To understand the role of clathrin-mediated endocytosis in the internalization of normal cellular prion protein (PrPc) in neuronal cells, N2a cells were depleted of clathrin by RNA interference. PrPc internalization via the constitutive endocytic pathway in the absence of Cu2+ and the stimulated pathway in the presence of Cu2+ were measured in both control and clathrin-depleted cells. Depletion of clathrin had almost no effect on the internalization of PrPc either in the presence or absence of Cu2+, in contrast to the marked reduction observed in transferrin uptake. By contrast, the internalization of PrPc was inhibited by the raft-disrupting drugs filipin and nystatin, and by the dominant-negative dynamin-1 mutant dynamin-1 K44A, both in the presence and absence of Cu2+. The internalized PrPc was found to colocalize with cargo that traffic in the Arf6 pathway and in large vacuoles in cells expressing the Arf6 dominant-active mutant. These results show that PrPc is internalized in a clathrin-independent pathway that is associated with Arf6.
Normal cellular prion protein (PrPc) is a glycosylphosphatidylinositol (GPI)-anchored glycoprotein that is expressed predominantly in the brain, as well as in the spinal cord and at lower levels in leukocytes and some peripheral tissues (Pauly and Harris, 1998; Prusiner, 1998; Weissmann and Flechsig, 2003). Prion diseases are caused by the conversion of PrPc into a protease-resistant misfolded isoform (PrPsc). The prion diseases include transmissible spongiform encephalopathies such as scrapie in sheep and goats, bovine spongiform encephalopathy, and Creutzfeldt-Jakob disease in humans (Prusiner, 1998). Formation of PrPsc in all of these disorders occurs when nerve cells are exposed to PrPsc, which then converts PrPc to aggregated deposits of PrPsc. In scrapie, it is not yet clear whether a loss of function of PrPc or a gain of function by PrPsc is responsible for the neuronal loss and spongiform changes in the brain. However, the observation that clinical symptoms occur without any obvious scrapie deposits (Collinge et al., 1990; Medori et al., 1992) suggests that it is the loss of normal PrPc function, not the formation of PrPsc deposits, that causes prion disease (Aguzzi et al., 1997).
To understand the pathogenesis of scrapie, it is important to know the mode of PrPc internalization in the cell. The biological role of PrPc internalization is unknown, although it is stimulated by Cu2+ (Pauly and Harris, 1998). Like many GPI-anchored proteins, PrPc is found clustered in the sphingomyelin- and cholesterol-rich domains of the membrane known as lipid rafts (Taylor and Hooper, 2006). Although PrPc is present in rafts, it has been widely accepted that, at least in neuronal cells, PrPc is internalized via clathrin-coated pits. Support for this view has come from multiple studies using different cell models. The clathrin-dependent pathway for PrPc internalization was first proposed by the Harris laboratory using a mouse N2a cell line stably transfected with chicken PrPc (Shyng et al., 1994). Their evidence that PrPc was internalized via clathrin-coated pits came from immunogold labeling of PrPc in clathrin-coated pits, from blocking PrPc internalization by hypertonic sucrose, which disrupts clathrin lattices, and from the detection of PrPc in clathrin-coated-vesicle preparations from chicken brain.
Additional evidence that PrPc is internalized via clathrin-coated pits has come from kinetic studies using primary neurons. In these studies, the Morris laboratory found that the rate of PrPc internalization was similar to that of transferrin, a cargo that is known to be internalized by clathrin-mediated endocytosis (Sunyach et al., 2003). They also found, by immunogold labeling, that endogenous PrPc was localized to clathrin-coated pits in N2a cells. Studies, primarily from the Hooper laboratory, using the human neuronal cell line SHY5Y stably expressing mouse PrPc have also supported the clathrin-dependent internalization of PrPc in the presence of Cu2+ (Taylor et al., 2005). This group found that the Cu2+-stimulated internalization of PrPc was blocked by the drug tyrphostin A23, which also blocks internalization of the transferrin receptor. More recently, the Taylor laboratory showed that depleting cells of the LRP1 receptor, a scavenger receptor that is internalized via clathrin-mediated endocytosis, blocks Cu2+-stimulated internalization of PrPc (Taylor and Hooper, 2007). However, it is not clear that this effect of LRP1 knockdown is due to a decrease in receptor internalization because LRP1 has also been shown to affect the biosynthetic trafficking of PrPc (Parkyn et al., 2008).
