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 γ (PPARγ). This abundance of calreticulin subsequently negatively regulates the expression of PPARγ, lipoprotein lipase, CCAAT enhancer–binding protein α, 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.

Introduction

Obesity is rapidly becoming a worldwide epidemic, leading to numerous medical conditions such as type 2 diabetes and cardiovascular disease. The development of obesity is marked by both an increase in adipocyte size and number as a consequence of differentiation of the stem cells or preadipocytes into mature adipocytes (for review see Gesta et al., 2007). Spontaneous differentiation of embryonic stem (ES) cells into adipocytes in culture is a rare event; thus, for ES cells to commit toward the adipocyte lineage, it is necessary to pretreat cells with retinoic acid (RA), a ligand of the nuclear receptor retinoid X receptor (RXR), during the first phase of differentiation (Dani et al., 1997). RXR in response to RA heterodimerizes with peroxisome proliferator–activated receptor (PPAR), resulting in activation of genes involved in terminal differentiation and lipid metabolism (Gregoire et al., 1998). PPARγ2, CCAAT enhancer–binding protein α (C/EBPα), 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 PPARγ2, C/EBPα, 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.

Results

Calreticulin deficiency promotes adipogenesis

Fig. 1 A shows that the adipogenic potential of ES cells, as measured by oil red O staining at differentiation day 20 (D20), was ∼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). 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.

Figure 1.

Adipogenesis increases in calreticulin-deficient EBs. Oil red O is a lysochrome used for staining triglycerides and some lipoproteins, thus being useful in visualizing the degree of adipogenesis. (A) At D20, calreticulin-deficient ES cells (G45crt−/− and L7crt−/−) stained more extensively with oil red O than WT cells (P < 0.01; n = 6) and made more adipocyte colonies (P < 0.01; n = 5) (B). (C) The ES cells lacking calreticulin (G45crt−/− and L7crt−/−) showed an increase in expression of PPARγ2, C/EBPα (by Western blotting), and aP2 (by PCR) compared with the WT or L7 cells. Relative protein levels were determined by normalizing the calreticulin, PPARγ2, and C/EBPα to GAPDH, and aP2 mRNA levels were normalized with the housekeeping gene L32. (D) Quantitative analysis of the relative expression of adipogenic markers at D20 (P < 0.01; n = 6). (E) Oil red O staining and Western blot analysis with anticalreticulin (CRT) antibodies of 3T3-L1 preadipocytes and 3T3-L1 cells overexpressing calreticulin (3T3-L1 + CRT). 3T3-L1 cell fibroblasts overexpressing calreticulin showed reduced oil red O staining. (F) RT-PCR showing that adipogenic marker expression decreased in 3T3-L1 + CRT cells overexpressing calreticulin compared with control 3T3-L1 cells. Error bars represent SD.

Figure 1.

Adipogenesis increases in calreticulin-deficient EBs. Oil red O is a lysochrome used for staining triglycerides and some lipoproteins, thus being useful in visualizing the degree of adipogenesis. (A) At D20, calreticulin-deficient ES cells (G45crt−/− and L7crt−/−) stained more extensively with oil red O than WT cells (P < 0.01; n = 6) and made more adipocyte colonies (P < 0.01; n = 5) (B). (C) The ES cells lacking calreticulin (G45crt−/− and L7crt−/−) showed an increase in expression of PPARγ2, C/EBPα (by Western blotting), and aP2 (by PCR) compared with the WT or L7 cells. Relative protein levels were determined by normalizing the calreticulin, PPARγ2, and C/EBPα to GAPDH, and aP2 mRNA levels were normalized with the housekeeping gene L32. (D) Quantitative analysis of the relative expression of adipogenic markers at D20 (P < 0.01; n = 6). (E) Oil red O staining and Western blot analysis with anticalreticulin (CRT) antibodies of 3T3-L1 preadipocytes and 3T3-L1 cells overexpressing calreticulin (3T3-L1 + CRT). 3T3-L1 cell fibroblasts overexpressing calreticulin showed reduced oil red O staining. (F) RT-PCR showing that adipogenic marker expression decreased in 3T3-L1 + CRT cells overexpressing calreticulin compared with control 3T3-L1 cells. Error bars represent SD.

Next, we examined the expression of both early (C/EBPα and PPARγ2) and late (aP2) adipogenic markers. The abundance of PPARγ2 and C/EBPα 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, PPARγ2, C/EBPα, 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 PPARγ2 and C/EBPα remained persistently low (Fig. 2, B and C). In contrast, in calreticulin-deficient (G45crt−/−) cells after induction of adipogenesis with RA, PPARγ2 and C/EBPα 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.

Figure 2.

Calreticulin, PPARγ2, and C/EBPα levels increase over the 20-d differentiation time line, whereas overexpression of calreticulin inhibits adipogenesis. (A–C) Calreticulin (CRT), PPARγ2, and C/EBPα expression was examined in WT and G45crt−/− cells by Western blotting over 20 d of adipocyte differentiation as described in Materials and methods. The arrows represent the addition of retinoic acid (RA) or insulin (Ins) and TIT (triiodothyronine). (A) 10−7 M RA treatment of cells during the first 3 d of differentiation induced calreticulin expression in WT cells. (B and C) Expression of PPARγ2 and C/EBPα in RA-treated crt−/− cells gradually increased, whereas their levels in WT cells remained low over the 20-d differentiation. Standard error bars in A–C are smaller than the symbols. RA treatment up-regulates calreticulin in 3T3-L1 cells. (D) Western blot analysis with anti-HA antibodies of 3T3-L1 preadipocytes (3T3-L1 control) and 3T3-L1 cells overexpressing HA-tagged calreticulin (3T3-L1 + CRT) show a marked increase in calreticulin levels after RA treatment. (E) 3T3-L1 cells treated with RA show a decrease in oil red O staining, whereas overexpression of calreticulin reduced the staining further. Error bars represent SD.

Figure 2.

