|Home | About | Journals | Submit | Contact Us | Français|
Chylomicron output by the intestine is proportional to intestinal phosphatidylcholine (PC) delivery. Using five different variations of PC delivery to the intestine, we found that lyso-phosphatidylcholine (lyso-PC), the absorbed form of PC, concentrations in the cytosol (0 to 0.45 nM) were proportional to the input rate. The activity of protein kinase C (PKC)ζ, which controls prechylomicron output rate by the endoplasmic reticulum (ER), correlated with the lyso-PC concentration suggesting that it may be a PKCζ activator. Using recombinant PKCζ, the Km for lyso-PC activation was 1.49 nM and the Vmax 1.12 nM, more than the maximal lyso-PC concentration in cytosol, 0.45 nM. Among the phospholipids and their lyso derivatives, lyso-PC was the most potent activator of PKCζ and the only one whose cytosolic concentration suggested that it could be a physiological activator because other phospholipid concentrations were negligible. PKCζ was on the surface of the dietary fatty acid transport vesicle, the caveolin-1-containing endocytic vesicle. Once activated, PKCζ, eluted off the vesicle. A conformational change in PKCζ on activation was suggested by limited proteolysis. We conclude that PKCζ on activation changes its conformation resulting in elution from its vesicle. The downstream effect of dietary PC is to activate PKCζ, resulting in greater chylomicron output by the ER.
Because obesity and its attendant health issues are a major health problem not only in the United States but also throughout the developed world, potential mechanisms for its mitigation are of increasing importance. One possibility is to block absorbed lipid export from the intestine. The intestine takes dietary FAs and sn-2-monoacylglcyerols (MAGs), the lipolytic products of dietary fat, and converts them to triacylglycerols (TAGs) at the level of the endoplasmic reticulum (ER). These TAGs are incorporated into the intestinal-specific TAG-rich lipoprotein prechylomicron for export from the lumen of the ER to the Golgi. This export step is performed by a specialized transport vesicle, the prechylomicron transport vesicle (PCTV) (1). The exit of prechylomicrons from the ER in PCTV is the rate-limiting step in the transit of dietary lipid across the enterocyte (2, 3). Inhibiting PCTV formation or its fission step from the ER could potentially block absorbed lipid at the level of the enterocyte. The molecular events associated with PCTV formation and their targeting to the Golgi have been recently elucidated (1, 4, 5). This knowledge could lead to specific targets for inhibition of this process.
Recent data suggest that the ER export step for prechylomicrons is potentially regulatable by control of the amount of liver fatty acid binding protein (FABP) 1 able to bind to the ER. We have shown that FABP1 can organize a set of four proteins that can select prechylomicrons as cargo and bud PCTV from the ER membrane (4, 6). We have further found that the binding of FABP1 to the ER to initiate budding is, in turn, controlled by its release from a four-membered cytosolic protein complex present in intestinal cytosol. In the absence of release from the complex, FABP1 cannot bind to the ER (7). The disruption of the complex is controlled by the phosphorylation of Secretion Associated, Ras Related GTPase 1B (Sar1b). We have identified the kinase performing this function as protein kinase C (PKC)ζ (8). On Sar1b phosphorylation, the heteroquatromer is completely disrupted, releasing FABP1 (7).
PKCζ is a member of a family of protein kinases, the atypical protein kinases, which do not require either diacylglycerol (DAG) or Ca2+ for their activation. Assays for PKCζ typically contain phosphatidylserine (PS) as activator. Because the amount of PS required for PKCζ activation is greater than that present in intestinal cytosol, we hypothesized that a different activator was operative. The hypothesis tested in this report is that lyso-phosphatidylcholine (lyso-PC) is the physiological activator of PKCζ in the intestine. We focused on lyso-PC because it has long been known that the amount of phosphatidylcholine (PC) delivered to the intestine correlates with chylomicron output into the lymph (9). Prior to its absorption, PC must be hydrolyzed to sn-1 lyso-PC by pancreatic phospholipase A2. In accord with this hypothesis, bile diversion or mdr2 knockout mice, which are not able to deliver PC to the bile, are associated with the lowest chylomicron output by the intestine (10, 11); whereas fat feeding, which increases biliary PC output (12), and PC supplementation both increase chylomicron output (13) as compared with chow-fed controls. These data were obtained using the same intraduodenal TAG input rates. In the past, the correlation between chylomicron output and PC delivery to the intestine has been thought to be due to the availability of PC for chylomicron surface formation. We provide evidence of an alternative mechanism in this report.