Although many studies have indicated that PrPc is internalized via clathrin-mediated endocytosis, several studies have suggested that PrPc is also internalized via cholesterol-enriched raft and/or caveolae-like domains; caveolae are a subset of lipid rafts containing the protein caveolin. Ying et al. found that endogenous PrPc was localized to caveolae in mouse N2a cells (Ying et al., 1992), although other laboratories have not even detected the presence of caveolin in N2a cells (Marella et al., 2002; Shyng et al., 1994). Immunogold labeling also localized PrPc to caveolae in a CHO cell line that stably expresses hamster PrPc (Peters et al., 2003), and endogenous PrPc has also been detected in lipid rafts, as well as in clathrin-coated pits, in N2a cells (Sunyach et al., 2003). In addition, PrPc has been found in detergent-insoluble extracts from N2a cells, which is indicative of its presence in raft and/or caveolae-like domains (Sunyach et al., 2003; Vey et al., 1996). These raft and/or caveolae structures seem to be important for PrPc trafficking, because their disruption by the use of sterol-binding agents, such as filipin and nystatin, inhibits Cu2+-stimulated PrPc internalization in human microglia cells and N2a cells overexpressing mouse PrPc (Marella et al., 2002; Peters et al., 2003). Although these results indicate the presence of PrPc in raft and/or caveolae structures, it has been suggested that PrPc normally occurs in rafts but then exits the rafts to be internalized via clathrin-coated pits (Morris et al., 2006; Sunyach et al., 2003).
In summary, although there is no consensus, it is generally considered that, at least in neuronal cells, PrPc occurs in rafts, but is then internalized via clathrin-mediated endocytosis. To investigate the importance of this latter pathway for the internalization of PrPc, we depleted N2a cells of clathrin using RNA interference and measured the internalization of the endogenous PrPc. Surprisingly, after clathrin depletion, confirmed by marked inhibition of transferrin uptake, we found no significant decrease in internalization of PrPc either in the presence or absence of Cu2+. By contrast, when rafts were disrupted by filipin or nystatin, PrPc internalization was blocked both in the presence and absence of Cu2+, whereas clathrin-mediated uptake of transferrin was unaffected. The internalized PrPc colocalized with cargo that was associated with the clathrin-independent Arf6-associated pathway and was present in large vacuoles in cells expressing the Arf6 dominant-active mutant. These results show that PrPc is internalized in a clathrin-independent pathway that is associated with Arf6.
