Calcium-dependent release of adenosine and uridine nucleotides from A549 cells

Extracellular nucleotides play an important role in lung defense, but the release mechanism and relative abundance of different nucleotide species secreted by lung epithelia are not well defined. In this study, to minimize cell surface hydrolysis, we used a low-volume, flow-through chamber and examined adenosine and uridine nucleotide concentrations in perfusate aliquots of human lung A549 cells challenged by 50% hypotonic shock. Adenosine triphosphate (ATP), adenosine diphosphate (ADP), adenosine monophosphate (AMP), and adenosine (Ado) were quantified in high-performance liquid chromatography (HPLC) analysis of fluorescent etheno derivatives, and uridine triphosphate (UTP) and uridine diphosphate (UDP) were measured using HPLC-coupled radioenzymatic assays. After the onset of hypotonic shock, ATP, ADP, UTP, and UDP in the perfusates increased markedly and peaked at approximately 2.5 min, followed by a gradual decay in the next 15–20 min; peak changes in Ado and AMP were relatively minor. The peak concentrations and fold increment (in parentheses) were: 34 ± 13 nM ATP (5.6), 11 ± 5 nM ADP (3.7), 3.3 ± 1.2 nM AMP (1.4), 23 ± 7 nM Ado (2.1), 21 nM UTP (>7), and 11 nM UDP (27). Nucleotide release was almost completely abolished from cells loaded with the calcium chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA). Under isotonic conditions, elevation of intracellular calcium with the calcium ionophore ionomycin (5 μM, 3 min) also released nucleotides with kinetics and relative abundance as above, albeit less robust. ADP:ATP (1:3) and UDP:UTP (1:2) ratios in perfusates from stimulated cells were markedly higher than the cytosolic ratios of these species, suggesting that a nucleotide diphosphate (NDP)-rich compartment, e.g., the secretory pathway, contributed to nucleotide release. Laser confocal microscopy experiments illustrated increased FM1-43 uptake into the plasma membrane upon hypotonic shock or ionomycin treatment, consistent with enhanced vesicular exocytosis under these conditions. In summary, our results strongly suggest that calcium-dependent exocytosis is responsible, at least in most part, for adenosine and uridine nucleotide release from A549 cells.
Introduction
Extracellular nucleotides control a diverse range of physiological processes by interacting with a large group of cell-surface P2Y and P2X receptor families [1–3]. In the lungs, extracellular nucleotides regulate airway surface liquid homeostasis and mucociliary clearance by modulating epithelial ion and fluid transport as well as ciliary beating [4]. In alveoli, adenosine triphosphate (ATP) is a potent secretagogue that stimulates type II cell-surfactant secretion. Nucleotide levels on airway surfaces, measured in vitro and ex vivo, show dynamic changes due to the combination of basal and stimulated release and their rapid metabolism by several groups of membrane-associated and membrane-secreted soluble ectoenzymes. These ectoenzymes extend the signaling potential of ATP by converting it to adenosine (Ado), a ligand of the A1–3 family of G-protein-coupled receptors [1, 4–7].
Nucleotide release is stimulated by cell mechanical perturbations, such as shear stress, membrane stretch, medium change, hyposmotic swelling, and hypoxia [1, 8]. It is now recognized that mechanosensitive ATP release occurs from healthy cells via physiological processes, which do not involve cell damage. Numerous reports have suggested that the cystic fibrosis transmembrane conductance regulator (CFTR) and other members of the superfamily of ATP-binding cassette-transport proteins serve as a conductive pathway for ATP release or regulate an associated ATP channel. However, other groups using patch-clamp, lipid-bilayer and luminometry techniques have not found any detectable CFTR-mediated or CFTR-regulated ATP release (reviewed in [1]). Volume-regulated anion channels (VRAC) and voltage-dependent anion channels (VDAC, porins or maxi-Cl− channels) are known to have considerable permeability to cations and large organic anions, and several laboratories have implicated these channels in cell-swelling-induced ATP release and the autocrine regulation of cell volume. However, recent studies have demonstrated that conductive pathways, including VRAC, VDAC, and stretch-activated channels, are not involved in cell-swelling-induced ATP release from A549 cells [1, 9, 10]. Finally, connexin hemichannels also have been proposed to mediate ATP release, but their role in this process in a physiological setting is not always clear [1].
Whereas conductive release mechanisms are still being debated, exocytotic ATP release is well established in excitatory cells, blood platelets, and chromaffin cells. Furthermore, there is growing experimental evidence supporting such a release mechanism also by epithelial and other nonexcitatory cells. In particular, we recently demonstrated that cell-swelling-induced ATP release from A549 and 16HBE14o− epithelial cells and 3T3 fibroblasts is tightly correlated with intracellular Ca2+ elevations; it is abolished in cells loaded with the calcium (Ca2+) chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA) or by low temperature, strongly suggesting the involvement of Ca2+-dependent exocytosis [10]. Whether release of other nucleotides from epithelial cells also involves Ca2+-dependent exocytosis remains incompletely explored. Such studies, however, are complicated by nucleotide hydrolysis at the airway surface, which makes it difficult to assess the magnitude and relative abundance of different nucleotide species released. Therefore, we used a flow-through chamber to minimize cell surface hydrolysis.
We found that hypotonic shock markedly increased ATP, adenosine diphosphate (ADP), uridine triphosphate (UTP), and uridine diphosphate (UDP) concentrations in perfusates, which peaked at approximately 2.5 min. Nucleotide release was almost completely abolished from cells loaded with BAPTA and, under isotonic conditions, could be evoked by elevation of intracellular calcium with the calcium ionophore ionomycin. High nucleotide diphosphate (NDP) concentrations in perfusates of stimulated cells suggested that an NDP-rich compartment, e.g., the secretory pathway, contributed to this release. Together with real-time FM1-43 fluorescence experiments, our results strongly indicate that calcium-dependent exocytosis is a major mechanism of adenosine and uridine nucleotide release from A549 cells.
Materials and methods
Cells
Human lung carcinoma A549 cells were grown in Dulbecco’s modified eagles medium (DMEM) supplemented with 10% fetal bovine serum (FBS), 2 mM L-glutamine, 56 U/ml penicillin-G and 56 μg/ml streptomycin sulfate. All constituents of the culture media were from GIBCO-BRL (Burlington, ON, Canada). ATP efflux was measured from cell monolayers grown to confluency on 24 × 60-mm glass coverslips. Fura-2 calcium imaging and FM1-43 microscopy experiments were performed on cells grown on circular 15-mm diameter no. 1 glass coverslips.
Nucleotide efflux assay
To measure nucleotide efflux during hypotonic challenge with high temporal resolution, we used a custom-designed, low-volume (300 μl), flow-through chamber, as described previously [10]. Briefly, 24 × 60-mm glass coverslips with confluent cell monolayers of ∼500 cells/mm2 were mounted in the chamber and perfused with warm (37°C/in-line heater; Warner Instrument, Hamden, CT, USA) Ringer solution at the rate of 1.3 ml/min. The Ringer solution contained (in mM): 110.5 NaCl, 24 NaHCO3, 1.3 KCl, 1 MgCl2, 1 CaCl2, 2.5 Na2HPO4, 2.5 KH2PO4, 1.2 K2HPO4, and 10 glucose; pH 7.4 was maintained by bubbling with 5% CO2. After an equilibration period in isotonic solution (5–15 min), a 50% hypotonic solution was applied, and the perfusate was continuously collected for 30-s intervals during the initial burst of ATP secretion (0–5 min) and during 1 min elsewhere. Nucleotide concentrations in the samples were evaluated by high-performance liquid chromatography (HPLC0 analysis, as described below. For studies in absence of extracellular calcium, CaCl2 was omitted, and the solutions were supplemented with 0.1 mM ethyleneglycoltetraacetic acid (EGTA). The 50% hypotonic solution was prepared by appropriate reduction of salt concentration, whereas divalent cation concentrations were kept constant.
HPLC quantification of adenine and uridine nucleotides
UTP concentrations were quantified by UDP-glucose pyrophosphorylase-based reaction [11]. Briefly, 100-μl samples were incubated in the presence of 0.5 U/ml UDPglucose pyrophosphorylase, 0.5 U/ml inorganic pyrophosphatase, 1.6 mM CaCl2, 2 mM MgCl2, 25 mM hydroxyethylpiperazine ethanesulfonic acid (HEPES) (pH 7.4), and ∼100,000 cpm 1 μM [14C]glucose-1P. Incubations lasted 1 h at 30°C. Reactions were terminated by heating the samples at 95°C for 2 min. Conversion of [14C]glucose-1P to [14C]UTP was determined by HPLC (Shimadzu) via a Nova Pack C18 column and ion-pairing mobile phase. Radioactivity was measured on-line with a Packard Flo-One detector. Assay sensitivity was 1 pmol (3 nM in 100-μl samples).
UDP was quantitatively phosphorylated in the presence of [γ32P]ATP using nucleoside diphosphokinase (NDPK). Briefly, 100-μl samples containing 0.1 U/ml NDPK, 0.1 μCi 60 nM [γ32P]ATP, 1.6 mM CaCl2, 2 mM MgCl2, and 25 mM HEPES (pH 7.4) were incubated for 5 min at 30°C. Reactions were terminated by heating the samples at 95°C for 2 min. The resulting conversion of [γ32P]ATP to [γ32P]UTP was monitored by HPLC [12]. A calibration curve employing known amounts of UDP was chartered in parallel during each assay. This assay allows UDP quantification with sensitivity of 20 fmol (0.2 nM in 100-μl samples).
For etheno (ɛ) derivatization, samples (200 μl) were derivatized for 30 min at 72°C in the presence of 1.0 M chloroacetaldehyde and 25 mM Na2HPO4 (pH 4.0). The resulting fluorescent 1,N6-ethenoadenine derivatives were analyzed by HLPC (Waters) in a Hamilton PRP-×100 anion exchange column, as described previously [4]. ɛ-ATP, ɛ-ADP, ɛ-AMP, and ɛ-Ado were quantified with a sensitivity of 200 fmol (1 nM in 200-μl samples).
