The effect of normal cellular prion protein (PrPC) on abnormal protein aggregation was examined by transfecting huntingtin fragments (Htt) into SN56 neuronal-derived cells depleted of PrPC by RNA interference. PrPC depletion caused an increase in both the number of cells containing granules and the number of apoptotic cells. Consistent with the increase in Htt aggregation, PrPC depletion caused an decrease in proteasome activity and a decrease in the activities of cellular defense enzymes compared with control cells whereas reactive oxygen species (ROS) increased more than threefold. Therefore, PrPC may protect against Htt toxicity in neuronal cells by increasing cellular defense proteins, decreasing ROS and increasing proteasome activity thereby increasing Htt degradation. Depletion of endogenous PrPC in non-neuronal Caco-2 and HT-29 cells did not affect ROS levels or proteasome activity suggesting that only in neuronal cells does PrPC confer protection against Htt toxicity. The protective effect of PrPC was further evident in that overexpression of mouse PrPC in SN56 cells transfected with Htt caused a decrease in both the number of cells with Htt granules and the number of apoptotic cells, whereas there was no effect of PrPC expression in non-neuronal NIH3T3 or CHO cells. Finally, in chronically scrapie (PrPSc)-infected cells, ROS increased more than twofold while proteasome activity was decreased compared to control cells. Although this could be a direct effect of PrPSc, it is also possible that, since PrPC specifically prevents pathological protein aggregation in neuronal cells, partial loss of PrPC itself increases PrPSc aggregation.
The normal cellular prion protein (PrPC) is a glycosylphosphatidylinositol-anchored glycoprotein that is predominantly expressed in the brain (Prusiner, 1998; Weissmann and Flechsig, 2003). In prion diseases, the protease-resistant misfolded scrapie isoform of prion protein (PrPSc) is the causative agent of transmissible spongiform encephalopathies, which are neurodegenerative disorders that include scrapie in sheep and goats, bovine spongiform encephalopathies, chronic wasting disease in deer and elk and Creutzfeldt-Jakob disease in humans (Prusiner, 1998). In all of these disorders, exposure of nerve cells to PrPSc converts PrPC to aggregated deposits of PrPSc. There have been numerous models proposed for the neuronal cell loss and spongiform changes in the brain that occur in scrapie, but it is still not clear whether this pathology is due to a loss of functional PrPC or only to a gain of function by PrPSc. Clinical symptoms can occur without any obvious scrapie deposits (Collinge et al., 1990; Medori et al., 1992), which has led to the suggestion that the loss of normal PrPC function, not formation of PrPSc deposits, causes prion disease (Aguzzi and Weissmann, 1997). Unfortunately, the normal function of PrPC is unknown, although its conservation in many different species suggests that it plays a prominent role in a basic physiological process. It has been reported that PrPC functions in cell survival, signal transduction, cell adhesion, copper-dependent antioxidant activity, and copper uptake and sequestration (Roucou and LeBlanc, 2005). Although PrPC knockout mice are healthy, the brains of these mice were found to have reduced levels of cell defense enzymes activity, such as catalase, and increased levels of oxidative stress markers (Klamt et al., 2001; Brown and Besinger, 1998; Brown et al., 1997b; Sakudo et al., 2005; Wong et al., 2001; Wong et al., 2000; Wong et al., 1999). Similarly, tissue cultures of nerve cells derived from the PrPC knockout mouse are less viable and more susceptible to oxidative damage and toxicity caused by agents such as copper and hydrogen peroxide than cells expressing wild-type PrPC (Brown et al., 1997a; Kuwahara et al., 1999). PrPC was hypothesized to act as an antioxidant (Brown and Besinger, 1998; Wong et al., 1999), but recent studies have established that PrPC has no superoxide dismutase activity either in vivo or in vitro (Hutter et al., 2003; Jones et al., 2005). Since numerous studies suggest that, under stress conditions, PrPC has a neuroprotective effect, this raises the question as to whether the neurodegenerative defects observed in scrapie-infected mice are aggravated by the loss of PrPC, as well as the build up of PrPSc amyloid plaque. In fact, neurons from both PrPC knockout mice and scrapie-infected animals show similar changes in neurophysiological function (Colling et al., 1996; Collinge et al., 1994; Jefferys et al., 1994; Johnston et al., 1997; Manson et al., 1995) and biochemical properties (Keshet et al., 1999; Ovadia et al., 1996). Furthermore, altered neuronal excitability can predispose individuals to neuronal damage and death (Leist and Nicotera, 1998) so it is possible that loss of PrPC function contributes to scrapie pathogenesis in this way. However, contrary to the idea that neuropathology is caused by loss of PrPC function, Collinge and coworkers found that there was no effect on neuronal survival when PrPC was knocked out from a 10-week-old mouse (Mallucci et al., 2002). Moreover, by disrupting the prion gene in a scrapie-infected mouse, they reversed the spongiosis, cognitive defects and neurological disfunction caused by scrapie (Mallucci et al., 2003; Mallucci et al., 2007).