Effect of clathrin depletion on the internalization of PrPc
It was previously shown that the rates of PrPc and transferrin internalization were the same in primary neurons in the absence of Cu2+ (Sunyach et al., 2003). This suggested that PrPc was internalized via clathrin-mediated endocytosis, the pathway for transferrin internalization. Because N2a cells have been used as a model to study PrPc trafficking and scrapie propagation, we examined whether transferrin and PrPc are internalized at the same rate in N2a cells. After loading the plasma membrane with either Alexa-Fluor-488–transferrin or anti-PrP Fab at 4°C for 30 minutes, cells were incubated for varying times at 37°C both in the presence and absence of Cu2+. The fluorescence intensity of PrPc and transferrin in the cell was measured to determine the rate of internalization. The data, plotted in Fig. 1 as fraction internalized versus time, shows that Cu2+ did not appreciably affect the kinetics of internalization. The major effect of Cu2+ was that it changed the distribution of PrPc in the cell. In the presence of Cu2+, the internal and external pools of PrPc became equal at steady state, which indicates that the in and out rates are equal. Because steady state was reached with a half-life of 5 minutes in the presence of Cu2+, half-lives of 10 minutes were calculated for both internalization and externalization of PrPc. In the absence of Cu2+, the internal pool was only one-quarter the size of the external pool at steady state (compared with 50% in the presence of Cu2+), which is consistent with the stimulatory role of Cu2+ on PrPc internalization (Pauly and Harris, 1998). Because steady state was reached with a half-life of about 10 minutes in the absence of Cu2+, half-lives of 48 and 12 minutes were calculated for the internalization and externalization of PrPc, respectively. As for transferrin, it was internalized at about the same rate in both the presence and absence of Cu2+ (Fig. 1B). Because more than 80% of the transferrin receptor was found in the internal pool, the half-life to reach steady state, 5 minutes, is essentially the half-life of internalization. These results show that, in N2a cells, transferrin is internalized about tenfold faster than PrPc in the absence of Cu2+, unlike in primary sensory neurons in which these rates were similar (Sunyach et al., 2003). In the presence of Cu2+, the in rate of transferrin internalization is only twice that of PrPc in N2a cells, but similar measurements have not been made in primary neurons.
These results suggest that, at least in N2a cells in the absence of Cu2+, transferrin and PrPc are not internalized via the same pathway because their internalization rates differ by about one order of magnitude. Therefore, to directly test whether, like transferrin, PrPc is internalized via clathrin-mediated endocytosis, N2a cells were depleted of clathrin using RNA interference. Knock down of clathrin has been used to identify cargos that are internalized via clathrin-mediated endocytosis while, at the same time, helping to elucidate alternative internalization pathways. N2a cells were depleted of clathrin using two different siRNA oligonucleotides, resulting in a 95% reduction in the clathrin levels of the siRNA-treated cells compared with control cells (supplementary material Fig. S1).
The effect of clathrin depletion on PrPc internalization was first measured in the absence of Cu2+. In these experiments, PrPc and transferrin internalization were measured concomitantly because transferrin internalization is a way to monitor clathrin-mediated endocytosis. In all of our experiments, internalization of PrPc was measured using anti-PrP Fab antibodies, whereas fluorescently labeled transferrin was used to measure transferrin uptake. Fluorescent images of PrPc and transferrin internalization, along with clathrin immunostaining, are shown for control cells in Fig. 2A-C. The clathrin-depleted and control cells were imaged using identical confocal settings to enable direct comparison of fluorescent intensity. Consistent with the western blot analysis, the clathrin-depleted cells (Fig. 2D-F) showed a marked reduction in clathrin staining at the trans-Golgi network and a concomitant decrease in transferrin uptake. These results showed that clathrin-siRNA treatment effectively blocks clathrin-mediated endocytosis. Surprisingly, although transferrin uptake was blocked, the clathrin-depleted cells did not show a significant reduction in PrPc internalization.
In order to quantify the pool of internalized PrPc, the staining protocol was modified to enable us to clearly resolve internal from surface-bound PrPc. The surface bound anti-PrP Fab was stained with a Cy5-conjugated secondary antibody prior to cell permeabilization, whereas the internal pool of PrPc was labeled with a rhodamine-conjugated secondary antibody after permeabilization. Fig. 3A shows immunostaining of PrPc in control and clathrin-depleted cells using this method, along with transferrin uptake. The surface PrPc outlining the cell is now clearly resolved from the internal pool. These images confirm that clathrin depletion caused marked inhibition of transferrin uptake, while not significantly affecting PrPc internalization.
