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Am J Physiol Endocrinol Metab 292: E952-E963, 2007. First published November 28, 2006; doi:10.1152/ajpendo.00559.2006 Free Article
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Transgenic overexpression of protein targeting to glycogen markedly increases adipocytic glycogen storage in mice

Michael J. Jurczak, Arpad M. Danos, Victoria R. Rehrmann, Margaret B. Allison, Cynthia C. Greenberg, and Matthew J. Brady

Section of Endocrinology, Diabetes and Metabolism, Department of Medicine, and Committee on Molecular Metabolism and Nutrition, The University of Chicago, Chicago, Illinois

Submitted 13 October 2006 ; accepted in final form 26 November 2006


   ABSTRACT
TOP
 ABSTRACT
MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Adipocytes express the rate-limiting enzymes required for glycogen metabolism and increase glycogen synthesis in response to insulin. However, the physiological function of adipocytic glycogen in vivo is unclear, due in part to the low absolute levels and the apparent biophysical constraints of adipocyte morphology on glycogen accumulation. To further study the regulation of glycogen metabolism in adipose tissue, transgenic mice were generated that overexpressed the protein phosphatase-1 (PP1) glycogen-targeting subunit (PTG) driven by the adipocyte fatty acid binding protein (aP2) promoter. Exogenous PTG was detected in gonadal, perirenal, and brown fat depots, but it was not detected in any other tissue examined. PTG overexpression resulted in a modest redistribution of PP1 to glycogen particles, corresponding to a threefold increase in the glycogen synthase activity ratio. Glycogen synthase protein levels were also increased twofold, resulting in a combined greater than sixfold enhancement of basal glycogen synthase specific activity. Adipocytic glycogen levels were increased 200- to 400-fold in transgenic animals, and this increase was maintained to 1 yr of age. In contrast, lipid metabolism in transgenic adipose tissue was not significantly altered, as assessed by lipogenic rates, weight gain on normal or high-fat diets, or circulating free fatty acid levels after a fast. However, circulating and adipocytic leptin levels were doubled in transgenic animals, whereas adiponectin expression was unchanged. Cumulatively, these data indicate that murine adipocytes are capable of storing far higher levels of glycogen than previously reported. Furthermore, these results were obtained by overexpression of an endogenous adipocytic protein, suggesting that mechanisms may exist in vivo to maintain adipocytic glycogen storage at a physiological set point.

insulin; glycogen synthesis; lipogenesis; protein phosphatase-1; targeting subunit

CIRCULATING ENERGY SOURCES are maintained in a narrow homeostatic range across a wide variety of physiological conditions through the coordinate regulation of energy uptake, consumption, storage, and release by the principal metabolic tissues, namely adipose tissue, liver, and skeletal muscle. During times of energy deficit, adipose tissue is a primary site for energy provision via the hydrolysis of stored triglyceride to release free fatty acid (FFA) for ATP production and glycerol for hepatic gluconeogenesis. Conversely, following a meal, dietary lipid is stored by adipose tissue, and glucose is disposed of as glycogen by the skeletal muscle and to a lesser extent by the liver. Alternately, glucose can be utilized postprandially by the liver and adipocytes for de novo lipogenesis and long-term storage as triglyceride. However, adipocytes also store glucose as glycogen, albeit at substantially lower rates than in skeletal muscle and liver, so the physiological role of adipocytic glycogen metabolism remains unclear.

In addition to its role in lipid metabolism, adipose tissue functions as an endocrine organ through the secretion of a number of factors collectively known as adipokines. One such factor, leptin, was first identified as the single gene defect responsible for the obesity and hyperphagia seen in the ob/ob mouse (73), and has since been shown to act centrally at the hypothalamus to regulate feeding behavior (11, 62, 63) and peripherally to promote energy consumption (19, 48, 74). Circulating leptin levels are directly related to adiposity in both rodents and humans (1, 10, 26, 40), but they are also subject to regulation in response to feeding and fasting (32, 66), contributing to the daily cycle of serum leptin values (61). Although relatively little glucose is taken up by the fat cell compared with skeletal muscle or liver (5, 41), insulin-stimulated glucose metabolism synergistically regulates leptin secretion (45, 46, 56), possibly via transcriptional and/or posttranscriptional mechanisms (35, 44). However, the metabolic fate(s) of glucose within the adipocyte responsible for the regulated synthesis and release of leptin remains unclear.

Since the 1960s, investigators have sought to determine the major pathways of glucose metabolism within the fat cell. Experiments conducted with primary rat adipocytes demonstrated that 80–85% of glucose metabolized by the fat cell was oxidized to CO2 or converted to glycerol or fatty acid components of triglyceride, 5–15% was catabolized to lactate (12, 39), and ~2–3% was stored as glycogen (16, 43). Interestingly, at a constant insulin concentration, low concentrations of glucose were preferentially metabolized to glycogen (36), and glucose conversion to glycogen by the adipocyte was more sensitive to insulin than either CO2 production or triglyceride synthesis (43). These experiments led to the hypothesis that glucose entering the fat cell is first used to replenish glycogen stores, but this pathway is saturated quickly and glucose is subsequently shunted to triglyceride synthesis and ATP production (21, 43).

Glycogen metabolism is regulated by two key enzymes: glycogen synthase, which catalyzes the addition of glucose to the glycogen chain, and glycogen phosphorylase, which catalyzes the breakdown of glycogen to release glucose-1-phosphate. Insulin promotes glycogen synthesis through the coordinate dephosphorylation of glycogen synthase and phosphorylase, resulting in enzymatic activation and inactivation, respectively. Although the precise molecular mechanisms by which insulin regulates glycogen-metabolizing enzymes are very complex and remain controversial, the localized activation of protein phosphatase-1 (PP1) at glycogen particles plays an important role (8). Five proteins have been described that target protein phosphatase-1 (PP1) to glycogen chains (2, 13, 14, 47, 57, 64, 65). One such protein, termed protein targeting to glycogen (PTG), was first identified from 3T3-L1 adipocytes (57) and remains the only PP1 glycogen-targeting subunit reported to be expressed in adipocytes. Importantly, as reported by several groups, overexpression of PTG in a variety of cell types in vitro and in rodent liver in vivo markedly enhanced cellular glycogen levels (17, 20, 22, 24, 37, 50, 57, 72).

