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Protein and Vesicle Trafficking, Cytoskeleton
1Department of Pharmacological and Physiological Science, Saint Louis University School of Medicine, St. Louis, Missouri, and 2Department of Neurobiology, Pharmacology and Physiology, University of Chicago, Chicago, Illinois
Submitted 8 September 2006 ; accepted in final form 1 October 2007
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ABSTRACT |
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 TOP ABSTRACT MATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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50%. Although expression levels of tyrosine hydroxylase, the rate-limiting enzyme in the dopamine synthesis pathway, was unaffected, expression of the vesicular monoamine transporter 1 was reduced by 50%. To evaluate whether NPY and CAs are found within the same vesicles and whether syt I is found localized to each of these NPY- and CA-containing vesicles, we used immunocytochemistry to determine that syt I colocalized with large dense core vesicles, with NPY, and with CAs. Furthermore, both CAs and NPY colocalized with one another and with large dense core vesicles. Electron micrographs show that large dense core vesicles are synthesized and available for release in cells that lack syt I. These results are consistent with syt I regulating differential release of transmitters. neuropeptide Y; catecholamines; adenosine 5'-triphosphate; dopamine; norepinephrine
The existence of such a wide range of differential release mechanisms for sympathetic neurotransmitters suggests that differential regulation of transmitters may occur at the vesicle level by specific proteins. Although many proteins are found on vesicles, one family of proteins includes the synaptotagmins (syt) that are each encoded by different genes. Syt I is well-established as the Ca2+ sensor that triggers evoked release of transmitter from vesicles in neurons and neuroendocrine cells (7, 15, 19), and other members of this family such as syt II, III, VII, and IX may also share a role in regulated release of transmitter (21–24, 32, 47, 61). Syt isoforms are categorized by their ability to bind phospholipids in the presence of Ca2+, their affinity for Ca2+, and whether they are found in neurons and on vesicle membranes or are ubiquitously expressed (for review, see Refs. 3, 55). Subtle differences between each syt isoform provides a large range of possible functions within cells, made even more complex by the ability of many syt isoforms to form homo- and hetero-oligomers that sense Ca2+ with a different affinity (24, 25). In this study, we were interested to determine whether syt I mediates the release of each of the sympathetic transmitters, NE, NPY, and ATP.
We used an immortalized cell line derived from a pheochromocytoma tumor of the rat adrenal gland, the PC12 cell (28). When PC12 cells are treated with NGF, they develop flattened, nonspherical cell bodies and long, branching neurites (28). Similar to neurons, PC12 cells require and express the same secretory machinery to support regulated, Ca2+-dependent vesicle release as neuronal synapses (6, 29, 35, 68). They have two types of vesicles, large dense core vesicles (LDCVs) and small, synaptic-like microvesicles that, after NGF treatment, are redistributed from the cell body to the processes (6, 29). The cells release catecholamines (CAs), dopamine (DA) and NE, as well as the transmitters NPY, ATP, and acetylcholine from vesicles in a Ca2+-dependent manner (6, 29, 35, 68). PC12 cells, like other neuroendocrine and neuronal cells, require the three main proteins that comprise the soluble N-ethylmaleimide-sensitive factor attachment protein receptor (SNARE) complex to release vesicles. The SNARE complex consists of the vesicular protein synaptobrevin 2 (also called VAMP-2) and two plasma membrane proteins, 1) syntaxin 1A and 2) synaptosome-associated protein of 25 kDa (SNAP-25) (4, 5, 56). Additionally, NGF-treated PC12 cells express two syt isoforms in greater abundance, syt I and IX, than two much lesser abundant isoforms, syt III and VII (44). These model secretory cells share many similar requirements for Ca2+-dependent transmitter release as neurons.
We also chose to use a model secretory cell because it is amenable to stable transfection and long-term incorporation of plasmid DNA. We generated a PC12 cell line that continuously expresses a short hairpin RNA (shRNA) designed to specifically silence expression of syt I. This plasmid-based method of RNA interference (RNAi) (33, 57) was accomplished by stably incorporating a plasmid to knockdown syt I in a homogenous population of cells. Establishment and characterization of the stable knockdown of syt I was previously described by us (44). Briefly, syt I expression was abolished, whereas syt IX and SNARE proteins were unaffected. Our previous study also showed that evoked release of CAs was reduced by 50%, but not abolished, using carbon-fiber amperometry and HPLC. The reduction of stimulated CA release in the syt I knockdown cells strongly suggests that although syt I is clearly involved in CA release, it is not the only protein to regulate stimulated release in neuronally differentiated PC12 cells, and another protein likely plays a role in regulated release of CAs.