FM1-43 studies
Bulk exocytosis was quantified in A549 cells as changes in the fluorescence intensity of FM1-43 incorporated into the plasma membrane [13, 14] and recorded by real-time confocal microscopy. Cells were washed with Hank’s balanced salt solution + 20 mM HEPES + 2 mM MgCl2 and 1.6 mM CaCl2 (HBSSH) or with HBSSH devoid of calcium (HBSSH 0 Ca). They were mounted onto the stage of a Leica SP2 AOBS confocal microscope equipped with HCX Apo L63x NA 0.9 immersion Leica lens and a 488-nm Argon laser. Experiments were initiated by incubation of the cells with 3 μM FM1-43. Cells were incubated for at least 10 min, and hypotonic shock was applied by decreasing salt concentration 33% while maintaining calcium, magnesium, and FM1-43 concentrations constant. Alternatively, cells incubated with FM1-43 in HBSSH or HBSSH 0 Ca were stimulated with 5- to 10-μM ionomycin. Real-time recording was performed by laser scanning in thexz axes with a galvostage, initially every 10 s and then every 30 s for the time periods indicated in the figures. Overall fluorescence intensity changes associated with the plasma membrane were estimated by measuring the intensity value associated with each pixel through time. The entire apical membrane compartment displayed in a confocal plane and five random regions of basolateral and subapical domains were analyzed, normalized to basal values (time = 0), and averaged for each region. Cell swelling was estimated as a change of cell height in thexz plane at different time points, normalized to basal values, and averaged.
Fura-2 calcium measurements
To load Fura-2, cells were incubated (1 h, 37°C, 5% CO2) in physiological solution containing 25 μM Fura-2-AM + 0.02% Pluronic F127 and 2.5 mM probenecid. This was followed by 30 min deesterification period in physiological solution containing probenecid. The physiological saline solution consisted of (in mM): 140 NaCl, 5 KCl, 1 MgCl2, 1 CaCl2, 10 glucose and 10 HEPES, pH 7.4, adjusted with sodium hydroxide (NaOH). Fifty percent hypotonic medium was prepared by reducing the salt concentration while keeping divalent cation concentration constant. For calcium imaging, coverslips with Fura-2-loaded cells were mounted in the imaging/perfusion chamber attached to the heated platform (Warner Instruments) on the stage of an inverted microscope (Nikon TE300). The cells were exposed to alternate (200 ms) illumination at 340 nm and 380 nm with a high-pressure mercury lamp (100 W) via interference filters (Chroma Technology, Brattleboro, VT, USA) mounted on a filter wheel (Sutter Lambda 10-C, Sutter Instrument, Novato, CA, USA) and a dichroic mirror (510/540 nm, Chroma Technology). Fluorescence images were recorded at 15- to 60-s intervals with the digital camera and stored for later analysis.
Chemicals
For calcium-imaging experiments, Fura-2-AM was obtained from Molecular Probes, Invitrogen Corp. (Burlington, ON, Canada). Probenicid, Pluronic F127 and all other reagents were from Sigma Aldrich (Oakville, ON, Canada).
Results
Kinetics of nucleotide release
Figure 1 shows an example of the time course of nucleotide release induced by 50% hypotonic shock. For clarity, the release of adenine and uridine nucleotides appears on separate graphs: a and b, respectively. The kinetics of release were remarkably similar for all nucleotides. After the onset of hypotonic shock, nucleotide concentration increased rapidly, peaking at ∼2.5 min, followed by a gradual decay in the next 10–15 min. The average peak values from several separate experiments are shown in Fig. 1c. Interestingly, ATP was the major species at the peak of stimulated release, whereas for basal release, Ado was the predominant species. The rank order of nucleotide abundance at the peak was: ATP > Ado ≥ UTP > ADP ≈ UDP > AMP. Relative increases of nucleotide conconcentrations at the peak were also the highest for nucleotide triphosphates (NTPs) and nucleotide diphosphates (NDPs) (ATP 5.6-fold, ADP 3.7-fold, UTP >7-fold, UDP 27-fold), whereas the increase was smaller for AMP and Ado (1.4-fold and 2.1-fold, respectively).
Fig. 1Transient nucleotide release from A549 cells induced by 50% hypotonic shock. Time course of adenosine (a) and uridine (b) nucleotide release observed in response to 50% hypotonic shock. A representative experiment is shown out of four performed under the same conditions. Hypotonic shock was applied at t = 0 min and was preceded by 15-min equilibration in isotonic solution. Basal (t = 0 min) and peak (t = 2.5 min) nucleotide concentrations detected in perfusates (c). Average values [± standard deviation (SD)] are from three to four experiments, such as in a and b, except for uridine triphosphate (UTP) and uridine diphosphate (UDP), which are from a single experiment. * Due to the limited sensitivity of UTP evaluations (≈3 nM), the basal level of UTP was found to be below the detection limit and was not shown
Role of intracellular calcium ([Ca+2]i)
To investigate the role of intracellular calcium ([Ca+2]i) in adenine nucleotide release, we tested the effect of the Ca2+ ionophore ionomycin. In the absence of hypotonic shock, exposure to 5 μM ionomycin for 3 min induced transient nucleotide release from A549 cells (Fig. 2a). This release reached a peak at approximately 1.5 min, i.e., slightly earlier and at somewhat lesser absolute peak amplitude compared with that induced by 50% hypotonic shock. Otherwise, the kinetics and relative nucleotide abundance were similar for both stimuli. Almost complete inhibition of swelling-induced adenine nucleotide release was observed for A549 cells loaded with the Ca2+ chelator BAPTA, compared with control untreated cells of the same batch tested in parallel experiments (Fig. 2b, c, respectively). Fura-2 fluorescence [Ca+2]i measurements confirmed a dramatic reduction of the [Ca+2]i response to 50% hypotonic shock in BAPTA-loaded cells (Fig. 2d). These experiments demonstrate that elevation of [Ca+2]i was required to trigger adenine nucleotide release from A549 epithelial cells.
Fig. 2Effect of intracellular calcium ([Ca2+i]) modulators on nucleotide release. (a). In the absence of hypotonic shock, application of 5 μM ionomycin resulted in transient nucleotide release with kinetics similar to that induced by hypotonic shock. An example, out of two similar experiments, is shown. Loading A549 cells with the Ca2+ chelator 1,2-bis(2-aminophenoxy)ethane-N,N,N',N'-tetraacetic acid (BAPTA) almost completely inhibited hypotonic-stress-induced nucleotide release compared with controls that were run in parallel (b and c, respectively) (n = 2). Effect of BAPTA on the [Ca2+]i response (d). The two traces represent changes of the Fura-2 fluorescence ratio at two excitation wavelengths, λ340/λ380, in response to 50% hypotonic shock, applied at t = 0 min, in control and BAPTA-loaded cells. Note that the rapid peak of the [Ca2+]i response was abolished in BAPTA-loaded cells. Similar responses were observed in four experiments
FM1-43 fluorescence changes implicate vesicular exocytosis
Strong calcium dependence of nucleotide release may indicate the involvement of vesicular exocytosis. To further explore this possibility, bulk exocytosis was examined by real-time confocal microscopy in cells bathed in FM1-43. Fluorescence intensity associated with the plasma membrane increased rapidly by incorporation of the soluble probe into the membrane and remained almost constant after 5 min in nonstimulated cells. Plasma-membrane-associated fluorescence intensity rose rapidly (∼20 s) with hypotonic shock stimulation. This change was accompanied by increased cell volume seen as increment of cell height (Fig. 3a, b). Cell volume peaked at about 3 min, followed by a regulatory volume decrease in the next 10–15 min (data not shown). A similar plasma-membrane-associated FM1-43 fluorescence intensity increase was observed in the absence of hypotonic shock when the cells were stimulated with ionomycin in the presence but not in the absence of calcium in the bathing solution (Fig. 4). These data indicate that both hypotonic-shock- and ionomycin-stimulated bulk exocytosis in A549 cells.
Fig. 3Effect of hypotonic shock on FM1-43 surface membrane staining. Confocal FM1-43 fluorescence images of confluent A549 cells after 20 min of dye loading in isotonic solution, followed by 50% hypotonic shock stimulation (a). The images depict the xz scans of the cell monolayer at different time points. Note the increase in cell size and fluorescence intensity consistent with hypotonic-shock-induced cell swelling and heightened bulk exocytosis. Time course of FM1-43 plasma membrane fluorescence and cell height changes during hypotonic shock (b)Fig. 4Effect of ionomycin on FM1-43 surface-membrane staining. Confocal FM1-43 fluorescence images of confluent A549 cells prior to and after 4-min exposure to 5 μM ionomycin in the presence of extracellular calcium. (a). The images show the xz scans of the cell monolayer at different time points. Note the increase in fluorescence intensity after ionomycin stimulation, indicating enhanced exocytosis. Time course of FM1-43 plasma membrane fluorescence changes of unstimulated cells (Δ CON) and cells during ionomycin treatment in the presence (▪ ION) or absence (• ION 0 Ca) of extracellular calcium (b)
Discussion
In this study, we used the flow-through chamber and an etheno derivatization measurement technique to examine the kinetics of nucleotide release from A549 lung epithelial cells induced by hypotonic shock. We demonstrated that ATP, ADP, AMP, Ado, UTP, and UDP appeared in the perfusates with the same kinetics, peaking at ∼2.5 min (Fig. 1), which coincided with the peak of [Ca+2]i elevation evoked by hypotonic shock (Fig. 2d). Similar tight temporal correlation between the [Ca+2]i signal and ATP release, measured by luciferase-luciferin luminescence assay, was reported previously for A549, 16HBE14o− epithelial cells and NIH3T3 fibroblasts [10]. Such a close temporal association suggests that hypotonic-shock-induced elevation of [Ca+2]i is a trigger for the release of nucleotides. This was supported by the strong inhibitory effect of BAPTA-AM and the induction of nucleotide release in the absence of hypotonic shock solely by the rise of [Ca+2]i with ionomycin. These data demonstrate tight Ca2+ dependence of nucleotide release, and point to a Ca2+-dependent exocytotic mechanism. It should be noted that the bulk of [Ca+2]i changes shown in Fig. 2d likely represent superposition of spatially separated, distinct [Ca+2]i responses, one evoked directly by hypotonic shock and the other due to autocrine/paracrine actions of the released nucleotides on P2Y receptors. The latter effects might be minimized, at least partially, by continuous perfusion, which reduced nucleotide concentration in the bulk of the chamber far below the IC50 of P2Y receptor activation; however, this possibility requires further study.
