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Endocrinology, doi:10.1210/en.2008-0096
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Endocrinology Vol. 149, No. 10 4948-4957
Copyright © 2008 by The Endocrine Society

Prolactin Secretion Sites Contain Syntaxin-1 and Differ from Ganglioside Monosialic Acid Rafts in Rat Lactotrophs

Paula P. Gonçalves1, Matjaz Stenovec1, Helena H. Chowdhury, Sonja Grilc, Marko Kreft and Robert Zorec

Centro de Estudos do Ambiente e do Mar (P.P.G.), Departamento de Biologia, Universidade de Aveiro, 3810-193 Aveiro, Portugal; and Celica Biomedical Center (M.S., H.H.C., S.G., M.K., R.Z.), Technology Park Ljubljana and Laboratory of Neuroendocrinology-Molecular Cell Physiology (M.S., H.H.C., S.G., M.K., R.Z.), Faculty of Medicine, University of Ljubljana, 1000 Ljubljana, Slovenia

Address all correspondence and requests for reprints to: Robert Zorec, Ph.D., Professor, Member Academia Europaea and The Slovenian Academy of Sciences and Arts, Laboratory of Neuroendocrinology-Molecular Cell Physiology, Institute of Pathophysiology, Faculty of Medicine, University of Ljubljana; Zaloska 4, 1000 Ljubljana, Slovenia. E-mail: robert.zorec{at}mf.uni-lj.si.


   Abstract
Top
 Abstract
Introduction
 Materials and Methods
 Results
 Discussion
 References
 
In neuroendocrine cells, discharge of hormones follows the fusion of exocytotic vesicles with the plasma membrane at confined sites; however, the molecular nature of these distinct sites remains poorly understood. We studied intact pituitary lactotrophs and plasma membrane lawns by confocal microscopy in conjunction with antibodies against rat prolactin (rPRL), soluble N-ethylmaleimide-sensitive factor-attachment protein receptor (SNARE) proteins (syntaxin-1 and synaptobrevin-2,) and fluorescent cholera toxin subunit B (CT-B), a marker of ganglioside monosialic acid (GM1) lipid rafts, to examine 1) whether rPRL vesicles discharge cargo at GM1 rafts, 2) whether discharging rPRL vesicles interact with SNAREs, and 3) to examine the overlap of GM1 rafts, rPRL, and syntaxin-1 sites in plasma membrane lawns. In intact cells, immunofluorescently labeled rPRL poorly colocalized (<6%) with CT-B. In conditions favoring endocytotic trafficking, vesicle SNARE synaptobrevin-2 modestly colocalized (35%) with CT-B, whereas it highly colocalized (58%) with retrieved rPRL. Although partial mixing between rPRL and CT-B intracellular trafficking pathways is likely, our results indicated that rPRL discharge involves interactions with plasma membrane SNAREs, but not with GM1 rafts. In support of this, the plasma membrane SNARE syntaxin-1 poorly colocalized with CT-B (<5%), whereas it highly colocalized (75%) with rPRL in inside-out plasma membrane lawns. Spontaneous and stimulated rPRL discharge in live lactotrophs is thus associated with plasma membrane sites enriched with SNARE proteins, however, spatially confined to plasma membrane areas other than GM1 rafts.


   Introduction
Top
 Abstract
 Introduction
Materials and Methods
 Results
 Discussion
 References
 
RAFTS ARE LATERAL domains in the plasma membrane that contain a specific repertoire of lipids and proteins (1), which are thought to be formed by cholesterol and sphingolipids that pack tightly into gel-like microdomains. Several types of rafts exist within the same plasma membrane and may be involved in cell signaling (2). Because the first copurification of soluble N-ethylmaleimide-sensitive factor-attachment protein receptor (SNARE) proteins with detergent-resistant membranes (DRMs) (3), the presence of SNARE proteins within lipid rafts has been anticipated, given that DRMs have been considered as analogs of lipid rafts.

Vesicle fusion with the plasma membrane apparently requires the formation of a SNARE complex between proteins in the two interacting membranes. Chamberlain and collaborators (4, 5, 6) reported the presence of syntaxin-1, synaptosome-associated proteins (SNAPs), and synaptobrevin in DRMs extracted with Triton from PC12 (pheochromocytoma 12) cells and adipocytes, which indicates a role for lipid rafts either as plasma membrane platforms for docking and fusion of vesicles in regulated exocytosis (7) or as negative regulators of neuronal exocytosis (6). A direct involvement of lipid rafts in the regulation of calcium-dependent exocytosis was also suggested in PC12 cells and in pancreatic β-cells, where SNARE proteins have been demonstrated to be enriched in DRMs. Depletion of membrane cholesterol in β-cells resulted in enhanced exocytotic events and insulin release (8), whereas cholesterol depletion in PC12 cells inhibited exocytosis and dopamine release (4, 9). Other investigators found SNAREs excluded (syntaxin-2), equally distributed between raft and nonraft fractions (syntaxin-4 and the vesicle-associated membrane proteins, VAMP-8 and VAMP-2), or selectively enriched in rafts (syntaxin-3 and SNAP-23) of RBL (rat basophilic leukemia) mast cells (10).

Recent fluorescent microscopic studies on SNARE proteins and lipid rafts in fixed cells and plasma membrane sheets seem to indicate that both are clustered in different plasma membrane microdomains. Yet, disintegration of lipid rafts by cholesterol depletion induces impairment of exocytosis. Cholesterol-depleted hippocampal cultures and hippocampal cultures from Niemann-Pick type C1-deficient mice show a remarkable overall augmentation of spontaneous vesicle recycling and concomitant decrease in evoked neurotransmission (11).

