabolite concentrations can vary across species and
take time to manifest in vivo, there may be cases when
clinically-relevant metabolites have not been identified
or adequately evaluated during nonclinical safety stud-
ies. This situation may occur because the metabolite(s)
being formed in humans are absent in the animal test
species (unique human metabolite) or because the
metabolite is present at much higher levels in humans
(major metabolite) than in the species used during stan-
dard toxicity testing. As such, access to long-term culture
systems that maintain the relevant biotransformation
machinery for prolonged periods will greatly improve
our ability to identify and test relevant metabolites prior
to clinical testing.
Identification and toxicity profiling of relevant circu-
lating metabolites in humans can be very challenging
currently, especially using microsomes or pooled sus-
pensions of primary human hepatocytes, especially with
low-turnover compounds (McGinnity et al., 2004; Obach
et al., 2008; Obach, 2009). Long-term organotypic culture
systems make possible the examination of metabolite
production over longer periods of time as well as the
opportunity to examine their role in the initiation of toxic
events. In some cases, low levels of circulating metabo-
lites and not the parent compound are the cause of direct
or indirect toxicity to target cells. Most in vitro systems,
especially short-term cell-based models, do not generate
or provide a complete picture of the types and amounts
of important metabolites that may be generated in vivo in
humans. In many cases, it can be due to the lack of meta-
bolic capacity of the in vitro system, but in other cases it
can be the lack of physiologic context or exposure time.
Long-term advanced culture models, especially those
that retain the full complement of phase 1 and 2 enzyme
profiles at near physiologic levels, as well as those that
incorporate other cell types into the configuration, are
more likely to provide relevant profile of metabolites, if
not the corresponding kinetic and temporal patterns
under which they appear over time in vivo.
Organotypic liver culture models 535
© 2012 Informa Healthcare USA, Inc.
The ability to assess metabolism by examining com-
pounds in the effluent from the culture systems could
be coupled with other bioanalytical data to evaluate the
fidelity between the in vivo and in vitro pathways. A well-
designed liver bioreactor could function similar to an
isolated-perfused liver system and provide useful infor-
mation on the first-pass metabolism and disposition of
compounds (Bessems et al., 2006). Analysis of metabo-
lites produced in a bioreactor might also serve to bench-
mark expected metabolic pathways. Evaluation of the
fidelity of the bioreactor and new organotypic systems
could be verified by assessing metabolite profiles with
specific test compounds using prototype compounds
whose metabolism had already been well-studied in
vivo. In addition, coupled bioreactors containing cells
representing different tissue types could theoretically
reproduce physiologically-relevant tissue exposure pat-
terns of parent compound and metabolites (Li, 2009).
6.7 Toxicity testing and computational modeling for
human risk assessment
An NRC report, “Toxicity Testing in the 21st Century:
A Vision and A Strategy” , discussed challenges for
contemporary toxicity testing for chemicals in commerce
other than drugs (National Research Council, 2007;
Krewski et al., 2010). The goals of proposed changes
were to increase the speed of testing, enhance human
relevance, provide better information on modes of
action, reduce numbers of animals used and their degree
of suffering, greatly enhance coverage of chemicals in
commerce, and reduce costs. The vision was to conduct
most toxicity tests in vitro using human cells or cell lines
by evaluating perturbations of toxicity pathways that are
simply normal biological signaling pathways. Today,
simple cellular systems or molecular assays can produce
results with astonishingly high-throughput – many
thousands of tests per day. The NRC report discussed
tools for interpreting in vitro results for risk assessment
– i.e. computational systems biology models of pathways
and pharmacokinetic models to equate concentrations
active in vitro with exposure expected to lead to these
concentrations in human populations. However, there
remain significant questions about the relationship of
the in vitro responses and overt toxicity in intact animals.
In initial studies with compounds with extensive in
vivo testing results, the US EPA ToxCast™ program has
compared in vitro signals from multiple HTS (high
throughput screening) assays with known toxicity test
results to determine whether the HTS assay results are
predictive on responses in animals (Shah et al., 2011).