Consistent with a Ca2+-dependent exocytotic mechanism, FM1-43 fluorescence studies revealed enhanced incorporation of the probe into the plasma membrane induced by hypotonic shock or ionomycin treatment. Increased plasma membrane-associated FM1-43 fluorescence intensity was a measure of the cumulative amount of membrane added by exocytosis [13]. FM1-43 fluorescence increased rapidly during the first 20 s of hypotonic shock (Fig. 3). Interestingly, the initial fluorescence increment exceeded the cell-surface increase estimated from xz confocal scans. In our previous study using a dual-image 3D cell reconstruction technique [15], we observed only an 11–30% surface increase of single substrate-attached A549 cells swollen in 50% hypotonic solution [10]. This difference could be attributed in part to different methods of cell-height and surface evaluation in that study, as well as differences in swelling responses of single isolated cells compared with confluent cell monolayer. However, a similar discrepancy between FM1-43 fluorescence, cell-surface and membrane electrical capacitance has been also reported by others, e.g., with pituitary lactotrophs, where the dense granules docked into the plasma membrane were intensely stained by FM1-43 in addition to the membrane added to the cell surface [16]. Therefore, the divergence of fluorescence and the cell-surface changes seen in our study with A549 cells may also result in part from FM1-43 staining of the lipophilic content of fused vesicles in addition to the staining of fused vesicular membranes. The FM1-43 dye is often employed to fluorescently marked surfactant-containing lamellar bodies in alveolar type II (ATII) cells [17, 18] and A549 cells, a model of ATII cells, also contain granules enriched in surface-active phospholipids [19]. However, the exact contribution of such granules to increased FM1-43 fluorescence in stimulated A549 cells was out of the scope of the present investigation and will require future direct study.
Vesicle exocytosis may be mechanistically associated with nucleotide release in two ways: nucleotides can be delivered to the extracellular medium as cargo molecules within exocytotic vesicles or, alternatively, secreted from the cytosol via vesicle-associated nucleotide-conducting channels or transporters transiently expressed at the plasma membrane as a consequence of vesicle-plasma membrane fusion. Analysis of the relative abundance of different nucleotide species that appear in the extracellular medium may help to distinguish between these two mechanisms. ATP and UDP sugars accumulate in the lumen of the secretory pathway up to 20- to 50-fold above their cytosolic concentrations, and luminal utilization of these molecules generates ADP and UDP. UDP is the major uridine nucleotide detected in ER/Golgi fractions [20] and, eventually, a UDP-selective apyrase (UDPase) converts UDP to urodine monophosphate (UMP) [21]. ER/Golgi ADP and UMP are exchanged for cytosolic ATP and UDP glucose via specific transporters [22–25]. As intraluminal nucleotides are not subject to the mechanisms that retrieve resident ER/Golgi proteins, they are predicted to remain within trafficking vesicles and to be released to the extracellular space from the secretory pathway. Our results suggest that this mechanism likely accounted for UDP release from A549 cells. Figure 1c illustrates that hypotonic stimulation promoted a sharp increase in UDP levels, with a UTP:UDP concentration ratio of 2:1. As the cytosolic UTP:UDP ratio is >10:1 [26], our results suggest, at least in part, a vesicular rather than a cytosolic source of UDP and, as a corollary, of ATP and ADP.
In summary, our results provide strong evidence for Ca2+-dependent vesicular exocytosis as a major mechanism of adenosine and uridine nucleotide release from A549 epithelial cells induced by hypotonic stress. Part of this release involves vesicles of the protein secretory pathway. Further investigations are needed to clarify the origin and contribution of other vesicular pools as well as the mechanisms and sources of intracellular Ca2+ elevations that evoke nucleotide secretion.

hypotonic shock
exocytosis
lung epithelial cells
ca2+ dependence
nucleotide secretion


Identification of Genetic and Epigenetic Variations in a Rat Model for Neurodevelopmental Disorders

A combination of genetic variations, epimutations and environmental factors may be involved in the etiology of complex neurodevelopmental disorders like schizophrenia. To study such disorders, we use apomorphine-unsusceptible (APO-UNSUS) Wistar rats and their phenotypic counterpart apomorphine-susceptible (APO-SUS) rats that display a complex phenotype remarkably similar to that of schizophrenic patients. As the molecular basis of the APO-SUS/UNSUS rat model, we recently identified a genomic rearrangement of the Aph-1b gene. Here, we discovered between the two rat lines differences other than the Aph-1b gene defect, including a remarkable cluster of genetic variations, two variants corresponding to topoisomerase II-based recombination hot spots and an epigenetic (DNA methylation) difference in cerebellum and (hypo)thalamic but not hippocampal genomic DNA. Furthermore, genetic variations were found to correlate with the degree of apomorphine susceptibility in unselected Wistar rats. Together, the results show that a number of genetic and epigenetic differences exist between the APO-SUS and -UNSUS rat genomes, raising the possibility that in addition to the Aph-1b gene defect the newly identified variations may also contribute to the complex APO-SUS phenotype.
Introduction
Schizophrenia is a neurodevelopmental disorder affecting nearly 1% of the world’s population (Jablensky et al. 1987), and is characterized by positive and negative symptoms (Kay and Opler 1987). The aetiology of schizophrenia and other related disorders, such as schizoaffective and bipolar disorder, is still unclear. Twin, family and adoption studies have suggested that complex interactions at the genetic and environmental level underlie the aetiology of schizophrenia (Gottesman 1991). It is thought that gene variations by themselves do not result in schizophrenia, but they can establish a predisposition status that, when combined with environmental stressors, may lead to schizophrenia pathogenesis. Numerous environmental factors, such as viral infections (Mednick et al. 1988), insufficient folate and methionine levels (Regland 2005), or repeated psychological stress (Goldstein 1987), can influence brain development of prenatal or early postnatal individuals with a genetic predisposition for neuropsychiatric disorders. Due to the heterogeneity in genetic and environmental interactions, most of the genes and pathways for schizophrenia and for other complex disorders are still unknown.
To get insight into the gene (or genes) that may be involved in schizophrenia pathogenesis, a rat model was developed with schizophrenia-like features. This model was based on the behavioural response of Wistar rats to the dopamine agonist apomorphine (Cools et al. 1990). The apomorphine-susceptible (APO-SUS) rat line displayed many features of psychopathology, with similar disturbances at the behavioural, physiological, endocrinological and pharmacological level as seen in schizophrenics (Ellenbroek and Cools 2002). For example, APO-SUS rats have a reduced prepulse inhibition and latent inhibition (Ellenbroek et al. 1995), display a higher plasma release of adrenocorticotropin (ACTH) and corticosteroids in response to novelty (Rots et al. 1995), are more sensitive to dopamimetic drugs (Ellenbroek et al. 2000), and have a higher susceptibility to inflammatory and infectious diseases when compared to apomorphine-unsusceptible (APO-UNSUS) rats (Kavelaars et al. 1997). We therefore wondered about the molecular-genetic basis underlying the APO-SUS/-UNSUS rat model and recently identified a genetic difference between the two rat lines (Coolen et al. 2005). Whereas APO-UNSUS rats harbour three gene copies of the γ-secretase component Aph-1b, APO-SUS rats have only one or two copies. This gene-dosage imbalance was due to an unequal crossing over event (nonallelic homologous recombination) between two direct repeats (a segmental duplication) within the Aph-1b locus. In addition, we observed a direct link between the Aph-1b genotypes and a number of phenotypic APO-SUS and -UNSUS characteristics (Coolen et al. 2005). Approximately 10 years after developing the APO-SUS and -UNSUS lines a second, independent breeding procedure was started that resulted in rats with features similar to those displayed by the original APO-SUS and -UNSUS rat lines (Ellenbroek and Cools 2002). Interestingly, the replicated rat lines also resulted in APO-UNSUS rats with three Aph-1b gene copy numbers and APO-SUS rats with only one or two gene copies (Coolen et al. 2005).
In the present study, we wondered whether genetic variations other than the Aph-1b gene-dosage imbalance may be present between the APO-SUS and -UNSUS rats, and whether epigenetic factors may be involved as well. Epigenetics has been defined as heritable changes in gene expression that do not occur by changes in the DNA sequence, but by modifications in DNA methylation and chromatin remodeling (Wolffe and Matzke 1999), or, in its widest sense, as any change in an organism that is not due to genetic factors (Van de Vijver et al. 2002). Increasing evidence suggests that epigenetic modifications play a role in disease susceptibility (reviewed by Jirtle and Skinner 2007). We used the arbitrarily primed-polymerase chain reaction (AP-PCR) fingerprinting technique (Welsh and McClelland 1990) to analyse the genomes and epigenomes (DNA methylation) of the APO-SUS and -UNSUS rats. Comparison of the AP-PCR fingerprints generated from the genomic DNAs of the two rat lines revealed genetic as well as epigenetic alterations and we conclude that, besides in the Aph-1b locus, a number of other variations are present in the APO-SUS and -UNSUS genomes and epigenomes.
Materials and methods
Experimental animals
The generation of the APO-SUS and -UNSUS rat lines with a high or low susceptibility for apomorphine, respectively, has been described previously (Cools et al. 1990). The present experiments were performed with male APO-SUS and -UNSUS rats belonging to the 32nd (original lines) and 18th (replicate lines) generation. At post-natal day 60 (PND60), APO-SUS and -UNSUS rats were sacrificed and the hippocampus, cerebellum and the combined thalamus/hypothalamus (further denoted as (hypo)thalamus) were isolated. To establish their apomorphine susceptibility, unselected male Wistar rats of the Nijmegen outbred population (PND60) were injected with apomorphine (1.5 mg/kg s.c.) and gnawing scores were measured in a gnawing box for 45 min, as described previously (Cools et al. 1990). Immediately following the measurements, the rats were sacrificed and the same tissues (hippocampus, cerebellum and (hypo)thalamus) were removed. All rats were bred and reared in the Central Animal Facility of the Radboud University Nijmegen under approved animal protocols and in accordance with institutional guidelines.
