**Contents**




### **B ´arbara Pereira da Silva, Nikolai Kolba, H ´ercia Stampini Duarte Martino, Jonathan Hart and Elad Tako**



### **About the Editor**

**Elad Tako** holds degrees in animal science (B.S.), endocrinology (M.S.), and physiology/nutrigenomics (Ph.D.), with previous appointments at the Hebrew University of Jerusalem, North Carolina State University, and Cornell University. As an Associate Professor with the Department of Food Science at Cornell University, Dr. Tako's research focuses on various aspects of trace mineral deficiencies, emphasizing molecular, physiological and nutritional factors and practices that influence the intestinal micronutrient absorption. With over 100 peer-reviewed publications and presentations, he leads a research team focused on understanding the interactions between dietary factors, physiological and molecular biomarkers, the microbiome, and intestinal functionality. His research accomplishments include the development of the Gallus gallus intra-amniotic administration procedure, and establishing recognized approaches for using animal models within mineral bioavailability and intestinal absorption screening processes. He has also developed a zinc status physiological blood biomarker (red blood cell Linoleic Acid: Dihomo–Linolenic Acid Ratio), and molecular tissue biomarkers to assess the effect of dietary mineral deficiencies on intestinal functionality, and how micronutrients dietary deficiencies alter gut microbiota composition and function.

### *Editorial* **Dietary Plant-Origin Bio-Active Compounds, Intestinal Functionality, and Microbiome**

#### **Elad Tako**

Department of Food Science, Cornell University, Stocking Hall, Ithaca, NY 14853-7201, USA; et79@cornell.edu Received: 5 October 2020; Accepted: 17 October 2020; Published: 22 October 2020

**Abstract:** In recent years, plant-origin bio-active compounds in foods (staple crops, fruit, vegetables, and others) have been gaining interest, and processes to consider them for public health recommendations are being presented and discussed in the literature. However, at times, it may be challenging to demonstrate causality, and there often is not a single compound–single effect relationship. Furthermore, it was suggested that health benefits may be due to metabolites produced by the host or gut microbiome rather than the food constituent per se. Over the years, compounds that were investigated were shown to increase gut microbial diversity, improve endothelial function, improve cognitive function, reduce bone loss, and many others. More recently, an additional and significant body of evidence further demonstrated the nutritional role and potential effects that plant-origin bio-active compounds might have on intestinal functionality (specifically the duodenal brush border membrane, morphology, and the abundance of health-promoting bacterial populations). Hence, the special issue "Dietary Plant-Origin Bio-Active Compounds, Intestinal Functionality, and Microbiome" comprises 11 peer-reviewed papers on the most recent evidence regarding the potential dietary intake and effects of plant-origin bio-active compounds on intestinal functionality, primarily in the context of brush border functional proteins (enzymes and transporters), mineral (and other nutrients) dietary bioavailability, and the intestinal microbiome. Original contributions and literature reviews further demonstrated the potential dietary relevance that plant bio-active compounds hold in human health and development. This editorial provides a brief and concise overview that addresses and summarizes the content of the *Dietary Plant-Origin Bio-Active Compounds, Intestinal Functionality, and Microbiome* special issue.

**Keywords:** plant origin; bio-active compounds; intestine; microbiome

The purpose of the current special issue is to further expand and add research knowledge of the vital role dietary plant-origin bio-active compounds hold in various nutrition-related physiological and metabolic pathways. In addition, the purpose is to further contribute to the knowledge regarding the relationship between plant-origin bio-active compounds, the intestinal morphology and functionality, and potential effects on the intestinal microbiome.

Plant-based diets contain a plethora of metabolites that may impact on health and disease prevention. Most are focused on the potential bioactivity and nutritional relevance of several classes of phytochemicals, such as polyphenols, flavonoids, carotenoids, phyto-estrogens, and frucrooligo-saccharides [1]. These compounds are found in fruit, vegetables, and herbs [2]. Daily intakes of some of these compounds may exceed 100 mg. Moreover, intestinal bacterial activity may transform complex compounds such as anthocyanins, procyanidins, and isoflavones into simple phenolic metabolites [3]. The colon is thus a rich source of potentially active phenolic acids that may impact both locally and systemically on gut health. Furthermore, non-digestible fiber (prebiotics) are dietary substrates that selectively promote proliferation and/or activity of health-promoting bacterial populations in the colon [4]. Prebiotics, such as inulin, raffinose, and stachyose, have a proven ability to promote the abundance of intestinal bacterial populations, which may provide

additional health benefits to the host [5–10]. Furthermore, various pulse seed soluble (fiber) extracts are responsible for improving gastrointestinal motility, intestinal functionality and morphology, and mineral absorption [9,11]. Studies have indicated that the consumption of seed-origin soluble extracts can up-regulate the expression of brush border membrane (BBM) proteins that contribute for digestion and absorption of nutrients. The soluble extracts can positively affect intestinal health by increasing the mucus production, goblet cells number/diameter, villus surface area, and crypt depth [9,10]. These functional and morphological effects appear to occur due to the increased motility of the digestive tract, leading to hyperplasia and/or hypertrophy of muscle cells. Plant-origin soluble extracts may act, directly or indirectly, as a factor that increases mineral solubility and, therefore, dietary bioavailability. This occurs due to fiber fermentation and bacterial production of short chain fatty acids (SCFA) that reduces intestinal pH, inhibits the growth of potentially pathogenic bacterial population and increases the solubility and, therefore, absorption of minerals. The SCFA can increase the proliferation of epithelial cells, which, in return, increase the absorptive surface area, which contributes to the absorption of nutrients [12,13]. Several phenolic acids and other phytochemicals affect the expression and activity of enzymes involved in the production of inflammatory mediators of pathways thought to be important in the development of gut disorders including colon cancer. However, it is still unclear as to which of these compounds are beneficial to gut health. Hence, the aim of the current special issue is to further explore the interactions between dietary plant-origin bio-active compounds, their potential effects on the intestinal bacterial populations, and overall intestinal functionality and gut health.

