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Article |
Calreticulin inhibits commitment to adipocyte differentiation
Correspondence to Michal Opas: m.opas{at}utoronto.ca
Calreticulin, an endoplasmic reticulum (ER) resident protein, affects many critical cellular functions, including protein folding and calcium homeostasis. Using embryonic stem cells and 3T3-L1 preadipocytes, we show that calreticulin modulates adipogenesis. We find that calreticulin-deficient cells show increased potency for adipogenesis when compared with wild-type or calreticulin-overexpressing cells. In the highly adipogenic crt–/– cells, the ER lumenal calcium concentration was reduced. Increasing the ER lumenal calcium concentration led to a decrease in adipogenesis. In calreticulin-deficient cells, the calmodulin–Ca2+/calmodulin-dependent protein kinase II (CaMKII) pathway was up-regulated, and inhibition of CaMKII reduced adipogenesis. Calreticulin inhibits adipogenesis via a negative feedback mechanism whereby the expression of calreticulin is initially up-regulated by peroxisome proliferator–activated receptor
(PPAR). This abundance of calreticulin subsequently negatively regulates the expression of PPAR, lipoprotein lipase, CCAAT enhancer–binding protein 
, and aP2. Thus, calreticulin appears to function as a Ca2+-dependent molecular switch that regulates commitment to adipocyte differentiation by preventing the expression and transcriptional activation of critical proadipogenic transcription factors.
Abbreviations used in this paper: CaMKII, Ca2+/calmodulin-dependent protein kinase II; C/EBP, CCAAT enhancer–binding protein; ChIP, chromatin immunoprecipitation; CREB, cAMP response element binding; EB, embryoid body; EMSA, electrophoretic mobility shift assay; ES, embryonic stem; GAPDH, glyceraldehyde 3-phosphate dehydrogenase; PPAR, peroxisome proliferator–activated receptor; PPRE, peroxisome proliferator responsive element; RA, retinoic acid; RXR, retinoid X receptor; WT, wild type.
© 2008 Szabo et al. This article is distributed under the terms of an Attribution–Noncommercial–Share Alike–No Mirror Sites license for the first six months after the publication date (see http://www.jcb.org/misc/terms.shtml). After six months it is available under a Creative Commons License (Attribution–Noncommercial–Share Alike 3.0 Unported license, as described at http://creativecommons.org/licenses/by-nc-sa/3.0/).
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Introduction |
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2, CCAAT enhancer–binding protein (C/EBP), and aP2 are crucial for the development of adipose cells (Rosen et al., 2000). The process of adipogenesis may also be influenced by different extrinsic factor and intracellular signaling pathways (Rosen and MacDougald, 2006), yet very little is known about the role of Ca2+ homeostasis in adipogenesis. Increased Ca2+ levels lead to inhibition of adipocyte differentiation, which is accompanied by the decrease or, in some cases, the loss of PPAR2, C/EBP, and aP2 expression (Jensen et al., 2004; Serlachius and Andersson, 2004; Zhang et al., 2007). Thapsigargin, an inhibitor of the sarco/ER Ca2+-ATPase, inhibits adipocyte differentiation in 3T3-L1 preadipocytes (Ntambi and Takova, 1996; Shi et al., 2000), suggesting a role for ER stores and ER Ca2+-binding proteins in the modulation of adipogenesis. Calreticulin is a major Ca2+-buffering protein in the lumen of the ER, which also acts as a molecular chaperone and modulator of gene expression (Bedard et al., 2005). The two major functions of calreticulin, chaperoning and Ca2+ buffering, are confined to specific protein domains. The N+P domain of calreticulin forms a folding module, and the acidic C-terminal C domain binds and buffers Ca2+ with high capacity (Baksh and Michalak, 1991; Nakamura et al., 2001). Calreticulin deficiency is embryonic lethal (Mesaeli et al., 1999), and calreticulin-deficient cells have a reduced capacity for Ca2+ storage within the ER and impaired agonist-stimulated Ca2+ release from ER stores, whereas cells overexpressing calreticulin have higher levels of lumenal ER Ca2+ and a larger pool of releasable Ca2+ (Nakamura et al., 2001). Calreticulin's impact on protein folding and Ca2+ homeostasis have been implicated in several signaling pathways (Bedard et al., 2005), thus affecting gene expression and, consequently, the behavior of individual cells and cell communities. In this study, we show that calreticulin may act as a Ca2+-dependent molecular switch that negatively regulates commitment to adipocyte differentiation by preventing the expression and transcriptional activation of critical proadipogenic transcription factors.
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Results |
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30-fold higher in the absence of calreticulin.