Arbitrarily primed-PCR
Genomic DNAs were isolated from hippocampus, cerebellum and (hypo)thalamus using standard procedures involving the use of proteinase K and phenol extraction. Two micrograms of genomic DNA were digested with 20 units of RsaI, 20 units RsaI in combination with the methylation-sensitive enzyme HpaII, or 20 units RsaI and MspI (MBI Fermentas) in a total volume of 40 μl at 37°C for 16 h. HpaII does not cut DNA if the internal cytosine (CCGG) is methylated, whereas MspI is insensitive to DNA methylation. Using such combinations of methylation-sensitive and -insensitive enzymes allows genome-wide screening for differences at the genetic level (single-nucleotide polymorphisms––SNPs, duplications, insertions, deletions and recombinations) as well as the epigenetic (DNA methylation) level. Restriction enzymes were heat inactivated by incubating the reactions at 65°C for 20 min. Digested DNA (100 ng) was amplified using AP-PCR (Welsh and McClelland 1990) with a single primer. PCRs were performed in a total volume of 25 μl containing 10 mM Tris-HCl (pH 8.3), 2.5 mM MgCl2, 50 mM KCl, 0.001% gelatin, 0.25% Nonidet P-40, 0.25% Tween-20, 200 μM each of the four deoxynucleotide triphosphates, ∼1 μCi of [α-32P]dCTP (3000 Ci/mmol, Amersham Corp.), 25 pmol of primer (AP-1: 5′-AACCCTCACCCTAACCCCGG-3′, AP-7: 5′-AACCCTCACCCTAAGGCGCG-3′, AP-777: 5′-CACTCCTCTACAAGGTGCCG-3′ or Topo: 5′-GCCTCCTTGCAGGTCTTT-3′), and 0.8 units of Taq polymerase (MBI Fermentas). Reactions were carried out in a thermal cycler (Perkin-Elmer) with five cycles of low stringency (94°C for 30 s, 40°C for 60 s, 72°C for 1.5 min), followed by 30 cycles of high stringency (94°C for 15 s, 55°C for 15 s, 72°C for 1 min). Two microliters of the PCR products were analysed on high-resolution 5% polyacrylamide gels under denaturing conditions (7 M urea) for 4–4.5 h at 70 W. Gels were dried and radiolabelled DNA was visualized by autoradiography at −70°C (CEA AB, Sweden).
Cloning and sequencing of AP-PCR fragments
AP-PCR fragments generated from APO-SUS and -UNSUS rat genomic DNAs were excised from the dried gels and incubated in 50 μl MilliQ at 80°C for 10 min. The eluted DNA (two microliters) was reamplified with the same primer as used for the AP-PCR to generate sufficient amounts of template for subsequent cloning. The reactions were carried out for 40 cycles of 94°C for 1 min, 55°C for 30 s, 72°C for 1 min, under the same conditions as described in the AP-PCR protocol (except that [α-32P]dCTP was not included). The PCR products were purified, cloned into the pGEM-T easy vector (Promega) and sequenced with a T7 or Sp6 primer according to the manufacturer’s instructions using the ABI310 machine (Applied Biosystems).
Sequencing and genotyping of chromosomal region 9q22
A 1948-bp fragment that harbours the nucleotide sequence corresponding to product 3 was obtained by PCR on genomic DNA derived from (hypo)thalamic tissue of an APO-UNSUS rat using forward primer 5′-GGGAAGCAACGCATCCTG-3′ and reverse primer 5′-CATATCAAAGCACCAAGTCCACAG-3′. The DNA was subsequently purified and directly sequenced using the ABI310 machine (Applied Biosystems). Genotyping of chromosomal region 9q22 was performed with PCR using primers specific for either the APO-SUS or APO-UNSUS genomic sequence. Briefly, PCRs were performed in a total volume of 20 μl containing 50 ng genomic DNA, 10 mM Tris-HCl (pH 8.3), 2.5 mM MgCl2, 50 mM KCl, 0.001% gelatin, 0.25% Nonidet P-40, 0.25% Tween-20, 200 μM each of the four deoxynucleotide triphosphates, 0.6 μM of each primer (FW: 5′-AACACTTGGACTCATTCTCACTGG-[G (SUS) or T (UNSUS)]-3′ and RV: 5′-CCTGGATGGAATGTTGACAC-[C (SUS) or T (UNSUS)]-3′), and 0.8 units of Taq polymerase (MBI Fermentas). Reactions were carried out at 94°C for 60 s, 58°C for 60 s and 72°C for 60 s for 35 cycles. Products were analysed on a 1% agarose gel.
Quantification and statistics
Quantification of AP-PCR products was performed using the Labworks 4.0 program (UVP BioImaging Systems, Cambridge, UK) and statistical evaluation was performed by means of an unpaired Student’s t-test.
Results
AP-PCR DNA fingerprint patterns of the APO-SUS and APO-UNSUS rat genomes and epigenomes
In order to identify differences between the genomes and epigenomes of APO-SUS and -UNSUS rats, we performed a comparative analysis of fingerprints of AP-PCR products generated from genomic DNAs of the two rat lines. Initially, genomic DNAs isolated from APO-SUS and -UNSUS (hypo)thalamic tissue and digested with RsaI in combination with the methylation-sensitive restriction enzyme HpaII (CCGG) was analysed using arbitrary primers AP-1, AP-7 or AP-777. These primers were selected from a total set of ten primers because they gave fingerprints with reproducible and discrete products (data not shown). Typical AP-PCR fingerprints obtained with the three selected arbitrary primers are shown in Fig. 1. With each arbitrary primer ∼30 chromosomal fragments were reproducibly amplified. DNAs digested with RsaI and the methylation-insensitive enzyme MspI served as controls to determine whether the observed differences were due to a differential methylation of the CCGG sequence or a genetic polymorphism in this sequence. AP-PCR analysis with primer AP-1 revealed 16 products corresponding to fragments without an HpaII site (“genetic fragments”) and 23 products corresponding to fragments containing an HpaII site (“epigenetic fragments”). Analysis with AP-PCR primer AP-7 showed 17 genetic and 11 epigenetic fragments, and with AP-777 primer 22 genetic and 6 epigenetic fragments.
Fig. 1AP-PCR analysis of genomic DNAs from APO-UNSUS (U) and APO-SUS (S) (hypo)thalamus. AP-PCR was performed with primers AP-1, AP-7 and AP-777 using genomic DNAs digested with RsaI and HpaII (H) or RsaI and MspI (M) as templates. Epigenetic products (methylation-sensitive and thus absent in the MspI lanes) are indicated by closed arrows and genetic products (methylation-insensitive and thus present in the MspI lanes) by open arrows
Genetic variations between the APO-SUS and APO-UNSUS rat genomes
Comparison of the genetic fingerprints generated with primers AP-1, -7 and -777 revealed three reproducible variations between the genomic DNAs from the original (F32) APO-SUS and -UNSUS rats, designated products 1, 2 and 3 (Fig. 2a). Product 1 was less prominent in the APO-SUS than in the APO-UNSUS rats, product 2 was found only in APO-SUS, while the level of product 3 was higher in the APO-SUS than -UNSUS rats. Interestingly, the three products were also present in the replicate (F18) lines and at the same levels, indicating that the replication of the APO-SUS and -UNSUS lines had resulted in a similar genotypic distribution. Next, digestions using RsaI in combination with MspI were used to examine whether the observed differences were due to a genetic or an epigenetic alteration. Following digestion with RsaI and MspI, products 1, 2 and 3 were still found, indicating that the presence of the three AP-PCR products was due to genetic differences (Fig. 2b).
Fig. 2Genetic variations in genomic DNAs from APO-SUS (S) and APO-UNSUS (U) (hypo)thalamus. Rats were from the original (F32) or the replicate (F18) rat lines. (A) Products generated by AP-PCR using primers AP-1 (product 1), AP-7 (product 2) or AP-777 (product 3) on genomic DNAs digested with the methylation-insensitive enzyme RsaI and the methylation-sensitive enzyme HpaII. ND = not determined. (B) Products 1, 2 and 3 generated by AP-PCR on genomic DNAs digested with RsaI and HpaII (H) or with the two methylation-insensitive enzymes RsaI and MspI (M). MspI products served as controls to determine whether products 1, 2 and 3 were due to differential methylation or to a genetic polymorphism. The fact that products 1, 2 and 3 were still present following MspI digestion indicates that they represent products without an HpaII site (“genetic fragments”). (C) Products generated by AP-PCR using a topoisomerase II binding site consensus sequence (5′-GCCTCCTTGCAGGTCTTT-3′) on genomic DNAs digested with the methylation-insensitive enzymes EcoRI (product 4) or MboI (product 5). Arrows indicate increased amounts of the AP-PCR products
We previously discovered that the Aph-1b gene-dosage imbalance between the APO-SUS and -UNSUS rats is the result of a DNA recombination event between the two Aph-1b genes. Furthermore, we identified the region in which the recombination occurred, namely in a region of 1106 nucleotides that is identical between the two genes and encompasses exon 5 (Coolen et al. 2005). In the present study, we decided to examine in detail the site of recombination and found a topoisomerase II binding site (5′-ACCCACCTGCTGGTGTCC-3′) in the DNA region harbouring the recombination site. Topoisomerase II binding sites (with the vertebrate consensus sequence 5′-RNYNNCNNGYNGKTNYNY-3′) (Spitzner and Muller 1988) are known to be hotspots where DNA recombination events occur easily (Craig and Nash 1983). We therefore wondered whether other topoisomerase II binding sites could have led to additional differences between the APO-SUS and -UNSUS rat genomes. Interestingly, using a primer based on the topoisomerase II binding site consensus for PCR analysis of genomic DNAs digested with EcoRI or MboI revealed two differences between the APO-SUS (n = 3) and -UNSUS (n = 2) rat genomes of the original (F32) lines, designated products 4 and 5 (Fig. 2c). Product 4 was present in APO-SUS but not in APO-UNSUS rat genomic DNA. In the replicate APO-SUS and -UNSUS lines (F18), the genomes of two APO-UNSUS rats did also not contain product 4, whereas it was present in two of the four APO-SUS rats tested. Product 5 was present in three of the four APO-UNSUS rats examined (in both the original and the replicate lines), whereas it was not observed in the seven APO-SUS rats tested (Fig. 2c).
Epigenetic variations between the APO-SUS and APO-UNSUS rat genomes
We then wondered whether, besides the five genetic variations, also epigenetic variations would be present between the APO-SUS and -UNSUS rat lines. AP-PCR analysis using primer AP-1 on RsaI- and HpaII-digested genomic DNAs from APO-SUS and -UNSUS (hypo)thalamus revealed one epigenetic variation, designated the E1-product (Fig. 3a). The difference was observed in both the original APO-SUS and -UNSUS rats as well as the replicated lines. An ∼1.4-fold reduced amount of the E1-product was observed in APO-SUS when compared with APO-UNSUS genomic DNAs (n = 12, P < 0.05). Since DNA methylation may be tissue specific, we decided to examine the E1-product in two other brain tissues. The level of the E1-product was ∼2-fold reduced in genomic DNAs from the cerebellum of APO-SUS compared to APO-UNSUS rats (n = 4, P < 0.01), while no difference in the E1-levels was found in the hippocampus of the APO-SUS and -UNSUS rats (n = 4).