In the present study, we measured release of two CAs, DA and NE, and also the transmitters NPY and ATP to determine whether the release of each transmitter is regulated similarly by syt I. To understand whether it is likely that vesicles express syt I on CA- and NPY-containing vesicles or whether it is likely that CA and NPY are common to one population of vesicle, we used immunocytochemistry to evaluate colocalization of proteins that serve as markers for LDCVs, CA-containing vesicles, and NPY-containing vesicles. We show that 1) syt I is colocalized with CAs and NPY as well as with LDCVs, 2) CAs are colocalized with NPY, and 3) CAs and NPY are colocalized with LDCVs.
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MATERIALS AND METHODS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Fig. 3B at a dilution of 1:300 (Invitrogen, Carlsbad, CA). TO-PRO-3 iodine, a far red fluorescent nuclei counterstain, was used at a dilution of 1:300 in Figs. 5 and 7 (Molecular Probes). Secondary fluorescent antibodies that were used included Rhodamine Red-X-conjugated AffiniPure donkey anti-rabbit IgG (Jackson ImmunoResearch Laboratories), Zenon rabbit or goat IgG labeling kits with Alexa Fluor 488 (Molecular Probes), and Alexa Fluor 488 goat anti-mouse IgG or Alexa Fluor 568 donkey anti-goat IgG (Molecular Probes).
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CA analysis. Cells were plated to six-well tissue culture dishes and cultured to 70–80% confluency. On the day of the experiment, cells were incubated for 15 min with HBSS at 37°C and shaken gently. HBSS solutions were replaced with either 1 ml of fresh HBSS for basal transmitter release or 50 mM K+-buffered saline that contained, in mM, 50 KCl, 87 NaCl, 1 MgCl2, 5 CaCl2, 10 D-glucose, and 12 HEPES (pH 7.3) for stimulated transmitter release for 5 or 15 min (see RESULTS) at 37°C and shaken gently. The incubation medium was removed from each well and centrifuged at 1,000 g for 4 min. Supernatants were used for measurement of released CAs, and the pellets were combined with the cells remaining in the well for measurement of total CA content. Each well of cells was treated with perchloric acid (0.1 N) for 5 min, scraped, and transferred to the pelleted cellular debris from the first spin. Cells were sonicated for 3 short bursts and centrifuged at 8,500 g for 10 min. Endogenous CAs (DA and NE) in acidified release medium or cell extracts were identified and quantified by HPLC with electrochemical detection as previously published (14, 16). The system consists of a Varian Pro-Star solvent delivery system and a model 9090 autosampler (Varian) coupled to a C18 column and an ESA Coulochem II detector. Separations were performed isocratically using a filtered and degassed mobile phase consisting of 12% methanol, 0.1 M sodium phosphate, 0.2 mM sodium octyl sulfate, and 0.1 mM EDTA, adjusted to pH 2.7 with phosphoric acid. A computer was used to collect and store the chromatograms that were analyzed with Varian Star software.
Purine analysis. Cells and samples were prepared and treated as they were for CA measurements. Purines were analyzed according to a method described previously (39). Chloroacetaldehyde was used to form fluorescent 1,N6-ethenopurine (E-purine) analogs, which can be simultaneously separated from the same sample by reverse-phase HPLC and quantified by fluorescence detection. Chloroacetaldehyde (50 µl; synthesized according to the modified method by Secrist et al., Ref. 52) was added to each cell sample, which was then incubated in a dry bath at 80°C for 40 min. The reaction was stopped by placing the samples on ice. Samples were then analyzed by HPLC fluorometric detection. Separation of purine compounds was achieved on a reverse-phase C18 column. A dual buffer gradient system was used to separate and elute the purines from the column by gradually increasing the concentration of buffer B while decreasing the concentration of buffer A (Varian 9010 solvent delivery system). Buffer A (pH 6.0) consisted of 0.1 M phosphate buffer, and buffer B (pH 6.0) consisted of 75% 0.1 M phosphate buffer and 25% methanol. The fluorescent purine derivatives were detected at an excitation wavelength of 300 nm and an emission wavelength of 420 nm (Varian 9070 Fluorescence Detector). Identification of the purine peaks was achieved by comparison of retention times of purine standards. A computer was used to collect and store the chromatograms that were analyzed with Varian Star software.