The number of adipocyte colonies in calreticulin-deficient embryoid bodies (EBs) was approximately ninefold higher than that in the calreticulin-containing EBs at D20 (Fig. 1 B). This was irrespective of whether the calreticulin gene was removed by homologous recombination (G45crt–/– cells) or by Cre recombinase–mediated excision (L7crt–/– cells; Fig. S1 A, available at http://www.jcb.org/cgi/content/full/jcb.200712078/DC1). Adipogenesis in ES cells was RA and insulin dependent and required an initial 3-d exposure to RA followed by insulin treatment (unpublished data). Microscopically, by Nile red staining, lipid droplets in the wild-type (WT) adipocytes were indistinguishable from those in calreticulin-deficient cells (Fig. S1 B), indicating that the gross morphology of lipid stores was not affected in the absence of calreticulin. Nile red is a red-emitting fluorescent lysochrome (Greenspan et al., 1985) that can be used as a fluorescent alternative to oil red O lipid visualization (Fowler and Greenspan, 1985). Fig. S2 shows the nucleotide sequence of the calreticulin promoter.
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and PPAR2) and late (aP2) adipogenic markers. The abundance of PPAR2 and C/EBP adipogenesis markers was markedly increased at D20 in G45crt–/– and L7crt–/– cells compared with WT (Fig. 1, C and D). The aP2 mRNA levels were undetectable in extracts from the WT and L7 cells, whereas they were high at D20 in extracts from calreticulin-deficient cells (Fig. 1, C and D). It should be stressed that in Fig. 1 C as well as in subsequent figures, adipogenic markers in calreticulin-containing cells are barely discernible. This is caused by a large disparity in the markers' abundance between calreticulin-deficient and calreticulin-containing cells, which makes it difficult to visualize all of them when equally loaded. Modulation of the expression of calreticulin also impacted adipogenesis of 3T3-L1 preadipocytes, a commonly used model for adipogenesis (Otto and Lane, 2005). Increased expression of calreticulin in 3T3-L1 preadipocytes inhibited their adipogenesis, as indicated by oil red O staining (Fig. 1 E). In agreement with ES cell results, molecular markers of adipogenesis (lipoprotein lipase, aP2, PPAR2, C/EBP, and C/EBPβ) were all down-regulated in 3T3-L1 cells overexpressing calreticulin (Fig. 1 F).
Upon induction of adipogenesis with RA, the abundance of calreticulin increased dramatically in the WT ES cells (Fig. 2 A), whereas the abundance of PPAR2 and C/EBP remained persistently low (Fig. 2, B and C).
In contrast, in calreticulin-deficient (G45crt–/–) cells after induction of adipogenesis with RA, PPAR2 and C/EBP levels steadily increased over 20 d of differentiation (Fig. 2, B and C). Similar to the WT ES cells, the 3T3-L1 preadipocytes and 3T3-L1 cells overexpressing calreticulin showed an increase in calreticulin abundance and reduced adipogenic potential upon RA treatment (Fig. 2, D and E). We concluded that the increased expression of calreticulin plays a negative regulatory role during adipogenesis.
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2, C/EBP, and aP2 remained nearly undetectable in extracts from WT and L7 cells (Fig. 3, C and E). However, ionomycin significantly decreased the abundance of PPAR2, C/EBP, and aP2 levels in extracts from calreticulin-deficient cell lines (G45crt–/– and L7crt–/–) at D20 (Fig. 3, C and E).
Modulation of adipogenesis by functional modules of calreticulin
Calreticulin has two structural and functional domains (Nakamura et al., 2001); one responsible for chaperoning, and another for Ca2+ buffering (Fig. 4 A).
To determine which of calreticulin's functions (domains) may be involved in the modulation of adipogenesis, two ES cell lines expressing single functional modules of calreticulin in calreticulin-deficient ES cells (G45crt–/–) were created and tested for adipogenic potential. Because the C domain of calreticulin cannot be stably maintained, it was fused to the P domain of calreticulin (Nakamura et al., 2001). Expression of the domains was tracked using anti-HA antibodies (Fig. 4 A).
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To assess whether the Ca2+ content of intracellular Ca2+ stores is modified in crt–/– cells and calreticulin-deficient cells expressing Ca2+ handling the P+C domain of calreticulin, we performed equilibrium loading experiments with 45Ca2+. Cells were cultured for 54 h in the regular culture medium containing 10 mCi/ml 45Ca2+. The total cellular Ca2+ content was then calculated based on the cell-associated radioactivity and on the specific activity of Ca2+ in the culture medium. We used thapsigargin, an inhibitor of sarco/ER Ca2+-ATPase, to measure the amount of Ca2+ associated with ER-exchangeable intracellular Ca2+ stores in either ES (Fig. 4 F) or 3T3-L1 (Fig. 4 H) cells. To assess the residual amount of Ca2+ contained within thapsigargin-insensitive lumenal Ca2+ stores, we used the Ca2+ ionophore ionomycin (Fig. 4, F and H). Measurement of ionomycin- and thapsigargin-induced Ca2+ discharge from the ER indicated that the WT ES cells and calreticulin-deficient ES cells expressing the P+C domain had higher [Ca2+]ER and [Ca2+]Tot, respectively, compared with the crt–/– ES cells (Fig. 4 F). To provide yet another measure of ER-releasable Ca2+, we measured thapsigargin-induced Ca2+ discharge from the ER versus cytosolic [Ca2+] using the Ca2+-sensitive fluorescent dye fura-2-AM under conditions preventing dye sequestration into the ER (Mery et al., 1996). Fig. 4 G shows that calreticulin-containing WT ES cells and calreticulin-deficient ES cells expressing the P+C domain had higher [Ca2+]ER and [Ca2+]Cyto, respectively, in comparison to the crt–/– ES cells. Similar to the ES cells, overexpression of calreticulin or the P+C domains in the 3T3-L1 preadipocytes also resulted in increased [Ca2+]ER and [Ca2+]Tot compared with control 3T3-L1 cells (Fig. 4 H). Finally, as ionomycin may be inactive in releasing Ca2+ from acidic intracellular compartments, we added the sodium proton ionophore monensin. Monensin-induced Ca2+ release was very small (unpublished data), suggesting that WT, crt–/– cells, and cells expressing the N+P domain did not contain substantial quantities of Ca2+ stored within acidic compartments. Thus, we conclude that the effects of calreticulin on adipogenesis may be mediated by calreticulin-dependent changes in intracellular Ca2+.