In this work, we addressed the question whether ganglioside monosialic acid (GM1) lipid rafts serve as membrane platforms for rat prolactin (rPRL) vesicle discharge in live lactotrophs, because current models suggest that in addition to SNARE protein interactions and Ca2+ dynamics, bilayer curvature and phosphoinositide metabolism represent key features for the exo-/endocytotic cycle of secretory vesicles (12, 13). GM1 is an order-preferring lipid that is currently taken as a raft marker because it can be easily visualized with the cell-binding subunit B of cholera toxin (CT-B) that interacts specifically with the ganglioside GM1 (14). Lactotrophs, rPRL-secreting cells from the anterior lobe of the pituitary gland, provide an ideal model to study regulated secretion of vesicle cargo mediated by distinct exocytotic mechanisms (full fusion, kiss-and-run, pulsing fusion pore, and compound exocytosis), because single exocytotic events can be observed with fluorescence microscopy in real time (15, 16, 17, 18, 19). Several years ago, it was also shown that cultured lactrotophs secrete rPRL in response to depolarization and require micromolar cytosolic Ca2+ (20, 21, 22). Moreover, G proteins close to the site of granule fusion inhibit intracellular Ca2+-dependent exocytosis of rPRL, whereas G proteins facilitate the translocation of secretory granules to the fusion sites (20). More recently, it was shown that in addition to full fusion that leads to a rapid vesicle depletion followed by synchronized endocytosis of clathrin-coated vesicles (23), discharge of subquantal contents of rPRL occurs through exocytotic fusion pores, probably due to kinetic constraints of opening and closing of the fusion pore (18, 19).

Here we show that the lactotroph plasma membrane exhibits discrete sites of rPRL release, which are distinct from the GM1 rafts, however, closely resembling the distribution of syntaxin-1 clusters, suggesting that the local environment of SNARE proteins is likely important for rPRL secretion.


   Materials and Methods
Top
 Abstract
 Introduction
 Materials and Methods
Results
 Discussion
 References
 
Solutions
Extracellular solution contained (in mM) 130 NaCl, 5 KCl, 8 CaCl2, 1 MgCl2, 10 D-glucose, and 10 HEPES/NaOH (pH 7.2). K+-enriched solution contained (in mM) 35 NaCl, 100 KCl, 8 CaCl2, 1 MgCl2, 10 D-glucose, and 10 HEPES/NaOH (pH 7.2). Chemicals were obtained from Sigma (Darmstadt, Germany) and were of highest grade of purity available.

Cultures and microscopy
Pituitaries were obtained after decapitation from male Wistar rats and enriched for lactotrophs (24). The care for experimental animals was in accordance with International Guiding Principles for Biomedical Research Involving Animals developed by the Council for International Organizations of Medical Sciences and Directive on Conditions for issue of License for Animal Experiments for Scientific Research Purposes (Official Gazette of the RS, No.40/85 and 22/87). Cell-loaded coverslips were placed in a recording chamber on the inverted confocal microscope (Zeiss LSM 510, Jena, Germany) and supplied with 400 µl extracellular solution. In some experiments, the extracellular solution contained 4 µM FM 4-64 (Invitrogen, Leiden, The Netherlands). Fluorescent images were acquired by a plan-apochromatic oil immersion objective (x63, 1.4 NA), using 488-nm Ar-Ion and 543-nm He-Ne laser excitation. To spectrally separate the emission of immunofluorescence-labeled rPRL, syntaxin-1, synaptobrevin-2, fluorescent CT-B, and FM 4-64, which were used in various double-labeling experiments, the band-pass 505- to 530-nm and long-pass 560-nm emission filters were used, respectively.

Evaluation of cell viability
Initially, an independent set of assays was undertaken to evaluate cell viability after treatment with decreased temperature and CT-B staining (Invitrogen). After overnight incubation, the cells were incubated 30 min in DMEM either at 37 C or at 4 C and in DMEM containing 1 µg/ml CT-B at 4 C. Cell viability was fluorometrically assessed within 2 h after particular treatment by application of 2 µM calcein acetoxymethyl ester and 4 µM ethidium homodimer-1. Live cells were labeled by calcein acetoxymethyl ester, a fluorogenic esterase substrate that is hydrolyzed intracellularly to a green fluorescent product, whereas dead cells were stained by ethidium homodimer-1, a high-affinity, red fluorescent nucleic acid stain that is able to pass through only the compromised membranes of dead cells. Both labels were applied in PBS for 30 min at room temperature (RT). Confocal and differential interference contrast (DIC) images of live (green fluorescent) and dead (red fluorescent) cells were acquired, and the proportion of live cells was calculated for each set of images. The statistical significance between the proportions of live cells after particular treatment was tested by using the Student’s t test. The results show that after a 30-min exposure to DMEM at either 37 C or at 4 C and to DMEM with 1 µg/ml CT-B at 4 C, the viability of cells was very high (90.1 ± 4.0%, n = 1081; 78.4 ± 6.4%, n = 561; and 86.2 ± 9.2%, n = 601) and comparable after various treatments. Thus, the transient decrease in temperature and binding of fluorescent CT-B to the cell surface did not influence the viability of cells significantly.

Immunocytochemistry of live cells
Cells were washed briefly in extracellular solution and stimulated either in 1 ml extracellular solution containing 50 µM TRH or K+-enriched solution for 5 min at RT, as indicated in the figure legend. Then, the cells were washed briefly with 3% BSA/PBS and incubated for 10 min at RT in 3% BSA/PBS containing guinea pig anti-rPRL antibody (1:1000; Dr. A. F. Parlow, The National Hormone and Peptide Program, Torrance, CA). Subsequently, the cells were washed three times with 3% BSA/PBS and exposed to Alexa Fluor 546-conjugated anti-guinea pig IgG (1:600; Invitrogen) for 20 min at RT. Cells were washed three times with 3% BSA/PBS and once with DMEM and subjected to CT-B staining. Briefly, the CT-B labeled by Alexa Fluor 488 (1:1000; Invitrogen) was applied for 10 min at 4 C. The cells were washed twice by chilled PBS and then subjected to application of anti-CT-B antibody (1:200) for 15 min at 4 C. The cells were washed twice by chilled PBS and transferred into extracellular solution for microscopic examination. In some experiments, the double-labeled cells were further incubated in DMEM at 37 C for 1 h or overnight and examined on the next day. In some assays, 4 µM FM 4-64 was applied either in K+-enriched solution during cell stimulation or in extracellular solution during microscopic observation.