Calreticulin, PPARγ2, and C/EBPα levels increase over the 20-d differentiation time line, whereas overexpression of calreticulin inhibits adipogenesis. (A–C) Calreticulin (CRT), PPARγ2, and C/EBPα expression was examined in WT and G45crt−/− cells by Western blotting over 20 d of adipocyte differentiation as described in Materials and methods. The arrows represent the addition of retinoic acid (RA) or insulin (Ins) and TIT (triiodothyronine). (A) 10−7 M RA treatment of cells during the first 3 d of differentiation induced calreticulin expression in WT cells. (B and C) Expression of PPARγ2 and C/EBPα in RA-treated crt−/− cells gradually increased, whereas their levels in WT cells remained low over the 20-d differentiation. Standard error bars in A–C are smaller than the symbols. RA treatment up-regulates calreticulin in 3T3-L1 cells. (D) Western blot analysis with anti-HA antibodies of 3T3-L1 preadipocytes (3T3-L1 control) and 3T3-L1 cells overexpressing HA-tagged calreticulin (3T3-L1 + CRT) show a marked increase in calreticulin levels after RA treatment. (E) 3T3-L1 cells treated with RA show a decrease in oil red O staining, whereas overexpression of calreticulin reduced the staining further. Error bars represent SD.

Changes in the intracellular Ca2+ concentration affect adipogenesis

One important function of calreticulin is modulation of Ca2+ homeostasis (Nakamura et al., 2001; Bedard et al., 2005). Thus, we next examined whether calreticulin-dependent changes in Ca2+ homeostasis might be responsible for modulation of adipogenesis. We used BAPTA-AM, a membrane-permeable Ca2+ chelator (Tsien, 1980), to lower cytosolic Ca2+ concentration ([Ca2+]Cyto) as well as ionomycin, a Ca2+ ionophore (Bergling et al., 1998), to increase [Ca2+]Cyto. Treatment of cells with BAPTA from 20 min to 2 h produced exactly the same results (unpublished data), which implies that BAPTA was not secreted from the cells. BAPTA-AM–treated WT and L7 cells showed a dramatic increase in oil red O staining, indicating that chelation of cytoplasmic Ca2+ promoted adipogenesis in these cells (Fig. 3, A and B). There was no significant increase in oil red O staining in BAPTA-AM–treated calreticulin-deficient ES cells (G45crt−/− or L7crt−/−; Fig. 3, A and B). In contrast, after treatment with ionomycin, there was no detectable oil red O–positive adipocytes derived from WT and L7 cells, whereas differentiation of both calreticulin-deficient cell lines (G45crt−/− and L7crt−/−) produced an extremely low number of oil red O–positive cells (Fig. 3, A and B).

Figure 3.

Changes in intracellular Ca2+ concentration affect adipogenesis. BAPTA-AM, a membrane-permeable Ca2+ chelator, was used to lower cytosolic Ca2+ concentration, whereas a Ca2+ ionophore, ionomycin, was used to increase it. (A) WT and L7 ES cells showed increased oil red O staining after treatment with BAPTA-AM, whereas no substantial change in the staining was discernible in calreticulin-deficient ES cells (G45crt−/− and L7crt−/−). Ionomycin treatment decreased oil red O staining in calreticulin-deficient cells. (B) Colorimetric assay of oil red O staining in the ES cells treated as in A (n = 6). (C) Treatment with BAPTA-AM increased the expression of PPARγ2, C/EBPα, and aP2 in WT or L7 ES cell lines. Conversely, ionomycin treatment decreased the adipogenic marker expression in calreticulin-deficient (G45crt−/− and L7crt−/−) cells. (D) Quantitative analysis of relative adipogenic marker expression after BAPTA-AM treatment (P < 0.01; n = 6). (E) Quantitative analysis of relative adipogenic marker expression after ionomycin treatment (P < 0.01; n = 6). Error bars represent SD.

Figure 3.

Changes in intracellular Ca2+ concentration affect adipogenesis. BAPTA-AM, a membrane-permeable Ca2+ chelator, was used to lower cytosolic Ca2+ concentration, whereas a Ca2+ ionophore, ionomycin, was used to increase it. (A) WT and L7 ES cells showed increased oil red O staining after treatment with BAPTA-AM, whereas no substantial change in the staining was discernible in calreticulin-deficient ES cells (G45crt−/− and L7crt−/−). Ionomycin treatment decreased oil red O staining in calreticulin-deficient cells. (B) Colorimetric assay of oil red O staining in the ES cells treated as in A (n = 6). (C) Treatment with BAPTA-AM increased the expression of PPARγ2, C/EBPα, and aP2 in WT or L7 ES cell lines. Conversely, ionomycin treatment decreased the adipogenic marker expression in calreticulin-deficient (G45crt−/− and L7crt−/−) cells. (D) Quantitative analysis of relative adipogenic marker expression after BAPTA-AM treatment (P < 0.01; n = 6). (E) Quantitative analysis of relative adipogenic marker expression after ionomycin treatment (P < 0.01; n = 6). Error bars represent SD.

In agreement with oil red O staining results, BAPTA-AM treatment significantly increased the abundance of all three adipogenic markers in adipocytes derived from WT and L7 cells containing calreticulin but did not have a significant effect on adipocytes derived from calreticulin-deficient (G45crt−/− and L7crt−/−) ES cells (Fig. 3, C and D). In the presence of ionomycin, the levels of PPARγ2, C/EBPα, and aP2 remained nearly undetectable in extracts from WT and L7 cells (Fig. 3, C and E). However, ionomycin significantly decreased the abundance of PPARγ2, C/EBPα, 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).

Figure 4.