[3H]oleic acid (9.2 Ci/mM) and [3H]PC (32.7 Ci/mM) were obtained from Perkin Elmer Life Sciences (Waltham, MA). Lyso-PC, lyso-phosphatidylserine (lyso-PS), lyso-phosphatidylinositol (lyso-PI), and lyso-phosphatidylethanolamine (lyso-PE) were purchased from Avanti Polar Lipids (Alabaster, AL). Iodixanol (OptiPrep) was purchased from Sigma (Sigma Chemical Co., St. Louis, MO). ECL reagents were procured from GE Healthcare (Fairfield, CT). Protease inhibitor cocktail tablets were obtained from Roche Applied Sciences (Indianapolis, IN). Immunoblot reagents were purchased from Bio-Rad (Hercules, CA). Other biochemicals used were of analytical grade from Sigma (Sigma Chemical Co.) or local companies. Male Sprague Dawley rats, 150–250 g were purchased from Harlan Laboratories (Indianapolis, IN). Rat chow containing 23% fat (w/w, composition 21% milk fat, 2% soybean oil) in pellet form was obtained from Harlan Laboratories.
Rabbit polyclonal antibodies against CD36, caveolin-1, PKCζ, and goat polyclonal antibodies against intestinal alkaline phosphatase were all purchased from Santa Cruz Biotechnology (Santa Cruz, CA). PKCζ substrate-biotinylated was obtained from Santa Cruz Biotechnology. Biotinylated Anti-Rabbit IgG (H+L) antibody was purchased from KPL (Gaithersburg, MD). Streptavidin, HRP conjugate was procured from EMD Millipore Co. (Billerica, MA). Anti-rabbit IgG conjugated with agarose beads was purchased from Sigma Chemical Co. Goat anti-rabbit IgG and goat anti-mouse IgG conjugated with HRP were also procured from Sigma. PKCζ was purchased from Enzo Life Sciences Inc. (Farmingdale, NY).
Intraduodenal infusion of male Sprague-Dawley rats (200–300 g) was performed as described previously (14). In brief, five groups (A to E) of rats participated in the experiments. All the animals were maintained on a chow diet (4% fat, w/w) except group D, which was placed on a high-fat diet (23% fat, w/w) for 2 weeks prior to the experiment. All rats received duodenal cannulas (PE 50) 1 day prior to the experiments and were infused intraduodenally with 0.15 M NaCl, 0.3 M KC1, and 5% glucose at 3 ml/h overnight. Group A rats had their bile duct cannulated (PE 10) in addition to their duodenal cannulas. The next day, the infusion for groups A and B was changed to 0.15 M NaCl. The infusions for groups C and D were changed to 30 mM triolein (TO), 10 mM taurocholate (TC), 0.15 M NaCl, and 10 mM Tris-HC1 (pH 7.4). Group E was given the same TO infusion supplemented with 2 mM PC. All infusions were at 4.5 ml/h for 4 h and were supplemented with [3H]oleate TO (10 × 106 dpm/ml). A summary of the models used is presented in Table 1. After the 4 h infusion was completed, all of the groups were treated identically. The rats were given an overdose of pentobarbital, and the proximal one-half of the small intestine was removed, flushed with ice-cold 0.15 M NaC1, and placed in iced saline.
Enterocytes from the proximal half of rat small intestine were isolated as described previously (1). In brief, enterocytes were isolated from intestinal villi (15) homogenized using a Parr bomb, and the cytosol was isolated by centrifugation. The isolated cytosol was dialyzed against buffer A [0.25 M sucrose, 30 mM HEPES (pH 7.2), 30 mM KCl, 5 mM MgCl2, 5 mM CaCl2, 2 mM DTT] overnight at 4°C. This cytosol was concentrated on a Centricon filter (Amicon, Beverly, MA) with a 10 kDa cutoff to 20 mg of protein/ml.