Adipose tissue has profound effects on whole body energy homeostasis through the regulated storage and mobilization of triglyceride and secretion of adipokines that influence feeding behavior and insulin sensitivity in other tissues. However, the potential importance of glycogen metabolism in adipocyte function has not received extensive attention. To further investigate the impact of altering adipocytic glycogen stores, we generated a transgenic mouse model overexpressing PTG in adipose tissue. We found that elevated PTG expression markedly increased glucose flux into the glycogen synthetic pathway, indicating that adipose tissue is capable of storing far higher levels of glycogen. In contrast, there were no detectable changes in triglyceride synthesis or mobilization in transgenic animals, indicating that adipocytic glycogen metabolism can be modulated in vivo independently of changes in lipid metabolism.


   MATERIALS AND METHODS
TOP
 ABSTRACT
 MATERIALS AND METHODS
RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Transgenic mouse generation. The PTG DNA sequence (Genbank U89924 [GenBank] ) was subcloned into a pBluescript II SK(+) vector containing a 5.4-kB adipocyte fatty acid binding protein (aP2) promoter (gift of Dr. Reed Graves), which had been used to obtain transgenic protein overexpression in adipose tissue (9). PTG was PCR-amplified to add an NH2-terminal FLAG-tag sequence (DYKDDDDK) and PvuII and SmaI restriction enzyme cut sites on the 5' and 3' ends, respectively. The PTG sequence was inserted downstream of the aP2 promoter using the SmaI restriction site. Blunt-end ligation of PTG downstream of the aP2 promoter eliminated the 5' PTG SmaI sequence. The 3' PTG SmaI site was retained and used to insert the poly(A) sequence (gift of Dr. Sally Radovick, Johns Hopkins University). The completed aP2-PTG-poly(A) construct was confirmed by sequencing. The pBluescript II SK(+) vector containing the PTG transgene was linearized with SalI and NotI, yielding a 7.4-kB construct (Fig. 1A) that was then microinjected into the pronuclei of CD-1 mice by the University of Chicago Diabetes Research and Training Center Transgenic/ES Cell Core. Potential founders from multiple litters were screened by Southern blotting (Fig. 1B) as previously described (71). Briefly, DNA was isolated from tail clips with a DNeasy kit (Qiagen) and then BamHI digested for resolution on agarose gel and transferred to Gene Screen Plus Hybridization Membrane (Perkin Elmer Life Sciences). Endogenous PTG is flanked by BamHI cut sites such that restriction digestion with BamHI produces a 7-kB fragment, whereas digestion of exogenous PTG yields a 3.7-kB fragment due to novel BamHI cut sites within the aP2 promoter and poly A tail. Bands were visualized with [32P]dCTP-labeled probe (American Radiolabeled Chemicals) homologous to the first 500 base pairs of the single PTG exon (Fig. 1B). Multiple founders were identified, and two males were selected for propagation of independent lines for characterization. DNA samples from tail clips of subsequent litters were screened by PCR with primers homologous to the FLAG insert (sense TAAGGTTCCTTCACAAAGATC) and poly A tail (antisense GACCTATAAAGACGACGACGACAAA) that amplified a 1-kB fragment, and with primers specific to the endogenous PTG exon (sense GCCATGAGGATTTGCTTG; antisense TCATTTGGGTGCCACCTG) that amplified an 800-bp fragment in all samples (data not shown).


Figure 1
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  		Fig. 1. Generation of the (PTG) driven by the adipocyte fatty acid binding protein (aP2)-protein targeting to glycogen (PTG) transgenic mouse line. A: transgene design showing introduction of novel BamHI (B) cut sites with the addition of the aP2 promoter and poly(A) tail on either end of FLAG (F)-tagged PTG. B: Southern blot analysis of genomic DNA from offspring of a transgenic male and wild-type female; DNA was digested with BamHI and hybridized with a probe homologous to the first 500 bp of PTG shown in A. Both endogenous (7-kb product) and exogenous (3.7-kb product) PTG exons were detected with this probe such that 1 band was detected for wild-type mice (Wt) and 2 bands were observed for transgenic mice (Tg).

 
Mouse treatment and care. The transgene was maintained on the CD-1 background throughout the study. CD-1 transgenic and wild-type mice from each line were allowed free access to a standard chow diet and water and maintained on a 12:12-h light-dark cycle under specific pathogen-free conditions in the Carlson Barrier Facility at the University of Chicago. All the procedures involving mice were approved by the University of Chicago Institutional Animal Care and Use Committee. After weaning at 3 wk of age, tail clippings were taken for genotyping by PCR, and mice were weighed weekly. At 2–3 and 6 mo of age, male mice were killed by CO2 narcosis for the studies described throughout. Brain, kidney, liver, skeletal and cardiac muscle, and gonadal, perirenal, and visceral fat were harvested, frozen on dry ice and stored at –80°C for future study. Blood was collected by cardiac puncture and centrifuged at 2,300 g, 4°C for 5 min to obtain serum.

For the high-fat-diet study, wild-type and transgenic 2-mo-old male mice were fed either the standard chow diet or a high-fat diet (45% of calories from fat; Research Diets) for 4 wk. Animals were weighed three times a week over the course of 4 wk, and food consumption was calculated over 3-day intervals for each of the 4 wk. Fed blood glucose was measured weekly by tail bleeds using an Ascensia Elite XL glucometer (Bayer). Glucose tolerance tests were performed at 2 and 4 wk after fasting by injecting 2 g/kg dextrose intraperitoneally. Glucose levels were measured from tail bleeds at 0, 15, 30, 60, and 120 min. After 4 wk, animals were killed, and tissue and serum were collected as described above. For fasting studies, animals were fasted overnight. Body weight and blood glucose were recorded pre- and postfast, and animals were killed for collection of serum and metabolic tissue as described above.

RNA and protein analysis. For real-time quantitative RT-PCR, RNA was isolated from 200 mg of gonadal fat of wild-type and transgenic mice using TRIzol, and RT-PCR was performed to produce cDNA with an iScript cDNA synthesis kit (Bio-Rad). Real-time quantitative RT-PCR was performed as previously described (23) with an iCycler and MyiQ software for data analysis (Bio-Rad) and primers for 18S (sense GCTGGAATTACCGCGGCT; antisense CGGCTACCACATCCAAGGA) or PTG (sense CCTTCCAGAAGAACCAGC; antisense CTCAGTTGGAATGACACG). Cycling parameters were as follows: 3 min at 95°C, followed by 40 cycles of 10 s at 95°C, 30 s at 60°C, and 30 s at 72°C. Glycogen synthase mRNA was measured as previously described (54).