In the present study, we have further examined whether knocking out PrPC contributes to a loss of function under stress conditions by examining the effect of PrPC depletion on protein aggregation. We used RNA interference (RNAi) to deplete endogenous PrPC from neuronal-derived tissue culture cell lines that were also transfected with HttQ103. Our results show that there is an increase in HttQ103 aggregation in PrPC-depleted cells. In addition, we found that PrPC may protect against Htt-induced toxicity possibly by increasing cellular defense enzymes, decreasing reactive oxygen species (ROS) and thereby increasing proteasome activity. Interestingly these effects of PrPC on ROS and proteasome activity are specific for nerve cells and do not occur in non-nerve cells even if these cells normally express PrPC.
Two different oligonucleotide sequences were used to knock down PrPC in mouse neuronal cells. Sequence 1 is within the coding region and sequence 2 is in the 3′ UTR. Fig. 1 shows western blots against PrPC of the cell lysates and lysates that were immunoprecipitated with the anti-prion before and after depleting PrPC from SN56 cells. The immunoprecipitated lysate, which has a much higher concentration of PrPC protein, shows multiple bands on the western blot as a result of the different glycosylated forms of PrPC, which are not visible at lower concentrations. From the quantification of the western blots, both sequences reduced PrPC by more than 90% following 2 days of transfection of SN56 cells with oligonucleotides. Similar levels of PrPC depletion were measured 3 days following transfection with the siRNA oligonucleotides (data not shown). As expected, PrPC levels were not affected by transfection of a scrambled sequence of oligonucleotide 1. Throughout this study, oligonucleotide sequences 1 and 2 produced the same phenotype. However, cells depleted of PrPC with sequence 2 could be rescued by expressing PrPC because unlike sequence 1, this sequence is in the UTR region of the message.
Since PrPC has been reported to be neuroprotective (Kuwahara et al., 1999; Roucou et al., 2005; Roucou et al., 2003), we investigated whether PrPC confers protection against Htt aggregation. Both control and PrPC-depleted SN56 cells were transfected with GFP-Htt constructs. Routinely, the day after transfecting with oligonucleotides, the cells were transfected with the Htt constructs. The phenotype of the cells was analyzed 2 days later or 72 hours after transfection of the siRNA. We used both HttQ25, which normally does not form granules, and HttQ103, which forms granules and is toxic to the cell. As expected, there was no aggregation of HttQ25 either in the presence or absence of PrPC (Fig. 2A). However, compared with cells only transfected with HttQ103 or scramble vector, cells depleted of PrPC with sequence 2 caused a marked increase in the number of cells with granules of HttQ103 (Fig. 2A). This effect could be partially reversed by expressing mouse PrPC.
