Adipose tissue gene expression profiling reveals distinct molecular pathways that define visceral adiposity in offspring of maternal protein-restricted rats

Haiyan Guan,1 Edith Arany,2 Jonathan P. van Beek,1 Astrid Chamson-Reig,2 Sandra Thyssen,2 David J. Hill,2,3 and Kaiping Yang1,3

Departments of 1Obstetrics and Gynaecology 2Medicine 3Physiology and Pharmacology, Canadian Institutes of Health Research Group in Fetal and Neonatal Health and Development, Children's Health Research Institute and Lawson Health Research Institute, University of Western Ontario, London, Ontario, Canada

Submitted 30 September 2004 ; accepted in final form 19 November 2004


    ABSTRACT
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
There is increasing evidence that poor early growth confers an increased risk of type 2 diabetes, hypertension, and other features of the metabolic syndrome in later life. We hypothesized that this may result from poor nutrition during early life exerting permanent effects on the structure and function of key metabolic organ systems. To study the long-term impact of early-life undernutrition on susceptibility to visceral adiposity, we used a rat model of maternal protein restriction (MPR) in which dams were fed a low-protein diet (containing 8% instead of 20% protein in control diet) throughout pregnancy and lactation. MPR offspring were born smaller than controls (offspring of dams on control diet) and in adulthood developed visceral adiposity. We compared the pattern of gene expression in visceral adipose tissue (VAT) between MPR offspring and controls with Affymetrix rat expression arrays. Of the total number of genes and expressed sequence tags analyzed (15,923 probe sets), 9,790 (61.5%) were expressed in VAT. We identified 650 transcripts as differentially expressed ≥1.5-fold in the VAT of MPR offspring. Gene ontology analysis revealed a global upregulation of genes involved in carbohydrate, lipid, and protein metabolism. A number of genes involved in adipocyte differentiation, angiogenesis, and extracellular matrix remodeling were also upregulated. However, in marked contrast to other rodent models of obesity, the expression of a large number of genes associated with inflammation was reduced in this rat model. Thus visceral adiposity in this early-life programmed rat model is marked by dynamic changes in the transcriptional profile of VAT. Our data provide new insights into the molecular mechanisms that underlie the early-life programming of visceral adiposity.

visceral adipose tissue; maternal protein restriction; DNA microarray


OBESITY IS A SERIOUS MEDICAL PROBLEM not only because it substantially impairs quality of life but also because it increases the risk of hypertension, type 2 diabetes, coronary heart disease, sleeping disorders, and cancers (43). There is strong evidence for a genetic component to human obesity (28). Multiple systems regulate energy homeostasis (35, 44), and a number of genes associated with human obesity have been identified (19), yet the genetic component of this condition cannot explain the dramatic increase in the prevalence of obesity in recent years.

A large number of epidemiological studies have revealed a strong statistical association between poor fetal growth and the subsequent development of type 2 diabetes, hypertension, and obesity, visceral obesity in particular (54). These observations were made initially by Barker et al. in England (3a) but have now been reproduced in a large number of populations worldwide. These findings have led to the "fetal origins" hypothesis, which states that an adverse intrauterine environment programs or imprints the development of fetal tissues, permanently determining physiological responses and ultimately producing dysfunction and disease (60). However, the molecular mechanisms that underpin this relationship remain elusive.

In an attempt to provide a conceptual framework to begin to explain these observations, the thrifty-phenotype hypothesis has been proposed (24), which postulates that fetal development is sensitive to the nutritional environment. When it is poor, an adaptive response is initiated to optimize the growth of certain organs, such as the brain, at the expense of others, such as peripheral organs. These adaptations serve to improve chances of fetal survival. They also lead to an altered postnatal metabolism, which enhances postnatal survival under conditions of intermittent and poor nutrition. However, these adaptations would become detrimental if postnatal nutrition were adequate or overabundant. The detrimental consequences may include increased risk of developing obesity and the associated metabolic diseases. Indeed, epidemiological studies in the human have suggested that adults who were growth restricted in utero (occurs in 5–10% pregnancies) have increased body fat and in particular increased visceral fat (52, 61).

A wide variety of animal models have been established to test the fetal origins hypothesis (55). Among them, the maternal protein restriction (MPR) rat model has been used extensively to study the long-term impact of an adverse intrauterine environment on susceptibility to hypertension, insulin resistance, type 2 diabetes, and other metabolic diseases (55). In this model, rat dams are subjected to a low-protein diet containing 8% instead of 20% protein (control diet) throughout pregnancy and lactation. The MPR offspring are known to exhibit low birth weight, the characteristic feature of intrauterine growth restriction, and become diabetic, insulin resistant, and hypertensive in adulthood (56). Other models of fetal programming also predispose to the development of these metabolic disorders, including maternal caloric restriction, uterine artery ligation, and excessive fetal glucocorticoid exposure (55). It is remarkable that these different insults in fetal life produce the same detrimental consequences that occur in adulthood, suggesting that a common mechanism may underlie the early-life programming of the adult diseases.

However, few studies have focused on the long-term impact of an adverse intrauterine environment on the susceptibility to obesity (57) and none on the early-life programming of adipose tissue gene expression. Therefore, the aims of the present study were to determine, using the MPR rat model, 1) whether poor early nutrition led to the development of visceral adiposity and 2) whether the phenotypic changes in fat mass were associated with alterations in the transcriptional profile of visceral adipose tissue (VAT).


    MATERIALS AND METHODS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Animals and dietary manipulations. Virgin female Wistar rats (initial weight 240–260 g) were purchased from Charles River Laboratories (Wilmington, MA) and bred in our animal care facility (Lawson Health Research Institute). They were housed individually and maintained at 22°C on a 12:12-h light-dark cycle. They were mated, and day 0 of gestation was set as the day on which vaginal plugs were expelled. Pregnant rats were fed a diet containing 20% protein (control) or an isocaloric diet containing 8% protein (low protein) throughout pregnancy and lactation. The composition and source of the diets are described in detail in Table 1. There were no differences in litter size or sex ratio between the control and protein-restricted groups. Litters from four control and four protein-restricted dams were followed in this study. At 3 days of age, litters were randomly reduced to eight pups, thus ensuring a standard litter size per dam. At 21 days of age, all pups were weaned onto a 20% protein diet. For simplicity, the two groups of offspring will be termed control and MPR rats. For consistency, only male offspring were used for the study because early-life programming is known to occur in a sexually dimorphic manner (41), which was not the focus of this study. At 130 days of age, male offspring were killed and their visceral fat pads (composed of mesenteric, omental, and retroperitoneal fat masses) isolated, weighed, frozen on dry ice, and stored at –80°C until use. This age was chosen because these animals were monitored for a separate study, after weaning, for their status of insulin homeostasis and found to exhibit insulin resistance for the first time (data not shown).