Immunocytochemistry of fixed cells
Before fixation, the cells were stimulated by K+-enriched solution, labeled for rPRL at the plasma membrane surface or GM1 rafts as described above, and incubated overnight in DMEM at 37 C. Then the cells were washed once with 1 ml PBS and fixed in paraformaldehyde (4% in PBS) for 15 min. Cells were permeabilized with 4% paraformaldehyde, 0.1% Triton X-100 for 10 min and washed four times with PBS. Subsequently, mouse serum containing anti-synaptobrevin-2 antibody (1:2000; Synaptic Systems, Goettingen, Germany) was applied onto lactotrophs that were further incubated for 2 h at 37 C. Cells were washed four times with PBS and exposed to the respective secondary antibody solution, Alexa Fluor 546-conjugated antimouse IgG (1:600) or Alexa Fluor 488-conjugated antimouse IgG (1:600). Cells were washed four times with PBS, and the preparation was finally treated with Slow Fade Light Antifade Kit (Invitrogen). In double immunolabeling of fixed cells, the cells were initially treated by a serum containing guinea pig anti-rPRL antibody (1:1000; Dr. A. F. Parlow, The National Hormone and Peptide Program) and incubated overnight at 4 C. Then cells were washed four times with PBS and exposed to respective secondary antibody solution, an Alexa Fluor 546-conjugated anti-guinea pig IgG (1:600; Invitrogen). Afterward, the cells were labeled by anti-synaptobrevin-2 antibody as described above.

Immunocytochemistry of inside-out plasma membrane lawns
Cells were washed briefly in extracellular solution and stimulated in 1 ml K+-enriched solution for 5 min at RT, as indicated in the figure legend. Then, the cells were transferred into 10 ml ice-cold distilled water and sheared by manual shaking for 10 min. For initial evaluation, the membrane lawns were visualized by the application of 4 µM FM 4-64 in extracellular solution. Coverslip-loaded inside-out plasma membrane lawns were briefly washed in DMEM and labeled first by the CT-B as described above. Then the lawns were washed with 3% BSA/PBS and incubated in 3% BSA/PBS containing mouse anti-syntaxin-1 antibody (1:200; Synaptic Systems) for 10 min at RT. The lawns were further washed three times with 3% BSA/PBS and exposed to Alexa Fluor 546-conjugated antimouse IgG (1:600; Invitrogen) for 20 min at RT. Basal membrane lawns were finally washed three times with 3% BSA/PBS and transferred into extracellular solution for microscopic examination.

Fluorescence colocalization and spot-size analysis
Colocalization analysis in double-labeled cells (rPRL/CT-B, FM 4-64/CT-B, rPRL/ synaptobrevin-2, CT-B/synaptobrevin-2 and syntaxin-1/CT-B, rPRL/syntaxin-1), was performed on exported TIFF files by ColocAna software (Celica, Ljubljana, Slovenia). A total of 839 images acquired in 275 cells was analyzed. Briefly, the program counted all red, all green, and all colocalized pixels within the image. The threshold for colocalized pixel count was set to be at least 20% of maximal green and red fluorescence intensity, respectively. The proportion of colocalization between green and red fluorescence was expressed as percentage of colocalized pixels in respect to either all green or all red pixels. Statistical differences between fractions of colocalization were assessed by Student’s t test.

The number of fluorescent rPRL spots was determined in LSM files with the WCIF ImageJ program. Each rPRL spot represented either a single or a multitude of rPRL cores discharged to the cell surface and fluorescently labeled by antibodies. The minimum spot size taken to identify the individual rPRL core was six nearby located pixels, thus the minimum surface covered by a fluorescent spot was 0.015 µm2. For the sake of simplicity, each fluorescent spot exceeding 0.300 µm2 was considered as a compound rPRL core, although some spots potentially reflect contribution of several individual rPRL cores positioned in close vicinity to each other. The total number, size, and the ratio between single and compound fluorescent spots were determined for nonstimulated and stimulated cells, and statistical difference was assessed by Student’s t test. Similar analysis was performed in inside-out plasma membrane lawns derived from nonstimulated and stimulated cells, where fluorescent spots larger than three pixels (0.0075 µm2) were considered as individual syntaxin-1 clusters.

To access fluorescence from rPRL and CT-B in spatially separated regions of the cells, the images were exported to TIFF files and processed by a custom-written Matlab software (Math Works, Natick, MA) that separated each confocal image into the image of the cell periphery and the image of the cell interior. Eight to 15 markers were manually set to the cell perimeter, and the software interpolated the curve between the markers. To obtain the image of the cell interior, the software exported the image of the central region limited by the curve, which was displaced 0.75 µm from the cell perimeter toward the cell interior. This was achieved by eroding 15 pixels (pixel size 0.05 x 0.05 µm) from the marked perimeter curve. The complementing image was exported as the image of the cell periphery. The separated pools of images were analyzed by ColocAna software (Celica) to determine region specific colocalization of rPRL and CT-B fluorescence.


   Results
Top
 Abstract
 Introduction
 Materials and Methods
 Results
Discussion
 References
 
rPRL and CT-B stain discrete domains of the lactotroph plasma membrane
To demonstrate discrete plasma membrane zones of hormone release in rPRL-secreting cells, live lactotrophs were exposed to antibody against rPRL and then to the red fluorescent secondary antibody. This resulted in the appearance of bright fluorescent spots on the cell perimeter (Fig. 1AGo); each spot likely reflected a wash-resistant staining of individual rPRL core surfaced at the cell plasma membrane (15). Fluorescent spots were not observed in cells exposed only to the fluorescent secondary antibody. The mean spot size was 120 pixels (pixel size 0.05 x 0.05 µm), representing an average spot area of 0.299 ± 0.030 µm2 (mean ±SEM; n = 49). These discrete spots likely represent structures with dimensions similar to rPRL-containing vesicles distorted by the point spread function. A spherical rPRL vesicle with a diameter of 250 nm theoretically covers an area of 0.049 µm2, which is below the resolution limit of the optical microscope. A similar, spot-like fluorescence appearance of rPRL vesicles was previously observed in stimulated lactotrophs stained by FM 1-43 and FM 4-64 (15, 18). FM-dyes are widely used to study exo-/endocytosis, vesicle trafficking, and organelle organization in living eukaryotic cells, given that these amphiphilic styryl fluorescent dyes are virtually nonfluorescent in water but show a dramatic increase in the quantum yield and thus increase in fluorescence emission upon incorporation into the cell membranes or any other hydrophobic environment (25).