Expression of the P+C domains of calreticulin reduces adipogenesis. (A) A schematic of the structural and functional calreticulin domains as well as Western blot analysis (with anti-HA antibodies) of G45crt−/− ES cells expressing the N+P and P+C domain of calreticulin. (B) Oil red O staining in WT, G45crt−/−, and G45crt−/− ES cells expressing P+C and N+P domains. G45crt−/−cells expressing the P+C domain had decreased staining compared with the G45crt−/− cells. When the cells were treated with 50 nM BAPTA-AM, both the P+C domain–expressing and WT ES cells showed a dramatic increase in oil red O staining. The N+P domain–expressing ES cells had oil red O staining similar to that of the G45crt−/− ES cells. When the cells were treated with 500 nM ionomycin, both the N+P domain–containing and G45crt−/− ES cells showed a decrease in oil red O staining. (C) Expression of the P+C domain in 3T3-L1 cells (3T3-L1 + [P+C]) reduced oil red O staining. Because 3T3-L1 is used as a control, it is always counted as 1. Expression of the P+C domain was confirmed using HA antibody. (D) Adipogenic marker expression was decreased in the P+C domain–containing G45crt−/− cells when compared with the G45crt−/− cells (P < 0.01; n = 6) and was comparable to that in WT cell levels (P > 0.05). BAPTA-AM treatment increased adipogenic marker expression in all cell lines (n = 6). (E) Adipogenic marker expression was higher in the N+P domain–expressing G45crt−/− cells than in the WT ES cells (P < 0.01; n = 6) and was similar to that in G45crt−/− cell levels (P > 0.05). Ionomycin treatment reduced adipogenic marker expression in all cell lines (n = 6). (F) 45Ca2+-loaded ES cells were treated with thapsigargin or ionomycin to measure ER-releasable and total [Ca2+], respectively. WT and P+C domain–expressing G45crt−/− cells had higher [Ca2+]ER and [Ca2+]Tot than the G45crt−/− cells (n = 3). (G) Fura-2-AM–loaded ES cells were treated with thapsigargin to measure ER-releasable and cytosolic [Ca2+]. WT and P+C domain–expressing G45crt−/− cells had higher [Ca2+]ER and higher cytosolic [Ca2+] than the G45crt−/− cells (n = 3). (H) 45Ca2+-loaded 3T3-L1 cells were treated with thapsigargin or ionomycin to measure [Ca2+]ER and [Ca2+]Tot levels. 3T3-L1 + CRT cells (overexpressing calreticulin) and 3T3-L1 expressing the P+C domain had higher [Ca2+]Tot and [Ca2+]ER than the control 3T3-L1 cells (n = 3). Error bars represent SD.

Figure 4.

Expression of the P+C domains of calreticulin reduces adipogenesis. (A) A schematic of the structural and functional calreticulin domains as well as Western blot analysis (with anti-HA antibodies) of G45crt−/− ES cells expressing the N+P and P+C domain of calreticulin. (B) Oil red O staining in WT, G45crt−/−, and G45crt−/− ES cells expressing P+C and N+P domains. G45crt−/−cells expressing the P+C domain had decreased staining compared with the G45crt−/− cells. When the cells were treated with 50 nM BAPTA-AM, both the P+C domain–expressing and WT ES cells showed a dramatic increase in oil red O staining. The N+P domain–expressing ES cells had oil red O staining similar to that of the G45crt−/− ES cells. When the cells were treated with 500 nM ionomycin, both the N+P domain–containing and G45crt−/− ES cells showed a decrease in oil red O staining. (C) Expression of the P+C domain in 3T3-L1 cells (3T3-L1 + [P+C]) reduced oil red O staining. Because 3T3-L1 is used as a control, it is always counted as 1. Expression of the P+C domain was confirmed using HA antibody. (D) Adipogenic marker expression was decreased in the P+C domain–containing G45crt−/− cells when compared with the G45crt−/− cells (P < 0.01; n = 6) and was comparable to that in WT cell levels (P > 0.05). BAPTA-AM treatment increased adipogenic marker expression in all cell lines (n = 6). (E) Adipogenic marker expression was higher in the N+P domain–expressing G45crt−/− cells than in the WT ES cells (P < 0.01; n = 6) and was similar to that in G45crt−/− cell levels (P > 0.05). Ionomycin treatment reduced adipogenic marker expression in all cell lines (n = 6). (F) 45Ca2+-loaded ES cells were treated with thapsigargin or ionomycin to measure ER-releasable and total [Ca2+], respectively. WT and P+C domain–expressing G45crt−/− cells had higher [Ca2+]ER and [Ca2+]Tot than the G45crt−/− cells (n = 3). (G) Fura-2-AM–loaded ES cells were treated with thapsigargin to measure ER-releasable and cytosolic [Ca2+]. WT and P+C domain–expressing G45crt−/− cells had higher [Ca2+]ER and higher cytosolic [Ca2+] than the G45crt−/− cells (n = 3). (H) 45Ca2+-loaded 3T3-L1 cells were treated with thapsigargin or ionomycin to measure [Ca2+]ER and [Ca2+]Tot levels. 3T3-L1 + CRT cells (overexpressing calreticulin) and 3T3-L1 expressing the P+C domain had higher [Ca2+]Tot and [Ca2+]ER than the control 3T3-L1 cells (n = 3). Error bars represent SD.

Fig. 4 B shows that reexpression of the P+C domain in calreticulin-deficient cells inhibited adipogenesis (Fig. 4, B and D). Overexpression of the P+C domain in 3T3-L1 preadipocytes also resulted in reduced adipogenesis (Fig. 4 C). Treatment of the P+C domain–expressing crt−/− ES cells with BAPTA-AM restored their adipogenic potential (Fig. 4, B and D). In contrast, expression of the N+P domain in crt−/− ES cells had no significant effect on their ability to differentiate into adipocytes, and they maintained a phenotype identical to calreticulin-deficient cells (Fig. 4 B). This was further supported by BATPA-AM and ionomycin experiments (Fig. 4), indicating that the chaperone function of calreticulin was not involved in the modulation of adipogenesis. Therefore, crt−/− cells expressing the N+P domain of calreticulin displayed features resembling those of crt−/− ES cells, whereas crt−/− cells expressing the Ca2+-buffering region of the protein (P+C domain) had properties characteristic of those of WT cells, indicating that calreticulin modulates adipogenesis via its Ca2+ homeostatic function.

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 PPARγ2

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 PPARγ transcriptional complex. Upon RA-dependent induction of adipogenesis, RXR and PPARγ form a transcriptionally active complex. The calreticulin promoter contains two PPARγ-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 PPARγ transcription factor may regulate the calreticulin gene. To test this, NIH3T3 fibroblasts were cotransfected with PPARγ expression vector and RXRα 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). Fig. 5 B shows that PPARγ induced luciferase activity in NIH3T3 cells. Cells transfected with promoterless control plasmids showed no detectable luciferase activity (unpublished data). This finding indicates that PPARγ activates transcription of the calreticulin gene.

Figure 5.