To analyze the effect of clathrin depletion on PrPc and transferrin internalization, more than 150 control and clathrin-depleted cells were randomly imaged. The average intensities of PrPc and transferrin in the recycling endosome were then measured in each cell. In Fig. 3B, the values measured in the clathrin-depleted cells are normalized to the control values. This graph shows that there was no significant difference in the average intensity of PrPc in the recycling endosome between control and clathrin-depleted cells, whereas transferrin uptake was reduced by 85% in clathrin-depleted cells compared with controls. Cells were also depleted using another clathrin-siRNA oligonucleotide (oligonucleotide #2) to ensure that the particular siRNA oligonucleotide used was not having an effect on PrPc internalization. With oligonucleotide #2, not only was the data analyzed using the average intensity of the recycling endosome, the internalization of PrPc and transferrin was also analyzed using MetaMorph software to determine the total internal pools of PrPc and transferrin from confocal z-stack cell images. The results from the three-dimensional analysis were not significantly different from the two-dimensional analysis, i.e. clathrin depletion did not significantly affect the extent of PrPc internalization, but markedly inhibited transferrin uptake (Fig. 3B, lanes 5-8). Therefore, constitutive internalization of PrPc apparently occurs via a clathrin-independent pathway.
Similar experiments were carried out in the presence of Cu2+, which has been shown to stimulate the internalization of PrPc (Pauly and Harris, 1998). Fig. 4A shows the immunostaining of surface and internal pools of PrPc, along with transferrin uptake, in cells that were incubated in the presence of Cu2+. Comparison of PrPc localization in the presence and absence of Cu2+ showed that, in addition to a larger internal pool of PrPc in the presence of Cu2+, the PrPc on the plasma membrane also had a patchier appearance compared with in the absence of Cu2+. Just as we observed in the absence of Cu2+, clathrin depletion of the N2a cells did not significantly affect internalization of PrPc (Fig. 4Ac-f), although, as expected, the clathrin-depleted cells showed a marked reduction in transferrin uptake. Again, we used two different siRNA oligonucleotides to deplete clathrin and two slightly different methods to measure the internalization of proteins. This quantitative analysis, shown in Fig. 4B, confirmed that clathrin depletion did not significantly affect the Cu2+-stimulated PrPc endocytic pathway, although it markedly inhibited transferrin uptake. Therefore, PrPc is internalized via a clathrin-independent pathway both in the presence and absence of Cu2+.
All of the above PrPc-internalization studies examining the effect of clathrin depletion were performed using the D13 anti-PrP Fab. To validate these results further, PrPc internalization was remeasured using a different anti-PrP Fab, one made from the AH6 anti-PrP mAb. This latter antibody recognizes residues 159-174, whereas the D13 Fab recognizes residues 95-105 (Novitskaya et al., 2006). Data obtained with the AH6 Fab confirms that neither constitutive nor Cu2+-stimulated internalization of PrPc was significantly affected by blocking clathrin-mediated endocytosis (Table 1).
Characterization of the alternative pathway for PrPc internalization
To better characterize the clathrin-independent pathway for internalizing PrPc, we examined the effect of disrupting lipid rafts on PrPc internalization. Lipid rafts were disrupted by treating cells with either filipin or nystatin, drugs that disrupt rafts by sequestering cholesterol while only having a limited effect on clathrin-mediated endocytosis (Orlandi and Fishman, 1998; Ricci et al., 2000). When the cells were incubated with nystatin, PrPc internalization was blocked both in the presence and absence of Cu2+ (Fig. 5A). By contrast, transferrin uptake was not significantly affected by nystatin treatment. Quantitative analysis of PrPc and transferrin internalization in nystatin-treated cells showed a marked inhibition of PrPc internalization and only a slight decrease in transferrin endocytosis (Fig. 5B). Similar results were obtained by treating the cells with filipin (Fig. 5B). In agreement with these data, Marella et al. found that, in the presence of Cu2+, nystatin and filipin blocked internalization of PrPc, but did not block transferrin uptake (Marella et al., 2002). Therefore, disruption of raft organization inhibited PrPc internalization both in the presence and absence of Cu2+. In addition to being dependent on rafts, PrPc internalization was found to be dynamin dependent both in the presence and absence of Cu2+. When cells were transfected with GFP constructs of either wild-type dynamin-1 or the dominant-negative mutant of dynamin-1, dynamin-1 (K44A), the latter inhibited internalization of both PrPc and transferrin in the presence and absence of Cu2+ (Table 2, supplementary material Fig. S2A). Consistent with these results, incubation of the cells with the dynamin inhibitor dynasore also inhibited PrPc internalization both in the presence and absence of Cu2+ (Table 2, supplemental material Fig. S2B). Therefore, whether or not Cu2+ is present, the internalization of PrPc in N2a cells is raft- and dynamin-dependent, but clathrin-independent.