NPY analysis. Cells and samples were prepared and treated similarly for basal and stimulated release of NPY as for CAs except cells were stimulated for 15 min to evoke measurable NPY release as previously determined for the optimal time course for stimulated release (13). NPY in the samples was purified with Sep-Pak C18 columns (Peninsula Laboratories) and measured by an enzyme immunoassay kit (Peninsula Laboratories). The 96-well plate was read by a PowerWaveX plate reader (Biotek Instruments; Winooski, VT), and the calculation of sample value was analyzed by KC Junior software (Biotek Instruments).
Electron microscopy. Cells were pelleted by centrifugation and fixed with 3.5% glutaraldehyde in 0.1 mol/l sodium cacodylate buffer (pH 7.25) containing 5% sucrose and 2 mmol/l calcium chloride for 16 h at 4°C. Cell pellets were washed in 0.1 mol/l sodium cacodylate buffer containing 5% sucrose (this and all subsequent steps up to polymerization were at room temperature) and postfixed in 1% osmium tetroxide in 0.1 mol/l sodium cacodylate buffer containing 5% sucrose for 3 h. Cell pellets were then washed twice in distilled water, incubated for 1 h in 2% aqueous uranyl acetate, dehydrated through graded ethanols to 100%, rinsed twice in propylene oxide, and infiltrated with a 1:1 mixture of Polybed resin (Polysciences, Warrington, PA) and propylene oxide for 3 h. The cell pellets were then incubated in fresh Polybed resin for 3 h, transferred to Beem capsules filled with fresh resin, and polymerized overnight at 70°C. Ultrathin sections (0.05 µm) were cut with a diamond knife using a Reichert Ultracut E ultramicrotome (Depew, NY), collected on 200-mesh copper grids, poststained with uranyl acetate and lead citrate, and viewed and photographed with a JEOL (Peabody, MA) 100CX transmission electron microscope.
Immunoblot analysis. Experiments were conducted as described by us (44). Briefly, lysates were prepared from cells grown to confluency, protein lysate concentration was determined, and total lysates were electrophoresed on a 10% NuPAGE Bis-Tris gel (Invitrogen), transferred to polyvinylidene difluoride membrane (Millipore, Billerica, MA), and blocked overnight. The membrane was incubated with primary antibody (1:1,000 dilution, except TH was 1:50,000), followed by a horseradish peroxidase-conjugated secondary antibody used at dilutions that ranged from 1:30,000 to 1:50,000 (Santa Cruz). Immunoreactive bands were detected and visualized with ECL Advance reagent (GE Healthcare, Piscataway, NJ), exposed to X-ray film, and developed. The blots were stripped and reprobed for β-actin (1:1,000 dilution). All immunoblots are representative of three or more independent experiments. Quantitative analysis of the immunoblots was performed with ImageJ (http://rsb.info.nih.gov/ij/) and normalized to arbitrary densitometry units of β-actin.
Immunocytochemistry. Cells were replated to collagen-treated Permanox chamber slides and fixed 2 days later with 2% paraformaldehyde. The cells were permeabilized with 0.1% Triton X-100, blocked with 2 mg/ml BSA, incubated with primary antibody, washed, and then incubated with a secondary antibody. Coverslips were mounted with Vectashield (Vector Laboratories, Burlingame, CA) and sealed. When the Zenon staining kit was used, the above protocol was followed for the initial primary and secondary antibody incubations, but an additional staining step was done with the Zenon-prepared antibody. The primary antibody was incubated with the Zenon reagent, followed by a blocking agent to bind any remaining Zenon reagent as per the instructions (Molecular Probes). The cells were incubated in this mixture for 1 h. Nuclei were stained with TO-PRO-3, and the cells were mounted with Vectashield. Images were acquired with a scanning confocal microscope (MRC 1024; Bio-Rad, Hercules, CA) with a 2x or 3x zoom on a x60 oil immersion lens (NA 1.4). Control experiments were performed without antibodies and with primary and secondary antibody alone to test for autofluorescence and nonspecific binding. In the dual imaging experiments, additional controls were performed with secondary antibody added to the primary antibody that it should not recognize. Results are representative of 3–10 independent experiments from 3 or more passages of cells.