PPARγ activates the calreticulin gene. (A) A schematic representation of reporter plasmids used in this study. The calreticulin promoter (pLCC0) contains two PPRE sites: PPRE1 (−1,944 bp) and PPRE2 (−590 bp). Two promoter deletion constructs (pLC1 and pLC2) and two promoter constructs, one containing a mutation of PPRE1 (pLC0mt1) and the other containing a mutation of PPRE2 (pLC0mt2), were used to identify PPRE sites activated by PPARγ on the calreticulin promoter. (B) To analyze the effect of PPARγ and RXRα on the calreticulin promoter, NIH3T3 cells were transiently cotransfected with the PPARγ and RXRα expression plasmids with a luciferase reporter gene controlled by the calreticulin promoter as indicated in A. Individual controls were obtained for each expression plasmid, and data were plotted relative to that control (luciferase/β-galactosidase). The data shown are mean ± SD (error bars) of three independent experiments. (C) EMSA analysis was performed to demonstrate the binding of PPARγ to the PPRE1 and PPRE2 elements on calreticulin promoter. The positions of PPARγ–RXRα–DNA complexes are indicated. White lines indicate that intervening lanes have been spliced out. (D) Supershift EMSA analysis of PPARγ binding to the PPRE1 and PPRE2 elements on calreticulin promoter using anti-HA tag antibody was used to show the binding of transcriptional factor to the DNA element. Mutated PPRE1 and PPRE2 oligonuleotide show no interaction with PPARγ. Arrows mark PPARγ complexes with either PPRE1 or PPRE2. (E) ChIP analysis of a putative PPARγ-binding site in the mouse calreticulin promoter was used to show binding of PPARγ to the endogenous calreticulin promoter. Lane 1 shows input DNA, lane 2 is a negative control without antibody treatment, and lane 3 shows ChIP with antibodies.

Figure 5.

PPARγ activates the calreticulin gene. (A) A schematic representation of reporter plasmids used in this study. The calreticulin promoter (pLCC0) contains two PPRE sites: PPRE1 (−1,944 bp) and PPRE2 (−590 bp). Two promoter deletion constructs (pLC1 and pLC2) and two promoter constructs, one containing a mutation of PPRE1 (pLC0mt1) and the other containing a mutation of PPRE2 (pLC0mt2), were used to identify PPRE sites activated by PPARγ on the calreticulin promoter. (B) To analyze the effect of PPARγ and RXRα on the calreticulin promoter, NIH3T3 cells were transiently cotransfected with the PPARγ and RXRα expression plasmids with a luciferase reporter gene controlled by the calreticulin promoter as indicated in A. Individual controls were obtained for each expression plasmid, and data were plotted relative to that control (luciferase/β-galactosidase). The data shown are mean ± SD (error bars) of three independent experiments. (C) EMSA analysis was performed to demonstrate the binding of PPARγ to the PPRE1 and PPRE2 elements on calreticulin promoter. The positions of PPARγ–RXRα–DNA complexes are indicated. White lines indicate that intervening lanes have been spliced out. (D) Supershift EMSA analysis of PPARγ binding to the PPRE1 and PPRE2 elements on calreticulin promoter using anti-HA tag antibody was used to show the binding of transcriptional factor to the DNA element. Mutated PPRE1 and PPRE2 oligonuleotide show no interaction with PPARγ. Arrows mark PPARγ complexes with either PPRE1 or PPRE2. (E) ChIP analysis of a putative PPARγ-binding site in the mouse calreticulin promoter was used to show binding of PPARγ to the endogenous calreticulin promoter. Lane 1 shows input DNA, lane 2 is a negative control without antibody treatment, and lane 3 shows ChIP with antibodies.

To identify the PPARγ-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 PPARγ-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 PPARγ activation of the promoter (Fig. 5, B and C). Thus, upon induction of adipogenesis with RA, RXR and PPARγ 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 PPARγ, 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 PPARγ–RXRα complex bound to PPRE1 and PPRE2. This DNA–protein interaction was only observed in the presence of RXRα, indicating that PPARγ is only functional once complexed with RXRα (Fig. 5 C). The specificity of the PPRE and PPARγ–RXRα 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 PPARγ. ChIP analysis indicated that PPARγ bound to both PPRE sites 1 and 2 on the calreticulin promoter (Fig. 5 E). We concluded that the PPARγ–RXRα complex binds to calreticulin PPRE1 and PPRE2 sites and activates the calreticulin gene.

PPARγ2, along with C/EBPα, directly affect gene transcription in the nucleus (Rosen, 2005; Rosen and MacDougald, 2006). Given that PPARγ2 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), PPARγ2 was distinctly nuclear (Fig. 6 A). However, in the WT ES cells, PPARγ2 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/EBPα was also investigated in the ES cells, and it showed an essentially identical pattern to that observed for PPARγ2 (unpublished data).

Figure 6.

PPARγ2 localization in calreticulin-deficient G45 and calreticulin-expressing CGR8 ES cells. At D20 of differentiation, the cells were labeled for PPARγ2 and propidium iodide (PI) to identify the nuclei as described in Materials and methods. (A) In the calreticulin-deficient cells, PPARγ2 (green) colocalized with propidium iodide (red). (B) In the calreticulin-containing cells, labeling for PPARγ2 was barely detectable and perinuclear. Bar, 25 μm.

Figure 6.

PPARγ2 localization in calreticulin-deficient G45 and calreticulin-expressing CGR8 ES cells. At D20 of differentiation, the cells were labeled for PPARγ2 and propidium iodide (PI) to identify the nuclei as described in Materials and methods. (A) In the calreticulin-deficient cells, PPARγ2 (green) colocalized with propidium iodide (red). (B) In the calreticulin-containing cells, labeling for PPARγ2 was barely detectable and perinuclear. Bar, 25 μm.

Calmodulin/CaMKII, CREB, and calcineurin during adipogenesis

Calmodulin–Ca2+/calmodulin-dependent protein kinase II (CaMKII) and cAMP response element binding (CREB pathways affect adipogenesis (Wang et al., 1997; Reusch et al., 2000). Western blot analysis with anti-Thr286–phosphorylated CaMKII, anticalmodulin, and anti-Ser133–phosphorylated CREB antibodies showed that their expression was higher in calreticulin-deficient G45crt−/− and L7crt−/− cells than in WT and L7 cells at the end of the 20 d of differentiation (Fig. 7, A and B). These results indicate that CaMKII and CREB play a positive role during adipogenesis of crt−/− cells and that they are up-regulated in the absence of calreticulin.

Figure 7.