For protein analysis, immunoblotting was performed as previously described (23). Briefly, tissues were lysed with a glass Dounce homogenizer in homogenization buffer [50 mM HEPES (pH 7.4), 150 mM NaCl, 1 mM EDTA, 10 mM NaF, 10% glycerol, and protease inhibitors] and then centrifuged at 500 g, 4°C for 5 min to obtain a postnuclear supernatant (PNS) fraction. Protein concentrations were determined by the Bradford method for normalization of protein loading. Immunoprecipitations were performed by incubating lysates with anti-FLAG monoclonal M2 antibodies coupled to agarose beads (Sigma) for 2 h at 4°C. Beads were washed three times with homogenization buffer and analyzed by immunoblotting (34) with anti-PTG antibodies (C-20 and N-19; Santa Cruz Biotechnologies). Insulin signaling experiments were conducted using primary adipocytes isolated from 2-mo-old animals as described below. Cells were incubated at 37°C in supplemented Krebs-Ringer bicarbonate-HEPES (KRBH) with or without 10 nM insulin for 10 min with shaking, placed on ice and then washed 3 times with KRBH before lysis.

For the fractionation experiments, the PNS fraction was centrifuged at 4°C, 100,000 g for 30–90 min to obtain a glycogen-enriched pellet fraction and cytosolic fraction. The pellet fractions were then resuspended in one-half the cytosolic volume of homogenization buffer with a 23-gauge needle for immunoblot analysis. Source of antibodies were the following: anti-glycogen phosphorylase and anti-phosphophosphorylase antibodies were generated and affinity purified as described (20); anti-glycogen synthase and anti-adiponectin were from Chemicon; anti-GLUT4 was from Alpha Diagnostics International; anti-phosphoglycogen synthase (Ser641), anti-phospho-Akt (Ser473), and anti-Akt were from Cell Signaling Technology; anti-pan PP1, anti-insulin receptor (IRbeta; C-19), and horseradish peroxidase-conjugated bovine anti-goat were from Santa Cruz Biotechnology; and anti-leptin was from Bio-Vendor (Candler, NC). Immunoblots were developed with the appropriate horseradish peroxidase-conjugated IgG (Bio-Rad) and enhanced chemiluminescence reagents (GE Healthcare Biosciences). Serum insulin and leptin were measured by Linco Diagnostic Services by radioimmunoassay.

Primary adipocyte preparation. Gonadal fat pads were minced in Krebs-Ringer buffer (3.5 ml/g wet weight) supplemented with 25 mM HEPES (pH 7.4), 4% bovine serum albumin, 5 mM glucose, 100 nM (–)-N6-(2-phenyl-isopropyl) adenosine, and 1 mg/ml type II collagenase (Sigma). Samples were incubated at 37°C for 40 min and mixed every 5 min. Isolated primary adipocytes were then transferred to 15-ml conical tubes and centrifuged at 150 rpm for 30 s. Liquid was removed from beneath the floating adipocyte layer, and the cells were washed three times with buffer.

Metabolic and enzymatic assays. PP1 and glycogen synthase activities were determined in vitro as previously described (24, 34) using primary adipocyte lysates. For glucose storage measurements, 100 µl of packed primary adipocytes from wild-type or transgenic mice were diluted with 100 µl KRBH containing 2 µCi D-[14C]glucose (MP Biomedicals) and 4 µg/ml adenosine deaminase with or without 10 nM insulin in triplicate. Cells were incubated at 37°C, 95% O2-5% CO2 for 60 min with shaking every 5 min. Following incubation, 90 µl of cells were added to 900 µl of PBS, and lipids were extracted with 4 ml Betafluor (National Diagnostics) for determination of lipogenesis rates. Ninety microliters of the remaining cells from each condition were added to 1 ml of 30% KOH to determine [14C]glucose incorporation into glycogen (24).

Glycogen and lipid determinations. Tissues were homogenized and glycogen was measured as described previously (24). Because of the large differences in adipocytic glycogen storage, ~0.6 mg of protein from gonadal fat pad lysate was spotted on 37-mm glass-fiber (GF/A) filters (Whatman) for transgenic mice and 2.0 mg for wild-type mice. Serum triglyceride and FFA were measured with a triglyceride determination kit (Sigma) according to the manufacturer's instructions.

Histology. Tissues were fixed overnight with a Carnoy's fixative (6:3:1; absolute ethanol-chloroform-glacial acetic acid) and embedded in paraffin and cross-sectioned by the Animal Resources Center at the University of Chicago. Deparaffined sections were stained with standard hematoxylin and eosin for general cell morphology and with a periodic acid Schiff (PAS) stain for glycogen. PAS staining was used to visualize neutral hexose sugars by first treating samples with periodic acid (HIO4·2H2O) to oxidize glycols to aldehydes. The aldehydes then reacted with and restored a pink color to a fuschin-sulfurous acid (Schiff reagent) dye, allowing for localization of polysaccharides such as glycogen.

Statistical analysis. The data shown represent means ± SE. Data were compared using unpaired, two-tailed Student's t-tests, and analyses were performed with Microsoft Excel XP. A P value of ≤0.05 was considered statistically significant.


   RESULTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
DISCUSSION
 GRANTS
 REFERENCES
 
aP2-driven overexpression of PTG in adipose tissue of transgenic mice. Previously, transient overexpression of the PP1 glycogen-targeting subunit PTG had been shown to increase intracellular glycogen levels in a number of cell types and rodent livers (17, 20, 22, 24, 37, 50, 57, 72). PTG, originally identified in the 3T3-L1 adipocyte cell line, is the only reported PP1 glycogen-targeting subunit expressed in adipose tissue (57). To investigate the role of glycogen metabolism in adipocyte function in vivo, PTG was subcloned downstream of the aP2 promoter for the generation of mouse lines overexpressing PTG in adipose tissue. Two independent lines were randomly chosen for further study. Litter sizes and offspring from transgenic matings appeared normal, and inheritance of the transgene from a heterozygous parent occurred at the expected Mendelian ratio.

Analysis of PTG mRNA by quantitative RT-PCR with RNA isolated from the epididymal fat pads of wild-type and transgenic mice demonstrated a 4.3-cycle decrease to threshold, corresponding to an ~20-fold increase in PTG mRNA expression (Fig. 2A). To determine the effects of increased PTG message on protein expression, lysates were prepared from epididymal fat pads for analysis by anti-PTG immunoblotting. PTG protein levels were readily detectable in adipocytic lysates from transgenic mice, whereas endogenous PTG was undetectable with the commercially available antibodies (Fig. 2B). Immunoprecipitation of adipocytic lysates using anti-FLAG antibody followed by anti-PTG immunoblotting confirmed that the increase in PTG protein was due to the overexpression of exogenous PTG (Fig. 2B). Exogenous PTG was also detected in perirenal and brown fat by Western blotting, but in no other tissue including brain, kidney, liver, and skeletal and cardiac muscle, confirming tissue-specific overexpression (data not shown). Similar results were obtained for each independent line regarding PTG mRNA, protein levels, and glycogen accumulation (see below) such that one line was selected for more extensive characterization.