Colocalization analysis. Pairs of fluorescent images (1 for each antibody) were acquired sequentially on the confocal microscope as grayscale and converted to green and red channels. A threshold or background for each image was set by an automatic algorithm without user intervention and subtracted before analysis. A qualitative analysis of antibody colocalization was carried out using ImageJ. Overlap of the red/green images were visualized in merged images as yellow pixels, and areas of overlap were considered colocalized. The extent of colocalization was analyzed using the Manders' coefficient plug-in for ImageJ as described on the web site for the Wright Cell Imaging Facility at University of Toronto (http://www.uhnresearch.ca/facilities/wcif/imagej/colour_analysis.htm#coloc_coeff). Manders' coefficient ranges from 0 to 1, with 0 indicating low colocalization and 1 indicating high colocalization. If there was a significant difference between the means for the control cells and the experimental cells, the mean Manders' coefficient for each antibody pair is given in the text as mean ± SE; if there is no difference, only the Manders' coefficient is given for the control PC12 cells. As a negative control, we experimentally determined the amount of colocalization calculated from two images that are unlikely to have overlap: cells stained with both a nuclei stain and an antibody to syt I, a nonnuclear protein (Fig. 3B). This control allows an estimation of the level of colocalization between two stains that occurs due to the resolution of the microscope, the objectives, and the levels of initial light intensity set by the experimenter for the two channels.
Statistical analysis. All data are displayed as means ± SE. A Student's t-test was used to compare significance, except in the case of comparison between multiple syt I knockdown cell lines when a one-way ANOVA comparison of the means was performed followed by a Bonferroni post-test.
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RESULTS |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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50%. This was confirmed with initial HPLC measurements of both DA and NE (44). In the current study, we extended the HPLC measurements of DA and NE and also measured stimulated NPY release and purine release. NGF-treated cells were stimulated with high K+ (50 mM) for 5 min (purines and CAs) or 15 min (NPY). Basal and stimulated release of both DA and NE were reduced by 51% and 46%, respectively, in syt I knockdown cells compared with control cells (Fig. 1, A and B). Detectable release of NPY from dense core vesicles requires a somewhat longer stimulation than used for release of CAs, perhaps reflecting a requirement for a different amount of Ca2+ influx, and possibly different Ca2+ sensors. Therefore, we stimulated the cells for 15 min rather than 5 min with high K+ to evoke detectable NPY release. Figure 1C shows that stimulated release of NPY from syt I knockdown cells is completely abolished compared with control cells. To be certain that the differences were not due to the length of time required for stimulation with high K+, we measured release of NE in response to 15 min of stimulation with high K+ for comparison to NPY stimulation for 15 min. Basal and stimulated release of NE at 15 min was not different than NE release measured after 5 min of stimulation. Similar to the 5-min stimulation, stimulated release of NE in the syt I knockdown cells was reduced by 55% following 15 min of high K+ stimulation compared with control cells. These results indicate that the length of stimulation time is not responsible for the differences between the abolished NPY release and a 50% reduction in CA release.
To demonstrate that the differences between a partial reduction of CA release and completely abolished NPY release were not due to a random variation in our stable syt I knockdown cell line, we isolated a second cell line that stably expressed the syt I RNAi plasmid as determined by PCR from genomic DNA (methods described in Ref. 44). This second shRNA-syt I cell line expressed syt I at 20% (± 0.07% SE; n = 4), the amount expressed in control cells. Similar to the shRNA-syt I cell line shown in Fig. 1 with abolished expression of syt I, the line that expressed 20% syt I also had abolished NPY release and reduced, but not abolished, fractional release of DA (Table 1). These results indicate that differential release of the transmitters is due to RNAi-induced knockdown of syt I and not to simple clonal variation among the PC12 cell lines. All of the experiments described below were performed with the shRNA-syt I cells that have abolished syt I expression.
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The results shown in Fig. 1 indicate that there is a differential release of transmitter from vesicles and that syt I may be responsible for regulating stimulated release of NPY and purines, but may only be partially responsible for regulating the release of CAs. How this differential release of transmitter is regulated at the individual vesicle is not clear from these results. To determine whether the effects of syt I knockdown are acting specifically on the regulated release of vesicles and not due to off-target effects of RNAi, which results in knockdown of protein(s) other than the intended target, we performed a number of control experiments.