Calmodulin, CaMKII, and CREB expression is up-regulated in calreticulin-deficient ES cells. (A) The G45crt−/− and L7crt−/− cells had increased calmodulin, phosphorylated CaMKII (Thr286), and phosphorylated CREB (pSer133) expression when compared with WT and L7 cells. (B) Quantitative analysis of the expression of calmodulin, CaMKII, and CREB (P < 0.01; n = 6). (C) 10 μM KN-62 and KN-93 reduced oil red O staining in calreticulin-null (G45crt−/− and L7crt−/−) cells. Treatment of cells with KN-92, an inactive analogue of KN-93, had no effect on oil red O staining. (D) Colorimetric analysis showed a significant decrease in oil red O staining in calreticulin-null cells upon CaMKII inhibition (P < 0.01; n = 6). (E) Western blot analysis of PPARγ2, C/EBPα, and aP2 after inhibition of CaMKII showed lower expression of adipogenic markers than in the untreated cells. Treatment of cells with KN-92 had no effect on PPARγ2 and C/EBPα expression. (F) Quantitative analysis of the expression of PPARγ2, C/EBPα, and aP2 levels after CaMKII inhibition (P < 0.01; n = 6). Calcineurin activity is higher in WT ES cells than in calreticulin-deficient cells. (G) At D20, calcineurin activity is higher in WT ES cells than either in calreticulin-deficient (G45crt−/−) cells or undifferentiated ES cells at D0 (n = 4; P < 0.01). (H) Inhibition of calcineurin with cyclosporin A (CsA) increased oil red O staining in 3T3-L1 cells, whereas expression of constitutively active calcineurin decreased oil red O staining. (I) Colorimetric assay of oil red O staining in the 3T3-L1 cells treated as in H (n = 6). 3T3-L1, control; 3T3-L1 + CsA, CsA-treated preadipocytes; 3T3-L1 + CaN, 3T3-L1 cells overexpressing constitutively active calcineurin (CaN). Error bars represent SD.

Figure 7.

Calmodulin, CaMKII, and CREB expression is up-regulated in calreticulin-deficient ES cells. (A) The G45crt−/− and L7crt−/− cells had increased calmodulin, phosphorylated CaMKII (Thr286), and phosphorylated CREB (pSer133) expression when compared with WT and L7 cells. (B) Quantitative analysis of the expression of calmodulin, CaMKII, and CREB (P < 0.01; n = 6). (C) 10 μM KN-62 and KN-93 reduced oil red O staining in calreticulin-null (G45crt−/− and L7crt−/−) cells. Treatment of cells with KN-92, an inactive analogue of KN-93, had no effect on oil red O staining. (D) Colorimetric analysis showed a significant decrease in oil red O staining in calreticulin-null cells upon CaMKII inhibition (P < 0.01; n = 6). (E) Western blot analysis of PPARγ2, C/EBPα, and aP2 after inhibition of CaMKII showed lower expression of adipogenic markers than in the untreated cells. Treatment of cells with KN-92 had no effect on PPARγ2 and C/EBPα expression. (F) Quantitative analysis of the expression of PPARγ2, C/EBPα, and aP2 levels after CaMKII inhibition (P < 0.01; n = 6). Calcineurin activity is higher in WT ES cells than in calreticulin-deficient cells. (G) At D20, calcineurin activity is higher in WT ES cells than either in calreticulin-deficient (G45crt−/−) cells or undifferentiated ES cells at D0 (n = 4; P < 0.01). (H) Inhibition of calcineurin with cyclosporin A (CsA) increased oil red O staining in 3T3-L1 cells, whereas expression of constitutively active calcineurin decreased oil red O staining. (I) Colorimetric assay of oil red O staining in the 3T3-L1 cells treated as in H (n = 6). 3T3-L1, control; 3T3-L1 + CsA, CsA-treated preadipocytes; 3T3-L1 + CaN, 3T3-L1 cells overexpressing constitutively active calcineurin (CaN). Error bars represent SD.

To elucidate the role of the CaMKII pathway, we used specific inhibitors of CaMKII, KN-62 and KN-93 (Hidaka and Kobayashi, 1994). Inhibition of CaMKII in G45crt−/− and L7crt−/− cells with either KN-62 or KN-93 decreased adipogenesis (Fig. 7, C and D). Inhibition of CaMKII also resulted in the decreased expression of PPARγ2, C/EBPα, 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).

Discussion

In this study, we show that calreticulin affects adipocyte differentiation from either ES cells or 3T3-L1 preadipocytes. The absence of calreticulin sways ES cell differentiation toward the adipocyte lineage. This choice of fate is reversed to the WT phenotype by overexpression of calreticulin or by the Ca2+-buffering domain of calreticulin but not by expression of the chaperoning domain of the protein. These effects of calreticulin are likely mediated through its Ca2+ regulatory function. In comparison to WT cells, calreticulin-deficient ES cells have reduced ER Ca2+ storage capacity, resulting in reduced ER Ca2+ release upon stimulation. Expression of full-length calreticulin or the calreticulin Ca2+-buffering domain in ES cells or 3T3-L1 cells increases ER Ca2+ content and reduces the commitment of these cells to adipogenesis. In WT cells, chelating cytoplasmic [Ca2+] enhances adipogenesis, and, conversely, increasing cytoplasmic [Ca2+] inhibits adipogenesis. These Ca2+ effects may negatively affect Ca2+-dependent transcriptional pathways that are required for adipogenesis. Calreticulin exerts its Ca2+ effects within the first 3 d of differentiation, as manipulation of intracellular [Ca2+] at later stages of differentiation (i.e., D7 or D9) did not alter the respective adipogenic potential of ES cells (unpublished data). This is consistent with what has been done to elicit calcium-dependent effects during ES cell differentiation by Li et al. (2002), who used 100 nm ionomycin at D4, and Grey et al. (2005), who used 100 nm ionomycin at D6. Interestingly, Li et al. (2002) showed that ionomycin could be used at concentrations of 10–1,000 nM at days 4–7 with the same effects. Moreover, as calreticulin levels were increasing during the first 3 d of differentiation, PPARγ2 and C/EBPα 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 PPARγ2, C/EBPα, 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/EBPα regulation and, thus, indirectly in PPAR regulation. CaMKII activates CREB, which, in turn, activates C/EBPα (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 PPARγ (Paez-Pereda et al., 2005; Park et al., 2007), indicating that the CaMKII pathway may be critical during adipogenesis. Once C/EBPα is expressed, PPARγ and C/EBPα 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 PPARγ activity through a negative feedback mechanism

PPARγ2 and C/EBPα are crucial transcription factors during adipogenesis (Rosen, 2005); however, although PPARγ2 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 PPARγ2 and C/EBPα 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 PPARγ–RXR complex involves PPARγ transcriptional activation of the calreticulin gene. PPARγ is a potent transcriptional activator of the calreticulin gene as a result of its direct interaction with the calreticulin promoter. Thus, PPARγ can up-regulate calreticulin, and, conversely, calreticulin can inhibit its activity. We also found an inverse relationship between calreticulin and PPARγ2 expression upon induction of adipogenesis with RA. RA induces the expression of calreticulin but reduces the expression of PPARγ2. This supports the notion of a hierarchical process in which transcriptional activation of the calreticulin gene by PPARγ is the early event followed by calreticulin modulation of PPARγ transcriptional activity. These data provide evidence for a previously unrecognized negative feedback loop whereby PPARγ regulates the expression of calreticulin, and calreticulin modulates PPARγ2 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.