Quantification of the granules in the SN56 cells (Fig. 2A, open bars) shows that 48 hours after transfection with HttQ103, 60% of the PrPC-depleted cells had HttQ103 granules whereas only 25% of the control cells had granules. This increase in the number of cells with granules was observed using both oligonucleotide 1 and 2. To insure that the observed phenotype was due to depletion of PrPC, cells depleted of PrPC with oligonucleotide 2 were partially rescued by transfecting with a plasmid expressing mouse PrPC. As expected, we could not rescue the phenotype generated with oligonucleotide 1 because it is in the prion coding region and therefore it inhibits expression of the plasmid PrPC along with the endogenous protein (data not shown). Remarkably, the extent of HttQ103 aggregation in the PrPC-depleted cells was similar to that obtained when HttQ103-transfected SN56 cells were treated with the proteasome inhibitor, lactacystin (Fig. 2B, lane 6). Essentially, the same results were obtained with N2a cells (gray bars). Therefore, PrPC depletion caused increase aggregation of HttQ103 in the neuronal cell lines, SN56 and N2a.
To ensure that the difference in the level of aggregation was not due to differences in expression levels of HttQ103, western blot analysis was performed on the cell lysates of the transfected cells. Similar levels of expression of HttQ103 were obtained in lysates from control cells, PrPC-depleted cells, and the PrPC-rescued cells (Fig. 2D). Moreover, PrPC depletion did not significantly affect the expression of Hsp70, although HttQ103 expression did cause a significant increase in Hsp70 levels under all conditions (Fig. 2E).
Given that PrPC depletion increased Htt aggregation, we examined whether depletion affected cell viability. Consistent with the results from the mouse PrPC knockout studies, PrPC depletion had no effect on either cell viability or apoptosis as measured by caspase-3 activity (Fig. 3). There was also no effect of HttQ25 expression on viability and caspase-3 activity in PrPC-depleted SN56 cells. As expected, transfection with HttQ103 alone caused a marked decrease in viability and an increase in caspase-3 activity. Interestingly, in HttQ103-expressing cells, PrPC depletion caused a further decrease in cell viability and increase in caspase-3 activity, which could be partially rescued by expression of mouse PrPC. Therefore, PrPC functions in neuronal cells to reduce HttQ103 aggregation and increase viability of the cells expressing HttQ103.
One possible mechanism for these observed phenotypes is that PrPC depletion causes these effects indirectly by reducing proteasome activity. As shown in Fig. 4, this is indeed the case. PrPC depletion causes a 40% decrease in proteasome activity. The expression of HttQ25 has no effect on proteasome activity, whereas expression of HttQ103 caused a 40% decrease in proteasome activity, in agreement with other studies (Jana et al., 2001; Nishitoh et al., 2002; Rangone et al., 2005). When HttQ103 was transfected in PrPC-depleted cells, there was a further reduction in proteasome activity. Specifically, when HttQ103 was expressed in PrPC-depleted cells, the proteasome activity was reduced to 15% of the control cells. Therefore, both HttQ103 expression and PrPC depletion caused a reduction in proteasome activity and the two effects appear additive. The marked reduction in proteasome activity in PrPC-depleted cells probably caused the marked increase in HttQ103 aggregation, similar to that observed when proteasome activity was inhibited with lactacystin.
Since oxidative stress causes a reduction in proteasome activity (Obin et al., 1998; Reinheckel et al., 2000), we measured whether PrPC depletion causes an increase in ROS levels. First, using fluorescence imaging we determined whether the ROS level was higher in PrPC-depleted cells. SiRNA transfections were performed using oligonucleotides conjugated to a fluorophore to enable us to visualize the transfected cells (Fig. 5Aa,c). Compared with transfection with scramble oligonucleotide, the ROS fluorescence intensity was much greater in cells transfected with oligonucleotide 1 than with the scramble oligonucleotide (Fig. 5Ab,d). The ROS fluorescence intensity of cells transfected with the scramble vector was not significantly different from that of the non-transfected cells (see cells with asterisks). Quantification of the ROS levels in the SN56 cells by FACs analysis showed that PrPC depletion caused more than a threefold increase in ROS levels compared with control cells (Fig. 5B). Expression of HttQ103 also caused about a fourfold increase in ROS, in agreement with previous studies (Solans et al., 2006; Wyttenbach et al., 2002). PrPC-depleted cells transfected with HttQ103 showed a sevenfold increase in ROS levels, so again, the effects of PrPC depletion and HttQ103 appears additive.