Syt I knockdown cells have reduced levels of the vesicle membrane transporter, but expression of TH is unaltered.
We used immunodetection methods to evaluate whether the reduction in transmitter release is specifically due to a lack of syt I or to an aberration in the synthesis or packaging pathways of the CAs. As a marker for CAs, we used antibodies against TH, the rate-limiting enzyme in the CA synthesis pathway and VMAT1, the vesicular transport protein for packaging CAs. Figure 2A shows that one of the primary transporters responsible for transporting DA into vesicles, VMAT1, is reduced by 50% in knockdown compared with control cells. Figure 2A shows an immunoblot for VMAT1 expressed in lysates made from control, syt I knockdown, and CT cells. The blot was stripped and reprobed for β-actin. Figure 2B shows a plot of the average (n = 4) expression of VMAT1 normalized to arbitrary densitometry units of β-actin. Syt I knockdown cells show a reduction in expression of VMAT1 compared with control cells. If the reduction in VMAT1 results in lower intravesicular concentrations of the CAs, this could explain the reduction of stimulated release of CAs in the syt I knockdown cells, but it cannot explain the differential component of CA release that does not require syt I for release.
To determine whether the synthesis of CAs has been affected by lack of syt I, we tested whether the cells had altered expression of TH, the rate-limiting enzyme that converts tyrosine to L-DOPA. Figure 2C shows a blot for TH and β-actin. Three independent experiments show little or no change in TH expression (data not shown). Immunocytochemistry experiments shown in Fig. 2D confirm that neither cellular expression nor localization of TH was altered in syt I knockdown cells. We also measured the total amount of CA per milligram of cellular protein and found that there were no differences in the total CA content among any of the cell lines (data not shown). Therefore, it appears that CA synthesis is normal in the syt I knockdown cells, although the reduction in VMAT1 may lead to an altered distribution of CA between the cytosol and secretory vesicles. However, it is also possible that the reduction in VMAT1 slows the transport of CAs into vesicles but does not alter the ultimate intravesicular content of CAs.
Neither LDCVs nor NPY are altered in expression or cellular localization in the syt I knockdown cells compared with control cells. PC12 cells have two types of vesicles, LDCVs and small clear vesicles, often equated with synaptic vesicles (called SLMVs). NPY is reported to be packaged only in LDCVs with chromogranins (CgA and CgB; Refs. 37, 51, 54, 67). On the other hand, CAs are thought to be found in both types of vesicles (1, 2, 46, 67; see, however, Ref. 6). Figure 2E shows an immunoblot probed for CgA. Figure 2F shows a plot of the average (n = 4) expression of CgA normalized to arbitrary densitometry units of β-actin. There is no significant difference in CgA expression between cell types.
Because mature NPY is a small peptide (36 amino acids), we are not able to detect NPY on an immunoblot as the resolution of the gel limits the ability to detect a peptide less than 10 kDa. However, NPY can be visualized with an antibody with confocal imaging after cells are prepared for immunocytochemistry. The NPY antibody we used is the accepted antibody used by researchers in the NPY field (personal communication from Dr. William Colmers, University of Alberta; see Refs. 20, 34, 49). To evaluate colocalization of CgA and NPY, we performed single cell immunocytochemistry with antibodies against CgA, a marker for LDCVs, and NPY. We evaluated visually whether the antigens were colocalized or altered compared with control cells. Each image was background subtracted, pseudo-colored red or green, and merged. Figure 3A shows the CgA stain as a red image and the NPY stain as a green image. Control and syt I knockdown cells stained similarly for the two proteins. When the red/green images are merged to show overlap of CgA and NPY, the merged image produces yellow pixels where the images overlap, thus showing that LDCVs and NPY overlap and are thus colocalized (merged panel of Fig. 3A). Manders' coefficient of colocalization was used to estimate the relative correlation of the two images. This coefficient provides a measure of colocalization of the two images, a value between 0–1. A value that is closer to 1 represents a higher level of colocalization, whereas a value closer to 0 represents less colocalization. In the merged panels of Fig. 3A, there is similar amount of overlap between the two proteins. The average Manders' coefficient for control cells was 0.65 ± 0.04 (n = 8), not significantly different than either the CT or the syt I knockdown cells.