Fig. 3Epigenetic variation in genomic DNAs from APO-SUS (S) and APO-UNSUS (U) (hypo)thalamus. Rats were from the original (F32) or the replicate (F18) rat lines. (A) Product E1 generated by AP-PCR using primer AP-1 on genomic DNAs digested with RsaI and HpaII. Arrow indicates a representative example of an AP-PCR product that did not show variable amounts using primer AP-1 (randomly chosen out of 38 products) and was used for normalization. (B) The AP-PCR E1-product generated from genomic DNAs from cerebellum, hippocampus and (hypo)thalamus. All rats were from different nests. The products indicated by the arrow were used for normalisation. Amounts for the epigenetic E1-product were significantly different between the APO-SUS and APO-UNSUS rats in the cerebellum (**P < 0.01; n = 4, plus s.e.m.) and (hypo)thalamus (*P < 0.05; n = 12, plus s.e.m.), but not in the hippocampus (n = 4, plus s.e.m.)
Chromosomal localizations of the genetic and epigenetic variations between the APO-SUS and -UNSUS rat genomes
To identify the locations of the genetic and epigenetic variations within the APO-SUS and -UNSUS rat genomes, AP-PCR fragments 1, 2, 3 and E1 were excised from the gel and each fragment was reamplified with the primer used for the original AP-PCR reaction. DNA sequence analysis of the amplified PCR fragments and database searches with the obtained nucleotide sequences revealed the chromosomal localisations of the four fragments; product 1 was located on chromosome 19q11, ∼3.3 kb upstream of the first exon of the GAIP-interacting protein, C terminus (GIPC) gene; product 2 was part of a repeat sequence located on chromosome 2q34; product 3 was located in the first intron of the myosin 1b gene on chromosome 9q22; the epigenetic variation E1 was located on chromosome 6q31, downstream of the Jun dimerization protein 2 (NP_446346.1) and upstream of the ATF-like basic leucine zipper transcriptional factor B-ATF (SF-HT-activated gene 2; XP_216745.2). Since product 3 was localized within a gene, we decided to analyse this chromosomal region in more detail. Sequence analysis of the DNA region corresponding to AP-PCR fragment 3 and its surrounding region revealed a remarkably high number of genetic variations: 10 of the 1948 base pairs analysed were different between the APO-SUS and -UNSUS genomes (Fig. 4). Comparison of the nucleotide sequences of the APO-SUS and -UNSUS DNA regions with the corresponding database sequence (geneID: 117057) revealed a 100% identity between the database and the APO-UNSUS sequences, indicating that the APO-SUS genome has diverged from the database sequence. Next, more APO-SUS and -UNSUS rats were genotyped for this region. All APO-UNSUS rats tested (n = 5) indeed showed the database sequence, whereas the APO-SUS rats tested (n = 5) all contained the relatively high number of variations in this chromosomal region.
Fig. 4Nucleotide sequence of chromosomal region 9q22, corresponding to product 3, in APO-SUS and APO-UNSUS rats. APO-SUS/-UNSUS genomic variations are indicated between brackets; the first nucleotide represents the APO-UNSUS sequence, the second nucleotide the APO-SUS sequence. The nucleotide sequences corresponding to the annealing sites of arbitrary primer AP-777 are underlined
The newly identified genetic and epigenetic variations, and apomorphine susceptibility in Wistar rats
We wondered whether the molecular-genetic difference between the APO-SUS and -UNSUS rats (the Aph-1b gene-dosage imbalance) would also be present in the Nijmegen outbred population of Wistar rats, or if the imbalance was generated during the breeding of the rat lines. PCR analysis of the Aph-1b locus in the 50 Wistar rats examined revealed in all cases the presence of three copies of the Aph-1b gene, suggesting that the reduction of Aph-1b copies in the APO-SUS rats had been induced during the breeding of the rats.
We next wondered whether the newly identified genetic and epigenetic variations in the APO-SUS and -UNSUS genomes were also induced during the breeding of the two rat lines, or if these variants are already present in Wistar rats. For this purpose, we performed AP-PCR analysis of (hypo)thalamic genomic DNAs from Wistar rats using primers AP-1, -7 and -777. The fingerprints revealed the presence of the APO-SUS as well as the -UNSUS variants of the genetic products 1, 2 and 3 (n = 8) and the epigenetic E1-product (n = 4) in Wistar rats (Fig. 5), indicating that the newly identified variations were not induced during breeding of the APO-SUS and -UNSUS lines, but were already present in the original Wistar population.
Fig. 5AP-PCR analysis (products 1, 2 and E1) and specific PCR (product 3) analysis of genomic DNAs from Wistar rats with low or high apomorphine susceptibility. (A) The presence or absence of products 1, 2 and 3 (genetic differences) was analysed in genomic DNAs from the (hypo)thalamus of Wistar rats with low (<10 gnaws in 45 min) or high (>500 gnaws in 45 min) apomorphine susceptibility. U: APO-UNSUS, S: APO-SUS. Lower panel: the amounts of the three products present in the Wistar rats were compared with the amounts found in the APO-SUS and -UNSUS rats. (■) the genotype of the genetic products 1, 2 or 3 in the Wistar rats with low apomorphine susceptibility is similar to the products 1, 2 and 3 genotype of the APO-UNSUS rats; (□) the genotype in the Wistar rats with low apomorphine susceptibility deviates from the APO-UNSUS genotype;
() the genotype in the Wistar rats with high apomorphine susceptibility is comparable with the genotype in the APO-SUS rat;
() the genotype in the Wistar rats with high apomorphine susceptibility deviates from the genotype in the APO-SUS rats. (B) AP-PCR analysis of the epigenetic E1-product (E1) on genomic DNAs from cerebellum and hippocampus of four Wistar rats. The product indicated by the arrow is an example of a product that did not show variable amounts and was used for normalisation of the E1-product. The amounts of the E1-products generated from the cerebellum or hippocampus were not different between the two Wistar rats with low gnawing scores (<10 gnaws in 45 min) and the two Wistar rats with high gnawing scores (>500 gnaws in 45 min)
To study whether the presence or absence of the genetic AP-PCR products observed in the APO-SUS and -UNSUS rats (products 1, 2 and 3) was linked to the apomorphine susceptibility of Wistar rats, we examined a group of five rats with low susceptibility to apomorphine (<10 gnaws per 45 min) and a highly susceptible group of three rats (>500 gnaws per 45 min). Product 1, which was less abundant in APO-SUS than -UNSUS rats, was not present in the three Wistar rats with high gnawing scores, but was also not found in two of the five Wistar rats with low gnawing scores. Of the eight rats examined, the APO-SUS-specific product 2 was found in two Wistar rats with a high and only one rat with a low degree of apomorphine susceptibility. The DNA region corresponding to product 3 was found twice as the APO-UNSUS sequence in the group consisting of the low-apomorphine-susceptible Wistar rats and once in the group of the high-apomorphine-susceptible rats (Fig. 5a). These results indicate that none of the three products was directly linked to the apomorphine susceptibility of the Wistar rats. However, apomorphine susceptibility may not be the result of only a single genetic variation, but rather of multiple genetic and epigenetic alterations. Interestingly, 67% of the three genetic products present in the Wistar rats with low apomorphine susceptibility were also found in the APO-UNSUS genome, and 78% of the three products in Wistar rats with high apomorphine susceptibility were APO-SUS-specific variants (Fig. 5a), indicating that a combination of the three genetic products may well be linked to apomorphine susceptibility. To study the link between the epigenetic E1-product and apomorphine susceptibility, two Wistar rats with low gnawing scores and two with high gnawing scores were examined for the amount of the E1-product in the cerebellum. Similar amounts of the E1-product as detected in APO-UNSUS rats were found in one Wistar rat with a low and in one Wistar rat with a high gnawing score. The amount of the E1-product detected in APO-SUS rats was found in one Wistar rat with low and in one Wistar rat with high apomorphine susceptibility, indicating that no direct link exists between the E1-product and apomorphine susceptibility. As observed in the APO-SUS and -UNSUS rats, no variation was detected in the level of the E1-product in hippocampal genomic DNAs from the four Wistar rats tested (Fig. 5b).
Discussion
In this study, we investigated the genetic and epigenetic background of the phenotypically well-characterized APO-SUS and -UNSUS rats. Unravelling the molecular basis of this rat model may help in our understanding of complex human neurodevelopmental disorders, since many of the characteristics of the APO-SUS rat line are also observed in schizophrenic patients (Ellenbroek and Cools 2002). We recently identified a gene-dosage imbalance in the Aph-1b locus of the APO-SUS and -UNSUS rats, leading to a reduced expression of the Aph-1b gene in APO-SUS rats and a segregation with a number of behavioural parameters (Coolen et al. 2005). Here, we used AP-PCR analysis as an approach to explore the presence of any other alterations in the two genomes and epigenomes. We indeed identified additional genetic and epigenetic variations, indicating that the Aph-1b locus might not be solely responsible for the observed phenotypes of the two rat lines and suggesting a multi-genetic and -epigenetic origin of the differences observed between the APO-SUS and -UNSUS rats. Importantly, we found similar AP-PCR patterns in both the original and replicate APO-SUS and -UNSUS lines. It is therefore highly unlikely that the observed variations between the APO-SUS and -UNSUS rat genomes were simply due to coincidence.
Detailed analysis of one of the newly identified genetic alterations (in the DNA region corresponding to product 3) revealed in APO-SUS rats a cluster of variations in the myosin 1b gene. Hence, this cluster appears to be a hotspot for genetic instability. Product 1 was located upstream of the GIPC gene encoding a protein interacting with membrane-associated and transmembrane proteins, including the dopamine receptors D2 and D3 (Jeanneteau et al. 2004). The genetic variation in this locus may therefore contribute to the differences in apomorphine susceptibility between the APO-SUS and -UNSUS rat lines. We also identified genetic alterations using a primer corresponding to a topoisomerase II binding site consensus. The rationale for this study was based on our present finding that a topoisomerase II binding site was located at the recombination site in the Aph-1b locus. The two newly identified, topoisomerase II-based genetic variations might point to a more general role for topoisomerase II binding sites in psychopathological mechanisms. We hypothesize that at these sites environmental factors, such as stress during early development, may cause an increase in the incidence of recombination and other mutagenic events, leading to brain dysfunction and affected behavior. Besides the genetic differences, one tissue-specific epigenetic variation was found between the APO-SUS and -UNSUS epigenomes. At present it is not clear what, if any, functional consequence should be attributed to the decrease in the methylation status of this CpG in the (hypo)thalamus and cerebellum, but not in the hippocampus, of APO-SUS relative to APO-UNSUS rats.