Caveolin-1-containing endocytic vesicles (CEVs) were isolated as described previously (16). In brief, cytosol (1 mg protein) was treated with 1% Triton X-100. The treated cytosol was placed at the bottom of an OptiPrep step gradient (0 to 35%), and the gradient was centrifuged for 20 h at 4°C. The gradient was resolved by collecting nine 1 ml fractions using a pipette starting at the top of the gradient. One hundred microliters from each fraction was collected for radioactivity determination. The proteins in each fraction were precipitated with TCA, washed twice with cold acetone, and suspended in Laemmli’s buffer. The proteins were separated by SDS-PAGE and identified by immunoblot using specific antibodies as indicated.
Nonfasting rats were anesthetized and gavaged with 0.5 ml corn oil containing [3H]PC (12 × 106 dpm). Thirty minutes later, the rats were euthanized and the proximal half intestines removed. Enterocytes were isolated, and cytosol was obtained as described above. CEVs were isolated by OptiPrep gradient. Ninety-two percent of the cytosolic radioactivity was in lyso-PC.
Phospholipids were extracted from cytosol by chloroform-methanol extraction. Two milliliters of CHCl3/methanol (2:1) was added to 100 μl of cytosol and agitated vigorously. The extracts were centrifuged at 9000 g for 5 min at 4°C. Five hundred microliters of 0.9% NaCl was added to the bottom layer, mixed vigorously, and recentrifuged as described above. The chloroform layer was collected and evaporated under nitrogen for quantitation.
Phospholipids in cytosol were first quantified as described by McHowat and Corr (17). In brief, samples were injected onto an Ascentis Si 5-μm HPLC (25 cm × 4.6 mm) column (Supelco Analytical, Sigma Chemical Co.) and eluted with a mobile phase of hexane-isopropyl alcohol-water (45:50:5) at a flow rate of 1.0 ml/min. Absorbance was measured at 203 nm. Samples were also quantitated radiometrically using [14C]acidic anhydride (10 mCi/mM; ARC, St. Louis, MO). Samples were treated as suggested by Wientzek et al. (18). In brief, the lipids from cytosol were extracted as above, and the lyso-PC was isolated from a silicic acid column using chloroform-methanol (1:9) after other lipids had been removed by passing progressively more methanol in chloroform over the column. The lyso-PC was treated with [14C]acidic anhydride (20,000 dpm), and the acetylated lyso-PC was separated by TLC. Its radioactivity was determined and compared with known quantities of lyso-PC treated similarly.
PKCζ activity in cytosol was quantified using a modified method described by Calbiochem (EMD Millipore Co.). In brief, to bind biotinylated anti-rabbit antibody to agarose beads, we incubated 10 μl biotinylated anti-rabbit antibody with 100 μl of anti-rabbit agarose beads for 3 h at 4°C. After incubation, the beads were washed three times with PBS. Recombinant PKCζ (rPKCζ) in 100 μl of buffer B (250 mM Tris-HCl, pH 7.0, 30 mM MgCl2, 5 mM EDTA, 10 mM EGTA, 50 mM β-mercaptoethanol), 15 μl of 1 mM ATP, 15 μl of 0 to 40 nM PS or 0 to 10 nM of lyso-PC, and 65 μl water were incubated with 2 μl of biotinylated PKCζ substrate for 5 min at 37°C. The reaction was stopped with 500 μl of cold PBS. The reaction mixture was added to anti-rabbit agarose beads as described above for 3 h at 4°C. After incubation, the beads were washed with cold PBS three times and suspended in 400 μl of PBS. Five microliters of HRP-conjugated streptavidin was added to the suspension and incubated for 2 h at 4°C, washed three times with cold PBS, followed by the addition of 100 μl of substrate (5 mg o-phenylenediamine in 5 ml of H2O2), and incubated for 5 min at room temperature. The reaction was stopped with 100 μl of 20% H3PO4, and 2 ml of PBS was added. The supernatant was obtained by centrifugation and read at 492 nm.