Other possibilities for comparisons across platforms
are from liver cells in suspension, to 2-D cultures, and
on to 3-D, organotypic cultures. Due to the longer-term
stability of 3-D cultures, assays can examine both ini-
tial targets and more integrated responses requiring
immune-cell activation, proliferation/mito-suppression,
fat accumulation, and adaptation over weeks of expo-
sure. These newer liver culture models should provide an
intermediate platform for assessing the ability of in vitro
test results to predict in life responses. The throughput
with organotypic platforms will be moderate to low, but
results from these assays could help ground more rel-
evant in vitro test systems against in vivo studies.
In addition to modeling cellular responses, more
integrated, virtual liver initiatives exist in both North
America and the EU (Shah and Wambaugh, 2010);
strategy_032309.pdf; http://www.virtual-liver.de/). The
overall concept with the US EPA Virtual Liver Project is
to predict liver toxicity using mathematical models that
span the spectrum from initial molecular targets, activa-
tion of key signaling pathways, alteration in biological
signaling networks and finally expressions of organ-level
and organism-level toxicity. The virtual liver project,
perhaps more specifically than the HTS efforts, exam-
ines the relationships of specific toxicity pathways and
adverse outcomes. The progress in developing 3-D liver
cultures should synergize virtual tissue efforts. For com-
pounds and pathways with known responses, the new
cultures should provide more mechanisms-based assays
for comparisons with existing toxicity results. For some
limited set of unknowns that lack in vivo results, the
organotypic cultures provide an opportunity to look at
the longer-term exposures and tease out a wider variety
of more integrated responses arising from cultures with
multiple cell types and by the ability to examine adaptive
responses occurring after initial tissue alterations from
target pathway activation.
Dose-response modeling of pathway assays will
depend on the ability to map and model the molecular
circuitry of pathway targets (Bhattacharya et al., 2011).
Empirical dose-response behaviors from perturbation of
the underlying biology of the circuitry would be collected
by conducting multipoint dose-response assessments.
Computational systems biology (CSB) modeling of the
pathway circuitry provides tools for calculating the differ-
ential dose-response. The core signaling processes in the
pathways include the cellular components involved in
signal recognition and the larger network through which
the initial perturbation propagates, eventually leading to
changes sufficiently large to suggest adverse potential.
For conducting experiments that will provide use-
ful data sets for computational modeling using in vitro
toxicity test systems, it will likely be necessary to develop
a co-culture system or a microfluidic system that main-
tains metabolism, recirculation, continuous addition of
test compound and ongoing loss from the culture system.