This monograph, based on a special issue of Nutrients, contains 11 manuscripts—1 review and 10 original publications—that reflect the wide spectrum of currently conducted research in the field of dietary plant-origin bio-active compounds, intestinal functionality, and microbiome. The manuscripts in this special issue collection include contributors and researchers from multiple countries, including USA, Canada, Australia, Brazil, Poland, Finland, Belgium, Netherlands, and Spain. The presented manuscripts cover a wide variety and range of topics in the field of dietary plant-origin bio-active compounds, intestinal functionality, and microbiome, with emphasis on diet and intestinal well-being and compositions of fecal microbiota and short chain fatty acids in oat and by using subjects with celiac disease or gluten sensitivity [14]. The demonstration of low phytate peas (Pisum sativum L.)-based diets improve iron status, gut microbiome, and brush border membrane functionality in vivo (*Gallus gallus*) [15]. The presentation of novel non-digestible, carrot-derived polysaccharide (cRG-I) and how it selectively modulates the human gut microbiota while promoting gut barrier integrity (an integrated in vitro approach) [16]. The in vitro evaluation of prebiotic properties of a commercial artichoke inflorescence extract revealed bifidogenic effects [17]. The discussion of possible protective effects of traumatic acid (TA) on the cancerous effect of mesotrione [18]. Is Acrylamide as a harmful as we think? A new look at the impact of Acrylamide on the viability of beneficial intestinal bacteria of the genus *Lactobacillus* [19] The biological activity of new cichoric acid–metal complexes in bacterial strains, yeast-like fungi, and human cell cultures in vitro [20]. The presentation of how soluble extracts from chia seed (*Salvia hispanica* L.) affect brush border membrane functionality, morphology and intestinal bacterial populations in vivo (*Gallus gallus*) [21]. The fructose consumption by adult rats exposed to dexamethasone in utero changes the phenotype of intestinal epithelial cells and exacerbates intestinal gluconeogenesis [22]. Alterations in the intestinal morphology, gut microbiota, and trace mineral status following intra-amniotic administration (Gallus gallus) of teff (Eragrostis tef) seed extracts [23]. Non-dairy fermented beverages as potential carriers to ensure probiotics, prebiotics, and bio-active compounds arrival to the gut and their health benefits [24]. These wide spectra of topics further demonstrate the importance and relevance of dietary plant-origin bio-active compounds, and their effects on intestinal functionality and microbiome.

This special issue and collection of manuscripts is a useful summary of progress in various areas related to dietary plant-origin bio-active compounds, intestinal functionality, and microbiome. It also points to additional research needs, including recommendations for future research in the field, and to better understand the dietary role that dietary plant-origin bio-active compounds hold and regarding human nutrition and overall health.

**Funding:** This research received no external funding

**Conflicts of Interest:** The author declares no conflict of interest.

#### **References**


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*Article*

## **Fructose Consumption by Adult Rats Exposed to Dexamethasone In Utero Changes the Phenotype of Intestinal Epithelial Cells and Exacerbates Intestinal Gluconeogenesis**

**Gizela A. Pereira 1, Frhancielly S. Sodré 1, Gilson M. Murata 1, Andressa G. Amaral 1, Tanyara B. Payolla 1, Carolina V. Campos 2, Fabio T. Sato 1, Gabriel F. Anhê <sup>2</sup> and Silvana Bordin 1,\***


Received: 3 September 2020; Accepted: 2 October 2020; Published: 7 October 2020

**Abstract:** Fructose consumption by rodents modulates both hepatic and intestinal lipid metabolism and gluconeogenesis. We have previously demonstrated that in utero exposure to dexamethasone (DEX) interacts with fructose consumption during adult life to exacerbate hepatic steatosis in rats. The aim of this study was to clarify if adult rats born to DEX-treated mothers would display differences in intestinal gluconeogenesis after excessive fructose intake. To address this issue, female Wistar rats were treated with DEX during pregnancy and control (CTL) mothers were kept untreated. Adult offspring born to CTL and DEX-treated mothers were assigned to receive either tap water (Control-Standard Chow (CTL-SC) and Dexamethasone-Standard Chow (DEX-SC)) or 10% fructose in the drinking water (CTL-fructose and DEX-fructose). Fructose consumption lasted for 80 days. All rats were subjected to a 40 h fasting before sample collection. We found that DEX-fructose rats have increased glucose and reduced lactate in the portal blood. Jejunum samples of DEX-fructose rats have enhanced phosphoenolpyruvate carboxykinase (PEPCK) expression and activity, higher facilitated glucose transporter member 2 (GLUT2) and facilitated glucose transporter member 5 (GLUT5) content, and increased villous height, crypt depth, and proliferating cell nuclear antigen (PCNA) staining. The current data reveal that rats born to DEX-treated mothers that consume fructose during adult life have increased intestinal gluconeogenesis while recapitulating metabolic and morphological features of the neonatal jejunum phenotype.

**Keywords:** intrauterine growth restriction (IUGR); fructose; dexamethasone; intestinal gluconeogenesis

#### **1. Introduction**

The consumption of fructose-sweetened beverages has significantly increased during the last decades and a great number of observational studies have associated this nutritional habit with increased cardiometabolic risk [1]. In accordance with this hypothesis, experimental studies have described that rats consuming high amounts of fructose or sucrose develop glucose intolerance and increased hepatic gluconeogenesis [2–4].

The mechanisms underlying the metabolic effects of excessive fructose intake rely on its hepatic as well as its intestinal metabolism [5]. The small intestine absorbs fructose through a facilitated glucose

transporter member 5 (GLUT5)-dependent mechanism and partially metabolizes it into lactate, glucose, and fatty acids that are sequentially secreted to the portal circulation [6,7]. However, intestinal fructose metabolism capacity is limited and when high amounts of fructose are consumed, a considerable fraction reaches the liver [7]. Hepatic metabolism of fructose, in turn, produces glyceraldehyde-3-phosphate that is driven to either gluconeogenesis or de novo lipogenesis (DNLG) [8].