Functional relationship between calreticulin and PPAR2
The results so far suggested that calreticulin plays a modulatory role during adipogenesis. We next wanted to determine whether there was a functional relationship between calreticulin and the PPAR transcriptional complex. Upon RA-dependent induction of adipogenesis, RXR and PPAR form a transcriptionally active complex. The calreticulin promoter contains two PPAR-binding sites termed peroxisome proliferator responsive elements (PPREs); one is found at –1,944 bp (designated PPRE1), and the other is found at –590 bp (designated PPRE2; Fig. 5 A), suggesting that the PPAR transcription factor may regulate the calreticulin gene.
To test this, NIH3T3 fibroblasts were cotransfected with PPAR expression vector and RXR expression vector (pLC0, pLC1, pLC2, pLC0mt1, and pLC0mt2) or luciferase reporter gene vectors (pCL2 and pCL3; Fig. 5 A) under control of the calreticulin promoter. pSVβ-galactosidase was used as an internal control. In the pLC0 vector, luciferase was controlled by 2.1 kb of the calreticulin promoter containing both PPRE sites (Fig. 5 A and Fig. S2, available at http://www.jcb.org/cgi/content/full/jcb.200712078/DC1). Fig. 5 B shows that PPAR induced luciferase activity in NIH3T3 cells. Cells transfected with promoterless control plasmids showed no detectable luciferase activity (unpublished data). This finding indicates that PPAR activates transcription of the calreticulin gene.
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-binding site on the calreticulin promoter, we performed reporter gene assay with calreticulin promoter deletions and mutations (Fig. 5, B and C). Deletion of PPRE1 and PPRE2 sites (pLC2 vector) completely abolished PPAR-dependent activation of the calreticulin promoter (Fig. 5 B). Deletion of PPRE site 2 at –1,944 bp (Fig. 5 A) or mutation of PPRE site 1 or 2 (Fig. 5 A) had no effect on PPAR activation of the promoter (Fig. 5, B and C). Thus, upon induction of adipogenesis with RA, RXR and PPAR form a complex, which binds PPRE sites 1 and 2 on the calreticulin promoter and transcriptionally activates the calreticulin gene.
To further elucidate a physical interaction between the calreticulin promoter and PPAR, an electrophoretic mobility shift assay (EMSA) was performed (Fig. 5 C). Synthetic oligodeoxynucleotides corresponding to PPRE sites 1 and 2 were used along with a positive control probe, an ideal PPAR site. Fig. 5 C shows that the PPAR–RXR complex bound to PPRE1 and PPRE2. This DNA–protein interaction was only observed in the presence of RXR, indicating that PPAR is only functional once complexed with RXR (Fig. 5 C). The specificity of the PPRE and PPAR–RXR binding was confirmed by reduced signal intensity after the addition of 30-fold excess of cold probe (Fig. 5 C). The lesser intensity in PPRE1 and PPRE2 complexes (Fig. 5 C, lanes 4 and 6) when compared with complexes containing the ideal PPRE site (Fig. 5 C, lanes 1, 3, and 10) is likely the result of the different nucleotide sequences of these probes. Specificity of the binding was further confirmed by supershift EMSA and mutation of both PPRE sites (Fig. 5 D). Chromatin immunoprecipitation (ChIP) was also performed to determine whether there was a direct interaction between the calreticulin promoter and PPAR. ChIP analysis indicated that PPAR bound to both PPRE sites 1 and 2 on the calreticulin promoter (Fig. 5 E). We concluded that the PPAR–RXR complex binds to calreticulin PPRE1 and PPRE2 sites and activates the calreticulin gene.
PPAR2, along with C/EBP, directly affect gene transcription in the nucleus (Rosen, 2005; Rosen and MacDougald, 2006). Given that PPAR2 is both necessary and sufficient for adipogenesis (Rosen et al., 2000), we determined its spatial expression during adipogenesis in our ES cell system (Fig. 6).