Materials And Methods

Cell culture and adipocyte differentiation

G45 crt−/− ES cells and the WT ES cell line (CGR8) were derived from J1 129/Sv mice. To generate the L7 ES cell line, J1 129/Sv ES cells were electroporated with a targeting vector containing the calreticulin gene with the loxP site inserted in introns 1 and 2 (Fig. S1 A) and maintained on mitomycin C–treated G-418–resistant mouse embryonic fibroblast feeder cells (Mesaeli et al., 1999). Recombinant clones were selected with 0.2 mg/ml G418 and 2 mM gancyclovir. Several hundred colonies were picked after 10 d in selection medium and expanded. The L7 clone was selected based on Southern blot analysis and Western blot analysis with anticalreticulin antibodies (Mesaeli et al., 1999). Cre recombinase with additional mutant estrogen receptor ligand-binding domains (Cre-ER and MerCreMer) was used to excise exons 2–4 from the calreticulin gene (Fig. S1 A). In brief, L7 ES cells were stably transfected with Cre recombinase mutant expression vector pANMerCreMER (a gift from J.D. Molkentin, University of Cincinnati, Cincinnati, OH) to generate the L7-Cre cell line. To generate L7crt−/−, 10 μM 4-hydroxytamoxifen was added to promote nuclear translocation and consequently activation of Cre recombinase. L7crt−/− contained a truncated and silenced calreticulin gene with interrupted reading frame and no expression of the protein (Fig. 1 C). Calreticulin-deficient ES (G45crt−/−) cell lines expressing N+P and P+C domains of the protein were also created and designated G45crt−/− N+P and G45crt−/− P+C.

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-rRxRα and pSG5-mPPARγ2 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), PPARγ, and RXRα 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-PPARγ2 (Sigma-Aldrich and Santa Cruz Biotechnology, Inc.) at 1:1,000 dilution; anticalreticulin (Opas et al., 1991) at 1:300 dilution; anti-C/EBPα (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 PPARγ, forward primer 5′-ATCAAGTTCAAACATATCACC-3′ and reverse primer 5′-TTGTCTTGGATGTCCTCGATG-3′; for C/EBPα, 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 RXRα and PPARγ 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 γ-[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-PPARγ 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 × 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 γ-[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-PPARγ2 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.

© 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/).

E. Szabo and Y. Qiu contributed equally to this paper.

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.

Acknowledgments

The superb technical help of Ewa Dziak is greatly appreciated. We thank Dr. L. Guo for generating the targeting vector with the calreticulin gene flanked by loxP and thank R. Rachubinski and J. Molkentin for plasmid DNA.

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.