View this table:
[in this window]
[in a new window]
 
Table 1. Composition of the diets

 
Histology. Paraffin-embedded sections (5 µm) of fat pad from control and MPR rats were stained with Oil red O. Sections were examined under a standard microscope, and the photomicrographs were captured at x20 magnification.

RNA extraction. Total RNA was extracted from homogenized adipose tissues with TRIzol (Invitrogen Life Technologies, Burlington, ON, Canada) and subsequently purified by RNeasy Mini Kit (Qiagen, Mississauga, ON, Canada) coupled with on-column DNase digestion with the RNase-Free DNase Set (Qiagen) according to the manufacturer's instructions.

DNA microarray and data analysis. A total of four arrays (Affymetrix Rat Expression Array RAE230A; Affymetrix, Santa Clara, CA) were conducted using total RNA samples from two control and two MPR rats. Microarrays were performed at the London Regional Genomics Centre (London, ON, Canada) following the standard procedures as outlined in the Affymetrix GeneChip Expression Analysis Technical Manual. The complete data set was submitted to the National Center for Biotechnology Information's Gene Expression Omnibus (GEO; http://www.ncbi.nlm.nih.gov/geo/) database (acc. no. GSE1813 [NCBI GEO] ).

Expression values for the Affymetrix GeneChip data were globally normalized to a preset value and analyzed using GeneSpring version 5.0 software (Silicon Genetics, Redwood City, CA). Values for the mean expression level for each gene were calculated for the control and MPR microarray data sets. Candidate genes were selected by a combination of absolute analysis (the absolute call was present on one of the two control arrays and on one of the two MPR arrays for genes considered to exhibit decreased and increased expression, respectively) and comparison analysis (displayed >1.5-fold increase or decrease from the control to MPR animals). A total of 15,923 data sets were sorted according to the following stringent criteria. To reduce false positives, data sets were excluded if 1) the raw values displayed >10-fold difference between the two data sets in either group (17 datasets were excluded); 2) one of the raw values in the two MPR datasets was less than the mean raw value of the two control data sets for genes exhibiting increased expression (15 data sets were excluded); and 3) one of the raw values in the two control data sets was less than the mean raw value of the MPR data sets for genes exhibiting decreased expression (14 data sets were excluded). This resulted in 650 genes being identified as differentially expressed in MPR offspring compared with the controls.

Real-time quantitative RT-PCR. Real-time quantitative RT-PCR (qRT-PCR) was used to study the expression of seven representative genes identified as differentially expressed (displayed variable degrees of change ranging from –4.7- to +3.3-fold) by the DNA microarray. In addition, transcript levels of two genes that displayed no change in expression with DNA microarray were also determined. Another important consideration for choosing these genes for qRT-PCR was their established role in adipogenesis, angiogenesis, or obesity. qRT-PCR was performed on the same total RNA samples as those used in the DNA microarray study and on additional RNA samples from four control and four MPR rats. Thus a total of six RNA samples per group were subjected to a two-step customer-designed SYBR Green I chemistry-based qRT-PCR, as described previously (71).

Briefly, 1 µg of total RNA was reverse transcribed in a total volume of 20 µl with the High Capacity Complementary Deoxyribonucleic Acid (cDNA) Archive Kit (Applied Biosystems), following the manufacturer's instructions. For every RT reaction set, one RNA sample was set up without reverse transcriptase enzyme to provide a negative control. Gene-specific primers (detailed in Table 2) were designed using Primer Express Software (Applied Biosystems, Foster City, CA), and the optimal concentrations for each gene were determined empirically. All primers were purchased from Sigma Genosys (Oakville, ON, Canada). The SYBR Green I assay was performed with the SYBR Green PCR Master Mix (Applied Biosystems) and a modified universal thermal cycling condition (2 min at 50°C and 10 min at 95°C, followed by 40 cycles of 10 s each at 95, 60, and 72°C) with the standard disassociation/melting parameters (15 s each at 95, 60, and 95°C) on the ABI Prism 7900HT Sequence Detection System (Applied Biosystems). The specificity of the SYBR Green I assay was verified by performing a melting curve analysis and by subsequent sequencing of the PCR products.


View this table:
[in this window]
[in a new window]
 
Table 2. Primer sequences for quantitative RT-PCR

 
Levels of 28S rRNA (housekeeping gene) and target mRNAs in each RNA sample were quantified by the relative standard curve method (Applied Biosystems). Briefly, standard curves for 28S rRNA and each target gene were generated by performing a dilution series of a mixed cDNA pool. For each RNA sample, the amount of target mRNA relative to that of 28S rRNA was obtained. For each target gene, fold changes in the MPR group compared with the control were then calculated and expressed as means ± SE. Data were analyzed by a standard t-test, and significance was set at P < 0.05. All calculations were performed using SPSS software version 9.0 (Chicago, IL).


    RESULTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
MPR leads to growth restriction and visceral adiposity. The male offspring of dams fed a low-protein (8%) diet were significantly smaller at birth than offspring of control dams fed a diet containing 20% protein (5.01 ± 0.15 vs. 6.40 ± 0.15 g, P < 0.01). At 130 days of age, the mean body weight of MPR male offspring was slightly lower than that of the controls, but the weight of visceral fat mass as well as the ratio of visceral fat to body weight were significantly higher (Fig. 1), indicative of visceral adiposity. As shown in Fig. 2, the sizes of adipocytes were similar between control and MPR rats, suggesting that visceral adiposity in this rat model was primarily a result of hyperplasia.



View larger version (10K):
[in this window]
[in a new window]
 
Fig. 1. Effects of maternal protein restriction (MPR) on body weight (A), visceral fat weight (B), and the ratio of visceral fat to body weight (C) of male offspring rats at 130 days of age. Data are expressed as means ± SE; n = 6–9 rats. *P < 0.05 and **P < 0.01 compared with controls (male offspring of rat dams on control diet).

 


View larger version (119K):
[in this window]
[in a new window]
 
Fig. 2. Fat pad histology. Paraffin-embedded sections of fat pad from control and MPR rats at 130 days of age were stained with Oil red O. Representative photomicrographs are shown.

 
Overview of changes in VAT gene expression. To determine whether MPR alters the transcriptional profile of VAT and to gain insight into the identity of genes and molecular pathways involved in the pathogenesis of visceral adiposity, DNA microarray experiments were conducted on VAT derived from MPR and control rats. Of the 15,923 genes and expressed sequence tags (ESTs) analyzed (hereafter referred to as genes), 9,790 (61.5%) were expressed at sufficient levels for detection in VAT. MPR altered the expression of 650 genes (>1.5-fold induction or repression) in VAT. Of these, 376 are known genes, including ESTs displaying sequence homology to known genes (Table 3). Among these 376 genes, 57 are of unknown function. However, 87 genes were associated with metabolism, including carbohydrate, lipid, protein, and other metabolic processes. Importantly, 78% (68 of 87 genes) were upregulated by MPR (Table 3). Moreover, 50 genes were classified as involved in inflammation, all but two of which (96%) were downregulated by MPR. In addition, 9 of 10 genes involved in cell cycle regulation were also suppressed. A total of 120 genes were differentially expressed twofold or more in the MPR group (Tables 4 and 5).