Figure 1
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  		FIG. 1. Spot-like appearance and non-overlapping punctate distribution of fluorescently labeled rPRL and GM1 rafts in plasma membrane of live lactotrophs. A, Confocal and DIC images of secreted rPRL visualized by anti-rPRL antibody and fluorescent secondary antibody (red); B, confocal and DIC images of GM1-enriched lipid rafts visualized by fluorescent CT-B conjugate (green); C, z-stacked confocal images of a lactotroph double-labeled with anti-rPRL antibody and fluorescent secondary antibody (red) and CT-B conjugate (green) taken 1–3 µm from the cell bottom plane (scale bar, 5 µm); D, fluorescence statistics (mean ±SEM) in nonstimulated (CON), TRH-stimulated (TRH), and high-K+-stimulated (K+) cells double labeled with anti-rPRL antibody and fluorescent secondary antibody (left) and CT-B conjugate (right), where bars denote rPRL fluorescence (black bars) and CT-B fluorescence (white bars) determined in respect to the total fluorescence acquired in the individual image. Numbers above bars indicate the number of images analyzed. *, P < 0.05; **, P < 0.02 denotes statistical differences from control (no stimulation).

 
To visualize GM1 rafts at the cell plasma membrane, live lactotrophs were exposed to fluorescently labeled CT-B and then to the anti-CT-B antibody. GM1 rafts mostly appeared as fluorescent puncta and sometimes as longer fluorescent bands covering the cell surface (Fig. 1BGo).

rPRL surface deposits are distinct from GM1 membrane domains
To examine whether rPRL vesicles fuse with and discharge their cargo by regulated exocytosis at plasma membrane sites containing GM1 rafts, the cells were stimulated by TRH and by K+-enriched solution and labeled by anti-rPRL antibody and fluorescent secondary antibody and subsequently by fluorescent CT-B conjugate. The analysis of pixel colocalization in confocal z-stack images of a fluorescently labeled cell (Fig. 1CGo) revealed very weak overlapping of rPRL and CT-B fluorescence (4.8 ± 0.6%, n = 68, TRH stimulation; 3.7 ± 0.3%, n = 102, K+-induced depolarization). The z-stack images showed apparent optical overlap of GM1-containing lipid rafts and rPRL vesicles with neighboring red rPRL and green CT-B spots clearly separated at successive confocal planes (Fig. 1CGo). In fact, a clear assessment of the overlap between rPRL and CT-B fluorescence was not possible within each optical plane, due to diffraction-limited resolution of the objective lens, as previously reported by Lang et al. (9).

Stimulation by either TRH or K+-enriched solution induced a significant increase of the proportion of rPRL fluorescence determined in respect to the total fluorescence acquired in the individual image, whereas even a slight decline of CT-B fluorescence has been observed after stimulation (Fig. 1DGo). The enhancement of rPRL labeling seems to reflect that the mode and/or the intensity of secretion differs in nonstimulated and in stimulated cells. The fraction of fluorescently labeled rPRL and its relative staining area per equatorial plane of individual cells was higher in stimulated cells (32.4 ± 1.8%, n = 68, and 33.5 ± 1.3%, n = 102, in response to stimulation by TRH and by K+-enriched solution, respectively) than in nonstimulated cells (24.9 ± 3.1%, n = 26). The number of fluorescent spots, likely reflecting the number of release sites, per equatorial plane was 7.38 ± 0.94 in cells stimulated by K+-enriched solution [range, 4–12 with 28% apparent compound secretory events consisting of two or more rPRL cores (see Materials and Methods); n = 145], 4.53 ± 0.23 in cells stimulated by TRH (range, 2–10 with 34% apparent compound secretory events; n = 86), and 6.00 ± 0.47 in nonstimulated cells (range, 4–7 with 19% apparent compound secretory events; n = 36). Thereby, an outstanding enhancement of the proportion of apparent compound events has been observed upon stimulation by either TRH or by K+-enriched solution.

To further verify that secreting rPRL vesicles in nonstimulated and in stimulated cells discharge their cargo at sites different from GM1 rafts, an alternative approach with FM 4-64 was employed to label exocytotic vesicles (Fig. 2Go). First, lactotroph vesicles that entered either spontaneous or stimulus-dependent fusion with the plasma membrane were labeled by 5 min incubation in either extracellular solution or K+-enriched solution containing 4 µM FM 4-64, and GM1 rafts were then labeled by CT-B conjugate in the same cells in the absence of FM 4-64. Stimulation by K+-enriched solution did not significantly (P = 0.32) alter the cell size; the cell diameter was 13.9 ± 0.9 µm in nonstimulated cells (n = 4) vs. 12.9 ± 0.4 µm in stimulated cells (n = 16).


Figure 2
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  		FIG. 2. Distinct labeling of vesicles by FM 4-64 and GM1 lipid rafts. Equatorial plane confocal images of FM 4-64 (red) and CT-B conjugate (green) fluorescence in resting cell and in cell subjected to membrane depolarization. A, Resting cell labeled with CT-B where FM 4-64 was added into the extracellular bath during microscopic observation to stain the entire plasma membrane of the cell; B, resting cell where FM 4-64 was added into extracellular solution but was extensively washed out from the bath before CT-B labeling to confine FM 4-64 staining mainly to endocytic vesicles; C, stimulated cell where FM 4-64 was added into the K+-enriched solution and then washed out before CT-B labeling to confine FM 4-64 staining mainly to endocytic vesicles (scale bar, 5 µm); D, fluorescence colocalization plot displays a fraction of colocalized pixels in respect to CT-B pixels. Vertical bars denote fluorescence statistics (mean ±SEM) measured in nonstimulated cells with FM 4-64 present (A) and with FM 4-64 washed (B) and in stimulated cells with FM 4-64 washed after stimulation (C). Numbers above bars indicate the number of images analyzed. ****, P < 0.001 denotes statistical difference from control (FM 4-64 in the incubation bath).