References

References
Abdel-Ghany, M., K. el-Gendy, S. Zhang, and E. Racker.
1990
. Control of src kinase activity by activators, inhibitors, and substrate chaperones.
Proc. Natl. Acad. Sci. USA.
87
:
7061
–7065.
Baksh, S., and M. Michalak.
1991
. Expression of calreticulin in Escherichia coli and identification of its Ca2+ binding domains.
J. Biol. Chem.
266
:
21458
–21465.
Bedard, K., E. Szabo, M. Michalak, and M. Opas.
2005
. Cellular functions of endoplasmic reticulum chaperones calreticulin, calnexin, and ERp57.
Int. Rev. Cytol.
245
:
91
–121.
Benaim, G., and A. Villalobo.
2002
. Phosphorylation of calmodulin. Functional implications.
Eur. J. Biochem.
269
:
3619
–3631.
Bergling, S., R.E. Dolmetsch, R.S. Lewis, and J. Keizer.
1998
. A fluorometric method for estimating the calcium content of internal stores.
Cell Calcium.
23
:
251
–259.
Bradford, M.M.
1976
. A rapid and sensitive method for the quantitation of microgram quantities of protein utilizing the principle of protein-dye binding.
Anal. Biochem.
72
:
248
–254.
Burns, K., B. Duggan, E.A. Atkinson, K.S. Famulski, M. Nemer, R.C. Bleackley, and M. Michalak.
1994
. Modulation of gene expression by calreticulin binding to the glucocorticoid receptor.
Nature.
367
:
476
–480.
Clark, R.A., S.L. Li, D.W. Pearson, K.G. Leidal, J.R. Clark, G.M. Denning, R. Reddick, K.H. Krause, and A.J. Valente.
2002
. Regulation of calreticulin expression during induction of differentiation in human myeloid cells. Evidence for remodeling of the endoplasmic reticulum.
J. Biol. Chem.
277
:
32369
–32378.
Corti, C., L.E. Leclerc, M. Quadroni, H. Schmid, I. Durussel, J. Cox, H.P. Dainese, P. James, and E. Carafoli.
1999
. Tyrosine phosphorylation modulates the interaction of calmodulin with its target proteins.
Eur. J. Biochem.
262
:
790
–802.
Dani, C., A.G. Smith, S. Dessolin, P. Leroy, L. Staccini, P. Villageois, C. Darimont, and G. Ailhaud.
1997
. Differentiation of embryonic stem cells into adipocytes in vitro.
J. Cell Sci.
110
:
1279
–1285.
Dedhar, S., P.S. Rennie, M. Shago, C.-Y. Leung-Hagesteijn, H. Yang, J. Filmus, R.G. Hawley, N. Bruchovsky, H. Cheng, R.J. Matusik, and V. Giguère.
1994
. Inhibition of nuclear hormone receptor activity by calreticulin.
Nature.
367
:
480
–483.
Fadel, M.P., E. Dziak, C.M. Lo, J. Ferrier, N. Mesaeli, M. Michalak, and M. Opas.
1999
. Calreticulin affects focal contact-dependent but not close contact-dependent cell-substratum adhesion.
J. Biol. Chem.
274
:
15085
–15094.
Fadel, M.P., M. Szewczenko-Pawlikowski, P. Leclerc, E. Dziak, J.M. Symonds, O. Blaschuk, M. Michalak, and M. Opas.
2001
. Calreticulin affects beta-catenin associated pathways.
J. Biol. Chem.
276
:
27083
–27089.
Fajas, L., V. Egler, R. Reiter, J. Hansen, K. Kristiansen, M.B. Debril, S. Miard, and J. Auwerx.
2002
. The retinoblastoma-histone deacetylase 3 complex inhibits PPARgamma and adipocyte differentiation.
Dev. Cell.
3
:
903
–910.
Fowler, S.D., and P. Greenspan.
1985
. Application of Nile red, a fluorescent hydrophobic probe, for the detection of neutral lipid deposits in tissue sections: comparison with oil red O.
J. Histochem. Cytochem.
33
:
833
–836.
Fox, K.E., D.M. Fankell, P.F. Erickson, S.M. Majka, J.T. Crossno Jr., and D.J. Klemm.
2006
. Depletion of cAMP-response element-binding protein/ATF1 inhibits adipogenic conversion of 3T3-L1 cells ectopically expressing CCAAT/enhancer-binding protein (C/EBP) alpha, C/EBP beta, or PPAR gamma 2.
J. Biol. Chem.
281
:
40341
–40353.
Fruman, D.A., P.E. Mather, S.J. Burakoff, and B.E. Bierer.
1992
. Correlation of calcineurin phosphatase activity and programmed cell death in murine T cell hybridomas.
Eur. J. Immunol.
22
:
2513
–2517.
Fukami, Y., T. Nakamura, A. Nakayama, and T. Kanehisa.
1986
. Phosphorylation of tyrosine residues of calmodulin in Rous sarcoma virus-transformed cells.
Proc. Natl. Acad. Sci. USA.
83
:
4190
–4193.
Gesta, S., Y.H. Tseng, and C.R. Kahn.
2007
. Developmental origin of fat: tracking obesity to its source.
Cell.
131
:
242
–256.
Greenspan, P., E.P. Mayer, and S.D. Fowler.
1985
. Nile red: a selective fluorescent stain for intracellular lipid droplets.
J. Cell Biol.
100
:
965
–973.
Gregoire, F.M., C.M. Smas, and H.S. Sul.
1998
. Understanding adipocyte differentiation.
Physiol. Rev.
78
:
783
–809.
Grey, C., A. Mery, and M. Puceat.
2005
. Fine-tuning in Ca2+ homeostasis underlies progression of cardiomyopathy in myocytes derived from genetically modified embryonic stem cells.
Hum. Mol. Genet.
14
:
1367
–1377.
Guo, L., J. Lynch, K. Nakamura, L. Fliegel, H. Kasahara, S. Izumo, I. Komuro, L.B. Agellon, and M. Michalak.
2001
. COUP-TF1 antagonizes Nkx2.5-mediated activation of the calreticulin gene during cardiac development.
J. Biol. Chem.
276
:
2797
–2801.
Guo, L., K. Nakamura, J. Lynch, M. Opas, E.N. Olson, L.B. Agellon, and M. Michalak.
2002
. Cardiac-specific expression of calcineurin reverses embryonic lethality in calreticulin-deficient mouse.
J. Biol. Chem.
277
:
50776
–50779.
Hidaka, H., and R. Kobayashi.
1994
. Pharmacology of protein kinase inhibitors.
Essays Biochem.
28
:
73
–97.
Jensen, B., M.C. Farach-Carson, E. Kenaley, and K.A. Akanbi.
2004
. High extracellular calcium attenuates adipogenesis in 3T3-L1 preadipocytes.
Exp. Cell Res.
301
:
280
–292.
Kennell, J.A., and O.A. MacDougald.
2005
. Wnt signaling inhibits adipogenesis through beta-catenin-dependent and -independent mechanisms.
J. Biol. Chem.
280
:
24004
–24010.
Klemm, D.J., J.W. Leitner, P. Watson, A. Nesterova, J.E. Reusch, M.L. Goalstone, and B. Draznin.
2001
. Insulin-induced adipocyte differentiation. Activation of CREB rescues adipogenesis from the arrest caused by inhibition of prenylation.
J. Biol. Chem.
276
:
28430
–28435.
Li, J., M. Puceat, C. Perez-Terzic, A. Mery, K. Nakamura, M. Michalak, K.H. Krause, and M.E. Jaconi.
2002
. Calreticulin reveals a critical Ca2+ checkpoint in cardiac myofibrillogenesis.
J. Cell Biol.
158
:
103
–113.
Lynch, J., and M. Michalak.
2003
. Calreticulin is an upstream regulator of calcineurin.
Biochem. Biophys. Res. Commun.
311
:
1173