Table 1 shows that the increase in ROS in PrPC-depleted cells was due to a reduction in antioxidant enzyme activities. Compared to control cells, PrPC-depleted cells showed a marked reduction in the activities of SOD, catalase and glutathione reductase. This reduction was partially rescued when the cells were transfected with mouse PrPC vector. Therefore, PrPC depletion caused a decrease in antioxidant activity, which in turn increased ROS levels thus causing decreased proteasome activity.
To determine whether PrPC functions to protect other cell types that endogenously express PrPC, Caco-2 and HT-29 cells, two human colonic adenocarcinoma cell lines, were depleted of PrPC. As shown in Fig. 6, these cell lines endogenously express PrPC, with HT-29 cells expressing much higher levels of PrPC than the Caco-2 cells (Garmy et al., 2006). By using siRNA oligomers made against human PrPC, we achieved at least a 90% reduction of PrPC in both Caco-2 and HT-29 cells (Fig. 6A). In contrast to neuronal cells, depletion of PrPC from both intestinal cell lines did not significantly affect either proteasome activity or ROS levels (Fig. 6B,C). These results suggest that PrPC confers protection only on neuronal cells.
The protective effect of PrPC in neuronal cells was further evident when mouse PrPC was overexpressed in SN56 cells. Overexpression of PrPC caused a reduction in the percentage of cells with HttQ103 granules and an increase in proteasome activity (Fig. 7). Specifically, when mouse PrPC was overexpressed in SN56 cells, HttQ103 granules decreased from 20% to 10%. Similarly, in SN56 cells overexpressing PrPC showed that HttQ103 only caused a 10% decrease in proteasome activity compared to the 40% decrease that occurred in SN56 cells just transfected with HttQ103. Consistent with the lack of protection conferred by PrPC on non-neuronal Caco-2 and HT-29 cells, there was no effect of PrPC expression on the percentage of cells with either HttQ103 granules or proteasome activity in the non-neuronal cell lines NIH3T3 and CHO. Therefore, consistent with our finding that PrPC only confers protection on neuronal cells, expression of PrPC is not protective against Htt-induced toxicity in non-neuronal cells that do not express endogenous PrPC.
Finally we examined whether the presence of the scrapie form of prion, PrPSc affected the aggregation properties of HttQ103. As shown in Fig. 8A, comparison of uninfected and scrapie-infected SN56 cells (ScSN56) showed that 40% of the infected cells had HttQ103 granules compared to 25% of the infected cells. ScSN56 and Sn56 cells had similar viability, probably due to the low levels of scrapie in most of the ScSN56 cells. However, HttQ103 had a different effect on the viability of SN56 and ScSN56 cells. Expression of HttQ103 caused a 40% reduction in the viability of the SN56 cells whereas it caused a 60% reduction in the viability of the ScSN56 cells. Although it had no effect on viability, PrPSc alone caused an increase in ROS and a corresponding decrease in proteasome activity, and HttQ103 caused further changes. Cells with both PrPSc and HttQ103 showed a sixfold increase in ROS activity compared to a twofold increase with PrPSc alone and a fourfold increase with HttQ103 alone (Fig. 8C). Similarly, cells with both PrPSc and HttQ103 showed a 60% decrease in proteasome activity compared to a 25% decrease with PrPSc alone and a 40% decrease in proteasome activity with HttQ103 alone (Fig. 8D). These data show that PrPSc further reduces proteasome activity beyond the reduction caused by HttQ103 expression alone, an effect that could explain the increase in the percentage of cells with HttQ103 granules and the decrease in cell viability in ScSN56 cells expressing HttQ103. PrPSc could cause these effects directly by contributing to the total amount of aggregated protein in the cell, or indirectly by decreasing the amount of active PrPC, or both.