View this table:
[in this window]
[in a new window]
 
Table 3. Overview of MPR-induced changes in VAT gene expression as determined by DNA microarray

 

View this table:
[in this window]
[in a new window]
 
Table 4. Genes with increased expression in MPR rat VAT

 

View this table:
[in this window]
[in a new window]
 
Table 5. Genes with decreased expression in MPR rat VAT

 
MPR increases expression of genes associated with carbohydrate metabolism. The expression of several genes involved in carbohydrate metabolism was upregulated by MPR (Table 3). Expression of the gene encoding glucose transporter-4 (GLUT4, +1.6-fold), which plays a key role in glucose uptake by adipocytes (12), was upregulated. MPR also increased the expression of glucose-6-phosphate isomerase (+1.6-fold) and phosphofructokinase (+1.5-fold), two important regulators of the glycolytic pathway (65). The expression of both malic enzyme (+1.8-fold) and malate dehydrogenase (+1.6-fold), two critical enzymes responsible for linking glycolysis with the tricarboxylic acid (TCA) cycle (65), was upregulated by MPR.

MPR increases expression of genes associated with lipid metabolism. Several genes involved in lipogenesis or adipocyte differentiation were upregulated by MPR (Table 4). Expression of fatty acid synthase (FAS, +3.3-fold), a key lipogenic enzyme (40), was upregulated by MPR. Moreover, MPR also increased the expression of other lipogenic enzymes, including glycerol-3-phosphate dehydrogenase [G3PDH, +1.9-fold (65)], acetyl-CoA carboxylase [ACC, +1.7-fold (65)], and the enzyme that catalyzes the rate-limiting step in lipogenesis, stearoyl-CoA desaturase [SCD, +2.6-fold (50)]. In addition to linking glycolysis with the TCA cycle, malic enzyme (+1.8-fold) also plays an important role in generating NADPH necessary for lipogenesis (65). It is well known that ANG (ANG) II is involved in lipogenesis (2), and the intracellular level of ANG II can be augmented either by the increased conversion of ANG I to ANG II or by the decreased inactivation of ANG II. It was remarkable that MPR not only upregulated the expression of chymase-1 (+2.0-fold), an enzyme involved in the conversion of ANG I to ANG II (15), but also downregulated the expression of leucyl-specific aminopeptidase (–1.8-fold), an enzyme that inactivates ANG II (73). Thus the expression of genes altered by MPR suggested the promotion of lipogenesis.

However, the expression of sterol regulatory element-binding protein-1c/adipocyte determination differentiation-dependent factor 1 (SREBP-1c/ADD1), a critical transcription factor involved in positively regulating a number of lipogenic enzymes (30), was not altered by MPR, suggesting that factors other than SREBP-1c may be responsible for the increased expression of lipogenic enzymes in the MPR adipose tissue. One such factor may be Spot-14, a gene known to activate the expression of lipogenic enzymes (14) and the expression of which (+1.6-fold) was upregulated by MPR.

In addition to upregulating genes associated with lipogenesis, MPR also enhanced the expression of genes involved in the uptake of free fatty acids (FFA) from circulation, including very low-density lipoprotein (VLDL) receptor [+1.6-fold (21)] and LDL-related protein-1 [+1.6-fold (6)]. Lipoprotein lipase, an enzyme responsible for hydrolyzing circulating lipids to produce FFA available for cellular uptake (74), and hormone-sensitive lipase, which plays a vital role in the mobilization of FFA from adipose tissue by controlling the rate of lipolysis of the stored triglycerides (23), were expressed abundantly in VAT, but MPR did not influence their expression. Digestion of dietary fat occurs through the activation of two enzymes: gastric lipases, secreted in the stomach, and pancreatic lipases, secreted in the duodenum. Interestingly, expression of pancreatic lipase [+4.6-fold (16)] and its activating cofactor, colipase [+7.0-fold (72)], was markedly upregulated by MPR. The expression of these two genes is novel to VAT, where they may play a role in facilitating the uptake of lipids from circulation and/or in cellular lipolysis. MPR also increased the expression of enzymes involved in cholesterol synthesis, sterol C5-desaturase [+1.6-fold (49)] and isopentenyl-diphosphate {Delta}-isomerase [+1.5-fold (5)]. Thus both cholesterol uptake and synthesis appeared to be enhanced by MPR.

MPR alters expression of genes associated with protein metabolism and other metabolic processes. Numerous pancreatic protein metabolizing enzymes, whose expression is novel to adipose tissue, including pancreatic cationic trypsinogen, pancreatic trypsin-1 (70), chymotrypsin-like chymotrypsin B (4), and carboxypeptidase B (13), were upregulated by MPR (Table 4). These pancreatic secreted enzymes normally function to digest proteins in the small intestine. In adipose tissue they may be involved in extracellular matrix (ECM) remodeling, as demonstrated in various carcinomas (59). MPR also altered the expression of enzymes involved in a variety of other metabolic processes, including nucleic acid metabolism, steroid hormone metabolism, and drug detoxification (Table 3).

MPR increases expression of genes associated with cellular proliferation and differentiation. MPR increased the expression of a number of growth factors, which are known to influence cellular proliferation and differentiation. These included transforming growth factor-{alpha} [TGF-{alpha}, +2.3-fold (66)], bone morphogenetic protein-3 [BMP3, +1.9-fold (3)], connective tissue growth factor [CTGF, +1.8-fold (48)], insulin-like growth factor II [IGF-II, +1.5-fold (39)], and fibroblast growth factors 7 and 21 [+1.8- and +1.5-fold, respectively (53)]. Moreover, several factors involved in adipocyte differentiation were upregulated by MPR, including MAP kinase phosphatase-1 [MKP-1, +2.2-fold (64)] as well as transcription factors CCAAT box enhancer-binding protein (C/EBP)-{delta} and C/EBP-{beta} [+1.9- and +1.6-fold, respectively (22)].