A raft-dependent dynamin-dependent pathway would typically suggest that PrPc is internalized via a caveolae-raft internalization pathway. However, the Harris laboratory reported that N2a cells contain trace amounts of caveolin-1 (Shyng et al., 1994) and we confirmed their finding using western blot analysis to show that the level of caveolin-1 in N2a cells was barely detectable compared with the level in HeLa cells (supplemental material Fig. S3A). To confirm that the internalization of PrPc does not occur via caveolae, cells were transiently transfected with GFP-labeled caveolin-1 with a mutation in tyrosine 14 (Y14F). Because phosphorylation of tyrosine 14 in caveolin is essential for caveolae internalization (del Pozo et al., 2005; Orlichenko et al., 2006), mutation of this tyrosine residue inhibits internalization of caveolin-associated rafts. In fact, we found that expression of this mutant caveolin-1 had no effect on PrPc internalization either in the presence or absence of Cu2+ (supplemental material Fig. S3B). By contrast, PrPc internalization was reduced by about 50% when actin was depolymerized by cytochalasin D.
Because nystatin and filipin have been shown to block internalization of cargo that traffic via the Arf6-regulated pathway (Naslavsky et al., 2003), we examined whether PrPc colocalizes with other cargo that traffic via this pathway. After internalizing PrPc, the cells were immunostained to detect CD98, a glycoprotein that traffics via the Arf6-associated pathway (Eyster et al., 2009). Both in the presence and absence of Cu2+, internalized PrPc was found to colocalize with CD98 in vacuolar structures that are frequently found in N2a cells (Fig. 6A). As expected, no internalized transferrin was associated with PrPc in these decorated vacuolar structures (supplemental material Fig. S4). We next determined whether cargo that traffics via the Arf6 pathway and is internalized at the same time as PrPc also colocalizes with PrPc. For this experiment, we followed the internalization of MHC-I, another protein whose trafficking via the Arf6 pathway has been well characterized (Naslavsky et al., 2003). Cells were preloaded with anti-PrP and anti-MHC-I antibodies, followed by incubation at 37°C for 30 minutes. Following staining of these proteins with secondary antibodies, MHC-I and PrPc were found to be colocalized on the same vacuolar structures both in the presence and absence of Cu2+ (Fig. 6B).
To further validate that PrPc trafficking is associated with the Arf6 pathway, N2a cells were transfected with the dominant-positive Arf6 mutant Arf6Q67L. This mutant has been shown to drive the formation of vacuoles that trap cargo that enters cells via a clathrin-independent pathway prior to the cargo merging with the clathrin pathway (Donaldson et al., 2009). If trafficking of PrPc was regulated by Arf6, then PrPc would be found on the large vacuoles that develop in cells expressing Arf6Q67L (Brown et al., 2001). As shown in Fig. 7A, both in the presence and absence of Cu2+, internalized PrPc indeed decorated large vacuoles that co-stained for CD98. Moreover, when antibodies against PrPc and MHC-I were used to simultaneously examine the uptake of these proteins, PrPc and MHC-I were colocalized on large vacuoles both in the presence and absence of Cu2+ (Fig. 7B). PrPc was found localized to these large vacuoles even in cells incubated with the dynamin inhibitor dynasore.