Earlier microarray analysis of mRNA expression in the hippocampus of the APO-SUS and -UNSUS rats (∼7000 full-length sequences and ∼1000 EST clusters) revealed that only Aph-1b was differentially expressed (Coolen et al. 2005). The newly identified variations open the possibility that for an explanation of the background of the rat model one has to consider more genes that operate together in a multi-genetic and -epigenetic setting with several susceptibility loci. Thus, based on our present findings, more than one locus may be responsible for the complex phenotype of APO-SUS rats. Recently, it has been established that the contribution of genetic modifiers is also of importance for the outcome of a phenotype, since they can modulate the severity of the affected phenotype and the phenotypic characteristics without having a clear effect on the normal situation (Nadeau 2001; Nadeau and Topol 2006). Hence, as part of the genetic and epigenetic background of the APO-SUS and -UNSUS rat lines, our newly identified variants can be genetic and epigenetic modifiers influencing the phenotypic expression of the model. Insight into the genetic and epigenetic background may provide diagnostic tools, and clues for mechanisms and pathways to explain complex disorders.
Besides the multiple variations in the APO-SUS and -UNSUS rat genomes and epigenomes, Aph-1b will presumably be a major player in the development of the complex APO-SUS phenotype, since it has a broad cellular effect via tissue-specific cleavage of many different substrates (Coolen et al. 2006b) and is functional already during early development (Coolen et al. 2006a). Remarkably, however, the number of Aph-1b gene copies was not linked to apomorphine susceptibility in Wistar rats. Whereas their apomorphine susceptibility varied, all unselected Wistar rats tested harboured three copies of the Aph-1b gene. Therefore, genetic and epigenetic factors other than Aph-1b will likely contribute to the susceptibility for apomorphine. We now indeed found a correlation between the newly identified genetic variations and the apomorphine susceptibility in the Wistar population, confirming our hypothesis that apomorphine susceptibility is caused by a number of genetic and epigenetic factors. A combination of the newly identified variations may have thus initially contributed to the degree of apomorphine susceptibility in the original Wistar rat population. Subsequent apomorphine injections to determine the susceptibility for the drug during the breeding of the APO-SUS and -UNSUS lines may have acted as an environmental stressor, triggering the Aph-1b recombination event at the topoisomerase II binding site only in rats with a high susceptibility for apomorphine. The induced gene-dosage imbalance of the Aph-1b gene, probably in combination with other genetic or epigenetic factors, could then have led to the complex phenotype observed in the APO-SUS rats.
In conclusion, the present findings suggest that psychopathological disturbances may be the result of multiple genetic as well as epigenetic factors. We infer that our newly identified variations are susceptibility loci for schizophrenia-like features in the rat and may give new insights into the genetic and epigenetic background of complex neurodevelopmental disorders.

genetic and epigenetic variations
neurodevelopmental disorders
aph-1b
ap-pcr
apo-sus rat model


Keratinocyte serum-free medium maintains long-term liver gene expression and function in cultured rat hepatocytes by preventing the loss of liver-enriched transcription factors

Freshly isolated hepatocytes rapidly lose their differentiated properties when placed in culture. Therefore, production of a simple culture system for maintaining the phenotype of hepatocytes in culture would greatly facilitate their study. Our aim was to identify conditions that could maintain the differentiated properties of hepatocytes for up to 28 days of culture. Adult rat hepatocytes were isolated and attached in Williams’ medium E containing 10% serum. The medium was changed to either fresh Williams’ medium E or keratinocyte serum-free medium supplemented with dexamethasone, epidermal growth factor and pituitary gland extract. The hepatic phenotype was then analysed using RT-PCR, immunohistochemistry, Western blotting and assays of liver function. Cells cultured in keratinocyte serum-free medium supplemented with dexamethasone, epidermal growth factor and pituitary gland extract maintained their phenotype for 3–4 weeks, based on expression of liver proteins, ureagensis and response to xenobiotics. In contrast, hepatocytes cultured in Williams’ medium E rapidly lost the expression of liver proteins after 3 days. Cells cultured in keratinocyte serum-free medium supplemented with dexamethasone, epidermal growth factor and pituitary gland extract maintained their expression of liver-enriched transcription factors (C/EBPα and β, HNF4α and RXRα) while expression was either lost or reduced in cells cultured in Williams’ medium E. These results suggest that keratinocyte serum-free medium supplemented with dexamethasone, epidermal growth factor and pituitary gland extract can maintain the hepatic phenotype for a prolonged period and that this is probably related to the continued expression of the liver-enriched transcription factors.
1
Introduction
The primary culture of hepatocytes from rodents constitutes an attractive model system for the study of liver function. However, a major limitation to such a model is the rapid and irreversible loss of differentiated hepatic functions in culture. De-differentiation is reflected not only in decreased liver-specific functions, but there is also an alteration of morphology: the cells flatten, depolarize, and lose many of the surface characteristics of normal hepatocytes in vivo. The mechanisms responsible for the loss of differentiated properties probably involve the downregulation of transcription factors involved in liver-specific gene expression (Padgham et al., 1993). The loss of differentiated functions has been attributed to the change in environmental conditions (extracellular matrix, hormonal conditions) following cell isolation (Padgham & Paine, 1993). The loss of the differentiated phenotype is most apparent in the rapid decline of total cytochrome P450s (CYPs) after isolation and in culture, particularly the CYP3A1 isoform (Padgham et al., 1993). Studies of liver function are therefore generally confined to the first few days of culture and this precludes longer-term studies. Conventional approaches to maintaining the differentiated properties of isolated hepatocytes in culture include supplementation of the medium with hormones such as dexamethasone (Dex) (Agius, Chowdhury, & Alberti, 1986; Enat et al., 1984), co-factors such as nicotinamide, pyruvate, DMSO and phenobarbital (LeCluyse, Bullock, Parkinson, & Hochman, 1996; Waxman, Morrissey, Naik, & Jauregui, 1990); the application of extracellular matrix components (Bissell, Arenson, Maher, & Roll, 1987; Gomez-Lechon et al., 1998) and co-culture with non-parenchymal epithelial cell-types (Rogiers & Vercruysse, 1993; Vallette et al., 1998).
In the present study we have introduced a new medium for the long-term culture of differentiated hepatocytes. This is keratinocyte serum-free medium (KSFM). It was tested both alone, and in combination with supplements (dexamethasone, EGF and pituitary gland extract), in comparison with the standard Williams’ medium E, and was used to maintain rat hepatocytes in culture for up to 28 days. We performed immunohistochemical, Western blotting and RT-PCR analysis of the cells under the different culture conditions. We chose hepatic markers which are known to be either rapidly switched off after hepatocyte isolation or are important for detoxification. Expression of a particular gene or protein is normally taken as an indication of intact liver function. However, it is difficult to know simply from expression of a gene or protein that the associated function remains intact. For this reason we also assayed for hepatocyte ureagenesis, glycogen synthesis and response to xenobiotics. Lastly, to gain some mechanistic insight we determined the expression of several liver-enriched transcription factors. Our results suggest that KSFM, in combination with dexamethasone, EGF and pituitary gland extract can maintain the liver phenotype for between 21 and 28 days and it is likely that the ability to do this depends on the sustained expression of liver-enriched transcription factors.
2
Materials and methods
2.1
Materials
Collagenase was obtained from Worthington Biochemical Corporation, dexamethasone, penicillin/streptomycin antibiotics, l-glutamine, Williams’ medium E and phenobarbital were purchased from Sigma Chemical Co. (St. Louis, MO). Fungizone (amphotericin B) and Keratinocyte serum-free medium (KSFM) and supplements, recombinant human epidermal growth factor and bovine pituitary extract at final concentrations of 5 ng/ml and 50 μg/ml, respectively, were obtained from GIBCO™. Sources of other chemicals and media were as described previously (Shen, Slack, & Tosh, 2000; Tosh, Shen, & Slack, 2002).
2.2
Rat hepatocyte isolation and culture
Male Wistar rats (270–330 g) were obtained from the Animal House of the University of Bath. Hepatocytes were isolated by the two-step collagenase perfusion technique as described previously (Tosh, Alberti, & Agius, 1988). Calcium-free EDTA perfusion medium was freshly prepared with the following composition: 0.05% (w/v) KCl (Fisher Scientific) in 10 mM 4-(2-hydroxyethyl)-1-piperazineethanesulfonic acid (HEPES, Sigma) buffer, 5 mM d-glucose (Sigma), 200 μM EDTA (Sigma) and 1/1000 v/v phenol red (Sigma) all in sterile phosphate buffered saline (PBS, pH7.4) The liver was perfused via the portal vein for approximately 10 min at a flow rate of 30–35 ml/min and then replaced with a collagenase-containing perfusion medium [composition: 20 mM HEPES buffer, 5 mM d-glucose, 1 mM CaCl2 (Fisher Scientific), 0.001% (v/v) phenol red (Sigma) all in sterile PBS (pH 7.4) and contained collagenase II (0.33 mg/ml, w/v)]. Approximately 150 ml was delivered to the liver for 10–15 min (with recirculation) at a flow rate of 30–35 ml/min. Following collagenase perfusion, the liver was removed and hepatocytes dissociated using fine forceps. The cells were filtered through a 70 μm filter. Cells were washed three times (50 × g for 2 min) in medium [20 mM HEPES buffer, 5 mM d-glucose, 1 mM CaCl2 (Fisher scientific), 0.001% (v/v) phenol red (Sigma) all in sterile saline at pH 7.4]. Using trypan blue (Sigma), the cell viability was determined to be approximately 85%. Approximately 3–5 × 105 rat hepatocytes were initially placed in 35 mm culture dishes in hepatocyte attachment medium (Williams’ medium E containing 10 U/ml penicillin, 100 μg/ml streptomycin,  2mM l-glutamine, and 10% foetal bovine serum). After 6–8 h, the attachment medium was removed and replaced with either serum-free Williams’ medium E or KSFM medium. Both media contained 10 U/ml penicillin, 100 μg/ml streptomycin, 2 mM l-glutamine, 50 μg/ml gentamicin, 100 ng/ml Fungizone with or without the indicated supplements. The composition with supplements was as follows: Williams’ medium E only (W), Williams’ medium E plus 1 μM Dex (WD), Williams’ medium E plus 5 ng/ml EGF (WE), Williams’ medium E plus 1 μM Dex plus 50 μg/ml pituitary gland extract (WDP), Williams’ medium E plus 1 μM Dex plus 5 ng/ml EGF (WDE), Williams’ medium E plus supplements (both 5 ng/ml EGF and 50 μg/ml pituitary gland extract) (WS) and Williams’ medium E plus supplements (5 ng/ml EGF and 50 μg/ml pituitary gland extract) plus 1 μM Dex (WDS); KSFM (K), KSFM plus 1 μM Dex (KD), KSFM plus 50 μg/ml pituitary gland extract (KP), KSFM plus 5 ng/ml EGF (KE), KSFM plus supplement (KS) and KSFM plus supplement plus 1 μM Dex (KDS).