MPR alters expression of genes associated with ECM remodeling. MPR increased the expression of matrix metalloproteinase (MMP)-24 (+1.7-fold), which functions to activate MMP-2 (62), an enzyme known to play an important role in adipose tissue ECM remodeling. The expression of the ECM proteoglycans tetranectin [+8.7-fold (76)] and fibromodulin [+1.8-fold (32)], which are involved in cell-cell adhesion and participate in the assembly of the ECM, was upregulated by MPR. There was also a marked downregulation of genes encoding ECM components, including osteopontin (–5.2-fold), fibrillin (–4.0-fold), glycoprotein nmb (–3.6-fold), cadherin-22 (–2.7-fold), and type-1 collagen (–1.8-fold) (59). In addition, expression of cystatin N [–8.6-fold (1)], which functions as an endogenous inhibitor of the lysosomal cysteine proteinases, was profoundly downregulated.

Interestingly, expression of genes encoding a range of proteins associated with cytoskeletal structure of cells was upregulated by MPR. These included actin (+7.6-fold), myosin light chain-1 (+2.1-fold), myosin light chain-2 (+1.7-fold), tropomyosin (+1.8-fold), integrin (+1.9-fold), and the integrin-complexing protein Tspan-2 (+1.5-fold). Moreover, MPR increased the expression of several neuron-related structural proteins, including myelin protein zero (+4.5-fold), proteolipid protein (+2.1-fold), and limbic system-associated membrane protein (+1.9-fold). Therefore, these complex alterations in the expression of many ECM proteins and enzymes were indicative of increased ECM remodeling activities in the adipose tissue of MPR animals.

MPR enhances expression of angiogenic factors. Accumulating evidence suggests that adipose tissue growth/expansion is dependent on angiogenesis. The expression of several proangiogenic factors was upregulated by MPR. These factors included leptin [+1.9-fold (7)], CTGF [+1.8-fold (8)], cysteine-rich protein-61 [CYR61, +1.8-fold (8)], and dermatopontin [+1.6-fold (51)]. In addition, MPR increased the expression of chymase-1 (+2.0-fold), a proangiogenic enzyme involved in the conversion of ANG I to ANG II (15). The angiogenic effects of ANG II in adipose tissue were likely further enhanced in MPR animals, since the expression of leucyl-specific aminopeptidase (–1.8-fold), an extracellular enzyme that inactivates ANG II (73), was decreased. In contrast, MPR caused downregulation of the two antiangiogenic factors, F-spondin [–2.6-fold (69)] and properdin [–2.6-fold (42]). Thus MPR altered the pattern of adipose tissue gene expression in favor of angiogenesis.

MPR decreases expression of genes associated with cell-cycle regulation. Bcl-2-related protein-A1 is involved in regulating cell-cycle progression, and its expression (–2.8-fold) was repressed by MPR. In addition, several other genes involved in regulating the cell cycle include mitogen-inducible gene 2 (–2.7-fold), cyclin-dependent kinase (–2.3-fold) and cell division cycle 25B (–1.5-fold) were also downregulated by MPR.

MPR suppresses expression of genes involved in inflammation. The most notable class of genes downregulated by MPR was that associated with inflammation (Tables 3 and 5). The expression of several immunoglobulin and antigen genes was decreased (Table 5). Glycosylation-dependent cell adhesion molecule 1 (Glycam-1) is an endothelial cell glycoprotein that functions as an adhesive ligand for the lymphocyte adhesion molecule, L-selectin. Glycam-1 (–6.5-fold) expression was downregulated dramatically by MPR. Expression of 12-lipoxygenase, one of the key enzymes involved in arachidonic acid metabolism, was decreased more than fourfold (Table 5). Expression of two inflammatory proteins, S100 calcium-binding protein-A8 (–2.7-fold) and S100 calcium-binding protein-A9 (–2.2-fold), which form functional heterodimers, was reduced by MPR. The expression of sialophorin (–3.3-fold), a member of a ligand receptor complex involved in T cell activation, was also downregulated. Moreover, MPR downregulated the expression of complement component 3A (–1.8-fold) and 8 (–1.5-fold), which are members of the complement immunity pathway.

Only three inflammation-related genes were upregulated by MPR, and they were NF-{kappa}B inhibitor-{alpha} (I{kappa}B{alpha}, +2.1-fold), FK506-binding protein-2 (+1.9-fold), and CD1D (+1.8-fold). Two of the three upregulated genes, I{kappa}B{alpha} and FK506-binding protein-2, are negative regulators of inflammatory processes. Thus MPR altered the expression of genes associated with inflammation in a manner compatible with a suppressed inflammatory state.

Confirmation of microarray data by qRT-PCR. To validate the DNA microarray data, we determined the expression of several candidate genes that exhibited increased or decreased expression by microarray by use of qRT-PCR. In addition, the expression of two genes that displayed no change in expression by microarray was also examined. qRT-PCR confirmed that the data from the gene chips were robust and that in many cases the magnitude of fold changes was underestimated on the microarrays (Fig. 3).



View larger version (33K):
[in this window]
[in a new window]
 
Fig. 3. Levels of mRNA of selected representative genes (as indicated in A–I) quantified by quantitative RT-PCR. See text for definitions. Data are expressed as means ± SE; n = 6 rats. *P < 0.05 and **P < 0.01 compared with controls (male offspring of rat dams on control diet). For comparison, fold changes as determined by DNA microarray are shown in brackets.

 

    DISCUSSION
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
Accumulating evidence suggests that poor early growth is associated with an increased risk of developing insulin resistance, type 2 diabetes, and hypertension in later life (25, 29). Although it is generally accepted that obesity is required for the full expression of this thrifty phenotype (56), very few studies have focused on the long-term impact of poor early growth on the susceptibility to obesity in adult life. Ozanne et al. (57) recently reported the long-term effects of poor early nutrition on weight gain and the development of obesity in response to a highly palatable cafeteria-style diet. They showed that MPR mice at defined time windows not only had long-term (programmed) effects on weight gain but also altered the response to an obesity-inducing cafeteria-style diet and, hence, the susceptibility to obesity in later life. Mice who were growth restricted in utero but underwent rapid postnatal catch-up growth were heavier than control offspring throughout the study. These animals also gained more weight than the control offspring in response to the obesity-inducing diet. In marked contrast, mice that were growth restricted during lactation remained permanently smaller than controls. Moreover, when these mice were weaned onto the obesity-inducing diet, they showed no additional weight gain compared with their littermates on a control diet. These findings suggest that nutritional influences, both pre- and postnatal, can exert long-term (programmed) effects on body weight homoeostasis in later life. However, the molecular mechanisms that underlie this programming effect of poor early nutrition are unknown.

In the present study, we used a similar rat model of MPR in which dams were fed a low-protein diet throughout pregnancy and lactation (33, 58). We chose to study specifically visceral adiposity/obesity because it is the best predictor of the metabolic syndrome (45), which is known to occur in this rat model (55). We demonstrate that MPR predisposes the male offspring to visceral adiposity but not overweight in early adult life. Moreover, we show that MPR dramatically alters the transcriptional profile of VAT.