Besides its metabolism to intermediates that feed gluconeogenesis and DNLG, fructose was described to modulate the expression of key metabolic genes [5]. Fructose consumption increased the expression of GLUT5 and gluconeogenic enzymes glucose-6-phosphatae (G6Pase) and fructose-1,6-bisphosphatase (FBP1) in the small intestine [9,10]. Excessive fructose intake was also reported to increase the hepatic expression of the gluconeogenesis enzymes G6Pase, FBP1, and phosphoenolpyruvate carboxykinase (PEPCK) [11–14], and the DNLG enzymes acetyl-CoA carboxylase (ACC), fatty acid synthase (FAS), and stearoyl-CoA desaturase-1 (SCD1) [15].

We have recently demonstrated that the modulation of key metabolic genes induced by fructose in the liver can interact with other factors, such as birth weight. Fructose supplementation of rats born small due to maternal treatment with dexamethasone (DEX) induced an expected increase in the expression of PEPCK, FAS, and ACC, but failed to increase the expression of genes involved in very low density lipoproteins (VLDL) assembly and secretion, leading to an exacerbation in hepatic steatosis [16].

In addition to low birth weight, in utero exposure to DEX is recognized to program the energy metabolism during adult life. Offspring born to DEX-treated mothers develop glucose intolerance and increased hepatic PEPCK expression as soon as the 21st day of life [17]. Additionally, pancreatic postnatal development of rats born to DEX-treated mothers is hallmarked by lower pancreatic b-cell mass and higher pancreatic a-cell mass and glucagon levels [18]. Treatment of pregnant mice with DEX was also described to epigenetically impair brown adipose tissue (BAT) thermogenesis and energy expenditure in the offspring, leading to increased adiposity and insulin resistance [19].

Aside from the liver, several studies have reported the expression of G6Pase and PEPCK in the small intestine of humans and rats, deeming this organ relevant in endogenous glucose production (EGP) [20–22]. Significant contribution of the small intestine to EGP is particularly relevant after a fasting period of at least 40 h, the period of time necessary for an increase in both G6Pase and PEPCK in the jejunum [23–25].

The present study has been undertaken to evaluate if prenatal exposure to DEX and excessive consumption of fructose during adult life could interact to modulate small intestine gluconeogenesis. To achieve this aim, we have evaluated key enzymatic and biochemical end-points indicative of intestinal gluconeogenesis as well as morphological aspects of the jejunum in 40 h fasted rats born to DEX-treated mothers and/or exposed to liquid fructose during adulthood.

#### **2. Materials and Methods**

#### *2.1. Experimental Design and Diet*

Eight-week-old nulliparous Wistar rats were acquired from the Animal Breeding Center at the Institute of Biomedical Sciences, University of Sao Paulo (Protocol # 5367250619). The animals were housed and mated with male rats as previously described [16].

After mating, pregnant rats were randomly assigned to receive 0.1 mg/kg/day dexamethasone (DEX) diluted in the drinking water from the 14th to the 19th day of pregnancy or remain untreated (CTL). On the 80th day of life, male offspring of DEX-treated and CTL dams were divided into two additional groups that were either kept with tap water or 10% fructose (w/v) solution ad libitum for the next 80 days. All offspring received standard chow ad libitum from the weaning to the 160th day of life.

The different groups were thereafter designed as follows: offspring born to CTL mothers that received only standard chow (SC) and tap water during adult life (CTL-SC); offspring born to DEX-treated mothers that received only SC and tap water during adult life (DEX-SC); offspring born to CTL mothers that received SC plus 10% fructose during adult life (CTL-fructose); and offspring born to DEX-treated mothers that received SC plus 10% fructose during adult life (DEX-fructose). On the 160th day of life, the animals were subjected to a 40 h fasting before euthanasia. During fasting, standard chow was removed and 10% fructose was replaced by tap water.

#### *2.2. Pyruvate Tolerance Test (PTT)*

Rats were fasted for 40 h and a 20% sodium pyruvate solution was injected intraperitoneal (i.p.) at a dosage of 2 g/kg of body mass. Glucose concentration was determined in blood extracted from the tail before (0 min) and 15, 30, 60, and 90 min after pyruvate injection (tail blood samples were chosen as representative of systemic blood). The area under the curve (AUC) of tail blood glucose levels vs. time was calculated using each individual baseline (basal glycemia) to estimate whole-body gluconeogenesis. We also collected portal blood samples at the end of the PTT (90 min after pyruvate challenge) to estimate the ability of the small intestine to convert pyruvate into glucose.

#### *2.3. Tissue Sampling and Preparation*

The rats were anesthetized with isoflurane. Proper level of anesthesia was assured by loss of pedal reflex. The abdominal cavity then was opened and portal blood was punctured. Euthanasia was performed by rupture of the diaphragm (with surgical scissors) followed by immediate cardiac puncture of the systemic blood. Systemic and portal blood samples were collected in EDTA-coated tubes and centrifuged at 2000 rpm for 20 min at 4 ◦C. Plasma samples were removed and used for biochemical analysis.

Intestinal segments were harvested from the proximal jejunum (5 cm beyond the ligament of Treitz), opened at the mesenteric border, pinned flat on a cork mat, and gently washed in ice-cold 0.1 M phosphate-buffered saline solution (PBS). The time elapsed between euthanasia and jejunum samples harvesting was approximately 2 min. The segments were transversally cut into two samples. In one sample, the mucosa was scraped and frozen under −80 ◦C for subsequent molecular and biochemical analyses (described below).

The second jejunum sample was immediately fixed with 10% buffered formaldehyde for 24 h, dehydrated in alcohol, diaphanized in xylol, and embedded in paraffin. Nonserial longitudinal sections (5–7 μm thick) were subjected to hematoxylin–eosin (HE) staining for morphometric analysis. In some rats, the entire small intestine was dissected and its length (cm) was measured and expressed as the relative length to tibia length [26].