On D20 of differentiation in the crt–/– ES cells (which exhibit increased adipogenesis), PPAR2 was distinctly nuclear (Fig. 6 A). However, in the WT ES cells, PPAR2 was barely detectable and appeared cytosolic (Fig. 6 B). Given that the WT ES cells show reduced adipogenesis and, thus, only sparse adipocyte colonies, Fig. 6 B represents a region of low adipogenic potential that predominates in WT outgrowths. Localization of C/EBP was also investigated in the ES cells, and it showed an essentially identical pattern to that observed for PPAR2 (unpublished data).
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2, C/EBP, and aP2 in all cell lines, with the most dramatic effects in calreticulin-deficient cells (Fig. 7, E and F). KN-92, an inactive analogue of KN-93, did not alter oil red O staining (Fig. 7, C and D) or adipogenic marker expression in either crt–/– or calreticulin-containing ES cells (Fig. 7, E and F). These data suggest that the calmodulin–CaMKII pathway plays an important role during adipogenesis from ES cells. Calcineurin is a negative regulator of adipogenesis (Neal and Clipstone, 2002; Kennell and MacDougald, 2005) and is known to affect calcineurin activity (Guo et al., 2002; Lynch et al., 2005). On D20, calcineurin activity was significantly higher in the WT ES cells than in the calreticulin-deficient cells (Fig. 7 G). In the 3T3-L1 preadipocytes, inhibition of calcineurin with cyclosporin-A promoted adipogenesis (Fig. 7, H and I), whereas constitutive expression of activated calcineurin decreased adipogenesis (Fig. 7, H and I). These data give support for the role of calcineurin in adipogenesis of both ES cells and 3T3-L1 preadipocytes. The present findings are consistent with earlier studies relating activation of calcineurin to the presence of calreticulin (Lynch and Michalak, 2003; Lynch et al., 2005).
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Discussion |
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2 and C/EBP levels were decreasing, implying that the effects of calreticulin fall within the commitment and/or initial stages of adipogenesis. We conclude that calreticulin may act as a Ca2+-dependent molecular switch that negatively regulates commitment to adipocyte differentiation by down-regulating the expression and transcriptional activation of critical proadipogenic transcription factors.
Ca2+-handling module of calreticulin inhibits adipogenesis
An important finding of this study is that calreticulin, at least in part, exerts its effects on adipogenesis via its role as a modulator of Ca2+ homeostasis. Ca2+ has previously been shown to affect adipogenesis of 3T3-L1 preadipocytes (Shi et al., 2000; Jensen et al., 2004). For example, 3T3-L1 cells exposed to elevated external Ca2+ levels accumulated little or no cytoplasmic lipids and showed the diminished expression of PPAR2, C/EBP, and aP2 (Jensen et al., 2004). Increasing cytoplasmic Ca2+ levels by thapsigargin also inhibited early stages of adipogenesis (Shi et al., 2000), all pointing out the importance of Ca2+ on adipogenesis. Indeed, we show here that an increase in intracellular Ca2+ concentration leads to a decrease in adipocyte differentiation. Conversely, decreasing intracellular Ca2+ concentrations promotes adipogenesis of calreticulin-deficient ES cells. The relationship between calreticulin expression, intracellular Ca2+ concentration, and adipogenesis implies that calreticulin exerts its effects on adipogenesis via its Ca2+ homeostatic function. However, the formal proof comes from experiments in which functional modules of calreticulin were expressed in calreticulin-deficient ES cells. Upon expression of calreticulin's Ca2+-buffering P+C domain, adipogenesis was halted, whereas expression of the N+P domain had no effect on the progression of adipogenesis.
Our findings that calreticulin affects the commitment to adipocyte differentiation are in line with reports that calreticulin-dependent Ca2+ signaling also affects several aspects of the differentiation of cardiomyocytes (Li et al., 2002; Grey et al., 2005;Puceat and Jaconi, 2005) and human myeloid cells (Clark et al., 2002). Interestingly, diminished intracellular Ca2+ stores attenuate cardiomyogenesis but promote adipogenesis and differentiation of myeloid cells. Combined, these findings imply that calreticulin and ER Ca2+ must play important roles in a variety of differentiation pathways. Moreover, our data regarding timing of the [Ca2+] manipulations indicate that in adipogenesis, as in cardiomyogenesis (Li et al., 2002), there is a calreticulin-regulated calcium-sensitive step referred to as a checkpoint by Li et al. (2002).