A long-standing question in the prion field is whether the loss of PrPC from scrapie-infected nerve cells contributes to the neuropathology of the disease. To investigate this question, we examined whether PrPC depletion affects Htt aggregation. Our results showed that PrPC depletion caused a marked increase in HttQ103 aggregation in both N2A and SN56 neuronal cell lines. The increase in the fraction of cells with Htt granules that occurred after PrPC depletion was similar to the increase that occurred after treatment of the cells with the proteasome inhibitor, lactacystin. Consistent with this observation, the proteasome activity of PrPC-depleted cells expressing HttQ103 was only 15% of that in control cells whereas it was 60% of the control activity in the absence of HttQ103 expression. Thus when the cells are stressed, there is a further decrease in proteasome function in PrPC-depleted neuronal cells.
Depletion of PrPC from neuronal cells also caused a reduction in the activity of antioxidant enzymes. However, despite this reduction in antioxidant enzymes and proteasome activity, there is no obvious phenotype caused by PrPC depletion in the absence of stress. Expression of HttQ25, had no effect on the PrPC-depleted cells. However, when the cells were stressed by expression of HttQ103, the PrPC-depleted cells showed a significant loss of viability and a marked increase in ROS levels. Our results are in agreement with the study of Klamt et al. (Klamt et al., 2001) in which an imbalance in antioxidant defense was found in PrPC-knockout mice. Specifically, oxidative damage to lipids and proteins was much higher in the knockout mice, and the activities of SOD and catalase were reduced.
Interestingly, we found that scrapie-infected SN56 cells had properties similar to PrPC-depleted cells. The scrapie infected SN56 cells showed increased HttQ103 aggregation, decreased proteasome activity, and increased ROS levels. In agreement with our results, scrapie-infected hypothalamic neuronal GT1 cells displayed a higher sensitivity to oxidative stress than non-infected cells, as well as a decrease in viability when subjected to stress (Milhavet et al., 2000). An increase in ROS levels was also found in scrapie-infected N2a cells (Fernaeus et al., 2005). It is not clear whether these effects of scrapie infection are due to the scrapie aggregation itself or whether it is also due to a reduction in the level of PrPC. Recent results from the Collinge laboratory showed that disruption of the prion gene in scrapie-infected mice reversed any morphological, neurological or behavioral defects due to scrapie infection (Malluci et al., 2002; Malluci et al., 2007). This shows that scrapie pathology can be reversed by removing PrPC from the cells. However, it is still possible that when scrapie aggregates are present, their effects are worsened by the absence of normal PrPC function.
Although we found that PrPC was protective in neuronal cells, it did not confer protection on non-neuronal cells. Depleting PrPC from two human epithelial cell lines, Caco-2 and HT-29, which like neuronal cells express PrPC endogenously, had no effect on proteasome activity and ROS levels. Furthermore, overexpression of PrPC reduced Htt aggregation in neuronal cells, but had no effect in either HeLa or CHO cells. Other labs have found that expressing PrPC in breast carcinoma MCF-7 cells inhibited the proapoptotic Bax conformational change (Roucou et al., 2005) and necrosis factor alpha-induced cell death (Diarra-Mehrpour et al., 2004), but there is no evidence that PrPC expression affected either ROS levels or proteasome activity in these cells.
The neuro-specific protective effect of PrPC suggests that the signaling pathway activated by PrPC only occurs in neurons. Many proteins have been reported to bind to PrPC, including Sti1, N-CAM, mNOS, APLP1, BL-2 and synapsin (Sakudo et al., 2006) and could be involved in the neuro-specific signaling pathway even if they are not only expressed in nerves. In addition, in a recent model proposed by the Harris laboratory to explain the toxic effects of the truncated PrPC protein (Δ105-125) on mouse viability, they suggested that there is a receptor on the outer surface of nerve cells that normally binds intact PrPC (Li et al., 2007). This putative receptor could be involved in the neuro-specific signaling pathway activated by PrPC.