To determine whether other cargos are found on these large vacuolar structures, the internalization of cholera toxin B and transferrin was examined in cells expressing Arf6Q67L. Both in the presence and absence of Cu2+, cholera toxin B was found to be colocalized with CD98 on these large vacuolar structures (Fig. 8). These results indicate that the trafficking of cholera toxin B is associated with the Arf6 pathway in N2a cells; this result has also been reported for the trafficking of cholera toxin B in HeLa cells (Naslavsky et al., 2004). As expected, there was no significant colocalization of internalized transferrin with CD98-immunostained vacuoles (Fig. 8). Therefore, the Arf6Q67L vacuoles trap PrPc and cholera toxin B, but not transferrin. These results show that PrPc is internalized via a clathrin-independent pathway that is associated with Arf6 in N2a cells.
To determine whether clathrin-mediated endocytosis is essential for internalizing PrPc, RNA interference was used to deplete clathrin in N2a cells. This technique has been extensively used to identify the role of clathrin in the internalization of various cargos. Several studies have indicated that PrPc is internalized via clathrin-mediated endocytosis in neuronal cells (Shyng et al., 1994; Sunyach et al., 2003; Taylor et al., 2005). Therefore, we expected that clathrin depletion would block PrPc internalization. Surprisingly, both in the presence and absence of Cu2+, the same amount of PrPc was internalized in cells treated with clathrin siRNA as in control cells. Although blocking the clathrin-dependent pathway did not significantly affect PrPc internalization, disrupting lipid rafts with nystatin or filipin markedly inhibited the internalization of PrPc both in the presence and absence of Cu2+ without significantly affecting transferrin uptake. These results suggest that, at least in N2a cells, both in the presence and absence of Cu2+, endogenous PrPc is internalized via a clathrin-independent pathway.
Our results are in agreement with the study from the Chabry laboratory showing inhibition of PrPc internalization by filipin, which disrupts rafts, but not by chloroperazine, which blocks clathrin-mediated endocytosis (Marella et al., 2002). Conversely, our findings disagree with a number of other studies that suggest that PrPc is internalized via clathrin-mediated endocytosis (Shyng et al., 1994; Taylor et al., 2005; Sunyach et al., 2003). This disagreement might partly be due to differences in cell types, as well as to differences in trafficking of endogenous PrPc and stably expressed PrPc. In addition, instead of using pharmacological agents with ill-defined targets to block clathrin, our study blocked clathrin internalization using siRNA, which specifically inhibits clathrin-mediated endocytosis. Finally, support for the view that PrPc is internalized via clathrin-mediated endocytosis has come from electron-microscopic images showing immunogold-labeled PrPc in clathrin-coated pits (Shyng et al., 1994; Sunyach et al., 2003), but these images do not provide a measure of the extent to which PrPc traffics via this pathway. In fact, electron microscopy of the immunolabeled PrPc found that greater than 80% of the labeled PrPc was in detergent-resistant membranes (Morris et al., 2006).
PrPc belongs to the family of GPI-anchored proteins, which are typically internalized via clathrin-independent pathways including a caveolin-pathway of uptake, flotillin-1 pathway, a Cdc42-regulated pathway and an Arf6-regulated pathway (Donaldson et al., 2009; Glebov et al., 2006; Mayor and Riezman, 2004). Our results using the dominant-active Arf6 mutant, in combination with our results using clathrin siRNA, show that PrPc traffics via a clathrin-independent Arf6-associated pathway. In addition to the presence of PrPc in Arf6Q67L vacuoles, the internalized PrPc colocalized with CD98 and MHC-I, proteins that traffic via the Arf6-regulated pathway (Eyster et al., 2009; Naslavsky et al., 2003). Typically, cargo that traffics via the Arf6 pathway is dynamin independent (Donaldson et al., 2009), but perhaps this is different in neuronal cells, which express both neuronal-specific dynamin-1 as well as dynamin-2.