Immunodepletion was performed by immunoadsorption (1). Briefly, cytosol (1 mg) was incubated with 20 μl of anti-PKCζ rabbit polyclonal antibodies at 4°C for 4 h, and then anti-rabbit IgG conjugated with agarose beads was added. The mixture was slowly agitated for 4 h at 4°C. The antibody-protein complexes were removed by centrifugation. Successful depletion of PKCζ from cytosol was obtained by three rounds of immunodepletion and confirmed by immunoblotting.
Fifty micrograms of CEV was incubated with 6 nM of lyso-PC in 100 μl of buffer C (250 mM Tris-HCl, pH 7.0, 30 mM MgCl2, 5 mM EDTA, 10 mM EGTA, 50 mM β-mercaptoethanol), 15 μl of 1 mM ATP, and 65 μl water for 30 min at 4°C. The reaction was stopped by adding 500 μl of cold PBS. CEVs were reisolated by OptiPrep step gradient as described above. Repopulation of CEVs by activated PKCζ was performed by the incubation of 500 μg PKCζ depleted cytosol with 1 nM of activated rPKCζ (incubated with 6 nM lyso-PC) in a total volume of 500 μl buffer D (0.25 M sucrose, 30 mM HEPES, pH 7.2, 30 mM KCl, 5 mM MgCl2, 5 mM CaCl2, 2 mM DTT) with an ATP regenerating system (1 mM ATP, 5 mM phosphocreatine, and 5 U creatine phosphokinase) for 30 min at 4°C. CEVs were reisolated using an OptiPrep gradient. The proteins from each fraction were separated by SDS-PAGE and identified by immunoblot using PKCζ antibodies.
Native and lyso-PC-activated PKCζ were subjected to tryptic digestion. Digestions were carried out at 37°C for 0, 2, 4, 8 min by using an enzyme-protein mass ratio of 1:200. Reactions were terminated by adding Laemmle’s buffer and boiled for 5 min. The proteolytic digestion products were resolved on SDS-PAGE followed by silver staining.
Cytosol (1 mg) was incubated with 20 μl of specific rabbit antibodies at 4°C for 4 h, anti-rabbit IgG conjugated with agarose beads was added, and the mixture was stirred slowly at 4°C overnight. The beads were removed by centrifugation, washed 10 times with cold PBS, and resuspended in Laemmli’s buffer for immunoblot analysis.
Proteins were separated by SDS-PAGE and then transblotted to nitrocellulose membranes (Bio-Rad). After incubation with specific primary antibodies and peroxidase-conjugated secondary antibodies, labeled proteins were detected using ECL (Amersham Biosciences) and Biomax film (Eastman Kodak, Rochester, NY).
TAG radioactivity was determined by liquid scintillation as described (16). Phospholipids were similarly measured.
Because we planned to greatly vary the PC input rate into the intestine, we first wished to be certain that the variation either in the delivery of PC to the intestinal lumen in our models or the flux of lyso-PC into the intestinal cytosol did not affect the mechanism of FA absorption. Absorbed FA enters the cytosol on a CEV and appears in the detergent resistant membrane (DRM) fraction of an OptiPrep gradient (16). Despite the wide variation in the PC intake (0–9 µmol/h), the FA tracer, [3H]oleate, remained in the DRM in cytosol, consistent with our prior observations (Fig. 1A–E, Table 1). Under all experimental conditions, caveolin-1, CD36, and PKCζ tracked with the [3H]oleate in the DRM. We have previously shown that the detergent soluble fraction in the OptiPrep gradient contains the expected clathrin (16).
These data suggest the possibility that lyso-PC would also be associated with CEV. To test this, rats were gavaged with corn oil supplemented with PC and [3H]PC. After 30 min, the intestine was harvested, and the cytosolic fraction was obtained. Ninety-two percent of the total cytosolic dpm were in lyso-PC. To determine the percentage of the lyso-PC dpm in CEV, we placed the cytosol on an OptiPrep gradient. We found that the majority (69%) of the lyso-PC in cytosol was in the DRM fraction as suggested by its distribution in the OptiPrep gradient. These data are consistent with the lyso-PC being in the CEV (Fig. 2).