#### *2.4. Analysis of Blood Parameters*

Plasma glucose, triglycerides, lactate, and cholesterol determinations were performed using commercially available kits (Labtest Diagnóstica SA, Lagoa Santa, MG, Brazil).

#### *2.5. Enzymatic Activity*

The activities of phosphoenolpyruvate carboxykinase (PEPCK; EC 4.1.1.32), glucose-6-phosphatase (G6Pase; EC 3.1.3.9), and hexokinase (HK; EC 2.7.1.1) were measured using spectrophotometric assays at 340 nm, following standard methods described elsewhere [27–29]. Protein concentration of each sample was determined by the Bradford method, and enzymatic activities were normalized by protein content.

#### *2.6. Molecular Analyses*

Scraped mucosal cells were processed for both qPCR and Western blotting (WB), as previously described [16]. The nitrocellulose membranes for WB were stained with Ponceau S before incubation with the primary antibodies. The stained membranes were allowed to dry at room temperature, scanned, and subjected to optical density (OD) quantification. All the lanes (from the top to the bottom) of labeled proteins were scanned to better represent the total amount of protein actually loaded in the gel. Subsequently, these values were applied to normalize the OD data of the target proteins detected in the respective membranes. This method was validated as an appropriate loading control [30]. The primary

antibodies used were as follows: anti-GLUT2 (cat. # sc-9117) from Santa Cruz Biotechnology (Santa Cruz, CA, USA) and anti-GLUT5 (cat. # IM-0292) from Rhea Biotech (Campinas, SP, Brazil).

Total RNA was extracted using QIAzol reagent and used for reverse transcription with random primers for the analysis of mRNA expression. The primer sequences and accession numbers were as follows: *G6pc* (NM\_013098) 5 -ACCTTCTTCCTGTTTGGTTTCGC-3 and 5 -CGGTACATGCTGGAGTTGAGGG-3 ; *Pck1* (NM\_198780) 5 -TGGTCTGGACTTCTCTGCCAAG-3 and 5 -AATGATGACCGTCTTGCTTTCG-3 ; *Ggt1* (NM\_053840) 5 -ACCCGACTTCATCGCTGTG-3 and 5 -GCATGTTCTCCAGAGTCCCAC-3 ; *Rpl37a* (X14069) 5 -CAAGAAGGTCGGGATCGTCG-3 ; and 5 -ACCAGGCAAGTCTCAGGAGGTG-3 . Values of mRNA expression were normalized using the internal control gene *Rpl37a*. Fold changes were calculated by the 2−ΔDDCT method.

#### *2.7. Morphometric Analysis of the Jejunum Wall*

The morphometric analyses were performed blindly using AxioVision Release 4.8-SP2 software (Carl Zeiss Microscopy, Jena, Germany) and consisted of the evaluation of the villus height and crypt depth. The height of each villus was measured from the top of the villus to the crypt transition, and the crypt depth was defined as the invagination between two villi. These analyses were performed in five fields at 100× magnification from different jejunum regions, including 2–3 villus/crypt per field, totaling 10–15 villus/crypt per animal [31].

#### *2.8. Immunohistochemical Evaluation of Cell Proliferation*

Antigen retrieval was performed in citric acid (10 mM, pH 6.0) at 95 ◦C for 40 min, followed by cooling for 30 min. After antigen retrieval, sections were incubated with 3% hydrogen peroxide diluted in methanol for 30 min to quench endogenous peroxidase, then rinsed with deionized water followed by PBS, pH 7.4. Sections then were incubated with 6% defatted milk for 30 min, at 37 ◦C, to block nonspecific staining. The anti-PCNA monoclonal antibody, from Dako-Agilent (Santa Clara, CA, USA; Cat. No. M0879), was diluted 1:1000 in PBS plus 1% bovine serum albumin (BSA) and incubated on sections overnight at 4 ◦C. All sections were washed three times in PBS for 5 min each time, then incubated with secondary antibody conjugated with horseradish peroxidase labeled polymer (EnVision + Dual link System-HRP) for 30 min at room temperature. 3,3 -diaminobenzidine (DAB) was used to visualize the antigen/antibody complex and the specimens were then lightly counterstained with hematoxylin. Negative control samples were performed by substituting the primary antibody with antibody diluent. We analyzed 10 crypts from each section under a 400× magnification [32]. Cell proliferation rate was expressed as percentage of PCNA-positive cells.

#### *2.9. Statistical Analyses*

Comparisons were performed using two-way ANOVA, followed by a Tukey's multiple comparison test. The two factors considered for the two-way ANOVA were in utero exposure to DEX (either exposed or not) and treatment of 10% fructose during adulthood (either treated or not). When making comparisons between two groups, the unpaired Student's *t*-test was used. Statistical analyses were conducted using GraphPad Prism software version 8.4.3 (GraphPad Software, Inc., San Diego, CA, USA). All results are presented as the means ± standard error of the mean (SEM). Results with *p* values lower than 0.05 were considered significant.

#### **3. Results**

#### *3.1. Consumption of Fructose by Rats Born to DEX-Treated Mothers Modifies Body Composition but Does Not Modulate Small Intestine Length*

As reported by us in previous studies [16,33], rats born to DEX-treated mothers displayed reduced birth weight (18% lower than CTL; *p* = 0.033) (Figure 1A). The body weights of the 40 h fasted offspring with 160 days of age were influenced by both in utero exposure to DEX and treatment with 10% fructose (*p* < 0.0001 and *p* = 0.0002, respectively). The post hoc analysis revealed that DEX-SC were lighter (9%; *p* < 0.01) while CTL-fructose were heavier (10%; *p* < 0.001) when compared to age-matched CTL-SC. In addition, DEX-fructose rats were lighter than the SC-fructose group (13%; *p* < 0.0001) (Figure 1B).