–1179.
Lynch, J., L. Guo, P. Gelebart, K. Chilibeck, J. Xu, J.D. Molkentin, L.B. Agellon, and M. Michalak.
2005
. Calreticulin signals upstream of calcineurin and MEF2C in a critical Ca2+-dependent signaling cascade.
J. Cell Biol.
170
:
37
–47.
Mery, L., N. Mesaeli, M. Michalak, M. Opas, D.P. Lew, and K.-H. Krause.
1996
. Overexpression of calreticulin increases intracellular Ca2+-storage and decreases store-operated Ca2+ influx.
J. Biol. Chem.
271
:
9332
–9339.
Mesaeli, N., K. Nakamura, E. Zvaritch, P. Dickie, E. Dziak, K.H. Krause, M. Opas, D.H. MacLennan, and M. Michalak.
1999
. Calreticulin is essential for cardiac development.
J. Cell Biol.
144
:
857
–868.
Meyer, T., P.I. Hanson, L. Stryer, and H. Schulman.
1992
. Calmodulin trapping by calcium-calmodulin-dependent protein kinase.
Science.
256
:
1199
–1202.
Nakamura, K., E. Bossy-Wetzel, K. Burns, M.P. Fadel, M. Lozyk, I.S. Goping, M. Opas, R.C. Bleackley, D.R. Green, and M. Michalak.
2000
. Changes in endoplasmic reticulum luminal environment affect cell sensitivity to apoptosis.
J. Cell Biol.
150
:
731
–740.
Nakamura, K., A. Zuppini, S. Arnaudeau, J. Lynch, I. Ahsan, R. Krause, S. Papp, H. De Smedt, J.B. Parys, W. Muller-Esterl, et al.
2001
. Functional specialization of calreticulin domains.
J. Cell Biol.
154
:
961
–972.
Neal, J.W., and N.A. Clipstone.
2002
. Calcineurin mediates the calcium-dependent inhibition of adipocyte differentiation in 3T3-L1 cells.
J. Biol. Chem.
277
:
49776
–49781.
Ntambi, J.M., and T. Takova.
1996
. Role of Ca2+ in the early stages of murine adipocyte differentiation as evidenced by calcium mobilizing agents.
Differentiation.
60
:
151
–158.
Opas, M., E. Dziak, L. Fliegel, and M. Michalak.
1991
. Regulation of expression and intracellular distribution of calreticulin, a major calcium binding protein of nonmuscle cells.
J. Cell. Physiol.
149
:
160
–171.
Otto, T.C., and M.D. Lane.
2005
. Adipose development: from stem cell to adipocyte.
Crit. Rev. Biochem. Mol. Biol.
40
:
229
–242.
Paez-Pereda, M., D. Giacomini, C. Echenique, G.K. Stalla, F. Holsboer, and E. Arzt.
2005
. Signaling processes in tumoral neuroendocrine pituitary cells as potential targets for therapeutic drugs.
Curr. Drug Targets Immune Endocr. Metabol. Disord.
5
:
259
–267.
Papp, S., M.P. Fadel, H. Kim, C.A. McCulloch, and M. Opas.
2007
. Calreticulin affects fibronectin-based cell-substratum adhesion via the regulation of c-src activity.
J. Biol. Chem.
282
:
16585
–16598.
Park, M.J., M.Y. Lee, J.H. Choi, H.K. Cho, Y.H. Choi, U.S. Yang, and J. Cheong.
2007
. Phosphorylation of the large subunit of replication factor C is associated with adipocyte differentiation.
FEBS J.
274
:
1235
–1245.
Puceat, M., and M. Jaconi.
2005
. Ca(2+) signalling in cardiogenesis.
Cell Calcium.
38
:
383
–389.
Reusch, J.E., and D.J. Klemm.
2002
. Inhibition of cAMP-response element-binding protein activity decreases protein kinase B/Akt expression in 3T3-L1 adipocytes and induces apoptosis.
J. Biol. Chem.
277
:
1426
–1432.
Reusch, J.E., L.A. Colton, and D.J. Klemm.
2000
. CREB activation induces adipogenesis in 3T3-L1 cells.
Mol. Cell. Biol.
20
:
1008
–1020.
Rosen, E.D.
2005
. The transcriptional basis of adipocyte development.
Prostaglandins Leukot. Essent. Fatty Acids.
73
:
31
–34.
Rosen, E.D., and O.A. MacDougald.
2006
. Adipocyte differentiation from the inside out.
Nat. Rev. Mol. Cell Biol.
7
:
885
–896.
Rosen, E.D., C.J. Walkey, P. Puigserver, and B.M. Spiegelman.
2000
. Transcriptional regulation of adipogenesis.
Genes Dev.
14
:
1293
–1307.
Rosen, E.D., C.H. Hsu, X. Wang, S. Sakai, M.W. Freeman, F.J. Gonzalez, and B.M. Spiegelman.
2002
. C/EBPalpha induces adipogenesis through PPARgamma: a unified pathway.
Genes Dev.
16
:
22
–26.
Serlachius, M., and L.C. Andersson.
2004
. Upregulated expression of stanniocalcin-1 during adipogenesis.
Exp. Cell Res.
296
:
256
–264.
Shi, H., Y.D. Halvorsen, P.N. Ellis, W.O. Wilkison, and M.B. Zemel.
2000
. Role of intracellular calcium in human adipocyte differentiation.
Physiol. Genomics.
3
:
75
–82.
Szabo, E., S. Papp, and M. Opas.
2007
. Differential calreticulin expression affects focal contacts via the calmodulin/Camk II pathway.
J. Cell. Physiol.
213
:
269
–277.
Tsien, R.Y.
1980
. New calcium indicators and buffers with high selectivity against magnesium and protons: design, synthesis, and properties of prototype structures.
Biochemistry.
19
:
2396
–2404.
Vankoningsloo, S., A. De Pauw, A. Houbion, S. Tejerina, C. Demazy, F. de Longueville, V. Bertholet, P. Renard, J. Remacle, P. Holvoet, M. Raes, and T. Arnould.
2006
. CREB activation induced by mitochondrial dysfunction triggers triglyceride accumulation in 3T3-L1 preadipocytes.
J. Cell Sci.
119
:
1266
–1282.
Wang, H., M.S. Goligorsky, and C.C. Malbon.
1997
. Temporal activation of Ca2+-calmodulin-sensitive protein kinase type II is obligate for adipogenesis.
J. Biol. Chem.
272
:
1817
–1821.
Wang, Z., B. Zhang, M. Wang, and B.I. Carr.
2003
. Persistent ERK phosphorylation negatively regulates cAMP response element-binding protein (CREB) activity via recruitment of CREB-binding protein to pp90RSK.
J. Biol. Chem.
278
:
11138
–11144.
Waser, M., N. Mesaeli, C. Spencer, and M. Michalak.
1997
. Regulation of calreticulin gene expression by calcium.
J. Cell Biol.
138
:
547
–557.
Winrow, C.J., K.S. Miyata, S.L. Marcus, K. Burns, M. Michalak, J.P. Capone, and R.A. Rachubinski.
1995
. Calreticulin modulates the in vitro DNA binding but not the in vivo transcriptional activation by peroxisome proliferator- activated receptor/retinoid X receptor heterodimers.
Mol. Cell. Endocrinol.
111
:
175
–179.
Zhang, J.W., D.J. Klemm, C. Vinson, and M.D. Lane.
2004
. Role of CREB in transcriptional regulation of CCAAT/enhancer-binding protein beta gene during adipogenesis.
J. Biol. Chem.
279
:
4471
–4478.
Zhang, L.L., L.D. Yan, L.Q. Ma, Z.D. Luo, T.B. Cao, J. Zhong, Z.C. Yan, L.J. Wang, Z.G. Zhao, S.J. Zhu, et al.
2007
. Activation of transient receptor potential vanilloid type-1 channel prevents adipogenesis and obesity.
Circ. Res.
100
:
1063
–1070.