The cellular development associated with adipose tissue expansion involves adipocyte hypertrophy (increase in cell size) and/or adipocyte hyperplasia (increase in cell number) (26). Hypertrophy is the result of excess triglyceride accumulation in preexisting adipocytes due to a positive energy balance (energy intake in excess of energy expenditure). Hyperplasia, also known as adipogenesis, results from the recruitment of new preadipocytes from progenitor cells in adipose tissue. In addition, it involves the proliferation of preadipocytes and their differentiation into lipid filled mature adipocytes.

Two processes contribute to lipid accumulation in adipocytes, increased lipogenesis and reduced lipolysis. The increased expression of genes encoding a number of lipogenic enzymes (e.g., FAS, G3PDH, ACC, and SCD) in the MPR adipose tissue suggested that enhanced lipogenesis was one of the factors contributing to visceral adiposity in this rat model. Furthermore, an enhanced ability to take up FAA from circulation resulting from increased expression of VLDL receptor and its functionally related protein, as well as an increased capacity for cholesterol synthesis as a result of increased expression of genes coding for enzymes involved in the biosynthesis of cholesterol, also likely contributed to the excess adipose tissue expansion in MPR rats.

In addition, increases in the expression of genes encoding several growth factors known to be involved in regulating cell proliferation and differentiation suggested that an enhanced adipogenesis program was likely a contributing factor in the MPR-induced programming of visceral adiposity. The observed upregulation of MKP-1, a recently identified factor essential for in vitro adipocyte differentiation (64), as well as increased expression of the key transcription factors C/EBP-{delta} and C/EBP-{beta} involved in promoting adipocyte differentiation, are consistent with this notion. It was shown previously that a large number of cell cycle-related genes were repressed during in vitro adipogenesis (68). It is interesting to note that an equally extensive list of genes associated with cell cycle regulation was downregulated in adipose tissue of MPR rats, adding further support to the notion that adipogenesis was enhanced in this rat model. Moreover, there were no apparent differences in the size of adipocytes between control and MPR rats. Collectively, these findings suggested that the MPR-induced visceral adiposity was primarily a result of adipocyte hyperplasia.

Adipocyte differentiation in vitro involves remodeling of the ECM (22). ECM remodeling is a complex and dynamic process that involves alterations in the expression and activity of ECM remodeling enzymes and the subsequent modification to ECM components. MMP-2 is known to be upregulated in obesity and plays a critical role in adipose tissue ECM remodeling (11, 38). Expression of MMP-24, an enzyme involved in the activation of MMP-2 (62), was increased in MPR rats. Furthermore, MPR upregulated the expression of tetranectin, a plasminogen-binding protein that has been suggested to play an important role in tissue remodeling due to its ability to stimulate plasminogen activation and its expression in developing tissues such as the bone and muscle (31). In addition, expression of genes encoding a large number of cellular structural proteins and pancreatic proteases was also upregulated in MPR rats. Therefore, active adipose tissue remodeling was likely a major factor contributing to the development of visceral adiposity in this rat model.

In addition to promoting adipocyte differentiation, ECM remodeling is also involved in angiogenesis (10). Recent evidence suggests that adipose tissue expansion is angiogenesis dependent (63). It is believed that the angiogenic switch is tightly controlled by a balance between pro- and antiangiogenic factors (17). Thus angiogenesis occurs in a tissue when levels of proangiogenic factors are increased while those of antiangiogenic factors are reduced (18). In the present study, we observed coordinated upregulation of angiogenic factors (leptin, TGF-{alpha}, CTGF, CYR61, dermatopontin, and chymase-1) and downregulation of antiangiogenic factors (leucyl-specific aminopeptidase, F-spondin and properdin) in MPR rats. This suggested that angiogenesis likely played a key role in the pathogenesis of visceral adiposity in this rat model. It is noteworthy that the majority of these factors whose expression was altered by MPR have not been described previously in adipose tissue, thus attesting to the utility of this functional genomics approach in identifying new players involved in adiposity/obesity.

In several rodent models of obesity, adipose tissue gene expression profiling has revealed a global upregulation of genes associated with inflammation (46, 47, 67). Adipocytes are capable of expressing inflammatory genes (34). In addition, inflammatory cells such as macrophages have been demonstrated to be present in higher numbers in adipose tissue in obesity (75, 77). It was surprising that a large number of genes involved in inflammatory processes were downregulated in this rat model. Although the precise reasons for this disparity are not apparent at the present time, it is tempting to speculate that the suppressed inflammatory state in the MPR adipose tissue may represent an earlier event occurring during the development of adiposity/obesity, because it is believed that the increased inflammatory response is likely a result of obesity (77). Consequently, it has been suggested that this enhanced inflammatory response may link obesity to its associated metabolic diseases (36). However, the pathophysiological significance of a suppressed inflammation state within the adipose tissue remains to be determined.

Interestingly, several of the changes in the transcriptional profile of VAT as a result of MPR are similar to those observed in white adipose tissue following long-term caloric restriction (27). These include enhanced expression of genes involved in regulating lipogenesis and adipocyte differentiation as well as a global downregulation of genes associated with inflammatory processes. This similarity between the two distinct models suggests that MPR and caloric restriction alter adipose tissue gene activity in a similar way, thus underscoring the significance of our present findings utilizing this rat model in understanding the long-term effects of poor early nutrition on susceptibility to visceral adiposity/obesity in later life.

It is noteworthy that the present study, like many others in the literature (20, 37, 46, 47, 67), used RNA samples derived from adipose tissue for gene expression profile analysis. This kind of study design is distinct (in the objectives and, hence, the interpretations of the results) from those in which RNA extracts prepared from a pure population of adipocytes was utilized (68). Consequently, many of the alterations in adipose tissue gene expression that we observed in this study could be explained by a change in the ratio of adipocytes to other cell types between control and MPR rats. Obviously, the validity of this contention requires future scrutiny.

In conclusion, we demonstrate that maternal protein restriction during pregnancy and lactation programs the susceptibility to visceral adiposity in the adult rat offspring, and that the coordinated upregulation of molecular pathways involved in adipogenesis and angiogenesis likely plays a key role in this process (Fig. 4).



View larger version (21K):
[in this window]
[in a new window]
 
Fig. 4. Schematic overview depicting key differentially expressed genes and postulated molecular pathways that may be involved in the pathogenesis of visceral adiposity in this rat model. For full explanation and definitions, see RESULTS and DISCUSSION.

 

    GRANTS
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 
This work was supported by grants from the Canadian Institutes of Health Research.


    ACKNOWLEDGMENTS
 
We thank Brenda Strutt for performing fat tissue histology.