Whatever the nature of the neuro-specific receptor that interacts with PrPC, it is clear that nerve cells respond to signaling triggered by PrPC by increasing cellular defense enzymes. Several pathways implicated in PrPC signaling are consistent with the increase in antioxidant enzymatic activities. A recent study showed that there is a reduction in AKT signaling in PrPC knockout mice compared to control mice (Weise et al., 2006). Similarly, attachment of PrPC-fusion proteins to monocytes caused an increase in AKT and ERK1 and ERK2 signaling (Krebs et al., 2006). Consistent with these observations there is an increase in phosphatidylinositol 3-kinase signaling in PrPC-expressing N2a cells (Vassallo et al., 2005). These signaling pathways promote cell survival and are perhaps responsible for the neuroprotective effect of PrPC on cell signaling. Ultimately, these neuroprotective pathways are not only regulated via phosphorylation but also by activation of the transcription factor, nuclear factor-κB, which is a central regulator of immunity, inflammation and cell survival.
A diagram of the activation of PrPC that we observed in neuronal cells is shown in Fig. 9. In this model, the deleterious effect of Htt aggregation caused by increasing ROS activity is mitigated by the action of PrPC, which reduces ROS. This in turn increases proteasome activity and reduces Htt aggregation. Interestingly, in a transgenic mouse model of amyotrophic lateral sclerosis, the PrPC protein was specifically repressed when the G85R SOD mutant was overexpressed, but overexpression of wild-type SOD had no effect on PrPC (Dupuis et al., 2002). This suggests that there may be a feedback mechanism that actually downregulates PrPC when cells are under stress, which in turn exacerbates the stress on the cells. There has been no parallel study in mouse models of Huntington disease to determine whether PrPC levels are reduced in animals overexpressing Htt with expanded polyglutamine repeats.
In conclusion, PrPC provides protection against protein aggregation and this protection is neuronal specific. It may be that neurons are particularly sensitive to damage from aggregated proteins and therefore have a specific regulatory pathway that protects against this damage. By maintaining ROS levels, PrPC protects the cell from a reduction in proteasome activity, thereby helping to prevent protein aggregation. As decreased proteasome activity has been implicated in several neurodegenerative disorders and PrPC specifically increases proteasome activity in neuronal cells, it will be of interest in the future to investigate the protective role that PrPC plays in neurodegenerative diseases caused by protein aggregation.
Materials and Methods
The SN56 cells were a generous gift from Bruce Wainer (Department of Pathology, Emory University School of Medicine, Atlanta, GA). The chronically infected ScSN56 cell line infected with the Chandler strain of scrapie was a generous gift from Byron Caughey (RML, Hamilton, MT). SN56 and ScSN56 cells were cultured as described previously (Baron et al., 2006). N2A (mouse neuroblastama cell line), NIH-3T3 (mouse fibroblast cell line) and Caco-2 and HT-29 (human intestinal cell lines) were obtained from the American Type Culture Collection (Manassas, VA). Cells were maintained in Dulbecco's modified Eagle's medium (DMEM), 10% fetal bovine serum, 2 mM glutamine, and 1% penicillin-streptomycin in 75 cm2 culture bottles in a 5% CO2 atmosphere at 37°C. CHO cells were cultured and seeded as described previously (Yim et al., 2005).
The following antibodies were used: anti-GFP polyclonal antibody (Abcam, Cambridge, UK), anti-Hsp70 monoclonal antibody (BD Transduction Laboratory, San Josa, CA), anti-β-actin polyclonal antibody (Abcam), anti-prion monoclonal antibodies D13 (InPro Corp., San Francisco, CA) and SAF70 (Cayman, Ann Arbor, MI). All secondary antibodies were from Jackson ImmunoResearch Laboratory (West Grove, PA).
RNAi, western blot, immunoprecipitation and filter retardation assays
The following oligonucleotides used in RNAi experiments were from Dharmacon (Chicago, IL): mouse prion 1 (nucleotides 299-320), mouse prion 2 (nucleotides 1278-1396), human prion 1(nucleotides 502-522) and human prion 2 (nucleotides 1034-1054), scrambled vectors of mouse and human prion sequence 1. Identical siRNA oligonucleotides were made that were conjugated to Dye547-3′ (Dharmacon, Chicago, IL) to indicate siRNA-transfected cells. These RNA olignucleotides were transiently transfected into SN56 and Caco-2 cell using Lipofectamine 2000 reagent (Invitrogen, San Diego, CA). Control cells were incubated with Lipofectamine under identical conditions as in the RNAi experiments.