As a negative control, we evaluated the level of colocalization of two proteins that are not expected to exhibit colocalization. Figure 3B shows poor colocalization of cytoplasmic syt I and a nuclear stain. In this panel, control PC12 cells (not treated with NGF) were stained as for the CgA/NPY dual antibody experiment in Fig. 3A. The merge of the red/green images shows predominately red syt I localized to the cytoplasm and green stain localized to the nucleus. The merge shows little or no yellow pixels at the interface between the nuclei and the cytoplasm. Manders' coefficient is nearer to 0, being 0.28 for this particular example. On average, the negative control of syt I costained with nuclei gave an average Manders' coefficient of 0.31 ± 0.014 (n = 8). Compared with this negative control, CgA and NPY are found colocalized in the cytoplasm, not in nuclei. These results indicate that NPY colocalizes with LDCVs in syt I knockdown cells similar to the control cells. Therefore, abolished release of stimulated NPY is unlikely due to lack of synthesized NPY or lack of synthesis of LDCVs that contain NPY.
Electron microscopy confirms that LDCVs are synthesized in syt I knockdown cells. A possible explanation for the reduction in VMAT1 is that the syt I knockdown cells have fewer secretory vesicles. To explore this possibility, we prepared cells for electron microscopy to confirm that LDCVs are synthesized and unaltered in their relative distribution with the cells and hence are available for release from syt I knockdown cells compared with control cells. Representative micrographs of control (top) and syt I knockdown cells (bottom) are shown in Fig. 4. The thin section of cells shows a nucleus (N) at the uppermost portion of each figure, numerous mitochondria (mit), and other typical cellular organelles including vesicles. Both types of cells have predominantly LDCVs (closed arrowheads) and fewer SLMVs (open arrowheads). The CT cells do not differ ultrastructurally from either the control or syt I knockdown cells (data not shown). Thus the reduction of stimulated NPY release cannot be attributed to a lack of LDCVs or physically sequestered LDCVs. Both cell types have multiple LDCVs that would be ready, presumably, for release following stimulation.
Syt I is colocalized with CAs, NPY, and LDCVs. The differential effects of syt I knockdown on CA and NPY stimulated release results presented above raise the question whether syt I is expressed on both CA-containing and NPY-containing vesicles. To evaluate colocalization of syt I with 1) CAs, 2) NPY, and 3) LDCVs, we used single cell immunocytochemistry with specific antibodies to VMAT1, the vesicle membrane-bound transporter for monoamines, as a marker for CAs, NPY, and CgA as a marker of LDCVs. We performed similar experiments as described above for colocalization of CgA and NPY.
Confocal imaging of VMAT1 and syt I show that syt I is highly colocalized with CA-containing vesicles. Figure 5 shows VMAT1 as a red image and syt I as a green image (control and CT cells). The cells were stained for nuclei (blue), and the right shows the merged image of all three panels. Manders' coefficient is 0.88 for the control cell and 0.83 for the CT cell. These figures represent the average of the experiments (n = 7 for each cell type) that were not significantly different from one another. Unlike the control cells, syt I knockdown cells have had syt I protein expression knocked down and did not exhibit any colocalization with CgA. Manders' coefficient for this particular example was 0.04. (Note that the reduction in VMAT1 expression shown in the syt I knockdown cells from the immunoblot analysis in Fig. 2 is not readily apparent in Fig. 5 because immunocytochemistry is not quantitative per cell for content).
Similar colocalization experiments were performed to evaluate whether syt I colocalized with NPY and with LDCVs. Images of colocalization for NPY and syt I are shown in Fig. 6A and of CgA and syt I in Fig. 6B. For each set of experiments, only the results from the control images are shown because they are similar to those shown in Fig. 5 for both the CT cells and the syt I knockdown cells. Syt I and NPY are colocalized. On average, Manders' coefficient of colocalization for the control cells was 0.77 ± 0.03 (n = 10) and 0.61 ± 0.04 (n = 7) for the CT cells, values that are significantly different (P < 0.01). Although the coefficients are statistically different, the reason for this difference is not apparent from the images, and the CT value of 0.61 is significantly different than the negative control for no colocalization, 0.31 (Fig. 3B).