    FOOTNOTES
 

Address for reprint requests and other correspondence: K. Yang, Children's Health Research Institute, Rm. A5-132, Victoria Research Laboratories- Westminster Campus, 800 Commissioners Road East, London, ON, Canada N6A 4G5 (E-mail: kyang{at}uwo.ca)

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.


    REFERENCES
 TOP
 ABSTRACT
 MATERIALS AND METHODS
 RESULTS
 DISCUSSION
 GRANTS
 REFERENCES
 

  1. Abrahamson M, Alvarez-Fernandez M, and Nathanson CM. Cystatins. Biochem Soc Symp: 179–199, 2003.
  2. Ailhaud G, Teboul M, and Massiera F. Angogen, adipocyte differentiation and fat mass enlargement. Curr Opin Clin Nutr Metab Care 5: 385–389, 2002.[CrossRef][ISI][Medline]
  3. Bahamonde ME and Lyons KM. BMP3: to be or not to be a BMP. J Bone Joint Surg Am 83-A, Suppl 1: S56–S62, 2001.
  4. Barker DJ. Fetal origins of coronary heart disease. BMJ 311: 171–174, 1995.[Free Full Text]
  5. Bell GI, Quinto C, Quiroga M, Valenzuela P, Craik CS, and Rutter WJ. Isolation and sequence of a rat chymotrypsin B gene. J Biol Chem 259: 14265–14270, 1984.[Abstract/Free Full Text]
  6. Biardi L and Krisans SK. Compartmentalization of cholesterol biosynthesis. Conversion of mevalonate to farnesyl diphosphate occurs in the peroxisomes. J Biol Chem 271: 1784–1788, 1996.[Abstract/Free Full Text]
  7. Boucher P, Gotthardt M, Li WP, Anderson RG, and Herz J. LRP: role in vascular wall integrity and protection from atherosclerosis. Science 300: 329–332, 2003.[Abstract/Free Full Text]
  8. Bouloumie A, Drexler HC, Lafontan M, and Busse R. Leptin, the product of Ob gene, promotes angiogenesis. Circ Res 83: 1059–1066, 1998.[Abstract/Free Full Text]
  9. Brigstock DR. Regulation of angiogenesis and endothelial cell function by connective tissue growth factor (CTGF) and cysteine-rich 61 (CYR61). Angiogenesis 5: 153–165, 2002.[CrossRef][Medline]
  10. Byrne CD and Phillips DI. Fetal origins of adult disease: epidemiology and mechanisms. J Clin Pathol 53: 822–828, 2000.[Abstract/Free Full Text]
  11. Chang C and Werb Z. The many faces of metalloproteases: cell growth, invasion, angiogenesis and metastasis. Trends Cell Biol 11: S37–43, 2001.[CrossRef][ISI][Medline]
  12. Chavey C, Mari B, Monthouel MN, Bonnafous S, ANGlard P, Van Obberghen E, and Tartare-Deckert S. Matrix metalloproteinases are differentially expressed in adipose tissue during obesity and modulate adipocyte differentiation. J Biol Chem 278: 11888–11896, xxxx.
  13. Chiappe De Cingolani GE and Caldiz CI. Insulin resistance and GLUT-4 glucose transporter in adipocytes from hypertensive rats. Metabolism 53: 382–387, 2004.[CrossRef][ISI][Medline]
  14. Clauser E, Gardell SJ, Craik CS, MacDonald RJ, and Rutter WJ. Structural characterization of the rat carboxypeptidase A1 and B genes. Comparative analysis of the rat carboxypeptidase gene family. J Biol Chem 263: 17837–17845, 1988.[Abstract/Free Full Text]
  15. Compe E, de Sousa G, Francois K, Roche R, Rahmani R, Torresani J, Raymondjean M, and Planells R. Spot 14 protein interacts and co-operates with chicken ovalbumin upstream promoter-transcription factor 1 in the transcription of the L-type pyruvate kinase gene through a specificity protein 1 (Sp1) binding site. Biochem J 358: 175–183, 2001.[CrossRef][ISI][Medline]
  16. Dell'Italia LJ and Husain A. Dissecting the role of chymase in ANG II formation and heart and blood vessel diseases. Curr Opin Cardiol 17: 374–379, 2002.[CrossRef][ISI][Medline]
  17. Figarella C, De Caro A, Leupold D, and Poley JR. Congenital pancreatic lipase deficiency. J Pediatr 96: 412–416, 1980.[ISI][Medline]
  18. Folkman J. Angiogenesis in cancer, vascular, rheumatoid and other disease. Nat Med 1: 27–31, 1995.[CrossRef][ISI][Medline]
  19. Folkman J. Fundamental concepts of the angiogenic process. Curr Mol Med 3: 643–651, 2003.[CrossRef][ISI][Medline]
  20. Froguel P and Boutin P. Genetics of pathways regulating body weight in the development of obesity in humans. Exp Biol Med (Maywood) 226: 991–996, 2001.[Abstract/Free Full Text]
  21. Gomez-Ambrosi J, Catalan V, Diez-Caballero A, Martinez-Cruz LA, Gil MJ, Garcia-Foncillas J, Cienfuegos JA, Salvador J, Mato JM, and Fruhbeck G. Gene expression profile of omental adipose tissue in human obesity. FASEB J 18: 215–217, 2004.[Abstract/Free Full Text]
  22. Goudriaan JR, Tacken PJ, Dahlmans VE, Gijbels MJ, van Dijk KW, Havekes LM, and Jong MC. Protection from obesity in mice lacking the VLDL receptor. Arterioscler Thromb Vasc Biol 21: 1488–1493, 2001.[Abstract/Free Full Text]
  23. Gregoire FM, Smas CM, and Sul HS. Understanding adipocyte differentiation. Physiol Rev 78: 783–809, 1998.[Abstract/Free Full Text]
  24. Haemmerle G, Zimmermann R, Hayn M, Theussl C, Waeg G, Wagner E, Sattler W, Magin TM, Wagner EF, and Zechner R. Hormone-sensitive lipase deficiency in mice causes diglyceride accumulation in adipose tissue, muscle, and testis. J Biol Chem 277: 4806–4815, 2002.[Abstract/Free Full Text]
  25. Hales CN and Barker DJ. Type 2 (non-insulin-dependent) diabetes mellitus: the thrifty phenotype hypothesis. Diabetologia 35: 595–601, 1992.[CrossRef][ISI][Medline]
  26. Hales CN and Ozanne SE. For debate: fetal and early postnatal growth restriction lead to diabetes, the metabolic syndrome and renal failure. Diabetologia 46: 1013–1019, 2003.[CrossRef][ISI][Medline]
  27. Hausman DB, DiGirolamo M, Bartness TJ, Hausman GJ, and Martin RJ. The biology of white adipocyte proliferation. Obes Rev 2: 239–254, 2001.[CrossRef][Medline]