It would be predicted that altering the intake of lyso-PC from the intestinal lumen would be reflected in the concentration of lyso-PC in the cytosol. To see if this were so, we measured the lyso-PC concentration in intestinal cytosol in each of our models by HPLC at 203 nm (Fig. 3A). To determine whether the signal at 203 nm was due solely to lyso-PC, we performed LC/MS/MS on a sample (supplementary Fig. 1). The spectrum obtained confirmed that the signal at 203 nm was lyso-PC. Because absorption at 203 nm may in part be due to artifacts, we also performed a radiometric assay on each sample. Lyso-PC was isolated by silicic acid column chromatography and acetylated using [14C]acetic anhydride. The acetylated lyso-PC was separated by TLC, and its radioactivity was determined. Similarly treated lyso-PC standards were used for quantitation. The results of this alternative assay (Fig. 3B) agreed closely (5–20% greater) with the quantitation using 203 nm (Fig. 3A).
As expected, the lyso-PC concentration in the cytosol varied with the input rate (Fig. 3A, B). Because most of the PC delivered to the intestine comes from bile (19), bile diversion should result in a reduction in cytosolic lyso-PC over bile intact rats. In agreement with this expectation, only negligible amounts of lyso-PC were found in this model (Fig. 3A, B). By contrast, the maximal amount of lyso-PC was found when PC was included in the intraduodenal TO infusion, 0.46 nM (Fig. 3A, B). We also analyzed the cytosol for PC, PS, phosphatidylethanolamine, and phosphatidylinositol, but only negligible amounts were found (data not shown). Similarly, the lyso derivatives of each phospholipid except PC were present in very small amounts (data not shown).
Our major interest was whether the changes in cytosolic lyso-PC were associated with alterations in PKCζ activity. To determine PKCζ activity in our models, we first wished to know the range of cytosolic protein concentrations associated with rates of PKCζ activity proportional to the PKCζ added (Fig. 4A). All subsequent PKCζ activity measurements used protein concentrations of 40 µg or less. As shown in Fig. 4B, PKCζ activity closely tracked the concentration of lyso-PC in the cytosol (Fig. 3). The lowest activity was found in the bile diversion model, and the highest activity was found when PC was included in the intraduodenal TO infusion model with a 34-fold difference between them. These changes in enzyme activity were not due to alterations in the quantity of PKCζ as judged by immunoblot (Fig. 4C).
The correlation of lyso-PC concentrations in the cytosol with PKCζ activity suggested that lyso-PC might be an activator of PKCζ. To directly test this, we incubated rPKCζ with ATP and increasing amounts of lyso-PC. The results (Fig. 5A) show a robust response in PKCζ activity on the addition of lyso-PC suggesting that lyso-PC is a potent PKCζ activator. Importantly, the calculated Km of 1.49 nM for lyso-PC activation is above the values found for lyso-PC concentrations in the cytosol in our models. This implies that changes in lyso-PC concentrations in our models would have a direct effect on PKCζ activity confirming the data in Fig. 4. Because PS is a well-known activator of PKCζ, we also tested PKCζ activity with increasing amounts of PS. As expected, PS also activated PKCζ (Fig. 5B) but with a reduced potency as compared with lyso-PC with a Km of 41 nM, 28-fold greater than lyso-PC. Importantly, the concentration of PS in our models was so low that it would not be expected to be an effective activator of PKCζ (data not shown).
Because lyso-PC is a detergent, we wondered if this property activated PKCζ or if there were a more specific interaction between the two. To test this, we used both SDS and Triton X-100 (1%), a concentration similar to what was used to isolate CEV. Both detergents were ineffective as activators (0.03% and 0.09% the activity of lyso-PC stimulated PKCζ, respectively). These results were not due to denaturation of PKCζ, because the enzyme maintains its ability to be activated by lyso-PC even after exposure to 1% Triton X-100. Finally, we considered the possibility that lyso-PC bound to PKCζ nonspecifically. To determine whether this were so, we immunoprecipitated PKCζ, Sar1b, vesicle associated membrane protein (VAMP)7, and IgG from intestinal cytosol using cytosol isolated from rats gavaged with corn oil, PC, and [3H]PC. At the end of three rounds of immunodepletion, when anti-PKCζ antibody was used, 75% of the [3H]PC had been removed from the cytosol but only 13% of the dpm when Sar1b was immunodepleted, 12% using anti-VAMP7 antibodies, and 9% using IgG. In sum, these data support the thesis that lyso-PC specifically binds to PKCζ and activates it.