**Figure 1.** Morphometrical parameters of rats exposed to dexamethasone (DEX) in utero. Body weight was measured at birth (**A**) and at the end of treatment (**B**) in nonfasted rats. Fasted rats were euthanized at the end of the 80th day of fructose consumption, and the small intestine length relative to tibia length (**C**), and fat pads (mesenteric, (**D**); epidydimal, (**E**); retroperitoneal, (**F**)) and liver (**G**) masses relative to body weight were also measured. Results are presented as mean ± standard error of the mean (S.E.M.). \* *p* < 0.05, \*\* *p* < 0.01 (*n* = 10–20). Offspring born to control (CTL) mothers that received only standard chow (SC) and tap water during adult life (CTL-SC); offspring born to DEX-treated mothers that received only SC and tap water during adult life (DEX-SC); offspring born to CTL mothers that received SC plus 10% fructose during adult life (CTL-fructose); and offspring born to DEX-treated mothers that received SC plus 10% fructose during adult life (DEX-fructose).

Small intestine length, relative to tibia length, was influenced by in utero exposure to DEX (*p* = 0.0035). The post hoc analysis revealed that both DEX-SC and DEX-fructose had shorter small intestine when compared to CTL-SC (respectively 8% and 7% shorter; *p* < 0.05) (Figure 1C).

Mesenteric adiposity was influenced by the factor in utero exposure to DEX in the 40 h fasted offspring (*p* = 0.0264). However, the post hoc analysis revealed a specific increase of mesenteric adiposity in 40 h fasted DEX-SC (42% higher than CTL-SC; *p* < 0.05) (Figure 1D). The treatments had no effect on epididymal adiposity (Figure 1E). Retroperitoneal adiposity of the 40 h fasted adult offspring presented changes that were similar to those seen for mesenteric adiposity. The post hoc analysis revealed a specific increase of retroperitoneal adiposity of the 40 h fasted DEX-SC (50% higher than CTL-SC; *p* < 0.01) (Figure 1F). The relative weight of the liver was not affected by the treatments (Figure 1G).

#### *3.2. Biochemical Changes Detected in Rats Born to DEX-Treated Mothers That Consume Fructose during Adulthood Indicates Increased Intestinal Gluconeogenesis*

Both in utero exposure to DEX and treatment with 10% fructose during adulthood affected systemic glucose levels after a 40 h fasting (*p* = 0.046 and *p* = 0.039, respectively). However, the post hoc analysis revealed that systemic glucose levels were increased exclusively in 40 h fasted DEX-fructose (28% higher than CTL-SC; *p* < 0.05) (Figure 2A). Similarly, in utero exposure to DEX and treatment with 10% fructose during adulthood influenced systemic triglyceride levels after a 40 h fast (*p* = 0.0025 and *p* < 0.0001, respectively). In regard to this, the post hoc analysis indicated that systemic triglyceride levels were increased in 40 h fasted DEX-fructose when compared to CTL-SC, DEX-SC, and CTL-fructose (respectively 108%, 62%, and 36%; *p* < 0.0001, *p* < 0.001, and *p* < 0.05). Systemic triglycerides were also increased in 40 h fasted CTL-fructose (52% higher than CTL-SC; *p* < 0.05) (Figure 2B). Total systemic cholesterol levels after 40 h fasting were not altered in any of the four groups studied (Figure 2C).

Both in utero exposure to DEX and treatment with 10% fructose during adulthood also modified portal glucose levels after a 40 h fasting (*p* < 0.0001 and *p* = 0.045, respectively). Our post hoc analysis revealed that portal glucose levels were increased in 40 h fasted DEX-fructose (144% higher than CTL-SC and 83% higher than CTL-fructose; *p* < 0.0001 and *p* < 0.01). The portal glucose levels were also increased in 40 h fasted DEX-SC (102% higher than CTL-SC; *p* < 0.05) (Figure 2D). Portal lactate levels detected after a 40 h fasting were only influenced by in utero exposure to DEX (*p* < 0.05). The post hoc analysis revealed a specific reduction of portal lactate levels in 40 h fasted DEX-fructose rats (26% lower than CTL-SC; *p* < 0.05) (Figure 2E).

**Figure 2.** *Cont*.

**Figure 2.** Effects of fructose on biochemical parameters of systemic and portal blood in rats exposed to dexamethasone (DEX) in utero. Systemic blood samples were collected to measure glucose (**A**), triacylglycerol (**B**), and total cholesterol (**C**). Portal hepatic vein samples were collected to measure glucose (**D**) and lactate (**E**). Results are presented as mean ± standard error of the mean (S.E.M.). \* *p* < 0.05, \*\* *p* < 0.01, \*\*\* *p* < 0.001, \*\*\*\* *p* < 0.0001 (*n* = 8–12). Offspring born to control (CTL) mothers that received only standard chow (SC) and tap water during adult life (CTL-SC); offspring born to DEX-treated mothers that received only SC and tap water during adult life (DEX-SC); offspring born to CTL mothers that received SC plus 10% fructose during adult life (CTL-fructose); and offspring born to DEX-treated mothers that received SC plus 10% fructose during adult life (DEX-fructose).

#### *3.3. Rats Born to DEX-Treated Mothers That Consume Fructose during Adulthood Display Increased Portal Glucose Levels after Challenge with Exogenous Pyruvate*

We next performed the pyruvate tolerance test with the attempt to clarify if the higher portal glucose levels seen in 40 h fasted DEX-fructose rats were due to increased gluconeogenesis. The glucose levels in tail blood samples were assessed at different time points after pyruvate injection (Figure 3A). Treatment with 10% fructose during adulthood influenced the area under the curve (AUC) values in 40 h fasted rats (*p* < 0.0001). Our post hoc analysis indicated that whole-body gluconeogenesis is increased in fructose-treated rats irrespective of maternal treatment with DEX. This can be concluded because the AUC values of both CTL-fructose and DEX-fructose rats were similar to each other and higher than those of CTL-SC (respectively 174% and 227% higher; *p* = 0.0227 and *p* = 0.0016) (Figure 3B).