Syt I is also colocalized with LDCVs. On average, syt I and CgA have Manders' coefficients of 0.81 ± 0.03 (n = 5) for control cells and 0.75 ± 0.03 (n = 7), values that are not statistically significant. As expected, the syt I knockdown cells do not express syt I and therefore exhibit very little colocalization of proteins similar to the images in Fig. 5. Thus syt I is colocalized with LDCVs, and syt I is found expressed on vesicles that contain both NPY and CAs. The similarity in colocalization of syt I with NPY and CAs is unlikely to explain how release of NPY is abolished in cells that lack syt I yet only affects CA release by 50%. There is not a well-defined small vesicle marker, and so we are not able to rule out the possibility that syt I is expressed on small vesicles or that CAs are also released from small vesicles.
CAs are found colocalized with both NPY and LDCVs. One possible explanation for the abolished release of NPY is that the LDCVs release predominantly NPY and, to lesser extent, CAs. Others have reported that CAs are released from both LDCVs and SLMVs in PC12 cells (40). To determine whether the LDCVs are colocalized with NPY to a greater extent than with CAs, we performed immunocytochemistry with antibodies for NPY and VMAT1, as well as CgA and VMAT1, and compared the results with those of Fig. 3. Control, syt I knockdown, and CT cells stained similarly for each antibody. Because the images were all similar visually, Fig. 7 shows images from only the control cells. Figure 7A shows that CAs are colocalized with NPY. The average Manders' coefficient for the control and CT cells were not significantly different, 0.96 ± 0.01 (n = 7) and 0.93 ± 0.01 (n = 6), respectively. However, both control and CT cells were significantly different (P < 0.05) from the average Manders' coefficient for the syt I knockdown cells even though the average value (0.82 ± 0.03, n = 12) was still quite highly colocalized and very similar visually to the merged panel for the control cell shown in Fig. 7A. Figure 7B shows that LDCVs are colocalized with CAs. On average, control cells had a Manders' coefficient of 0.62 ± 0.02 (n = 12), and neither CT cells nor syt I knockdown cells were significantly different. From these data and those of Fig. 3, NPY and CAs are colocalized in LDCVs to a similar level, 0.65 for CgA/NPY and 0.62 for CgA/VMAT1. NPY and CAs are highly colocalized with values for Manders' coefficient that are as near to 1 as any dual antibody pair we have ever used. Therefore, the colocalization data for NPY and CAs with LDCVs does not readily explain the differential release of NPY over that of the CAs.
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DISCUSSION |
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TOPABSTRACTMATERIALS AND METHODSRESULTSDISCUSSIONREFERENCES |
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Differential release of transmitters from sympathetic neurons is well-established (for review, see Ref. 45). NE, NPY, and ATP are coreleased from the same sympathetic neuron; however, different release mechanisms exist for each transmitter. Evidence to support these differential release mechanisms stem from observations made that prejunctional activation of each receptor type by its own transmitter has different effects on the release of each of the other transmitters (9, 10, 18, 31, 36). Results in PC12 cells (66), adrenal chromaffin cells (12, 26), sympathetic neurons (17, 42), as well as pancreatic β-cells (8) also support the hypothesis of differential regulation of transmitter release. Vesicular release components such as the SNARE proteins and Ca2+ sensors are likely candidates for regulation of differential transmitter release. In support of this hypothesis, our results show that syt I regulates the release of all NPY and purines but is only partially responsible for CA release. Additional experiments with a second knockdown cell line confirmed that differential release is due to specific effects of RNAi reduction of syt I and not to clonal variations of the stable cells.
In a recent study, stable knockdown of both syt I and IX in PC12 cells resulted in completely abolished release of LDCVs, whereas if only one or the other syt isoform was expressed, LDCV release remained (43). Furthermore, a second study using RNAi to knockdown expression of syt I and IX showed that the fusion pore is not opening to release the LDCVs and shifts full fusion events to more kiss-and-run events (69). In contrast, an earlier study with transient RNAi knockdown of syt I in PC12 cells resulted in no change in stimulated NPY release; however, a 50% reduction of syt IX expression resulted in partial reduction of NPY release (21). Our results show that NPY release is completely inhibited by stable knockdown of syt I expression. The stably transfected cells used in our study and that of Lynch and Martin (43) may explain, at least in part, our experimental differences with the study using transiently transfected cells (21) in that stably transfected cells provide a homogenous cell population that removes variability in transfection efficiency and in knockdown with the same plasmid between different stable cell lines (unpublished observations).