Figure 2
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  		Fig. 2. Overexpression of PTG increases PTG mRNA and protein levels in adipose tissue. A: RNA was extracted from epididymal fat pads from wild-type and transgenic 2-mo-old male animals, and PTG levels were determined by real-time quantitative RT-PCR. The number of cycles needed to reach threshold are shown in the left panel, and the corresponding change in message levels is depicted in the right panel. Data represent the mean of 3 animals per genotype. B: gonadal fat pad lysates were prepared from 3 animals for each genotype, and subjected to anti-FLAG immunoprecipitation (IP) followed by anti({alpha})-PTG immunoblotting. Left panel: lysate before IP; right panel: IP pellet. Results are representative of 5 independent experiments.

 
Phenotypic analysis of aP2-PTG transgenic mice. Initial observations made in 2-, 3- and 6-mo-old wild-type and transgenic male animals revealed no striking differences in a number of broad parameters (Table 1). Similar observations were made for females (data not shown). There was no significant difference in body weight for wild-type and transgenic mice at 2–3 and 6 mo of age. Gonadal fat pad weight and food consumption were also approximately equal at both time points, as were blood glucose concentrations, serum triglyceride levels, and FFA values (Table 1). No evidence of abnormalities or change in tissue weight was observed during a gross anatomic survey of gonadal, perirenal, and visceral fat, liver, kidney, heart, and skeletal muscle performed during dissection. Additionally, no change in liver or skeletal muscle glycogen content was detected when measured directly or by PAS staining (data not shown). Lastly, primary adipocytes isolated from wild-type and transgenic animals were morphologically similar under light microscopy (Fig. 3A).


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  		Table 1. Characterization of wild-type and transgenic male mice

 

Figure 3
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  		Fig. 3. Glycogen storage is markedly increased in transgenic mice. A: primary adipocytes were isolated by collagenase digestion and diluted in Krebs-Ringer bicarbonate-HEPES (KRBH) for plating on silicone coated slides for viewing with a light microscope. B: glycogen storage was measured from gonadal fat pads from 2- and 6-mo-old wild-type and transgenic mice. Glycogen levels are expressed as micrograms of glycogen per milligram of protein. Data represent the mean ± SE of 3–4 and 4–8 mice for each genotype at 2 and 6 mo, respectively. C: histological analysis of gonadal fat pads from 2-mo-old animals was performed using periodic acid Schiff (PAS) staining for glycogen. Images shown are at x40 magnification and representative of observations made in 3 mice per genotype. ***P < 0.001 for wild-type vs. transgenic mice.

 
PTG overexpression markedly increases adipocytic glycogen storage. The effects of PTG overexpression on adipocytic glycogen stores were then examined. Epididymal fat pads were excised from 2- and 6-mo-old animals for determination of total glycogen storage. In wild-type mice, glycogen levels in adipose tissue were low, ranging from ~0.5 to 1.5 µg/mg protein (Fig. 3B). In parallel with the observed increases in PTG mRNA and protein levels, total glycogen storage was significantly increased in both lines of the transgenic mice, reaching levels as high as 200 µg/mg protein, or 100–400 times more glycogen than in wild-type mice (Fig. 3B). Changes in gonadal fat glycogen storage were equivalent in transgenic female mice (data not shown) such that future characterization focused on male animals.

The large increases in glycogen storage were surprising given the low glycogen levels normally present in adipose tissue (16, 43). In addition, glycogen is a spatially inefficient means of storing energy in that it comprises long branching, hydrophobic chains that require hydration by 3–4 g of water/g of glycogen. Furthermore, adipocytes possess a highly restricted intracellular space due to the densely packed lipid droplet. Thus, we investigated the localization of glycogen particles in the transgenic adipose tissue. Gonadal fat pads from 2-mo-old male wild-type and transgenic mice were excised and fixed overnight for embedding and cross-sectioning in paraffin. Sections were analyzed for general cell morphology (data not shown) and by PAS stain for visualization of intracellular glycogen. Two distinct differences in adipose tissue taken from transgenic mice were observed. First, PAS staining of glycogen in gonadal fat confirmed the marked increase in total cellular glycogen levels in transgenic fat pads, where glycogen was detected as a dark pink coloration that appeared at or near the plasma membrane (Fig. 3C). Second, adipocytes from transgenic mice appeared wrinkled, most likely resulting from the dehydration step of the fixation process, which would strip the water molecules hydrating the glycogen chains, presumably leading to the collapse of the plasma membranes.

Glycogen-targeted PP1 activity is elevated in transgenic mice. Glycogen synthesis and net glycogen storage represent the summation of a number of extracellular signals that regulate glucose uptake and metabolism and glycogen synthase and phosphorylase activities. Previous work demonstrated that bidirectional modulation of PTG expression in 3T3-L1 adipocytes specifically affected PP1 activity directed against enzymes involved in glycogen metabolism (23, 24). To initially investigate the mechanism by which PTG overexpression led to enhanced glycogen storage in the adipose tissue of transgenic mice, we determined PP1 protein levels, intracellular localization, and in vitro activity. Cell lysates were prepared from gonadal fat pads from three transgenic and three wild-type mice, and cellular levels were determined by immunoblotting. As shown in Fig. 4A, there was no detectable difference in PP1 protein expression between the two genotypes. In contrast, PP1 activity against 32P-labeled glycogen phosphorylase was elevated twofold in transgenic lysates (Fig. 4A), paralleling results obtained upon PTG overexpression in 3T3-L1 adipocytes (24).


Figure 4
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  		Fig. 4. Adipocytic protein phosphatase-1 (PP1) activity is increased in transgenic animals. A: PP1 activity was measured in vitro with gonadal fat pad lysates and 32P-labeled phosphorylase as substrate. Released [32P]phosphate levels in the supernatant were determined by scintillation counting. Activity measurements were made in triplicate for 4 wild-type and 8 transgenic mice, and data shown are means ± SE. PP1 protein levels were determined by anti-PP1 immunoblotting of gonadal fat pad lysates. 100 µg of cellular protein were analyzed. B: gonadal fat pad lysates from wild-type (Wt) and transgenic (Tg) animals were subjected to differential centrifugation to obtain a post-nuclear spin supernatant fraction (PNS), cytosolic supernatant fraction (Cytosol), and glycogen-enriched pellet fraction (GEP). The pellet fraction was then resuspended in homogenization buffer, using half the volume of the cytosolic supernatant fraction. Equal volumes of each fraction were then analyzed by anti-PP1 and anti-PTG immunoblotting. Data shown are representative of observations made in 3 mice per genotype. **P < 0.01.