CaMKII and calcineurin activities are crucial for adipogenesis
Both CaMKII and calcineurin are Ca2+ dependent and have been implicated in affecting adipogenesis (Wang et al., 1997; Neal and Clipstone, 2002; Kennell and MacDougald, 2005). The CaMKII pathway is involved in C/EBP regulation and, thus, indirectly in PPAR regulation. CaMKII activates CREB, which, in turn, activates C/EBP (Fajas et al., 2002; Wang et al., 2003; Zhang et al., 2004). CREB plays a role in adipogenesis, and, indeed, expression of dominant-negative CREB blocks adipogenesis (Reusch et al., 2000; Klemm et al., 2001; Reusch and Klemm, 2002; Fox et al., 2006; Vankoningsloo et al., 2006). CaMKII is also involved in the activation of PPAR (Paez-Pereda et al., 2005; Park et al., 2007), indicating that the CaMKII pathway may be critical during adipogenesis. Once C/EBP is expressed, PPAR and C/EBP regulate each other, thereby promoting adipogenesis (Rosen, 2005). Here, we show that increased levels of threonine-phosphorylated CaMKII in calreticulin-deficient cells correlate with their increased adipogenesis, and inhibition of CaMKII attenuates adipogenesis in these cells. In calreticulin-deficient cells, in addition to increased CaMKII activity, calmodulin and CREB are also up-regulated, thus explaining the elevated adipogenesis in these cells.
Given that [Ca2+]ER is lower in calreticulin-deficient cells, the question arises as to how CaMKII activity could be increased in these cells. In calreticulin-deficient cells, the level of tyrosine phosphorylation, including that of c-Src (Papp et al., 2007), is higher than in WT cells (Fadel et al., 1999, 2001; Szabo et al., 2007). Activated c-Src phosphorylates calmodulin on tyrosine 99, thereby increasing the affinity of calmodulin for CaMKII (Abdel-Ghany et al., 1990; Benaim and Villalobo, 2002). This phosphorylation event is inhibited by high Ca2+ concentration (Fukami et al., 1986). Tyrosine-phosphorylated calmodulin effectively activates CaMKII (Corti et al., 1999). In addition, after initial activation, CaMKII becomes autophosphorylated, remaining active even though its activators are removed (Meyer et al., 1992). Therefore, CaMKII may remain active and promote adipogenesis in calreticulin-deficient cells even under reduced intracellular Ca2+ concentration.
We have found that endogenous calcineurin activity was significantly higher in calreticulin-expressing cells that have reduced adipogenic potential compared with the calreticulin-deficient cells that are highly adipogenic. This is in agreement with previous studies on calcineurin-dependent inhibition of adipogenesis (Neal and Clipstone, 2002; Kennell and MacDougald, 2005). Thus, we propose that although the calcineurin pathway inhibits adipogenesis, the Ca2+-independent CaMKII pathway might be responsible for the promotion of adipogenesis in the absence of calreticulin observed here.
Calreticulin modulates PPAR activity through a negative feedback mechanism
PPAR2 and C/EBP are crucial transcription factors during adipogenesis (Rosen, 2005); however, although PPAR2 is necessary for adipogenesis to take place (Rosen et al., 2000), C/EBP has a more accessory role during this process (Rosen et al., 2002). We showed here that PPAR2 and C/EBP are down-regulated in cells overexpressing calreticulin. Therefore, calreticulin may act as a transcriptional regulator of PPAR, whereby it inhibits the binding of PPAR/RXR to PPRE in the promoter region of the target genes, thus inhibiting transcription activation by peroxisome proliferators and by fatty acids (Burns et al., 1994; Dedhar et al., 1994; Winrow et al., 1995). Here, we show that the initial process that precedes the effects of calreticulin on the activity of the PPAR–RXR complex involves PPAR transcriptional activation of the calreticulin gene. PPAR is a potent transcriptional activator of the calreticulin gene as a result of its direct interaction with the calreticulin promoter. Thus, PPAR can up-regulate calreticulin, and, conversely, calreticulin can inhibit its activity. We also found an inverse relationship between calreticulin and PPAR2 expression upon induction of adipogenesis with RA. RA induces the expression of calreticulin but reduces the expression of PPAR2. This supports the notion of a hierarchical process in which transcriptional activation of the calreticulin gene by PPAR is the early event followed by calreticulin modulation of PPAR transcriptional activity. These data provide evidence for a previously unrecognized negative feedback loop whereby PPAR regulates the expression of calreticulin, and calreticulin modulates PPAR2 activity and expression. This is likely one important mechanism whereby calreticulin and ER Ca2+ homeostasis regulate adipogenesis.
In conclusion, for the first time, we show an essential role for organellar Ca2+ in adipogenic regulation. The presence of calreticulin inhibits adipogenesis through its effects on Ca2+ homeostasis, causing the indirect regulation of crucial pathways affecting adipogenesis, such as the calcineurin and CaMKII pathways. Although the CaMKII pathway is important for adipogenic differentiation, the calcineurin pathway may be involved in other choices of fate of ES cells. Finally, we show that the expression of calreticulin is tightly regulated during adipogenesis, and any modification of this expression influences adipogenesis such that aberrant calreticulin expression may lead to a variety of cellular pathologies, including obesity and diabetes.