2.3
Immunostaining and antisera
For immunofluorescence staining, cells cultured on glass coverslips were rinsed with PBS to remove any excess medium and then fixed with 4% PFA for 20–30 min. The cells were washed twice in PBS and kept at 4 °C prior to staining. Cells were immunostained as described previously (Shen et al., 2000). The coverslips were mounted with Gel/Mount aqueous mounting medium (Biomeda, Foster City, CA). The primary and secondary antibodies used for immunofluorescence staining are listed in Table 1. For the detection of connexin 32 protein, the cells were fixed in acetone/methanol (1:1, v/v) then incubated in 1× citrate buffer (Lab Vision Corporation, CA) at 37 °C for 30 min before blocking and staining.
2.4
Western blotting
Protein extracts from cultured rat hepatocytes were prepared by lysing pre-washed cells with lysis buffer [20 mM HEPES (pH 7.6), 150 mM NaCl, 1 mM EDTA, 2 mM dithiothreitol (DTT) and 1% (v/v) Triton X-100] containing a 1/100 dilution of protease inhibitor cocktail (Sigma) for 10 min on ice. The cell lysates were then centrifuged at 13,000 rpm at 4 °C and supernatants were stored at −80 °C until further use. The protein concentration of the supernatant was determined using the Bio-Rad protein assay reagent according to the manufacturer's instructions. Total protein (10–15 μg) was mixed with same volume of 2× sample loading buffer [125 mM Tris–HCl (pH 6.8), 4% (v/v) SDS, 20% (v/v) glycerol, 0.2 mM DTT and 0.02% (v/v) bromophenol blue] and denatured by heating to 100 °C for 5 min. The proteins were then separated in 10% or 15% Criterion™ pre-cast Tris–HCl polyacrylamide gel (Bio-Rad) and transferred onto BioTraceNT® nitrocellulose membrane (Pall Corporation, Pensacola, FL). The membrane was blocked with 5% (v/v) non-fat milk in 0.1% (v/v) PBS–Tween20 (Tween-20, was obtained from Sigma) at 4 °C overnight. The membrane was probed with primary and secondary antibodies (listed in Table 2) at room temperature for 2 and 1 h, respectively. Antibodies were diluted in 3% (v/v) non-fat milk in 0.1% (v/v) PBS–Tween and are listed Table 2. The signals were detected with the ECL™ Western blotting analysis system (Amersham) and developed on Hyperfilm™ (Amersham).
2.5
Qualitative and real-time RT-PCR
Total RNA was extracted from cultured rat hepatocytes using TRI reagent (Sigma, Poole, UK). The measurement of total extracted RNAs and reverse transcription were carried out as described previously (Li, Horb, Tosh, & Slack, 2005). Polymerase chain reactions containing the mixture of the same concentration of cDNA, 1.1X ReddyMix™ PCR Master Mix (ABgene, Surrey, UK) and 50 ng sense/antisense primers (listed in Table 3) were processed in a DNA thermal cycler using the following conditions: denatured at 94 °C for 1 min, amplification at 58 °C for 1 min and elongation at 72 °C for 1 min for indicated cycles. The samples were analysed in 1.2% agarose electrophoresis with 1 kb ladder marker (Invitrogen Life Technologies).
To obtain quantitative results, we performed real-time PCR using the Lightcycler system (Roche Diagnostics). The calibrators were the cDNA reverse transcribed from adult rat liver mRNA. The same amount of cDNA from calibrator and experimental samples were mixed with 1XSYBR@ Green Taq ReadyMix™ (Sigma), 50 ng sense and antisense primers and transferred into Lightcycler capillaries (Roche Diagnostics). The following conditions were used for the amplification of PCR products: denaturation at 95 °C for 30 s; amplification at 58 °C for 5 s followed by 72 °C for 20 s for 40–50 cycles; cool-down at 40 °C for 2 min. The fluorescence signal was detected at the same time point of each cycle. Data were presented as the normalised ratio, which is the target/reference ratio of the sample divided by the target/reference ratio of the calibrator by Lightcycler Relative Quantification software.
2.6
Periodic acid–Schiff's (PAS) staining
PAS staining was performed to detect glycogen. The cells were seeded onto coverslips and cultured for up to 4 weeks in KDS medium. The cells were then incubated with KDS plus 25 mM glucose for 24 h, fixed with 4% PFA and then permeabilised with 1% Triton X-100 at room temperature for 20–25 min. The cells were washed with tap water for 1–2 min and transferred to 1% periodic acid solution for 30 min. Next, the cells were washed with running tap water for 3 min and then incubated in the Schiff's reagent at room temperature for 30 min to develop. After washing in running tap water for 10 min, the slides were mounted in Gel/Mount mounting medium.
2.7
Urea cycle assays
Two colorimetric assays, designed to detect secretion of urea and activity of arginase, were performed on the cultured rat hepatocytes. The urea assay was based on a previously published protocol (Meng, Zhang, & Wu, 2004). Briefly, 100 μl of urea standards (0–50 μg/ml) and culture media collected over 24 h were incubated with 300 μl urease buffered solution (Sigma) at room temperature for 20 min. 600 μl of phenol nitroprusside, 600 μl of alkaline hypochlorite (both from Sigma) and 3 ml distilled water were then added, gently mixed and incubated at room temperature for 30 min. The absorbance of standards and samples were then measured at an optical density (OD) of 630 nm. Urea production was expressed as the amount accumulated in 24 h per culture dish (μg/dish/day).
The arginase assay was based on Corraliza, Campo, Soler, and Modolell (1994). Cultured rat hepatocytes were lysed in 0.1% (v/v) Triton X-100 containing a 1/100 dilution of protease inhibitor cocktail and shaken for 30 min. The lysate was then mixed with same volume of 25 mM Tris–HCl (pH 7)/5 mM MnCl2 and the enzyme was activated by incubation at 56 °C for 10 min. Activated lysate (25 μl) was then incubated with 25 μl of 0.5 M l-arginine at 37 °C for 1 h. The samples and the urea standards (0–500 μg/ml) were then incubated with 400 μl of an acid solution comprised of H2SO4:H3PO4:H2O at a ratio of 1:3:7 and 25 μl 9% (w/v, dissolved in 100% ethanol) iso-nitro-propiophenone (Sigma) at 100 °C for 45 min. The urea production was detected at an OD of 540 nm. For urea assay, the secreted urea was presented as the amount of urea per dish per day. The urea production from the hydrolysis of arginine by arginase was taken as a proportional representation of endogenous arginase activity in the arginase assay experiments. The results were normalised with total cellular protein and shown as the amount of urea produced per mg protein per day (μg urea production/mg total protein/day).
2.8
Image collection
Fluorescent images were collected using a Zeiss LSM 510 confocal microscope and figures compiled using Adobe Photoshop 7.0. For cell counting and PAS staining experiments, numbers of random fields were selected using the 40× objective lens of a Leica DMRB microscope. The cell numbers were visualised by 4,6-diamidino-2-phenylindole (DAPI) staining. Coverslips were incubated with DAPI (500 ng/ml) in PBS for 30 min at room temperature before being mounted on to slides in Gel/Mount mounting medium.
2.9
Statistical analysis
The data were expressed as mean ± standard deviation. Comparison of individual treatments was conducted using Student's t-test or one-way ANOVA analysis with Fisher's LSD pairwise comparison.
3
Results
3.1
Expression of hepatic differentiation markers in rat hepatocytes cultured in Williams’ medium E and KSFM
Despite numerous attempts, it has been very difficult to maintain hepatocytes in a well-differentiated state for more than a few days in culture without significantly changing the culture conditions. We therefore sought to develop a protocol for long-term maintenance of hepatic functions in vitro by the use of a simple medium. Several groups previously used KSFM for culture of oesophageal keratinocytes, corneal epithelial cells and foreskin keratinocytes (Andl et al., 2003; Chen, Chang, Lee, Javier, & Azar, 2002; Vallette et al., 1998). Moreover, Katsura et al. (2002) previously used KSFM medium along with serum to culture adult human hepatocytes (but in the absence of the EGF and PGE supplements). We tested the KSFM medium for maintaining the liver phenotype in primary rat hepatocytes in the absence and presence of EGF and pituitary gland extract. Adult rat hepatocytes isolated by the collagenase perfusion technique were allowed to attach in Williams’ medium E containing 10% serum and maintained for 6–10 h before changing to a serum-free defined Williams’ medium E or KSFM supplemented with Dex and/or EGF and pituitary gland extract. The initial measurements were taken for cultures at 72 h. Albumin expression was maintained in cells cultured in KSFM [either with Dex or EGF/pituitary gland extract (referred to as KD and KS, respectively, see Section 2)] in comparison to cells cultured in WD or WS (Fig. 1A). We then compared expression of other liver-specific markers including serum proteins (α1-antitrypsin, haptoglobin and transferrin), liver-enriched transcription factors (C/EBPα, C/EBPβ, HNF-4α and RXRα), enzymes associated with ammonia detoxification (GS and CPS) and Phase II metabolism (UDP-glucuronosyltransferase, UGT) in WS and KS. It was noted that KS maintained the hepatocyte heterogeneity in terms of populations of perivenous type GS and periportal type CPS-expressing cells (Fig. 1B). The results at 72 h showed that all the liver markers examined were maintained to a greater extent in KSFM-cultured conditions in comparison to Williams’ medium E (Fig. 1B and C).
To test the active component in the supplements, we added EGF and pituitary gland extract separately to the Williams’ medium E (Fig. 1C). Compared to EGF, pituitary gland extract had a greater effect on transferrin expression after 72 h of culture. However, we could not maintain the cells for more than 4–5 days, either with one or both the supplements. This result suggests that the KSFM medium itself must contribute to the maintenance of the hepatic phenotype, and it is not just an effect of supplements.