 
To determine whether overexpression of PTG impacted the cellular distribution of PP1, homogenized gonadal fat pads were fractionated by differential centrifugation. Lysates were centrifuged at 500 g to obtain a PNS fraction, followed by ultracentrifugation at 100,000 g for up to 90 min to separate the cytosolic supernatant fraction from the crude glycogen-enriched pellet. Immunoblot analysis of each fraction demonstrated that PP1 levels in the PNS and cytosol were unchanged (Fig. 4B, top panel, lane 1 vs. 2 and 3 vs. 4), while PP1 present in the glycogen-enriched pellet was moderately increased in transgenic mice (Fig. 4B, top panel, lane 5 vs. 6). Furthermore, exogenous PTG was largely recovered in the glycogen-enriched pellet (Fig. 4B, top panel, lane 6), although detectable amounts were found in the cytosolic fraction. Potentially, a portion of PTG was not bound to glycogen because of high levels of protein overexpression or because it was released on cellular homogenization. Because other cellular substructures and organelles are present in the high-speed particulate fraction, the pellet fraction was treated with amylase to digest glycogen and centrifuged again at 100,000 g to identify glycogen-bound proteins released into the supernatant. Under these conditions, all of the PTG and most of the PP1 were released (data not shown), confirming that these proteins in the crude pellet were largely associated with glycogen. Unfortunately, we have exhausted the anti-PTG antibody that previously recognized glycogen-bound PTG in 3T3-L1 adipocytes (23), and commercially available antibodies were unable to detect endogenous adipocytic PTG even in the glycogen-containing fraction.

PTG overexpression increases both active and total glycogen synthase activities. An increase in glycogen-targeted PP1 activity could potentially alter cellular glycogen synthase and/or phosphorylase activities. First, glycogen synthase activity in adipocytic lysates from wild-type and transgenic mice was measured in vitro, in the presence and absence of the allosteric activator glucose-6-phosphate (G-6-P); activity measured in its absence is indicative of active glycogen synthase present within the cell, whereas activity measured in its presence corresponds to total glycogen synthase activity. In the absence of G-6-P, glycogen synthase activity was six times greater in transgenic than in wild-type mice (Fig. 5A), suggesting increased dephosphorylation and/or expression of glycogen synthase. In the presence of G-6-P, total glycogen synthase activity was also significantly increased in transgenic mice (Fig. 5A; P < 0.01), indicating an increase in enzyme protein levels. Immunoblotting of gonadal fat pad lysates confirmed that glycogen synthase levels were indeed elevated in transgenic mice (Fig. 5B). The apparent increase by immunoblotting was greater than results from in vitro activity assays (Fig. 5A vs. 5B). However, this apparent discrepancy may be explained by the finding that the anti-glycogen synthase antibody used recognized the dephosphorylated form of the enzyme more strongly (53). Thus the increase in immunoreactive protein is reflective of both elevated glycogen synthase protein levels and enhanced dephosphorylation on PTG overexpression in transgenic adipose tissue. The changes in glycogen synthase expression most likely result from enhanced protein stabilization secondary to increased glycogen levels, because quantitative real-time RT-PCR detected no significant difference in adipocytic glycogen synthase mRNA levels between the two genotypes (data not shown). There was also an increase in the amount of phosphoglycogen synthase detected in lysates from transgenic adipocytes (Fig. 5B). However, this modest change is most likely reflective of the larger increase in total enzyme levels resulting from PTG overexpression. Furthermore, the activity ratio for glycogen synthase was markedly increased (0.55 vs. 0.18, transgenic vs. wild type, respectively), corresponding to increased protein dephosphorylation.


Figure 5
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  		Fig. 5. PTG overexpression alters the subcellular distribution and activity of glycogen-metabolizing enzymes. A: glycogen synthase (GS) activity in gonadal fat pad lysates was measured in vitro. Activity was determined in the absence (active) and presence (total) of 10 mM glucose-6-phosphate. Each condition was performed in triplicate for 3 mice per genotype, and data shown are means ± SE. B: enzyme levels and phosphorylation state were determined by immunoblotting with anti-GS, anti-phospho-GS, anti-glycogen phosphorylase (GP), and anti-phospho-GP antibodies using gonadal fat pad lysates. One hundred micrograms of protein were analyzed for each sample. C: GS and GP distribution were determined by immunoblotting with gonadal fat pad lysates following the same centrifugation steps described in Fig. 4B. Immunoblotting data shown are representative of 3 independent determinations. **P < 0.01.

 
Next, the cellular distribution of glycogen synthase was determined by differential centrifugation of homogenized gonadal fat from wild-type and transgenic mice. Interestingly, overexpression of PTG led to a more pronounced redistribution of glycogen synthase than of PP1 within the cell (Fig. 5C, top panel vs. 4B, top panel). In wild-type mice, following the first 100,000 g centrifugation, a greater percentage of glycogen synthase was found in the cytosolic fraction than in the glycogen-containing pellet fraction (Fig. 5C, top panel, lane 3 vs. 5). In transgenic animals, however, the opposite was true, with most of the glycogen synthase detected in the glycogen-enriched pellet (Fig. 5C, top panel, lane 4 vs. 6). Thus, PTG overexpression enhanced glycogen synthase activity via direct and indirect mechanisms: dephosphorylation, increased protein expression, and intracellular redistribution of the enzyme, all of which would be expected to increase glycogen synthesis rates.

Glycogen phosphorylase protein levels and cellular distribution are modulated in transgenic mice. The effects of PTG overexpression on adipocytic glycogen phosphorylase were examined in parallel. In gonadal fat pad lysates from transgenic mice, glycogen phosphorylase protein levels were modestly elevated compared with wild type, although not as dramatically as glycogen synthase (Fig. 5B). Additionally, phosphophosphorylase levels increased in parallel with total enzyme levels. Despite these slight changes in glycogen phosphorylase protein levels and phosphorylation state, no significant difference in enzyme activity assayed in vitro was detected due to the extremely low enzymatic activities in adipocytic lysates from both genotypes (data not shown). Finally, phosphorylase distribution in wild-type and transgenic adipocytes was determined. Overexpression of PTG had a greater effect on phosphorylase localization compared with PP1 or glycogen synthase, as there was an almost complete redistribution of glycogen phosphorylase from the cytosolic to glycogen-enriched pellet fraction in transgenic adipocyte lysates (Fig. 5C, bottom panel). These data suggested that the dramatic increase in glycogen stores in transgenic adipose tissue resulted in a compensatory change in phosphorylase localization, which may have inhibited further glycogen accumulation.