  28. Higami Y, Pugh TD, Page GP, Allison DB, Prolla TA, and Weindruch R. Adipose tissue energy metabolism: altered gene expression profile of mice subjected to long-term caloric restriction. FASEB J 18: 415–417, 2004.[Abstract/Free Full Text]
  29. Hofbauer KG. Molecular pathways to obesity. Int J Obes Relat Metab Disord 26: S18–S27, 2002.
  30. Holemans K, Aerts L, and Van Assche FA. Fetal growth restriction and consequences for the offspring in animal models. J Soc Gynecol Investig 10: 392–399, 2003.[CrossRef][ISI][Medline]
  31. Horton JD, Goldstein JL, and Brown MS. SREBPs: activators of the complete program of cholesterol and fatty acid synthesis in the liver. J Clin Invest 109: 1125–1131, 2002.[Free Full Text]
  32. Iba K, Durkin ME, Johnsen L, Hunziker E, Damgaard-Pedersen K, Zhang H, Engvall E, Albrechtsen R, and Wewer UM. Mice with a targeted deletion of the tetranectin gene exhibit a spinal deformity. Mol Cell Biol 21: 7817–7825, 2001.[Abstract/Free Full Text]
  33. Jepsen KJ, Wu F, Peragallo JH, Paul J, Roberts L, Ezura Y, Oldberg A, Birk DE, and Chakravarti S. A syndrome of joint laxity and impaired tendon integrity in lumican- and fibromodulin-deficient mice. J Biol Chem 277: 35532–35540, 2002.[Abstract/Free Full Text]
  34. Joanette EA, Reusens B, Arany E, Thyssen S, Remacle RC, and Hill DJ. Low-protein diet during early life causes a reduction in the frequency of cells immunopositive for nestin and CD34 in both pancreatic ducts and islets in the rat. Endocrinology 145: 3004–3013, 2004.[Abstract/Free Full Text]
  35. Kershaw EE and Flier JS. Adipose tissue as an endocrine organ. J Clin Endocrinol Metab 89: 2548–2556, 2004.[Abstract/Free Full Text]
  36. Krysiak R, Okopien B, Belowski D, Madej A, and Herman ZS. Recent insights into body weight control: from physiology to pathology. J Pept Sci 7: 571–578, 2001.[CrossRef][ISI][Medline]
  37. Lehrke M and Lazar MA. Inflamed about obesity. Nat Med 10: 126–127, 2004.[CrossRef][ISI][Medline]
  38. Li J, Yu X, Pan W, and Unger RH. Gene expression profile of rat adipose tissue at the onset of high-fat-diet obesity. Am J Physiol Endocrinol Metab 282: E1334–E1341, 2002.[Abstract/Free Full Text]
  39. Lijnen HR, Maquoi E, Holvoet P, Mertens A, Lupu F, Morange P, Alessi MC, and Juhan-Vague I. Adipose tissue expression of gelatinases in mouse models of obesity. Thromb Haemost 85: 1111–1116, 2001.[ISI][Medline]
  40. Liu Q, Yan H, Dawes NJ, Mottino GA, Frank JS, and Zhu H. Insulin-like growth factor II induces DNA synthesis in fetal ventricular myocytes in vitro. Circ Res 79: 716–726, 1996.[Abstract/Free Full Text]
  41. Loftus TM, Jaworsky DE, Frehywot GL, Townsend CA, Ronnett GV, Lane MD, and Kuhajda FP. Reduced food intake and body weight in mice treated with fatty acid synthase inhibitors. Science 288: 2379–2381, 2000.[Abstract/Free Full Text]
  42. Matthews SG. Early programming of the hypothalamo-pituitary-adrenal axis. Trends Endocrinol Metab 13: 373–380, 2002.[CrossRef][ISI][Medline]
  43. Maves KK and Weiler JM. Properdin: approaching four decades of research. Immunol Res 12: 233–243, 1993.[ISI][Medline]
  44. Mokdad AH, Ford ES, Bowman BA, Dietz WH, Vinicor F, Bales VS, and Marks JS. Prevalence of obesity, diabetes, and obesity-related health risk factors, 2001. JAMA 289: 76–79, 2003.[Abstract/Free Full Text]
  45. Montague CT, Farooqi IS, Whitehead JP, Soos MA, Rau H, Wareham NJ, Sewter CP, Digby JE, Mohammed SN, Hurst JA, Cheetham CH, Earley AR, Barnett AH, Prins JB, and O'Rahilly S. Congenital leptin deficiency is associated with severe early-onset obesity in humans. Nature 387: 903–908, 1997.[CrossRef][ISI][Medline]
  46. Montague CT and O'Rahilly S. The perils of portliness: causes and consequences of visceral adiposity. Diabetes 49: 883–888, 2000.[Abstract]
  47. Moraes RC, Blondet A, Birkenkamp-Demtroeder K, Tirard J, Orntoft TF, Gertler A, Durand P, Naville D, and Begeot M. Study of the alteration of gene expression in adipose tissue of diet-induced obese mice by microarray and reverse transcription-polymerase chain reaction analyses. Endocrinology 144: 4773–4782, 2003.[Abstract/Free Full Text]
  48. Nadler ST, Stoehr JP, Schueler KL, Tanimoto G, Yandell BS, and Attie AD. The expression of adipogenic genes is decreased in obesity and diabetes mellitus. Proc Natl Acad Sci USA 97: 11371–11376, 2000.[Abstract/Free Full Text]
  49. Nakanishi T, Yamaai T, Asano M, Nawachi K, Suzuki M, Sugimoto T, and Takigawa M. Overexpression of connective tissue growth factor/hypertrophic chondrocyte-specific gene product 24 decreases bone density in adult mice and induces dwarfism. Biochem Biophys Res Commun 281: 678–681, 2001.[CrossRef][ISI][Medline]
  50. Nishi S, Nishino H, and Ishibashi T. cDNA cloning of the mammalian sterol C5-desaturase and the expression in yeast mutant. Biochim Biophys Acta 1490: 106–108, 2000.[ISI][Medline]
  51. Ntambi JM, Miyazaki M, Stoehr JP, Lan H, Kendziorski CM, Yandell BS, Song Y, Cohen P, Friedman JM, and Attie AD. Loss of stearoyl-CoA desaturase-1 function protects mice against adiposity. Proc Natl Acad Sci USA 99: 11482–11486, 2002.[Abstract/Free Full Text]
  52. Okamoto O, Fujiwara S, Abe M, and Sato Y. Dermatopontin interacts with transforming growth factor beta and enhances its biological activity. Biochem J 337: 537–541, 1999.[CrossRef][ISI][Medline]
  53. Oken E and Gillman MW. Fetal origins of obesity. Obes Res 11: 496–506, 2003.[Abstract/Free Full Text]