 
Similar to results of fluorescent rPRL and CT-B double staining (Fig. 1Go), a rather low colocalization between FM 4-64 and CT-B fluorescence in nonstimulated cells (18.5 ± 3.0%; n = 18; Fig. 2Go, B and D) and in stimulated cells (5.7 ± 1.2%; n = 29; Fig. 2Go, C and D) was observed. The fluorescence colocalization after either cell treatment differed significantly from the positive controls, where cells were continuously bathed in extracellular solution containing 4 µM FM 4-64 (69.7 ± 3.1%; n = 14; Fig. 2Go, A and D). The increased colocalization of FM 4-64 and CT-B fluorescence in comparison with that of rPRL and CT-B, particularly under resting conditions, potentially reflects less specific labeling of membrane compartments by FM 4-64. In addition to rPRL vesicles that fused with, and were subsequently retrieved from the plasma membrane, other endocytic membrane compartments like sintaxin-6-positive intracellular membranes, early endosomes, the endosomal recycling compartments used by transferrin (17), and the membranes constitutively cycling between the plasma membrane and the cell interior, likely picked up FM dye. Interestingly, relatively higher colocalization between FM 4-64 with CT-B fluorescence in nonstimulated vs. stimulated cells was observed (Fig. 2DGo). If these membrane sites would be equally labeled by both CT-B and FM 4-64, one would expect an unchanged colocalization in resting and stimulated conditions, if the markers co-retrieved, which was not observed. These results may indicate that after membrane depolarization, endocytosis mainly favors retrieval of exocytotic vesicles that were fused to the plasma membrane in response to stimulation.

Retrieval of rPRL- and CT-B-labeled plasma membrane domains
To study whether the retrieval of rPRL and CT-B labeled plasma membrane domains proceeds along distinct pathways, the cells were stimulated by K+-enriched solution during 5 min, and colocalization analysis of rPRL and CT-B fluorescence signals was performed in cells after moderate (1 h) or prolonged (14 h) incubation at 37 C (Fig. 3Go).


Figure 3
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  		FIG. 3. Depolarizing stimulation of cells predominately influences the labeling of rPRL, whereas subsequent prolonged cell incubation at 37 C favors endocytotic internalization of fluorescently labeled rPRL cores that were engaged in exoctyotic discharge. A, Equatorial plane confocal images in nonstimulated cell (CON), in stimulated cell (STIM), in stimulated cell after 1 h incubation at 37 C (pSTIM 1 h), and in stimulated cell after 14 h incubation at 37 C (pSTIM 14 h), double labeled by anti-rPRL antibody and fluorescent secondary antibody (red, left) and CT-B conjugate (green, middle). Merged fluorescence is displayed in the images on the right. Note the larger number of fluorescently labeled rPRL spots on the perimeter of stimulated cell (see STIM) and the larger number of internalized rPRL spots in stimulated cells after 14 h incubation at 37 C (see pSTIM 14 h). Scale bar, 5 µm. B, The corresponding fluorescence statistics (mean ±SEM), where bars denote fraction of colocalized pixels in respect to the total fluorescence acquired in the individual image. Numbers indicate the number of images analyzed; ****, P < 0.001 denotes statistical difference from control (no stimulation).

 
As it can be observed in Fig. 3Go, where colocalization analysis of rPRL and CT-B fluorescence was undertaken in control and stimulated conditions (Fig. 3AGo), the colocalization of signals remained very low in all cells analyzed either at rest (1.9 ± 0.2%; n = 26) or after stimulation (3.7 ± 0.3%; n = 102) (Fig. 3BGo). Notably, the level of colocalization between rPRL and CT-B fluorescence was low after 1 h exposure to 37 C (2.8 ± 0.3%; n = 73) but significantly increased after 14 h exposure to 37 C (8.8 ± 1.4%; n = 35). The mean area of rPRL spots remained similar when considering the entire period of observation (resting, stimulation, and poststimulation periods). The mean spot size under resting conditions was 0.205 ± 0.028 µm2 (n = 49) and somewhat resembles the mean spot size after stimulation (0.299 ± 0.030 µm2; n = 49) and during the poststimulation period (0.216 ± 0.020 µm2, n = 71, and 0.306 ± 0.043 µm2, n = 70, after 1 and 14 h, respectively).

It is interesting to note that the exposure to 37 C seems to favor internalization of labeled rPRL and CT-B, because live cells also displayed fluorescent red and green spots in the deeper regions of the cytoplasm (Fig. 3AGo, pSTIM 1 h, pSTIM 14 h), although the predominant number of fluorescent spots was still confined to the cell surface. This observation clearly indicates that vesicles with fluorescently labeled rPRL cores that initially surfaced at the plasma membrane sites distinct to GM1-enriched lipid domains were in part subsequently recaptured to traffic toward the central cytoplasm, consistent with previous report (17). Thereby, we further analyzed colocalization of rPRL and CT-B fluorescence separately at the cell periphery and in the cell interior (Fig. 4AGo).


Figure 4
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  		FIG. 4. Internalization of rPRL cores and GM1 lipid rafts after cell stimulation and subsequent incubation at 37 C. Data displayed in Fig. 3Go were reanalyzed to assess fluorescence from rPRL and CT-B in the periphery and in the interior of the cell. A, Schematic representation of the manual position of the markers (black spots) delimiting the cell perimeter (large circle) and splitting the image into the image of the cell periphery (p) and the image of the cell interior (i) (see explanation in Materials and Methods); B, fluorescence colocalization plot displays the fraction of colocalized pixels in respect to all pixels (gray bars) in the interior of stimulated cells (STIM), of stimulated cells after 1 h incubation at 37 C (pSTIM 1 h), and of stimulated cells after 14 h incubation at 37 C (pSTIM 14 h), double labeled by anti-rPRL antibody and CT-B conjugate; C, The corresponding fluorescence statistics (mean ± SEM), where bars denote rPRL fluorescence (black bars) and CT-B fluorescence (white bars) determined in respect to the total fluorescence acquired in the cell interior; D, the corresponding fluorescence statistics (mean ± SEM), where bars denote rPRL fluorescence (black bars) and CT-B fluorescence (white bars) determined in respect to the total fluorescence acquired in the cell periphery. **, P < 0.02; ***, P < 0.01; ****, P < 0.001 denote statistical difference from control (stimulation).