Overexpression of PTG enhances partitioning of glucose into glycogen vs. lipid. The impact of markedly altering adipocytic glycogen storage on glucose flux into glycogen vs. lipid stores was measured. Primary adipocytes were incubated with [14C]glucose with or without insulin for 60 min and then divided into two equal aliquots, one for determination of glucose incorporation into glycogen and the other for incorporation into lipid. For both wild-type and transgenic animals, the majority of glucose stored under both basal and insulin-stimulated conditions was stored as lipid (97 and 98% wild type, 84 and 73% transgenic; Fig. 6A). These results are in stark contrast to glucose partitioning experiments in 3T3-L1 adipocytes, in which >85% of the glucose was stored as glycogen and <15% was stored as lipid (24). Overexpression of PTG significantly increased the amount of [14C]glucose recovered as glycogen in the absence and presence of insulin compared with wild-type controls (Fig. 6A; P < 0.01). Basal levels of glycogen synthesis were ~4 times greater in transgenic mice, and insulin-stimulated rates were elevated 16-fold. Interestingly, increasing glucose partitioning to glycogen appeared to reduce the amount of glucose being stored as lipid, although the difference was not significant compared with wild-type animals (Fig. 6A; P = 0.23 basal, P = 0.32 insulin). Furthermore, total glucose storage (calculated as the sum of glucose incorporated into glycogen and triglyceride) under these conditions was also unchanged.


Figure 6
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  		Fig. 6. Glycogen synthesis is increased in adipose tissue of transgenic mice. A: partitioning of glucose into glycogen and triglyceride was determined by incubating isolated primary adipocytes from wild-type and transgenic mice with [14C]glucose with or without 10 nM insulin for 60 min. Each aliquot of cells was then split in half for determination of glucose incorporation into glycogen and lipids in parallel. Each condition was investigated in triplicate, and data shown are means ± SE. Data shown are the average of 4 independent experiments. B: lysates were prepared from 3 animals of each genotype, and protein levels were determined by immunoblotting with anti-insulin receptor (IRbeta) and anti-GLUT4 antibodies. 100 µg protein were analyzed. For signaling studies, primary adipocytes from 2 animals for each genotype were incubated with (+) or without (–) 10 nM insulin for 10 min at 37°C. Immunoblots were probed with anti-phospho-Akt antibody, stripped, and then reprobed with anti-Akt antibody. **P < 0.01.

 
Glycogen synthesis and glucose conversion to triglyceride are both dependent on glucose transport into the adipocyte via the insulin-responsive glucose transporter, GLUT4. Immunoblot analyses of gonadal fat pad lysates revealed no change in IRbeta, GLUT4, or Akt protein levels (Fig. 6B). Furthermore, insulin treatment of primary adipocytes from wild-type and transgenic mice demonstrated no striking difference in basal or insulin stimulated phospho-Akt (Fig. 6B).

Adipocytic function is maintained in transgenic animals during fasting and high-fat feeding. Adipocytes store and release energy through the hormonal regulation of lipolysis and lipogenesis. To determine the effects of glycogen accumulation upon PTG overexpression on lipolysis and lipogenesis in vivo, two independent studies were conducted. First, 2-mo-old animals were fasted overnight or given free access to food. Both wild-type and transgenic fasted animals experienced an ~10% reduction in body weight, or a loss of 3.9 g and 4.0 g, respectively (Fig. 7A). Fed and fasted blood glucose, FFA, and insulin levels did not differ significantly between groups (Fig. 7, B and C), indicating that glycogen hyperaccumulation did not alter the adipocytic response to hormonal stimuli and mobilization of energy stores during a fast. Glycogen levels in the gonadal fat of transgenic animals dropped after overnight fasting (Fig. 7D), with the change approaching statistical significance, indicating that the adipocytic glycogen stores could be mobilized. In contrast, adenoviral-mediated overexpression of PTG in rat livers in vivo or cultured primary hepatocytes prevented glycogen mobilization in response to fasting or forskolin treatment (20, 50). This apparent disparity may reflect differences in cell type and/or potential compensatory changes in the transgenic adipocytes upon chronic PTG overexpression vs. acute upregulation in hepatocytes.


Figure 7
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  		Fig. 7. Fasting of wild-type and transgenic mice results in similar weight loss and changes in sera parameters. A: wild-type and transgenic male mice were fasted overnight, and they were weighed pre- and posttreatment. B: blood glucose was measured pre- and postfast using a glucometer, and blood was obtained from a tail bleed. C: circulating FFA (mM) and insulin (ng/ml) values were measured from sera as described in MATERIALS AND METHODS. D: gonadal fat pad glycogen content was measured in fasted and ad libitum-fed animals for each genotype, reported here as µg glycogen per mg protein. Values are means ± SE for 9 mice for each genotype.

 
In the second series of studies, 2-mo-old wild-type and transgenic mice were randomly chosen and raised on a high-fat or control chow diet for 4 wk. Both wild-type and transgenic mice on the high-fat diet gained ~25% of their initial body weight (Fig. 8A). Food consumption did not differ between wild-type and transgenic mice on either diet (Fig. 8B), and changes in blood glucose, FFA, and insulin were also similar (data not shown). Furthermore, glucose tolerance tests performed at the completion of the high-fat feeding demonstrated that both transgenic and wild-type animals developed glucose intolerance to a similar degree (Fig. 8C). Cumulatively, the in vitro and in vivo data indicated that the elevation of adipocytic glycogen stores in transgenic animals did not affect acute lipogenic rates, lipid accumulation during high-fat feeding, or mobilization of triglyceride by a prolonged fast, suggesting potential compartmentalization of energy storage as glycogen and lipid in adipose tissue.


Figure 8
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  		Fig. 8. High-fat feeding induces comparable weight gain and glucose intolerance in wild-type and transgenic mice. A: wild-type (Wt) and transgenic (Tg) mice were fed either a chow or high-fat diet for 4 wk, and changes in body weight were recorded every 3 days, reported here as weekly changes. Left panel, wild-type chow vs. wild-type high-fat; right panel, transgenic chow vs. transgenic high-fat. B: mice were maintained 3–4 animals per cage, and food was weighed every 3 days over 4 wk, reported here as grams consumed per mouse per day (g/m/d). C: glucose tolerance tests were performed after 4 wk of the dietary intervention as described in MATERIALS AND METHODS. Blood glucose values were significantly greater (P < 0.05) at all times points following injection for high-fat- vs. chow-fed animals for both genotypes. Values are means ± SE for 7–11 mice per genotype per condition. **P < 0.01. ***P < 0.001.