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Materials and methods |
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Adipocyte differentiation was performed by hanging drop method. In brief, 1,000 ES cells in 20-µl drops were placed on the lids of tissue culture dishes for 2 d. The differentiation medium contained high glucose DME (Multicell; Wisent) supplemented with 15% FBS (Multicell; Wisent), sodium pyruvate, L-glutamine, minimal essential medium nonessential amino acids, and β-mercaptoethanol. After 2 d, the aggregated ES cells formed EBs, which were floated in differentiation medium supplemented with 10–7 M RA for an additional 3 d. The EBs were then plated onto tissue culture dishes coated with 0.1% gelatin (in PBS) in differentiation medium supplemented with 1 µg/ml insulin and 2 nM triiodothyronine. The differentiation medium was changed every 2 d for the duration of differentiation (15 d). The EBs were then collected for Western blot analysis, RNA isolation, or staining with oil red O.
Mouse embryonic fibroblasts were isolated from calreticulin-deficient (K42) and WT embryos (K41) as previously described (Nakamura et al., 2000). The 3T3-L1 preadipocyte cells were cultured in DME supplemented with 10% bovine growth serum at 37°C. For adipocyte differentiation, 2-d postconfluent cells were cultured for 2 d in differentiation medium containing 1 µM dexamethasone, 0.5 mM methylisobutylxanthine, 5 µg/ml insulin, and 0.5 µM rosiglitazone (Cayman Chemical). Afterward, the 2-d insulin (5 µg/ml) and 0.5 µM rosiglitazone were added. At day 9 of differentiation, the cells were harvested for Western blot analysis, RNA isolation, or staining with oil red O.
Plasmid DNA and transfections
pSG5-rRxR and pSG5-mPPAR2 were gifts from R. Rachubinski (University of Alberta, Edmonton, Alberta, Canada). Plasmid DNA was purified using Mega plasmid preparation and columns (QIAGEN) as recommended by the manufacturer. Expression vectors encoding the HA-tagged N+P domain (pcDNA3.1_Zeo-CRTNPd-1) and P+C domain (pcDNA3.1_Zeo-CRTPCd-3) of calreticulin were previously described (Nakamura et al., 2001). ES cells were electroporated with 30 µg/cuvette of pcDNA3.1_Zeo-CRTNPd-1 or pcDNA3.1_Zeo-CRTPCd-3 expression vector by electroporation (1,500 V/cm; 25 IxF), and Zeocin (30 µg/ml)-resistant clones were isolated. Expression of the recombinant N+P and P+C domains was monitored with anti-HA antibodies (Nakamura et al., 2001).
3T3-L1 cells were stably transfected with pcDNA3.1-CRT (encoding HA-tagged full-length calreticulin), pcDNA3.1_Zeo-CRTNPd-1, or pcDNA3.1_Zeo-CRTPCd-3 using Lipofectamine reagent (Invitrogen) according to the manufacturer's instructions. 3T3-L1 clones expressing full-length calreticulin or calreticulin domains were selected with 50 µg/ml Zeocin. Expression of recombinant proteins was monitored with anti-HA antibodies (Nakamura et al., 2001).
Reporter gene assay
3T3-NIH cells were cotransfected with reporter plasmid containing the calreticulin promoter, deletion of the calreticulin promoter, or the calreticulin promoter with mutations of PPRE1 or PPRE2 (pLC0, pLC1, pLC2, pLC0mt1, and pLC0mt2), PPAR, and RXR expression vectors. pLC1 and pLC2 plasmids encode the luciferase reporter gene under the control of 1.7-kb and 0.4-kb calreticulin promoters, respectively (Waser et al., 1997). pLC0 encoded the luciferase reporter gene under control of the 2.1-kb calreticulin promoter (Waser et al., 1997). To generate pLC0mt1 and pLC0mt2 plasmids, site-directed mutagenesis of PPRE1 and PPRE2 of the calreticulin promoter, respectively, was performed using the QuikChange Site-Directed Mutagenesis kit (Stratagene). Specifically, PPRE1 (AGGTCAGAGGACA) was mutated to AGGctcGAGGAtc, whereas PPRE2 (TGGCCCTTGACCT) was mutated to ccGggCTgctCCc (lowercase letters are mutated nucleotides). After 48 h, cells were harvested in a lysis buffer containing 100 mM Tris, pH 7.8, 0.5% NP-40, and 0.5 mM DTT. Luciferase and β-galactosidase activity were measured as described previously (Waser et al., 1997).
Inhibitor studies
At the floating stage (days 3–5 of differentiation), EBs were incubated with KN-62 CaMK inhibitor or its inactive analogue (KN-93; Hidaka and Kobayashi, 1994) at concentrations of 10 µM for 2 h during the 3 d of EB flotation stage. The optimal inhibitory concentrations of KN-62 and KN-93 were determined to be in the range of 10 to 15 µM. Under these conditions, the drugs had no effect on cell survival/proliferation. For inhibition of calcineurin, 3T3-L1 preadipocytes were incubated with 1 µg/ml cyclosporin-A. The cells were then allowed to differentiate for an additional 15 d and were harvested for Western blot analysis, RNA isolation, or staining with oil red O.