**Figure 3.** *Cont*.

**Figure 3.** Whole-body and intestinal use of pyruvate as a gluconeogenesis substrate by rats exposed to dexamethasone (DEX) in utero and treated with fructose during adult life. The 40 h fasted rats received an i.p. injection containing sodium pyruvate. The blood from the tail was collected before and 15, 30, 60, and 90 min after intraperitoneal (i.p.) injection for glucose measurements (**A**) and the area under the curve (AUC) was calculated above each individual baseline (**B**). Glucose levels were also measured in portal blood at the end of the pyruvate tolerance test (PTT) (**C**). Results are presented as mean ± standard error of the mean (S.E.M.) \* *p* < 0.05, \*\* *p* < 0.01, \*\*\*\* *p* < 0.0001 (*n* = 8). Offspring born to control (CTL) mothers that received only standard chow (SC) and tap water during adult life (CTL-SC); offspring born to DEX-treated mothers that received only SC and tap water during adult life (DEX-SC); offspring born to CTL mothers that received SC plus 10% fructose during adult life (CTL-fructose); and offspring born to DEX-treated mothers that received SC plus 10% fructose during adult life (DEX-fructose).

Glucose levels in portal blood 90 min after challenge with pyruvate were influenced by both in utero exposure to DEX and treatment with 10% fructose during adulthood (*p* = 0.0043 and *p* = 0.0001, respectively). In contrast to the changes in whole-body gluconeogenesis, our post hoc analysis revealed increased portal glucose levels after challenge with pyruvate exclusively in DEX-fructose rats (65% higher than CTL-SC, 26% higher than CTL-fructose, and 39% higher than DEX-SC; *p* < 0.001, *p* = 0.0301, and *p* = 0.0055) (Figure 3C).

#### *3.4. Rats Born to DEX-Treated Mothers That Consume Fructose during Adulthood Display Increased PEPCK Expression and Activity in the Jejunum*

The expression of *G6pc* (the gene that encodes G6Pase) in the jejunum was not affected in any of the four groups after a 40 h fasting (Figure 4A). The activity of G6Pase in the jejunum of the 40 h fasted rats was influenced by in utero exposure to DEX (*p* = 0.035) but no specific differences were found in the post hoc analysis (Figure 4B).

The expression of *Pck1* (the gene that encodes PEPCK) in the jejunum of the 40 h fasted rats was modulated by in utero exposure to DEX (*p* = 0.005). In this case, our post hoc analysis indicated a marked increase of *Pck1* expression in the jejunum of the 40 h DEX-fructose (88% higher than CTL-SC; *p* < 0.05) (Figure 4C). As with changes in expression, PEPCK activity in the jejunum of the 40 h fasted rats was regulated by in utero exposure to DEX (*p* = 0.043). Our post hoc analysis indicated a specific increase of PEPCK activity in the jejunum of the 40 h fasted DEX-fructose group (14% higher than CTL-SC; *p* < 0.05) (Figure 4D). The expression of fructose 1,6-bisphosphatase (*Fbp1*) was also evaluated but no differences were found among the groups (data not shown).

**Figure 4.** Expression and activity of enzymes involved in intestinal gluconeogenesis. Scraped epithelium of jejunum fragments was isolated and processed for qPCR detection of G6pc (**A**) and Pck1 (**C**) gene expression, as well as for maximum activities of the corresponding enzymes G6Pase (**B**) and PEPCK (**D**). Results are presented as mean ± standard error of the mean (S.E.M.). \* *p* < 0.05 (*n* = 6–12). Offspring born to control (CTL) mothers that received only standard chow (SC) and tap water during adult life (CTL-SC); offspring born to dexamethasone (DEX)-treated mothers that received only SC and tap water during adult life (DEX-SC); offspring born to CTL mothers that received SC plus 10% fructose during adult life (CTL-fructose); and offspring born to DEX-treated mothers that received SC plus 10% fructose during adult life (DEX-fructose).

#### *3.5. Rats Born to DEX-Treated Mothers That Consume Fructose during Adulthood Display Reduced HK Activity and Increased GLUT5 and GLUT2 Expression in the Jejunum*

Both in utero exposure to DEX and treatment with 10% fructose during adulthood influenced hexokinase (HK) activity in the jejunum of the 40 h fasted rats (*p* = 0.0362 and *p* = 0.0004, respectively). The post hoc analysis revealed that reductions in HK activity in the jejunum of the 40 h fasted rats were specific for the DEX-fructose group (46% lower than CTL-SC, 51% lower than DEX-SC, and 44% lower than CTL-fructose; *p* < 0.001, *p* < 0.0001, and *p* < 0.01) (Figure 5A).

The expression of glutathione S-transferase 1 (*Gtt1*) in the jejunum of the 40 h fasted rats was influenced by in utero exposure to DEX (*p* < 0.0001). The post hoc analysis revealed that 40 h fasted DEX-SC rats had increased expression of *Gtt1* in the jejunum (90% higher than CTL-SC; *p* < 0.05). Increased expression of *Gtt1* in the jejunum was also found in 40 h fasted DEX-fructose (170% higher than CTL-fructose and 156% higher than CTL-SC; *p* < 0.001) (Figure 5B).

The content of facilitated glucose transporter member 2 (GLUT2) in the jejunum of the 40 h fasted rats was altered by treatment with 10% fructose during adulthood (*p* = 0.0017). The post hoc analysis revealed that the increase of GLUT2 content in the jejunum of the 40 h fasted rats was specific for DEX-fructose (60% higher than CTL-SC and 73% higher than DEXA-SC; *p* < 0.01) (Figure 5C).

The content of GLUT5 in the jejunum of the 40 h fasted rats was also influenced by in utero exposure to DEX (*p* = 0.0299). The post hoc analysis revealed a specific increase of GLUT5 content in the jejunum of the 40 h fasted DEX-fructose (75% higher than CTL-fructose; *p* < 0.05) (Figure 5D).