 
Increased circulating leptin levels in transgenic mice. Several lines of evidence suggest that postprandial changes in circulating insulin induce leptin synthesis and secretion (3, 7, 35). Furthermore, insulin-stimulated glucose metabolism synergistically increases leptin expression (45, 46, 70). To determine whether altering the partitioning of glucose between energy storage sites within the adipocyte impacted adipokine expression, gonadal fat pad lysates were analyzed by immunoblotting with anti-adiponectin and anti-leptin antibodies on the same membrane. The expression of the adiponectin was similar between the two mouse lines (Fig. 9A). In contrast, leptin protein content was increased two- to threefold in gonadal fat pad lysates from transgenic mice (Fig. 9A). To explore these changes further, serum leptin levels were measured by radioimmunoassay. In wild-type animals, leptin levels were 6 ng/ml in agreement with previously published reports ranging from 3–6 ng/ml (33, 51, 52, 58). Interestingly, a significant increase in circulating leptin levels in 2-mo-old male transgenic mice was observed (Fig. 9B; P < 0.01), in agreement with the immunoblotting results (Fig. 9A). These results indicate that enhancement of adipocytic glucose flux into glycogen exerted specific effects on the leptin expression profile, although the molecular mechanism underlying this change is unknown. Despite a doubling in circulating leptin levels in the transgenic animals, there was no detectable change in either food consumption or weight gain (Fig. 8, A and B), indicating that the leptin elevation in the transgenic animals was not profound enough to affect feeding behavior or energy utilization. Additionally, high-fat feeding resulted in a substantial and comparable increase in circulating leptin in both genotypes (Fig. 9C), indicating that the differences in basal leptin secretion were not preserved upon expansion of adipose mass.


Figure 9
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  		Fig. 9. Overexpression of PTG increases circulating leptin and protein levels in adipose tissue. A: samples were resolved by PAGE and transferred to nitrocellulose for anti-leptin and anti-adiponectin immunoblotting on the same membrane. One hundred micrograms of protein were analyzed. B: 2-mo-old wild-type and transgenic mice were killed, and blood was collected by cardiac puncture. Serum was isolated by centrifugation, and leptin was measured by radioimmunoassay. Values are means ± SE of 6–7 mice for each genotype. C: sera collected during the high-fat-diet study was analyzed for leptin content by radioimmunoassay. Values are means ± SE for 10–11 mice for each condition. **P < 0.01.

 

   DISCUSSION
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 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
GRANTS
 REFERENCES
 
While glycogen storage is an important energy reserve for skeletal muscle contraction and increased hepatic glucose output, its function in adipose tissue has not been widely addressed. In rodents, skeletal muscle and liver are the two primary sites of glycogen storage, where ~3 and 40 µg of glycogen are stored per milligram protein, respectively (30, 31, 38, 42, 49, 67, 68). In contrast, adipocytic glycogen storage in wild-type mice ranged from 0.5 to 1.5 µg/mg protein in this study. Because the total protein content of adipose tissue is also far less than skeletal muscle or liver, adipocytic glycogen represents a very small portion of whole body glycogen stores. Glycogen is composed of long branching chains of glucose monomers, which are a spatially inefficient means of storing energy, especially compared with the densely packed lipid droplet. Because the intracellular space of an adipocyte is highly restricted due to the presence of the large lipid droplet, biophysical restrictions were thought to be a major constraint on adipocytic glycogen accumulation. However, why the fat cell would retain the ability to synthesize glycogen is currently unknown, as was whether an adipocyte could support higher levels of glycogen storage.

In this study, the PP1 glycogen-targeting subunit PTG was overexpressed using the aP2 promoter in transgenic mice as a means of altering glycogen levels in adipose tissue. Previous work demonstrated that transient PTG overexpression in cell lines and rat liver in vivo markedly increased glycogen stores (4, 24, 37, 50), but this is the first report of a transgenic mouse line stably overexpressing PTG. At 6 mo of age, PTG overexpression in adipose tissue resulted in a 400-fold increase in glycogen levels, demonstrating a long-term alteration in glycogen metabolism resulting from the genetic manipulation of an endogenous adipocytic protein. Interestingly, glycogen supercompensation in skeletal muscle following exercise is associated with transient insulin resistance and reduced glycogen synthetic rates (28, 29), which persist until glycogen levels have returned to a physiological set point. In contrast, no major changes in insulin signaling in primary adipocytes from transgenic mice were observed. Also, acute rates of basal and insulin-stimulated glycogen synthesis were elevated in primary adipocytes from transgenic animals, and glycogen breakdown in response to an overnight fast was detected, indicating that glycogen stores had not reached a static ceiling, but were instead dynamically regulated and that a new physiological set point may have been established.

Histological examination demonstrated that the transgenic adipocytes accommodated the enhanced glycogen levels through deposition at the outer edges of the cell, without altering the normal rounded appearance of an intact adipocyte. Several observations from this study support the conclusion that glycogen hyperaccumulation did not impair general adipocyte function. First, transgenic animals were born in the expected Mendelian ratios, developed and gained weight normally, possessed the same fat mass as wild-type controls and showed no changes in circulating glucose, triglyceride, or FFA levels. High-fat-diet and fasting studies designed to specifically challenge the adipocyte demonstrated that lipogenesis and lipolysis were not affected in vivo. Furthermore, partitioning experiments with radiolabeled glucose showed that acute glucose flux into triglyceride, as well as total glucose storage, were not impacted by increased flux of glucose into glycogen, potentially suggesting distinct pathways of glucose metabolism into lipid and glycogen.