The microfluidic, body-on-a-chip design has the poten-
tial for creating custom in vitro toxicity evaluations for
multiple cells plated onto different parts of the microflu-
idic plate (Maguire et al., 2009). This system, which was
designed based on PBPK model structures developed
by Shuler and colleagues, requires more development,
especially to move from a laboratory research device to
low- to medium-throughput (Esch et al., 2011). Another
useful variation would be to have a hepatic bioreactor
536 E. L. LeCluyse et al.
Critical Reviews in Toxicology
with diverted flow to multiple chambers with various
other cell types for in vitro testing of metabolites. The
cells would have continuous flow of the bioreactor fluid
and the effluent from the culture plates could be col-
lected and re-circulated to the bioreactor. While these
designs are not yet readily available, they are techni-
cally within reach (Maguire et al., 2009; Novik et al.,
2010) and a number of new initiatives have been cre-
ated to develop a microfluidic ‘human-on-a-chip’ plat-
form (e.g. Defense Advanced Research Projects Agency
(DARPA), Microphysiological Systems, Broad Agency
7 Conclusions and future directions
The challenges that face the scientific community for
meeting the vision and standards for relevant in vitro
toxicity testing set by industrial, academic and regula-
tory demands are significant. A coordinated effort from
many scientific disciplines will be required to design and
create a more sustainable organotypic culture system of
the liver. The challenges are clear for retaining the native
configuration and phenotype of important cell types
along with the local hemodynamic conditions observed
in vivo. Material scientists, engineers, toxicologists and
biologists alike will be required to capture the respective
cell and tissue biology with current state-of-the-art mate-
rials and microfluidic platforms. Despite the scientific
and technical hurdles that must be overcome, substantial
progress has been made in recent years and the newer
hepatic culture technologies have begun to incorporate
more of the specific features that restore and maintain
phenotypic architecture and gene expression profiles
With the increased knowledge of the molecular and
cellular factors that determine hepatic structure and
function in vivo, improved incubation and cultivation
techniques have greatly expanded the utility and number
of applications for hepatocytes for toxicity testing. We
now know that the critical elements of matrix chemistry,
cell–cell interactions, and soluble media components
are interrelated and clearly dependent upon one
another for achieving optimal expression of hepatic
structure and function in vitro. In the liver, the specific
cellular niche, localized extracellular matrix chemistry,
and large number of soluble factors in the plasma and
interstitial fluid are equally important in regulating gene
expression and cell phenotype. Clearly, it is difficult
to duplicate exactly the dynamic environment of the
systemic and portal blood flow without incorporating a
corresponding dynamic in vitro culture environment. In
addition, the specific workflow and throughput demands
of a particular application will greatly affect the culture
conditions employed during the course of compound
testing and therefore the quality and relevance of the
corresponding data generated.
Each of the modifications discussed in this review
is subject to functional and logistical limitations. For
example, in the case of co-cultures, the presence of mul-
tiple cell types can complicate the analysis of drug extrac-
tion and metabolism. Moreover, additional experiments
must usually be run to determine the particular activity
of interest in the co-incubated cell lines themselves
to determine contaminating activity. Addition of high
concentrations of exogenous chemical agents for solu-
bilization (e.g. DMSO, alcohols) can lead to altered drug
metabolism due to induction of, or competition for, drug
metabolizing pathways. Cultures maintained on complex
substrata or sandwiched between two layers of extracel-
lular matrix are not amenable to transfection with DNA
constructs which limits the kinds of studies that can be
performed to examine the regulation of gene expression
(Pasco and Fagan, 1989).
The utility of any hepatic culture system for pharma-
cological and toxicological studies must also be consid-
ered in light of the architecture and function of the liver
as a whole. There are a number of metabolic differences
between periportal and perivenous hepatocytes in the
mammalian liver resulting from zonal differences in the
activity of several enzymes, and possibly from morpho-
logical differences as well (see section “Basic anatomy
and physiology of the liver”). The metabolic heteroge-
neity across regions of the liver lobule is thought to be a
function of the location in the microcirculation and may
be related to inherent gradients of oxygen, hormones,
metabolites, and matrix composition. Indeed, there are
distinct forms of hepatotoxicity that occur due to these
zonal differences in the gene expression patterns and
biochemical pathways of the respective cell types. As
such, in vitro model systems are likely to mimic only one
particular microenvironment at a time because control
of the dynamic differences in matrix chemistry, gene
expression profiles and gradients of soluble factors and
substrates is complex and beyond reasonable technolog-
ical expectations for the near future. However, it may be
possible to engineer consecutive organotypic cultures to
mimic sequential periportal, mid-zonal, and pericentral
conditions, or a single culture device that recapitulate
decreasing oxygen tensions across the perfusion flow
Another caveat to performing in vitro studies on any
isolated organ system, regardless of the level of engineer-
ing and sophistication, is that it does not adequately
address the complexities of the effects on the liver derived
from other areas of the body, such as delivery of portal
contents (e.g. lipids, endotoxins, gut-altered metabolites)
and humoral influences that may affect liver function
and blood flow secondary to chemical-induced liver
injury. Recapitulation within an isolated culture device
of the microenvironments and interactions of the various
liver cell types of the intact liver will not result alone in
a full reproduction and corresponding understanding of
the action of a xenobiotic on the liver as presented to an
animal or human in vivo.