Ca2+ studies and measurements
The 45Ca2+ measurements as well as [Ca2+] measurements using the Ca2+-sensitive fluorescent dye fura-2-AM were performed as previously described in detail by Mery et al. (1996). The ionomycin and BAPTA-AM treatment regimen was performed as previously described by Li et al. (2002) and Grey et al. (2005) with the following modifications: to obtain the optimal functional output, three consecutive pulse treatments were used for either ionomycin or BAPTA-AM. To chelate cytoplasmic Ca2+, floating EBs were incubated with 50 nM BAPTA-AM for 30 min at days 3–5 of differentiation. To increase cytoplasmic Ca2+, the EBs were incubated for 2 h with 500 nM ionomycin in the same manner. Exposure to ionomycin or BAPTA-AM exerts a significant change even after 1 d of treatment, but optimal results were obtained with three pulses of treatment on three consecutive days of differentiation. The cells were then allowed to differentiate on gelatinized (0.01%) tissue culture dishes for 15 d, after which they were collected for Western blot analysis, RNA isolation, or staining with oil red O.
SDS-PAGE and Western blot analysis
Cells were harvested in a lysis buffer containing 50 mM Tris-HCl, pH 8.0, 120 mM NaCl, and 0.5% NP-40. Protein concentration was determined by the method of Bradford (Bradford, 1976). Protein samples (10 µg per lane) were separated by SDS-PAGE and transferred to nitrocellulose membrane (Nakamura et al., 2001). Western blot analysis was performed with the following antibodies: anti-PPAR2 (Sigma-Aldrich and Santa Cruz Biotechnology, Inc.) at 1:1,000 dilution; anticalreticulin (Opas et al., 1991) at 1:300 dilution; anti-C/EBP (Santa Cruz Biotechnology, Inc.) at 1:1,000 dilution; anti–glyceraldehyde 3-phosphate dehydrogenase (GAPDH; Labfrontiers) at 1:5,000 dilution; anti-CaMKII (Affinity BioReagents) at 1:1,000 dilution; antiphospho-CaMKII Thr286 (Invitrogen) at 1:500 dilution; anticalmodulin (Affinity BioReagents) at 1:1000 dilution; and anti-CREB pSer133 (Sigma-Aldrich) at 1:1,000 dilution. All secondary antibodies were horseradish peroxidase conjugated (Jackson Immunochemicals) and were used at 1:100,000 dilution. Immunoreactive bands were detected with a chemiluminescence ECL Western blotting system (GE Healthcare). Western blots were normalized by using anti-GAPDH antibodies. Relative mRNA levels were normalized to the housekeeping gene L32. Bands were quantified using ImageJ software (National Institutes of Health).
RNA preparation and RT-PCR analysis
Total RNA was isolated from cells grown in 10-cm tissue culture dishes using TRIzol reagent (Invitrogen) according to the manufacturer's instructions. 1 µg RNA was used for synthesis of cDNA followed by RT-PCR using Superscript II (Invitrogen) according to the manufacturer's protocol. The following primers were used for PCR analysis: for PPAR, forward primer 5'-ATCAAGTTCAAACATATCACC-3' and reverse primer 5'-TTGTCTTGGATGTCCTCGATG-3'; for C/EBP, reverse primer 5'-CGCAAGAGCCGAGATAAAGC-3' and forward primer 5'-GCGGTCATTGTCACTGGTCA-3'; for aP2, reverse primer 5'-CATCAGCGTAAATGGGGATT-3' and forward primer 5'-TCGACTTTCCATCCCACTTC-3'; and for L32, reverse primer 5'-CATGGCTGCCCTTCGGCCTC-3' and forward primer 5'-CATTCTCTTCGCTGCGTAGCC-3'. PCR products were separated in 1.5% agarose gel. The mRNA levels were normalized using L32 as the housekeeping gene, and relative mRNA levels were quantified using ImageJ software.
Oil red O staining
Before staining with oil red O, cells were washed twice with PBS, fixed with 10% formaldehyde for 15 min at room temperature, and washed twice with distilled water and once with 70% isopropanol. Next, cells were stained for 1 h at room temperature with filtered oil red O at a ratio of 60% oil red O stock solution (0.5% wt/vol in isopropanol) to 40% distilled water. The cells were washed twice with distilled water, twice with PBS, and examined under a light microscope. An invertoscope (Diaphot; Nikon) equipped with a 10/0.25 DL dry plan Apochromatic objective (Nikon) was used for imaging at room temperature. A camera (Coolpix 4500; Nikon) was used for image acquisition. For quantitative analysis, oil red O was extracted with 5 ml isopropanol for 2 min, and optical density of each sample was determined at 540 nm.