To address the mechanism by which PTG overexpression led to glycogen accumulation, analyses of protein levels, localization, and activity states of the relevant glycogen regulatory enzymes were conducted. PTG overexpression did not affect PP1 expression but caused a moderate increase in PP1 levels bound to glycogen, which in turn correlated with an increase in PP1 activity directed against glycogen phosphorylase in vitro. Additionally, basal and insulin-stimulated glycogen synthase activity ratios were persistently elevated in transgenic adipose tissue, reflective of increased PP1-mediated dephosphorylation of glycogen synthase. Chronically elevated glycogen stores led to a secondary increase in glycogen synthase levels, presumably through increased protein stabilization because mRNA levels were unchanged. In contrast, expression of glycogen phosphorylase was much more modestly altered in the transgenic adipose tissue. PTG overexpression also increased the percentage of cellular glycogen synthase and phosphorylase recovered in the glycogen-enriched pellet fractions, most likely again a secondary effect of the enhanced glycogen stores. Transient overexpression of PTG in 3T3-L1 adipocytes also increased glycogen synthase dephosphorylation and activity but without any change in total enzyme levels (24). The differences may be due to the 2- to 3-day overexpression in cells vs. the 2-mo overexpression in the animals. Additionally, knockdown of endogenous PTG in 3T3-L1 adipocytes with short hairpin RNA revealed that glycogen phosphorylase rather than glycogen synthase is the principal cellular substrate of the PTG-PP1 complex (23). Thus, the alterations in glycogen synthase dephosphorylation in the transgenic animals may not be fully reflective of endogenous PTG action in adipose tissue in vivo.

Overexpression of PTG resulting in elevated adipocytic glycogen storage occurred in conjunction with a twofold increase in circulating leptin. The effect appeared specific, because leptin protein levels in gonadal fat were also increased and adiponectin did not change. Long-term circulating leptin levels have been shown to be directly proportional to adiposity in both humans and rodents (18, 40), whereas the acute mechanisms responsible for the regulated synthesis and secretion during fasting and refeeding are more complex. Several lines of study have demonstrated that insulin regulates leptin via posttranscriptional mechanisms, including increased synthesis and secretion, as well as by modulating tissue storage and turnover (3, 7, 35, 59). Work directed at determining the nutrient or signal responsible for the regulation of leptin in response to feeding has implicated the insulin-stimulated metabolism of glucose, as opposed to a direct effect of insulin (6, 15, 45, 46). Because fat pad masses and body weight were unchanged in transgenic mice, the observed differences in leptin levels were more likely due to changes in glucose flux into the adipocyte than to alterations in adiposity. It is interesting that gonadal fat pad leptin content was elevated in transgenic mice, suggesting increased biosynthesis, but testing this supposition will require future study.

Despite the observed increase in circulating leptin, no difference in feeding behavior or body weight in the transgenic mice was found. In previous work, daily injections of recombinant leptin for 28 days produced no significant changes in body weight, adiposity, or blood glucose or insulin levels in wild-type mice, even at the highest dosage (55). Only when leptin was administered by constant infusion for 6 days, or as twice daily, high-dosage injections, was a modest but significant weight loss observed in wild-type mice (25, 55). In studies where transgenic overexpression of the ob gene resulted in a lean phenotype, circulating leptin levels were as high as 30, 60 and 100 ng/ml (33, 51, 52), suggesting that the increases observed here (11.4 ± 1.2 ng/ml) were too modest to have a profound effect on feeding behavior.

Although the results presented demonstrate that adipose tissue can sustain far higher levels of glycogen that normally seen in vivo, the physiological role of endogenous glycogen in adipocyte function remains uncertain. During fasting, it has been estimated that ~30–40% of the FFA released from triglyceride is retained and recycled back to triglyceride in adipose tissue from rodents and humans (27, 69). This reesterification is dependent on a supply of glycerol-3-phosphate, either from glucose via glycolysis or from pyruvate via an often overlooked pathway, termed glyceroneogenesis (reviewed in Ref. 60). The onset of glyceroneogenesis in adipose tissue seems to occur later during fasting as a break on FFA release and hence the onset of ketogenesis by the liver. Potentially, the earlier production of glycerol-3-phosphate for reesterification of FFA during fasting could be derived from intracellular glucose or glucose released from glycogen. Thus, increasing adipocytic glycogen storage might be expected to increase FFA recycling and reduce the amount of FFA released during a fast. However, despite markedly increased cellular levels of glycogen in transgenic adipose tissue and subsequent mobilization during fasting conditions, no difference in circulating FFA levels were detected between the two genotypes following a prolonged fast. Further studies are required to examine the precise ratio of glycerol to FFA released from transgenic vs. control adipocytes in vitro, and the cellular fate of the glucose-1-phosphate released from glycogen to fully address this potential scenario. A second hypothesis regarding the function of adipocytic glycogen storage was that glycogen might serve as a short-term store of three carbon precursors for the correct pairing of one molecule of glycerol-3-phosphate and three FFAs for triglyceride synthesis during lipogenesis. Thus, by enhancing the available pool of glycogen and the supply of FFA, triglyceride synthesis might be expected to increase in the described transgenic model. However, this was not the case because wild-type and transgenic animals raised on a high-fat diet gained the same amount of total weight and fat mass, and acute basal and insulin-stimulated lipogenic rates were similar.

In summary, overexpression of PTG in adipose tissues resulted in a dramatic increase in glycogen stores that persisted for up to 1 yr. It is important to note that the transgenic lines in this study were generated by overexpression of an endogenous adipocytic protein. Thus, these results suggest that PTG expression may be regulated in adipocytes to ensure the relatively low glycogen levels observed in wild-type adipose tissue. These data further demonstrate that adipocytes are physically capable of storing far greater levels of glycogen than normally occurs in vivo, and surprisingly that adipocytes are able to function normally despite this significant metabolic change. Therefore, it appears that the required glycogen stores in adipose tissue are low and that further accumulation did not significantly impact adipose tissue development or function. To conclusively determine the physiological role of adipocytic glycogen storage, future studies that employ gene-silencing technologies to disrupt adipocytic glycogen synthesis would be required.


   GRANTS
TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
REFERENCES
 
This work was supported by a Career Development Award from the American Diabetes Association, National Institutes of Health (NIH) Grant R01 DK-064772 and a Pilot and Feasibility Award from the University of Chicago Diabetes Research and Training Center (to M. J. Brady), and a predoctoral fellowship from NIH Research Training Grant in Digestive Diseases and Nutrition T32 DK-07074 (to M. J. Jurczak). Work done through the University of Chicago Transgenic/ES Cell Core was supported by the Diabetes Research Training Center (P60 DK-20595).


   ACKNOWLEDGMENTS
 
We thank Linda Degenstein for assistance and advice throughout the generation of the transgenic line and Kevin Gibbs for excellent technical assistance and management of the mouse colony.


   FOOTNOTES
 

Address for reprint requests and other correspondence: M. J. Brady, Dept. of Medicine, Univ. of Chicago, MC1027, 5841 S. Maryland Ave., Chicago, IL 60637 (e-mail: mbrady{at}medicine.bsd.uchicago.edu)

The costs of publication of this article were defrayed in part by the payment of page charges. The article must therefore be hereby marked "advertisement" in accordance with 18 U.S.C. Section 1734 solely to indicate this fact.


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