It is important to recognize that many cargos are internalized via multiple pathways. For example, similar to PrPc in our study, the β-amyloid peptide is internalized in a clathrin-independent, dynamin-dependent and raft-dependent pathway in the absence of apoliproprotein E (Saavedra et al., 2007). By contrast, in the presence of apoliproprotein E, β-amyloid peptide is internalized via clathrin-mediated endocytosis (Kang et al., 2000). Furthermore, even though there is disagreement on whether PrPc is internalized via a clathrin-dependent or -independent pathway, it is important to note that both of these pathways converge either directly or indirectly on the early endosome (Donaldson et al., 2009; Kirkham et al., 2005). It has previously been shown that PrPc first traffics to the early endosome (Magalhaes et al., 2002; Pimpinelli et al., 2005) and then to the late endosome (Peters et al., 2003; Pimpinelli et al., 2005). Ultimately, it is then either degraded in the lysosome or recycled back to the plasma membrane (Borchelt et al., 1992).
Although the simplest way of interpreting our studies that used clathrin siRNA is that the majority of PrPc does not enter the cells via clathrin-mediated endocytosis, it is possible that, in N2a cells, PrPc is internalized via multiple pathways. Perhaps when clathrin is depleted, a clathrin-independent raft pathway is upregulated. Compensatory mechanisms of trafficking were previously observed when a dominant-negative dynamin-1 mutant, dynamin-1 K44A, was expressed in cells (Damke et al., 1994). Therefore, it is possible that a clathrin-independent pathway compensates for the clathrin-mediated endocytosis when clathrin is depleted. As for the inhibition of PrPc internalization that occurs in the presence of clathrin when the raft pathway is disrupted, this might occur if PrPc must be present in intact rafts before it can shift over to clathrin-coated pits to be internalized via clathrin-mediated endocytosis (Taylor and Hooper, 2006). One way to test this rather complicated model would be to isolate the carrier protein that PrPc binds to during internalization. A recent report from the Hooper laboratory suggested that the LRP1 receptor was the carrier for PrPc (Taylor and Hooper, 2007), but a more recent study from the Morris laboratory suggested that the primary role of this receptor was to transport PrPc to the plasma membrane rather than act as a carrier for internalization of PrPc (Parkyn et al., 2008). Isolation of the PrPc carrier protein might finally provide a definitive answer as to how PrPc is internalized in N2a cells and primary neurons.
Materials and Methods
Cell culture and transfection
The N2a mouse neuroblastoma cell line was purchased from ATCC (American Type Culture Collection). N2a cells were grown at 37°C in a humidified incubator under 95% air, 5% CO2 in a MEM (Mediatech, Herndon, VA) supplemented with 10% FBS, 100 IU/ml penicillin, 200 μg/ml streptomycin and nonessential amino acids (Invitrogen, Carlsbad, CA). Cells, cultured for 24 hours in complete medium without antibiotics, were transfected with the following constructs using Lipofectamine 2000 (Invitrogen, Carlsbad, CA): GFP-labeled wild-type dynamin-1 and K44A dynamin-1 (gift from Pietro De Camilli, Yale University, New Haven, CT), Arf6Q67L (gift from Julie Donaldson, NHLBI, NIH, Bethesda, MD), and GFP-labeled caveolin-1 and GFP-labeled mutant Y14F caveolin-1 (gift from Mark A. McNiven, Mayo Clinic, Rochester, MN).