**Figure 5.** In utero dexamethasone (DEX) exposure alters glucose metabolism and phenotypic features of the jejunum epithelia. Scraped epithelium of jejunum fragments was isolated and processed for measurement of maximum hexokinase activity (**A**), expression of glutathione S-transferase 1 (Ggt1) by quantitative polymerase chain reaction (qPCR) (**B**), and Western blot of facilitated glucose transporter member 2 (GLUT2) (**C**) and facilitated glucose transporter member 5 (GLUT5) (**D**). Results are presented as mean ± standard error of the mean (S.E.M.). \* *p* < 0.05, \*\*\* *p* < 0.001, \*\*\*\* *p* < 0.0001 (*n* = 6–12). Offspring born to control (CTL) mothers that received only standard chow (SC) and tap water during adult life (CTL-SC); offspring born to DEX-treated mothers that received only SC and tap water during adult life (DEX-SC); offspring born to CTL mothers that received SC plus 10% fructose during adult life (CTL-fructose); and offspring born to DEX-treated mothers that received SC plus 10% fructose during adult life (DEX-fructose).

#### *3.6. Rats Born to DEX-Treated Mothers That Consume Fructose during Adulthood Display Morphological Changes in the Jejunum Epithelium*

We have also assessed the mean crypt depth and the mean villous height, two aspects of the jejunum epithelium that are vital for its absorptive capacity. Images of the HE-stained jejunum sections are shown from each of the four different groups of 40 h fasted adult offspring (Figure 6A–D).

**Figure 6.** Morphometric analysis of the jejunum wall. The figure shows representative sections of villus and crypts of the four experimental groups (**A**–**D**). The height of each villus was measured from the top of the villus to the crypt transition (**E**), and the crypt depth was defined as the invagination between two villi (**F**). Results are presented as mean ± standard error of the mean (S.E.M.). \* *p* < 0.05, \*\* *p* < 0.01 (*n* = 5). Offspring born to control (CTL) mothers that received only standard chow (SC) and tap water during adult life (CTL-SC); offspring born to dexamethasone (DEX)-treated mothers that received only SC and tap water during adult life (DEX-SC); offspring born to CTL mothers that received SC plus 10% fructose during adult life (CTL-fructose); and offspring born to DEX-treated mothers that received SC plus 10% fructose during adult life (DEX-fructose).

The mean crypt depth in the jejunum epithelium of 40 h fasted rats was modulated by in utero exposure to DEX (*p* = 0.0119). On the other hand, the post hoc analysis revealed that crypt depth was only increased in the jejunum epithelium of 40 h fasted DEX-fructose rats (25% higher than DEX-SC and 43% higher than CTL-fructose; *p* < 0.05 and *p* < 0.01) (Figure 6E).

Villous height was influenced by treatment with 10% fructose during adulthood (*p* = 0.0017). Similar to crypt depth, the post hoc analysis revealed that villous height was only increased in the jejunum epithelium of 40 h fasted DEX-fructose (27% higher than CTL-SC and 39% higher than DEX-SC; *p* < 0.05 and *p* < 0.01) (Figure 6F).

The proliferative potential in the jejunum epithelium was evaluated by assessing the relative number of PCNA-positive cells. Representative images of the jejunum sections stained with anti-PCNA antibody are shown from each of the four different groups of 40 h fasted adult offspring (Figure 7A–D). The hematoxylin-counterstained section that served as a negative control (by omission of the primary antibody) is shown in Figure 7E.

**Figure 7.** Jejunum was removed for immunohistochemical detection of proliferating cell nuclear antigen (PCNA). The figure shows representative sections of PCNA staining (**A**–**D**) and negative control sample (**E**). Sections were used to calculate the percentage of PCNA-positive cells in crypt cells (**F**). Results are presented as mean ± standard error of the mean (S.E.M.). \*\*\*\* *p* < 0.0001 (*n* = 5). Offspring born to control (CTL) mothers that received only standard chow (SC) and tap water during adult life (CTL-SC); offspring born to dexamethasone (DEX)-treated mothers that received only SC and tap water during adult life (DEX-SC); offspring born to CTL mothers that received SC plus 10% fructose during adult life (CTL-fructose); and offspring born to DEX-treated mothers that received SC plus 10% fructose during adult life (DEX-fructose).

Both factors, in utero exposure to DEX and treatment with 10% fructose during adulthood, influenced cell proliferation rate in the jejunum epithelium of the 40 h fasted rats (*p* = 0.0158 and *p* < 0.0001, respectively). The post hoc analysis revealed a particular increase in the cell proliferation rate of the jejunum epithelium of 40 h fasted DEX-fructose (48% higher than CTL-SC, 78% higher than DEX-SC, and 43% higher than CTL-fructose; *p* < 0.0001) (Figure 7F).

#### **4. Discussion**

In utero exposure to DEX is well known for programming metabolic changes in the adult offspring of rats. The metabolic imprinting caused by excessive exposure to DEX during fetal life is hallmarked by glucose intolerance, increased whole-body gluconeogenesis, and upregulation of PEPCK expression in the liver [17,33,34]. Recently, we have also described that in utero exposure to DEX exacerbates hepatic steatosis caused by fructose consumption during adult life [16]. The present study further contributes to this topic by revealing that rats born to DEX-treated mothers present exacerbated intestinal gluconeogenesis after consuming excessive fructose during adult life.

Changes in key endpoints support the above claim; increased PEPCK expression and activity in the jejunum and increased portal glucose levels were detected after a 40 h fasting in rats born to DEX-treated mothers that consumed fructose during adult life. An additional finding that supports the proposition that in utero exposure to DEX increases gluconeogenesis capacity is the higher portal glucose levels detected in DEX-fructose rats 90 min after the challenge with pyruvate, a known gluconeogenesis substrate. The 40 h fasting that preceded our sample collection and the PTT was performed because it has been previously described that intestinal gluconeogenesis does not significantly occur during shorter periods of food deprivation [25].