 
We observed that the enhancement of colocalization of rPRL and CT-B fluorescence during exposure of live lactotrophs to 37 C was mostly evident in the cell interior. In the cell interior, the overlapping of rPRL and CT-B staining increased over time from 1.3 ± 0.3% after depolarization to 17.8 ± 4.5% 14 h after depolarization (Fig. 4BGo). Accordingly, fluorescence colocalization of rPRL and CT-B remained very low at the cell periphery. These results suggest that trafficking pathways of retrieved rPRL and GM1 rafts potentially merge at some point after endocytotic retrieval.

To verify that fluorescent spots in the cell interior likely reflect internalization of rPRL and CT-B from the plasma membrane, both fluorescence signals were analyzed separately in the cell interior (Fig. 4CGo) and in the cell periphery (Fig. 4DGo) and expressed as a percentage of total fluorescence. In fact, the relative amount of rPRL labeling in the cell interior increased significantly after cell stimulation, reaching a half-maximal value (~50%) after prolonged exposure (14 h) (Fig. 4CGo). Simultaneously, a proportional decrease of fluorescently labeled rPRL spots was observed at the cell periphery (Fig. 4DGo). CT-B staining, which remained mainly at the level of plasma membrane over the period of observation (Fig. 4DGo), was also internalized, although at a slower rate in comparison with endocytotic retrieval of fluorescently labeled rPRL (Fig. 4CGo). This indicates that 1) initially, the retrieved rPRL and the lipid rafts are kept within separate endocytotic pathways, and 2) at some later stage after retrieval, the vesicle cargo and the plasma membrane constituents underwent partial and simultaneous coprocessing within the same subcellular organelle.

SNARE proteins preferentially colocalize with rPRL deposits
Given that partial internalization of rPRL and CT-B fluorescence was observed, the cells labeled by rPRL and CT-B were then subjected to classical double immunostaining, where colocalization of rPRL and CT-B with synaptobrevin-2, a vesicle membrane SNARE, was examined (Fig. 5Go). The analysis of fluorescence colocalization revealed that synaptobrevin-2 colocalizes significantly less with GM1 rafts labeled by CT-B (35.8 ± 1.9%, n = 133; Fig. 5AGo) than with rPRL vesicles (57.7 ± 1.5%, n = 100; P < 0.001). As further confirmed by double immunostaining in fixed cells, a relatively larger number of rPRL vesicles contain synaptobrevin-2 (77.2 ± 0.9%, n = 103; Fig. 5BGo), which may favor SNARE-dependent fusion of vesicles with the plasma membrane (26, 27, 28). The relatively high proportion of anti-synaptobrevin-2 and CT-B fluorescence colocalization potentially indicates interactions of vSNAREs (vesicle membrane SNAP receptors) with constituents of GM1 rafts. Perhaps, synaptobrevin-2-containing vesicles may play a role in the delivery of matter onto the GM1 rafts. Alternatively, synaptobrevin-2 may escape from the fused vesicle membrane into the plasma membrane once the vesicle is fused fully with the plasma membrane (29).


Figure 5
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  		FIG. 5. Synaptobrevin-2, a vesicle membrane SNARE, confined more to rPRL vesicles than to GM1 rafts. Confocal images are shown of cells stimulated by K+-enriched solution and incubated overnight at 37 C. A, Cell labeled by anti-synaptobrevin-2 (Syb2) antibody and fluorescent secondary antibody (red) and by fluorescent CT-B conjugate (green); B, cell labeled by anti-rPRL antibody and fluorescent secondary antibody (red) and by anti-synaptobrevin-2 antibody and fluorescent secondary antibody (green). Scale bar, 5 µm.

 
To further verify that SNAREs appear less likely at GM1 rafts, we tested whether tSNAREs (target membrane SNAP receptors) colocalize with the GM1 rafts (Fig. 6Go). Thus, further immunolabeling was performed by anti-syntaxin-1 antibody and fluorescent secondary antibody and by fluorescent CT-B conjugate. Due to inaccessibility of the plasma membrane SNAREs to the extracellularly applied antibodies, the cells were osmotically challenged to obtain plasma membrane lawns with the cytosolic face exposed to the extracellular space and then immunolabeled by antibodies (Fig. 6AGo). Initially, the integrity of plasma membrane lawns was visualized by the styryl dye FM 4-64 (Fig. 6BGo). Stained membrane lawns appeared flat and decorated with fluorescent syntaxin-1 and CT-B spots of various fluorescence intensities (Fig. 6Go). The apparent plasma membrane lawn thickness was <1 µm as revealed by z-stacked acquisition of confocal images (Fig. 6BGo, bottom images). To verify that membrane lawns derived from nonstimulated and stimulated cells were of approximately the same size, the minimal and maximal diameter of lawns was measured. The minimal diameter was 9.2 ± 0.5 µm (n = 22) vs. 10.1 ± 0.3 µm (n = 45) in lawns derived from nonstimulated and stimulated cells, respectively. The maximal diameter was 13.6 ± 0.5 µm (n = 22) vs. 13.8 ± 0.4 µm (n = 45) in lawns derived from nonstimulated and stimulated cells, respectively, not significantly different (P = 0.21).