EMSA
Full-length RXR and PPAR and luciferase control proteins were synthesized using a coupled transcription and translation reticulocyte system (Promega; Guo et al., 2001). Synthetic oligodeoxynucleotides corresponding to PPAR site 1 (PPRE1; 5'-AGTGTGGAGGTCAGAGGACACCGGCTC-3') and PPAR site 2 (PPRE2; 5'-TCCTGGCTGGCCCTTGACCTTATCCTG-3') in the calreticulin promoter were used. An ideal PPRE oligodeoxynucleotide sequence, 5'-CAAAACTAGGTCAAAGGTCAAGGCATC-3', was used as a positive control. Underlined residues correspond to the PPAR-binding site. Deoxyoligonucleotides were labeled with -[32P]ATP (GE Healthcare) using T4 polynucleotide kinase. EMSA was performed as described previously (Guo et al., 2001). For supershift analysis, anti-HA tag antibodies were used.
ChIP assay
ChIP assay was performed as described previously (Lynch et al., 2005). In brief, NIH-3T3 cells were transiently transfected with HA-PPAR and cross-linked in 1% formaldehyde at room temperature for 20 min. Cells were then lysed with the Extract-N-Amp kit (Sigma-Aldrich) according to the manufacturer's instructions. Chromatin was sheared by sonication followed by centrifugation for 10 min. Supernatants were precleared with protein A–Sepharose beads for 1 h at 4°C. Immunoprecipitation was performed with mouse anti-HA antibodies at 4°C overnight. DNA was purified and analyzed by PCR using the following primers: PPRE site 1, forward primer 5'-TGTGTCTGAAGACAGCTACAGTG-3' and reverse primer 5'-GCAGCAGGAGAAAAGAAGAGAG-3'; PPRE site 2, forward primer 5'-CTCTATGGCCTGAACAACTGTG-3' and reverse primer 5'-TGGTCAGAGGGAAGAAGTAAGG-3'
Calcineurin activity assay
Calcineurin activity assay was performed as previously described (Fruman et al., 1992). In brief, a peptide corresponding to the regulatory domain of protein kinase A (Sigma-Aldrich) was used as the substrate in an in vitro dephosphorylation assay (RII peptide). 1.0 x 106 cells were lysed in 50 ml hypotonic lysis buffer containing 50 mM Tris, pH 7.5, 0.1 mM EGTA, 1 mM EDTA, 250 mM DTT, and protease inhibitors. 20 ml of lysate was added to 5 mM -[32P]–labeled RII peptide. The reaction was performed for 20 min at 30°C in the presence of 0.5 mM okadaic acid with a total reaction volume of 60 ml. The released phosphate reported the activity of calcineurin and was expressed in picomoles of phosphate released per milligrams of total protein.
Immunofluorescence and Nile red staining
Cells on coverslips were fixed in 3.7% formaldehyde in PBS for 10 min. After washing (three times for 5 min) in PBS, the cells were permeabilized with 0.1% Triton X-100 in buffer containing 100 mM Pipes, pH 6.9, 1 mM EGTA, and 4% (wt/vol) polyethylene glycol 8000 for 2 min, washed three times for 5 min in PBS, and incubated with goat polyclonal anti-PPAR2 antibody (diluted 1:50 in PBS; Santa Cruz Biotechnology, Inc.) for 30 min at room temperature. After washing three times for 5 min in PBS, the cells were incubated with the secondary antibody for 30 min at room temperature. The secondary antibody was FITC-conjugated donkey anti–goat IgG(H+L) diluted 1:50 in PBS. The cells were then incubated with 0.2 mM ribonuclease A (Sigma-Aldrich) for 2 h at room temperature to digest RNA and washed in PBS (three times for 5 min) followed by incubation with 1 µl/ml propidium iodide in PBS for 30 min to identify nuclei. For Nile red staining, the dye stock solution of 0.5 mg/ml in acetone was diluted in a glycerol/PBS mixture (0.05 ml of Nile red stock solution in 1 ml of 3:1 glycerol/PBS). Cells were fixed and permeabilized as for immunofluorescence and incubated with Nile red working solution for 10 min. After the final wash (three times for 5 min in PBS), the slides were mounted in fluorescent mounting medium (to prevent photobleaching; Dako). A confocal fluorescence microscope (MRC 600; Bio-Rad Laboratories) equipped with a 60/1.40 plan Apochromatic oil immersion objective (Nikon) and a krypton/argon laser was used for fluorescence imaging at room temperature. COMOS software (Bio-Rad Laboratories) was used for image acquisition.
Statistical analysis
Differences between mean values for different treatments were calculated using analysis of variance or two-tailed unpaired t test (where applicable) and were considered to be significant at P < 0.05 and P < 0.01.
Online supplemental material
Fig. S1 shows genomic configuration of the calreticulin gene. Fig. S2 shows nucleotide sequence of the calreticulin promoter. Online supplemental material is available at http://www.jcb.org/cgi/content/full/jcb.200712078/DC1.
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Acknowledgments |
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M. Opas is a member of the Heart and Stroke/Richard Lewar Centre of Excellence. This work was supported by grants from the Canadian Institutes of Health Research (CIHR; 36384) and the Heart and Stroke Foundation of Ontario (T6181) to M. Opas and CIHR grants 15415 and 53050 to M. Michalak.
Submitted: 14 December 2007
Accepted: 16 June 2008
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