With these limitations in mind, the latest 3-D, organo-
typic culture technologies and platforms offer valuable
Organotypic liver culture models 537
© 2012 Informa Healthcare USA, Inc.
alternatives to examine many issues relevant to toxicity
testing of drugs and other xenobiotics. In many respects,
these newer models of the liver represent the only in vitro
systems with which to conduct long-term toxicity testing
under well-defined conditions. Thus, they allow extended
studies of chemical interactions on cellular systems at
physiologically-relevant exposure levels. Whereas, other
in vitro model systems (e.g. liver slices, cell suspensions,
2-D static cultures) are limited by the short duration that
hepatocytes under these conditions retain acceptable
viability and liver-specific functions. Other advantages of
these advanced models include a reduction in the num-
ber of laboratory animals required for chemical and drug
testing due to the longevity of the systems and the ability
to repeat studies or conduct wash-out experiments using
the same system.
The development of three-dimensional tissue
engineering and microtechnology has narrowed the
gap between in vivo animal models and in vitro HTS
assays (Mazzoleni et al., 2009; Pampaloni et al., 2009;
Yang et al., 2009). Cells in microenvironments receive
signals from many different cell types and sources,
and certain pathways may only be recapitulated in a
3-D multicellular environment (Nirmalanandhan and
Sittampalam, 2009; Pampaloni et al., 2009; Mazzoleni
et al., 2009). Liver spheroids, which are an example
of a human tissue organoid, display more in vivo-like
responses than two-dimensional (2-D) counterparts (Lee
et al., 2009a; Lee et al., 2009b). Many tissues have already
been successfully engineered into 3-D format, including
the liver and cardiovascular tissues (Nirmalanandhan
and Sittampalam, 2009; Pampaloni et al., 2009; Hastings
et al., 2009; Hastings et al., 2007). The in vitro organotypic
model systems highlighted in this article are just a few
examples of the surrogate culture systems for human liver
cells that are viable and functional for several weeks. The
combination of stem cells, partially differentiated stem
cell systems, and 3-D tissue culture engineering should
greatly accelerate progress toward more effective toxicity
testing by providing the necessary renewable resources to
generate the human cells and tissues required to meet the
future demands for surrogate model systems. In addition,
the promise of pluripotent stem cells, if achieved, could
provide a renewable bank of cells representing different
genotypes and phenotypes including those that have
been associated with idiosyncratic drug-induced liver
injury. The improvements in the 3-D organotypic culture
platforms should provide the relevant context within
which to place the cells for greater predictive power and
As a final note, we find ourselves at a pivotal point
in time to advance the field of in vitro toxicology and
to address complex chemico-biological relationships
that underlie both reproducible and idiosyncratic toxic
responses that continue to plague both the chemical and
pharmaceutical industries. The significant progress being
made on advanced cell culture technologies is encourag-
ing for the eventual creation and employment of a more
predictive surrogate model of human liver. From our per-
spective, the opportunities for more rapid development
of improved in vitro ADME methodologies in general
are particularly timely. The technology to support these
initiatives in conjunction with the relevant scientific
knowledge and expertise is continuing to mature while
the needs within toxicity testing for both drugs and com-
mercial chemicals are continuing to grow. In addition, the
advances in stem cell biology should eventually allow the
development of custom bioreactors with more relevant
cellular composition, phenotypes and configurations.
With these additional improvements, the future biore-
actor systems will allow investigators to utilize them as
both metabolite generators and model systems to explore
the modes of action for hepatotoxicity and biological
responses to molecules. Future enhancements in these
areas should continue to prove valuable for the devel-
opment of more predictive in vitro surrogate models of
human toxicity, especially for pathways affecting the liver.