To more directly confirm the relationship between PKCζ activation and lyso-PC concentrations in cytosol, we related PKCζ activation to lyso-PC concentration. This relationship revealed a typical saturation curve (Fig. 5C). We utilized the initial rate portion of the curve to delineate a mathematical relationship between the two variables (Fig. 5D). This would suggest that for every additional 1 nM of lyso-PC in the cytosol, PKCζ is ~20% activated.
When lyso-PC was directly tested against PS and ceramide as an activator of PKCζ, lyso-PC was more effective than either phospholipid even though 7-fold more PS and 16-fold more ceramide than lyso-PC were used in the assay (Fig. 6A). Importantly, FA (oleate) was also ineffective as an activator of PKCζ. We also tested lyso-PC against the lyso derivatives of other phospholipids. Although each was an effective activator of PKCζ, lyso-PC was the most potent despite the fact that the other lyso derivatives were used at least at 20-fold greater concentrations (Fig. 6B). Because in vivo the concentrations of the lyso compounds except for lyso-PC were very low in cytosol, none of these would be expected to be effective in vivo PKCζ stimulators. Further, we found that the concentration of ceramide in cytosol did not vary under changing PC input rates into the intestine (Fig. 6C).
We next wished to see what effect exposure of PKCζ to lyso-PC had on its binding to CEV. The data presented in Fig. 7A suggest that the greater the concentration of lyso-PC in cytosol, the more PKCζ eluted from CEV. The data are consistent with the hypothesis that activation of PKCζ detaches PKCζ from CEV allowing it to diffuse in the cytosol. By contrast, other components of CEV such as caveolin-1 (Fig. 7B) and CD36 (Fig. 7C) were relatively unaffected by changes in lyso-PC concentrations and remained with the CEV. These data confirm the specificity of the effect of lyso-PC on PKCζ with respect to CEV binding.
To further confirm the effect of PKCζ activation on PKCζ binding to CEV, we tested whether PKCζ exposed to lyso-PC would bind to CEV. As shown in Fig. 7D, native CEV is replete with PKCζ (bar 1) as expected under native conditions, but on exposure to lyso-PC, CEV was nearly completely depleted of PKCζ (bar 2). Importantly, rPKCζ, activated by lyso-PC, does not repopulate CEV (bar 3). In sum, these data would suggest that activated PKCζ elutes from CEV in a vectorial fashion.
The elution of PKCζ from CEV on its activation suggested the possibility that activation induces a conformational change in the kinase. To test this idea, we used limited proteolysis using trypsin on native and lyso-PC-activated rPKCζ (Fig. 8A). PKCζ was incubated with trypsin from 0 to 8 min as shown below the gel for both the native and activated species. Native PKCζ resisted proteolysis significantly more than activated PKCζ. Native PKCζ’s silver stain signal was extinguished at 8 min of incubation as compared with 4 min for activated PKCζ. Proteolytic fragments can be seen at lower Mr than PKCζ at each time of incubation except for T = 0. Fig. 8B, C show a graphical presentation of the densitometry readings (Gel DocX) at each incubation time as a percentage of the density reading at T = 0 for nonactivated PKCζ (Fig. 8B) and activated PKCζ (Fig. 8C). The increase in the rapidity of proteolysis on activation of PKCζ is clearly evident.
Although CD36, PKCζ, and caveolin-1 were each present on CEV, we next wished to determine whether they were interactive in vivo. To test this, we performed a series of immunoprecipitation experiments (Fig. 9) using cytosol from PC-infused rats (see Materials and Methods). Each of the proteins studied, CD36, PKCζ, and caveolin-1, was found to immunoprecipitate the other two. When the reverse immunoprecipitations were performed, an equally strong signal was obtained confirming the specificity of the reaction.