Figure 6
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  		FIG. 6. Fluorescently labeled syntaxin-1 clusters segregate from GM1 lipid rafts but colocalize highly with rPRL in plasma membrane lawns derived from lactotrophs. A, Confocal images of membrane lawns derived from nonstimulated cells double labeled by anti-syntaxin-1 (Syx1) antibody and fluorescent secondary antibody (red) and by fluorescent CT-B conjugate (green). B, The same membrane lawn as in A displayed at various z-positions (top) and completely visualized by 4 µM FM 4-64 (bottom). Note that thickness of basal membrane lawn is apparently less than 1 µm. C, Confocal images of membrane lawns derived from stimulated cells double labeled by anti-Syx1 antibody and fluorescent secondary antibody (red) and by fluorescent CT-B conjugate (green). Scale bar, 5 µm. D, Confocal images of membrane lawns derived from stimulated cells double labeled by anti-Syx1 antibody and fluorescent secondary antibody (green) and by anti-rPRL antibody and fluorescent secondary antibody (red). Note high apparent colocalization between Syx1 clusters and rPRL spots in membrane lawn. Scale bar, 5 µm. E, The fluorescence colocalization statistics (mean ±SEM), where bars denote fraction of colocalized Syx1 pixels in respect to the CT-B and rPRL fluorescence. Numbers indicate the number of images analyzed. *, P < 0.05; ****, P < 0.001 denote statistical difference from control (no stimulation).

 
The analysis revealed a weak colocalization of CT-B and anti-syntaxin-1 fluorescence: 6.2 ± 0.7% (n = 58) in plasma membrane lawns derived from resting cells and significantly smaller (4.2 ± 0.6%, n = 51; P < 0.05) in those derived from stimulated cells. This contrasts to previous observations, where membrane depolarization induced an increase between CT-B and rPRL fluorescence overlap (Fig. 3Go). However, anti-rPRL fluorescence was strongly colocalized to anti-syntaxin-1 fluorescence (74.7 ± 2.4%, n = 29) in membrane lawns derived from stimulated cells. The amount of syntaxin-1 in plasma membrane lawns was rather low in comparison with the amount of GM1 rafts (Fig. 6Go, A and C) as confirmed by the fraction of CT-B and syntaxin-1 fluorescence in respect to total fluorescence. The CT-B fluorescence covered approximately four times more surface than the fluorescence of labeled syntaxin-1 (84.5 ± 1.9 vs. 20.1 ± 2.1% in membrane lawns derived from resting cells, n = 58; 87.9 ± 1.3 vs. 15.5 ± 1.6% in membrane lawns derived from stimulated cells, n = 51). The slight, statistically nonsignificant reduction of syntaxin-1 fluorescence in response to membrane depolarization influenced also the apparent increase of CT-B fluorescence in respect to the total fluorescence acquired in lawns derived from stimulated cells. Interestingly, membrane depolarization significantly changed the size of syntaxin-1 spots from 0.0363 ± 0.0007 µm2 (n = 710 from 21 membrane lawns) to 0.0412 ± 0.0006 µm2 (n = 1654 from 50 membrane lawns; P < 0.02), suggesting that stimulation favored the enlargement of syntaxin-1 clusters. Actually, it has been previously shown that sequence-specific interactions do occur in the transmembrane segments of syntaxin that allow homo- and heterodimerization of syntaxin molecules (30) and its relative enrichment at the fusion sites (31).


   Discussion
Top
 Abstract
 Introduction
 Materials and Methods
 Results
 Discussion
References
 
Lactotrophs discharge rPRL at sites distinct from GM1 lipid rafts
We used confocal microscopy to distinguish and characterize sites of rPRL release and GM1 rafts in plasma membrane of live cultured rat lactotrophs. Given that both lipid rafts and exocytotic vesicles are dynamic membrane structures and clustering of elemental rafts seems to occur in response to stimulation at least in T cells (32), the GM1 rafts and discharged rPRL deposits were compared in nonstimulated and stimulated conditions.

Cell stimulation significantly influenced the probability of secretory activity (Fig. 1Go), as observed previously (16). Increase in rPRL surface deposits was not accompanied by a significant CT-B fluorescence change (Fig. 1Go), indicating that plasma membrane GM1 rafts are apparently not influenced by addition of exocytotic vesicle membrane. This does not mean that GM1 rafts are static structures but is consistent with the view that the sites of rPRL release are spatially distinct from GM1 rafts. Accordingly, the undertaken colocalization analysis of rPRL and CT-B fluorescence denote very low (<5%) colocalization of signals in all cells analyzed either at rest or after stimulation by TRH or K+-enriched solution. Weaker rPRL staining observed in nonstimulated cells may potentially result from a slower discharging process, likely affected by a narrow open fusion pore previously observed in spontaneous elementary exocytotic events, in contrast to rapid stimulated hormone discharge (18).

Retrieved rPRL vesicles and CT-B-labeled domains may coalesce within the same intracellular compartments deeper in the cytoplasm
Recent updates to the action mechanism of cholera toxin include elegant studies that look at the toxin mobility in live cell membranes, using fluorescence correlation spectroscopy (33). GM1-bound membrane-localized CT-B exhibits very low cytoskeleton-dependent mobility (<10–9 cm2/sec) and is slowly taken up as endocytotic cargo into small vesicles until it reaches the Golgi apparatus. Although we found weak colocalization of immunolabeled rPRL with CT-B after 14 h incubation of lactotrophs after brief stimulation (Figs. 3BGo and 4BGo), membranous compartments labeled with CT-B and rPRL remained undoubtedly separated up to 60 min incubation at 37 C (Fig. 3BGo), which seems to be sufficient for endocytotic vesicles to fuse with Golgi cisternae in rat anterior pituitary cells (34). Meanwhile, overlabeling of the vSNARE (vesicle membrane SNAP receptor) protein synaptobrevin-2 in fixed and permeabilized lactotrophs after CT-B endocytosis (14 h incubation at 37 C) showed weak fluorescence colocalization between the two molecular entities (Fig. 5Go), supporting the earlier prediction that during trafficking, GM1-enriched domains might mix with other membrane components, losing their ordered state (35). Consistent with this result, the colocalization between rPRL and CT-B in live cells showed a significant, time-dependent increase after membrane depolarization (Fig. 3BGo). Furthermore, the appearance of rPRL/synaptobrevin-2 and CT-B/synaptobrevin-2 double-labeled structures after prolonged incubation at 37 C (Fig. 5Go) provides evidence for partial mixing between both rPRL and CT-B intracellular trafficking pathways during later stages after retrieval of molecules from the plasma membrane (Fig. 4Go).