Internalization of PrPc
N2a cells, grown on eight-well-chamber glass slides (Nalge Nunc International, Rochester, NY), were washed twice with phosphate buffer saline and then incubated at 4°C for 30 minutes with 2.5 μg/ml anti-PrP Fab antibody. Transferrin (10 μg/ml), either Alexa-Fluor-488-conjugated transferrin (Invitrogen, Carlsbad, CA) or Cy5-conjugated transferrin (Jackson ImmunoResearch, West Grove, PA), was routinely added to the cells, followed by incubation of the cells for 30 minutes at 37°C prior to fixation. To follow cholera-toxin-B internalization, rhodamine-labeled cholera toxin B (Sigma) was added at 1 μg/ml. In the experiments in the presence of Cu2+, 400 μM CuSO4 was added to the cells when they were incubated at 37°C. In the lipid-raft-disruption experiments, cells were first treated nystatin (50 μg/ml) and filipin (5 μg/ml) (Sigma-Aldrich, St Louis, MO) in complete medium at 37°C for 30 minutes. Cells continued to be incubated with the drugs during the preincubation at 4°C and incubation at 37°C. To distinguish between internal and surface-bound PrPc, fixed cells were first immunostained with Cy5-conjugated secondary anti-Fab antibody and then, following permeabilization, were immunostained with a secondary anti-Fab antibody conjugated to a different fluorophore to detect internalized PrPc. To depolymerize actin, 1 μM cytochlasin D (Sigma) was added to N2a cells for 30 minutes at 37°C. To inhibit dynamin, 10 μM dynasore (Sigma) was added to cells at 37°C for 30 minutes.
The anti-PrP Fab antibody used was either D13 Fab (InPro, San Francisco, CA) or AH6 Fab, made from the AH6 antibody (TSE Resource Center, Berkshire, UK) using the Fab preparation kit (Pierce, Rockford, IL). Cells were immunostained for clathrin with X22 antibody (Affinity BioReagents, Golden, CO), GFP antibody (Abcam) and Alexa-Fluor-488-conjugated CD98 antibody (AbD Serotec, Raleigh, NC). Immunoblots were probed with anti-clathrin-heavy-chain mAb clone 23 (BD Biosciences, San Jose, CA), β-actin (Abcam, Cambridge MA), caveolin-1 (Santa Cruz Biotechnology, Santa Cruz, CA), glycerol 3-phosphate dehydrogenase (Novus Biologicals, Littleton, CO) and major histocompatibility protein class I (MHC-I) (from Natalie Porat-Shoram, NHLBI, NIH, Bethesda, MD). Fluorescent secondary antibodies were from Jackson ImmunoResearch and Invitrogen.
Immunofluorescence microscopy and analysis
Images were obtained using the LSM510 confocal microscope (Zeiss, Thornwood, NY) using Plan-Apochromat 63×/1.4 objective. In comparing control and either siRNA- or drug-treated cells, identical confocal settings were used to enable direct comparison between images. Analysis of the internalized PrPc and transferrin was performed using MetaMorph software (Molecular Devices, Sunnyvale, CA).
RNA interference of clathrin heavy chain was performed using siRNAs (Dharmacon, Chicago, IL) to the following sequence: 5′-UCAGAAGAAUUGCUGCUUAUU-3′ (oligonucleotide #1) and 5′-UAAUCCAAUUCGAAGACCAAUUU-3′ (oligonucleotide #2). Lipofectamine RNAiMAX (Invitrogen) was used to transfect the siRNA oligonucleotides using both the forward and reverse method to transfect cells as described in the manufacturer's transfection protocol. To determine cellular levels of clathrin, cell lysates were run on 4-12% gels (Invitrogen), followed by transferring the proteins to nitrocellulose for immunoblotting. Protein bands were detected using the Odyssey infrared detection system (Li-Cor Bioscience, Lincoln, NE) or chemiluminescent substrate (Pierce, cat. no. 34080) followed by densitometer imaging (ChemiImager, Alpha Innotech).
We thank Julie Donaldson and members of the Donaldson laboratory for helpful discussions.
Supplementary material available online at http://jcs.biologists.org/cgi/content/full/122/22/4062/DC1
- Accepted August 24, 2009.
- © The Company of Biologists Limited 2009