Finally, we wished to know whether PKCζ, once eluted from CEV, could bind to its cytosolic substrate, Sar1b. This interaction was confirmed by the co-immunoprecipitation experiments shown in Fig. 9. For these experiments, cytosol isolated from PC-infused rats (see Materials and Methods) was used to ensure that most PKCζ was eluted off CEV and was available for potential binding to Sar1b. As shown, the strongest binding to Sar1b was with PKCζ.
The present studies were undertaken to test the hypothesis that the physiological activator of PKCζ in the intestine is lyso-PC. This thesis was based on prior work that showed that chylomicron output into the lymph by the intestine was directly proportional to the amount of PC delivered to the intestine (10, 11, 13). Because the ability of the intestine to export chylomicrons presumably depends on the rate of budding of the PCTV, we thought it likely that lyso-PC, the absorbed hydrolytic product of dietary and biliary PC, would be an activator of PKCζ. This supposition was supported by our preliminary findings on ER-PCTV budding activity using bile-diverted and PC-infused rats (20), and the fact that in the absence of PKCζ, no ER-PCTV budding occurred (7, 8). Additionally, we felt that if we could correlate cytosolic lyso-PC concentrations with PCTV budding, it would more closely tie PCTV budding to chylomicron export by the intestine.
We have used primary isolates of intestinal absorptive cells for these studies. Although intestinal cell models such as the Caco2 cell and the IPEC cell have been used to study lipid absorption and transport, none of them are as efficient as intestinal cells in vivo in the transport of dietary fat. The cell models transport <10% of their TAG content, whereas rat intestine is able to turn over 60% of its TAG content per hour (13). There are a number of potential explanations for this. Caco2 cells have greatly reduced monoacylglycerol acyltransferase (MGAT) activity (21). Acylation of MAG by MGAT provides the major source of DAG substrate for TAG synthesis. TAG synthesized from MAG is more likely to be incorporated into chylomicron-TAG than if it is synthesized via the Kennedy pathway (13, 22). Further, the data reported here were obtained in intestinal epithelial cells that were exposed to lipid solely on their apical surface. Clear differences have been described in the metabolism of FAs absorbed from the basolateral membrane as compared with the apical membrane of this cell type (23). The proportion of absorbed FA utilized for TAG synthesis versus phospholipid synthesis or conversion to CO2 is greater when the FA is presented from the apical versus the basolateral side of the membrane (24, 25). Further, studies using cell models yield differing results when performed in whole animals: apoAIV overexpression is associated with increased TAG output by IPEC cells (26, 27) but not in whole animal experiments (28). Caco2 cells also lack FABP2 that is thought to have a transport role for FA intracellularly (29). Finally, and most importantly, Caco2 cells appear to require a coat protein II mechanism for TAG to exit the ER (30, 31), whereas rat intestinal cells do not (1). In sum, these data provide a strong rationale for using whole intestine to study the molecular details of fat absorption.
In the present study, we found that lyso-PC is a physiological activator of PKCζ. PKCζ is a member of the PKC superfamily of protein kinases, in particular it is an atypical PKC kinase. The atypical PKC’s, specifically PKCζ, are stimulated by a variety of agents such as PS, arachidonic acid (32, 33), phosphatidic acid (PA), lyso-PA, phosphatidylinositol, and phosphatidylglycerol (33) but not DAG or Ca2+. In intestinal cells, we show here that lyso-PC is a potent stimulator over a broad range of activator concentrations. By contrast, in prior experiments lyso-PC has been shown to be a concentration-dependent stimulator/inhibitor of PKCζ activity (32), to be a modest stimulator in a nondose ranging study (34), or to require PS for PKCζ activity to be expressed (35).
In this study, we focused on potential stimulators of PKCζ activity that could be derived from the diet. The major dietary lipids presented for absorption are FAs and MAGs derived from the lipolysis of TAG by pancreatic triacylglycerol lipase and lyso-phospholipids derived from the phospholipolysis of the phospholipids by pancreatic phospholipase A2. The most prominent dietary phospholipid is PC, most of which comes from bile (19). The most prominent lyso-phospholipid identified in intestinal cytosol was lyso-PC, the majority of which was found in the CEV fraction using chow-fed rats. Ceramide was also considered as a potential activator because of its generation from sphingomyelin by sphingomyelinase located on the brush border of enterocytes (36, 37). Because ceramide concentrations did not vary in the cytosol over a wide range of activity changes in PKCζ and was found not to be a potent activator of PKC, it is unlikely that ceramide is a physiologically important PKCζ activator under the conditions of our experiments.