  54. Ornitz DM and Itoh N. Fibroblast growth factors. Genome Biol 2: Reviews 3005, 2001.
  55. Osmond C and Barker DJ. Fetal, infant, and childhood growth are predictors of coronary heart disease, diabetes, and hypertension in adult men and women. Environ Health Perspect 108: 545–553, 2000.[ISI][Medline]
  56. Ozanne SE. Metabolic programming in animals. Br Med Bull 60: 143–152, 2001.[Abstract/Free Full Text]
  57. Ozanne SE and Hales CN. Early programming of glucose-insulin metabolism. Trends Endocrinol Metab 13: 368–373, 2002.[CrossRef][ISI][Medline]
  58. Ozanne SE, Lewis R, Jennings BJ, and Hales CN. Early programming of weight gain in mice prevents the induction of obesity by a highly palatable diet. Clin Sci (Lond) 106: 141–145, 2004.[Medline]
  59. Petrik J, Reusens B, Arany E, Remacle C, Coelho C, Hoet JJ, and Hill DJ. A low protein diet alters the balance of islet cell replication and apoptosis in the fetal and neonatal rat and is associated with a reduced pancreatic expression of insulin-like growth factor-II. Endocrinology 140: 4861–4873, 1999.[Abstract/Free Full Text]
  60. Pupa SM, Menard S, Forti S, and Tagliabue E. New insights into the role of extracellular matrix during tumor onset and progression. J Cell Physiol 192: 259–267, 2002.[CrossRef][ISI][Medline]
  61. Purdy LP and Metzger BE. Influences of the intrauterine metabolic environment on adult disease: what may we infer from size at birth? Diabetologia 39: 1126–1130, 1996.[CrossRef][ISI][Medline]
  62. Rogers I. The influence of birthweight and intrauterine environment on adiposity and fat distribution in later life. Int J Obes Relat Metab Disord 27: 755–777, 2003.[CrossRef][Medline]
  63. Romanic AM, Burns-Kurtis CL, Ao Z, Arleth AJ, and Ohlstein EH. Upregulated expression of human membrane type-5 matrix metalloproteinase in kidneys from diabetic patients. Am J Physiol Renal Physiol 281: F309–F317, 2001.[Abstract/Free Full Text]
  64. Rupnick MA, Panigrahy D, Zhang CY, Dallabrida SM, Lowell BB, Langer R, and Folkman MJ. Adipose tissue mass can be regulated through the vasculature. Proc Natl Acad Sci USA 99: 10730–10735, 2002.[Abstract/Free Full Text]
  65. Sakaue H, Ogawa W, Nakamura T, Mori T, Nakamura K, and Kasuga M. Role of MAP kinase phosphatase-1 (MKP-1) in adipocyte differentiation. J Biol Chem 20: 20, 2004.
  66. Salway JG. Metabolism at a Glance. Malden, MA: Blackwell, 2004.
  67. Serrero G and Lepak N. Endocrine and paracrine negative regulators of adipose differentiation. Int J Obes Relat Metab Disord 20: S58–S64, 1996.[Medline]
  68. Soukas A, Cohen P, Socci ND, and Friedman JM. Leptin-specific patterns of gene expression in white adipose tissue. Genes Dev 14: 963–980, 2000.[Abstract/Free Full Text]
  69. Soukas A, Socci ND, Saatkamp BD, Novelli S, and Friedman JM. Distinct transcriptional profiles of adipogenesis in vivo and in vitro. J Biol Chem 276: 34167–34174, 2001.[Abstract/Free Full Text]
  70. Terai Y, Abe M, Miyamoto K, Koike M, Yamasaki M, Ueda M, Ueki M, and Sato Y. Vascular smooth muscle cell growth-promoting factor/F-spondin inhibits angiogenesis via the blockade of integrin alphavbeta3 on vascular endothelial cells. J Cell Physiol 188: 394–402, 2001.[CrossRef][ISI][Medline]
  71. Towatari T, Ide M, Ohba K, Chiba Y, Murakami M, Shiota M, Kawachi M, Yamada H, and Kido H. Identification of ectopic anionic trypsin I in rat lungs potentiating pneumotropic virus infectivity and increased enzyme level after virus infection. Eur J Biochem 269: 2613–2621, 2002.[Abstract/Free Full Text]
  72. Van Beek J, Guan H, Julan L, and Yang K. Glucocorticoid stimulates the expression of 11 beta-hydroxysteroid dehydrogenase type 2 in cultured human placental trophoblast cells. J Clin Endocrinol Metab 89: 5614–5621, 2004.[Abstract/Free Full Text]
  73. Van Tilbeurgh H, Bezzine S, Cambillau C, Verger R, and Carriere F. Colipase: structure and interaction with pancreatic lipase. Biochim Biophys Acta 1441: 173–184, 1999.[ISI][Medline]
  74. Watanabe Y, Shibata K, Kikkawa F, Kajiyama H, Ino K, Hattori A, Tsujimoto M, and Mizutani S. Adipocyte-derived leucine aminopeptidase suppresses angiogenesis in human endometrial carcinoma via renin-ANG system. Clin Cancer Res 9: 6497–6503, 2003.[Abstract/Free Full Text]
  75. Weinstock PH, Bisgaier CL, Aalto-Setala K, Radner H, Ramakrishnan R, Levak-Frank S, Essenburg AD, Zechner R, and Breslow JL. Severe hypertriglyceridemia, reduced high density lipoprotein, and neonatal death in lipoprotein lipase knockout mice. Mild hypertriglyceridemia with impaired very low density lipoprotein clearance in heterozygotes. J Clin Invest 96: 2555–2568, 1995.[ISI][Medline]
  76. Weisberg SP, McCann D, Desai M, Rosenbaum M, Leibel RL, and Ferrante AW Jr. Obesity is associated with macrophage accumulation in adipose tissue. J Clin Invest 112: 1796–1808, 2003.[Abstract/Free Full Text]
  77. Wewer UM, Iba K, Durkin ME, Nielsen FC, Loechel F, Gilpin BJ, Kuang W, Engvall E, and Albrechtsen R. Tetranectin is a novel marker for myogenesis during embryonic development, muscle regeneration, and muscle cell differentiation in vitro. Dev Biol 200: 247–259, 1998.[CrossRef][ISI][Medline]
  78. Xu H, Barnes GT, Yang Q, Tan G, Yang D, Chou CJ, Sole J, Nichols A, Ross JS, Tartaglia LA, and Chen H. Chronic inflammation in fat plays a crucial role in the development of obesity-related insulin resistance. J Clin Invest 112: 1821–1830, 2003.[Abstract/Free Full Text]