The authors thank Philip Lee (CellASIC), Salman Khetani
(Hepregen), Martin Yarmush and Eric Novik (Hµrel), and
Dawn Applegate (RegenMed) for providing background
information and materials for respective technology
platforms represented in Section “Advanced organotypic
culture technologies” of this manuscript. Figure 5E was
used with permission from the Royal Society of Chemistry
(RSC) (Domansky et al., 2010). The authors thank Dr. Lola
Reid, University of North Carolina at Chapel Hill, for the
illustration used in Figure 7, and John Jackson and Karli
E. Stephenson of Life Technologies for their assistance
with preparation of figures and final editing.
Declaration of interest
The authors’ affiliations are as shown on the cover page.
The contribution of ELL and MEA in this work was
supported by the Long-Range Research Initiative (LRI)
of the American Chemistry Council (ACC). The authors
alone are responsible for the content and writing of the
paper and have no conflicts of interest to declare.
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ADME, absorption, distribution, metabolism, excretion;
AE2, anion exchange protein 2; AhR, aryl hydrocarbon
receptor; AQP, aquaporin protein; APAP, acetamino-
phen; APC, antigen-presenting cells; AUC, area under
the curve; CAR, constitutively active receptor; CCL21,
chemokine (C-C motif) ligand 21; CCR5, C-C chemokine
receptor type 5; CFTR, cystic fibrosis transmembrane
conductance regulator; CINC-1, cytokine-induced neu-
trophil chemoattractant-1; CS-PG, chondroitin sulfate
proteoglycans; CTGF, connective tissue growth factor;
CYP, cytochrome P450; DILI, drug-induced liver injury;
ECM, extracellular matrix; EGF, epidermal growth factor;
ET-1, endothelin-1; FGF, fibroblast growth factor; GAG,
glycosaminoglycan; GGT, γ-glutamyltranspeptidase;
GSH, reduced glutathione; HC, hepatocytes; HCI, high-
content imaging; HGF, hepatocyte growth factor; HMGB-
1, high-mobility group box-1; HPC, hepatic progenitor
cells; HP-PG, heparin proteoglycans; HSC, hepatic stel-
late cells; HS-PG, heparan/heparin-sulfate proteogly-
cans; HTS, high-throughput screening; IGF-I and II,
Insulin-like growth factor I and II; IHL, intrahepatic
lymphocytes; lhx2, LIM homeobox gene 2; IL, interleu-
kin; iPSC, induced pluripotent stem cells; IVIVE, in vitro-
in vivo extrapolation; KC, Kupffer cells; LC-MS, liquid
chromatography-mass spectroscopy; LPS, lipopolysac-
charide; LSEC, liver sinusoidal endothelial cells; MAPC,
multipotent adult progenitor cells; MAPK, MAPKK,
mitogen-activated protein kinases; 3MC, 3-methyl-
cholanthrene; M-CSF, macrophage colony-stimulating
factor; MCP, monocyte chemotactic peptide; MHC,
major histocompatibility complex; MIP-2, macrophage
inflammatory protein-2; MOA, mode of action; NAPQI,
N-acetyl-p-benzoquinoneimine; NKC, natural killer cells;
NPC, nonparenchymal cells; MSC, mesenchymal stem
cells; PAPS, 3’-phosphoadenosine-5’-phosphosulfate;
PB, phenobarbital; PBPK, physiologically-based phar-
macokinetic; PDGF, platelet-derived growth factor; PG,
proteoglycans; PLT, platelets; PMN, polymorphonuclear
leukocytes; PPAR, peroxisome proliferator-activated
receptor; PSC, pluripotent stem cells; PXR, pregnane X
receptor; QSAR, quantitative structure–activity relation-
ships; RANTES, regulated on activation normal T-cell
expressed and secreted; RES, reticuloendothelial system;
RLEC, rat liver epithelial cells; SLC4A2, solute carrier
family 4 member 2; TAT, tyrosine aminotransferase; TGF-
α, transforming growth factor α; TLR, toll-like receptors;
TNF-α, tumor necrosis factor α; TGF-α, transforming
growth factor α; αSMA, alpha-smooth muscle actin;
UDPGA, uridine 5′-diphospho-glucuronic acid; UDP-GT,
Uridine 5′-diphospho-glucuronosyl transferase.