To quantify protein, lysates were run on SDS-PAGE gels (Invitrogen) and then western blot analysis was performed. PrPC was detected by immunoblotting using SAF-70 anti-prion antibody, Htt was detected using an anti-GFP antibody, and Hsp70 was detected using an anti-Hsp70 antibody. The protein bands were detected using chemiluminescent substrate (Pierce, Rockford, IL) and analyzed using the ChemiImager densitometer (Alpha Innotech Corp., San Leandro, CA). The linear range of the western blot analysis was determined by loading from 5 μg to 200 μg of cell lysates. Quantification of the western blots established linearity over a 10-fold range of protein concentration. PrPC was immunoprecipitated by mixing the cell lysate (500 μg) with 4 μg of D13 anti-prion antibody followed by the addition of 50 μl of protein A-Sepharose 4 Fast Flow (GE Health Science, Piscataway, NJ). After incubating at 4°C, the Sepharose beads were collected by centrifugation at 14,000 g, washed with PBS and resuspended in 50 μl of 2× Laemmli sample buffer. Protein concentration of the lysates was measured using the Bradford reagent (Bio-Rad, Hercules, CA). For the filter retardation assay, 20 μg of denatured protein samples were filtered through a 0.2 μm cellulose acetate membrane (Adventec MFS, Dublin, CA) using a dot-blot filtration unit (Bio-Rad).
Plasmids and transfection
The Htt polyglutamine constructs (HttQ25 and HttQ103) containing glutamine repeat expansion had an EGFP-tag on its C-terminal (Zeng et al., 2004). A mouse prion gene with the hamster epitope tag (a gift from S. Priola, Rocky Mountain Lab, MT) and a hamster prion gene (a gift from D. Ramanujan Hegde, NIH, Bethesda, MD) was subcloned using HindIII and XhoI restriction sites into pCDNA4 vector (Invitrogen). Cells were seeded and transfected with Htt plasmids as described previously (Zeng et al., 2004). At 48 hours after transfection, cells on coverslips were fixed with 4% paraformaldehyde and 3 μg/ml 4′,6-diamidino-2-phenylindole (Invitrogen) in 1× PBS at room temperature for 20 minutes and mounted.
Microscopy and immunostaining
Cells grown on two-well chamber slides (Labtek, NY) were imaged with a 40×, 1.4 NA objective using a Zeiss LSM 510 confocal microscope (Jena, Germany). The argon and helium lasers were used to excite at 488 nm and 543nm, respectively. Apoptosis was measured by fixing and staining the cells with cleaved caspase-3 antibody (Cell Signaling Technology, Danver, MA). Cells were counted as aggregate positive if one or several granules were visible within a cell. GFP-positive cells (500 per experiment) were counted in multiple random visual fields on each slide.
Measurement of proteasome activity, antioxidant activities, reactive oxygen species (ROS) and cell toxicity
Proteasome activity was measured from the fluorescence intensity of amino-methyl coumarin (AMC)-conjugated to the chymotrypsin peptide substrate LLVY, using a commercial activity assay kit (Chemicon, Temecula, CA). Cleavage products were measured using the Spectra max Gemini fluorescence plate reader (Molecular Devices Co., Sunyvale, CA). The activities of superoxide dismutase (SOD; Chemicon), catalase (Invitrogen) and glutathione reductase (Cayman) were measured in the cell homogenates. To measure ROS levels, cells were stained with DCF-DA or red CC-1 (Invitrogen). For live cells, FACS analysis was performed on the FACS Calibur instrument (Becton Dickinson, Franklin Lakes, NJ) and for fixed cells, images were obtained on the Zeiss LSM 510 confocal microscope. Cell viability was assessed using a 3-[4,5-dimethylthiazol-2-yl]-2,5-diphenyltetrazolium bromide (MTT)-based colorimetric assay kit (Invitrogen).
All data are an average of at least three independent experiments. Student's t-test was used to assess the statistical significance of differences.
- Accepted May 17, 2007.
- © The Company of Biologists Limited 2007