In our previous study, we (44) determined that the syt I stable knockdown cells did not show any changes, either reduction or upregulation, of endogenous levels of the other expressed syt isoforms, syt II, III, VII, or IX. Furthermore, when we transiently transfected a plasmid that coded for human syt I that was refractory to the RNAi, the number of amperometry events were rescued to control levels. Technical difficulties with low transfection efficiency and NPY sensitivity for the ELISA detection have not allowed us to show rescue of the NPY release. However, because our (44) previous results showed rescue of CA release with human syt I in individual syt I knockdown cells, it is unlikely that our present NPY release results are due to off-target RNAi effects of expression of other syt isoforms. It is clear from our results that syt I is necessary for NPY release. In support of this, our results not only confirm that NPY is exclusively packaged in LDCVs (37, 54), but is also colocalized with syt I in those vesicles.
Although still controversial, CAs are reported to be packaged in both LDCVs and SLMVs (1, 2, 46, 67). Our release studies indicate that only 50% of the releasable pool of CA is dependent on syt I, even though our marker for CAs, VMAT1, is colocalized with syt I. To account for the 50% of CA release that remains in syt I knockdown cells, one explanation is that CA-containing vesicles are not exclusively dependent on syt I as a Ca2+ sensor for regulated release. Other possible candidates for a Ca2+ sensor include one of the additional syt isoforms endogenously expressed by the cells such as syt IX or one or more of the less abundant isoforms, syt III and VII, or even an entirely different protein that can function as a Ca2+ sensor. However, the fact that NPY, CAs, and syt I are colocalized in our studies suggests an alternative explanation. There are at least two modes of vesicular release: full fusion where the vesicle membrane fuses fully to release the entire contents of the vesicle, and kiss-and-run fusion where a fusion pore opens, allowing only partial release of the vesicle contents. Previously, it has been shown that these two modes of release are dependent on whether syt binds Ca2+ (60). Our data supports a role for syt I in controlling full fusion vesicle release. Therefore, if the decision for a vesicle to release all or none of the dense core content lies, at least in part, with the Ca2+ sensor proteins such as the syt isoforms (58, 60), lack of syt I may not allow LDCVs to undergo full fusion events. LDCVs have a "halo" region that surrounds the core region of the vesicle. Evidence suggests that a proportion of vesicular CA content is located in the halo region of the vesicle (53). Therefore, during kiss-and-run, only the contents of the halo would be released. If, in the absence of syt I, full fusion cannot occur, then NPY, a core component, would not be expected to be released. However, any CAs in the halo region could be released, thus providing a possible explanation for the remaining release of CAs we observe in the absence of syt I.
Our results of partial inhibition of CA release in the absence of syt I agree with our previous findings (44) of CA release measured by amperometry from single cells. In that study, syt I knockdown cells exhibited reduced number of secretion events (44), consistent with reduced stimulated CA release shown in this study. Quantitative kinetic analysis of the events in syt I knockdown cells showed that amplitude of the spike events was reduced to 50% of control cells. In that study, we analyzed the kinetic parameters of the spike events, and they point to two main possibilities. The first is that the open-pore time of the vesicle fusion process is reduced, affecting either the size of the open pore or the length of time that transmitter is free to diffuse from the vesicle. The second possibility is that vesicles are smaller in syt I knockdown cells, resulting in a reduced volume of stored transmitter. Kinetic analysis of amperometry events in syt I knockdown cells is most consistent with the first possibility (44). Furthermore, overexpression of syt I causes the fusion pore to remain open for prolonged durations (59).
Of interest was the finding that VMAT1 expression was reduced in the syt I knockdown cells compared with control cells. A plausible explanation is that syt I may function as a trafficking protein for VMAT1 to vesicle membranes. Very little if any direct evidence has been published to date suggesting that syt I acts as a trafficking protein, although other members of the syt family have been implicated in protein trafficking (30, 48). A second possible explanation for the reduction in VMAT1 expression may be due to an off-target effect of the RNAi, although this seems unlikely as none of the other protein expression levels evaluated (SNARE proteins, syt isoforms, TH, or CgA) in this study or in a previous study (44) were affected by syt I knockdown. Regardless of the reasons as to why VMAT1 is reduced, 50% of the remaining CAs is released upon stimulation in the absence of syt I. Moreover, reduction in expressed VMAT1 did not alter the total CA content in syt I knockdown compared with control cells. Taken together, these results support a role for syt I in the differential regulation of transmitter release.
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ACKNOWLEDGMENTS |
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FOOTNOTES |
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The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.
* W. H. Roden and J. B. Papke contributed equally to this work. 
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