The new data clearly confirm coupling of GM1-enriched lipid raft trafficking and trafficking of endocytotic vesicles during late recycling. Hence, the observation of lower colocalization of CT-B with rPRL than with FM 4-64, which was even more pronounced under resting conditions (Fig. 2Go), is consistent with reports on the presence of raft resident lipids and proteins in intracellular membranous intermediates (36, 37). In fixed and permeabilized lactotrophs, a partial overlap of CT-B and synaptobrevin-2 staining was observed (Fig. 5AGo), which seems to mainly result from trafficking of components that cycle with trans-Golgi network and plasma membrane during regulated and constitutive exocytotic pathways (38, 39). Actually, a small but statistically significant increase of CT-B overlap with rPRL was observed after prolonged cell incubation (Figs. 3BGo and 4AGo), which favored internalization of both GM1 rafts and secretory vesicles that retained rPRL (Fig. 4Go, C and D), suggesting connections between the two recycling pathways. Bauer et al. (17) have shown that nonsecreted, internalized rPRL traffics through the same early endosomal recycling compartment used by transferrin, whose receptors have been considered nonraft proteins that track the clathrin-mediated endocytotic pathway (40).

rPRL is discharged at plasma membrane sites enriched in syntaxin-1
Presently, it is not clear whether molecular composition of exocytotic sites differ in a cell type-selective manner. Data related to localization of SNARE proteins in lipid rafts were obtained mostly in subcellular fractions and immortalized cell lines (41, 42, 43), whereas our experiments were conducted in live differentiated cells. Although such differences may help explain putative cell-specific patterns of releasing sites, different methodologies likely add to the diversity of results obtained by different groups (41, 44). Most investigators imaged the association between lipid rafts and SNARE proteins in fixed permeabilized cells, because the intracellular localization of SNARE protein epitopes renders anti-SNARE antibody application to intact cells impossible. In this work, we prepared inside-out basal membrane lawns from live lactotrophs, which allow antibody access to both leaflets of the plasma membrane, and performed copatching experiments to address whether syntaxin-1 is localized to GM1 rafts and whether rPRL is discharged at sites enriched in syntaxin-1 (Fig. 6Go).

Because syntaxin-1 was not localized in GM1 rafts in resting and in stimulated lactotrophs, our results entirely agree with above discussed segregation of labeled rPRL deposits from GM1 rafts (Fig. 3Go). Thereby, our data argue against de novo formation of CT-B-labeled lipid rafts at the vesicle docking sites in stimulated neuroendocrine cells, as previously proposed by Chasserot-Golaz et al. (45). These investigators observed very high colocalization between dopamine release sites, assessed with dopamine β-hydroxylase, and cholera toxin at the cell surface upon stimulation of chromaffin cells with nicotine. Attractively, Ohara-Imaizumi and collaborators (46) achieved a conclusion matching that of our results. They observed that insulin granules are preferentially docked to and fuse with intact plasma membrane where syntaxin-1 cluster are located, which in turn do not colocalize with flotillin-1-enriched lipid rafts in MIN6 (mouse insulinoma 6) β-cells. Moreover, in inside-out basal membrane lawns prepared from PC12 cells, secretory vesicles appear also to preferentially dock and fuse at syntaxin clusters that are distinct from common cholesterol-dependent lipid rafts, because they are detergent soluble and do not copatch with the lipid raft marker Thy-1, a 25- to 37-kDa glycosylphosphatidylinositol-anchored cell surface protein expressed in various cell types (9).

Although we did not concentrate on how retained, nonsecreted rPRL is recycled in lactotrophs, it seems that vesicles, which had undergone transient reversible fusion, are retrieved with cargo via clathrin coating from nonraft domains and at late stages of recycling coalescence with endocytotic vesicles derived from lipid rafts, probably within synaptobrevin-2-containing vesicles. In agreement with this hypothesis, we found negligible colocalization of syntaxin-1 and CT-B when the spatial microlocalization of the plasma membrane SNARE protein and GM1 rafts was examined in inside-out membrane lawns prepared from resting and stimulated lactotrophs (Fig. 6Go). Further examination of microcolocalization of the plasma membrane SNARE syntaxin-1 and fluorescent CT-B clearly revealed distinct sites enriched in syntaxin-1 not overlapping with GM1 rafts (Fig. 6Go, A and C), whereby immunolabeled rPRL highly colocalized with basal membrane sites enriched in syntaxin-1 (Fig. 6DGo). Thereby, this study also provides several significant findings concerning the localization of the so-called SNARE complex (27), which corresponds to the most broadly accepted core machinery that is associated with membrane fusion (47, 48, 49), despite the molecular basis of different forms of exocytosis remaining controversial.

Taken together, our results not only suggest that rPRL-releasing sites coincide with plasma membrane microdomains enriched in syntaxin-1 but also that GM1 rafts are not portions of plasma membrane where rPRL discharge occurs. The segregation of rPRL-releasing sites from GM1 rafts was confirmed in nonstimulated and in stimulated cells, albeit the appearance of rPRL/synaptobrevin-2 and CT-B/synaptobrevin-2 double-labeled structures providing evidence for virtual partial mixing between both rPRL and CT-B intracellular trafficking pathways.


   Acknowledgments
 
We thank Dr. Maja Potokar for assistance in immunocytochemical experiments.


   Footnotes
 
This work was supported by the Grants P3 310 381, Z3 7476 1683, and BI-PT/06-07-002 from the Ministry of Higher Education, Sciences, and Technology of the Republic of Slovenia and by a grant from the Gabinete de Relações Internacionais da Ciência e do Ensino Superior (GRICES) of the Portuguese Ministry of Sciences, Technology, and High Education.

Disclosure Statement: The authors have nothing to disclose.

First Published Online June 12, 2008

1 P.P.G. and M.S. contributed equally to this work. Back

Abbreviations: CT-B, Cholera toxin subunit B; DIC, differential interference contrast; DRM, detergent-resistant membrane; GM1, ganglioside monosialic acid; rPRL, rat prolactin; RT, room temperature; SNAP, synaptosome-associated protein; SNARE, soluble N-ethylmaleimide-sensitive factor-attachment protein receptor.

Received January 22, 2008.

Accepted for publication June 5, 2008.


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 Materials and Methods
 Results
 Discussion
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