Another finding that supports the notion that intestinal gluconeogenesis is increased in rats born to DEX-treated mothers that consume fructose during adulthood is the increase in GLUT2 content in the jejunum of DEX-fructose rats, with parallel reduction in HK activity. GLUT2 is classically recognized for mediating the basolateral transport of glucose to the capillary vessels that feed the portal vein [35]. Our interpretation is that enterocytes of DEX-fructose rats have increased ability to synthesize glucose de novo (due to increased PEPCK) and release the newly synthesized glucose to the portal bloodstream (due to increased GLUT2 expression). Lower HK activity in the enterocytes of the DEX-fructose rats may contribute to the increased intestinal glucose release by reducing their rate of conversion of newly synthesized glucose back to glucose-6-phosphate.

In parallel with the above-mentioned biochemical changes that evidence increased intestinal gluconeogenesis, DEX-fructose rats exhibited lower portal lactate levels. This is particularly relevant because 10% fructose solution was replaced by water during the 40 h fasting. Thus, the current experiments do not support the notion that in utero exposure to DEX increases intestinal conversion of fructose into glucose but instead indicate that the jejunum of 40 h fasted adult DEX-fructose rats may increase the use of lactate as substrate for gluconeogenesis.

Interestingly, previous studies have reported that the small intestine of the fasted adult rat preferentially uses glutamine and glycerol, instead of lactate, as substrates for gluconeogenesis [23]. On the other hand, the small intestine of suckling rats is able to convert lactate into glucose [36]. Considering this, our data suggest that the small intestine of the offspring born to DEX-treated mothers that chronically consume fructose during adult life preserves a metabolic feature of the newborn small intestine.

Although it is challenging to presume the functional relevance of the increased GLUT5 in the jejunum of DEX-fructose rats after a 40 h fasting, this particular result reinforces the proposition that the small intestine of the adult offspring born to DEX-treated mothers preserves phenotypic features of the newborn after chronic exposure to fructose. Supporting this suggestion, it was previously demonstrated that DEX exacerbates GLUT5 expression induced by fructose in samples of small intestine of neonatal rats [37]. It is important to note that in utero exposure to DEX alone is not sufficient to stimulate GLUT5 content in the jejunum of the offspring. Such findings have also been previously reported in other studies [38] and support the notion that the two factors together (both in utero exposure to DEX and fructose consumption) seem to be necessary for enhancing jejunal GLUT5 content.

With regard to the morphological impact of DEX on the small intestine early in life, it has been previously reported that the lactating pups treated with DEX during lactation exhibit transitory changes in the small intestine epithelial architecture. Pups treated with DEX between the 11th and the 21st days of life display increased villous height and crypt depth in the jejunum soon after weaning. These changes are no longer detected by the age of 50 days [39]. Hence, we conclude that aside from the metabolic/biochemical features, morphological changes transiently described in the lactating pups exposed to DEX during early life are sustained in the adult offspring born to DEX-treated mothers only after consumption of excessive fructose.

We have also found that the frequency of PCNA-positive cells in the jejunum epithelial surface, a parameter that spontaneously reduces in the jejunum of the adult rat as the age advances beyond 90 days of life [40], is exacerbated in the 160-day-old DEX-fructose rats. Instead, *Ggt1* expression, an enzyme that plays a crucial role in de novo synthesis of intracellular GSH and ROS removal [41], is increased in the jejunum of adult rats born to DEX-treated mothers, irrespective of fructose consumption. Notably, GGT1 inhibition was associated with increased apoptosis in smooth muscle cells [42]. Thus, intestinal epithelial cells of the adult DEX-fructose offspring are unique in such a way that they combine long-term pro-proliferative and antiapoptotic adaptations.

Another interesting metabolic feature exhibited in the 40 h fasted DEX-fructose rats is the increased circulating triglyceride levels. Although we are not able to discern the hepatic or the intestinal origin of the lipoproteins that contribute to this phenomenon, it is important to take into account that intestinal production of triglyceride-enriched lipoprotein accounts for up to 40% of the triglycerides in fasting rats [43]. Moreover, chronic consumption of a fructose-enriched diet was described to increase the intestinal production of chylomicrons during fasting periods [44].

In summary, the present study supports the proposition that consumption of fructose by adult rats exposed to DEX during fetal life leads to an exacerbation in intestinal gluconeogenesis and retention of morphological features in the jejunum that are commonly found during neonatal life. These data provide a new mechanism to explain the increased prevalence of metabolic disturbances in humans that are born with low birth weight.

**Author Contributions:** Conceptualization, S.B. and G.A.P.; methodology, G.A.P., F.S.S., G.M.M., A.G.A., T.B.P., C.V.C., and F.T.S.; validation, G.A.P., F.S.S., G.M.M., and A.G.A; formal analysis, G.A.P., G.M.M., and A.G.A.; investigation, G.A.P. and F.S.S.; resources, S.B. and G.F.A.; data curation, S.B., G.M.M., and A.G.A.; writing—original draft preparation, G.A.P., A.G.A., and G.F.A.; writing—review and editing, S.B. and G.F.A.; supervision, S.B.; funding acquisition, S.B. and G.F.A. All authors have read and agreed to the published version of the manuscript.

**Funding:** This study was supported by the Research Foundation of the State of Sao Paulo (FAPESP Grants 2013/07607-8 and 2019/03196-0) and the National Council of Research (CNPq).

**Acknowledgments:** We acknowledge the technical support of Mariana M. Onari, Tiffany. B. Watanabe, Lais O.C. Lima, and Amanda M.S. Silva. We also thank Charles Serpellone Nash for carefully reviewing the manuscript.

**Conflicts of Interest:** The authors declare no conflicts of interest, financial or otherwise, associated with this article. The authors are responsible for the writing and content of the article.

#### **References**


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