PKCζ may be activated by three different mechanisms, phosphorylation of T410 by phosphatidylinositol-3-phosphate/P DK-1, autophosphorylation of T560, and a conformational change in the kinase resulting in relief of pseudosubstrate inhibition (38). In the current experiments, ATP was required for activation, and a conformational change was observed in the kinase on activation by lyso-PC resulting in us being unable to differentiate between autophosphorylation of T560 and the presumed relief of inhibition induced by the pseudosubstrate on binding to lyso-PC.
On activation, PKCζ is usually considered to move from the cytosol to a specific binding site such as tight junctions (39, 40), signaling molecules (40), or caveolae (41). By contrast, we show in the intestine that PKCζ elutes from CEV on activation. One possibility for this is that PKCζ activation most likely takes place on the surface of the CEV inducing a conformational change in PKCζ as we show here. This structural alteration results in PKCζ eluting from the surface of the CEV, enabling it to diffuse in the cytosol quicker and more easily find its substrate, Sar1b, than if it were on CEV. Once bound to Sar1b, it phosphorylates a threonine (7). The consequence of Sar1b phosphorylation is to free FABP1 from its cytosolic heterotetramer (7), enabling it to bind to the ER and organize the PCTV budding complex (4, 6, 7).
It should be noted that the major actors in the activation of PKCζ are all present on the surface of CEV. Because of this constricted geographic location, small amounts of activator could potentially have significant effects on its target as we show here with lyso-PC and PKCζ. Further, the Km of lyso-PC activation of PKCζ and the measured concentration of lyso-PC in the cytosol in our various models would suggest that in each model, the amount of lyso-PC present would directly affect PKCζ activation.
The results of our limited proteolysis experiments suggest that PKCζ bound to CEV is in a more globular form (molten globule) in which many tryptic susceptible hydrolytic sites are hidden. By contrast, on activation by lyso-PC, PKCζ presumably assumes a more open configuration resulting in more tryptic attackable sites becoming available.
Our current and past data lead us to propose a new feed forward theory of dietary fat absorption (Fig. 10). This theory would suggest that dietary FAs (MAG) enter the apical membrane of enterocytes via caveolae and are endocytosed as CEVs in proportion to the quantity of absorbed FAs (16). We propose that the CEVs are targeted to the ER where they unload their FA and MAG (not shown to be present in CEV) cargo. The FAs and MAG are converted to TAG in the ER, in preparation for their incorporation into prechylomicrons. On the surface of CEV is lyso-PC that has been absorbed from the lumen derived mostly from dietary (biliary) PC and PKCζ. On the surface of the CEV, lyso-PC activates PKCζ, which elutes from the vesicle, binds to, and phosphorylates Sar1b as a member of an FABP1-containing heteroquatromer in intestinal cytosol (7). As a complex member, FABP1 cannot bind to the ER (7). Sar1b phosphorylation disrupts the complex, releasing FABP1, which can now bind to the ER membrane. On binding, FABP1 organizes the PCTV budding complex generating the vesicle that transports the prechylomicron to the Golgi (4, 6). The result of the proposed mechanism is that the ability of the ER to generate and transport prechylomicrons is controlled, at least in part, by the amount of dietary FA delivered to the ER as substrate for chylomicron TAG. This is consistent with the more FAs absorbed, the more CEVs are produced (16). This would provide a potential mechanism whereby the amount of dietary FA could correlate with PCTV production. Although luminal PC availability also controls chylomicron and PCTV output (9, 11, 13, 20, 42), it should be recognized that the more lipid ingested, the greater the gall bladder contraction and thus PC delivery to the intestine. Thus, the amount of dietary lipid and PC delivery to the intestine is mutually supportive.
[S]The online version of this article (available at http://www.jlr.org) contains supplementary data in the form of one figure.