Vimentin is a cytoskeletal intermediate filament protein the expression of which has been associated with the de-differentiation of hepatocytes (Blaheta et al., 1998). We found that this was expressed in Williams’ medium E cultured isolated rat hepatocytes but was absent in KSFM-cultured cells after a 72-h culture period (Fig. 1D). This suggests that KSFM is preventing de-differentiation as well as maintaining differentiated functions.
Connexin 32 (Cx32) is the predominant gap junction protein expressed in hepatocytes (Kumar & Gilula, 1986; Nicholson et al., 1987). Gap junction intracellular communication plays an important role in regulating cell survival and apoptosis (Krysko, Leybaert, Vandenabeele, & D’Herde, 2005), cell differentiation and proliferation (Cheng et al., 2004) and tumourigenesis (Luebeck, Buchmann, Schneider, Moolgavkar, & Schwarz, 2005). We examined the expression of Cx32 in rat hepatocytes cultured for 72 h in Williams’ medium E or KSFM media. As shown in Fig. 2, gap junction-like arrays of Cx32 were only detected in hepatocytes cultured in KS and KDS (Fig. 2).
3.2
Rat hepatocytes cultured in KDS medium retain their differentiated properties for up to 28 days
Although the results for 3-day cultures with KSFM were encouraging, we wanted to know whether the medium could maintain the hepatic phenotype for longer culture periods. We began by examining the survival of hepatocytes under different culture conditions. Equal numbers of cells were seeded and cultured in six different KSFM culture conditions. The cell number at 24 h of culture was set as 100% and the percentages of hepatocytes at 1, 2 and 3 weeks were determined (Fig. 3A). In all cases there was a progressive loss of cells but the cells cultured in KDS exhibited the best survival rates, and we decided to use this combination for subsequent experiments. In addition, we also analysed the hepatic phenotype in the living cells by two different approaches. The first involved counting the number of DAPI-positive cells that also express liver proteins (Fig. 3B) or by immunostaining for the liver protein transferrin and collecting the corresponding transmitted light image to show the distribution of the cells (Fig. 3C). The results confirmed that more than 90% of the cells expressed liver proteins over 2 or 4 weeks of culture (Fig. 3B and C). Lastly, to determine whether KSFM prevents outgrowth of mesenchymal cells in long-term cultures, we co-stained for vimentin and transferrin after 2 and 4 weeks of KDS culture (Fig. 3C). At both time points we found only occasional vimentin positive cell (see insert in Fig. 3C) suggesting the KDS does indeed prevent outgrowth of mesenchymal cells.
We determined the expression of liver markers in KDS-cultured rat hepatocytes using RT-PCR (Fig. 4A), Western blotting (Fig. 4B) and immunostaining (Fig. 4C and D) and compared the results to fresh liver and freshly isolated hepatocytes (designated ‘pre’ and ‘post’). For RT-PCR, the primers were designed towards messages of detoxification enzymes and enzymes involved in amino acid metabolism. All the transcripts were highly expressed in KDS-cultured rat hepatocytes up to 3 weeks and most were still present after 4 weeks of culture (Fig. 4A). Using Western blotting, hepatic proteins including haptoglobin and the liver-enriched transcription factor hepatocyte nuclear factor 1α (HNF1α) were maintained without diminution for at least 2 weeks (Fig. 4B). Furthermore, we also demonstrated that the expression of several liver proteins was maintained for at least 3 weeks. These include apolipoprotein B (ApoB), transferrin, albumin and Cyp2E1, C/EBPα, RXRα and HNF4α (Fig. 4C and D). In general, the expression of both transcription factors and differentiation products was similar to those in fresh liver.
3.3
Glycogen storage and urea cycle activity is preserved in KSFM-cultured rat hepatocytes
To test whether hepatocytes cultured under different conditions maintained their functional capacity, we examined the potential to store glycogen and perform ureagenesis. We were able to detect glycogen in hepatocytes cultured with KDS for at least 4 weeks of culture (Fig. 5). We also investigated ureagenesis following culture of hepatocytes under KDS conditions. Assays were performed from 24 h up to 28 days of culture using either the detection of secreted urea (Fig. 6B), or by measurement of arginase activity (Fig. 6A). The results showed that both arginase activity and urea synthesis was generally maintained and are comparable to previously published values (Kang, Berthiaume, Nath, & Yarmush, 2004).
3.4
Hepatic metabolising enzymes are induced by phenobarbital treatment in KDS-cultured hepatocytes
Phenobarbital is widely used as an inducer of Phase I and II metabolising enzymes in liver. Following treatment by phenobarbital, the transcription of phase I (cytochrome P450s) and phase II (UGT) detoxification enzymes are activated through the binding of CAR-RXR (CAR, constitutive active receptor; RXR, 9-cis-retinoic acid receptor) heterodimers with the cis-acting element (DR4 motifs, such as NR1 and NR2 domain on the enhancer of Cyp2b10 gene) of target genes (Kakizaki et al., 2003). We initially examined the time course of expression of CAR and PXR (pregnane X receptor, an additional important nuclear receptor accounting for detoxification in hepatocytes) by RT-PCR in KDS-treated cells (Fig. 7A). The levels were well-maintained for 3 weeks and then started to drop off between 3 and 4 weeks of culture.
To examine the effect of phenobarbital on KDS-cultured rat hepatocytes, we analysed the mRNA expression of UGT, Cyp2B12, Cyp3A1 and Cyp7A1 by real-time RT-PCR. Rat hepatocytes were cultured for 7 or 14 days in KDS and then treated with and without phenobarbital for 3 days (total of 10 or 17 days culture). Fig. 7 shows that all the genes, including UGT and cytochrome P450s in the phenobarbital-treated cells, were upregulated 2–10 fold in comparison with control cells. The result demonstrates that hepatocytes cultured with KDS remain responsive to xenobiotics.
4
Discussion
Much effort has been devoted in the past in attempting to develop conditions suitable for the in vitro culture of rodent hepatocytes. Some culture media have been described for sustaining the differentiated state of hepatocytes but they only maintain function in the short term, i.e., 7 days or less (Michalopoulos, Bowen, Mule, & Luo, 2003; Miyazaki et al., 1998). Alternative approaches to the resolving the problem for human liver includes isolating and maintaining embryonic liver cells in culture or adult human hepatocytes (Lazaro et al., 2003; Runge et al., 2000). Our work here shows that KSFM medium with dexamethasone, EGF and pituitary gland extract is able to maintain the differentiated properties of rat hepatocytes for between 21 and 28 days. This statement is based on the sustained expression of liver-specific proteins (including liver-enriched transcription factors), the ability to induce drug metabolising enzymes, store glycogen and produce urea. The main benefit of KSFM medium is that it is relatively simple in comparison with some of the culture conditions reported by other groups. In previous studies, various supplements have been utilised to maintain hepatocytes in culture. These include growth factors (EGF and hepatocyte growth factor) (Michalopoulos et al., 2003; Miyazaki et al., 1998), differentiation-promoting chemicals (nicotinamide and DMSO) (Mitaka, 1998), metal ions (ferrous iron, copper, manganese or zinc) (Kojima et al., 1996), and hormones (e.g. glucocorticoid) (Shelly, Tynan, Schmid, Schutz, & Yeoh, 1989). In contrast, the present medium is relatively simple to prepare it is commercially available and the only supplements required are dexamethasone, EGF and pituitary gland extract. It is also worth noting that extracellular matrices (Rana, Mischoulon, Xie, Bucher, & Farmer, 1994) are not required. We have cultured hepatocytes in KDS media on collagen, fibronectin, matrigel and glass for 2 weeks and then immunostained for albumin. There were no differences showing that the KDS medium alone is sufficient to maintain albumin expression.
Liver-enriched transcription factors such as HNF1α, HNF4α and C/EBPα are important in the control of hepatic differentiation (Costa, Kalinichenko, Holterman, & Wang, 2003). Indeed over-expression of HNF4α by adenoviral infection prolongs the expression of liver-specific genes, cell viability and liver functions such as ammonia metabolism in cultured rat hepatocytes (Naiki, Nagaki, Asano, Kimata, & Moriwaki, 2005). In addition, C/EBPβ has been shown to be a master regulator for the formation of transdifferentiated hepatocytes from pancreatic-derived cells (Kurash, Shen, & Tosh, 2004; Shen et al., 2000; Tosh et al., 2002). When we examined the expression of the liver-enriched transcription factors we found that C/EBPα and β, HNF4α and RXRα expression were reduced within 3 days of culture with Williams’ medium E but were maintained when the cells were cultured in KDS. This observation suggests that loss of liver-enriched transcription factors is probably the cause of the loss of the differentiated hepatic phenotype.
Recently, it was suggested that loss of connexin expression might be used as a prognostic marker for hepatocellular carcinoma (Sheen et al., 2004). The presence of connexin proteins in liver is associated with normal liver function. The induction of Cx32 and Cx26 proteins in DMSO-containing primary mouse hepatocytes cultures was also observed and confirmed the importance of connexin proteins in differentiating hepatocytes (Stoehr & Isom, 2003). In the present study, Cx32 was shown to be expressed in hepatocytes maintained in KDS culture medium for up to 3 weeks (Fig. 4C). The persistent expression of Cx32 may provide further evidence of the efficacy of KDS as a medium for primary hepatocyte culture.
Although the KDS mixture does not constitute a defined medium, because of uncertainty about the precise composition of the pituitary extract, our results do show that KSFM, the EGF and the PGE all have a part to play in the maintenance of differentiated hepatocytes in culture. We note that PGE alone has previously been shown to maintain the differentiation status of mammary carcinoma cells (Kano-Sueoka et al., 1979; Platica et al., 1992).
In summary, we describe a simple, novel system for maintaining rat hepatocytes in culture and show that the differentiated phenotype can be maintained for long periods in culture. This method may provide a standard system for studies on drug metabolism and toxicity testing without the problem of overgrowth of mesenchymal cells. Primary hepatocytes are a better model for the liver in vitro than hepatoma cell lines, but they also provide a cheaper alternative and a more controllable system than the intact animal. The method may also have potential for reducing the number of animals used for this type of work. Finally, we believe that this long-term differentiated culture method could also pave the way for bioengineering applications such as the development of a bioartificial liver.

rt-pcr, reverse transcription polymerase chain reaction